Cilia: Model Organisms and Intraflagellar Transport, Volume 93 (Methods in Cell Biology)

Series Editors Leslie Wilson Department of Molecular, Cellular and Developmental Biology University of California Santa...

Author: Stephen M. King | Gregory J Pazour

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Series Editors Leslie Wilson Department of Molecular, Cellular and Developmental Biology University of California Santa Barbara, California

Paul Matsudaira Department of Biological Sciences National University of Singapore Singapore

Methods in Cell Biology VOLUME 93 Cilia: Model Organisms and Intraflagellar Transport

Edited by

Stephen M. King Department of Molecular, Microbial and Structural Biology University of Connecticut Health Center Farmington, Connecticut

Gregory J. Pazour Program in Molecular Medicine University of Massachusetts Medical School Worcester, Massachusetts

AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

Academic Press is an imprint of Elsevier 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 32, Jamestown Road, London NW1 7BY, UK Linacre House, Jordan Hill, Oxford OX2 8DP, UK First edition 2009 Copyright © 2009 Elsevier Inc. All rights reserved No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made ISBN–13: 978-0-12-381377-0 ISSN: 0091-679X For information on all Academic Press publications visit our website at elsevierdirect.com

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CONTRIBUTORS

Numbers in parentheses indicate the pages on which the authors’ contributions begin.

Seyda Acar (235), Molecular and Cell Biology, University of California, Davis, California 95616 Philippe Bastin (59), Trypanosome Cell Biology Unit, Institut Pasteur and CNRS, Paris, France Robert H. Behal (179), Department of Microbiology, Molecular Biology and ?Biochemistry and the Center for Reproductive Biology, LSS142, University of Idaho, Moscow, Idaho 83844-3052 Joseph C. Besharse (219), Department of Cell Biology, Neurobiology and Anatomy, Medical College of Wisconsin, Milwaukee, Wisconsin 53226-0509 Tatiana Y. Besschetnova (331), Renal Division, Department of Medicine, Brigham and Women’s Hospital and Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115 Ewelina Betleja (179), Department of Microbiology, Molecular Biology and ?Biochemistry and the Center for Reproductive Biology, LSS142, University of Idaho, Moscow, Idaho 83844-3052 Oliver E. Blacque (267), School of Biomolecular and Biomedical Science, UCD Conway Institute, University College Dublin, Belfield, Dublin 4, Ireland Heriberto Cerutti (99), School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska, Lincoln, Nebraska 68588 Douglas G. Cole (179), Department of Microbiology, Molecular Biology and ? Biochemistry and the Center for Reproductive Biology, LSS142, University of Idaho, Moscow, Idaho 83844-3052 Drashti Dave (1), Department of Cellular Biology, University of Georgia, Athens, Georgia 30605 William Dentler (145), Department of Molecular Biosciences, University of Kansas, Lawrence, Kansas Dennis Diener (111), Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, Connecticut 06520 Iain Drummond (197), Departments of Medicine and Genetics, Harvard Medical School and Massachusetts General Hospital, Charlestown, Massachusetts 02129 Susan K. Dutcher (121), Department of Genetics, Washington University School of Medicine, St. Louis, Missouri 63110 Jonathan T. Eggenschwiler (347), Department of Molecular Biology, Princeton University, Lewis Thomas Labs, Princeton, New Jersey 08544 Benjamin D. Engel (157), Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, California 94158 xi

xii

Contributors

James Evans (235), Molecular and Cell Biology, University of California, Davis, California 95616 Jacek Gaertig (1), Department of Cellular Biology, University of Georgia, Athens, Georgia 30605 Limin Hao (235), Molecular and Cell Biology, University of California, Davis, California 95616 Kent L. Hill (21), Department of Microbiology, Immunology, and Molecular Genetics, University of California Los Angeles, Los Angeles, CA-90095, United States of America, and Molecular Biology Institute, University of California Los Angeles, Los Angeles, CA-90095, United States of America Mitsuo Ikebe (157) Department of Physiology, University of Physiology, University of Massachusetts Medical School, Worcester, Massachusetts 01655 Peter N. Inglis (267), Department of Molecular Biology and Biochemistry, Simon Fraser University, 8888 University Drive, Burnaby, BC V5A 1S6, Canada Christine Insinna (219), Department of Cell Biology, Neurobiology and Anatomy, Medical College of Wisconsin, Milwaukee, Wisconsin 53226-0509 Carlo Iomini (121), Department of Developmental and Regenerative Biology, Mt. Sinai School of Medicine, New York, New York 10023 Daria Julkowska (59), Trypanosome Cell Biology Unit, Institut Pasteur and CNRS, Paris, France Robert A. Kesterson (305), Department of Genetics, University of Alabama at Birmingham, Birmingham, Alabama 35294 Eun-Jeong Kim (99), School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska, Lincoln, Nebraska 68588 Stephen M. King (81), Department of Molecular, Microbial & Structural Biology, University of Connecticut Health Center, Farmington, Connecticut 06030-3305 Hyuk W. Ko (347), Department of Molecular Biology, Princeton University, Lewis Thomas Labs, Princeton, New Jersey 08544 Karl-Ferdinand Lechtreck (157), Department of Cell Biology, University of Massachusetts Medical School, Worcester, Massachusetts 01655 Michel R. Leroux (267), Department of Molecular Biology and Biochemistry, Simon Fraser University, 8888 University Drive, Burnaby, BC V5A 1S6, Canada Brian A. Link (219), Department of Cell Biology, Neurobiology and Anatomy, Medical College of Wisconsin, Milwaukee, Wisconsin 53226-0509 Aimin Liu (347), Department of Biology, Eberly College of Science, The Pennsylvania State University, 201 Life Science Building, University Park, Pennsylvania 16802 Miguel A. Lopez (21), Department of Microbiology, Immunology, and Molecular Genetics, University of California Los Angeles, Los Angeles, CA-90095, United States of America Katherine Luby-Phelps (219), Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, Texas 75390-9039 Wallace F. Marshall (157), Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, California 94158

Contributors

xiii Amber K. O’Connor (305), Department of Cell Biology, University of Alabama at Birmingham, Birmingham, Alabama 35294 Michael Oberholzer (21), Department of Microbiology, Immunology, and Molecular Genetics, University of California Los Angeles, Los Angeles, CA-90095, United States of America Guangshuo Ou (235), Department of Cellular and Molecular Pharmacology, University of California, San Francisco, California 94107 Ramila S. Patel-King (81), Department of Molecular, Microbial & Structural Biology, University of Connecticut Health Center, Farmington, Connecticut 06030-3305 Mary E Porter (145), Department of Genetics, Cell Biology, and Development, University of Minnesota, Minneapolis, Minnesota 55455 Katherine S. Ralston* (21), Department of Microbiology, Immunology, and Molecular Genetics, University of California Los Angeles, Los Angeles, CA-90095, United States of America. Panteleimon Rompolas (81), Department of Molecular, Microbial & Structural Biology, University of Connecticut Health Center, Farmington, Connecticut 06030-3305 Barnali Roy (331), Renal Division, Department of Medicine, Brigham and Women’s Hospital and Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115 Tsuyoshi Sakai (157), Department of Physiology, University of Massachusetts Medical School, Worcester, Massachusetts 01655 Jonathan M. Scholey (235), Molecular and Cell Biology, University of California, Davis, California 95616 Jagesh V. Shah (331), Renal Division, Department of Medicine, Brigham and Women’s Hospital and Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115 Jacob E. Till (121), Department of Genetics, Washington University School of Medicine, St. Louis, Missouri 63110 Kristyn VanderWaal (145), Department of Genetics, Cell Biology, and Development, University of Minnesota, Minneapolis, Minnesota George B. Witman (157), Department of Cell Biology, University of Massachusetts Medical School, Worcester, Massachusetts 01655 Dorota Wloga (1), Department of Cellular Biology, University of Georgia, Athens, Georgia 30605 Bradley K. Yoder (305), Department of Cell Biology, University of Alabama at Birmingham, Birmingham, Alabama 35294

*

Current address Department of Medicine, Division of Infectious Diseases and International Health, University of Virginia Health Sciences Center, Charlottesville, VA-22908, United States of America.

PREFACE

Cilia and flagella have long been the subject of intense study and a previous volume of Methods in Cell Biology dedicated to this organelle was published in 1995. However, in the 15 years since that publication, interest in the organelle has dramatically increased as it has come to be appreciated that these tiny structures play fundamental roles in the development and health of mammals and are vital for vertebrates to perceive their environment and respond to it. In humans the list of ciliary diseases, or ciliopathies, has grown tremendously since the publication of the previous volume. In 1995 the field recognized that cilia and flagella played critical roles in male fertility and respiratory disease and were recognized as being important in the determination of left–right asymmetry of vertebrates but the mechanism was not known. In addition, it was known that the senses of vision and smell depended on receptors localized to modified cilia. It is now appreciated that ciliary defects underlie a wide range of human diseases. These include polycystic kidney disease (PKD), nephronophthisis, Bardet– Biedl syndrome (BBS), Meckel–Gruber syndrome, Joubert syndrome, Jeune syndrome, and short rib-polydactyly syndrome that are thought to result from defects in primary cilia. Other diseases such as male infertility, hydrocephaly, juvenile myoclonic epilepsy, primary ciliary dyskinesia, Kartagener’s syndrome, and left-right asymmetry defects of the heart are thought to result from defects in motile cilia. In addition, anosmia and blindness can derive from dysfunction of the highly specialized sensory cilia of the olfactory epithelium and retina. It is clear from studies in mouse that this collection of diseases is just the tip of the iceberg for ciliary disorders of man. Eukaryotic cilia and flagella are complex organelles composed of hundreds of different proteins. This complexity likely reflects the diverse motility and sensory roles played by these organelles. The motility functions of cilia have long been recognized and in mammals these are important for moving mucus in the lungs, moving cerebrospinal fluid in the brain, and propelling the male gametes. The sensory functions are less well known but include roles in olfaction in the nose and light detection in the eye. In addition, nearly every cell type in vertebrate organisms is ciliated by nonmotile primary cilia that are thought to sense the extracellular environment. The proteins of the cilium are organized around a microtubule-based cytoskeleton termed the axoneme and a specialized domain of the plasma membrane that covers the axoneme. The ciliary membrane is contiguous with the plasma membrane of the cell but is a separate domain containing a unique set of proteins, many of which play roles in sensory perception. The axonemes of motile cilia typically have a 9 þ 2 arrangement of microtubules while nonmotile sensory and primary cilia typically have a 9 þ 0 arrangement. These microtubules serve as scaffolding to bind and organize the multitude of proteins needed to carry out the motility and sensory functions of cilia. The microtubules of the axoneme are templated from a centriole at xv

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the center of the centrosome. When the cell is ciliated, the centriole (which is now called a basal body) and centrosome remain at the base of the cilium. The centrosome is best known for its role in organizing the cytoskeleton and also is postulated to be an important control center of the cell, integrating signals that regulate morphology, migration, and proliferation. With the explosion of interest in cilia, the model organisms available to study cilia and flagella have grown much more diverse, and the techniques available for assessing cilia structure and function have become more sophisticated. In these three volumes, we have asked top researchers in the field to provide methods used in their laboratories to study cilia and flagella. Cilia: Structure and Motility, Volume 91, focuses on general methods to study these organelles covering microscopic techniques for both structural analysis and detailing motility parameters, as well as biochemical approaches to define protein–protein associations and complexes. Cilia: Motors and Regulation, Volume 92, focuses on techniques for studying dynein structure and function and the varied mechanisms by which these motor complexes are regulated. Cilia: Model Organisms and Intraflagellar Transport, Volume 93, focuses on the methods for studying intraflagellar transport which is required for assembly of the organelle and provides general approaches for studying this and other cilia-related phenomena in all of the major model organisms that are currently being used to study cilia and flagella.

CHAPTER 1

Manipulating Ciliary Protein-Encoding Genes in Tetrahymena thermophila Drashti Dave, Dorota Wloga, and Jacek Gaertig Department of Cellular Biology, University of Georgia, Athens, Georgia 30602

Abstract I. Introduction II. Designing the Targeting Fragment for Gene Disruption III. Biolistic Transformation Protocol (Germline Gene Knockout) A. Reagents and Materials for Biolistic Transformation B. Digestion and Purification of Targeting Fragment DNA C. Preparation of Tetrahymena Cells for Biolistic Transformation D. Biolistic Transformation IV. Construction of Gene Knockout Heterokaryons A. Identification of Putative Germline Transformants (G1) B. Sexual Maturation and Assortment of Drug Resistance from the Macronucleus of G1 Transformants C. Testing for the Germline Presence of the Disrupted Allele D. Round I Genomic Exclusion Cross to Produce G2 Synclones E. Isolation of Heterokaryon Cells with Different Mating Types Among G2 Synclones F. Testing the Putative Heterokaryons and Bringing the Gene Disruption Phenotype to Expression by a Heterokaryon Heterokaryon Cross V. Disruption of Multiple Genes in the Germline A. Disruption of Multiple Genes with Different Selectable Markers B. Disruption of Multiple Genes with a Single Selectable Marker VI. Somatic (Macronuclear) Gene Knockout VII. Phenotypic Rescue by Introduction of a Wild-type Gene into a Knockout Strain A. Rescue of Mating Heterokaryons B. Rescue of Vegetative Paralyzed Knockout Cells Acknowledgments References

METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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Abstract Tetrahymena thermophila has emerged as an excellent protist model for studies on cilia that are based on reverse genetic approaches. In Tetrahymena, genes can be routinely disrupted by the DNA homologous recombination. We present established protocols for the manipulation of genes in either the germline micronucleus or the somatic macronucleus. A detailed protocol is provided for the construction of heterokaryon strains that carry a gene disruption only in the micronucleus. Heterokaryon strain can be propagated like wild-type cells, and ciliary phenotypes can be expressed on demand by mating. We describe methods that can be used for disruption of multiple genes. We include protocols for the generation and maintenance of Tetrahymena cells that either lack cilia or have paralyzed cilia.

I. Introduction Tetrahymena is a relatively large cell, covered with about 500 cilia (in the G1 phase of the cell cycle) that are required for cell motility, phagocytosis, and mating. The majority of cilia are arranged in about 20 longitudinal rows that generate cell motility, while about 100 cilia form oral membranelles that sweep food into the oral cavity. Tetrahymena is an established genetic model that is particularly suitable for reverse genetic approaches, based on its exclusively homologous DNA recombination activity. Tetrahymena has two distinct nuclei: the germline diploid micronucleus and the somatic polyploid macronucleus. The vegetative phenotype is controlled by transcription of genes in the macronucleus, while the micronuclear genome serves as a transcriptionally silent germline. During the vegetative life cycle, Tetrahymena cells divide by binary fission. The micronucleus divides by mitosis while the macronucleus divides by amitosis, a nuclear fission that involves random segregation of homologous chromosomes to daughter nuclei. Two starved Tetrahymena cells with different mating types can undergo conjugation, a developmental process during which the micronucleus undergoes meiosis, a postmeiotic mitosis that produces two pronuclei, reciprocal exchange and fusion of pronuclei, and two mitotic divisions of the zygotic micronucleus. The micronuclei that are products of divisions of the zygotic micronucleus differentiate into new macronuclei and micronuclei while the old macronucleus is degraded (Fig. 1). For a review of Tetrahymena as a cell biological model see Frankel (2000). The genome of Tetrahymena encodes a large number of conserved ciliary proteins (Eisen et al., 2006). Proteomes of cilia (Smith et al., 2005) and basal bodies (Kilburn et al., 2007) have recently been examined. Using the exclusively homologous DNA recombination activity, genes can be routinely disrupted either in the micronucleus (germline gene knockout) (Cassidy-Hanley et al., 1997; Hai and Gorovsky, 1997) or in the macronucleus (somatic gene knockout) (Shen et al., 1995). A number of ciliary mutants defective in either assembly or motility of cilia have been constructed by gene

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1. Manipulating Ciliary Protein-Encoding Genes

(A)

(B)

(C)

(F)

(E)

(D)

(J) (G)

(H)

(I)

Phenotypic assortment

Fig. 1 A scheme illustrating a conjugation between Tetrahymena thermophila strains having different alleles. (A–C) Pairing and meiosis. (D) Postmeiotic mitosis. (E–F) Reciprocal exchange and fusion of pronuclei. (G) Postzygotic divisions. (H) Differentiation of new micronuclei and macronuclei and degradation of the old macronucleus. (I) After pair separation and the first vegetative life cycle, the progeny cells are heterozygous in both the micronucleus and the macronucleus. (J) During the subsequent 60–80 generations, the macronuclei become homozygous for one of the two alleles by phenotypic assortment.

disruptions (Angus et al., 2001; Beales et al., 2007; Brown et al., 1999b, 2003; Hennessey et al., 2002; Liu et al., 2004; Rajagopalan et al., 2009; Sharma et al., 2007; Tsao and Gorovsky, 2008a,b; Williams et al., 2006; Wood et al., 2007). To evaluate the function of a gene, often it is sufficient to disrupt its locus in the macronucleus. A somatic (macronuclear) gene knockout is a straightforward procedure. The macronucleus has about 45 copies of each protein-coding gene in the G1 phase of the cell cycle. The elimination of all gene copies is possible by taking advantage of the phenomenon of phenotypic assortment (Orias and Flacks, 1975). During amitosis, chromosomes are segregated randomly to daughter nuclei. As a result, an initially heterozygous macronucleus, after 60–80 generations, becomes homozygous for one of the two alleles (Doerder et al., 1992). In a somatic gene knockout experiment, initially one or a few copies are replaced by a disrupted drugresistant allele using DNA homologous recombination. Next, replacement of the remaining wild-type copies is promoted by the propagation of transformants under increasing selective drug pressure that favors retention of the disrupted allele.

Gene disruption (A)

(B)

(C)

(F)

(E)

(D)

(G)

(H)

(I) Phenotypic assortment

(L)

(K)

(J)

(M)

(N)

(O)

(Q)

(P)

Star strain

Heterokaryons

Gene knockout cells (R)

Fig. 2 A scheme illustrating construction of knockout heterokaryons. (A–H) Construction of heterozygous G1 transformants by biolistic bombardment of mating cells. (I) G1 transformants that have lost the drug-resistant allele from the macronucleus are obtained by phenotypic assortment. (J–N) A cross of an assorted G1 transformant to a star strain (round I genomic exclusion) yields G2 cells (heterokaryons) that are homozygous for the disrupted locus in the micronucleus. (O–R) A standard conjugation between two heterokaryons produces progeny that are homozygous for the disrupted allele both in the micronucleus and in the macronucleus.

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The germline gene disruption approach, while more time consuming, is preferred for studies of essential genes, such as those encoding important basal body components (Culver et al., 2009; Shang et al., 2002a) or ciliary proteins that also have essential cell body functions (Sharma et al., 2007) or when multiple gene disruptions need to be combined. To introduce a gene into the micronucleus, a targeting fragment is transfected into mating cells, at the time when the micronucleus undergoes meiotic prophase (Cassidy-Hanley et al., 1997). Using specialized crosses, heterokaryon strains are constructed that are homozygous for a disrupted gene in the micronucleus and have wild-type alleles in the macronucleus (Hai et al., 1999). The heterokaryons are propagated vegetatively as wild-type cells, and the gene disruption phenotype is expressed by mating that produces progeny cells with a new macronucleus (Fig. 2).

II. Designing the Targeting Fragment for Gene Disruption To prepare a targeting plasmid, two fragments of the targeted locus are subcloned on either side of a selectable marker cassette. In the past we have used the neo2 cassette that expresses the neomycin phosphotransferase coding region between the HHF4 (histone H4) gene promoter and the BTU2 transcriptional terminator (Gaertig et al., 1994a). In transformed Tetrahymena cells, neo2 produces a relatively low level of resistance to paromomycin (pm). If the goal is to produce a germline gene disruption, the use of neo2 reduces the background of somatic transformants. A modified cassette that utilizes a cadmium-inducible MTT1 promoter and produces a robust resistance to pm (neo3) is available (Shang et al., 2002b). A similar neo4 cassette has the coding sequence that is codon-optimized for Tetrahymena (Mochizuki, 2008). The bsr cassette expresses the blasticidin S (bs) resistance gene (Brown et al., 1999b). Most recently, a new marker cassette was developed by J. Bowen and M. Gorovsky (University of Rochester), rpl29-K40M, based on the coding region of RPL29 that confers resistance to cycloheximide (cy) (Yao and Yao, 1991) and is expressed under the MTT1 promoter (J. Bowen and M. Gorovsky, personal communication). The frequency of DNA homologous recombination decreases with the increased size of the targeted region. Typically for germline gene disruption, we attempt to delete the entire coding region only if it has the size of 1 kb or less. Most often, the targeted part is limited to a portion of the coding region that includes an important functional domain or the first few exons including the start codon. Larger fragments can be successfully deleted using a somatic approach. In our hands, for a germline gene disruption, the flanking homology fragments should be at least 3 kb of total length (typically 3–4 kb, 1.5–2 on each side). We place the selectable cassette in a reverse transcriptional orientation in reference to the target locus, with the intention of avoiding transcriptional occlusion by the promoter of the targeted gene. During construction of the targeting fragment, after transformation, we incubate Escherichia coli at lower temperatures (30°C instead of standard 37°C) to improve stability of AT-rich noncoding sequences.

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III. Biolistic Transformation Protocol (Germline Gene Knockout) The PDS1000/He biolistic microparticle delivery system (BioRad, Hercules, CA) is assembled and used with strict adherence to the manufacturer’s instructions. PDS1000/ He is connected to a vacuum pump (Precision D150) and a tank with ultrahigh quality pressurized helium. The protocols for biolistic transformation and heterokaryon construction are based on the original methods published by Hai et al. (1999) and Bruns and Cassidy-Hanley (2000a). Researchers not familiar with the cultivation and genetics of Tetrahymena may consult additional sources (Bruns and Cassidy-Hanley, 2000b; Hamilton and Orias, 2000; Orias et al., 2000). A. Reagents and Materials for Biolistic Transformation

1. Microcarrier Suspension 1. Add 1 ml of 100% ethanol to the original tube with 0.6 nm gold microcarriers (250 mg, BioRad #1652262) and vortex on high setting for 3–4 min. 2. Immediately distribute the suspension into four 1.5-ml Eppendorf tubes (250 µl per tube). 3. Add 750 µl of 100% ethanol per tube and vortex for 1 min on the highest setting. 4. Spin tubes in a microcentrifuge (10,000 g, 1 min) and remove the supernatant (should be clear). 5. Add 1 ml of 100% ethanol per tube and repeat step 4 ( twice). 6. Add 1 ml of sterile water per tube, vortex for 3 min, centrifuge as above, and discard the supernatant (repeat twice). 7. Suspend the pellets in 1 ml of 50% sterile glycerol per tube; make 100 µl aliquots and store at –20°C.

2. Other Reagents and Materials Needed for Biolistic Transformation 1. Spermidine (Sigma-Aldrich, St. Louis, MO, S0266). Dilute to 0.1 M. Filter sterilize, make 1 ml aliquots, and store at –20°C. 2. 2.5 M CaCl2 (50 ml; filter sterilize), 70% ethanol (50 ml), and 100% ethanol (50 ml)—keep all refrigerated. 3. SPP medium (Gorovsky, 1973): 1% proteose peptone, 0.2% glucose, 0.1% yeast extract, and 0.003% EDTA ferric sodium salt. 4. MEPP medium (Orias and Rasmussen, 1976). Prepare 1 l of 2% proteose peptone (autoclave), 2.5 mg/ml folinic acid (25 mg in 10 ml, filter sterilize), 0.2 M sodium citrate (3 g in 50 ml, filter sterilize), and 0.1 M FeCl3, 3 mM CuSO4 (1.35 g of FeCl3 and 22 mg of CuS04 in 50 ml, filter sterilize). Make MEPPA shortly before use by mixing the solutions in the following order (scale up as needed): 50 ml of 2% proteose peptone, 20 µl of folinic acid stock, 0.5 ml of sodium citrate stock, 0.5 ml of FeCl3 and CuSO4 stock, and 1 ml of an antibiotic/antimycotic mix.

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5. 10 mM Tris–HCl, pH 7.5, autoclaved. 6. Antibiotic/antimycotic mix (Invitrogen, Carlsbad, CA, #15240–104). Use at 1:100 dilution with SPP (SPPA) and 1:50 with MEPP (MEPPA). 7. Selective drug stock solutions (all stored in aliquots at –20°C): pm (Sigma, P9297) 100 mg/ml in water, 6-methylpurine (mp) (Sigma M6502) 15 mg/ml in water, cy (Sigma, C7690) 15 mg/ml in ethanol, bs (Invivogen, San Diego, CA, ant-bl-1) 10 mg/ml in water, and CdCl2 5 mg/ml in water. 8. Biolistic supplies (BioRad): rupture discs 900 psi, stopping screens, macrocarriers, and macrocarrier holders. 9. Circular 9 cm Whatman #1 filters (sterilize by autoclaving in a glass Petri dish).

B. Digestion and Purification of Targeting Fragment DNA 1. Digest 80 µg of plasmid DNA (purified with the Qiagen kit 12143) with restriction enzymes (800 µl total volume) to separate the targeting fragment from the plasmid backbone (there is no need to gel purify). 2. Clean the digested DNA using the QIAquick gel purification kit (Qiagen, Valencia, CA 28706) according to the instructions for “purification of a PCR product,” using PB binding buffer followed by two washes with PE buffer. Elute the digested DNA in 25 µl of water. We clean DNA in four 200 µl (20 µg of DNA) aliquots and elute each aliquot in 25 µl of water.

C. Preparation of Tetrahymena Cells for Biolistic Transformation Strains are available from the Tetrahymena Stock Center (http://tetrahymena.vet. cornell.edu). 1. Start 100 ml cultures of CU428 (mpr1-1/mpr1-1, mp-s, mt VII) and B2086 (þ/þ, mt II) strains in SPPA medium with an inoculum from feeder cultures to obtain populations in the mid log phase (~2–3 105 cells/ml) in the morning on the next day. Grow at 30°C with shaking at 160 rpm. The generation time is ~3 h for CU428 and ~4 h for B2086 strains. 2. Next morning, centrifuge cells in four 50-ml conical centrifuge tubes at ~1100 g, discard the supernatant, suspend each pellet in 50 ml of 10 mM Tris–HCl (pH 7.5), and centrifuge again. Discard the supernatant and suspend each strain in 100 ml of 10 mM Tris–HCl (pH 7.5). Add 1 ml of the antibiotic/antimycotic mix and incubate (without shaking) at 30°C overnight (18–24 h) in 500-ml Erlenmayer flasks. 3. Next day in the morning, adjust the cell concentration of each strain to 2 105 cells/ ml in 100 ml. 4. Our experience is that the most optimal periods for germline targeting are between 2–2.5 and 3.5–4 h after strain mixing. To obtain mating cells within these two periods, mix 25 ml of starved B2086 with 25 ml of starved CU428 cells, place in a 0.5 to 1 l sterile Erlenmayer flask, and incubate without shaking at 30°C. Mix

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additional (25 þ 25 ml) volumes of starved cells (and incubate in separate flasks) 30, 90, and 120 min later. 5. Incubate mating cells at 30°C for 2 h (after mixing the last sample of cells). The above time points are used for a typical sample that gives 80% of cells in pairs 1 h after mixing. If there is a delay in pairing, change the incubation time accordingly.

D. Biolistic Transformation

1. Preparation of PDS1000/He and Culture Media 1. Clean the laminar flow hood surface, the inside surfaces of the PDS1000/He chamber, and the removable parts: target platform, rupture disc retaining cap (RDRC), and macrocarrier assembly (MA) with 70% ethanol. 2. Place the macrocarrier holders, catching screens, 900-psi rupture discs, and macrocarriers (1 of each per shot, 4 total) in 70% ethanol inside covered 10-cm Petri dishes. Two hours before shooting remove all items (except rupture disks) and let dry inside the hood under UV. 3. Prepare four 250 ml Erlenmeyer flasks with 50 ml of 10 mM Tris-HCl, pH 7.5 with antibiotic/antimycotic mix at 1:100. Place the flasks inside the laminar flow hood.

2. Coating Gold Microcarriers with DNA The following procedure applies to one sample of DNA per shot. Prepare four samples simultaneously. Particles are handled in a cold room until ready for transfer onto macrocarriers. 1. Add 25 µl of digested DNA (20 µg) to a 0.5-ml Eppendorf tube with a 100-µl aliquot of suspended 0.6-µm gold microcarriers. Vortex at full speed for 10 s. 2. Add 125 µl of 2.5 M CaCl2 and vortex for 10 s. 3. Add 65 µl of 0.1 M spermidine and vortex for 10 min (on a tube platform). 4. Place a macrocarrier inside a macrocarrier holder and keep inside the laminar flow hood until coated microcarriers are ready. 5. Briefly centrifuge the tube with DNA/microcarriers (1200 g, ~3500 rpm in a microcentrifuge) and remove the supernatant. 6. Add 200 µl of 70% ethanol, suspend by pipetting, and vortex for 1 min. Pulsecentrifuge and remove the supernatant. 7. Repeat step 6 with 100% ethanol. 8. Add 15 µl of 100% ethanol to the pellet. Place the tube inside a water bath sonicator (Branson 1210) for ~20 s. 9. Immediately suspend the pellet by pipetting and spread the content in the center of the macrocarrier (the particles should form a ~1 cm ring). 10. Leave the loaded macrocarrier/holder in a desiccator (a small plastic box with bottom covered with a layer of silica gel desiccant) until ready to shoot.

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3. Starting the Biolistic Gun PDS1000/He 1. Open the main gas valve on the helium tank by turning counterclockwise until the right gauge on the helium pressure regulator (HPR) shows 2000 psi. 2. Turn the adjustment screw of the HPR clockwise until the left gauge on the HPR shows 1100 psi. 3. Turn the vacuum pump and PDS1000/He on. 4. Using forceps, load a 900-psi rupture disc (taken from 70% ethanol) into the bottom of the RDRC (ensure a proper fit). Screw the loaded RDRC onto the end of the acceleration tube inside the gun chamber. Tighten the RDRC to proper torque using the torque wrench that came with PDS1000/He. 5. Load a stopping screen into the fixed nest of the MA (inside the fixed nest, the stopping screen support is placed at the bottom position so that the macrocarrier flying distance during the shot is 16 mm). 6. Invert and place the loaded macrocarrier holder over the stopping screen inside the MA. Screw on the MA cover lid. Insert the MA into the second groove from the top within the gun chamber (the “gap distance” between the MA cover and the bottom of RDRC should be 1/40). 7. Place a sterile 9-cm circular Whatman #1 filter inside the cover of a 10-cm Petri plate. Wet the filter with 1 ml of 10 mM Tris–HCl, pH 7.5 (make sure that the filter is flat and without air bubbles). Place the Petri plate cover with the filter inside the target platform.

4. Concentration of Cells Immediately Before Shooting After the gun has been set up precisely at 2 h after strain mixing, combine all mating cells in a 1-l Erlenmayer flask. Prepare cells in 50-ml portions before each shot (see below).

5. Biolistic Shooting 1. Centrifuge 50 ml of pooled mating cells in a 50 ml conical centrifuge tube for 3 min at 1100 g. 2. Gently suspend the pellet in 0.5 ml of 10 mM Tris–HCl (pH 7.5) and transfer cells onto the center of the wet filter paper. 3. Insert the target platform with cells into the second groove from the bottom inside the gun chamber (12 cm target distance) and close the door. 4. Press the “vacuum” switch until the gauge on top of the gun reads 27 mmHg and move the switch to the “hold” position. 5. Press and hold the “fire” switch—a popping noise will be heard when the pressure reaches ~900 psi and the rupture disc breaks. 6. Vent the vacuum, open the chamber door, and remove the target platform.

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7. Using sterile forceps place the filter paper inside an Erlenmeyer flask with 50 ml of 10 mM Tris-HCl pH 7.5 with antibiotic/antimycotic mix at 1:100. Collect the residue of cells from the Petri plate by rinsing with some medium from the flask. 8. Discard the used rupture disc, macrocarrier, and stopping screen. 9. Repeat the shooting three times with the remaining samples of mating cells. If using different DNA samples, wipe the inside surfaces of the gun with 70% ethanol.

6. Turning the Gun Off 1. Close the main gas valve on the helium tank by turning it clockwise. Set the vacuum to ~25 mmHg. 2. Hold the “fire” switch until both gauges of the HPR read 0. 3. Turn the adjustment screw of the HPR counterclockwise until it turns easily. 4. Vent the vacuum. 5. Turn off PDS1000/He and the vacuum pump.

7. Selection of Cells After Shooting 1. After all four shots are completed, place the flasks with cells at 30°C for 12–18 h without shaking. Typically, we incubate cells for 16–18 h but in some cases starting selection at 12 h produced more transformants. 2. Spin cells down at 1100 g for 3 min and suspend in 50 ml of SPPA (with 2 μg/ml of cd for MTT1-driven selectable cassettes). Incubate for 2 h at 30°C and add the selective drug. For neo cassettes, pm is used at 70–100 µg/ml. For bsr, use 60 µg/ml bs. For rpl29-K40M use 12 µg/ml cy. 3. Transfer cells into 96-well plates (flat bottom) at 100 µl per well using a multichannel pipette. Incubate plates in a moist box at 30°C. Screen for wells containing drug-resistant cells 3–4 days after plating. If the bsr cassette is used, the original plates are replicated onto new plates with 30 µl per well of SPPA with 60 µg/ml bs and screened on the next day. bs-resistant (r) cells grow vigorously and swim rapidly while bs-sensitive (s) cells are swollen and grow and divide slowly.

IV. Construction of Gene Knockout Heterokaryons Below we provide a protocol for a typical case of a gene disrupted with neo3 that confers resistance to pm þ cd. A. Identification of Putative Germline Transformants (G1) Typically, after 2–4 days of selection in 70–100 µg/ml pm þ 2 µg/ml cd, 80–100% of wells have growing cells. Most of these wells contain only somatic transformants (derived from nonmating cells). The CU428 cells used for germline transformation have the mpr1-1/mpr1-1 dominant marker in the micronucleus conferring 6-mp

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resistance. Consequently, germline transformants, as the progeny of conjugation between CU428 and B2086, are mp-r, while somatic transformants are mp-s because these cells do not develop a new macronucleus. 1. To identify rare germline transformants (G1 cells, for generation 1) replicate pm þ cd plates onto plates with 15 µg/ml mp in SPPA (50 µl per well) using a 96pin microplate replicator. To clean and sterilize the replicator between different plates, immerse the replicator into water, dry briefly by touching a paper towel, immerse the replicator in acetone, and let dry for a few seconds. 2. On the next day, identify wells containing mp – r cells. Confirm that these cells can grow on media containing pm þ cd þ mp. B. Sexual Maturation and Assortment of Drug Resistance from the Macronucleus of G1 Transformants 1. Establish two to four sublines from each G1 transformant population by single-cell isolations into drops of SPPA (25 µl) on a Petri dish. If the number of (pm þ cd-r/mp-r) G1 transformants is high, we do not make sublines but propagate each G1 population. We expand 10–12 sublines (or the original G1 populations) as one row on a 96-well microtiter plate at 30°C and transferred daily (1 µl into 100 µl of fresh media). 2. After 12–16 transfers, G1 cells should be sexually mature and at least some of them are likely to have lost the pm þ cd resistance by phenotypic assortment. To identify G1 sublines that contain pm þ cd-s cells, make eight single-cell isolations in SPPA drops for each G1 subline. Drops of SPPA (~25 µl) are arranged as a 6 8 grid on a 10-cm Petri dish with spacing that fits the pattern of a 96-well microtiter plate. For each G1 subline, eight isolates represent one column in the grid. We prepare pipettes by melting a glass tube (Kimble, Vineland, NJ, 34,500 99) over the Bunsen burner and pulling at both ends (outside the flame) once the glass becomes soft. A second pull is usually needed to obtain a pipette tip with a proper diameter. The pipette is inserted into the end of an aspirator tube with a mouthpiece (Sigma, A5177-5EA) and operated under a dissecting scope. Grow cells in drops for 2 days at 30°C in a moist box and replicate onto a microtiter plate containing 50 µl/well of SPPA with 1 µg/ml cd. Incubate the plates for 1 h at 30°C and add 50 µl of 200 µg/ml pm, 1 µg/ml cd/well. On the next day, screen the replicated plates and pick up four clones of G1 that are pm þ cd-s (from the corresponding nonselected drops). Preferably each G1 clone should come from a different original G1 transformant. C. Testing for the Germline Presence of the Disrupted Allele 1. Grow each of the 4 pm þ cd-s G1 clones by inoculating 20 µl into 5 ml of SPP in a 15-ml conical centrifuge tube. Also, start a 5-ml culture of CU427 (chx1-1/chx1-1, cy-s, mt VI) strain (homozygous in the micronucleus for the chx-1 allele conferring cy resistance). Keep tubes in a horizontal position to increase aeration and incubate without shaking at 30°C.

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2. On the next morning, transfer 4 ml of each culture into a new 15-ml centrifuge tube; keep the remaining 1 ml of culture at þ15°C. Centrifuge (1100 g, 3 min), discard the supernatant, add 4 ml of 10 mM Tris–HCl (pH 7.5), centrifuge again, and suspend cells in 4 ml of the same buffer. If the cell density appears lower than ~1–2 105 cells/ml, suspend in a smaller volume accordingly (1–3 ml). Add 1:100 volume of the antibiotic/antimycotic mix. Incubate tubes overnight in a horizontal position without shaking. 3. On the next day, mix 50 µl of each starved G1 clone with 50 µl of starved CU427 cells in a 96-well microtiter flat bottom plate. Pairs should start forming after 1–2 h. After 6–8 h, isolate single pairs into drops of SPPA (~25 µl) arranged in a 6 8 grid pattern on a 10-cm Petri plate. Make 24 isolations (half of a grid) for each G1 CU427 cross. If pairs fail to form between CU427 cells and one or more of the G1 populations, these cells may have the same mating type as CU427 (request a strain similar to CU427 with a different mating type from the Tetrahymena Stock Center). 4. Grow G1 CU427 progeny synclones (populations containing cells derived from both exconjugants) for 48 h at 30°C and test for resistance to cy 12 µg/ml and pm 100 µg/ml þ cd 1 µg/ml as follows—Replicate drops onto: (1) 96-well microtiter plates with 50 µl/well of 12 µg/ml of cy in SPPA and (2) 96-well microtiter plates with 50 µl/well of 1 µg/ml cd in SPPA. Incubate plates for 1–2 h at 30°C and add 50 µl of pm 200 µg/ml þ 1 µg/ml cd to the plate containing SPPA with cd. Incubate the plates overnight at 30°C and score for resistance. For a typical heterozygous germline transformant, among the cy-r synclones, about 50% are pm þ cd-r. Sometimes we observe more of the cy-r/pm þ cd-s progeny than expected based on a Mendelian segregation. This is likely because in the developing macronucleus, the neo3 coding region is recognized as a foreign sequence and eliminated by an RNAi-based mechanism that normally deletes micronucleus-specific sequences (Yao and Chao, 2005). Occasionally, the G1 subline, despite the original pm þ cd and mp resistance, lacks the disrupted allele in the micronucleus (e.g., due to a chromosome loss during vegetative propagation or when the targeting fragment replaces the gene directly in the new macronucleus) and fails to give any cy-r and pm þ cd-r progeny in a cross to CU427. In such a case, analyze sublines from independent G1 transformants if available.

D. Round I Genomic Exclusion Cross to Produce G2 Synclones A cross to a * (star) strain is used to make the micronuclear genome of the G1 transformant homozygous for the disrupted gene (round I genomic exclusion). A * strain has an aneuploid micronucleus that is not capable of completing meiosis during conjugation (Allen, 1967). When a fertile cell mates to a * cell, the micronucleus of the fertile cell undergoes meiosis, a postmeiotic mitosis, and unilateral pronuclear transfer to the cytoplasm of the star strain. Following the transfer, pronuclei in both partners undergo endoreduplication and the diploid number of chromosomes is restored (Fig. 2,

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steps J-O). The conjugants of the star cross separate early without developing new macronuclei. The procedure described here is used to perform a round I genomic exclusion cross and an outcross to identify G2 cells that are homozygous for the disrupted gene in the micronucleus. 1. Grow one to four G1 sublines (identified earlier as carrying a disrupted allele in the germline, 1 ml of which was stored at þ15°C) in 5-ml SPP cultures at 30°C overnight. Also grow a 5 ml culture of A*III (or B*VII) strain. Induce matings of each G1 subline with A*III using the technique described in IV.C.2–3 and isolate 48 pairs into drops for each cross. Incubate Petri plates with drops at 30°C for 2 days. 2. On the same day, start a culture of CU427 cells and grow overnight to a density of 2–3 105 cells/ml. 3. On the next day, starve the 50 ml culture of CU427 by washing with 10 mM Tris– HCl (pH 7.5). Add the antibiotic/antimycotic mix (0.5 ml) and leave starving overnight at 30°C without shaking. 4. On the following day, adjust the cell concentration of starved CU427 to 2 105 cells/ml. Distribute CU427 cells, 50 µl per well on a 96-well microtiter plate (for four G1 sublines, two plates are needed). Also prepare plates with 100 µl/well SPPA. Replicate the G1 A*III progeny (G2 synclones) from drops onto plates containing: (1) SPPA (100 µl/well) and (2) starved CU427 cells (50 µl per well). Store the SPPA plates at 15°C (these are backup plates containing G2 synclones). Incubate the plates containing G2 synclones CU427 cells at 30°C overnight. Pairs should start forming at the bottom of wells after 6–8 h. 5. On the next day (16–24 h later), add 50 µl/well of SPPA with 2 µg/ml cd to the G2 synclone CU427 plates. Incubate for 7–8 h at 30°C. Add 50 µl/well of 300 µg/ml pm, 36 µg/ml cy, and 1 µg/ml cd. Incubate plates at 30°C for 1–2 days. Score plates for pm þ cd þ cy resistance. E. Isolation of Heterokaryon Cells with Different Mating Types Among G2 Synclones The wells identified in the previous step contain G2 synclones with a mixture of heterokaryons with two different mating types. Heterokaryons with single mating types are re-isolated by the following steps: 1. Pick up cells from four G2 synclone wells (from the SPPA backup) that gave pm þ cd þ cy resistance in a cross to CU427 and start 5-ml cultures to grow overnight. 2. On the next day, transfer 4 ml of each G2 synclone culture into a new 15-ml centrifuge tube, wash by centrifugation with 10 mM Tris (pH 7.5), and suspend in 2–4 ml of the same buffer so that the cell density is 1–2 105 cells/ml. Transfer 50 µl of each starving G2 synclone into wells of a 96-well plate at 30°C. 3. About 3–4 h later, pairs will start to form. For each G2 synclone, pick up four early pairs and gently isolate into drops of SPPA (one pair per drop). Incubate drops at room temperature for 30–60 min. During this period, pairs will separate because early addition of food reverses commitment for conjugation. For each drop, pick up one early exconjugant cell and transfer to an adjacent empty drop of SPPA.

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4. On the next day, inoculate 5-ml tube cultures with exconjugant clones. Establish pairs of clones that originate from four G2 G2 pairs (eight clones total). Heterokaryon clones derived from the same G2 pair have isogenic micronuclei. Thus, progeny of such mating heterokaryons will have a nearly identical macronuclear genome [except for alternative processing events that are known to occur in the developing macronucleus (White and Allen, 1986)]. F. Testing the Putative Heterokaryons and Bringing the Gene Disruption Phenotype to Expression by a Heterokaryon Heterokaryon Cross 1. You should now have four pairs of putative (micronucleus-isogenic) heterokaryon clones. To test whether these clones are homozygous for the disrupted drugresistant allele in the micronucleus, cross each to CU427, isolate 48 single pairs in drops, grow, and test for resistance to cy and pm þ cd (separately) by replica plating. Among the cy-r progeny, all or nearly all progeny should be pm þ cd-r. A high incidence of cy-s progeny or lack of viability in a CU427 outcross indicates that the heterokaryon strain is deficient in fertility, possibly due to micronuclear aneuploidy. Fertility can be increased by crossing the heterokaryon to a wild-type strain, maturing heterozygotes, and repeating the round I genomic exclusion to isolate a new generation of heterokaryons. 2. To reveal the gene disruption phenotype, cross a pair of heterokaryons that are derived from the same G2 synclone. Isolate 48 single pairs of mating heterokaryons into drops of MEPPA. This medium bypasses the requirement for cilia-dependent phagocytosis (Orias and Rasmussen, 1976) and supports growth of cells lacking functional cilia (Brown et al., 1999b). A typical outcome of a mutation that prevents the assembly of cilia is the appearance of paralyzed cells that are thin, lack food vacuoles, and are multinucleated. Tetrahymena cells use ciliary motility to sever the cytoplasmic bridge at the end of cytokinesis (rotokinesis), and the presence of an excessive number of nuclei is a result of cytokinesis failures (in 50–60% of cells) (Brown et al., 1999a). Paralyzed cells can be grown in larger volumes of MEPPA with shaking (at 160 rpm) that provides a sufficient mechanical force for abscission (Brown et al., 2003). Note that among the heterokaryon heterokaryon progeny, usually several drops contain apparently wild-type cells. These cells are derived from abortive conjugation that is associated with retention of the original macronucleus and are pm þ cd-s. A high frequency of such progeny indicates low fertility of heterokaryons. The fertility can be improved by an outcross to a wildtype strain followed by another round I genomic exclusion.

V. Disruption of Multiple Genes in the Germline Multiple disrupted genes can be combined in one strain by crosses. This is facilitated by the apparently high level of meiotic recombination in Tetrahymena (McCoy, 1977). A disruption of multiple genes is needed in studies on gene products that are encoded

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by multiple paralogous genes. For example, the function of the kinesin-2 heavy chain in anterograde IFT is provided redundantly by the KIN1 and KIN2 genes (Brown et al., 1999b). To delete multiple genes in one strain, we disrupt each paralog gene separately and combine the disrupted alleles by crosses.

A. Disruption of Multiple Genes with Different Selectable Markers As a typical example, a heterokaryon with a neo3 marker is crossed to a heterokaryon with an rpl29-K40M marker. The double drug resistance is lost by phenotypic assortment. Next, four drug-sensitive heterozygotes are crossed to A*III, and wells containing synclones with both gene disruptions are identified by a cross to starved B2086 (wild-type) cells following the procedure outlines in Section IV.D. and selection of progeny using pm, cd, and cy. To generate a triple knockout clone, cross an appropriate double knockout heterokaryon to single knockout heterokaryon. If all three cassette types are used (neo, cy, bsr) all drugs can be combined together at the stage of selection of (triple heterozygote A*) B2086 progeny.

B. Disruption of Multiple Genes with a Single Selectable Marker If the goal is to combine two disruptions that contain the same marker (e.g., neo3), the double heterozygote synclones that gave pm þ cd resistance in a cross to CU427 (IV.D) are screened for the presence of two neo3-interrupted genes by PCR genotyping as follows: 1. Select pm þ cd-r (double heterozygote A*III) B2086 progeny for 2–3 days at 30°C. Pick up 10–20 drug-resistant progeny and inoculate 5-ml SPPA cultures. Grow overnight in SPP with pm þ cd. Take 1.5 ml of each culture and extract total genomic DNA as follows: Genomic DNA Miniprep [based on a protocol by Martin Zillmann, personal communication (Gaertig et al., 1994b)]. 2. Spin down 1.5 ml cells (1–3 105 cells/ml) in a microcentrifuge for 30 s. 3. Remove the supernatant. Add 0.7 ml of the urea extraction buffer, mix gently by inverting tubes two to three times, and incubate for up to 4 min at room temperature. The urea buffer contains 42% urea, 0.35 M NaCl, 0.01 M Tris (pH 7.4), 0.01 M EDTA, and 1% SDS. Make a 2 buffer stock without urea and store at room temperature. To prepare the working solution, add 4.2 g urea to 5 ml of the 2 buffer stock and adjust the volume to 10 ml with water. 4. Extract the lysate twice with an equal volume of phenol : chloroform : isoamyl alcohol (25:24:1) and once with chloroform : isoamyl alcohol (24:1). Invert tubes to mix, but do not vortex. Each time centrifuge tubes in a microcentrifuge at 16,000 g for 5 min at room temperature. Collect the upper phase after each extraction.

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5. Add 0.15 ml of 5 M NaCl (usually to ~0.5 ml of recovered upper phase). 6. Add an equal volume (usually 0.65 ml) of isopropyl alcohol to precipitate DNA, mix gently for 1 min, and spin down at 6,000 g for 2 min in a microcentrifuge. 7. Wash the DNA pellet with 0.5 ml of 70% ethanol, remove as much of the supernatant as possible and air dry the pellet. 8. Suspend in 40 µl of water and add 1 µl RNAse A (10 mg/ml). Incubate at 55°C for 3–4 h or at 37°C overnight. 9. Use PCR to amplify each genomic DNA with a combination of primers that amplify the junction between the selectable marker and the region of the gene outside of the targeted segment. Identify the (double heterozygote A*III) B2086 progeny that carries both disrupted alleles. Pick two to four synclone populations from the corresponding backup plates in SPPA and identify heterokaryons with different mating types as described in Section IV.E. The procedure can be repeated to disrupt and combine additional loci. We have managed to eliminate a family of six paralogous genes in one strain using only two markers (Wloga et al., 2009).

