The Nucleus

The Nucleus A Subject Collection from C o ld Spring H arbor Perspectives in Biology Articles online at www.cshperspecti...

Author: David L. Spector | Tom Misteli (editors)

92 downloads 1596 Views 25MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

The Nucleus A Subject Collection from C o ld Spring H arbor Perspectives in Biology Articles online at www.cshperspectives.org A ll rights reserved < 2 0 1 1 by C old Spring Harbor Laboratory Press, Cold Spring Harbor, N ew York Printed in the United States o f America Publisher

|ohn Inglis

Acquisition Editor

Richard Sever

Project M anager

M aryliz Dickerson

Permissions C oordinator

Carol Brown

Production Editor

Kaaren Kockenmeister

Cover Designer

Denise Weiss

Front cover artwork: Diagram illustrating the organization of the eukaryotic nucleus. Depicted are chromatin (orange), enclosed by and anchored to the nuclear envelope (gray), and transcrib ing RNPs (purple), w hich after processing are exported through nuclear pore complexes (green and blue). (Illustration courtesy o f M ichael P. Rout. The Rockefeller University.)

Library o f Congress Cataloging-in-Publication Data

The nucleus/edited by Tom Mistelli, David L. Spector, p. cm. Includes bibliographical references and index. ISBN 978-0-87969-894-2 (hardcover: alk. paper) 1. C ell nuclei. I. Mistelli, Tom. II. Spector, David L. III. Title. Q H 5 9 5 .N 8 5 2011 571,6'6--dc22 2010039936 10

9

8

7

6

5

4

3

2

1

A ll W orld W id e W eb addresses are accurate to the best of our knowledge at the time of printing. Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Cold Spring Harbor Laboratory Press, provided that the appropriate fee is paid directly to the Copyright Clearance Center (CCC). Write or call CC C at 222 Rosewood Drive, Danvers, M A 01923 (978-750-8400) for information about fees and regulations. Prior to photocopying items for educational classroom use, contact C C C at the above address. Additional information on C C C can be obtained at C C C O nline at http://www.copyright.com/. A ll Cold Spring Harbor Laboratory Press publications may be ordered directly from Cold Spring Harbor Laboratory Press, 500 Sunnyside Blvd., Woodbury', New York 11797-2924. Phone: 1-800843-4388 in Continental U.S. and Canada. A ll other locations: (516) 422-4100. FAX: (516) 4224097. E-mail: cshpress
Contents

Preface, ix

OVERVIEW The Nucleus Introduced, 1 Thoru Pederson

THE NU CLEA R PERIPHERY The Nuclear Envelope, 17 M a r lin W. Helzer

Nuclear Lamins, 33 Thomas Dechat, Stephen A. Adam , Pekka Taimen, Takeshi Shimi, and Robert D. Goldman

Lamin-binding Proteins, 55 Katherine I.. Wilson a n d Roland Foisner

The Nuclear Pore Complex and Nuclear Transport, 73 Susan R. Wente a n d M ich a e l P. Rout

C H R O M O S O M ES & CHROM ATIN Chromosome Territories, 93 Thomas Cremer and M a rio n Cremer

Gene Pbsitioning, 115 Carm elo Ferrai, Ines Jesus de Castro, Liron Lavitas, M ita Chotalia, a n d Ana Pombo

Chromatin Higher-order Structure and Dynamics, 133 Christopher L Woodcock and Rajarshi P. Ghosh

Nuclear Organization and Dosage Compensation, 159 Jennifer C. Chow and F.dith Heard

The Budding Yeast Nucleus, 175 Angela Taddei, Heiko Schober, and Susan M . Gasser

Nuclear Functions of Actin, 195 Neus Visa and Piergiorgio Percipalle

NU CLEA R BODIES The Nucleolus, 209 Thoru Pederson

Nuclear Speckles, 225 D av id I.. Speclor and Angus I. Lam ond

The Cajal Body and Histone Locus Body, 237 Zehra Nizam i, Svetlana Deryiisheva, and Joseph G. G a ll

PM L Nuclear Bodies, 249 Valerie Lallemand-Breitenbach and Hugues de The

The Perinucleolar Compartment, 267 C allie Pollock and Sui H uang

Rjraspeckles, 277 Archa H . Fox and Angus I. Lam ond

Nuclear Stress Bodies, 291 Giuseppe Biam onti a n d Claire Vourc’h

Orphan Nuclear Bodies, 303 M a r ia Carmo-Fonseca, M a ria T. Berciano, a n d M iguel Lafarga

Biogenesis of Nuclear Bodies, 315 M iroslav D u n d r and Tom M is le li

F U N C T IO N A L ORGANIZATION Organization of Transcription, 331 Lyubomira Chakalova a n d Peter Fraser

Organization of D N A Replication, 347 Vadim O. Chagin, Jeffrey II. Stear, a n d M . Cristina Cardoso

D N A Damage Response, 361 Giuseppina G ig lia -M a ri, Angelika Zotter, and Wint Vermeulen

RN A Processing and Export, 381 Sam i Hocine, Robert H . Singer, a n d D avid G riinw ald

N UCLEAR ARCHITECTURE IN DIFFERENTIATION A N D DISEASE Diseases of the Nuclear Envelope, 401 H ow ard J. Worman, Cecilia Ostlund, a n d Yuexia Wang

Higher-order Genome Organization in Human Disease, 419 Tom M isteli

Nuclear Ataxias, 437 I Iarry T. O rr

Developments in RN A Splicing and Disease, 449 M ich ael G. Poulos, Ranjan Batra, Konstantinos Charizanis, and M a urice S. Swanson

APPENDIX Tables of Data on the Nucleus, 463 (Adapted by kind permission of Federation o f American Societies for Experimental Biology [FASEB] from B iological Handbooks. I. Cell Biology (Altman P.L. and Katz DD; 1976], Bethesda, Maryland.)

Index, 507

The Nucleus Introduced Thoru Pederson Program in Cell and Developmental Dynamics, Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts 01605 Correspondence: [email protected]

Now is an opportune moment to address the confluence of cell biological form and function that is the nucleus. Its arrival is especially timely because the recognition that the nucleus is extremely dynamic has now been solidly established as a paradigm shift over the past two decades, and also because we now see on the horizon numerous ways in which organization itself, including gene location and possibly self-organizing bodies, underlies nuclear functions.

“We have entered the cell, the Mansion of our birth, and started the inventory of our acquired wealth.” —Albert Claude

hen I first read that morsel from Albert Claude’s 1974 Nobel Prize lecture it seemed Solomonic wisdom, as it indeed was. Though he was referring to cell biology en toto, the study of the nucleus was then at a tipping point and new advances were just at hand. Since then, the nucleus field has literally nucleated and we are now at a position to both admire the recent past and register excitement about the present and where the nucleus field may be headed.

W

THE NUCLEUS DISCOVERED

We cannot know who first saw the nucleus but we do know that the father of optical

microscopy, Antony van Leeuwenhoeck, did so with amphibian and avian erythrocytes in 1710 and that in 1781 Felice Fontana did so as well in eel skin cells. More definitive accounts followed by Franz Bauer, who in 1802 sketched orchid cells and pointed out the nucleus (Bauer 1830 –1838), as well as by Jan Purkyneˇ, who described it as the vesicula germanitiva in chicken oocytes (Purkyneˇ 1825), and Robert Brown who observed it in a variety of plant cells (Brown 1829 – 1832), earning additional fame for coining the term “nucleus” (for excellent accounts of these early descriptions of the nucleus see Gall 1996; Harris 1999). Of course, these early observations did not ascribe particular significance to this structure, the given name simply conveying its central location. Later, the nucleus was increasingly observed and became, with some prescience, a key tenet of the cell theory. The nucleus remained a rather lonely item in the eukaryotic cell’s parts list for many

Editors: David L. Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved. Advanced Online Article. Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a000521

T. Pederson

decades, until the discoveries of discrete cytoplasmic entities, e.g., mitochondria, the Golgi apparatus etc. (reviewed in Wilson 1925). THE NUCLEUS INHERITED

We do not know how and when the genome of an ancestral cell first became encased in a primitive nucleus. We have no evidence that cells living in the RNA world ever had a membrane (or any other structure) around the genome, i.e., that they ever became nucleate. Once a ribozyme RNA replicase arose, anything would have been possible including the emergence of ribozymes with lipid biosynthetic activities. Enthalpy-favored or free energy-driven events could then have led to stabilizing selection of RNA-lipid affinities and on from there. A cottage industry of experiments on the interactions of lipids with RNA has emerged in the chemical biology field in the past decade but the significance of these studies to prebiotic systems and the earliest cells remains speculative. As for the advent of the DNA world, and of eukaryotes, a major concept is that a prokaryote organism was invaded by another, nonnucleated cell, setting up an endosymbiotic relationship, with the entering organism’s outer membrane seeding what would become the nuclear envelope. The major proponent of this plausible idea has also suggested that this hypothetical invader also brought in a centriole, the forerunner of what we know as the centriole/ basal body in extant eukaryotes (Margulis et al. 2000). We can not play the videotape of life on Earth backwards and although we can reconstruct some things with a degree of empirical confidence, albeit amidst debate (reviewed by Misteli 2001a; Poole and Penny 2001; Rotte and Martin 2001), or speculation (e.g., Lake 2009), when it comes to how the nucleus arrived, we just do not know. THE NUCLEUS ENVELOPED

Notwithstanding the uncertainty of its evolutionary origin, the nucleus is bounded by a double membrane, the nuclear envelope, which in many cells is contiguous with the endoplasmic

reticulum. The frequently observed intimacy of the nuclear envelope with the endoplasmic reticulum has been often under-appreciated, particularly as it bears on the isolation of nuclei and issues of resulting purity. There is also growing interest in how nuclear membrane proteins may be integrators of nuclear and cytoplasmic organization and dynamics (e.g., King et al. 2008; Roux et al. 2009; Starr 2009). A seminal finding was that the nuclear envelope contains pores (reviewed by Gall 1964; 1967), which have now been defined in considerable compositional and structural detail (e.g., Alber et al. 2007; Fernandez-Martinez and Rout 2009). A somewhat less familiar but equally important area of investigation has revealed that the nuclear envelope harbors a signal transduction system of its own, featuring players in common with the plasma membrane, for example the lipid-linked inositol trisphosphate system (Martelli et al. 1991; for reviews see Divecha et al. 1993; Cocco et al. 2009; Barton et al. 2010). THE NUCLEUS DIVERSIFIED

However the nucleus arose, it went on to display a variety of organizations. These range from the highly condensed nuclei of mature erythrocytes in nonmammalian vertebrates to the bimorphic nuclei in almost all ciliates. In the latter organisms a micronucleus perpetuates the genome whereas a macronucleus contains DNA fragments that represent only a fraction of the organism’s genome complexity and which encode the RNAs and proteins needed for vegetative life. It was a particular feature of this genomic strategy, namely the macronuclear amplification of the ribosomal RNA genes (Gall 1974), that led to the discovery of the telomere DNA sequence (Blackburn and Gall 1978). It was also in these ciliates that selfsplicing RNA was discovered and in which the era of chromatin epigenetic marks was launched (reviewed in Pederson 2010). THE NUCLEUS VIEWED

Beyond early observations made by bright field microscopy, the staining method developed by

The Nucleus, Introduced

Robert Feulgen, which attaches a dye to aciddepurinated DNA, was a major tool in advancing the DNA ¼ gene theory, based on studies of the DNA content of haploid versus diploid cells by a graduate student, Hewson Swift, in the laboratory of Arthur Pollister at Columbia University, and concurrent ones by Hans Ris in the laboratory of Alfred Mirsky at the Rockefeller Institute (reviewed by Pederson 2005). Although biochemical measurements of the DNA contents of germ versus somatic cells had revealed a twofold difference, these findings mostly remained in the biochemical community and did not have as much impact as might have been expected. It was the Feulgen cytophotometry results that helped catalyze the idea that DNA is genes. This was, to borrow part of Winston Churchill’s famous phrase—“the end of the beginning” (Avery et al. 1944). Meanwhile, phase contrast microscopy had beautifully revealed the interphase nucleus and mitotic chromosomes, the latter not “nuclear” in the strictest sense. Electron microscopy led to the visualization of the double nuclear membrane and nuclear pores (Gall 1964; 1967), the tripartite structure of the nucleolus (Bernhard et al. 1952), the nuclear lamina (Fawcett 1966), and subsequently the observation of chromatin n-bodies (Olins and Olins 1974), later re-named nucleosomes (Oudet et al. 1975). An intriguing nuclear structure first observed by the Spanish neuroanatomist Santiago Ramon y Cajal (Cajal 1903, 1910) underwent a renaissance of interest when one of its protein components was identified by Eng Tan and colleagues (Andrade et al. 1991; Raska et al. 1991). Initially termed the coiled body, a campaign in 1999 by Joseph Gall resulted in a consensus to rename this structure the Cajal body. Gall has made some of the most seminal advances in our understanding of this nuclear body (for reviews see Gall 2000; Handwerger et al. 2006; Gall 2009; Nizami et al. 2010). Similarly, in the 1990s the nucleolus, interchromatin granule clusters (a.k.a. nuclear speckles) and other nuclear bodies also underwent advances in molecular definition (reviewed by Spector 1993; Pederson 2002a; Spector 2001; 2006).

THE NUCLEUS ISOLATED

Using pus-soaked bandages from a local hospital, Friedrich Miescher discovered DNA in the late 1860s. As described in a richly detailed historical account (Portugal and Cohen 1977), Miescher wrote a manuscript and submitted it to a top journal but had to sit by and wait while the editor (and his former mentor), Felix Hoppe-Seyler, checked the findings in his own lab, resulting in the publication of two concurrent papers (Miescher 1871; Hoppe-Seyler 1871). Over the next 90 years, efforts to isolate nuclei did not go very far. In the 1960s Alfred Dounce at the University of Rochester, Van Potter and Conrad Elvehjem at the University of Wisconsin, and Philip Siekevitz and George Palade at the Rockefeller Institute pioneered the isolation of nuclei from animal tissue. (The Rochester and Wisconsin investigators had the homogenizers they developed named for them.) Important advances in isolating nuclei from plant tissue were made at approximately the same time by James Bonner’s group at Caltech. Later, groups at Baylor College of Medicine (Harris Busch), Rockefeller University (Alfred Mirsky) and Albert Einstein College of Medicine (Sheldon Penman) published refined methods. As is the case for most cell fractionation methods, none of these proved to be perfect but they were major advances nonetheless. The fact that the endoplasmic reticulum and the nuclear envelope are contiguous in many cells is but one example of this challenge, and the tendency of the cytoplasmic intermediate filament system to often collapse upon the nucleus during cell fractionation is yet another. The pioneer methods of nuclear isolation employed sucrose concentrations in which the tissue homogenate was exposed to hydrodynamic forces and/or sucrose density differences that caused cytoplasmic elements and adherent endoplasmic reticulum to separate from the nuclei. The most popular of the early methods (Chauveau et al. 1956) was subsequently refined in important ways to further minimize cytoplasmic contamination (Maggio et al. 1963; Blobel and Potter 1966), with these two latter advances constituting the gold standard of

T. Pederson

nuclear isolation for many years, and still today for many tissues. As regards methods aimed at single-cell studies, an important method (Penman 1966) involved swelling cells in a hypotonic buffer so that the expanded cell volume would collide with the hydrostatic shear forces delivered by a 0.0015-inch clearance stainless steel device, patterned after ones introduced by Dounce. Detergent-based methods were also introduced (Traub et al. 1964). These typically employed Triton X-100 or Royal Dutch Shell’s “nonionic P-40” (a.k.a. NP-40). These breach the plasma membrane and, depending on cell type, also strip away the endoplasmic reticulum. In some cases, NP-40 collapses some of the plasma membrane onto the nucleus, so that nuclei prepared by this method may not always be as pure as some investigators have assumed. For some cells, the main advantage of the NP-40 method is less the nuclear fraction, but the relatively high purity of the ribosomes that are released (Borun et al. 1967). All methods of nuclear isolation must be monitored both for what remains attached from the cytoplasm (or is taken up from it), as well as what is lost from within. Various markers have been used to assess the depletion of cytoplasmic material in isolated nuclei, based on the presumption that the true intracellular locations of such markers are known to some degree of certainty. In one study, HeLa cell nuclei were isolated in a buffer consisting of a previous cytoplasmic fraction from 3H-leucine-labeled cells (Bhorjee and Pederson 1972), thus allowing the level of cytoplasmic protein contamination of the nuclei to be readily estimated. Issues of nuclear purity loomed large in early studies claiming that protein synthesis occurs in the isolated nuclei (and thus presumably within nuclei in vivo) but this work was challenged on several grounds (reviewed by Goldstein 1970; Pederson 1976). More recently this issue has resurfaced (e.g., Iborra et al. 2001, although this study also included intact cell experiments). Various controls for nuclear purity have been better but still not sufficient to quiet all doubts as to the source of apparent protein synthesis in isolated nuclei (for reviews

see Pederson 2001a; Dahlberg et al. 2003; Nathanson et al. 2003). Meanwhile, less attention has been devoted to analyzing not what is adsorbed to nuclei as opposed to what is lost from nuclei during various isolation methods. There have been other instructive guideposts over the years for judging the purity of nuclear fractions. An instructive vignette is the saga of the biosynthesis of the U-rich spliceosomal small nuclear RNAs. We now know that they are exported to the cytoplasm as 30 extended precursors, there to be trimmed, 50 -cap hypermethylated and assembled with various proteins before re-entry into the nucleus as functional snRNPs. Although many studies had pointed to this pathway at varying degrees of cogency, it was the use of two particular nuclear isolation methods, each with exceptionally strong credentials, that led to acceptance of this pathway of snRNP biogenesis (Gurney and Eliceiri 1980; Zieve et al. 1988). This was further solidified by the demonstration that exogenous U2 snRNA precursor molecules introduced into the cytoplasm of mammalian cells enter this pathway and become 30 trimmed, cap hypermethylated, and assembled into snRNPs, followed by nuclear uptake (Kleinschmidt and Pederson 1990). Another example of how the critical issue of nuclear purity was addressed arose in a study in which the total sequence complexity of nuclear versus cytoplasmic RNA was measured (Holland et al. 1980). Because the nuclear RNA was anticipated to have a higher sequence complexity than cytoplasmic RNA, contamination of the cytoplasmic fraction by leaked nuclear RNA loomed large. Accordingly, an experiment was undertaken in which the cells were labeled with an RNA tracer for a very short time (2 min), far too brief for any labeled transcripts to be exported to the cytoplasm in vivo, before cell fractionation. The label was followed throughout the isolation of the cytoplasmic fraction and polyribosome-associated mRNA, which revealed that no more than 0.03% of the nuclear RNA synthesized in the previous 2 min was present in the final mRNA preparation (Holland et al. 1980). Given that these

The Nucleus, Introduced

kinds of controls are conceptually obvious and experimentally quite facile, it is surprising how rarely they have been employed in studies in which knowing the purity of a nuclear or cytoplasmic fraction is essential to weighing the findings. Beyond the isolation of nuclei in bulk, there is the gold standard of manual isolation of nuclei from material that affords this opportunity (e.g., Duryee 1954; later refined and reviewed by Lund and Paine 1990; Paine et al. 1992). In a recent application of this approach, frog oocyte nuclei were manually isolated under oil and the relative densities of the nucleolus versus surrounding nucleoplasmic bodies were determined by differential interference light microscopy, with results that defied most expectations (Handwerger et al. 2005). For most of the cells and tissues from which one wants to isolate nuclei, manual isolation is of course out of the question. But it is worth emphasizing that what has been learned from occasional studies employing manually isolated nuclei is likely to be a more reliable guide to understanding nuclear organization and function than has sometimes been realized by those members of the nucleus research community whose work is based on bulk nuclear isolation. A general point to be made is that although isolated nuclei served many important experimental uses during the modern era of research on the nucleus, their role in advancing the biochemistry of gene expression was surprisingly short-lived and quite limited. Cell-free systems based on isolated nuclei played only transitory and rather minor roles in the areas of DNA replication, transcription, and mRNA processing—usually giving way to more efficient soluble systems in short order. An exception was the use of isolated nuclei to allow RNA polymerase II to continue transcription and thus, by hybridization analysis of the extended chains, determine the boundaries of a transcription unit (Weber et al. 1977). In contrast to these limited roles of isolated nuclei in the investigation of gene expression, a very different cell-free system derived from frog eggs (Newport and Forbes 1987) was of monumental importance in advancing our understanding of chromatin

and nuclear envelope assembly and the control of cell-cycle progression. THE NUCLEUS COMPOSED

In 1976, the Federation of American Societies of Experimental Biology (FASEB) heroically collated all available information on the “properties of cells” and I was chosen to organize and edit the chapter on the nucleus. At the suggestion of the editors, and with FASEB’s permission, we are republishing here the tables of data on the nucleus from the original volume (Altman and Katz 1976) (see supplemental data online). These tables speak to painstaking analytical work carried out in a time gone by and rarely pursued today. But let there be no mistake, the concentrations of protein or RNA in various preparations of isolated nuclei, or extracts derived from them, can be a critical factor in the interpretation of certain kinds of studies. What is needed now to complement these important compositional data is additional biophysical information (reviewed in Pederson 2002a). One important step in this direction is that the fluid viscosity of the interchromatin space has now been estimated, based on diffusion coefficients of reporter molecules, to be approximately one-fifth that in water (Wachsmuth et al. 2000). The considerations of fluid viscosity and free versus anomalous diffusion are critical to the understanding of intranuclear transport and nuclear body dynamics (Wachsmuth et al. 2000). In terms of chemical kinetics one would also want to know the water concentration in the nucleus. Only then could one know, or at least estimate, the ionic strength of the solvent phase and the true concentrations of solutes. The importance of these biophysical parameters is conveyed by the fact that only a relatively small change in the intracellular ion concentration elicits a dramatic mitosis-like state, which is completely reversible (Robbins et al. 1970). Polyamines are abundant constituents of the nucleus in most cells and yet their possible functions are rarely considered. Iron is a surprisingly abundant component of the interphase nucleus and mitotic chromosomes and is essential for DNA

T. Pederson

replication (Robbins and Pederson, 1970). Many compositional issues such as these are rarely considered in current research on the nucleus and yet they can be extremely determinative. But, happily, there are steps in this direction, as biophysicists increasingly join the nucleus research community (for an “interdisciplinary” review see O’Brien et al. 2003). THE NUCLEUS DECONSTRUCTED

Major advances in biology were made by teasing a cell or its parts out, as from the squid giant axon, leading to the discovery of kinesin, or by glycerinating rabbit muscle to discover the biochemistry of its contraction. The major conceptual discovery about the nucleus, that it contains the genome, did not involve its dissection. But a desire to know its parts naturally arose in due course. Early attempts to isolate chromatin were made by the laboratories of Alfred Mirsky (Rockefeller) and James Bonner (Caltech) but these fractions were of dubious enrichment and the major advances in the field of chromatin and chromosome structure came from in situ studies (reviewed by DuPraw 1970). One of the first, convincing isolations of a subnuclear component was that of the nucleolus from starfish oocytes (Vincent 1955). Later, three groups went further. Palade and Siekevitz at Rockefeller purified guinea pig liver nuclei, subjected them to sonication and isolated nucleolar and nucleoplasmic fractions (Maggio et al. 1963). The demonstrated degree of fractionation was impressive and this work constituted one of the most important advances in the nucleus field at the time. Meanwhile, the laboratory of Harris Busch at Baylor College of Medicine developed a similar, sonication-based method for isolating nucleoli from rat hepatoma cells (Muramatsu et al. 1963) and subsequently this group, notably Ramachandra Reddy, exploited this method to identify a number of small nucleolar and nucleoplasmic RNAs long before their functions became known (for a review see Reddy and Busch 1988). Shortly thereafter, Sheldon Penman developed a method to resolve a HeLa cell nuclear fraction into nucleoli and nucleoplasm (Penman 1966).

His method involved the use of DNAse and high ionic strength and produced nucleolar and nucleoplasmic fractions that were less native than those obtained by the sonication method, although it led to spectacular advances in our understanding of RNA biosynthesis. More recently, mammalian and plant cell nucleoli have been purified and subjected to extensive proteomic analyses (Andersen et al. 2002; Scherl et al. 2002; Pendle et al. 2005; Hinsby et al. 2006; for reviews see Dundr and Misteli 2002; Pederson 2002b; Coute´ et al. 2006 ; Leung et al. 2006). Nucleoplasmic bodies known as interchromatin granule clusters (a.k.a. “speckles”) were isolated by David Spector and colleagues (Mintz et al. 1999) and later subjected by this group to proteomics analysis (Saitoh et al. 2004; reviewed in Lamond and Spector 2003). Meanwhile, Cajal bodies (reviewed by Gall 2000, 2009; Nizami et al. 2010) were isolated by Angus Lamond’s laboratory (Lam et al. 2002). An exciting aspect of this recent period was that these nuclear bodies were being analyzed in such molecular detail at the very time the dynamics of their components were being observed in live cell studies, as will be discussed below. Another nuclear structure appears in some cells, the perinucleolar compartment (Huang et al. 1997; 1998), and its presence has recently been found to have promising prognostic value in the staging of breast carcinoma (Kamath et al. 2005). It is a discoid, caplike structure intimately attached to the nucleolus and has thus resisted isolation so far. There presently are no clues to its function, beyond its accretion of certain small RNAs transcribed by RNA polymerase III (Matera et al. 1995; Weng et al. 2003). THE NUCLEUS TERRITORIALIZED

A major development was the discovery that interphase chromosomes occupy specific locations (Zorn et al. 1979; Cremer et al. 1982), which to some extent had been anticipated from classical studies of the arrangement of chromosomes in the metaphase plate (Rabl 1885) and by the actual coining of the term “chromosome territory” (Boveri 1909). The

The Nucleus, Introduced

modern concept of chromosome territories was promptly adopted in certain sectors of the human genetics and radiation biology communities but took more than two decades to gain broader traction (reviewed by Cremer and Cremer 2001; 2006), now with appreciated relevance to chromosomal translocations (see Pederson 2003 for a review), genome evolution (Tanabe et al. 2002; Foster and Bridger 2005) and the entire issue of how chromosome location relates to gene density and/or expression. This latter area has been hyperactive in the past few years (reviewed by Spector 2003; Pederson 2004; Gilbert et al. 2005; Sprout et al. 2005; Cremer et al. 2006; Akhtar and Gasser 2007; Misteli et al. 2007; Sexton et al. 2007; Takizawa et al. 2008; Towbin et al. 2009; Deniaud and Bickmore 2009) but the results have been surprisingly variable and confounding (for a particularly lucid summary see the Introduction in Meaburn et al. 2008). A distinct possibility that has arisen recently is that gene positioning is self-organizing, based not on expression per se but on the potential for sets of genes to be coregulated (Rajapakse et al. 2009; reviewed by Misteli 2009). A clinical feature of chromosome territoriality is the fact that the extreme cytological manifestation of gene repression, i.e., heterochromatin, has long served as a key landmark for pathologists, both in its extent and location. But an exciting new dimension of diagnostic potential lies in studies, mentioned earlier, of intranuclear gene locations in relation to not only reciprocal translocations (Roix et al. 2003; reviewed in Pederson 2003) but in more recent work on gene repositioning in solid tumors (e.g., Meaburn et al. 2009). The nucleus is territorialized not only with respect to the locations of the chromosomes themselves but also to a considerable extent with respect to the layout of expressed versus silenced genomic regions. This aspect of the nuclear structure field has recently taken one particularly surprising turn, reminding us that we are still in the early days. In most cells, heterochromatin is located at the nuclear periphery or in close approximation to the nucleolus. Remarkably, it has recently been reported that

in the rod photoreceptor cell nuclei of nocturnal animals, the heterochromatin is coalesced into a large central domain in the nucleus with the euchromatin displaced to more peripheral locations, with plausible speculations as to the efficiency of light transmission and retinal harvesting (Solovei et al. 2009; reviewed by Ragoczy and Groudine 2009). Another important recent development has been the introduction of methods to capture the interchromosome regions that lie in closest juxtaposition (Dekker et al. 2002; reviewed by Dostie and Dekker 2007). With this new methodology the field of genome organization is moving to a 4-D spatialtemporal registration, which is itself likely to be determinative of phenotype. The extraordinary packing density of interphase chromatin has been recently investigated with “Hi-C”, a powerful variation of the chromosome conformation capture methodology (LiebermanAiden et al. 2009; for a review see Langowski 2010).

THE NUCLEUS NOT LIKELY MATRIXED

Breaking up nuclei with sonication is one thing, extracting them with salt is another. The latter was a major effort of many labs in the 1950s and thereafter, most notably for isolating and characterizing histones. But others (e.g., Zbarsky and Georgiev 1959) employed graded salt extractions to fractionate total nuclear components. This yielded successive fractions of differing composition, not unexpected because the zwitterion concept of protein net charge had long been discovered by Arne Tiselius and advanced by John Edsall. There was therefore no reason to believe that cell components such as nuclei, mitochondria, or ribosomes would not release different sets of proteins as the ionic strength is elevated. Few cell biologists were comfortable with assigning a residue of the nucleus any relationship to the in vivo situation. But in 1974 this nuclear residue got renamed a “nuclear matrix” (Berezney and Coffey 1974) and later work gave rise to the idea that chromatin and nascent RNA are attached to it.

T. Pederson

The notion took hold in some quarters and gained a degree of traction as all sorts of entities were observed to be present in this fraction. But there was never any proof of the in vivo existence of such a structure nor is there proof today (for detailed reviews see Pederson 1998; 2000) and many leaders in the field of nuclear structure and function question the nuclear matrix concept. The consensual skepticism as to a nucleuswide, arborized network of filaments extending throughout the nucleoplasm certainly does not rule out the existence of short-range structural motifs. THE NUCLEUS IN DIVISION

The current focus of the cell division field on chromosome capture by kinetochore– microtubule interactions, how anaphase works mechanistically and the operation of mitotic checkpoints is of course well justified. But it is also possible that this emphasis may be missing one of the most intriguing of all events—the formation of daughter nuclei. Precursors of the nucleolus, nuclear envelope, and nuclear lamina congress, mature, and assemble in “daughter to be” cells as early as anaphase and definitively in telophase. This presently subsidiary effort in the cell division field needs more momentum as it is no less intriguing than anaphase, and likely involves very different mechanisms such as the free energy of DNA–lipid affinities and other chemical phenomena that are unique to telophase and the assembly of daughter nuclei, seemingly via pathways whose memory had not been erased from the previous mitosis. THE NUCLEUS IMAGED

In his detailed recipes, the histochemistry pioneer A.G. Everson Pearse described various ways to stain cells, including the nucleus (Pearse 1961). I tried many of these methods as a student including the aforementioned Feulgen reaction. The combination of methyl green and pyronin ended up as my favorite, with which I first saw nucleoli, to become a longstanding interest. At the end of the classical era, immunostaining and susbsequently the

advent of GFP advanced the localization of nuclear proteins and bodies, and the discovery of in situ nucleic acid hybridization (Gall and Pardue 1969) made possible the localization of both genes and RNAs. These methods, especially when powerfully used in combination, have made it possible to even demarcate subregions of nuclear domains formerly thought to be relatively homogenous, such as the granular component of the nucleolus, which now appears to be a landscape of distinct molecular zones rather than a uniform lawn of nascent ribosomes (Politz et al. 2002; 2005). Of course, these light microscopy advances have left ample room for the continuing application of electron microscopy. Particular progress has been made in the past two decades with respect to the ultrastructural analysis of both nuclear pore complexes, as mentioned earlier, as well as a defined messenger RNP, sometimes the two caught together (Mehlin et al. 1992). THE NUCLEUS IN MOTION, WITHIN

There was never much doubt that, around the less mobile chromosomes and nucleoli, molecules roam the nucleus. This emerged from numerous studies in which tagged molecules were microinjected into the nucleus and observed to display high mobility (e.g., Wang et al. 1990). Subsequently it became possible to express fluorescently tagged nuclear proteins and observe their dynamics. The first such study examined the dynamics of interchromatin granule clusters (Misteli et al. 1997) which revealed these nucleoplasmic bodies to be more dynamic than had been anticipated. Then, a new wave of studies appeared resulting from the application of the method of fluorescence recovery after photobleaching (FRAP). In the first of these, FRAP was employed to study the mobility of the DNA repair machinery tagged with GFP (Houtsmuller et al. 1999). This paper represented both a technical and conceptual milestone and soon thereafter FRAP was applied to several other nuclear proteins in a boomlet of important studies (Kruhlak et al. 2000; Lever et al. 2000; Misteli et al. 2000; Phair and Misteli 2000; Boisvert et al. 2001; Chen and

The Nucleus, Introduced

Huang 2001; reviewed in Pederson 2000b; Misteli 2001b; Pederson 2001b; Phair and Misteli 2001). These revealed much more rapid and/ or more extensive dynamics than would have been anticipated from either earlier in vitro work, or from the apparent stasis of certain nuclear bodies, constituting a true paradigm shift in the nucleus field (reviewed by Misteli 2001b). Even the nuclear lamina, which had long been viewed as one of the most stable structures in the nucleus, was found to undergo dynamic exchange of subunits, leading to an appreciable nucleoplasmic concentration (Moir et al. 2000; reviewed by Dechat et al. 2008) and there is increasing evidence that the dynamic lamins have functions while in the nucleoplasm (Malhas et al. 2007; Lee et al. 2009; reviewed by Shimi et al. 2008; Shumaker et al. 2008), and it now appears that nucleoporins do as well (Capelson et al. 2010; Kalverda et al. 2010). This wave of FRAP studies addressed the dynamics of protein constituents of nuclear bodies and was, of course, based on GFP. But GFP also enabled investigation of the movements of nuclear bodies and their constituents. Studies of the movements of chromosomes (Marshall et al. 1997; reviewed by Gasser 2002), promyelocytic leukemia (PML) bodies (Muratani et al. 2002) and Cajal bodies (Platani et al. 2002) were the first of these. The Marshall et al. study was ahead of its time and it was amusing to recall the incredulity expressed by some that interphase chromosomes, relatively giant structures, are moving, and with no dependence on metabolic energy. But others properly anticipated that there is of course no reason the chromosomes would not display this microscopic biophysical property, viz. the manifestation of the kinetic energy of any particle. Although GFP enabled these studies (for reviews see Misteli et al. 1997; Eils et al. 2000; Pederson 2000b; Misteli 2001b; Pederson 2001b; Chubb and Bickmore 2003; Dundr et al. 2002; Gasser 2002), investigating the intranuclear movements of the other major nuclear species, RNA, required different innovation. The fact that microinjection of a pre-mRNAs

or small nucleolar RNAs into the nucleus of mammalian cells resulted in localization with sites known to contain splicing machinery or the nucleoli, respectively (Wang et al. 1990; Jacobson et al. 1995; Jacobson and Pederson 1998; reviewed in Pederson 2001c) showed that introduced RNA is mobile. Further innovations led to tagging of endogenous RNA and these studies confirmed that poly(A) RNA is highly diffusive in the nucleus (Politz et al. 1998; 1999; reviewed in Politz and Pederson 2000; Pederson 2001c), and subsequent studies revealed, importantly, that this was true of specific mRNAs (Singh et al. 1999; Shav-Tal et al. 2004) and also 28S ribosomal RNA (Politz et al. 2003). These approaches also made it possible to investigate the live cell dynamics of RNA in relation to specific intranuclear structures, e.g., interchromatin granules (Politz et al. 2006). An increasing number of single-molecule level studies of nuclear molecular dynamics have appeared recently (Dange et al. 2008; Gru¨nwald et al. 2008; Siebrasse et al. 2009) and, together with revolutionary advances in the spatial resolution of diffraction-limited optical microscopy (for a brief review see Pederson 2006), one senses the dawning of a new era as systems biology enters the nucleus, or vice-versa (reviewed by Gorski and Misteli 2005). Another major advance was the introduction of a method to tag specific chromatin regions via the binding of GFP-tagged lac repressor to an integrated tandem array of the lac operator (Robinett et al. 1996), which in turn allowed the visualization in living cells of gene activation at discrete loci (Tsukamoto et al. 2000; Janicki et al. 2004) and the dynamics of transcription factors (Becker et al. 2002; Karpova et al. 2008; Sprouse et al. 2008). As mentioned earlier, the movements of interphase chromosomes have implications for the statistics of reciprocal exchanges between interphase chromosomes, with disease relevance (Roix et al. 2003; reviewed by Pederson 2003). Nucleoli also move to the extent that the chromosomes which contain the repeated rRNA genes are mobile. Diffusion, by definition, arises from the thermal energy inherent in the particle itself. However, the possibility that gene repositioning may be mediated by a process

T. Pederson

that uses metabolic energy has recently been the topic of initial studies (and debate). There is a growing body of evidence for actin and myosin in the nucleus, a field that has moved past its initial uncertainties and is now addressing functions (reviewed by Pederson and Aebi 2002; 2005; Pederson 2008). In particular, three recent studies have implicated actin-based processes in gene repositioning (Chuang et al. 2006; Dundr et al. 2007; Hu et al. 2008). This emerging concept is among the most deserving of vigilance in the nucleus field at this time. THE NUCLEUS THAT AWAITS US

Speculation is always risky. One thing is clear— the question of how nuclear bodies arise in the first place (reviewed by Misteli 2001c) has taken on a new dimension because of the discovery that a Cajal body can assemble when key components are experimentally addressed to a specific nuclear site (Kaiser et al. 2008). Other recent studies raise the possibility that noncoding, nucleus-retained RNAs may play roles in nuclear architecture (Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009; reviewed by Wilusz et al. 2009). Also on the present horizon are stem cell issues that call for increasing input from experts on the nucleus. Stem cells have been hyped but it is early days and the nucleus cell biology community has much to offer, both by science and skepticism (for the latter perspective see Lander 2009). The essence of “stem-cellness” is an asymmetric descent of phenotypic potential vs. maintaining a continual seed of replenishment. So the two daughter cells have to be profoundly different but it is a difference we presently grasp very dimly. This is as deep a problem in cell biology as there is today, and there are already encouraging signs of progress (e.g., Parnell and Stillman 2008). Another field that is ripe for the intervention of nucleus experts is somatic nuclear transfer, in which it is the reaction of the introduced nucleus to the maternal environment, known to be dominant from classical studies, that sets in motion the reprogrammed developmental events. What are the molecules that underlie this powerful influence of the egg cytoplasm?

Finally, there can be no doubt that the increasing ability to study nuclear dynamics and function in living cells, now reaching single molecule level detection, constitutes one of the most powerful advances. The recent breaking of the classical diffraction limit of optical microscopy brings the nucleus into nanoscale range. The possible coaptation of these breakthrough approaches with instructive cell types and model systems stirs excitement for what lies ahead. The nucleus research community is living in very interesting times.

CODA

The assumption underlying this collection on nuclear structure and function is that the nucleus has reached a stage of enabling coherence as part of the epistemological structure of modern biological science. However, there are likely to be many things about the nucleus that we don’t yet know and may not know anytime soon. We can only hope that what the geneticist J.B.S. Haldane posited on the cosmos will prove not to be true for the nucleus: “Now, my suspicion is that the universe is not only queerer than we suppose, but queerer than we can suppose.” If we appropriately bear in mind that the nucleus may be more complicated than we may have once thought, and yet just may be knowable, then this very belief may empower us and our students and successors to penetrate the subject’s awaiting depths, the next of which now beckon. There is every reason to believe in this program. So let us be of good cheer.

ACKNOWLEDGMENTS

I am indebted to Joseph Gall for characteristically insightful suggestions on the manuscript. Cited work from the author’s laboratory was funded by grants from the National Institutes of Health (GM-21595, GM-23914 and GM-60551), the National Science Foundation (MCB-0445841), the American Cancer Society (CD-126), and the G. Harold and Leila Y. Mathers Foundation. Some important ideas

The Nucleus, Introduced

that stirred in the modern era of the nucleus field were catalyzed by a series of symposia convened by the author at the Marine Biological Laboratory, Woods Hole, supported by a grant from the Grass Foundation and the late Ellen Grass. REFERENCES Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprato A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007. The molecular architecture of the nuclear pore complex. Nature 450: 695– 701. Altman PL, Katz DD. 1976. Biological Handbooks. I. Cell Biology. Federation of American Societies for Experimental Biology, Bethesda, Maryland. Andersen JS, Lyon CE, Fox AH, Leung AKL, Lam YW, Steen H, Mann M, Lamond AI. 2002. Directed proteomic analysis of the human nucleolus. Curr Biol 12: 1 –11. Andrade LEC, Chan EKL, Rasˇka I, Peebles CL, Roos G, Tan EM. 1991. Human autoantibody to a novel protein of the nuclear coiled body: immunological characterization and cDNA cloning of p80-coilin. J Exp Med 173: 1407– 1419. Avery OT, MacLeod CM, McCarty M. 1944. Studies on the chemical nature of the substance inducing transformation of pneumococcal types. I. Induction of transformation by a deoxyribonucleic acid fraction isolated from pneumococcus type III. J Exp Med 79: 137–158. Barton CA, Laishram RS, Anderson RA. 2010. Nuclear phosphoinositides: a signaling enigma wrapped in a compartmental conundrum. Trends Cell Biol 20: 25–35. Bauer F. 1830–1838. Illustrations of orchidaceous plants Ridegway, London. Becker M, Baumann C, John S, Walker DA, Vigneron M, McNally JG, Hager GL. 2002. Dynamic behavior of transcription factors on a natural promoter in living cells. EMBO reports 3: 1188– 1194. Berezney R, Coffey DS. 1974. Identification of a nuclear protein matrix. Biochem Biophys Res Commun 60: 1410– 1417. Bernhard W, Haguenau F, Oberling C. 1952. L’ultrastructure du nucleole de quelques cellules animals, re´ve´le´e par le microscope e´lectronique. Experientia 8: 58– 63. Bhorjee JS, Pederson T. 1972. Nonhistone chromosomal proteins in synchronized HeLa cells. Proc Natl Acad Sci 69: 3345–3349. Blobel G, Potter VR. 1966. Nuclei from rat liver: isolation method that combines purity with high yield. Science 154: 1662–1665. Blackburn EH, Gall JG. 1978. A tandemly repeated sequence at the termini of the extrachromosomal ribosomal RNA genes in Tetrahymena. J Mol Biol 120: 33–53. Boisvert FM, Kruhlak MJ, Box AK, Hendzel MJ, BazettJones DP. 2001. The transcription coactivator CBP is a dynamic component of the promyelocytic leukemia nuclear body. J Cell Biol 152: 1089–1106.

Borun TW, Scharff MD, Robbins E. 1967. Preparation of mammalian polyribosomes with the detergent Nonidet-P40. Biochim Biophys Acta 149: 302– 304. Boveri T. 1909. Die blastomerenkern von Ascaris megalocephala und die theorie der chromosomenindividualita¨t. Arch Zellforsch 3: 181– 268. Brown R. 1829– 1832. Observations on the organs and mode of fecundation in Orchideae and Asclepiadeae. Trans Linn Soc Lond 16: 685– 746. Cajal SRy. 1903. Un sencillo metodo de coloracion selectiva del reticulo protoplasmatico y sus efectos en los diversos organos nerviosos de vertebrados e invertebrados. Trab Lab Invest Biol Univ Madrid 2: 129– 221. Cajal SRy. 1910. El nu´cleo de las ce´lulas piramidales del cerebro humano y del algunos mamiferos. Trab Lab Invest Biol Univ Madrid 8: 27–62. Capelson M, Liang Y, Schulte R, Mair W, Wagner U, Hetzer MW. 2010. Chromatin-bound nuclear pore components regulate gene expression in higher eukaryotes. Cell 140: 372 –383. Chauveau J, Moule Y, Rouiller C. 1956. Isolation of pure and unaltered liver nuclei: morphology and biochemical composition. Exp Cell Res 11: 317– 321. Chen D, Huang S. 2001. Nucleolar components involved in ribosome biogenesis cycle between the nucleolus and nucleoplasm in interphase cells. J Cell Biol 153: 169– 176. Chuang CH, Carpenter AE, Fuchsova B, Johnson T, de Lanerolle P, Belmont AS. 2006. Long-range directional movement of an interphase chromosome site. Curr Biol 16: 825– 831. Chubb JR, Bickmore WA. 2003. Considering nuclear compartmentalization in the light of nuclear dynamics. Cell 112: 403–406. Clemson CM, Hutchinson JN, Sara SA, Ensminger AW, Fox AH, Chess A, Lawrence JB. 2009. An architectural role for a nuclear noncoding RNA: NETA1 RNA is essential for the structure of paraspeckles. Mol Cell 33: 717– 726. Cocco L, Faenza I, Follo MY, Billi AM, Ramazzotti G, Papa V, Martelli AM, Manzoli L. 2009. Nuclear inositides: Pl-PLC signaling in cell growth, differentiation and pathology. Adv Enzyme Reg 49: 2 –10. Coute´ Y, Burgess JA, Diaz JJ, Chichester C, Lisacek F, Greco A, Sanchez JC. 2006. Deciphering the human nucleolar proteome. Mass Spect Rev 25: 215– 234. Cremer T, Cremer C. 2001. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat Rev Genet 2: 292–301. Cremer T, Cremer C. 2006. Rise, fall and resurrection of chromosome territories: a historical perspective. Part II: Fall and resurrection of chromosome territories during the 1950s and 1960s. Part III: Chromosome territories and the functional nuclear architecture: experiments and models from the 1990s to the present. Eur J Histochem 50: 223 –272. Cremer T, Cremer M, Dietzel S, Muller S, Solovei I, Fakan S. 2006. Chromosome territories—a functional nuclear landscape. Curr Opin Cell Biol 18: 307 –316. Cremer T, Cremer C, Schenider T, Baumann H, Hens L, Kirsch-Volders M. 1982. Analysis of chromosome positions in the interphase nucleus of Chinese hamster cells

T. Pederson by laser-UV-microirradiation experiments. Hum Genet 62: 201 –209. Dahlberg JE, Lund E, Goodwin EB. 2003. Nuclear translation: what is the evidence? RNA 9: 1 –8. Dange T, Gru¨nwald D, Gru¨nwald A, Peters R, Kubitscheck U. 2008. Autonomy and robustness of translocation through the nuclear pore complex: a single-molecule study. J Cell Biol 183: 77– 86. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832 –853. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306–1311. Deniaud E, Bickmore WA. 2009. Transcription and the nuclear periphery: edge of darkness? Curr Opin Genet Dev 19: 187 –191. Divecha N, Banfic´ F, Irvine RF. 1993. Inositides and the nucleus and inositides in the nucleus. Cell 74: 405– 407. Dostie J, Dekker J. 2007. Mapping networks of physical interactions between genomic elements using 5C technology. Nat Protocols 2: 988 –1002. Dundr M, McNally JG, Cohen J, Misteli T. 2002. Quantitation of GFP-fusion proteins in single living cells. J Struct Biol 140: 92–99. Dundr M, Misteli T. 2002. Nucleolomics: an inventory of the nucleolus. Mol Cell 9: 5 –7. Dundr M, Ospina JK, Sung MH, John S, Upender M, Ried T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095– 1103. DuPraw EJ. 1970. DNA and Chromosomes Holt, Rinehart & Winston, New York. Duryee WR. 1954. Microdissection studies on human ovarian eggs. Trans NY Acad Sci 17: 103– 108. Eils R, Gerlich D, Tvarusko´ W, Spector DL, Misteli T. 2000. Quantitative imaging of pre-mRNA splicing factors in living cells. Mol Biol Cell 11: 413– 418. Fawcett DW. 1966. On the occurrence of a fibrous lamina on the inner aspect of the nuclear envelope in certain cells of vertebrates. Am J Anat 119: 129 –145. Fernandez-Martinez J, Rout MP. 2009. Nuclear pore complex biogenesis. Curr Opin Cell Biol 21: 603– 612. Foster HA, Bridger JM. 2005. The genome and the nucleus: a marriage made by evolution. Chromosoma 114: 212 –229. Gall JG. 1964. Electron microscopy of the nuclear envelope. Protoplasmatologia 5: 4 –25. Gall JG. 1967. Octagonal nuclear pores. J Cell Biol 32: 391 – 399. Gall JG. 1974. Free ribosomal RNA genes in the macronucleus of Tetrahymena. Proc Natl Acad Sci USA 71: 3078– 3081. Gall JG. 1996. A pictorial history. Views of the cell. American Society for Cell Biology, Bethesda, MD. Gall JG. 2000. Cajal bodies: the first 100 years. Ann Rev Cell Dev Biol 16: 273– 300. Gall JG. 2009. Chromosome odds and ends. Ann Rev Cell Dev Biol, in press.

Gall JG, Pardue ML. 1969. Formation and detection of RNA-DNA hybrid molecules in cytological preparations. Proc Natl Acad Sci 63: 378– 383. Gasser SM. 2002. Visualizing chromatin dynamics in interphase nuclei. Science 296: 1412–1416. Gilbert N, Gilchrist S, Bickmore WA. 2005. Chromatin organization in the mammalian nucleus. Int Rev Cytol 242: 283–336. Goldstein L. 1970. On the question of protein synthesis by cell nuclei. Adv Cell Biol 1: 187–210. Gorski S, Misteli T. 2005. Systems biology in the cell nucleus. J Cell Sci 118: 4083– 4092. Gru¨nwald D, Martin RM, Buschmann V, Bazett-Jones DP, Leonhardt H, Kubitscheck U, Cardoso MC. 2008. Probing intranuclear environments at the single-molecule level of detection. Biophys J 94: 2847–2858. Gurney T, Eliceiri GL. 1980. Intracellular distribution of low molecular weight RNA species in HeLa cells. J Cell Biol 87: 398– 403. Handwerger KE, Cordero JA, Gall JG. 2005. Cajal bodies, nucleoli, and speckles in the Xenopus oocyte nucleus have a low-density, sponge-like structure. Mol Biol Cell 16: 202– 211. Handwerger KE, Gall JG. 2006. Subnuclear organelles: new insights into form and function. Trends Cell Biol 16: 19– 26. Harris H. 1999. The birth of the cell. Yale University Press, New Haven. Hinsby AM, Kiemer L, Karlberg EO, Lage K, Fausbøll A, Juncker AS, Andersen JS, Mann M, Brunak S. 2006. A wiring of the human nucleolus. Mol Cell 22: 285– 295. Holland CA, Mayrand SH, Pederson T. 1980. Sequence complexity of nuclear and messenger RNA in HeLa cells. J Mol Biol 138: 755 –778. ¨ ber die chemische zusammensetHoppe-Seyler F. 1871. U zung des eiters. Hoppe-Seyler’s Medizinisch-Chemischen Untersuchungen 4: 486–501. Houtsmuller AB, Rademakers S, Nigg AL, Hoogstraten D, Hoeijmakers JHJ, Vermuelen W. 1999. Action of DNA repair endonuclease ERCC1/XPF in living cells. Science 284: 958–961. Hu Q, Kwon YS, Nunez E, Cardamone MD, Hutt KR, Ohgi KA, Garcia-Bassets I, Rose DW, Glass CK, Rosenfeld MG, et al. 2008. Enhancing nuclear receptor-induced transcription requires nuclear motor and LSD1-dependent gene networking in interchromatin granules. Proc Natl Acad Sci USA 105: 19199– 19204. Huang S, Derrinck TJ, Ellisman MH, Spector DL. 1997. The dynamic organization of the perinucleolar compartment in the cell nucleus. J Cell Biol 137: 965– 974. Huang S, Derrinck TJ, Ellisman MH, Spector DL. 1998. The perinucleolar compartment and transcription. J Cell Biol 143: 35–47. Iborra FJ, Jackson DA, Cook PR. 2001. Coupled transcription and translation within nuclei of mammalian cells. Science 293: 1139– 1142. Jacobson MR, Cao LC, Wang YL, Pederson T. 1995. Dynamic localization of RNase MRP RNA in the nucleolus observed by fluorescent RNA cytochemistry in living cells. J Cell Biol 131: 1649–1658.

The Nucleus, Introduced Jacobson MR, Pederson T. 1998. A 7-methylguanosine cap commits U3 and U8 small nuclear RNAs to the nucleolar localization pathway. Nucleic Acids Res 26: 756– 760. Janicki SM, Tsukamoto T, Salghetti SE, Tansey WP, Sachidanandam R, Prasanth KV, Ried T, Shav-Tal Y, Bertrand E, Singer RH, et al. 2004. From silencing to gene expression: real-time analysis in single cells. Cell 116: 683– 698. Kaiser TE, Intine RV, Dundr M. 2008. De novo formation of a subnuclear body. Science 322: 1713– 1717. Kalverda B, Pickersgill H, Shloma VV, Fornerod M. 2010. Nucleoporins directly stimulate expression of developmental and cell-cycle genes inside the nucleoplasm. Cell 140: 360– 371. Kamath RV, Thor AD, Wang C, Edgerton SM, Slusarczyk A, Leary DJ, Wang J, Wiley EL, Jovanovic B, Wu Q, et al. 2005. Perinucleolar compartment prevalence has an independent prognostic value for breast cancer. Cancer Res 65: 246– 253. Karpova TS, Kim MJ, Spriel C, Nalley K, Stasevich TJ, Kherrouche Z, Heliot L, McNally JG. 2008. Concurrent fast and slow cycling of a transcriptional activator at an endogenous promoter. Science 319: 466– 469. King MC, Drivas TG, Blobel G. 2008. A network of nuclear envelope membrane proteins linking centromeres to microtubules. Cell 134: 427–438. Kleinschmidt AM, Pederson T. 1990. RNA processing and ribonucleoprotein assembly studied in vivo by RNA transfection. Proc Natl Acad Sci 87: 1283–1287. Kruhlak MJ, Lever MA, Fischle W, Verdin E, Bazaett-Jones DP, Hendzel MJ. 2000. Reduced mobility of the alternate splicing factor (ASF) through the nucleoplasm and steady state speckle compartments. J Cell Biol 150: 41–51. Lake JA. 2009. Evidence of an early prokaryotic endosymbiosis. Nature 460: 967– 971. Lam YW, Lyon CE, Lamond AI. 2002. Large-scale isolation of Cajal bodies from HeLa cells. Mol Biol Cell 13: 2461– 2473. Lamond AI, Spector DL. 2003. Nuclear speckles: a model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605–612. Lander AD. 2009. The ‘stem cell’ concept: is it holding us back? J Biol 8: 70. Langowski J. 2010. Chromosome conformation by crosslinking: polymer physics matters. Nucleus 1: 13–16. Lee DC, Welton KL, Smith ED, Kennedy BK. 2009. A-type nuclear lamins act as transcriptional repressors when targeted to promoters. Exp Cell Res 315: 996 –1007. Lever MA, Th’ng JP, Sun X, Hendzel MJ. 2000. Rapid exchange of histone H1.1 on chromatin in living cells. Nature 408: 873–876. Leung AK, Trinkle-Mullcahy L, Lam YW, Andersen JS, Mann M, Lamond AI. 2006. NOPdb: nucleolar proteome database. Nucleic Acid Res 34: D218–220. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289 –293. Lund E, Paine P. 1990. Nonaqueous isolation of transcriptionally active nuclei from Xenopus oocytes. Methods Enzymol 181: 36–43.

Maggio R, Siekevitz P, Palade GE. 1963. Studies on isolated nuclei. I. Isolation and chemical characterization of a nuclear fraction from guinea pig liver. J Cell Biol 18: 267 –291. Maggio R, Siekevitz P, Palade GE. 1963. Studies on isolated nuclei. II. Isolation and chemical characterization of nucleolar and nucleoplasmic subfractions. J Cell Biol 18: 292– 312. Malhas A, Lee CF, Sanders R, Saunders NJ, Vaux DJ. 2007. Defects in lamin B1 expression or processing affect interphase chromosome position and gene expression. J Cell Biol 176: 593 –603. Margulis L, Dolan MF, Guerrero R. 2000. The chimeric eukaryote: origin of the nucleus from the karyomastigont in amitochondriate protists. Proc Natl Acad Sci 97: 6954–6959. Marshall WF, Straight A, Marko JF, Swedlow J, Dernburg A, Belmont A, Murray AW, Agard DA, Sedat JW. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930–939. Martelli AM, Gilmour RS, Bertagnolo V, Neri LM, Manzoli L, Cocco L. 1991. Nuclear localization and signalling activity of phosphoinositidase Cb in Swiss 3T3 cells. Nature 358: 242 –244. Matera AG, Frey MR, Margelot K, Wolin SL. 1995. A perinucleolar compartment contains several RNA polymerase III transcripts as well as the polypyrimidine tract binding protein, hnRNP I. J Cell Biol 129: 1181–1193. Meaburn KJ, Newbold RF, Bridger JM. 2008. Positioning of human chromosomes in murine cell hybrids according to synteny. Chromosoma 117: 579– 591. Meaburn KJ, Gugla PR, Khan S, Lockett SJ, Misteli T. 2009. Disease-specific gene repositioning in breast cancer. J Cell Biol 187: 801 –812. Mehlin H, Daneholt B, Skoglund U. 1992. Translocation of a specific premessenger ribonucleoprotein particle through the nuclear pore studied with electron microscope tomography. Cell 15: 605– 613. ¨ ber die chemische zusammensetzung Miescher F. 1871. U der eiterzellen. Hoppe-Seyler’s Medizinisch-Chemischen Untersuchungen 4: 441–460. Mintz PJ, Patterson SD, Neuwald AF, Spahr CS, Spector DL. 1999. Purification and biochemical characterization of interchromatin granule clusters. EMBO J 18: 4308–4320. Misteli T, Gunjan A, Hock R, Bustin M, Brown DT. 2000. Dynamic binding of histone H1 to chromatin in living cells. Nature 408: 877–881. Misteli T. 2001a. Where the nucleus comes from. Trends Cell Biol 11: 149. Misteli T. 2001b. Protein dynamics: implications for nuclear architecture and gene expression. Science 291: 843– 847. Misteli T. 2001c. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Misteli T. 2007. Beyond the sequence: cellular organization of genome function. Cell 128: 787– 800. Misteli T. 2009. Self-organization in the genome. Proc Natl Acad Sci USA 106: 6885– 6886. Misteli T, Ca´ceres JF, Spector DL. 1997. The dynamics of a pre-mRNA splicing factor in living cells. Nature 387: 523 –527.

T. Pederson Moir RD, Yoon M, Khuon S, Goldman RD. 2000. Nuclear lamins A and B1: different pathways of assembly during nuclear envelope formation in living cells. J Cell Biol 151: 1155–1168. Muramatsu M, Smetana K, Busch H. 1963. Quantitative aspects of isolation of nucleoli of the Walker carcinosarcoma and liver of the rat. Cancer Res 23: 510– 518. Muratani M, Gerlich D, Janicki SM, Gebhard M, Eils R, Spector DL. 2002. Metabolic-energy-dependent movement of PML bodies within the mammalian cell nucleus. Nature Cell Biol 4: 106 –110. Nathanson L, Xia T, Deutscher MP. 2003. Nuclear protein synthesis: a re-evaluation. RNA 9: 9– 13. Newport JW, Forbes DJ. 1987. The nucleus: structure, function, and dynamics. Ann Rev Biochem 56: 535–565. Nizami Z, Deryusheva S, Gall J. 2010. The Cajal body and histone locus body. Cold Spring Harb Perspect Biol 2: a000653. O’Brien TP, Bult CJ, Cremer C, Grunze M, Knowles BB, Langowski J, McNally J, Pederson T, Politz JC, Pombo A, et al. 2003. Genome function and nuclear architecture: from gene expression to nanoscience. Genome Res 13: 1029– 1041. Olins AL, Olins DE. 1974. Spheroid chromatin units (n bodies). Science 183: 330 –332. Paine PL, Johnson ME, Lau YT, Tluczek LJM, Miller DS. 1992. The oocyte nucleus isolated in oil retains in vivo structure and functions. Biotechniques 13: 238 –245. Parnell EJ, Stillman DJ. 2008. Getting a transcription factor to only one nucleus following mitosis. PLoS Biol 6: 1829– 1832. Pearse AGE. 1961. Histochemistry. 2nd edition, Little, Brown and Co., Boston. Pederson T. 1976. Cellular aspects of histone synthesis. In Protein synthesis (ed. E.H. McConkey), Vol. 2: pp. 69– 123, Marcel Dekker, New York. Pederson T. 1998. Thinking about a nuclear matrix. J Mol Biol 277: 147–159. Pederson T. 2000a. Half a century of “the nuclear matrix”. Mol Biol Cell 11: 799– 805. Pederson T. 2000b. Diffusional protein transport within the nucleus: a message in the medium. Nature Cell Biol 2: E73– 74. Pederson T. 2001a. Is the nucleus in need of translation? Trends Cell Biol 11: 395–397. Pederson T. 2001b. Protein mobility within the nucleus— what are the right moves? Cell 104: 635–638. Pederson T. 2001c. Fluorescent RNA cytochemistry: tracking gene transcripts in living cells. Nucleic Acids Res 29: 1013– 1016. Pederson T. 2002a. Dynamics and genome-centricity of inter-chromatin domains in the nucleus. Nat Cell Biol 4: E287– E291. Pederson T. 2002b. Proteomics of the nucleolus: more proteins, more functions? Trends Biochem. Sci 27: 111 –112. Pederson T. 2003. Gene territories and cancer. Nature Genetics 34: 242–243. Pederson T. 2004. The spatial organization of the genome in mammalian cells. Curr Opin Genet Dev 14: 203–209.

Pederson T. 2005. In memoriam- Hans Ris. Chromosoma 114: 307– 309. Pederson T. 2006. Diffraction at the Brandeburg g(r)ate. EMBO reports 7: 1202–1205. Pederson T. 2008. As functional nuclear actin comes into view, is it globular, filamentous, or both? J Cell Biol 180: 1061– 1064. Pederson T. 2010. An olympian protozoan. Nucleus 1: 1– 3. Pederson T, Aebi U. 2002. Actin in the nucleus: what form and what for? J Struct Biol 140: 3 –9. Pederson T, Aebi U. 2005. Nuclear actin extends, with no contraction in sight. Mol Biol Cell 16: 5055– 5060. Pendle AF, Clark GP, Boon R, Lewandowska D, Lam YW, Andersen J, Mann M, Lamond AI, Brown JW, Shaw PJ. 2005. Proteomic analysis of the Arabidopsis nucleolus suggest novel nucleolar functions. Mol Biol Cell 16: 260–269. Penman S. 1966. RNA metabolism in the Hela cell nucleus. J Mol Biol 17: 117– 130. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604 –609. Phair RD, Misteli T. 2001. Kinetic modeling approaches to in vivo imaging. Nat Rev Mol Cell Biol 2: 898 –907. Platani M, Goldberg I, Lamond AI, Swedlow JR. 2002. Cajal body dynamics and association with chromatin are ATP-dependent. Nat Cell. Biol 4: 502– 508. Politz JCR, Pederson T. 2000. Review: movement of mRNA from transcription site to nuclear pores. J Struc Biol 129: 252 –257. Politz JC, Lewandowski LB, Pederson T. 2002. Signal recognition particle RNA localization within the nucleolus differs from the classical sites of ribosome synthesis. J Cell Biol 159: 411– 418. Politz JCR, Tuft RA, Pederson T. 2003. Diffusion-based transport of nascent ribosomes in the nucleus. Mol Biol Cell 14: 4805– 4812. Politz JC, Brown ES, Wolf DE, Pederson T. 1998. Intranuclear diffusion and hybridization state of oligonucleotides measured by fluorescence correlation spectroscopy in living cells. Proc Natl Acad Sci 95: 6043–6048. Politz JCR, Polena I, Trask I, Bazett-Jones DP, Pederson T. 2005. A nonribosomal landscape in the nucleolus revealed by the stem cell protein nucleostemin. Mol Biol Cell 16: 3401– 4310. Politz JC, Tuft RA, Pederson T, Singer RH. 1999. Movement of nuclear poly(A) RNA throughout the interchromatin space in living cells. Curr Biol 9: 285– 291. Politz JCR, Tuft RA, Prasanth KV, Baudendistel N, Fograt KE, Lifshitz LM, Langowski J, Spector DL, Pederson T. 2006. Rapid, diffusional shuttling of poly(A) RNA between nuclear speckles and the nucleoplasm. Mol Biol Cell 17: 1239–1249. Poole A, Penny D. 2001. Does endo-symbiosis explain the origin of the nucleus? Nat Cell Biol 3: E173– E174. Portugal FH, Cohen JS. 1977. ACentury of DNA. MIT Press, Cambridge, MA. Purkyneˇ JE. 1825. Symbolae ad ovi avium historiam ante incubationem. 1st ed. Vratislaviae. Reprinted 1948: Opera selecta. Tomus 1, Opera facultatis medicae universitatis carolinae pragensis. ¨ ber zelltheilung. Morph Jb 10: 214–330. Rabl C. 1885. U

The Nucleus, Introduced Ragoczy T, Groudine M. 2009. The nucleus inside outthrough a rod darkly. Cell 137: 205–207. Rajapakse I, Perlman MD, Scalzo D, Kooperberg C, Groudine M, Kosak ST. 2009. The emergence of lineage-specific chromosomal topologies from coordinate gene regulation. Proc Natl Acad Sci USA 106: 6679– 6684. Rasˇka I, Andrade LEC, Ochs RL, Chan EKL, Chang CM, Roos G, Tan EM. 1991. Immunological and ultrastructural studies of the nuclear coiled body with autoimmune antibodies. Exp Cell Res 195: 27–37. Reddy R, Busch H. 1988. Small nuclear RNAs: sequences, structure and modifications. In Structure and function of major and minor small nuclear ribonucleoprotein particles M.L. Birnstiel, ed. Springer-Verlag, Berlin, pp. 1–37. Robinett CC, Straight A, Li G, Willhelm C, Sudlow G, Murray A, Belmont AS. 1996. In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. J Cell Biol 135: 1685–1700. Robbins E, Pederson T. 1970. Iron: its intracellular localization and possible role in cell division. Proc Natl Acad Sci USA 66: 1244– 1251. Robbins E, Pederson T, Klein P. 1970. Comparison of mitotic phenomena and effects induced by hypertonic solutions in HeLa cells. J Cell Biol 44: 400 –416. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in lymphomas. Nat Genetics 34: 287– 291. Rotte C, Martin W. 2001. Does endo-symbiosis explain the origin of the nucleus? Nat Cell Biol 3: E173–E174. Roux KJ, Crisp ML, Liu Q, Kim D, Kozlov S, Stewart CL, Burke B. 2009. Nesprin 4 is an outer nuclear membrane protein that can induce kinesin-mediated cell polarization. Proc Natl Acad USA 106: 2194– 2199. Saitoh N, Spahr CS, Patterson SD, Bubulya P, Neuwald AF, Spector DL. 2004. Proteomic analysis of interchromatin granule clusters. Mol Biol Cell 15: 3876–3890. Sasaki YTF, Ideue T, Sano M, Mituyama T, Hirose T. 2009. MEN e/b noncoding RNAs are essential for structural integrity of nuclear paraspeckles. Proc Natl Acad Sci USA 106: 2525– 2530. Scherl A, Coute´ Y, Deon C, Calle K, Kindbeiter K, Sanchez JD, Greco A, Hochstrasser D, Diaz JJ. 2002. Functional proteomic analysis of human nucleolus. Mol Biol Cell 13: 4100–4109. Sexton T, Schober H, Fraser P, Gasser SM. 2007. Gene regulation through nuclear organization. Nat Struc. Mol Biol 14: 1049–1055. Shav-Tal Y, Darzacq X, Shenoy SM, Fusco D, Janicki SM, Spector DL, Singer RH. 2004. Dynamics of single mRNPs in nuclei of living cells. Science 304: 1797– 1800. Shimi T, Pfleghaar K, Kojima S, Pack CG, Solovei I, Goldman AE, Adam SA, Shumaker DK, Kinjo M, Cremer T, et al. 2008. The A- and B-type nuclear lamin networks: microdomains involved in chromatin organization and transcription. Genes Dev 22: 3409– 3421. Shumaker DK, Solimando L, Sengupta K, Shimi T, Adam SA, Grunwald A, Strelkov SV, Aebi U, Cardoso MC, Goldman RD. 2008. The highly conserved nuclear lamin

Ig-fold binds to PCNA: its role in DNA replication. J Cell Biol 181: 269– 280. Siebrasse JP, Veith R, Dobay A, Leonhardt H, Daneholt B, Kubitscheck U. 2009. Discontinuous movement of mRNP particles in nucleoplasmic regions devoid of chromatin. Proc Natl Acad Sci 105: 20291–20296. Singh OP, Bjo¨rkroth B, Masich S, Wieslander L, Daneholt B. 1999. The intranuclear movement of Balbiani ring premessenger ribonucleoprotein particles. Exp Cell Res 251: 135–146. Solovei I, Kreysing M, Lanctoˆt C, Ko¨sem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell 137: 356 –368. Spector DL. 1993. Macromolecular domains within the cell nucleus. Ann Rev Cell Biol 9: 265–315. Spector DL. 2001. Nuclear domains. J Cell Sci 114: 2891– 2893. Spector DL. 2003. The dynamics of chromosome organization and gene regulation. Ann Rev Biochem 72: 573 –608. Spector DL. 2006. SnapShot: cellular bodies. Cell 127: 1070. Sprouse RO, Karpova TS, Mueller F, Dasgupta A, McNally JG, Auble DT. 2008. Regulation of TATA-binding protein dynamics in living yeast cells. Proc Natl Acad Sci USA 105: 13304– 13308. Sprout D, Gilbert N, Bickmore WA. 2005. The role of chromatin structure in regulating the expression of clustered genes. Nat Rev Genet 6: 775 –781. Starr DA. 2009. A nuclear-envelope bridge positions nuclei and moves chromosomes. J Cell Sci 122: 577–586. Sunwoo H, Dinger ME, Wilusz JE, Amaral PP, Mattick JS, Spector DL. 2009. MEN e/b nuclear-retained noncoding RNAs are up-regulated upon muscle differentiation and are essential components of paraspeckles. Genome Res 19: 347 –359. Takizawa T, Meaburn KJ, Misteli T. 2008. The meaning of gene positioning. Cell 135: 9– 13. Tanabe H, Muller S, Neusser M, von Hase J, Calcagno E, Cremer M, Solovei I, Cremer C, Cremer T. 2002. Evolutionary conservation of chromosome territory arrangements in cell nuclei from higher primates. Proc Natl Acad Sci USA 99: 4424– 4429. Towbin BD, Meister P, Gasser SM. 2009. The nuclear envelope- a scaffold for silencing? Curr Opin Genet Dev 19: 180– 186. Traub A, Kaufmann E, Ginzburg-Tietz Y. 1964. Studies on nuclear ribosomes. I. Association of DPN-pyrophosphorylase with nuclear ribosomes in normal and neoplastic tissues. Exp Cell Res 34: 371–383. Tsukamoto T, Hashiguchi N, Janicki SM, Tumbar T, Belmont AS, Spector DL. 2000. Visualization of gene activity in living cells. Nat Cell Biol 2: 871 –878. Vincent WS. 1955. Structure and chemistry of nucleoli. Int Rev Cytol 4: 269–298. Wachsmuth M, Waldeck W, Langowski J. 2000. Anomalous diffusion of fluorescent probes inside living cell nuclei investigated by spatially-resolved fluorescence correlation spectroscopy. J Mol Biol 298: 677– 689.

T. Pederson Wang J, Cao LG, Wang YL, Pederson T. 1990. Localization of pre-messenger RNA at discrete nuclear sites. Proc Natl Acad Sci 88: 7391–7395. Weber J, Jelinek W, Darnell JE. 1977. The definition of large viral transcription unit late in Ad2 infection of HeLa cells: mapping of nascent RNA molecules labeled in isolated nuclei. Cell 10: 611– 616. Weng C, Politz JC, Pederson T, Huang S. 2003. RNA polymerase III transcripts and the PTB protein are essential for the integrity of the perinucleolar compartment. Mol Biol Cell 14: 2425– 2435. Wilson EB. 1925. The cell in development and heredity. 3rd edition. Macmillan, New York, NY, pp. 28–57.

Wilusz JE, Sunwoo H, Spector DL. 2009. Long noncoding RNAs: functional surprises from the RNA world. Genes Dev 23: 1494–1504. Zbarsky IB, Georgiev GP. 1959. Cytological characteristics of protein and nucleoprotein fractions of cell nuclei. Biochim Biophys Acta 32: 301 –302. Zieve GW, Sauterer RA, Feeney RJ. 1988. Newly synthesized small nuclear RNAs appear transiently in the cytoplasm. J Mol Biol 199: 259– 267. Zorn C, Cremer C, Cremer T, Zimmer J. 1979. Unscheduled DNA synthesis after partial UV irradiation of the cell nucleus: distribution in interphase and metaphase. Exp Cell Res 124: 111 –119.

The Nuclear Envelope Martin W. Hetzer Salk Institute for Biological Studies, Molecular and Cell Biology Laboratory, La Jolla, California 92037 Correspondence: [email protected]

The nuclear envelope (NE) is a highly regulated membrane barrier that separates the nucleus from the cytoplasm in eukaryotic cells. It contains a large number of different proteins that have been implicated in chromatin organization and gene regulation. Although the nuclear membrane enables complex levels of gene expression, it also poses a challenge when it comes to cell division. To allow access of the mitotic spindle to chromatin, the nucleus of metazoans must completely disassemble during mitosis, generating the need to re-establish the nuclear compartment at the end of each cell division. Here, I summarize our current understanding of the dynamic remodeling of the NE during the cell cycle.

he NE, a hallmark of eukaryotic cells, is a highly organized double membrane that encloses the nuclear genome (Kite 1913). Early electron microscopy (EM) images revealed that the inner (INM) and outer nuclear membranes (ONM) are continuous with the endoplasmic reticulum (ER) (Watson 1955). Despite the lipid continuity between the NE and the ER, both ONM and INM are comprised of diverse groups of proteins that are typically not enriched in the ER (Hetzer et al. 2005) (Table 1). The first group consists of 30 different polypeptides, called nucleoporins or Nups, which form the 40 – 70 MD nuclear pore complexes (NPCs) (Tran and Wente 2006; D’Angelo and Hetzer 2008). NPCs are aqueous channels that show eightfold rotational symmetry with an outer diameter of 100 nm and a central transport channel measuring 40 nm in diameter, through which bidirectional exchange of proteins, RNA, and

T

ribonucleoprotein complexes between the nucleoplasm and cytoplasm occurs (Beck et al. 2004; Beck et al. 2007; Terry et al. 2007). A subset of Nups is stably embedded in the NE, forming a scaffold structure or NPC core (Rabut et al. 2004; D’Angelo et al. 2009), which is thought to stabilize the highly curved and energetically unfavorable pore membrane (Alber et al. 2007; Boehmer et al. 2008). This core includes the Nup107/160 complex (Nup84 complex in yeast) and the Nup205 complex (yeast Nup170), which together constitute 50% of the entire NPC (Fig. 1) (Brohawn et al. 2009). Attached to this scaffold are peripheral Nups, many of which contain phenylalanine-glycine (FG) rich repeats that establish a permeability barrier and also mediate active, receptor-dependent transport across the NE (Peters 2009). A second group of NE proteins, specifically localizes to the INM (Fig. 1) (Schirmer and Gerace 2005).

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000539 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000539

M.W. Hetzer Table 1. List of NE proteins. NPC

INM

ONM

Lamina

Nup35

LBR

Nup37

Lap1

Nup43

Lap2b

Nup50

Emerin

Lamin A Lamin B1 Lamin B2 Lamin C

Nup54 Nup58/45 Nup62 Nup75 Nup88 Nup96

MAN1 Nurim NET8 NET38 NET56 LEM2/ NET25 NET9 NET32 NET37 Sun1 Sun2 LUMA 60 NETs

Nesprin-3/ Net35 Syne/Myne/ Nesprin 1 Nesprin-2a and b Syne/ Nesprin-2G Samp1

Nup98 Nup107 Nup133 Nup153 Nup155 Nup160 Nup188 Nup205 Nup214 Nup358/ RanBP2 Sec13R Seh1 Pom121 Ndc1 Gp210 Tpr Rae1 Aladin Nlp1/hCG1

The nuclear pore complex (NPC) contains 30 nucleoporins (Nups). Nup98 and Nup96 are synthesized as a single polypeptide that becomes autoproteolytically cleaved to give rise to Nup98 and Nup96. Inner nuclear membrane (INM) proteins: lamin B receptor (LBR), lamin-associated protein (LAP), Nuclear Envelope Transmembrane protein (NET). More than 60 NETs have no assigned function. Outer nuclear membrane (ONM): spectrin repeat containing nuclear envelope protein (syne), spindle-associated membrane protein 1 (Samp1). Lamina: lamin A and C are products of alternative splicing.

Although most of these .60 integral membrane proteins (also referred to as NE transmembrane proteins or NETs [Schirmer et al. 2003]) remain largely uncharacterized, interaction with lamins (see later) and chromatin have been shown for some of them, such as lamin B receptor (LBR), lamina-associated polypeptide (LAP) 1, LAP2, emerin, and MAN1 (Akhtar and Gasser 2007; Dorner et al. 2007; Schirmer and Foisner 2007). It is becoming increasingly clear that INM proteins play vital and diverse roles in nuclear function such as chromatin organization, gene expression, and DNA metabolism (Mattout et al. 2006; Heessen and Fornerod 2007; Reddy et al. 2008). Importantly, improper localization and function of INM proteins have been linked to numerous human diseases, which has sparked considerable interest in NE biology over the last decade (Vlcek and Foisner 2007; Worman and Bonne 2007; Neilan 2009). A third class of NE proteins specifically resides in the ONM (Fig. 1). This diverse group of integral membrane proteins shares a small KASH (Klarsicht, ANC-1, Syne Homolgy) domain, which has been shown to interact with Sad1p/UNC-84 (SUN)-domain proteins of the inner nuclear membrane within the periplasmic space of the NE (Starr and Han 2003; Wilhelmsen et al. 2006). Two other related ONM proteins, nuclear envelope spectrin repeat (nesprin)-1 and -2, have been shown to directly interact with the actin cytoskeleton through their amino-terminal actin-binding domain (ABD) (Wilhelmsen et al. 2005). These ONM proteins are implicated in nuclear positioning that is essential for processes such as cell polarization, pronuclear migration, and the organization of syncitia (Fridkin et al. 2009). In addition, ONM and INM proteins form “bridges” across the perinuclear space that might be involved in separating the two NE membrane leaflets at an even distance of 50 nm (Voeltz and Prinz 2007). These lumenal proteinaceous bridges could establish physical connections between the cytoskeleton and chromatin,

Organization of the Nuclear Membrane

Cytoplasm

Actin, intermediate filaments

BAF

BAF

nesprin

nesprin

ONM

SUN

nurim ?

emerin

?

LEM2

HP1

LAP2

LBR

BAF

NETs

MAN1

ER lumen (PNS)

SUN

NPC

INM

Lamina (lamin A/C and B)

Chromatin Nucleoplasm

Figure 1. Topology of the NE. Inner and outer nuclear membranes (INM and ONM, respectively) are separated

by the ER lumen or perinuclear space (PNS). The nuclear lamina interacts with NE proteins and chromatin. INM proteins link the NE to chromatin and the lamina. ONM proteins provide a connection from the nucleus to the cytoskeleton. The lamin B receptor (LBR) interacts both with B-type lamins and chromatin-associated heterochromatin protein 1 (HP1) in conjunction with core histones. Members of the LEM (lamina-associated protein 2 [LAP2], emerin, MAN1)-domain family ( pink) bind to lamins and interact with chromatin through barrier-to-autointegration factor (BAF). SUN proteins (SUN 1 and 2) interact with nesprins in the ONM, thereby forming so-called LINC complexes that establish connections to actin and intermediate filaments in the cytoplasm. Nurim is a multi-pass membrane protein with unknown function. Proteomic approaches have identified 60 putative transmembrane proteins (NETs), most of which remain uncharacterized.

which might be relevant for transcription, replication, and DNA repair mechanisms (Tzur et al. 2006; Stewart et al. 2007). The final group of NE proteins constitutes the lamina, a meshwork of intermediate filaments that is composed of A- and B-type lamins (Gruenbaum et al. 2000). Although the lamina has been shown to be critical for nuclear stability, particularly in tissues that are exposed to mechanical forces such as muscle fibers (Cohen et al. 2008), it has become clear that lamins also play major roles in chromatin function and gene expression (Gruenbaum et al. 2005; Dechat et al. 2008; Reddy et al. 2008). Similar to INM proteins, mutations in lamins are linked to a large number of diverse human diseases (Mounkes et al. 2003; Muchir and Worman 2004; Mattout et al. 2006) and to aging (Gruenbaum et al.

2005; Scaffidi and Misteli 2006), highlighting the crucial role of the NE protein network for normal cell function. In summary, the NE fulfills a critical role in shielding the genome from cytoplasmic components, but also represents a highly specialized membrane that provides anchoring sites for chromatin and the cytoskeleton (D’Angelo and Hetzer 2006). NE REMODELING IN DIVIDING CELLS

Although a membrane-enclosed nuclear genome can be found in all eukaryotes, there is a critical difference in the cell-cycle dependent dynamics of the NE between “lower” eukaryotes (e.g., yeast and filamentous fungi) and metazoa (i.e., “higher” eukaryotes). The former undergo

M.W. Hetzer

closed mitosis, where spindle microtubules can either form inside the nucleus or are able to penetrate an intact nuclear membrane (Heywood 1978; Byers 1981; Ribeiro et al. 2002). In contrast, the NE of metazoan cells completely disintegrates during cell division to allow the mitotic spindle to access chromosomes (Kutay and Hetzer 2008). As a consequence, every dividing cell has to reform the NE and re-establish the identity of the nuclear compartment (Hetzer et al. 2005; Anderson and Hetzer 2008b; Guttinger et al. 2009). NE remodeling in proliferating cells is a highly dynamic process that involves a vast number of molecular players (Figs. 2,3). By the end of interphase in G2, the nuclei have duplicated their genome, doubled the number of NPCs, and increased the surface area of the NE. The surrounding ER network is continuous with the NE, but not enriched in NE proteins. The entry of mitosis, i.e., prophase, is marked by NE breakdown (NEBD) and the loss of the nucleo-cytoplasmic compartmentalization (Burke and Ellenberg 2002). Between NEBD

G2 ER sheets

and early anaphase, when chromosomes align in the metaphase plate and subsequently segregate, chromatin is essentially free of membranes (Puhka et al. 2007; Anderson and Hetzer 2008a). During these cell-cycle stages, the majority of soluble NE proteins are distributed throughout the cytoplasm and transmembrane NE proteins reside in the mitotic ER (Ellenberg et al. 1997; Anderson and Hetzer 2007; Puhka et al. 2007) (Fig 2). In anaphase, ER membranes begin to reassociate with and rapidly enclose the chromatin mass (Anderson and Hetzer 2008a). Chromatin association of a subset of Nups, decondensation of chromatin, and the assembly of new NPCs occurs concomitantly with NE formation (Anderson and Hetzer 2008c). At the end of cell division, the NE has reformed as a closed membrane barrier and re-establishes the nuclear compartment by enabling selective nucleo-cytoplasmic transport (Dultz et al. 2008). After its formation, the NE expands and undergoes additional structural changes necessary for cell-cycle progression and transcription, including the assembly of new

NEBD in prophase

ER tubules

Metaphase

Dispersal of NE components into ER

Mitotic spindle NE/ER membranes

Chromatin

Chromatin

NPC

Chromatin

Intact NE with NPCs

Disassembled NPCs

Nups at kinetochores

Figure 2. Schematic illustration of NE breakdown. In G2, the cell nucleus has completed DNA replication and

NPC duplication. The NE (dark green), which is continuous with the ER network (green), encloses the chromosomes (blue). NPCs (red/blue) mediate nuclear transport. When cells enter mitosis, NPCs disassemble and the NE gets reabsorbed into the ER, which at this stage is composed of tubules. NPC components are dispersed into the cytoplasm and NE proteins are partitioned into the ER (dashed green lines). Centrosomes (orange dots) move to the NE and microtubules ( purple) participate in the rupturing of the NE. In metaphase, a subset of NPC components has associated with kinetochores and the spindle is established. At this stage, chromosomes at the metaphase plate are devoid of membranes.

Organization of the Nuclear Membrane Anaphase Re-attaching NE/ER membranes

Telophase NE flattening and sealing

G1 NE expansion

Chromatin Chromatin

Chromatin

Chromatin-bound Nups

Assembling NPCs

Nuclear transport

Figure 3. Schematic illustration of NE reformation around segregated chromosomes in one of the two daughter

cells. In anaphase, ER membranes associate with chromatin (red arrows) and a subset of nucleoporins associates with chromatin. Additional membrane tubules and NE proteins are recruited to the chromatin surface and mediate NE flattening. At this stage, most NE proteins are cleared from the ER network. In late anaphase/ early telophase, a closed NE has formed with pores being assembled in a step-wise manner. Once pores become transport competent, the NE expands and cells move into G1.

NPCs (Winey et al. 1997; D’Angelo et al. 2006) (Fig. 3).

BREAKING DOWN THE NUCLEUS

Live cell imaging of mammalian cells has revealed that NEBD, which occurs rapidly within a few minutes in prophase (Lenart et al. 2003), occurs in a step-wise fashion. One of the initial events is the selective loss of Nups from the NPCs (Terasaki et al. 2001; Dultz et al. 2008; Katsani et al. 2008). Nup98, a dynamic NPC component (Griffis et al. 2002), is released before the rest of the NPC components, which disassemble synchronously (Dultz et al. 2008). Collective dispersal of Nups can also be observed in Drosophila, suggesting that pore disassembly is evolutionarily conserved (Katsani et al. 2008). Biochemical evidence suggests that the NPC disassembles into stable subcomplexes and not necessarily into individual polypeptides (Miller and Forbes 2000; Harel et al. 2003b; Walther et al. 2003a), which might stabilize the nucleoporins and allow more rapid reformation of NPCs at the end of mitosis.

Unfortunately, we do not have a clear picture of the biochemical nature of the disassembled NPC components. Recent evidence suggests that some nuclear pore components are degraded in a proteasome-dependent manner during mitosis, a process that might regulate total pore numbers. Furthermore, in the same study, Nup43 has been shown to be absent from the mitotic Nup107/160 complex, which therefore seems to differ from its counterpart in interphase (Chakraborty et al. 2008).

THE ROLE OF KINASES IN NE BREAKDOWN

NPC disassembly is critical for early stages of NEBD, such as disrupting the permeability barrier and allowing the influx of molecules, which are thought to be critical for further disassembly steps. For example, protein kinases like cyclindependent kinase 1(Cdk1/CycB1) may require access to proteins of the INM and lamins for proper mitotic regulation (Wu et al. 1998). That is consistent with the idea that the bulk of lamin disassembly occurs after the NPCs are permeabilized (Lee et al. 2000; Beaudouin

M.W. Hetzer

et al. 2002). As the lamina is disrupted, many INM proteins lose their anchor and are released into the ER (Ellenberg et al. 1997; Yang et al. 1997; Puhka et al. 2007; Anderson et al. 2009a; Lu et al. 2009). One of the critical events during NEBD is the hyperphosphorylation of NE proteins that is thought to disrupt protein complexes and/ or play a role in the activation of factors involved in this process. Several kinases, such as Cdk1/ CycB1, protein kinase C (PKC), Nima, and Aurora A, have been implicated in NEBD (Collas 1999; Burke and Ellenberg 2002; Gong et al. 2007; Portier et al. 2007). The key kinase seems to be Cdk1, which phosphorylates lamins and Nups, including the Nup107/160 complex, Nup93 subcomplex, Nup53, Nup98, Ndc1, and gp210 (Guttinger et al. 2009). The disassembly of INM proteins, such as Lap2a and b and LBR, seem to depend on Cdk1 (Courvalin et al. 1992; Dechat et al. 1998; Dreger et al. 1999). Several other kinases have been shown to participate in NEBD including PKC (Goss et al. 1994), Aurora A (Portier et al. 2007), and polo-like kinase (PLK1) (Chase et al. 2000); however, their targets remain to be determined. Lamin B is the only known target of PKC during NEBD (Goss et al. 1994). In this context, it is worth mentioning that NPCs in organisms with closed mitosis are also partially disassembled. For instance, in Aspergillus nidulans, 14 Nups are released, leaving a partial NPC that allows the entry of the Cdk1/ cyclin B complex (Osmani et al. 2006). This partial disassembly requires the kinases Nima and Cdk1 (De Souza et al. 2004). Thus, structural reorganization of the NPC during mitosis might be an evolutionarily conserved feature shared by all eukaryotes. MEMBRANE DYNAMICS DURING NEBD

An interesting aspect of NEBD was recently revealed by the finding that the nuclear membranes are actively participating in the process. Work in the nematode Caenorhabditis elegans has shown that depletion of reticulons and the GTPase Rab5 inhibits NEBD (Audhya et al. 2007). The reticulons are part of a family

of membrane-associated proteins that together with a distinct class of membrane-bending proteins like DP1 have been shown to tubulate the ER (Voeltz et al. 2006). These results suggest that active reshaping of the ER might be a critical step in NEBD. Two nucleoporins have also been implicated in this process; however, their roles remain unclear. The transmembrane nucleoporin gp210 is enriched in its phosphorylated form at the NE just before NEBD occurs (Galy et al. 2008). The mechanism by which gp210 contributes to the disassembly of NPCs and the lamina remains to be characterized. Depletion of Nup153, a dynamic component of the nuclear basket, inhibits NEBD and was shown to recruit the COPI complex, which mediates retrograde transport from the Golgi to the ER, to the NE (Liu et al. 2003). What roles Nup153 and the COPI complex play in this process has yet to be elucidated. New data suggests that lipid synthesis itself might play a role in NE dynamics. The knock down of lipin, a conserved phosphatidic acid (PA) phosphatase that catalyzesthe dephosphorylation of PA to yield diacylglycerol (DAG) (Siniossoglou 2009), causes defects in NEBD, abnormal chromosome segregation, and irregular nuclear morphology in C. elegans. Interestingly, down-regulation of nematode Lpin-1 was required for disassembly of the nuclear lamina during late NEBD, suggesting that the Lpin-1 requirement appears to be separable from the effect of Lpin-1 on the peripheral ER (Golden et al. 2009; Gorjanacz and Mattaj 2009). Although phosphorylation is clearly a key event in NEBD, interactions of the NE with microtubules are thought to generate mechanical forces that assist in rupturing the nuclear membrane and presumably the lamina in a dynein-mediated process (Beaudouin et al. 2002; Salina et al. 2002; Muhlhausser and Kutay 2007). This process might also involve the small GTPase Ran, which has been shown to regulate microtubule dynamics during NEBD, potentially uncovering yet another potential mitotic function of Ran (Muhlhausser and Kutay 2007). However, it should be kept in mind that nuclear disassembly can also occur in the absence of microtubules in vitro (Lenart

Organization of the Nuclear Membrane

et al. 2003), indicating that dynein-dependent rupturing of the NE may not be essential. STRUCTURE AND FUNCTION OF NE COMPONENTS DURING MITOSIS

As mentioned above, chromosomes are essentially membrane-free between metaphase and anaphase. EM and fluorescence microscopy has provided compelling evidence that the NE is reabsorbed into the ER (Ellenberg et al. 1997; Anderson and Hetzer 2007; Anderson and Hetzer 2008a; Anderson et al. 2009a; Lu et al. 2009). For instance, fluorescence timelapse microscopy in living cells has shown that NE proteins such as the nucleoporins gp210, Ndc1, and Pom121 are partitioned into the mitotic ER (Ellenberg et al. 1997; Puhka et al. 2007). Likewise, INM proteins can be found in the ER during mitosis (Puhka et al. 2007). The topology of the mitotic ER, however, remains controversial. Three-dimensional modeling of the ER by electron tomography suggested that the mitotic ER remains an intact network of membrane tubules that is essentially free of sheets (Puhka et al. 2007). This is supported by recent data suggesting that the intrinsic propensity of the ER to oscillate between tubules and sheets is used during mitosis and affects the fate of the NE during the mitosis of C. elegans (Audhya et al. 2007). However, compelling data from nematodes and mammalian cells challenges the view of an entirely tubular ER and instead suggest that the ER is largely composed of membrane sheets. For instance, it was shown that the rough ER of metaphase HeLa cells appears to be exclusively cisternal and concentrates at the cell cortex, often following the contours of the plasma membrane (McCullough and Lucocq 2005). In a study using three-dimensional (3D) reconstructions of living cells, the ER appeared entirely formed of cisternae and very few tubules were observed (Lu et al. 2009). Similar results were obtained in C. elegans (Poteryaev et al. 2005). Determining the exact nature of the ER and the disassembled NE components remain critical future goals, because this has important implications for NE formation.

NE PROTEINS HAVE MITOTIC FUNCTIONS

Although the dynamic organization of the ER membranes remains elusive, it has become clear that disassembled NE components fulfill critical functions in mitosis. For instance, the multimeric Nup107-160 complex, which in interphase is essential for pore assembly and function (D’Angelo et al. 2006), and Nup358, which shows SUMO E3 ligase activity that modifies RanGAP, Ran’s GTPase activating protein (Joseph et al. 2002; Pichler et al. 2002), can both be detected in association with kinetochores in mitosis. In addition, ELYS/MEL-28 and Nup107-160 are also found with spindle poles and proximal microtubules (Loiodice et al. 2004; Galy et al. 2006; Rasala et al. 2006). This association is not critical in Drosophila, suggesting that the role of Nups in kinetochore function is a relatively late evolutionary event (Katsani et al. 2008). However, in mammalian cells, the absence of the Nup107-160 complex perturbs bipolar spindle formation (Orjalo et al. 2006), and inhibits Nup358 recruitment to microtubule-bound kinetochores (Salina et al. 2003; Joseph et al. 2004). Recently it was shown that the kinesin-binding domain (KBD) of Nup358 associates with kinesin-1, KIF5B/ KIF5C. Interestingly, the KBD stimulates the ATPase activity of KIF5B in the presence of microtubules, thereby increasing its activity (Cho et al. 2009). The Nup107-160 complex might have an independent role at kinetochores because in a Xenopus in vitro spindle assembly assay, the Nup358/SUMO-RanGAP complex does not associate with kinetochores (Arnaoutov and Dasso 2005). Because mNup133 (a mNup107160 member) and CENP-F (a kinetochore protein) interact with each other, this function might be linked to the dynein partners Nde1 and Nde1l, which also bind CENP-F (Vergnolle and Taylor 2007; Zuccolo et al. 2007). Additional roles for other NE proteins in cell-cycle and mitotic progression have also been reported. Depletion of lamin B, a type V intermediate-filament protein and a component of the nuclear lamina, also results in mitotic spindle defects (Zheng and Tsai 2006).

M.W. Hetzer

Interestingly, the spindle-associated lamin B appears to be present in a membranous, matrixlike network and seems to facilitate spindle microtubule organization in a dynein-dependent manner (Tsai et al. 2006; Ma et al. 2009). Although the mechanistic details of spindlematrix function with respect to lamin B remain to be determined, these results contribute to the emerging paradigm for structural components of the NE having roles in mitosis. This idea is further supported by the recent finding of spindle-associated membrane protein 1 (Samp1), which during interphase is localized to the inner nuclear membrane, specifically localized to the polar regions of the mitotic spindle (Buch et al. 2009). Depletion of Samp1 expression resulted in separation of centrosomes from the NE, indicating that it is functionally connected to the cytoskeleton. This provides evidence for the existence of interactions between mitotic ER membranes with the spindle. RE-ESTABLISHING ORDER: REFORMATION OF THE NE

NE formation has been studied in various cellfree systems, cell types, and organisms (Burke and Ellenberg 2002; Hetzer et al. 2005; Anderson and Hetzer 2008c). Because membranes are typically delicate structures that are easily disrupted during cell fractionation and fixation, it is easy to see how data obtained from different systems lead to the postulation of sometimes opposing models. There are essentially two different ideas about how the NE is formed: (1) by vesicle fusion and (2) by reshaping of ER into NE sheets. Based on biochemical data and EM observations, it was initially proposed that the NE fragments into NE vesicles (Collas and Courvalin 2000). This idea is derived from cell-free systems mainly of Xenopus, starfish, and Drosophila (Lohka and Masui 1983; Newport 1987). Frog egg extracts have the advantage of containing large stockpiles of disassembled NE components (Newport and Spann 1987) and nuclei can be assembled within 60 minutes in a test tube by mixing a chromatin source, cytosol, and membranes. These artificial nuclei are

capable of nucleocytoplasmic transport, DNA replication, as well as NEBD (Sheehan et al. 1988; Gant and Wilson 1997; Hetzer et al. 2002). The idea of vesicle fusion as the principal mechanism of NE formation was further supported by findings that GTPgS, a nonhydrolyzable GTP analog, blocks NE formation and results in assembly intermediates covered with chromatin-bound vesicles (Boman et al. 1992; Hetzer et al. 2001). Interestingly, depletion of the small GTPase Ran mimicked the effect of GTPgS addition (Hetzer et al. 2000; Zhang and Clarke 2000). Because the Ran exchange factor RCC1 (Ohtsubo et al. 1989) is stably associated with chromatin, it was proposed that high levels of RanGTP are generated around chromatin that provide a spatial signal for chromatin-associated processes such as NE formation and NPC assembly (Bilbao-Cortes et al. 2002; Hetzer et al. 2002; Hutchins et al. 2004). It was subsequently shown that Ran releases the nuclear transport receptor Importin b (Nachury et al. 2001; Harel et al. 2003a; Harel and Forbes 2004) from nucleoporins and thereby triggers NPC assembly (Harel et al. 2003a; Walther et al. 2003b). A similar mechanism might be required for the formation of a closed NE, but the targets of Importin b remain unknown. Corroborating the in vitro data, Ran has been shown to be required for NE formation in C. elegans (Askjaer et al. 2002) and yeast (Ryan et al. 2003). Although egg extracts provide a unique experimental system to study ER and NE reconstitution, it is important to realize that the assembly reactions are initiated with isolated membranes, which are in a highly fragmented state that does not represent the intact in vivo organization of the mitotic ER. Recent evidence shows that an intact NE can also form from preformed ER in vitro (Anderson and Hetzer 2007). Strikingly, nuclear assembly from a preformed ER is insensitive to fusion inhibitors such as GTPgS, ATPgS, or antibodies that inhibit the function of the AAA-ATPase p97. In addition, factors that are essential for ER tubule formation, such as ATP (Dreier and Rapoport 2000) and reticulons (Voeltz et al. 2006), only block nuclear assembly if added

Organization of the Nuclear Membrane

before the ER network is organized (Anderson and Hetzer 2007). The results strongly support the notion that the ER is the source of NE membranes. According to this idea, the observed inhibition of nuclear assembly by fusion inhibitors and the role of SNARE proteins in NE formation are likely to be an indirect effect of blocked ER reconstitution (Baur et al. 2007). Although these experiments show that the NE can form from an ER network in vitro, they do not address the question of how NE formation occurs in intact cells. High resolution imaging in intact cells has revealed ER tubules contacting chromatin at early stages of NE formation. Shortly after contact, the rapid coating of chromatin by ER membranes can be observed (Anderson and Hetzer 2007). Recent data suggest that endogenous concentrations of NE-promoting transmembrane proteins are ratelimiting for nuclear assembly. Because NE formation is also affected by endogenous levels of the ER-shaping reticulon proteins that slow NE formation, these findings suggest a tug of war between reticulons and their membrane-curving activity and NE proteins, which promote membrane attachment and spreading around chromatin (Anderson and Hetzer 2008a). The massive membrane-restructuring event that results in the formation of the sheet-like NE involves functionally diverse groups of NE proteins that collaborate during mitosis to tether membranes to the chromatin surface and thereby drive NE formation (Anderson et al. 2009b). Recent findings that DNA-binding activity of some INM proteins is required for NE formation in vitro (Ulbert et al. 2006) and that membrane sheets formed efficiently on protein-free immobilized DNA in vitro support this idea. In addition, NE formation also seems to involve the ability of several INM proteins to bind to chromatin factors. For example, it has been shown that the integral INM protein LBR, which binds to heterochromatin-binding protein 1 (HP1), is required for targeting and anchoring NE membranes to chromatin in vitro (Collas et al. 1996; Pyrpasopoulou et al. 1996). In a similar fashion, the barrier-to-autointegration factor (BAF), a chromatin-binding protein (Segura-Totten et al. 2002) and its kinase Vrk have recently been shown to play a direct role in

NE formation by recruiting LEM domain proteins to chromatin (Gorjanacz et al. 2007). Vrk seems to be a key regulator in this process because BAF phosphorylation reduces chromatin binding and interactions with LEM domain proteins such as emerin (Bengtsson and Wilson 2004; Hirano et al. 2005). Thus, NE formation is likely to involve a complex interplay of transmembrane NE proteins distributed into mitotic ER with the reorganizing chromatin. ROLE OF CHROMATIN IN NE FORMATION

Recent studies of cell-free nuclear assembly systems suggest that initial ER/chromatin contacts are mediated via tubule ends. In a second step, these tethered ER tubules are reorganized into flat nuclear membrane sheets by DNA-binding-NE-specific membrane proteins (Ulbert et al. 2006; Anderson and Hetzer 2007). In cells, the ER appears to be largely cisternal (Lu et al. 2009) and thus it is likely that pre-existing membrane sheets are directly tethered to the chromatin surface in vivo. This is consistent with the finding that overexpression of reticulons delays NE formation, whereas their knock down accelerates NE formation (Anderson and Hetzer 2008a). During NE formation, chromatin undergoes a series of conformational changes, from a state of maximal chromatin compaction in late anaphase, right before NE formation (MoraBermudez et al. 2007), to transcription- and replication-competent decondensation in interphase. Anaphase compaction requires the kinesin-like DNA binding protein (Kid), which ensures the formation of a compact chromosome cluster during anaphase and the properenclosure of the segregated chromatin mass into a single nucleus (Ohsugi et al. 2008). Interestingly, Kid loading onto anaphase chromosomes is dependent on Importin b (Tahara et al. 2008), adding to the growing number of mitotic processes regulated by this transport receptor. Recently, a compelling case has been made that mechanistically links NE formation and chromatin decondensation. The hexameric ATPase Cdc48/p97, which was implicated in membrane fusion (Kondo et al. 1997) and

M.W. Hetzer

ubiquitin-dependent processes (Ye et al. 2003), extracts Aurora B from chromatin, which results in its inactivation and subsequent chromatin decondensation. Interestingly, the inhibition of Cdc48/p97 blocked NE formation, suggesting that chromatin decondensation is required for NE formation (Ramadan et al. 2007), possibly by opening chromatin structure. It is tempting to speculate that this open chromatin structure provides binding sites for INM proteins and therefore drives NE formation. Concomitantly, with the coating of chromatin by the NE, NPCs assemble in the reforming nuclei. This fascinating example of protein selfassembly is coordinated by the stepwise recruitment of a subset of NPC proteins to chromatin (Dultz et al. 2008) and some progress has been made in determining how this process occurs. A protein called Mel-28/ELYS, which was identified in a screen in C. elegans for factors involved in pronuclear formation, is critical for the association of the Nup107/160 complex to chromatin (Rasala et al. 2006; Franz et al. 2007). Because Mel-28/ELYS contains an AT-hook domain (Kimura et al. 2002), a likely scenario is that this step occurs by direct binding to DNA. Interestingly, RanGTP stimulates Mel-28/ELYS recruitment (Franz et al. 2007), presumably by releasing Importin b from one of the Nup107/ 160/ELys components (Hetzer et al. 2005). Although other nucleoporins have been implicated in pore assembly, their exact role remains to be determined. For instance, a complex of Nup53 and Nup155 has recently been shown to be essential for NE formation in nematodes and vertebrates (Franz et al. 2005; HawrylukGara et al. 2008); however, how the membraneassociated Nup53 coordinates interactions between chromatin, membranes, and soluble Nup155 remains unclear. A group of nucleoporins have been shown to be essential for pore assembly (Hetzer et al. 2005), the largest subcomplex being the Nup107/160 complex, whose depletion results in NPC-free nuclear membranes (Harel et al. 2003b; Walther et al. 2003a). In vertebrates, two transmembrane nucleoporins have also been shown to participate in NE formation, Ndc1 and POM121 (Mansfeld et al. 2006; Stavru

et al. 2006; Anderson et al. 2009b). The third known transmembrane Nup, gp210, is not expressed in all cell types and thus is unlikely to be essential for pore assembly (Eriksson et al. 2004). Whereas the molecular role of these scaffold Nups is unclear, an interesting link between POM121 and the Nup107/160 complex has been made and nuclear membrane formation might actually be linked to pore assembly by a poorly understood checkpoint by which the Nup107/160 complex “senses” nuclear membranes (Antonin et al. 2005). REMODELING THE INTERPHASE NE

In order for cells to properly progress through multiple cell divisions, the number of NPCs doubles in interphase (Maul et al. 1972). It is unclear if the increase simply reflects the necessity to double pores for daughter cells of the next division cycle, or whether an increase in NPC number is important for interphase cellcycle progression, e.g., efficient replication or transcription. Interphase nuclear pore assembly is particularly interesting because NPC formation occurs from both sides of the NE (D’Angelo et al. 2006). Thus, the question arises, what is the mechanism of communication between the double membranes? Interestingly, Nup133, a member of the Nup107/160 complex, contains an ALPS-like motif including an amphipathic a-helical domain that has been shown to act as a membrane curvature sensor in vitro (Drin et al. 2007). It is possible that this domain is involved in targeting the Nup107/160 complex to membranes during NE formation. A prediction from this idea is that the membrane hole is formed before Nup107/160 is recruited. Whereas the fusion of INM and ONM remains elusive in mammalian cells, significant progress has been made in deciphering this process in yeast. Genetic studies in S. cerevisiae revealed a network of protein– protein interactions that appears to initiate NPC formation. The nucleoporins Nup59/53 and the integral pore membrane nucleoporins Pom152 and Pom34 tether Nup170 and a third integral membrane nucleoporin Ndc1 to the NE at new assembly sites (Onischenko et al. 2009). Furthermore, depletion

Organization of the Nuclear Membrane

of Nup170 and its homolog Nup157 causes the accumulation of NPC-like structures in the INM and at cytoplasmic foci rather than properly localized to nuclear pores spanning the NE. It is interesting to note that the yeast reticulons and Yop1 not only display genetic interactions with the Poms, but also seem to play an essential role in the formation of new NPCs (Dawson et al. 2009). Similar to the depletion of Nup170, in the absence of reticulons, NPClike intermediates also accumulate in an aberrant manner in the INM and ONM of the NE. The current idea is that a complex consisting of the three membrane-spanning nucleoporins Pom152, Pom34, and Ndc1 marks a new assembly site by joining or juxtaposing ONM and INM, which is consistent with the observation that NPC assembly proceeds from both sides of the NE (D’Angelo et al. 2006). According to this idea, Nup53/59 and the reticulons Yop1/ Dbp1 are recruited to the nuclear membrane and inserted into the outer leaflet of the membrane using monotopic membrane insertion domains, inducing or stabilizing local curvature to the membrane. However, this idea was recently challenged on theoretical grounds, arguing that reticulons induce positive membrane curvature (e.g., giving rise to the convex surface of a tubule), but not negative curvature, which is likely to occur during NPC assembly. Instead, it was proposed that reticulons might participate in the local enrichment of negative curvatureinducing proteins involved in NE formation (Antonin 2009). Whether transmembrane nucleoporins, reticulons, oran unidentified machinery induces the fusion of INM and ONM during NPC assembly remains an open question. During interphase, the NE expands considerably and thus requires the supply of additional nuclear membrane and proteins. In vitro nuclear expansion is blocked by disrupting the connection of nuclei with the peripheral ER (D’Angelo et al. 2006), suggesting that membranes feed into the ONM via connections with ER tubules. Growth of the INM requires passage of membrane components through the fusion sites with the ONM at the NPCs. The current view is that INM proteins are retained once they reach the INM by interactions

with either the lamina or chromatin. How INM proteins are targeted in metazoa is less clear. One model suggested that ATP-driven changes in nucleoporin interactions might allow membrane proteins to travel across the NPC (Ohba et al. 2004). In yeast, integral INM proteins have been shown to directly interact with specific nucleoporins and transport receptors to promote their movement past the NPC (King et al. 2006). By the end of interphase, the NE has undergone major changes in protein composition and is ready to break down at the onset of mitosis to start a new life cycle in the two daughter cells. During interphase, NE connections with the cytoskeleton have to withstand external forces. Recent work has suggested that SUN and KASH domain proteins play key roles in this process (Adam 2001). KASH domain proteins interact with centrosomes/SPBs or cytoskeletal elements in the cytoplasm and bind to SUN domain proteins in the INM to connect the NE to structures on the outside of the nucleus. The nuclear lamina could serve in metazoa to distribute the forces across a broader area (Stewart et al. 2007) (Roux and Burke 2007). CONCLUDING REMARKS

The NE is well known for its role in shielding the nuclear genome from cytoplasmic components and mediating nucleocytoplasmic transport. Less well understood is its dynamic behavior in dividing cells and its role in the distribution of the genetic material. Furthermore, it has become clear that the NE might actively control the organization of the genome and directly regulate gene expression. This opens some interesting opportunities for future research activities. For instance, the nature of most NE proteinchromatin interactions remain uncharacterized and many of the INM proteins are expressed in a tissue-specific manner. This raises the important question of whether NE composition is a critical component of cell fate determination. If NE proteins control chromatin organization, then is there a link between NE formation and nuclear organization? In other words, are the NE-chromatin contact points that are established

M.W. Hetzer

in anaphase/telophase maintained in interphase? Because many NE proteins are linked to a large number of diseases, answers to these questions are likely to have a direct impact on human health.

ACKNOWLEDGMENTS

I would like to thank members in my laboratory, in particular, Maximiliano D’Angelo, Roberta Schulte, and Jesse Vargas for critically reading the manuscript. This work was supported by the National Institute of General Medical Sciences (award number R01GM073994).

REFERENCES Adam SA. 2001. The nuclear pore complex. Genome Biol 2: REVIEWS0007. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007. The molecular architecture of the nuclear pore complex. Nature 450: 695 –701. Anderson DJ, Hetzer MW. 2007. Nuclear envelope formation by chromatin-mediated reorganization of the endoplasmic reticulum. Nat Cell Biol 9: 1160–1166. Anderson DJ, Hetzer MW. 2008a. Reshaping of the endoplasmic reticulum limits the rate for nuclear envelope formation. J Cell Biol 182: 911 –924. Anderson DJ, Hetzer MW. 2008b. Shaping the endoplasmic reticulum into the nuclear envelope. J Cell Sci 121: 137–142. Anderson DJ, Hetzer MW. 2008c. The life cycle of the metazoan nuclear envelope. Curr Opin Cell Biol 20: 386 –392. Anderson DJ, Vargas J, Hisao J, Hetzer MW. 2009a. Recruitment of functionally distinct membrane proteins to chromatin mediates nuclear envelope formation in vivo. J Cell Biol 186: 183 –191. Anderson DJ, Vargas JD, Hsiao JP, Hetzer MW. 2009b. Recruitment of functionally distinct membrane proteins to chromatin mediates nuclear envelope formation in vivo. J Cell Biol 186: 183 –191. Antonin W. 2009. Nuclear envelope: Membrane bending for pore formation? Curr Biol 19: R410–412. Antonin W, Franz C, Haselmann U, Antony C, Mattaj IW. 2005. The integral membrane nucleoporin pom121 functionally links nuclear pore complex assembly and nuclear envelope formation. Mol Cell 17: 83–92. Arnaoutov A, Dasso M. 2005. Ran-GTP regulates kinetochore attachment in somatic cells. Cell Cycle 4: 1161–1165. Askjaer P, Galy V, Hannak E, Mattaj IW. 2002. Ran GTPase cycle and importins a and b are essential for spindle formation and nuclear envelope assembly in living Caenorhabditis elegans embryos. Mol Biol Cell 13: 4355–4370.

Audhya A, Desai A, Oegema K. 2007. A role for Rab5 in structuring the endoplasmic reticulum. J Cell Biol 178: 43– 56. Baur T, Ramadan K, Schlundt A, Kartenbeck J, Meyer HH. 2007. NSF- and SNARE-mediated membrane fusion is required for nuclear envelope formation and completion of nuclear pore complex assembly in Xenopus laevis egg extracts. J Cell Sci 120: 2895– 2903. Beaudouin J, Gerlich D, Daigle N, Eils R, Ellenberg J. 2002. Nuclear envelope breakdown proceeds by microtubuleinduced tearing of the lamina. Cell 108: 83– 96. Beck M, Forster F, Ecke M, Plitzko JM, Melchior F, Gerisch G, Baumeister W, Medalia O. 2004. Nuclear pore complex structure and dynamics revealed by cryoelectron tomography. Science 306: 1387–1390. Beck M, Lucic V, Forster F, Baumeister W, Medalia O. 2007. Snapshots of nuclear pore complexes in action captured by cryo-electron tomography. Nature 449: 611– 615. Bengtsson L, Wilson KL. 2004. Multiple and surprising new functions for emerin, a nuclear membrane protein. Curr Opin Cell Biol 16: 73–79. Bilbao-Cortes D, Hetzer M, Langst G, Becker PB, Mattaj IW. 2002. Ran binds to chromatin by two distinct mechanisms. Curr Biol 12: 1151– 1156. Boehmer T, Jeudy S, Berke IC, Schwartz TU. 2008. Structural and functional studies of Nup107/Nup133 interaction and its implications for the architecture of the nuclear pore complex. Mol Cell 30: 721 –731. Boman AL, Delannoy MR, Wilson KL. 1992. GTP hydrolysis is required for vesicle fusion during nuclear envelope assembly in vitro. J Cell Biol 116: 281 –294. Brohawn SG, Partridge JR, Whittle JR, Schwartz TU. 2009. The nuclear pore complex has entered the atomic age. Structure 17: 1156–1168. Buch C, Lindberg R, Figueroa R, Gudise S, Onischenko E, Hallberg E. 2009. An integral protein of the inner nuclear membrane localizes to the mitotic spindle in mammalian cells. J Cell Sci 122: 2100– 2107. Burke B, Ellenberg J. 2002. Remodelling the walls of the nucleus. Nat Rev Mol Cell Biol 3: 487– 497. Byers B. 1981. Cytology of the yeast life cycle. In Molecular Biology of the Yeast Saccharomyces. In Life Cycle and Inheritance (ed. Strathern E.W.J.a.J.R.B. J. N. Strathern), pp. 59– 96. Cold Spring Harbor Laboratory Press, NY. Chakraborty P, Wang Y, Wei JH, van Deursen J, Yu H, Malureanu L, Dasso M, Forbes DJ, Levy DE, Seemann J, et al. 2008. Nucleoporin levels regulate cell cycle progression and phase-specific gene expression. Dev Cell 15: 657– 667. Chase D, Serafinas C, Ashcroft N, Kosinski M, Longo D, Ferris DK, Golden A. 2000. The polo-like kinase PLK-1 is required for nuclear envelope breakdown and the completion of meiosis in Caenorhabditis elegans. Genesis 26: 26– 41. Cho KI, Yi H, Desai R, Hand AR, Haas AL, Ferreira PA. 2009. RANBP2 is an allosteric activator of the conventional kinesin-1 motor protein, KIF5B, in a minimal cell-free system. EMBO Rep 10: 480 –486. Cohen TV, Hernandez L, Stewart CL. 2008. Functions of the nuclear envelope and lamina in development and disease. Biochem Soc Trans 36: 1329–1334.

Organization of the Nuclear Membrane Collas P. 1999. Sequential PKC- and Cdc2-mediated phosphorylation events elicit zebrafish nuclear envelope disassembly. J Cell Sci 112: 977– 987. Collas I, Courvalin JC. 2000. Sorting nuclear membrane proteins at mitosis. Trends Cell Biol 10: 5 –8. Collas P, Courvalin JC, Poccia D. 1996. Targeting of membranes to sea urchin sperm chromatin is mediated by a lamin B receptor-like integral membrane protein. J Cell Biol 135: 1715– 1725. Courvalin JC, Segil N, Blobel G, Worman HJ. 1992. The lamin B receptor of the inner nuclear membrane undergoes mitosis-specific phosphorylation and is a substrate for p34cdc2-type protein kinase. J Biol Chem 267: 19035–19038. D’Angelo MA, Hetzer MW. 2006. The role of the nuclear envelope in cellular organization. Cell Mol Life Sci 63: 316–332. D’Angelo MA, Hetzer MW. 2008. Structure, dynamics and function of nuclear pore complexes. Trends Cell Biol 18: 456–466. D’Angelo MA, Anderson DJ, Richard E, Hetzer MW. 2006. Nuclear pores form de novo from both sides of the nuclear envelope. Science 312: 440–443. D’Angelo MA, Raices M, Panowski SH, Hetzer MW. 2009. Age-dependent deterioration of nuclear pore complexes causes a loss of nuclear integrity in postmitotic cells. Cell 136: 284– 295. Dawson TR, Lazarus MD, Hetzer MW, Wente SR. 2009. ER membrane-bending proteins are necessary for de novo nuclear pore formation. J Cell Biol 184: 659–675. De Souza CP, Osmani AH, Hashmi SB, Osmani SA. 2004. Partial nuclear pore complex disassembly during closed mitosis in Aspergillus nidulans. Curr Biol 14: 1973– 1984. Dechat T, Gotzmann J, Stockinger A, Harris CA, Talle MA, Siekierka JJ, Foisner R. 1998. Detergent-salt resistance of LAP2a in interphase nuclei and phosphorylationdependent association with chromosomes early in nuclear assembly implies functions in nuclear structure dynamics. EMBO J 17: 4887– 4902. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832 –853. Dorner D, Gotzmann J, Foisner R. 2007. Nucleoplasmic lamins and their interaction partners, LAP2a, Rb, and BAF, in transcriptional regulation. FEBS J 274: 1362– 1373. Dreier L, Rapoport TA. 2000. In vitro formation of the endoplasmic reticulum occurs independently of microtubules by a controlled fusion reaction. J Cell Biol 148: 883–898. Dreger M, Otto H, Neubauer G, Mann M, Hucho F. 1999. Identification of phosphorylation sites in native lamina-associated polypeptide 2 b. Biochemistry 38: 9426– 9434. Drin G, Casella JF, Gautier R, Boehmer T, Schwartz TU, Antonny B. 2007. A general amphipathic a-helical motif for sensing membrane curvature. Nat Struct Mol Biol 14: 138–146. Dultz E, Zanin E, Wurzenberger C, Braun M, Rabut G, Sironi L, Ellenberg J. 2008. Systematic kinetic analysis of

mitotic dis- and reassembly of the nuclear pore in living cells. J Cell Biol 180: 857 –865. Ellenberg J, Siggia ED, Moreira JE, Smith CL, Presley JF, Worman HJ, Lippincott-Schwartz J. 1997. Nuclear membrane dynamics and reassembly in living cells: Targeting of an inner nuclear membrane protein in interphase and mitosis. J Cell Biology 138: 1193–1206. Eriksson C, Rustum C, Hallberg E. 2004. Dynamic properties of nuclear pore complex proteins in gp210 deficient cells. FEBS Lett 572: 261 –265. Franz C, Askjaer P, Antonin W, Iglesias CL, Haselmann U, Schelder M, de Marco A, Wilm M, Antony C, Mattaj IW. 2005. Nup155 regulates nuclear envelope and nuclear pore complex formation in nematodes and vertebrates. EMBO J 24: 3519–3531. Franz C, Walczak R, Yavuz S, Santarella R, Gentzel M, Askjaer P, Galy V, Hetzer M, Mattaj IW, Antonin W. 2007. MEL-28/ELYS is required for the recruitment of nucleoporins to chromatin and postmitotic nuclear pore complex assembly. EMBO Rep 8: 165– 172. Fridkin A, Penkner A, Jantsch V, Gruenbaum Y. 2009. SUN-domain and KASH-domain proteins during development, meiosis and disease. Cell Mol Life Sci 66: 1518–1533. Galy V, Antonin W, Jaedicke A, Sachse M, Santarella R, Haselmann U, Mattaj I. 2008. A role for gp210 in mitotic nuclear-envelope breakdown. J Cell Sci 121: 317– 328. Galy V, Askjaer P, Franz C, Lopez-Iglesias C, Mattaj IW. 2006. MEL-28, a novel nuclear-envelope and kinetochore protein essential for zygotic nuclear-envelope assembly in C. elegans. Curr Biol 16: 1748–1756. Gant TM, Wilson KL. 1997. Nuclear assembly. Annu Rev Cell Dev Biol 13: 669 –695. Golden A, Liu J, Cohen-Fix O. 2009. Inactivation of the C. elegans lipin homolog leads to ER disorganization and to defects in the breakdown and reassembly of the nuclear envelope. J Cell Sci 122: 1970–1978. Gong D, Pomerening JR, Myers JW, Gustavsson C, Jones JT, Hahn AT, Meyer T, Ferrell JE Jr. 2007. Cyclin A2 regulates nuclear-envelope breakdown and the nuclear accumulation of cyclin B1. Curr Biol 17: 85– 91. Gorjanacz M, Mattaj IW. 2009. Lipin is required for efficient breakdown of the nuclear envelope in Caenorhabditis elegans. J Cell Sci 122: 1963–1969. Gorjanacz M, Klerkx EP, Galy V, Santarella R, Lopez-Iglesias C, Askjaer P, Mattaj IW. 2007. Caenorhabditis elegans BAF-1 and its kinase VRK-1 participate directly in postmitotic nuclear envelope assembly. EMBO J 26: 132– 143. Goss VL, Hocevar BA, Thompson LJ, Stratton CA, Burns DJ, Fields AP. 1994. Identification of nuclear bII protein kinase C as a mitotic lamin kinase. J Biol Chem 269: 19074– 19080. Griffis ER, Altan N, Lippincott-Schwartz J, Powers MA. 2002. Nup98 is a mobile nucleoporin with transcriptiondependent dynamics. Mol Biol Cell 13: 1282– 1297. Gruenbaum Y, Margalit A, Goldman RD, Shumaker DK, Wilson KL. 2005. The nuclear lamina comes of age. Nat Rev Mol Cell Biol 6: 21–31. Gruenbaum Y, Wilson KL, Harel A, Goldberg M, Cohen M. 2000. Review: nuclear lamins– structural proteins with fundamental functions. J Struct Biol 129: 313–323.

M.W. Hetzer Guttinger S, Laurell E, Kutay U. 2009. Orchestrating nuclear envelope disassembly and reassembly during mitosis. Nat Rev Mol Cell Biol 10: 178– 191. Harel A, Forbes DJ. 2004. Importin b: Conducting a much larger cellular symphony. Mol Cell 16: 319 –330. Harel A, Chan RC, Lachish-Zalait A, Zimmerman E, Elbaum M, Forbes DJ. 2003a. Importin b negatively regulates nuclear membrane fusion and nuclear pore complex assembly. Mol Biol Cell 14: 4387–4396. Harel A, Orjalo AV, Vincent T, Lachish-Zalait A, Vasu S, Shah S, Zimmerman E, Elbaum M, Forbes DJ. 2003b. Removal of a single pore subcomplex results in vertebrate nuclei devoid of nuclear pores. Mol Cell 11: 853–864. Hawryluk-Gara LA, Platani M, Santarella R, Wozniak RW, Mattaj IW. 2008. Nup53 is required for nuclear envelope and nuclear pore complex assembly. Mol Biol Cell 10.1091/mbc.E07-08-0820. Heessen S, Fornerod M. 2007. The inner nuclear envelope as a transcription factor resting place. EMBO Rep 8: 914–919. Hetzer M, Gruss OJ, Mattaj IW. 2002. The Ran GTPase as a markerof chromosome position in spindle formation and nuclear envelope assembly. Nat Cell Biol 4: E177– 184. Hetzer M, Walther TC, Mattaj IW. 2005. Pushing the envelope: Structure, function, and dynamics of the nuclear periphery. Annu Rev Cell Dev Biol 21: 367– 380. Hetzer M, Bilbao-Cortes D, Walther TC, Gruss OJ, Mattaj IW. 2000. GTP hydrolysis by Ran is required for nuclear envelope assembly. Mol Cell 5: 1013–1024. Hetzer M, Meyer HH, Walther TC, Bilbao-Cortes D, Warren G, Mattaj IW. 2001. Distinct AAA-ATPase p97 complexes function in discrete steps of nuclear assembly. Nat Cell Biol 3: 1086–1091. Heywood P. 1978. Ultrastructure of mitosis in the chloromonadophycean alga Vacuolaria virescens. J Cell Sci 31: 37–51. Hirano Y, Segawa M, Ouchi FS, Yamakawa Y, Furukawa K, Takeyasu K, Horigome T. 2005. Dissociation of emerin from barrier-to-autointegration factor is regulated through mitotic phosphorylation of emerin in a Xenopus egg cell-free system. J Biol Chem 280: 39925–39933. Hutchins JR, Moore WJ, Hood FE, Wilson JS, Andrews PD, Swedlow JR, Clarke PR. 2004. Phosphorylation regulates the dynamic interaction of RCC1 with chromosomes during mitosis. Curr Biol 14: 1099– 1104. Joseph J, Liu ST, Jablonski SA, Yen TJ, Dasso M. 2004. The RanGAP1-RanBP2 complex is essential for microtubulekinetochore interactions in vivo. Curr Biol 14: 611 –617. Joseph J, Tan SH, Karpova TS, McNally JG, Dasso M. 2002. SUMO-1 targets RanGAP1 to kinetochores and mitotic spindles. J Cell Biol 156: 595– 602. Katsani KR, Karess RE, Dostatni N, Doye V. 2008. In vivo dynamics of Drosophila nuclear envelope components. Mol Biol Cell 19: 3652–3666. Kimura N, Takizawa M, Okita K, Natori O, Igarashi K, Ueno M, Nakashima K, Nobuhisa I, Taga T. 2002. Identification of a novel transcription factor, ELYS, expressed predominantly in mouse foetal haematopoietic tissues. Genes Cells 7: 435 –446.

King MC, Lusk CP, Blobel G. 2006. Karyopherin-mediated import of integral inner nuclear membrane proteins. Nature 442: 1003– 1007. Kite GL. 1913. The relative permeability of the surface and interior portions of the cytoplasm of animal and plant cells. Biol Bull 25: 1–7. Kondo H, Rabouille C, Newman R, Levine TP, Pappin D, Freemont P, Warren G. 1997. p47 is a cofactor for p97-mediated membrane fusion. Nature 388: 75– 78. Kutay U, Hetzer MW. 2008. Reorganization of the nuclear envelope during open mitosis. Curr Opin Cell Biol 20: 669 –677. Lee KK, Gruenbaum Y, Spann P, Liu J, Wilson KL. 2000. C. elegans nuclear envelope proteins emerin, MAN1, lamin, and nucleoporins reveal unique timing of nuclear envelope breakdown during mitosis. Mol Biol Cell 11: 3089–3099. Lenart P, Rabut G, Daigle N, Hand AR, Terasaki M, Ellenberg J. 2003. Nuclear envelope breakdown in starfish oocytes proceeds by partial NPC disassembly followed by a rapidly spreading fenestration of nuclear membranes. J Cell Biol 160: 1055– 1068. Liu J, Prunuske AJ, Fager AM, Ullman KS. 2003. The COPI complex functions in nuclear envelope breakdown and is recruited by the nucleoporin Nup153. Dev Cell 5: 487 –498. Lohka MJ, Masui Y. 1983. Formation in vitro of sperm pronuclei and mitotic chromosomes induced by amphibian ooplasmic components. Science 220: 719–721. Loiodice I, Alves A, Rabut G, Van Overbeek M, Ellenberg J, Sibarita JB, Doye V. 2004. The entire Nup107–160 complex, including three new members, is targeted as one entity to kinetochores in mitosis. Mol Biol Cell 15: 3333–3344. Lu L, Ladinsky MS, Kirchhausen T. 2009. Cisternal organization of the endoplasmic reticulum during mitosis. Mol Biol Cell 15: 3471– 3480. Ma L, Tsai MY, Wang S, Lu B, Chen R, Iii JR, Zhu X, Zheng Y. 2009. Requirement for Nudel and dynein for assembly of the lamin B spindle matrix. Nat Cell Biol 11: 247–256. Mansfeld J, Guttinger S, Hawryluk-Gara LA, Pante N, Mall M, Galy V, Haselmann U, Muhlhausser P, Wozniak RW, Mattaj IW, et al. 2006. The conserved transmembrane nucleoporin NDC1 is required for nuclear pore complex assembly in vertebrate cells. Mol Cell 22: 93–103. Mattout A, Dechat T, Adam SA, Goldman RD, Gruenbaum Y. 2006. Nuclear lamins, diseases and aging. Curr Opin Cell Biol 18: 335– 341. Maul GG, Maul HM, Scogna JE, Lieberman MW, Stein GS, Hsu BY, Borun TW. 1972. Time sequence of nuclear pore formation in phytohemagglutinin-stimulated lymphocytes and in HeLa cells during the cell cycle. J Cell Biol 55: 433– 447. McCullough S, Lucocq J. 2005. Endoplasmic reticulum positioning and partitioning in mitotic HeLa cells. J Anat 206: 415– 425. Miller BR, Forbes DJ. 2000. Purification of the vertebrate nuclear pore complex by biochemical criteria. Traffic 1: 941 –951. Mora-Bermudez F, Gerlich D, Ellenberg J. 2007. Maximal chromosome compaction occurs by axial shortening in

Organization of the Nuclear Membrane anaphase and depends on Aurora kinase. Nat Cell Biol 9: 822–831. Mounkes L, Kozlov S, Burke B, Stewart CL. 2003. The laminopathies: Nuclear structure meets disease. Curr Opin Genet Dev 13: 223 –230. Muchir A, Worman HJ. 2004. The nuclear envelope and human disease. Physiology 19: 309–314. Muhlhausser P, Kutay U. 2007. An in vitro nuclear disassembly system reveals a role for the RanGTPase system and microtubule-dependent steps in nuclear envelope breakdown. J Cell Biol 178: 595– 610. Nachury MV, Maresca TJ, Salmon WC, Waterman-Storer CM, Heald R, Weis K. 2001. Importin b is a mitotic target of the small GTPase Ran in spindle assembly. Cell 104: 95–106. Neilan EG. 2009. Laminopathies, other progeroid disorders, and aging: Common pathogenic themes and possible treatments. Am J Med Genet A 149A: 563– 566. Newport J. 1987. Nuclear reconstitution in vitro: Stages of assembly around protein-free DNA. Cell 48: 205 –217. Newport J, Spann T. 1987. Disassembly of the nucleus in mitotic extracts: Membrane vesicularization, lamin disassembly, and chromosome condensation are independent processes. Cell 48: 219– 230. Ohba T, Schirmer EC, Nishimoto T, Gerace L. 2004. Energyand temperature-dependent transport of integral proteins to the inner nuclear membrane via the nuclear pore. J Cell Biol 167: 1051–1062. Ohsugi M, Adachi K, Horai R, Kakuta S, Sudo K, Kotaki H, Tokai-Nishizumi N, Sagara H, Iwakura Y, Yamamoto T. 2008. Kid-mediated chromosome compaction ensures proper nuclear envelope formation. Cell 132: 771–782. Ohtsubo M, Okazaki H, Nishimoto T. 1989. The RCC1 protein, a regulator for the onset of chromosome condensation locates in the nucleus and binds to DNA. J Cell Biol 109: 1389– 1397. Onischenko E, Stanton LH, Madrid AS, Kieselbach T, Weis K. 2009. Role of the Ndc1 interaction network in yeast nuclear pore complex assembly and maintenance. J Cell Biol 185: 475–491. Orjalo AV, Arnaoutov A, Shen Z, Boyarchuk Y, Zeitlin SG, Fontoura B, Briggs S, Dasso M, Forbes DJ. 2006. The Nup107–160 nucleoporin complex is required for correct bipolar spindle assembly. Mol Biol Cell 17: 3806– 3818. Osmani AH, Davies J, Liu HL, Nile A, Osmani SA. 2006. Systematic deletion and mitotic localization of the nuclear pore complex proteins of Aspergillus nidulans. Mol Biol Cell 17: 4946–4961. Peters R. 2009. Translocation through the nuclear pore: Kaps pave the way. Bioessays 31: 466– 477. Pichler A, Gast A, Seeler JS, Dejean A, Melchior F. 2002. The nucleoporin RanBP2 has SUMO1 E3 ligase activity. Cell 108: 109– 120. Portier N, Audhya A, Maddox PS, Green RA, Dammermann A, Desai A, Oegema K. 2007. A microtubule-independent role for centrosomes and aurora a in nuclear envelope breakdown. Dev Cell 12: 515– 529. Poteryaev D, Squirrell JM, Campbell JM, White JG, Spang A. 2005. Involvement of the actin cytoskeleton and

homotypic membrane fusion in ER dynamics in Caenorhabditis elegans. Mol Biol Cell 16: 2139–2153. Puhka M, Vihinen H, Joensuu M, Jokitalo E. 2007. Endoplasmic reticulum remains continuous and undergoes sheet-to-tubule transformation during cell division in mammalian cells. J Cell Biol 179: 895 –909. Pyrpasopoulou A, Meier J, Maison C, Simos G, Georgatos SD. 1996. The lamin B receptor (LBR) provides essential chromatin docking sites at the nuclear envelope. EMBO J 15: 7108–7119. Rabut G, Doye V, Ellenberg J. 2004. Mapping the dynamic organization of the nuclear pore complex inside single living cells. Nat Cell Biol 6: 1114– 1121. Ramadan K, Bruderer R, Spiga FM, Popp O, Baur T, Gotta M, Meyer HH. 2007. Cdc48/p97 promotes reformation of the nucleus by extracting the kinase Aurora B from chromatin. Nature 450: 1258–1262. Rasala BA, Orjalo AV, Shen Z, Briggs S, Forbes DJ. 2006. ELYS is a dual nucleoporin/kinetochore protein required for nuclear pore assembly and proper cell division. Proc Natl Acad Sci 103: 17801– 17806. Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243 –247. Ribeiro KC, Pereira-Neves A, Benchimol M. 2002. The mitotic spindle and associated membranes in the closed mitosis of trichomonads. Biol Cell 94: 157– 172. Roux KJ, Burke B. 2007. Nuclear envelope defects in muscular dystrophy. Biochim Biophys Acta 1772: 118– 127. Ryan KJ, McCaffery JM, Wente SR. 2003. The Ran GTPase cycle is required for yeast nuclear pore complex assembly. J Cell Biol 160: 1041–1053. Salina D, Bodoor K, Eckley DM, Schroer TA, Rattner JB, Burke B. 2002. Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell 108: 97–107. Salina D, Enarson P, Rattner JB, Burke B. 2003. Nup358 integrates nuclear envelope breakdown with kinetochore assembly. J Cell Biol 162: 991–1001. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059– 1063. Schirmer EC, Foisner R. 2007. Proteins that associate with lamins: Many faces, many functions. Exp Cell Res 313: 2167–2179. Schirmer EC, Gerace L. 2005. The nuclear membrane proteome: Extending the envelope. Trends Biochem Sci 30: 551– 558. Schirmer EC, Florens L, Guan T, Yates JR 3rd, Gerace L. 2003. Nuclear membrane proteins with potential disease links found by subtractive proteomics. Science 301: 1380–1382. Segura-Totten M, Kowalski AK, Craigie R, Wilson KL. 2002. Barrier-to-autointegration factor: Major roles in chromatin decondensation and nuclear assembly. J Cell Biol 158: 475–485. Sheehan MA, Mills AD, Sleeman AM, Laskey RA, Blow JJ. 1988. Steps in the assembly of replication-competent nuclei in a cell-free system from Xenopus eggs. J Cell Biol 106: 1 –12. Siniossoglou S. 2009. Lipins, Lipids and Nuclear Envelope Structure. Traffic 10: 1181– 1187.

M.W. Hetzer Starr DA, Han M. 2003. ANChors away: an actin based mechanism of nuclear positioning. J Cell Sci 116: 211–216. Stavru F, Hulsmann BB, Spang A, Hartmann E, Cordes VC, Gorlich D. 2006. NDC1: A crucial membrane-integral nucleoporin of metazoan nuclear pore complexes. J Cell Biol 173: 509–519. Stewart CL, Roux KJ, Burke B. 2007. Blurring the boundary: The nuclear envelope extends its reach. Science 318: 1408– 1412. Tahara K, Takagi M, Ohsugi M, Sone T, Nishiumi F, Maeshima K, Horiuchi Y, Tokai-Nishizumi N, Imamoto F, Yamamoto T, et al. 2008. Importin-b and the small guanosine triphosphatase Ran mediate chromosome loading of the human chromokinesin Kid. J Cell Biol 180: 493–506. Terasaki M, Campagnola P, Rolls MM, Stein PA, Ellenberg J, Hinkle B, Slepchenko B. 2001. A new model for nuclear envelope breakdown. Mol Biol Cell 12: 503–510. Terry LJ, Shows EB, Wente SR. 2007. Crossing the nuclear envelope: Hierarchical regulation of nucleocytoplasmic transport. Science 318: 1412–1416. Tran EJ, Wente SR. 2006. Dynamic nuclear pore complexes: Life on the edge. Cell 125: 1041–1053. Tsai MY, Wang S, Heidinger JM, Shumaker DK, Adam SA, Goldman RD, Zheng Y. 2006. A mitotic lamin B matrix induced by RanGTP required for spindle assembly. Science 311: 1887–1893. Tzur YB, Wilson KL, Gruenbaum Y. 2006. SUN-domain proteins: ’Velcro’ that links the nucleoskeleton to the cytoskeleton. Nat Rev Mol Cell Biol 7: 782– 788. Ulbert S, Platani M, Boue S, Mattaj IW. 2006. Direct membrane protein-DNA interactions required early in nuclear envelope assembly. J Cell Biol 173: 469– 476. Vergnolle MA, Taylor SS. 2007. Cenp-F links kinetochores to Ndel1/Nde1/Lis1/dynein microtubule motor complexes. Curr Biol 17: 1173–1179. Vlcek S, Foisner R. 2007. Lamins and lamin-associated proteins in aging and disease. Curr Opin Cell Biol 19: 298–304. Voeltz GK, Prinz WA. 2007. Sheets, ribbons and tubules— how organelles get their shape. Nat Rev Mol Cell Biol 8: 258–264. Voeltz GK, Prinz WA, Shibata Y, Rist JM, Rapoport TA. 2006. A class of membrane proteins shaping the tubular endoplasmic reticulum. Cell 124: 573– 586. Walther TC, Alves A, Pickersgill H, Loiodice I, Hetzer M, Galy V, Hulsmann BB, Kocher T, Wilm M, Allen T, et al.

2003a. The conserved Nup107– 160 complex is critical for nuclear pore complex assembly. Cell 113: 195– 206. Walther TC, Askjaer P, Gentzel M, Habermann A, Griffiths G, Wilm M, Mattaj IW, Hetzer M. 2003b. RanGTP mediates nuclear pore complex assembly. Nature 424: 689 –694. Watson ML. 1955. The nuclear envelope; its structure and relation to cytoplasmic membranes. J Biophys Biochem Cytol 1: 257– 270. Wilhelmsen K, Ketema M, Truong H, Sonnenberg A. 2006. KASH-domain proteins in nuclear migration, anchorage and other processes. J Cell Sci 119: 5021–5029. Wilhelmsen K, Litjens SH, Kuikman I, Tshimbalanga N, Janssen H, van den Bout I, Raymond K, Sonnenberg A. 2005. Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. J Cell Biol 171: 799– 810. Winey M, Yarar D, Giddings TH Jr, Mastronarde DN. 1997. Nuclear pore complex number and distribution throughout the Saccharomyces cerevisiae cell cycle by three-dimensional reconstruction from electron micrographs of nuclear envelopes. Mol Biol Cell 8: 2119–2132. Worman HJ, Bonne G. 2007. “Laminopathies”: Awide spectrum of human diseases. Exp Cell Res 313: 2121– 2133. Wu L, Osmani SA, Mirabito PM. 1998. A role for NIMA in the nuclear localization of cyclin B in Aspergillus nidulans. J Cell Biol 141: 1575– 1587. Yang L, Guan T, Gerace L. 1997. Integral membrane proteins of the nuclear envelope are dispersed throughout the endoplasmic reticulum during mitosis. J Cell Biol 137: 1199–1210. Ye Y, Meyer HH, Rapoport TA. 2003. Function of the p97Ufd1-Npl4 complex in retrotranslocation from the ER to the cytosol: Dual recognition of nonubiquitinated polypeptide segments and polyubiquitin chains. J Cell Biol 162: 71– 84. Zhang C, Clarke PR. 2000. Chromatin-independent nuclear envelope assembly induced by Ran GTPase in Xenopus egg extracts. Science 288: 1429–1432. Zheng Y, Tsai MY. 2006. The mitotic spindle matrix: A fibromembranous lamin connection. Cell Cycle 5: 2345–2347. Zuccolo M, Alves A, Galy V, Bolhy S, Formstecher E, Racine V, Sibarita JB, Fukagawa T, Shiekhattar R, Yen T, et al. 2007. The human Nup107–160 nuclear pore subcomplex contributes to proper kinetochore functions. EMBO J 26: 1853–1864.

Nuclear Lamins Thomas Dechat1, Stephen A. Adam1, Pekka Taimen2, Takeshi Shimi1, and Robert D. Goldman1 1

Department of Cell and Molecular Biology, Feinberg School of Medicine, Northwestern University, Chicago, Illinois 60611

2

Department of Pathology, University of Turku, Kiinamyllynkatu 10 20520 Turku, Finland

Correspondence: [email protected]

The nuclear lamins are type V intermediate filament proteins that are critically important for the structural properties of the nucleus. In addition, they are involved in the regulation of numerous nuclear processes, including DNA replication, transcription and chromatin organization. The developmentally regulated expression of lamins suggests that they are involved in cellular differentiation. Their assembly dynamic properties throughout the cell cycle, particularly in mitosis, are influenced by posttranslational modifications. Lamins may regulate nuclear functions by direct interactions with chromatin and determining the spatial organization of chromosomes within the nuclear space. They may also regulate chromatin functions by interacting with factors that epigenetically modify the chromatin or directly regulate replication or transcription.

filamentous layer located between the inner nuclear membrane (INM) and peripheral heterochromatin was evident even in early electron microscopic studies of vertebrate cell nuclei (Fawcett 1966). This layer, later termed the nuclear lamina, is also found to be closely associated with nuclear pore complexes (NPCs) and contains three major structurally related polypeptides (Aaronson and Blobel 1975). These proteins are named nuclear lamins A, B, and C according to their molecular weights (Gerace and Blobel 1980). Further biochemical characterization and cDNA cloning of the nuclear lamins classifies them as type V intermediate filament proteins (Goldman et al. 1986; McKeon et al. 1986).

A

STRUCTURE AND BIOCHEMICAL PROPERTIES OF NUCLEAR LAMINS Lamin Isoforms and Expression Patterns

Lamins are present in all metazoans examined to date ranging from hydra to human, but are not found in unicellular organisms and plants (Cohen et al. 2001; Melcer et al. 2007). Extensive characterization in several model organisms including humans, mice, frogs, fruit flies and nematodes shows that their properties are shared across species (Melcer et al. 2007; Dechat et al. 2008b). Based on their sequence homologies, expression patterns, structural features, and biochemical and dynamic properties, lamins are subdivided into A- and B-types. All

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000547 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000547

T. Dechat et al.

metazoans express at least one B-type lamin. Typically, invertebrates have only a single lamin gene of the B-type, with some exceptions such as Drosophila, which expresses one B-type (lamin Dm0) and one A-type lamin (lamin C) encoded by two distinct genes. Most vertebrates have one A-type lamin and two B-type lamin genes except for Xenopus, which has three B-type genes (Melcer et al. 2007). The two major isoforms of vertebrate A-type lamins, lamins A and C, are derived from a single gene (LMNA) by alternative splicing (see Fig. 1) (Lin and Worman 1993). In some vertebrates, alternative splicing can also produce two less abundant isoforms, lamins AD10 and C2, from LMNA (Nakajima and Abe 1995; Machiels et al. 1996). Lamins B1 and B2 are the two major B-type lamins in most vertebrates. They are encoded by the LMNB1 and LMNB2 genes, respectively (Peter et al. 1989; Vorburger et al. 1989). The latter also encodes the minor isoform lamin B3 (Furukawa and Hotta 1993). Although at least one B-type lamin is expressed in all cells throughout development, the expression of A-type lamins is developmentally regulated (Benavente et al. 1985; Schatten et al. 1985; Lehner et al. 1987). During mouse development, lamins A and C are not expressed until days 10 –12 of mouse embryogenesis and then primarily in primordial muscle cells (Stewart and Burke 1987; Rober et al. 1989). Lamin A/C expression in other organs does not occur until after birth (Rober et al. 1989). Cells of hematopoietic lineage express only B-type lamins (Guilly et al. 1990; Rober et al. 1990). Similar patterns of expression of A and B-type lamins take place during the developmental α-helical rod L12

L1 Lamin A

Lamin C

1A

1A

1B

1B

progression of other vertebrates (Benavente et al. 1985; Lehner et al. 1987; Prather et al. 1989) and Drosophila (Frasch et al. 1988; Riemer et al. 1995). The regulated expression of A- and B-type lamins is also evident during differentiation of stem cells in culture. For example, undifferentiated human and mouse embryonic stem (ES) cells lack lamins A and C, but express lamins B1 and B2 (Constantinescu et al. 2006). The minor mammalian isoforms, lamins C2 and B3, are expressed exclusively in germ cells (Furukawa and Hotta 1993; Machiels et al. 1995; Nakajima and Abe 1995), whereas small amounts of lamin AD10 appear to be present in a variety of cell types (Machiels et al. 1995). The importance of the developmental regulation of lamin expression is evident from studies in LMNA knockout (LMNA 2/2 ) and LMNB1 mutant (LMNB D/D) mice. In mice null for the A-type lamins, no obvious embryonic defects can be detected, but these animals show severe postnatal growth retardation and muscular dystrophy (Sullivan et al. 1999). On the other hand, mice with an insertional mutation in LMNB1 develop defects in their lungs and bones during embryogenesis and die at birth, even though they continue to express lamin B2 (Vergnes et al. 2004). Structure and Assembly of the Nuclear Lamins

The nuclear lamins have the typical tripartite structure of intermediate filament (IF) proteins, consisting of a highly a-helical central rod domain flanked by a short globular amino-terminal “head” domain and a longer carboxy-terminal NLS

lg-fold

L2 2A

2A

2B 389

417 436

544

389

417 436

544 572

646

2B

Figure 1. Structure of nuclear lamins. Schematic drawing of mature lamin A and lamin C polypeptide chains.

The lamin structure consists of a short amino terminal head domain, a central a-helical rod domain (red), and the carboxy-terminal domain containing the NLS and the Ig-fold (blue; the nine b-strands of the Ig-fold motif are depicted). Modified, with permission, from (Dechat et al. 2008b).

Nuclear Lamins

“tail” domain (Parry et al. 1986). The central rod domain is composed of four subhelical regions comprised of heptad repeats and designated as coil 1A, 1B, 2A, and 2B. These individual “coils” are separated from each other by three short linker segments, L1, L12, and L2, of which L12 is the most flexible (Parry et al. 1986). Although the head domain appears to be unstructured, the tail domain contains a highly conserved structural motif similar to a type s immunoglobulin fold (Ig-fold) (Dhe-Paganon et al. 2002; Krimm et al. 2002). A nuclear localization signal (NLS), required for transport into the nucleus, is present in all lamins between the carboxy-terminal end of the central rod domain and the Ig-fold (Loewinger and McKeon 1988) (Fig. 1). In vitro, lamins self-assemble into higher order structures through a series of steps. The first step involves the formation of a coiled-coil dimer by the in register and in parallel association of two a-helical rod domains into a left handed superhelix (Heitlinger et al. 1991) (Fig. 2A). Next, the lamin dimers associate in a head-to-tail fashion (Heitlinger et al. 1991; Stuurman et al. 1996) and these polarized arrays interact in an antiparallel fashion to form apolar tetrameric protofilaments (Heitlinger et al. 1991) (Fig. 2B). The interaction of four lamin protofilaments leads to the formation of 10-nm filaments (Heitlinger et al. 1991; Stuurman et al. 1996; Ben-Harush et al. 2009) (Fig. 2C,D). Although the assembly of some invertebrate lamins appears to terminate at this stage of the assembly process, vertebrate lamins continue to assemble into well ordered paracrystalline arrays in vitro (Zackroff and Goldman 1979; Heitlinger et al. 1991; Stuurman et al. 1996; Ben-Harush et al. 2009) (Fig. 2E). The assembly of lamin proteins differs from their cytoplasmic counterparts in several ways. For example, lamin dimers associate by polar “head-to-tail” interactions to form protofilaments, which is very different from the halfstaggered, antiparallel side-by-side association of cytoplasmic IF dimers, which assemble into 10 nm filaments (Strelkov et al. 2004). This difference is at least in part explained by the crystal structure of the coiled-coil dimer of the lamin A coil 2B region (Strelkov et al. 2004). Although

Figure 2. Assembly of the nuclear lamins in vitro. Lamins self assemble to form dimers (A) which then join to form linear head-to-tail polymers ( protofilaments) (B). Bar ¼ 100 nm; electron micrograph of rotary shadowed chicken lamin B2. These protofilaments further assemble into “beaded” filaments or fibers (C ) which in turn associate laterally into thicker fibers (D), and eventually into paracrystalline arrays (E); C,D are negatively stained electron microscope preparations. Reprinted from Stuurman et al. (1998) with permission from Elsevier.

this structure appears to be similar to the overall structure of the homologous segment of the cytoplasmic IF protein vimentin, the distribution of charged residues varies causing significant changes in the patterns of intra- and interhelical salt bridges. Additionally, vertebrate lamins have six extra heptads in the central rod domain (McKeon et al. 1986), which may also help to explain assembly differences between cytoplasmic IF and lamins. The central rod domains of some invertebrates lamins contain fewer extra heptads and these lamins stop assembling into higher order structures at the 10-nm (IF) stage of assembly, similar to cytoplasmic IF proteins (Ben-Harush et al. 2009). Posttranslational Processing and Modifications of the Nuclear Lamins

Lamins A, B1, and B2 are expressed as prelamins that require extensive posttranslational modifications of their carboxy-terminal – CAAX box

T. Dechat et al.

to become mature lamins (Rusinol and Sinensky 2006; Davies et al. 2009). Modification of the – CAAX box takes place in a highly regulated temporal sequence starting with the farnesylation of the cysteine residue (Farnsworth et al. 1990; Lutz et al. 1992) by a farnesyltransferase (Zhang and Casey 1996). This modification initiates a sequence of processing steps (Beck et al. 1990) beginning with the removal of the – AAX by a CAAX prenyl protease. Two members of this zinc metalloproteinase family have been identified in humans: Rce1 (Ras-converting enzyme 1) and Zmpste24 (Zinc metalloprotease related to the STE24 homolog in yeast), also known as FACE1 (farnesylated-proteins converting enzyme)(Boyartchuk et al. 1997; Leung et al. 2001; Corrigan et al. 2005); followed by carboxymethylation of the carboxy-terminal

cysteine by isoprenylcysteine carboxyl methyltransferase (Icmt) (Winter-Vann and Casey 2005). In contrast to B-type lamins, which remain permanently farnesylated and carboxymethylated, an additional 15 amino acids are removed from the carboxyl terminus of farnesylated/carboxymethylated prelamin A by Zmpste24/FACE1 (Corrigan et al. 2005). This final processing step results in the production of mature lamin A lacking the carboxy-terminal farnesyl- and carboxymethyl-modifications (Fig. 3). This sequence of lamin processing steps is highly regulated and each step depends on the preceding modification (Kilic et al. 1997). Lamin C, which is 74 residues shorter than mature lamin A, does not possess a – CAAX box and therefore is not farnesylated or otherwise modified (see Fig. 1). The precise location

Pre-lamin CAAX-COOH Farnesyltransferase S

CAAX-COOH AAX endopeptidase S

Carboxylmethyltransferase

C-COOH

S

C-COCH3 Zmpste24/FACE1 S

C-COCH3 417 436

544 Mature lamin A

646

545 415 438 Mature lamin B1

583

S

C-COCH3 577 435 470 Mature lamin B2

617

Figure 3. Posttranslational processing of the carboxyl terminus of prelamins A, B1, and B2. Processing takes place in a series of steps: (1) addition of a farnesyl group to the cysteine residue of the –CAAX box of pre-lamin A, prelamin B1 and prelamin B2 by a farnesyltransferase; (2) removal of the last three residues (2AAX) by an AAX endopeptidase; (3) methylation of the terminal carboxylic acid group (2COOH) by a carboxyl methyltransferase; (4) removal of the carboxyl terminal 15 amino acids of lamin A with the farnesyl attached by the metalloprotease Zmpste24/FACE1. This last proteolysis step does not occur on B-type lamins and therefore they remain farnesylated. Modified, with permission, from Dechat et al. (2008b).

Nuclear Lamins

of the intracellular post-translational processing sites for the lamins remains largely unknown. Although the enzymes required for processing the carboxyl terminus of the lamins are present in both the inner nuclear membrane and endoplasmic reticulum membranes, the complete processing of prelamin A can occur when the protein is confined either to the cytoplasm or to the nucleus (Barrowman et al. 2008). Even though the prelamins can be processed in the cytoplasm, it is more likely that all lamin processing occurs in the nucleus (Lutz et al. 1992), because lamins are rapidly transported into the nucleus after translation in the cytoplasm (Lehner et al. 1986). In addition to farnesylation and carboxymethylation, lamins are also posttranslationally modified by phosphorylation (Ottaviano and Gerace 1985), sumoylation (Zhang and Sarge 2008), ADP-ribosylation (Adolph 1987), and possibly by glycosylation (Ferraro et al. 1989). Phosphorylation of serine and threonine residues proximal to the NLS by protein kinase C is known to play a role in the regulated import of lamins into the nucleus (Hennekes et al. 1993; Leukel and Jost 1995). At the onset of mitosis, the phosphorylation of lamins at specific sites by cyclin-dependent kinase (Cdk) 1 and protein kinase C (PKC) is required to drive disassembly of the lamina (Gerace and Blobel 1980; Dessev and Goldman 1988; Dessev et al. 1988; Dessev et al. 1989; Heald and McKeon 1990; Peter et al. 1990; Ward and Kirschner 1990; Dessev et al. 1991; Molloy and Little 1992; Goss et al. 1994; Collas 1999). Subsequently, dephosphorylation of the mitotic sites by protein phosphatase 1a is required for lamin/lamina assembly during the telophase/early G1 transition (Thompson et al. 1997). Lamin Structure and Dynamic Properties within the Lamina and Nucleoplasm

Very little is known about the formation, composition, and structure of lamin polymers in vivo. The structures of the lamins within the lamina that have been described to date range from a regular meshwork of 10 – 15 nm filaments observed in Xenopus oocyte germinal

vesicles and in sperm pronuclei assembled in vitro (Aebi et al. 1986; Zhang and Casey 1996; Goldberg et al. 2008a) (Fig. 4A), to a more irregular filamentous meshwork seen in mammalian cells (Belmont et al. 1993; Schermelleh et al. 2008; Shimi et al. 2008) (Fig. 4B,C). These meshworks have long been assumed to be stable structures based on the resistance of the nuclear lamins within the lamina to extraction in detergent containing high salt solutions, and the propensity of purified lamins to polymerize into higher order insoluble structures in vitro at relatively low critical concentrations (Aebi et al. 1986; Lourim and Lin 1989; Glass and Gerace 1990). In further support of this assumption, measurements of lamin mobility by fluorescence recovery after photobleaching (FRAP) reveal that GFP-tagged A- and B-type lamins do not appreciably exchange subunits within the lamina for up to several hours throughout most of interphase (Broers et al. 1999; Moir et al. 2000; Dahl et al. 2006). Recently it has become apparent that the Aand B-type lamins form separate filamentous networks in the lamina. These structures can be identified by both high resolution light microscopy (Schermelleh et al. 2008; Shimi et al. 2008) and whole mount electron microscopy in lamin B containing Xenopus oocyte nuclei ectopically expressing A-type lamins (Goldberg et al. 2008b). Even though the A-type and B-type lamin networks appear to be mainly separate structures (Shimi et al. 2008), there is evidence that the individual networks overlap and interact to varying degrees. Evidence for interaction between these two lamin meshworks is based on studies using fluorescence resonance energy transfer (FRET) in combination with time domain fluorescence lifetime imaging and high resolution confocal immunofluorescence (Moir et al. 2000; Delbarre et al. 2006; Shimi et al. 2008). Other evidence supporting the interactions between the A and B type lamins is derived from studies of nuclei in which either A or B type lamins are perturbed. Fibroblasts derived from LMNA 2/2 mouse embryos (MEFs) or a limb girdle muscular dystrophy patient with a homozygous nonsense mutation in LMNA

T. Dechat et al.

Figure 4. The nuclear lamins form a meshwork of fil-

aments within the lamina. (A) Spread nuclear envelope from Xenopus oocytes after detergent extraction and preparation for transmission electron microscopy by freeze-drying/unidirectional metal shadowing. The micrograph shows the nuclear lamina meshwork partially studded with nuclear pore complexes. (Inset) Higher-magnification view of a particularly wellpreserved area clearly shows the near-tetragonal lamina meshwork. Bars, 1 mm. Reprinted from Stuurman et al. (1998), with permission from Elsevier. (B) Structured illumination microsopy (SIM) reveals that there is an irregular meshwork of nuclear lamin B as revealed by immunofluorescence (green). This preparation is also stained with antibodies directed against nuclear pores and is stained with DAPI for DNA/ chromatin. Pores (red), DAPI (blue). From Schermelleh et al. (2008). Reprinted with permission from AAAS. (C) Confocal immunofluorescence localization of lamin A/C (green) and lamin B1 (red) in HeLa cells. Lamins A/C and B1 in a single nucleus are seen in an equatorial section (left panels) and the nuclear surface (right panels). The areas indicated by white squares in the top panels are enlarged fivefold in the lower panels. These images demonstrate that lamins form mainly separate networks with some overlapping regions. Bar, 5 mm. Adapted, with permission, from Shimi et al. (2008).

contain abnormally shaped nuclei containing blebs or lobules (Sullivan et al. 1999; Muchir et al. 2003). A significant number of these blebbed regions lack B-type lamins and NPCs. In addition, nuclei in LMNB D/D MEFs are also highly lobulated, showing a dramatic increase in the mesh size of the lamin A/C network (Vergnes et al. 2004). HeLa cells depleted of lamin B1 by shRNA silencing frequently develop extensively enlarged lamin A/C meshworks and nuclear envelope blebs which lack lamin B2 and NPCs (Shimi et al. 2008). Many of these nuclear phenotypes are also found in cells endogenously or ectopically expressing either point mutations or truncations of lamins A/C (Ostlund et al. 2001; Vaughan et al. 2001; Vigouroux et al. 2001; Bechert et al. 2003; Favreau et al. 2003; Muchir et al. 2003; Goldman et al. 2004). It is also noteworthy that lamin A is more mobile in the nuclear lamina in nuclei with decreased amounts of lamin B1 (Tang et al. 2008). The A- and B-type lamins have different disassembly and assembly properties during mitosis as revealed by biochemical fractionation, immunofluorescence, and live cell imaging analysis, further supporting the idea that the two types of lamins form separate networks in the lamina. As described earlier, depolymerization of the lamins is regulated by mitotic kinases and reassembly requires dephosphorylation by protein phosphatase 1a (Thompson et al. 1997; Ito et al. 2007). When the nuclear envelope is disassembled during late prophase, A-type lamins become dispersed throughout the cytoplasm in an apparently freely diffusible state whereas the majority of B-type lamins remain associated with the nuclear membranes, which appear to be mainly dispersed into the endoplasmic reticulum (Gerace and Blobel 1980; Stick et al. 1988). This association of B-type lamins with membranes during mitosis appears to be mainly attributable to their permanently farnesylated state (Rusinol and Sinensky 2006). Further support for differences in the properties of the A- and B-type lamins is derived from studies revealing their spatial and temporal order of assembly into the nuclear lamina at anaphase/telophase and in the early stages of

Nuclear Lamins

G1. For example, in mouse keratinocytes and hamster cells, lamin B1 accumulates along the entire periphery of the decondensing daughter chromosomes and assembles into a relatively stable polymer by mid- to late- telophase (Broers et al. 1999; Moir et al. 2000). In these cells, lamin A begins to accumulate within the nucleus mainly after the major components of the nuclear envelope including NPCs are assembled in daughter cells in late telophase. In HeLa cells, however, a small fraction of A-type lamins associates with chromosomes much earlier than in mouse cells, first assembling in the central or “core” region of chromosomes in close association with kinetochores (Dechat et al. 2004; Dechat et al. 2007; Haraguchi et al. 2008) (Fig. 5). The remainder of the A-type lamins is then transported into the nucleus after formation of an intact nuclear envelope. Later, other elements of the nuclear envelope begin to assemble in daughter cells (Dechat et al. 2004; Dechat et al. 2007; Haraguchi et al. 2008). Although the major fraction of the various lamin isoforms is associated with the nuclear lamina, these proteins are also present

throughout the nucleoplasm, especially in interphase (Goldman et al. 1992; Lutz et al. 1992). These nucleoplasmic lamins are likely to have different functions, for example in DNA replication and transcription (Dechat et al. 2008b), and a small fraction may represent assembly intermediates that are subsequently incorporated into the nuclear lamina (Goldman et al. 1992). Interestingly, B-type lamins in the nucleoplasm appear to be relatively static, similar to those in the lamina, whereas the nucleoplasmic A-type lamins are much more dynamic. These differences in the properties of nucleoplasmic A- and B-type lamins are supported experimentally by their biochemical extractability and their dynamic properties as determined in vivo by FRAP analyses and fluorescence correlation spectroscopy (FCS) (Broers et al. 1999; Moir et al. 2000; Shimi et al. 2008). For example, FCS measurements of GFP-tagged lamins A/C and lamins B1/B2 showed that nucleoplasmic lamins A/C are highly mobile, whereas lamin B1 and lamin B2 are mainly immobile (Shimi et al. 2008). This latter observation suggests that nucleoplasmic B-type lamins are either assembled into some type of

Figure 5. Association of lamin A with chromosomes during mitosis. HeLa cells expressing GFP-lamin A were

followed by time-lapse microscopy from the metaphase/anaphase transition (far left panels) into early G1 (far right panels). GFP-lamin A first associates with the core regions of chromosomes during telophase and spreads to cover the entire chromatin surface by early G1. DIC images of the same series are shown in the bottom row (Dechat et al. 2007).

T. Dechat et al.

structure or are tightly associated with other unknown immobile structural components. Interestingly, decreasing the amount of nucleoplasmic lamin B1 by shRNA silencing increases the rate of mobility for a large fraction of nucleoplasmic lamin A (Shimi et al. 2008). Taken together, these FCS results suggest that, as in the case of the lamina, the A- and B-type lamins form separate yet interacting structures within the nucleoplasm. Obviously, it will be of great interest in the future to determine the specific functions of these nucleoplasmic lamins.

FUNCTIONS OF THE LAMINS Regulation of Nuclear Shape and Mechanical Stability

It is becoming increasingly obvious that some of the functions of nuclear lamins are analogous to those of cytoskeletal intermediate filament proteins, which are known to be involved in the determination and maintenance of cell shape and mechanical properties (Goldman et al. 2008). In support of this, the nuclei of LMNA 2/2 MEFs display increased deformability and impaired viability under mechanical strain compared to the nuclei in control MEFs (Houben et al. 2007). Increased nuclear deformability is also observed in human ES cells lacking A-type lamins as compared to nuclei in differentiated cells expressing the A-type lamins (Pajerowski et al. 2007). In addition, cells either deficient in lamins or expressing mutant lamin proteins often contain misshapen nuclei (Ostlund et al. 2001; Vaughan et al. 2001; Vigouroux et al. 2001; Bechert et al. 2003; Favreau et al. 2003; Muchir et al. 2003; Goldman et al. 2004). Quantitative rheological measurements, particle tracking methods and differential interference contrast microscopy reveal that in vitro reconstituted lamin B1 networks are extremely porous, with elastic stiffness providing resistance to shear deformations (Panorchan et al. 2004a; Panorchan et al. 2004b). Interestingly, measurements made by atomic force microscopy show that germinal vesicles isolated from Xenopus oocytes, which normally contain only B-type lamins, display a significant

increase in stiffness on the ectopic expression of lamin A (Schape et al. 2009). Nuclear Lamins and the Regulation of Chromatin Positioning and Gene Expression

Lamins may regulate transcription by organizing chromatin into active and inactive domains. For example, electron microscopic and light microscopic observations show that there is a close association between peripherally localized heterochromatin and the nuclear lamina (Fawcett 1966; Paddy et al. 1990) (Fig. 6). This suggests that lamins may be involved in anchoring or organizing interphase chromosomes (Sullivan et al. 1999; Goldman et al. 2004; Nikolova et al. 2004; Galiova et al. 2008). Additional support for this idea comes from biochemical experiments showing that lamins A/C bind to mitotic chromosomes and to polynucleosomal particles in vitro (Burke 1990; Glass and Gerace 1990; Yuan et al. 1991). The interaction of lamins with chromatin could be mediated either by their direct binding to histones (Taniura et al. 1995; Goldberg et al. 1999; Mattout et al. 2007), or to specific DNA sequences in the matrix attachment/scaffold-associated regions (MARs/ SARs) (Luderus et al. 1992; Luderus et al. 1994; Baricheva et al. 1996; Zhao et al. 1996). The Role of Lamins in Mitosis

In addition to nuclear disassembly, several lines of evidence suggest a direct role for lamins in nuclear assembly following mitosis. The

Figure 6. The nuclear lamina is a fibrous network (fibrous lamina) located between peripheral heterochromatin and the inner nuclear membrane. A transmission electron micrograph of a thin section showing a portion of a smooth muscle nucleus in guinea pig epedidymis. Adapted from Fawcett (1966), copyright 1995, Wiley-Liss, Inc. Reprinted with permission of John Wiley & Sons, Inc. American Journal of Anatomy, Vol. 119, No. 1, pg. 140.

Nuclear Lamins

microinjection of lamin antibodies into mitotic cells causes arrest in a telophase-like state with the chromosomes remaining condensed (Benavente and Krohne 1986). In addition, decreased expression of lamin Dm0 in Drosophila inhibits nuclear membrane assembly and causes an enrichment of NPCs in cytoplasmic annulate lamellae (Lenz-Bohme et al. 1997). In Caenorhabditis elegans, the down-regulation of the single lamin leads to a loss of chromosomes, defects in chromosome separation into daughter cell nuclei, and abnormal condensation of chromatin (Liu et al. 2000). In vitro nuclear assembly in frog egg extracts can be inhibited by addition of the C-terminal tail fragment of lamin B3, in part because of the Ig-fold motif in the lamin tail, which can inhibit lamin polymerization in vitro (Shumaker et al. 2005). Lastly, the expression of the mutant lamin A which causes Hutchinson-Gilford Progeria Syndrome (HGPS) leads to defects in cell division including a delay in cytokinesis, delayed completion of nuclear reassembly at the end of mitosis and an increase in mitotic cells (Cao et al. 2007; Dechat et al. 2007). B-type lamins also play a role in the formation of the mitotic spindle. In cultured cells, a diffuse localization of B-type lamins can be observed associated with the mitotic spindle (Georgatos et al. 1997; Maison et al. 1997; Moir et al. 2000; Beaudouin et al. 2002; Tsai et al. 2006). In frog egg extracts, a mitotic spindle can be induced to form around added DNA or on beads coated with a variety of factors important for mitotic spindle formation (Tsai and Zheng 2005). In these lysates, lamin B3, the major lamin in frog eggs, is required to organize a “spindle matrix” structure. This matrix contains membrane vesicles and proteins known to regulate the assembly and dynamic properties of the microtubules comprising the mitotic spindle (Tsai et al. 2006). Depletion of lamin B3 from the egg extracts with antibodies or the addition of dominant negative fragments of lamin B3 known to disrupt lamin assembly, also disrupt the formation of the mitotic spindle. Furthermore, lamin B3 is associated with several spindle assembly factors within the spindle matrix such as

NuMA and Nudel and it appears to play an important role in regulating the activity of these proteins during mitosis (Tsai et al. 2006; Ma et al. 2009).

The Role of Lamins in DNA Replication and Repair

Several lines of evidence suggest that lamins play a role in DNA replication. In cultured cells, sites of DNA replication can be visualized as discrete early or late replication foci in the nucleoplasm. These foci can be detected by incorporation of bromodeoxyuridine into sites of replication (Moir et al. 1994) or by immunolocalization of replication-associated proteins such as proliferating cell nuclear antigen (PCNA) (Shumaker et al. 2008). It has been shown that lamin B1 colocalizes with these replication foci during late S phase in mouse 3T3 cells (Moir et al. 1994) and that lamins A/C are present at sites of early replication in normal human fibroblasts (Kennedy et al. 2000). Furthermore, DNA replication in Xenopus nuclei assembled in vitro is inhibited by depletion of lamin B3 or by the addition of a dominant-negative fragment of lamin B3, which drives the disassembly of the endogenous lamin network (Lopez-Soler et al. 2001; Shumaker et al. 2005; Shumaker et al. 2008). There is evidence that the lamins play a direct role in regulating replication by their binding to DNA replication factors such as PCNA. The lamin binding site for PCNA resides within the highly conserved Ig-fold motif located in the carboxyl terminus of both the A- and B-type lamins (Shumaker et al. 2008). In addition, there is a close association between lamin B3 and PCNA at the surface of sperm head chromatin during the earliest stages of nuclear assembly in Xenopus interphase egg extracts (Shumaker et al. 2008). Nuclear lamins have also been implicated in DNA repair, although the mechanistic basis of this remains unclear. For example, the expression of disease-causing mutant lamins impairs the formation of DNA repair foci (Liu et al. 2005; Manju et al. 2006). Furthermore, genetic instability because of defects in telomere

T. Dechat et al.

function and DNA repair has been implicated in progeria, the premature aging syndrome (Gonzalez-Suarez et al. 2009a; Gonzalez-Suarez et al. 2009b).

The Role of Lamins in Transcription

Several lines of experimental evidence support the possibility that lamins play a role in transcription. An amino-terminally deleted dominant negative lamin A, which disassembles lamin networks, specifically inhibits RNA pol II activity in hamster cells or in nuclei isolated from Xenopus embryos (Spann et al. 2002). In addition, over-expression of lamins A/C or silencing of lamin B1 leads to a significant inhibition of pol II transcription in HeLa cells (Kumaran and Spector 2008; Shimi et al. 2008; Tang et al. 2008). Lamins also associate with several transcription factors leading to the suggestion that they are involved in specific regulatory pathways (Heessen and Fornerod 2007; Andres and Gonzalez 2009). For example, the association of Oct-1 with lamin B1 at the nuclear envelope appears to be important for the oxidative stress response in MEFs, as lamin B1 deficiency leads to a dysregulation of Oct-1-dependent genes and to an increase in reactive oxygen species (Malhas et al. 2009). Lamins A/C also interact with the transcription factors c-Fos, MOK2, and sterol response element binding protein 1 (SREBP1) (Dreuillet et al. 2002; Lloyd et al. 2002; Ivorra et al. 2006; Dreuillet et al. 2008; Harper et al. 2009). In the case of c-Fos, its interaction with lamins A/C at the nuclear envelope appear to suppress AP-1 (activating protein 1) binding to DNA and transcriptional activity in an extracellular signal-regulated kinase (ERK) 1/2 activity-dependent fashion (Ivorra et al. 2006; Gonzalez et al. 2008). In addition to regulating transcription factor function by direct interaction, lamins are also associated with transcription factors indirectly via several lamin-binding proteins including emerin, LAP2b and pRb. Although the evidence supporting a role for the lamins in regulating transcription is convincing, it remains unclear

whether this regulation involves direct or indirect binding to transcription factors. Lamins as Regulators of the Positioning and Organization of Chromatin

The positioning of genes near or in contact with the nuclear lamina may be a mechanism for modulating gene expression. Inactive genes and gene-poor chromatin are frequently found in association with the lamina region, which suggests that the nuclear lamina may be a transcriptionally silent microdomain (Boyle et al. 2001; Cremer et al. 2001). Furthermore, lamins may play a role in establishing this microdomain by directly interacting with chromatin. The interactions of lamins with chromatin/ DNA at the nuclear lamina can be identified in more detail by DamID labeling (Pickersgill et al. 2006). In this technique, an expression vector encoding a potential chromatin/DNA binding protein fused to the Escherichia coli enzyme DNA adenine methyl transferase (Dam) is expressed in cells. DNA methylated by the lamin Dam fusion protein is then analyzed by sequencing. In Drosophila Kc cells, lamin Dm0 seems to mainly associate with transcriptionally inactive, mid-to-late replicating genome regions, lacking active histone marks and enriched in large intergenic regions (Pickersgill et al. 2006). In human lung fibroblasts lamin B1-associated domains (LADs) are present along chromosomes in a distinct pattern interspersed with “lamin B1 poor” regions (Guelen et al. 2008). These LADs are 1 Mb in size, are mostly heterochromatic. Genes present within LADs are 5–10-fold less active than genes outside LADs. The lamina appears to have gene or chromatin-specific effects on activation or repression of transcription. Evidence for the lamina constituting a repressive environment for transcription comes from studies localizing inactive gene loci to the nuclear periphery (Kosak et al. 2002; Zink et al. 2004; Williams et al. 2006). For example, the inactive Ig-heavy and Ig-k loci are preferentially positioned at the nuclear periphery in non-B-cell lineages, but are centrally located in B-cell nuclei where

Nuclear Lamins

they are actively engaged in transcription (Kosak et al. 2002). In addition, transcriptionally inactive testis-specific gene clusters are frequently associated with lamin Dm0 in the lamina of Drosophila S2 cells. Upon downregulation of lamin Dm0, these gene clusters relocate from the lamina to the nuclear interior and become transcriptionally active (Shevelyov et al. 2009). Artificial tethering of a reporter gene to the nuclear envelope using the membrane spanning domain of the inner nuclear membrane protein emerin results in the transcriptional inactivation of the gene in mouse cells (Reddy et al. 2008). However, the relocalization of chromosomes from the nuclear interior to the nuclear periphery using the inner membrane protein Lap2b inactivates some genes, but not others (Finlan et al. 2008). The idea that the nuclear lamina is not repressive to all transcription is further supported by the finding that targeting a genetic locus to the nuclear lamina by artificial tethering to lamin B1 does not interfere with its transcriptional activation (Kumaran and Spector 2008). In addition, lamin A can act as a transcriptional repressor in mammalian cells and in yeast when artificially targeted to specific promoters (Lee et al. 2009). This latter finding is intriguing because yeast do not contain lamin genes. These somewhat conflicting results demonstrate that more research is required to determine the role of lamins in transcription both in the region of the lamina and throughout the nucleoplasm. Lamins are Involved in the Epigenetic Regulation of Chromatin

The dramatic loss of peripheral heterochromatin in the nuclei of cells expressing one of the mutant forms of lamin A, progerin, that causes the premature aging disorder HGPS emphasizes the important role of lamins in the modification and/or the organization of chromatin (Shumaker et al. 2006) (Fig. 7). A partial loss of peripheral heterochromatin is also seen in LMNA 2/2 MEFs (Sullivan et al. 1999; Nikolova et al. 2004). Furthermore, down-regulation of lamin B1 in HeLa cells results in lamin A/C-rich and lamin B2 deficient nuclear blebs

or microdomains characterized by the absence of heterochromatin (Shimi et al. 2008). These changes in chromatin organization related to lamin expression are reflected in alterations in histone modifications including reductions in trimethylated H3K9, trimethylated H3K27 and an increase in trimethylation of H4K20 in HGPS cells (Scaffidi and Misteli 2006; Shumaker et al. 2006; Shimi et al. 2008) (Fig. 7). These studies demonstrate that the interaction of chromatin with lamins, whether direct or indirect, has strong effects on the epigenetic modification of histones, and that laminassociated microdomains in the nucleus might regulate modification of chromatin (Pegoraro et al. 2009). One potential link between lamins and chromatin modification may involve the tumor suppressor ING1. This protein, which interacts with histone acetyltransferases and histone deacetylases, appears to be stabilized and targeted to the nucleus by its interaction with A-type lamins (Han et al. 2008). HGPS cells have reduced ING1 levels and the expression of ING1 protein lacking its lamin interacting domain causes a loss of peripheral heterochromatin in normal cells. Lamins and Interphase Chromosome Organization

The organization of chromosome territories and domains is influenced by the expression of lamins. For example, in LMNB D/D MEFs and in cells derived from patients suffering from some types of laminopathies, gene-poor chromosome 18 is positioned away from its normal location at the nuclear periphery toward the nuclear interior (Malhas et al. 2007; Meaburn et al. 2007). Furthermore, in HeLa cells silenced for the expression of lamin B1, the lamin A/C rich and lamin B2-deficient nuclear envelope blebs that form are associated predominantly with gene-rich chromosome regions (Shimi et al. 2008) (Fig. 8). Together, these findings suggest that A-type lamins are preferentially associated with gene-rich chromatin regions and B-type lamins are preferentially associated with gene-poor regions of chromosomes. It must be emphasized that these studies are of a

T. Dechat et al. Figure 7. The association of the nuclear lamina with

chromatin in normal and progeria patients’ cells. (A) Thin section electron micrograph of a late passage blebbed nucleus in an HGPS patient’s skin fibroblast (left and center panels) and normal human foreskin fibroblasts (right panel). A high-magnification view of the nuclear envelope in a normal human foreskin fibroblast shows a normal array of heterochromatin adjacent to the nuclear envelope, making any lamina structure difficult to detect (right panel). A higher-magnification view of a HGPS cell showing a loss of peripheral heterochromatin and a prominent electron-dense lamina region associated with the inner nuclear envelope membrane (the nucleus is to the left in center and right panels) (Goldman et al. 2004). (B) Alterations of histone methylation patterns in HGPS fibroblasts. Normal and HGPS fibroblasts from female donors were double-labeled with antibodies against lamins A/C (red) and trimethylated Lys 9 in histone H3 (H3K9me3), Lys 27 in histone H3 (H3K27me3), or Lys 20 in histone H4 (H4K20me3) (all green). Note the decrease of H3K9me3 and H3K27me3 and the increase of H4K20me3 in the lobulated HGPS nuclei compared with normal nuclei. The decrease in H3K27me3 is best observed at the inactive X chromosome, which is normally enriched in this histone modification (see arrowhead in center left panel). Bars, 10 mm. Reprinted, with permission, from Dechat et al. (2008b).

preliminary nature and that much more work is required to establish the precise relationships between the different types of lamins and gene rich/gene poor chromosome domains within the interphase nucleus. Lamins may also play a role in the localization and function of centromeres and telomeres in the nucleus. Centromeres are typically arranged near the nuclear periphery in an interphase nucleus (Solovei et al. 2004). However, the lamin A/C-rich, lamin B2-deficient nuclear envelope blebs that form in lamin B1-silenced cells are devoid of centromeres, suggesting that B-type lamins are involved in anchoring these

heterochromatic structures at the nuclear lamina (Shimi et al. 2008). Telomeres are also influenced by the composition and structure of the lamina, because in LMNA 2/2 MEFs, the distribution, length and structure of telomeres is altered leading to increased genomic instability compared to control MEFs (Gonzalez-Suarez et al. 2009a). Human fibroblasts ectopically expressing HGPS lamin A mutant proteins show rapid telomere shortening and an accelerated replicative senescence phenotype (McClintock et al. 2006; Huang et al. 2008). Furthermore, the proliferative defects seen in normal dermal fibroblasts expressing progerin can be overcome

Nuclear Lamins

Figure 8. Localization of specific gene rich chromosomal regions in nuclear blebs in LB1-silenced HeLa cells. Chromosomes are detected by fluorescence in situ hybridization. DNA is counterstained with Hoechst 33258 (blue). Chromosome 6p (or 19), and chromosome 6q (or 18) are shown in green and red, respectively. Bars, 5 mm. Adapted, with permission, from Shimi et al. (2008).

by expression of the catalytic subunit of telomerase or the inactivation of p53 (Kudlow et al. 2008). Other support for the involvement of lamins in telomere function comes from the finding that the D4Z4 human subtelomeric repeat can localize an adjacent telomere to the nuclear periphery probably by interacting with A-type lamins and CTCF (CCCTC-binding factor) (Ottaviani et al. 2009a; Ottaviani et al. 2009b). The D4Z4 repeat acts as a CTCF and A-type lamin-dependent transcriptional insulator, suggesting that nuclear lamin architecture may organize specific regions of chromatin and influence gene expression (Ottaviani et al. 2009a). Strong support for the role of the lamins in interphase chromosome organization comes from a study of cells obtained from patients with a form of atypical HGPS (Taimen et al. 2009b). Unlike the most common form of HGPS, which results in only defective lamin A, these patients have a mutation resulting in the expression of a point mutation, E145K, in the 1B segment of the central rod domain of both lamins A and C (see Fig. 1). The cells from these patients have a flower-shaped nucleus with centrally clustered centromeres and abnormal peripherally displaced telomeres (Fig. 9). This abnormal configuration of chromosomes requires a round of cell division and persists throughout interphase. The multilobulated

Figure 9. Mislocalization of heterochromatin and centromeres in HGPS E145K cells. Control and E145K patient cells were stained with anti-lamin A (red), CREST antiserum (green), and Hoechst (white). Maximum projections of series of z-sections spanning the entire nucleus and side projections are shown. Centromeres are clustered in the central region of the nucleus in E145K cells, while they are either closely associated with the peripheral lamina region or elsewhere in the nucleoplasm in control cells. Inset shows an example of the close association of one centromere with the lamina in a single confocal section. In the right hand panels, side projections from merged images are shown. Centromeres are distributed throughout the nuclei in normal cells. In E145K cells centromeres are clustered in the middle of one region of the nucleus (Taimen et al. 2009a).

nuclei are therefore unlikely to correctly establish normal interphase chromosome territory arrangements. Both the abnormal nuclear shapes and chromosome configurations in E145K cells appear to be due to the aberrant assembly of the nuclear lamina by the mutant lamin A. In vitro analysis of E145K lamin A polymerization shows that this mutant lamin forms disorganized higher-order structures, a defect not seen in the more common form of progeria resulting from a truncation near the carboxyl terminus (Taimen et al. 2009b). Lamins in Cell Proliferation and Differentiation

Deficiencies in A-type lamins or the expression of mutant A-type lamins both lead to an array of proliferative defects because of cell cycle

T. Dechat et al.

acceleration, cell cycle arrest and/or premature senescence (Verstraeten et al. 2007; Dechat et al. 2008a). There is evidence that lamins A/ C are involved in the regulation of the pRb/ E2F pathway responsible for the G1/S cell cycle transition. One current model suggests that pRb in its suppressive hypophosphorylated state is in a complex with lamins A/C and LAP2a, and that the loss of lamins A/C leads to the degradation of pRb (Ozaki et al. 1994; Markiewicz et al. 2002; Johnson et al. 2004). In addition to pRb, cyclin D3, another critical regulator of G1 progression, interacts directly with lamins A/C (Mariappan et al. 2007). The findings that lamins are involved in the regulation of cell cycle progression and that they are differentially expressed during development, suggest that they also function in the regulation of cell differentiation. In support of this, during myoblast differentiation there is a decrease in the soluble nucleoplasmic lamin A/C pool, most likely in a pRb and cyclin D3dependent manner (Muralikrishna et al. 2001; Mariappan and Parnaik 2005). Furthermore, the down-regulation of lamins A/C or the expression of lamin A/C mutants associated with Autosomal Dominant Emery Dreifuss Muscular Dystrophy (AD-EDMD) lead to impaired myoblast differentiation possibly by disrupting the Rb-MyoD pathway (Bakay et al. 2006; Frock et al. 2006). As further evidence for the importance of the lamin A/pRb interaction, the expression of either an AD-EDMD lamin A mutant protein or the HGPS lamin A mutant protein, progerin, results in the inhibition of the phosphorylation of pRb (Markiewicz et al. 2005; Dechat et al. 2007). In the case of the AD-EDMD mutation, a further decrease in pRb levels along with the failure to hyperphosphorylate the available pRB inhibits myoblast differentiation (Favreau et al. 2004; Kandert et al. 2009). In adipocyte differentiation, the overexpression of either wild-type lamin A or a lamin A mutant associated with familial partial lipodystrophy impairs the potential of mouse fibroblasts to differentiate into mature adipocytes; and LMNA 2/2 mouse embryonic fibroblasts differentiate more readily into fat-containing cells compared to control cells

(Boguslavsky et al. 2006). There is also evidence that lamins A/C regulate adipocyte differentiation in a complex with the inner nuclear membrane protein emerin, by influencing the nucleocytoplasmic distribution of b-catenin (Tilgner et al. 2009). In addition, lamins A/C appear to be involved in osteoblast differentiation, as silencing their expression causes impaired osteoblastogenesis and accelerated osteoclastogenesis in human bone marrow stromal cells (Akter et al. 2009; Rauner et al. 2009). Lamin A may also be involved in adult stem cell differentiation as the expression of progerin causes defects mouse stem cell populations (Espada et al. 2008) in the differentiation potential of human mesenchymal stem cells, probably by affecting the Notch-signaling pathway (Espada et al. 2008; Scaffidi and Misteli 2008). Another aspect of the involvement of the lamins in differentiation is related to aging. In this regard, progerin can be detected in cells from healthy individuals and the protein appears to accumulate with increasing age (McClintock et al. 2006; Scaffidi and Misteli 2006; Cao et al. 2007; Rodriguez et al. 2009). Furthermore, dermal fibroblasts from older individuals (Scaffidi and Misteli 2006), and cells in aged C. elegans (Haithcock et al. 2005) display changes in nuclear shape similar to those found in premature aging. The Lamins are Connected to the Cytoskeleton

There is evidence that the nuclear lamins are connected to the cytoskeleton through a complex of proteins called the LINC complex (linker of nucleoskeleton and cytoskeleton) (Crisp et al. 2006; Ketema et al. 2007; StewartHutchinson et al. 2008; Burke and Roux 2009). In this complex the integral proteins of the inner nuclear membrane, Sun1 and Sun2, interact with the outer nuclear membrane proteins, nesprin-1, nesprin-2, and nesprin-3a,in the luminal space between the inner and outer nuclear membranes. Although the Sun proteins bind to the lamins (Crisp et al. 2006), the nesprins are associated with both the microfilament and intermediate filament cytoskeletons

Nuclear Lamins

via direct actin binding and plectin (Tzur et al. 2006). Another lamin binding protein of the inner nuclear membrane, emerin, binds directly to nesprin isoforms, providing additional linkages between the lamins and the cytoskeleton (Wheeler et al. 2007; Zhang et al. 2007). These interactions between the nuclear lamina and the cytoskeleton may have important functional significance. For example, nuclei in fibroblasts deficient in lamin B1 rotate at a higher frequency than nuclei in control cells (Ji et al. 2007). LMNA 2/2 MEFs, on the other hand, are reported to show a decrease in mechanical stiffness, defects in mechanotransduction resulting in impaired strain-induced signaling, defects in cell polarization and cell migration, and a disturbed organization of microfilaments, vimentin IF networks and microtubules (Houben et al. 2007; Lee et al. 2007; Dahl et al. 2008; Houben et al. 2009). CONCLUSIONS AND FUTURE PERSPECTIVES

Over the past decade it has become clear that the nuclear lamins are one of the key players in determining nuclear architecture and function. There is growing evidence that these Type V IF proteins provide a filamentous scaffold built from interconnecting A- and B-type lamin networks that pervade the entire nucleus. This scaffold not only determines the shape and mechanical properties of the nucleus, but also serves as a docking site for chromatin and for numerous proteins involved in chromatin organization and various nuclear functions. Besides the extensive lamin structures located within the lamina, smaller and more dynamic lamin polymers appear to be components of large protein complexes known to be involved in a wide range of nuclear housekeeping functions such as DNA replication, DNA repair and RNA PolII transcription. We have learned much about the nuclear lamins from the analysis of the changes in nuclear form and function that take place in cells from patients suffering from the remarkably large number of diseases attributable to hundreds of mutations in the LMNA gene.

However, the precise roles that normal lamins play in nuclear functions remain largely unknown. To gain more insights into their specific functions we need a better understanding of their normal structure at the highest levels of resolution. To date only a few subdomains of the lamins have been shown to be amenable to analysis by X-ray diffraction, because of the well known difficulties inherent in crystallizing IF proteins. Other challenges for future studies include determining the steps in lamin polymerization and depolymerization in vivo and determining how complex networks of the Aand B-type lamins are assembled and interact with each other both at the nuclear periphery and within the nucleoplasm throughout the cell cycle. Finally, it will be of great interest to determine the different roles of the A- and B-type lamins in determining the overall architecture of the nucleus and the mechanisms involved in their linkages to other nuclear structures, such as the nuclear membrane and pore complexes.

ACKNOWLEDGMENTS

Work in the Goldman lab on nuclear lamins is funded by the National Institute on Aging and the National Cancer Institute. PT received a fellowship from Sigrid Juselius Foundation, Orion-Farmos Research Foundation, Cancer Society of Southwestern Finland, and Finnish Cultural Foundation.

REFERENCES Aaronson RP, Blobel G. 1975. Isolation of nuclear pore complexes in association with a lamina. Proc Natl Acad Sci 72: 1007–1011. Adolph KW. 1987. ADPribosylation of nuclear proteins labeled with [3H]adenosine: changes during the HeLa cycle. Biochim Biophys Acta 909: 222– 230. Aebi U, Cohn J, Buhle L, Gerace L. 1986. The nuclear lamina is a meshwork of intermediate-type filaments. Nature 323: 560–564. Akter R, Rivas D, Geneau G, Drissi H, Duque G. 2009. Effect of lamin A/C knockdown on osteoblast differentiation and function. J Bone Miner Res 24: 283 –293. Andres V, Gonzalez JM. 2009. Role of A-type lamins in signaling, transcription, and chromatin organization. J Cell Biol 187: 945 –957.

T. Dechat et al. Bakay M, Wang Z, Melcon G, Schiltz L, Xuan J, Zhao P, Sartorelli V, Seo J, Pegoraro E, Angelini C, et al. 2006. Nuclear envelope dystrophies show a transcriptional fingerprint suggesting disruption of Rb-MyoD pathways in muscle regeneration. Brain 129: 996– 1013. Baricheva EA, Berrios M, Bogachev SS, Borisevich IV, Lapik ER, Sharakhov IV, Stuurman N, Fisher PA. 1996. DNA from Drosophila melanogaster b-heterochromatin binds specifically to nuclear lamins in vitro and the nuclear envelope in situ. Gene 171: 171– 176. Barrowman J, Hamblet C, George CM, Michaelis S. 2008. Analysis of prelamin A biogenesis reveals the nucleus to be a CaaX processing compartment. Mol Biol Cell 19: 5398– 5408. Beaudouin J, Gerlich D, Daigle N, Eils R, Ellenberg J. 2002. Nuclear envelope breakdown proceeds by microtubule-induced tearing of the lamina. Cell 108: 83–96. Bechert K, Lagos-Quintana M, Harborth J, Weber K, Osborn M. 2003. Effects of expressing lamin A mutant protein causing Emery-Dreifuss muscular dystrophy and familial partial lipodystrophy in HeLa cells. Exp Cell Res 286: 75–86. Beck LA, Hosick TJ, Sinensky M. 1990. Isoprenylation is required for the processing of the lamin A precursor. J Cell Biol 110: 1489–1499. Belmont AS, Zhai Y, Thilenius A. 1993. Lamin B distribution and association with peripheral chromatin revealed by optical sectioning and electron microscopy tomography. J Cell Biol 123: 1671– 1685. Ben-Harush K, Wiesel N, Frenkiel-Krispin D, Moeller D, Soreq E, Aebi U, Herrmann H, Gruenbaum Y, Medalia O. 2009. The supramolecular organization of the C elegans nuclear lamin filament. J Mol Biol 386: 1392– 1402. Benavente R, Krohne G. 1986. Involvement of nuclear lamins in postmitotic reorganization of chromatin as demonstrated by microinjection of lamin antibodies. J Cell Biol 103: 1847– 1854. Benavente R, Krohne G, Franke WW. 1985. Cell typespecific expression of nuclear lamina proteins during development of Xenopus laevis. Cell 41: 177– 190. Boguslavsky RL, Stewart CL, Worman HJ. 2006. Nuclear lamin A inhibits adipocyte differentiation: implications for Dunnigan-type familial partial lipodystrophy. Hum Mol Genet 15: 653– 663. Boyartchuk VL, Ashby MN, Rine J. 1997. Modulation of Ras and a-factor function by carboxyl-terminal proteolysis. Science 275: 1796–1800. Boyle S, Gilchrist S, Bridger JM, Mahy NL, Ellis JA, Bickmore WA. 2001. The spatial organization of human chromosomes within the nuclei of normal and emerinmutant cells. Hum Mol Genet 10: 211– 219. Broers JL, Machiels BM, van Eys GJ, Kuijpers HJ, Manders EM, van Driel R, Ramaekers FC. 1999. Dynamics of the nuclear lamina as monitored by GFP-tagged A-type lamins. J Cell Sci 112: 3463–3475. Burke B. 1990. On the cell-free association of lamins A and C with metaphase chromosomes. Exp Cell Res 186: 169–176. Burke B, Roux KJ. 2009. Nuclei take a position: Managing nuclear location. Dev Cell 17: 587–597.

Cao K, Capell BC, Erdos MR, Djabali K, Collins FS. 2007. A lamin A protein isoform overexpressed in HutchinsonGilford progeria syndrome interferes with mitosis in progeria and normal cells. Proc Natl Acad Sci 104: 4949–4954. Cohen M, Lee KK, Wilson KL, Gruenbaum Y. 2001. Transcriptional repression, apoptosis, human disease and the functional evolution of the nuclear lamina. Trends Biochem Sci 26: 41–47. Collas P. 1999. Sequential PKC- and Cdc2-mediated phosphorylation events elicit zebrafish nuclear envelope disassembly. J Cell Sci 112: 977–987. Constantinescu D, Gray HL, Sammak PJ, Schatten GP, Csoka AB. 2006. Lamin A/C expression is a marker of mouse and human embryonic stem cell differentiation. Stem Cells 24: 177–185. Corrigan DP, Kuszczak D, Rusinol AE, Thewke DP, Hrycyna CA, Michaelis S, Sinensky MS. 2005. Prelamin A endoproteolytic processing in vitro by recombinant Zmpste24. Biochem J 387: 129– 138. Cremer M, von Hase J, Volm T, Brero A, Kreth G, Walter J, Fischer C, Solovei I, Cremer C, Cremer T. 2001. Nonrandom radial higher-order chromatin arrangements in nuclei of diploid human cells. Chromosome Res 9: 541 –567. Crisp M, Liu Q, Roux K, Rattner JB, Shanahan C, Burke B, Stahl PD, Hodzic D. 2006. Coupling of the nucleus and cytoplasm: Role of the LINC complex. J Cell Biol 172: 41– 53. Dahl KN, Ribeiro AJ, Lammerding J. 2008. Nuclear shape, mechanics, and mechanotransduction. Circ Res 102: 1307–1318. Dahl KN, Scaffidi P, Islam MF, Yodh AG, Wilson KL, Misteli T. 2006. Distinct structural and mechanical properties of the nuclear lamina in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci 103: 10271– 10276. Davies BS, Fong LG, Yang SH, Coffinier C, Young SG. 2009. The posttranslational processing of prelamin A and disease. Annu Rev Genomics Hum Genet 10: 153– 174. Dechat T, Adam SA, Goldman RD. 2008a. Nuclear lamins and chromatin: When structure meets function. Adv Enzyme Regul 49: 157 –166. Dechat T, Gajewski A, Korbei B, Gerlich D, Daigle N, Haraguchi T, Furukawa K, Ellenberg J, Foisner R. 2004. LAP2a and BAF transiently localize to telomeres and specific regions on chromatin during nuclear assembly. J Cell Sci 117: 6117– 6128. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008b. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832– 853. Dechat T, Shimi T, Adam SA, Rusinol AE, Andres DA, Spielmann HP, Sinensky MS, Goldman RD. 2007. Alterations in mitosis and cell cycle progression caused by a mutant lamin A known to accelerate human aging. Proc Natl Acad Sci 104: 4955– 4960. Delbarre E, Tramier M, Coppey-Moisan M, Gaillard C, Courvalin JC, Buendia B. 2006. The truncated prelamin A in Hutchinson-Gilford progeria syndrome alters segregation of A-type and B-type lamin homopolymers. Hum Mol Genet 15: 1113– 1122.

Nuclear Lamins Dessev G, Goldman R. 1988. Meiotic breakdown of nuclear envelope in oocytes of Spisula solidissima involves phosphorylation and release of nuclear lamin. Dev Biol 130: 543–550. Dessev G, Iovcheva-Dessev C, Bischoff JR, Beach D, Goldman R. 1991. A complex containing p34cdc2 and cyclin B phosphorylates the nuclear lamin and disassembles nuclei of clam oocytes in vitro. J Cell Biol 112: 523 –533. Dessev G, Iovcheva C, Tasheva B, Goldman R. 1988. Protein kinase activity associated with the nuclear lamina. Proc Natl Acad Sci 85: 2994– 2998. Dessev G, Palazzo R, Rebhun L, Goldman R. 1989. Disassembly of the nuclear envelope of spisula oocytes in a cellfree system. Dev Biol 131: 496– 504. Dhe-Paganon S, Werner ED, Chi YI, Shoelson SE. 2002. Structure of the globular tail of nuclear lamin. J Biol Chem 277: 17381– 17384. Dreuillet C, Harper M, Tillit J, Kress M, Ernoult-Lange M. 2008. Mislocalization of human transcription factor MOK2 in the presence of pathogenic mutations of lamin A/C. Biol Cell 100: 51– 61. Dreuillet C, Tillit J, Kress M, Ernoult-Lange M. 2002. In vivo and in vitro interaction between human transcription factor MOK2 and nuclear lamin A/C. Nucleic Acids Res 30: 4634–4642. Espada J, Varela I, Flores I, Ugalde AP, Cadinanos J, Pendas AM, Stewart CL, Tryggvason K, Blasco MA, Freije JM, et al. 2008. Nuclear envelope defects cause stem cell dysfunction in premature-aging mice. J Cell Biol 181: 27– 35. Farnsworth CC, Gelb MH, Glomset JA. 1990. Identification of geranylgeranyl-modified proteins in HeLa cells. Science 247: 320– 322. Favreau C, Dubosclard E, Ostlund C, Vigouroux C, Capeau J, Wehnert M, Higuet D, Worman HJ, Courvalin JC, Buendia B. 2003. Expression of lamin A mutated in the carboxyl-terminal tail generates an aberrant nuclear phenotype similar to that observed in cells from patients with Dunnigan-type partial lipodystrophy and EmeryDreifuss muscular dystrophy. Exp Cell Res 282: 14– 23. Favreau C, Higuet D, Courvalin JC, Buendia B. 2004. Expression of a mutant lamin A that causes EmeryDreifuss muscular dystrophy inhibits in vitro differentiation of C2C12 myoblasts. Mol Cell Biol 24: 1481–1492. Fawcett DW. 1966. On the occurrence of a fibrous lamina on the inner aspect of the nuclear envelope in certain cells of vertebrates. Am J Anat 119: 129 –145. Ferraro A, Eufemi M, Cervoni L, Marinetti R, Turano C. 1989. Glycosylated forms of nuclear lamins. FEBS Lett 257: 241– 246. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Frasch M, Paddy M, Saumweber H. 1988. Developmental and mitotic behaviour of two novel groups of nuclear envelope antigens of Drosophila melanogaster. J Cell Sci 90: 247 –263. Frock RL, Kudlow BA, Evans AM, Jameson SA, Hauschka SD, Kennedy BK. 2006. Lamin A/C and emerin are critical for skeletal muscle satellite cell differentiation. Genes Dev 20: 486 –500.

Furukawa K, Hotta Y. 1993. cDNA cloning of a germ cell specific lamin B3 from mouse spermatocytes and analysis of its function by ectopic expression in somatic cells. Embo J 12: 97–106. Galiova G, Bartova E, Raska I, Krejci J, Kozubek S. 2008. Chromatin changes induced by lamin A/C deficiency and the histone deacetylase inhibitor trichostatin A. Eur J Cell Biol 87: 291– 303. Georgatos SD, Pyrpasopoulou A, Theodoropoulos PA. 1997. Nuclear envelope breakdown in mammalian cells involves stepwise lamina disassembly and microtubuledrive deformation of the nuclear membrane. J Cell Sci 110: 2129– 2140. Gerace L, Blobel G. 1980. The nuclear envelope lamina is reversibly depolymerized during mitosis. Cell 19: 277 –287. Glass JR, Gerace L. 1990. Lamins A and C bind and assemble at the surface of mitotic chromosomes. J Cell Biol 111: 1047–1057. Goldberg M, Harel A, Brandeis M, Rechsteiner T, Richmond TJ, Weiss AM, Gruenbaum Y. 1999. The tail domain of lamin Dm0 binds histones H2A and H2B. Proc Natl Acad Sci 96: 2852–2857. Goldberg MW, Fiserova J, Huttenlauch I, Stick R. 2008a. A new model for nuclear lamina organization. Biochem Soc Trans 36: 1339– 1343. Goldberg MW, Huttenlauch I, Hutchison CJ, Stick R. 2008b. Filaments made from A- and B-type lamins differ in structure and organization. J Cell Sci 121: 215 –225. Goldman RD, Grin B, Mendez MG, Kuczmarski ER. 2008. Intermediate filaments: versatile building blocks of cell structure. Curr Opin Cell Biol 20: 28–34. Goldman AE, Maul G, Steinert PM, Yang HY, Goldman RD. 1986. Keratin-like proteins that coisolate with intermediate filaments of BHK-21 cells are nuclear lamins. Proc Natl Acad Sci 83: 3839– 3843. Goldman AE, Moir RD, Montag-Lowy M, Stewart M, Goldman RD. 1992. Pathway of incorporation of microinjected lamin A into the nuclear envelope. J Cell Biol 119: 725–735. Goldman RD, Shumaker DK, Erdos MR, Eriksson M, Goldman AE, Gordon LB, Gruenbaum Y, Khuon S, Mendez M, Varga R, et al. 2004. Accumulation of mutant lamin A causes progressive changes in nuclear architecture in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci 101: 8963– 8968. Gonzalez JM, Navarro-Puche A, Casar B, Crespo P, Andres V. 2008. Fast regulation of AP-1 activity through interaction of lamin A/C, ERK1/2, and c-Fos at the nuclear envelope. J Cell Biol 183: 653–666. Gonzalez-Suarez I, Redwood AB, Gonzalo S. 2009a. Loss of A-type lamins and genomic instability. Cell Cycle 8: 3860–3865. Gonzalez-Suarez I, Redwood AB, Perkins SM, Vermolen B, Lichtensztejin D, Grotsky DA, Morgado-Palacin L, Gapud EJ, Sleckman BP, Sullivan T, et al. 2009b. Novel roles for A-type lamins in telomere biology and the DNA damage response pathway. Embo J 28: 2414–2427. Goss VL, Hocevar BA, Thompson LJ, Stratton CA, Burns DJ, Fields AP. 1994. Identification of nuclear b II protein

T. Dechat et al. kinase C as a mitotic lamin kinase. J Biol Chem 269: 19074–19080. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W, et al. 2008. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453: 948–951. Guilly MN, Kolb JP, Gosti F, Godeau F, Courvalin JC. 1990. Lamins A and C are not expressed at early stages of human lymphocyte differentiation. Exp Cell Res 189: 145–147. Haithcock E, Dayani Y, Neufeld E, Zahand AJ, Feinstein N, Mattout A, Gruenbaum Y, Liu J. 2005. Age-related changes of nuclear architecture in Caenorhabditis elegans. Proc Natl Acad Sci USA 102: 16690–16695. Han X, Feng X, Rattner JB, Smith H, Bose P, Suzuki K, Soliman MA, Scott MS, Burke BE, Riabowol K. 2008. Tethering by lamin A stabilizes and targets the ING1 tumour suppressor. Nat Cell Biol 10: 1333–1340. Haraguchi T, Kojidani T, Koujin T, Shimi T, Osakada H, Mori C, Yamamoto A, Hiraoka Y. 2008. Live cell imaging and electron microscopy reveal dynamic processes of BAF-directed nuclear envelope assembly. J Cell Sci 121: 2540– 2554. Harper M, Tillit J, Kress M, Ernoult-Lange M. 2009. Phosphorylation-dependent binding of human transcription factor MOK2 to lamin A/C. Febs J 276: 3137– 3147. Heald R, McKeon F. 1990. Mutations of phosphorylation sites in lamin A that prevent nuclear lamina disassembly in mitosis. Cell 61: 579– 589. Heessen S, Fornerod M. 2007. The inner nuclear envelope as a transcription factor resting place. EMBO Rep 8: 914–919. Heitlinger E, Peter M, Haner M, Lustig A, Aebi U, Nigg EA. 1991. Expression of chicken lamin B2 in Escherichia coli: Characterization of its structure, assembly, and molecular interactions. J Cell Biol 113: 485 –495. Hennekes H, Peter M, Weber K, Nigg EA. 1993. Phosphorylation on protein kinase C sites inhibits nuclear import of lamin B2. J Cell Biol 120: 1293– 1304. Houben F, Ramaekers FC, Snoeckx LH, Broers JL. 2007. Role of nuclear lamina-cytoskeleton interactions in the maintenance of cellular strength. Biochim Biophys Acta 1773: 675–686. Houben F, Willems CH, Declercq IL, Hochstenbach K, Kamps MA, Snoeckx LH, Ramaekers FC, Broers JL. 2009. Disturbed nuclear orientation and cellular migration in A-type lamin deficient cells. Biochim Biophys Acta 1793: 312 –324. Huang S, Risques RA, Martin GM, Rabinovitch PS, Oshima J. 2008. Accelerated telomere shortening and replicative senescence in human fibroblasts overexpressing mutant and wild-type lamin A. Exp Cell Res 314: 82–91. Ito H, Koyama Y, Takano M, Ishii K, Maeno M, Furukawa K, Horigome T. 2007. Nuclear envelope precursor vesicle targeting to chromatin is stimulated by protein phosphatase 1 in Xenopus egg extracts. Exp Cell Res 313: 1897– 1910. Ivorra C, Kubicek M, Gonzalez JM, Sanz-Gonzalez SM, Alvarez-Barrientos A, O’Connor JE, Burke B, Andres V.

2006. A mechanism of AP-1 suppression through interaction of c-Fos with lamin A/C. Genes Dev 20: 307–320. Ji JY, Lee RT, Vergnes L, Fong LG, Stewart CL, Reue K, Young SG, Zhang Q, Shanahan CM, Lammerding J. 2007. Cell nuclei spin in the absence of lamin b1. J Biol Chem 282: 20015– 20026. Johnson BR, Nitta RT, Frock RL, Mounkes L, Barbie DA, Stewart CL, Harlow E, Kennedy BK. 2004. A-type lamins regulate retinoblastoma protein function by promoting subnuclear localization and preventing proteasomal degradation. Proc Natl Acad Sci 101: 9677–9682. Kandert S, Wehnert M, Muller CR, Buendia B, Dabauvalle MC. 2009. Impaired nuclear functions lead to increased senescence and inefficient differentiation in human myoblasts with a dominant p.R545C mutation in the LMNA gene. Eur J Cell Biol 88: 593– 608. Kennedy BK, Barbie DA, Classon M, Dyson N, Harlow E. 2000. Nuclear organization of DNA replication in primary mammalian cells. Genes Dev 14: 2855–2868. Ketema M, Wilhelmsen K, Kuikman I, Janssen H, Hodzic D, Sonnenberg A. 2007. Requirements for the localization of nesprin-3 at the nuclear envelope and its interaction with plectin. J Cell Sci 120: 3384–3394. Kilic F, Dalton MB, Burrell SK, Mayer JP, Patterson SD, Sinensky M. 1997. In vitro assay and characterization of the farnesylation-dependent prelamin A endoprotease. J Biol Chem 272: 5298–5304. Kosak ST, Skok JA, Medina KL, Riblet R, Le Beau MM, Fisher AG, Singh H. 2002. Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296: 158 –162. Krimm I, Ostlund C, Gilquin B, Couprie J, Hossenlopp P, Mornon JP, Bonne G, Courvalin JC, Worman HJ, ZinnJustin S. 2002. The Ig-like structure of the C-terminal domain of lamin A/C, mutated in muscular dystrophies, cardiomyopathy, and partial lipodystrophy. Structure 10: 811 –823. Kudlow BA, Stanfel MN, Burtner CR, Johnston ED, Kennedy BK. 2008. Suppression of proliferative defects associated with processing-defective lamin A mutants by hTERT or inactivation of p53. Mol Biol Cell 19: 5238–5248. Kumaran RI, Spector DL. 2008. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol 180: 51–65. Lee JS, Hale CM, Panorchan P, Khatau SB, George JP, Tseng Y, Stewart CL, Hodzic D, Wirtz D. 2007. Nuclear lamin A/C deficiency induces defects in cell mechanics, polarization, and migration. Biophys J 93: 2542–2552. Lee DC, Welton KL, Smith ED, Kennedy BK. 2009. A-type nuclear lamins act as transcriptional repressors when targeted to promoters. Exp Cell Res 315: 996– 1007. Lehner CF, Furstenberger G, Eppenberger HM, Nigg EA. 1986. Biogenesis of the nuclear lamina: in vivo synthesis and processing of nuclear protein precursors. Proc Natl Acad Sci 83: 2096–2099. Lehner CF, Stick R, Eppenberger HM, Nigg EA. 1987. Differential expression of nuclear lamin proteins during chicken development. J Cell Biol 105: 577– 587. Lenz-Bohme B, Wismar J, Fuchs S, Reifegerste R, Buchner E, Betz H, Schmitt B. 1997. Insertional mutation of the

Nuclear Lamins Drosophila nuclear lamin Dm0 gene results in defective nuclear envelopes, clustering of nuclear pore complexes, and accumulation of annulate lamellae. J Cell Biol 137: 1001– 1016. Leukel M, Jost E. 1995. Two conserved serines in the nuclear localization signal flanking region are involved in the nuclear targeting of human lamin A. Eur J Cell Biol 68: 133–142. Leung GK, Schmidt WK, Bergo MO, Gavino B, Wong DH, Tam A, Ashby MN, Michaelis S, Young SG. 2001. Biochemical studies of Zmpste24-deficient mice. J Biol Chem 276: 29051– 29058. Lin F, Worman HJ. 1993. Structural organization of the human gene encoding nuclear lamin A and nuclear lamin C. J Biol Chem 268: 16321–16326. Liu J, Rolef Ben-Shahar T, Riemer D, Treinin M, Spann P, Weber K, Fire A, Gruenbaum Y. 2000. Essential roles for Caenorhabditis elegans lamin gene in nuclear organization, cell cycle progression, and spatial organization of nuclear pore complexes. Mol Biol Cell 11: 3937–3947. Liu B, Wang J, Chan KM, Tjia WM, Deng W, Guan X, Huang JD, Li KM, Chau PY, Chen DJ, et al. 2005. Genomic instability in laminopathy-based premature aging. Nat Med 11: 780 –785. Lloyd DJ, Trembath RC, Shackleton S. 2002. A novel interaction between lamin A and SREBP1: Implications for partial lipodystrophy and other laminopathies. Hum Mol Genet 11: 769 –777. Loewinger L, McKeon F. 1988. Mutations in the nuclear lamin proteins resulting in their aberrant assembly in the cytoplasm. Embo J 7: 2301– 2309. Lopez-Soler RI, Moir RD, Spann TP, Stick R, Goldman RD. 2001. A role for nuclear lamins in nuclear envelope assembly. J Cell Biol 154: 61–70. Lourim D, Lin JJ. 1989. Expression of nuclear lamin A and muscle-specific proteins in differentiating muscle cells in ovo and in vitro. J Cell Biol 109: 495– 504. Luderus ME, de Graaf A, Mattia E, den Blaauwen JL, Grande MA, de Jong L, van Driel R. 1992. Binding of matrix attachment regions to lamin B1. Cell 70: 949– 959. Luderus ME, den Blaauwen JL, de Smit OJ, Compton DA, van Driel R. 1994. Binding of matrix attachment regions to lamin polymers involves single-stranded regions and the minor groove. Mol Cell Biol 14: 6297– 6305. Lutz RJ, Trujillo MA, Denham KS, Wenger L, Sinensky M. 1992. Nucleoplasmic localization of prelamin A: Implications for prenylation-dependent lamin A assembly into the nuclear lamina. Proc Natl Acad Sci 89: 3000–3004. Ma L, Tsai MY, Wang S, Lu B, Chen R, Iii JR, Zhu X, Zheng Y. 2009. Requirement for Nudel and dynein for assembly of the lamin B spindle matrix. Nat Cell Biol 11: 247– 256. Machiels BM, Broers JL, Raymond Y, de Ley L, Kuijpers HJ, Caberg NE, Ramaekers FC. 1995. Abnormal A-type lamin organization in a human lung carcinoma cell line. Eur J Cell Biol 67: 328 –335. Machiels BM, Henfling ME, Schutte B, van Engeland M, Broers JL, Ramaekers FC. 1996. Subcellular localization of proteasomes in apoptotic lung tumor cells and persistence as compared to intermediate filaments. Eur J Cell Biol 70: 250– 259.

Maison C, Pyrpasopoulou A, Theodoropoulos PA, Georgatos SD. 1997. The inner nuclear membrane protein LAP1 forms a native complex with B-type lamins and partitions with spindle-associated mitotic vesicles. Embo J 16: 4839–4850. Malhas AN, Lee CF, Vaux DJ. 2009. Lamin B1 controls oxidative stress responses via Oct-1. J Cell Biol 184: 45– 55. Malhas A, Lee CF, Sanders R, Saunders NJ, Vaux DJ. 2007. Defects in lamin B1 expression or processing affect interphase chromosome position and gene expression. J Cell Biol 176: 593 –603. Manju K, Muralikrishna B, Parnaik VK. 2006. Expression of disease-causing lamin A mutants impairs the formation of DNA repair foci. J Cell Sci 119: 2704–2714. Mariappan I, Parnaik VK. 2005. Sequestration of pRb by cyclin D3 causes intranuclear reorganization of lamin A/C during muscle cell differentiation. Mol Biol Cell 16: 1948–1960. Mariappan I, Gurung R, Thanumalayan S, Parnaik VK. 2007. Identification of cyclin D3 as a new interaction partner of lamin A/C. Biochem Biophys Res Commun 355: 981–985. Markiewicz E, Ledran M, Hutchison CJ. 2005. Remodelling of the nuclear lamina and nucleoskeleton is required for skeletal muscle differentiation in vitro. J Cell Sci 118: 409 –420. Markiewicz E, Dechat T, Foisner R, Quinlan RA, Hutchison CJ. 2002. Lamin A/C binding protein LAP2alpha is required for nuclear anchorage of retinoblastoma protein. Mol Biol Cell 13: 4401– 4413. Mattout A, Goldberg M, Tzur Y, Margalit A, Gruenbaum Y. 2007. Specific and conserved sequences in D melanogaster and C elegans lamins and histone H2A mediate the attachment of lamins to chromosomes. J Cell Sci 120: 77–85. McClintock D, Gordon LB, Djabali K. 2006. HutchinsonGilford progeria mutant lamin A primarily targets human vascular cells as detected by an anti-Lamin A G608G antibody. Proc Natl Acad Sci 103: 2154– 2159. McKeon FD, Kirschner MW, Caput D. 1986. Homologies in both primary and secondary structure between nuclear envelope and intermediate filament proteins. Nature 319: 463–468. Meaburn KJ, Cabuy E, Bonne G, Levy N, Morris GE, Novelli G, Kill IR, Bridger JM. 2007. Primary laminopathy fibroblasts display altered genome organization and apoptosis. Aging Cell 6: 139– 153. Melcer S, Gruenbaum Y, Krohne G. 2007. Invertebrate lamins. Exp Cell Res 313: 2157– 2166. Moir RD, Montag-Lowy M, Goldman RD. 1994. Dynamic properties of nuclear lamins: lamin B is associated with sites of DNA replication. J Cell Biol 125: 1201–1212. Moir RD, Yoon M, Khuon S, Goldman RD. 2000. Nuclear lamins A and B1: Different pathways of assembly during nuclear envelope formation in living cells. J Cell Biol 151: 1155–1168. Molloy S, Little M. 1992. p34cdc2 kinase-mediated release of lamins from nuclear ghosts is inhibited by cAMPdependent protein kinase. Exp Cell Res 201: 494– 499. Muchir A, van Engelen BG, Lammens M, Mislow JM, McNally E, Schwartz K, Bonne G. 2003. Nuclear envelope

T. Dechat et al. alterations in fibroblasts from LGMD1B patients carrying nonsense Y259X heterozygous or homozygous mutation in lamin A/C gene. Exp Cell Res 291: 352 –362. Muralikrishna B, Dhawan J, Rangaraj N, Parnaik VK. 2001. Distinct changes in intranuclear lamin A/C organization during myoblast differentiation. J Cell Sci 114: 4001– 4011. Nakajima N, Abe K. 1995. Genomic structure of the mouse A-type lamin gene locus encoding somatic and germ cellspecific lamins. FEBS Lett 365: 108– 114. Nikolova V, Leimena C, McMahon AC, Tan JC, Chandar S, Jogia D, Kesteven SH, Michalicek J, Otway R, Verheyen F, et al. 2004. Defects in nuclear structure and function promote dilated cardiomyopathy in lamin A/C-deficient mice. J Clin Invest 113: 357– 369. Ostlund C, Bonne G, Schwartz K, Worman HJ. 2001. Properties of lamin A mutants found in Emery-Dreifuss muscular dystrophy, cardiomyopathy and Dunnigan-type partial lipodystrophy. J Cell Sci 114: 4435–4445. Ottaviani A, Rival-Gervier S, Boussouar A, Foerster AM, Rondier D, Sacconi S, Desnuelle C, Gilson E, Magdinier F. 2009a. The D4Z4 macrosatellite repeat acts as a CTCF and A-type lamins-dependent insulator in facio-scapulohumeral dystrophy. PLoS Genet 5: e1000394. Ottaviani A, Schluth-Bolard C, Rival-Gervier S, Boussouar A, Rondier D, Foerster AM, Morere J, Bauwens S, Gazzo S, Callet-Bauchu E, et al. 2009b. Identification of a perinuclear positioning element in human subtelomeres that requires A-type lamins and CTCF. Embo J 28: 2428– 2436. Ottaviano Y, Gerace L. 1985. Phosphorylation of the nuclear lamins during interphase and mitosis. J Biol Chem 260: 624–632. Ozaki T, Saijo M, Murakami K, Enomoto H, Taya Y, Sakiyama S. 1994. Complex formation between lamin A and the retinoblastoma gene product: Identification of the domain on lamin A required for its interaction. Oncogene 9: 2649–2653. Paddy MR, Belmont AS, Saumweber H, Agard DA, Sedat JW. 1990. Interphase nuclear envelope lamins form a discontinuous network that interacts with only a fraction of the chromatin in the nuclear periphery. Cell 62: 89–106. Pajerowski JD, Dahl KN, Zhong FL, Sammak PJ, Discher DE. 2007. Physical plasticity of the nucleus in stem cell differentiation. Proc Natl Acad Sci 104: 15619–15624. Panorchan P, Wirtz D, Tseng Y. 2004b. Structure-function relationship of biological gels revealed by multipleparticle tracking and differential interference contrast microscopy: The case of human lamin networks. Phys Rev E Stat Nonlin Soft Matter Phys 70: 041906. Panorchan P, Schafer BW, Wirtz D, Tseng Y. 2004a. Nuclear envelope breakdown requires overcoming the mechanical integrity of the nuclear lamina. J Biol Chem 279: 43462– 43467. Parry DA, Conway JF, Steinert PM. 1986. Structural studies on lamin. Similarities and differences between lamin and intermediate-filament proteins. Biochem J 238: 305 –308. Pegoraro G, Kubben N, Wickert U, Gohler H, Hoffmann K, Misteli T. 2009. Ageing-related chromatin defects through loss of the NURD complex. Nat Cell Biol 11: 1261– 1267.