VI. Somatic (Macronuclear) Gene Knockout Somatic transformants can be isolated as a byproduct of the germline transformation experiment as described above. If the goal is limited to somatic transformation, the same protocol as for the germline knockout is used (Section IV), except that only the CU428 strain is used and the mating steps are omitted. Transformant wells are identified and 10–12 are propagated to promote complete allelic replacement by phenotypic assortment as follows: 1. For a neo3-disrupted allele, select somatic transformants in SPPA with 100 µg/ml pm þ 2.5 µg/ml cd. 2. Take cells from 12 positive wells, transfer onto 100 µl of fresh media with 100 µg/ml pm þ 0.5 µg/ml cd. Transfer cells every 1–2 days (1 µl into 100 µl fresh media) on a microtiter 96-well plate, while gradually increasing the pm concentration. If cells grow very slowly after transfer, go back to the previous lower drug concentration and continue to grow cells. 3. After 12–24 transfers, make single-cell isolations (48 drops per transformant) in MEPPA. If the targeted gene is needed for either ciliary assembly or motility of cilia, on the next day, some drops will contain most or all cells that are paralyzed or nearly paralyzed. Re-isolate populations with paralyzed cells several times and grow in MEPPA. If after several re-isolations, there is no evidence of phenotype reversal, it is safe to assume that all copies of the endogenous genes have been eliminated. This can be confirmed by performing western blots or RT-PCR.

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VII. Phenotypic Rescue by Introduction of a Wild-type Gene into a Knockout Strain A phenotypic rescue can be done by transforming the gene knockout strain with a fragment containing a functional intact (or epitope-tagged) gene. A rescuing fragment can be targeted either into the native locus or into another nonessential locus (by adding appropriate flanking sequences on the sides of the targeting fragment). Reintroduction of the rescuing fragment into another nonessential locus is preferred as a proof that the mutant phenotype is caused by loss of function of the targeted gene. This is because a genomic integration of the selectable marker cassette could affect adjacent genes. Moreover, nested genes with overlapping or even embedded ORFs on the opposite strands of genomic DNA have been reported (Zweifel et al., 2009). For targeting into a third locus, we typically use the BTU1 locus, which encodes a nonessential b-tubulin (Xia et al., 2000). Typically, the rescuing coding region operates under the MTT1 promoter. Alternatively, a rescue can be done by reintroduction of the functional coding region into the native locus (that replaces the disrupted copy). This approach is preferred for placing an epitope tag and localization studies, since the levels of expression of the tagged protein are expected to be nearly normal. Two approaches can be used for introduction of the rescuing fragment: (1) rescue of mating heterokaryons and (2) rescue of vegetatively growing knockout cells.

A. Rescue of Mating Heterokaryons Grow and starve heterokaryon strains and follow the general protocol for biolistic transformation, except that the shooting is done between 10 and 24 h after mixing of two heterokaryon strains. In one case, the 24 h time point gave the highest rescue frequency (Hai et al., 1999). After shooting, select cells on SPPA with pm þ cd (for neo3) and screen for plates with drug-resistant cells that also recover motility. The selection will allow for survival of progeny of mating heterokaryons that express the knockout allele and at the same time express the rescuing transgene.

B. Rescue of Vegetative Paralyzed Knockout Cells Grow paralyzed or slowly moving knockout cells in MEPPA (50 ml) to a density of 2 105 cells/ml, wash cells with 10 mM Tris–HCl, pH 7.5, subject to biolistic transformation with a rescuing fragment, place on 96-well plates, and screen for cell that regained normal motility after 2–3 days. If the rescuing fragment has a coding region that operates under the MTT1 promoter, cd can be added to the selection media after transformation (e.g., 0.5–1 µg/ml). However, in most of the cases we have studied, rescues have occurred without exogenous cadmium, apparently because the basal level of MTT1 promoter activity is sufficient for expression of ciliary genes.

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Acknowledgments We thank Jody Bowen and Martin Gorovsky (University of Rochester) and Donna Cassidy-Hanley (Cornell University) for sharing lab protocols. We acknowledge the support of the National Science Foundation (grant MBC-033965).

References Allen, S.L. (1967). Genomic exclusion: A rapid means for inducing homozygous diploid lines in Tetrahymena pyriformis, syngen 1. Science 155, 575–577. Angus, S.P., Edelmann, R.E., and Pennock, D.G. (2001). Targeted gene knockout of inner arm 1 in Tetrahymena thermophila. Eur. J. Cell Biol. 80, 486–497. Beales, P.L., Bland, E., Tobin, J.L., Bacchelli, C., Tuysuz, B., Hill, J., Rix, S., Pearson, C.G., Kai, M., Hartley, J., Johnson, C., Irving, M., et al., (2007). IFT80, which encodes a conserved intraflagellar transport protein, is mutated in Jeune asphyxiating thoracic dystrophy. Nat. Genet. 39, 727–729. Brown, J.M., Fine, N.A., Pandiyan, G., Thazhath, R., and Gaertig, J. (2003). Hypoxia regulates assembly of cilia in suppressors of Tetrahymena lacking an intraflagellar transport subunit gene. Mol. Biol. Cell 14, 3192–3207. Brown, J.M., Hardin, C., and Gaertig, J. (1999a). Rotokinesis, a novel phenomenon of cell locomotionassisted cytokinesis in the ciliate Tetrahymena thermophila. Int. Cell Biol. Rep. 23, 841–848. Brown, J.M., Marsala, C., Kosoy, R., and Gaertig, J. (1999b). Kinesin-II is preferentially targeted to assembling cilia and is required for ciliogenesis and normal cytokinesis in Tetrahymena. Mol. Biol. Cell 10, 3081–3096. Bruns, P.J., and Cassidy-Hanley, D. (2000a). Biolistic transformation of macro- and micronuclei. Methods Cell Biol. 62, 501–512. Bruns, P.J., and Cassidy-Hanley, D. (2000b). Methods for genetic analysis. Methods Cell Biol. 62, 229–240. Cassidy-Hanley, D., Bowen, J., Lee, J., Cole, E.S., VerPlank, L.A., Gaertig, J., Gorovsky, M.A., and Bruns, P.J. (1997). Germline and somatic transformation of mating Tetrahymena thermophila by particle bombardment. Genetics 146, 135–147. Culver, B.P., Meehl, J.B., Giddings, T.H., Jr., and Winey, M. (2009). The two SAS-6 homologs in Tetrahymena thermophila have distinct functions in basal body assembly. Mol. Biol. Cell 20, 1865–1877. Doerder, F.P., Deak, J.C., and Lief, J.H. (1992). Rate of phenotypic assortment in Tetrahymena thermophila. Dev. Genet. 13, 126–132. Eisen, J.A., Coyne, R.S., Wu, M., Wu, D., Thiagarajan, M., Wortman, J.R., Badger, J.H., Ren, Q., Amedeo, P., Jones, K.M., Tallon, L.J., Delcher , et al., (2006). Macronuclear genome sequence of the ciliate Tetrahymena thermophila, a model eukaryote. PLoS Biol. 4(9): e286. Frankel, J. (2000). Cell biology of Tetrahymena. Methods Cell Biol. 62, 27–125. Gaertig, J., Gu, L., Hai, B., and Gorovsky, M.A. (1994a). High frequency vector-mediated transformation and gene replacement in Tetrahymena. Nucleic Acids Res. 22, 5391–5398. Gaertig, J., Thatcher, T.H., Gu, L., and Gorovsky, M.A. (1994b). Electroporation-mediated replacement of a positively and negatively selectable b-tubulin gene in Tetrahymena thermophila. Proc. Natl. Acad. Sci. USA 91, 4549–4553. Gorovsky, M.A. (1973). Macro- and micronuclei of Tetrahymena pyriformis: A model system for studying the structure and function of eukaryotic nuclei. J. Protozool. 20, 19–25. Hai, B., Gaertig, J., and Gorovsky, M.A. (1999). Knockout heterokaryons enable facile mutagenic analysis of essential genes in Tetrahymena. Methods Cell Biol. 62, 513–531. Hai, B., and Gorovsky, M.A. (1997). Germ-line knockout heterokaryons of an essential alpha-tubulin gene enable high-frequency gene replacement and a test of gene transfer from somatic to germ-line nuclei in Tetrahymena thermophila. Proc. Natl. Acad. Sci. USA 94, 1310–1315. Hamilton, E.P., and Orias, E. (2000). Genetic crosses: Setting up crosses, testing progeny, and isolating phenotypic assortants. Methods Cell Biol. 62, 219–228.

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Hennessey, T.M., Kim, D.Y., Oberski, D.J., Hard, R., Rankin, S.A., and Pennock, D.G. (2002). Inner arm dynein 1 is essential for Caþþ-dependent ciliary reversals in Tetrahymena thermophila. Cell Motil. Cytoskeleton 53, 281–288. Kilburn, C.L., Pearson, C.G., Romijn, E.P., Meehl, J.B., Giddings, T.H., Jr., Culver, B.P., Yates, J.R., 3rd., and Winey, M. (2007). New Tetrahymena basal body protein components identify basal body domain structure. J. Cell Biol. 178, 905–912. Liu, S., Hard, R., Rankin, S., Hennessey, T., and Pennock, D.G. (2004). Disruption of genes encoding predicted inner arm dynein heavy chains causes motility phenotypes in Tetrahymena. Cell Motil. Cytoskeleton 59, 201–214. McCoy, J.W. (1977). Linkage and genetic map length in Tetrahymena thermophila. Genetics 87, 421–439. Mochizuki, K. (2008). High efficiency transformation of Tetrahymena using a codon-optimized neomycin resistance gene. Gene 425, 79–83. Orias, E., and Flacks, M. (1975). Macronuclear genetics of Tetrahymena. I. Random distribution of macronuclear genecopies in T. pyriformis, syngen 1. Genetics 79, 187–206. Orias, E., Hamilton, E.P., and Orias, J.D. (2000). Tetrahymena as a laboratory organism: Useful strains, cell culture, and cell line maintenance. Methods Cell Biol. 62, 189–211. Orias, E., and Rasmussen, L. (1976). Dual capacity for nutrient uptake in Tetrahymena. IV. Growth without food vacuoles. Exp. Cell Res. 102, 127–137. Rajagopalan, V., Subramanian, A., Wilkes, D.E., Pennock, D.G., and Asai, D.J. (2009). Dynein-2 affects the regulation of ciliary length but is not required for ciliogenesis in Tetrahymena thermophila. Mol. Biol. Cell 20, 708–720. Shang, Y., Li, B., and Gorovsky, M.A. (2002a). Tetrahymena thermophila contains a conventional gamma tubulin that is differentially required for the maintenance of different micotubule organizing centers. J. Cell Biol. 158, 1195–1206. Shang, Y., Song, X., Bowen, J., Corstanje, R., Gao, Y., Gaertig, J., and Gorovsky, M.A. (2002b). A robust inducible-repressible promoter greatly facilitates gene knockouts, conditional expression, and overexpression of homologous and heterologous genes in Tetrahymena thermophila. Proc. Natl. Acad. Sci. USA 99, 3734–3739. Sharma, N., Bryant, J., Wloga, D., Donaldson, R., Davis, R.C., Jerka-Dziadosz, M., and Gaertig, J. (2007). Katanin regulates dynamics of microtubules and biogenesis of motile cilia. J. Cell Biol. 178, 1065–1079. Shen, X., Yu, L., Weir, J.W., and Gorovsky, M.A. (1995). Linker histones are not essential and affect chromatin condensation in vivo. Cell 82, 47–56. Smith, J.C., Northey, J.G., Garg, J., Pearlman, R.E., and Siu, K.W. (2005). Robust method for proteome analysis by MS/MS using an entire translated genome: Demonstration on the ciliome of Tetrahymena thermophila. J. Proteome Res. 4, 909–919. Tsao, C.C., and Gorovsky, M.A. (2008a). Different effects of Tetrahymena IFT172 domains on anterograde and retrograde intraflagellar transport. Mol. Biol. Cell 19, 1450–1461. Tsao, C.C., and Gorovsky, M.A. (2008b). Tetrahymena IFT122A is not essential for cilia assembly but plays a role in returning IFT proteins from the ciliary tip to the cell body. J. Cell Sci. 121, 428–436. White, T.C., and Allen, S.L. (1986). Alternative processing of sequences during macronuclear development in Tetrahymena thermophila. J. Protozool. 33, 30–38. Williams, N.E., Tsao, C.C., Bowen, J., Hehman, G.L., Williams, R.J., and Frankel, J. (2006). The actin gene ACT1 is required for phagocytosis, motility, and cell separation of Tetrahymena thermophila. Eukaryot. Cell 5, 555–567. Wloga, D., Webster, D., Rogowski, K., Bré, M.H., Levilliers, N., Jerka-Dziadosz, M., Janke, C., Dougan, S.T., and Gaertig, J. (2009). TTLL3 is a tubulin glycine ligase that regulates the assembly of cilia. Dev. Cell 16, 867–876. Wood, C.R., Hard, R., and Hennessey, T.M. (2007). Targeted gene disruption of dynein heavy chain 7 of Tetrahymena thermophila results in altered ciliary waveform and reduced swim speed. J. Cell Sci. 120, 3075–3085. Xia, L., Hai, B., Gao, Y., Burnette, D., Thazhath, R., Duan, J., Bré, M.-H., Levilliers, N., Gorovsky, M.A., and Gaertig, J. (2000). Polyglycylation of tubulin is essential and affects cell motility and division in Tetrahymena thermophila. J. Cell Biol. 149, 1097–1106.

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Drashti Dave et al. Yao, M.C., and Chao, J.L. (2005). RNA-guided DNA deletion in Tetrahymena: an RNAi-based mechanism for programmed genome rearrangements. Annu. Rev. Genet. 39, 537–559. Yao, M.-C., and Yao, C.-H. (1991). Transformation of Tetrahymena to cycloheximide resistance with a ribosomal protein gene through sequence replacement. Proc. Natl. Acad. Sci. USA 88, 9493–9497. Zweifel, E., Smith, J., Romero, D., Giddings, T.H., Jr., Winey, M., Honts, J., Dahlseid, J., Schneider, B., and Cole, E.S. (2009). Nested genes CDA12 and CDA13 encode proteins associated with membrane trafficking in the ciliate Tetrahymena thermophila. Eukaryot. Cell. 8, 899–912.

CHAPTER 2

Approaches for Functional Analysis of Flagellar Proteins in African Trypanosomes Michael Oberholzer1, Miguel A. Lopez1, Katherine S. Ralston1,3, and Kent L. Hill1,2 1 Department of Microbiology, Immunology, and Molecular Genetics, University of California Los Angeles, Los Angeles, CA-90095, United States of America 2 Molecular Biology Institute, University of California Los Angeles, Los Angeles, CA-90095, United States of America 3 Current address Department of Medicine, Division of Infectious Diseases and International Health, University of Virginia Health Sciences Center, Charlottesville, VA-22908, United States of America

Abstract I. Introduction II. Rationale III. Methods A. Culture Methods B. Transfection C. Protocols for Flagellum Skeleton Isolation D. Assays for Trypanosome Motility E. Genetic Approaches to Study Individual Components of the T. brucei Flagellum IV. Materials A. Culture Medium B. Transfection Reagents C. Antibiotic Concentrations for Selection (See Table II) D. Buffers for Isolation of Extracted Flagellum Skeletons (Robinson et al., 1991) E. Plasmids (See Table III) F. Online Resources V. Summary and Outlook Acknowledgments References METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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978-0-12-381377-0 DOI: 10.1016/S0091-679X(08)93002-8

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Abstract The eukaryotic flagellum is a highly conserved organelle serving motility, sensory, and transport functions. Although genetic, genomic, and proteomic studies have led to the identification of hundreds of flagellar and putative flagellar proteins, precisely how these proteins function individually and collectively to drive flagellum motility and other functions remains to be determined. In this chapter we provide an overview of tools and approaches available for studying flagellum protein function in the protozoan parasite Trypanosoma brucei. We begin by outlining techniques for in vitro cultivation of both T. brucei life cycle stages, as well as transfection protocols for the delivery of DNA constructs. We then describe specific assays used to assess flagellum function including flagellum preparation and quantitative motility assays. We conclude the chapter with a description of molecular genetic approaches for manipulating gene function. In summary, the availability of potent molecular tools, as well as the health and economic relevance of T. brucei as a pathogen, combine to make the parasite an attractive and integral experimental system for the functional analysis of flagellar proteins.

I. Introduction The eukaryotic flagellum, aka cilium1, is a ubiquitous organelle present on most differentiated mammalian cells and on many single-celled microbes. The flagellum performs motility, transport, and sensory functions and is essential for normal human development and physiology (Badano et al., 2006; Ibanez-Tallon et al., 2003; Satir and Christensen, 2008). Flagella are also required for many important human pathogens, including the causative agents of malaria, African sleeping sickness, leishmaniasis, epidemic diarrhea, and trichomoniasis (Ferguson, 2002; Vlachou et al., 2006). Together, these diseases are responsible for morbidity and mortality in several hundred million people worldwide. Therefore, in addition to providing insight into the operation of molecular motors and principles of cell biology, studies of the mechanisms of flagellum protein function are directly relevant to understanding mechanisms of heritable and infectious human diseases. In this chapter we provide an overview of tools and approaches available for studying flagellum protein function in the protozoan parasite Trypanosoma brucei.

1

Flagellum and cilium are interchangeable terms for the same eukaryotic organelle and distinct from the bacterial flagellum.

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II. Rationale African trypanosomes, for example, T. brucei2, are protozoan parasites that are the causative agent of African trypanosomiasis, commonly referred to as African sleeping sickness in humans. These parasites are devastating human and animal pathogens that cause significant human mortality and limit economic development in sub-Saharan Africa. Trypanosomes are transmitted between mammalian hosts through the bite of a tsetse fly and parasite motility is important in both hosts (Kohl and Bastin, 2005; Ralston and Hill, 2008). Trypanosome motility is driven by a single flagellum that emerges from the flagellar pocket at the cell posterior and wraps around the cell body as it extends to the cell anterior (Fig. 1) (Ralston et al., 2009). The propulsive beat is a tractile beat that progresses tip-to-base and drives the cell forward with the flagellum tip leading (Walker, 1961). The flagellum is laterally attached to the cell body along most of its length, causing the entire cell to undulate and rotate in an auger-like fashion as the flagellum beats (http://www.elsevierdirect.com/companions/ 9780123749734, Supplementary Movie 1, adapted from Ralston et al. 2006, with permission). This distinctive motility gives the genus its name, from the Greek trypanon (“auger”) and soma (“body”) (Gruby, 1843).

2 3

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2 4 3 1 (A)

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Fig. 1 Trypanosoma brucei cell and flagellum structure. (A) Scanning electron microscopy image of PCF T. brucei. (1) flagellum, (2) flagellar pocket, (3) cell body posterior end, (4) flagellar tip, corresponds to cell anterior. (B) Cross-section of PCF T. brucei cell as imaged by transmission electron microscopy. (1) flagellar membrane, (2) axoneme, (3) paraflagellar rod, (4) Intraflagellar transport particle, (5) cell body membrane. Subpellicular microtubules (not labeled) are visible in cross section in cell body. Adapted from Ralston et al. (2009), with permission.

2

We use the term trypanosome to refer to Trypanosoma brucei, the topic of the present review, although related Trypanosoma species, most notably T. congelense are also responsible for disease in wild and domestic animals.

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The T. brucei flagellum contains a conserved 9 þ 2 axoneme, as well as a unique paraaxonemal structure known as the paraflagellar rod (PFR) (Fig. 1). The function of the PFR is not fully understood, but it is required for normal motility (Bastin et al., 1998) and is hypothesized to play structural (Maga and LeBowitz, 1999) and regulatory roles, since it contains components of calcium and cAMP signaling pathways in addition to proteins with nucleotide transfer domains (Oberholzer et al., 2007a,b; Portman et al., 2009; Pullen et al., 2004). Trypanosomes are extracellular parasites and thus are presumed to be dependent on their own flagellum-mediated motility to migrate through host tissues. The trypanosome flagellum is also a critical host–parasite interface postulated to provide sensory functions (Tetley and Vickerman, 1985; Vassella et al., 2009) and recent functional studies indicate that the flagellum and/or flagellar motility are required for cell morphogenesis, cell division, and evasion of host immune defenses (Broadhead et al., 2006; Kohl et al., 2003; Ralston et al., 2006). As such, the flagellum and flagellar motility have emerged as attractive drug targets in T. brucei and understanding mechanisms of flagellum protein function in these parasites is critical for exploiting this possibility. Genetic screens, biochemical, genomic, and proteomic analyses in several organisms have led to the identification of hundreds of flagellar and putative flagellar proteins (Broadhead et al., 2006; Li et al., 2004; Ostrowski et al., 2002; Pazour, 2004; Pazour et al., 2005; Smith et al., 2005). A key challenge now is to determine where and how these proteins function individually and collectively to drive flagellar motility and other flagellum functions. In addition to its relevance to public health and economic development in some of the poorest places in the world, the molecular tools available in T. brucei make it an excellent experimental system in which to study flagellum protein function. Both life cycle stages are easily cultured in vitro in semidefined medium and amenable to biochemical and cell biological analyses. T. brucei also offers facile systems for reverse genetics, including heritable and inducible RNAi, targeted gene knockouts, stable and inducible expression of recombinant proteins, as well as systems for forward genetic screens (Leal et al., 2004; Morris et al., 2002). Highly efficient homologous recombination allows epitope and tandem affinity purification tags to be readily integrated into any endogenous locus using a single-step PCR approach (Oberholzer et al., 2006; Schimanski et al., 2005; Shen et al., 2001). The genome is sequenced and annotated and the absence of introns makes gene identification simple and reliable (Berriman et al., 2005). These features combine to make T. brucei a valuable experimental system with tools and approaches for functional analysis of flagellar proteins that are complementary to those available in other organisms.

III. Methods The methods section is organized in three parts. Sections III.A and III.B describe basic culture and transfection methodologies. Sections III.C and III.D describe specific assays used to assess flagellum function, for example, flagellum preparation

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and motility assays. In Section III.E, we describe molecular genetic approaches for manipulating gene function.

A. Culture Methods Insect-form (procyclic culture-form, “PCF”) and bloodstream-form (“BSF”) trypanosomes can be maintained in semi-defined media in suspension culture. Both life cycle forms can also be maintained on semi-solid agarose plates (Carruthers and Cross, 1992), although the current review is restricted to methods employing suspension cultures.

1. Procyclic Form (PCF) Two basic media formulations, SM (Cunningham, 1977) or SDM-79 (Brun and Schonenberger, 1979), are used to cultivate PCF trypanosomes in suspension cultures. The recipe for SM is provided in Section IV, while the SDM-79 recipe can be found in Brun and Schonenberger (1979). Work in the authors’ laboratory employs SM exclusively and specific growth rates might vary for SDM-79. Cultures are maintained at 27°C with or without 5% CO2. The precise advantage provided by CO2 supplementation is unclear, but is postulated to be via pH effects [http:// tryps.rockefeller.edu/] and in our experience generally provides for faster cell doubling and facilitates growth at low cell densities. If desired, phenol red can be added to monitor pH of the medium, though this is not required and should be avoided if sedimentation assays using optical density (Section III.D) are performed. The most widely used cell line employed for laboratory studies, 29–13, is derived from the 427 strain and engineered to encode a T7 polymerase and Tet repressor for regulated gene expression (Wirtz et al., 1999). This strain has a doubling time of 8– 9 h in SM medium, with logarithmic growth between ~1 106 and 1 107 cells/ml. For routine maintenance, cultures are diluted ~1:100–1:300 every 2–3 days. For best results in transfections, cultures should be maintained in log phase for several generations, achieved by daily 1:5–1:10 dilution.

2. Bloodstream Form BSF 427-derived trypanosomes encoding T7 polymerase and Tet repressor for inducible gene expression (Wirtz et al., 1999) are most commonly used for laboratory studies. BSF cells are grown in suspension cultures using HMI-9 medium (Hesse et al., 1995) containing 10–15% FBS. The recipe for HMI-9 is provided in Section IV. BSF suspension cultures are maintained at 37°C with 5% CO2. Doubling times are 6–8 h and logarithmic growth is between ~2 105 and 3 106 cells/ ml. Cultures are diluted as for PCF cultures, though additional care should be taken to avoid overgrowth beyond log phase, because BSF cultures die quickly upon reaching stationary phase.

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3. Freezing and Thawing PCF and BSF cells can be frozen in culture media, including FBS and containing 10% glycerol (“freezing medium”), and stored at –80°C for several months or in liquid nitrogen for several years. Trypanosomes should be frozen from a healthy logarithmic phase culture. Cells are harvested by centrifugation at 2500 g for 5 min. The cell pellet is washed once in prewarmed culture medium and resuspended gently in prewarmed freezing medium using 1/2–1/10 of the initially harvested volume, giving a final density of ~2 107 cells/ml. Freezing medium may be supplemented with 10% conditioned medium and increased FBS concentration to 15%. Conditioned medium is obtained by harvesting a late log phase culture by centrifugation at 2500 g for 5 min and filtering the supernatant through a 0.22 µm filter. Conditioned medium can be stored for several weeks at 4°C or frozen at –20°C. Cells in freezing medium are aliquoted (0.5–1.0 ml) into appropriate cryotubes with rubber O-ring, for example, Sarstedt, 72.694.006, then frozen slowly by placing them into a styrofoam tube rack at –80°C. Stock viability should be checked by thawing a few days after freezing, prior to transfer to permanent storage. Thawing of trypanosomes is done as quickly as possible, for example, in a water bath at 27°C for PCF or 37°C for BSF. Thawed cells are diluted 1/10 into fresh, prewarmed culture medium and placed in the incubator. If desired, cells can be washed once in prewarmed culture medium to wash out the glycerol. Generally, if the culture survives the first 24 h after thawing, it is viable for future, though it should be maintained for an additional week prior to use in experiments. B. Transfection PCF and BSF cells can be transiently or stably transfected by electroporation with linear and circular plasmid DNA constructs (e.g., Hill et al., 1999; Lee and Van der Ploeg, 1990; Zomerdijk et al., 1990). PCR products can also be used (Arhin et al., 2004; Oberholzer et al., 2006; Shen et al., 2001). Transfection efficiency using standard electroporation systems is 10–3–10–6 for PCF and 107108 for BSF (Burkard et al., 2007). Several labs have reported a sharp increase (70–1000-fold) in stable transfection efficiency of BSF trypanosomes when using the Amaxa Nucleofector® system (http:// www.lonzabio.com/technology.html) (Burkard et al., 2007), which employs preset programs and “cell-type specific” confidential transfection solutions. Further increases in BSF transfection efficiency have been reported when DNA double strand breaks are introduced in the target cells (Glover and Horn, 2009). These new developments are very promising, particularly for genome-wide analyses in BSF cells. However, the current discussions are restricted to standard electroporation conditions used in the authors’ laboratory. Alternate protocols and solutions for transfection may be obtained from online sources, for example, G. Cross lab (http://tryps.rockefeller.edu/).

1. DNA Preparation Any standard high-quality plasmid preparation system, for example, Qiagen®, etc., is appropriate for transfection. Care should be taken to ensure sterile technique is

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employed. Miniprepped DNA can be used as well, although care should be taken that it is of good quality. Circular plasmids (25–100 µg) are used for transient transfection. Linearized plasmids or DNA constructs (5–20 µg) are used for stable transfection.

2. PCF Transfection Cells in mid log phase, for example, 4–7 106 cells/ml, are harvested by centrifugation at 2500 g for 5 min, washed once with room temperature Electroporation Media (“EM,” see recipe in Section IV), then resuspended in EM to a final concentration of 5 107 cells/ml. A 0.45 ml aliquot of cells (2.25 107 cells) is added to a 0.4 cm BioRad® electroporation cuvette containing 100 µl of DNA in water. Circular DNA (50– 100 µg) is used for transient transfection and 5–20 µg of linear DNA for stable transfection. Cells and DNA are mixed by gently pipeting up and down, avoiding air bubbles. Cell–DNA mix is then electroporated with two successive pulses, separated by a 10 s interval, in a Gene Pulser II (BioRad®) electroporator using the following settings: 1500 V, 25 µF. The time constant should be between 0.5 and 0.7 ms. For transient transfections, 0.3 ml of electroporated cells are transferred to 5 ml fresh medium and monitored 12–36 h post-transfection for phenotype. For stable transfections, following electroporation, cells are immediately transferred into 12 ml prewarmed culture medium and aliquotted (0.5 ml) into each well of a 24-well culture plate. Following an overnight incubation, antibiotic for selection is added as 0.5 ml of culture medium containing 2 the amount of required antibiotic to each well. The amount of antibiotic required for selection is summarized in Section IV.C. Monitor cells daily and dilute as necessary in presence of drug until mock transfected samples die out and transfected cells are growing well, generally 1–2 weeks post-transfection. Each well represents a “pool” of clones, which can then be isolated by limiting dilution (Klingbeil et al., 2002). Alternatively, one can perform limiting dilution immediately after adding drug selection. Clones or pools are tested by Southern blot, northern blot, qRT-PCR, protein expression, etc… as appropriate, to confirm presence of transfected DNA construct.

3. Notes and Limitations Best results are obtained using cells that have been maintained in log-phase for several generations, achieved by daily 1:5–1:10 dilutions. Transient transfection efficiencies range from 25 to 80%, depending on the reporter, for example, tubulin dsRNA constructs provide a more sensitive readout than GFP expression. Stable transfection efficiencies are in the range reported by others (Burkard et al., 2007), that is, 103104. Electroporation with instruments other than the BioRad Gene Pulser II® are also effective, though optimal conditions should be determined empirically.

4. BSF Transfection Cells in mid log phase (5 1053 106 cells/ml) are harvested and washed in “EM,” then resuspended at 2 107 cells/ml. A 0.45 ml aliquot of cells (0.9 107 cells) is added

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to a 0.4 cm BioRad® electroporation cuvette containing 100 µl of linearized DNA (20 ug) in water. Cells and DNA are mixed by gently pipeting up and down, avoiding air bubbles. Electroporation is done with one pulse in a Gene Pulser II (BioRad®) electroporator using the following settings: 1500 V, 25 µF. The time constant should be between 0.5 and 0.7 ms. Following electroporation, cells are immediately transferred into 12 ml prewarmed culture medium and aliquotted (0.5 ml) into a 24-well culture plate. Following an overnight incubation, antibiotic for selection is added by adding 0.5 ml of culture medium containing 2 the amount of the required antibiotic (Table II) to each well. Monitor cells daily and dilute as necessary in presence of drug until mock transfected samples die out and transfected cells are growing well, generally 5–7 days post-transfection. In general, one can expect to see 1–5 wells from a 24-well plate grow out. Each well represents a “pool” of clones, which can then be isolated by limiting dilution. Note that since the stable transfection efficiency is much lower than for PCF cells, cells in each “pool” are quite likely clonal. Clones or pools are tested by Southern blot, northern blot, qRT-PCR, protein expression, etc… as appropriate, to confirm presence of transfected DNA construct. The efficiency for stable transfection using this protocol is 106107. We have not had experience with transient transfection of BSF cells. C. Protocols for Flagellum Skeleton Isolation The trypanosome flagellum is laterally attached to the cell body along most of its length (Fig. 1), making isolation of intact, that is, membrane-enclosed, flagella challenging. Progress is being made employing the Fla1 mutant (LaCount et al., 2002) in which the lateral connections are disrupted, but current analyses have been restricted to detergent- and salt-extracted samples lacking the flagellar membrane and soluble matrix (Robinson et al., 1991). For the current discussion, we refer to such preparations as “flagellum skeletons,” which include the axoneme, PFR, basal body, and some components of the flagellum attachment zone. A simple and reliable two-step preparation for flagellum skeletons has been published previously (Robinson et al., 1991; Schneider et al., 1987) and is widely utilized (e.g., Broadhead et al., 2006; Hart et al., 2009; Hill et al., 2000; Portman et al., 2009). Briefly, this method consists of two basic steps: (i) Non-ionic detergent extraction of whole cells to isolate the cytoskeleton, including flagellum skeleton; followed by (ii) high salt, for example, 0.5–1.0 M NaCl, extraction on ice to depolymerize subpellicular microtubules, leaving the flagellar skeleton intact. In some cases, subpellicular microtubules are depolymerized with 1 mM Ca2þ instead of 1 M NaCl (Robinson et al., 1991). At each step insoluble material, that is, cytoskeleton or flagellum skeleton, is separated from solubilized material by centrifugation. Treatment with DNase removes the mitochondrial genome associated with the flagellar basal body, and greatly improves the solubility of isolated flagellum skeletons. What remains is the flagellum skeleton, containing extracted axoneme, PFR, basal body, and flagellum attachment zone components (Robinson et al., 1991). Two independent proteomic analyses of such extracted flagella identified a total of 524 potential flagellar proteins (Broadhead et al., 2006; Hart et al., 2009).

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(A) 0.5 M NaCl

(B) 0.15 M NaCl

Fig. 2 T. brucei flagellum skeleton. Cross-section of detergent and salt-extracted flagellar skeletons imaged by transmission electron microcopy. (A) Extraction performed using 0.5 M NaCl, (B) Extraction performed using 0.15 M NaCl.

As with other organisms (Dutcher et al., 1984) high salt extraction, for example, 0.5 M NaCl and above, perturbs flagellum ultrastructure in T. brucei (Fig. 2A). Commonly outer dyneins are missing and central pair microtubules are compromised. Modification of the procedure to reduce salt concentration and employ a single extraction step yields flagellum skeletons with improved ultrastructure (Fig. 2B). For this one-step preparation, trypanosomes are harvested at 2000 g for 5 min, washed in PBS, then resuspended at 2 108 cells/ml in cold PMN buffer [10 mM NaPO4 (pH7.2), 1 mM MgCl2, and 150 mM NaCl) (Robinson et al., 1991] containing 1% NP-40, DNAse, and protease inhibitors. The sample is incubated on ice for 10 min to depolymerize subpellicular microtubules. Addition of Ca2þ (Robinson et al., 1991) sometimes resulted in buffer precipitation and was therefore omitted. Intact flagellum skeletons are harvested by centrifugation at 16,000 g for 10 min, the supernatant is aspirated and flagellar skeletons are washed in cold PMN as above. This sample can be used for biochemical, structural, cell biological or proteomic, etc… analyses. Proteomic analysis of the resulting improved flagellum skeleton preparation identified 868 putative flagellar proteins (KH, unpublished), including 76.5% of the proteins identified in earlier analyses.

1. Notes and Limitations The one-step, low salt preparation above generally yields samples with improved ultrastructure versus high-salt extraction, but there are still samples in the preparation

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with structural defects. Hence, there is still room for improvement. Coupling flagellum preparation with mutants provides a powerful approach for identification of proteins making up specific flagellum substructures. For example, the protein composition of the PFR was partially determined using an elegant approach that employs quantitative proteomic analysis (iTRAQ and 2D DIGE) of detergent extracted flagella from control cells and mutants lacking the PFR (Portman et al., 2009).

D. Assays for Trypanosome Motility A variety of assays are available to assess trypanosome motility, either at the single cell or population level. These assays provide readouts for studying the effects of knockdown, knockout, or mutation of flagellar proteins.

1. Sedimentation Assay Motility of wild-type PCF trypanosomes enables them to remain relatively evenly distributed in suspension cultures over time, while motility mutants sediment (Fig. 3A). This provides for a convenient motility assay simply by measuring the optical density at 600 nm (OD600) over time (Bastin et al., 1999; Branche et al., 2006; Ralston et al., 2006). For sedimentation assays, a culture grown to a density of 5 106 cells/ml is aliquotted into four cuvettes and incubated under standard growth conditions. We use 1 ml size BioRad® disposable cuvettes (BioRad® part # 223-9955). The optical density at 600 nm (OD600) is then measured every 2 h for 12 h. In order to distinguish a motility defect from a growth defect, two cuvettes are left undisturbed, whereas the other two cuvettes are resuspended at each time point. The DOD600 is calculated by subtracting the OD600 of the resuspended samples from that of the undisturbed samples and plotted versus time. This DOD600 plot is then compared to the control population. An example of the assay is given in Fig. 3B (adapted from Ralston et al., 2006). Sedimentation can also be done reiteratively to enrich for motility mutants (Section III.E.3).

2. Notes and Limitations Care should be taken to keep test and control samples at equivalent cell densities, because motility can change as a function of cell density. In our hands, BSF cells do not generally remain evenly suspended, so sedimentation assays have not worked well for assessing motility of this life cycle stage. Note that the sedimentation assay provides a convenient and quantitative assessment of motility, but applies only populations and is “low-resolution.” Mutants with sedimentation phenotypes are therefore followed through more detailed analyses by video microscopy (see below).

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(A)

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Fig. 3 Assays for trypanosome motility. (A) Cartoon illustration depicting sedimentation assays. Cells with wild type motility remain relatively evenly suspended (on left), where as motility mutants sediment (on right). Degree of sedimentation is quantitated using OD600 (arrow). (B) Example dataset derived from a sedimentation assay (adapted from Ralston et al., 2006). Depletion of RSP3 by Tet-inducible RNAi (RSP3 þ Tet) cause cells to sediment, while uninduced cells, expressing wt level of RSP3 (RSP3-Tet) remain suspended over time. Adapted from Ralston et al. (2006), with permission. (C) Illustration depicting a motility chamber. (D) Example dataset derived from a motility trace assay (adapted from Hutchings et al., 2002). Open circles represent cell position at 1 s intervals, arrowheads indicate ending position. Uninduced cells, expressing wt level of Trypanin (-Tet) exhibit robust motility, whereas Trypain depleted cells (þTet) fail to exhibit productive forward translocation. Respective western blots (WB) against Trypanin or tubulin as a loading control show protein levels. Adapted from Hutchings et al. (2002), with permission.

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(C)

Cover glass Microscope slide Tape (double sided) (D)

(17s) (25s) (25s)

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3. Motility Traces A second method for assessing motility is to track movement of individual cells in a population (Baron et al., 2007; Bastin et al., 1999; Broadhead et al., 2006; Gadelha et al., 2007; Hutchings et al., 2002). This method employs a motility chamber for digital video microscopy analysis (Fig. 3C) (Baron et al., 2007). Each motility chamber is comprised

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33

of a glass slide and glass coverslip held together by 50-µm-thick (approximately two cell lengths) double-sided tape. To prevent cell adherence, the slide and coverslip are precoated in 0.25% poly-L-glutamate (Gadelha et al., 2007) as described here: (1) Clean slide and coverslip with 6 N nitric acid for 10 min. (2) Rinse thoroughly with double-distilled water, rinse with ethanol (abs.), and dry using compressed gas duster. (3) Coat slide and coverslip with poly-L-glutamic acid (poly-E) sodium salt (cat. no. P4886, Sigma-Aldrich, St. Lous, MO) in PBS. Incubate for 20 min. (4) Rinse off excess poly-E with doubledistilled water, rinse with ethanol (abs.), and dry using compressed gas duster. To mimimize capillary flow, the edges of the coverslip are sealed with Vaseline® following the application of cells. Cells are grown to a density of 5 106 cells/ml and resuspended into fresh, prewarmed medium at a final concentration of 1 106 cells/ml. Next, the cells are applied to motility chamber and viewed under dark-field illumination, using a 10 objective. Generally, 30 s of digital video from separate regions on each slide are captured for PCF and 2 min for BSF. For a camera, typically any good quality hand-held video camera will suffice and can be mounted directly to the microscope light path using appropriate adaptors available from the microscope vendor. Digital video is captured directly to the computer hard drive using third-party video capture software, for example, Adobe Photoshop Premier Elements®. Digital videos, either as avi files or as image stacks, are then analyzed using motility tracking software. We have used Metamorph® from Molecular Devices, CA, USA (Baron et al., 2007), which offers a robust suite of programs that can be tailored to individual preferences for motility analyses. Cell movements can be readily quantitated and traces generated automatically (Fig. 3D, adapted from Hutchings et al., 2002). Cell movements are analyzed for average velocity, burst velocity, distance migrated, and directionality of each cell within a video, etc…

4. Video Microscopy Analysis of Flagellar Beat To characterize subtleties of flagellar beating and more rigorously describe motility phenotypes, video-microscopy of individual cells at high magnification is employed. Cells are placed in a motility chamber (see above) and imaged using 63 or 100 oil-immersion objectives. The use of differential-interference contrast (DIC) optics aids in viewing of the flagellum. In some cases, it can be useful to image at high resolution using dark field optics, which enhances outline features of the cell. Alternatively, cells can be imaged directly in the tissue culture flask using a 60 long distance objective. Depending on the optics of the microscope used, this approach can give very highresolution and high-contrast videos even with out oil-immersion objectives or DIC optics (see, e.g., http://www.elsevierdirect.com/companions/9780123749734, Supplementary Movie 1, from Ralston et al. 2006, with permission). Flagellum movements can further be quantified via video microscopy (Branche et al., 2006; Ralston et al., 2006). This approach provides a high-resolution means whereby the beat defects of individual mutants can be quantitatively assessed. Additionally, through the use of this approach, one can distinguish the severity in paralysis between different motility mutants and can also describe phenotypes characteristic to more severe motility defects.

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5. Notes and Limitations Although the above-mentioned strategies offer a way by which to characterize motility defects at an individual cell level, they are not without limitations. To begin with, although video microscopy captured at conventional speeds (30 fps) can effectively yield information about the gross number of beats carried out by a flagellum, the specific mechanics of that beat remain unappreciated in T. brucei, owing to highly erratic and three-dimensional beating of the flagellum, together with the challenges posed by lateral attachment of the flagellum to the cell body. In other kinetoplastids, the use of high-speed imaging has yielded novel data sets that have been used to more rigorously describe the mechanisms driving biological functions (Gadelha et al., 2007; Sugrue et al., 1988). A second concern is the fact that the flagellar beat in T. brucei is three-dimensional and bending can occur independently in the distal and proximal portions of the flagellum, it becomes impossible to reliably measure the frequency and amplitude of a single waveform along the length of the entire flagellum. Lastly, when conducting motility assays, one should take care to be aware of cell density, as cells swim faster when entering stationary phase and appear to be slower when in cytokinesis (examples of application and related references: See references in text).

E. Genetic Approaches to Study Individual Components of the T. brucei Flagellum Molecular genetic tools based on homologous recombination, controllable promoters, RNAi, and a suite of plasmid systems combine to provide complementary approaches for investigating flagellum protein function in T. brucei. Here we outline these approaches, which employ the simple and straightforward methodologies presented in earlier sections. Where relevant, we list a few published examples, though this is not a comprehensive listing.

1. Genetic Tools Based on Homologous Recombination African trypanosomes exhibit highly efficient homologous recombination and this is exploited to target DNA constructs to specific sites in the parasite genome (Clayton, 1999; Lee and Van der Ploeg, 1990; Li et al., 1996; Zomerdijk et al., 1990). The approach can be used to “knockout” a target gene, or to introduce an altered transgene at its endogenous locus, that is, “knock-in.” The altered transgene carries specific point mutations or an epitope tag. Recombination efficiency varies depending on the length of the homology region and is generally higher in procyclic PCF form versus BSF cells. Homology regions can be as short as 40– 80 bps (Arhin et al., 2004; Oberholzer et al., 2006; Shen et al., 2001), although the length required varies from locus to locus and a few hundred base pairs are generally used for increased reliability.

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a. Gene Disruption by Homologous Recombination Constructs for gene deletion are constructed by flanking a selectable marker in a plasmid vector with sequences corresponding to the 50 untranslated region (UTR) and 30 UTR of the targeted gene. This construct is than excised from the plasmid backbone by restriction digest and stably transfected into trypanosomes (Section III.B). The integrated construct employs endogenous expression signals, for example, splice acceptor and poly-adenylation. In PCF trypanosomes, where stable transfection efficiency is highest, constructs for gene disruption can be generated by a fast, one-step PCR-based approach using long (~100 bp) primers containing the homology region in their 50 ends (Arhin et al., 2004). Efficiency of integration into the targeted locus is reduced relative to employing longer flanking sequences and the choice of using the PCR-based or classical, cloning-based approach is decided on a case-by-case basis. Disruption of multiple genes by homologous recombination is limited to the available resistance markers. The recent development of Cre-lox system for trypanosomes opens new possibilities for the generation of novel selection markers and multiple gene deletions in the same cell line (Scahill et al., 2008). Examples of application and related references: One-step PCR-based disruption of genes (Arhin et al., 2004) Multiple gene disruption in the same cell line (Haenni et al., 2006; Vassella et al., 2009) Cre-lox system for T. brucei (Scahill et al., 2008) b. In Situ Tagging of Genes by a “knock-in” Approach Homologous recombination is also exploited to introduce small epitope or fluorescent protein tags at endogenous chromosomal loci, a procedure referred to as “in situ tagging” (Fig. 4) (Kelly et al., 2007; Oberholzer et al., 2006; Schimanski et al., 2005; Shen et al., 2001). The epitope or fluorescent protein tag can be introduced at either end of the target gene by choosing the appropriate “tagging cassette,” consisting of a tag, a spacer DNA element (intergenic region, igr) and an antibiotic resistance marker for selection (Fig. 4 and http://www.elsevierdirect.com/companions/9780123749734, Supplementary Fig. 1). The tagging cassette is flanked by homology regions, for homologous recombination into the target gene locus, using one of two approaches: either a two-step cloning procedure or a single-step PCR amplification (Fig. 4). Both approaches employ the same set of pMOTag tagging plasmids (Oberholzer et al., 2006), which contain a tag, selectable marker, intergenic sequences to direct RNA processing, and multicloning sites (Fig. 4). For C-terminal tagging, the 30 end of the target open reading frame (without stop codon) is fused in-frame to the 50 end of the tag and the target gene’s 30 UTR is cloned downstream of the drug resistance marker (Fig. 4). For N-terminal tagging, the target gene’s 50 UTR is fused upstream of the drug resistance marker and the 50 end of the target open reading frame is fused in frame to the 30 end of the tag (Kelly et al., 2007; Schimanski et al., 2005; Shen et al., 2001) (Suppl. Fig. 1). The tagging cassette, either excised from the plasmid backbone using restriction enzymes or obtained as a PCR product, is stably transfected into recipient cells as outlined in Section III.B. Resulting tagged lines are analyzed as desired. Generally, a single copy

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MCS tag

igr

DrugR

MCS

pMOTag

PCR or cloning

tag

igr

DrugR

Integration

5’UTR

ORF

5’UTR

ORF

wt

3’UTR tag

DrugR

igr

3’UTR

tagged

Protein expression

N

protein

Forward tagging primer (homologous to target ORF)

tag

igr

C

Intergenic region of alpha-beta tubulin

Reverse tagging primer (homologous to target 3’ UTR)

5’UTR

5’ untranslated region of target gene

tag

Tag with in frame stop codon

3’UTR

3’ untranslated region of target gene

DrugR

Antibiotic resistance cassette

ORF

Open reading frame of target gene

MCS

Multicloning site

Fig. 4 Schematic Illustration outlining procedure for C-terminal in situ tagging of T. brucei genes via “knock-in” approach using the pMOTag vector series (Oberholzer et al., 2006). The tagging-integration construct is generated using PCR with chimeric primers including target gene ORF (blue) or 30 UTR (gray) at the 50 end and pMOTag sequences at the 30 prime end. Alternatively, the integration construct is generated by cloning ORF and UTR sequences into the pMOTag multicloning sites (MCS). The tagging-integration construct is integrated at the endogenous chromosomal locus by homologous recombination. Analogous vectors are used for tagging at the N-terminus (Suppl. Fig. 1).

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of the tagging cassette integrates into one allele of the target gene. If desired, tagging of both alleles or a different gene is easily accomplished by employing an additional selectable drug marker and selecting under both drugs (Zhao et al., 2008). The genomic landscape at the site of integration and the number of integration events can be verified by PCR and Southern blot analysis. A wide variety of epitope or fluorescent protein tags are available (Table II), including tags for protein visualization in fixed or live cells, tandem affinity purification, or analysis of protein–protein interaction. DNA elements encoding different epitopes have been synthesized for optimum codon usage in trypanosomes (HA, c-Myc, Flag; Oberholzer et al., 2006). The most widely used epitope tags are hemagglutinin HA, c-Myc, and TY1 tag for protein localization and PTP tag for affinity purification of protein complexes (Table I). Most of the antibodies used for the detection of these tags are commercially available, although some cross-react with endogenous trypanosome proteins in western blotting and immunofluorescence microcopy (see Table I). The highest signal intensity with minimal cross-reactivity has been observed when using the 3 c-Myc tag in combination with the monoclonal antibody 9E10 (Santa Cruz) (Oberholzer et al., 2006; Cross lab, unpublished). Besides small epitope tags, various fluorescent proteins (GFP, eYFP, mCherry, …) are widely used for in situ tagging of genes in trypanosomes (Kelly et al., 2007; Li et al., 2008; Umeyama and Wang, 2008). A list of published and unpublished vectors for in situ tagging of genes via the use of various epitope and fluorescent protein tags is provided in the supplemental Material (http://www.elsevierdirect.com/companions/9780123749734, Supplementary Table 1I). Other in situ Tagging Systems. The in situ tagging system developed by the Tschudi lab (Shen et al., 2001) as well as the pMOTag vectors (Oberholzer et al., 2006) discussed above employ two regions of homology to direct integration at a specific locus. An alternate in situ tagging strategy uses a single region of homology to direct integration (Kelly et al., 2007; Schimanski et al., 2005). The advantage of this second system is that only one homology region has to be cloned, thus simplifying the cloning approach. The disadvantages are as follows: (i) a truncated version of the tagged gene is left behind in the genome and (ii) the plasmid backbone also integrates into the genome. This approach thus generates cell lines with one wild-type allele, one tagged allele, and a third truncated allele, which might cause unforeseen complications. Nonetheless, this tagging system has been extremely successful and has been utilized in several instances (Nguyen et al., 2006; Portman et al., 2009; Schimanski et al., 2005; Umeyama and Wang, 2008), particularly for incorporating tags that enable tandem affinity purification. Considerations and Limitations. In situ tagging has two main advantages over traditional, plasmid-based tagging methods. First, because the tagged copy is under the control of the endogenous promoter, one avoids several of the potential risks associated with expressing an extra copy of a gene from a heterologous promoter. Second, large quantities of the tagging construct can be generated via a single-step PCR, thus

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Table I Most Commonly Used Epitope Tags for In Situ Tagging of T. brucei Genes Tag HA 3 HA

Tag sequence (*C-terminal stop)

Antibody (source)

Antibody specificity

Selected references

YPYDVPDYA* YPYDVPDYAYPYDVPDYAYPYDVPDYA*

HA.11 monoclonal mouse (Covance) HA Y-11 polyclonal rabbit (Santa Cruz) HA monoclonal mouse 12CA5 (Roche)

Highly specific in WB and IF

Oberholzer et al., 2007b

Background in WB and IF. Needs pre-absorbtion Background in WB and IF

Siegel et al. Genes and Development, 2009 Zamudio et al., 2009 Umeyama and Wang, 2008

EQKLISEEDL* GGRSRSEEQKLISEEDLLRSE QKLISEDLLRSEEQKLISEEDLL*

Anti-c-Myc monoclonal mouse 9E10 (Santa Cruz, Biotechnology)

Highly specific in WB and IF

Zhao et al., 2008 Oberholzer et al., 2007b Kunz et al., 2009

TY1

EVHTNQDPLD*

BB2, monoclonal mouse (Prof. K. Gull, Oxford)

Highly specific in WB and IF

Bastin et al., 1996 Portman et al., 2009 Lowell et al., 2004

PTP

ProtC-TEV-ProtA-ProtA

Anti-ProtC monoclonal HPC4 (Roche), ProtA-specific PAP reagent (Sigma-Aldrich)

Schimanski et al., 2005 Zamudio et al., 2009 Nguyen et al., 2006

Michael Oberholzer et al.

c-Myc 3 c-Myc

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circumventing multiple sublconing steps and expediting the tagging process. Replacement of the target gene’s endogenous 30 UTR by the intergenic region of the a–b tubulin locus might alter expression level relative to endogenous levels and this should be taken into consideration. Gene expression through the life cycle of trypanosomes is regulated largely at the post-transcriptional level (e.g., Urwyler et al., 2007); therefore care should be taken to understand the developmentally regulated expression profile of the gene targeted for in situ tagging. Next-generation tagging cassettes may overcome this limitation by taking advantage of the CRE recombinase-based system developed for T. brucei (Barrett et al., 2004; Scahill et al., 2008). In this case, loxP sites flanking the igr and selectable marker of the tagging cassette allow these sequences to be excised, placing the target gene back under control of the endogenous UTR. Such a in situ tagging strategy would introduce a tag at the 30 end of a gene leaving only a LoxP-site, constituting a 34 bp “scar” between the stop codon and the endogenous 30 UTR. The CRE recombinase system was integrated into the pMOTag in situ tagging vectors series by the research group of Prof. G. Cross, but its use remains to be tested (Cross lab homepage for vector details, unpublished). Examples of applications and related references: Vectors for in situ tagging of T. brucei genes (Kelly et al., 2007; Oberholzer et al., 2006; Shen et al., 2001) In situ tagging of T. brucei genes with epitope tags (Li et al., 2008; Oberholzer et al., 2007b; Portman et al., 2009; Zhao et al., 2008) In situ tagging of T. brucei genes with fluorescent protein tags (Li et al., 2008; Umeyama and Wang, 2008) In situ tagging of T. brucei genes with PTP tag for affinity purification of protein complexes (Nguyen et al., 2006; Schimanski et al., 2005; Zamudio et al., 2009)

2. Genetic Tools Based on Controllable Promoters T. brucei offers a variety of integrble plasmids for constitutive and tetracyclineinducible expression of genes and RNAi constructs (LaCount et al., 2002; Wang et al., 2000; Wirtz and Clayton, 1995; Wirtz et al., 1999). Tet-inducible systems are particularly useful for studying protein localization as a function of cell cycle, analyzing proteins harboring specific point mutations (see Section III.E.4), and for investigating function of essential genes (Broadhead et al., 2006; LaCount et al., 2002; Oberholzer et al., 2007b; Ralston et al., 2006). The key for the Tet-regulated system is a modified cell line, which expresses a Tet repressor and a T7 polymerase (Wirtz et al., 1999). Construction of this cell line was keystone development for functional analysis in T. brucei, with the initial publication (Wirtz et al., 1999) being cited 382 times in the Pubmed database as of July 2009. a. Plasmids for Inducible Expression Plasmid details vary, but they all have three key features: (1) a Tet operator sequence to control the transgene promoter; (2) a drug resistance marker to enable selection of stable transfectants; (3) sequences

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homologous to a silent chromosomal region to allow stable integration. The selectable marker is either located downstream from the inducible promoter and transcribed with the transgene (tandem arrangement), or oriented divergently from the Tet-inducible transgene and transcribed by a separate, constitutive promoter (Fig. 5) (Wirtz et al., 1999). Divergent promoters are generally preferred, as this arrangement enables more complete silencing of the transgene in the absence of tetracycline. For protein expression, the plasmid backbone includes 50 and 30 untranslated regions to direct posttransciptional processing and translation. The constructs are targeted to transcriptionally silent loci to ensure no transgene expression in the absence of tetracycline. Most frequently the spacer region of the rDNA locus or the silent regions of the minichromosomes are used (Wickstead et al., 2002; Wirtz et al., 1999). Two widely used vectors that served as templates, physically and conceptually, for the various inducible systems are pLEW100 and pLEW82 (Wirtz et al., 1999). Their overall structure and modules serve as an example for Tet-inducible expression systems (Fig. 5). Sequences of the vectors are accessible on the homepage of the research group of Prof. G. Cross (http://tryps.rockefeller.edu/). Considerations and Limitations. of a transgene from ectopic sites.

Several factors contribute to the expression level

(i) The design of the vector used for expression, including promoter identity, promoter orientation, and nature of the UTR sequences flanking the transgene. Any given construct might also be subject to life cycle stage-specific regulation, and hence this should be considered when choosing the recipient cell line. (ii) The nature of the protein expressed from the construct can also influence the final expression levels. Namely, in our experience cells are able to more readily accommodate high expression levels of proteins that are endogenously highly expressed. For example cells expressing GFP-tagged paraflagellar rod 2 protein (PFR2) (Schlaeppi et al., 1989) can be readily visualized in live fluorescent video microscopy (Fig. 5B and http://www.elsevierdirect.com/companions/ 9780123749734, Supplementary Movie 2, MO unpublished), whereas less abundant proteins, such as trypanin-GFP or CMF19-GFP (two flagellar proteins) require exposure times in excess of 1 s (Baron et al., 2007), despite using the same expression vector pLEW100, same cell-line and same induction conditions. A limitation of Tet-regulated expression system can be leakiness of the vector in the uninduced state. The extent of leakiness varies and has not been systematically studied for a large number of genes, though it would primarily only be problematic for expression of toxic gene products or dsRNA. Examples of application and related references: Cell line expressing Tet repressor and T7 polymerase. Describing of pLEW100 and pLEW82 vectors (Wirtz et al., 1999) Selected applications of pLEW100 and pLEW82 (Baron et al., 2007; Boothroyd et al., 2009; Grandgenett et al., 2007; Selvapandiyan et al., 2007; Umeyama and Wang, 2008)

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b. Plasmids for Inducible RNA Interference (RNAi) T. brucei is exceptionally well suited to analyze gene function using RNA interference (RNAi), as first demonstrated by the research group of Prof. E. Ullu (Ngo et al., 1998) and subsequently used to tremendous success (e.g., LaCount and Donelson, 2001; Motyka and Englund, 2004; Ullu et al., 2004; Wang et al., 2000). RNAi in T. brucei is potent, specific and can be achieved with long or short dsRNA triggers

(A) NotI, EcorV

NotI, EcorV

MCS MCS

rDNA spacer

Drug

R

rDNA spacer

NotI, EcorV

NotI, EcorV

Drug

rDNA spacer

R

Tet operator controlled PARP promoter

MCS MCS

rDNA spacer

rDNA spacer

3’ processing element

Tet operator controlled T7 polymerse promoter

5’ processing element

Drug

R

PLEW82

Non-transcribed spacer of the rDNA locus

T7 polymerase promoter

T7 polymerase terminater

MCS MCS

pLEW100

Antibiotic resistance cassette

Multicloning site

Fig. 5 Plasmids for inducible gene expression. (A) Illustration of two vectors (pLEW100 and pLEW82) (Wirtz et al. 1999), commonly used for inducible expression in T. brucei. (B) Example of a PCF cell line expressing a GFP-tagged flagellum protein (PFR2) through the use of pLEW100. Top panels, fluorescence image of a Tet induced live cell. On bottom, series of images showing the movement of the GFP-PFR2 expressing cell line under fluorescence microscopy (see Suppl. Movie 2, MO unpublished). Cells are costained with a live DNA dye, CFSE to visualize nucleus and kinetoplast. Movie is recorded using a 63 objective (recorded and played back at 30 fps). Timestamps given in ms.

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(B)

overlay

GFP

0

33

66

99

132

165

198

231

264

297

Fig. 5 (Continued) (See Plate no. 1 in the Color Plate Section.)

(Ngo et al., 1998). The T. brucei genome is sequenced, lack of introns makes gene identification simple and reliable, and web-based algorithms provide for automated identification and design of primers to amplify the appropriate nonredundant sequences in any target gene (Redmond et al., 2003). These features, coupled with the tightly regulated Tet-inducible promoter systems described above, allow for facile construction of cell lines in which target gene expression is ablated by heritable and inducible RNAi (Bastin et al., 2000; LaCount et al., 2000; LaCount et al., 2002; Shi et al., 2000; Wang et al., 2000). RNAi knockdown of an endogenous mRNA requires production of dsRNA homologous to a portion of the target mRNA (Bosher and Labouesse, 2000). In T. brucei, this is commonly achieved by placing a single copy of the target between opposing, “head-to-head,” promoters (Fig. 6A) (LaCount et al., 2000; Wang et al., 2000). Alternatively, inverted copies of the target sequence are placed downstream of a single promoter (Bastin et al., 2000; Hutchings et al., 2002; Oberholzer et al., 2007b). It is

43

2. Approaches for Functional Analysis

(A)

5’ target

3’ NotI, EcorV

NotI, EcorV

DrugR

MCS

rDNA spacer

rDNA spacer

p2T7

+ Tet

3’ 5’

5’ 3’

dsRNA

RNAi mediated degradation of target mRNA

(B)

DNA library NotI, EcorV

NotI, EcorV

DrugR

MCS

rDNA spacer

rDNA spacer

pZJM RNAi libary

+ Tet

5’ 3’

3’ 5’

dsRNA

RNAi mediated degradation of target mRNA

Ribosomal RNA promoter (red), T7 polymerase promoter (blue)

Mutant screen

rDNA spacer Non-transcribed spacer of the rDNA locus

3’ processing element

“Head-to-head” Tet operator controlled T7 polymerse promoters

5’ processing element

T7 polymerase terminator MCS

Multicloning site

DrugR

Antibiotic resistance cassette

Fig. 6 Inducible RNA interference (RNAi) in T. brucei. (A) Illustration of a widely used inducible RNAi vector (p2T7), LaCount et al. (2002). (B) Illustration of a forward genetic screen through the use of a Tetinducible RNAi library, Wang et al. (2002). See text for details.

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also possible to simultaneously knockdown two or more genes by cloning the target sequences in tandem between opposing promoters, thereby allowing for assessment of genetic interactions (Ralston et al., 2006). Details on individual systems for generating dsRNA to induce RNAi in trypanosomes are reviewed elsewhere (LaCount and Donelson, 2001; Ullu et al., 2002). The head-to-head system is used most commonly, owing to simplicity of cloning. The vectors used most commonly for head-to-head systems are p2T7 and pZJM (LaCount et al., 2002; Wang et al., 2000). Each contains a multicloning site between opposing, tetracycline-inducible promoters, together with a selectable marker and sequences to direct homologous integration at the silent rRNA spacer locus. Appropriate target sequence and primers are identified using the online web-based RNAit application (Redmond et al., 2003), then the target sequence is PCRamplified and inserted into the multicloning site using standard methods. The resulting plasmid is linearized within the rDNA homology sequence and transfected into PCF or BSF trypanosomes as described above. Considerations and Limitations. Tet-inducible RNAi in T. brucei is robust and powerful and has been utilized successfully in hundreds of applications. A potential limitation can be that the RNAi construct may be “leaky” in the uniduced state, therefore expressing a small amount of dsRNA without induction by Tet. This can be problematic when targeting an essential gene, although it has not posed a significant limitation in practice, and selective pressures enrich for more completely silenced constructs. RNAi over extended time periods, for example, more than a week, should be monitored closely, as this can select for cells that downregulate the RNAi machinery. In large part, this limitation is overcome by utilizing inducible rather than constitutive promoters to drive expression of the RNAi construct. By definition, RNAi phenotypes result from loss of the target protein. Hence, if the target protein is part of a complex, as are most flagellar proteins, one must consider whether the entire complex might be destabilized and, if so, whether the phenotype results from loss of the target protein per se, or loss of the entire complex. Recent developments (see Section III.E.4) provide approaches to overcome this limitation through heterologous expression of the RNAi target gene harboring point mutations, thereby disrupting function without altering protein expression and assembly. Examples of application and related references: Description of p2T7 and pZJM RNAi vectors (LaCount et al., 2000; LaCount et al., 2002; Wang et al., 2000) Selected applications of inducible RNAi (Absalon et al., 2008; Jetton et al., 2009; Oberholzer et al., 2007b; Ralston and Hill, 2006; Zhao et al., 2008)

3. Forward Genetic Screens using RNAi Libraries Forward genetic screens are technically challenging to perform in trypanosomes, owing primarily to the fact that experimentally accessible life cycle stages are diploid. Recently, the research group of Paul Englund described a novel

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45

method for conducting forward genetic screens in T. brucei (Englund et al., 2005; Morris et al., 2002). This method employs a tetracycline-inducible RNAi library that contains random T. brucei DNA fragments between two opposing, tetracycline-inducible promoters in a plasmid that can be stably integrated into the genome (Fig. 6B). The library was introduced into PCF 29–13 cells to provide approximately five-fold genome coverage (Morris et al., 2002). Induction with Tet produces a distinct dsRNA in each cell that downregulates expression of the target gene. From this heterogeneous population of “mutated” (tetracycline induced) cells, mutants are isolated by screening for the desired phenotype (e.g., Morris et al., 2002). Alternatively, the library can be sorted and individual clones induced and screened independently (e.g., Zhao et al., 2008). The target gene is identified by PCR amplification using primers flanking the DNA insert in the RNAi plasmid, and then tested independently. The dominant nature of RNAi circumvents limitations posed by diploidy in trypanosomes and the RNAi plasmid provides a simple and rapid means of identifying the target gene. The approach has been successfully used to identify genes regulating surface protein expression, targets of the trypanosome drug tubericidine, proteins linking the kinetoplast to the flagellum basal body and regulators of the cell cycle (Drew et al., 2003; Monnerat et al., 2009; Morris et al., 2002; Zhao et al., 2008). Although RNAi library screening has not yet been applied for isolation of motility mutants, the approach is readily adaptable to motility mutant screens, for example, through reiterative rounds of sedimentation (Section III.D) to enrich for motility mutants. Forward genetic screens employing transposon mutagenesis have also been established (Leal et al., 2004). While promising, this methodology has yet to be implemented widely and is not discussed here. a. Considerations and Limitations Currently, the RNAi library approach has only been used in the PCF life cycle stage. Until recently, BSF trypanosome transfection efficiencies were too low to enable construction of a library with sufficient coverage. However, the recent development of a novel high-efficiency transfection protocol for BSF trypanosomes (Burkard et al., 2007), see Section III.B, now make it feasible to generate RNAi libraries in BSF cells. As always, coverage of the library needs to be considered large or multicopy genes are biasedly screened over smaller, single copy genes. This is due to the fact that larger genes, as well as multicopy genes, are represented with greater frequency as a consequence of the methodology employed to generate the RNAi library. As the approach becomes more widely utilized, new and improved libraries will evolve. As with any library-based approach, the extent of genome coverage is based on the quality of the library and there is also a risk of losing library complexity the more it is amplified. Likewise, as with any mutant screen, care must be put into the development and implementation of the screening strategy, as this largely determines the effectiveness of the screen. A potential limitation of the approach is that the RNAi vector can integrate in a genomic locus other than the intended rDNA locus, leading to unexpected mis-

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regulation of flanking genes (Motyka et al., 2004), particularly if the target fragment happens to contain a site for the restriction enzyme used to linearize the plasmid. For this reason, each target gene identified should be tested independently to confirm the phenotype. Note that even in the event of apparent mis-integration, the mis-regulated gene(s) responsible for the phenotype can be identified by careful analysis of the mutant through restriction mapping and PCR to identify the site of vector integration, combined with northern blotting or qRTPCR to assess flanking gene expression levels (Motyka et al., 2004). Examples of application and related references: Description and applications of the pZJM RNAi library (Drew et al., 2003; Monnerat et al., 2009; Morris et al., 2002; Zhao et al., 2008)

4. Combination of RNAi and Heterologous Expression for Structure–Function Studies An inherent limitation to simple RNAi or gene knockout approaches to assess gene function is that the phenotypes observed reflect loss of the target protein. If the protein is part of a complex, as is generally the case for flagellar proteins, this can cause loss of other subunits or the entire complex, thereby confounding or limiting interpretation. Such cases provide validation of flagellum protein identification and offer information about proteins required for assembly of flagellum sub-structures, but offer little or no information about specific functions of the target protein. Indeed, despite the expansion of inventories of flagellar proteins through genetic, genomic, and proteomic studies, knowledge about how these proteins function and details on key amino acids and domains is severely limited. In order to fill this gap in knowledge of flagellum biology, we recently developed an inducible RNAi-based system for structure–function analysis of T. brucei flagellar proteins (Ralston and Hill, unpublished). Details on the system and its application will be published elsewhere, so we provide an outline of the approach here. The system employs dual vectors to allow for inducible knockdown of an endogenous gene, with simultaneous induction of a heterologous copy harboring specific point mutations (Fig. 7). The “knockdown vector” targets the 30 UTR of the endogenous gene so that the coding sequence can still be expressed in the RNAi background. The heterologous copy of the target protein carries a different 30 UTR so that it is not targeted for knockdown, and an epitope tag to distinguish it from the endogenous protein. Using this system, we identified amino acids necessary for function of the outer arm dynein light chain 1 (LC1) protein (Ralston and Hill, unpublished). a. Considerations and Limitations Important for the success of this approach is that the heterologous copy of the target gene be expressed at a level that is reasonable, compared to wild-type levels. If antibodies are available, this can be tested empirically. In any case, a wild-type copy of the target gene should be expressed using the system, in order to validate that functional rescue of the knockdown is achieved, before testing site mutants. The system also requires

47

2. Approaches for Functional Analysis ORF

3’UTR

target gene

NotI, EcorV

NotI, EcorV

DrugR

3’UTR

rDNA spacer

rDNA spacer

p2T7 3’UTR RNAi vector

NotI, EcorV

NotI, EcorV

rDNA spacer

**

ORF (mutated)

pKR10 expression

DrugR

3xHA

rDNA spacer

vector

+ Tet

loss of endogenous protein

ORF

**

ORF (mutated)

3xHA

expression of mutated tagged protein

Ribosomal RNA promoter (red), T7 polymerase promoter (blue)

rDNA spacer

Non-transcribed spacer of the rDNA locus

3’ processing element

“Head-to-head” Tet operator controlled T7 polymerse promoters

5’ processing element Tet operator controlled PARP promotor DrugR

Antibiotic resistance cassette

T7 polymerase terminator

Fig. 7 Combination of RNAi and heterologous expression for structure–function studies. In the example, knockdown of the target gene is achieved by RNAi against the gene’s 30 UTR. A tagged copy of the target gene, WT, or carrying directed point mutations is expressed simultaneously from a different vector.

that the 30 UTR is effective at ablating expression of the endogenous target gene, without significantly affecting the gene downstream of the target. This is tested using quantitative, real-time reverse transcriptase PCR. In our experience we have

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Michael Oberholzer et al.

found both criteria to be met for all genes so far tested. As with any epitope tagging analysis, care must be taken to ensure that the tag does not interfere with protein localization, assembly, or function. Heterologous expression of a nonmutated, tagged copy of the target to test for rescue is done to address this question. Examples of application: Ralston et al. in preparation

IV. Materials A. Culture Medium Cunningham’s culture media (“SM” medium) for cultivating PCF trypanosomes (Cunningham, 1977) Inorganic salts I. NaH2PO41H2O (Fisher #S369. FW = 137.99) II. A. MgCl26H2O (Fisher #M33. FW = 203.31) B. MgSO47 H2O (Fisher #M63. FW = 246.48) C. KCl (anhydrous) (Fisher #P217. FW = 74.56 III. CaCl22H2O (BDH #10070. FW = 147.02)

4 liter 2,120 mg 12,160 mg 14,800 mg 11,920 mg 600 mg

Sugars D-Glucose D-Fructose Sucrose

2,800 mg 1,600 mg 1,600 mg

(Fisher #D16. FW = 180.16) (Fisher #L95 FW = 180.16) (B.Mann. #100168 FW = 342.3)

Organic acids L-Malic a-Ketoglutaric Fumaric Succinic cis-Aconitate (3 mM final) Na-Pyruvate (1 mM final)

Vitamins BME vitamins (100 stock) Pen-Strep (100 stock) Phenol Red (0.5% stock)

4 liter 2,680 mg 1,480 mg 220 mg 240 mg 2,088 40 ml (100 mM Stock)

8 ml 40 ml 16 ml

(M-1000 FW = 134.1) (K-1128 FW = 146.1) (Fisher #A120 FW = 116.07) (MCB Rgts. #SX1040 FW = 118.09) (A-3412 FW = 174.1) (Gibco MEM #06304)

(B-6891 stored) (Gibco #15140-122) (Sigma, ßP-0290, optionala)

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2. Approaches for Functional Analysis

Amino acids 1. B-Alanine 2. DL-Alanine 3. L-Arginine 4. L-Asparagine 1 H2O 5. L-Aspartic acid 6. L-Cysteine-HCl H2O 7. L-Cystine 8. L-Glutamic acid 9. L-Glutamine 10. Glycine 11. L-Histidine 12. DL-Isoleucine 13. L-Leucine 14. L-Lysine 15. DL-Methionine 16. L-Phenylalanine 17. L-Proline 18. DL-Serine 19. L-Taurine 20. DL-Threonine 21. L-Tryptophan 22. L-Tyrosine 23. DL-Valine

Cat. Sigma # A-7752 A-7502 A-3909 A-8381 A-9256 Fisher BP-376 C-8767 G-1251 G-1517 Gibco 1106-038 H-8000 I-2627 L-8000 L-5626 M-2768 P-2126 P-0380 S-4375 T-0625 T-1520 Gibco 2110-010 T-3754 V-0375

F.W.: (g/mol)

4l

89.09 89.09 210.7 150.1 133.1 175.63 240.3 147.1 146.1 75.07 155.2 131.2 131.2 182.7 149.2 165.2 115.1 105.1 125.1 119.1 204.0 181.2 117.2

8,000 mg 4,360 mg 1,760 mg 960 mg 440 mg 320 mg 120 mg 1,000 mg 6,560 mg 480 mg 640 mg 360 mg 360 mg 600 mg 800 mg 800 mg 27,600 mg 800 mg 1,080 mg 400 mg 400 mg 800 mg 840 mg

1. Mix 23 amino acids in ~2.6 l H2O and stir at room temperature (RT) overnight to dissolve. 2. Mix independently, inorganic salts I, II, and III in 280, 160, and 160 ml H2O, respectively. 3. Mix sugars plus organic acids in 560 ml. 4. Combine all ingredients, including vitamins. 5. Adjust pH to 7.4 with 10 N NaOH (5–8 ml/l) and bring to 4 l final volume. 6. Filter sterilize and store in 1 l aliquots at 4°C. 7. Add HI-FCS to 10% before use. a Phenol red is optional and should be omitted if doing sedimentation assays.

SDM-79 medium for PCF trypanosomes (Brun and Schonenberger, 1979) HMI-9 media for cultivating BSF trypanosomes (Hesse et al., 1995)

Reagent 1 2 3 4 5

Iscove’s MDM, liquid Hypoxanthine Bathocuproine Disulfonate L-Cysteine H2O 2-mercaptoethanol

Cat number Gibco #12440-053 Sigma #H9377 Sigma #B1125 Fisher BP376-100 Sigma #M6250

FW 1 Liquid 136.1 g/mol 564.5 g/mol 175.6 g/mol 14.3 M

Final

For 500 ml

1 1 mM 50 uM 1.5 mM 0.2 mM

500 ml bottle 68 mg 14.1 mg 91 mg 7.2 ul

(Continued )

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Michael Oberholzer et al.

(Continued ) Reagent 6 7 8

Thymidine Na-Pyruvate Pen/Strep (þ 29.2 mg/ml L-Gln)

Cat number

FW

Sigma #T9250 242.2 g/mol Gibco #11360-070 100 mM Gibco #10378-016 10,000 µ/ml

Final 0.16 mM 1 mM 100 µ/ml

For 500 ml 19.5 mg 5 ml 5 ml

1. Dissolve all reagents in 500 ml of prewarmed (37°C) Iscove’s MDM by stirring at RT. This can be done right in the Iscove’s MDM bottle. Be sure all powders are dissolved. 2. If necessary, adjust pH to 7.4 with 1 N NaOH. 3. Filter sterilize (0.22 um filter unit) and store at 4°C. 4. Add heat-inactivated fetal calf serum to 15% prior to use.

Alternatively, HMI-9 can be made from powdered IMDM (Gibco #12200-036; 17.7 g/pk) as a starting point.

Reagent

Cat number

FW

Final

For 1 l “1 ”

1 Iscove’s MDM, powder 1 NaHCO3

17.7 g/pk 84.01 g/mol

1X = 1LTR 36 mM

1 pkg 3.024 g

2 3 4 5 6 7 8

136.1 g/mol 564.5 g/mol 175.6 g/mol 14.3 M 242.2 g/mol 100 mM 10,000 u/ml

1 mM 50 uM 1.5 mM 0.2 mM 0.16 mM 1 mM 100 u/ml

136 mg 28.2 mg 182 mg 14.4 ul 39 mg 10 ml 10 ml

Gibco #12200-036 EM sciences #SX0320-1 Hypoxanthine Sigma #H9377 Bathocuproine disulfonate Sigma #B1125 L-Cysteine H2O Fisher BP376-100 2-mercaptoethanol Sigma #M6250 Thymidine Sigma #T9250 Na-Pyruvate Gibco #11360-070 Pen/Strep (þ 29.2 mg/ml L-Gln) Gibco #10378-016

1. Dissolve pkg of IMDM in 800 ml warm MiliQ H2O 2. Add 3.024 g NaHCO3 while stirring at RT. 3. Add remaining ingredients while stirring at RT. Be sure all powders are dissolved. 4. If necessary, adjust pH to 7.4 with 1 N NaOH. 5. Bring to 1 l with H2O 6. Filter sterilize (0.22 um filter unit) and store at 4°C. 7. Add heat-inactivated fetal calf serum to 15% prior to use.

B. Transfection Reagents 1. BioRad® Gene Pulser II gene pulser. 2. BioRad® electroporation cuvettes 0.4 cm. 3. Electroporation medium (“EM”) (Hill et al., 1999)

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2. Approaches for Functional Analysis

Cytomix for trypanosome transfections: 120 mM KCl, 0.15 mM CaCl2, 10 mM K2HPO4 (anhydrous) (pH 7.6), 25 mM HEPES, 2 mM EDTA, and 5 mM MgCl2 Adjust to pH 7.6. Phosphate-buffered sucrose: 277 mM Sucrose, 16 mM KPO4 (pH 7.4), 1 mM MgCl2. 1. Combine cytomix and phosphate-buffered sucrose 3:1 for final EM.

C. Antibiotic Concentrations for Selection (See Table II) Table II Concentrations of Antibiotics Most Commonly Used for Selection of PCF and BSF Trypanosomes PCF

BSF

Antibiotic

Final concentration (µg/ml)

Final concentration (µg/ml)

G418

15a,b,c

Phleomycin

2.5a,b,c

Hygromycin

25–50a,b,c

Puromycin Blasticidin

1b,c 10b

2.5a 1–3b 15c 2.5a 1–2.5b 5c 5a 4–5b 50c 0.1b,c 5b

Note: Concentration may vary dependent on the expression level of the antibiotic resistance marker, for example, plasmid versus in situ expression, choice of promoter and UTR, etc… a Wirtz et al., 1999. b Homepage of Prof. G. Cross (http://tryps.rockefeller.edu/trypsru2). c Hill lab.

D. Buffers for Isolation of Extracted Flagellum Skeletons (Robinson et al., 1991) PMN: 10 mM NaPO4 (pH 7.2), 1 mM MgCl2, and 150 mM NaCl NP40: add to PMN to 1% final, as needed PEME: 0.1 M PIPES pH6.9, 2 mM EGTA, 1mM MgSO4, 0.1 mM EDTA E. Plasmids (See Table III)

F. Online Resources Trypanosome Genome website at Sanger “GeneDB,” http://www.genedb.org/ Trypanosome Genome Resource, “TriTryp DB,” http://beta.tritrypdb.org/tritrypdb1.1/

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Table III Most Commonly Used Vectors Name

Description and application

pLEW100 (Fig. 5) All purpose inducible expression using divergent promoters pLEW82 (Fig. 5) All purpose inducible expression using a single promoter pKR10 (Fig. 7) pLEW100 modified to include HA tag and MCS for N-terminal or C-terminal tagging pZJM (Fig. 6b) All purpose RNAi vector with opposing, Tet-inducible promoters p2T7 (Fig. 6a) All purpose RNAi vector with opposing, Tet-inducible promoters pMOTag series Series of vectors for in situ tagging (Fig. 4) with epitope, fluorescent protein (See Suppl. Table 1 for complete list of pMOTag variations and related vector systems) pC-PTP-Neo, pN- Plasmids for in situ tagging with PTP PTP-Puro tandem affinity purification tag

Antibiotic resistance markera

References

Phleo

Wirtz et al., 1999

Phleo

Wirtz et al., 1999

Puro

Ralston and Hill, unpublished

Phleo

Wang et al., 2000

Phleo

LaCount et al., 2002

G418, Puro, Hygro, Phleo, Blast

Oberholzer et al., 2006, http://www.izb.unibe. ch/res/seebeck/index. php

G418, Puro

Schimanski et al., 2005

Abbreviations: Phleo, phleomycin resistance; G418, G418 resistance; Puro, puromycin resistance; Hygro, hygromycin resistance; Blast, blasticidin resistance. Many additional plasmids and variations exist. Plasmids listed are those commonly used and for which the authors’ laboratory has direct experience. a For most plasmids, several different antibiotic resistance markers are available.

Trypanofan RNAi resource page and RNAit algorithm for selecting RNAi targets and primers, “Trypanofan,” http://trypanofan.path.cam.ac.uk/trypanofan/main/ Useful lab web pages: Prof. Prof. Prof. Prof. Prof. Prof.

G. Cross, http://tryps.rockefeller.edu/ C. Clayton, http://www.zmbh.uni-heidelberg.de/Clayton/default.shtml M. Carrington, http://web.me.com/mc115/mclab/resources.html D. Horn, http://homepages.lshtm.ac.uk/~ipmbdhor/dhhome.htm Thomas Seebeck, http://www.izb.unibe.ch/res/seebeck/index.php L. Simpson, http://dna.kdna.ucla.edu/simpsonlab

V. Summary and Outlook T. brucei offers a robust array of molecular genetic tools for functional analysis of flagellar proteins, as well as approaches for forward genetics. In particular,

2. Approaches for Functional Analysis

53

efficient homologous recombination and broad utility of heritable and inducible RNAi combine to provide tools complementary to those available in other systems. The ability to simultaneously ablate expression of an endogenous gene and replace it with a heterologous copy carrying directed point mutations will be useful for moving deeper into understanding how individual proteins work, particularly as the era of flagellum protein identification comes to a close and efforts shift to functional analysis. As useful as the systems are for functional analysis in T. brucei, limitations, and therefore opportunities, remain. For example, in situ tagging requires replacement of the target gene’s 30 UTR with a heterologous UTR, potentially resulting in altered expression. The recent development of a CRE recombinase-based system in T. brucei (Scahill et al., 2008) should overcome this limitation in next-generation tagging cassettes. A second area of need is further development of forward genetic screening approaches. The RNAi library has circumvented many limitations and should be applied more broadly, while construction of additional libraries should continue to improve genome coverage. The mariner transposon mutagenesis approach (Leal et al., 2004) also warrants more attention. Regarding flagellum fractionation, lateral attachment of the flagellum to the cell body poses significant challenges for isolating membrane-enclosed flagella. Advances are being made by employing mutants that disrupt lateral flagellum attachment and this approach is likely to yield results in the near future. The erratic and three-dimensional nature of T. brucei motility has so far limited quantitative descriptions of flagellar beating and cell movement. High-speed and three-dimensional video microscopy methods will need to be applied to overcome these limitations.

Acknowledgments We are grateful to all of our colleagues who have contributed to the development of systems and tools for analysis of gene function in T. brucei and apologize to those whose work was not cited. Work in the authors’ laboratory is supported by grants from the NIH-NIAID (AI052348), Burroughs Wellcome Fund, the Beckman Young Investigator program, and the Ellison Medical Foundation. ML is the recipient of an NIH-NRSA (GM07185). KR is the recipient of an NIH-NRSA (GM07104) and a Dissertation Year Fellowship from the UCLA graduate division. MO is the recipient of a Swiss National Science Foundation Fellowship.

References Absalon, S., Blisnick, T., Kohl, L., Toutirais, G., Dore, G., Julkowska, D., Tavenet, A., and Bastin, P. (2008). Intraflagellar transport and functional analysis of genes required for flagellum formation in trypanosomes. Mol. Biol. Cell 19(3), 929–944. Arhin, G.K., Shen, S., Ullu, E., and Tschudi, C. (2004). A PCR-based method for gene deletion and protein tagging in Trypanosoma brucei. Methods Mol. Biol. 270, 277–286. Badano, J.L., Mitsuma, N., Beales, P.L., and Katsanis, N.(2006). The ciliopathies: An emerging class of human genetic disorders. Annu. Rev. Genomics Hum. Genet. 7, 125–148. Baron, D.M., Kabututu, Z.P., and Hill, K.L. (2007). Stuck in reverse: Loss of LC1 in Trypanosoma brucei disrupts outer dynein arms and leads to reverse flagellar beat and backward movement. J. Cell Sci. 120(Pt 9), 1513–1520.

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Michael Oberholzer et al. Baron, D.M., Ralston, K.S., Kabututu, Z.P., and Hill, K.L. (2007). Functional genomics in Trypanosoma brucei identifies evolutionarily conserved components of motile flagella. J. Cell Sci. 120(Pt 3), 478–491. Barrett, B., LaCount, D.J., and Donelson, J.E. (2004). Trypanosoma brucei: A first-generation CRE-loxP site-specific recombination system. Exp. Parasitol. 106(1–2), 37–44. Bastin, P., Ellis, K., Kohl, L., and Gull, K. (2000). Flagellum ontogeny in trypanosomes studied via an inherited and regulated RNA interference system. J. Cell Sci. 113(Pt 18), 3321–3328. Bastin, P., Pullen, T.J., Sherwin, T., and Gull, K. (1999). Protein transport and flagellum assembly dynamics revealed by analysis of the paralysed trypanosome mutant snl-1. J. Cell Sci. 112(Pt 21), 3769–3777. Bastin, P., Sherwin, T., and Gull, K. (1998). Paraflagellar rod is vital for trypanosome motility. Nature 391(6667), 548. Bastin, P., Bagherzadeh, Z., Matthews, K. R. and Gull, K. (1996). Mol. Biochem. Parasitol. 77, 235–239. Berriman, M., et al. (2005). The genome of the African trypanosome Trypanosoma brucei. Science 309(5733), 416–422. Boothroyd, C.E., Dreesen, O., Leonova, T., Ly, K.I., Figueiredo, L.M., Cross, G.A., and Papavasiliou, F.N. (2009). A yeast-endonuclease-generated DNA break induces antigenic switching in Trypanosoma brucei. Nature 459(7244), 278–281. Bosher, J.M., and Labouesse, M. (2000). RNA interference: Genetic wand and genetic watchdog. Nat. Cell Biol. 2(2), E31–36. Branche, C., Kohl, L., Toutirais, G., Buisson, J., Cosson, J., and Bastin, P. (2006). Conserved and specific functions of axoneme components in trypanosome motility. J. Cell Sci. 119(Pt 16), 3443–3455. Broadhead, R., Dawe, H.R., Farr, H., Griffiths, S., Hart, S.R., Portman, N., Shaw, M.K., Ginger, M.L., Gaskell, S.J., McKean, P.G., and Gull, K. (2006). Flagellar motility is required for the viability of the bloodstream trypanosome. Nature 440(7081), 224–227. Brun, R., and Schonenberger, M. (1979). Cultivation and in vitro cloning or procyclic culture forms of Trypanosoma brucei in a semi-defined medium. Short communication. Acta Trop. 36(3), 289–292. Burkard, G., Fragoso, C.M., and Roditi, I. (2007). Highly efficient stable transformation of bloodstream forms of Trypanosoma brucei. Mol. Biochem. Parasitol. 153(2), 220–223. Carruthers, V.B., and Cross, G.A. (1992). High-efficiency clonal growth of bloodstream- and insect-form Trypanosoma brucei on agarose plates. Proc. Natl. Acad. Sci. U.S.A. 89(18), 8818–8821. Clayton, C.E. (1999). Genetic manipulation of kinetoplastida. Parasitol. Today 15(9), 372–378. Cunningham, I. (1977). New culture medium for maintenance of tsetse tissues and growth of trypanosomatids. J. Protozool. 24(2), 325–329. Drew, M.E., Morris, J.C., Wang, Z., Wells, L., Sanchez, M., Landfear, S.M., and Englund, P.T. (2003). The adenosine analog tubercidin inhibits glycolysis in Trypanosoma brucei as revealed by an RNA interference library. J. Biol. Chem. 278(47), 46596–46600. Dutcher, S.K., Huang, B., and Luck, D.J. (1984). Genetic dissection of the central pair microtubules of the flagella of Chlamydomonas reinhardtii. J. Cell Biol. 98(1), 229–236. Englund, P.T., Agbo, E.E., Lindsay, M.E., Liu, B., Liu, Y., Motyka, S.A., Yildirir, G., and Zhao, Z. (2005). RNAi libraries and kinetoplast DNA. Biochem. Soc. Trans. 33(Pt 6), 1409–1412. Ferguson, D.J. (2002). Toxoplasm? gondii and sex: Essential or optional extra? Trends Parasitol. 18(8), 355–359. Gadelha, C., Wickstead, B., and Gull, K. (2007). Flagellar and ciliary beating in trypanosome motility. Cell Motil. Cytoskeleton 64(8), 629–643. Glover, L., and Horn, D. (2009). Site-specific DNA double-strand breaks greatly increase stable transformation efficiency in Trypanosoma brucei. Mol. Biochem. Parasitol. 166(2), 194–197. Grandgenett, P.M., Otsu, K., Wilson, H.R., Wilson, M.E., and Donelson, J.E. (2007). A function for a specific zinc metalloprotease of African trypanosomes. PLoS Pathog. 3(10), 1432–1445. Gruby, M. (1843). Recherches et observations sur une nouvelle espèce d’hématozoaire, Trypanosoma sanguinis. C. R. Acad. Sci. Paris 17, 1134–1336. Haenni, S., Renggli, C.K., Fragoso, C.M., Oberle, M., and Roditi, I. (2006). The procyclin-associated genes of Trypanosoma brucei are not essential for cyclical transmission by tsetse. Mol. Biochem. Parasitol. 150 (2), 144–156.

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Hart, S.R., Lau, K.W., Hao, Z., Broadhead, R., Portman, N., Huhmer, A., Gull, K., McKean, P.G., Hubbard, S.J., and Gaskell, S.J. (2009). Analysis of the trypanosome flagellar proteome using a combined electron transfer/collisionally activated dissociation strategy. J. Am. Soc. Mass Spectrom. 20(2), 167–175. Hesse, F., Selzer, P.M., Muhlstadt, K., and Duszenko, M. (1995). A novel cultivation technique for long-term maintenance of bloodstream form trypanosomes in vitro. Mol. Biochem. Parasitol. 70(1–2), 157–166. Hill, K.L., Hutchings, N.R., Russell, D.G., and Donelson, J.E. (1999). A novel protein targeting domain directs proteins to the anterior cytoplasmic face of the flagellar pocket in African trypanosomes. J. Cell Sci. 112(Pt 18), 3091–3101. Hill, K.L., Hutchings, N.R., Grandgenett, P.M., and Donelson, J.E. (2000). T lymphocyte-triggering factor of African trypanosomes is associated with the flagellar fraction of the cytoskeleton and represents a new family of proteins that are present in several divergent eukaryotes. J. Biol. Chem. 275(50), 39369–39378. Hutchings, N.R., Donelson, J.E., and Hill, K.L. (2002). Trypanin is a cytoskeletal linker protein and is required for cell motility in African trypanosomes. J. Cell Biol. 156(5), 867–877. Ibanez-Tallon, I., Heintz, N., and Omran, H. (2003). To beat or not to beat: Roles of cilia in development and disease. Hum. Mol. Genet. 12(Spec No 1), R27–35. Jetton, N., Rothberg, K.G., Hubbard, J.G., Wise, J., Li, Y., Ball, H.L., and Ruben, L. (2009). The cell cycle as a therapeutic target against Trypanosoma brucei: Hesperadin inhibits Aurora kinase-1 and blocks mitotic progression in bloodstream forms. Mol. Microbiol. 72(2), 442–458. Kelly, S., Reed, J., Kramer, S., Ellis, L., Webb, H., Sunter, J., Salje, J., Marinsek, N., Gull, K., Wickstead, B., and Carrington, M. (2007). Functional genomics in Trypanosoma brucei: A collection of vectors for the expression of tagged proteins from endogenous and ectopic gene loci. Mol. Biochem. Parasitol. 154(1), 103–109. Klingbeil, M.M., Motyka, S.A., and Englund, P.T. (2002). Multiple mitochondrial DNA polymerases in Trypanosoma brucei. Mol. Cell 10(1), 175–186. Kohl, L., and Bastin, P. (2005). The flagellum of trypanosomes. Int. Rev. Cytol. 244, 227–285. Kohl, L., Robinson, D., and Bastin, P. (2003). Novel roles for the flagellum in cell morphogenesis and cytokinesis of trypanosomes. EMBO J. 22(20), 5336–5346. Kunz, S., Luginbuehl, E. and Seebeck, T. (2009). PLoS Negl Trop Dis 3, e455. LaCount, D.J., Barrett, B., and Donelson, J.E. (2002). Trypanosoma brucei FLA1 is required for flagellum attachment and cytokinesis. J. Biol. Chem. 277(20), 17580–17588. LaCount, D.J., Bruse, S., Hill, K.L., and Donelson, J.E. (2000). Double-stranded RNA interference in Trypanosoma brucei using head-to-head promoters. Mol. Biochem. Parasitol. 111(1), 67–76. LaCount, D.J., and Donelson, J.E. (2001). RNA interference in African trypanosomes. Protist 152(2), 103–111. Leal, S., Acosta-Serrano, A., Morris, J., and Cross, G.A. (2004). Transposon mutagenesis of Trypanosoma brucei identifies glycosylation mutants resistant to concanavalin A. J. Biol. Chem. 279(28), 28979–28988. Lee, M.G., and Van der Ploeg, L.H. (1990). Homologous recombination and stable transfection in the parasitic protozoan Trypanosoma brucei. Science 250(4987), 1583–1587. Li, J.B., et al. (2004). Comparative genomics identifies a flagellar and basal body proteome that includes the BBS5 human disease gene. Cell 117(4), 541–552. Li, F., Hua, S.B., Wang, C.C., and Gottesdiener, K.M. (1996). Procyclic Trypanosoma brucei cell lines deficient in ornithine decarboxylase activity. Mol. Biochem. Parasitol. 78(1–2), 227–236. Li, Z., Umeyama, T., and Wang, C.C. (2008). The chromosomal passenger complex and a mitotic kinesin interact with the Tousled-like kinase in trypanosomes to regulate mitosis and cytokinesis. PLoS ONE 3(11), e3814. Lowell, J. E. and Cross, G. A. (2004). J. Cell. Sci. 117, 5937–5947. Maga, J.A., and LeBowitz, J.H. (1999). Unravelling the kinetoplastid paraflagellar rod. Trends Cell Biol. 9 (10), 409–413. Monnerat, S., Clucas, C., Brown, E., Mottram, J.C., and Hammarton, T.C. (2009). Searching for novel cell cycle regulators in Trypanosoma brucei with an RNA interference screen. BMC Res. Notes 2, 46. Morris, J.C., Wang, Z., Drew, M.E., and Englund, P.T. (2002). Glycolysis modulates trypanosome glycoprotein expression as revealed by an RNAi library. EMBO J. 21(17), 4429–4438.

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Michael Oberholzer et al. Motyka, S.A., and Englund, P.T. (2004). RNA interference for analysis of gene function in trypanosomatids. Curr. Opin. Microbiol. 7(4), 362–368. Motyka, S.A., Zhao, Z., Gull, K., and Englund, P.T. (2004). Integration of pZJM library plasmids into unexpected locations in the Trypanosoma brucei genome. Mol. Biochem. Parasitol. 134(1), 163–167. Ngo, H., Tschudi, C., Gull, K., and Ullu, E. (1998). Double-stranded RNA induces mRNA degradation in Trypanosoma brucei. Proc. Natl. Acad. Sci. U.S.A. 95(25), 14687–14692. Nguyen, T.N., Schimanski, B., Zahn, A., Klumpp, B., and Gunzl, A. (2006). Purification of an eight subunit RNA polymerase I complex in Trypanosoma brucei. Mol. Biochem. Parasitol. 149(1), 27–37. Oberholzer, M., Bregy, P., Marti, G., Minca, M., Peier, M., and Seebeck, T. (2007a). Trypanosomes and mammalian sperm: One of a kind? Trends Parasitol. 23(2), 71–77. Oberholzer, M., Marti, G., Baresic, M., Kunz, S., Hemphill, A., and Seebeck, T. (2007b). The Trypanosoma brucei cAMP phosphodiesterases TbrPDEB1 and TbrPDEB2: Flagellar enzymes that are essential for parasite virulence. FASEB J. 21(3), 720–731. Oberholzer, M., Morand, S., Kunz, S., and Seebeck, T. (2006). A vector series for rapid PCR-mediated Cterminal in situ tagging of Trypanosoma brucei genes. Mol. Biochem. Parasitol. 145(1), 117–120. Ostrowski, L.E., Blackburn, K., Radde, K.M., Moyer, M.B., Schlatzer, D.M., Moseley, A., and Boucher, R. C. (2002). A proteomic analysis of human cilia: Identification of novel components. Mol. Cell Proteomics 1(6), 451–465. Pazour, G.J. (2004). Comparative genomics: Prediction of the ciliary and basal body proteome. Curr. Biol. 14(14), R575–577. Pazour, G.J., Agrin, N., Leszyk, J., and Witman, G.B. (2005). Proteomic analysis of a eukaryotic cilium. J. Cell Biol. 170(1), 103–113. Portman, N., Lacomble, S., Thomas, B., McKean, P.G., and Gull, K. (2009). Combining RNA interference mutants and comparative proteomics to identify protein components and dependences in a eukaryotic flagellum. J. Biol. Chem. 284(9), 5610–5619. Pullen, T.J., Ginger, M.L., Gaskell, S.J., and Gull, K. (2004). Protein targeting of an unusual, evolutionarily conserved adenylate kinase to a eukaryotic flagellum. Mol. Biol. Cell 15(7), 3257–3265. Ralston, K.S., and Hill, K.L. (2006). Trypanin, a component of the flagellar Dynein regulatory complex, is essential in bloodstream form African trypanosomes. PLoS Pathog. 2(9), e101. Ralston, K.S., and Hill, K.L. (2008). The flagellum of Trypanosoma brucei: New tricks from an old dog. Int. J. Parasitol. 38(8–9), 869–884. Ralston, K.S., Lerner, A.G., Diener, D.R., and Hill, K.L. (2006). Flagellar motility contributes to cytokinesis in Trypanosoma brucei and is modulated by an evolutionarily conserved dynein regulatory system. Eukaryotic Cell 5(4), 696–711. Ralston, K.S., Kabututu, Z.P., Melehani, J.H., Oberholzer, M., and Hill, K.L. (2009). The Trypanosoma brucei flagellum: Moving parasites in new directions. Annu. Rev. Microbiol. 63, 335–362. Redmond, S., Vadivelu, J., and Field, M.C. (2003). RNAit: An automated web-based tool for the selection of RNAi targets in Trypanosoma brucei. Mol. Biochem. Parasitol. 128(1), 115–118. Robinson, D., Beattie, P., Sherwin, T., and Gull, K. (1991). Microtubules, tubulin, and microtubuleassociated proteins of trypanosomes. Methods Enzymol. 196, 285–299. Satir, P., and Christensen, S.T. (2008). Structure and function of mammalian cilia. Histochem. Cell Biol. 129(6), 687–693. Scahill, M.D., Pastar, I., and Cross, G.A. (2008). CRE recombinase-based positive-negative selection systems for genetic manipulation in Trypanosoma brucei. Mol. Biochem. Parasitol. 157(1), 73–82. Schimanski, B., Nguyen, T.N., and Gunzl, A. (2005). Highly efficient tandem affinity purification of trypanosome protein complexes based on a novel epitope combination. Eukaryotic Cell 4(11), 1942–1950. Schlaeppi, K., Deflorin, J., and Seebeck, T. (1989). The major component of the paraflagellar rod of Trypanosoma brucei is a helical protein that is encoded by two identical, tandemly linked genes. J. Cell Biol. 109(4 Pt 1), 1695–1709. Schneider, A., Sherwin, T., Sasse, R., Russell, D.G., Gull, K., and Seebeck, T. (1987). Subpellicular and flagellar microtubules of Trypanosoma brucei brucei contain the same alpha-tubulin isoforms. J. Cell Biol. 104(3), 431–438.