Peter M, Kitten GT, Lehner CF, Vorburger K, Bailer SM, Maridor G, Nigg EA. 1989. Cloning and sequencing of cDNA clones encoding chicken lamins A and B1 and comparison of the primary structures of vertebrate Aand B-type lamins. J Mol Biol 208: 393– 404. Peter M, Nakagawa J, Doree M, Labbe JC, Nigg EA. 1990. In vitro disassembly of the nuclear lamina and M phase-specific phosphorylation of lamins by cdc2 kinase. Cell 61: 591–602. Pickersgill H, Kalverda B, de Wit E, Talhout W, Fornerod M, van Steensel B. 2006. Characterization of the Drosophila melanogaster genome at the nuclear lamina. Nat Genet 38: 1005–1014. Prather RS, Sims MM, Maul GG, First NL, Schatten G. 1989. Nuclear lamin antigens are developmentally regulated during porcine and bovine embryogenesis. Biol Reprod 41: 123– 132. Rauner M, Sipos W, Goettsch C, Wutzl A, Foisner R, Pietschmann P, Hofbauer LC. 2009. Inhibition of lamin A/C attenuates osteoblast differentiation and enhances RANKL-dependent osteoclastogenesis. J Bone Miner Res 24: 78–86. Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243 –247. Riemer D, Stuurman N, Berrios M, Hunter C, Fisher PA, Weber K. 1995. Expression of Drosophila lamin C is developmentally regulated: Analogies with vertebrate A-type lamins. J Cell Sci 108: 3189– 3198. Rober RA, Weber K, Osborn M. 1989. Differential timing of nuclear lamin A/C expression in the various organs of the mouse embryo and the young animal: A developmental study. Development 105: 365–378. Rober RA, Sauter H, Weber K, Osborn M. 1990. Cells of the cellular immune and hemopoietic system of the mouse lack lamins A/C: distinction versus other somatic cells. J Cell Sci 95: 587–598. Rodriguez S, Coppede F, Sagelius H, Eriksson M. 2009. Increased expression of the Hutchinson-Gilford progeria syndrome truncated lamin A transcript during cell aging. Eur J Hum Genet 17: 928– 937. Rusinol AE, Sinensky MS. 2006. Farnesylated lamins, progeroid syndromes and farnesyl transferase inhibitors. J Cell Sci 119: 3265– 3272. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059–1063. Scaffidi P, Misteli T. 2008. Lamin A-dependent misregulation of adult stem cells associated with accelerated ageing. Nat Cell Biol 10: 452 –459. Schape J, Prausse S, Radmacher M, Stick R. 2009. Influence of lamin A on the mechanical properties of amphibian oocyte nuclei measured by atomic force microscopy. Biophys J 96: 4319– 4325. Schatten G, Maul GG, Schatten H, Chaly N, Simerly C, Balczon R, Brown DL. 1985. Nuclear lamins and peripheral nuclear antigens during fertilization and embryogenesis in mice and sea urchins. Proc Natl Acad Sci 82: 4727– 4731. Schermelleh L, Carlton PM, Haase S, Shao L, Winoto L, Kner P, Burke B, Cardoso MC, Agard DA, Gustafsson MG, et al. 2008. Subdiffraction multicolor imaging of

Nuclear Lamins the nuclear periphery with 3D structured illumination microscopy. Science 320: 1332– 1336. Shevelyov YY, Lavrov SA, Mikhaylova LM, Nurminsky ID, Kulathinal RJ, Egorova KS, Rozovsky YM, Nurminsky DI. 2009. The B-type lamin is required for somatic repression of testis-specific gene clusters. Proc Natl Acad Sci 106: 3282–3287. Shimi T, Pfleghaar K, Kojima S, Pack CG, Solovei I, Goldman AE, Adam SA, Shumaker DK, Kinjo M, Cremer T, et al. 2008. The A- and B-type nuclear lamin networks: microdomains involved in chromatin organization and transcription. Genes Dev 22: 3409– 3421. Shumaker DK, Dechat T, Kohlmaier A, Adam SA, Bozovsky MR, Erdos MR, Eriksson M, Goldman AE, Khuon S, Collins FS, et al. 2006. Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci 103: 8703– 8708. Shumaker DK, Lopez-Soler RI, Adam SA, Herrmann H, Moir RD, Spann TP, Goldman RD. 2005. Functions and dysfunctions of the nuclear lamin Ig-fold domain in nuclear assembly, growth, and Emery-Dreifuss muscular dystrophy. Proc Natl Acad Sci USA 102: 15494–15499. Shumaker DK, Solimando L, Sengupta K, Shimi T, Adam SA, Grunwald A, Strelkov SV, Aebi U, Cardoso MC, Goldman RD. 2008. The highly conserved nuclear lamin Ig-fold binds to PCNA: Its role in DNA replication. J Cell Biol 181: 269 –280. Solovei I, Schermelleh L, During K, Engelhardt A, Stein S, Cremer C, Cremer T. 2004. Differences in centromere positioning of cycling and postmitotic human cell types. Chromosoma 112: 410 –423. Spann TP, Goldman AE, Wang C, Huang S, Goldman RD. 2002. Alteration of nuclear lamin organization inhibits RNA polymerase II-dependent transcription. J Cell Biol 156: 603– 608. Stewart-Hutchinson PJ, Hale CM, Wirtz D, Hodzic D. 2008. Structural requirements for the assembly of LINC complexes and their function in cellular mechanical stiffness. Exp Cell Res 314: 1892– 1905. Stewart C, Burke B. 1987. Teratocarcinoma stem cells and early mouse embryos contain only a single major lamin polypeptide closely resembling lamin B. Cell 51: 383–392. Stick R, Angres B, Lehner CF, Nigg EA. 1988. The fates of chicken nuclear lamin proteins during mitosis: Evidence for a reversible redistribution of lamin B2 between inner nuclear membrane and elements of the endoplasmic reticulum. J Cell Biol 107: 397 –406. Strelkov SV, Schumacher J, Burkhard P, Aebi U, Herrmann H. 2004. Crystal structure of the human lamin A coil 2B dimer: Implications for the head-to-tail association of nuclear lamins. J Mol Biol 343: 1067– 1080. Stuurman N, Heins S, Aebi U. 1998. Nuclear lamins: Their structure, assembly, and interactions. J Struct Biol 122: 42–66. Stuurman N, Sasse B, Fisher PA. 1996. Intermediate filament protein polymerization: Molecular analysis of Drosophila nuclear lamin head-to-tail binding. J Struct Biol 117: 1 –15. Sullivan T, Escalante-Alcalde D, Bhatt H, Anver M, Bhat N, Nagashima K, Stewart CL, Burke B. 1999. Loss of A-type

lamin expression compromises nuclear envelope integrity leading to muscular dystrophy. J Cell Biol 147: 913 –920. Taimen P, Pfleghaar K, Shimi T, Moller D, Ben-Harush K, Erdos MR, Adam SA, Herrmann H, Medalia O, Collins FS, et al. 2009a. A progeria mutation reveals functions for lamin A in nuclear assembly, architecture, and chromosome organization. Proc Natl Acad Sci 106: 20788– 20793. Taimen P, Pfleghaar K, Shimi T, Moller D, Ben-Harush K, Erdos MR, Adam SA, Herrmann H, Medalia O, Collins FS, et al. 2009b. A progeria mutation reveals functions for lamin A in nuclear assembly, architecture, and chromosome organization. Proc Natl Acad Sci 106: 20788– 20793. Tang CW, Maya-Mendoza A, Martin C, Zeng K, Chen S, Feret D, Wilson SA, Jackson DA. 2008. The integrity of a lamin-B1-dependent nucleoskeleton is a fundamental determinant of RNA synthesis in human cells. J Cell Sci 121: 1014– 1024. Taniura H, Glass C, Gerace L. 1995. A chromatin binding site in the tail domain of nuclear lamins that interacts with core histones. J Cell Biol 131: 33– 44. Thompson LJ, Bollen M, Fields AP. 1997. Identification of protein phosphatase 1 as a mitotic lamin phosphatase. J Biol Chem 272: 29693– 29697. Tilgner K, Wojciechowicz K, Jahoda C, Hutchison C, Markiewicz E. 2009. Dynamic complexes of A-type lamins and emerin influence adipogenic capacity of the cell via nucleocytoplasmic distribution of beta-catenin. J Cell Sci 122: 401–413. Tsai MY, Zheng Y. 2005. Aurora A kinase-coated beads function as microtubule-organizing centers and enhance RanGTP-induced spindle assembly. Curr Biol 15: 2156–2163. Tsai MY, Wang S, Heidinger JM, Shumaker DK, Adam SA, Goldman RD, Zheng Y. 2006. A mitotic lamin B matrix induced by RanGTP required for spindle assembly. Science 311: 1887– 1893. Tzur YB, Wilson KL, Gruenbaum Y. 2006. SUN-domain proteins: ‘Velcro’ that links the nucleoskeleton to the cytoskeleton. Nat Rev Mol Cell Biol 7: 782–788. Vaughan A, Alvarez-Reyes M, Bridger JM, Broers JL, Ramaekers FC, Wehnert M, Morris GE, Whitfield WGF, Hutchison CJ. 2001. Both emerin and lamin C depend on lamin A for localization at the nuclear envelope. J Cell Sci 114: 2577– 2590. Vergnes L, Peterfy M, Bergo MO, Young SG, Reue K. 2004. Lamin B1 is required for mouse development and nuclear integrity. Proc Natl Acad Sci 101: 10428– 10433. Verstraeten VL, Broers JL, Ramaekers FC, van Steensel MA. 2007. The nuclear envelope, a key structure in cellular integrity and gene expression. Curr Med Chem 14: 1231–1248. Vigouroux C, Auclair M, Dubosclard E, Pouchelet M, Capeau J, Courvalin JC, Buendia B. 2001. Nuclear envelope disorganization in fibroblasts from lipodystrophic patients with heterozygous R482Q/W mutations in the lamin A/C gene. J Cell Sci 114: 4459– 4468. Vorburger K, Kitten GT, Nigg EA. 1989. Modification of nuclear lamin proteins by a mevalonic acid derivative

T. Dechat et al. occurs in reticulocyte lysates and requires the cysteine residue of the C-terminal CXXM motif. Embo J 8: 4007– 4013. Ward GE, Kirschner MW. 1990. Identification of cell cycle-regulated phosphorylation sites on nuclear lamin C Cell 61: 561– 577. Wheeler MA, Davies JD, Zhang Q, Emerson LJ, Hunt J, Shanahan CM, Ellis JA. 2007. Distinct functional domains in nesprin-1a and nesprin-2b bind directly to emerin and both interactions are disrupted in X-linked Emery-Dreifuss muscular dystrophy. Exp Cell Res 313: 2845– 2857. Williams RR, Azuara V, Perry P, Sauer S, Dvorkina M, Jorgensen H, Roix J, McQueen P, Misteli T, Merkenschlager M, et al. 2006. Neural induction promotes largescale chromatin reorganisation of the Mash1 locus. J Cell Sci 119: 132–140. Winter-Vann AM, Casey PJ. 2005. Post-prenylationprocessing enzymes as new targets in oncogenesis. Nat Rev Cancer 5: 405– 412. Yuan J, Simos G, Blobel G, Georgatos SD. 1991. Binding of lamin A to polynucleosomes. J Biol Chem 266: 9211– 9215.

Zackroff RV, Goldman RD. 1979. In vitro assembly of intermediate filaments from baby hamster kidney (BHK-21) cells. Proc Natl Acad Sci 76: 6226–6230. Zhang FL, Casey PJ. 1996. Protein prenylation: Molecular mechanisms and functional consequences. Annu Rev Biochem 65: 241– 269. Zhang YQ, Sarge KD. 2008. Sumoylation regulates lamin A function and is lost in lamin A mutants associated with familial cardiomyopathies. J Cell Biol 182: 35– 39. Zhang Q, Bethmann C, Worth NF, Davies JD, Wasner C, Feuer A, Ragnauth CD, Yi Q, Mellad JA, Warren DT, et al. 2007. Nesprin-1 and -2 are involved in the pathogenesis of Emery Dreifuss muscular dystrophy and are critical for nuclear envelope integrity. Hum Mol Genet 16: 2816–2833. Zhao K, Harel A, Stuurman N, Guedalia D, Gruenbaum Y. 1996. Binding of matrix attachment regions to nuclear lamin is mediated by the rod domain and depends on the lamin polymerization state. FEBS Lett 380: 161– 164. Zink D, Amaral MD, Englmann A, Lang S, Clarke LA, Rudolph C, Alt F, Luther K, Braz C, Sadoni N, et al. 2004. Transcription-dependent spatial arrangements of CFTR and adjacent genes in human cell nuclei. J Cell Biol 166: 815 –825.

Lamin-binding Proteins Katherine L. Wilson1 and Roland Foisner2 1

Department of Cell Biology, Johns Hopkins University School of Medicine, Baltimore, Maryland 21205

2

Max F. Perutz Laboratories, Department of Medical Biochemistry, Medical University of Vienna, Vienna 1030

Correspondence: [email protected]

A- and B-type lamins are the major intermediate filaments of the nucleus. Lamins engage in a plethora of stable and transient interactions, near the inner nuclear membrane and throughout the nucleus. Lamin-binding proteins serve an amazingly diverse range of functions. Numerous inner-membrane proteins help anchor lamin filaments to the nuclear envelope, serving as part of the nuclear “lamina” network that is essential for nuclear architecture and integrity. Certain lamin-binding proteins of the inner membrane bind partners in the outer membrane and mechanically link lamins to the cytoskeleton. Inside the nucleus, lamin-binding proteins appear to serve as the “adaptors” by which the lamina organizes chromatin, influences gene expression and epigenetic regulation, and modulates signaling pathways. Transient interactions of lamins with key components of the transcription and replication machinery may provide an additional level of regulation or support to these essential events.

he eukaryotic cell nucleus is a complex membrane-bounded organelle that houses, organizes, and regulates the genome. The nucleus is structurally organized into functional domains, one of which is the nuclear envelope (NE). The NE has two concentric membranes, named the “inner” and “outer” nuclear membranes (INM and ONM, respectively). These membranes are separated by a 30– 50 nm lumen, and fuse to form holes ( pores) occupied by nuclear pore complexes (NPCs), which mediate active and passive movement of molecules between the cytoplasm and nucleoplasm (Gruenbaum et al. 2005; Stewart et al. 2007). The NE and its lumen are continuous with the endoplasmic reticulum (ER) and share many

T

ER functions. However, the INM and ONM are also each structurally and functionally unique, because of specific enrichments for distinct integral membrane proteins (Schirmer and Gerace 2005; Schirmer and Foisner 2007). In mammals, the INM in particular appears to be populated by over 50 different membrane proteins, most of which are uncharacterized. Among characterized INM proteins, most can bind directly to A- or B-type lamins, or both. A- and B-type lamins polymerize to form separate networks of nuclear intermediate filaments that concentrate near the INM in metazoans (Dechat et al. 2008). Many INM proteins are localized by binding directly or indirectly to lamin filaments. This network of filaments and

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000554 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000554

K.L. Wilson and R. Foisner

lamin-binding proteins at the INM is known as the “peripheral lamina.” B-type lamins are essential for cell viability and development, whereas A-type lamins arose later in evolution and are nonessential. Lamins are important structurally and as “scaffolds” for many other proteins and complexes in the nucleus. Lamin filaments are important for the assembly, structure, shape, and mechanical stability of metazoan nuclei. There is also growing evidence that lamins regulate chromatin organization and gene expression, and influence signaling (Gruenbaum et al. 2005; Dechat et al. 2008). These functions involve a full spectrum of biochemical interactions between lamins, chromatin, and a variety of partners, including regulatory proteins responsive to external and intrinsic signals (Fig. 1). The pace of discovering new lamin-binding proteins has nearly

overwhelmed our capacity to characterize them. Each new partner has the potential to provide fresh insight into the structure and regulation of the nucleoskeleton and its relationships with the genome. Further motivation for research in this area comes with the discovery that an increasing number of human diseases are linked to defects in lamins or lamin-binding proteins. THE CONCEPT OF LAMINA-ASSOCIATED POLYPEPTIDES

The term lamina-associated-polypeptide (LAP) was coined to define a novel group of nuclear proteins that cofractionated with lamins and required high concentrations of monovalent salts and nonionic detergents to extract them from nuclei (Senior and Gerace 1988; Foisner

Cytoskeleton NPC ONM INM

Reinforce nucleoskeleton

Link nucleoskeleton to cytoskeleton

Regulate signaling

NPC anchoring assembly

Chromatin tethering

Retention at INM Lamin filaments

Gene regulation

Lamin assembly

Regulate signaling/ transcription activity

Lamin-binding proteins

Figure 1. Overview of known roles for lamin-binding proteins. Many lamin-binding proteins located in the

INM, on chromatin, and in the nucleoplasm are thought to have mechanical and structural roles, such as reinforcing the nucleoskeleton, interlinking the nucleoskeleton and cytoskeleton, anchoring NPCs, and tethering chromatin to the nuclear envelope. Others regulate signaling or transcription. Many lamin-binding proteins require lamins for their correct localization, whereas others regulate or facilitate lamin assembly.

Lamin-binding Proteins

and Gerace 1993). The first characterized LAPs were all integral INM proteins: LAP1 (Martin et al. 1995) and LAP2 (Harris et al. 1994; Furukawa et al. 1995; Berger et al. 1996) each has one transmembrane domain and is expressed as multiple isoforms, whereas the so-called lamin B receptor (LBR) has eight transmembrane domains (Worman et al. 1990). These proteins all bind lamins directly in vitro (Worman et al. 1988; Foisner and Gerace 1993; Furukawa et al. 1998). Since then, around 80 additional putative NE membrane proteins have been discovered (Schirmer et al. 2005; Schirmer and Foisner 2007). Most fit the original LAP criterion (resistance to biochemical extraction) and localize at the NE, but are untested for binding to lamins. The LAP concept expanded as lamins (Moir et al. 2000b; Naetar et al. 2008) and lamin-binding proteins including Narf (Barton and Worman 1999), LAP2a (Naetar et al. 2008), and Lco1 (Vlcek et al. 2004; Naetar et al. 2008) were discovered in the nuclear interior. Laminbinding proteins can have widely differing, and differentially regulated, affinities for lamins that might influence their biochemical extraction. Some proteins are known to bind lamins directly and relatively stably at the NE or in the nuclear interior. By contrast, other proteins bind lamins more transiently, probably as a means of regulating their activities; these partners include PCNA (Shumaker et al. 2008), c-Fos (Gonzalez et al. 2008), retinoblastoma (Rb) (Johnson et al. 2004; Pekovic et al. 2007), and Oct-1 (Malhas et al. 2009). LEM-DOMAIN PROTEINS AND BAF: TETHERING CHROMATIN TO THE NUCLEOSKELETON

One prominent family of lamin-binding proteins involved in nuclear architecture and chromatin organization share the LEM (LAP2, Emerin, MAN) domain, an 45-residue motif that folds as two a-helices (Laguri et al. 2001). The LEM domain binds a conserved metazoan chromatin protein named Barrier to Autointegration Factor, BAF (Furukawa 1999; Cai et al. 2001; Lee et al. 2001; Shumaker et al. 2001; Shimi et al. 2004; Cai et al. 2007). BAF, a mobile

lamin-binding protein, can “bridge” DNA and interacts with histones (Margalit et al. 2007). Most, but not all, LEM proteins are integral INM proteins with one or two transmembrane domains (Fig. 2). Mammals have four genes encoding characterized LEM proteins, LAP2 (a, b, and other isoforms), MAN1, emerin (Lin et al. 2000), and LEM2/NET25 (Schirmer et al. 2003; Brachner et al. 2005; Chen et al. 2006; Ulbert et al. 2006), and three uncharacterized genes encoding predicted nonmembrane proteins, LEM3, LEM4, and LEM5 (Lee and Wilson 2004), known respectively as ANKLE1, ANKLE2, and LEMD1. Three LEM proteins (emerin, LEM2, and LEM3) are conserved in Caenorhabditis elegans (Lee et al. 2000; Gruenbaum et al. 2002; Liu et al. 2003). Four are expressed in Drosophila: MAN1 (Wagner et al. 2006), otefin (Goldberg et al. 1998; Jiang et al. 2008), and Bocksbeutel a and b(Wagner et al. 2004). The functions and interactions of LEM proteins, lamins, and BAF are strongly conserved between mammals, nematodes, and flies, suggesting they have fundamental roles in the nucleus. Some LEM-domain proteins have additional domains that bind DNA, or other chromatin proteins. For example, LAP2b can bind HA95, a chromatin protein involved in DNA replication (Martins et al. 2003). All isoforms of LAP2 have a second “LEM-like” domain that binds DNA, rather than BAF (Cai et al. 2001; Laguri et al. 2001). Similarly, MAN1 and probably LEM2 binds DNA directly via a carboxyterminal “winged helix” domain (Caputo et al. 2006). Furthermore, all tested LEM proteins bind either A- or B-type lamins, or both, directly (Clements et al. 2000; Lee et al. 2001; Sakaki et al. 2001; Brachner et al. 2005; Mansharamani and Wilson 2005), and some require this interaction to localize at the NE (Sullivan et al. 1999; Vaughan et al. 2001; Brachner et al. 2005). Lamins, in turn, also directly bind DNA (Shoeman and Traub 1990; Luderus et al. 1992; Stierle et al. 2003) and histones (Taniura et al. 1995; Mattout et al. 2007). Thus, a complex network of interactions between lamins, LEM proteins, BAF, and, most likely, other INM proteins is involved in anchoring chromatin to the NE and lamins.

K.L. Wilson and R. Foisner

Ce-Emerin Dm-Bocksbeutel Dm-Otefin h-Emerin

Ce-LEM2 Dm-MAN1 h-MAN1 h-LEM2

h-LAP2-beta

ONM INM Lamina

HA95 BAF

HDAC3

BAF

BAF

BAF

Rb

Gcl

Gcl Lmo7 Btf β-catenin

Gcl Smad

h-LAP2-alpha Lamin BAF

LEM

LEM-like

Winged Helix

Figure 2. LEM-domain proteins organize chromatin and regulate signaling and transcription. Domain

organizations of characterized LEM proteins; the LEM motif confers direct binding to barrier to autointegration factor (BAF). All LEM proteins can bind A- and/or B-type lamins. Some, including human (h-)emerin, C. elegans (Ce-) emerin, D. melanogaster (Dm-) bocksbeutel a, and Dm-Otefin, have one transmembrane (TM) domain. LAP2 proteins, expressed only in vertebrates, also have a DNA-binding “LEM-like” domain. Others (e.g., hLem2, Ce-Lem2, hMAN1, and Dm-MAN1) have two TM domains and a carboxy-terminal Winged Helix domain that binds DNA. Some, including hLAP2a (shown) and three uncharacterized LEM proteins (not shown), have no TM domain. LEM proteins have additional functional domains; e.g., LAP2b partners include replication protein HA95 and HDAC3; LAP2b recruits HDAC3 to the NE and contributes to epigenetic regulation. Emerin binds many transcription factors including GCL, Lmo7, Btf, and b-catenin. GCL also binds hLAP2b and hMAN1, suggesting functional overlap. MAN1 is unique in binding regulatory Smads, and inhibits BMP and TGF-b signaling.

LEM proteins might appear to function redundantly in chromatin organization because there are so many of them, and functional inactivation of any single tested LEM gene does not grossly disrupt chromatin organization. Functional overlap is supported by C. elegans studies, which showed that double-knockdown of two LEM proteins (Ce-emerin and Ce-lem2) caused embryonic lethality at the 100-cell stage, when embryonic gene expression normally begins, with phenotypes that included aneuploidy, grossly defective chromatin organization and nuclear structure, and failure to assemble nuclei after mitosis (Liu et al. 2003). Nearly identical phenotypes are caused by down-regulating

either Ce-lamin or BAF alone (Gruenbaum et al. 2005; Margalit et al. 2005a), strongly suggesting that BAF and LEM-domain proteins are key components of nuclear architecture. Nevertheless, despite potential redundancies, eliminating just one LEM protein, for example emerin in mice, can affect nuclear structure: Emerin-null skeletal muscle nuclei are fragile (Ozawa et al. 2006), and show significantly increased autophagic degradation of structurally aberrant regions of the nucleus (Park et al. 2009a). Lamin mutations also cause obvious defects in nuclear shape and chromatin organization (Sabatelli et al. 2001; Scaffidi and Misteli 2006; Shumaker et al. 2006; Wang

Lamin-binding Proteins

et al. 2006; Hakelien et al. 2008; Park et al. 2009b), as well as enhanced autophagic degradation of nuclei (Park et al. 2009a). Further evidence for the physiological relevance of both lamins and LEM proteins in chromatin organization comes from recent genomewide studies to identify lamina-bound DNA. The analysis of methylated DNA in cultured Drosophila cells expressing lamin B1 fused to DNA methyltransferase (Dam), revealed 500 genes in close contact with the lamina (Pickersgill et al. 2006). In human fibroblasts, this strategy revealed both lamin B1 and emerin in close contact with .1300 sharply defined domains in the genome, most of which were flanked either by binding sites for the insulator protein CTCF, or by E2F/DP-regulated promoters (Guelen et al. 2008). Interestingly, at least two INM LEM proteins, LAP2b and emerin, are implicated in the regulation of E2F/DP promoters (Nili et al. 2001; Holaska and Wilson 2006), and both CTCF and A-type lamins are required for the insulator function of a subtelomeric array of D4Z4 repeats, shortening of which cause Facio-Scapulo-Humeral Dystrophy (FSHD) (Ottaviani et al. 2009). Future application of genomewide “close contact” studies to other LEM proteins, and other types of lamin-binding proteins, may provide much-needed insight into their relationships with chromosomes. Interestingly, in proliferating cells, these “relationships” might be established actively while nuclei reassemble during mitotic anaphase and telophase. Nuclear assembly involves a complex series of interactions between chromatin, membranes, INM and ONM proteins, lamins, and other mitotically solubilized components, the mechanisms and regulation of which remain poorly understood (Margalit et al. 2005b; Anderson and Hetzer 2008; Guttinger et al. 2009). However, protein – protein interactions, for example between BAF, lamins, and LEM proteins, are required because nuclear assembly fails when one component is missing, as shown in C. elegans studies (Margalit et al. 2005b), and also fails in HeLa cells that express emerin and BAF mutants with impaired interactions (Haraguchi et al. 2001).

LINC COMPLEXES LINK THE NUCLEAR INTERIOR TO THE CYTOPLASM

Many independent studies in yeast, worms, flies, and mammals converged on a major discovery: Membrane proteins of the NE mechanically interlink the nucleoskeleton and cytoskeleton. These “LINC” complexes (Crisp et al. 2006) enable the regulated, cytoplasmic-motor-driven movement of the entire nucleus to new positions within the cell, and the movement of entire NE-tethered chromosomes to new positions along the NE (for reviews, see Tzur et al. 2006b; Wilhelmsen et al. 2006; Starr 2009). In addition, gene-regulatory responses to external force must now be assumed to potentially involve not only indirect signaling from the cell surface, but also direct signaling via mechanotransducing complexes at the NE. Considered at the simplest level, LINC complexes consist of lamin-binding SUN-domain proteins in the INM that interact in the NE lumen with KASH-domain proteins (Fig. 3) anchored in the ONM (Hodzic et al. 2004; Padmakumar et al. 2005; Crisp et al. 2006; Haque et al. 2006); KASH proteins also bind directly or indirectly to cytoplasmic actin, microtubules, centrosomes, or intermediate filaments (Zhang et al. 2001; Zhen et al. 2002; Padmakumar et al. 2004; Wilhelmsen et al. 2005; McGee et al. 2006; Roux et al. 2009). It is important to note that at least two of the many KASH-domain protein isoforms (nesprin-1a and nesprin-2b) are INM proteins that bind directly to lamins and emerin (Mislow et al. 2002a; Mislow et al. 2002b; Zhang et al. 2005). SUN-domain proteins, conserved from yeast to humans, are defined by a 120-residue motif located in the lumenal space of the NE; the name SUN was based on S. pombe Sad1 and D. melanogaster UNC-84 (Malone et al. 1999). C. elegans and D. melanogaster each have two SUN genes: UNC84 and germ-cell-specific matefin/SUN1 in C. elegans, and Klaroid and CG6589 in Drosophila. Mammals have four: SUN1 and SUN2, and testis-specific genes SUN3 and Spag4 (reviewed in Starr 2009). Human SUN1 (hSUN1) and hSUN2 each have a coiled-coil domain near the SUN domain that

K.L. Wilson and R. Foisner

KASH-domain proteins Ce-ANC-1 Nesprin-1/2

h-Nesprin 3

Ce-ZYG-12 Dm-Klarsicht

Ce-ZYG-12

Intermediate Filament Actin

Centrosomes

Microtubule

Plectin Dynein Dynein

ONM INM Him-8 ?

Lamina h-Nesprin 1/2 Ce-UNC84 h-SUN1/2

Ce-UNC84 Dm-Klaroid

Meiotic chromosome h-SUN1/2

Ce-Matefin h-SUN 1

SUN-domain proteins

SUN domain

KASH domain

Actin binding domain

Figure 3. SUN- and KASH-domain proteins interact in the NE lumen to form LINC (Links Nucleoskeleton and

Cytoskeleton) complexes. SUN-domain proteins span the INM and dimerize; each nucleoplasmic domain binds lamins and each SUN-domain in the NE lumen binds the KASH domain of an ONM-embedded KASH protein, which binds actin, centrosomes/microtubules, or plectin/intermediate filaments in the cytoplasm. Only one KASH domain partner of a SUN dimer is depicted. SUN proteins also bind, potentially via Him-8, the pairing centers of meiotic chromosomes. Many additional nesprin isoforms reside elsewhere, including the INM; e.g., human nesprins-1a and -2b in the INM bind lamins and emerin (not shown).

mediates dimerization and is also involved in binding the KASH domain; their exposed nucleoplasmic amino-terminal domain binds lamins (reviewed by Tzur et al. 2006b; Starr 2009). KASH- (Klarsicht, ANC-1, Syne homology) domain nomenclature is confusing because these proteins were identified in diverse species before their conservation was recognized. Mammalian KASH proteins are now known as “nesprins.” The KASH domain comprises a single carboxy-terminal transmembrane domain followed by a 35-residue lumenal domain. ONM-localized KASH proteins require the interaction with SUN-domain proteins for retention at the ONM. C. elegans encodes

three known KASH-domain proteins (ANC-1, UNC-83, and ZYG-12) (Starr and Han 2002; Malone et al. 2003; McGee et al. 2006) and D. melanogaster encodes the two proteins Klarsicht and MSP 300 (Patterson et al. 2004; Xie and Fischer 2008). Nesprin isoform complexity is much higher in mammals, which have four known genes: Nesprin-1 (SYNE1), nesprin-2 (SYNE2), nesprin-3, and nesprin-4. The Nesprin-1 and -2 genes are alternatively transcribed and alternatively spliced to produce more than 12 protein isoforms each, from small to enormous (.1 MDa). Some isoforms lack the KASH domain and are not membrane-localized, whereas others localize at the INM or even the Golgi complex (Starr and Fischer 2005; Wilhelmsen et al.

Lamin-binding Proteins

2006). All nesprins have multiple “spectrin repeat” domains that confer an extended configuration, but can also mediate specific protein – protein interactions. For example, specific spectrin repeat domains in the small, INM-localized isoform nesprin-1a mediate direct binding to lamins, emerin, and other nesprin-1a molecules (Mislow et al. 2002a). The largest isoforms of nesprin-1 and -2 bind actin (Zhen et al. 2002; Padmakumar et al. 2004). Nesprin-3 binds plectin, a cytoskeletal protein that binds cytoplasmic intermediate filaments (Wilhelmsen et al. 2005), whereas nesprin-4 binds kinesin 1, a plus-enddirected microtubule-dependent motor, and may be involved in dislocalizing the centrosome and Golgi membranes away from the nucleus in epithelial cells (Roux et al. 2009). LINC complexes formed by SUN- and KASHdomain proteins are known to determine the spacing between the INM and ONM (Crisp et al. 2006), position NPCs (Liu et al. 2007), and control nuclear size and NE architecture (Luke et al. 2008). Different combinations of SUN- and KASH-domain proteins mediate different functions (Fig. 3), including forcetransfer across the NE, and the stiffness of the cytoskeleton (Stewart-Hutchinson et al. 2008). In C. elegans, the SUN protein UNC84 anchors nuclei to cytoplasmic actin filaments viathe KASHdomain protein ANC-1 (Malone et al. 1999; Starr and Han 2002), mediates movement of nuclei via a different KASH partner (UNC-83) (Starr et al. 2001), and attaches centrosomes to the NE via a third KASH partner (ZYG-12) and the dynein subunit DLI-1 (Malone et al. 2003) (Fig. 3). The other C. elegans SUN-domain protein, matefin/SUN1, is also required to recruit the proapoptotic protein CED-4 to the NE (Tzur et al. 2006a), implying additional roles for LINC complexes in death signaling. In Drosophila, LINC complexes mediate centrosome attachment to the NE, and nuclear migration in developing photoreceptor cells (Patterson et al. 2004; Kracklauer et al. 2007). In mice, deletions in the nesprin-1 or nesprin-2 genes, or overexpression of the KASH-domain, which destroys endogenous LINC complexes, disrupts nuclear organization, results in failure of nuclei to cluster at neuromuscular junctions (Grady et al. 2005;

Zhang et al. 2007), and can cause muscular dystrophy (Puckelwartz et al. 2009). Amazingly, LINC complexes can also transmit mechanical force from the cytoskeleton through the NE to move chromosomes, forexample during meiotic prophase, when telomeres attach to the NE and cluster to form bouquet-like structures that facilitate chromosome pairing (Chikashige et al. 2007). In a C. elegans matefin/ SUN1 mutant strain, chromosome reorganization is disrupted in early meiosis, and ZYG-12 “patches” do not form on the NE (Penkner et al. 2007). This suggests meiotic chromosome clustering and homolog recognition requires both matefin/SUN1 and ZYG-12 (a KASH-domain protein), potentially via HIM-8-mediated attachment of pairing centers to matefin/SUN1 (Penkner et al. 2007). Similarly, mammalian SUN1 and SUN2 (Ding et al. 2007; Schmitt et al. 2007) are telomere-associated between the leptotene and diplotene stages of meiosis, and SUN1-knockout mice have both impaired NE-telomere association and impaired homolog pairing (Ding et al. 2007). The meiotic functions of mammalian SUN proteins do not require A-type lamins (Schmitt et al. 2007). Potential roles for B-type lamin(s) remain to be tested in mammals, and seem likely because in C. elegans the nuclear migration and nuclear anchoring functions of LINC complexes require the one (B-type) lamin expressed in this organism (Lee et al. 2002). However, similar SUN- and KASH-dependent pathways exist in S. pombe and S. cerevisiae, which do not have lamins (Starr 2009). It is unknown how the functions and interactions of SUN and KASH proteins are regulated (Starr 2009). One interesting possibility is that LINC complexes might be regulated by proteins located in the NE lumen. This possibility is suggested by Torsin A, a so-called “AAAþ ATPase” enzyme located in the NE/ER lumen that is required to localize nesprin (Nery et al. 2008). REGULATION OF SIGNALING AND DIFFERENTIATION BY LAMIN-BINDING PROTEINS

The plasma membrane contains many structural complexes (e.g., focal adhesions and dystroglycan

K.L. Wilson and R. Foisner

complex) that can be dynamically regulated and generate signals to which cells respond in a tissue-specific manner, in many cases at the level of gene regulation. There is increasing evidence that this paradigm might also apply to laminassociated protein complexes at the NE, and in the nucleoplasm. The concept of “lamin-linked dynamic signaling” is supported by studies on several LEM-domain proteins. For example, LAP2b, the largest INM-localized LAP2 isoform, binds lamin B specifically, and also directly interacts with transcription regulators (e.g., GCL) and epigenetic modifiers including HDAC3, and contributes to transcriptional repression (Nili et al. 2001; Somech et al. 2005). Interestingly, another INM-localized LEM protein, MAN1, directly binds and inhibits R-Smads and functions as a major regulator of BMP- and TGF-b-signaling in early vertebrate development (Osada et al. 2003; Raju et al. 2003; Lin et al. 2005; Mansharamani and Wilson 2005; Pan et al. 2005; Cohen et al. 2007). Roles for lamin-binding proteins in TGF-b-signaling are also reported in Drosophila: The INM-localized LEM protein Otefin binds directly to Medea (Smad-4) and they physically tether their repressed target locus, bam (“bag of marbles”), at the NE (Jiang et al. 2008). The INM protein emerin also supports the paradigm of complex regulation of protein – protein interactions and signaling at the NE. Emerin directly binds more than ten known partners, suggesting a degree of biochemical complexity at the NE that rivals many cell surface structures. Among these partners are structural proteins (nesprin-1a, nesprin-2b, actin, nuclear myosin 1c, lamins, and tubulin), other INM proteins (MAN1 and LUMA), proteins involved in signaling, transcription, mRNA splicing (b-catenin, GCL, Lmo7, Btf, and YT521-B), and of course, BAF (Lee et al. 2001; Mislow et al. 2002a; Holaska et al. 2003; Haraguchi et al. 2004; Holaska et al. 2004; Mansharamani and Wilson 2005; Holaska et al. 2006; Markiewicz et al. 2006; Wheeler et al. 2007; Bengtsson and Otto 2008) (for reviews, see Bengtsson and Wilson 2004; Wagner and Krohne 2007). The biochemical purification of emerin-associated

protein complexes from HeLa cell nuclei suggests emerin can form at least six distinct multiprotein complexes (Holaska and Wilson 2007). Some putative complexes included primarily architectural components (e.g., lamins, actin, nuclear myosin 1c, and nuclear aII-spectrin), whereas others included chromatin and gene regulators (e.g., histones, Lmo7, Rb, and components of the Nuclear Co-Repressor [NCoR] complex). Emerin may also be involved in sensing and responding to mechanical tension at the NE (Lammerding et al. 2005). The interaction of Lmo7 (Holaska et al. 2006) and b-catenin (Markiewicz et al. 2006) with emerin regulates their nuclear export and represses their activity. These regulatory functions may be important for muscle and adipocyte differentiation, respectively (Holaska et al. 2006; Tilgner et al. 2009). It is still unclear how emerin regulates the nuclear export of these factors. Emerin was also reported to localize in the ONM and mediate centrosome-NE attachment (Salpingidou et al. 2007). How emerin “knows” when to assemble or disassemble particular complexes, and how its localization in the INM versus ONM is regulated, are important open questions. Growing evidence that emerin is phosphorylated suggests mechanisms by which different kinases and signaling pathways might directly control its binding to specific partners, including BAF (Hirano et al. 2005; Roberts et al. 2006; Schlosser et al. 2006; Luo et al. 2009). REGULATION OF SIGNALING AND PROLIFERATION CONTROL BY LAMIN-BINDING PROTEINS IN THE NUCLEAR INTERIOR

The largest LAP2 isoform, LAP2a, lacks a transmembrane domain, localizes to the nuclear interior, and specifically binds A-type lamins (Dechat et al. 2000; Vlcek et al. 2002; Dechat et al. 2004). In proliferating cells, LAP2a is essential and sufficient to target and retain a subset of lamins within the nuclear interior (Naetar et al. 2007; Naetar et al. 2008). Nucleoplasmic lamins are stable, but more mobile than peripheral lamins, suggesting interior lamins assemble

Lamin-binding Proteins

differently or bind to chromatin (Moir et al. 2000b). LAP2a and lamin A also bind directly to tumor suppressor Rb (Ozaki et al. 1994; Markiewicz et al. 2002; Pekovic et al. 2007), and LAP2a is involved in Rb-mediated repression of E2F/Rb target genes (Dorner et al. 2006) and Rb-dependent proliferation control in tissue progenitor stem cells (Naetar et al. 2008). The physiological relevance of intranuclear LAP2a-lamin complexes in Rb regulation is supported by studies showing destabilization and proteolytic degradation (Johnson et al. 2004; Nitta et al. 2006; Nitta et al. 2007) and impaired phosphorylation-dependent repressor activity control (Van Berlo et al. 2005) of Rb in cells lacking A-type lamins. “FAMILIAR” REGULATORY PROTEINS AND ENZYMES DISCOVERED TO BIND LAMINS

In addition to Rb, other known transcription factors and regulators have been discovered to bind A- and/or B-type lamins directly. This binding, whether transient or stable in vivo, suggests these proteins—and pathways they represent—either require or are influenced by lamins. These lamin-binding proteins include cyclin D3 (Mariappan et al. 2007), transcription factors cFos, Oct-1, SREBP1, MOK2, and tumor suppressor ING1 (Han et al. 2008), an enzyme named 12(S)-lipoxygenase (12[S] LOX) (Tang et al. 2000), and at least two kinases: protein kinase C a (PKCa) and JIL-1 kinase (Bao et al. 2005) (reviewed in Zastrow et al. 2004; Vlcek and Foisner 2007). Binding to lamin B1 sequesters transcription factor Oct-1 at the NE, away from promoters; in lamin B1-deficient cells, Oct-1-dependent genes that respond to oxidative stress and reactive oxygen species are misregulated (Malhas et al. 2009). Similarly, c-Fos, an early response transcription factor, is sequestered at the NE by A-type lamins (Ivorra et al. 2006) and during mitogen activated protein (MAP) kinase signaling, activated ERK1/2 interacts with lamin A and c-Fos at the NE to rapidly release c-Fos and facilitate cell proliferation (Gonzalez et al. 2008). Deregulation of this pathway in emerin and lamin A-deficient cells might up-regulate ERK-mediated signaling, a

known phenotype of certain lamin- or emerindeficient cells (Muchir et al. 2007; Muchir et al. 2009). Lamin A is also reported to affect Notch signaling, required to differentiate mesenchymal stem cells, by tethering SKIP, a coactivator of Notch-dependent target genes (Scaffidi and Misteli 2008). A better understanding of these interactions is sorely needed because it might begin to illuminate how and why nuclear architecture defects, for example in accelerated aging or other diseases linked to mutations in lamins and lamin-binding proteins, might disrupt signaling and gene regulation pathways. The nuclear import receptor, importin a, is also a lamin-binding protein (Adam et al. 2008). Importin a inhibits lamin assembly in vitro, suggesting it prevents newly synthesized lamins from assembling in the cytoplasm; this interaction is disrupted by Ran-GTP, found at high levels only within nuclei, releasing the assembly block once lamins are in the nucleus. PCNA, the Sliding Clamp “Ring” for DNA Replication, Binds Lamins

Lamins are also involved in DNA replication. Replication is a structural challenge, because replication complexes must traverse each huge chromosome, including regions tethered as heterochromatin to the NE. DNA replication complexes are anchored to nucleoskeletal structures (Hozak et al. 1993). During replication, the NE-anchored heterochromatin becomes disconnected, moves to replication complexes, and is then retethered at the NE (Li et al. 1998). DNA replication arrests at the elongation phase in nuclei subjected to lamina network disruption by “dominant –negative” lamin fragments (Spann et al. 1997; Moir et al. 2000a). In a breakthrough for this understudied area, PCNA ( proliferating cell nuclear antigen), which forms a sliding clamp “ring” ahead of the replication fork, was found to bind lamins directly (Shumaker et al. 2008). Further study of PCNA, the first but perhaps not the only lamin-binding component of DNA replication machinery, might shed light on how and why this fundamental process came to depend on the nucleoskeleton and lamins in metazoans.

K.L. Wilson and R. Foisner

LAMIN-BINDING PROTEINS AND HUMAN DISEASE

The diversity and physiological relevance of lamin-binding proteins is reflected in the growing number of human diseases linked to genes encoding lamins or lamin-binding proteins (Broers et al. 2006; Capell and Collins 2006; Worman and Bonne 2007). These diseases are known collectively as “laminopathies” or “envelopathies.” More than 12 overlapping or distinct syndromes including autosomal dominant Emery-Dreifuss muscular dystrophy (ADEDMD) are caused by mutations in the gene encoding A-type lamins (LMNA). In several cases, the same or clinically indistinguishable diseases can also be caused by mutations in one or more lamin-binding proteins. For example, EDMD is also caused by X-linked recessive loss of emerin (Bione et al. 1994), or dominant missense mutations in SYNE1 or SYNE2, encoding nesprin-1 and nesprin-2, respectively (Zhang et al. 2007; Puckelwartz et al. 2009). These similar pathologies suggest the disruption of a functional complex(es) that requires products of all four genes (Gotzmann and Foisner 2006). On the other hand, mutations in emerin can also cause other syndromes including Limb-Girdle muscular dystrophy, cardiomyopathy with conduction defects, or familial atrial fibrillation (Ben Yaou et al. 2007; Ura et al. 2007; Karst et al. 2008). “Classical” EDMD is characterized by progressive muscle wasting, contractures of tendons, and cardiomyopathy with ventricular conduction system defects (Bione et al. 1994; Manilal et al. 1996). Most X-linked EDMD patients are functionally null for emerin because of instability of the mutated protein or, more rarely, failed INM retention of mutant emerin proteins. In many AD-EDMD patients with LMNA mutations, emerin drifts into the ER and is functionally lost from the NE (Broers et al. 2006; Worman and Bonne 2007); however, with LMNA mutations, one must assume many additional INM or ONM proteins similarly mislocalize, if their retention at the NE requires A-type lamins. Gene-expression profiling of muscle biopsies from EDMD patients (Bakay et al. 2006) and emerin-down-regulated mouse cells (Melcon

et al. 2006) suggested shared defects in the expression of genes regulated by Rb1 and MyoD, which are required for muscle stem cell regulation and muscle differentiation (Gotzmann and Foisner 2006). A mutation in LAP2a that disrupts binding to lamin A is known to cause dilated cardiomyopathy (Taylor et al. 2005). Whether this mutation might also impair Rb activity, or reflect a novel role of LAP2a, is unknown. Additional disease mechanisms are suggested by the impaired organization and function of the neuromuscular junction in lamin-null mice and AD-EDMD patients (Mejat et al. 2009), and evidence that emerin is regulated by Her2 signaling (Tifft et al. 2009). “Signaling” Laminopathies Linked to MAN1 and LBR

Heterozygous loss-of-function mutations in MAN1 (LEMD3) cause osteopoikilosis, Buschke-Ollendorff syndrome, and melorheostosis, characterized by increased bone density (Hellemans et al. 2004). This pathology may result from impaired MAN1 regulation of TGF-b (Smad) signaling (discussed above), which is important for bone development. LBR, the INM-localized sterol reductase that also binds lamin B, is required for nuclei to change shape and reorganize chromatin in differentiating neutrophils (Hoffmann et al. 2002). LBR is proposed to associate with heterochromatic under-acetylated chromatin (Polioudaki et al. 2001; Makatsori et al. 2004) through binding to heterochromatin protein 1 (HP1) (Ye and Worman 1996). Heterozygous LBR mutations cause Pelger-Huet anomaly, a benign autosomal dominant syndrome characterized by abnormal nuclear shape and chromatin organization in blood granulocytes (Hoffmann et al. 2002). Greater loss of LBR protein correlates with more severe phenotypes, including developmental delay, epilepsy, and skeletal abnormalities (Hoffmann et al. 2007). Mutations that significantly reduce the sterol reductase activity of LBR are linked to autosomal recessive Greenberg’s skeletal dysplasia, characterized by lethal skeletal and visceral anomalies (Waterham et al. 2003).

Lamin-binding Proteins

Central Nervous System (CNS) Laminopathies

Brain pathologies with genetic links to lamins or lamin-binding proteins were noticeably missing, until recently. Surprisingly, the duplication of LMNB1, encoding lamin B1, causes autosomal dominant leukodystrophy, a slowly progressive disorder characterized by widespread demyelination of the CNS (Padiath et al. 2006; Brussino et al. 2009). Proper regulation of lamin B1 expression is crucial for oligodendrocyte development and myelination (Lin and Fu 2009). The lamin B1-binding proteins and pathways that underlie these functions in the brain are open questions. Primary dystonia is a CNS-based autosomal-dominant movement disorder caused by mutations in the AAAþATPase torsin A. This brain pathology was traced to the aberrantly tight binding of the mutated torsin A protein to the lumenal domain of LAP1 (Naismith et al. 2004; Goodchild and Dauer 2005), an INM protein that binds A- and B-type lamins. Wild-type torsin A is recruited to the NE by LAP1 when ATP is bound, whereas torsin A distributes throughout the ER lumen in the absence of ATP; the immobilization of mutant torsin A in the NE lumen correlates with cytoskeletal protein accumulation near the NE and disrupted cytoskeletal dynamics, neurite extension, and cell adhesion (Hewett et al. 2006). In addition to causing EDMD, other mutations in nesprin-1 (SYNE1) can cause autosomal-recessive cerebellar ataxia (Gros-Louis et al. 2007), characterized by impaired walking. Although this pathology might reflect impaired function of a brain-specific postsynaptic nesprin-1 isoform, parallel disruption of nuclear isoforms cannot be ruled out.

CONCLUDING REMARKS

Lamin-binding proteins are an emerging class of proteins that may explain how chromosomes are organized, how nuclei are assembled after mitosis or moved to new positions in the cell, and how transcription, replication, signaling, and many other activities are supported by lamin filaments and the nucleoskeleton. Further

exploration of this understudied area of cell biology may lead to improved and potentially therapeutic understanding of many human diseases. ACKNOWLEDGMENTS

We gratefully acknowledge grant support from the Austrian Science Research Fund (FWF P17871) and the EURO-Laminopathies research project of the European Commission (Contract LSHM-CT-2005-018690) to R.F. and National Institutes of Health Directors Bridge Award (2R56GM08646-13A1) to K.L.W. REFERENCES Adam SA, Sengupta K, Goldman RD. 2008. Regulation of nuclear lamin polymerization by importin a. J Biol Chem 283: 8462– 8468. Anderson DJ, Hetzer MW. 2008. Shaping the endoplasmic reticulum into the nuclear envelope. J Cell Sci 121: 137 –142. Bakay M, Wang Z, Melcon G, Schiltz L, Xuan J, Zhao P, Sartorelli V, Seo J, Pegoraro E, Angelini C, et al. 2006. Nuclear envelope dystrophies show a transcriptional fingerprint suggesting disruption of Rb-MyoD pathways in muscle regeneration. Brain 129: 996 –1013. Bao X, Zhang W, Krencik R, Deng H, Wang Y, Girton J, Johansen J, Johansen KM. 2005. The JIL-1 kinase interacts with lamin Dm0 and regulates nuclear lamina morphology of Drosophila nurse cells. J Cell Sci 118: 5079–5087. Barton RM, Worman HJ. 1999. Prenylated prelamin A interacts with Narf, a novel nuclear protein. J Biol Chem 274: 30008– 30018. Ben Yaou R, Toutain A, Arimura T, Demay L, Massart C, Peccate C, Muchir A, Llense S, Deburgrave N, Leturcq F, et al. 2007. Multitissular involvement in a family with LMNA and EMD mutations: Role of digenic mechanism? Neurology 68: 1883–1894. Bengtsson L, Otto H. 2008. LUMA interacts with emerin and influences its distribution at the inner nuclear membrane. J Cell Sci 121: 536 –548. Bengtsson L, Wilson KL. 2004. Multiple and surprising new functions for emerin, a nuclear membrane protein. Curr Opin Cell Biol 16: 73–79. Berger R, Theodor L, Shoham J, Gokkel E, Brok-Simoni F, Avraham KB, Copeland NG, Jenkins NA, Rechavi G, Simon AJ. 1996. The characterization and localization of the mouse thymopoietin/lamina- associated polypeptide 2 gene and its alternatively spliced products. Genome Res 6: 361– 370. Bione S, Maestrini E, Rivella S, Mancini M, Regis S, Romeo G, Toniolo D. 1994. Identification of a novel X-linked gene responsible for Emery-Dreifuss muscular dystrophy. Nature Genet 8: 323–327.

K.L. Wilson and R. Foisner

Brachner A, Reipert S, Foisner R, Gotzmann J. 2005. LEM2 is a novel MAN1-related inner nuclear membrane protein associated with A-type lamins. J Cell Sci 118: 5797– 5810. Broers JL, Ramaekers FC, Bonne G, Yaou RB, Hutchison CJ. 2006. Nuclear lamins: Laminopathies and their role in premature ageing. Physiol Rev 86: 967 –1008. Brussino A, Vaula G, Cagnoli C, Mauro A, Pradotto L, Daniele D, Di Gregorio E, Barberis M, Arduino C, Squadrone S, et al. 2009. A novel family with Lamin B1 duplication associated with adult-onset leucoencephalopathy. J Neurol Neurosurg Psychiatry 80: 237–240. Cai M, Huang Y, Ghirlando R, Wilson KL, Craigie R, Clore GM. 2001. Solution structure of the constant region of nuclear envelope protein LAP2 reveals two LEM-domain structures: One binds BAF and the other binds DNA. EMBO J 20: 4399– 4407. Cai M, Huang Y, Suh JY, Louis JM, Ghirlando R, Craigie R, Clore GM. 2007. Solution NMR structure of the barrierto-autointegration factor-Emerin complex. J Biol Chem 282: 14525– 14535. Capell BC, Collins FS. 2006. Human laminopathies: Nuclei gone genetically awry. Nat Rev Genet 7: 940– 952. Caputo S, Couprie J, Duband-Goulet I, Konde E, Lin F, Braud S, Gondry M, Gilquin B, Worman HJ, Zinn-Justin S. 2006. The carboxyl-terminal nucleoplasmic region of MAN1 exhibits a DNA binding winged helix domain. J Biol Chem 281: 18208–18215. Chen IH, Huber M, Guan T, Bubeck A, Gerace L. 2006. Nuclear envelope transmembrane proteins (NETs) that are up-regulated during myogenesis. BMC Cell Biol 7: 38. Chikashige Y, Haraguchi T, Hiraoka Y. 2007. Another way to move chromosomes. Chromosoma 116: 497–505. Clements L, Manilal S, Love DR, Morris GE. 2000. Direct interaction between emerin and lamin A. Biochem Biophys Res Commun 267: 709 –714. Cohen TV, Kosti O, Stewart CL. 2007. The nuclear envelope protein MAN1 regulates TGFb signaling and vasculogenesis in the embryonic yolk sac. Development 134: 1385– 1395. Crisp M, Liu Q, Roux K, Rattner JB, Shanahan C, Burke B, Stahl PD, Hodzic D. 2006. Coupling of the nucleus and cytoplasm: Role of the LINC complex. J Cell Biol 172: 41–53. Dechat T, Gajewski A, Korbei B, Gerlich D, Daigle N, Haraguchi T, Furukawa K, Ellenberg J, Foisner R. 2004. LAP2a and BAF transiently localize to telomeres and specific regions on chromatin during nuclear assembly. J Cell Sci 117: 6117–6128. Dechat T, Korbei B, Vaughan OA, Vlcek S, Hutchison CJ, Foisner R. 2000. Lamina-associated polypeptide 2a binds intranuclear A-type lamins. J Cell Sci 113: 3473–3484. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832–853. Ding X, Xu R, Yu J, Xu T, Zhuang Y, Han M. 2007. SUN1 is required for telomere attachment to nuclear envelope and gametogenesis in mice. Dev Cell 12: 863– 872.

Dorner D, Vlcek S, Foeger N, Gajewski A, Makolm C, Gotzmann J, Hutchison CJ, Foisner R. 2006. Lamina-associated polypeptide 2a regulates cell cycle progression and differentiation via the retinoblastoma-E2F pathway. J Cell Biol 173: 83–93. Foisner R, Gerace L. 1993. Integral membrane proteins of the nuclear envelope interact with lamins and chromosomes, and binding is modulated by mitotic phosphorylation. Cell 73: 1267–1279. Furukawa K. 1999. LAP2 binding protein 1 (L2BP1/BAF) is a candidate mediator of LAP2- chromatin interaction. J Cell Sci 112: 2485–2492. Furukawa K, Fritze CE, Gerace L. 1998. The major nuclear envelope targeting domain of LAP2 coincides with its lamin binding region but is distinct from its chromatin interaction domain. J Biol Chem 273: 4213–4219. Furukawa K, Pante N, Aebi U, Gerace L. 1995. Cloning of a cDNA for lamina-associated polypeptide 2 (LAP2) and identification of regions that specify targeting to the nuclear envelope. EMBO J 14: 1626– 1636. Goldberg M, Lu H, Stuurman N, Ashery-Padan R, Weiss AM, Yu J, Bhattacharyya D, Fisher PA, Gruenbaum Y, Wolfner MF. 1998. Interactions among Drosophila nuclear envelope proteins lamin, otefin, and YA. Mol Cell Biol 18: 4315– 4323. Gonzalez JM, Navarro-Puche A, Casar B, Crespo P, Andres V. 2008. Fast regulation of AP-1 activity through interaction of lamin A/C, ERK1/2, and c-Fos at the nuclear envelope. J Cell Biol 183: 653– 666. Goodchild RE, Dauer WT. 2005. The AAAþ protein torsinA interacts with a conserved domain present in LAP1 and a novel ER protein. J Cell Biol 168: 855– 862. Gotzmann J, Foisner R. 2006. A-type lamin complexes and regenerative potential: A step towards understanding laminopathic diseases? Histochem Cell Biol 125: 33–41. Grady RM, Starr DA, Ackerman GL, Sanes JR, Han M. 2005. Syne proteins anchor muscle nuclei at the neuromuscular junction. Proc Natl Acad Sci 102: 4359–4364. Gros-Louis F, Dupre N, Dion P, Fox MA, Laurent S, Verreault S, Sanes JR, Bouchard JP, Rouleau GA. 2007. Mutations in SYNE1 lead to a newly discovered form of autosomal recessive cerebellar ataxia. Nat Genet 39: 80– 85. Gruenbaum Y, Lee KK, Liu J, Cohen M, Wilson KL. 2002. The expression, lamin-dependent localization and RNAi depletion phenotype for emerin in C. elegans. J Cell Sci 115: 923 –929. Gruenbaum Y, Margalit A, Goldman RD, Shumaker DK, Wilson KL. 2005. The nuclear lamina comes of age. Nat Rev Mol Cell Biol 6: 21–31. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W, et al. 2008. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453: 948–951. Guttinger S, Laurell E, Kutay U. 2009. Orchestrating nuclear envelope disassembly and reassembly during mitosis. Nat Rev Mol Cell Biol 10: 178 –191. Hakelien AM, Delbarre E, Gaustad KG, Buendia B, Collas P. 2008. Expression of the myodystrophic R453W mutation

Lamin-binding Proteins

of lamin A in C2C12 myoblasts causes promoter-specific and global epigenetic defects. Exp Cell Res 314: 1869– 1880. Han X, Feng X, Rattner JB, Smith H, Bose P, Suzuki K, Soliman MA, Scott MS, Burke BE, Riabowol K. 2008. Tethering by lamin A stabilizes and targets the ING1 tumour suppressor. Nat Cell Biol 10: 1333–1340. Haque F, Lloyd DJ, Smallwood DT, Dent CL, Shanahan CM, Fry AM, Trembath RC, Shackleton S. 2006. SUN1 interacts with nuclear lamin A and cytoplasmic nesprins to provide a physical connection between the nuclear lamina and the cytoskeleton. Mol Cell Biol 26: 3738–3751. Haraguchi T, Holaska JM, Yamane M, Koujin T, Hashiguchi N, Mori C, Wilson KL, Hiraoka Y. 2004. Emerin binding to Btf, a death-promoting transcriptional repressor, is disrupted by a missense mutation that causes EmeryDreifuss muscular dystrophy. Eur J Biochem 271: 1035– 1045. Haraguchi T, Koujin T, Segura-Totten M, Lee KK, Matsuoka Y, Yoneda Y, Wilson KL, Hiraoka Y. 2001. BAF is required for emerin assembly into the reforming nuclear envelope. J Cell Sci 114: 4575–4585. Harris CA, Andryuk PJ, Cline S, Chan HK, Natarajan A, Siekierka JJ, Goldstein G. 1994. Three distinct human thymopoietins are derived from alternatively spliced mRNAs. Proc Natl Acad Sci 91: 6283–6287. Hellemans J, Preobrazhenska O, Willaert A, Debeer P, Verdonk PC, Costa T, Janssens K, Menten B, Van Roy N, Vermeulen SJ, et al. 2004. Loss-of-function mutations in LEMD3 result in osteopoikilosis, Buschke-Ollendorff syndrome and melorheostosis. Nat Genet 36: 1213– 1218. Hewett JW, Zeng J, Niland BP, Bragg DC, Breakefield XO. 2006. Dystonia-causing mutant torsinA inhibits cell adhesion and neurite extension through interference with cytoskeletal dynamics. Neurobiol Dis 22: 98–111. Hirano Y, Segawa M, Ouchi FS, Yamakawa Y, Furukawa K, Takeyasu K, Horigome T. 2005. Dissociation of emerin from barrier-to-autointegration factor is regulated through mitotic phosphorylation of emerin in a Xenopus egg cell-free system. J Biol Chem 280: 39925–39933. Hodzic DM, Yeater DB, Bengtsson L, Otto H, Stahl PD. 2004. Sun2 is a novel mammalian inner nuclear membrane protein. J Biol Chem 279: 25805– 25812. Hoffmann K, Dreger CK, Olins AL, Olins DE, Shultz LD, Lucke B, Karl H, Kaps R, Muller D, Vaya A, et al. 2002. Mutations in the gene encoding the lamin B receptor produce an altered nuclear morphology in granulocytes (Pelger-Huet anomaly). Nat Genet 31: 410– 414. Hoffmann K, Sperling K, Olins AL, Olins DE. 2007. The granulocyte nucleus and lamin B receptor: Avoiding the ovoid. Chromosoma 116: 227– 235. Holaska JM, Wilson KL. 2006. Multiple roles for emerin: Implications for Emery-Dreifuss muscular dystrophy. Anat Rec A Discov Mol Cell Evol Biol 288: 676– 680. Holaska JM, Wilson KL. 2007. An emerin “proteome”: Purification of distinct emerin-containing complexes from HeLa cells suggests molecular basis for diverse roles including gene regulation, mRNA splicing, signaling, mechanosensing, and nuclear architecture. Biochemistry 46: 8897–8908. Holaska JM, Kowalski AK, Wilson KL. 2004. Emerin caps the pointed end of actin filaments: Evidence for an actin

cortical network at the nuclear inner membrane. PLoS Biol 2: E231. Holaska JM, Rais-Bahrami S, Wilson KL. 2006. Lmo7 is an emerin-binding protein that regulates the transcription of emerin and many other muscle-relevant genes. Hum Mol Genet 15: 3459– 3472. Holaska JM, Lee KK, Kowalski AK, Wilson KL. 2003. Transcriptional repressor germ cell-less (GCL) and barrier to autointegration factor (BAF) compete for binding to emerin in vitro. J Biol Chem 278: 6969–6975. Hozak P, Hassan AB, Jackson DA, Cook PR. 1993. Visualization of replication factories attached to nucleoskeleton. Cell 73: 361–373. Ivorra C, Kubicek M, Gonzalez JM, Sanz-Gonzalez SM, Alvarez-Barrientos A, O’Connor JE, Burke B, Andres V. 2006. A mechanism of AP-1 suppression through interaction of c-Fos with lamin A/C. Genes Dev 20: 307–320. Jiang X, Xia L, Chen D, Yang Y, Huang H, Yang L, Zhao Q, Shen L, Wang J. 2008. Otefin, a nuclear membrane protein, determines the fate of germline stem cells in Drosophila via interaction with Smad complexes. Dev Cell 14: 494 –506. Johnson BR, Nitta RT, Frock RL, Mounkes L, Barbie DA, Stewart CL, Harlow E, Kennedy BK. 2004. A-type lamins regulate retinoblastoma protein function by promoting subnuclear localization and preventing proteasomal degradation. Proc Natl Acad Sci 101: 9677– 9682. Karst ML, Herron KJ, Olson TM. 2008. X-linked nonsyndromic sinus node dysfunction and atrial fibrillation caused by emerin mutation. J Cardiovasc Electrophysiol 19: 510– 515. Kracklauer MP, Banks SM, Xie X, Wu Y, Fischer JA. 2007. Drosophila klaroid encodes a SUN domain protein required for Klarsicht localization to the nuclear envelope and nuclear migration in the eye. Fly (Austin) 1: 75–85. Laguri C, Gilquin B, Wolff N, Romi-Lebrun R, Courchay K, Callebaut I, Worman HJ, Zinn-Justin S. 2001. Structural characterization of the LEM motif common to three human inner nuclear membrane proteins. Structure 9: 503 –511. Lammerding J, Hsiao J, Schulze PC, Kozlov S, Stewart CL, Lee RT. 2005. Abnormal nuclear shape and impaired mechanotransduction in emerin-deficient cells. J Cell Biol 170: 781–791. Lee KK, Wilson KL. 2004. All in the family: Evidence for four new LEM-domain proteins Lem2 (NET-25), Lem3, Lem4 and Lem5 in the human genome. Symp Soc Exp Biol: 329–339. Lee KK, Gruenbaum Y, Spann P, Liu J, Wilson KL. 2000. C. elegans nuclear envelope proteins emerin, MAN1, lamin, and nucleoporins reveal unique timing of nuclear envelope breakdown during mitosis. Mol Biol Cell 11: 3089–3099. Lee KK, Haraguchi T, Lee RS, Koujin T, Hiraoka Y, Wilson KL. 2001. Distinct functional domains in emerin bind lamin A and DNA-bridging protein BAF. J Cell Sci 114: 4567–4573. Lee KK, Starr D, Cohen M, Liu J, Han M, Wilson KL, Gruenbaum Y. 2002. Lamin-dependent localization of UNC-84, a protein required for nuclear migration in Caenorhabditis elegans. Mol Biol Cell 13: 892– 901.

K.L. Wilson and R. Foisner

Li G, Sudlow G, Belmont AS. 1998. Interphase cell cycle dynamics of a late-replicating, heterochromatic homogeneously staining region: Precise choreography of condensation/decondensation and nuclear positioning. J Cell Biol 140: 975–989. Lin ST, Fu YH. 2009. miR-23 regulation of lamin B1 is crucial for oligodendrocyte development and myelination. Dis Model Mech 2: 178 –188. Lin F, Blake DL, Callebaut I, Skerjanc IS, Holmer L, McBurney MW, Paulin-Levasseur M, Worman HJ. 2000. MAN1, an inner nuclear membrane protein that shares the LEM domain with lamina-associated polypeptide 2 and emerin. J Biol Chem 275: 4840– 4847. Lin F, Morrison JM, Wu W, Worman HJ. 2005. MAN1, an integral protein of the inner nuclear membrane, binds Smad2 and Smad3 and antagonizes transforming growth factor-b signaling. Hum Mol Genet 14: 437 –445. Liu J, Lee KK, Segura-Totten M, Neufeld E, Wilson KL, Gruenbaum Y. 2003. MAN1 and emerin have overlapping function(s) essential for chromosome segregation and cell division in Caenorhabditis elegans. Proc Natl Acad Sci 100: 4598–4603. Liu Q, Pante N, Misteli T, Elsagga M, Crisp M, Hodzic D, Burke B, Roux KJ. 2007. Functional association of Sun1 with nuclear pore complexes. J Cell Biol 178: 785–798. Luderus ME, de Graaf A, Mattia E, den Blaauwen JL, Grande MA, de Jong L, van Driel R. 1992. Binding of matrix attachment regions to lamin B1. Cell 70: 949 –959. Luke Y, Zaim H, Karakesisoglou I, Jaeger VM, Sellin L, Lu W, Schneider M, Neumann S, Beijer A, Munck M, et al. 2008. Nesprin-2 Giant (NUANCE) maintains nuclear envelope architecture and composition in skin. J Cell Sci 121: 1887–1898. Luo Y, Yang C, Jin C, Xie R, Wang F, McKeehan WL. 2009. Novel phosphotyrosine targets of FGFR2IIIb signaling. Cell Signal 21: 1370– 1378. Makatsori D, Kourmouli N, Polioudaki H, Shultz LD, McLean K, Theodoropoulos PA, Singh PB, Georgatos SD. 2004. The inner nuclear membrane protein lamin B receptor forms distinct microdomains and links epigenetically marked chromatin to the nuclear envelope. J Biol Chem 279: 25567–25573. Malhas AN, Lee CF, Vaux DJ. 2009. Lamin B1 controls oxidative stress responses via Oct-1. J Cell Biol 184: 45–55. Malone CJ, Fixsen WD, Horvitz HR, Han M. 1999. UNC-84 localizes to the nuclear envelope and is required for nuclear migration and anchoring during C. elegans development. Development 126: 3171– 3181. Malone CJ, Misner L, Le Bot N, Tsai MC, Campbell JM, Ahringer J, White JG. 2003. The C. elegans hook protein, ZYG-12, mediates the essential attachment between the centrosome and nucleus. Cell 115: 825 –836. Manilal S, Nguyen TM, Sewry CA, Morris GE. 1996. The Emery-Dreifuss muscular dystrophy protein, emerin, is a nuclear membrane protein. Hum Mol Genet 5: 801– 808. Mansharamani M, Wilson KL. 2005. Direct binding of nuclear membrane protein MAN1 to emerin in vitro and two modes of binding to barrier-to-autointegration factor. J Biol Chem 280: 13863–13870.

Margalit A, Brachner A, Gotzmann J, Foisner R, Gruenbaum Y. 2007. Barrier-to-autointegration factor—a BAFfling little protein. Trends Cell Biol 17: 202–208. Margalit A, Segura-Totten M, Gruenbaum Y, Wilson KL. 2005a. Barrier-to-autointegration factor is required to segregate and enclose chromosomes within the nuclear envelope and assemble the nuclear lamina. Proc Natl Acad Sci 102: 3290– 3295. Margalit A, Vlcek S, Gruenbaum Y, Foisner R. 2005b. Breaking and making of the nuclear envelope. J Cell Biochem 95: 454– 465. Mariappan I, Gurung R, Thanumalayan S, Parnaik VK. 2007. Identification of cyclin D3 as a new interaction partner of lamin A/C. Biochem Biophys Res Commun 355: 981–985. Markiewicz E, Dechat T, Foisner R, Quinlan RA, Hutchison CJ. 2002. Lamin A/C binding protein LAP2a is required for nuclear anchorage of retinoblastoma protein. Mol Biol Cell 13: 4401–4413. Markiewicz E, Tilgner K, Barker N, van de Wetering M, Clevers H, Dorobek M, Hausmanowa-Petrusewicz I, Ramaekers FC, Broers JL, Blankesteijn WM, et al. 2006. The inner nuclear membrane protein emerin regulates b-catenin activity by restricting its accumulation in the nucleus. EMBO J 25: 3275–3285. Martin L, Crimaudo C, Gerace L. 1995. cDNA cloning and characterization of lamina-associated polypeptide 1C (LAP1C), an integral protein of the inner nuclear membrane. J Biol Chem 270: 8822–8828. Martins S, Eikvar S, Furukawa K, Collas P. 2003. HA95 and LAP2 b mediate a novel chromatin-nuclear envelope interaction implicated in initiation of DNA replication. J Cell Biol 160: 177– 188. Mattout A, Goldberg M, Tzur Y, Margalit A, Gruenbaum Y. 2007. Specific and conserved sequences in D. melanogaster and C. elegans lamins and histone H2A mediate the attachment of lamins to chromosomes. J Cell Sci 120: 77– 85. McGee MD, Rillo R, Anderson AS, Starr DA. 2006. UNC-83 is a KASH protein required for nuclear migration and is recruited to the outer nuclear membrane by a physical interaction with the SUN protein UNC-84. Mol Biol Cell 17: 1790– 1801. Mejat A, Decostre V, Li J, Renou L, Kesari A, Hantai D, Stewart CL, Xiao X, Hoffman E, Bonne G, et al. 2009. Lamin A/ C-mediated neuromuscular junction defects in EmeryDreifuss muscular dystrophy. J Cell Biol 184: 31– 44. Melcon G, Kozlov S, Cutler DA, Sullivan T, Hernandez L, Zhao P, Mitchell S, Nader G, Bakay M, Rottman JN, et al. 2006. Loss of emerin at the nuclear envelope disrupts the Rb1/E2F and MyoD pathways during muscle regeneration. Hum Mol Genet 15: 637– 651. Mislow JM, Holaska JM, Kim MS, Lee KK, Segura-Totten M, Wilson KL, McNally EM. 2002a. Nesprin-1a self-associates and binds directly to emerin and lamin A in vitro. FEBS Lett 525: 135– 140. Mislow JM, Kim MS, Davis DB, McNally EM. 2002b. Myne-1, a spectrin repeat transmembrane protein of the myocyte inner nuclear membrane, interacts with lamin A/C. J Cell Sci 115: 61–70. Moir RD, Spann TP, Herrmann H, Goldman RD. 2000a. Disruption of nuclear lamin organization blocks the

Lamin-binding Proteins

elongation phase of DNA replication. J Cell Biol 149: 1179– 1192. Moir RD, Yoon M, Khuon S, Goldman RD. 2000b. Nuclear lamins A and B1: different pathways of assembly during nuclear envelope formation in living cells. J Cell Biol 151: 1155–1168. Muchir A, Pavlidis P, Bonne G, Hayashi YK, Worman HJ. 2007. Activation of MAPK in hearts of EMD null mice: Similarities between mouse models of X-linked and autosomal dominant Emery Dreifuss muscular dystrophy. Hum Mol Genet 16: 1884–1895. Muchir A, Shan J, Bonne G, Lehnart SE, Worman HJ. 2009. Inhibition of extracellular signal-regulated kinase signaling to prevent cardiomyopathy caused by mutation in the gene encoding A-type lamins. Hum Mol Genet 18: 241–247. Naetar N, Hutter S, Dorner D, Dechat T, Korbei B, Gotzmann J, Beug H, Foisner R. 2007. LAP2a-binding protein LINT-25 is a novel chromatin-associated protein involved in cell cycle exit. J Cell Sci 120: 737– 747. Naetar N, Korbei B, Kozlov S, Kerenyi MA, Dorner D, Kral R, Gotic I, Fuchs P, Cohen TV, Bittner R, et al. 2008. Loss of nucleoplasmic LAP2a-lamin A complexes causes erythroid and epidermal progenitor hyperproliferation. Nat Cell Biol 10: 1341– 1348. Naismith TV, Heuser JE, Breakefield XO, Hanson PI. 2004. TorsinA in the nuclear envelope. Proc Natl Acad Sci 101: 7612–7617. Nery FC, Zeng J, Niland BP, Hewett J, Farley J, Irimia D, Li Y, Wiche G, Sonnenberg A, Breakefield XO. 2008. TorsinA binds the KASH domain of nesprins and participates in linkage between nuclear envelope and cytoskeleton. J Cell Sci 121: 3476– 3486. Nili E, Cojocaru GS, Kalma Y, Ginsberg D, Copeland NG, Gilbert DJ, Jenkins NA, Berger R, Shaklai S, Amariglio N, et al. 2001. Nuclear membrane protein LAP2b mediates transcriptional repression alone and together with its binding partner GCL (germ-cell-less). J Cell Sci 114: 3297– 3307. Nitta RT, Smith CL, Kennedy BK. 2007. Evidence that proteasome-dependent degradation of the retinoblastoma protein in cells lacking A-type lamins occurs independently of gankyrin and MDM2. PLoS ONE 2: e963. Nitta RT, Jameson SA, Kudlow BA, Conlan LA, Kennedy BK. 2006. Stabilization of the retinoblastoma protein by A-type nuclear lamins is required for INK4A-mediated cell cycle arrest. Mol Cell Biol 26: 5360– 5372. Osada S, Ohmori SY, Taira M. 2003. XMAN1, an inner nuclear membrane protein, antagonizes BMP signaling by interacting with Smad1 in Xenopus embryos. Development 130: 1783– 1794. Ottaviani A, Rival-Gervier S, Boussouar A, Foerster AM, Rondier D, Sacconi S, Desnuelle C, Gilson E, Magdinier F. 2009. The D4Z4 macrosatellite repeat acts as a CTCF and A-type lamins-dependent insulator in facioscapulo-humeral dystrophy. PLoS Genet 5: e1000394. Ozaki T, Saijo M, Murakami K, Enomoto H, Taya Y, Sakiyama S. 1994. Complex formation between lamin A and the retinoblastoma gene product: Identification of the domain on lamin A required for its interaction. Oncogene 9: 2649– 2653.

Ozawa R, Hayashi YK, Ogawa M, Kurokawa R, Matsumoto H, Noguchi S, Nonaka I, Nishino I. 2006. Emerinlacking mice show minimal motor and cardiac dysfunctions with nuclear-associated vacuoles. Am J Pathol 168: 907–917. Padiath QS, Saigoh K, Schiffmann R, Asahara H, Yamada T, Koeppen A, Hogan K, Ptacek LJ, Fu YH. 2006. Lamin B1 duplications cause autosomal dominant leukodystrophy. Nat Genet 38: 1114–1123. Padmakumar VC, Abraham S, Braune S, Noegel AA, Tunggal B, Karakesisoglou I, Korenbaum E. 2004. Enaptin, a giant actin-binding protein, is an element of the nuclear membrane and the actin cytoskeleton. Exp Cell Res 295: 330 –339. Padmakumar VC, Libotte T, Lu W, Zaim H, Abraham S, Noegel AA, Gotzmann J, Foisner R, Karakesisoglou I. 2005. The inner nuclear membrane protein Sun1 mediates the anchorage of Nesprin-2 to the nuclear envelope. J Cell Sci 118: 3419– 3430. Pan D, Estevez-Salmeron LD, Stroschein SL, Zhu X, He J, Zhou S, Luo K. 2005. The integral inner nuclear membrane protein MAN1 physically interacts with the R-Smad proteins to repress signaling by the transforming growth factor-b superfamily of cytokines. J Biol Chem 280: 15992–16001. Park YE, Hayashi YK, Bonne G, Arimura T, Noguchi S, Nonaka I, Nishino I. 2009a. Autophagic degradation of nuclear components in mammalian cells. Autophagy 5: 795 –804. Park YE, Hayashi YK, Goto K, Komaki H, Hayashi Y, Inuzuka T, Noguchi S, Nonaka I, Nishino I. 2009b. Nuclear changes in skeletal muscle extend to satellite cells in autosomal dominant Emery-Dreifuss muscular dystrophy/ limb-girdle muscular dystrophy 1B. Neuromuscul Disord 19: 29–36. Patterson K, Molofsky AB, Robinson C, Acosta S, Cater C, Fischer JA. 2004. The functions of Klarsicht and nuclear lamin in developmentally regulated nuclear migrations of photoreceptor cells in the Drosophila eye. Mol Biol Cell 15: 600–610. Pekovic V, Harborth J, Broers JL, Ramaekers FC, van Engelen B, Lammens M, von Zglinicki T, Foisner R, Hutchison C, Markiewicz E. 2007. Nucleoplasmic LAP2a-lamin A complexes are required to maintain a proliferative state in human fibroblasts. J Cell Biol 176: 163 –172. Penkner A, Tang L, Novatchkova M, Ladurner M, Fridkin A, Gruenbaum Y, Schweizer D, Loidl J, Jantsch V. 2007. The nuclear envelope protein Matefin/SUN-1 is required for homologous pairing in C. elegans meiosis. Dev Cell 12: 873 –885. Pickersgill H, Kalverda B, de Wit E, Talhout W, Fornerod M, van Steensel B. 2006. Characterization of the Drosophila melanogaster genome at the nuclear lamina. Nat Genet 38: 1005–1014. Polioudaki H, Kourmouli N, Drosou V, Bakou A, Theodoropoulos PA, Singh PB, Giannakouros T, Georgatos SD. 2001. Histones H3/H4 form a tight complex with the inner nuclear membrane protein LBR and heterochromatin protein 1. EMBO Rep 2: 920 –925. Puckelwartz MJ, Kessler E, Zhang Y, Hodzic D, Randles KN, Morris G, Earley JU, Hadhazy M, Holaska JM, Mewborn

K.L. Wilson and R. Foisner

SK, et al. 2009. Disruption of nesprin-1 produces an Emery Dreifuss muscular dystrophy-like phenotype in mice. Hum Mol Genet 18: 607 –620. Raju GP, Dimova N, Klein PS, Huang HC. 2003. SANE, a novel LEM domain protein, regulates bone morphogenetic protein signaling through interaction with Smad1. J Biol Chem 278: 428–437. Roberts RC, Sutherland-Smith AJ, Wheeler MA, Jensen ON, Emerson LJ, Spiliotis II, Tate CG, Kendrick-Jones J, Ellis JA. 2006. The Emery-Dreifuss muscular dystrophy associated-protein emerin is phosphorylated on serine 49 by protein kinase A. Febs J 273: 4562–4575. Roux KJ, Crisp ML, Liu Q, Kim D, Kozlov S, Stewart CL, Burke B. 2009. Nesprin 4 is an outer nuclear membrane protein that can induce kinesin-mediated cell polarization. Proc Natl Acad Sci 106: 2194–2199. Sabatelli P, Lattanzi G, Ognibene A, Columbaro M, Capanni C, Merlini L, Maraldi NM, Squarzoni S. 2001. Nuclear alterations in autosomal-dominant Emery-Dreifuss muscular dystrophy. Muscle Nerve 24: 826–829. Sakaki M, Koike H, Takahashi N, Sasagawa N, Tomioka S, Arahata K, Ishiura S. 2001. Interaction between emerin and nuclear lamins. J. Biochem 129: 321 –327. Salpingidou G, Smertenko A, Hausmanowa-Petrucewicz I, Hussey PJ, Hutchison CJ. 2007. A novel role for the nuclear membrane protein emerin in association of the centrosome to the outer nuclear membrane. J Cell Biol 178: 897– 904. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059– 1063. Scaffidi P, Misteli T. 2008. Lamin A-dependent misregulation of adult stem cells associated with accelerated ageing. Nat Cell Biol 10: 452–459. Schirmer EC, Foisner R. 2007. Proteins that associate with lamins: Many faces, many functions. Exp Cell Res 313: 2167– 2179. Schirmer EC, Gerace L. 2005. The nuclear membrane proteome: Extending the envelope. Trends Biochem Sci 30: 551–558. Schirmer EC, Florens L, Guan T, Yates JR 3rd, Gerace L. 2003. Nuclear membrane proteins with potential disease links found by subtractive proteomics. Science 301: 1380– 1382. Schirmer EC, Florens L, Guan T, Yates JR 3rd, Gerace L. 2005. Identification of novel integral membrane proteins of the nuclear envelope with potential disease links using subtractive proteomics. Novartis Found. Symp 264: 63–76; discussion 76– 80, 227– 230. Schlosser A, Amanchy R, Otto H. 2006. Identification of tyrosine-phosphorylation sites in the nuclear membrane protein emerin. FEBS J 273: 3204– 3215. Schmitt J, Benavente R, Hodzic D, Hoog C, Stewart CL, Alsheimer M. 2007. Transmembrane protein Sun2 is involved in tethering mammalian meiotic telomeres to the nuclear envelope. Proc Natl Acad Sci 104: 7426– 7431. Senior A, Gerace L. 1988. Integral membrane proteins specific to the inner nuclear membrane and associated with the nuclear lamina. J Cell Biol 107: 2029– 2036. Shimi T, Koujin T, Segura-Totten M, Wilson KL, Haraguchi T, Hiraoka Y. 2004. Dynamic interaction between BAF

and emerin revealed by FRAP, FLIP, and FRET analyses in living HeLa cells. J Struct Biol 147: 31–41. Shoeman RL, Traub P. 1990. The in vitro DNA-binding properties of purified nuclear lamin proteins and vimentin. J Biol Chem 265: 9055–9061. Shumaker DK, Dechat T, Kohlmaier A, Adam SA, Bozovsky MR, Erdos MR, Eriksson M, Goldman AE, Khuon S, Collins FS, et al. 2006. Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci 103: 8703– 8708. Shumaker DK, Lee KK, Tanhehco YC, Craigie R, Wilson KL. 2001. LAP2 binds to BAF.DNA complexes: requirement for the LEM domain and modulation by variable regions. EMBO J 20: 1754–1764. Shumaker DK, Solimando L, Sengupta K, Shimi T, Adam SA, Grunwald A, Strelkov SV, Aebi U, Cardoso MC, Goldman RD. 2008. The highly conserved nuclear lamin Ig-fold binds to PCNA: Its role in DNA replication. J Cell Biol 181: 269– 280. Somech R, Shaklai S, Geller O, Amariglio N, Simon AJ, Rechavi G, Gal-Yam EN. 2005. The nuclear-envelope protein and transcriptional repressor LAP2b interacts with HDAC3 at the nuclear periphery, and induces histone H4 deacetylation. J Cell Sci 118: 4017–4025. Spann TP, Moir RD, Goldman AE, Stick R, Goldman RD. 1997. Disruption of nuclear lamin organization alters the distribution of replication factors and inhibits DNA synthesis. J Cell Biol 136: 1201– 1212. Starr DA. 2009. A nuclear-envelope bridge positions nuclei and moves chromosomes. J Cell Sci 122: 577–586. Starr DA, Fischer JA. 2005. KASH ’n Karry: The KASH domain family of cargo-specific cytoskeletal adaptor proteins. Bioessays 27: 1136– 1146. Starr DA, Han M. 2002. Role of ANC-1 in tethering nuclei to the actin cytoskeleton. Science 298: 406–409. Starr DA, Hermann GJ, Malone CJ, Fixsen W, Priess JR, Horvitz HR, Han M. 2001. unc-83 encodes a novel component of the nuclear envelope and is essential for proper nuclear migration. Development 128: 5039–5050. Stewart CL, Roux KJ, Burke B. 2007. Blurring the boundary: The nuclear envelope extends its reach. Science 318: 1408–1412. Stewart-Hutchinson PJ, Hale CM, Wirtz D, Hodzic D. 2008. Structural requirements for the assembly of LINC complexes and their function in cellular mechanical stiffness. Exp Cell Res 314: 1892–1905. Stierle V, Couprie J, Ostlund C, Krimm I, Zinn-Justin S, Hossenlopp P, Worman HJ, Courvalin JC, DubandGoulet I. 2003. The carboxyl-terminal region common to lamins A and C contains a DNA binding domain. Biochemistry 42: 4819– 4828. Sullivan T, Escalante-Alcalde D, Bhatt H, Anver M, Bhat N, Nagashima K, Stewart CL, Burke B. 1999. Loss of A-type lamin expression compromises nuclear envelope integrity leading to muscular dystrophy. J Cell Biol 147: 913 –920. Tang K, Finley RL Jr, Nie D, Honn KV. 2000. Identification of 12-lipoxygenase interaction with cellular proteins by yeast two-hybrid screening. Biochemistry 39: 3185–3191.

Lamin-binding Proteins

Taniura H, Glass C, Gerace L. 1995. A chromatin binding site in the tail domain of nuclear lamins that interacts with core histones. J Cell Biol 131: 33–44. Taylor MR, Slavov D, Gajewski A, Vlcek S, Ku L, Fain PR, Carniel E, Di Lenarda A, Sinagra G, Boucek MM, et al. 2005. Thymopoietin (lamina-associated polypeptide 2) gene mutation associated with dilated cardiomyopathy. Hum Mutat 26: 566 –574. Tifft KE, Bradbury KA, Wilson KL. 2009. Tyrosine phosphorylation of nuclear-membrane protein emerin by Src, Abl and other kinases. J Cell Sci 122: 3780–3790. Tilgner K, Wojciechowicz K, Jahoda C, Hutchison C, Markiewicz E. 2009. Dynamic complexes of A-type lamins and emerin influence adipogenic capacity of the cell via nucleocytoplasmic distribution of b-catenin. J Cell Sci 122: 401– 413. Tzur YB, Wilson KL, Gruenbaum Y. 2006b. SUN-domain proteins: ’Velcro’ that links the nucleoskeleton to the cytoskeleton. Nat Rev Mol Cell Biol 7: 782– 788. Tzur YB, Margalit A, Melamed-Book N, Gruenbaum Y. 2006a. Matefin/SUN-1 is a nuclear envelope receptor for CED-4 during Caenorhabditis elegans apoptosis. Proc Natl Acad Sci 103: 13397– 13402. Ulbert S, Antonin W, Platani M, Mattaj IW. 2006. The inner nuclear membrane protein Lem2 is critical for normal nuclear envelope morphology. FEBS Lett 580: 6435– 6441. Ura S, Hayashi YK, Goto K, Astejada MN, Murakami T, Nagato M, Ohta S, Daimon Y, Takekawa H, Hirata K, et al. 2007. Limb-girdle muscular dystrophy due to emerin gene mutations. Arch Neurol 64: 1038–1041. Van Berlo JH, Voncken JW, Kubben N, Broers JL, Duisters R, van Leeuwen RE, Crijns HJ, Ramaekers FC, Hutchison CJ, Pinto YM. 2005. A-type lamins are essential for TGF-b1 induced PP2A to dephosphorylate transcription factors. Hum Mol Genet 14: 2839–2849. Vaughan A, Alvarez-Reyes M, Bridger JM, Broers JL, Ramaekers FC, Wehnert M, Morris GE, Whitfield WGF, Hutchison CJ. 2001. Both emerin and lamin C depend on lamin A for localization at the nuclear envelope. J Cell Sci 114: 2577–2590. Vlcek S, Foisner R. 2007. A-type lamin networks in light of laminopathic diseases. Biochim Biophys Acta 1773: 661–674. Vlcek S, Foisner R, Wilson KL. 2004. Lco1 is a novel widely expressed lamin-binding protein in the nuclear interior. Exp Cell Res 298: 499 –511. Vlcek S, Korbei B, Foisner R. 2002. Distinct functions of the unique C terminus of LAP2a in cell proliferation and nuclear assembly. J Biol Chem 277: 18898–18907. Wagner N, Krohne G. 2007. LEM-Domain proteins: New insights into lamin-interacting proteins. Int Rev Cytol 261: 1 –46. Wagner N, Schmitt J, Krohne G. 2004. Two novel LEMdomain proteins are splice products of the annotated Drosophila melanogaster gene CG9424 (Bocksbeutel). Eur J Cell Biol 82: 605– 616. Wagner N, Kagermeier B, Loserth S, Krohne G. 2006. The Drosophila melanogaster LEM-domain protein MAN1. Eur J Cell Biol 85: 91–105.

Wang Y, Herron AJ, Worman HJ. 2006. Pathology and nuclear abnormalities in hearts of transgenic mice expressing M371K lamin A encoded by an LMNA mutation causing Emery-Dreifuss muscular dystrophy. Hum Mol Genet 15: 2479–2489. Waterham HR, Koster J, Mooyer P, Noort Gv G, Kelley RI, Wilcox WR, Wanders RJ, Hennekam RC, Oosterwijk JC. 2003. Autosomal recessive HEM/Greenberg skeletal dysplasia is caused by 3 b-hydroxysterol delta 14-reductase deficiency due to mutations in the lamin B receptor gene. Am J Hum Genet 72: 1013–1017. Wheeler MA, Davies JD, Zhang Q, Emerson LJ, Hunt J, Shanahan CM, Ellis JA. 2007. Distinct functional domains in nesprin-1a and nesprin-2b bind directly to emerin and both interactions are disrupted in X-linked Emery-Dreifuss muscular dystrophy. Exp Cell Res 313: 2845–2857. Wilhelmsen K, Ketema M, Truong H, Sonnenberg A. 2006. KASH-domain proteins in nuclear migration, anchorage and other processes. J Cell Sci 119: 5021–5029. Wilhelmsen K, Litjens SH, Kuikman I, Tshimbalanga N, Janssen H, van den Bout I, Raymond K, Sonnenberg A. 2005. Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. J Cell Biol 171: 799– 810. Worman HJ, Bonne G. 2007. “Laminopathies”: Awide spectrum of human diseases. Exp Cell Res 313: 2121– 2133. Worman HJ, Evans CD, Blobel G. 1990. The lamin B receptor of the nuclear envelope inner membrane: A polytopic protein with eight potential transmembrane domains. J Cell Biol 111: 1535–1542. Worman HJ, Yuan J, Blobel G, Georgatos SD. 1988. A lamin B receptor in the nuclear envelope. Proc Natl Acad Sci 85: 8531–8534. Xie X, Fischer JA. 2008. On the roles of the Drosophila KASH domain proteins Msp-300 and Klarsicht. Fly 2: 74– 81. Ye Q, Worman HJ. 1996. Interaction between an integral protein of the nuclear envelope inner membrane and human chromodomain proteins homologous to Drosophila HP1. J Biol Chem 271: 14653–14656. Zastrow MS, Vlcek S, Wilson KL. 2004. Proteins that bind A-type lamins: Integrating isolated clues. J Cell Sci 117: 979 –987. Zhang Q, Ragnauth CD, Skepper JN, Worth NF, Warren DT, Roberts RG, Weissberg PL, Ellis JA, Shanahan CM. 2005. Nesprin-2 is a multi-isomeric protein that binds lamin and emerin at the nuclear envelope and forms a subcellular network in skeletal muscle. J Cell Sci 118: 673 –687. Zhang Q, Skepper JN, Yang F, Davies JD, Hegyi L, Roberts RG, Weissberg PL, Ellis JA, Shanahan CM. 2001. Nesprins: A novel family of spectrin-repeat-containing proteins that localize to the nuclear membrane in multiple tissues. J Cell Sci 114: 4485– 4498. Zhang X, Xu R, Zhu B, Yang X, Ding X, Duan S, Xu T, Zhuang Y, Han M. 2007. Syne-1 and Syne-2 play crucial roles in myonuclear anchorage and motor neuron innervation. Development 134: 901 –908. Zhen YY, Libotte T, Munck M, Noegel AA, Korenbaum E. 2002. NUANCE, a giant protein connecting the nucleus and actin cytoskeleton. J Cell Sci 115: 3207– 3222.

The Nuclear Pore Complex and Nuclear Transport Susan R. Wente1 and Michael P. Rout2 1

Department of Cell and Developmental Biology, Vanderbilt University Medical Center, Nashville, Tennessee 37232

2

Laboratory of Cellular and Structural Biology, The Rockefeller University, New York, New York 10065

Correspondence: [email protected] and [email protected]

Internal membrane bound structures sequester all genetic material in eukaryotic cells. The most prominent of these structures is the nucleus, which is bounded by a double membrane termed the nuclear envelope (NE). Though this NE separates the nucleoplasm and genetic material within the nucleus from the surrounding cytoplasm, it is studded throughout with portals called nuclear pore complexes (NPCs). The NPC is a highly selective, bidirectional transporter for a tremendous range of protein and ribonucleoprotein cargoes. All the while the NPC must prevent the passage of nonspecific macromolecules, yet allow the free diffusion of water, sugars, and ions. These many types of nuclear transport are regulated at multiple stages, and the NPC carries binding sites for many of the proteins that modulate and modify the cargoes as they pass across the NE. Assembly, maintenance, and repair of the NPC must somehow occur while maintaining the integrity of the NE. Finally, the NPC appears to be an anchor for localization of many nuclear processes, including gene activation and cell cycle regulation. All these requirements demonstrate the complex design of the NPC and the integral role it plays in key cellular processes.

axonomically speaking, all life on earth falls into one of two fundamental groups, the prokaryotes and the eukaryotes. The prokaryotes, the first group to evolve, are single cell organisms bounded by a single membrane. About 1.5 billion years later, a series of evolutionary innovations led to the emergence of eukaryotes. Eukaryotes have multiple inner membrane structures that allow for compartmentalization within the cell, and therefore differentiation of the cell and regulation within it. Ultimately, the greater cellular complexity of

T

eukaryotes allowed them to adopt a multicellular lifestyle, as seen in the plants, fungi and animals of today (reviewed in Field and Dacks 2009). Internal membrane bound structures sequester all genetic material in eukaryotic cells. The most prominent of these structures, which gives the eukaryotes their Greek-rooted name, is the nucleus—the central “kernel” (gr. “karyo-”) of the cell. The nucleus is bounded by a double membrane termed the nuclear envelope (NE), which separates the nucleoplasm and genetic

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000562 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000562

S.R. Wente and M.P. Rout

material from the surrounding cytoplasm. However the genetic material in the nucleus is not totally isolated from the rest of the cell. Studded throughout the NE are portals called nuclear pore complexes (NPCs). The NPC is a highly selective, bidirectional transporter for a tremendous range of cargoes. Going into the nucleus, these cargoes include inner nuclear membrane proteins and all the proteins in the nucleoplasm. Going out are RNA-associated proteins that are assembled into ribosomal subunits or messenger ribonucleoproteins (mRNPs). Once transported, the NPC must ensure these cargos are retained in their respective nuclear and cytoplasmic compartments. All the while the NPC must prevent the passage of nonspecific macromolecules, yet allow the free diffusion of water, sugars, and ions. These many types of nuclear transport are regulated at multiple stages, providing a powerful extra level of cellular control that is not necessary in prokaryotes. Assembly, maintenance, and repair of the NPC must somehow occur while maintaining the integrity of the NE. Finally, the NPC appears to be an anchor for localization of many nuclear processes, including gene activation and cell cycle regulation (reviewed in Ahmed and Brickner 2007; Hetzer and Wente 2009). All these requirements demonstrate the complex design of the NPC and the integral role it plays in key cellular processes. STRUCTURE OF THE NPC: SET UP OF THE MACHINE

The specifications of the NPC’s transport machinery represent a huge engineering challenge for evolution. No transitional forms of this elaborate transport system have yet been found in modern day organisms to reveal how it evolved. However, recent clues show that the NPC itself retains in its core a fossil of its ancient origins, indicating that the same mechanism that generated the internal membranes of eukaryotes might also have been responsible for the NPCs and the transport machinery. In the electron microscope, the NPC appears as a complex cylindrical structure with strong octagonal symmetry, measuring some

100 – 150 nm in diameter and 50 – 70 nm in thickness depending on the organism (reviewed in Wente 2000; Lim et al. 2008). This overall appearance seems broadly conserved throughout all eukaryotes. The two membranes of the NE, the outer and inner membranes, join only in a specialized, sharply curved piece of “pore membrane” that forms a grommet in the NE within which the NPC sits. Within each NPC is a core structure containing eight spokes surrounding a central tube. This central hole (30 nm diameter and 50 nm long) is where the nucleoplasm connects to the cytoplasm and where macromolecular exchange occurs. Peripheral filaments are attached to the core, filling the central hole as well as emanating into the nucleoplasm and cytoplasm. These filaments form a basket-like structure on the nuclear side of the NPC (Fig. 1). One can envision the NPC as being comprised of layers of interacting proteins, starting with the core structure, moving outwards through its peripheral filaments, and then to associating clouds of soluble transport factors and peripherally associating protein complexes in the nucleus and cytoplasm (Rout and Aitchison 2001). These protein interactions can occur on radically different time scales. Some proteins form relatively permanent associations with the core structure, and so are termed nuclear pore complex components or “nucleoporins” (“Nups”). Other proteins associate transiently with the NPC, either constantly cycling on and off or attaching only at particular times in the cell’s life cycle. The NPC is covered in binding sites for these transiently associating proteins. Because the NPC is neither a motor nor an enzyme, the interactions provided by its binding sites wholly define the function of the NPC. Recent work, mainly in the yeast Saccharomyces cerevisiae and in vertebrates, has begun to elucidate the molecular architecture of the NPC (Rout et al. 2000; Cronshaw et al. 2002; Alber et al. 2007b). Given its large size, the main body of the NPC comprises a surprisingly small number of 30 unique proteins (Table 1). However, because of the NPC’s eightfold symmetry, these Nups are each present in

Nuclear Pore Complexes and Nuclear Transport Central tube

Core scaffold

Outer ring Inner ring

Lumenal ring

Cytoplasmic filaments Cytoplasmic FG nups Symmetric FG nups Nuclear envelope

Pore membrane

Linker nups

Nucleoplasmic FG nups Nuclear basket

Figure 1. Major structural features of the NPC (based on the architectural map of Alber et al. (2007b); see Table 1

and main text for details).

multiple copies (usually 16 per NPC) resulting in around 400 polypeptides for each NPC in every eukaryote (Rout et al. 2000; Cronshaw et al. 2002; DeGrasse et al. 2009). Further redundancy is evident from the recent mapping of the yeast NPC. Indeed, the NPC’s structure is modular, consisting of a few highly repetitive protein fold types (Devos et al. 2006; Alber et al. 2007b; DeGrasse et al. 2009). This suggests that the bulk of the NPC’s structure has evolved through multiple duplications of a small precursor set of genes encoding just a handful of progenitor Nups.

To understand its evolutionary origins, the NPC of the highly divergent Trypanosoma was recently characterized (DeGrasse et al. 2009). Despite significant divergence in primary structure, the Trypanosome NPC consists mainly of the motifs and domains found in vertebrate and yeast NPCs, indicating on a molecular level that the basic structural components of the NPC are conserved across all eukaryotes. Importantly, this also strongly implies that the last common eukaryotic ancestor had many features in common with contemporary NPCs, and perhaps provided a key adaptive advantage

Table 1. Nucleoporin homologs of yeast and vertebrates NPC substructure

Outer Ring

Inner Ring

Cytoplasmic FG Nups and Filaments Lumenal Ring Symmetric FG Nups Linker Nups Nucleoplasmic FG Nups and Filaments

Yeast components

Vertebrate components

Nup84 subcomplex (Nup84, Nup85, Nup120, Nup133, Nup145C, Sec13, Seh1) Nup170 subcomplex (Nup170, Nup157, Nup188, Nup192, Nup59, Nup53) Nup159, Nup42

Nup107-160 complex (Nup160, Nup133, Nup107, Nup96, Nup75, Seh1, Sec13, Aladin, Nup43, Nup37) Nup155 subcomplex (Nup155, Nup205, Nup188, Nup35)

Ndc1, Pom152, Pom34 Nsp1, Nup57, Nup49, Nup145N, Nup116, Nup100 Nup82, Nic96 Nup1, Nup60, Mlp1, Mlp2

Gp210, Ndc1, Pom121 Nup62, Nup58/45, Nup54, Nup98

Nup358, Nup214, Nlp1

Nup88, Nup93 Nup153, Tpr

S.R. Wente and M.P. Rout

for this organism that has been retained, little changed, ever since. The structural proteins making up the bulk of the spokes and rings give the NPC its shape and strength (Fig. 1). These core proteins of the NPC also maintain the stability of the nuclear envelope and facilitate the bending of the pore membrane into the inner and outer NE membranes. The most equatorial rings, termed the inner rings, are comprised of the Nup170 complex (yeast) or Nup155 complex (vertebrates) (Aitchison et al. 1995; Grandi et al. 1997; Miller et al. 2000) (Fig. 1). The inner rings are sandwiched between the outer rings, which are comprised of the Nup84 complex (yeast) or Nup107 complex (vertebrates) (Table 1) (Siniossoglou et al. 1996; Fontoura et al. 1999; Siniossoglou et al. 2000; Belgareh et al. 2001; Vasu et al. 2001). Together, these Nup complexes form a scaffold that hugs the curved surface of the pore membrane and helps form the central tube through which macromolecular exchange occurs (Alber et al. 2007a; Alber et al. 2007b). Nups in the core scaffold represent roughly half the mass of the whole NPC and are composed almost entirely of either b-propeller folds, a-solenoid folds, or a distinct arrangement of both in an amino-terminal b-propeller followed by a carboxy-terminal a-solenoid fold. The core scaffold of all eukaryotes appears to retain this basic fold composition (Devos et al. 2004; Devos et al. 2006; DeGrasse et al. 2009). Strikingly, there are similarities between the structures of the core NPC scaffold curving around the pore membrane and other membrane-associated complexes such as clathrin/ adaptin, COPI, and COPII (Fig. 1) (Devos et al. 2004; Devos et al. 2006). Clathrin/adaptin is involved in coat-mediated endocytosis at the plasma membrane, and COPI and COPII are responsible for coat-mediated vesicular transport between the plasma membrane and endomembrane systems such as the Golgi and ER. Indeed, the similarities between core scaffold Nups and coating complexes have been borne out in numerous crystallographic studies (Berke et al. 2004; Hsia et al. 2007; Brohawn et al. 2008; Debler et al. 2008; Brohawn et al. 2009; Leksa et al. 2009; Seo et al. 2009; Whittle and Schwartz

2009), although nearly 2 billion years of evolution have made it difficult at first glance to recognize the common origin of these two groups. However, their common b-propeller and helixturn-helix repeat structure is still unmistakable (Brohawn et al. 2008; Field and Dacks 2009). In NPCs the “coat” comprises the core scaffold of the NPC, where—analogous to the curved membrane of a vesicle being stabilized by a COP or clathrin coat—it stabilizes the curved pore membrane. These similarities also give a tantalizing glimpse into the deep evolutionary origins of eukaryotes. It seems early protoeukaryotes distinguished themselves from their prokaryotes by acquiring a membrane-curving protein module, the “proto-coatomer” (likely composed of a simple b-propeller/a-solenoid protein), that allowed them to mold their plasma membranes into internal compartments. Modern eukaryotes diversified this module into many specialized membrane coating complexes, accounting for the evolution of their internal membrane systems (Devos et al. 2004; Devos et al. 2006). The framework of the NPC serves two key transport purposes: to form a barrier of defined permeability within the pore, and to facilitate transport of selected macromolecules across it. Both processes are dependent on the correct positioning of critical Nups in the NPC architecture (Radu et al. 1995; Strawn et al. 2004; Liu and Stewart 2005). Attached to the inside face of the NPC core scaffold, facing the central tube’s cavity, are groups of nucleoporins termed “linker nucleoporins” (Fig. 1). Together with the inner ring, these seem to form most of the attachment sites for a last set of nucleoporins, termed “FG Nups” (Alber et al. 2007b). These FG Nups, named for their phenylalanine-glycine repeats, are the direct mediators of nucleocytoplasmic transport (Radu et al. 1995; Strawn et al. 2004; Liu and Stewart 2005) (see the following section). The core NPC scaffold is connected to a set of integral membrane proteins, which form an outer luminal ring in the NE lumen and anchor the NPC into the NE (Nehrbass et al. 1996; Alber et al. 2007a; Alber et al. 2007b) (Fig. 1). Oddly, the membrane nucleoporins seem poorly

Nuclear Pore Complexes and Nuclear Transport

conserved—if at all—across the eukaryotes. The fact that all the currently known pore membrane proteins in Aspergillus nidulans can seemingly be dispensed with for NPC function and assembly might indicate that there are not strong pressures for their conservation, and that there are other membrane proteins that can serve the role (Liu et al. 2009). This fact also sets up a quandary—if most or all of the NPC’s presumed membrane anchors are dispensible, how then is the NPC reliably anchored to the membrane? Several groups are seeking the answer to this question (Hetzer and Wente 2009). OPERATION OF THE MACHINE: THE SOLUBLE PHASE

Understanding the transport machine requires resolving both its barrier and binding activities. How the NPC machine balances both of these selective functions has been a challenging mystery. Studies of fluorescently labeled sized dextrans or gold particles microinjected into cells (Feldherr and Akin 1997; Keminer and Peters 1999) have defined the practical permeability limits of the NPC, showing that under physiological time scales, macromolecules greater than 40 kDa in size do not show any measureable redistribution between the nucleus and cytoplasm and thus, no movement through the NPC. Conversely, metal ions, small metabolites, and molecules less than 40 kDa in mass or 5 nm in diameter can pass relatively freely. NPC permeability is altered in several yeast nup mutants, pinpointing NPC structural elements, including core scaffold components, that are critical to the assembly or maintenance of this barrier (Shulga et al. 2000; Denning et al. 2001; Shulga and Goldfarb 2003; Strawn et al. 2004; Patel et al. 2007). Larger macromolecules overcome this permeability barrier by interacting either directly with the NPC themselves or through soluble transport factors. These macromolecules account for a tremendous variety of cargo including proteins, tRNAs, ribosomal subunits, and viral particles (reviewed in Macara 2001). Overall, the NPC is capable of transporting cargo up to 39 nm in diameter. This is on par with the size of the ribosomal subunits

and viral capsids that are known to move as intact complexes (Pante and Kann 2002). Macromolecules larger than this can still be translocated across the NPC, including mRNPs (mRNAs coated with RNA-binding proteins) with masses reaching several hundred thousand daltons. EM images of Balbiani ring mRNP particles associated with the NPC show the posttranscriptional 50 nm mRNA–protein particles to rearrange into rodlike structures, decreasing their maximum diameter to 25 nm (Mehlin et al. 1992). Thus, cargoes above a limiting diameter must rearrange to pass through the selective barrier of the NPC (Daneholt 2001). A transport signal and a shuttling receptor for that transport signal are the minimal requirements for any facilitated translocation (reviewed in Mattaj and Englmeier 1998; Pemberton and Paschal 2005). The targeting of proteins into or out of the nucleus requires specific amino acid sequence spans, termed nuclear localization sequences (NLSs) or nuclear export sequences (NESs). All the information required to target a protein to the nucleus is within these short sequences. In fact, fusion of an NLS to a nonnuclear protein is sufficient to mediate its transport and import to the nucleus (Goldfarb et al. 1986). For proteins, there are many distinct types of NLSs and NESs. For example, the classical NLS (cNLS) is the simple five amino acid peptide KKKRK, necessary and sufficient for targeting its attached protein to the nucleus (Goldfarb et al. 1986), whereas many proteins carry a more complex “bipartite” NLS consisting of two clusters of basic amino acids, separated by a spacer of roughly 10 amino acids (Dingwall et al. 1988). However, the full spectrum of sequences recognized by each transport receptor has not yet been carefully and fully defined. The most in depth analysis of NLS structural recognition by a transport receptor and extrapolation to predicting cargoes on a broader genome level has only been reported for one transport receptor (Lee et al. 2006). The key parameters defining an NLS or NES include critical tests for necessity and sufficiency in the endogenous protein. Importantly, some proteins undergo dynamic cycles of nuclear import and export and harbor both NLSs and

S.R. Wente and M.P. Rout

NESs. This can increase the complexity of identifying the signals. Moreover, the recognition and accessibility of the signals can be controlled by signaling, cell cycle, and developmental events (reviewed in Weis 2003; Terry et al. 2007). During NPC translocation, soluble transport factors are required to either bring cargo to the NPC or modulate cargo translocation across the NPC. Most of these soluble transport factors come from the family of proteins known

as the karyopherins (Table 2). The karyopherins (also called importins, exportins, and tranportins) were the first family of shuttling transport factors discovered. Fourteen karyopherin family members are found in Saccharomyces cerevisiae whereas at least 20 have been found in metazoans (reviewed in Fried and Kutay 2003; Pemberton and Paschal 2005). Most karyopherins bind their cargoes directly. However, in some cases an adaptor protein is needed in addition to the

Table 2. Karyopherin transport factors of yeast and vertebrates S. cerevisiae Karyopherins

Vertebrate Karyopherins

Kap95

Importin-b1

Kap104

Transportin or Transportin 2

Kap108/Sxm1

Importin 8

Kap109/Cse1 Kap111/Mtr10

CAS Transportin SR1 or SR2

Kap114

Importin 9

Kap119/Nmd5

Importin 7

Kap120 Kap121/Pse1

HsRanBP11 Importin 5/Importin b3/RanBP5

Kap122/Pdr6 Kap123

Importin 4

Kap127/Los1 Kap142/Msn5

Exportin-t Exportin 5

Crm1/Xpo1

Importin 13 CRM1/Exportin 1

— — —

Exportin 4 Exportin 6 Exportin 7/RanBP16

Examples of Cargo(s): (v) – vertebrate, (sc) – S. cerevisiae

Imports via sc-Kap60/v-importin-a adaptor proteins with cNLS; Imports via v-Snurportin the UsnRNPs; with no adaptor, imports v-cargo SREBP-2, HIV Rev, HIV TAT, cyclin B Imports sc-cargo – Nab2, Hrp1; v-cargo – PY-NLS proteins, mRNA-binding proteins, histones, ribosomal proteins Imports sc-cargo – Lhp1, ribosomal proteins; v-cargo – SRP19, Smad Imports sc-cargo – Kap60/Srp1; v-cargo – importin as Imports sc-cargo – Npl3, tRNAs; v-cargo – SR proteins, HuR Imports sc-cargo – TBP, histones, Nap1, Sua7; v-cargo – histones, ribosomal proteins Imports sc-cargo – Hog1, Crz1, Dst1, ribosomal proteins, histones; v-cargo – Smad, ERK, GR, ribosomal proteins Imports sc-cargo – Rpf1 Imports sc-cargo – Yra1, Spo12, Ste12, Yap1, Pho4, histones, ribosomal proteins; v-cargo – histones, ribosomal proteins Imports sc-cargo – Toa1 and Toa2, TFIIA Imports sc-cargo – SRP proteins, histones, ribosomal proteins; v-cargo - Transition Protein 2, histones, ribosomal protein S3a Exports tRNAs sc-cargo – imports replication protein A; exports Pho4, Crz1, Cdh1; v-cargo - exports pre-miRNA v-cargo – imports UBC9, Y14; exports eIF1A Exports proteins with leucine-rich NES, 60S ribosomal subunits (via NMD3 adaptor), 40S ribosomal subunits v-cargo – imports SOX2, SRY; exports Smad3, eIF5A Exports profilin, actin Exports p50-RhoGAP

Based on references cited within and adapted from Tran et al. 2007a and DeGrasse et al. 2009.

Nuclear Pore Complexes and Nuclear Transport

karyopherin to recognize signals. Not only do karyopherins have a cargo-binding domain, they also have an NPC-binding domain(s) as well as a binding domain at the amino-terminus for the small Ras-like GTPase Ran (see the following paragraph) (reviewed in Macara 2001; Harel and Forbes 2004). Overall, karyopherin family members share only modest sequence homology, with the greatest similarity being within their Ran-binding domains (Gorlichet al. 1997). However, a hallmark architecture within the karyopherins, as determined by recent highresolution structural studies, is the tandem HEAT-repeat fold formed by antiparallel helices connected by a short turn (reviewed in Conti and Izaurralde 2001). The HEAT-repeats arrange to form a superhelical structure, similar to a snail’s shell. This folding is reminiscent of the helix-turn-helix repeats found in the NPC’s core scaffold proteins. This similarity raises the intriguing possibility that karyopherins diverged from a common structure involved in both stationary and soluble phases of transport.

Import cycle

The association and dissociation of a karyopherin-cargo complex is regulated by direct binding of the small GTPase Ran (Fig. 2) (reviewed in Fried and Kutay 2003; Madrid and Weis 2006; Cook et al. 2007). In vitro binding studies show that import complexes are dissociated by RanGTP binding. Conversely, export complexes are formed via RanGTP association (Rexach and Blobel 1995; Floer and Blobel 1996; Chi and Adam 1997; Floer et al. 1997; Kutay et al. 1997a; Kutay et al. 1997b; Nakielny et al. 1999). Based on the localizations of the Ran GTPase activating protein (RanGAP) in the cytoplasm and the Ran guanine nucleotide exchange factor (RanGEF) in the nucleoplasm, cytoplasmic Ran is primarily in the GDP-bound state whereas nucleoplasmic Ran is kept primarily in the GTP-bound state (Fig. 2). The gradient formed from these localizations has been elegantly demonstrated by imaging fluorescence resonance energy transfer-based biosensors (Kalab et al. 2002). The RanGTP gradient across the two faces of the NPC is essential

Export cycle

NLS cargo

T

Import karyopherin

T

D

D

RanGDP

RanGDP

RanGTP

RanGTP T T

T T

T

T

Export karyopherin NES cargo

Figure 2. The nuclear transport cycle for karyopherins and their cargos. See main text for details.

S.R. Wente and M.P. Rout

for establishing the directionality of karyopherin-mediated transport. The pathway for karyopherin-mediated translocation is well described (reviewed in Weis 2003; Terry et al. 2007) (Fig. 2). For import, a specific karyopherin recognizes its cognate cargo in the cytoplasm where RanGTP levels are low. The karyopherin mediates the binding of the import complex to the NPC and facilitates translocation through the NPC. Once the complex moves through the NPC, release and dissociation of the karyopherin-cargo complex are stimulated by RanGTP in the nucleus. The karyopherin bound to RanGTP is then recycled back to the cytoplasm. Finally, GTP hydrolysis of Ran on the cytoplasmic side frees the karyopherin to interact with a second cargo molecule for further cycles of transport. Overall, Ran decreases the affinity of the karyopherin for its cargo (reviewed in Macara 2001; Cook et al. 2007). For export complexes, an analogous process occurs, but in this case, RanGTP binding increases the affinity of the karyopherin b for the export cargo. For example, for the exporting karyopherin Crm1 and an export cargo SPN1 (snurportin 1 adaptor for UsnRNPs), the Crm1 affinities for RanGTP and SPN1 in the ternary RanGTP-Crm1-SPN1 complex are increased 1000-fold (Paraskeva et al. 1999; Monecke et al. 2009). Actual movement through the NPC does not require energy input. The Ran affinity switches provide the energy for efficient cargo delivery and release. The only possible exception to this rule involves the import of large cargoes, where the presence of Ran and hydrolyzable GTP may be required for the import of cargoes .500 kDa in vitro (Lyman et al. 2002). It was also originally thought that an individual karyopherin was adapted for either import or export, but not both. However, there are now documented examples of karyopherins functioning in both import and export, although with different cargoes in each direction (Yoshida and Blobel 2001). In addition to protein import and export, karyopherins can also transport RNAs. For example, the karyopherins Crm1 and exportin-t mediate the export of uridine-rich small nuclear RNAs (U snRNAs) and tRNAs, respectively

(Simos et al. 2002; Rodriguez 2004). Transport is accomplished via direct binding of karyopherins to RNA or to signal sequences within the protein components of the RNP complexes. For example, Crm1 does not bind UsnRNAs directly and requires the adaptor PHAX that binds the cap-complex on the RNA (Ohno et al. 2000). However, exportin-t directly interacts with tRNAs (Arts et al. 1998; Hellmuth et al. 1998; Kutay et al. 1998; Lipowsky et al. 1999). Karyopherins are also involved in the export of some viral RNAs, including their mRNAs (Carmody and Wente 2009). However, the primary mRNP export transport receptor is a nonkaryopherin designated Mex67 in yeast and NXF1 in metazoans, which heterodimerizes with a protein termed respectively Mtr2 or p15/Nxt1 (Erkmann and Kutay 2004). Even though Mex67 is unrelated in sequence and structure to the karyopherin family, it has all the requirements of a transport receptor: cargo binding, nucleocytoplasmic shuttling, and NPC-binding. The stoichiometry of the Mex67-Mtr2 heterodimer per transported mRNP is unknown; Mex67-Mtr2 either is recruited directly to the mRNA or interacts cotranscriptionally with proteins of the mRNP assembly (Erkmann and Kutay 2004; Carmody and Wente 2009). Like karyopherins, Mex67Mtr2 heterodimers bind directly to FG Nups, although it seems they prefer different subsets of FG Nups to their karyopherin counterparts, which might reflect how the karyopherin mediated transport pathways and mRNPexport pathways are kept apart at the NPC (Strawn et al. 2001; Terry et al. 2007; Terry and Wente 2007). As mRNP is a major source of traffic across the NPC, it is interesting that most of this transit is facilitated by non-karyopherin carriers. Because Ran is not utilized to establish a gradient, directionality in the mRNA export pathway is conferred by proteins that modify the mRNPs as they cross the NPC. Chief among these is the protein Dbp5. Dbp5 is a member of the SF2 helicase superfamily of RNAdependent ATPases (Snay-Hodge et al. 1998; Tseng et al. 1998) and carries a DEAD/H-box sequence motif. Such DEAD-box proteins are

Nuclear Pore Complexes and Nuclear Transport

involved in the alteration of RNA structure, such as the facilitation of RNA duplex formation and unwinding or aiding the association and dissociation of RNA-binding proteins (Jankowsky et al. 2001; Fairman et al. 2004; Yang and Jankowsky 2006). Much like the GDP/GTP triggered switches for Ran, DEADbox proteins potentially use ADP/ATP for nucleotide-dependent conformational switches (Henn et al. 2002; Tran et al. 2007b; Henn et al. 2008; Fan et al. 2009). Dbp5 ATPase activity is activated by Gle1, a protein that exchanges between the nucleus and cytoplasm with a docking site on the cytoplasmic filaments of the NPC (Cole and Scarcelli 2006). Gle1 activation of Dbp5 in turn requires the soluble molecule inositol hexakisphosphate (IP6) to increase the binding affinity of Dbp5 to RNA (Alcazar-Roman et al. 2006; Weirich et al. 2006). This raises the possibility that IP6 is acting as a signaling molecule to somehow regulate the mRNA export machinery (York et al. 1999). The Dbp5 on the cytoplasmic side of the NPC triggers the release of mRNP proteins such as Mex67 and Nab2, effectively remodeling the mRNP protein composition as they exit the NPC as well as imparting directionality to mRNP transport (Tran et al. 2007b). There also seems to be a checkpoint at the NPC basket, to ensure that mRNAs have been correctly spliced and packaged into mRNPs immediately before export. SUMO is a ubiquitin-like molecule that can be attached to proteins as a signal or modulator of their function. The SUMO protease, Ulp1, is retained at the NPC via its interaction with the nuclear basket component Mlp/Tpr, and it has been suggested that sumoylation of mRNPs acts to signal that they are correctly assembled. Conversely, the desumoylation of incorrectly spliced mRNPs at the nuclear basket prevents their export and instead targets them for degradation (Lewis et al. 2007; Xu et al. 2008). Other nonkaryopherin carriers for NPC translocation exist with the same capacity to dock at various NPC sites via the FG Nups. Ntf2/p10 is a carrier for RanGDP importing Ran to the nucleus. Ntf2/p10 also uses the localization of RanGEF in the nucleus to create

a differential affinity for RanGDP versus RanGTP (reviewed in Pemberton and Paschal 2005; Cook et al. 2007). There are also some proteins with karyopherin-like HEAT-repeats that require no receptor and translocate independently of a carrier. The driving force for their accumulation in the nucleus is their affinity for binding targets in the nucleoplasm (Fagotto et al. 1998). Perhaps the most abundant nuclear transport cargos in many cells are the 60S and 40S ribosomal subunits, on the way from synthesis in the nucleolus to form mature ribosomes in the cytoplasm. Export of ribosomal subunits is still poorly understood, and it seems that unlike the case of a single karyopherin bound to each protein cargo molecule as described earlier, several transport receptors must chaperone each ribosomal subunit across the NPC. The earliest identified ribosomal exporter was Crm1 (Fornerod et al. 1997; Fukuda et al. 1997; Stade et al. 1997). For export of the 60S subunit, the NES is provided by the export adaptor Nmd3 (Johnson et al. 2002). Nmd3 binds reversibly to the 60S subunit while the latter is being assembled in the nucleus. After export into the cytoplasm, a GTPase called Lsg1 appears to mediate the release of Nmd3 from the 60S ribosomal subunit (Zemp and Kutay 2007). In yeast the Mex67-Mtr2 heterodimer has also been suggested to serve as an export receptor for the 60S subunit (Yao et al. 2007). Although more karyopherin-like proteins have recently been identified that play a role in ribosomal export (Oeffinger et al. 2004), other nonkaryopherin receptors like Arx1 have been found, though Arx1 still interacts with the same NPC docking sites as karyopherins (Bradatsch et al. 2007). Why might so many transport factors be required to ferry the ribosomal subunits out of the nucleus? One reason might be explained by the analogy of maneuvering ships out of the harbor. One tugboat is all that is required to move a small barge quickly and efficiently, but on its own does not have the power to move an ocean liner, which instead needs many tugboats together to take it safely out of harbor. Similarly, as the efficiency of passage through the NPC decreases with cargo size

S.R. Wente and M.P. Rout

(see the preceding section) but increases with binding strength to FG Nups (see the following section), many transport factors are needed to carry large cargos across the NPC. Regardless, the fundamental mechanism for transport in all these cases of different carriers is still dependent on the same features of the NPC machine. OPERATION OF THE NPC: THE “FG” TRANSLOCATION MACHINERY

Although the cargo binding mechanisms of the transport factors discussed earlier vary, almost all binding sites with the NPC are found in the same class of Nups. These Nups are collectively termed FG Nups because they contain “FG repeat regions.” The NPC core framework provides the correct positioning of these FG Nups so that they flank and fill the central tube. Approximately a third of all Nups contain FG repeat regions, which consist of multiple small hydrophobic clusters containing an FG (Phe-Gly) dipeptide (usually FG, FXFG, or GLFG) separated by 20 – 70 residue hydrophilic linkers (reviewed in Rout and Wente 1994; Allen et al. 2001; Tran and Wente 2006). Binding of these repeats is cooperative, with two to four repeats associating with each transport factor molecule (Bayliss et al. 2002; Grant et al. 2003; Isgro and Schulten 2005; Liu and Stewart 2005). With 5– 50 repeats per FG Nup and 200 FG Nups per NPC, there is a potential for .1000 transport factor binding sites per NPC. Imaging of the trajectories of a single translocating molecule through the NPC is consistent with movement between multiple binding sites ( presumably the many FG repeats) within the NPC (Yang et al. 2004; Kubitscheck et al. 2005; Yang and Musser 2006). FG repeat regions are natively disordered and having no secondary structure they instead form filaments that can diffusively writhe around their attachment sites at the NPC. Although some will be diffuse random coils, others may have enough internal cohesion to form more compact “molten globules” (Denning et al. 2003; Lim et al. 2006; Patel et al. 2007; Krishnan et al. 2008). By forming a dense meshwork in and around the central tube, the FG repeats appear to set

up an entropic potential that excludes macromolecules from their vicinity, while permitting the approach of small molecules such as water and nucleotides (Lim et al. 2006). However, karyopherins overcome this barrier as they bind FG Nups and, passing from FG Nup to FG Nup, diffuse readily through the NPC. Indeed, this model is supported by the fact that a simple device with similar properties—a nanotube coated with FG Nups—replicated many of the features of nuclear transport (Jovanovic-Talisman et al. 2009). In terms of the Gibb’s free energy required to cross the NPC, the enthalpy for binding the FG Nups cancels out the entropic barrier of the central tube region, flattening the energy landscape and thus lowering the activation energy of translocation across the NE. This is analogous to how an enzyme or catalyst lowers the activation energy of the transition state permitting a substrate to convert to product. This concept has been termed “virtual gating” (Rout et al. 2000; Rout et al. 2003). Nevertheless, despite much work over the last few years, how exactly this gating works at the molecular level remains unclear and several models have attempted to explain it. One possibility is that the natively unfolded FG Nups could act as a “polymer brush” where rapid diffusion-driven movements of their unfolded domains would sweep away macromolecules from their vicinity. Although a central tube filled with such brushes would exclude nonbinding macromolecules, transport factors would overcome this exclusion by binding to the FG repeats and so pass through the NPC (Rout et al. 2000; Rout et al. 2003; Lim et al. 2006). In another model, regions of FG repeats may collapse upon binding transport factors, as indicated by atomic force microscopy data (Lim et al. 2007a; Lim et al. 2007b). As they pass through the central tube, transport factors would open up their own passageway through the meshwork of FG repeats. A different view is provided by the “saturated” model (Frey et al. 2006; Frey and Gorlich 2007), originally the “hydrophobic gel” model (Ribbeck and Gorlich 2002). Here it is suggested that the phenylalanines in the FG repeat regions are crosslinked with each other through their Phe residues to

Nuclear Pore Complexes and Nuclear Transport

form a dense gel of FG repeat filaments. Transport factors pass through this gel by binding the FG repeats, and in doing so they dissolve the crosslinks. Other macromolecules are therefore excluded because they cannot do this. A more recent “reduction in dimensionality”-model posits that the FG repeat regions form a layer coating the inner walls of the central tube (Peters 2005). Although transport factors enter this layer through binding giving them full access to the tube’s volume, nonbinding molecules can only pass through the narrow FG Nupfree middle. Recently, work has emerged that suggests while crossing the NPC, transport factors not only carry cargoes but also enhance the selectivity of the NPC by competing away nonspecific macromolecules, indicating that the transport factors themselves are also important components of the selectivity barrier (Timney et al. 2006; Zilman et al. 2007; Jovanovic-Talisman et al. 2009). Not all of these ideas are mutually exclusive, and “hybrid” models have also been proposed (Strawn et al. 2004; Patel et al. 2007; Krishnan et al. 2008). FG Nup repeat regions seem to fall into several classes on the basis of their charge density and FG repeat motif type. Moreover, these classes seem to have somewhat different physical behaviors (Krishnan et al. 2008). Each class of FG Nup appears to be preferred by only certain kinds of transport factor. Thus, these different FG Nup classes might serve to mediate multiple, simultaneous, functionally independent transport pathways through the NPC (reviewed in Tran and Wente 2006). Supporting evidence for this has come from in vivo studies using target combinations of different FG region deletions. Certain deletion combinations can affect passage of one transport factor through the NPC without affecting another (Strawn et al. 2004; Terry and Wente 2007). FG Nups organization around the NPC can be categorized as either symmetric or asymmetric. “Symmetric” FG Nups are found on both sides of the NPC closely packed around and within the central tube. “Asymmetric” FG Nups are found primarily on either the nuclear or cytoplasmic side only, and significantly further from the tube (Rout et al. 2000; Alber et al.

2007b). Deletion mutagenesis experiments in yeast have shown that the FG regions of asymmetric Nups are not essential for transport (Strawn et al. 2004; Zeitler and Weis 2004). Instead, they may guide the directionality of transport-factor/cargo complexes by providing a high-affinity binding platform at the far end of a transport factor’s route through the NPC. A transport factor would be drawn to this site and prevented by it from returning through the NPC until Ran terminates the transport reaction, in this way increasing the efficiency of transport (Rout et al. 2003; Strawn et al. 2004; Zilman et al. 2007). REGULATORS OF TRANSLOCATION: THE NON-FG DETERMINANTS

Unidirectional transport through the NPC can be viewed as being driven by the formation of gradients, in which cargos move from a “source” to a “sink.” Karyopherin-cargo complexes form on one side of the NPC and diffuse down their concentration gradient to the other side, where their concentration is low because they are disassembled there. This “source and sink” process also drives other forms of nucleocytoplasmic transport, such as the attachment of RNA binding proteins on one side of the NPC and their removal on the other side during RNP export (reviewed in Rout et al. 2003; Stewart 2007). The NPC on its own does not enforce directionality (Nachury and Weis 1999). Single molecule studies show the transport receptor-cargo complexes seem to move randomly within the NPC channel until they exit either face of the NPC (Yang et al. 2004). Thus, in a physiological context, other NPC events and non-FG Nup binding sites are required to provide transport efficiency, regulation, and directionality (Fig. 3). As detailed earlier, there are specific factors for mediating transport directionality and cargo release: The RanGTPase for karyopherins and the Gle1-IP6 activated Dbp5 for mRNA export. These use non-FG NPC docking sites that increase their local concentration at their respective sites of action. Dbp5 and Gle1 bind to distinct but structurally juxtaposed Nups on the cytoplasmic filaments. In budding yeast,

S.R. Wente and M.P. Rout Transport factors & cargos

Transport modulators

Nuclear organization

NLS cargo Import karyopherin

SUMOylation/ RanGAP complex

Membrane cargo

Gle1/1P6/Dbp5 complex Ran binding sites

Active gene tethered at NPC

SUMO peptidase

Exporting RNP

Ran Export karyopherin

RNP proofreading complex

NES cargo Mex67 mRNP export complex

Ran binding sites

Microtubule attachment NPC assembly factors Lamina and Mlp/Tpr tethering of NPC Mad proteins

Mobile nup

Figure 3. The main NPC interactors: left, transport factors and their cargos, center, modulators of RNP export,

karyopherins or Ran, right, cytoskeletal, nucleoskeletal, and chromosomal elements.

Dbp5 interacts with Nup159 (Hodge et al. 1999; Schmitt et al. 1999) and Gle1 with Nup42 (Murphy and Wente 1996; Strahm et al. 1999), with both sets of binding partners fully conserved in metazoans. The NPC can also help to regulate the GTP-bound state of Ran in its vicinity. Thus, mammalian cells have binding sites for sumoylated-RanGAP on the cytoplasmic filament Nup358, likely to help enhance the dissociation of exporting karyopherin complexes as they exit the NPC (Mahajan et al. 1997; Matunis et al. 1998). In organisms like plants that lack Nup358, unique NE associated proteins facilitate the localized anchoring of RanGAP (Xu et al. 2007; Zhao et al. 2008), so it seems that regulating the GTP-bound state in the immediate vicinity of the NPC is important for transport efficiency in many eukaryotes. Although the FG domains do not form a gradient of docking sites in the translocation channel, there are non-FG high affinity docking sites located at terminal transport steps that influence efficiency. In particular, two Nups on the opposing cytoplasmic and nuclear faces provide a virtual smorgasbord of binding sites for different factors that regulate cell physiology: Nup358 and Nup153, respectively (Fig. 3). In metazoan cells, the cytoplasmic facing Nup358 provides a high affinity binding site for the mRNA export factor NXF1 on the cytoplasmic face of the NPC (Forler et al. 2004). Similarly, Nup153 provides a high affinity binding site for an importing karyopherin on the nuclear face of the NPC (Moroianu et al. 1997).

Although both have FG domains it is the nonFG docking sites that are functionally distinguishing. Nup358 was originally identified as a RanGTP binding protein (Wu et al. 1995; Yokoyama et al. 1995) and binds sumoylated RanGAP. However, it also has a domain with E3 SUMO1 ligase activity and docks the sumolylation cofactor enzyme Ubc9 to facilitate the addition of SUMO to targets directly at the cytoplasmic filaments (Pichler et al. 2002; Reverter and Lima 2005). Nup358 also harbors a cyclophilin A homologous domain that associates with subunits of the 26S proteasome and might modulate ubiquitin-proteasome function (Wu et al. 1995; Yi et al. 2007). The amino-terminal domain of Nup358 allows interaction with microtubules during interphase as well as with kinetochores in mitosis (Salina et al. 2003; Joseph et al. 2004; Joseph and Dasso 2008). On the nuclear side of the NPC, Nup153 has a Zn finger Ran binding domain (Nakielny et al. 1999; Higa et al. 2007) attracting Ran to the vicinity of the NPC to regulate karyopherinbased transport events at its nuclear face. In sharp contrast to the Nup358-SUMO1 ligase activity, Nup153 actually binds a SUMO1/ sentrin/SMT3 specific peptidase 2, perhaps to help desumolyate misassembled mRNPs (Hang and Dasso 2002; Zhang et al. 2002). Nup153 also possesses a novel single-stranded RNA binding domain (Ball et al. 2004) that is perhaps concomitant with a function in RNP export. Nup153 also plays a role in recruiting the

Nuclear Pore Complexes and Nuclear Transport

COPI coatomer complex, possibly to mediate NPC mitotic disassembly (Liu et al. 2003). Taken together, Nup358 and Nup153 are but two examples of an emerging paradigm for multifunctional nucleoporins. They demonstrate that the NPC serves as a scaffold for a diversity of processes that are distinct from those strictly associated with simply moving a cargo from one side of the NPC to the other. Transport can also be modulated at the level of the transport factors and their cargos. A common method is to regulate the binding strength or accessibility of an NLS or NES in a cargo for its cognate karyopherin. This often involves phosphorylation in or near the signal sequence of the cargo. Another method involves modulating the level of transport machinery components, usually particular subtypes of karyopherin. Using this method, transport of particular proteins recognized by that karyopherin is prevented until the karyopherin itself is expressed (reviewed in Poon and Jans 2005). THE NPC AS A NUCLEAR COMPONENT

Although the most dynamic process associated with the NPC is nucleocytoplasmic transport, the NPC should not be regarded as a static structure. New NPCs are assembled de novo during interphase (D’Angelo et al. 2006) and, in higher eukaryotes at least, the NPC is reversibly disassembled during mitosis (reviewed in Hetzer et al. 2005; Fernandez-Martinez and Rout 2009). There is emerging evidence for multiple nonnucleoporin accessory factors that play roles in NPC biogenesis and maintenance (Fig. 3). This includes proteins that regulate membrane composition and membrane dynamics such as Acc1 and Apq12, respectively (Schneiter et al. 1996; Scarcelli et al. 2007). Changes in the lipid composition or membrane fluidity can impact NPC structure and localization. Chromatin associated proteins, such as ELYS, are required specifically for postmitotic assembly of NPCs (Franz et al. 2007). Recent studies have found several discrete roles for the reticulon (Rtn) family of membrane-associated proteins in NE disassembly and reassembly during open mitosis. The Rtns

are critical for regulating transitions between the endoplasmic reticulum tubular network and the NE sheet (Anderson and Hetzer 2008). However, during interphase, an Rtn1 pool at the NE facilitates de novo NPC assembly (Dawson et al. 2009). This most likely is to provide a transient stabilizer of the highly convex region of the pore membrane during early steps in its formation. It is likely that additional non-NPC accessory factors will continue to be revealed, and this is an active area of research that has been well covered in several recent reviews (Antonin et al. 2008; D’Angelo and Hetzer 2008; Kutay and Hetzer 2008; Lim et al. 2008). Even after its assembly, many of the interactions holding the bulk of the NPC together are actually surprisingly transient. Upon completion of NPC construction, some Nups remain stably associated, others turn over promptly, and some shuttle rapidly between the nucleoplasm and cytoplasm, spending little time on the NPC itself (Rabut et al. 2004; Dilworth et al. 2001; Dilworth et al. 2005). For example, there are multiple roles for Nups in kinetochore function and mitosis, as well as in the maintenance of the boundaries between silent and active chromatin (Roux and Burke 2006; Kalverda et al. 2008; Lim et al. 2008). Despite these processes requiring Nups, none appear to occur at the NPC itself. Indeed, recent work underscores how a great number of Nup-chromatin interactions occur within the nuclear interior and away from the NPC itself; these interactions have also been shown to be crucial in controlling gene expression during cell differentiation in higher eukaryotes (Capelson et al. 2010; Kalverda et al. 2010). Conversely, the NPC can act as a reservoir for cell cycle proteins when they are inactive. One example, the Mad proteins, are docked to the NPC in interphase but are released when they function in mitosis (Iouket al. 2002; Scott et al. 2005; Lee et al. 2008). The NPC also plays an important role in the organization of internal nuclear architecture (Fig. 3) and is used as an attachment site for such nuclear peripheral proteins as lamins and yMlp/vTpr. An exciting recent discovery is that certain inducible genes carry specific sequences

S.R. Wente and M.P. Rout

that can cause them to associate with the NPC’s vicinity upon transcriptional activation; once there, they retain a “memory” of their active state for some time after they cease transcription, leaving the genes poised for rapid reactivation (Ahmed et al. 2010). This tethering of induced genes to the NPC might also be also aided by factors required for posttranscriptional mRNP processing and export (Dieppois et al. 2006; Chekanova et al. 2008; Rougemaille et al. 2008). CONCLUSIONS

Our structural and functional understanding of the nuclear transport machinery has blossomed in the past decade, revealing new and unexpected aspects about the NPC and its interacting transport factors. Perhaps most surprising has been the discovery of the evolutionary connections between the NPC and the vesicular trafficking apparatus, the unusual nature of the FG Nup-mediated selective barrier, and the increasing range of connectivity between the NPC and global aspects of cellular physiology and gene expression. Yet as is so often the case, the solving of one question has raised many others. Many of the unanswered questions regarding the mechanisms and regulation of the nuclear transport machinery present exciting challenges for the years ahead. ACKNOWLEDGMENTS

We acknowledge that many of the primary references for critical studies are included in reviews cited, and appreciate our colleagues’ consideration in this limitation. We thank Renee Dawson and Kelli Mullin for assistance with manuscript preparation. This work was supported by funding from the National Institutes of Health (R01 GM51219 to S. R. W. and R01 GM062427 and R01 GM071329 to M. P. R.). REFERENCES Ahmed S, Brickner JH. 2007. Regulation and epigenetic control of transcription at the nuclear periphery. Trends Genet 23: 396 –402. Ahmed S, Brickner DG, Light WH, Cajigas I, McDonough M, Froyshteter AB, Volpe T, Brickner JH. 2010. DNA zip codes control an ancient mechanism for gene

targeting to the nuclear periphery. Nat Cell Biol 12: 111 –118. Aitchison JD, Rout MP, Marelli M, Blobel G, Wozniak RW. 1995. Two novel related yeast nucleoporins Nup170p and Nup157p: Complementation with the vertebrate homologue Nup155p and functional interactions with the yeast nuclear pore-membrane protein Pom152p. J Cell Biol 131: 1133–1148. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007a. Determining the architectures of macromolecular assemblies. Nature 450: 683 –694. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007b. The molecular architecture of the nuclear pore complex. Nature 450: 695– 701. Alcazar-Roman AR, Tran EJ, Guo S, Wente SR. 2006. Inositol hexakisphosphate and Gle1 activate the DEAD-box protein Dbp5 for nuclear mRNA export. Nat Cell Biol 8: 711– 716. Allen NP, Huang L, Burlingame A, Rexach M. 2001. Proteomic analysis of nucleoporin interacting proteins. J Biol Chem 276: 29268– 29274. Anderson DJ, Hetzer MW. 2008. Reshaping of the endoplasmic reticulum limits the rate for nuclear envelope formation. J Cell Biol 182: 911– 924. Antonin W, Mattaj IW. 2005. Nuclear pore complexes: Round the bend? Nat Cell Biol 7: 10– 12. Antonin W, Ellenberg J, Dultz E. 2008. Nuclear pore complex assembly through the cell cycle: Regulation and membrane organization. FEBS Lett 582: 2004– 2016. Arts GJ, Kuersten S, Romby P, Ehresmann B, Mattaj IW. 1998. The role of exportin-t in selective nuclear export of mature tRNAs. EMBO J 17: 7430– 7441. Ball JR, Dimaano C, Ullman KS. 2004. The RNA binding domain within the nucleoporin Nup153 associates preferentially with single-stranded RNA. RNA 10: 19– 27. Bayliss R, Leung SW, Baker RP, Quimby BB, Corbett AH, Stewart M. 2002. Structural basis for the interaction between NTF2 and nucleoporin FxFG repeats. EMBO J 21: 2843–2853. Belgareh N, Rabut G, Bai SW, van Overbeek M, Beaudouin J, Daigle N, Zatsepina OV, Pasteau F, Labas V, FromontRacine M, et al. 2001. An evolutionarily conserved NPC subcomplex, which redistributes in part to kinetochores in mammalian cells. J Cell Biol 154: 1147–1160. Berke IC, Boehmer T, Blobel G, Schwartz TU. 2004. Structural and functional analysis of Nup133 domains reveals modular building blocks of the nuclear pore complex. J Cell Biol 167: 591– 597. Bradatsch B, Katahira J, Kowalinski E, Bange G, Yao W, Sekimoto T, Baumgartel V, Boese G, Bassler J, Wild K, et al. 2007. Arx1 Functions as an Unorthodox Nuclear Export Receptor for the 60S Preribosomal Subunit. Mol Cell 27: 767–779. Brohawn SG, Leksa NC, Spear ED, Rajashankar KR, Schwartz TU. 2008. Structural evidence for common ancestry of the nuclear pore complex and vesicle coats. Science 322: 1369– 1373.

Nuclear Pore Complexes and Nuclear Transport Brohawn SG, Partridge JR, Whittle JR, Schwartz TU. 2009. The nuclear pore complex has entered the atomic age. Structure 17: 1156– 1168. Capelson M, Liang Y, Schulte R, Mair W, Wagner U, Hetzer MW. 2010. Chromatin-bound nuclear pore components regulate gene expression in higher eukaryotes. Cell 140: 372–383. Carmody SR, Wente SR. 2009. mRNA nuclear export at a glance. J Cell Sci 122: 1933– 1937. Chekanova JA, Abruzzi KC, Rosbash M, Belostotsky DA. 2008. Sus1, Sac3, and Thp1 mediate post-transcriptional tethering of active genes to the nuclear rim as well as to non-nascent mRNP. RNA 14: 66–77. Chi NC, Adam SA. 1997. Functional domains in nuclear import factor p97 for binding the nuclear localization sequence receptor and the nuclear pore. Mol Biol Cell 8: 945–956. Cole CN, Scarcelli JJ. 2006. Transport of messenger RNA from the nucleus to the cytoplasm. Curr Opin Cell Biol 18: 299 –306. Conti E, Izaurralde E. 2001. Nucleocytoplasmic transport enters the atomic age. Curr Opin Cell Biol 13: 310–319. Cook A, Bono F, Jinek M, Conti E. 2007. Structural biology of nucleocytoplasmic transport. Annu Rev Biochem 76: 647–671. Cronshaw JM, Krutchinsky AN, Zhang W, Chait BT, Matunis MJ. 2002. Proteomic analysis of the mammalian nuclear pore complex. J Cell Biol 158: 915– 927. D’Angelo MA, Hetzer MW. 2008. Structure, dynamics and function of nuclear pore complexes. Trends Cell Biol 18: 456–466. D’Angelo MA, Anderson DJ, Richard E, Hetzer MW. 2006. Nuclear pores form de novo from both sides of the nuclear envelope. Science 312: 440–443. Daneholt B. 2001. Assembly and transport of a premessenger RNP particle. Proc Natl Acad Sci 98: 7012– 7017. Dawson TR, Lazarus MD, Hetzer MW, Wente SR. 2009. ER membrane-bending proteins are necessary for de novo nuclear pore formation. J Cell Biol 184: 659–675. Debler EW, Ma Y, Seo HS, Hsia KC, Noriega TR, Blobel G, Hoelz A. 2008. A fence-like coat for the nuclear pore membrane. Mol Cell 32: 815 –826. DeGrasse JA, DuBois KN, Devos D, Siegel TN, Sali A, Field MC, Rout MP, Chait BT. 2009. Evidence for a shared nuclear pore complex architecture that is conserved from the last common eukaryotic ancestor. Mol Cell Proteomics 8: 2119–2130. Denning D, Mykytka B, Allen NP, Huang L, Al B, Rexach M. 2001. The nucleoporin Nup60p functions as a Gsp1p-GTP-sensitive tether for Nup2p at the nuclear pore complex. J Cell Biol 154: 937–950. Denning DP, Patel SS, Uversky V, Fink AL, Rexach M. 2003. Disorder in the nuclear pore complex: The FG repeat regions of nucleoporins are natively unfolded. Proc Natl Acad Sci 100: 2450–2455. Devos D, Dokudovskaya S, Alber F, Williams R, Chait BT, Sali A, Rout MP. 2004. Components of coated vesicles and nuclear pore complexes share a common molecular architecture. PLoS Biol 2: e380. Devos D, Dokudovskaya S, Williams R, Alber F, Eswar N, Chait BT, Rout MP, Sali A. 2006. Simple fold composition

and modular architecture of the nuclear pore complex. Proc Natl Acad Sci 103: 2172– 2177. Dieppois G, Iglesias N, Stutz F. 2006. Cotranscriptional recruitment to the mRNA export receptor Mex67p contributes to nuclear pore anchoring of activated genes. Mol Cell Biol 26: 7858– 7870. Dilworth DJ, Suprapto A, Padovan JC, Chait BT, Wozniak RW, Rout MP, Aitchison JD. 2001. Nup2p dynamically associates with the distal regions of the yeast nuclear pore complex. J Cell Biol 153: 1465–1478. Dilworth DJ, Tackett AJ, Rogers RS, Yi EC, Christmas RH, Smith JJ, Siegel AF, Chait BT, Wozniak RW, Aitchison JD. 2005. The mobile nucleoporin Nup2p and chromatin-bound Prp20p function in endogenous NPC-mediated transcriptional control. J Cell Biol 171: 955 –965. Dingwall C, Robbins J, Dilworth SM, Roberts B, Richardson WD. 1988. The nucleoplasmin nuclear location sequence is larger and more complex than that of SV-40 large T antigen. J Cell Biol 107: 841 –849. Erkmann JA, Kutay U. 2004. Nuclear export of mRNA: From the site of transcription to the cytoplasm. Exp Cell Res 296: 12–20. Fagotto F, Gluck U, Gumbiner BM. 1998. Nuclear localization signal-independent and importin/karyopherinindependent nuclear import of b-catenin. Curr Biol 8: 181 –190. Fairman ME, Maroney PA, Wang W, Bowers HA, Gollnick P, Nilsen TW, Jankowsky E. 2004. Protein displacement by DExH/D "RNA helicases" without duplex unwinding. Science 304: 730–734. Fan JS, Cheng Z, Zhang J, Noble C, Zhou Z, Song H, Yang D. 2009. Solution and crystal structures of mRNA exporter Dbp5p and its interaction with nucleotides. J Mol Biol 388: 1– 10. Feldherr CM, Akin D. 1997. The location of the transport gate in the nuclear pore complex. J Cell Sci 110: 3065–3070. Fernandez-Martinez J, Rout MP. 2009. Nuclear pore complex biogenesis. Curr Opin Cell Biol 21: 603 –612. Field MC, Dacks JB. 2009. First and last ancestors: Reconstructing evolution of the endomembrane system with ESCRTs, vesicle coat proteins, and nuclear pore complexes. Curr Opin Cell Biol 21: 4 –13. Floer M, Blobel G. 1996. The nuclear transport factor karyopherin b binds stoichiometrically to Ran-GTP and inhibits the Ran GTPase activating protein. J Biol Chem 271: 5313–5316. Floer M, Blobel G, Rexach M. 1997. Disassembly of RanGTP-karyopherin b complex, an intermediate in nuclear protein import. J Biol Chem 272: 19538– 19546. Fontoura BM, Blobel G, Matunis MJ. 1999. A conserved biogenesis pathway for nucleoporins: Proteolytic processing of a 186-kilodalton precursor generates Nup98 and the novel nucleoporin, Nup96. J Cell Biol 144: 1097–1112. Forler D, Rabut G, Ciccarelli FD, Herold A, Kocher T, Niggeweg R, Bork P, Ellenberg J, Izaurralde E. 2004. RanBP2/Nup358 provides a major binding site for NXF1-p15 dimers at the nuclear pore complex and functions in nuclear mRNA export. Mol Cell Biol 24: 1155–1167.

S.R. Wente and M.P. Rout Fornerod M, Ohno M, Yoshida M, Mattaj IW. 1997. CRM1 is an export receptor for leucine-rich nuclear export signals. Cell 90: 1051–1060. Franz C, Walczak R, Yavuz S, Santarella R, Gentzel M, Askjaer P, Galy V, Hetzer M, Mattaj IW, Antonin W. 2007. MEL-28/ELYS is required for the recruitment of nucleoporins to chromatin and postmitotic nuclear pore complex assembly. EMBO Rep 8: 165 –172. Frey S, Gorlich D. 2007. A saturated FG-repeat hydrogel can reproduce the permeability properties of nuclear pore complexes. Cell 130: 512 –523. Frey S, Richter RP, Gorlich D. 2006. FG-rich repeats of nuclear pore proteins form a three-dimensional meshwork with hydrogel-like properties. Science 314: 815– 817. Fried H, Kutay U. 2003. Nucleocytoplasmic transport: Taking an inventory. Cell Mol Life Sci 60: 1659– 1688. Fukuda M, Asano S, Nakamura T, Adachi M, Yoshida M, Yanagida M, Nishida E. 1997. CRM1 is responsible for intracellular transport mediated by the nuclear export signal. Nature 390: 308 –311. Goldfarb DS, Gariepy J, Schoolnik G, Kornberg RD. 1986. Synthetic peptides as nuclear localization signals. Nature 322: 641– 644. Gorlich D, Dabrowski M, Bischoff FR, Kutay U, Bork P, Hartmann E, Prehn S, Izaurralde E. 1997. A novel class of RanGTP binding proteins. J Cell Biol 138: 65–80. Grandi P, Dang T, Pane N, Shevchenko A, Mann M, Forbes D, Hurt E. 1997. Nup93, a vertebrate homologue of yeast Nic96p, forms a complex with a novel 205-kDa protein and is required for correct nuclear pore assembly. Mol Biol Cell 8: 2017– 2038. Grant RP, Neuhaus D, Stewart M. 2003. Structural basis for the interaction between the Tap/NXF1 UBA domain and FG nucleoporins at 1A resolution. J Mol Biol 326: 849–858. Hang J, Dasso M. 2002. Association of the human SUMO-1 protease SENP2 with the nuclear pore. J Biol Chem 277: 19961–19966. Harel A, Forbes DJ. 2004. Importin b: Conducting a much larger cellular symphony. Mol Cell 16: 319 –330. Hellmuth K, Lau DM, Bischoff FR, Kunzler M, Hurt E, Simos G. 1998. Yeast Los1p has properties of an exportinlike nucleocytoplasmic transport factor for tRNA. Mol Cell Biol 18: 6374– 6386. Henn A, Cao W, Hackney DD, De La Cruz EM. 2008. The ATPase cycle mechanism of the DEAD-box rRNA helicase, DbpA. J Mol Biol 377: 193 –205. Henn A, Shi SP, Zarivach R, Ben-Zeev E, Sagi I. 2002. The RNA helicase DbpA exhibits a markedly different conformation in the ADP-bound state when compared with the ATP- or RNA-bound states. J Biol Chem 277: 46559–46565. Hetzer MW, Wente SR. 2009. Border control at the nucleus: Biogenesis and organization of the nuclear membrane and pore complexes. Dev Cell 17: 606– 616. Hetzer MW, Walther TC, Mattaj IW. 2005. Pushing the envelope: Structure, function, and dynamics of the nuclear periphery. Annu Rev Cell Dev Biol 21: 347– 380.

Higa MM, Alam SL, Sundquist WI, Ullman KS. 2007. Molecular characterization of the Ran-binding zinc finger domain of Nup153. J Biol Chem 282: 17090–17100. Hodge CA, Colot HV, Stafford P, Cole CN. 1999. Rat8p/ Dbp5p is a shuttling transport factor that interacts with Rat7p/Nup159p and Gle1p and suppresses the mRNA export defect of xpo1–1 cells. EMBO J 18: 5778– 5788. Hsia KC, Stavropoulos P, Blobel G, Hoelz A. 2007. Architecture of a coat for the nuclear pore membrane. Cell 131: 1313–1326. Iouk T, Kerscher O, Scott RJ, Basrai MA, Wozniak RW. 2002. The yeast nuclear pore complex functionally interacts with components of the spindle assembly checkpoint. J Cell Biol 159: 807– 819. Isgro TA, Schulten K. 2005. Binding dynamics of isolated nucleoporin repeat regions to importin-b. Structure 13: 1869–1879. Jankowsky E, Gross CH, Shuman S, Pyle AM. 2001. Active disruption of an RNA-protein interactionby a DExH/D RNA helicase. Science 291: 121–125. Johnson AW, Lund E, Dahlberg J. 2002. Nuclear export of ribosomal subunits. Trends Biochem Sci 27: 580– 585. Joseph J, Dasso M. 2008. The nucleoporin Nup358 associates with and regulates interphase microtubules. FEBS Lett 582: 190– 196. Joseph J, Liu ST, Jablonski SA, Yen TJ, Dasso M. 2004. The RanGAP1-RanBP2 complex is essential for microtubulekinetochore interactions in vivo. Curr Biol 14: 611– 617. Jovanovic-Talisman T, Tetenbaum-Novatt J, McKenney AS, Zilman A, Peters R, Rout MP, Chait BT. 2009. Artificial nanopores that mimic the transport selectivity of the nuclear pore complex. Nature 457: 1023–1027. Kalab P, Weis K, Heald R. 2002. Visualization of a Ran-GTP gradient in interphase and mitotic Xenopus egg extracts. Science 295: 2452– 2456. Kalverda B, Roling MD, Fornerod M. 2008. Chromatin organization in relation to the nuclear periphery. FEBS Lett 582: 2017–2022. Kalverda B, Pickersgill H, Shloma VV, Fornerod M. 2010. Nucleoporins Directly Stimulate Expression of Developmental and Cell-Cycle Genes Inside the Nucleoplasm. Cell 140: 360 –371. Keminer O, Peters R. 1999. Permeability of single nuclear pores. Biophys J 77: 217– 228. Krishnan VV, Lau EY, Yamada J, Denning DP, Patel SS, Colvin ME, Rexach MF. 2008. Intramolecular cohesion of coils mediated by phenylalanine–glycine motifs in the natively unfolded domain of a nucleoporin. PLoS Comput Biol 4: e1000145. Kubitscheck U, Grunwald D, Hoekstra A, Rohleder D, Kues T, Siebrasse JP, Peters R. 2005. Nuclear transport of single molecules: Dwell times at the nuclear pore complex. J Cell Biol 168: 233 –243. Kutay U, Hetzer MW. 2008. Reorganization of the nuclear envelope during open mitosis. Curr Opin Cell Biol 20: 669 –677. Kutay U, Bischoff FR, Kostka S, Kraft R, Gorlich D. 1997a. Export of importin a from the nucleus is mediated by a specific nuclear transport factor. Cell 90: 1061–1071. Kutay U, Izaurralde E, Bischoff FR, Mattaj IW, Gorlich D. 1997b. Dominant-negative mutants of importin-b block

Nuclear Pore Complexes and Nuclear Transport multiple pathways of import and export through the nuclear pore complex. EMBO J 16: 1153–1163. Kutay U, Lipowsky G, Izaurralde E, Bischoff FR, Schwarzmaier P, Hartmann E, Gorlich D. 1998. Identification of a tRNA-specific nuclear export receptor. Mol Cell 1: 359–369. Lee BJ, Cansizoglu AE, Suel KE, Louis TH, Zhang Z, Chook YM. 2006. Rules for nuclear localization sequence recognition by karyopherin b 2. Cell 126: 543– 558. Lee SH, Sterling H, Burlingame A, McCormick F. 2008. Tpr directly binds to Mad1 and Mad2 and is important for the Mad1-Mad2-mediated mitotic spindle checkpoint. Genes Dev 22: 2926–2931. Leksa NC, Brohawn SG, Schwartz TU. 2009. The structure of the scaffold nucleoporin Nup120 reveals a new and unexpected domain architecture. Structure 17: 1082– 1091. Lewis A, Felberbaum R, Hochstrasser M. 2007. A nuclear envelope protein linking nuclear pore basket assembly, SUMO protease regulation, and mRNA surveillance. J Cell Biol 178: 813 –827. Lim RY, Fahrenkrog B, Koser J, Schwarz-Herion K, Deng J, Aebi U. 2007a. Nanomechanical basis of selective gating by the nuclear pore complex. Science 318: 640– 643. Lim RY, Huang NP, Koser J, Deng J, Lau KH, SchwarzHerion K, Fahrenkrog B, Aebi U. 2006. Flexible phenylalanine-glycine nucleoporins as entropic barriers to nucleocytoplasmic transport. Proc Natl Acad Sci 103: 9512– 9517. Lim RY, Koser J, Huang NP, Schwarz-Herion K, Aebi U. 2007b. Nanomechanical interactions of phenylalanineglycine nucleoporins studied by single molecule forcevolume spectroscopy. J Struct Biol 159: 277 –289. Lim RY, Ullman KS, Fahrenkrog B. 2008. Biology and biophysics of the nuclear pore complex and its components. Int Rev Cell Mol Biol 267: 299–342. Lipowsky G, Bischoff FR, Izaurralde E, Kutay U, Schafer S, Gross HJ, Beier H, Gorlich D. 1999. Coordination of tRNA nuclear export with processing of tRNA. Rna 5: 539–549. Liu SM, Stewart M. 2005. Structural basis for the highaffinity binding of nucleoporin Nup1p to the Saccharomyces cerevisiae importin-b homologue, Kap95p. J Mol Biol 349: 515–525. Liu HL, De Souza CP, Osmani AH, Osmani SA. 2009. The three fungal transmembrane nuclear pore complex proteins of Aspergillus nidulans are dispensable in the presence of an intact An-Nup84–120 complex. Mol Biol Cell 20: 616 –630. Liu J, Prunuske AJ, Fager AM, Ullman KS. 2003. The COPI complex functions in nuclear envelope breakdown and is recruited by the nucleoporin Nup153. Dev Cell 5: 487–498. Lyman SK, Guan T, Bednenko J, Wodrich H, Gerace L. 2002. Influence of cargo size on Ran and energy requirements for nuclear protein import. J Cell Biol 159: 55– 67. Macara IG. 2001. Transport into and out of the nucleus. Microbiol Mol Biol Rev 65: 570– 594. Madrid AS, Weis K. 2006. Nuclear transport is becoming crystal clear. Chromosoma 115: 98–109. Mahajan R, Delphin C, Guan T, Gerace L, Melchior F. 1997. A small ubiquitin-related polypeptide involved in

targeting RanGAP1 to nuclear pore complex protein RanBP2. Cell 88: 97–107. Mattaj IW, Englmeier L. 1998. Nucleocytoplasmic transport: The soluble phase. Annu Rev Biochem 67: 265– 306. Matunis MJ, Wu J, Blobel G. 1998. SUMO-1 modification and its role in targeting the Ran GTPase-activating protein, RanGAP1, to the nuclear pore complex. J Cell Biol 140: 499–509. Mehlin H, Daneholt B, Skoglund U. 1992. Translocation of a specific premessenger ribonucleoprotein particle through the nuclear pore studied with electron microscope tomography. Cell 69: 605– 613. Miller BR, Powers M, Park M, Fischer W, Forbes DJ. 2000. Identification of a new vertebrate nucleoporin, Nup188, with the use of a novel organelle trap assay. Mol Biol Cell 11: 3381– 3396. Monecke T, Guttler T, Neumann P, Dickmanns A, Gorlich D, Ficner R. 2009. Crystal structure of the nuclear export receptor CRM1 in complex with Snurportin1 and RanGTP. Science 324: 1087–1091. Moroianu J, Blobel G, Radu A. 1997. RanGTP-mediated nuclear export of karyopherin a involves its interaction with the nucleoporin Nup153. Proc Natl Acad Sci 94: 9699–9704. Murphy R, Wente SR. 1996. An RNA-export mediator with an essential nuclear export signal. Nature 383: 357– 360. Nachury MV, Weis K. 1999. The direction of transport through the nuclear pore can be inverted. Proc Natl Acad Sci 96: 9622–9627. Nakielny S, Shaikh S, Burke B, Dreyfuss G. 1999. Nup153 is an M9-containing mobile nucleoporin with a novel Ran-binding domain. EMBO J 18: 1982–1995. Nehrbass U, Rout MP, Maguire S, Blobel G, Wozniak RW. 1996. The yeast nucleoporin Nup188p interacts genetically and physically with the core structures of the nuclear pore complex. J Cell Biol 133: 1153–1162. Oeffinger M, Dlakic M, Tollervey D. 2004. A pre-ribosomeassociated HEAT-repeat protein is required for export of both ribosomal subunits. Genes Dev 18: 196 –209. Ohno M, Segref A, Bachi A, Wilm M, Mattaj IW. 2000. PHAX, a mediator of U snRNA nuclear export whose activity is regulated by phosphorylation. Cell 101: 187 –198. Pante N, Kann M. 2002. Nuclear pore complex is able to transport macromolecules with diameters of about 39 nm. Mol Biol Cell 13: 425– 434. Paraskeva E, Izaurralde E, Bischoff FR, Huber J, Kutay U, Hartmann E, Luhrmann R, Gorlich D. 1999. CRM1mediated recycling of snurportin 1 to the cytoplasm. J Cell Biol 145: 255– 264. Patel SS, Belmont BJ, Sante JM, Rexach MF. 2007. Natively unfolded nucleoporins gate protein diffusion across the nuclear pore complex. Cell 129: 83–96. Pemberton LF, Paschal BM. 2005. Mechanisms of receptormediated nuclear import and nuclear export. Traffic 6: 187 –198. Peters R. 2005. Translocation through the nuclear pore complex: Selectivity and speed by reduction-of-dimensionality. Traffic 6: 421– 427.

S.R. Wente and M.P. Rout Pichler A, Gast A, Seeler JS, Dejean A, Melchior F. 2002. The nucleoporin RanBP2 has SUMO1 E3 ligase activity. Cell 108: 109– 120. Poon IK, Jans DA. 2005. Regulation of nuclear transport: Central role in development and transformation? Traffic 6: 173 –186. Rabut G, Doye V, Ellenberg J. 2004. Mapping the dynamic organization of the nuclear pore complex inside single living cells. Nat Cell Biol 6: 1114– 1121. Radu A, Moore MS, Blobel G. 1995. The peptide repeat domain of nucleoporin Nup98 functions as a docking site in transport across the nuclear pore complex. Cell 81: 215 –222. Reverter D, Lima CD. 2005. Insights into E3 ligase activity revealed by a SUMO-RanGAP1-Ubc9-Nup358 complex. Nature 435: 687–692. Rexach M, Blobel G. 1995. Protein import into nuclei: Association and dissociation reactions involving transport substrate, transport factors, and nucleoporins. Cell 83: 683–692. Ribbeck K, Gorlich D. 2002. The permeability barrier of nuclear pore complexes appears to operate via hydrophobic exclusion. EMBO J 21: 2664– 2671. Rodriguez MS, Dargemont C, Stutz F. 2004. Nuclear export of RNA. Biol Cell 96: 639–655. Rougemaille M, Dieppois G, Kisseleva-Romanova E, Gudipati RK, Lemoine S, Blugeon C, Boulay J, Jensen TH, Stutz F, Devaux F, et al. 2008. THO/Sub2p functions to coordinate 30 -end processing with gene-nuclear pore association. Cell 135: 308 –321. Rout MP, Wente SR. 1994. Pores for thought: Nuclear pore complex proteins. Trends Cell Biol 4: 357– 365. Rout MP, Aitchison JD. 2001. The nuclear pore complex as a transport machine. J Biol Chem 276: 16593– 16596. Rout MP, Aitchison JD, Magnasco MO, Chait BT. 2003. Virtual gating and nuclear transport: The hole picture. Trends Cell Biol 13: 622–628. Rout MP, Aitchison JD, Suprapto A, Hjertaas K, Zhao Y, Chait BT. 2000. The yeast nuclear pore complex: Composition, architecture, and transport mechanism. J Cell Biol 148: 635– 651. Roux KJ, Burke B. 2006. From pore to kinetochore and back: Regulating envelope assembly. Dev Cell 11: 276– 278. Salina D, Enarson P, Rattner JB, Burke B. 2003. Nup358 integrates nuclear envelope breakdown with kinetochore assembly. J Cell Biol 162: 991– 1001. Scarcelli JJ, Hodge CA, Cole CN. 2007. The yeast integral membrane protein Apq12 potentially links membrane dynamics to assembly of nuclear pore complexes. J Cell Biol 178: 799–812. Schmitt C, von Kobbe C, Bachi A, Pante N, Rodrigues JP, Boscheron C, Rigaut G, Wilm M, Seraphin B, CarmoFonseca M, et al. 1999. Dbp5, a DEAD-box protein required for mRNA export, is recruited to the cytoplasmic fibrils of nuclear pore complex via a conserved interaction with CAN/Nup159p. EMBO J 18: 4332–4347. Schneiter R, Hitomi M, Ivessa AS, Fasch EV, Kohlwein SD, Tartakoff AM. 1996. A yeast acetyl coenzyme A carboxylase mutant links very-long-chain fatty acid synthesis to the structure and function of the nuclear membrane-pore complex. Mol Cell Biol 16: 7161–7172.

Scott RJ, Lusk CP, Dilworth DJ, Aitchison JD, Wozniak RW. 2005. Interactions between Mad1p and the nuclear transport machinery in the yeast Saccharomyces cerevisiae. Mol Biol Cell 16: 4362– 4374. Seo HS, Ma Y, Debler EW, Wacker D, Kutik S, Blobel G, Hoelz A. 2009. Structural and functional analysis of Nup120 suggests ring formation of the Nup84 complex. Proc Natl Acad Sci 106: 14281–14286. Shulga N, Goldfarb DS. 2003. Binding dynamics of structural nucleoporins govern nuclear pore complex permeability and may mediate channel gating. Mol Cell Biol 23: 534 –542. Shulga N, Mosammaparast N, Wozniak R, Goldfarb DS. 2000. Yeast nucleoporins involved in passive nuclear envelope permeability. J Cell Biol 149: 1027–1038. Simos G, Grosshans H, Hurt E. 2002. Nuclear export of tRNA. Results Probl Cell Differ 35: 115– 131. Siniossoglou S, Lutzmann M, Santos-Rosa H, Leonard K, Mueller S, Aebi U, Hurt E. 2000. Structure and assembly of the Nup84p complex. J Cell Biol 149: 41– 54. Siniossoglou S, Wimmer C, Rieger M, Doye V, Tekotte H, Weise C, Emig S, Segref A, Hurt EC. 1996. A novel complex of nucleoporins, which includes Sec13p and a Sec13p homolog, is essential for normal nuclear pores. Cell 84: 265–275. Snay-Hodge CA, Colot HV, Goldstein AL, Cole CN. 1998. Dbp5p/Rat8p is a yeast nuclear pore-associated DEADbox protein essential for RNA export. EMBO J 17: 2663–2676. Stade K, Ford CS, Guthrie C, Weis K. 1997. Exportin 1 (Crm1p) is an essential nuclear export factor. Cell 90: 1041–1050. Stewart M. 2007. Ratcheting mRNA out of the nucleus. Mol Cell 25: 327–330. Strahm Y, Fahrenkrog B, Zenklusen D, Rychner E, Kantor J, Rosbach M, Stutz F. 1999. The RNA export factor Gle1p is located on the cytoplasmic fibrils of the NPC and physically interacts with the FG-nucleoporin Rip1p, the DEAD-box protein Rat8p/Dbp5p and a new protein Ymr 255p. EMBO J 18: 5761– 5777. Strawn LA, Shen T, Wente SR. 2001. The GLFG regions of Nup116p and Nup100p serve as binding sites for both Kap95p and Mex67p at the nuclear pore complex. J Biol Chem 276: 6445– 6452. Strawn LA, Shen T, Shulga N, Goldfarb DS, Wente SR. 2004. Minimal nuclear pore complexes define FG repeat domains essential for transport. Nat Cell Biol 6: 197– 206. Terry LJ, Wente SR. 2007. Nuclear mRNA export requires specific FG nucleoporins for translocation through the nuclear pore complex. J Cell Biol 178: 1121– 1132. Terry LJ, Shows EB, Wente SR. 2007. Crossing the nuclear envelope: Hierarchical regulation of nucleocytoplasmic transport. Science 318: 1412– 1416. Timney BL, Tetenbaum-Novatt J, Agate DS, Williams R, Zhang W, Chait BT, Rout MP. 2006. Simple kinetic relationships and nonspecific competition govern nuclear import rates in vivo. J Cell Biol 175: 579– 593. Tran EJ, Wente SR. 2006. Dynamic nuclear pore complexes: Life on the edge. Cell 125: 1041– 1053. Tran EJ, Bolger TA, Wente SR. 2007a. SnapShot: Nuclear transport. Cell 131: 420.

Nuclear Pore Complexes and Nuclear Transport Tran EJ, Zhou Y, Corbett AH, Wente SR. 2007b. The DEADbox protein Dbp5 controls mRNA export by triggering specific RNA:Protein remodeling events. Mol Cell 28: 850–859. Tseng SS, Weaver PL, Liu Y, Hitomi M, Tartakoff AM, Chang TH. 1998. Dbp5p, a cytosolic RNA helicase, is required for poly(A)þ RNA export. EMBO J 17: 2651–2662. Vasu S, Shah S, Orjalo A, Park M, Fischer WH, Forbes DJ. 2001. Novel vertebrate nucleoporins Nup133 and Nup160 play a role in mRNA export. J Cell Biol 155: 339–354. Weirich CS, Erzberger JP, Flick JS, Berger JM, Thorner J, Weis K. 2006. Activation of the DExD/H-box protein Dbp5 by the nuclear-pore protein Gle1 and its coactivator InsP6 is required for mRNA export. Nat Cell Biol 8: 668–676. Weis K. 2003. Regulating Access to the Genome. Nucleocytoplasmic Transport throughout the Cell Cycle. Cell 112: 441–451. Wente SR. 2000. Gatekeepers of the nucleus. Science 288: 1374– 1377. Whittle JR, Schwartz TU. 2009. Architectural nucleoporins Nup157/170 and Nup133 are structurally related and descend from a second ancestral element. J Biol Chem Wu J, Matunis MJ, Kraemer D, Blobel G, Coutavas E. 1995. Nup358, a cytoplasmically exposed nucleoporin with peptide repeats, Ran-GTP binding sites, zinc fingers, a cyclophilin A homologous domain, and a leucine-rich region. J Biol Chem 270: 14209–14213. Xu XM, Meulia T, Meier I. 2007. Anchorage of plant RanGAP to the nuclear envelope involves novel nuclearpore-associated proteins. Curr Biol 17: 1157–1163. Xu Z, Lam LS, Lam LH, Chau SF, Ng TB, Au SW. 2008. Molecular basis of the redox regulation of SUMO proteases: A protective mechanism of intermolecular disulfide linkage against irreversible sulfhydryl oxidation. FASEB J 22: 127–137. Yang W, Musser SM. 2006. Nuclear import time and transport efficiency depend on importin b concentration. J Cell Biol 174: 951 –961.

Yang Q, Jankowsky E. 2006. The DEAD-box protein Ded1 unwinds RNA duplexes by a mode distinct from translocating helicases. Nat Struct Mol Biol 13: 981–986. Yang W, Gelles J, Musser SM. 2004. Imaging of singlemolecule translocation through nuclear pore complexes. Proc Natl Acad Sci 101: 12887–12892. Yao W, Roser D, Kohler A, Bradatsch B, Bassler J, Hurt E. 2007. Nuclear export of ribosomal 60S subunits by the general mRNA export receptor Mex67-Mtr2. Mol Cell 26: 51–62. Yi H, Friedman JL, Ferreira PA. 2007. The cyclophilin-like domain of Ran-binding protein-2 modulates selectively the activity of the ubiquitin-proteasome system and protein biogenesis. J Biol Chem 282: 34770– 34778. Yokoyama N, Hayashi N, Seki T, Pante N, Ohba T, Nishii K, Kuma K, Hayashida T, Miyata T, Aebi U, et al. 1995. A giant nucleopore protein that binds Ran/TC4. Nature 376: 184–188. York JD, Odom AR, Murphy R, Ives EB, Wente SR. 1999. A phospholipase C-dependent inositol polyphosphate kinase pathway required for efficient messenger RNA export. Science 285: 96– 100. Yoshida K, Blobel G. 2001. The karyopherin Kap142p/ Msn5p mediates nuclear import and nuclear export of different cargo proteins. J Cell Biol 152: 729–740. Zeitler B, Weis K. 2004. The FG-repeat asymmetry of the nuclear pore complex is dispensible for bulk nucleocytoplasmic transport in vivo. J Cell Biol 167: 583–590. Zemp I, Kutay U. 2007. Nuclear export and cytoplasmic maturation of ribosomal subunits. FEBS Lett 581: 2783–2793. Zhang H, Saitoh H, Matunis MJ. 2002. Enzymes of the SUMO modification pathway localize to filaments of the nuclear pore complex. Mol Cell Biol 22: 6498–6508. Zhao Q, Brkljacic J, Meier I. 2008. Two distinct interacting classes of nuclear envelope-associated coiled-coil proteins are required for the tissue-specific nuclear envelope targeting of Arabidopsis RanGAP. Plant Cell 20: 1639– 1651. Zilman A, Di Talia S, Chait BT, Rout MP, Magnasco MO. 2007. Efficiency, selectivity, and robustness of nucleocytoplasmic transport. PLoS Comput Biol 3: e125.

Chromosome Territories Thomas Cremer1,2 and Marion Cremer1 1

Biozentrum, Department of Biology II (Chair of Anthropology and Human Genetics), Ludwig-MaximiliansUniversity, Grosshadernerstrasse 2, 82152 Martinsried, Germany

2

Munich Center for Integrated Protein Sciences (CIPSM), 81377 Munich, Germany

Correspondence: [email protected]

Chromosome territories (CTs) constitute a major feature of nuclear architecture. In a brief statement, the possible contribution of nuclear architecture studies to the field of epigenomics is considered, followed by a historical account of the CT concept and the final compelling experimental evidence of a territorial organization of chromosomes in all eukaryotes studied to date. Present knowledge of nonrandom CT arrangements, of the internal CT architecture, and of structural interactions with other CTs is provided as well as the dynamics of CT arrangements during cell cycle and postmitotic terminal differentiation. The article concludes with a discussion of open questions and new experimental strategies to answer them.

mpressive progress has been achieved during the last decade with regard to the functional implications of DNA methylation, histone modifications, and chromatin remodeling events for gene regulation (Fuks 2005; Kouzarides 2007; Maier et al. 2008; Jiang and Pugh 2009). It has, however, also become obvious that decoding the chromatin language does not suffice to fully understand the ways in which the diploid genome contributes to the formation of the different epigenomes present in the various cell types of a multicellular organism. Different epigenomes and their functional implications also depend on differences in higher-order chromatin organization and nuclear architecture at large. Epigenomic research aims

I

for an integrated understanding of the structural and functional aspects of epigenetics with nuclear architecture during the differentiation of toti- or pluripotent cells to functionally distinct cell types. The territorial organization of chromosomes in interphase (chromosome territories, CTs) constitutes a basic feature of nuclear architecture. This article starts with a brief historical account of the CT concept and the compelling experimental evidence in favor of a territorial organization of chromosomes in all eukaryotes studied to date. A survey of what is presently known about nonrandom arrangements of CTs, about changes of such arrangements in cycling cells as a result of internal or external

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a003889 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a003889

T. Cremer and M. Cremer

influences and about the internal architecture of CTs and their structural interactions with each other is provided. The article concludes with a discussion of open questions on CT organization and new experimental strategies to answer them.

ORIGIN OF THE CHROMOSOME TERRITORY CONCEPT

Since the late 19th century, an uncounted number of microscopic studies has appeared on numerous aspects of nuclear structure and on the observation of mitotic chromosomes. A territorial organization of interphase chromosomes was first suggested for animal cell nuclei by Carl Rabl (Rabl 1885) (Fig. 1A,B), but it was Theodor Boveri who introduced the term chromosome territory (CT) in his seminal studies of blastomere stages of the horse roundworm Parascaris equorum or Ascaris megalocephala, as the species was called at his time (Boveri 1909). The worm exists in two varieties, one with two pairs (Ascaris bivalens), the other with one pair of germ line chromosomes (Ascaris univalens). Boveri argued that each chromosome visible in mitosis retains its individuality during interphase and occupies a distinct part of the nuclear space. As can be seen in his drawings, Boveri was able to distinguish chromosome ends sticking out in protrusions of prophase nuclei (Fig. 1C) and used these protrusions as markers for the nuclear position of the asserted CTs in interphase nuclei (Fig. 1D). In fixed two-cell embryos, Boveri noted that their nuclear topography was strikingly similar during interphase and prophase, when chromosomes became visible as distinct entities. In four cell embryos, however, he typically observed two pairs of nuclei each carrying distinctly different protrusion patterns (Fig. 1D). His ingenious speculative talent led him to the following predictions on chromosome arrangements in Ascaris nuclei during the first steps of postzygotic development (for a comprehensive review, see Cremer and Cremer

2006a): (I) CTorder is stably maintained during interphase; (II) Chromosome neighborhood patterns change from prophase to metaphase; and (III) New chromosome neighborhood arrangements established in the metaphase plate are conserved to a considerable extent throughout anaphase and telophase, resulting in rather symmetrical arrangements of CTs in the two daughter nuclei. Different opinions on the structural organization of CTs were put forward in the early days of the 20th century. Eduard Strasburger (1905) published a colored cartoon claiming that CTs are built up from little chromatin clumps (Fig. 1E), whereas Theodor Boveri discussed a sponge-like CT structure built up from networks of anastomizing chromatin bundles (Fig. 1F). Despite light microscopic evidence in favor of chromosome territories at least in some species and cell types (for review see Stack et al. 1977) the weight of electron microscopic evidence established since the 1950s apparently argued for an unraveling of chromosomes in interphase nuclei into intermingling chromatin fibers of 10 – 30 nm in diameter with no sign of individual chromosomes. As a consequence, the concept of chromosome territories fell into oblivion or was even considered to be experimentally disproved (Wischnitzer 1973). During the 1970s and 1980s, most researchers seemed content with the assumption that the nucleus is filled with intermingling chromatin fibers and loops like a dish of spaghetti, an assumption widely reflected by textbooks of cell biology.

EXPERIMENTAL EVIDENCE FOR CHROMOSOME TERRITORIES

Stephen M. Stack, David B. Brown, and William C. Dewey were among the few researchers in the 1970s that still adhered to Boveri’s seemingly outdated concept (Stack et al. 1977; for review, see Cremer and Cremer 2006b). They squashed fixed cells from Allium cepa root tips, as well as Chinese hamster cells in acetic acid, and

Chromosome Territories

subjected the air-dried cells to salt solutions containing NaOH, and finally performed a Giemsa staining. This treatment yielded clumps of condensed chromatin arguably reflecting interphase chromosomes (Fig. 2A). The authors concluded that “chromosomes . . . remain in distinct domains throughout interphase.” Our group obtained early evidence for CTs in nuclei of diploid Chinese hamster cells with the help of laser-UV-microirradiation experiments (Zorn et al. 1976; Zorn et al. 1979; Cremer et al. 1982a; Cremer et al. 1982b; for review, see Cremer and Cremer 2006b; Meaburn and Misteli 2007). A laser microbeam (l ¼ 257 nm) was used to induce UV-damaged DNA within a small part of the nucleus. It was predicted that DNA damage inflicted within a small volume of the nucleus would yield different results depending on how chromosomes were arranged. Figure 2B exemplifies the experimental rationale with the example of woolen threads assembled within a “nuclear” space (Cremer et al. 1982a). Each thread reflects a chromatin fiber constituting an individual chromosome. In case of threads distributed throughout the whole nuclear space (upper panel), the “damage” label would become scattered over many threads. If individual threads occupy distinct territories (lower panel), localized label would mark only a small subset of threads. This experimental rationale was realized in the following way (Fig. 2C – E): Nuclei of living cells were microirradiated in G1. Locally damaged DNA was pulse-labeled with 3H-thymidine reflecting unscheduled DNA synthesis during excision repair of DNA photolesions. 3H-thymidine incorporation was detected by autoradiography in nuclei fixed immediately after the pulse (Fig. 2C) or in metaphase spreads prepared from cells that were allowed to proceed to the next mitosis (Fig. 2D). Alternatively, microirradiated DNA was visualized by immunostaining with antibodies raised against UV-damaged DNA (Cremer et al. 1984b) (Fig. 2E). As predicted by the CT concept, both approaches clearly showed that microirradiation of a small part of the nucleus damaged only a small

subset of the mitotic chromosome complement (Fig. 2D). The labeled parts of mitotic chromosomes revealed the UV damaged segments of neighboring chromosome territories hit by the microbeam during the preceding interphase. Consistent with this result, microirradiation of a small part of the metaphase plate of a living cell yielded a mirror-like pattern of distinctly labeled domains in the resulting daughter nuclei (Fig. 2F). These experiments provided the first compelling, although still indirect, evidence for the existence of chromosome territories. The direct visualization of individual CTs was made possible by in situ hybridization techniques developed during the mid 1980s. Initial experiments were performed with total human genomic DNA on cell hybrids that contained only one or few human chromosomes in a mouse or hamster genome complement (Manuelidis 1985; Schardin et al. 1985) (Fig. 3A). The achievement of chromosome sorting by flow cytometry of fluorescently labeled mitotic chromosomes (Cremer et al. 1984a; Gray et al. 1987; Fawcett et al. 1994) enabled the generation of chromosome specific painting probes for a large number of species. Subsequent amplification of DNA by cloning in bacterial vectors or by universal PCR (Telenius et al. 1992) and novel techniques for the suppression of ubiquitous repetitive sequences by COT-1 DNA (Cremer et al. 1988; Pinkel et al. 1988; Lichter et al. 1988a; Lichter et al. 1988b) or by depletion of these sequences from the respective probes (Bolzer et al. 1999) made it possible to delineate individual chromosomes in metaphase plates (Fig. 3B) and their territories in the interphase nucleus (Fig. 3C – F,K). To study the spatial arrangement of CTs, elaborate three-dimensional (3D) FISH protocols were developed (Cremer et al. 2008). 3D FISH in combination with light optical serial sectioning of nuclei by laser confocal microscopy and 3D image reconstruction has become the method of choice for studies of higher-order arrangements of CTs. The increasing availability of DNA probes for specific subchromosomal regions from chromosome arms down to bands

T. Cremer and M. Cremer

Figure 1. Early concepts in favor of a territorial organization of chromosomes in interphase nuclei. (A–B) Carl

Rabl´s hypothetical view (Rabl 1885) of a territorial chromosome arrangement in the interphase nucleus was based on studies of Proteus and Salamandra, in particular on epithelial cells from Salamandra maculata larvae. (A) Side view; supposed CTs are built up from primary chromatin threads (left side), from which (Continued )

Chromosome Territories

and single gene loci enabled studies of CT substructures (Fig. 3G – J). NONRANDOM ARRANGEMENTS OF CHROMOSOME TERRITORIES AND CHROMOSOMAL SUBREGIONS

Higher-order chromatin arrangements may primarily reflect geometrical constraints, which obviously affect the 3D distribution of larger and smaller CTs crowded together in the nuclear space (Neusser et al. 2007). Occasionally, however, proximity patterns assembled by chance, may have provided functional advantageous and consequently were favored by natural selection. The search for nonrandom chromatin assemblies, the mechanisms responsible for their formation and their functional implications is one of the major goals of nuclear architecture research. This search is still in its beginning. Methodological Problems in Distinguishing Random and Nonrandom Higher-Order Chromatin Arrangements

An overview on methods and problems for the spatial quantitative analysis of fluorescently labeled CTs and other nuclear structures is provided in Ronneberger et al. (2008). To decide about the random or nonrandom distribution

of a given target, such as a CT, chromosomal subregion, or gene, it is important to define a single or sometimes multiple 3D reference points, which represent the target in question in the nuclear space. For a painted CT, its intensity gravity center can be chosen as a single reference point or the CT surface may be used to define multiple reference points. Next, proper reference structures must be defined to decide whether the chosen target is distributed randomly or nonrandomly with respect to them. For reference structures, one can choose other chromatin targets or distinct nuclear structures, e.g., the nuclear lamina, nucleoli, or splicing speckles. Two cases of nonrandom arrangements of specific chromatin targets are considered here: nonrandom radial arrangements and nonrandom neighborhood arrangements, also referred to as proximity patterns. For studies of the radial arrangement, the nuclear space can be divided into a number of concentric shells with equal volume. 3D distances measured in a random sample of nuclei between the fluorescence intensity gravity center of a given labeled target and either the nuclear center or the closest point of the nuclear lamina provide information about its preferred radial nuclear position. Alternatively, all voxels contributing to a given target and reference structure, respectively, may be used to measure 3D

Figure 1. (Continued ). secondary and tertiary threads branch out and form a chromatin network (right side). Spindle attachment sites, now known as centromeres, congress at one site of the nucleus (top, Rabl’s Polfeld), whereas the telomeres cluster at the opposite site (bottom, Rabl’s Gegenpolseite). (B) View from above on the Polfeld. (C) Drawings made by Theodor Boveri (1909) from two Ascaris megalocephala univalens embryos fixed during prophase of the two-cell stage. Arrows point to nuclear protrusions containing the distal parts of the two germ line chromosomes. (D) Boveri’s drawing of a fixed four-cell embryo shows two pairs of cells with a distinctly different arrangement of these protrusions in interphase nuclei. Boveri argued that the two upper and two lower cells, respectively, represent daughter cells and explained the strikingly different nuclear protrusion patterns observed in the two cell pairs as a result of chromosome movements during prometaphase. (E) Eduard Strasburger’s colored model view of a tissue cell nucleus from the plant Galtonia candicans (Strasburger 1905). Chromosome territories supposedly are built up from higher-order chromatin foci delineated in red and blue. (F ) The white and gray shaded bundles in Boveri’s sketch from 1909 reflect two neighboring CTs with sponge-like structures built up from networks of anastomizing chromatin bundles. The continuous lines reflect Rabl’s primary threads, and the dotted line depicts the possibility of a rare pathological situation, in which secondary chromatin threads from one CT encompass the primary thread of the other, a situation possibly leading to an exchange of chromatin material between the chromosomes or a segregation failure during the next mitosis.

T. Cremer and M. Cremer

B

A

C

D 2

E

2 1

Figure 2. Experimental evidence for a territorial organization of interphase chromosomes. (A) Giemsa stained interphase nucleus of a fixed Chinese hamster cell (CHO line) reveals chromatin clumps likely representing individual CTs (reprinted with permission from Stack et al. 1977). (B) Experimental rationale of laser-UVmicrobeam experiments to distinguish between a nonterritorial (upper row) and a territorial (bottom row) chromosome arrangement in cell nuclei (compare Cremer et al. 1982a). (C) Autoradiograph of a diploid Chinese Hamster cell. The nucleus of the living cell was microirradiated in G1, pulse-labeled with 3H thymidine and fixed immediately thereafter. The arrow points to a cluster of silver grains detected over the site of microirradiation. (D) Metaphase spread from the same experiment obtained about 40 hours after microirradiation. One chromosome 1 and one chromosome 2 are intensely marked with silver grains, indicating that the microbeam hit the respective territories during interphase, whereas their homologs are unlabeled, arguing against the spatial association of the homolog territories. (E) Immunocytochemical identification of microirradiated DNA (arrows) in a pair of Chinese hamster daughter nuclei fixed around 4 hours after microirradiation of a small part of a metaphase plate. The mirror-like distribution of microirradiated chromatin in the two daughter nuclei argues for a similar arrangement of chromosome territories (reprinted from Cremer et al. [1984b] with permission).

distances. An equal frequency of voxels for a given chromatin target in all nuclear shells strongly argues for its random radial nuclear interior or peripheral arrangement of the target.

Nonrandom proximity patterns between sets of targets are indicated by 3D distance measurements, which are significantly smaller than expected in case of their random distribution

Chromosome Territories

in the nuclear space. It is important to note that nonrandom proximity patterns between genes located at distant sites of the same chromosome or on different chromosomes provide hints for but do not prove functional implications. For example, an apparently nonrandom proximity pattern of certain targets may simply reflect their nonrandom radial distribution in an interior nuclear shell, resulting in significantly smaller mean distances and a significantly higher frequency of contacts compared with targets randomly arranged in a peripheral nuclear shell. The importance of clearly defining targets and reference structures for which possible random or nonrandom arrangements are analyzed, can hardly be overemphasized. Certain targets may be arranged highly nonrandomly in the nuclear space, but still entirely at random in an internal nuclear shell or with respect to other more limited reference volumes or other reference structures. Before a detected nonrandom proximity pattern between certain targets gives rise to speculations about functional interactions, it should always be shown that this pattern cannot be simply explained as a consequence of a nonrandom radial distribution. As another caveat, it should be noted that the size of samples of nuclei subjected to a detailed 3D analysis is typically small (in the order of 20 – 50). It is of utmost importance to avoid a biased selection of such nuclei.

Radial Nuclear Arrangements of Chromosome Territories and Chromosome Subregions

Early evidence for a nonrandom radial distribution of entire CTs was based on (3D) FISH experiments in human lymphocyte nuclei using chromosome painting probes for human (HSA) chromosome 19, the chromosome with the highest gene density and for HSA 18, a gene poor chromosome. HSA 19 CTs were consistently found in the interior of human lymphocyte nuclei and of numerous other cell

types, whereas the territories of the gene poor HSA 18 were located at the nuclear periphery (Fig. 3C) (Croft et al. 1999; Cremer et al. 2001; Cremer et al. 2003). These observations of a gene density correlated radial arrangement in the nucleus were completed and confirmed by analyses comprising all human chromosomes (Boyle et al. 2001). An evolutionary comparison of lymphoblastoid cells from various primate species showed that this nonrandom radial nuclear distribution has been evolutionary conserved despite major evolutionary chromosome rearrangements. Orthologous segments of HSA 19 are positioned in the nuclear interior, whereas segments corresponding to human chromosome 18 locate at the nuclear periphery (Fig. 3D) (Tanabe et al. 2002). Nonrandom radial nuclear arrangements of CTs, depending on their gene density, were also observed in rodents (Mayer et al. 2005; Neusser et al. 2007), cattle (Koehler et al. 2009), and birds (Habermann et al. 2001). Recently, it was shown in bovine preimplantion embryos that this difference was not yet present in nuclei of early blastomere stages. Its first appearance correlated with major genome activation and was fully established in blastocysts (Koehler et al. 2009) (Fig. 3E). A multicolor 3D FISH approach was established for diploid human fibroblasts (46,XY) and allowed the colorful discrimination of the 22 pairs of autosmal CTs and the CTs of the two sex chromosomes (Bolzer et al. 2005) (Fig. 3F). For the flat-ellipsoid fibroblast nuclei, a radial arrangement of CTs, mainly according to chromosome size or DNA content, was shown in contrast to the clearly gene density correlated radial pattern observed in spherically shaped nuclei, such as nuclei in lymphocytes. Still, gene density correlated patterns on the subchromosomal level were also present in fibroblast nuclei, as shown by the preference of gene dense, Alu sequence-rich chromatin in the nuclear interior. 3D FISH experiments using sets of BAC clones with inserts from gene dense and gene poor chromosome segments confirmed such a distinct gene density correlated radial

T. Cremer and M. Cremer

A

D

B

#18 (5 genes/Mbp) #19 (20 genes/Mbp)

C

Human

E

Orangutan

6

F

White handed gibbon 3

6

1

6 Y

3

Y

3 6

2

1 1

1 2

G

H

2

I

Mb 0.5 1.0 1.5 2.0

11p15.5

J

K

Xa Xa Xi

Xa

Xi

Xi

Figure 3. Direct evidence for chromosome territories (CTs) by in situ hybridization experiments. (A) In situ

hybridization with biotinylated human genomic DNA of a Chinese hamster human hybrid cell line carrying a single human X chromosome reveals the specifically labeled human X chromosome (arrow) in metaphase spreads (top; other chromosomes are Giemsa stained) and its respective human X territory in interphase nuclei (bottom); for details, see Schardin et al. (1985). (B) Visualization of individual chromosomes in a human (HSA) metaphase plate (chr. HSA18 in red, chr. HSA 19 in green) after fluorescence in situ hybridization (FISH) using labeled chromosome painting probes. (C) A single light optical mid-section through the nucleus of a human lymphoblastoid cell after 3D FISH with the (Continued )

Chromosome Territories

nuclear arrangement (Fig. 3G,H). Present data provide strong evidence that the local gene density within windows of 2 – 10 Mb is apparently a strong and likely pivotal key player for the radial position of chromatin in the nucleus (Kozubek et al. 2002; Cremer et al. 2003; Murmann et al. 2005; Kupper et al. 2007). Other parameters, in particular transcriptional activity, replication timing, and GC content have also been

correlated with nonrandom radial nuclear arrangements of CTs and chromosomal subregions (Mayer et al. 2005; Federico et al. 2006; Goetze et al. 2007; Grasser et al. 2008; Hepperger et al. 2008). However, the interdependence of these parameters within a given DNA segment makes it difficult to dissect their impact on radial chromatin distribution in the nucleus (Kupper et al. 2007). The observation

Figure 3. (Continued ). same painting probes shows a HSA 19 CT (green) in the nuclear center and a HSA 18 CT

(red) at the periphery; for details, see Tanabe et al. (2002). (D) Top: Idiogramatic illustration of primate chromosomes or subchromosomal regions from the Orangutan ( pongus pygmaeus, PPY, middle) and white handed gibbon (Hylobates lar, HLA, right) orthologous to human chromosomes 18 (red) and 19 (green). Note the pronounced chromosomal rearrangements between the human and the HLA karyotype: The arm of a large HLA chromosome is orthologous to HSA 18, whereas four HSA 19 orthologous chromosome segments are distributed on three other large HLA chromosomes. Bottom: 3D reconstructions of representative HSA (left), PPY (middle), and HLA (right) lymphoblastoid cell nuclei reveal the same nonrandom radial nuclear positions of orthologous gene dense HSA 19 (green) and gene poor chromatin HSA 18 (red); for details, including quantitative evaluation, see Tanabe et al. (2002). (E) Painted CTs of the gene dense chromosome BTA 19 (green) and gene poor BTA 20 chromosome (red) in a domestic cattle (Bos taurus, BTA) embryo during blastocyst stage. Left: Maximum intensity projections of painted CTs in DAPI stained nuclei (blue). Right: 3D reconstructions of nuclei of the same embryo from a different perspective. Gene dense BTA 19 chromatin is preferentially distributed in the nuclear interior and gene poor BTA 20 chromatin at the nuclear periphery: for details, including quantitative evaluation, see Koehler et al. (2009). (F ) Simultaneous delineation of all chromosomes in a human fibroblast nucleus (left) and a prometaphase rosette (right) by multi-color FISH. Light optical mid-sections with false color representation of all CTs and prometaphase chromosomes, respectively, are shown. Examples of individual CTs and mitotic chromosomes are denoted with their respective karyotypic number; for details, see Bolzer et al. (2005). (G) Partial 3D reconstruction of a human lymphocyte nucleus after 3D-FISH of two differently labeled sets of BAC clones from HSA 12 carrying sequences from several gene-dense (green) and several gene-poor chromosome segments (red), respectively. This nucleus illustrates two neighboring HSA 12 CTs with distinct gene density correlated radial nuclear arrangement. (H ) Left: 3D reconstruction of a single painted HSA 12 territory (blue) showing the distinctly different, polarized arrangements of these two sets of BAC-clones. Right: 3D reconstruction of a HSA 12 CT recorded after 3D FISH with two differentially labeled sets of BAC clones containing sequences from highly expressed (green) and repressed genes (red), respectively, shows that active and silent genes are distributed throughout the CT; for details, see Kupper et al. (2007). (I ) Multicolor 3D FISH to a human fibroblast reveals the two HSA 11 CTs (blue) together with the particular gene dense region 11p15.5 (green, yellow, and red). For the delineation of this region, 15 BACs were used and differentially labeled as shown in the inset. Z-projections of light optical serial sections illustrate different shapes of this region: a finger-like chromatin protrusion in the lower CT and a much more compact shape in the upper CT; for details, see Albiez et al. (2006). (J ) Top: Two human X chromosomes in a human fibroblast metaphase plate are shown after multicolor FISH with four differentially labeled BAC pools representing four segments from qter to pter (q-arm: green, blue; p-arm: yellow, red). Bottom: Projections of light optical sections through the Xa- and Xi-territory of a human fibroblast nucleus following 3D FISH with the same BAC pools show four separate domains of these segments within the Xa- and Xi-territory (images courtesy of Kathrin Teller, Univ of Munich). (K) Left: Painting of the Xa- and Xi-territory in a female human fibroblast nucleus (46,XX) exemplifies the different shape and painting intensity of the two X-territories. The Xi-territory was independently identified by Barr body staining (not shown). Right: intensity profiles of enlarged images of the Xi- and Xa-territory. The color code white, yellow, and red reflects high, medium, and low intensities (images courtesy of Irina Solovei, Univ of Munich).

T. Cremer and M. Cremer

that radial positions of specific gene regions can differ significantly between cell types argues for additional cell-type-specific factors (Hepperger et al. 2008). The picture gets even more complex when we consider movements toward the nuclear interior shown for individual genes on transcriptional activation (see later). These movements emphasize that radial nuclear positions of genes do not solely depend on the local DNA sequence environment, where a given gene is embedded. Proximity Patterns of Chromosome Territories

Whereas little evidence for nonrandom neighborhood/proximity patterns was found between specific homologous and heterologous chromosomes in nuclei of human fibroblasts (Bolzer et al. 2005), nonrandom proximity patterns for smaller subsets of nonhomologous CTs were described by Misteli and coworkers in cell nuclei from various mouse tissues (Parada et al. 2002; Roix et al. 2003; Parada et al. 2004) and in human lymphocytes (Brianna Caddle et al. 2007; Khalil et al. 2007). These proximity patterns, however, were of a rather probabilistic nature, i.e., their presence was shown by an excess of certain heterologous (and occasionally also homologous) CTs in a population of nuclei, not as an event consistently observed in each nucleus. Differences of statistically preferred proximity patterns were recently also described for CT subsets in different fibroblast-derived cell lines (Zeitz et al. 2009). It should be emphasized that a pronounced cell-to-cell variation of CT neighborhood arrangements was found in all cell types studied so far. Neither functional implications of cell type or cell-line-specific probabilistic neighborhood arrangements nor mechanisms responsible for their establishment are presently understood. Such studies resemble snap-shots of individuals present in a room. A single snapshot does not allow conclusions about the dynamic behavior of these individuals. Accordingly, a decision whether certain CTs are permanently close to each other, have met before or will meet after the snap-shot was taken, would

require many snap-shots of well defined cell types in fixed samples at the same stage of interphase or postmitotic terminal differentiation, which are presently not available.

CT STRUCTURE, SHAPES, AND PLASTICITY

Apparently, 1 Mb-chromatin domains, first detected in S-phase nuclei as replication foci (Ma et al. 1998) and later shown to be persistent higher-order chromatin structures, are basic structural units, which build up CTs (Jackson and Pombo 1998; Visser and Aten 1999; Berezney et al. 2005; Albiez et al. 2006). To date, neither their ultrastructural organization nor the packaging of chromatin connecting these domains has been fully clarified. In addition, chromonema fibers may play an important role in the higher-order organization of CTs (Belmont and Bruce 1994) and their structural interconnection to global chromatin networks (Albiez et al. 2006). Single 1-Mb domains are likely built up from smaller loop domains, whereas larger chromatin clumps may be composed of clusters of 1-Mb domains. CTs visualized by 3D FISH appear as structures with manifold shapes composed of higher-order chromatin domains (Dietzel et al. 1998; Khalil et al. 2007; Kupper et al. 2007) (Fig. 3H–K). Separate arm domains were disclosed by painting of chromosome arms or parts of them in human cell nuclei (Dietzel et al. 1998) (Fig. 3K). The outer surface of an individual CTapparently does not generally provide a particular compartment for gene dense and/or transcriptionally active chromatin as originally suggested (Zirbel et al. 1993). A few chromatin regions with particular high gene density and/or transcriptional activity, such as the 11p15.5 segment (Fig. 3I), the MHC and EDC loci, or the HOX gene cluster have been consistently found looping out as protrusions from the core territory (Volpi et al. 2000; Williams et al. 2002; Chambeyron et al. 2005; Kupper et al. 2007). On a more global level, however, gene-dense and/or highly expressed sequences were found equally distributed throughout their respective territories (Mahy et al. 2002; Kupper et al. 2007)

Chromosome Territories

(Fig. 3H). The dissociation of gene localization relative to their CTs and gene regulation was shown for the murine Hoxd locus during differentiation and development: Although both decondensation and movement outside of the CT occur during gene activation in ES cells and the tail bud of the embryo, in the limb bud, gene activation and chromatin condensation occur without any looping out from the CT (Morey et al. 2007). This topic is described in detail in Heard and Bickmore (2007) and Morey et al. (2009). A particular staining intensity of the inactive X chromosome (Xi) with DNA-specific dyes was observed decades ago (so called Barr body [Lyon 1962]), indicating a higher compactness in Xi compared with the active X chromosome (Xa) or other autosomes (Fig. 3K, L). Previous studies showed distinct differences between Xi and Xa in shape and surface structure but surprisingly little differences in volume (Eils et al. 1996). In Xi, Clemson et al. (2006) observed a preferential positioning of genes —irrespective of their transcriptional activity— at the nuclear periphery, whereas repetitive sequences were found rather in the nuclear interior. Such a distinct radial arrangement was not reported for Xa, possibly because of larger invaginations in Xa. At large, however, structural differences between Xi and Xa on the subchromosomal level are poorly defined to date, although the CTs X provide an excellent model system for structure/function relationships in homologous chromosomes with different functional allocation. Detailed knowledge of the folding structure (conformation) of small contiguous chromatin segments within a given CT is limited so far to a few regions in autosomes and further studies are definitely required before a comprehensive picture can be drawn on the internal structure and local conformation of CTs. Shopland et al. (2006) analyzed the 3D structure of a highly conserved 4.3-Mb region on mouse chromosome 14 containing four clusters of genes separated by gene “deserts.” In Drosophila melanogaster, gene dense and gene poor segments within a 7-Mb region of chromosome 2 were described to form spatially segregated

clusters of different stability and folding structure (Boutanaev et al. 2005), and Goetze et al. (2007) showed for several cell lines that ridges (contiguous genomic regions harboring adjacent genes with high ubiquitous transcriptional activity) are in general less condensed and more irregularly shaped than “antiridges”, arguing that the structure of these two types of genomic domains is largely independent of tissuespecific variations in gene expression. MateosLangerak et al. (2009) provide data on the folding structure of subregions of CT 1 and 11 that are in accordance with a suggested random loop model with 10 – 30 loops/Mb. The relevance of 3D FISH studies for conclusions regarding the fine structure of CTs in vivo has to consider some technical issues. Differences in the fixation procedure of cells, amplification, and labeling efficiency of chromosome painting probes may have an impact on CT delineation. The necessary suppression or depletion of interspersed repetitive sequences prevents a complete visualization of CTs even under optimal hybridization conditions. The most crucial step for CT fine structure occurs during the denaturation of the nuclear DNA, a step necessary for successful probe hybridization (Solovei et al. 2002). These limitations emphasize the need for the visualization of CTs in living cells under conditions which do not interfere with chromatin or other nuclear functions (Zink et al. 1998; Walter et al. 2003). Although urgently required, in vivo approaches have their own technical problems, such as an interference of in vivo labeling strategies with the function of labeled chromatin and phototoxicity effects during prolonged observation periods. Different approaches have been established for the quantitative assessment of CT conformation. These include density, volume, roundness, and smoothness factors, as well as principal component analyses, which are also potentially useful to assess CT orientation within the nucleus (Eils et al. 1996; Roix et al. 2003; Ronneberger et al. 2008). The subjective choice of threshold setting for confocal image stacks influences the determination of CT borders and accordingly of measured CT volumes. Low threshold settings can result in the inclusion of

T. Cremer and M. Cremer

background, leading to overestimation of CT volumes and apparent chromatin intermingling between neighboring CTs, whereas high thresholds can lead to the visual loss of fine DNA structures looping out from the chromatin bulk of a given territory. STABILITY AND CHANGES OF CT ARRANGEMENTS IN CYCLING CELLS

The question of to what extent a given proximity pattern established between CTs during a given interphase may be transmitted through mitosis to the next interphase has remained controversial (Gerlich et al. 2003; Walter et al. 2003; Thomson et al. 2004; Kalmarova et al. 2008). These studies were based on live-cell approaches using transgenic cell lines with either FP-tagged or photoconvertible tagged histones. This allowed following up fluorescently labeled chromatin through mitosis into the next interphase. Present evidence argues for the stability of a given CT neighborhood arrangement once established at the onset of interphase until the next prophase, whereas chromosome movements during prometaphase necessary to establish the metaphase plate can lead to major changes of side-by-side chromosome arrangements in the metaphase plate compared with the side-by-side arrangements of the respective CTs during the preceding interphase (Walter et al. 2003; Cvackova et al. 2009). DYNAMICS OF CT ARRANGEMENTS DURING POSTMITOTIC CELL DIFFERENTIATION AND IN TERMINALLY DIFFERENTIATED CELLS

In a seminal investigation, Barr and Bertram described that on electric stimulation of cat motor neurons, a “nucleolar satellite” (now known as the Barr body) moved from its usual position adjacent to the nucleolus toward the nuclear membrane within a time course of several days (Barr and Bertram 1949). Borden and Manuelidis (1988) showed a pronounced repositioning of the human X-territories in neurons of both males and females in electrophysiologically

defined seizure foci. Repositioning of CTs during cellular differentiation of murine cerebellar Purkinje neurons was described by Martou and De Boni (2000) in terms of changes of centromere positions that adopted their final position around day 5 post partum. A striking example of CT reorganization during terminal differentiation was recently shown by Solovei et al. (2009) in a study of the mammalian retina. In mammals adapted to nocturnal life, all heterochromatin becomes located in the nuclear interior during postmitotic terminal differentiation of rod cells, whereas all euchromatin is shifted toward the nuclear periphery. This transformation starts around day 6 post partum and takes several weeks for completion. Rod cells of mammals with diurnal life styles do not show such a chromatin reorganization. Their nuclei reveal the conventional pattern with heterochromatin enriched at the nuclear periphery and around the nucleoli, whereas euchromatin is mostly distributed in the nuclear interior. This global nuclear reorganization in rod cells of nocturnal species necessitates a profound reorganization of radial chromatin arrangements rather than a change of CT proximity patterns. Unexpectedly, the inverted pattern of rod cell nuclei in nocturnal mammals reflects an adaptation to vision in low light conditions. Because of the somewhat higher refractive index of heterochromatin compared with less condensed euchromatin, inverted nuclei act as microlenses, which help to channel photons to the photoreceptors. This may be the first example in which a specific adaptive advantage of a celltype-specific nuclear architecture could be shown. Interestingly, this functional specialty shows to what extent the nuclear architecture can be modified under an overriding selective pressure. DYNAMICS AND INTERACTIONS OF SPECIFIC GENE LOCI LOCATED ON THE SAME OR ON DIFFERENT CTs

This question is particularly important with respect to long-range chromatin movements

Chromosome Territories

involved in the congression of coregulated genes. Live-cell experiments performed with cultured mammalian and Drosophila cells showed locally constrained movements of subchromosomal domains (Abney et al. 1997; Marshall et al. 1997; Bornfleth et al. 1999; Edelmann et al. 2001). The typical prevalence of heterochromatin localized at the lamina and the observation of silenced genes in this peripheral nuclear subcompartment has supported the concept that the nuclear periphery is a largely repressive environment for transcription, and vice versa, the nuclear interior a compartment for transcriptional activity (Schneider and Grosschedl 2007). This would require corresponding movements of gene loci in correlation to their (cell type) correlated transcriptional activity. Recent studies, however, support a more complex picture (Taddei et al. 2006; Akhtar and Gasser 2007; Deniaud and Bickmore 2009). Several groups succeeded to tether specific chromatin segments to the nuclear envelope in living cells. They found that some genes were suppressed when closely associated with the envelope, but that others were not (Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). Of particular interest are hints that a longrange spatial nuclear convergence of genes, which are located many megabases apart in cis, i.e., on the same chromosome, or trans, i.e., on different chromosomes, might be involved in mechanisms of gene activation or silencing (Zuckerkandl and Cavalli 2007; Bartkuhn and Renkawitz 2008). This phenomenon has been referred to as “gene kissing” (Kioussis 2005) or “chromosome kissing” (Cavalli 2007). As an early example, LaSalle and Lalande (1996) presented 3D FISH evidence for the transient spatial association of the AS/PWS loci during late S phase. These loci comprise the genes involved in two imprinting disorders, the Angelman syndrome and the Prader–Willi syndrome. The authors argued that transient “kissing” between the two loci is required for maintaining opposite imprints in cycling cells. This specific case of “kissing,” however, could not be confirmed in a later study (Teller et al. 2007).

A “nonmicroscopic” biochemical approach with a great potential for the disclosure of spatial interactions of specific genomic loci is the technique of “chromosome conformation capture” (3C) introduced by Dekker et al. (2002) and its recent extension to 4C (chromosome conformation capture-on-chip [Simonis et al. 2006] and “circular chromosome conformation capture” [Zhao et al. 2006]), respectively. With this “high throughput” approach, it has become possible to determine genome wide nonrandom spatial interactions between specific DNA segments in cis and trans to a reference locus on a given CT (for review, see Dekker 2008). This method is based on in situ formaldehyde cross-linking of proximal DNA– protein interactions with a distance of several A˚ngstro¨m, subsequent fragmentation of cross linked DNA, recircularization (ligation) and amplification, and finally identification of these products by microarray techniques. To date, numerous spatial interactions of CTs were reported in cis and trans (Ling et al. 2006; Lomvardas et al. 2006; Babu et al. 2008). These assays demand strict controls to escape the danger of false positive or negative findings (Simonis et al. 2007). By combining this proximity based ligation assay with massive parallel sequencing (Hi-C), the construction of spatial proximity maps of an entire genome down to a resolution level of 1 Mb was recently achieved as shown for a human lymphoblastoid cell line (LiebermanAiden et al. 2009). These maps confirmed the presence of chromosome territories, the spatial proximity of small, gene dense chromosomes, and the spatial segregation of open and closed chromatin, a parameter that strongly correlates with gene density (Gilbert et al. 2004). At the megabase scale, a chromatin conformation consistent with a fractal globule was suggested. High throughput assays with a superior resolution and microscopic approaches are complementary to further elucidate CT conformation and spatial interactions in trans (Simonis and de Laat 2008). Ligation techniques to date require millions of cells and their principle excludes an application at the single cell level. More important, microscopic approaches are

T. Cremer and M. Cremer

the only way to reveal the entire structure of nuclear components and to determine their topography with respect to each other. MODELS OF NUCLEAR ARCHITECTURE: OPEN QUESTIONS AND EXPERIMENTAL STRATEGIES TO ANSWER THEM

The territorial organization of interphase chromosomes is now generally accepted as a basic principle of nuclear organization in both animals (Cremer and Cremer 2001) and plants (Shaw et al. 2002; Pecinka et al. 2004; Berr et al. 2006), and may even hold for single-cell eukaryotes, such as budding and fission yeast (Bystricky et al. 2005; Molnar and Kleckner 2008). However, severe limitations in our present knowledge of the functional architecture of CTs and the nucleus at large become obvious when we consider currently proposed models (Fig. 4). The chromosome territory-interchromatin compartment (CT-IC) model (Fig. 4A,B) argues that nuclei are built up from two principal components, chromosome territories (CTs) and the interchromatin compartment (IC). Individual CTs form an interconnected higherorder chromatin network (Visser et al. 2000; Albiez et al. 2006). According to the CT-IC model, this chromatin network is spatially associated with a second contiguous 3D spatial network, the interchromatin compartment (IC), which was observed both at the light and electron microscopic level (Albiez et al. 2006; Rouquette et al. 2009). The IC concept asserts a DNA free or at least largely free, contiguous space of channels, which start at the nuclear pores and expand as larger channels and lacunes between the higher-order chromatin network described earlier. The IC harbors splicing speckles and a variety of nonchromatin nuclear bodies (Verschure et al. 1999; Visser et al. 2000; Albiez et al. 2006). The IC concept evolved from the interchromosomal domain (ICD) concept originally proposed by Peter Lichter and co-workers (Zirbel et al. 1993), who defined the ICD as a network-like space expanding mainly around CTs with little penetration into the CT interior (Cremer et al. 1993).

Supposedly, genes were preferentially transcribed in a region of decondensed chromatin at the CT periphery and RNA transcripts would be directly released into the ICD compartment. This concept was supported by a series of studies from this group (Bridger et al. 1998; Reichenzeller et al. 2000; Bridger et al. 2005; Richter et al. 2005). Accumulating evidence for genes transcribed both outside and in the interior of CTs (Cmarko et al. 1999; Verschure et al. 1999; Mahy et al. 2002; Kupper et al. 2007) is consistent with electron microscopic evidence for a (nearly) network-like DNA free space both outside and inside CTs (Visser et al. 2000; Rouquette et al. 2009). Based on this evidence, the hypothetical CT structure suggested by the CT-IC model can be compared with a sponge of chromatin permeated by intraterritorial IC channels (Fig. 4B). The entire IC is separated from the more condensed interior of chromatin domains and/or higher-order chromatin fibers by a thin (,200 nm) layer of rather decondensed chromatin, termed the perichromatin region (PR) (Fakan and van Driel 2007). EM evidence has supported the view that the PR topographically represents the utmost periphery of a given chromatin domain bordering the IC and functionally represents the major nuclear subcompartment for transcription, cotranscriptional RNA splicing (Fakan and Bernhard 1971; Cmarko et al. 1999; Trentani et al. 2003), as well as DNA replication (Jaunin and Fakan 2002) and possibly also DNA repair (Solimando et al. 2009). Transcription yields perichromatin fibrils, which are generated in the PR as nascent pre-mRNA transcripts of single genes complexed with hnRNPs and are served by nearby speckles with factors for cotranscriptional splicing. The PR concept is a decisive part of the CT-IC model but has been disregarded by proponents of other models (Dehghani et al. 2005; Branco and Pombo 2006; Fraser and Bickmore 2007; Alberts et al. 2008; Fedorova and Zink 2008; Fedorova and Zink 2009). The “lattice” model of interphase chromatin proposed by Dehghani et al. (2005) suggests a lattice-like network of 10- and 30-nm fibers. This structure yields a porous organization of chromatin with

Chromosome Territories

A

C

1 Mb chromatin domains

Perichromatin region

IC

5 P

CTs

B Chromosome territory

D Speckle

IC Genes

Chromatin domain

IC

PF Genes

Interchromatin compartment

Figure 4. Different models of nuclear architecture. (A) Chromosome territory-interchromatin compartment

(CT-IC) model (for description, see text). (B) Hypothetical view of the functional nuclear architecture according to the CT-IC model. Chromatin domains are considered the major constituents of a CT. The IC expands between these domains as a rather DNA free nuclear compartment carrying splicing speckles and nuclear bodies. The width of the IC space is highly variable depending on Brownian movements of chromatin domains and allowing transient contacts of domain surfaces in cis and trans. During ongoing transcription, genes are at least partially decondensed at any given time into the perichromatin region (PR) located at the domain periphery. Perichromatin fibrils (PF) are generated there. Each PF carries a nascent transcript (green) from a different gene. White dots with a line symbolize RNA Pol II molecules with their CTD domain, which may play a role in the structural organization of splicing events. Splicing speckles located in the IC provide the splicing factors to PFs, which also represent the structures in which cotranscriptional splicing occurs. (C) According to the interchromatin network (ICN) model (Branco and Pombo 2006), intermingling chromatin fibers/loops from the same CT, as well as from neighboring CTs, can make contact in cis and trans. Blue dots represent sites of intrachromosomal and interchromosomal contacts with unknown composition. Although there is extensive space between chromatin fibers/loops, this space should not be confused with the functional relationship of the IC and PR predicted by the CT-IC model. (D) Model suggested by Fraser and Bickmore (Fraser and Bickmore 2007, figure reprinted with permission from Macmillan Publishers Ltd). These authors review evidence arguing for the colocalization of genes in the nucleus for expression or coregulation. Transcription factories (dark pink) can recruit genes in cis and trans located on decondensed chromatin loops that extend outside chromosome territories. The pale pink area on the left represents a splicing-factor enriched speckle. The blue circle exemplifies an interaction for coregulation in trans, which can occur between regulatory elements and/or gene loci.

fibers intermingling at the borders of neighboring CTs. The interchromatin network (ICN) model (Fig. 4C) (Branco and Pombo 2006) predicts that chromatin fibers and loops intermingle in a rather uniform way both in the interior of individual CTs and between differentially labeled neighboring CTs, making any

distinction between the interior or periphery of distinct chromatin domains functionally meaningless. In this interchromatin network, loops may expand from one CT to meet loops from another CT. Arguably, active genes on decondensed chromatin loops that extend outside chromosome territories can colocalize both

T. Cremer and M. Cremer

in cis in so called expression hubs (Kosak and Groudine 2004) or be transcribed at preassembled transcription factories (Fig. 4D) (Fraser and Bickmore 2007). Some models emphasize the functional importance of giant chromatin loops, which emanate from chromosome territories (Chubb and Bickmore 2003; Fraser and Bickmore 2007). Supposedly, giant loops may even expand across the nuclear space (Alberts et al. 2008). According to these models, giant loops may carry genes to even very remote sites in the nuclear space for coregulation in an expression hub (Kosak and Groudine 2004). Alternatively, giant loops may transport genes to a remote repressive nuclear compartment (Alberts et al. 2008). The lack of quantitative rigor with respect to the predicted numbers, length, and compaction levels of such fibers and their 3D distribution has limited the usefulness of the models described earlier. To overcome this limitation, chromatin polymer models have been developed, which make experimentally testable, quantitative predictions about functionally important features of the nuclear architecture, such as the expected size distribution of chromatin loops and chromatin compaction levels (van den Engh et al. 1992; Munkel and Langowski 1998; Munkel et al. 1999). A recent chromatin polymer model has assumed a broad range of loop sizes (Mateos-Langerak et al. 2009). None of the present models for functional nuclear architecture is fully supported by compelling experimental evidence. In particular, two problems need urgent clarification: 1) the possible speed and extent of chromatin movements driven by Brownian motion in case of random walks and by unknown mechanisms in case of directed chromatin movements; 2) the topography of the major nuclear functions and the validation or experimental falsifications of the functional marriage between the IC and the PR as predicted by the CT-IC model (Fig. 4B) but not by other models (Fig. 4C, D). Although estimates of the number of transcription factories vary widely, the number of genes transcribed at any given time seems to be much larger. Accordingly, it has been suggested that a single transcription factory is able to

transcribe several genes simultaneously in cis and trans and that an extraordinarily large part of the genome passes through the limited number of transcription factories in a cell nucleus (Chakalova et al. 2005; Sutherland and Bickmore 2009). As a consequence, one would also expect an extraordinary mobility of chromatin in the nucleus. In contrast, the concept of perichromatin fibrils argues that most transcription takes place in the PR and that simultaneous transcription in a single factory is rather the exception than the rule, if this concept can finally be proven (Sutherland and Bickmore 2009). On the basis of FISH experiments with differentially labeled chromosome painting probes on cryosections (140 – 180 nm) from cell nuclei, Branco and Pombo (2006) detected zones of color overlap between pained CTs (Fig. 5A– C), which they interpreted as the result of intermingling chromatin fibers. Evidence for intermingling was substantiated by transmission electron microscopy with colloidal gold particles of different size for “intermingling” CTs. In control experiments, Branco and Pombo showed that the positions of gold grains reflecting the location of histone H2B molecules did not significantly change when cryosections were studied before and after mock FISH, excluding the possibility that chromatin intermingling was an artifact of chromatin denaturation. Examination of published electron micrographs does not, however, allow a morphological orientation in the section with regard to chromatin domains and other nuclear structures. Controversial issues both with respect to CT structure and the interchromatin space are in part because of the limited resolution of present light microscopy techniques and the difficulties to obtain 3D data sets by electron microscopy. Recent developments of laser-based light microscopic techniques with ultra-high resolution has now opened possibilities for light optical nanoscopy, which will help to overcome these limitations (Hell 2007; Schermelleh et al 2008; Gunkel et al. 2009). The combination of a focused ion beam with high resolution scanning electron microscopy is an additional, very promising tool for 3D analysis of chromosome architecture (Schroeder-Reiter et al. 2009).

Chromosome Territories

A

B

C

D

E

H 1

1

3

3

2

2

4

4

5

5

F

G

I 1

1

3

4 2

4 2 5

Figure 5. Experimental evidence against and in favor of an interchromatin space. Fluorescence microscopic

images of a painted HSA 5 CT (A) and HSA 7 CT (B) in a cryosection of approximately 150 nm thickness from a human lymphocyte nucleus (Branco and Pombo 2006). (C ) Overlap of images A and B shows the two CTs in different colors. The area of assumed intermingling between the two CTs is delineated by the yellow line. (D –G) Topographical relationships between double minute (DM) chromosomes carrying active MYCN genes (red) and the painted 3q-arm domain (green) observed in a nucleus of the human neuroblastoma cell HDN-16 (for details, see Solovei et al. 2000). (D) Laser confocal section. DMs denoted by 1 –5 are located both at the periphery of the 3q-arm domain and in channel-like invaginations of the interchromatin compartment (IC) penetrating into the interior of the arm domain. (E) IC invaginations are emphasized in this gray image of the same 3q-arm domain section. Numbers indicate the location of DMs 1 – 5 in D. (F–G) 3D reconstruction of the entire 3q-domain (green) with all associated DMs (red). (F) top-view and (G) section through the reconstructed domain. (H,I) 3D reconstruction of a rat liver cell nucleus specifically stained for DNA and imaged by Serial Block-Face—Scanning Electron Microscopy (Rouquette et al. 2009). (H ) A reconstructed mid part of the nucleus with 250-nm thickness shows a strongly inhomogeneous distribution of DNA in higher-order chromatin clusters (gray) and the wide mostly DNA free interchromatin compartment (white) expanding between these clusters. (I ) Reconstruction of the major part of the nucleus. The dotted line indicates the removal of the nuclear periphery exposed to the viewer. Here, the nuclear volume seems to be entirely filled with chromatin, most likely—as we suggest—as a consequence of the sponge-like organization of CTs built up from interconnected chromatin domains/bundles permeated by the IC.

T. Cremer and M. Cremer

ACKNOWLEDGMENTS

This work was supported by DFG (Cr 59/28-1) and CIPSM. The authors are grateful to the many colleagues of the group of the Cremer laboratory who contributed with their experiments, ideas, and enthusiasm to unravel the nature of CTs.

REFERENCES Abney JR, Cutler B, Fillbach ML, Axelrod D, Scalettar BA. 1997. Chromatin dynamics in interphase nuclei and its implications for nuclear structure. J Cell Biol 137: 1459– 1468. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P. 2008. Molecular biology of the cell. Garland Science, New York. Albiez H, Cremer M, Tiberi C, Vecchio L, Schermelleh L, Dittrich S, Kupper K, Joffe B, Thormeyer T, von Hase J, et al. 2006. Chromatin domains and the interchromatin compartment form structurally defined and functionally interacting nuclear networks. Chromosome Res 14: 707 – 733. Babu DA, Chakrabarti SK, Garmey JC, Mirmira RG. 2008. Pdx1 and b2/NeuroD1 participate in a transcriptional complex that mediates short-range DNA looping at the insulin gene. J Biol Chem 283: 8164– 8172. Barr ML, Bertram EG. 1949. A morphological distinction between neurons of the male and female, and the behavior of the nucleolar satellite during accelerated nucleoprotein synthesis. Nature 163: 676–677. Bartkuhn M, Renkawitz R. 2008. Long range chromatin interactions involved in gene regulation. Biochim Biophys Acta 1783: 2161–2166. Belmont AS, Bruce K. 1994. Visualization of G1 chromosomes: A folded, twisted, supercoiled chromonema model of interphase chromatid structure. J Cell Biol 127: 287– 302. Berezney R, Malyavantham KS, Pliss A, Bhattacharya S, Acharya R. 2005. Spatio-temporal dynamics of genomic organization and function in the mammalian cell nucleus. Adv Enzyme Regul 45: 17– 26. Berr A, Pecinka A, Meister A, Kreth G, Fuchs J, Blattner FR, Lysak MA, Schubert I. 2006. Chromosome arrangement and nuclear architecture but not centromeric sequences are conserved between Arabidopsis thaliana and Arabidopsis lyrata. Plant J 48: 771 –783. Bolzer A, Craig JM, Cremer T, Speicher MR. 1999. A complete set of repeat-depleted, PCR-amplifiable, human chromosome-specific painting probes. Cytogenet Cell Genet 84: 233 –240. Bolzer A, Kreth G, Solovei I, Koehler D, Saracoglu K, Fauth C, Muller S, Eils R, Cremer C, Speicher MR, et al. 2005. Three-dimensional maps of all chromosomes in human male fibroblast nuclei and prometaphase rosettes. PLoS Biol 3: e157.

Borden J, Manuelidis L. 1988. Movement of the X chromosome in epilepsy. Science 242: 1687– 1691. Bornfleth H, Edelmann P, Zink D, Cremer T, Cremer C. 1999. Quantitative motion analysis of subchromosomal foci in living cells using four-dimensional microscopy. Biophys J 77: 2871–2886. Boutanaev AM, Mikhaylova LM, Nurminsky DI. 2005. The pattern of chromosome folding in interphase is outlined by the linear gene density profile. Mol Cell Biol 25: 8379–8386. Boveri T. 1909. Die Blastomerenkerne von Ascaris megalocephala und die Theorie der Chromosomenindividualita¨t. Arch Zellforsch 3: 181– 268. Boyle S, Gilchrist S, Bridger JM, Mahy NL, Ellis JA, Bickmore WA. 2001. The spatial organization of human chromosomes within the nuclei of normal and emerinmutant cells. Hum Mol Genet 10: 211 –219. Branco MR, Pombo A. 2006. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol 4: e138. Brianna Caddle L, Grant JL, Szatkiewicz J, van Hase J, Shirley BJ, Bewersdorf J, Cremer C, Arneodo A, Khalil A, Mills KD. 2007. Chromosome neighborhood composition determines translocation outcomes after exposure to high-dose radiation in primary cells. Chromosome Res 15: 1061–1073. Bridger JM, Herrmann H, Munkel C, Lichter P. 1998. Identification of an interchromosomal compartment by polymerization of nuclear-targeted vimentin. J Cell Sci 111: 1241– 1253. Bridger JM, Kalla C, Wodrich H, Weitz S, King JA, Khazaie K, Krausslich HG, Lichter P. 2005. Nuclear RNAs confined to a reticular compartment between chromosome territories. Exp Cell Res 302: 180–193. Bystricky K, Laroche T, van Houwe G, Blaszczyk M, Gasser SM. 2005. Chromosome looping in yeast: telomere pairing and coordinated movement reflect anchoring efficiency and territorial organization. J Cell Biol 168: 375 –387. Cavalli G. 2007. Chromosome kissing. Curr Opin Genet Dev 17: 443– 450. Chakalova L, Debrand E, Mitchell JA, Osborne CS, Fraser P. 2005. Replication and transcription: shaping the landscape of the genome. Nat Rev Genet 6: 669–677. Chambeyron S, Da Silva NR, Lawson KA, Bickmore WA. 2005. Nuclear re-organisation of the Hoxb complex during mouse embryonic development. Development 132: 2215–2223. Chubb JR, Bickmore WA. 2003. Considering nuclear compartmentalization in the light of nuclear dynamics. Cell 112: 403–406. Clemson CM, Hall LL, Byron M, McNeil J, Lawrence JB. 2006. The X chromosome is organized into a gene-rich outer rim and an internal core containing silenced nongenic sequences. Proc Natl Acad Sci 103: 7688– 7693. Cmarko D, Verschure PJ, Martin TE, Dahmus ME, Krause S, Fu XD, van Driel R, Fakan S. 1999. Ultrastructural analysis of transcription and splicing in the cell nucleus after bromo-UTP microinjection. Mol Biol Cell 10: 211 –223.

Chromosome Territories Cremer C, Rappold G, Gray JW, Muller CR, Ropers HH. 1984a. Preparative dual-beam sorting of the human Y chromosome and in situ hybridization of cloned DNA probes. Cytometry 5: 572 –579. Cremer M, Grasser F, Lanctot C, Muller S, Neusser M, Zinner R, Solovei I, Cremer T. 2008. Multicolor 3D fluorescence in situ hybridization for imaging interphase chromosomes. Methods Mol Biol 463: 205 –239. Cremer M, Kupper K, Wagler B, Wizelman L, von Hase J, Weiland Y, Kreja L, Diebold J, Speicher MR, Cremer T. 2003. Inheritance of gene density-related higher order chromatin arrangements in normal and tumor cell nuclei. J Cell Biol 162: 809–820. Cremer M, von Hase J, Volm T, Brero A, Kreth G, Walter J, Fischer C, Solovei I, Cremer C, Cremer T. 2001. Nonrandom radial higher-order chromatin arrangements in nuclei of diploid human cells. Chromosome Res 9: 541 – 567. Cremer T, Baumann H, Nakanishi K, Cremer C. 1984b. Correlation between interphase and metaphase chromosome arrangements as studied by laser-uv-microbeam experiments. Chromosomes Today 8: 203– 212. Cremer T, Cremer C. 2001. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat Rev Genet 2: 292– 301. Cremer T, Cremer C. 2006a. Rise, fall and resurrection of chromosome territories: a historical perspective. Part I. The rise of chromosome territories. Eur J Histochem 50: 161 –176. Cremer T, Cremer C. 2006b. Rise, fall and resurrection of chromosome territories: a historical perspective. Part II. Fall and resurrection of chromosome territories during the 1950s to 1980s. Part III. Chromosome territories and the functional nuclear architecture: experiments and models from the 1990s to the present. Eur J Histochem 50: 223–272. Cremer T, Cremer C, Baumann H, Luedtke EK, Sperling K, Teuber V, Zorn C. 1982a. Rabl’s model of the interphase chromosome arrangement tested in Chinese hamster cells by premature chromosome condensation and laser-UV-microbeam experiments. Hum Genet 60: 46– 56. Cremer T, Cremer C, Schneider T, Baumann H, Hens L, Kirsch-Volders M. 1982b. Analysis of chromosome positions in the interphase nucleus of Chinese hamster cells by laser-UV-microirradiation experiments. Hum Genet 62: 201 –209. Cremer T, Kurz A, Zirbel R, Dietzel S, Rinke B, Schrock E, Speicher MR, Mathieu U, Jauch A, Emmerich P, et al. 1993. Role of chromosome territories in the functional compartmentalization of the cell nucleus. Cold Spring Harb Symp Quant Biol 58: 777– 792. Cremer T, Lichter P, Borden J, Ward DC, Manuelidis L. 1988. Detection of chromosome aberrations in metaphase and interphase tumor cells by in situ hybridization using chromosome-specific library probes. Hum Genet 80: 235–246. Croft JA, Bridger JM, Boyle S, Perry P, Teague P, Bickmore WA. 1999. Differences in the localization and morphology of chromosomes in the human nucleus. J Cell Biol 145: 1119–1131.

Cvackova Z, Masata M, Stanek D, Fidlerova H, Raska I. 2009. Chromatin position in human HepG2 cells: although being non-random, significantly changed in daughter cells. J Struct Biol 165: 107– 117. Dehghani H, Dellaire G, Bazett-Jones DP. 2005. Organization of chromatin in the interphase mammalian cell. Micron 36: 95–108. Dekker J. 2008. Gene regulation in the third dimension. Science 319: 1793– 1794. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306– 1311. Deniaud E, Bickmore WA. 2009. Transcription and the nuclear periphery: Edge of darkness? Curr Opin Genet Dev 19: 187– 191. Dietzel S, Jauch A, Kienle D, Qu G, Holtgreve-Grez H, Eils R, Munkel C, Bittner M, Meltzer PS, Trent JM, et al. 1998. Separate and variably shaped chromosome arm domains are disclosed by chromosome arm painting in human cell nuclei. Chromosome Res 6: 25– 33. Edelmann P, Bornfleth H, Zink D, Cremer T, Cremer C. 2001. Morphology and dynamics of chromosome territories in living cells. Biochim Biophys Acta 1551: M29–39. Eils R, Dietzel S, Bertin E, Schrock E, Speicher MR, Ried T, Robert-Nicoud M, Cremer C, Cremer T. 1996. Threedimensional reconstruction of painted human interphase chromosomes: Active and inactive X chromosome territories have similar volumes but differ in shape and surface structure. J Cell Biol 135: 1427– 1440. Fakan S, Bernhard W. 1971. Localisation of rapidly and slowly labelled nuclear RNA as visualized by high resolution autoradiography. Exp Cell Res 67: 129 –141. Fakan S, van Driel R. 2007. The perichromatin region: A functional compartment in the nucleus that determines large-scale chromatin folding. Semin Cell Dev Biol 18: 676 –681. Fawcett JJ, Longmire JL, Martin JC, Deaven LL, Cram LS. 1994. Large-scale chromosome sorting. Methods Cell Biol 42: 319 –330. Federico C, Scavo C, Cantarella CD, Motta S, Saccone S, Bernardi G. 2006. Gene-rich and gene-poor chromosomal regions have different locations in the interphase nuclei of cold-blooded vertebrates. Chromosoma 115: 123– 128. Fedorova E, Zink D. 2008. Nuclear architecture and gene regulation. Biochim Biophys Acta 1783: 2174–2184. Fedorova E, Zink D. 2009. Nuclear genome organization: common themes and individual patterns. Curr Opin Genet Dev 19: 166– 171. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Fraser P, Bickmore W. 2007. Nuclear organization of the genome and the potential for gene regulation. Nature 447: 413–417. Fuks F. 2005. DNA methylation and histone modifications: teaming up to silence genes. Curr Opin Genet Dev 15: 490 –495. Gerlich D, Beaudouin J, Kalbfuss B, Daigle N, Eils R, Ellenberg J. 2003. Global chromosome positions are transmitted through mitosis in mammalian cells. Cell 112: 751 –764.

T. Cremer and M. Cremer Gilbert N, Boyle S, Fiegler H, Woodfine K, Carter NP, Bickmore WA. 2004. Chromatin architecture of the human genome: gene-rich domains are enriched in open chromatin fibers. Cell 118: 555–566. Goetze S, Mateos-Langerak J, Gierman HJ, de Leeuw W, Giromus O, Indemans MH, Koster J, Ondrej V, Versteeg R, van Driel R. 2007. The three-dimensional structure of human interphase chromosomes is related to the transcriptome map. Mol Cell Biol 27: 4475–4487. Grasser F, Neusser M, Fiegler H, Thormeyer T, Cremer M, Carter NP, Cremer T, Muller S. 2008. Replicationtiming-correlated spatial chromatin arrangements in cancer and in primate interphase nuclei. J Cell Sci 121: 1876– 1886. Gray JW, Dean PN, Fuscoe JC, Peters DC, Trask BJ, van den Engh GJ, Van Dilla MA. 1987. High-speed chromosome sorting. Science 238: 323 –329. Gunkel M, Erdel F, Rippe K, Lemmer P, Kaufmann R, Hormann C, Amberger R, Cremer C. 2009. Dual color localization microscopy of cellular nanostructures. Biotechnol J 4: 927 –938. Habermann FA, Cremer M, Walter J, Kreth G, von Hase J, Bauer K, Wienberg J, Cremer C, Cremer T, Solovei I. 2001. Arrangements of macro- and microchromosomes in chicken cells. Chromosome Res 9: 569–584. Heard E, Bickmore W. 2007. The ins and outs of gene regulation and chromosome territory organisation. Curr Opin Cell Biol 19: 311 –316. Hell SW. 2007. Far-field optical nanoscopy. Science 316: 1153– 1158. Hepperger C, Mannes A, Merz J, Peters J, Dietzel S. 2008. Three-dimensional positioning of genes in mouse cell nuclei. Chromosoma 117: 535–551. Jackson DA, Pombo A. 1998. Replicon clusters are stable units of chromosome structure: evidence that nuclear organization contributes to the efficient activation and propagation of S phase in human cells. J Cell Biol 140: 1285– 1295. Jaunin F, Fakan S. 2002. DNA replication and nuclear architecture. J Cell Biochem 85: 1 –9. Jiang C, Pugh BF. 2009. Nucleosome positioning and gene regulation: advances through genomics. Nat Rev Genet 10: 161 –172. Kalmarova M, Smirnov E, Kovacik L, Popov A, Raska I. 2008. Positioning of the NOR-bearing chromosomes in relation to nucleoli in daughter cells after mitosis. Physiol Res 57: 421– 425. Khalil A, Grant JL, Caddle LB, Atzema E, Mills KD, Arneodo A. 2007. Chromosome territories have a highly nonspherical morphology and nonrandom positioning. Chromosome Res 15: 899 –916. Kioussis D. 2005. Gene regulation: Kissing chromosomes. Nature 435: 579–580. Koehler D, Zakhartchenko V, Froenicke L, Stone G, Stanyon R, Wolf E, Cremer T, Brero A. 2009. Changes of higher order chromatin arrangements during major genome activation in bovine preimplantation embryos. Exp Cell Res 315: 2053–2063. Kosak ST, Groudine M. 2004. Form follows function: The genomic organization of cellular differentiation. Genes Dev 18: 1371–1384.

Kouzarides T. 2007. Chromatin modifications and their function. Cell 128: 693 –705. Kozubek S, Lukasova E, Jirsova P, Koutna I, Kozubek M, Ganova A, Bartova E, Falk M, Pasekova R. 2002. 3D Structure of the human genome: order in randomness. Chromosoma 111: 321– 331. Kumaran RI, Spector DL. 2008. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol 180: 51–65. Kupper K, Kolbl A, Biener D, Dittrich S, von Hase J, Thormeyer T, Fiegler H, Carter NP, Speicher MR, Cremer T, et al. 2007. Radial chromatin positioning is shaped by local gene density, not by gene expression. Chromosoma 116: 285–306. LaSalle JM, Lalande M. 1996. Homologous association of oppositely imprinted chromosomal domains. Science 272: 725–728. Lichter P, Cremer T, Borden J, Manuelidis L, Ward DC. 1988a. Delineation of individual human chromosomes in metaphase and interphase cells by in situ suppression hybridization using recombinant DNA libraries. Hum Genet 80: 224– 234. Lichter P, Cremer T, Tang CJ, Watkins PC, Manuelidis L, Ward DC. 1988b. Rapid detection of human chromosome 21 aberrations by in situ hybridization. Proc Natl Acad Sci 85: 9664–9668. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289– 293. Ling JQ, Li T, Hu JF, Vu TH, Chen HL, Qiu XW, Cherry AM, Hoffman AR. 2006. CTCF mediates interchromosomal colocalization between Igf2/H19 and Wsb1/Nf1. Science 312: 269–272. Lomvardas S, Barnea G, Pisapia DJ, Mendelsohn M, Kirkland J, Axel R. 2006. Interchromosomal interactions and olfactory receptor choice. Cell 126: 403 –413. Lyon MF. 1962. Sex chromatin and gene action in the mammalian X-chromosome. Am J Hum Genet 14: 135 –148. Ma H, Samarabandu J, Devdhar RS, Acharya R, Cheng PC, Meng C, Berezney R. 1998. Spatial and temporal dynamics of DNA replication sites in mammalian cells. J Cell Biol 143: 1415–1425. Mahy NL, Perry PE, Gilchrist S, Baldock RA, Bickmore WA. 2002. Spatial organization of active and inactive genes and noncoding DNA within chromosome territories. J Cell Biol 157: 579– 589. Maier VK, Chioda M, Becker PB. 2008. ATP-dependent chromatosome remodeling. Biol Chem 389: 345– 352. Manuelidis L. 1985. Individual interphase chromosome domains revealed by in situ hybridization. Hum Genet 71: 288– 293. Marshall WF, Straight A, Marko JF, Swedlow J, Dernburg A, Belmont A, Murray AW, Agard DA, Sedat JW. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930–939. Martou G, De Boni U. 2000. Nuclear topology of murine, cerebellar Purkinje neurons: changes as a function of development. Exp Cell Res 256: 131– 139.

Chromosome Territories Mateos-Langerak J, Bohn M, de Leeuw W, Giromus O, Manders EM, Verschure PJ, Indemans MH, Gierman HJ, Heermann DW, van Driel R, et al. 2009. Spatially confined folding of chromatin in the interphase nucleus. Proc Natl Acad Sci 106: 3812– 3817. Mayer R, Brero A, von Hase J, Schroeder T, Cremer T, Dietzel S. 2005. Common themes and cell type specific variations of higher order chromatin arrangements in the mouse. BMC Cell Biol 6: 44. Meaburn KJ, Misteli T. 2007. Cell biology: chromosome territories. Nature 445: 379 –781. Molnar M, Kleckner N. 2008. Examination of interchromosomal interactions in vegetatively growing diploid Schizosaccharomyces pombe cells by Cre/loxP site-specific recombination. Genetics 178: 99– 112. Morey C, Da Silva NR, Perry P, Bickmore WA. 2007. Nuclear reorganisation and chromatin decondensation are conserved, but distinct, mechanisms linked to Hox gene activation. Development 134: 909–919. Morey C, Kress C, Bickmore WA. 2009. Lack of bystander activation shows that localization exterior to chromosome territories is not sufficient to up-regulate gene expression. Genome Res 19: 1184–1194. Munkel C, Eils R, Dietzel S, Zink D, Mehring C, Wedemann G, Cremer T, Langowski J. 1999. Compartmentalization of interphase chromosomes observed in simulation and experiment. J Mol Biol 285: 1053– 1065. Munkel C, Langowski J. 1998. Chromosome structure predicted by a polymer model. Phys Rev E57: 5888– 5896. Murmann AE, Gao J, Encinosa M, Gautier M, Peter ME, Eils R, Lichter P, Rowley JD. 2005. Local gene density predicts the spatial position of genetic loci in the interphase nucleus. Exp Cell Res 311: 14–26. Neusser M, Schubel V, Koch A, Cremer T, Muller S. 2007. Evolutionarily conserved, cell type and species-specific higher order chromatin arrangements in interphase nuclei of primates. Chromosoma 116: 307 –320. Parada LA, McQueen PG, Misteli T. 2004. Tissue-specific spatial organization of genomes. Genome Biol 5: R44. Parada LA, McQueen PG, Munson PJ, Misteli T. 2002. Conservation of relative chromosome positioning in normal and cancer cells. Curr Biol 12: 1692–1697. Pecinka A, Schubert V, Meister A, Kreth G, Klatte M, Lysak MA, Fuchs J, Schubert I. 2004. Chromosome territory arrangement and homologous pairing in nuclei of Arabidopsis thaliana are predominantly random except for NOR-bearing chromosomes. Chromosoma 113: 258–269. Pinkel D, Landegent J, Collins C, Fuscoe J, Segraves R, Lucas J, Gray J. 1988. Fluorescence in situ hybridization with human chromosome-specific libraries: detection of trisomy 21 and translocations of chromosome 4. Proc Natl Acad Sci 85: 9138–9142. ¨ ber Zelltheilung. Morph Jb 10: 214– 330. Rabl C. 1885. U Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243–247. Reichenzeller M, Burzlaff A, Lichter P, Herrmann H. 2000. In vivo observation of a nuclear channel-like system: evidence for a distinct interchromosomal domain compartment in interphase cells. J Struct Biol 129: 175 –185.

Richter K, Reichenzeller M, Gorisch SM, Schmidt U, Scheuermann MO, Herrmann H, Lichter P. 2005. Characterization of a nuclear compartment shared by nuclear bodies applying ectopic protein expression and correlative light and electron microscopy. Exp Cell Res 303: 128 –137. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287– 291. Ronneberger O, Baddeley D, Scheipl F, Verveer PJ, Burkhardt H, Cremer C, Fahrmeir L, Cremer T, Joffe B. 2008. Spatial quantitative analysis of fluorescently labeled nuclear structures: Problems, methods, pitfalls. Chromosome Res 16: 523– 562. Rouquette J, Genoud C, Vazquez-Nin GH, Kraus B, Cremer T, Fakan S. 2009. Revealing the high-resolution threedimensional network of chromatin and interchromatin space: A novel electron-microscopic approach to reconstructing nuclear architecture. Chromosome Res 17: 801 –810. Schardin M, Cremer T, Hager HD, Lang M. 1985. Specific staining of human chromosomes in Chinese hamster x man hybrid cell lines demonstrates interphase chromosome territories. Hum Genet 71: 281– 287. Schermelleh L, Carlton PM, Haase S, Shao L, Winoto L, Kner P, Burke B, Cardoso MC, Agard DA, Gustafsson MG, et al. 2008. Subdiffraction multicolor imaging of the nuclear periphery with 3D structured illumination microscopy. Science 320: 1332–1336. Schneider R, Grosschedl R. 2007. Dynamics and interplay of nuclear architecture, genome organization, and gene expression. Genes Dev 21: 3027–3043. Schroeder-Reiter E, Perez-Willard F, Zeile U, Wanner G. 2009. Focused ion beam (FIB) combined with high resolution scanning electron microscopy: A promising tool for 3D analysis of chromosome architecture. J Struct Biol 165: 97– 106. Shaw PJ, Abranches R, Paula Santos A, Beven AF, Stoger E, Wegel E, Gonza´lez-Melendi P. 2002. The architecture of interphase chromosomes and nucleolar transcription sites in plants. J Struct Biol 140: 31–38. Shopland LS, Lynch CR, Peterson KA, Thornton K, Kepper N, Hase J, Stein S, Vincent S, Molloy KR, Kreth G, et al. 2006. Folding and organization of a contiguous chromosome region according to the gene distribution pattern in primary genomic sequence. J Cell Biol 174: 27– 38. Simonis M, de Laat W. 2008. FISH-eyed and genome-wide views on the spatial organisation of gene expression. Biochim Biophys Acta 1783: 2052– 2060. Simonis M, Klous P, Splinter E, Moshkin Y, Willemsen R, de Wit E, van Steensel B, de Laat W. 2006. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nat Genet 38: 1348–1354. Simonis M, Kooren J, de Laat W. 2007. An evaluation of 3C-based methods to capture DNA interactions. Nat Methods 4: 895– 901. Solimando L, Luijsterburg MS, Vecchio L, Vermeulen W, van Driel R, Fakan S. 2009. Spatial organization of nucleotide excision repair proteins after UV-induced DNA damage in the human cell nucleus. J Cell Sci 122: 83– 91.

T. Cremer and M. Cremer Solovei I, Cavallo A, Schermelleh L, Jaunin F, Scasselati C, Cmarko D, Cremer C, Fakan S, Cremer T. 2002. Spatial preservation of nuclear chromatin architecture during three-dimensional fluorescence in situ hybridization (3D-FISH). Exp Cell Res 276: 10–23. Stack SM, Brown DB, Dewey WC. 1977. Visualization of interphase chromosomes. J Cell Sci 26: 281 –299. Solovei I, Kreysing M, Lanctot C, Kosem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell 137: 356– 368. Strasburger E. 1905. Die stofflichen Grundlagen der Vererbung im organischen Reich. Gustav Fischer, Jena, Germany. Sutherland H, Bickmore WA. 2009. Transcription factories: gene expression in unions? Nat Rev Genet 10: 457–466. Taddei A, Van Houwe G, Hediger F, Kalck V, Cubizolles F, Schober H, Gasser SM. 2006. Nuclear pore association confers optimal expression levels for an inducible yeast gene. Nature 441: 774– 778. Tanabe H, Muller S, Neusser M, von Hase J, Calcagno E, Cremer M, Solovei I, Cremer C, Cremer T. 2002. Evolutionary conservation of chromosome territory arrangements in cell nuclei from higher primates. Proc Natl Acad Sci 99: 4424–4429. Telenius H, Pelmear AH, Tunnacliffe A, Carter NP, Behmel A, Ferguson-Smith MA, Nordenskjold M, Pfragner R, Ponder BA. 1992. Cytogenetic analysis by chromosome painting using DOP-PCR amplified flow-sorted chromosomes. Genes Chromosomes Cancer 4: 257 –263. Teller K, Solovei I, Buiting K, Horsthemke B, Cremer T. 2007. Maintenance of imprinting and nuclear architecture in cycling cells. Proc Natl Acad Sci 104: 14970–14975. Thomson I, Gilchrist S, Bickmore WA, Chubb JR. 2004. The radial positioning of chromatin is not inherited through mitosis but is established de novo in early G1. Curr Biol 14: 166 –172. Trentani A, Testillano PS, Risueno MC, Biggiogera M. 2003. Visualization of transcription sites at the electron microscope. Eur J Histochem 47: 195– 200. Van den Engh G, Sachs R, Trask BJ. 1992. Estimating genomic distance from DNA sequence location in cell nuclei by a random walk model. Science 257: 1410– 1412. Verschure PJ, van Der Kraan I, Manders EM, van Driel R. 1999. Spatial relationship between transcription sites and chromosome territories. J Cell Biol 147: 13–24. Visser AE, Aten JA. 1999. Chromosomes as well as chromosomal subdomains constitute distinct units in interphase nuclei. J Cell Sci 112: 3353–3360.

Visser AE, Jaunin F, Fakan S, Aten JA. 2000. High resolution analysis of interphase chromosome domains. J Cell Sci 113: 2585– 2593. Volpi EV, Chevret E, Jones T, Vatcheva R, Williamson J, Beck S, Campbell RD, Goldsworthy M, Powis SH, Ragoussis J, et al. 2000. Large-scale chromatin organization of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei. J Cell Sci 113: 1565–1576. Walter J, Schermelleh L, Cremer M, Tashiro S, Cremer T. 2003. Chromosome order in HeLa cells changes during mitosis and early G1, but is stably maintained during subsequent interphase stages. J Cell Biol 160: 685 –697. Williams RR, Broad S, Sheer D, Ragoussis J. 2002. Subchromosomal positioning of the epidermal differentiation complex (EDC) in keratinocyte and lymphoblast interphase nuclei. Exp Cell Res 272: 163–175. Wischnitzer S. 1973. The submicroscopic morphology of the interphase nucleus. Int Rev Cytol 34: 1– 48. Zeitz MJ, Mukherjee L, Bhattacharya S, Xu J, Berezney R. 2009. A probabilistic model for the arrangement of a subset of human chromosome territories in WI38 Human fibroblasts. J Cell Physiol 221: 120–129. Zhao Z, Tavoosidana G, Sjolinder M, Gondor A, Mariano P, Wang S, Kanduri C, Lezcano M, Sandhu KS, Singh U, et al. 2006. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat Genet 38: 1341–1347. Zink D, Cremer T, Saffrich R, Fischer R, Trendelenburg MF, Ansorge W, Stelzer EH. 1998. Structure and dynamics of human interphase chromosome territories in vivo. Hum Genet 102: 241– 251. Zirbel RM, Mathieu UR, Kurz A, Cremer T, Lichter P. 1993. Evidence for a nuclear compartment of transcription and splicing located at chromosome domain boundaries. Chromosome Res 1: 93–106. Zorn C, Cremer C, Cremer T, Zimmer J. 1979. Unscheduled DNA synthesis after partial UV irradiation of the cell nucleus. Distribution in interphase and metaphase. Exp Cell Res 124: 111 –119. Zorn C, Cremer T, Cremer C, Zimmer J. 1976. Laser UV microirradiation of interphase nuclei and post-treatment with caffeine. A new approach to establish the arrangement of interphase chromosomes. Hum Genet 35: 83–89. Zuckerkandl E, Cavalli G. 2007. Combinatorial epigenetics, “junk DNA”, and the evolution of complex organisms. Gene 390: 232 –242.

Gene Positioning Carmelo Ferrai, Ineˆs Jesus de Castro, Liron Lavitas, Mita Chotalia, and Ana Pombo Genome Function Group, MRC Clinical Sciences Centre, Imperial College School of Medicine, Hammersmith Hospital Campus, Du Cane Road, London W12 0NN, United Kingdom Correspondence: [email protected]

Eukaryotic gene expression is an intricate multistep process, regulated within the cell nucleus through the activation or repression of RNA synthesis, processing, cytoplasmic export, and translation into protein. The major regulators of gene expression are chromatin remodeling and transcription machineries that are locally recruited to genes. However, enzymatic activities that act on genes are not ubiquitously distributed throughout the nucleoplasm, but limited to specific and spatially defined foci that promote preferred higher-order chromatin arrangements. The positioning of genes within the nuclear landscape relative to specific functional landmarks plays an important role in gene regulation and disease.

egulation of gene expression is vital to establishing the metabolic activities in every cell in all organisms, and allows their adaptation to both stress and developmental processes. In humans, more than 20,000 genes and at least an equal number of noncoding transcription units are distributed among the 22 pairs of autosomes and two sex chromosomes. The physical organization of chromosomes within the nucleus occurs in a hierarchical manner. At the most basic level of organization, DNAwinds around octamers of histone molecules leading to the formation of nucleosomes and adjacent nucleosomes are linked via DNA. Nucleosome arrays have an inherent propensity to form compacted, filamentous structures with the extent of condensation proposed to have regulatory potential. Condensed nucleosome arrays form fibers with a diameter of approximately 30 nm (Dorigo et al. 2004), a conformation in which the accessibility of DNA may be

R

restricted. Chromatin condensation can result in masking of regulatory sequences thereby affecting transcription (Cairns 2005). Decondensed, open chromatin is thought to be more accessible to factors necessary for the activation and transcription of genes (Cosma 2002; Sproul et al. 2005), and is also segregated toward the nuclear interior in most cell types (Solovei et al. 2009). Important processes that affect the condensation status of chromatin include the covalent modification of histone tails within nucleosomes and the repositioning of nucleosomes along the DNA fiber (Cosma 2002). At higher levels of chromatin organization, the three-dimensional arrangement of genes in the nuclear space also influences gene expression. Radial positioning of genes mostly correlates with states of expression, with active genes often located in the interior and silent genes at the periphery (Takizawa et al. 2008a; Takizawa et al. 2008b). However, radial

Editors: David L. Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000588 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000588

C. Ferrai et al.

positioning of genes is unlikely to be a major determinant of gene expression per se, as chromatin regulator complexes do not display radial gradients of abundance, but are instead homogenously distributed or preferentially localized in nuclear subcompartments. The association of inactive genes with the nuclear periphery, which is enriched for transcriptionally silent heterochromatin in most cell types, suggests a molecular mechanism for gene repression. For example, the human CFTR gene associates with peripheral heterochromatin in cell types where the locus is silent, but occupies more internal positions in cell types where CFTR is expressed (Zink et al. 2004). In contrast, the highly conserved murine CFTR homologue and adjacent genes are more centrally positioned irrespective of their expression (Sadoni et al. 2008), possibly because of chromosomal arrangement constraints outside of the Cftr locus. A correlation between gene silencing and positioning at the nuclear periphery has also been shown for several differentiationspecific genes in various cell systems (Brown et al. 1999; Skok et al. 2001; Kosak et al. 2002; Kim et al. 2004; Williams et al. 2006). More recently, the silencing effect of direct associations to the nuclear lamina have been investigated by artificial tethering of reporter genes (Andrulis et al. 1998; Finlan et al. 2008; Reddy et al. 2008). Many genes are inactivated in response to an induced association to the lamina, despite other genes remaining active; these conflicting responses suggest that individual genes have differential sensitivity to the silencing mechanisms at the periphery. Gene neighborhoods constitute a major determinant of gene positioning as each gene is not free to establish its preferred position, but responds to the properties of neighboring genes in the same chromosome. Chromosomes occupy discrete territories within the interphase nucleus in preferred, yet nondeterministic positions that correlate with cell type-specific patterns of gene expression (Cremer et al. 1982; Parada and Misteli 2002). Painting of whole chromosomes in human activated lymphocytes, at high-resolution using a fluorescence in situ hybridization (FISH) approach on

ultrathin cryosections, shows that 20% of the nuclear volume contains intermingled chromatin from neighboring chromosomes, and that a specific inducible locus (MHCII) can be positioned within other CTs (Branco and Pombo 2006). The close spatial proximity between chromosomes has cell type specificity and correlates with increased translocation probability (Roix et al. 2003; Kuroda et al. 2004; Osborne et al. 2007). Chromosome positioning depends on the physical constraints of packaging large chromatin polymers within a small nuclear volume, and is influenced by cell-type specific gene expression programs. In spherical nuclei, such as lymphocytes, the radial positioning of chromosomes correlates with their gene density, with gene-dense and gene-poor chromosomes positioned centrally or at the periphery, respectively (Boyle et al. 2001; Branco et al. 2008). In cells with flat nonspherical nuclei, such as fibroblasts, the size of the chromosome correlates with the radial position, with smaller chromosomes occupying central positions of the nucleus and larger chromosomes positioned toward the periphery independently of gene density (Bolzer et al. 2005). Gene expression is generally controlled by the combinatorial binding of a multitude of transcription factors to specific DNA sequences, which can be located either immediately upstream of protein coding sequences (at promoters) or at considerable distances, upstream or downstream, from the gene. For example, the locus-control region (LCR) at the murine b-globin cluster is located 60 kb upstream of the adult bmajor globin gene (Hbb1) (Dean 2006). Activating transcription factors act either by promoting interactions between transcription start sites and RNA polymerase (RNAP) enzymes or associated transcription machineries, or by recruiting chromatin modifiers that establish local open chromatin states, which are permissive for transcription (Szutorisz et al. 2005). The complex three-dimensional network of chromatin interactions likely reflects the transcriptional program of the cell, in which genes assume “on” or “off ” states according to their local chromatin associations and to their

Gene Positioning

positioning within specific nuclear neighborhoods with activating or silencing features, respectively (Fig. 1) (Dillon 2006; Misteli 2007). GENE POSITIONING AT THE LOCAL LEVEL: CHROMATIN INTERACTIONS BETWEEN REGULATORY ELEMENTS

The transcriptional activation of genes is often described by the simple binding of transcription factors to specific regulatory sequences at variable distances from transcription start sites, followed by the recruitment of RNAP and associated regulatory factors. Novel methodological approaches have enabled the identification of long-range chromatin interactions between regulatory elements and the genes they regulate across the genome, thereby permitting the study of gene regulation in three dimensions. A major breakthrough in this area was the development of the chromosome conformation capture (3C) method, a molecular approach first used to study the three-dimensional folding of Saccharomyces cerevisiae chromosome III (Dekker et al. 2002). In the 3C technique, chromatin is extracted from whole nuclei after chemical cross-linking of DNA-protein complexes and digestion with restriction enzymes or sonication. Intermolecular DNA ligation at low DNA concentration promotes ligation between protein-crosslinked DNA fragments because of their local proximity (Dekker et al. 2002). Interaction partners can then be identified using primers. 3C-based technologies have been extensively used to examine the mechanism of gene regulation at the b-globin gene cluster where the LCR and additional hypersensitive sites form an active chromatin hub with the Hbb1 promoter in a tissue-specific manner (Tolhuis et al. 2002). The conformation of the active hub changes throughout erythrocyte development, in agreement with gene activity at the locus (Palstra et al. 2003), and is dependent on specific regulatory factors (Drissen et al. 2004; Vakoc et al. 2005; Splinter et al. 2006). These findings illustrate that intrachromosomal associations, driven by specific interactions between chromatin binding factors, are an

additional feature of gene regulation. 3C-based approaches have also been widely used to show long-range chromatin interactions at other loci, specifically between distal enhancers and gene promoters (Eivazova and Aune 2004; Spilianakis and Flavell 2004; Liu and Garrard 2005; Amano et al. 2009), but also between promoter and terminator sequences of the same gene, dependent on RNAPII (O’Sullivan et al. 2004; Tan-Wong et al. 2009). Chromatin looping has also been observed at silencers, insulators and boundary elements, which are thought to limit the interaction of enhancers to their target gene promoters. For example, differentially methylated regions (DMRs), involved in the control of parental specific gene expression, establish specific loop structures that may partition differentially expressed genes into distinct loops of active versus silent domains (Murrell et al. 2004). Cell-type specific chromatin interactions mediated by chromatin-associated proteins such as Polycomb repressor complexes, SATB1 and 2, CTCF, and cohesins, have also been identified. For example, cohesin and CTCF binding sites at the developmentally regulated cytokine locus IFNg, in human TH1 lymphocytes, correlate with chromatin interactions in cis, although CTCF is not sufficient to maintain these interactions in the absence of cohesin (Hadjur et al. 2009). GENE POSITIONING AT HIGHER LEVEL: CHROMATIN INTERACTIONS BETWEEN DISTAL GENES

Chromatin interactions between functionally related genes on the same or different chromosomes have also been described by both 3Cbased approaches and by single-cell imaging using FISH. The most extensively studied example is that of the murine b-globin gene, Hbb1, which has been shown to interact with other erythroid-specific genes, Eraf and Uros that are located approximately 40 Mb downstream of Hbb1 on chromosome 7 (Osborne et al. 2004; Simonis et al. 2006). Although it remains unclear whether such interactions are necessary for the correct temporal onset of erythroid

C. Ferrai et al. PcG bodies

Splicing speckles

Centromeric clusters

Cajal bodies

RNAPII factories

Chromosome intermingling Nucleolus

Chromosome territories

Nuclear lamina

PML bodies

Gene

Cajal bodies

PML bodies

RNAPII factories

PcG bodies

Nuclear lamina

Splicing speckles

Centromeric clusters

Nuclear pore

Figure 1. Nuclear subcompartments. An illustration of the mammalian cell nucleus showing nuclear domains and features of gene positioning (orange bar) that have been identified thus far. The nuclear positioning of genes and their associations with different nuclear landmarks are implicated in gene activation and gene repression as discussed in the text. DNA counterstain in blue where applicable. Centromeric clusters. Human CD2 transgenes (green) colocalize with a centromeric cluster (red) in nonexpressing T cells from hCD2 transgenic mice (line 1.3A14). Reprinted by permission from (Hiragami-Hamada et al. 2009).

Gene Positioning

transcription, they appear to involve transcription factories (Osborne et al. 2004; Schoenfelder et al. 2010) or the larger splicing speckles (Brown et al. 2006; Brown et al. 2008; Hu et al. 2009). Both compartments are associated with phosphorylated forms of RNAPII (Xie et al. 2006), but transcription factories are active sites of transcription containing many active RNAPs and associated nascent transcripts (Iborra et al. 1996; Jackson et al. 1998; Pombo et al. 1999), whereas splicing speckles are enriched for a pool of splicing factors and RNAPII complexes that remain stably associated upon transcriptional inhibition (Hall et al. 2006; Xie et al. 2006; Lawrence and Clemson 2008). Association of speckles with highly active genes or with genes that have complex splicing reactions (Xing et al. 1995; Jolly et al. 1999; Smith et al. 1999; Johnson et al. 2000; Shopland et al. 2002; Shopland et al. 2003; Hall et al. 2006; Smith et al. 2007) suggests that gene proximity to splicing speckles may increase the efficiency of RNA splicing and thereby influence the levels of gene expression. Other interchromosomal associations between genes, which correlate with either gene activation or repression, have been described in many different systems (Harmon and Sedat 2005; Spilianakis et al. 2005; Bacher et al. 2006; Ling et al. 2006; Lomvardas et al. 2006; Xu et al. 2006; Apostolou and Thanos 2008; Noordermeer et al. 2008; Schoenfelder et al. 2010). The initial 3C technique revealed many

interesting long-range interactions but was limited to the analyses of a small number of sequences requiring prior knowledge of both interacting baits. Developments of the 3C technique, including circular chromosomal conformation capture (Zhao et al. 2006) and chromosome conformation capture-on-chip (Simonis et al. 2006) (both called 4C), use inverse PCR primers to amplify circular DNA obtained by the ligation of two proximal segments of chromatin. These extended applications of 3C technology enable the simultaneous identification of many interactions using single known baits. Unbiased genome-wide strategies have also emerged. In yeast high-throughput sequencing of 3C-ligation products and bioinformatic identification of ligated fragments has revealed novel chromatin arrangements (Rodley et al. 2009). Two independent approaches in mammalian systems, Hi-C (Lieberman-Aiden et al. 2009) and chromatin interaction analysis by paired-end tag sequencing (ChIA-PET) (Fullwood et al. 2009) have allowed unbiased identification of ligation products by tagging the ligation junction and then enriching the region using affinity purification. In contrast to Hi-C, the ChIA-PET approach enriches 3C products associated with a specific protein of interest thereby identifying protein-based interchromatin networks. One inherent limitation of the 3C-derived approaches is its population-based analysis

Figure 1. (Continued ) Splicing speckles. The C3/C4 genomic region of murine chromosome 8 (red) associates

with splicing speckles (green) in fetal liver cells. Reprinted by permission from (Noordermeer et al. 2008). PcG bodies. Bxd gene (red) positioned at PcG bodies (green) in the posterior part of wild-type stage 5-8 hour Drosophila embryos examined by 2D-FISH. Reprinted by permission from Macmillan Publishers Ltd: Nature Cell Biology (Lanzuolo et al. 2007), copyright (2007). Chromosome intermingling. Intermingling between chromosomes 5 (green) and 7 (red) in human lymphocytes. Reproduced with permission from (Branco and Pombo 2006). Chromosome territories. uPA locus (green) is positioned inside of its chromosome territory (CT; chromosome 10, red) in HepG2 cells. Reproduced with permission from (Ferrai et al. 2010). Nuclear lamina. Igh locus (green) interaction with the nuclear lamina (LMNB1, red) in NIH3T3. Reprinted by permission from Macmillan Publishers Ltd: Nature (Reddy et al. 2008), copyright (2008). PML bodies. MHCII region (red) is proximal to PML bodies (green) in control fibroblasts. Reproduced with permission from (Shiels et al. 2001). Cajal bodies. Simultaneous hybridization of RNU1, RNU2, RNU3, RNU7, RNU12, HIST1, and HIST2 loci show several simultaneous gene interactions with Cajal bodies (red). Reprinted from (Frey et al. 1999). Copyright (1999), with permission from Elsevier. RNAPII factories. Association of hCD2 transgenes with RNAPII-S2p in T cells from hCD2 transgenic mouse line 1.3B (SQ Xie, R Festenstein, A Pombo, unpubl.).

C. Ferrai et al.

that gives rise to average maps of chromatin interactions unlikely to exist in an individual cell. High resolution analyses of physical proximity between 3C partners using cryo-FISH (Simonis et al. 2006) or confocal microscopy (Lieberman-Aiden et al. 2009; Schoenfelder et al. 2010) show remarkably low frequencies of association at distances below 500 nm, which are not consistent with close molecular interactions. Chromatin associations detected by 3C may not always be directly related to regulatory interactions, but may be inherent to chromosome organization and/or promoted by independent coassociation to large nuclear structures. Cytological approaches that measure the frequency of locus-specific interactions at the single cell level remain an important complement to molecular 3C approaches, albeit limited to the analyses of few sequences. GENE POSITIONING RELATIVE TO SITES OF TRANSCRIPTION: TRANSCRIPTION FACTORIES

Different classes of genes are transcribed by the three nuclear RNAPs type I, II, or III: RNAPI transcribes the 45S ribosomal RNA genes tandemly repeated in the genome, RNAPII transcribes messenger RNAs and small structural RNAs, and RNAPIII transcribes the 5S rRNA, transfer RNA and other small structural RNA genes. Visualization of transcriptional activity after pulse-labeling of tagged nucleotides showed that genes are active in distinct subnuclear domains termed transcription factories (Jackson et al. 1993; Wansink et al. 1993). The limited number of transcription sites per nucleus as compared with the larger number of active transcription units suggests that many active RNAPs and associated transcription units are present in each transcription factory (Iborra et al. 1996; Jackson et al. 1998; Kimura et al. 1999; Pombo et al. 1999). Evidence that most active genes are associated with only one or a few active RNAPs at any given time led to the idea that each transcription factory simultaneously transcribes more than one gene (Cook 1999). Support for this model

initially came from observations that Hbb1, Eraf, and Uros genes, separated by several Mb, coassociate in a shared factory when transcribed (Osborne et al. 2004). A recent analysis of a larger number of genes in the same cell type shows proximity between Hbb1 and additional genes separated by large genomic regions within the same chromosome and between genes from different chromosomes, albeit specific associations are not required for transcriptional activity of the genes analyzed (Schoenfelder et al. 2010). Other studies also support the functional specialization of transcription factories, depending on the type of RNAP (Pombo et al. 1999), gene structure (Xu and Cook 2008) (Schoenfelder et al. 2010), or activation state of RNAPII (Ferrai et al. 2010).

GENE POSITIONING RELATIVE TO NUCLEOPLASMIC SUBCOMPARTMENTS Splicing Speckles

Splicing speckles are major subnuclear compartments characterized by an accumulation of components of the splicing machinery, polyAþ RNA and numerous mRNA metabolic factors, as well as RNAPII (Hall et al. 2006; Xie et al. 2006; Lawrence and Clemson 2008). Several splicing speckle domains (10 – 30) are typically present in each nucleus, measuring 0.5 – 3 mm in diameter and occupying 5% –10% of the nucleoplasmic volume (Spector 1993; Fay et al. 1997; Xie et al. 2006). Subchromosomal R bands, containing 210 Mb blocks of gene-rich genomic sequence, are frequently found associated with speckles (Shopland et al. 2003). A subset of genes such as heat-shock genes, erythroid specific genes, collagen genes, and muscle specific genes preferentially associate with SC35 domains when active (Xing et al. 1995; Jolly et al. 1999; Smith et al. 1999; Johnson et al. 2000; Shopland et al. 2002; Shopland et al. 2003; Hall et al. 2006; Smith et al. 2007). In vivo studies using multicopy BAC transgenes, containing the inducible Hsp70 gene tagged with Lac operator sequences and visualized on GFP-LacI expression, show increased association of the transgene with

Gene Positioning

splicing speckles within 30 min of heat shock activation (Hu et al. 2009). Coassociation of genes that share the same pathway has been observed at speckles in skeletal muscle and in erythroid cells. The musclespecific genes, myf-4, MyoD, and cMyHC, important for skeletal muscle differentiation, localize to SC35 domains in differentiated muscle cells, but not in the undifferentiated satellite myoblasts or fibroblasts (Moen et al. 2004). Erythroid genes, Hbb, Hba, Eraf, Slc4a1, and Gata1, located on four different murine chromosomes, have also been found nonrandomly positioned relative to common splicing speckles (Brown et al. 2008). Importantly, the association with speckles of genes that have complex splicing reactions and the accumulation of unspliced RNAs at these sites (Johnson et al. 2000; Smith et al. 2007) suggests that gene associations with speckles may influence the yield of mature mRNAs. Cajal Bodies and PML Bodies

Cajal bodies are nucleoplasmic structures, typically one to six per nucleus, with diameters of 0.1 – 1 mm (Gall 2003). They are involved in the biogenesis of several classes of spliceosomal small nuclear ribonucleoprotein particles (snRNPs) and are enriched in RNAP subunits, specific basal transcription factors, and RNA processing factors. U2 snRNA genes associate within Cajal bodies in a mechanism that depends on the U2 gene promoter and is thought to be important for the maturation of newly synthesized U2 snRNAs (Frey et al. 1999; Smith and Lawrence 2000; Frey and Matera 2001). In vivo imaging of U2 snRNA arrays showed that they only associate with Cajal bodies upon induction. The mechanism involves long-range movements toward relatively stably positioned Cajal bodies and is thought to be mediated by actin polymerization (Dundr et al. 2007). Cajal bodies contain the U7 snRNP, a key component of the histone mRNA 30 -end formation machinery (Frey and Matera 1995). Although replication-dependent histone genes that are cell-cycle regulated, including HIST1

and HIST2 multigene clusters, associate with Cajal bodies, their transcriptional activity is not dependent on this association, which appears to be mediated by the nascent transcripts (Jacobs et al. 1999; Schul et al. 1999; Liu et al. 2006; Stanek and Neugebauer 2006). In contrast, cell cycle-independent histone genes, such as H3.3A, which are constitutively active and produce polyadenylated transcripts, do not associate with Cajal bodies (Shopland et al. 2001). Promyelocytic leukemia nuclear bodies (PML) are nuclear multiprotein domains, (Five to 30 per nucleus) with diameters between 0.1 – 1.0 mm that can vary in relation to the cell cycle stage and cell type (Bernardi and Pandolfi 2007). PML bodies contain many proteins and have been implicated in multiple regulatory pathways, from the control of cell proliferation to DNA repair or apoptosis, and in viral infection. PML bodies colocalize with the TP53 locus (tumor protein 53 gene) in Jurkat cells, raising the possibility that PML bodies play a role in regulating TP53 transcription (Sun et al. 2003). Interaction with PML bodies has also been detected in living cells (Tsukamoto et al. 2000), but ongoing transcription appeared independent of PML body association (Tsukamoto et al. 2000). Furthermore, the analysis of multiple genomic regions showed that PML bodies associate with genomic regions characterized by high transcriptional activity (Wang et al. 2004). However, detection of active alleles by RNA-FISH revealed that transcriptional activity is independent of gene association with PML bodies, and PML knock-down does not alter transcription levels (Wang et al. 2004), suggesting that expression does not require association with PML bodies. Polycomb Bodies

Polycomb group (PcG) complexes are involved in heritable gene repression by epigenetic chromatin modifications and by regulating the nuclear organization of their target genes (Schuettengruber et al. 2007). In mammalian and Drosophila cells, several PcG proteins have a nonhomogeneous distribution in the

C. Ferrai et al.

nucleoplasm being enriched in discrete foci called PcG bodies (Saurin et al. 1998; Orlando 2003), which can be preferentially localized near centromeric heterochromatin (Saurin et al. 1998). Whole-mount FISH in Drosophila melanogaster embryos showed that Polycomb response elements (PREs) such as Fab-7 or bxd of the inactive homeotic bithorax complex (BX-C) colocalize with PcG bodies in more than 80% of the anterior nuclei (Lanzuolo et al. 2007). In the posterior part of the embryo, inactive PREs (bxd) remain predominately bound to PcG bodies, whereas active elements (Fab-7) clearly lose contact with these domains, supporting the idea that gene association with PcG bodies is important for gene silencing. The pairing of two Fab-7 PREs from different genomic regions, as well as the coassociation of three PREs (Fab-7, Mcp, and bxd) within the BX-C Hox gene locus, occur within shared PcG bodies (Grimaud et al. 2006; Lanzuolo et al. 2007). A coassociation of several PcG regulated genes in PcG bodies is also suggested by the existence of fewer PcG bodies than PcG target genes identified by genome-wide chromatin immunoprecipitation (ChIP) (Sexton et al. 2009), further supporting the idea that repositioning of genomic regions with respect to nuclear compartments may be important for the regulation of gene expression. ChIP analysis of a major Polycomb protein, Enhancer of Zeste (Ezh2), combined with 3C showed that Ezh2 is present at intra- and interchromosomal hubs that are associated with gene silencing (Tiwari et al. 2008a; Tiwari et al. 2008b). Centromeric Heterochromatin Regions

Gene associations with centromeric heterochromatin regions also correlate with gene silencing. Brown and colleagues analyzed the nuclear position of several genes (CD2, l5, CD8a, Rag, Sox1 and Hoxb4-b9) relative to centromeric regions in mature B lymphocytes, before and following mitogenic CD40mediated activation (Brown et al. 1999). Genes that are not expressed in mature B cells (l5, CD8a, Rag, Sox1, and Hoxb4-b9) were not associated with centromeric chromatin in resting

mature B cells, but became juxtaposed to these domains in cycling mature B cells after mitogenic stimulation. The CD2 gene, which is widely expressed in mature B lymphocytes, remained distant from centromeric domains irrespectively of stimulation. These findings indicate that some inactive genes become associated with centromeric heterochromatic regions during cell division. Analyses of Rag and TdT genes, expressed in CD4þCD8þ thymocytes but down-regulated on T-cell receptor engagement, showed repositioning of these genes to centromeric regions during induced differentiation, resulting in heritable silencing (Brown et al. 1999). Surprisingly, transient silencing of Rag and TdT in a transformed thymocyte cell line was reversible and was not accompanied by repositioning of these genes to centromeric regions, implicating that silencing is initiated through a more complex mechanism. Genes also reposition to centromeric regions on allelic exclusion during B-cell maturation (Skok et al. 2001). In immature B lymphocytes, IgH and IgL alleles are not associated with centromeric regions, but become associated with these regions upon allelic exclusion which leads to B cells expressing functional receptors encoded by a single IgH allele and one of four IgL alleles. The repositioning of genes to centromeric regions is proposed to be mediated through a DNA binding protein, Ikaros/Lyf-1, which is important for lymphocyte development (Brown et al. 1997; Brown et al. 1999). Nevertheless, transgenes integrated in centromeric regions, are not always silenced (Sabbattini et al. 2001). The mouse l5 and VpreB1 genes are expressed in pro- and pre-B cells but silenced in immature and mature B cells, where they are relocated to centromeric regions. A l5-VpreB1-transgene with a mutated Ikaros binding site was released from Ikaros-mediated silencing (Sabbattini et al. 2001). Interestingly, the integration of the mutated l5-VpreB1transgene into centromeric heterochromatin region does not restore silencing at the mature B-cell stage (Sabbattini et al. 2001), showing that transcription can take place in centromeric regions and Ikaros’ pivotal role in mediating l5-VpreB1-silencing on B-cell differentiation.

Gene Positioning

Nuclear Lamina

The nuclear lamina underlays the inner nuclear membrane and consists of type V intermediate filament proteins called nuclear lamins (Dechat et al. 2008). Markers of gene repression, such as enrichment for hypoacetylated chromatin (Sadoni et al. 1999), depletion of both RNAPII and the open chromatin histone modification H3K4me2 (dimethylation of lysine 4 of histone 3) are associated with the nuclear lamina (Luco et al. 2008). Genome-lamina interactions in human occur through more than 1,300 sharply defined large chromatin domains, referred to as lamina associated domains (LADs) (Guelen et al. 2008). These domains are 0.1 – 10 Mb in size in human fibroblasts, are enriched five- 10-fold for genes with lower expression levels than genes outside LADs and are enriched in repressive histone marks, H3K27me3 (trimethylation of lysine 27 on histone 3) and H3K9me2 (dimethylation of lysine 9 on histone 3), but are depleted of the active histone mark H3K4me2. Dynamic repositioning of some genes to the lamina correlates with changes in gene expression. Activation of genes including Hbb during differentiation of mouse erythroid cells (Ragoczy et al. 2006), IgH and IgK in murine B-cell differentiation (Kosak et al. 2002) or Mash1 during murine neuronal differentiation (Williams et al. 2006), correlates with their separation from the repressive lamina environment. Interestingly, some genes initiate transcription while at the nuclear lamina before internal repositioning (Hbb) (Ragoczy et al. 2006) or remain in close proximity to the nuclear periphery upon activation (IFNg) (Hewitt et al. 2004) indicating that the peripheral gene positioning per se is not restrictive for transcription. Induced tethering of reporter genes to the nuclear lamina has shown in yeast and cultured mammalian cells the relevance of physical gene associations with the nuclear periphery for transcriptional activity (Andrulis et al. 1998; Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). Although some genes were repressed by the interaction with the nuclear periphery (Andrulis et al. 1998; Finlan et al.

2008; Reddy et al. 2008), others remained fully transcriptionally competent (Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). Finlan and colleagues showed that the mechanisms of repression at the nuclear periphery could be alleviated by inhibition of histone deacetylase (HDAC) activity without affecting gene association with the lamina (Finlan et al. 2008), indicating the importance of HDACs in promoting gene repression at the lamina. In all three mammalian studies, transgene repositioning required the passage through mitosis (Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). In summary, gene repression at the nuclear lamina is therefore not a general phenomenon but gene specific, depending on multiple parameters such as transcription factor accessibility, promoter strength, existence of insulator elements and pre-existing chromatin marks, which may counteract the mechanisms of transcription repression at the nuclear lamina. Gene silencing mechanisms that correlate with chromatin association to the nuclear periphery have also been observed in lower eukaryotes (Akhtar and Gasser 2007; Towbin et al. 2009). In yeast nuclei, which do not have a nuclear lamina, repressive proteins such as Sir proteins (Sir2, Sir3, and Sir4) are known to interact with subtelomeric heterochromatic regions and such interactions are mostly confined to foci localized at the nuclear envelope. Transcription-dependent associations of specific groups of genes with nuclear pores have also been described from yeast to mammals, suggesting that the nuclear periphery promotes distinct functional environments compatible with both gene repression and activation. CHROMATIN DYNAMICS

Live cell imaging in interphase nuclei shows that chromatin movements occur within a radius of 0.5 – 1.0 mm, which given the large size of the mammalian nucleus (10 mm diameter) suggests that a given locus is constrained to a very small fraction of the total nuclear volume (Abney et al. 1997; Chubb et al. 2002; Walter et al. 2003). Larger chromatin movements

C. Ferrai et al.

(.2 mm) were observed in early G1 phase of the cell cycle, suggesting that chromatin repositioning occurs during the re-establishment of interphase nuclear architecture (Walter et al. 2003). The importance and complexity of chromatin mobility in gene expression is highlighted in examples of gene looping out of their own chromosome territories (CTs) upon gene induction, such as observed for MHCII, EDC and Hox clusters and for the uPA gene (Volpi et al. 2000; Chambeyron and Bickmore 2004; Williams et al. 2006; Ferrai et al. 2010). The repositioning of Hox clusters B and D relative to their CTs and the condensation level of clusters have been studied during retinoic acid-induced differentiation of murine embryonic stem (ES) cells and in embryos (Chambeyron and Bickmore 2004; Chambeyron et al. 2005; Morey et al. 2007). Hox clusters extrude outside their territories and decondense concomitantly with expression. However, locus decondensation occurs independently of locus position relative to its CT and the HoxD locus is preferentially positioned at the CT interior in limb bud embryo cells (embryonic day 9.5) where it is active (Morey et al. 2007), suggesting that locus positioning relative to the CTmay not always be important for gene expression upon induction. RNA-FISH experiments have shown that in the induced state active alleles locate inside, at the edge, and outside of the CT, with a preferential positioning of inactive alleles within the CT (Morey et al. 2009). A more recent analysis of the inducible uPA gene in human HepG2 cells shows that it relocates out of the CT on activation with phorbol esters within 3 hours of treatment (Ferrai et al. 2010). In the active state, uPA gene activity is independent of its position in relation to its CT. Unexpectedly, before tetradecanoyl phorbol acetate (TPA) induction of the gene, the majority of uninduced uPA genes positioned inside the CT are silenced, whereas a small proportion of uPA loci located outside of the CT are more likely to be transcribed. Collectively, these studies suggest that locus positioning inside the CT correlates in some situations with gene silencing, despite the presence of active RNAPII and transcription occurring throughout the volume

of CTs (Abranches et al. 1998; Verschure et al. 1999; Verschure et al. 2003; Branco and Pombo 2006; Ferrai et al. 2010). ALTERED GENE POSITIONING IN DISEASE

Nuclear architecture and chromatin structure are important features for the regulation of gene expression, and can be altered in diseases. Unbalanced levels or mutations of lamin proteins or emerin, an integral protein of the nuclear envelope, are linked to a group of rare genetic disorders called nuclear envelopathies (Webster et al. 2009). Well known nuclear envelopathies such as Hutchinson-Gilford progeria syndrome (HGPS; premature aging) and Emery-Dreifuss muscular dystrophy (EDMD) are associated with mutations of the lamin A/C or emerin genes and are often characterized by alterations in nuclear morphology that affect chromatin organization and may compromise gene expression (Fig. 2A) (Webster et al. 2009). Studies in primary fibroblasts of several nuclear envelopathies (including HGPS and EDMD) show the repositioning of chromosomes 13 and 18 toward the nuclear interior (Fig. 2B) (Meaburn et al. 2007). Repositioning of chromosome 18 to the nuclear interior and up-regulation of genes within this chromosome are also observed after lamin B1 knockout in mouse embryonic fibroblasts (Malhas et al. 2007). Aberrant chromosome distribution in envelopathies only affects specific tissues and cell lineages (Meaburn et al. 2007), as it is not observed in cell types such as cultured lymphoblastoid cells derived from patients with emerin and lamin A mutations (Boyle et al. 2001; Meaburn et al. 2005). Repositioning of genes, small chromatin regions, or CTs is also seen in other diseases, such as facioscapulohumeral muscular dystrophy (FSHD) (Masny et al. 2004), immunodeficiency centromeric instability facial anomalities (ICF) (Matarazzo et al. 2007), epilepsy (Borden and Manuelidis 1988), or in male factor infertility (Finch et al. 2008). ICF syndrome is caused by mutations in DNA methyltransferase 3B (DNMT3B) and is associated with DNA

Gene Positioning

A

Abnormal nuclei in progeria

B

Chromosome repositioning in progeria

C

Gene repositioning in ICF

D

Gene repositioning in cancer

Figure 2. Nuclear organization defects in disease. (A) Abnormal nuclear shape in Hutchinson-Gilford Progeria

Syndrome. Nuclei in patients with progeria show an abnormal morphology (right) contrasting with the regular nuclei shape found in healthy individuals (left). Reproduced with permission from (Scaffidi et al. 2005). (B) Chromosome repositioning in Hutchinson-Gilford Progeria Syndrome. Interior repositioning of human chromosome 13 (green, arrows) in a HGPS fibroblast cell line (right), in comparison to a control cell line (left). Proliferation marker pKi-67 (red). Reproduced with permission from (Meaburn et al. 2007). (C ) Gene repositioning in immunodeficiency centromeric instability facial anomalies (ICF). In control male cells (left), the SYBL1 (red, arrows) inactive allele (Y chromosome) is localized inside or at the edge of the Y CT, whereas in ICF cells (right) it loops out of its CT and escapes silencing. Reproduced with permission from (Matarazzo et al. 2007). Copyright (2007) National Academy of Sciences. (D) Gene repositioning in early mammary tumorigenesis. Position of the cancer AKT1 locus (red, arrows) before (left) and after (right) oncogenic activation of an epithelial cell line. After activation there is a radial shift in the locus position toward the periphery. Reproduced with permission from (Meaburn and Misteli 2008).

hypomethylation at specific genomic sites, such as the SYBL1 gene promoter which becomes derepressed. Interestingly, in ICF male and female patients, the SYBL1 inactive allele, located on the Y and inactive X chromosome, respectively, relocates outside of its respective CT, recapitulating the position of the active alleles (Fig. 2C) (Matarazzo et al. 2007). In females, but not in males, both inactive SYBL1 and its silent neighbor SPRY3 loop out from the CT, suggesting that the reorganization extends beyond the derepressed, hypomethylated gene (Matarazzo et al. 2007). Altered chromosome positioning has also been associated with numerous types of cancer

in which altered gene expression patterns lead to malignant transformation of the cell. Chronic leukemias are associated with enlarged nuclei in lymphoid cells (Rozycka et al. 1988). Abnormal relocation of chromosome 18 from the nuclear periphery to the interior has been documented in several types of tumor cell lines, including melanoma-derived line, cervix carcinoma, colon carcinoma, Hodgkin-derived cell line and colon carcinoma metastasis cells (Cremer et al. 2003). A radial shift of chromosome 8 and a significant change in its shape was observed in pancreatic neoplastic tissue as well as a radial shift of centromeric regions of chromosome 17 and HER2 domains in tumor

C. Ferrai et al.

breast tissues (Wiech et al. 2005). Individual gene loci are also repositioned during early tumorigenesis. In vitro induction of tumorigenesis in early breast cancer showed altered positioning of a set of cancer-associated genes such as AKT1, BCL2, ERB2, and VEGF loci although no correlation was found between radial redistribution and gene activity level (Fig. 2D; (Meaburn and Misteli 2008). The altered positioning of genes has recently been analyzed in invasive breast cancers toward devising prognosis markers of cancer. Eight genes (HES5, ERBB2, MYC, FOSL2, HSP90AA1, AKT1, TGFB3, and CSF1R) were robustly repositioned in breast cancer affected patients in comparison to normal tissues (Meaburn et al. 2009). The origin of chromosomal rearrangements has long been assumed to be a random process, in which a DNA break in one chromosome results in stochastic joining with a DNA segment of another chromosome by defective DNA repair. However, recurrent translocation partners in specific types of cancer have hinted for a contribution of proximal nuclear positioning of the loci involved. Chromosomes that are known to preferentially translocate in specific cell types are more often associated in the nucleus (Roix et al. 2003; Brianna Caddle et al. 2007). Furthermore, the extent of intermingling between specific pairs of chromosomes correlates with the frequency of translocations between the same chromosomes observed after ionizing radiation (Branco and Pombo 2006). Analyses of single genes close to or at translocation breakpoints have also shown that their nuclear positioning is more proximal in cell types where the translocation is observed (Neves et al. 1999; Nikiforova et al. 2000; Roix et al. 2003; Kuroda et al. 2004; Osborne et al. 2007). More recently, studies on anaplastic large cell lymphoma, often associated with t(2;5) translocations, and prostate cancer, with frequent translocations between TMPRSS2 and ERG or ETV1, have shown a direct correlation between increased gene proximity upon transcription induction and translocation (Lin et al. 2009; Mani et al. 2009; Mathas et al. 2009). Considered together, these observations strongly support the notion that physical

proximity of genomic regions, influenced by states of gene activity and cell-type specific genome architecture, can predispose translocations that are characteristic of specific cell types and cancers. CONCLUDING REMARKS

The absence of “membrane-enclosed” subcompartments in the eukaryotic cell nucleus led to early notions that molecules and processes were randomly distributed within this cellular compartment. Work in past decades has shown many features of the structural and functional organization of the eukaryotic cell nucleus. Most nuclear events do not occur throughout the nucleoplasm, but are limited to specific and spatially defined sites (Cook 1999; Misteli 2007). The nuclear volume contains morphological distinct higher-order chromatin domains, such as CTs and condensed heterochromatin, and a number of proteinaceous subcompartments including transcription factories, nuclear lamina, the nucleolus, speckles, and Cajal bodies (Handwerger and Gall 2006). The physically distinct nature of each compartment not only contributes to spatial partitioning of the nuclear space, but also creates distinct functional subdomains within the nucleus. The mechanisms by which spatial positioning of the genome contributes to functional regulation are far from understood, and represent one of the biggest challenges in modern molecular and cell biology. One possibility is that subcompartmentalization of the nucleus influences multiple parameters including transcription factor accessibility, promoter strength, existence of insulator elements and pre-existing chromatin marks, which cumulatively lead to the correct expression signature unique for each cell type. ACKNOWLEDGMENTS

We thank Sheila Q. Xie, Andre´ Mo¨ller and Miguel R. Branco for help researching literature for this article, and the Medical Research Council (UK) for support. C.F. was a recipient of a Wellcome Trust Value in People (VIP) Award.

Gene Positioning

I.J.C. is supported by Fundac¸a˜o para a Cieˆncia e a Tecnologia (Portugal). REFERENCES Abney JR, Cutler B, Fillbach ML, Axelrod D, Scalettar BA. 1997. Chromatin dynamics in interphase nuclei and its implications for nuclear structure. J Cell Biol 137: 1459– 1468. Abranches R, Beven AF, Aragon-Alcaide L, Shaw PJ. 1998. Transcription sites are not correlated with chromosome territories in wheat nuclei. J Cell Biol 143: 5–12. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Amano T, Sagai T, Tanabe H, Mizushina Y, Nakazawa H, Shiroishi T. 2009. Chromosomal dynamics at the Shh locus: Limb bud-specific differential regulation of competence and active transcription. Dev Cell 16: 47–57. Andrulis ED, Neiman AM, Zappulla DC, Sternglanz R. 1998. Perinuclear localization of chromatin facilitates transcriptional silencing. Nature 394: 592 –595. Apostolou E, Thanos D. 2008. Virus Infection Induces NF-kB-dependent interchromosomal associations mediating monoallelic IFN-bgene expression. Cell 134: 85–96. Bacher CP, Guggiari M, Brors B, Augui S, Clerc P, Avner P, Eils R, Heard E. 2006. Transient colocalization of X-inactivation centres accompanies the initiation of X inactivation. Nat Cell Biol 8: 293 –299. Bernardi R, Pandolfi PP. 2007. Structure, dynamics and functions of promyelocytic leukaemia nuclear bodies. Nat Rev Mol Cell Biol 8: 1006–1016. Bolzer A, Kreth G, Solovei I, Koehler D, Saracoglu K, Fauth C, Muller S, Eils R, Cremer C, Speicher MR, et al. 2005. Three-dimensional maps of all chromosomes in human male fibroblast nuclei and prometaphase rosettes. PLoS Biol 3: e157. Borden J, Manuelidis L. 1988. Movement of the X chromosome in epilepsy. Science 242: 1687–1691. Boyle S, Gilchrist S, Bridger JM, Mahy NL, Ellis JA, Bickmore WA. 2001. The spatial organization of human chromosomes within the nuclei of normal and emerinmutant cells. Hum Mol Genet 10: 211– 219. Branco MR, Pombo A. 2006. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol 4: e138. Branco MR, Branco T, Ramirez F, Pombo A. 2008. Changes in chromosome organization during PHA-activation of resting human lymphocytes measured by cryo-FISH. Chromosome Res 16: 413 –426. Brianna Caddle L, Grant JL, Szatkiewicz J, van Hase J, Shirley BJ, Bewersdorf J, Cremer C, Arneodo A, Khalil A, Mills KD. 2007. Chromosome neighborhood composition determines translocation outcomes after exposure to high-dose radiation in primary cells. Chromosome Res 15: 1061–1073. Brown KE, Baxter J, Graf D, Merkenschlager M, Fisher AG. 1999. Dynamic repositioning of genes in the nucleus of

lymphocytes preparing for cell division. Mol Cell 3: 207 –217. Brown JM, Green J, das Neves RP, Wallace HA, Smith AJ, Hughes J, Gray N, Taylor S, Wood WG, Higgs DR, et al. 2008. Association between active genes occurs at nuclear speckles and is modulated by chromatin environment. J Cell Biol 182: 1083–1097. Brown KE, Guest SS, Smale ST, Hahm K, Merkenschlager M, Fisher AG. 1997. Association of transcriptionally silent genes with Ikaros complexes at centromeric heterochromatin. Cell 91: 845–854. Brown JM, Leach J, Reittie JE, Atzberger A, Lee-Prudhoe J, Wood WG, Higgs DR, Iborra FJ, Buckle VJ. 2006. Coregulated human globin genes are frequently in spatial proximity when active. J Cell Biol 172: 177– 187. Cairns BR. 2005. Chromatin remodeling complexes: Strength in diversity, precision through specialization. Curr Opin Genet Dev 15: 185– 190. Chambeyron S, Bickmore WA. 2004. Chromatin decondensation and nuclear reorganization of the HoxB locus upon induction of transcription. Genes Dev 18: 1119–1130. Chambeyron S, Da Silva NR, Lawson KA, Bickmore WA. 2005. Nuclear re-organisation of the Hoxb complex during mouse embryonic development. Development 132: 2215– 2223. Chubb JR, Boyle S, Perry P, Bickmore WA. 2002. Chromatin motion is constrained by association with nuclear compartments in human cells. Curr Biol 12: 439–445. Cook PR. 1999. The organization of replication and transcription. Science 284: 1790–1795. Cosma MP. 2002. Ordered recruitment: Gene-specific mechanism of transcription activation. Mol Cell 10: 227 –236. Cremer T, Cremer C, Baumann H, Luedtke EK, Sperling K, Teuber V, Zorn C. 1982. Rabl’s model of the interphase chromosome arrangement tested in Chinese hamster cells by premature chromosome condensation and laser-UV-microbeam experiments. Hum Genet 60: 46– 56. Cremer M, Kupper K, Wagler B, Wizelman L, von Hase J, Weiland Y, Kreja L, Diebold J, Speicher MR, Cremer T. 2003. Inheritance of gene density-related higher order chromatin arrangements in normal and tumor cell nuclei. J Cell Biol 162: 809 –820. Dean A. 2006. On a chromosome far, far away: LCRs and gene expression. Trends Genet 22: 38–45. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832– 853. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306– 1311. Dillon N. 2006. Gene regulation and large-scale chromatin organization in the nucleus. Chromosome Res 14: 117 –126. Dorigo B, Schalch T, Kulangara A, Duda S, Schroeder RR, Richmond TJ. 2004. Nucleosome arrays reveal the two-start organization of the chromatin fiber. Science 306: 1571– 1573.

C. Ferrai et al. Drissen R, Palstra RJ, Gillemans N, Splinter E, Grosveld F, Philipsen S, de Laat W. 2004. The active spatial organization of the b-globin locus requires the transcription factor EKLF. Genes Dev 18: 2485–2490. Dundr M, Ospina JK, Sung MH, John S, Upender M, Ried T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095– 1103. Eivazova ER, Aune TM. 2004. Dynamic alterations in the conformation of the Ifng gene region during T helper cell differentiation. Proc Natl Acad Sci U S A 101: 251–256. Fay FS, Taneja KL, Shenoy S, Lifshitz L, Singer RH. 1997. Quantitative digital analysis of diffuse and concentrated nuclear distributions of nascent transcripts, SC35 and poly(A). Exp Cell Res 231: 27– 37. Ferrai C, Xie SQ, Luraghi P, Munari D, Ramirez F, Branco MR, Pombo A, Crippa MP. 2010. Poised transcription factories prime silent uPA gene prior to activation. PLoS Biol 8: e1000270. Finch KA, Fonseka KG, Abogrein A, Ioannou D, Handyside AH, Thornhill AR, Hickson N, Griffin DK. 2008. Nuclear organization in human sperm: Preliminary evidence for altered sex chromosome centromere position in infertile males. Hum Reprod 23: 1263–1270. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Frey MR, Matera AG. 1995. Coiled bodies contain U7 small nuclear RNA and associate with specific DNA sequences in interphase human cells. Proc Natl Acad Sci U S A 92: 5915– 5919. Frey MR, Matera AG. 2001. RNA-mediated interaction of Cajal bodies and U2 snRNA genes. J Cell Biol 154: 499–509. Frey MR, Bailey AD, Weiner AM, Matera AG. 1999. Association of snRNA genes with coiled bodies is mediated by nascent snRNA transcripts. Curr Biol 9: 126 –135. Fullwood MJ, Liu MH, Pan YF, Liu J, Xu H, Mohamed YB, Orlov YL, Velkov S, Ho A, Mei PH, et al. 2009. An oestrogen-receptor-a-bound human chromatin interactome. Nature 462: 58–64. Gall JG. 2003. The centennial of the Cajal body. Nat Rev Mol Cell Biol 4: 975–980. Grimaud C, Bantignies F, Pal-Bhadra M, Ghana P, Bhadra U, Cavalli G. 2006. RNAi components are required for nuclear clustering of Polycomb group response elements. Cell 124: 957– 971. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W, et al. 2008. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453: 948 –951. Hadjur S, Williams LM, Ryan NK, Cobb BS, Sexton T, Fraser P, Fisher AG, Merkenschlager M. 2009. Cohesins form chromosomal cis-interactions at the developmentally regulated IFNG locus. Nature 460: 410–413. Hall LL, Smith KP, Byron M, Lawrence JB. 2006. Molecular anatomy of a speckle. Anat Rec A Discov Mol Cell Evol Biol 288: 664– 675.

Handwerger KE, Gall JG. 2006. Subnuclear organelles: New insights into form and function. Trends Cell Biol 16: 19– 26. Harmon B, Sedat J. 2005. Cell-by-cell dissection of gene expression and chromosomal interactions reveals consequences of nuclear reorganization. PLoS Biol 3: e67. Hewitt SL, High FA, Reiner SL, Fisher AG, Merkenschlager M. 2004. Nuclear repositioning marks the selective exclusion of lineage-inappropriate transcription factor loci during T helper cell differentiation. Eur J Immunol 34: 3604–3613. Hiragami-Hamada K, Xie SQ, Saveliev A, Uribe-Lewis S, Pombo A, Festenstein R. 2009. The molecular basis for stability of heterochromatin-mediated silencing in mammals. Epigenetics Chromatin 2: 14. Hu Y, Kireev I, Plutz M, Ashourian N, Belmont AS. 2009. Large-scale chromatin structure of inducible genes: Transcription on a condensed, linear template. J Cell Biol 185: 87– 100. Iborra FJ, Pombo A, Jackson DA, Cook PR. 1996. Active RNA polymerases are localized within discrete transcription “factories” in human nuclei. J Cell Sci 109: 1427–1436. Jackson DA, Hassan AB, Errington RJ, Cook PR. 1993. Visualization of focal sites of transcription within human nuclei. Embo J 12: 1059– 1065. Jackson DA, Iborra FJ, Manders EM, Cook PR. 1998. Numbers and organization of RNA polymerases, nascent transcripts, and transcription units in HeLa nuclei. Mol Biol Cell 9: 1523– 1536. Jacobs EY, Frey MR, Wu W, Ingledue TC, Gebuhr TC, Gao L, Marzluff WF, Matera AG. 1999. Coiled bodies preferentially associate with U4, U11, and U12 small nuclear RNA genes in interphase HeLa cells but not with U6 and U7 genes. Mol Biol Cell 10: 1653– 1663. Johnson C, Primorac D, McKinstry M, McNeil J, Rowe D, Lawrence JB. 2000. Tracking COL1A1 RNA in osteogenesis imperfecta. splice-defective transcripts initiate transport from the gene but are retained within the SC35 domain. J Cell Biol 150: 417– 432. Jolly C, Vourc’h C, Robert-Nicoud M, Morimoto RI. 1999. Intron-independent association of splicing factors with active genes. J Cell Biol 145: 1133– 1143. Kim SH, McQueen PG, Lichtman MK, Shevach EM, Parada LA, Misteli T. 2004. Spatial genome organization during T-cell differentiation. Cytogenet Genome Res 105: 292 –301. Kimura H, Tao Y, Roeder RG, Cook PR. 1999. Quantitation of RNA polymerase II and its transcription factors in an HeLa cell: Little soluble holoenzyme but significant amounts of polymerases attached to the nuclear substructure. Mol Cell Biol 19: 5383– 5392. Kosak ST, Skok JA, Medina KL, Riblet R, Le Beau MM, Fisher AG, Singh H. 2002. Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296: 158 –162. Kumaran RI, Spector DL. 2008. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol 180: 51–65. Kuroda M, Tanabe H, Yoshida K, Oikawa K, Saito A, Kiyuna T, Mizusawa H, Mukai K. 2004. Alteration of

Gene Positioning chromosome positioning during adipocyte differentiation. J Cell Sci 117: 5897– 5903. Lanzuolo C, Roure V, Dekker J, Bantignies F, Orlando V. 2007. Polycomb response elements mediate the formation of chromosome higher-order structures in the bithorax complex. Nat Cell Biol 9: 1167– 1174. Lawrence JB, Clemson CM. 2008. Gene associations: True romance or chance meeting in a nuclear neighborhood? J Cell Biol 182: 1035– 1038. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289 –293. Lin C, Yang L, Tanasa B, Hutt K, Ju BG, Ohgi K, Zhang J, Rose DW, Fu XD, Glass CK, et al. 2009. Nuclear receptorinduced chromosomal proximity and DNA breaks underlie specific translocations in cancer. Cell 139: 1069– 1083. Ling JQ, Li T, Hu JF, Vu TH, Chen HL, Qiu XW, Cherry AM, Hoffman AR. 2006. CTCF mediates interchromosomal colocalization between Igf2/H19 and Wsb1/Nf1. Science 312: 269– 272. Liu Z, Garrard WT. 2005. Long-range interactions between three transcriptional enhancers, active Vk gene promoters, and a 30 boundary sequence spanning 46 kilobases. Mol Cell Biol 25: 3220–3231. Liu JL, Murphy C, Buszczak M, Clatterbuck S, Goodman R, Gall JG. 2006. The Drosophila melanogaster Cajal body. J Cell Biol 172: 875 –884. Lomvardas S, Barnea G, Pisapia DJ, Mendelsohn M, Kirkland J, Axel R. 2006. Interchromosomal interactions and olfactory receptor choice. Cell 126: 403– 413. Luco RF, Maestro MA, Sadoni N, Zink D, Ferrer J. 2008. Targeted deficiency of the transcriptional activator Hnf1a alters subnuclear positioning of its genomic targets. PLoS Genet 4: e1000079. Malhas A, Lee CF, Sanders R, Saunders NJ, Vaux DJ. 2007. Defects in lamin B1 expression or processing affect interphase chromosome position and gene expression. J Cell Biol 176: 593–603. Mani RS, Tomlins SA, Callahan K, Ghosh A, Nyati MK, Varambally S, Palanisamy N, Chinnaiyan AM. 2009. Induced chromosomal proximity and gene fusions in prostate cancer. Science 326: 1230. Masny PS, Bengtsson U, Chung SA, Martin JH, van Engelen B, van der Maarel SM, Winokur ST. 2004. Localization of 4q35.2 to the nuclear periphery: Is FSHD a nuclear envelope disease? Hum Mol Genet 13: 1857–1871. Matarazzo MR, Boyle S, D’Esposito M, Bickmore WA. 2007. Chromosome territory reorganization in a human disease with altered DNA methylation. Proc Natl Acad Sci U S A 104: 16546– 16551. Mathas S, Kreher S, Meaburn KJ, Johrens K, Lamprecht B, Assaf C, Sterry W, Kadin ME, Daibata M, Joos S, et al. 2009. Gene deregulation and spatial genome reorganization near breakpoints prior to formation of translocations in anaplastic large cell lymphoma. Proc Natl Acad Sci U S A 106: 5831–5836.

Meaburn KJ, Misteli T. 2008. Locus-specific and activity-independent gene repositioning during early tumorigenesis. J Cell Biol 180: 39– 50. Meaburn KJ, Cabuy E, Bonne G, Levy N, Morris GE, Novelli G, Kill IR, Bridger JM. 2007. Primary laminopathy fibroblasts display altered genome organization and apoptosis. Aging Cell 6: 139– 153. Meaburn KJ, Gudla PR, Khan S, Lockett SJ, Misteli T. 2009. Disease-specific gene repositioning in breast cancer. J Cell Biol 187: 801 –812. Meaburn KJ, Levy N, Toniolo D, Bridger JM. 2005. Chromosome positioning is largely unaffected in lymphoblastoid cell lines containing emerin or A-type lamin mutations. Biochem Soc Trans 33: 1438–1440. Misteli T. 2007. Beyond the sequence: Cellular organization of genome function. Cell 128: 787– 800. Moen PTJr, Johnson CV, Byron M, Shopland LS, de la Serna IL, Imbalzano AN, Lawrence JB. 2004. Repositioning of muscle-specific genes relative to the periphery of SC-35 domains during skeletal myogenesis. Mol Biol Cell 15: 197 –206. Morey C, Kress C, Bickmore WA. 2009. Lack of bystander activation shows that localization exterior to chromosome territories is not sufficient to up-regulate gene expression. Genome Res 19: 1184– 1194. Morey C, Da Silva NR, Perry P, Bickmore WA. 2007. Nuclear reorganisation and chromatin decondensation are conserved, but distinct, mechanisms linked to Hox gene activation. Development 134: 909 –919. Murrell A, Heeson S, Reik W. 2004. Interaction between differentially methylated regions partitions the imprinted genes Igf2 and H19 into parent-specific chromatin loops. Nat Genet 36: 889 –893. Neves H, Ramos C, da Silva MG, Parreira A, Parreira L. 1999. The nuclear topography of ABL, BCR, PML, and RARa genes: Evidence for gene proximity in specific phases of the cell cycle and stages of hematopoietic differentiation. Blood 93: 1197–1207. Nikiforova MN, Stringer JR, Blough R, Medvedovic M, Fagin JA, Nikiforov YE. 2000. Proximity of chromosomal loci that participate in radiation-induced rearrangements in human cells. Science 290: 138– 141. Noordermeer D, Branco MR, Splinter E, Klous P, van Ijcken W, Swagemakers S, Koutsourakis M, van der Spek P, Pombo A, de Laat W. 2008. Transcription and chromatin organization of a housekeeping gene cluster containing an integrated b-globin locus control region. PLoS Genet 4: e1000016. O’Sullivan JM, Tan-Wong SM, Morillon A, Lee B, Coles J, Mellor J, Proudfoot NJ. 2004. Gene loops juxtapose promoters and terminators in yeast. Nat Genet 36: 1014–1018. Orlando V. 2003. Polycomb, epigenomes, and control of cell identity. Cell 112: 599–606. Osborne CS, Chakalova L, Brown KE, Carter D, Horton A, Debrand E, Goyenechea B, Mitchell JA, Lopes S, Reik W, et al. 2004. Active genes dynamically colocalize to shared sites of ongoing transcription. Nat Genet 36: 1065–1071. Osborne CS, Chakalova L, Mitchell JA, Horton A, Wood AL, Bolland DJ, Corcoran AE, Fraser P. 2007. Myc

C. Ferrai et al. dynamically and preferentially relocates to a transcription factory occupied by Igh. PLoS Biol 5: e192. Palstra RJ, Tolhuis B, Splinter E, Nijmeijer R, Grosveld F, de Laat W. 2003. The b-globin nuclear compartment in development and erythroid differentiation. Nat Genet 35: 190 –194. Parada L, Misteli T. 2002. Chromosome positioning in the interphase nucleus. Trends Cell Biol 12: 425– 432. Pombo A, Jackson DA, Hollinshead M, Wang Z, Roeder RG, Cook PR. 1999. Regional specialization in human nuclei: Visualization of discrete sites of transcription by RNA polymerase III. Embo J 18: 2241– 2253. Ragoczy T, Bender MA, Telling A, Byron R, Groudine M. 2006. The locus control region is required for association of the murine b-globin locus with engaged transcription factories during erythroid maturation. Genes Dev 20: 1447– 1457. Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243–247. Rodley CD, Bertels F, Jones B, O’Sullivan JM. 2009. Global identification of yeast chromosome interactions using Genome conformation capture. Fungal Genet Biol 46: 879–886. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287 –291. Rozycka M, Sawicki W, Traczyk Z, Bem W, Strojny P. 1988. Analysis of chromatin pattern in blood lymphocytes of healthy donors and in lymphoid cells of patients with chronic lymphocytic leukaemia. J Clin Pathol 41: 504–509. Sabbattini P, Lundgren M, Georgiou A, Chow C, Warnes G, Dillon N. 2001. Binding of Ikaros to the lambda5 promoter silences transcription through a mechanism that does not require heterochromatin formation. Embo J 20: 2812–2822. Sadoni N, Langer S, Fauth C, Bernardi G, Cremer T, Turner BM, Zink D. 1999. Nuclear organization of mammalian genomes. Polar chromosome territories build up functionally distinct higher order compartments. J Cell Biol 146: 1211–1226. Sadoni N, Targosz BS, Englmann A, Fesser S, Koch J, Schindelhauer D, Zink D. 2008. Transcription-dependent spatial arrangements of CFTR and conserved adjacent loci are not conserved in human and murine nuclei. Chromosoma 117: 381 –397. Saurin AJ, Shiels C, Williamson J, Satijn DP, Otte AP, Sheer D, Freemont PS. 1998. The human polycomb group complex associates with pericentromeric heterochromatin to form a novel nuclear domain. J Cell Biol 142: 887– 898. Scaffidi P, Gordon L, Misteli T. 2005. The cell nucleus and aging: Tantalizing clues and hopeful promises. PLoS Biol 3: e395. Schoenfelder S, Sexton T, Chakalova L, Cope NF, Horton A, Andrews S, Kurukuti S, Mitchell JA, Umlauf D, Dimitrova DS, et al. 2010. Preferential associations between co-regulated genes reveal a transcriptional interactome in erythroid cells. Nat Genet 42: 53–61.

Schuettengruber B, Chourrout D, Vervoort M, Leblanc B, Cavalli G. 2007. Genome regulation by polycomb and trithorax proteins. Cell 128: 735– 745. Schul W, van Der Kraan I, Matera AG, van Driel R, de Jong L. 1999. Nuclear domains enriched in RNA 30 -processing factors associate with coiled bodies and histone genes in a cell cycle-dependent manner. Mol Biol Cell 10: 3815–3824. Sexton T, Bantignies F, Cavalli G. 2009. Genomic interactions: Chromatin loops and gene meeting points in transcriptional regulation. Semin Cell Dev Biol. 20: 849–55. Shiels C, Islam SA, Vatcheva R, Sasieni P, Sternberg MJ, Freemont PS, Sheer D. 2001. PML bodies associate specifically with the MHC gene cluster in interphase nuclei. J Cell Sci 114: 3705– 3716. Shopland LS, Johnson CV, Lawrence JB. 2002. Evidence that all SC-35 domains contain mRNAs and that transcripts can be structurally constrained within these domains. J Struct Biol 140: 131– 139. Shopland LS, Byron M, Stein JL, Lian JB, Stein GS, Lawrence JB. 2001. Replication-dependent histone gene expression is related to Cajal body (CB) association but does not require sustained CB contact. Mol Biol Cell 12: 565– 576. Shopland LS, Johnson CV, Byron M, McNeil J, Lawrence JB. 2003. Clustering of multiple specific genes and gene-rich R-bands around SC-35 domains: Evidence for local euchromatic neighborhoods. J Cell Biol 162: 981– 990. Simonis M, Klous P, Splinter E, Moshkin Y, Willemsen R, de Wit E, van Steensel B, de Laat W. 2006. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nat Genet 38: 1348–1354. Skok JA, Brown KE, Azuara V, Caparros ML, Baxter J, Takacs K, Dillon N, Gray D, Perry RP, Merkenschlager M, et al. 2001. Nonequivalent nuclear location of immunoglobulin alleles in B lymphocytes. Nat Immunol 2: 848 –854. Smith KP, Lawrence JB. 2000. Interactions of U2 gene loci and their nuclear transcripts with Cajal (coiled) bodies: Evidence for PreU2 within Cajal bodies. Mol Biol Cell 11: 2987–2998. Smith KP, Byron M, Johnson C, Xing Y, Lawrence JB. 2007. Defining early steps in mRNA transport: Mutant mRNA in myotonic dystrophy type I is blocked at entry into SC-35 domains. J Cell Biol 178: 951 –964. Smith KP, Moen PT, Wydner KL, Coleman JR, Lawrence JB. 1999. Processing of endogenous pre-mRNAs in association with SC-35 domains is gene specific. J Cell Biol 144: 617–629. Solovei I, Kreysing M, Lanctot C, Kosem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell 137: 356 –368. Spector DL. 1993. Macromolecular domains within the cell nucleus. Annu Rev Cell Biol 9: 265– 315. Spilianakis CG, Flavell RA. 2004. Long-range intrachromosomal interactions in the T helper type 2 cytokine locus. Nat Immunol 5: 1017–1027. Spilianakis CG, Lalioti MD, Town T, Lee GR, Flavell RA. 2005. Interchromosomal associations between alternatively expressed loci. Nature 435: 637–645.

Gene Positioning Splinter E, Heath H, Kooren J, Palstra RJ, Klous P, Grosveld F, Galjart N, de Laat W. 2006. CTCF mediates long-range chromatin looping and local histone modification in the b-globin locus. Genes Dev 20: 2349– 2354. Sproul D, Gilbert N, Bickmore WA. 2005. The role of chromatin structure in regulating the expression of clustered genes. Nat Rev Genet 6: 775– 781. Stanek D, Neugebauer KM. 2006. The Cajal body: A meeting place for spliceosomal snRNPs in the nuclear maze. Chromosoma 115: 343 –354. Sun Y, Durrin LK, Krontiris TG. 2003. Specific interaction of PML bodies with the TP53 locus in Jurkat interphase nuclei. Genomics 82: 250–252. Szutorisz H, Dillon N, Tora L. 2005. The role of enhancers as centres for general transcription factor recruitment. Trends Biochem Sci 30: 593 –599. Takizawa T, Meaburn KJ, Misteli T. 2008b. The meaning of gene positioning. Cell 135: 9 –13. Takizawa T, Gudla PR, Guo L, Lockett S, Misteli T. 2008a. Allele-specific nuclear positioning of the monoallelically expressed astrocyte marker GFAP. Genes Dev 22: 489–498. Tan-Wong SM, Wijayatilake HD, Proudfoot NJ. 2009. Gene loops function to maintain transcriptional memory through interaction with the nuclear pore complex. Genes Dev 23: 2610–2624. Tiwari VK, Cope L, McGarvey KM, Ohm JE, Baylin SB. 2008a. A novel 6C assay uncovers Polycomb-mediated higher order chromatin conformations. Genome Res 18: 1171– 1179. Tiwari VK, McGarvey KM, Licchesi JD, Ohm JE, Herman JG, Schubeler D, Baylin SB. 2008b. PcG proteins, DNA methylation, and gene repression by chromatin looping. PLoS Biol 6: 2911–2927. Tolhuis B, Palstra RJ, Splinter E, Grosveld F, de Laat W. 2002. Looping and interaction between hypersensitive sites in the active b-globin locus. Mol Cell 10: 1453–1465. Towbin BD, Meister P, Gasser SM. 2009. The nuclear envelope–a scaffold for silencing? Curr Opin Genet Dev 19: 180–186. Tsukamoto T, Hashiguchi N, Janicki SM, Tumbar T, Belmont AS, Spector DL. 2000. Visualization of gene activity in living cells. Nat Cell Biol 2: 871–878. Vakoc CR, Letting DL, Gheldof N, Sawado T, Bender MA, Groudine M, Weiss MJ, Dekker J, Blobel GA. 2005. Proximity among distant regulatory elements at the b-globin locus requires GATA-1 and FOG-1. Mol Cell 17: 453 –462. Verschure PJ, van der Kraan I, Manders EM, Hoogstraten D, Houtsmuller AB, van Driel R. 2003. Condensed chromatin domains in the mammalian nucleus are accessible to large macromolecules. EMBO Rep 4: 861– 866. Verschure PJ, van Der Kraan I, Manders EM, van Driel R. 1999. Spatial relationship between transcription sites and chromosome territories. J Cell Biol 147: 13–24. Volpi EV, Chevret E, Jones T, Vatcheva R, Williamson J, Beck S, Campbell RD, Goldsworthy M, Powis SH, Ragoussis J,

et al. 2000. Large-scale chromatin organization of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei. J Cell Sci 113: 1565–1576. Walter J, Schermelleh L, Cremer M, Tashiro S, Cremer T. 2003. Chromosome order in HeLa cells changes during mitosis and early G1, but is stably maintained during subsequent interphase stages. J Cell Biol 160: 685–697. Wang J, Shiels C, Sasieni P, Wu PJ, Islam SA, Freemont PS, Sheer D. 2004. Promyelocytic leukemia nuclear bodies associate with transcriptionally active genomic regions. J Cell Biol 164: 515– 526. Wansink DG, Schul W, van der Kraan I, van Steensel B, van Driel R, de Jong L. 1993. Fluorescent labeling of nascent RNA reveals transcription by RNA polymerase II in domains scattered throughout the nucleus. J Cell Biol 122: 283–293. Webster M, Witkin KL, Cohen-Fix O. 2009. Sizing up the nucleus: Nuclear shape, size and nuclear-envelope assembly. J Cell Sci 122: 1477– 1486. Wiech T, Timme S, Riede F, Stein S, Schuricke M, Cremer C, Werner M, Hausmann M, Walch A. 2005. Human archival tissues provide a valuable source for the analysis of spatial genome organization. Histochem Cell Biol 123: 229 –238. Williams RR, Azuara V, Perry P, Sauer S, Dvorkina M, Jorgensen H, Roix J, McQueen P, Misteli T, Merkenschlager M, et al. 2006. Neural induction promotes large-scale chromatin reorganisation of the Mash1 locus. J Cell Sci 119: 132–140. Xie SQ, Martin S, Guillot PV, Bentley DL, Pombo A. 2006. Splicing speckles are not reservoirs of RNA polymerase II, but contain an inactive form, phosphorylated on serine2 residues of the C-terminal domain. Mol Biol Cell 17: 1723–1733. Xing Y, Johnson CV, Moen PTJr, McNeil JA, Lawrence J. 1995. Nonrandom gene organization: Structural arrangements of specific pre-mRNA transcription and splicing with SC-35 domains. J Cell Biol 131: 1635– 1647. Xu M, Cook PR. 2008. Similar active genes cluster in specialized transcription factories. J Cell Biol 181: 615 –623. Xu N, Tsai CL, Lee JT. 2006. Transient homologous chromosome pairing marks the onset of X inactivation. Science 311: 1149– 1152. Zhao Z, Tavoosidana G, Sjolinder M, Gondor A, Mariano P, Wang S, Kanduri C, Lezcano M, Sandhu KS, Singh U, et al. 2006. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat Genet 38: 1341–1347. Zink D, Amaral MD, Englmann A, Lang S, Clarke LA, Rudolph C, Alt F, Luther K, Braz C, Sadoni N, et al. 2004. Transcription-dependent spatial arrangements of CFTR and adjacent genes in human cell nuclei. J Cell Biol 166: 815 –825.

Chromatin Higher-order Structure and Dynamics Christopher L. Woodcock1,2 and Rajarshi P. Ghosh2 1

Biology Department, University of Massachusetts, Amherst, Massachusetts 01003

2

Program in Molecular and Cellular Biology, University of Massachusetts, Amherst, Massachusetts 01003

Correspondence: [email protected]

The primary role of the nucleus as an information storage, retrieval, and replication site requires the physical organization and compaction of meters of DNA. Although it has been clear for many years that nucleosomes constitute the first level of chromatin compaction, this contributes a relatively small fraction of the condensation needed to fit the typical genome into an interphase nucleus or set of metaphase chromosomes, indicating that there are additional “higher order” levels of chromatin condensation. Identifying these levels, their interrelationships, and the principles that govern their occurrence has been a challenging and much discussed problem. In this article, we focus on recent experimental advances and the emerging evidence indicating that structural plasticity and chromatin dynamics play dominant roles in genome organization. We also discuss novel approaches likely to yield important insights in the near future, and suggest research areas that merit further study.

s the cell’s primary information storage, retrieval, and duplication organelle, the nucleus, by analogy with human-created information repositories (libraries and computer discs), might be expected to be highly structured. However, chromatin within the living nucleus appears to be organized in a quite unstructured manner, and the rules that define or encode the principles of chromatin organization have been difficult to decipher. These frustrations are reflected in some reviews of chromatin organization with titles such as “Chromatin higher-order structure: chasing a mirage?” (van Holde and Zlatanova 1995), “Higher-order structures of chromatin: the elusive 30 nm fiber” (Tremethick 2007), and “Chromatin fiber structure: where is the problem now?” (van Holde and Zlatanova 2007). In this article, we discuss what is known about chromatin structure, and

A

consider the prospects of improving our understanding in the near future. In recent years, work on chromatin organization has increasingly focused on the many dynamic aspects of chromatin, which contribute to its structural and functional plasticity, and it is becoming clear that dynamics plays a crucially important functional role, perhaps contributing to nuclear selforganization (Misteli 2001). CHROMATIN HIGHER-ORDER STRUCTURE

In the context of chromatin, “higher-order structure” may be defined as any assemblage of nucleosomes that assumes a reproducible conformation in 3D space. The most obvious chromatin higher-order structure is the mitotic/ meiotic chromosome in which the DNA is compacted some 10,000- to 20,000-fold. Metaphase

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000596 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000596

C.L. Woodcock and R.P. Ghosh

chromosomes have characteristic shapes, banding patterns, and locations of specific genes. Although chromosomes have a consistent structure at the light microscope level in terms of longitudinal positioning of bands and genes, this consistency is evidently modulated by an intrinsic variability in longitudinal position of up to 0.3 mm (Strukov and Belmont 2009). However, these authors reported that there was no consistency in the axial positioning of loci, suggesting that chromosome architecture involves considerable plasticity at some level or levels of folding. Understanding how chains of nucleosomes are folded in the creation of mitotic chromosomes and how they are arranged in interphase continues to be an exciting yet technically challenging endeavor. The concept of primary, secondary, tertiary, and quaternary structures used for proteins can also be usefully applied to chromatin structural hierarchies (Woodcock and Dimitrov 2001), with the beads-on-a-string organization of nucleosomes and linker DNA constituting the primary structure, and arrangements resulting from interactions between nucleosomes giving rise to secondary structures. Thus, the chromatin equivalent of protein secondary structure might involve interactions analogous to those leading to a-helices and b-sheets. Unlike proteins that consist of sequences of 20 amino acids, chromatin consists of a repeating chain of more-or-less identical nucleosomes, and thus might be predicted to form highly “ordered” secondary structures. Note that the term “ordered” can be used in the sense that a crystal is highly ordered and also used to describe hierarchical levels where the concept is more akin to its meaning in “orders of magnitude.” Although in one sense, the basic nucleosome-linker DNA unit comprising the primary structure of chromatin is simple, there are a number of potentially variable parameters that contribute to complexity. There is structural uncertainty even at the level of the nucleosome core particle (NCP). Although the structure of the NCP is well established at the atomic level (Luger et al. 1997), portions of the four core histones are not seen in crystals because of their evident mobility. For example, the unstructured

terminal regions that are extremely important in modulating chromatin structure are not seen in X-ray data. The length of linker DNA between NCPs varies not only between species, but also between tissues of the same organism, and within a single nucleus (van Holde 1989). Mean linker lengths (defined as the nucleosome repeat length minus the 145 bp of DNA protected in NCPs) range from a low value of 20 bp in budding yeast to 75 bp in echinoderm sperm. For a typical vertebrate nucleus the mean linker length is 35 bp. The rich literature concerning the locations of nucleosome positions on specific DNA sequences, reveals instances, especially in the upstream control regions of genes, in which nucleosomes are preferentially located or excluded, and this positioning is often important in regulating transcription (Simpson 1991; Jiang and Pugh 2009). For some DNA sequences, the preference for nucleosomes to be present at a particular location is retained when the nucleosomes are reconstituted on the DNA in vitro. In addition to linker DNA length, nucleosomes vary in their complement of core histone variants and histone postsynthetic modifications. All these variations are likely to impact chromatin secondary structures. The obvious experimental approaches to determine chromatin secondary and higherorder structures using light and electron microscopy yield disappointingly little information, largely because nucleosomes and linker DNA cannot be adequately resolved in the compact chromatin that occurs in the nucleus. A recent study using cryo-electron microscopy of thin sections of chromosomes vitrified in vivo, and therefore expected to faithfully reflect the native structure, provides an instructive example. Despite careful image processing of the micrographs, there was no evidence for any higher-order structure, leading the authors to suggest that the chromatin forms a “molten” mass similar to that assumed by certain polymers, and that in this form, the size, shape and trajectories of any higher-order structures would not be resolved (Eltsov et al. 2008). This may well be the case, but nevertheless, the arrangements of arrays of nucleosomes in

Chromatin Higher-order Structure and Dynamics

3D space are to some extent constrained and nonrandom, and important information would be obtained if it were possible to trace the paths of chains of nucleosomes through the nucleus. The generally amorphous appearance of chromatin in thin sections contrasts sharply with the appearance of cytoskeletal elements such as microtubules, which can be resolved in cells even when in compact bundles. This striking difference supports the concept that tightly packed chains of nucleosomes readily interdigitate with each other, leading to the rather featureless appearance of chromatin in EM images (Woodcock and Horowitz 1995; Woodcock 2006). A new technique for preparing frozen hydrated material that involves thinning of samples by milling with a focused ion beam (Marko et al. 2007, 2008) that does not involve the compression and possible local heating of cryosectioning is currently under development in several laboratories, and it will be most interesting to see whether this method provides improved structural information when applied to nuclear structures. The 30-nm Fiber

In the absence of clear conclusions from the examination of nuclei and chromosomes in situ, many investigators have focused on isolated chromatin. Through the use of nucleases that cut the linker DNA between nucleosomes, it is possible to isolate polynucleosomes and study their properties under defined conditions in solution. Early work clearly established that the compaction state of polynucleosomes is highly dependent on the ionic milieu. Naked DNA, being highly charged, is self-repulsive in low ionic strength buffers—the self-repulsion can be reduced or eliminated in the presence of cations, with divalent and polyvalent cations being especially effective. Chromatin behaves similarly to DNA in this respect because the DNA negative charge is not fully neutralized by histones. The response of chains of nucleosomes to changes in the ionic environment can be satisfactorily modeled (Clark and Kimura 1990; Arya and Schlick 2006), suggesting that our basic understanding of the response of

chromatin to changes in the ionic environment is robust. Electron microscopy of isolated polynucleosomes clearly reveals an open beads-on-a-string conformation in low salt and a progressive compaction that occurs as the ionic strength is raised. There is general agreement that an early stage in compaction is the formation of a fiber 30 nm in diameter, and that, at least for polynucleosomes in vitro, this constitutes a bona fide chromatin secondary structure. However, the arrangement of nucleosomes and linker DNA within isolated 30 nm fibers has been difficult to study and remains controversial. Careful examination of isolated 30 nm fibers has failed to reveal any consistent clear-cut arrangement of nucleosomes. The realization that native nucleosomes are not identical, but, as described earlier, vary in several parameters, suggested that these could be the source of the equivocal findings regarding 30-nm fiber organization. To eliminate these potential problems, investigators turned to fully defined artificial polynucleosomes reconstituted onto DNA containing regularly spaced nucleosome localization sequences (Simpson et al. 1985). A major achievement using this approach was the successful crystallization and structure determination by X-ray diffraction of a tetranucleosome (Schlach et al. 2005). This showed unequivocally that, at least for this particular construct, exposed to crystallization-promoting conditions that included 90-mM Mgþþ , the complex assumed a zigzag arrangement, with contacts between nucleosomes 1 and 3 and between 2 and 4, with the linker extending between them (Fig. 1). The work clearly established the predominant internucleosome interactions that result in a specific secondary structure for this particular chromatin construct in this ionic environment. It is not yet clear how far the zigzag structure can be extrapolated to long chromatin with heterogeneity in histone content and linker length. Nevertheless, it does provide an important context in which to view earlier results. For example, the interpretation of the first EM studies of isolated compact 30-nm fibers in terms of a simple solenoidal organization in

C.L. Woodcock and R.P. Ghosh

N2

N1'

N1

N2'

Figure 1. The DNA path of a tetranucleosome as

determined by X-ray diffraction. The structure consists of two stacks of nucleosomes, with linker DNA passing back and forth between them. Thus, the primary interactions occur between alternate rather than adjacent nucleosomes along the DNA strand, creating a zigzag architecture. From Schlach et al. 2005.

which linker DNA followed the helical trajectory established in the nucleosome is probably incorrect, whereas subsequent suggestions involving some species of zigzag are probably closer to the mark. THE ROLE OF HISTONE H1

In addition to the four “core” histones present as an octameric unit in the nucleosomes, a fifth histone, H1 or linker histone, is present in most nuclei. As its name implies linker histone is associated with linker DNA, and provides partial nuclease protection for 20 bp of linker DNA. H1 contains a conserved globular region and extended amino- and carboxy-termini, the latter being rich in lysines and able to interact strongly with DNA. Although the exact location of H1 in chromatin remains controversial, it is clear that it plays a crucial role in promoting chromatin higher-order structure. H1-containing chromatin shows a distinct structural motif in which the entering and exiting linker DNA segments are brought together, perhaps promoting an overall zigzag arrangement (Bednar et al. 1998). Arrays of nucleosomes depleted in H1 do not readily form 30-nm fibers in vitro, and it is likely that loss of the charge neutralization effected by H1 is at least partly responsible. The crystals of tetranucleosomes discussed above were created without H1—and to date, no publications of the X-ray structure of

H1-containing mono- or oligo-nucleosomes have appeared. Modest depletion of H1 from cells (achieved for the mouse by knocking out several of the six somatic H1 variants) results in reductions in nucleosome repeat length in some tissues whereas severe depletion is fatal for both mice and Drosophila (Fan et al. 2003; Lu et al. 2009). H1 depletion also inhibits the proper folding of chromosomes at mitosis (Maresca and Heald 2006). Interestingly, there is a strong linear relationship between H1 content and nucleosome repeat length in vivo (Woodcock et al. 2006), a feature that will act to maintain the electrostatic balance between DNA and histones. CHROMATIN FIBERS IN NUCLEI

Thin sections of typical nuclei reveal a coarse differentiation between strongly staining heterochromatin (see the following for more on this term) adjacent to the nuclear periphery and nucleoli, and more weakly staining euchromatin dispersed throughout. Little internal structural detail can be resolved in either component. There are, however, a few special cases that are more informative. Echinoderm sperm nuclei contain uniformly distributed and highly compact chromatin, but on controlled reduction in ionic strength, the chromatin swells, revealing a mass of 30-nm fibers. Although the fibers appear to be quite uniform in diameter, tomographic 3D reconstructions reveal an internal structure that consists of an irregular arrangement of nucleosomes and linker DNA but with a distinctive underlying zigzag motif (Horowitz et al. 1994). Sections of the transcriptionally inert chicken erythrocyte nucleus also reveal a uniform compaction of chromatin, which resolves into irregularly structured 30-nm fibers when allowed to swell slightly by, for example, lowering the ionic strength. In contrast, 30-nm fibers are rarely seen in more typical nuclei from cycling cells, even after controlled swelling (Woodcock and Horowitz, 1995). These special cases in which the 30-nm organization is revealed on swelling have several common features, including uniform core histone modifications, a greater

Chromatin Higher-order Structure and Dynamics

abundance of linker histone (H1) relative to core histones, the presence of H1 variants with more positive charges and unusually long linker DNA. It is not clear why these properties lead to the appearance of distinct 30-nm fibers in nucleo, but an attractive possibility is that these conditions favor intrafiber over interfiber interactions. NUCLEOSOME-NUCLEOSOME INTERACTIONS IN ISOLATED CHROMATIN FIBERS

One approach to understanding chromatin fiber structure is to identify nucleosome-nucleosome interactions within the compact fiber. As noted, this cannot be achieved by direct EM observation of compact fibers, but ways to examine this have been devised. One study used the premise that the face-to-face nucleosome contacts that occur in crystals of mononucleosomes (Luger et al. 1997) would likely be seen in compact chromatin. The contacts involved the positively charged N-terminus of histone H4 and the “acidic patch” that includes a portion of histone H2A. Dorigo et al. (2004) created H4 and H2A constructs with a cysteine in the putative contact region and reconstituted them into nucleosomal arrays. The arrays were then placed in conditions that promoted compaction and formation of S-S crosslinks between close cysteines. If the basic architecture of the array were solenoidal, crosslinking would result in a simple stack-of-coins conformation. In contrast, a zigzag arrangement would result in the creation of a ladder-like structure with the linker DNA segments forming the rungs. When cross-linking was initiated, the arrays clearly adopted the ladder-like structure, strongly supporting a zigzag architecture, albeit under somewhat artificial conditions. An approach that does not restrict internucleosome interactions to a specific histone-histone contact involves exposing nucleosomal arrays or whole cells to controlled formaldehyde crosslinking such that a few of the nucleosome-nucleosome contacts become covalently linked. Subsequently, the arrays are allowed to disperse in low salt and imaged by electron microscopy. With

his technique, termed EMANIC (EM-assisted nucleosome interaction capture) (Grigoryev et al. 2009), decondensed arrays appear in the open beads-on-a-string conformation (Fig. 2A), whereas crosslinked sites are seen as touching nucleosomes. Analysis of the crosslinking pattern of defined nucleosome arrays with H1 present once again reveals a predominantly zigzag pattern (Fig. 2 D, E). With H1 and 1 mM MgCl2 together, some crosslinking is also seen between adjacent nucleosomes, suggesting these conditions favor a mixed, heteromorphic fiber architecture (Fig. 2F, G) in which the zigzag dominates, but is interrupted by the type of nucleosome-nucleosome interaction expected for a solenoidal architecture. Interestingly, computer modeling predicts heteromorphic fibers, and also shows that this arrangement allows greater compaction as it reduces the crowding of linker DNAs in the fiber interior (Grigoryev et al. 2009), and is more consistent with the higher packing ratios seen in compact chromatin (Daban 2000; 2003). Although there is substantial support for a basic propensity of chromatin to form zigzag higher-order structures, results leading to alternate architectures have been presented. EM examination of long reconstituted nucleosomal arrays containing linker histone reveals remarkably uniform structures interpreted as consisting of solenoids in which adjacent gyres interdigitate (Robinson et al. 2006). This arrangement also increases the nucleosome packing density, and may thus better match the calculated in vivo packing. Additional studies of long nucleosomal arrays have led to the conclusion that the architecture is strongly influenced by linker length (Routh et al. 2008), increasing the potential complexity of chromatin structure in vivo, where varied linker DNA lengths are the norm. MECHANICAL PROPERTIES OF ISOLATED CHROMATIN FIBERS

The development of sophisticated instruments using “optical tweezers” has made it possible to take an isolated chromatin fiber, anchor one end, and pull on the other end while

C.L. Woodcock and R.P. Ghosh

A

Crosslinking

Unfolding and EM

i+/–1

i+/–2

B

C

D

D’ 2 8

E

F

G

H

I

I’ 2

2

Figure 2. EMANIC analysis of internucleosomal interactions. (A) Scheme of the EMANIC procedure. The two

models for the structure of the chromatin 30-nm fiber, namely solenoid (Upper) and zigzag (Lower), lead to dominant i + 1 and i + 2 internucleosome interactions, respectively. (B – D) EM of nucleosome reconstitutes crosslinked in low salt without linker histone show few crosslinks. (E-I) With H1 present, +2 interactions predominate. D0 and I0 diagram the nucleosome arrays corresponding to the adjacent EM images. From Grigoryev et al. 2009.

measuring the force/length relationship. The applied force is expected first to break the weak internucleosomal bonds holding the fiber together, and ultimately, the much stronger intranucleosomal bonds. For a fiber with a zigzag arrangement, the breaking of individual nucleosome-nucleosome interactions is predicted to produce jumps in length with a consistent increment which would not be observed with solenoid type structures. Analysis of force/length curves for isolated chromatin fibers from chicken erythrocyte nuclei led to the conclusion that a zigzag arrangement predominated (Cui and Bustamante 2000). However, force/length data from a more recent study using reconstituted nucleosomal arrays (Kruithof et al. 2009) were interpreted as supporting a solenoidal architecture. The differences in starting material and ionic conditions used in these studies makes it difficult to evaluate the findings. In the future, studies encompassing a broader range of conditions and

chromatin substrates should provide more definitive information from this potentially valuable technique. INSIGHTS FROM COMPUTER MODELING

As noted earlier, computer modeling was consistent with the empirical finding of a heteromorphic fiber architecture. Current “mesoscale” modeling programs are supplied with sophisticated structural and chemical models of the nucleosome, H1 (if desired) and linker DNA, as well as the selected ionic environment (Arya and Schlick 2006). The changes in structure are then followed over a large number of small increments of time, and the final structures analyzed. As expected, the stochastic nature of the interactions involved lead to different final structures with each run of the program. However, consistent motifs emerge: the final structures are fibers, rather than spherical aggregates of nucleosomes, and, although zigzag

Chromatin Higher-order Structure and Dynamics

arrangements of nucleosomes and linker DNA predominate, they may be interspersed with the bent linker motif characteristic of simple helical arrangements of nucleosomes. Modeling results are dependent on the accuracy of the input data, and computational limitations still preclude modeling at the atomic level. Nevertheless, this approach has the advantage of being able to sample a wide range of conditions, informing the selection of conditions for empirical examination with “real” chromatin. As computational speed continues to increase, modeling is likely to lead to more penetrating insights into chromatin architecture. LOCAL EVENTS THAT MAY MODULATE CHROMATIN FIBER ARCHITECTURE

Much of recent work on chromatin secondary structure has featured fully defined nucleosomal arrays in which recombinant histones are assembled in vitro onto DNA containing strong nucleosome positioning sequences. Yet, as noted, even with highly uniform arrays, results have led to differing conclusions regarding the underlying architecture. In the nucleus, local effects of linker DNA length variability, core histone variants, and postsynthetic histone modifications are likely to contribute even more irregularity to secondary structures. Additional complexity is introduced by the presence of nonhistone proteins known collectively as Chromatin Architectural Proteins (CAPs) that influence chromatin conformation (Luger and Hansen 2005). In addition to the “major” core histones, synthesized primarily in S phase and deposited at replication forks, there are numerous histone variants encoded by separate genes, which are often synthesized constitutively at low levels and incorporated differently. Some of these have been shown to play a critical role in establishing the local properties of chromatin in which they are embedded (Santenard and Torres-Padilla 2009). An important concept in this regard is the distinction between the major variant that is synthesized during S phase and deposited as newly replicated chromatin, and minor variants. It has been proposed that the

incorporation of different H3 variants serves as a “bar code” (Hake and Allis 2006) contributing to the establishment of the local functional potential. Of the several variants of H2A, H2A.Z has been particularly well studied and shown to modulate chromatin higher-order structure. The X-ray structure of nucleosomes containing H2A.Z (Suto et al. 2000) reveals an enlargement of the ‘acidic patch’ at the nucleosome surface that appears to strengthen nucleosome-nucleosome interactions, and contribute to the highly compact chromatin surrounding centromeres (Greaves et al. 2007). Another major source of chromatin variation is the large number of postsynthetic modifications of the core histones and H1 (Kouzarides 2007). They fall into two major categories, those that alter the charge on the histone, and those that establish binding sites for other proteins. Acetylation of lysines results in a reduction of the histone positive charge and weakens the DNA-histone interaction leading to a less compact chromatin secondary structure. The balance in activity between histone acetyl transferases (HATs) and histone deacetylases (HDACs) governs the acetylation status of a given region of chromatin. In general, hyperacetylation is the hallmark of active chromatin, whereas hypoacetylation is seen in repressed chromatin. A well-studied example is the acetylation of H4 at lysine 16. The loss of a positive charge at this site is likely to weaken the internucleosome interaction between the amino-terminus of H4 and the acidic patch on H2A discussed above. Indeed, the effect of this modification is to loosen the compaction of chromatin secondary structures (ShogrenKnaak et al. 2006; Fischle et al. 2003). An example of a modification that establishes a binding site is the methylation of H3 at lysine 9. The modification creates a binding site for a domain of the HP1 protein (Jacobs and Khorasanizadeh 2002). As discussed below, HP1 binding is associated with the formation and propagation of compact heterochromatin. Work on postsynthetic histone modifications has led to the widely discussed concept that, collectively, they constitute a “histone code” that defines the local structural and functional potential

C.L. Woodcock and R.P. Ghosh

of a region of chromatin (Jenuwein and Allis 2001). Chromatin architectural proteins, as their name implies, do impact chromatin structure, mostly by increasing the compaction state (Luger and Hansen 2005). This appears to be their sole common feature, because otherwise they are highly diverse in both structure and in chromatin binding mechanism. Binding may be restricted to specific DNA sequences (Polycomb), methylated DNA (MeCP2), or specific histone modifications (HP1). Only in a few cases, such as the SIR family of DNA-binding regulatory proteins is a detailed account of their molecular interactions with chromatin and with each other emerging (Buchberger et al. 2008). Some CAPs contain more than one DNA/chromatin binding sites, allowing them to act as inter- and/or intrafiber bridges, and thus promote chromatin compaction. Chromatin remodeling complexes may be considered as a distinct class of CAPs. Several families of these large multicomponent complexes have been identified. In vitro, they have the ability to restructure, eject, or move nucleosomes in an ATP-dependent manner (Clapier and Cairns 2009). This type of activity in vivo will likely influence chromatin secondary structure, and, by altering the local positioning of nucleosomes, modulate chromatin folding. An interesting case has been reported of the combined effect of a histone variant (H2A.Z), a histone modification (H3K9Me), and a CAP (HP1), all of which act to increase chromatin compaction (Fan et al. 2004). Thus, the proposed histone code based on histone modifications may need to be expanded to include synergistic effects of histone variants. A novel function in maintaining the pluripotential “open” state of chromatin for the remodeling factor Chd1 (Gaspar-Maia et al. 2009) may presage new insights into the roles of these complexes. A scenario of chromatin secondary structure that, in our opinion, appears to fit all the current data involves chains of nucleosomes that are sufficiently close packed in vivo to interdigitate, effectively merging with each other and preventing individual fibers being seen by

microscopy. Nevertheless, the overall trajectory of a chain of nucleosomes traces out an irregular 30-nm fiber within the nucleus. Fragmentation of chromatin by limited nuclease digestion followed by elution from nuclei results in individual fibers becoming separated, and available for microscopic and biochemical characterization. Irregularity dominates the architecture of isolated native chromatin fibers, reflecting the heterogeneity in linker DNA length, and composition of histone variants, histone postsynthetic modification, and CAPs. The end result is likely a hybrid (heteromorphic) organization with a predominantly zigzag architecture interspersed with other structural motifs. Most of these sources of irregularity are dynamic, changing at different time scales. Thus, it is probably futile to expect that chromatin secondary structure can be defined with the precision expected from studies of other biological fibers such as microtubules or Tobacco Mosaic Virus. Thus, seeking a defined secondary structure for chromatin may indeed be “Chasing a Mirage” (van Holde and Zlatanova 1995). Structural Order Beyond the 30-nm Fiber— Electron Microscopy Results

The lack of definitive information on chromatin secondary structure complicates the question of whether there is a bona fide set of hierarchical structures above the 30-nm fiber. Although there is no doubt that some higher levels of organization are present—the metaphase chromosome being an obvious example—precisely what form they take and whether they are truly hierarchical is still a matter of intensive study and debate. Much of this work uses the light microscope and has been limited in resolution to 250 nm by the properties of light. The impact of a 250-nm resolution limit is well illustrated in a study that considers the possible structural arrangement of large genetic loci (400 kb) by matching the predictions of a number of models for large-scale chromatin structures with actual observations (Muller et al. 2004). Somewhat surprisingly, at the resolution attainable with a high quality confocal

Chromatin Higher-order Structure and Dynamics

microscope, all models fitted the data equally well! The higher resolution of the EM, coupled with tomography, and applied to thin sections has revealed short regions of fiber-like structures with a variety of diameters, none particularly consonant with a hierarchical series based on a 30-nm organization (Konig et al. 2007; Belmont et al. 1989, 1994). More recently, however, significant improvements in technique have been made using an in vivo immuno-gold technique that promises to lead to important insights (Kireev et al. 2008). The introduction by Belmont and colleagues of techniques for constructing cell lines containing large foreign DNA segments that are “self identifying” through an ingenious use of the lac operator/repressor system (Belmont et al. 1999) has had a major impact within the field and is widely used. A recent adaptation of the method involving the intracellular injection of antibodies specific for the introduced chromatin followed by gold decoration of the antibodies, thin sectioning and EM observation reveals clear-cut cylindrical fibers as can be seen in the stereo pair in Figure 3 (Kireev et al. 2008). The fibers ranged in diameter from 120 nm to 170 nm with different mean values in two cell types examined. Only rarely were individual 30-nm-like fibers seen, and, as the authors note, it remains to be seen whether the thick fibers constitute one level in a hierarchical folding series. Hopefully, further exploitation of the technique will lead to long-awaited progress in our understanding of chromatin organization in situ. Structural Order Beyond the 30-nm fiber— Light Microscopy Results

The ability to identify specific segments of chromatin using light microscopy has provided important insights into interphase chromatin organization. However, in evaluating the impressive body of information obtained with the light microscope, it is important to bear in mind the preparative method used in each case, because this may significantly modulate the outcome of an investigation, especially when close to the resolution limit. Studies

Figure 3. Stereo pair of a section of nucleus in which a

large region of chromosome is decorated with gold particles. Clear fiber-like structures of the order of 100 nm are seen. Scale is 500 nm. From Kireeva et al., 2008.

with living cells are likely to provide the most reliable data, whereas the use of fluorescence in situ hybridization (FISH) requires a denaturation step which may introduce structural perturbations. Muller et al. (2004) reported that 400 kb segments of nontranscribed chromatin occupy roughly spherical volumes with a diameter not much greater than the resolution limit of the microscope. A 400 kb length of DNA translates into 2000 nucleosomes, and, if present as a linear 30 nm fiber with 10 nucleosomes per 10 nm length would have a length of 2 mm. Thus, in this case, an untranscribed region of chromatin must be extremely tightly packed. However, an underlying fiber-like organization below the resolution of the light microscope cannot be excluded (Tumbar et al. 1999). It will be very interesting to see if the higher resolution offered by subdiffraction light microscopy will reveal any internal structure in these compact regions. The situation changes dramatically on transcription, when loci typically expand, often becoming linear and adopting a beaded form. In an informative study, Shopland et al. (2006) examined the intranuclear distribution of a 4 Mb segment of mouse chromosome 14 that included four gene clusters separated by gene “deserts.” By labeling the gene clusters and deserts with different colored fluorescent probes, it was possible to trace the conformation of this large segment of chromatin (Fig. 4). Four patterns occurred in roughly

C.L. Woodcock and R.P. Ghosh

Figure 4. (A) A large locus consisting of 4-Mbp region containing regions of gene “deserts” (red fluorescence)

and gene clusters (green fluorescence) is seen in the nucleus in multiple configurations (C ). In general, gene deserts are more closely associated with the heterochromatin at the nuclear periphery (B). Scale is 1 mm. From Shopland et al. 2006.

equal proportions, ranging from linear “striped” arrangements to more compact clusters. Interestingly, the inactive segments were often clustered and oriented closer to the nuclear periphery than the gene-rich segments, in keeping with the preferential location of inactive chromatin in peripheral heterochromatin. HETEROCHROMATIN AND EUCHROMATIN

The terms heterochromatin and euchromatin refer to states of compaction and transcriptional potential rather than categories of chromatin higher-order structure per se. Nevertheless, they are used ubiquitously in the literature and merit a brief mention. Heterochromatin was originally defined well before the discovery of DNA as regions of nuclei that stained strongly with basic dyes (Heitz, 1928) and, together with its counterpart euchromatin, has provided a useful qualitative indication of chromatin compaction state. In general, heterochromatin tends to be located at the nuclear periphery, where specific interactions with the envelope may occur and often forms blocks surrounding the nucleolus. These distinctions can be clearly seen in light and electron micrographs. Transcription is largely

confined to euchromatin, and it is interesting to note that Heitz presciently suggested a functional difference between the two forms. Today, the term heterochromatin is more loosely applied, and is often extended to include transcriptionally silent regions of chromatin regardless of their staining properties. An important distinction is made between constitutive and facultative heterochromatin. Constitutive heterochromatin is always compact, and tends to be enriched in repetitive, gene-poor, and late replicating DNA sequences, whereas facultative heterochromatin can reversibly undergo transitions from a compact, transcriptionally inactive state to become more open, and transcriptionally competent. In a recent review, Trojer and Reinberg (2007) suggested that facultative heterochromatin be molecularly defined as condensed, transcriptionally silent chromatin regions that decondense and allow transcription within temporal, spatial, or parental/heritable contexts. During embryogenesis, for example, the amount of facultative heterochromatin increases as unwanted sets of genes are progressively shut down until at maturity, a cell expresses only the genes appropriate for that tissue. The reverse occurs when,

Chromatin Higher-order Structure and Dynamics

for example, differentiated cells are reprogrammed to become stem cells. These events are typically accompanied by profound changes in histone variants, histone modifications, and the presence of CAPS. An important feature of heterochromatin is its propensity to “spread” to adjacent regions of chromatin (Liaw and Lustig 2006; Hines et al. 2009). A recent comparison of properties of heterochromatin and euchromatin as diffusion barriers has yielded interesting and provocative results (Bancaud et al. 2009). Measurements of diffusion constants of large polymers within these nuclear compartments confirmed that heterochromatin constitutes a more crowded environment, leading to the more efficient trapping of chromatin binding proteins such as histone H1. The crowding effect was also suggested to assist in the maintenance of heterochromatin. Further, kinetic analyses indicated an anomalous component of diffusion that was interpreted in terms of a fractal chromatin organization at spatial scales below 100 nm. This indicates a physical arrangement more similar to the “molten globule” state suggested by the lack of structure in cryo sections than to the classic 30-nm fiber. The ubiquitous location of heterochromatin at the nuclear periphery and association with the nuclear lamina and nuclear envelope suggests that this location is both structurally and functionally important. It was therefore surprising to read that, in the rod photoreceptor cells of nocturnal, but not diurnal, mammals, heterochromatin is concentrated in the center of nuclei (Solovei et al. 2009). The authors postulate that with this arrangement, nuclei act as collecting lenses, providing additional sensitivity in very low light environments. This unexpected finding underscores the plasticity of chromatin organization. CHROMATIN LOOPS

The concept of large scale chromatin loops is important in many scenarios of chromatin function. The generic term “chromatin loop” is used for a variety of phenomena that play important roles in nuclear organization and

function, but otherwise may have little in common. Further, it is not at all clear whether the various loop phenomena that have been reported constitute distinct level(s) of chromatin higher-order structure (Kadauke and Blobel 2009). It is clear, however, that loop phenomena vary in terms of their stability. For example, enhancer-promoter loops that facilitate transcription are transitory, dynamic events, whereas other types of loops appear to be more stable. An early observation that has stood the test of time is the formation of a DNA “halo” around isolated nuclei exposed to a mild treatment that releases histones (Cook and Brazell 1975). The halos were shown to consist of supercoiled DNA, suggesting that the DNA twisting that occurs when histones are removed from nucleosomes is preserved by being anchored in the residual nuclear structure. This finding is also consistent with the concept of an insoluble proteinaceous nuclear matrix or karyoskeleton to which chromatin loops are anchored by DNA sequences referred to as matrix attachment regions (MARs). However, because it has not been possible to define its composition and structure, the matrix remains a useful working concept rather than a well-accepted structure in the same sense as the cytoskeleton (Pederson 2000). A somewhat different perspective involves the “looping out” from chromosome territories of large segments of genome that are related in terms of transcriptional activity. The supposition that these different loop phenomena reflect the same underlying chromatin organization was called into question by a surprising recent finding that halo diameter was related to the spacing of origins of replication during the previous S phase (Courbet et al. 2008). Under conditions of rapid replication, origins were widely spaced, leading to large halos, whereas slow replication triggered the firing of additional replication origins, and led to smaller halos. Halos can also be generated from metaphase chromosomes, and giant meiotic lampbrush chromosomes provide a particularly compelling example of large chromatin loops and their relation to transcriptionally coupled genes. Large-scale chromatin loops are featured in some models of chromatin and

C.L. Woodcock and R.P. Ghosh

chromosome organization, but how they are integrated into chromatin/nuclear structure and whether they constitute a distinct hierarchical level of chromatin organization remains to be established. The interaction between enhancers and promoters probably represents an unrelated manifestation of chromatin looping that is critical for transcriptional activity. Enhancer sequences are often many Kb distant from promoters, and may be located upstream, downstream, or on a different chromosome. There seems now to be a consensus that physical interaction between enhancer and promoter is necessary to initiate transcription (Visel et al. 2009), but how widely separated loci are brought into contact has remained an open question. One possibility, supported by recent data (Nolis et al. 2009) is that sequence-specific transcription factors bind to both enhancer and promoter, and also bind strongly with each other. In support of this hypothesis, “decoy” transcription factor binding sites placed between a promoter and its enhancer blocked transcription, suggesting that they trapped the enhancer in unproductive loops (Nolis et al. 2009). Finally, the recently introduced technique of chromosome conformation capture (3C) and related methods, which allow mapping of physical chromatin interactions in vivo, is providing growing evidence for physical interactions between distant loci other than enhancerpromoter juxtapositions (Gondor and Ohlssen 2009). Hints that there must be mechanisms for bringing specific loci together has come from the common occurrence of some chromosomal translocations, especially those leading to human diseases. For example, Roix et al. (2003) showed that genes associated with chromosome translocations leading to human lymphomas tend to be in physical proximity and located toward the nuclear interior. The data also suggested that the phenomenon was dependent on the “higherorder spatial organization” of the genome rather than the sequences of the genes involved. Human prostate cancer offers a system for tracking the physical proximity of two loci. The TMPRSS and ERG genes are 3MB apart on human chromosome 21, but become fused in 50% of

prostate cancer cases, apparently in response to hormone levels. Using FISH to follow the locations of the two genes, Mani et al. (2009) showed that, upon hormone treatment, the two genes became physically close in a significant proportion of cells. The mechanism(s) involved in this large-scale motion remain unknown. Another recent study of intranuclear chromatin associations capitalizing on massively-parallel sequencing has yielded important insights into the large scale organization of chromatin (LiebermanAiden et al. 2009). Although the technique in its current form is limited to megabase scale resolution, the data clearly indicated that active and inactive chromatin occupy different domains within chromosome territories. Further, at this level of resolution, chromatin conformation was consistent with a knot-free, fractal globule organization. In the representation of their conclusions, Lieberman-Aiden et al. show modeled nuclei with chromatin modeled as worm-like structures of uniform diameter, and it is important to appreciate that these structures are not intended to correspond to a specific level of chromatin higher-order structure. It is interesting to note that whereas this study indicated a fractal-like organization at very large scales, unrelated work on the accessibility of differentially packed chromatin suggested a fractal conformation for chromatin below 10 nm (Bancaud et al. 2009). Hopefully, these two sets of findings obtained through widely divergent experimental approaches, augur a future in which chromatin organization can be studied effectively at many different levels. The study of chromatin long range organization using 3C and related techniques (Rusk, 2009) is clearly at an early stage in development, and promises to provide important insights into the mechanisms that bring distant loci together and their relationship to chromosome territories and chromatin dynamics. The Metaphase Chromosome

The metaphase chromosome, in which the DNA is compacted some 10,000- to 20,000fold, is the one consistent manifestation of chromatin higher-order structure. It is therefore sobering to reflect that despite a great deal of

Chromatin Higher-order Structure and Dynamics

research, there is still no widely accepted model for the internal organization of this critical cellular component. Andrew Belmont, who has contributed seminal work in the field, has aptly applied the quotation “a riddle, wrapped in a mystery, inside an enigma” to illustrate the complexities of metaphase chromosome structure. There are several features that must be accounted for in any viable model. For a given species, the diameter of chromosome arms at metaphase is quite constant, although there are wide variations in chromosome diameter and number from species to species. The width constancy appears to be independent of DNA sequence. This is seen clearly in cases in which a chromosome bears a large insert, either as a response of the cell to a genetic defect that is overcome by accumulating multiple tandem repeats of the mutated gene (e.g., Sullivan and Bickmore 2000), or by insertion of large tandemly repeated loci. Typically, these inserts, which appear at metaphase as homogeneously staining regions (HSRs) on account of their lack of band/interband organization, have a uniform diameter that closely matches that of the rest of the chromosomes. This indicates that something other than DNA sequence is establishing the underlying folding pattern that governs chromosome diameter. It is well-established that, with the exception of polytene systems, chromosomes are unineme, in that a single strand of DNA, albeit multiply folded, extends from one end (telomere) to the other (reviewed in Gall 1981). Although for a particular chromosome, metaphase banding patterns and gene locations are consistent from cell to cell, the spatial resolution of light microscopy provides very limited structural information in the context of a chromosome arm that may have a diameter of 500 nm. There is, however, a great deal of information relevant to the overall architecture of metaphase chromosomes. An early and striking result was derived from treating chromosomes to remove histones, which, like similar treatments of nuclei, revealed a “halo” of DNA loops surrounding a denser core with a size and shape comparable to the starting chromosome (Earnshaw and Laemmli

1983). The core structures were termed “scaffolds” and it was postulated that they formed the structural basis of chromosome architecture. DNase treatment could be used to isolate the scaffolds, which contained specific DNA sequences (scaffold-attachment regions or SARS) and were enriched for a few proteins thought to be essential for scaffold, and hence chromosome structural integrity. Avery fruitful approach to chromosome structure that allows experimental manipulation uses amphibian egg extracts, which can be manipulated to recapitulate, in sperm nuclei, the essential cell cycle events of DNA replication, chromosome formation and mitosis (Lohka and Masui 1983). Importantly, the system allows the roles of individual components to be explored by their addition to or removal from the extract. Support for the scaffold hypothesis was provided by the observation that, in chromosomes formed in vitro in egg extracts, some components isolated from scaffolds were distributed axially at metaphase rather than uniformly. However, subsequent experiments established that the situation was more complex. For example, topoisomerase II could be removed from chromosomes assembled in vitro without compromising their structure (Hirano and Mitchison 1993), but was essential for proper chromosome assembly and separation of sister chromatids at anaphase. Further, genetic screens for chromosome segregation mutants revealed a family of “structural maintenance of chromosomes” (SMC) proteins (Hirano 2006). Also identified were two complexes, cohesin and condensin that contain SMCs and other proteins and, in additiontootherfunctions,appear tobeinvolved in sister chromatin adhesion and chromosome condensation (Hagstrom and Meyer 2003). Condensin is consistently seen to have an axial distribution in chromosomes, and in its absence or mutation, proper condensation fails. A completely different approach, in which micromechanical properties of chromosomes are measured, has introduced a dramatically different aspect of their structure. Importantly, these results have had the salutatory effect of requiring fresh thinking to reconcile the seemingly disparate sets of data. The strategy is to

C.L. Woodcock and R.P. Ghosh

isolate chromosomes either from mitotic cells or egg extracts, hold each end with micropipettes, pull on one end, and measure the force needed for a given length increase (Houchmandzadeh et al. 1997; Poirier and Marko 2003). The environment of the chromosome can be altered by modulating the buffer with cations etc., and the effects of nucleases or proteases can be determined. The results show that chromosomes exhibit a remarkable degree of elasticity (Fig. 5), repeatedly returning to the original length after being stretched fivefold. The general conclusion from several studies is that mitotic chromosomes consists of a network or gel in which individual chromatin fibers are connected by crosslinking elements, presumably proteins (Fig. 6). It was also possible to estimate a mean distance between crosslinks of 15 Kbp by applying restriction enzymes with different cutting frequencies to chromosomes before stretching (Poirier and Marko, 2002). The estimated distance between crosslinks is close to that calculated for the chromatin loops associated with chromosome scaffolds (Gasser et al. 1986). One surprising feature of the work was the large difference in flexibility between “native” chromosomes extracted from cells by micromanipulation and those assembled in egg extracts in vitro. The latter were almost two orders of magnitude more flexible, suggesting that their non-DNA components were thin, rigid, and highly extensible (Houchmandzadeh et al. 1997) In its simplest form, the network model seems to be consistent with the lack of consistent large-scale structures seen in cryo-sections of HeLa cell chromosomes (Eltsov et al. 2008) or in EM tomograms of chromosomes formed in Xenopus egg extracts (Ko¨nig et al. 2007), but at odds with hierarchical models of chromosome structure, which would predict that stretching a chromosome would result in the unfolding of hierarchical levels with stepwise reductions in chromosome diameter. The network model also seems to contradict the many light microscopic observations of an axial localization of key proteins, especially condensins known to be involved in chromosome compaction.

Figure 5. Time sequence showing the extension of a metaphase chromosome in which the two ends are anchored by micropipettes. The diameter remains quite constant throughout. From Marko 2008.

Recently, however, models that account for most of the apparently disparate observations have been put forward. An architecture based on observations of early chromosome condensation that involves hierarchical folding with an axial “glue” provided by condensin (Fig. 7) has been proposed by Kireeva et al. (2004), and a more elaborate system with specific locations and roles for topoisomerase II, cohesin and condensin has been put forward by Marko (2007). In the case of the model in Figure 7, a delicate balance between the forces maintaining the 200– 250 nm and 500 – 750 nm fibers would Linker protein

30 nm chromatin fiber

Mitotic chromosome

Extensible net model of mitotic chromosome structure derived from force-extension measurements. From Poirier and Marko 2002.

Figure

6.

Chromatin Higher-order Structure and Dynamics

A

30 nm

B

100–130 nm

200–250 nm 500–750 nm

Figure 7. Model of chromosome formation that incorporates the concept of a central axis enriched in condensins (red dots) and irregularly folded chromatin fibers. From Kireeva et al. 2004.

be needed to produce the stretching effect shown in Figure 5. As noted by Belmont (2006), many questions remain unsolved, and there are probably additional proteins, levels of complexity, and redundant mechanisms to be discovered before the complete story of chromosome architecture, formation and maintenance is uncovered. Indeed, a new activity, RCA (regulator of chromosome architecture), that can condense chromosomes in the absence of condensins has recently been proposed (Vagnarelli et al. 2006). To date, no satisfactory explanation of the constant diameter of chromosomes has emerged. Thus, the simple-appearing metaphase chromosome is far from simple, and fully understanding its composition, formation, maintenance and architecture will likely remain a challenge for some time. LAMPBRUSH AND POLYTENE CHROMOSOMES

These two special and quite rare manifestations of chromatin provide some insight into the

surprisingly wide range of nuclear organizations that “work.” Lampbrush chromosomes (LBCs) are largely confined to meiosis in oocytes, especially avian and amphibian, but are also present in the large unicellular alga Acetabularia. They appear to be an adaptation for maximizing transcriptional output to serve thousands of cells in the developing embryo and the macroscopic cell body in Acetabularia. Light microscopy reveals an axial core from which loops of varying size extend. Loops contain transcription units that can be seen in the electron microscope to be loaded with RNA polymerases and continuously transcribing. Although the loops are the most conspicuous features of LBCs, they represent only a small fraction of the genome, the bulk forming the compact chromatin of the axial cores. The fact that sperm nuclei exposed to extract from oocytes at the lampbrush state develop into haploid LBCs (Gall and Murphey 1998) indicates that the signals needed to transform a chromosome into the lampbrush state evidently reside in the oocyte. A key question is whether this loop organization is unique to lampbrush chromosomes or whether it offers an especially clear example of a general organizing principle for transcriptionally active chromatin. Although the growing evidence for the ubiquitous organization of chromatin into large scale loops containing functionally related genes is consistent with the latter interpretation, there is also compelling evidence transcription of “normal” chromatin is organized quite differently (Hu et al. 2009). Polytene chromosomes are formed by repeated mitoses without nuclear division and consist of genomes precisely aligned in trans. They are especially prominent in dipteran larva, with 1024 copies of the genome being present in the third instar of Drosophila. In the light microscope, a unique pattern of bands and interbands is seen, allowing the locations of individual genes to be mapped cytologically. It has long been recognized that at the sites of transcriptionally active genes, “puffs” appear as the chromatin is locally decondensed. Disappointingly, polytene chromosomes have not yielded much insights into chromatin organization in general.

C.L. Woodcock and R.P. Ghosh

In the electron microscope, their alignment appears far from regular, and it is as difficult to recognize and follow individual chromatin fibers as in normal diploid nuclei. Polytene chromosomes may also be used to infer the structural organization of interphase chromatin. For example, a component that, when depleted or mutated, results in disorganization of polytene chromosomes, may be involved in sister chromatid adhesion. Polyteny is also seen in the first stages of the development of the nurse cells surrounding the Drosophila ovary. As nurse cells mature, the polytene alignment breaks down, and the nuclei become polyploid. Surprisingly, in flies with mutations of putative condensin 2 components, the polytene chromosomes fail to disperse at the appropriate time, whereas the same mutations induce dispersion of salivary gland chromosomes (Hartl et al. 2008). These findings may lead to further insights into the complex events controlling polytene chromosome integrity. Both lampbrush and polytene chromosomes dispel the idea that transcription is obligatorily confined to “factories” that contain RNA polymerases and the multiple other factors necessary for RNA synthesis and processing (Cook 1999). Observation of lampbrush loops clearly indicates that it is the polymerases that move along the loops rather than vice versa. Similarly, it seems very unlikely that DNA replication in polytene chromosomes occurs at replication factories through which DNA is translated (Cook 1999). These considerations do not lessen the potential importance of transcription and replication factories in general, but again emphasize the remarkable plasticity of nuclear organization. CHROMATIN DYNAMICS

The profound structural rearrangements that chromatin must undergo during the cell cycle underscores one aspect of the dynamic nature of chromatin. There is, in addition, compelling evidence of an important role for local structural changes in compaction and/or position of specific genetic loci likely to be of critical importance in their transcriptional regulation.

In fact, most of the functionally important characteristics of chromatin show dynamic behavior in the sense of time-dependent changes. These include the mobility of many chromatinbinding components, including chromatin architectural proteins (Phair and Misteli, 2000), and the status of posttranslational modifications, especially of the core histones. Many of these phenomena are likely to impact local and global chromatin conformation and thus modulate higher-order structure. The study of chromatin dynamics in all of its manifestations has opened up exciting new perspectives—indeed, a literature search reveals over 60 reviews with titles containing the terms “chromatin” and “dynamics” (e.g., Huebner and Spector 2010). Here, we focus on aspects of dynamics that are likely to impact higher-order structures. All involve changes in location and/ or shape of specific genetic loci that are thought to have important functional implications. As the field of chromatin dynamics has developed, it has become increasingly clear that a number of underlying physical and molecular phenomena are involved, and that to understand them fully, information at many different levels is required. For a given locus these may include the temporal and spatial scales of observed movements, their energy dependence, and their location, both within the nucleus and within their specific chromosome territory. One important goal of current research is to be able to relate these changes in location/shape to the underlying physical changes in chromatin higher-order structure. For movements that are correlated with changes in transcriptional activity, it is also important to determine whether a change in location is a prerequisite for, or a consequence of, the altered level of RNA synthesis. Advances in light microscopy techniques over the past 25 years have allowed chromatin dynamics to be examined with steadily improving temporal and spatial resolution, and has enabled three principal types of dynamic change to be recognized (Soutoglou and Misteli 2007). These differ in the extent of motion, their energy dependence, and the time scales involved (Fig. 8). Early studies tracking changes in

Chromatin Higher-order Structure and Dynamics

location of large segments of chromatin and whole chromosome territories indicated that, when corrected for nuclear rotation, the regions were essentially immobile over distances .0.4 mm (Cremer et al. 1982, Diboni and Mintz 1986, Shelby et al. 1996). However, more recent work combining higher spatial resolution with in vivo labeling of smaller defined segments of chromatin, revealed a more complex scenario with different regions of chromatin showing very different mobilities. For example, a segment of budding yeast chromatin located near a centromere was found to show constrained random walk diffusive motion with a confinement radius of 0.3 mm (Marshall et al. 1997), that was independent of the metabolic state of the cell. The rate of mobility of this region of chromatin was calculated to be approximately three orders of magnitude lower than expected for free DNA of similar length (Marshall et al. 1997), leading to the suggestion that the locus was tethered within the nucleus, perhaps to the nuclear envelope or some internal structure. A detailed study of chromatin

mobility in Drosophila spermatocytes using loci labeled with the lac repressor system recorded relatively large movements during short time intervals in early nuclei. Movements of this magnitude would be expected to result in displacements of over 4 mm in 1 h and cover the entire 11 – 12 mm diameter of a nucleus in 6– 7 h (Vazquez et al. 2001). However, when long time periods were examined, it was clear that loci were constrained within a volume approximately equivalent to a chromosome territory. Another important finding was that spermatocytes in late G2 stage and approaching meiosis showed greatly reduced chromatin mobility, suggesting that some form of additional tethering precedes chromosome condensation. A comparison of the mobility of different chromosomal sites in yeast also revealed striking locus- and cell cycle-dependent differences in chromatin dynamics (Heun et al. 2001). Some chromatin regions showed occasional large (0.5 mm) movements over time periods as short as 10 seconds that were inhibited in ATPdepleted cells and thus dependent on the

Short-range motion -Locally constrainemd within ~1μm -Fast (seconds) -Random direction -ATP dependent -Frequent -Occurs by default

Long-range motion -Several micrometers -Slow (minutes) -Directed -ATP/motor dependent -Rare -In response to physiological cues (cell cycle)

Genome Reorganization -Several micrometers -Slow (hours, days) -Global changes -Likely requires cell division -In response to physiological cues (differentiation, development)

Figure 8. The different forms of large-scale motion that contribute to chromatin dynamics. From Soutoglou and

Misteli 2007.

C.L. Woodcock and R.P. Ghosh

metabolic state of the cell. The fact that the magnitude of these large movements is similar for organisms with widely different nuclear volumes has important structural and functional implications. For example, in budding yeast, which has a nuclear diameter of 2 mm, movement within a radius of 0.5 mm would allow access to a large portion of the nuclear volume. In contrast, in a typical mammalian nucleus of 10 mm in diameter this motion would explore only a thousandth of the nuclear volume. In yeast, the effectively large displacements of loci within nuclei may promote the observed high recombination frequency, which may require substantial intermingling among different regions of the genome (Gasser 2002). Observations of “free” ectopic regions of chromatin, exemplified by large (15 kb) circular plasmids in yeast underscore the context dependence of chromatin motion. An early study using a centromere-containing plasmid reported very limited movements, similar to those observed for centromeric regions in intact chromosomes (Marshall et al. 1997). However, more recently, Gartenberg et al. (2004) using plasmids designed to be either transcriptionally silent or competent, found that the active plasmids showed unconstrained movement, whereas the silent ones were strongly constrained and preferentially located near the nuclear envelope. Also relevant to the factors controlling intranuclear mobility is the finding that whereas double-stranded DNA breaks tend to stay together through the repair process (Kruhlak et al. 2006), the absence of a critical repair factor leads to long-range movements of the broken ends (Downs et al. 2004). A general pattern emerging from numerous studies using different organisms is that all chromatin loci show constrained Brownian motion with rather similar diffusion constants, but a wide range of confinement volumes. A strong relationship between the confinement volume, and distance from the nuclear envelope has also been established, with loci closer to the envelope being more constrained. The periphery of the nucleus proximal to the nuclear envelope is considered a transcriptionally repressive environment, establishing a correlation between

transcriptional silencing and limited intranuclear mobility (Marshall 2002). The underlying mechanism for this correlated behavior is not clear, but there is evidence that the constrained motion results from physical tethering to the nuclear lamina (Heun et al. 2001; Chubb et al. 2002; Gartenberg et al. 2004). The higher spatial resolution (20 nm) of two-photon microscopy, together with a temporal resolution of 30 msec has revealed new levels of complexity for constrained diffusive motion of chromatin (Levi et al. 2005). In these experiments, GFP-labeled repeats inserted in CHO cells showed periods of rapid constrained diffusive motion alternating with occasional energy-dependent curvilinear leaps of 150 nm that lasted only 0.3 – 2.0 second. The energy-dependence of these leaps suggested that they may need the activity of chromatin remodeling complexes or other ATP-requiring changes in chromatin organization that could lead to events such as the decondensation of compact 30-nm fibers resulting in the rapid changes in location. Another important outcome of this study pertinent to the extent of physical coupling between neighboring loci on the same chromosome was the finding that loci 1– 2 mm apart in the nucleus moved independently (Levi et al. 2005). A similar conclusion was reached by Hu et al. (2009) who showed that for a 2 Mb region of chromatin that appeared as a linear cluster of beads in the light microscope, the individual beads moved independently. These findings suggest that neighboring loci are not structurally tied together by, for example, being anchored to an underlying nonchromatin structure. As it becomes possible to study yet closer regions of chromatin, there will be a point at which movement of the neighbors is correlated, and defining this point will be very useful in being able to infer the effective chromatin compaction level. Thus, although the accumulating data clearly demonstrate that chromatin in the interphase nucleus undergoes constant dynamic reorganization through constrained diffusive motion, the level(s) of chromatin higher-order structure that participate in this phenomenon remain to be determined.

Chromatin Higher-order Structure and Dynamics

Some hints of the mechanism whereby loci undergo directed movement have emerged from work on an inserted transgene that typically resides in peripheral heterochromatin, but tends to relocate toward the center of the nucleus within 1– 2 hour of transcriptional up-regulation (Chuang et al. 2006). Importantly, this movement was blocked by the expression of a mutant nuclear myosin I and by a nonpolymerizable mutant of actin. This intriguing finding should be viewed in the context that several families of actin and actinrelated proteins (ARPs) are found in the nucleus, many of which are components of chromatin remodeling and histone modification complexes (Chen and Shen 2007). Also, it has been shown that the BAF remodeling complex binds to the ends and branch points of actin filaments in a PIP2-dependent manner (Rando et al. 2002). Unlike their cytoplasmic counterparts, nuclear actins and ARPs tend to occur as monomers rather than the filamentous structures, and it is therefore curious that it was an actin polymerization mutant that was defective in long range movement (Chuang et al. 2006). Determining the mechanism by which actin/myosin influences the mobility and directionality of loci is clearly an important future goal. CHROMATIN DYNAMICS AND TRANSCRIPTION

Studies of the spatial locations and transcriptional competence of loci with respect to their chromosome territories has provided some important insights. For example, the ANT2 gene located on the X chromosome is found in the interior of the territory of the inactive homolog, but peripherally on the active homolog (Dietzel et al. 1999). In contrast, the related ANT3 gene, which is located on the pseudoautosomal region of the X chromosome was peripherally located on the territory of both active and inactive homologs. This pattern of inactive genes located inside chromosome territories, and relocating to the periphery, perhaps by a “looping out” mechanism requiring a dramatic relaxation of local chromatin structure,

has been widely observed (Chambeyron and Bickmore 2005; Chambeyron et al. 2005; Volpi et al. 2000). Other studies have documented large scale movements of loci undergoing changes in transcriptional status as part of a developmental program. Loci were found to relocate to specific regions of the nucleus rather than locations within territories. An informative example is provided by the nuclear reorganization that accompanies differentiation of mouse lymphocytes (Brown et al. 1997, 1999). Here, lymphoid genes targeted for shutdown become relocated close to the heterochromatic clusters of pericentromeric chromatin characteristic of murine cells. Importantly, in this case, transcriptional repression appears to precede movement, suggesting that relocation is not a prerequisite for silencing. The relocation of genes destined for transcriptional up-regulation away from centromeric chromatin (Schu¨beler et al. 2000) or the nuclear periphery (Kosak et al. 2002) has also been documented. Although there is a consensus regarding the correlation of intranuclear and intraterritory location with transcription, the chromatin conformational changes that accompany switches in transcriptional status are still far from clear. On the one hand, it seems unlikely that the bulky transcription machinery could access DNA and operate in compact chromatin, and in this respect, the concept of the chromatin looping out from compact domains to facilitate transcription is attractive. Indeed, transcriptional activity in LBC loops and polytene puffs is demonstrably unfolded, in some cases to the beads-on-a-string primary structure. Although there is general agreement that segments of chromatin are more compact when transcriptionally active than when silenced (e.g., Muller et al. 2001), the extent of this decondensation is unlikely to be as great as in LBC loops. A recent study designed to examine the compaction state of active and inactive chromatin in interphase nuclei has indicated quite small transcription-related changes in higher-order structure, implying that transcription can proceed in quite compact chromatin (Hu et al. 2009). Labeled 2 Mb chromatin domains designed to support a level of transcriptional

C.L. Woodcock and R.P. Ghosh

activity typical of nuclear genes were inserted into cells, and tracked over time. In the uninduced state, the arrays assumed a linear, often beaded conformation, with lengths indicative of DNA packing ratios of over 1000:1. On transcriptional induction, the arrays retained their linear, beaded conformation, and showed only a modest 1.5- to threefold increase in length. No evidence was seen for large-scale looping or unfolding to the primary beads-on-a-string level of chromatin structure. Can transcription proceed in a compact structure that is over 10-times more compact than a 30-nm fiber? In terms of access, several studies have shown that large macromolecules can penetrate quite compact chromatin (Verschure et al. 2003; Chen et al. 2005; Bancaud et al. 2009), perhaps as a function of the dynamic mobility of chromatin. Also, in a simple in vitro system, conformational fluctuations allowed direct access to the central nucleosome in a compact trinucleosome (Poirier et al. 2009). It is also possible that the binding of transcription factors could be followed by extensive, local unfolding, undetectable at the resolution of the light microscope. The many facets of chromatin dynamics span orders of magnitude both in the spatial and temporal domains. Placing this complexity in the context of the level(s) of chromatin higher-order structure that are modulated during changes in transcriptional status remains an important challenge in understanding nuclear function. The difficulty is compounded by the current uncertainty regarding chromatin organization above the 30-nm fiber level. Nevertheless, continuing improvements in the spatial resolution with which specific loci can be imaged should soon begin to narrow the possible interpretations in terms of the level(s) of chromatin structure that could be involved. For example, as it becomes possible to image closer and closer neighboring loci on the same chromatin and follow their individual dynamics, there should be a point at which coupled motion will be observed. Once that point is reached, it will become possible to combine knowledge of the physical properties of arrays of nucleosomes with the observed dynamics

and decide whether, for example, the data are consistent with a change from a compact to a decondensed 30-nm fiber. New strategies that allow “super” resolution in light microscopy will soon be in common use and may open the way to this type of correlative work. CONCLUDING REMARKS

The past few years have witnessed remarkable progress in our understanding of chromatin higher-order structure, and this rapid trajectory can be expected to continue. There are some areas of research that seem especially poised to generate significant new insights in the near future, and others that have been overly neglected. Approaches that are likely to provide important new information in the near future include the application of “super resolution” light microscopy, and further exploration of the role of RNAi and other small and long nuclear RNAs (for a review see Wilusz et al. 2009). A plethora of strategies for extending the resolution limitation of the light microscope well beyond the “diffraction limit” is maturing and some have been implemented commercially (Huang et al. 2009). Techniques capable of generating high resolution 3D images of the nuclear interior using principles such as PALM and STED will be especially suitable for studies of nuclei and chromatin. However, as experience with electron microscopy shows, high resolution alone may not be informative, especially for compact chromatin. Progress in understanding the paths of putative 30-nm fibers in nuclei will require both high resolution and a contrasting technique that highlights a single stretch of DNA/chromatin. One application of an interference-based subdiffraction technique that was focused on the nuclear envelope showed the remarkable improvement that can be expected (Schermelleh et al. 2008), but little additional detail was seen in peripheral heterochromatin. It is also important that, as resolution improves further, preparative methods that provide a level of preservation consistent with the spatial resolution of the imaging system. Such methods should be capable of

Chromatin Higher-order Structure and Dynamics

preserving membranous and cytoskeletal elements, as judged by electron microscopy. Since the discovery of RNAi and other small nuclear RNAs, their importance in a host of nuclear functions has been shown. Among these functions is a role in the formation of the constitutive heterochromatin that flanks centromeres (Verdel et al. 2004; Moazed et al. 2006; Li et al. 2009). The process involves an RNAi-containing complex (RNA-induced transcriptional silencing, RITS) that binds to specific DNA sequences via siRNA-directed base pairing, and also to histone H3 methylated at lysine 9, a heterochromatin-specific histone modification. In combination, these processes promote the spreading of heterochromatin. It seems very likely that, as the roles of small nuclear RNAs become more fully understood, they will be found to impact other aspects of chromatin higher-order structure. In addition, the identification and characterization of long noncoding RNAs that are retained in the nucleus is beginning to emerge as an important field that will add to our understanding of gene expression (see Wilusz et al. 2009 for a recent review). In the category of neglected areas that merit further study are the ionic composition of nuclei, chromosomes, and chromatin, and the perhaps related issue of the impact of intranuclear molecular “crowding” (Hancock 2004a). Given the dramatic effects of changes in conformation that isolated chromatin undergoes in response to changes in the ionic environment, it is surprising that so little is known about the intranuclear ionic environment. It is generally assumed that “physiological” salt includes 100 mM monovalent ions, and a smaller concentration (1 – 2 mM) of divalent cations. Buffers for work with isolated nuclei and chromatin in solution typically include Naþ or Kþ for the monovalent cation, Mgþþ for the divalent cation, and Cl2 for the anion. It is also well established that polyamines, especially spermidine (4þ) and spermine (8þ) are ubiquitous nuclear components with potent chromatin compaction properties. Very little is known about the intranuclear distributions and concentrations of these charged species, making it

difficult to have confidence that in vitro experiments and modeling studies are based on milieus that accurately reflect conditions in the nucleus. An isolated report of the distribution of cations in nuclei and isolated mitotic chromosomes using secondary ion mass spectrometry (Strick et al. 2001) concluded that during the transition from interphase to mitosis, Caþþ and Mgþþ concentrations increased three- to four-fold, reaching 18 mM and 11 mM respectively, whereas isolated chromosomes contained almost twice these amounts. Interestingly, Caþþ , but not Mgþþ showed an axial enrichment in chromosomes. Of the monovalent ions, nuclear Naþ concentrations were close to those measured in the cytosol, whereas Kþ was clearly enriched in both interphase and mitosis. It is not clear what the effect would be of using these remarkably high divalent ion levels in nuclear buffers, and it is unfortunate that the technique has not been further exploited using a wider range of ionic conditions. An area of research that has received little attention until recently, but may have a farreaching impact on our understanding of chromatin function and dynamics relates to the highly crowded nuclear milieu (Hancock 2004a; Richter et al. 2008; Miyoshi and Sigimoto 2008; Bancaud et al. 2009). It is clear that the extremely high concentrations of macromolecules (of the order of 100 mg/mL) that are present in the nucleus have important thermodynamic implications (Minton, 2001). These concentrations cannot be effectively recapitulated in vitro, with the consequence that experiments in solution may not reflect the events that occur in nucleo. In one set of experiments, when isolated nuclei were allowed to swell in low ionic strength medium, nucleoli and PML bodies disassembled and nucleolar transcription was dramatically reduced (Hancock 2004b). However, after adding inert macromolecules ( polyethylene glycol or dextran) to 12% w/v, the compartments reassembled, and nucleolar transcription recovered. Interestingly, this experiment also showed that it was not the lack of ions per se that caused disassembly. A fuller understanding of these effects, as well as the

C.L. Woodcock and R.P. Ghosh

intranuclear levels and roles of ions is likely to expand our understanding of chromatin and nuclear organization. This snapshot in time of chromatin higherorder structure and dynamics reveals an exciting period in research, with rapid advances being made on many fronts. As the different interweaving threads begin to coalesce, we can anticipate a much more comprehensive understanding of the structural and functional organization of chromatin in the context of its nuclear location. An overriding theme is that chromatin and its higher-order structures feature plasticity and dynamics rather than rigidity. How this is possible in what must at some level be a self-organizing system (Misteli 2001) remains a key open question.

REFERENCES Arya G, Schlick T. 2006. Role of histone tails in chromatin folding revealed by a mesoscopic oligonucleosome model. Proc Natl Acad Sci 103: 16236– 16241. Bancaud A, Huet S, Daigle N, Mozziconacci J, Beaudouin J, Ellenberg J. 2009. Molecular crowding affects diffusion and binding of nuclear proteins in heterochromatin and reveals the fractal organization of chromatin. EMBO J doi:10.1038/emboj.2009.340. Bednar J, Horowitz RA, Grigoryev SA, Carruthers LM, Hansen JC, Koster AJ, Woodcock CL. 1998. Nucleosomes, linker DNA, and linker histone form a unique structural motif that directs the higher-order folding and compaction of chromatin. Proc Natl Acad Sci 95: 14173–14178. Belmont AS. 2006. Mitotic chromosome structure and condensation. Curr Opin Cell Biol 18: 632 –638. Belmont AS, Bruce K. 1994. Visualization of G1 chromosomes: A folded, twisted, supercoiled chromonema model of interphase chromatid structure. J Cell Biol 127: 287– 302. Belmont AS, Braunfeld MB, Sedat JW, Agard DA. 1989. Large-scale chromatin structural domains within mitotic and interphase chromosomes in vivo and in vitro. Chromosoma 98: 129–43. Belmont AS, Li G, Sudlow G, Robinett C. 1999. Visualization of large-scale chromatin structure and dynamics using the lac operator/lac repressor reporter system. Methods Cell Biol 58: 203–222. Brown KE, Baxter J, Graf D, Merkenschlager M, Fisher AG. 1999. Dynamic repositioning of genes in the nucleus of lymphocytes preparing for cell division. Mol Cell 3: 207–17. Brown KE, Guest SS, Smale ST, Hahm K, Merkenschlager M, Fisher AG. 1997. Association of transcriptionally silent genes with Ikaros complexes at centromeric heterochromatin. Cell 91: 845– 54.

Buchberger JR, Onishi M, Li G, Seebacher J, Rudner AD, Gygi SP, Moazed D. 2008. Sir3-nucleosome interactions in spreading of silent chromatin in Saccharomyces cerevisiae. Mol Cell Biol 28: 6903–6918. Chambeyron S, Bickmore WA. 2004. Chromatin decondensation and nuclear reorganization of the HoxB locus upon induction of transcription. Genes Dev 18: 1119–1130. Chambeyron S, Da Silva NR, Lawson KA, Bickmore WA. 2005. Nuclear re-organisation of the Hoxb complex during mouse embryonic development. Development 132: 2215–2223. Chen M, Shen X. 2007. Nuclear actin and actin-related proteins in protein dynamics. Curr Opin Cell Biol 19: 326 –330. Chen D, Dundr M, Wang C, Leung A, Lamond A, Misteli T, Huang S. 2005. Condensed mitotic chromatin is accessible to transcription factors and chromatin structural proteins. J Cell Biol 168: 41–54. Cremer T, Cremer C, Baumann H, Luedtke EK, Sperling K, Teuber V, Zorn C. 1982. Rabl’s model of the interphase chromosome arrangement tested in Chinese hamster cells by premature chromosome condensation and laser-UV-microbeam experiments. Hum Genet 60: 46– 56. Chuang CH, Carpenter AE, Fuchsova B, Johnson T, de Lanerolle P, Belmont AS. 2006. Long-range directional movement of an interphase chromosome site. Curr Biol 16: 825– 31. Chubb JR, Boyle S, Perry P, Bickmore WA. 2002. Chromatin motion is constrained by association with nuclear compartments in human cells. Curr Biol 12: 439–445. Clapier CR, Cairns BR. 2009. The biology of chromatin remodeling complexes. Annu Rev Biochem 78: 273– 304. Clark DJ, Kimura T. 1990. Electrostatic mechanism of chromatin folding. J Mol Biol 211: 883– 96. Cook PR. 1999. The organization of transcription and replication. Science 284: 1790–1795. Cook PR, Bazell IA. 1975. Supercoils in human DNA. J Cell Sci 19: 261– 279. Courbet S, Gay S, Arnoult N, Wronka G, Anglana M, Brison O, Debatisse M. 2008. Replication fork movement sets chromatin loop size and origin choice in mammalian cells. Nature 455: 557–560. Cui Y, Bustamante C. 2000. Pulling a single chromatin fiber reveals the forces that maintain its higher-order structure. Proc Natl Acad Sci 97: 127– 32. Daban J-R. 2000. Physical constraints in the condensation of eukaryotic chromosomes. Local concentration of DNA versus linear packing ratio in higher order chromatin structures. Biochemistry 39: 3861–3866. Daban J-R. 2003. High concentration of DNA in condensed chromatin. Biochem Cell Biol 81: 91–99 DiBoni U, Mintz AH. 1986. Curvilinear, 3-dimensional motion of chromatin domains and nucleoli in neuronal interphase nuclei. Science 234: 863– 866. Dietzel S, Schiebel K, Little G, Edelmann P, Rappold GA, Eils R, Cremer C, Cremer T. 1999. The 3D positioning of ANT2 and ANT3 genes within female X chromosome territories correlates with gene activity. Exp Cell Res 252: 363–375.

Chromatin Higher-order Structure and Dynamics Dorigo B, Schlach T, Kulangara A, Duda S, Schroeder RR, Richmond TJ. 2004. Nucleosome arrays reveal the two-start organization of the chromatin fiber. Science 306: 1571–1573. Downs JA, Jackson SP. 2004. A means to a DNA end: The many roles of Ku. Nature Rev Mol Cell Biol 5: 367 –378. Earnshaw WC, Laemmli UK. 1983. Architecture of metaphase chromosomes and chromosome scaffolds. J Cell Biol 96: 84–93. Eltsov M, Maclellan KM, Maeshima K, Frangakis AS, Dubochet J. 2008. Analysis of cryo-electron microscopy images does not support the existence of 30-nm chromatin fibers in mitotic chromosomes in situ. Proc Natl Acad Sci 105: 19732–19737. Fan Y, Nikitina T, Morin-Kensicki EM, Zhao J, Magnuson TR, Woodcock CL, Skoultchi AI. 2003. H1 linker histones are essential for mouse development and affect nucleosome spacing in vivo. Mol Cell Biol 23: 4559–72. Fan JY, Rangasamy D, Luger K, Tremethick DJ. 2004. H2A.Z alters the nucleosome surface to promote HP1alphamediated chromatin fiber folding. Mol Cell 16: 655– 61. Fischle W, Wang Y, Jacobs SA, Kim Y, Allis CD, Khourasanizadeh S. 2003. Molecular basis for the discrimination of repressive methyl-lysine marks in histone H3 by Polycomb and HP1 chromodomains. Genes Dev 17: 1870– 1881. Gall JG. 1981. Chromosome structure and the C-value paradox. J Cell Biol 91: 3s– 14s. Gall J, Murphy C. 1998. Pictures in cell biology. Lampbrush chromosomes from sperm chromatin. Trends Cell Biol 8: 207. Gartenberg MR, Neumann FR, Laroche T, Blaszczyk M, Gasser SM. 2004. Sir-mediated repression can occur independently of chromosomal and subnuclear contexts. Cell 119: 955– 67. Gaspar-Maia A, Alajem A, Polesso F, Sridharan R, Mason MJ, Heidersbach A, Ramalho-Santos J, McManus MT, Plath K, Meshorer E, et al. 2009. Chd1 regulates open chromatin and pluripotency of embryonic stem cells. Nature 460: 863–8. Gasser SM, 2002, Visualizing chromatin dynamics in interphase nuclei. Science 296: 1412– 1416. Gasser SM, Laroche T, Falquet J, Boy de la Tour E, Laemmli UK. 1986. Metaphase chromosome structure. Involvement of topoisomerase II. J Mol Biol 188: 613 –29. Gondor A, Ohlsson R. 2009. Chromosome cross-talk in three dimensions. Nature 461: 199 –205. Greaves IK, Rangasamy D, Ridgway P, Tremethick DJ. 2007. H2A.Z contributes to the unique 3D structure of the centromere. Proc Natl Acad Sci 104: 525–530. Grigoryev SA, Arya G, Correll S, Woodcock CL, Schlick T. 2009. Evidence for heteromorphic chromatin fibers from analysis of nucleosome interactions. Proc Natl Acad Sci 106: 13317– 22. Hagstrom KA, Meyer BJ. 2003. Condensin and cohesin: More than chromosome compactor and glue. Nat Rev Genet 4: 520 –534. Hake SB, Allis CD. 2006. Histone H3 variants and their potential role in indexing mammalian genomes: The “H3 barcode hypothesis.” Proc Natl Acad Sci 103: 6428– 35.

Hancock R. 2004a. A role for molecular crowding effects in the assembly and function of compartments in the nucleus. J Struct Biol 146: 281 –290. Hancock R. 2004b. Internal organisation of the nucleus: Assembly of compartments by macromolecular crowding and the nuclear matrix model. Biol Cell 96: 595–601. Hartl TA, Smith HF, Bosco G. 2008. Chromosome alignment and transvection are antagonized by condensin II. Science. 322: 1384–7. Heitz E. 1928. Das Heterochromatin der Moose, 1, Jahrb Wiss Bot 69: 762– 818. Heun P, Laroche T, Shimada K, Furrer P, Gasser SM. 2001. Chromosome dynamics in the yeast interphase nucleus. Science 294: 2181– 2186. Hines KA, Cryderman DE, Flannery KM, Yang H, Vitalini MW, Hazelrigg T, Mizzen CA, Wallrath L. 2009. Domains of HP1 Required for Drosophila melanogaster heterochromatin spreading. Genetics. 182: 967– 77. Hirano T. 2006. At the heart of the chromosome: SMC proteins in action. Nat Rev Mol Cell Biol 7: 311 –322. Hirano T, Mitchison. 1993. Topoisomerase II does not play a scaffolding role in the organization of mitotic chromosomes assembled in Xenopus egg extracts. J Cell Biol 120: 601–612. Horowitz RA, Agard DA, Sedat JM, Woodcock CL. 1994. The three dimensional archeitecture of chromatin in situ: Electron tomography reveals fibers composed of a continuously variable zigzag nucleosomal ribbon. J Cell Biol 125: 1 –10. Houchmandzadeh B, Marko JF, Chatenay D, Libchaber A. 1997. Elasticity and structure of eukaryote chromosomes studied by micromanipulation and micropipette aspiration. J Cell Biol 139: 1– 12. Hu Y, Kireev I, Plutz M, Ashourian N, Belmont AS. 2009. Large-scale chromatin structure of inducible genes: Transcription on a condensed, linear template. J Cell Biol 185: 87– 100. Huang B, Bates M, Zhuang X. 2009. Super-resolution fluorescence microscopy. Annu Rev Biochem 78: 993– 1016. Huebner MR, Spector DL. 2010. Chromatin dynamics. In Press. Jacobs SA, Khorasanizadeh S. 2002. Structure of the HP1 chromodomain bound to a lysine 9-methylated histone H3 tail. Science 295: 2080–2083. Jenuwein T, Allis CD. 2001. Translating the histone code. Science 293: 1074– 80. Jiang C, Pugh BF. 2009. Nucleosome positioning and gene regulation: Advances through genomics. Nat Rev Genet 10: 161– 72. Kadauke S, Blobel GA. 2009. Chromatin loops in gene regulation. Biochim Biophys Acta 1789: 17–25. Kireev I, Lakonishok M, Liu W, Joshi VN, Powell R, Belmont AS. 2008. In vivo immunogold labeling confirms largescale chromatin folding motifs. Nat Methods 5: 311– 313. Kireeva N, Lakonishok M, Kireev I, Hirano T, Belmont AS. 2004. Visualization of early chromosome condensation: A hierarchical folding, axial glue model of chromosome structure. J Cell Biol 166: 775– 785. Ko¨nig P, Braunfeld MB, Sedat JW, Agard DA. 2007. The three-dimensional structure of in vitro reconstituted

C.L. Woodcock and R.P. Ghosh Xenopus laevis chromosomes by EM tomography. Chromosoma 116: 349 –372. Kosak ST, Skok JA, Medina KL, Riblet R, Le Beau MM, Fisher AG, Singh H. 2002. Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296: 158– 62. Kouzarides T. 2007. Chromatin modifications and their function. Cell 128: 693–705. Kruhlak MJ, Celeste A, Nussenzweig A. 2006. Spatiotemporal dynamics of chromatin containing DNA breaks. Cell Cycle 5: 1910–1912. Kruithof M, Chien FT, Routh A, Logie C, Rhodes D, van Noort J. 2009. Single-molecule force spectroscopy reveals a highly compliant helical folding for the 30-nm chromatin fiber. Nat Struct Mol Biol 16: 534– 40. Li H, Motamedi MR, Yip C K, Wang Z, Waltz T, Patel DJ, Moazed D. 2009. An Alpha Motif at Tas3 C Terminus Mediates RITS cis Spreading and Promotes Heterochromatic Gene Silencing. Mol Cell 34: 155– 167. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289 –93. Levi V, Ruan Q, Plutz M, Belmont AS, Gratton E. 2005, Chromatin dynamics in interphase cells revealed by tracking in a two-photon excitation microscope. Biophys J 89: 4275–85. Liaw H, Lustig AJ. 2006. Sir3 C-terminal domain involvement in the initiation and spreading of heterochromatin. Mol Cell Biol 26: 7616–7631. Lohka MJ, Masui Y. 1983. Formation in vitro of sperm pronuclei and mitotic chromosomes induced by amphibian ooplasmic components. Science 220: 719– 721. Lu X, Wontakal SN, Emelyanov AV, Morcillo P, Konev AY, Fyodorov DV, Skoultchi AI. 2009. Linker histone H1 is essential for Drosophila development, the establishment of pericentric heterochromatin, and a normal polytene chromosome structure. Genes Dev 23: 452– 65. Luger K, Hansen JC. 2005. Nucleosome and chromatin fiber dynamics. Curr Opin Struct Biol 15: 188 –196. Luger K, Mader AW, Richmond RK, Sargent DF, Richmond TJ. 1997. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389: 251–60. Mani RS, Tomlins SA, Callahan K, Ghosh A, Nyati MK, Varambally S, Palanisamy N, Chinnaiyan AM. 2009. Induced chromosomal proximity and gene fusions in prostate cancer. Science 326: 1230. Marko JF. 2008. Micromechanical studies of mitotic chromosomes. Chromosome Res 16: 469– 497. Marko M, Hsieh C, Salmon N, Rodriguez M, Frank J, Manella C. 2008. FIB milling of frozen hydrated cells and tissues for TEM cryo-tomography. Microsc Microanal 13: 968CD. Marko M, Hsieh C, Schalek R, Frank J, Mannella C. 2007. Focused-ion-beam thinning of frozen-hydrated biological specimens for cryo-electron microscopy. Nat Methods 4: 215 –7. Maresca TJ, Heald R. 2006. The long and the short of it: Linker histone H1 is required for metaphase chromosome compaction. Cell Cycle 5: 589– 91.

Marshall WF. 2002. Order and disorder in the cell nucleus. Curr Biol 12: R185–R192. Marshall WF, Straight A, Marko JF, Swedlow J, Dernberg A, Belmont A, Murray AW, Agard DA, Sedat J. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930–939. Minton AP. 2001. The influence of macromolecular crowding and macromolecular confinement on biological reactions in physiological media. J Biol Chem 276: 10577– 10580. Misteli T. 2001. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Miyoshi D, Sigimoto N. 2008. Molecular crowding effects on structure and stability of DNA. Biochimie 90: 1040–1051. Moazed D, Bu¨hler M, Buker SM, Colmenares SU, Gerace EL, Gerber SA, Hong EJ, Motamedi MR, Verdel A, Ville´n J, Gygi SP. 2006. Studies on the mechanism of RNAidependent heterochromatin assembly. Cold Spring Harb Symp Quant Biol 71: 461–471. Muller WG, Rieder D, Kreth G, Cremer C, Trajanoski Z, McNally JG. 2004. Generic features of tertiary chromatin structure as detected in natural chromosomes. Mol Cell Biol 24: 9359– 9370. Muller WG, Walker D, Hager G, McNally J. 2001. Large-scale chromatin decondensation and recondensation regulated by transcription from a natural promoter. J Cell Biol 154: 33– 48. Nolis IK, McKay DJ, Mantouvalou E, Lomvaradas S, Merika M, Thanos D. 2009. Transcription factors mediate longrange enhancer-promoter interactions. Proc Natl Sci 106: 20222–7. Pederson T. 2000. Half a century of “the nuclear matrix.” Mol Biol Cell 11: 799– 805. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian nucleus. Nature 404: 604–609. Poirier MG, Marko JF. 2002. Mitotic chromosomes are chromatin networks without a mechanically contiguous protein scaffold. Proc Natl Acad Sci 99: 15393– 15397. Poirier MG, Marko JF. 2003. Micromechanical studies of mitotic chromosomes. Curr Top Dev Biol 55: 75–141. Poirier MG, Oh E, Tims HS, Widom J. 2009. Dynamics and function of compact nucleosome arrays. Nat Struct Mol Biol 16: 938 –44. Rando OJ, Zhao K, Janmey P, Crabtree GR. 2002. Phosphatidylinositol-dependent actin filament binding by the SWI/SNF-like BAF chromatin remodeling complex. Proc Natl Acad Sci 99: 2824– 2829. Richter K, Nessling M, Lichter P. 2008. Macromolecular crowding and its potential impact on nuclear function. Biochim Biophys Acta 1783: 2100–2107. Robinson PJ, Fairall L, Huynh VA, Rhodes D. 2006. EM measurements define the dimensions of the “30-nm” chromatin fiber: Evidence for a compact, interdigitated structure. Proc Natl Acad Sci 103: 6506– 6511. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287– 291. Routh A, Sandin S, Rhodes D. 2008. Nucleosome repeat length and linker histone stoichiometry determine chromatin fiber structure. Proc Natl Acad Sci 105: 8872–8877.

Chromatin Higher-order Structure and Dynamics Rusk N. 2009. When ChIA PETs meet Hi-C. Nature Methods 6: 863. Santenard A, Torres-Padilla ME. 2009. Epigenetic reprogramming in mammalian reproduction: Contribution from histone variants. Epigenetics 4: 80–4. Schermelleh L, Carlton PM, Haase S, Shao L, Winoto L, Kner P, Burke B, Cardoso MC, Agard DA, Gustafsson MG, et al. 2008. Subdiffraction multicolor imaging of the nuclear periphery with 3D structured illumination microscopy. Science 320: 1332– 1336. Schlach T, Duda S, Sargent DF, Richmond TJ. 2005. X-ray structure of a tetranucleosome and its implications for the chromatin fibre. Nature 436: 138– 141. Schu¨beler D, Francastel C, Cimbora DM, Reik A, Martin DI, Groudine M. 2000. Nuclear localization and histone acetylation: A pathway for chromatin opening and transcriptional activation of the human beta-globin locus. Genes Dev 14: 940 –50. Shelby RD, Hahn KM, Sullivan KF. 1996. Dynamic elastic behavior of alpha satellite domains visualized in living cells. J Cell Biol 135: 545–557. Simpson RT. 1991. Nucleosome positioning: Occurrence, mechanisms, and functional consequences. Prog Nucleic Acid Res Mol Biol 40: 143 –84. Shogren-Knaak M, Ishii H, Sun JM, Pazin MJ, Davie JR, Peterson CL. 2006. Histone H4-K16 acetylation controls chromatin structure and protein interactions. Science 311: 844– 847. Shopland LS, Lynch CR, Peterson KA, Thornton K, Kepper N, Hase J, Stein S, Vincent S, Molloy KR, Kreth G, et al. 2006. Folding and organization of a contiguous chromosome region according to the gene distribution pattern in primary genomic sequence. J Cell Biol 174: 27–38. Simpson RT, Thoma F, Brubaker J. 1985. Chromatin reconstituted from tandemly repeated cloned DNA fragments and core histones: A model system for study of higher order structure. Cell 42: 799 –808. Solovei I, Kreysing M, Lanctoˆt C, Ko¨sem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell. 137: 356 –68. Soutoglou E, Misteli T. 2007. Mobility and immobility of chromatin in transcription and genome stability. Curr Opin Genet Dev 17: 435 –4. Strick R, Strissel PL, Gavrilov K, Levi-Setti R. 2001. Cation-chromatin binding as shown by ion microscopy is essential for the structural integrity of chromosomes. J Cell Biol 155: 899 –910. Strukov YG, Belmont AS. 2009. Mitotic chromosome structure: Reproducibility of folding and symmetry between sister chromatids. Biophys J 96: 1617–28. Sullivan BA, Bickmore WA. 2000. Unusual chromosome architecture and behaviour at an HSR. Chromosoma 109: 181– 9.

Suto RK, Clarkson MJ, Tremethick DJ, Luger K. 2000. Crystal structure of a nucleosome core particle containing the histone variant H2A.Z. Nature Struct Biol 7: 1121–1124. Tremethick DJ. 2007. Higher-order structures of chromatin: The elusive 30 nm fiber. Cell 128: 651– 654. Trojer P, Reinberg D. 2007. Facultative heterochromatin: Is there a molecular signature. Mol Cell 28: 1 –12. Tumbar T, Sudlow G, Belmont AS. 1999. Large-scale chromatin unfolding and remodeling induced by VP16 acidic activation domain. J Cell Biol 145: 1341– 54. Vagnarelli P, Hudson DF, Ribeiro SA, Trinkle-Mulcahy L, Spence JM, Lai F, Farr CJ, Lamond AI, Earnshaw WC. 2006. Condensin and Repo-Man-PP1 co-operate in the regulation of chromosome architecture during mitosis. Nat Cell Biol 8: 1133– 1142. van Holde K. 1989. Chromatin. Springer Verlag, New York. van Holde K, Zlatanova J. 1995. Chromatin structure: Chasing a mirage? J Biol Chem 270: 8373–8376. van Holde K, Zlatanova J. 2007. Chromatin fiber structure: Where is the problem now? Sem Cell Dev Biol 18: 651– 658. Vazquez J, Bemont AS, Sedat JW. 2001. Multiple regimes of constrained chromosome motion are regulated in the interphase Drosophila nucleus. Curr Biol 11: 1227–1239. Verdel A, Jia S, Gerber S, Sugiyama T, Gygi S, Grewal SI, Moazed D. 2004. RNAi-mediated targeting of heterochromatin by the RITS complex. Science 303: 672– 676. Verschure PJ, van der Kraan I, Manders EM, Hoogstraten D, Houtsmuller AB, van Driel R. 2003. Condensed chromatin domains in the mammalian nucleus are accessible to large macromolecules. EMBO Rep 4: 861 –866. Visel A, Rubin EM, Pennacchio LA. 2009. Genomic views of distant-acting enhancers. Nature 461: 199– 205. Volpi EV, Chevret E, Jones T, Vatcheva R, Williamson J, Beck S, Campbell RD, Goldsworthy M, Powis SH, Ragoussis J, et al. 2000. Large-scale chromatin organization of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei. J Cell Sci 113: 1565–2576. Wilusz JE, Sunwoo H, Spector DL. 2009. Long non-coding RNAs: Functional surprises from the RNA world. Genes Dev 23: 1494–1504. Woodcock CL. 2006. Chromatin architecture. Curr Opin Struct Biol 16: 213 –220. Woodcock CL, Dimitrov S. 2001. Higher-order structure of chromatin and chromosomes. Curr Opin Genet Dev 11: 130 –135. Woodcock CL, Horowitz RA. 1995. Chromatin organization re-viewed. Trends Cell Biol 5: 272– 277. Woodcock CL, Skoultchi AI, Fan Y. 2006. Role of linker histone in chromatin structure and function: H1 stoichiometry and nucleosome repeat length. Chromosome Res 14: 17–25.

Nuclear Organization and Dosage Compensation Jennifer C. Chow and Edith Heard Mammalian Developmental Epigenetics Group, Institut Curie, CNRS UMR3215, INSERM U934, Paris, F-75248 France Correspondence: [email protected]

Dosage compensation is a strategy to deal with the imbalance of sex chromosomal gene products relative to autosomes and also between the sexes. The mechanisms that ensure dosage compensation for X-chromosome activity have been extensively studied in mammals, worms, and flies. Although each entails very different mechanisms to equalize the dose of X-linked genes between the sexes, they all involve the co-ordinate regulation of hundreds of genes specifically on the sex chromosomes and not the autosomes. In addition to chromatin modifications and changes in higher order chromatin structure, nuclear organization is emerging as an important component of these chromosome-wide processes and in the specific targeting of dosage compensation complexes to the sex chromosomes. Preferential localization within the nucleus and 3D organization are thought to contribute to the differential treatment of two identical homologs within the same nucleus, as well as to the chromosome-wide spread and stable maintenance of heterochromatin.

pecies that have evolved a chromosomally based sex determination system face the problem of dosage compensation of sex chromosomes between males and females. As the non-recombining chromosome degenerates, the heterogametic sex is left with only one functional copy of genes on the sex chromosomes compared with the double dose that exists in the homogametic sex. Diverse dosage compensation strategies have evolved in different species to deal with this problem and have been most extensively studied in flies (D. melanogaster), worms (C. elegans), and mammals. In flies, males (XY) up-regulate their single X to achieve expression levels equivalent to the two

S

copies present in females (reviewed in Gelbart and Kuroda 2009). Increased transcriptional output of the X chromosome is also observed in worms (Gupta et al. 2006) and mammals (Nguyen and Disteche 2006; Lin et al. 2007). However, in these latter cases, hyper-expression is not specific to the heterogametic sex, as it occurs in both males and females, necessitating secondary compensation mechanisms in females who would otherwise over-express Xlinked genes. In C. elegans, hermaphrodites (XX) down-regulate transcription from both their X chromosomes by half, whereas in female mammals, one of the two X chromosomes is chosen for inactivation (reviewed in Vicoso

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000604 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000604

J.C. Chow and E. Heard

and Bachtrog 2009). Therefore, it is through the co-ordinate regulation of hundreds of genes across the chromosome that these compensation mechanisms restore not only the balance between sexes but also the ratio of gene dosage between the sex chromosomes and the autosomes. How dosage compensation complexes are targeted specifically to the sex chromosomes on a chromosome-wide scale is still not fully understood in any of these organisms. Changes in chromatin structure and higher-order chromosome architecture clearly participate, but it is also becoming increasingly clear that in all of these dosage compensation systems, nuclear organization may play a role.

NUCLEAR ORGANIZATION AND DOSAGE COMPENSATION

The eukaryotic nucleus is a highly organized organelle consisting of multiple compartments. In addition to more stable structures, such as the nuclear envelope and the nucleolus, it also contains numerous specialized functional compartments, where processes such as RNA transcription and processing, as well as DNA replication and repair, take place (reviewed in Spector 2003; Misteli 2007). Chromosomes themselves also exhibit a distinct 3D organization and can adopt non-random positions within the nucleus, both with respect to each other and with respect to specific nuclear landmarks. It is becoming clear that this spatial organization likely plays an important role in the regulation and expression of the genome and is believed to be a key factor in the chromosomewide regulation that occurs during dosage compensation. In particular, the interphase organization of X-chromosome sequences may facilitate their differential activity compared to autosomes or between the sexes. Similarly, preferential localization in specialized nuclear subcompartments may contribute to specific processes that affect the sex chromosomes more than the autosomes. In the case of mammals, nuclear compartmentalization is likely to facilitate the differential treatment of the two X chromosomes within the same nucleus.

Drosophila melanogaster

In Drosophila, dosage compensation is mediated by a ribonucleoprotein complex that binds hundreds of sites on the single X chromosome in males, and induces a two-fold up-regulation of X-chromosomal genes (Fig. 1A). The complex, known as the male-specific lethal (MSL) complex, is made up of five protein subunits (MSL1, MSL2, MSL3, MOF, MLE) and two non-coding RNAs (roX1, roX2) (reviewed in Gelbart and Kuroda 2009; Hallacli and Akhtar 2009). It can only form in male cells, due to the male-specific expression of the male-specificlethal 2 (MSL2) protein. MSL2 is required to stabilize components of the complex such as MSL1 and roX RNAs. MOF and MLE are enzymatic components, MLE being an RNA/DNA helicase and MOF a histone acetyltransferase, responsible for acetylation of histone H4 at lysine 16 (H4K16Ac). The H4K16Ac chromatin modification is believed to be a key component in the transcriptional up-regulation across the X chromosome in males (Bone et al. 1994). MSL1 and MSL2 form the core of the complex and are thought to be the main targeting components; however, the full complex is required for complete localization to the X chromosome (Gelbart and Kuroda 2009). Targeting to the X chromosome is thought to occur through co-transcriptional assembly of the complex at the sites of roX1 and roX2 transcription on the X chromosome (Kelley et al. 1999). From these assembly sites, the complex then spreads to other high affinity binding sites on the X or chromatin entry sites (CES), and is thought to occur, at least partially, through the recognition of specific DNA sequence motifs (Alekseyenko et al. 2008; Straub et al. 2008) (Fig. 1A). Nevertheless, the identified DNA motifs alone are unable to unequivocally predict MSL complex binding since many motifs are not associated with MSL binding; therefore, other factors must also be involved. It has been proposed that nuclear organization and higher order chromatin structure may play a role in this regard, perhaps facilitating the spread of the complexes to the actively transcribed portions of the chromosome. In fact, the X chromosome has been found

Nuclear Organization and Dosage Compensation

A

D. melanogaster

Males: Two-fold up-regulation of single X chromosome

Females

mRNA more rapidly processed or exported?

Nuclear pore

Nuclear pore

MSL complex

X

roX1

Clustering of MSL complexbound high affinity sites may aid in the spread to lower affinity sites

roX2

Y High affinity sites Low affinity sites Lowest affinity sites

roX RNA

roX 1,2 loci

Transcriptionally active genes roX 1 and 2 high affinity chromatin entry sites High affinity sites Low affinity sites Lowest affinity sites MSL complex H4K16 acetylated chromatin

B

C. elegans

Hermaphrodites: Down-regulation of both X chromosomes by half Nuclear membrane

Males Nuclear membrane

DCC complex

X X High affinity sites (rex sites) Low affinity sites (dox sites)

C

DCC-binding to high affinity sites and spread to low affinity sites leads to global chromosome-wide repression

Mammals

Females Inactive X

Xi Xa

Xic Xic

High affinity sites (rex sites) Low affinity sites (dox sites) Genes Dosage compensation complex

Inactivated genes are recruited into the compartment

Unknown Xist RNA “entry” sites (some genes?)

Genes that Waystations/booster elements (repeats, including LINEs - facilitators of repetitive escape remain Active LINE1 core formation) elements facilitate external Waystations/booster elements (young, active LINEs) silencing

Nuclear membrane

Active X

Xist RNA Xist RNA creates a silent compartment made up of silent LINE1 elements and other repeats

RNA Pol II

Figure 1. Diverse strategies for dosage compensation in different organisms. (A) In D. melanogaster, males (XY)

up-regulate two-fold their single X chromosome. The dosage compensation complex has been found to be associated with nuclear pore proteins, which may indicate a participation of nuclear localization in this upregulation. (B) C. elegans hermaphrodites (XX) down-regulate both X chromosomes by one half. This may be linked to a particular organization of the two Xs with the dosage compensation machinery. (C) Female mammals inactivate one of their two X chromosomes. The X chromosomes often reside at the nuclear periphery or at the nucleolus. The inactive X is organized as a silent, repetitive compartment by Xist RNA, into which genes are located as they become silenced.

to adopt a male-specific 3D conformation with “clustering” of the high affinity MSL complex binding sites occurring in males, but not females (Grimaud and Becker 2009). In addition, MSL proteins co-purify with components of the nuclear pore complex, Nup153 and Megator (Mtor), and RNAi knockdowns have shown

that these factors are essential for Msl-mediated dosage compensation in male cells (Mendjan et al. 2006). Although the nuclear periphery is often associated with heterochromatin and transcriptional silencing, there are sub-regions at the nuclear envelope, particularly at nuclear pores, that are associated with transcriptional

J.C. Chow and E. Heard

activity (Taddei et al. 2004; Akhtar and Gasser 2007). Recently, the genome-wide chromatin binding profiles of Nup153 and Megator have been reported (Vaquerizas et al. 2010). These Nucleoporin-Associated Regions (NARs) represent regions of open chromatin as they are enriched in markers for active transcription such as RNA polymerase II and histone H4K16 acetylation. Interestingly, the male hyperactive X chromosome is particularly enriched in NARs and relies on the interaction with nucleoporins for its peripheral localization (Vaquerizas et al. 2010). The link between the MSL complex, nuclear pore components, and the nuclear periphery suggests that the X chromosome in males specifically localizes to nuclear sub-domains through its interactions with the nuclear pore and that this might contribute to its two-fold up-regulation. Thus, the enhanced expression of the X chromosome in males may be not only transcriptional via chromatin-mediated effects, but also posttranscriptional, perhaps through the more rapid processing and export of messenger RNA, thanks to the association with the nuclear pore. Caenorhabditis elegans

In C. elegans, dosage compensation involves a two-fold reduction in X-chromosomal gene activity in hermaphrodites (Fig. 1B). The worm dosage compensation complex (DCC) consists of a hermaphrodite-specific core of SDC (sex determination and dosage compensation) proteins, required for the stabilization and targeting of the complex to the X chromosome, as well as a condensin subcomplex, which somehow induces the chromatin changes necessary for dosage compensation. How the DCC imposes a precise two-fold down-regulation of X-linked gene expression in a chromosomewide manner remains unclear. The DCC is targeted to X chromosomes by initial binding to a number of recruiting elements, followed by dispersal or spreading to secondary sites. However, DCC binding does not correlate with DCCmediated repression, implying that the complex acts in a chromosome-wide manner, rather than on a gene-by-gene basis. Indeed, it has been

proposed that the DCC does not repress genes by direct binding, but rather acts at a distance and may involve looping of chromatin fibers to bring regulated genes to the vicinity of DCC bound sites (Jans et al. 2009). The implication of condensins suggests that X-chromosome down-regulation may involve similar mechanisms to mitotic chromosome condensation (Chuang et al. 1996; Lieb et al. 1996; Lieb et al. 1998; Tsai et al. 2008; Csankovszki et al. 2009). The DCC may alter higher order chromosome organization by inducing partial chromosome condensation at interphase, and this might in turn lead to decreased transcription. This is supported by the fact that the DCC condensin subunit differs from the mitotic condensin I complex by only a single subunit. Although the exact molecular mechanisms of condensin-mediated X chromosome down-regulation are still unclear, the consensus is that this process must involve some higher-order reorganization of the X chromosomes within the nucleus. Whether nuclear compartmentalization or localization also plays a role remains to be seen. NUCLEAR ORGANIZATION OF THE INACTIVE X CHROMOSOME IN FEMALE MAMMALS

In mammals, the presence of an inactive X (Xi) chromosome in female cells was first noted at the cytological level by Barr in 1949 as a heteropycnotic body that was often at the nuclear periphery or within the perinucleolar region (Barr and Bertram 1949). These initial observations were already suggestive of a possible role for nuclear compartmentalization in the formation/maintenance of the inactive state (Comings 1968). The nuclear and perinucleolar compartments are indeed often associated with heterochromatin (de Wit and van Steensel 2009), and in the case of the mammalian X chromosome, this might promote the inactive state by facilitating access to heterochromatic factors or restricting access to transcription factors. The nucleolar localization of the Xi (Zhang et al. 2007; Rego et al. 2008) has been proposed to be important for faithful replication of the Xi’s epigenetic state, as the nucleolar

Nuclear Organization and Dosage Compensation

periphery is enriched in factors required for replication of heterochromatin, such as Snf2h (Zhang et al. 2007). In addition to its preferential locations in the nucleus, the inactive X chromosome also appears to adopt a very distinctive 3D organization. In both mouse and humans, fluorescence in situ hybridization (FISH) studies have shown that the inactive X chromosome is coated by the non-coding Xist RNA, which triggers the initiation of X inactivation. Xist induces global changes in chromatin structure, and also mediates changes in the global organization of the chromosome, both of which are thought to play roles in chromosome-wide silencing. Xist RNA initially coats a central core of the X-chromosome territory that is made up largely of repetitive DNA (Chaumeil et al. 2006; Chow et al. 2010). X-linked genes are mostly excluded from this inner region, particularly at the early stages of inactivation and instead reside outside, or at the periphery, of this repetitive compartment (Chaumeil et al. 2006). However, as X inactivation proceeds and genes become silenced, they move into the Xist RNA coated, silent compartment. There is thus a progressive nuclear reorganization of the X chromosome as it becomes inactivated and heterochromatic, ultimately resulting in a condensed structure with a repetitive core and silent genes embedded into its periphery (Clemson et al. 2006) (Figs. 1C, 2A). The degree to which this particular 3D organization of the Xi is due to a compartmentalizing function of Xist RNA, or to the chromatin changes that are induced during X chromosome inactivation (XCI) or to the repetitive nature of the mammalian X chromosome, still remains to be seen. HETEROGENEOUS HETEROCHROMATIN ON THE INACTIVE X CHROMOSOME

Inactivation of the X chromosome involves the acquisition of many chromatin changes that clearly distinguish it from its active homolog. Immunofluorescence studies in mitotic and/or interphase cells show that several histone marks typical of heterochromatin appear to be enriched on the Xi, including H3K27me3, H3K9me2, H3K9me3, H4K20me1, H2AK119Ub, and the

histone variant macroH2A (reviewed in Heard and Disteche 2006). Conversely, euchromatic marks such as H3K4me2/3 and H3 and H4 acetylated lysines are largely depleted. However, these modifications are not uniformly distributed across the chromosome, suggesting that the inactive X chromosome is made up of several different types of heterochromatin. In human somatic cell lines, at least two different heterochromatic states have been characterized by immunofluorescence: One is defined by the presence of XIST RNA, macroH2A, H3K27me3, H4K20me1, and H2AK119Ub, and the other is defined by features more reminiscent of constitutive heterochromatin, including later replication timing, H3K9me3, H4K20me3, and association with HP1. These different heterochromatin domains appear to remain spatially distinct from each other, both during metaphase and interphase (Chadwick and Willard 2004; Chadwick 2007). Ultra-structural analyses of the H3K27me3-enriched portion of the Xi by light and electron microscopy in mouse and human fibroblasts (Rego et al. 2008) have shown that this portion of the heterochromatic X has a distinctive structure, being made up of densely packed fibers with intervening spaces and a mean chromatin compaction higher than euchromatin, but less than constitutive, H3K9me3-enriched heterochromatin (Fig. 2B, C). The significance of these different types of heterochromatin and whether they reflect differences in the epigenetic mechanisms underlying the initiation and/or maintenance of the inactive state is still unknown. The relative proportions of these two heterochromatic signatures varies considerably between species. In mouse, enrichment of HP1 and H3K9me3 on the Xi is much less distinctive than has been described in humans (Peters et al. 2002; Kohlmaier et al. 2004). Nevertheless, it should be noted that in both of these species, the inactive X appears to be similarly organized in the nucleus, with genes surrounding a silent repetitive core (Chaumeil et al. 2006; Clemson et al. 2006). In marsupials, although the Xi is clearly depleted for active marks as on the mouse and human Xi (Koina et al. 2009; Mahadevaiah et al. 2009), the degree of enrichment on the Xi

J.C. Chow and E. Heard

A

5 μm

Xi

Xi

Xi

Xi *Xa

*Xa

1 μm

*Xa

LINE1 DNA Lamp2,G6pdX, AtrX,Fmr1, Huwe1 DNA

LINE1 DNA Xist RNA

LINE1 DNA Lamp2,G6pdX, AtrX,Fmr1, Huwe1 DNA Xist RNA

B

nu

1 μm

2 μm 2 μm

C Nu

1 μm S11

S11

Figure 2. Nuclear organization of the inactive X chromosome in female mammals. (A) Combined RNA-DNA

fluorescence in-situ hybridization in day 8 differentiated female ES cells shows the distinctive organization of the silent inactive X (Xi) compared to the active X (Xa). Both the Xa and the Xi often reside at the periphery of the nucleus. However, on the Xi, Xist up-regulation and coating results in the formation of a transcriptionally silent compartment made up of a central core of repetitive sequences, including LINE1 elements. Genes transcriptionally silenced at this stage (Lamp2, G6pdX, AtrX, Fmr1, and Huwe1) have been recruited into the compartment. All images shown are single slices taken from the same Z plane. (B,C ) Electron micrographs of single 200nm sections from female WI-38 human fibroblasts. The inactive X chromosome (arrow) has an ultrastructure distinct from the chromocenters (arrowheads), adopting a looser, less compact organization. Adapted, with permission, from Rego et al. 2008, Journal of Cell Science.

for histone modifications characterizing heterochromatin(H3K9me2,H3K27me3,H3K9me3, H4K20me3) is less clear-cut, with some studies reporting no enrichment on the Xi at metaphase (Koina et al. 2009) and others reporting some enrichment of H3K27me3 at interphase (Mahadevaiah et al. 2009). Given that marsupials lack the XIST gene (Duret et al. 2006), differences in histone modification patterns on the

Xi might be expected, since Xist is required for the recruitment of many of the chromatin marks in mouse. The situation in monotremes appears to be even more complex with females having five pairs of XX chromosomes and males having five XY pairs. Although there does appear to be some degree of dosage compensation, no systematic analysis has been performed to determine the heterochromatic status of the

Nuclear Organization and Dosage Compensation

X chromosomes in females, and dosage compensation appears to be largely incomplete and variable between genes (Grutzner et al. 2004; Rens et al. 2004; Deakin et al. 2008). A more detailed view of the chromatin content of the eutherian inactive X chromosome is starting to be obtained by higher resolution epigenomic analyses, such as ChIP-chip and allele-specific ChIP (Brinkman et al. 2006; Valley et al. 2006; Valley and Willard 2006; Chadwick 2007; Marks et al. 2009; Mietton et al. 2009). ChIP-chip for macroH2A1 in mouse liver cells reveals an approximately 1.5-fold enrichment in females versus males. As this enrichment is distributed uniformly across the entire X chromosome, this implies that macroH2A1 may influence global chromatin structure, rather than directly inhibiting transcription at promoters of genes (Mietton et al. 2009). The distribution of H3K27me3 has also been examined on the inactivating X chromosome in differentiating mouse ES cells by ChIP-Seq (Marks et al. 2009). Allele-specific mapping of ChIP-seq tags showed that the H3K27me3 mark is deposited on the inactive X chromosome in female ES cells, particularly in gene-rich, active regions, which is in agreement with the results observed in human somatic cells by immunofluorescence (Chadwick 2007; Marks et al. 2009). DNA methylation of promoters of X-linked genes is another hallmark of XCI in eutherian somatic cells. Global microarray analyses involving methylated DNA immunoprecipitation (MeDIP) were used to assess the DNA methylation status of the Xi relative to the active X (Xa) in human primary cells. This study revealed that although CpG islands are hypermethylated on the Xi, the overall levels of methylation on the X chromosome are in fact lower in females compared to males, especially in gene-poor regions (Weber et al. 2005). Hellman and Chess (2007) analyzed the DNA methylation status of more than 1000 X-linked loci and found that on the active X, DNA methylation is concentrated in gene bodies, confirming previous studies (for review, see Heard et al. 1997). Although the function of methylation within transcribed genes is unknown, it is a general feature of active

genes in many organisms (for example, Zhang et al. 2006). The lack of gene-body DNA methylation on the inactive X chromosome at least partly explains its globally hypomethylated status, with only promoters of genes being hypermethylated. In summary, the inactive X chromosome contains many different epigenetic marks, with some variations in distribution between different mammals. The degree to which the different signatures on the Xi reflect differences in the higher-order structure and/or nuclear organization of the inactive X should become more clear with the advent of techniques such as chromosome conformation capture, which measures physical interaction within the chromatin fiber (Simonis et al. 2007; van Berkum and Dekker 2009). INITIATING X INACTIVATION: XIST RNA REGULATION AND FUNCTION

The evolution of the Xist gene is believed to have been a key event in the evolution of the stable, chromosome-wide X-inactivation process that exists in eutherian mammals. Xist RNA coating of the prospective inactive X chromosome appears to be the key trigger for the X-inactivation cascade. Deletion analyses have shown that Xist is essential for silencing and likely has a role at multiple levels in the process (Penny et al. 1996; Marahrens et al. 1997; Csankovszki et al. 1999; Wutz et al. 2002; Hoki et al. 2009). The regulation of Xist during development is a complex process, ensured by multiple long-range cis elements and trans-acting factors. Xist is in fact located in a region of the X chromosome known as the X-inactivation center (Xic) that ensures the appropriate initiation of X inactivation in female cells (reviewed in Barakat et al. 2010) (Fig. 3). The Xic contains sequence elements that ensure that Xist is only up-regulated in XX and not XY cells. The Xic also ensures that in XX cells, Xist up-regulation occurs on only one and not both X chromosomes. Recent insights into this process have revealed that the presence of two or more copies of certain X-linked loci can activate Xist. This may work at several levels. For example, a double dose of the X-linked Rnf12

J.C. Chow and E. Heard Xic mediated initiation of XCI: • Sensing and counting X chromosome number: trigger Xist up-regulation • Choosing one X for inactivation

Undifferentiated female ES cells

Monoallelic Xist up-regulation

Xic

Xa Xa Pluripotency factors Rnf12

Xpr

+

Xist Tsix Xite

Rnf12

Xpr

?+

+

Pluripotency factors

= trans interactions

Xist

Xist

Tsix Xite

Rnf12

Xpr

Tsix Xite

?–

–

= Xist RNA

Differentiated female ES cells

• Xist RNA accumulation and coating of the X in cis: exclusion of RNA pol II, silencing of repeats

Xa

Xi

• Appearance of macro H2A on Xi • Smchd1 recruitment to Xi? • DNA methylation of promoters of X-linked genes (hypomethylation of gene bodies)

• Histone modifications enriched on Xi (H3K9me2, H3K27me3, H4K20me1, etc) • Shift to late replication timing • Young LINE expression on Xi • Progressive spread of silencing onto all parts of the X (facilitated by LINEs?)

• Xist RNA silent compartment • Loss of active histone marks (H3K4me2/me3, H3Ac, H4Ac) • Gene silencing and relocation into Xist RNA compartment (via SATB1/2?) • Recruitment of PcG complexes

Spread and progressive locking in of heterochromatic state

Figure 3. Random X chromosome inactivation is a multi-stage process. During the initiation stage, develop-

mental factors as well as a complex interplay of elements within the X inactivation center (Xic) ensure that monoallelic Xist up-regulation occurs only when more than one X chromosome is present per diploid autosome set. Once up-regulated, Xist RNA coats the chromosome in cis and triggers chromosome-wide gene silencing. The spread of silencing occurs progressively with the acquisition of many different epigenetic changes that together serve to create an extremely stable, heterochromatic state.

gene, which encodes a ubiquitin ligase, can trigger Xist up-regulation (Jonkers et al. 2009). Trans interactions ( pairing) between Xics may also be involved in triggering the expression of Xist and in ensuring its monoallelic regulation in female cells (Bacher et al. 2006; Xu et al. 2006; Augui et al. 2007). These transient Xic pairing events may facilitate the coordination of reciprocal Xist expression patterns on

the future active and inactive X chromosomes during the initiation of X inactivation (Augui et al. 2007; Nicodemi and Prisco 2007; Xu et al. 2007). Thus, the initiation of monoallelic Xist expression in female cells involves a dynamic series of events at the Xic loci, integrating developmental triggers, dosage-dependent proteins factors, interactions in trans, and noncoding RNAs (Fig. 3).

Nuclear Organization and Dosage Compensation

Once Xist RNA has become monoallelically up-regulated (at around day 2 in differentiating female ES ells), it begins the process of chromosome-wide gene silencing. Xist RNA induces X inactivation during an early differentiation time window, and recent studies have implicated the SATB1 and SATB2 (Special AT-rich Binding Protein) proteins in enabling Xist function during this early differentiation time window and also in some cancer cells (Agrelo et al. 2009). The 17,000–19,000nt-long, spliced, poly-adenylated Xist transcript consists of multiple different functional domains (Wutz et al. 2002). The most conserved region of Xist is a set of tandem repeats at the 50 end of the gene, called the Arepeats, that are required for its gene silencing function (Wutz et al. 2002; Hoki et al. 2009). Inducible Xist transgenes deleted for A-repeats are unable to induce gene silencing, but can still coat the chromosome in cis and recruit polycomb group proteins, which are involved in the recruitment of repressive chromatin changes including H3K27me3 to the X chromosome (Wutz et al. 2002; Plath et al. 2003; Kohlmaier et al. 2004). Thus, other portions of Xist RNA are required for its capacity to coat the X chromosome in cis and to recruit polycomb and macroH2A (Wutz et al. 2002). The Xist A-repeat region also produces an independent 1.6-kb transcript, RepA, and although its function is not entirely clear, this transcript may play a role in regulating Xist itself (Zhao et al. 2008; Hoki et al. 2009). Although Xist RNA may be involved in directly or indirectly recruiting chromatin-modifying enzymes, including polycomb group proteins, so far there is no evidence that this accounts for its gene-silencing role. However, there is increasing evidence pointing to Xist RNA having an architectural role that may be important for the specific 3D-organization of the inactive X (Chaumeil et al. 2006; Clemson et al. 2006; Rego et al. 2008), which may impact both the initiation and maintenance of the inactive state. One of the earliest events following Xist up-regulation and coating of the future inactive X chromosome is the formation of a nuclear compartment that is depleted for RNA polymerase, transcription factors, and

euchromaticmarks(Chaumeiletal.2006).Based on the analysis of Xist and PcG mutant ES cell lines, gene silencing and the recruitment of polycomb group proteins can be functionally dissociated from the formation of the Xist silent nuclear compartment (Plath et al. 2003; Kohlmaier et al. 2004; Chaumeil et al. 2006; Schoeftner et al. 2006). The silent compartment initially encompassed by Xist RNA is in fact largely constituted of X-chromosome repeat sequences, rather than genes. Prior to their inactivation, X-linked genes actually reside outside or at the periphery of this Xist RNA compartment. As differentiation proceeds and X inactivation occurs, genes move into this Xist RNA domain as they become silenced (Chaumeil et al. 2006). The mechanisms underlying this relocalization of genes are currently unknown. The Xist A-repeat region, which is essential for gene silencing, may have a role since genes do not become relocated into the silent repetitive compartment in Xist A-repeat mutants. Furthermore, given the implication of the SATB1 and SATB2 factors in Xist RNA’s competence to induce gene silencing during early differentiation (Agrelo et al. 2009), and the fact that these DNA-binding proteins facilitate the organization of chromatin structure during T-cell development (Cai et al. 2006), they may similarly play a role in gene relocation into the silent Xist repetitive compartment during X inactivation. So far, no physical association between Xist A-repeats and SATB1/2 has been demonstrated, but their window of action and their known function in nuclear organization make them key candidates for a role in Xist-mediated gene silencing and relocation. Importantly, recent findings demonstrate that Xist RNA can actually induce gene silencing outside an early developmental context, in some cancer cells (Hall et al. 2002; Chow et al. 2007), and in adult progenitor stem cells (Savarese et al. 2006). Agrelo et al. (2009) show that this reacquired capacity for Xist-induced silencing in tumor cells is associated with the re-expression of the SATB1 protein. This finding reinforces the notion that the nuclear and epigenetic plasticity found in some cancer cells may be similar to that found during early embryogenesis.

J.C. Chow and E. Heard

SPREADING INACTIVATION ACROSS THE X CHROMOSOME

Early studies using inducible Xist genes in male ES cells demonstrated that Xist had a developmentally restricted function, limited to the first few days of differentiation (Wutz and Jaenisch 2000). X-inactivation was thus originally thought to be a rather concerted process, occurring in the space of just a few cell divisions. However, recent studies in mouse pre-implantation embryos (Patrat et al. 2009) and differentiating female ES cells (Lin et al. 2007; Chow et al. 2010) have revealed that inactivation of the X chromosome in its entirety can take much longer. Some genes are silenced very quickly (in a matter of days or even hours after Xist RNA accumulation), while others are silenced much later (days, or .1 week), well outside of the time window in which Xist RNA is thought to act (Lin et al. 2007; Patrat et al. 2009). This suggests either that Xist’s silencing function is not limited to early differentiation for some loci, or else that the changes induced by Xist are propagated more slowly into some regions compared to others. Such heterogeneity suggests that the kinetics of inactivation are controlled by regionspecific processes, presumably due to different sequence environments. In addition, several X-linked genes are known to escape the X-inactivation process either partly or fully (Yang et al. 2010). Some of these genes (e.g., Jarid1c/Kdm5c) have the capacity to escape from X inactivation, whatever their location on the X chromosome (Li and Carrel 2008), implying that they have specific features preventing them from being efficiently silenced. Importantly, all escapees tend to be located outside the Xist RNA compartment, suggesting that part of the mechanism enabling escape may be linked to their capacity to resist internalization into the silent, repetitive compartment. However, the exact sequence and/or chromatin signatures that underlie the capacity of some genes to escape and those that underlie the variability in X-inactivation kinetics between genes, are still not well understood. Identifying such genomic and/ or chromatin features has turned out to be a challenge since multiple actors are likely to be

at work and account for the diversity of X-inactivation profiles. Genes that can escape from X inactivation may be flanked by insulator elements that help form domains that prevent the spread of heterochromatinization and/or recruitment of a gene into the silent repetitive Xist compartment. CTCF-binding sites have been hypothesized to play such a role as they have been identified between inactivation and escape domains (Filippova et al. 2005). Sequences that facilitate the propagation of X inactivation have also been proposed to exist, termed “way-stations” or “booster elements” (Gartler and Riggs 1983). Lyon proposed that possible candidates for such booster elements could include non-LTR retrotransposon LINE1 repeats, as they are enriched on the X chromosome and their density correlates well with the spread of silencing into autosomal sequences in X: autosome translocations (Lyon 1998). Furthermore, LINE1 elements are depleted in the immediate vicinity of escape genes (Bailey et al. 2000; Carrel et al. 2006). More recently, it has been shown that LINEs may facilitate the propagation of X inactivation at at least two different levels. Truncated, ancient LINEs may participate in nucleating the heterochromatic repetitive compartment induced by Xist RNA early in X inactivation (Chow et al. 2010). On the other hand, younger, more active LINEs, which are actually transcribed from the inactive X during development, may participate in the local propagation of X inactivation into regions that would otherwise be prone to escape (Chow et al. 2010). One such region that has been studied in detail contains the Huwe1 gene, which has a full length LINE at its 30 end. Huwe1 displays very slow X-inactivation kinetics, initiating only after day 2 of differentiation (Patrat et al. 2009; Chow et al. 2010). The slow silencing of Huwe1 may be due to its proximity to an escapee, Jarid1c, which might render its surrounding region, including Huwe1, more resistant to X inactivation. Huwe1 is, however, eventually inactivated, possibly due to the presence of nearby fulllength LINE1 elements. Intriguingly, these full length LINEs are transcribed from the inactive X chromosome during the exact window of time that X inactivation is occurring and their

Nuclear Organization and Dosage Compensation

transcription is accompanied by the production of small RNAs (Chow et al. 2010) (Fig. 4). In the case of Huwe1, it has been proposed that transient LINE1 transcription may drive antisense transcription into the gene and this may help to induce its silencing. Antisense transcription has increasingly emerged as an important regulator of gene expression in a diverse number of ways (Su et al. 2010). Active LINE1 elements/ promoters may therefore represent a significant source of antisense transcription and play regulatory roles in the genome, particularly on the X chromosome, where they are enriched, at least in eutherian mammals. In summary, LINEs may participate in generating heterochromatin in more than one way, with young, full-length

LINEs acting as local boosters to facilitate silencing of certain regions of the X chromosome; for example, by driving the transient production of natural antisense transcripts on the X chromosome undergoing inactivation; while older, truncated LINEs may facilitate creation of the silent repetitive core of the X chromosome, into which genes are recruited as they are inactivated (Chaumeil et al. 2006; Chow et al. 2010). Although it is clear that Xist RNA is essential for the recruitment and propagation of chromatin modifications as well as the specific reorganization of the chromosome, how these changes result in later changes, such as the shift to late replication timing, macroH2A enrichment,

Day 2 differentiation

Undifferentiated ES cells (Active X)

Xist up-regulation triggers the formation of a transcriptionally silent repetitive compartment

Genes

Day 4-8 differentiation Repeats Full length active L1 elements Recruitment of silencing factors

Huwe1

Xist RNA

Heterochromatinization? Methylation?

Active L1 element

kdm5C/Jarid1C

Spread of silencing? Small RNA pathway? Spread?

Huwe1

Active L1 element

Some regions are more resistant to silencing and are subject to slower silencing kinetics (e.g., near escape genes?) Differentiated ES cells (Inactive X)

Boundary elements

Kdm5C/Jarid1C Full length L1 elements may help to spread silencing into these more resistant regions.

Figure 4. Diverse, region-specific elements contribute to the spread and stability of silencing and result in

the heterogeneous inactivation kinetics observed across the chromosome. The X-linked Huwe1 gene is Xinactivated relatively late during development. This resistance to silencing may be due to the influence of the nearby escapee domain containing the Jarid1C/Kdm5c gene. The presence of a full length LINE1 element 30 of Huwe1 may help to eventually spread silencing into the Huwe1 gene.

J.C. Chow and E. Heard

and DNA methylation at promoters, remains unclear. A recent new player in X inactivation that may be involved in some of the downstream changes is the Smchd1 protein (Blewitt et al. 2008). Smchd1 mutant female embryos display both placental and extra-embryonic defects and die prior to stage E13.5. This relatively late lethality is suggestive of a role in the maintenance, rather than the initiation of XCI. Indeed, promoter DNA hypomethylation on the Xi was detected in mutants, suggesting that Smchd1 may affect the DNA methylation deposition step (Blewitt et al. 2008). Smchd1 is an SMC-like protein with homology to components of cohesin and condensin complexes. Interestingly, condensins are implicated in the down-regulation of both X chromosomes in C. elegans hermaphrodites (Chuang et al. 1994), providing an exciting potential link between these two forms of dosage compensation.

and chromatin patterns found for different regions of the inactive X chromosome suggest that there may be more than one way of inducing heterochromatin during X inactivation. Furthermore, different mammals exploit different mechanisms to achieve chromosome-wide silencing, with variations in chromatin marks. The extent to which nuclear organization and spatial segregation of the X chromosomes represent a universal feature of dosage compensation strategies will remain an interesting question for the future. ACKNOWLEDGMENTS

Our work is supported by grants from the CNRS, FRM (Equipe FRM), ANR, and Canceropole (GepiG), as well as HEROIC (Highthroughput Epigenetic Regulatory Organization in Chromatin), an integrated project funded by the European Union, and the ERC.

CONCLUSIONS AND PERSPECTIVES

Although very different mechanisms have evolved in different organisms to ensure chromosomewide gene regulation during dosage compensation, a common theme is the necessity for recruitment of dosage compensation complexes across the length of the X chromosome in a concerted fashion. In flies and worms, specific recruitment or entry sites are beginning to be identified, and should help to unravel the exact mechanisms by which the dosage compensation complexes are recruited and spread in these organisms. However, chromatin structure, noncoding RNAs, and nuclear organization may also play important roles. Mammals present the unusual situation whereby two genetically identical X chromosomes have to be differentially treated within the same nucleus and nuclear organization may contribute to this process. Coating of one of the two X chromosomes by the non-coding Xist RNA triggers formation of a silent nuclear compartment, into which genes are recruited as they become inactivated, as well as the recruitment of repressive complexes. The cis-limited action of Xist RNA still remains mysterious, as do the proteins or sequences to which it binds. The diversity of X-inactivation kinetics

REFERENCES Agrelo R, Souabni A, Novatchkova M, Haslinger C, Leeb M, Komnenovic V, Kishimoto H, Gresh L, Kohwi-Shigematsu T, Kenner L, et al. 2009. SATB1 defines the developmental context for gene silencing by Xist in lymphoma and embryonic cells. Dev Cell 16: 507 –516. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507– 517. Alekseyenko AA, Peng S, Larschan E, Gorchakov AA, Lee OK, Kharchenko P, McGrath SD, Wang CI, Mardis ER, Park PJ, et al. 2008. A sequence motif within chromatin entry sites directs MSL establishment on the Drosophila X chromosome. Cell 134: 599 –609. Augui S, Filion GJ, Huart S, Nora E, Guggiari M, Maresca M, Stewart AF, Heard E. 2007. Sensing X chromosome pairs before X inactivation via a novel X-pairing region of the Xic. Science 318: 1632– 1636. Bacher CP, Guggiari M, Brors B, Augui S, Clerc P, Avner P, Eils R, Heard E. 2006. Transient colocalization of Xinactivation centres accompanies the initiation of X inactivation. Nat Cell Biol 8: 293–299. Bailey JA, Carrel L, Chakravarti A, Eichler EE. 2000. Molecular evidence for a relationship between LINE-1 elements and X chromosome inactivation: the Lyon repeat hypothesis. Proc Natl Acad Sci U S A 97: 6634– 6639. Barakat TS, Jonkers I, Monkhorst K, Gribnau J. 2010. X-changing information on X inactivation. Exp Cell Res 316: 679–687. Barr ML, Bertram EG. 1949. A morphological distinction between neurones of the male and female, and the behaviour of the nucleolar satellite during accelerated nucleoprotein synthesis. Nature 163: 676.

Nuclear Organization and Dosage Compensation Blewitt ME, Gendrel AV, Pang Z, Sparrow DB, Whitelaw N, Craig JM, Apedaile A, Hilton DJ, Dunwoodie SL, Brockdorff N, et al. 2008. SmcHD1, containing a structuralmaintenance-of-chromosomes hinge domain, has a critical role in X inactivation. Nat Genet 40: 663 –669. Bone JR, Lavender J, Richman R, Palmer MJ, Turner BM, Kuroda MI. 1994. Acetylated histone H4 on the male X chromosome is associated with dosage compensation in Drosophila. Genes Dev 8: 96–104. Brinkman AB, Roelofsen T, Pennings SW, Martens JH, Jenuwein T, Stunnenberg HG. 2006. Histone modification patterns associated with the human X chromosome. EMBO Rep 7: 628 –634. Cai S, Lee CC, Kohwi-Shigematsu T. 2006. SATB1 packages densely looped, transcriptionally active chromatin for coordinated expression of cytokine genes. Nat Genet 38: 1278–1288. Carrel L, Park C, Tyekucheva S, Dunn J, Chiaromonte F, Makova KD. 2006. Genomic environment predicts expression patterns on the human inactive X chromosome. PLoS Genet 2: e151. Chadwick BP. 2007. Variation in Xi chromatin organization and correlation of the H3K27me3 chromatin territories to transcribed sequences by microarray analysis. Chromosoma 116: 147 –157. Chadwick BP, Willard HF. 2004. Multiple spatially distinct types of facultative heterochromatin on the human inactive X chromosome. Proc Natl Acad Sci U S A 101: 17450–17455. Chaumeil J, Le Baccon P, Wutz A, Heard E. 2006. A novel role for Xist RNA in the formation of a repressive nuclear compartment into which genes are recruited when silenced. Genes Dev 20: 2223–2237. Chow JC, Ciaudo C, Fazzari MJ, Mise N, Servant N, Glass JL, Attreed M, Avner P, Wutz A, Barillot E, et al. 2010. LINE-1 activity in facultative heterochromatin formation during X chromosome inactivation. Cell 141: 956 –969. Chow JC, Hall LL, Baldry SE, Thorogood NP, Lawrence JB, Brown CJ. 2007. Inducible XIST-dependent X-chromosome inactivation in human somatic cells is reversible. Proc Natl Acad Sci U S A 104: 10104– 10109. Chuang PT, Albertson DG, Meyer BJ. 1994. DPY-27:a chromosome condensation protein homolog that regulates C. elegans dosage compensation through association with the X chromosome. Cell 79: 459– 474. Chuang PT, Lieb JD, Meyer BJ. 1996. Sex-specific assembly of a dosage compensation complex on the nematode X chromosome. Science 274: 1736–1739. Clemson CM, Hall LL, Byron M, McNeil J, Lawrence JB. 2006. The X chromosome is organized into a gene-rich outer rim and an internal core containing silenced nongenic sequences. Proc Natl Acad Sci U S A 103: 7688– 7693. Comings DE. 1968. The rationale for an ordered arrangement of chromatin in the interphase nucleus. Am J Hum Genet 20: 440– 460. Csankovszki G, Collette K, Spahl K, Carey J, Snyder M, Petty E, Patel U, Tabuchi T, Liu H, McLeod I, et al. 2009. Three distinct condensin complexes control C. elegans chromosome dynamics. Curr Biol 19: 9 –19.