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Selvapandiyan, A., Kumar, P., Morris, J.C., Salisbury, J.L., Wang, C.C., and Nakhasi, H.L. (2007). Centrin1 is required for organelle segregation and cytokinesis in Trypanosoma brucei. Mol. Biol. Cell 18(9), 3290–3301. Shen, S., Arhin, G.K., Ullu, E., and Tschudi, C. (2001). In vivo epitope tagging of Trypanosoma brucei genes using a one step PCR-based strategy. Mol. Biochem. Parasitol. 113(1), 171–173. Shi, H., Djikeng, A., Mark, T., Wirtz, E., Tschudi, C., and Ullu, E. (2000). Genetic interference in Trypanosoma brucei by heritable and inducible double-stranded RNA. RNA 6(7), 1069–1076. Siegel, T.N., Hekstra, D.R., Kemp, L.E., Figueiredo, L.M., Lowell, J.E., Fenyo, D., Wang, X., Dewell, S., and Cross, G.A. (2009). Genes Dev. 23, 1063–1076. Smith, J.C., Northey, J.G., Garg, J., Pearlman, R.E., and Siu, K.W. (2005). Robust method for proteome analysis by MS/MS using an entire translated genome: Demonstration on the ciliome of Tetrahymena thermophila. J. Proteome Res. 4(3), 909–919. Sugrue, P., Hirons, M.R., Adam, J.U., and Holwill, M.E. (1988). Flagellar wave reversal in the kinetoplastid flagellate Crithidia oncopelti. Biol. Cell 63(2), 127–131. Tetley, L., and Vickerman, K. (1985). Differentiation in Trypanosoma brucei: Host–parasite cell junctions and their persistence during acquisition of the variable antigen coat. J. Cell Sci. 74, 1–19. Ullu, E., Tschudi, C., and Chakraborty, T. (2004). RNA interference in protozoan parasites. Cell Microbiol. 6(6), 509–519. Ullu, E., Djikeng, A., Shi, H., and Tschudi, C. (2002). RNA interference: Advances and questions. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 357(1417), 65–70. Umeyama, T., and Wang, C.C. (2008). Polo-like kinase is expressed in S/G2/M phase and associated with the flagellum attachment zone in both procyclic and bloodstream forms of Trypanosoma brucei. Eukaryotic Cell 7(9), 1582–1590. Urwyler, S., Studer, E., Renggli, C.K., and Roditi, I. (2007). A family of stage-specific alanine-rich proteins on the surface of epimastigote forms of Trypanosoma brucei. Mol. Microbiol. 63(1), 218–228. Vassella, E., Oberle, M., Urwyler, S., Renggli, C.K., Studer, E., Hemphill, A., Fragoso, C., Butikofer, P., Brun, R., and Roditi, I. (2009). Major surface glycoproteins of insect forms of Trypanosoma brucei are not essential for cyclical transmission by tsetse. PLoS ONE 4(2), e4493. Vlachou, D., Schlegelmilch, T., Runn, E., Mendes, A., and Kafatos, F.C. (2006). The developmental migration of Plasmodium in mosquitoes. Curr. Opin. Genet. Dev. 16(4), 384–391. Walker, P.J. (1961). Organization of function in trypanosome flagella. Nature 189, 1017–1018. Wang, Z., Morris, J.C., Drew, M.E., and Englund, P.T. (2000). Inhibition of Trypanosoma brucei gene expression by RNA interference using an integratable vector with opposing T7 promoters. J. Biol. Chem. 275(51), 40174–40179. Wickstead, B., Ersfeld, K., and Gull, K. (2002). Targeting of a tetracycline-inducible expression system to the transcriptionally silent minichromosomes of Trypanosoma brucei. Mol. Biochem. Parasitol. 125(1–2), 211–216. Wirtz, E., and Clayton, C. (1995). Inducible gene expression in trypanosomes mediated by a prokaryotic repressor. Science 268(5214), 1179–1183. Wirtz, E., Leal, S., Ochatt, C., and Cross, G.A. (1999). A tightly regulated inducible expression system for conditional gene knock-outs and dominant-negative genetics in Trypanosoma brucei. Mol. Biochem. Parasitol. 99(1), 89–101. Zamudio, J.R., Mittra, B., Chattopadhyay, A., Wohlschlegel, J.A., Sturm, N.R., and Campbell, D.A. (2009). Trypanosoma brucei spliced leader RNA maturation by the cap 1 2’-O-ribose methyltransferase and SLA1 H/ACA snoRNA pseudouridine synthase complex. Mol. Cell Biol. 29(5), 1202–1211. Zhao, Z., Lindsay, M.E., Roy Chowdhury, A., Robinson, D.R., and Englund, P.T. (2008). p166, a link between the trypanosome mitochondrial DNA and flagellum, mediates genome segregation. EMBO J. 27 (1), 143–154. Zomerdijk, J.C., Ouellette, M., ten Asbroek, A.L., Kieft, R., Bommer, A.M., Clayton, C.E., and Borst, P. (1990). The promoter for a variant surface glycoprotein gene expression site in Trypanosoma brucei. EMBO J. 9(9), 2791–2801.

CHAPTER 3

Tools for Analyzing Intraflagellar Transport in Trypanosomes Daria Julkowska and Philippe Bastin Trypanosome Cell Biology Unit, Institut Pasteur and CNRS, Paris, France

Abstract I. Introduction II. Methodology B. Cell Culture B. In Silico Analysis of the Genome C. Genetic Tools for IFT Gene Silencing, Expression, and Tagging of IFT Proteins D. Visualization of IFT and IFT-Related Phenotypes III. Perspectives Acknowledgments References

Abstract African trypanosomes are evolutionary-divergent eukaryotes responsible for sleeping sickness. They duplicate their single flagellum while maintaining the old one, providing a unique model to examine mature and elongating flagella in the same cell. Like in most eukaryotes, the trypanosome flagellum is constructed by addition of novel subunits at its distal end via the action of intraflagellar transport (IFT). Almost all genes encoding IFT proteins and motors are conserved in trypanosomes and related species, with only a few exceptions. A dozen of IFT genes have been functionally investigated in this organism, thanks to the potent reverse genetic tools available. Several alternative techniques to trigger RNAi are accessible, either transient RNAi by transfection of long doublestranded RNA or by generation of clonal cell lines able to express long double-stranded RNA under the control of tetracycline-inducible promoters. In addition, we provide a series of techniques to investigate cellular phenotypes in trypanosomes where expression of IFT genes has been silenced. In this chapter, we describe different methods for tagging and expression of IFT proteins in trypanosomes and for visualizing IFT in live cells. METHODS IN CELL BIOLOGY, VOL. 93 Copyright Ó 2009 Elsevier Inc. All rights reserved.

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978-0-12-381377-0 DOI: 10.1016/S0091-679X(08)93003-X

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I. Introduction Trypanosoma brucei is a unicellular flagellated parasite causing sleeping sickness, a fatal tropical disease. Trypanosomes are found in the bloodstream of various mammalian hosts where they proliferate as extracellular parasites. They are transmitted by the bite of the tsetse fly in which they undergo a complex series of differentiation, migration, and proliferation steps. Development is accompanied by changes in morphology, biochemistry, cell cycle status, and expression of major surface markers. Trypanosomes and related parasites have been extensively studied as they are the causative agents of severe tropical diseases for which no vaccine is currently available. Moreover, due to their early evolutionary divergence, these parasites also developed original molecular and cellular features that made them interesting model organisms. For example, RNA editing and the GPI anchor were first discovered in African trypanosomes (Ferguson, 1988). Trypanosomes belong to the order of kinetoplastids that are characterized by the presence of the kinetoplast—a DNA-containing “granule” located within the single elongated mitochondrion and associated with the flagellar basal body (Kohl and Bastin, 2005; Ralston and Hill, 2008). They possess a single flagellum that presents several original features (Fig. 1). First, it is attached to the cell body for most of its length with the exception of its distal tip. Second, flagellum movement is initiated from

Fig. 1 (A) EM cross-section through the flagellum of a procyclic T. brucei cell showing the classic axoneme made of nine doublets of peripheral microtubules carrying dynein arms and radial spokes and of the lattice-like paraflagellar rod (PFR) structure. An IFT particle is indicated by the arrow. Reproduced from Absalon et al. (2008b). (B) Longitudinal section through the base of the flagellum of a procyclic T. brucei cell revealing the basal body (BB), the transition region (TR), the anterior (A), and posterior (P) sides of the flagellar pocket and the tight junctions that mark the collar region (arrows). Reproduced from Absalon et al. (2008a).

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the distal end, dragging the cell forward but also twisting the cell body. The flagellum wave can also be reversed, resulting in changes of orientation and backward motility. Third, the flagellum contains an extra-axonemal structure called the paraflagellar rod (PFR), a lattice-like structure that is physically linked to the axoneme (Fig. 1A) and that is required for flagellum motility. Fourth, the flagellum emerges from the flagellar pocket, an invagination of the cell membrane that is also the sole site for endocytosis or exocytosis (Fig. 1B). Fifth, the cell assembles its new flagellum while maintaining the old one. This means that at the later stages of the cell cycle, trypanosomes possess two flagella: a long one that is fully assembled and a shorter one that is under construction. Therefore, it is possible to compare mature and elongating flagella in the same cell. Last but not least, the flagellum is essential for trypanosome survival both in vitro and in vivo (Broadhead et al., 2006; Griffiths et al., 2007; Ralston and Hill, 2006; Ralston et al., 2006) and hence represents a potential drug target for the fight against sleeping sickness. Genome analysis has revealed that almost all the genes involved in intraflagellar transport (IFT) are conserved in trypanosomes (Table I) (Berriman et al., 2005; Briggs et al., 2004; Kohl and Bastin, 2005) with a few exceptions. Two genes encoding motor subunits of kinesin II are present; however, the kinesin-associated protein (KAP) identified in numerous flagellated eukaryotic organisms is absent from all Trypanosoma and Leishmania subspecies, suggesting that the kinesin motor is dimeric (heterodimer or two separate homodimers). Two separate genes are found for the dynein heavy chain (DHC1b) motor that share only 60% similarity at the amino acid levels. This feature is also conserved in all Trypanosomatid species and not encountered in other eukaryotes studied so far. Among IFT proteins, candidate homologues have been identified by BLASTP with high E-values with the exception of IFT43 and DYF11 that were not confirmed by reciprocal analysis. Experimental validation will be required to confirm the involvement of these two candidate proteins in IFT. All IFT proteins studied to date have been localized to the basal body and the flagellum compartment, with possibly some protein in the cell body (Absalon et al., 2008b; Adhiambo et al., 2009). Live video microscopy demonstrated that IFT is active in both the old and the new flagellum (Absalon et al., 2008b), showing its involvement in maintenance and construction, respectively. Individual RNAi targeting of a dozen of IFT genes revealed that only construction of the new flagellum is affected, presumably because RNAi targets mRNA and not existing proteins that were synthesized before the onset of RNAi (Kohl et al., 2003; Absalon et al., 2007, 2008b; Adhiambo et al., 2009; Davidge et al., 2006). These data suggest that the old and the new flagellum are separate entities. In all cases, the axoneme was either very short or absent, and no PFR structure could be recognized. It now remains to be understood whether the PFR is assembled via IFT or whether a distinct transport system is necessary. These experiments confirmed the specific involvement of several IFT proteins in anterograde or retrograde transport. Inhibition of anterograde transport blocks flagellum formation (Absalon et al., 2008b), and only a new basal body, shorter than usual, is formed (Absalon et al., 2008a), whereas interfering with retrograde transport results in the formation of very short flagella (less than 2 µm) filled with IFT material

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Table I IFT Genes in Trypanosoma brucei Gene name Kinesin motor KIF3A KIF3B KAP Dynein motor DHC2.2 DHC2.1 LC8 D1bLIC IFT complex A IFT43 IFT122A IFT122B IFT139 IFT140 PIFTF6 IFT Complex B IFT20 IFT25 IFT27 IFT46 IFT52 IFT57 IFT74 IFT80 IFT81 IFT88 IFT172 PIFTA1 PIFTB2 PIFTC3 DYF11 RABL5

Other name

GeneDB

Reference

KLP11/FLA8 KLP20/FLA10 KAP1/FLA3

Tb927.5.2090 Tb11.01.5490 Missing

n.d. Kohl and Bastin (2005)

CHE3/DHC2 CHE3/DHC2 FLA14 FAP133 XBX1/D2LIC

Tb11.02.0030 Tb927.4.560 Tb11.03.0815 Tb927.3.5540 Tb11.01.8570

Kohl et al. (2003) n.d. n.d. Blisnick et al., unpublished Blisnick et al., unpublished

DAF10 IFTA1/OSEG4 THM1 CHE11 IFT144/DYF2

Tb927.2.4530 Tb10.70.1660 Tb927.5.3030 Tb09.160.5670 Tb10.61.2260 Tb11.03.0880

n.d. Absalon et al. (2008b) Absalon et al. (2008b) Absalon et al. (2008b) Absalon et al. (2008b)

Tb927.6.3290 Tb11.01.4880 Tb927.3.5550 Tb927.6.3100 Tb10.61.1590 Tb10.26.0670 Tb927.7.3370 Tb10.61.1560 Tb10.70.5020 Tb11.55.0006 Tb10.70.6920 Tb927.3.1110 Tb927.3.5490 Tb927.3.3000 Tb10.70.7880 Tb11.01.8590

Absalon et al. (2007) n.d. Huet et al., unpublished n.d. Absalon et al. (2008b) Absalon et al. (2008b) n.d. Absalon et al. (2008b), Davidge et al. (2006) n.d. Kohl et al. (2003) Absalon et al. (2008b) Absalon et al. (2008b) Absalon et al. (2008b) Absalon et al. (2008b) n.d. Adhiambo et al. (2009)

RABL4 DYF6 OSM6 CHE13/Hippi CHE2 OSM5/Tg737 OSM1 DYF3 DYF1/fleer DYF13 DYF11/MIP-T3 IFTA2/IFT22

n.d., not done.

(Absalon et al., 2008b). Remarkably, the membrane keeps on elongating despite the absence of an axoneme or microtubules, showing that membrane formation is not completely dependent on axoneme construction (Absalon et al., 2008a; Davidge et al., 2006). It should be noted that the small G protein RABL5/IFT22, despite being found associated to the B complex for anterograde transport in Chlamydomonas, is actually necessary for retrograde IFT (Adhiambo et al., 2009). However, RABL5 is not conserved in every flagellated species (e.g., a gene encoding RABL5 is missing from most insect genomes) and is not involved in cilium formation in Caenorhabditis elegans, although it is transported along the axoneme (Schafer, et al., 2006). This could

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reflect peculiar interspecies differences and justifies the use of multiple models to investigate function of IFT-related proteins. Several molecular and cellular tools are available to investigate the trypanosome flagellum, including easy culture systems, a full genome sequence (including for several related species), and potent reverse genetics. In this chapter, we will describe several techniques for functional studies in trypanosomes, with special emphasis on IFT and analysis of flagellum-related phenotypes.

II. Methodology B. Cell Culture Trypanosomes exhibit at least nine different life-cycle stages in vivo, four of which are actually proliferating, the others being blocked in the G0 phase of their cell cycle. In the salivary glands, T. brucei does proliferate while being attached to the microvilli of the epithelial cells. Curiously, this adhesion takes place via the flagellum whose membrane produces intricate extensions with electron-dense structures resembling hemi-desmosomes (Tetley and Vickerman, 1985). This life-cycle stage has not been successfully maintained in culture so far. The second proliferative stage is called the long slender form and is encountered in the bloodstream of mammalian hosts. It can be injected into rodents via different routes (intraperitoneal or intravenous) and can be grown in culture. Parasites are grown in suspension in HMI9 medium (Hirumi and Hirumi, 1989) and do not need a feeder layer of cells. HMI9 is prepared by dissolution of 1 prepacked Iscove’s Mod. Dulbecco’s medium (GIBCO/BRL Cat. No. 42200-022) in 1 l of water, followed by the addition of 15.1 g of NaHCO3. The solution is filtersterilized and stored for several weeks/months at 4°C. Before use, it needs to be supplemented with 10 ml of each of the followings: 100 bathocuproine stock (dissolve 546 mg in 200 ml H2O, sterile filtrate, and store at 4°C), 100 thymidine stock (dissolve 780 mg in 200 ml H2O, filter-sterilized, and store at 4°C), 100 ß-mercaptoethanol stock (add 280 µl ß-mercaptoethanol in 200 ml H2O, filtersterilized, and store at 4°C), hypoxanthine stock (dissolve 2.72 g in 200 ml 0.1 M NaOH, filter-sterilized, and store in 10-ml aliquots at 20°C), and freshly prepared L-cysteine (18.2 mg/ml) before addition of 10% fetal calf serum (heat-inactivated 1 h at 56°C). Bloodstream slender cells are grown at 37°C with 5% CO2 and cell density must not exceed 8 105 cells per ml but strong dilutions are very well tolerated. The third proliferative stage is termed the procyclic form (PCF) and is encountered in the peritrophic space of the midgut of the tsetse fly. This life-cycle stage can also be grown in suspension in semidefined medium 79 (SDM79) (Brun and Schonenberger, 1979) or in Cunningham medium (Cunningham, 1977). In Europe, this semidefined medium can be purchased from PAA Laboratories, Les Mureaux, France with specific adjustments. The powder is dissolved in 4.5 l of water before addition of 10 g of NaHCO3. The pH is adjusted to 7.3 with 2 N NaOH, the medium is filter-sterilized and stored as 450-ml aliquots at 4°C. Before use, the medium is supplemented with 7.5 mg hemin/l (stock

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solution 2.5 mg/ml in 0.05 N NaOH) and 10% fetal calf serum. In both cases, batches of serum should be tested for compatibility with trypanosomes and for absence of tetracycline if one is using a tetracycline-inducible expression system. Procyclic trypanosomes are grown at 27°C and maintained between 3 and 10 millions cells per ml. They do not tolerate strong dilutions without addition of conditioned medium or larger amounts of serum. B. In Silico Analysis of the Genome The full 26-megabase nuclear genome sequence of T. brucei was published in 2005 (Berriman et al., 2005). It contains 11 diploid megabase chromosomes of similar architecture, with the housekeeping genes located internally and with the gene families devoted to host invasion and interaction at the subtelomeric region (Berriman et al., 2002). It encodes 9068 predicted genes of which 908 were classified as pseudogenes and ~1700 as T. brucei-specific genes. The genome of kinetoplastids represents a peculiar physical architecture of transcription units. Adjacent genes tend to map on the same coding strand and can be depicted as a succession of long, nonoverlapping gene clusters, where all predicted genes share the same transcriptional orientation. These directional gene clusters are devoid of introns and are transcribed as a polycistron that is subsequently trans-spliced and polyadenylated (Campbell et al., 2003). Genes in clusters are not functionally related and are organized randomly in contrast to bacteria where adjacent genes often form functionally related operating units. Most of the protein-coding genes are transcribed by RNA polymerase II (RNAP II) that is recruited and, as far as it is known, initiates transcription without the presence of a specific promoter (Clayton, 2002). The exceptions are two loci—a bloodstream form (BSF) variant surface glycoprotein (VSG) at subtelomeric regions and the intrachromosomal EP/GPEET procyclin that are both transcribed by RNA polymerase I (Sherman et al., 1991). These gene families encode the most abundant and major surface protein for each specific stage of the life cycle. T. brucei genome data together with that from other kinetoplastids Trypanosoma cruzi, Leishmania major, Leishmania braziliensis, and Leishmania infantum and ongoing genome sequencing projects for Trypanosoma vivax, T. brucei gambiense, and T. brucei congolense can be accessed via several on-line databases. A common initiative of The Welcome Trust Sanger Institute and the Institute for Genome Research led to the creation of GeneDB (www.genedb.org) and trypanosomatid functional analysis network (TrypanoFAN, www.trypanofan.org) projects. GeneDB aims to develop and maintain database resources for a diverse range of organisms, including access to DNA and protein sequences with direct connection to associated BLAST servers, gene ontology annotation, predicted protein properties, similarity information, summary of up-to-date protein domain(s) and motif searches and useful literature links (Hertz-Fowler et al., 2004). TrypanoFAN is a program that is using the information from the T. brucei genome project to produce a research resource and systematic collection of plasmids and strains used for gene inactivation by RNAi (Subramaniam et al., 2006). It contains simple and standardized functional characterization of

3. Tools for Analyzing Intraflagellar Transport in Trypanosomes

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different mutants and, among many other functions, allows identification of primers for production of RNAi constructs taking into account eventual cross-talk between related gene products. Lately, the TriTrypDB data base (http://TriTrypDB.org) has become available. This database integrates genomic and functional genomic data for pathogens of the family Trypanosomatidae, including organisms in both the Leishmania and the Trypanosoma genera. Importantly, users can directly add comments and annotations. TriTrypDB and its continued development are possible through the collaborative efforts between EuPathDB, GeneDB, and scientists at the Seattle Biomedical Research Institute (SBRI). Genome scan and the selection of protein candidates for IFT can be performed according to several criteria: (1) conservation of the genes whose products are involved in flagellum formation by comparison with the genomes of different ciliated (e.g., Chlamydomonas, Drosophila, Caenorhabditis, Plasmodium, and Homo sapiens) and nonciliated (Arabidopsis, Saccharomyces, and Dictyostelium) species; (2) comparison with the genomes of organisms assembling their flagella using the IFT machinery (Chlamydomonas, Homo sapiens, Drosophila, and Caenorhabditis) and the genome of Plasmodium subspecies, where flagella are found only in the male gametes and are assembled in the cytoplasm (Avidor-Reiss et al., 2004). In the sequenced genome of all four Plasmodium subspecies, IFT genes are absent in contrast to genes encoding axonemal protein components such as dyneins and central pair proteins. Finally, the third criterion is the comparison of the protein sequences and presence of domains often found in IFT proteins, such as coiled-coil, WD40 domains, or TPR motifs (Cole, 2003). Information present in the databases described above allowed us to identify T. brucei genes coding for homologues of proteins involved in IFT (Kohl et al., 2003) and novel genes putatively involved in IFT (Absalon et al., 2008b) (see also Introduction, Table I). C. Genetic Tools for IFT Gene Silencing, Expression, and Tagging of IFT Proteins RNA interference (RNAi) is a cellular process in which the presence of a doublestranded RNA (dsRNA) of a given gene leads to destruction of mRNA of homologous sequence, specifically inhibiting the expression of the corresponding gene. This posttranscriptional gene silencing process involves several cellular proteins and is well conserved in eukaryotes including T. brucei (Cerutti and Casas-Mollano, 2006). Curiously, in the family of kinetoplastids, the RNAi machinery is missing from Trypanosoma cruzi and most Leishmania species with the exception of L. braziliensis (Peacock, et al., 2007). The process is initiated by the action of Dicer, a multidomain RNaseIII (Shi et al., 2006) that generates short interfering RNAs (siRNA) by the cleavage of dsRNA into small ~25 nucleotide fragments. The siRNAs are separated into single strands and integrated into an active RNA-induced silencing complex (RISC) that contains the Argonaute endonuclease (Durand-Dubief and Bastin, 2003; Shi et al., 2004). Argonaute cleaves the target mRNA complementary to siRNA that is bound in the RISC complex. The mRNA is degraded by exo- and endonucleases and the translation process can no longer take place.

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In trypanosomes, RNAi silencing (gene knockdown) can be performed either by transfection of cells directly with in vitro-synthesized dsRNA (Ngo et al., 1998) or by use of specific vectors allowing expression of dsRNA in the cell (Bastin et al., 2000; Bringaud et al., 2000; LaCount et al., 2000; Shi et al., 2000; Wang et al., 2000). The RNAi response is activated almost immediately after electroporation of parasites with exogenous dsRNA, and its effects can be observed during the following hours, usually for at least one cell generation. However, silencing does not persist and the electroporation itself kills about half of the cell population, thereby complicating the interpretation. A nice alternative is transfection with plasmids expressing dsRNA under the control of a tetracycline-inducible promoter. The plasmid integrates by homologous recombination into chromosomal DNA, exactly within the region of homology where it was linearized. This results in a stable cell line, where the plasmid sequence will be maintained and duplicated during each cell division. The presence of a specific tetracycline-inducible promoter allows tight control of dsRNA production and kinetic analysis of the knockdown phenotype, even in the case of essential genes.

1. Expression Vectors for dsRNA in Trypanosomes The expression of dsRNA must be strictly controlled and therefore needs the use of an inducible promoter. As mentioned before, most of the trypanosome genes are transcribed by continuous run-through of the RNAP II as polycistronic products without any particular promoter sequence. Two exceptions are the genes encoding the cell surface proteins procyclin (PCF) or VSG (BSF) whose expression is dependent on RNA polymerase I activity. Both promoters have been mapped and their sequence is available (Sherman et al., 1991). To generate an inducible expression system, the EP procyclin promoter was modified by insertion of two TetO operator sequences creating potential TetR repressor binding sites. The Tn10-encoded tetR gene, modified for expression in eukaryotes, was introduced into the tubulin locus of T. brucei generating stable cell line expressing the tetracycline repressor. This new version of an EP procyclin promoter was used for the creation of plasmids allowing controllable production of dsRNA when transfected into TetRþ cell line (Wirtz and Clayton, 1995). However, due to the use of the EP promoter, the expression of these constructs was still dependent on the life cycle and limited to the PCF. To expand reverse genetic approaches in T. brucei, the well-known T7 phage polymerase and promoter system together with the tetracycline operator of Escherichia coli were used (Wirtz and Clayton, 1995; Wirtz et al., 1998). In trypanosomes, possibly due to a more open chromatin conformation and accessibility of DNA template, T7 transcription driven from the T7 promoter is highly processive (Wirtz et al., 1998). In order to obtain the level of expression similar to that mediated by endogenous RNAP II, the wild type T7 promoter sequence was modified by the introduction of point mutations and TetO sequence for inducible expression (Wirtz et al., 1998). To exploit the new T7 polymerase-dependent system, two cell lines, 2913 (procyclic) and 9013 (bloodstream), were generated (Wirtz et al., 1999). Both of them express constitutively the T7 polymerase and the TetR repressor. These cell lines can be transfected with

67

3. Tools for Analyzing Intraflagellar Transport in Trypanosomes Notl

(A)

(B)

BamHI Eam1105l

Notl pT7

Ter

rDNA spacer

rDNA spacer

6000 1000 AMP

6000

6371 bps

Ter

2000

pT7 Kpnl

p2T7TAblue Ter

5000

6740 bps 2000

4000 3000

1000

pT7

pZJM

5000

Gene fragment

Eam1105l Xhol

BLE

gene fragment

pT7 Xhol

AMP 4000

3000

Hygro ALD polyA

Ter pT7 HindIII

Mlul

BamHI

Fig. 2 Maps of the “RNAi vectors” pZJM (A) and p2T7TAblue (B) used for expression of dsRNA in T. brucei. AMP, ampicillin; BLE, phleomycin; HYGRO, hygromycin, antibiotic resistance genes; T7 transcription terminators—Ter (black bars); T7 promoters—pT7 (black arrows); TetO-tetracycline operators; gene fragment; rDNA spacer -rDNA intergenic region.

plasmids bearing the gene of interest under the control of T7 promoters that are controlled by tetracycline repressor and tetracycline inducible (see below). Two families of vectors (Fig. 2) are now being commonly used for RNAi knockdown in T. brucei: pZJM (Wang et al., 2000) and p2T7TAblue (Alsford et al., 2005; LaCount et al., 2000). In both of them a PCR-amplified gene fragment is ligated in a single step between two opposing tetracycline-inducible T7 promoters necessary to generate sense and antisense RNA. The TetO operators were placed immediately downstream of each T7 promoter to allow regulation of their activity by the TetR. Dual T7 transcription terminators were positioned outside the T7 promoters to prevent transcription of the neighboring genes by T7 RNA polymerase. As most of the trypanosome genome is transcribed by RNAP II, it is of prime importance to integrate the plasmid into a silent region to avoid run-through transcription of the reporting constructs. Two such regions are commonly used: the intergenic rDNA region (inverted orientation to rRNA transcription) and the 177 bp-repeats situated on minichromosomes (Wickstead et al., 2002). Integration of the plasmids into nontranscribed region of the T. brucei genome results in the establishment of a stable cell line. In the pZJM plasmid the gene fragment is cloned between unique XhoI and HindIII restriction sites. P2T7TAblue possesses an additional restriction site for Eam1105I that generates 30 -overhanging T-ends, which allow PCR products with a 30 A-extension to be cloned directly. Plasmids are linearized at the NotI site and integrated into rDNA loci in procyclic 2913 or bloodstream 9013 parasites. p2T7TAblue vector is now a tool of choice for establishing collections of RNAi mutants in BSF parasites in the TrypanoFAN program (trypanofan.org).

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Another alternative for dsRNA expression are stem-loop vectors where the fragment of the targeted gene is cloned in two inverted copies linked by a stuffer. This results in the production of dsRNA in the form of a hairpin structure (Bastin et al., 2000; Bringaud et al., 2000; Wang et al., 2000). The efficiency of this approach is highly effective for gene silencing in trypanosomes, but the risk of cross RNAi is higher and the cloning procedure is more time consuming than in the case of dual promoter systems (Durand-Dubief et al., 2003; Wang et al., 2000). The long dsRNA at least 200 are transcribed upon addition of tetracycline that releases the repressor from the T7 or the EP procyclin promoters, and intermolecular or intramolecular dsRNA is formed upon annealing of complementary RNA. These molecules are subsequently processed into multiple siRNA that trigger RNAi. Using the pZJM vector, we were able to silence the expression of several IFT (IFT122, IFT140, IFT20, IFT52, IFT55, IFT80, IFT88, IFT172, DHC1b) and IFTrelated (PIFTA1, PIFTB2, PIFTC3, PIFTF6, PIFTD4, RABL5) genes and to demonstrate their importance in flagellum construction (Absalon et al., 2007, 2008b; Adhiambo et al., 2009; Davidge et al., 2006; Kohl et al., 2003). The knockdown efficiency was monitored at the RNA level by semi-quantitative RT-PCR. Total RNA was extracted from tetracycline-induced cells and purified with Trizol. At least one of the primers was designed to be outside the region of dsRNA to avoid amplification of RNA deriving from dsRNA trigger. A series of RT-PCR with different concentrations of RNA template was performed to obtain a linearity curve. This is essential to establish a range in which the quantity of RNA used for the RT-PCR reaction will allow detection of changes in the amount of RNA (Fig. 3). In all cases RNAi silencing leads to the inhibition of formation of the new (A)

(C) CDS

3’UTR

AAAA

30 100 ng 0 ng

Signal intensity

(B) 20

10

0

0

20

40 60 RNA (ng)

80

100

Fig. 3 RT-PCR amplification of IFT172 carried out using different amounts of total RNA from wild-type trypanosomes as a template. (A) Amplification strategy. (B) Agarose gel showing amplification products. Respective amounts of RNA were 100, 50, 25, 12.5, 6.3, 3.1, and 0 (from left to right). (C) Diagram showing the relationship between signal intensity and amount of RNA used. The linear range is found between 3 and 15 ng of template RNA.

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3. Tools for Analyzing Intraflagellar Transport in Trypanosomes

(A)

(B) IFT122 IFT140

Flagellated cells (%)

100 80

80

60

60

40

40

20

20

0

0

24

48

72 96 Time (h)

120

IFT52 IFT55 IFT80 IFT88 IFT172

100

0

0

24

48

72 96 Time (h)

120

Fig. 4 RNA silencing of IFT genes. (A) Inactivation of genes encoding proteins of complex A (retrograde IFT). (B) Inactivation of genes encoding proteins belonging to complex B (anterograde IFT). Graphs represent the proportion of flagellated cells left in the culture after the indicated days of RNAi silencing. Reproduced from Absalon et al. (2008b).

flagellum and finally to the emergence of cells with a very short flagellum (complex A) or non-flagellated cells (complex B). Trypanosomes are difficult to synchronize and RNAi induction may produce different effects depending on the cycle stage. It is essential to examine cells throughout the induction. We noted significant differences in the rate of emergence of cells with a defective flagellum within complex A and complex B mutants (Fig. 4). However, these were not due to variable efficiency of RNA degradation, as kinetics of RNA silencing are very reproducible and reach their maximum (~90%) after 48 h (Fig. 5), but rather these likely reflect differences at the protein level (Absalon et al., 2008b).

2. Vectors for IFT Protein Tagging and Protein Fusions Experiments for protein–protein interactions, purification of protein complexes, and in vivo visualization of IFT particles become essential to understand the mechanisms responsible for flagellum growth and maintenance. Assembly of IFT complexes, cargo loading and release events, speed of IFT particles or their exchange at the base, and tip of the flagellum are still to be fully elucidated. Many of these questions could be solved by the use of proteins fused either to fluorescent marker like GFP or tagged with a specific peptide. Here we present two vectors that allow constitutive/inducible expression of epitope tagged/GFP fused proteins or endogenous protein tagging for physiological level protein expression. The pHD430 plasmid was originally created in Christine Clayton’s laboratory (Wirtz and Clayton, 1995) for inducible gene expression mediated by a prokaryotic TetR repressor in T. brucei. As mentioned before, the procyclin promoter was modified by the insertion of two TetO operator sequences for a tight, tetracycline inducible,

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Control

Target

IFT172/control relative abundance

IFT172RNAi

80 60 40 20 0

0

24

48

72

96

IFT172RNAi

100

120

0

Target Control

0

24

48

72

96

120

PIFTD4/control relative abundance

PIFTD4RNAi

48 72 24 96 Induction time (h)

120

PIFTD4RNAi

100 80 60 40 20 0

0

96 24 48 72 Induction time (h)

120

Fig. 5 RT-PCR on total RNA extracted from IFT172RNAi and PIFTD4RNAi cell lines at successive days during the course of RNAi silencing. PIFTD4RNAi shows similar kinetics for RNAi silencing to IFT172RNAi mutant but presents a very slow and low-penetrance phenotype. Reproduced from Absalon et al. (2008b).

transcriptional control in cell lines expressing the TetR repressor (Wirtz et al., 1999). The EP promoter also allows expression of genes of interest in wild-type cells that are processed by RNA polymerase I. In these conditions, its activity is comparable to that of the operatorless parent promoter (Wirtz and Clayton, 1995). Figure 6A represents a modified version of the pHD430 vector used in our laboratory. In this example the gene IFT52 is fused to one encoding the GFP fluorescent marker and placed under control of the inducible EP promoter. Plasmid is linearized by NotI and inserted in the rDNA spacer region. It is noteworthy that the transcription of the phleomycin resistance gene is also dependent on the procyclin promoter. Therefore, in cell lines bearing the TetR repressor, tetracycline must always be added when using phleomycin. This construct allows production of GFP-tagged IFT proteins that can be visualized either directly in live cells or using anti-GFP antibodies in immunofluorescence assays (Fig. 7A). GFP fusions can be also exploited in western blotting, protein purification, and immunoprecipitation experiments. Placing the gene under control of an inducible promoter enables a precise control of gene expression that can be very useful in case of toxicity of the gene product. It can also be exploited in studies of kinetics of different processes like assembly of protein complexes or flagellum elongation (Bastin et al., 1999). However, it can lead to overexpression, and the generic 30 -UTR sequence provided by the vector often does not correspond with the 30 -UTR of the actual gene. Given the absence of transcriptional control, the 30 -UTR often controls the stability, turn-over, and translation of mRNA

71

3. Tools for Analyzing Intraflagellar Transport in Trypanosomes (A)

(B)

Hindlll Clal

Notl EcoRV

Kpnl

EPprocyclin Ti Hindlll Ndel

rDNA spacer

Nsil

PFRAinter

PIFTC3 7000

6000

1000

AMP 6000

7664 bps

2000

5000

pPCPFR PIFTC3-Tag

Xbal

5000

epitope Tag

6633 bps 2000

PFRCinter

AMP 3000

4000

4000 3’UTRactin

1000

eGFP

pHD430eGFP-IFT52

3000 Puro

IFT52

Nhel BamHi Mlul Clal Ascl

BLE

Sall

3’UTR tubulin 5’UTRactin Srfl

3’UTRaldo BamHI

Mlul BstXl

Fig. 6 The pHD430 vector for inducible protein expression and the pPCPFR vector for constitutive expression of genes. AMP, ampicillin; BLE, phleomycin; PURO, puromycin antibiotic resistance genes, EPTi—modified inducible procyclin promoter; IFT52, PIFTC3—genes of interest; rDNA—locus of integration; PFR1inter, PFR2inter, PFR1, and PFR2—intergenic region fragments. The pPCPFR vector can be integrated in the PFR2 intergenic locus (NsiI digest), in the tubulin locus (BstXI digest), or in the gene of interest itself, leading to endogenous tagging.

(A)

(B)

Fig. 7 Expression of the GFP::IFT52 fusion protein with the pHD430 vector (A) or of the GFP::RABL5 fusion protein in the pPCPFR after integration in the PFR2 locus. GFP was visualized using anti-GFP antibody after methanol fixation (A) or direct GFP visualization. (A) A strong expression level can be achieved with pHD430 but sometimes with over-expression and accumulation of cytoplasmic material. (B) When using the pPCPFR expression vector, the amount of GFP::RABL5 is weaker but more homogenous. The cells were double-stained with the monoclonal antibody MAb22 to reveal the proximal region of the basal body (red). For color viewing please refer to Adhiambo et al. (2009). (See Plate no. 2 in the Color Plate Section.)

(Clayton and Hotz, 1996). To express the amount of proteins close to that produced in normal conditions, endogenous tagging of the gene is required. In this case, a gene fragment is selected according to the presence of a unique restriction site in its sequence, which is necessary for targeting site-specific recombination in the gene

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locus on the chromosome. It is cloned within the plasmid and can be fused to fluorescent markers or an epitope tag. Tagging of flagellar proteins allows detailed observation of protein turnover and localization during the parasite life cycle. For this purpose we built the pPCPFR vector (Fig. 6B). To tag the gene of interest, the Ty1 epitope has become a popular choice in trypanosomes but also in other parasites such as Leishmania and Toxoplasma. The Ty-tag is a 10-amino-acid sequence from the immunologically well-characterized major structural protein of the Ty1 virus-like particle of Saccharomyces cerevisiae (Brookman et al., 1995). The epitope is recognized by two different monoclonal antibodies, BB2 (IgG1) and TYG5 (IgA), that were tested in a variety of immunological conditions and do not cross-react with trypanosome proteins (Bastin et al., 1996). Thanks to polymerase II run-through activity, an efficient expression of the gene of interest can also be achieved by the integration of a plasmid at any of three loci present in the pPCPFR vector; the PFR1 and PFR2 intergenic regions, and the 30 -UTR of tubulin. The pPCPFR plasmid was used with success for tagging several IFT proteins (Adhiambo et al., 2009) (Fig. 7B).

3. Nucleofection: A Novel Transfection Technique for Efficient Plasmid Integration Usually introduction of vector DNA into parasite cells is obtained by a traditional electroporation technique (ten Asbroek et al., 1990). A high number of cells (~3 107) is suspended in Cytomix buffer (120 mM KCl, 0.15 mM CaCl2, 10 mM KH2PO4, 25 mM Hepes, 2 mM EDTA, 5 mM MgCl2, pH 7.6) and electroporated by one (BSF) or two (PCF) successive electro-pulses of 1.5 kV with a 25 µF capacitance to allow entry of the plasmid DNA (ten Asbroek et al., 1990). For procyclic trypanosomes, this method can give efficiencies of 103 to 106 (McCulloch et al., 2004; ten Asbroek et al., 1990) but is much less effective in BSFs, producing stable transformants with a frequency of only 107 to 108. Recently, an improved method was developed by Lonza (www.amaxa.com). The Amaxa Nucleofector® apparatus and its nucleofection kit containing cell-type-specific solution allow transfection of bloodstream parasites with an efficiency level increased to ~104 (Burkard et al., 2007). However, for procyclic trypanosomes, no significant differences in the number of transformants were reported when the two methods were compared (Burkard et al., 2007). From our experience (B. Rotureau and I. Subota, unpublished data), nucleofection is also more efficient in PCFs especially, for procyclic cells derived from bloodstream pleomorphic cell lines such as AnTat 1.1. Usually the traditional transfection generates heterogenous populations of clones that subsequently need to be subcloned. To obtain a clonal population, cells are diluted into 96-well plates to obtain 1 cell per well in specific conditioning medium that enables growth of a single procyclic trypanosome. Interestingly, with a similar starting number of parasites and an additional step of serial dilution of freshly “nucleofected” cells (Gresch et al., 2004; Hamm et al., 2002), it is possible to obtain more homogenous populations with a more uniform level of protein expression.

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D. Visualization of IFT and IFT-Related Phenotypes Immunofluorescence is a technique that enables localization of proteins in cells using specific antibodies. The protocols presented below were specifically adapted for imaging of IFT proteins in trypanosomes.

1. Methanol Fixation of Cells Procyclic cells at a concentration of 3–8 106/ml are spun down at 700 rcf, washed in PBS, and spread on poly-L-lysine-coated slides. When the sample is dry, cells are fixed in methanol at –20°C for 30 s to 5 min maximum and rehydrated for 15 min in PBS. The time of fixation depends on the target of the antibody. We noticed that fixation in methanol for longer than 5 min leads to a loss of signal with anti-IFT antibodies, in agreement with the fact that IFT proteins are not tightly attached to the flagellar skeleton as they need to shuttle along microtubules. In contrast, visualization of proteins in skeletal structures, such as the axoneme or paraflagellar rod (PFR), can accommodate longer fixation times.

2. Paraformaldehyde Fixation of Cells A 4% (w/v) solution of paraformaldehyde (PFA) in PBS is prepared fresh before each experiment. All incubation steps are performed in a humid chamber to avoid drying. Cells are washed in PBS and settled on poly-L-lysine-coated slides for 15 min. The PFA solution is added, slides are incubated for 10 min, and washed in PBS for 5 min. Cells may also be postfixed with methanol for ~30 s after PFA treatment, which has the double advantage of both permeabilizing the cells and also flattening them, facilitating observation. Alternatively, cell permeabilization may be performed by incubation with mild detergent for 10 min. For this purpose 0.1–1% Nonidet P-40 or 5% N-octyl-glucoside are used. Again, the choice of the detergent and its concentration should be adjusted according to peculiarities of each antibody. The samples are washed gently several times in PBS and blocked for 1 h in PBS containing 0.1–1% bovine serum albumin (BSA) (filtered before use).

3. Detergent Extraction of Cytoskeletons Cells are prepared and settled on a slide as described above. Samples are briefly exposed to 1% Nonidet P-40 (~1 min) in spindle stabilization buffer (4 M glycerol, 10 mM PIPES, pH 6.5, 10 mM MgCl2, 5 mM EGTA) and immediately rinsed with PBS to remove the excess detergent. Finally cytoskeletons are fixed in methanol at –20°C for 5 min or in 4% PFA for 10 min.

4. Immunofluorescence Assay After methanol/PFA fixation, rehydration in PBS, and eventual blocking, slides are incubated for 1 h with the appropriate dilution of primary antibodies in PBS-containing

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Daria Julkowska and Philippe Bastin

(A)

(B)

(C)

Fig. 8 Immunofluorescence assay on wild-type cells using anti-IFT172 monoclonal IgG antibody. (A) fixation in methanol. The signal is present in the region of the basal body and along the flagellum. (B) Fixation in PFA. Signal is weak and punctate, which may be explained by weak penetration of the antibody into the cell body and flagellum for this type of fixation. (C) Detergent extraction of cytoskeletons. The signal is still present in the basal body region and proximal part of the flagellum, suggesting stronger attachment of IFT172 protein in these regions of the flagellum. (See Plate no. 3 in the Color Plate Section.)

0.1% BSA (w/v), washed four times for 5 min in PBS, and incubated for 45 min with the secondary antibodies in a humid chamber. After a second series of washings (again 4 5 min with PBS), cells are stained with DAPI for visualization of the kinetoplast and nuclear DNA content. A drop of the Prolong Gold (Invitrogen-Molecular Probes, Eugene, OR, USA) antifading agent is applied to the slides before mounting the cover slip. Figure 8 shows an immunofluorescence experiment using the anti-IFT172 antibody with the three types of cell fixation. Table II presents primary and secondary antibodies useful for visualisation of IFT proteins, flagellum, basal body, and flagellar pocket structures. It is possible to perform double staining with two antibodies of the same species of origin. In that case, the procedure of incubation with antibodies (the primary/secondary couple) is repeated twice with an additional blocking step of 45 min in 0.1% BSA in PBS between the two series of incubations (Fig. 9). It is advisable to apply the “weakest” antibody first, as it

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3. Tools for Analyzing Intraflagellar Transport in Trypanosomes

Table II List of Antibodies Frequently Used for Analysis of IFT, Flagella, Basal Bodies, and Flagellar Pocket Name

Type

Species

Subclass

IFT Anti-IFT172

MAb

Mouse

IgG1

S S

Mouse Mouse

MAb MAb MAb MAb

Mouse Mouse Mouse Mouse

IgG2a IgG1 IgG1 IgG2b

MAb MAb

Mouse Rat

IgG2b IgG2a

Basal bodies MAb22 Anti-TBBC

MAb S

Mouse Rabbit

IgM

Flagellar pocket Anti-CRAM Anti-BILBO1

S S

Rabbit Mouse

Anti-PIFTF6/IFT144 Anti-RABL5 Flagellar structures MAb25 L8C4 L13D6 KMX-1 TAT1 YL1/2

Specificity

Immunogen origin

IFT172

Recombinant

PIFTF6/IFT144 RABL5

Recombinant Recombinant

Axonemal protein PFR2 PFR1þPFR2 Beta-tubulin

T. brucei T. brucei T. brucei Physarum polycephalum Alpha-tubulin T. brucei Tyrosinated alpha- T. brucei tubulin

Reference

T. Blisnick et al. (unpublished data) Adhiambo et al. (2009) Adhiambo et al. (2009) Pradel et al. (2006) Kohl et al. (1999) Kohl et al. (1999) Birkett et al. (1985) Woods et al. (1989) Kilmartin et al. (1982)

Basal body/TAC TBBC

T. brucei T. brucei

Bonhivers et al. (2008) Dilbeck et al. (1999)

CRAM FP collar

T. brucei Recombinant

Lee et al. (1990) Bonhivers et al. (2008)

TBBC, trypanosome basal body component; CRAM, cystein-rich acidic membrane; TAC, tripartite attachment complex (structure linking the basal body to the kinetoplast); S, serum; MAb, monoclonal antibody.

Fig. 9 Immunofluorescence assay with two mouse polyclonal antibodies. Antibodies target two different IFT proteins, IFT172 and RABL5. Cells were fixed in methanol. First incubation was performed with the anti-IFT172 antibody and revealed by the secondary anti-mouse coupled to Alexa 488 (green). Subsequently, slides were blocked for 45 min with 0.1% BSA in PBS, followed by a second incubation with anti-RABL5 and secondary anti-mouse Cy3 antibody (red). For color viewing please refer to Adhiambo et al. (2009). (See Plate no. 4 in the Color Plate Section.)

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will be easier to block compared to antibodies producing strong signal (e.g., against tubulin or PFR proteins).

5. Observation of IFT Particle Movement in the Flagellum The movement of IFT proteins fused to a fluorescent marker along the flagellum can be visualized in live parasites provided that cells are immobilized. This can be achieved by mixing a small amount of cells with 3% low melting point (LMP) agarose (BioRad, Hercules, CA, USA), melted previously at 37°C in 1% PBS and applying the mixture immediately to a slide. A cover slip is put and pressed gently to help agarose spreading. Cells are observed at 1000 magnification usually with an inverted DMR Leica microscope, and images are directly recorded with a Cohu 460LI camera connected to a DVD recorder or captured with a Cool Snap HQ numerical camera (Roper Scientific, Trenton, NJ, USA) using IPLab software. Using this video microscopy technique we were able to monitor the movement of IFT52 protein fused to GFP in the flagellum (Absalon et al., 2008b). Fluorescent moving particles were detected in both old and new flagellum. Both anterograde and retrograde events could be distinguished (see Movie 1, supplementary data; http://www.elsevierdirect.com/companions/9780123749734).

6. Scanning Electron Microscopy Scanning electron microscopy (SEM) observation of intact cells or of detergentextracted cytoskeletons is a powerful tool to analyze IFTRNAi mutant phenotypes (Fig. 10). Cells (107) are washed in PBS and fixed in 1 ml of 2.5% glutaraldehyde in PBS or cacodylate (0.1 M, pH 7.4) buffer for a minimum of 30 min. For preparation of cytoskeletons, cells are first settled on the slide for 30 min, treated with 1% Triton X100 or 1% Nonidet P40 for 10 min at 4°C, and washed three times in PBS prior to glutaraldehyde fixation. Subsequently samples are washed three times in cacodylate buffer and postfixed in 1% OsO4 for 1 h. After brief washing in water, cells are dehydrated in successive ethanol baths (25, 50, 75, 95%) for 5 min each time and finally twice in 100% ethanol for 10 min. After these dehydration steps, samples are critical-point dried (Emitech K850 or Balzers Union CPD30) and coated with a 15-nm layer of gold or palladium (Jeol JFC-1200 or Gatam Ion Beam Coater 681). Observations are performed with a Jeol 840A or a Jeol JM6700 F scanning microscope.