Csankovszki G, Panning B, Bates B, Pehrson JR, Jaenisch R. 1999. Conditional deletion of Xist disrupts histone macroH2A localization but not maintenance of X inactivation. Nat Genet 22: 323– 324. de Wit E, van Steensel B. 2009. Chromatin domains in higher eukaryotes: insights from genome-wide mapping studies. Chromosoma 118: 25–36. Deakin JE, Hore TA, Koina E, Marshall Graves JA. 2008. The status of dosage compensation in the multiple X chromosomes of the platypus. PLoS Genet 4: e1000140. Duret L, Chureau C, Samain S, Weissenbach J, Avner P. 2006. The Xist RNA gene evolved in eutherians by pseudogenization of a protein-coding gene. Science 312: 1653–1655. Filippova GN, Cheng MK, Moore JM, Truong JP, Hu YJ, Nguyen DK, Tsuchiya KD, Disteche CM. 2005. Boundaries between chromosomal domains of X inactivation and escape bind CTCF and lack CpG methylation during early development. Dev Cell 8: 31– 42. Gartler SM, Riggs AD. 1983. Mammalian X-chromosome inactivation. Annu Rev Genet 17: 155– 190. Gelbart ME, Kuroda MI. 2009. Drosophila dosage compensation: a complex voyage to the X chromosome. Development 136: 1399–1410. Grimaud C, Becker PB. 2009. The dosage compensation complex shapes the conformation of the X chromosome in Drosophila. Genes Dev 23: 2490–2495. Grutzner F, Rens W, Tsend-Ayush E, El-Mogharbel N, O’Brien PC, Jones RC, Ferguson-Smith MA, Marshall Graves JA. 2004. In the platypus a meiotic chain of ten sex chromosomes shares genes with the bird Z and mammal X chromosomes. Nature 432: 913 –917. Gupta V, Parisi M, Sturgill D, Nuttall R, Doctolero M, Dudko OK, Malley JD, Eastman PS, Oliver B. 2006. Global analysis of X-chromosome dosage compensation. J Biol 5: 3. Hall LL, Byron M, Sakai K, Carrel L, Willard HF, Lawrence JB. 2002. An ectopic human XIST gene can induce chromosome inactivation in postdifferentiation human HT-1080 cells. Proc Natl Acad Sci U S A 99: 8677–8682. Hallacli E, Akhtar A. 2009. X chromosomal regulation in flies: when less is more. Chromosome Res 17: 603–619. Heard E, Clerc P, Avner P. 1997. X-chromosome inactivation in mammals. Annu Rev Genet 31: 571– 610. Heard E, Disteche CM. 2006. Dosage compensation in mammals: fine-tuning the expression of the X chromosome. Genes Dev 20: 1848– 1867. Hellman A, Chess A. 2007. Gene body-specific methylation on the active X chromosome. Science 315: 1141–1143. Hoki Y, Kimura N, Kanbayashi M, Amakawa Y, Ohhata T, Sasaki H, Sado T. 2009. A proximal conserved repeat in the Xist gene is essential as a genomic element for X-inactivation in mouse. Development 136: 139 –146. Jans J, Gladden JM, Ralston EJ, Pickle CS, Michel AH, Pferdehirt RR, Eisen MB, Meyer BJ. 2009. A condensin-like dosage compensation complex acts at a distance to control expression throughout the genome. Genes Dev 23: 602 –618. Jonkers I, Barakat TS, Achame EM, Monkhorst K, Kenter A, Rentmeester E, Grosveld F, Grootegoed JA, Gribnau J. 2009. RNF12 is an X-Encoded dose-dependent activator of X chromosome inactivation. Cell 139: 999– 1011.

J.C. Chow and E. Heard Kelley RL, Meller VH, Gordadze PR, Roman G, Davis RL, Kuroda MI. 1999. Epigenetic spreading of the Drosophila dosage compensation complex from roX RNA genes into flanking chromatin. Cell 98: 513– 522. Kohlmaier A, Savarese F, Lachner M, Martens J, Jenuwein T, Wutz A. 2004. A chromosomal memory triggered by Xist regulates histone methylation in X inactivation. PLoS Biol 2: E171. Koina E, Chaumeil J, Greaves IK, Tremethick DJ, Graves JA. 2009. Specific patterns of histone marks accompany X chromosome inactivation in a marsupial. Chromosome Res 17: 115– 126. Li N, Carrel L. 2008. Escape from X chromosome inactivation is an intrinsic property of the Jarid1c locus. Proc Natl Acad Sci U S A 105: 17055– 17060. Lieb JD, Albrecht MR, Chuang PT, Meyer BJ. 1998. MIX-1: an essential component of the C. elegans mitotic machinery executes X chromosome dosage compensation. Cell 92: 265 –277. Lieb JD, Capowski EE, Meneely P, Meyer BJ. 1996. DPY-26, a link between dosage compensation and meiotic chromosome segregation in the nematode. Science 274: 1732– 1736. Lin H, Gupta V, Vermilyea MD, Falciani F, Lee JT, O’Neill LP, Turner BM. 2007. Dosage compensation in the mouse balances up-regulation and silencing of X-linked genes. PLoS Biol 5: e326. Lyon MF. 1998. X-chromosome inactivation: a repeat hypothesis. Cytogenet Cell Genet 80: 133– 137. Mahadevaiah SK, Royo H, VandeBerg JL, McCarrey JR, Mackay S, Turner JM. 2009. Key features of the X inactivation process are conserved between marsupials and eutherians. Curr Biol 19: 1478–1484. Marahrens Y, Panning B, Dausman J, Strauss W, Jaenisch R. 1997. Xist-deficient mice are defective in dosage compensation but not spermatogenesis. Genes Dev 11: 156 –166. Marks H, Chow JC, Denissov S, Francoijs KJ, Brockdorff N, Heard E, Stunnenberg HG. 2009. High-resolution analysis of epigenetic changes associated with X inactivation. Genome Res 19: 1361– 1373. Mendjan S, Taipale M, Kind J, Holz H, Gebhardt P, Schelder M, Vermeulen M, Buscaino A, Duncan K, Mueller J, et al. 2006. Nuclear pore components are involved in the transcriptional regulation of dosage compensation in Drosophila. Mol Cell 21: 811– 823. Mietton F, Sengupta AK, Molla A, Picchi G, Barral S, Heliot L, Grange T, Wutz A, Dimitrov S. 2009. Weak but uniform enrichment of the histone variant macroH2A1 along the inactive X chromosome. Mol Cell Biol 29: 150–156. Misteli T. 2007. Beyond the sequence: cellular organization of genome function. Cell 128: 787 –800. Nguyen DK, Disteche CM. 2006. Dosage compensation of the active X chromosome in mammals. Nat Genet 38: 47–53. Nicodemi M, Prisco A. 2007. Symmetry-breaking model for X-chromosome inactivation. Phys Rev Lett 98: 108104. Patrat C, Okamoto I, Diabangouaya P, Vialon V, Le Baccon P, Chow J, Heard E. 2009. Dynamic changes in paternal Xchromosome activity during imprinted X-chromosome inactivation in mice. Proc Natl Acad Sci U S A 106: 5198– 5203.

Penny GD, Kay GF, Sheardown SA, Rastan S, Brockdorff N. 1996. Requirement for Xist in X chromosome inactivation. Nature 379: 131 –137. Peters AH, Mermoud JE, O’Carroll D, Pagani M, Schweizer D, Brockdorff N, Jenuwein T. 2002. Histone H3 lysine 9 methylation is an epigenetic imprint of facultative heterochromatin. Nat Genet 30: 77–80. Plath K, Fang J, Mlynarczyk-Evans SK, Cao R, Worringer KA, Wang H, de la Cruz CC, Otte AP, Panning B, Zhang Y. 2003. Role of histone H3 lysine 27 methylation in X inactivation. Science 300: 131– 135. Rego A, Sinclair PB, Tao W, Kireev I, Belmont AS. 2008. The facultative heterochromatin of the inactive X chromosome has a distinctive condensed ultrastructure. J Cell Sci 121: 1119– 1127. Rens W, Grutzner F, O’Brien PC, Fairclough H, Graves JA, Ferguson-Smith MA. 2004. Resolution and evolution of the duck-billed platypus karyotype with an X1Y1X2Y2X3Y3X4Y4X5Y5 male sex chromosome constitution. Proc Natl Acad Sci U S A 101: 16257–16261. Savarese F, Flahndorfer K, Jaenisch R, Busslinger M, Wutz A. 2006. Hematopoietic precursor cells transiently reestablish permissiveness for X inactivation. Mol Cell Biol 26: 7167–7177. Schoeftner S, Sengupta AK, Kubicek S, Mechtler K, Spahn L, Koseki H, Jenuwein T, Wutz A. 2006. Recruitment of PRC1 function at the initiation of X inactivation independent of PRC2 and silencing. EMBO J 25: 3110–3122. Simonis M, Kooren J, de Laat W. 2007. An evaluation of 3C-based methods to capture DNA interactions. Nat Methods 4: 895– 901. Spector DL. 2003. The dynamics of chromosome organization and gene regulation. Annu Rev Biochem 72: 573–608. Straub T, Grimaud C, Gilfillan GD, Mitterweger A, Becker PB. 2008. The chromosomal high-affinity binding sites for the Drosophila dosage compensation complex. PLoS Genet 4: e1000302. Su WY, Xiong H, Fang JY. 2010. Natural antisense transcripts regulate gene expression in an epigenetic manner. Biochem Biophys Res Commun 396: 177–181. Taddei A, Hediger F, Neumann FR, Gasser SM. 2004. The function of nuclear architecture: a genetic approach. Annu Rev Genet 38: 305 –345. Tsai CJ, Mets DG, Albrecht MR, Nix P, Chan A, Meyer BJ. 2008. Meiotic crossover number and distribution are regulated by a dosage compensation protein that resembles a condensin subunit. Genes Dev 22: 194– 211. Valley CM, Pertz LM, Balakumaran BS, Willard HF. 2006. Chromosome-wide, allele-specific analysis of the histone code on the human X chromosome. Hum Mol Genet 15: 2335–2347. Valley CM, Willard HF. 2006. Genomic and epigenomic approaches to the study of X chromosome inactivation. Curr Opin Genet Dev 16: 240– 245. van Berkum NL, Dekker J. 2009. Determining spatial chromatin organization of large genomic regions using 5C technology. Methods Mol Biol 567: 189–213. Vaquerizas JM, Suyama R, Kind J, Miura K, Luscombe NM, Akhtar A. 2010. Nuclear pore proteins nup153 and megator define transcriptionally active regions in the Drosophila genome. PLoS Genet 6: e1000846.

Nuclear Organization and Dosage Compensation Vicoso B, Bachtrog D. 2009. Progress and prospects toward our understanding of the evolution of dosage compensation. Chromosome Res 17: 585 –602. Weber M, Davies JJ, Wittig D, Oakeley EJ, Haase M, Lam WL, Schubeler D. 2005. Chromosome-wide and promoter-specific analyses identify sites of differential DNA methylation in normal and transformed human cells. Nat Genet 37: 853– 862. Wutz A, Jaenisch R. 2000. A shift from reversible to irreversible X inactivation is triggered during ES cell differentiation. Mol Cell 5: 695 –705. Wutz A, Rasmussen TP, Jaenisch R. 2002. Chromosomal silencing and localization are mediated by different domains of Xist RNA. Nat Genet 30: 167 –174. Xu N, Donohoe ME, Silva SS, Lee JT. 2007. Evidence that homologous X-chromosome pairing requires transcription and Ctcf protein. Nat Genet 39: 1390– 1396.

Xu N, Tsai CL, Lee JT. 2006. Transient homologous chromosome pairing marks the onset of X inactivation. Science 311: 1149– 1152. Yang F, Babak T, Shendure J, Disteche CM. 2010. Global survey of escape from X inactivation by RNA-sequencing in mouse. Genome Res 20: 614 –622. Zhang LF, Huynh KD, Lee JT. 2007. Perinucleolar targeting of the inactive X during S phase: evidence for a role in the maintenance of silencing. Cell 129: 693– 706. Zhang X, Yazaki J, Sundaresan A, Cokus S, Chan SW, Chen H, Henderson IR, Shinn P, Pellegrini M, Jacobsen SE, et al. 2006. Genome-wide high-resolution mapping and functional analysis of DNA methylation in arabidopsis. Cell 126: 1189– 1201. Zhao J, Sun BK, Erwin JA, Song JJ, Lee JT. 2008. Polycomb proteins targeted by a short repeat RNA to the mouse X chromosome. Science 322: 750– 756.

The Budding Yeast Nucleus Angela Taddei1,2, Heiko Schober3, and Susan M. Gasser3 1

UMR 218, Centre National de la Recherche Scientifique, 26 rue d’Ulm, 75231 Paris Cedex 05, France

2

Institut Curie-Section de Recherche, 26 rue d’Ulm, 75231 Paris Cedex 05, France

3

Friedrich Miescher Institute for Biomedical Research, Maulbeerstrasse 66, CH-4058 Basel, Switzerland

Correspondence: [email protected]

The budding yeast nucleus, like those of other eukaryotic species, is highly organized with respect to both chromosomal sequences and enzymatic activities. At the nuclear periphery interactions of nuclear pores with chromatin, mRNA, and transport factors promote efficient gene expression, whereas centromeres, telomeres, and silent chromatin are clustered and anchored away from pores. Internal nuclear organization appears to be function-dependent, reflecting localized sites for tRNA transcription, rDNA transcription, ribosome assembly, and DNA repair. Recent advances have identified new proteins involved in the positioning of chromatin and have allowed testing of the functional role of higher-order chromatin organization. The unequal distribution of silent information regulatory factors and histone modifying enzymes, which arises in part from the juxtaposition of telomeric repeats, has been shown to influence chromatin-mediated transcriptional repression. Other localization events suppress unwanted recombination. These findings highlight the contribution budding yeast genetics and cytology have made to dissecting the functional role of nuclear structure.

ith 16 chromosomes and a closed mitosis, the budding yeast nucleus provides an ideal model system for understanding structure– function relationships within one of the cell’s most complex compartments. Eukaryotic chromatin has an intrinsic higher-order structure based on the folding of the nucleosomal fiber. Yet beyond the intrinsic folding of the 10- and 30-nm fibers, sequence-determined subchromosomal domains and entire chromosomes assume non-random positions in the nucleus. Here we discuss a few general principles of nuclear organization. The first is an order imposed by enzymatic function. Examples include replication foci, splicing centers,

W

or sites of DNA repair by homologous recombination (Lamond and Spector 2003). The second is a global territorial organization that stems from a nonoverlapping positioning of entire chromosomes within the nucleus (Cremer et al. 2006). The third principle reflects binding characteristics of particular sequence elements, such as simple repeats, and the factors they recruit to assemble unique higher-order levels of chromatin folding (Gasser et al. 2004). The most straightforward way to show the functional importance of nuclear and chromosomal structure is to identify the proteins and sequences involved and to mutate those elements. Any resulting alteration in nuclear

Editors: David L. Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000612 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000612

A. Taddei, H. Schober, and S.M. Gasser

function might stem from the change in organization. These loss-of-function approaches then need to be coupled with gain-of-function assays that restore or promote a particular function or spatial organization. For these manipulations budding yeast is an ideal organism. Yeast has extremely efficient homologous recombination machinery that allows for targeted gene disruption, mutagenesis, chromosomal modification, and in vivo tagging of both DNA and proteins for live imaging. These techniques can be coupled with powerful biochemical and molecular genetic techniques, many of which give genome-wide read-outs. Such approaches have provided basic insights into the relationship of nuclear structure and function (e.g., Andrulis et al. 1998; Taddei et al. 2004; Taddei et al. 2006). Unfortunately, some elements of nuclear organization present in multicellular organisms are missing in yeast. Having a closed mitosis means that disassembly and reassembly of the yeast nuclear envelope does not occur during the cell cycle, although yeast has less pronounced changes in its nuclear membrane during cell division. Coupled with the yeast cell’s closed mitosis is a lack of nuclear intermediate filament proteins or lamins, which form a rigid network underlying the inner nuclear membrane (INM). The nuclearlamina stabilizes nuclear shape and help anchor interphase chromatin (Gruenbaum et al. 2005). Yeast cells also lack many of the well-characterized subnuclear “bodies” with the exception of a nucleolar domain that functions in snoRNA maturation (Verheggen et al. 2001) and which appears to be equivalent to Cajal bodies (Gall, 2000). Despite lacking detectable splicing compartments (Misteli, 2000) or PML ( promyelocytic leukemia) (Zhong et al. 2000) bodies, yeast supports many of the activities coordinated by these compartments, and recent evidence suggests that the yeast nuclear pore provides a platform for events related to gene expression and double strand break processing (Akhtar and Gasser 2007; Taddei 2007; Nagai et al. 2008). Other aspects of subnuclear organization, such as the distribution of chromatin within the nucleus, are conserved from yeast to man.

NUCLEAR ORGANIZATION Nuclear Envelope Associated Proteins and the Nuclear Pore Complex

The nucleus is delimited by a double membrane called the nuclear envelope (NE) that is contiguous with the endoplasmic reticulum. The NE separates chromatin from the cytoplasm and provides anchorage for various nuclear and chromosomal structures, including the spindle pole body, and separate clusters of centromeres and telomeres. Trafficking between the nucleoplasm and the cytoplasm occurs through approximately 200 nuclear pore complexes (NPCs), which allow the free diffusion of small molecules, whereas regulating the transport of macromolecules. NPCs also provide a platform for mRNA transcription and quality control, as well as its export. These proteinaceous assemblies of approximately 50 MDa contain 456 nucleoporins of 30 different types (D’Angelo and Hetzer 2008). Each pore is a doughnut-shaped structure with eightfold symmetry around a central channel. Flexible protein filaments emanate from the core into both the cytoplasm and nucleoplasm, providing binding sites for both transport proteins and chromatin. A detailed map for the relative position of each nucleoporin was calculated based on multiple molecular, biochemical, and structural data revealing a strongly modular structure (Alber et al. 2007). Beside nucleoporins, many yeast proteins were shown by imaging techniques to be associated with the nuclear envelope (Huh et al. 2003). Of particular interest are the integral proteins of the inner nuclear membrane (INM) (Lusk et al. 2007) including: Doa10, a RING domain containing protein that targets nuclear proteins for degradation, Mps3, a member of the SUN (Sad1, UNC-84) family that is a shared component of the INM and of the spindle pole body (SPB; Jaspersen et al. 2002), and Helix – extension – helix-1 and -2 (Src1 or Heh1 and Heh2) (King et al. 2006), which are orthologs of the mammalian lamin associated protein MAN1. These proteins interact with the nuclear lamina in mammals, as well as with chromatin through a small protein called BAF1. Although yeast lack the lamins, the roles of SUN-domain

The Budding Yeast Nucleus

and Man1 family proteins in chromosome anchorage seem to be conserved (Grund et al. 2008). In addition, an INM-associated protein called Esc1 (Enhancer of silent chromatin 1) shares features with lamina components: Esc1 binds but does not span the inner nuclear membrane, and anchors silent chromatin through a component of the silencer complex, Sir4 (Taddei et al. 2004). Intriguingly, overexpression of Esc1 induces INM expansion (Hattier et al. 2007). Given their intimate proximity, it is not surprising that functional cross-talk among INM proteins such as Esc1 and the nuclear pore basket proteins Nup60, Mlp1 and Mlp2 (Therizols et al. 2006; Lewis et al. 2007; Palancade et al. 2007) has been detected. Chromosome Folding

Observations of chromosome folding were originally recorded by one of the founders of the field of cytology, Carl Rabl (Rabl 1885), who described the fold-back conformation of anaphase chromosomes in spotted salamander larvae. In the so-called Rabl conformation a chromosome folds back at its centromere, such that telomeres are juxtaposed (Fig. 1). Today we know that this conformation results from the attachment of microtubules to kinetochores, which assemble at centromeric chromatin, and which lead the movement of chromosomes as chromosomes are pulled by microtubules into daughter cells. The telomeres, on the other hand, lag passively behind. Evidence that yeast chromosomes assume a Rabl conformation came initially from the work of Loidl and coworkers, who used fluorescent in situ hybridization (FISH) to identify chromosomal landmarks such as centromeres and telomeres (Jin et al. 2000). Using chromosome conformation capture (3C) to model chromosome conformation, the Kleckner lab predicted that yeast Chr III folds as a contorted ring, with a strong bend near the centromere and the telomeres in close proximity to each other (Dekker et al. 2002). Gasser and co-workers confirmed and extended these observations showing that not only Chr III, but other small metacentric chromosomes, such as Chr VI,

form small loops in vivo with closely juxtaposed right and left telomeres. In contrast, telomeres on chromosome arms of vastly different lengths did not interact (Bystricky et al. 2005; Schober et al. 2008). Importantly, these studies showed that the Rabl configuration is not unique to anaphase in yeast, but persists throughout the cell cycle. Using a genome-wide conformation capture approach, in which spatial chromosome interactions can be mapped, O’Sullivan and co-workers have recently shown that yeast chromosome placement is not random, i.e., some chromosomes were found never to interact, whereas others showed favored interactions (Rodley et al. 2009). Furthermore, this method detected intra-chromosomal loops, much like those described by microscopic analysis for chromosomes 3 and 6 (Bystricky et al. 2005; Schober et al. 2008). Consistently, a new imaging and data analysis approach revealed a strong confinement of several loci into “gene territories” reflecting the architectural constraints imposed on chromosomes by nuclear structures (i.e., centromere attachments to the SPB and telomere anchoring to the NE). Importantly, this organization can be significantly remodeled upon transcriptional activation (Berger et al. 2008). Chromatin Dynamics

Chromatin in living cells is subjected to constant motion, which can be best described as a constrained random walk (Marshall et al. 1997) (Fig. 2). Rapid time-lapse imaging led to the distinction of at least two types of motion in yeast: small random movements (,0.2 mm within 1.5 s) that occur constantly, as well as larger, more directional movements (i.e., .0.5 mm in a 10.5 s interval) (Heun et al. 2001). The smaller movements are observed for internal loci as well as for peripheral silent domains, although the active loci in the nuclear lumen make large movements more frequently (i.e., a large movement is .0.5mm within 10.5 s) (Heun et al. 2001). Intriguingly, changes in cellular energy levels because of depletion of glucose or the addition of protonophores to deplete membrane potential, abolish large movements

A. Taddei, H. Schober, and S.M. Gasser

A

B Telophase 23

Nucleolus

Interphase TEL

CEN

TEL

C a

Telomeres + HM

b

SBP

c TelR, TelL

SBP

Figure 1. Chromosomes have distinct orientation within the yeast interphase nucleus, which results from their

Rabl configuration at anaphase: centromeres cluster and lead the way into daughter nuclei, with telomeres following behind. Telomeres of equal-length chromosome arms tend to cluster together in yeast interphase nuclei. (A) In 1885, Carl Rabl drew the folded conformation of anaphase chromosomes in spotted salamander larvae, in which anaphase chromosomes fold back on themselves because of attachment of the centromere to microtubules through the kinetochore. Centromeres lead the way as chromosomes are actively pulled into daughter cells. (B) The Rabl organization persists in the interphase yeast nucleus. (C ) Confocal fluorescence images show (a) the clustering of yeast centromeres (green by FISH) near the spindle pole body (SPB, red) or in the larger image the nucleolus (red) opposite the SPB (anti-Spc42, white); (b) yeast telomeric foci are labeled with anti-Sir4 protein and FISH for the silent HM loci in yeast; (c) visualization of the centromere and right and left telomeres of Chr VI through Tel6R-CFP-lacO, Tel6L-YFP-TetR, and centromere staining (white).

(Marshall et al. 1997; Heun et al. 2001). This suggests that interphase chromatin movement at least responds to ATP levels. Recent work suggests that these most likely stem from the action of chromatin remodelers, rather than RNA or DNA polymerases. Indeed, highly transcribed galactose-inducible genes that become tethered at nuclear pores show less mobility, despite their high level of transcription (Taddei et al. 2006; Cabal et al. 2006). Moreover, chromatin movement decreases, rather than increases, as cells enter S phase and DNA polymerases are active (Heun et al. 2001). Given that the mobility of a genomic locus is constrained by its continuity

with the rest of the chromosome (Gartenberg et al. 2004), it is likely that the S-phase associated drop in mobility reflects the association of DNA in replication factories (Kitamura et al. 2006). DNA BASED COMPARTMENTS: NUCLEOLUS, TELOMERES, tRNA The Nucleolus

The most evident subnuclear compartment is the nucleolus, a crescent-shaped structure abutting the nuclear envelope and occupying roughly one third of the nucleus opposite the spindle pole body (Yang et al. 1989; Bystricky

The Budding Yeast Nucleus Active loci, mid-chromosome XIV

Induced HXK1 gene at pore

Figure 2. Rapid time-lapse imaging of GFP-lacI tagged chromosomal loci in yeast cells bearing GFP-Nup49,

allows 3D tracking of an internal locus (Gartenberg et al. 2004). 3D stacks are taken at 1.5 s intervals and the spatial rendering and alignment is performed with Imaris (Bitplane, Zu¨rich). Tracks show movement over 7.5 min (300 stacks). To the left is a region of Chr IVX that shows low level transcription upon activation by glucose. To the right is shown the movement of the induced HXK1, a subtelomeric gene that is strongly induced in the absence of glucose (Taddei et al. 2006). Reductions in radius of constraint and diffusion coefficients are monitored for the pore-associated gene.

et al. 2005). This subcompartment is the site of RNA pol I-mediated rDNA transcription and ribosome subunit assembly and can be seen as a factory dedicated to ribosome biogenesis. Its morphology is strongly influenced by the cell growth rate, probably as a result of adapting the rate of ribosome production to the needs of the cell (Oakes et al. 1993; Powers and Walter, 1999). The nucleolus is a prime example of a functional organization principle, for it compartmentalizes different steps of ribosome biogenesis although the nucleolar machineries are in permanent exchange with the nucleoplasm and other nuclear bodies. The nucleolus further generates a domain of retention/sequestration of molecules that are normally active only outside the nucleolus (Sirri et al. 2008). In budding yeast, rRNA is encoded in 100– 200 tandem repeats. Each repeat unit is 9.1 kb in size and yields a 35S precursor rRNA, transcribed by RNA polymerase I and a 5S rRNA, transcribed by RNA polymerase III. The 5S unit is surrounded by two intergenic spacers, IGS1 and IGS2 (Fig. 3). Within this spacer there is a so-called polar replication fork barrier (RFB), a recombination enhancer (RE), a RNA

polymerase I transcription initiation region (TIR) and an origin of replication (ARS, from autonomously replicating sequence). This extended array of tandem repeats serves as an ideal template for homologous recombination, which is highly efficient in budding yeast. However, suppression of this recombination and the ensuing stability of the yeast rDNA array, are absolutely critical for cell growth and survival. By stabilizing the rDNA repeats the cell avoids or delays replicative senescence (Sinclair and Guarente 1997). The mechanisms that suppress recombination within the rDNA array involve both local nucleosomal organization mediated by Sir2 (Gottlieb and Esposito 1989; Bryk et al. 1997; Fritze et al. 1997; Smith and Boeke 1997) and long-range chromatin structure that involves tethering at the nuclear envelope. tRNA genes

Although the 274 Pol III-transcribed tRNA genes are scattered throughout the yeast genome, many of these are clustered close to the nucleolus (Thompson et al. 2003; Wang et al.

A. Taddei, H. Schober, and S.M. Gasser 9.1 kb 35S

35S

RE

35S

5S

35S

ARS

RFB IGS1

TIR IGS2

Figure 3. Organization of the rDNA repeats. The 9.1 kb unit encodes the 35S precursor for all the 25S, 18S, and

5.8S rRNA and the 5S rRNA. The 35S is transcribed by RNA polI whereas the 5S is transcribed by RNA pol III. The 9.1 kb unit is repeated up to 200 times on chr XII. Other abbreviations are as follows: IGS, intergenic spacer; RE recombination enhancer; RFB, Replication fork block, TIR, PolI transcription initiation region, open circle—origin of DNA replication.

2005). This association could create an environment that fosters the coregulation of transcription by RNA pol III and RNA pol I, given that the other major RNA pol III transcript is the 5S rRNA, which is part of the rDNA repeat (Fig. 3). Intriguingly, some tRNA genes remain associated with the nucleolus throughout the cell cycle, although they are not present on Chr XII, which contains the rDNA. This unusual positioning depends both on microtubules and on the action of condensin (Haeusler et al. 2008). Given the high number of RNA pol III-transcribed genes, their positioning is likely to affect the spatial organization of a large portion of the genome. Intriguingly, RNA pol II genes in the vicinity of tRNA genes become silenced through a phenomenon known as tRNA-gene mediated gene silencing (tgm) (Wang et al. 2005). Consistent with a link between nucleolar tethering and tRNAassociated silencing, it was shown that conditional ablation of condensin subunits leads to a coordinate loss of both nucleolar tRNA clustering and tgm (Haeusler et al. 2008). We note, however, that not all active tRNA genes relocate to the nucleolus. It remains to be determined what characterizes those that do show nucleolar association. Telomere Foci: Assemblies of Repetitive DNA and Silencing Factors

Another type of subnuclear compartment stems from the clustering of the 32 yeast telomeres

into three to eight foci, which are mainly found at the nuclear periphery. These foci are unfavorable for RNA polII-driven transcription. The budding yeast telomeric repeat consist of 250 – 300 base pairs of irregular tandem repeats (called TG1-3) (Shampay et al. 1984). A crucial feature of telomeric DNA is the 30 overhang of the G-rich strand, which is 10 – 15bp in length (Larrivee et al. 2004). Toward the end of S phase after completion of replication, an extended G-rich overhang is generated, providing a template for the action of telomerase, a conserved ribonucleoprotein complex with reverse transcriptase activity (reviewed in Zakian 1996; Hug and Lingner 2006). The protein subunits of this complex were identified in a screen for “ever shorter telomeres” and are called Est1, Est2, and Est3 (Lundblad and Szostak 1989; Lendvay et al. 1996), whereas the RNA moiety that templates the extension of TG repeats is called Tlc1 (Singer and Gottschling 1994). Tlc1 and Est2 form the catalytic core of telomerase, whereas Est1 and Est3 are auxiliary subunits that are dispensable for in vitro activity (Lingner et al. 1997). The end-binding factor yKu interacts with telomerase and helps recruit it to telomeres (Stellwagen et al. 2003), as does the ssDNA binding factor Cdc13 (reviewed in Fisher et al. 2004). Unlike this association with ssDNA, the Repressor Activator protein 1 binds the doublestranded telomeric repeat (Rap1) (Shore and Nasmyth 1987). Rap1 is a conserved factor with a double Myb-like domain that mediates

The Budding Yeast Nucleus

high affinity binding to its consensus within the TG1-3 repeat. The Rap1 carboxyl terminus is a binding site for silencing factors Sir3 and Sir4 (Moretti et al. 1994; Marcand et al. 1997; Wotton and Shore 1997), which form a stochiometric complex with the NAD-dependent histone deacetylase Sir2, to mediate transcriptional repression in subtelomeric zones (Aparicio et al. 1991; Martino et al. 2009). The same site binds Rif1, which together with Rif2, antagonizes Sir4 interaction, and feeds back to limit telomerase activity (Wotton and Shore 1997). To prevent spreading of silent chromatin, subtelomeric regions contain boundaries, which are characterized by the presence of acetylated histone variant Htz1 (or H2A.Z) (Meneghini et al. 2003; Babiarz et al. 2006). In addition to the immediate zone of SIR-mediated repression, characterized by nucleosomes that lack detectable acetylation or methylation, there is a subtelomeric region called HAST for (Hda1affected subtelomeric) that is characterized by a continuous stretch of Hda1-deacetylated chromatin that extends 10 – 25 kb inwards from the telomeric repeat (Robyr et al. 2002). Thus telomeres are not only designed to ensure end replication, but they nucleate special domains with respect to SIR protein spreading and histone modifications.

MECHANISMS UNDERLYING NUCLEAR COMPARTMENTATION Anchoring of DNA at the Nuclear Envelope

The positioning of chromosomes within the nucleus depends on reversible interactions of chromosomal landmarks with structural features of the nucleus, such as the spindle pole body (SPB) and the inner nuclear envelope. Centromeres are held in a cluster near the SPB through short microtubules that persist through interphase (Jin et al. 2000; Bystricky et al. 2004). The tethering of telomeres in perinuclear foci is achieved by two redundant pathways that require Sir4, a silencing factor, and the yKu70/yKu80 heterodimer (Hediger et al. 2002; Taddei et al. 2004). Sir4 anchors repressed chromatin to the NE through its

partitioning and anchoring domain (PAD, aa 950 to 1262). PAD specifically binds Esc1, a low-abundance acidic protein associated exclusively with the inner face of the NE (Andrulis et al. 2002; Gartenberg et al. 2004; Taddei et al. 2004). By electron microscopy it was shown that Esc1 localizes in patches along the nuclear membrane independently of Sir4 and is excluded from nuclear pores (Taddei et al. 2004). Although yKu80 and Sir4 interact, a mutant allele of yKu80 that loses interaction with Sir4 (yKu80-4) (Taddei et al. 2004; Roy et al. 2004) can still tether chromatin to the NE (Taddei et al. 2004). Given that this occurs in strains deleted for the genes encoding Sir4 and Esc1, it was proposed that yKu should bind at least one additional membrane-bound factor. This is achieved indirectly through the ability of yKu to bind telomerase (Schober et al. 2009). Intriguingly, the requirements of the Sir4independent yKu tethering pathway varied with phases of the cell cycle: yKu80 tethering was dependent on yKu70 in G1-phase but not in S-phase cells (Taddei et al. 2004). Conversely, the domain of yKu80 that binds a stem-loop in the telomerase RNA Tlc1, was found to be necessary for telomere anchoring in S-phase, but not in G1-phase cells (Schober et al. 2009). The S-phase specific anchoring by yKu not only required Tlc1, but Est1 as well, which forms a complex with the catalytic subunit of telomerase, Est2. It was shown that a targeted Est2 fusion protein can anchor DNA to the nuclear perimeter in an Est1-dependent manner, and that Est1 interacts with an integral nuclear membrane protein, Mps3 (Fig. 4) (Schober et al. 2009; Jaspersen et al. 2002; Antoniacci et al. 2007). An acidic amino-terminal domain of this SUN-domain family member extends into the nucleoplasm where it contacts Est1 to tether telomerase-bound telomeres. Yet Mps3 also appears to contribute to an alternative telomere anchoring pathway mediated by Sir4 (Bupp et al. 2007). In both cases Mps3dependent anchoring is specific to S-phase cells. The budding yeast Mps3 also organizes the SPB (Jaspersen et al. 2002; Nishikawa et al.

A. Taddei, H. Schober, and S.M. Gasser Nuclear envelope

Nuclear pore

Mps3

Esc1 X?

G1

G1

S phase

G1 S phase

Est1 Sir4

Est3

Rap1

yKu

Tlc1 TG(1–6)

yKu Est2

Nucleosomes

Figure 4. Parallel mechanisms lead to yeast telomere attachment at the nuclear envelope. At different stages of the

cell cycle the telomere associated proteins mediate different contacts with inner nuclear membrane components. The Sir4-PAD domain binds the Esc1 C terminus, as well as yKu80 and Mps3. yKu80 binds telomerase, which also associates with Mps3 in S phase through Est1. There is an unidentified anchor for yKu in G1 phase that is neither Esc1- nor Mps3-dependent.

2003), yet by deletion of its N-terminal domain, the role of Mps3 in chromatin tethering could be separated from its essential role in spindle pole body organization (Bupp et al. 2007). Importantly, antagonism of the yKu-telomerase-Mps3 interaction led to hyper-recombination among telomeres, suggesting that this mechanism protects replicating ends from unequal strand invasion (Schober et al. 2009). This may be particularly relevant in S phase. Finally, it has been shown that the yKu-mediated peripheral positioning is switched off after DNA replication, leading to the dislodgment of telomeres from the nuclear envelope after replication (Ebrahimi and Donaldson 2008). Such release may facilitate mitotic separation of replicated telomeres. Besides these pathways other proteins such as Asf1, Rtt109, Esc2, and Ctf18 have been implicated yeast telomers anchoring, although it could not be determined whether their effects were direct or indirect (Hiraga et al. 2006, 2008). Indeed, mutations in proteins of the nuclear pore such as Mlp1, Mlp2, or Nup133 were shown to interfere with mRNA export and cell cycle progression, which could indirectly affect perinuclear organization.

Trans-Association of Chromatin Loci

The high concentration of macromolecules within the nucleoplasm (between 100 and 400 mg/ml) produces volume exclusion effects that enhance attractive interactions among macromolecules, a phenomenon known as macromolecular crowding. This effect has been proposed to favor the formation of compartments by driving bulky components into structurally compact organizations (Richter et al. 2007). Combination of these effects with specific intermolecular interactions and attachment to nuclear structures such as the NE could account for the formation of subnuclear compartments. One example of a functional compartment is provided by telomere clusters onto which silencing factors concentrate. The bifunctional role of Sir4 as a mediator of repression and an anchor for silent chromatin provides a mechanism for the self-organization of repressive compartments. In vivo, a nonsilent telomere can be localized at the NE through interaction with yKu, and thus be brought into proximity of other telomeres which generate a zone enriched for Sir proteins. Recruitment to this

The Budding Yeast Nucleus

compartment results in a higher probability that a subtelomeric gene gets repressed. Once Sir-dependent repression is established, the Sir4 anchoring pathway ensures that the silent chromatin stays perinuclear. In this way, the formation of silent chromatin is self-reinforcing, thanks to its ability to bind to the same sites as telomeres. Intriguingly, telomeres are not evenly distributed at the nuclear periphery but rather form discrete foci. We cannot rule out that telomere clustering arises in part from volume exclusion effects and molecular crowding (Iborra 2007), but it has also been shown that Sir proteins interact with themselves and with each other, establishing trans-interactions between telomeres. Recent work has shown that interactions between silent domains depends not only on silencing proteins Sir2, Sir3, and Sir4, but also on Sir1 and Esc2, two proteins involved in establishment of silencing at HM loci (Miele et al. 2009). Trans-interactions were not dependent on yKu or Esc1, suggesting that the mechanisms of NE-tethering and telomere clustering can be at least partially separated. Nonetheless, the self-organization of heterochromatin into perinuclear foci illustrates a self-perpetuating mechanism that applies to other chromatin-based domains of epigenetic character.

FUNCTIONAL CONSEQUENCES OF NUCLEAR ORGANIZATION ON GENE TRANSCRIPTION Nuclear Organization and Gene Silencing

Chromatin-based compartments like those discussed earlier not only concentrate factors where they are needed, but also keep factors away from sites where they would interfere with other types of regulation (Taddei et al. 2009). Specifically, the clustering of telomeres leads to the sequestration of SIRs, which was shown both to favor subtelomeric repression and to prevent promiscuous effects on a distinct subset of promoters (Taddei et al. 2009). Essential to such a phenomenon is the fact that the Sir3 protein level, estimated at 1400 copies per cell

(Ghaemmaghami et al. 2003), is both tightly regulated and limiting for the spread of silent chromatin (Renauld et al. 1993). Sir4 abundance is also tightly regulated, and low level overexpression of Sir4 actually disrupts telomeric repression. It has been shown that each Rap1 tail and each nucleosome in the silent subtelomeric domain binds one SIR complex comprising Sir2, Sir3 and Sir4 in precisely equal molar ratio (Martino et al. 2009). This means that each telomere of 16 Rap1 binding sites (Gilson et al. 1993) and 18 subtelomeric nucleosomes (Renauld et al. 1993; Strahl-Bolsinger et al. 1997), provides binding sites for 30 –40 SIR complexes. Because each haploid yeast cell has 32 telomeres clustered in three to eight telomeric foci, a focus contains several hundred potential SIR binding sites. The clustering of telomeric repeats could thus create a “sink” for limiting amounts of Sir proteins, and could account for the strikingly unequal distribution of this repressive complex within the nucleoplasm (Gotta et al. 1996). Confirming the functional consequences of this unequal distribution of Sir proteins, it was found that silencer-nucleated repression is highly sensitive to the distance of the reporter from a telomere (Renauld et al. 1993; Stavenhagen and Zakian 1994; Thompson et al. 1994; Maillet et al. 1996; Marcand et al. 1996). Reciprocally, the tethering of a weakened HMR silencer to the NE favored the repression of a reporter gene (Andrulis et al. 1998). Importantly, this effect depends on the ability of telomere clusters to concentrate SIR factors at the nuclear envelope (Mondoux et al. 2007; Taddei et al. 2009). Thus, it appears that transcriptional silencing is not inherent to position, but rather requires facilitated access to a local high concentration of SIR proteins. Confirming this, when telomere anchoring is impaired, by deletion of YKU70 and ESC1, transcription is affected differently at different loci: genes at internal loci flanked by silencers show enhanced repression, whereas telomere-proximal repression is lost (Maillet et al. 2001; Taddei et al. 2009). In conclusion, the efficiency with which silent chromatin is formed depends on spatial concern,

A. Taddei, H. Schober, and S.M. Gasser

such as the local concentration of SIR proteins, and the strength of cis-acting nucleation elements. In apparent contradiction with this interpretation is the absence of a correlation between the colocalization of a subtelomeric gene in telomere clusters (visualized as Rap1-GFP foci) and the efficiency of TPE (Mondoux et al. 2007). This may be because of the dynamic behavior of individual telomeres, which move into and out of Rap1 and SIR foci rapidly, presumably without loss of their epigenetic status, because they retain SIR factors (Schober et al. 2008). Indeed, a repressed ring of chromatin maintains a stable state of repression, even when it is excised from its subtelomeric chromosomal context, suggesting that a high local concentration of SIR factors is dispensable once silencing is established (Gartenberg et al. 2004). The promiscuous repression of nontelomeric genes by released SIR factors, was analyzed genome-wide using a computational program that maps putative transcriptional factor sites. The affected promoters carry specific transcription factor motifs for either Abf1 or the PAC factors, Pdf1 and Pdf2, which are part of the Rpd3L histone deacetylase complex (RNA Polymerase A and C promoters; Dequard-Chablat et al. 1991; Zhu et al. 2009). A related motif, RRPE (ribosomal RNA processing element; Hughes et al. 2000) also correlated weakly with promiscuous SIR regulation. Intriguingly, these elements are primarily bound to promoters of genes expressing enzymes and proteins involved in ribosome biogenesis. Down-regulation may be seen as a global attempt to suppress growth, and perhaps favor a stress survival pathway. These results make a forceful argument that changes in the spatial distribution of repetitive sequences that bind silencing factors can regulate patterns of gene expression genome-wide. The cell may exploit such a mechanism by controlling SIR complex release, an event that can respond rapidly to environmental insult (Martin et al. 1999; Ai et al. 2002). Consistently, SIR dispersion, or modulation of TPE, have been observed in the presence of various forms of stress (Stone and Pillus 1996; Martin et al.

1999; McAinsh et al. 1999; Mills et al. 1999; Ray and Runge 1999; Bi et al. 2004; Mercier et al. 2005), stress-induced redistribution of SIR proteins may derepress subtelomeric genes required for use of alternative carbon sources, and simultaneously contribute to the downregulation of genes involved in ribosome biogenesis. Nuclear Organization and Inducible Gene Expression

Over the last years nuclear pore complexes (NPCs) emerged as a major player in organizing gene activity. First, the nucleoporin Nup2 was shown to exert a strong boundary activity that can block the spread of heterochromatin when targeted on both sides of a reporter gene (Ishii et al. 2002). In this case, NPC tethering was proposed to establish a protected chromatin domain by creating a small DNA loop. Subsequently, a series of genome-wide studies identified nuclear pore components to be associated with highly active genes (Casolari et al. 2004, 2005; Schmid et al. 2006). Furthermore, specific inducible genes (INO1, HXK1, GAL1, GAL2, HSP104) were shown to associate with the nuclear periphery upon activation (Akhtar and Gasser 2007; Taddei 2007) (Fig. 5). Although many active genes seem to be associated with the NE, a stable interaction is not an obligate feature of gene activity (Casolari et al. 2004) and activation of the same promoter by different pathways or with different 30 UTR can alter a gene’s position within the nucleus (Abruzzi et al. 2006; Taddei et al. 2006). How specific genes associate with the NPC is still unclear and each step of mRNA production, maturation and export has been implicated at some level (Taddei 2007 for review). If multiple steps contribute to the stable association of active genes to NPCs, the importance of each individual step to NPC anchoring is probably specific for each gene. One possible scenario is that promoter-bound factors such as SAGA and Sus1 promote an initial, transient contact of the gene with pore proteins. This may be later stabilized by factors recruited for transcript termination, processing, quality

The Budding Yeast Nucleus Telomere clusters

A

Transcription compartments On Off

Off

Nucleolus

On

On

SPB

Foci of silent information regulators stem from anchored telomeres (Gotta et al. 1996) and favor repression (Andrulis et al. 1998) Boundary elements and stress-induced genes bind nuclear pores (Ishii et al. 2002; Casolari et al., 2004), in some cases increasing mRNA level (Brickner and Walter, 2004; Taddei et al. 2006)

On/Off Normal promoter-based Pol IIdependent transcription RNA Pol III (tRNA) transcription at the nucleolar edge (Thompson et al. 2003)

NPC Boundary

NPC

DNA Repair compartments

B Mps3

Mps3 binds Est1-telomerase to suppress tel-tel recombination (Oza et al. 2009; Schober et al. 2009)

NPC Heh1

Canonical HR-mediated repair (Bystricky et al. 2009) Nucleolus : exclusion of recombination proteins (Torres-Rosell et al. 2007) Nup84/Slx5/Slx8 regulates a switch to alternative repair pathways and telomerase-independent telomere maintenance (Azam et al. 2006; Nagai et al. 2008)

Heh1 NPC

Heh1 (Man1 homologue) suppresses rDNA recombination by tethering (Mekhail et al. 2008)

Figure 5. Nuclear subcompartments of yeast. (A) Transcription subcompartments within the yeast nucleus.

Telomere clustering favors repression through SIR factors whereas pore association correlates with induced expression of certain genes and boundary function. (B) DNA repair compartments within the yeast nucleus. The nucleoplasm is the site of Rad52-mediated recombination whereas sequestration at Mps3 or nuclear pores either suppresses recombination between telomeres or processes a DSB for alternative repair pathways. For the rDNA, binding to the INM protein Heh1 (Src1) prevents rDNA recombination.

control and export. These multiple anchors could favor the formation of active gene loops connecting promoters and 30 sequences, thus improving the recycling of polymerases and elongation efficiency (O’Sullivan et al. 2004). Gene-NPC association might be particularly important for inducible genes such as

galactose- and heat-shock controlled promoters for which NPC association involves the histone acetyltransferase SAGA complex. This complex is connected to the mRNA export machinery by one of its components, Sus1, which is also part of the Sac3-Thp1-Cdc31 complex that binds the NPC through Nup1. The SAGA

A. Taddei, H. Schober, and S.M. Gasser

complex is generally required for the induction of stress-responsive genes, suggesting a rationale for gene relocalization: Stress-induced genes are likely to require rapid and high level expression and export, which could be facilitated by their positioning at pores. Although, mutations affecting gene-NPC association show normal activation level for GAL1 (Cabal et al. 2006) it is still possible that the kinetics of induction are affected in these mutants. Importantly, association with the nuclear periphery has been shown to enhance the activation of at least some genes (Brickner and Walter 2004; Menon et al. 2005; Taddei et al. 2006) and could provide an additional layer of regulation for fine-tuning expression levels. Gene-NPC association has also been proposed to reflect an epigenetic mark that enables past events to be “remembered” (Brickner et al. 2007). It was reported that both the GAL1 and INO1 genes remain associated with the NPC for several generations after the inducing agent is removed. These genes appeared to be more rapidly re-activated in a pore-bound state than after a long-term repression. However, this mechanism has been called into question because other mechanisms involving cytoplasmic factors also can account for the rapid reactivation of these genes (Zacharioudakis et al. 2007). The mechanism through which NPC-association can influence expression is still unclear. In 1985, Blobel proposed the “gene gating” hypothesis according to which a “circumscribed space subjacent to the NPC and extending into the interior of the nucleus was envisioned to serve as the locale where transcription and much of the co- and posttranscriptional processing would occur” (Blobel 1985). Thus, the NPC was proposed to be a scaffold to build an assembly line favoring the coordination of the different processes that occur at an active gene. Because of their eightfold symmetry, each pore may accommodate multiple genes, forming a factory for highly efficiency transcription, in which factors that promote initiation or elongation can concentrate. The existence of internal transcription factories has been proposed in differentiation mammalian nuclei or in cultured

cells to coordinate expression of coregulated genes (Sexton et al. 2007). Such internal transcription factories might also exist in yeast although it remains unclear what provides the scaffolding for promoter recruitment. The Dual Role of the Nuclear Envelope

Given its role in the creation of repressive telomeric compartments and its ability to support, if not promote, high-level gene induction, one can conclude that the nuclear periphery has a dual role in regulating gene expression. This dual role is corroborated by the structural organization of the nuclear envelope. Indeed, microscopy studies show that repressive and activating compartments at the nuclear periphery coexist but do not overlap, as visualized by the positioning of subtelomeric domains in-between nuclear pores (Taddei et al. 2004). This proximity between repressive (telomere clusters) and activating compartments (NPCs) could favor the efficiency and reversibility of gene induction. This might be especially relevant for subtelomeric genes, which are mainly involved in the usage of alternative carbon sources and have to be induced only under specific growth conditions (Fabre et al. 2005). Consistently, increasing the association of the HXK1 subtelomeric gene with the nuclear periphery improves both its repression on glucose medium and its activation in the absence of glucose (Taddei et al. 2006). FUNCTIONAL CONSEQUENCES OF NUCLEAR ORGANIZATION ON GENOME STABILITY

Although these studies indicate important functions for yeast nuclear structure in the regulation of gene expression, recent work suggests that DNA repair may also respond to spatial cues in the nucleus. The most obvious elements that organize DNA repair are foci that form at double strand breaks (DSBs) as they are processed for homologous recombination (HR). This can be followed through the binding of fluorescently tagged Rad52, a protein that facilitate Rad51-dependent strand invasion, preceding

The Budding Yeast Nucleus

the formation of a Holliday junction (New et al. 1998). A Rad52-containing HR center is able to recruit more than one DSB (Lisby et al. 2003), suggesting that there is a self-recognition event that allows breaks to congregate. Foci of Rad52, whether spontaneous or induced by DNA damage, are strongly enriched in the nuclear interior (Bystricky et al. 2009). Given that sister chromatid exchange is the dominant pathway for homologous recombination in mitotically dividing cells (Kadyk and Hartwell 1992), the internal enrichment of Rad52 foci argues that most sister chromatid exchange occurs in the nuclear interior. Indeed, when the HMR or HML sequences are available as donors for the cleaved MATa locus, MAT stays internal and recruits the appropriate donor, e.g., HMLa, from a more peripheral position for gene conversion (Nagai et al. 2008; Bystricky et al. 2009). The inward shift is only seen for the appropriate donor locus, and thus is matingtype dependent. Pore Association of Persistent DSBs, Collapsed Forks and Critically Short Telomeres

If a donor sequence is not available, and the HO endonuclease is continuously expressed, the cut at the MAT locus persists. Such irreparable breaks are seen to shift to the nuclear periphery in a process dependent on Mec1 kinase and the histone variant Htz1 (Nagai et al. 2008; Kalocsay et al. 2009; Oza et al. 2009). Several lines of evidence implicate a nuclear pore subcomplex as the binding site, although the INM protein Mps3 has also been implicated transiently in the relocation process (Nagai et al. 2008; Kalocsay et al. 2009; Oza et al. 2009). Alternatively one might argue that the breaks move nonspecifically to the NE because crosslinking to both nuclear pores and Mps3, was reported (Nagai et al. 2008; Kalocsay et al. 2009; Oza et al. 2009). Links between DNA damage and nuclear pores was supported by an epistasis-miniarray profile (E-MAP) analysis (Collins et al. 2007b), in which the Nup84 subcomplex of the nuclear pore, containing Nup133, Nup120 and

Nup84, was clustered with a conserved SUMOdependent ubiquitin ligase complex, Slx5/Slx8. The Slx5/Slx8 complex plays an important role in the maintenance of genome integrity particularly during DNA replication (Perry et al. 2008). In addition, Slx5, which bears two SUMO interacting motifs, interacts physically with components of the proteasome (Collins et al. 2007a). Tellingly, there is an accumulation of SUMOylated proteins in cells lacking either Slx5 or Slx8 (Wang et al. 2006; Xie et al. 2007), suggesting that the Slx5/Slx8 complex normally helps the proteasome degrade SUMOylated proteins through its ubiquitin ligase activity (Wang et al. 2006; Xie et al. 2007). Indeed, an accumulation of SUMO conjugates has been shown to be responsible for the DNA damage sensitivity of the slx8 mutant in fission yeast (Prudden et al. 2007). Mutation of any component of the Nup84 complex, i.e., Nup84, Nup120, or Nup133 nucleoporins, renders cells hypersensitive to DNA damage, such as irradiation, MMS, bleomycin and HU (Bennett et al. 2001; Loeillet et al. 2005; Therizols et al. 2006). The same is true for mutation of Slx5 or Slx8 (Nagai et al. 2008). Furthermore, mutants of the Nup84 complex have been shown to be synthetically lethal with mutants of DNA repair, recombination and replication genes (Loeillet et al. 2005; Pan et al. 2006; Collins et al. 2007b; Nagai et al. 2008). A direct involvement of nuclear pores in DNA repair was supported by the fact that Nup84 and Slx5/Slx8 colocalize and coimmunoprecipitate (Nagai et al. 2008). ChIP assays have shown that irreparable DSBs and collapsed, but not stalled, replication forks, becomes associated with the Nup84 – Slx5/Slx8 complex. Moreover, the rate of spontaneous gene conversion could be enhanced by tethering the recipient site of recombination to the nuclear periphery. The enhancement of DNA repair was dependent on Slx8 and Nup84 proteins, suggesting that Slx5/Slx8 - Nup84 complex facilitates at least one kind of strand invasion event (Nagai et al. 2008). Short telomeres have also been reported to associate with the nuclear pore complex (Khadaroo et al. 2009).

A. Taddei, H. Schober, and S.M. Gasser

Importantly, pore binding is not detected if DSBs can be readily repaired by HR (Nagai et al. 2008). This suggests that only lesions that are refractive to DNA repair by canonical HR pathways associate stably with pores. At the pore a repair intermediate may be shunted to alternative recombination and repair pathways through steps that are controlled by SUMO recognition, ubiquitylation, and degradation, in other words through Slx5/Slx8 and the proteasome. The yeast proteasome was also shown by ChIP to be recruited to irreparable DSBs, and was genetically shown to be required for recombination-mediated modes of DNA repair (Krogan et al. 2004). In mammals, the proteasome has been implicated in the differential choice of recombination pathways; namely the up-regulation of gene-conversion at the expense of single-strand annealing (Gudmundsdottir et al. 2007). Consistently, Slx5/Slx8 has been shown to counteract Rad51-independent recombination, such as single-strand annealing and the break induced replication (Burgess et al. 2007). An increased dependence on Rad51-independent recombination pathways favors deleterious genomic rearrangements, such as gene amplifications, deletions and translocations (Haber and Debatisse 2006). Thus, we suggest the possibility that the pore-associated DNA repair pathway mediated by Slx5/Slx8 complex helps counteract error-prone Rad51-independent recombination, whereas up-regulating alternative pathways for recovery. Alternative pathways may be particularly important at collapsed replication forks and at telomeres. Which pathways are used, will depend on the targets of Slx5/Slx8 action. Based on E-MAP data likely pathways will include Pol32, Rad27, Est1, and Srs2 (Nagai et al. 2008). Intriguingly, the outgrowth of type II survivors of telomerase inhibition, which requires TG repeat amplification by recombination, requires not only Rad52 and Sgs1, but Slx5 and Slx8 (Azam et al. 2006; McEachern and Haber 2006). Given the involvement of the Slx5/8 E3 ligase and proteasome, it seems likely that shifting the mode of DNA repair at these loci may require the degradation of an inhibitor, one that could be

the target of SUMO-directed ubiquitination by Slx5/Slx8 (Collins et al. 2007a; Wang et al. 2006; Xie et al. 2007). Finally, there may be crosstalk between Mps3 and these pathways, because the mps3D75-150 mutant, which fails to bind DSBs and telomeres, delayed recombinational repair of DSBs and decreased gross chromosomal rearrangements in either a slx5 deletion or in the pif1-m2 mutant, which abrogates the negative regulation of telomerase activity by the Pif1 helicase (Oza et al. 2009). Consistently, overexpression of the Mps3N domain increased telomere exchange (Schober et al. 2009). Regulation of Recombination at the rDNA Locus

In budding yeast, the maintenance of rDNA repeat number is important for cell growth as well as for preventing premature senescence. It has been proposed that HR helps to maintain rDNA copy number (Smith 1974). Indeed, expansion or contraction of rDNA repeats appears to stem from HR, which follows on Fob1induced replication fork blocking (Kobayashi et al. 1998). Although several sequence elements that stimulate recombination are found within the rDNA (Keil and Roeder 1984; Kobayashi et al. 1998), the frequency of HR is nonetheless significantly lower than what one would expect for a repetitive array in yeast. This argues that recombination is actively suppressed at the rDNA locus to maintain rDNA repeat homeostasis. Interestingly, components of the HR machinery, namely Rad52, Rad51, Rad55, and Rad59, are excluded from the nucleolus, although sensors of DSBs, such as Mre11 and Rfa1 are present (Torres-Rosell et al. 2007). Furthermore, a single DSB in the rDNA induced by the I-SceI endonuclease shifts away from the nucleolus when Rad52 was recruited, suggesting repair of rDNA by HR can only take place outside of the nucleolus. Nucleolar exclusion of Rad52 foci requires the Smc5/Smc6 complex, which harbors E3 SUMO ligase activity, and requires SUMOylation of Rad52. Given that mutants that impair nucleolar exclusion of

The Budding Yeast Nucleus

HR, i.e., mutants of Smc5 or Smc6, show rDNA hyper-recombination and unequal sister chromatid exchange, one can conclude that the extrusion of HR proteins from the nucleolus is functionally important for genome stability (Burgess et al. 2007; Torres-Rosell et al. 2007). Although it is well-established that the yeast nucleolus abuts the nuclear periphery (Oakes et al. 1998), the functional significance of this localization has been poorly understood. Recent work shows that the perinuclear anchoring of rDNA repeats by the inner nuclear membrane proteins Heh1(also called Src1) and Nur1, contributes to the maintenance of rDNA repeat stability (Mekhail et al. 2008). Heh1 shares homology with human Man1 (King et al. 2006), an INM protein containing a conserved LEM (LAP-Emerin-Man1) domain. Deletion of Heh1 or Nur1 releases rDNA from the nuclear periphery, and like loss of Smc5/ Smc6, this release correlates with increase unequal sister chromatid exchange. Importantly, unlike Sir2, which also functions to reduce unequal sister chromatid exchange (Gottlieb and Esposito 1989), neither Heh1 nor Nur1 is required for rDNA silencing. The artificial tethering of repeats to the INM through a Sir2-Heh1 fusion suppresses rDNA instability in the absence of Lrs4, a protein required for peripheral tethering of the rDNA. Thus, the peripheral tethering appears to limit access of the repetitive arrays to the machinery mediating recombination and sister chromatid exchange (Mekhail et al. 2008).

adapts to changes in carbon source or encounters other conditions of metabolic stress. These changes reinforce the notion of “cell type-” or “condition-” specific levels of nuclear organization. Most importantly, yeast cells allow us to test for subtle changes in the efficiency of replication, DNA repair and transcription, as a result of changes in nuclear structure. Although the field is still at an early stage, the number of genes recognized as regulating aspects of nuclear structure increases constantly, and both microscopy and genomic assays improve at staggering rates. These are forceful arguments for pursuing yeast as a model in which to test the relationship between nuclear structure and function. ACKNOWLEDGMENTS

We would like to acknowledge critical reading by V. Dion and H. Ferreira, and many fruitful discussions with the Gasser laboratory. We thank S. Nagai for the DNA repair figure and K. Bystricky images in Figure 1. S.M.G. acknowledges support of the Novartis Research Foundation, the National Center for Competence in Research, “Frontiers-in-Genetics” and the EU NOE Epigenome. A.T. is supported by the French “Agence Nationale pour la Recherche” (ANR), and received funding from the European Research Council under the European Community’s Seventh Framework Programme (FP7/2007-2013 Grant Agreement no. [210508]). REFERENCES

CONCLUDING REMARKS

The yeast nucleus provides a readily manipulated genetic system for the analysis of nuclear organization. Its main advantage is that yeast provides ample means to test both gain and loss of function mutations on nuclear organization and genetic function. Data show clearly that there are zones that regulate DNA repair (Fig. 5B) just as there are zones that influence transcription (Fig. 5A). The one drawback of yeast is that it does not differentiate into highly differentiated cell types. It does, however, show changes in nuclear organization as the culture

Abruzzi KC, Belostotsky DA, Chekanova JA, Dower K, Rosbash M. 2006. 30 -end formation signals modulate the association of genes with the nuclear periphery as well as mRNP dot formation. EMBO J 25: 4253–4262. Ai W, Bertram PG, Tsang CK, Chan TF, Zheng XF. 2002. Regulation of subtelomeric silencing during stress response. Mol Cell 10: 1295–1305. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507– 17. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007. The molecular architecture of the nuclear pore complex. Nature 450: 695– 701. Andrulis ED, Neiman AM, Zappulla DC, Sternglanz R. 1998. Perinuclear localization of chromatin facilitates transcriptional silencing. Nature 394: 592– 595.

A. Taddei, H. Schober, and S.M. Gasser Andrulis ED, Zappulla DC, Ansari A, Perrod S, Laiosa CV, Gartenberg MR, Sternglanz R. 2002. Esc1, a nuclear periphery protein required for Sir4-based plasmid anchoring and partitioning. Mol Cell Biol 22: 8292– 8301. Antoniacci LM, Kenna MA, Skibbens RV. 2007. The nuclear envelope and spindle pole body-associated Mps3 protein bind telomere regulators and function in telomere clustering. Cell Cycle 6: 75– 79. Aparicio OM, Billington BL, Gottschling DE. 1991. Modifiers of position effect are shared between telomeric and silent mating-type loci in S. cerevisiae. Cell 66: 1279– 1287. Azam M, Lee JY, Abraham V, Chanoux R, Schoenly KA, Johnson FB. 2006. Evidence that the S.cerevisiae Sgs1 protein facilitates recombinational repair of telomeres during senescence. Nucleic Acids Res 34: 506 –516. Babiarz JE, Halley JE, Rine J. 2006. Telomeric heterochromatin boundaries require NuA4-dependent acetylation of histone variant H2A.Z in Saccharomyces cerevisiae. Genes Dev 20: 700 –710. Bennett CB, Lewis LK, Karthikeyan G, Lobachev KS, Jin YH, Sterling JF, Snipe JR, Resnick MA. 2001. Genes required for ionizing radiation resistance in yeast. Nat Genet 29: 426–434. Berger AB, Cabal GG, Fabre E, Duong T, Buc H, Nehrbass U, Olivo-Marin JC, Gadal O, Zimmer C. 2008. Highresolution statistical mapping reveals gene territories in live yeast. Nature Methods 5: 1031–1037. Bi X, Yu Q, Sandmeier JJ, Elizondo S. 2004. Regulation of transcriptional silencing in yeast by growth temperature. J Mol Biol 344: 893 –905. Blobel G. 1985. Gene gating: A hypothesis. Proc Natl Acad Sci 82: 8527–8529. Brickner JH, Walter P. 2004. Gene recruitment of the activated INO1 locus to the nuclear membrane. PLoS Biol 2: e342. Brickner DG, Cajigas I, Fondufe-Mittendorf Y, Ahmed S, Lee PC, Widom J, Brickner JH. 2007. H2A.Z-mediated localization of genes at the nuclear periphery confers epigenetic memory of previous transcriptional state. PLoS Biol 5: e81. Bryk M, Banerjee M, Murphy M, Knudsen KE, Garfinkel DJ, Curcio MJ. 1997. Transcriptional silencing of Ty1 elements in the RDN1 locus of yeast. Genes Dev 11: 255–269. Bupp JM, Martin AE, Stensrud ES, Jaspersen SL. 2007. Telomere anchoring at the nuclear periphery requires the budding yeast Sad1-UNC-84 domain protein Mps3. J Cell Biol 179: 845 –854. Burgess RC, Rahman S, Lisby M, Rothstein R, Zhao X. 2007. The Slx5-Slx8 complex affects sumoylation of DNA repair proteins and negatively regulates recombination. Mol Cell Biol 27: 6153–6162. Bystricky K, Heun P, Gehlen L, Langowski J, Gasser SM. 2004. Long-range compaction and flexibility of interphase chromatin in budding yeast analyzed by highresolution imaging techniques. Proc Natl Acad Sci U S A 101: 16495– 16500. Bystricky K, Laroche T, van Houwe G, Blaszczyk M, Gasser SM. 2005. Chromosome looping in yeast: telomere pairing and coordinated movement reflect anchoring

efficiency and territorial organization. J Cell Biol 168: 375 –387. Bystricky K, Van Attikum H, Montiel MD, Dion V, Gehlen L, Gasser SM. 2009. Regulation of nuclear positioning and dynamics of the silent mating type loci by the yeast Ku70/80 complex. Mol Cell Biol 29: 835 –848. Cabal GG, Genovesio A, Rodriguez-Navarro S, Zimmer C, Gadal O, Lesne A, Buc H, Feuerbach-Fournier F, OlivoMarin JC, Hurt EC, et al. 2006. SAGA interacting factors confine sub-diffusion of transcribed genes to the nuclear envelope. Nature 441: 770– 773. Casolari JM, Brown CR, Drubin DA, Rando OJ, Silver PA. 2005. Developmentally induced changes in transcriptional program alter spatial organization across chromosomes Genes Dev 19: 1188–98. Casolari JM, Brown CR, Komili S, West J, Hieronymus H, Silver PA. 2004. Genome-wide localization of the nuclear transport machinery couples transcriptional status and nuclear organization. Cell 117: 427– 439. Collins SR, Kemmeren P, Zhao XC, Greenblatt JF, Spencer F, Holstege FC, Weissman JS, Krogan NJ. 2007a. Toward a comprehensive atlas of the physical interactome of Saccharomyces cerevisiae. Mol Cell Proteomics 6: 439–450. Collins SR, Miller KM, Maas NL, Roguev A, Fillingham J, Chu CS, Schuldiner M, Gebbia M, Recht J, Shales M et al. 2007b. Functional dissection of protein complexes involved in yeast chromosome biology using a genetic interaction map. Nature 446: 806 –810. Cremer T, Cremer M, Dietzel S, Mu¨ller S, Solovei I, Fakan S. 2006. Chromosome territories – a functional nuclear landscape. Curr Opin Cell Biol 18: 307 –316. D’Angelo MA, Hetzer MW. 2008. Structure, dynamics and function of nuclear pore complexes. Trends Cell Biol 18: 456 –466. Dequard-Chablat M, Riva M, Carles C, Sentenac A. 1991. RPC19, the gene for a subunit common to yeast RNA polymerases A (I) and C (III). J Biol Chem 266: 15300– 15307. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306– 1311. Ebrahimi H, Donaldson AD. 2008. Release of yeast telomeres from the nuclear periphery is triggered by replication and maintained by suppression of Ku-mediated anchoring. Genes Dev 22: 3363–3374. Fabre E, Muller H, Therizols P, Lafontaine I, Dujon B, Fairhead C. 2005. Comparative genomics in hemiascomycete yeasts: Evolution of sex, silencing, and subtelomeres. Mol Biol Evol 22: 856–873. Fisher TS, Taggart AK, Zakian VA. 2004. Cell cycledependent regulation of yeast telomerase by Ku. Nat Struct Mol Biol 11: 1198– 1205. Fritze CE, Verschueren K, Strich R, Easton Esposito R. 1997. Direct evidence for SIR2 modulation of chromatin structure in yeast rDNA. EMBO J 16: 6495–6509. Gall JG. 2000. Cajal bodies: The first 100 years. Annu Rev Cell Dev Biol 16: 273 –300. Gasser SM, Hediger F, Taddei A, Neumann FR, Gartenberg MR 2004. The function of telomere clustering in yeast: The Circe effect. Cold Spring Harb Symp Quant Biol 69: 327 –337.

The Budding Yeast Nucleus Gartenberg MR, Neumann FR, Laroche T, Blaszczyk M, Gasser SM. 2004. Sir-mediated repression can occur independently of chromosomal and subnuclear contexts. Cell 119: 955– 967. Ghaemmaghami S, Huh WK, Bower K, Howson RW, Belle A, Dephoure N, O’Shea EK, Weissman JS. 2003. Global analysis of protein expression in yeast. Nature 425: 737–741. Gilson E, Roberge M, Giraldo R, Rhodes D, Gasser SM. 1993. Distortion of the DNA double helix by RAP1 at silencers and multiple telomeric binding sites. J Mol Biol 231: 293–310. Gotta M, Laroche T, Formenton A, Maillet L, Scherthan H, Gasser SM. 1996. The clustering of telomeres and colocalization with Rap1, Sir3, and Sir4 proteins in wild-type Saccharomyces cerevisiae. J Cell Biol 134: 1349–1363. Gottlieb S, Esposito RE. 1989. A new role for a yeast transcriptional silencer gene, SIR2, in regulation of recombination in ribosomal DNA. Cell 56: 771 –776. Gruenbaum Y, Margalit A, Goldman RD, Shumaker DK, Wilson KL. 2005. The nuclear lamina comes of age. Nat Rev Mol Cell Biol. 6: 21–31. Grund SE, Fischer T, Cabal GG, Antunez O, Perez-Ortin JE, Hurt E. 2008. The inner nuclear membrane protein Src1 associates with subtelomeric genes and alters their regulated gene expression. J Cell Biol 182: 897– 910. Gudmundsdottir K, Lord CJ, Ashworth A. 2007. The proteasome is involved in determining differential utilization of double-strand break repair pathways. Oncogene 26: 7601–7606. Haber JE, Debatisse M. 2006. Gene amplification: Yeast takes a turn. Cell 125: 1237–1240. Haeusler RA, Pratt-Hyatt M, Good PD, Gipson TA, Engelke DR. 2008. Clustering of yeast tRNA genes is mediated by specific association of condensin with tRNA gene transcription complexes. Genes Dev 22: 2204– 2214. Hattier T, Andrulis ED, Tartakoff AM. 2007. Immobility, inheritance and plasticity of shape of the yeast nucleus. BMC Cell Biol 8: 47. Hediger F, Neumann FR, Van Houwe G, Dubrana K, Gasser SM. 2002. Live imaging of telomeres: yKu and Sir proteins define redundant telomere-anchoring pathways in yeast. Curr Biol 12: 2076– 2089. Heun P, Laroche T, Shimada K, Furrer P, Gasser SM. 2001. Chromosome dynamics in the yeast interphase nucleus. Science 294: 2181–2186. Hiraga S, Botsios S, Donaldson AD. 2008. Histone H3 lysine 56 acetylation by Rtt109 is crucial for chromosome positioning. J Cell Biol 183: 641–651. Hiraga S, Robertson ED, Donaldson AD. 2006. The Ctf18 RFC-like complex positions yeast telomeres but does not specify their replication time. EMBO J 25: 1505– 1514. Hug N, Lingner J. 2006. Telomere length homeostasis. Chromosoma. 115: 413– 25. Hughes JD, Estep PW, Tavazoie S, Church GM. 2000. Computational identification of cis-regulatory elements associated with groups of functionally related genes in Saccharomyces cerevisiae. J Mol Biol 296: 1205–1214.

Huh WK, Falvo JV, Gerke LC, Carroll AS, Howson RW, Weissman JS, O’Shea EK. 2003. Global analysis of protein localization in budding yeast. Nature 425: 686 –691. Iborra FJ. 2007. Can visco-elastic phase separation, macromolecular crowding and colloidal physics explain nuclear organisation? Theor Biol Med Model 4: 15. Ishii K, Arib G, Lin C, Van Houwe G, Laemmli UK. 2002. Chromatin boundaries in budding yeast: The nuclear pore connection. Cell 109: 551 –562. Jaspersen SL, Giddings THJr, Winey M. 2002. Mps3p is a novel component of the yeast spindle pole body that interacts with the yeast centrin homologue Cdc31p. J Cell Biol 159: 945– 956. Jin QW, Fuchs J, Loidl J. 2000. Centromere clustering is a major determinant of yeast interphase nuclear organization. J Cell Sci 113: 1903–1912. Kadyk LC, Hartwell LH. 1992. Sister chromatids are preferred over homologs as substrates for recombinational repair in Saccharomyces cerevisiae. Genetics 132: 387 –402. Kalocsay M, Hiller NJ, Jentsch S. 2009. Chromosome-wide Rad51 spreading and SUMO-H2A.Z-dependent chromosome fixation in response to a persistent DNA doublestrand break. Mol Cell 33: 335 –343. Keil RL, Roeder GS. 1984. Cis-acting, recombinationstimulating activity in a fragment of the ribosomal DNA of S. cerevisiae. Cell 39: 377– 386. Khadaroo B, Teixeira MT, Luciano P, Eckert-Boulet N, Germann SM, Simon MN, Gallina I, Abdallah P, Gilson E, Ge´li V, et al. 2009. The DNA damage response at eroded telomeres and tethering to the nuclear pore complex. Nat Cell Biol Jul 13. [Epub ahead of print] King MC, Lusk CP, Blobel G. 2006. Karyopherin-mediated import of integral inner nuclear membrane proteins. Nature 442: 1003– 1007. Kitamura E, Blow JJ, Tanaka TU. 2006. Live-cell imaging reveals replication of individual replicons in eukaryotic replication factories. Cell, 125: 1297– 1308. Kobayashi T, Heck DJ, Nomura M, Horiuchi T. 1998. Expansion and contraction of ribosomal DNA repeats in Saccharomyces cerevisiae: Requirement of replication fork blocking (Fob1) protein and the role of RNA polymerase I. Genes Dev 12: 3821–3830. Krogan N, Lam MHY, Fillingham J, Keogh MC, Gebbia M, Li J, Datta N, Cagney G, Buratowski S, Emili A, et al. 2004. Proteasome involvement in the repair of DNA doublestrand breaks. Molecular Cell 16: 1027– 1034. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Mol Cell Biol 4: 605–612. Larrivee M, LeBel C, Wellinger RJ. 2004. The generation of proper constitutive G-tails on yeast telomeres is dependent on the MRX complex. Genes Dev 18: 1391–1396. Lendvay TS, Morris DK, Sah J, Balasubramanian B, Lundblad V. 1996. Senescence mutants of Saccharomyces cerevisiae with a defect in telomere replication identify three additional EST genes. Genetics 144: 1399– 1412. Lewis A, Felberbaum R, Hochstrasser M. 2007. A nuclear envelope protein linking nuclear pore basket assembly, SUMO protease regulation, and mRNA surveillance. J Cell Biol 178: 813– 27.

A. Taddei, H. Schober, and S.M. Gasser Lingner J, Cech TR, Hughes TR, Lundblad V. 1997. Three ever shorter telomere (EST) genes are dispensable for in vitro yeast telomerase activity. Proc Natl Acad Sci 94: 11190–11195. Lisby M, Mortensen UH, Rothstein R. 2003. Colocalization of multiple DNA double-strand breaks at a single Rad52 repair centre. Nat Cell Biol 5: 572– 577. Loeillet S, Palancade B, Cartron M, Thierry A, Richard GF, Dujon B, Doye V, Nicolas A. 2005. Genetic network interactions among replication, repair and nuclear pore deficiencies in yeast. DNA Repair (Amst) 4: 459– 468. Lundblad V, Szostak JW. 1989. A mutant with a defect in telomere elongation leads to senescence in yeast. Cell 57: 633 –643. Lusk CP, Blobel G, King MC. 2007. Highway to the inner nuclear membrane: rules for the road. Nat Rev Mol Cell Biol 8: 414– 420. Maillet L, Boscheron C, Gotta M, Marcand S, Gilson E, Gasser SM. 1996. Evidence for silencing compartments within the yeast nucleus: A role for telomere proximity and Sir protein concentration in silencer-mediated repression. Genes Dev 10: 1796– 1811. Maillet L, Gaden F, Brevet V, Fourel G, Martin SG, Dubrana K, Gasser SM, Gilson E. 2001. Ku-deficient yeast strains exhibit alternative states of silencing competence. EMBO Rep 2: 203 –210. Marcand S, Gilson E, Shore D. 1997. A protein-counting mechanism for telomere length regulation in yeast. Science 275: 986– 990. Marcand S, Buck SW, Moretti P, Gilson E, Shore D. 1996. Silencing of genes at nontelomeric sites in yeast is controlled by sequestration of silencing factors at telomeres by Rap 1 protein. Genes Dev 10: 1297– 1309. Marshall WF, Straight A, Marko JF, Swedlow J, Dernburg A, Belmont A, Murray AW, Agard DA, Sedat JW. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930– 939. Martin SG, Laroche T, Suka N, Grunstein M, Gasser SM. 1999. Relocalization of telomeric Ku and SIR proteins in response to DNA strand breaks in yeast. Cell 97: 621–633. Martino F, Kueng S, Robinson P, Tsai-Pflugfelder M, van Leeuwen F, Ziegler M, Cubizolles F, Cockell MM, Rhodes D, Gasser SM. 2009. Reconstitution of yeast silent chromatin: Multiple contact sites and O-AADPR binding load SIR complexes onto nucleosomes in vitro. Mol Cell 33: 323 –334. McAinsh AD, Scott-Drew S, Murray JA, Jackson SP. 1999. DNA damage triggers disruption of telomeric silencing and Mec1p-dependent relocation of Sir3p. Curr Biol 9: 963–966. McEachern MJ, Haber JE. 2006. Break-induced replication and recombinational telomere elongation in yeast. Annu Rev Biochem 75: 111–135. Mekhail K, Seebacher J, Gygi SP, Moazed D. 2008. Role for perinuclear chromosome tethering in maintenance of genome stability. Nature 456: 667 –670. Meneghini MD, Wu M, Madhani HD. 2003. Conserved histone variant H2A.Z protects euchromatin from the ectopic spread of silent heterochromatin. Cell 112: 725–736.

Menon BB, Sarma NJ, Pasula S, Deminoff SJ, Willis KA, Barbara KE, Andrews B, Santangelo GM. 2005. Reverse recruitment: the Nup84 nuclear pore subcomplex mediates Rap1/Gcr1/Gcr2 transcriptional activation. Proc Natl Acad Sci 102: 5749–5754. Mercier G, Berthault N, Touleimat N, Kepes F, Fourel G, Gilson E, Dutreix M. 2005. A haploid-specific transcriptional response to irradiation in Saccharomyces cerevisiae. Nucleic Acids Res 33: 6635–6643. Miele A, Bystricky K, Dekker J. 2009. Yeast silent mating type loci form heterochromatic clusters through silencer protein-dependent long-range interactions. PLoS Genet 5: e1000478. Epub 2009 May 8. Mills K, Sinclair D, Guarente L. 1999. MEC1-dependent redistribution of the Sir3 silencing protein from telomeres to DNA double-strand breaks. Cell 97: 609 –620. Misteli T. 2000. Cell biology of transcription and pre-mRNA splicing: Nuclear architecture meets nuclear function. J Cell Sci 113: 1841– 1849. Mondoux MA, Scaife JG, Zakian VA. 2007. Differential nuclear localization does not determine the silencing status of Saccharomyces cerevisiae telomeres. Genetics 177: 2019–2029. Moretti P, Freeman K, Coodly L, Shore D. 1994. Evidence that a complex of SIR proteins interacts with the silencer and telomere-binding protein RAP1. Genes Dev 8: 2257–2269. Nagai S, Dubrana K, Tsai-Pflugfelder M, Davidson MB, Roberts TM, Brown GW, Varela E, Hediger F, Gasser SM, Krogan NJ. 2008. Functional targeting of DNA damage to a nuclear pore-associated SUMO-dependent ubiquitin ligase. Science 322: 597– 602. New JH, Sugiyama T, Zaitseva E, Kowalczykowski SC. 1998. Rad52 protein stimulates DNA strand exchange by Rad51 and replication protein A. Nature 391: 407– 410. Nishikawa S, Terazawa Y, Nakayama T, Hirata A, Makio T, Endo T. 2003. Nep98p is a component of the yeast spindle pole body and essential for nuclear division and fusion. J Biol Chem 278: 9938–9943. Oakes M, Aris JP, Brockenbrough JS, Wai H, Vu L, Nomura M. 1998. Mutational analysis of the structure and localization of the nucleolus in the yeast Saccharomyces cerevisiae. J Cell Biology 143: 23– 34. Oakes M, Nogi Y, Clark MW, Nomura M. 1993. Structural alterations of the nucleolus in mutants of Saccharomyces cerevisiae defective in RNA polymerase I. Mol Cell Biol 13: 2441–2455. O’Sullivan JM, Tan-Wong SM, Morillon A, Lee B, Coles J, Mellor J, Proudfoot NJ. 2004. Gene loops juxtapose promoters and terminators in yeast. Nat Genet 36: 1014–1018. Oza P, Jaspersen SL, Miele A, Dekker J, Peterson CL. 2009. Mechanisms that regulate localization of a DNA doublestrand break to the nuclear periphery. Genes Dev 23: 912 –927. Palancade B, Liu X, Garcia-Rubio M, Aguilera A, Zhao X, Doye V. 2007. Nucleoporins prevent DNA damage accumulation by modulating Ulp1-dependent sumoylation processes. Mol Biol Cell 18: 2912– 2923.

The Budding Yeast Nucleus Pan X, Ye P, Yuan DS, Wang X, Bader JS, Boeke JD. 2006. A DNA integrity network in the yeast Saccharomyces cerevisiae. Cell 124: 1069–1081. Perry JJ, Tainer JA, Boddy MN. 2008. A SIM-ultaneous role for SUMO and ubiquitin. Trends Biochem Sci 33: 201–208. Powers T, Walter P. 1999. Regulation of ribosome biogenesis by the rapamycin-sensitive TOR-signaling pathway in Saccharomyces cerevisiae. Mol Biol Cell 10: 987–1000. Prudden J, Pebernard S, Raffa G, Slavin DA, Perry JJ, Tainer JA, McGowan CH, Boddy MN. 2007. SUMO-targeted ubiquitin ligases in genome stability. EMBO J 26: 4089– 4101. ¨ ber Zelltheilung. Morphologisches Jahrbuch Rabl C. 1885. U 10: 214 –330. Ray A, Runge KW. 1999. Varying the number of telomerebound proteins does not alter telomere length in tel1D cells. Proc Natl Acad Sci U S A 96: 15044–15049. Renauld H, Aparicio OM, Zierath PD, Billington BL, Chhablani SK, Gottschling DE. 1993. Silent domains are assembled continuously from the telomere and are defined by promoter distance and strength, and by SIR3 dosage. Genes Dev 7: 1133– 1145. Richter K, Nessling M, Lichter P. 2007. Experimental evidence for the influence of molecular crowding on nuclear architecture. J Cell Sci 120: 1673–1680. Robyr D, Suka Y, Xenarios I, Kurdistani SK, Wang A, Suka N, Grunstein M. 2002. Microarray deacetylation maps determine genome-wide functions for yeast histone deacetylases. Cell 109: 437 –446. Rodley CD, Bertels F, Jones B, O’Sullivan JM. 2009. Global identification of yeast chromosome interactions using Genome conformation capture. Fungal Genet Biol 46: 879–886. Roy R, Meier B, McAinsh AD, Feldmann HM, Jackson SP. 2004. Separation-of-function mutants of yeast Ku80 reveal a Yku80p-Sir4p interaction involved in telomeric silencing. J Biol Chem 279: 86–94. Schmid M, Arib G, Laemmli C, Nishikawa J, Durussel T, Laemmli UK. 2006. Nup-PI: the nucleopore-promoter interaction of genes in yeast. Mol Cell 21: 379 –91. Schober H, Ferreira H, Kalck V, Gehlen LR, Gasser SM. 2009. Yeast telomerase and the SUN domain protein Mps3 anchor telomeres and repress subtelomeric recombination. Genes Dev 23: 928– 938. Schober H, Kalck V, Vega-Palas MA, Van Houwe G, Sage D, Unser M, Gartenberg MR, Gasser SM. 2008. Controlled exchange of chromosomal arms reveals principles driving telomere interactions in yeast. Genome Res 18: 261 –271. Sexton T, Umlauf D, Kurukuti S, Fraser P. 2007. The role of transcription factories in large-scale structure and dynamics of interphase chromatin. Semin Cell Dev Biol 18: 691 –697. Shampay J, Szostak JW, Blackburn EH. 1984. DNA sequences of telomeres maintained in yeast. Nature 310: 154–157. Shore D, Nasmyth K. 1987. Purification and cloning of a DNA binding protein from yeast that binds to both silencer and activator elements. Cell 51: 721– 732. Sinclair DA, Guarente L. 1997. Extrachromosomal rDNA circles –a cause of aging in yeast. Cell 91: 1033– 1042.

Singer MS, Gottschling DE. 1994. TLC1: template RNA component of Saccharomyces cerevisiae telomerase. Science 266: 404–409. Sirri V, Urcuqui-Inchima S, Roussel P, Hernandez-Verdun D. 2008. Nucleolus: The fascinating nuclear body. Histochem Cell Biol 129: 13– 31. Smith GP. 1974. Unequal crossover and the evolution of multigene families. Cold Spring Harb Symp Quant Biol 38: 507– 513. Smith JS, Boeke JD. 1997. An unusual form of transcriptional silencing in yeast ribosomal DNA. Genes Dev 11: 241 –254. Stavenhagen JB, Zakian VA. 1994. Internal tracts of telomeric DNA act as silencers in Saccharomyces cerevisiae. Genes Dev 8: 1411–1422. Stellwagen AE, Haimberger ZW, Veatch JR, Gottschling DE. 2003. Ku interacts with telomerase RNA to promote telomere addition at native and broken chromosome ends. Genes Dev 17: 2384– 2395. Stone EM, Pillus L. 1996. Activation of an MAP kinase cascade leads to Sir3p hyperphosphorylation and strengthens transcriptional silencing. J Cell Biol 135: 571– 583. Strahl-Bolsinger S, Hecht A, Luo K, Grunstein M. 1997. SIR2 and SIR4 interactions differ in core and extended telomeric heterochromatin in yeast. Genes Dev 11: 83– 93. Taddei A. 2007. Active genes at the nuclear pore complex. Curr Opin Cell Biol 19: 305– 310. Taddei A, Hediger F, Neumann FR, Bauer C, Gasser SM. 2004. Separation of silencing from perinuclear anchoring functions in yeast Ku80, Sir4 and Esc1 proteins. EMBO J 23: 1301–1312. Taddei A, Van Houwe G, Hediger F, Kalck V, Cubizolles F, Schober H, Gasser SM. 2006. Nuclear pore association confers optimal expression levels for an inducible yeast gene. Nature 441: 774–778. Taddei A, Van Houwe G, Nagai S, Erb I, van Nimwegen E, Gasser SM. 2009. The functional importance of telomere clustering: global changes in gene expression result from SIR factor dispersion. Genome Res 19: 611– 625. Therizols P, Fairhead C, Cabal GG, Genovesio A, OlivoMarin JC, Dujon B, Fabre E. 2006. Telomere tethering at the nuclear periphery is essential for efficient DNA double strand break repair in subtelomeric region. J Cell Biol 172: 189– 199. Thompson JS, Johnson LM, Grunstein M. 1994. Specific repression of the yeast silent mating locus HMR by an adjacent telomere. Mol Cell Biol 14: 446 –455. Thompson M, Haeusler RA, Good PD, Engelke DR. 2003. Nucleolar clustering of dispersed tRNA genes. Science 302: 1399– 1401. Torres-Rosell J, Sunjevaric I, De Piccoli G, Sacher M, EckertBoulet N, Reid R, Jentsch S, Rothstein R, Aragon L, Lisby M. 2007. The Smc5-Smc6 complex and SUMO modification of Rad52 regulates recombinational repair at the ribosomal gene locus. Nat Cell Biol 9: 923– 931. Verheggen C, Mouaikel J, Thiry M, Blanchard JM, Tollervey D, Bordonne R, Lafontaine DL, Bertrand E. 2001. Box C/ D small nucleolar RNA trafficking involves small nucleolar RNP proteins, nucleolar factors and a novel nuclear domain. EMBO J 20: 5480–5490.

A. Taddei, H. Schober, and S.M. Gasser Wang Z, Jones GM, Prelich G. 2006. Genetic analysis connects SLX5 and SLX8 to the SUMO pathway in Saccharomyces cerevisiae. Genetics 172: 1499– 1509. Wang L, Haeusler RA, Good PD, Thompson M, Nagar S, Engelke DR. 2005. Silencing near tRNA genes requires nucleolar localization. J Biol Chem 280: 8637–8639. Wotton D, Shore D. 1997. A novel Rap1p-interacting factor, Rif2p, cooperates with Rif1p to regulate telomere length in Saccharomyces cerevisiae. Genes Dev 11: 748– 760. Xie Y, Kerscher O, Kroetz MB, McConchie HF, Sung P, Hochstrasser M. 2007. The yeast Hex3.Slx8 heterodimer is a ubiquitin ligase stimulated by substrate sumoylation. J Biol Chem 282: 34176–34184. Yang CH, Lambie EJ, Hardin J, Craft J, Snyder M. 1989. Higher order structure is present in the yeast nucleus: autoantibody probes demonstrate that the nucleolus

lies opposite the spindle pole body. Chromosoma 98: 123 –128. Zacharioudakis I, Gligoris T, Tzamarias D. 2007. A yeast catabolic enzyme controls transcriptional memory. Curr Biol 17: 2041–2046. Zakian VA. 1996. Structure, function, and replication of Saccharomyces cerevisiae telomeres. Annu Rev Genet 30: 141 –72. Zhong S, Salomoni P, Pandolfi PP. 2000. The transcriptional role of PML and the nuclear body. Nat Cell Biol 2:, E85–90. Zhu C, Byers KJ, McCord RP, Shi Z, Berger MF, Newburger DE, Saulrieta K, Smith Z, Shah MV, Radhakrishnan M et al. 2009. High-resolution DNA-binding specificity analysis of yeast transcription factors. Genome Res 19: 556 –566.

Nuclear Functions of Actin Neus Visa1 and Piergiorgio Percipalle2 1

Department of Molecular Biology & Functional Genomics, Stockholm University, SE-106 91, Stockholm, Sweden

2

Department of Cell and Molecular Biology, Karolinska Institutet, SE-171 77, Stockholm, Sweden

Correspondence: [email protected] and [email protected]

Actin participates in several essential processes in the cell nucleus. Even though the presence of actin in the nucleus was proposed more than 30 years ago, nuclear processes that require actin have been only recently identified. Actin is part of chromatin remodeling complexes; it is associated with the transcription machineries; it becomes incorporated into newly synthesized ribonucleoproteins; and it influences long-range chromatin organization. As in the cytoplasm, nuclear actin works in conjunction with different types of actin-binding proteins that regulate actin function and bridge interactions between actin and other nuclear components.

ctin is a highly conserved protein of approximately 42 kDa found in all eukaryotic cells. It is a major component of the cytoskeleton and plays fundamental roles in essential biological processes such as determining cell shape, cell migration, and intracellular trafficking. The functions of actin in the cytoplasm are intrinsically coupled to the dynamics of actin polymerization, which is a tightly regulated process that responds to extracellular signals (reviewed by Moustakas and Heldin 2008; Papakonstanti and Stournaras 2008). Early studies raised the possibility that actin is also present in the cell nucleus and is implicated in the expression of protein-coding genes (Scheer et al. 1984; Egly et al. 1984). However, the existence of the so-called “nuclear actin” was

A

initially met with massive skepticism (reviewed by Pederson and Aebi 2002). Biochemists could not rule out contamination artifacts in nuclear preparations because of the high abundance of actin in the cytoplasm, and microscopists could not visualize in the cell nucleus the conspicuous actin filaments that are commonly observed in the cytoplasm. Nevertheless, research performed in the last 10 years has provided convincing evidence for the existence of actin in the cell nucleus and for the involvement of actin in fundamental nuclear processes. Actin is part of the chromatin remodeling complex; it is associated with the transcription machineries; it associates with newly synthesized ribonucleoproteins; and it influences long-range chromatin organization.

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000620 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000620

N. Visa and P. Percipalle

ACTIN AND CHROMATIN REMODELING

Actin participates in gene expression as a component of chromatin-modifying complexes. Early findings by Crabtree and coworkers revealed that actin interacts with Brg1, the ATPase subunit of the BAF (Brg or Brm Associated Factors) SWI/SNF-like chromatin remodeling complex (Zhao et al. 1998). Since then, b-actin and a considerable number of actin-related proteins (ARPs) have been identified as components of different types of chromatin remodeling and histone acetyltransferase (HAT) complexes in a wide range of organisms from yeast to man (reviewed by Olave et al. 2002; Chen and Shen 2007; Farrants 2008). A central question that has not been fully answered concerns the mechanism(s) by which actin and ARPs contribute to chromatin remodeling. Not all chromatin remodeling complexes contain actin or ARPs, which indicates that these proteins are not essential for chromatin remodeling per se. The fact that actin and ARPs often bind to the ATPase subunit of the chromatin remodeling complexes suggests that actin and ARPs work as allosteric regulators of chromatin remodeling (Blessing et al. 2004). In agreement with this possibility, Zhao et al. (1998) showed that actin is needed for the full ATPase activity of the BAF complex. Actin also mediates or facilitates the binding of BAF to chromatin (Zhao et al. 1998). The ability of Brg1 to bind actin filaments in vitro (Rando et al. 2002) and the association of actin with the RNA polymerase (see the following discussion) suggest that actin recruits BAF to transcribed genes by bridging the interaction between the chromatin remodeling complex and the transcription machinery. In other cases, the binding of ARPs to core histones might bridge the interaction (reviewed by Blessing et al. 2004). Independently of its function in chromatin remodeling, actin participates in the regulation of a subset of genes that code for components of the actin cytoskeleton and that are activated by the serum response factor (SRF) (reviewed by Farrants 2008; Vartiainen 2008). This function of actin is mediated by MAL, a transcriptional coactivator of SRF that can sense variations in

the cellular concentration of G-actin (Miralles et al. 2003). MAL is an actin-binding protein that shuttles continuously between the nucleus and the cytoplasm, and only the actin-bound form of MAL is efficiently exported from the nucleus. Actin regulates both the nucleocytoplasmic transport of MAL and the ability of the MAL-SRF complex to activate SRF target genes. Serum stimulation reduces the rate of MAL nuclear export. Under these conditions, MAL and SRF associate with the SRF target promoters, and the SRF target genes become activated upon disruption of the MAL-actin interaction (Vartiainen et al. 2007). ACTIN IN ASSOCIATION WITH THE TRANSCRIPTION MACHINERY

Actin is associated with all three RNA polymerases, Pol I, II, and III (Fomproix and Percipalle 2004; Philimonenko et al. 2004; Hofmann et al. 2004; Hu et al. 2004; Kukalev et al. 2005). The actin – polymerase interaction has not been characterized at the structural level. Three Pol III subunits have been detected in association with actin in coimmunoprecipitation experiments (Hu et al. 2004). It has been proposed that two of them, RPABC2 and RPABC3, build the actin-binding site, because these subunits are common to all three polymerases and are located close to each other at the surface of the polymerase (Cramer et al. 2001; Hu et al. 2004). Coimmunoprecipitation experiments also showed that actin can interact with the largest Pol I subunit and with the carboxy-terminal domain (CTD) of the largest Pol II subunit (Fomproix and Percipalle 2004; Kukalev et al. 2005), which provides additional actin-binding sites specific for these polymerases. The interaction of actin with the RNA polymerases is functionally relevant. Anti-actin antibodies inhibit transcription by Pol I and Pol II in vivo and in vitro (Hofmann et al. 2004; Philimonenko et al. 2004), and actin restores the full activity of partially purified, inactive Pol III preparations in vitro (Hu et al. 2004). These observations indicate that actin associates with RNA polymerases that are engaged in transcription.

Nuclear Functions of Actin

The inhibitory effect of anti-actin antibodies on Pol I and II, as well as the ability of actin to activate Pol III, have been observed in vitro on naked DNA (Hofmann et al. 2004; Philimonenko et al. 2004; Hu et al. 2004), which suggests that actin is involved in basal transcription independently of its role in chromatin regulation. These observations have led to the proposal that actin is required for the activity of the basal transcription machinery. However, we are far from understanding how actin contributes to the transcription process and whether all three RNA polymerases use actin in the same manner. A relevant question is whether actin is required for transcription initiation or elongation. Actin occupancy at promoters of Pol I and Pol II genes and copurification of actin with preinitiation complexes (PIC) suggest that actin is required for the assembly of transcription-competent polymerases (Hofmann et al. 2004; Philimonenko et al. 2004; Percipalle et al. 2006; Obrdlik et al. 2008; Louvet and Percipalle 2009). However, in vitro rDNA transcription assays that discriminate between transcription initiation (defined operationally by the synthesis of the initial trinucleotide) and elongation show that actin and nuclear myosin 1 (NM1) are needed for Pol I transcription elongation (Philimonenko et al. 2004; Percipalle et al. 2006). An important contribution toward understanding the role of actin in association with the transcription machinery was provided by the finding that NM1 is also intimately associated with Pol I and Pol II (Pestic-Dragovich et al. 2000; Fomproix and Percipalle 2004; Hofmann et al. 2006a). NM1 is a short-tailed monomeric myosin that acts as an actindependent ATPase. NM1 was found on active nucleolar transcription sites of both interphase and mitotic HeLa cells (Fig. 1) (Fomproix and Percipalle 2004), and microinjection of anti-NM1 antibodies into living cells inhibited rRNA synthesis (Philimonenko et al. 2004). Furthermore, depletion of either actin or NM1 from transcription extracts repressed the ability of the extracts to support transcription in vitro (Hofmann et al. 2004; Philimonenko et al.

2004). The facts that endogenous actin and NM1 associate with promoter and coding regions of rRNA genes, that actin and NM1 are associated with the transcription machinery, and that impairment of the myosin ATPase activity down-regulates Pol I transcription led to the proposal that an actomyosin activity is required for Pol I transcription (Fomproix and Percipalle 2004; Philimonenko et al. 2004; Percipalle et al. 2006). A recent study by Ye and coworkers (2008) provided further experimental support to this hypothesis. Actin mutants that are deficient in polymerization failed to associate with Pol I and did not support Pol I transcription. NM1 mutants that were defective in ATP-binding, actin-binding, or calmodulinbinding failed to associate with the rDNA and with Pol I and finally, the association of actin and NM1 with Pol I was regulated by ATP hydrolysis (Ye et al. 2008). These observations have enabled a model to be proposed in which the actin-NM1 complex acts as a molecular motor that helps the transcription machinery slide along the rDNA (Ye et al. 2008). Such a motor should interact with the transcription machinery itself and with the DNA, directly or indirectly (Fig. 2A). The actin-Pol I interaction is well-documented. The direct interaction between NM1 and DNA in vitro (Hofmann et al. 2006b) might explain how the actin-NM1 complex fastens to the DNA. ACTIN IN NASCENT TRANSCRIPTS

Actin not only interacts with the transcription machineries, it also associates with the nascent transcripts (Percipalle et al. 2001; Percipalle et al. 2002). Immunohistochemistry experiments performed in the dipteran C. tentans provided initial insights into the cotranscriptional binding of actin to nascent transcripts, because antibodies to actin labeled many loci in polytene chromosome preparations but failed to label after treatment of the chromosomes with RNase A (Percipalle et al. 2001). Immunoelectron microscopy experiments on ultrathin sections of C. tentans salivary glands showed that actin was preferentially associated with the distal region of the active transcription unit,

N. Visa and P. Percipalle

Actin

Fibrillarin

Merge

NM1

Fibrillarin

Merge

NM1 pre-serum

Fibrillarin

Merge

Figure 1. Actin and NM1 localize to mammalian nucleoli. Homogeneous preparations of nucleoli from HeLa

cells were analyzed by immunofluorescence labeling and confocal microscopy using rabbit polyclonal antibodies to actin (scale bar, 2 mm) and NM1 (scale bar, 2 mm) or corresponding preimmune sera (scale bar, 1 mm). All the specimens were coimmunostained with a mouse monoclonal antibody to fibrillarin as marker for nucleoli.

away from the promoter (Percipalle et al. 2001). Later on, anti-actin antibodies coprecipitated coding regions of Pol I and Pol II genes in chromatin immunoprecipitation experiments (Hofmann et al. 2004; Philimonenko et al. 2004; Obrdlik et al. 2008), emphasizing the presence of actin along active genes. The connection between actin and RNAwas further shown at the molecular level by chromatin RNA immunoprecipitation assays, in which it was possible to analyze the cotranscriptional association of protein factors with nascent RNA (Obrdlik and Percipalle 2009; Obrdlik et al. 2008). Specific heterogeneous nuclear ribonucleoproteins (hnRNPs) bind actin and mediate its association with RNA. Actin-associated hnRNP proteins were discovered mainly by resolving

nuclear extracts or RNP preparations with DNase I affinity chromatography. In both C. tentans and mammals, a significant fraction of actin-associated hnRNPs belong to the A/B type family. The hnRNP A/B proteins contain two conserved RNA recognition motifs (RRMs) flanked at the amino terminus by an acidic domain and at the carboxyl terminus by a divergent module for protein– protein interactions, termed the “auxiliary domain” (Krecic and Swanson 1999; Dreyfuss et al. 2002). In C. tentans, actin is associated with the shuttling mRNA-binding protein hrp36 (Visa et al. 1996), whereas in mammals, in which the repertoire of A/B-type hnRNPs is somewhat larger (Krecic and Swanson 1999; Dreyfuss et al. 2002), actin associates with several proteins, including

Nuclear Functions of Actin

A

5¢

5¢

NM 1

Chromatin remodeling complex Actin Actin Actin Actin

Pol Actin

NM 1

Actin

Pol

Actin Actin

B

5¢

hnRNP proteins

5¢

Chromatin Actin remodeling complex 65/U HAT

5¢

Pol

Pol

Pol

Figure 2. Two possible ways by which actin contributes to transcription elongation. (A) An actomyosin motor

helps the transcription machinery slide along the DNA. The architecture of the motor is not well understood but we envisage that the actin-NM1 complex makes contact with both the transcription machinery and the DNA/ chromatin. Two possible configurations are depicted in the figure. (B) Actin participates in the recruitment of histone modifiers to protein-coding genes. Actin binds to hnRNP proteins and becomes incorporated into the nascent pre-mRNPs. Actin forms a complex with specific hnRNP adaptor proteins such as Hrp65 in Chironomus tentans and hnRNP U in mammals, and recruits HATs that acetylate histones and facilitate transcription elongation. The HATs might be components of larger chromatin remodeling complexes that might also establish direct contacts with actin.

hnRNP A2, hnRNPA3, and the transport mediator CArG-box binding factor (CBF-A) (Percipalle et al. 2002; Raju et al. 2008). Another actin-associated hnRNP protein of C. tentans, hrp65, shares strong homology with the human proteins PSF (Patton et al. 1993), p54nrb/NonO (Dong et al. 1993), PSP1 (Fox et al. 2002), and with the Drosophila NonA/BJ6 (Jones and Rubin 1990; von Besser et al. 1990). These proteins do not belong to the A/B-type and they are characterized by a central domain of about 320 amino acids

termed “DBHS” (Drosophila behavior and human splicing). This domain is evolutionarily conserved. The DBHS domain comprises two RRMs and an additional carboxy-terminal stretch of approximately 100 amino acids that mediates dimerization or oligomerization among DBHS proteins (Kiesler et al. 2003). The association of DBHS proteins with actin is conserved in mammals; the human PSF and NonO proteins have been identified as components of a large nuclear protein complex together with actin, Pol II, and N-WASP, a key

N. Visa and P. Percipalle

regulator of cytoplasmic microfilaments that is also present in the cell nucleus (Wu et al. 2006). Actin associates with hnRNP U/SAF-A (Scaffold attachment factor A), a large 120kDa tripartite protein with ATP-binding activity in mammals. The amino terminus of this protein displays DNA-binding activity, the carboxyl terminus contains two RRMs for RNA binding, and the central domain displays Pol II-binding activity (Kukalev et al. 2005). Some of the actin-associated hnRNP proteins identified so far interact directly with actin, whereas others are part of actin-containing complexes and are not in direct contact with actin. The C. tentans hrp65-2 (hrp65 isoform 2) and the human hnRNP U proteins interact with actin through a conserved motif (Percipalle et al. 2003; Kukalev et al. 2005). Remarkably, inhibition of the actin-hrp65-2 or actin-hnRNP U interactions in living cells inhibits transcription elongation by Pol II as monitored by incorporation of labeled nucleotide precursors in run-on assays (Percipalle et al. 2003; Kukalev et al. 2005). Based on these results, we proposed a regulatory mechanism in which actin facilitates the cross talk between chromatin, elongating polymerase, and nascent RNA (Louvet and Percipalle 2009). How do actin-hnRNP complexes facilitate transcription elongation? Experiments in insects and mammals suggest that specific actinhnRNP complexes facilitate the elongation phase by recruiting transcriptional coactivators to remodel the nucleosome barrier imposed by chromatin (Percipalle and Visa 2006; Miralles and Visa 2006; Louvet and Percipalle 2009). Actin-hrp65-2 and actin-hnRNP U complexes mediate the recruitment of the HATs p2D10 and PCAF, respectively, to transcribed genes (Fig. 2B) (Sjo¨linder et al. 2005; Obrdlik et al. 2008). The disruption of these actin-hnRNP interactions in vivo reduced the level of histone acetylation in chromatin, blocked Pol II transcription, and affected HAT occupancy along active genes (Sjo¨linder et al. 2005; Kukalev et al. 2005; Obrdlik et al. 2008). These results indicate that the integrity of specific actin-hnRNP

interactions is required for coactivator recruitment to active genes. The topology of the actin complex is not known. However, an antibody specific for the actin-binding site on hnRNP U coprecipitates endogenous hnRNP U and PCAF but fails to precipitate actin in mammals (Obrdlik et al. 2008). This leads us to favor the idea that hnRNP U is able to interact simultaneously with both PCAF and actin. There is also recent evidence that the HAT PCAF and BRG1, the ATPase subunit of the SWI/ SNF chromatin remodeling complex, are sequentially recruited to the promoter of the myogenin gene and are both involved in gene activation (Li et al. 2007). Actin is also a component of the SWI/SNF complex (reviewed in Farrants 2008). We speculate, therefore, that during elongation of nascent pre-mRNA, the actin-RNP complexes work as modules that facilitate recruitment of HATs and chromatin remodeling complexes to active genes (see Fig. 2B). Studies by Obrdlik and coworkers (2008) revealed that the interplay between actin and the nascent RNP also involves interaction with the RNA polymerase in some cases. During elongation, the carboxy-terminal domain (CTD) of the largest subunit of Pol II is subjected to a series of covalent modifications that modulate recruitment of transcriptional coactivators. The amino acid residues Ser2 and Ser5 within the conserved CTD heptapeptide repeats become combinatorially phosphorylated and the different phosphorylation states represent hallmarks for individual transcription steps (Meinhart et al. 2005). Therefore, if specific actin-hnRNP complexes are required for efficient elongation, it is likely that they are cotranscriptionally associated with specific CTD phosphorylation states. Recent evidence shows that this is the case. Actin together with hnRNP U associates only with Ser2-phosphorylated heptapeptide repeats or with heptapeptide repeats that are phosphorylated on both Ser5 and Ser2, and fails to interact with unphosphorylated heptapeptide repeats or with those repeats that are phosphorylated on Ser5 only (Obrdlik et al. 2008). These findings

Nuclear Functions of Actin

support the idea that specific actin-hnRNP complexes physically associate with the elongating Pol II, a view that is corroborated by evidence that PSF and p54nrb, mammalian homologs of the C. tentans hrp65 protein, also interact with actin and with the CTD (Emili et al. 2002; Wu et al. 2006). As discussed earlier, actin and NM1 are required for transcription elongation by Pol I, and NM1 also appears to mediate the association of specific chromatin-modifying components to the rDNA genes. NM1 is a core component of the B-WICH chromatin remodeling complex together with the Williams syndrome transcription factor (WSTF) and the ATPase SNF2 h (Percipalle et al. 2006), and this complex turned out to also contain precursor rRNA (Cavellan et al. 2006). Remarkably, inhibition of endogenous NM1 or WSTF in abortive initiation assays did not affect the formation of initial pre-rRNA transcripts but affected the synthesis of the entire transcript when chromatin templates were used in run-on transcription assays (Philimonenko et al. 2004; Percipalle et al. 2006). These results suggest that the core components of the B-WICH complex, including NM1, are required after transcription initiation, and a model has been proposed in which the dynamic interaction between actin and NM1 is required for local recruitment of the B-WICH complex to active ¨ stlund Farrants 2006; genes (Percipalle and O Louvet and Percipalle 2009). The previous considerations suggest that actin modulates the elongation of nascent pre-mRNAs and pre-rRNAs by controlling the cotranscriptional association of chromatinmodifying components with the active genes. This view is of particular interest in the case of Pol I transcription, where active rRNA genes are believed to be devoid of nucleosomes. For protein-coding genes, it is intriguing that actin together with hnRNP U and the HAT PCAF associate with the CTD in a phosphorylated Ser2dependent manner (Obrdlik et al. 2008), an observation that strengthens the hypothesis that the coupling of actin to the elongating Pol II facilitates the modifications of chromatin structure.

ACTIN BEYOND TRANSCRIPTION

The studies performed in C. tentans showed that actin is cotranscriptionally added to nascent pre-mRNA, remains incorporated in mature RNPs released into the nucleoplasmic milieu, and accompanies the mRNA all the way to polysomes (Percipalle et al. 2001). There is no evidence of nuclear actin-based motors promoting directional movement of RNP particles or even preribosomal subunits (Singh et al. 1999; Politz et al. 2003; Siebrasse et al. 2008). However, actin is associated with mature mRNPs and with pre-60S subunits, it decorates pore-linked filaments that project inward into the nucleoplasm from the nuclear pore basket, and it has been implicated in controlling the export of viral mRNA transcripts (Percipalle et al. 2001, 2002; Hofmann et al. 2001; Kiseleva et al. 2004; Oeffinger et al. 2007). Also, NM1 is loaded into preribosomes and involved in the export-competent pre-60S subunits (Obrdlik et al. 2009). In addition, NM1 accompanies preribosomal subunits in transit to the nuclear envelope (Cisterna et al. 2007; Obrdlik et al. 2009), is present at the nuclear envelope (Holaska et al. 2007), and has been recently incorporated in the export of small ribosomal subunits (Cisterna et al. 2009). Finally, there is recent evidence that a complex containing rRNA and NM1 is associated with the RNA-binding protein Nup153 at the nuclear pore (Obrdlik et al. 2009), which suggests that NM1 accompanies the rRNA from the gene to the nuclear pore complex. These observations suggest that the synergy between actin and NM1 plays a more general role, and is not limited to the gene. We favor the idea that actin and NM1 contribute to the maturation and assembly of export-competent mRNPs and rRNPs by promoting the remodeling of RNP structures through dynamic protein–protein interactions (reviewed in Percipalle et al. 2009).

REGULATION OF ACTIN POLYMERIZATION

The presence of actin in the cell nucleus raises the possibility that there are nuclear-based mechanisms regulating actin polymerization

N. Visa and P. Percipalle

(Pederson 2008; Louvet and Percipalle 2009). It has been suggested that nuclear actin coexists in monomeric (G) or short oligomeric forms and in a polymeric form (Louvet and Percipalle 2009; Gieni & Hendzel 2009). It is unclear how transitions between these forms occur, but dynamic alterations in actin polymerization may be instrumental in modulating the function of actin in RNA biogenesis. Canonical actin filaments, which are commonly seen in the cytoplasm by immunofluorescence methods, have not been convincingly visualized in the cell nucleus. However, many of the factors that are known to control cytoplasmic actin polymerization and stabilize filamentous F-actin structures have been detected in the cell nucleus. N-WASP and the ARP2/3 complexes cooperate in the nucleation and branching of new actin filaments. These proteins are present in the cell nucleus, where they are implicated in the elongation of nascent mRNA transcripts (Wu et al. 2006; Yoo et al. 2007). More recently, other proteins that facilitate cytoplasmic actin nucleation independently of N-WASP and ARP2/3 have been discovered in the nucleus, including the forminlike mDia proteins and JMY (Miki et al. 2008; Zuchero et al. 2009). These findings indeed suggest the presence of polymeric actin in the nucleus, and it is likely that there are several independent mechanisms that regulate nuclear actin polymerization, as is the case in the cytoplasm. It is possible that the need to have several mechanisms for actin polymerization reflects the multiple roles that actin plays in the cell nucleus. This may or may not be the case, but it is clear that nuclear actin assembled into higher order structures is rather dynamic with a high turnover (McDonald et al. 2006), which indicates that a pool of monomeric actin is also present in the nucleus. This pool is probably necessary to feed dynamic polymeric actin structures. The existence of monomeric actin was indirectly suggested by the discovery of G-actin binding proteins such as cofilin, profilin, b-thymosin, and a gelsolin-like protein in the cell nucleus (Prendergast et al. 1991; Pendleton et al. 2003; Skare et al. 2003; Huff et al. 2004). Actin can be copurified with hnRNP U and

the Pol II machinery using DNase I affinity chromatography (Kukalev et al. 2005; Obrdlik et al. 2008), which has higher affinity for monomeric than for filamentous actin (Zechel 1980). In addition, monoclonal antibodies designed to recognize epitopes exclusively available in monomeric actin or actin dimers revealed significant immunostaining of the cell nucleus (reviewed in Jokusch et al. 2006). There is also evidence that monomeric or short oligomeric actin is directly involved in nuclear functions such as the role of monomeric actin in the regulation of SRF-dependent transcription in complex with the coactivator MAL (Vartiainen et al. 2007). The G-actin binding protein profilin affects transcription of rRNA genes, and drugs that inhibit actin polymerization, such as cytochalasin D and latrunculin B, inhibit the transcription of rDNA genes in vivo and in vitro, whereas drugs that favor actin assembly do not influence rDNA transcription (Ye et al. 2008). Furthermore, a recent study indicated that actin polymerization takes place in retinoic acid (RA)-induced HoxB transcription (Ferrai et al. 2009). These observations suggest that polymeric actin is functional in the cell nucleus. Because N-WASP and ARP2/3 are important during premRNA elongation, and an actin-NM1 complex is clearly implicated in transcription elongation by Pol I, we favour the idea that both monomeric and polymeric actin are present along active genes, and that cotranscriptional transitions in the actin polymerization states result in the existence of different actin forms associated with different parts of the gene. What is the polymerization state of actin in RNPs? Ultrastructural analysis has not revealed canonical filamentous actin structures attached to mature RNPs (reviewed in Percipalle 2009), which hints at an intrinsic mechanism that keeps actin from polymerizing as soon as it is assembled into RNPs. An equally intriguing possibility is that actin undergoes covalent modifications that do not favor the establishment of polymers. Some of these ideas are speculative, but they emphasize the importance of regulating actin polymerization for nuclear function.

Nuclear Functions of Actin

LONG-RANGE MOVEMENT

Most of our current knowledge of nuclear actin originates from research in the field of chromatin regulation and transcription. However, recent studies point to the possibility that actin plays a role also in the large-scale organization of the genome. The eukaryotic nucleus is highly compartmentalized and, in spite of the lack of membrane-bound organelles, most nuclear components show specific distributions restricted to certain nuclear domains (reviewed by Lamond and Spector 2003; Schneider and Grosschedl 2007). The functional compartmentalization of the nucleus is reflected in the fact that different types of sequences occupy different positions in the nucleus. For example, gene-rich chromosome regions tend to occupy internal positions, whereas gene-poor sequences are associated with the nuclear periphery, and repressed genes are often associated with heterochromatic regions, whereas active genes are usually located near nuclear speckles (reviewed by Spector 2003). What is most interesting is that the position of a gene locus can change in response to transcriptional activation (Parada and Misteli 2002) and that the transcriptional activity of a gene is affected by its position in the nucleus (Finlan et al. 2008). Chuang and coworkers (2006) visualized the dynamics of gene repositioning by constructing cell lines that carry tandem gene arrays under the control of an inducible promoter. When the expression of the array was activated, the entire locus relocated from the cell periphery to the nuclear interior. Unidirectional trajectories were recorded over distances of 1 – 5 mm, which suggests that an active, directed mechanism for long-range chromatin movement is present. Interestingly, the expression of mutant actin or NM1 with defects in polymerization and actin-binding inhibited the relocation of the array on transcriptional activation (Chuang et al. 2006). Dundr and coworkers (2007) used a similar experimental design based on the use of synthetic inducible arrays to study the association of Cajal bodies (CBs) with U2 snRNA genes in human cells. After transcription had been induced, the U2 snRNA gene array moved

over long distances inside the nucleus (2 – 3 mm) and became stably associated with CBs. Also in this case, the expression of a nonpolymerizable actin mutant inhibited the movement of the U2snRNA locus (Dundr et al. 2007). In a third study, Hu and coworkers identified long-range, estrogen-induced interactions among genes located in different chromosomes. The interactions were lost after treatment of the cells with drugs that either blocked actin polymerization or inhibited actin depolymerization. Depletion of NM1 by RNAi or nuclear injection of antibodies against NMI also blocked the estrogen-induced interactions (Hu et al. 2008). Moreover, the inhibitory effect of the anti-NM1 antibodies could be reversed by the expression of wild-type NM1, but not by the expression of NM1 mutants that were defective in actin binding or lacked ATPase activity (Hu et al. 2008). The results reveal that the dynamics of actin in the cell nucleus affect gene positioning and large-scale chromatin organization, in a direct or indirect manner. The molecular mechanisms underlying these long-range chromatin movements are still unclear. The type of movement observed could be explained by active transport mechanisms based on actomyosin motors. This would imply the existence of mechanisms for the regulation of actin dynamics at micrometer scale inside the nucleus. If this is the case, further work must characterize such an actin large-scale regulatory network and elucidate the mechanisms that impose directionality inside the nucleus. Alternatively, and considering the fact that the observed relocations take place after transcription activation, the actin dependence could be at the transcriptional level and the chromatin movements could be a consequence of chromatin decondensation. This model, however, does not provide a satisfactory explanation of the fact that the U2 snRNA genes are targeted to relatively stable positions in the nucleus, nor does it provide any basis for the directionality of the movements. Further work is needed to understand the molecular mechanisms of large-scale chromatin relocation and to answer the questions raised by the discovery of gene relocation events. We anticipate that the

N. Visa and P. Percipalle

answers to these questions will provide a fundamental advance in our understanding of the dynamics of the eukaryotic genome.

activity of actin in gene expression is also sensitive to extracellular stimuli.

ACKNOWLEDGMENTS CONCLUDING REMARKS

There is an increasing body of evidence that supports the idea that actin plays a role in controlling multiple phases of gene transcription as a component of chromatin-remodeling complexes and RNP particles, and that it is closely associated with all RNA polymerases. Two possible models by which actin may regulate transcription elongation are illustrated in Figure 2. One model proposes the existence of an actin-NM1 motor, whereas the other one emphasizes the role of actin-RNP complexes in recruiting chromatin modifiers to transcribed genes. Both models are based on solid experimental data and are not mutually exclusive. We do not know whether both mechanisms act in all types of genes. Support for the existence of an actin-NM1 motor comes mostly from studies of Pol I transcription, whereas the recruitment of HATs mediated by RNPs has been mostly documented for protein-coding genes. It is possible that the way in which actin acts in transcription is different for different types of genes. After many years of debate about the existence of actin in the nucleus, it is now clear that actin is an abundant nuclear protein and many independent studies support the idea that it is involved in gene expression. An interesting question is whether the nuclear functions of actin are linked to—or regulated by—the dynamics of actin in the cytoplasm. The actin cytoskeleton is highly dynamic and reacts to a variety of extracellular signals (for recent reviews, see Moustakas and Heldin 2008; Papakonstanti and Stournaras 2008), and there is evidence that the cytoplasmic and actin pools are not independent of each other (Vartiainen et al. 2007). It is therefore tempting to suggest that actin acts as a sensor of extracellular signals with the ability to transduce the signals to the genome and modulate gene expression. The question remains whether the overall

¨ stlund Farrants for We thank Ann-Kristin O critical reading of the manuscript and George Farrants for language editing. Our work is supported by grants from the Swedish Research Council (Vetenskapsra˚det), the Swedish Cancer Society (Cancerfonden), and the European Science Foundation (Eurocores Programme RNAQuality).

REFERENCES Blessing CA, Ugrinova GT, Goodson HV. 2004. Actin and ARPs: Action in the nucleus. Trends Cell Biol 14: 435 –442. ¨ stlund Farrants AK. 2006. Cavellan E, Asp P, Percipalle P, O The chromatin remodelling complex WSTF-SNF2 h interacts with several nuclear proteins in transcription. J Biol Chem 281: 16264– 16271. Chen M, Shen X. 2007. Nuclear actin and actin-related proteins in chromatin dynamics. Curr Opin Cell Biol 19: 326 –330. Chuang CH, Carpenter AE, Fuchsova B, Johnson T, de Lanerolle P, Belmont AS. 2006. Long-range directional movement of an interphase chromosome site. Curr Biol 16: 825– 831. Cisterna B, Malatesta M, Dieker J, Muller S, Prosperi E, Biggiogera M. 2009. An active mechanism flanks and modulates the export of the small ribosomal subunits. Histochem Cell Biol 131: 743– 753. Cisterna B, Necchi D, Prosperi E, Biggiogera M. 2006. Small ribosomal subunits associate with nuclear myosin and actin in transit to the nuclear pores. FASEB J 20: 1901–1903. Cramer P, Bushnell DA, Kornberg RD. 2001. Structural basis of transcription: RNA polymerase II at 2.8 angstrom resolution. Science 292: 1863– 1876. Dong B, Horowitz DS, Kobayashi R, Krainer AR. 1993. Purification and cDNA cloning of HeLa cell p54nrb, a nuclear protein with two RNA recognition motifs and extensive homology to human splicing factor PSF and Drosophila NONA/BJ6. Nucl Acids Res 21: 4085–4092. Dreyfuss G, Kim VN, Kataoka N. 2002. Messenger-RNAbinding proteins and the messages they carry. Nat Rev Mol Cell Biol 3: 195–205. Dundr M, Ospina JK, Sung M-H, John S, Upender M, Reid T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo, J. Cell Biol 179: 1095–1103. Egly JM, Miyamoto NG, Moncollin V, Chambon P. 1984. Is actin a transcription initiation factor for RNA polymerase B? EMBO J 3: 2363– 2371.

Nuclear Functions of Actin Emili A, Shales M, McCracken S, Xie W, Tucker PW, Kobayashi R, Blencowe BJ, Ingles CJ. 2002. Splicing and transcription-associated proteins PSF and p54nrb/nonO bind to the RNA polymerase II CTD. RNA 8: 1102– 1111. Farrants AK. 2008. Chromatin remodelling and actin organisation. FEBS Lett 582: 2041–2050. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Fomproix N, Percipalle P. 2004. An actin-myosin complex on actively transcribing genes. Exp Cell Res 294: 140 –148. Fox AH, Lam YW, Leung AK, Lyon CE, Andersen J, Mann M, Lamond AI. 2002. Paraspeckles: A novel nuclear domain. Curr Biol 12: 13–25. Ferrai C, Naum-Onganı´a G, Longobardi E, Palazzolo M, Disanza A, Diaz VM, Crippa MP, Scita G, Blasi F. 2009. Induction of HoxB transcription by retinoic acid requires actin polymerization. Mol Biol Cell May 28. [Epub ahead of print]. Gieni RS, Hendzel MJ. 2009. Actin dynamics and functions in the interphase nucleus: Moving toward an understanding of nuclear polymeric actin. Biochem Cell Biol 87: 283 –306. Harata M, Oma Y, Mizuno S, Jiang YW, Stillman DJ, Wintersberger U. 1999. The nuclear actin-related protein of Saccharomyces cerevisiae, Act3p/Arp4, interacts with core histones. Mol Biol Cell 10: 2595–2605. Hofmann WA, Johnson T, Klapczynski M, Fan J-L, de Lanerolle P. 2006b. From transcription to transport: Emerging roles for nuclear myosin I. Biochem Cell Biol 84: 418 –426. Hofmann WA, Reichart B, Ewald A, Muller E, Schmitt I, Stauber RH, Lottspeich F, Jockusch BM, Scheer U, Hauber J, et al. 2001. Cofactor requirements for nuclear export of Rev response element (RRE)- and constitutive transport element (CTE)-containing retroviral RNAs. An unexpected role for actin. J. Cell Biol 152: 895– 910. Hofmann WA, Stojiljkovic L, Fuchsova B, Vargas GM, Mavrommatis E, Philimonenko V, Kysela K, Goodrich JA, Lessard JL, Hope TJ, et al. 2004. Actin is part of preinitiation complexes and is necessary for transcription by RNA polymerase II. Nat Cell Biol 6: 1094–1101. Hofmann WA, Vargas GM, Ramchandran R, Stojiljkovic L, Goodrich JA, de Lanerolle P. 2006a. Nuclear myosin I is necessary for the formation of the first phosphodiester bond during transcription initiation by RNA polymerase II. J Cell Biochem 99: 1001–1009. Holaska JM, Wilson KL. 2007. An emerin “proteome”: Purification of distinct emerin-containing complexes from HeLa cells suggests molecular basis for diverse roles including gene regulation, mRNA splicing, signaling, mechanosensing, and nuclear architecture. Biochemistry 46: 8897–8908. Hu P, Wu S, Hernandez N. 2004. A role for b-actin in RNA polymerase III transcription. Genes Dev 18: 3010– 3005. Hu Q, Kwon YS, Nunez E, Cardamone MD, Hutt KR, Ohgi KA, Garcia-Bassets I, Rose DW, Glass CK, Rosenfeld MG, et al. 2008. Enhancing nuclear receptor-induced transcription requires nuclear motor and LSD1-dependent gene networking in interchromatin granules. Proc Natl Acad Sci 105: 19199– 19204.

Huff T, Rosorius O, Otto AM, Muller CS, Ballweber E, Hannappel E, Mannherz HG. 2004. Nuclear localisation of the G-actin sequestering peptide thymosin b4. J Cell Sci 117: 5333– 5341. Jockusch BM, Schoenenberger CA, Stetefeld J, Aebi U. 2006. Tracking down the different forms of nuclear actin. TRENDS Cell Biol 16: 391–396. Jones KR, Rubin GM. 1990. Molecular analysis of no-on-transient A, a gene required for normal vision in Drosophila. Neuron 4: 711 –723. Kiesler E, Miralles F, Ostlund Farrants AK, Visa N. 2003. The Hrp65 self-interaction is mediated by an evolutionarily conserved domain and is required for nuclear import of Hrp65 isoforms that lack a nuclear localization signal. J Cell Sci 116: 3949– 3956. Kiseleva E, Drummond SP, Goldberg MW, Rutherford SA, Allen TD, Wilson KL. 2004. Actin- and protein-4.1containing filaments link nuclear pore complexes to subnuclear organelles in Xenopus oocyte nuclei. J Cell Sci 117: 2481– 2490. Krecic AM, Swanson MS. 1999. hnRNP complexes: Composition, structure, and function. Curr Opin Cell Biol 11: 363 –371. Kukalev A, Nord Y, Palmberg C, Bergman T, Percipalle P. 2005. Actin and hnRNP U cooperate for productive transcription by RNA polymerase II. Nat Struct Mol Biol 12: 238 –244. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605– 612. Li ZY, Yang J, Gao X, Lu JY, Zhang Y, Wang K, Cheng MB, Wu NH, Zhang Y, Wu Z, et al. 2007. Sequential recruitment of PCAF and BRG1 contributes to myogenin activation in 12-O-tetradecanoylphorbol-13-acetate-induced early differentiation of rhabdomyosarcoma-derived cells. J Biol Chem 282: 18872– 18878. Louvet E, Percipalle P. 2009. Actin and myosin in gene transcription. Int Rev Cell Mol Biol 272: 107–147. McDonald D, Carrero G, Andrin C, de Vries G, Hendzel MJ. 2006. Nucleoplasmic b-actin exists in a dynamic equilibrium between low-mobility polymeric species and rapidly diffusing populations. J Cell Biol 172: 541– 552. Meinhart A, Kamenski T, Hoeppner S, Baumli S, Cramer P. 2005. A structural perspective of CTD function. Genes Dev 19: 1401–1415. Miki T, Okawa K, Sekimoto T, Yoneda Y, Watanabe S, Ishizaki T, Narumiya S. 2008. mDia2 shuttles between the nucleus and the cytoplasm through the importin-{a}/ {b}- and CRM1-mediated nuclear transport mechanism. J Biol Chem 284: 5753–5762. Miralles F, Posern G, Zaromytidou AI, Treisman R. 2003. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113: 329 –342. Moustakas A, Heldin CH. 2008. Dynamic control of TGF-b signaling and its links to the cytoskeleton. FEBS Lett 582: 2051–2065. Obrdlik A, Kukalev A, Louvet E, Farrants AK, Caputo L, Percipalle P. 2008. The histone acetyltransferase PCAF associates with actin and hnRNP U for RNA polymerase II transcription. Mol Cell Biol 28: 6342–6357. Obrdlik A, Louvet E, Kukalev A, Naschekin D, Kiseleva E, Fahrenkrog B, Percipalle P. 2009. Nuclear myosin 1 is in

N. Visa and P. Percipalle complex with mature rRNA transcripts and associates with the nuclear pore complex. FASEB J, 2009 Sept 3 [Epub ahead of print] PMID: 19729515. Oeffinger M, Wei KE, Rogers R, DeGrasse JA, Chait BT, Aitchison JD, Rout MP. 2007. Comprehensive analysis of diverse ribonucleoprotein complexes. Nat Methods 4: 951–956. Olave IA, Reck-Peterson SL, Crabtree GR. 2002. Nuclear actin and actin-related proteins in chromatin remodelling. Annu Rev Biochem 71: 755–781. Papakonstanti EA, Stournaras C. 2008. Cell responses regulated by early reorganization of actin cytoskeleton. FEBS Lett 582: 2120– 2127. Parada L, Misteli T. 2002. Chromosome positioning in the interphase nucleus. Trends Cell Biol 12: 425– 432. Patton JG, Porro EB, Galceran J, Tempst P, Nadal-Ginard B. 1993. Cloning and characterization of PSF, a novel pre-mRNA splicing factor. Genes Dev 7: 393– 406. Pederson T. 2008. As functional nuclear actin comes into view, is it globular, filamentous, or both? J Cell Biol 180: 1061–1064. Pederson T, Aebi U. 2002. Actin in the nucleus: what form and what for? J Struct Biol 140: 3– 9. Pendleton A, Pope B, Weeds A, Koffer A. 2003. Latrunculin B or ATP depletion induces cofilin-dependent translocation of actin into nuclei of mast cells. J Biol Chem 278: 14394–14400. Percipalle P. 2009. The long journey of actin and actin-associated proteins from gene to polysomes. Cell Mol Life Sci 66: 2151–2165. Percipalle P, Obrdlik A. 2009. Analysis of nascent RNA transcripts by chromatin RNA immunoprecipitation. Meth Mol Biol 567: 215– 235. Percipalle P, Visa N. 2006. Molecular functions of nuclear actin in transcription. J Cell Biol 172: 967– 971. ¨ stlund Farrants AK. 2006. Chromatin remodPercipalle P, O elling and transcription: Be-WICHed by nuclear myosin 1. Curr Opin Cell Biol 18: 267 –274. Percipalle P, Fomproix N, Cavella´n E, Voit R, Reimer G, Kru¨ger T, Thyberg J, Scheer U, Grummt I, Farrants AK. 2006. The chromatin remodelling complex WSTFSNFh interacts with nuclear myosin 1 and has a role in RNA polymerase I transcription. EMBO Rep 7: 525 –530. Percipalle P, Fomproix N, Kylberg K, Miralles F, Bjorkroth B, Daneholt B, Visa N. 2003. An actin-ribonucleoprotein interaction is involved in transcription by RNA polymerase II. Proc Natl Acad Sci 100: 6475–6480. Percipalle P, Jonsson A, Nashchekin D, Karlsson C, Bergman T, Guialis A, Daneholt B. 2002. Nuclear actin is associated with a specific subset of hnRNP A/B-type proteins. Nucleic Acids Res 30: 1725– 1734. Percipalle P, Zhao J, Pope B, Weeds A, Lindberg U, Daneholt B. 2001. Actin bound to the heterogeneous nuclear ribonucleoprotein hrp36 is associated with Balbiani ring mRNA from the gene to polysomes. J Cell Biol 153: 229–236. Pestic-Dragovich L, Stojiljkovic L, Philimonenko AA, Nowak G, Ke Y, Settlage RE, Shabanowitz J, Hunt DF, Hozak P, de Lanerolle P. 2000. A myosin I isoform in the nucleus. Science 290: 337– 341.

Philimonenko VV, Zhao J, Iben S, Dingova H, Kysela K, Kahle M, Zentgraf H, Hofmann WA, de Lanerolle P, Hozak P, et al. 2004. Nuclear actin and myosin I are required for RNA polymerase I transcription. Nat Cell Biol 6: 1165– 1171. Politz JC, Tuft RA, Pederson T. 2003. Diffusion-based transport of nascent ribosomes in the nucleus. Mol Biol Cell 14: 4805–4812. Prendergast GC, Ziff EB. 1991. Mbh 1: a novel gelsolin/ severin-related protein which binds actin in vitro and exhibits nuclear localization in vivo. EMBO J 10: 757– 766. Raju CS, Go¨ritz C, Nord Y, Hermanson O, Lopez-Iglesias C, Visa N, Castelo-Branco G, Percipalle P. 2008. In cultured oligodendrocytes the A/B-type hnRNP CBF-A accompanies MBP mRNA bound to mRNA trafficking sequences. Mol Biol Cell 19: 3008– 3018. Rando OJ, Zhao K, Janmey P, Crabtree GR. 2002. Phospatidylinositol-dependent actin filament binding by the SWI/SNF-like BAF chromatin remodelling complex. Proc Natl Acad Sci 99: 2824– 2829. Scheer U, Hinssen H, Franke WW, Jockusch BM. 1984. Microinjection of actin-binding proteins and actin antibodies demonstrates involvement of nuclear actin in transcription of lampbrush chromosomes. Cell 39: 111 –122. Schneider R, Grosschedl R. 2007. Dynamics and interplay of nuclear architecture, genome organization, and gene expression. Genes Dev 21: 3027–3043. Siebrasse JP, Veith R, Dobay A, Leonhardt H, Daneholt B, Kubitscheck U. 2008. Discontinuous movement of mRNP particles in nucleoplasmic regions devoid of chromatin. Proc Natl Acad Sci 105: 20291–20296. Singh OP, Bjo¨rkroth B, Masich S, Wieslander L, Daneholt B. 1999. The intranuclear movement of Balbiani ring premessenger ribonucleoprotein particles. Exp Cell Res 251: 135–146. ¨ stlund Farrants Sjo¨linder M, Bjo¨rk P, Soderberg E, Sabri N, O AK, Visa N. 2005. The growing pre-mRNA recruits actin and chromatin-modifying factors to transcriptionally active genes. Genes Dev 19: 1871– 1884. Skare P, Kreivi JP, Bergstro¨m A, Karlsson R. 2003. Profilin I colocalizes with speckles and Cajal bodies: A possible role in pre-mRNA splicing. Exp Cell Res 286: 12–21. Spector DL. 2003. The dynamics of chromosome organization and gene regulation. Annu Rev Biochem 72: 573–608. Vartiainen MK. 2008. Nuclear actin dynamics– from form to function. FEBS Lett 582: 2033–2040. Vartiainen MK, Guettler S, Larijani B, Treisman R. 2007. Nuclear actin regulates dynamic subcellular localization and activity of the SRF cofactor MAL. Science 316: 1749–1752. Visa N, Alzhanova-Ericsson AT, Sun X, Kiseleva E, Bjo¨rkroth B, Wurtz T, Daneholt B. 1996. A pre-mRNA-binding protein accompanies the RNA from the gene through the nuclear pores and into polysomes. Cell 84: 253– 264. von Besser H, Schnabel P, Wieland C, Fritz E, Stanewsky R, Saumweber H. 1990. The puff-specific Drosophila protein Bj6, encoded by the gene no-on transient A, shows homology to RNA-binding proteins. Chromosoma 100: 37– 47.

Nuclear Functions of Actin Wu X, Yoo Y, Okuhama NN, Tucker PW, Liu G, Guan JL. 2006. Regulation of RNA-polymerase-II-dependent transcription by N-WASP and its nuclear-binding partners. Nat Cell Biol 8: 756– 763. Ye J, Zhao J, Hoffmann-Rohrer U, Grummt I. 2008. Nuclear myosin I acts in concert with polymeric actin to drive RNA polymerase I transcription. Genes Dev 22: 322–330. Yoo Y, Wu X, Guan JL. 2007. A novel role of the actinnucleating Arp2/3 complex in the regulation of RNA polymerase II-dependent transcription. J Biol Chem 282: 7616– 7623.

Zhao K, Wang W, Rando OJ, Xue Y, Swiderek K, Kuo A, Crabtree GR. 1998. Rapid and phosphoinositoldependent binding of the SWI/SNF-like BAF complex to chromatin after T lymphocyte receptor signaling. Cell 95: 625–636. Zechel K. 1980. Isolation of polymerization competent cytoplasmic actin by affinity chromatography on immobilized DNase I using formamide as eluant. Eur J Biochem 110: 343–348. Zuchero JB, Coutts AS, Quinlan ME, Thangue NB, Mullins RD. 2009. p53-cofactor JMY is a multifunctional actin nucleation factor. Nat Cell Biol 11: 451–459.

The Nucleolus Thoru Pederson Program in Cell and Developmental Dynamics, Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts 01605 Correspondence: [email protected]

When cells are observed by phase contrast microscopy, nucleoli are among the most conspicuous structures. The nucleolus was formally described between 1835 and 1839, but it was another century before it was discovered to be associated with a specific chromosomal locus, thus defining it as a cytogenetic entity. Nucleoli were first isolated in the 1950s, from starfish oocytes. Then, in the early 1960s, a boomlet of studies led to one of the epochal discoveries in the modern era of genetics and cell biology: that the nucleolus is the site of ribosomal RNA synthesis and nascent ribosome assembly. This epistemologically repositioned the nucleolus as not merely an aspect of nuclear anatomy but rather as a cytological manifestation of gene action—a major heuristic advance. Indeed, the finding that the nucleolus is the seat of ribosome production constitutes one of the most vivid confluences of form and function in the history of cell biology. This account presents the nucleolus in both historical and contemporary perspectives. The modern era has brought the unanticipated discovery that the nucleolus is plurifunctional, constituting a paradigm shift.

FIRST SIGHTING

t is likely that some of the few lucky enough to have a microscope in the 18th century saw the nucleolus if they examined thin specimens of tissue in the mode of illumination that later became known as bright field, a century before phase contrast was discovered, for which a Nobel Prize, rare in microscopy, was conferred on Frits Zernike in 1953 (Fig. 1). The first properly documented accounts of the nucleolus were made independently by Wagner (1835) and Valentin (1836, 1839). Beyond occasional studies in which nucleoli were mentioned in passing in the context of broader cytological work, no significant literature on the nucleolus ensued for another half century. Then,

I

a monumental monograph on the nucleolus was published by Montgomery (1898), with an astonishing 346 hand-drawn color figures of nuclei and nucleoli from a vast array of biological material. But as comprehensive and elegant a piece of scholarship as it was, this treatise is primarily known to us today not because of what it revealed the nucleolus to be at the time, but because of what the nucleolus later became. THE NUCLEOLUS BECOMES PART OF THE KARYOTYPE

For three decades after Montgomery’s treatise, there was little momentum until the discovery that the nucleolus arises at a specific chromosomal locus (Heitz 1931; McClintock 1934). While

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved. Advanced Online Article. Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a000638

1

T. Pederson

Figure 1. Looking at the nucleolus. Nucleoli observed in HeLa cells (left) and isolated HeLa cell nuclei (right) by differential interference microscopy. (Images courtesy of David L. Spector [left] and Thoru Pederson [right].)

the intimacy of associations between nucleoli and heterochromatin had previously been noted, the independent findings of Heitz and McClintock established that the nucleolus actually forms at a discrete locus, termed the nucleolus organizer by McClintock. The importance of this advance cannot be overstated because it meant that rather than being mere nuclear anatomy, nucleoli are cytogenetic entities. From this perspective, the astonishing diversity of nucleolar shapes and sizes recorded by Montgomery began to be sensed as reflecting differences in the chromosomal sites at which they arise. The discovery of the nucleolus organizer meant that, whatever its function(s), the nucleolus is stationed at a genetic locus, although it would still be another decade before the discovery that the genetic material is DNA. MONTEVIDEO: A HIGH ALTITUDE VIEW OF THE NUCLEOLUS

We have all attended meetings, and the luckiest of us know that sometimes, when the muses smile, we are witnesses to a renaissance. It just happened that leading up to a meeting on the nucleolus in Montevideo, Uruguay, in December 1965, several new findings had been made or were just about to be submitted for publication. Donald Brown presented a brilliant

experiment, done with John Gurdon (Brown and Gurdon 1964), showing that anucleoate Xenopus embryos arrest in development, plausibly due to the inability of these embryos to make new ribosomes when the maternal stockpile becomes limiting (Fig. 2). Brown also reported the existence of amplified nucleoli in the germinal vesicle (nucleus) of Xenopus oocytes (a finding that had been made contemporaneously [Brown and Dawid 1968; Gall 1968]). Papers on the isolation of nucleoli were given by Walter Vincent (from starfish oocytes), and by Rachele Maggio and Harris Busch (from guinea pig and rat liver, respectively). The biosynthesis of rRNA via large precursor molecules was reported in talks by Joseph Gall, Sheldon Penman, Georgii Georgiev, and Robert Perry. But the most important discovery announced at the meeting was that reported in the talks by Max Birnstiel and Ferrucio Ritossa; namely, that nucleic acid hybridization revealed that DNA complementary to rRNA resides in the nucleolus, which, together with the results of Brown and Gurdon (1964), ushered in a new age in the history of the nucleolus. The proceedings of this conference, including the lively discussion exchanges after each talk (Vincent and Miller 1966), together with an exceptionally insightful synopsis of the meeting (Perry 1966), constitute a definitive archive of this exciting moment in the field.

The Nucleolus

Figure 2. A portal to the modern era: the dependence of ribosomal RNA synthesis on the nucleolus. Anucleolate

(left) or wild-type (right) Xenopus laevis embryos at the neurula stage were incubated with C14-labeled carbon dioxide and RNA was extracted 20 hours later, when both groups of embryos were still morphologically and physiologically indistinguishable. Shown are sucrose gradient sedimentation profiles of 28S and 18S rRNA and transfer RNA. This finding was a keystone in establishing the role of the nucleolus in the biosynthesis of ribosomes. (Reproduced from Brown DC and Gurdon JB 1964. Proc Natl Acad Sci USA 51: 139– 146, by kind permission of Donald D. Brown, Carnegie Institution for Science, Washington, D.C.)

COMING DOWN FROM THE MOUNTAIN

Oscar Miller had presented EM pictures of spread nucleolar “cores” and stretched nucleolar DNA at the meeting, but they were his first attempts and not particularly revealing. But later, he and his talented research assistant Barbara Beatty showed the world what these genes really look like in full transcriptional action (Miller and Beatty 1969). These pictures earned their rightful place as among the most iconic of any in the history of cytology and cell biology. Incisive studies on the synthesis and processing of ribosomal RNA introduced by Sheldon Penman at the Montevideo meeting were subsequently refined by him and independently by the laboratory of James Darnell (reviewed by Lewin 1980). In contrast, the isolation of nucleoli presented at the meeting was to await several decades for further advances. In his summary of the Montevideo conference, the Edinburgh embryologist C.H.

Waddington said: “The nucleolus probably should not be considered a relatively simple organelle with a single function, comparable to a machine tool turning out a particular part of an automobile. It is not just ‘the organelle where the cell manufactures ribosomes.’ It is rather a structure through which materials of several different kinds are flowing, comparable more to a whole production line than to a single machine tool.” One cannot imagine a more prescient view. As we shall see, every atom of his statement has been borne out in subsequent research on the nucleolus.

INTO THE MODERN ERA

In the 1970s and 1980s, the nucleolus field addressed the details of ribosome biosynthesis. One axis was a troubled one: the goal of reconciling the stages of ribosome synthesis with the classically defined subcompartments of the

T. Pederson

nucleolus. Just as one commentator famously described (to every Latin student centuries later) the Roman conquest territory that would later become France, “Gallia est omnis divisa in partes tres” (Caesar, 40– 50s BC), the nucleolus is also tripartite. Its three classical regions are defined by the different appearance of intranucleolar regions when viewed by electron microscopy (Fig. 3). These are the fibrillar centers, the surrounding dense fibrillar component, and the granular component. Few in the field anticipated the controversies that would ensue when several labs tried, logically enough, to link the sites of rRNA transcription and processing and assembly into nascent ribosomes with these three EM-defined nucleolar zones. These debates were waged with intense controversy and, in a few cases, frank boorishness on the part of some in this raging controversy on the floor at international meetings. In hindsight, this had to do with how short a pulse label one could introduce to tag nascent rRNA. Thankfully, recent advances have settled the issue. A key step in this resolution was a reconciliation of the “Christmas tree” EM images of spread nascent ribosomal RNA (Miller and Beatty 1969) with the high resolution detection of these genes and

Figure 3. The tripartite organization of the nucleolus.

Electron micrograph of a mouse fibroblast nucleolus. (f ) fibrillar center, (d) dense fibrillar component, (g) granular component, (arrows) perinucleolar heterochromatin, ( ) denotes the presence of dense fibrillar component material within the fibrillar center, which is occasionally observed. (Reprinted from Trends in Cell Biol 13: 517 –525, Rasˇka, I. # 2003, with permission from Elsevier.)

their transcripts in nucleoli in situ (Koberna et al. 2002; for further discussion of the earlier controversy and its resolution, see Rasˇka et al. 2006). Recent work has indicated that one of the three nucleolar regions, the granular component, is itself composed of at least two distinct molecular domains. Meanwhile, the covalent steps in rRNA processing continued to be explored, revealing different nuances in yeast versus higher eukaryotes superimposed on a conserved foundation (reviewed by Fatica and Tollervey 2002). But another important axis was also emerging in the 1970s. The laboratory of Harris Busch, notably Ramachandra Reddy, came upon and sequenced several small nucleolar RNAs. One of these, U3 small nucleolar RNA, was subsequently shown to be base-paired with nucleolar rRNA in vivo (Calvet and Pederson 1981) and later was demonstrated to function in the initial cleavage of the 45S rRNA precursor (Kass et al. 1990). All of these studies were done, of course, with deproteinized nucleolar RNA. Soon thereafter, isolated nucleoli were subjected to various extraction conditions designed to preserve nascent rRNA– protein complexes, resulting in the characterization of the ribonucleoprotein forms of the rRNA precursor and processing intermediates that had previously been defined with deproteinized RNA (Warner and Soeiro 1967; Pederson and Kumar 1971; Kumar and Warner 1972). Jumping ahead, we now enjoy a far more detailed picture of the covalent steps of rRNA maturation (Fatica and Tollervey 2002) as well as numerous accessory factors that support this pathway but are not part of the final ribosomes (reviewed by Granneman and Baserga 2004). Among the latter are more than 100 small nucleolar RNAs that guide the extensive site-specific nucleotide modifications (ribose 20 -O-methylation and pseudouridine formation) of rRNA during its processing (reviewed by Bachellerie et al. 2002). Like the initial rRNA precursor, processing intermediates, and products themselves, all these small nucleloar RNAs are complexed with proteins, thus instilling in the nucleolar landscape a high concentration of both RNA and protein, a point to which we shall return.

The Nucleolus

A PARADIGM SHIFT

By the 1960s and into the 1970s, the notion that the nucleolus is the seat of ribosome synthesis was as cogently established as anything in the modern canon of the molecular and cell biology of eukaryotes. Then, something unexpected happened. In my laboratory, we had been studying the traffic of fluorescently labeled small nucleolar RNAs after microinjection into the nuclei of cultured mammalian cells and finding them to move into nucleoli, as expected, but extending this to define nucleolar targeting elements in these RNAs (Jacobson et al. 1995; 1997; Jacobson and Pederson 1998a). In the case of U3 and U8 (Jacobson and Pederson 1998a), we planned to expand the findings but first decided to submit an initial report to convey our novel experimental approach (reviewed in Pederson 2001a), something quite new at the time for the nucleolus, though we had previously applied it to study pre-mRNA traffic to nucleoplasmic domains (Wang et al. 1991). While writing this paper (Jacobson and Pederson 1998a), it occurred to us that we had only used a single RNA as a negative control. Although it did not display nucleolar localization after microinjection, we decided to try a second small RNA that presumably would also not localize in nucleoli, thus amplifying our case that the observed nucleolar localizations of U3 and U8 were specific. My post-doc Marty Jacobson suggested that we try the signal recognition particle RNA, a 300-nt pol III transcript, not only to fulfill the need for another negative control but also because, to his credit, he realized nothing was known about the nuclear phase of SRP biosynthesis. To our surprise, SRP RNA displayed as rapid and as quantitative a nucleolar localization as U3 and U8 snoRNAs. But looking into this further, we saw that SRP RNA trafficked into nucleoli only transiently. U3 and U8 on the other hand remained stably localized in nucleoli, as did other small nucleolar RNAs we had begun to study. SRP RNA then moved out into the cytoplasm, where it became associated with the endoplasmic reticulum, its expected final localization (Jacobson and Pederson 1998b).

These findings on SRP RNA, together with a number of other contemporaneous results that were stirring, led to the formulation of the “plurifunctional nucleolus” hypothesis (Pederson 1998a). Subsequently, the link between the nucleolus and signal recognition particle biosynthesis was made in yeast (Ciufo and Brown 2000; Grosshans et al. 2001), and confirmed and extended in mammalian cells (Politz et al. 2000) (Fig. 4) and in Xenopus oocytes (Sommerville et al. 2005). A speculative essay suggested how a common nucleolar assembly site might have co-evolved for the four translational ribonucleoproteins (the two ribosomal subunits, the 5S rRNA– protein complex, and the SRP) (Pederson and Politz 2000). Notwithstanding how it came about in the evolution of eukaryotes, the fact that the nucleolus is a site of more than ribosome synthesis had clearly arrived once its role in the biosynthesis of the SRP had been discovered. Also embedded in the plurifunctional nucleolus hypothesis were provocative results linking the nucleolus to cellcycle progression (Pederson 1998a,b). Both the SRP and cell-cycle aspects of the hypothesis were welcomed into reviews on the nucleolus (Scheer and Hock 1999; Olson et al. 2000; Olson et al. 2002; Visintin and Amon 2000), and the concept has continued to be well received (Alberts et al. 2002; Rasˇka et al. 2006; Boisvert et al. 2007). THE PARTS LIST, AND SEGUEING INTO THE CELL CYCLE

When nucleoli were first isolated (Vincent 1952), only assays for total protein and RNA, as well as tests for various enzymatic activities, were available (the latter scoring only one—acid phosphatase). Jumping ahead four and a half decades, we now speak of specific nucleolar molecules. As mentioned earlier, progress on chronicling all of the small RNAs present in nucleoli had proceeded through the 1980s and 1990s and on into the 2000 decade. As for nucleolar proteins (beyond the ribosomal structural proteins), early accounts (e.g., Soeiro and Basile 1973) were followed by numerous reports of individual, non-ribosomal proteins observed

T. Pederson

Figure 4. The nucleolus and signal recognition particle biosynthesis. Nucleolar localization of GFP-tagged

SRP19 (left) and SRP68 (right) proteins in rat NRK cells. Cytoplasmic localization is also observed, as expected from the known steady-state localization of the signal recognition particle. Similar results were obtained with GFP-tagged SRP72 protein, whereas the SRP54 protein assembles with the nascent signal recognition particle in the cytoplasm (Politz et al. 2000; Sommerville et al. 2005). Reproduced from Politz et al. 2000, with permission from the Proceedings of the National Academy of Sciences.

either in isolated nucleoli or, more typically, by immunofluorescence or GFP-tagged protein localization. These were useful findings on each protein’s individual merit but obviously did not reveal the molecular breadth of the nucleolar protein landscape. In 2002, two groups refined the classical nucleolar isolation methods (notably that developed by Maggio et al. 1963) and prepared high purity nucleoli from HeLa cells followed by proteomics (Andersen et al. 2002; Scherl et al. 2002; reviewed by Pederson 2002 and Leung et al. 2003). These two contemporaneous proteomics studies revealed, in combination, a total of approximately 350 proteins. Excluding ribosomal structural proteins, there were many that had been or would be later linked to ribosome biosynthesis as chaperone-like mediators not ending up in mature ribosomes. This finding expanded and put more molecular definition on earlier research, showing that nucleolar preribosomal particles have a higher protein:RNA mass ratio than mature ribosomes (Liau and Perry 1969; Pederson and Kumar 1971; Kumar and Warner 1972). Many others of the observed nucleolar proteins were ones involved in DNA replication or repair, cell-cycle progression, or its control, validating this aspect of the initial

plurifunctional nucleolus hypothesis (Pederson 1998a) and findings being made at the time that had suggested such a link (reviewed by Pederson 1998b; Garcia and Pillus 1999; Vinistin and Amon 2000). Similar findings were later made in a proteomics analysis of purified nucleoli from Arabidopsis (Pendle et al. 2005). Beyond proteomic analysis, and its emphasis on nonribosomal proteins, there has been recent work on ribosomal proteins themselves that has brought surprises. The first of these implicated ubiquitinylation, including polyubiquitinylation, of ribosomal proteins in the pathway of ribosome synthesis (Stavreva et al. 2006), a most unanticipated, counterintuitive result and one not immediately reconcilable with the “standard model” of ribosome biosynthesis. The second study revealed a surprisingly large pool of unassembled ribosomal proteins, exceeding the level needed to support ribosome assembly on the number of rRNA transcripts being produced (Lam et al. 2007). Obviously, one senses the likely possibility of a link between these two sets of observations. These findings also lead us to bear in mind the possibility that there may be further surprises ahead regarding the established lore of the nucleolar ribosome biosynthetic pathway, just as

The Nucleolus

has been the case with respect to the nonribosomal aspects of nucleolar organization and function in the 12 years since the plurifunctional nucleolus hypothesis was advanced (Pederson 1998a). THE NUCLEOLUS IS NEITHER AS STATIC NOR AS COMPACT AS FIRST THOUGHT

Between 2000 and 2001, a number of studies based on fluorescence recovery after photobleaching (FRAP), in one case employing as well the complementary method of fluorescence loss in photobleaching (FLIP), brought about a complete revision in thinking about the stasis of several nuclear proteins (for reviews, see Pederson 2000; 2001b; Leung and Lamond 2003). In the case of the nucleolar proteins studied, namely fibrillarin (Phair and Misteli 2000) and selected ribosomal structural proteins (Chen and Huang 2001), the dynamics were faster than would have been anticipated by many (perhaps most) nucleolus investigators. Fibrillarin functions as an rRNA modification enzyme in the dense fibrillar component and therefore might have been assumed to either be stably associated with the nuclelous or at least display a long residence time. A nucleolar stasis of ribosomal structural proteins was an even more entrenched notion, as they would have been assumed to enter nucleoli in stoichiometric amounts to assemble with nascent rRNA and yet they were found to be rapidly shuttling in and out of nucleoli. As mentioned above, later work revealed that ribosomal structural proteins undergo ubiquitinylation and degradation on the assembly pathway (Stavreva et al. 2006) and also enter the nucleus in suprastoichiometric levels (Lam et al. 2007). Clearly, the FRAP results (Chen and Huang 2001) reflected, or at least were significantly influenced by, these later-recognized aspects of ribosomal protein instability, whereas the fast exchange of fibrillarin (Phair and Misteli 2000) was, and remains, even more surprising. It has been more difficult to study the dynamics of nucleolar RNAs. The fact that fluorescent snoRNAs accumulate in nucleoli after microinjection into the nucleus does not address this

question since the initial localization may either be filling unoccupied sites or replacing those formerly occupied by egressing snoRNAs. Stated differently, snoRNAs are always at high concentration in the nucleolus so the concentration of the putative molecules that have moved into the nucleoplasm in any experiment that examines the steady-state would be very low. Only methods like FRAP or FLIP can capture this and these are not applicable to RNA. However, endogenous RNAs can be tagged with fluorescent probes (Politz et al. 1998), including photoactivatable ones (Politz 1999; Politz et al. 2005). In a study using this approach to track 28S rRNA out of the nucleolus (Politz et al. 2003), we noted that some signal occasionally returned to nucleoli. We pondered that there is probably no barrier to a completed ribosomal subunit occasionally gaining access back into the nucleolus (which is surprisingly porous, vide infra) before it engages the nuclear export machinery. Notwithstanding this observation, how dynamic or static nucleolar RNAs are remains an open and important question. How compact a bundle of mass is the nucleolus? Its appearance in differential interference microscopy (Fig. 1) indicates that it has higher mass per unit volume than the surrounding nucleolplasm, but how much higher? Fortunately, differential interference microscopy can be used in a quantitative mode. Its application to the nucleus of Xenopus oocytes revealed that the nucleoli are only about twice as dense as the surrounding nucleoplasm (Handwerger et al. 2005; discussed in Pederson 2010a.) In retrospect, it may be that a preferentially high affinity of some nucleolar proteins for the heavy metal atoms (uranium, osmium, tungsten) used in electron microscopy has led to a false perception of a higher ( protein) density in the nucleolus, or in some of its regions, than is actually the case. The use of basic dyes in classical cytological work on the nucleolus that are taken preferentially by polyanions such as RNA, or silver stains that bind avidly to certain nucleolar proteins, may have contributed to this notion from cytochemistry. In any case, similar studies to those done in the nucleus of Xenopus oocyte nuclei are now needed in mammalian cells, where

T. Pederson

there are more data on nucleolar protein dynamics that can be used as parameters in computational models. A first step has been taken in this direction, in which the percolation of ovalbumin (molecular weight 43,000) in the nucleolus and nucleoplasm was measured in a mammalian cell (Spell and Kubitscheck 2009), revealing that this protein had the same mobility in both compartments. One is also reminded here that the use of photo-activatable GFP-tagged nucleolar proteins in such studies could be very revealing, as has been their use in studying another protein’s dynamics in the Xenopus oocyte nucleus (Deryusheva and Gall 2004). A GROWING FAMILY OF CELL-CYCLE REGULATORS IN THE NUCLEOLUS

The nucleolus proteomics studies revealed numerous cell-cycle-related proteins among the residents (or visitors). A great many studies over the past decade have brought further insights into the nucleolar transit of cell-cycle regulatory proteins. These good endeavors have now engendered a large literature, much of which is more pertinent to cell-cycle progression control in general, whereas only a subset of this emerging work relates to the nucleolus; hence the brief, selective coverage that follows. Before embarking on the relationship between nucleolar activity and cell-cycle progression, a caveat needs to be registered. There is no doubt that when ribosome production is impaired or limiting, cell growth slows or ceases. The obligatory relationship between ribosome biosynthesis and cell growth rate was established ´´ le Maaloe and Karl for bacteria in studies by O Lark in the 1960s, and later was confirmed for eukaryotes. We can plausibly surmise that new ribosomes do count for a cell’s (or organism’s) future because selection would disfavor cells, embryos, or adult organisms that are slow to build new proteins for either cell division (a necessary inheritance for G1 progeny) or for sustained vigor of tissues and organs. That said, we here take up the question of whether the nucleolus might play roles in cell-cycle progression beyond ribosome biosynthesis (see also

Pederson 2007). This is not an idea that was pondered in the classical era, when ribosome production, protein synthesis rates, and cell growth were seen as the operation of a continuous, forward- and back-regulated production logic circuit. The point of departure was the finding that the action of two cell-cycle regulators, p53 and Mdm2, are modulated by nucleolar sequestration (Weber et al. 1999; Kulikov et al. 2010). Another key finding was that p53 is stabilized upon DNA damage-triggered nucleolar disruption or elicited by other cellular stressors (Rubbi and Milner 2003). Subsequent work has defined an extensive interactome of nucleolus-nucleoplasmic shuttling cell-cycle progression proteins and it now appears that the action of several is based on their dynamic interplay between the nucleolus and the nucleoplasm. An instructive example is nucleostemin (reviewed by Ma and Pederson 2008a; Pederson and Tsai 2009). This protein was discovered in a study of neural stem cells in the subventricular zone of the adult rat brain (Tsai and McKay 2002). Nucleostemin is a p53-interactive protein (Tsai and McKay 2002; Ma and Pederson 2007) and the initial idea (Tsai and McKay 2005; reviewed by Misteli 2005) was that when it shuttles from the nucleolus into the nucleoplasm, it binds to and disables p53, and thus drives the cell cycle. But the continuing challenge in this field is to reconcile a protein’s intranuclear location with function. For example, although p53 is mostly nucleoplasmic, the small nucleolar fraction of p53 has different binding partners, measured by bimolecular fluorescence complemenatation (see Ma and Pederson 2008b), than those with which it consorts in the nucleoplasm (Pederson, unpubl.). It is usually assumed that the highest intranuclear concentration of a protein is where it executes its function. This assumption may be suspect in some cases, and in the case of nucleostemin, the jury is out as to whether it enacts its cell-cycledriving function in the nucleolus (at high concentration) or in the nucleoplasm (at low concentration). A protein’s concentration in a region of the cell may not be a quantitative signal for its action, which should be kept in mind

The Nucleolus

as systems biology approaches now arrive in cell (and nuclear) biology. THE PERINUCLEOLAR COMPARTMENT: ANOTHER LINK BETWEEN THE NUCLEOLUS AND GROWTH CONTROL?

In some cells, there is a discoid structure situated on the nucleolus, covering a portion of its surface as a “cap.” Termed the perinucleolar compartment (PNC), it was discovered on the basis of its high concentration of a particular hnRNP protein (Ghetti et al. 1992; Matera et al. 1995; for reviews, see Huang 2000 and Pollock and Huang 2010). The PNC accretes, but is not the transcription site of, a number of small RNAs (Matera et al. 1995; Wang et al. 2003). The fact that PNCs are not seen in all cells but are noted particularly in mammalian transformed cell lines, tumor-derived cell lines, and tumor biopsy specimens suggests a key link between their appearance and tumor initiation and/or progression (see Pollock and Huang 2010). Although this is an encouraging new tool for the pathologist, the PNC’s harboring of certain proteins and RNAs presently offers no clues whatsoever as to why it appears more frequently in tumor cells. Clearly, further work on the PNC is warranted. In fact, it is the only presently known link between the nucleolus (in the form of a nucleolus-vicinal cytological entity) and a very high morbidity/mortality human disease, breast carcinoma. THE NUCLEOLUS AND VIRAL REPLICATION, STEM CELL BIOLOGY, AND CELLULAR SENESCENCE: ONLY ZEPHYRS SO FAR

Beyond cell-cycle progression and growth control, the past decade has seen a surge of attention to the nucleolus in three other fields: viral replication, stem cell biology, and cellular senescence. These studies include a number of provocative findings, briefly discussed below, but the objective summary statement is that a mechanistic link to the nucleolus has not been made. Regarding viral replication, some viruses display a nucleolar tropism, but the functional

significance of this is generally not well understood. A surprising recent finding was that the nucleolus and Cajal bodies are required for the establishment of a systemic infection by a plant virus (Kim et al. 2007). In an intriguing retroviral therapeutic development, the fact that unspliced HIV RNA exits the nucleus via the nucleolus was exploited to design a nucleolustargeting inhibitor of viral replication (Michienzi et al. 2002). The nucleolus has also been implicated in the replication of herpes viruses (Boyne and Whitehouse 2006.) Yet another recent case is the finding that the replication of the Dependovirus AAV2 (adenovirus-associated virus 2) takes place in the nucleolus (Sonntag et al. 2010). AAV2 is among the smallest animal cell viruses known (25 nm in diameter) and its assembly might thus be feasible within the relatively porous nucleolus, yet the biological significance of this nucleolar tropism is presently unclear (reviewed by Pederson 2010a). The connection of the nucleolus to stem cell biology has received some attention, with a possible link hinted at both by classical and contemporary studies (e.g., Gonda et al. 2003; reviewed by Misteli 2003; Fle´chon 2006). Thus, when a somatic nucleus from an adult vertebrate animal is placed into an enucleated egg, development can proceed up to a tadpole in the case of amphibians or even birth in the case of mammals. Much attention has been placed on the effects of the egg cytoplasm on the implanted nucleus in terms of epigenetic marks and the redirected program of gene expression. But, something else also happens. In the nucleolus of the adult, somatic nucleus transiently disappears and then comes back into view a few hours later. This intriguing behavior of the nucleolus in somatic nucleus transfer experiments has not been explored to the extent it should be. If properly investigated, this is likely to reveal profound clues as to what happens when newly combined nuclei and cytoplasm become molecularly familiar with themselves. Another potential link between the nucleolus and stem cell biology has arisen from studies of the transcription factor Hand1, whose localization within and outside the nucleolus acts as a trigger for cell fate determination (Martindill

T. Pederson

et al. 2007). Another engaging recent study has implicated a U3 small nucleolar RNA-bound protein, Wcd, in a neural stem cell niche of Drosophila embryos (Fichelson et al. 2009). Wcd displays asymmetric distribution to daughter cells and thus has “stem cellness” written all over it. One hopes that these groups and/or other nucleolus investigators are pursuing these two findings, for they are among the most intriguing links between the nucleolus and stem cell biology to have been reported. Studies linking the nucleolus to cellular senescence round out this body of “conceptually nucleolus-related” research in that, once again, some provocative clues have been uncovered, but there is no sense at present of a unifying concept or mechanism. For example, an influential study in yeast reported an acceleration of aging (whatever this means in fungi) in mutants displaying nucleolar fragmentation (Sinclair et al. 1997; reviewed in Guarente 1997). And, a number of studies have made a connection between the nucleolus and Werner syndrome, an inherited human-aging-like condition (Marciniak et al. 1998; Szekely et al. 2000; von Kobbe and Bohr 2002; Kyng et al. 2003), and this seems the best case so far for a role of the nucleolus in cellular senescence, though still far from being linked to aging of a higher eukaryotic organism. A QUADRIPARTITE NUCLEOLUS, OR MORE?

After discovering that the SRP assembly occurs in the nucleolus (Jacobson and Pederson 1998b; Politz et al. 2000; Sommerville et al. 2005), the question arose as to the intranucleolar site. Probing for both SRP RNA and 28S rRNA in dual color in situ hybridization experiments revealed incomplete overlap, although both signals resided in the granular component (Politz et al. 2002). Given this evidence for molecular segregation even within a given nucleolar subcompartment, we reasoned that probing for a molecule not involved in either ribosome or SRP biosynthesis would be the logical next step and chose nucleostemin. The results conformed and amplified the notion that the nucleolar granular component is molecularly territorialized (Politz et al. 2005). But, the most important

finding in this study came from the application of electron spectroscopic imaging, which revealed the granular component to be an interspersed array of RNA-rich and RNA-deficient particles, rather than a uniform lawn of nascent ribosomes, as would have been anticipated (Fig. 5). It remains to be seen how many subdivisions exist within the granular component, or within the dense fibrillar component for that matter. The probes and optical resolution are now at hand to address this further. TO RESIST OR REVISIT HERESY? DOES THE NUCLEOLUS PARTICIPATE IN MESSENGER RNA BIOSYNTHESIS?

Beyond the possible roles of the nucleolus in cell-cycle control, viral replication, stem cell biology, and cellular senescence, there is another potential nucleolus function that warrants consideration, both for its historical precedence and its now reasonable plausibility based on a number of studies. This is the possibility that the nucleolus is involved, somehow, in the production of messenger RNA. In the 1960s and 1970s, Henry Harris at the University of Oxford posited that the nucleolus was a way station for mRNA export from the nucleus (see Harris 1974). He had been a pioneer of somatic cell genetics, unsullied as such, but was also a constitutional contrarian. He had become something of an enfant terrible outside his immediate field by challenging the thenemerging (and prevailing) views on the meaning of high molecular weight nuclear RNA as regards the pathway of eukaryotic messenger RNA biosynthesis (at a time when the connection was admittedly still wobbly). But, in contrast to his mRNA biosynthesis contrarianism, Harris’ idea that the nucleolus might be involved in mRNA export was based on results, not mere speculation. He had fused chicken erythrocytes, containing a condensed, inactive nucleus, with HeLa cells and found that chicken-specific proteins appeared in these heterokaryons only after the erythrocyte nucleus had undergone chromatin decondensation and the appearance of nucleoli. From this, he elaborated, and for years promulgated, the idea that messenger RNA must

The Nucleolus

Figure 5. The nucleolar granular component contains both RNA-rich and RNA-deficient particles. Electron spectroscopic imaging (ESI) is a mode of energy loss measurement in electron microscopy that can be tuned to record beam collisions with orbital electrons of nitrogen or phosphorus in the specimen. Using the empirical chemical formulas of protein and nucleic acid with respect to these two elements, ESI allows one to construct a map of protein-rich versus nucleic-acid-rich objects. Shown is an ESI image from a portion of a human neuroblastoma cell nucleus spanning a region of the nucleoplasm and part of a nucleolus. The protein-rich and nucleicacid-rich regions have been colored blue and yellow, respectively. (DCh) decondensed chromatin, (CCh) condensed chromatin, (PML) promyelocytic leukemia body, (GC) granular component of the nucleolus. The PML body (entirely blue, i.e., protein) serves as a valuable internal control, as it is known not to contain appreciable nucleic acid. The granular component of the nucleolus, once thought to be a uniform zone of nascent ribosomes, contains an interspersed landscape of nucleic-acid-rich versus nucleic-acid-deficient particles. ESI cannot distinguish between DNA and RNA, but since several independent methods have detected no DNA in the nucleolar granular component, the nucleic-acid-rich particles are plausibly nascent ribosomes, perhaps with a contribution from assembling signal recognition particles, whereas the nucleic-acid-deficient, protein-rich particles may be heterotypic complexes of cell-cycle regulatory proteins, as discussed. (Reprinted from Politz JC et al. 2005, Mol Biol Cell 16: 3401–3410.)

obligatorily pass through the nucleolus. The contemporaneous work of Penman, Darnell, and other groups dispelled this notion for mRNA as a population, en masse, but in a few corners of the field the possibility lingered that

a subset of mRNAs might visit the nucleolus prior to export. Subsequently, there were reports of specific mRNAs in the nucleolus (reviewed in Pederson 1998a) and a recent study (Kim et al. 2009) has rekindled these earlier observations and carried them into the modern era of methodology. In addition, several microRNAs have now been localized in the nucleolus (Politz et al. 2006; Politz et al. 2009) and there is increasing evidence that some microRNAs may be derived from snoRNAs (discussed in Politz et al. 2009; see also Taft et al. 2009). There is also recent evidence for transfer RNA-derived microRNAs (reviewed by Pederson 2010b), and given that the nucleolus has been implicated in tRNA biosynthesis, this is yet another fascinating possibility for a link between the nucleolus and mRNA regulation via microRNAs. Although it is counterintuitive that negative regulators of mRNA function would be associating with nascent mRNAs before the latter even leave the nucleus, intuition has often been an unreliable docent in the field of gene expression. The possible role of the nucleolus in mRNA export and/or microRNA function and dynamics certainly warrants further investigation. NEW FRONTIERS: WHAT LIES BENEATH

Among the remaining issues, the classical phenomenon of nucleolar dominance stands ready for renewed analysis, as has been happening productively (Earley et al. 2010; reviewed by Costa-Nunes et al. 2010). The role of intergenic transcripts in the control of ribosomal RNA transcription (Mayer et al. 2006; Santoro et al. 2010) is another fertile area of current investigation (and likely intersects with nucleolar dominance). Moreover, there is growing evidence that a gene’s positioning or repositioning near or at the nucleolus can regulate its expression (e.g., Royo et al. 2009) and that the rDNA can itself impact the genome globally (e.g., Parades and Maggert 2009). Understanding the process of nucleolar assembly is also a high priority and although there has been substantial progress on this from a cell biological perspective (reviewed by Hernandez-Verdun et al. 2002), a new system

T. Pederson

(Prieto and McStay 2007, 2008) promises molecular detail. A role for the nucleolus has recently emerged in studies of how a postsynaptic density protein of dendritic spines shuttles back to the cell body after neurotransmitter activation (reviewed by Richer and Fallon 2007), raising the possibility of an entire new field linking the nucleolus to neurobiology. Also foremost on the agenda is continued investigation of the extent to which the nucleolus directs the cell cycle (Pederson 2007), as well as the clinical utility of the PNC. A link between the nucleolus, p53, and innate immunity in C. elegans (Fuhrman et al. 2009) is also among the most provocative recent findings and warrants immediate exploration both in nematodes and in a wider phyletic range. Also standing before us is the entire question of mRNA and microRNA traffic through the nucleolus and its functional meaning. Yet, other new vistas on the nucleolus horizon will almost certainly appear in due course, but meanwhile, the agenda is both full and promising. CONCLUSION

The nucleolus is unique in that the discovery of one of its functions took far longer, namely more than a century, than was the case for most other cell components. Another half-century has brought the realization that the nucleolus is also the site of signal recognition particle biosynthesis, serves as a regulatory zone of cell-cycle progression mediators, and is a locus of mRNA and microRNA traffic, the functional meaning of which remains to be discovered. The nucleolus is now known to be a more dynamic domain of nuclear organization than once thought, and with functions even beyond those first grandly recognized atop a Uruguay mountain in 1965. ACKNOWLEDGMENTS

Cited work from the author’s laboratory was supported by grants R01 GM-21595 and R01 GM60551 from the National Institutes of Health, grant MCB-0445841 from the National Science Foundation, and grant RGP0031 from the Human Frontier Scientific Program Organization

(with co-investigators Jan Ellenberg, Angus Lamond, and Matthias Mann). Marty Jacobson in the author’s lab played a key role in the signal recognition particle work that was a catalyst in formulating the plurifunctional nucleolus hypothesis. Joan Politz creatively advanced and confirmed this hypothesis in her SRP work, amidst other pioneer research she led on mRNA and rRNA intranuclear dynamics in the author’s lab. I am greatly indebted to them both. I am also grateful to Joseph Gall, Susan Gerbi, and Masayasu Nomura for their longstanding support of my research on the nucleolus, encouraging my efforts into unsettled and unexplored terrain. REFERENCES Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P. 2002. Molecular Biology of the Cell, 4th edition. Garland Science, New York. p. 332. Andersen JS, Lyon CE, Fox AH, Leung AK, Lam YW, Steen H, Mann M, Lamond AI. 2002. Directed proteomic analysis of the human nucleolus. Curr Biol 12: 1 –11. Bachellerie JP, Cavaille´ J, Hu¨ttenhofer A. 2002. The expanding RNA world. Biochimie 84: 775–790. Boisvert FM, van Koningsbruggen S, Navascue´s J. Lamond AI. 2007. The multifunctional nucleolus. Nat Rev Mol Cell Biol 8: 574– 585. Boyne JR, Whitehouse A. 2006. Nucleolar trafficking is essential for nuclear export of intronless herpesvirus mRNA. Proc Natl Acad Sci USA 103: 15190– 15195. Brown DD, Gurdon JB. 1964. Absence of rRNA synthesis in the anucleolate mutant of X. laevis. Proc Natl Acad Sci USA 51: 139 –146. Brown DD, Dawid IG. 1968. Specific gene amplication in oocytes. Oocyte nuclei contain extrachromosomal replicas of the genes for ribosomal RNA. Science 160: 272– 280. Caesar J. (40-50s BC). De Bello Gallico I: 1. Calvet JP, Pederson T. 1981. Base-pairing interactions between snRNAs and nuclear RNA precursors as revealed by psoralen cross-linking in vivo. Cell 26: 363 –370. Chen D. Huang S. 2001. Nucleolar components involved in ribosome biogenesis cycle between the nucleolus and nucleoplasm in interphase cells. J Cell Biol 153: 169– 176. Ciufo LF, Brown JD. 2000. Nuclear export of yeast signal recognition particle lacking Srp54p by the Xpo1p/Crm1p NES-dependent pathway. Curr Biol 10: 1256– 1264. Costa-Nunes P, Pontes O, Preuss SB. Pikaard CS. 2010. Extra views on RNA-dependent DNA methylation and MBD6dependent heterochromatin formation in nucleolar dominance. Nucleus 1: 1– 6. Deryusheva S, Gall JG. 2004. Dynamics of coilin in Cajal bodies of the Xenopus germinal vesicle. Proc Natl Acad Sci USA 101: 4810–4814.

The Nucleolus Earley KW, Pontvianne F, Wierzbicki AT, Blevins T, Tucker S, Costa-Nunes P, Pontes O, Pikaard CS. 2010. Mechanisms of HDA6-mediated rRNA gene silencing; suppression of intergenic pol II transcription and differential effects on maintenance versus siRNA-directed cytosine methylation. Genes Dev 24: 1119–1132. Fatica A, Tollervey D. 2002. Making ribosomes. Curr Opin Cell Biol 14: 313 –318. Fichelson P, Moch C, Ivanovitch K, Martin C, Sidor CM, Lepesant J-A, Bellaiche Y. Huynh J-R. 2009. Live-imaging of single stem cells within their niche reveals that a U3 snoRNP component segregates asymmetrically and is required for self-renewal in Drosophila. Nat. Cell Biol 11: 685–693. Fle´chon JE. 2006. Analysis of the nucleolar compartment of the nucleus as an indicator of nuclear reprogramming after nuclear transfer. Meth Mol Biol 325: 225 –238. Fuhrman LE, Goel AK, Smith J, Shianna KV, Aballay A. 2009. Nucleolar proteins suppress Caenorhabditis elegans innate immunity by inhibiting p53/CEP-1. PLoS Genetics 5: e1000657 Gall JG. 1968. Differential synthesis of the genes for ribosomal RNA during amphibian oogenesis. Proc Natl Acad Sci USA 60: 553– 560. Garcia SN, Pillus L. 1999. Net results of nucleolar dynamics. Cell 97: 825– 828. Ghetti A, Pinol-Roma S, Michael WM, Morandi C. Dreyfuss G. 1992. hnRNP I, the polypyrimidine tract-binding protein: distinct nuclear localization and association with hnRNAs. Nucleic Acids Res 20: 3671– 3678. Gonda K, Fowler J, Katoku-Kikyo N, Haroldson J, Wudel J, Kikyo N. 2003. Reversible disassembly of somatic nucleoli by the germ cell proteins FRGY2a and FRGY2b. Nat Cell Biol 5: 205–210. Granneman S, Baserga SJ. 2004. Ribosome biogensis: of knobs and RNA processing. Exp Cell Res 296: 43– 50. Grosshans H, Deinhert K, Hurt E, Somos G. 2001. Biogenesis of the signal recognition particle (SRP) involves import of SRP proteins into the nucleolus, assembly with the SRP RNA, and Xpo1p-mediated export. J Cell Biol 153: 745– 761. Guarente L. 1997. Link between aging and the nucleolus. Genes Dev 11: 2449–2455. Handwerger KE, Cordero JA, Gall JG. 2005. Cajal bodies, nucleoli and speckles have a low-density, sponge-like structure. Mol Biol Cell 16: 202– 211. Harris H. 1974. Nucleus and Cytoplasm 3rd edition. Clarendon Press, Oxford. Heitz E. 1931. Nukleolar und chromosomen in der gattung. Vicia Planta 15: 495–505. Hernandez-Verdun D, Roussel P. Ge´brane-Youne`s J. 2002. Emerging concepts of nucleolar assembly. J Cell Sci 115: 2265– 2270. Huang S. 2000. Perinucleolar structures. J Struct Biol 129: 233–240. Jacobson MR, Cao LG, Wang YL, Pederson T. 1995. Dynamic localization of RNase MRP RNA in the nucleolus observed by fluorescent RNA cytochemistry in living cells. J Cell Biol 131: 1649– 1658.

Jacobson MR, Cao LG, Taneja K, Singer RH, Wang YL, Pederson T. 1997. Nuclear domains of the RNA subunit of RNase P. J Cell Sci 110: 829 –837. Jacobson MR, Pederson T. 1998a. A 7-methylguanosine cap commits U3 and U8 small nuclear RNAs to the nucleolar localization pathway. Nucleic Acids Res 26: 756–760. Jacobson MR, Pederson T. 1998b. Localization of signal particle RNA in the nucleolus of mammalian cells. Proc Natl Acad Sci USA 95: 7981– 7986. Kass S, Tyc K, Steitz JA, Sollner-Webb B. 1990. The U3 small nucleolar ribonucleoprotein functions in the first step of pre-ribosomal RNA processing. Cell 60: 897–908. Kim SH, Koroleva OA, Lewandowska D, Pendle AF, Clark GP, Simpson CG, Shaw PJ, Brown JW. 2009. Aberrant mRNA transcripts and the nonsense-mediated decay proteins UPF2 and UPF3 are enriched in the Arabidopsis nucleolus. Plant Cell 21: 2045– 2057. Kim SH, Ryabov EV, Kalinina NO, Rakitina DV, Gillespie T, MacFarlane S, Haupt S, Brown JWS, Taliansky M. 2007. Cajal bodies and the nucleolus are required for a plant virus systemic infection. EMBO J 26: 2169–2179. Koberna K, Malinsky J, Pliss A, Masata M, Vecerova J, Fiolova M, Bednar J, Rasˇka I. 2002. Ribosomal genes in focus: new transcripts label the dense fibrillar components and form clusters indicative of “Christmas trees”. J Cell Biol 157: 743– 748. Kulikov R, Letienne J, Kaur M, Grossman SR, Arts J, Blattner C. 2010. Mdm2 facilitates the association of p53 with the proteasome. Proc Natl Acad Sci USA 107: 10038– 10043. Kumar A, Warner JR. 1972. Characterization of ribosomal precursor particles from HeLa cell nucleoli. J Mol Biol 63: 233– 246. Kyng KJ, May A, Kø´vraa S, Bohr VA. 2003. Gene expression profiling in Werner syndrome closely resembles that of normal aging. Proc Natl Acad Sci USA 100: 12259– 12264. Lam YW, Lamond AI, Mann M, Andersen JS. 2007. Analysis of nucleolar protein dynamics reveals the nuclear degradation of ribosomal proteins. Curr Biol 17: 749 –760. Leung AKL, Andersen JS, Mann M, Lamond AI. 2003a. Bioinformatic analysis of the nucleolus. Biochem J 376: 553 –569. Leung AKL, Lamond AI. 2003. The dynamics of the nucleolus. Crit Rev Euk Gene Expr 13: 39– 54. Lewin B. 1980. Gene Expression. Volume 2. Eukaryotic Chromosomes. 2nd edition. John Wiley & Sons, New York. Liau MC, Perry RP. 1969. Ribosome precursor particles in nucleoli. J Cell Biol 42: 272– 283. Ma H, Pederson T. 2007. Depletion of the nucleolar protein nucleostemin causes G1 cell cycle arrest via the p53 pathway. Mol Biol Cell 18: 2630–2635. Ma H, Pederson T. 2008a. Nucleostemin: a multiplex regulator of cell-cycle progression. Trends Cell Biol 18: 575– 579. Ma H, Pederson T. 2008b. Nucleophosphmin is a binding partner of nucleostemin in human osteosarcoma cells. Mol Biol Cell 19: 2870– 2875. Maggio R, Siekevitz P, Palade GE. 1963. Studies on isolated nuclei. I. Isolation and chemical composition of nucleolar and nucleoplasmic subfractions. J Cell Biol 18: 293 –312.

T. Pederson Marciniak RA, Lombard DB, Johnson FB, Guarente L. 1998. Nucleolar localization of the Werner syndrome protein in human cells. Proc Natl Acad Sci USA 95: 6887–6892. Martindill DMJ, Risebro CA, Smart N, Franco-Viseras MDM, Rosario CO, Swallow CJ, Dennis JW, Roliry PR. 2007. Nucleolar release of Hand1 acts as a molecular switch to determine cell fate. Nat Cell Biol 9: 1131– 1141. Matera AG, Frey MR, Margelot K, Wolin SL. 1995. A perinucleolar compartment contains several RNA polymerase III transcripts as well as the polypyrimidine tract-binding protein. J Cell Biol 129: 1181– 1193. Mayer C, Schmitz KM, Li J, Grummt I, Santoro R. 2006. Intergenic transcripts regulate the epigenetic state of rRNA genes. Mol Cell 22: 351– 361. McClintock B. 1934. The relationship of a particular chromosomal element to the development of the nucleoli in Zea mays. Z. Zellforsch Mikrosk 21: 294– 398. Michienzi A, Li S, Zaia JS, Rossi JJ. 2002. A nucleolar TAR decoy inhibitor of HIV-1 replication. Proc Natl Acad Sci USA 99: 14047–14052. Miller OL Jr, Beatty BR. 1969. Visualization of nucleolar genes. Science 164: 955– 957. Misteli T. 2003. A nucleolar disappearing act in somatic cloning. Nat Cell Biol 5; 183 –184. Misteli T. 2005. Going in GTP cycles in the nucleolus. J Cell Biol 168: 177–178. Montgomery TH. 1898. Comparative cytological studies, with especial regard to the morphology of the nucleolus. J Morphol 15: 265– 582. Olson MO, Dundr M, Szebeni A. 2000. The nucleolus: an old factory with unexpected capabilities. Trends Cell Biol 10: 189– 196. Olson MO, Hingorani K, Szebeni A. 2002. Conventional and nonconventional roles of the nucleolus. Int Rev Cytol 219: 199– 266. Parades S, Maggert KA. 2009. Ribosomal DNA contributes to global chromatin regulation. Proc Natl Acad Sci USA 106: 17829– 17834. Pederson T. 1998a. The plurifunctional nucleolus. Nucleic Acids Res 26: 3871–3876. Pederson T. 1998b. Growth factors in the nucleolus? J Cell Biol 143: 279–281. Pederson T. 2000. Diffusional protein transport within the nucleus: a message in the medium. Nat Cell Biol 2: E73– E74. Pederson T. 2001a. Fluorescent RNA cytochemistry: tracking gene transcripts in living cells. Nucleic Acids Res 29: 1013– 1016. Pederson T. 2001b. Protein mobility within the nucleuswhat are the right moves? Cell 104: 635–638. Pederson T. 2002. Proteomics of the nucleolus: more proteins, more functions? Trends Biochem Sci 27: 111– 112. Pederson T. 2007. Ribosomal protein mutations in Diamond-Blackfan anemia: might they operate upstream of protein synthesis? The FASEB J 21: 3442– 3445. Pederson T. 2010a. “Compact” nuclear domains: reconsidering the nucleolus. Nucleus 1: 444– 445. Pederson T. 2010b. Regulatory derived from transfer RNA? RNA 16: 1865– 1869.

Pederson T, Kumar A. 1971. Relationship between protein synthesis and ribosome assembly in HeLa cells. J Mol Biol 61: 655 –668. Pederson T, Politz JC. 2000. The nucleolus and the four ribonucleoproteins of translation. J Cell Biol 148: 1091–1095. Pederson T, Tsai RYL. 2009. In search of nonribosomal protein function and regulation. J Cell Biol 184: 771– 776. Pendle AF, Clark GP, Boon R, Lewandowska D, Lam YW, Andersen J, Mann M, Lamond AI, Brown JW, Shaw PJ. 2005. Proteomic analysis of the Arabidopsis nucleolus suggests novel nucleolar functions. Mol Biol Cell 16: 260– 269. Perry RP. 1966. Nucleolus: Structure and function. Science 153: 214–219. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604 –609. Politz JC. 1999. Use of caged fluorochromes to track macromolecular movement in living cells. Trends Cell Biol 9: 284 –287. Politz JC, Browne ES, Wolf DE, Pederson T. 1998. Intranuclear diffusion and hybridization state of oligonucleotides measured by fluorescence correlation spectroscopy. Proc Natl Acad Sci USA 95: 6043– 6048. Politz JCR, Hogan EM, Pederson T. 2009. MicroRNAs with a nucleolar location. RNA 15: 1705– 1715. Politz JC, Lewandowski LB, Pederson T. 2002. Signal recognition particle RNA localization within the nucleolus differs from the classical sites of ribosome synthesis. J Cell Biol 159: 411 –418. Politz JCR, Polena I, Trask I, Bazett-Jones DP, Pederson T. 2005. A nonribosomal landscape in the nucleolus revealed by the stem cell protein nucleostemin. Mol Biol Cell 16: 3401– 3410. Politz JC, Tuft RA, Pederson T. 2003. Diffusion-based transport of nascent ribosomes in the nucleus. Mol Biol Cell 14: 4805–4812. Politz JCR, Tuft RA, Pederson T. 2005. Photoactivationbased labeling and in vivo tracking of RNA molecules in the nucleus. Live Cell Imaging: A Laboratory Manual Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Politz JC, Yarovoi S, Kilroy S, Gowda K, Zwieb C, Pederson T. 2000. Signal recognition particle components in the nucleolus. Proc Natl Acad Sci USA 97: 55–60. Politz JCR, Zhang F, Pederson T. 2006. MicroRNA-206 colocalizes with ribosome-rich regions in both the nucleolus and cytoplasm of rat myogenic cells. Proc Natl Acad Sci USA 103: 18957– 18962. Pollock C, Huang S. 2010. The perinucleolar compartment. Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect. a000679. Prieto J-L, McStay B. 2007. Recruitment of factors linking transcription and processing of pre-rRNA to NOR chromatin is UBF-dependent and occurs independent of transcription in human cells. Genes Dev 21: 2041–2054. Prieto J-L, McStay B. 2008. Pseudo-NORs; a novel method for studying nucleoli. Biochim et Biophys Acta 1783: 2116–2123. Rasˇka I. 2003. Oldies but goldies: searching for Christmas trees within the nucleolar architecture. Trends Cell Biol 13: 517– 525.

The Nucleolus Rasˇka I, Shaw PJ, Cmarko D. 2006. Structure and function of the nucleolus in the spotlight. Curr Opin Cell Biol 18: 325–334. Richter J.D. Fallon JR. 2007. Synapses go nucle(ol)ar. Nat Neurosci 10: 399–400. Royo F, Paz N, Espinosa L, McQueen PG, Vello´n L, Parada LA. 2009. Spatial link between nucleoli and expression of the ZacI gene. Chromosoma 118: 711– 722. Rubbi CP, Milner J. 2003. Disruption of the nucleolus mediates stabilization of p53 in response to DNA damage and other stressors. EMBO J 22: 6068– 6077. Santoro R, Schmitz K-M, Sandoval J, Grummt I. 2010. Intergenic transcripts originating from a subclass of ribosomal DNA repeats silence ribosomal RNA genes in trans. EMBO rep 11: 52– 58. Scheer UHock R. 1999. Structure and function of the nucleolus. Curr Opin Cell Biol 11: 385 –390. Scherl A, Coute´ Y, De´on C, Calle´ A, Kindbeiter K, Sanchez JC, Greco A, Hochstrasser D, Diaz JJ. 2002. Functional proteomic analysis of human nucleolus. Mol Biol Cell 13: 4100–4109. Sinclair DA, Milles K, Guarente L. 1997. Accelerated aging and nucleolar fragmentation in yeast sgs1 mutants. Science 277: 1313– 1316. Soeiro R, Basile C. 1973. Non-ribosomal nucleolar proteins in HeLa cells. J Mol Biol 79: 507 –519. Sommerville J, Brumwell CL, Politz JCR, Pederson T. 2005. Signal recognition particle assembly in relation to the function of amplified nucleoli of Xenopus oocytes. J Cell Sci 118: 1299– 1307. Sonntag F, Schmidt K, Kleinschmidt JA. 2010. A viral assembly factor promotes AAV2 capsid assembly in the nucleolus. Proc Natl Acad Sci USA 107: 10220– 10225. Spell J, Kubitscheck U. 2009. Single ovalbumin molecules exploring nucleoplasm and nucleoli of living cell nuclei. Biochim Biophys Acta 1803: 396 –404. Stavreva DA, Kawasaki M, Dundr M, Koberna K, Muller WG, Tsujimura-Takahashi T, Komatsu W, Hayano T, Isobe T, Rasˇka I, et al. 2006. Potential roles for ubiquitin and the proteasome during ribosome biogenesis. Mol Cell Biol 26: 5131– 5145.

Szekely AM, Chen Y-H, Zhang C, Oshima J, Weissman SM. 2000. Werner protein recruits DNA polymerase d to the nucleolus. Proc Natl Acad Sci USA 97: 11365– 11370. Taft RJ, Glazov EA, Lassmann T, Hayashizaki Y, Carninci P, Mattick JS. 2009. Small RNAs derived from snoRNAs. RNA 15: 1233–1240. Tsai RY, McKay RD. 2002. A nucleolar mechanism controlling cell proliferation in stem cells and cancer cells. Genes Dev 16: 2991–3003. Tsai RY, McKay RD. 2005. A multistep, GTP-driven mechanism driving the dynamic cycling of nucleostemin. J Cell Biol 168: 179 –184. Valentin G. 1836. Repertorium fu¨r Anatomie und Physiologie 1: 1 –293. Verlag von Veit und Comp. Berlin. Valentin G. 1839. Repertorium fu¨r Anatomie und Physiologie 4: 1 –275. Verlag von Veit und Comp. Berlin. Vincent WS. 1952. The isolation and chemical properties of the nucleoli of starfish oocytes. Proc Natl Acad Sci USA 38: 139– 145. Vincent WS, Miller OL. 1966. International symposium on the nucleolus. Its structure and function. Montevideo, Uruguay. J Natl Cancer Inst Monogr 23: 1– 610. Vinistin R, Amon A. 2000. The nucleolus: the magician’s hat for cell cycle tricks. Curr Opin Cell Biol 12: 372– 377. von Kobbe C, Bohr VA. 2002. A nucleolar targetting sequence in the Werner syndrome protein resides within residues 949-1092. J Cell Sci 115: 3901–3907. Wagner R. 1835. Einige Bemerkungen und Fragen u¨ber das Keimbla¨schen (vesicular germinativa). Mu¨ller’s Archiv Anat Physiol Wissenschaft Med 373– 377. Wang J, Cao LG, Wang YL, Pederson T. 1991. Localization of pre-messenger RNA at discrete nuclear sites. Proc Natl Acad Sci 88: 7391–7395. Wang C, Politz JC, Pederson T, Huang S. 2003. RNA polymerase III transcripts and the PTB protein are essential for the integrity of the perinucleolar compartment. Mol Biol Cell 14: 2425–2435. Warner JR, Soeiro R. 1967. Nascent ribosomes from HeLa cells. Proc Natl Acad Sci USA 58: 1984– 1990. Weber JD, Taylor LJ, Roussel MF, Sherr CJ, Bar-Sagi D. 1999. Nucleolar Arf sequesters Mdm2 and activates p53. Nat Cell Biol 1: 20– 26.

Nuclear Speckles David L. Spector1 and Angus I. Lamond2 1

Cold Spring Harbor Laboratory, One Bungtown Road, Cold Spring Harbor, New York 11724

2

Wellcome Trust Centre for Gene Regulation and Expression, College of Life Sciences, University of Dundee, MSI/WTB/JBC Complex Dow Street, Dundee DD1 5EH, United Kingdom

Correspondence: [email protected], [email protected]

Nuclear speckles, also known as interchromatin granule clusters, are nuclear domains enriched in pre-mRNA splicing factors, located in the interchromatin regions of the nucleoplasm of mammalian cells. When observed by immunofluorescence microscopy, they usually appear as 20– 50 irregularly shaped structures that vary in size. Speckles are dynamic structures, and their constituents can exchange continuously with the nucleoplasm and other nuclear locations, including active transcription sites. Studies on the composition, structure, and dynamics of speckles have provided an important paradigm for understanding the functional organization of the nucleus and the dynamics of the gene expression machinery.

he mammalian cell nucleus is a highly compartmentalized yet extremely dynamic organelle (reviewed in Misteli 2001a; Spector 2006; Zhao et al. 2009). Many nuclear factors are localized in distinct structures, such as speckles, paraspeckles, nucleoli, Cajal bodies, polycomb bodies, and promyelocytic leukemia bodies and show punctate staining patterns when analyzed by indirect immunofluorescence microscopy (reviewed in Lamond et al. 1998; Spector 2001; Spector 2006). In mammalian cells the pre-mRNA splicing machinery, including small nuclear ribonucleoprotein particles (snRNPs), spliceosome subunits, and other non-snRNP protein splicing factors, shows a punctate nuclear localization pattern that is usually termed “a speckled pattern” but has also been referred to as “SC35 domains (Wansink et al. 1993)” or “splicing factor

T

compartments (Phair et al. 2000)” (Figs. 1 and 2). The first detailed description of the nuclear domains that we presently refer to as nuclear speckles was reported by Santiago Ramo´n y Cajal in 1910 (Ramo´n y Cajal 1910; reviewed in Lafarga et al. 2009). Ramo´n y Cajal used acid aniline stains to identify structures he referred to as “grumos hialinas” (literally “translucent clumps”). The term “speckles” was first put forth in 1961 by J. Swanson Beck (Beck 1961) upon examination of rat liver sections immunolabeled with the serum of individuals with autoimmune disorders. Although the connection was not made at the time, these speckles had been identified two years earlier by Hewson Swift (Swift 1959) at the electron microscopic level and called interchromatin particles. Swift observed that these particles were not randomly distributed but that they occurred in localized

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2011 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000646 Cite this article as Cold Spring Harb Perspect Biol 2011;3:a000646

D.L. Spector and A.I. Lamond

Figure 1. Speckles form in the interchromatin space.

HeLa cells showing splicing factors localized in a speckled pattern as well as being diffusely distributed throughout the nucleoplasm. Bar ¼ 5 mm.

“clouds,” and cytochemical analysis indicated that they contained RNA (Swift 1959). However, the first link between pre-mRNA splicing and nuclear speckles or interchromatin granule clusters came from an examination of the distribution of snRNPs using anti-splicing factor-specific antibodies, demonstrating a speckled distribution pattern of snRNPs in cell nuclei (Lerner et al. 1981; Perraud et al. 1979; Spector et al. 1983). It is now clear that much of the punctate localization of splicing factors observed by immunofluorescence microscopy corresponds to the presence of these factors in nuclear speckles of variable size and irregular shape that are revealed by electron microscopy as interchromatin granule clusters (IGCs) (Fig. 3). IGCs range in size from one to several micrometers in diameter and are composed of 20 – 25 nm granules that are connected in places by a thin fibril resulting in a beaded chain appearance (Thiry 1995b). These structures can be observed by electron microscopy without antibody labeling (Thiry 1995b). We will define “speckles” here specifically as the IGC component of the

splicing factor labeling pattern, and distinguish this from other nuclear structures, including perichromatin fibrils and Cajal bodies, which also contain splicing factors (reviewed in Fakan 1994; Spector 1993). For some of the speckle components a speckle targeting signal has been identified. The arginine/serine-rich domain (RS domain) of some SR pre-mRNA splicing factors has been shown to be necessary and sufficient for the targeting of these factors to nuclear speckles (Caceres et al. 1997; Hedley et al. 1995; Li et al. 1991). In addition, the threonine-proline repeats of SF3b1 (Eilbracht et al. 2001) and the forkhead-associated domain in NIPP1 (Jagiello et al. 2000) have also been implicated in speckle-targeting. Most recently, Salichs et al. (Salichs et al. 2009) performed a genomewide analysis of homopolymeric histidine tracts and identified 86 human proteins that contain stretches of five or more histidines. Of the 22

Figure 2. Structured illumination microscopy, using the OMX system (Applied Precision, Issaqua, Washington), of a HeLa cell expressing SC35-EYFP. At 100 nm resolution substructure can be observed within speckles. In addition, the diffuse population of SC35-EYFP is resolved as a granular distribution. Projection of twelve 0.125 mm optical sections through the center of a nucleus encompassing 1.5 mm. Image provided by Zsolt Lazar and R. Ileng Kumaran. Bar ¼ 2 mm.

Nuclear Speckles

studies using an anti-splicing factor antibody to confirm the localization of any novel factors to nuclear speckles. STRUCTURE AND LOCATION OF SPECKLES

Figure 3. Nuclear speckles are equivalent to inter-

chromatin granule clusters. Immunoelectron microscopy using a primary antibody against SC35 and a secondary antibody conjugated to 15 nm colloidal gold. IGCs are composed of a series of particles measuring 20 –25 nm in diameter that are connected in places by a thin fibril resulting in a beaded chain appearance. Bar ¼ 500 nm.

that were nuclear localized, 15 were shown to be present in nuclear speckles. Based on these data, the polyHis-repeats were proposed to act as a speckle-targeting signal that functions by acting as an interaction surface for resident nuclear speckle constituents. Interestingly, these targeting signals rely mostly on charge effects, being basic protein regions. Interestingly, structures similar to nuclear speckles have been identified in the amphibian oocyte nucleus (Gall et al. 1999) and in Drosophila melanogaster embryos when transcription increases upon cellularization during cycle 14 (Segalat et al. 1992), but not in yeast (Potashkin et al. 1990). Importantly, not all nuclear proteins that show a speckle-like labeling pattern by immunofluorescence microscopy localize to IGCs. For example, the ER repeat protein YT521-B localizes in a speckled-like distribution that corresponds to YT bodies (Nayler et al. 2000), whereas PSPC1 localizes in approximately 5– 20 punctate interchromatin structures termed “paraspeckles,” which are also distinct from nuclear speckles (Fox et al. 2002; Fox and Lamond 2010). Therefore, it is essential to perform double-label immunofluorescence

As determined by both light and electron microscopy, the clusters of interchromatin granules that constitute speckles form throughout the nucleoplasm in regions containing little or no DNA (Thiry 1995b). Although they apparently contain few, if any, genes speckles are often observed close to highly active transcription sites. This suggests that they likely have a functional relationship with gene expression, and some specific genes have been reported to preferentially localize near speckles (Brown et al. 2008; Huang et al. 1991; Johnson et al. 2000; Moen et al. 2004; Smith et al. 1999; Xing et al. 1993; Xing et al. 1995), although this does not appear to be obligatory for transcription/premRNA splicing. Interestingly, Shopland et al. (Shopland et al. 2003) found that gene-rich chromosomal regions (R-bands) are more frequently found along the edge of nuclear speckles than gene-poor regions (G-bands). In addition, coordinately expressed active genes can be found in association with the same nuclear speckle. Based on these findings, Shopland et al. (2003) suggested that nuclear speckles act as functional centers that organize active genes on their periphery to form euchromatic neighborhoods. Several lines of evidence point to speckles acting as storage/assembly/modification compartments that can supply splicing factors to active transcription sites (reviewed in Lamond and Spector 2003). For example, a series of high resolution pulse-labeling experiments analyzed at the electron microscopic level, studying the incorporation of either tritiated uridine or Br-UTP after short pulses, have shown that nascent pre-mRNA is predominantly localized outside of nuclear speckles (IGCs) in fibrillar structures, 3– 5 nm in diameter, which are termed perichromatin fibrils (PFs) (Cmarko et al. 1999; Fakan et al. 1971; Fakan et al. 1978; Monneron et al. 1969). It is likely that most of the cotranscriptional splicing

D.L. Spector and A.I. Lamond

is associated with these PFs, rather than within IGCs. PFs can occur both on the periphery of IGCs and in nucleoplasmic regions away from IGCs (Fakan 1994). Some apparent discrepancies in the literature concerning the possible direct role of speckles as splicing sites may have arisen because the PFs can show a close topological relationship with the periphery of IGCs. Using the fluorescence microscope it is difficult to distinguish these PFs from the IGCs. In addition, as highly expressed genes will recruit a significant amount of pre-mRNA splicing factors (Huang et al. 1996), these regions of highly active transcription will be indistinguishable from IGCs at the fluorescence microscopy level. Although many in the field do not view speckles as direct transcription/pre-mRNA splicing centers, others suggest that they may have a more direct role relating to the splicing and transport of pre-mRNA (reviewed in Hall et al. 2006; Melcak et al. 2000; Shopland et al. 2002; Wei et al. 1999). COMPOSITION OF SPECKLES

Many pre-mRNA splicing factors, including snRNPs and SR proteins (Fu 1995), have been localized to nuclear speckles by either immunofluorescence, fluorescent protein-tagging, and/ or immunoelectron microscopy. In fact, this speckled localization pattern is highly diagnostic for proteins involved in pre-mRNA splicing. In addition, several kinases (Clk/STY, hPRP4, and PSKHI) (Brede et al. 2002; Colwill et al. 1996; Ko et al. 2001; Kojima et al. 2001; SaccoBubulya et al. 2002) and phosphatases (PP1) (Trinkle-Mulcahy et al. 1999; Trinkle-Mulcahy et al. 2001) that phosphorylate/dephosphorylate components of the splicing machinery have also been localized to nuclear speckles. This supports the idea that speckles may be involved in regulating the pool of factors that are accessible to the transcription/pre-mRNA processing machinery (reviewed in Misteli et al. 1997b). The protein composition of nuclear speckles has been assessed by proteomic analysis of an enriched IGC fraction purified from mouse liver nuclei. This approach identified 146 known proteins, as well as numerous uncharacterized

proteins (Mintz et al. 1999; Saitoh et al. 2004). The proteomic information, together with additional localization studies, has revealed that speckles contain many other proteins apart from pre-mRNA splicing factors. Of particular interest is the localization of transcription factors (Larsson et al. 1995; Mortillaro et al. 1996; Zeng et al. 1997), 30 -end RNA processing factors (Krause et al. 1994; Schul et al. 1998), eukaryotic translation initiation factor eIF4E (Dostie et al. 2000), eif4AIII, a protein involved in translation inhibition (Li et al. 1999), and structural proteins (Jagatheesan et al. 1999; Nakayasu et al. 1984; Sharma et al. 2010). Consistent with these findings, proteomic analyses of in vitro assembled spliceosomes indicate that they may also contain transcription and 30 -end RNA processing factors, together with splicing factors, in a higher order complex (Rappsilber et al. 2002; Zhou et al. 2002). However, they do not contain factors primarily involved in ribosome subunit biogenesis or tRNA production, and the protein composition of speckles underlines their close relationship with mechanisms of gene expression by RNA Pol II. Although transcription does not take place within the majority of nuclear speckles (Cmarko et al. 1999) and DNA is not localized to these nuclear regions (reviewed in Thiry 1995b), a population of the serine-2-phosphorylated form of the RNA polymerase II (RNAPII) large subunit (LS) that is involved in elongation has been localized to these regions by immunofluorescence microscopy (Bregman et al. 1995; Mortillaro et al. 1996). In addition, biochemical characterization of the IGC proteome has identified several subunits of RNAPII (Mintz et al. 1999; Saitoh et al. 2004), supporting the presence of a pool of RNAPII in speckles. However, other studies have not observed an enrichment of RNAPII in speckles (Grande et al. 1997; Kimura et al. 2002; Zeng et al. 1997) and it is not present in B snurposomes (Doyle et al. 2002). The Cdk9-cyclin T1 complex, also known as TAK/P-TEFb, is thought to be involved in transcriptional elongation via phosphorylation of the RNAPII LS (reviewed in Price 2000). This complex was found diffusely distributed throughout the nucleoplasm, but not in nucleoli

Nuclear Speckles

(Herrmann et al. 2001). In addition, a significant overlap between cyclin T1 and nuclear speckles was observed. However, although Cdk9 was present in the vicinity of nuclear speckles, the degree of overlap was limited (Herrmann et al. 2001; Matera et al. 1993). Further evidence for a link between transcription and speckles comes from observations on FBI-1, a cellular POZ-domain-containing protein that binds to the HIV-1 long terminal repeat and associates with the HIV-1 transactivator protein Tat (Pessler et al. 1997). FBI-1 has been found to partially colocalize with Tat and its cellular cofactor, P-TEFb at nuclear speckles (Pendergrast et al. 2002). In addition, the nucleosome binding protein HMG-17, which can alter the structure of chromatin and enhance transcription, has been localized in a similar pattern to FBI-1 (Hock et al. 1998). Therefore, although little or no transcription takes place in nuclear speckles, a subset of proteins involved in this process are associated with these nuclear regions in addition to being present at transcription sites. Although it is currently unclear what determines the subset of transcription factors that are localized to nuclear speckles, their presence may relate to the assembly of higher-order complexes and/or to regulatory steps affecting either the modification state or accessibility of specific transcription factors. In addition to transcription factors, a population of poly(A)þ RNA has been localized to nuclear speckles (Carter et al. 1991; Huang et al. 1994; Visa et al. 1993). This population of poly(A)þ RNA does not chase to the cytoplasm when transcription is blocked with aamanitin, as would be expected if these species represented nascent mRNA (Huang et al. 1994). Interestingly, Hutchinson et al. (Hutchinson et al. 2007) identified MALAT1 (metastasis-associated lung adenocarcinoma transcript 1), a long nuclear retained noncoding RNA to be enriched in nuclear speckles (reviewed in Wilusz et al. 2009). Recent studies have implicated MALAT1 in the recruitment of SR splicing factors from nuclear speckles to sites of transcription (Bernard et al. 2010) and in the regulation of alternative splicing by modulating SF2/ ASF phosphorylation (Tripathi et al. 2010).

Although an underlying scaffold that would serve as a platform on which to organize IGCs has thus far not been identified (Sacco-Bubulya et al. 2002), several proteins with possible structural roles in the nucleus, such as a population of lamin A (Jagatheesan et al. 1999) and snRNP-associated actin (Nakayasu et al. 1984), have been detected in nuclear speckles. However, another study failed to detect an alteration in nuclear speckles in LMNA-/- cells (Vecerova et al. 2004). In addition to actin, phosphatidylinositol (4,5)-bisphosphate [PtdIns(4,5)P2], a lipid that regulates actin-binding proteins (Zhao et al. 1998), as well as multiple phosphatidylinositol phosphate kinase (PIPK) isoforms, have also been localized to nuclear speckles (Boronenkov et al. 1998). Recently, Sharma et al. (Sharma et al. 2010) have implicated the 2530amino acid Son protein in the organization of nuclear speckles. Son contains a concentrated region of multiple tandem repeat sequences including multiple serine-rich repeats and an RS domain. Based on RNAi depletion experiments, Son was proposed to act as a scaffolding protein for RNA processing factors in nuclear speckles (Sharma et al. 2010). DYNAMICS OF NUCLEAR SPECKLES

Speckles are dynamic structures; their size, shape, and number can vary, both between different cell types and within a cell type, according to the levels of gene expression and in response to signals that influence the pools of active splicing and transcription factors available. When transcription is halted, either by the use of inhibitors, or as a result of heat shock, splicing factors accumulate predominantly in enlarged, rounded speckles (Melcak et al. 2000; Spector et al. 1991; Spector et al. 1983). The fact that nuclear speckles become round and increase in size upon transcriptional inhibition supports the view that speckles may function in the storage/assembly/modification of splicing factors, and that they are not direct sites of splicing. Furthermore, when expression of introncontaining genes increases (Huang et al. 1996; Misteli et al. 1997a), or during viral infection when transcription levels are high (Bridge et al.

D.L. Spector and A.I. Lamond

1995; Jime´nez-Garcı´a et al. 1993), the accumulation of splicing factors in speckles is reduced, and they redistribute to nucleoplasmic transcription sites. Individual speckle components can therefore shuttle continually between speckles and active gene loci. Speckles are also dynamically regulated during mitosis. The movement of factors into and out of speckles can be directly visualized by fluorescence microscopy as fluctuations in the shape and intensity of speckles in live cells expressing splicing factor/fluorescent protein fusions (Misteli et al. 1997a). Speckles in such cells show transcription-dependent peripheral movements, although individual speckles remain in their neighborhoods. Photobleaching techniques have also been used to measure the flux of some speckle components and have shown that their exchange rate is very rapid (Kruhlak et al. 2000; Phair et al. 2000). Complete recovery for GFP-SF2/ASF (a member of the SR-family of pre-mRNA splicing factors), after photobleaching of the fluorescence signal in speckles, was apparent in approximately 30 seconds with half recovery in approximately 3–5 seconds. The movement rates for splicing factors through the nucleoplasm were measured to be slow compared with free GFP; this reduction in movement was proposed to result from multiple transient interactions of splicing factors with nuclear binding sites, both within and outside of speckles. Kinetic modeling indicated that the maximal mean residence time for GFP-SF2/ASF in speckles was less than 50 seconds (Phair et al. 2000). It is a remarkable feature of nuclear organization that at steady-state the overall structure of speckles, as well as other nuclear domains, persists despite the large flux of their components. THE SPECKLE CELL CYCLE

Upon entry into mitosis and following breakdown of the nuclear envelope/lamina, proteins associated with nuclear speckles become diffusely distributed throughout the cytoplasm (Ferreira et al. 1994; Reuter et al. 1985; Spector et al. 1986; Thiry 1995a). During metaphase, these proteins continue to localize in a diffuse cytoplasmic pattern and also accumulate within

one to three small structures called mitotic interchromatin granules (MIGs) (Ferreira et al. 1994; Leser et al. 1989; Prasanth et al. 2003; Verheijen et al. 1986). MIGs appear to be structurally analogous to IGCs (Leser et al. 1989; Thiry 1993; Thiry 1995a). As mitosis progresses from anaphase to early telophase, the MIGs increase in number and size. During mid- to late-telophase and after re-formation of the nuclear envelope/lamina, pre-mRNA splicing factors enter daughter nuclei and, concomitantly, their localization in MIGs decreases, demonstrating that these factors are recycled from the cytoplasmic MIGs into daughter nuclei (Prasanth et al. 2003). Live-cell studies have indicated that the majority of these factors enter daughter nuclei within 10 minutes (Prasanth et al. 2003). Although MIGs have been proposed to be the mitotic equivalent of nuclear speckles (Ferreira et al. 1994; Leser et al. 1989; Thiry 1995a; Thiry 1995b), their function in mitotic cells is unclear. In telophase cells, some MIGs were found to be in close proximity to the newly formed nuclear envelope (Prasanth et al. 2003; Thiry 1995a). This close proximity of MIGs to the nuclear periphery and the disappearance of MIGs in late telophase cells with the concomitant appearance of IGCs in daughter nuclei have suggested that the MIGs might be directly transported into the nuclei (Leser et al. 1989; Thiry 1995a). However, colocalization of SF2/ ASF and a hyperphosphorylated form of RNAPII LS in MIGs of late telophase cells has suggested that this may not be the case. For example, SF2/ASF and other pre-mRNA processing factors were shown to enter daughter nuclei while a subpopulation of SC35 and RNAPII LS remained in MIGs until G1, demonstrating that various components of MIGs are differentially released for subsequent entry into daughter nuclei (Prasanth et al. 2003). Further support for differential release of factors from MIGs comes from an earlier study that reported the nuclear import of snRNPs while cytoplasmic MIGs were still labeled with anti-SR protein and anti-SC35 antibodies (Ferreira et al. 1994). Based on these findings, it was suggested that MIGs may play important roles

Nuclear Speckles

either in the modification of the components of the splicing machinery before their nuclear entry, or as enriched populations of these factors, allowing for protein-protein interactions to occur between subsets of proteins before their nuclear entry (Prasanth et al. 2003). Interestingly, splicing factors were shown to be competent for pre-mRNA splicing immediately upon entry into daughter nuclei (Prasanth et al. 2003), supporting the possibility that MIGs may be responsible for splicing factor modification, allowing for immediate targeting of modified ( phosphorylated) pre-mRNA processing complexes to transcription sites in telophase nuclei. Because daughter nuclei late in telophase have not yet assembled nuclear speckles, cytoplasmic MIGs are likely to function as their counterparts to provide competent premRNA splicing factors to the initial sites of transcription in newly formed nuclei (Prasanth et al. 2003). Perhaps splicing factors are released from MIGs via hyperphosphorylation, as has been shown for their release from nuclear speckles in interphase nuclei. SPECKLE BIOGENESIS

Nuclear speckles are one of the most prominent nuclear compartments, and their study has served as a paradigm for understanding the biogenesis of nuclear bodies. Most evidence points to the fact that nuclear speckles form through a process of self-assembly (reviewed in Misteli 2001b) whereby transient macromolecular interactions form the basis of speckle morphogenesis. Under steady-state conditions, the respective rates of association and disassociation of individual speckle components will define their exchange rates and the sizes of their bound and soluble pools in the nucleus. Regulatory mechanisms can influence these association and/or disassociation rates, thereby changing the fraction of bound and soluble speckle components in response to specific cellular signals. In this view, the entry of splicing factors into late-telophase nuclei results in an association of a subset of these factors with initial transcription/pre-mRNA processing sites (Prasanth et al. 2003). As the population of factors

increases, there is an increased probability of protein-protein interactions among those factors not engaged in transcription/pre-mRNA processing, resulting in the formation of nuclear speckles. These initial speckles appear to form predominantly in nucleoplasmic regions that are devoid of chromosome territories and/or other nuclear organelles. They may initiate either at random locations, or in the vicinity of genes that are transcribed at high levels during the telophase/G1 transition. Interestingly, in this regard Brown et al. (Brown et al. 2008) examined the position of various erythroid genes in erythroblasts and found that the majority of associations between erythroid genes occurred at a nuclear speckle. Interestingly, the associations were predominantly observed in regard to active genes. Based on their findings, the authors proposed that active genes can be brought into close proximity by the nucleation of splicing factors into nuclear speckles. The size and shape of interphase speckles is a reflection of the steady-state dynamics of protein constituents that are both arriving at and leaving from these structures (Kruhlak et al. 2000; Phair et al. 2000). Although photobleaching analyses have indicated rapid recovery kinetics of splicing factors in speckles, consistent with a diffusion-based process (Kruhlak et al. 2000; Phair et al. 2000), the relative size of speckles remains constant throughout interphase. In fact, the incubation of permeabilized cells with a nuclear extract containing an ATPregenerating system maintains transcriptional activity and does not result in a loss of speckles (Misteli et al. 1996), nor does simple treatment of unfixed cells with detergent (Spector et al. 1992). The observed basal exchange rate may be directly related to the maintenance of speckles, rather than indirectly related to an involvement in transcriptional/pre-mRNA processing events. In addition, the irregular shape of individual nuclear speckles in interphase nuclei may result from a nonuniform release and/or delivery of factors, related to the location of active genes in their vicinity (Misteli et al. 1997a). Consistent with this possibility, either upon inhibition of RNAPII transcription by a-amanitin

D.L. Spector and A.I. Lamond

(Spector et al. 1993), or inhibition of pre-mRNA splicing using an antisense approach (O’Keefe et al. 1994), speckles tend to round-up, suggesting a uniform exchange rate of factors in all directions. This exchange rate of speckle factors may be regulated through phosphorylation/dephosphorylation events. For example, phosphorylation of the RS domain of SR splicing factors has been shown to be necessary for recruitment of SR proteins from nuclear speckles to sites of transcription/pre-mRNA processing (Misteli et al. 1998) and for their association with the forming spliceosome (Mermoud et al. 1994). Several kinases (i.e., Clk/STY [Colwill et al. 1996; Sacco-Bubulya et al. 2002] and hPRP4 [Kojima et al. 2001]) involved in this phosphorylation, as well as a kinase proposed to be involved in the phosphorylation of the carboxyterminal domain of RNAPII LS in vitro (Ko et al. 2001), have been localized to nuclear speckles, leaving open the possibility that phosphorylation/dephosphorylation plays a role in determining the basal rate of factor exchange. However, in addition to the basal activities, an additional level of control can be exerted by modulating phosphorylation events. For example, the rapid induction either of a gene (Huang et al. 1996), or group of genes such as during viral infection (Bridge et al. 1995; Jime´nezGarcı´a et al. 1993), can result in an increased outward flow of factors from speckles. An extreme example of this can be observed upon overexpression of Clk/STY kinase, or addition of SRPK1 kinase to permeabilized cells (Gui et al. 1994a; Gui et al. 1994b), which results in the complete redistribution of splicing factors from speckles to the diffuse nuclear pool (Colwill et al. 1996; Sacco-Bubulya et al. 2002). Interestingly, expression of a mutant form of Clk/STY that lacks its catalytic activity resulted in an increased accumulation of factors in highly concentrated foci on the periphery of speckles, possibly a reflection of their inability to be released (Sacco-Bubulya et al. 2002). Consistent with this observation, the addition of kinase inhibitors to cells resulted in an inhibition of the dynamic movements on the periphery of speckles (Misteli et al. 1997a). Also,

protein phosphatase 1 (PP1) inhibitors resulted in enlarged irregularly shaped speckles with less well defined edges, probably resulting from the inability of factors to be released from PFs, on the periphery of IGCs, also consistent with a modulating effect on the exchange rate (Misteli et al. 1996). In summary, a basal exchange rate of factors, coupled with a mechanism to modulate this rate (that is, providing a stimulus-induced burst), ensures that the needed factors, in the correct phosphorylation state, are available to pre-mRNA transcripts at the sites of transcription. In addition, such a mechanism ensures that a significant population of factors, which are not functionally needed, are sequestered out of the soluble nuclear pool, in this case in nuclear speckles, representing a basic mechanism for the organization of nonmembranebound nuclear organelles. CONCLUSIONS

Nuclear speckles are organelles located in the interchromatin nuclear space and are among the most widely studied nuclear domains. They are best known for accumulating high local concentrations of snRNPs and other non-snRNP protein splicing factors. However, the presence in speckles of many other factors involved in mRNA production by RNA polymerase II further supports their intimate relationship with gene expression. Although most speckles apparently do not contain DNA in an analogous way to the rRNA gene repeats within nucleoli, nonetheless highly expressed genes can be found associated with speckles, consistent with an important role for speckles in coordinating the supply and/or recycling of pre-mRNA processing and transcription factors. The analysis of nuclear speckles has helped to establish some key paradigms and principles for the dynamic assembly of membrane-free organelles in the nucleus. We anticipate that future work will refine our understanding of speckle composition and of mechanisms involved in targeting proteins to speckles and regulating their formation. It is likely that building a detailed model of gene expression in vivo will require further

Nuclear Speckles

characterization of nuclear speckles to define how the complex events required for transcription and RNA processing are efficiently coordinated within the nucleus.

ACKNOWLEDGMENTS

We thank members of the Spector and Lamond groups for helpful comments. Angus Lamond is a Wellcome Trust Principal Research Fellow. David L. Spector is funded by NIGMS/NIH 42694, NIH/NCI 5PO1CA013106-38, and NIH/ EY 18244.

REFERENCES Beck JS. 1961. Variations in the morphological patterns of “autoimmune” nuclear fluorescence. Lancet 1: 1203– 1205. Bernard D, Prasanth KV, Tripathi V, Colasse S, Nakamura T, Xuan Z, Zhang MQ, Sedal F, Jourdren L, Coulpier F, et al. 2010. A long nuclear retained no-coding RNA regulates synaptogenesis by modulating gene expression. EMBO J doi:10.1038/emboj.2010.199. Boronenkov IV, Loijens JC, Umeda M, Anderson RA. 1998. Phosphoinositide signaling pathways in nuclei are associated with nuclear speckles containing pre-mRNA processing factors. Mol Biol Cell 9: 3547– 3560. Brede G, Solheim J, Prydz H. 2002. PSKH1, a novel splice factor compartment-associated serine kinase. Nucleic Acids Res 30: 5301–5309. Bregman DB, Du L, van der Zee S, Warren SL. 1995. Transcription-dependent redistribution of the large subunit of RNA polymerase II to discrete nuclear domains. J Cell Biol 129: 287 –298. Bridge E, Xia D-X, Carmo-Fonseca M, Cardinali B, Lamond AI, Pettersson U. 1995. Dynamic organization of splicing factors in adenovirus-infected cells. J Virol 69: 281 –290. Brown JM, Green J, das Neves RP, Wallace HA, Smith AJ, Hughes J, Gray N, Taylor S, Wood WG, Higgs DR, et al. 2008. Association between active genes occurs at nuclear speckles and is modulated by chromatin environment. J Cell Biol 182: 1083–1097. Caceres JF, Misteli T, Screaton GR, Spector DL, Krainer AR. 1997. Role of the modular domains of SR proteins in subnuclear localization and alternative splicing specificity. J Cell Biol 138: 225 –238. Carter KC, Taneja KL, Lawrence JB. 1991. Discrete nuclear domains of poly(A) RNA and their relationship to the functional organization of the nucleus. J Cell Biol 115: 1191– 1202. Cmarko D, Verschure PJ, Martin TE, Dahmus ME, Krause S, Fu XD, van Driel R, Fakan S. 1999. Ultrastructural analysis of transcription and splicing in the cell nucleus after bromo-UTP microinjection. Mol Biol Cell 10: 211–223.

Colwill K, Pawson T, Andrews B, Prasad J, Manley JL, Bell JC, Duncan PI. 1996. The Clk/Sty protein kinase phosphorylates splicing factors and regulates their intranuclear distribution. EMBO J 15: 265 –275. Dostie J, Lejbkowicz F, Sonenberg N. 2000. Nuclear eukaryotic initiation factor 4E (eIF4E) colocalizes with splicing factors in speckles. J Cell Biol 148: 239– 247. Doyle O, Corden JL, Murphy C, Gall JG. 2002. The distribution of RNA polymerase II largest subunit (RPB1) in the Xenopus germinal vesicle. J Struct Biol 140: 154 –166. Eilbracht J, Schmidt-Zachmann MS. 2001. Identification of a sequence element directing a protein to nuclear speckles. Proc Natl Acad Sci 98: 3849–3854. Fakan S. 1994. Perichromatin fibrils are in situ forms of nascent transcripts. Trends Cell Biol 4: 86–90. Fakan S, Bernhard W. 1971. Localisation of rapidly and slowly labelled nuclear RNA as visualized by high resolution autoradiography. Exp Cell Res 67: 129 –141. Fakan S, Nobis P. 1978. Ultrastructural localization of transcription sites and of RNA distribution during the cell cycle of synchronized CHO cells. Exp Cell Res 113: 327 –337. Ferreira JA, Carmo-Fonseca M, Lamond AI. 1994. Differential interaction of splicing snRNPs with coiled bodies and interchromatin granules during mitosis and assembly of daughter cell nuclei. J Cell Biol 126: 11– 23. Fox AH, Lam YW, Leung AK, Lyon CE, Andersen J, Mann M, Lamond AI. 2002. Paraspeckles: A novel nuclear domain. Curr Biol 12: 13– 25. Fox AH, Lamond AI. 2010. Paraspeckles. Cold Spring Harb Perspect Biol doi:10.1101/cshperspect.a000687. Fu X-D. 1995. The superfamily of arginine/serine-rich splicing factors. RNA 1: 663– 680. Gall JG, Bellini M, Wu Z, Murphy C. 1999. Assembly of the nuclear transcription and processing machinery: Cajal bodies (coiled bodies) and transcriptosomes. Mol Biol Cell 10: 4385– 4402. Grande MA, van der Kraan I, de Jong L, van Driel R. 1997. Nuclear distribution of transcription factors in relation to sites of transcription and RNA polymerase II. J Cell Sci 110: 1781– 1791. Gui JF, Lane WS, Fu X-D. 1994a. A serine kinase regulates intracellular localization of splicing factors in the cell cycle. Nature 369: 678 –682. Gui JF, Tronchere H, Chandler SD, Fu X-D. 1994b. Purification and characterization of a kinase specific for the serine- and arginine-rich pre-mRNA splicing factors. Proc Natl Acad Sci 91: 10824–10828. Hall LL, Smith KP, Byron M, Lawrence JB. 2006. Molecular anatomy of a speckle. Anat Rec A Discov Mol Cell Evol Biol 288: 664–675. Hedley ML, Amrein H, Maniatis T. 1995. An amino acid sequence motif sufficient for subnuclear localization of an arginine/serine rich splicing factor. Proc Natl Acad Sci 92: 11524–11528. Herrmann CH, Mancini MA. 2001. The Cdk9 and cyclin T subunits of TAK/P-TEFb localize to splicing factor-rich nuclear speckle regions. J Cell Sci 114: 1491– 1503. Hock R, Wilde F, Scheer U, Bustin M. 1998. Dynamic relocation of chromosomal protein HMG-17 in the nucleus

D.L. Spector and A.I. Lamond is dependent on transcriptional activity. EMBO J 17: 6992– 7001. Huang S, Spector DL. 1991. Nascent pre-mRNA transcripts are associated with nuclear regions enriched in splicing factors. Genes Dev 5: 2288– 2302. Huang S, Spector DL. 1996. Intron-dependent recruitment of pre-mRNA splicing factors to sites of transcription. J Cell Biol 131: 719 –732. Huang S, Deerinck MH, Ellisman MH, Spector DL. 1994. In vivo analysis of the stability and transport of nuclear poly(A)þ RNA. J Cell Biol 126: 877 –899. Hutchinson JN, Ensminger AW, Clemson CM, Lynch CR, Lawrence JB, Chess A. 2007. A screen for nuclear transcripts identifies two linked noncoding RNAs associated with SC35 splicing domains. BMC Genomics 8: 39. Jagatheesan G, Thanumalayan S, Muralikrishna B, Rangaraj N, Karande AA, Parnaik VK. 1999. Colocalization of intranuclear lamin foci with RNA splicing factors. J Cell Sci 112: 4651–4661. Jagiello I, Van Eynde A, Vulsteke V, Beullens M, Boudrez A, Keppens S, Stalmans W, Bollen M. 2000. Nuclear and subnuclear targeting sequences of the protein phosphatase-1 regulator NIPP1. J Cell Sci 21: 3761–3768. Jime´nez-Garcı´a LF, Spector DL. 1993. In vivo evidence that transcription and splicing are coordinated by a recruiting mechanism. Cell 73: 47– 59. Johnson C, Primorac D, McKinstry M, McNeil J, Rowe D, Lawrence JB. 2000. Tracking COL1A1 RNA in osteogenesis imperfecta. splice-defective transcripts initiate transport from the gene but are retained within the SC35 domain. J Cell Biol 150: 417 –432. Kimura H, Sugaya K, Cook PR. 2002. The transcription cycle of RNA polymerase II in living cells. J Cell Biol 159: 777– 782. Ko TK, Kelly E, Pines J. 2001. CrkRS: A novel conserved Cdc2-related protein kinase that colocalises with SC35 speckles. J Cell Sci 114: 2591– 2603. Kojima T, Zama T, Wada K, Onogi H, Hagiwara M. 2001. Cloning of human PRP4 reveals interaction with Clk1. J Biol Chem 276: 32247–32256. Krause S, Fakan S, Weis K, Wahle E. 1994. Immunodetection of Poly(A) Binding Protein II in the Cell Nucleus. Exp Cell Res 214: 75–82. Kruhlak MJ, Lever MA, Fischle W, Verdin E, Bazett-Jones DP, Hendzel MJ. 2000. Reduced mobility of the alternate splicing factor (ASF) through the nucleoplasm and steady state speckle compartments. J Cell Biol 150: 41–51. Lafarga M, Casafont I, Bengoechea R, Tapia O, Berciano MT. 2009. Cajal’s contribution to the knowledge of the neuronal cell nucleus.. Chromosoma 118: 437–443. Lamond AI, Earnshaw WC. 1998. Structure and function in the nucleus. Science 280: 547– 553. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605 –612. Larsson SH, Charlieu JP, Miyagawa K, Engelkamp D, Rassoulzadegan M, Ross A, Cuzin F, van Heyningen V, Hastie ND. 1995. Subnuclear localization of WT1 in splicing or transcription factor domains is regulated by alternative splicing. Cell 81: 391– 401. Lerner EA, Lerner MR, Janeway CA, Steitz JA. 1981. Monoclonal antibodies to nucleic acid-containing cellular

constituents: Probes for molecular biology and autoimmune disease. Proc Natl Acad Sci 78: 2737–2741. Leser GP, Fakan S, Martin TE. 1989. Ultrastructural distribution of ribonucleoprotein complexes duirng mitosis. snRNP antigens are contained in mitotic granule clusters. Eur J Cell Biol 50: 376–389. Li H, Bingham PM. 1991. Arginine/Serine-rich domains of the su(wa) and tra RNA processing regulators target proteins to a subnuclear compartment implicated in splicing. Cell 67: 335– 342. Li Q, Imataka H, Morino S, Rogers GW Jr, Richter-Cook NJ, Merrick WC, Sonenberg N. 1999. Eukaryotic translation initiation factor 4AIII (eIF4AIII) is functionally distinct from eIF4AI and eIF4AII. Mol Cell Biol 19: 7336– 7346. Matera AG, Ward DC. 1993. Nucleoplasmic organization of small nuclear ribonucleoproteins in cultured human cells. J Cell Biol 121: 715 –727. Melcak I, Cermanova S, Jirsova K, Koberna K, Malinsky J, Raska I. 2000. Nuclear pre-mRNA compartmentalization: Trafficking of released transcripts to splicing factor reservoirs. Mol Biol Cell 11: 497– 510. Mermoud JE, Cohen PTW, Lamond AI. 1994. Regulation of mammalian spliceosome assembly by a protein phosphorylation mechanism. EMBO J 13: 5679– 5688. Mintz PJ, Patterson SD, Neuwald AF, Spahr CS, Spector DL. 1999. Purification and biochemical characterization of interchromatin granule clusters. Embo J 18: 4308–4320. Misteli T. 2001a. Protein dynamics: Implications for nuclear architecture and gene expression. Science 291: 843– 847. Misteli T. 2001b. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Misteli T, Spector DL. 1996. Serine/threonine phosphatase 1 modulates the subnuclear distribution of pre-mRNA splicing factors. Mol Biol Cell 7: 1559– 1572. Misteli T, Spector DL. 1997b. Protein phosphorylation and the nuclear organization of pre-mRNA splicing. Trends Cell Biol 7: 135– 138. Misteli T, Ca´ceres JF, Spector DL. 1997a. The dynamics of a pre-mRNA splicing factor in living cells. Nature 387: 523 –527. Misteli T, Caceres JF, Clement JQ, Krainer AR, Wilkinson MF, Spector DL. 1998. Serine phosphorylation of SR proteins is required for their recruitment to sites of transcription in vivo. J Cell Biol 143: 297– 307. Moen PT Jr, Johnson CV, Byron M, Shopland LS, de la Serna IL, Imbalzano AN, Lawrence JB. 2004. Repositioning of muscle-specific genes relative to the periphery of SC-35 domains during skeletal myogenesis. Mol Biol Cell 15: 197 –206. Monneron A, Bernhard W. 1969. Fine structural organization of the interphase nucleus in some mammalian cells. J Ultrastruct Res 27: 266– 288. Mortillaro MJ, Blencowe BJ, Wei X, Nakayasu H, Du L, Warren SL, Sharp PA, Berezny R. 1996. A hyperphosphorylated form of the large subunit of RNA polymerase II is associated with splicing complexes and the nuclear matrix. Proc Natl Acad Sci 93: 8253–8257. Nakayasu H, Ueda K. 1984. Small nuclear RNA-protein complex anchors on the actin filaments in bovine lymphocyte nuclear matrix. Cell Struct Funct 9: 317 –325.

Nuclear Speckles Nayler O, Hartmann AM, Stamm S. 2000. The ER repeat protein YT521-B localizes to a novel subnuclear compartment. J Cell Biol 150: 949 –962. O’Keefe RT, Mayeda A, Sadowski CL, Krainer AR, Spector DL. 1994. Disruption of pre-mRNA splicing in-vivo results in reorganization of splicing factors. J Cell Biol 124: 249– 260. Pendergrast PS, Wang C, Hernandez N, Huang S. 2002. FBI-1 Can Stimulate HIV-1 Tat Activity and Is Targeted to a Novel Subnuclear Domain that Includes the Tat-P-TEFb-containing Nuclear Speckles. Mol Biol Cell 13: 915 –929. Perraud M, Gioud M, Monier JC. 1979. Intranuclear structures recognized by autoantibodies against ribonucleoproteins: Study on monkey kidney cells in culture using immunofluorescent techniques and immunoelectron microscopy [in French]. Ann Immunol 130: 635 –647. Pessler F, Pendergrast PS, Hernandez N. 1997. Purification and characterization of FBI-1, a cellular factor that binds to the human immunodeficiency virus type 1 inducer of short transcripts. Mol Cell Biol 17: 3786–3798. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604–609. Potashkin JA, Derby RJ, Spector DL. 1990. Differential distribution of factors involved in pre-mRNA processing in the yeast cell nucleus. Mol Cell Biochem 10: 3524– 3534. Prasanth KV, Sacco-Bubulya P, Prasanth SG, Spector DL. 2003. Sequential entry of components of gene expression machinery into daughter nuclei. Molec Biol Cell 14: 1043– 1057. Price DH. 2000. P-TEFb, a cyclin-dependent kinase controlling elongation by RNA polymerase II. Mol Cell Biol 20: 2629– 2634. Ramo´n y Cajal S. 1910. El nucleo de las celulas piramidales del cerebro humano y de algunos mamiferos. Trab Lab Invest Biol 8: 27–62. Rappsilber J, Ryder U, Lamond AI, Mann M. 2002. Largescale proteomic analysis of the human spliceosome. Genome Res 12: 1231– 1245. Reuter R, Appel B, Rinke J, Lu¨hrmann R. 1985. Localization and structure of snRNPs during mitosis. Immunofluorescent and biochemical studies. Exp Cell Res 159: 63– 79. Sacco-Bubulya P, Spector DL. 2002. Disassembly of interchromatin granule clusters alters the coordination of transcription and pre-mRNA splicing. J Cell Biol 156: 425–436. Saitoh N, Spahr CS, Patterson SD, Bubulya P, Neuwald AF, Spector DL. 2004. Proteomic analysis of interchromatin granule clusters. Mol Biol Cell 15: 3876–3890. Salichs E, Ledda A, Mularoni L, Alba MM, de la Luna S. 2009. Genome-wide analysis of histidine repeats reveals their role in the localization of human proteins to the nuclear speckles compartment. PLoS Genet 5: e1000397. Schul W, van Driel R, de Jong L. 1998. A subset of poly(A) polymerase is concentrated at sites of RNA synthesis and is associated with domains enriched in splicing factors and poly(A) RNA. Exp Cell Res 238: 1 –12. Segalat L, Lepesant JA. 1992. Spatial distribution of the Sm antigen in Drosophila early embryos. Biol Cell 75: 181– 185.

Sharma A, Takata H, Shibahara K, Bubulya A, Bubulya PA. 2010. Son is essential for nuclear speckle organization and cell cycle progression. Mol Biol Cell 21: 650– 663. Shopland LS, Johnson CV, Lawrence JB. 2002. Evidence that all SC-35 domains contain mRNAs and that transcripts can be structurally constrained within these domains. J Struct Biol 140: 131– 139. Shopland LS, Johnson CV, Byron M, McNeil J, Lawrence JB. 2003. Clustering of multiple specific genes and generich R-bands around SC-35 domains: Evidence for local euchromatic neighborhoods. J Cell Biol 162: 981– 990. Smith KP, Moen PT, Wydner KL, Coleman JR, Lawrence JB. 1999. Processing of endogenous pre-mRNAs in association with SC-35 domains is gene specific. J Cell Biol 144: 617–629. Spector DL. 1993. Macromolecular domains within the cell nucleus. Annu Rev Cell Biol 9: 265– 315. Spector DL. 2001. Nuclear bodies. J Cell Sci 114: 2891– 2893. Spector DL. 2006. SnapShot: Cellular bodies. Cell 127: 1071. Spector DL, Smith HC. 1986. Redistribution of U-snRNPs during mitosis. Exp Cell Res 163: 87–94. Spector DL, Fu X-D, Maniatis T. 1991. Associations between distinct pre-mRNA splicing components and the cell nucleus. EMBO J 10: 3467–3481. Spector DL, Lark G, Huang S. 1992. Differences in snRNP localization between transformed and nontransformed cells. Mol Biol Cell 3: 555– 569. Spector DL, O’Keefe RT, Jime´nez-Garcı´a LF. 1993. Dynamics of transcription and pre-mRNA splicing within the mammalian cell nucleus. Cold Spring Harb Symp Quant Biol 58: 799 –805. Spector DL, Schrier WH, Busch H. 1983. Immunoelectron microscopic localization of snRNPs. Biol Cell 49: 1 –10. Swift H. 1959. Studies on nuclear fine structure. Brookhaven Symp Biol 12: 134– 152. Thiry M. 1993. Differential location of nucleic acids within interchromatin granule clusters. Eur J Cell Biol 62: 259 –269. Thiry M. 1995a. Behavior of interchromatin granules during the cell cycle. European J Cell Biol 68: 14–24. Thiry M. 1995b. The interchromatin granules. Histol Histopathol 10: 1035– 1045. Trinkle-Mulcahy L, Sleeman JE, Lamond AI. 2001. Dynamic targeting of protein phosphatase 1 within the nuclei of living mammalian cells. J Cell Sci 114: 4219– 4228. Trinkle-Mulcahy L, Ajuh P, Prescott A, Claverie-Martin F, Cohen S, Lamond AI, Cohen P. 1999. Nuclear organisation of NIPP1, a regulatory subunit of protein phosphatase 1 that associates with pre-mRNA splicing factors. J Cell Sci 112: 157 –168. Tripathi V, Ellis J, Shen Z, Song D, Freier SM, Bennett CF, Sharma A, Bubulya PA, Blencowe BJ, Prasanth SG, et al. 2010. Nuclear-retained non-coding RNA regulates alternative splicing by modulating SR splicing factor phosphorylation. Mol Cell advance online publication doi:101016/jmolcel201008011. Vecerova J, Koberna K, Malinsky J, Soutoglou E, Sullivan T, Stewart CL, Raska I, Misteli T. 2004. Formation of nuclear

D.L. Spector and A.I. Lamond splicing factor compartments is independent of lamins A/C. Mol Biol Cell 15: 4904– 4910. Verheijen R, Kuijpers H, Vooijs P, Van Venrooij W, Ramaekers F. 1986. Distribution of the 70K U1 RNA-associated protein during interphase and mitosis. Correlation with other U RNP particles and proteins of the nuclear matrix. J Cell Sci 86: 173– 190. Visa N, Puvion-Dutilleul F, Harper F, Bachellerie J-P, Puvion E. 1993. Intranuclear distribution of poly A RNA determined by electron microscope in situ hybridization. Exp Cell Res 208: 19– 34. Wansink DG, Schul W, van der Kraan I, van Steensel B, van Driel R, de Jong L. 1993. Fluorescent labeling of nascent RNA reveals transcription by RNA polymerase II in domains scattered throughout the nucleus. J Cell Biol 122: 283– 293. Wei X, Somanathan S, Samarabandu J, Berezney R. 1999. Three-dimensional visualization of transcription sites and their association with splicing factor-rich nuclear speckles. J Cell Biol 146: 543– 558. Wilusz JE, Sunwoo H, Spector DL. 2009. Long noncoding RNAs: Functional surprises from the RNA world. Genes Dev 23: 1494–1504.

Xing Y, Johnson CV, Dobner PR, Lawrence JB. 1993. Higher level organization of individual gene transcription and RNA splicing. Science 259: 1326–1330. Xing Y, Johnson CV, Moen PT, McNeil JA, Lawrence JB. 1995. Nonrandom gene organization: Structural arrangements of specific pre-mRNA transcription and splicing with SC-35 domains. J Cell Biol 131: 1635–1647. Zeng C, Kim E, Warren SL, Berget SM. 1997. Dynamic relocation of transcription and splicing factors dependent upon transcriptional activity. EMBO J 16: 1401– 1412. Zhao R, Bodnar MS, Spector DL. 2009. Nuclear neighborhoods and gene expression. Curr Opin Genet Dev 19: 172 –179. Zhao K, Wang W, Rando OJ, Xue Y, Swiderek K, Kuo A, Crabtree GR. 1998. Rapid and phosphoinositol-dependent binding of the SWI/SNF-like BAF complex to chromatin after T lymphocyte receptor signaling. Cell 95: 625 –636. Zhou Z, Licklider LJ, Gygi SP, Reed R. 2002. Comprehensive proteomic analysis of the human spliceosome. Nature 419: 182–185.

The Cajal Body and Histone Locus Body Zehra Nizami, Svetlana Deryusheva, and Joseph G. Gall Department of Embryology, Carnegie Institution for Science, Baltimore, Maryland 21218 Correspondence: [email protected]

The Cajal body (CB) is a nuclear organelle present in all eukaryotes that have been carefully studied. It is identified by the signature protein coilin and by CB-specific RNAs (scaRNAs). CBs contain high concentrations of splicing small nuclear ribonucleoproteins (snRNPs) and other RNA processing factors, suggesting that they are sites for assembly and/or posttranscriptional modification of the splicing machinery of the nucleus. The histone locus body (HLB) contains factors required for processing histone pre-mRNAs. As its name implies, the HLB is associated with the genes that code for histones, suggesting that it may function to concentrate processing factors at their site of action. CBs and HLBs are present throughout the interphase of the cell cycle, but disappear during mitosis. The biogenesis of CBs shows the features of a self-organizing structure.

he CB and the HLB can be considered together for several reasons. First, the canonical marker for the CB, coilin, occurs at high concentration in some HLBs. Second, CBs and HLBs are often physically associated, suggesting some type of interaction between the two. Finally, the very large CBs of the amphibian oocyte, long a favorite for studies of CB function, share properties of both HLBs and CBs.

T

CAJAL BODIES

As the current name suggests, the CB was first described by Ramon y Cajal, the great Spanish neuroanatomist, who in 1906 shared the Nobel Prize with Camillo Golgi for studies on the cellular architecture of the nervous system. Using a silver impregnation technique, Cajal found a small round body within the nuclei of various nerve cells, which he called the accessory body

(cuerpo accessorio) (Cajal 1903; Cajal 1910). His remarkable morphological studies also included observations on splicing speckles, nucleoli, and the nuclear envelope (Lafarga et al. 2009). Bodies that we now know are related to Cajal’s cuerpo accessorio were discovered independently in organisms as diverse as mammals, amphibians, insects, and plants. They were given equally diverse names—coiled bodies in mouse, rat, and human cells (Monneron and Bernhard 1969), Binnenko¨rper or endobodies in insects (Bier et al. 1967), and nucleolus associated bodies in plants (Chamberland and Lafontaine 1993). Order was brought to this somewhat chaotic field with the discovery of a protein named coilin, after its discovery in the coiled bodies of HeLa cells (Andrade et al. 1991; Raska et al. 1991). Antibodies against coilin turned out to be good markers for coiled bodies in vertebrate cells and

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000653 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000653

Z. Nizami, S. Deryusheva, and J.G. Gall

even cross-reacted with the nucleolus associated bodies of the pea, Pisum sativum (Beven et al. 1995). It thus became clear that homologous nuclear organelles existed in a wide range of eukaryotes (Figs. 1 and 2). To recognize this commonality and to regularize the terminology, the name Cajal body was adopted as a general term for a nuclear body that contains coilin (Gall et al. 1999).

Despite its usefulness as a marker for the CB, relatively little is known about coilin as a protein, particularly what biochemical function(s) it might play in the CB. Coilin binds to the survival motor neuron (SMN) protein and to various Sm and Lsm proteins, suggesting that it may function in assembly or modification of snRNPs (Hebert and Matera 2000; Hebert et al. 2001; Xu et al. 2005). There is strong genetic evidence from the mouse, Arabidopsis, and Drosophila that coilin is required for formation of CBs. Knockout of the coilin gene in the mouse leads to a semi-lethal phenotype. Some homozygous individuals die as embryos, and those adults that do survive have significant fertility and fecundity defects (Walker et al. 2009). Cultured cells derived from the knockouts do not have typical CBs. Instead, they show three types of “residual” bodies, each of which contains a subset of CB components (Tucker et al. 2001; Ja´dy et al. 2003). In Arabidopsis the no cajal body 1 (ncb-1) mutant involves a single base substitution in the coilin gene, although it is not certain that the mutant is completely null for coilin (Collier et al. 2006). Homozygous ncb-1 plants are fully viable, but CBs are not detectable with antibodies against other CB components (U2B00 and fibrillarin) or by electron microscopy. In Drosophila two independent null mutants for coilin are fully

Coilin

After its discovery in HeLa cells, coilin quickly became the signature marker for CBs in mammalian cells. Human and mouse coilin are similar in size, MW ¼ 62.6 and 62.3 kDa respectively, and their overall sequence identity is high (Andrade et al. 1991; Tucker et al. 2000). Xenopus coilin is slightly smaller, 59.6 kDa, and its sequence diverges considerably from that of the two mammalian proteins (Tuma et al. 1993). Outside the vertebrates it becomes increasingly difficult to recognize coilin by sequence comparison. What are clearly orthologs of coilin have been described from Arabidopsis (Collier et al. 2006) and Drosophila (Liu et al. 2009), but so far coilin has not been identified in Caenorhabditis, Saccharomyces, or other important nonvertebrate model organisms.

A

B

Figure 1. CBs in a HeLa cell nucleus. (A) DIC image showing four prominent nucleoli and three faintly

detectable CBs (arrowheads). (B) The same nucleus immunostained with antibodies against coilin (green) and fibrillarin (red). The three CBs in this nucleus appear yellow because they stain with both antibodies. The nucleoli stain only for fibrillarin. Fibrillarin is the methyltransferase for 20 -O-methylation of rRNA in the nucleoli and snRNAs in the CBs. Modified from Gall (2003) with permission from Nature Reviews. Bar ¼ 10 mm.

Cajal Body and Histone Locus Body

A

B

Figure 2. A cell from a protoplast of Arabidopsis after

transfection with a construct of U2B00 -mRFP, a component of the U2 snRNP that is enriched in the CB. (A) Fluorescence image showing two brightly labeled CBs in the nucleus. Each is associated with a large, unstained nucleolus. (B) DIC image of the same cells. The nucleoli are now visible (arrowheads) but the CBs are not. Bar ¼ 10 mm. Image kindly provided by Zdravko Lorkovic, Medical University of Vienna, Austria.

viable as homozygotes (Beumer et al. 2008; Liu et al. 2009). Cells from coilin-null flies lack CBs detectable by immunostaining or in situ hybridization for several typical CB components. Thus, in the three organisms in which coilin mutations have been studied, coilin is required for normal CB formation, but neither coilin nor a typical CB is essential for viability. Despite the long-standing use of coilin as a unique marker for CBs, and the critical role that coilin plays in maintaining structural integrity of CBs, we now know that coilin can also occur in HLBs, as discussed later in this article. Small Nuclear Ribonucleoproteins (SnRNPs)

As soon as CBs were identified by immunostaining with antibodies against coilin, it became easy to use other antibodies and in situ hybridization probes to make a catalog of common CB components. It was quickly realized that CBs contain a variety of proteins and RNAs involved in RNA processing, particularly the splicing snRNAs (U1, U2, U4, U5, and U6) and their associated proteins (Lamond and CarmoFonseca 1993; Matera and Ward 1993; Spector 1993). Because splicing itself does not occur in CBs, it was suggested that CBs play some role in the assembly or modification of the splicing snRNPs before their recruitment to the chromosomes. The biogenesis of splicing snRNPs

is a complex process involving both nuclear and cytoplasmic steps (Will and Lu¨hrmann 2001; Patel and Bellini 2008). In brief, transcription of the snRNAs occurs in the nucleus followed by their export to the cytoplasm. In the cytoplasm the monomethyl guanosine cap at the 50 end becomes trimethylated, and each snRNA is assembled into a complex with seven conserved Sm proteins. Finally, the assembled snRNPs are imported back into the nucleus. Because snRNAs in the CB are associated with snRNP proteins and have a trimethylguanosine cap, it is assumed that they have already returned from the cytoplasm. This conclusion is reinforced by kinetic studies showing that newly imported snRNPs travel first to the CB (Sleeman and Lamond 1999) and only later appear in the speckles (interchromatin granule clusters) and finally on the chromosomes, where splicing itself takes place. Modification of specific nucleotides on the snRNAs probably occurs in the CB. It is less clear how much assembly of the splicing machinery takes place in the CB. A role for CBs has been suggested in the final steps of U2 snRNP biogenesis (Nesic et al. 2004) and a particularly compelling case has been made that assembly of the U4/U6-U5 tri-snRNP occurs in the CB (Stanek et al. 2003; Schaffert et al. 2004). Evidence has also been presented that snRNPs recycle through CBs (Stanek et al. 2008). The splicing snRNPs very probably pass from CBs to the nuclear speckles (interchromatin granule clusters) on their way to the sites of RNA synthesis and splicing on the chromosomes. However, the extent to which individual snRNPs are organized into higher complexes in the speckles is not known (Lamond and Spector 2003). A recent study on the amphibian oocyte suggests that snRNPs may be recruited to lampbrush chromosomes independently of their assembly into mature spliceosomes (Patel et al. 2007). If this is generally true for other cells, then CBs may have only a limited role in assembly of snRNPs into higher order complexes. Small CB-associated RNAs (scaRNAs)

An important advance in the understanding of CB function came with the discovery of

Z. Nizami, S. Deryusheva, and J.G. Gall

CB-specific RNAs (scaRNAs). Sca-RNAs are closely related to small nucleolar RNAs (snoRNAs) in both structure and function. Both share the so-called box C/D and box H/ ACA motifs and both are involved in the posttranscriptional modification of other RNAs. Box C/D snoRNAs act as guides for the insertion of 20 -O-methyl groups on specific ribose moieties of rRNA, whereas the box H/ACA class mediates the conversion of specific uridines to pseudouridine (Bachellerie and Cavaille 1997; Tollervey and Kiss 1997). Fibrillarin is the methyl transferase and dyskerin/NAP57/CBF5 is the pseudouridine synthase; each is complexed with three additional proteins to form the active enzyme. ScaRNAs carry out similar reactions on snRNAs. The first discovered and the best-studied scaRNA is U85 (Darzacq et al. 2002; Kiss et al. 2002). It is an unusual guide RNA that mediates two modifications: 20 O-methylation of C45 and pseudouridylation of U46 in human U5 snRNA (C46 and U47 in Drosophila). Cell fractionation and in situ hybridization experiments demonstrate that U85 scaRNA is strongly localized in CBs in both HeLa cells and Drosophila (Figs. 3A and 4B). This localization contrasts with that of its substrate, U5 snRNA, which is concentrated in CBs, but like other snRNAs is widely distributed throughout the nucleus. The localization of U85 and other scaRNAs in the CB differs from the localization of most box C/D and box H/ ACA guide RNAs, which concentrate in the nucleolus. It has been shown that CB localization in vertebrate cells is dependent on a relatively short consensus sequence termed the CAB box (Richard et al. 2003). A related but somewhat different motif has been described for Drosophila scaRNAs. The CAB box of both human and Drosophila scaRNAs binds a conserved WD40-repeat protein, which is required for their CB localization (Tycowski et al. 2009). The specific localization of scaRNAs in the CB suggests that methylation and pseudouridylation of snRNAs occur in the CB, after import of the assembled snRNPs into the nucleus. This hypothesis is strongly supported by experiments in cultured cells showing that artificial substrates of scaRNAs became modified when

U5 + U85

SDMA + Lsm11

A

C

B

D

Figure 3. Malpighian tubule nuclei of Drosophila after

fluorescent in situ hybridization (A and B) or immunostaining (C and D) for CB components. (A) A nucleus from a wild type fly showing two CBs. The CBs appear yellow because they contain both U85 scaRNA (red) and U5 snRNA (green). There is a high level of U5 throughout the nucleoplasm. (B) A nucleus from a coilin-null fly hybridized as in A. A CB is no longer detectable, although U5 label in the nucleoplasm is unaffected. (C) A nucleus from a wild type fly showing a CB immunostained for symmetric dimethylarginine, a marker for snRNP proteins (green), and an HLB labeled for Lsm11, a component of the U7 snRNP (red). (D) A nucleus from a coilin-null fly labeled as in C. A CB is no longer detectable, but the HLB is apparently unaffected. Modified from Liu et al. (2009) with permission from Molecular Biology of the Cell. Bar ¼ 10 mm in A and B, 5 mm in C and D.

targeted to the CB but not when targeted to the nucleolus (Ja´dy et al. 2003). This hypothesis is also consistent with the well-known concentration of fibrillarin in the CB (Fig. 1). At the same time, it is unlikely that modification of snRNAs is limited to CBs, because coilin-null flies, which lack CBs (Fig. 3 B,D; Fig. 4C), have normal levels of scaRNAs, and all of their snRNAs are correctly modified (Deryusheva and Gall 2009). It seems probable that scaRNAs and other CB components normally exist in the nucleoplasm as macromolecular complexes that are too small to be resolved individually by conventional light microscopy. Coilin is essential

Cajal Body and Histone Locus Body

A scaRNA of unusual interest is the RNA component of telomerase, the enzyme responsible for maintaining the ends or telomeres of eukaryotic chromosomes. Telomerase RNA has been shown by in situ hybridization in CBs in human cancer cell lines (Zhu et al. 2003; Ja´dy et al. 2004), but it is at low or undetectable levels in noncancer cells. Telomerase RNA has a box H/ACA motif as well as a CAB box. As with other scaRNAs, the CAB box is required for CB localization (Theimer et al. 2007). The same WD40-repeat protein that binds other scaRNPs is part of the human telomerase holoenzyme and is required for telomere synthesis in HeLa cells (Venteicher et al. 2009).

A

B

D

C

E

F

CBs, Gems, and the SMN Protein

Figure 4. CBs and HLBs in Drosophila. (A) Each

nucleus of the ejaculatory duct of the male fly usually has a single CB, immunostained green with an antibody against coilin, and a single HLB, immunostained red with an antibody against Lsm11. Often the two bodies are close to one another or touching. Bar ¼ 5 mm. (B and C) In Drosophila and many other insects, highly polyploid nurse cells supply proteins and RNAs to the transcriptionally silent oocyte. Their nuclei display prominent CBs and HLBs, here shown after double in situ hybridization for U85 scaRNA (red) and U7 snRNA (green). (B) shows a stage 8 egg chamber of a wild type fly. C shows the same stage from a coilin-null fly. CBs are not detectable in the mutant, whereas HLBs appear unaffected. Bar ¼ 5 mm. (D–F) Nurse cell nuclei from a stage 11 egg chamber showing prominent HLBs that contain coilin. Immunostained with antibodies against Lsm11 (D, red) and coilin (E, green). The overlay in F shows colocalization of Lsm11 and coilin. Modified from Liu et al. (2009) with permission from Molecular Biology of the Cell. Bar ¼ 20 mm.

for these complexes to assemble into detectable CBs, but assembly of such bodies is not required for function, at least not for the scaRNAdependent modification of splicing snRNAs.

An unusually interesting component of CBs is the survival motor neuron protein (SMN). When the intracellular localization of SMN was first studied by immunofluorescence, the protein was seen throughout the cytoplasm and in a nuclear body similar in size to the CB and often close to it. For this reason the body was called the “Gemini of the CB” or simply gem (Liu and Dreyfuss 1996). By chance the HeLa line in which the discovery was made is unusual: In other human cell lines, including different HeLa strains, in primary neurons, and in Drosophila, SMN is colocalized with coilin in the CB (Matera and Frey 1998; Carvalho et al. 1999; Liu et al. 2006). Hence SMN can generally be treated as an important component of the CB, not as a marker for a separate nuclear body. As the name implies, SMN is required in mammals for proper functioning of motor neurons, especially those of the spinal cord. In the mouse and Drosophila, null mutations in the single-copy smn gene are lethal (Miguel-Aliaga et al. 2000; Chan et al. 2003; Rajendra et al. 2007). The situation in humans is somewhat different, because there are two copies of the gene, one of which has an altered splice site that leads to inefficient processing of its transcript. Without going into the somewhat complex genetics of the human smn gene, suffice it to say that mutations often lead to the condition

Z. Nizami, S. Deryusheva, and J.G. Gall

known as spinal muscular atrophy (SMA). SMA occurs in approximately 1 in 6000 newborns and results in early death. Biochemical studies show that SMN in vertebrate cells occurs in a macromolecular complex called the assemblysome, consisting of SMN itself, seven gemins, and several other factors. This complex functions in the cytoplasm as the chaperone for assembly of the splicing snRNAs with the seven-member Sm ring (Meister et al. 2002; Pellizzoni et al. 2002; Shpargel and Matera 2005; Battle et al. 2006). SMN accompanies the assembled snRNPs on their return to the nucleus (Narayanan et al. 2004), but whether SMN has a specific nuclear function in the CB is not known. Expression of GFP-tagged human SMN in budding yeast led to specific localization of the probe in a small structure inside the nucleolus, which the authors called the nucleolar body (Verheggen et al. 2001; Verheggen et al. 2002). Some steps in the maturation of U3 snoRNA also occur in this body. The association with the nucleolus, the targeting of SMN, and the maturation of U3 all suggest that the yeast nucleolar body is equivalent to the CB of more complex eukaryotes. Association of CBs with Specific Gene Loci

Because nucleoli are associated with a specific locus or loci on chromosomes, it is natural to ask whether similar associations exist for CBs and other nuclear organelles. In the case of the CB, there is no evidence that transcription occurs in the body itself and so there is no reason to suppose that a CB corresponds to a highly active gene locus like the nucleolus. Nevertheless, CBs might form at specific loci or travel there by some unknown mechanism as a means of bringing needed factors to those loci. Such a relationship is suggested by the fact that CBs in cultured vertebrate cells show preferential association with snRNA gene loci. CBs in these cells associate not only with U1, U2, and U4 gene clusters but also with the loci of the minor U11 and U12 snRNAs (Smith et al. 1995; Frey et al. 1999; Jacobs et al. 1999; Frey and Matera 2001). Matera suggested that snRNAs in the

CB somehow exert a feedback regulation on snRNA transcription at these gene loci. Whatever the reason for the association, the relationship between CBs and snRNA loci is both dynamic and dependent on transcription, as shown in a recent experimental analysis (Dundr et al. 2007). An array of inducible U2 snRNA genes was cotransfected into cultured cells along with fluorescently labeled coilin. So long as the U2 array was transcriptionally silent, the array showed no specific relationship to CBs in the same nucleus. However, upon induction of transcription, the U2 array moved toward the closest CB and eventually came in contact with it. This remarkable movement was disrupted in the presence of a dominant negative mutant of b-actin, suggesting a role for nuclear actin in repositioning chromosomal loci in response to transcriptional activation. Another special relationship exists between CBs and the telomeres. During most of the cell cycle telomerase RNA is demonstrable only in CBs. However, during S phase, when telomeres are elongated, telomerase RNA can also be detected in a subset of telomeres (Ja´dy et al. 2006; Tomlinson et al. 2006; Tomlinson et al. 2008). These telomeres contain other components of the telomerase holoenzyme, which do not normally accumulate there. In addition, CBs were seen to make transient associations with telomeres during the S phase. These results suggest that specific interactions occur between CBs and telomeres during telomere elongation. The functional relevance remains to be determined. HISTONE LOCUS BODIES

The HLB was described several times in Drosophila and human cells before a clear distinction was made between it and the CB (Frey and Matera 1995; Calvi et al. 1998; Liu et al. 2000; Ma et al. 2000). The existence of two separate bodies within the same nucleus was shown in experiments designed to identify the Drosophila CB in tissues of the fly (Liu et al. 2006). Because Drosophila coilin was not available at that time, our laboratory decided to use two other probes that we thought would serve equally well: the U7

Cajal Body and Histone Locus Body

snRNP and U85 scaRNA. Based on studies of the Xenopus oocyte, the U7 snRNP seemed to be a good marker for the CB (Wu and Gall 1993), and U85 scaRNA had been shown to label the CB in Drosophila S2 cells (Darzacq et al. 2002). To our surprise we found that the U7 snRNP and U85 scaRNA detected two independent nuclear organelles in various tissues of the fly (Fig. 3C,D; Fig. 4A,B). The organelle that contained U85 scaRNA was designated the CB, based on the presence of other common CB components. Because the organelle that contained the U7 snRNP was invariably associated with the histone genes on chromosome 2, we called it the HLB. The CB and HLB lie close together or actually touch one another in many Drosophila nuclei, suggesting some still unknown functional relationship between the two. Studies from other laboratories soon showed that HLBs could be identified as organelles distinct from CBs in the nuclei of cultured mammalian cells (Bongiorno-Borbone et al. 2008; Ghule et al. 2008). Most of the factors that have been shown in HLBs are known to be involved in processing of histone pre-mRNAs. The genes coding for the major histones do not contain introns and hence their pre-mRNAs are not spliced. However, the pre-mRNAs carry a 30 extension that is cut off before export to the cytoplasm (reviewed in Marzluff 2005). The site of cleavage is determined by the U7 snRNP acting in concert with a variety of other factors (Dominski et al. 2005; Kolev and Steitz 2005; Sullivan et al. 2009). In addition to U7 snRNA and the two U7 specific snRNP proteins Lsm10 and Lsm11, a number of other proteins have recently been detected in HLBs, including stem-loop binding protein (SLBP), FLASH, p220NPAT, negative elongation factor (NELF), symplekin, and the unknown protein recognized by mAb MPM-2 (Narita et al. 2007; White et al. 2007; Bongiorno-Borbone et al. 2008; Ghule et al. 2008; Ghule et al. 2009; Sullivan et al. 2009; Yang et al. 2009). The behavior of CBs and HLBs during oogenesis in Drosophila turned out to be particularly interesting. The polyploid nurse cell nuclei in the youngest egg chambers usually

display a single CB and a single HLB. As oogenesis proceeds, the number of HLBs increases until about stage 8, when there are usually 8 – 16 HLBs. As the nurse cells continue to grow, their CBs break up into tiny fragments that eventually disappear. In contrast, the HLBs retain their size and number and begin to accumulate coilin. After the nurse cells have dumped their cytoplasm into the oocyte and their nuclei begin to degenerate, the HLBs are especially prominent. They display the usual HLB markers like the U7 snRNP, but now stain for coilin as brightly as typical CBs in other cells (Fig. 4D – F). The net result is that at the end of oogenesis nurse cell nuclei lack typical CBs but have prominent coilin-containing HLBs. AMPHIBIAN OOCYTE NUCLEI: CBs OR HLBs?

The giant oocyte nucleus or germinal vesicle (GV) of amphibians such as Xenopus houses several thousand extrachromosomal bodies. Among these are 50 – 100 bodies originally called spheres on the basis of their shape, but now referred to as CBs, because they contain a high concentration of coilin (Fig. 5). Although most of these bodies “float” freely in the nucleoplasm, a few are attached to the lampbrush chromosomes at the histone gene loci (Gall et al. 1981; Callan et al. 1991). There is no question that the free and attached bodies are equivalent structures, because they are identical in all features of morphology and molecular composition, including a high concentration of the U7 snRNP and other factors involved in histone pre-mRNA processing (Wu and Gall 1993; Abbott et al. 1999). Thus, by composition and association with the histone genes, these amphibian oocyte bodies are HLBs. It is tempting to equate them with the late-stage HLBs of the Drosophila ovary, which also have a high concentration of coilin. If this analogy is carried further, one might expect to find both CBs and HLBs in early amphibian oocytes, with loss of the CBs during oogenesis. The distribution of scaRNAs should shed light on this issue, because they are not in the HLBs of Drosophila, and hence may provide a more

Z. Nizami, S. Deryusheva, and J.G. Gall

A

B

C

D

Figure 5. A single large nuclear body from a Xenopus

germinal vesicle after immunostaining for coilin (green) and trimethylguanosine (red). Because of their high concentration of coilin, these bodies have been described as CBs. However, they contain the U7 snRNP and some are attached to the histone gene loci, making them HLBs by definition. (A) DIC image showing one speckle inside the CB/HLB and three speckles attached to its periphery. The irregular object to the lower left is an extrachromosomal nucleolus. (B) Green channel, showing that coilin is limited to the CB/HLB, being absent from the speckles. (C) Red channel showing trimethylguanosine at high concentration in the CB/HLB and lower concentration in the speckles. (D) Overlay of B and C. Although trimethylguanosine occurs in the nucleolus (caps on U3 and U8 snoRNAs), its concentration is too low to detect in this image. Modified from Bellini and Gall (1998) with permission from Molecular Biology of the Cell. Bar ¼ 5 mm.

definitive marker to distinguish the two types of bodies. We know that the Xenopus early oocyte contains several types of nuclear bodies (besides the amplified nucleoli), but it is not yet clear what happens to them during the midstages of oogenesis.

DYNAMICS AND ASSEMBLY OF CBs AND HLBs

The rate at which components of a cellular structure turn over can be a useful indicator of metabolic activity and ultimately of function.

Broadly speaking, one can distinguish storage particles, whose components are expected to be relatively immobile, and metabolically active bodies that exchange molecules with their environment on a short time scale. Various studies, especially those involving fluorescence recovery after photobleaching (FRAP), suggest that CBs and HLBs fall into the latter category. FRAP studies performed on CBs in cultured cells show rapid exchange of coilin, fibrillarin, and other CB components between the body and the nucleoplasm, with half-times for recovery of the order of a few minutes (Snaar et al. 2000; Sleeman et al. 2003; Dundr et al. 2004). Somewhat slower kinetics were found for coilin, U7 snRNA, and the TATA-binding protein in the large CBs/HLBs of Xenopus oocytes, of the order of 30 min (Handwerger et al. 2003; Deryusheva and Gall 2004). These studies suggest that macromolecules have relatively short residence times inside CBs and HLBs, consistent with these bodies having metabolic functions independent of simple storage. All subcellular organelles show a certain degree of Brownian movement, depending primarily on their size and the viscosity of the surrounding medium. CBs show Brownian motion, but careful time-lapse measurements made on CBs in both mammalian and plant cells show more complex behavior as well, including anomalous diffusion and associations with chromatin (Boudonck et al. 1999; Platani et al. 2000; Carmo-Fonseca et al. 2002; Platani et al. 2002). Occasionally CBs have been seen to move in a directed manner, even traversing a nucleolus (Boudonck et al. 1999). Despite considerable knowledge about the composition of CBs and HLBs, it remains unclear what function is served by the concentration of their components in structures large enough to be seen by conventional microscopy. Because coilin null mutants either lack CBs entirely (Drosophila and Arabidopsis) or show unusual distribution of CB components, one cannot argue that CBs are absolutely essential for nuclear physiology. A possible scenario is that CBs bring macromolecular components together at higher concentrations than

Cajal Body and Histone Locus Body

ordinarily found in the nucleus, thus favoring reactions that otherwise might proceed at unacceptably slow rates. Alternatively, one could argue that CBs might serve to sequester splicing components and hence act to regulate the rate of RNA processing in the cell. In a sense, it is easier to speculate about the function of HLBs. In this case, factors that process histone pre-mRNAs are concentrated at the sites where the pre-mRNAs are transcribed. Whether the bodies are the actual sites of processing is not known. HLBs are found throughout the cell cycle in Drosophila (Liu et al. 2006; White et al. 2007), even though histone pre-mRNA transcription and processing is limited to the S phase. Without knowing the biological role of nuclear bodies in detail, one can still ask how such bodies assemble and disassemble during the cell cycle. Two broad categories of explanation can be distinguished, one being that a body must be built step by step, using cellular machinery in an energy-dependent fashion, the other being that a body will nucleate spontaneously when and if its macromolecular components occur at high enough concentration (Misteli 2001). A recent study provides strong support for this second self-assembly mechanism. In an experiment designed to ask whether coilin can act as a nucleating site for CB formation, Dundr and his colleagues made a construct in which GFP-coilin was linked to the Lac repressor and then transfected into cells that carried an array of 250 repeats of the Lac operator at a specific chromosome site (Kaiser et al. 2008). As expected, an accumulation of GFP-coilin appeared at the site of the Lac operator array. Remarkably, however, this accumulation not only had the size and shape of a typical CB but it contained a variety of other CB components, including SMN, snRNP proteins, and U85 scaRNA. Thus a genuine CB had formed at the chromosomal locus, morphologically and compositionally identical to other CBs found elsewhere in the same nucleus. To test whether coilin was required for the nucleation process, 17 other CB components were targeted one by one to the Lac operator. In each case a CB was formed essentially

identical to the one formed by coilin. In fact, several other components were more efficient than coilin in terms of the fraction of transfected cells that displayed a CB. These experiments demonstrate clearly that formation of a CB is not a step-wise process involving the hierarchical addition of components starting, for instance, with coilin. Instead assembly of the CB has the hallmarks of a self-organizing system, taking place whenever the concentration of one of the major macromolecules, or macromolecular complexes, is high enough. The process continues, sweeping up other components until some still undefined end point is reached.

CONCLUDING REMARKS

In the 18 years since the identification of coilin as a marker for the CB, this nuclear organelle has gone from an almost completely neglected curiosity to a well-studied nuclear body whose involvement in snRNP biogenesis has been clearly shown. Because CBs are found in all eukaryotes that have been carefully examined, it is probable that new and unexpected roles for them will be uncovered in the near future, when additional genetic and molecular studies are undertaken. The relationship of the newly recognized HLB to the CB remains to be worked out. As additional organisms are examined, it seems likely that the HLB, like the CB, will prove to have a wide phylogenetic distribution. Because of its intimate relationship with the histone genes, the HLB promises to give new insight into the control of histone pre-mRNA processing during the cell cycle.

ACKNOWLEDGMENTS

We thank former members of our laboratory for their contributions to the understanding of CB and HLB structure and function. Original research cited here was supported by National Institutes of Health grant GM 33397 from the National Institute for General Medical Studies. JGG is American Cancer Society Professor of Developmental Genetics.

Z. Nizami, S. Deryusheva, and J.G. Gall

REFERENCES Abbott J, Marzluff WF, Gall JG. 1999. The stem loop binding protein (SLBP1) is present in coiled bodies of the Xenopus germinal vesicle. Mol Biol Cell 10: 487 –499. Andrade LEC, Chan EKL, Raska I, Peebles CL, Roos G, Tan EM. 1991. Human autoantibody to a novel protein of the nuclear coiled body: Immunological characterization and cDNA cloning of p80-coilin. J Exp Med 173: 1407– 1419. Bachellerie J-P, Cavaille J. 1997. Guiding ribose methylation of rRNA. Trends Biochem Sci 22: 257– 261. Battle DJ, Kasim M, Yong J, Lotti F, Lau CK, Mouaikel J, Zhang Z, Han K, Wan L, Dreyfuss G. 2006. The SMN complex: An assembly machine for RNPs. Cold Spring Harb Symp Quant Biol 71: 313– 320. Bellini M, Gall JG. 1998. Coilin can form a complex with the U7 small nuclear ribonucleoprotein. Mol Biol Cell 9: 2987– 3001. Beumer KJ, Trautman JK, Bozas A, Liu J-L, Rutter J, Gall JG, Carroll D. 2008. Efficient gene targeting in Drosophila by direct embryo injection with zinc-finger nucleases. Proc Natl Acad Sci USA 105: 19821– 19826. Beven AF, Simpson GG, Brown JWS, Shaw PJ. 1995. The organization of spliceosomal components in the nuclei of higher plants. J Cell Sci 108: 509–518. Bier K, Kunz W, Ribbert D. 1967. Struktur und Funktion der Oocytenchromosomen und Nukleolen sowie der ExtraDNS wa¨hrend der Oogenese panoistischer und meroistischer Insekten. Chromosoma 23: 214– 254. Bongiorno-Borbone L, De Cola A, Vernole P, Finos L, Barcaroli D, Knight RA, Melino G, De Laurenzi V. 2008. FLASH and NPAT positive but not Coilin positive Cajal Bodies correlate with cell ploidy. Cell Cycle 7: 2357– 2367. Boudonck K, Dolan L, Shaw PJ. 1999. The movement of coiled bodies visualized in living plant cells by the green fluorescent protein. Mol Biol Cell 10: 2297–2307. Cajal SRy. 1903. Un sencillo metodo de coloracion seletiva del reticulo protoplasmatico y sus efectos en los diversos organos nerviosos de vertebrados e invertebrados. Trab Lab Invest Biol Univ Madrid 2: 129 –221. Cajal SRy. 1910. El nu´cleo de las ce´lulas piramidales del cerebro humano y de algunos mamiferos. Trab Lab Invest Biol Univ Madrid 8: 27– 62. Callan HG, Gall JG, Murphy C. 1991. Histone genes are located at the sphere loci of Xenopus lampbrush chromosomes. Chromosoma 101: 245– 251. Calvi BR, Lilly MA, Spradling AC. 1998. Cell cycle control of chorion gene amplification. Genes Dev 12: 734– 744. Carmo-Fonseca M, Platani M, Swedlow JR. 2002. Macromolecular mobility inside the cell nucleus. Trends Cell Biol 12: 491– 495. Carvalho T, Almeida F, Calapez A, Lafarga M, Berciano MT, Carmo-Fonseca M. 1999. The spinal muscular atrophy disease gene product, SMN: A link between snRNP biogenesis and the Cajal (coiled) body. J Cell Biol 147: 715–727. Chamberland H, Lafontaine JG. 1993. Localization of snRNP antigens in nucleolus-associated bodies: Study of plant interphase nuclei by confocal and electron microscopy. Chromosoma 102: 220– 226.

Chan YB, Miguel-Aliaga I, Franks C, Thomas N, Trulzsch B, Sattelle DB, Davies KE, van den Heuvel M. 2003. Neuromuscular defects in a Drosophila survival motor neuron gene mutant. Hum Mol Genet 12: 1367– 1376. Collier S, Pendle A, Boudonck K, van Rij T, Dolan L, Shaw P. 2006. A distant coilin homologue is required for the formation of Cajal bodies in Arabidopsis. Mol Biol Cell 17: 2942–2951. Darzacq X, Ja´dy BE, Verheggen C, Kiss AM, Bertrand E, Kiss T. 2002. Cajal body-specific small nuclear RNAs: A novel class of 20 -O-methylation and pseudouridylation guide RNAs. EMBO J 21: 2746–2756. Deryusheva S, Gall JG. 2004. Dynamics of coilin in Cajal bodies of the Xenopus germinal vesicle. Proc Natl Acad Sci USA 101: 4810–4814. Deryusheva S, Gall JG. 2009. Small Cajal body-specific RNAs of Drosophila function in the absence of Cajal bodies. Mol Biol Cell 20: 5250–5259. Dominski Z, Yang XC, Marzluff WF. 2005. The polyadenylation factor CPSF-73 is Involved in histone-pre-mRNA processing. Cell 123: 37– 48. Dundr M, Hebert MD, Karpova TS, Stanek D, Xu H, Shpargel KB, Meier UT, Neugebauer KM, Matera AG, Misteli T. 2004. In vivo kinetics of Cajal body components. J Cell Biol 164: 831 –842. Dundr M, Ospina JK, Sung MH, John S, Upender M, Ried T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095–1103. Frey MR, Matera AG. 1995. Coiled bodies contain U7 small nuclear RNA and associate with specific DNA sequences in interphase human cells. Proc Natl Acad Sci USA 92: 5915–5919. Frey MR, Matera AG. 2001. RNA-mediated interaction of Cajal bodies and U2 snRNA genes. J Cell Biol 154: 499 –509. Frey MR, Bailey AD, Weiner AM, Matera AG. 1999. Association of snRNA genes with coiled bodies is mediated by nascent snRNA transcripts. Current Biol 9: 126– 135. Gall JG. 2003. The centennial of the Cajal body. Nature Rev Mol Cell Biol 4: 975–980. Gall JG, Bellini M, Wu Z, Murphy C. 1999. Assembly of the nuclear transcription and processing machinery: Cajal bodies (coiled bodies) and transcriptosomes. Mol Biol Cell 10: 4385– 4402. Gall JG, Stephenson EC, Erba HP, Diaz MO, BarsacchiPilone G. 1981. Histone genes are located at the sphere loci of newt lampbrush chromosomes. Chromosoma 84: 159 –171. Ghule PN, Dominski Z, Lian JB, Stein JL, van Wijnen AJ, Stein GS. 2009. The subnuclear organization of histone gene regulatory proteins and 3’ end processing factors of normal somatic and embryonic stem cells is compromised in selected human cancer cell types. J Cell Physiol 220: 129–135. Ghule PN, Dominski Z, Yang XC, Marzluff WF, Becker KA, Harper JW, Lian JB, Stein JL, van Wijnen AJ, Stein GS. 2008. Staged assembly of histone gene expression machinery at subnuclear foci in the abbreviated cell cycle of human embryonic stem cells. Proc Natl Acad Sci USA 105: 16964–16969.

Cajal Body and Histone Locus Body Handwerger KE, Murphy C, Gall JG. 2003. Steady-state dynamics of Cajal body components in the Xenopus germinal vesicle. J Cell Biol 160: 495 –504. Hebert MD, Matera AG. 2000. Self-association of coilin reveals a common theme in nuclear body localization. Mol Biol Cell 11: 4159–4171. Hebert MD, Szymczyk PW, Shpargel KB, Matera AG. 2001. Coilin forms the bridge between Cajal bodies and SMN, the spinal muscular atrophy protein. Genes Dev 15: 2720– 2729. Jacobs EY, Frey MR, Wu W, Ingledue TC, Gebuhr TC, Gao L, Marzluff WF, Matera AG. 1999. Coiled bodies preferentially associate with U4, U11 and U12 small nuclear RNA genes in interphase HeLa cells but not with U6 and U7 genes. Mol Biol Cell 10: 1653–1663. Ja´dy BE, Bertrand E, Kiss T. 2004. Human telomerase RNA and box H/ACA scaRNAs share a common Cajal bodyspecific localization signal. J Cell Biol 164: 647–652. Ja´dy BE, Darzacq X, Tucker KE, Matera AG, Bertrand E, Kiss T. 2003. Modification of Sm small nuclear RNAs occurs in the nucleoplasmic Cajal body following import from the cytoplasm. EMBO J 22: 1878–1888. Ja´dy BE, Richard P, Bertrand E, Kiss T. 2006. Cell cycledependent recruitment of telomerase RNA and Cajal bodies to human telomeres. Mol Biol Cell 17: 944– 954. Kaiser TE, Intine RV, Dundr M. 2008. De novo formation of a subnuclear body. Science 322: 1713– 1717. Kiss AM, Ja´dy BE, Darzacq X, Verheggen C, Bertrand E, Kiss T. 2002. A Cajal body-specific pseudouridylation guide RNA is composed on two box H/ACA sno-RNA-like domains. Nucleic Acids Res 30: 4643–4649. Kolev NG, Steitz JA. 2005. Symplekin and multiple other polyadenylation factors participate in 3’-end maturation of histone mRNAs. Genes Dev 19: 2583– 2592. Lafarga M, Casafont I, Bengoechea R, Tapia O, Berciano MT. 2009. Cajal’s contribution to the knowledge of the neuronal cell nucleus. Chromosoma 118: 437– 443. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nature Rev Mol Cell Biol 4: 604– 612. Lamond AI, Carmo-Fonseca M. 1993. Localisation of splicing snRNPs in mammalian cells. Mol Biol Reports 18: 127–133. Liu Q, Dreyfuss G. 1996. A novel nuclear structure containing the survival of motor neurons protein. EMBO J 15: 3555– 3565. Liu J-L, Hebert MD, Ye Y, Templeton DJ, Kung H-J, Matera AG. 2000. Cell cycle-dependent localization of the CDK2-cyclin E complex in Cajal (coiled) bodies. J Cell Sci 113: 1543–1552. Liu J-L, Murphy C, Buszczak M, Clatterbuck S, Goodman R, Gall JG. 2006. The Drosophila melanogaster Cajal body. J Cell Biol 172: 875 –884. Liu J-L, Wu Z, Nizami Z, Deryusheva S, Rajendra TK, Gao H, Beumer KJ, Carroll D, Matera AG, Gall JG. 2009. Coilin is essential for Cajal body organization in Drosophila melanogaster. Mol Biol Cell 20: 1661–1670. Ma T, Van Tine BA, Wei Y, Garrett MD, Nelson D, Adams PD, Wang J, Qin J, Chow LT, Harper JW. 2000. Cell cycle-regulated phosphorylation of p220NPAT by cyclin

E/Cdk2 in Cajal bodies promotes histone gene transcription. Genes Dev 14: 2298– 2313. Marzluff WF. 2005. Metazoan replication-dependent histone mRNAs: A distinct set of RNA polymerase II transcripts. Curr Opin Cell Biol 17: 274–280. Matera AG, Frey MR. 1998. Coiled bodies and gems: Janus or Gemini? Amer J Hum Gen 63: 317–321. Matera AG, Ward DC. 1993. Nucleoplasmic organization of small nuclear ribonucleoproteins in cultured human cells. J Cell Biol 121: 715 –727. Meister G, Eggert C, Fischer U. 2002. SMN-mediated assembly of RNPs: A complex story. Trends Cell Biol 12: 472 –478. Miguel-Aliaga I, Chan YB, Davies KE, van den Heuvel M. 2000. Disruption of SMN function by ectopic expression of the human SMN gene in Drosophila. FEBS Lett 486: 99– 102. Misteli T. 2001. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Monneron A, Bernhard W. 1969. Fine structural organization of the interphase nucleus in some mammalian cells. J Ultrastruct Res 27: 266– 288. Narayanan U, Achsel T, Lu¨hrmann R, Matera AG. 2004. Coupled in vitro import of U snRNPs and SMN, the spinal muscular atrophy protein. Mol Cell 16: 223 –234. Narita T, Yung TM, Yamamoto J, Tsuboi Y, Tanabe H, Tanaka K, Yamaguchi Y, Handa H. 2007. NELF interacts with CBC and participates in 30 end processing of replicationdependent histone mRNAs. Mol Cell 26: 349 –365. Nesic D, Tanackovic G, Kramer A. 2004. A role for Cajal bodies in the final steps of U2 snRNP biogenesis. J Cell Sci 117: 4423– 4433. Patel SB, Bellini M. 2008. The assembly of a spliceosomal small nuclear ribonucleoprotein particle. Nucleic Acids Res 36: 6482– 6493. Patel SB, Novikova N, Bellini M. 2007. Splicing-independent recruitment of spliceosomal small nuclear RNPs to nascent RNA polymerase II transcripts. J Cell Biol 178: 937 –949. Pellizzoni L. 2007. Chaperoning ribonucleoprotein biogenesis in health and disease. EMBO Rep 8: 340– 345. Pellizzoni L, Yong J, Dreyfuss G. 2002. Essential role for the SMN complex in the specificity of snRNP assembly. Science 298: 1775– 1779. Platani M, Goldberg I, Lamond AI, Swedlow JR. 2002. Cajal body dynamics and association with chromatin are ATPdependent. Nature Cell Biol 4: 502–508. Platani M, Goldberg I, Swedlow JR, Lamond A. 2000. In vivo analysis of Cajal body movement, separation, and joining in live human cells. J Cell Biol 151: 1561–1574. Rajendra TK, Gonsalvez GB, Walker MP, Shpargel KB, Salz HK, Matera AG. 2007. A Drosophila melanogaster model of spinal muscular atrophy reveals a function for SMN in striated muscle. J Cell Biol 176: 831 –841. Raska I, Andrade LEC, Ochs RL, Chan EKL, Chang C-M, Roos G, Tan EM. 1991. Immunological and ultrastructural studies of the nuclear coiled body with autoimmune antibodies. Exp Cell Res 195: 27– 37. Richard P, Darzacq X, Bertrand E, Ja´dy BE, Verheggen C, Kiss T. 2003. A common sequence motif determines the

Z. Nizami, S. Deryusheva, and J.G. Gall Cajal body-specific localisation of box H/ACA scaRNAs. EMBO J 22: 4283– 4293. Schaffert N, Hossbach M, Heintzmann R, Achsel T, Lu¨hrmann R. 2004. RNAi knockdown of hPrp31 leads to an accumulation of U4/U6 di-snRNPs in Cajal bodies. EMBO J 23: 3000– 3009. Shpargel KB, Matera AG. 2005. Gemin proteins are required for efficient assembly of Sm-class ribonucleoproteins. Proc Natl Acad Sci USA 102: 17372–17377. Sleeman JE, Lamond AI. 1999. Newly assembled snRNPs associate with coiled bodies before speckles, suggesting a nuclear snRNP maturation pathway. Current Biol 9: 1065– 1074. Sleeman JE, Trinkle-Mulcahy L, Prescott AR, Ogg SC, Lamond AI. 2003. Cajal body proteins SMN and coilin show differential dynamic behaviour in vivo. J Cell Sci 116: 2039–2050. Smith KP, Carter KC, Johnson CV, Lawrence JB. 1995. U2 and U1 snRNA gene loci associate with coiled bodies. J Cell Biochem 59: 473– 485. Snaar S, Wiesmeijer K, Jochemsen A, Tanke H, Dirks R. 2000. Mutational analysis of fibrillarin and its mobility in living human cells. J Cell Biol 151: 653– 662. Spector DL. 1993. Nuclear organization of pre-mRNA processing. Curr Opin Cell Biol 5: 442 –447. Stanek D, Pridalova-Hnilicova J, Novotny I, Huranova M, Blazikova M, Wen X, Sapra AK, Neugebauer KM. 2008. Spliceosomal small nuclear ribonucleoprotein particles repeatedly cycle through Cajal bodies. Mol Biol Cell 19: 2534– 2543. Stanek D, Rader SD, Klingauf M, Neugebauer KM. 2003. Targeting of U4/U6 small nuclear RNP assembly factor SART3/p110 to Cajal bodies. J Cell Biol 160: 505–516. Sullivan KD, Steiniger M, Marzluff WF. 2009. A core complex of CPSF73, CPSF100, and Symplekin may form two different cleavage factors for processing of poly(A) and histone mRNAs. Mol Cell 34: 322– 332. Theimer CA, Ja´dy BE, Chim N, Richard P, Breece KE, Kiss T, Feigon J. 2007. Structural and functional characterization of human telomerase RNA processing and Cajal body localization signals. Mol Cell 27: 869–881. Tollervey D, Kiss T. 1997. Function and synthesis of small nucleolar RNAs. Curr Opin Cell Biol 9: 337 –342. Tomlinson RL, Abreu EB, Ziegler T, Ly H, Counter CM, Terns RM, Terns MP. 2008. Telomerase reverse transcriptase is required for the localization of telomerase RNA to Cajal bodies and telomeres in human cancer cells. Mol Biol Cell 19: 3793–3800. Tomlinson RL, Ziegler TD, Supakorndej T, Terns RM, Terns MP. 2006. Cell cycle-regulated trafficking of human telomerase to telomeres. Mol Biol Cell 17: 955 –965.

Tucker KE, Berciano MT, Jacobs EY, LePage DF, Shpargel KB, Rossire JJ, Chan EKL, Lafarga M, Conlon RA, Matera AG. 2001. Residual Cajal bodies in coilin knockout mice fail to recruit Sm snRNPs and SMN, the spinal muscular atrophy gene product. J Cell Biol 154: 293 –307. Tucker KE, Massello LK, Gao L, Barber TJ, Hebert MD, Chan EKL, Matera AG. 2000. Structure and characterization of the murine p80 coilin gene, Coil. J Struct Biol 129: 269 –277. Tuma RS, Stolk JA, Roth MB. 1993. Identification and characterization of a sphere organelle protein. J Cell Biol 122: 767 –773. Tycowski KT, Shu MD, Kukoyi A, Steitz JA. 2009. A conserved WD40 protein binds the Cajal body localization signal of scaRNP particles. Mol Cell 34: 47– 57. Venteicher AS, Abreu EB, Meng Z, McCann KE, Terns RM, Veenstra TD, Terns MP, Artandi SE. 2009. A human telomerase holoenzyme protein required for Cajal body localization and telomere synthesis. Science 323: 644– 648. Verheggen C, Lafontaine DLJ, Samarsky D, Mouaikel J, Blanchard J-M, Bordonne´ R, Bertrand E. 2002. Mammalian and yeast U3 snoRNPs are matured in specific and related nuclear compartments. EMBO J 21: 2736–2745. Verheggen C, Mouaikel J, Thiry M, Blanchard J-M, Tollervey D, Bordonne´ R, Lafontaine DLJ, Bertrand E. 2001. Box C/D small nucleolar RNA trafficking involves small nucleolar RNP proteins, nucleolar factors and a novel nuclear domain. EMBO J 20: 5480–5490. Walker MP, Tian L, Matera AG. 2009. Reduced viability, fertility and fecundity in mice lacking the Cajal body marker protein, coilin. PLoS One 4: e6171. White AE, Leslie ME, Calvi BR, Marzluff WF, Duronio RJ. 2007. Developmental and cell cycle regulation of the Drosophila histone locus body. Mol Biol Cell 18: 2491–2502. Will CL, Lu¨hrmann R. 2001. Spliceosomal UsnRNP biogenesis, structure and function. Curr Opin Cell Biol 13: 290 –301. Wu C-HH, Gall JG. 1993. U7 small nuclear RNA in C snurposomes of the Xenopus germinal vesicle. Proc Natl Acad Sci USA 90: 6257–6259. Xu H, Pillai RS, Azzouz TN, Shpargel KB, Kambach C, Hebert MD, Schu¨mperli D, Matera AG. 2005. The Cterminal domain of coilin interacts with Sm proteins and U snRNPs. Chromosoma 114: 155 –166. Yang XC, Burch BD, Yan Y, Marzluff WF, Dominski Z. 2009. FLASH, a proapoptotic protein involved in activation of caspase-8, is essential for 30 end processing of histone premRNAs. Mol Cell 36: 267– 278. Zhu Y, Tomlinson RL, Lukowiak AA, Terns RM, Terns MP. 2003. Telomerase RNA accumulates in Cajal bodies in human cancer cells. Mol Biol Cell 15: 81– 90.

PML Nuclear Bodies Vale´rie Lallemand-Breitenbach1 and Hugues de The´1,2 1

INSERM/CNRS/Universite´ Paris Diderot/Institut Universitaire He´matologie U944/ UMR7212, Laboratoire associe´ de la Ligue Nationale contre le Cancer, Hoˆpital St. Louis, 1, Av. C. Vellefaux 75475 Paris Cedex 10, France

2

Service de Biochimie, Hoˆpital St. Louis, 1, Av. C. Vellefaux 75475 Paris Cedex 10, France

Correspondence: [email protected]

PML nuclear bodies are matrix-associated domains that recruit an astonishing variety of seemingly unrelated proteins. Since their discovery in the early 1960s, PML bodies have fascinated cell biologists because of their beauty and their tight association with cellular disorders. The identification of PML, a gene involved in an oncogenic chromosomal translocation, as the key organizer of these domains drew instant interest onto them. The multiple levels of PML body regulation by a specific posttranslational modification, sumoylation, have raised several unsolved issues. Functionally, PML bodies may sequester, modify or degrade partner proteins, but in many ways, PML bodies still constitute an enigma.

PML BODIES, AN INTRODUCTION

ML nuclear bodies (NBs) are spheres of 0.1 – 1.0 mm in diameter found in most cell-lines and many tissues. They belong to the nuclear matrix, an ill-defined super-structure of the nucleus proposed to anchor and regulate many nuclear functions, including DNA replication, transcription, or epigenetic silencing (Stuurman et al. 1990). The PML protein is the key organizer of these domains that recruits an ever-growing number of proteins, whose only common known feature to date is their ability to be sumoylated (Bernardi and Pandolfi 2007). PML and NBs were proposed to fine-tune a wide variety of processes, through facilitation of partner protein posttranslational modifications (notably sumoylation itself ) resulting in partner sequestration, activation, or degradation. Several

P

NBs subtypes have been defined on morphological bases, which all contain an electron-dense shell and an inner core. PML NBs came to the forefront with the observation that the oncogenic PML/RARA protein disrupts them in a treatment-reversible manner (Daniel et al. 1993; Dyck et al. 1994; Koken et al. 1994; Weis et al. 1994; Zhu et al. 1997). PML NBs are regulated by cellular stress: viral infection (Everett 2006), DNA-damage, transformation (Koken et al. 1995; Terris et al. 1995; Gurrieri et al. 2004), and oxidative stress (Yamada et al. 2001a; Villagra et al. 2006). Moreover, transcription of PML and several genes encoding partner proteins is dramatically enhanced by interferons. Yet, pml-/- mice, which cannot assemble NBs, develop normally and live well, demonstrating that NBs are dispensable for most basic biological functions.

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000661 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000661

V. Lallemand-Breitenbach and H. de The´

Nevertheless, recent data has implicated PML in the control of cellular senescence and stem cell self-renewal, extending the fields of investigation of PML function (Ferbeyre et al. 2000; Pearson et al. 2000; Salomoni and Pandolfi 2002; Ito et al. 2008).

HISTORICAL OVERVIEW

As for several other nuclear structures, electron microscopy and auto-antibodies were the two founding fathers of the field. Work from several pioneers in the early 1960s noted the presence of dense spherical objects by electron microscopy (de The´ et al. 1960). Two classes were morphologically described: Empty ones (fibrillar) and granular ones, with an inner microgranular material proposed to be ribonucleoproteins (Fig. 1). PML NBs were later observed by immunofluorescence using autoimmune sera from primary billiary cirrhosis patients. These allowed the identification of the first NB-associated protein, SP100 (Szostecki et al. 1990), and

an initial characterization of these structures (Ascoli and Maul 1991). Identification of an anti-nuclear matrix antibody, which labeled the same structures as SP100 (Stuurman et al. 1992), drew the first link between these bodies and the nuclear matrix. The localization of PML, a protein fused to the retinoic acid receptor a(RARA) in the PML/RARA oncoprotein of acute promyelocytic leukemia (APL), to the same nuclear dots as SP100, renewed interest for these domains. The observation that in APL cells PML/RARA disrupted these domains drew instant excitement from the scientific community (Daniel et al. 1993; Dyck et al. 1994; Koken et al. 1994; Weis et al. 1994). Moreover, PML bodies were restored by two different anti-APL therapies, retinoic acid and arsenic trioxide, later shown to trigger PML/RARA degradation (Quignon et al. 1997; Zhu et al. 2001), identifying the first striking parallel between the status of the bodies and that of the cell. Many further studies showed that PML bodies altered in stress conditions, notably viral infections, heat shock, and

Figure 1. Different types of PML bodies. Immuno-gold electron microscopy of PML NBs in CHO cell line stably

overexpressing PML. (A, B, C) “Classical” PML NBs. PML is distributed on a dense electron shell (A, B) or on light halo (C ), that can contain a microgranular inner core (B, C) or not (A). (D, E) PML body in arsenic trioxide-treated cell. Note the absence of the inner core and the enrolled fibrillar aspect of the body (D). (F ) PML-II targets the inner membrane region of the nuclear envelope in pml-/- MEF cells. Images courtesy of Edmond and Francine Puvion (CNRS, Villejuif, France).

PML Nuclear Bodies

exposure to heavy metals (Everett 2001; Dellaire and Bazett-Jones 2004; Everett 2006). PML bodies recruit an ever-increasing number of partner proteins (now in the range of 100), one of the most studied being DAXX, a potent repressor of transcription and modulator of apoptosis. Critically, PML is the actual organizer of the bodies (Ishov et al. 1999; Zhong et al. 2000a; Lallemand-Breitenbach et al. 2001). Among these recruited proteins, one deserves a special mention: an ubiquitin-like protein named SUMO, as PML conjugation by SUMO plays a critical role in recruitment of partners, many of which are sumoylated themselves. Recent studies have focused on more fundamental cell-biology issues: Dynamics of the bodies (Eskiw et al. 2003, Muratani et al., 2002, Chen et al., 2008), relation to other nuclear components (Wang et al. 2004; Batty et al. 2009; Russell et al. 2009), mode of assembly (Shen et al. 2006), and determinants of partner recruitment and actual function in cellulo and in vivo. THE PML PROTEINS

PML is a member of the TRIM/RBCC family of proteins, many members of which are ubiquitin ligases that generate subcellular structures through autoassembly (Reymond et al. 2001; Meroni and Diez-Roux 2005). Transcription of the PML gene is tightly controlled by interferons a/b or g, but also by p53 (Stadler et al. 1995; de Stanchina et al. 2004), which both yield a dramatic increase in the number and the size of the bodies. PML harbors an amino-terminal RING finger that directly binds the SUMO E2 ligase UBC9 (Duprez et al. 1999), two RINGlike domains, the B boxes (Tao et al. 2008), and a coiled-coil mediating homodimerization (Kastner et al. 1992). For other RBCC/TRIM family members, partner binding-specificity often relies on the carboxyl terminus. For PML, a variety of carboxy-terminal domains generated by alternative splicing yield isoforms (Jensen et al. 2001). When expressing single PML isoform in pml-/- cells, distinct types of PML NBs were observed, implying that isoformspecific sequences contact different nuclear constituents that influence morphogenesis (Beech

Table 1. PML isoforms with specific localizations. PML isoform: Specific localization in pml -/- cells:

PML I PML II PML IV PML V PML VII

Nucleolar upon stress, cytoplasm Fibrillar/nuclear envelope (lamina?) Nucleolar upon stress Thick shell/nuclear matrix tethering Cytoplasmic/early endosomes

The expression of PML isoforms in pml-/- cells reveals specific localizations, suggesting that the carboxy-terminus drives interactions with specific unidentified partners. From (Beech et al. 2005; Condemine et al. 2006; WeidtkampPeters et al. 2008).

et al. 2005; Condemine et al. 2006; WeidtkampPeters et al. 2008) (Table 1). Yet, because of the coiled-coil, all endogenous isoforms colocalize. The most abundant (but perhaps least studied) isoform, PML-I, harbors an exonuclease-III domain, that targets PML to nucleolar caps in stressed or senescent cells (Condemine et al. 2007). In addition to the nuclear localization signal (NLS) present in all PML isoforms, PML-I harbors a nuclear export signal (NES) that allows shuttling of all isoforms between the two compartments through heterodimer formation (Henderson and Eleftheriou 2000; Beech et al. 2005; Condemine et al. 2006). The most extensively studied isoform, PML-IV, induces senescence in primary human fibroblasts (Bischof et al. 2002) and apoptosis in many other cellular settings, at least in part through p53 activation (Guo et al. 2000). PML undergoes several critical posttranslational modifications, notably phosphorylation and sumoylation. PML sumoylation has been implicated in NB-morphogenesis. DNA damageorstress-activated kinases like ATM, ATR, CHK2, HIPK2, CK2, or ERK phosphorylate PML, possibly regulating PML stability, NB biogenesis and partner association (Engelhardt et al. 2003; Hayakawa and Privalsky 2004; Scaglioni et al. 2006; Gresko et al. 2009) and contributing to DNA repair or apoptosis control. MORE THAN ONE PML BODY!

It is generally assumed that “PML NBs” designates a single object. However, there is considerable evidence that PML bodies are diverse.

V. Lallemand-Breitenbach and H. de The´

Indeed, PML aggregates onto different sites to create an unsuspected repertoire of PML-accumulating domains in response to a variety of stresses (Eskiw et al. 2003; Bernardi and Pandolfi 2007). Before engaging in the catalog of structures that contain PML, one has to realize that contrarily to the common appreciation, the overwhelming majority of the PML protein pool in a cell is actually not NB-bound. In most cell lines, even in vivo, more than 90% of PML has a diffuse nuclear localization, not associated with the nuclear matrix or NBs (Lallemand-Breitenbach et al. 2001) (P. Hemmerich, personal communication). The most extensively studied factor modulating PML distribution has been arsenic trioxide (see later discussion), although several DNA-damage-activated kinases are also

important. Stress-induced aggregation may promote aggregation of typical NBs or conversely disperse them into microspeckles (see Table 2). The differences among these bodies can be based on morphology or content, yielding a much more dynamic view of PML than previously thought. Structure of the Classical Body

The classical PML body is a spherical object with a diameter of 0.1 – 1 mm, which may or may not have a micro-granular centre. These bodies, from five to 15 per nucleus in cell-lines, are mostly proteinaceous in nature and do not in general contain RNA or DNA (Boisvert et al. 2000). PML forms the outer shell of the

Table 2. PML NBs are sensitive to cellular stress. PML NB phenotypes

SUMO colocalization

Cell types

Ref

IFNs

Increased number and size

Increased

All

Stadler, 1995

As2O3

Large PML shell, decreased number

Increased

All

Zhu, 1997 LallemandBreitenbach, 2001

CdCl2/heat shock

Dispersed micro-bodies

No

All

Eskiw, 2003

Nucleolus

Yes

Primary

Condemine, 2007 Mattsson K, 2001

Increased number and size

Increased

Transformed

LallemandBreitenbach, 2001 Everett, 1999

Large, Peri-nucleolar

?

Primary þ transformed

JanderovaRossmeislova, 2007 Condemine, 2007

Dispersed micro-bodies

Yes

Transformed

Eskiw, 2004

Large, perinucleolar

Yes

Primary transformed

Bernardi, 2004 Condemine, 2007 Kurki, 2003

Yes

Primary transformed

Salomoni, 2005, Condemine, 2007 Eskiw, 2003 Eskiw, 2004

Stress

Proteasome Inhibition

Actinomycin-D

Doxo, IRgamma, UVc DNA Damage

UVc, alkylators, staurosporine DNase

Dispersed micro-bodies

List of exogenous agents that perturb PML NB organization; the status of PML sumoylation followed by NB localization is indicated. The phenotype of PML NBs is dependent on the type of stress and cellular context (transformed cell lines or primary cells).

PML Nuclear Bodies PML

PML dimerization

PML NB nucleation

PML sumoylation

Mature PML NBs

As2O3

SIM (SUMO interaction motif) SIM-containing partners

SUMO Sumoylated partners

Partners containing both a SIM and SUMO

Figure 2. Schematic representation of PML NB biogenesis. PML proteins first dimerize through the RBCC

domains and then multimerize to nucleate NBs. PML sumoylation then leads to organization in spherical body. SIM-containing or sumoylated partners (or both) are recruited by the SUMO or SIM of PML into the inner core of the body.

bodies and partners are usually inside, which is easily shown after PML/partner overexpression (Guiochon-Mantel et al. 1995) (Figs. 1, 2). Like several other bodies, PML NBs are present in the interchromosomal space (Bridger et al. 1998), likely explaining why they are often found close or adjacent to other bodies (Wang et al. 2004; Sun et al. 2005; Batty et al. 2009; Russell et al. 2009). Although devoid of DNA, PML NBs may be associated with some specific chromosomal loci, like MHC class I gene cluster region for which PML NBs were proposed to modulate chromatin architecture and transcription (Shiels et al. 2001; Wang et al. 2004; Kumar et al. 2007). One elegant study has found a PML body constantly juxtaposed to a repressor locus, underscoring the links with transcriptional regulation (Tsukamoto et al. 2000). Conversely, chromatin changes occurring during transcription or the cell cycle may modulate PML NBs structure and number (Eskiw et al. 2004; Wang et al. 2004;

Ching et al. 2005). PML NBs are profoundly modified during many virus infections (Everett 2006). They may, for example, accumulate viral genomes at their periphery or within their central core during infection of quiescent cells (Everett et al. 2007). PML NBs, Telomeres and DNA Damage

Alternative lengthening of telomeres (ALT)-associated PML bodies (APBs) are larger structures observed in cell-lines that do not express telomerase and maintain telomere length by homologous recombination. APBs contain two types of DNA double-strand break repair and homologous recombination factors, the Rad50/Mre11/NBS1 complex and Rad51/Rad52, together with the replication factor A (RPA), the helicase BLM and the telomeric repeat-binding factors TRF1 and TRF2 (Wu et al. 2000; Wu et al. 2003). APBs harbor actively replicating telomeres in the S/G2

V. Lallemand-Breitenbach and H. de The´

phase of the cell cycle. Almost all these proteins are sumoylated. Association of the SUMO ligase SMC5/6 complex with APBs is required for TFR1/2 sumoylation and cell survival (Potts and Yu 2007). Recent studies have argued for a role of PML in facilitating these processes (Jiang et al. 2007; Draskovic et al. 2009). PML is phosphorylated by several DNAdamage activated kinases, including ATM, ATR, HIPK2, or CHK2. Several studies have implicated PML in DNA-repair, through the recruitment to or release from NBs of many proteins like BLM (Bischof et al. 2001), the Mre11 complex (Carbone et al. 2002), WRN, or TOPBP1 (Dellaire and Bazett-Jones 2004). Some of these proteins are localized in NBs in unstressed conditions, whereas others are only associated with NBs after DNA-damage. Although NBs partially overlap with gH2AX foci, their possible localization on the damage sites is still disputed (Boe et al. 2006). Initial studies localized NBs with BLM on BrdU-positive putative DNA repairsites afterirradiation (Bischof et al. 2001). Yet, in later reports, PML-NBs were proposed to retain BLM away from DSB through BLM sumoylation, which could be released by phosphorylation (Eladad et al. 2005; Rao et al. 2005). In response to DNA damage, PML NBs appear to sense the damaged sites, yielding an increased number of microbodies by a fission mechanism. The exact role of PML (bystander or actor) in theses diverse processes is still unclear. Yet, pml-/- cells have a very high rate of sister chromatin exchange (Zhong et al. 1999), implying that PML regulates at least some aspects of homologous recombination.

have observed this granular core in a subset of PML-labeled structures (Fig. 1). The nature of this micro-granular core deserves a renewed exploration. Some studies showed the presence of nascent RNA within a subset of the bodies (LaMorte et al. 1998; Fuchsova et al. 2002; Kiesslich et al. 2002), whereas others did not (Boisvert et al. 2000). Finally, a series of papers has linked PML bodies and mRNA export and translation through eukaryotic translation initiation factor 4E (Lai and Borden 2000; Cohen et al. 2001; Culjkovic et al. 2006; Borden and Culjkovic 2009). Heterogeneity of the results may reflect analysis of different cellular systems, but these issues clearly deserve additional studies. In primary or senescent cells, others and we have observed larger structures that contain nucleolar elements, senescence-associated NBs (SANBs) (Condemine et al. 2007; JanderovaRossmeislova et al. 2007). Interestingly, in this setting, NB-associated proteins are not present within the inner core, but directly on the PML shell (Condemine et al. 2007). Formation of these domains requires PML-I, which contains a nucleolar targeting domain, possibly the exonuclease-III homology domain (Condemine et al. 2007). The nucleolar constituent recognized by this domain and the consequence of PML delocalization for nucleolar function are unknown. Yet, upon genotoxic stress or inhibition of transcription, ATR-mediated PML phosphorylation relocalizes PML to nucleolar caps, where PML was proposed to sequester the p53 ubiquitin ligase MDM2 (Bernardi et al. 2004; Shav-Tal et al. 2005) (Table 2).

Nucleolus and RNAs

Other Subnuclear Compartments

Electron microscopy studies noted that a subset of bodies contains a granular core, typical for the presence of ribonucleoproteins (Dupuy-Coin et al. 1972). Some of these granular bodies were proposed to bud from the nucleolus (DupuyCoin and Bouteille 1972). This most likely reflects the presence of specific RNA-binding proteins and is often observed in hormone-stimulated primary cells (Vagner-Capodano et al. 1982). What determines the presence of the microgranular core inside PML NBs is unknown. We

Studies performed in human embryonic stem cells (HESC) have revealed a further unsuspected heterogeneity in PML NBs (Butler et al. 2009). PML NBs assemble in “rosettes” surrounding DNA centromeres or are distributed in tracks bridging two centromeres. These tracks can cross the nucleus and were frequently observed filing lamina gaps at the nuclear envelope. These PML structures do not contain SUMO, SP100, or DAXX proteins and their role in stem cell fate remains to be explored. The

PML Nuclear Bodies

PML-II isoform is associated with the inner membrane of nuclearenvelope in somatic cells (Condemine et al. 2006) (Fig. 1). NBs are associated with centromeres in proteasome inhibitor-treated cells in the G2 phase of the cell cycle (Everett et al. 1999a). Finally, NBs containing pericentromeric satellite DNA together with HP1, BRCA1, ATRX, and DAXX were proposed to play a role in the re-establishment of the condensed heterochromatic state of late-replicating satellite DNA in ICF syndrome and possibly in normal cells (Luciani et al. 2006). Cytoplasmic PML

Among PML isoforms some are devoid of NLS and yield purely cytoplasmic bodies, when expressed in pml-/- cells. Similarly, because PML-I has a nuclear export signal, it forms some cytoplasmic PML-labeled bodies (Condemine et al. 2006). Cytoplasmic PML was proposed to regulate TGFb signaling by controlling the association of Smad2/3 with SARA and accumulation of SARA and TGFb-receptor in the early endosomes (Lin et al. 2004) (Table 1). NB DYNAMICS Basal Status

NBs have been the focus of several studies using imaging technologies such as FRET or FRAP (Tsukamoto et al. 2000). These have shown that PML is a stable component of the bodies and that partner proteins are more mobile, although they are transiently retained in NBs (Boisvert et al. 2001; Weidtkamp-Peters et al. 2008). The exchange rates of the different PML isoforms between NBs and nucleoplasm showed a clear difference for PML-V, which forms peculiar thick-shelled NBs and might anchor NBs onto the nuclear matrix (Condemine et al. 2006; Weidtkamp-Peters et al. 2008). The bodies themselves are not very mobile, although fusions and fissions can be observed through the cell cycle progression (see later discussion). Some bodies labeled by GFP-SP100 are smaller and more dynamic than typical PML NBs (Muratani et al. 2002; Wiesmeijer et al. 2002).

Telomeres quickly get in and out of APBs, whereas their PML shell is quite immobile (Molenaar et al. 2003; Jegou et al. 2009), similar to that of SANBs (Condemine et al. 2007). When interpreting these studies, one has to remember that some were performed in transiently transfected cells where posttranslational modifications, which are essential determinants of partner residence, are unlikely to be complete. Cell-Cycle Analysis

Analyses of NBs during the cell cycle have provided evidence for duplication by a fission mechanism during S phase (Dellaire et al. 2006c) and have dissected NB reformation during the M/G1 transition (Dellaire et al. 2006b; Chen et al. 2008). During mitosis, PML proteins remain aggregated, but are phosphorylated, become desumoylated and release partners (Everett et al. 1999b). Before nuclear membrane breakdown in prometaphase, PML NBs lose their chromatin-tethering, resulting in increased mobility (Chen et al. 2008). Yet, FRAP performed during mitosis showed that PML proteins did not exchange, demonstrating that mitotic bodies are highly stable (Dellaire et al. 2006c). PML associates with nuclear membranes and nucleoporins during mitosis, facilitating reformation of the nuclear envelope during the telophase/G1 transition (PuvionDutilleul et al. 1995; Jul-Larsen et al. 2009). Finally, during telophase to G1 transition, SP100 and DAXX re-enter the nucleus and then bind preformed NBs, SP100 first and later DAXX, consistent with the critical role of PML in NB-nucleation. Stress Response

Apart from arsenic trioxide that promotes NB formation (see below), heat shock or heavy metals induce reversible NB fragmentation by budding of highly mobile micro-bodies that are devoid of SUMO and most partners (Table 2). In contrast, transcriptional inhibition or DNA damage induce formation of microspeckles or nucleolar caps depending on the type of cells (transformed or primary), with a maintained

V. Lallemand-Breitenbach and H. de The´

SUMO-colocalization (Table 2) (Eskiw et al. 2003; Nefkens et al. 2003; Dellaire et al. 2006a). Single cell studies after stress recovery have shown that the initial size, location, and number of NBs was somehow restored, suggesting that NBs may assemble on predetermined sites. A full exploration of the role of PML phosphorylation for the regulation of PML mobility and NB formation would be important to clarify these issues. IS SUMO THE NB-ASSEMBLY GLUE?

SUMO is conjugated to target proteins on the side chain of lysine residues, creating a branched peptide and significantly changing the binding properties of the protein. SUMO has been implicated in multiple pathways, mostly as a regulator of protein interactions (Hay 2005). Conjugated SUMO may interact with a short motif, known as the SUMO interaction motif (SIM) (Minty et al. 2000; Hecker et al. 2006). SUMO-1 was first identified as a PML partner in a two-hybrid screen (Boddy et al. 1996). PML was shown to be sumoylated on three lysine residues: K65 in the RING finger, K160 in the B1 box, and K490 in the NLS. PML also contains a SIM. Accordingly, intermolecular interactions between the PML SUMO and SIM were proposed to underlie NB biogenesis (Matunis et al. 2006; Shen et al. 2006). Sumoylation-defective Ubc9-/- cells indeed show NB defects (Nacerddine et al. 2005). Moreover, because most partner proteins associated with classical PML NBs are sumoylated and many contain a SIM, SIM/SUMO interactions may also account for partner recruitment as well as sequestration. Studies on NB-biogenesis have been greatly facilitated by arsenic trioxide. Arsenic trioxide, a very potent therapeutic agent in APL (Zhu et al. 2002; Kogan 2009), was first shown to target PML/RARA for degradation through its PML moiety (Chen et al. 1997; Zhu et al. 1997; Muller et al. 1998; Shao et al. 1998). Arsenic trioxide is the only factor that regulates the partitioning of PML between the nucleoplasm and the nuclear matrix, promoting in a sequential manner NB-formation, PML sumoylation, partner recruitment, and PML degradation

(Lallemand-Breitenbach et al. 2001; LallemandBreitenbach et al. 2008) (Fig. 2). Critically, PML sumoylation on K160/K65 follows matrix-targeting and is required for partner recruitment (Ishovet al. 1999; Zhong et al. 2000a; LallemandBreitenbach et al. 2001). Yet, howarsenic trioxide initiates the SUMO-independent transfer from the soluble diffuse nucleoplasmic form toward the insoluble matrix fraction and why matrix association is followed by sumoylation of these two sites remains unexplained to date. SUMO/SIM interactions were initially proposed to underlie both formation of the PML mesh and recruitment of partners. Although appealing, this model has been significantly challenged by recent evidence. Specific PML isoforms that do not harbor the SIM yield normal bodies (Weidtkamp-Peters et al. 2008). Conversely, analysis of several partner proteins has shown that their SIM is essential, whereas their sumoylation is dispensable for NBtargeting (Takahashi et al. 2005; Lin et al. 2006; Cho et al. 2009). Taken together, SIM/ SUMO are unlikely to play a fundamental role in PML aggregation to create the PML mesh, but may be critical for partner recruitment. WHAT ARE THE FUNCTIONS OF NBs?

PML influences or regulates key processes such as transcription, apoptosis, senescence, response to DNA-damage or resistance to micro-organisms, which have all been extensively reviewed elsewhere (Salomoni and Pandolfi 2002; Everett 2006; Bernardi and Pandolfi 2007; Bernardi et al. 2008). Two novelties have emerged: (1) the protective role of PML, DAXX, and SP100 against many viral infections (Everett et al. 2006; Tavalai et al. 2006; Everett et al. 2008; Tavalai et al. 2008; Kyratsous and Silverstein 2009), at least in the absence of the viral proteins that disrupt NBs; (2) the role of PML in normal orcancer stem cell fate (Ito et al. 2008; Li et al. 2009; Regad et al. 2009). PML-enforced stem cell self-renewal may rely on its ability to modulate the AKT pathway through regulation of AKT phosphorylation or through the localization of its regulator PTEN (Trotman et al. 2006; Song et al. 2008; Ito et al. 2009). TR2, a modulator of the critical Oct4

PML Nuclear Bodies

stem cell transcriptional regulator, is dependent on PML for its sumoylation (Park et al. 2007; Gupta et al. 2008). The importance of PML in “stemness” highlights the distinctly uncommon morphologyof NBs in ES cells (Butleret al. 2009) and questions the nature of the PML isoforms that they actually express. How are these functions actually achieved? The current models envision PML as glue, whose major function is to recruit and concentrate partners within NBs. Recruitment of partners together with many protein-modifying enzymes, could, in principle, enhance posttranslational modifications, yielding activation, sequestration or degradation. There is evidence to support all three of these processes (Fig. 3). Partner Posttranslation Modifications

Perhaps the most studied posttranslational modifications have been those on the tumor

suppressor p53. A striking finding has been the concentration of p53-modifying enzymes (CBP, HDM2, HIPK2, and HAUSP) within NBs. PML-enhanced acetylation, sumoylation, and phosphorylation occurring in NBs all appear to enhance p53 function. In that respect pml-/- cells are distinctly impaired in their ability to undergo senescence (Ferbeyre et al. 2000; Pearson et al. 2000; de Stanchina et al. 2004), whereas conversely PML-IVoverexpression triggers senescence by a pathway involving both p53 and Rb (Bischof et al. 2002; Mallette et al. 2004; Bischof et al. 2005). The complexity of the multiple mechanisms through which PML enhances p53 function and some apparent contradictions in the published data plea for a renewed exploration of this critical issue in primary cells, using siRNA or gene excision rather than PML/partner overexpression. Kinase activity may be affected by translocation into NBs: PP2A dephosphorylates p-Akt

Apoptosis/ senescence

Post-translational modifications PML

Ub Poly-Q proteins Transcription control

Degradation

Viral proteins

Sequestration

SUMO

PML Daxx Sp100 TDG PIASy HIPK2 LEF1

Phospho Akt ChK2 P53

Regulation of activity Stem cell maintenance

Acetyl p53

DNA repair /ALT

Figure 3. A general function for PML NBs: Integrated posttranslational protein modifications? PML NBs regulate posttranslational modifications of partner proteins like sumoylation, ubiquitination, but also phosphorylation or acetylation. These modifications regulate a wide variety of partners, leading to modulation of biological processes like transcription, apoptosis/senescence, DNA repair, or ALT and stem cell self-renewal.

V. Lallemand-Breitenbach and H. de The´

within PML NBs (Trotman et al. 2006), whereas PP1A was proposed to dephosphorylate RB (Regad et al. 2009). In contrast, recruitment into NBs may favor auto-phosphorylation of some kinases, such as CHK2 (Yang et al. 2002; Yang et al. 2006). There are also some indications that PML can directly enhance global protein sumoylation in yeast (Quimby et al. 2006) and NBs were proposed to enhance sumoylation of specific partner proteins (Park et al. 2007). Because many NB-associated proteins contain a SIM, this may enhance partner sequestration within NBs.

that silences proliferation-associated genes, senescence-associated heterochromatin foci (SAHF), is one of the first known senescenceassociated events. The latter is initiated by the concentration of two histone chaperones HIRA and ASF1a, together with the heterochromatin protein HP1, within PML NBs (Zhang et al. 2005; Ye et al. 2007a; Ye et al. 2007b). What initiates their translocation into NBs and how transient concentration of heterochromatin-associated histones and their chaperones into NBs later favor SAHF formation is not understood. Partner Degradation

Partner Sequestration

Sequestration or “depot” was the first proposed function of PML NBs (Negorev and Maul 2001). This sequestration is evident by the relative accumulation of the nucleoplasmic and the NB-associated form of PML-partners, which very significantly varies between individual partners and levels of PML expression, as well as sumoylation. A well-studied sequestered partner is DAXX, a potent repressor that partitions between sumoylated proteins, including PML and many transcription factors. Sequestration of DAXX by NB-associated, sumoylated PML releases transcriptional repression by DNA-bound sumoylated transcription factors (Li et al. 2000; Lehembre et al. 2001; Lin et al. 2003; Lin et al. 2006). Sequestration of DAXX also regulates apoptosis. In particular, DAXX enhances Fas-triggered caspase activation, possibly through release of ASK1 kinase activity (Torii et al. 1999; Zhong et al. 2000b; Salomoni and Khelifi 2006), although opposite results have been observed in other cell-types (Meinecke et al. 2007). Sequestration of DAXX may also modulate other critical regulators, such as HAUSP, ultimately deubiquitinating the PTEN tumor suppressor (Song et al. 2008). In any case, DAXX, which exerts pro- or anti-apoptotic functions, is a critical partner of PML involved in many of it properties (Maul et al. 2000). A related situation is the concentration of histones and their chaperones in senescent cells. Formation of a specific type of heterochromatin

Several unstable proteins have been localized to PML bodies (Anton et al. 1999; Smith et al. 2004), whereas proteins that are impaired in their degradation aggregate with PML, SUMO, and ubiquitin (Yamada et al. 2001b). NBs concentrate proteasomes and ubiquitin (Fabunmi et al. 2001; Lallemand-Breitenbach et al. 2001; Lafarga et al. 2002; Lallemand-Breitenbach et al. 2008). Arsenic trioxide-induced PML degradation has linked PML bodies and protein degradation. Arsenic trioxide triggers an initial sumoylation of K160, followed by proteasomedependent degradation, a very controversial proposal at the time this was discovered (Zhu et al. 1997; Lallemand-Breitenbach et al. 2001). Yet, identification of a yeast SUMO-dependent ubiquitin ligase (Geoffroy and Hay 2009) and realization that its human ortholog RNF4 localizes to NBs (Hakli et al. 2005) led to renewed interest in these findings. The initial arsenic trioxide-induced K160 sumoylation triggers a secondary polyubiquitination by RNF4 and proteasome-mediated degradation in NBs (Lallemand-Breitenbach et al. 2008; Tatham et al. 2008). There is also some evidence for partner degradation within NBs (Kitagawa et al. 2006; Qin et al. 2006; St-Germain et al. 2008), but how general this is and what the specific signals are that promote partner degradation within NBs is currently unknown. Note that for the same pathway, a combination of those different mechanisms may cooperate. For example, modulation of p53 function

PML Nuclear Bodies

may involve both enhanced p53 modifications and HDM2 sequestration. Similarly, regulation of AKT function involves both dephosphorylation in NBs and sequestration of the DAXX/ HAUSP regulators of PTEN activity, to control the critical AKT/TOR/FOXO pathway for stem cell fate (Ito et al. 2009). CONCLUDING REMARKS AND OPEN QUESTIONS

Because PML can aggregate with so many cellular components, forced overexpression can yield many artifacts, notably by titration of partners or aggregation of unstable proteins (Takahashi et al. 2004; Bernardi and Pandolfi 2007). Paradoxically, there are only few studies in which PML functions were investigated in stable transfectants or in pml-/- cells or in which siRNA were used to inactivate the endogenous PML proteins. Even these experimental approaches preclude assessing the role of the bodies per se, compared with that of the diffuse nucleoplasmic fraction of PML. For example, PMLIV-initiated senescence is unaffected by the viral protein CMV IE1, which disrupts NBs (Ahn and Hayward 1997; Bischof et al. 2002). Separating the PML NB functions from those of unassembled PML may be difficult until ad hoc mutants are described. Another critical issue concerns isoforms. The vast majority of studies have used the PML-IV isoform, which is expressed at very low levels in most cell lines (Stuurman et al. 1992; Condemine et al. 2006), questioning their actual relevance. Finally, future studies will have to address the paradoxical critical role of PML in multiple key processes and the surprisingly modest phenotype of pml-/- mice. Perhaps one clue to this issue is that stress response has not been fully explored in these animals, despite some important contributions (Wang et al. 1998) and that stem cell defects may have relatively few manifestations in adults. The PML field has been extremely active, yielding over the past years a large number of remarkable results. Yet, some very fundamental issues remain to be explained. What is the biochemical and physiological basis for NB-

assembly? What is the micro-granular core? What are the specific functions of different isoforms? Why are so many NB-associated proteins transcriptionally controlled by interferons, including PML itself (Stadler et al. 1995; Gro¨tzinger et al. 1996; Engelhardt et al. 2001; Fabunmi et al. 2001)? These issues will certainly be clarified in the years to come, maintaining a vivid and exciting field. ACKNOWLEDGMENTS

We apologize to our many colleagues whose work could not be cited because of space restrictions. We warmly thank Edmond and Francine Puvion for their electron microscopy images and many discussions at various stages of this project, Stan Fakan for helpful comments, Morgane Le Bras and Julien Ablain for critical reading of the manuscript. Work in the laboratory is supported by Ligue Nationale contre le Cancer, INSERM, CNRS, University Paris Diderot, Institut Universitaire de France, Institut National du Cancer and Canceropole programs. REFERENCES Ahn JH, Hayward GS. 1997. The major immediate-early proteins IE1 and IE2 of human cytomegalovirus colocalize with and disrupt PML-associated nuclear bodies at very early times in infected permissive cells. J Virol 71: 4599–4613. Anton LC, Schubert U, Bacik I, Princiotta MF, Wearsch PA, Gibbs J, Day P, Realini C, Rechsteiner M, Bennink J, et al. 1999. Intracellular localization of proteasomal degradation of a viral antigen. J Cell Biol 146: 113 –124. Ascoli CA, Maul GG. 1991. Identification of a novel nuclear domain. J Cell Biol 112: 785– 795. Batty E, Jensen K, Freemont P. 2009. PML nuclear bodies and their spatial relationships in the mammalian cell nucleus. Front Biosci 14: 1182– 1196. Beech SJ, Lethbridge KJ, Killick N, McGlincy N, Leppard KN. 2005. Isoforms of the promyelocytic leukemia protein differ in their effects on ND10 organization. Exp Cell Res 307: 109 –117. Bernardi R, Pandolfi PP. 2007. Structure, dynamics and functions of promyelocytic leukaemia nuclear bodies. Nat Rev Mol Cell Biol 8: 1006–1016. Bernardi R, Papa A, Pandolfi PP. 2008. Regulation of apoptosis by PML and the PML-NBs. Oncogene 27: 6299–6312. Bernardi R, Scaglioni PP, Bergmann S, Horn HF, Vousden KH, Pandolfi PP. 2004. PML regulates p53 stability by

V. Lallemand-Breitenbach and H. de The´ sequestering Mdm2 to the nucleolus. Nat Cell Biol 6: 665–672. Bischof O, Kim SH, Irving J, Beresten S, Ellis NA, Campisi J. 2001. Regulation and localization of the bloom syndrome protein in response to dna damage. J Cell Biol 153: 367–380. Bischof O, Kirsh O, Pearson M, Itahana K, Pelicci PG, Dejean A. 2002. Deconstructing PML-induced premature senescence. Embo J 21: 3358–3369. Bischof O, Nacerddine K, Dejean A. 2005. Human papillomavirus oncoprotein E7 targets the promyelocytic leukemia protein and circumvents cellular senescence via the Rb and p53 tumor suppressor pathways. Mol Cell Biol 25: 1013–1024. Boddy MN, Howe K, Etkin LD, Solomon E, Freemont PS. 1996. PIC1, a novel ubiquitin-like protein which interacts with the PML component of a multiprotein complex that is disrupted in acute promyelocytic leukaemia. Oncogene 13: 971 –982. Boe SO, Haave M, Jul-Larsen A, Grudic A, Bjerkvig R, Lonning PE. 2006. Promyelocytic leukemia nuclear bodies are predetermined processing sites for damaged DNA. J Cell Sci 119: 3284– 3295. Boisvert FM, Hendzel MJ, Bazett-Jones DP. 2000. Promyelocytic leukemia (PML) nuclear bodies are protein structures that do not accumulate RNA. J Cell Biol 148: 283–292. Boisvert FM, Kruhlak MJ, Box AK, Hendzel MJ, BazettJones DP. 2001. The transcription coactivator CBP is a dynamic component of the promyelocytic leukemia nuclear body. J Cell Biol 152: 1099–1106. Borden KL, Culjkovic B. 2009. Perspectives in PML: A unifying framework for PML function. Front Biosci 14: 497–509. Bridger J, Herrmann H, Munkel C, Lichter P. 1998. Identification of an interchromosomal compartment by polymerization of nuclear-targeted vimentin. J Cell Sci 111: 1241– 1253. Butler JT, Hall LL, Smith KP, Lawrence JB. 2009. Changing nuclear landscape and unique PML structures during early epigenetic transitions of human embryonic stem cells. J Cell Biochem 107: 609 –621. Carbone R, Pearson M, Minucci S, Pelicci PG. 2002. PML NBs associate with the hMre11 complex and p53 at sites of irradiation induced DNA damage. Oncogene 21: 1633– 1640. Chen YC, Kappel C, Beaudouin J, Eils R, Spector DL. 2008. Live cell dynamics of promyelocytic leukemia nuclear bodies upon entry into and exit from mitosis. Mol Biol Cell 19: 3147–3162. Chen GQ, Shi XG, Tang W, Xiong SM, Zhu J, Cai X, Han ZG, Ni JH, Shi GY, Jia PM, et al. 1997. Use of arsenic trioxide (As2O3) in the treatment of acute promyelocytic leukemia (APL): I. As2O3 exerts dose-dependent dual effects on APL cells. Blood 89: 3345–3353. Ching RW, Dellaire G, Eskiw CH, Bazett-Jones DP. 2005. PML bodies: A meeting place for genomic loci? J Cell Sci 118: 847– 854. Cho G, Lim Y, Golden JA. 2009. SUMO interaction motifs in SIZN1 are required for PML-NB localization and for

transcriptional activation. J Biol Chem 284: 19592– 19600. Cohen N, Sharma M, Kentsis A, Perez JM, Strudwick S, Borden KL. 2001. PML RING suppresses oncogenic transformation by reducing the affinity of eIF4E for mRNA. Embo J 20: 4547–4559. Condemine W, Takahashi Y, Le Bras M, de The H. 2007. A nucleolar targeting signal in PML-I addresses PML to nucleolar caps in stressed or senescent cells. J Cell Sci 120: 3219– 3227. Condemine W, Takahashi Y, Zhu J, Puvion-Dutilleul F, Guegan S, Janin A, de The H. 2006. Characterization of endogenous human promyelocytic leukemia isoforms. Cancer Res 66: 6192–6198. Culjkovic B, Topisirovic I, Skrabanek L, Ruiz-Gutierrez M, Borden KL. 2006. eIF4E is a central node of an RNA regulon that governs cellular proliferation. J Cell Biol 175: 415 –426. Daniel M-T, Koken M, Romagne´ O, Barbey S, Bazarbachi A, Stadler M, Guillemin M-C, Degos L, Chomienne C, de The´ H. 1993. PML protein expression in hematopoietic and acute promyelocytic leukemia cells. Blood 82: 1858–1867. de Stanchina E, Querido E, Narita M, Davuluri RV, Pandolfi PP, Ferbeyre G, Lowe SW. 2004. PML is a direct p53 target that modulates p53 effector functions. Mol Cell 13: 523 –535. de The´ G, Rivie`re M, Bernhard W. 1960. Examen au microscope e´lectronique de la tumeur VX2 du lapin domestique de´rive´e du papillome de Shope. Bull Cancer 47: 570 –584. Dellaire G, Bazett-Jones DP. 2004. PML nuclear bodies: Dynamic sensors of DNA damage and cellular stress. Bioessays 26: 963– 977. Dellaire G, Ching RW, Ahmed K, Jalali F, Tse KC, Bristow RG, Bazett-Jones DP. 2006a. Promyelocytic leukemia nuclear bodies behave as DNA damage sensors whose response to DNA double-strand breaks is regulated by NBS1 and the kinases ATM, Chk2, and ATR. J Cell Biol 175: 55–66. Dellaire G, Ching RW, Dehghani H, Ren Y, Bazett-Jones DP. 2006b. The number of PML nuclear bodies increases in early S phase by a fission mechanism. J Cell Sci 119: 1026–1033. Dellaire G, Eskiw CH, Dehghani H, Ching RW, Bazett-Jones DP. 2006c. Mitotic accumulations of PML protein contribute to the re-establishment of PML nuclear bodies in G1. J Cell Sci 119: 1034–1042. Draskovic I, Arnoult N, Steiner V, Bacchetti S, Lomonte P, Londono-Vallejo A. 2009. Probing PML body function in ALT cells reveals spatiotemporal requirements for telomere recombination. Proc Natl Acad Sci 106: 15726– 15731. Duprez E, Saurin AJ, Desterro JM, Lallemand-Breitenbach V, Howe K, Boddy MN, Solomon E, de The´ H, Hay RT, Freemont PS. 1999. SUMO-1 modification of the acute promyelocytic leukaemia protein PML: Implications for nuclear localisation. J Cell Sci 112: 381– 393. Dupuy-Coin AM, Bouteille M. 1972. Developmental pathway of granular and beaded nuclear bodies from nucleoli. J Ultrastruct Res 40: 55–67.

PML Nuclear Bodies Dupuy-Coin AM, Kalifat SR, Bouteille M. 1972. Nuclear bodies as proteinaceous structures containing ribonucleoproteins. J Ultrastruct Res 38: 174 –187. Dyck JA, Maul GG, Miller WH, Chen JD, Kakizuka A, Evans RM. 1994. A novel macromolecular structure is a target of the promyelocyte- retinoic acid receptor oncoprotein. Cell 76: 333– 343. Eladad S, Ye TZ, Hu P, Leversha M, Beresten S, Matunis MJ, Ellis NA. 2005. Intra-nuclear trafficking of the BLM helicase to DNA damage-induced foci is regulated by SUMO modification. Hum Mol Genet 14: 1351–1365. Engelhardt OG, Boutell C, Orr A, Ullrich E, Haller O, Everett RD. 2003. The homeodomain-interacting kinase PKM (HIPK-2) modifies ND10 through both its kinase domain and a SUMO-1 interaction motif and alters the posttranslational modification of PML. Exp Cell Res 283: 36–50. Engelhardt OG, Ullrich E, Kochs G, Haller O. 2001. Interferon-induced antiviral Mx1 GTPase is associated with components of the SUMO-1 system and promyelocytic leukemia protein nuclear bodies. Exp Cell Res 271: 286–295. Eskiw CH, Dellaire G, Bazett-Jones DP. 2004. Chromatin contributes to structural integrity of promyelocytic leukemia bodies through a SUMO-1-independent mechanism. J Biol Chem 279: 9577–9585. Eskiw CH, Dellaire G, Mymryk JS, Bazett-Jones DP. 2003. Size, position and dynamic behavior of PML nuclear bodies following cell stress as a paradigm for supramolecular trafficking and assembly. J Cell Sci 116: 4455– 4466. Everett RD. 2001. DNAviruses and viral proteins that interact with PML nuclear bodies. Oncogene 20: 7266– 7273. Everett RD. 2006. Interactions between DNA viruses, ND10 and the DNA damage response. Cell Microbiol 8: 365–374. Everett RD, Earnshaw WC, Pluta AF, Sternsdorf T, Ainsztein AM, Carmena M, Ruchaud S, Hsu WL, Orr A. 1999a. A dynamic connection between centromeres and ND10 proteins. J Cell Sci 112: 3443– 3454. Everett RD, Lomonte P, Sternsdorf T, van Driel R, Orr A. 1999b. Cell cycle regulation of PML modification and ND10 composition. J Cell Sci 112: 4581–4588. Everett RD, Murray J, Orr A, Preston CM. 2007. Herpes simplex virus type 1 genomes are associated with ND10 nuclear substructures in quiescently infected human fibroblasts. J Virol 81: 10991–11004. Everett RD, Parada C, Gripon P, Sirma H, Orr A. 2008. Replication of ICP0-null mutant herpes simplex virus type 1 is restricted by both PML and Sp100. J Virol 82: 2661– 2672. Everett RD, Rechter S, Papior P, Tavalai N, Stamminger T, Orr A. 2006. PML contributes to a cellular mechanism of repression of herpes simplex virus type 1 infection that is inactivated by ICP0. J Virol 80: 7995–8005. Fabunmi RP, Wigley WC, Thomas PJ, DeMartino GN. 2001. Interferon g regulates accumulation of the proteasome activator PA28 and immunoproteasomes at nuclear PML bodies. J Cell Sci 114: 29–36. Ferbeyre G, de Stanchina E, Querido E, Baptiste N, Prives C, Lowe SW. 2000. PML is induced by oncogenic ras and

promotes premature senescence. Genes Dev 14: 2015–2027. Fuchsova B, Novak P, Kafkova J, Hozak P. 2002. Nuclear DNA helicase II is recruited to IFN-a-activated transcription sites at PML nuclear bodies. J Cell Biol 158: 463 –473. Geoffroy MC, Hay RT. 2009. An additional role for SUMO in ubiquitin-mediated proteolysis. Nat Rev Mol Cell Biol 10: 564 –568. Gresko E, Ritterhoff S, Sevilla-Perez J, Roscic A, Frobius K, Kotevic I, Vichalkovski A, Hess D, Hemmings BA, Schmitz ML. 2009. PML tumor suppressor is regulated by HIPK2-mediated phosphorylation in response to DNA damage. Oncogene 28: 698 –708. Gro¨tzinger T, Jensen K, Will H. 1996. The interferon (IFN)stimulated gene Sp100 promoter contains an IFN-g activation site and an imperfect IFN-stimulated response element which mediate type I IFN inducibility. J Biol Chem 271: 25253. Guiochon-Mantel A, Savouret JF, Quignon F, Delabre K, Milgrom E, de The´ H. 1995. Effect of PML and PML-RAR on the transactivation properties and subcellular distribution of steroid hormone receptors. Mol Endocrinol 9: 1791–1803. Guo A, Salomoni P, Luo J, Shih A, Zhong S, Gu W, Pandolfi PP. 2000. The function of PML in p53-dependent apoptosis. Nat Cell Biol 2: 730– 736. Gupta P, Ho PC, Huq MM, Ha SG, Park SW, Khan AA, Tsai NP, Wei LN. 2008. Retinoic acid-stimulated sequential phosphorylation, PML recruitment, and SUMOylation of nuclear receptor TR2 to suppress Oct4 expression. Proc Natl Acad Sci 105: 11424–11429. Gurrieri C, Capodieci P, Bernardi R, Scaglioni PP, Nafa K, Rush LJ, Verbel DA, Cordon-Cardo C, Pandolfi PP. 2004. Loss of the tumor suppressor PML in human cancers of multiple histologic origins. J Natl Cancer Inst 96: 269 –279. Hakli M, Karvonen U, Janne OA, Palvimo JJ. 2005. SUMO-1 promotes association of SNURF (RNF4) with PML nuclear bodies. Exp Cell Res 304: 224 –233. Hay RT. 2005. SUMO: A history of modification. Mol Cell 18: 1– 12. Hayakawa F, Privalsky ML. 2004. Phosphorylation of PML by mitogen-activated protein kinases plays a key role in arsenic trioxide-mediated apoptosis. Cancer Cell 5: 389 –401. Hecker CM, Rabiller M, Haglund K, Bayer P, Dikic I. 2006. Specification of SUMO1- and SUMO2-interacting Motifs. J Biol Chem 281: 16117– 16127. Henderson BR, Eleftheriou A. 2000. A comparison of the activity, sequence specificity, and CRM1-dependence of different nuclear export signals. Exp Cell Res 256: 213 –224. Ishov AM, Sotnikov AG, Negorev D, Vladimirova OV, Neff N, Kamitani T, Yeh E, Strauss J, Maul G. 1999. PML is critical for ND10 formation and recruits the PML-interacting protein Daxx to this nuclear structure when modified by SUMO-1. J Cell Biol 147: 221 –234. Ito K, Bernardi R, Pandolfi PP. 2009. A novel signaling network as a critical rheostat for the biology and

V. Lallemand-Breitenbach and H. de The´ maintenance of the normal stem cell and the cancerinitiating cell. Curr Opin Genet Dev 19: 51– 59. Ito K, Bernardi R, Morotti A, Matsuoka S, Saglio G, Ikeda Y, Rosenblatt J, Avigan DE, Teruya-Feldstein J, Pandolfi PP. 2008. PML targeting eradicates quiescent leukaemiainitiating cells. Nature 453: 1072– 1078. Janderova-Rossmeislova L, Novakova Z, Vlasakova J, Philimonenko V, Hozak P, Hodny Z. 2007. PML protein association with specific nucleolar structures differs in normal, tumor and senescent human cells. J Struct Biol 159: 56–70. Jegou T, Chung I, Heuvelman G, Wachsmuth M, Gorisch SM, Greulich-Bode KM, Boukamp P, Lichter P, Rippe K. 2009. Dynamics of telomeres and promyelocytic leukemia nuclear bodies in a telomerase-negative human cell line. Mol Biol Cell 20: 2070–2082. Jensen K, Shiels C, Freemont PS. 2001. PML protein isoforms and the RBCC/TRIM motif. Oncogene 20: 7223– 7233. Jiang WQ, Zhong ZH, Henson JD, Reddel RR. 2007. Identification of candidate alternative lengthening of telomeres genes by methionine restriction and RNA interference. Oncogene 26: 4635–4647. Jul-Larsen A, Grudic A, Bjerkvig R, Boe SO. 2009. Cell-cycle regulation and dynamics of cytoplasmic compartments containing the promyelocytic leukemia protein and nucleoporins. J Cell Sci 122: 1201– 1210. Kastner P, Perez A, Lutz Y, Rochette-Egly C, Gaub M-P, Durand B, Lanotte M, Berger R, Chambon P. 1992. Structure, localization and transcriptional properties of two classes of retinoic acid receptor a fusion proteins in acute promyelocytic leukemia (APL): Structural similarities with a new family of oncoproteins. EMBO J 11: 629 –642. Kiesslich A, von Mikecz A, Hemmerich P. 2002. Cell cycledependent association of PML bodies with sites of active transcription in nuclei of mammalian cells. J Struct Biol 140: 167– 179. Kitagawa D, Kajiho H, Negishi T, Ura S, Watanabe T, Wada T, Ichijo H, Katada T, Nishina H. 2006. Release of RASSF1C from the nucleus by Daxx degradation links DNA damage and SAPK/JNK activation. EMBO J 25: 3286–3297. Kogan SC. 2009. Curing APL: Differentiation or destruction? Cancer Cell 15: 7– 8. Koken MHM, Linares-Cruz G, Quignon F, Viron A, Chelbi-Alix MK, Sobczak-The´pot J, Juhlin L, Degos L, Calvo F, de The´ H. 1995. The PML growth-suppressor has an altered expression in human oncogenesis. Oncogene 10: 1315–1324. Koken MHM, Puvion-Dutilleul F, Guillemin MC, Viron A, Linares-Cruz G, Stuurman N, de Jong L, Szostecki C, Calvo F, Chomienne C, et al. 1994. The t(15;17) translocation alters a nuclear body in a RA-reversible fashion. EMBO J 13: 1073– 1083. Kumar PP, Bischof O, Purbey PK, Notani D, Urlaub H, Dejean A, Galande S. 2007. Functional interaction between PML and SATB1 regulates chromatin-loop architecture and transcription of the MHC class I locus. Nat Cell Biol 9: 45–56. Kyratsous CA, Silverstein SJ. 2009. Components of nuclear domain 10 bodies regulate varicella-zoster virus replication. J Virol 83: 4262– 4274.

Lafarga M, Berciano MT, Pena E, Mayo I, Castano JG, Bohmann D, Rodrigues JP, Tavanez JP, Carmo-Fonseca M. 2002. Clastosome: A subtype of nuclear body enriched in 19S and 20S proteasomes, ubiquitin, and protein substrates of proteasome. Mol Biol Cell 13: 2771–2782. Lai HK, Borden KL. 2000. The promyelocytic leukemia (PML) protein suppresses cyclin D1 protein production by altering the nuclear cytoplasmic distribution of cyclin D1 mRNA. Oncogene 19: 1623– 1634. Lallemand-Breitenbach V, Jeanne M, Benhenda S, Nasr R, Lei M, Peres L, Zhou J, Zhu J, Raught B, de The´ H. 2008. Arsenic degrades PML or PML-RARa through a SUMO-triggered RNF4/ubiquitin-mediated pathway. Nat Cell Biol 10: 547 –555. Lallemand-Breitenbach V, Zhu J, Puvion F, Koken M, Honore N, Doubeikovsky A, Duprez E, Pandolfi PP, Puvion E, Freemont P, et al. 2001. Role of promyelocytic leukemia (PML) sumolation in nuclear body formation, 11S proteasome recruitment, and As(2)O(3)-induced PML or PML/retinoic acid receptor a degradation. J Exp Med 193: 1361– 1372. LaMorte VJ, Dyck JA, Ochs RL, Evans RM. 1998. Localization of nascent RNA and CREB binding protein with the PML-containing nuclear body. Proc Natl Acad Sci 95: 4991–4996. Lehembre F, Muller S, Pandolfi PP, Dejean A. 2001. Regulation of Pax3 transcriptional activity by SUMO-1modified PML. Oncogene 20: 1– 9. Li W, Ferguson BJ, Khaled WT, Tevendale M, Stingl J, Poli V, Rich T, Salomoni P, Watson CJ. 2009. PML depletion disrupts normal mammary gland development and skews the composition of the mammary luminal cell progenitor pool. Proc Natl Acad Sci 106: 4725– 4730. Li H, Leo C, Zhu J, Wu X, O’Neil J, Park EJ, Chen JD. 2000. Sequestration and inhibition of daxx-mediated transcriptional repression by PML. Mol Cell Biol 20: 1784–1796. Lin HK, Bergmann S, Pandolfi PP. 2004. Cytoplasmic PML function in TGF-b signalling. Nature 431: 205–211. Lin DY, Huang YS, Jeng JC, Kuo HY, Chang CC, Chao TT, Ho CC, Chen YC, Lin TP, Fang HI, et al. 2006. Role of SUMO-interacting motif in Daxx SUMO modification, subnuclear localization, and repression of sumoylated transcription factors. Mol Cell 24: 341–354. Lin DY, Lai MZ, Ann DK, Shih HM. 2003. Promyelocytic leukemia protein (PML) functions as a glucocorticoid receptor co-activator by sequestering Daxx to the PML oncogenic domains (PODs) to enhance its transactivation potential. J Biol Chem 278: 15958–15965. Luciani JJ, Depetris D, Usson Y, Metzler-Guillemain C, Mignon-Ravix C, Mitchell MJ, Megarbane A, Sarda P, Sirma H, Moncla A, et al. 2006. PML nuclear bodies are highly organised DNA-protein structures with a function in heterochromatin remodelling at the G2 phase. J Cell Sci 119: 2518– 2531. Mallette FA, Goumard S, Gaumont-Leclerc MF, Moiseeva O, Ferbeyre G. 2004. Human fibroblasts require the Rb family of tumor suppressors, but not p53, for PMLinduced senescence. Oncogene 23: 91– 99. Matunis MJ, Zhang XD, Ellis NA. 2006. SUMO: The glue that binds. Dev Cell 11: 596 –597.

PML Nuclear Bodies Maul GG, Negorev D, Bell P, Ishov AM. 2000. Review: Properties and assembly mechanisms of ND10, PML bodies, or PODs. J Struct Biol 129: 278 –287. Meinecke I, Cinski A, Baier A, Peters MA, Dankbar B, Wille A, Drynda A, Mendoza H, Gay RE, Hay RT, et al. 2007. Modification of nuclear PML protein by SUMO-1 regulates Fas-induced apoptosis in rheumatoid arthritis synovial fibroblasts. Proc Natl Acad Sci 104: 5073– 5078. Meroni G, Diez-Roux G. 2005. TRIM/RBCC, a novel class of ‘single protein RING finger’ E3 ubiquitin ligases. Bioessays 27: 1147– 1157. Minty A, Dumont X, Kaghad M, Caput D. 2000. Covalent modification of p73a by SUMO-1. Two-hybrid screening with p73 identifies novel SUMO-1-interacting proteins and a SUMO-1 interaction motif. J Biol Chem 275: 36316–36323. Molenaar C, Wiesmeijer K, Verwoerd NP, Khazen S, Eils R, Tanke HJ, Dirks RW. 2003. Visualizing telomere dynamics in living mammalian cells using PNA probes. EMBO J 22: 6631–6641. Muller S, Matunis MJ, Dejean A. 1998. Conjugation with the ubiquitin-related modifier SUMO-1 regulates the partitioning of PML within the nucleus. EMBO J 17: 61– 70. Muratani M, Gerlich D, Janicki SM, Gebhard M, Eils R, Spector DL. 2002. Metabolic-energy-dependent movement of PML bodies within the mammalian cell nucleus. Nat Cell Biol 4: 106– 110. Nacerddine K, Lehembre F, Bhaumik M, Artus J, CohenTannoudji M, Babinet C, Pandolfi PP, Dejean A. 2005. The SUMO pathway is essential for nuclear integrity and chromosome segregation in mice. Dev Cell 9: 769–779. Nefkens I, Negorev DG, Ishov AM, Michaelson JS, Yeh ET, Tanguay RM, Muller WE, Maul GG. 2003. Heat shock and Cd2þ exposure regulate PML and Daxx release from ND10 by independent mechanisms that modify the induction of heat-shock proteins 70 and 25 differently. J Cell Sci 116: 513– 524. Negorev D, Maul GG. 2001. Cellular proteins localized at and interacting within ND10/PML nuclear bodies/ PODs suggest functions of a nuclear depot. Oncogene 20: 7234–7242. Park SW, Hu X, Gupta P, Lin YP, Ha SG, Wei LN. 2007. SUMOylation of Tr2 orphan receptor involves Pml and fine-tunes Oct4 expression in stem cells. Nat Struct Mol Biol 14: 68–75. Pearson M, Carbone R, Sebastiani C, Cioce M, Fagioli M, Saito S, Higashimoto Y, Appella E, Minucci S, Pandolfi PP, et al. 2000. PML regulates p53 acetylation and premature senescence induced by oncogenic Ras. Nature 406: 207–210. Potts PR, Yu H. 2007. The SMC5/6 complex maintains telomere length in ALT cancer cells through SUMOylation of telomere-binding proteins. Nat Struct Mol Biol 14: 581–590. Puvion-Dutilleul F, Chelbi-Alix MK, Koken M, Quignon F, Puvion E, de The´ H. 1995. Adenovirus infection induces rearrangements in the intranuclear distribution of the nuclear body-associated PML protein. Exp Cell Res 218: 9– 16. Qin Q, Inatome R, Hotta A, Kojima M, Yamamura H, Hirai H, Yoshizawa T, Tanaka H, Fukami K, Yanagi S. 2006. A

novel GTPase, CRAG, mediates promyelocytic leukemia protein-associated nuclear body formation and degradation of expanded polyglutamine protein. J Cell Biol 172: 497 –504. Quignon F, Chen Z, de The´ H. 1997. Retinoic acid and arsenic: towards oncogene targeted treatments of acute promyelocytic leukaemia. Biochim Biophys Acta 1333: M53– M61. Quimby BB, Yong-Gonzalez V, Anan T, Strunnikov AV, Dasso M. 2006. The promyelocytic leukemia protein stimulates SUMO conjugation in yeast. Oncogene 25: 2999–3005. Rao VA, Fan AM, Meng L, Doe CF, North PS, Hickson ID, Pommier Y. 2005. Phosphorylation of BLM, dissociation from topoisomerase IIIa, and colocalization with g-H2AX after topoisomerase I-induced replication damage. Mol Cell Biol 25: 8925– 8937. Regad T, Bellodi C, Nicotera P, Salomoni P. 2009. The tumor suppressor Pml regulates cell fate in the developing neocortex. Nat Neurosci 12: 132– 140. Reymond A, Meroni G, Fantozzi A, Merla G, Cairo S, Luzi L, Riganelli D, Zanaria E, Messali S, Cainarca S, et al. 2001. The tripartite motif family identifies cell compartments. Embo J 20: 2140–2151. Russell RA, Adams NM, Stephens DA, Batty E, Jensen K, Freemont PS. 2009. Segmentation of fluorescence microscopy images for quantitative analysis of cell nuclear architecture. Biophys J 96: 3379–3389. Salomoni P, Khelifi AF. 2006. Daxx: Death or survival protein? Trends Cell Biol 16: 97–104. Salomoni P, Pandolfi PP. 2002. The role of PML in tumor suppression. Cell 108: 165– 170. Scaglioni PP, Yung TM, Cai LF, Erdjument-Bromage H, Kaufman AJ, Singh B, Teruya-Feldstein J, Tempst P, Pandolfi PP. 2006. A CK2-dependent mechanism for degradation of the PML tumor suppressor. Cell 126: 269– 283. Shao W, Fanelli M, Ferrara FF, Riccioni R, Rosenauer A, Davison K, Lamph WW, Waxman S, Pelicci PG, Lo Coco F, et al. 1998. Arsenic trioxide as an inducer of apoptosis and loss of PML/RARa protein in acute promyelocytic leukemia cells. J Natl Cancer Inst 90: 124– 133. Shav-Tal Y, Blechman J, Darzacq X, Montagna C, Dye BT, Patton JG, Singer RH, Zipori D. 2005. Dynamic sorting of nuclearcomponents into distinct nucleolar caps during transcriptional inhibition. Mol Biol Cell 16: 2395– 2413. Shen TH, Lin HK, Scaglioni PP, Yung TM, Pandolfi PP. 2006. The mechanisms of PML-nuclear body formation. Mol Cell 24: 331–339. Shiels C, Islam SA, Vatcheva R, Sasieni P, Sternberg MJ, Freemont PS, Sheer D. 2001. PML bodies associate specifically with the MHC gene cluster in interphase nuclei. J Cell Sci 114: 3705– 3716. Smith KP, Byron M, O’Connell BC, Tam R, Schorl C, Guney I, Hall LL, Agrawal P, Sedivy JM, Lawrence JB. 2004. c-Myc localization within the nucleus: evidence for association with the PML nuclear body. J Cell Biochem 93: 1282–1296. Song MS, Salmena L, Carracedo A, Egia A, Lo-Coco F, Teruya-Feldstein J, Pandolfi PP. 2008. The deubiquitinylation and localization of PTEN are regulated by a HAUSP-PML network. Nature 455: 813– 817.

V. Lallemand-Breitenbach and H. de The´ St-Germain JR, Chen J, Li Q. 2008. Involvement of PML nuclear bodies in CBP degradation through the ubiquitin-proteasome pathway. Epigenetics 3: 342– 349. Stadler M, Chelbi-Alix MK, Koken MHM, Venturini L, Lee C, Saı¨b A, Quignon F, Pelicano L, Guillemin M-C, Schindler C, et al. 1995. Transcriptional induction of the PML growth suppressor gene by interferons is mediated through an ISRE and a GAS element. Oncogene 11: 2565–2573. Stuurman N, de Graaf A, Floore A, Josso A, Humbel B, de Jong L, van Driel R. 1992. A monoclonal antibody recognizing nuclear matrix-associated nuclear bodies. J Cell Sci 101: 773– 784. Stuurman N, Meijne AML, van der Pol AJ, de Jong L, van Driel R, van Renswoude J. 1990. The nuclear matrix from cells of different origin. Evidence for a common set of matrix proteins. J Biol Chem 265: 5460–5465. Sun J, Xu H, Subramony SH, Hebert MD. 2005. Interactions between coilin and PIASy partially link Cajal bodies to PML bodies. J Cell Sci 118: 4995–5003. Szostecki C, Guldner H, Netter H, Will H. 1990. Isolation and characterization of cDNA encoding a human nuclear antigen predominantly rrecognized by autoantibodies of patients with primary biliary cirrhosis. J Immunol 145: 4338– 4347. Takahashi H, Hatakeyama S, Saitoh H, Nakayama KI. 2005. Noncovalent SUMO-1 binding activity of thymine DNA glycosylase (TDG) is required for its SUMO-1 modification and colocalization with the promyelocytic leukemia protein. J Biol Chem 280: 5611–5621. Takahashi Y, Lallemand-Breitenbach V, Zhu J, de The´ H. 2004. PML nuclear bodies and apoptosis. Oncogene 23: 2819– 2824. Tao H, Simmons BN, Singireddy S, Jakkidi M, Short KM, Cox TC, Massiah MA. 2008. Structure of the MID1 tandem B-boxes reveals an interaction reminiscent of intermolecular ring heterodimers. Biochemistry 47: 2450– 2457. Tatham MH, Geoffroy MC, Shen L, Plechanovova A, Hattersley N, Jaffray EG, Palvimo JJ, Hay RT. 2008. RNF4 is a poly-SUMO-specific E3 ubiquitin ligase required for arsenic-induced PML degradation. Nat Cell Biol 10: 538–546. Tavalai N, Papior P, Rechter S, Leis M, Stamminger T. 2006. Evidence for a role of the cellular ND10 protein PML in mediating intrinsic immunity against human cytomegalovirus infections. J Virol 80: 8006–8018. Tavalai N, Papior P, Rechter S, Stamminger T. 2008. Nuclear domain 10 components promyelocytic leukemia protein and hDaxx independently contribute to an intrinsic antiviral defense against human cytomegalovirus infection. J Virol 82: 126–137. Terris B, Baldin V, Dubois S, Degott C, Flejou JF, Henin D, Dejean A. 1995. PML nuclear bodies are general targets for inflammation and cell proliferation. Cancer Res 55: 1590– 1597. Torii S, Egan DA, Evans RA, Reed JC. 1999. Human Daxx regulates Fas-induced apoptosis from nuclear PML oncogenic domains (PODs). EMBO J 18: 6037– 6049. Trotman LC, Alimonti A, Scaglioni PP, Koutcher JA, Cordon-Cardo C, Pandolfi PP. 2006. Identification of a

tumour suppressor network opposing nuclear Akt function. Nature 441: 523 –527. Tsukamoto T, Hashiguchi N, Janicki SM, Tumbar T, Belmont AS, Spector DL. 2000. Visualization of gene activity in living cells. Nat Cell Biol 2: 871 –878. Vagner-Capodano AM, Bouteille M, Stahl A. 1982. Nucleolar ribonucleoprotein release into the nucleoplasm as nuclear bodies in cultured thyrotropin-stimulated thyroid cells: Autoradiographic kinetics. J Ultrastruct Res 78: 13–25. Villagra NT, Navascues J, Casafont I, Val-Bernal JF, Lafarga M, Berciano MT. 2006. The PML-nuclear inclusion of human supraoptic neurons: A new compartment with SUMO-1- and ubiquitin-proteasome-associated domains. Neurobiol Dis 21: 181– 193. Wang Z-G, Ruggero D, Ronchetti S, Zhong S, Gaboli M, Rivi R, Pandolfi PP. 1998. PML is essential for multiple apoptotic pathways. Nature Genet 20: 266 –272. Wang J, Shiels C, Sasieni P, Wu PJ, Islam SA, Freemont PS, Sheer D. 2004. Promyelocytic leukemia nuclear bodies associate with transcriptionally active genomic regions. J Cell Biol 164: 515– 526. Weidtkamp-Peters S, Lenser T, Negorev D, Gerstner N, Hofmann TG, Schwanitz G, Hoischen C, Maul G, Dittrich P, Hemmerich P. 2008. Dynamics of component exchange at PML nuclear bodies. J Cell Sci 121: 2731–2743. Weis K, Rambaud S, Lavau C, Jansen J, Carvalho T, Carmo-Fonseca M, Lamond A, Dejean A. 1994. Retinoic acid regulates aberrant nuclear localization of PML/ RARa in acute promyelocytic leukemia cells. Cell 76: 345 –356. Wiesmeijer K, Molenaar C, Bekeer IM, Tanke HJ, Dirks RW. 2002. Mobile foci of Sp100 do not contain PML: PML bodies are immobile but PML and Sp100 proteins are not. J Struct Biol 140: 180– 188. Wu G, Lee WH, Chen PL. 2000. NBS1 and TRF1 colocalize at promyelocytic leukemia bodies during late S/G2 phases in immortalized telomerase-negative cells. Implication of NBS1 in alternative lengthening of telomeres. J Biol Chem 275: 30618– 30622. Wu G, Jiang X, Lee WH, Chen PL. 2003. Assembly of functional ALT-associated promyelocytic leukemia bodies requires Nijmegen Breakage Syndrome 1. Cancer Res 63: 2589–2595. Yamada M, Sato T, Shimohata T, Hayashi S, Igarashi S, Tsuji S, Takahashi H. 2001a. Interaction between neuronal intranuclear inclusions and promyelocytic leukemia protein nuclear and coiled bodies in CAG repeat diseases. Am J Pathol 159: 1785– 1795. Yamada M, Wood JD, Shimohata T, Hayashi S, Tsuji S, Ross CA, Takahashi H. 2001b. Widespread occurrence of intranuclear atrophin-1 accumulation in the central nervous system neurons of patients with dentatorubralpallidoluysian atrophy. Ann Neurol 49: 14– 23. Yang S, Jeong JH, Brown AL, Lee CH, Pandolfi PP, Chung JH, Kim MK. 2006. Promyelocytic leukemia activates Chk2 by mediating Chk2 autophosphorylation. J Biol Chem 281: 26645–26654. Yang S, Kuo C, Bisi JE, Kim MK. 2002. PML-dependent apoptosis after DNA damage is regulated by the checkpoint kinase hCds1/Chk2. Nat Cell Biol 4: 865 –870.

PML Nuclear Bodies Ye X, Zerlanko B, Kennedy A, Banumathy G, Zhang R, Adams PD. 2007a. Downregulation of Wnt signaling is a trigger for formation of facultative heterochromatin and onset of cell senescence in primary human cells. Mol Cell 27: 183–196. Ye X, Zerlanko B, Zhang R, Somaiah N, Lipinski M, Salomoni P, Adams PD. 2007b. Definition of pRB- and p53-dependent and -independent steps in HIRA/ASF1amediated formation of senescence-associated heterochromatin foci. Mol Cell Biol 27: 2452–2465. Zhang R, Poustovoitov MV, Ye X, Santos HA, Chen W, Daganzo SM, Erzberger JP, Serebriiskii IG, Canutescu AA, Dunbrack RL, et al. 2005. Formation of MacroH2Acontaining senescence-associated heterochromatin foci and senescence driven by ASF1a and HIRA. Dev Cell 8: 19–30. Zhong S, Hu P, Ye TZ, Stan R, Ellis NA, Pandolfi PP. 1999. A role for PML and the nuclear body in genomic stability. Oncogene 18: 7941–7947.

Zhong S, Muller S, Ronchetti S, Freemont PS, Dejean A, Pandolfi PP. 2000a. Role of SUMO-1-modified PML in nuclear body formation. Blood 95: 2748– 2752. Zhong S, Salomoni P, Ronchetti S, Guo A, Ruggero D, Pandolfi PP. 2000b. Promyelocytic leukemia protein (PML) and Daxx participate in a novel nuclear pathway for apoptosis. J Exp Med 191: 631– 639. Zhu J, Lallemand-Breitenbach V, de The´ H. 2001. Pathways of retinoic acid- or arsenic trioxide-induced PML/RARa catabolism, role of oncogene degradation in disease remission. Oncogene 20: 7257–7265. Zhu J, Chen Z, Lallemand-Breitenbach V, de The´ H. 2002. How acute promyelocytic leukemia revived arsenic. Nature Reviews on Cancer 2: 705–713. Zhu J, Koken MHM, Quignon F, Chelbi-Alix MK, Degos L, Wang ZY, Chen Z, de The´ H. 1997. Arsenic-induced PML targeting onto nuclear bodies: implications for the treatment of acute promyelocytic leukemia. Proc Natl Acad Sci 94: 3978–3983.

The Perinucleolar Compartment Callie Pollock and Sui Huang Northwestern University Feinberg School of Medicine, Chicago, Illinois 60611 Correspondence: [email protected]

The perinucleolar compartment (PNC) is a subnuclear body characterized by its location to the periphery of the nucleolus. The PNC is a dynamic structure and is highly enriched in RNA-binding proteins and pol III RNA. The structural stability of the PNC is dependent on continuous pol III transcription and the presence of key proteins. The PNC is associated with malignancy both in vitro and in vivo and its presence positively correlates with metastatic capacity, making it a potential cancer marker. Recent studies also suggest an association between the PNC and a specific DNA locus, and ongoing PNC research continues to focus on determining the structure and function of the PNC to understand its role in cancer. This article summarizes the current understanding of PNC structure and function with an emphasis on the association of PNC and malignancy.

he cell nucleus is a complex and highly organized organelle that is responsible for many critical cellular functions, including housing and replicating DNA, transcribing DNA into RNA, RNA processing and splicing, macromolecular trafficking between the nucleus and cytoplasm, and regulating gene expression. Increasing evidence shows that these functions are not randomly distributed in the nucleus, but rather are highly compartmentalized into a higher order three-dimensional organization. Such organization is represented by nonmembrane-bound nuclear domains and nuclear bodies, which can be identified and visualized through various molecular probes and resolved by microscopy. The compartmentalization within the nucleus may reflect the highly efficient spatial and temporal organization of essential molecular complexes for specific functions (reviewed in Zimber et al. 2004).

T

Nucleoli, Cajal bodies (Morris 2008), promyelocytic leukemia (PML) bodies (Bernardi and Pandolfi 2007), nuclear speckles (Lamond and Spector 2003), and perinucleolar compartments (PNCs) are among the many nuclear compartments that have distinct nuclear localization patterns. As ongoing research examines the structure and function of these bodies, their functional relevance is being revealed. For example, nucleoli are involved in the biogenesis of ribosomes; nuclear speckles are involved in pre-mRNA processing and metabolism; Cajal bodies are indicated in the assembly and trafficking of snRNP, snoRNP, and telomere functions; and PML as well as PNC are associated with malignancy. This article summarizes the current understanding of the structure and function of one nuclear body, the PNC, and its role in the malignant phenotype.

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000679 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000679

C. Pollock and S. Huang

PNC STRUCTURE

The perinucleolar compartment (PNC) is a unique subnuclear structure that was originally described during the characterization of the hnRNP I/PTB ( polypyrimidine tract binding) protein, in which the protein was shown to localize at a perinucleolar focus (Ghetti et al. 1992). The PNC is irregularly shaped, ranging from .25 to 4 mm in length, and is associated with the periphery of the nucleolus (Matera et al. 1995; Huang et al. 1997). Electron microscopic analysis of HeLa cells reveals that the PNC is composed of multiple thick, electron dense strands, each measuring approximately 80 – 180 nm in diameter. The PNC is physically associated with the nucleolus while remaining structurally distinct (Fig. 1) (Huang et al. 1997). A three-dimensional computer reconstruction of electron microscopic images from serially sectioned HeLa cell nuclei shows that the PNC forms a reticulated mesh on the nucleolar surface (Huang et al. 1998). Time lapse microscopy of cells expressing green fluorescent protein-tagged polypyrimidine tract-binding protein (GFP-PTB) shows that the PNC is a dynamic structure that moves discretely along the nucleolar periphery over time (Table 1) (Huang et al. 1997). PNC structure is maintained throughout interphase, disassembled during mitosis in conjunction with the disassociation of nucleoli, and is reassembled at late telophase as nucleoli begin to assemble. The PNC is commonly heritable from mother to daughter cells, and the PNCs

PTB

CUG-BP

Overlay

Dapi

of daughter cells are of similar shape, size, and number (Huang et al. 1997). MOLECULAR COMPONENTS OF THE PNC

Although the composition of the PNC has not been fully characterized, a number of proteins and RNAs show enrichment in the PNC. All known PNC-associated proteins are RNAbinding proteins that are implicated in pol II RNA metabolism. The proteins include CUG binding protein (CUG-BP) (Timchenko et al. 1996), Polypyrimidine tract-binding protein (PTB) (Ghetti et al. 1992), KH-type splicing regulatory protein (KSRP) (Hall et al. 2004), Raver1 (Huttelmaier et al. 2001), Raver2 (Kleinhenz et al. 2005), Rod1, and nucleolin (Kopp and Huang 2005). Interestingly, the RNAs identified in the PNC so far are small, noncoding RNAs transcribed by pol III, which is surprising because pol II RNAs commonly interact with the PNC proteins. RNAs that show enrichment in the PNC include RNase mitochondrial processing (MRP) RNA, ribonuclease P (RNase P) RNA, hY (1,2,5) RNA (Matera et al. 1995), Alu RNA, and signal recognition particle (SRP) (7SL) RNA (Wang et al. 2003). PNC-associated Proteins

Proteins of diverse cellular functions are enriched in the PNC. What these proteins have in common is their primary implication in the processing of pol II RNAs. For example, the first protein identified in the PNC, PTB, is involved

Nucleolus PNC

Figure 1. PNC localization. A specific PTB monoclonal antibody, SH54, immunolabels the perinucleolar

compartment (PNC) in green, and CUG-BP is immunolabeled in red. In the far right panel, an electron microscopic image shows the PNC at the periphery of the nucleolus. Bar, 30 mm.

The PNC Table 1. Protein and RNA components of the PNC Component Name

Protein components CUG-BP KSRP Nucleolin PTB Raver1/2 ROD1 RNA Components Alu RNA hY RNA MRP RNA RNase P RNA SRP RNA

Brief Description

Binds polyadenylated RNA and is involved in alternative splicing, deadenylation, and mytonic dystrophy (Timchenko et al. 1996; Savkur et al. 2001; Ho et al. 2005) Involved in alternative splicing, splicing regulation, and RNA decay (Min et al. 1997; Nechama et al. 2008) Enriched in the nucleolus and PNC, plays a role in rDNA transcription and RNA processing (Ginisty et al. 1999; Tuteja and Tuteja 1998) Protein involved in pre-mRNA splicing, alternative splicing, RNA polyadenylation, and translation (Lou et al. 1996; Stoneley and Willis 2004; Wagner and Garcia-Blanco 2001) RNA binding, nuclear RNPs that interact with PTB (Kleinhenz et al. 2005) Shares homology with PTB and is a homolog to yeast regulator of differentiation (Yamamoto et al. 1999) Forms Alu complex that is implicated in regulation of transcription and translation (Hasler and Strub 2006a; Hasler and Strub 2006b) Interacts with Ro protein to form Ro ribonucleoproteins (RNPs) (Wolin and Steitz 1984) RNA portion of the MRP RNP, which plays a role in mitochondrial DNA replication and preribosomal RNA processing (van Eenennaam et al. 2000) RNA component of RNase P complex, involved in maturation of 50 end of tRNA (van Eenennaam et al. 2000) Part of the signal recognition particle, implicated in secretory protein transport and elongation arrest (Wolin and Walter 1989)

in pre-mRNA splicing (Wagner and GarciaBlanco 2001), alternative splicing, and regulation of RNA polyadenylation (Lou et al. 1996) and translation (Stoneley and Willis 2004). Another protein enriched in the PNC, CUG-BP, is known for its role in myotonic dystrophy neuromuscular disease, and it is also involved in alternative splicing and deadenylation (Timchenko et al. 1996; Savkur et al. 2001; Ho et al. 2005). KSRP has been implicated in alternative splicing and regulation of splicing (Min et al. 1997; Nechama et al. 2008). The enrichment of RNA processing proteins in the PNC suggests a possible role for the PNC in RNA metabolism. However, not all pre-mRNA processing factors, such as SR proteins, are enriched in the PNC. In addition, many attempts to localize pre- or mRNA have failed to observe any enrichment of pol II RNA in the PNC. Although the PNC is physically associated with the nucleolus, many nucleolar proteins are not enriched in the PNC. The one nucleolar protein found enriched in the PNC is nucleolin (Kopp and Huang 2005).

PNC-associated RNAs

A subset of pol III transcripts are enriched in the PNC. RNase P and MRP RNA (Matera et al. 1995) are two highly conserved components of sequence-specific endoribonucleases. As part of the endoribonuclease complexes, RNase P is involved in tRNA maturation, and RNase MRP is implicated in pre-ribosomal RNA processing and mitochondrial DNA replication (van Eenennaam et al. 2000). Furthermore, RNase P and MRP RNA are structurally related to each other and share protein binding partners in their functional endoribonuclease complexes. It is not known if this similarity is involved in the coenrichment of these RNAs to the PNC. Other PNC-enriched RNAs include hY RNA, an abundant RNA that associates with the Ro protein and may be involved in DNA replication (Wolin and Steitz 1984; Christov et al. 2006), and SRP RNA, an RNA component of the signal recognition particle, which functions in secretory protein transport and elongation arrest (Wolin and Walter 1989; Grudnik et al. 2009).

C. Pollock and S. Huang

Alu RNA also localizes to the PNC and is known to associate with the signal recognition particle (Hasler and Strub 2006b). Although only pol III transcripts have been detected in the PNC, not all pol III transcripts are enriched in the PNC. In situ hybridization of many pol III transcripts including U6, tRNA, and 7SK do not show localization to the PNC (Matera et al. 1995; Pollock and Huang 2009). Formation of the PNC

The PNC is enriched with pol III transcribed RNAs and RNA binding proteins that are primarily implicated in the metabolism of pol II RNAs. The enrichment of the RNA and proteins in the PNC does not appear to be due to overexpression of these components because overexpression of the protein components (PTB or CUG-BP) alone or with PNC-associated RNAs (MRP RNA and RNase P RNA) does not induce the formation of the PNC in primary or immortalized cells (Kopp and Huang 2005). Many of the molecular components of the PNC have been identified and the interactions amongst the known PNC-associated components are beginning to be investigated. It is known that some of the PNC-associated proteins interact with one another and the pol III RNAs that enrich in the PNC. For example, PTB interacts with Raver1 (Huttelmaier et al. 2001), nucleolin (Singh et al. 2004), and some of the hY RNAs (Fabini et al. 2001). It is not known, however, if these interactions occur within the PNC or elsewhere in the cell. PNC AND RNA TRANSCRIPTION

RNA is crucial to PNC stability. This importance of RNA was first noticed when RNase, but not DNase, treatment eliminated PNC in permeabilized cells (Huang et al. 1998). A 5 minute pulse-labeling of Br-U showed that a majority of PNC-associated RNA is newly synthesized (Huang et al. 1998). To determine the origin of PNC-enriched RNA, pol I, II, or III transcriptional activities were selectively inhibited. Initial experiments using actinomycin D at a low concentration that preferentially

inhibits pol I transcription showed that PNC structure is disrupted by the treatment. However, a later experiment using cycloheximide treatment, a protein translation inhibitor that also inhibits pol I transcription (5 h at 100 mg/mL) (Higashi et al. 1968), did not induce PNC disassembly or affect labeled nucleotide enrichment in the PNC, demonstrating that pol I transcription is not important to PNC structure. The sensitivity of the PNC to actinomycin D is most likely because of the multiple modes of action of the drug, including its DNA interaction, which directly affects PNC structure (see below). Additionally, pol I transcripts (i.e., 28S and 18S RNAs) or prerRNA processing factors, such as fibrillarin, are not detected in the PNC (Matera et al. 1995; Huang et al. 1997; Kopp and Huang 2005). These results suggest that PNC-enriched RNAs are not likely to be of pol I origin. Nor is it likely that the primary source of PNCenriched RNA is from RNA pol II because treatment of a-amanitin, a drug that selectively and specifically inhibits pol II polymerase, does not induce PNC dissociation (Huang et al. 1998). In addition, in situ hybridization to several pre-mRNAs did not show coenrichment of these RNA to the PNC (Hall et al. 2004). However, it can not be excluded, at this point, that a small subset of pol I and II transcripts may associate with the PNC. The structural integrity of the PNC is dependent on pol III transcription. Injection of tagetin, a specific inhibitor of polymerase III, causes disassembly of the PNC within 2 hours (Wang et al. 2003). Furthermore, the continuous production of pol III transcripts, not just pol III activity, is important for PNC structure. Overexpression of RNase MRP RNA from a pol II promoter was able to partially overcome the PNC instability caused by pol III inhibition (Wang et al. 2003). PNC-enriched pol III RNAs are most likely newly transcribed. A 5 minute pulse-label with BrU shows that newly synthesized RNAs are coenriched in the PNC with the RNA binding proteins (Huang et al. 1998). In addition, during initial pol III inhibition, PNC-associated MRP RNA rapidly dissociates, whereas mature pol III RNAs in

The PNC

their functional complexes remain unaltered. For example, nucleolar-associated functional MRP RNA remains in the nucleolus at similar time points (Wang et al. 2003). These results reveal that the pol III transcripts enriched in the PNC are not likely to be in their mature, functional complexes, and these RNAs are necessary for the maintenance of PNC structure. Although the PNC is enriched with newly transcribed pol III RNAs, it is not the site of transcription for these RNAs. In situ hybridization of genes encoding four PNC-associated RNAs shows no spatial proximity to the PNC (Matera et al. 1995; Kopp and Huang 2005). In addition, it is also unlikely that the PNC acts as an assembly point for the newly transcribed RNAs into their respective RNP complexes, as the protein subunits of these complexes are not detected to be similarly enriched in the PNC (Hall et al. 2004; Kopp and Huang 2005). Although the specific function of the PNC remains unidentified, the enrichment of newly synthesized pol III RNAs and RNA binding proteins suggests a role in RNA metabolism. Several potential functions of the PNC can be envisioned. First, the PNC may represent a transitional depot between the newly synthesized pol III RNA and the assembly of their final functional complex. Second, the PNC may be a site of RNA processing. Third, the PNC could act as a site of assembly for RNA trafficking complexes for movement to their functional destinations, including those in the nucleus or in the cytoplasm. Fourth, the PNC may be a site for degradation of excess pol III RNAs that are produced because of the pol III deregulation often observed in cancer cells. Because live cell studies have shown that PTB and CUG-BP shuttle in and out of PNCs rapidly (Huang et al. 1997), the association of these RNAs with the PNC is most likely dynamic rather than simple aggregation or long-term storage. PNC ASSOCIATION WITH DNA

Initial characterization of the PNC showed that daughter cells inherit PNCs often arranged as mirror images from their parent cells (Huang et al. 1997). This heritable nature of the PNC

suggested a connection between the structure and DNA, and prompted further investigation into the association. More recent chemical and cell biology experiments show that PNC stability is reliant on DNA integrity, and the PNC is associated with a DNA locus. Chemical Biology Evidence

To better understand the structure and function of the PNC, a screen was designed to identify compounds that disassemble the PNC, and through this study a large array of genome-toxic drugs were found to dissociate PNCs (Norton et al. 2009). These drugs work through three distinct modes of action: pol III transcription inhibition, DNA damage, or both. A pol III promoter assay in cells treated with various compounds that influence pol III transcription shows that PNC prevalence reduction corresponds to the level of transcription inhibition induced by the drugs (Norton et al. 2009), which is consistent with a previous finding that pol III transcription is essential for the PNC integrity (Wang et al. 2003). To determine the mechanism by which DNA damage disassembles PNCs, the role of DNA damage repair on PNC structure was evaluated. Blocking the repair response pathway through either specific inhibitors or siRNA to the DNA damage signaling kinases ATM, ATR, and Chk2 does not block the reduction of the PNC after DNA damage, but does block the recovery of the structure after DNA damage (Norton et al. 2009). This suggests that the loss of PNC stability is a direct result of DNA damage, not an effect of the DNA damage response pathway. Additionally, not all types of DNA damage disrupt the PNC. Although DNA intercalators, topisomerase inhibitors, UV treatment, and cross linkers reduce PNC, double strand or single strand breakers, reactive oxygen species, or alkalators have no impact on PNC structure. The effective PNC disassembly compounds all are able to disrupt the base-pairing capacity of DNA, suggesting that the integrity of DNA is critical to the PNC structure. Furthermore, treatment with a histone deacetylase inhibitor (TSA), which favors an open

C. Pollock and S. Huang

chromatin conformation, also changes the PNC from a dense, round structure into an extended fibril structure. This finding indicates that the PNC-associated locus is responsive to the epigenetic regulations (Norton et al. 2009). Cell Biology Evidence

one or more PNC) is consistently low (,5%) in normal cell lines and immortalized cell lines (0% – 6%) derived from multiple human and mouse tissue types including stromal, endothelial, hematopoietic, or embryonic stem cells. PNC prevalence heterogeneously increases (15% to near 100%) in cancer cell lines. PNCs form in carcinomas, blastomas, and sarcomas, but PNC prevalence is not significantly elevated in malignant cell lines of hematopoietic origin. This suggests that the PNC selectively forms in malignant cells derived from solid tumor tissues (Norton et al. 2008).

Cell biological studies, using several approaches, further support the association of the PNC with a DNA locus. When a conditional mutant of the cell cycle kinase Cdk1 is grown at a nonpermissive condition, DNA undergoes continuous replication without cell division. The number of PNC per cell increases in parallel with the number of replication cycles in these endoreplicating cells, indicating that the number of PNC per cell is directly linked to the copy number of the DNA (Norton et al. 2009). Examination of synchronized HeLa cells shows that during S phase, PNCs generally split into a doublet predominantly at 4 – 6 hours into replication, but become one again during G2 phase. This is consistent with the behavior of a DNA locus during replication and implies that the PNC-associated locus replicates at mid S phase (Norton et al. 2009). Together with the finding that the PNC extends into a fibril structure on treatment with TSA, these studies firmly indicate the association of the PNC with a specific DNA locus. Although recent studies suggest a specific PNC – DNA association, the exact locus with which the PNC interacts has yet to be determined. It is possible that the DNA locus may represent the transcription site for a previously unidentified PNC-associated RNA, or alternatively the PNC could nucleate on a specific locus and regulate expression. Research is ongoing to determine the PNC associating DNA locus.

The strong correlation of PNC prevalence with malignancy in vitro prompted in vivo investigations. PNC prevalence was evaluated in paraffin-embedded human breast cancer tissues in varying clinical stages. Histological samples of normal breast tissue, primary tumors, affected lymph nodes, and distant metastasis were scored for PNC prevalence. The results showed that PNC prevalence is 0% in normal breast tissue, but it increases in the primary tumor (47.9%) and affected lymph nodes (76.3%), and reaches near 100% in distant metastases (Kamath et al. 2005). Additionally, high PNC prevalence in primary tumors of stage I patients positively correlates with disease relapse in a case-matched study, and it is predictive of survival in a retrospective 17-year followup study (Kamath et al. 2005). The correlation between PNC prevalence and metastasis suggests that the PNC-containing cells have a metastatic advantage over non-PNC-containing cells. The formation of the PNC may reflect key changes during transformation that are associated with metastatic capability.

PNC AND TRANSFORMATION

PNC and Metastasis

Initial characterization of the PNC showed that the PNC is predominantly present in cancer cells (Huang et al. 1997). A more complete investigation of over 50 cancerous and normal cell lines confirmed this trend (Norton et al. 2008). PNC prevalence (% cells containing

To address the association of PNC prevalence and metastasis, cell lines of varying metastatic potential have been examined. Studies using a well characterized prostate cancer metastasis model (Pettaway et al. 1996) show that PC-3M cells, which are enriched for metastatic capacity,

In Vivo Studies

The PNC

have a much higher PNC prevalence (85%) than the parental PC-3 heterogenous population (4%). Furthermore, additional rounds of selection for metastatic capacity led to a further increase in PNC prevalence. For example, the PC-3M LN4 cell line, which undergoes an additional four rounds of selection for metastatic capacity, has a PNC prevalence of 98% and abnormally large PNCs. In contrast, PC-3M Pro 4 cells, which are PC-3M cells that have gone through four rounds of selection for a lack of metastatic capacity, show a marked reduction in PNC size and a small reduction in PNC prevalence (71%) compared with PC-3M cells (85%) (Norton et al. 2008). Moreover, when the PNC prevalence is adjusted to the percentage of cells with PNCs greater than 2.2 mm, it correlates very closely with the metastatic behavior of these cells. These observations in cells of the same origin, but of varying metastatic capacities, further confirm that PNC prevalence reflects the metastatic capability of cancer cells. The association of PNC with metastatic capacity is not limited to prostate cancer. To evaluate the association of PNC prevalence with metastatic behavior in another system, two cell lines have been created from a single patient: one from a primary melanoma tumor and one from a distant metastasis. PNC prevalence increases dramatically in cell lines derived from distant metastases over those derived from the primary tumor, and the same results are observed in a similar experiment using colorectal cell lines (Norton et al. 2008). Additionally, PNC prevalence has been examined in cell lines overexpressing the breast cancer related metastatic suppressor protein (BRMS). BRMS is a chromatin remodeling protein that suppresses the ability of breast cancer cells to metastasize (Samant et al. 2000). Stable overexpression of this protein in two breast cancer cell lines significantly reduces metastasis compared with the parental cells when injected into a nude mouse (Samant et al. 2000). The PNC prevalence in the BRMS overexpressing cell lines is significantly lower than in the parental cells, further confirming the association of the PNC with metastatic cells (Norton et al. 2009).

The unique association of the PNC with metastatic cancer cells from solid tumors suggests that it forms in connection with cellular conditions specific to these cells. Multiple traits common to cancer cells, such as increased proliferation and glycolysis, have been examined for an association with PNC prevalence. Results show that PNC prevalence is not affected by cell proliferation. HeLa cells grown in serum-free media show a significantly reduced growth rate, but PNC prevalence does not change. High glucose or low glucose growth conditions do not alter PNC prevalence, which shows the glycolysis state does not influence PNC formation. Furthermore, inducing changes in differentiation status of cancer cells does not affect PNC prevalence, and PNCs are not present in embryonic stem cells (Norton et al. 2008). These results support the idea that PNC formation selectively associates with metastatic behavior. Potential of Clinical Applications for the PNC

Recent studies propose the use of the PNC as a cancer marker for solid tumors. The PNC is an easily scored, multicomponent structure that reflects the complex cellular conditions that develop in the later stages of malignant transformation. The PNC is unique in that it is selectively associated with metastasis without known links to proliferation, glycolysis, or differentiation state. Selective and specific tumor markers could help make appropriate treatment decisions for many cancer patients. Currently, histological PNC prevalence scoring in tissue samples is being refined to improve its reproducibility and reliability as a marker. The PNC has also been presented as an anticancer drug discovery marker. The PNC is associated with the metastatic capacity of cells, and therefore PNC elimination could indicate a shift of cellular behavior from a higher malignant toward a more benign phenotype. In this way, the efficacy of drugs as anticancer agents could be screened for their ability to reduce PNC prevalence. Clinically used cancer drugs and experimental cancer drugs have been screened and a large number of these drugs

C. Pollock and S. Huang

effectively reduce PNC prevalence (Norton et al. 2009). This validates the potential use of PNC prevalence reduction as a phenotypic screening strategy to identify novel compounds that selectively inhibit malignant cells. Compounds that eliminate the PNC not only have potential to be developed into novel drugs, but can also be used as chemical biology tools to help understand the structure and function of the PNC and its functional significance in cancer cells. PNC FUNCTION IN MALIGNANCY

The function of the PNC in malignancy remains to be investigated. However, the close association with cells of metastatic capability suggests that the PNC forms in response to the transformation process, possibly allowing cells to best adapt to a new physiological condition. In other words, the PNC may form as the consequence of transformation to better maintain or further promote the transformed phenotype. Thus, PNCs could play an active role in regulation of gene expression in these cells. As PNCs have been shown to be enriched in newly synthesized pol III RNA, the function of the PNC may be tied to the regulation of pol III RNA expression in malignant cells. The link of Pol III deregulation to cancer was first observed in mouse melanoma cells, where pol III is hyperactive (Schwartz et al. 1974), and more recent investigations support these findings in human cell lines and tissues (Liebhaber et al. 1978; Winter et al. 2000). A recent study showed that the overexpression of the pol III transcripts, tRNA or 5S rRNA alone, is sufficient to cause increased proliferation and oncogenic transformation (Marshall et al. 2008). These findings suggest that increased pol III transcription not only correlates with malignancy, but may play a direct role in the development or progression of cancer. Although PNC-enriched pol III transcripts have not been shown to have a direct role in the development of cancer, the concentration of pol III transcripts in the PNC could be indicative of the deregulation of these RNA in cancer cells. The nucleation of pol III transcripts at the PNC may be involved

in regulating the function of these RNAs in cancer progression or maintenance. Studies are ongoing to investigate the complexes that pol III RNAs form in association with the PNC and their functional relevance to malignancy. There are a number of additional possibilities regarding the potential functions of PNC formation in malignancy. First, yet to be identified factors in the PNC might be altered in solid tumor cells. The increased expression or altered function of these components may lead to nucleation of the RNAs and RNA-binding proteins in the PNC, which could directly influence the availability of PNC-associated RNA or proteins in cancer cells. Second, the PNC may play a role in nucleolar function, which is known to be deregulated in malignant cells (reviewed in Ruggero and Pandolfi 2003). The PNC is in close spatial proximity to the nucleolus, and the PNC is enriched in components (i.e., Nucleolin, PTB, RNase MRP RNA, and RNase P RNA) that are involved in rDNA transcription and ribosome synthesis. Studies are underway to determine the role of the PNC in malignancy. SUMMARY

Although the function of the PNC is not yet fully understood, significant progress has been made in the characterization of the PNC in recent years. The PNC is a unique and dynamic nuclear body that selectively associates with metastatic capacity in a broad array of solid tumor tissues. The PNC is enriched in both RNA-binding proteins and a subset of newly synthesized pol III transcripts. Furthermore, the PNC is nucleated on a DNA locus or loci, and the integrity of DNA is critical for the PNC structure. The current working model is that the PNC is associated with and may represent changes in novel molecular complexes that are involved in the metabolism of newly synthesized pol III RNA during malignancy. Nucleation of the PNC on a DNA locus may be directly involved in the regulation of gene expression at that locus. Studies are underway to identify and characterize novel pol III-RNA complexes and the DNA locus they nucleate

The PNC

on to form the PNC. These investigations will help elucidate the role of the PNC and increase our knowledge of basic cancer biology. ACKNOWLEDGMENTS

S.H. is funded by R01 GM078555-01A1 and C.P. is funded by the T32 CA080621-06A2 training grant. REFERENCES Bernardi R, Pandolfi PP. 2007. Structure, dynamics and functions of promyelocytic leukaemia nuclear bodies. Nat Rev Mol Cell Biol 8: 1006–1016. Christov CP, Gardiner TJ, Szuts D, Krude T. 2006. Functional requirement of noncoding Y RNAs for human chromosomal DNA replication. Mol Cell Biol 26: 6993– 7004. Fabini G, Raijmakers R, Hayer S, Fouraux MA, Pruijn GJ, Steiner G. 2001. The heterogeneous nuclear ribonucleoproteins I and K interact with a subset of the ro ribonucleoprotein-associated Y RNAs in vitro and in vivo. J Biol Chem 276: 20711–20718. Ghetti A, Pinol-Roma S, Michael WM, Morandi C, Dreyfuss G. 1992. hnRNP I, the polypyrimidine tract-binding protein: Distinct nuclear localization and association with hnRNAs. Nucleic Acids Res 20: 3671– 3678. Grudnik P, Bange G, Sinning I. 2009. Protein targeting by the signal recognition particle. Biol Chem 390: 775 –782. Hall MP, Huang S, Black DL. 2004. Differentiation-induced colocalization of the KH-type splicing regulatory protein with polypyrimidine tract binding protein and the c-src pre-mRNA. Mol Biol Cell 15: 774 –786. Hasler J, Strub K. 2006a. Alu elements as regulators of gene expression. Nucleic Acids Res 34: 5491–5497. Hasler J, Strub K. 2006b. Alu RNP and Alu RNA regulate translation initiation in vitro. Nucleic Acids Res 34: 2374– 2385. Higashi K, Matsuhisa T, Kitao A, Sakamoto Y. 1968. Selective suppression of nucleolar RNA metabolism in the absence of protein synthesis. Biochim Biophys Acta 166: 388 –393. Ho TH, Bundman D, Armstrong DL, Cooper TA. 2005. Transgenic mice expressing CUG-BP1 reproduce splicing mis-regulation observed in myotonic dystrophy. Hum Mol Genet 14: 1539–1547. Huang S, Deerinck TJ, Ellisman MH, Spector DL. 1997. The dynamic organization of the perinucleolar compartment in the cell nucleus. J Cell Biol 137: 965 –974. Huang S, Deerinck TJ, Ellisman MH, Spector DL. 1998. The perinucleolar compartment and transcription. J Cell Biol 143: 35–47. Huttelmaier S, Illenberger S, Grosheva I, Rudiger M, Singer RH, Jockusch BM. 2001. Raver1, a dual compartment protein, is a ligand for PTB/hnRNPI and microfilament attachment proteins. J Cell Biol 155: 775– 786. Kamath RV, Thor AD, Wang C, Edgerton SM, Slusarczyk A, Leary DJ, Wang J, Wiley EL, Jovanovic B, Wu Q, et al. 2005. Perinucleolar compartment prevalence has an

independent prognostic value for breast cancer. Cancer Res 65: 246–253. Kleinhenz B, Fabienke M, Swiniarski S, Wittenmayer N, Kirsch J, Jockusch BM, Arnold HH, Illenberger S. 2005. Raver2, a new member of the hnRNP family. FEBS Lett 579: 4254– 4258. Kopp K, Huang S. 2005. Perinucleolar compartment and transformation. J Cell Biochem 95: 217 –225. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605– 612. Liebhaber SA, Wolf S, Schlessinger D. 1978. Differences in rRNA metabolism of primary and SV40-transformed human fibroblasts. Cell 13: 121– 127. Lou H, Gagel RF, Berget SM. 1996. An intron enhancer recognized by splicing factors activates polyadenylation. Genes Dev 10: 208– 219. Marshall L, Kenneth NS, White RJ. 2008. Elevated tRNA(iMet) synthesis can drive cell proliferation and oncogenic transformation. Cell 133: 78–89. Matera AG, Frey MR, Margelot K, Wolin SL. 1995. A perinucleolar compartment contains several RNA polymerase III transcripts as well as the polypyrimidine tract-binding protein, hnRNP I. J Cell Biol 129: 1181–1193. Min H, Turck CW, Nikolic JM, Black DL. 1997. A new regulatory protein, KSRP, mediates exon inclusion through an intronic splicing enhancer. Genes Dev 11: 1023–1036. Morris GE. 2008. The Cajal body. Biochim Biophys Acta 1783: 2108– 2115. Nechama M, Ben-Dov IZ, Briata P, Gherzi R, Naveh-Many T. 2008. The mRNA decay promoting factor K-homology splicing regulator protein post-transcriptionally determines parathyroid hormone mRNA levels. FASEB J 22: 3458–3468. Norton JT, Pollock CB, Wang C, Schink JC, Kim JJ, Huang S. 2008. Perinucleolar compartment prevalence is a phenotypic pancancer marker of malignancy. 113: 861 –869. Norton JT, Wang C, Gjidoda A, Henry RW, Huang S. 2009. The perinucleolar compartment is directly associated with DNA. J Biol Chem 284: 4090– 4101. Pettaway CA, Pathak S, Greene G, Ramirez E, Wilson MR, Killion JJ, Fidler IJ. 1996. Selection of highly metastatic variants of different human prostatic carcinomas using orthotopic implantation in nude mice. Clin Cancer Res 2: 1627–1636. Pollock C, Huang S. 2009. The perinucleolar compartment. J Cell Biochem 107: 189–193. Ruggero D, Pandolfi PP. 2003. Does the ribosome translate cancer? Nat Rev Cancer 3: 179– 192. Samant RS, Seraj MJ, Saunders MM, Sakamaki TS, Shevde LA, Harms JF, Leonard TO, Goldberg SF, Budgeon L, Meehan WJ, et al. 2000. Analysis of mechanisms underlying BRMS1 suppression of metastasis. Clin Exp Metastasis 18: 683–693. Savkur RS, Philips AV, Cooper TA. 2001. Aberrant regulation of insulin receptor alternative splicing is associated with insulin resistance in myotonic dystrophy. Nat Genet 29: 40– 47. Schwartz LB, Sklar VE, Jaehning JA, Weinmann R, Roeder RG. 1974. Isolation and partial characterization of

C. Pollock and S. Huang the multiple forms of deoxyribonucleic acid-dependent ribonucleic acid polymerase in the mouse myeloma, MOPC 315. J Biol Chem 249: 5889–5897. Singh K, Laughlin J, Kosinski PA, Covey LR. 2004. Nucleolin is a second component of the CD154 mRNA stability complex that regulates mRNA turnover in activated T cells. J Immunol 173: 976– 985. Stoneley M, Willis AE. 2004. Cellular internal ribosome entry segments: Structures, trans-acting factors and regulation of gene expression. Oncogene 23: 3200– 3207. Timchenko LT, Miller JW, Timchenko NA, DeVore DR, Datar KV, Lin L, Roberts R, Caskey CT, Swanson MS. 1996. Identification of a (CUG)n triplet repeat RNA-binding protein and its expression in myotonic dystrophy. Nucleic Acids Res 24: 4407–4414. van Eenennaam H, Jarrous N, van Venrooij WJ, Pruijn GJ. 2000. Architecture and function of the human endonucleases RNase P and RNase MRP. IUBMB Life 49: 265–272. Wagner EJ, Garcia-Blanco MA. 2001. Polypyrimidine tract binding protein antagonizes exon definition. Mol Cell Biol 21: 3281– 3288.

Wang C, Politz JC, Pederson T, Huang S. 2003. RNA polymerase III transcripts and the PTB protein are essential for the integrity of the perinucleolar compartment. Mol Biol Cell 14: 2425–2435. Winter AG, Sourvinos G, Allison SJ, Tosh K, Scott PH, Spandidos DA, White RJ. 2000. RNA polymerase III transcription factor TFIIIC2 is overexpressed in ovarian tumors. Proc Natl Acad Sci 97: 12619–12624. Wolin SL, Steitz JA. 1984. The Ro small cytoplasmic ribonucleoproteins: Identification of the antigenic protein and its binding site on the Ro RNAs. Proc Natl Acad Sci 81: 1996–2000. Wolin SL, Walter P. 1989. Signal recognition particle mediates a transient elongation arrest of preprolactin in reticulocyte lysate. J Cell Biol 109: 2617– 2622. Yamamoto H, Tsukahara K, Kanaoka Y, Jinno S, Okayama H. 1999. Isolation of a mammalian homologue of a fission yeast differentiation regulator. Mol Cell Biol 19: 3829–3841. Zimber A, Nguyen QD, Gespach C. 2004. Nuclear bodies and compartments: Functional roles and cellular signalling in health and disease. Cell Signal 16: 1085–1104.

Paraspeckles Archa H. Fox1 and Angus I. Lamond2 1

Western Australian Institute for Medical Research and Centre For Medical Research, University of Western Australia, Crawley 6009 Western Australia, Australia

2

Wellcome Trust Centre for Gene Regulation & Expression, College of Life Sciences, University of Dundee DUNDEE DD1 5EH UK

Correspondence: [email protected]

Paraspeckles are a relatively new class of subnuclear bodies found in the interchromatin space of mammalian cells. They are RNA-protein structures formed by the interaction between a long nonprotein-coding RNA species, NEAT1/Men 1/b, and members of the DBHS (Drosophila Behavior Human Splicing) family of proteins: P54NRB/NONO, PSPC1, and PSF/SFPQ. Paraspeckles are critical to the control of gene expression through the nuclear retention of RNA containing double-stranded RNA regions that have been subject to adenosine-to-inosine editing. Through this mechanism paraspeckles and their components may ultimately have a role in controlling gene expression during many cellular processes including differentiation, viral infection, and stress responses.

DISCOVERY OF PARASPECKLES

he cell nucleus is a large and complex cellular organelle with an intricate internal organization that is still not fully characterized. One feature of nuclear organization is the presence of distinct subnuclear bodies, each of which contain specific nuclear proteins and nucleic acids (Platani and Lamond 2004). Most subnuclear bodies reside in the interchromatin space, including Cajal bodies, PML bodies, and nuclear speckles, enriched in splicing factors (Lamond and Spector 2003). Paraspeckles are one of the most recent subnuclear bodies identified, discovered in 2002 as part of a study to better understand the full biological role of the nucleolus. In a mass spectrometry based proteomic analysis of purified

T

human nucleoli, 271 proteins were identified, 30% of which were novel (Andersen et al. 2002). A follow up analysis on one of these newly identified novel proteins, showed that it was not enriched in nucleoli, but instead was found diffusely distributed within the nucleoplasm as well as concentrated in 5 – 20 subnuclear foci (Fox et al. 2002). Colocalization studies showed that these foci neither coincided, nor directly overlapped, with markers for any previously known subnuclear structure. The foci were thus named “paraspeckles” because they were observed in the interchromatin space near to, yet distinct from, nuclear speckles (Fig. 1). The novel protein that localized to these structures was subsequently named “Paraspeckle Protein 1” (PSPC1).

Editors: Tom Misteli and David Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000687 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000687

A.H. Fox and A. Lamond

B

A

D

C

DBHS sequence conservation Protein–protein interaction

P54NRB PSF PSPC1

RRM1

RRM2

NOPS Coiled coil

RRM1

RRM2

NOPS Coiled coil

RRM1

RRM2

NOPS Coiled coil

Minimal region of PSPC1 needed for paraspeckle targeting

Figure 1. Paraspeckles seen with fluorescent and electron microscopy. (A) Combined DIC and fluorescence

micrograph of a HeLa cell stained with anti-PSPC1 to show paraspeckles (green) and B23 nucleolar marker (red). (B) Fluorescence micrograph of a section through a HeLa cell stained with anti-PSPC1 (green), anti-SC35 (red), and DAPI (blue) to show the relationship between paraspeckles and nuclear speckles. (C) Transmission electron micrograph of sections of HeLa cells immuno-gold labelled with anti-PSPC1: image kindly provided by Sylvie Souquere and Gerard Pierron (Villejuif, France). Scale bars in A–B, 10 mm, scale bar in C, 0.5 mm. (D) The DBHS protein family showing domain structure and indicating regions involved in paraspeckle biology.

Considering that PSPC1 was first identified in a proteomic screen for nucleolar factors, it was initially surprising that localization studies did not detect it accumulated in nucleoli. However, exploring the dynamic nature of PSPC1 revealed its nucleolar relationship. When cells were treated with drugs that inhibit RNA Polymerase II (Pol II) transcription, PSPC1 relocalized to perinucleolar cap structures (Fox et al. 2002 and Fig. 2). Perinucleolar enrichment of PSPC1 was also observed in newly divided cells that had not recommenced transcription following cell division (Fox et al. 2005). Thus PSPC1 is found in paraspeckles in transcriptionally active cells, and perinucleolar caps in cells that are not actively transcribing Pol II genes. Further, photodynamic studies showed that under normal conditions PSPC1 molecules continually traffic through nucleoli, despite

their steady-state enrichment within paraspeckles, explaining their presence in the nucleolar proteome (Fox et al. 2002). PARASPECKLE CHARACTERIZATION

Paraspeckles are small, irregularly sized, and unevenly distributed subnuclear bodies. Depending on the cell type, paraspeckles number between 5 and 20 foci per nucleus (for example, HeLa contain 13– 17 foci/nucleus and NIH3T3 5 – 10 foci/nucleus, Clemson et al. 2009). EM studies and fluorescent images show a paraspeckle size range of 0.5 – 1 mm in diameter, and they have an irregular, sausagelike shape (Cardinale et al. 2007 and Fig. 1C). Transmission EM of cells labeled with paraspeckle markers show labeling of distinct nuclear structures, rich in RNA (Prasanth et al. 2005; Cardinale et al.

Paraspeckles

A Nucleolus

Peri-nucleolar caps

B Contain fibrillarin, p80coilin, Pol I factors

Contain paraspeckle proteins and other Pol II factors

Nucleolar body

Figure 2. Perinucleolar caps observed with RNA Pol

II transcription inhibition. (A) Combined DIC and fluorescence micrograph of HeLa cells following 4 h treatment with Actinomycin D to inhibit RNA Pol II transcription. Nucleolar morphology changes under these conditions, to create a nucleolar body and a number of perinucleolar caps (arrow). Cells were transfected with a plasmid expressing YFP-PSPC1 (green), that localizes to perinucleolar caps under these conditions (large arrow); scale bar, 5 mm. (B) Perinucleolar caps form upon inhibition of RNA Pol II transcription, see text and (Shav-tal et al. 2005).

2007, Fig. 1C). These EM structures labeled with paraspeckle markers correspond, at least partly, to the Interchromatin Granule Associated Zones (IGAZ – Visa et al. 1993). IGAZ are electron dense fibrillar regions closely aligned to interchromatin granules (nuclear speckles) with unknown function. IGAZ are reported to contain both U1 RNA and coilin, although there is no evidence of colocalization between paraspeckles and either of these molecules at the level of fluorescence (Fox et al. 2002).

Thus far, paraspeckles are only evident in mammalian nuclei. Within mammalian tissues and cells, paraspeckles are wide-spread: The majority of mouse and human cell lines and tissues examined contain paraspeckles, including transformed and primary cell lines, embryonic fibroblasts, and tumorigenic biopsies (Fox et al. 2002; Prasanth et al. 2005; Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009 and unpublished data). Interestingly, human embryonic stem cells (hESC) are so far the only mammalian cell type that are reported not to contain paraspeckles (Chen and Carmichael, 2009). Orthologs of the core paraspeckle protein components are found in other vertebrate and invertebrate species, however, a noncoding RNA (ncRNA) NEAT1, which is essential for paraspeckle formation (see below) is specific to mammals, likely explaining the restriction of paraspeckles to this class. Paraspeckles are observed within the interchromatin space, sandwiched between larger nuclear speckles and chromatin. Current evidence suggests paraspeckles do not directly overlap with sites of active transcription, as they do not contain newly made pulse-labeled Br-UTP containing transcripts, however paraspeckles may still form in association with some active genes (see below) (Fox et al. 2002; Xie et al. 2006). Quantitation of staining on ultrathin sections of labeled cells shows that paraspeckles contain inactive RNA Polymerase II (Pol II), whereas the newly made RNA and active RNA Pol II reside on the edge of paraspeckles (Xie et al. 2006). The functional relationship between paraspeckles and the nucleolus is yet to be fully elucidated, but the observed cycling of PSPC1 between paraspeckles and nucleoli, and the localization of paraspeckle proteins to perinucleolar caps when RNA Pol II transcription is inhibited, suggests that it may mediate some form of regulatory cross talk. Many other proteins involved in Pol II transcription are also observed within perinucleolar caps in cells where Pol II transcription is inactive (Shav-Tal et al. 2005). These proteins appear to segregate into distinct cap structures: large “dark” perinucleolar caps containing the paraspeckle proteins, as well as

A.H. Fox and A. Lamond

several other RNA-binding proteins; and a number of smaller “light” caps containing fibrillarin, coilin, and RNA Pol I components (Fig. 2) (Shav-Tal et al. 2005).

nuclear ncRNA, NEAT1, is an architectural paraspeckle component essential for their formation and maintenance. DBHS Protein Family

PARASPECKLE COMPONENTS

A small number of protein and RNA components are known to be enriched in paraspeckles (Table 1). Most of the paraspeckle proteins have roles in Pol II transcription and/or RNA processing; including the members of the DBHS protein family, several transcription factors, a cotranscriptional splicing factor and a 30 RNA cleavage factor. There are two specific paraspeckle RNA components, each belong to different classes: Ctn is regulated within paraspeckles, and is implicated in the control of gene expression by RNA nuclear retention, whereas the abundant

The core protein components of paraspeckles are the three mammalian members of the DBHS protein family: PSF/SFPQ, NONO/P54NRB, and PSPC1. Both endogenous and tagged forms of these proteins are found localized within the nucleoplasm as well as paraspeckles in mammalian cells. Of the three DBHS proteins, PSPC1 has most often been used as the marker for paraspeckles, as it has a lower background nucleoplasmic signal in many cell types than P54NRB and PSF. The three proteins have 50% sequence identity within two amino-terminal RNA binding motifs and a carboxy-terminal coiled coil domain (Fig. 1D). The DBHS proteins appear to play a key role in the structural integrity of

Table 1. Paraspeckle components Name

Synonyms

Proteins P54NRB

NONO, NMT55, NRB54

PSF

SFPQ

PSPC1 CoAAa CFIM68

PSP1 PSP2, RBM14, SIP, SYTIP1 CPSF6, HPBRII-4

SOX9b WTXa WT1(þKTS)a

SRA1

BCL11Aa, b RNA Pol II

CTIP1, ZNF856

RNA NEAT1

Ctn a

WAGR

Men 1/b, VINC-1

Comments

DBHS; required for paraspeckle integrity in HeLa DBHS; required for paraspeckle integrity in HeLa DBHS Transcriptional/splicing coregulator Cleavage factor. Also found in nuclear speckles Developmental transcription factor Wilms tumor, tumor suppressor Wilms tumor, partial colocalization with paraspeckles Zinc finger transcription factor Also found associated with chromatin and nuclear speckles Long noncoding RNA found in mammals; required for paraspeckle integrity in HeLa Mouse-specific, alternative transcript produced from mCAT2 locus

Paraspeckle localization reference

Fox et al. 2002 Prasanth et al. 2005 Fox et al. 2002 Fox et al. 2002 Dettwiler et al. 2004 Hata et al. 2008 Rivera et al. 2009 Dutton et al. 2006 Liu et al. 2006 Xie et al. 2006

Clemson et al. 2009; Sunwoo et al. 2009; Sasaki et al. 2009 Prasanth et al. 2005

Only overexpressed proteins have been assessed for localization in paraspeckles. Further studies need to address if these proteins are genuine paraspeckle components, or are retargeting DBHS proteins into different subnuclear locations. b

Paraspeckles

paraspeckles, because knockdown of the abundant proteins P54NRB or PSF in HeLa cells results in a loss of paraspeckles (Sasaki et al. 2009). Within HeLa cells, PSPC1 is much less abundant than PSF and P54NRB (Fox et al. 2005), and knockdown of PSPC1 does not affect paraspeckles in these cells (Sasaki et al. 2009). Reciprocal protein–protein interactions have been reported between members of this family, and it is likely in vivo they are found as either homo or heterodimers (Myojin et al. 2004; Fox et al. 2005). PSPC1 and P54NRB interact via their coiled-coil domains, and, given the sequence similarity, it is likely that PSF also interacts through this domain. Similarly, Hrp65, the Chironomus tentans DBHS homolog homodimerizes through its coiled-coil domain (Kiesler et al. 2003). RNA-binding domains, as well as protein – protein interaction domains are required for members of the DBHS family to be targeted to paraspeckles. The minimal fragment of PSPC1 that is targeted to paraspeckles contains at least one RRM motif, in addition to its coiledcoil domain (Fox et al. 2005 and Fig. 1D). In contrast, neither RNA recognition motif (RRM) is required for PSPC1 to be targeted to perinucleolar caps on transcription inhibition. Within the DBHS family, the specificity of RRMs for paraspeckle targeting varies, as, unlike PSPC1, which can use either one of its two RRMs to be targeted to paraspeckles, only the second RRM of PSF is sufficient to target PSF to subnuclear foci (Dye and Patton 2001). These findings raise the possibility that subtle variations in affinities for different RNA targets may allow each DBHS protein to target different RNA species to paraspeckles. DBHS Family Member Functions

Members of the DBHS family of proteins have been shown to bind both double- and singlestranded DNA and RNA and have been copurified with numerous different complexes, leading to the label of “multifunctional nuclear proteins” for this protein family (reviewed in Shav-Tal and Zipori 2002). The paraspeckle connection to the known DBHS functions has not been addressed in most cases.

The DBHS family proteins are involved in many aspects of RNA production and processing including transcription initiation, transcriptional termination, and splicing (Patton et al. 1993; Yang et al. 1993; Peng et al. 2002; Kameoka et al. 2004; Kaneko et al. 2007; Ito et al. 2008). Beyond RNA production, PSF and P54NRB also play a role in RNA surveillance: Binding and retaining hyper-adenosine-toinosine (A-to-I) edited RNA within the nucleus (Zhang and Carmichael 2001). Nuclear A-to-I edited RNA is generated when double-stranded RNA-dependent adenosine deaminases target long (optimally 100bp) double-stranded RNA and randomly convert up to 50% of the adenosines to inosines. In what is thought to have originated as an antiviral mechanism, the resulting inosine-containing-RNA can be retained in the nucleus, rather than being exported to the cytoplasm (Zhang and Carmichael 2001). DBHS proteins may also have roles in the cytoplasm. PSF and P54NRB were isolated as part of an RNA-transport granule in dendrites (Kanai et al. 2004) and both proteins have CRM1-independent nuclear export signals, in addition to nuclear retention motifs, however only P54NRB actively shuttles into the cytoplasm (Zolotukhin et al. 2003). With roles in constitutive processes such as splicing and transcription, the functional implications for the DBHS proteins are wideranging. One interesting example implicates P54NRB in the control of mammalian circadian rhythms. P54NRB is required for circadian rhythm maintenance via association with the PERIOD-1 protein, an essential component of negative feedback in mammalian circadian rhythm (Brown et al. 2005). Similarly in Drosophila melanogaster, mutants of the DBHS protein, NonA, are nearly arrhythmic, indicating a conserved role in circadian rhythm regulation for this protein family (Brown et al. 2005). Other Paraspeckle Proteins

The current definition of a paraspeckle protein is one that colocalizes in subnuclear foci with DBHS proteins, however, the localization pattern is not always identical. For example, the

A.H. Fox and A. Lamond

cleavage factor CFIm is observed in paraspeckles, but is in addition found within nuclear speckles (Dettwiler et al. 2004), and RNA Pol II is partially localized to paraspeckles and also to chromatin and nuclear speckles (Xie et al. 2006). It should be noted that several of the proteins reported to localize in paraspeckles have only been examined following overexpression, namely BCL11A, WTX, WT1(þKTS), and COAA (Fox et al. 2002; Dutton et al. 2006; Liu et al. 2006; Rivera et al. 2009), therefore these proteins cannot be considered bona fide paraspeckle components until the localization of the endogenous factors has been examined. Additional paraspeckle proteins identified to date are either transcription factors or transcriptional coregulators. CoAA (PSP2/RBM14) is a transcriptional coactivator that regulates steroid receptor-mediated transcription and alternative RNA splicing (Iwasaki et al. 2001; Auboeuf et al. 2004). An alternatively spliced isoform of CoAA, termed CoAM, acts as a dominant negative antagonistic corepressor of CoAA. Interestingly, there is evidence that PSF and P54NRB act to influence which splice variant is generated from the CoAA gene (Yang et al. 2007). RBM4, the mammalian homolog of the Drosophila melanogaster Lark protein (an essential component of circadian rhythm regulation and early development in Drosophila), forms a fusion with CoAA, via putative trans-splicing events (Brooks et al. 2009). This finding suggests another possible link between paraspeckles and proteins involved in the control of circadian rhythms. Another transcriptional coregulator and tumor suppressor, WTX, colocalizes with P54NRB in paraspeckles and coactivates WT1 transcription (Rivera et al. 2009). Interestingly, an isoform of WT1 also colocalizes with some, but not all, paraspeckles (Dutton et al. 2006). Sox9 is a transcription factor that plays an essential role in bone formation and interacts with P54NRB (Hata et al. 2008). Whilst SOX9 is reported to be a paraspeckle component, its overexpression results in altered subnuclear localization of P54NRB and PSPC1, suggesting that it may instead be re-targeting these proteins out of paraspeckles to different subnuclear sites (Hata et al. 2008). BCL11A, a transcription

factor involved in B cell lymphoma/leukemia, colocalizes with PSPC1 and P54NRB in subnuclear foci (Liu et al. 2006). Again, these foci do not resemble paraspeckles—suggesting the proteins have been directed out of paraspeckles to new complexes. Supporting this argument, in contrast to paraspeckles observed with PSPC1 and P54NRB, RNAase treatment of BCL11A foci does not lead to their dissolution, instead, DNAase treatment results in a diffuse BCL11A signal (Liu et al. 2006). Mammalian cleavage factor I (CF Im) is an essential factor that is required for the first step in pre-mRNA 30 end processing, and localizes to both paraspeckles and nuclear speckles (Dettwiler et al. 2004; Cardinale et al. 2007). The largest subunit of the CF Im heterodimer contains an amino-terminal RRM and a carboxy-terminal charged arginine- and serinerich “RS” domain, common to splicing factors. Interestingly, the RS domain is sufficient for nuclear speckle localization, whereas both the RS and RRM domains are required for paraspeckle targeting (Cardinale et al. 2007). Because the RRM domain of CF Im has been reported to mediate protein – protein interactions (Dettwiler et al. 2004), it is likely CF Im is targeted to these two compartments via interaction with different subsets of proteins. PARASPECKLE RNAs

When we first identified paraspeckles, although only protein components were known, we proposed that their function would most likely be linked to some aspect of RNA, such as processing or transport, because several lines of evidence indicated that paraspeckles were RNAprotein bodies. First, paraspeckles were disrupted when cells were treated with RNAse (Fox et al. 2005; Prasanth et al. 2005); second, all of the major paraspeckle proteins contained RNA binding motifs and had previously described functions in RNA processing; thirdly, PSPC1 required its RNA binding domain for paraspeckle targeting (Fox et al. 2005). Finally, it was shown that paraspeckles disassemble without active RNA Pol II transcription and subsequently reassemble when transcription is restored.

Paraspeckles

Two paraspeckle-associated RNA species have since been identified, each providing clues to paraspeckle formation and function. Ctn RNA

The first RNA found that specifically localized to paraspeckles was termed Ctn, described by Spector and colleagues (Prasanth et al. 2005). Ctn is a mouse-specific, nuclear-enriched, spliced poly(Aþ) transcript that is generated from the mCAT2 gene locus. Ctn contains all of the coding exons of the CAT2 (cationic amino acid transporter 2) protein. However, compared with the canonical mCAT2, it is generated from a different promoter, and has a distal poly(Aþ) site resulting in a much longer 30 untranslated region (UTR). RNA-FISH with Ctn-specific probes showed that it was nucleoplasmic as well as localizing to paraspeckles in several cell types. Why then is Ctn not subject to the usual nuclear export fate of most mRNAs? The answer lies in the long 30 UTR of Ctn, which contains A-to-I edited stretches of RNA. The editing takes place on inverted repeat elements that form long dsRNA regions. Given that P54NRB was known to preferentially bind inosineRNA and retain it in the nucleus (Zhang and Carmichael 2001), the hypothesis was that DBHS proteins would mediate the paraspeckleand nuclear retention of Ctn. Consistent with this, both PSPC1 and P54NRB were shown to associate with Ctn in vivo (Prasanth et al. 2005). The nuclear retention of Ctn is linked to the control of gene expression of mCAT2: a protein in the nitric oxide response pathway, needed for wound healing and defense against infection. A variety of stress signals were shown to trigger the cleavage of the long 30 UTR from Ctn, resulting in lower nuclear levels of Ctn, with a concomitant rise in mCAT2 levels in the cytoplasm, and increased mCAT2 protein production. This gene expression control system results in a pulse of protein expression, and effectively provides the cell with a rapid increase in the production of protein upon the signal being received (Fig. 3). Although Ctn is mouse-specific, analysis of the human transcriptome has indicated that the

control of gene expression by nuclear retention likely also occurs in humans: as many as 50% of all genes may produce transcripts with extended 30 UTRs that use a distal poly(Aþ) site (Iseli et al. 2002). Moreover, the repeat elements that are A-to-I edited in Ctn are amongst the most abundant repetitive elements in mammalian genomes, and a large proportion of these elements are A-to-I edited in humans (Levanon et al. 2005; Chen and Carmichael 2008). A recent study found 333 genes in the human genome with inverted Alu repeat elements in their 30 UTR (Alu being the most common family of human repeat elements) (Chen et al. 2008). Amplified inverted Alus from two of these genes were both capable of mediating RNA nuclear retention of a reporter gene (Chen et al. 2008). Moreover, as with Ctn, DBHS protein interaction plays a role in the retention of the reporter RNA, within subnuclear foci that partly overlap paraspeckles, as defined by exogenously expressed P54NRB (Chen et al. 2008). Additionally, because there is at least one other example of a nuclear-retained RNA that does not have inverted repeats (Kay et al. 2005), it is also likely that other RNA elements may mediate nuclear retention. The exploration of the RNA nuclear retention mechanism for controlling gene expression has only just begun. One issue is the need for a large scale study of nuclear enriched coding and ncRNA transcripts to determine how prevalent nuclear retention is, and if all nuclear retained RNAs may be associated with paraspeckles or analogous structures. Another area that will be important to examine is the release of RNA from nuclear retention in paraspeckles. A recent bioinformatic analysis has begun to address this by determining that sequence databases contain hundreds of examples of mRNA transcripts in which the genome-encoded inverted repeats have been excised, an event presumably linked with release of these mRNAs from nuclear retention (Osenberg et al. 2009). NEAT1/Men 1/b RNA

We have previously speculated that a paraspeckle-specific RNA species was required for

A.H. Fox and A. Lamond

Cytoplasm

Nucleus Gene locus or plasmid

ORF

ORF AAAA

AAAA

RNA + inverted repeat elements

Export

‘Normal’ mRNA

A-to-I Editing Embryonic stem cells: No paraspeckles

Nuclear retention

Translation

Paraspeckle

Export

P54NRB AAAA

For Ctn: ?

Cleavage A???

Export Translation

Signal

Response

Stress

Figure 3. Gene expression by nuclear retention. RNA transcripts containing double-stranded RNA regions

(formed by inverted repeat elements) are subject to A-to-I editing and retained in the nucleus and within paraspeckles. This mechanism has been shown for several endogenous genes, as well as exogenously expressed reporter genes (Prasanth et al. 2005; Chen et al. 2008; Chen and Carmichael, 2009). Nuclear retention of hyper-edited RNA is linked to the formation of paraspeckles, and does not occur efficiently in human embryonic stem cells that lack paraspeckles. In the case of Ctn, stress signals mediate cleavage of the 30 UTR and release of the RNA from the nucleus (Prasanth et al. 2005).

paraspeckle formation, however Ctn could not fulfill this role: first, because it is mouse-specific and second, because knockdown of Ctn does not disrupt paraspeckles (Prasanth et al. 2005). In 2009, three different groups reported the discovery that NEAT1, a long nuclear ncRNA, is essential for paraspeckle structural integrity and formation (Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009).

It is now known, for mammals in particular, that the majority of the genome is transcribed, to generate both protein-coding and nonprotein coding RNA (Carninci et al. 2005). NcRNAs may arise from transcribed introns of protein-coding genes, or may be antisense to them, or may be transcribed from their own bona fide gene loci between protein-coding genes (Mercer et al. 2009). In recent years the

Paraspeckles

explosion of information related to the identity and function of small ncRNAs has been evident, however, we are only beginning to understand the varying roles that long ncRNAs (.200nt) may be playing in the cell (Prasanth and Spector 2007). In 2007 a study aimed at identifying nuclear ncRNA species described two abundant, ubiquitously expressed Nuclear Enriched Autosomal noncoding Transcripts termed NEAT1 (also known as Men1/b or VINC-1 ) and NEAT2 (also known as MALAT-1) (Hutchinson et al. 2007). RNA-FISH showed that MALAT-1 localized to nuclear speckles, whereas NEAT1 was observed within subnuclear foci found near nuclear speckles, shown to be paraspeckles. Both NEAT1 and MALAT-1 ncRNAs are produced by RNA Pol II, independently of protein-coding genes, and their genes are conserved syntenically a short distance apart in mammalian genomes (Fig. 4A). Two variant transcripts of NEAT1 are transcribed, NEAT1_v1 and NEAT1_v2 ( previously referred to as Men1 and Menb), these share approx 3– 4 kb of sequence at the 50 end that precisely delineates NEAT1_v1, with the longer isoform, NEAT1_v2, containing an additional approximately 20kb of RNA (Fig. 4A). Both NEAT1_v2 RNA, as well as MALAT-1 RNA, are cleaved very close to their 30 ends to produce an unusual small tRNA-like molecule (Wilusz et al. 2008; Sunwoo et al. 2009). Interestingly, bioinformatic analysis, although not exhaustive, has not found any other ncRNAs with this unusual 30 end, suggesting it could be specific to this duo of long nuclear ncRNAs (Wilusz et al. 2008). NEAT1 is found within paraspeckles in a variety of different cell lines in both mouse and human cells (Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009). An essential role for NEAT1 in paraspeckle integrity was shown by knockdown of NEAT1 in cultured As of 27/7/09 the official HUGO gene nomenclature symbol for this ncRNA is NEAT1 and the HUGO gene name is “Nuclear ParaspEckle Assembly Transcript 1 (nonprotein coding).” HUGO does not have jurisdiction on nomenclature of transcript variants, however, in line with the HUGO recommendation, we propose the use of NEAT1_v1 for the 3.7kb variant and NEAT1_v2 for the 23kb variant.

cells, resulting in the loss of paraspeckles, as judged by loss of DBHS protein localization to subnuclear foci (Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009). Moreover, paraspeckles do not re-form following reversible transcription inhibition in the presence of the NEAT1 knockdown, suggesting NEAT1 is also required for paraspeckle formation (Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009). Further evidence suggests that NEAT1 is the nucleating factor for paraspeckles, at least in NIH3T3 cells, as stable overexpression of NEAT1_v1 results in an increase in paraspeckle number (Clemson et al. 2009). NEAT1 ncRNA is not only needed for paraspeckle integrity, but there is evidence that paraspeckles form near to the NEAT1 gene itself. Combined DNA and RNA FISH showed that cells in early G1 have the first paraspeckles forming close to the NEAT1 gene, and in interphase, clusters of paraspeckles are observed close to the NEAT1 gene locus (Clemson et al. 2009). As with Ctn, it appears that interaction with DBHS proteins plays a role in NEAT1 paraspeckle localization, as coimmunoprecipitation experiments with each of the DBHS proteins pulled out NEAT1 to varying levels of enrichment (Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009). The precise NEAT1 RNA binding site for the DBHS proteins has not been established, although recombinant DBHS proteins can bind the NEAT1_v1 in vitro (Clemson et al. 2009), and paraspeckles can persist solely in the presence of NEAT1_v1 (Sunwoo et al. 2009), other evidence suggests that the final 10kb of human NEAT1_v2 may contain the necessary sequence for paraspeckle formation in vivo (Sasaki et al. 2009). Interestingly, in contrast to Ctn, NEAT1 is not A-to-I edited, indicating that the molecular interactions of the DBHS proteins to each RNA may differ. Moreover, the NEAT1 genes in mouse and human vary greatly in sequence, suggesting that likely RNA structure, rather than primary sequence, is conserved. Changes in NEAT1 RNA levels can inform about paraspeckle function, because NEAT1 is linked to controlling the formation of paraspeckles. A relevant example of this is the recent

A.H. Fox and A. Lamond

A

Human Chr 11 (q13.1)

55kb

NEAT1_v2

MALAT-1

NEAT1_v1

Mouse Chr 19 (qA)

20kb

NEAT1_v2

MALAT-1

NEAT1_v1

B

C NEAT1 locus (red)

NEAT1 RNA

P54NRB

Paraspeckles (green)

Linescan P54NRB NEAT1

Figure 4. Paraspeckles contain NEAT1 ncRNA and form near to the NEAT1 gene. (A) NEAT1 and MALAT-1 gene loci on human chromosome 11 q13.1 and mouse chromosome 19qA. Two transcripts are produced from the NEAT1 gene, 3.7kb and 23kb in humans, and 3 kb and approximately 20 kb in mouse. (B) RNA-FISH with probes to NEAT1 ncRNA (green) and immunofluorescence against P54NRB (red) colocalizing in paraspeckles. The line scan is taken over the line indicated in the merged image (lower panels). (C ) A HeLa cell in interphase with combined NEAT1 RNA-FISH to mark paraspeckles (green) and chromosome 11 q13.1 DNA-FISH (red). Panels B and C are reproduced from (Clemson et al. 2009) with permission. Scale bar in B, 10 mm, in panel C, 5 mm.

finding that human embryonic stem cells have little or no detectable NEAT1 RNA and do not show paraspeckles (Chen and Carmichael, 2009). However, hESCs do express all three DBHS proteins, and hESCs also express mRNAs containing A-to-I edited inverted repeats. Critically, within hESCs these A-to-I edited RNAs are not exclusively retained in the nucleus, instead they also appear in the cytoplasm. When the hESCs are induced to differentiate, NEAT1 is expressed and paraspeckles appear.

Importantly, the appearance of paraspeckles is linked to an increase in the efficiency of nuclear retention of the A-to-I edited mRNAs, suggesting that it is NEAT1 forming the paraspeckle structures themselves that are critical for mediating the nuclear retention mechanism. This study highlights the importance of organising the DBHS proteins into paraspeckles for RNA nuclear retention, but also reveals a clear link between the formation of paraspeckles and cellular differentiation. A role for paraspeckles in

Paraspeckles

Paraspeckle Formation and Function

The model for paraspeckle formation begins with the production of NEAT1 transcripts in daughter nuclei following cell division. Once transcribed, NEAT1 molecules form complexes with DBHS proteins, generally before the RNA has had a chance to diffuse far away from its gene locus (Fig. 5). The finished paraspeckle likely consists of multiple copies of NEAT1 RNA-DBHS protein complexes, which form a structural scaffold that is nevertheless dynamic, in that individual DBHS protein molecules in paraspeckles can exchange with a pool of DBHS proteins in the nucleoplasm. It is possible that both the known oligomerization propensity of the DBHS proteins, as well as possible RNARNA intramolecular interactions in NEAT1 ncRNA both contribute to the paraspeckle structural lattice. Without the production of NEAT1 RNA, paraspeckles fail to form, explaining why paraspeckles are not observed when all RNA Pol II transcription is inhibited, or in cell types that do not express NEAT1. Conversely, without abundant DBHS proteins, paraspeckles are also not observed. Throughout interphase, some paraspeckles are observed closely associated with the NEAT1 gene, however, it is not

Ca p

Splicing speckle

CTNRNA

Splicing speckle

Paraspeckle

A AAA

CONCLUSIONS

Nucleolus

II Pol

differentiation was first suggested by Sunwoo et al. (2009), who found that NEAT1 was upregulated when a cultured myoblast cell-line was induced to differentiate into myotubes, with a corresponding increase in paraspeckle size. Another example of varying NEAT1 RNA levels was reported in an earlier study that found levels of VINC-1 RNA (subsequently found to correspond to NEAT1) increased in the central nervous system of mice infected with Japanese encephalitis or rabies viruses (Saha et al. 2006). Although changes in paraspeckles in this experimental model are yet to be evaluated, these viruses may trigger an increase in paraspeckle size and number, raising the possibility that certain viruses may use paraspeckles for their processing, or that paraspeckles are part of a viral defense mechanism.

DBHS proteins ?

Other I-RNA

Neat1 RNA Chromatin

Figure 5. Model of paraspeckle formation. A paraspeckle forms near the NEAT1 gene locus within the interchromatin space, abutting a nuclear speckle. The paraspeckle is formed via interactions between the NEAT1 RNA and DBHS proteins. Additional RNA species regulated within paraspeckles and elsewhere in the nucleus, such as A-to-I edited mRNA, are likely recruited via interaction with DBHS proteins, and traffic through the paraspeckle. Under steady-state conditions the DBHS proteins are dynamic, and exchange between paraspeckles, the nucleoplasm and the nucleolus. When RNA Pol II transcription is inhibited, DBHS proteins accumulate at perinucleolar caps.

known if these are exclusively newly formed paraspeckles, nor is it known what mechanism may be holding the paraspeckles close to the locus. A major function of paraspeckles likely relates to their other known RNA component, namely the mRNA containing dsRNA structures, generally formed by inverted repeats, and A-to-I edited. The RNA nuclear retention mechanism may be involved in many cellular processes such as stress responses, viral infection and circadian rhythm maintenance. Perhaps most significantly, given the link between paraspeckles and two different models of differentiation, it is likely that paraspeckles are playing a part in the reprogramming of a cell that takes place with differentiation, possibly by

A.H. Fox and A. Lamond

altering the expression of key proteins via RNA nuclear retention. The corollary of this is that the absence of both NEAT1 and paraspeckles can potentially be used as a marker for pluripotency (Chen and Carmichael 2009). As yet, the roles of non-DBHS proteins known to localize to paraspeckles have not been addressed within the context of paraspeckle function. Some proteins could be involved in the control of gene expression through nuclear retention, for example, CFIm may be the cleavage factor mediating release of RNA, whereas others proteins with roles in transcription may be involved in recruiting paraspeckles to certain active gene loci. Nuclear RNA and Disease

As yet there are no examples of diseases caused by the absence or presence of paraspeckles, however, there are several RNA dominant diseases associated with the production of toxic nuclear RNAs (Osborne and Thornton 2006). An example is myotonic muscular dystrophy, in which RNA transcribed from mutated genes with expanded CTG-repeats is retained in the nucleus within subnuclear foci. The repeats form RNA hairpins that are bound by the muscleblind-like family of proteins. The toxicity arises as the muscleblind-like proteins are effectively sequestered in the foci and are no longer able to carry out their roles as modulators of alternative splicing (O’Rourke and Swanson 2009). These CUG-repeat foci do not colocalize with paraspeckles in patient-derived cells (Clemson et al. 2009). However, the parallels to paraspeckles are striking: specific RNAprotein interactions, retention of RNA in the interchromosome space and splicing functions of the proteins. These parallels suggest that the cell has common themes in nuclear retention of RNA that are apparent in both normal cell function and disease. Paraspeckles as a Paradigm for a Class of Subnuclear Bodies

Given that the complexity of the mammalian transcriptome is only just beginning to be

explored, including the full extent of long ncRNA production, it is exciting to speculate that paraspeckles will not be the only example of a subnuclear structure formed around a long ncRNA. Indeed, there already exist examples of long ncRNA localizing to unique subnuclear foci, although in these cases no corresponding protein partners are known (Royo et al. 2007; Sone et al. 2007). One interesting possibility is that the IGAZ (structures labeled in the EM by paraspeckle markers) may not be solely composed of paraspeckles, but may also include other subnuclear bodies akin to paraspeckles, each containing distinct structural ncRNAs, specific RNA-binding proteins and having different species of RNA regulated/ retained within them. The lack of identified marker ncRNAs and proteins for these bodies may have so far prevented their detection and characterization. Further studies on long ncRNAs and their subcellular localization will in future provide a fuller picture of these structures and their roles in the cell. CONCLUDING REMARKS

Paraspeckles were identified as recently as 2002, making them one of the ‘youngest’ nuclear structures known. However, already in the short time since their discovery, much has been learned about their composition, formation and function. Paraspeckles are the first, but likely not the last example of a subnuclear body that forms dependent on a long ncRNA, whose function, at least at this point, appears to be to form this subcellular structure. Paraspeckles are also critical for a novel mechanism for controlling gene expression, i.e., the nuclear retention of otherwise translation-competent RNA. We suggest that many key molecules may be regulated in this manner and discovering their identity and functions will be of great interest in the years to come. ACKNOWLEDGMENTS

We thank all those authors who have agreed for their published work to appear in figures within this review. We would also like to thank Charles

Paraspeckles

Bond (University of WA, Australia), Sam Swift and Silvana van Koningsbruggen (University of Dundee, UK) for help generating figures and Sylvie Souquere and Gerard Pierron, (Villejuif, France) for providing TEM images. AIL is a Wellcome Trust Principal Research Fellow. AHF is funded by the National Health & Medical Research Council, Australia. REFERENCES Andersen JS, Lyon CE, Fox AH, Leung AKL, Lam YW, Steen H, Mann M, Lamond AI. 2002. Directed proteomic analysis of the human nucleolus. Current Biology 12: 1– 11. Auboeuf D, Dowhan DH, Li X, Larkin K, Ko L, Berget SM, O’Malley BW. 2004. CoAA, a nuclear receptor coactivator protein at the interface of transcriptional coactivation and RNA splicing. Mol Cell Biol 24: 442–453. Brooks YS, Wang G, Yang Z, Smith KK, Bieberich E, Ko L. 2009. Functional pre-mRNATrans-splicing of coactivator CoAA and corepressor RBM4 during stem/progenitor cell differentiation. J Biol Chem 284: 18033– 18046. Brown SA, Ripperger J, Kadener S, Fleury-Olela F, Vilbois F, Rosbash M, Schibler U. 2005. PERIOD1-associated proteins modulate the negative limb of the mammalian circadian oscillator. Science 308: 693–696. Cardinale S, Cisterna B, Bonetti P, Aringhieri C, Biggiogera M, Barabino SM. 2007. Subnuclear localization and dynamics of the Pre-mRNA 3’ end processing factor mammalian cleavage factor I 68-kDa subunit. Mol Biol Cell 18: 1282–1292. Carninci P, Kasukawa T, Katayama S, Gough J, Frith MC, Maeda N, Oyama R, Ravasi T, Lenhard B, Wells C, et al. 2005. The transcriptional landscape of the mammalian genome. Science 309: 1559–1563. Chen LL, Carmichael GG. 2008. Gene regulation by SINES and inosines: Biological consequences of A-to-I editing of Alu element inverted repeats. Cell Cycle 7: 3294– 3301. Chen LL, Carmichael GG. 2009. Altered nuclear retention of mRNAs containing inverted repeats in human embryonic stem cells: Functional role of a nuclear noncoding RNA. Mol Cell DOI: 101016/jmolcel200906027 Chen LL, DeCerbo JN, Carmichael GG. 2008. Alu elementmediated gene silencing. EMBO J 27: 1694–1705. Clemson CM, Hutchinson JN, Sara SA, Ensminger AW, Fox AH, Chess A, Lawrence JB. 2009. An architectural role for a nuclear noncoding RNA: NEAT1 RNA is essential for the structure of paraspeckles. Mol Cell 33: 717 –726. Cremer T, Cremer M, Dietzel S, Muller S, Solovei I, Fakan S. 2006. Chromosome territories–a functional nuclear landscape. Curr Opin Cell Biol 18: 307– 316. Dettwiler S, Aringhieri C, Cardinale S, Keller W, Barabino SM. 2004. Distinct sequence motifs within the 68-kDa subunit of cleavage factor Im mediate RNA binding, protein-protein interactions, and subcellular localization. J Biol Chem 279: 35788–35797. Dutton JR, Lahiri D, Ward A. 2006. Different isoforms of the Wilms’ tumour protein WT1 have distinct patterns of

distribution and trafficking within the nucleus. Cell Prolif 39: 519– 535. Dye BT, Patton JG. 2001. An RNA recognition motif (RRM) is required for the localization of PTB-associated splicing factor (PSF) to subnuclear speckles. Exp Cell Res 263: 131 –144. Fox AH, Bond CS, Lamond AI. 2005. P54nrb forms a heterodimer with PSP1 that localizes to paraspeckles in an RNA-dependent manner. Mol Biol Cell 16: 5304– 5315. Fox AH, Lam YW, Leung AK, Lyon CE, Andersen J, Mann M, Lamond AI. 2002. Paraspeckles: A novel nuclear domain. Curr Biol 12: 13– 25. Hata K, Nishimura R, Muramatsu S, Matsuda A, Matsubara T, Amano K, Ikeda F, Harley VR, Yoneda T. 2008. Paraspeckle protein p54nrb links Sox9-mediated transcription with RNA processing during chondrogenesis in mice. J Clin Invest 118: 3098– 3108. Hutchinson JN, Ensminger AW, Clemson CM, Lynch CR, Lawrence JB, Chess A. 2007. A screen for nuclear transcripts identifies two linked noncoding RNAs associated with SC35 splicing domains. BMC Genomics 8: 39. Iseli C, Stevenson BJ, de Souza SJ, Samaia HB, Camargo AA, Buetow KH, Strausberg RL, Simpson AJ, Bucher P, Jongeneel CV. 2002. Long-range heterogeneity at the 3’ ends of human mRNAs. Genome Res 12: 1068–1074. Ito T, Watanabe H, Yamamichi N, Kondo S, Tando T, Haraguchi T, Mizutani T, Sakurai K, Fujita S, Izumi T, et al. 2008. Brm transactivates the telomerase reverse transcriptase (TERT) gene and modulates the splicing patterns of its transcripts in concert with p54(nrb). Biochem J 411: 201 –209. Iwasaki T, Chin WW, Ko L. 2001. Identification and characterization of RRM-containing coactivator activator (CoAA) as TRBP-interacting protein, and its splice variant as a coactivator modulator (CoAM). J Biol Chem 276: 33375–33383. Kameoka S, Duque P, Konarska MM. 2004. p54(nrb) associates with the 5’ splice site within large transcription/ splicing complexes. EMBO J 23: 1782– 1791. Kanai Y, Dohmae N, Hirokawa N. 2004. Kinesin transports RNA: Isolation and characterization of an RNAtransporting granule. Neuron 43: 513 –525. Kaneko S, Rozenblatt-Rosen O, Meyerson M, Manley JL. 2007. The multifunctional protein p54nrb/PSF recruits the exonuclease XRN2 to facilitate pre-mRNA 3’ processing and transcription termination. Genes Dev 21: 1779–1789. Kay RA, Ellis IR, Jones SJ, Perrier S, Florence MM, Schor AM, Schor SL. 2005. The expression of migration stimulating factor, a potent oncofetal cytokine, is uniquely controlled by 3’-untranslated region-dependent nuclear sequestration of its precursor messenger RNA. Cancer Res 65: 10742–10749. Kiesler E, Miralles F, Ostlund Farrants AK, Visa N. 2003. The Hrp65 self-interaction is mediated by an evolutionarily conserved domain and is required for nuclear import of Hrp65 isoforms that lack a nuclear localization signal. J Cell Sci 116: 3949– 3956. Lamond AI Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605 –612.