III. Perspectives Thanks to the numerous genetic and cellular tools developed over the last 10 years; trypanosomes have become attractive models to study the function of novel genes coding for flagellar proteins, especially those involved in IFT. Moreover, trypanosomes are fascinating organisms to study. Their flagellum, combining conserved and original

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1 µm

(B)

1 µm (A)

(C)

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Fig. 10 Scanning electron micrographs showing different phenotypes of IFT mutants. (A) WT cells modified from Absalon et al, 2008a; (B) IFT88RNAi (complex B) mutant showing short cells without a flagellum; (C) IFT140RNAi (complex A) mutant cells with a short dilated flagellum accumulating IFT particles reproduced from Absalon et al, 2008b. Scale bars are indicated.

features, will write another exciting page in the story of IFT. Indeed, understanding the mechanisms that govern the dramatic changes in flagellum length and positioning during the parasite cycle (Vickerman, 1985) is a challenge that will require understanding of how IFT is finely tuned. Recently, we have been able to monitor parasites expressing a fluorescent flagellar protein during the infection of tsetse flies (B. Rotureau and P. Bastin, unpublished data). Similar experiments where fluorescent IFT proteins are expressed open the road to real in vivo studies of the dynamics of the flagellum during the life cycle of trypanosomes and should bring exciting insight into IFT regulation. Acknowledgments D. J. was funded by ANR grants and a Roux fellowship. Work in the authors’ laboratory is funded by the Institut Pasteur, the CNRS, and ANR grants. We thank Sabrina Absalon for providing RT-PCR calibration data for Fig. 3 and the immunofluorescence image for Fig. 7A, and Thierry Blisnick for immunofluorescence and electron microscopy images for Figs. 9 and 10. We also thank Brice Rotureau and Ines Subota for critical review of this article.

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References Absalon, S., Kohl, L., Branche, C., Blisnick, T., Toutirais, G., Rusconi, F., et al. (2007). Basal body positioning is controlled by flagellum formation in Trypanosoma brucei. PLoS One 2(5), e437. Absalon, S., Blisnick, T., Bonhivers, M., Kohl, L., Cayet, N., Toutirais, G., et al. (2008a). Flagellum elongation is required for correct structure, orientation and function of the flagellar pocket in Trypanosoma brucei. J. Cell Sci. 121(Pt. 22), 3704–3716. Absalon, S., Blisnick, T., Kohl, L., Toutirais, G., Dore, G., Julkowska, D., et al. (2008b). Intraflagellar transport and functional analysis of genes required for flagellum formation in trypanosomes. Mol. Biol. Cell 19(3), 929–944. Adhiambo, C., Blisnick, T., Toutirais, G., Delannoy, E., and Bastin, P. (2009). A novel function for the atypical small G protein Rab-like 5 in the assembly of the trypanosome flagellum. J. Cell Sci. 122(Pt. 6), 834–841. Alsford, S., Kawahara, T., Glover, L., and Horn, D. (2005). Tagging a T. brucei RRNA locus improves stable transfection efficiency and circumvents inducible expression position effects. Mol. Biochem. Parasitol. 144(2), 142–148. Archer, S.K. (2009). Standard culture medium allows clonal dilution of Trypanosoma brucei procyclic cells after auto-conditioning. Mol. Biochem. Parasitol. 164(1), 100–103. Avidor-Reiss, T., Maer, A.M., Koundakjian, E., Polyanovsky, A., Keil, T., Subramaniam, S., et al. (2004). Decoding cilia function: Defining specialized genes required for compartmentalized cilia biogenesis. Cell 117(4), 527–539. Bastin, P., Bagherzadeh, Z., Matthews, K.R., and Gull, K. (1996). A novel epitope tag system to study protein targeting and organelle biogenesis in Trypanosoma brucei. Mol. Biochem. Parasitol. 77(2), 235–239. Bastin, P., MacRae, T.H., Francis, S.B., Matthews, K.R., and Gull, K. (1999). Flagellar morphogenesis: Protein targeting and assembly in the paraflagellar rod of trypanosomes. Mol. Cell Biol. 19(12), 8191–8200. Bastin, P., Ellis, K., Kohl, L., and Gull, K. (2000). Flagellum ontogeny in trypanosomes studied via an inherited and regulated RNA interference system. J. Cell Sci. 113(Pt. 18), 3321–3328. Benne, R., et al. (1986). Major transcript of the frameshifted coxII gene from trypanosome mitochondria contains four nucleotides that are not encoded in the DNA. Cell 46(6), 819–826. Berriman, M., Hall, N., Sheader, K., Bringaud, F., Tiwari, B., Isobe, T., et al. (2002). The architecture of variant surface glycoprotein gene expression sites in Trypanosoma brucei. Mol. Biochem. Parasitol. 122 (2), 131–140. Berriman, M., Ghedin, E., Hertz-Fowler, C., Blandin, G., Renauld, H., Bartholomeu, D.C., et al. (2005). The genome of the African trypanosome Trypanosoma brucei. Science 309(5733), 416–422. Birkett, C.R., Foster, K.E., Johnson, L., and Gull, K. (1985). Use of monoclonal antibodies to analyse the expression of a multi-tubulin family. FEBS Lett. 187, 211–218. Bonhivers, M., Landrein, N., Decossas, M., and Robinson, D.R. (2008). A monoclonal antibody marker for the exclusion-zone filaments of Trypanosoma brucei. Parasit Vectors 1(1), 21. Bonhivers, M., Nowacki, S., Landrein, N., and Robinson, D.R. (2008). Biogenesis of the trypanosome endoexocytotic organelle is cytoskeleton mediated. PLoS Biol. 6(5), e105. Boothroyd, J.C. (1985). Antigenic variation in African trypanosomes. Annu. Rev. Microbiol. 39, 475–502. Briggs, L.J., Davidge, J.A., Wickstead, B., Ginger, M.L., and Gull, K. (2004). More than one way to build a flagellum: Comparative genomics of parasitic protozoa. Curr. Biol. 14(15), R611–R612. Bringaud, F., Robinson, D.R., Barradeau, S., Biteau, N., Baltz, D., and Baltz, T. (2000). Characterization and disruption of a new Trypanosoma brucei repetitive flagellum protein, using double-stranded RNA inhibition. Mol. Biochem. Parasitol. 111(2), 283–297. Broadhead, R., Dawe, H.R., Farr, H., Griffiths, S., Hart, S.R., Portman, N., et al. (2006). Flagellar motility is required for the viability of the bloodstream trypanosome. Nature 440(7081), 224–227. Brookman, J.L., Stott, A.J., Cheeseman, P.J., Burns, N.R., Adams, S.E., Kingsman, A.J., et al. (1995). An immunological analysis of Ty1 virus-like particle structure. Virology 207(1), 59–67.

3. Tools for Analyzing Intraflagellar Transport in Trypanosomes

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Brun, R., and Schonenberger, M. (1979). Cultivation and in vitro cloning or procyclic culture forms of Trypanosoma brucei in a semi-defined medium. Short communication. Acta Trop. 36(3), 289–292. Burkard, G., Fragoso, C.M., and Roditi, I. (2007). Highly efficient stable transformation of bloodstream forms of Trypanosoma brucei. Mol. Biochem. Parasitol. 153(2), 220–223. Campbell, D.A., Thomas, S., and Sturm, N.R. (2003). Transcription in kinetoplastid protozoa: Why be normal?. Microbes Infect. 5(13), 1231–1240. Cerutti, H., and Casas-Mollano, J.A. (2006). On the origin and functions of RNA-mediated silencing: From protists to man. Curr. Genet. 50(2), 81–99. Clayton, C.E. (2002). Life without transcriptional control? From fly to man and back again. EMBO J. 21(8), 1881–1888. Clayton, C., and Hotz, H.R. (1996). Post-transcriptional control of PARP gene expression. Mol. Biochem. Parasitol. 77(1), 1–6. Cole, D.G. (2003). The intraflagellar transport machinery of Chlamydomonas reinhardtii. Traffic 4(7), 435–442. Cunningham, I. (1977). New culture medium for maintenance of tsetse tissues and growth of trypanosomatids. J. Protozool. 24(2), 325–329. Davidge, J.A., Chambers, E., Dickinson, H.A., Towers, K., Ginger, M.L., McKean, P.G., et al. (2006). Trypanosome IFT mutants provide insight into the motor location for mobility of the flagella connector and flagellar membrane formation. J. Cell. Sci. 119(Pt. 19), 3935–3943. Dilbeck, V., Berberof, M., Van Cauwenberge, A., Alexandre, H., and Pays, E. (1999). Characterization of a coiled coil protein present in the basal body of Trypanosoma brucei. J. Cell Sci. 112(Pt. 24), 4687–4694. Durand-Dubief, M., and Bastin, P. (2003). TbAGO1, an argonaute protein required for RNA interference, is involved in mitosis and chromosome segregation in Trypanosoma brucei. BMC Biol. 1, 2. Durand-Dubief, M., Kohl, L., and Bastin, P. (2003). Efficiency and specificity of RNA interference generated by intra- and intermolecular double stranded RNA in Trypanosoma brucei. Mol. Biochem. Parasitol. 129 (1), 11–21. Gresch, O., Engel, F.B., Nesic, D., Tran, T.T., England, H.M., Hickman, E.S., et al. (2004). New non-viral method for gene transfer into primary cells. Methods 33(2), 151–163. Griffiths, S., Portman, N., Taylor, P.R., Gordon, S., Ginger, M.L., and Gull, K. (2007). RNA interference mutant induction in vivo demonstrates the essential nature of trypanosome flagellar function during mammalian infection. Eukaryot. Cell 6(7), 1248–1250. Hamm, A., Krott, N., Breibach, I., Blindt, R., and Bosserhoff, A.K. (2002). Efficient transfection method for primary cells. Tissue Eng. 8(2), 235–245. Hertz-Fowler, C., Peacock, C.S., Wood, V., Aslett, M., Kerhornou, A., Mooney, P., et al. (2004). GeneDB: A resource for prokaryotic and eukaryotic organisms. Nucleic Acids Res. 32(Database issue), D339–D343. Hirumi, H., and Hirumi, K. (1989). Continuous cultivation of Trypanosoma brucei blood stream forms in a medium containing a low concentration of serum protein without feeder cell layers. J. Parasitol. 75(6), 985–989. Kilmartin, J.V., Wright, B., and Milstein, C. (1982). Rat monoclonal antitubulin antibodies derived by using a new nonsecreting rat cell line. J. Cell Biol. 93(3), 576–582. Kohl, L., and Bastin, P. (2005). The flagellum of trypanosomes. Int. Rev. Cytol. 244, 227–285. Kohl, L., Robinson, D., and Bastin, P. (2003). Novel roles for the flagellum in cell morphogenesis and cytokinesis of trypanosomes. EMBO J. 22(20), 5336–5346. Kohl, L., Sherwin, T., and Gull, K. (1999). Assembly of the paraflagellar rod and the flagellum attachment zone complex during the Trypanosoma brucei cell cycle. J. Eukaryot. Microbiol. 46(2), 105–109. LaCount, D.J., Bruse, S., Hill, K.L., and Donelson, J.E. (2000). Double-stranded RNA interference in Trypanosoma brucei using head-to-head promoters. Mol. Biochem. Parasitol. 111(1), 67–76. Lee, M.G., Bihain, B.E., Russell, D.G., Deckelbaum, R.J., Van der Ploeg, L.H. (1990). Characterization of a cDNA encoding a cysteine-rich cell surface protein located in the flagellar pocket of the protozoan Trypanosoma brucei. Mol. Cell Biol. 10(9), 4506–4517. McCulloch, R., Vassella, E., Burton, P., Boshart, M., and Barry, J.D. (2004). Transformation of monomorphic and pleomorphic Trypanosoma brucei. Methods Mol. Biol. 262, 53–86.

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CHAPTER 4

Schmidtea mediterranea: A Model System for Analysis of Motile Cilia Panteleimon Rompolas, Ramila S. Patel-King, and Stephen M. King Department of Molecular, Microbial & Structural Biology, University of Connecticut Health Center, Farmington, Connecticut 06030-3305

Abstract I. Introduction II. Materials and Methods A. Maintaining and Expanding a Healthy Population of Schmidtea mediterranea B. RNAi of Ciliary Genes in Schmidtea mediterranea C. Imaging of Planarian Cilia D. Motility Assays III. Discussion IV. Summary Acknowledgments References

Abstract Cilia are cellular organelles that appeared early in the evolution of eukaryotes. These structures and the pool of about 600 genes involved in their assembly and function are highly conserved in organisms as distant as single-cell protists, like Chlamydomonas reinhardtti, and humans (Silflow and Lefebvre, 2001). A significant body of work on the biology of cilia has been produced over the years, with the help of powerful model organisms including C. reinhardtti, Caenorhabditis elegans, sea urchins, and mice. However, specific limitations of these systems, especially regarding the ability to efficiently study gene loss-of-function, warrant the search for a new model organism to study cilia and cilia-based motility. Schmidtea mediterranea is a species of planarian (Class: Tubellaria) with a well-defined monostratified ciliated epithelium, which METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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contributes to the motility of the organism, in addition to other more specialized ciliary structures. The use of S. mediterranea as an experimental system to study stem cell biology and regeneration has led to a recently sequenced genome and to the development of a wide array of powerful tools including the ability to inhibit gene expression via RNA interference. In addition, we have developed and describe here a number of methods for analyzing motile cilia in S. mediterranea. Overall, S. mediterranea is a highly versatile, easy to maintain, and genetically tractable organism that provides a powerful alternative model system for the study of motile cilia.

I. Introduction Planarians have been studied extensively over the last century for their regenerative properties and more recently as a model for stem cell biology. Planarians are triploblastic, acoelomate, and bilaterally symmetrical metazoans that belong to the class Tubellaria. Together with flukes (Trematoda) and tapeworms (Cestoda) they form the phylum of Platyhelminthes; however, in contrast to the other two members of the group, planarians are free-living nonparasitic organisms most often found in rivers, streams, and ponds. Their body consists of tissues that are derived from all three basic germ layers (endoderm, mesoderm, and ectoderm) and presents with only one opening that leads to a three-branched digestive system but no anus. In addition, planarians have a simple central nervous system (CNS) that includes brain ganglia and two nerve cords that bilaterally span the length of the body and are connected by a set of commissural neurons (Cebria et al., 2002). Specialized receptors, with the most prominent being a set of eye spots in the head region, are used to effectively respond to environmental stimuli. Planarians are hermaphrodites, possessing testes, ovaries, and copulatory apparatus and have the capacity for sexual reproduction through crossfertilization (Newmark and Sanchez Alvarado, 2002; Sanchez Alvarado, 2003). Planarians have no apparent circulatory or respiratory systems; however, they do possess muscles which line the inner wall of the body in longitudinal, diagonal, and circular orientations. The mesenchyme of the planarian body is populated with numerous undifferentiated pluripotent cells, known as neoblasts. Neoblasts account for 25–30% of all cells in the planarian body and every tissue is ultimately derived from these cells. The capacity of this cell type to self-renew, rapidly proliferate, and migrate to sites of injury gives planarians the unmatched ability to regenerate. This attribute has made planarians the model of choice for the study of stem cell biology and regeneration. Schmidtea mediterranea (Fig. 1, upper panel) has recently emerged as the planarian species of choice for scientific research and its genome has been sequenced to 11.6 coverage. The sexual strain of S. mediterranea that was used for sequencing has a stable diploid genome consisting of eight chromosomes (4.8 108 bp). A wide array of tools has also been developed for the genetic manipulation of S. mediterranea including inhibition of gene expression via RNA interference (RNAi) (Sanchez Alvarado and Newmark, 1999) and exogenous gene expression (Gonzalez-Estevez et al., 2003). For more

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Fig. 1

Schmidtea mediterranea cilia. A typical S. mediterranea adult flatworm has an average length of 10–20 mm and displays bilateral body symmetry and cephalization. e, eyespots; au, auricles; ph, pharynx. Motile cilia are found on the entire ventral surface of the planarian body. In a dorsal view, cilia are mostly visible on the lateral sides of the head region.

general information on the biology and regeneration of planarians, see Reddien and Sanchez Alvarado (2004) and Saló (2006). Schmidtea mediterranea like other planarians is able to move using a characteristic gliding motion in which the animal displays little or no muscular effort. It has long been thought that this mode of locomotion is mediated via motile cilia (Fig. 1, lower panels) which populate the ventral surface of the planarian body and are in contact with the substrate. Indeed, the planarian ventral epidermis consists of a monostratified multiciliated epithelium. Between the ciliated cells of the epidermis are located gland cells that secrete mucus which forms a substrate on which the animal is able to glide. Using scanning electron microscopy (SEM) we studied the appearance and distribution

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of the cilia in S. mediterranea. We found that the entire ventral epithelium consists of cells that are largely ciliated. However, the number of cilia appears much larger around the head compared to other regions of the body. In contrast, the dorsal surface is almost completely devoid of cilia with only small loci that appear to house clusters of cilia. The cilia lining the ventral surface of the S. mediterranea body are about 10 µm in length and are motile. Using high-speed video microscopy we measured the beat frequency of the motile cilia that are easily visible on the lateral sides of the head region and found that they beat at ~20 Hz. Cross-section transmission electron microscopy (TEM) images of planarian motile cilia reveals the typical axoneme structure with a 9 þ 2 microtubule arrangement and other structures found in motile cilia from other organisms including radial spokes and outer and inner dynein arms. Other types of ciliary structures with sensory and secretory roles are also present in planarians. For example, flame cells which are specialized excretory cells have continuously beating cilia. Comparative analysis of the S. mediterranea genome revealed a great number of highly conserved genes with known ciliary function, including genes present in humans. In this chapter we describe general methods for keeping and maintaining S. mediterranea flatworms in the laboratory and ways to study gene expression via RNAi. In addition, we discuss several protocols that we have developed in our laboratory for the analysis of cilia and cilia-based motility using S. mediterranea as a model organism.

II. Materials and Methods A. Maintaining and Expanding a Healthy Population of Schmidtea mediterranea Planarians are most often found in freshwater ponds, streams, and springs usually hiding in dark places during the day. A colony of S. mediterranea can be easily maintained and expanded in the laboratory if appropriate conditions that closely resemble their natural habitat are met. Unfortunately, as of now there are no commercial vendors or a central facility for distributing S. mediterranea clones. However, a founding colony can be obtained from the expanding number of laboratories that are using S. mediterranea as a model organism for their research.

1. General Culture Conditions Planarians can be kept in food-grade plastic containers (Rubbermaid or Ziploc) of appropriate size, filled with culture medium to about half of the container’s total capacity. A 2-l rectangular container can accommodate up to 500 small- to mediumsize flatworms (1–10 mm in length) but the size of the colony should be carefully monitored to avoid stress from overcrowding. New containers can be rinsed with deionized water and wiped clean but use of soap or other detergents may leave residues harmful to the animals and therefore should be avoided. Planarian colonies should be stored in the dark, although occasional exposure to direct light (e.g., during feeding)

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will not harm them. It is also best if the lids of their containers are left half open to permit air circulation; this will allow sufficient gas exchange thus maintaining appropriate levels of dissolved oxygen in the culture medium. In addition, S. mediterranea is best maintained at temperatures ranging between 17 and 20°C. Although planarians can tolerate a greater range of temperatures, during long-term storage, keeping the temperature below 20°C discourages growth of bacteria or other pathogens that can contribute to fouling of the culture medium. An appropriate place that meets all of the above culture conditions for S. mediterranea is a temperature-controlled incubator with sufficient air flow.

2. Culture Medium The appropriate culture medium for S. mediterranea species is a 1 solution of Montjuïch salts (1.6 mM NaCl, 1.0 mM CaCl2, 1.0 mM MgSO4, 0.1 mM MgCl2, 0.1 mM KCl, 1.2 mM NaHCO3) (Cebria and Newmark, 2005). This solution can be prepared as a 5 or 10 concentrate and kept at 4°C for up to a month. However, since there is no active buffering system in this medium, the pH may change over time and some salts may precipitate. Therefore, the conditions should be carefully monitored and the medium should be made fresh when necessary.

3. Feeding and Cleaning The best type of food for planarians maintained for experimental purposes is calf liver. To prepare the food, fresh organic calf liver is obtained, thoroughly rinsed with tap water, and cleaned of excess fatty and connective tissues. The liver is then cut in small pieces and homogenized using a blender. If necessary, the homogenate can be passed through a food strainer to remove any remaining chunks of connective tissue. The liver puree is then aliquoted in 15-ml Falcon tubes and stored at –80°C for up to a year or until used. Planarians are fed at least once a week or more frequently if we are aiming to expand the colony. The aliquot of liver paste is first thawed and a transfer pipette is used to place a small amount at the bottom of the container. Many times the liver will float to the surface and away from the reach of worms; therefore, it is best that some of the medium is first removed from the container before feeding. Within 1 h the worms should have finished eating and swum away from the food appearing lighter in color from the ingested liver. Any food residues are then removed from the container using a 10-ml disposable pipette before the rest of the soiled medium is carefully poured off to a waste beaker. Using a 15-ml pipette, squirt some fresh medium to rinse the bottom of the container and to force all worms to detach and gather in one corner. Carefully discard the rinse and then use a Kimwipe to wipe off the remaining food debris as well as any mucus or other residues from the sides and bottom of the container. Fill the container with fresh medium and place it back in the incubator. This cleaning regimen can be repeated as necessary and should be sufficient to keep the colony clean and healthy.

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4. Expanding the Colony There are two different strains of S. mediterranea: a sexual and an asexual strain which is a result of a chromosome translocation. Animals of the sexual strain although hermaphrodites will cross-fertilize and deposit eggs that hatch after approximately 3 weeks to reveal young juvenile worms of about 1 mm in length. In contrast, asexual animals can only reproduce by fission, in which case the flatworm will stretch its body until a fission event occurs resulting in the front three quarters of the body separating from a small tail piece; each body part will then individually regenerate to give two new flatworms. Although both strains of S. mediterranea have the capacity to reproduce when healthy and well-fed it may be easier and faster to cut the larger flatworms in order to expand the colony with genetically identical animals. In this case planarians are fed several times per week in order to grow faster in size. Individuals that have reached a suitable size (10–20 mm in length) are collected 3 days after the last feeding session and placed in a separate container with fresh medium. Each flatworm is cut with a scalpel into two to three pieces which are then left in the incubator for 2–3 weeks to regenerate into full animals. During this period the worms are not fed and the medium is changed regularly and the container cleaned of tissue debris that results from the regeneration process.

5. Troubleshooting Although S. mediterranea and planarians in general are relatively hardy animals, able to tolerate a great range of environmental conditions in addition to starvation and injury, sometimes the colony can be faced with some serious problems. Issues most commonly occur because of poor quality of the culture medium or because of a pathogen infestation. Signs of a stressed colony may include: (1) oversecretion of mucus causing the flatworms to attach firmly to the substrate, (2) animals do not eat and stay on the sides of the container and close to the surface of the medium, (3) lesions are visible usually in the head region and the colony develops a strong foul odor, and (4) planarians are curled and lay on their side and in unusual positions. To help a colony recover from such a stressed state, the containers should be thoroughly cleaned and the medium should be replaced with fresh several times per week. During this period the flatworms should not be fed to discourage the further growth of protozoa and bacteria. In order to fight secondary bacterial infections, the culture medium can be supplemented with gentamicin (50 µg/ ml). Finally, caution should be exercised by handling the sick cultures separately from the healthy ones in order to avoid cross-contamination. B. RNAi of Ciliary Genes in Schmidtea mediterranea

1. Bioinformatics Sequencing of the S. mediterranea genome is currently under way at the Washington University Genome Sequencing Center (http://genome.wustl.edu/genome.cgi?GENOME=Schmidtea%20mediterranea) and has achieved 11.6 coverage. The clonal inbred line utilized for genome sequencing was developed in the laboratory of

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Dr Sánchez-Alvarado and is a derivative of a sexual strain collected by Dr Maria Pala on the Mediterranean island of Corsica. Blast searches of S. mediterranea (taxid: 79327) genes are available through the National Center for Biotechnology Information (NCBI) (http://blast.ncbi.nlm.nih.gov/Blast.cgi). In addition, the S. mediterranea genome database (SmedGD—http://smedgd.neuro.utah.edu/) introduces valuable tools for genomic analysis as well as information about gene expression and established RNAi phenotypes (Robb et al., 2008). Extensive express sequence tag (EST) libraries have also been developed and are publicly available (Sanchez Alvarado et al., 2002; Zayas et al., 2005). Since S. mediterranea is a relatively new experimental model organism and in order to avoid the confusion experienced in other model systems from multiple names given to single genes, specific guidelines have been published for assigning gene and protein names as well as for describing RNAi experiments (Reddien et al., 2008). In brief, S. mediterranea gene names are lowercase, italicized, and with the prefix “Smed” followed by a dash before the conventionally used name (e.g., Smed-genex). Proteins are assigned the name of the planarian gene from which they are derived but using nonitalics, uppercase fonts (e.g., SMED-GENEX). Finally, RNAi experiments are denoted with the suffix (RNAi) following the name of the specific gene that is being targeted (e.g., Smed-genex(RNAi)).

2. Synthesis of dsRNA Gene expression can be inhibited in planarians as in other organisms via doublestranded RNA (dsRNA)-mediated RNAi (Fire et al., 1998; Sanchez Alvarado and Newmark, 1999). dsRNA for inhibition of gene expression is synthesized using a vector (pL4440) with two opposing T7 RNA polymerase promoters that flank a multiple cloning site (Timmons and Fire, 1998). Alternatively, the pDONRdT7 vector can be used which is derived from pL4440 but also contains two class I, T7 transcriptional terminators that reportedly give better overall expression of dsRNA (Reddien et al., 2005). pDONRdT7 also utilizes a modified Gateway cloning strategy (Invitrogen, Carlsbad, CA, USA) to facilitate the transfer of the cDNA sequence. A cDNA fragment of about 200–600 bp is usually sufficient to effectively suppress gene expression. The resulting plasmid is then used to transform HT115 (DE3) Escherichia coli cells that are deficient in RNaseIII, which prevents the degradation of the newly synthesized dsRNA (Timmons et al., 2001).

3. Delivery of dsRNA via Bacterial Feeding A starter culture of the transformed cells is inoculated in 4 ml of 2xYT medium, supplemented with the appropriate antibiotics and grown at 37°C for 16 h. The culture is then diluted 1:10 in fresh, prewarmed 2xYT medium and grown to OD595 = 0.4. Expression of dsRNA is induced with 0.1–1 mM isopropyl thiogalactoside (IPTG) at 37°C for 2–3 h. A pellet corresponding to 2 ml of the induced bacterial culture is mixed with 50 µl of liver homogenate that was previously diluted 1:1 with distilled water. 1 µl of food dye (McCormick, Sparks, MD, USA) is also added to the artificial food mix to monitor the course of the feeding (Fig. 2). This quantity is usually enough to feed a group of 10 averagesized flatworms. As the food is digested within the course of the next few days, bacteria are

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Fig. 2 RNAi in S. mediterranea. Inhibition of gene expression is possible in S. mediterranea via dsRNAmediated RNAi. A small cDNA sequence is placed in a vector between two flanking T7 promoters and expression of dsRNA is induced in Escherichia coli cells. Bacterial pellets are mixed with liver homogenate and food dye, and used to feed a group of flatworms (A, B). Planarians almost immediately commence feeding and their food intake can be monitored by the change in the body color (C, D). The inset in panel C shows a flatworm protruding its pharynx (white) and feeding on the artificial food mix.

degraded and the dsRNA is released and absorbed by cells throughout the body. This regimen is then repeated every 2 days or until sufficient suppression of gene expression is achieved. RNAi-induced phenotypes usually start to appear after 2 weeks from initiating the bacterial feeding; however, the actual timing and levels of inhibition will ultimately depend on the actual gene. The same set of parameters (induction time and temperature, concentration of IPTG, type of media, etc.) that can give variable results in protein expression using an inducible T7 promoter can be also optimized in this system. Glycerol stocks of transformed HT115 (DE3) cells can be stored at –80°C in order to be used again in future experiments; however, freshly transformed cells tend to give the best results.

4. Evaluation of Gene Expression Semiquantitative reverse transcription polymerase chain reaction (RT-PCR) can be used to assess the level of gene knockdown after performing RNAi in planarians. In

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this case two to three animals of about 10–20 mm in length are removed from the group and total RNA is isolated using Trizol (Invitrogen) following the manufacturer’s recommendations. An amount of 0.5–1 µg of purified planarian RNA is used to set up a 10-µl firststrand cDNA synthesis reaction with cloned avian myeloblastosis virus (AMV) reverse transcriptase (New England Biolabs, Ipswich, MA, USA), following the instructions provided by the manufacturer. PCR amplification using primers specific for the gene of interest is performed for 20, 25, and 30 cycles (depending on the abundance of the transcript) and the samples resolved on agarose gels and stained with ethidium bromide. Alternatively, gene expression levels can be monitored via northern blot analysis or using optimized methods for whole-mount in situ hybridization (Pearson et al., 2009).

C. Imaging of Planarian Cilia

1. Immobilization and Fixation A major challenge when sacrificing and fixing planarians is that they have a tendency to contract, swell, swivel, or curl their body when killed with conventional fixatives, while also secreting large quantities of mucus which collectively may hinder the visualization of cilia. A very effective method for simultaneously relaxing and fixing planarians while preventing the secretion of mucus was described by Dawar (1973). The relaxant formula consists of 1% HNO3, 0.8325% formaldehyde, and 50 mM MgSO4. The animals should be placed directly into the relaxant solution at room temperature. Initially, the flatworms may contract and twist but eventually their body flattens and relaxes until they are completely immobilized by the fixative. Fixation is complete within 24 h at which point the mucus is almost completely dissolved from the surface of the epidermis. After treatment with the relaxant, other fixatives may be used to preserve specific ultrastructures before samples are processed for optical or electron microscopy.

2. Immunofluorescence In evaluating the loss-of-function effects of ciliary genes in planarians, it is crucial to be able to visualize the levels and distribution of their respective proteins on the cilia. Unfortunately, since the use of S. mediterranea for experimental purposes is still in its infancy a very limited number of antibodies raised against planarian proteins are currently available. However, a great number of antibodies generated against highly conserved proteins from other species were found to crossreact with epitopes in S. mediterranea tissues (Robb and Sanchez Alvarado, 2002). We have used an antibody against a-tubulin (clone B-5-1-2, Sigma, St. Louis, MO, USA) in order to visualize the cilia on the planarian body by means of immunofluorescence. Whole animals are first killed and fixed for 16 h in relaxant fixative, briefly washed with phosphate-buffered saline (PBS), pH 7.2, and postfixed with freshly made 4% paraformaldehyde in PBS, pH 7.2. After fixation, samples are washed with three changes of PBS and permeabilized with 1% Igepal CA-630 (Sigma, St. Louis, MO, USA) in PBS, pH 7.2, for 10 min. After briefly rinsing with

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PBS, samples are treated for 1 h at room temperature with blocking buffer containing 3% normal goat serum, 1% bovine serum albumin (BSA), 1% cold water fish skin gelatin, 0.1% Igepal CA-630, and 0.05% Tween-20 in PBS pH 7.2. All antibodies used for immunofluorescence are diluted in PBS buffer containing 1% BSA, 0.1% cold-waterfish gelatin, and 0.05% Tween-20. Samples are treated with a 1:100 dilution of primary antibody against a-tubulin (clone B-5-1-2, Sigma) for 16 h at 4°C, washed with four changes of PBS and incubated with Alexa Fluor 488-conjugated antimouse secondary antibody (Invitrogen, Carlsbad, CA, USA) for 1 h. After washing briefly with PBS the samples are mounted whole on glass slides using a glycerol-based mounting medium containing 1,4-diazabicyclo(2,2,2)octane (DABCO) (Sigma, St. Louis, MO, USA) as an antifade agent and silicone spacers to preserve the integrity of the body before placing a coverslip on top. Stained cilia on the lateral edges of the head region or on the ventral surface can be visualized by means of confocal microscopy (Fig. 3).

3. Scanning Electron Microscopy Planarians prepared for electron microscopy are first killed in relaxant fixative at room temperature. When they are completely immobilized the relaxant fixative is replaced once with fresh and then the samples are left at room temperature for 16–24 h during which time the mucus that can hinder the visualization of the cilia should be completely removed. The relaxant fixative is replaced with two changes of 2.5% glutaraldehyde in 0.1 M sodium cacodylate, pH 7.4 (EM Sciences, Hatfield, PA, USA), and the samples are fixed for 16– 24 h at 4°C. Next, the samples are briefly washed with four changes of 0.1 M sodium cacodylate, pH 7.4, and placed in 1% osmium tetroxide for 1 h in the dark, at room temperature. Afterward, the samples are washed again with four changes of 0.1 M sodium cacodylate, pH 7.4, and dehydrated through a series of 10, 25, 50, 70, 85, 95, and 100% ethanol solutions. Dehydrated worms are dried at the critical point (Autosamdri-815, Series A, Tousimis Research Corp., Rockville, MD, USA) and mounted with carbon tape and colloidal silver paint. Samples are sputter-coated (Cressington 208 HR Sputter Coater, Ted Pella, Redding, CA, USA) before being placed in the scanning electron microscope (Quanta 200 FEG SEM, FEI, Hillsboro, OR, USA) for imaging (Fig. 4). For more detailed information regarding SEM, see Chapter 4 by SanAgustin et al., in volume 91 of this series. D. Motility Assays

1. Imaging of Live Cilia In the S. mediterranea body as in other planarians, the ratio of cilia to total surface area is relatively high. However, the great majority of the cilia are present on the ventral surface and in contact with the substrate. This, combined with the fact that the animal is in constant motion, especially when confined and illuminated, makes the imaging of live cilia a challenging venture. Under the microscope the most visible cilia on intact flatworms are those on the lateral sides of the head region (Fig. 1, lower panels). It is possible to image the beating of these cilia using differential interference contrast (DIC) microscopy, coupled with a high-frame-rate digital camera (Fig. 5). In

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Fig. 3 Immunofluorescence microscopy of S. mediterranea cilia. Planarians are killed in relaxant fixative, postfixed with paraformaldehyde, treated with antibody against a-tubulin, and the dorsal surface is visualized by wide-field (A) and confocal (B–E) microscopy. Cilia are specifically stained with the a-tubulin antibody (C–E). The bright spots visible on the dorsal surface and particularly around the edges are sites of mucus secretion which tend to absorb the fluorescent probes (A).

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Fig. 4 Electron microscopy of S. mediterranea cilia. Planarians are killed in relaxant fixative, postfixed with glutaraldehyde and osmium tetroxide, and examined by SEM. The entire ventral surface of the planarian body consists of a highly ciliated epithelium. The middle and right panels show magnified views of central (V) and side (S) regions of the planarian ventral surface. The inset on the right of the far right panel shows a cross section of two S. mediterranea cilia visualized by TEM.

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Fig. 5 Measurement of S. mediterranea ciliary beat frequency. A live animal was placed on a glass slide and confined in place using silicone spacers and a coverslip. The beat frequency of these cilia was measured by high-speed video microscopy capturing 500 frames/s under DIC optics. This panel shows sequential frames (4 ms apart) from a decompiled movie. The cilia complete a full beat cycle in 44–60 ms (~20 Hz).

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this case a live animal is placed within a small drop of planarian medium on a glass slide and confined in place using silicone spacers and a coverslip (alternatively, two layers of parafilm may be used as spacers without danger of squashing an averagesized flatworm). Using DIC optics, under 60 magnification, focus on the sides of the head region in order to image the beating of the cilia. The shallow depth of field combined with the constant movement of the animal may hinder attempts to focus on the cilia. On some occasions it may be helpful to cut a flatworm and use only the head portion for live imaging, since that is the region where most of the motile cilia can be seen. In addition, this will minimize the effect of the constant muscle movement which hinders efforts to focus on the beating cilia. Unfortunately, as of now, there is no reliable method for immobilizing planarians without obstructing the beating of the cilia.

2. Measuring Gliding Motility It has long been hypothesized that the beating of cilia contributes to the gliding motility of planarians. In this case mutations that affect the structure or function of cilia should have an impact on the motility of the entire organism. It is possible to study such effects by measuring the gliding velocity of planarians. To do this we have used a high-resolution digital video camera with a lens that is able to focus on a relative short distance (MiniVID, LW Scientific, Lawrenceville, GA, USA). A group of live animals is placed in a clean plastic container (a Ziploc container or a Petri dish is usually sufficient), with fresh medium and left there for at least 24 h to acclimate. The flatworms are then placed under the camera so that the entire container is included in the optical field. A mild fluorescent illumination can be used while taking care not to create any glare for the camera. Planarians may not move without a stimulus therefore before starting to record the movie the animals are carefully agitated by squirting a small stream of medium using a transfer pipette. A series of short movies is recorded and the distance covered by each individual flatworm is measured. We utilize Metamorph (MDS Analytical Technologies, Sunnyvale, CA, USA) as it allows us to track the flatworms and calculate their average gliding velocity (Fig. 6).

III. Discussion To test whether S. mediterranea might provide a useful model for the analysis of motile cilia, we used RNAi to inhibit the expression of genes that are essential for cilia formation and maintenance. Specifically, we made RNAi vectors against Smed-ift88 and Smed-kif3a. ift88 encodes a protein that is a component of intraflagellar transport (IFT) complex B, and kif3a encodes one motor unit of the kinesin that powers anterograde IFT; both genes are essential for cilia biogenesis and mutations that affect the normal levels of their respective proteins result in short or diminished cilia

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Fig. 6 Measurement of S. mediterranea gliding velocity. Groups of flatworms are placed in separate containers and their movement is monitored by digital video recording (left panels). The individual tracks are analyzed and the average gliding velocity is measured using Metamorph software (right panels). Gliding velocity is considerably lower in Smed-ift88(RNAi) flatworms compared to controls. (See Plate no. 5 in the Color Plate Section.)

(Marszalek et al., 1999; Pazour et al., 2000). Both Smed-ift88 and Smed-kif3a exhibit a high degree of sequence homology compared to their respective human genes. Groups of planarians were fed three times a week with bacteria expressing Smed-ift88(RNAi), Smed-kif3a(RNAi), and the empty pL4440 vector, respectively, over a period of 3 weeks. After that period three animals were removed from each group in order to estimate expression levels via semiquantitative RT-PCR. We found that the expression levels of Smed-ift88 and Smed-kif3a were significantly lower

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compared to that of the control. The rest of the flatworms were used to analyze cilia formation and the potential impact on the organism’s motility. Microscopic analysis revealed a significant decrease in the number and overall length of cilia present on the ventral surface of the Smed-ift88(RNAi) and Smed-kif3a(RNAi) animals while cilia on the control flatworms remained largely unchanged. These results were in agreement with what has been reported in other organisms and consistent with the predicted role of these two genes in planarian ciliogenesis. Next, we investigated the impact of compromised cilia formation on the motility of Smed-ift88(RNAi) and Smed-kif3a(RNAi) animals. To calculate the average gliding speed, we measured the distance that flatworms in each group covered over a period of time. We found that the average velocity of Smed-ift88(RNAi) and Smed-kif3a(RNAi) animals was significantly lower compared to the control group. Most importantly, when we examined individual animals from the Smed-ift88 (RNAi) and Smed-kif3a(RNAi) groups more closely, we found that the means by which they moved was significantly different compared to the control or wild type. Specifically, both Smed-ift88(RNAi) and Smed-kif3a(RNAi) animals displayed no gliding motility and relied almost exclusively on peristaltic movements of their muscles, essentially squeezing themselves along the substrate. Our results show that defects on cilia structure and/or motility have detrimental effects on the ability of the organism to move in its environment and provide direct evidence that cilia are required for gliding-based locomotion in planarians. Finally, in both Smedift88(RNAi) and Smed-kif3a(RNAi) animals, the diminished cilia had no apparent effect on viability. However we did observe frequent fission events in motilitycompromised planarians, which we attributed to the increasing stress on the body due to the repeating crawling movements. Overall, our results suggest that S. mediterranea is powerful model for identifying genes that play a role in cilia formation and cilia-based motility.

IV. Summary Schmidtea mediterranea is a very attractive model organism for studying the biology of cilia and cilia-based motility. Advantages of using S. mediterranea include: (1) easy access to an abundance of ciliary structures; (2) the low cost to keep, maintain, and expand a sufficient number of animals in the laboratory; (3) a recently sequenced genome; and (4) established tools and methods for loss-ofgene-function studies via RNAi. In addition, we have developed and described in this chapter a number of powerful assays for evaluating cilia structure and motility.

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Acknowledgments We would like to thank Dr B. R. Gravely for providing us with a founding colony of Schmidtea mediterranea flatworms and Dr M. Smielewska for useful discussions. We would also like to thank Dr G. Pazour and J. Follit for their help with the SEM.

References Cebria, F., Nakazawa, M., Mineta, K., Ikeo, K., Gojobori, T., and Agata, K. (2002). Dissecting planarian central nervous system regeneration by the expression of neural-specific genes. Dev. Growth Differ. 44, 135–146. Cebria, F., and Newmark, P.A. (2005). Planarian homologs of netrin and netrin receptor are required for proper regeneration of the central nervous system and the maintenance of nervous system architecture. Development 132, 3691–3703. Dawar, B.L. (1973). A combined relaxing agent and fixative for triclads (planarians). Biotech. Histochem. 48, 93–94. Fire, A., Xu, S., Montgomery, M.K., Kostas, S.A., Driver, S.E., and Mello, C.C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811. Gonzalez-Estevez, C., Momose, T., Gehring, W.J., and Salo, E. (2003). Transgenic planarian lines obtained by electroporation using transposon-derived vectors and an eye-specific GFP marker. Proc. Natl. Acad. Sci. USA 100, 14046–140451. Marszalek, J.R., Ruiz-Lozano, P., Roberts, E., Chien, K.R., and Goldstein, L.S.B. (1999). Situs inversus and embryonic ciliary morphogenesis defects in mouse mutants lacking the KIF3A subunit of kinesin-II. Proc. Natl. Acad. Sci. USA 96, 5043–5048. Newmark, P.A., and Sanchez Alvarado, A. (2002). Not your father’s planarian: A classic model enters the era of functional genomics. Nat. Rev. Genet. 3, 210–219. Pazour, G.J., Dickert, B.L., Vucica, Y., Seeley, E.S., Rosenbaum, J.L., Witman, G.B., and Cole, D.G. (2000). Chlamydomonas IFT88 and its mouse homologue, polycystic kidney disease gene tg737, are required for assembly of cilia and flagella. J. Cell Biol. 151, 709–718. Pearson, B.J., Eisenhoffer, G.T., Gurley, K.A., Rink, J.C., Miller, D.E., and Sánchez Alvarado, A. (2009). Formaldehyde-based whole-mount in situ hybridization method for planarians. Dev. Dyn. 238, 443–450. Reddien, P.W., Bermange, A.L., Murfitt, K.J., Jennings, J.R., and Sanchez Alvarado, A. (2005). Identification of genes needed for regeneration, stem cell function, and tissue homeostasis by systematic gene perturbation in planaria. Dev. Cell 8, 635–649. Reddien, P.W., Newmark, P.A., and Alvarado, A.S. (2008). Gene nomenclature guidelines for the planarian Schmidtea mediterranea. Dev. Dyn. 237, 3099–3101. Reddien, P.W., and Sanchez Alvarado, A. (2004). Fundamentals of planarian regeneration. Annu. Rev. Cell Dev. Biol. 20, 725–757. Robb, S.M., Ross, E., and Sanchez Alvarado, A. (2008). SmedGD: The Schmidtea mediterranea genome database. Nucleic Acids Res. 36, D599–D606. Robb, S.M., and Sanchez Alvarado, A. (2002). Identification of immunological reagents for use in the study of freshwater planarians by means of whole-mount immunofluorescence and confocal microscopy. Genesis: J. Genet Dev. 32, 293–298. Saló, E. (2006). The power of regeneration and the stem-cell kingdom: Freshwater planarians (Platyhelminthes). Bioessays 28, 546–559. Sanchez Alvarado, A. (2003). The freshwater planarian Schmidtea mediterranea: Embryogenesis, stem cells and regeneration. Curr. Opin. Genet. Dev. 13, 438–444. Sanchez Alvarado, A., and Newmark, P.A. (1999). Double-stranded RNA specifically disrupts gene expression during planarian regeneration. Proc. Natl. Acad. Sci. USA 96, 5049–5054. Sanchez Alvarado, A., Newmark, P.A., Robb, S.M., and Juste, R. (2002). The Schmidtea mediterranea database as a molecular resource for studying platyhelminthes, stem cells and regeneration. Development 129, 5659–5665.

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CHAPTER 5

Targeted Gene Silencing by RNA Interference in Chlamydomonas Eun-Jeong Kim and Heriberto Cerutti School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska, Lincoln, Nebraska 68588

Abstract I. Introduction II. Materials and Methods A. Plasmid Construction B. Culture Conditions, Cell Transformation, and Selection Procedures C. RNA Analyses D. Reverse Transcriptase-PCR Analyses III. Results and Discussion IV. Summary Acknowledgments References

Abstract Small RNA-guided gene silencing is an evolutionarily conserved process that operates by a variety of molecular mechanisms and plays an essential role in developmental pathways and defense responses against genomic parasites in eukaryotes. Double-stranded RNA (dsRNA) triggered posttranscriptional gene silencing, termed RNA interference (RNAi), is also becoming a powerful tool for reverse genetics studies. Stable RNAi, induced by the expression of long dsRNAs or duplex small RNAs from genome-integrated transgenes, has been achieved in multiple organisms, including the green alga Chlamydomonas reinhardtii. However, the level of gene repression is often quite variable, depending on the type of construct, transgene copy number, site of integration, and target gene. Moreover, unintended transcripts partly complementary to a trigger dsRNA can also be silenced, making METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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difficult the interpretation of observed phenotypes. To obviate some of these problems we have developed a tandem inverted repeat system that consistently induces cosilencing of a gene with a selectable RNAi-induced phenotype (encoding tryptophan synthase b-subunit) and any other (nonessential) gene of interest. In addition, to circumvent off-target effects, for each tested gene, RNAi lines are generated with at least two transgenes, homologous to distinct and nonoverlapping sequences of the target transcript. A common phenotype among these independent RNAi strains is expected to result from suppression of expression of the gene of interest. We demonstrate this approach for the characterization of a gene of unknown function in Chlamydomonas, encoding a predicted exoribonuclease with weak similarity to 30 hExo/ERI-1.

I. Introduction RNA-mediated silencing is an evolutionarily conserved mechanism(s) by which double-stranded RNA (dsRNA) induces the inactivation of cognate sequences in eukaryotes, although dsRNA is also known to participate in the activation of gene expression (Carthew and Sontheimer, 2009; Cerutti and Casas-Mollano, 2006; Ghildiyal and Zamore, 2009; Grimm, 2009). The role of long dsRNA in triggering repression was initially characterized in Caenorhabditis elegans and termed RNA interference (RNAi) (Fire et al., 1998). However, silencing phenomena had already been described in multiple species and the connection to dsRNA helped to unify several, apparently disparate, processes including posttranscriptional RNA degradation, translational repression, transcriptional gene silencing via heterochromatin formation and/or DNA methylation, DNA elimination, and meiotic silencing by unpaired DNA (Baulcombe, 2004; Cerutti and Casas-Mollano, 2006; Matzke and Birchler, 2005). In plants, animals, and the unicellular green alga Chlamydomonas reinhardtii, the RNAi machinery is also involved in the production of microRNAs (miRNAs), small RNAs resulting from the processing of genome encoded imperfect RNA hairpins, which play a role in developmental regulation and other processes (Bartel, 2004; Carthew and Sontheimer, 2009; Casas-Mollano et al., 2008; Ghildiyal and Zamore, 2009; Molnar et al., 2007; Zhao et al., 2007). The most extensively characterized dsRNA-mediated mechanism is transcript silencing triggered by small interfering RNAs (siRNAs). In this process, long dsRNAs are processed into siRNAs (20–25 nt in length) by an RNaseIII-like endonuclease, named Dicer (Carthew and Sontheimer, 2009; Ghildiyal and Zamore, 2009). The siRNAs are then incorporated into a multiprotein complex, the RNA-induced silencing complex (RISC), where they associate with Argonaute (AGO) polypeptides (Carthew and Sontheimer, 2009; Ghildiyal and Zamore, 2009). Recent evidence suggests that a siRNA duplex is loaded into RISC and then AGO cleaves one of the siRNA strands (the passenger strand) triggering its dissociation from the complex (Carthew and Sontheimer, 2009; Ghildiyal and Zamore, 2009). Activated RISC then uses the

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remaining single-stranded siRNA as a guide to identify homologous RNAs, ultimately causing transcript degradation or translation repression (Carthew and Sontheimer, 2009; Ghildiyal and Zamore, 2009). In slightly over a decade, RNAi has evolved from a fascinating biological phenomenon into a powerful experimental tool for reverse genetics studies and for inducing phenotypic changes for practical applications (Grimm, 2009; Hebert et al., 2008). RNAi can be used to suppress the expression of genes of interest by introduction into cells or whole organisms of exogenously synthesized dsRNAs/siRNAs or nonintegrative dsRNA/siRNA-producing viral vectors or plasmids (Grimm, 2009; Hebert et al., 2008; Waterhouse and Helliwell, 2003). However, the silencing effects of these exogenous nucleic acids are generally transient. Stable and heritable RNAi has also been developed in a variety of eukaryotes, including Chlamydomonas (Grimm, 2009; Rohr et al., 2004; Schroda, 2006; Waterhouse and Helliwell, 2003). Common approaches rely on the production of antisense RNA or dsRNA by transcription from genome-integrated transgenes (Rohr et al., 2004; Schroda, 2006; Waterhouse and Helliwell, 2003). Yet, the level of silencing triggered by these transgenes is often variable and several (many) individual lines need to be molecularly characterized for suppression of a certain gene before potential phenotypic defects can be evaluated. More recently, transgenes mimicking the structure of miRNA precursors have been exploited to produce artificial miRNAs (amiRNAs) targeting genes of interest (Grimm, 2009; Molnar et al., 2009; Zeng et al., 2002; Zhao et al., 2009). In C. reinhardtii, the expression of amiRNAs appears to be more stable than that of long dsRNAs produced from inverted repeat (IR) transgenes but, in some cases, high levels of amiRNAs do not lead to target mRNA suppression (Molnar et al., 2009; Zhao et al., 2009). An additional issue with all RNAi approaches is that of target specificity since unintended transcripts, partly complementary to the trigger dsRNA, siRNA, or amiRNA, may also be silenced, a phenomenon termed “off-target” effects (Grimm, 2009; Kulkarni et al., 2006; Xu et al., 2006). To overcome some of these problems, we have developed a tandem inverted repeat (TIR)–RNAi system that allows for the direct selection of effective transgenic RNAi lines and the simultaneous silencing of two unrelated genes (Fig. 1A; Rohr et al., 2004). In Chlamydomonas, tryptophan synthase b-subunit (encoded by the MAA7 gene) converts the indole analog 5-fluoroindole (5-FI) into the toxic tryptophan analog 5-fluorotryptophan (Palombella and Dutcher, 1998). RNAi-mediated suppression of MAA7 leads to strains resistant to 5-FI (Rohr et al., 2004). Thus, selection for this RNAi-induced phenotype permits the identification of transgenic lines showing effective interference of any (nonessential) gene cotargeted with MAA7, as part of the same dsRNA trigger synthesized from a TIR (Fig. 1A; Rohr et al., 2004). In addition, for each gene of interest, RNAi lines are generated with at least two independent transgenes, homologous to distinct and nonoverlapping sequences of the target transcript. A common phenotype among these strains is expected to result from suppression of expression of the desired gene rather than off-target effects. We demonstrate here this approach for the in vivo characterization of a predicted exonuclease (EXO) in Chlamydomonas.