A.H. Fox and A. Lamond Levanon K, Eisenberg E, Rechavi G, Levanon EY. 2005. Letter from the editor: Adenosine-to-inosine RNA editing in Alu repeats in the human genome. EMBO Rep 6: 831–835. Liu H, Ippolito GC, Wall JK, Niu T, Probst L, Lee BS, Pulford K, Banham AH, Stockwin L, Shaffer AL, et al. 2006. Functional studies of BCL11A: Characterization of the conserved BCL11A-XL splice variant and its interaction with BCL6 in nuclear paraspeckles of germinal center B cells. Mol Cancer 5: 18. Mercer TR, Dinger ME, Mattick JS. 2009. Long non-coding RNAs: Insights into functions. Nat Rev Genet 10: 155–159. Myojin R, Kuwahara S, Yasaki T, Matsunaga T, Sakurai T, Kimura M, Uesugi S, Kurihara Y. 2004. Expression and functional significance of mouse paraspeckle protein 1 on spermatogenesis. Biol Reprod 71: 926– 932. O’Rourke JR Swanson MS. 2009. Mechanisms of RNAmediated disease. J Biol Chem 284: 7419–7423. Osborne RJ Thornton CA. 2006. RNA-dominant diseases. Hum Mol Genet 15 Spec No 2: R162– 169. Osenberg S, Dominissini D, Rechavi G, Eisenberg E. 2009. Widespread cleavage of A-to-I hyperediting substrates. RNA 15: 1632– 1639. Patton JG, Porro EB, Galceran J, Tempst P, Nadal-Ginard B. 1993. Cloning and characterization of PSF, a novel premRNA splicing factor. Genes Dev 7: 393 –406. Peng R, Dye BT, Perez I, Barnard DC, Thompson AB, Patton JG. 2002. PSF and p54nrb bind a conserved stem in U5 snRNA. RNA 8: 1334–1347. Platani M, Lamond AI. 2004. Nuclear organisation and subnuclear bodies. Prog Mol Subcell Biol 35: 1 –22. Prasanth KV, Prasanth SG, Xuan Z, Hearn S, Freier SM, Bennett CF, Zhang MQ, Spector DL. 2005. Regulating gene expression through RNA nuclear retention. Cell 123: 249– 263. Prasanth KV, Spector DL. 2007. Eukaryotic regulatory RNAs: An answer to the ‘genome complexity’ conundrum. Genes Dev 21: 11–42. Rivera MN, Kim WJ, Wells J, Stone A, Burger A, Coffman EJ, Zhang J, Haber DA. 2009. The tumor suppressor WTX shuttles to the nucleus and modulates WT1 activity. Proc Natl Acad Sci 106: 8338–8343. Royo H, Basyuk E, Marty V, Marques M, Bertrand E, Cavaille J. 2007. Bsr, a nuclear-retained RNAwith monoallelic expression. Mol Biol Cell 18: 2817–2827. Saha S, Murthy S, Rangarajan PN. 2006. Identification and characterization of a virus-inducible non-coding RNA in mouse brain. J Gen Virol 87: 1991– 1995. Sasaki YT, Ideue T, Sano M, Mituyama T, Hirose T. 2009. MEN1/b noncoding RNAs are essential for structural

integrity of nuclear paraspeckles. Proc Natl Acad Sci 106: 2525– 2530. Shav-Tal Y Zipori D. 2002. PSF and p54(nrb)/NonO– multi-functional nuclear proteins. FEBS Lett 531: 109 –114. Shav-Tal Y, Blechman J, Darzacq X, Montagna C, Dye BT, Patton JG, Singer RH, Zipori D. 2005. Dynamic sorting of nuclear components into distinct nucleolar caps during transcriptional inhibition. Mol Biol Cell 16: 2395–2413. Sone M, Hayashi T, Tarui H, Agata K, Takeichi M, Nakagawa S. 2007. The mRNA-like noncoding RNA Gomafu constitutes a novel nuclear domain in a subset of neurons. J Cell Sci 120: 2498– 2506. Sunwoo H, Dinger ME, Wilusz JE, Amaral PP, Mattick JS, Spector DL. 2009. MEN 1/b nuclear-retained noncoding RNAs are up-regulated upon muscle differentiation and are essential components of paraspeckles. Genome Res 19: 347 –359. Visa N, Puvion-Dutilleul F, Bachellerie JP, Puvion E. 1993. Intranuclear distribution of U1 and U2 snRNAs visualized by high resolution in situ hybridization: Revelation of a novel compartment containing U1 but not U2 snRNA in HeLa cells. Eur J Cell Biol 60: 308– 321. Wilusz JE, Freier SM, Spector DL. 2008. 3’ end processing of a long nuclear-retained noncoding RNA yields a tRNAlike cytoplasmic RNA. Cell 135: 919 –932. Xie SQ, Martin S, Guillot PV, Bentley DL, Pombo A. 2006. Splicing speckles are not reservoirs of RNA polymerase II, but contain an inactive form, phosphorylated on serine2 residues of the C-terminal domain. Mol Biol Cell 17: 1723–1733. Yang YS, Hanke JH, Carayannopoulos L, Craft CM, Capra JD, Tucker PW. 1993. NonO, a non-POU-domaincontaining, octamer-binding protein, is the mammalian homolog of Drosophila nonAdiss. Mol Cell Biol 13: 5593–5603. Yang Z, Sui Y, Xiong S, Liour SS, Phillips AC, Ko L. 2007. Switched alternative splicing of oncogene CoAA during embryonal carcinoma stem cell differentiation. Nucleic Acids Res 35: 1919– 1932. Zhang Z Carmichael GG. 2001. The fate of dsRNA in the nucleus: A p54(nrb)-containing complex mediates the nuclear retention of promiscuously A-to-I edited RNAs. Cell 106: 465 –475. Zolotukhin AS, Michalowski D, Bear J, Smulevitch SV, Traish AM, Peng R, Patton J, Shatsky IN, Felber BK. 2003. PSF acts through the human immunodeficiency virus type 1 mRNA instability elements to regulate virus expression. Mol Cell Biol 23: 6618–6630.

Nuclear Stress Bodies Giuseppe Biamonti1 and Claire Vourc’h2 1

Istituto di Genetica Molecolare. CNR; Via Abbiategrasso 207. 27100 Pavia, Italy

2

Universite´ Joseph Fourier; INSERM; Institut Albert Bonniot U823, La Tronche BP170, 38042 Grenoble cedex 9, France

Correspondence: [email protected] and [email protected]

Nuclear stress bodies (nSBs) are unique subnuclear organelles which form in response to heat shock. They are initiated through a direct interaction between heat shock transcription factor 1 (HSF1) and pericentric tandem repeats of satellite III sequences and correspond to active transcription sites for noncoding satellite III transcripts. Given their unusual features, nSBs are distinct from other known transcription sites. In stressed cells, they are thought to participate in rapid, transient, and global reprogramming of gene expression through different types of mechanisms including chromatin remodeling and trapping of transcription and splicing factors. The analysis of these atypical and intriguing structures uncovers new facets of the relationship between nuclear organization and nuclear function.

uclear stress bodies (or nSBs) were discovered in the late 1980s and very soon after were associated with cellular response to stress agents (Mahl et al. 1989, Sarge et al. 1993). They are transient subnuclear organelles clearly distinct from other nuclear bodies (Cotto et al. 1997). High-resolution electron microscopy analysis has revealed the peculiar and complex organization of nSBs that appear as highly electron-dense structures frequently adjacent to chromatin blocks. The electron-dense core structure consists of a large number of perichromatin granules (PGs) and is surrounded by individual PGs that seem to enter or exit the central core (Chiodi et al. 2000). The function of nSBs is still largely unknown, however, it is commonly accepted that they correspond to highly packed forms of ribonucleoprotein

N

complexes. nSBs are rarely detectable in unstressed cells; their number drastically increases after heat shock as if specific processes involved in the production and/or maturation of specific RNAs were altered in stressed cells. Another distinguishing feature of nSBs is their specificity for human and primate cells (Denegri et al. 2002). Further molecular characterization proved that nSBs originate from the unexpected transcription of large pericentromeric heterochromatic blocks triggered by transcription factors involved in the cell response to stress (Jolly et al. 2004, Rizzi et al. 2004). Intriguingly, these RNAs remain close to the sites of transcription and probably exert their function by recruiting specific factors and affecting chromatin organization. Thus nSBs are at the convergence of several important aspects of

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000695 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000695

G. Biamonti and C. Vourc’h

cell biology such as the epigenetic control of gene expression, noncoding RNAs, and control of RNA splicing activities (reviewed in Biamonti 2004; Jolly and Lakhotia 2006; Eymery et al. 2009a). THE DAWN OF NUCLEAR STRESS BODIES: THE TWO FACES OF MOLECULAR JANUS STRUCTURES

Even a limited increase of the growth temperature of a few degrees Celsius, referred to as “heat shock,” induces in all cell types and organisms, a series of functional and morphological alterations, which are gradually reversed over a period of several hours once the physiological temperature is restored. Heat shock leads to an immediate and almost complete block of important cellular processes such as DNA replication and transcription. At the posttranscriptional level, heat shock transiently inhibits pre-mRNA splicing, nucleo-cytoplasmic transport and translation (Morimoto and Santoro 1998). The mechanisms underlying splicing inhibition are still poorly understood. In 1989, Mahl et al. proposed that heat shock could act by perturbing the integrity of ribonucleoprotein complexes, i.e., the substrates of splicing and of nuclear export. Upon thermal stress, a subset of hnRNP (heterogeneous ribonucleoprotein particles), which are the main protein constituents of ribonucleoprotein complexes, were seen to be recruited to specific nuclear sites which, as visualized by electron microscopy, appear to be enriched in highly packed forms of ribonucleoprotein complexes called “perichromatin granules” (PGs) (Mahl et al. 1989). This was the first report of what we now know as nuclear stress bodies or nSBs. A few years later, another protein, heat shock factor 1 (HSF1), was shown to form a small number of nuclear granules after different types of stress conditions (Sarge et al. 1993). The cellular response to adverse environmental and physiological conditions such as heat shock, or an exposure to amino acid analogs, heavy metals, oxidative stress, anti-inflammatory drugs, or arachidonic acid, leads to a rapid and transient activation of genes encoding heat

shock proteins (hsps) and molecular chaperones (reviewed in Lindquist 1986 and in Christians et al. 2002). Stress-induced transcription is regulated by a family of heat shock transcription factors (HSF). In vertebrates, four members of the HSF gene family (HSFs 1-4) have been characterized (reviewed in Pirkkala et al. 2001), each mediating the response to distinct forms of cellular stress, including HSF1, which responds to the classical inducers of the heat shock response. Whereas in unstressed cells HSF1 is maintained unbound to DNA, after heat shock, it undergoes reversible oligomerization into a DNA binding competent trimer. Two distinct mechanisms, involving negative regulatory domains and phosphorylation, cooperate to control the activity of this factor (reviewed in Cotto and Morimoto 1999). In higher eukaryotes, HSF1 trimers appear within minutes of activation and bind to specific heat shock elements (HSE) in the promoters of heat shock genes (hsp genes). In 1993, Sarge et al. showed that full activation of HSF1, induced by heat shock, cadmium sulfate or by the amino acid analog L-azetidine-2-carboxylic acid, also results in the accumulation of this factor into a small number (four to six) of nuclear granules with a maximum diameter of 2– 2.5 mm (Sarge et al. 1993) (Fig. 1). Intriguingly, these granules were described only in monkey and human cells and were not observed in rodent cells (reviewed in Jolly and Lakhotia 2006). HSF1 granules were also characterized independently (Cotto et al. 1997; Jolly et al. 1997) as novel entities, distinct from other subnuclear compartments. Their kinetics of formation and disappearance depends both on the nature and on the severity (duration, and concentration or intensity) of the stressing agent. More importantly, the number of bodies correlates with cell ploidy. On the basis of this latter finding it was suggested that HSF1 granules could be assembled on specific chromosomal targets and may represent stressdependent transcription sites. However, the lack of colocalization with the classical hsp genes, such as hsp70 and hsp90, initially argued against this model (Jolly et al. 1997). A few years later the hypothesis was revitalized by the in vivo analysis of HSF1 granules with HSF1-GFP

HS

NHS

nSBs and Nuclear Function

DNA

αHSF1

α acLys

Merge

Figure 1. nSBs visualized with anti HSF1 and anti

acetylated lysine antibodies. In unstressed cells (NHS), HSF1 (green labeling) displays a diffuse nucleocytoplasmic distribution. Upon heat-shock (HS), HSF1 is redistributed to a few nuclear foci, also known as stress granules, which form primarily on the 9q12 loci (indicated with arrow heads). An average of three copies of chromosome 9 are presentinHelacells. Inheat-shocked cells, the 9q12 locus is enriched in acetylated histones. The binding of HSF1 is followed by the formation of acetylated foci, the recruitment of RNA polymerase II, the subsequent transcription of sat III repeats mainly into “G rich” transcripts, and the ultimate recruitment of splicing factors. (Bar scale ¼ 5 mm).

(Jolly et al. 1999a). This analysis showed that successive, short rounds of heat shock, induced cycles of assembly/disassembly of HSF1 granules. Interestingly, granules always formed in the same nuclear positions as if bound to an underlying immobile matrix. These experiments, therefore, raised the question about the nature of the chromosomal targets involved. Soon after the first description of HSF1 granules, several studies by the group of Biamonti made the link between these structures and the nuclear bodies containing RNA binding proteins, which had previously been identified by Fuchs et al. (Mahl et al. 1989). These studies started with the characterization of a novel hnRNP protein called hnRNP A1 interacting protein—HAP, identified by others as Saf-B, scaffold attachment factor B (Renz and Fackelmayer 1996), or as HET, the Hsp27-ERE-TATAbinding protein (Oesterreich et al. 1997). After a mild heat shock, HAP/Saf-B is recruited to a small number of nuclear bodies. Importantly, HAP bodies coincide with HSF1 granules, highlighting the double nature of these bodies containing both transcription factors and proteins involved in pre-mRNA metabolism (Weighardt et al. 1999). We now know that HSF1 granules

and HAP bodies define two different functional states, partially overlapping in time, of what we now call nuclear stress bodies or nSBs (reviewed in Biamonti 2004). Although HSF1 granules form at the onset of the heat shock response and rapidly disappear during the recovery period following heat shock, HAP bodies become visible after 1 hour of mild heat shock, with a maximum size reached after 3 hours of recovery. The formation of HAP bodies requires ongoing transcription (Weighardt et al. 1999) and these structures are sensitive to RNAse treatment (Chiodi et al. 2000), suggesting that RNA is a major component of nSBs. The importance of RNA in the assembly of nSBs was further supported by the ultrastructural analysis of nSBs (Chiodi et al. 2000), which appeared as clusters of perichromatin granules surrounded by compact chromatin (Charlier et al. 2009). PGs in the bodies are specifically labeled by antibodies against hnRNP HAP and contain nascent bromouridine-labeled RNA. These structures are clearly distinct from nuclear speckles, where a number of pre-mRNA processing factors also accumulate. A subset of splicing factors of the SR family, including SF2/ASF, SRp30 and 9G8 are efficiently recruited to nSBs, whereas the distribution of other members of the same family, including SC35, the standard marker of nuclear splicing speckles, is not affected by stress (Jolly et al. 1999b; Denegri et al. 2001). THE COMING OF AGE OF NUCLEAR STRESS BODIES

Several studies clarified the nature of nSBs, unveiling unexpected links with the epigenetic organization of large chromosomal blocks and with noncoding RNAs. Nuclear Stress Bodies Assemble on Blocks of Satellite III DNA

In 2002 C. Jolly and C. Vourc’h directly addressed the nature of the chromosomal targets on which nSBs are assembled (Jolly et al. 2002). By investigating the distribution of HSF1 on metaphase chromosome spreads they found

G. Biamonti and C. Vourc’h

that, in all heat-shocked cells, HSF1 binds to the extended pericentric heterochromatic q11-q12 region of human chromosome 9. This region, also known as secondary constriction or 9qh region, is a large block of heterochromatin primarily composed of long tandem arrays of Sat III repeats (Jones et al. 1973). Although the Sat III consensus sequence does not contain canonical HSE elements, purified HSF1 binds to a genomic Sat III fragment (Grady et al. 1992) from human chromosome 9 but not to other satellite repetitive sequences (i.e., Sat II or a-satellite) in in vitro assays (Jolly et al. 2004), demonstrating the existence of direct interaction ofHSF1 with Sat IIIsequences. Intriguingly, HSF1 binding does not require preliminary stressinduced chromatin reorganization because an HSF1 mutant, deleted in its carboxy-terminal trans-activation domain, constitutively binds to the 9qh region in unstressed cells (Jolly et al. 2004). Meanwhile, the group of Biamonti exploited a completely different strategy to identify chromosomal regions involved in the assembly of nSBs (Denegri et al. 2002). Taking advantage of the fact that nSBs occur in human but not in rodent cells they used human-hamster somatic cell hybrids to identify human chromosomes that direct the assemblyof nSBs in hamsters cells. In addition to chromosome 9, which represents the primary target of nSBs formation, they identified two other human chromosomes, 12 and 15, that are positive in this assay. With the same approach they narrowed the region of chromosome 9 required for the formation of nSBs to the 9q12 band identified by Jolly and Vourc’h. This chromosomal band, therefore, acts as the recruiting for nSBs. Transcriptional Activation of Pericentromeric Heterochromatin

Altogether these findings paved the way for more work on this subject, which brought researchers to the crossroads between epigenetics and noncoding RNAs. Most of the data available at that time suggested that nSBs could be large transcription factories. However, this hypothesis was in conflict with the fact that the 9q12 region

was described as a region of noncoding constitutive, and therefore transcriptionally inactive, heterochromatin (Kokalj-Vokac et al. 1993). The idea of a transcription factory gained momentum, however, when it was shown that nSBs were enriched in acetylated histones (Fig. 1), an epigenetic mark of transcriptionally active chromatin, and did not contain typical heterochromatin markers such as HP1 proteins or histone H3 tri-methylated on lysine 9 (Rizzi et al. 2004, Jolly et al. 2004). Moreover, it was also observed that HSF1 binding, through the corecruitment of the histone acetyl transferase CREB binding protein (CBP), initiated a series of events involving chromatin remodeling, the recruitment of RNA pol II, but not of polI or III, and culminated with the production of Sat III transcripts, which bound to several splicing factors (Jollyet al. 2004, Rizzi et al. 2004, Metz et al. 2004). Nuclear Stress Bodies are Part of a General Response to Stress

Theexpression of Sat IIIsequencesin heat-shocked cells now represents one of the best-documented examples of transcriptional activation of pericentric heterochromatin in metazoan cells. In addition to heat shock, the expression of Sat III sequences can also be induced by the amino acid analog azetidine and by a variety of physical (UV-light) and chemical (Cadmium sulfate) stressors known to activate HSF1 (Valgardsdottir et al. 2008, Sengupta et al. 2009), indicating that this event is part of the gene expression program controlled by HSF1. The kinetics and the extent of the induction vary with the nature of the stress agent, probably reflecting the robustness of HSF1 activation. Interestingly, in all cases Sat III RNAs remain associated with nSBs as they do in heat-shocked cells (Jolly et al. 2004) and are undetectable in the cytoplasm (Jolly et al. 2004; Valgardsdottir et al. 2005), as if these molecules exerted their action on chromatin. HSF2, an isoform of HSF1 that forms heterotrimers with HSF1 and modulates its activity (Sandqvist et al. 2009), is also present in nSBs where it binds to DNA in an HSF1-dependent manner (Alastalo et al. 2003). Depletion of

nSBs and Nuclear Function

HSF2 leads to an increase of the heat induced transcription of Sat III sequences whereas, intriguingly, elevated HSF2 expression, mimicking what is observed in development, activates Sat III transcription in unstressed cells (Sandqvist et al. 2009). Although the functional implication of the HSF2/HSF1 interaction is still largely undefined, this result clearly indicates that Sat III expression may play a role in various physiological settings. This idea is also suggested by the observation that another transcription factor, the tonicity enhancer binding protein (TonEBP), also directs the formation of nSBs and the transcription of Sat III sequences in response to hyper-osmotic stress (Valgardsdottir et al. 2008). Notably, TonEBP is physiologically crucial for the formation and function of kidney protecting cells in the renal inner medullafrom extraordinarily high levels of NaCl and urea. Putative binding sites for this factor are present in the Sat III sequence. Thus, distinct signaling pathways elicited by different stress inducers and acting through different transcription factors lead to the production of Sat III RNAs (Valgardsdottir et al. 2008). This is suggestive of an active role of Sat III sequences in the cell response to stress. Interestingly, besides environmental stress, other physiological and pathological conditions lead to the activation of pericentric sequences in human cells; however, in none of these cases has the presence of nSB-like structures been documented. Frequently, the expression is linked to a change in the epigenetic organization of Sat III sequences and pericentric regions in general. For instance, expression of Sat III sequences is observed in cells treated with 5-Azacytidin, a potent inhibitor of DNA methylation (Eymery et al. 2009b). The expression of pericentric transcripts also occurs during replicative senescence at late passages of both primary fibroblasts and cancer cells (Enukashvily et al. 2007; Eymery et al. 2009b). The fact that an alteration of the heterochromatin structure may favor an accumulation of Sat III transcripts is also suggested by observations made in fibroblasts from patients affected by the Hutchinson – Gilford progeria syndrome (HPGS), in which a complete loss of the heterochromatic marks is accompanied by the expression of

chromosome 9 specific Sat III sequences (Shumaker et al. 2006). Another example is expression of Sat III transcripts in human testis, suggesting that Sat III transcripts may be somehow involved in the differentiation of germinal cells (Jehan et al. 2007, Sandqvist et al. 2009). These findings, together with a recent study showing expression of Sat III transcripts in embryonic cells (Faulkner et al. 2009), potentially link the expression of Sat III sequences to developmental programs. The occurrence of transcription in normally silent portions of the genome, regardless of the formation of specialized nuclear structures, raises a number of questions about the role of pericentric heterochromatin and the impact of their transcriptional awakening. Does activation of Sat III arrays participate in global stress-induced, genome-wide down regulation of genome expression through transient sequestration of transcription factors? Alternatively, does activation of Sat III facilitate transcription of nearby genes through cis-acting effects or by creating nuclear domains in which genes escaping heat-induced transcriptional repression would relocate? Is gene expression also affected by transient targeting of specific splicing factors to Sat III RNAs? Finally, why do noncoding Sat III RNAs remain associated with, or in proximity to, arrays of SatIII sequences in the human genome? As discussed later, an answer might be that transcripts of pericentric origin are known to play a role in the establishment and maintenance of heterochromatin structure. Addressing these questions will be a major subject of future investigation. THE ELUSIVE NONCODING SATELLITE III RNAs

The function of the SatIII transcripts in nSBs is unknown but they do have several intriguing properties. First, heat shock drastically induces the strong expression of polyadenylated Sat III RNAs, mainly corresponding to the G-rich strand (the presence of GGAAT repeats in Sat III transcripts imposes a difference in the G/C content between the two complementary DNA strands) (Jolly et al. 2004; Valgardsdottir et al. 2005 and 2008). It is worth noticing that

G. Biamonti and C. Vourc’h

transcription of Sat III sequences occurs even in unstressed cells (Valgardsdottir et al. 2008). Because of the repetitive nature of the transcripts and uncertainties about their size (see later dicussion), it is impossible to assess the level of Sat III RNA molecules, which, however, appears to be low. The expression of Sat III sequences in unstressed cells may change our view of the role of pericentric heterochromatin, and is consistent with the observation that a subset of Sat III DNA sequences exists in an open, transcriptionally permissive state of chromatin in human cells (Gilbert et al. 2004). Another important aspect is the fact that the kinetics of accumulation of Sat III transcripts depend on cell type and the nature of the stressing agent. After mild heat shock (1 hour at 42 8C) the level of these RNAs peaks at 2– 3 hour of recovery. However, because of their stability, the level of Sat III transcripts are still higher than in unstressed cells after one day of recovery (Jolly et al. 2004 and Valgardsdottir et al. 2008). Unlike protein coding transcripts, Sat III transcripts always remain associated with—or in close proximity to— the locus from which they originate (Jolly et al. 2004). Discrepancies exist between estimates of the length of the Sat III transcripts. According to some authors (Jolly et al. 2004), Sat III RNAs are larger than 10 Kb whereas others (Rizzi et al. 2004) have observed that Sat III RNAs have a broad size distribution with most of the molecules falling between 5 and 2 kb. The possibility that shorter transcripts could derive from long precursors through post-transcriptional processing, i.e., splicing, remains to be established. The level of the expression induction of these sequences has also recently been measured by real time RT-PCR (Valgardsdottir et al. 2008), showing an induction between 10,000and 100,000-fold for G-rich and 50-fold for C-rich Sat III sequences, and a peak at 2 h of recovery. Because of the repetitive nature of Sat III sequences, the actual induction of Sat III RNA molecules is certainly lower than this value. Because of our lack of knowledge concerning the structure of the repetitive Sat III arrays, it is not yet clear whether the production

of Sat III RNAs is driven by a canonical promoter. It is also conceivable that transcripts start randomly in the proximity of HSF1 molecules bound to HSE elements. Finally, nothing is known so far about the fate of these transcripts and the machinery involved in their degradation. AN OPEN CHALLENGE: FINDING (A) FUNCTION(S) FOR nSBs AND SAT III TRANSCRIPTS

Sat III sequences have appeared late in evolution (Jarmuz et al. 2007). They are specific to the Hominoidea superfamily and are present on most human chromosomes. Their recent evolution accounts for the fact that Sat III RNAs and nSBs, or similar recruiting centers for RNA processing factors, are not found in rodent cells. This raises questions about the function of these sequences and suggests that other noncoding RNAs may play similar functions in different species. Interestingly, structures comparable to nSBs were found in Drosophila cells and called v-speckles (Prasanth et al. 2000). Similarly to Sat III RNA, v-speckles control the dynamics of pre-mRNA processing factors in heatshocked cells. This evolutionary convergence of otherwise different systems (v-transcripts derive from a single copy gene rather than from repetitive satellite elements) indicates that nSBs could be involved in splicing regulation. Molecular Traps for Transcription and Splicing Factors?

nSBs may be viewed as transcription factories comprising a natural amplification of RNA polII promoters. Although the number of transcription units is not yet defined, both the extent of the Sat III arrays and the size of HSF1 foci suggest that thousands of transcriptional units may be simultaneously activated. Thus, the massive concentration of factors involved in the activation of Sat III sequences such as RNA polII or the histone acetyl transferase CBP may result in the transient sequestration of transcription factors from the surrounding nucleoplasm. This is in agreement with the

nSBs and Nuclear Function

observation that, in heat-shocked cells, the formation of nSBs is accompanied by a global deacetylation of chromatin in the rest of the nucleus (Fritah et al. 2009). A similar hypothesis can be proposed for factors involved in pre-mRNA processing. Sat III RNAs are stable components of nSBs and mediate the recruitment of a number of proteins involved in premRNA maturation (Denegri et al. 2001; Metz et al. 2004). Some RNA binding proteins are recruited through protein-RNA interactions (Chiodi et al. 2004). The recruitment of the splicing factor SF2/ASF is mediated by the RRM2 (RNA recognition motif )-domain, which is critical for its activity in alternative splicing (Metz et al. 2004, Chiodi et al. 2004), whereas other RNA binding proteins, such as hnRNP HAP, are recruited through protein – protein interactions (Denegri et al. 2001). Notably, although not accumulating in nSBs, other essential components of the splicing machinery, i.e., the snRNPs, are associated with Sat III RNAs (Metz et al. 2004). This is in line with the hypothesis that Sat III RNAs undergo at least some steps in the splicing reaction. In this context it is worth recalling that the splicing reaction is either blocked or delayed by thermal stress (Yost and Lindquist 1986; Bond 1988; reviewed in Bond 2006). It is intriguing that nSBs correspond to large clusters of PGs. Although the nature of PGs has not yet been elucidated, it has been suggested that they could contain aberrant RNAs blocked at early stages of maturation and engaged in degradation (Cervera 1979; Puvion and Viron 1981). Disruption of RNA maturation at an early stage of the process, as occurring after heat shock or in the presence of transcription inhibitors, could block RNA on its DNA matrix, leading to the formation of PGs. Although the heat shock response correlates with a global shut-down of transcription and with an alteration of splicing functions (Yost and Lindquist 1986; Bond 1988; reviewed in Bond 2006), it is not entirely clear whether it affects the majority of pre-mRNAs, whether all transcripts are affected to a similar degree, or whether heat shock targets only specific subsets

of pre-mRNAs. Whatever the extent of this phenomenon, the activation of Sat III sequences could contribute to the shutdown or reprogramming of gene expression. Alternative splicing affects more than 90% of cellular transcripts. Splicing profiles are controlled by the relative abundance of antagonistic hnRNP and SR proteins. It is plausible that Sat III RNAs, by sequestering specific RNA binding proteins into nSBs, may shift splicing decisions, as for instance toward the synthesis of molecules involved in the cell defense to stress (Fig. 2A). nSBs are not the only nuclear bodies whose assembly involves a specific RNA molecule. This is also the case of paraspeckles, which are subnuclear foci found adjacent to nuclear splicing speckles, that contribute to regulation of gene expression by trapping adenosine to inosine (A to I) hyperedited RNA within the nucleus (reviewed in Bond and Fox 2009). Similarly to nSBs, formation of paraspeckles involves the interaction of the NEAT1 (also known as MEN-1/b or VINC-1) noncoding RNA (ncRNA) with several RNA binding proteins, i.e., members of the DBHS (Drosophila melanogaster behavior, human splicing) protein family, consisting of PSPC1, P54NRB/NONO, or PSF/SFPQ. As for nSBs, the RNA moiety (NEAT1 ncRNA) is essential for the maintenance of the body and is also the nucleating factor (Sasaki et al. 2009, Clemson et al. 2009, Chen and Carmicael 2009, Sunwood et al. 2009). Indeed, paraspeckles form in early G1 near to the NEAT1 gene locus and are often found clustered near the NEAT1 gene in interphase. There are also important differences between the two bodies. Contrary to heterogeneous Sat III RNAs, NEAT1 is a well-characterized transcript encoded by a single-copy gene locus. Moreover, nSBs remain always associated or in close proximity with the sites of transcription of Sat III RNAs whereas paraspeckles can leave NEAT1 transcription sites to associate with nuclear speckles. The mechanisms allowing the existence of paraspeckles at distance from the sites of NEAT1 transcription are still unknown.

G. Biamonti and C. Vourc’h

A

B Sat III repeats HSf1

nSB

nSB

C Sat III repeats

Adjacent loci

HSf1

TFs

Figure 2. Schematic illustration of the possible roles of nSBs in heat-shocked cells. nSBs are thought to play a role

in the cellular response to stress and cell protection. Three main hypotheses have been proposed which are not mutually exclusive: (A) Control of transcription and splicing activities. Upon heat-shock, sat III sequences and transcripts are thought to play a role in the control of transcriptional and splicing activities through sequestration of transcription and splicing factors (both represented as dots). Transient trapping of these factors could contribute to the shutdown or reprogramming of gene expression. It is also plausible that Sat III RNAs, by sequestering specific RNA-binding proteins into nSBs, may influence splicing decisions toward the synthesis of molecules involved in the cell defense to stress. (B) Regeneration of heterochromatin structure. In fission yeast, transcripts from pericentric regions play a role in the formation and maintenance of heterochromatin. In human cells, sat III transcripts may also play a role in protecting heterochromatic pericentric regions following heat-shock, either as long RNA molecules or as small RNA molecules generated by the RNAi machinery. (C) Transcriptional de-repression of genes located in the vicinity of nSBs through position effects. Loss of epigenetic repressive marks (red flags) at the 9q12 locus following heat shock could abolish the transcriptional repression exerted by pericentric heterochromatin on the activity of promoter genes present in cis (here visualized in brown and green) or possibly in trans (not shown) through chromatin opening and binding of transcription factors (TF).

nSBs and Nuclear Function

A Role in Heterochromatin Assembly and/or Maintenance?

RNA appears to have a major role in the assembly of heterochromatin (Maison et al. 2002; Muchardt et al. 2002). Two mechanisms have been so far identified through which RNA may act. In Schizosaccharomyces pombe, small-sized pericentric transcripts, generated through the processing of longer RNAs by the endoribonuclease Dicer, target the RITS (RNA induced transcriptional silencing) complex to complementary DNA sequences and direct the assembly of heterochromatin (Verdel et al. 2004; Buhler et al. 2006). One can speculate that, similarly, Sat III transcripts may be processed into small dsRNA, which would then be recruited to the human RITS complex (reviewed in Eymery et al. 2009a). In support of this hypothesis, in a chicken – human cell hybrid containing human chromosome 21, a loss of Dicer leads to the accumulation of pericentric specific transcripts and results in cell death and premature sister chromatin separation (Fukagawa et al. 2004). Moreover, real time RT-PCR indicates that, both in unstressed and heat shock cells, the level of C-rich Sat III RNAs is drastically lower than that of the complementary G-rich molecules (Valgardsdottir et al. 2008). This difference in complementary transcripts clearly argues against the existence of long double stranded Sat III RNAs that, as in S. pombe, would then be processed by Dicer. Moreover, so far, no evidence of short Sat III RNAs in human cells has been found. The establishment of heterochromatin can also involve long noncoding RNAs through still poorly understood mechanisms. This is the case for the Xist transcript involved in female X chromosome inactivation (reviewed in Heard 2004). Because Sat III transcripts are stable transcripts that remain associated with the 9q12 regions even through the G2/M transition (Jolly et al., 2004), one can speculate that long Sat III RNAs may be involved in a similar process. Finally, it has been suggested that Sat III RNAs may have a role in stabilizing chromosomal regions that are prone to instability and rearrangements (Bartlett et al. 1988; Lamszus et al.

1999; reviewed in Robertson and Wolffe 2000 and in Duker 2002; Reshmi-Skarja et al. 2003) (Fig. 2B). Impact of nSBs on Nuclear Organization

The transcriptional activation of pericentric sequences could profoundly affect the functional organization of the cell nucleus. Pericentric chromatin is thought to influence the expression of genes, either on the same chromosome or within the nuclear volume, through cis or transmechanisms. Several examples have been reported in the literature in which gene inactivation is indeed associated with repositioning of repressed genes in the vicinity of these large blocks of heterochromatin. Based on these observations a model has been proposed in which repositioning of repressive genes in the vicinity of heterochromatin would be necessary for the maintenance of their repressed status through a position effect mechanism (reviewed in Fisher et al. 2002; Francastel et al. 2001; Zhimulev and Belyaeva 2003). It is conceivable that transcriptional activation of Sat III sequences may impact on the activity of other genes associated either on the same chromosome or in the nuclear space. This could be one of the multiple ways by which stress may induce a transient reprogramming of gene expression profiles (Fig. 2C). CONCLUDING REMARKS

Although nSBs have not revealed all of their nature and function, a picture emerges today of a large center for the recruitment of transcription and splicing factors, involved in the global control of gene expression. If this hypothesis proves to be correct, it would represent a new remarkable illustration of the ingenuity of the mechanisms implemented by the cell to quickly adapt to environmental changes, and ensure its survival. Our poor knowledge of the structure of Sat III encoding regions and related transcripts still represents a barrier to functional analysis. However, combination of in situ approaches to determine the kinetics of formation and disappearance of nSBs, coupled

G. Biamonti and C. Vourc’h

with epigenetic and transcriptomic genomewide approaches can now be performed to determine the impact of nSBs formation on global changes of gene activity in the course of the stress response. Answering this question represents an important and exciting challenge for the next decade. ACKOWLEDGMENTS

We would like to thank Dr S.Pison-Rousseaux for her helpful suggestions. G. Biamonti is supported by grants from AIRC, Cariplo Foundation and from European Union (EURASNET) Network of Excellence on Alternative Splicing (EURASNET). C. Vourc’h is supported by grants from the Institut National du Cancer (EPISTRESS project), by Cance´ropoˆle Lyon Auvergne Rhoˆne Alpes (EpiPro, EpiMed) and by ARC (grant#3449). REFERENCES Alastalo T-P, Hellesuo M, Sandqvist A, Hietakangas V, Kallio M, Sistonen L. 2003. Formation of nuclear stress granules involves HSF2 and coincides with the nucleolar localization of Hsp70. J Cell Sci 116: 3557–3570. Bartlett JM, Watters AD, Ballantine SA, Going JJ, Grigor KM, Cooke TG. 1998. Is chromosome 9 a marker of disease recurrence in transitional cell carcinoma of the urinary bladder? Br J Cancer 7: 2193–2198. Biamonti G. 2004. Nuclear stress bodies: A heterochromatin affair? Nat Rev Mol Cell Biol 5: 493– 498. Bond U. 1988. Heat shock but not other stress inducers leads to the disruption of a sub-set of snRNPs and inhibition of in vitro splicing in HeLa cells. EMBO J 7: 3509– 3518. Erratum in EMBO J 7: 4020. Bond U. 2006. Stressed out! Effects of environmental stress on mRNA metabolism. FEMS Yeast Res 6: 160– 170. Bond CS, Fox AH. 2009. Paraspeckles; nuclear bodies on long noncoding RNA. J Cell Biol 186: 637–644. Buhler M, Verdel A, Moazed D. 2006. Tethering RITS to a nascent transcript initiates RNAi- and heterochromatindependent gene silencing. Cell 125: 873 –886. Cervera J. 1979. Effects of thermic shock on HEp-2 cells. II. Inhibition of induction of perichromatin granules by cordycepin and actinomycin D. J Ultrastruct Res 66: 182–189. Charlier C, Lamaye F, Thelen N, Thiry M. 2009. Ultrastructural detection of nucleic acids within heat shockinduced perichromatin granules of HeLa cells by cytochemical and immunocytological methods. J Struct Biol 166: 329– 336. Chen LL, Camichael GG. 2009. Altered nuclear retention of mRNAs containing inverted repeats in human

embryonic stem cells: Functional role of a nuclear noncoding RNA. Mol Cell 35: 467– 478. Chiodi I, Biggiogera M, Denegri M, Corioni M, Weighardt F, Cobianchi F, Riva S, Biamonti G. 2000. Structure and dynamics of hnRNP-labelled nuclear bodies induced by stress treatments. J Cell Sci 113: 4043–4053. Chiodi I, Corioni M, Giordano M, Valgardsdottir R, Ghigna C, Cobianchi F, Xu RM, Riva S, Biamonti G. 2004. RNA recognition motif 2 directs the recruitment of SF2/ASF to nuclear stress bodies. Nucleic Acids Res 32: 4127–4136. Clemson CM, Hutchinson JN, Sara SA, Ensminger AW, Fox AH, Chess A, Lawrence JB. 2009. An architectural role for a nuclear noncoding RNA: NEAT1 RNA is essential for the structure of paraspeckles. Mol Cell 33: 717– 726. Cotto JJ, Morimoto RI. 1999. Stress-induced activation of the heat-shock response: Cell and molecular biology of heat-shock factors. Biochem Soc Symp 64: 105– 118. Cotto J, Fox S, Morimoto RI. 1997. HSF1 granules: A novel stress-induced nuclear compartment of human cells. J Cell Sci 110: 2925– 2934. Christians ES, Yan LJ, Benjamin IJ. 2002. Heat shock factor 1 and heat shock proteins: Critical partners in protection against acute cell injury. Crit Care Med 30: S43 –50. Denegri M, Chiodi I, Corioni M, Cobianchi F, Riva S, Biamonti G. 2001. Stress-induced nuclear bodies are sites of accumulation of pre-mRNA processing factors. Mol Biol Cell 12: 3502–3514. Denegri M, Moralli D, Rocchi M, Biggiogera M, Raimondi R, Cobianchi F, De Carli L, Riva S, Biamonti G. 2002. Human chromosomes 9, 12, and 15 contain the nucleation sites of stress-induced nuclear bodies. Mol Biol Cell 13: 2069–2079. Duker NJ. 2002. Chromosome breakage syndromes and cancer. Am J Med Genet 115: 125– 129. Eymery A, Callanan M, Vourc’h C. 2009a. The secret message of heterochromatin: New insights into the mechanisms and function of centromeric and pericentric repeat sequence transcription. Int J Dev Biol 53: 259– 268. Eymery A, Horard B, El Atifi-Borel M, Fourel G, Berger F, Vitte AL, Van den Broeck A, Brambilla E, Fournier A, Callanan M, et al. 2009b. A transcriptomic analysis of human centromeric and pericentric sequences in normal and tumor cells. Nucleic Acids Res 37: 6340–6354. Enukashvily NI, Donev R, Waisertreiger IS, Podgornaya OI. 2007. Human chromosome 1 satellite 3 DNA is decondensed, demethylated and transcribed in senescent cells and in A431 epithelial carcinoma cells. Cytogenet Genome Res 118: 42–54. Faulkner GJ, Kimura Y, Daub CO, Wani S, Plessy C, Irvine KM, Schroder K, Cloonan N, Steptoe AL, Lassmann T, et al. 2009. The regulated retrotransposon transcriptome of mammalian cells. Nat Genet 41: 563–571. Fisher AG, Merkenschlager M. 2002. Gene silencing, cell fate and nuclear organisation. Curr Opin Genet Dev 12: 193 –197. Francastel C, Magis W, Groudine M. 2001. Nuclear relocation of a transactivator subunit precedes target gene activation. Proc Natl Acad Sci 98: 12120– 12125. Fritah S, Col E, Boyault C, Govin J, Sadoul K, Chiocca S, Christians E, Khochbin S, Jolly C, Vourc’h C. 2009.

nSBs and Nuclear Function Heat Shock Factor 1 controls genome-wide acetylation in heat-shocked cells. Mol Biol Cell 20: 4976–4984. Fukagawa T, Nogami M, Yoshikawa M, Ikeno M, Okazaki T, Takami Y, Nakayama T, Oshimura M. 2004. Dicer is essential for formation of the heterochromatin structure in vertebrate cells. Nat Cell Biol 6: 784 –791. Gilbert N, Boyle S, Fiegler H, Woodfine K, Carter NP, Bickmore WA. 2004. Chromatin architecture of the human genome: Gene-rich domains are enriched in open chromatin fibers. Cell 118: 555–566. Grady DL, Ratliff RL, Robinson DL, McCanlies EC, Meyne J, Moyzis RK. 1992. Highly conserved repetitive DNA sequences are present at human centromeres. Proc Natl Acad Sci 89: 1695–1699. Heard E. 2004. Recent advances in X-chromosome inactivation. Curr Opin Cell Biol 16: 247 –255. Jarmuz M, Glotzbach CD, Bailey KA, Bandyopadhyay R, Shaffer LG. 2007. The Evolution of satellite III DNA subfamilies among primates. Am J Hum Genet 80: 495 –501. Jehan Z, Vallinayagam S, Tiwari S, Pradhan S, Singh L, Suresh A, Reddy HM, Ahuja YR, Jesudasan RA. 2007. Novel noncoding RNA from human Y distal heterochromatic block (Yq12) generates testis-specific chimeric CDC2L2. Genome Res 17: 433– 440. Jolly C, Lakhotia SC. 2006. Human sat III and Drosophila hsr omega transcripts: A common paradigm for regulation of nuclear RNA processing in stressed cells. Nucleic Acids Res 34: 5508–5514. Jolly C, Usson Y, Morimoto RI. 1999a. Rapid and reversible relocalization of heat shock factor 1 within seconds to nuclear stress granules. Proc Natl Acad Sci 96: 6769– 6774. Jolly C, Konecny L, Grady DL, Kutskova YA, Cotto JJ, Morimoto RI, Vourc’h C. 2002. In vivo binding of active heat shock transcription factor 1 to human chromosome 9 heterochromatin during stress. J Cell Biol 156: 775 –781. Jolly C, Metz A, Govin J, Vigneron M, Turner BM, Khochbin S, Vourc’h C. 2004. Stress-induced transcription of satellite III repeats. J Cell Biol 164: 25–33. Jolly C, Morimoto RI, Robert-Nicoud M, Vourc’h C. 1997. HSF1 transcription factor concentrates in nuclear foci during heat shock: Relationship with transcription sites. J Cell Sci 110: 2935–2941. Jolly C, Vourc’h C, Robert-Nicoud M, Morimoto RI. 1999b. Intron-independent association of splicing factors with active genes. J Cell Biol 145: 1133–1143. Jones KW, Prosser J, Corneo G, Ginelli E. 1973. The chromosomal location of human satellite DNA 3. Chromosoma 42: 445 –451. Kokalj-Vokac N, Almeida A, Viegas-Pe´quignot E, Jeanpierre M, Malfoy B, Dutrillaux B. 1993. Specific induction of uncoiling and recombination by azacytidine in classical satellite-containing constitutive heterochromatin. Cytogenet Cell Genet 63: 11– 15. Lamszus KL, Kluwe J, Matschke H, Meissner H, Lass R, Westphal M. 1999. Allelic losses at 1p, 9q, 10q, 14q and 22q in the progressionof aggressive meningiomas and undifferentiated meningeal sarcomas. Cancer Genet Cytogenet 110: 103–110. Lindquist S. 1986. The heat-shock response. Annu Rev Biochem 55: 1151–1191.

Mahl P, Lutz Y, Puvion E, Fuchs JP. 1989. Rapid effect of heat shock on two heterogeneous nuclear ribonucleoproteinassociated antigens in HeLa cells. J Cell Biol 109: 1921–1935. Maison C, Bailly D, Peters AH, Quivy JP, Roche D, Taddei A, Lachner M, Jenuwein T, Almouzni G. 2002. Higher-order structure in pericentric heterochromatin involves a distinct pattern of histone modification and an RNA component. Nat Genet 30: 329–334. Metz A, Soret J, Vourc’h C, Tazi J, Jolly C. 2004. A key role for stress-induced satellite III transcripts in the relocalization of splicing factors into nuclear stress granules. J Cell Sci 117: 4551– 4558. Misteli T. 2007. Beyond the sequence: Cellular organization of genome function. Cell 128: 787– 800. Morimoto RI, Santoro MG. 1998. Stress-inducible responses and heat shock proteins: New pharmacologic targets for cytoprotection. Nat Biotechnol 16: 833 –838. Muchardt C, Guilleme M, Seeler JS, Trouche D, Dejean A, Yaniv M. 2002. Coordinated methyl and RNA binding is required for heterochromatin localization of mammalian HP1a. EMBO Rep 3: 975 –981. Oesterreich S, Lee AV, Sullivan TM, Samuel SK, Davie JR, Fuqua SA. 1997. Novel nuclear matrix protein HET binds to and influences activity of the HSP27 promoter in human breast cancer cells. J Cell Biochem 67: 275– 286. Pirkkala L, Nykanen P, Sistonen L. 2001. Roles of the heat shock transcription factors in regulation of the heat shock response and beyond. FASEB J 7: 1118– 1131. Prasanth KV, Rajendra TK, Lal AK, Lakhotia SC. 2000. Omega speckles – a novel class of nuclear speckles containing hnRNPs associated with noncoding hsr-omega RNA in Drosophila. J Cell Sci 113: 3485– 3497. Puvion E, Viron A. 1981. In situ structural and functional relationships between chromatin pattern and RNP structures involved in non-nucleolar chromatin transcription. J Ultrastruct Res 74: 351– 360. Renz A, Fackelmayer FO. 1996. Purification and molecular cloning of the scaffold attachment factor B (SAF-B), a novel human nuclear protein that specifically binds to S/MAR-DNA. Nucleic Acids Res 24: 843– 849. Reshmi-Skarja S, Huebner A, Handschug K, Fineglod DN, Clark AJL, Gollin SM. 2003. Chromosomal fragility in patients with triple A syndrome. Am J Med Genet 117A: 30– 36. Rizzi N, Denegri M, Chiodi I, Corioni M, Valgardsdottir R, Cobianchi F, Riva S, Biamonti G. 2004. Transcriptional activation of a constitutive heterochromatic domain of the human genome in response to heat shock. Mol Biol Cell 15: 543–551. Robertson KD, Wolffe AP. 2000. DNA methylation in health and disease. Nat Rev Genet 1: 11–19. Sandqvist A, Bjork JK, Akerfelt M, Chitikova Z, Grichine A, Vourc’h C, Jolly C, Salminen TA, Nymalm Y, Sistonen L. 2009. Heterotrimerization of heat-shock factors 1 and 2 provides a transcriptional switch in response to distinct stimuli. Mol Biol Cell 20: 1340–1347. Sarge KD, Murphy SP, Morimoto RI. 1993. Activation of heat shock gene transcription by heat shock factor 1 involves oligomerization, acquisition of DNA-binding

G. Biamonti and C. Vourc’h activity, and nuclear localization and can occur in the absence of stress. Mol Cell Biol 13: 1392– 1407. Sasaki YT, Ideue T, Sano M, Mituyama T, Hirose T. 2009. MENepsilon/b noncoding RNAs are essential for structural integrity of nuclear paraspeckles. Proc Natl Acad Sci 106: 2525–2530. Sengupta S, Parihar R, Ganesh S. 2009. Satellite III noncoding RNAs show distinct and stress-specific patterns of induction. Biochem Biophys Res Commun 382: 102–107. Shumaker DK, Dechat T, Kohlmaier A, Adam SA, Bozovsky MR, Erdos MR, Eriksson M, Goldman AE, Khuon S, Collins FS, et al. 2006. Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci 103: 8703– 8708. Sunwoo H, Dinger ME, Wilusz JE, Amaral PP, Mattick JS, Spector DL. 2009. MEN epsilon/b nuclear-retained noncoding RNAs are up-regulated upon muscle differentiation and are essential components of paraspeckles. Genome Res 2009 19: 347– 359. Valgardsdottir R, Chiodi I, Giordano M, Cobianchi F, Riva S, Biamonti G. 2005. Structural and functional

characterization of noncoding repetitive RNAs transcribed in stressed human cells. Mol Biol Cell 16: 2597–2604. Valgardsdottir R, Chiodi I, Giordano M, Rossi A, Bazzini S, Ghigna C, Riva S, Biamonti G. 2008. Transcription of Satellite III non-coding RNAs is a general stress response in human cells. Nucleic Acids Res 36: 423 –434. Verdel A, Jia S, Gerber S, Sugiyama T, Gygi S, Grewal SI, Moazed D. 2004. RNAi-mediated targeting of heterochromatin by the RITS complex. Science 303: 672 – 676. Weighardt F, Cobianchi F, Cartegni L, Chiodi I, Villa A, Riva S, Biamonti G. 1999. A novel hnRNP protein (HAP/ SAF-B) enters a subset of hnRNP complexes and relocates in nuclear granules in response to heat shock. J Cell Sci 112: 1465– 1476. Yost HJ, Lindquist S. 1986. RNA splicing is interrupted by heat shock and is rescued by heat shock proteins synthesis. Cell 45: 185–193. Zhimulev IF, Belyaeva ES. 2003. Intercalary heterochromatin and genetic silencing. Bioessays 25: 1040–1051.

Orphan Nuclear Bodies Maria Carmo-Fonseca1, Maria T. Berciano2, and Miguel Lafarga2 1

Instituto de Medicina Molecular, Faculdade de Medicina, Universidade de Lisboa, 1649-028 Lisboa, Portugal

2

Department of Anatomy and Cell Biology, and “Centro de Investigacion Biomedica en Red sobre Enfermedades Neurodegenerativas (CIBERNED)”, University of Cantabria, Santander 39011, Spain

Correspondence: [email protected]

Orphan nuclear bodies are defined as nonchromatin nuclear compartments that have been less well studied compared with other well-characterized structures in the nucleus. Nuclear bodies have traditionally been thought of as uniform distinct entities depending on the protein “markers” they contain. However, it is becoming increasingly apparent that nuclear bodies enriched in different sets of transcriptional regulators share a link to the ubiquitin-proteasome and SUMO-conjugation pathways. An emerging concept is that some orphan nuclear bodies might act as sites of protein modification by SUMO and/or proteasomal degradation of ubiquitin-tagged proteins. By defining a specialized environment for protein modification and degradation, orphan nuclear bodies may increase the capacity of cells to survive under varying environmental conditions.

nuclear body is a region within the nucleus that is morphologically distinct from its surroundings when observed by transmission electron microscopy (reviewed in Spector 2006). This excludes structures that are only detected upon overexpression of tagged nuclear proteins and most likely result from nonphysiological aggregation of excess protein. On the basis of their structure observed at the electron microscopic level, nuclear bodies have been classified as either simple or complex (Bouteille et al. 1974). The simple nuclear bodies are small (0.2 – 0.5 mm), round, compact and finely fibrillar, whereas the complex nuclear bodies are larger (0.2 –1.2 mm), heterogeneous in shape and texture, and enveloped by a peripheral capsule, which gives them a doughnut-shaped appearance. Orphan nuclear

A

bodies are primarily proteinaceous structures enriched in either transcriptional regulators or RNA-binding proteins (Table 1). Although it was initially thought that these structures might act as specialized compartments for gene expression, most genes and RNA species targeted by the protein factors enriched in nuclear bodies show a widespread distribution throughout the nucleoplasm. Moreover, nuclear bodies contain only a fraction of the total pool of transcriptional regulators and RNA-binding proteins in the nucleus, and many of these factors are in constant flux in and out of the bodies. A breakthrough in this field was the recent discovery that transcription factors enriched in nuclear bodies are modified by SUMO and/or ubiquitin, and that SUMO-conjugation plays an important role in nuclear body assembly

Editors: Tom Misteli and David Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000703 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000703

M. Carmo-Fonseca et al. Table 1. Orphan nuclear bodies

Nuclear body Clastosome

Cleavage body

OPT domain

Polycomb body

Sam68 body

SUMO body

Description

Reference

Concentrates the proteasomal 20S and 19S complexes, and ubiquitin conjugates. Detected predominantly when the activity of the proteasome is stimulated, disassembles upon proteasomal inhibition. The number per cell ranges between 0 and 3, and the diameter is 0.2 –1.2 mm. Enriched in cleavage factors CstF 64 kDa and CPSF 100 kDa, and DEAD box protein DDX1. Detected predominantly during S phase, is not affected by transcription inhibition. The number per cell ranges between 1 and 4, and the diameter is 0.2 –1.0 mm. Enriched in transcription factors Oct1 and PTF. Partial colocalization with transcription sites. Detected predominantly during late G1 phase, disassembles upon transcription inhibition. The number per cell ranges between 1 and 3, and the diameter is 1.0 –1.5 mm. Enriched in PcG proteins in Drosophila embryos and human cells. Concentrates human RING1, BMI1, HPC, and some associate with pericentromeric heterochromatin. The number per cell ranges between 12 and 16, and the diameter is 0.3 –1.0 mm. Concentrates Sam68 and Sam68-like proteins SLM-1 and SLM-2. Disassembles upon transcription inhibition. Most likely enriched in RNA. The number per cell ranges between 2 and 5, and the diameter is 0.6 –1.0 mm. Enriched in SUMO-1 and SUMO- conjugating enzyme Ubc9. Concentrates transcription factors pCREB, CBP, c-Jun. The number per cell ranges between 1 and 3, and the diameter is 1 –3 mm.

(reviewed in Heun 2007). It is therefore possible that some nuclear bodies act as protein modification and degradation centers. THE CLASTOSOME: A NUCLEAR BODY LINKED TO THE UBIQUITIN-PROTEASOME SYSTEM

Proteins in cells and organisms exist in a dynamic state with individual half-lives ranging from minutes to years. This implies that protein degradation must be highly selective and tightly

Lafarga et al. 2002

Schul et al. 1996; Ble´oo et al. 2001; Li et al. 2006

Pombo et al. 1998

Buchenau et al. 1998; Saurin et al. 1998

Chen et al. 1999

Navascue´s et al. 2007

controlled (reviewed in Ravid and Hochstrasser 2008). Eukaryotic cells have two major mechanisms for protein degradation: lysosomes and proteasomes. Although exogenous particles and endogenous cellular organelles are targeted for digestion in lysosomes, the majority of intracellular proteins are degraded by proteasomes (reviewed in Murata et al. 2009). Proteasomes degrade short-lived regulatory proteins and thereby control cellular processes such as signal transduction, cell cycle, and gene transcription. Proteasomes additionally clear misfolded and

Orphan Nuclear Bodies

aggregated proteins from the cell and produce some of the peptides to be displayed at the cell surface as part of the adaptive immune response. A fundamental question about intracellular proteolysis is how specific proteins are recognized by the proteolytic machinery. In eukaryotic cells, most substrate proteins are targeted to the proteasome by the covalent attachment of many copies of the small protein ubiquitin. Once a protein has been tagged by polyubiquitination, it is recognized by the proteasome, unfolded and then degraded (Fig. 1). However, ubiquitination can function independently of the proteasome and proteasomes can degrade some proteins without their prior modification by ubiquitin (Ravid and Hochstrasser 2008). The proteasome is an approximately 2.5MDa (26S) protein machine composed of two

subcomplexes: a 20S barrel-shaped core that carries the catalytic activity, and a flanking regulatory 19S particle that recognizes and unfolds protein substrates (reviewed in Cheng 2009). The 20S core particle consists of four stacked rings that form a cavity at the center of the particle where the peptidolytic active sites are sequestered. Substrates gain access to this cavity through narrow pores present at either axial end of the core particle. These pores are normally closed and require a mechanism of gate opening carried out by the 19S regulatory particle. One regulatory particle may associate with either axial end of the core particle. The regulatory particle contains ATPases presumed to function in substrate unfolding, and ubiquitin chain receptors responsible for recognition of the substrate proteins (Fig. 1). Proteasomes were first shown to localize to the nucleus by Franke and colleagues (Hugle

Ub Ub Ub Ub

Regulatory particle Gated pore

Active sites

Core particle

Gated pore

Regulatory particle

Figure 1. The 26S proteasome is composed of one core particle (20S) and one or two regulatory particles (19S).

Proteins destined for degradation are initially attached to ubiquitin polymers. After this covalent modification, the substrate protein is able to bind (either directly or via adaptor proteins) to the 19S regulatory complex. Then, the protein is unfolded by ATPases that encircle the entrance of the 20S catalytic core, and the polyubiquitin chain is removed by proteasome-associated deubiquitylating enzymes. Finally, the unfolded protein is translocated into the central proteolytic chamber, where it is cleaved into short peptides.

M. Carmo-Fonseca et al.

et al. 1983; Kleinschmidt et al. 1983). Subsequent studies confirmed that proteasomes, as well as components of the ubiquitin-proteasome system occur both in the nucleus and in the cytoplasm. Although some nuclear proteins, such as p53, are first exported from the nucleus and then degraded in the cytoplasm, several lines of evidence indicate that proteins can be ubiquitylated and degraded by the proteasome within the nucleus (Tao et al. 2005; Rockel et al. 2005). Immunofluorescence studies using antibodies specific for the 20S and 19S proteasomal subcomplexes reveal that although proteasomes are most often diffusely distributed throughout the nucleoplasm, they occasionally concentrate in discrete structures (Fig. 2). By electron microscopy, these structures correspond to previously described complex, ring-shaped

19S & Ub

19S

A

B

20S & Nup

20S

C

D

nuclear bodies (Bouteille et al. 1974). Doublelabeling experiments further show the presence of ubiquitin-conjugates, c-Jun and c-Fos, two short-lived transcription factors degraded by the ubiquitin-proteasome system, in the proteasome-containing nuclear bodies (Lafarga et al. 2002). Proteasome-containing nuclear bodies form in response to stimuli that activate proteasome-dependent proteolysis and disappear when proteasome function is inhibited. Taken together, these results suggest that proteasomecontaining nuclear bodies are sites of protein degradation, and the name clastosome (from the Greek klastos, broken and soma, body) was coined to refer to this nuclear domain (Lafarga et al. 2002). Clastosomes are defined as nuclear bodies of irregular shape and size that contain both catalytic and regulatory subunits of the

20S

E

Figure 2. The clastosome is a nuclear body enriched in proteasomes. (A,B) Colocalization of

ubiquitin-conjugates (A, red staining) and 19S proteasomal complexes (A and B, green staining) in a human neuron mechanically isolated from dorsal root ganglia obtained from an autopsy of a patient without any diagnosed neurological disorder. Bar, 5 mm. (C, D) Several clastosomes are observed in the nucleus of a neurosecretory neuron isolated from rat hypothalamus after osmotic stress; double-immunofluorescence with antibodies specific to 20S proteasomal complexes (C and D, red staining) and a nucleoporin (C, green staining). (E) Immunogold labeling with antibodies directed against the 20S proteasome reveals a doughnutor ring-shaped nuclear body. Bar, 300 nm.

Orphan Nuclear Bodies

proteasome and ubiquitylated proteasome substrates. Clastosomes are normally absent or scarce in most cell types, indicating that these nuclear bodies are not essential for proteasome function. Most likely, protein substrates tagged for proteolysis are rapidly encountered, recognized and degraded by the diffusely distributed proteasomes in the nucleoplasm. Clastosomes become prominent when cells are exposed to stimuli that suddenly increase the levels of proteins targeted for degradation, for example, inducible expression of c-fos by serum, or stress. Clastosomes may therefore form as a result of an overload of protein substrates. Clastosomes assemble transiently and revert after the stimulus, consistent with the view that they accumulate substrates queuing up for proteolysis. Moreover, clastosomes are not detected in cells treated with specific proteasome inhibitors, arguing that clastosome assembly requires proteasomal activity. Clastosomes may also act by sequestering proteasomes, thereby controlling the availability of proteolytic machines for nuclear protein degradation. Recent studies suggest that regulated protein degradation by the ubiquitin proteasome system is an important modulator of neuronal function (reviewed in Tai et al. 2008). In particular, the localization of proteasomes to synapses is controlled by synaptic stimulation, thus providing a mechanism for local protein turnover (Bingol and Schuman 2006). The observation that clastosomes are widespread in glial cells and many types of brain neurons (A´dori et al. 2006; Baltrons et al. 2008) raises the possibility that these nuclear bodies participate in distinctive features of protein turnover in the nervous system. The importance of proteolysis in neurons is further underlined by the presence in a wide spectrum of neurodegenerative diseases of pathological protein aggregates that lead to a severe impairment of the ubiquitin-proteasome system (Bennett et al. 2005). Although the presence of ubiquitin and proteasomes in inclusion bodies formed by disease-associated protein aggregates suggests a link with dysfunction of the ubiquitin-proteasome system (Janer et al. 2006), the mechanism by which accumulation of protein aggregates in inclusion bodies

contributes to disease remains unknown (see Orr 2010). THE PROTEASOME IS A COMPONENT OF BOTH CLASTOSOMES AND PML NUCLEAR BODIES

The PML protein colocalizes with ubiquitin and the proteasome in nuclear bodies (Zhong et al. 2000; Lallemand-Breitenbach et al. 2001; Lafarga et al. 2002). Recent studies show that PML can be modified by SUMO and this triggers its polyubiquitylation and proteasomal degradation (Tatham et al. 2008; LallemandBreitenbach et al. 2008). According to the model proposed by De The´ and colleagues (Lallemand-Breitenbach et al. 2001), after PML protein modification, proteasomes are recruited to primary PML bodies giving rise to mature PML nuclear bodies, which have a characteristic shell- or ring-like morphology by electron microscopy and are therefore indistinguishable from clastosomes. An alternative view is that modified PML protein is recruited to clastosomes (Fig. 3). As clastosomes form independently of PML protein (Lafarga et al. 2002), we favor the later model. Indeed, clastosomes can be seen in the nucleus containing distinct proteasome substrates, suggesting diversity among this type of nuclear bodies (Fig. 3). Clearly, it is crucial to determine what nucleates de novo assembly of a clastosome: Is it the proteasome, the protein substrate or a complex of proteasomes with associated substrates? Moreover, it is becoming increasingly apparent that the cell’s population of proteasomes may actually represent a diverse group of functionally distinct members (Hanna et al. 2007; Hanna and Finley 2007). An exciting speculation is that clastosomes result from particular proteasome configurations that display novel and unique properties, distinct from the “canonical” forms present throughout the nucleoplasm. THE POLYCOMB BODY: A TRANSCRIPTIONAL REPRESSOR DOMAIN?

Polycomb group (PcG) proteins are well-conserved, essential regulatory factors that bind

M. Carmo-Fonseca et al.

20S & PML

20S

A

B

19S

19S & c-fos

C

D

E

SUMO

Primary PML body

Ub

Clastosome

Figure 3. Clastosomes concentrate protein substrates for proteasomal degradation. (A, B) The panels depict a

human neuron mechanically isolated from dorsal root ganglia obtained from an autopsy of a patient without any diagnosed neurological disorder, double-labeled with anti-PML (B, green staining) and anti-20S proteasomal complexes (A and B, red staining). Note that only one of the multiple PML bodies concentrates proteasomes (arrowhead) and that one body enriched in proteasomes does not contain PML (arrow). (C, D) Colocalization of 19S proteasomal complexes (C and D, red staining) and c-Fos (D, green staining) in the nucleus of a neurosecretory neuron isolated from rat hypothalamus after osmotic stress. Bar, 5 mm. (E) Hypothetical model for PML protein traffic through PML bodies and clastosomes. PML (blue spheres) distributes diffusely in the nucleoplasm and associates transiently with primary PML bodies. Upon modification by SUMO and ubiquitin, the modified PML proteins (red spheres) are recruited to clastosomes for degradation.

to specific DNA regions in target genes and repress their transcription. PcG proteins maintain the silenced state of developmental genes, including homeotic genes and genes involved in mammalian embryonic stem cell differentiation (Schuettengruber et al. 2007). PcG proteins were first described in Drosophila as factors responsible for maintaining the

transcriptionally repressed state of Hox/homeotic genes throughout development. A growing number of vertebrate genes related to the Drosophila PcG proteins have recently been identified. PcG proteins exist in at least two distinct complexes termed Polycomb repressive complex 1 (PRC1) and Polycomb repressive complex 2 (PRC2). Members of PRC1, including

Orphan Nuclear Bodies

the HPH1/HPH2, Bmi1, HPC2 and RING1 gene products are diffusely distributed in the nucleus and in addition colocalize in both Drosophila and mammalian cells as large nuclear domains termed Polycomb or PcG bodies (Buchenau et al. 1998; Saurin et al. 1998). In Drosophila, several genes that are repressed by PcG proteins localize to Polycomb bodies, suggesting that these domains correspond to sites of gene silencing (Grimaud et al. 2006). However, the number of Polycomb bodies is orders of magnitude smaller than the number of genetic loci targeted by PcG proteins. This implies that either multiple gene targets are associated with each Polycomb body or that gene silencing by PcG proteins can occur outside Polycomb bodies. Although endogenous homeotic genes appear to cluster at PcG bodies by a mechanism that requires components of the RNA interference machinery (Grimaud et al. 2006), it remains unknown whether assembly of a Polycomb body is necessary for the functional association of PcG proteins with chromatin. POLYCOMB AND PML NUCLEAR BODIES: THE SUMO CONNECTION

The small ubiquitin-like modifier (SUMO) is an ubiquitin-related protein that can be covalently conjugated to a variety of protein substrates, altering the properties of the modified proteins. SUMO conjugation is essential for cell and organism viability, from yeast to mammals (Nacerddine et al. 2005), and affects many biological processes, including cell cycle progression, maintenance of genome integrity, and transcription (reviewed in Hay 2005). SUMO substrates include protein components of both PML and Polycomb nuclear bodies and recent evidence implicates the SUMOylation pathway as an important player in the assembly of these domains (Heun 2007). The PML protein and the PML body components HIPK2, Daxx, and Sp100 are modified by SUMO. Moreover, SUMO conjugation is necessary for formation of PML bodies and for recruitment of HIPK2, Daxx, and Sp100 into the bodies (reviewed in Hay 2005). SUMO-

modified HIPK2 represses the activity of bound transcription factors by associating with Groucho corepressor and HDAC1, and SUMO modification of Sp100 enhances its interaction with heterochromatin protein 1 (HP1), a well known initiator of repressive domains in chromatin. The mechanism of SUMO conjugation involves a cascade of events catalysed by an activating enzyme E1, a conjugating enzyme E2 (known as Ubc9), and a protein ligase E3 (Hay 2005). There are at least three types of SUMO E3 ligases in cells: the protein inhibitor of activated signal transducer and activator of transcription (PIAS) family, the nuclear pore complex protein RanBP2/Nup358, and the Polycomb group protein Pc2. One of the substrates of Pc2 is the carboxy-terminal binding protein (CtBP) transcriptional corepressor (Kagey et al. 2003). It has been proposed that Pc2 functions to recruit Ubc9 and CtBP to PcG bodies, where SUMO modification of CtBP occurs (Kagey et al. 2004). More recently, the zinc finger protein CTCF was also shown to be recruited to Polycomb bodies and to be modified by SUMO (MacPherson et al. 2009). Intriguingly, only a small proportion of the total cellular pool of a given transcription factor is modified by SUMO. Yet, the entire population is functionally repressed. The following model was proposed to accommodate these observations: Upon conjugation to SUMO, a transcription factor is incorporated into a repression complex located in a nuclear body; even if SUMO is removed from the transcription factor at a later stage by specific proteases, the protein would still be retained in the repression complex (Hay 2005). Consistent with this view, PML and Polycomb nuclear bodies might result from the assembly of SUMO-dependent repression complexes. A variation of this hypothesis postulates that a SUMO-modified transcription factor recruits an enzyme capable of posttranslationally modifying chromatin. SUMO could then be removed from the transcription factor, whereas transcriptional repression would be maintained by the chromatin modification (Hay 2005). According to this alternative view, PML and Polycomb nuclear bodies could be sumoylation centers, from

M. Carmo-Fonseca et al.

which SUMO-modified transcription factors would be released. Consistent with the model that nuclear bodies can act as sites for SUMOconjugation, the SUMO-conjugating enzyme Ubc9 is highly enriched in these structures (Navascue´s et al. 2007) (Fig. 4). SEVERAL TRANSCRIPTIONAL REGULATORS LOCALIZE TO NUCLEAR BODIES

Many transcription factors associated with RNA polymerase II and III activity appear uniformly distributed throughout the nucleoplasm. In addition, some of these factors are found concentrated in larger nuclear domains, at least in some cell lines. A well characterized example is the so-called OPT domain, a region of approximately 1.3 mm in diameter that concentrates transcription factors Oct1 and PTF

A

(Pombo et al. 1998; Grande et al. 1997). The OPT domain is dynamic during the cell cycle: It appears during G1 phase and disappears in S phase. The OPT domain is also transcriptiondependent: the transcription inhibitor DRB inhibits its formation and hastens its disappearance (Pombo et al. 1998). Although it has been proposed that the OPT domain might play a role in clustering genes on different chromosomes to a region where the appropriate transcription factors are concentrated (Pombo et al. 1998), as yet there is no evidence to support this view. In particular, genes that contain PTF and Oct1-binding sites have not been observed to colocalize in the domain. Alternatively, the OPT domain may contain intermediary complexes of transcription factors that form independently of their binding to regulatory sequences in target genes. Consistent with this

C

nu

nu

B

D

Figure 4. SUMO-1 localizes to nuclear bodies in neuron-like UR61 cells. (A) Electron microscopy reveals the

presence of a round body (arrow) in the nucleus (nu, nucleolus). Bar, 1 mm. (B) Immunogold labeling of a nuclear body using antibodies specific for SUMO-1 (an arrow points to an adjacent doughnut-shaped structure that is most probably a clastosome). Bar, 300 nm. (C) The distribution of SUMO-1 is detected in green by immunofluore-scence; the nucleolus is blue (immunofluorescence with antifibrillarin antibody) and the cell periphery is red (Rhodamine-Phalloidin Staining). (D) Co-localization of GFP-SUMO-1 (green) and SUMO E2 conjugating enzyme DsRed-Ubc9 (red) in nuclear bodies; the cytoplasm is stained blue (immunofluo-rescence with anti-SMN antibody). Bar, 5 mm.

Orphan Nuclear Bodies

view, nuclear domains enriched in transcription factors could form as a result of protein modification, namely SUMOylation, as suggested for PML bodies and more recently, for a novel type of nuclear body that concentrates KRABzinc finger transcriptional regulators (Briers et al. 2009). RNA-BINDING PROTEINS IN NUCLEAR BODIES

Members of three major classes of RNA-binding proteins localize to orphan nuclear bodies. These include Sam68 and the related SLM-1 and SLM-2 proteins of the signal transduction and activation of RNA (STAR) family, two subunits of the cleavage stimulation factor (CstF) and the cleavage and polyadenylation specificity factor (CPSF) components of the mRNA 30 -end processing machinery, and DDX1, a member of the DEAD box protein family of RNA helicases. Sam68 belongs to the STAR family of proteins characterized by a KH (hnRNP K homology) domain embedded in a highly conserved region called GSG (GRP33/Sam68/GLD1) domain, which is required for homodimerization and sequence-specific RNA binding. Sam68 is a substrate for Src tyrosine kinases and is therefore considered a strong candidate to integrate signal transduction pathways and RNA metabolism, particularly alternative splicing (Paronetto et al. 2007). Although Sam68 can translocate to the cytoplasm and associate with specific RNAs in polysomes (Grange et al. 2009; Paronetto et al. 2009), the protein is predominantly detected in the nucleus (Fig. 5). Sam68 localizes diffusely in the nucleoplasm but additionally concentrates in a few prominent structures termed Sam68 nuclear bodies or SNBs (Chen et al. 1999). SNBs are dynamic structures that disassemble during mitosis and after treatment of cells with transcriptional inhibitors. Occasionally they appear adjacent to the nucleolus (Fig. 5) or to a Cajal body. Electron microscopic studies show that SNBs contain nucleic acids, presumably RNA, and targeting of Sam68 to these structures involves the highly conserved GSG protein domain (Chen et al. 1999). Thus, SNBs might form as a result of

A

B

C

Figure 5. The RNA-binding protein Sam68 localizes to nuclear bodies in both Hela cells (A) and rat neurons (B, C). Sam 68 is detected in green by immunofluorescence labeling. Nucleoli and cytoplasm are detected in red ( propidium iodide staining). RNA splicing speckles are detected in blue (immunofluorescence with an antibody directed against the 2, 2, 7trimethylguanosine cap structure of snRNAs). Bar, 10 mm.

Sam68 self-assembly and binding to specific RNAs in the nucleus. Although the identity of such potential scaffold RNAs remains to be determined, they are expected to be short-lived because SNBs disassemble within a few hours after transcription inhibition (Chen et al. 1999). Recently, Sam68 was reported to be modified by SUMO (Babic et al. 2006), and a pending question is whether SUMOylation is related to formation of SNBs. CPSF and CstF are multiprotein complexes involved in formation of the 30 -ends of most mRNAs (reviewed in Danckwardt et al. 2008). The CPSF 100 kDa subunit and the CstF 64 kDa subunits are diffusely distributed throughout the nucleoplasm and additionally concentrate in a few bright foci termed cleavage bodies (Schul et al. 1996). Cleavage bodies are primarily observed during S phase, apparently do not contain RNA, and are not affected

M. Carmo-Fonseca et al.

by inhibitors of RNA transcription (Li et al. 2006). Cleavage bodies often concentrate the DEAD box protein DDX1 (Ble´oo et al. 2001) and are frequently found adjacent to Cajal bodies (Li 2006). CONCLUDING REMARKS

Unlike the nucleolus, most types of orphan nuclear bodies are not detected in all cell types. It is therefore unlikely that these structures play an essential role in the nucleus. In general, orphan nuclear bodies are dynamic structures that assemble and disassemble at specific stages of the cell cycle or in response to changes in environmental conditions. A fundamental question is whether bodies form under these specific conditions as a result of aggregation of excessive protein that is not used, or contribute to increase the capacity of cells to survive under varying environmental stresses. Noteworthy, SUMOylation of the Caenorhabditis elegans PcG protein SOP-2 is required for both its localization to nuclear bodies and its physiological repression of Hox genes (Zhang et al. 2004). In further agreement with the view that nuclear bodies are functionally relevant, several lines of evidence implicate SUMOylation in assembly of PML nuclear bodies and maintenance of cell viability (Heun 2007). Taken together, these results suggest that some nuclear bodies might represent sites of SUMO-conjugation and/or protein degradation by the ubiquitin-proteasome system, whereas others are more likely associated with transcription regulation and RNA metabolism. Further studies are needed to understand precisely what roles these different types of bodies play in the nucleus. ACKNOWLEDGMENTS

We thank our colleagues Joa˜o Ferreira and Jose´ Rino for stimulating discussions and help in some figures. Our laboratories are supported by grants from Fundac¸a˜o para a Cieˆncia e Tecnologia, Portugal (PTDC/BIA-BCM/101575/ 2008), the European Commission (LSHG-CT2005-518238 and MRTN-CT-2006-035733),

“Direccio´n General de Investigacion” (BFU200800175, Spain) and CIBERNED (CB06/05/ 0037, Spain). REFERENCES A´dori C, Lo¨w P, Moszkovkin G, Bagdy G, La´szlo´ L, Kova´cs GG. 2006. Subcellular distribution of components of the ubiquitin-proteasome system in non-diseased human and rat brain. J Histochem Cytochem 54: 263– 267. Babic I, Cherry E, Fujita DJ. 2006. SUMO modification of Sam68 enhances its ability to repress cyclin D1 expression and inhibits its ability to induce apoptosis. Oncogene 25: 4955–4964. Baltrons MA, Pifarre´ P, Berciano MT, Lafarga M, Garcı´a A. 2008. LPS-induced down-regulation of NO-sensitive guanylyl cyclase in astrocytes occurs by proteasomal degradation in clastosomes. Mol Cell Neurosci 37: 494– 506. Bennett EJ, Bence NF, Jayakumar R, Kopito RR. 2005. Global impairment of the ubiquitin-proteasome system by nuclear or cytoplasmic protein aggregates precedes inclusion body formation. Mol Cell 17: 351– 365. Bingol B, Schuman EM. 2006. Activity-dependent dynamics and sequestration of proteasomes in dentritic spines. Nature 441: 1144– 1148. Ble´oo S, Sun X, Hendzel M, Rowe JM, Packer M, Godbout R. 2001. Association of human DEAD box protein DDX1 with a cleavage stimulation factor involved in 30 -end processing of pre-mRNA. Mol Biol Cell 12: 3046– 3059. Bouteille M, Laval M, Dupuy-Coin AM. 1974. Localization of nuclear functions as revealed by ultrastructural autoradiography and cytochemistry. In The cell nucleus, (ed. H. Busch), Vol. I, pp. 3 –71. Academic Press, New York. Briers S, Crawford C, Bickmore WA, Sutherland HG. 2009. KRAB zinc-finger proteins localize to novel KAP1containing foci that are adjacent to PML nuclear bodies. J Cell Sci 122: 937 –946. Buchenau P, Hodgson J, Strutt H, Arndt-Jovin DJ. 1998. The distribution of polycomb-group proteins during cell division and development in Drosophila embryos: impact on models for silencing. J Cell Biol 141: 469–481. Chen T, Boisvert F-M, Bazett-Jones DP, Richard S. 1999. A role for the GSG domain in localizing Sam68 to novel nuclear structures in cancer cell lines. Mol Biol Cell 10: 3015–3033. Cheng Y. 2009. Toward an atomic model of the 26S proteasome. Curr Opin Struct Biol 19: 203– 208. Danckwardt S, Hentze MW, Kulozik AE. 2008. 30 end mRNA processing: Molecular mechanisms and implications for health and disease. EMBO J 27: 482 –498. Grande MA, van der Kran I, de Jong L, van Driel R. 1997. Nuclear distribution of transcription factors in relation to sites of transcription and RNA polymerase II. J Cell Sci 110: 1781– 1791. Grimaud C, Bantignies F, Pal-Bhadra M, Ghana P, Bhadra U, Cavalli G. 2006. RNAi components are required for nuclear clustering of Polycomb group response elements. Cell 124: 957 –971. Grange J, Belly A, Dupas S, Trembleau A, Sadoul R., Goldberg Y. 2009. Specific interaction between San68 and

Orphan Nuclear Bodies neuronal mRNAs: Implication for the activity-dependent biosynthesis of elongation factor eEF1A. J Neurosci Res 87: 12–25. Hanna J, Finley D. 2007. A proteassome for all occasions. FEBS Lett 581: 2854– 2861. Hanna J, Meides A, Zhang DP, Finley D. 2007. A ubiquitin stress response induces altered proteasome composition. Cell 129: 747– 759. Hay RT. 2005. SUMO: a history of modification. Mol Cell 18: 1– 12. Heun P. 2007. SUMOrganization of the nucleus. Curr Opin Cell Biol 19: 350 –355. Hugle B, Kleinschmidt JA, Franke WW. 1983. The 22S cylinder particles of Xenopus laevis. II. Immunological characterization and localization of their proteins in tissues and cultured cells. Eur J Cell Biol 32: 157 –163. Janer A, Martin E, Muriel M-P, Latouche M, Fujigasaki H, Ruberg M, Brice A, Trottier Y, Sittler A. 2006. PML clastosomes prevent nuclear accumulation of mutant ataxin-7 and other polyglutamine proteins. J Cell Biol 174: 65– 76. Kagey MH, Melhuish TA, Wotton D. 2003. The polycomb protein Pc2 is a SUMO E3. Cell 113: 127 –137. Kagey MH, Melhuish TA, Powers SE, Wotton D. 2004. Multiple activities contribute to Pc2 E3 function. EMBO J 24: 108–119. Kleinschmidt JA, Hugle B, Grund C, Franke WW. 1983. The 22S cylinder particles of Xenopus laevis. I. Biochemical and electron microscopic characterization. Eur J Cell Biol 32: 143– 156. Lafarga L, Berciano MT, Pena E, Mayo I, Castan˜o JG, Bohmann D, Rodrigues JP, Tavanez JP, Carmo-Fonseca M. 2002. Clastosome: a subtype of nuclear body enriched in 19S and 20S proteasomes, ubiquitin, and protein substrates of proteasome. Mol Biol Cell 13: 2771– 2782. Lallemand-Breitenbach V, Zhu J, Puvion F, Koken M, Honore´ N, Doubeikovsky A, Duprez E, Pandolfi PP, Puvion E, Freemenot P, et al. 2001. Role of promyelocytic leukemia (PML) sumolation in nuclear body formation, 11S proteasome recruitment, and As2O3-induced PML or PML/retinoic acid receptor a degradation. J Exp Med 193: 1361–1371. Lallemand-Breitenbach V, Jeanne M, Benhenda S, Nasr R, Lei M, Peres L, Zhou J, Zhu J, Raught B, de The´ H. 2008. Arsenic degrades PML or PML-RARalpha through a SUMO-triggered RNF4/ubiquitin-mediated pathway. Nat Cell Biol 10: 547–555. Li L, Roy K, Katyal S, Sun X, Ble´oo S, Godbout R. 2006. Dynamic nature of cleavage bodies and their spatial relationship to DDX1 bodies, Cajal bodies and Gems. Mol Biol Cell 17: 1126– 1140. MacPherson MJ, Beatty LG, Zhou W, Du M, Sadowski PD. 2009. The CTCF insulator protein is posttranslationally modified by SUMO. Mol Cell Biol 29: 714 –725. Murata S, Yashiroda H, Tanaka K. 2009. Molecular mechanisms of proteasome assembly. Nat Rev Mol Cell Biol 10: 104 –115. Nacerddine K, Lehembre F, Bhaumik M, Artus J, CohenTannoudji M, Babinet C, Pandolfi PP, Dejean A. 2005. The SUMO pathway is essential for nuclear integrity and chromosome segregation in mice. Dev Cell 9: 769–779.

Navascue´s J, Bengoechea R, Tapia O, Vaque´ JP, Lafarga M, Berciano MT. 2007. Characterization of a new SUMO-1 nuclear body (SNB) enriched in pCREB, CBP, c-Jun in neuron-like UR61 cells. Chromosoma 116: 441– 451. Orr HT. 2010. Nuclear ataxias. Cold Spring Harb Perspect Biol 2: a000786. Paronetto MP, Achsel T, Massiello A, Chalfant CE, Sette C. 2007. The RNA-binding protein Sam68 modulates the alternative splicing of Bcl-x. J Cell Biol 176: 929– 939. Paronetto MP, Messina V, Bianchi E, Barchi M, Vogel G, Moretti C, Palombi F, Stefanini M, Geremia R, Richard S, et al. 2009. Sam68 regulates translation of target mRNAs in male germ cells, necessary for mouse spermatogenesis. J Cell Biol 185: 235 –249. Pombo A, Cuello P, Schul W, Yoon J-B, Roeder RG, Cook PR, Murphy S. 1998. Regional and temporal specialization in the nucleus: a transcriptionally-active nuclear domain rich in PTF, Oct1 and PIKA antigens associates with specific chromosomes early in the cell cycle. EMBO J 17: 1768–1778. Ravid T, Hochstrasser M. 2008. Degradation signal diversity in the ubiquitin-proteasome system. Nat Rev Mol Cell Biol 9: 679–690. Rockel TD, Stuhlmann D, von Mikecz A. 2005 Proteasomes degrade proteins in focal subdomains of the human cell nucleus. J Cell Sci 118: 5231– 5242. Saurin AJ, Shiels C, Williamson J, Satijn DP, Otte AP, Sheer D, Freemont PS. 1998. The human polycomb group complex associates with pericentromeric heterochromatin to form a novel nuclear domain. J Cell Biol 142: 887–898. Schuettengruber B, Chourrout D, Vervoort M, Leblanc B, Cavalli G. 2007. Genome regulation by Polycomb and Trithorax proteins. Cell 128: 735– 745. Schul W, Groenhout B, Koberna K, Takagaki Y, Jenny A, Manders EMM, Raska I, van Driel R, de Jong L. 1996. The RNA 30 cleavage factors CstF 64 kDa and CPSF 100 kDa are concentrated in nuclear domains closely associated with coiled bodies and newly synthesized RNA. EMBO J 15: 2883–2892. Spector DL. 2006. SnapShot: cellular bodies. Cell 127: 1070–1070.e1. Tai HC, Schuman EM. 2008. Ubiquitin, the proteasome and protein degradation in neuronal function and dysfunction. Nat Rev Neurosci 9: 826–838. Tao LZ, Cheung AY, Nibau C, Wu HM. 2005. RAC GTPases in tobacco and Arabidopsis mediate auxin-induced formation of proteolytically active nuclear protein bodies that contain AUX/IAA proteins. Plant Cell 17: 2369– 2383. Tatham MH, Geoffroy MC, Shen L, Plechanovova A, Hattersley N, Jaffray EG, Palvimo JJ, Hay RT. 2008. RNF4 is a poly-SUMO-specific E3 ubiquitin ligase required for arsenic-induced PML degradation. Nat Cell Biol 10: 538 –546. Zhang H, Smolen GA, Palmer R, Christoforu A, van den Heuvel S, Haber DA. 2004. SUMO modification is required for in vivo Hox gene regulation by the Caenorhabditis elegans Polycomb group protein SOP-2. Nat Genet 36: 507– 511. Zhong S, Salomoni P, Pandolfi PP. 2000. The transcriptional role of PML and the nuclear body. Nat Cell Biol 2: E85–E90.

Biogenesis of Nuclear Bodies Miroslav Dundr1 and Tom Misteli2 1

Department of Cell Biology, Rosalind Franklin University of Medicine and Science, North Chicago, Ilinois 60064

2

National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892

Correspondence: [email protected]; [email protected]

The nucleus is unique amongst cellular organelles in that it contains a myriad of discrete suborganelles. These nuclear bodies are morphologically and molecularly distinct entities, and they host specific nuclear processes. Although the mode of biogenesis appears to differ widely between individual nuclear bodies, several common design principles are emerging, particularly, the ability of nuclear bodies to form de novo, a role of RNA as a structural element and self-organization as a mode of formation. The controlled biogenesis of nuclear bodies is essential for faithful maintenance of nuclear architecture during the cell cycle and is an important part of cellular responses to intra- and extracellular events.

he mammalian cell nucleus contains a multitude of discrete suborganelles, referred to as nuclear bodies or nuclear compartments (reviewed in Dundr and Misteli 2001; Spector 2001; Lamond and Spector 2003; Handwerger and Gall 2006; Zhao et al. 2009). These bodies are an essential part of the nuclear landscape as they compartmentalize the nuclear space and create distinct environments within the nucleus (reviewed in Misteli 2007). Many nuclear bodies carry out specific nuclear functions, such as the synthesis and processing of pre-ribosomal RNA in the nucleolus, the storage and assembly of spliceosomal components in nuclear speckles, or the retention of RNA molecules in paraspeckles. The mechanisms by which nuclear bodies contribute to function are highly diverse. In some cases, a nuclear body may be host to a particular activity such as transcription; in other cases, a nuclear body

T

seems to act indirectly by regulating the local concentration of its components in the nucleoplasm. In many ways, nuclear bodies are similar to conventional cellular organelles in the cytoplasm. Like cytoplasmic organelles, they contain a specific set of resident proteins, which defines each structure molecularly. Although many nuclear bodies are spherical in shape, most can be characterized based on their unique morphology, particularly when analyzed by electron microscopy and by their nuclear distribution patterns. However, in stark contrast to conventional cytoplasmic organelles, nuclear bodies are not delineated by lipid membranes, and their structural integrity appears to be entirely mediated by protein – protein and possibly protein – RNA interactions. The absence of a demarcating lipid membrane points to unique mechanisms of biogenesis.

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000711 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000711

M. Dundr and T. Misteli

The mechanisms of nuclear body biogenesis are physiologically important for two reasons. First, proper biogenesis is essential for their faithful inheritance during cell division. Many nuclear bodies disassemble as cells enter mitosis and must then efficiently and rapidly reassemble at the end of mitosis to ensure full functionality of the daughter nuclei. Second, nuclear bodies may form in response to specific cellular activities. Coordinated disassembly and efficient biogenesis of nuclear bodies is thus critical for maintenance of cellular integrity, homeostasis, and responsiveness. NUCLEAR BODIES ARE DYNAMIC STEADY-STATE STRUCTURES

Nuclear bodies are a morphologically and molecularly highly diverse group of cellular structures (Table 1; Fig. 1A– C). At one end of the spectrum is the nucleolus, which is often found in less than 3 copies, and often as a single organelle, and may reach a size up to several micrometers. At the other end of the spectrum, Cajal bodies are rarely larger than 1.5 mm, and as many as 10 can be found in a cell. Size, shape, and number of a given nuclear body are often dependent on cell type, are variable between tissues, and may change in response to cellular conditions. Despite their diverse morphological appearances, nuclear bodies share several common features of organization (reviewed in Misteli 2001, 2007; Lamond and Spector 2003; Handwerger and Gall 2006; Bernardi and Pandolfi 2007; Pederson and Tsai 2009). A common property of nuclear bodies is their overall structural stability. When followed by time-lapse microscopy, individual nuclear bodies persist during the entire interphase between entry into G1 phase and exit from G2 phase. During this time, the bodies undergo dynamic motions within the nucleus, whose extent is generally inversely proportional to their size (Platani et al. 2000; Muratani et al. 2002; Dundr et al. 2007). The large bodies such as nucleoli and nuclear speckles, typically 2 – 3 mm in diameter, are largely immobile and only undergo limited local motion (Misteli et al. 1997a; Kruhlak et al. 2000; Olson and

Dundr 2005). The smaller bodies such as Cajal bodies or PML nuclear bodies, with an average size of 500 nm –1 mm, undergo more extensive intranuclear motion and frequent fusion and fission events (Platani et al. 2002; Dellaire et al. 2006; Dundr et al. 2007; Chen et al. 2008). Despite the overall structural stability, nuclear bodies are, however, intrinsically highly dynamic structures (Phair and Misteli 2000; Handwerger and Gall 2006). The vast majority of nuclear body components are proteins that are enriched in a nuclear body but they are also present, albeit at lower concentration, in the surrounding nucleoplasm (reviewed in Dundr and Misteli 2001). Key insights into the nature of nuclear bodies comes from photobleaching experiments that have demonstrated rapid dynamic exchange of a large number of the major nuclear body components with the surrounding nucleoplasm (Kruhlak et al. 2000; Phair and Misteli 2000; Snaar et al. 2000; Chen and Huang 2001; WeidtkampPeters et al. 2008). For example, the major nucleolar component fibrillarin or the marker Cajal body component coilin have residence times on the order of tens of seconds and several minutes, respectively, and it can be calculated that the entire pool of a given nuclear body protein is completely replaced within a few minutes (Phair and Misteli 2000; Dundr et al. 2004). The absence of a delineating lipid boundary to distinguish nuclear bodies from their environment indicates that nuclear bodies are accumulations of proteins and RNAs and that the morphological appearance of a given nuclear body is largely determined by the sum of interactions of its components within the body. At present, no strictly architectural protein components of nuclear bodies are known. However, a role for dedicated protein – protein interactions in the structural maintenance of nuclear bodies is hinted at by the fact that several prominent nuclear body proteins have selfassociation properties (Hebert and Matera 2000). The major Cajal body protein coilin contains a self-interacting N-terminal domain, the PML protein forms homo-oligomers and the SMN protein, which is a major constituent of the gem nuclear bodies, oligomerizes readily

Table 1. Biogenesis and inheritance of nuclear bodies. Putative functions

Nucleolus

Ribosome biogenesis

Nuclear speckles Nuclear stress bodies Histone locus body

Storage and recycling of splicing factors Regulation of transcription and splicing under stress Histone gene synthesis

Cajal body

Typical Typical Defining components Size (mm) Number

RNA pol I machinery rRNA processing/ ribosome assembly factors Pre-mRNA splicing factors HSF1, HAP NPAT, FLASH,U7 snRNP

Biogenesis, maturation and Coilin, SMN recycling of small RNAs PML nuclear Regulation of genome stability, PML protein body DNA repair, control of transcription, viral defense Paraspeckles mRNA regulation, RNA editing NEAT1/MEN1/ bncRNAs PSP1, p54nrb/NONO Perinucleolar Post-transcriptional regulation PTB compartment of a subset of pol III RNAs

Activity dependence

Nucleation template

De novo formation

Mode of Inheritance

3–8

1 –4

þ

DNA/RNA

þ

disassembly, persist as mitotic nucleoli derived foci

2–3

20–50

þ

DNA/RNA

þ

1 –2

2 –6

þ

RNA

Unknown

Disassemble/ reassembly Unknown

0.2 –1.2

2 –4

þ

DNA/RNA

Unknown

0.2 –1.5

1 –10

2

DNA/RNA

þ

0.1 –1

10 –30

2

DNA/RNA

þ

0.2 –1

2 –20

þ

RNA

Unknown

Mitotic paraspeckles

0.2 –1

1 –2

þ

RNA

Unknown

Disassembly/ reassembly

Disassemble in metaphase, reform in G1/ S-phase Disassembly into mitotic CBs Mitotic PML accumulations

M. Dundr and T. Misteli

A

B

C

protein/RNA structures, which form by dynamic interactions of their components in the nucleoplasm (Fig. 1D). NUCLEAR BODY MAINTENANCE

D

kon = koff

kon >> koff

kon << koff

Figure 1. Structure and maintenance of nuclear

bodies. The appearance and number of nuclear bodies is highly variable. Prominent examples include (A) the nucleolus (RNA polymerase I subunit RPA43 in green, rRNA processing enzyme fibrillarin in red); (B) nuclear speckles (splicing factor SC35 in red; DNA in blue); and (C) Cajal bodies (coilin in red) and histone locus bodies (histone genes transcription factor NPAT in green), which are closely associated with each other (arrow). Inset: higher magnification view. Scale bars: 2 mm. (D) Nuclear bodies are dynamic, steady-state structures exposed to a continuous flow of material in and out. The ratio of on/off rate determines the size, and possibly the shape, of the body. Minor changes in on/off-rate ratio can strongly influence nuclear body morphology.

with itself (Hebert and Matera 2000; Kentsis et al. 2002; Dundr et al. 2004; Morse et al. 2007). No covalent interactions between nuclear body components have, however, been identified to date as determinants of nuclear body structure, and it seems likely that most intrabody interactions are noncovalent and weak. Taken together, the overall structural stability and the rapid exchange of individual components make nuclear bodies dynamic steady-state

The steady-state appearance of nuclear bodies is maintained by the continuous flow of molecules into and out of the body (Fig. 1D). The size, and possibly shape, of nuclear bodies is likely determined by the balance of on-rate relative to the off-rate of its components. An increase in on-rate or a drop in the off-rate will lead to an increase in size, whereas a decrease in on-rate or an increase in off-rate will lead to shrinkage (Fig. 1D). The molecular mechanisms that determine on/off rates of individual nuclear body components are poorly understood but likely involve post-translational modifications. This is best exemplified by SR protein splicing factors, which are major constituents of the nuclear speckles, which serve to store and assemble pre-mRNA splicing components. Several speckle components undergo a cycle of phosphorylation that coincides with their association with speckles (reviewed in Misteli 1997b). Many SR protein splicing factors exist in a hypo-phosphorylated state inside of speckles, where they are the target for speckle-resident SR-protein kinases of the Clk family. Hyper-phosphorylation of SR proteins then makes them competent to participate in the splicing reaction and at the same time leads to their dissociation from speckles. During the splicing reaction, the SR proteins are dephosphorylated, re-establishing their affinity for speckles, where they are recycled for a new round of splicing (reviewed in Misteli 1997b). A similar cyclical mechanism to control nuclear body association has been characterized for the nucleolar protein nucleostemin, which contains a GTP-binding domain that mediates nucleolar retention (Tsai and McKay 2005; Pederson and Tsai 2009). It is likely that similar, yet uncharacterized, mechanisms play key roles in controlling the association and dissociation, and thus shape and size, of many other nuclear bodies. Control of nuclear body number is poorly understood. Even the number of nucleoli, which

Nuclear Body Biogenesis

form around a fixed number of nucleolar organizing regions consisting of constant numbers of rDNA repeats on a defined number of chromosomes, varies greatly in different tissues and cell types. Some initial insight into number control comes from analysis of the Cajal body, where the marker protein coilin is capable of regulating the number of Cajal bodies per cell. This involves C-terminal phosphoserine residues that regulate its self-interaction activity (Shpargel et al. 2003; Hearst et al. 2009). Truncation or mutation of these phosphorylation sites leads to alteration in the number of Cajal bodies. In addition, expression of heterologous coilin proteins in different cellular backgrounds results in deregulation of Cajal body numbers. For example, mouse coilin forms numerous foci in human HeLa cells, but when expressed in mouse cells, the number of Cajal bodies does not increase. When frog coilin is expressed in either cell type, the number of Cajal bodies is highly up-regulated. In contrast, human coilin is unable to form foci in cells derived from coilin knockout mice (Shpargel et al. 2003). Size and number of nuclear bodies are not fixed but are dependent on physiological conditions. The number of nucleoli correlates positively with cell proliferation. For example, in lymphocytes with high proteosynthetic activity, and thus a high requirement for ribosomal RNA (rRNA), nucleolar size increases (HernandezVerdun 2006). Cellular stress, such as inflammation and oncogenic stimulation, cause an increase in the number and size of PML nuclear bodies along with increased expression of PML protein (reviewed in Bernardi and Pandolfi 2007). Coilin in primary cells is hyperphosphorylated on C-terminal phosphoserine residues that reduce its self-interaction activity, and these cells lack Cajal bodies. In contrast, in transformed cells, which have a large number of Cajal bodies (CBs), coilin is hypo-phosphorylated and able to self-associate (Hearst et al. 2009). It seems that the phosphorylation status of coilin has an impact on Cajal body formation, and coilin is likely a target of a yet uncharacterized signal transduction pathway that might control Cajal body formation (Hearst et al. 2009).

ACTIVITY-DEPENDENT AND ACTIVITY-INDEPENDENT NUCLEAR BODIES

Nuclear bodies may be divided into two classes based on their mode of formation: activitydependent nuclear bodies and activity-independent nuclear bodies. This classification is at this point somewhat tenuous, since unknown nuclear activities may be responsible for formation of a nuclear body assumed to form in an activity-independent fashion (Fig. 2). The former class includes bodies that form at sites of a particular nuclear function, such as transcription, and whose morphological appearance is strictly dependent on this activity (Table I). The prototypical example for activity-dependent bodies is the nucleolus, which forms in response to transcriptional activity of clustered ribosomal DNA genes located on multiple chromosomes that cluster in the nucleolus (Shaw and Jordan 1995; Boisvert et al. 2007). The strict dependence of the morphological appearance of the nucleolus on ongoing transcription is evident by its rapid structural reorganization upon inhibition of rDNA transcription (Olson and Dundr 2005). Conversely, introduction of exogenous rDNA genes on plasmids into the nucleus is sufficient for the formation of new nucleoli (Oakes et al. 1998). Other activity-dependent nuclear bodies include the histone locus body, which forms around histone gene

A Activity dependent

On site formation

B Activity independent

Recruitment

Figure 2. Classification of nuclear bodies based on activity. Nuclear bodies may either form (A) associated with a nuclear function, such as transcription (yellow), making their appearance dependent on the ongoing activity or (B) they may form independently of a function and subsequently associate with a particular nuclear site, such as an active gene locus (yellow).

M. Dundr and T. Misteli

clusters in response to transcriptional activation of replication-dependent histone genes during S-phase (Bongiorno-Borbone et al. 2008), nuclear stress bodies that form at sites of satellite III repeat transcription in response to heat shock (Jolly et al. 2002), and nuclear speckles that undergo dramatic morphological changes by rounding up upon inhibition of transcription and they rapidly re-form at sites of heat shockactivated genes (reviewed in Misteli 2007; Hu et al. 2009). On the other hand, there are nuclear bodies that seem to form without the requirement for a particular nuclear activity. Such apparently activity-independent bodies form in the nucleoplasm and may subsequently associate with particular nuclear locations (Table 1). Activity-independent bodies include PML nuclear bodies and Cajal bodies. Just like activity-dependent bodies, these structures are often associated with particular locations in the nucleus and sometimes with specific gene loci; however, these bodies probably do not form at these sites but rather are either recruited to them or activated genes are repositioned to associate with the preformed bodies (Carmo-Fonseca 2007). The clearest example of such recruitment comes from analysis of the interactions of the U2 snRNA gene cluster with the Cajal body (Dundr et al. 2007). Upon activation of the U2 snRNA genes, they undergo directed, actin-dependent motion to associate with a preformed Cajal body in the vicinity (Dundr et al. 2007). NUCLEAR BODY TEMPLATES AND NUCLEATORS

Formation of a nuclear body requires an initial nucleation event. Nucleation serves to immobilize some of the freely diffusible key components of the body and to provide a template to recruit additional building blocks. In the case of activity-associated nuclear bodies, this function is carried out by the activities that drive the nuclear body formation. Examples for well-defined nucleation events are the nucleolus, where accumulation of nucleolar proteins on ribosomal DNA and pre-ribosomal RNA leads to the formation of the nucleolus or the

recruitment of the histone pre-mRNA 30 -end processing factors in histone locus bodies. Immobilization of proteins in this way likely leads to recruitment of some of their interaction partners and to increased concentration of these proteins at the site of activity, ultimately resulting in the formation of a morphologically distinct suborganelle. In the case of activity-independent bodies, other nucleators must exist. These architectural elements may either be structural scaffold proteins or RNAs. Such nucleators are likely fairly abundant components of a given nuclear body, and they should be necessary and sufficient for the formation of a nuclear body. No such nucleators of activity-independent nuclear bodies have been identified to date. A possibility is that putative nucleators of activity-independent bodies are part of an immobile nuclear matrix. However, this scenario is unlikely since most nuclear bodies are mobile and thus not permanently tethered to an underlying structure (Platani et al. 2002; Muratani et al. 2002; Go¨risch et al. 2004). The identification of nuclear body nucleators will be critically important for a full understanding of nuclear body biogenesis and function. DE-NOVO FORMATION OF NBs

A critical test for whether a cellular structure is autonomous and self-sustaining is whether it can form de novo. Several nuclear bodies form de novo either under physiological or experimental conditions. The nucleolus can be formed de novo by introducing extra rDNA minigenes on plasmids into a cell (Nierras et al. 1997; Oakes et al. 1998). Apparently functional mini-nucleoli with a tendency to coalesce into a single, larger nucleolus rapidly form around the ribosomal genes (Oakes et al. 1998). In addition, a large number of small nucleoli are formed in Xenopus oocytes as thousands of extrachromosomal rDNA gene copies are amplified during oogenesis (Mais et al. 2002). Similarly, nuclear speckles can form de novo upon re-activation of global transcription after inhibition. After reversal of a global transcription block, speckles expand

Nuclear Body Biogenesis

from a condensed state and take on their typical irregular shape and new speckles form, associated with re-activated genes (Misteli et al. 1997; Shopland et al. 2003). De novo formation is not limited to activity-dependent nuclear bodies. During viral infection, PML nuclear bodies form rapidly. Upon entry of a viral genome into the nucleus, several major components of the PML nuclear body, including PML protein, Sp100, and Daxx, rapidly associate with the foreign genome and form what appears to be fully functional PML nuclear bodies (Everett and Murray 2005). The formation of PML nuclear bodies at sites of viral transcription has been proposed to be a first-line antiviral defense, possibly via PML-mediated modulation of the interferon response (Everett and Chelbi-Alix 2007). A similarly physiological and possibly protective de novo formation event occurs in response to acute cellular stress. Heat shock induces the transcription of the normally silent satellite III repeats on human chromosome 9 (Jolly et al. 2002). Transcriptional activation of these sequences leads to accumulation of various RNAbinding proteins and several SR protein splicing factors to form de novo a morphologically distinct nuclear stress body (Denegri et al. 2001; Metz et al. 2004) The strongest evidence for de novo formation of nuclear bodies comes from studies on the Cajal body. Transient overexpression of Cajal body components or the SMN protein, which is enriched in Cajal-body-associated structures called gems, leads to formation of Cajal bodies in cells that usually do not contain them (Sleeman et al. 2001). More direct evidence for de novo formation of Cajal bodies comes from tethering experiments in which components of the Cajal body were artificially immobilized on chromatin. Immobilization of individual Cajal body components invariably resulted in formation of a fully functional and morphologically intact Cajal body (Kaiser et al. 2008). Since the site of tethering was random and did not represent an active gene locus, these results demonstrate the capacity of Cajal bodies to form de novo and in a DNA sequence-independent manner in vivo.

A ROLE FOR RNA IN NUCLEAR BODY FORMATION

RNA is emerging as an important structural component of nuclear bodies. It may serve two roles in their biogenesis. For one, nascent RNAs may act as templates for the formation of nuclear bodies. Most activity-dependent nuclear bodies form around sites of transcription such as the nucleolus around rDNA, the histone locus bodies near histone gene clusters, or nuclear stress granules in response to activation of satellite III repeats upon heat stress. The nascent RNA transcripts generated at these loci act as a template to attract and retain RNA-interacting proteins, leading to the formation of a nuclear body. An additional role of RNA in nuclear body biogenesis is its function as a dedicated architectural element. Many non-coding RNAs (ncRNAs) play important functions in nuclear RNA metabolic complexes such as tRNAs, snRNAs, or rRNAs. It had long been hypothesized that ncRNAs may also contribute to nuclear architecture (Huang and Spector 1996; Prasanth et al. 2005). This proposal was based on the finding that poly(A)-RNAs are retained for extended periods of time in nuclear speckles after inhibition of transcription, at a time when all exported RNAs should have left the nucleus (Huang and Spector 1996). The recent identification of the first architectural RNAs confirms the role of ncRNAs as structural elements of the nucleus. The NEAT1 (also known as MEN-1/b), a long stable nuclearretained polyadenylated ncRNA (Chen and Carmichael 2009; Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009; reviewed in Bond and Fox 2009), was found to be essential for the formation of paraspeckles. While other protein components of paraspeckles are also located diffusely throughout the nucleoplasm, NEAT1 is found almost exclusively in paraspeckles. Knockdown of these ncRNAs by antisense or RNAi induces the disruption of paraspeckles. Interestingly, knockdown of paraspeckle-specific proteins, p54nrb/NONO and PSF, which interact with NEAT1, induce similar paraspeckle disintegration, indicating

M. Dundr and T. Misteli

that NEAT1 – paraspeckle protein interactions are crucial for paraspeckle integrity (Chen and Carmichael 2009; Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009). Interestingly, paraspeckles are not present in human embryonic stem cells (hESCs) due to the lack of NEAT1 expression (Chen and Carmichael 2009). Expression of NEAT1 is activated when hESCs are induced to differentiate into trophoblasts, which is accompanied by the appearance of paraspeckles. Thus, it seems that paraspeckle formation is linked to the loss of pluripotency in hESCs. Moreover, NEAT1 is strongly upregulated during muscle differentiation concomitant with an increase in the number of paraspeckles in differentiated myotubes, possibly pointing to a role in physiological control of paraspeckle number (Sunwoo et al. 2009). These findings make it likely that ncRNAs play a key role as structural components in nuclear bodies and open the door for identification of architectural ncRNAs in other nuclear bodies. NUCLEAR BODY ASSEMBLY PATHWAYS

In theory, NBs can be assembled by two distinct mechanisms: (1) The biogenesis may involve a tightly controlled sequence of assembly steps or (2) it may occur via the largely stochastic and random interaction of individual components without a strict hierarchical order of assembly (Fig. 3). Recent experiments in which

Cajal body components were irreversibly tethered to chromatin to assess their ability to nucleate a Cajal body have tested these possibilities and the results support a stochastic assembly model. Immobilization of individual CB components to an engineered random site in the genome was shown to be sufficient to initiate the formation of morphologically normal and functional Cajal bodies de novo (Kaiser et al. 2008). De novo Cajal bodies formed after tethering of structural components such as coilin or SMN, which are required for formation, since their depletion by RNAi abolishes Cajal body biogenesis. Importantly, Cajal body formation can also be initiated by other functional groups of Cajal body components, including spliceosomal snRNPs, the SMN protein complex, small nucleolar RNPs, and small Cajalbody specific RNPs, indicating the absence of a requirement for a strict sequence of assembly events. While these experiments used the activity-independent Cajal body as a model system, it remains to be seen whether formation of activity-dependent nuclear bodies have a stricter requirement for sequential assembly steps. A ROLE FOR SELF-ORGANIZATION IN NUCLEAR BODY BIOGENESIS

Several features of nuclear bodies point to a role for self-organization in their biogenesis (reviewed in Misteli 2001, 2007). First, the intrinsically

Sequential ordered assembly

Stochastic self-organization Nuclear body

Figure 3. Theoretical nuclear body assembly pathways. Assembly of nuclear bodies may either involve a hierarch-

ical and ordered assembly pathway in which assembly occurs by association of nuclear body components in a strictly defined sequence of steps or, alternatively, in a stochastic manner in which components assemble in random order.

Nuclear Body Biogenesis

dynamic nature of all nuclear bodies in which a stable structure is formed by the dynamic exchange and interplay of multiple factors is a hallmark of self-organized structures. The dynamic nature of interactions within the body permits the rapid, combinatorial exploration of various binding partners to achieve the most stable network of interactions. The nature of this network ultimately determines the size and possibly the shape of the structure. The existence of nuclear bodies as dynamic steady-state structures combines two opposing needs of a nuclear body: the requirement for a stable, permanent structural entity and the requirement to respond rapidly to changing environmental conditions by changing its architecture. The dynamic nature of interactions facilitates rapid reorganization of the nuclear body structure, for example in response to environmental cues, by altering the interactions of a few nuclear body components. These relatively small changes have the potential to lead to large changes in structure. A second indicator of the self-organizing nature of nuclear bodies is the finding that de novo assembly of Cajal bodies does not follow a strict and linear assembly pathway but rather that many, in fact the majority, of Cajal body components can trigger full assembly of the body (Kaiser et al. 2008). Although this observation alone does not demonstrate a role for self-organization, it is consistent with it. Despite the absence of experimental data to test the assembly pathways for other nuclear bodies, it is very likely that most activity-dependent nuclear bodies follow similar assembly principles once they are nucleated by an upstream component, often a nascent RNA such as the case for the nucleolus or the histone locus body. A further hallmark of the self-organizing nature of nuclear bodies is their limited size. Self-organizing systems often are characterized by size criticality in which a structure does not grow beyond a certain size. This limiting size is set by intrinsic properties of the system, such as the combination of interactions within the structure or the flow of material through the structure, but it is generally independent of concentration. Concentration-independent size regulation is found in at least some nuclear

bodies. Cajal bodies do not grow beyond a certain size when nucleated by overexpression of large amounts of components; neither do de novo-formed Cajal bodies nucleated by tethering grow beyond their typical size despite an excess of building components in the nucleus (Sleeman et al. 2001; Kaiser et al. 2008). Similarly, the size of most other nuclear bodies falls within awell-defined range and may only change in response to cellular cues, which likely affect the flow and the interactions of proteins within the structure, thus changing the intrinsic critical size of the body. Self-organization is likely a driving force in nuclear body biogenesis. Similar properties such as a high rate of flow, size-criticality, and a high degree of plasticity are also hallmarks of other cellular compartments defined by membranes, such as the Golgi complex or the endosomal system, as well as of other cellular structures, such as the actin cytoskeleton. It is therefore likely that the self-organizing nature of nuclear bodies is not the exception but rather the rule in cellular biogenesis, and insights from nuclear body biogenesis will be applicable to understanding cellular biogenesis in general. MITOTIC INHERITANCE OF NUCLEAR BODIES

The assembly and disassembly of nuclear bodies is of critical relevance for their inheritance during cell division. Some nuclear bodies that are present in high copy number persist through cell division in the form of mitotic bodies leading to the equal segregation of their components between daughter cells based on their stochastic distribution throughout the cell volume. Other nuclear bodies, including the activity-associated bodies, disassemble during cell division and must be rapidly re-assembled as cells enter G1 phase. Nucleolus

At the onset of mitosis, when transcription is blocked by the mitosis-dependent phosphorylation of RNA polymerase I transcription factors and processing components, the nucleolus

M. Dundr and T. Misteli

breaks down (Gautier et al. 1992; Leung et al. 2004; Boisvert et al. 2007). By the time cells reach prophase, processing of pre-rRNA is also inhibited, leading to accumulation of unprocessed and partially processed pre-rRNAs with many rRNA processing components in the periphery of condensed chromosomes and dispersed throughout the cytoplasm (Dundr and Olson 1998; Van Hooser et al. 2005). During anaphase, pre-rRNA processing intermediates with associated processing components appear in numerous highly mobile nucleolus-derived foci (NDFs) scattered throughout the cytoplasm (Dundr et al. 2000). These structures are the mitotic remnants of the interphase nucleolus. Once transcription resumes in early telophase, pre-rRNA intermediates with associated processing components are released from disintegrating NDFs and the surface of chromosomes. After the release they accumulate in the prenucleolar bodies (PNBs) in the nucleoplasm of newly formed daughter nuclei. As chromosomes decondense in late telophase, these components are sequentially released from PNBs to re-build nucleoli around transcriptionally active nucleolar organizing regions (NORs). Thus, pre-rRNA intermediates and processing components are recycled from one cell cycle to the next (Dundr et al. 2000; Savino et al. 2001; Boisvert et al. 2007; Prieto and McStay 2008). Moreover, the re-formation of nucleoli after mitosis is associated with resumption of RNA polymerase I transcription since microinjection of antiRNA polymerase I antibodies into mitotic cells blocks nucleolus re-formation (Benavente et al. 1987). In addition, nucleolar re-formation depends on resumption of pre-rRNA processing. Inactivation of phosphorylation activity of cyclin B/Cdk1 in metaphase by the specific inhibitor roscovitine leads to reactivation of RNA polymerase I transcription but not pre-rRNA processing (Sirri et al. 2000). This strongly indicates that these two events are reactivated in a coordinated manner. Thus, NORs at the end of mitosis need to sequester components of both the transcription and processing machineries prior to the onset of transcription. This conclusion is supported by analysis of pseudoNORs, engineered arrays containing multiple

enhancer elements of ribosomal gene stably integrated in human cells (Prieto and McStay 2008). These pseudo-NORs are capable of triggering the formation of a nuclear structure without RNA polymerase I transcription but they only accumulate the RNA polymerase I transcription machinery plus factors required for early events in pre-rRNA maturation, but not later processing factors (Prieto and McStay 2007; 2008). The differential effects of early and late steps in rRNA synthesis on nucleolus morphology indicate that they make independent and additive contributions to nucleolar structure. Nuclear Speckles

During early mitosis, nuclear speckles disassemble and their components scatter throughout the cytoplasm. Reassembly of nuclear speckles occurs in telophase when components aggregate into cytoplasmic speckles, referred to as mitotic interchromatin granule clusters (Ferreira et al. 1994; Prasanth et al. 2003). Once the nuclear envelope is formed in telophase, the RNA polymerase II transcription machinery enters the nucleus followed by pre-mRNA splicing components and other pre-mRNA processing factors (Prasanth et al. 2003; Bubulya et al. 2004). In late telophase, SR splicing factors accumulate briefly in nucleolus associated patches (NAPs) around active nucleolar organizing regions, whereas spliceosomal snRNPs are enriched in other regions of newly formed daughter nuclei. NAPs form very transiently and disintegrate when newly formed nuclear speckles appear in G1 phase. SR splicing proteins are hypo-phosphorylated in NAPs, which may be necessary for SR protein modification for their activation to a splicing-competent state, prior to their targeting to newly formed nuclear speckles (Bubulya et al. 2004). These findings indicate that the pre-mRNA splicing machinery is recruited to transcription sites upon its entry into daughter nuclei, and nuclear speckles form in an activity-dependent manner in early G1 phase in response to the establishment of a steady level of RNA polymerase II transcription.

Nuclear Body Biogenesis

Paraspeckles

Cajal Body

Formation of paraspeckles is initiated after cell division. Once paraspeckles are formed, they remain stable throughout the cell cycle and persist in mitosis until anaphase, when they scatter throughout the cell (Fox et al. 2005). Disappearance of cytoplasmic paraspeckles in early telophase, when daughter nuclei are formed, is coordinated with the nuclear entry of PSP1, a marker protein of paraspeckles. Individual paraspeckles begin to form in early G1 phase until approximately 1 hour after RNA transcription is initiated (Fox et al. 2005). Importantly, blocking the initiation of RNA polymerase II transcription by the inhibitor DRB at the end of mitosis prevents formation of paraspeckles, suggesting that these structures are dependent on ongoing transcription (Fox et al. 2005; Clemson et al. 2009). The earliest paraspeckles are often positioned next to the NEAT1 gene locus and an increase in NEAT1 expression by ectopic plasmids leads to a corresponding increase in the number of paraspeckles, which is essentially unchanged upon PSP1 overexpression (Clemson et al. 2009). However, eventually additional paraspeckles form, and they appear to be nonrandomly distributed throughout the nucleoplasm, suggesting their association with specific, yet unidentified, gene loci.

The number of Cajal bodies differs during the cell cycle and is maximal in G1/S phase. Cajal bodies are frequently associated with snRNA genes and histone genes in a transcriptiondependent manner (Frey and Matera 1995, 2001; Cioce and Lamond 2005; Matera and Shpargel 2006). At the onset of mitosis, Cajal bodies remain intact, and they scatter throughout the cytoplasm without physical contact with condensed chromosomes. The number and size of mitotic Cajal bodies remain approximately unchanged from metaphase to telophase. Importantly, mitotic Cajal bodies contain spliceosomal snRNPs through the course of mitosis even without transcription. Once the nuclear envelope is formed in daughter cells during early telophase, mitotic Cajal bodies disintegrate and coilin rapidly enters the nuclear interior, where it is initially diffusely distributed. Newly re-formed Cajal bodies appear in daughter cells later in G1 phase once transcription is fully established (Carmo-Fonseca et al. 1993; Ferreira et al. 1994).

Histone Locus Body

Histone locus bodies are nuclear bodies that form around the replication-dependent histone gene clusters, which are only active in S phase (Marzluff et al. 2008). Histone locus bodies persist until early prometaphase and disintegrate in metaphase until telophase. When diploid cells enter G1 phase, two histone locus bodies are formed and in S phase their number increases to four, reflecting the number of histone gene clusters (Zhao et al. 2000; Bongiorno-Borbone et al. 2008). It is neither known in mammalian cells whether histone locus bodies form synchronously with Cajal bodies, nor, whether histone locus bodies and Cajal bodies form as distinct nuclear bodies and then fuse or whether they assemble as a joint structure.

PML Nuclear Body

As cells enter mitosis, the number of PML nuclear bodies decreases by aggregation into larger Mitotic Accumulations of the PML Protein (MAPPs) (Dellaire et al. 2006; Chen et al. 2008). The PML protein, a structural component of PML nuclear bodies, is de-SUMOylated during mitosis, which affects its ability to recruit PML nuclear body components and likely influences MAPP formation (Dellaire et al. 2006). MAPPs become more mobile by relaxing their tethering to chromatin regions, and they progressively lose key components such as Sp100, Daxx or SUMO-1, from prophase to telophase. A subset of MAPPs physically interacts with mitotic chromosomes, which likely provide a source of PML protein for PML nuclear body formation in newly forming daughter nuclei. Upon exit from mitosis, after the nuclear envelope is fully reformed, Sp100 and Daxx enter daughter nuclei first before their targeting into newly formed PML nuclear

M. Dundr and T. Misteli

bodies in early G1 phase (Chen et al. 2008). The loss of MAPPs from the cytoplasm and the increase in PML nuclear body number indicates that components of MAPPs are being incorporated into PML bodies even though a large portion of PML aggregates persist in the cytoplasm during G1 phase, slowly decreasing in number over time (Dellaire et al. 2006; Chen et al. 2008). PHYSIOLOGICAL BIOGENESIS OF NUCLEAR BODIES

Biogenesis of nuclear bodies is important during various physiological responses. The de novo biogenesis of heat-shock bodies is a dramatic example of formation of nuclear bodies as part of a specific cellular response. Cells respond to stressful conditions by widespread inhibition of transcription and activation of specific heat-shock gene expression programs. One of the cellular events during heat shock is the transcriptional activation of tandem arrays of repetitive noncoding satellite III (sat III) repeats located on chromosome 9q12 (Valgardsdottir et al. 2008). Upon induction, the sat III transcripts remain associated with the locus and act as scaffolds for recruitment of several transcription and splicing factors to form nuclear stress bodies (Denegri et al. 2001; Alastalo et al. 2003). Their formation is strictly dependent on the presence of sat III transcripts, and they rapidly disassemble upon return to physiological temperature (Metz et al. 2004). It has been proposed that formation of nuclear stress bodies provides a dynamic means to sequester and release specific protein factors in normal and/or stressed cells and that sat III transcripts may be involved in modulating epigenetic changes during the heat-shock response (Biamonti 2004). PML nuclear body formation has been implicated in responses to viral infection (Everett and Chelbi-Alix 2007). DNAviruses commonly localize and replicate their genomes in proximity to PML nuclear bodies (reviewed in Maul 1998). In addition, PML bodies form de novo at the sites of viral genome deposition in the nucleus (Everett and Murray 2005). Association of

sites of viral DNA replication with PML nuclear bodies indicates that these nuclear bodies may enhance viral DNA replication (Sourvinos and Everett 2002). The fact that many viral proteins accumulate in PML nuclear bodies and then mediate PML nuclear body disruption suggests antiviral activity associated with these domains. In fact, failure to disrupt PML nuclear bodies during infection with HSV-1 virus inhibits viral DNA replication (Burkham et al. 2001). In line with an antiviral role of PML nuclear body biogenesis, exogenous expression of PML isoform III or PML VI can impede infection by a number of RNA and DNA viruses (Everett and Chelbi-Alix 2007) and upon treatment of cells with interferon, which is produced in response to viral infection. PML nuclear bodies undergo a dramatic increase in size and number (Lavau et al. 1995), possibly as a consequence of interferon induced expression of several PML-nuclear body components, including PML itself (ChelbiAlix et al. 1995; Fabunmi et al. 2001). Although not well-characterized for most nuclear bodies, their appearance also changes during development. For example, in Drosophila, nurse nuclei of the youngest egg chambers contain a single HLB and a single CB, but as the egg chamber grows, the number of HLBs increases to 8– 16 HLBs with one or two prominent CBs (Liu at al. 2009). In older egg chambers, CBs break up into multiple small bodies that eventually disappear, but the number and size of HLBs remain the same and they start to accumulate coilin. Thus, at the end of oogenesis, nurse cell nuclei contain prominent HLBs positive for coilin but lack typical CBs. It seems likely that most nuclear bodies undergo changes in shape and size as organisms develop. CONCLUSIONS

Nuclear bodies are prominent features in the nuclear landscape. They carry out important nuclear functions and contribute to cellular responses. While their functions are gradually being uncovered, their biogenesis is poorly understood. Elucidation of the molecular mechanisms for the maintenance and formation is of great interest not only as part of ongoing

Nuclear Body Biogenesis

efforts to understand the role of nuclear architecture in genome function, but also because nuclear bodies are unique amongst cellular organelles in that they are devoid of defining membrane, yet they are distinct and stable structures. Insights into their biogenesis will likely reveal how higher-order protein aggregates form in cells and might have implications for the assembly of large macromolecular complexes and other cellular organelles. It is highly probable that the next few years will bring dramatic progress in our still rudimentary understanding of nuclear body biogenesis. The availability of new experimental tools promises to explore the molecular basis of nuclear-body architecture. Proteomic analysis of nuclear bodies will yield a thorough understanding of the composition of each body. Using these component lists, the role of each can be systematically probed by RNAi-based approaches, which will identify the components necessary for nuclear body establishment and maintenance. Conversely, the recently developed tethering approaches allow for the first time testing of what protein, and possibly RNA, components are sufficient to form nuclear bodies. The combination of these tools should yield a thorough picture of the mechanisms and principles involved in nuclear body formation. In parallel, an even greater challenge is the full understanding of the physiological role of nuclear bodies and their formation. This question is now also amenable to experimental investigation using similar tools in systems of differentiation and disease. Although the biogenesis of nuclear bodies is still mysterious, there is little doubt that this area is likely one with the most potential for future discoveries—and surprises.

REFERENCES Alastalo TP, Hellesuo M, Sandqvist A, Hietakangas V, Kallio M, Sistonen L. 2003. Formation of nuclear stress granules involves HSF2 and coincides with the nucleolar localization of Hsp70. J Cell Sci 116: 3557–3570. Benavente R, Rose KM, Reimer G, Hu¨gle-Do¨rr B, Scheer U. 1987. Inhibition of nucleolar reformation after microinjection of antibodies to RNA polymerase I into mitotic cells. J Cell Biol 105: 1483– 1491.

Bernardi R, Pandolfi PP. 2007. Structure, dynamics and functions of promyelocytic leukemia nuclear bodies. Nat Rev Mol Cell Biol 8: 1006–1016. Biamonti G. 2004. Nuclear stress bodies: a heterochromatin affair? Nat Rev Mol Cell Biol 5: 493–498. Bongiorno-Borbone L, De Cola A, Vernole P, Finos L, Barcaroli D, Knight RA, Melino G, De Laurenzi V. 2008. FLASH and NPAT positive but not Coilin positive Cajal Bodies correlate with cell ploidy. Cell Cycle 7: 2357–2367. Boisvert FM, van Koningsbruggen S, Navascue´s J, Lamond AI. 2007. The multifunctional nucleolus. Nat Rev Mol Cell Biol 8: 574– 585. Bond CS, Fox AH. 2009. Paraspeckles: nuclear bodies built on long noncoding RNA. J Cell Biol 186: 637 –644. Bubulya PA, Prasanth KV, Deerinck TJ, Gerlich D, Beaudouin J, Ellisman MH, Ellenberg J, Spector DL. 2004. Hypophosphorylated SR splicing factors transiently localize around active nucleolar organizing regions in telophase daughter nuclei. J Cell Biol 167: 51–63. Burkham J, Coen DM, Hwang CB, Weller SK. 2001. Interactions of herpes simplex virus type 1 with ND10 and recruitment of PML to replication compartments. J Virol 75: 2353–2367. Carmo-Fonseca M, Ferreira J, Lamond AI. 1993. Assembly of snRNP-containing coiled bodies is regulated in interphase and mitosis– evidence that the coiled body is a kinetic nuclear structure. J Cell Biol 120: 841 –852. Carmo-Fonseca M. 2007. How genes find their way inside the cell nucleus. J Cell Biol 179: 1093–1094. Chelbi-Alix MK, Pelicano L, Quignon F, Koken MH, Venturini L, Stadler M, Pavlovic J, Degos L, de The´ H. 1995. Induction of the PML protein by interferons in normal and APL cells. Leukemia 9: 2027– 2033. Chen D, Huang S. 2001. Nucleolar components involved in ribosome biogenesis cycle between the nucleolus and nucleoplasm in interphase cells. J Cell Biol 153: 169 –176. Chen L-L, Carmichael GG. 2009. Altered nuclear retention of mRNAs containing inverted repeats in human embryonic stem cells: functional role of a nuclear noncoding RNA. Mol Cell 35: 467 –478. Chen YC, Kappel C, Beaudouin J, Eils R, Spector DL. 2008. Live cell dynamics of promyelocytic leukemia nuclear bodies upon entry into and exit from mitosis. Mol Biol Cell 19: 3147– 3162. Cioce M, Lamond AI. 2005. Cajal bodies: a long history of discovery. Annu Rev Cell Dev Biol 21: 105– 131. Clemson CM, Hutchinson JN, Sara SA, Ensminger AW, Fox AH, Chess A, Lawrence JB. 2009. An architectural role for a nuclear noncoding RNA: NEAT1 RNA is essential for the structure of paraspeckles. Mol Cell 33: 717– 726. Dellaire G, Eskiw CH, Dehghani H, Ching RW, Bazett-Jones DP. 2006. Mitotic accumulations of PML protein contribute to the re-establishment of PML nuclear bodies in G1. J Cell Sci 119: 1034–1042. Denegri M, Chiodi I, Corioni M, Cobianchi F, Riva S, Biamonti G. 2001. Stress-induced nuclear bodies are sites of accumulation of pre-mRNA processing factors. Mol Biol Cell 12: 3502–3514.

M. Dundr and T. Misteli Dundr M, Olson MO. 1998. Partially processed pre-rRNA is preserved in association with processing components in nucleolus-derived foci during mitosis. Mol Biol Cell 9: 2407– 2422. Dundr M, Misteli T, Olson MO. 2000. The dynamics of postmitotic reassembly of the nucleolus. J Cell Biol 150: 433–446. Dundr M, Misteli M. 2001. Functional architecture in the cell nucleus. Biochem J 356: 297 –310. Dundr M, Hebert MD, Karpova TS, Stanek D, Xu H, Shpargel KB, Meier UT, Neugebauer KM, Matera AG, Misteli T. 2004. In vivo kinetics of Cajal body components. J Cell Biol 164: 831–842. Dundr M, Ospina JK, Sung M-H, John S, Upender M, Ried T, Hager G, Matera GA. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095– 1103. Everett RD, Murray J. 2005. ND10 components relocate to sites associated with herpes simplex virus type 1 nucleoprotein complexes during virus infection. J Virol 79: 5078– 5089. Everett RD, Chelbi-Alix MK. 2007. PML and PML nuclear bodies: implications in antiviral defence. Biochimie 89: 819–830. Fabunmi RP, Wigley WC, Thomas PJ, DeMartino GN. 2001. Interferon gamma regulates accumulation of the proteasome activator PA28 and immunoproteasomes at nuclear PML bodies. J Cell Sci 114: 29–36. Ferreira JA, Carmo-Fonseca M, Lamond AI. 1994. Differential interaction of splicing snRNPs with coiled bodies and interchromatin granules during mitosis and assembly of daughter cell nuclei. J Cell Biol 126: 11–23. Fox AH, Bond CS, Lamond AI. 2005. p54nrb forms a heterodimer with PSP1 that localizes to paraspeckles in an RNA-dependent manner. Mol Biol Cell 16: 5304–5315. Frey MR, Matera AG. 1995. Coiled bodies contain U7 small nuclear RNA and associate with specific DNA sequences in interphase human cells. Proc Natl Acad Sci U S A 92: 5915– 5919. Frey MR, Matera AG. 2001. RNA-mediated interaction of Cajal bodies and U2 snRNA genes. J Cell Biol 154: 499–509. Gautier T, Robert-Nicoud M, Guilly MN, HernandezVerdun D. 1992. Relocation of nucleolar proteins around chromosomes at mitosis. A study by confocal laser scanning microscopy. J Cell Sci 102: 729– 737. Go¨risch SM, Wachsmuth M, Ittrich C, Bacher CP, Rippe K, Lichter P. 2004. Nuclear body movement is determined by chromatin accessibility and dynamics. Proc Natl Acad Sci U S A 101: 13221– 13226. Handwerger KE, Gall JG. 2006. Subnuclear organelles: new insights into form and function. Trends Cell Biol 16: 19–26. Hearst SM, Gilder AS, Negi SS, Davis MD, George EM, Whittom AA, Toyota CG, Husedzinovic A, Gruss OJ, Hebert MD. 2009. Cajal-body formation correlates with differential coilin phosphorylation in primary and transformed cell lines. J Cell Sci 122: 1872–1881. Hebert MD, Matera AG. 2000. Self-association of coilin reveals a common theme in nuclear body localization. Mol Biol Cell 11: 4159–4171.

Hernandez-Verdun D. 2006. The nucleolus: a model for the organization of nuclear functions. Histochem. Cell Biol 126: 135–148. Hu Y, Kireev I, Plutz M, Ashourian N, Belmont AS. 2009. Large-scale chromatin structure of inducible genes: transcription on a condensed, linear template. J Cell Biol 185: 87– 100. Huang S, Spector DL. 1996. Intron-dependent recruitment of pre-mRNA splicing factors to sites of transcription. J Cell Biol 133: 719– 732. Jolly C, Konecny L, Grady DL, Kutskova YA, Cotto JJ, Morimoto RI, Vourc’h C. 2002. In vivo binding of active heat shock transcription factor 1 to human chromosome 9 heterochromatin during stress. J Cell Biol 156: 775 –781. Kaiser TE, Intine RV, Dundr M. 2008. De novo formation of a subnuclear body. Science 322: 1713–1717. Kentsis A, Gordon RE, Borden KL. 2002. Control of biochemical reactions through supramolecular RING domain self-assembly. Proc Natl Acad Sci U S A 99: 15404– 15409. Kruhlak MJ, Lever MA, Fischle W, Verdin E, Bazett-Jones DP, Hendzel MJ. 2000. Reduced mobility of the alternate splicing factor (ASF) through the nucleoplasm and steady state speckle compartments. J Cell Biol 150: 41–51. Lamond AI, Spector DL. 2003. Nuclear speckles: a model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605 –612. Lavau C, Marchio A, Fagioli M, Jansen J, Falini B, Lebon P, Grosveld F, Pandolfi PP, Pelicci PG, Dejean A. 1995. The acute promyelocytic leukaemia-associated PML gene is induced by interferon. Oncogene 11: 871 –876. Leung AK, Gerlich D, Miller G, Lyon C, Lam YW, Lleres D, Daigle N, Zomerdijk J, Ellenberg J, Lamond AI. 2004. Quantitative kinetic analysis of nucleolar breakdown and reassembly during mitosis in live human cells. J Cell Biol 166: 787– 800. Liu J-L, Wu Z, Nizami Z, Deryusheva S, Rajendra TK, Beumer KJ, Gao H, Matera AG, Carroll D, Gall JG. 2009. Coilin is essential for Cajal body organization in Drosophila melanogaster. Mol Cell Biol 20: 1661– 1670. Mais C, McStay B, Scheer U. 2002. On the formation of amplified nucleoli during early Xenopus oogenesis. J Struct Biol 140: 214 –226. Marzluff WF, Wagner EJ, Duronio RJ. 2008. Metabolism and regulation of canonical histone mRNAs: life without a poly(A) tail. Nat Rev Genet 9: 843–854. Matera AG, Shpargel KB. 2006. Pumping RNA: nuclear bodybuilding along the RNP pipeline. Curr Opin Cell Biol 18: 317 –324. Maul GG. 1998. Nuclear domain 10, the site of DNA virus transcription and replication. Bioessays 20: 660 –667. Metz A, Soret J, Vourc’h C, Tazi J, Jolly C. 2004. A key role for stress-induced satellite III transcripts in the relocalization of splicing factors into nuclear stress granules. J Cell Sci 117: 4551– 4558. Misteli T, Caceres JF, Spector DL. 1997a The dynamics of a pre-mRNA splicing factor in living cells. Nature 387: 523 –527. Misteli TD, Spector L 1997b. Protein phosphorylation and the nuclear organization of pre-mRNA splicing. Trends in Cell Biol 7: 135 –138.

Nuclear Body Biogenesis Misteli T. 2001. The concept of self-organization in cellular architecture. J Cell Biol 155: 181 –185. Misteli T. 2007. Beyond the sequence: cellular organization of genome function. Cell 128: 787 –800. Morse R, Shaw DJ, Todd AG, Young PJ. 2007. Targeting of SMN to Cajal bodies is mediated by self-association. Hum Mol Genet 16: 2349–2358. Muratani M, Gerlich D, Janicki SM, Gebhard M, Eils R, Spector DL. 2002. Metabolic-energy-dependent movement of PML bodies within the mammalian cell nucleus. Nat Cell Biol 4: 106– 110. Nierras CR, Liebman SW, Warner JR. 1997. Does Saccharomyces need an organized nucleolus? Chromosoma 105: 444–451. Oakes M, Aris JP, Brockenbrough JS, Wai H, Vu L, Nomura M. 1998. Mutational analysis of the structure and localization of the nucleolus in the yeast Saccharomyces cerevisiae. J Cell Biol 143: 23–34. Olson MO, Dundr M. 2005. The moving parts of the nucleolus. Histochem. Cell Biol 123: 203 –216. Pederson T, Tsai RY. 2009. In search of nonribosomal nucleolar protein function and regulation. J Cell Biol 184: 771–776. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604–609. Platani M, Goldberg I, Swedlow JR, Lamond AI. 2000. In vivo analysis of Cajal body movement, separation, and joining in live human cells. J Cell Biol 151: 1561– 1574. Platani M, Goldberg I, Lamond AI, Swedlow JR. 2002. Cajal body dynamics and association with chromatin are ATPdependent. Nat Cell Biol 4: 502–508. Prasanth KV, Sacco-Bubulya PA, Prasanth SG, Spector DL. 2003. Sequential entry of components of the gene expression machinery into daughter nuclei. Mol Biol Cell 14: 1043– 1057. Prasanth KV, Prasanth SG, Xuan Z, Hearn S, Freier SM, Bennett CF, Zhang MQ, Spector DL. 2005. Regulating gene expression through RNA nuclear retention. Cell 123: 249–263. Prieto JL, McStay B. 2007. Recruitment of factors linking transcription and processing of pre-rRNA to NOR chromatin is UBF-dependent and occurs independent of transcription in human cells. Genes Dev 21: 2041– 2054. Prieto JL, McStay B. 2008. Pseudo-NORs: a novel model for studying nucleoli. Biochim Biophys Acta 1783: 2116– 2123. Sunwoo H, Dinger ME, Wilusz JE, Amaral PP, Mattick JS, Spector DL. 2009. MEN epsilon/beta nuclear-retained non-coding RNAs are up-regulated upon muscle differentiation and are essential components of paraspeckles. Genome Res 19: 347–359. Sasaki YT, Ideue T, Sano M, Mituyama T, Hirose T. 2009. MENepsilon/beta noncoding RNAs are essential for

structural integrity of nuclear paraspeckles. Proc Natl Acad Sci U S A 106: 2525– 2530. Savino TM, Ge´brane-Youne`s J, De Mey J, Sibarita JB, Hernandez-Verdun D. 2001. Nucleolar assembly of the rRNA processing machinery in living cells. J Cell Biol 153: 1097– 1110. Shaw PJ, Jordan EG. 1995. The nucleolus. Annu Rev Cell Dev Biol 11: 93– 121. Shopland LS, Johnson CV, Byron M, McNeil J, Lawrence JB. 2003. Clustering of multiple specific genes and gene-rich R-bands around SC-35 domains: evidence for local euchromatic neighborhoods. J Cell Biol 162: 981– 990. Shpargel KB, Ospina JK, Tucker KE, Matera AG, Hebert MD. 2003. Control of Cajal body number is mediated by the coilin C-terminus. J Cell Sci 116: 303 –312. Sirri V, Hernandez-Verdun D, Roussel P. 2002. Cyclindependent kinases govern formation and maintenance of the nucleolus. J Cell Biol 156: 969–981. Sleeman JE, Ajuh P, Lamond AI. 2001. snRNP protein expression enhances the formation of Cajal bodies containing p80-coilin and SMN. J Cell Sci 114: 4407– 4419. Snaar S, Wiesmeijer K, Jochemsen AG, Tanke HJ, Dirks RW. 2000. Mutational analysis of fibrillarin and its mobility in living human cells. J Cell Biol 151: 653–662. Sourvinos G, Everett RD. 2002. Visualization of parental HSV-1 genomes and replication compartments in association with ND10 in live infected cells. EMBO J 21: 4989– 4997. Spector DL. 2001. Nuclear domains. J Cell Sci 114: 2891– 2893. Tsai RY, McKay RD. 2005. A multistep, GTP-driven mechanism controlling the dynamic cycling of nucleostemin. J Cell Biol 168: 179– 184. Valgardsdottir R, Chiodi I, Giordano M, Rossi A, Bazzini S, Ghigna C, Riva S, Biamonti G. 2008. Transcription of Satellite III non-coding RNAs is a general stress response in human cells. Nucleic Acids Res 36: 423–434. Van Hooser AA, Yuh P, Heald R. 2005. The perichromosomal layer. Chromosoma 114: 377– 388. Weidtkamp-Peters S, Lenser T, Negorev D, Gerstner N, Hofmann TG, Schwanitz G, Hoischen C, Maul G, Dittrich P, Hemmerich P. 2008. Dynamics of component exchange at PML nuclear bodies. J Cell Sci 121: 2731– 2743. Zhao J, Kennedy BK, Lawrence BD, Barbie DA, Matera AG, Fletcher JA, Harlow E. 2000. NPAT links cyclin E-Cdk2 to the regulation of replication-dependent histone gene transcription. Genes Dev 14: 2283– 2297. Zhao R, Bodnar MS, Spector DL. 2009. Nuclear neighborhoods and gene expression. Curr Opin Genet Dev 19: 172 –179.

Organization of Transcription Lyubomira Chakalova1,2 and Peter Fraser1 1

Laboratory of Chromatin and Gene Expression, The Babraham Institute, Babraham Research Campus, Cambridge, CB22 3AT, United Kingdom

2

Research Centre for Genetic Engineering and Biotechnology, Macedonian Academy of Sciences and Arts, Skopje 1000, Republic of Macedonia

Correspondence: [email protected]

Investigations into the organization of transcription have their origins in cell biology. Early studies characterized nascent transcription in relation to discernable nuclear structures and components. Advances in light microscopy, immunofluorescence, and in situ hybridization helped to begin the difficult task of naming the countless individual players and components of transcription and placing them in context. With the completion of mammalian genome sequences, the seemingly boundless task of understanding transcription of the genome became finite and began a new period of rapid advance. Here we focus on the organization of transcription in mammals drawing upon information from lower organisms where necessary. The emerging picture is one of a highly organized nucleus with specific conformations of the genome adapted for tissue-specific programs of transcription and gene expression.

uch of what is known about eukaryotic transcription is dominated by decades of advances in in vitro biochemistry with wholecell extracts, or subfractionated and recombinant proteins on purified DNA templates. These reductionist approaches have lead to seminal findings describing the basic DNA sequence regulatory elements and enzymatic machinery of transcription. More recent research incorporating genetic approaches has added to the complexity of transcription, involving literally hundreds of factors, cofactors, remodeling complexes, histone modifiers, and elongation-, splicing- and termination-factors required for or associated with a single transcriptional event. Furthermore the discovery of, sometimes

M

distant, sequence elements required for regulated transcription of some genes has added to the intricacy of the transcriptional process that occurs in vivo. Though there is still much to learn, the difficult task of integrating this information and placing it in the context of the nucleus is gathering momentum. The widely held view of transcriptional mechanics, of the RNA polymerase complex sliding along a template to generate a transcript is also dominated by biochemistry. Textbooks are full of descriptions of promoter bound factors recruiting RNA polymerase, which initiates transcription before sliding along the transcription unit. Indeed, single molecules of prokaryotic RNA polymerase have been visualized in

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000729 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000729

L. Chakalova and P. Fraser

vitro, sliding along a fixed DNA template during a one-dimensional diffusional search for a promoter (Kabata et al. 1993; Guthold et al. 1999; Harada et al. 1999) or during transcription (Schafer et al. 1991; Wang et al. 1998; Guthold et al. 1999; Davenport et al. 2000). However, which molecule actually moves, the polymerase or the DNA depends on which is fixed (Iborra et al. 1996b). Several studies have shown that an anchored polymerase generates considerable pulling force on a DNA template, rotating the double helix in a clockwise manner as it threads the strand through the protein during transcription (Kabata et al. 1993; Wang et al. 1998; Guthold et al. 1999). It is impossible to know from these in vitro studies what actually happens in vivo. Although the answer to this question may seem trivial, it has profound implications for our understanding of transcription and genome function, and can only be answered by examining evidence of transcription in the nucleus. Fakan and colleagues have studied nascent transcripts at high resolution for decades (Fakan and Bernhard 1971; Fakan et al. 1976; Fakan 2004). They found that nascent RNA is located in perichromatin fibrils (PF). PF are structures observed using electron microscopy by specific contrasting methods, and most often located in the perichromatin region, the boundaries between condensed and decondensed chromatin (Fakan and Bernhard 1971). Early autoradiographic studies combined with later findings indicated that PF are the in situ form of nascent RNA complexed with processing factors (Nash et al. 1975; Fakan et al. 1976; Cremer et al. 2004). Seminal studies on transcription in mammalian nuclei were carried out by Jackson and Cook (Jackson et al. 1981; Jackson and Cook 1985). They uniformly labeled DNA of HeLa cells with 14C in vivo, and encapsulated them in agarose beads before a short incubation with [3H]uridine to label nascent RNA. The cells were then lysed in an isotonic solution and chromatin was digested with a restriction enzyme or DNase followed by electrophoresis to remove the digested chromatin. They found that over 90% of nascent RNA is retained in

the beads after as much as 98% of the DNA/ chromatin had been removed. The chromatin released from the beads by electrophoresis was assayed further and found to be considerably larger than an RNAPII holocomplex, suggesting that nonattached transcribing complexes should have been released with the chromatin. However, they found that 60% of the original RNA polymerase activity was retained in the beads after loss of 75% of the chromatin. These results suggested a model whereby newly synthesized RNA and the transcriptional machinery are retained in the nucleus by a structure that is resistant to nuclease digestion. Indeed analysis of the DNA retained in the beads indicated that it was enriched in active genes. Jackson and Cook proposed the involvement of a “nucleoskeleton” in the process of transcription, suggesting attachment of the polymerase complex, and arguing against the concept of free RNA polymerase complexes tracking DNA templates. These were controversial studies in their time, and the merits and conclusions are still debated today. Though Jackson and Cook were careful to maintain physiological conditions during their experiments (Jackson and Cook 1985), critics argued that the system employed may have created an artificial network of proteins that retained active gene sequences. TRANSCRIPTION OCCURS IN FACTORIES

These studies were expanded and advanced by Jackson et al. (1993), and particularly Wansink et al. (1993) in which transcription sites were visualized by indirect immunofluorescence after pulse-labeling or microinjecting cells with halogenated ribonucleoside precursors. The use of in vivo labeling methods excluded potential artifacts, which may have been introduced by the earlier run-on procedures. These studies revealed that nascent transcripts were not equally distributed throughout the nucleoplasm, but that transcription occurred in a limited number of discrete sites or foci that were sensitive to transcriptional inhibitors. An important observation was that increased labeling times did not result in detection of more foci, only increased intensity suggesting that all sites of

Organization of Transcription

transcription were being detected. Estimates of the number of transcription foci vary between several hundred and several thousand per nucleus depending on the cell type (Jackson et al. 1993; Iborra et al. 1996a; Pombo et al. 1999; Osborne et al. 2004). The total number of sites appeared to be several times lower than the number of active transcription units suggesting that each site contained multiple genes (Jackson et al. 1993; Jackson et al. 1998). The term “transcription factories” was coined (Iborra et al. 1996a) to reflect the potential gathering of several transcription units to each factory in similarity to replication factories, which form during the S phase of the cell cycle, each accommodating several replicons at once. Fakan and colleagues have often observed that immunogold detection of nascent RNAs after short labeling is most often represented by individual gold particles associated with PF (Cmarko et al. 1999). This was taken as evidence that transcription takes place in individual localized sites rather than in factories. However quantitative estimates based on immunogold labeling on the surface of EM sections is tenuous. Though PF and transcription factories have not been linked directly, probably because with the various imaging techniques used one can only see what one attempts to detect, it seems likely that PFs do represent nascent transcripts (Fakan 1994) produced at focal RNAPII factories (Cmarko et al. 1999). Both groups (Jackson et al. 1993; Wansink et al. 1993), investigated the localization of splicing components relative to nascent transcription sites. Large, intensely labeled SC-35 domains localized close to some nascent transcription sites but did not overlap with them, whereas some colocalization was observed between transcription sites and weak SC-35 foci (Wansink et al. 1993). These observations are consistent with suggestions that large SC-35 domains are storage sites containing no transcriptional activity (Iborra et al. 1996a; Pombo and Cook 1996; Fay et al. 1997), whereas small SC-35 foci may result from recruitment of splicing components to transcription sites forcotranscriptional splicing (Huang and Spector 1996; Misteli et al. 1997; Lamond and Spector 2003).

Building on previous work, Cook and colleagues employed electron microscopy (EM) to study nascent transcripts in pulse labeled cells (Iborra et al. 1996a) using indirect labeling with immuno-gold particles. They found that gold particles marking nascent transcripts appeared as clusters, which increased in particle number with increased labeling times suggesting that the clusters marked synthetic sites. Once again, the total number of clusters did not change with increasing labeling times, which they interpreted as evidence that the technique was sensitive enough to detect all sites. Importantly, they found that the diameter of nascent transcript clusters remained fairly constant (at 75 nm) regardless of the labeling time and increased elongation, even during extended chase periods. Iborra et al. (1996a) suggest that this result is contrary to models in which the polymerase slides along the template during transcription. They argue that such a polymerase tracking mechanism would result in nascent transcripts occupying an increased volume with increasing elongation. Instead they proposed their data to be consistent with a model in which the DNA template slides through a polymerase that is immobilized in a factory. This has been one of the most difficult concepts for many in the transcription field to accept, and perhaps rightly so because it contradicts the textbook view that holds the gene as the central scaffold to which the transcription complex is recruited. Interestingly, in the case of DNA replication, it is widely accepted that DNA is drawn through and extruded from discrete replication factories containing many active polymerases (Berezney et al. 2000). Though this is not proof that a similar mechanism occurs during transcription, it appears to be the most plausible given the available evidence in favor of this concept and relative lack of evidence to the contrary. Iborra et al. (1996a) also examined the relationship between nascent transcript sites and RNAPII by combining detection of nascent RNA and RNAPII with immno-gold particles of different sizes. Gold particles marking RNAPII were also found in discrete clusters throughout the nucleus and averaged 56 nm in diameter. They showed that nascent transcript

L. Chakalova and P. Fraser

gold clusters were intimately associated with RNAPII clusters but did not overlap exactly. The centers of nascent transcript clusters and RNAPII clusters were shifted relative to each other by an average of 24 nm, consistent with two overlapping zones, one rich in transcripts, and the other rich in RNA polymerase II. Simultaneous immunofluorescent detection of RNAPII and nascent transcripts combined with confocal microscopy confirmed this association between focal RNAPII sites and transcriptional activity (Grande et al. 1997). RNAPII-containing foci colocalized with sites of nascent transcript labeling and vice versa, however, there were also many sites strongly labeled for RNAPII that contained little or no BrUTP label, and vice versa. With hindsight, this might be taken as one of the first hints of specialized polymerase factories and the possibility that not all factories are equally active (see below). What actually constitutes the factory is a question open to interpretation. Nascent transcripts are undoubtedly the product but the enzymatic machinery is in essence the factory. The most recent and probably most accurate measurements of transcription factories size employed electron spectroscopic imaging (ESI) (Eskiw et al. 2008). Rather than measure the size of a cluster of gold particles, ESI records atomic signatures (Fig. 1). Transcription factories appear as large proteinacious (nitrogen rich) structures with an average diameter of 87 nm. Chromatin and nascent transcripts with associated RNPs appear as fibrous, relatively phosphorus-rich structures located at the periphery of factories. A GFP-tagged form of the human catalytic subunit of RNAPII (Sugaya et al. 2000) was used to monitor RNAPII kinetics in living cells by fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) (Kimura et al. 2002; Hieda et al. 2005). GFP-RNAPII dynamics were consistent with the existence of at least two populations of RNAPII in nuclei. Most of the tagged RNAPII (75 – 80% of the pool) had a very short recovery time on the order of seconds characteristic of rapid free diffusion as has been seen for other

A

ch

B

*

ch

* * *

*

*

* * IG

* * *

*

* *

*

*

* * * 100 nm

Figure 1. (A) Schematic diagram of transcription of multiple genes at a nuclear RNAPII transcription factory. RNAPII factory shown as central blue circle with three transcribing genes and their associated transcription factors (small colored circles). Nascent transcripts are shown in red, chromatin is dark blue, and splicing components are depicted as small black circles with orange halo. (B) Electron spectroscopic imaging of HeLa cell nucleus. Phosphorous-rich structures are colored red and nitrogen green. Arrows point to nitrogen-rich transcription factory. White dots are immunogold detection of BrdU pulse-labeled nascent transcripts. Asterisks outline a small region of interchromatin granules (IG) and ch denotes regions of relatively compact chromatin. Image courtesy of Dr. Christopher Eskiw.

nuclear factors. The transcriptionally engaged fraction (20 –25%) had a considerably longer recovery time (t1/2 approx. 20 min) consistent with relative immobilization. The latter fraction could be further subdivided into initiating and elongating subfractions of decreasing turnover rates and differential sensitivity to transcriptional inhibitors of initiation and elongation. These results suggested that RNAPII and most

Organization of Transcription

likely other factory components undergo continual but relatively slow dynamic exchange with freely diffusing components, but that the position of the factories themselves may be more stable. Under conditions in which new initiation is specifically and globally inhibited (Allen et al. 2004; Espinoza et al. 2004; Mariner et al. 2008), it was found that expressed genes disengage from transcription factories but focal sites of Ser5-RNAPII were still present after 30 min (Mitchell and Fraser 2008). These findings suggest that factories are not simply aggregates of RNAPII on active genes but appear to be genuine subnuclear compartments. Consistent with this model, inhibition with drugs which freeze or slow the elongating polymerase (Palstra et al. 2008), did not lead to genome reorganization. HOW MANY FACTORIES ARE THERE?

The number of transcription sites per cell nucleus has been determined in a number of ways in many different cell and tissue types. The earliest methods used nascent transcript labeling in commonly used cultured cell lines such as HeLa and other fibroblastic cells and determined that there were 100 – 500 sites of nascent RNA per nucleus (Jackson et al. 1993; Wansink et al. 1993). In those days, the most reliable estimates put the number of eukaryotic genes at over 100,000 with the number of expressed or active genes in a given cell type around 10,000– 30,000, far greater than the number of RNA synthesis sites. Wansink et al. (1993) suggested two possible reasons to explain this discrepancy. Because few eukaryotic genes are transcribed at relatively high rates and most are transcribed at low rates, the small number of nascent RNA sites could be caused by detection of only the most highly transcribed genes, and a failure to detect the bulk of active genes making few nascent transcripts. In this scenario there would be no evidence in favor of a factory model because each gene could be transcribing in isolation. Another possibility, particularly favored by Jackson and Cook was that each transcription site was occupied by a number of actively transcribed genes. Over the

subsequent decade, refinements in nascent transcript labeling and detection have led to improved estimates of the number of factories per nucleus (Martin and Pombo 2003). Approximately 2100 (Iborra et al. 1996a) and 2400 (Jackson et al. 1998) nascent transcript sites were detected respectively in HeLa nuclei and an equivalent number of RNAPII sites were observed (Iborra et al. 1996a). Fay et al. (1997) had similar results but noted considerable variations in the number of nascent transcript sites (849 – 3888) in individual human fibroblasts. Pombo et al. (1999) using cryosectioning found evidence for 10,000 nucleoplasmic factories in HeLa cells with separate factories for RNAPII and RNAPIII; approximately 8000 RNAPII and 2000 RNAPIII factories. As the number of transcription sites increased, estimates of the number of eukaryotic genes in the genome and the number of active genes in a particular cell type decreased. However, the new appreciation of the extent of nongenic noncoding transcription has again increased the total number of transcription units and may now in fact exceed even the early estimates of gene numbers. So we are left with the realization that the number of active transcription units greatly outweighs the number of transcription sites per cell nucleus even with the highest estimates of factory numbers from HeLa cells. Some of the reports detailing the number of factories per nucleus make claims of highly sensitive techniques enabling the detection of all nascent transcription sites. Part of the variation in factory numbers observed may be because of differences in the ability to detect weak nascent RNA sites, but it also seems that there may be substantial variation within a cell type and between different cell types. In many cases HeLa or fibroblastic cells, which flatten out in culture were used, resulting in a substantial increase in nuclear diameter and nuclear volume when compared to cells with spherical nuclei. Osborne et al. (2004) used immunofluorescence to detect nuclear sites with high concentrations of the active form of RNAPII phosphorylated on Serine 5 (Ser5-RNAPII). They found approximately 2000 Ser5-RNAPII sites in the extended and flattened nuclei of

L. Chakalova and P. Fraser

mouse embryo fibroblast, consistent with previous nascent transcript labeling studies in fibroblastic cells (Fig. 2). In contrast, they found that erythroblasts, B-cells, T-cells, and fetal brain cells, which have spherical nuclei with significantly smaller radii and nuclear volumes, have dramatically fewer Ser5-RNAPII sites (100 – 300 per nucleus). Though the claim of detecting all RNAPII transcription sites was not made, the RNAPII detection threshold used by two different groups was sufficient to show that 90 – 99% of nascent RNA FISH signals for a variety of different genes overlapped with RNAPII factories (Osborne et al. 2004; Ragoczy et al. 2006; Osborne et al. 2007; Schoenfelder et al. 2010). Thus, although some RNAPII sites may have been missed because of the RNAPII detection threshold, the fact that nearly all nascent gene transcript signals colocalize with the RNAPII sites detected suggest that the majority of sites were detected. These results support the concept that all gene transcription occurs in RNAPII factories, consistent with conclusions from nascent transcript labeling studies. The large differences in factory numbers seen in nuclei from tissues versus cells grown on a surface appear genuine and may be a consequence of a reduced potential for intrachromosomal and especially interchromosomal sharing of factories in flattened cell nuclei. More factories may be required as an adaptation to service the same number of genes in flattened nuclei.

10 um

Figure 2. Maximum intensity projections of Ser5-

RNAPII factories in splenic B cell (left) and primary mouse embryo fibroblast (right) nuclei. From Osborne et al., 2004.

Interestingly, changes in cell and nuclear morphology have been demonstrated to result in widespread changes in gene expression (Dalby et al. 2007; Chang and Hughes-Fulford 2009). HIGHER-ORDER CHROMATIN FOLDING, TRANSCRIPTION AND SHARED RNAPII SITES

Since the discovery of transcriptional enhancers, which function to increase gene expression from remote genomic positions and independent of orientation to gene promoters, debate has centered on their mechanism. Most now agree that long-range gene control by remote enhancers and locus control regions involves direct interaction between chromatin at the enhancer sites and gene local regulatory elements (Carter et al. 2002; Tolhuis et al. 2002). It appears that DNA binding factors, transcription factors, and associated factors are required to either create an appropriate chromatin structure for distal elements to engage in interactions or mediate interactions by acting as bridging molecules (Drissen et al. 2004; Vakoc et al. 2005; Kurukuti et al. 2006; Splinter et al. 2006; Song et al. 2007; Majumder et al. 2008; Hadjur et al. 2009). Though we are still in the dark as to what specific transcriptional advantage such folding endows to a particular gene locus, there is little doubt that higher-order chromatin folding plays a critical role in transcriptional regulation over genomic distances of up to a megabase or more (Chakalova et al. 2005a; Kleinjan and van Heyningen 2005). One line of reasoning suggests that the increased local concentration of transcription factors brought into proximity of the promoter by the distal enhancer results in increased recruitment of (or to) the transcriptional machinery. A logical extension then leads to the question of what effect associations or clustering of distal actively transcribed genes and their complexed enhancers at transcription factories would have? A critical tenet of the transcription factory model is that each site has the potential to be occupied by a number of actively transcribed genes or transcription units. Cook (2002) proposed that transcription factories might form

Organization of Transcription

by the aggregation of a local cluster of transcriptionally active genes and their associated polymerases. Logic suggested that active genes in proximity in the primary DNA sequence would be more likely to engage in the same factory than active genes separated by long stretches of inactive genomic sequence. This may be true, but the original demonstration that actively transcribed genes could indeed share the same factory greatly exceeded this expectation. Osborne et al. (2004) showed using primary transcript RNA FISH and immunofluorescence (RNA FISH) for Ser5-RNAPII that genes separated by 25 – 40 megabases of chromosomal DNA could share factories at remarkably high frequencies in mouse erythroid progenitors. Chromosome conformation capture (3C) (Dekker et al. 2002) was used to verify the spatial association between distal transcribed sequences. It was also apparent that active genes on separate chromosomes could share factories but appeared to do so at reduced frequency compared to linked transcribed genes. These results, coupled with a realization of the transcriptional behavior of “active” genes in vivo lead to some important conclusions. Several lines of evidence suggest that transcription of “active” genes in a particular cell or tissue type is not continuous. Genes appear to be transcribed in bursts or pulses of transcriptional activity separated by variable periods of inactivity (Chubb et al. 2006; Raj et al. 2006). Similarly, RNA FISH for most “active” genes results in signals for only a portion of alleles across a population of expressing cell, with individual cells displaying none, one, or two actively transcribed alleles (Wijgerde et al. 1995; Levsky et al. 2002; Osborne et al. 2004; Osborne et al. 2007), There are a handful of so called “super genes,” which appear to defy this general rule (Fraser 2006). For example, the a- and b-globin genes (Hba and Hbb) in erythroid cells (Wijgerde et al. 1995) and the immunoglobulin genes (Igh, Igk, and Igl) in B-cells (Bolland et al. 2004; Osborne et al. 2007) tend to be continually or constitutively active with nearly all alleles in an interphase population associated with transcription factories (Osborne et al. 2004; Osborne et al. 2007). For other “expressed

genes,” which appear to be less frequently transcribed, inactive alleles were located away from RNAPII factories. Thus the high degree of colocalization between transcribed genes suggested that upon activation a significantly high proportion of newly transcribed genes join preestablished transcription sites containing other active genes rather than assemble their own transcription site de novo. This was further investigated by studying the dynamics of the immediate early genes Fos and Myc in resting splenic mouse B-cells (Osborne et al. 2007). Most alleles for these genes are transcriptionally inactive and located away from factories in resting cells. However, in cells induced for 5 min, the opposite is found, most Fos and Myc alleles are associated with factories and transcriptionally active. A high percentage of the newly active Fos alleles associated with the same factory as the constitutively transcribed Igh locus located approximately 30 Mb away on chromosome 12. Most interestingly, 25% of the newly active Myc alleles from chromosome 15 were also found to be associated with the same factory as the Igh locus. This appeared to be a highly preferential interchromosomal association because RNA FISH analyses of many other genes in trans had significantly lower interchromosomal association frequencies with Igh. Previous studies had shown that chromosomes 12 and 15 are preferred neighbors in mouse B-cells (Roix et al. 2003; Parada et al. 2004) and that a significant degree of intermingling occurs between chromosome territories (Branco and Pombo 2006). That MYC and IGH come into proximity in B-cells is without doubt, because the two genes are the most common translocation partners in Burkitt’s Lymphoma in humans and plasmacytomas in mouse (Hecht and Aster 2000; Potter 2003). Osborne’s work showed that induction of MYC correlated with a measurable shift in position of MYC alleles towards IGH alleles, indicating that activation of transcription involves short-range chromatin movements over distances of 0.5 – 1.5 microns to access a factory, and suggesting that preferential interchromosomal transcription factory coassociations could be involved in the mechanism of chromosomal translocations. Others

L. Chakalova and P. Fraser

have also documented long-range intra- and interchromosomal interactions between loci or coassociations with specific nuclear subcompartments (Spilianakis et al. 2005; Bacher et al. 2006; Brown et al. 2006; Ling et al. 2006; Lomvardas et al. 2006; Simonis et al. 2006; Wurtele and Chartrand 2006; Xu et al. 2006; Zhao et al. 2006; Apostolou and Thanos 2008; Brown et al. 2008; Hu et al. 2008) and in many cases correlated clustering or proximity with effects on gene expression. This remains a very controversial area, and even those who promote such concepts, in many cases cannot agree on how, where and to what effect proximity in nuclear space has a functional role.

CHROMATIN DYNAMICS AND TRANSCRIPTION

Constrained Brownian motion of chromatin could in theory account for the rapid, shortrange chromatin movement (Chakalova et al. 2005a) of induced Myc alleles toward the Igh factory mentioned earlier. The Myc gene in resting B-cells is not transcriptionally naı¨ve. Myc expression is involved in stimulating proliferation of immature B-cells and would have been induced at various points in B-cell differentiation concurrent with Igh transcription. However, that is not to say that active or directed processes are not involved in reorganization of chromatin and transcription in vivo. More dramatic chromatin movements may be involved, and have been suggested in the initial activation of a locus from a completely silent state. Ground-breaking live cell studies (Chuang et al. 2006) reported dramatic vectorial movements of chromatin in response to targeting a transcriptional activator to a silent transgene array. Migration occurred 1 –2 h after targeting at speeds of up to 0.9 microns/min over distances of 1– 5 microns. Rapid vectorial movements appeared to be punctuated by brief periods of randomdiffusionalmotion.Chuang etal.(2006) showed that repositioning did not require transcription and did not appear to involve extensive decondensation of the transgene array structure. Actin and nuclear myosin were

required for directed movements and the analysis of an actin point mutant defective in actin polymerization suggested that filamentous, Factin may be involved. Dundr et al. (2007) also showed actin-dependent, long-range repositioning using a tagged array of inducible U2 snRNA genes. Again, hours rather than minutes after induction, they observed vectorial movement of the transgene array toward relatively stably positioned Cajal bodies that are normally found in association with snRNAs and histone gene loci. Expression of a dominant negative mutant of b-actin markedly inhibited repositioning, supporting a role for nuclear actin in long-range chromatin movements. Large-scale chromatin movement has not been seen in every instance of transcriptional activation in live cells. Kumaran and Spector (Kumaran and Spector 2008) targeted a transgene array to the peripheral nuclear lamina. Transcriptional induction resulted in activation at the periphery with a clear increase in local RNAPII concentration. It was not clear whether the transgenes used local factories, which increased in size to accommodate the 200-copy array, or whether RNAPII was recruited to each promoter in the array de novo. However long-range movements were not observed. Similarly, Yao et al. (Yao et al. 2007) observed recruitment of polymerase components to heat shock loci in live cells on Drosophila polytene chromosomes creating characteristic puffs. Apart from local swelling, movement of the heat shock loci on the giant polytene chromosomes was not observed upon activation of transcription. Clearly live cell imaging holds the most potential for understanding the dynamic relationships between genes, transcriptional components and transcription factories. The studies of Kumaran and Spector (2008) and Yao et al. (2007) are probably the best available evidence in favor of the textbook model of transcription, in which RNAPII is recruited to gene promoters. However, aside from their obvious use, it remains to be seen whether the hundreds of gene copies in transgene arrays or giant polytene chromosomes are good models that accurately reflect the behavior of an isolated single copy gene in a diploid nucleus. It

Organization of Transcription

is possible that these localized multi-gene assemblies are in themselves super factories that alter the dynamic exchange of polymerase components (Kimura et al. 2002) to create a steady state structure that is several times larger than the average transcription factory.

TRANSCRIPTIONAL INTERACTOME AND TRANSCRIPTION HOTSPOTS

The number of transcription units sharing a factory at any one time has been estimated to be between two and 30 in HeLa cell nuclei (Jackson et al. 1998; Pombo et al. 1999). Mouse erythroid precursors, which have a few hundred factories, approximately 13,000 expressed alleles (6500 active genes assuming that most are bi-allelically expressed) with approximately half actively transcribing at any given moment are estimated to hold about 10– 30 genes per factory (Schoenfelder et al. 2010). Obviously the large amount of nongenic transcription must be factored into this calculation, but most studies suggest that a substantial fraction of these transcripts occur in the vicinity of active genes (Kapranov et al. 2007) and in the case of the globin genes are found in a relatively small fraction of the erythroid cells concomitant with gene transcription (Ashe et al. 1997; Gribnau et al. 2000; Chakalova et al. 2005b; Miles et al. 2007). The observation that preferential interchromosomal gene associations at factories can occur raises the possibility that the combination of genes at a particular factory could be meaningful in terms of transcriptional output of the assembled genes. Recent studies examining the intranuclear localization of transcriptionally active mini-chromosomes suggested that similarly regulated genes cluster at a limited number of factories. Xu and Cook (2008) used expression constructs carrying transcription units driven by different promoters. The multicopy constructs were assembled into nucleosomes in cells and the resulting mini-chromosomes underwent rounds of transcription and replication. RNA immuno-FISH experiments showed that transcriptionally active copies of mini-chromosomes clustered at a subset of the available

transcription factories. Moreover, the minichromosome appeared to share transcription sites with endogenous genes suggesting the introduced constructs were nonrandomly accommodated by host sites. The authors went on to cotransfect pairs of different constructs to test whether plasmids containing functionally similar genes and regulatory elements would share nuclear factories more often than functionally unrelated transcription units. Firstly, mini-chromosomes carrying RNAPI, II, or III transcription units separated into different factories, specialized in RNAPI, II, or III transcription, respectively. RNAPI genes acquired nucleolar localization, whereas RNAPII and III plasmids were spread in nonoverlapping foci throughout the nucleoplasm. These results were in agreement with earlier findings showing separate RNAPII and RNAPIII nucleoplasmic transcription sites (Pombo et al. 1999), and supported the view that the mini-chromosomes mimicked the behavior of endogenous chromosomal loci. Secondly, evidence for specialization was found within the RNAPII factory subclass. Minichromosomes carrying different genes driven by identical RNAPII promoters co-occupied the same foci at high frequencies. Conversely, plasmids carrying different promoters were mostly transcribed in separate factories. These preferences extended to endogenous genes too, because an edogenous gene was more likely to be in close proximity to its plasmid-borne counterpart than an unrelated gene. Taken together, these results suggested that factories are not functionally equal, and may be specialized to transcribe similarly regulated genes. However, important questions remain: Does the tendency for factory preference or specialization result in a non-random organization of native or chromosomally integrated transcription units? Schoenfelder et al. (2010) answered these questions in two ways. First they observed the nuclear localization of ectopically integrated, transcriptionally active human HBB transgenes in erythroid cells from several different lines of transgenic mice. They found that the HBB transgene locus had a three- to 15-fold preference to be transcribed in the same factory as the endogenous mouse Hbb genes compared

L. Chakalova and P. Fraser

to the endogenous Hba locus. The fact that the ectopically integrated transgene in all six lines tested showed preferential interchromosomal associations, in one case occurring in up to 35% of cells, suggested that preferential factory co-associations had the potential to reorganize the genome in the nucleus. They next developed an enhanced 4C assay to screen the entire genome for genes sharing transcription factories with the Hbb or Hba genes. Supported by extensive FISH and 3C analyses they revealed two overlapping transcription networks, each involving hundreds of preferred transcription partners in cis and trans. Genes regulated by the erythroid-specific transcription factor, Klf1 (erythroid Kruppel-like factor) were overrepresented in the globin transcription networks suggesting that coregulated genes preferentially transcribed in shared factories. Klf1 was previously shown to be required for high-level transcription of the Hbb gene in definitive erythroid cells (Nuez et al. 1995; Perkins et al. 1995) and also appears to contribute to Hba regulation (Shyu et al. 2006; Vernimmen et al. 2007). Schoenfelder et al. (2010) used immunofluorescence to show that, Klf1 localized to 30-40 small foci in erythroid nuclei and that nearly all overlapped with Ser5-RNAPII foci. These results suggested the possibility that a subset of factories were specialized because of an increased concentration of a particular transcription factor; a phenomena seen before for other factors (Grande et al. 1997). Actively transcribed alleles of Klf1-regulated genes were more often found in association with factories containing high levels of Klf1 suggesting that they had a higher probability of transcription when associated with a specialized factory. Furthermore, they found that many Klf1-regulated genes were preferentially clustered at these sites. This data provided strong evidence for functional specialization of transcription factories, and suggested that individual factories could become hotspots for optimal transcription of a subset of coregulated genes. There seemed to be no specific requirement for the globin genes to transcribe in the same factory with any other particular gene, but that in most, if not all, erythroid cells the globin genes associated with a varied

subset of other Klf1 regulated genes (Schoenfelder et al. 2010).

PULLING SOME STRINGS

The emerging evidence provides strong support for the transcription factory model where distant genes in cis and trans nonrandomly associate during transcription. A key feature of this model as originally proposed by Cook is that transcribed genes are reeled through factories by the relatively immobilized polymerase while extruding nascent RNA, rather than the polymerase tracking along the template. Cook and colleagues recently tested this prediction by assessing nuclear proximity of two distal, rapidly induced genes, one long and one short. SAMD4A is 221 kb long and located approximately 50 Mb from TNFAIP2—a relatively short gene of 11 kb. Both genes are activated simultaneously by tumor necrosis factor a(TNFa). Induction of SAMD4A has been shown to result in a fairly synchronous wave of transcription and RNAPII traversing the transcription unit over a 70-min period (Wada et al. 2009). The 3C assay was used to investigate long-range associations over this time period between the short, repeatedly transcribed TNFAIP2 gene and several regions along the length of SAMD4A. Before induction there was no evidence for long-range association between these genes, but upon activation they found that proximity between TNFAIP2 and the SAMD4A promoter was rapidly induced. As the wave of RNAPII moved along SAMD4A, contacts between the promoter and TNFAIP2 were lost in favor of new contacts between TNFAIP2 and successive downstream regions of SAMD4A. In other words, it appeared that the long gene was sliding past the distal short gene with a temporal pattern that matched the progress of the transcribing polymerase on the long gene. These results showing that different parts of transcription units are brought into proximity are consistent with the idea that transcribed genes are reeled through factories and very difficult to reconcile with the textbook model of RNAPII sliding along a relatively immobile template.

Organization of Transcription

CONCLUDING REMARKS

The realization that the active form of RNAPII is compartmentalized in the nucleus into a finite number of discrete transcription factories and that actively transcribed genes are non-randomly organized around these sites has widespread implications in understanding nuclear organization of the genome and genome function. This turns the classical view of gene transcription on its head, and with this new understanding come new insights and new questions. Rather than the requirement for near simultaneous recruitment and assembly of hundreds of factors required for regulated activation of transcription at the site of individual genes, this model suggests that many transcriptional components are preassembled into active sites which genes migrate to for transcription. There is still of course ample scope for regulatory factors and transcription components being recruited to genes while outside factories. For example, transcription factors, cofactors, chromatin remodeling complexes, and various histone modifying activities may act as licensing factors that affect genes before factory entry to prepare them for productive engagements with an RNAPII complex. The hypophosphorylated or initial binding form of RNAPII may also engage genes outside factories. The chance or directed coassociation of coregulated genes at a factory may favor re-initiation of those genes through sharing of dynamically binding or interacting factors, which will in turn stabilize the presence of those genes at the factory for as long as it takes to transcribe them. This may then become a favorable place for other coregulated genes to alight leading to a dynamically self-organizing hotspot of transcription for a specific subgroup or network of genes. Within the factory, epigenetic modifications may also be laid down or re-affirmed in transcribed regions, which may further promote re-initiation and/or serve as a memory for the activation state. Obviously chromatin mobility is a potentially important component of this new understanding. Factors or activities that compact chromatin or tether them to “repressive” nuclear domains may serve to restrict mobility

and access to transcription factories. Genes or alleles not able to reach one of their hotspots may disengage after relative few rounds of transcription at a generic factory because of a relatively low local concentration of specific factors required for re-initiation and stabilization at a factory. There is still much to do and many of the important questions will best be answered by live cell experiments on single copy or endogenous genes. For example, the transcriptional period of individual genes needs to be assessed. Are factory encounters transient and highly dynamic, potentially reshuffling at every transcriptional cycle, or once established, do active alleles/genes remain associated with factories throughout a cell cycle. It is likely the answers will vary widely for different genes and may encompass the entire range, potentially influenced by the character of long-range regulatory elements that individual active genes are complexed with as well as other transcription units they may encounter in factories. Also, how does the genome end up in such highly-organized, tissue-specific conformations that allow specific subgroups of genes the opportunity to cluster at specialized factories? There is controversy surrounding evidence suggesting chromosome positions may be partially inherited through mitosis (Gerlich et al. 2003; Walter et al. 2003; Cvackova et al. 2009), and that in very early G1 phase genome reorganization is more dramatic than in the rest of interphase (Dimitrova and Gilbert 1999). It appears likely that mitosis offers an essential opportunity for large-scale, though imperfect, genome reorganization which may be further refined in early G1. If the organized associations of co-regulated genes are beneficial then we may expect considerable pressure to be exerted on the primary organization of the genome, such that individual genes are optimally placed to take advantage of their eventual folding in 3 dimensions in the various tissues in which they will be expressed. These are exciting times and the advent of new technologies coupled with deep sequencing along with advanced microscopy hold great promise for further exciting advances in understanding the relationships between genome

L. Chakalova and P. Fraser

organization and transcriptional control as well as other genome functions. And hopefully, future textbooks will reflect these important aspects of the transcriptional mechanism so that the next generation of genome investigators will be more adequately prepared to advance the understanding of transcription inside the complex world of the nucleus.

ACKNOWLEDGMENTS

We would like to thank in particular C. Eskiw and other members of the Laboratory of Chromatin and Gene Expression for many helpful discussions. We are also grateful to A. Papantonis and P. Cook for generously sharing unpublished data.

REFERENCES Allen TA, Von Kaenel S, Goodrich JA, Kugel JF. 2004. The SINE-encoded mouse B2 RNA represses mRNA transcription in response to heat shock. Nat Struct Mol Biol 11: 816– 821. Apostolou E, Thanos D. 2008. Virus Infection Induces NF-kB-dependent interchromosomal associations mediating monoallelic IFN-b gene expression. Cell 134: 85–96. Ashe HL, Monks J, Wijgerde M, Fraser P, Proudfoot NJ. 1997. Intergenic transcription and transinduction of the human b-globin locus. Genes Dev 11: 2494– 2509. Bacher CP, Guggiari M, Brors B, Augui S, Clerc P, Avner P, Eils R, Heard E. 2006. Transient colocalization of X-inactivation centres accompanies the initiation of X inactivation. Nat Cell Biol 8: 293 –299. Berezney R, Dubey DD, Huberman JA. 2000. Heterogeneity of eukaryotic replicons, replicon clusters, and replication foci. Chromosoma 108: 471– 484. Bolland DJ, Wood AL, Johnston CM, Bunting SF, Morgan G, Chakalova L, Fraser PJ, Corcoran AE. 2004. Antisense intergenic transcription in V(D)J recombination. Nat Immunol 5: 630– 637. Branco MR, Pombo A. 2006. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biology 4: e138. Brown JM, Green J, das Neves RP, Wallace HA, Smith AJ, Hughes J, Gray N, Taylor S, Wood WG, Higgs DR, et al. 2008. Association between active genes occurs at nuclear speckles and is modulated by chromatin environment. J Cell Biol 182: 1083– 1097. Brown JM, Leach J, Reittie JE, Atzberger A, Lee-Prudhoe J, Wood WG, Higgs DR, Iborra FJ, Buckle VJ. 2006. Coregulated human globin genes are frequently in spatial proximity when active. J Cell Biol 172: 177– 187.

Carter D, Chakalova L, Osborne CS, Dai YF, Fraser P. 2002. Long-range chromatin regulatory interactions in vivo. Nat Genet 32: 623 –626. Chakalova L, Debrand E, Mitchell JA, Osborne CS, Fraser P. 2005a. Replication and transcription: Shaping the landscape of the genome. Nat Rev Genet 6: 669–677. Chakalova L, Osborne CS, Dai YF, Goyenechea B, Metaxotou-Mavromati A, Kattamis A, Kattamis C, Fraser P. 2005b. The Corfu db thalassemia deletion disrupts g-globin gene silencing and reveals post-transcriptional regulation of HbF expression. Blood 105: 2154–2160. Chang TT, Hughes-Fulford M. 2009. Monolayer and spheroid culture of human liver hepatocellular carcinoma cell line cells demonstrate distinct global gene expression patterns and functional phenotypes. Tissue Eng Part A 15: 559 –567. Chuang CH, Carpenter AE, Fuchsova B, Johnson T, de Lanerolle P, Belmont AS. 2006. Long-range directional movement of an interphase chromosome site. Curr Biol 16: 825– 831. Chubb JR, Trcek T, Shenoy SM, Singer RH. 2006. Transcriptional pulsing of a developmental gene. Curr Biol 16: 1018–1025. Cmarko D, Verschure PJ, Martin TE, Dahmus ME, Krause S, Fu XD, van Driel R, Fakan S. 1999. Ultrastructural analysis of transcription and splicing in the cell nucleus after bromo-UTP microinjection. Mol Biol Cell 10: 211 –223. Cook PR. 2002. Predicting three-dimensional genome structure from transcriptional activity. Nat Genet 32: 347 –352. Cremer T, Kupper K, Dietzel S, Fakan S. 2004. Higher order chromatin architecture in the cell nucleus: On the way from structure to function. Biol Cell 96: 555– 567. Cvackova´ Z, Masata M, Stane˘k D, Fidlerova´ H, Raska I. 2009. Chromatin position in human HepG2 cells: Although being non-random, significantly changed in daughter cells. J Struct Biol. 165:107–117. Dalby MJ, Gadegaard N, Herzyk P, Sutherland D, Agheli H, Wilkinson CD, Curtis AS. 2007. Nanomechanotransduction and interphase nuclear organization influence on genomic control. J Cell Biochem 102: 1234–1244. Davenport RJ, Wuite GJ, Landick R, Bustamante C. 2000. Single-molecule study of transcriptional pausing and arrest by E. coli RNA polymerase. Science 287: 2497– 2500. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306– 1311. Dimitrova DS, Gilbert DM. 1999. The spatial position and replication timing of chromosomal domains are both established in early G1 phase. Mol Cell 4: 983– 993. Drissen R, Palstra RJ, Gillemans N, Splinter E, Grosveld F, Philipsen S, de Laat W. 2004. The active spatial organization of the b-globin locus requires the transcription factor EKLF. Genes Dev 18: 2485– 2490. Dundr M, Ospina JK, Sung MH, John S, Upender M, Ried T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095–1103. Eskiw CH, Rapp A, Carter DR, Cook PR. 2008. RNA polymerase II activity is located on the surface of protein-rich transcription factories. J Cell Sci 121: 1999–2007.

Organization of Transcription Espinoza CA, Allen TA, Hieb AR, Kugel JF, Goodrich JA. 2004. B2 RNA binds directly to RNA polymerase II to repress transcript synthesis. Nat Struct Mol Biol 11: 822–829. Fakan S. 1994. Perichromatin fibrils are in situ forms of nascent transcripts. Trends Cell Biol 4: 86–90. Fakan S. 2004. The functional architecture of the nucleus as analysed by ultrastructural cytochemistry. Histochem Cell Biol 122: 83–93. Fakan S, Bernhard W. 1971. Localisation of rapidly and slowly labelled nuclear RNA as visualized by high resolution autoradiography. Exp Cell Res 67: 129–141. Fakan S, Puvion E, Sphor G. 1976. Localization and characterization of newly synthesized nuclear RNA in isolate rat hepatocytes. Exp Cell Res 99: 155 –164. Fay FS, Taneja KL, Shenoy S, Lifshitz L, Singer RH. 1997. Quantitative digital analysis of diffuse and concentrated nuclear distributions of nascent transcripts, SC35 and poly(A). Exp Cell Res 231: 27– 37. Fraser P. 2006. Transcriptional control thrown for a loop. Curr Opin Genet Dev 16: 490 –495. Gerlich D, Beaudouin J, Kalbfuss B, Daigle N, Eils R, Ellenberg J. 2003. Global chromosome positions are transmitted through mitosis in mammalian cells. Cell 112: 751–764. Grande MA, van der Kraan I, de Jong L, van Driel R. 1997. Nuclear distribution of transcription factors in relation to sites of transcription and RNA polymerase II. J Cell Sci 110: 1781–1791. Gribnau J, Diderich K, Pruzina S, Calzolari R, Fraser P. 2000. Intergenic transcription and developmental remodeling of chromatin subdomains in the human b-globin locus. Mol Cell 5: 377–386. Guthold M, Zhu X, Rivetti C, Yang G, Thomson NH, Kasas S, Hansma HG, Smith B, Hansma PK, Bustamante C. 1999. Direct observation of one-dimensional diffusion and transcription by Escherichia coli RNA polymerase. Biophys J 77: 2284– 2294. Hadjur S, Williams LM, Ryan NK, Cobb BS, Sexton T, Fraser P, Fisher AG, Merkenschlager M. 2009. Cohesins form chromosomal cis-interactions at the developmentally regulated IFNG locus. Nature 460: 410–413. Harada Y, Funatsu T, Murakami K, Nonoyama Y, Ishihama A, Yanagida T. 1999. Single-molecule imaging of RNA polymerase-DNA interactions in real time. Biophys J 76: 709–715. Hecht JL, Aster JC. 2000. Molecular biology of Burkitt’s lymphoma. J Clin Oncol 18: 3707–3721. Hieda M, Winstanley H, Maini P, Iborra FJ, Cook PR. 2005. Different populations of RNA polymerase II in living mammalian cells. Chromosome Res 13: 135–144. Hu Q, Kwon YS, Nunez E, Cardamone MD, Hutt KR, Ohgi KA, Garcia-Bassets I, Rose DW, Glass CK, Rosenfeld MG, et al. 2008. Enhancing nuclear receptor-induced transcription requires nuclear motor and LSD1-dependent gene networking in interchromatin granules. Proc Natl Acad Sci 105: 19199– 19204. Huang S, Spector DL. 1996. Dynamic organization of premRNA splicing factors. J Cell Biochem 62: 191 –197. Iborra FJ, Pombo A, Jackson DA, Cook PR. 1996a. Active RNA polymerases are localized within discrete

transcription “factories” in human nuclei. J Cell Sci 109: 1427– 1436. Iborra FJ, Pombo A, McManus J, Jackson DA, Cook PR. 1996b. The topology of transcription by immobilized polymerases. Exp Cell Res 229: 167–173. Jackson DA, Cook PR. 1985. Transcription occurs at a nucleoskeleton. EMBO J 4: 919– 925. Jackson DA, McCready SJ, Cook PR. 1981. RNA is synthesized at the nuclear cage. Nature 292: 552– 555. Jackson DA, Hassan AB, Errington RJ, Cook PR. 1993. Visualization of focal sites of transcription within human nuclei. Embo J 12: 1059– 1065. Jackson DA, Iborra FJ, Manders EM, Cook PR. 1998. Numbers and organization of RNA polymerases, nascent transcripts, and transcription units in HeLa nuclei. Mol Biol Cell 9: 1523–1536. Kabata H, Kurosawa O, Arai I, Washizu M, Margarson SA, Glass RE, Shimamoto N. 1993. Visualization of single molecules of RNA polymerase sliding along DNA. Science 262: 1561– 1563. Kapranov P, Willingham AT, Gingeras TR. 2007. Genomewide transcription and the implications for genomic organization. Nat Rev Genet 8: 413– 423. Kimura H, Sugaya K, Cook PR. 2002. The transcription cycle of RNA polymerase II in living cells. J Cell Biol 159: 777–782. Kleinjan DA, van Heyningen V. 2005. Long-range control of gene expression: Emerging mechanisms and disruption in disease. Am J Hum Genet 76: 8– 32. Kumaran RI, Spector DL. 2008. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol 180: 51–65. Kurukuti S, Tiwari VK, Tavoosidana G, Pugacheva E, Murrell A, Zhao Z, Lobanenkov V, Reik W, Ohlsson R. 2006. CTCF binding at the H19 imprinting control region mediates maternally inherited higher-order chromatin conformation to restrict enhancer access to Igf2. Proc Natl Acad Sci 103: 10684–10689. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605– 612. Levsky JM, Shenoy SM, Pezo RC, Singer RH. 2002. Singlecell gene expression profiling. Science 297: 836– 840. Ling JQ, Li T, Hu JF, Vu TH, Chen HL, Qiu XW, Cherry AM, Hoffman AR. 2006. CTCF mediates interchromosomal colocalization between Igf2/H19 and Wsb1/Nf1. Science 312: 269–272. Lomvardas S, Barnea G, Pisapia DJ, Mendelsohn M, Kirkland J, Axel R. 2006. Interchromosomal interactions and olfactory receptor choice. Cell 126: 403 –413. Majumder P, Gomez JA, Chadwick BP, Boss JM. 2008. The insulator factor CTCF controls MHC class II gene expression and is required for the formation of long-distance chromatin interactions. J Exp Med 205: 785 –798. Mariner PD, Walters RD, Espinoza CA, Drullinger LF, Wagner SD, Kugel JF, Goodrich JA. 2008. Human Alu RNA is a modular transacting repressor of mRNA transcription during heat shock. Mol Cell 29: 499– 509. Martin S, Pombo A. 2003. Transcription factories: Quantitative studies of nanostructures in the mammalian nucleus. Chromosome Res 11: 461– 470.

L. Chakalova and P. Fraser Miles J, Mitchell JA, Chakalova L, Goyenechea B, Osborne CS, O’Neill L, Tanimoto K, Engel JD, Fraser P. 2007. Intergenic transcription, cell-cycle and the developmentally regulated epigenetic profile of the human b-globin locus. PLoS One 2: e630. Misteli T, Caceres JF, Spector DL. 1997. The dynamics of a pre-mRNA splicing factor in living cells. Nature 387: 523–527. Mitchell JA, Fraser P. 2008. Transcription factories are nuclear subcompartments that remain in the absence of transcription. Genes Dev 22: 20–25. Nash RE, Puvion E, Bernhard W. 1975. Perichromatin fibrils as components of rapidly labeled extranucleolar RNA. J Ultrastruct Res 53: 395 –405. Nuez B, Michalovich D, Bygrave A, Ploemacher R, Grosveld F. 1995. Defective haematopoiesis in fetal liver resulting from inactivation of the EKLF gene. Nature 375: 316– 318. Osborne CS, Chakalova L, Brown KE, Carter D, Horton A, Debrand E, Goyenechea B, Mitchell JA, Lopes S, Reik W, et al. 2004. Active genes dynamically colocalize to shared sites of ongoing transcription. Nat Genet 36: 1065– 1071. Osborne CS, Chakalova L, Mitchell JA, Horton A, Wood AL, Bolland DJ, Corcoran AE, Fraser P. 2007. Myc dynamically and preferentially relocates to a transcription factory occupied by igh. PLoS Biol 5: e192. Palstra RJ, Simonis M, Klous P, Brasset E, Eijkelkamp B, de Laat W. 2008. Maintenance of long-range DNA interactions after inhibition of ongoing RNA polymerase II transcription. PLoS One3: e1661. Parada LA, McQueen PG, Misteli T. 2004. Tissue-specific spatial organization of genomes. Genome Biol 5: R44. Perkins AC, Sharpe AH, Orkin SH. 1995. Lethal b-thalassaemia in mice lacking the erythroid CACCC-transcription factor EKLF. Nature 375: 318–322. Pombo A, Cook PR. 1996. The localization of sites containing nascent RNA and splicing factors. Exp Cell Res 229: 201–203. Pombo A, Jackson DA, Hollinshead M, Wang Z, Roeder RG, Cook PR. 1999. Regional specialization in human nuclei: Visualization of discrete sites of transcription by RNA polymerase III. EMBO J 18: 2241– 2253. Potter M. 2003. Neoplastic development in plasma cells. Immunol Rev 194: 177–195. Ragoczy T, Bender MA, Telling A, Byron R, Groudine M. 2006. The locus control region is required for association of the murine b-globin locus with engaged transcription factories during erythroid maturation. Genes Dev 20: 1447– 1457. Raj A, Peskin CS, Tranchina D, Vargas DY, Tyagi S. 2006. Stochastic mRNA synthesis in mammalian cells. PLoS Biol 4: e309. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287 –291. Schafer DA, Gelles J, Sheetz MP, Landick R. 1991. Transcription by single molecules of RNA polymerase observed by light microscopy. Nature 352: 444– 448. Schoenfelder S, Sexton T, Chakalova L, Cope NF, Horton A, Andrews S, Kurukuti S, Mitchell JA, Umlauf D, Dimitrova DS, et al. 2010. Preferential associations between

co-regulated genes reveal a transcriptional interactome in erythroid cells. Nat Genet 42:53– 61. Shyu YC, Wen SC, Lee TL, Chen X, Hsu CT, Chen H, Chen RL, Hwang JL, Shen CK. 2006. Chromatin-binding in vivo of the erythroid kruppel-like factor, EKLF, in the murine globin loci. Cell Res 16: 347– 355. Simonis M, Klous P, Splinter E, Moshkin Y, Willemsen R, de Wit E, van Steensel B, de Laat W. 2006. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nat Genet 38: 1348–1354. Song SH, Hou C, Dean A. 2007. A positive role for NLI/ Ldb1 in long-range b-globin locus control region function. Mol Cell 28: 810– 822. Spilianakis CG, Lalioti MD, Town T, Lee GR, Flavell RA. 2005. Interchromosomal associations between alternatively expressed loci. Nature 435: 637–645. Splinter E, Heath H, Kooren J, Palstra RJ, Klous P, Grosveld F, Galjart N, de Laat W. 2006. CTCF mediates long-range chromatin looping and local histone modification in the b-globin locus. Genes Dev 20: 2349–2354. Sugaya K, Vigneron M, Cook PR. 2000. Mammalian cell lines expressing functional RNA polymerase II tagged with the green fluorescent protein. J Cell Sci 113: 2679– 2683. Tolhuis B, Palstra RJ, Splinter E, Grosveld F, de Laat W. 2002. Looping and interaction between hypersensitive sites in the active b-globin locus. Mol Cell 10: 1453–1465. Vakoc CR, Letting DL, Gheldof N, Sawado T, Bender MA, Groudine M, Weiss MJ, Dekker J, Blobel GA. 2005. Proximity among distant regulatory elements at the b-globin locus requires GATA-1 and FOG-1. Mol Cell 17: 453– 462. Vernimmen D, De Gobbi M, Sloane-Stanley JA, Wood WG, Higgs DR. 2007. Long-range chromosomal interactions regulate the timing of the transition between poised and active gene expression. Embo J 26: 2041–2051. Wada Y, Ohta Y, Xu M, Tsutsumi S, Minami T, Inoue K, Komura D, Kitakami J, Oshida N, Papantonis A, et al. 2009. Awave of nascent transcription on activated human genes. Proc Natl Acad Sci 106:18357– 18361. Walter J, Schermelleh L, Cremer M, Tashiro S, Cremer T. 2003. Chromosome order in HeLa cells changes during mitosis and early G1, but is stably maintained during subsequent interphase stages. J Cell Biol 160:685– 697. Wang MD, Schnitzer MJ, Yin H, Landick R, Gelles J, Block SM. 1998. Force and velocity measured for single molecules of RNA polymerase. Science 282: 902 –907. Wansink DG, Schul W, van der Kraan I, van Steensel B, van Driel R, de Jong L. 1993. Fluorescent labeling of nascent RNA reveals transcription by RNA polymerase II in domains scattered throughout the nucleus. J Cell Biol 122: 283–293. Wijgerde M, Grosveld F, Fraser P. 1995. Transcription complex stability and chromatin dynamics in vivo. Nature 377: 209–213. Wurtele H, Chartrand P. 2006. Genome-wide scanning of HoxB1-associated loci in mouse ES cells using an openended Chromosome Conformation Capture methodology. Chromosome Res 14: 477 –495.

Organization of Transcription Xu M, Cook PR. 2008. Similar active genes cluster in specialized transcription factories. J Cell Biol 181: 615–623. Xu N, Tsai CL, Lee JT. 2006. Transient homologous chromosome pairing marks the onset of X inactivation. Science 311: 1149–1152. Yao J, Ardehali MB, Fecko CJ, Webb WW, Lis JT. 2007. Intranuclear distribution and local dynamics of RNA

polymerase II during transcription activation. Mol Cell 28: 978– 990. Zhao Z, Tavoosidana G, Sjolinder M, Gondor A, Mariano P, Wang S, Kanduri C, Lezcano M, Sandhu KS, Singh U, et al. 2006. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat Genet 38: 1341–1347.

Organization of DNA Replication Vadim O. Chagin1, Jeffrey H. Stear2, and M. Cristina Cardoso1 1

Department of Biology, Technische Universita¨t Darmstadt, 64287 Darmstadt, Germany

2

Institute for Biology, Humboldt University, 10115 Berlin, Germany

Correspondence: [email protected]

The discovery of the DNA double helix structure half a century ago immediately suggested a mechanism for its duplication by semi-conservative copying of the nucleotide sequence into two DNA daughter strands. Shortly after, a second fundamental step toward the elucidation of the mechanism of DNA replication was taken with the isolation of the first enzyme able to polymerize DNA from a template. In the subsequent years, the basic mechanism of DNA replication and its enzymatic machinery components were elucidated, mostly through genetic approaches and in vitro biochemistry. Most recently, the spatial and temporal organization of the DNA replication process in vivo within the context of chromatin and inside the intact cell are finally beginning to be elucidated. On the one hand, recent advances in genome-wide high throughput techniques are providing a new wave of information on the progression of genome replication at high spatial resolution. On the other hand, novel superresolution microscopy techniques are just starting to give us the first glimpses of how DNA replication is organized within the context of single intact cells with high spatial resolution. The integration of these data with time lapse microscopy analysis will give us the ability to film and dissect the replication of the genome in situ and in real time.

ollowing the elucidation of the DNA double helix structure by Watson and Crick (Watson and Crick 1953) and the isolation of the first DNA polymerase by Kornberg (Kornberg 1960), a fundamental biological question has been how genomes are duplicated prior to cell division. In the 50 years since these seminal discoveries, the basic mechanisms of DNA replication have been established by a powerful combination of genetics and in vitro biochemistry. From this body of work, it is clear that the fundamental features and components of DNA replication are well conserved throughout evolution from

F

bacteria to mammals. A comprehensive list of proteins that are involved in the process and their activities has been compiled, and it is now possible to provide a relatively detailed description of the DNA replication machinery on a molecular level (Fig. 1) (Perumal et al. 2009). However, genomic DNA in eukaryotic cells is hierarchically packed within the nucleus and genome duplication requires the concerted effort of many thousands of individual replication units. As such, an equally important question is how DNA replication is coordinated in space and time across the entire genome within the living cell.

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000737 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000737

V.O. Chagin, J.H. Stear, and M.C. Cardoso

DNA Polymerase

Clamp loader

Helicase

Topoisomerase

PCNA

DNA Ligase Primase

Endonucleases

SSB proteins RNAseH

Figure 1. Schematic outline of the DNA replication fork and molecular components and enzymatic activities of

the replisome. PCNA, proliferating cell nuclear antigen (also termed DNA polymerase clamp or processivity factor). SSB, single strand DNA binding proteins.

DNA REPLICATION: THE BASICS

Every cycling cell needs to duplicate its genome before it divides. This entails not only duplicating precisely and completely its genetic information, which is the focus of this article, but also restoring its epigenetic information to build up the differentiated chromatin structures. After cells divide, the so-called prereplication complexes including the DNA helicase complex (MCM 2-7) assemble onto the DNA throughout the G1 phase (Blow and Dutta 2005). At the end of G1, input from the cell cycle machinery via the cell-cycle dependent kinases triggers initiation of DNA replication at discrete sites, known as origins of replication, scattered throughout the genome. Each origin fires once per cell cycle and their spacing must ensure that the entire genome is replicated during the S phase. Early results from cell fusion experiments indicated that replicated DNA differed from unreplicated DNA because it was not permissive for replication unless it passed through one mitotic division (Rao and Johnson 1970). Subsequent experiments using cell-free replication assays further refined this idea and a model was proposed whereby replication origins were “licensed” to replicate at late mitosis and G1,

and the “license” was removed as the DNA was replicated in S phase (Blow and Laskey 1988). Detailed biochemical evidence is now available and indicates that preventing rebinding of the MCM complex to DNA is the key to avoid rereplication (Blow and Dutta 2005). In bacteria and lower eukaryotes, replication origins are defined by specific DNA sequences; whereas in metazoans the defining characteristics of these sites remain less clear (Robinson and Bell 2005; Aladjem 2007; Hamlin et al. 2008). At each origin, replication proteins are assembled, which then duplicate one segment (replicon) of the genome in a processive manner (Jacob and Brenner 1963; Jacob 1993). The active units of DNA replication, consisting of the replication machinery (also called replisome), the existing DNA template, and the nascent DNA strand, are referred to as replication forks. Following initiation of DNA synthesis, replication forks proceed bidirectionally from the origin, unwinding the genomic DNA as they traverse the chromosome (Huberman and Riggs 1968). At the molecular level, replication of a genomic DNA template involves at least two different DNA polymerases, a DNA primase, a DNA helicase, a single strand DNA binding (SSB) protein complex (replication protein A, RPA), DNA

DNA Replication

topoisomerases, a clamp loading complex (replication factor C, RFC), and a DNA polymerase clamp or processivity factor (proliferating cell nuclear antigen, PCNA). Furthermore, the inherent polarity of the DNA synthesis reaction (50 to 30 direction) necessitates discontinuous duplication of the lagging strand in addition to the continuous leading-strand synthesis. This process involves the synthesis of short (180–200 bp in eukaryotic cells) DNA fragments known as Okazaki fragments (Okazaki et al. 1968) that are subsequently processed and ligated together by a number of additional enzymes, including the flap endonucleases (FEN-1) and DNA ligase I (Hubscher and Seo 2001). To explain how the synthesis of both strands is coordinated, an asymmetric dimer of DNA polymerases and associated factors has been proposed first for prokaryotic DNA replication (McHenry 1988) and subsequently extended for eukaryotes (Tsurimoto and Stillman 1989). In order that the dimeric polymerases can extend the two antiparallel strands at the same rate and in the same direction, looping back of the lagging strand into the replisome (“trombone” model) has been postulated, with recycling of the lagging-strand polymerase from the end of one Okazaki fragment to the next RNA primer forming a priming loop (Sinha et al. 1980; Pandey et al. 2009). This situation suggests that the replication machinery at a given replication fork likely consists of at least two functional (sub)modules, one responsible for leadingstrand synthesis, and the other for lagging-strand synthesis (Fig. 1). Biochemical evidence points to the existence of large preformed multiprotein replication complexes that contain all activities (Noguchi et al. 1983; Tom et al. 1996). However, evidence from live-cell microscopy analysis points to short time interactions between the individual components, suggesting highly dynamic complexes (Sporbert et al. 2002; Sporbert et al. 2005; Schermelleh et al. 2007; Gorisch et al. 2008). DNA REPLICATION: IN VIVO

Investigating how the distinct activities required for DNA synthesis are organized within the cell nucleus relatesto thelarger issueofunderstanding

how DNA replication is regulated on a cellular level. A cell must duplicate its entire genome once and only once every time it divides. Therefore, in the broadest sense, investigating the global regulation of DNA replication can be distilled into the question of how the activity of individual replication units is coordinated throughout a cell cycle. Thisrather daunting problem canbe broken down into more specific questions. How is replication propagated along the chromosomes? How does the cell ensure that the whole genome gets replicated? Where along the chromosome does replication begin? Thesequestions were originally addressed in bacteria, where the replication program proceeds in a rather straightforward fashion with genetically well defined replication origins, which fire once per cell cycle (Mott and Berger 2007). However, the much larger size and complexity of eukaryotic genomes impose additional difficulties on the organization of DNA replication. Hence, eukaryotic DNA replication represents a more complex situation that remains poorly understood. The confusion centers around two seemingly contradictory observations. First, a number of studies have clearly shown that DNA replication follows a defined, temporal progression (Sparvoli et al. 1994; Cardoso et al. 1997; Jackson and Pombo 1998; Ma et al. 1998; Dimitrova and Gilbert 2000; Leonhardt et al. 2000; Sadoni et al. 2004; Easwaran et al. 2005). Actively transcribed, euchromatic regions of the genome tend to be duplicated early in S phase, whereas heterochromatin, which is more condensed and often transcriptionally silent, replicates late in S phase (Fig. 2 and Movie 1 online at http://cshperspectives.cshlp.org/). This phenomenon was originally described by observing replication along Giemsa stained chromosomes and correlating DNA synthesis with banding patterns (Drouin et al. 1990). Second, eukaryotic replication origins fire in a stochastic fashion throughout S phase (Dijkwel et al. 2002; Patel et al. 2006). Therefore, the distribution of active origins, and thus replication initiation, changes between each cell cycle. Given the random nature of replication origin firing, it is hard to understand how a cell can maintain the temporal progression of replication. Reconciling

V.O. Chagin, J.H. Stear, and M.C. Cardoso G1 03:15

SE 11:45

SE

SM 13:45

SM

SL 17:45

G2 20:00

M 21:45

SL

Figure 2. Temporal progression of genome replication. Snapshots of a time-lapse confocal microscopy movie of

DNA replication throughout the cell cycle in human HeLa cells stably expressing GFP-tagged PCNA. Times are indicated in hours:minutes and cell cycle phases as G1/SE (S early)/SM (S mid)/SL (S late)/G2/M. Scale bar, 5 mm. During early S phase, small foci distributed throughout the nucleus and mostly corresponding to euchromatic genomic regions are duplicated. Subsequently, the DNA replication machinery loads at perinuclear and perinucleolar heterochromatin regions followed at later times by large constitutive heterochromatic chromosomal regions. This temporal replication program is recapitulated at each cell cycle. See also Movie 1 online at http://cshperspectives.cshlp.org/.

these data acquired at very different spatial resolution levels (from hundreds of base pairs in stretched DNA fibers and by two-dimensional gel electrophoresis analysis to megabase chromatin domains in whole cells in situ) presents a significant hurdle to our understanding of how replication proceeds in eukaryotic cells. It should be noted that the budding yeast Saccharomyces cerevisiae represents an exception to the standard eukaryotic strategy for genome duplication. Similar to bacteria, S. cerevisiae possess well-defined replication origin sequences that can fire at a very efficient rate during S phase, leading to a very homogenous pattern of DNA replication (Fangman and Brewer 1991; Gilbert 2001). Furthermore, genome-wide analysis of replication initiation indicates no bias for sites of active transcription and no observable delay for any distinct regions of the genome (Raghuraman et al. 2001). For these reasons, it

is perhaps misleading to generalize conclusions obtained with this system when contemplating eukaryotic DNA replication timing. Therefore, this article focuses on how DNA replication is regulated within the nucleus of metazoan systems. GENOMIC APPROACHES TO STUDYING REPLICATION TIMING

The recent development of genomic assays has for the first time permitted a genome-wide examination of replication timing in populations of eukaryotic cells. For example, microarray analysis has been successfully used in a variety of systems to generate genome-wide profiles of replication timing. First performed in S. cerevisiae, and more recently in S. pombe, Drosophila melanogaster, and Homo sapien cells (Raghuraman et al. 2001; Schubeler et al. 2002;

DNA Replication

Watanabe et al. 2002; White et al. 2004; Woodfine et al. 2004; Jeon et al. 2005; Eshaghi et al. 2007; Hiratani et al. 2008; Watanabe et al. 2008), this technique usually requires the synchronization of a population of cells at the G1/S boundary or their flow cytometric sorting based on increasing DNA content through S phase. Following release into S phase, the accumulation of newly synthesized DNA over time is measured by hybridization to DNA arrays. Alternatively, isolated nascent DNA can be directly sequenced using novel high-throughput deep sequencing techniques. A summary of these studies is compiled in Table 1. DNA combing is another technique that has recently been used to systematically investigate replication initiation and elongation at the level of single DNA fibers. Long, individual DNA molecules are stripped of proteins, uniformly stretched across a glass surface, and examined by standard fluorescence microscopy (Bensimon et al. 1994). By pulse-labeling cells with nucleotides prior to this treatment, it is possible to directly examine sites of DNA synthesis (Pasero et al. 2002; Anglana et al. 2003). Originally used with radioactively labeled nucleotides to accurately measure the rate of replication fork progression (Huberman and Riggs 1968), more recent studies have used this approach to compare the efficiency of origin firing between early and late replicating regions (Patel et al. 2006). The application of these methods has, for the first time, permitted the examination of how DNA replication proceeds at the genomic level. The availability of several complete genome sequences has further allowed in silico evaluation of the correlation of DNA replication with structural and functional genomic features. For example, whereas a relationship between transcriptional activity and replication timing had been suggested by earlier reports analyzing single genes (Gilbert 2002), microarray studies investigating the timing of DNA replication along D. melanogasterand human chromosomes convincingly linked these processes on a genome-wide level (Schubeleret al. 2002; Woodfine et al. 2004; Jeon et al. 2005). Specifically, early replicating regions displayed a strong correlation with gene rich areas that possessed a high

GC content and contained actively transcribed genes. In fact, a high-resolution human genome analysis revealed that boundaries between regions with different GC content (the so-called isochores) correlated with borders between DNA replication timing zones (Costantini and Bernardi 2008). In addition, recent studies also showed that a substantial portion of the genome (approx. 60%) does not replicate until much later in S phase (Eshaghi et al. 2007). The dynamics of origin firing has also been examined by genome-wide analysis. Consistent with earlier reports, this process was found to be stochastic, leading to a random distribution of replication initiation sites across the chromosomes (Patel et al. 2006). It was also shown that early S phase origins are quite inefficient at initiating replication. In contrast, the relative firing efficiency of late S phase origins, although still occurring in a random manner, was significantly higher (Eshaghi et al. 2007). This could reflect the fact that less DNA is “licensed” to replicate at later S phase stages leading to a seemingly higher firing efficiency. Alternatively, assuming recycling of a factor limiting replication initiation—firing propensity redistribution—has also allowed reasonable modeling of S phase based on stochastic firing of individual origins (Lygeros et al. 2008). Cyclin-dependent kinases were suggested as factors limiting initiation. These factors are recycled to late-firing origins, thus increasing the probability of their activation. Together, these results have led to the “increasing efficiency model,” which has the potential to explain many of the questions surrounding the regulation of DNA replication. The idea centers around the hypothesis that as a cell progresses through S phase, the overall efficiency with which the remaining fraction of available origins initiates replication increases (Lucas et al. 2000; Hyrien et al. 2003). An appealing aspect of this model is that it accounts for how cells can avoid the problem of gaps in DNA replication. Furthermore, by assuming that different regions of the genome possess variable efficiencies of origin firing, it can also explain how a cell could maintain stochastic firing of replication origins and still replicate its DNA in an ordered fashion (Rhind 2006).

V.O. Chagin, J.H. Stear, and M.C. Cardoso Table 1. Genomic studies of DNA replication. Genome duplication characteristic

Species

Genome segment investigated

Reference

Timing of replication Chromosome bands/ isochores; GC/AT content

Human Human Human

Mouse Human Human

Chromosome 22 MN1/PITPNB region of Chromosome 22 Whole genome (at 1 Mbp resolution), Chromosomes 22q and 6 Whole genome ENCODE regions Chromosomes 21q, 11q and 6

(White et al. 2004) (Schmegner et al. 2007) (Woodfine et al. 2004; Woodfine et al. 2005) (Farkash-Amar et al. 2008) (Karnani et al. 2007) (Costantini and Bernardi 2008)

Alu/LINE repeats content

Human

Whole genome (at 1 Mbp resolution), Chromosomes 22q and 6

(Woodfine et al. 2004; Woodfine et al. 2005)

Transcription activity; Coding sequences

Human Human

Chromosome 22 Whole genome (at 1 Mbp resolution), Chromosomes 22q and 6 1,589 genes in Chromosomes 21 and 22 ENCODE regions Whole genome

(White et al. 2004) (Woodfine et al. 2004; Woodfine et al. 2005)

Human Human Mouse embryonic

(Jeon et al. 2005) (Karnani et al. 2007) (Hiratani et al. 2008)

House keeping/Tissue specific genes

Mouse

Whole genome

(Farkash-Amar et al. 2008)

H3K4 and H3K9 methylation; H3K9 and H3K14 acetylation; H3K4 and H3K36 trimethylation

Human Human

ENCODE regions MYC, LMNB2, HBB genes, FRAXA region, 1075 kbp of Chromosome 22 Whole genome

(Karnani et al. 2007) (Lucas et al. 2007)

ENCODE regions

(Cadoret et al. 2008; Necsulea et al. 2009)

1,589 genes in Chromosomes 21 and 22 ENCODE regions Whole genome

(Jeon et al. 2005)

Mouse embryonic Human

(Hiratani et al. 2008)

Mode of replication Asynchronous replication of 9– 60% of the sequences throughout S-phase

Human

Acute transition between individual replication timing zones

Human Human

ENCODE regions MN1/PITPNB region of Chromosome 22

(Karnani et al. 2007) (Schmegner et al. 2007)

Gradient transition between individual replication timing zones with large replicons locating at the boundaries of replication timing zones

Human

Chromosomes11q and 21q

Mouse embryonic

Whole genome

(Watanabe et al. 2008; Watanabe et al. 2009) (Hiratani et al. 2008)

Human Mouse

(Karnani et al. 2007) (Farkash-Amar et al. 2008)

Continued

DNA Replication Table 1. Continued Genome duplication characteristic

Species

Genome segment investigated

Reference

X inactivation independent CpG-associated Ori activity

Mouse

X chromosome

(Gomez and Brockdorff 2004)

Correlation of Ori activity with promoter regions and gene density

Mouse embryonic

10 Mbp on chromosome X and 3 (Sequeira-Mendes et al. 2009)

Replicon clusters with interorigin distance 100 kbp

Mouse embryonic Human

10 Mbp on chromosome X and 3 (Sequeira-Mendes et al. 2009)

Ori/Replicon distribution

Human Single large replicons (300 kbp)

Human

Human Mouse Human Human

MYC, LMNB2, HBB genes, FRAXA region, 1075 kbp of Chromosome 22 Chromosomes 21q, 11q and 6

(Lucas et al. 2007)

MYC, LMNB2, HBB genes, FRAXA region, 1075 kbp of Chromosome 22 ENCODE regions Whole genome Chromosomes 21q, 11q and 6 Chromosomes11q and 21q

(Lucas et al. 2007)

(Costantini and Bernardi 2008)

(Cadoret et al. 2008) (Farkash-Amar et al. 2008) (Costantini and Bernardi 2008) (Watanabe et al. 2008; Watanabe et al. 2009)

ENCODE – ENCyclopedia Of DNA Elements represent regions covering 1% (30Mb) of human genome.

In general, this proposal provides a satisfactory explanation for how replication timing is modulated in eukaryotic nuclei. However, two very important questions remain. First, how does the firing efficiency of replication origins increase over the course of S phase? Second, what determines the inherent replication efficiency of a given genomic region? Regarding the first question, one model centers on the concept of polymerase recycling, whereby a fixed number of DNA polymerase complexes are available to the cell. At the onset of S phase, only the most efficient and/or accessible origins have a chance of initiating replication. As the genome becomes duplicated, the number of potential origins decreases, thereby increasing the probability that they will fire. The answer to the second question remains less clear, but alterations in chromatin structure are likely to play a central role. The likelihood of a connection between chromatin structure and replication timing has been well established (Donaldson 2005).

This discussion is most often framed by the correlation between the open chromatin structure present at transcriptionally active regions and the fact that these sites are often replicated early in S phase. In the simplest view, a relaxed chromatin structure results in more accessible genomic DNA, which leads to more efficient binding by both transcription and replication factors. There has been some discussion as to whether replication or transcription plays a causal role in this relationship (i.e., whether early replicating sites “define” regions of transcription or vice versa) but no conclusive results have been obtained. One shortcoming of the genomic methods described earlier is that whereas replication timing profiles certainly reflect the influence of chromatin structure, it is difficult to backtrack and directly examine the nature of this chromatin. This is due in part to the ensemble/pooled nature of genomic analysis, as well as the fact that neither assay permits a particularly detailed examination of nuclear structure.

V.O. Chagin, J.H. Stear, and M.C. Cardoso

CELLULAR ORGANIZATION OF REPLICATION

Chromosomes and chromosomal domains are nonrandomly organized within eukaryotic nuclei and their topology is thought to have functional significance (Cremer and Cremer 2001; Kumaran et al. 2008; Takizawa et al. 2008). Microscopic inspection of nuclei is a powerful approach to investigate the spatiotemporal organization of replication within the context of nuclear architecture. Ongoing DNA synthesis provides a way to directly detect the nuclear sites of DNA replication after introducing labeled nucleotides into the cells. Initially, radioactive thymidine was used (Milner 1969) and later, with the development of antibodies specifically detecting halogenated thymidine analogs (Gratzner 1982; Aten et al. 1992), immunofluorescence analysis of nuclear replication structures became reality (Nakamura et al. 1986; Nakayasu and Berezney 1989; O’Keefe et al. 1992). More recently, live-cell microscopy analysis of replication progression was made possible by introducing fluorescently conjugated nucleotides (Schermelleh et al. 2001) or by expression of fluorescent replication factors (Cardoso et al. 1997; Leonhardt et al. 2000). With these approaches, DNA replication was found to occur at subnuclear sites called replication foci, which accumulate numerous DNA replication factors and cell cycle proteins (Cardoso et al. 1993; Cardoso et al. 1997). These foci show distinct patterns of localization over the course of S phase; as such, studying their composition and dynamics enables an examination of how DNA replication is regulated on a cellular level. Time-lapse microscopy of living mammalian cells over the course of an entire cell cycle (Fig. 2 and Movie 1 online at http:// cshperspectives.cshlp.org/) has shown that early in S phase, immediately following the onset of DNA replication, a multitude of small replication foci are distributed throughout the nucleus. During mid S phase, the replication foci are uniformly larger, and are distributed around the periphery of the nucleoli and nuclear envelope. Finally, at the end of S phase, the sites of replication have been consolidated into a small number

of very large foci. In addition to reflecting the stage of S phase progression, the pattern of replication foci also correlates with the nature and topologyof the chromatin that is being replicated. For example, early replication patterns represent actively transcribed euchromatin, whereas late replication patterns are associated with heterochromatic regions. Thus, this type of microscopic analysis permits the simultaneous visualization of both replication dynamics and chromatin structure in a single cell basis. The role of chromatin modifications and structural rearrangements in replication organization is yet to be established. Some experimental evidence suggests a role of chromatin remodeling and assembly factors in facilitating replication through heterochromatin domains (Collins et al. 2002; Quivy et al. 2008). However, the impact of histone modifications is less clear. For example, disruption of histone H3 lysine 9 trimethylation, the typical epigenetic mark for heterochromatin, does not significantly affect the late replication of mouse chromocenters (Wu et al. 2006). Other histone modifications, such as phosphorylation of the linker histone H1 by Cdk2, have been proposed to play a role in the large-scale decondensation of chromatin associated with replication (Alexandrow and Hamlin 2005). Examining the dynamics of the replication machinery during S phase has in addition provided a detailed view of how DNA replication proceeds on a cellular level. The combination of time-lapse microscopy with fluorescence photobleaching/activation indicated that the processivity of the replication machinery is built on transient interactions of various replication enzymes with a stable core consisting of the processivity factor PCNA (Sporbert et al. 2002; Sporbert et al. 2005; Schermelleh et al. 2007; Gorisch et al. 2008). The latter stayed associated with the DNA and upon photobleaching no recovery was measured for periods of over 10 minutes. When assembly of PCNA was measured, it was found to occur at sites adjacent to the ones previously labeled. This indicated that once replication is completed at a given site, a new replication focus assembles de novo at a neighboring site (Sporbert et al. 2002; Sadoni et al. 2004). Consistent data has

DNA Replication

been reported using double nucleotide pulsechase-pulse experiments, whereby sequentially replicated DNA is labeled by two consecutive pulses of modified nucleotides (Manders et al. 1996; Jackson and Pombo 1998). Thus, replication of a specific genomic region facilitates subsequent loading of new replication factors at neighboring sites. One interpretation of this result is that the act of replication induces local changes in chromatin condensation, which in turn promotes the access/recruitment of replication factors and the initiation of additional replication cycles. Replication would begin at sites with an “open” chromatin conformation, similar to what has previously been proposed. As a result of these initial replication events, the chromatin at adjacent regions, which normally would not support replication initiation, would begin to decondense, leading to an increased probability that origins present at these sites would fire. It is tempting to suggest that the activity of replication helicases such as the MCM proteins would promote local chromatin decondensation. However, it is equally feasible that the mere act of DNA

polymerization is sufficient to induce such changes. This propagating chromatin fiber decondensation can be visualized as analogous to pulling on a shoelace. On a genome-wide scale, this model, which we refer to as the domino model, leads to a simple, self-propagating mode by which the entire chromosomes become fully duplicated by simply spreading the replication process using the “nearest neighbor” principle (Fig. 3). This hypothesis also dovetails nicely with the increasing efficiency model, in that it provides a physical basis for why origin efficiency would increase over the course of S phase. The increased efficiency of late origins is not because of polymerase recycling; rather it can be accounted for by the fact that as a cell progresses through S phase, it becomes more and more likely that any given region of the genome is proximal to a site of DNA replication. Therefore, even origins contained in hyper-condensed, late replicating regions such as the heterochromatin become accessible as more and more of the surrounding chromatin undergoes replication.

A

B G1

Early S

Mid S

Late S G2

Figure 3. Model for the progression of genome replication—domino model. (A) One replicon cluster (red)

initiates replication at early S phase and DNA synthesis proceeds bidirectionally (replication bubbles). This activates initiation of neighboring replicon clusters (blue), which in turn activate replication at later replicon clusters (green) until the whole chromosome (see scheme below) is fully duplicated in G2 phase. (B) Mouse primary fibroblast cell was pulse labeled for 30 minutes with the nucleotide analogue IdU (early S, red) followed by 3 hours chase, another 30 minutes pulse labeling with CldU (mid S, blue) and 3 hours chase, fixation and detection of replication sites with an antibody to DNA ligase I (late S, green). The micrograph represents a mid optical section of the late S phase fibroblast cell depicting the overlay of the three sequential replication labels and the spatiotemporal progression of genome replication.

V.O. Chagin, J.H. Stear, and M.C. Cardoso

FUTURE DIRECTIONS

e.g., many DNA replication factors are shared with DNA repair pathways (the same is also the case between repair and transcription factors) is far more economic for the cell when dealing with similar tasks. On the other hand, this can be a dangerous strategy as mutations on single factors will have pleiotropic effects and factors can get exhausted and become rate limiting. Finally, understanding the DNA replication process will require the ability to connect data from genomic studies with data from single cells in a unified coherent model. Closing this (temporal and spatial) gap is becoming reality with the advent of novel super-resolution nanoscopy techniques. In fact, recent analysis of DNA replication in intact cells provided much increased resolution and consequently much higher numbers of replication foci throughout S phase (Baddeley et al. 2009). In the near future, a major goal of the field will be to visualize and to characterize in full detail single replicons in intact cells, which were previously only identified in stretched DNA fibers (Fig. 4). The

Despite significant advances in the characterization of the process of DNA replication, several basic questions remain unanswered. For example, it is still not fully evident how propagation of DNA replication over chromatin is coordinated with other nuclear processes. In other words, how is duplication of the (epi)genome once and only once per cell cycle achieved with high precision in a highly variable environment including parallel transcription, repair, and other DNA metabolic activities, and on such a differentiated template as chromatin? How are chromatin epistates maintained at every cell cycle? Is transcriptional activity or chromatin structure determining replication timing or are they rather determined by the time they are replicated during S phase? Interdependency of all these nuclear processes is also influenced by the fact that multiple molecular components acting on DNA metabolism are shared. On the one hand, the fact that,

CLSM

3D-SIM

* * ?

(See facing page for legend)

DNA Replication

integration of these data with time lapse analysis will give us the ability to film and dissect the replication of the genome in situ and in real time. ACKNOWLEDGMENTS

We are indebted to Robert M. Martin for the artwork in Figure 1 and to Corella Casas Delucchi for the DNA fibers in Figure 4. We also thank all the past and present members of our laboratory for their many contributions along the years. Last but not least, we thank our many collaborators, which have made our work possible and enjoyable. Our research has been supported by grants of the Deutsche Forschungsgemeinschaft and the Volkswagen foundation. We regret that because of space constraints, we had to eliminate many important and relevant citations. REFERENCES Aladjem MI. 2007. Replication in context: Dynamic shownion of DNA replication patterns in metazoans. Nat Rev Genet 8: 588 –600. Alexandrow MG, Hamlin JL. 2005. Chromatin decondensation in S-phase involves recruitment of Cdk2 by Cdc45 and histone H1 phosphorylation. J Cell Biol 168: 875–886. Anglana M, Apiou F, Bensimon A, Debatisse M. 2003. Dynamics of DNA replication in mammalian somatic cells: Nucleotide pool modulates origin choice and interorigin spacing. Cell 114: 385 –394. Aten JA, Bakker PJ, Stap J, Boschman GA, Veenhof CH. 1992. DNA double labelling with IdUrd and CldUrd for

spatial and temporal analysis of cell proliferation and DNA replication. Histochem J 24: 251– 259. Baddeley D, Chagin VO, Schermelleh L, Martin S, Pombo A, Carlton PM, Gahl A, Domaing P, Birk U, Leonhardt H, et al. 2009. Measurement of replication structures at the nanometer scale using super-resolution light microscopy. Nucleic Acids Res doi: 10.1093/nar/gkp901. Bensimon A, Simon A, Chiffaudel A, Croquette V, Heslot F, Bensimon D. 1994. Alignment and sensitive detection of DNA by a moving interface. Science 265: 2096– 2098. Blow JJ, Dutta A. 2005. Preventing re-replication of chromosomal DNA. Nat Rev Mol Cell Biol 6: 476– 486. Blow JJ, Laskey RA. 1988. A role for the nuclear envelope in controlling DNA replication within the cell cycle. Nature 332: 546–548. Cadoret JC, Meisch F, Hassan-Zadeh V, Luyten I, Guillet C, Duret L, Quesneville H, Prioleau MN. 2008. Genomewide studies highlight indirect links between human replication origins and gene regulation. Proc Natl Acad Sci 105: 15837–15842. Cardoso MC, Joseph C, Rahn HP, Reusch R, Nadal-Ginard B, Leonhardt H. 1997. Mapping and use of a sequence that targets DNA ligase I to sites of DNA replication in vivo. J Cell Biol 139: 579– 587. Cardoso MC, Leonhardt H, Nadal-Ginard B. 1993. Reversal of terminal differentiation and control of DNA replication: Cyclin A and Cdk2 specifically localize at subnuclear sites of DNA replication. Cell 74: 979 –992. Collins N, Poot RA, Kukimoto I, Garcia-Jimenez C, Dellaire G, Varga-Weisz PD. 2002. An ACF1-ISWI chromatinremodeling complex is required for DNA replication through heterochromatin. Nat Genet 32: 627– 632. Costantini M, Bernardi G. 2008. Replication timing, chromosomal bands, and isochores. Proc Natl Acad Sci 105: 3433–3437. Cremer T, Cremer C. 2001. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat Rev Genet 2: 292–301. Dijkwel PA, Wang S, Hamlin JL. 2002. Initiation sites are distributed at frequent intervals in the Chinese hamster dihydrofolate reductase origin of replication but are used with very different efficiencies. Mol Cell Biol 22: 3053–3065.

Figure 4. Connecting replicon analysis on DNA fibers to whole cell in situ replication foci. Human HeLa cells

expressing GFP-tagged PCNA were imaged using either conventional confocal laser scanning microscopy (CLSM) or super-resolution 3D-structured illumination microscopy (3D-SIM). Shown are maximal intensity projections of the full 3D image stacks illustrating the spatial organization of replication foci in late S phase. The inset shows a twofold magnification of the areas containing replicating heterochromatic regions marked by an asterisk in the image. Scale bar, 2 mm. Even in these condensed heterochromatin domains, the much larger number of individual replicating sites revealed by the increased resolution of the 3D-SIM technique is apparent. Below the cell images, combed DNA fibers are shown with a cartoon representation of nucleotide labeled replicon units overlayed in green. The same replicons are shown underneath with the vertical arrows indicating the origin of replication of each replicon. Several of the replicons are closed together in clusters (underlined by green lines), which initiate replication coordinately. By comparing labeling of replicons in combed DNA fibers and the numbers of replication foci counted by conventional light microscopy, it was concluded that one replication focus corresponds to a spatially organized cluster of replicons, as indicated by the dashed lines. Using 3D-SIM and other super-resolution nanoscopy techniques, much higher numbers of replication foci are resolved. This will soon allow us to measure in detail the characteristics of individual replicons that were formerly visualized only on stretched DNA fibers.

V.O. Chagin, J.H. Stear, and M.C. Cardoso Dimitrova DS, Gilbert DM. 2000. Temporally coordinated assembly and disassembly of replication factories in the absence of DNA synthesis. Nat Cell Biol 2: 686– 694. Donaldson AD. 2005. Shaping time: Chromatin structure and the DNA replication programme. Trends Genet 21: 444–449. Drouin R, Lemieux N, Richer CL. 1990. Analysis of DNA replication during S-phase by means of dynamic chromosome banding at high resolution. Chromosoma 99: 273–280. Easwaran HP, Leonhardt H, Cardoso MC. 2005. Cell cycle markers for live cell analyses. Cell Cycle 4: 453– 455. Eshaghi M, Karuturi RK, Li J, Chu Z, Liu ET, Liu J. 2007. Global profiling of DNA replication timing and efficiency reveals that efficient replication/firing occurs late during S-phase in S. pombe. PLoS ONE 2: e722. Fangman WL, Brewer BJ. 1991. Activation of replication origins within yeast chromosomes. Annu Rev Cell Biol 7: 375–402. Farkash-Amar S, Lipson D, Polten A, Goren A, Helmstetter C, Yakhini Z, Simon I. 2008. Global organization of replication time zones of the mouse genome. Genome Res 18: 1562– 1570. Gilbert DM. 2001. Making sense of eukaryotic DNA replication origins. Science 294: 96–100. Gilbert DM. 2002. Replication timing and transcriptional control: Beyond cause and effect. Curr Opin Cell Biol 14: 377 –383. Gomez M, Brockdorff N. 2004. Heterochromatin on the inactive X chromosome delays replication timing without affecting origin usage. Proc Natl Acad Sci 101: 6923– 6928. Gorisch SM, Sporbert A, Stear JH, Grunewald I, Nowak D, Warbrick E, Leonhardt H, Cardoso MC. 2008. Uncoupling the replication machinery: Replication fork progression in the absence of processive DNA synthesis. Cell Cycle 7: 1983–1990. Gratzner HG. 1982. Monoclonal antibody to 5-bromo- and 5-iododeoxyuridine: A new reagent for detection of DNA replication. Science 218: 474–475. Hamlin JL, Mesner LD, Lar O, Torres R, Chodaparambil SV, Wang L. 2008. A revisionist replicon model for higher eukaryotic genomes. J Cell Biochem 105: 321–329. Hiratani I, Ryba T, Itoh M, Yokochi T, Schwaiger M, Chang CW, Lyou Y, Townes TM, Schubeler D, Gilbert DM. 2008. Global reorganization of replication domains during embryonic stem cell differentiation. PLoS Biol 6: e245. Huberman JA, Riggs AD. 1968. On the mechanism of DNA replication in mammalian chromosomes. J Mol Biol 32: 327–341. Hubscher U, Seo YS. 2001. Replication of the lagging strand: A concert of at least 23 polypeptides. Mol Cells 12: 149–157. Hyrien O, Marheineke K, Goldar A. 2003. Paradoxes of eukaryotic DNA replication: MCM proteins and the random completion problem. Bioessays 25: 116– 125. Jackson DA, Pombo A. 1998. Replicon clusters are stable units of chromosome structure: Evidence that nuclear organization contributes to the efficient activation and propagation of S phase in human cells. J Cell Biol 140: 1285– 1295.

Jacob F. 1993. The replicon: Thirty years later. Cold Spring Harb Symp Quant Biol 58: 383 –387. Jacob F, Brenner S. 1963. On the regulation of DNA synthesis in bacteria: The hypothesis of the replicon. C R Hebd Seances Acad Sci 256: 298–300. Jeon Y, Bekiranov S, Karnani N, Kapranov P, Ghosh S, MacAlpine D, Lee C, Hwang DS, Gingeras TR, Dutta A. 2005. Temporal profile of replication of human chromosomes. Proc Natl Acad Sci 102: 6419– 6424. Karnani N, Taylor C, Malhotra A, Dutta A. 2007. Pan-S replication patterns and chromosomal domains defined by genome-tiling arrays of ENCODE genomic areas. Genome Res 17: 865 –876. Kornberg A. 1960. Biologic synthesis of deoxyribonucleic acid. Science 131: 1503–1508. Kumaran RI, Thakar R, Spector DL. 2008. Chromatin dynamics and gene positioning. Cell 132: 929–934. Leonhardt H, Rahn HP, Weinzierl P, Sporbert A, Cremer T, Zink D, Cardoso MC. 2000. Dynamics of DNA replication factories in living cells. J Cell Biol 149: 271– 280. Lucas I, Chevrier-Miller M, Sogo JM, Hyrien O. 2000. Mechanisms ensuring rapid and complete DNA replication despite random initiation in Xenopus early embryos. J Mol Biol 296: 769– 786. Lucas I, Palakodeti A, Jiang Y, Young DJ, Jiang N, Fernald AA, Le Beau MM. 2007. High-throughput mapping of origins of replication in human cells. EMBO Rep 8: 770 –777. Lygeros J, Koutroumpas K, Dimopoulos S, Legouras I, Kouretas P, Heichinger C, Nurse P, Lygerou Z. 2008. Stochastic hybrid modeling of DNA replication across a complete genome. Proc Natl Acad Sci 105: 12295– 12300. Ma H, Samarabandu J, Devdhar RS, Acharya R, Cheng P, Meng C, Berezney R. 1998. Spatial and temporal dynamics of DNA replication sites in mammalian cells. J Cell Biol 143: 1415–1425. Manders EM, Stap J, Strackee J, van Driel R, Aten JA. 1996. Dynamic behavior of DNA replication domains. Exp Cell Res 226: 328–335. McHenry CS. 1988. DNA polymerase III holoenzyme of Escherichia coli. Annu Rev Biochem 57: 519– 550. Milner GR. 1969. Changes in chromatin structure during interphase in human normoblasts. Nature 221: 71–72. Mott ML, Berger JM. 2007. DNA replication initiation: mechanisms and regulation in bacteria. Nat Rev Microbiol 5: 343– 354. Nakamura H, Morita T, Sato C. 1986. Structural organizations of replicon domains during DNA synthetic phase in the mammalian nucleus. Exp Cell Res 165: 291 –297. Nakayasu H, Berezney R. 1989. Mapping replicational sites in the eucaryotic cell nucleus. J Cell Biol 108: 1– 11. Necsulea A, Guillet C, Cadoret JC, Prioleau MN, Duret L. 2009. The relationship between DNA replication and human genome organization. Mol Biol Evol 26: 729– 741. Noguchi H, Prem veer Reddy G, Pardee AB. 1983. Rapid incorporation of label from ribonucleoside disphosphates into DNA by a cell-free high molecular weight fraction from animal cell nuclei. Cell 32: 443– 451. O’Keefe RT, Henderson SC, Spector DL. 1992. Dynamic organization of DNA replication in mammalian cell

DNA Replication nuclei: Spatially and temporally defined replication of chromosome-specific alpha-satellite DNA sequences. J Cell Biol 116: 1095– 1110. Okazaki R, Okazaki T, Sakabe K, Sugimoto K, Sugino A. 1968. Mechanism of DNA chain growth. I. Possible discontinuity and unusual secondary structure of newly synthesized chains. Proc Natl Acad Sci 59: 598– 605. Pandey M, Syed S, Donmez I, Patel G, Ha T, Patel SS. 2009. Coordinating DNA replication by means of priming loop and differential synthesis rate. Nature 462: 940 –943. Pasero P, Bensimon A, Schwob E. 2002. Single-molecule analysis reveals clustering and epigenetic regulation of replication origins at the yeast rDNA locus. Genes Dev 16: 2479–2484. Patel PK, Arcangioli B, Baker SP, Bensimon A, Rhind N. 2006. DNA replication origins fire stochastically in fission yeast. Mol Biol Cell 17: 308– 316. Perumal SK, Yue H, Hu Z, Spiering MM, Benkovic SJ. 2009. Single-molecule studies of DNA replisome function. Biochim Biophys Acta doi: 10.1016/j.bbapap.2009.07.022. Quivy JP, Ge´rard A, Cook AJL, Roche D, Almouzni G. 2008. The HP1–p150/CAF-1 interaction is required for pericentric heterochromatin replication and S-phase progression in mouse cells. Nat Structural Mol Biol 15: 972 –979. Raghuraman MK, Winzeler EA, Collingwood D, Hunt S, Wodicka L, Conway A, Lockhart DJ, Davis RW, Brewer BJ, Fangman WL. 2001. Replication dynamics of the yeast genome. Science 294: 115 –121. Rao PN, Johnson RT. 1970. Mammalian cell fusion: Studies on the regulation of DNA synthesis and mitosis. Nature 225: 159– 164. Rhind N. 2006. DNA replication timing: Random thoughts about origin firing. Nat Cell Biol 8: 1313–1316. Robinson NP, Bell SD. 2005. Origins of DNA replication in the three domains of life. FEBS J 272: 3757–3766. Sadoni N, Cardoso MC, Stelzer EH, Leonhardt H, Zink D. 2004. Stable chromosomal units determine the spatial and temporal organization of DNA replication. J Cell Sci 117: 5353–5365. Schermelleh L, Haemmer A, Spada F, Rosing N, Meilinger D, Rothbauer U, Cardoso MC, Leonhardt H. 2007. Dynamics of Dnmt1 interaction with the replication machinery and its role in postreplicative maintenance of DNA methylation. Nucleic Acids Res 35: 4301–4312. Schermelleh L, Solovei I, Zink D, Cremer T. 2001. Two-color fluorescence labeling of early and mid-to-late replicating chromatin in living cells. Chromosome Res 9: 77– 80. Schmegner C, Hameister H, Vogel W, Assum G. 2007. Isochores and replication time zones: A perfect match. Cytogenet Genome Res 116: 167– 172. Schubeler D, Scalzo D, Kooperberg C, van Steensel B, Delrow J, Groudine M. 2002. Genome-wide DNA replication profile for Drosophila melanogaster: a link between transcription and replication timing. Nat Genet 32: 438–442. Sequeira-Mendes J, Diaz-Uriarte R, Apedaile A, Huntley D, Brockdorff N, Gomez M. 2009. Transcription initiation activity sets replication origin efficiency in mammalian cells. PLoS Genet 5: e1000446.

Sinha NK, Morris CF, Alberts BM. 1980. Efficient in vitro replication of double-stranded DNA templates by a purified T4 bacteriophage replication system. J Biol Chem 255: 4290– 4293. Sparvoli E, Levi M, Rossi E. 1994. Replicon clusters may form structurally stable complexes of chromatin and chromosomes. J Cell Sci 107: 3097– 3103. Sporbert A, Domaing P, Leonhardt H, Cardoso MC. 2005. PCNA acts as a stationary loading platform for transiently interacting Okazaki fragment maturation proteins. Nucleic Acids Res 33: 3521–3528. Sporbert A, Gahl A, Ankerhold R, Leonhardt H, Cardoso MC. 2002. DNA polymerase clamp shows little turnover at established replication sites but sequential de novo assembly at adjacent origin clusters. Mol Cell 10: 1355–1365. Takizawa T, Meaburn KJ, Misteli T. 2008. The meaning of gene positioning. Cell 135: 9– 13. Tom TD, Malkas LH, Hickey RJ. 1996. Identification of multiprotein complexes containing DNA replication factors by native immunoblotting of HeLa cell protein preparations with T-antigen-dependent SV40 DNA replication activity. J Cell Biochem 63: 259–267. Tsurimoto T, Stillman B. 1989. Multiple replication factors augment DNA synthesis by the two eukaryotic DNA polymerases, a and d. Embo J 8: 3883–3889. Watanabe Y, Abe T, Ikemura T, Maekawa M. 2009. Relationships between replication timing and GC content of cancer-related genes on human chromosomes 11q and 21q. Gene 433: 26– 31. Watanabe Y, Fujiyama A, Ichiba Y, Hattori M, Yada T, Sakaki Y, Ikemura T. 2002. Chromosome-wide assessment of replication timing for human chromosomes 11q and 21q: Disease-related genes in timing-switch regions. Human Mol Gen 11: 13–21. Watanabe Y, Shibata K, Ikemura T, Maekawa M. 2008. Replication timing of extremely large genes on human chromosomes 11q and 21q. Gene 421: 74– 80. Watson JD, Crick FH. 1953. Molecular structure of nucleic acids; a structure for deoxyribose nucleic acid. Nature 171: 737–738. White EJ, Emanuelsson O, Scalzo D, Royce T, Kosak S, Oakeley EJ, Weissman S, Gerstein M, Groudine M, Snyder M, Schubeler D. 2004. DNA replication-timing analysis of human chromosome 22 at high resolution and different developmentalstates. Proc Natl Acad Sci 101: 17771–17776. Woodfine K, Beare DM, Ichimura K, Debernardi S, Mungall AJ, Fiegler H, Collins VP, Carter NP, Dunham I. 2005. Replication timing of human chromosome 6. Cell Cycle 4: 172– 176. Woodfine K, Fiegler H, Beare DM, Collins JE, McCann OT, Young BD, Debernardi S, Mott R, Dunham I, Carter NP. 2004. Replication timing of the human genome. Hum Mol Genet 13: 191 –202. Wu R, Singh PB, Gilbert DM. 2006. Uncoupling global and fine-tuning replication timing determinants for mouse pericentric heterochromatin. J Cell Biol 174: 185– 194.

DNA Damage Response Giuseppina Giglia-Mari1,2,3, Angelika Zotter1,4, and Wim Vermeulen1 1

Department of Genetics, Erasmus University Medical Center, Dr Molewaterplein 50, 3015 GE Rotterdam, The Netherlands

2

Department of Cancer Biology, Institute of Pharmacology and Structural Biology, CNRS, 205 route de Narbonne, 31077 Toulouse, France

3

Universite´ de Toulouse Paul Sabatier, 31000, Toulouse, France

4

Core Research Laboratory, Istituto Toscano Tumori, Villa delle Rose, Via Cosimo il Vecchio 2, 50139 Firenze, Italy

Correspondence: [email protected]

Structural changes to DNA severely affect its functions, such as replication and transcription, and play a major role in age-related diseases and cancer. A complicated and entangled network of DNA damage response (DDR) mechanisms, including multiple DNA repair pathways, damage tolerance processes, and cell-cycle checkpoints safeguard genomic integrity. Like transcription and replication, DDR is a chromatin-associated process that is generally tightly controlled in time and space. As DNA damage can occur at any time on any genomic location, a specialized spatio-temporal orchestration of this defense apparatus is required.

elective advantage by random mutations in the genetic material has driven evolution of terrestrial life. Despite this obvious advantage for biological diversity, genome instability has in most cases adverse effects on organismal life. Preservation of genomic integrity is a prerequisite for proper cell function and faithful transmission of the genome to progeny. However, environmental factors and the chemical properties of DNA do not guarantee lifelong stability and proper functioning of the genome. Genomic insults arise from side effects of DNA metabolizing processes, such as replication errors, uncontrolled recombination, off-target mutation induction by somatic hypermutation

S

during antigen production, and inaccurate VDJ recombination (Liu and Schatz 2009; Mahaney et al. 2009). The biggest genomic burden is, however, induced by processes that directly damage DNA. DNA lesions are derived from three main sources (Lindahl 1993; Friedberg et al. 2006): environmental agents such as ultraviolet light, ionizing radiation, and numerous genotoxic chemicals; reactive oxygen species (ROS) generated by respiration and lipid peroxidation; and spontaneous hydrolysis of nucleotide residues, inducing abasic sites and deamination of C, A, G, or 5methyl-C. It is estimated that each cell is confronted with approximately 104 – 105 lesions per day, indicating that clearance of genomic injuries constitutes

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved. Advanced Online Article. Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a000745

G. Giglia-Mari, A. Zotter, and W. Vermeulen

a demanding task to maintain proper genome function. Essential genome processes, such as transcription and replication, are severely affected by DNA lesions. Replication over damaged DNA induces mutations, which may initiate and propagate carcinogenesis. Acute effects arise when lesions block transcription causing cellular senescence or apoptosis, resulting in damageinduced accelerated aging (Mitchell et al. 2003; Akbari and Krokan 2008; Sinclair and Oberdoerffer 2009). THE DNA DAMAGE RESPONSE

To deal with the fundamental problem of genomic erosion, a sophisticated network of DNA damage-response (DDR) systems has evolved. These include a set of DNA repair mechanisms, damage tolerance processes, and cell-cycle

checkpoint pathways. The biological significance of a functional DDR for human health is clearly illustrated by the severe consequences of inherited defects in DDR factors resulting in various diseases, including immune deficiency, neurological degeneration, premature aging, and severe cancer susceptibility (Hoeijmakers 2001; Hoeijmakers 2009). DNA Repair Mechanisms

The heart of the cellular defense against DNA injuries is formed by a variety of DNA repair mechanisms (Hoeijmakers 2001; Hoeijmakers 2009), each with their own damage specificity (Table 1). Together, they are able to remove the vast majority of injuries from the genome. The simplest solution that emerged in evolution is the direct reversal of lesions by specialized activities, such as photolyases that selectively

Table 1. Induction of DNA lesions and corresponding repair pathway. Lesion (1)

CPD, 6-4PP Bulky adducts(2) Intrastrand crosslinks 8-oxo-dG(3) Thymineglycol(3) N7-Alkyl-dG, N3-Alkyl-dA O6-Alkyl-dG 5-methyl-dC Uracil, (Hypo)Xanthine Abasic site Single-strand breaks Double-strand breaks Tyrosyl-30 DNA(8) Mismatches Small insertion/deletions Interstrand crosslinks

Cause

Repair process(es)

Sunlight Food, cigarette smoke Chemotherapy (e.g., Cis-Pt) ROS(4), respiration ROS(4), respiration Food, pollutants Food, pollutants DNMT(6) Spontaneous deamination Spontaneous hydrolysis Ionizing radiation, ROS Ionizing radiation, ROS, VDJ-rec Topo-I inhibition, ROS Replication errors Replication slippage Chemotherapy

NER NER NER BER BER BER DR(5), BER? BER/AID-BER/NER?(7) BER BER Ligation, BER HR, NHEJ SSBR MMR MMR ICLR/ HR?

1. CPD: cyclobutane pyrimidine dimer; 6-4 PP: 6-4 pyrimidine-pyrimidone photo-product. 2. A large group of chemicals conjugated to bases that cause DNA helix destabilization such as: Benzo[a]pyrene (a polycylic aromatic hydrocarbon); Aflatoxins (present in fungal food contaminations); and Nitrosamines (tobacco smoke). 3. A large group of different oxidation products affecting either the base or the phosphate-sugar backbone of which 8-oxo-dG is the most abundant. 4. ROS: reactive oxygen species, produced as side-product of respiration/metabolism and ionizing radiation. 5. DR: direct reversal, involving the suicide enzyme MGMT. 6. DNMT: DNA methyltransferase, functions in epigenetic gene-expression control (e.g., at CpG islands). 7. The mechanism of 5-Me-C repair/conversion is a matter of debate. Recently, a GADD45a-dependent NER reaction was suggested (Barreto et al. 2007). 8. Proteolytic degradation of conjugated Topo-I to 30 DNA termini creates tyrosyl-30 DNA bonds, resolved by TDP1 (El-Khamisy et al. 2009).

DNA Repair

reverse UV-induced DNA damage (Weber 2005) and the suicide enzyme O6-methylguanine transferase (MGMT) that transfers the methyl group from DNA by covalently coupling it to an internal cysteine residue of MGMT, thereby destroying the enzymatic activity (Friedberg et al. 2006). Photolyases are not conserved into the mammalian branch and mammals have to rely on a more complex mechanism to remove UV injuries: nucleotide excision repair (NER) (see below). Base Excision Repair (BER)

Bases with small chemical alterations that do not strongly disturb the DNA double-helix structure are substrates for Base Excision Repair (BER) (Almeida and Sobol 2007; Hegde et al. 2008) (Table 1). These damages, or group of lesions, are targeted by lesion-specific DNA glycosylases that both recognize and remove the damaged base from the sugar-phosphate backbone. The resulting abasic (AP) site is incised by AP-endonucleases and the single nucleotide gap is filled-in by the BER-specific DNA polymerase b and finally sealed by the XRCC1/Ligase III complex. Single strand breaks (SSBs) are repaired by a specialized BER mechanism, designated single-strand break repair (SSBR). The abundant nuclear protein Poly-ADP-RibosePolymerase (PARP) is rapidly activated by SSBs and causes auto-poly-ADP-ribosylation, which recruits the XRCC1/ligase III complex as well as end-processing enzymes such as aprataxin (Gueven et al. 2004) and TDP1 (tyrosyl-DNAphosphodiesterase) to create ligatable DNA ends (Caldecott 2007; El-Khamisy et al. 2009). Nucleotide Excision Repair (NER)

NER removes a broad spectrum of single-strand lesions that cause local helix-destabilization (Table 1). NER is a complex multi-step process, involving the concerted action of at least 25 different polypeptides (Hoeijmakers 1993; Gillet and Scharer 2006) (Fig. 1). Two different modes of damage detection are operational in NER: transcription-coupled NER (TC-NER), which efficiently removes transcription-stalling

lesions and allows quick resumption of transcription (Bohr et al. 1986; Hanawalt 1994), and global genome NER (GG-NER), which localizes lesions anywhere in the genome. In TC-NER, damage sensing is performed by the stalled RNA polymerase, and the Cockayne syndrome factors A and B (CSA and CSB) play essential roles in TC-NER complex assembly (Fousteri et al. 2006; Fousteri and Mullenders 2008). Lesion discrimination in GG-NER is executed by the concerted action of two complexes: XPC/hHR23B (Masutani et al. 1994) and UVDDB (DDB1 and DDB2/XPE) (Chu and Chang 1988; Keeney et al. 1994; Sugasawa et al. 2009). The subsequent steps of TC-NER and GG-NER converge into a common mechanism in which first the NER/basal transcription factor TFIIH (Egly 2001) is recruited (Yokoi et al. 2000; Volker et al. 2001). The bi-directional helicase of TFIIH opens the damaged DNA segment over a stretch of approximately 30 nucleotides (Sugasawa et al. 2009). The unwound DNA is stabilized by XPA and RPA (Replication Protein A) that also orient (de Laat et al. 1998) the two structure-specific endonucleases XPG (O’Donovan et al. 1994) and the ERCC1-XPF complex (Sijbers et al. 1996), which respectively incise the damaged strand 30 and 50 with respect to the lesion. The resulting 25–30 nucleotide single strand gap is filled in by normal DNA replication proteins, including replication factor C (RFC), PCNA, RPA, and the DNA polymerases d, 1, or k (Ogi et al. 2010). Finally, the gap is sealed by DNA ligases I or III, dependent on the proliferation status of the cell (Moser et al. 2007) (Fig. 1). DNA Double-Strand Break Repair (DSBR)

Lesions that are substrates for NER and BER are located in one of the strands of DNA and are removed in a “cut-and-patch”-mechanism. In both cases, the undamaged complementary strand serves as a faithful template for the repair of the damaged strand. Some damaging agents, however, affect both strands, such as ionizing radiation that induces DNA double-strand breaks (DSBs) and agents that produce inter-strand cross-links (ISCLs) (Table 1). These lesions are extremely cytotoxic because they are more

G. Giglia-Mari, A. Zotter, and W. Vermeulen Transcription-coupled NER

Global-Genome NER

Chemicals/ UV-irradiation RNAP2 9 UV-DDB or

??

or

XPC

1a

??

Nascent RNA CSB 8

1b

H2A/H2B TFIIH

XPG

CSA-complex H3/H4

2a

7

2b RPA

CAF1

Helix unwinding Ligase 1/3

XPA 6

3

PCNA XPF/ERCC1

4

DNA polymerase δ/ε/κ

RFC

5

Figure 1. Molecular mechanism of nucleotide excision repair (NER). Bulky DNA lesions (e.g., UV-induced

photo-products and chemical conjugates to nucleotides) that destabilize the DNA double-helix are targeted by NER. Damage recognition is performed by transcription-coupled NER (TC-NER) and Global Genome NER (GG-NER). It is suggested that prior to damage recognition, chromatin has to be modified. Lesions in the transcribed strand of active genes are detected by the elongating RNA polymerase II (RNAP2) and stabilize the interaction with CSB (step 1b). Within GG-NER, lesions are recognized by the UV-DDB and the XPC complexes (step 1a). These intermediates load transcription factor TFIIH together with the endonuclease XPG (steps 2a and 2b). In TC-NER, CSA is also recruited to modify and reposition lesion-stalled RNAP2 (step 2b). After the two modes of lesion detection, the two processes merge into a common pathway of NER factor assembly by recruiting XPA and replication protein A (RPA) (step 3). This NER-intermediate loads and properly orients the structure-specific endonuclease ERCC1/XPF complex (step 4). After dual incision by XPG (30 from the lesion) and ERCC1/XPF (50 from the lesion), a single-strand of 25– 29 nucleotides is created (step 5). XPG is likely involved in recruiting the sliding clamp PCNA, which is loaded by RFC and forms the platform for the gap-filling DNA polymerases d, 1, or k (step 6). Each of these polymerases has been found to participate in NERdependent gap-filling. PCNA or RFC are likely also involved in recruiting the ligases (i.e., Ligase I and Ligase III/ XRCC1, depending on the proliferation capacity of the cell) to seal the nick (step 7). PCNA also plays a role in attracting the histone-chaperone CAF1 (step 8) to restore the chromatin structure after repair (stage 9).

difficult to repair as the cell cannot rely on simply copying the information from the undamaged strand. Two distinct pathways, homologous recombination (HR) and nonhomologous end-joining (NHEJ), repair DSBs (Cahill et al. 2006; Wyman and Kanaar 2006; Helleday et al. 2007). The division of tasks between these repair mechanisms is mainly determined by the phase of the cell cycle. As

HR requires a homologous sister chromatid, it acts exclusively in S- and G2-phase. In contrast, post-mitotic cells and cycling cells in G1 phase have to seal DSBs by NHEJ. Within NHEJ, breaks are quickly recognized by the Ku70/Ku80 hetero-dimer that activates the PI3-kinase DNA-PK and sets the scene for subsequent recruitment of the Artemis nuclease and the MRE11/Rad50/NBS1 (MRN) protein

DNA Repair

complex. These proteins are involved in DNA end-processing, preceding ligation performed by the XRCC4/LigaseIV complex (Weterings and van Gent 2004; Burma et al. 2006; van Gent and van der Burg 2007). During DNA end-processing, loss or changes of a few nucleotides may occur. For this reason, NHEJ, although it very rapidly seals DSBs, is an errorprone repair process. However, when cells do have a homologous template, as in the S- and G2-phase of the cycle, DSBs can be repaired by HR. Homologous recombination is initiated by binding of the MRN complex to a DSB and functions to hold the broken pieces together (de Jager et al. 2001) and provides the structural bases for the CtIP nuclease. The MRN-CtIP complex catalyzes end resection at the break in concert with exonuclease I (EXO1) (Limbo et al. 2007; Sartori et al. 2007; Takeda et al. 2007). Subsequently, RPA binds to the newly created singlestrand region and through a complicated handoff mechanism, the RPA-filament is exchanged into a RAD51 nucleo-protein filament. This RAD51-filament is crucial for strand invasion into the homologous sister, creating a temporarily triplex-DNA structure in which strand exchange occurs (Wyman et al. 2004). The molecular details of these complex transactions are as yet enigmatic, although genetic studies have revealed a whole list of proteins that play an important role in these transactions (Lisby and Rothstein 2009). The biggest challenge within HR-driven DSBR is, however, the question on how homologous regions are identified within the complex nuclear environment. DNA Damage Tolerance

Persisting lesions not removed by any of the repair mechanism will interfere with DNA replication. Lesion-stalled replication forks can lead to highly cytotoxic DSBs and require a prompt response. At least two DNA damage tolerance mechanisms have evolved: translesion synthesis (TLS) and recombination-dependent daughter-strand gap repair (DSGR) (Scully et al. 2000; Li et al. 2002). These processes do not actually remove lesions, but serve as a

temporary solution to overcome stalled DNA replication machines. Upon lesion-induced replication blockage, the regular high-fidelity DNA polymerases ( pold/1 or a) are temporarily exchanged with translesion polymerase (pol z-k) (Friedberg et al. 2005; Lehmann 2006) to synthesize across the lesion. Although TLS can circumvent lesion-induced replication stalling, the reduced fidelity of the alternative polymerases causes generally enhanced mutagenesis. Damage Signaling

To create an extended time window to allow completion of lesion removal prior to replication or cell division, damage sensing is linked to an intricate signal transduction cascade that induces cell cycle arrest (Bartek et al. 2007; Callegari and Kelly 2007). Depending on the nature of the DNA injury and the phase of the cell cycle in which the lesion is encountered, the cell cycle can be arrested at the G1/S transition, within the S-phase, or at the G2/M transition (Zhou and Elledge 2000). Alternatively, when too many injuries are encountered, apoptosis is triggered in order to protect the organism from potentially harmful cells (Bernstein et al. 2002). The phosphatidylinositol 3-kinase (PI3) ATM (Ataxia Telangiectasia mutated) is directly recruited and activated by the DSB-recognizing protein complex MRN. This initiating kinase transduces phosphorylation to a high number of adapter/transducer proteins, carrying the ATM-consensus sequence (Matsuoka et al. 2007). Finally, downstream effector kinases, such as the checkpoint kinase Chk2, are activated (Falck et al. 2002). Bulky lesions cause replication collapse that induces single-strand DNA by the retraction of the replication fork. RPA binds to ssDNA and recruits ATR (ATMrelated) via its association with ATRIP (ATR interacting protein) and activates the checkpoint protein Chk1 (Tibbetts et al. 2000; Chen and Sanchez 2004). RPA covered ssDNA also triggers the Rad17-dependent loading of the RAD9-HUS1-RAD1 (9-1-1 complex), which is an important transducer of checkpoint activation upon DNA damage (Smits et al. 2010). A third PI3 kinase, DNA-PK (DNA-dependent

G. Giglia-Mari, A. Zotter, and W. Vermeulen

protein kinase, composed of its catalytic subunit DNA-PKcs and a regulatory Ku70/80 heterodimer), is also activated by IR-induced DSBs. DNA-PKcs is essential for NHEJ in higher eukaryotes (Burma et al. 2006) and additionally functions in telomere maintenance and induction of apoptosis (Burma and Chen 2004). In total, a complicated set of different emergency strategies are called into action when genomic insults are encountered. Although many of the individual players are identified and the downstream signaling cascades have been dissected, their respective interactions and communication is far from resolved. Intertwined DNA-transacting Processes

The different repair processes are generally considered separate entities. However, in recent years, it has become clear that most of these DDR processes are part of an intricate network with significant overlap, often sharing specific essential components. Several DDR factors appeared to act in diverse DNA maintenance systems. One typical example is the hetero-dimeric ERCC1/XPF complex. This structure-specific endonuclease was originally identified as the nuclease that incises 50 of the DNA lesions within NER (Westerveld et al. 1984; Sijbers et al. 1996). Further analysis revealed additional functions for this complex in HR (Adair et al. 2000; Niedernhofer et al. 2001), interstrand cross-link repair (De Silva et al. 2000; Niedernhofer et al. 2004), and telomere maintenance (Zhu et al. 2003). Besides overlap between distinct repair processes, DDR is also linked to other essential DNA transacting mechanisms, such as transcription and replication. A prime example of such a link is the tight connection between NER and transcription, illustrated by the existence of a specialized transcription-coupled NER pathway (TC-NER) (Bohr et al. 1985; Fousteri and Mullenders 2008). The chromatin remodeling protein Cockayne Syndrome B (CSB) (Citterio et al. 2000) is essential for TC-NER and is implicated in transcription elongation (van den Boom et al. 2004). Moreover, the basal RNA polymerase II transcription factor TFIIH

is also a pivotal factor in NER (Fig. 1) (Schaeffer et al. 1993; Drapkin et al. 1994; Egly 2001; Hoogstraten et al. 2002). Finally, the essential replication factor A (RPA) is implicated in basically all DDR mechanisms, including NER, HR, and damage signaling. Different strategies are used to control the multi-functionality of these factors: (1) distinct spatial organization, (2) incorporation into diverse functional complexes, and/or (3) dynamic sharing of these components. Regulation of pleiotropic functionality of proteins is commonly achieved by distinct posttranslational modifications (PTMs). Within DDR, different PTMs were identified, ranging from phosphorylation, acetylation, methylation, neddylation, mono- and poly-ubiquitylation, and sumoylation to poly-ADP-ribosylation (Harper and Elledge 2007; Huen and Chen 2008). One of the most common PTMs involved in DDR is differential phosphorylation, mainly driven by the ATM, ATR, and DNA-PKcs kinases (Matsuoka et al. 2007). Recent research indicates also that differential ubiquitination plays an important role in DDR regulation (Bergink et al. 2007; Harper and Elledge 2007; Reed and Gillette 2007; Huen and Chen 2008; Alpi and Patel 2009; Panier and Durocher 2009).

STRUCTURAL AND FUNCTIONAL ORGANIZATION OF DDR Chromatin and DDR

The nucleus is highly structured and functionally compartmentalized in part due to areas of various degrees of chromatin compaction, creating possible obstacles for DDR factor accessibility. Decompaction and subsequent restoration of the starting chromatin structure in conjunction with DDR thus creates another level of complexity in genome maintenance regulation. Chromatin-associated processes such as transcription, replication, and DNA repair are regulated by a complex set of structural changes in chromatin (Groth et al. 2007; Li et al. 2007). Control of chromatin functions and its compaction occurs by at least four known processes: (1) active ATP-consuming

DNA Repair

remodeling machines of the SWI/SNF-superfamily of DNA-dependent ATPases (NevesCosta and Varga-Weisz 2006; Saha et al. 2006) that slide or physically evict core histones or entire nucleosomes from active sites; (2) incorporation of diverse histone variants by histone chaperones (Loyola and Almouzni 2007; Altaf et al. 2009); (3) differential binding of abundant non-core histone proteins, such as the linker histone H1, the family of high mobility group proteins (HMG), or different isoforms of the hetero-chromatin protein 1 (HP1); and (4) covalent modifications or PTMs of the core histones, such as acetylation, methylation, phosphorylation, and ubiquitylation (He and Lehming 2003). Accordingly, recently a large number of chromatin modifications and remodeling events were shown to be linked to DDR (Groth et al. 2007; Dinant et al. 2008; Misteli and Soutoglou 2009; Nag and Smerdon 2009; van Attikum and Gasser 2009). Access, Repair, and Restore

Despite increasing knowledge of the role of chromatin in DDR, a general mode of action or detailed mechanistic insight is lacking. Already in 1991 a hypothetical three step model for DNA repair in chromatin was postulated (Smerdon 1991), the so-called “ARR-model,” for Access, Repair, and Restore, based on analogy to transcription regulation in chromatin. In this model, it was postulated that chromatin remodeling would be required to provide “access” of damage-recognition factors to initiate “DNA repair” and, when the job is finished, “restoration” of the chromatin structure. Since then, clear indications have been found that the H3/H4 chaperone CAF1, likely in conjunction with Asf1 (anti-silencing function 1), is implicated in restoring chromatin after NER (Green and Almouzni 2002; Mello et al. 2002; Polo et al. 2006). Additional studies implicated Asf1 and FACT (facilitating transcription factor) (Chen et al. 2008; Heo et al. 2008) in histone exchange near DSBs. However, less direct evidence for chromatin remodeling factors for the first step (access) was found. Although several chromatin remodelers, such as INO80

(Downs et al. 2000; van Attikum et al. 2004), facilitate DDR factor recruitment by moving histones away from the break, they appear to act after the initial damage recognition. In addition, chromatin modifications in yeast occur after UV-irradiation by Gcn5-induced H3 acetylation and are dependent on the Swi/Snf DNA translocase Rad16/Rad7/Abf1. Despite the more open chromatin structure by hyperacetylation, it is not directly clear whether this modification facilitates the recruitment of the Rad4 (yeast ortholog of XPC) DNA damage recognition protein (Waters et al. 2009). Phosphorylation of the Histone H2A Variant H2AX

The most prominent DDR-associated covalent histone modification is the phosphorylation of the histone H2A-variant H2AX in response to DNA damage by the checkpoint kinases ATM, ATR, and DNA-PKcs (Rogakou et al. 1998; O’Driscoll et al. 2003; Falck et al. 2005). H2AX is incorporated into approximately 5% – 25% of histone octamers, although its phosphorylation (gH2AX) is constrained to microscopically discernable structures, the ionizing irradiation-induced foci (IRIF) (Fig. 2). Phosphorylation of H2AX is a relatively early event after damage, immediately following MRN binding and ATM activation. These gH2AX foci co-localize with most of the DSB-associated DDR factors (see below) and are thought to serve as docking sites for recruiting and retaining DNA repair and signaling factors to DSBs. g-H2AX spreads over several megabases around DSBs and appears condensed into IRIFs (Rogakou et al. 1999), suggesting a dominant structural role in DSB-DDR. Surprisingly, however, although mice lacking H2AX are radiation-sensitive and exhibit several features associated with defective DDR, they are only partially defective in DSB repair and are not fully compromised in checkpoint activation (Celeste et al. 2002). This notion argues, contrary to expectation, that this impressive structural organization into large molecular assemblies only makes the DDR process more efficient but is not essential for DDR.

G. Giglia-Mari, A. Zotter, and W. Vermeulen

A

γ-irradiation

MDC1-GFP

Anti-γH2AX

B

Filter UV-irr.

XPC-GFP

Anti-CPD

Figure 2. Localization of DSBR and NER factors. Typical examples of subnuclear distributions of MDC1, an important factor involved in an early step of DNA double-strand break repair via homologous recombination (A) and XPC, one of the damage-recognizing proteins in NER (B). (A) GFP signal (upper panel) derived from MDC1-GFP stably expressed in U2OS cells, 2 hours after g-irradiation and fixed with paraformaldehyde, showing the accumulation of MDC1 in typical ionizing-radiation induced foci (IRIF), counter stained with anti-g-H2AX antibodies (lower panel). (B) NER factors do not accumulate in nuclear foci after DNA damage induction. To allow live cell analysis of NER factor kinetics, local UVdamage infliction through a micro-porous filter is performed (see Fig. 3B,C). XPC-GFP expressed in human fibroblasts (upper panel) accumulates at local UV-damaged sub-nuclear areas as recognized by anti-CPD (the major UV-induced DNA lesion) antibodies (lower panel).

UV-light also induces g-H2AX, although in this case the modification is homogenously distributed throughout the nucleus, with the exception of cells in S-phase (O’Driscoll et al. 2003; Hanasoge and Ljungman 2007; Stiff et al. 2008). H2AX phosphorylation upon UV in non-S-phase cells depends on ATR and active processing of the lesion by the NER machinery (O’Driscoll et al. 2003; Marti et al. 2006), suggesting that NER-intermediates trigger this response. The notion that g-H2AX formation occurs in response to NER and that NER is proficient in H2AX-deficient cells, suggests that this modification mainly plays a role in checkpoint activation during UV lesion repair.

DDR in Higher Order Chromatin Structure

Within mammalian cells, chromatin comes in different flavors, classified into compacted and often transcription-silent heterochromatin and the more open transcriptionally active euchromatin. Chromatin compaction is, however, dynamic and varies within different nuclear areas, throughout the different phases of the cell cycle and between different somatic cell types. Obviously, higher order packaging beyond the basic nucleosomal level will raise further accessibility problems. The versatile NER pathway removes lesions throughout the genome, although repair of photo-lesions in nucleosomal templates is repaired slower in vitro compared to naked DNA (Nag and Smerdon 2009). Evidence for an inhibitory effect of higher order chromatin structure on NER in vivo is restricted to yeast studies. Yeast mutants for the histone acetyltransferase Gcn5, which affect chromatin packaging of specific loci, exhibit significant reduced UV-lesion repair on these silenced loci (Waters et al. 2009). Evidence for reduced repair kinetics of DSBs in compact heterochromatin was recently provided (Goodarzi et al. 2008). Repair in these areas requires the release of the heterochromatin-associated and transcriptional co-repressor protein KAP1 (KAP-associated protein 1) from these compact regions. This release appeared to depend on phosphorylation by ATM, providing evidence for a direct role of this important DSB PI3-kinase in chromatin decompaction to support DDR. ATM-dependent repair in heterochromatin can also be alleviated by depletion of the heterochromatic factor HDAC1/2 and simultaneous depletion of the three isoforms (a, b, and g) of HP1 (heterochromatin protein 1). The implication of this protein in DDR has recently led to controversy in the DDR literature (Ball and Yokomori 2009), as one group describes a DNA damage-induced release of phosphorylated HP1 from H3K9me (Histon H3, lysine 9 methylation) (Ayoub et al. 2008), while Luijsterburg and co-workers report recruitment of HP1 to damaged sites (Luijsterburg et al. 2009). Ayoub showed an initial release of phosphorylated HP1 followed by a subsequent

DNA Repair

spreading to neighboring chromatin. This discrepancy is difficult to explain, besides possible difference in DNA damage induction and slight differences in kinetic measurements. A possible explanation for the apparent initial disappearance prior to the observed accumulation at damaged sites might be the sudden highly localized damage induction. The extreme high local concentration of light might in addition to DNA damage also induce chromatin-protein damage. The next wave of HP1 accumulation reflects then the more physiological response to DNA damage induction. Despite the conflicting data and interpretations, it is clear that compaction of chromatin and HP1 play an as yet not entirely understood role in DDR, as disruption of HP1 orthologs in C. elegans induces a diverse spectrum of DNA damage sensitivities (Ball and Yokomori 2009; Luijsterburg et al. 2009). Tools to Analyze DDR in Living Cells

The dynamic interactions with chromatin and the multiple engagements of DDR factors indicate that analysis of each of the separate processes in vitro is not sufficient to fully uncover mechanistic details, and demands cellular biological approaches. The possibility to genetically tag proteins with the autofluorescent protein GFP has revolutionized cell biology (Tsien and Miyawaki 1998). The simultaneous technological advances in microscopy and development of quantitative fluorescent measurements and sophisticated photo-bleaching procedures (White and Stelzer 1999; Houtsmuller and Vermeulen 2001; LippincottSchwartz et al. 2001) have provided spectacular new insights into the regulation and dynamic organization of chromatin-associated processes (Houtsmuller et al. 1999; Phair and Misteli 2000). In particular, the development of several systems to locally introduce DNA damage or immobilize DDR factors in cultured living cells has been beneficial (Figs. 2 and 3): (1) irradiation through a filter or mask that partly shield the cells (Nelms et al. 1998; Katsumi et al. 2001; Mone et al. 2001); (2) micro-beam laser irradiation, with or without photo-sensitizers, at sub-nuclear areas (Cremer et al. 1980; Tashiro

et al. 2000; Lukas et al. 2003; Meldrum et al. 2003; Lan et al. 2004; Dinant et al. 2007); (3) guided a-particle and heavy iron radiation (Jakob et al. 2003; Aten et al. 2004; Hauptner et al. 2004); (4) integration of rare-cutting endonucleases (Lisby et al. 2004; Rodrigue et al. 2006; Soutoglou et al. 2007); and (5) DDR protein tethering to specific integrated amplified sequences (Soutoglou and Misteli 2008). Organization of DNA Double-Strand Break Response

In situ studies revealed that next to g-H2AX, a high number of DDR proteins relocalize into IRIF foci upon genomic stress (Bekker-Jensen et al. 2006). This is particularly pronounced for proteins implicated in the repair and signaling of DSBs by homologous recombination (HR) (Fig. 2). A systematic analysis of the spatial distribution of DSB-DDR factors using a method to locally introduce DSBs in cultured living cells (Lukas et al. 2003) resulted in a localization classification of DDR factors (Bekker-Jensen et al. 2006). In this procedure, cells were cultured in the presence of photo-sensitizing nucleotideanalogs (Iodo-deoxyuridine) prior to microbeam laser irradiation with 337 nm that induces DSB in a user-defined sub-nuclear area (Lukas et al. 2003). Several subclasses of repair proteins were found based on their recruitment properties: (1) The major checkpoint mediators, such as Mdc1 (mediator of DNA damage checkpoint protein 1), 53BP ( p53 binding protein), BRCA1 (breast cancer protein 1), ATM, and the MRN complex, co-localize in IRIF with g-H2AXdecorated chromatin, termed “DSB-flanking chromatin”. Assembly of these proteins at the DSB-flanking chromatin appeared to occur throughout the cell cycle. It has been estimated that these foci contain several hundred copies of each of the participating DDR factors. (2) Another group of DSB-activated proteins assembles in much smaller ssDNA micro-compartments that are most likely formed by 50 resections at DSB, an important HR intermediate. These “microfoci” are only formed in S- and G2-phase cells and typically accumulate next to RPA, factors directly involved in HR repair, like

G. Giglia-Mari, A. Zotter, and W. Vermeulen

A

B

XPC-GFP

Hoechst

Merge

TFIIH-GFP

Hoechst

Merge

GFP-XPA

Hoechst

Merge

D

E

Relative fluorescence

C

0

Time (s)

Figure 3. Live cell analysis of NER. (A) Distribution of three different NER factors tagged with GFP in living cells; DNA is stained by the DNA stain Hoechst. The damage recognition factor XPC is concentrated in nuclear areas (top panel) that also contain high DNA concentrations when the XPC-GFP (Hoogstraten et al. 2008) is expressed in mouse-embryonal fibroblasts that exhibit the species-specific dense DNA-containing areas. This inhomogeneous distribution contrasts to other NER factors such as XPA (Rademakers et al. 2003), which are homogenously distributed (lower panel) and the repair/transcription factor TFIIH that is enriched in nuclei (Hoogstraten et al. 2002). (B) Schematic cartoon of the procedure to locally inflict UV-damage in living cultured cells by irradiation through a microporous filter (Volker et al. 2001). (C) Local accumulation of XPB-GFP (TFIIH subunit) in UV-damaged areas used to determine the dwell time of this NER factor in the damaged area by fluorescence recovery after photo bleaching (FRAP). (D) Human fibroblasts expressing XPC-GFP locally damaged at the indicated position ( purple flash, middle panel) by UV-C laser (Dinant et al. 2007). The right panel shows a clear accumulation of this protein as soon as 30 seconds after irradiation. (E) Schematic representation of the quantification of NER factor time-dependent accumulation at local UV-damage in living cells to determine the assembly kinetics of these factors within the chromatin-bound NER complex.

Rad51, Rad52, BRCA1, and FANCD2 (Fanconi anemia complementation group D2). They also contain the ssDNA-activated checkpoint kinase ATR and the 9-1-1 complex (Warmerdam et al. 2009). Also MRN and BRCA1 were found in these microfoci independent of g-H2AX or Mdc1, while their recruitment to DSB-flanking chromatin is dependent on these proteins. (3) Several DSB repair factors, particularly those involved in NHEJ, such as DNA-PKcs and Ku70/80, could not be found to re-localize

into microscopically discernible foci. It is likely that this process proceeds much faster than HR and that NHEJ factors do not need to be loaded in such large molecular assemblies to execute their function. However, using procedures that introduce high local concentrations of breaks in living cells, with the aid of multi-photon micro-beam laser irradiation, microscopically discernible accumulations of NHEJ could be found (Mari et al. 2006; Uematsu et al. 2007). These accumulations likely reflect high local

DNA Repair

concentrations of breaks and repair factors rather than a specific chromatin structure. (4) Other factors implicated in DSB processing do not exhibit discernible accumulation at sites of damage, since these proteins are omnipresent on chromatin and simply get post-translationally modified at or near breaks. One of them is Smc1 (Structural Maintenance of Chromosomes 1), a structural component of the cohesin complex required for sister chromatid cohesion during S-phase and also implicated in DSB repair. Smc1 is phosphorylated on Serine 957 (a canonical ATM target site) by ATM and ATR after exposure to a broad array of stimuli including IR, HU, and UV-light (Kim et al. 2002). (5) While many of the DDR factors are recruited or retained at the site of damage, proteins like the effector kinases Chk1 and Chk2 are released from chromatin in response to DNA damage. Activated checkpoint proteins distribute through to nucleus to activate soluble pan-nuclear targets such as p53 and Cdc25A (Kastan and Bartek 2004). Also, these effectors, crucial for efficient DNA damageinduced gene expression ( p53) and cell-cycle arrest (Cdc25A, p53), do not accumulate at DNA damage sites. Dynamics and Function of IRIF

One obvious question is: What is the function of IRIFs? Although they are certainly associated with DSBs, this seemingly easy question is, however, difficult to answer, and different models can be envisaged; for example, foci may (1) represent sites of active DSB repair or (2) sites refractory or difficult to repair. Determining the dynamics of these structures might shed some light. Dwell time measurements of HR proteins in IRIFs have revealed a highly dynamic interaction of some of the factors (Rad54 and Rad52) with these apparent long-lasting structures (Essers et al. 2002). The more structural protein Rad51 that forms nucleo-protein filaments exhibits much longer residence times in these foci. Real time imaging in living cells of GFPtagged DSB-DDR protein distribution in response to local damage induction allowed determination of the assembly kinetics of the

different factors (Bekker-Jensen et al. 2005). One of the most striking findings in these studies is that assembly occurs in two kinetically separable waves, i.e., an immediate loading of, for example, MRN and MDC1, followed by a second wave of loading of, for example, 53BP1 and BRCA1. This secondary, slower wave has been suggested to retain and concentrate the repair factors near the insult. It is surprising to note that particularly the DNA repair proteins (Rad51, Rad54, etc.) are only found in the micro-foci, whereas DDR proteins implicated in damage recognition and signaling appear to accumulate in larger structures. Recently, it was shown that H2A and H2AX ubiquitination occurs in response to DSB and that these modified histones as well as the enzymes RNF8, Ubc13, RNF168, and HERC2 accumulate in large foci (Huen et al. 2007; Mailand et al. 2007; Doil et al. 2009; Stewart et al. 2009; Bekker-Jensen et al. 2010). This histone modification appears to play an important role in recruiting the signaling proteins 53BP1 and BRCA1. It seems that the larger structures are particularly important for transducing and amplifying damage signaling. Despite intensive research, the exact molecular function of IRIFs remains enigmatic. The most popular model is that IRIFs serve to locally concentrate the enzymes required for DSB. If that is indeed their prime function, it is surprising to note that such huge amounts of activities are required. The high number of proteins and the long-lasting presence of foci argue that the reaction catalyzed by these enzymes is inefficient. In light of this reasoning, the option that foci represent breaks refractory or difficult to break remains open. Recently, a hint toward the possible molecular function of the large chromatin depositions was revealed by directly targeting DDR factors to specific artificial genomic positions in the absence of actual lesions (Soutoglou and Misteli 2008). Immobilizing repair factors to chromatin elicits a damage signaling response without the actual presence of DNA damage. These data suggest that prolonged binding of repair factors is sufficient to trigger, sustain, and amplify the DNA damage signaling.

G. Giglia-Mari, A. Zotter, and W. Vermeulen

Another interesting debate in the field with respect to structure and nuclear distribution of DSBs is on the choreography of DSBs in the nuclear space and the issue of how ends of different breaks find each other. Chromosomal translocations are initiated by DSBs and it has been shown that translocations between different chromosomes occur in a cell-type specific manner (Meaburn et al. 2007), a phenomenon likely driven by the non-random spatial organization of the genome (Roix et al. 2003; Lanctot et al. 2007; Meaburn and Misteli 2007). These observations favor a so-called “contact-first” model, i.e., that chromosome fibers should be in close proximity to allow translocations, as opposed to a “breakage-first” model in which breaks are mobile and roam the nucleus for interactions. Soutoglou et al. developed an elegant procedure to investigate this enigma by specifically generating a single DSB, using a specific endonuclease site located between two repetitive sequences of distinct repressor binding sites, which can be visualized by different fluorescently-tagged repressors (Soutoglou et al. 2007). With this procedure, very limited movement over time of the DSB was observed, thus supporting the “contact-first” model. In contrast, however, Aten et al. found evidence for the “breakage-first” model, using a-partical tracks to inflict DSB (Aten et al. 2004), in which they observed limited movement and fusion of foci. These contradictory findings might be derived from the different experimental procedures and cell-cycle phase in which the analyses are performed. In addition, both models may not be mutually exclusive and both processes may play a role in the process of chromosomal translocations. Organization of Nucleotide Excision Repair

In the absence of DNA damage, NER factors are generally homogenously distributed throughout the nucleoplasm (Fig. 3A); however, XPC and TFIIH are exceptions to this rule. XPC appears to co-localize with dense or high DNA concentrations (Hoogstraten et al. 2008) and TFIIH is enriched in the nucleolus (Hoogstraten et al. 2002). Unlike DSB repair, NER

factors do not exhibit re-localization into microscopically discernible subnuclear structures upon DNA damage induction, making it difficult to unravel the structural organization of NER-dependent damage response. Despite the absence of microscopically discernable repair foci within NER, live cell studies on NER proteins were, however, the first to reveal the highly dynamic character and mobility of chromatin-transacting proteins in mammalian cells (Houtsmuller et al. 1999). GFP-based studies showed that the NER-specific 50 -endonuclease ERCC1/XPF (Houtsmuller et al. 1999), the damaged DNA binding proteins DDB2 (Luijsterburg et al. 2007) and XPC (Hoogstraten et al. 2008), the damage verification factor XPA (Rademakers et al. 2003), the 30 endonuclease XPG (Zotter et al. 2006), and the multifunctional TFIIH complex (Hoogstraten et al. 2002; Giglia-Mari et al. 2006) each move with their own unique rate through the nucleus. This notion contrasts to an earlier model, based on isolation of NER factors from cell nuclei, in which it was postulated that an assembly or complex of most NER-factors, i.e., the so-called “nucleotide excision repairosome,” forms the functional unit within NER (Svejstrup et al. 1995). Further application of cell lines stably expressing these biologically active GFP-tagged NER factors have allowed detailed analysis of the kinetic properties of each of these factors when actively engaged in NER. The development of a procedure to locally inflict NER-specific DNA damage in mammalian cells at the single cell level, using UV-C light irradiation through a microporous filter (Katsumi et al. 2001; Mone et al. 2001) (Figs. 2 and 3B) and later by the development of UV-C laser microirradiation set-up (Dinant et al. 2007) (Fig. 3D), provided detailed insight into how the different NER factors assemble into NER complexes (Mone et al. 2004; Zotter et al. 2006; Luijsterburg et al. 2007; Alekseev et al. 2008; Hoogstraten et al. 2008; Dinant et al. 2009; Nishi et al. 2009) (Fig. 3E). Additional FRAP studies on a series of NER factors, using different doses of UV (correlating with different concentrations of photo-lesions, which are a prime target for NER), variable repair times and in cell

DNA Repair

lines with distinct NER-efficiencies further provided insight into the kinetic framework of NER in living mammalian cells (Politi et al. 2005; Luijsterburg et al. 2010). Most of the NER factors, with the exception of the DNA damage sensor XPC (Hoogstraten et al. 2008; Nishi et al. 2009), freely diffuse through the nuclear space and only assemble into functional repair complexes at the site of the damage. Advanced modeling based on NER kinetic studies favored a model of kinetic-proofreading to achieve high specificity of lesion recognition by proteins with a relatively low discrimination of damaged sites versus non-damaged DNA (Luijsterburg et al. 2010). NER and Damage Signaling

In spite of detailed knowledge on the NER mechanism, the connection with UV-induced DNA damage signaling is less well characterized. The ATR kinase and loading of the 9-1-1 complex involving the RAD17 clamp-loader certainly play a role in UV-damage signaling (Niida and Nakanishi 2006). However, the confounding effect of UV-induced replication stress (Zou and Elledge 2003; Falck et al. 2005) makes it difficult to disentangle NER-related signaling from replication stress-induced signaling. Nevertheless, a direct relationship between NER and checkpoint signaling was identified (Giannattasio et al. 2004). In addition, NERdependent ATR activation and H2AX phosphorylation occurred in non S-phase cells (O’Driscoll et al. 2003; Hanasoge and Ljungman 2007), likely caused by ssDNA-containing NER-intermediates (Stiff et al. 2008). NERprocessing and ATR are also required for UVinduced H2A ubiquitination (Bergink et al. 2006). A similar chromatin mark was found in response to DSBs (Huen et al. 2007; Ikura et al. 2007; Mailand et al. 2007; Nicassio et al. 2007). Strikingly, the enzymes involved in DSBinduced H2A ubiquitination, such as UBC13 and RNF8, were also responsible for the NERdependent H2A-ubiquitination (Marteijn et al. 2009). This UV-induced chromatin mark further triggers the recruitment of MDC1, BRCA1, and 53BP1, factors previously known

to function in DSB-induced DDR. These findings suggest highly conserved chromatin modification and loading of signaling factors between entirely distinct DDR pathways, DSB repair, and NER. This notion further corroborates the suggestion that large-scale chromatin modifications in response to DNA damage and local concentration of DDR factors play an important function in damage signal maintenance and amplification (Soutoglou and Misteli 2008; Marteijn et al. 2009). Dynamic Organization in Somatic Cells

It is important to keep in mind that all described live cell studies in DNA repair have been conducted on cultured cells. Cultured cells are under constant stress (e.g., atmospheric oxygen) and usually in a highly replicative status. Moreover, physiological processes critically depend on the cellular context or micro-environment (cell-cell contacts with neighboring cells, extracellular matrix, etc.). Within larger animals, more than 90% of the somatic cells are in a non-proliferative status, thus making extrapolations to the actual in vivo situation even more delicate. To acquire an integral view on DDR in different post-mitotic highly differentiated cells, knock-in mouse models expressing endogenously fluorescently tagged crucial proteins have been generated. In the first example of such a mouse model, the yellow variant of GFP was fused to the XPB subunit of the repair/ transcription factor TFIIH, by targeted integration into the endogenous Xpb gene locus (Giglia-Mari et al. 2009). Previous studies in cultured cells showed that TFIIH interacts for a few seconds with transcription initiation sites (Hoogstraten et al. 2002). A similar dynamic behavior was observed in highly proliferative cells in mouse tissue, e.g., skin keratinocytes (Giglia-Mari et al. 2009). Surprisingly, transcription-dependent chromatin binding takes on the order of minutes/hours in post-mitotic cells, such as neurons. This suggests that a wellknown and extensively studied cellular pathway, such as transcription, can have a completely different dynamic organization in different cells. The mechanistic reason for this dynamic

G. Giglia-Mari, A. Zotter, and W. Vermeulen

behavioral change remains enigmatic and will be the next challenge to reveal. In view of these observations, it remains questionable whether current concepts of DDR functioning are applicable to all cell types and tissues. Is a keratinocyte repairing DNA damage differently than a neuron? Do all DDR factors play similar functions in different cells? Are there also development- and differentiationdriven variations in DDR, and if so, how are they regulated? Intriguingly, differential repair kinetics and damage sensitivities have been found in somatic cultured cells and embryonic stem cells (ESC) (de Waard et al. 2008). Part of these differences can be attributed to a more open chromatin structure in ESC, as further reduction of chromatin compaction by reducing the amount of the linker histone H1 increased the damage response (Murga et al. 2007). CONCLUSION

With the availability of protein tagging technology and advanced confocal imaging, spectacular novel insight in the dynamic interplay of DDR factors with damaged DNA has been gained. These studies have revealed a general minimal model of freely diffusing constituents that assemble in a stochastic fashion with damaged DNA to create dynamic assemblies of multiple factors at these sites to finally exert their function (Dinant et al. 2009; Luijsterburg et al. 2010). This view of the dynamic organization of complex pathways in the mammalian cell nucleus has challenged the current textbook models that give the impression of stable structures containing large complexes, in which all constituents are present at all times. Live cells studies on DDR have revealed that pathways intermingle and share components. Controlling this complex interplay requires perfect coordination in time and space of functions to ensure stability and maintenance of functions. But differently from man-made machines, the dynamic organization of nuclear functions is not the result of a predefined master plan, but, fascinatingly, is the result of a long evolution process selecting for a subtle mix of

stochastic diffusion and protein affinities for optimal performance. Dynamic studies in living cells and, recently, in living animals, allow us to study repair mechanisms in action. Together with the current “omics” approaches ( proteomic, genomic and transcription arrays, deep-sequencing, etc.) and the emerging systems biological procedures, these new tools and techniques provide tremendous opportunities to reach a full understanding of DDR, the biological consequences of inefficient DDR in patients and in the general population, in cancer protection, and in agerelated diseases.

REFERENCES Adair GM, Rolig RL, Moore-Faver D, Zabelshansky M, Wilson JH, Nairn RS. 2000. Role of ERCC1 in removal of long non-homologous tails during targeted homologous recombination. Embo J 19: 5552–5561. Akbari M, Krokan HE. 2008. Cytotoxicity and mutagenicity of endogenous DNA base lesions as potential cause of human aging. Mech Age Dev 129: 353–365. Alekseev S, Luijsterburg MS, Pines A, Geverts B, Mari PO, Giglia-Mari G, Lans H, Houtsmuller AB, Mullenders LH, Hoeijmakers JH, et al. 2008. Cellular concentrations of DDB2 regulate dynamic binding of DDB1 at UV-induced DNA damage. Mol Cell Biol 28: 7402– 7413. Almeida KH, Sobol RW. 2007. A unified view of base excision repair: lesion-dependent protein complexes regulated by post-translational modification. DNA Repair 6: 695 –711. Alpi AF, Patel KJ. 2009. Monoubiquitylation in the Fanconi anemia DNA damage response pathway. DNA Repair 8: 430 –435. Altaf M, Auger A, Covic M, Cote J. 2009. Connection between histone H2Avariants and chromatin remodeling complexes. Biochem Cell Biol 87: 35–50. Aten JA, Stap J, Krawczyk PM, van Oven CH, Hoebe RA, Essers J, Kanaar R. 2004. Dynamics of DNA doublestrand breaks revealed by clustering of damaged chromosome domains. Science 303: 92– 95. Ayoub N, Jeyasekharan AD, Bernal JA, Venkitaraman AR. 2008. HP1-beta mobilization promotes chromatin changes that initiate the DNA damage response. Nature 453: 682–686. Ball AR Jr, Yokomori K. 2009. Revisiting the role of heterochromatin protein 1 in DNA repair. J Cell Biol 185: 573 –575. Barreto G, Schafer A, Marhold J, Stach D, Swaminathan SK, Handa V, Doderlein G, Maltry N, Wu W, Lyko F, et al. 2007. Gadd45a promotes epigenetic gene activation by repair-mediated DNA demethylation. Nature 445: 671 –675.

DNA Repair Bartek J, Bartkova J, Lukas J. 2007. DNA damage signalling guards against activated oncogenes and tumour progression. Oncogene 26: 7773–7779. Bekker-Jensen S, Lukas C, Kitagawa R, Melander F, Kastan MB, Bartek J, Lukas J. 2006. Spatial organization of the mammalian genome surveillance machinery in response to DNA strand breaks. J Cell Biol 173: 195– 206. Bekker-Jensen S, Lukas C, Melander F, Bartek J, Lukas J. 2005. Dynamic assembly and sustained retention of 53BP1 at the sites of DNA damage are controlled by Mdc1/NFBD1. J Cell Biol 170: 201– 211. Bekker-Jensen S, Rendtlew Danielsen J, Fugger K, Gromova I, Nerstedt A, Lukas C, Bartek J, Lukas J, Mailand N. 2010. HERC2 coordinates ubiquitin-dependent assembly of DNA repair factors on damaged chromosomes. Nat Cell Biol 12: 80– 86. Bergink S, Jaspers NG, Vermeulen W. 2007. Regulation of UV-induced DNA damage response by ubiquitylation. DNA Repair 6: 1231– 1242. Bergink S, Salomons FA, Hoogstraten D, Groothuis TA, de Waard H, Wu J, Yuan L, Citterio E, Houtsmuller AB, Neefjes J, et al. 2006. DNA damage triggers nucleotide excision repair-dependent monoubiquitylation of histone H2A. Genes Dev 20: 1343–1352. Bernstein C, Bernstein H, Payne CM, Garewal H. 2002. DNA repair/pro-apoptotic dual-role proteins in five major DNA repair pathways: fail-safe protection against carcinogenesis. Mut Res 511: 145– 178. Bohr VA, Okumoto DS, Hanawalt PC. 1986. Survival of UV-irradiated mammalian cells correlates with efficient DNA repair in an essential gene. Proc Natl Acad Sci 83: 3830– 3833. Bohr VA, Smith CA, Okumoto DS, Hanawalt PC. 1985. DNA repair in an active gene: removal of pyrimidine dimers from the DHFR gene of CHO cells is much more efficient than in the genome overall. Cell 40: 359–369. Burma S, Chen BP, Chen DJ. 2006. Role of non-homologous end joining (NHEJ) in maintaining genomic integrity. DNA Repair 5: 1042– 1048. Burma S, Chen DJ. 2004. Role of DNA-PK in the cellular response to DNA double-strand breaks. DNA Repair 3: 909–918. Cahill D, Connor B, Carney JP. 2006. Mechanisms of eukaryotic DNA double strand break repair. Front Biosci 11: 1958–1976. Caldecott KW. 2007. Mammalian single-strand break repair: mechanisms and links with chromatin. DNA Repair 6: 443–453. Callegari AJ, Kelly TJ. 2007. Shedding light on the DNA damage checkpoint. Cell Cycle 6: 660 –666. Celeste A, Petersen S, Romanienko PJ, Fernandez-Capetillo O, Chen HT, Sedelnikova OA, Reina-San-Martin B, Coppola V, Meffre E, Difilippantonio MJ, et al. 2002. Genomic instability in mice lacking histone H2AX. Science 296: 922– 927. Chen CC, Carson JJ, Feser J, Tamburini B, Zabaronick S, Linger J, Tyler JK. 2008. Acetylated lysine 56 on histone H3 drives chromatin assembly after repair and signals for the completion of repair. Cell 134: 231–243.

Chen Y, Sanchez Y. 2004. Chk1 in the DNA damage response: conserved roles from yeasts to mammals. DNA Repair 3: 1025–1032. Chu G, Chang E. 1988. Xeroderma pigmentosum group E cells lack a nuclear factor that binds to damaged DNA. Science 242: 564–567. Citterio E, Van Den Boom V, Schnitzler G, Kanaar R, Bonte E, Kingston RE, Hoeijmakers JH, Vermeulen W. 2000. ATP-Dependent Chromatin Remodeling by the Cockayne Syndrome B DNA Repair-Transcription-Coupling Factor. Mol Cell Biol 20: 7643–7653. Cremer C, Cremer T, Fukuda M, Nakanishi K. 1980. Detection of laser– UV microirradiation-induced DNA photolesions by immunofluorescent staining. Hum Genet 54: 107 –110. de Jager M, van Noort J, van Gent DC, Dekker C, Kanaar R, Wyman C. 2001. Human Rad50/Mre11 is a flexible complex that can tether DNA ends. Mol Cell 8: 1129–1135. de Laat WL, Appeldoorn E, Sugasawa K, Weterings E, Jaspers NG, Hoeijmakers JH. 1998. DNA-binding polarity of human replication protein A positions nucleases in nucleotide excision repair. Genes Dev 12: 2598– 2609. de Silva IU, McHugh PJ, Clingen PH, Hartley JA. 2000. Defining the roles of nucleotide excision repair and recombination in the repair of DNA interstrand crosslinks in mammalian cells. Mol Cell Biol 20: 7980– 7990. de Waard H, Sonneveld E, de Wit J, Esveldt-van Lange R, Hoeijmakers JH, Vrieling H, van der Horst GT. 2008. Cell-type-specific consequences of nucleotide excision repair deficiencies: Embryonic stem cells versus fibroblasts. DNA Repair 7: 1659– 1669. Dinant C, de Jager M, Essers J, van Cappellen WA, Kanaar R, Houtsmuller AB, Vermeulen W. 2007. Activation of multiple DNA repair pathways by sub-nuclear damage induction methods. J Cell Sci 120: 2731–2740. Dinant C, Houtsmuller AB, Vermeulen W. 2008. Chromatin structure and DNA damage repair. Epig & C 1: 9. Dinant C, Luijsterburg MS, Hofer T, von Bornstaedt G, Vermeulen W, Houtsmuller AB, van Driel R. 2009. Assembly of multiprotein complexes that control genome function. J Cell Biol 185: 21–26. Doil C, Mailand N, Bekker-Jensen S, Menard P, Larsen DH, Pepperkok R, Ellenberg J, Panier S, Durocher D, Bartek J, et al. 2009. RNF168 binds and amplifies ubiquitin conjugates on damaged chromosomes to allow accumulation of repair proteins. Cell 136: 435– 446. Downs JA, Lowndes NF, Jackson SP. 2000. A role for Saccharomyces cerevisiae histone H2A in DNA repair. Nature 408: 1001– 1004. Drapkin R, Reardon JT, Ansari A, Huang JC, Zawel L, Ahn K, Sancar A, Reinberg D. 1994. Dual role of TFIIH in DNA excision repair and in transcription by RNA polymerase II. Nature 368: 769–772. Egly JM. 2001. The 14th Datta Lecture. TFIIH: from transcription to clinic. FEBS Lett 498: 124– 128. El-Khamisy SF, Katyal S, Patel P, Ju L, McKinnon PJ, Caldecott KW. 2009. Synergistic decrease of DNA singlestrand break repair rates in mouse neural cells lacking both Tdp1 and aprataxin. DNA Repair 8: 760–766.

G. Giglia-Mari, A. Zotter, and W. Vermeulen Essers J, Houtsmuller AB, van Veelen L, Paulusma C, Nigg AL, Pastink A, Vermeulen W, Hoeijmakers JH, Kanaar R. 2002. Nuclear dynamics of RAD52 group homologous recombination proteins in response to DNA damage. Embo J 21: 2030– 2037. Falck J, Coates J, Jackson SP. 2005. Conserved modes of recruitment of ATM, ATR and DNA-PKcs to sites of DNA damage. Nature 434: 605– 611. Falck J, Petrini JH, Williams BR, Lukas J, Bartek J. 2002. The DNA damage-dependent intra-S phase checkpoint is regulated by parallel pathways. Nat Genet 30: 290– 294. Fousteri M, Mullenders LH. 2008. Transcription-coupled nucleotide excision repair in mammalian cells: molecular mechanisms and biological effects. Cell Res 18: 73–84. Fousteri M, Vermeulen W, van Zeeland AA, Mullenders LH. 2006. Cockayne syndrome A and B proteins differentially regulate recruitment of chromatin remodeling and repair factors to stalled RNA polymerase II in vivo. Mol Cell 23: 471–482. Friedberg E, Walker G, Siede W, Wood R, Schultz R, Ellenberg T. 2006. DNA Repair and Mutagenesis. ASM Press, Washington, DC. Friedberg EC, Lehmann AR, Fuchs RP. 2005. Trading places: how do DNA polymerases switch during translesion DNA synthesis? Mol Cell 18: 499–505. Giannattasio M, Lazzaro F, Longhese MP, Plevani P, MuziFalconi M. 2004. Physical and functional interactions between nucleotide excision repair and DNA damage checkpoint. Embo J 23: 429–438. Giglia-Mari G, Miquel C, Theil AF, Mari PO, Hoogstraten D, Ng JM, Dinant C, Hoeijmakers JH, Vermeulen W. 2006. Dynamic interaction of TTDAwith TFIIH is stabilized by nucleotide excision repair in living cells. PLoS Biology 4: e156. Giglia-Mari G, Theil AF, Mari PO, Mourgues S, Nonnekens J, Andrieux L, de Wit J, Miquel C, Wijgers N, Maas A, et al. 2009. Differentiation Driven Changes in the Dynamic Organization of Basal Transcription Initiation. PLoS Biology 7: e1000220. Gillet LC, Scharer OD. 2006. Molecular mechanisms of mammalian global genome nucleotide excision repair. Chem Rev 106: 253– 276. Goodarzi AA, Noon AT, Deckbar D, Ziv Y, Shiloh Y, Lobrich M, Jeggo PA. 2008. ATM signaling facilitates repair of DNA double-strand breaks associated with heterochromatin. Mol Cell 31: 167 –177. Green CM, Almouzni G. 2002. When repair meets chromatin. First in series on chromatin dynamics. EMBO Rep 3: 28–33. Groth A, Rocha W, Verreault A, Almouzni G. 2007. Chromatin challenges during DNA replication and repair. Cell 128: 721– 733. Gueven N, Becherel OJ, Kijas AW, Chen P, Howe O, Rudolph JH, Gatti R, Date H, Onodera O, Taucher-Scholz G, et al. 2004. Aprataxin, a novel protein that protects against genotoxic stress. Hum Mol Gen 13: 1081–1093. Hanasoge S, Ljungman M. 2007. H2AX phosphorylation after UV irradiation is triggered by DNA repair intermediates and is mediated by the ATR kinase. Carcinogenesis 28: 2298–2304.

Hanawalt PC. 1994. Transcription-coupled repair and human disease. Science 266: 1957–1958. Harper JW, Elledge SJ. 2007. The DNA damage response: ten years after. Mol Cell 28: 739–745. Hauptner A, Dietzel S, Drexler GA, Reichart P, Krucken R, Cremer T, Friedl AA, Dollinger G. 2004. Microirradiation of cells with energetic heavy ions. Radiat Environ Biophys 42: 237– 245. He H, Lehming N. 2003. Global effects of histone modifications. Brief Funct Genomic Proteomic 2: 234– 243. Hegde ML, Hazra TK, Mitra S. 2008. Early steps in the DNA base excision/single-strand interruption repair pathway in mammalian cells. Cell Res 18: 27–47. Helleday T, Lo J, van Gent DC, Engelward BP. 2007. DNA double-strand break repair: from mechanistic understanding to cancer treatment. DNA Repair 6: 923– 935. Heo K, Kim H, Choi SH, Choi J, Kim K, Gu J, Lieber MR, Yang AS, An W. 2008. FACT-mediated exchange of histone variant H2AX regulated by phosphorylation of H2AX and ADP-ribosylation of Spt16. Mol Cell 30: 86– 97. Hoeijmakers JH. 2001. Genome maintenance mechanisms for preventing cancer. Nature 411: 366–374. Hoeijmakers JH. 2009. DNA damage, aging, and cancer. N Engl J Med 361: 1475– 1485. Hoeijmakers JHJ. 1993. Nucleotide excision repair II: from yeast to mammals. Trends in Gen 9: 211 –217. Hoogstraten D, Bergink S, Ng JM, Verbiest VH, Luijsterburg MS, Geverts B, Raams A, Dinant C, Hoeijmakers JH, Vermeulen W, et al. 2008. Versatile DNA damage detection by the global genome nucleotide excision repair protein XPC. J Cell Sci 121: 2850– 2859. Hoogstraten D, Nigg AL, Heath H, Mullenders LH, van Driel R, Hoeijmakers JH, Vermeulen W, Houtsmuller AB. 2002. Rapid Switching of TFIIH between RNA Polymerase I and II Transcription and DNA Repair In Vivo. Mol Cell 10: 1163–1174. Houtsmuller AB, Rademakers S, Nigg AL, Hoogstraten D, Hoeijmakers JHJ, Vermeulen W. 1999. Action of DNA repair endonuclease ERCC1/XPF in living cells. Science 284: 958–961. Houtsmuller AB, Vermeulen W. 2001. Macromolecular dynamics in living cell nuclei revealed by fluorescence redistribution after photobleaching. Histochem Cell Biol 115: 13–21. Huen MS, Chen J. 2008. The DNA damage response pathways: at the crossroad of protein modifications. Cell Res 18: 8– 16. Huen MS, Grant R, Manke I, Minn K, Yu X, Yaffe MB, Chen J. 2007. RNF8 transduces the DNA-damage signal via histone ubiquitylation and checkpoint protein assembly. Cell 131: 901 –914. Ikura T, Tashiro S, Kakino A, Shima H, Jacob N, Amunugama R, Yoder K, Izumi S, Kuraoka I, Tanaka K, et al. 2007. DNA damage-dependent acetylation and ubiquitination of H2AX enhances chromatin dynamics. Mol Cell Biol 27: 7028– 7040. Jakob B, Scholz M, Taucher-Scholz G. 2003. Biological imaging of heavy charged-particle tracks. Radiat Res 159: 676–684.

DNA Repair Kastan MB, Bartek J. 2004. Cell-cycle checkpoints and cancer. Nature 432: 316– 323. Katsumi S, Kobayashi N, Imoto K, Nakagawa A, Yamashina Y, Muramatsu T, Shirai T, Miyagawa S, Sugiura S, Hanaoka F, et al. 2001. In situ visualization of ultraviolet-light-induced DNA damage repair in locally irradiated human fibroblasts. J Inves Derma 117: 1156– 1161. Keeney S, Eker APM, Brody T, Vermeulen W, Bootsma D, Hoeijmakers JHJ, Linn S. 1994. correction of the DNA repair defect in xeroderma pigmentosum group E by injection of a DNA damage-binding protein. Proc Natl Acad Sci 91: 4053–4056. Kim JS, Krasieva TB, LaMorte V, Taylor AM, Yokomori K. 2002. Specific recruitment of human cohesin to laserinduced DNA damage. J Biol Chem 277: 45149–45153. Lan L, Nakajima S, Oohata Y, Takao M, Okano S, Masutani M, Wilson SH, Yasui A. 2004. In situ analysis of repair processes for oxidative DNA damage in mammalian cells. Proc Natl Acad Sci 101: 13738– 13743. Lanctot C, Cheutin T, Cremer M, Cavalli G, Cremer T. 2007. Dynamic genome architecture in the nuclear space: regulation of gene expression in three dimensions. Nat Rev Genet 8: 104 –115. Lehmann AR. 2006. Translesion synthesis in mammalian cells. Exp Cell Res 312: 2673– 2676. Li B, Carey M, Workman JL. 2007. The role of chromatin during transcription. Cell 128: 707– 719. Li Z, Xiao W, McCormick JJ, Maher VM. 2002. Identification of a protein essential for a major pathway used by human cells to avoid UV- induced DNA damage. Proc Natl Acad Sci 99: 4459– 4464. Limbo O, Chahwan C, Yamada Y, de Bruin RA, Wittenberg C, Russell P. 2007. Ctp1 is a cell-cycle-regulated protein that functions with Mre11 complex to control doublestrand break repair by homologous recombination. Mol Cell 28: 134 –146. Lindahl T. 1993. Instability and decay of the primary structure of DNA. Nature 362: 709– 715. Lippincott-Schwartz J, Snapp E, Kenworthy A. 2001. Studying protein dynamics in living cells. Nature Reviews 2: 444–456. Lisby M, Barlow JH, Burgess RC, Rothstein R. 2004. Choreography of the DNA damage response: spatiotemporal relationships among checkpoint and repair proteins. Cell 118: 699– 713. Lisby M, Rothstein R. 2009. Choreography of recombination proteins during the DNA damage response. DNA Repair 8: 1068– 1076. Liu M, Schatz DG. 2009. Balancing AID and DNA repair during somatic hypermutation. Trends in Imm 30: 173–181. Loyola A, Almouzni G. 2007. Marking histone H3 variants: how, when and why? Trends in Bioch Sci 32: 425 –433. Luijsterburg MS, Dinant C, Lans H, Stap J, Wiernasz E, Lagerwerf S, Warmerdam DO, Lindh M, Brink MC, Dobrucki JW, et al. 2009. Heterochromatin protein 1 is recruited to various types of DNA damage. J Cell Biol 185: 577– 586. Luijsterburg MS, Goedhart J, Moser J, Kool H, Geverts B, Houtsmuller AB, Mullenders LH, Vermeulen W, van

Driel R. 2007. Dynamic in vivo interaction of DDB2 E3 ubiquitin ligase with UV-damaged DNA is independent of damage-recognition protein XPC. J Cell Sci 120: 2706–2716. Luijsterburg MS, von Bornstaedt G, Gourdin AM, Politi AZ, Mone MJ, Warmerdam DO, Goedhart J, Vermeulen W, van Driel R, Hofer T. 2010. Stochastic and reversible assembly of a multiprotein DNA repair complex ensures accurate target site recognition and efficient repair. J Cell Biol 189: 445 –463. Lukas C, Falck J, Bartkova J, Bartek J, Lukas J. 2003. Distinct spatiotemporal dynamics of mammalian checkpoint regulators induced by DNA damage. Nat Cell Biol 5: 255 –260. Mahaney BL, Meek K, Lees-Miller SP. 2009. Repair of ionizing radiation-induced DNA double-strand breaks by non-homologous end-joining. Biochem J 417: 639– 650. Mailand N, Bekker-Jensen S, Faustrup H, Melander F, Bartek J, Lukas C, Lukas J. 2007. RNF8 ubiquitylates histones at DNA double-strand breaks and promotes assembly of repair proteins. Cell 131: 887– 900. Mari P-O, Florea BI, Persengiev SP, Verkaik NS, Bruggenwirth HT, Modesti M, Giglia-Mari G, Bezstarosti K, Demmers JAA, Luider TM, et al. 2006. Dynamic assembly of end-joining complexes requires interaction between Ku70/80 and XRCC4. Proc Natl Acad Sci 103: 18597– 18602. Marteijn JA, Bekker-Jensen S, Mailand N, Lans H, Schwertman P, Gourdin AM, Dantuma NP, Lukas J, Vermeulen W. 2009. Nucleotide Excision Repair-induced H2A ubiquitination is dependent on MDC1 and RNF8 and reveals a universal DNA damage response. J Cell Biol 186: 835 –847. Marti TM, Hefner E, Feeney L, Natale V, Cleaver JE. 2006. H2AX phosphorylation within the G1 phase after UV irradiation depends on nucleotide excision repair and not DNA double-strand breaks. Proc Natl Acad Sci 103: 9891–9896. Masutani C, Sugasawa K, Yanagisawa J, Sonoyama T, Ui M, Enomoto T, Takio K, Tanaka K, van der Spek PJ, Bootsma D, et al. 1994. Purification and cloning of a nucleotide excision repair complex involving the xeroderma pigmentosum group C protein and a human homolog of yeast RAD23. EMBO J 13: 1831–1843. Matsuoka S, Ballif BA, Smogorzewska A, McDonald ER, 3rd Hurov, Luo J, Bakalarski CE, Zhao Z, Solimini N, Lerenthal Y, et al. 2007. ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science 316: 1160– 1166. Meaburn KJ, Misteli T. 2007. Cell biology: chromosome territories. Nature 445: 379– 781. Meaburn KJ, Misteli T, Soutoglou E. 2007. Spatial genome organization in the formation of chromosomal translocations. Semin Cancer Biol 17: 80– 90. Meldrum RA, Botchway SW, Wharton CW, Hirst GJ. 2003. Nanoscale spatial induction of ultraviolet photoproducts in cellular DNA by three-photon near-infrared absorption. EMBO Rep 4: 1144–1149. Mello JA, Sillje HH, Roche DM, Kirschner DB, Nigg EA, Almouzni G. 2002. Human Asf1 and CAF-1 interact and synergize in a repair-coupled nucleosome assembly pathway. EMBO Rep 3: 329–334.

G. Giglia-Mari, A. Zotter, and W. Vermeulen Misteli T, Soutoglou E. 2009. The emerging role of nuclear architecture in DNA repair and genome maintenance. Nature Reviews 10: 243 –254. Mitchell JR, Hoeijmakers JH, Niedernhofer LJ. 2003. Divide and conquer: nucleotide excision repair battles cancer and ageing. Curr Opin Cell Biol 15: 232– 240. Mone MJ, Bernas T, Dinant C, Goedvree FA, Manders EM, Volker M, Houtsmuller AB, Hoeijmakers JH, Vermeulen W, van Driel R. 2004. In vivo dynamics of chromatinassociated complex formation in mammalian nucleotide excision repair. Proc Natl Acad Sci 101: 15933–15937. Mone MJ, Volker M, Nikaido O, Mullenders LH, van Zeeland AA, Verschure PJ, Manders EM, van Driel R. 2001. Local UV-induced DNA damage in cell nuclei results in local transcription inhibition. EMBO Rep 2: 1013– 1017. Moser J, Kool H, Giakzidis I, Caldecott K, Mullenders LH, Fousteri MI. 2007. Sealing of chromosomal DNA nicks during nucleotide excision repair requires XRCC1 and DNA ligase III alpha in a cell-cycle-specific manner. Mol Cell 27: 311 –323. Murga M, Jaco I, Fan Y, Soria R, Martinez-Pastor B, Cuadrado M, Yang SM, Blasco MA, Skoultchi AI, Fernandez-Capetillo O. 2007. Global chromatin compaction limits the strength of the DNA damage response. J Cell Biol 178: 1101– 1108. Nag R, Smerdon MJ. 2009. Altering the chromatin landscape for nucleotide excision repair. Mut Res 682: 13–20. Nelms BE, Maser RS, MacKay JF, Lagally MG, Petrini JH. 1998. In situ visualization of DNA double-strand break repair in human fibroblasts. Science 280: 590– 592. Neves-Costa A, Varga-Weisz P. 2006. The roles of chromatin remodelling factors in replication. Results Probl Cell Differ 41: 91– 107. Nicassio F, Corrado N, Vissers JH, Areces LB, Bergink S, Marteijn JA, Geverts B, Houtsmuller AB, Vermeulen W, Di Fiore PP, et al. 2007. Human USP3 is a chromatin modifier required for S phase progression and genome stability. Curr Biol 17: 1972–1977. Niedernhofer LJ, Essers J, Weeda G, Beverloo B, de Wit J, Muijtjens M, Odijk H, Hoeijmakers JH, Kanaar R. 2001. The structure-specific endonuclease Ercc1-Xpf is required for targeted gene replacement in embryonic stem cells. Embo J 20: 6540– 6549. Niedernhofer LJ, Odijk H, Budzowska M, van Drunen E, Maas A, Theil AF, de Wit J, Jaspers NG, Beverloo HB, Hoeijmakers JH, et al. 2004. The structure-specific endonuclease Ercc1-Xpf is required to resolve DNA interstrand cross-link-induced double-strand breaks. Mol Cell Biol 24: 5776– 5787. Niida H, Nakanishi M. 2006. DNA damage checkpoints in mammals. Mutagenesis 21: 3 –9. Nishi R, Alekseev S, Dinant C, Hoogstraten D, Houtsmuller AB, Hoeijmakers JH, Vermeulen W, Hanaoka F, Sugasawa K. 2009. UV-DDB-dependent regulation of nucleotide excision repair kinetics in living cells. DNA Repair 8: 767–776. O’Donovan A, Davies AA, Moggs JG, West SC, Wood RD. 1994. XPG endonuclease makes the 30 incision in human DNA nucleotide excision repair. Nature 371: 432–435.

O’Driscoll M, Ruiz-Perez VL, Woods CG, Jeggo PA, Goodship JA. 2003. A splicing mutation affecting expression of ataxia-telangiectasia and Rad3-related protein (ATR) results in Seckel syndrome. Nat Genet 33: 497–501. Ogi T, Limsirichaikul S, Overmeer RM, Volker M, Takenaka K, Cloney R, Nakazawa Y, Niimi A, Miki Y, Jaspers NG, et al. 2010. Three DNA polymerases, recruited by different mechanisms, carry out NER repair synthesis in human cells. Mol Cell 37: 714 –727. Panier S, Durocher D. 2009. Regulatory ubiquitylation in response to DNA double-strand breaks. DNA Repair 8: 436 –443. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604 –609. Politi A, Mone MJ, Houtsmuller AB, Hoogstraten D, Vermeulen W, Heinrich R, van Driel R. 2005. Mathematical modeling of nucleotide excision repair reveals efficiency of sequential assembly strategies. Mol Cell 19: 679–690. Polo SE, Roche D, Almouzni G. 2006. New histone incorporation marks sites of UV repair in human cells. Cell 127: 481 –493. Rademakers S, Volker M, Hoogstraten D, Nigg AL, Mone MJ, Van Zeeland AA, Hoeijmakers JH, Houtsmuller AB, Vermeulen W. 2003. Xeroderma pigmentosum group A protein loads as a separate factor onto DNA lesions. Mol Cell Biol 23: 5755– 5767. Reed SH, Gillette TG. 2007. Nucleotide excision repair and the ubiquitin proteasome pathway– do all roads lead to Rome? DNA Repair 6: 149 –156. Rodrigue A, Lafrance M, Gauthier MC, McDonald D, Hendzel M, West SC, Jasin M, Masson JY. 2006. Interplay between human DNA repair proteins at a unique doublestrand break in vivo. EMBO J 25: 222–231. Rogakou EP, Boon C, Redon C, Bonner WM. 1999. Megabase chromatin domains involved in DNA double-strand breaks in vivo. J Cell Biol 146: 905– 916. Rogakou EP, Pilch DR, Orr AH, Ivanova VS, Bonner WM. 1998. DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 273: 5858–5868. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287– 291. Saha A, Wittmeyer J, Cairns BR. 2006. Mechanisms for nucleosome movement by ATP-dependent chromatin remodeling complexes. Results Probl Cell Differ 41: 127 –148. Sartori AA, Lukas C, Coates J, Mistrik M, Fu S, Bartek J, Baer R, Lukas J, Jackson SP. 2007. Human CtIP promotes DNA end resection. Nature 450: 509– 514. Schaeffer L, Roy R, Humbert S, Moncollin V, Vermeulen W, Hoeijmakers JHJ, Chambon P, Egly J. 1993. DNA repair helicase: a component of BTF2 (TFIIH) basic transcription factor. Science 260: 58–63. Scully R, Puget N, Vlasakova K. 2000. DNA polymerase stalling, sister chromatid recombination and the BRCA genes. Oncogene 19: 6176–6183. Sijbers AM, De Laat WL, Ariza RR, Biggerstaff M, Wei YF, Moggs JG, Carter KC, Shell BK, Evans E, De Jong MC, et al. 1996. Xeroderma pigmentosum group F caused

DNA Repair by a defect in a structure-specific DNA repair endonuclease. Cell 86: 811–822. Sinclair DA, Oberdoerffer P. 2009. The ageing epigenome: damaged beyond repair? Age Res Rev 8: 189 –198. Smerdon MJ. 1991. DNA repair and the role of chromatin structure. Curr Opin Cell Biol 3: 422–428. Smits VA, Warmerdam DO, Martin Y, Freire R. 2010. Mechanisms of ATR-mediated checkpoint signalling. Front Biosci 15: 840– 853. Soutoglou E, Dorn JF, Sengupta K, Jasin M, Nussenzweig A, Ried T, Danuser G, Misteli T. 2007. Positional stability of single double-strand breaks in mammalian cells. Nat Cell Biol 9: 675– 682. Soutoglou E, Misteli T. 2008. Activation of the cellular DNA damage response in the absence of DNA lesions. Science 320: 1507–1510. Stewart GS, Panier S, Townsend K, Al-Hakim AK, Kolas NK, Miller ES, Nakada S, Ylanko J, Olivarius S, Mendez M, et al. 2009. The RIDDLE syndrome protein mediates a ubiquitin-dependent signaling cascade at sites of DNA damage. Cell 136: 420– 434. Stiff T, Cerosaletti K, Concannon P, O’Driscoll M, Jeggo PA. 2008. Replication independent ATR signalling leads to G2/M arrest requiring Nbs1, 53BP1 and MDC1. Hum Mol Gen 17: 3247–3253. Sugasawa K, Akagi J, Nishi R, Iwai S, Hanaoka F. 2009. Twostep recognition of DNA damage for mammalian nucleotide excision repair: Directional binding of the XPC complex and DNA strand scanning. Mol Cell 36: 642 –653. Svejstrup JQ, Wang Z, Feaver WJ, Wu X, Bushnell DA, Donahue TF, Friedberg EC, Kornberg RD. 1995. Different forms of TFIIH for transcription and DNA repair: holo-TFIIH and a nucleotide excision repairosome. Cell 80: 21– 28. Takeda S, Nakamura K, Taniguchi Y, Paull TT. 2007. Ctp1/ CtIP and the MRN complex collaborate in the initial steps of homologous recombination. Mol Cell 28: 351 –352. Tashiro S, Walter J, Shinohara A, Kamada N, Cremer T. 2000. Rad51 accumulation at sites of DNA damage and in postreplicative chromatin. J Cell Biol 150: 283– 291. Tibbetts RS, Cortez D, Brumbaugh KM, Scully R, Livingston D, Elledge SJ, Abraham RT. 2000. Functional interactions between BRCA1 and the checkpoint kinase ATR during genotoxic stress. Genes Dev 14: 2989– 3002. Tsien RY, Miyawaki A. 1998. Seeing the machinery of live cells. Science 280: 1954–1955. Uematsu N, Weterings E, Yano K, Morotomi-Yano K, Jakob B, Taucher-Scholz G, Mari PO, van Gent DC, Chen BP, Chen DJ. 2007. Autophosphorylation of DNA-PKCS regulates its dynamics at DNA double-strand breaks. J Cell Biol 177: 219–229. van Attikum H, Fritsch O, Hohn B, Gasser SM. 2004. Recruitment of the INO80 complex by H2A phosphorylation links ATP-dependent chromatin remodeling with DNA double-strand break repair. Cell 119: 777 –788. van Attikum H, Gasser SM. 2009. Crosstalk between histone modifications during the DNA damage response. Trends Cell Biol 19: 207 –217.

van den Boom V, Citterio E, Hoogstraten D, Zotter A, Egly JM, van Cappellen WA, Hoeijmakers JH, Houtsmuller AB, Vermeulen W. 2004. DNA damage stabilizes interaction of CSB with the transcription elongation machinery. J Cell Biol 166: 27–36. van Gent DC, van der Burg M. 2007. Non-homologous endjoining, a sticky affair. Oncogene 26: 7731–7740. Volker M, Mone´ MJ, Karmakar P, Hoffen A, Schul W, Vermeulen W, Hoeijmakers JHJ, van Driel R, Zeeland AA, Mullenders LHF. 2001. Sequential Assembly of the Nucleotide Excision Repair Factors In Vivo. Mol Cell 8: 213 –224. Warmerdam DO, Freire R, Kanaar R, Smits VA. 2009. Cell cycle-dependent processing of DNA lesions controls localization of Rad9 to sites of genotoxic stress. Cell Cycle 8: 1765–1774. Waters R, Teng Y, Yu Y, Yu S, Reed SH. 2009. Tilting at windmills? The nucleotide excision repair of chromosomal DNA. DNA Repair 8: 146– 152. Weber S. 2005. Light-driven enzymatic catalysis of DNA repair: a review of recent biophysical studies on photolyase. Biochim Biophys Acta 1707: 1– 23. Westerveld A, Hoeijmakers JH, van Duin M, de Wit J, Odijk H, Pastink A, Wood RD, Bootsma D. 1984. Molecular cloning of a human DNA repair gene. Nature 310: 425– 429. Weterings E, van Gent DC. 2004. The mechanism of nonhomologous end-joining: a synopsis of synapsis. DNA Repair 3: 1425–1435. White J, Stelzer E. 1999. Photobleaching GFP reveals protein dynamics inside live cells. Trends Cell Biol 9: 61–65. Wyman C, Kanaar R. 2006. DNA double-strand break repair: all’s well that ends well. Annu Rev Genet 40: 363– 383. Wyman C, Ristic D, Kanaar R. 2004. Homologous recombination-mediated double-strand break repair. DNA Repair 3: 827– 833. Yokoi M, Masutani C, Maekawa T, Sugasawa K, Ohkuma Y, Hanaoka F. 2000. The xeroderma pigmentosum group C protein complex XPC-HR23B plays an important role in the recruitment of transcription factor IIH to damaged DNA. J Biol Chem 275: 9870–9875. Zhou BB, Elledge SJ. 2000. The DNA damage response: putting checkpoints in perspective. Nature 408: 433 –439. Zhu XD, Niedernhofer L, Kuster B, Mann M, Hoeijmakers JH, de Lange T. 2003. ERCC1/XPF removes the 3’ overhang from uncapped telomeres and represses formation of telomeric DNA-containing double minute chromosomes. Mol Cell 12: 1489– 1498. Zotter A, Luijsterburg MS, Warmerdam DO, Ibrahim S, Nigg A, van Cappellen WA, Hoeijmakers JH, van Driel R, Vermeulen W, Houtsmuller AB. 2006. Recruitment of the nucleotide excision repair endonuclease XPG to sites of UV-induced dna damage depends on functional TFIIH. Mol Cell Biol 26: 8868–8879. Zou L, Elledge SJ. 2003. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300: 1542– 1548.

RNA Processing and Export Sami Hocine1, Robert H. Singer1, and David Gru¨nwald2 1

Department for Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, New York 10461

2

Department of Bionanoscience, Delft University of Technology

Correspondence: [email protected]

Messenger RNAs undergo 5’ capping, splicing, 3’-end processing, and export before translation in the cytoplasm. It has become clear that these mRNA processing events are tightly coupled and have a profound effect on the fate of the resulting transcript. This processing is represented by modifications of the pre-mRNA and loading of various protein factors. The sum of protein factors that stay with the mRNA as a result of processing is modified over the life of the transcript, conferring significant regulation to its expression.

essenger RNA (mRNA) transcripts are extensively processed before export. 50 capping, splicing, and 30 -end processing represent nuclear processes that are large determinants of the fate of a transcript. As mRNA processing events involve different cellular machinery (Fig. 1), RNA sequences, and have different consequences for target mRNAs, these processes were long seen to be independent of one another. It has become clear over the last decade, however, that these events are integrated and coordinated in space and time (Schroeder et al. 2000; reviewed in Bentley 2002; reviewed in Moore and Proudfoot 2009). Nuclear processing steps require a large set of proteins, many of which are loaded onto the transcript as a result of processing, adding a layer of regulatory information that can affect export, localization, translation, and stability of the transcript. Examples of such proteins include the exon-junction complex (EJC), left behind from splicing, and the THO/TREX complex,

M

loaded during elongation. Indeed, the highly integrated nature of nuclear mRNA processing adds a new level of complexity to our picture of gene regulation. The availability of genetic fluorescent tags and sophisticated microscopy technology adds a dynamic component to this picture, providing spatial and temporal information and highlighting how nuclear structure might regulate gene expression (reviewed in Gorski et al. 2006; reviewed in Moore and Proudfoot 2009). Transcription, the major contributor to RNA biogenesis, takes place under constraints of an anisotropic nuclear landscape that is highly structured (chromatin, distinct nuclear bodies, etc.) and dynamic (gene mobility, diffusive factors, genomic reorganization during cell cycle progression, etc.) (Yao et al. 2008). The availability of increasingly sensitive equipment and fluorescent markers has made it possible to intensively interrogate transcriptional dynamics (Fig. 2) (Becker et al. 2002; Janicki

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000752 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000752

Initiation

S. Hocine, R.H. Singer, and D. Gru¨nwald

Promoter Pol II Transcription factors

CTDP Ser 5

5′ Capping RNA triphosphotase Guanylyltransferase Methyltransferase

Elongation

Cap binding protein

Ser 2

Splicing Splice sites Splicesome/U snRNPs

Termination

EJC SR proteins hnRNP proteins THO/TREX Export factors

3′ End-processing Cleavage sites Cleavage/Poly(A) factors Poly(A) polymerase Poly(A) binding protein

Release & export Nuclear pores Export factors

Figure 1. mRNA processing is tightly coupled to tran-

model for transcription complex recruitment (Gorski et al. 2008; reviewed in Darzacq et al. 2009). Modulation of transcriptional speed and processivity is suggested to be a way of regulating gene expression (Darzacq et al. 2007; reviewed in Core and Lis 2008; Core et al. 2008). These techniques have revealed highly inefficient transcription by RNA polymerase II (Pol II) and stochastic assembly of transcription complexes (Darzacq et al. 2007; Gorski et al. 2008), and both assembly and processivity can be regarded as rate limiting steps of in vivo transcription (reviewed in Core and Lis 2008). A novel way to look at gene expression has been recently presented by analyzing transcript amounts in individual cells on the single molecule level. Here the mean expression level and its variation are accessible, leading to a detailed understanding of variability in gene expression within a population (Zenklusen et al. 2008). Time lapse experiments add further information concerning the expression mode of individual genes, providing insights into differences between constitutive expression and bursts in different species (reviewed in Larson et al. 2009). These experimental approaches allow for in depth characterization of transcriptional dynamics (reviewed in Darzacq et al. 2009; reviewed in Larson et al. 2009), and are likely to provide greater insight into downstream processes.

scription. The phosphorylation state of the C-terminal domain (CTD) of RNA Polymerase II (Pol II) is given on the right in relation to major steps of transcription. Listed are functional processing sequences (red), components of processing machinery (blue) and factors that are loaded onto the transcript as a result of processing (green).

TRANSCRIPTION SETS THE STAGE FOR PROCESSING

et al. 2004; reviewed in Shav-Tal et al. 2004b; Darzacq et al. 2007; Yao et al. 2007). Such approaches provide a new perspective on mRNA metabolism that has been mostly based on biochemical data. For instance, whereas preformed transcription complexes are sufficient for in vitro transcription, live cell imaging experiments show that different transcription factors show a wide range of dwell times at the promoter and suggest a link between transcription complex assembly dynamics and transcriptional output, consistent with a subunit assembly

Transcription, particularly the carboxy-terminal domain (CTD) of RNA Pol II, contributes significantly to the integration of nuclear mRNA processing (Fig. 3). The CTD is an essential domain of the largest RNA Pol II subunit, composed of conserved YSPTSPS heptad repeats that are subject to reversible phosphorylation. It is well established that the CTD functions in transcription, and it has an equally important role in mRNA processing. The CTD interacts with a large number of protein factors, and among the protein domains shown to show preferential CTD binding are: CTD interacting domains (CIDs), WW domains,

RNA Processing and Export

A

YSP P PTS PS

Pol II

P YS P

YSP P PTS PS

P TSP

S

256 lac O

Pol II

Pol II

Promoter

YSP P PTS PS

Pol II

YSP P PTS PS

Pol II

YSP P PTS PS

24 MS2 repeats

CFP-SKL

YSP P PTS PS

Pol II

Pol II

Intron Intron

5′

3′

Pol II elongation (3.3 kb)

B

C

D

MS2 labeled RNA transcription (2.3 kb)

N RFP-LacI

E

P P YSPTSPS

CFP-Lacl

H

P P YSPTSPS

GFP-MS2

S

CFP-Lacl

P P YSPTSPS

GFP-MS2

3′ Cy3

X

W

Merge

Y

Merge

H5

L

M

CFP-LacI RFP-LacI

Merge

U

J V

K

5′ Cy3

T

Merge

H14

I

RFP-LacI

Q

G R

RFP-LacI

P

Merge

V15

F

O

YFP–Pol II

Exon CY3

Intron Cy5

Merge

Merge

Figure 2. In vivo detection of transcription using fluorescence microscopy. Schematic shows a reporter cassette

that is integrated as gene arrays into the genome. RFP-LacI labels reveal array locus and immunofluorescence shows that RNA Pol II with three distinct phosphorylation states is recruited to these active transcription sites (B – M). Similarly, the gene array, visualized by CFP-LacI, colocalizes with both MS2-tagged mRNAs, seen by GFP, and mRNA FISH probes targeting exonic and intronic regions (N – Y). (Reprinted, with permission, from Darzacq et al. 2007 [# 2007 Macmillan Publishers, Ltd.].)

and FF domains (Verdecia et al. 2000; Smith et al. 2004; Noble et al. 2005). Serine 5 phosphorylation of the CTD occurs when RNA Pol II is at the 50 end of the gene and is mediated by the TFIIH-associated kinase CDK7 (Kin28 in yeast) as transcription initiates. Serine 2 phosphorylation is mediated by PTEFb (CTDK1 in yeast) as the processive RNA Pol II elongates through the body of the gene (Komarnitsky et al. 2000; Peterlin and Price 2006). These phosphorylation marks are critical to the proper progression of transcription and are required for coordinating processing events

(reviewed in Hirose and Manley 2000). Indeed, the CTD has been shown to bind over 100 different yeast proteins in its phosphorylated state (Phatnani et al. 2004). It can adopt different conformations depending on phosphorylation patterns, protein interactions, and proline isomerization via peptidyl-prolyl cis/trans isomerases (PPIases) (reviewed in Hirose and Ohkuma 2007). In this way, the CTD functions as a recruitment scaffold for different processing factors throughout transcription, thereby integrating processing events in time. Furthermore, changes in chromatin structure on

S. Hocine, R.H. Singer, and D. Gru¨nwald Poised polymerase

Promoter

Ser5P

Early elongation

H3K4Me3

H3K36Me3

Promoter

Ser5P

Ser2P

Cap

Elongation coupled to splicing Promoter

Splicing factors

Intron

Intron Cotranscriptionally recruited splicing factors

Figure 3. Depiction of several events that coincide with the switch from a promoter-engaged “poised” RNA pol-

ymerase to an elongating productive RNA polymerase (shown in purple). Histone modifications and changing CTD phosphorylation states are known to be associated with this switch. Furthermore, transcription is intimately coupled to mRNA processing events such as 50 capping and splicing. (Reprinted, with permission, from Moore and Proudfoot 2009 [# 2009, Elsevier].)

transcriptional activation are likely to contribute to mRNA processing through gene positioning, or relocation at or near nuclear pores, and gene looping, or formation of DNA loops in which 50 and 30 ends make contact (reviewed in Moore and Proudfoot 2009). Recent evidence from studies using galactose genes in yeast showed that gene positioning can be a further regulatory step, and an interaction has been shown between transcription-dependent complexes (e.g TREX/TREX-2), chromatin remodeling complexes (e.g. SAGA), and nuclear pores (reviewed in Blobel 1985; Cabal et al. 2006; Klockner et al. 2009; reviewed in Moore and Proudfoot 2009). Export however, is likely to depend on successful maturation of the mRNA, and whereas in yeast gene gating to nuclear pores is suggested, mRNAs of other genes reach nuclear pores in a diffusive manner. Studies in mammalian cell systems have not supported gene gating, as fluorescence correlation spectroscopy (FCS), FRAP and single particle tracking consistently suggest diffusive behavior (Politz et al. 1998; Shav-Tal et al.

2004a; Grunwald et al. 2006; Politz et al. 2006; Braga et al. 2007; Siebrasse et al. 2008). In addition, high resolution studies of the nuclear periphery led to the conclusion that DNA is absent at the nuclear basket of nuclear pores (Schermelleh et al. 2008). Thus, transcription, and the associated machinery, serves to localize processing factors to the appropriate place on the nascent mRNA, stimulate and coordinate processing and possibly establish functionally significant chromatin conformations. CAPPING

The first processing event an mRNA undergoes is 50 -end capping which requires three enzymatic activities: RNA triphosphatase, guanylyltransferase and 7-methyltransferase (reviewed in Shuman 2001). Occurring early in transcription after RNA Pol II has transcribed the first 25– 30 nucleotides, the RNA triphosphatase first acts on the terminal nucleotide to remove the g-phosphate. The guanylyltransferase then transfers GMP from GTP to form

RNA Processing and Export

GpppN, which is subsequently methylated. In mammals, a bifunctional capping enzyme includes both amino-terminal RNA triphosphate activity and carboxy-terminal guanylyltransferase activity. In yeast, the RNA triphosphatase (Cet1) and separate guanylyltransferase (Ceg1) form a heterodimeric capping enzyme. In both cases, a separate methyltransferase (Abd1 in yeast) is required to methylate the guanine at the N7 position.

to drugs that disrupt elongation and show decreased transcription through promoter-proximal pause sites (Kim et al. 2004a). Furthermore, yeast capping enzyme subunits influence RNA Pol II occupancy at the 50 end and may regulate transcription reinitiation (Myers et al. 2002; Schroeder et al. 2004). Taken together, it seems that transcription complexes are held at the promoter until capping occurs, after which the polymerase switches into an elongating mode.

TRANSCRIPTION AND CAPPING ARE TIGHTLY COUPLED

CONSEQUENCES OF SUCCESSFUL CAPPING

Mammalian capping enzyme binds directly to the elongating RNA Pol II with a phosphorylated CTD (termed RNA Pol II0) through its guanylyltransferase domain (Yue et al. 1997), thereby coupling capping to the early stage of transcription. Yeast capping enzyme subunits also bind directly and independently to RNA Pol II0, and this interaction is dependent on Kin28, the subunit of TFIIH responsible for serine 5 phosphorylation (Rodriguez et al. 2000). The loss of serine 5 phosphorylation during transcription correlates with the release of the capping enzyme, which is believed to occur before the nascent transcript is 500 nucleotides long (reviewed in Zorio and Bentley 2004). Ceg1 guanylylation activity is inhibited by the phosphorylated CTD but is restored and enhanced by Cet1, and this allosteric regulation may represent a means to temporally coordinate guanylylation and triphosphatase activities (Cho et al. 1998; Ho et al. 1998). In mammals, CTD binding to the guanylyltransferase has an allosteric affect, causing a twofold increase in affinity of guanylyltransferase for GTP (Ho and Shuman 1999). CTD phosphorylation that accompanies the transition from initiation to elongation has a clear impact on capping and allows communication between the transcriptional machinery and capping enzymes. It seems that capping also has a direct impact on transcription. Recent evidence indicates that capping enzymes can relieve transcriptional repression, suggesting an additional role in promoter clearance (Mandal et al. 2004). Temperature sensitive ceg1 yeast mutants are sensitive

Capping of transcripts confers stability. In yeast as well as mammals, capping helps protect the transcript from 50 !30 exonucleases present in both the nucleus and cytoplasm (Hsu and Stevens 1993; Walther et al. 1998). The 50 !30 degradation pathway involves deadenylation followed by rapid decapping by Dcp1/Dcp2, and degradation by the processive 50 !30 exonuclease, Xrn1 (Hsu and Stevens 1993; Muhlrad et al. 1995). The cap is also important in mediating mRNA recruitment to ribosomes. The protein complex eIF4F recognizes the cap before translation, and facilitates circularization of mRNAs via an interaction with polyA-binding protein (PAB1), thereby aiding in translation reinitiation and enhancing protein synthesis (Tarun and Sachs 1996; Wakiyama et al. 2000). Transcripts engaged in translation are protected from degradation suggesting competition between translation and degradation (reviewed in Jacobson and Peltz 1996). Finally, depletion of CBC (cap-binding complex) from HeLa cell extracts inhibits the endonucleolytic cleavage step of 30 -end formation, reduces the stability of poly(A) cleavage complexes, and disrupts communication between 50 and 30 ends (Flaherty et al. 1997). SPLICING

The precise removal of noncoding intervening sequences, or introns, from many pre-mRNAs is a required process for proper protein expression. In both yeast and mammals, this reaction is catalyzed by the spliceosome, consisting of

S. Hocine, R.H. Singer, and D. Gru¨nwald

the U1, U2, U4, U5 and U6 small nuclear RNPs (snRNPs) in conjunction with a large number of additional proteins (reviewed in Stark and Luhrmann 2006). Within the spliceosome, a series of RNA– RNA, RNA– protein, and protein – protein interactions is needed to identify and remove intronic regions and join exons, producing a mature transcript (reviewed in Collins and Guthrie 2000). The mature spliceosome carries out splicing through two transesterification reactions. First, the 20 -OH of a branch point nucleotide performs a nucleolytic attack on the first nucleotide of the intron, forming a lariat intermediate. Second, the 30 OH from the free exon performs a nucleolytic attack on the last nucleotide of the intron, thereby joining exons and releasing the lariat intron. Intron identification relies on certain sequences, including the 50 splice site, branch point (and downstream polypyrimidine tract) and 30 splice site. In yeast, splice sites are easily identified, and although only 3% of genes contain single short introns, they account for more than 25% of cellular mRNAs (Ares et al. 1999; Lopez and Seraphin 1999; reviewed in Barrass and Beggs 2003). In mammals, however, splice sites are less clear, and many genes contains multiple introns that vary from a few hundred to hundreds of thousands of nucleotides (Lander et al. 2001). The presence of putative splice sites in higher eukaryotes does not necessarily lead to selection of these sites by the spliceosome. Flanking pre-mRNA regulatory elements, including intronic and exonic splicing enhancers or silencers, bind trans-acting regulatory factors that enhance or repress snRNP recruitment to splice sites. Generally, exonic splicing enhancers are bound by Serine/Arginine-rich (SR) proteins, whereas exonic splicing silencers are bound by heterogenous nuclear ribonucleoprotein (hnRNP) proteins (reviewed in Cartegni et al. 2002; reviewed in Singh and Valcarcel 2005). Therefore in higher eukaryotes, it is the cumulative effect of multiple factors that modulates splice site selection. As a result, 92%–94% of human transcripts are subject to alternative splicing, representing an important source of diversity in gene expression with serious implications for health and disease

(reviewed in Nissim-Rafinia and Kerem 2005; Wang et al. 2008). Conversely, in yeast, SR proteins do not appear to have a significant role in splicing, consistent with the absence of exonic splicing enhancers (reviewed in Wahl et al. 2009). The spliceosome is rich in proteins, containing approximately 125 different proteins (more than two-thirds of its mass), and spliceosome assembly is characterized by a remarkable exchange of components from one step to the next (reviewed in Wahl et al. 2009). Although RNA-RNA base pairing interactions are critical to the precise recognition of splice sites, they are generally weak and require additional proteins for enhanced stabilization. DExD/H-type RNAdependent ATPases/helicases have long been implicated in rearrangements within the spliceosome, and many are conserved between yeast and humans. These proteins act at discrete stages of splicing including single-strand RNA translocation, strand annealing, and protein displacement (reviewed in Pyle 2008). Human spliceosomes also contain several PPIases that are absent in yeast, though the role of these proteins in splicing is not well understood. Similar to DExD/H-type RNA-dependent ATPases/ helicases, they are recruited at discrete stages of splicing and are thought to be involved in at least one protein conformational switching event (reviewed in Wahl et al. 2009). Furthermore, post-translational modification of splicing factors and spliceosomal proteins may act as switches to allow fine tuning of the spliceosome (Bellare et al. 2008; reviewed in Wahl et al. 2009). The nature of interactions during such a tightly regulated protein-rich process is not very well documented and may be best studied using in vivo imaging techniques. For example, one recent study (Fig. 4) employed FRET-FLIM and revealed for the first time that different complexes of splicing factors show differential distributions in live cell nuclei (Ellis et al. 2008). SPLICING LEAVES A MARK

Mature mRNAs are occupied by a number of different proteins that determine their fate in many ways, and several of these associations are

RNA Processing and Export

A

EGFP-U1 70K

mCherry-C1

EGFP-U1 70K

mCherry-SF2/ASF

τm lifetime (ps)

2300 5

2125

τm lifetime (ps)

DRB, 25 μg/ml, 2 h

EGFP-U1 70K

τm lifetime (ps)

mCherry-C1

τm lifetime (ps)

mCherry-SF2/ASF

2125

70

FRET %

15 FRET %

FRET amplitude %

15 20

80

ρ = 0.81

C 14 12 FRET efficiency %

2250 5

FRET amplitude %

12.5 30

2250 5

2125 EGFP-U1 70K

12.5 FRET %

2300 5

2125

B

FRET %

ρ = 0.66 *ρ = 0.018

10 8 6 4 2 0

K K SF K 35 K SF K SF K SF K SF m) K SF 70 70 2/A 70 AF 70 2/A 70 2/A s) 70 2/A 70 2/A las 70 2/A 1 1 1 1 1 2 F F U C1 U U U SF le U1 F op U1 SF ) U1 SF -U -S -U -S P- rryP- rryP- rry- eck FP- ry-S cle P- rry- asm FP- ryF FP erry s) F FP erry F F r u e p l e e e er EG Ch EG Ch ckle EG Ch EG Ch EG Ch B (s EG Che B (n EG Ch leop EG Ch B m m pe m m m DR m uc m m R R (s + (n +D +D

Figure 4. Spatial mapping of the interaction of U1 70K with SF2/ASF in vivo. Shown are confocal images of cells transfected with EGFP-U1 70K and cotransfected with either mCherry-C1 or mCherry-SF2/ASF. Mean fluorescence lifetime (in picoseconds) and percentages of FRET efficiency and FRET amplitude are shown ( pseudocolor) in these same cells (A). The same experiment was repeated in the presence of 25 mg/ml DBR for 2 hours before imaging (B). FRET efficiencies calculated from FLIM measurements for the interaction of SF2/ ASF with U1 70K (C). # Ellis et al., 2008. J. Cell Biol. 181: 921–934.

S. Hocine, R.H. Singer, and D. Gru¨nwald

splice-dependent. As mentioned, various proteins associate cotranscriptionally and accompany the packaged mRNA into the cytoplasm where they can direct localization, translation and decay. Shuttling SR proteins specifically serve as mRNP binding sites for export factors, and the phosphorylation state of these proteins confers export competence (reviewed in Huang and Steitz 2005; reviewed in Moore and Proudfoot 2009). The THO/TREX complex, which functions in transcription and export, associates with spliced mRNAs at the 50 -most exon (Cheng et al. 2006). THO/TREX recruitment is enhanced by splicing, and promotes rapid export (Valencia et al. 2008). In mammals, REF/Aly and UAP56 (homologs of yeast Yra1 and Sub2), are recruited as a consequence of splicing and have a role in aiding export. Perhaps the most studied splice-dependent mark is the exon junction complex, or EJC. EJCs are stably deposited 20 – 24 nucleotides upstream of exon-exon junctions late in splicing (Le Hir et al. 2000). Interestingly, spliced mRNAs appear to have greater translational efficiency than their cDNA counterparts (reviewed in Le Hir et al. 2003). Aside from their role in nonsense mediated decay, EJCs appear to directly enhance translation initiation. Although there are several proposed mechanisms by which EJCs do this, they ultimately serve to promote the pioneer round of translation (reviewed in Moore and Proudfoot 2009). Finally, a number of DEAD-box proteins have recently been identified as associating with mRNAs in a splicedependent manner, and these are believed to influence many aspects of mRNA metabolism (Merz et al. 2007; reviewed in Rosner and Rinkevich 2007). Taken together, it is clear that spliced mRNAs carry with them numerous protein marks that indicate their splicing history and have important downstream effects. LINKS BETWEEN TRANSCRIPTION AND SPLICING

Initial work focusing on the link between transcription and splicing suggested that splicing occurs cotranscriptionally and factors involved in splicing colocalize with transcription sites

(Beyer and Osheim 1988; Zhang et al. 1994). Similar to capping, the CTD of RNA Pol II has an important role in splicing. CTD truncation causes inefficient splicing in mammalian cells and inhibition of colocalization of splicing factors with transcription sites (McCracken et al. 1997; Misteli and Spector 1999). Furthermore, RNA Pol II0 has been shown to physically associate with splicing factors that do not bind RNA Pol IIA (RNA Pol II with an unphosphorylated CTD), suggesting this interaction depends on the phosphorylation state of the CTD. Both anti-CTD antibodies and CTD peptides can inhibit splicing in vitro, and expression of phosphorylated CTD peptides has a similar effect on mammalian cells in vivo (Yuryev et al. 1996; Du and Warren 1997). RNA Pol II0 enhances splicing in vitro, whereas RNA Pol IIA has an inhibitory effect (Hirose et al. 1999). This is believed to result from RNA Pol II0-dependent stimulation of the early steps of spliceosome assembly, possibly by facilitating the binding of snRNPs to the nascent transcript. Therefore, it seems that CTD phosphorylation can act to recruit splicing factors to the nascent transcript to ensure rapid and accurate splicing. Transcription is also linked to splicing in ways independent of the CTD. Promoter identity and expression levels of certain SR proteins are known to affect alternative splicing (Cramer et al. 1997). Furthermore, in both yeast and mammals, disruption of RNA Pol II elongation markedly shifts the balance of alternatively spliced isoforms (de la Mata et al. 2003; Howe et al. 2003). This is consistent with a “first come first served” model in which elongation rate regulates splice site selection, as 50 splice sites are more likely to pair with newly transcribed 30 splice sites. Additionally, the transcriptional coactivator, p52, is known to interact with SF2/ASF and stimulates splicing (Ge et al. 1998), suggesting that transcriptional machinery can modulate splicing factor recruitment. Finally, splicing can also have an impact on transcription. The presence of introns can confer increased transcriptional efficiency, possibly through increase initiation rates (Brinster et al. 1988). Recruitment of snRNPs by TAT-SF1, an elongation factor that associates with P-TEFb,

RNA Processing and Export

enhances elongation and this effect is dependent on the presence of functional splice signals (Fong and Zhou 2001; Kameoka et al. 2004). Reminiscent of the effect of capping on transcription, certain splicing factors have also been shown to promote elongation (Lin et al. 2008). LINKS BETWEEN CAPPING AND SPLICING

Capping also has a role in splicing. The 50 cap structure increases splicing efficiency in mammalian cell extracts and in vivo (Konarska et al. 1984; Inoue et al. 1989). Depletion of CBC in cell extracts prevents spliceosome assembly at an early step in complex formation (Izaurralde et al. 1994). In yeast, CBC interacts with components of the earliest identified splicing complexes (Colot et al. 1996). Similarly, in mammals, successful capping and CBC recruitment is implicated in U1 small nuclear ribonucleoproteins (snRNP) recognition of the 50 splice site, but this effect is specific to mRNAs with only one intron (Lewis et al. 1996). QUALITY CONTROL OF SPLICING

Splicing defects can lead to potentially harmful protein variants. mRNAs are subject to quality control in the nucleus resulting in the prevention of export of splicing-defective transcripts. To date, several mutations in cis (conserved splice sites) and in trans (spliceosome components required for 1st or 2nd step catalysis) yield drastic reductions in mature mRNA without a corresponding increase in unspliced premRNAs (reviewed in Staley and Guthrie 1998; Bousquet-Antonelli et al. 2000). Mature mRNA levels are largely restored in these same mutants when degradation is inhibited, suggesting that spliceosomes are able to act successfully on these substrates if they are not quickly destroyed. Thus, quality control acts on a number of different splice-defective pre-mRNAs, and degradation is in direct competition with productive splicing. This sort of kinetic competition is exemplified by the finding that decreased ATP hydrolysis of the DExD/Htype RNA-dependent ATPases, Prp16, leads to

productive splicing of pre-mRNAs harboring mutant branch points that would normally be discarded (Burgess and Guthrie 1993). This same principal has been extended to other splice site mutations and members of the DExD/H-type RNA-dependent ATPases/helicases family (Mayas et al. 2006). By coupling spliceosomal rearrangements with irreversible ATP hydrolysis, these proteins ensure that splice-aberrant mRNAs which are unable to complete splicing within a time frame dictated by ATP hydrolysis rates, are discarded (reviewed in Villa et al. 2008). Quality control takes place in the nucleus, can act at numerous stages in the splicing process and serves to commit unspliced mRNAs to degradation pathways. One such pathway involves both 30 ! 50 degradation by the exosome and 50 ! 30 degradation by the nuclear exonuclease Rat1 (BousquetAntonelli et al. 2000). Other degradation pathways include Dbr1-mediated debranching of aberrant lariat-intermediates followed by export and cytoplasmic degradation, and Rnt1mediated endonucleolytic cleavage of unspliced pre-mRNAs followed by nuclear degradation (Danin-Kreiselman et al. 2003; Hilleren and Parker 2003). An additional quality control mechanism involves spliceosome-dependent nuclear retention of unspliced transcripts. A number of different proteins seem to be involved in anchoring these transcripts at the nuclear side of the nuclear pore complex, and retention may be regulated by desumoylation (reviewed in Dziembowski et al. 2004; Galy et al. 2004; Palancade et al. 2005; Lewis et al. 2007). Furthermore, splice-defective mRNAs are also known to be retained at the site of transcription (Custodio et al. 1999; reviewed in Custodio and Carmo-Fonseca 2001). Thus, the cell has evolved various ways to ensure that unspliced transcripts do not leak into the cytoplasm. Finally, nonsense mediated decay (NMD) represents a highly specific form of quality control that extends to higher eukaryotes. The presence of an EJC downstream of a stop codon triggers degradation of translating mRNAs (reviewed in Stalder and Muhlemann 2008). Splice-dependent EJC deposition can increase the translational efficiency of normal

S. Hocine, R.H. Singer, and D. Gru¨nwald

mRNAs while ensuring rapid degradation of aberrant mRNAs.

3’-END PROCESSING

The final step of transcription is endonucleolytic cleavage which occurs 10 – 30 nucleotides downstream of a signal sequence (conserved AAUAAA sequence in mammals or an AU-rich sequence in yeast), followed by poly(A) addition at the 30 end (reviewed in Proudfoot 2004). The poly(A) tail is similar to the cap in that it is important for the stability and translational efficiency of the mRNA (Drummond et al. 1985). Cleavage requires multiple proteins, including cleavage/polyadenylation specificity factor (CPSF), cleavage stimulation factor (CstF), and two cleavage factors (CFIm and CFIIm) in mammals, or cleavage-polyadenylation factor (CPF) and two cleavage factors (CF1A and CF1B) in yeast. Poly(A) polymerase (PAP) then adds the poly(A) tail to the 30 -OH that is exposed on cleavage (reviewed in Proudfoot 2004). CPSF and CstF are highly conserved between yeast and humans and are required for both cleavage and polyadenylation (reviewed in Shatkin and Manley 2000). CPSF recognizes RNA and facilitates PAP recruitment. The endonuclease responsible for cleavage in mammals is CPSF-73 and Ydh1 in yeast (Ryan et al. 2004; Mandel et al. 2006). CstF recognizes U/ GU-rich elements found in the mRNA and is directly involved in polyadenylation. The conserved AAUAAA sequence and downstream U/GU site comprise the core poly(A) element, although additional auxiliary elements can influence polyadenylation efficiency (Gil and Proudfoot 1984; Sadofsky and Alwine 1984; Russnak 1991; Bagga et al. 1995). Cleavage is closely coupled to poly(A) tail synthesis, which also requires PAB1. A number of other factors also participate in 30 -end processing, and many do not share homologs in other systems (reviewed in Shatkin and Manley 2000). As with splicing, transcripts can be alternatively polyadenylated, thereby altering stability, localization or transport. It is estimated that

more than half of the genes in the human genome are subject to such alternative 30 -end processing, generating isoforms that differ in 30 UTR length or encoding different proteins altogether (Tian et al. 2005). Alternative polyadenylation can be tissue specific, may be coupled to alternative splicing and can have implications for health and disease (Peterson and Perry 1989; Beaudoing and Gautheret 2001; Tian et al. 2005; Lu et al. 2007; reviewed in Danckwardt et al. 2008). In fact, many human genes contain multiple potential 30 -end cleavage sites, and appropriate site selection is achieved by alternate mechanisms, representing an additional layer of complexity in the regulation of gene expression (reviewed in Wilusz and Spector 2010). It is important to note that the mechanism and machinery responsible for alternative polyadenylation remain unclear. TRANSCRIPTION AND 30 -END PROCESSING INFLUENCE EACH OTHER

Many factors involved in 30 -end processing have been shown to interact with the CTD, including CstF subunits. Purified CTD can stimulate the cleavage step and is needed for processing in reconstituted reactions (Hirose and Manley 1998). As with 50 capping, the specific CTD phosphorylation pattern is important in 30 end processing. In this case, loss of serine 2 phosphorylation in Ctk1 (yeast) or Cdk9 (Drosophila) mutants leads to a defect in 30 -end processing, likely resulting from improper or inefficient recruitment of processing factors (reviewed in Hirose and Ohkuma 2007). Furthermore, several yeast proteins involved in 30 end processing preferentially bind the CTD phosphorylated at serine 2, which may ensure processing occurs as the polymerase reaches the end of a gene. In mammalian cells, unlike yeast, the CTD is also required for cleavage (Licatalosi et al. 2002). Once again, the CTD appears to mediate coupling between transcription and 30 -end processing. Although transcription elongation continues for quite some distance after the poly(A) signal, transcription termination and 30 -end

RNA Processing and Export

processing are intimately coupled. This was supported by the finding that termination requires functional poly(A) signals (reviewed in Hirose and Manley 2000). There are multiple ways in which 30 -end processing may be coupled to transcription termination: † In the antiterminator model, extrusion of the

poly(A) signal from the polymerase triggers a change in the factors associated, possibly releasing elongation factors or recruiting termination factors (reviewed in Buratowski 2005). The PAF and TREX complexes represent good candidates for such a model, and have been shown to cross-link throughout the gene up to the poly(A) signal, in which they are largely absent (Kim et al. 2004b). The absence of these and other factors may switch the polymerase into a nonprocessive mode, releasing it from the DNA template.

snoRNAs (reviewed in Proudfoot et al. 2002). As described, CTD phosphorylation patterns are linked to processing events, and Ssu72, in conjunction with Pta1 (another component of yeast CPF), also appears to have phosphatase activity specific for serine 5 (Krishnamurthy et al. 2004). Additional genetic and physical interactions have been described between the transcriptional machinery (TFIIB and Sub1) and the 30 -end processing factors Ssu72 and Rna15 (Sun and Hampsey 1996; Wu et al. 1999; Calvo and Manley 2005). Interactions between factors located on opposite ends of genes is likely facilitated by the formation of gene loops, and TFIIB, Ssu72 and Pta1 all appear to have a role in this (Singh and Hampsey 2007). Overall, it seems that the transcriptional machinery, DNA template, nascent mRNA and 30 -end processing machinery are in constant communication for processing.

† The torpedo model of termination has

gained recent support in both yeast and mammals (reviewed in Buratowski 2005). According to this model, the 50 ! 30 exonucleases Rat1 (Xrn2 in mammals) is recruited to the 30 end of the mRNA, in which it interacts with Rtt103, which is known to contact the CTD phosphorylated at serine 2. Rat1/ Xrn1 degrades the downstream product, eventually catching up with and triggering the release of RNA Pol II (Kim et al. 2004c; West et al. 2004). Transcriptional pause sites positioned downstream of the poly(A) signal also seem to be important for 30 -end processing and termination of a number of mammalian genes, reestablishing the theme of kinetic coupling between transcription and processing (Yonaha and Proudfoot 2000). Although 30 -end processing machinery is enriched at the 30 end of genes, certain factors can be found at or near promoters toward the 50 end. For example, CPSF can be recruited to promoters through an association with TFIID (Dantonel et al. 1997). Ssu72, a component of yeast CPF, seems to function at many stages, including transcription initiation, 30 -end processing and termination of certain mRNAs and

QUALITY CONTROL OF 30 -END PROCESSING

Evidence in yeast shows that transcripts undergoing aberrant 30 -end processing are disposed of. Defects associated with pap1-1 mutants are suppressed by deletion of the exosomal subunit Rrp6, which is known to interact with both PAP and the export factor Npl3 (Burkard and Butler 2000). Hypoadenylated mRNAs are retained in the nucleus at the site of transcription, and these transcripts are stabilized and exported in the absence of Rrp6 (Hilleren et al. 2001). In rna14 and rna15 mutant strains, defects in termination lead to readthrough transcripts and aberrant polyadenylation. Deletion of Rrp6 stabilizes the aberrantly polyadenylated population whereas depletion of Rrp41 stabilizes the population of long readthrough transcripts (Libri et al. 2002; Torchet et al. 2002). Therefore, different components of the exosome may have evolved specialized roles in mRNA surveillance, ensuring rapid degradation of transcripts that possess aberrant 30 ends. Additionally, THO complex or sub2 mutants show defects in 30 end formation, reduced mRNA levels and retention at the site of transcription. Codeletion of Rrp6 and TRAMP components restores mRNA

S. Hocine, R.H. Singer, and D. Gru¨nwald

levels, although the retention effect only requires Rrp6 (Libri et al. 2002; Rougemaille et al. 2007). These observations may suggest kinetic competition between 30 end formation and degradation. Transcripts that do not quickly and efficiently undergo 30 -end processing are exposed to the exosome, and mutation of exosome components may allow more time for defective 30 -end processing machinery to function. Finally, recent evidence suggests that nuclear mRNP assembly factors are involved in releasing the 30 -end processing machinery from the transcript after polyadenylation (Qu et al. 2009). This may be a way to temporally coordinate 30 -end formation, mRNP maturation and export. EXPORT OF mRNA

Transport through the nuclear pore complex (NPC) represents the link between the nucleus and cytoplasm. Several studies have investigated mRNA mobility in the nucleoplasm and have revealed probabilistic movement of mRNAs with diffusion coefficients between 0.03 mm2/s and 4 mm2/s (Politz et al. 1998; Shav-Tal et al. 2004a; Braga et al. 2007; Siebrasse et al. 2008). In pulse-chase experiments, mRNA was found in the cytoplasm within 20 min after transcription (Lewin 1980). However, singlemolecule tracking experiments suggest that transit through the NPC is significantly faster, on the order of fractions of a second. Different forms of RNA have been observed in close proximity to the nuclear envelope in electron micrographs (reviewed in Franke and Scheer 1974). Detailed resolution of individual mRNPs and how they move through nuclear pores is mainly derived from work using mRNPs of Balbiani Ring (BR) genes in salivary gland cells of Chironomus (Mehlin et al. 1992; Kiseleva et al. 1998). These large mRNPs, 50 nm in diameter, have been visualized interacting with nuclear pores by electron microscopy. Because of their size, these mRNPs unfold at the NPC and show directional translocation through the NPC, beginning with the 50 end (reviewed in Daneholt 2001). Hypothetical sequences for distinct steps in the export process have been assembled

from EM series (Kiseleva et al. 1998). mRNA export likely involves distinct docking, translocation and release steps from the NPC, analogous to a ratchet model that includes a specific function for the mRNA associated DEAD box helicase DBP5 (reviewed in Stewart 2007). Direct interaction of DPB5 with Nup214, a cytoplasmic component of the NPC, has been shown (Napetschnig et al. 2009; von Moeller et al. 2009) and DPB5 is thought to promote export factor release and eventual reorganization of the mRNA (reviewed in Iglesias and Stutz 2008). DBP5 localization, the timing and location of loading onto the transcript, and how many export factors are actually attached to any individual transcript remains unclear (Zhao et al. 2002; Estruch and Cole 2003; Lund and Guthrie 2005; von Moeller et al. 2009). Discrepancy in the number of BRmRNPs observed on the nuclear and cytoplasmic surfaces of nuclear pores in EM studies have been interpreted as uncoupled asynchronous functions of the export process, which should result in a waiting step during transport. Simultaneously, it was concluded that translocation through the central channel of nuclear pores is probably fast compared to the docking step on the nuclear surface of the pore (Kiseleva et al. 1998). Biochemical studies indicates that mRNA export competence is directly linked to transcription (reviewed in Kohler and Hurt 2007; reviewed in Hurt and Silver 2008; reviewed in Iglesias and Stutz 2008; reviewed in Carmody and Wente 2009; reviewed in Moore and Proudfoot 2009). Recently, a link between actin and transcription has been suggested. Actin (1) can be detected as a component of pre-mRNP complexes, (2) binds transcription factors, (3) is involved in chromatin remodeling, and (4) associates directly to RNA polymerases (reviewed in Miralles and Visa 2006). Interestingly, actin may also have a role in export, as it has been observed to associate cotranscriptionally with BR-mRNPs and remains associated throughout export (Percipalle et al. 2001). Furthermore, the nuclear export receptor exportin-6 shows specificity for profilin-actin, suggesting an additional role as an adaptor for export of certain mRNAs

RNA Processing and Export

(Stuven et al. 2003; reviewed in Miralles and Visa 2006). Export is mediated by protein factors associated with the mRNA, and mRNAs that do not carry the necessary adaptor and export factors are retained in the nucleus (reviewed in Iglesias and Stutz 2008). Most mRNAs seem to export via a TAP (Mex67 in yeast)-dependent pathway. TAP is not a member of the karyopherin family and does not rely on the GTPase Ran, which mediates nuclear import (Segref et al. 1997; reviewed in Macara 2001; reviewed in Dreyfuss et al. 2002). Export factors do not recognize the mRNA directly, but rather through adapters such as Aly/REF (Yra1 in yeast), which is necessary for export (Stutz et al. 2000; Zhou et al. 2000; Gatfield et al. 2001; Le Hir et al. 2001). However, CBP80, a component of the CBC, has been implicated in mediating contacts between transport receptors and mRNA (Hamm and Mattaj 1990; Cheng et al. 2006). Aly/REF has been identified to be loaded in a splice-dependent manner as part of the EJC (Le Hir et al. 2001). Complex regulation of Aly/REF links mRNA export to cell cycle progression (Zhou et al. 2000; Okada et al. 2008; reviewed in Okada and Ye 2009) and Yra1, has been linked to S-phase entry (Swaminathan et al. 2007). Conversely, in Drosophila and Caenorhabditis elegans, mRNA export is Aly/REF independent (Gatfield and Izaurralde 2002; Longman et al. 2003). The transcription-export complex (TREX) exemplifies the tight coupling between transcription and export. Recruitment of the TREX complex is coupled to the transcription machinery in yeast but associated with the splicing machinery in metazoans (Masuda et al. 2005; Cheng et al. 2006). In metazoans, the TREX complex was initially thought to be part of the EJC (Gatfield et al. 2001; Le Hir et al. 2001) but is now known to be recruited to the 50 end independently, in a splicing and 50 cap-dependent manner (Masuda et al. 2005; Cheng et al. 2006). In yeast, molecular machinery serves to dock transcribing genes to nuclear pores, resulting in “gene gating.” SAGA, involved in histone modification and DNA remodeling, has also been shown to interact with transcription

factors (Grant et al. 1997; Larschan and Winston 2001; reviewed in Daniel and Grant 2007). Components of SAGA and TREX2 complexes are required for gene gating of the GAL locus following activation (Cabal et al. 2006; Wilmes and Guthrie 2009). Sus1, one such component, promotes NPC docking and export (reviewed in Blobel 1985; Jani et al. 2009; Klockner et al. 2009; Wilmes and Guthrie 2009). Sus1 is also involved in transcription elongation and may prevent harmful DNA:RNA hybrids during transcription (Klockner et al. 2009). Interactions have been shown for both TREX and TREX2 complexes with Swt1, an endonuclease that interacts with nuclear pores and is involved in mRNA quality control (Skruzny et al. 2009). Although major pathways for mRNA export have been identified, based on interspecies variations and the number of transport factors and cofactors involved, additional pathways are likely to exist. For example, export of certain viral and other mRNAs depend on CRM1 as an export factor (Ohno et al. 2000; reviewed in Dreyfuss et al. 2002; reviewed in Iglesias and Stutz 2008). The extent to which processing and export might be regulated in a species- and differentiation-dependent manner currently remains unclear. Dynamic regulation of mRNA processing factors is a relatively new question, and it is still unclear to what degree mRNA export is regulated at the level of individual nuclear pores. A dynamic view of how mRNA transitions through the nuclear pore is lacking, but recent developments in mRNA labeling and imaging technology may provide the opportunity to fill this gap in the near future. CONCLUDING REMARKS

As discussed, a wealth of ensemble biochemical studies has provided great molecular and mechanistic insight into mRNA processing, leading to further questions that will require in vivo imaging approaches. Several recent studies highlight the importance of such techniques in gaining quantitative information on fundamental aspects of mRNA processing events, with spatial and temporal resolution.

S. Hocine, R.H. Singer, and D. Gru¨nwald

For example, the nucleus is the site of numerous essential cellular processes that are likely coordinated through highly organized and compartmentalized nuclear bodies with distinct functions, as observed using cellular imaging (reviewed in Misteli 2007). Nuclear bodies lack membranes and are highly dynamic yet steady-state structures. This represents a more advanced view from the original idea of nuclear factories for transcription and replication, in which core components localized to static discrete foci (reviewed in Iborra et al. 1996). This environment facilitates gene expression at multiple levels, including chromatin accessibility, transcriptional control, integration of processing events, stringent quality control, export of mature mRNPs to the cytoplasm and translation. Recent work suggests that Cajal body formation does not require specific gene loci and can initiate from any Cajal body protein, supporting a self-assembling model for nuclear bodies (Kaiser et al. 2008; reviewed in Misteli 2008). Additionally, the long-standing question of whether differentially spliced transcripts recruit distinct sets of basal pre-mRNA splicing factors has recently been addressed. Quantitative single-cell imaging has shown the first in vivo evidence of differential association of pre-mRNA splicing factors with alternatively spliced transcripts, supporting a stochastic model of alternative splicing which would predict that combinatorial sets of splicing factors contribute to splicing outcome (Mabon and Misteli 2005). FRET and FLIM techniques have been applied to investigate interactions between SR proteins and splicing components. Unlike biochemical methods, FRET can be used to study interactions in living cells, with minimal perturbation to the highly structured and dynamic nuclear environment (reviewed in Wouters et al. 2001; Ellis et al. 2008). Such an approach has revealed individual interactions that occur in the presence of RNA Pol II inhibitors, suggesting they are not exclusively cotranscriptional. FRAP analysis suggests that processing factors are highly dynamic and are exchanged between nuclear bodies and other nuclear locations in a matter of seconds (Phair and Misteli 2000; reviewed in Lamond and

Spector 2003). Both FRET and FRAP have been used to study the localization and association of SF1 and U2AF. The mobility of these proteins is correlated with their ability to interact with each other, and they are believed to interact in what are described as extraspliceosomal complexes that form before and persist after spliceosome assembly (Rino et al. 2008). The development of real-time, single-molecule imaging techniques provides an especially exciting and promising opportunity to probe in vivo realities, reconciling molecular and mechanistic details within a kinetic and spatial context. One such example involves single particle tracking of U1 snRNP within the nucleus (see Fig. 5), revealing both mobile and transiently immobile

Figure 5. In vivo trajectories of single U1 snRNPs within the nucleus of HeLa cells. Fluorescently labeled native U1 snRNPs were microinjected to visualize and track single molecules, recorded at 200 Hz. SF2/ASF-GFP was transiently expressed to distinguish mobile and transiently immobilized U1 snRNP particles within the nucleoplasm and speckles, outlined in gray (A). A 8 mm2 area from A is broken down into a short image sequence displaying a single trajectory over time (B). Grunwald et al. 2006, # 2006 by The American Society for Cell Biology.

RNA Processing and Export

states (Grunwald et al. 2006). Single molecule imaging makes the range of mobility states over a population of molecules immediately apparent, and changes in the behavior of any individual molecule during observation can be assessed. A major lesson from these studies is that “the mobility” of a given molecule is more likely a mixed population of different states.

ACKNOWLEDGMENTS

This work was supported by National Institutes of Health grant EB2060 to R.H.S. and a DFG fellowship (DG 3388) to D.G. The authors would like to thank S. J. Orenstein, Drs. A. Joseph and V. de Turris for critical reading of the manuscript.

REFERENCES Ares M Jr, Grate L, Pauling MH. 1999. A handful of introncontaining genes produces the lion’s share of yeast mRNA. RNA 5: 1138– 1139. Bagga PS, Ford LP, Chen F, Wilusz J. 1995. The G-rich auxiliary downstream element has distinct sequence and position requirements and mediates efficient 30 end premRNA processing through a trans-acting factor. Nucleic Acids Res 23: 1625–1631. Barrass JD, Beggs JD. 2003. Splicing goes global. Trends Genet 19: 295 –298. Beaudoing E, Gautheret D. 2001. Identification of alternate polyadenylation sites and analysis of their tissue distribution using EST data. Genome Res 11: 1520–1526. Becker M, Baumann C, John S, Walker DA, Vigneron M, McNally JG, Hager GL. 2002. Dynamic behavior of transcription factors on a natural promoter in living cells. EMBO Rep 3: 1188– 1194. Bellare P, Small EC, Huang X, Wohlschlegel JA, Staley JP, Sontheimer EJ. 2008. A role for ubiquitin in the spliceosome assembly pathway. Nat Struct Mol Biol 15: 444 –451. Bentley D. 2002. The mRNA assembly line: Transcription and processing machines in the same factory. Curr Opin Cell Biol 14: 336 –342. Beyer AL, Osheim YN. 1988. Splice site selection, rate of splicing, and alternative splicing on nascent transcripts. Genes Dev 2: 754 –765. Blobel G. 1985. Gene gating: A hypothesis. Proc Natl Acad Sci 82: 8527–8529. Bousquet-Antonelli C, Presutti C, Tollervey D. 2000. Identification of a regulated pathway for nuclear pre-mRNA turnover. Cell 102: 765 –775. Braga J, McNally JG, Carmo-Fonseca M. 2007. A reactiondiffusion model to study RNA motion by quantitative fluorescence recovery after photobleaching. Biophys J 92: 2694–2703.

Brinster RL, Allen JM, Behringer RR, Gelinas RE, Palmiter RD. 1988. Introns increase transcriptional efficiency in transgenic mice. Proc Natl Acad Sci 85: 836 –840. Buratowski S. 2005. Connections between mRNA 30 end processing and transcription termination. Curr Opin Cell Biol 17: 257– 261. Burgess SM, Guthrie C. 1993. A mechanism to enhance mRNA splicing fidelity: The RNA-dependent ATPase Prp16 governs usage of a discard pathway for aberrant lariat intermediates. Cell 73: 1377– 1391. Burkard KT, Butler JS. 2000. A nuclear 30 -50 exonuclease involved in mRNA degradation interacts with Poly(A) polymerase and the hnRNA protein Npl3p. Mol Cell Biol 20: 604 –616. Cabal GG, Genovesio A, Rodriguez-Navarro S, Zimmer C, Gadal O, Lesne A, Buc H, Feuerbach-Fournier F, OlivoMarin JC, Hurt EC, et al. 2006. SAGA interacting factors confine sub-diffusion of transcribed genes to the nuclear envelope. Nature 441: 770– 773. Calvo O, Manley JL. 2005. The transcriptional coactivator PC4/Sub1 has multiple functions in RNA polymerase II transcription. EMBO J 24: 1009– 1020. Carmody SR, Wente SR. 2009. mRNA nuclear export at a glance. J Cell Sci 122: 1933– 1937. Cartegni L, Chew SL, Krainer AR. 2002. Listening to silence and understanding nonsense: Exonic mutations that affect splicing. Nat Rev Genet 3: 285– 298. Cheng H, Dufu K, Lee CS, Hsu JL, Dias A, Reed R. 2006. Human mRNA export machinery recruited to the 50 end of mRNA. Cell 127: 1389– 1400. Cho EJ, Rodriguez CR, Takagi T, Buratowski S. 1998. Allosteric interactions between capping enzyme subunits and the RNA polymerase II carboxy-terminal domain. Genes Dev 12: 3482–3487. Collins CA, Guthrie C. 2000. The question remains: Is the spliceosome a ribozyme? Nat Struct Biol 7: 850 –854. Colot HV, Stutz F, Rosbash M. 1996. The yeast splicing factor Mud13p is a commitment complex component and corresponds to CBP20, the small subunit of the nuclear cap-binding complex. Genes Dev 10: 1699– 1708. Core LJ, Lis JT. 2008. Transcription regulation through promoter-proximal pausing of RNA polymerase II. Science 319: 1791– 1792. Core LJ, Waterfall JJ, Lis JT. 2008. Nascent RNA sequencing reveals widespread pausing and divergent initiation at human promoters. Science 322: 1845– 1848. Cramer P, Pesce CG, Baralle FE, Kornblihtt AR. 1997. Functional association between promoter structure and transcript alternative splicing. Proc Natl Acad Sci 94: 11456– 11460. Custodio N, Carmo-Fonseca M. 2001. Quality control of gene expression in the nucleus. J Cell Mol Med 5: 267 –275. Custodio N, Carmo-Fonseca M, Geraghty F, Pereira HS, Grosveld F, Antoniou M. 1999. Inefficient processing impairs release of RNA from the site of transcription. EMBO J 18: 2855–2866. Danckwardt S, Hentze MW, Kulozik AE. 2008. 30 end mRNA processing: Molecular mechanisms and implications for health and disease. EMBO J 27: 482 –498.

S. Hocine, R.H. Singer, and D. Gru¨nwald Daneholt B. 2001. Assembly and transport of a premessenger RNP particle. Proceedings of the National Academy of Sciences of the United States of America 98: 7012–7017. Daniel JA, Grant PA. 2007. Multi-tasking on chromatin with the SAGA coactivator complexes. Mutat Res 618: 135–148. Danin-Kreiselman M, Lee CY, Chanfreau G. 2003. RNAse III-mediated degradation of unspliced pre-mRNAs and lariat introns. Mol Cell 11: 1279– 1289. Dantonel JC, Murthy KG, Manley JL, Tora L. 1997. Transcription factor TFIID recruits factor CPSF for formation of 30 end of mRNA. Nature 389: 399 –402. Darzacq X, Shav-Tal Y, de Turris V, Brody Y, Shenoy SM, Phair RD, Singer RH. 2007. In vivo dynamics of RNA polymerase II transcription. Nat Struct Mol Biol 14: 796–806. Darzacq X, Yao J, Larson DR, Causse SZ, Bosanac L, de Turris V, Ruda VM, Lionnet T, Zenklusen D, Guglielmi B, et al. 2009. Imaging transcription in living cells. Annu Rev Biophys 38: 173– 196. de la Mata M, Alonso CR, Kadener S, Fededa JP, Blaustein M, Pelisch F, Cramer P, Bentley D, Kornblihtt AR. 2003. A slow RNA polymerase II affects alternative splicing in vivo. Mol Cell 12: 525– 532. Dreyfuss G, Kim VN, Kataoka N. 2002. Messenger-RNAbinding proteins and the messages they carry. Nat Rev Mol Cell Biol 3: 195– 205. Drummond DR, Armstrong J, Colman A. 1985. The effect of capping and polyadenylation on the stability, movement and translation of synthetic messenger RNAs in Xenopus oocytes. Nucleic Acids Res 13: 7375–7394. Du L, Warren SL. 1997. A functional interaction between the carboxy-terminal domain of RNA polymerase II and premRNA splicing. J Cell Biol 136: 5– 18. Dziembowski A, Ventura AP, Rutz B, Caspary F, Faux C, Halgand F, Laprevote O, Seraphin B. 2004. Proteomic analysis identifies a new complex required for nuclear pre-mRNA retention and splicing. EMBO J 23: 4847– 4856. Ellis JD, Lleres D, Denegri M, Lamond AI, Caceres JF. 2008. Spatial mapping of splicing factor complexes involved in exon and intron definition. J Cell Biol 181: 921– 934. Estruch F, Cole CN. 2003. An early function during transcription for the yeast mRNA export factor Dbp5p/ Rat8p suggested by its genetic and physical interactions with transcription factor IIH components. Mol Biol Cell 14: 1664–1676. Flaherty SM, Fortes P, Izaurralde E, Mattaj IW, Gilmartin GM. 1997. Participation of the nuclear cap binding complex in pre-mRNA 30 processing. Proc Natl Acad Sci 94: 11893–11898. Fong YW, Zhou Q. 2001. Stimulatory effect of splicing factors on transcriptional elongation. Nature 414: 929 –933. Franke WW, Scheer U. 1974. Pathways of nucleocytoplasmic translocation of ribonucleoproteins. Symp Soc Exp Biol 28: 249 –282. Galy V, Gadal O, Fromont-Racine M, Romano A, Jacquier A, Nehrbass U. 2004. Nuclear retention of unspliced mRNAs in yeast is mediated by perinuclear Mlp1. Cell 116: 63–73.

Gatfield D, Izaurralde E. 2002. REF1/Aly and the additional exon junction complex proteins are dispensable for nuclear mRNA export. J Cell Biol 159: 579– 588. Gatfield D, Le Hir H, Schmitt C, Braun IC, Kocher T, Wilm M, Izaurralde E. 2001. The DExH/D box protein HEL/ UAP56 is essential for mRNA nuclear export in Drosophila. Curr Biol 11: 1716–1721. Ge H, Si Y, Wolffe AP. 1998. A novel transcriptional coactivator, p52, functionally interacts with the essential splicing factor ASF/SF2. Mol Cell 2: 751 –759. Gil A, Proudfoot NJ. 1984. A sequence downstream of AAUAAA is required for rabbit b-globin mRNA 30 -end formation. Nature 312: 473– 474. Gorski SA, Dundr M, Misteli T. 2006. The road much traveled: Trafficking in the cell nucleus. Curr Opin Cell Biol 18: 284– 290. Gorski SA, Snyder SK, John S, Grummt I, Misteli T. 2008. Modulation of RNA polymerase assembly dynamics in transcriptional regulation. Mol Cell 30: 486 –497. Grant PA, Duggan L, Cote J, Roberts SM, Brownell JE, Candau R, Ohba R, Owen-Hughes T, Allis CD, Winston F, et al. 1997. Yeast Gcn5 functions in two multisubunit complexes to acetylate nucleosomal histones: Characterization of an Ada complex and the SAGA (Spt/Ada) complex. Genes Dev 11: 1640– 1650. Grunwald D, Spottke B, Buschmann V, Kubitscheck U. 2006. Intranuclear binding kinetics and mobility of single native U1 snRNP particles in living cells. Mol Biol Cell 17: 5017–5027. Hamm J, Mattaj IW. 1990. Monomethylated cap structures facilitate RNA export from the nucleus. Cell 63: 109– 118. Hilleren PJ, Parker R. 2003. Cytoplasmic degradation of splice-defective pre-mRNAs and intermediates. Mol Cell 12: 1453–1465. Hilleren P, McCarthy T, Rosbash M, Parker R, Jensen TH. 2001. Quality control of mRNA 30 -end processing is linked to the nuclear exosome. Nature 413: 538– 542. Hirose Y, Manley JL. 1998. RNA polymerase II is an essential mRNA polyadenylation factor. Nature 395: 93– 96. Hirose Y, Manley JL. 2000. RNA polymerase II and the integration of nuclear events. Genes Dev 14: 1415–1429. Hirose Y, Ohkuma Y. 2007. Phosphorylation of the Cterminal domain of RNA polymerase II plays central roles in the integrated events of eucaryotic gene expression. J Biochem 141: 601– 608. Hirose Y, Tacke R, Manley JL. 1999. Phosphorylated RNA polymerase II stimulates pre-mRNA splicing. Genes Dev 13: 1234–1239. Ho CK, Shuman S. 1999. Distinct roles for CTD Ser-2 and Ser-5 phosphorylation in the recruitment and allosteric activation of mammalian mRNA capping enzyme. Mol Cell 3: 405–411. Ho CK, Schwer B, Shuman S. 1998. Genetic, physical, and functional interactions between the triphosphatase and guanylyltransferase components of the yeast mRNA capping apparatus. Mol Cell Biol 18: 5189– 5198. Howe KJ, Kane CM, Ares MJr 2003. Perturbation of transcription elongation influences the fidelity of internal exon inclusion in Saccharomyces cerevisiae. RNA 9: 993 –1006.

RNA Processing and Export Hsu CL, Stevens A. 1993. Yeast cells lacking 50 !30 exoribonuclease 1 contain mRNA species that are poly(A) deficient and partially lack the 50 cap structure. Mol Cell Biol 13: 4826– 4835. Huang Y, Steitz JA. 2005. SRprises along a messenger’s journey. Mol Cell 17: 613 –615. Hurt JA, Silver PA. 2008. mRNA nuclear export and human disease. Dis Model Mech 1: 103– 108. Iborra FJ, Pombo A, McManus J, Jackson DA, Cook PR. 1996. The topology of transcription by immobilized polymerases. Exp Cell Res 229: 167– 173. Iglesias N, Stutz F. 2008. Regulation of mRNP dynamics along the export pathway. FEBS Lett 582: 1987– 1996. Inoue K, Ohno M, Sakamoto H, Shimura Y. 1989. Effect of the cap structure on pre-mRNA splicing in Xenopus oocyte nuclei. Genes Dev 3: 1472– 1479. Izaurralde E, Lewis J, McGuigan C, Jankowska M, Darzynkiewicz E, Mattaj IW. 1994. A nuclear cap binding protein complex involved in pre-mRNA splicing. Cell 78: 657–668. Jacobson A, Peltz SW. 1996. Interrelationships of the pathways of mRNA decay and translation in eukaryotic cells. Annu Rev Biochem 65: 693– 739. Jani D, Lutz S, Marshall NJ, Fischer T, Kohler A, Ellisdon AM, Hurt E, Stewart M. 2009. Sus1, Cdc31, and the Sac3 CID region form a conserved interaction platform that promotes nuclear pore association and mRNA export. Mol Cell 33: 727– 737. Janicki SM, Tsukamoto T, Salghetti SE, Tansey WP, Sachidanandam R, Prasanth KV, Ried T, Shav-Tal Y, Bertrand E, Singer RH, et al. 2004. From silencing to gene expression: Real-time analysis in single cells. Cell 116: 683 –698. Kaiser TE, Intine RV, Dundr M. 2008. De novo formation of a subnuclear body. Science 322: 1713– 1717. Kameoka S, Duque P, Konarska MM. 2004. p54(nrb) associates with the 50 splice site within large transcription/ splicing complexes. EMBO J 23: 1782–1791. Kim HJ, Jeong SH, Heo JH, Jeong SJ, Kim ST, Youn HD, Han JW, Lee HW, Cho EJ. 2004a. mRNA capping enzyme activity is coupled to an early transcription elongation. Mol Cell Biol 24: 6184–6193. Kim M, Ahn SH, Krogan NJ, Greenblatt JF, Buratowski S. 2004b. Transitions in RNA polymerase II elongation complexes at the 30 ends of genes. EMBO J 23: 354 –364. Kim M, Krogan NJ, Vasiljeva L, Rando OJ, Nedea E, Greenblatt JF, Buratowski S. 2004c. The yeast Rat1 exonuclease promotes transcription termination by RNA polymerase II. Nature 432: 517 –522. Kiseleva E, Goldberg MW, Allen TD, Akey CW. 1998. Active nuclear pore complexes in Chironomus: Visualization of transporter configurations related to mRNP export. Journal of Cell Science 111: 223– 236. Klockner C, Schneider M, Lutz S, Jani D, Kressler D, Stewart M, Hurt E, Kohler A. 2009. Mutational uncoupling of the role of Sus1 in nuclear pore complex targeting of an mRNA export complex and histone H2B deubiquitination. J Biol Chem 284: 12049–12056. Kohler A, Hurt E. 2007. Exporting RNA from the nucleus to the cytoplasm. Nat Rev Mol Cell Biol 8: 761– 773. Komarnitsky P, Cho EJ, Buratowski S. 2000. Different phosphorylated forms of RNA polymerase II and associated

mRNA processing factors during transcription. Genes Dev 14: 2452–2460. Konarska MM, Padgett RA, Sharp PA. 1984. Recognition of cap structure in splicing in vitro of mRNA precursors. Cell 38: 731–736. Krishnamurthy S, He X, Reyes-Reyes M, Moore C, Hampsey M. 2004. Ssu72 Is an RNA polymerase II CTD phosphatase. Mol Cell 14: 387 –394. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605– 612. Lander ES, Linton LM, Birren B, Nusbaum C, Zody MC, Baldwin J, Devon K, Dewar K, Doyle M, FitzHugh W, et al. 2001. Initial sequencing and analysis of the human genome. Nature 409: 860–921. Larschan E, Winston F. 2001. The S. cerevisiae SAGA complex functions in vivo as a coactivator for transcriptional activation by Gal4. Genes Dev 15: 1946–1956. Larson DR, Singer RH, Zenklusen D. 2009. A single molecule view of gene expression. Trends Cell Biol 19: 630 –637. Le Hir H, Gatfield D, Izaurralde E, Moore MJ. 2001. The exon-exon junction complex provides a binding platform for factors involved in mRNA export and nonsensemediated mRNA decay. EMBO J 20: 4987– 4997. Le Hir H, Izaurralde E, Maquat LE, Moore MJ. 2000. The spliceosome deposits multiple proteins 20-24 nucleotides upstream of mRNA exon-exon junctions. EMBO J 19: 6860–6869. Le Hir H, Nott A, Moore MJ. 2003. How introns influence and enhance eukaryotic gene expression. Trends Biochem Sci 28: 215– 220. Lewin B. 1980. Eucaryotic Chromosomes. in Gene Expression, pp. 730 –761. John Wiley & Sons, New York. Lewis A, Felberbaum R, Hochstrasser M. 2007. A nuclear envelope protein linking nuclear pore basket assembly, SUMO protease regulation, and mRNA surveillance. J Cell Biol 178: 813– 827. Lewis JD, Izaurralde E, Jarmolowski A, McGuigan C, Mattaj IW. 1996. A nuclear cap-binding complex facilitates association of U1 snRNP with the cap-proximal 50 splice site. Genes Dev 10: 1683–1698. Libri D, Dower K, Boulay J, Thomsen R, Rosbash M, Jensen TH. 2002. Interactions between mRNA export commitment, 30 -end quality control, and nuclear degradation. Mol Cell Biol 22: 8254– 8266. Licatalosi DD, Geiger G, Minet M, Schroeder S, Cilli K, McNeil JB, Bentley DL. 2002. Functional interaction of yeast pre-mRNA 30 end processing factors with RNA polymerase II. Mol Cell 9: 1101–1111. Lin S, Coutinho-Mansfield G, Wang D, Pandit S, Fu XD. 2008. The splicing factor SC35 has an active role in transcriptional elongation. Nat Struct Mol Biol 15: 819– 826. Longman D, Johnstone IL, Caceres JF. 2003. The Ref/Aly proteins are dispensable for mRNA export and development in Caenorhabditis elegans. Rna 9: 881– 891. Lopez PJ, Seraphin B. 1999. Genomic-scale quantitative analysis of yeast pre-mRNA splicing: Implications for splice-site recognition. RNA 5: 1135– 1137. Lu H, Huang X, Zhang L, Guo Y, Cheng H, Zhou R. 2007. Multiple alternative splicing of mouse Dmrt1 during

S. Hocine, R.H. Singer, and D. Gru¨nwald gonadal differentiation. Biochem Biophys Res Commun 352: 630– 634. Lund MK, Guthrie C. 2005. The DEAD-box protein Dbp5p is required to dissociate Mex67p from exported mRNPs at the nuclear rim. Mol Cell 20: 645 –651. Mabon SA, Misteli T. 2005. Differential recruitment of premRNA splicing factors to alternatively spliced transcripts in vivo. PLoS Biol 3: e374. Macara IG. 2001. Transport into and out of the nucleus. Microbiol Mol Biol Rev 65: 570– 594, table of contents. Mandal SS, Chu C, Wada T, Handa H, Shatkin AJ, Reinberg D. 2004. Functional interactions of RNA-capping enzyme with factors that positively and negatively regulate promoter escape by RNA polymerase II. Proc Natl Acad Sci 101: 7572–7577. Mandel CR, Kaneko S, Zhang H, Gebauer D, Vethantham V, Manley JL, Tong L. 2006. Polyadenylation factor CPSF-73 is the pre-mRNA 30 -end-processing endonuclease. Nature 444: 953–956. Masuda S, Das R, Cheng H, Hurt E, Dorman N, Reed R. 2005. Recruitment of the human TREX complex to mRNA during splicing. Genes Dev 19: 1512–1517. Mayas RM, Maita H, Staley JP. 2006. Exon ligation is proofread by the DExD/H-box ATPase Prp22p. Nat Struct Mol Biol 13: 482– 490. McCracken S, Fong N, Yankulov K, Ballantyne S, Pan G, Greenblatt J, Patterson SD, Wickens M, Bentley DL. 1997. The C-terminal domain of RNA polymerase II couples mRNA processing to transcription. Nature 385: 357–361. Mehlin H, Daneholt B, Skoglund U. 1992. Translocation of a Specific Premessenger Ribonucleoprotein Particle through the Nuclear-Pore Studied with Electron-Microscope Tomography. Cell 69: 605 –613. Merz C, Urlaub H, Will CL, Luhrmann R. 2007. Protein composition of human mRNPs spliced in vitro and differential requirements for mRNP protein recruitment. RNA 13: 116 –128. Miralles F, Visa N. 2006. Actin in transcription and transcription regulation. Curr Opin Cell Biol 18: 261– 266. Misteli T. 2007. Beyond the sequence: Cellular organization of genome function. Cell 128: 787 –800. Misteli T. 2008. Cell biology: Nuclear order out of chaos. Nature 456: 333–334. Misteli T, Spector DL. 1999. RNA polymerase II targets premRNA splicing factors to transcription sites in vivo. Mol Cell 3: 697– 705. Moore MJ, Proudfoot NJ. 2009. Pre-mRNA processing reaches back to transcription and ahead to translation. Cell 136: 688– 700. Muhlrad D, Decker CJ, Parker R. 1995. Turnover mechanisms of the stable yeast PGK1 mRNA. Mol Cell Biol 15: 2145–2156. Myers LC, Lacomis L, Erdjument-Bromage H, Tempst P. 2002. The yeast capping enzyme represses RNA polymerase II transcription. Mol Cell 10: 883–894. Napetschnig J, Kassube SA, Debler EW, Wong RW, Blobel Gn, Hoelz A. 2009. Structural and functional analysis of the interaction between the nucleoporin Nup214 and the DEAD-box helicase Ddx19. Proceedings of the National Academy of Sciences 106: 3089– 3094.

Nissim-Rafinia M, Kerem B. 2005. The splicing machinery is a genetic modifier of disease severity. Trends Genet 21: 480 –483. Noble CG, Hollingworth D, Martin SR, Ennis-Adeniran V, Smerdon SJ, Kelly G, Taylor IA, Ramos A. 2005. Key features of the interaction between Pcf11 CID and RNA polymerase II CTD. Nat Struct Mol Biol 12: 144–151. Ohno M, Segref A, Bachi A, Wilm M, Mattaj IW. 2000. PHAX, a mediator of U snRNA nuclear export whose activity is regulated by phosphorylation. Cell 101: 187 –198. Okada M, Ye K. 2009. Nuclear phosphoinositide signaling regulates messenger RNA export. RNA Biol 6: 12– 16. Okada M, Jang SW, Ye K. 2008. Akt phosphorylation and nuclear phosphoinositide association mediate mRNA export and cell proliferation activities by ALY. Proc Natl Acad Sci 105: 8649– 8654. Palancade B, Zuccolo M, Loeillet S, Nicolas A, Doye V. 2005. Pml39, a novel protein of the nuclear periphery required for nuclear retention of improper messenger ribonucleoparticles. Mol Biol Cell 16: 5258–5268. Percipalle P, Zhao J, Pope B, Weeds A, Lindberg U, Daneholt B. 2001. Actin bound to the heterogeneous nuclear ribonucleoprotein hrp36 is associated with Balbiani ring mRNA from the gene to polysomes. J Cell Biol 153: 229 –236. Peterlin BM, Price DH. 2006. Controlling the elongation phase of transcription with P-TEFb. Mol Cell 23: 297 –305. Peterson ML, Perry RP. 1989. The regulated production of mu m and mu s mRNA is dependent on the relative efficiencies of mu s poly(A) site usage and the c mu 4-to-M1 splice. Mol Cell Biol 9: 726–738. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604 –609. Phatnani HP, Jones JC, Greenleaf AL. 2004. Expanding the functional repertoire of CTD kinase I and RNA polymerase II: Novel phosphoCTD-associating proteins in the yeast proteome. Biochemistry 43: 15702– 15719. Politz JC, Browne ES, Wolf DE, Pederson T. 1998. Intranuclear diffusion and hybridization state of oligonucleotides measured by fluorescence correlation spectroscopy in living cells. Proc Natl Acad Sci 95: 6043–6048. Politz JC, Tuft RA, Prasanth KV, Baudendistel N, Fogarty KE, Lifshitz LM, Langowski J, Spector DL, Pederson T. 2006. Rapid, diffusional shuttling of poly(A) RNA between nuclear speckles and the nucleoplasm. Mol Biol Cell 17: 1239–1249. Proudfoot N. 2004. New perspectives on connecting messenger RNA 30 end formation to transcription. Curr Opin Cell Biol 16: 272– 278. Proudfoot NJ, Furger A, Dye MJ. 2002. Integrating mRNA processing with transcription. Cell 108: 501– 512. Pyle AM. 2008. Translocation and unwinding mechanisms of RNA and DNA helicases. Annu Rev Biophys 37: 317 –336. Qu X, Lykke-Andersen S, Nasser T, Saguez C, Bertrand E, Jensen TH, Moore C. 2009. Assembly of an exportcompetent mRNP is needed for efficient release of the 30 end processing complex after polyadenylation. Mol Cell Biol 29: 5327–5338.

RNA Processing and Export Rino J, Desterro JM, Pacheco TR, Gadella TWJr, CarmoFonseca M. 2008. Splicing factors SF1 and U2AF associate in extraspliceosomal complexes. Mol Cell Biol 28: 3045– 3057. Rodriguez CR, Cho EJ, Keogh MC, Moore CL, Greenleaf AL, Buratowski S. 2000. Kin28, the TFIIH-associated carboxy-terminal domain kinase, facilitates the recruitment of mRNA processing machinery to RNA polymerase II. Mol Cell Biol 20: 104– 112. Rosner A, Rinkevich B. 2007. The DDX3 subfamily of the DEAD box helicases: Divergent roles as unveiled by studying different organisms and in vitro assays. Curr Med Chem 14: 2517–2525. Rougemaille M, Gudipati RK, Olesen JR, Thomsen R, Seraphin B, Libri D, Jensen TH. 2007. Dissecting mechanisms of nuclear mRNA surveillance in THO/sub2 complex mutants. EMBO J 26: 2317– 2326. Russnak RH. 1991. Regulation of polyadenylation in hepatitis B viruses: Stimulation by the upstream activating signal PS1 is orientation-dependent, distance-independent, and additive. Nucleic Acids Res 19: 6449–6456. Ryan K, Calvo O, Manley JL. 2004. Evidence that polyadenylation factor CPSF-73 is the mRNA 30 processing endonuclease. RNA 10: 565–573. Sadofsky M, Alwine JC. 1984. Sequences on the 30 side of hexanucleotide AAUAAA affect efficiency of cleavage at the polyadenylation site. Mol Cell Biol 4: 1460–1468. Schermelleh L, Carlton PM, Haase S, Shao L, Winoto L, Kner P, Burke B, Cardoso MC, Agard DA, Gustafsson MG, et al. 2008. Subdiffraction multicolor imaging of the nuclear periphery with 3D structured illumination microscopy. Science 320: 1332– 1336. Schroeder SC, Schwer B, Shuman S, Bentley D. 2000. Dynamic association of capping enzymes with transcribing RNA polymerase II. Genes Dev 14: 2435–2440. Schroeder SC, Zorio DA, Schwer B, Shuman S, Bentley D. 2004. A function of yeast mRNA cap methyltransferase, Abd1, in transcription by RNA polymerase II. Mol Cell 13: 377 –387. Segref A, Sharma K, Doye V, Hellwig A, Huber J, Luhrmann R, Hurt E. 1997. Mex67p, a novel factor for nuclear mRNA export, binds to both poly(A)þ RNA and nuclear pores. EMBO J 16: 3256–3271. Shatkin AJ, Manley JL. 2000. The ends of the affair: Capping and polyadenylation. Nat Struct Biol 7: 838 –842. Shav-Tal Y, Singer RH, Darzacq X. 2004b. Imaging gene expression in single living cells. Nat Rev Mol Cell Biol 5: 855–861. Shav-Tal Y, Darzacq X, Shenoy SM, Fusco D, Janicki SM, Spector DL, Singer RH. 2004a. Dynamics of single mRNPs in nuclei of living cells. Science 304: 1797– 1800. Shuman S. 2001. Structure, mechanism, and evolution of the mRNA capping apparatus. Prog Nucleic Acid Res Mol Biol 66: 1 –40. Siebrasse JP, Veith R, Dobay A, Leonhardt H, Daneholt B, Kubitscheck U. 2008. Discontinuous movement of mRNP particles in nucleoplasmic regions devoid of chromatin. Proc Nat Acad Sci 105: 20291– 20296. Singh R, Valcarcel J. 2005. Building specificity with nonspecific RNA-binding proteins. Nat Struct Mol Biol 12: 645–653.

Singh BN, Hampsey M. 2007. A transcription-independent role for TFIIB in gene looping. Mol Cell 27: 806–816. Skruzny M, Schneider C, Racz A, Weng J, Tollervey D, Hurt E. 2009. An endoribonuclease functionally linked to perinuclear mRNP quality control associates with the nuclear pore complexes. PLoS Biol 7: e8. Smith MJ, Kulkarni S, Pawson T. 2004. FF domains of CA150 bind transcription and splicing factors through multiple weak interactions. Mol Cell Biol 24: 9274– 9285. Stalder L, Muhlemann O. 2008. The meaning of nonsense. Trends Cell Biol 18: 315 –321. Staley JP, Guthrie C. 1998. Mechanical devices of the spliceosome: motors, clocks, springs, and things. Cell 92: 315 –326. Stark H, Luhrmann R. 2006. Cryo-electron microscopy of spliceosomal components. Annu Rev Biophys Biomol Struct 35: 435–457. Stewart M. 2007. Ratcheting mRNA out of the nucleus. Mol Cell 25: 327–330. Stutz F, Bachi A, Doerks T, Braun IC, Seraphin B, Wilm M, Bork P, Izaurralde E. 2000. REF, an evolutionary conserved family of hnRNP-like proteins, interacts with TAP/Mex67p and participates in mRNA nuclear export. Rna 6: 638 –650. Stuven T, Hartmann E, Gorlich D. 2003. Exportin 6: A novel nuclear export receptor that is specific for profilin.actin complexes. EMBO J 22: 5928– 5940. Sun ZW, Hampsey M. 1996. Synthetic enhancement of a TFIIB defect by a mutation in SSU72, an essential yeast gene encoding a novel protein that affects transcription start site selection in vivo. Mol Cell Biol 16: 1557– 1566. Swaminathan S, Kile AC, MacDonald EM, Koepp DM. 2007. Yra1 is required for S phase entry and affects Dia2 binding to replication origins. Mol Cell Biol 27: 4674– 4684. Tarun SZJr, Sachs AB. 1996. Association of the yeast poly(A) tail binding protein with translation initiation factor eIF-4G. EMBO J 15: 7168–7177. Tian B, Hu J, Zhang H, Lutz CS. 2005. A large-scale analysis of mRNA polyadenylation of human and mouse genes. Nucleic Acids Res 33: 201– 212. Torchet C, Bousquet-Antonelli C, Milligan L, Thompson E, Kufel J, Tollervey D. 2002. Processing of 30 -extended readthrough transcripts by the exosome can generate functional mRNAs. Mol Cell 9: 1285–1296. Valencia P, Dias AP, Reed R. 2008. Splicing promotes rapid and efficient mRNA export in mammalian cells. Proc Natl Acad Sci 105: 3386–3391. Verdecia MA, Bowman ME, Lu KP, Hunter T, Noel JP. 2000. Structural basis for phosphoserine-proline recognition by group IV WW domains. Nat Struct Biol 7: 639–643. Villa T, Rougemaille M, Libri D. 2008. Nuclear quality control of RNA polymerase II ribonucleoproteins in yeast: Tilting the balance to shape the transcriptome. Biochim Biophys Acta 1779: 524– 531. von Moeller H, Basquin C, Conti E. 2009. The mRNA export protein DBP5 binds RNA and the cytoplasmic nucleoporin NUP214 in a mutually exclusive manner. Nat Struct Mol Biol 16: 247– 254. Wahl MC, Will CL, Luhrmann R. 2009. The spliceosome: Design principles of a dynamic RNP machine. Cell 136: 701 –718.

S. Hocine, R.H. Singer, and D. Gru¨nwald Wakiyama M, Imataka H, Sonenberg N. 2000. Interaction of eIF4G with poly(A)-binding protein stimulates translation and is critical for Xenopus oocyte maturation. Curr Biol 10: 1147– 1150. Walther TN, Wittop Koning TH, Schumperli D, Muller B. 1998. A 50 -30 exonuclease activity involved in forming the 30 products of histone pre-mRNA processing in vitro. RNA 4: 1034– 1046. Wang ET, Sandberg R, Luo S, Khrebtukova I, Zhang L, Mayr C, Kingsmore SF, Schroth GP, Burge CB. 2008. Alternative isoform regulation in human tissue transcriptomes. Nature 456: 470–476. West S, Gromak N, Proudfoot NJ. 2004. Human 50 ! 30 exonuclease Xrn2 promotes transcription termination at co-transcriptional cleavage sites. Nature 432: 522– 525. Wilmes GM, Guthrie C. 2009. Getting to the gate: Crystallization of a Sac3(CID):Sus1:Cdc31 complex. Mol Cell 33: 671–672. Wilusz JE, Spector DL. 2010. An unexpected ending: Noncanonical 30 end processing mechanisms. RNA 16: 259–266. Wouters FS, Verveer PJ, Bastiaens PI. 2001. Imaging biochemistry inside cells. Trends Cell Biol 11: 203– 211. Wu WH, Pinto I, Chen BS, Hampsey M. 1999. Mutational analysis of yeast TFIIB. A functional relationship between Ssu72 and Sub1/Tsp1 defined by allele-specific interactions with TFIIB. Genetics 153: 643–652. Yao J, Ardehali MB, Fecko CJ, Webb WW, Lis JT. 2007. Intranuclear distribution and local dynamics of RNA polymerase II during transcription activation. Mol Cell 28: 978–990.

Yao J, Zobeck KL, Lis JT, Webb WW. 2008. Imaging transcription dynamics at endogenous genes in living Drosophila tissues. Methods 45: 233–241. Yonaha M, Proudfoot NJ. 2000. Transcriptional termination and coupled polyadenylation in vitro. EMBO J 19: 3770–3777. Yue Z, Maldonado E, Pillutla R, Cho H, Reinberg D, Shatkin AJ. 1997. Mammalian capping enzyme complements mutant Saccharomyces cerevisiae lacking mRNA guanylyltransferase and selectively binds the elongating form of RNA polymerase II. Proc Natl Acad Sci 94: 12898– 12903. Yuryev A, Patturajan M, Litingtung Y, Joshi RV, Gentile C, Gebara M, Corden JL. 1996. The C-terminal domain of the largest subunit of RNA polymerase II interacts with a novel set of serine/arginine-rich proteins. Proc Natl Acad Sci 93: 6975–6980. Zenklusen D, Larson DR, Singer RH. 2008. Single-RNA counting reveals alternative modes of gene expression in yeast. Nat Struct Mol Biol 15: 1263–1271. Zhang G, Taneja KL, Singer RH, Green MR. 1994. Localization of pre-mRNA splicing in mammalian nuclei. Nature 372: 809–812. Zhao J, Jin SB, Bjorkroth B, Wieslander L, Daneholt B. 2002. The mRNA export factor Dbp5 is associated with Balbiani ring mRNP from gene to cytoplasm. EMBO J 21: 1177–1187. Zhou Z, Luo MJ, Straesser K, Katahira J, Hurt E, Reed R. 2000. The protein Aly links pre-messenger-RNA splicing to nuclear export in metazoans. Nature 407: 401 –405. Zorio DA, Bentley DL. 2004. The link between mRNA processing and transcription: communication works both ways. Exp Cell Res 296: 91–97.

Diseases of the Nuclear Envelope ¨ stlund, and Yuexia Wang Howard J. Worman, Cecilia O Department of Medicine and Department of Pathology and Cell Biology, College of Physicians and Surgeons, Columbia University, New York, New York 10032 Correspondence: [email protected]

In the past decade, a wide range of fascinating monogenic diseases have been linked to mutations in the LMNA gene, which encodes the A-type nuclear lamins, intermediate filament proteins of the nuclear envelope. These diseases include dilated cardiomyopathy with variable muscular dystrophy, Dunnigan-type familial partial lipodystrophy, a CharcotMarie-Tooth type 2 disease, mandibuloacral dysplasia, and Hutchinson-Gilford progeria syndrome. Several diseases are also caused by mutations in genes encoding B-type lamins and proteins that associate with the nuclear lamina. Studies of these so-called laminopathies or nuclear envelopathies, some of which phenocopy common human disorders, are providing clues about functions of the nuclear envelope and insights into disease pathogenesis and human aging.

utations in LMNA encoding the A-type lamins cause a group of human disorders often collectively called laminopathies. The major A-type lamins, lamin A and lamin C, arise by alternative splicing of the LMNA pre-mRNA and are expressed in virtually all differentiated somatic cells. Although the A-type lamins are widely expressed, LMNA mutations are responsible for at least a dozen different clinically defined disorders with tissue-selective abnormalities. Mutations in genes encoding B-type lamins and lamin-associated proteins, most of which are similarly expressed in almost all somatic cells, also cause tissue-selective diseases. Research on the laminopathies has provided novel clues about nuclear envelope function. Recent studies have begun to shed light on how alterations in the nuclear envelope could

M

explain disease pathogenesis. Along with basic research on nuclear structure, the nuclear lamins, and lamina-associated proteins, clinical research on the laminopathies will contribute to a complete understanding of the functions of the nuclear envelope in normal physiology and in human pathology. LMNA: ONE GENE, MANY DISEASES

George Beadle and Edward Tatum (Beadle and Tatum 1941) proposed what became known as the “one gene-one enzyme” hypothesis and was later modified to the “one gene-onepolypeptide” hypothesis. The premise underlying this hypothesis was that genes act through the production of polypeptides, with each gene producing a single polypeptide functioning in

Editors: Tom Misteli and David Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000760 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000760

¨ stlund, and Y. Wang H.J. Worman, C. O

a particular step in a metabolic pathway or other cellular process. A corollary of this hypothesis, which formed the foundation of most early studies using positional cloning to identify disease genes, was the “one gene-one disease” principle. We now know that this is not correct and perhaps the best example that disproves this principle is LMNA. LMNA encoding the A-type lamins was characterized in 1993 and subsequently mapped to chromosome 1q21.2-q21.3 (Lin and Worman 1993; Wydner et al. 1996). The first human disease identified by positional cloning to be caused by LMNA mutations was autosomal dominant Emery-Dreifuss muscular dystrophy (Bonne et al. 1999). Rare compound heterozygous mutations in LMNA causing recessively inherited Emery-Dreifuss muscular dystrophy were described shortly thereafter (Raffaele di Barletta et al. 2000). Patients with Emery-Dreifuss muscular dystrophy classically have early contractures of the elbows, Achilles tendons, and posterior neck, rigidity of the spine, slowly progressive muscle weakness in the upper arms and lower legs, and dilated cardiomyopathy with an early onset atrioventricular conduction block (Emery 2000; Muchir and Worman 2007). Soon after LMNA mutations were shown to cause Emery-Dreifuss muscular dystrophy, mutations in this gene were shown to cause other dominantly inherited diseases affecting primarily striated muscle, including dilated cardiomyopathy 1A (Fatkin et al. 1999) and limb girdle muscular dystrophy type 1B (Muchir et al. 2000). Like Emery-Dreifuss muscular dystrophy, these conditions have a predominant dilated cardiomyopathy with early onset atrioventricular conduction block. In dilated cardiomyopathy 1A, skeletal muscle is minimally affected or unaffected. In limb-girdle muscular dystrophy, the distribution of skeletal muscle involvement is primarily around the shoulders and hips with sparing of the distal extremities. Most subjects with limb-girdle muscular dystrophy do not have joint contractures characteristic of classical Emery-Dreifuss muscular dystrophy. It was originally proposed that Emery-Dreifuss muscular dystrophy, dilated

cardiomyopathy 1A, and limb-girdle muscular dystrophy type 1B resulted from different mutations in LMNA. However, the phenotypic variability is most likely because of the influence of modifier genes or environmental factors. This is suggested by intrafamilial variability and phenotypic overlap in patients with LMNA mutations and muscle disease (Bonne et al. 2000). Within a single family, one affected individual can be diagnosed with isolated cardiomyopathy, another with Emery-Dreifuss muscular dystrophy, and others with limb-girdle muscular dystrophy (Brodsky et al. 2000). Based on the combined phenotypic and genetic data, dilated cardiomyopathy with variable skeletal muscle involvement, a phrase used by Brodsky et al. (2000), is a very appropriate descriptor of the striated muscle disease caused by LMNA mutations. Although most LMNA mutations causing muscle disorders present during childhood or early adulthood, rare subjects present with congenital muscular dystrophy (Quijano-Roy et al. 2008). Congenital muscular dystrophy has an earlier onset and more severe phenotype than the later-onset muscle disorders caused by LMNA mutations. Most cases of LMNAassociated congenital muscular dystrophy are caused by de novo mutations but cases of germinal mosaicism have also been identified (Makri et al. 2009). Although some of the LMNA mutations causing congenital muscular dystrophy appear to be unique, others have been reported in patients with the later-onset myopathies. A unique LMNA splice site mutation has also been associated with a heart-hand syndrome, which is characterized by the association of congenital cardiac disease and limb deformities (Renou et al. 2008). After LMNA mutations were shown to cause striated muscle diseases, a surprising discovery was made regarding another monogenic disease affecting different tissues. In 1998, the genetic locus for Dunnigan-type familial partial lipodystrophy had been mapped using positional cloning to chromosome 1q21-22 (Jackson et al. 1998; Peters et al. 1998). Lipodystrophies are a group of disorders characterized by the absence or reduction of subcutaneous adipose

Laminopathies and Aging

tissue. Patients with Dunnigan-type familial partial lipodystrophy, a dominantly inherited disorder, are born with normal fat distribution but after the onset of puberty there is regional loss of fat from the extremities associated with insulin resistance and frequently diabetes mellitus (Dunnigan et al. 1974). Knowing that the genetic locus for this disease was at chromosome 1q21-22, Cao and Hegele (2000) hypothesized that the analogy between the regional muscle wasting in autosomal dominant EmeryDreifuss muscular dystrophy and the regional adipocyte degeneration in this disease made LMNA a candidate gene. They identified a novel missense mutation in exon 8 leading to a R482Q amino-acid substitution, which cosegregated with the lipodystrophy phenotype in five Canadian families. At around the same time, the two groups that had mapped the disease to chromosome 1q21-22 performed finer mapping and identified the LMNA R482Q and other mutations in exon 8 leading to amino-acid substitutions (Shackleton et al. 2000; Speckman et al. 2000). Missense mutations in exon 11 of LMNA leading to R582H and R584H aminoacid substitutions in lamin A, but not lamin C, were further identified in some atypical cases (Speckman et al. 2000; Vigouroux et al. 2000). Subsequently, there have been a few reports of patients with other LMNA mutations with atypical lipodystrophy syndromes, sometimes in combination with muscle abnormalities (Vigouroux and Capeau 2005). By 2000, the positional cloners had clearly shown that mutations in LMNA cause two quite different diseases: dilated cardiomyopathy with variable muscular dystrophy and partial lipodystrophy. However, the situation soon became more complicated when just a couple of years later De Sandre-Giovannoli et al. (2002) performed homozygosity mapping in inbred Algerian families with an autosomal recessive form of Charcot-Marie-Tooth disease type 2, linked it to chromosome 1q21.2-q21.3, and identified a LMNA mutation leading to the R298C amino-acid substitution. Subjects with Charcot-Marie-Tooth disease type 2 diseases, including the subtype caused by LMNA mutation, have slight or absent reduction of nerve

conduction velocities, loss of large myelinated fibers, and axonal degeneration (Chaouch et al. 2003). Affected individuals with the R298C mutation have variable severity and progression of disease, suggesting that modifier genes influence the phenotype of peripheral neuropathy caused by LMNA mutation (Tazir et al. 2004). Later in 2002, Novelli et al. (2002) hypothesized that LMNA mutations might cause mandibuloacral dysplasia, a rare autosomal recessive disorder in which subjects have an undersized jaw, underdeveloped clavicles, other congenital bone abnormalities, and partial lipodystrophy. They studied five consanguineous Italian families and identified a homozygous LMNA missense mutation causing a R527H amino-acid substitution that was shared by all affected patients. Subsequent subjects have been described with homozygous LMNA mutations causing R527C or A529V amino-acid substitutions (Agarwal et al. 2008; Garg et al. 2005). A compound heterozygous subject for the LMNA R527H and a V440M mutation with some features of mandibuloacral dysplasia, lack of muscle strength, and decreased muscle tone has also been reported (Lombardi et al. 2007). Hutchinson-Gilford progeria syndrome, first described over a century ago, is a rare disease with features of accelerated or premature aging (Hutchinson 1886; Gilford 1904; McKusick 1952; DeBusk 1972). Individuals with this autosomal dominant sporadic syndrome generally die in the second decade of life from myocardial infarction or stroke (DeBusk 1972; Merideth et al. 2008). Other prominent phenotypic features are sclerotic skin, joint contractures, prominent eyes, an undersized jaw, decreased subcutaneous fat, alopecia, skin dimpling and mottling, prominent vasculature in the skin, fingertip tufting, and growth impairment (Merideth et al. 2008). In 2003, Francis Collins and colleagues localized the responsible gene to chromosome 1q by observing two cases in which this chromosomal region was from the same parent and one case with a six-megabase paternal interstitial deletion (Eriksson et al. 2003). They then showed that 18 out of 20 classical cases of Hutchinson-Gilford progeria

¨ stlund, and Y. Wang H.J. Worman, C. O

had a de novo G608G (nucleotide 1824 C.T) mutation within exon 11 of LMNA and another case with a G608S (nucleotide 1822 G.A) mutation (Eriksson et al. 2003), a finding that was simultaneously reported by De SandreGiovannoli et al. (2003) and then confirmed by Cao and Hegele (2003). These mutations activate a cryptic splice donor site resulting in the synthesis of a protein with 50 amino acids deleted near the carboxyl terminus of prelamin A. This truncated prelamin A variant is not appropriately processed to lamin A (see below). Other LMNA missense mutations not generating abnormal RNA splicing within exon 11 have also been reported in variant progeroid syndromes (Chen et al. 2003; Csoka et al. 2004; Verstraeten et al. 2006). Mandibuloacral dysplasia caused by LMNA mutations, as discussed previously, also has progeroid features. In summary, genetic studies since the late 1990s have shown that mutations in LMNA cause about a dozen clinical disorders with different names (Table 1). These can more broadly Table 1. Mutations in LMNA cause several distinct clinical diseases predominantly affecting striated muscle, adipose, and peripheral nerve, or give a progeria phenotype

Striated Muscle Autosomal dominant (and rarely recessive) Emery-Dreifuss muscular dystrophy Cardiomyopathy dilated 1A Limb-girdle muscular dystrophy type 1B Congenital muscular dystrophy “Heart-hand” syndrome Adipose Tissue Dunnigan-type familial partial lipodystrophy Lipoatrophy with diabetes and other features of insulin resistance Atypical lipodystrophy syndromes Mandibuloacral dysplasia Peripheral Nerve Charcot-Marie-Tooth disease type 2B1 Progeria Phenotype Hutchinson-Gilford progeria syndrome Atypical Werner Syndrome Variant progeroid disorders Mandibuloacral Dysplasia Mandibuloacral dysplasia has features of lipodystrophy and progeria

be classified into diseases affecting predominantly (1) striated muscle, (2) adipose tissue, (3) peripheral nerve, or (4) multiple tissues resulting in progeroid phenotypes. These mostly tissue-selective disorders occur even though A-type lamins are intermediate filament protein components of the nuclear lamina in virtually all differentiated somatic cells. OTHER LAMINOPATHIES/NUCLEAR ENVELOPATHIES

Monogenic diseases resulting from mutations in genes encoding B-type lamins and proteins that are directly or indirectly associated with the nuclear lamina are also sometimes referred to as laminopathies or nuclear envelopathies (Table 2). Emery-Dreifuss muscular dystrophy was shown to be inherited in an X-linked manner years before autosomal inheritance was described (Emery and Dreifuss 1966). In its classical presentation, the X-linked inherited disease phenocopies the autosomal form. In 1994, Daniella Toniolo and colleagues reported that the gene responsible for X-linked EmeryDreifuss muscular dystrophy encoded a monotopic transmembrane protein expressed in virtually all cells that the authors named emerin (Bione et al. 1994). They extended their initial findings to more patients (Bione et al. 1995). Soon after, emerin was shown to be a protein of the inner nuclear membrane (Manilal et al. 1996; Nagano et al. 1996). Emerin was further shown to depend on A-type lamins for its localization to the nuclear envelope and to directly interact with lamins (Clements et al. 2000; Fairley et al. 1999; Sullivan et al. 1999). The clinical spectrum of disease resulting from mutations in EMD encoding emerin is actually wider than the classical Emery-Dreifuss phenotype and includes a limb-girdle muscular dystrophy, cardiomyopathy with minimal muscle or joint involvement, and various intermittent forms (Astejada et al. 2007). Lamin B receptor is an integral protein of the inner nuclear membrane that binds to B-type lamins (Worman et al. 1988). It has a basically charged, nucleoplasmic, amino-terminal domain that binds to lamins, DNA, and

Laminopathies and Aging Table 2. Diseases caused by mutations in genes encoding B-type lamins or proteins associated with the nuclear lamina Gene

Protein

EMD LBR

emerin lamin B receptor

LEMD3 SYNE1 TMPO

MAN1 nesprin-1 lamina-associated polypeptide 2 torsinA lamin B1 lamin B2 prelamin A endoprotease

TOR1A LMNB1 LMNB2 ZMPSTE24

Disease

cardiomyopathy with muscular dystrophy Pelger-Hue¨t anomaly (heterozygous) Greenberg skeletal dysplasia (homozygous) Sclerosing bone dysplasias cerebellar ataxia cardiomyopathy DYT1 dystonia adult-onset autosomal dominant leukodystrophy acquired partial lipodystrophy restrictive dermopathy and progeroid disorders

Reported in two affected individuals in a single family without gene sequencing from unaffected individuals in the same family.

chromatin proteins followed by a stretch of eight putative transmembrane segments that has high sequence similarity to sterol reductases (Worman et al. 1990; Ye and Worman 1994, 1996; Holmer et al. 1998). Heterozygous mutations in LBR that lead to reduced expression of the protein cause Pelger-Hue¨t anomaly, a benign autosomal dominant disorder characterized by abnormal nuclear shape and chromatin organization in blood neutrophils (Hoffmann et al. 2002). In contrast, homozygous LBR mutations lead to severe developmental abnormalities or are lethal in utero (Hoffmann et al. 2002; Oosterwijk et al. 2003). In one case, a homozygous mutation leading to production of a truncated protein lacking the carboxyl-terminal 82 amino acids was reported to cause hydrops-ectopic calcification-“moth-eaten” or Greenberg skeletal dysplasia, a lethal disorder, which was associated with loss of detectable sterol D14-reductase activity (Waterham et al. 2003). It appears that depending on the amount of expression and the affected functional domains of the protein, the phenotypes resulting from mutations in LBR can range from a benign alteration in neutrophil nuclear morphology to death in utero. MAN1 is an integral inner nuclear membrane protein with two transmembrane segments and two nucleoplasmic domains (Lin et al. 2000). The nucleoplasmic domain preceding the first transmembrane segment has been

reported to bind to A-type lamins, B-type lamins, and emerin (Mansharamani and Wilson, 2005). The nucleoplasmic domain following the second transmembrane segment binds to regulatory-Smads and DNA (Hellemans et al. 2004; Lin et al. 2005; Pan et al. 2005; Caputo et al. 2006). Heterozygous loss-of-function mutations in LEMD3 encoding MAN1 cause osteopoikilosis, Buschke-Ollendorff syndrome, and nonsporadic melorheostosis, sclerosing bone dysplasias that sometimes have hyperproliferative skin abnormalities. These phenotypes are likely associated with enhanced transforming growth factor-b and bone morphogenic protein signaling, the effects of which are mediated by regulatory-Smads. SYNE1 encodes nesprin-1, a protein with several isoforms that arise by alternative RNA splicing. Depending on their size, nesprin-1 isoform may localize to the inner or outer nuclear membrane. Larger nesprin-1 isoforms localize to the outer nuclear membrane and interact in the perinuclear space with Sun proteins, integral proteins of the inner nuclear membrane that bind to lamins, forming a complex connecting the nucleus to the cytoskeleton (Crisp et al. 2006). Homozygous mutations in SYNE1 have been shown to cause an autosomal recessive cerebellar ataxia specifically affecting a part of the brain (Gros-Louis et al. 2007). In one large family, however, a SYNE1 mutation was shown to cosegregate with autosomal

¨ stlund, and Y. Wang H.J. Worman, C. O

recessive arthrogryposis, a disease characterized by bilateral clubfoot, decreased fetal movements, and delayed motor milestones with progressive motor decline after the first decade (Attali et al. 2009). Mutations in genes encoding other proteins that interact directly or indirectly with lamins also cause tissue-selective human diseases. Polymorphisms in the gene encoding laminaassociated polypeptide 2 have been identified in two individuals with cardiomyopathy in a single family; however, sequencing of the gene in unaffected family members was not reported (Taylor et al. 2005). DYT1 dystonia is a central nervous system movement disorder in which sustained muscle contractions lead to twisting and repetitive movements or abnormal postures; it is caused by an in-frame deletion in TOR1A encoding torsinA that leads to loss of a glutamic acid residue (DE302/3) from the protein (Ozelius et al. 1997). TorsinA is an AAAþ ATPase of the endoplasmic reticulum that interacts with the luminal domain of lamina-associated polypeptide 1, an integral inner nuclear membrane protein that interacts with lamins (Goodchild and Dauer 2005). The torsinA DE302/3 variant concentrates in the perinuclear space relative to the bulk endoplasmic reticulum (Gonzalez-Alegre and Paulson 2004; Goodchild and Dauer 2004; Naismith et al. 2004), where it appears to selectively disrupt the structure of the nuclear envelopes of neurons (Goodchild et al. 2005). Mutations in genes other than LMNA but directly affecting lamins also cause diseases. ZMPSTE24 is an endoprotease responsible for the processing of prelamin A to lamin A (see later). Loss-of-function mutations in ZMPSTE24 cause autosomal recessive restrictive dermopathy, a neonatal lethal progeroid disorder (Navarro et al. 2005). Compound heterozygous mutations in ZMPSTE24 also cause progeroid disorders with some cases being diagnosed as mandibuloacral dysplasia (Agarwal et al. 2003; Shackleton et al. 2005). Duplication of LMNB1 encoding lamin B1 leading to increased expression of the protein causes adult onset autosomal dominant leukodystrophy, a slowly progressive neurological disorder characterized

by symmetrical widespread myelin loss in the central nervous system (Padiath et al. 2006). Heterozygous mutations or polymorphisms in LMNB2 encoding lamin B2 have been also reported in patients with acquired partial lipodystrophy (Hegele et al. 2006). Overall, a number of human disorders have been described that are caused by mutations in genes encoding lamins and associated nuclear envelope proteins. This number is likely to continue to grow over time. Achalsia-Addisonianism-alacrima syndrome, familial atrial fibrillation, infantile bilateral striatal necrosis, and infection-triggered acute necrotizing encephalopathy have also been shown to result from mutations in genes encoding proteins of the nuclear pore complex, a major component of the nuclear envelope that mediates nucleocytoplasmic transport (Tullio-Pelet et al. 2000; Cronshaw and Matunis 2003; Basel-Vanagaite et al. 2006; Zhang et al. 2008; Neilson et al. 2009). NUCLEAR ENVELOPE FUNCTION AND DISEASE PATHOGENESIS

Studies of laminopathies have provided insights into novel functions of the nuclear envelope. Perhaps most significantly, these studies strongly suggest that the intermediate filament nuclear lamina, although serving as a structural support for the nuclear membranes, must have additional functions. Because very different disease phenotypes can result from alterations in lamins, the nuclear lamina likely has cell-type and tissue-selective properties. Like all intermediate filament proteins, A-type lamins have a relatively small head domain, a conserved a-helical rod domain, and a tail domain. Lamins A and C are identical for the first 566 amino acids, sharing the head, rod, and first portion of the tail domain, with lamin C having six unique carboxy-terminal amino acids and prelamin A having 98 unique carboxy-terminal residues. Examination of the predominant genotype-phenotype correlations for alterations in lamin A structure strongly suggests that different domains of the proteins have different tissue-selective functions (Fig. 1). Most mutations that cause striated muscle

Laminopathies and Aging

CMT FPLD

Myopathies

Head domain Rod domain

lg-like fold

Tail domain Δ

CaaX ZMPSTE24 cleavage site MAD HGPS

Figure 1. Schematic diagram of a prelamin A molecule with mutations causing laminopathies indicated. Mutations

causing Dunnigan-type famililal partial lipodystrophy (FPLD, dark blue asterisks), Charcot-Marie-Tooth disease (CMT, orange asterisk), mandibuloacral dysplasia (MAD, light blue asterisks), or Hutchinson-Gilford progeria syndrome (HGPS, purple asterisk) are found in specific areas of the molecule, whereas mutations causing myopathies (red asterisks) are found throughout. Mutation data are from Leiden Muscular Dystrophy pages (http://www.dmd.nl/lmna_home.html). Photographs are reproduced with permission from Elsevier (Chaouch et al. 2003; Emery 2000; Novelli et al. 2002), Macmillan Publishers Ltd (Peters et al. 1998; Towbin and Bowles 2002), and the American Association for the Advancement of Science (De Sandre-Giovannoli et al. 2003).

diseases lead to amino-acid substitutions, small deletions, splice site alterations, or truncations throughout lamins A and C. Approximately 90% of the mutations that cause Dunnigan-type familial partial lipodystrophy generate aminoacid substitutions within an immunoglobulintype fold in the tails of lamins A and C. Most mutations causing peripheral neuropathy lead to the R298C substitution in the rod domain and most mutations causing mandibuloacral dysplasia generate amino-acid substitutions at or very near amino-acid residue 527 in the

immunoglobulin-type fold. Classical HutchinsonGilford progeria syndrome is caused by mutations within exon 11 of LMNA, leading to an in frame deletion of 50 amino acids from the tail of prelamin A. The distribution of LMNA mutations causing striated muscle diseases suggests that these lead to a defect in overall lamina structure in cells. Homozygous Lmna knockout mice develop regional skeletal and cardiac muscle abnormalities within the first 2 months of life and their heterozygous littermates develop

¨ stlund, and Y. Wang H.J. Worman, C. O

cardiomyopathy with conduction block at much older ages (Sullivan et al. 1999; Wolf et al. 2008). Heterozygous mutations leading to significant lamin truncations in humans also cause striated muscle disease (Bonne et al. 1999; Jakobs et al. 2001; MacLeod et al. 2003). Expression of missense lamin A variants that cause muscle disease disrupts the structure of the nuclear lamina and leads to morphological alterations similar to those in cells from which ¨ stlund et al. A-type lamins are depleted (O 2001; Raharjo et al. 2001). Transgenic expression of a lamin A variant encoded by a LMNA mutation that causes cardiomyopathy and muscular dystrophy in humans alters lamina structure and induces severe heart disease in mice (Wang et al. 2006). These observations suggest that muscle disease results either from a partial loss of A-type lamins or expression of variants that “dominantly interfere” with the overall structure and function of the nuclear lamina. It remains generally accepted that one function of the lamina is to provide structural support to the nuclear envelope. One hypothesis for the pathogenesis of striated muscle disease is that a defective lamina fails to properly carry out this support function. Fibroblasts from Lmna knockout mice, transfected cells that express lamin A variants and fibroblasts from human subjects with LMNA mutations and muscle diseases, all have abnormal nuclear morphology at the light microscopy level ¨ stlund et al. 2001; Raharjo (Sullivan et al. 1999; O et al. 2001; Muchir et al. 2004). Fibroblasts from Lmna knockout mice also have decreased mechanical stiffness (Broers et al. 2004; Lammerding et al. 2004; Lee et al. 2007). Abnormalities in the nuclear lamina could even potentially affect cytoskeleton functions, as the lamina is connected to cytoplasmic actin via the nesprin isoforms and Sun proteins that span the perinuclear space (Crisp et al. 2006). Depletion of A-type lamins indeed disrupts cytoskeletal processes such as cellular migration and nuclear positioning (Lee et al. 2007; Houben et al. 2009). In hearts and to a lesser extent in skeletal muscle from Lmna H222P knockin mice, a model of Emery-Dreifuss muscular dystrophy,

there is abnormal activation of the MAP kinases ERK1/2 and JNK (Muchir et al. 2007b). These MAP kinases are activated by mechanical stress in cardiomyocytes (Baines and Molkentin 2005). Chronically increased ERK and JNK activation is detrimental to hearts and treatment of Lmna H222P knockin mice with an inhibitor of ERK signaling prevents development of cardiomyopathy (Muchir et al. 2009). Altered nuclear envelope elasticity is also caused by loss of emerin, which binds to A-type lamins, and this could contribute to increased nuclear fragility in humans subjects with mutations in EDM and striated muscle disease (Rowat et al. 2006). ERK is also abnormally activated in hearts of mice lacking emerin (Muchir et al. 2007a). Structural alterations in the nuclear envelope and connected cytoskeleton resulting from LMNA or EDM mutations may therefore make cells such as cardiomyocytes highly susceptible to damage by recurrent mechanical stress, leading to the activation of stress-response pathways that are further detrimental to cells over time. Data from subjects with Dunnigan-type familial partial lipodystrophy suggest that the immunoglobulin-type fold in the tail of A-type lamins has specific functions in adipose cells. LMNA mutations responsible for approximately 90% of cases lead to amino-acid substitutions that decrease the positive charge of a solvent-exposed surface on the immunoglobulin-type fold but are not predicted to alter the three-dimensional structure of lamins (Dhe-Paganon et al. 2002; Krimm et al. 2002). However, they affect the ability of the immunoglobulin-type fold of A-type lamins to bind to DNA (Stierle´ et al. 2003) and could potentially alter binding of a protein important in adipocyte differentiation or survival. These mutations have in fact been shown to decrease an interaction between lamin A and SREBP1 (Lloyd et al. 2002); however, the physiological consequences of this interaction remain to be shown. LMNA mutations that cause Dunnigantype familial partial lipodystrophy may result in a “gain of function,” as overexpression of either lamin A with a causative amino-acid change or wild-type lamin A both block differentiation of preadipocytes in adipocytes in vitro

Laminopathies and Aging

(Boguslavsky et al. 2006) and deficiency of A-type lamins does not cause lipodystrophy in mice (Cutler et al. 2002). The immunoglobulintype fold of A-type lamins may therefore negatively regulate adipocyte differentiation, or survival “gain of function” may cause partial lipodystrophy. Mandibuloacral dysplasia, which has partial lipodystrophy as a predominant feature, also results from amino-acid substitutions in the immunoglobulin-type fold; however, there are other abnormalities affecting different tissues in this autosomal recessive disease requiring inheritance of two mutant LMNA alleles. Perhaps the best example of a laminopathy providing insights into nuclear envelope protein function is what progeroid disorders have taught us about the need for proper prelamin A processing. Since the early 1980s, it has been recognized that lamin A is synthesized as a precursor molecule prelamin A (Gerace et al. 1984). Prelamin A contains a CaaX motif at its carboxyl terminus, a sequence known to initiate three sequential chemical reactions (Clarke 1992; Zhang and Casey 1996; Young et al. 2005). In the first reaction, the cysteine of the carboxy-terminal CaaX motif is farnesylated

RD

CaaX

Farnesyltransferase

by protein farnesyltransferase. In the second reaction, the – aaX is clipped off. In the third, the farnesylcysteine is methylated by isoprenylcysteine carboxyl methyltransferase. The role of these three reactions in prelamin A processing is shown in Figure 2. Initially, the groups of Klaus Weber and Michael Sinensky showed that processing of prelamin A resulted from cleavage of an isoprenylated, specifically farnesylated, polypeptide 15 amino acids away from the carboxy-terminal cysteine (Weber et al. 1989; Beck et al. 1990; Sinensky et al. 1994). Sinensky’s group then characterized a farnesylation-dependent prelamin A endoprotease activity in cells (Kilic et al. 1997). In 2002, the groups of Stephen Young and Carlos Lo´pez-Otı´n reported that mice deficient in zinc metalloproteinase ZMPSTE24 were defective in the processing of prelamin A to lamin A (Bergo et al. 2002; Penda´s et al. 2002). In 2005, Sinensky’s group confirmed in vitro using recombinant ZMPSTE24 that this endoprotease clips the – aaX from prelamin A and catalyzes the second cleavage that removes the remaining 15 carboxyl-terminal amino acids (Corrigan et al. 2005). Removal of the – aaX from prelamin A is likely

WT

CaaX

Farnesyltransferase CaaX

RCE1

CaaX

C

CaaX ZMPSTE24 or RCE1

C Isoprenylcysteine carboxyl methyltransferase COCH3

COCH3

CaaX

Farnesyltransferase

ZMPSTE24 or RCE1

Isoprenylcysteine carboxyl methyltransferase

HGPS

C Isoprenylcysteine carboxyl methyltransferase COCH3

ZMPSTE24 Mature lamin A

Figure 2. Processing of prelamin A to mature lamin A in wild-type (WT) cells occurs in several steps, described in

the text (middle column). In restrictive dermopathy (RD), the ZMPSTE24 enzyme is nonfunctioning, resulting in accumulation of farnesylated prelamin A (left column). In Hutchinson-Gilford progeria syndrome (HGPS), the second cleavage site for ZMPSTE24 is deleted, resulting in accumulation of a truncated form of farnesylated prelamin A (right column).

¨ stlund, and Y. Wang H.J. Worman, C. O

redundantly catalyzed by the enzyme RCE1 (Young et al. 2005). The LMNA mutation causing HutchinsonGilford progeria syndrome that activates a cryptic splice site in exon 11 leads to an in frame deletion of 50 amino acids that contains the second ZMPSTE24 endoproteolytic site in prelamin A (De Sandre-Giovannoli et al. 2003; Eriksson et al. 2003). As a result, a farnesylated, truncated prelamin A variant that cannot be properly processed accumulates in cells (Fig. 2). Loss of ZMPSTE24 activity leads to accumulation of unprocessed, farnesylated prelamin A (Fig. 2), which causes restrictive dermopathy and other progeroid disorders that have clinical overlap with Hutchinson-Gilford progeria syndrome. Zmpste24 knockout mice have progeroid features that overlap with mice having a targeted knockin mutation of Lmna that causes Hutchinson-Gilford progeria syndrome (Yang et al. 2006). Blocking farnesylation of the truncated prelamin A or unprocessed prelamin A in these mice with chemical inhibitors improves the mouse phenotypes (Fong et al. 2006; Yang et al. 2006; Varela et al. 2008). Similarly, heterozygosity for Lmna deficiency eliminates the progeria-like phenotypes in Zmpste24 knockout mice (Fong et al. 2004). These results indicate that accumulation of farnesylated prelamin A or the truncated variant plays a key role in the pathogenesis of the progeroid phenotype. It is less clear how accumulation of farnesylated prelamin A polypeptides lead to progeria phenotypes. Cultured cells expressing these proteins have microscopic abnormalities in nuclear morphology and blocking protein farnesyltransferase activity significantly reverses these abnormalities (Eriksson et al. 2003; De Sandre-Giovannoli et al. 2003; Bridger and Kill 2004; Goldman et al. 2004; Paradisi et al. 2005; Yang et al. 2005; Capell et al. 2005; Glynn and Glover 2005; Mallampalli et al. 2005; Toth et al. 2005; Young et al. 2005; Wang et al. 2008). These nuclear morphological abnormalities are associated with reduced deformability of the lamina and increased stiffness of the nucleus (Dahl et al. 2006; Verstraeten et al. 2008). However, alterations in nuclear morphology and nuclear mechanics are not unique to progeroid

syndromes and occur as a result of LMNA mutations that cause other laminopathies. Expression of farnesylated prelamin A or the truncated variant in Hutchinson-Gilford progeria syndrome perturb DNA damage response and repair, leading to genomic instability (Liu et al. 2005), but inhibition of protein farnesylation does not appear to reduce DNA double-strand breaks or damage checkpoint signaling (Liu et al. 2006). Cells accumulating these isoprenylated A-type lamins also have altered signaling pathways involved in regulating stem cell behavior (Espada et al. 2008; Scaffidi and Misteli 2008). Although accumulation of isoprenylated prelamin A polypeptides is important in progeria pathogenesis, genetic studies show that it is not the entire explanation. Some atypical progeroid disorders resulting from LMNA mutations are not associated with accumulation of prelamin A (Verstraeten et al. 2006). Furthermore, Yang et al. (2008) elegantly showed that expression in mice of a nonfarnesylated variant of the truncated prelamin A in HutchinsonGilford progeria syndrome nonetheless causes a progeroid-like phenotype, albeit less severe than that in mice expressing the farnesylated protein. Hence, alterations in A-type lamins other than accumulation of isoprenylated forms can lead to the same cellular defects that give rise to progeroid phenotypes. CONCLUDING REMARKS

Are studies of the rare monogenic laminopathies relevant to common human diseases and physiological aging? Some data suggest that this might indeed be the case. As patients with familial dilated cardiomyopathies have been screened for genetic causes at some medical centers, LMNA mutations have been shown to be responsible for approximately ten percent of all cases and a third with atrioventricular conduction block (Arbustini et al. 2002; Taylor et al. 2003; van Tintelen et al. 2007; Parks et al. 2008). Compared with other dilated cardiomyopathies, those caused by LMNA mutations are associated with the rapid development of heart failure, early life-threatening arrhythmias, and

Laminopathies and Aging

sudden death (Be´cane et al. 2000; Taylor et al. 2003; van Berlo et al. 2005; Pasotti et al. 2008). Hence, screening for LMNA mutations as part of the clinical routine could provide information that leads to early placement of a pacemaker and an implantable cardioverter defibrillator to prevent sudden death (Meune et al. 2006). Given that mutations in genes encoding nuclear envelope proteins cause rare monogenic diseases, it is possible that polymorphic variants of the same genes predispose or contribute quantitatively to the development of common diseases. As LMNA mutations cause rare lipodystrophy disorders, several groups have examined if LMNA polymorphisms contribute to the development of common disorders. Although not completely conclusive, some studies suggest that polymorphic variations in LMNA may predispose to insulin resistance, diabetes mellitus, and metabolic syndrome (Duesing et al. 2008; Steinle et al. 2004; Wegner et al. 2007; Owen et al. 2007; Mesa et al. 2007; Murase et al. 2002). Hence, subtle alterations in A-type lamins may contribute to the pathogenesis of diseases that are endemic in the developed world. One could similarly hypothesize that polymorphisms in genes encoding other nuclear envelope proteins could contribute to other common diseases. For example, subtle alterations in MAN1, loss of function of which causes sclerosing bone dysplasias, could hypothetically contribute to the development of osteoporosis. The involvement of A-type lamins in the pathogenesis of progeroid syndromes has raised interest about their role in physiological aging. The abnormal RNA splicing occurring as a result of the LMNA mutations that cause Hutchinson-Gilford progeria syndrome takes place at very low levels in normal cells (McClintock et al. 2007; Scaffidi and Misteli 2006). One study has shown that the truncated prelamin A that results from this RNA splicing event accumulates in dermal fibroblasts and keratinocytes in older individuals (McClintock et al. 2007). Fibroblasts from older normal subjects have also been reported to show defects similar to those in cells from subjects with Hutchinson-Gilford progeria syndrome, such as abnormal nuclear morphology, increased

DNA damage, and changes in histone modifications (Scaffidi and Misteli 2006). These findings support a hypothesis that low-level expression of the truncated prelamin A generated as a result of the LMNA mutation that causes HutchinsonGilford progeria syndrome may contribute to aspects of physiological aging. In closing, a word of caution is warranted about extrapolating data from laminopathies to common diseases. For example, children with Hutchinson-Gilford progeria syndrome have normal cognitive and other brain functions, whereas central nervous system degeneration is a major feature of normal human aging. Elevated blood levels of total cholesterol, C-reactive protein, and low density lipoprotein do not appear to contribute to the accelerated vascular occlusive disease in HutchinsonGilford progeria syndrome (Gordon et al. 2005) and affected vessels show pathological features that are unusual for typical atherosclerosis (Stehbens et al. 1999). Hence, HutchinsonGilford progeria syndrome may not be a perfectly accurate model to understand some of the major complications of physiological human aging. Similarly, other laminopathies that share phenotypic features with common human disorders may have different underlying pathogenic mechanisms. Nonetheless, research on this fascinating group of rare diseases is clearly providing clues about fundamental functions of the nuclear envelope as well as relevant insights into cellular processes that must be at least partly involved in certain aspects of common diseases and aging. ACKNOWLEDGMENTS

The authors were supported by grants from the National Institutes of Health (AR048997, NS059352, and AG025240) and a grant from the Muscular Dystrophy Association (MDA4287) to H.J. Worman. REFERENCES Agarwal AK, Fryns JP, Auchus RJ, Garg A. 2003. Zinc metalloproteinase, ZMPSTE24, is mutated in mandibuloacral dysplasia. Hum Mol Genet 12: 1995– 2001.

¨ stlund, and Y. Wang H.J. Worman, C. O Agarwal AK, Kazachkova I, Ten S, Garg A. 2008. Severe mandibuloacral dysplasia-associated lipodystrophy and progeria in a young girl with a novel homozygous Arg527Cys LMNA mutation. J Clin Endocrinol Metab 93: 4617–4623. Arbustini E, Pilotto A, Repetto A, Grasso M, Negri A, Diegoli M, Campana C, Scelsi L, Baldini E, Gavazzi A, Tavazzi L. 2002. Autosomal dominant dilated cardiomyopathy with atrioventricular block: A lamin A/C defect-related disease. J Am Coll Cardiol 39: 981– 990. Astejada MN, Goto K, Nagano A, Ura S, Noguchi S, Nonaka I, Nishino I, Hayashi YK. 2007. Emerinopathy and laminopathy clinical, pathological and molecular features of muscular dystrophy with nuclear envelopathy in Japan. Acta Myol 26: 159–164. Attali R, Warwar N, Israel A, Gurt I, McNally E, Puckelwartz M, Glick B, Nevo Y, Ben-Neriah Z, Melki J. 2009. Mutation of SYNE-1, encoding an essential component of the nuclear lamina, is responsible for autosomal recessive arthrogryposis. Hum Mol Genet 18: 3462– 3469. Baines CP, Molkentin JD. 2005. STRESS signaling pathways that modulate cardiac myocyte apoptosis. J Mol Cell Cardiol 38: 47–62. Basel-Vanagaite L, Muncher L, Straussberg R, PasmanikChor M, Yahav M, Rainshtein L, Walsh CA, Magal N, Taub E, Drasinover V, et al. 2006. Mutated nup62 causes autosomal recessive infantile bilateral striatal necrosis. Ann Neurol 60: 214– 222. Beadle GW, Tatum EL. 1941. Genetic control of biochemical reactions in Neurospora. Proc Natl Acad Sci 27: 499 –506. Be´cane HM, Bonne G, Varnous S, Muchir A, Ortega V, Hammouda EH, Urtizberea JA, Lavergne T, Fardeau M, Eymard B, et al. 2000. High incidence of sudden death with conduction system and myocardial disease due to lamins A and C gene mutation. Pacing Clin Electrophysiol 23: 1661–1666. Beck LA, Hosick TJ, Sinensky M. 1990. Isoprenylation is required for the processing of the lamin A precursor. J Cell Biol 110: 1489– 1499. Bergo MO, Gavino B, Ross J, Schmidt WK, Hong C, Kendall LV, Mohr A, Meta M, Genant H, Jiang Y, et al. 2002. Zmpste24 deficiency in mice causes spontaneous bone fractures, muscle weakness, and a prelamin A processing defect. Proc Natl Acad Sci 99: 13049– 13054. Bione S, Maestrini E, Rivella S, Mancini M, Regis S, Romeo G, Toniolo D. 1994. Identification of a novel X-linked gene responsible for Emery-Dreifuss muscular dystrophy. Nat Genet 8: 323– 327. Bione S, Small K, Aksmanovic VM, D’Urso M, Ciccodicola A, Merlini L, Morandi L, Kress W, Yates JR, Warren ST, et al. 1995. Identification of new mutations in the EmeryDreifuss muscular dystrophy gene and evidence for genetic heterogeneity of the disease. Hum Mol Genet 4: 1859– 1863. Boguslavsky RL, Stewart CL, Worman HJ. 2006. Nuclear lamin A inhibits adipocyte differentiation: Implications for Dunnigan-type familial partial lipodystrophy. Hum Mol Genet 15: 653– 663. Bonne G, Di Barletta MR, Varnous S, Be´cane HM, Hammouda EH, Merlini L, Muntoni F, Greenberg CR, Gary F, Urtizberea JA, et al. 1999. Mutations in the gene encoding

lamin A/C cause autosomal dominant Emery-Dreifuss muscular dystrophy. Nat Genet 21: 285 –288. Bonne G, Mercuri E, Muchir A, Urtizberea A, Be´cane HM, Recan D, Merlini L, Wehnert M, Boor R, Reuner U, et al. 2000. Clinical and molecular genetic spectrum of autosomal dominant Emery-Dreifuss muscular dystrophy due to mutations of the lamin A/C gene. Ann Neurol 48: 170– 180. Bridger JM, Kill IR. 2004. Aging of Hutchinson –Gilford progeria syndrome fibroblasts is characterised by hyperproliferation and increased apoptosis. Exp Gerontol 39: 717 –724. Brodsky GL, Muntoni F, Miocic S, Sinagra G, Sewry C, Mestroni L. 2000. Lamin A/C gene mutation associated with dilated cardiomyopathy with variable skeletal muscle involvement. Circulation 101: 473–476. Broers JL, Peeters EA, Kuijpers HJ, Endert J, Bouten CV, Oomens CW, Baaijens FP, Ramaekers FC. 2004. Decreased mechanical stiffness in LMNA-/- cells is caused by defective nucleo-cytoskeletal integrity: Implications for the development of laminopathies. Hum Mol Genet 13: 2567– 2580. Cao H, Hegele RA. 2000. Nuclear lamin A/C R482Q mutation in Canadian kindreds with Dunnigan-type familial partial lipodystrophy. Hum Mol Genet 9: 109 –112. Cao H, Hegele RA. 2003. LMNA is mutated in HutchinsonGilford progeria (MIM 176670) but not in WiedemannRautenstrauch progeroid syndrome (MIM 264090). J Hum Genet 48: 271 –274. Capell BC, Erdos MR, Madigan JP, Fiordalisi JJ, Varga R, Conneely KN, Gordon LB, Der CJ, Cox AD, Collins FS. 2005. Inhibiting farnesylation of progerin prevents the characteristic nuclear blebbing of Hutchinson– Gilford progeria syndrome. Proc Natl Acad Sci 102: 12879– 12884. Caputo S, Couprie J, Duband-Goulet I, Konde´ E, Lin F, Braud S, Gondry M, Gilquin B, Worman HJ, Zinn-Justin S. 2006. The carboxyl-terminal nucleoplasmic region of MAN1 exhibits a DNA binding winged helix domain. J Biol Chem 281: 18208– 18215. Chaouch M, Allal Y, De Sandre-Giovannoli A, Vallat JM, Amer-el-Khedoud A, Kassouri N, Chaouch A, Sindou P, Hammadouche T, Tazir M, et al. 2003. The phenotypic manifestations of autosomal recessive axonal Charcot-Marie-Tooth due to a mutation in Lamin A/C gene. Neuromuscul Disord 13: 60– 67. Chen L, Lee L, Kudlow BA, Dos Santos HG, Sletvold O, Shafeghati Y, Botha EG, Garg A, Hanson NB, Martin GM, et al. 2003. LMNA mutations in atypical Werner’s syndrome. Lancet 362: 440– 445. Clarke S. 1992. Protein isoprenylation and methylation at carboxyl-terminal cysteine residues. Annu Rev Biochem 61: 355– 386. Clements L, Manilal S, Love DR, Morris GE. 2000. Direct interaction between emerin and lamin A. Biochem Biophys Res Commun 267: 709–714. Corrigan DP, Kuszczak D, Rusinol AE, Thewke DP, Hrycyna CA, Michaelis S, Sinensky MS. 2005. Prelamin A endoproteolytic processing in vitro by recombinant Zmpste24. Biochem J 387: 129– 138.