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Fig. 1 TIR–RNAi system for silencing of the EXO gene in Chlamydomonas. (A) Model of the TIR–RNAi system designed to target simultaneously the MAA7 and EXO transcripts. The subcellular localization of long dsRNA processing by Dicer in Chlamydomonas is presently hypothetical. Target mRNA cleavage is the depicted outcome of RNAi but the machinery can also operate by translation repression. (B) Diagram of the EXO precursor messenger RNA (excluding 50 and 30 untranslated regions), with exons indicated by white boxes. The annealing sites of primers used for RT-PCR amplification are shown underneath the exons. (C) Diagram depicting the domain architecture of the EXO protein. Exonuc_X-T, exonuclease, RNase T/DNA polymerase III domain; NLS, binary nuclear localization signal; zf-GRF, zinc finger motif named after three conserved residues in the center of the domain alignment.

II. Materials and Methods A. Plasmid Construction For generating an IR transgene targeting the EXO gene (protein ID 407657; http:// genomeportal.jgi-psf.org/Chlre4/Chlre4.home.html), a 580-bp fragment corresponding to the 30 end of its coding sequence was amplified by reverse transcription polymerase

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chain reaction (RT-PCR), using as template total RNA isolated from the CC-124 Chlamydomonas strain, with primers Exo-2 (50 -AAGCAGCCATGGTGGCAAG-30 ) and Exo-4 (50 -TACACACAAACACACGCTGACG-30 ) (Fig. 1B). This PCR product was digested with SmaI/MluI to generate a 435-bp fragment that was cloned by blunt-end ligation into the MluI site of a pSTBlue-1 vector (Novagen, Madison, WI, USA), containing a 200-bp DNA spacer previously inserted into the EcoRV site (Rohr et al., 2004). Another copy of the 435-bp SmaI/MluI fragment was then inserted by blunt-end ligation into the HincII site of the generated vector. This way, two identical EXO segments were ligated flanking the DNA spacer. The orientation of the cloned fragments was examined by digestion with PvuII, which cuts only once within the 435 bp EXO sequence, and a plasmid containing the inserts in inverted orientation was verified by DNA sequencing. Standard protocols were used for RT-PCR, restriction enzyme digestion, generation of blunt ends with T4 DNA polymerase, and DNA ligation (Sambrook and Russell, 2001). The EXO3 IR cassette was then excised by digestion with PstI/HindIII and inserted, by blunt-end ligation, into the EcoRI sites of the MAA7/X IR vector (Rohr et al., 2004) to generate the MAA7/EXO3 IR transgene. This construct contains a TIR with homology to the MAA7 and the EXO genes so that transcription leads to the production of a hairpin-loop-shaped RNA that can be processed by the RNAi machinery into siRNAs targeting both MAA7 and EXO transcripts (Fig. 1A). The MAA7/X IR vector contains, in addition to a MAA7 IR, an engineered aphVIII gene, encoding aminoglycoside 30 -phosphotransferase (Sizova et al., 2001), conferring resistance to paromomycin for selection purposes (Rohr et al., 2004). A similar strategy was used to construct the MAA7/EXO5 IR transgene, which is homologous to the middle region of the EXO coding sequence. Briefly, a 490-bp fragment was amplified by RT-PCR with primers Exo-10 (50 -CCTGTTCACACAGCGCTACAA30 ) and Exo-7 (50 -ATCGCTCCTGCTCCTGCTC-30 ) (Fig. 1B). This PCR product was then cloned, by blunt-end ligation, in forward and reverse orientations into the MluI and HincII sites of the pSTBlue-1 vector, flanking the 200-bp DNA spacer (Rohr et al., 2004). The EXO5 IR cassette was then excised by digestion with PstI/ HindIII and cloned, by blunt-end ligation, into the EcoRI sites of the MAA7/X IR vector (Rohr et al., 2004) to generate the MAA7/EXO5 IR transgene. The EXO sequences used to build the IR constructs were chosen because they are distinct and display very low homology (no contiguous stretch longer than 19 nt) to nontarget genes in BLAST searches of the Chlamydomonas genome (http://genomeportal.jgipsf.org/Chlre4/Chlre4.home.html), in order to minimize the possibility of off-target effects (Kulkarni et al., 2006). B. Culture Conditions, Cell Transformation, and Selection Procedures Chlamydomonas reinhardtii cells were grown in Tris–acetate–phosphate (TAP) medium (Harris, 1989). Plasmid DNA linearized with SacI, cleaving upstream from the transgenic promoter, was used in all transformation experiments. The wild type strain CC-124 (mt–) (Harris, 1989) was transformed by the glass beads procedure (Kindle, 1990)

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and allowed to recover for 2 days, to permit induction of RNAi, before plating under selective conditions. MAA7/EXO3 IR or MAA7/EXO5 IR transformants were selected on TAP medium containing 10 µg/ml paromomycin and 7.5 µM 5-FI (Sigma, St. Louis, MO, USA). Plates were incubated under dim lights (~50 µmol/m2/s photosynthetically active radiation) to lessen the light-induced degradation of 5-FI (Palombella and Dutcher, 1998). In our previous system, the selective medium was supplemented with 1.5 mM L-tryptophan (Rohr et al., 2004). However, addition of this amino acid was found to be unnecessary for survival or normal growth of the Chlamydomonas transformants. As a negative control, CC-124 was also transformed with the MAA7/X IR vector (Rohr et al., 2004), which only contains the IR transgene designed to induce RNAi of MAA7. All isolated transgenic strains were kept under constant selective pressure to circumvent any potential loss of activity due to silencing of the integrated IR transgenes. For phenotypic analyses, cells grown to logarithmic phase in liquid TAP medium were serially diluted, spotted on plates of the appropriate media (TAP or TAP containing 7.5 µM 5-FI), and incubated for 10–15 days under dim lights (Zhang et al., 2002). C. RNA Analyses Total cell RNA was purified with TRI Reagent (Molecular Research Center, Cincinnati, OH, USA), according to the manufacturer’s instructions. The isolated RNA was separated by agarose/formaldehyde gel electrophoresis, blotted onto nylon membranes, and hybridized with 32P-labeled probes specific for ACT1 (encoding actin) or the coding sequence of histone H2A (Sambrook and Russell, 2001; Wu-Scharf et al., 2000; Zhang et al., 2002). D. Reverse Transcriptase-PCR Analyses TRI Reagent isolated total RNA was treated with DNase I (RNase-free, Ambion, Austin, TX, USA) to remove contaminating DNA and reverse transcription reactions were carried out as previously described (Carninci et al., 1998). The synthesized cDNA was then used as a template in standard PCR reactions (Sambrook and Russell, 2001; WuScharf et al., 2000). The numbers of cycles showing a linear relationship between input RNA and the final product were determined in preliminary experiments. Primers were designed to match exonic sequences flanking one or more introns, to distinguish contaminating PCR products generated by the amplification of any remaining DNA. Controls also included the use as template of reactions without RT and verification of PCR products by hybridization with specific probes (data not shown). The PCR conditions for amplification of ACT1 were 23 cycles at 93°C for 30 s, at 55°C for 30 s, and at 71°C for 60 s. The EXO PCR product was amplified by 32 cycles under the same conditions. Aliquots (5-µl) of each RT-PCR reaction were resolved on 2% agarose gels and visualized by ethidium bromide staining. The primer sequences were as follows: Exo-5, 50 AACCTGAAGCGCCTGTTCAC-30 , EXO-11, 50 -GCTGCTCCCTGAAGCTGT CT-30 ; ACT-cod-F (50 -GACATCCGCAAGGACCTCTAC-30 ), and ACT-cod-R 0 (5 -GATCCACATTTGCTGGAAGGT-30 ).

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III. Results and Discussion The predicted Chlamydomonas EXO protein belongs to the DnaQ-H superfamily of 30 to 50 exonucleases, with a DEDDh motif that is characteristic of the 30 hExo/ERI-1 subfamily (Kupsco et al., 2006). The 30 hExo exoribonuclease was initially identified as a candidate regulator of histone mRNA degradation in mammalian cells (Dominski et al., 2003), but its in vivo role is not clear since the decay of histone transcripts involves similar pathways to those required for the degradation of poly(A) mRNAs after deadenylation (Marzluff et al., 2008). Intriguingly, the closest C. elegans and Schizosaccharomyces pombe homologs of 30 hExo, named ERI-1, have been implicated as negative regulators of RNAi and in 5.8S rRNA processing (Gabel and Ruvkun, 2008; Iida et al., 2006; Kennedy et al., 2004). Another member of this subfamily, Drosophila melanogaster Snipper, efficiently degrades structured, dsRNA, and DNA substrates as long as there exists a 30 overhang of a few nucleotides to initiate decay (Kupsco et al., 2006). However, the in vivo function of Snipper remains unknown since Snp mutant flies are viable and display no obvious phenotypic abnormalities (Kupsco et al., 2006). The human Snipper homolog, named exonuclease domain containing 1 (ExoD1), is produced in two isoforms (Gene ID 112479): a shorter one similar in structure to Snipper and a longer one that includes, in addition to the exonuclease domain, a zinc finger of the GRF (Glycine-Arginine-Phenylalanine) type. Thus, exonucleases belonging to the 30 hExo/ERI-1 subfamily appear to show activity toward a diverse array of substrates and play a variety of roles. Chlamydomonas EXO is most similar in sequence and domain structure (Fig. 1C) to the long human ExoD1 isoform and lacks a nucleic acid-binding SAP (after SAF-A/B, Acinus and PIAS) motif, typical of ERI-1 proteins. We used the TIR–RNAi system (Fig. 1A) as a useful tool to gain insight into the function of the uncharacterized EXO protein. In addition, we were interested in examining the possibility that EXO might act as a negative regulator of RNAi in Chlamydomonas and its suppression might, therefore, lead to strains with enhanced RNAi. We constructed two tandem IR transgenes targeting simultaneously the 30 untranslated region of the MAA7 mRNA as well as distinct, nonoverlapping regions of the EXO transcript: the MAA7/EXO3 IR transgene produces dsRNA homologous to the 30 end of the EXO coding sequence (Fig. 1B, region flanked by primers 2 and 4) and the MAA7/EXO5 IR transgene generates dsRNA matching the middle of the EXO coding sequence (Fig. 1B, region flanked by primers 10 and 7). Chlamydomonas cells resistant to 5-FI were isolated by transformation with either the MAA7/EXO3 IR transgene (Fig. 2A) or the MAA7/ EXO5 IR transgene (data not shown). Interestingly, transgenic lines transformed with these TIRs grew slower than the wild-type CC-124 on solid or in liquid TAP medium (Fig. 2A and data not shown). By contrast, Chlamydomonas strains containing an IR transgene targeting exclusively the MAA7 gene can also survive on medium containing 5-FI but are not affected in their growth rate (Fig. 2A, Maa7-IR5; Rohr et al., 2004). The transgenic lines transformed with the MAA7/EXO IR containing plasmids and resistant to 5-FI also showed reduced levels of the EXO transcript in semiquantitative RT-PCR assays (Fig. 2B), consistent with RNAi-mediated suppression of both the

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Fig. 2 Silencing of the EXO gene by tandem IR transgenes in transformants selected on 5-FI-containing medium. (A) Growth and survival of the indicated strains on TAP medium without (upper panel) or with (lower panel) 7.5 µM of 5-FI. CC-124, untransformed wild-type strain; Exo3-IR5 and Exo3-IR8, CC-124 transformed with a tandem IR transgene inducing simultaneous RNAi of the MAA7 (encoding tryptophan synthase b-subunit) and the EXO genes; Maa7-IR5, CC-124 transformed with an IR transgene designed to induce RNAi of MAA7. (B) Semiquantitative RT-PCR analysis of EXO gene expression in the indicated strains. Amplification of ACT1 (encoding actin) transcripts is shown as an input control. Reactions using RNA not treated with reverse transcriptase as the template were employed as a negative control (data not shown). Exo5-IR7 and Exo5-IR11, CC-124 transformed with a tandem IR transgene inducing simultaneous RNAi of the MAA7 and the EXO genes but targeting a different region of the EXO transcript than the MAA7/ EXO3 IR (see text for details). (C) Northern blot analysis of total cell RNA sequentially hybridized with 32 P-labeled PCR products corresponding to the coding sequence of histone H2A (upper panel), to evaluate the degree of mRNA reduction, or the coding sequence of ACT1 (lower panel), to assess the amount of sample loaded per lane.

MAA7 and EXO genes. This observation applied to strains generated with the MAA7/ EXO3 IR vector (Fig. 2B, Exo3-IR5 and Exo3-IR8) as well as those generated with the MAA7/EXO5 IR transgene (Fig. 2B, Exo5-IR7 and Exo5-IR11). However, the degree of EXO gene suppression varied among the different transgenic lines. In contrast, the Maa7-IR5 strain, containing the IR transgene designed to downregulate exclusively MAA7, did not display any decrease in EXO mRNA levels when compared with the wild type (Fig. 2B). Thus, the TIR–RNAi system, as previously demonstrated for several unrelated genes (Rohr et al., 2004), allowed the recovery of effective EXOsuppressed RNAi strains by selection for MAA7 silencing in medium containing 5-FI. The ERI-1 exonuclease has been implicated in the degradation of siRNA duplexes with two-nucleotide 30 overhangs, reducing the efficiency of RNAi (Kennedy et al., 2004), and in endogenous small RNA pathways in C. elegans (Lee et al., 2006). Thus, we tested whether Chlamydomonas EXO might act as an RNAi regulator in the transgenic lines. However, the EXO-suppressed RNAi strains did not show any change in the levels of several endogenous small RNAs when compared with the wild-type CC-124 (data not shown). The mammalian homolog of ERI-1, 30 hExo has been

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proposed to play a role, although presently undefined, in histone mRNA metabolism (Dominski et al., 2003; Yang et al., 2009). Interestingly, when we examined the Chlamydomonas lines containing the MAA7/EXO IR transgenes for histone H2A transcript amounts they displayed a significant reduction in comparison with the control strains (Fig. 2C). Most core histone genes in Chlamydomonas, similarly to those in metazoans, have a highly conserved palindromic sequence (that can fold into a stem–loop structure) at their 30 end, within a short distance of the stop codon (Fabry et al., 1995). Replicationdependent histone mRNAs lack a poly(A) tail in metazoans and their 30 end is formed by endonucleolytic cleavage very near the 30 stem–loop sequence (Marzluff et al., 2008). A single protein, the stem–loop-binding protein (SLBP), binds to this conserved hairpin structure and participates in many aspects of histone mRNA metabolism, including 30 end processing (Marzluff et al., 2008). Knockdown of SLBP results in reduced histone transcript levels, nuclear retention of histone mRNA, and defects in mammalian cell division (Sullivan et al., 2009). These phenotypes are very similar to those of the Chlamydomonas EXO RNAi strains, although we did not examine the possible nuclear retention of histone transcripts. Nonetheless, it is tempting to speculate that the EXO protein, like mammalian SLBP, may be required for the maturation of the 30 end of histone mRNAs in Chlamydomonas. In its absence, improperly processed histone transcripts may be unstable and subject to degradation by RNA quality control mechanisms (Doma and Parker, 2007). A number of approaches have been successfully used for RNA-mediated suppression of gene expression in Chlamydomonas, including genome-integrated transgenes generating long dsRNAs or amiRNAs (Molnar et al., 2009; Rohr et al., 2004; Schroda, 2006; Zhao et al., 2009). IR transgenes producing long dsRNAs have the disadvantage that they may be prone to self-silencing at the transcriptional level (Rohr et al., 2004; Yamasaki et al., 2008), although this may be partly avoided by expression from an inducible promoter (Koblenz and Lechtreck, 2005). In addition, long dsRNAs can be processed into a complex and heterogeneous population of siRNAs with the possibility that some of these small RNAs will match unintended transcripts and cause off-target effects (Grimm, 2009; Kulkarni et al., 2006; Xu et al., 2006). In contrast, amiRNA transgenes appear to be more stable in their expression and, since each amiRNA precursor gives rise to a single small RNA species, they can be optimized to avoid off-target effects (Molnar et al., 2009; Zhao et al., 2009). However, there are also disadvantages with the amiRNA approach since many aspects of small RNA processing, regulation, and miRNA action remain unknown in Chlamydomonas (CasasMollano et al., 2008). Thus, designed amiRNAs may not be processed as intended or the unique site where an amiRNA would associate on a target transcript may not be accessible, for instance, because of secondary structure formation (Grimm, 2009; Molnar et al., 2009). This may result in the accumulation of ineffective amiRNAs with certain transgenes and require careful design of more than one amiRNA construct for any given target gene (Molnar et al., 2009). The TIR–RNAi system obviates some of these issues. In addition, a recently reported PCR strategy can greatly simplify the construction of IR transgenes (Pawloski

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et al., 2005). Transcriptional self-silencing and other problems associated with genomic transgene integration (Rohr et al., 2004) are avoided by directly selecting for an RNAi-induced phenotype. Moreover, this also allows the recovery of effective RNAi strains against any (nonessential) gene cotargeted with the selectable MAA7 marker. Conceptually, a similar result can be achieved with dimeric amiRNA precursors (Molnar et al., 2009; Zhao et al., 2009). However, the production of long dsRNAs simplifies the design of RNAi transgenes because of the increased probability, relative to a single amiRNA, that at least some of the processed siRNAs will be assembled into a functional RISC and will have accessibility to the target transcript. The greater risk of off-target effects when using long dsRNA can be circumvented by silencing each gene of interest with at least two transgenes, homologous to distinct and nonoverlapping sequences of the transcript. The finding of similar phenotypes in several independent RNAi strains generated with different transgenes increases confidence that any phenotypic abnormalities are the consequence of downregulation of the intended gene. Moreover, there should be a clear correlation between target transcript and/or protein depletion and phenotypic severity. As demonstrated here for Chlamydomonas EXO, this approach can be used for the characterization of genes of unknown function.

IV. Summary RNAi is rapidly becoming a standard method for experimental and practical gene silencing. In the unicellular green alga Chlamydomonas reinhardtii, work from many groups has generated a number of tools for effective RNAi, including the TIR system described here. Both IR transgenes, producing long dsRNA, as well as amiRNA transgenes, generating unique small RNA molecules, have been successfully used for targeted gene silencing. However, potential drawbacks to this promising technology have also become apparent, such as a possible lack of specificity resulting in off-target effects. Moreover, relatively little is known about the role of the RNAi machinery in gene regulation and other processes in Chlamydomonas and the consequences that overexpression of transgenic RNAi triggers may have on these endogenous mechanisms. This emphasizes the need for careful design of RNAi experiments, including appropriate negative controls and the use of multiple amiRNA or long dsRNA species for each target gene. Whenever possible, data from RNAi-mediated knockdown should also be validated with alternative approaches. In addition, although throughout this chapter we have accentuated reduction in target transcript level as the expected outcome of RNAi, recent evidence suggests that RNAi may also operate by translational repression in Chlamydomonas requiring the assessment of protein amounts in transgenic RNAi lines. Ultimately, despite its limitations, RNAi provides a powerful method for uncovering gene function in organisms where gene targeting by homologous recombination is not practical.

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Acknowledgments This work was supported by a grant from the National Institutes of Health (GM62915) to H.C. We also acknowledge the support of the Nebraska EPSCoR program.

References Bartel, D.P. (2004). MicroRNAs: Genomics, biogenesis, mechanism, and function. Cell 116, 282–297. Baulcombe, D. (2004). RNA silencing in plants. Nature 431, 356–363. Carninci, P., Nishiyama, Y., Westover, A., Itoh, M., Nagaoka, S., Sasaki, N., Okazaki, Y., Muramatsu, M., and Hayashizaki, Y. (1998). Thermostabilization and thermoactivation of thermolabile enzymes by trehalose and its application for the synthesis of full length cDNA. Proc. Natl. Acad. Sci. USA 95, 520–524. Carthew, R.W., and Sontheimer, E.J. (2009). Silence from within: Endogenous siRNAs and miRNAs. Cell 136, 642–655. Casas-Mollano, J.A., Rohr, J., Kim, E.J., Balassa, E., van Dijk, K., and Cerutti, H. (2008). Diversification of the core RNA interference machinery in Chlamydomonas reinhardtii and the role of DCL1 in transposon silencing. Genetics 179, 69–81. Cerutti, H., and Casas-Mollano, J.A. (2006). On the origin and functions of RNA-mediated silencing: From protists to man. Curr. Genet. 50, 81–99. Doma, M.K., and Parker, R. (2007). RNA quality control in eukaryotes. Cell 131, 660–668. Dominski, Z., Yang, X.-C., Kaygun, H., Dadlez, M., and Marzluff, W.F. (2003). A 30 exonuclease that specifically interacts with the 30 end of histone mRNA. Mol. Cell 12, 295–305. Fabry, S., Muller, K., Lindauer, A., Park, P.B., Cornelius, T., and Schmitt, R. (1995). The organization, structure and regulatory elements of Chlamydomonas histone genes reveal features linking plant and animal genes. Curr. Genet. 28, 333–345. Fire, A., Xu, S., Montgomery, M.K., Kostas, S.A., Driver, S.E., and Mello, C.C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811. Gabel, H.W., and Ruvkun, G. (2008). The exonuclease ERI-1 has a conserved dual role in 5.8S rRNA processing and RNAi. Nat. Struct. Mol. Biol. 15, 531–533. Ghildiyal, M., and Zamore, P.D. (2009). Small silencing RNAs: An expanding universe. Nat. Rev. Genet. 10, 94–108. Grimm, D. (2009). Small silencing RNAs: State-of-the-art. Adv. Drug Deliv. Rev. 61, 672–703. Harris, E.H. (1989). “The Chlamydomonas Sourcebook—A Comprehensive Guide to Biology and Laboratory Use.” Academic Press, San Diego. Hebert, C.G., Valdes, J.J., and Bentley, W.F. (2008). Beyond silencing—engineering applications of RNA interference and antisense technology for altering cellular phenotype. Curr. Opin. Biotechnol. 19, 500–505. Iida, T., Kawaguchi, R., and Nakayama, J. (2006). Conserved ribonuclease, Eri1, negatively regulates heterochromatin assembly in fission yeast. Curr. Biol. 16, 1459–1464. Kennedy, S., Wang, D., and Ruvkun, G. (2004). A conserved siRNA-degrading RNase negatively regulates RNA interference in C. elegans. Nature 427, 645–649. Kindle, K.L. (1990). High frequency nuclear transformation of Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA 87, 1228–1232. Koblenz, B., and Lechtreck, K.F. (2005). The NIT1 promoter allows inducible and reversible silencing of centrin in Chlamydomonas reinhardtii. Eukaryot. Cell 4, 1959–1962. Kulkarni, M.M., Booker, M., Silver, S.J., Friedman, A., Hong, P., Perrimon, N., and Mathey-Prevot, B. (2006). Evidence of off-target effects associated with long dsRNAs in Drosophila melanogaster cell-based assays. Nat. Methods 3, 833–838. Kupsco, J.M., Wu, M.-J., Marzluff, W.F., Thapar, R., and Duronio, R.J. (2006). Genetic and biochemical characterization of Drosophila Snipper: A promiscuous member of the metazoan 30 hExo/ERI-1 family of 30 to 50 exonucleases. RNA 12, 2103–2117.

110

Eun-Jeong Kim and Heriberto Cerutti Lee, R.C., Hammell, C.M., and Ambros, V. (2006). Interacting endogenous and exogenous RNAi pathways in Caenorhabditis elegans. RNA 12, 589–597. Marzluff, W.F., Wagner, E.J., and Duronio, R.J. (2008). Metabolism and regulation of canonical histone mRNAs: Life without a poly(A) tail. Nat. Rev. Genet. 9, 843–854. Matzke, M.A., and Birchler, J.A. (2005). RNAi-mediated pathways in the nucleus. Nat. Rev. Genet. 6, 24–35. Molnar, A., Schwach, F., Studholme, D.J., Thuenemann, E.C., and Baulcombe, D.C. (2007). miRNAs control gene expression in the single-cell alga Chlamydomonas reinhardtii. Nature 447, 1126–1129. Molnar, A., Bassett, A., Thuenemann, E., Schwach, F., Karkare, S., Ossowski, S., Weigel, D., and Baulcombe, D. (2009). Highly specific gene silencing by artificial microRNAs in the unicellular alga Chlamydomonas reinhardtii. Plant J. 58, 165–174. Palombella, A.L., and Dutcher, S.K. (1998). Identification of the gene encoding the tryptophan synthase bsubunit from Chlamydomonas reinhardtii. Plant Physiol. 117, 455–464. Pawloski, L.C., Deal, R.B., McKinney, E.C., Burgos-Rivera, B., and Meagher, R.B. (2005). Inverted repeat PCR for the rapid assembly of constructs to induce RNA interference. Plant Cell Physiol. 46, 1872–1878. Rohr, J., Sarkar, N., Balenger, S., Jeong, B.-R., and Cerutti, H. (2004). Tandem inverted repeat system for selection of effective transgenic RNAi strains in Chlamydomonas. Plant J. 40, 611–621. Sambrook, J., and Russell, D.W. (2001). “Molecular Cloning—A Laboratory Manual.” Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Schroda, M. (2006). RNA silencing in Chlamydomonas: Mechanisms and tools. Curr. Genet. 49, 69–84. Sizova, I., Fuhrmann, M., and Hegemann, P.A. (2001). Streptomyces rimosus aph VIII gene coding for a new type phosphotransferase provides stable antibiotic resistance to Chlamydomonas reinhardtii. Gene 277, 221–229. Sullivan, K.D., Mullen, T.E., Marzluff, W.F., and Wagner, E.J. (2009). Knockdown of SLBP results in nuclear retention of histone mRNA. RNA 15, 459–472. Waterhouse, P.M., and Helliwell, C.A. (2003). Exploring plant genomes by RNA-induced gene silencing. Nat. Rev. Genet. 4, 29–38. Wu-Scharf, D., Jeong, B.-r., Zhang, C., and Cerutti, H. (2000). Transgene and transposon silencing in Chlamydomonas reinhardtii by a DEAH-box RNA helicase. Science 290, 1159–1162. Xu, P., Zhang, Y., Kang, L., Roossinck, M.J., and Mysore, K.S. (2006). Computational estimation and experimental verification of off-target silencing during posttranscriptional gene silencing in plants. Plant Physiol. 142, 429–440. Yamasaki, T., Miyasaka, H., and Ohama, T. (2008). Unstable RNAi effects through epigenetic silencing of an inverted repeat transgene in Chlamydomonas reinhardtii. Genetics 180, 1927–1944. Yang, X.C., Torres, M.P., Marzluff, W.F., and Dominski, Z. (2009). Three proteins of the U7-specific Sm ring function as the molecular ruler to determine the site of 30 -end processing in mammalian histone premRNA. Mol. Cell Biol. 29, 4045–4056. Zeng, T., Wagner, E.J., and Cullen, B.R. (2002). Both natural and designed micro RNAs can inhibit the expression of cognate mRNAs when expressed in human cells. Mol. Cell 9, 1327–1333. Zhang, C., Wu-Scharf, D., Jeong, B.-R., and Cerutti, H. (2002). A WD40-repeat containing protein, similar to a fungal co-repressor, is required for transcriptional gene silencing in Chlamydomonas. Plant J. 31, 25–36. Zhao, T., Li, G., Mi, S., Li, S., Hannon, G.J., Wang, X.J., and Qi, Y. (2007). A complex system of small RNAs in the unicellular green alga Chlamydomonas reinhardtii. Genes Dev. 21, 1190–1203. Zhao, T., Wang, W., Bai, X., and Qi, Y. (2009). Gene silencing by artificial microRNAs in Chlamydomonas. Plant J. 58, 157–164.

CHAPTER 6

Analysis of Cargo Transport by IFT and GFP Imaging of IFT in Chlamydomonas Dennis Diener Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, Connecticut 06520

Abstract I. Introduction II. Methods for GFP Imaging of IFT Proteins A. General Considerations in Tagging IFT Proteins B. Preparing Cells for Observing Movement of GFP-Tagged Proteins C. Illumination D. Optics and Image Capture III. IFT Cargo A. Analysis of IFT Cargo by Sucrose Density Gradients B. Materials C. Procedure Acknowledgments References

Abstract Chlamydomonas reinhardtii is the organism in which intraflagellar transport (IFT) was first visualized and in which the composition of IFT particles was originally elucidated. As the universality of IFT among ciliated/flagellated cells was uncovered, the diversity of organisms used to study IFT has grown. Still, because of the ease of isolation of flagella from Chlamydomonas and the battery of temperature-sensitive mutants affecting IFT proteins and motors, this unicellular alga remains the principal model for biochemical studies of IFT motors and cargo; furthermore, the long, exposed flagella of this cell are ideally suited for observing IFT in real time with GFP-tagged components of IFT. METHODS IN CELL BIOLOGY, VOL. 93 Copyright Ó 2009 Elsevier Inc. All rights reserved.

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I. Introduction IFT was first observed by video-enhanced DIC light microscopy in flagella of Chlamydomonas as the movement of varicosities along the length of the flagella [Kozminski et al. (1993); see Kozminski (1995) for methods]. It was in Caenorhabditis elegans, however, that IFT was first visualized using GFP-tagged proteins (Orozco et al., 1999). Before that time, expression of foreign genes, including genes encoding GFP-tagged protein, in Chlamydomonas had been problematic, but in that same year a GFP gene was synthesized using the Chlamydomonas codon bias and incorporating mutations known to accelerate processing, increase fluorescence, and produce a single excitation peak at 489 nm in the encoded GFP (Fuhrmann et al., 1999). This engineered GFP was successfully expressed in Chlamydomonas and has since been used to tag several flagellar proteins including a subunit of the anterograde IFT motor (Mueller et al., 2005), an IFT protein (Qin et al., 2007), and a potential membrane cargo protein (Huang et al., 2007). The steps required to image the movement of IFT particles in the flagella of Chlamydomonas are similar to those required for imaging of any GFP-tagged protein: genetically tagging and expressing the protein of interest in the cell, exciting the GFP, and imaging the fluorescent signal. Some of these steps will only be dealt with in a general way here, with more emphasis on some challenges specific to imaging GFP in Chlamydomonas flagella. Whereas the movement of IFT components tagged with GFP has been visualized, the movement of cargo has been more elusive. CrPKD2, the Chlamydomonas orthologue of polycystin-2, a component of ciliary and flagellar membranes that plays a role in the development of polycystic kidney disease in humans, has been tagged with GFP and seen to move in an IFT-dependent manner (Huang et al., 2007), but whether it is transported directly by IFT is not certain. Components of the axoneme—dynein arms, radial spokes, proteins of the central pair complex and tubulin itself—make up a major class of putative cargo carried by IFT; yet to date no one has directly visualized any of these cargos moving by IFT. Instead, less direct biochemical methods have been used to study transport of these proteins into and out of flagella. One axonemal component whose behavior within the flagellar matrix has been studied biochemically is the radial spoke. The intact radial spoke attached to the flagellar axoneme is a complex of 23 proteins that sediments at 20 S (Yang et al., 2001, 2006). In the cell body, however, the more prominent form of the spoke is a smaller complex composed of a subset of RSPs that sediment at 12 S (Qin et al., 2004). Both complexes are present in the soluble matrix fraction of the flagella: the 12 S complex on its way to the tip to assemble with other RSPs onto the axoneme; and the 20 S complex as a product of axonemal turnover, returning to the cell body. Analysis of flagellar matrix proteins in sucrose gradients for the presence of these two complexes can be used to differentiate anterograde from retrograde IFT cargo. Such analysis combined with various temperature-sensitive IFT mutants can be used to examine how deficiencies in the IFT machinery affect retrograde and anterograde movement of axonemal precursors. Methods will be described to analyze flagellar extracts on sucrose gradients to test for the presence of these complexes.

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II. Methods for GFP Imaging of IFT Proteins A. General Considerations in Tagging IFT Proteins In the best of all worlds one could track the movement of an intracellular protein in its native state. Preferably a protein tag would have no effect on the activity or distribution of the protein and the tagged protein would be expressed at the same level as the endogenous protein that it replaces; but even if these standards are not completely met, a lot can be learned through GFP tagging. Frequently tags are placed at either the N- or C-terminus of the protein by use of a suitable vector. Our laboratory has designed such a vector (Huang et al., 2009), which contains two tandem sequences encoding GFP separated by two flexible linkers. As diagrammed in Fig. 1, judicious placement of unique restriction sites allows the 50 or 30 GFP sequence to be replaced by the coding sequence of choice, resulting in the encoded protein being fused at its C- or N-terminus to GFP via a flexible linker. Using this vector, transcription is driven by the PsaD promoter, which can transcribe cDNAs as well as genomic constructs in Chlamydomonas. A potential drawback of this vector is that to ensure the fidelity of expression of the tagged gene, use of its native promoter may be preferable. To maintain the proper level of expression, ideally one would rescue a null mutant with a genomic clone encoding the tagged protein. In this way expression is driven by the native promoter and the tagged protein does not have to compete with the endogenous protein. This approach also has the advantage that rescue of the mutant phenotype ensures that the tagged protein is fully functional. Though ideal, null mutants are not necessary for satisfactory tagging of an IFT protein and observing its movement in flagella: the KAP subunit of the FLA10 heterotrimeric kinesin, which drives anterograde IFT, FLA3, has been successfully tagged with GFP in the temperature-sensitive mutant fla3-1 (Mueller et al., 2005); and the small GTPase IFT27 has been tagged with GFP and expressed in a wild-type background (Qin et al., 2007). In both cases the tagged protein was imaged moving in flagella at rates characteristic of IFT.

GASGQGASGADIGASGQGASGA

PPsaD

Ndel

Fig. 1

GFP

GFP EcoR V

PHSP70A-RSC2::aphVIII EcoR I

The vector designed for tagging Chlamydomonas genes with GFP, pHK85, has two copies of the GFP gene separated by two tandem linker sequences. The first or second GFP can be replaced by a cDNA or genomic sequence encoding a protein to be tagged with GFP at either its C- or N-terminus, respectively. Transcription is driven by the PsaD promoter and is terminated by the PsaD 30 genomic sequence. The plasmid also encodes aphVIII as a selectable marker in Chlamydomonas.

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B. Preparing Cells for Observing Movement of GFP-Tagged Proteins Motile Chlamydomonas often adhere to a coverslip by their flagella and remain quiescent, but they are unlikely to remain stationary in the light required for GFP imaging; therefore, in practice, paralyzed flagella are much more cooperative for making videos of GFP in flagella. Kozminski (1995) has detailed the advantages of using various paralyzed mutants and using mechanical or chemical means to paralyze flagella for observing IFT by video-enhanced DIC microscopy. For this purpose paralyzed flagella that remain flaccid are essential, so they can attach to a coverslip. Such flagella are found in motility mutants including pf1 (Kozminski et al., 1995) and pf15 (Iomini et al., 2001). The optical requirements for observing IFT by immunofluorescence are not as stringent as for DIC and stiff, paralyzed flagella, such as are found in pf18, are also suitable for observing the movement of intraflagellar GFP-tagged proteins if the cells are immobilized in 0.75–1% low melt agarose. One consideration when beginning a project of GFP tagging an IFT protein is, therefore, whether the recipient cells should be a paralyzed strain to facilitate the subsequent analysis of movement. When not complementing an IFT mutant with the tagged construct, using a paralyzed mutant as the recipient strain obviates the need for crossing the transformed strain. If transforming a flagellaless IFT mutant crossing the IFT mutant would be easier after transformation, when expression of the tagged protein has restored flagella to the transformant. Although paralyzed mutants make the imaging of IFT more convenient, imaging can also be done in motile strains using a variety of methods to inhibit flagellar motility. We induce flagellar paralysis using 20 mM LiCl or 20 mM sodium pyrophosphate, which do not affect the rate or frequency of IFT (Dentler, 2005). Unlike sodium pyrophosphate, which causes flagellar shortening (Lefebvre et al., 1978), the flagella actually elongate in LiCl (Nakamura et al., 1987). We can obtain good videos of IFT27::GFP in cells with motile flagella with the following protocol. Pipette 3 µl of cells (~2 106 cells/ml) onto an 18 18-mm coverslip and add 1.2 µl of 200 mM LiCl. Pipette 8 µl of 1.5% low melt agarose maintained at 37°C onto the cells and gently mix it into the cell solution with the pipette tip. Carefully invert the coverslip onto a slide and gently tap it down so the solution covers the entire surface of the cover slip. Cells stabilized in the agar with straight, immotile flagella parallel to the coverslip can be located and used to visualize IFT. If the flagella begin to curl or swell at the tip as may occur after about 30 min, make a new slide. C. Illumination While mercury arc lamps can be used for exciting GFP and observing IFT in the flagella of living Chlamydomonas, an argon 488-nm laser excites GFP more effectively. Observing a GFP signal in the flagella is complicated by the overwhelming autofluorescence generated from the cell body, which can severely restrict the portion of the flagellum that can be clearly seen. To circumvent this problem we use a Mosaic digital illumination system (Photonics Instruments, St. Charles, IL) to control the area of the specimen that is illuminated. With this system we can illuminate only the flagella, thereby limiting the signal from the cell body. Another method of reducing

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the excessive fluorescence from the cell body is the use of confocal microscopy to limit detection of emitted light to a thin focal plane, as was done for KAP::GFP (Mueller et al., 2005), or TIRF microscopy in which only those GFP molecules very near the coverslip surface are excited (see Chapter 9 by Engel and Marshall, this volume). For the latter technique the flagella must be attached to the coverslip.

D. Optics and Image Capture

Distance

For imaging we use a Nikon Eclipse TE2000 inverted microscope equipped with a 100, 1.4 NA Plan Apo objective lens. Samples are illuminated with a 300 mW 488 nm wavelength argon laser (Argon Ion Laser; National Laser Co., Salt Lake City, Ut) operating at 38 mW. The light is further attenuated by a prism that transmits only 70% of the incident light to the specimen. Using a tungsten lamp, preparations are scanned for a cell with stationary, preferably straight flagella parallel to the focal plane. Once one is found the Mosaic (or MetaMorph, (Molecular Devices, Sunnyvale, Ca)) software is used to draw the area to be illuminated over one or both flagella, avoiding getting too close to the cell body. Illumination is switched to the laser and images are collected with a Cascade 512B CCD camera (Photometrics, Tucson, AZ) with a back-illuminated 512 512 pixel array. The camera is controlled by MetaMorph software (version 7.1), and typically movement of flagellar IFT27::GFP can be imaged with the camera operating at 5 MHz with 4 gain, and an exposure time of 50 ms. For a full frame image this results in a frame rate of 118 ms. Faster frame rates can be obtained by decreasing the exposure time; operating the camera at 10 MHz, which also utilizes on-chip electron multiplier (EM) gain; and/or by decreasing the vertical size of the frame to a minimum. An example of a video of IFT27::GFP in the flagella of Chlamydomonas is shown in the supplementary material (http://www.elsevierdirect.com/companions/9780123749734, supplementary video) and one frame from the video is shown in Fig. 2.

(A)

(B) Time

Fig. 2 Visualization of IFT27::GFP in the flagella of Chlamydomonas. (A) IFT27::GFP is visible in IFT trains (arrows) in the two flagella of the cell. A video of the IFT movement of this cell is presented in the supplementary material. (B) Traces of IFT trains are visible as lines on this kymograph of the video shown in the supplement. The downward pointing dotted line shows retrograde IFT and the upward pointed dotted line shows anterograde IFT.

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To analyze movement, kymographs are generated using the kymograph function of MetaMorph by tracing a line over the flagellum wide enough to cover all of the fluorescent IFT trains and creating the kymograph based on maximum intensity (Fig. 2). Movement rates are computed from the slope of individual traces once the measurements of height and width are converted from number of pixels to distance in microns and time in seconds.

III. IFT Cargo IFT cargo might be defined as molecules that are not part of the IFT machinery itself that are moved into, throughout, or out of the flagella by IFT. Evidence that axonemal proteins are cargo of IFT has been obtained through dikaryon rescue experiments (Piperno et al., 1996), sedimentation gradients (Piperno and Mead, 1997; Qin et al., 2004), immunoprecipitation (Qin et al., 2004), and genetic methods (Hou et al., 2007). Yet, to date, the membrane protein CrPKD2 is the only potential cargo whose movement in the flagella of Chlamydomonas actually has been measured to be the rate of IFT. A. Analysis of IFT Cargo by Sucrose Density Gradients Although we have not visualized radial spokes moving by IFT in flagella we can analyze factors that influence the accumulation of these putative cargos in the soluble fraction of the flagella. For example, the ratio of 12 S radial spoke complex entering flagella to 20 S complex exiting the flagella, increases during flagellar regeneration (Qin et al., 2004). This ratio is reversed (with relatively more 20 S exiting the flagella) when the flagella is resorbing due to inactivation of the anterograde motor in fla10 cells (Qin et al., 2004), which harbor a temperature sensitive mutation in the anterograde motor, FLA10. B. Materials HMDEK: 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (Hepes), pH 7.4, 5 mM MgSO4, 0.5 mM ethylene glycol bis(b-aminoethyl ether) N,N0 -tetraacetic acid EGTA, 1 mM dithiothreitol, 25 mM KCl. We include the following protease inhibitors in HMDEK: 2 µg/ml aprotinin, 20 µg/ml benzamidine; 1 µg/ml leupeptin, 1 µg/ml pepstatin, and 50 µg/ml trypsin inhibitor. HMDEK þ 10 or 30% sucrose. C. Procedure 1. Harvest flagella from 32 to 40 l of Chlamydomonas according to published protocols (King, 1995; Witman, 1986; Witman et al., 1972). Regardless of which protocol is used, do not add sucrose to the cells before deflagellation (add it just after deflagellation instead) because 4% sucrose can affect the ratio of 12 S and 20 S

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2.

3. 4. 5.

6. 7.

8.

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spoke complexes and increases the amount of IFT proteins in the flagella. We generally use the pH shock method of deflagellation and obtain 0.25–0.50 ml of packed flagella when they are ultimately collected in a microfuge tube. All steps following deflagellation are done on ice or at 4°C. Resuspend the flagellar pellet in an equal volume of HMDEK, add NP-40 to 0.5%, and incubate on ice for 10 min. Alternatively, the flagella can be frozen and thawed, releasing the matrix proteins but leaving much of the membrane in the axonemal pellet. Sediment the insoluble material in a microfuge for 10 min. Further clarify the supernatant at 100,000 g for 10 min. Load the clarified supernatant (less than 0.5 ml) onto a 12-ml 10–30% sucrose gradient in HMDEK. We pour gradients using a piston gradient former (Jule Inc., Milford, CT, USA) though satisfactory results can be obtained with an open air former. Centrifuge the gradients for 16 h at 178,000g (38,000 rpm) in an SW41Ti rotor (Beckman Coulter). Gradients are typically fractionated into 24–26 0.5-ml aliquots by pumping them from the bottom. A 25-µl capillary tube (Sigma) attached to 0.76-mm inner diameter pump tubing (Rainin) is inserted into the centrifuge tube. The tubing passes through a peristaltic pump and leads to a fraction collector. There may be a pellet in the bottom of the tube and if the capillary tube reaches the bottom of the tube fragments of the pellet may contaminate fractions. To avoid this problem put a piece of tape on the capillary tube to keep it suspended above the pellet. Separate 15 µl aliquots of each fraction in an SDS-PAGE gel, transfer to blotting paper, and probe for a radial spoke protein.

The gradient profile should have two peaks of the radial spoke complexes at 12 S and 20 S. There may be some smaller complex or monomer as well. Standards should be used to calculate S-values: BSA (4.4 S), aldolase (7.35 S), catalase (11.3 S), and thyroglobulin (19.4 S) on parallel, or in the same, gradient. The 12-ml gradients fractionated into 24 fractions offers good resolution, but if this is not required, larger fractions can be collected or smaller gradients can be run. For example, 2-ml gradients can be run in 4 h to analyze 0.1 ml of soluble flagellar protein (Pan and Snell, 2003). Biochemistry cannot directly show movement of molecules within the flagella so GFP imaging of axonemal precursors moving in the flagella at rates comparable to IFT would be a welcome advance. We have expressed RSP3::GFP in pf14 cells, but initial efforts to see its movement in the flagellum were hampered by the signal from the abundant copies of RSP3::GFP already attached to the axoneme. This problem may be solved by watching tagged spokes enter spokeless flagella following mating of the tagged strain with spokeless pf14 cells. Alternatively, photobleaching the tagged protein present on the axoneme or photoactivating a tagged protein inside the cell may allow precursors in transit to the flagellar tip to be seen.

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Acknowledgments I thank Dr. Kaiyao Huang for help with the microscopic techniques described here and for suggestions on the manuscript and Dr. Joel Rosenbaum for comments on the text and support of this work (National Institutes of Health grant GM014642).

References Dentler, W. (2005). Intraflagellar transport (IFT) during assembly and disassembly of Chlamydomonas flagella. J. Cell Biol. 170, 649–659. Fuhrmann, M., Oertel, W., and Hegemann, P. (1999). A synthetic gene coding for the green fluorescent protein (GFP) is a versatile reporter in Chlamydomonas reinhardtii. Plant J. 19, 353–361. Hou, Y., Qin, H., Follit, J., Pazour, G.J., Rosenbaum, J.L., and Witman, G.B. (2007). Functional analysis of an individual IFT protein: IFT46 is required for transport of outer dynein arms into flagella. J. Cell Biol. 176, 653–665. Huang, K., Diener, D., Mitchell, A., Pazour, G.J., Witman, G.B., and Rosenbaum, J.L. (2007). Function and dynamics of PKD2 in Chlamydomonas reinhardtii flagella. J. Cell Biol. 179, 501–514. Huang, K., Diener, D.R., and Rosenbaum, J.L. (2009). The ubiquitin conjugation system is involved in the disassembly of cilia and flagella. J. Cell Biol. 186, 601–613. Iomini, C., Bebaev-Khaimov, V., Sassaroli, M., and Piperno, G. (2001). Protein particles in Chlamydomonas flagella undergo a transport cycle consisting of four phases. J. Cell Biol. 153, 480–491. King, S.M. (1995). Large-scale isolation of Chlamydomonas flagella. In: “Methods in Cell Biology” (W.L. Dentler and G.B. Witman, eds.), Vol. 47, pp. 9–12. Academic Press, San Diego. Kozminski, K.G. (1995). High-resolution imaging of flagella. In “Methods in Cell Biology”. (W.L. Dentler and G.B. Witman, eds.), Vol. 47, pp. 263–271. Academic Press, San Diego. Kozminski, K.G., Beech, P.L. and Rosenbaum, J.L. (1995). The Chlamydomonas kinesin-like protein FLA10 is involved in motility associated with the flagellar membrane. J. Cell Biol. 131, 1517–1527. Kozminski, K.G., Johnson, K.A., Forscher, P., and Rosenbaum, J.L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA. 90, 5519–5523. Lefebvre, P.A., Nordstrom, S.A., Moulder, J.E., and Rosenbaum, J.L. (1978). Flagellar elongation and shortening in Chlamydomonas IV. Effects of flagellar detachment, regeneration, and resorption on the induction of flagellar protein synthesis. J. Cell Biol. 78, 8–27. Mueller, J., Perrone, C.A., Bower, R., Cole, D.G., and Porter, M.E. (2005). The FLA3 KAP subunit is required for localization of kinesin-2 to the site of flagellar assembly and processive anterograde intraflagellar transport. Mol. Biol. Cell 16, 1341–1354. Nakamura, S., Takino, H., and Kojima, M. (1987). Effect of lithium on flagellar length in Chlamydomonas reinhardtii. Cell Struct. Funct. 12, 369–374. Orozco, J.T., Wedaman, K.P., Signor, D. Brown, H., Rose, L., and Scholey, J.M. (1999). Movement of motor and cargo along cilia. Nature 398, 674. Pan, J., and Snell, W. (2003). Kinesin II and regulated intraflagellar transport of Chlamydomonas aurora protein kinase. J. Cell Sci. 116, 2179–2186. Piperno, G., and Mead, K. (1997). Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella. Proc. Natl. Acad. Sci. USA 94, 4457–4462. Piperno, G., Mead, K., and Henderson, S. (1996). Inner dynein arms but not outer dynein arms require the activity of kinesin homologue protein KHP1FLA10 to reach the distal part of flagella in Chlamydomonas. J. Cell Biol. 133, 371–379. Qin, H., Diener, D.R., Geimer, S., Cole, D.G., and Rosenbaum, J.L. (2004). Intraflagellar transport (IFT) cargo: IFT transports flagellar precursors to the tip and turnover products to the cell body. J. Cell Biol. 164, 255–266. Qin, H., Wang, Z., Diener, D., and Rosenbaum, J. (2007). Intraflagellar transport protein 27 is a small G protein involved in cell-cycle control. Curr. Biol. 17, 193–202.

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Witman, G.B. (1986). Isolation of Chlamydomonas flagella and flagellar axonemes. Methods Enzymol. 134, 280–290. Witman, G.B., Carlson, K., Berliner, J., and Rosenbaum, J.L. (1972). Chlamydomonas flagella I. Isolation and electrophoretic analysis of microtubules, matrix, membranes, and mastigonemes. J. Cell Biol. 54, 507–539. Yang, P., Diener, D.R., Rosenbaum, J.L., and Sale, W.S. (2001). Localization of calmodulin and dynein light chain LC8 in flagellar radial spokes. J. Cell Biol. 153, 1315–1326. Yang, P., Diener, D.R., Yang, C., Kohno, T., Pazour, G.J., Dienes, J.M., Agrin, N.S., King, S.M., Sale, W.S., Kamiya, R., Rosenbaum, J.L., and Witman, G.B. (2006). Radial spoke proteins of Chlamydomonas flagella. J. Cell Sci. 119, 1165–1174.

CHAPTER 7

Genetic and Phenotypic Analysis of Flagellar Assembly Mutants in Chlamydomonas reinhardtii Carlo Iomini*, Jacob E. Till†, and Susan K. Dutcher† * Department of Developmental and Regenerative Biology, Mt. Sinai School of Medicine, New York, New York 10023 †

Department of Genetics, Washington University School of Medicine, St. Louis, Missouri 63110

Abstract I. Introduction II. Isolation of Aflagellate Strains III. Isolation of Conditional Flagellar Assembly (fla) Mutants A. Chemical Mutagenesis B. Solutions C. Day 1. Mutagenesis D. Day 2. Eliminate Swimming Cells at 32°C (First Cycle of Deflagellation) E. Day 3. Eliminate Nonconditional Nonswimming Cells F. Day 4. Culture Single Cells on Solid Medium G. Day 10. Screen of Single-Cell Colonies IV. Characterizing Intraflagellar Transport in fla Mutants A. VE-DIC Light Microscopy B. Digital Kymography V. Genetic Characterization A. Determining the Number of Genes Among a Collection of Mutants B. Linkage Analysis C. Chlamydomonas Transformation and SHIRT D. Reversion of Flagellar Mutants Using Ultraviolet Light E. Diploid Strain Construction and Analysis VI. Characterization of Genetic Interactions A. Synthetic Phenotypes B. Second-Site Noncomplementation C. Two-Copy Suppression References METHODS IN CELL BIOLOGY, VOL. 93 Copyright Ó 2009 Elsevier Inc. All rights reserved.

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Abstract Conditional mutants for flagellar assembly (fla) provide a useful tool to study intraflagellar transport (IFT) at the molecular level, and provide a unique set of tools to analyze cilia. The analysis of IFT phenotypes of fla mutants at the permissive temperature by a quantitative image analysis approach identified four distinct phases of the IFT cycle and directly demonstrated structural and functional remodeling of IFT particles at both axonemal extremities. In addition, the genetic analysis of fla mutants reveal interesting interactions among genes involved in flagellar assembly that help to provide information about the structure and function of IFT particles and their motors. This chapter provides protocols to isolate, characterize, and identify conditional Chlamydomonas flagellar assembly mutants and their genes and to test genetic interactions among proteins encoded by these genes.

I. Introduction Intraflagellar transport (IFT), the evolutionary conserved intracellular transport required for ciliary/flagellar assembly, maintenance, and signaling was first observed as a bidirectional movement of particles along Chlamydomonas flagella by using video-enhanced differential interference contrast (VE-DIC) light microscopy (Kozminski et al., 1993). Mutants in Chlamydomonas with conditional, temperaturesensitive defects in flagellar assembly (fla) retain flagella at the permissive temperature (21°C) but lose them at the restrictive temperature (32°C). Biochemical and microscopic analysis of fla10, which has a conditional mutation in the gene encoding one of the motor subunits of kinesin-2 (Walther et al., 1994), lead to the identification of IFT–protein complexes and uncovered the role of IFT in flagella-mediated signaling (Cole et al., 1998; Pan and Snell, 2002; Piperno and Mead, 1997; Piperno et al., 1996; Wang et al., 2006). Biochemical analysis of the Chlamydomonas flagella led to the identification of many of the proteins in the IFT particles and showed that they dissociate into two complexes called A and B (Cole et al., 1998; Piperno and Mead, 1997). Null alleles in Chlamydomonas IFT genes, which include bld1-1, bld1-2, and ift88-1, were instrumental in showing the essential role of IFT components in flagellar assembly (Brazelton et al., 2001; Pazour et al., 2000). Strains with these null alleles lack flagella and prevent a biochemical and microscopic approach to study the function of proteins encoded by their disrupted genes. Temperature-sensitive mutants for flagellar assembly (fla) provide a useful tool to study IFT at the molecular level, and give Chlamydomonas a unique set of tools to analyze cilia (Adams et al., 1982; Huang et al., 1977; Iomini et al., 2001; Lux and Dutcher, 1991; Piperno et al., 1998). The analysis of IFT phenotypes of fla mutants at the permissive temperature by a quantitative image analysis approach called kymography identified four distinct phases of the IFT cycle and directly demonstrated structural and functional remodeling of IFT particles at both axonemal extremities.

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Finally, fla mutants reveal unique genetic interactions among genes involved in flagellar assembly that help to provide information about the structure and function of IFT particles and their motors. This chapter provides protocols to isolate, characterize, and identify conditional Chlamydomonas mutants for flagellar assembly and to test genetic interactions among proteins encoded by these genes. This approach has helped to advance our understanding of ciliary biology by uncovering specific functions of individual components of the IFT machinery and providing new tools to identify new genes involved in IFT regulation.

II. Isolation of Aflagellate Strains Many, but not all aflagellate strains, have defects in IFT components and motors. Some of these mutants come from screens for cells that fail to mate, as flagellar proteins are required for recognition between cells of opposite mating types (Goodenough et al., 2007). For example, BLD1 that encodes IFT52 was identified in this screen (Brazelton et al., 2001). An alternative approach is based on the observation that exogenous DNA integrates into the Chlamydomonas genome nonhomologously (Tam and Lefebvre, 1993). Thus, it has been used extensively for insertional mutagenesis. Insertional mutants have yielded aflagellate or very short, stumpy flagellar strains. Screening Southern blots using probes to IFT proteins or motors has identified many of these proteins, which include ift46, dhc1b-1, stf1-1, stf1-2, and d1blic (Hou et al., 2004, 2007; Pazour et al., 1999; Porter et al., 1999), which assemble stumpy or short flagella. The selectable markers for insertional mutagenesis currently include ARG7, NIT1, BLE, and APHVIII, which confers arginine prototrophy, nitrate prototrophy, resistance to Zeocin, and resistance to paromomycin, respectively. The APHVIII gene has several advantages. It is a relatively small gene. Paromomycin is highly selective and is not mutagenic like Zeocin, which induces chromosome breaks if the transgene is not active immediately. Null alleles are useful in combination with conditional alleles for deciphering genetic interactions (Section VI). Thermal asymmetric interlaced (TAIL) and adaptor PCR are two methods to identify the sites of insertion without the need to use a candidate gene for Southern blots. 1. Prepare genomic DNA (1 µg) from insertional mutant strains and wild-type parental strains by previously described techniques (Johnson and Dutcher, 1991) and digest with PmlI (50 U) and PvuII (10 U) overnight (16–18 h) to obtain blunt-ended fragments 1–6 kb in length. 2. Verify the digestion by agarose gel (10 µl in a 2% gel) electrophoresis. The fragments are purified by column PCR purification and eluted with 30 µl H2O. 3. The adaptor (Table I) is prepared by annealing the plus and minus adaptor (25 µM each, in a thermocycler, set to decrease temperature from 95 to 4°C over 3 h (1°C every 2 min), and stored at 4°C.

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Table I Primers for Identifying the Insertion Site Primer name TPA1 TPA2 TPA3 TPA1R TPA2R TPA3R TPA4 TPA5 TPA6 TPA7 AD3 AD5 AD6 AD11 Minus adaptor Plus adaptor RIM3-1 RIM3-2 AP1 AP2

Primer sequence CGGGAGTTGTTTGTCAAGGT GTTTGTCAAGGTGGCAGCTC GATTCCCGTACCTCGTGTTGT ACCTTGACAAACAACTCCCG GAGCTGCCACCTTGACAAAC ACAACACGAGGTACGGGAATC CTCAGAAGAACTCGTCCAACAG CATCAGGTCCCTCAGAAGAACT GTAAAACGCCAGCTTTTCCTC GAACCACGGGTCCTCCTC WGTGNAGNANCANAGA STTGNTASTNCTNTGC WCAGNTGWTNGTNCTG NCASGAWAGNCSWCAA 50 phosphorylation – 50 – ACCAGCCCGG – –30 C7 spacer arm – 30 amino modifier 50 GTAATACGACTCACTATAGAGTACGCGTGGTCGACGGCCCGGGCTGGT CGGTATCGGAGGAAAAGCTG GCTGTTGGACGAGTTCTTCTG GTAATACGACTCACTATAGAGT ACTATAGAGTACGCGTGGT

4. The blunt-end genomic DNA fragments (3 µl) are ligated to the adaptor in a 20 µl reaction containing the adaptor (1.25 µM) and T4 DNA Ligase (400 U) in T4 DNA Ligase buffer. Incubate the reaction at 16°C for 16 h and inactivate the enzyme by incubation at 80°C for 20 min. Dilute the reaction to a total of 90 µl (with H2O) before being used in PCR reactions. 5. Two nested PCR reactions are conducted using primers RIM3-1 and TPA3R, then, RIM3-2 and TPA1R for the iPCR (Table I). Four different nested primer sets are used for the adaptor-mediated PCR: RIM3-1, AP1, then, RIM3-2, AP2; TPA1, AP1, then, TPA3, TPA2; TPA3R, AP1, then, TPA1R, AP2; and TPA4, AP1, then, TPA6, AP2. PCR reactions are carried out in a 50-µl volume, using 2 µl of the above DNA preparation mixture and consisting of 0.4 mM dNTPs, 5% (v/v) dimethyl sulfoxide (DMSO), 0.4 µM primers, 0.24 µl KLENTaq-LA, in supplied buffer was used. The reactions use a touchdown PCR protocol. 6. Products are sequenced to determine the location of the insertion site. Approximately 60% of the mutants tested gave insertions that can be verified by mapping.

III. Isolation of Conditional Flagellar Assembly (fla) Mutants The conditional class of fla mutants are isolated on the basis that they are motile at the permissive temperature of 21°C but lose their flagella at the restrictive temperature of 32°C. As is observed for many conditional mutants, fla mutants also display weaker,

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but specific mutant phenotypes at the permissive temperature that have been informative about the function of their gene products. The total number of independent fla mutants in Chlamydomonas is currently 22 in 17 different loci (Table II). Some of these affect IFT and its regulation, and others may affect templating or docking functions of the basal bodies. Given the complexity of the IFT machinery and its regulation, it is not surprising that the number of loci identified has not reached saturation. Only three of the loci (FLA8, FLA10, and FLA17) have more than one allele.

A. Chemical Mutagenesis The screen for fla mutants is based on the observation that aflagellated Chlamydomonas cells are unable to oppose gravity. When cells lacking flagella are grown in liquid culture they form a pellet. By contrast, wild-type cells swim and are distributed evenly throughout the culture medium. Because the nonswimmer phenotype could also derive from cells with paralyzed flagella, a secondary screen with light microscopy is needed to identify strains with paralyzed flagella. The fla mutants currently available were generated in three different screens (Adams et al., 1982; Huang et al., 1977; Iomini et al., 2001; Piperno et al., 1998).

B. Solutions

1. Stock Solution of N-methyl-N 0 -nitro-N Nitrosoguanidine Dissolve 2 mg/ml N-methyl-N 0 -nitro-N Nitrosoguanidine (MNNG) in 0.02 M citrate buffer at pH 5 prepared in fume hood. R medium for cell culturing is based Sager and Granick’s method as described previously (Harris, 1989). M-N/5 medium used in mutagenesis is described below. M-N/5 stock solutions 1. 10% sodium citrate 2. Trace elements For 1 l of stock: H3BO3 ZnSO47H2O MnSO4H2O CoCl26H2O CuSO45H2O NH4 molybdate4H2O 3. 1% FeCl36H2O 4. 4% CaCl22H2O 5. 10% MgSO47H2O 6. 10% K2HPO4 pH will be 7.8–8

For 500 ml 0.5 ml 1 ml 100 mg 100 mg 40 mg 20 mg 4 mg 15 mg 0.1 ml 0.1 ml 0.3 ml 2.6 ml

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Table II Phenotypes and Genes for the FLA Loci Allele

Gene

fla2-1 fla3-1 fla4 fla5 fla6 fla7 (fla10-1) Fla8-1 fla8-2 (fla1) fla9 519 fla10-14 544 fla10-15 fla11 fla12 fla13 fla15 fla16

FLA2 FLA3 FLA4 FLA5 LOST FLA10 FLA8 FLA8 FLA9 FLA10 FLA10 FLA11 FLA12 FLA13 FLA15 FLA17

Kinesin-2 motor subunit Kinesin-2 motor subunit Kinesin-2 motor subunit Unknown Kinesin-2 motor subunit Kinesin-2 motor subunit IFT172 Unknown Unknown IFT144 IFT139

fla17-1

FLA17

IFT139

fla17-2

FLA17

IFT139

fla18 fla21 fla24 fla27 fla28

FLA18 FLA21 FLA24 FLA27 FLA28

Unknown Unknown Unknown Unknown Unknown

c

Unknown Kinesin-associated protein Unknown Unknown

Mutation

F753L

N329K E21K F55S E24K Unknown L1615P

C1283R Deletion of exons 17–19 Deletion of exons 17–19 Deletion of exons 17–19

Linkageb group

Screenc

VI X X XI

H H H A

XIX XII/XIII XII/XIII XIX XIX XIX XIX XIX ND XIV VI

H A H A A A A A A P P

Miller et al. (2005) Lux and Dutcher (1991) Pedersen et al. (2005)

Reduced retrograde IFT

VI

P

Iomini et al. (2009)

Reduced retrograde IFT

VI

P

Iomini et al. (2009)

Reduced anterograde IFT No IFT defect Reduced retrograde IFT Reduced anterograde IFT Reduced anterograde IFT

ND ND VI ND ND

P P P P P

Permissive temperature phenotypea Reduced retrograde IFT Not tested No IFT defect No IFT defect Reduced anterograde IFT Reduced anterograde IFT Reduced anterograde IFT No IFT defect Not tested Not tested Reduced retrograde IFT No IFT defect Not tested Reduced retrograde IFT Reduced retrograde IFT

Permissive temperature phenotypes are from Iomini et al. (2001). b ND Map location is not determined. Indicates the screen that found the mutants: H, Huang et al. (1977); A, Adams et al. (1982); P, Piperno et al. (1998).

References

Mueller et al. (2005)

Walther et al. (1994) Miller et al. (2005) Miller et al. (2005)

Iomini et al. (2009) Iomini et al. (2009)

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a

Protein

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Solutions for deflagellation 0.5 N acetic acid 1 M KHCO3

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For 500 ml: 14.3 ml of glacial acetic in H2O For 500 ml: 50.06 g KHCO3 in H2O

C. Day 1. Mutagenesis 1. Grow one loop of Chlamydomonas wild-type cells 137þ (Strain CC-125) taken from a freshly streaked culture in two flasks containing 100 ml of R medium each for 3 days at 25°C under bright fluorescent light. 2. Collect cells by centrifugation at 350 g for 15 min at 21°C. 3. Resuspend each pellet in 20 ml of 0.02 M citrate buffer at pH 5, count cell number with a hemocytometer, and repeat centrifugation. Resuspend in 50 ml 0.02 M citrate buffer at pH 5 at 106 cells/ml in three 50-ml polypropylene conical tubes. Cultures need to be maintained with sterile conditions. 4. Add MNNG to final concentration of 1, 5, or 10 µg/ml, respectively, to tubes and incubate in the dark for 30 min at 25°C. 5. Collect all cells by centrifugation at 350 g for 15 min at 21°C and wash two times in 10 ml M-N/5 medium, resuspend cells to 106 cells/ml. Medium lacking nitrogen (N) induces gametogenesis and stops cell division. This prevents the isolation of identical, nonindependent alleles from the screening. To determine cell viability after exposure to MNNG, plate 100 µl of cells at the concentration of 5 103 cells/ml based on cell counts on an R plate and let plate grow for at least 3 days in bright light. Viability is 70, 60, and 45% for cell cultures treated with 1, 5, or 10 µg/ml of MNNG, respectively. 6. Incubate. cells exposed to the mutagenic agent overnight in bright light at 21°C. D. Day 2. Eliminate Swimming Cells at 32°C (First Cycle of Deflagellation) 1. Deflagellate cells by adding 0.5 N acetic acid to reduce the pH of the medium to 4.5. Quickly check a sample by phase microscopy to ensure that cells have lost their flagella than neutralize with 1 M KHCO3 solution to return the medium to a pH of 7. Do not leave cells for more than 1–2 min at pH 4.5. 2. Collect all cells by centrifugation at 350 g for 15 min at 21°C and discard the supernatant. 3. Resuspend cells in 30 ml of fresh M-N/5 medium prewarmed to 32°C in 50-ml polypropylene conical tubes. 4. Incubate the tubes at 32°C for 2 h to allow flagella to partially reassemble. 5. Centrifuge tubes at 230 g for 5 min to allow all deflagellated cells to settle at the bottom of the tube. Wrap tubes with aluminum foil so that light enters only from the top 2 cm of the tube and continue to incubate at 32°C for 2 h. Cells that have assembled flagella swim and phototax to the lighted part of the tube. Cells without flagella remain at the bottom in the unlit part of the tube. 6. Remove and discard supernatant by aspiration from the top of the tube using a 25-ml pipette and leave 5 ml of medium.

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7. Add 30 ml of fresh M-N/5 medium pre-warmed to 32°C, spin at 230 g for 5 min and incubate for 2 h at 32°C. 8. Repeat step 5. 9. Remove and discard all the supernatant aspiring from the top of the tube and resuspend the pellet in 7 ml of fresh M-N/5 at 21°C, transfer to 16-mm 125-mm sterile glass tubes with round bottom and incubate at 21°C in bright light overnight. E. Day 3. Eliminate Nonconditional Nonswimming Cells 1. Wrap tubes at 21°C in foil as described in step 5 and continue incubation at 21°C for 1 h to enrich for swimming cells. 2. Transfer the top 5 ml to a new tube. 3. Deflagellate and collect all cells by centrifugation at 350 g for 15 min at 21°C and discard the supernatant. 4. Resuspend cells in 5 ml of fresh M-N/5 medium prewarmed to 32°C using 16-mm 125-mm glass tubes. 5. Spin at 230 g for 5 min, wrap tubes with aluminum foil as described above and incubate at 32°C for 1 h. 6. Remove and discard all the supernatant aspiring from the top of the tube and resuspend the pellet in 3 mL of fresh M-N/5 at 21°C and incubate at 21°C in bright light overnight. F. Day 4. Culture Single Cells on Solid Medium 1. Wrap tube at 21°C in foil as described above and continue incubation at 21°C for 1 h and transfer most of the supernatant to a new tube. 2. Incubate at 32°C for 2 h, wrap tubes in foil, and continue incubation at 32°C for 1 h. 3. Remove all the supernatant and resuspend the pellet in 1 ml of fresh M-N/5 medium. 4. Adjust volume to obtain a suspension of 5 103 cells/ml and plate 100 µl per R plate. G. Day 10. Screen of Single-Cell Colonies 1. Incubate plates at 21°C for 4–6 days until colonies appear. Pick single colonies into 200 µl of R medium overnight at 21°C. 2. Screen for colonies that fail to swim after 4 h at 32°C. 3. Analyze cells under phase microscope for aflagellate versus paralyzed flagellar cells.

IV. Characterizing Intraflagellar Transport in fla Mutants An accurate quantitative analysis of IFT in Chlamydomonas can be achieved by applying an image analysis approach called kymography to VE-DIC recordings, which allows visualizing IFT particle trajectories in two dimensions as a diagram representing

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space (y) and time (x) (Fig. 1). When this approach is applied to the analysis of flagella of wild-type cells, we find IFT particles moving toward the distal end of flagella are larger in size, slower in velocity, and less numerous than IFT particles moving toward

Fig. 1 (A) Kymographs of flagella from pf15, pf15; fla27, pf15; fla18, pf15; fla24, and pf15; fla9 cells. Vertical (y) and horizontal (x) axes represent distance from cell body (µm) and time of observation (s), respectively. Black and white arrows indicate traces formed by anterograde or retrograde IFT particles, respectively. The IFT phase that is defective in each recombinant is indicated (see Table III for further details). The strain pf15; fla9 and other double mutants (pf15; fla4, pf15; fla5, and pf15; fla21) behave similarly to each, and are not significantly different from pf15 at 21°C and are labeled as unknown (?). Velocity, frequency ratio, and overall particle frequency can be obtained from these images. (B, C) Kymographs of pf15 flagella analyzed by different methods. Kymograph in B was generated using the approach described previously (Iomini et al., 2001: Piperno et al., 1998). The kymograph in C was generated with MetaMorph (see text). Kymograph B has had noise removed while kymograph C has not. Lower levels of noise facilitate detection of particles trajectories. A bifurcation of a trajectory indicating a possible split of one particle or a sudden change in velocity of one of two particles moving together is resolved in B but not in C (*). Recordings of pf15 in B and C, and pf15; fla27, pf15; fla18, and pf15; fla24 in A are available as Supplemental movies 1, 2, 3, and 4, respectively (http://www.elsevierdirect.com/companions/9780123749734). Panel A is reprinted from Iomini et al. (2001).

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Table III The Phases of Intraflagellar Transport Phase of IFT I: Particles bind a cargo and change in structure and/or size in a position close to the basal bodies. II: Particles and cargo are transported to the distal end of flagella by an anterograde motor. III: Particles release the cargo and change their structure and/or size at the distal end of the flagella. IV: Particles are transported to a region close to the basal bodies by a retrograde motor.

Predicted phenotype of phase-defective mutants Defects in subunits of the particles or basal bodies. Accumulation of particles at proximal end of flagella. Defects in subunits of the particles or kinesin-2. Defective anterograde velocity of IFT particles. Accumulation of particles at proximal end of flagella. Defects in subunits of the particles or components of the axoneme or matrix at the flagellar tip. Accumulation of particles at the distal end of flagella. Defects in subunits of the particles or cytoplasmic dynein. Defective retrograde velocity of IFT particles. Accumulation of particles at distal end or along flagella.

the proximal end of the flagella suggesting a model in which at least four distinct phases of the IFT are recognizable (Iomini et al., 2001). Kymographs obtained from the analysis of fla mutant flagella at the permissive temperature lead to the identification of different classes of mutants that show specific defects in one or two consecutive phases of the IFT that are indicative of the function of the defective gene (see Table III). A. VE-DIC Light Microscopy Since imaging of IFT is only possible on paralyzed and straight flagella, recombinant strains between each of the fla mutants and a paralyzed flagella (pf) mutant that affects the axonemal central complex (pf15) were used. 1. Cells to be analyzed by VE-DIC microscopy are resuspended in R or M-N/5 liquid medium and cultured from 3 to 16 h at 21°C to allow complete flagellar assembly. 2. Cells were transferred to medium containing 0.25% low melting temperature agarose cooled to 30°C (SeaPlaque). A drop of this cell suspension was than delivered, using a Pasteur pipette, into a chamber created by a #1 (22 mm square) glass coverslip and a microscope slide held together by two parallel strips of a Scotch double-side adhesive tape and immediately cooled on a 4°C metallic plate for about 1 min. 3. Higher concentrations of low melting temperature agarose (up to 0.75%) can be utilized to inhibit flagellar motility when analyzing strains not available in the pf background (Mueller et al., 2005). 4. VE-DIC of IFT in living cells was carried out using a Zeiss Axiovert 35 (Carl Zeiss, Inc., Oberkochen, Germany) microscope with a 1.4 NA condenser, a Pan-Neofluar 100/1.3 oil objective, and a 4 magnifier placed on the trinocular head in front of a video camera equipped with a Newvicon tube (Model C2400 Hamamatsu, Bridgewater, NJ).

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Transillumination light was provided by a mercury arc and filtered through a Zeiss standard green filter. Video images were acquired at a rate of 30 frames/s and stored directly on optical disks. 5. Calibration of pixel dimensions can be performed by use of a stage micrometer. In order to obtain an accurate angular alignment between the flagellar axis and the DIC axis, we mounted a home made rotating stage (similar to that used in polarizing microscopes) on the microscope. The slide was attached to the stage using grease and rotated so that the flagellum is perpendicular to the (x) axis and the cell body at the bottom of the image. Consequently, no reorientation of the images is required during image acquisition and analysis.

B. Digital Kymography Methods for obtaining kymographs were described previously (Iomini et al., 2001; Piperno et al., 1998). 1. Each image in a video sequence was read from the optical disc recorder and digitized by a frame grabber. A 7-pixel-wide light intensity profile, or linescan, along the flagellum was obtained using a built-in function of the Image-1 software package (Universal Imaging Corporation, Dowingtown, PA). 2. Noise arises from several sources that include the presence of uneven background, light intensity fluctuations, and digitization noise. Since the noise interferes with particle trajectory detection, the data were subjected to singular value decomposition and principal component analysis (Golub and Reinsch, 1970; Malinowski, 1991; Press et al., 1992). Custom software that applies these procedures to each linescan in an automated fashion was developed and is available upon request. It averages across the flagellum. Commercial software such as MetaMorph generates kymographs from image stacks as well. 3. A second method uses an 8-s analog video obtained with a 30 frames/s camera. It was converted to 240 digital frames using a media converter (Sony, San Diego, CA, DVMC-DA2). The image stack was opened and processed using MetaMorph (Universal Imaging Corporation). a. From the Stack menu, choose Kymograph. The Kymograph dialog box will appear. b. Select the source image stack with the Source Stack image selector. c. With the single line tool draw a line region across the area of interest in the image window. Because the Kymograph command draws the result image starting with the line region values in the topmost plane in the stack, you should verify that you are looking at the top plane in your stack; otherwise, your result image may appear to be inverted. d. Select the All Planes check box. e. Select the Line Width spin box and assign the value 7 or a value that cover the entire width of the flagellum. f. Select Average as a gray-scale value. g. Choose Create Kymograph and save the new image file.

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4. Figure 1B and C shows kymographs obtained with noise reduction as described in step 2 and without noise reduction by MetaMorph on the same data set. Although the noise due to light fluctuation is completely removed when singular value decomposition and principal component analysis are applied, the two plots are comparable in terms of detection of IFT particles trajectories. 5. In these plots, moving particles appeared as diagonal ridges or streaks, whose slope was proportional to their velocity. The velocity of each particle was calculated from the slope of a line drawn manually along each of the diagonal ridges. A lower limit for the value of the frequency of IFT, expressed in particles/second, was estimated from the ratio of the total number of particles detected to the total observation time, equal to the number of linescans divided by 30, the video frame rate. 6. Kymographs obtained as Tiff files were filtered using Photoshop CS3 (Adobe Corp., San Jose, CA) by applying Gaussian blur (0.7) and unsharp mask (298%, 2.7 pixel radius, 1 threshold) (Dentler, 2005).

V. Genetic Characterization A. Determining the Number of Genes Among a Collection of Mutants Few mutants in Chlamydomonas are tested for complementation in diploid strains (Ebersold, 1967). Instead, loci are generally defined by recombination mapping. 1. Once a mutant phenotype is likely to be caused by a single mutation because it shows 2:2 segregation of the mutant and wild-type phenotypes in tetrads or in random progeny, pairwise crosses can be performed (Dutcher, 1995). It is often useful to backcross a mutagenized strain three to four times before further characterization to remove unlinked mutations. 2. The phenotypes of progeny from pair-wise crosses are analyzed. Tight linkage indicates the mutants are likely to be in the same locus (no recombinants out of 30–50 progeny). Complementation in diploids can be performed (see Section V.E and Dutcher et al., 1988).

B. Linkage Analysis

1. Linkage Group Determination To determine linkage between candidate genes and flagellar assembly mutants or between a linkage group and flagellar assembly mutants, PCR-based markers can be used more efficiently than other phenotypic markers (Bowers et al., 2003; Kathir et al., 2003). DNA from progeny that pelleted at 32°C from tetrads of crosses of fla mutants CC-1952 was analyzed for segregation of the mutant allele with respect to polymorphic alleles. CC-1952, also known as S1D2, is a highly polymorphic strain (Gross et al., 1988) that has approximately one change per 80 bp (Vysotskaia et al.,

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2001). Markers developed by low coverage sequencing of the S1D2 strain has provided over 12,000 possible markers to use for mapping using PCR amplification of meiotic progeny from a cross of the mutant and CC-1952 (Rymarquis et al., 2005). There are three classes of markers. The derived cleaved amplified polymorphisms (dCAPS) markers are amplified PCR products that produce differences between the strains after digestion with a restriction enzyme. The simple sequence repeats (SSR) markers are amplified PCR products that produce products that differ in length that result from differences in the size of repetitive elements that include GT, CA, TA, TCC, TGA, and TGG. The STS markers are sequence-tagged sites that use three primers for their analysis. When mapping a new mutation, the first goal is to determine the linkage group. It is useful to determine if the mutant shows centromere linkage. Genes that are linked (<20 map units from the centromere) can be mapped with a set of primers that are linked to each of the 17 centromeres. If the gene is unlinked to its centromere, then it must be mapped to a linkage group using markers that are located in the middle of each of the 34 arms of the linkage groups.

2. Centromere Linkage 1. Isolate DNA from meiotic progeny from 10 tetrads that have scored for the flagellar phenotype. A very crude DNA preparation can be made in a 20-µl reaction. It contains 2 µl Vent Buffer (20 mM Tris–Cl pH 8.8, 10 mM KCl, 10 mM (NH4)SO4, 2 mM MgSO4, 0.1% Triton X-100 ), 1 µl proteinase K (20 mg/ml), 17 µl water, and ~105 cells. The mix is subjected to 58°C for 1 h and 95°C for 30 min. The mixture containing the DNA can be stored at 4°C for several months. A volume of 0.2 µl of the supernatant is used in 10-µl PCR reactions. 2. Perform PCR with a centromere-linked marker. HSP70A and TcTex1 are excellent markers as both are very close to their respective centromeres (<0.5 map units). If the fla mutant and the PCR marker are on different chromosomes, then the frequency of tetratype tetrads provides a way to determine the distance from FLA to its centromere. If the frequency is greater than 67%, then the FLA gene is unlinked to its centromere. If it is less, then the FLA gene is linked. The distance is described by the equation: Distance from FLA gene to its centromere = T 100/2 (number of tetrads). 3. The centromere markers and PCR conditions using Taq polymerase are shown in Table IV. 4. If the FLA gene is close (<20% tetratypes) to the centromere-linked markers, then a single marker for each chromosome can be used to detect linkage. Markers can be picked from various sources (Bowers et al., 2003; Kathir et al., 2003; Rymarquis et al., 2005). If the FLA gene is more than 20 map units from the centromere, then a marker from the middle of each arm can be used for mapping. Mapping to a chromosome can generally be accomplished using 40–50 meiotic progeny that can be collected from dissected tetrads or random progeny. 5. Once linkage is detected, the next goal is to find markers that are tightly linked to the FLA gene. On an average, 1 map unit or 1% recombination is about 100–105 kb in

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Table IV Centromere-Linked Primers and PCR Conditions Primer 1

Primer 2

TcTex1 ATT GAC GTC TCG ACC TCC TGT TGC GAA GAG GA ACC ATT TC HSP70A AGC TGC TGC AGG GCT GGT TGT CGG ACT TCT TC AGT AGG TC a

Anneal T (°C)

[Mg] (mM)

PCR product Enzyme (bp)

Band sizesa

55

3.5

400

HaeIII

400/500

55

2.7

400

HaeIII

180/200

The distinguishing band from the CC-125 parent is listed first and the CC-1952 parent is listed second.

length. Regions near the centromere or telomere appear to have less recombination. Mapping to an interval of 1–2 map units generally requires 300–500 progeny. This generates a small enough region to allow identification by transformation and rescue.

3. Linkage with Candidate Genes Candidate genes provide a useful way to identify the gene more quickly than mapping across the entire genome. In the identification of FLA8, biochemical data suggested that it was missing kinesin-2 (Cole et al., 1998). Thus, it was straightforward to map fla8 and fla1 strains with respect to the kinesin-2 subunits in the genome (Miller et al., 2005). The flagellar assembly mutants that show defective retrograde IFT are another example. Five mutants (fla2, fla15, fla16, fla17, and fla24) show decreased retrograde velocities, an increased ratio of anterograde to retrograde particles and an accumulation of complex B proteins in the flagella (Iomini et al., 2001). Sequences for complex A proteins were used to make dCAP markers and each were mapped relative to each of these mutants. 1. Design dCAPs markers. If there are no markers that have been developed for a gene of interest, the 30 UTR or stretches of repeats that are at least 100 bp long provide the easiest ways to develop a new marker. 2. Sets of primers that produce a product of 200–400 bp are chosen. 3. Each primer is subjected to BLAST to the JGI genome browser to ask if the primer is unique in the genome at an expected value of 1e–1. If there are multiple hits, then new primers need to be picked. 4. If there is no size difference between PCR products for the mutant and the CC-1952 strain is observed, six to eight restriction enzymes that have GC-rich recognition sites are tested for one that produces size differences between the two parental strains (Fig. 2). 5. Linkage is tested in 40–50 meiotic progeny from a cross of the mutant by CC-1952. The progeny are generally chosen to have the mutant phenotype. If linkage is found, then the candidate gene is sequenced from the mutant.

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1

2

3

4

5

6

7

8

9

M

P

S

Fig. 2 PCR products from meiotic progeny (1–9) of a cross of the mutant parent (P) by the CC-1952 (S1D1) strain. The primers span a repetitive GT region and the two strains have a size polymorphism. This marker is linked (9/9) to the mutant phenotype.

6. Reversion of the mutant phenotype can be used to provide further evidence (Rymarquis et al., 2005; Iomini et al., 2009) or transformation and rescue. C. Chlamydomonas Transformation and SHIRT This protocol is modified from other protocols (Kovar et al., 2002; Shimogawara et al., 1998). 1. Chlamydomonas cells were inoculated in 100 ml liquid R medium for 3 days under continuous illumination with gentle shaking until cells reached a concentration of ~5 106 cells/ml. 2. Cells were collected by centrifugation and treated with autolysin for 0.5 h at room temperature to remove cell walls (Dutcher, 1995; Harris, 1989). Sterility must be maintained throughout the protocol. 3. Autolysin-treated cells were chilled on ice for 10 min before collected by centrifugation at 4°C at 1000 g. Cells were gently resuspended on ice in R þ 100 mM mannitol to the final concentration of ~4 108 cells/ml. 4. A volume of 250 µl of cells (~1 108 cells) was used for transformation with 6 µg of bacterial artificial chromosome (BAC) DNA or 1 µg plasmid DNA with or without (the addition of 1 µg of pSI103, which confers resistance to paromomycin (Sizova et al., 2001), for cotransformation). 5. Cells and DNA were added to an electroporation cuvette (4-mm gap, Bio-Rad, Hercules, CA) and incubated in a 16°C water bath for 5 min before electroporation, which was performed in a Bio-Rad Gene Pulser II with the following setting: 0.75 kV, 25 µF, and 50 . Cells were electroporated with one pulse and incubated at room temperature for 10 min before transfer to 50 ml R þ 100 mM mannitol liquid medium and incubated overnight at room temperature with continuous illumination. 6. Cells were resuspended gently in 1 ml 25% cornstarch in R medium. 7. Cells are innoculated onto five R plates with 10 µg/ml paromomycin for cotransformation experiments. Colonies appear within 5–7 days at 25°C. 8. Without cotransformation, the cells are inoculated into 48 tubes (25 mm 150 mm) with 20-ml R liquid medium. In liquid medium, the upper 5–10 ml of medium is transferred to new R liquid 4–5 over the course of 2 weeks. Cells are transferred

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before they reach a density of 5 106 to prevent cells entering stationary phase where they become aflagellated. Once swimming cells are detected, 50 µl of medium to plated to solid medium for single colonies. A single colony that is able to swim at the restrictive temperature is selected for further study from each tube. 9. Each rescued transformant is crossed to a wild-type strain (CC-124) to determine if the rescue is extragenic.

1. SHIRT Analysis Because a BAC often contains many genes, rescue does not always reveal which gene is encoded by the locus of interest. One can subclone fragments of the BAC and transform with the smaller pieces. Another option is the Segregation of Heterozygosity in Rescued Transformants (SHIRT) method (Esparza, 2008). The SHIRT analysis provides a rapid method to determine which regions of BAC DNA are integrated nonhomologously into the genome and cosegregate with the rescued phenotype. It obviates the requirement for subcloning to determine the causative DNA for rescued transformants. It is rare in Chlamydomonas that the entire transforming DNA of a BAC or plasmid integrates into the genome, and independent transformants generally integrate nonidentical segments of the BAC or plasmid DNA (Dutcher and Trabuco, 1998; Myster et al., 1999). 1. For this analysis, multiple independent transformants are mated to the polymorphic strain CC-1952 and meiotic tetrad progeny are recovered. 2. Nonparental ditype (NPD) tetrads are identified that have two mutant progeny (fla) and two wild-type progeny (FLA BAC DNA). A dCAPS marker is made for each gene in the BAC used for transformation (Fig. 3). As discussed above, markers predicted by Rymarquis et al. (2005) are tested first. If no predictions are available, new primers can be designed as described above. 3. In the progeny, from the cross in step 1, there are three alleles of each gene. One copy will be from the CC-1952 parent, one from the inserted BAC, and one from the mutant parent. Since the BAC library was made from the same strain as the mutant parent, the alleles contributed by the BAC and the mutant parent will possess the same polymorphism while the allele contributed by the CC-1952 parent will be different. PCR is performed for each marker the progeny that show an NPD segregation pattern for the flagellar phenotype and the rescue (Fig. 3). 4. The two aflagellate progeny should have only a copy of the parental allele and the two swimming progeny have the BAC and CC-1952 alleles (Fig. 3; see Gene 2 diagram), if the gene is responsible for rescue. 5. If the gene from the BAC is not integrated into the genome, then the PCR products will resemble the Gene 3 example (Fig. 3). 6. If the gene from the BAC does not rescue, the pattern may resemble Gene 1 and not cosegregate (Fig. 3C). 7. DNA was isolated from four NPD tetrads for each transformant and the segregation of genes from the BAC determined. Figure 4 shows the pattern for rescue of the fla15 mutant.

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Fig. 3 Schematic diagram of SHIRT. SHIRT uses mapping and PCR-based markers to determine which parts of a BAC are responsible (cosegregate) for rescue of a mutant phenotype following transformation. (A) Diagram of three genes on a BAC from the mutant parent (black), the polymorphic mapping strain CC-1952 (red), and the BAC (blue). Primers for PCR are generally made to the 30 UTR of genes to be tested (arrows). (B) Bands of digested PCR products (dCAP markers) for the three genes from the three sources of DNA. The digested PCR products from the black and blue alleles cannot be distinguished from each other. (C) Three possible outcomes among NPD tetrads in which the mutant phenotype is observed in two of the four progeny (fla and þ). The pattern with Gene 1 suggests that the BAC DNA is not responsible for rescue. The hybrid (Black and Blue) band indicates that the band is amplified from both the mutant and the BAC allele. The pattern with Gene 2 is consistent with the BAC DNA providing rescue. Further proof requires additional NPD tetrads with this pattern. The pattern with Gene 3 suggests that it is not integrated into the genome and therefore not responsible for rescue. This figure is reprinted from Iomini et al. (2009). (See Plate no. 6 in the Color Plate Section.)

D. Reversion of Flagellar Mutants Using Ultraviolet Light 1. Cultures of mutant cells were plated onto R medium at a density of 2000 colonies per plate and grown for 4 days at 21°C. They were subjected to ultraviolet irradiation (70 J) and left in the dark for 18 h to prevent photoreactivation. An ultraviolet lamp that has been calibrated or a Stratagene UV Stratalinker 1800 La Jolla, CA can be used.

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fla15 Patent

FLA15 Patent

fla15

FLA15 FLA15 TG TG

fla15

433 380 299 144

Fig. 4 SHIRT analysis of the FLA15 transgene. PCR products from an NPD are shown. The parents (fla15 þ BAC; CC-1952) are in lanes 1 and 2. The four meiotic progeny are in lanes 3–6. Lanes 3 and 6 are from cells aflagellate at 32°C and the band is consistent with the mutant allele. Lanes 4 and 5 are from flagellated cells and the bands are consistent with the wild-type FLA15 allele from the CC-1952 parent and the BAC contributed transgene.

2. Approximately 40 mutagenized colonies are transferred to 20 ml of R medium in 25-mm 150-mm tubes and placed at 32°C. A volume of 5 ml of supernatant was transferred to new tubes after 3 days and this was repeated four times. 3. When swimming cells are visible, an aliquot with 3 102 cells is plated to achieve individual colonies and one colony from each tube with swimming cells is retained and scored for flagellar assembly. 4. Cross each swimming strain by wild-type cells to determine if it is intragenic (failure to recover the fla mutant phenotype) or extragenic (recovery of the fla mutant phenotype). 5. The intragenic revertants can be analyzed by a dCAPs marker if the mutant creates or destroys a restriction enzyme restriction site or by sequencing (Miller et al., 2005). 6. The outcome from reversion analysis is highly dependent on the starting allele and locus. This is illustrated by the outcomes of experiments with different genes and different alleles. Some alleles give only intragenic revertants or extragenic suppressors while others give both types of events (Table V). It appears to be allele specific and not locus specific. 7. Intragenic revertants can be used for correlating structure and function of the mutated protein (Miller et al., 2005). E. Diploid Strain Construction and Analysis Diploid strains are useful for determining if the mutations are recessive and can be selected by several different selective markers. Since less than 0.1% of zygotes become mitotically growing diploid strains, it is important to have a selection for them. 1. One selection uses the arg2 and arg7 alleles. These are alleles that show intra-allelic complementation in the arginosucinate lyase locus (ARG7). The two parents both carry an auxotrophic mutation and must be grown on medium containing arginine. Diploid strains are selected on medium lacking arginine. One disadvantage of this method arises from the acquisition of additional mutations that block further rounds of meiosis.

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Table V Outcomes for Isolating Revertants and Suppressors of fla Mutants Gene fla10-1 fla10-14 fla10-14a fla10-15 fla8-1 fla8-2 fla15-1 fla17-2 a

Number of intragenic events 181 22 19 13 15 35 56 24

Percent intragenic events 98.4 32 100 21 100 100 100 0

References Lux and Dutcher (1991) Lux and Dutcher (1991) Miller et al. (2005) Lux and Dutcher (1991) Miller et al. (2005) Miller et al. (2005) Iomini et al. (2009) Iomini et al. (2009)

Independent experiment.

2. A second set of selective markers is nit2, AC17 and NIT2, ac17 strains. The diploid strains are selected on minimal medium lacking acetate with 2 mM KNO2 instead of ammonium in the medium. This method is often easy as many of the chemically induced flagellar mutants have the nit2, AC17 genotype. It has one disadvantage in that the nit2 parent shows some growth on the medium. Isolation of the diploid strains requires a second round of streaking onto selective medium to remove the parental strain.

VI. Characterization of Genetic Interactions A. Synthetic Phenotypes Synthetic phenotypes are often used to build networks of interactions in many cellular processes (Ooi et al., 2006). Many double mutant combinations of hypomorphic alleles for genes that act in the same pathway show synthetic phenotypes, but mutants in two independent pathways can also underlie synthetic phenotypes. In some cases, these alternatives can be distinguished by comparison of null alleles. Double mutants between various conditional flagellar assembly mutants suggest that this is a useful method. 1. Construct haploid strains with two mutations by crossing two fla mutations. Each of the haploid fla mutant strains assemble flagella at 21°C but not at 32°C. 2. Check for phenotype in the progeny with two mutations. In NPD tetrads, two of the progeny will be wild type and swimming as they have both wild-type alleles. The other two progeny will have both fla mutants. In some examples, the double mutants have a phenotype at the permissive temperature (Table VI). The appearance of a synthetic phenotype with partial loss of function alleles indicates that the genes act in the same pathway (Hereford and Hartwell, 1974). 3. Additional conditional and null alleles would strengthen these arguments about the presence and absence of interactions (Iomini et al., 2009).

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Table VI Interpretations of Synthetic Phenotypes Phenotypic observation fla15; fla17 double mutants are aflagellate at 21°C fla15; fla11 double mutants are aflagellate at 21°C fla15; fla10 double mutants are unaffected and flagellate at 21°C

Statement

Conclusions

Mutants in two complex A proteins increase the severity of the mutant phenotype Mutants in a complex A and a complex B protein increase the severity of the mutant phenotype A mutant in complex A and kinesin-2 do not increase the severity of the phenotype at the permissive temperature

These proteins may interact in the IFT particle Complex A may interact with complex B through IFT172 and IFT144 Complex A and kinesin-2 do not interact

B. Second-Site Noncomplementation Second-site noncomplementation, also known as dominant enhancement, is defined as the lack of complementation between two recessive mutations in two unlinked loci (Hawley and Gilliland, 2006). It can arise via a number of mechanisms, which include the creation of a poisonous interaction complex or the reduction in the dose of a complex formed by the two gene products. 1. Noncomplementation requires that the mutations be recessive. Diploid strains heterozygous for fla mutants must be tested for dominance. For example, diploid strains heterozygous for the fla2-1, fla11-1, fla15-1, fla17-1, or fla24-1 alleles are able to swim at 21 and 32°C; all of these mutations are recessive to the wild-type allele. 2. Diploid strains that are heterozygous for all pair-wise combinations of fla mutations are constructed. For the retrograde defects (Iomini et al., 2009) and for suppressors of pf10 (Dutcher et al., 1988), many combinations show second-site noncomplementation. 3. Many of the double heterozygous diploid strains (fla15, FLA15, fla17, FLA17) show a more extreme phenotype than the homozygous diploid strains at 21°C. This suggests that IFT particles are more compromised in these combinations at the permissive temperature. Again this provides evidence for interactions. 4. The availability of both conditional and null alleles would help in the analysis of these interactions. If null alleles show the phenotype, then the noncomplementation cannot arise from poisonous interactions. It would be likely to reflect a strict requirement for equal dosage of the two genes. C. Two-Copy Suppression The presence of multiple copies of a gene or overexpression of a gene can suppress the phenotype of mutations in a noncognate gene in Saccharomyces cerevisiae and Schizosaccharomyces pombe (Forsburg, 2001). This observation has been termed

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multicopy suppression and has been used extensively to find genes that act in the same pathway. Dosage suppressors may occur because the protein at the increased dosage helps to stabilize the mutant protein or the increased dosage protein may act downstream to bypass the requirement for the mutant protein. Multicopy suppression has not been sought or observed extensively in Chlamydomonas. Recently, we have found examples of it. 1. To observe two-copy suppression, mutants that have been rescued by a cognate transgene are crossed with other fla mutants. The strains can be genotyped with dCAPs markers. For example, two copies of the wild-type IFT144 gene (FLA15) can rescue the fla17-1 mutant phenotype at 32°C. 2. The pattern of rescue by two wild-type copies of a gene may provide information about the number of copies of a protein in a complex. For example, two copies of the IFT144 gene partially rescue the phenotype of fla11-1 cells, but do not rescue the mutant phenotypes of the fla2, fla8, fla10, or fla24 strains (Iomini et al., 2009). Two copies of the wild-type IFT139 gene (FLA17) rescue the fla15-1 mutant phenotype. Two copies of the IFT139 gene do not rescue the fla2, fla8, fla10, fla11, or fla24 mutant phenotypes. 3. To demonstrate that the rescue requires two wild-type copies, rescue is tested in a double mutant where there cannot be two wild-type copies of the gene in question. For example, a single wild-type copy of IFT139 or IFT144 rescues the aflagellate phenotype of the fla15; fla17 double mutant at 21°C, but not at 32°C. One interpretation is that two-copy suppression results from stabilization of the mutant gene product in the presence of an excess in the gene product from the noncognate transgene product together with the endogenous copy of the gene. 4. If two-copy suppression is not reciprocal (two copies of gene 1 rescues mutant 2 and two copies of gene 2 rescues mutant 1), it may suggest that there is more than one copy of one of the proteins is present in each IFT particle. 5. The availability of null alleles would be useful to ask if two copies are sufficient or whether the interaction requires the mutant gene product.

References Adams, G.M., Huang, B., and Luck, D.J. (1982). Temperature-sensitive, assembly-defective flagella mutants of Chlamydomonas reinhardtii. Genetics 100, 579–586. Bowers, A.K., Keller, J.A., and Dutcher, S.K. (2003). Molecular markers for rapidly identifying candidate genes in Chlamydomonas reinhardtii. Ery1 and ery2 encode chloroplast ribosomal proteins. Genetics 164, 1345–1353. Brazelton, W.J., Amundsen, C.D., Silflow, C.D., and Lefebvre, P.A. (2001). The bld1 mutation identifies the Chlamydomonas osm-6 homolog as a gene required for flagellar assembly. Curr. Biol. 11, 1591–1594. Cole, D.G., Diener, D.R., Himelblau, A.L., Beech, P.L., Fuster, J.C., and Rosenbaum, J.L. (1998). Chlamydomonas kinesin-II-dependent intraflagellar transport (IFT): IFT particles contain proteins required for ciliary assembly in Caenorhabditis elegans sensory neurons. J. Cell Biol. 141, 993–1008. Dentler, W. (2005). Intraflagellar transport (IFT) during assembly and disassembly of Chlamydomonas flagella. J. Cell Biol. 170, 649–659.