Laminopathies and Aging Crisp M, Liu Q, Roux K, Rattner JB, Shanahan C, Burke B, Stahl PD, Hodzic D. 2006. Coupling of the nucleus and cytoplasm: Role of the LINC complex. J Cell Biol 172: 41–53. Cronshaw JM, Matunis MJ. 2003. The nuclear pore complex protein ALADIN is mislocalized in triple A syndrome. Proc Natl Acad Sci 100: 5823–5827. Csoka AB, Cao H, Sammak PJ, Constantinescu D, Schatten GP, Hegele RA. 2004. Novel lamin A/C gene (LMNA) mutations in atypical progeroid syndromes. J Med Genet 41: 304 –308. Cutler DA, Sullivan T, Marcus-Samuels B, Stewart CL, Reitman ML. 2002. Characterization of adiposity and metabolism in Lmna-deficient mice. Biochem Biophys Res Commun 291: 522 –527. Dahl KN, Scaffidi P, Islam MF, Yodh AG, Wilson KL, Misteli T. 2006. Distinct structural and mechanical properties of the nuclear lamina in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci 103: 10271–10276. DeBusk FL. 1972. The Hutchinson-Gilford progeria syndrome. J Pediat 80: 697–724. De Sandre-Giovannoli A, Bernard R, Cau P, Navarro C, Amiel J, Boccaccio I, Lyonnet S, Stewart CL, Munnich A, Le Merrer M, et al. 2003. Lamin a truncation in Hutchinson-Gilford progeria. Science 300: 2055. De Sandre-Giovannoli A, Chaouch M, Kozlov S, Vallat JM, Tazir M, Kassouri N, Szepetowski P, Hammadouche T, Vandenberghe A, Stewart CL, et al. 2002. Homozygous defects in LMNA, encoding lamin A/C nuclear-envelope proteins, cause autosomal recessive axonal neuropathy in human (Charcot-Marie-Tooth disorder type 2) and mouse. Am J Hum Genet 70: 726 –736. Dhe-Paganon S, Werner ED, Chi YI, Shoelson SE. 2002. Structure of the globular tail of nuclear lamin. J Biol Chem 277: 17381– 17384. Duesing K, Charpentier G, Marre M, Tichet J, Hercberg S, Froguel P, Gibson F. 2008. Evaluating the association of common LMNA variants with type 2 diabetes and quantitative metabolic phenotypes in French Europids. Diabetologia 51: 76– 81. Dunnigan MG, Cochrane MA, Kelly A, Scott JW. 1974. Familial lipoatrophic diabetes with dominant transmission. A new syndrome. Q J Med 43: 33– 48. Emery AE. 2000. Emery-Dreifuss muscular dystrophy—a 40 year retrospective. Neuromuscul Disord 10: 228–232. Emery AE, Dreifuss FE. 1966. Unusual type of benign x-linked muscular dystrophy. J Neurol Neurosurg Psychiatry 29: 338–342. Eriksson M, Brown WT, Gordon LB, Glynn MW, Singer J, Scott L, Erdos MR, Robbins CM, Moses TY, Berglund P, et al. 2003. Recurrent de novo point mutations in lamin A cause Hutchinson-Gilford progeria syndrome. Nature 423: 293– 298. Espada J, Varela I, Flores I, Ugalde AP, Cadin˜anos J, Penda´s AM, Stewart CL, Tryggvason K, Blasco MA, Freije JM, et al. 2008. Nuclear envelope defects cause stem cell dysfunction in premature-aging mice. J Cell Biol 181: 27– 35. Fairley EA, Kendrick-Jones J, Ellis JA. 1999. The EmeryDreifuss muscular dystrophy phenotype arises from aberrant targeting and binding of emerin at the inner nuclear membrane. J Cell Sci 112: 2571–2582.

Fatkin D, MacRae C, Sasaki T, Wolff MR, Porcu M, Frenneaux M, Atherton J, Vidaillet HJ Jr, Spudich S, De Girolami U, et al. 1999. Missense mutations in the rod domain of the lamin A/C gene as causes of dilated cardiomyopathy and conduction-system disease. N Engl J Med 341: 1715– 1724. Fong LG, Frost D, Meta M, Qiao X, Yang SH, Coffinier C, Young SG. 2006. A protein farnesyltransferase inhibitor ameliorates disease in a mouse model of progeria. Science 311: 1621– 1623. Fong LG, Ng JK, Meta M, Cote´ N, Yang SH, Stewart CL, Sullivan T, Burghardt A, Majumdar S, Reue K, et al. 2004. Heterozygosity for Lmna deficiency eliminates the progeria-like phenotypes in Zmpste24-deficient mice. Proc Natl Acad Sci 101: 18111–18116. Garg A, Cogulu O, Ozkinay F, Onay H, Agarwal AK. 2005. A novel homozygous Ala529Val LMNA mutation in Turkish patients with mandibuloacral dysplasia. J Clin Endocrinol Metab 90: 5259–5264. Gerace L, Comeau C, Benson M. 1984. Organization and modulation of nuclear lamina structure. J Cell Sci Suppl 1: 137– 160. Gilford H. 1904. Ateleiosis and progeria: Continuous youth and premature old age. Brit Med J 2: 914– 918. Glynn MW, Glover TW. 2005. Incomplete processing of mutant lamin A in Hutchinson–Gilford progeria leads to nuclear abnormalities, which are reversed by farnesyltransferase inhibition. Hum Mol Genet 14: 2959–2969. Goldman RD, Shumaker DK, Erdos MR, Eriksson M, Goldman AE, Gordon LB, Gruenbaum Y, Khuon S, Mendez M, Varga R, et al. 2004. Accumulation of mutant lamin A causes progressive changes in nuclear architecture in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci 101: 8963–8968. Gonzalez-Alegre P, Paulson HL. 2004. Aberrant cellular behavior of mutant torsinA implicates nuclear envelope dysfunction in DYT1 dystonia. J Neurosci 24: 2593–2601. Goodchild RE, Dauer WT. 2004. Mislocalization to the nuclear envelope: An effect of the dystonia-causing torsinA mutation. Proc Natl Acad Sci 101: 847– 852. Goodchild RE, Dauer WT. 2005. The AAAþ protein torsinA interacts with a conserved domain present in LAP1 and a novel ER protein. J Cell Biol 168: 855– 862. Goodchild RE, Kim CE, Dauer WT. 2005. Loss of the dystonia-associated protein torsinA selectively disrupts the neuronal nuclear envelope. Neuron 48: 923–932. Gordon LB, Harten IA, Patti ME, Lichtenstein AH. 2005. Reduced adiponectin and HDL cholesterol without elevated C-reactive protein: Clues to the biology of premature atherosclerosis in Hutchinson-Gilford Progeria Syndrome. J Pediatr 146: 336 –341. Gros-Louis F, Dupre´ N, Dion P, Fox MA, Laurent S, Verreault S, Sanes JR, Bouchard JP, Rouleau GA. 2007. Mutations in SYNE1 lead to a newly discovered form of autosomal recessive cerebellar ataxia. Nat Genet 39: 80– 85. Hegele RA, Cao H, Liu DM, Costain GA, Charlton-Menys V, Rodger NW, Durrington PN. 2006. Sequencing of the reannotated LMNB2 gene reveals novel mutations in patients with acquired partial lipodystrophy. Am J Hum Genet 79: 383– 389.

¨ stlund, and Y. Wang H.J. Worman, C. O Hellemans J, Preobrazhenska O, Willaert A, Debeer P, Verdonk PC, Costa T, Janssens K, Menten B, Van Roy N, Vermeulen SJ, et al. 2004. Loss-of-function mutations in LEMD3 result in osteopoikilosis, Buschke-Ollendorff syndrome and melorheostosis. Nat Genet 36: 1213– 1238. Hoffmann K, Dreger CK, Olins AL, Olins DE, Shultz LD, Lucke B, Karl H, Kaps R, Mu¨ller D, Vaya´ A, et al. 2002. Mutations in the gene encoding the lamin B receptor produce an altered nuclear morphology in granulocytes (Pelger-Hue¨t anomaly). Nat Genet 31: 410– 414. Holmer L, Pezhman A, Worman HJ. 1998. The human lamin B receptor/sterol reductase multigene family. Genomics 54: 469 –476. Houben F, Willems CH, Declercq IL, Hochstenbach K, Kamps MA, Snoeckx LH, Ramaekers FC, Broers JL. 2009. Disturbed nuclear orientation and cellular migration in A-type lamin deficient cells. Biochim Biophys Acta 1793: 312 –324. Hutchinson J. 1886. Case of congenital absence of hair, with atrophic condition of the skin and its appendages, in a boy whose mother had been almost wholly bald from alopecia areata from the age of six. Lancet I: 923. Jackson SN, Pinkney J, Bargiotta A, Veal CD, Howlett TA, McNally PG, Corral R, Johnson A, Trembath RC. 1998. A defect in the regional deposition of adipose tissue (partial lipodystrophy) is encoded by a gene at chromosome 1q. Am J Hum Genet 63: 534– 540. Jakobs PM, Hanson EL, Crispell KA, Toy W, Keegan H, Schilling K, Icenogle TB, Litt M, Hershberger RE. 2001. Novel lamin A/C mutations in two families with dilated cardiomyopathy and conduction system disease. J Card Fail 7: 249– 256. Kilic F, Dalton MB, Burrell SK, Mayer JP, Patterson SD, Sinensky M. 1997. In vitro assay and characterization of the farnesylation-dependent prelamin A endoprotease. J Biol Chem 272: 5298– 5304. ¨ stlund C, Gilquin B, Couprie J, Hossenlopp P, Krimm I, O Mornon JP, Bonne G, Courvalin JC, Worman HJ, Zinn-Justin S. 2002. The Ig-like structure of the C-terminal domain of lamin A/C, mutated in muscular dystrophies, cardiomyopathy, and partial lipodystrophy. Structure 10: 811 –823. Lammerding J, Schulze PC, Takahashi T, Kozlov S, Sullivan T, Kamm RD, Stewart CL, Lee RT. 2004. Lamin A/C deficiency causes defective nuclear mechanics and mechanotransduction. J Clin Invest 113: 370–378. Lee JS, Hale CM, Panorchan P, Khatau SB, George JP, Tseng Y, Stewart CL, Hodzic D, Wirtz D. 2007. Nuclear lamin A/C deficiency induces defects in cell mechanics, polarization, and migration. Biophys J 93: 2542– 2552. Lin F, Worman HJ. 1993. Structural organization of the human gene encoding nuclear lamin A and nuclear lamin C. J Biol Chem 268: 16321–16326. Lin F, Blake DL, Callebaut I, Skerjanc IS, Holmer L, McBurney MW, Paulin-Levasseur M, Worman HJ. 2000. MAN1, an inner nuclear membrane protein that shares the LEM domain with lamina-associated polypeptide 2 and emerin. J Biol Chem 275: 4840–4847. Lin F, Morrison JM, Wu W, Worman HJ. 2005. MAN1, an integral protein of the inner nuclear membrane, binds Smad2 and Smad3 and antagonizes transforming growth factor-b signaling. Hum Mol Genet 14: 437 –445.

Liu Y, Rusinol A, Sinensky M, Wang Y, Zou Y. 2006. DNA damage responses in progeroid syndromes arise from defective maturation of prelamin A. J Cell Sci 119: 4644–4649. Liu B, Wang J, Chan KM, Tjia WM, Deng W, Guan X, Huang JD, Li KM, Chau PY, Chen DJ, et al. 2005. Genomic instability in laminopathy-based premature aging. Nat Med 11: 780– 785. Lloyd DJ, Trembath RC, Shackleton S. 2002. A novel interaction between lamin A and SREBP 1: Implications for partial lipodystrophy and other laminopathies. Hum Mol Genet 11: 769 –777. Lombardi F, Gullotta F, Columbaro M, Filareto A, D’Adamo M, Vielle A, Guglielmi V, Nardone AM, Azzolini V, Grosso E, et al. 2007. Compound heterozygosity for mutations in LMNA in a patient with a myopathic and lipodystrophic mandibuloacral dysplasia type A phenotype. J Clin Endocrinol Metab 92: 4467–4471. MacLeod HM, Culley MR, Huber JM, McNally EM. 2003. Lamin A/C truncation in dilated cardiomyopathy with conduction disease. BMC Med Genet 4: 4. Makri S, Clarke NF, Richard P, Maugenre S, Demay L, Bonne G, Guicheney P. 2009. Germinal mosaicism for LMNA mimics autosomal recessive congenital muscular dystrophy. Neuromuscul Disord 19: 26– 28. Mallampalli MP, Huyer G, Bendale P, Gelb MH, Michaelis S. 2005. Inhibiting farnesylation reverses the nuclear morphology defect in a HeLa cell model for Hutchinson– Gilford progeria syndrome. Proc Natl Acad Sci 102: 14416– 14421. Manilal S, Nguyen TM, Sewry CA, Morris GE. 1996. The Emery-Dreifuss muscular dystrophy protein, emerin, is a nuclear membrane protein. Hum Mol Genet 5: 801 –808. Mansharamani M, Wilson KL. 2005. Direct binding of nuclear membrane protein MAN1 to emerin in vitro and two modes of binding to barrier-to-autointegration factor. J Biol Chem 280: 13863– 13870. McClintock D, Ratner D, Lokuge M, Owens DM, Gordon LB, Collins FS, Djabali K. 2007. The mutant form of lamin A that causes Hutchinson-Gilford progeria is a biomarker of cellular aging in human skin. PLoS One 2: e1269. McKusick VA. 1952. The clinical observations of Jonathan Hutchinson. Am J Syph Gonorrhea Vener Dis 36: 101 –126. Merideth MA, Gordon LB, Clauss S, Sachdev V, Smith AC, Perry MB, Brewer CC, Zalewski C, Kim HJ, Solomon B, et al. 2008. Phenotype and course of Hutchinson-Gilford progeria syndrome. N Engl J Med 358: 592– 604. Mesa JL, Loos RJ, Franks PW, Ong KK, Luan J, O’Rahilly S, Wareham NJ, Barroso I. 2007. Lamin A/C polymorphisms, type 2 diabetes, and the metabolic syndrome: Case-control and quantitative trait studies. Diabetes 56: 884 –889. Meune C, Van Berlo JH, Anselme F, Bonne G, Pinto YM, Duboc D. 2006. Primary prevention of sudden death in patients with lamin A/C gene mutations. N Engl J Med 354: 209–210. Muchir A, Bonne G, van der Kooi AJ, van Meegen M, Baas F, Bolhuis PA, de Visser M, Schwartz K. 2000. Identification of mutations in the gene encoding lamins A/C in

Laminopathies and Aging autosomal dominant limb girdle muscular dystrophy with atrioventricular conduction disturbances (LGMD1B). Hum Mol Genet 9: 1453–1459. Muchir A, Medioni J, Laluc M, Massart C, Arimura T, van der Kooi AJ, Desguerre I, Mayer M, Ferrer X, Briault S, et al. 2004. Nuclear envelope alterations in fibroblasts from patients with muscular dystrophy, cardiomyopathy, and partial lipodystrophy carrying lamin A/C gene mutations. Muscle Nerve 30: 444 –450. Muchir A, Worman HJ. 2007. Emery-Dreifuss muscular dystrophy. Curr Neurol Neurosci Rep 7: 78–83. Muchir A, Pavlidis P, Bonne G, Hayashi YK, Worman HJ. 2007a. Activation of MAPK in hearts of EMD null mice: Similarities between mouse models of X-linked and autosomal dominant Emery Dreifuss muscular dystrophy. Hum Mol Genet 16: 1884–1895. Muchir A, Pavlidis P, Decostre V, Herron AJ, Arimura T, Bonne G, Worman HJ. 2007b. Activation of MAPK pathways links LMNA mutations to cardiomyopathy in Emery-Dreifuss muscular dystrophy. J Clin Invest 117: 1282– 1293. Muchir A, Shan J, Bonne G, Lehnart SE, Worman HJ. 2009. Inhibition of extracellular signal-regulated kinase signaling to prevent cardiomyopathy caused by mutation in the gene encoding A-type lamins. Hum Mol Genet 18: 241–247. Murase Y, Yagi K, Katsuda Y, Asano A, Koizumi J, Mabuchi H. 2002. An LMNA variant is associated with dyslipidemia and insulin resistance in the Japanese. Metabolism 51: 1017–1021. Nagano A, Koga R, Ogawa M, Kurano Y, Kawada J, Okada R, Hayashi YK, Tsukahara T, Arahata K. 1996. Emerin deficiency at the nuclear membrane in patients with EmeryDreifuss muscular dystrophy. Nat Genet 12: 254–259. Naismith TV, Heuser JE, Breakefield XO, Hanson PI. 2004. TorsinA in the nuclear envelope. Proc Natl Acad Sci 101: 7612–7617. Navarro CL, Cadin˜anos J, De Sandre-Giovannoli A, Bernard R, Courrier S, Boccaccio I, Boyer A, Kleijer WJ, Wagner A, Giuliano F, et al. 2005. Loss of ZMPSTE24 (FACE-1) causes autosomal recessive restrictive dermopathy and accumulation of Lamin A precursors. Hum Mol Genet 14: 1503–1513. Neilson DE, Adams MD, Orr CM, Schelling DK, Eiben RM, Kerr DS, Anderson J, Bassuk AG, Bye AM, Childs AM, et al. 2009. Infection-triggered familial or recurrent cases of acute necrotizing encephalopathy caused by mutations in a component of the nuclear pore, RANBP2. Am J Hum Genet 84: 44– 51. Novelli G, Muchir A, Sangiuolo F, Helbling-Leclerc A, D’Apice MR, Massart C, Capon F, Sbraccia P, Federici M, Lauro R, et al. 2002. Mandibuloacral dysplasia is caused by a mutation in LMNA-encoding lamin A/C. Am J Hum Genet 71: 426 –431. Oosterwijk JC, Mansour S, van Noort G, Waterham HR, Hall CM, Hennekam RC. 2003. Congenital abnormalities reported in Pelger-Hue¨t homozygosity as compared to Greenberg/HEM dysplasia: highly variable expression of allelic phenotypes. J Med Genet 40: 937 –941. ¨ stlund C, Bonne G, Schwartz K, Worman HJ. 2001. PropO erties of lamin A mutants found in Emery-Dreifuss

muscular dystrophy, cardiomyopathy and Dunnigantype partial lipodystrophy. J Cell Sci 114: 4435– 4445. Owen KR, Groves CJ, Hanson RL, Knowler WC, Shuldiner AR, Elbein SC, Mitchell BD, Froguel P, Ng MC, Chan JC, et al. 2007. Common variation in the LMNA gene (encoding lamin A/C) and type 2 diabetes: Association analyses in 9,518 subjects. Diabetes 56: 879–883. Ozelius LJ, Hewett JW, Page CE, Bressman SB, Kramer PL, Shalish C, de Leon D, Brin MF, Raymond D, Corey DP, et al. 1997. The early-onset torsion dystonia gene (DYT1) encodes an ATP-binding protein. Nat Genet 17: 40– 48. Padiath QS, Saigoh K, Schiffmann R, Asahara H, Yamada T, Koeppen A, Hogan K, Pta´cek LJ, Fu YH. 2006. Lamin B1 duplications cause autosomal dominant leukodystrophy. Nat Genet 38: 1114–1123. Pan D, Este´vez-Salmero´n LD, Stroschein SL, Zhu X, He J, Zhou S, Luo K. 2005. The integral inner nuclear membrane protein MAN1 physically interacts with the R-Smad proteins to repress signaling by the transforming growth factor-{b} superfamily of cytokines. J Biol Chem 280: 15992–16001. Paradisi M, McClintock D, Boguslavsky RL, Pedicelli C, Worman HJ, Djabali K. 2005. Dermal fibroblasts in Hutchinson– Gilford progeria syndrome with the lamin A G608G mutation have dysmorphic nuclei and are hypersensitive to heat stress. BMC Cell Biol 6: 27. Parks SB, Kushner JD, Nauman D, Burgess D, Ludwigsen S, Peterson A, Li D, Jakobs P, Litt M, Porter CB, et al. 2008. Lamin A/C mutation analysis in a cohort of 324 unrelated patients with idiopathic or familial dilated cardiomyopathy. Am Heart J 156: 161 –169. Pasotti M, Klersy C, Pilotto A, Marziliano N, Rapezzi C, Serio A, Mannarino S, Gambarin F, Favalli V, Grasso M, et al. 2008. Long-term outcome and risk stratification in dilated cardiolaminopathies. J Am Coll Cardiol 52: 1250–1260. Penda´s AM, Zhou Z, Cadin˜anos J, Freije JM, Wang J, Hultenby K, Astudillo A, Wernerson A, Rodrı´guez F, Tryggvason K, et al. 2002. Defective prelamin A processing and muscular and adipocyte alterations in Zmpste24 metalloproteinase-deficient mice. Nat Genet 31: 94–99. Peters JM, Barnes R, Bennett L, Gitomer WM, Bowcock AM, Garg A. 1998. Localization of the gene for familial partial lipodystrophy (Dunnigan variety) to chromosome 1q21-22. Nat Genet 18: 292– 295. Quijano-Roy S, Mbieleu B, Bo¨nnemann CG, Jeannet PY, Colomer J, Clarke NF, Cuisset JM, Roper H, De Meirleir L, D’Amico A, et al. 2008. De novo LMNA mutations cause a new form of congenital muscular dystrophy. Ann Neurol 64: 177 –186. Raharjo WH, Enarson P, Sullivan T, Stewart CL, Burke B. 2001. Nuclear envelope defects associated with LMNA mutations cause dilated cardiomyopathy and EmeryDreifuss muscular dystrophy. J Cell Sci 114: 4447– 4457. Raffaele di Barletta M, Ricci E, Galluzzi G, Tonali P, Mora M, Morandi L, Romorini A, Voit T, Orstavik KH, Merlini L, et al. 2000. Different mutations in the LMNA gene cause autosomal dominant and autosomal recessive EmeryDreifuss muscular dystrophy. Am J Hum Genet 66: 1407–1412.

¨ stlund, and Y. Wang H.J. Worman, C. O Renou L, Stora S, Yaou RB, Volk M, Sinkovec M, Demay L, Richard P, Peterlin B, Bonne G. 2008. Heart-hand syndrome of Slovenian type: A new kind of laminopathy. J Med Genet 45: 666– 671. Rowat AC, Lammerding J, Ipsen JH. 2006. Mechanical properties of the cell nucleus and the effect of emerin deficiency. Biophys J 91: 4649–4664. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059– 1063. Scaffidi P, Misteli T. 2008. Lamin A-dependent misregulation of adult stem cells associated with accelerated ageing. Nat Cell Biol 10: 452–459. Shackleton S, Lloyd DJ, Jackson SN, Evans R, Niermeijer MF, Singh BM, Schmidt H, Brabant G, Kumar S, Durrington PN, et al. 2000. LMNA, encoding lamin A/C, is mutated in partial lipodystrophy. Nat Genet 24: 153 –156. Shackleton S, Smallwood DT, Clayton P, Wilson LC, Agarwal AK, Garg A, Trembath RC. 2005. Compound heterozygous ZMPSTE24 mutations reduce prelamin A processing and result in a severe progeroid phenotype. J Med Genet 42: e36. Sinensky M, Fantle K, Trujillo M, McLain T, Kupfer A, Dalton M. 1994. The processing pathway of prelamin A. J Cell Sci 107: 61–67. Speckman RA, Garg A, Du F, Bennett L, Veile R, Arioglu E, Taylor SI, Lovett M, Bowcock AM. 2000. Mutational and haplotype analyses of families with familial partial lipodystrophy (Dunnigan variety) reveal recurrent missense mutations in the globular C-terminal domain of lamin A/C. Am J Hum Genet 66: 1192–1198. Stehbens WE, Wakefield SJ, Gilbert-Barness E, Olson RE, Ackerman J. 1999. Histological and ultrastructural features of atherosclerosis in progeria. Cardiovasc Pathol 8: 29–39. Steinle NI, Kazlauskaite R, Imumorin IG, Hsueh WC, Pollin TI, O’Connell JR, Mitchell BD, Shuldiner AR. 2004. Variation in the lamin A/C gene: Associations with metabolic syndrome. Arterioscler Thromb Vasc Biol 24: 1708– 1713. ¨ stlund C, Krimm I, Zinn-Justin S, Stierle´ V, Couprie J, O Hossenlopp P, Worman HJ, Courvalin JC, Duband-Goulet I. 2003. The carboxyl-terminal region common to lamins A and C contains a DNA binding domain. Biochemistry 42: 4819–4828. Sullivan T, Escalante-Alcalde D, Bhatt H, Anver M, Bhat N, Nagashima K, Stewart CL, Burke B. 1999. Loss of A-type lamin expression compromises nuclear envelope integrity leading to muscular dystrophy. J Cell Biol 147: 913–920. Taylor MR, Fain PR, Sinagra G, Robinson ML, Robertson AD, Carniel E, Di Lenarda A, Bohlmeyer TJ, Ferguson DA, Brodsky GL, et al. 2003. Natural history of dilated cardiomyopathy due to lamin A/C gene mutations. J Am Coll Cardiol 41: 771 –780. Taylor MR, Slavov D, Gajewski A, Vlcek S, Ku L, Fain PR, Carniel E, Di Lenarda A, Sinagra G, Boucek MM, et al. Familial Cardiomyopathy Registry Research Group. 2005. Thymopoietin (lamina-associated polypeptide 2) gene mutation associated with dilated cardiomyopathy. Hum Mutat 26: 566– 574. Tazir M, Azzedine H, Assami S, Sindou P, Nouioua S, Zemmouri R, Hamadouche T, Chaouch M, Feingold J,

Vallat JM, et al. 2004. Phenotypic variability in autosomal recessive axonal Charcot-Marie-Tooth disease due to the R298C mutation in lamin A/C. Brain 127: 154 –163. Toth JI, Yang SH, Qiao X, Beigneux AP, Gelb MH, Moulson CL, Miner JH, Young SG, Fong LG. 2005. Blocking protein farnesyl transferase improves nuclear shape in fibroblasts from humans with progeroid syndromes. Proc Natl Acad Sci 102: 12873–12878. Towbin JA, Bowles NE. 2002. The failing heart. Nature 415: 227 –233. Tullio-Pelet A, Salomon R, Hadj-Rabia S, Mugnier C, de Laet MH, Chaouachi B, Bakiri F, Brottier P, Cattolico L, Penet C, et al. 2000. Mutant WD-repeat protein in triple-A syndrome. Nat Genet 26: 332–335. van Berlo JH, de Voogt WG, van der Kooi AJ, van Tintelen JP, Bonne G, Yaou RB, Duboc D, Rossenbacker T, Heidbu¨chel H, de Visser M, et al. 2005. Meta-analysis of clinical characteristics of 299 carriers of LMNA gene mutations: Do lamin A/C mutations portend a high risk of sudden death? J Mol Med 83: 79– 83. van Tintelen JP, Hofstra RM, Katerberg H, Rossenbacker T, Wiesfeld AC, du Marchie Sarvaas GJ, Wilde AA, van Langen IM, Nannenberg EA, van der Kooi AJ, et al. 2007. High yield of LMNA mutations in patients with dilated cardiomyopathy and/or conduction disease referred to cardiogenetics outpatient clinics. Am Heart J 154: 1130– 1139. Varela I, Pereira S, Ugalde AP, Navarro CL, Sua´rez MF, Cau P, Cadin˜anos J, Osorio FG, Foray N, Cobo J, et al. 2008. Combined treatment with statins and aminobisphosphonates extends longevity in a mouse model of human premature aging. Nat Med 14: 767–772. Verstraeten VL, Broers JLV, Steensel MAM, Zinn-Justin S, Ramaekers FCS, Steijlen PM, Kamps M, Kuijpers HJH, Merckx D, Smeets HJM, et al. 2006. Compound heterozygosity for mutations in LMNA causes a progeria syndrome without prelamin A accumulation. Hum Mol Genet 15: 2509–2522. Verstraeten VL, Ji JY, Cummings KS, Lee RT, Lammerding J. 2008. Increased mechanosensitivity and nuclear stiffness in Hutchinson-Gilford progeria cells: Effects of farnesyltransferase inhibitors. Aging Cell 7: 383– 393. Vigouroux C, Capeau J. 2005. A-type lamin-linked lipodystrophies. Novartis Found Symp 264: 166–177. Vigouroux C, Magre´ J, Vantyghem MC, Bourut C, Lascols O, Shackleton S, Lloyd DJ, Guerci B, Padova G, Valensi P, et al. 2000. Lamin A/C gene: Sex-determined expression of mutations in Dunnigan-type familial partial lipodystrophy and absence of coding mutations in congenital and acquired generalized lipoatrophy. Diabetes 49: 1958– 1962. Wang Y, Herron AJ, Worman HJ. 2006. Pathology and nuclear abnormalities in hearts of transgenic mice expressing M371K lamin A encoded by an LMNA mutation causing Emery-Dreifuss muscular dystrophy. Hum Mol Genet 15: 2479–2489. Wang Y, Panteleyev AA, Owens DM, Djabali K, Stewart CL, Worman HJ. 2008. Epidermal expression of the truncated prelamin A causing Hutchinson-Gilford progeria syndrome: Effects on keratinocytes, hair and skin. Hum Mol Genet 17: 2357– 2369.

Laminopathies and Aging Waterham HR, Koster J, Mooyer P, Noort Gv G, Kelley RI, Wilcox WR, Wanders RJ, Hennekam RC, Oosterwijk JC. 2003. Autosomal recessive HEM/Greenberg skeletal dysplasia is caused by 3 b-hydroxysterol d14-reductase deficiency due to mutations in the lamin B receptor gene. Am J Hum Genet 72: 1013– 1017. Weber K, Plessmann U, Traub P. 1989. Maturation of nuclear lamin A involves a specific carboxy-terminal trimming, which removes the polyisoprenylation site from the precursor; implications for the structure of the nuclear lamina. FEBS Lett 257: 411–414. Wegner L, Andersen G, Sparsø T, Grarup N, Glu¨mer C, Borch-Johnsen K, Jørgensen T, Hansen T, Pedersen O. 2007. Common variation in LMNA increases susceptibility to type 2 diabetes and associates with elevated fasting glycemia and estimates of body fat and height in the general population: Studies of 7,495 Danish whites. Diabetes 56: 694 –698. Wolf CM, Wang L, Alcalai R, Pizard A, Burgon PG, Ahmad F, Sherwood M, Branco DM, Wakimoto H, Fishman GI, et al. 2008. Lamin A/C haploinsufficiency causes dilated cardiomyopathy and apoptosistriggered cardiac conduction system disease. J Mol Cell Cardiol 44: 293–303. Worman HJ, Evans CD, Blobel G. 1990. The lamin B receptor of the nuclear envelope inner membrane: A polytopic protein with eight potential transmembrane domains. J Cell Biol 111: 1535–1542. Worman HJ, Yuan J, Blobel G, Georgatos SD. 1988. A lamin B receptor in the nuclear envelope. Proc Natl Acad Sci 85: 8531– 8534. Wydner KL, McNeil JA, Lin F, Worman HJ, Lawrence JB. 1996. Chromosomal assignment of human nuclear envelope protein genes LMNA, LMNB1, and LBR by fluorescence in situ hybridization. Genomics 32: 474– 478.

Yang SH, Andres DA, Spielmann HP, Young SG, Fong LG. 2008. Progerin elicits disease phenotypes of progeria in mice whether or not it is farnesylated. J Clin Invest 118: 3291–3300. Yang SH, Bergo MO, Toth JI, Qiao X, Hu Y, Sandoval S, Meta M, Bendale P, Gelb MH, Young SG, et al. 2005. Blocking protein farnesyltransferase improves nuclear blebbing in mouse fibroblasts with a targeted Hutchinson–Gilford progeria syndrome mutation. Proc Natl Acad Sci 102: 10291– 10296. Yang SH, Meta M, Qiao X, Frost D, Bauch J, Coffinier C, Majumdar S, Bergo MO, Young SG, Fong LG. 2006. A farnesyltransferase inhibitor improves disease phenotypes in mice with a Hutchinson-Gilford progeria syndrome mutation. J Clin Invest 116: 2115–2121. Ye Q, Worman HJ. 1994. Primary structure analysis and lamin B and DNA binding of human LBR, an integral protein of the nuclear envelope inner membrane. J Biol Chem 269: 11306–11311. Ye Q, Worman HJ. 1996. Interaction between an integral protein of the nuclear envelope inner membrane and human chromodomain proteins homologous to Drosophila HP1. J Biol Chem 271: 14653–14656. Young SG, Fong LG, Michaelis S. 2005. Prelamin A, Zmpste24, misshapen cell nuclei, and progeria—New evidence suggesting that protein farnesylation could be important for disease pathogenesis. J Lipid Res 46: 2531–2558. Zhang FL, Casey PJ. 1996. Protein prenylation: Molecular mechanisms and functional consequences. Annu Rev Biochem 65: 241– 269. Zhang X, Chen S, Yoo S, Chakrabarti S, Zhang T, Ke T, Oberti C, Yong SL, Fang F, Li L, et al. 2008. Mutation in nuclear pore component NUP155 leads to atrial fibrillation and early sudden cardiac death. Cell 135: 1017–1027.

Higher-order Genome Organization in Human Disease Tom Misteli National Cancer Institute, NIH, Bethesda, Maryland 20892 Correspondence: [email protected]

Genomes are organized into complex higher-order structures by folding of the DNA into chromatin fibers, chromosome domains, and ultimately chromosomes. The higher-order organization of genomes is functionally important for gene regulation and control of gene expression programs. Defects in how chromatin is globally organized are relevant for physiological and pathological processes. Mutations and transcriptional misregulation of several global genome organizers are linked to human diseases and global alterations in chromatin structure are emerging as key players in maintenance of genome stability, aging, and the formation of cancer translocations.

enomes in their native state are folded into complex higher-order structures. Hierarchical folding of DNA gives rise to chromatin fibers, chromosomes domains, and eventually chromosomes (reviewed in Felsenfeld and Groudine 2003; Belmont 2006; Woodcock 2006). At the lowest level, DNA is wrapped around an octamer of core histone proteins, which are the primary architectural elements of the chromatin fiber, to form a nucleosome. Multiple nucleosomes are linked by stretches of DNA, often occupied by a linker histone, into a beads-on-a string fiber of 10 nm in diameter. This primary fiber is then further compacted onto itself to form higher-order fibers of various diameters, although their precise geometry in vivo is unknown (reviewed in Belmont 2006; Woodcock 2006). At the next level of organization, the chromatin fiber folds into subchromosomal domains of 1 Mb in

G

size (reviewed in Cremer and Cremer 2001; Cremer et al. 2006). These subchromosomal domains in turn are folded to give rise to an interphase chromosome. In mammalian cells, the degree of fiber compaction from naked DNA to a chromosome is estimated to be on the order of 10,000-fold (reviewed in Belmont 2006). During interphase, chromosomes exist as chromosome territories (reviewed in Cremer and Cremer 2001; Meaburn and Misteli 2007; Misteli 2007). A chromosome territory is defined as the nuclear space taken up by the DNA of a given chromosome. The term “territory” refers to the fact that the occupied space is compact, typically roughly ovoid in shape, with a volume of about 2– 3 mm in diameter (reviewed in Cremer and Cremer 2001; Meaburn and Misteli 2007; Misteli 2007). The internal structure of chromosome territories is

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000794 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000794

T. Misteli

poorly understood but likely consists of a highly interconnected and branched network of channels, which form between looping chromatin fibers (reviewed in Cremer and Cremer 2001; Meaburn and Misteli 2007; Misteli 2007). This relatively open structure allows access of gene regulatory factors into the interior of chromosome territories. Although chromosome territories are discrete structures within the nucleus, neighboring chromosomes can overlap considerably and chromatin loops from one territory can easily invade the body of the neighboring territory (Visser et al. 2000; Branco and Pombo 2006). The higher-order organization of the genome into chromatin fibers and chromosomes is now known to critically contribute to gene regulation (reviewed in Fraser and Bickmore 2007; Lanctot et al. 2007). It is therefore not surprising that defects in higher-order chromatin and chromosome organization cause disease. There is an increasing list of diseases in which changes in histone modifications have been documented, although it is often not clear whether these alterations are a cause or merely a side-effect of the disease process. Furthermore, although many disease-related epigenetic changes have implicitly been assumed to result in changes in higher-order chromatin structure, it is often not clear whether the observed disease phenotypes are because of structural chromatin defects or because of altered gene expression caused by local changes in histone modifications at particular genes. This article focuses primarily on disease mechanisms that involve bona-fide structural defects in chromatin. ARCHITECTURAL CHROMATIN PROTEINS IN DISEASE

The higher-order structure of chromatin is mediated by a multitude of architectural chromatin proteins (reviewed in Woodcock 2006; Hock et al. 2007; Fanti and Pimpinelli 2008; Phillips and Corces 2009). Although some of these proteins, such as linker histones, highmobility group proteins, and heterochromatin protein 1, decorate the chromatin fiber extensively, others associate with the fiber only

locally and are often involved in formation of chromatin loops. Looping is a prominent and powerful mechanism of gene regulation because loops can bring together distantly located genome regions and in this way lead to physical proximity of a regulatory sequence and its target (reviewed in Misteli 2007; Phillips and Corces 2009). Several key architectural chromatin proteins, many involved in looping, have been implicated in disease. SATB1

SATB1 (Special AT-rich binding protein 1) is a thymocyte specific DNA-binding protein that was initially characterized because of its propensity to bind to AT-rich sequences that unwind in response to superhelical tension (Dickinson et al. 1992). This unique binding property already hinted at a potential structural role for SATB1 in higher-order genome organization. Such a role was confirmed by the finding that SATB1 forms a “cagelike” matrix within the nucleus of thymocytes and serves to anchor, via loop formation, a large number of genes whose activity is tightly regulated during T-cell differentiation (Cai et al. 2003). Regulation of several T-cell differentiation genes is dependent on SATB1 and its loss leads to dramatic differentiation defects. Although some of these effects are likely because of a role of SATB1 in recruiting histone modifying activities and chromatin remodeling complexes, some of its regulatory function has been attributed to its ability to promote formation of higher-order chromatin loops (Cai et al. 2006). For example, the activation of the T helper-cell cytokine locus 2 during T-cell differentiation requires the superposition of an upstream Locus Control Region (LCR) with its promoter region. SATB1 is required for this looping event and is found associated with the locus at what is likely the base of the loop (Cai et al. 2006). An unresolved issue is whether SATB1 acts itself as a structural scaffold for these loops or as an adaptor to an underlying structural component. SATB1 appears to be a critical player in cancer. The expression level of SATB1 correlates with poor prognosis in breast cancer and its

Genome Organization in Disease

expression induces aggressive growth of breast cells in vitro, whereas its depletion reverses cancer metastasis in animal models (Han et al. 2008). In metastatic breast cancer cells, SATB1 regulates a large number of genes predominantly involved in cell adhesion, cellular signaling, and cell-cycle regulation. SATB1 binds near many of these genes and its depletion leads to changes in their epigenetic makeup (Han et al. 2008). An attractive, although at present unproven, model is that SATB1 acts as a structural platform providing a base for chromatin loops and in this way tethers its target genes into a shared regulatory environment. Given that SATB1 is only expressed at relatively low levels in breast tissue, it will be interesting to determine whether the protein plays a similar role in other tumor types and whether it affects the same set of genes in other tumors. There are currently no reported mutations in SATB1 in cancer patients and it will be important to uncover the mechanism by which SATB1 is misregulated in cancer. CTCF

CTCF (CCCTC-binding factor) is a zinc-finger protein with a central, evolutionarily highly conserved, DNA-binding domain (reviewed in Phillips and Corces 2009). The protein is widely expressed, although levels and localization within the nucleus are tissue- and cell-type specific. Both loss of CTCF as well as its overexpression have severe and global effects on gene expression profiles and lead to physiological defects in development and differentiation. Increasing evidence suggests that CTCT acts mainly as an insulator protein blocking the action of enhancers on a gene target or to buffer genes from adjacent regulatory sequences (reviewed in Phillips and Corces 2009). Various observations suggest that CTCF exerts this insulator function via chromatin looping. CTCF forms homodimers and multimers on chromatin and CTCF-mediated chromatin loops have been shown for various genes including mouse b-globin, human MHC class II, and the imprinted H19/Igf2 locus (Kurukuti et al. 2006; Splinter et al. 2006; Majumder et al.

2008). Because of its ability to readily form loops and because CTCF is widely bound throughout the genome with between 15,000 – 25,000 binding sites (Xie et al. 2005; Heintzman et al. 2009), it is a strong candidate for a global genome organizer (reviewed in Phillips and Corces 2009). This role is particularly strongly supported by the finding that CTCF can also mediate interchromosomal interactions. Genomewide studies have indicated that the imprinted H19/Ig2 locus on human chromosome 11 physically interacts with regulatory elements on other chromosomes and that these interchromosomal interactions are critical for efficient imprinting (Ling et al. 2006; Zhao et al. 2006). CTCF associates preferentially with the repressed, maternal allele of H19/Igf2 in a DNA methylation-dependent manner and most of the interchromosomal interactions occur at the maternal allele and are dependent on CTCF, suggesting that the protein plays a key role in its repression (Yoon et al. 2007; Li et al. 2008). It thus appears that CTCF acts as a global genome organizer by forming intrachromosomal loops and interchromosomal interactions. CTCF has been implicated in various diseases. Trinucleotide repeat expansion diseases including Huntington’s disease, fragile X mental retardation, and myotonic dystrophy are caused by excessive lengthening of microsatellite repeat sequences (reviewed in Orr and Zoghbi 2007). For example, although a healthy individual has a stretch of fewer than 27 CAG repeats in their HTT gene, a Huntington’s disease patient has typically more than 35. The trigger for repeat expansion is unknown. Recent evidence suggests that CTCF might contribute to generating fragile sites within repeats, thus facilitating their expansion (Libby et al. 2008). Mutation of a CTCF-binding site near a repeat leads to increased genomic instability and increased repeat length, similar to that seen in disease situations. Although not tested, it is possible that interference with CTCF binding, either by mutation of its target site or mutations in an interacting partner, may contribute to trinucleotide repeat diseases (Libby et al. 2008). Defects in CTCF, and other genome organizers,

T. Misteli

may also play a key role in destabilizing expanded microsatellite repeats in other trinucleotide repeat diseases (reviewed in Dion and Wilson 2009). CTCF has also been suggested to play a role in two human syndromes, Silver-Russell (SRS) and Beckwith-Wiedeman Syndrome (BWS) (Sparago et al. 2004; Eggermann et al. 2008). BWS is a developmental disorder with variable clinical symptoms including increased frequency of tumors, macroglossia, and overgrowth. It is caused by aberrant activation of the normally maternally imprinted H19/Igf2 locus on chromosome 11. Several sporadic and familial mutations have been characterized in the H19/Igf2 region, which lead to the loss of CTCF binding sites, directly implicating CTCF in the disease (Sparago et al. 2004; Eggermann et al. 2008). Similarly, in SRS, a developmental disorder characterized by severe growth retardation and body asymmetry caused by activation of the paternal allele of Igf2, mutations in CTCF binding sites have been identified (Scho¨nherr et al. 2008). CTCF has also been implicated in cancer (Witcher and Emerson 2009). The tumor suppressor p16 is often up-regulated in cancer cells. This up-regulation coincides with a loss of the chromatin boundary definition around the locus leading to its activation and loss of chromatin structure in the vicinity of the locus (Witcher and Emerson 2009). CTCF associates with the boundaries of the transcriptionally active p16 gene, but not its inactive form, and p16 activity correlates with CTCF binding. In contrast, reduction of CTCF nearly ablates p16 expression (Witcher and Emerson 2009). Interestingly, the extent of CTCF binding in cancer cells is modulated by methylation of its binding site and cancer cells are well know to have global methylation defects. Although these observations strongly point to a role of CTCF in regulation of the tumor suppressor p16, other cancer relevant genes appear to be regulated in a similar fashion because silencing of the tumor suppressors RASSF1A and CDH1 also correlate with CTCF binding (Witcher and Emerson 2009). These observations allow for the possibility that CTCF plays a critical role in

establishing and maintaining the higher-order chromatin structure around p16 and other tumor suppressor genes and in this way contributes to tumor prevention (Witcher and Emerson 2009). Whether mutations in CTCF have an oncogenic effect is unknown. Cohesin

Cohesin is a multiprotein complex involved in establishment and maintenance of pairing of sister chromatids during DNA replication and into mitosis (reviewed in McNairn and Gerton 2008; Peters et al. 2008). The evolutionarily conserved complex consists of four core proteins, which are thought to form a ring around the replicated chromatids holding them together, and several accessory proteins involved in loading and maintenance of the complex (reviewed in McNairn and Gerton 2008; Peters et al. 2008). Although the traditional role of cohesin is in holding chromatids together, alternate roles are emerging. Cohesin is expressed at robust levels in postmitotic cells that do not have a requirement for chromatid cohesion. Furthermore, genome-wide mapping studies in several species show nonrandom association of cohesin with the genome and its distribution is responsive to ongoing transcription, suggesting its involvement in higher-order chromosome organization. In Saccharomyces cerevisiae cohesin is predominantly associated with intergenic regions and binds near convergently transcribed genes, as it does in Schizosaccharomyces pombe (Glynn et al. 2004; Lengronne et al. 2004; D’Ambrosio et al. 2008; Schmidt et al. 2009). In mammalian cells, cohesin colocalizes strikingly with CTCF in insulator regions (Parelho et al. 2008; Wendt et al. 2008) and in Drosophila melanogaster, the complex is generally found near highly transcribed regions of the genome (Misulovin et al. 2008). Further evidence for a role of cohesion in genome organization comes from the finding that a mutation in the cohesion subunit Smc1 leads to loss of boundary function at the silent mating type locus in yeast and mutations in the Scc2/Nipped-B subunit leads to promoter-enhancer defects in D. melanogaster (Donze et al. 1999; Rollins et al. 1999).

Genome Organization in Disease

Mutations in cohesin components are the cause of two major multisystem developmental disorders, Cornelia de Lange Syndrome (CLS) and Roberts Syndrome (RS) (Krantz et al. 2004; Tonkin et al. 2004; Vega et al. 2005). CLS is a dominant disorder with severe growth defects and mental retardation; RS is a rare recessive disorder characterized by pre- and postnatal growth deficiency and mental retardation. However, patient cells do not show chromatin cohesion or segregation defects, suggesting that the disease symptoms are related to the noncohesion functions of the cohesin complex (reviewed in Dorsett and Krantz 2009). A likely scenario is that mutations in cohesin components weaken the complex’s association with target sites throughout the genome and in this way interfere with its boundary and insulator function, possibly by cooperating with CTCF (reviewed in McNairn and Gerton 2008; Dorsett and Krantz 2009). Failure to properly define gene boundaries is predicted to result in widespread misexpression of genes and depletion of cohesin in human cells indeed leads to altered expression of several hundred genes (Wendt et al. 2008). Interestingly, cohesin is increasingly implicated in development via control of several developmentally regulated genes, possibly explaining the pre- and postnatal growth and development defects typically seen in CLS and RS (reviewed in Dorsett 2009). C-MYC

C-MYC is one of the prototypical oncogenes. It encodes a basic helix-loop-helix zipper transcription factor and plays a role in numerous critical events in cancer including proliferation, cell cycle regulation, and apoptosis (reviewed in Eilers and Eisenman 2008). It has generally been assumed that c-myc exerts its function by acting on a few selected target genes whose misregulation would largely explain the oncogene’s detrimental effects (reviewed in Eilers and Eisenman 2008). However, genome-wide mapping of c-myc binding sites has revealed that rather than binding to a few target genes, c-myc binds to as many as 15% of human genes and also associates with intergenic regions, suggesting

other functions for c-myc (Knoepfler 2007; reviewed in Eilers and Eisenman 2008). Several other observations also suggest that c-myc might have a more global function in higherorder chromatin structure. In knockout cells, neuronal stem cells show striking defects in nuclear architecture and chromatin organization including reduced nuclear size, aberrant nuclear morphology, and wide-spread chromatin condensation (Knoepfler et al. 2002; Knoepfler et al. 2006). Importantly, these changes appear to be independent of functional defects on proliferation and apoptosis, suggesting that they are not secondary in nature, further supported by the fact that acute disruption of c-myc recapitulates chromatin defects (Knoepfler et al. 2002; Knoepfler et al. 2006). At the molecular level, loss of c-myc results in reduction of histone H3 and H4 acetylation but a marked increase in trimethylation of histone H3 lysine 9, consistent with an increase in heterochromatin in these cells. These observations strongly suggest that c-myc is a regulator of global chromatin structure and contributes to maintaining euchromatin in an open, accessible state (Knoepfler et al. 2006; reviewed in Knoepfler 2007). The mechanism by which c-myc functions in higher-order chromatin organization is unclear. One model suggests that that the protein attracts histone modifying activities and chromatin remodeling machinery to its many binding sites spread throughout the genome, thus, exposing large portions of the genome to them (reviewed in Knoepfler 2007). Alternatively, the global effect of c-myc could be mediated by misregulation of a single, or a few, global chromatin modifiers. A candidate for this model is the histone acetyltransferase GCN5, which is a known target gene of c-myc (Knoepfler et al. 2006). Its up-regulation on loss of c-myc may contribute to some of the observed changes in histone modification and possibly chromatin structure. The involvement of cmyc in higher-order chromatin structure, and particularly its effect upon histone acetylation, is of potential clinical relevance because c-myc is misregulated in a wide range of tumors and modulators of histone modifications are

T. Misteli

increasingly explored as potential anticancer drugs. HP1

Heterochromatin protein 1 (HP1) is a prominent structural component of heterochromatin (Fanti and Pimpinelli 2008). The family of HP1 proteins is evolutionarily conserved and in higher eukaryotes exists as three isoforms: HP1a, b, and g. HP1 proteins bind specifically via their chromo-domain to trimethylated lysine 9 on histone H3, enriched in heterochromatin, but are also present interdispersed throughout the genome (reviewed in Fanti and Pimpinelli 2008). The protein is thought to crosslink multiple chromatin fibers by multimerization via its chromo-shadow domain thus leading to chromatin compaction. In addition to its structural role in heterochromatin, HP1 proteins can have both activating as well as repressive functions in gene expression. As silencing factors they are thought to condense chromatin in regulatory regions of target genes, particularly at promoters, thus preventing efficient transcription initiation. As activators, they might similarly create a particular chromatin environment conducive to transcription (reviewed in Fanti and Pimpinelli 2008). HP1 has been implicated in cancer in several ways. Reduced expression of various HP1 isoforms has been reported in multiple cancers including breast, brain, colon, and ovarian cancer (Dialynas et al. 2008). HP1 appears to be particularly important for cancer progression and the degree of HP1 loss correlates with poor prognosis, whereas increased HP1 levels correlate with reduced metastasis (Pomeroy et al. 2002; Ruginis et al. 2006). A direct causal effect of HP1 has been shown in breast cancer where experimental depletion of HP1a from poorly invasive cancer cells increased their invasion capacity, but expression of HP1a in highly invasive cells decreased their ability to do so (Kirschmann et al. 2000; Norwood et al. 2006). These observations suggest that HP1a is a metastasis suppressor. The mechanism by which HP1 mediates this effect is not fully elucidated, but appears to involve global genome

misregulation because a large number of genes with known connections to cancer and metastasis are affected by its depletion (Dialynas et al. 2008). HIGHER-ORDER CHROMATIN STRUCTURE IN DNA REPAIR AND TRANSLOCATIONS

The first line of cellular defense against genome damage is the DNA repair machinery, which recognizes DNA lesions and corrects them before lasting damage can be done. There is increasing evidence that higher-order chromatin structure plays a key role in the efficiency of DNA repair (Goodarzi et al. 2009; reviewed in Misteli and Soutoglou 2009). For one, chromatin remodeling events are a prominent part of the DNA repair cascade (reviewed in Downs et al. 2007; Misteli and Soutoglou 2009). Early on, after a double-strand break (DSB) occurs, the chromatin around the break site rapidly decondenses, presumably to facilitate access of the repair machinery to the break site. Later in the repair process, several ATP-dependent chromatin-remodeling machines, mostly of the SWI/SNF family, are recruited to the break to prepare the chromatin for resection and ligation (reviewed in Downs et al. 2007; Misteli and Soutoglou 2009). The requirement for chromatin remodeling machines in efficient repair implies that the structural properties of chromatin influence the progression of the repair process and possibly also affect the susceptibility of chromatin to insult (Fig. 1). Direct evidence for this comes from the finding that irradiation-induced DSBs located in heterochromatin are more slowly repaired than the same breaks in euchromatin (Goodarzi et al. 2008). Although no kinetic studies have been reported to date, the most likely reason for the delay is the inefficient recruitment of the repair machinery into highly compacted chromatin because of reduced accessibility of binding sites. However, there is also the possibility that DSBs in euchromatin and heterochromatin use different repair mechanisms altogether. In support of this scenario, the repair of DSB in heterochromatin is significantly more sensitive to the activity of the major DNA repair kinase ATM

Genome Organization in Disease Heterochromatin

Euchromatin

ATM

Architectural chromatin proteins Repair machinery

Repair machinery

Figure 1. Higher-order chromatin structure and DNA repair. (A) The condensation status of chromatin affects

DNA repair. If a double strand break occurs in more densely packed heterochromatin region, architectural proteins (green) such as HP1, linker histone H1, or HMG proteins, associated with these domains, prevent access of the DNA repair machinery (red) and must be removed, possibly via action of the ATM kinase. Upon removal, the DNA repair machinery can gain more immediate access to the DSBs. (B) In the less densely packed euchromatin regions, the repair machinery has freer access to the DSBs.

(Ataxia telangiectasia mutated) and slowly repaired, persistent DSBs are preferentially found within heterochromatin rather than in euchromatin (Goodarzi et al. 2009). Removal of several architectural chromatin proteins, including HP1, KAP-1, or HDAC1/2, alleviates the delay in ATM-mediated repair, suggesting that these heterochromatin components act as a barrier for efficient repair (Goodarzi et al. 2009) (Fig. 1). In line with this interpretation, HP1 is actively released from sites of DNA damage via specific phosphorylation of Thr51 and is only recruited at later time points, presumable when all critical repair factors have gained access to the DSB and normal chromatin structure needs to be re-established again (Ayoub et al. 2008; Luijsterburg et al. 2009). Further evidence that chromatin architecture affects the DNA response comes from

observations on the HMGN1 protein (Highmobility group protein 1). HMGN1 is a sequence-independent global DNA binding protein implicated in transcriptional control via regulation of histone modifications (Hock et al. 2007). Mice lacking HMGN1 have a defective DNA damage response including reduced activating phosphorylation of ATM and its downstream targets CHK1 and CHK2, two key cell-cycle checkpoint signaling kinases (Birger et al. 2003; Kim et al. 2009). HMGN1 is not preferentially recruited to DSB but still plays an important role there. Loss of the protein leads to precocious accumulation of ATM on chromatin even in the absence of a DSB (Birger et al. 2003; Kim et al. 2009). The association with ATM on chromatin is controlled by the global level of acetylation of histone H3 on K14, which is known to be affected by HMGN1 (Lim et al.

T. Misteli

2005). Further evidence for a critical role of chromatin structure in susceptibility to DNA damage and repair comes from analysis of the DNA damage response in embryonic stem cells, which are unique in that they have a strict requirement to maintain their genomes in pristine conditions because any genomic defect will be propagated as the stem cells differentiate into various tissues. ES cells lacking the linker histone H1, one of the key global architectural components of chromatin, are characterized by less compacted chromatin and, remarkably, have a heightened DNA damage response and are able to repair DSB more rapidly after irradiation (Fan et al. 2005; Murga et al. 2007). This is likely because of the global decondensation of chromatin in these cells since the effect can be mimicked by treatment of cells with the histone-deacetylase inhibitor trichostatin A, which also leads to chromatin decondensation (Murga et al. 2007). These observations suggest that chromatin structure directly affects susceptibility of the genome to damage and that different regions, such as heterochromatin and euchromatin, in the same genome respond differently to DNA damage. Defects in components involved in maintenance of higher-order chromatin structure are therefore candidates to promote mutations and therefore disease. Higher-order chromatin structure may not only be important for global susceptibility of the genome to DNA damage, but may also contribute to the formation of specific chromosomal translocations. This may occur by local changes in chromatin structure, which create local fragile sites making them highly susceptible to breakage. An intriguing observation in support of this scenario has been provided by analysis of the molecular mechanisms involved in formation of a chromosomal translocation involved in ALCL (anaplastic large cell lymphoma) (Mathas et al. 2009). Many ALCL patients carry a translocation between chromosomes 2 and 5 (t(2;5)( p23;q35)) leading to the synthesis of a fusion protein between nucleophosmin and ALK (anaplastic lymphoma kinase), constitutively activating the kinase. Intriguingly, a significant fraction of ALCL patients with indistinguishable disease

symptoms lack the t(2; 5) translocation, clearly indicating the fusion protein is not necessary nor causal for disease. In analyzing the genes near the break points on chromosomes 2 and 5, it was found that several genes adjacent to nucleophosmin and ALK were already highly expressed even in patients without the translocation (Mathas et al. 2009). Although not experimentally tested, these results suggest that the chromatin structure near the breakpoints is altered even in the absence of the translocation. Interestingly, patient cells lacking the t(2; 5) translocation were more highly susceptible to form the t(2; 5) translocation than control cells, suggesting that the transcriptional activation of the flanking genes, and presumably the altered chromatin structure in these regions, predisposed cells to formation of the translocation (Mathas et al. 2009). It is attractive to speculate, that the transcriptional activation of the break-point proximal regions were brought about by improper recruitment of transcription factors to these sites, activating them and altering the local chromatin increasing their susceptibility to breakage and translocation. In this model, the first step in formation of a chromosomal translocation is the aberrant activation of genome regions, leading to their decondensation. Direct support for this scenario comes from recent finding that translocation breakpoints in prostate cancer contain binding sites for androgen receptor (AR), a transcriptional activator. Upon treatment with androgens, AR binds to these sites leading to chromatin remodeling, which may increase the susceptibility to DNA damage (Lin et al. 2009). This model is also supported by the finding that active alleles, but not inactive ones, of the translocation prone MYC and IGH loci, whose translocation causes Burkitts lymphoma, often associate with each other in the threedimensional (3D) space of the nucleus, predisposing them for translocation (Roix et al. 2003; Osborne et al. 2007). It will be important to further test this model by determining whether other translocation breakpoints are also characterized by heightened transcription and what the molecular mechanisms in this cascade of events are.

Genome Organization in Disease

SPATIAL CHROMOSOME ORGANIZATION IN CANCER

Chromosomal abnormalities in the form of translocations and aneuploidy are a general hallmark of cancer cells. While aneuploidy is generated by unequal chromosome segregation during mitosis, translocations occur during interphase by illegitimate joining of two or multiple chromosomes containing persistently damaged DNA such as double-strand breaks (DSBs). Because the formation of a translocation requires the physical interaction of the involved chromosomes, the spatial arrangement of chromosomes, i.e. their localization relative to each other, in the nucleus may contribute to determining which chromosomes undergo translocations (reviewed in Misteli

2004; Meaburn et al. 2007). It has been proposed that translocations preferentially occur between neighboring chromosomes but not between more distantly located chromosomes (Fig. 2). This is a particularly intriguing, and important, possibility because it is now well established that genomes are nonrandomly distributed within the nuclear space in a cell-type and tissue-specific manner (reviewed in Cremer and Cremer 2001; Misteli 2007). A body of correlative evidence indeed suggests that the nonrandomness in spatial genome organization contributes to determining translocation partners (reviewed in Meaburn et al. 2007). A first hint for a role of spatial genome organization in the formation of translocations comes from the finding that in humans the acrocentric chromosomes, which contain nucleolar

No translocation

Chromosome intermingling

Formation of proximal DSBs

Illegitimate joining

Figure 2. Spatial organization of chromosomes in the formation of cancer translocations. Translocations

preferentially occur between proximally positioned chromosomes (red, green), and only rarely between distally located chromosomes (blue). Closely juxtaposed double-strand breaks (yellow stars) occurring at the interface between chromosomes create free chromosome ends, which may recombine to form a chromosome translocation by illegitimate joining.

T. Misteli

organizing regions (NORs) and cluster in nucleoli, undergo translocations to form Robertsonian translocations with higher frequency than many other chromosomes (Therman et al. 1989). This is presumably because of their tight packing in the nucleolus, increasing their probability of illegitimate joining any time two or more DSBs are not promptly repaired. Similarly, the translocation frequency of chromosomes lining the nuclear envelope is elevated compared with internally located chromosomes (Bickmore and Teague 2002). In addition to these general observations, many examples of correlations between physical proximity and translocation frequencies of specific chromosomes exist (reviewed in Meaburn et al. 2007). Mouse chromosomes MMU12, 14, and 15 are frequently involved in translocations in mouse lymphoma and these chromosomes are found in pairs or in triplet clusters in more than 60% of normal lymphocytes before the formation of a translocation, placing them in close spatial proximity and predisposing them to undergo translocations among each other once persistent DSBs occur (Parada et al. 2002). Furthermore, chromosomes MMU5 and MMU6, which are frequently involved in translocations in kidney, are neighbors in more than 40% of normal hepatocytes (Parada et al. 2004) and human chromosomes HSA12 and 16, frequently translocated in adipocyte cancers, are in close proximity in differentiated adipocytes (Kuroda et al. 2004). Translocations often occur between specific regions or genes of the two involved chromosomes and studies on the localization of the involved chromosomal breakpoints further strengthen the case for a role of spatial proximity in determining translocation frequency. The BCR and ABL genes, located on chromosomes 9 and 22, whose translocation leads to formation of a fusion protein, which is causal in chronic myeloid leukemia, are found in proximity to each other in normal hematopoietic cells at much higher frequency than would be expected based on their random distribution and significantly higher than observed for various other gene loci in the same nuclei (Lukasova et al. 1997; Neves et al. 1999; Bartova et al. 2000).

In B-cells, the PML and RARa genes, located on chromosomes 15 and 17, respectively, which translocate to cause promyelocytic leukemia, are in close proximity (Neves et al. 1999) and the MYC gene on chromosome 8 and the IGH locus on chromosome 14, whose translocation leads to Burkitts lymphoma, are frequently juxtaposed (Roix et al. 2003; Osborne et al. 2007). Importantly, two less frequent Burkitts lymphoma translocation partners, Igk on chromosome 11, and Igl on chromosome 22, show less frequent association with their translocation partner MYC correlating with their limited propensity to translocate with MYC (Roix et al. 2003). Proximity effects on translocation formation are not limited to gene loci located on different chromosomes but also apply to events on the same chromosome. The RET and the H4 genes are frequent translocation partners in thyroid tumors and are both located on chromosome 10 about 30 MB apart but are brought together in 3D space by looping of the chromosome fiber, thus juxtapositioning, and predisposing, them to undergo a mutual translocation resulting in an internal chromosome reversion (Nikiforova et al. 2000). In addition to these correlative observations, direct evidence for a role of proximity comes from irradiation experiments in cells from anaplastic large cell lymphoma (ALCL). When cells which contain nonrandom pairs of closely positioned chromosome 2 and 5 were irradiation to induce DSBs, these two chromosomes formed translocations with significantly higher frequency than in cells where they were distant from each other, suggesting that their proximity facilitated their translocation (Mathas et al. 2009). A role for proximity in determining translocation outcome is in line with the wellestablished fact that translocation frequencies differ among tissues (Mitelman et al. 2007). For example, although translocations between mouse chromosomes 12 and 14 are often associated with lymphoma, translocations between chromosomes 5 and 6 are most prominently found in cancers of the kidney. The tissue-specificity of these translocations is paralleled by tissue specific organization of the involved chromosomes whereby chromosomes 12 and 14 pair in

Genome Organization in Disease

lymphocytes but not in hepatocyes, but chromosomes 5 and 6 pair in hepatocytes and not lymphocytes. It seems that the preference in translocation frequency is thus a reflection of the tissue-specific arrangement of the involved translocation partners (Parada et al. 2004). The conclusion that the relative position of translocating regions is important in determining their interaction relies on the assumption that once DSBs form in a chromosome, the generated free ends are immobile and cannot widely roam the nuclear space to find other broken chromosome ends. Several lines of evidence indeed suggest that broken chromosome ends are immobile in the nucleus. Visualization of DSBs in chemically fixed and living cells, show that irradiated genome regions remain in place for extended periods of time (Nelms et al. 1998; Kruhlak et al. 2006). Furthermore, single fluorescently tagged DSBs induced by sitespecific cutting with a restriction endonuclease remained stationary within the nuclear space (Soutoglou et al. 2007). These experiments also revealed that the radius of motion of a DSB is limited to about 250 nm. The mechanisms responsible for DSB immobility are unknown but might involve tethering of DSBs to DNA repair foci. The picture emerging from these observations is that translocations occur preferentially between already closely positioned genome regions and do not involve an extensive search process during which DSBs on multiple chromosomes diffuse through the nucleus in search of other DSBs (Fig. 2). Translocations will only take place if two DSBs occur in chromatin of two adjacent, and likely intermingling, chromosomes and are not separated by more than 500 nm, corresponding to the combined radius of motion of the two DSBs (Fig. 2). This model is in line with the finding that translocation frequencies between chromosomes not only correlate with their relative position but with the degree of overlap between the chromatin of adjacent chromosomes (Branco and Pombo 2006). The genome is continuously exposed to external insults such as irradiation and chemicals, which lead to DNA damage and DSBs.

The immobility of DSBs likely acts as a protective mechanism to ensure genome stability and it explains the relatively low frequency by which translocations occur. It is estimated that the genome in an individual cell suffers several thousand DNA damage events daily, including DSBs, however, translocations occur at frequencies of 1027. If DSBs were highly mobile within the nucleus, translocation frequencies would be much higher because randomly occurring DSBs would be able to rapidly find other DSBs in the same genome to readily undergo translocations. The intrinsic immobility of DSBs in the nucleus prevents the juxtaposition of multiple broken chromosome ends and allows the DNA repair machinery to resolve the defect before a translocation partner can be found. In this sense, the higher-order organization of the chromosomes and the chromatin fiber, and their immobility, act as a guardian of the genome. HIGHER-ORDER CHROMATIN STRUCTURE IN AGING

Aging is characterized by several global changes in chromatin structure and function. Structurally, in aged cells several key architectural chromatin proteins, including the structural heterochromatin protein HP1, are lost because of posttranslational degradation (Scaffidi and Misteli 2006; Pegoraro et al. 2009). Similarly, in the premature aging disorder HutchinsonGilford Progeria Syndrome (HGPS), caused by a mutation in lamin A, one of the major architectural elements of the mammalian nucleus, the same chromatin proteins are lost and, most likely as a consequence of their absence, morphologically discernible heterochromatin blocks and peripheral heterochromatin are largely missing from HGPS patient cells (Goldman et al. 2004; Scaffidi and Misteli 2006). These defects are because of downregulation of the NUcleosome Remodeling and Deacetylase (NURD) chromatin remodeling complex (Pegoraro et al. 2009). Several components of NURD, including its catalytic histone deacetylase I (HDAC 1), are lost in normally aged and HGPS patient cells and removal

T. Misteli

of this complex from normal cells is sufficient to recapitulate most major aging-associated chromatin defects (Pegoraro et al. 2009). These observations are supported by the finding of reduced activity of several histone deacetylases in aged mice (Shen et al. 2008). Aging-related structural defects in chromatin are also seen in yeast S. cerevisiae where instability of ribosomal repeats in the nucleolus have been associated with aging (Sinclair and Guarente 1997) and in Caenorhabditis elegans where both internal and peripheral heterochromatin blocks are lost during aging (Haithcock et al. 2005). The aging-associated defects in chromatin structure have various functional consequences. To start with, aged genomes are characterized by increased DNA damage and high levels of persistent DNA breaks, possibly brought about by structural changes, which increase the susceptibility of the genome to damage. Furthermore, probably as a consequence of loss of pericentromeric heterochromatin structure, physiologically aged and premature aged cells express normally silenced heterochromatic satellite III repeats (Gaubatz and Cutler 1990; Shumaker et al. 2006). Physiologically aged and prematurely aged genomes also undergo wide-ranging changes in epigenetic modifications. Global DNA methylation is reduced in many aged mammalian tissues (Wilson and Jones 1983; Fraga and Esteller 2007) and several histone modifications are affected during aging. In particularly, and in line with the morphological changes in chromatin, the heterochromatinassociated trimethylation of histone H3 lysine 9 and the transcriptionally repressive trimethylation of histone H3 lysine 27 are largely lost in aged and prematurely aged cells (Scaffidi and Misteli 2006; Shumaker et al. 2006). Conversely, global trimethylation of H4 lysine 20 increases with age (Sarg et al. 2002). In S. cerevisiae, acetylation of H4 lysine 16 increases, and that of H3 lysine 56 decreases with age, whereas several other modifications are unchanged (Dang et al. 2009). The level of these acetylation events is dependent on the histone deacetylase Sir2 and is counteracted by the acetyltransferase Sas2 (Dang et al. 2009). Evidence that these changes in histone modifications are not merely

bystanders in the aging process but actively drive it comes from the finding that mutations in histones H3 and H4, which interfere with their acetylation on lysine 16 or lysine 56 prolong lifespan (Dang et al. 2009). Given these extensive changes in histone modifications, not surprisingly, aged cells show dramatic and global misregulation of gene expression. Although some of these changes are likely part of specific aging-related gene expression programs including inflammation and cellular stress responses, others likely occur largely stochastically because of random changes in epigenetic modifications and chromatin structure. The mechanisms that drive chromatin and epigenetic changes during aging are currently unknown (Fig. 3). It has been suggested that the epigenetic alterations are largely triggered by DNA damage (reviewed in Oberdoerffer and Sinclair 2007). In this scenario, randomly occurring DNA damage leads to chromatin remodeling and to redistribution of chromatin modifiers within the genome with modifiers being recruited away from their normal sites so that they can engage in the repair of the DNA damage and others being recruited to newly available sites. The redistribution of histone modifiers then leads to activation of aging-related gene expression programs and contribute to general, random misregulation of genes globally (Fig. 3) (reviewed in Oberdoerffer and Sinclair 2007). This model is supported by the observed redistribution of the histone deacetylase SIRT1, which has previously been implicated in aging, in response to DNA damage and its association with genes, which are deregulated during aging (Oberdoerffer et al. 2008). However, the model does not fully account for the observation that several epigenetic modifiers are lost, rather than redistributed within the genome, in normally and prematurely aged cells (Shen et al. 2008; Pegoraro et al. 2009). Furthermore, at least in the premature aging syndrome HGPS, DNA damage is a late event and is unlikely the trigger for the observed epigenetic changes because the loss of key chromatin proteins, such as NURD, occurs before DNA damage and is followed by changes in chromatin structure and epigenetic

Genome Organization in Disease Cellular stress

Chromatin remodeling

DNA damage

Histone modifier redistribution

Epigenetic status

Susceptibility

Aging-associated gene expression programs

Non-specific gene expression events

Organismal aging

Figure 3. Chromatin effects in aging. A complex network of interactions links chromatin structure to aging.

Cellular stress may directly induce changes in the epigenetic status of the genome leading to local and global chromatin remodeling, which in turn may make the genome more susceptible to DNA damage. Cellular stress may also cause DNA lesions itself. As part of the cellular response to these lesions, chromatin remodeling events occur and may lead to redistribution of epigenetic modifiers away from their regular binding sites and toward inappropriate targets, thus altering the epigenetic state of the genome. Alterations in global chromatin structure and epigenetic status lead to activation of gene expression programs including specific-aging associated programs such as activation of inflammation and cellular stress responses, but they likely also contribute to random misregulation of genes throughout the genome. These specific and nonspecific misregulation events likely act in a feed-back loop to further destabilize the epigenetic homeostasis of the aging genome.

modifications and only eventually the appearance of DNA damage (Pegoraro et al. 2009). An alternative model is that aging is primarily driven by steady loss of epigenetic modifiers brought about by continuous, often low level, exposure to cellular stressors (Fig. 3). The loss or aberrant function of epigenetic modifiers alters the makeup of the global epigenome and as a consequence global chromatin structure, including loss of heterochromatin regions. This loss of heterochromatin may make the genome more susceptible to DNA damage, which, in a feedback mechanism, triggers further global epigenetic changes including aging-associated changes in gene expression programs and random misregulation of genes globally (Fig. 3). These considerations make it clear that although the involvement of epigenetic modifications, chromatin structure and DNA damage are now clearly established as

important components of the aging process, the precise interdependencies of these events remain to be elucidated (Fig. 3). CONCLUSIONS

The organization of the genome into higherorder chromatin and chromosomes is one of the fundamental features of eukaryotic cells. Not surprisingly, proper organization of the genome is essential for faithful genome function and defects lead to disease. Structural chromatin defects may come about directly by mutations or misexpression of bona-fide architectural chromatin proteins or chromatin remodeling activities. Alternatively, chromatin structure may be compromised because of impaired function of histone modifying activities, which in turn lead to structural chromatin changes. The number of known diseases

T. Misteli

involving structural abnormalities of chromatin is likely to grow rapidly in the next few years. In many diseases involving gene misregulation, often assumed to be because of altered epigenetic modifications, chromatin structure has not been analyzed and it is possible that many “epigenetic” diseases are in reality diseases of chromatin structure. Furthermore, genome wide mapping of single-nucleotide polymorphisms is likely to reveal a significant number of novel mutations, which affect higher-order chromatin structure. In addition to chromatin structural defects leading to specific diseases, the importance of global higher-order chromatin organization as a protective means in the maintenance of genome integrity as a shield against the continued onslaught of DNA damaging agents is rapidly becoming clear. This function is emerging as an integral mechanism in pathological processes such as formation of cancer translocations and aging. Elucidation of the precise role and molecular mechanisms involved in maintaining chromatin structure and its role will be essential for our understanding of these all-important events. The involvement of higher-order chromatin structure, and nuclear architecture in general, in disease is a powerful example of how fundamental cellular processes are relevant for disease. We often think of the study of basic cellular function as a separate discipline from clinical research. However, we are now seeing a dramatic convergence of basic and disease research. As we are making progress in outlining at the molecular level the mechanisms underlying cellular function, links to, often unexpected, diseases are being discovered. On the other hand, the genomic tools, which have led to a dramatic increase in the rate of discovery of disease genes highlight the relevance of basic cellular machinery for disease because for each identified disease gene, its basic mechanisms of action must be elucidated to truly understand the disease. The combination of increasingly detailed insight into basic biological processes and the accelerated rate of disease gene discovery provides a powerful, and promising, forum for the development of novel diagnostic and therapeutic strategies in the future.

REFERENCES Ayoub N, Jeyasekharan AD, Bernal JA, Venkitaraman AR. 2008. HP1-b mobilization promotes chromatin changes that initiate the DNA damage response. Nature 453: 682 –686. Bartova E, Kozubek S, Kozubek M, Jirsova P, Lukasova E, Skalnikova M, Buchnickova K. 2000. The influence of the cell cycle, differentiation and irradiation on the nuclear location of the abl, bcr and c-myc genes in human leukemic cells. Leuk Res 24: 233 –241. Belmont AS. 2006. Mitotic chromosome structure and condensation. Curr Opin Cell Biol 18: 632 –638. Bickmore WA, Teague P. 2002. Influences of chromosome size, gene density and nuclear position on the frequency of constitutional translocations in the human population. Chromosome Res 10: 707– 715. Birger Y, West KL, Postnikov YV, Lim JH, Furusawa T, Wagner JP, Laufer CS, Kraemer KH, Bustin M. 2003. Chromosomal protein HMGN1 enhances the rate of DNA repair in chromatin. EMBO J 22: 1665– 1675. Branco MR, Pombo A. 2006. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol 4: e138. Cai S, Han HJ, Kohwi-Shigematsu T. 2003. Tissue-specific nuclear architecture and gene expression regulated by SATB1. Nat Genet 34: 42– 51. Cai S, Lee CC, Kohwi-Shigematsu T. 2006. SATB1 packages densely looped, transcriptionally active chromatin for coordinated expression of cytokine genes. Nat Genet 38: 1278–1288. Cremer T, Cremer C. 2001. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat Rev Genet 2: 292–301. Cremer T, Cremer M, Dietzel S, Muller S, Solovei I, Fakan S. 2006. Chromosome territories—a functional nuclear landscape. Curr Opin Cell Biol 18: 307 –316. D’Ambrosio C, Schmidt CK, Katou Y, Kelly G, Itoh T, Shirahige K, Uhlmann F. 2008. Identification of cis-acting sites for condensin loading onto budding yeast chromosomes. Genes Dev 22: 2215– 2227. Dang W, Steffen KK, Perry R, Dorsey JA, Johnson FB, Shilatifard A, Kaeberlein M, Kennedy BK, Berger SL. 2009. Histone H4 lysine 16 acetylation regulates cellular lifespan. Nature 459: 802– 807. Dialynas GK, Vitalini MW, Wallrath LL. 2008. Linking Heterochromatin Protein 1 (HP1) to cancer progression. Mutat Res 647: 13– 20. Dickinson LA, Joh T, Kohwi Y, Kohwi-Shigematsu T. 1992. A tissue-specific MAR/SAR DNA-binding protein with unusual binding site recognition. Cell 70: 631 –645. Dion V, Wilson JH. 2009. Instability and chromatin structure of expanded trinucleotide repeats. Trends Genet (in press). Donze D, Adams CR, Rine J, Kamakaka RT. 1999. The boundaries of the silenced HMR domain in Saccharomyces cerevisiae. Genes Dev 13: 698 –708. Dorsett D. 2009. Cohesin, gene expression and development: Lessons from Drosophila. Chromosome Res 17: 185 –200.

Genome Organization in Disease Dorsett D, Krantz ID. 2009. On the molecular etiology of Cornelia de Lange syndrome. Ann N Y Acad Sci 1151: 22–37. Downs JA, Nussenzweig MC, Nussenzweig A. 2007. Chromatin dynamics and the preservation of genetic information. Nature 447: 951–958. Eggermann T, Eggermann K, Schonherr N. 2008. Growth retardation versus overgrowth: Silver-Russell syndrome is genetically opposite to Beckwith-Wiedemann syndrome. Trends Genet 24: 195– 204. Eilers M, Eisenman RN. 2008. Myc’s broad reach. Genes Dev 22: 2755–2766. Fan Y, Nikitina T, Zhao J, Fleury TJ, Bhattacharyya R, Bouhassira EE, Stein A, Woodcock CL, Skoultchi AI. 2005. Histone H1 depletion in mammals alters global chromatin structure but causes specific changes in gene regulation. Cell 123: 1199– 1212. Fanti L, Pimpinelli S. 2008. HP1: A functionally multifaceted protein. Curr Opin Genet Dev 18: 169– 174. Felsenfeld G, Groudine M. 2003. Controlling the double helix. Nature 421: 448–453. Fraga MF, Esteller M. 2007. Epigenetics and aging: The targets and the marks. Trends Genet 23: 413– 418. Fraser P, Bickmore W. 2007. Nuclear organization of the genome and the potential for gene regulation. Nature 447: 413– 417. Gaubatz JW, Cutler RG. 1990. Mouse satellite DNA is transcribed in senescent cardiac muscle. J Biol Chem 265: 17753–17758. Glynn EF, Megee PC, Yu HG, Mistrot C, Unal E, Koshland DE, DeRisi JL, Gerton JL. 2004. Genome-wide mapping of the cohesin complex in the yeast Saccharomyces cerevisiae. PLoS Biol 2: E259. Goldman RD, Shumaker DK, Erdos MR, Eriksson M, Goldman AE, Gordon LB, Gruenbaum Y, Khuon S, Mendez M, Varga R, et al. 2004. Accumulation of mutant lamin A causes progressive changes in nuclear architecture in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci 101: 8963–8968. Goodarzi AA, Noon AT, Jeggo PA. 2009. The impact of heterochromatin on DSB repair. Biochem Soc Trans 37: 569–576. Goodarzi AA, Noon AT, Deckbar D, Ziv Y, Shiloh Y, Lobrich M, Jeggo PA. 2008. ATM signaling facilitates repair of DNA double-strand breaks associated with heterochromatin. Mol Cell 31: 167 –177. Haithcock E, Dayani Y, Neufeld E, Zahand AJ, Feinstein N, Mattout A, Gruenbaum Y, Liu J. 2005. Age-related changes of nuclear architecture in Caenorhabditis elegans. Proc Natl Acad Sci 102: 16690– 16695. Han HJ, Russo J, Kohwi Y, Kohwi-Shigematsu T. 2008. SATB1 reprogrammes gene expression to promote breast tumour growth and metastasis. Nature 452: 187– 193. Heintzman ND, Hon GC, Hawkins RD, Kheradpour P, Stark A, Harp LF, Ye Z, Lee LK, Stuart RK, Ching CW, et al. 2009. Histone modifications at human enhancers reflect global cell-type-specific gene expression. Nature 459: 108–112. Hock R, Furusawa T, Ueda T, Bustin M. 2007. HMG chromosomal proteins in development and disease. Trends Cell Biol 17: 72– 79.

Kim YC, Gerlitz G, Furusawa T, Catez F, Nussenzweig A, Oh KS, Kraemer KH, Shiloh Y, Bustin M. 2009. Activation of ATM depends on chromatin interactions occurring before induction of DNA damage. Nat Cell Biol 11: 92– 96. Kirschmann DA, Lininger RA, Gardner LM, Seftor EA, Odero VA, Ainsztein AM, Earnshaw WC, Wallrath LL, Hendrix MJ. 2000. Down-regulation of HP1Hsalpha expression is associated with the metastatic phenotype in breast cancer. Cancer Res 60: 3359–3363. Knoepfler PS. 2007. Myc goes global: New tricks for an old oncogene. Cancer Res 67: 5061– 5063. Knoepfler PS, Cheng PF, Eisenman RN. 2002. N-myc is essential during neurogenesis for the rapid expansion of progenitor cell populations and the inhibition of neuronal differentiation. Genes Dev 16: 2699– 2712. Knoepfler PS, Zhang XY, Cheng PF, Gafken PR, McMahon SB, Eisenman RN. 2006. Myc influences global chromatin structure. EMBO J 25: 2723–2734. Krantz ID, McCallum J, DeScipio C, Kaur M, Gillis LA, Yaeger D, Jukofsky L, Wasserman N, Bottani A, Morris CA, et al. 2004. Cornelia de Lange syndrome is caused by mutations in NIPBL, the human homolog of Drosophila melanogaster Nipped-B. Nat Genet 36: 631–635. Kruhlak MJ, Celeste A, Dellaire G, Fernandez-Capetillo O, Muller WG, McNally JG, Bazett-Jones DP, Nussenzweig A. 2006. Changes in chromatin structure and mobility in living cells at sites of DNA double-strand breaks. J Cell Biol 172: 823– 834. Kuroda M, Tanabe H, Yoshida K, Oikawa K, Saito A, Kiyuna T, Mizusawa H, Mukai K. 2004. Alteration of chromosome positioning during adipocyte differentiation. J Cell Sci 117: 5897–5903. Kurukuti S, Tiwari VK, Tavoosidana G, Pugacheva E, Murrell A, Zhao Z, Lobanenkov V, Reik W, Ohlsson R. 2006. CTCF binding at the H19 imprinting control region mediates maternally inherited higher-order chromatin conformation to restrict enhancer access to Igf2. Proc Natl Acad Sci 103: 10684– 10689. Lanctot C, Cheutin T, Cremer M, Cavalli G, Cremer T. 2007. Dynamic genome architecture in the nuclear space: Regulation of gene expression in three dimensions. Nat Rev Genet 8: 104– 115. Lengronne A, Katou Y, Mori S, Yokobayashi S, Kelly GP, Itoh T, Watanabe Y, Shirahige K, Uhlmann F. 2004. Cohesin relocation from sites of chromosomal loading to places of convergent transcription. Nature 430: 573 –578. Li T, Hu JF, Qiu X, Ling J, Chen H, Wang S, Hou A, Vu TH, Hoffman AR. 2008. CTCF regulates allelic expression of Igf2 by orchestrating a promoter-polycomb repressive complex 2 intrachromosomal loop. Mol Cell Biol 28: 6473–6482. Libby RT, Hagerman KA, Pineda VV, Lau R, Cho DH, Baccam SL, Axford MM, Cleary JD, Moore JM, Sopher BL, et al. 2008. CTCF cis-regulates trinucleotide repeat instability in an epigenetic manner: A novel basis for mutational hot spot determination. PLoS Genet 4: e1000257. Lim JH, West KL, Rubinstein Y, Bergel M, Postnikov YV, Bustin M. 2005. Chromosomal protein HMGN1 enhances the acetylation of lysine 14 in histone H3. EMBO J 24: 3038–3048.

T. Misteli Lin C, Yang L, Tanasa B, Hutt K, Ju B, Ohgi KA, Zhang J, Rose DW, Fu X-D, Glass CK, Rosenfeld MG. 2009. Nuclear receptor-induced chromosomal proximity and DNA breaks underlie specific translocations in cancer. Cell 139: 1069– 1083 Ling JQ, Li T, Hu JF, Vu TH, Chen HL, Qiu XW, Cherry AM, Hoffman AR. 2006. CTCF mediates interchromosomal colocalization between Igf2/H19 and Wsb1/Nf1. Science 312: 269– 272. Luijsterburg MS, Dinant C, Lans H, Stap J, Wiernasz E, Lagerwerf S, Warmerdam DO, Lindh M, Brink MC, Dobrucki JW, et al. 2009. Heterochromatin protein 1 is recruited to various types of DNA damage. J Cell Biol 185: 577– 586. Lukasova E, Kozubek S, Kozubek M, Kjeronska J, Ryznar L, Horakova J, Krahulcova E, Horneck G. 1997. Localisation and distance between ABL and BCR genes in interphase nuclei of bone marrow cells of control donors and patients with chronic myeloid leukaemia. Hum Genet 100: 525– 535. Majumder P, Gomez JA, Chadwick BP, Boss JM. 2008. The insulator factor CTCF controls MHC class II gene expression and is required for the formation of long-distance chromatin interactions. J Exp Med 205: 785–798. Mathas S, Kreher S, Meaburn KJ, Johrens K, Lamprecht B, Assaf C, Sterry W, Kadin ME, Daibata M, Joos S, et al. 2009. Gene deregulation and spatial genome reorganization near breakpoints prior to formation of translocations in anaplastic large cell lymphoma. Proc Natl Acad Sci 106: 5831–5836. McNairn AJ, Gerton JL. 2008. Cohesinopathies: One ring, many obligations. Mutat Res 647: 103–111. Meaburn KJ, Misteli T. 2007. Cell biology: Chromosome territories. Nature 445: 379 –781. Meaburn KJ, Misteli T, Soutoglou E. 2007. Spatial genome organization in the formation of chromosomal translocations. Semin Cancer Biol 17: 80–90. Misteli T. 2004. Spatial positioning: A new dimension in genome function. Cell 119: 153 –156. Misteli T. 2007. Beyond the sequence: Cellular organization of genome function. Cell 128: 787 –800. Misteli T, Soutoglou E. 2009. The emerging role of nuclear architecture in DNA repair and genome maintenance. Nat Rev Mol Cell Biol 10: 243– 254. Misulovin Z, Schwartz YB, Li XY, Kahn TG, Gause M, MacArthur S, Fay JC, Eisen MB, Pirrotta V, Biggin MD, Dorsett D. 2008. Association of cohesin and Nipped-B with transcriptionally active regions of the Drosophila melanogaster genome. Chromosoma 117: 89–102. Mitelman F, Johansson B, Mertens F. 2007. The impact of translocations and gene fusions on cancer causation. Nat Rev Cancer 7: 233– 245. Murga M, Jaco I, Fan Y, Soria R, Martinez-Pastor B, Cuadrado M, Yang SM, Blasco MA, Skoultchi AI, FernandezCapetillo O. 2007. Global chromatin compaction limits the strength of the DNA damage response. J Cell Biol 178: 1101–1108. Nelms BE, Maser RS, MacKay JF, Lagally MG, Petrini JH. 1998. In situ visualization of DNA double-strand break repair in human fibroblasts. Science 280: 590– 592.

Neves H, Ramos C, da Silva MG, Parreira A, Parreira L. 1999. The nuclear topography of ABL, BCR, PML, and RARalpha genes: evidence for gene proximity in specific phases of the cell cycle and stages of hematopoietic differentiation. Blood 93: 1197– 1207. Nikiforova MN, Stringer JR, Blough R, Medvedovic M, Fagin JA, Nikiforov YE. 2000. Proximity of chromosomal loci that participate in radiation-induced rearrangements in human cells. Science 290: 138– 141. Norwood LE, Moss TJ, Margaryan NV, Cook SL, Wright L, Seftor EA, Hendrix MJ, Kirschmann DA, Wallrath LL. 2006. A requirement for dimerization of HP1Hsalpha in suppression of breast cancer invasion. J Biol Chem 281: 18668–18676. Oberdoerffer P, Sinclair DA. 2007. The role of nuclear architecture in genomic instability and ageing. Nat Rev Mol Cell Biol 8: 692– 702. Oberdoerffer P, Michan S, McVay M, Mostoslavsky R, Vann J, Park SK, Hartlerode A, Stegmuller J, Hafner A, Loerch P, et al. 2008. SIRT1 redistribution on chromatin promotes genomic stability but alters gene expression during aging. Cell 135: 907– 918. Orr HT, Zoghbi HY. 2007. Trinucleotide repeat disorders. Annu Rev Neurosci 30: 575– 621. Osborne CS, Chakalova L, Mitchell JA, Horton A, Wood AL, Bolland DJ, Corcoran AE, Fraser P. 2007. Myc dynamically and preferentially relocates to a transcription factory occupied by Igh. PLoS Biol 5: e192. Parada L, McQueen P, Misteli T. 2004. Tissue-specific spatial organization of genomes. Genome Biology 7: R44. Parada L, McQueen P, Munson P, Misteli T. 2002. Conservation of relative chromosome positioning in normal and cancer cells. Curr Biol 12: 1692. Parelho V, Hadjur S, Spivakov M, Leleu M, Sauer S, Gregson HC, Jarmuz A, Canzonetta C, Webster Z, Nesterova T, et al. 2008. Cohesins functionally associate with CTCF on mammalian chromosome arms. Cell 132: 422– 433. Pegoraro G, Kubben L, Wickert U, Goehler H, Hoffman K, Misteli T. 2009. Aging-related chromatin defects via loss of the NURD complex. Nat Cell Biol in press. Peters JM, Tedeschi A, Schmitz J. 2008. The cohesin complex and its roles in chromosome biology. Genes Dev 22: 3089–3114. Phillips JE, Corces VG. 2009. CTCF: Master weaver of the genome. Cell 137: 1194– 1211. Pomeroy SL, Tamayo P, Gaasenbeek M, Sturla LM, Angelo M, McLaughlin ME, Kim JY, Goumnerova LC, Black PM, Lau C, et al. 2002. Prediction of central nervous system embryonal tumour outcome based on gene expression. Nature 415: 436– 442. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287– 291. Rollins RA, Morcillo P, Dorsett D. 1999. Nipped-B, a Drosophila homologue of chromosomal adherins, participates in activation by remote enhancers in the cut and Ultrabithorax genes. Genetics 152: 577– 593. Ruginis T, Taglia L, Matusiak D, Lee BS, Benya RV. 2006. Consequence of gastrin-releasing peptide receptor activation in a human colon cancer cell line: A proteomic approach. J Proteome Res 5: 1460– 1468.

Genome Organization in Disease Sarg B, Koutzamani E, Helliger W, Rundquist I, Lindner HH. 2002. Postsynthetic trimethylation of histone H4 at lysine 20 in mammalian tissues is associated with aging. J Biol Chem 277: 39195– 39201. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059– 1063. Schmidt CK, Brookes N, Uhlmann F. 2009. Conserved features of cohesin binding along fission yeast chromosomes. Genome Biol 10: R52. Scho¨nherr N, Binder G, Korsch E, Ka¨mmerer E, Wollmann HA, Eggermann T. 2008. Are H19 variants associated with Silver-Russell syndrome? J Pediatr Endocrinol Metab 21: 985 –993. Shen S, Liu A, Li J, Wolubah C, Casaccia-Bonnefil P. 2008. Epigenetic memory loss in aging oligodendrocytes in the corpus callosum. Neurobiol Aging 29: 452– 463. Shumaker DK, Dechat T, Kohlmaier A, Adam SA, Bozovsky MR, Erdos MR, Eriksson M, Goldman AE, Khuon S, Collins FS, et al. 2006. Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci 103: 8703– 8708. Sinclair DA, Guarente L. 1997. Extrachromosomal rDNA circles—a cause of aging in yeast. Cell 91: 1033– 1042. Soutoglou E, Dorn JF, Sengupta K, Jasin M, Nussenzweig A, Ried T, Danuser G, Misteli T. 2007. Positional stability of single double-strand breaks in mammalian cells. Nat Cell Biol 9: 675– 682. Sparago A, Cerrato F, Vernucci M, Ferrero GB, Silengo MC, Riccio A. 2004. Microdeletions in the human H19 DMR result in loss of IGF2 imprinting and BeckwithWiedemann syndrome. Nat Genet 36: 958– 960. Splinter E, Heath H, Kooren J, Palstra RJ, Klous P, Grosveld F, Galjart N, de Laat W. 2006. CTCF mediates long-range chromatin looping and local histone modification in the b-globin locus. Genes Dev 20: 2349– 2354. Therman E, Susman B, Denniston C. 1989. The nonrandom participation of human acrocentric chromosomes in Robertsonian translocations. Ann Hum Genet 53: 49– 65.

Tonkin ET, Wang TJ, Lisgo S, Bamshad MJ, Strachan T. 2004. NIPBL, encoding a homolog of fungal Scc2-type sister chromatid cohesion proteins and fly Nipped-B, is mutated in Cornelia de Lange syndrome. Nat Genet 36: 636 –641. Vega H, Waisfisz Q, Gordillo M, Sakai N, Yanagihara I, Yamada M, van Gosliga D, Kayserili H, Xu C, Ozono K, et al. 2005. Roberts syndrome is caused by mutations in ESCO2, a human homolog of yeast ECO1 that is essential for the establishment of sister chromatid cohesion. Nat Genet 37: 468– 470. Visser AE, Jaunin F, Fakan S, Aten JA. 2000. High resolution analysis of interphase chromosome domains. J Cell Sci 113: 2585– 2593. Wendt KS, Yoshida K, Itoh T, Bando M, Koch B, Schirghuber E, Tsutsumi S, Nagae G, Ishihara K, Mishiro T, et al. 2008. Cohesin mediates transcriptional insulation by CCCTC-binding factor. Nature 451: 796– 801. Wilson VL, Jones PA. 1983. DNA methylation decreases in aging but not in immortal cells. Science 220: 1055–1057. Witcher M, Emerson BM. 2009. Epigenetic silencing of the 16(INK4a) tumor suppressor is associated with loss of CTCF binding and a chromatin boundary. Mol Cell 34: 271 –284. Woodcock CL. 2006. Chromatin architecture. Curr Opin Struct Biol 16: 213 –220. Xie X, Lu J, Kulbokas EJ, Golub TR, Mootha V, Lindblad-Toh K, Lander ES, Kellis M. 2005. Systematic discovery of regulatory motifs in human promoters and 3’ UTRs by comparison of several mammals. Nature 434: 338–345. Yoon YS, Jeong S, Rong Q, Park KY, Chung JH, Pfeifer K. 2007. Analysis of the H19ICR insulator. Mol Cell Biol 27: 3499–3510. Zhao Z, Tavoosidana G, Sjolinder M, Gondor A, Mariano P, Wang S, Kanduri C, Lezcano M, Sandhu KS, et al. 2006. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat Genet 38: 1341–1347.

Nuclear Ataxias Harry T. Orr Institute of Translational Neuroscience, Department of Laboratory Medicine and Pathology, University of Minnesota, Minneapolis, Minnesota 55455 Correspondence: [email protected]

Historically basic neuroscience research has made several important contributions to the cell biology of the nucleus, in particular the elucidation of nuclear structures and compartments. As research progressed towards elucidating the mechanism of neurological disease at the cellular and molecular levels, it is now providing insight into the importance and basis of coordination of nuclear pathways within the nucleus and with other cellular compartments. Ataxias, lethal neurodegenerative diseases that are distinguished by a progressive loss of motor coordination, stem from disruption of nuclear function.

t seems fitting to take note that Ramon y Cajal, often regarded as the father of cellular neuroscience, also made major contributions to delineating nuclear compartments through his studies of the neuronal nucleus (Lafarga et al. 2009). The importance of this is work is highlighted by Cajal’s discovery of the nuclear accessory body now designated as the Cajal body. This nuclear structure is at center stage given its role in the biogenesis of snRNPs and snoRNPs (Gall 2003). Spinal muscular atrophy (SMA) is a genetic neurodegenerative disorder caused by a deficiency in survival of motor neuron 1 gene (SMN1). The SMN protein targets to Cajal bodies by a direct interaction with the Cajal body signature protein colin (Herbert et al. 2001). Ataxia, from Greek means literally a gross lack of coordination of movement, typically in

I

gait and limb. Although ataxia often results from degeneration in the cerebellum and associated connections, damage to any point along the neuroaxis controlling movement can lead to ataxia. This latter point provides the basis for the wide variety of genes that when mutated cause ataxia (Taroni and DiDonato 2004). There are three forms of ataxia for which evidence strongly supports a role of nuclear dysfunction in pathogenesis (Fig. 1). Among the dominantly inherited ataxias caused by expansions of glutamine tracts, are the spinocerebellar ataxia types 1 and 17. For each there is considerable evidence implicating misregulation of gene expression as being critical for disease. The third form of ataxia is the group of recessive ataxias caused by defects in DNA repair, most notably ataxia telangiectasia (AT).

Editors: David L. Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000786 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000786

H.T. Orr

SCA1

Transcription corepressorcoacitivators

Basal transcription complex

SCA17

RNA splicing complex

RNA polymerase

Single-strand break repair complex

AT AOA1 SCAN1

Figure 1. Overview of the Nuclear Ataxias. The different nuclear ataxias are positioned relative to the stage of

gene regulation thought to be impacted by each disease. SCA1—spinocerebellar ataxia type 1; SCA17—spimnocerebelllar ataxia type 17; AT—ataxia telangiectasia; AOA1—ataxia with ocular aprexia type 1; SCAN1 spinocerebellar ataxia and neuropathy 1.

COORDINATION IN THE NUCLEUS— SPINOCEREBELLAR ATAXIA TYPE 1 (SCA1)

SCA1 typically presents in middle age and progresses over 10 – 20 year to cause premature death. Juvenile as well as late-onset cases owing to larger or smaller CAG repeat expansions, respectively, are observed. Ataxia, tremor, and dysarthria (difficulty in articulating words) are common to the SCA1 clinical picture. Purkinje cell loss from the cerebellar cortex is a prominent pathological feature of SCA1. Loss of several brain stem neurons, including the inferior olive, is also a common feature. SCA1 is caused by expansion of a translated CAG repeat in a gene encoding the protein designated ataxin-1 (ATXN1) (Orr et al. 1993). Normal SCA1 alleles contain from six to 42 CAG repeats, encoding the amino acid glutamine, with those greater than 20 being interrupted with one to three CAT trinucleotides, encoding a histidine residue. Disease alleles, on the other hand, are pure CAG tracts ranging from 39 to 82 units. The length of the repeat tract is a major contributor to the age of disease onset.

The longer the repeat length on the mutant allele, the earlier is the age of onset. Individuals with 70 or more repeat units have a juvenile form of SCA1 whereas those containing mutant alleles with 40–50 repeats have an onset in the fourth or fifth decade of life. Mutant alleles also show germline instability such that in successive generations the repeat can expand further causing earlier onset of symptoms and increasing severity of disease in successive generations, a phenomenon known as anticipation. Genetic evidence consistently indicates that a gain-of-function mechanism of pathogenesis is critical for induction of SCA1. Although polyglutamine pathogenesis was initially thought to center on the polyglutamine aggregates and large inclusions (Ross and Poier 2004), more recently this has come under considerable question (Cummings et al. 1999; Klement et al. 1998; Saudou et al. 1998; Slow et al. 2005). Particularly in the case of SCA1, it is becoming increasingly apparent that the disorder is defined by the actions of the expanded polyglutamine tract in the context of ATXN1 (Gatchel and Zoghbi 2005, Orr 2001). Central to this idea is the

Nuclear Ataxias

concept that the normal function and interactions of ATXN1 are critical for defining the pathogenic pathway. It is further envisioned that expansion of the glutamine tract induces a change in protein conformation that in turn triggers alterations in its normal interactions with other cellular proteins. These altered interactions result in neuronal dysfunction, leading to neurodegeneration and neuronal loss. An important step in understanding SCA1 pathogenesis is the observation that in order for mutant ATXN1 to cause disease it had to enter the nucleus of Purkinje cells (Klement et al. 1998). Later studies revealed that wild-type ATXN1 has biochemical properties consistent with a role in the nucleus, and in some instances the normal function of ATXN1 are linked to pathogenesis. These normal activities of ATXN1 include the ability to bind RNA (Yue et al. 2001), to shuttle between the nucleus and cytoplasm (Irwin et al. 2005), and to interact with a variety of other nuclear proteins including several transcription factors (Lam et al. 2006; Okazawa et al. 2002; Serra et al. 2006; Tsai et al. 2004; Tsuda et al. 2005; Lim et al. 2008). Structurally, the carboxyl terminus of ATXN1 shares a 120-residue stretch with the HMG box transcription factor HBP1 (HMG box-containing protein-1) (Mushegian et al. 1997; de Chiara et al. 2003), designated as the AXH (ataxin-1/HBP1) domain. The genomes of Caenorhabditis elegans and Drosophila melanogaster encode small proteins consisting of essentially just an AXH domain, indicating that perhaps this region has the ability to fold independently (de Chiara et al. 2003). Analysis of the crystal structure of the AXH domain from ATXN1 showed it exists as a dimer in an OB (oligmer-binding)-fold (Chen et al. 2004). Proteins with an OB-fold are known to bind nucleic acids, particularly RNA. Thus, the AXH domain of ATXN1 is likely responsible for its RNA-binding property (Yue et al. 2001; de Chiara et al. 2003). It is worth noting that ATXN1 is SUMOylated on at least five lysine residues, two of which are in the AXH domain (Riley et al. 2005). SUMOylation of ATXN1 is inversely affected by the length of the polyglutamine tract and dependent on the ability of ATXN1 to be

localized to the nucleus. Most targets of SUMOylation are nuclear proteins, and many play a role in gene transcription (Seeler and Dejean 2003). In neurodegenerative diseases, understanding why specific sets of neurons are affected in spite of widespread expression of the disease protein remains one of the unresolved seminal issues. Interaction of ATXN1 with RORa, and the RORa coactivator Tip60 provides some insight into the enhanced susceptibility of Purkinje cells to SCA1. This work also challenges the concept that neurodegenerative diseases are unrelated to developmental abnormalities. Serra et al. (2006), using a conditional transgenic mouse model of SCA1 to delay the postnatal expression of mutant ATXN1 until after completion of cerebellar development, found that delayed postnatal expression of mutant ATXN1 led to a substantial reduction in disease severity in adults in comparison with early postnatal mutant ATXN1 expression. This result seems to be linked to a destabilization of RORa, a transcription factor critical for cerebellar development (Hamilton et al. 1996). In SCA1 mice, RORa was depleted and expression of genes controlled by RORa reduced. Furthermore, partial loss of RORa enhanced mutant ATXN1 pathogenicity. These studies suggest a basis for the Purkinje cell specificity of SCA1 because in the cerebellum these neurons are the only ones that express both ATXN1 and RORa. Another important step in dissecting ATXN1’s function in the nucleus was the realization that a major portion of wild type as well as mutant ATXN1 is located within large, soluble and relatively stable complexes (Lam et al. 2006). Moreover, substituting an Ala for the Ser at position 776 in ATXN1 decreases considerably the amount of ATXN1 in certain of these large complexes. The importance of this latter observation is linked to the previous demonstration that the S776 is a site where ATXN1 is phosphorylated and an A776 replacement abolishes Purkinje cell toxicity of mutant ATXN1 in vivo (Emamian et al. 2003). These data lead to the idea that mutant ATXN1 induced neuropathology stems from its assembly into native complexes with other nuclear

H.T. Orr

proteins and that this assembly might be regulated by the phosphorylation of ATXN1 at S776. More recently it was shown that glutamine expansion has opposing affects on different complexes (Lim et al. 2008). In complexes containing mutant ATXN1 and the RNA-binding motif protein 17, RBM17, glutamine expansion increased complex formation. Interestingly, SPF45 (the Drosophila homolog of RBM17) is a splicing factor that functions to regulate the second step of splicing by virtue of it having a role in defining the site of exon ligation (Lallena et al. 2002). In contrast to the situation with RBM17, the amount of mutant ATXN1 in complexes with the transcription repressor Capicua is decreased with glutamine expansion. Additional data suggest that the ATXN1Capicua complex is protective while its complex with RBM17 is neurotoxic. Thus, shifting the balance of cellular protein complexes contributes to neurodegeneration induced by mutant ATXN1. The data suggest a model with ATXN1 as a multifunctional regulatory protein in neurons where it acts to regulate and couple events along the multistep pathway of gene expression (Fig. 2). In the nucleus it is becoming increasingly apparent that transcription is coupled to the steps of RNA processing and transport including splicing, 30 -end formation and mRNA export from the nucleus (Wilkinson and Shyu 2001; Maniatis and Reed 2002; Reed 2003; Pandit et al. 2008). Functional coupling is facilitated by physical contacts between transcription and RNA processing molecular complexes such that the various steps in regulating are tethered to each other. Thus, ATXN1, which interacts with various regulators of transcription (e.g., Capicua and RORa/Tip60), RNA, as well as RNA-processing proteins (e.g., RBM17), is well positioned to function in the integration of transcription and RNA processing. Because the interaction of ATXN1 with RBM17 is modified by phosphorylation at S776 (Lim et al. 2008), it is reasonable to suggest that such a function under normal conditions would be dynamic and tightly regulated. Any disturbance in either the dynamics or regulation as a result of a mutation in ATXN1

would likely alter at several steps the expression of many critical genes. BASAL TRANSCRIPTION— SPINOCEREBELLAR ATAXIA TYPE 17

Perhaps no other neurodegenerative disorder exemplifies the paradox of selective neuropathology caused by mutations in a widely expressed protein more than does spinocerebellar ataxia type 17 (SCA17), which is caused by expansion of a glutamine tract located in the TATA-binding protein, TBP (Nakamura et al. 2001). TBP, also known as TFIID, is a wellstudied universal transcription factor essential for the function of all three nuclear RNA polymerases. Assembly of the transcription preinitiation complex, sufficient to drive a basal level of transcription, typically begins with TBP/TFIID binding to the TATA box found in the core promoter of many genes (Thomas and Chiang 2006). Yet, SCA17 is characterized by late onset of neurological symptoms that consists of progressive dementia, ataxia, and seizures. SCA17 pathology includes marked cerebellar atrophy and loss of Purkinje cells as well as less pronounced degeneration in other regions of the brain (Rolfs et al. 2003; Bruni et al. 2004). Another interesting question highlighted by TBP/TFIID is whether the polyglutamine tract has a normal function to regulate activity of the protein. It has been recognized for some time that polyglutamine or glutamine-rich stretches are present in many eukaryotic transcription factors, perhaps functioning as a transcription activation domain (Courey and Tjian 1988). The glutamine tract in wild-type alleles of human TBP/TFIID, like the glutamine stretch in other human polyglutamine proteins (Hardy and Orr 2006), is typically longer and much more polymorphic than in other species including nonhuman primates. This raises the interesting possibility that perhaps the glutamine tract of some proteins has evolved to assume an additional regulatory function. Evidence that this might be the case for TBP/TFIID was reported recently (Friedman et al. 2007). Previous work showed that dimerization of

Nuclear Ataxias

A

B Splicing RBM17 P ATXN1 P CIC

RORa / Tip60

Transcription

Figure 2. (A) ATXN1, a nuclear protein. A confocal image of Purkinje cells from a SCA1 transgenic mouse

expressing an expanded allele of the SCA1 gene. The ATXN1 protein (yellow) is within Purkinje cell nuclei. (B) A model depicting ataxin1 nuclear function. In this model coupling of transcription and RNA splicing is mediated by the interactions of ATXN1 with Capicua (CIC), RORa/Tip60, and RBM17. P Denotes ATXN1 interactions regulated by phosphorylation at Ser776.

TBP/TFIID functions to prevent the up-regulation of gene expression (Jackson-Fisher et al. 1999). Friedman and colleagues went on to show that the formation of inactive dimmers of TBP/TFIID, driven by the carboxy-terminal dimerization domain, was inversely related to the length of the glutamine tract at the amino terminus. These investigators further show that

the length of the glutamine stretch in TBP/ TFIID affects the extent to which it interacts with the general transcription factor TFIIB, with expanded glutamines enhancing interaction. They also examined the occupancy of TATA-containing promoters in the cerebellum of SCA17 transgenic mice expressing TBP with an expanded glutamine tract and found

H.T. Orr

selectively decreased occupancy at some but not all TATA-containing promoters. Notably, they found a decrease in the amount of both TBP/ TFIID and TFIIB at the promoter of the gene encoding the small heat shock protein HSPB1. Hspb1 expression is down-regulated in affected SCA17 mice and its overexpression reduces the toxic effects of mutant TBP. The recent demonstration that the polyglutamine region from huntingtin adopts multiple conformations (Kim et al. 2009), suggests that this region imparts considerable conformational flexibility to the native protein that likely has important functional consequences.

DNA REPAIR—ATAXIA TELANGIECTASIA (AT), ATAXIA WITH OCULAR APREXIA TYPE 1 (AOA1), AND SPINOCEREBELLAR ATAXIA AND NEUROPATHY (SCAN1)

In dividing cells, the link between defective DNA repair and unregulated cell division, i.e., cancer, dates back to studies on xeroderma pigmentosum published in 1968 by James E. Cleaver (Cleaver 1968). Postmitotic neurons, in contrast, respond to deficits in DNA repair with the induction of neuronal cell death that often presents clinically with ataxia (Rass et al. 2007). Among the ataxias associated with defects in DNA repair (Paulson and Miller 2005), recent studies on ataxia telangiectasia (AT), ataxia with ocular aprexia Type 1 (AOA1), and spinocerebellar ataxia and neuropathy-1 (SCAN1) suggest specific repair pathways and how their disruption may lead to neurodegeneration (Table 1).

AT is an autosomal recessive childhood disease characterized by progressive ataxia with cerebellar atrophy and loss of Purkinje cells as well as an increased instance of lymphoid cancer (Lavin 2008). The gene mutated in AT (Savitsky et al. 1995), AT mutated (ATM) encodes a Ser/ Thr kinase that associates with the Mre11/ Rad50/Nbs1 (MRN) complex that acts as a sensor of double strand breaks (DSBs) in DNA. ATM is critical for proper activation of the broad cellular response to DSBs. The recognition of DSBs by the MRN complex does not require ATM (Mirzoeva and Petrini 2003). Yet it is involved in the activation of ATM and as an adaptor for ATM-mediated phosphorylation of some of its substrates. Although numerous phosphorylation substrates for ATM are reported, perhaps the most notable substrate is p53 given its well characterized role in genome instability (Banin et al. 1998; Canman et al. 1998; Khanna et al. 1998). Somewhat paradoxically, although less than 20% of AT patients are reported to develop cancer, AT is a paradigm for linking DNA damage, cell signaling and cancer (Lavin 2008). On the other hand, efforts to understand the loss of Purkinje cells from the cerebellum, a hallmark of ATsuch that patients typically are wheelchairbound early in life, have proven to be much more intractable. One impediment to understanding the neurodegeneration seen in AT is the lack of a similar phenotype in Atm knockout mice. Although mice homozygous for a null allele of Atm show many signs of AT including extreme sensitivity to g-irradiation and lymphomas, the neurological deficits are relatively mild

Table 1. Ataxias associated with defects in DNA repair DNA damage repaired

Neurological SYMPTOMS

Disease

Gene

Ataxia telangiectasis (AT)

ATM

DSBs

Ataxia

Ataxia with oculomotor apraxia one1 (AOA1) Spinocerebellar ataxia with axonal neuropathy (SCAN1)

APTX

SSBs

TDP1

SSBs

Ataxia, OA, neuropathy Ataxia, neuropathy

Nonneurological symptoms

Predisposition to cancer, immunodeficiency, telangiectasia None None

Nuclear Ataxias

and, importantly, are not associated with cerebellar pathology (Barlow et al. 1996; Xu et al. 1996). Histological examination of Atm 2/2 mice show normal Purkinje cells and no evidence of any other brain abnormalities. These data suggest there is a missing link in the mouse. A physiological function of the MRN complex in the brain, with which ATM interacts in response to DNA damage, was demonstrated using a conditional knockout approach. The mouse homologue of human NBS1, a component of the MRN complex and the gene mutated in the AT-related disorder Nijmegan breakage syndrome (D’Amours and Jackson 2002), was selectively deleted in the CNS (Frappart et al. 2005). Loss of Nbn results in an ataxia similar to that in AT and the induction of apoptosis of postmitotic neurons of the cerebellum, including Purkinje cells. These results argue that the ATM-associated MRN complex has a role in maintaining neuronal integrity and suggest that loss of Nbn in CNS of the mouse may provide a model for the neurodegeneration seen in AT. Unlike AT, AOA1 has no symptoms outside of the nervous system, i.e. cancer. The autosomal recessive disorder AOA1 consists of early onset cerebellar ataxia. In addition AOA1 is characterized by sensorimotor neuropathy. The mutation causing AOA1 is in the gene encoding Aprataxin (Date et al. 2001). Aprataxin is a 342 amino acid nuclear/nucleolar protein that has three domains, an amino-terminal forkhead-associated domain, a central histidine triad (HIT) domain, and a carboxy-terminal zinc-finger domain. Aprataxin interacts with XRCC1 and XRCC4, subunits of the DNA ligase complexes involved in single-strand break (SSB) and DSB repair, respectively. These interactions plus the presence of a DNA-binding zinc-finger domain in Aprataxin hinted at an involvement in DNA repair. Importantly, most of the AOA1 mutations are found within or near the HIT domain of Aprataxin (Rass et al. 2007). Typically HIT domains are associated with nucleotide hydrolase and transferase activities (Brenner 2002). However, the nucleotide hydrolase activity of Aprataxin is two orders of magnitude lower than that seen for other hydrolases. More recent biochemical studies show that Aprataxin is a

DNA-specific adenylase with this activity being several orders of magnitude greater than its hydrolase activity (Ahel et al. 2006), indicating that it has the ability to scan the genome for adenylated DNA lesions. Like AOA1, SCAN1 symptoms are largely restricted to the nervous system. SCAN1 patients are affected with cerebellar ataxia and axonal sensorimotor neuropathy. In addition, they have mild hypercholesterolemia and hypoalbuminemia. The gene affected in SCAN1, TDP1, encodes tyrosyl-DNA-hosphopdiesterase 1 (Takashima et al. 2002). TDP1 is known to repaircovalent topoisomerase I-DNA adducts created by aborted topoisomerase I reactions. Removal of these adducts by TDP1 generates SSBs that are subsequently repaired by polynucleotidekinase300 phosphatase(PNKP). Besides interacting with PNKP, TDP1 was recently reported to be in a complex with XRCC1 and DNA-ligase III that repair SSBs (El-Khamisy et al. 2005). Accordingly, SCAN1 cells are unable to repairs SSBs induced by the drug camptothecin. Thus, TDP1 likely functions as a critical component of a complex involved in the repair of SSBs. AT, AOA1, and SCAN1 provide compelling evidence that neurons are especially sensitive to the presence of DNA lesions, in particular SSBs. Clearly, this sensitivity resides in some aspect or combination of features unique to neurons—with cerebellar Purkinje cells being particularly distinctive. The first point that typically comes to mine is that neurons are terminally differentiated postmitotic cells. Not being cycling cells neurons would not have the ability to repair lesions using replication-based mechanisms. Over the years, as one argument goes, unrepaired lesions accumulate in absence of ATM, Aprataxin, or TDP1. This accumulation might be enhanced because of the high metabolic rate of neurons and an exposure to oxidative DNA damage. Accumulated damage would in turn impede transcription eventually hitting functions critical for neuronal integrity. Although such factors likely contribute to toxicity, this scenario differs little from that proposed linking DNA repair defects to cancer where eventually the regulation of oncogene

H.T. Orr

and/or tumor suppressor gene expression is circumvented. Except in cycling cells the result is uncontrolled growth and in neurons it is death. Maybe the response of neurons to defects in DNA repair does reside in their being postmitotic and like cycling cells the eventual target of DNA lesions is regulation of the cell cycle. However, as highlighted recently by Herrup and Yang (2007), the fact that once a neuron leaves the region of neurogenesis it will never divide again could place strong pressure to constantly keep its cell cycle in check. Experimental evidence shows that if a neuron is induced to re-enter the cell cycle it dies. In mice null for the retinoblastoma protein, Rb, postmitotic neurons undergo cell death that is coupled with DNA synthesis and dependent on transcription factors E2F1 and p53 (Clarke et al. 1992; Jacks et al. 1992; Lee et al. 1992; Macleod et al. 1996; Tsai et al. 1998). Likewise, inactivation of Rb in neurons using a SV40 T antigen transgene induces rapid cell death associated with DNA synthesis and E2F1 (al-Ubaidi et al. 1992; Feddersen et al. 1992; 1995; 1997; Athanasiou et al. 1998). It is intriguing that AT, AOA1, and SCA1 are neurodegenerative diseases of childhood with a relatively rapid progression. These features of AT, AOA1, and SCAN1 are quite distinct from the late age of onset and slower progression typical of the dominant polyglutamine-based ataxias (Orr and Zoghbi 2007). Possibly these features are related mechanistically to the rapid course of cell death as seen in the mice with inactivation Rb in their Purkinje cells, i.e., reflecting misregulation of the cell cycle in a postmitotic neuron. CONCLUDING REMARKS

An unrelenting question concerning the nuclear ataxias (as well as for many other forms of ataxia and neurodegenerative disease) is why is there a very specific set of symptoms with a mutation in a gene that encodes a protein that is widely expressed in the brain and has what is considered to be a housekeeping function. In the case of SCA1, there is increasing evidence that mutant ATXN1 affects the expression and function of proteins more selectively expressed by

Purkinje cells, e.g., RORa, thereby altering pathways uniquely critical for their function (Serra et al. 2006). What about the other nuclear ataxias? Why does a polyglutamine expansion in the TATA binding protein, which has a role in regulating basal transcription of many genes in many cell types, cause late onset SCA17 that is highlighted by a prominent loss of cerebellar Purkinje cells? How does one explain a disruption in DNA repair leading to the loss of specific nondividing neurons, i.e., cerebellar Purkinje cells, in AT, AOA1, and SCAN1? Perhaps the answer lies in neuronal chromatin. TBP, mutated in SCA17, as well as components of the DNA repair machinery, affected in AT, AOA1, and SCAN1, all function upon interacting with DNA. Chromatin structure is emerging as an essential aspect in regulating access of proteins to DNA. Concurrent with their mitotic exit, neurons have a switch in the subunit composition in their chromatinremodeling complexes. This switch in subunit composition is critical for the proper regulation of dendritic development (Wu et al. 2007). Moreover, a distinct feature of cerebellar Purkinje cells is their large and euchromatic nuclei. Other neurons, such as cerebellar granule cells, manifest a more typical distribution of more densely packed heterochromatin. Epigenetic modification, e.g., DNA methylation, is another means by which chromatin structure is regulated. In light of the recent identification of a methylated nucleotide, 5hydroxymethylcytosine, unique to the brain and enriched in Purkinje cells this kind of regulation could provide another avenue for marking chromatin in these neurons (Kriaucionis and Heintz 2009). Thus, there are aspects of chromatin structure and remodeling unique to neurons as well as to specific populations of neurons that might very well affect the access pattern of DNA-binding proteins that could underlie the cellular specificity of pathology in SCA17, AT, AOA1, and SCAN1. Regardless, as the pathogenesis of these nuclear ataxias is elucidated a better understanding of how critical nuclear functions are regulated will be forthcoming.

Nuclear Ataxias

REFERENCES Ahel I, Rass U, El-Khamisy SF, Katyal S, Clements PM, McKinnon PJ, Caldecott KW, West SC. 2006. The neurodegenerative diseases protein aprataxin resolves abortive DNA ligation intermediates. Nature 443: 713 –716. Al-Ubaidi MR, Hollyfield JG, Overbeek PA, Baehr W. 1992. Photoreceptor degeneration induced by the expression of simian virus 40 large tumor antigen in the retina of transgenic mice. Proc Natl Acad Sci 89: 1194– 1198. Athanasiou MC, Yunis W, Coleman N, Ehlenfeldt R, Clark HB, Orr HT, Feddersen RM. 1998. The transcription factor E2F-1 in SV40 T antigen-induced cerebellar Purkinje cell degeneration. Mol Cell Neurosci 12: 16–28. Banin S, Moyal L, Shieh S, Taya Y, Anderson CW, Chessa L, Smorodinsky NI, Prives C, Reiss Y, Shiloh Y, et al. 1998. Enhanced phosphorylation of p53 by ATM in response to DNA damage. Science 281: 1674–1677. Barlow C, Hirotsune S, Paylor R, Liyanage M, Eckhaus M, Collins F, Shiloh Y, Crawley JN, Ried T, Tagle D, et al. 1996. Atm-deficient mice: a paradigm of ataxia telangiectasia. Cell 86: 159 –171. Brenner C. 2002. Hint, Fhit, and GalT: function, structure, evolution and mechanism of three branches of the histidine triad superfamily of nucleotide hydrolases and transferases. Biochem 41: 9003– 9014. Bruni AC, Takahashi-Fujigasaki J, Maltecca F, Foncin JF, Servadio A, Casari G, D’Adamo P, Maletta R, Curcio SA, De Michele G, et al. 2004. Behavioral disorder, dementia, ataxia, and rigidity in a large family with TATA-box binding protein mutation. Arch Neurol 61: 1314– 1320. Canman CE, Lim DS, Cimprich KA, Taya Y, Tamai K, Sakaguchi K, Appella E, Kastan MB, Siliciano JD. 1998. Activation of the ATM kinase by ionizing radiation and phosphorylation of p53. Science 281: 1677– 1679. Chen YW, Allen MD, Veprintsev DB, Lsˇwe J, Bycroft M. 2004. The structure of the AXH domain of spinocerebellar ataxin-1. J Biol Chem 278: 3758–3765. Clarke AR, Maandag ER, van Roon M, van der Lugt NM, van der Valk M, Hooper ML, Berns A, te Riele H. 1992. Requirement for a functional Rb-1 gene in murine development. Nature 359: 328–330. Cleaver JE. 1968. Defective DNA repair replication in xeroderma pigmentosum. Nature 218: 652–656. Courey AJ, Tjian R. 1988. Analysis of Sp1 in vivo reveals multiple transcriptional domains, including a novel glutamine-rich activation motif. Cell 55: 887– 898. Cummings CJ, Reinstein E, Sun Y, Antalffy B, Jiang Y-H, Ciechanover A, Orr HT, Beaudet AL, Zoghbi HY. 1999. Mutation of the E6-AP ubiquitin ligase reduces nuclear inclusion frequency while accelerating polyglutamineinduced pathology in SCA1 transgenic mice. Neuron 24: 879 –892. D’Amours D, Jackson SP. 2002. The Mrel 1 complex: at the crossroads of DNA repair and checkpoint signaling. Nat Rev Mol Cell Biol 3: 317– 327. Date H, Onodera O, Tanaka H, Iwabuchi K, Uekawa K, Igarashi S, Koike R, Hiroi T, Yuasa T, Awaya Y, et al. 2001. Early-onset ataxia with ocular motor apraxia and hypoalbuminemia is caused by mutations in a new HIT superfamily gene. Nat Genet 29: 184– 188.

de Chiara C, Giannini C, Adinolfi S, de Boer J, Guida S, Ramos A, et al. 2003. The AXH module: an independently folded domain common to ataxin-1 and HBP1. FEBS Letters 551: 107 –112. El-Khamisy SF, Saifi GM, Weinfeld M, Johansson F, Helleday T, Lupski JR, Caldecott KW. 2005. Defective DNA singlestrand break repair in spinocerebellar ataxia with axonal neuropathy-1. Nature 434: 108–113. Emamian ES, Kaytor MD, Duvick LA, Zu T, Tousey SK, Zoghbi HY, Clark HB, Orr HT. 2003. Serine 776 of ataxin-1 is critical for polyglutamine-induced disease in SCA1 transgenic mice. Neuron 38: 375– 387. Feddersen RM, Ehlenfeldt R, Yunis WS, Clark HB, Orr HT. 1992. Disrupted cerebellar development and progressive degeneration of Purkinje cells in SV40 T antigen transgenic mice. Neuron 9: 955– 966. Feddersen RM, Clark HB, Yunis WS, Orr HT. 1995. In vivo viability of postmitotic Purkinje neurons requires pRb family member function. Mol Cell Neurosci 6: 153– 167. Feddersen RM, Yunis WS, O’Donnell MA, Ebner TJ, Shen L, Iadecola C, Orr HT, Clark HB. 1997. Susceptibility to cell death induced by mutant SV40 T-antigen correlates with Purkinje neuron functional development. Mol Cell Neurosci 9: 42– 62. Frappart P-O, Tong W-M, Demuth I, Radovanovic I, Herceg Z, Aguzzi A, Digweed M, Wang Z-Q. 2005. An essential function for NBS1 in the prevention of ataxia and cerebellar defects. Nat Med 11: 538–544. Friedman MJ, Shah AG, Fang Z-H, Ward EG, Warren ST, Li S, Li X-J. 2007. Polyglutamine domain modulates TBPTFIIB interaction: Implications for its normal function and neurodegeneration. Nat Neurosci 10: 1519–1528. Gall JG. 2003. The centennial of the Cajal body. Nat Rev Mol Cell Bio 4: 975– 980. Gatchel JR, Zoghbi HY. 2005. Diseases of unstable repeat expansion: Mechanisms and common principals. Nat Rev Genet 6: 743– 755. Hamilton BA, Frankel WN, Kerrebrock AW, Hawkins TL, Fitz-Hugh W, Kusumi K, Russell LB, Mueller KL, van Berkel V, Birren B, et al. 1996. Disruption of the nuclear hormone receptor RORa in staggerer mice. Nature 379: 736 –739. Hardy J, Orr HT. 2006 The genetics of neurodegenerative diseases. J Neurochem 97: 1690–1699. Herbert MD, Szymczyk PW, Shpargel KB, Matera AG. 2001. Colin forms the bridge between Cajal bodies and SMN, the spinal muscular atrophy protein. Genes Dev 15: 2720–2729. Herrup K, Yang Y. 2007. Cell cycle regulation in the postmitotic neuron: Oxymoron or new biology? Nat Rev Neurosci 8: 368– 378. Irwin S, Vandelft M, Howell JL, Graczyk J, Orr HT, Truant R. 2005. RNA association and nucleocytoplasmic shuttling by ataxin-1. J Cell Sci 118: 233– 242. Jacks T, Fazeli A, Schmitt EM, Bronson RT, Goodell MA, Weinberg RA. 1992. Effects of an Rb mutation in the mouse. Nature 359: 295 –300. Jackson-Fisher AJ, Chitikila C, Mitra M, Pugh BF. 1999. A role for TBP dimerization in preventing unregulated gene expression. Mol Cell 3: 717– 727.

H.T. Orr Khanna KK, Keating KE, Kozlov S, Scott S, Gatei M, Hobson K, Taya Y, Gabrielli B, Chan D, Lees-Miller SP, et al. 1998. ATM associates with and phosphorylates p53: mapping the region of interaction. Nat Genet 20: 398– 400. Kim MW, Yogarany C, Kim SW, Otwinowski Z, Bezprozvanny I. 2009. Secondary structure of Huntingtin aminoterminal region. Structure, 17: 1205– 1212. Klement IA, Skinner PJ, Kaytor MD, Yi H, Hersch SM, Clark HB, Zoghbi HY, Orr HT. 1998. Ataxin-1 nuclear localization and aggregation: Role in polyglutamineinduced disease in SCA1 transgenic mice. Cell 95: 41–53. Kriaucionis S, Heintz N. 2009. The nuclear DNA base 5-hydroxymethylcytosine is present in Purkinje neurons and the brain. Science 324: 929 –930. Lafarga M, Casafont I, Bengoechea R, Tapia O, Berciano MT. 2009. Cajal’s contribution to the knowledge of the neuronal cell nucleus. Cromosoma 118: 437 –443. Lallena MJ, Chalmers KJ, Llamazares S, Lamond AI, Valc‡rcel J. 2002. Splicing regulation at the second catalytic step by sex-lethal involves 30 splice site recognition by SPF45. Cell 109: 285 –296. Lam YC, Bowman AB, Jafar-Nejad P, Lim J, Richman R, Fryer JD, Hyun E, Duvick LA, Orr HT, Botas J, et al. 2006. Mutant ATAXIN-1 interacts with the repressor Capicua in its native complex to cause SCA1 neuropathology. Cell 127: 1335–1347. Lavin MF. 2008. Ataxia-telangiectasia; from a rare disorder to a paradigm for cell signaling and cancer. Nat Rev Mol Cell Bio 9: 759– 769. Lee EY, Chang CY, Hu N, Wang YC, Lai CC, Herrup K, Lee WH, Bradley A. 1992. Mice deficient for Rb are nonviable and show defects in neurogenesis and haematopoiesis. Nature 329: 288–294. Lim J, Crespo-Barreto J, Jafar-Nejad P, Bowman AB, Richman R, Hill DE, Orr HT, Zoghbi HY. 2008. Opposing effects of polyglutamine expansion on native protein complexes contribute to SCA1. Nature 452: 713–719. Macleod KF, Hu Y, Jacks T. 1996. Loss of Rb activates both p53-dependen and independent cell death pathways in the developing mouse nervous system. EMBO J 15: 6178– 6188. Maniatis T, Reed R. 2002. An extensive network of coupling among gene expression machines. Nature 416: 499 –506. Matera AG, Frey MR. 1998. Nuclear structure ’98. Coiled bodies and gems: Janus or Gemini? Am J Hum Genet 63: 317 –321. Mirzoeva OK, Petrini JH. 2003. DNA replication-dependent nuclear dynamics of the Mre 11 complex. Mol Can Res 1: 207–218. Mushegian AR, Bassett DEJr, Boguski MS, Bork P, Koonin EV. 1997. Positionally cloned human disease genes: patterns of evolutionary conservation and functional motifs. Proc Nat Acad Sci 94: 5831– 5836. Nakamura K, Jeong SY, Uchihara T, Anno M, Nagashima K, Nagashima T, Ikeda S, Tsuji S, Kanazawa I. 2001. SCA17, a novel autosomal dominant cerebellar ataxia caused by an expanded polyglutamine in TATA-binding protein. Hum Mol Genet 10: 1441–1448.

Okazawa H, Rich T, Chang A, Lin X, waragai M, Kajikawa M, Enokido Y, Komuro A, Kato S, Shibata M, et al. 2002. Interaction between mutant ataxin-1 and PQBP-1 affects transcription and cell death. Neuron 34: 701 –713. Orr HT, Chung MY, Banfi S, Kwiatkowski TJJr, Servadio A, Beaudet AL, McCall AE, Duvick LA, Ranum LPW, Zoghbi HY. 1993. Expansion of an unstable trinucleotide CAG repeat in spinocerebellar ataxia type 1. Nat Genet 4: 221 –226. Orr HT. 2001. Beyond the Qs in the polyglutamine diseases? Genes Dev 15: 925– 932. Orr HT, Zoghbi HY. 2007. Trinucleotide repeat disorders. Ann Rev Neurosci 30: 575 –621. Pandit S, Wang D, Fu X-D. 2008. Functional integration of transcriptional and RNA processing machines. Curr Opin Cell Bio 20: 260– 265. Paulson HL, Miller VM. 2005. Breaks in coordination: DNA repair in inherited ataxia. Neuron 46: 845 –848. Rass U, Ahel I, West SC. 2007. Defective DNA repair and neurodegenerative disease. Cell 130: 991– 1004. Reed R. 2003. Coupling transcription, splicing and mRNA export. Curr Opin Cell Bio 15: 326–331. Riley BE, Zoghbi HY, Orr HT. 2005. SUMOylation of the polyglutamine protein, ataxin-1 is dependent on a functional nuclear localization signal. J Biol Chem 280: 21942– 21948. Rolfs A, Koeppen AH, Bauer I, Bauer P, Buhlmann S, Topka H, Schsˇls L, Riess O. 2003. Clinical features and neuropathology of autosomal dominant spioncerebellar ataxia (SCA17). Ann Neurol 54: 367–375. Ross CA, Poier MA. 2004. Protein aggregation and neurodegenerative disease. Nat Med S10 Saudou F, Finkbeiner S, Devys D, Greenberg ME. 1998. Huntingtin acts in the nucleus to induce apoptosis but death does not correlate with the formation of intranuclear inclusions. Cell 95: 55–66. Savitsky K, Bar-Shira A, Gilad S, Rotman G, Ziv Y, Vanagaite L, Tagle DA, Smith S, Uziel T, Sfez S, et al. 1995. A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science 268: 1749–1753. Seeler J-S, Dejean A. 2003. Nuclear and unclear functions of SUMO. Nat Rev Cell Bio 4: 690 –699. Serra HG, Duvick L, Zu T, Carlson K, Stevens S, Jorgensen N, Lysholm A, Burright E, Zoghbi HY, Clark HB, et al. 2006. RORa-mediated Purkinje cell development determines disease severity in adult SCA1 mice. Cell 127: 697 –708. Slow EJ, Graham RK, Osmand AP, Devon RS, Lu G, Deng Y, Pearson J, Vaid K, Bissada N, Wetzel R, et al. 2005. Absence of behavioral abnormalities and neurodegeneration in vivo despite widespread neuronal huntingtin inclusions. Proc Natl Acad Sci USA 102: 11402– 11407. Taroni F, DiDonato S. 2004. Pathways to motor incoordination: the inherited ataxias. Nat Rev Neurosci 5: 641 –656. Thomas MC, Chiang CM. 2006. The general transcription machinery and general cofactors. Crit Rev Biochem Mol Biol 41: 105 –178.

Nuclear Ataxias Tsai KY, Hu Y, Macleod KF, Crowley D, Yamasaki L, Jacks T. 1998. Mutation of E2f-1 suppresses apoptosis and inappropriate S phase entry and extends survival of Rb-deficient mouse embryos. Mol Cell 2: 293 –304. Tsai CC, Kao HY, Mitzutani A, Banayo E, Rajan H, McKeown M, Evans RM. 2004. Ataxin 1, a SCA1 neurodegenerative disorder protein, is functionally linked to the silencing mediator of retinoid and thyroid hormone receptors. Proc Natl Acad Sci USA 101: 4047– 4052. Tsuda H, Jafar-Nejad H, Patel AJ, Sun Y, Chen H-K, Rose MF, Venken KJT, Botas J, Orr HT, Bellen HJ, et al. 2005. The AXH domain in mammalian/Drosophila Ataxin-1 mediates neurodegeneration in spinocerebellar ataxia 1 through its interaction with Gfi-1/Senseless proteins. Cell 122: 633– 644.

Wilkinson MF, Shyu AB. 2001. Multifunctional regulatory proteins that control gene expression in both the nucleus and the cytoplasm. Bioessays 23: 775–787. Wu JI, Lessard J, Olave IA, Qiu Z, Ghosh A, Graef IA, Crabtree GR. 2007. Regulation of dendritic development by neuron-specific chromatin remodeling complexes. Neuron 56: 94– 108. Xu Y, Ashley T, Brainerd EE, Bronson RT, Meyn MS, Baltimore D. 1996. Targeted disruption of ATM leads to growth retardation, chromosomal fragmentation during meiosis, immune defects, and thymic lymphoma. Genes Dev 10: 2411–2422. Yue S, Serra H, Zoghbi HY, Orr HT. 2001. The SCA1 protein, ataxin-1, has RNA-binding activity that is inversely affected by the length of its polyglutamine tract. Hum Mol Genet 10: 25– 30.

Developments in RNA Splicing and Disease Michael G. Poulos, Ranjan Batra, Konstantinos Charizanis, and Maurice S. Swanson Department of Molecular Genetics and Microbiology and the Genetics Institute, University of Florida, College of Medicine, Gainesville, Florida 32611 Correspondence: [email protected]

Pre-mRNA processing, including 50 -end capping, splicing, editing, and polyadenylation, consists of a series of orchestrated and primarily cotranscriptional steps that ensure both the high fidelity and extreme diversity characteristic of eukaryotic gene expression. Alternative splicing and editing allow relatively small genomes to encode vast proteomic arrays while alternative 30 -end formation enables variations in mRNA localization, translation, and stability. Of course, this mechanistic complexity comes at a high price. Mutations in the myriad of RNA sequence elements that regulate mRNA biogenesis, as well as the trans-acting factors that act upon these sequences, underlie a number of human diseases. In this review, we focus on one of these key RNA processing steps, splicing, to highlight recent studies that describe both conventional and novel pathogenic mechanisms that underlie muscle and neurological diseases.

eep sequencing and microarray analyses of the human transcriptome have revealed that .90% of our genes undergo alternative splicing, which permits a limited genome to encode the vast proteomic repertoire required for the human interactome (Blencowe 2006; Wang and Burge 2008; Wang et al. 2008). From a biochemical viewpoint, the RNA splicing reaction is a relatively simple process that consists of two transesterification reactions (Black and Grabowski 2003; Konarska and Query 2005; House and Lynch 2008). However, this apparent simplicity is compounded by the fact that exons are generally buried within much larger introns and both authentic, and many more false, splice sites exist in genes. In addition, multiple alternative splicing modes are utilized in vivo (Fig. 1). Finding the majority

D

of these sites is the function of the major spliceosome, which is composed of five small nuclear ribonucleoprotein complexes (snRNPs) and 170 proteins (Deckert et al. 2006; Behzadnia et al. 2007; Bessonov et al. 2008; Wahl et al. 2009). Conserved sequence elements at exon-intron boundaries, the 50 and 30 splice sites of each exon, and the branchpoint facilitate spliceosome recruitment via base-pairing interactions with five spliceosome-associated snRNAs (U1, U2, U4/6, U5) (Fig. 2). A minor spliceosome, which recognizes variant splice sites and contains a unique set of snRNAs (U11, U12, U4atac, U6atac) in addition to U5, also exists in the nucleus (Will and Luhrmann 2005; Steitz et al. 2008). Spliceosomal recognition of these core elements is modulated by a myriad of additional sequence elements in

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved. Advanced Online Article. Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a000778

M.G. Poulos et al.

Alternative promoters

Retained intron

Cassette exon

Mutually exclusive cassettes

Alternative 5′ splice site

Alternative 3′ splice site

Alternative 3′ exons

U2 augaug ISE ISS

ynyu r y a

U2AF65 (y)n

hnRNP

SR

(A)n

BP

U1 u c cauuca

35 nag

(A)n

AG gu r agu

G ESE ESS

ISE ISS

Figure 1. RNA sequences and factors involved in the regulation of alternative splicing. The upper diagram shows

various patterns of alternative splicing using a 12-exon (black and colored boxes) gene that contains alternative promoters (arrows adjacent to turquoise boxes), constitutive exons (black), a retained intron (gray), a cassette and mutual exclusion exons (blue), alternative 50 and 30 splice sites (orange), and alternative 30 exons with polyadenylation sites (A)n. The lower diagram highlights the conserved RNA sequence elements at the branch point (BP, U2 binding site) and the 50 (U1 binding site) and 30 (U2AF heterodimer binding site) splice sites of an alternatively spliced cassette exon (blue box). Also shown are the less conserved exonic and intronic splicing enhancer (ESE, ISE) and silencer (ESS, ISS) elements that are recognized by multiple factors including the SR and hnRNP proteins.

both exons and introns that either activate (exonic splicing enhancer, ESE; intronic splicing enhancer, ISE) or repress (exonic splicing silencer, ESS; intronic splicing silencer, ISS) spliceosome recruitment by interacting with a variety of splicing factors, including serinearginine-rich (SR) proteins and heterogeneous nuclear ribonucleoproteins (hnRNPs) (Fig. 1) (Martinez-Contreras et al. 2007; Long and Caceres 2009). The density of these sites is astonishing, and current estimates suggest that splicing regulatory motifs comprise .75% of the nucleotides in an average exon (Chasin 2007). Both the importance and complexity of splicing regulation is typified by the expression of dystrophin, a 427 kDa protein encoded by the DMD gene composed of 79 exons and 8 promoters, in skeletal muscle (Ervasti 2007; Gurvich et al. 2008). Dystrophin plays an essential role in skeletal muscle cells, or myofibers, since it binds to both actin and b-dystroglycan and

thus provides a vital link between the actinmyosin contractile apparatus and the extracellular matrix (ECM). A number of single nucleotide mutations result in skeletal muscle degenerative diseases, such as the severe Duchenne (DMD) and the milder Becker (BMD) muscular dystrophies, as well as X-linked dilated cardiomyopathy. For Duchenne, mutations often disrupt the DMD open reading frame, sometimes by altering splicing enhancers and silencers, while the reading frame, and some dystrophin function, is mostly preserved in BMD. Although every tissue requires flexibility in gene expression, the significance of alternative splicing to human function is particularly striking in the central nervous system. For example, CACNA1H, which encodes a T-type low-voltage-activated calcium channel and is a candidate gene for idiopathic generalized epilepsies, is extensively alternatively spliced to potentially generate .4,000 mRNA variants

RNA Splicing Disease AG gu

a

ag G

Pre-mRNA U1

Intron

mRNA SNRNP200/hBrr2

U1

U2 U6

PRPF3/hPrp3 PRPF31/hPrp31 U1 U2

U6

Pre-spliceosome

U4

Post-spliceosomal complex

U4/U6.U5 tri-snRNP

U5

U2 U5

Step 2 PRPF8/hPrp8 SNRNP200/hBrr2

Pre-catalytic spliceosome

U1 U6 U4 U5 U2

U6

U2

U5

U1 U4

Step 1 U6 U2 U5

SNRNP200/hBrr2

Activated spliceosome

Figure 2. Spliceosome dynamics and retinopathy. The assembly/disassembly cycle of the spliceosome (adapted with permission from Wahl et al. 2009) during splicing with only the pre-mRNA (blue and yellow boxes, exons; thick black line, intron) and snRNP (grey) complexes illustrated to highlight the roles of U4-associated (PRPF3/ hPrp3, PRPF31/hPrp31) and U5-associated (PRPF8/hPrp8, SNRP200/hBrr2) proteins as well as several other steps that require the DExD/H-type RNA-dependent ATPase/helicase. Steps 1 and 2 refer to the first and second catalytic steps of splicing.

(Zhong et al. 2006). Therefore, it is not surprising that errors in splicing regulation have been linked to a number muscle and neurologic diseases and this number is likely to increase in the future. Since a number of excellent reviews have been published recently that survey how defects in splicing are associated with a broad range of human diseases (Licatalosi and Darnell 2006; Cooper et al. 2009; Tazi et al. 2009), this review focuses on only a few hereditary diseases in which mutations result in unusual pathogenic mechanisms, which have led to some intriguing insights into splicing regulatory mechanisms. Given the essential nature of the splicing

reaction in all tissues, we begin by examining the paradox of why mutations in select spliceosomal components cause retinal degeneration while loss of a snRNP assembly factor leads to a lower motor neuron disease and proximal muscle atrophy. In addition to the core spliceosome, numerous trans-acting factors recognize pre-mRNA sequence elements to orchestrate spliceosome recruitment and alternative splicing. Thus, we next review the roles of both tandem repeats and dispersed repetitive elements in modulating splicing regulation and the emerging roles of repeat polymorphisms and reiterative element mutations on the pathogenesis of specific diseases.

M.G. Poulos et al.

GHOSTS IN THE MACHINE

The spliceosome is a remarkably dynamic macromolecular machine that depends on the recognition of loose consensus sequence elements at splice sites and the branch point (Fig. 2) (Wahl et al. 2009). This splice site detection process is facilitated by a number of other splicing factors that promote snRNP recruitment (Wang and Burge 2008; Licatalosi and Darnell 2010). For splicing to occur, snRNPs and the pre-mRNA undergo a series of associationdissociation steps and conformational transitions and, although the splicing reaction involves only two catalytic steps, intron removal must be accomplished with great precision in all tissues. While splicing errors underlie a large number of human diseases, it is puzzling that some diseases, which are characterized by the degeneration of only certain cell types, are caused by mutations either in ubiquitously expressed spliceosome components or depletion of factors important for snRNP assembly.

The Spliceosome and Retinopathies

Autosomal dominant retinitis pigmentosa (adRP) is characterized by progressive degeneration of predominantly rod photoreceptor neurons resulting in visual impairment. Some forms of adRP are caused by mutations in four pre-mRNA processing factors (PRPF3, PRPF8, PRPF31, SNRNP200) that are components of the ubiquitously expressed U4/U6.U5 trisnRNP, which is recruited as a preassembled complex to form the functional spliceosome (Fig. 2) (Mordes et al. 2006; Wahl et al. 2009; Zhao et al. 2009). Another gene, PIM1-associated protein (PAP1), which encodes a protein that interacts with PRPF3 to regulate splicing, also causes adRP (Maita et al. 2004; Maita et al. 2005). Why do mutations in these specific genes cause a specific eye disease? One possible explanation relates to the unusual metabolic requirements of photoreceptor cells, including circadian oscillation of transcript levels (von Schantz et al. 1999), and the importance of tri-snRNP dynamics during splicing. Although remodeling of spliceosomal subunits is common

during splicing, the U4/U6.U5 tri-snRNP presents an extreme case since catalytic activation of the spliceosome requires disruption of the extensively base-paired U4/U6 duplex to allow U6 snRNA interactions with U2 and the premRNA (Wahl et al. 2009). Evidence that trisnRNP remodeling is particularly important for photoreceptors comes from recent reports that SNRNP200 mutations cause adRP (Li et al. 2009; Zhao et al. 2009). SNRNP200, also known as hBrr2, is a DExD/H-box protein that is required for both unwinding of the U4/U6 duplex and spliceosome disassembly. Interestingly, the SNRNP200 mutation associated with more severe adRP causes a larger defect in U4/U6 unwinding activity (Zhao et al. 2009). Of course, it is possible that PRPF3, PRPF8, PRPF31, SNRNP200, and PAP1 have nonsplicing functions that are particularly critical for retinal function, but this possibility seems less likely given their common association with the tri-snRNP. Both loss-of-function and gain-of-function models have been proposed to explain why mutations in multiple components of the tri-snRNP result in retinal disease (Mordes et al. 2006). For loss-of-function models, tri-snRNP-associated mutations result in haploinsufficiency that only cause overt disease in photoreceptors because these neurons are unusually sensitive to decrements in splicing function due to the high demand for specific proteins, such as rhodopsin (RHO mutations also cause adRP). For example, more than 40 PRPF31 mutations have been reported and the majority of these create a premature termination codon (PTC), transcript degradation via the nonsense-mediated decay (NMD) pathway, and reduced mutant allele expression (Rio Frio et al. 2008). Additionally, the incomplete penetrance noted for PRPF31 mutations may reflect variations in expression from the normal allele (Vithana et al. 2003; Rivolta et al. 2006). A potential problem with this haploinsufficiency model is that other neurons (e.g., Purkinje cells) and other tissues (e.g., regenerating muscle) also place considerable demands on transcription and RNA processing. Indeed, a recent study generated mice carrying either a human ADRP

RNA Splicing Disease

mutation in Prpf31 (Prpf31 A261P/þ knockin) or Prpf31 þ/2 heterozygous knockout mice. Although the mouse retina is enriched in rod photoreceptors compared to humans, these lines fail to model adRP and do not develop retinal degeneration, although the corresponding homozygous knockin and knockout mice are embryonic lethal (Bujakowska et al. 2009). In contrast, heterozygous Rho þ/2 mice, which express lower levels of rhodopsin, develop retinal degeneration (Lem et al. 1999). While this is a single study and may reflect functional differences in the rodent versus human retina, the failure to model adRP in mice will impede further testing of the haploinsufficiency model. A caveat is that Prpf31 A261P/þ knockin mice were generated on a mixed genetic background, so it is possible that mutant Prpf31 alleles will show a retinal phenotype when placed on a different background (Buchner et al. 2003). Additionally, mouse models should be generated for other adRP mutations, particularly Snrnp200. For gain-of-function models, mutant tri-snRNP proteins might alter the spliceosome so that it generates, or fails to efficiently proofread, incorrectly spliced mRNAs (Mordes et al. 2006; Zhao et al. 2009). Another pathogenic mechanism, characteristic of some neurodegenerative diseases, is that U4/U6.U5 tri-snRNP mutations lead to the synthesis of aggregation-prone proteins, which are particularly toxic to photoreceptors. snRNP Biogenesis and Motor Neuron Disease

The spliceosome requires the functions of multiple snRNPs, which contain a seven-member ring of Sm proteins that assemble onto the Sm site of each snRNA in the cytoplasm with the exception of U6. A critical factor in this assembly process in vivo is the survival of motor neurons gene 1 (SMN1), which is ubiquitously expressed in human tissues (Monani 2005; Burghes and Beattie 2009; Chari et al. 2009). In addition to SMN, this ATP-dependent SMN “assemblyosome” complex consists of several GEMINS (SIP1/GEMIN2, DDX20/ GEMIN3, GEMINS4-8) and STRAP/UNRIP

(Paushkin et al. 2002; Burghes and Beattie 2009). Loss of SMN1 expression, due to either mutation or deletion, causes the autosomal recessive disease spinal muscular atrophy (SMA), a leading cause of infantile mortality. SMA is associated with the degeneration of lower motor neurons in the ventral horn of the spinal cord and muscle atrophy, which ranges from severe Type I disease, characterized by an early (,6 months) age of onset and death (,2 years of age), to Type III disease, which manifests later during development (.18 months) and patients may have a normal lifespan (Monani 2005). In a cruel twist of fate, the human genome contains the nearly identical gene SMN2, but this copy fails to compensate for SMN1 loss due to a C ! T transition in SMN2 exon 7. This single nucleotide change blocks inclusion of exon 7 and results in inhibition of SMN protein oligomerization, and/or the generation of a C-terminal degradation signal, leading to low levels of functional protein (Monani 2005; Burghes and Beattie 2009; Cho and Dreyfuss 2010). Similar to adRP, both splicing and nonsplicing models have been proposed for SMA pathogenesis to explain how loss-of-function mutations in a ubiquitously expressed gene result in selective effects on specific cell populations. Splicing models emphasize the possibility that lower motor neurons may be particularly sensitive to SMA-associated declines in SMN activity and relative snRNP abundance, which causes mis-splicing of critical pre-mRNAs. In support of this possibility, there is a strong correlation between phenotypic severity, SMN levels, and snRNP assembly activity in mouse models of SMA (Gabanella et al. 2007). Loss of SMN in these models also causes cell-type specific changes in snRNA levels, with minor spliceosome snRNAs particularly affected in brain and spinal cord (Gabanella et al. 2007; Zhang et al. 2008). Exon microarrays have been used to detect widespread splicing abnormalities in the Smn 2/2 ; SMN2; SMND7 mouse SMA model, with a maximum lifespan of 14 days, at end-stage disease ( postnatal day 11, P11), which led to the suggestion that SMA is a pan-splicing disease (Zhang et al. 2008).

M.G. Poulos et al.

This analysis has been challenged since symptomatic mice were analyzed, suggesting that alternative splicing changes may represent secondary effects due to disease progression. Indeed, a comparison of presymptomatic ( postnatal day 1, P1), early-symptomatic (P7), and late-symptomatic (P13) mice showed that the majority of splicing changes in spinal cord were only observed in late-symptomatic mice (Baumer et al. 2009). Nevertheless, it is possible that many gene expression and RNA splicing changes have been missed in presymptomatic mice using microarrays so it will be important to re-examine these mice using RNASeq technologies. Efforts to develop effective SMA therapies have employed several elegant splicing-related strategies designed to enhance SMN2 exon 7 splicing levels (Khoo and Krainer 2009; Sendtner 2010). Prominent among these are approaches based on bifunctional antisense oligonucleotides (ASOs) or U7 snRNAs composed of sequences complementary to SMN2 exon 7 for targeting coupled to splicing enhancer (ASO-ESE, U7-ESE), or minimal RS domain peptide, motifs to increase exon 7 inclusion. Evidence that this strategy is effective in vivo has been provided recently. SMA model mice (Smn 2/2 ; hSMN2 þ/þ ), which also express a U7-ESE transgene, show a dramatic improvement in survival over Smn 2/2 ; hSMN2 þ/þ mice alone (Meyer et al. 2009). Importantly, extensive ASO tiling analyses across exon 7 and flanking introns 6 and 7 revealed additional splicing silencers, and ASO-mediated antisense masking of these sequence motifs increases exon 7 inclusion in several tissues of human SMN2 transgenic mice (Hua et al. 2007; Hua et al. 2008). Non-splicing functions of SMN have also been reported. Indeed, this protein has been proposed to be a master assembler of RNP complexes (Terns and Terns 2001). For example, the Sm-like (LSm) proteins also form ring structures that function in cytoplasmic RNA turnover and RNA– RNA interactions, but it is not known if SMN activity is essential for this assembly process (Wilusz and Wilusz 2005). Other proposed SMN functions include

transcriptional regulation and axonal mRNP transport and translation (Monani 2005; Burghes and Beattie 2009). REPETITIVE ELEMENTS AND SPLICING

Studies on trans-acting factor mutations have expanded our understanding of spliceosome assembly and function. However, many other diseases are caused by mutations in pre-mRNA sequence elements, and there is a growing appreciation for the importance of reiterative sequences in splicing regulation. Repetitive sequence elements comprise .50% of the human genome and are broadly divided into two categories, dispersed and tandem repeats (Richard et al. 2008). Dispersed repeats include tRNA genes, gene paralogs, and transposon/ transposon-derived elements such as longand short-interspersed elements (LINEs, SINEs) and LTR retrotransposons. Among these, transposable elements (TEs) may cause disease either by insertional mutagenesis or nonallelic homologous recombination between TEs (Belancio et al. 2009). An intriguing example of how these elements can affect splicing regulation is provided by the short interspersed element (SINE) Alu. Alu is a small (300 nt) primate-specific non-LTR retrotransposon that is the most abundant (.106 copies) repetitive element in the human genome (Keren et al. 2010). Alu elements are dimeric structures composed of left and right arms joined by an A-rich linker region and are preferentially located in introns in gene-rich regions often in the antisense orientation. During evolution, Alu mutations have resulted in the appearance of regulatory sequences that are recognized by the spliceosome so that these elements are sometimes “exonized” or included in the mRNA product as internal alternative exons (Lev-Maor et al. 2003; Keren et al. 2010). Other mutations can affect Alu-associated splicing events. For example, a C!T transition in intron 6 of the CTDP1 gene, which encodes a phosphatase associated with the C-terminal domain in RNA polymerase II, generates a 50 splice site while also activating an upstream cryptic 30 splice site in an antisense Alu element, leading

RNA Splicing Disease

to its inclusion and the autosomal recessive disorder congenital cataracts facial dysmorphism neuropathy (CCFDN) (Varon et al. 2003). Tandem repeats include gene tandems, rDNA arrays, and satellite DNAs. The latter group, specifically polymorphic microsatellites, plays a prominent role in hereditary neurological disease and splicing dysregulation. Aside from short repeats, the contraction of a larger repeat, the D4Z4 3.3 kb macrosatellite on chromosome 4q, also appears to underlie at least one muscular dystrophy, and recent studies suggest that contraction-induced alterations in chromatin structure and RNA processing may also be pathogenic.

Microsatellite-induced Disruption of Alternative Splicing

The human genome contains 253,000 di-/tri-/ tetra-nucleotide microsatellite repeats (Richard et al. 2008). These simple sequence repeats may contract or expand due to errors in DNA replication, recombination, and repair. When the latter occurs, transcription of some expanded repeats results in the synthesis of RNA hairpin structures, which are toxic because they affect the activities of several splicing factors. For the neuromuscular disease myotonic dystrophy (DM), the DMPK and CNBP/ZNF9 genes contain CTG and CCTG expansions, respectively. DM type 1 (DM1) is associated with DMPK (CTG)37 – .3,500, while type 2 disease (DM2) is caused by CNBP (CCTG)75 – 11,000, expansions. DM is an autosomal dominant disease that affects multiple systems including skeletal (myotonia, weakness/wasting) and heart (arrhythmias) muscles, the eye (subcapsular cataracts), and the brain (mental retardation, cerebral atrophy). Moreover, DM is both a congenital (CDM) and adult-onset degenerative disease. Curiously, CDM is only associated with CTGexp mutations in the DMPK gene. While the molecular basis for this unique association has not been delineated, one possibility that has not been examined is the potential for problematic transmission of very large CNBP CCTGexp (e.g., .1,000) alleles through the

germline, so disease-associated CNBP mutant alleles might only be generated postnatally. Upon transcription of the mutant DMPK and CNBP/ZNF9 genes, the corresponding CUG and CCUG expansion (CUGexp, CCUGexp) RNAs accumulate in nuclear, or ribonuclear, foci and inhibit the muscleblind-like (MBNL) proteins while also increasing CUGBP1/ ETR-3-like factor (CELF) levels (Fig. 3). Coordinate MBNL loss and CELF1 (formerly CUGBP1) up-regulation occurs because (C)CUGexp RNAs form RNA hairpins that sequester MBNL proteins while also activating protein kinase C (PKC), which results in elevated CELF1 steady state levels due to hyperphosphorylation. The MBNL and CELF proteins act antagonistically to control alternative splicing during development. MBNL1 promotes adult, while CELF1 increases fetal, splicing patterns for specific exons during the fetal to adult transition, so fetal isoform expression persists in adult DM tissues and causes characteristic disease manifestations. The timing of disease onset in adults could reflect the continued somatic expansion of these repeats, particularly in tissues with postmitotic cell populations (e.g., muscle, brain), so that a pathogenic threshold is crossed. Genome-wide expression and splice junction microarray analyses, as well as a crystal structure of two MBNL1 zinc finger-like domains (Zn3/4) bound to r(CGCUGU), indicate that MBNL1 preferentially recognizes a core YGCY motif consistent with its high affinity binding to CUGexp and CCUGexp RNAs (Castle et al. 2008; Teplova and Patel 2008; Du et al. 2010; Goers et al. 2010). In agreement with studies on the brain-specific Nova family of splicing regulators and the importance of binding site position relative to the regulated exon (Ule et al. 2006; Ule and Darnell 2007), binding of MBNL1 upstream of the 30 splice site promotes skipping of that alternative exon while binding downstream of the 50 splice site favors inclusion (Du et al. 2010). A similar scenario has been described for CELF proteins that recognize a UGUGU motif (Faustino and Cooper 2005; Castle et al. 2008; Daughters et al. 2009). CELF proteins contain three RNA recognition

M.G. Poulos et al. (CTG)5 DNA 14

15

Replication recombination repair

Errors

Normal DMPK gene

Transcription

(CUG)5

(A)n

(CTG)>50

Normal DMPK mRNA

MBNL1 recruitment 15 Mutant DMPK gene

MBNL1 CELF1 PKC

Adult splicing

G C U

P U

CELF1 Muscle CLCN1

C G G C

U

6

7a

7b

U

U

4

5

6

U U

C G

8

G C C G G C

U

Brain GRIN1

Transcription

U C G G C

U

Sequestered MBNL 1 U

C G G C C G G C

U

U U

C G C G

Fetal splicing

(A)n Mutant DMPK mRNA

Figure 3. RNA-mediated disease mechanism in myotonic dystrophy. Errors in DNA replication, recombination,

and repair may lead to an increase in a normal length (CTG)5 (small red box) repeat located in DMPK exon 15 to (CTG).50 expansions (large red box). Following transcription, these mutant mRNAs with CUGexp RNA hairpins sequester the MBNL proteins (red ovals) while also triggering activation of protein kinase C (PKC), which results in hyperphosphorylation (white P in black circle) of CELF1 (green oval). MBNL proteins are required during postnatal development to activate adult splicing patterns (red arrows) while CELF1 promotes fetal exon splicing (green arrows). Only specific exons are regulated by these two antagonistic splicing factors. For example, MBNL induces adult-specific skipping of CLCN1 exon 7a in muscle and GRIN1/NMDAR1 exon 5 in brain, while CELF1 promotes inclusion of these fetal exons. Open red and green arrowheads indicate that the precise roles of MBNL and CELF1 in GRIN1 pre-mRNA splicing have not been determined.

motifs (RRM1-3) and the C-terminal RRM of CELF1 forms specific contacts with the UGU trinucleotide (Tsuda et al. 2009). Both Mbnl1 knockout and CELF1 transgenic mouse models have been generated, which recapitulate some features of DM disease (Kanadia et al. 2003; Timchenko et al. 2004; Ho et al. 2005). Nevertheless, the relative contributions of MBNL1 and CELF1 misregulation to pathogenesis are

unclear, and the molecular etiology of congenital DM remains a mystery. The aberrant sequestration of splicing factors by another CG-rich microsatellite has been reported for the late-onset neurological disease fragile X-associated tremor ataxia syndrome (FXTAS). The primary features of FXTAS are action tremor and gait ataxia but cognitive decline, executive function deficits,

RNA Splicing Disease

and mild parkinsonism have also been noted (Hagerman et al. 2001; Grigsby et al. 2007). This neurodegenerative disorder, which was originally identified in older adult carriers of premutation alleles of the fragile X syndrome (FRAXA) mental retardation gene (FMR1), is caused by (CGG)55 – 200 expansions in the FMR1 50 UTR that are shorter than those observed for FRAXA-associated (CGG).230 repeats (Hagerman and Hagerman 2007). In neurons and astrocytes, transcripts from the mutant FMR1 gene accumulate in large ubiquitin-positive intranuclear inclusions, together with a number of other proteins. Interestingly, these toxic CGGexp RNAs are expressed in other cell types, including fibroblasts, and during embryonic development, which might explain the neurodevelopmental abnormalities observed in some premutation children (Garcia-Arocena et al. 2010). Several RNAbinding proteins, including CELF1 and hnRNP A2/B1, have been implicated in FXTAS pathogenesis, but a recent study indicates that the splicing factor Sam68, encoded by the KHDRBS1 gene, is particularly prone to sequestration by CGGexp RNAs (Sellier et al. 2010). Although this is a surprising finding because previous studies have indicated that Sam68 preferentially recognizes A- and U-rich sequences, the Sam68 CGGexp interaction region does not include the central KH RNA-binding domain. Instead, Sam68 interacts with CGGexp RNA via an N-terminal region important for protein – protein interactions, so recruitment to nuclear rCGGexp inclusions is probably mediated by another uncharacterized factor. In support of the hypothesis that Sam68 plays an important role in FXTAS, expression of this protein is required for neuronal differentiation of P19 cells and Khdrbs1 2/ – null mice develop motor coordination deficiencies, as indicated by rotarod and hindlimb beam walking tests, although cerebellar degeneration has not been reported (Lukong and Richard 2008; Chawla et al. 2009). Significantly, Sam68 depletion, either by sequestration in FXTAS brains or by shRNA-mediated knockdowns, results in the alternative splicing of some Sam68 target exons. Unlike DM, where specific mis-splicing events

have been linked to distinct pathological consequences (e.g., myotonia due to aberrant inclusion of CLCN1 exon 7a), it is not clear if FXTAS is a developmental splicing disease. To address this concern, splicing-sensitive microarray and RNA-Seq analyses should be performed to determine if a FXTAS-specific splicing signature emerges. Microsatellite expansions and the disruption of developmentally regulated splicing transitions have been implicated in other neurological diseases. The autosomal dominant spinocerebellar ataxias (SCAs) are a group of late-onset and progressive neurodegenerative diseases that are characterized by cerebellar atrophy resulting in ataxia (abnormal limb movement/coordination), tremor, and premature death (Carlson et al. 2009). SCA type 8 (SCA8) is particularly interesting since it is transmitted with reduced penetrance and caused by CTG†CAG repeat expansions in both coding (ATXN8 gene) and non-coding (ATXN8OS) regions since the locus is bidirectionally transcribed (Moseley et al. 2006). For ATXN8OS, CUGexp RNAs accumulate in ribonuclear foci that are detectable in the human SCA8 cerebellum, particularly in molecular layer interneurons and Bergmann glia. Ribonuclear foci are also detectable in a BAC transgenic mouse model for SCA8. These mice express a human (CTG†CAG)116 expansion and develop characteristic motor deficits possibly due to loss of cerebellar GABAergic inhibition. A recent study confirmed that expression of ATXN8OS CUGexp RNAs affects cerebellar inhibitory pathways by altering the activities of developmentally regulated splicing events controlled by CELF1 and MBNL1. Using the crosslinking/immunoprecipation protocol (CLIP), a number of brain RNA targets for mouse Cugbp1 were revealed including the GABA-A transporter 4, GAT4/GABT4A. Expression of CUGexp RNA, which was shown to be coordinately regulated by MBNL1 and CUGBP1, leads to persistence of the fetal GAT4/GABT4 splicing pattern, chronically elevated GAT4/GABT4 levels, and loss of GABAergic inhibition in the granular layer of the cerebellum (Daughters et al. 2009).

M.G. Poulos et al.

SCA type 31 (SCA31) has also been reported to be associated with a TGGAAexp insertion in newly indentified introns of the TK2 and BEAN genes, which partially overlap and are transcribed in opposite directions on chromosome 16q22.1 (Sato et al. 2009). Similar to DM, mutant SCA31 RNAs accumulate in ribonuclear foci, so this SCA may also be an RNAmediated disease, although these foci are detectable in Purkinje cells in SCA31. However, the splicing factors involved may be different since UGGAA repeats are not predicted to form stem-loop structures. A clue to the identity of these factors has emerged from studies on repetitive satellite III (SatIII) DNA repeats, which are composed of (GGAAT)n CAAC(C/ A)CGAGT repeats (Valgardsdottir et al. 2008). Thermal and chemical stresses trigger SatIII transcription and these RNAs concentrate in nuclear stress bodies (nSBs), which also contain splicing factors including SFRS1 (ASF/SF2). Interestingly, SFRS1 and SFRS9 (SRp30c) bind to (UGGAA)8 RNAs in vitro, so it is possible that sequestration of these splicing factors causes misregulation of alternative splicing in Purkinje cells and degeneration (Sato et al. 2009). In contrast to DM and SCA31, Friedreich ataxia (FRDA) is an autosomal recessive disease characterized by atrophy of primary sensory neurons in the dorsal root ganglia as well as hypertrophic cardiomyopathy (Pandolfo 2008). FRDA is caused by noncoding GAA expansions in the first intron of the FXN gene, which encodes the frataxin protein that is associated with the mitochondrial matrix and involved in iron homeostasis. Normal FXN alleles have up to 38 GAA repeats that expand to 70 to .1,000 repeats in FRDA. These GAA expansions are currently believed to be pathogenic because they inhibit transcription by altering DNA structure, by forming non-B DNA triplexes and “sticky” DNA, and by promoting chromatin condensation (Sakamoto et al. 1999; Saveliev et al. 2003). Recently, GAA expansions have also been shown to influence alternative splicing regulation. Insertion of a (GAA)100 expansion downstream of an alternative exon in a hybrid minigene splicing reporter resulted in position-dependent effects on exon

skipping, flanking intron retention, and splicing of a cryptic exon containing the (GAA)100 repeat (Baralle et al. 2008). More significantly, insertion of (GAA)100, but not a complementary (TTC)100 repeat, into a frataxin minigene reporter did not alter nascent transcript levels but did inhibit splicing of FXN intron 1. Thus, missplicing of FXN pre-mRNA may be a primary event that leads to decreased FXN mRNA levels. Satellite Contractions: Potential Effects on RNA Processing

While loss-of-function mutations alter cisacting splicing regulatory sequences in the DMD gene to cause exon skipping and frameshifting in DMD/BMD, toxic C(C)UGexp RNAs expressed in DM alter the activities of trans-acting splicing factors that control the expression of developmentally regulated isoforms. A third mechanism that affects splicing regulation may be important in the molecular etiology of another muscular dystrophy, facioscapulohumeral muscular dystrophy (FSHD). FSHD is caused by the contraction of a D4Z4 macrosatellite repeat array in the subtelomeric region of chromosome 4qter (van der Maarel et al. 2007; Tawil 2008). This 3.3 kb repeat is normally 11 – 100 D4Z4 units in length but contracts to 1– 10 repeats in affected individuals and is preferentially associated with the 4qA161 haplotype. Current position-effect models propose that repeat contraction leads to a decrease in D4Z4 epigenetic modifications resulting in increased expression of genes that are either centromeric, including the spliceosome-associated protein FRG1, or within the repeat, such as the toxic double homeobox protein DUX4. In support of the former proposal, transgenic mice overexpressing FRG1 10- to 50-fold develop several FSHD-associated muscle features including kyphosis and RNA splicing changes (Gabellini et al. 2006). However, microarray expression studies have challenged the possibility that FRG1 is overexpressed in FSHD muscle (Osborne et al. 2007; Klooster et al. 2009). Alternatively, contractionrelated changes in chromatin structure, or hypomethylation in the absence of contractions

RNA Splicing Disease

also known as phenotypic FSHD, may lead to changes in transcript abundance and/or processing from specific D4Z4 repeats that are toxic to muscle cells. Recent studies have focused on the most telomeric D4Z4 repeat, which together with allele-specific flanking sequences, produces transcripts that are processed into si/miRNAsized fragments as well as alternatively processed DUX4 RNAs that potentially antagonize myogenesis (Dixit et al. 2007; Bosnakovski et al. 2008; Bosnakovski et al. 2009; Snider et al. 2009). Future studies should determine if D4Z4, and perhaps other repetitive element, contractions activate transcription of locusspecific repeat arrays while coordinately altering transcript processing to generate pathogenic RNAs and/or proteins. Concluding Remarks

Studies designed to clarify the molecular basis of some puzzling hereditary neurological disorders have provided both surprising results and fertile territory for further exploration. The key question of why mutations in genes encoding broadly expressed factors target specific cell types, such as tri-snRNP-associated proteins in adRP and SMN1 in SMA, remains unresolved, but the link to splicing misregulation has been reinforced by recent reports. Dynamic mutations associated with several muscular dystrophies and ataxias have also highlighted a novel disease mechanism, RNA-mediated pathogenesis, and the critical roles of novel alternative splicing factors during development. Future developments should yield more surprises and strengthen the connection between aberrant RNA processing and disease. ACKNOWLEDGMENTS

Research in our lab is funded by the NIH (AR046799, NS048843, NS058901) and the MDA. REFERENCES Baralle M, Pastor T, Bussani E, Pagani F. 2008. Influence of Friedreich ataxia GAA noncoding repeat expansions on pre-mRNA processing. Am J Hum Genet 83: 77–88.

Baumer D, Lee S, Nicholson G, Davies JL, Parkinson NJ, Murray LM, Gillingwater TH, Ansorge O, Davies KE, Talbot K. 2009. Alternative splicing events are a late feature of pathology in a mouse model of spinal muscular atrophy. PLoS Genet 5: e1000773. Behzadnia N, Golas MM, Hartmuth K, Sander B, Kastner B, Deckert J, Dube P, Will CL, Urlaub H, Stark H, et al. 2007. Composition and three-dimensional EM structure of double affinity-purified, human prespliceosomal A complexes. EMBO J 26: 1737– 1748. Belancio VP, Deininger PL, Roy-Engel AM. 2009. LINE dancing in the human genome: transposable elements and disease. Genome Med 1: 97. Bessonov S, Anokhina M, Will CL, Urlaub H, Luhrmann R. 2008. Isolation of an active step I spliceosome and composition of its RNP core. Nature 452: 846 –850. Black DL, Grabowski PJ. 2003. Alternative pre-mRNA splicing and neuronal function. Prog Mol Subcell Biol 31: 187– 216. Blencowe BJ. 2006. Alternative splicing: new insights from global analyses. Cell 126: 37–47. Bosnakovski D, Daughters RS, Xu Z, Slack JM, Kyba M. 2009. Biphasic myopathic phenotype of mouse DUX, an ORF within conserved FSHD-related repeats. PLoS One 4: e7003. Bosnakovski D, Xu Z, Gang EJ, Galindo CL, Liu M, Simsek T, Garner HR, Agha-Mohammadi S, Tassin A, Coppee F, et al. 2008. An isogenetic myoblast expression screen identifies DUX4-mediated FSHD-associated molecular pathologies. EMBO J 27: 2766–2779. Buchner DA, Trudeau M, Meisler MH. 2003. SCNM1, a putative RNA splicing factor that modifies disease severity in mice. Science 301: 967–969. Bujakowska KM, Maubaret C, Chakarova CF, Tanimoto N, Beck SC, Fahl E, Humphries MM, Kenna P, Makarov E, Makarova O, et al. 2009. Study of gene targeted mouse models of splicing factor gene Prpf31 implicated in human autosomal dominant retinitis pigmentosa (RP). Invest Ophthalmol Vis Sci. 50: 5927–5933. Burghes AH, Beattie CE. 2009. Spinal muscular atrophy: why do low levels of survival motor neuron protein make motor neurons sick? Nat Rev Neurosci 10: 597– 609. Carlson KM, Andresen JM, Orr HT. 2009. Emerging pathogenic pathways in the spinocerebellar ataxias. Curr Opin Genet Dev 19: 247– 253. Castle JC, Zhang C, Shah JK, Kulkarni AV, Kalsotra A, Cooper TA, Johnson JM. 2008. Expression of 24,426 human alternative splicing events and predicted cis regulation in 48 tissues and cell lines. Nat Genet 40: 1416–1425. Chari A, Paknia E, Fischer U. 2009. The role of RNP biogenesis in spinal muscular atrophy. Curr Opin Cell Biol 21: 387 –393. Chasin LA. 2007. Searching for splicing motifs. Adv Exp Med Biol 623: 85– 106. Chawla G, Lin CH, Han A, Shiue L, Ares M Jr, Black DL. 2009. Sam68 regulates a set of alternatively spliced exons during neurogenesis. Mol Cell Biol 29: 201 –213. Cho S, Dreyfuss G. 2010. A degron created by SMN2 exon 7 skipping is a principal contributor to spinal muscular atrophy severity. Genes Dev 24: 438– 442.

M.G. Poulos et al. Cooper TA, Wan L, Dreyfuss G. 2009. RNA and disease. Cell 136: 777– 793. Daughters RS, Tuttle DL, Gao W, Ikeda Y, Moseley ML, Ebner TJ, Swanson MS, Ranum LP. 2009. RNA gain-of-function in spinocerebellar ataxia type 8. PLoS Genet 5: e1000600. Deckert J, Hartmuth K, Boehringer D, Behzadnia N, Will CL, Kastner B, Stark H, Urlaub H, Luhrmann R. 2006. Protein composition and electron microscopy structure of affinity-purified human spliceosomal B complexes isolated under physiological conditions. Mol Cell Biol 26: 5528–5543. Dixit M, Ansseau E, Tassin A, Winokur S, Shi R, Qian H, Sauvage S, Matteotti C, van Acker AM, Leo O, et al. 2007. DUX4, a candidate gene of facioscapulohumeral muscular dystrophy, encodes a transcriptional activator of PITX1. Proc Natl Acad Sci U S A 104: 18157–18162. Du H, Cline MS, Osborne RJ, Tuttle DL, Clark TA, Donohue JP, Hall MP, Shiue L, Swanson MS, Thornton CA, et al. 2010. Aberrant alternative splicing and extracellular matrix gene expression in mouse models of myotonic dystrophy. Nat Struct Mol Biol 17: 187– 193. Ervasti JM. 2007. Dystrophin, its interactions with other proteins, and implications for muscular dystrophy. Biochim Biophys Acta 1772: 108 –117. Faustino NA, Cooper TA. 2005. Identification of putative new splicing targets for ETR-3 using sequences identified by systematic evolution of ligands by exponential enrichment. Mol Cell Biol 25: 879– 887. Gabanella F, Butchbach ME, Saieva L, Carissimi C, Burghes AH, Pellizzoni L. 2007. Ribonucleoprotein assembly defects correlate with spinal muscular atrophy severity and preferentially affect a subset of spliceosomal snRNPs. PLoS One 2: e921. Gabellini D, D’Antona G, Moggio M, Prelle A, Zecca C, Adami R, Angeletti B, Ciscato P, Pellegrino MA, Bottinelli R, et al. 2006. Facioscapulohumeral muscular dystrophy in mice overexpressing FRG1. Nature 439: 973– 977. Garcia-Arocena D, Yang JE, Brouwer JR, Tassone F, Iwahashi C, Berry-Kravis EM, Goetz CG, Sumis AM, Zhou L, Nguyen DV, et al. 2010. Fibroblast phenotype in male carriers of FMR1 premutation alleles. Hum Mol Genet 19: 299–312. Goers ES, Purcell J, Voelker RB, Gates DP, Berglund JA. 2010. MBNL1 binds GC motifs embedded in pyrimidines to regulate alternative splicing. Nucleic Acids Res 38: 2467– 2484. Grigsby J, Brega AG, Leehey MA, Goodrich GK, Jacquemont S, Loesch DZ, Cogswell JB, Epstein J, Wilson R, Jardini T, et al. 2007. Impairment of executive cognitive functioning in males with fragile X-associated tremor/ataxia syndrome. Mov Disord 22: 645 –650. Gurvich OL, Tuohy TM, Howard MT, Finkel RS, Medne L, Anderson CB, Weiss RB, Wilton SD, Flanigan KM. 2008. DMD pseudoexon mutations: splicing efficiency, phenotype, and potential therapy. Ann Neurol 63: 81–89. Hagerman PJ, Hagerman RJ. 2007. Fragile X-associated tremor/ataxia syndrome– an older face of the fragile X gene. Nat Clin Pract Neurol 3: 107 –112. Hagerman RJ, Leehey M, Heinrichs W, Tassone F, Wilson R, Hills J, Grigsby J, Gage B, Hagerman PJ. 2001. Intention

tremor, parkinsonism, and generalized brain atrophy in male carriers of fragile X. Neurology 57: 127– 130. Ho TH, Bundman D, Armstrong DL, Cooper TA. 2005. Transgenic mice expressing CUG-BP1 reproduce splicing mis-regulation observed in myotonic dystrophy. Hum Mol Genet 14: 1539– 1547. House AE, Lynch KW. 2008. Regulation of alternative splicing: more than just the ABCs. J Biol Chem 283: 1217–1221. Hua Y, Vickers TA, Baker BF, Bennett CF, Krainer AR. 2007. Enhancement of SMN2 exon 7 inclusion by antisense oligonucleotides targeting the exon. PLoS Biol 5: e73. Hua Y, Vickers TA, Okunola HL, Bennett CF, Krainer AR. 2008. Antisense masking of an hnRNP A1/A2 intronic splicing silencer corrects SMN2 splicing in transgenic mice. Am J Hum Genet 82: 834– 848. Kanadia RN, Johnstone KA, Mankodi A, Lungu C, Thornton CA, Esson D, Timmers AM, Hauswirth WW, Swanson MS. 2003. A muscleblind knockout model for myotonic dystrophy. Science 302: 1978– 1980. Keren H, Lev-Maor G, Ast G. 2010. Alternative splicing and evolution: diversification, exon definition and function. Nat Rev Genet 11: 345– 355. Khoo B, Krainer AR. 2009. Splicing therapeutics in SMN2 and APOB. Curr Opin Mol Ther 11: 108– 115. Klooster R, Straasheijm K, Shah B, Sowden J, Frants R, Thornton C, Tawil R, van der Maarel S. 2009. Comprehensive expression analysis of FSHD candidate genes at the mRNA and protein level. Eur J Hum Genet 17: 1615–1624. Konarska MM, Query CC. 2005. Insights into the mechanisms of splicing: more lessons from the ribosome. Genes Dev 19: 2255–2260. Lem J, Krasnoperova NV, Calvert PD, Kosaras B, Cameron DA, Nicolo M, Makino CL, Sidman RL. 1999. Morphological, physiological, and biochemical changes in rhodopsin knockout mice. Proc Natl Acad Sci U S A 96: 736– 741. Lev-Maor G, Sorek R, Shomron N, Ast G. 2003. The birth of an alternatively spliced exon: 30 splice-site selection in Alu exons. Science 300: 1288–1291. Li N, Mei H, Macdonald IM, Jiao X, Hejtmancik F. 2009. Mutations in ASCC3L1 on chromosome 2q11.2 are associated with autosomal dominant retinitis pigmentosa in a Chinese Family. Invest Ophthalmol Vis Sci 51: 1036–1043 Licatalosi DD, Darnell RB. 2006. Splicing regulation in neurologic disease. Neuron 52: 93–101. Licatalosi DD, Darnell RB. 2010. RNA processing and its regulation: global insights into biological networks. Nat Rev Genet 11: 75–87. Long JC, Caceres JF. 2009. The SR protein family of splicing factors: master regulators of gene expression. Biochem J 417: 15–27. Lukong KE, Richard S. 2008. Motor coordination defects in mice deficient for the Sam68 RNA-binding protein. Behav Brain Res 189: 357– 363. Maita H, Kitaura H, Ariga H, Iguchi-Ariga SM. 2005. Association of PAP-1 and Prp3p, the products of causative genes of dominant retinitis pigmentosa, in the tri-snRNP complex. Exp Cell Res 302: 61–68.

RNA Splicing Disease Maita H, Kitaura H, Keen TJ, Inglehearn CF, Ariga H, Iguchi-Ariga SM. 2004. PAP-1, the mutated gene underlying the RP9 form of dominant retinitis pigmentosa, is a splicing factor. Exp Cell Res 300: 283– 296. Martinez-Contreras R, Cloutier P, Shkreta L, Fisette JF, Revil T, Chabot B. 2007. hnRNP proteins and splicing control. Adv Exp Med Biol 623: 123–147. Meyer K, Marquis J, Tru¨b J, Nlend RN, Verp S, Ruepp M-D, Imboden H, Barde I, Trono D, Schu¨mperli D. 2009. Rescue of a severe mouse model for spinal muscular atrophy by U7 snRNA-mediated splicing modulation. Hum Mol Genet 18: 546 –555. Monani UR. 2005. Spinal muscular atrophy: a deficiency in a ubiquitous protein; a motor neuron-specific disease. Neuron 48: 885–896. Mordes D, Luo X, Kar A, Kuo D, Xu L, Fushimi K, Yu G, Sternberg P Jr, Wu JY. 2006. Pre-mRNA splicing and retinitis pigmentosa. Mol Vis 12: 1259– 1271. Moseley ML, Zu T, Ikeda Y, Gao W, Mosemiller AK, Daughters RS, Chen G, Weatherspoon MR, Clark HB, Ebner TJ, et al. 2006. Bidirectional expression of CUG and CAG expansion transcripts and intranuclear polyglutamine inclusions in spinocerebellar ataxia type 8. Nat Genet 38: 758 –769. Osborne RJ, Welle S, Venance SL, Thornton CA, Tawil R. 2007. Expression profile of FSHD supports a link between retinal vasculopathy and muscular dystrophy. Neurology 68: 569 –577. Pandolfo M. 2008. Friedreich ataxia. Arch Neurol 65: 1296– 1303. Paushkin S, Gubitz AK, Massenet S, Dreyfuss G. 2002. The SMN complex, an assemblyosome of ribonucleoproteins. Curr Opin Cell Biol 14: 305 –312. Richard GF, Kerrest A, Dujon B. 2008. Comparative genomics and molecular dynamics of DNA repeats in eukaryotes. Microbiol Mol Biol Rev 72: 686– 727. Rio Frio T, Wade NM, Ransijn A, Berson EL, Beckmann JS, Rivolta C. 2008. Premature termination codons in PRPF31 cause retinitis pigmentosa via haploinsufficiency due to nonsense-mediated mRNA decay. J Clin Invest 118: 1519–1531. Rivolta C, McGee TL, Rio Frio T, Jensen RV, Berson EL, Dryja TP. 2006. Variation in retinitis pigmentosa-11 (PRPF31 or RP11) gene expression between symptomatic and asymptomatic patients with dominant RP11 mutations. Hum Mutat 27: 644 –653. Sakamoto N, Chastain PD, Parniewski P, Ohshima K, Pandolfo M, Griffith JD, Wells RD. 1999. Sticky DNA: selfassociation properties of long GAA.TTC repeats in R.R.Y triplex structures from Friedreich’s ataxia. Mol Cell 3: 465– 475. Sato N, Amino T, Kobayashi K, Asakawa S, Ishiguro T, Tsunemi T, Takahashi M, Matsuura T, Flanigan KM, Iwasaki S, et al. 2009. Spinocerebellar Ataxia Type 31 Is Associated with “Inserted” Penta-Nucleotide Repeats Containing (TGGAA)(n). Am J Hum Genet 85: 544 –557. Saveliev A, Everett C, Sharpe T, Webster Z, Festenstein R. 2003. DNA triplet repeats mediate heterochromatinprotein-1-sensitive variegated gene silencing. Nature 422: 909– 913.

Sellier C, Rau F, Liu Y, Tassone F, Hukema RK, Gattoni R, Schneider A, Richard S, Willemsen R, Elliott DJ, et al. 2010. Sam68 sequestration and partial loss of function are associated with splicing alterations in FXTAS patients. EMBO J 29: 1248–1261. Sendtner M. 2010. Therapy development in spinal muscular atrophy. Nat Neurosci 13: 795– 799. Snider L, Asawachaicharn A, Tyler AE, Geng LN, Petek LM, Maves L, Miller DG, Lemmers RJ, Winokur ST, Tawil R, et al. 2009. RNA transcripts, miRNA-sized fragments and proteins produced from D4Z4 units: new candidates for the pathophysiology of facioscapulohumeral dystrophy. Hum Mol Genet 18: 2414– 2430. Steitz JA, Dreyfuss G, Krainer AR, Lamond AI, Matera AG, Padgett RA. 2008. Where in the cell is the minor spliceosome? Proc Natl Acad Sci U S A 105: 8485–8486. Tawil R. 2008. Facioscapulohumeral muscular dystrophy. Neurotherapeutics 5: 601 –606. Tazi J, Bakkour N, Stamm S. 2009. Alternative splicing and disease. Biochim Biophys Acta 1792: 14–26. Teplova M, Patel DJ. 2008. Structural insights into RNA recognition by the alternative-splicing regulator muscleblind-like MBNL1. Nat Struct Mol Biol 15: 1343–1351. Terns MP, Terns RM. 2001. Macromolecular complexes: SMN–the master assembler. Curr Biol 11: R862–864. Timchenko NA, Patel R, Iakova P, Cai ZJ, Quan L, Timchenko LT. 2004. Overexpression of CUG triplet repeat-binding protein, CUGBP1, in mice inhibits myogenesis. J Biol Chem 279: 13129–13139. Tsuda K, Kuwasako K, Takahashi M, Someya T, Inoue M, Terada T, Kobayashi N, Shirouzu M, Kigawa T, Tanaka A, et al. 2009. Structural basis for the sequence-specific RNA-recognition mechanism of human CUG-BP1 RRM3. Nucleic Acids Res 37: 5151– 5166. Ule J, Darnell RB. 2007. Functional and mechanistic insights from genome-wide studies of splicing regulation in the brain. Adv Exp Med Biol 623: 148–160. Ule J, Stefani G, Mele A, Ruggiu M, Wang X, Taneri B, Gaasterland T, Blencowe BJ, Darnell RB. 2006. An RNA map predicting Nova-dependent splicing regulation. Nature 444: 580 –586. Valgardsdottir R, Chiodi I, Giordano M, Rossi A, Bazzini S, Ghigna C, Riva S, Biamonti G. 2008. Transcription of Satellite III non-coding RNAs is a general stress response in human cells. Nucleic Acids Res 36: 423 –434. van der Maarel SM, Frants RR, Padberg GW. 2007. Facioscapulohumeral muscular dystrophy. Biochim Biophys Acta 1772: 186–194. Varon R, Gooding R, Steglich C, Marns L, Angelicheva D, Yong KK, Ambrugger P, Reinhold A, Morar B, Baas F, et al. Partial deficiency of the C-terminal-domain phosphatase of RNA polymerase II is associated with congenital cataracts facial dysmorphism neuropathy syndrome. Nat Genet 35: 185 –189. Vithana EN, Abu-Safieh L, Pelosini L, Winchester E, Hornan D, Bird AC, Hunt DM, Bustin SA, Bhattacharya SS. 2003. Expression of PRPF31 mRNA in patients with autosomal dominant retinitis pigmentosa: a molecular clue for incomplete penetrance? Invest Ophthalmol Vis Sci 44: 4204– 4209.

M.G. Poulos et al. von Schantz M, Lucas RJ, Foster RG. 1999. Circadian oscillation of photopigment transcript levels in the mouse retina. Brain Res Mol Brain Res 72: 108 –114. Wahl MC, Will CL, Luhrmann R. 2009. The spliceosome: design principles of a dynamic RNP machine. Cell 136: 701–718. Wang ET, Sandberg R, Luo S, Khrebtukova I, Zhang L, Mayr C, Kingsmore SF, Schroth GP, Burge CB. 2008. Alternative isoform regulation in human tissue transcriptomes. Nature 456: 470–476. Wang Z, Burge CB. 2008. Splicing regulation: from a parts list of regulatory elements to an integrated splicing code. RNA 14: 802–813. Will CL, Luhrmann R. 2005. Splicing of a rare class of introns by the U12-dependent spliceosome. Biol Chem 386: 713– 724.

Wilusz CJ, Wilusz J. 2005. Eukaryotic Lsm proteins: lessons from bacteria. Nat Struct Mol Biol 12: 1031– 1036. Zhang Z, Lotti F, Dittmar K, Younis I, Wan L, Kasim M, Dreyfuss G. 2008. SMN deficiency causes tissue-specific perturbations in the repertoire of snRNAs and widespread defects in splicing. Cell 133: 585 –600. Zhao C, Bellur DL, Lu S, Zhao F, Grassi MA, Bowne SJ, Sullivan LS, Daiger SP, Chen LJ, Pang CP, et al. 2009. Autosomal-dominant retinitis pigmentosa caused by a mutation in SNRNP200, a gene required for unwinding of U4/U6 snRNAs. Am J Hum Genet 85: 617– 627. Zhong X, Liu JR, Kyle JW, Hanck DA, Agnew WS. 2006. A profile of alternative RNA splicing and transcript variation of CACNA1H, a human T-channel gene candidate for idiopathic generalized epilepsies. Hum Mol Genet 15: 1497–1512.

Index

A

Base excision repair (BER), 363

A b fl, 184

Bauer, Franz, I

Access, Repair, and Restore model, 367

BCL11A, 282

Achalsia-Addisonianism-alacrima syndrome, 406

BC L2 , 126 Becker muscular dystrophy (B M D ), 450, 458

Actin chromatin remodeling, 196

Beckwith-Wiedeman syndrome (BVVS), 422

functional overview, 195

BER. See Base excision repair

large-scale organization o f genome, 203-204

BI.M, 253-254

polymerization regulation, 201-202

B M D . See Becker muscular dystrophy

prospects for study, 204

Bonner, James, 6

transcription function

BRCA1, 370, 373

machinery association, 196—197

Breast cancer, gene positioning, 126

nascent transcript association, 197-201

B rg l, 196, 200

ribonucleoprotein association, 201

BRMS, 273

Actin-related proteins (ARPs), 196

Brown, Donald, 210

Aging

Brown, Robert, 1

chromatin higher-order structure, 429-431

Budding yeast. See Yeast nucleus, budding

nucleolus in cellular senescence, 218

Bushke-OUendorff syndrome, 405

А КТ, 126, 256-257

B-WASP, 199, 202

A I .C L See Anaplastic large cell lymphoma

B-WICF1 complex, 201

A L K , 426

BWS. See Beckwith-Wiedeman syndrome

Anaplastic large cell lymphoma (AI.CL), 426,428 Androgen receptor (AR), 426

С

A N T 2 , 151 A O A l. See Ataxia with ocular aprexia type 1

C A C N A 1H , 450

AR. See Androgen receptor

CAF1, 367

A R P2 /3 , 202

Cajal body (CB)

ARPs. See Actin-related proteins

activity independence, 320

A rx l, 81

amphibian oocytes, 243-244

A s fl, 182, 367

biogenesis, 317, 321

ASP I a, 258

coilin as marker, 238-239

AT. See Ataxia telangiectasia

de novo formation, 321

Ataxia telangiectasia (AT), 442- 443

discovery, 3,6, 237

Ataxia with ocular aprexia type 1 (A O A l), 443

dynamics and assembly, 244-245

Ataxin-1 (ATXN1), 438-440

gene loci, 242

A T M , 254, 365, 371, 424-425, 442-443

gene positioning, 121

ATR, 254, 365, 370-371, 373

maintenance, 318-319

ATRIP, 365

mode o f inheritance, 317, 325

ATXN 1. See Ataxin-1

prospects for study, 245

ATXN8, 457

small Cajal body-associated R N A , 239-241 small nuclear ribonucleoprotein association, 239 S M N , 241-242, 245

В

Cajal, Santiago Ramon, 3, 237, 437

BAF. See Barrier-lo-autoinlegration factor

Cancer chromosome organization, 427-429

BAF1, 176 Barrier-to-autointegration factor (BAF), 25, 57-59, 151, 1%

HP1 defects, 424 perinucleolar compartment

prospects for study, 152-154

Cancer (continued)

30-nm fiber, 135-136

malignancy function, 274 metastasis, 272-273

isolation studies, 6

therapeutic targeting, 273-274

lam in function epigenetic regulation, 43

transformation studies, 272

positioning and gene expression, 40

Capping, m R N A , 384-385 CAPs. See Chromatin architectural proteins

transcription. See Transcription

CB. See Cajal body

yeast

C D 2 , 122

dynamics, 177-178

C D 8 a , 122

trans-association o f chromatin loci, 182-183

C d c l3 , 180

Chromatin architectural proteins (CAPs), 139-140

Cdc25A, 370-371

Chromatin im m unoprecipitation (ChIP), 165

Cdc48, 26

Chromosome

C D H 1,422

lampbrush chromosome, 147-148

C d k l, 21-22,37

polytene chromosome, 147-148

Cdk9, 390

yeast folding, 177

C ELF1, 455-457

Chromosome territory (CT)

CF.NP-F, 23

cell cycle changes, 104

C F IA , 390

conformation, 102-103

C F IB , 390

experimental evidence, 94-97

CFIm , 282, 390

gene loci dynamics and interactions, 104-106

CFIIm , 390

gene positioning, 124

C F T R , 116

history o f study, 6 - 7 , 94

Charcot-Marie-Tooth disease, 403

nonrandom arrangements

СЫР. See Chrom atin immunoprecipitation

distinguishing random and nonrandom

C H K 1 , 371,425

arrangements, 97-99 proximity patterns, 102

C H K 2 , 254, 365, 371,425 Chrom atin

radial nuclear arrangements, 99, 101-102

aging and higher-order structure, 429-431 disease and mutations o f architectural proteins cohcsin, 422-423

plasticity, 103-104 prospects for study, 106-109 Clastosome

CTCF, 421-422

overview, 304-307

HP1, 424

proteasomc component, 307

c-Mvc, 423-424

C I.CN 1,457

SATB 1,420-421

Cleavage body, 304

D N A damage response organization, 366-369

C lk, 228, 232

D N A repair and higher-order structure, 424-427

CLS. See Cornelia de Lange syndrome

dynamics and transcription, 148-150

CNBP, 455

euchromatin, 142-143

C o A A , 282

heterochromatin, 142-143

C o A M , 282

higher-order structure

Cohesin, mutation and disease, 422-423

computer modeling, 138-139

C o ilin

electron microscopy, 140-141

Cajal body marker, 238-239, 322

fibers in nuclei, 136-137

structure and function, 238-239

h isto n eFU function, 136

Cornelia de Lange syndrome (CLS), 423

lampbrush chromosome, 147-148

CPF, 390-391

light microscopy, 141-142

CPSF, 311,390-391

local events in modulation, 139-140

C R M 1,393

loops, 143-147

C S F I R , 126

mechanical properties o f isolated fibers,

CstF, 311,390

137-138 metaphase chromosome, 14 ' nucleosom e-nudeosom e i isolated fibers, 137

CT. See Chromosome territory ’

CTCF, 45, 117, 168, 421-422 ns in

C T D P 1,454 C tfl8 , 182

overview, 133-135

C lk 1,390

polytene chromosome, 147-

C tn , 283

C U G -b in d in g protein, 268-269

Caetiorhabditis elegans studies, 162

C y cB l, 21-22

D rosophila rnelanogasler studies, 160-162

Cytochalasin D, 202

overview, 159-160 prospects for study, 170 Dosage compensation complex (D C C ), 162

D

Double-strand break repair (DSBR), 363-365, 369372, 428- 429

Dam. See D N A adenine methyltransferase Daughter-strand gap repair (DSGR), 365

DSBR. See Double-strand break repair

D A X X , 255, 258

DSGR. See Daughter-strand gap repair

Daxx, 325

Duchenne muscular dystrophy ( D M D ) , 450, 458

D B H S domain, 199

Dunnigan-type familial partial lipodystrophy, 408

D B H S proteins, 297

D U X 4 , 458

Dbp5, 80-81, 83-84

DYT1 dystonia, 406

D C C . See Dosage compensation complex D D R. See D N A damage response DD X1, 311-312

E

D H B S proteins, 280-281

E D M D . See Em ery-D reifus muscular dystrophy

Discovery, nucleus, 1- 2

Edsall, John, 7

D M D , 450

EJC. See Exon-junction complex

D M D . See Duchennc muscular dystrophy

E M A N IC , 138

D M P K , 455

E M D . See Emerin

D N A adenine methyltransferase (Dam), 42

Emerin (E M D ), 62, 404

D N A damage response (DDR)

Em ery-D reifus muscular dystrophy (E D M D ), 64-65,

intertwined DNA-lransacting processes, 366

124, 402-404

live cell studies, 369

Epilepsy, 124

organization

Eraf, 117

chromatin, 366-369

ER B 2, 126

double-strand break response, 369-372

ERBB2, 126

nucleotide excision repair, 372-373

ERCC1, 363, 366, 372

somatic cells, 373-374

E R G , 126

overview, 361 -362

E scl, 176, 181, 183

perinucleolar compartment, 271

Esc2, 182

prospects for study, 374

ESE. See Exonic splicing enhancer

repair pathways. See D N A repair

E stl, 180-181, 188

signaling, 365-366

E st2,180-181

tolerance to damage, 365

E st3,180

DNA-dependent protein kinase, 365-366

E T V 1 , 126

D N A ligasc I, 349

Euchromatin. See Chromatin

D N A ligase III, 363

Evolution, nucleus, 2

D N A ligase IV, 365

Exon-junction complex (FJC), 381, 388, 393

D N A methyltransferase, D N M T 3 B , 124

Exonic splicing enhancer (ESE), 450

D N A repair

Exonuclease I, 365

base excision repair, 363 chromatin higher-order structure, 424-427 diseases, 442-444

F

double-strand break repair, 363-365, 369-372

FACT, 367

nucleotide excision repair, 363, 372-373

FA N C D 2 , 370

overview, 362

Fascioscapulohumeral muscular dystrophy (FSHD), 124, 458

D N A replication cellular organization, 354-355

FBI-1,229

genomic approaches to tim ing studies, 350-353

F E N - 1,349

in vivo studies, 349-350

Feulgen reaction, 8

overview, 347-349

Fibrillarin, 240

prospects for study, 355-356

Fluorescence recovery after photobleaching (FRAP), history o f nucleus studies, 8 - 9

DoalO, 176 Dosage compensation. See also X chromosome

FM R 1.457

F O S L2 , 126

Heh2, 176

Fragile X-associated tremor ataxia syndrome (FXTAS), 456-457 Fragile X syndrome, 457

H ES5, 126

Heterochromatin. Sec Chromatin Heterochromatin-binding protein-1 (HP1), 25, 64,

FRAP. See Fluorescence recovery after photoblcaching

139-140, 163, 258, 294, 368- 369, 424

FRD A. See Friedreich ataxia

HGPS. See H u tch in son -G ilford Progeria Syndrome

FRG 1,458

HIPk2, 254

Friedreich ataxia (FRD A), 458

H IR A , 258

FSHD. See Fascioscapulohumeral muscular dystrophy

Histone deacetylase (H D A C ), 123, 429

F X N , 458

Histone H I , 136, 143

FXTAS. Sec Fragile X-associated t remor ataxia syndrome

Histone H2A, 139-140, 163,165, 367, 371 Histone H 2 A X , 367-371, 373 Histone H3, 139, 163-165, 368, 430

С

Histone H4, 139, 163, 430

G A B T 4 ,457

Histone locus body (HI.B)

G A L 1 , 184-185

activity dependence, 319-320

G A L 2 , 184

amphibian oocytes, 243-244

Gall, loseph, 1, 3

biogenesis, 317

У 5, 122

discovery, 242-243

GAT4, 457

dynamics and assembly, 244 -245

G C N S , 423

functions, 242-243

G K M IN proteins, 453

maintenance, 318-319

Gene positioning

mode o f inheritance, 317, 325

chromatin dynamics, 123-124 chromatin interactions

prospects for study, 245 Historical perspective, nucleus studies

between distal genes, 117, 119-120

chromosome territories, 6 - 7

between regulatory elements, 117

composition, 5 - 6

disease defects, 124-126

discovery, 1-2

nucleoplasmic subcompartments

division, 8

Cajal body, 121

envelope, 2

centromeric heterochromatin regions, 122

internal dynamics, 8-10

lamina, 123

isolation, 3 - 5

P M L nuclear body, 121

matrix, 7 - 8

Polycomb bodies, 121-122 splicing speckles, 120-121

microscopy, 2 -3 , 8 H LB . See Histone locus body

overview, 115-118

H M G N 1,425

prospects for study, 126

H M L , 187

transcription sites, 120

H M R , 187

G le l. 81, 83-84

Homogeneously staining regions (HSRs), 145

G urdon, John, 210

Hom ologous recombination (FIR), regulation at rD N A locus, 188-189 HoxB, 202

H

Hoxb4-b9, 122

H 4 ,428

H P 1. See Hctcrochromatin-binding protein-1

H 19, 421

H R. See Hom ologous recombination

H a n d l, 217

HSBP1, 442

HAP, 293

HSF1, 292-296

Harris, Henry, 218

HSF2, 294-295

HAST, 181

H S PI0 4 , 184

H b a, 340

Hsp7<), 120

H b b , 340

HSRs. See Homogeneously staining regions

H b b l, 117

H tz l, 181, 187

H B P I, 439

H u tch in son -G ilford Progeria Syndrome (HGPS), 41, 124, 295, 403, 410-411, 429-430

H D A C . See Histone deacetylase HEAT-repeat fold, 79, 81

H u w e l, 168

H e h l, 176, 189

H X K I , 184-185

diseases, 64-65 ICR See Immunodeficiency centromcric instability facial anomalies IC N. See Interchromatin network

functional overview, 55-56 L E M dom ain proteins, 57-59 L IN C complexes, 59-61

IGAZ. See Interchromatin granule associated zone

P C N A , 63

IGC. See Interchromatin granule cluster

regulation o f signaling and differentiation, 61-63

Igf2, 421 IG H , 337

dynamics within lamina and nucleoplasm, 37-40

Ig H , 122-123

functions cell proliferation and differentiation, 45-46

IgK, 123

chromatin

I g L 122

epigenetic regulation, 43

Immunodeficiency centromcric instability facial

positioning and gene expression, 40

anomalies (ICF), 124

cytoskeleton, 46-47

ING1.43 I N O I , 184-185

D N A replication and repair, 41 -42

Interchromatin granule associated zone (IGAZ), 279,

interphase chromosome organization, 43-45

288 Interchromatin granule cluster (IGC), 226-228

mitosis, 23-24, 40-41

Interchromatin network (ICN ), model, 107

nuclear shape and stability, 40

Interphase chromosome, lamins in organization, 43-45

transcription, 42 isoforms and expression patterns, 33-34

Intronic splicing enhancer (ISE), 450

nuclear pore complex interactions, 85-86

I RIF, 369, 371-372

posttranslational processing and modification, 35-37

Iron, nuclear content, 5 ISE. See Intronic splicing enhancer Isolation, historical perspective o f nuclear isolation methods, 3 - 6

prospects for study, 47 structure and assembly, 34-35 Lamin A (1.MNA) mutations and disease, 401 -404, 409-410 processing, 409

)

Lamin В receptor (LBR), 18,64 -65, 404 -405

Jacobsen, Marty, 213

Lamin B1 ( L M N B 1 ), mutations and disease, 406

Ja rid lc , 168

Lamin B1 -associated dom ain (LAD ), 42

JMY, 202

Lam in B2 (LM N B 2), mutations and disease, 406 Lamina, gene positioning, 123 Lamina-associated polypeptide (LAP)

К

LAP1, 18,65

К А Р 1,368, 425

history o f study, 56-57

Kaps. See Karyopherins

LAP2, 18,62-64

Karyopherins (Kaps), types and cargoes, 78-80

proliferation control, 62-63

K A SH dom ain proteins, 59-61

Lampbrush chromosome (LBC), 147-148, 151

K H D R B S I , 457

Lantrunculin B, 202

Kid, 25

LAP. See Lamina-associated polypeptide

KIF5B, 23

LBC. See Lampbrush chromosome

Kinesin-1, 23

LBR. See Lam in В receptor

Kinctochore, microtubule interactions, 8

LCR. See Locus-control region

KSRP, 268-269

L E M dom ain proteins, 57-59, 62

Ku70, 171, 183

LE M D 3 . See M A N 1

Ku80, 171

Lim b-girdle muscular dystrophy, 64 L IN C complex, 59-61 LIN E, 454

L

LIN E1, 168-169

LAD. See Lam in B1-associated domain

L M N A . See Lamin A

Lamin

L M N B 1 . See Lamin B1

binding proteins. See also Lamina-associated polypeptide BAF, 57-59

L M N B 2 . See Lamin B2 Locus-control region (LCR), 116-117 L s g l, 81

M

Nonhom ologous end-joining (N H EI), 364, 370

M A L , 196

N O N O . See P54NRB

M A L A T l, 229, 285

N PC . See Nuclear pore complex

Male-specific lethal complex (MSI.), 160-161

nSB. See Nuclear stress body

M A N I, 18,64, 176-177, 189,405

Ntf2, 81

M A PP, 325

Nuclear bodies. See also Cajal body; Histone locus body;

M A R s. See Matrix attachment regions

Nuclear speckles; Nuclear stress body;

M a sh I, 123

Nucleolus; Paraspeckles; Perinucleolar compartment; PM1. nuclear body

M AT, 187 M atrix attachment regions (M AR s), 143

activity dependence and independence, 319-320

M B N L 1 , 455-457 M C M 2-7, 348, 355

biogenesis

M D C I , 371, 373

assembly pathways, 322

M e c l, 187

dc novo formation, 320-321

M eCP2, 140

physiological biogenesis, 326

Medea, 62

prospects for study, 326-327

Mel-28, 26

R N A role, 321-322

M ex67,80

self-organization role, 322-323

M G M T , 363

templates and nudeators, 320

M IG . See M ito tic interchromatin granule

dynamics, 316, 318

M iller, Oscar, 211

maintenance, 318-319

Mirsky, Alfred, 6

mitotic inheritance, 323-326

M ito tic interchromatin granule (M IG ), 230-231

orphan nuclear bodies

M L E , 160

clastosome, 304-307

M lp l, 176

Polycomb body, 307-310

M lp2, 176

prospccts for study, 312

M O F, 160

R N A -binding proteins, 311 -312

Mps3, 176, 181-182, 187-188

transcriptional regulator association, 310-311

M r e l l, 188, 253, 364 M R N , 253, 364-365, 370-371, 443 M RP, 269-271 MSI.. See Male-specific lethal complex M tr2, 80 M Y C , 126, 337

types, 304 types, 317 Nuclear envelope (NE) breakdown kinases, 21-22

c-M yc, mutation and disease, 423-424

membrane dynamics, 22-23

M y o D , 121

overview, 21

M yotonic dystrophy, 455

diseases emerin mutations, 404 lamin A mutations, 401 -404,409-410

N

lamin В receptor mutations, 404-405

NARs. See Nucleoporin-assodated regions

lamin B1 mutations, 406

NBS1. See M R N

lamin B2 mutations, 406

N C R Sec Nucleosomc core particle

M A N 1 mutations, 405

N d c l, 22-23, 27

nesprin-1 mutations, 405-406

NK. See Nuclear envelope

pathogenesis, 406-410

N E A T I , 279, 283-288, 297, 321, 322, 325

prospects for study, 4 1 0 -4 1 1

N E A T 2. See M A L A T -l

torsin A mutations, 406

NER. See Nucleotide excision repair

Z M P S T E 2 4 mutations, 406

NES. See Nuclear export sequence Nesprins overview, 46, 65 S Y N E I mutations and disease, 405-406

formation chromatin role, 25-26 reformation, 24-25 history o f study, 2

N IPP1, 226

protein types, 17-18

N15. See Nuclear localization sequence

remodeling

N M 1 . See Nuclear myosin I

dividing cells, 19-21

Nm d3, 81

interphase, 26-27

structure and function during mitosis, 23-24

Nucleolus

topology, 17,19

biogenesis, 317

yeast

compact structure, 215-216

D N A anchoring, 181-182

composition, 213-215

proteins, 176-177

de novo formation, 320-321

transcription and nuclear organization, 186

discovery, 209

Nuclear export sequence (NES), 77-78

dynamics, 215-216

Nuclear localization sequence (N'LS), 77

functions

N udear matrix, history o f study, 7 -8

cellular senescence, 218

Nuclear myosin 1 (N M 1), 197, 201-204

stem cell biolog)', 217

Nuclear pore complex (N P C )

transcription, 218-219

assembly, 85

viral replication, 217

dynamics. See Nuclear envelope

history- o f study, 209-213

functional overview, 73-74

maintenance, 318-319

karyopherins, 78-80

mode o f inheritance, 317, 323-324

lam in interactions, 85-86

pcrinucleolar compartment, 217

nucleoporin homologs, 75

P M L nuclear body, 254

phenylalanine-glycine translocation machinery,

prospects for study, 219-220

83-84 regulation o f translocation, 83-85 R N A export, 392-393

regulators, 216-217 territories, 218 )vast, 178-179

structure, 74-77

Nucleoporin-associated regions (NARs), 162

transport cycle, 77-82

Nucleosome, n udeosom e - n u cleosome

yeast

interactions in isolated chromatin

association o f persistent double-strand breaks,

fibers, 137

collapsed forks, and short telomeres,

Nucleosome core particle (N C P ), 134

187-188

Nucleotide excision repair (NF.R), 363,

overview, 176-177 recombination regulation at r D N A locus, 188-189 Nudear speckles

372-373 Nup42, 84 Nup53, 22, 26 Nup59, 26

biogenesis, 231-232, 317

Nup60, 176

cell cycle changes, 230-231

Nup84, 17, 187

composition, 228-229

Nup93, 22

discovery, 225

Nup98, 22

dynamics, 229-230

N u p l0 7 , 17, 22-23, 26

localization, 227-228

N u p l2 0 , 187

maintenance, 318-319

N u p l3 3 , 187

mode o f inheritance, 317,324

N u p l5 3 , 84 - 85

overview, 225-227

N u p l5 5 , 26

prospects for study, 232-233

N u p l5 7 , 27

structure, 227

N u p l5 9 , 84

Nudear stress body (nSB)

N u p l6 0 , 17, 22-23, 26

assembly on satellite III D N A , 293-294

N u p l7 0 , 17

biogenesis, 317

Nup205, 17

heterochromatin functions, 299

Nup358, 84

history o f study, 291 -293

N u r l, 189

maintenance, 318-319

N U R D , 420

mode o f inheritance, 317

N x tl, 80

molecular trap function, 296-298 nuclear organization impact, 299 prospects for study, 299-300

О

satellite III R N A function, 295-296

0ctl.310

stress response, 294-295

O P T domain, 304,310

transcriptional activation o f pericentromeric

Osteopoikilosis, 405

hctcrochromatin, 294

Otcfin, 62

PM L-IV, 257,259

P

P M L nuclear body

p i 6, 422 p53, 257, 371,442

activity independence, 320

P54NRB, 280-282, 321

alternative lengthening o f telomeres-associated P M L bodies, 253

PA D domain, 181 PAF, 391

biogenesis, 253, 317

PAP, 390-391

cytoplasmic bodies, 255

PA P 1,452

de novo formation, 321

Paraspecldes

dynamics

biogenesis, 317, 321, 325

basal status, 255

composition

cell cycle changes, 255

proteins, 280-282 R N A , 282-287

stress response, 252,255-256 functions, 256-259

discovery, 277-278

gene positioning, 121

diseases, 288

history o f study, 250-251

formation, 287

kinases, 254

function, 287-288

maintenance, 318-319

maintenance, 318-319

mode o f inheritance, 317, 325-326

mode o f inheritance, 317

overview, 249-250

prospects for study, 288

partner proteins

structure, 278-280 P C N A . See Proliferating cell nuclear antigen

degradation, 258 - 259 posttranslational modifications, 257-258 sequestration, 258

P d fl, 184

prospects for study, 259

Pdf2, 184 Pederson, Т., 213

proteasome component, 307

Penman, Sheldon, 211

proteins, 251

Perichromatin fibril (PF), 332-333

structure o f classical body, 252-253

Perichromatin region (PR), model, 106

subnudear compartments, 254-255

Perinucleolar compartment (PN C )

sumoylation, 251, 256, 309-310

biogenesis, 317

types, 250-252 P N C . See Perinucleolar compartment

cancer malignancy function, 274

POI32, 188

metastasis, 272-273

Poly(A) tail. m R N A , 390-392

therapeutic targeting, 273-274

Polvcomb body

transformation studies, 272 components proteins, 268-269 R N A . 269-270

gene positioning, 121-122 overview, 304, 307-309 sumoylation, 309-310 Polytene chromosome, 147-148

D N A association evidence, 271-272

P O M 121,26

formation, 270

Pom34, 26

maintenance, 318-319

Pom 152, 26

mode o f inheritance, 317

PR. See Perichromatin region

overview, 217, 267

PRC1, 308- 309

prospects for study, 274-275

PRC2, 308-309

structure, 268

Prim ary dystonia, 65

transcription role, 270-271

Proliferating cell nuclear antigen (PC N A ), 41, 63,349, 354, 363

PERIOD-1,281 PF. See Perichromatin fibril

Prostate cancer, gene positioning, 126

Phenylalanine-glycine, translocation machinery,

Proteasome, nuclear body component, 305-307

83-84

P R P F 3 ,452

Phosphatidvlinositol phosphate kinase (PIPK), 229

PR PF 3 1 ,452-453

P ifl, 188

PRPF8, 452

PIPK. See Phosphatidylinositol phosphate kinase

PSF, 201,280-281

PLK 1 .2 2

PSK1I1, 228

P M L , 428

PSP1, 199, 325

PML-1,251

PSPC1, 237, 277-278, 280-281

P tal, 391

transcription coupling

PTB , 268-269

capping, 385

PTEFb, 383

overview, 382-383

P T E N , 258

poly(A) tail, 390-391 splicing, 388-389

P I T, 310

RNase P, 269-270 RN F8, 373

R

Roberts syndrome (RS), 423

Rad 17, 365

RPA. Sec Replication protein A

Rad27, 188

RPABC2, 196

Rad50, 253, 364

RPABC3, 196

Rad51, 186, 188, 253

Rpd3L, 184

Rad52, 186-188, 253, 371

Rrp6. 391-392

Rad54, 371

R rp41,391

Rad55, 188

RS. See Roberts syndrome

Rad59, 188

Rttl09, 182

Rag, 122

Ran, 22, 26,79-81,84 R a p l, 180-181, 184

S

R A R a , 428

SAF-A, 200

RASSF1A, 422

Saf-B, 293

R a il, 391

SAGA, 184-185, 384

RBCC, 251

SAHF, 258

RBM 17

Sam68 body, 304, 311

RBM 4, 282

S A M D 4 A , 340

RCA. See Regulator o f chromosome archileclure

SANB, 255

rD N A , yeasl

SARs. See Scaffold-attachment regions

genes, 179

SATB1, 117, 167, 420-421

recombination regulation, 188-189

SATB2, 117, 167

Regulator o f chromosome architecture (RCA), 147 Replication factor С (RFC), 349, 363

Satellite III D N A nuclear stress body assembly, 293-294 transcript function, 295-296

Replication protein A (RPA), 348, 363, 365

SCA. See Spinocerebellar ataxia

R E T, 428

Scaffold-attachment regions (SARs), 145

R fal, 188

SCAN. See Spinocerebellar ataxia and neuropathy

RFC. See Replication factor С

scaRNA. See Small Cajal body-associated R N A

R ifl, 181

SFRS 1,458

R ift. 181

SFRS9, 458

R N A polymerase. See a b o Transcription

Sgsl, 188

actin association, 196-198

Silver-Russell syndrome (SRS), 422

gene positioning, 116-117, 119-120

SINE, 454

nuclear speckle association, 228

Sir2, 183

R N A processing coupling, 382-383, 385

Sir3, 181, 183

yeast, 179-180

Sir4, 181, 183

R N A processing capping, 384-385

SIRT1, 430 Slx5, 187-188

export, 390-391

Slx8, 187-188

overview, 381-382

S M A . See Spinal muscular atrophy

poly(A) tail, 390-392

Small Cajal body-associated R N A (scaRNA), 239-241

prospects for study, 393-395

Small nuclear ribonudeoprotein (snRN P), Cajal body

quality control poly(A) tail, 391-392 splicing, 389-390

association, 239 gene loci, 242 Small nuclear R N A (snRNA), 121, 128, 130, 203, 239,

splicing

242, 320, 328, 338, 449-452

diseases, 449-459

S M C proteins, 145

overview, 385-390

Smc5, 188

Smc6, 188

TLS. See Translesion synthesis

Smchd 1, 170

T M P R S S 2 , 126

S M N , 241-242, 245, 322, 437

T N F A IP 2 , 340

S M N 1,453

T O P B 1,254

S M N 2 , 453-454

T O R I A. See Torsin A

SNF, 424

Torsin A (TO R 1A ), mutations and disease, 406

snRN A. See Small nuclear R N A

T R A M P , 391

snRNP. See Small nuclear ribonucleoproiein

Transcription

SNRNP200, 452

actin function

SOP-2, 312

machinery association, 196-197

S o x l, 122

nascent transcript association, 197-201 ribonucleoproiein association, 201

SP100, 255, 325 SPB. See Spindle pole body

chromatin dynamics and transcription, 338-339

Spc42, 178 Speckles. See Nuclear speckles; Paraspeckles

factories, 332-336

Spinal muscular atrophy (SM A), 242, 437-440, 453

interactome and hotspots, 339-340

Spindle pole body (SPB), 181

lamin function, 42

Spinocerebellar ataxia (SCA) type 1,438-440

nucleolus role, 218-219 orphan nuclear body regulation, 310-311

type 3,458

perinucleolar compartment role, 270-271

type 17,440-442

polymerization regulation, 201-202

Spinocerebellar ataxia and neuropathy (SCAN), 443 Splicing, m R N A diseases, 449-459 overv iew, 385-390

prospects for study, 341-342 R N A processing. See R N A processing shared R N A polymerase II sites, 336 - 338 Transfer R N A (tRN A), yeast genes, 179-180

Srcl, 176

Translesion synthesis (TLS), 365

SRF.BP 1,408

TRF.X, 381, 384, 388, 391, 393

SRF, 196

TR EX-2, 384, 393

SRS. See Silver-Russell syndrome

T R F I.2 5 3

Srs2, 188

TR F2, 253

SSB, 348

T R IM , 251

Ssu72, 391

tRNA. See Transfer R N A

STRAP, 453 S u b l, 391 S U M O , 81, 84, 187-188, 251, 256, 303, 309-310, 325, 439

U UBC13, 373

S U M O body, 304

Ubc9, 84

S U N dom ain proteins, 59-61

Uros, 117

Susl, 184-185 SWI, 424 S Y B L I , 125

SYNE1. See Nesprins

V VEGF, 126 V IN C - 1 . 287

T TAT-SF1, 388

W

TBP, 440-442

Wed, 218

TD P1.443

W R N , 254

Tdt, 122

VVT1, 282

Telomere, yeast, 180-181, 187-188

YVTX, 282

TFIIB, 391 TFIID , 391, 440 -442 TF1IH, 363, 366, 372 T H O , 381,388, 391 T ic l, 181-180 Tip60, 439

X X chromosome. See also Dosage compensation heterogeneous heterochromatin in inactive X chromosome, 163-165

nuclear organization o f inactive X chromosome in

genome s

female mammals, 162-163 prospects fo r study, 170

ind nuclear organization

DNA

« 6 - 187

nude.'

jm plcx association o f persistent

spreading inactivation, 168-170

strand breaks, collapsed forks,

Xist in inactivation, 165-167

•rt tdomcres, 187-188

Xic. See X inactivation center

recombiiw..>on regulation at rD N A locus,

X inactivation center (Xic), 165-166

188-189

Xist, 163, 165-167

nuclear envelope proteins, 176-177

XPB, 373

nuclear pore complex, 176-177

X P C , 363,372-373

nucleolus, 178-179

ХРГ, 363, 366, 372

prospects for study, 189

XPG , 372

telomere, 180-181

X R C C 1, 363, 365, 443

transcription and nuclear organization

X R C C 4, 443

gene silencing, 183-184

X r n l, 391

inducible gene expression, 184-186 nuclear envelope function, 186 transfer R N A genes, 179-180

Y

Y o p l, 27

Yeast nucleus, budding yeast

YT-521, 237

chromatin dynamics, 177-178 chromosome folding, 177 compartmentation mechanisms

Z

D N A anchoring at nuclear envelope, 181-182

Zemike, Frits, 209

trans-association o f chromatin loci, 182-183

Z M P S T E 2 4 , mutation and disease, 406, 409-410