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Carlo Iomini et al. Dutcher, S.K. (1995). Mating and tetrad analysis in Chlamydomonas reinhardtii. Methods Cell Biol. 47, 531–540. Dutcher, S.K., Gibbons, W., and Inwood, W.B. (1988). A genetic analysis of suppressors of the PF10 mutation in Chlamydomonas reinhardtii. Genetics 120, 965–976. Dutcher, S.K., and Trabuco, E.C. (1998). The UNI3 gene is required for assembly of basal bodies of Chlamydomonas and encodes delta-tubulin, a new member of the tubulin superfamily. Mol. Biol. Cell 9, 1293–1308. Ebersold, W.T. (1967). Chlamydomonas reinhardi: Heterozygous diploid strains. Science 157, 447–449. Esparza, J.M. (2008). Epsilon tubulin mutants affect basal body integrity, disrupt katanin localization and increase microtubule stability. Ph. D. Thesis. Washington University, St. Louis. Forsburg, S.L. (2001). The art and design of genetic screens: Yeast. Nat. Rev. Genet. 2, 659–668. Golub, G.H., and Reinsch, C. (1970). Singular value decomposition and least squares solutions. Numer. Math. 14, 403–420. Goodenough, U., Lin, H., and Lee, J.H. (2007). Sex determination in Chlamydomonas. Semin. Cell Dev. Biol. 18, 350–361. Gross, C.H., Ranum, L.P., and Lefebvre, P.A. (1988). Extensive restriction fragment length polymorphisms in a new isolate of Chlamydomonas reinhardtii. Curr. Genet. 13, 503–508. Harris, E.H. (1989). “The Chlamydomonas Sourcebook.” Academic Press, Inc., San Diego. Hawley, R.S., and Gilliland, W.D. (2006). Sometimes the result is not the answer: The truths and the lies that come from using the complementation test. Genetics 174, 5–15. Hereford, L.M., and Hartwell, L.H. (1974). Sequential gene function in the initiation of Saccharomyces cerevisiae DNA synthesis. J. Mol. Biol. 84, 445–461. Hou, Y., Pazour, G.J., and Witman, G.B. (2004). A dynein light intermediate chain, D1bLIC, is required for retrograde intraflagellar transport. Mol. Biol. Cell 15, 4382–4394. Hou, Y., Qin, H., Follit, J.A., Pazour, G.J., Rosenbaum, J.L., and Witman, G.B. (2007). Functional analysis of an individual IFT protein: IFT46 is required for transport of outer dynein arms into flagella. J. Cell Biol. 176, 653–665. Huang, B., Rifkin, M.R., and Luck, D.J. (1977). Temperature-sensitive mutations affecting flagellar assembly and function in Chlamydomonas reinhardtii. J. Cell Biol. 72, 67–85. Iomini, C., Babaev-Khaimov, V., Sassaroli, M., and Piperno, G. (2001). Protein particles in Chlamydomonas flagella undergo a transport cycle consisting of four phases. J. Cell Biol. 153, 13–24. Iomini, C., Li, L., Esparza, J.M., and Dutcher, S.K. (2009). Retrograde IFT mutants identify complex A proteins with multiple genetic interactions in Chlamydomonas reinhardtii. Genetics, 183, in press. Johnson, D.E., and Dutcher, S.K. (1991). Molecular studies of linkage group XIX of Chlamydomonas reinhardtii: Evidence against a basal body location. J. Cell Biol. 113, 339–346. Kathir, P., LaVoie, M., Brazelton, W.J., Haas, N.A., Lefebvre, P.A., and Silflow, C.D. (2003). Molecular map of the Chlamydomonas reinhardtii nuclear genome. Eukaryotic Cell 2, 362–379. Kovar, J.L., Zhang, J., Funke, R.P., and Weeks, D.P. (2002). Molecular analysis of the acetolactate synthase gene of Chlamydomonas reinhardtii and development of a genetically engineered gene as a dominant selectable marker for genetic transformation. Plant J. 29, 109–117. Kozminski, K.G., Johnson, K.A., Forscher, P., and Rosenbaum, J.L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523. Lux, F.G., III, and Dutcher, S.K. (1991). Genetic interactions at the FLA10 locus: Suppressors and synthetic phenotypes that affect the cell cycle and flagellar function in Chlamydomonas reinhardtii. Genetics 128, 549–561. Malinowski, E.R. (1991). “Factor Analysis in Chemistry.” John Wiley & Sons Inc., New York. Miller, M.S., Esparza, J.M., Lippa, A.M., Lux, F.G., III, Cole, D.G., and Dutcher, S.K. (2005). Mutant kinesin-2 motor subunits increase chromosome loss. Mol. Biol. Cell 16, 3810–3820. Mueller, J., Perrone, C.A., Bower, R., Cole, D.G., and Porter, M.E. (2005). The FLA3 KAP subunit is required for localization of kinesin-2 to the site of flagellar assembly and processive anterograde intraflagellar transport. Mol. Biol. Cell 16, 1341–1354. Myster, S.H., Knott, J.A., Wysocki, K.M., O’Toole, E., and Porter, M.E. (1999). Domains in the 1alpha dynein heavy chain required for inner arm assembly and flagellar motility in Chlamydomonas. J. Cell Biol. 146, 801–818.

7. Flagellar Assembly Mutants in Chlamydomonas reinhardtii

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Ooi, S.L., Pan, X., Peyser, B.D., Ye, P., Meluh, P.B., Yuan, D.S., Irizarry, R.A., Bader, J.S., Spencer, F.A., and Boeke, J.D. (2006). Global synthetic-lethality analysis and yeast functional profiling. Trends Genet. 22, 56–63. Pan, J., and Snell, W.J. (2002). Kinesin-II is required for flagellar sensory transduction during fertilization in Chlamydomonas. Mol. Biol. Cell 13, 1417–1426. Pazour, G.J., Dickert, B.L., Vucica, Y., Seeley, E.S., Rosenbaum, J.L., Witman, G.B., and Cole, D.G. (2000). Chlamydomonas IFT88 and its mouse homologue, polycystic kidney disease gene tg737, are required for assembly of cilia and flagella. J. Cell Biol. 151, 709–718. Pazour, G.J., Dickert, B.L., and Witman, G.B. (1999). The DHC1b (DHC2) isoform of cytoplasmic dynein is required for flagellar assembly. J. Cell Biol. 144, 473–481. Pedersen, L.B., Miller, M.S., Geimer, S., Leitch, J.M., Rosenbaum, J.L., and Cole, D.G. (2005). Chlamydomonas IFT172 is encoded by FLA11, interacts with CrEB1, and regulates IFT at the flagellar tip. Curr. Biol. 15, 262–266. Piperno, G., and Mead, K. (1997). Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella. Proc. Natl. Acad. Sci. USA 94, 4457–4462. Piperno, G., Mead, K., and Henderson, S. (1996). Inner dynein arms but not outer dynein arms require the activity of kinesin homologue protein KHP1(FLA10) to reach the distal part of flagella in Chlamydomonas. J. Cell Biol. 133, 371–379. Piperno, G., Siuda, E., Henderson, S., Segil, M., Vaananen, H., and Sassaroli, M. (1998). Distinct mutants of retrograde intraflagellar transport (IFT) share similar morphological and molecular defects. J. Cell Biol. 143, 1591–1601. Porter, M.E., Bower, R., Knott, J.A., Byrd, P., and Dentler, W. (1999). Cytoplasmic dynein heavy chain 1b is required for flagellar assembly in Chlamydomonas. Mol. Biol. Cell 10, 693–712. Press, W.H., Teukolski, S.A., Vetterling, W.T., and Flannery, B.P. (1992). “Numerical Recipes in C: The Art of Scientific Computing.” Cambridge University Press, Cambridge, UK. Rymarquis, L.A., Handley, J.M., Thomas, M., and Stern, D.B. (2005). Beyond complementation. Map-based cloning in Chlamydomonas reinhardtii. Plant Physiol. 137, 557–566. Shimogawara, K., Fujiwara, S., Grossman, A., and Usuda, H. (1998). High-efficiency transformation of Chlamydomonas reinhardtii by electroporation. Genetics 148, 1821–1828. Sizova, I., Fuhrmann, M., and Hegemann, P. (2001). A Streptomyces rimosus aphVIII gene coding for a new type phosphotransferase provides stable antibiotic resistance to Chlamydomonas reinhardtii. Gene 277, 221–229. Tam, L.W., and Lefebvre, P.A. (1993). Cloning of flagellar genes in Chlamydomonas reinhardtii by DNA insertional mutagenesis. Genetics 135, 375–384. Vysotskaia, V.S., Curtis, D.E., Voinov, A.V., Kathir, P., Silflow, C.D., and Lefebvre, P.A. (2001). Development and characterization of genome-wide single nucleotide polymorphism markers in the green alga Chlamydomonas reinhardtii. Plant Physiol. 127, 386–389. Walther, Z., Vashishtha, M., and Hall, J.L. (1994). The Chlamydomonas FLA10 gene encodes a novel kinesin-homologous protein. J. Cell Biol. 126, 175–188. Wang, Q., Pan, J., and Snell, W.J. (2006). Intraflagellar transport particles participate directly in ciliumgenerated signaling in Chlamydomonas. Cell 125, 549–562.

CHAPTER 8

Recording and Analyzing IFT in Chlamydomonas Flagella William Dentler*, Kristyn VanderWaal†, and Mary E Porter† *

Department of Molecular Biosciences, University of Kansas, Lawrence, Kansas 66045

† Department of Genetics, Cell Biology, and Development, University of Minnesota, Minneapolis, Minnesota 55455

Abstract I. Introduction II. Methods A. Slide Preparation B. Specimen Preparation C. Microscopy D. Recording IFT E. IFT analysis III. Summary Acknowledgments References

Abstract The transport of materials to and from the cell body and tips of eukaryotic flagella and cilia is carried out by a process called intraflagellar transport, or IFT. This process is essential for the assembly and maintenance of cilia and flagella: in the absence of IFT, cilia cannot assemble and, if IFT is arrested in ciliated cells, the cilia disassemble. The major IFT complex proteins and the major motor proteins, kinesin-2 and osm-3 (which transport particles from the cell body to ciliary tips) and cytoplasmic dynein 1b (which transports particles from ciliary tips to the cell body) have been identified. However, we have little understanding of the structure of the IFT particles, the cargo that these particles carry, how cargo is loaded and unloaded from the particles, or how METHODS IN CELL BIOLOGY, VOL. 93 Copyright Ó 2009 Elsevier Inc. All rights reserved.

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the motor proteins are regulated. The focus of this chapter is to provide methods to observe and quantify the movements of IFT particles in Chlamydomonas flagella. IFT movements can be visualized in paralyzed or partially arrested flagella using either differential interference contrast (IFT) microscopy or, in cells with fluorescently tagged IFT components, with fluorescence microscopy. Methods for recording IFT movements and analyzing movements using kymograms are described.

I. Introduction Intraflagellar transport (IFT) first was observed using a specialized differential interference contrast (DIC) microscope and recording system (Kozminski, 1995; Kozminski et al., 1993). The elegant movies of particle movement within flagella and use of the tsmutant fla10 to identify the anterograde motor for IFT (Kozminski et al., 1995) led to a new area in studies of cilia and flagella and the discovery of ciliary defects associated with a number of human diseases, or ciliopathies (Badano et al., 2006; Marshall, 2008). Analysis of the rapid bidirectional movement of IFT particles in 0.2-µm diameter flagella was made possible by the development of kymograms that displayed particle positions along the flagellum as a function of time (Iomini et al., 2001; Piperno et al., 1998). Although these techniques were major advances in the field, they required equipment and computer programs that were not generally available, so their application was limited. Recent improvements in microscope optics, recording equipment, and computer programs have made it possible for most investigators to visualize IFT at modest cost (Dentler, 2005; Mueller et al., 2005). These techniques permit the analysis of anterograde and retrograde IFT as well as the movement of particles along flagellar surfaces using high-resolution DIC (Fig. 1). With the development of green fluorescent protein (GFP)-tagged IFT components (Mueller et al., 2005; Snow et al., 2004) IFT can be observed using wide-field fluorescence microscopy (Fig. 2). The purpose of this chapter is to provide methods that can be used to observe, record, and measure IFT using commercially available equipment. It will be focused on observations of IFT in Chlamydomonas flagella, because the long flagella are ideal specimens to observe IFT. IFT can be observed using DIC and, with GFP-tagged IFT components, by fluorescence microscopy. The analytical methods described here also are useful for analyzing fluorescently labeled flagella, regardless of the organism from which images were captured.

II. Methods The most important decision is to choose cells with long and relatively immobile flagella. In our hands, pf18, a Chlamydomonas mutant that lacks central microtubules and has fairly straight and rigid flagella, is the easiest strain with which to observe IFT. The flagella attach to coverslips and IFT can be observed throughout the flagellum from the base, where flagella exit the cell wall, to the distal tips. Other paralyzed flagellar mutants to consider include pf16, which has an unstable C1 microtubule, and pf1, which lacks

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(C)

(A) (B)

(D)

Fig. 1 IFT particle movement analyzed using kymographs. (A) A flagellum with IFT particles (arrowheads) is shown to the left of “A” and a kymograph showing the movement of these particles is at the right of the flagellum. Anterograde particles move toward the flagellar tip (black arrows) and smaller retrograde particles move toward the base of the flagellum (white arrows). The white arrowhead points to a larger and slowly moving particle, likely moving along the flagellar surface. (B) Thin section of Chlamydomonas flagella showing IFT particles with higher magnification images shown in (C) and (D). It is not known if these are anterograde or retrograde particles.

DIC

GFP

Fig. 2 Comparison of IFT by DIC and wide-field fluorescence microscopy recorded with a digital camera. The vertical bar is 1 µm and the horizontal bar is 1 s.

spoke heads (Kozminski, 1995). For many experiments, motile cells need to be examined, and so methods to immobilize these cells also are described in this chapter.

A. Slide Preparation Slides and coverslips must be clean for optimal imaging. They can be cleaned by soaking in hot water and detergent followed by extensive washing with deionized water and air or oven drying. Alternatively, they can be cleaned by an overnight soak in

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6 N HCl followed by extensive washing in deionized water, rinsing in 95% EtOH, and air drying (Kozminski, 1995). For DIC, thin (#0) coverslips can be used as a slide and coverslip. An aluminum slide is cut from a 1-mm-thick aluminum sheet and an opening, ~7.5 cm 3 cm, is cut out for the coverslip. A clean 24-mm 60-mm No 0 coverslip (Gold Seal #3223) is attached to the bar by double stick tape and two small (~2 mm 20 mm) Parafilm spacers are placed on the large coverslip before mounting cells attached to a smaller coverslip (below) to avoid crushing and deflagellating cells. DIC and fluorescence microscopy also can be carried out using standard cleaned glass slides and #1 or 1.5 coverslips. If cells are mounted in agar (below), the spacers are not needed. B. Specimen Preparation Observation of IFT in paralyzed mutants is relatively straightforward and primarily requires patience. Observation of IFT in nonparalyzed cells requires mechanical or chemical methods to paralyze flagella. In each of these methods, cells are applied to coverslips and allowed to sit for 2–5 min. This promotes attachment of flagella to the coverslips, which is essential for observing IFT. Attachment of flagella to coverslips is facilitated by coating coverslips with 0.01–1 mg/ml poly-L-lysine (Sigma Corporation, St. Louis, MO P1274). Higher concentrations of polylysine will induce flagella to curl and to detach from cells (Bloodgood, 1981). Polylysine is applied to clean cover glasses and allowed to sit for 5 min to overnight. Then rinse coverslips with water and air dry. Apply cells to the coverslips, allow cells to attach, and then invert the coverslips over the larger coverslip or slide with Parafilm spacers. Withdraw as much of the medium as possible using filter paper and seal the slides with VALAP. Agarose can be used to mechanically immobilize motile cells. Kozminski (1995) formed a small agar chamber beneath the coverslip and placed cells in the chamber. We have had more success using 1–2% low EEO agarose (Fisher Scientific BP160) dissolved in culture medium and maintained at ~30°C. Apply the cells to a coverslip and allow them to attach for 2–5 min. Remove most of the media, add 8–10 µl of the agarose solution to the cells, invert over a slide, and seal with VALAP. VALAP is made by adding equal weights of Vaseline, lanolin, and paraffin to a beaker and heating to melt the mixture. VALAP can be stored indefinitely at room temperature. For use, melt VALAP and apply to the slide with a glass Pasteur pipette. Flatten the bead of VALAP with a warm spatula to minimize the possibility of contaminating the objective lens. Motile flagella can be paralyzed by adding 20 mM LiCl to the mounting medium. Although LiCl induces flagellar growth (Nakamura et al., 1987; Periz et al., 2007), little growth occurs in the time during which IFT is recorded and 20 mM LiCl does not induce changes in the rate or frequency of anterograde or retrograde IFT (Dentler, 2005). Used cautiously, addition of 10–20 mM LiCl can facilitate recording IFT in motile flagella.

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C. Microscopy

1. Differential Interference Contrast IFT was discovered using DIC microscopy and high-resolution objective and condenser lenses. Our best images are obtained using infinity-corrected optics and a Zeiss (Carl Zeiss, Thornwood, NY, USA) Axioplan 2ie upright microscope. Most flagella remain attached to the cover glass in the upright microscope but it is possible that an inverted microscope may maintain better attachment of flagella and better imaging. Full and even illumination of the back focal plane of the condenser lens is essential for high-resolution imaging. With fixed tube length optics, a mercury or xenon lamp and a fiber-optic scrambler was essential to obtain sufficient illumination to produce high-quality images (Kozminski, 1995; Piperno et al., 1998). In our experience, this intense light damages the cells even after short periods of illumination. Light transmission is greatly improved with newer infinity-corrected optics and we currently use a standard halogen lamp built into the microscope base. Together with a UV- and IR-absorbing filter and a green interference filter, cells can be observed for at least 3 h without any visible damage. High-resolution DIC requires objective and condenser lenses with numerical apertures of 1.4 or greater. Immersion oil must be used between the slide and condenser lens and between the coverslip and objective lens. The microscope must be properly aligned to produce full Koehler illumination. To record IFT the long axis of the flagellum should fill the camera field as much as possible. This requires 100 NA1.4 planapochromat (for DIC) or 100 NA 1.3 plan fluar (for fluorescence) objective lenses. The image should be projected on the camera face using a 1.6 Bertrand lens (Optovar), and an 8 eyepiece or a 1.2 TV tube magnifier. a. Specimen Orientation. Proper specimen orientation is essential. Normally, flagella are best observed when their long axis is perpendicular to the shear axis. For IFT, flagella must be positioned so that their long axis is parallel to the shear axis. The IFT particles then will move perpendicular to the shear axis and be shadowed. When properly oriented, the flagellum may be nearly invisible when observed through the microscope eyepieces but the increased contrast provided by the video camera and computer will make the flagellum and IFT visible on a monitor. While flagella in the proper orientation to the shear axis can be found by randomly scanning the slide, it is helpful to have a properly aligned rotating stage, so that the flagellum can be rotated to lie in the proper shear axis. For our Axioplan 2ie microscope, if a cell body is in the center of the field, the flagellum should point to ~10 o’clock for optimal orientation. Flagella are most easily selected by scanning by eye. To observe flagella, close the condenser aperture diaphragm and scan the field. When the specimen to be recorded is found, open the diaphragm to fill the back focal plane for maximum resolution, and view the flagella on the monitor.

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2. Fluorescence Microscopy IFT also can be observed in tissues or in protozoan flagella if IFT particles contain fluorescently tagged molecules (Mueller et al., 2005; Snow et al., 2004). Some microscopy facilities may have better equipment for fluorescence microscopy than they do for DIC, so capturing IFT using fluorescence microscopy should be considered. The major disadvantages of fluorescence microscopy include the requirement for a fluorescently tagged IFT particle or motor protein subunit and problems associated with photobleaching and phototoxicity. The primary requirements for microscope optics and specimen preparation are the same as described for DIC (above). To minimize photobleaching, first find and orient flagella with observable IFT using DIC. Then switch to fluorescent illumination and record a series of images. Phototoxicity can be a major problem and high levels of UV light can result in GFP-tagged IFT particles slowing down after 30 s. To minimize photobleaching and phototoxic effects, we use an X-cite light (CanadaExfo Photonics Solutions, Mississauga, Ontario, Canada) at an intensity of 75% or less to observe movements of GFP-tagged kinesinassociated protein subunits (Mueller et al., 2005). Using these conditions, IFT can be observed and recorded for up to 1 min. Others have successfully used laser illumination systems to observe movements of GFP-tagged IFT particles (Qin et al., 2007). Use of monochromatic light sources also may reduce phototoxicity. Preliminary reports on the use of TIRF microscopy to study IFT are promising (see Chapter 9 by Engel et al., this volume). Are the same particles observed using DIC and fluorescence microscopy? Comparisons of IFT recorded by fluorescence and DIC microscopy (Fig. 2) indicate that it is likely that the same anterograde particles are observed by both methods. However, the smaller retrograde IFT particles (Dentler, 2005; Iomini et al., 2001; Mueller et al., 2005) are more difficult to resolve using fluorescence microscopy, so many movies and kymographs must be examined to have a sufficient number of particle tracks to confidently assess the retrograde IFT rates using fluorescence microscopy. D. Recording IFT

1. Cameras A variety of cameras are available for microscopy, and each laboratory will likely have their own specific equipment. It is helpful to mount the camera on the microscope so that the camera can be rotated to orient the flagellum vertically on the screen. This reduces the size of the image to be captured, and reduces the memory required for image capture, and allows one to focus on IFT within the flagellum. Our most reliable camera for DIC observations is a Nuvicon Video Camera (NC-70, DAGE-MTI, Michigan City, IN, USA). The camera is sufficiently sensitive to record IFT using halogen lamps (above) and adjustments are adequate to produce highcontrast images. Additional image enhancement can be done by sending the video signal through an Avio Image processor, but to our knowledge, this equipment no

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longer is available. The Nuvicon camera must be used with a frame grabber card such as the Scion FG-7 (www.scioncorp.com), which can be used with Macintosh (OS10) and PC computers and Windows (Microsoft Corporation, Redmond, WA, USA). Digital cameras also can be used. We have had limited experience using an Orca ER (Hammamatsu Phototonics, Hammamatsu City, Japan) and find that it produces excellent images, depending on the capture software and computer buffering capacity. For fluorescence microscopy, a highly sensitive digital camera must be used along with the appropriate light source. The camera must be sufficiently sensitive to detect a fluorescent signal in a flagellum and be fast enough to record particles moving at IFT rates. We have had the best experience with a Rolera MGi EMCCD camera, Q-Imaging, Surry, British Columbia, Canada (www.qimaging.com). Standard GFP filter sets should be sufficient to image GFP-tagged IFT particles, although a Semrock GFP-3035B filter set (www.Semrock.com) provides significantly brighter GFP images than conventional filters. Be aware that the DIC images taken with digital cameras are not as crisp as those taken with the Nuvicon camera, but this is a trade-off between magnification and light intensity that must be made to collect both DIC and fluorescent images from the same camera.

2. Image capture Image capture can be carried out with a variety of cameras and programs. Two methods that we use are described below. We also have successfully used Slidebook software (http://www.intelligent-imaging.com/home.php) and a Hamamatsu Orca ER (www.hamamatsu.com) camera to capture images, but, for DIC, find the Nuvicon cameras to work slightly better than the digital cameras. It is helpful, although not essential, to have a second (live) monitor that can be used to view and focus the live image. The signal from the camera passes through the live monitor and then into the frame grabber in the computer, where the image appears on the computer monitor. a. Scion Image (Scion Corporation, Frederick, MD, USA). The least expensive method, in terms of equipment and software, is to use a Nuvicon video camera, Scion frame grabber, and a Macintosh or PC computer. The computer requirements are modest: most of our images were captured with a Macintosh 7600 computer with 512 MB of RAM and a Scion VG5 frame grabber. A more modern computer with additional memory and newer frame-grabber board will allow capture of larger image stacks. The Nuvicon camera and controller are relatively inexpensive and the Image J program is free (http://rsbweb.nih.gov/ij/). The following steps describe the capture process with the Scion VG5 frame grabber and Scion Image (http://www.scioncorp.com/frames/fr_technical_support.htm). For the newer FG7 frame grabber, software for Windows and Macintosh computers can be downloaded from Scion (http://www.scioncorp.com/frames/fr_technical_support.htm). Install the appropriate plug-ins into Image J before capturing image stacks using Image J.

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1. Place the specimen on the microscope stage, focus the specimen, adjust the condenser focus, and open the field and condenser diaphragms to achieve Koehler illumination. 2. Select cells with straight flagella attached to the coverslips. If possible, select cells that are oriented parallel to the DIC shear. Most of the time, flagella will need to be oriented by rotating the microscope stage to orient the flagellum parallel to the DIC shear. In a Zeiss Axioplan, orient the flagella to point to ~10 o’clock, or ~315° . 3. Open Scion Image and select Special>Camera(Live). 4. Optimize image contrast using the camera controller. 5. Adjust image contrast and the DIC slider to obtain the best image on the computer and on the live monitor. 6. Rotate the camera to orient the image of the flagellum so the flagellum is vertical on the monitor. 7. Select a rectangular area to select the flagellum. 8. Set up to capture a movie. Stacks>Make Movie (movies are simply stacks of images). a. Select the number of frames to be captured. We usually use 600 frames, depending on the amount of memory available. Record the number of frames and the time of recording for measurements. b. Select the frame rate— usually 30 frames/s 9. Press “OK.” The computer will set up a buffer for the frames and will start to collect the images. Some adjustment for focus may need to be made. This is easily done looking at the “live” monitor but can be done viewing the computer image. 10. When frames are captured, save the movie to a file. b. Metamorph (Molecular Devices, Downington, PA, USA). The following steps are for Metamorph Version 7.5.6. Some older versions of Metamorph have a bug in the kymograph program that can lead to errors in the estimation of distance and should be used with caution. 1. Open the program and select Acquire>Acquire>Show Live. 2. Select the Center Quad camera area to increase the camera’s frame rate capability. 3. Select and adjust the EM gain in the Special tab in the Acquire window. A good starting range is 5 MHz, at a value around 2000. 4. Select an exposure time so that the camera’s frame rate is 10–30 frames/s. Do not use binning, or the image quality will be too poor for analysis. 5. Set up to capture a movie. Select Acquire>Stream Acquisition. Then, select the number of frames and hit the acquire button to capture the movie. 6. After the movie is acquired, save the movie.

E. IFT analysis IFT particles rapidly move up and down the flagellum, so it is essential to visually separate anterograde and retrograde particles by preparing kymographs that reveal the location of each particle relative to the time recorded (Iomini et al., 2001). Anterograde particles,

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moving from base to tip are readily distinguished from retrograde particles, which move from the tip to the flagellar base. The size of the particles can be estimated from the thickness of the path. Kymographs are easily generated using Image J or Metamorph (version 7.5.6)

1. Kymograph Preparation a. Image J. 1. Individual kymographs 1. Open the captured movies (image stack) with Image J and rotate flagellar image stacks so that the flagella is vertical (Image>Rotate>Arbitrarily). 2. Use the rectangle tool to make a rectangular selection of the flagellum, cropping as close as possible to the flagellum. 3. Adjust contrast (Image>adjust>brightness/contrast). Set the maximum and minimum sliders to include the areas in the histogram. Click “Apply” to apply the contrast change to all frames in the image stack. 4. Reslice the image (Image>Stacks>Reslice). Select “Start At”: Left and click the Rotate 90 Degrees box. Then select “OK.” A “Reslice” image will open. This may be relatively faint, so the image can be greatly improved using “Z-project.” 5. Move the slider on the bottom of the Reslice window to select the frames that best show the IFT tracks. 6. With the Reslice window active, select Image>Stacks>Z project. Ignore the warning window and click OK. 7. Select start and stop slices. This can be ignored, if the flagellum is closely cropped, or the slices can be selected by observing the slices that show the best image as the slider in the Reslice window is moved. Select Average Intensity and click OK. 8. A kymogram showing anterograde and retrograde transport will appear in a new window, AVG_Reslice of (file name). 9. Save the kymogram as a tiff file (File>Save As …) 2. Kymograph montage To analyze IFT, collect as many movies as possible. Some will show better IFT than others. To compare all tracks in various experiments, a montage combining all kymograms from an experiment can be prepared using Photoshop (Adobe Systems, INC, San Jose, CA, USA). Kymograph images are then sharpened and enhanced to visualize the IFT tracks and to allow comparison among different experiments. 1. Open each kymograph, adjust contrast with “levels” (select Auto), and copy the kymograph to a single Photoshop page. When all kymograms are arranged on the page, flatten the image layers (Layer>Flatten) and crop the image. 2. To enhance the images: a. Select the entire montage and apply Gaussian Blur (Filter>Blur>Gaussian Blur). Select 0.7 pixels and press OK. b. Apply Unsharp Mask (Filter>Sharpen>Unsharp Mask). Select 298%, 2.7 pixels, threshold = 1 and press OK. This will provide a good set of kymograms in which IFT tracks are readily visible. 3. Save the final filtered montage as an uncompressed TIFF file for analysis with Image J.

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b. Metamorph. 1. Open the captured movie with Metamorph. 2. Adjust digital contrast to best see IFT (Display>Adjust Digital Contrast). 3. Set up to make the kymograph (Stack>Kymograph). 4. Draw a straight line using the line tool along the length of the flagellum. Change the line width in the Kymograph window to cover the width of the flagellum. 5. Select which frames to make the Kymograph from. All frames can be used if the “All frames” box is checked. 6. Select Create to make the kymograph. 7. Adjust digital contrast on the kymograph so that the IFT tracks are most clearly seen. 8. Save the kymograph as a tiff file (File>Save).

2. Kymograph Analysis IFT particle rates and frequencies are manually scored by analyzing kymograms with Image J or Metamorph software. For IFT rates, measure the angle of each anterograde (up and to the right) and each retrograde (down and to the right) track using the Image J angle tool or the Metamorph line tool. Save each measurement. Both Image J and Metamorph measurements can be opened in commonly used spreadsheets. Be certain to record the frame rate of the movie by dividing the total number of images (frames) by the recording time. The rate of movement is a function of the tangent of the measured angle of IFT movement and the measured length of the flagellum. The frequency (number of anterograde or retrograde tracks/time) is measured by counting the number of tracks and dividing by the time recorded. Estimation of IFT particle frequencies with the fluorescent images is somewhat more problematic due to potential bleaching of the signal over the time course of image capture. We typically use only the first 30 s of a recording to estimate particle frequencies.

III. Summary Recording and analyzing IFT is relatively simple and can be accomplished using commercially available cameras, computers, and software packages. Here we described two different packages but similar results can be obtained with other software. It is critical to have high-resolution optics, microscopes with high light throughput, and, for DIC, properly aligned the optics and flagellar orientation relative to the DIC shear. Of equal importance is patience. Frequently, one will start recording flagella exhibiting beautiful IFT only to find that the cell becomes camera-shy and glides out of the field of view. With patience and experience, however, IFT recordings are easily made.

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Acknowledgments Studies of IFT in our laboratories have been made possible by support from NIH (P20 RR016475 to WD) and the National Institutes of Health (GM55667 to MEP). Kristyn VanderWaal was also supported in part by a predoctoral fellowship from the American Heart Association, Midwest Affiliate (0715799Z), and a Grantin-Aid (20828) from the University of Minnesota Graduate School to M.E.P.

References Badano, J.L, Mitsuma, N., Beales, P.L., and Katsanis, N. (2006). The ciliopathies: An emerging class of human genetic disorders. Annu. Rev. Genomics Hum. Genet. 7, 125–148. Bloodgood, R.A. (1981). Flagella-dependent gliding motility in Chlamydomonas. Protoplasma 106, 183–192. Dentler, W.L. (2005). Intraflagellar transport (IFT) during assembly and disassembly of Chlamydomonas flagella. J. Cell Biol. 170(4), 649–659. Iomini, C., Babaev-Khaimovm V., Sassaroli, M., and Piperno, G. (2001). Protein particles in Chlamydomonas flagella undergo a transport cycle consisting of four phases. J. Cell Biol. 153(1), 13–24. Kozminski, K.G. (1995). High-resolution imaging of flagella. Methods Cell Biol. 47, 263–271. Kozminski, K.G., Beech, P.L., and Rosenbaum, J.L. (1995). The Chlamydomonas kinesin-like protein FPA10 is involved in motility associated with the flagellar membrane. J. Cell Biol. 131, 1517–1527. Kozminski, K.G., Johnson, K.A., Forscher, P., and Rosenbaum, J.L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523. Marshall, W.F. (2008). The cell biological basis of ciliary disease. J. Cell Biol. 180(1), 17–21. Mueller, J., Perrone, C.A., Bower, R., Cole, D.G., and Porter, M.E. (2005). The FLA3KAP subunit is required for the localization of kinesin-2 to the site of flagellar assembly and processive anterograde intraflagellar transport. Mol. Biol. Cell. 16(3), 1341–1354. Nakamura, S.,Tabino, H., and Kojima, M.K. (1987). Effect of lithium on flagellar length in Chlamydomonas reinhardtii. Cell Struct. Funct. 12, 369–374. Periz, G., Dharia, D., Miller, S.H., and Keller, L.R. (2007). Flagellar elongation and gene expression in Chlamydomonas reinhardtii. Eukaryot. Cell 6(8), 1411–1420. Piperno, G., Siuda, E., Henderson, S., Segil, M., Vaananen, H., and Sassaroli, M. (1998). Distinct mutants of retrograde intraflagellar transport (IFT) share similar morphological and molecular defects. J. Cell Biol. 143(6), 1591–1601. Qin, H., Wang, Z., Diener, D., and Rosenbaum, J. (2007). Intraflagellar transport protein 27 is a small G protein involved in cell-cycle control. Curr. Biol. 17(3), 193–202. Snow, J.J., Ou, G., Gunnarson, A.L., Regina, M., Walker, S., Zhou, H.M., Brust-Mascher, I., and Scholey, J.M. (2004). Two anterograde intraflagellar transport motors cooperate to build sensory cilia on C. elegans neurons. Nat. Cell Biol. 6, 1109–1113.

CHAPTER 9

Total Internal Reflection Fluorescence (TIRF) Microscopy of Chlamydomonas Flagella Benjamin D. Engel*, Karl-Ferdinand Lechtreck†, Tsuyoshi Sakai‡, Mitsuo Ikebe‡, George B. Witman†, and Wallace F. Marshall* * Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, California 94158 †

Department of Cell Biology, University of Massachusetts Medical School, Worcester, Massachusetts 01655

‡

Department of Physiology, University of Massachusetts Medical School, Worcester, Massachusetts 01655

Abstract I. Introduction II. Rationale: A History of Flagellar Microscopy in Chlamydomonas A. Phase Contrast and Dark Field: Early Insights into Flagellar Motilities B. DIC: The Discovery and Initial Characterization of IFT C. Fluorescence: Algae Meet GFP D. TIRF Microscopy: A Perfect Match for Chlamydomonas Flagella III. Materials and Methods: Technical Considerations of Chlamydomonas TIRF A. Slide Preparation and Cell Immobilization B. The Angle of Incidence and Depth of the TIRF Field C. GFP Labeling of Chlamydomonas Flagellar Proteins IV. Discussion: Future Prospects for Chlamydomonas TIRF A. Simultaneous Multicolor TIRF B. FRAP, Photoactivation, and Photoconversion C. Applications Beyond IFT V. Summary Acknowledgments References

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Abstract The eukaryotic flagellum is host to a variety of dynamic behaviors, including flagellar beating, the motility of glycoproteins in the flagellar membrane, and intraflagellar transport (IFT), the bidirectional traffic of protein particles between the flagellar base and tip. IFT is of particular interest, as it plays integral roles in flagellar length control, cell signaling, development, and human disease. However, our ability to understand dynamic flagellar processes such as IFT is limited in large part by the fidelity with which we can image these behaviors in living cells. This chapter introduces the application of total internal reflection fluorescence (TIRF) microscopy to visualize the flagella of Chlamydomonas reinhardtii. The advantages and challenges of TIRF are discussed in comparison to confocal and differential interference contrast techniques. This chapter also reviews current IFT insights gleaned from TIRF microscopy of Chlamydomonas and provides an outlook on the future of the technique, with particular emphasis on combining TIRF with other emerging imaging technologies.

I. Introduction Nearly every cell in the human body projects a primary cilium from its apical surface into the extracellular environment. Once thought to be a vestigial remnant from our days as single-celled organisms, the primary cilium is now appreciated both as a mechanosensory organelle and as a specialized compartment for signaling pathways such as Hedgehog and platelet - derived growth factor (PDGFRaa) (Huangfu et al., 2003; Schneider et al., 2005; Singla and Reiter, 2006). Meanwhile, ciliary defects have been implicated in a wide array of human disorders, from polycystic kidney disease to retinal degeneration and hydrocephalus (Bisgrove and Yost, 2006; Lechtreck et al., 2008; Pazour and Rosenbaum, 2002). For over 40 years, the humble green alga Chlamydomonas reinhardtii has carried the banner of flagellar biology. Chlamydomonas possesses a wealth of flagellar mutants, which have greatly informed our understanding of these organelles. The cell’s two flagella (identical to cilia in every way but their name) are readily purified for biochemical studies (King, 1995) and are amenable to numerous experimental perturbations, including regeneration after amputation (Lefebvre, 1995; Rosenbaum et al., 1969). However, one of this little alga’s greatest contributions to flagellar biology was the discovery and characterization of intraflagellar transport (IFT). IFT is indispensable for the assembly and maintenance of eukaryotic flagella. The only way for axonemal precursors to reach the site of flagellar assembly at the flagellar tip is to be carried there by large IFT particles, powered by the anterograde motor heterotrimeric kinesin-2. At the tip, IFT particles are remodeled and loaded with axonemal turnover products for their return trip to the cell body, driven by the retrograde motor cytoplasmic dynein-1b. The IFT machinery is highly evolutionarily conserved and many Chlamydomonas genes that encode IFT proteins are homologous to human disease genes (Pazour et al., 2000).

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IFT is a live-cell microscopy-defined phenomenon. Though IFT particles (originally dubbed “rafts” and recently renamed “trains”) had been observed in electron micrographs for decades, their function was not understood until IFT was visualized by DIC microscopy (Kozminski et al., 1993). Subsequent biochemistry studies have revealed much about the composition of IFT particles (Cole et al., 1998; Piperno and Mead, 1997), as well as their interactions with specific axonemal precursors (Hou et al., 2007; Qin et al., 2004). In addition, cryoelectron tomography studies have produced insights into the modular three-dimensional architecture of IFT particles (Pigino et al., 2009). However, since IFT is a description of the movement of proteins, live-cell microscopy remains an invaluable technique for understanding this important cellular behavior. In this paper, we briefly review the history of transmitted light and fluorescence microscopy in Chlamydomonas and then detail the promising new application of total internal reflection fluorescence (TIRF) microscopy to studying IFT and other dynamic flagellar processes.

II. Rationale: A History of Flagellar Microscopy in Chlamydomonas A. Phase Contrast and Dark Field: Early Insights into Flagellar Motilities Each new advance in imaging technology has yielded fresh perspectives on the numerous dynamic processes of Chlamydomonas flagella. Phase contrast microscopy was central to many of the early descriptions of Chlamydomonas flagellar behaviors, including the kinetics of flagellar regeneration and shortening under a variety of conditions (Rosenbaum et al., 1969). Decades later, these careful measurements still provide the foundation for investigations into flagellar length control. Phase contrast was also utilized to observe the dynamic bidirectional movements of polystyrene microspheres along the surface of the flagellar membrane (Bloodgood, 1977). This motility was later associated with the movements of flagellar glycoproteins and Chlamydomonas gliding motility, which occurs when the cell’s flagella adhere to a solid surface (Bloodgood, 1995). Dark-field microscopy, which provides enough contrast to visualize the dynamic instability of individual microtubules, was used to elegantly observe the rotation of the central pair microtubules in detergent-treated Chlamydomonas flagella (Kamiya, 1982). However, little was known about the traffic of proteins within eukaryotic flagella until researchers began examining Chlamydomonas with differential interference contrast (DIC) microscopy.

B. DIC: The Discovery and Initial Characterization of IFT IFT was first described by video-enhanced DIC microscopy in 1993 (Kozminski et al., 1993), and for the following decade, this was the only technique available for visualizing IFT in living Chlamydomonas cells. While the molecular identities of the

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IFT proteins were elucidated by biochemistry, these experiments were guided by DIC microscopy of temperature-sensitive mutants. Following identification of the fla10 mutant as a “kinesin-like protein” (Walther et al., 1994; later shown to be part of heterotrimeric kinesin-2 by Cole et al., 1998), DIC microscopy of fla10 at the restrictive temperature revealed that kinesin-2 was required for IFT (Kozminski et al., 1995a). Two groups took advantage of this observation to identify at least 15 IFT proteins that were depleted from fla10 flagella at the restrictive temperature (Cole et al., 1998; Piperno and Mead, 1997). Thus, microscopy steered the biochemical isolation of the IFT proteins, and throughout the subsequent years, DIC mutant analysis continued to shed light on the mechanisms of IFT. For instance, the loss of only retrograde trafficking events in fla15, fla16, and fla17 mutants correlated with the specific depletion of IFT complex A proteins from flagella, implying that complex A is exclusively required for retrograde transport (Piperno et al., 1998). While the wild-type speeds of anterograde and retrograde transport were reported with the initial description of IFT, detailed DIC kymograph analysis of speeds and frequencies enabled numerous fla mutants to be classified as defective in one or more stages of IFT: particle loading at the base, anterograde transport, particle turnaround at the tip, and retrograde transport (Iomini et al., 2001). DIC kymograph analysis has also revealed that the frequency of IFT is relatively constant in shortening, lengthening, and steadystate flagella (Dentler, 2005). Although DIC microscopy was invaluable for the discovery and initial characterization of IFT, the technique suffers from a few major limitations. The quality of IFT visualization is highly contingent on how flagella are oriented in relation to the axis of shear (parallel provides the greatest contrast). DIC quality also varies depending on how flagella are immobilized (certain paralyzed strains work better than others and mounting cells on agarose pads decreases resolution; Kozminski, 1995b). However, the greatest drawback of DIC is the inability to distinguish the traffic of specific proteins. Although combining DIC with temperature-sensitive mutants enabled the correlation of different IFT proteins with distinct IFT behaviors (Iomini et al., 2001; Piperno et al., 1998), these observations are indirect. In order to further tease apart the intricacies of IFT, it is necessary to examine the dynamics of specific IFT proteins and cargos through fluorescence microscopy. C. Fluorescence: Algae Meet GFP The first fluorescence imaging of IFT was achieved in Caenorhabditis elegans in 1999 (Orozco et al., 1999). In the last decade, tremendous work has gone into characterizing IFT in worms and nearly 20 IFT proteins have now been tagged with green fluorescent protein (GFP) (Blacque et al., 2006; Mukhopadhyay et al., 2007; Ou et al., 2007; Snow et al., 2004). By quantifying IFT speeds in several mutants, these worm studies provided insights into the coordination of different IFT proteins and anterograde motors (Ou et al., 2005). Efforts to visualize IFT with GFP in other organisms, including Chlamydomonas (Mueller et al., 2005; Qin et al., 2007), Trypanosoma (Absalon et al., 2008), and mammalian cells (Follit et al., 2006), lagged

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behind for several years but are currently all areas of active research. Due to its GCcodon bias, Chlamydomonas does not easily express many foreign proteins (Heitzer et al., 2007). At the same time that the first fluorescent IFT proteins were being put into worms, Chlamydomonas received a codon-optimized enhanced GFP that fortuitously eliminated a cryptic splice site from the original sequence (Fuhrmann et al., 1999). Six years later, KAP (kinesin-associated protein, the nonmotor subunit of heterotrimeric kinesin-2) became the first GFP-labeled IFT protein in Chlamydomonas (Mueller et al., 2005). Since then, additional IFT proteins have been tagged, including IFT27 (Qin et al., 2007), IFT20 (Lechtreck et al., 2009a), and BBS4 (Lechtreck et al., 2009a). The signaling protein CrPKD2 has also been labeled with GFP, revealing that ~10% of flagellar CrPKD2 undergoes anterograde transport, likely as an IFT cargo (Huang et al., 2007). While fluorescence imaging of IFT in Chlamydomonas has its share of hurdles (difficulties with protein expression, no homologous recombination, cells are autofluorescent and highly motile), there are significant advantages as well. At ~12 µm, wild-type Chlamydomonas flagella are twice as long as C. elegans sensory cilia and most mammalian primary cilia, enabling a more detailed analysis of protein traffic. Furthermore, Chlamydomonas flagella extend away from the cell body, which greatly enhances the signal-to-noise ratio by allowing the flagella to be imaged upon a dark background. In contrast, mammalian primary cilia must be viewed against the backdrop of a GFP-expressing cell. Finally, Chlamydomonas is endowed with a rich catalog of flagellar mutants, such as the kinesin-2 mutant fla10 (Huang et al., 1977; Walther et al., 1994), which can be mated to GFP-tagged strains to observe altered IFT dynamics. D. TIRF Microscopy: A Perfect Match for Chlamydomonas Flagella The imaging advantages of examining GFP-labeled flagellar proteins in Chlamydomonas become significantly pronounced with the addition of TIRF microscopy. While spinning disk confocal microscopy is superior to epifluorescence (less rapid GFP bleaching and better signal-to-noise by omitting out-of-focus light), TIRF provides even greater imaging fidelity at video rate speeds (compare Fig. 2B and C). Instead of passing the excitation laser through the sample like confocal microscopy, with TIRF the angle of incident light is increased until a critical angle is reached where all the light is reflected from the interface between the higher refractive index glass (n = 1.518) and lower refractive index aqueous medium (n = 1.33–1.38). Although no incident light penetrates the sample, it generates an electromagnetic field in the lower refractive index medium, known as the evanescent wave. This TIRF field has the same wavelength as the incident light and decays exponentially as it propagates from the interface, illuminating only the bottom few hundred nanometers of the sample (Axelrod, 2001). Chlamydomonas is an ideal specimen for TIRF microscopy due to its natural gliding behavior. When Chlamydomonas encounters a solid substrate such as the cover glass, it tightly adheres both flagella to the glass, orienting them 180° from each other

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(Bloodgood, 1995, 1977). Since the flagella are roughly 200 nm thick, they fit perfectly within the evanescent field, while the large autofluorescent cell body is excluded from the field of illumination (Fig. 1A). Multiple through-the-objective TIRF systems are commercially available, each equipped with high numerical aperture (NA) objectives that are capable of producing the increased angles of incident illumination required for TIRF (for the images in Fig. 2, we used a Nikon TE2000-E inverted scope equipped with a 100/1.49 NA TIRF objective, the exceptionally useful Nikon Perfect Focus System, and a cooled Photometrics QuantEM:512SC EMCCD camera with quantitative gain). By eliminating light pollution from the rest of the sample, TIRF achieves exceptional signal-to-noise, enabling the clear visualization of low abundance proteins, such as BBS4-GFP, that would otherwise be difficult to detect. We utilized this improved image fidelity to assay the protein content of IFT particles in regenerating Chlamydomonas flagella. With TIRF, we managed to quantify not only the speed and frequency of IFT events, but also the GFP intensity of these trafficking IFT particles (Engel et al., 2009). We found that the KAP-GFP and IFT27-GFP intensities of particles in short flagella were several times brighter than particles in long flagella, suggesting that particles in short flagella are assembled from more IFT proteins. We confirmed this disparity in protein content by counting the stepwise GFP bleaching events of particles in formaldehyde-fixed flagella exposed to TIRF illumination. Because TIRF has exceptional signal detection and slower photobleaching kinetics, many distinct GFP steps can be identified, especially when the bleach intensity plots are enhanced with an edge-preserving filter (Chung and Kennedy, 1991; Leake et al., 2006). Thus, by simply applying TIRF techniques to existing GFP-labeled Chlamydomonas strains, we were able to revise the balance-point model of flagellar length control (proposed in Marshall and Rosenbaum, 2001; Marshall et al., 2005), concluding that IFT particle size scales inversely with flagellar length. A clear advantage of fluorescence microscopy (and TIRF in particular) is the ability to follow the traffic of specific GFP-tagged flagellar proteins. While DIC primarily visualizes large, processive anterograde and retrograde particles (Fig. 2A; Dentler, 2005; Iomini et al., 2001; Kozminski et al., 1993), TIRF microscopy of Chlamydomonas flagella has revealed different classes of IFT behavior. KAP-GFP, IFT27-GFP, and IFT20-GFP all have large anterograde particles that move with the same frequency and speed as the anterograde particles seen in DIC (Fig. 2A–E). (http://www-elsevierdirect. com/Companions/9780123749734, see supplementary movies 1–3). However, while IFT27-GFP has processive retrograde transport similar to particles imaged by DIC (Fig. 2D), IFT20-GFP has slow, thick retrograde traces that frequently pause or change speeds (Fig. 2B and C). KAP-GFP, meanwhile, shows very few retrograde traces at all and has much higher background flagellar fluorescence than IFT27-GFP and IFT20-GFP, particularly toward the flagellar tip (Fig. 2E). Thus, the observation of only three GFPlabeled proteins has already begun to reveal several new intricacies of IFT. All three proteins likely travel out to the flagellar tip together at ~2 µm/s, in large processive anterograde particles that are clearly visible by DIC (Fig. 2F and G). At the tip, it is possible that some proteins, including the kinesin-2 anterograde motor and IFT20, which

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Fig. 1 Chlamy TIRF imaging setup and the effect of axoneme geometry on GFP fluorescence. (A) Imaging of Chlamydomonas via through-the-objective TIRF. Excitation light comes out of the objective at an incident angle that is greater than the critical angle. The light is totally internally reflected, creating an evanescent field on the other side of the interface. Both flagella adhere to the cover glass and are readily imaged, while the cell body remains outside of the TIRF field. (B) Due to the ~200-nm diameter of the axoneme, GFP molecules near the glass interface will be brighter and more likely to photobleach, while GFP proteins on the far side of the axoneme will produce a weaker signal, but are also less likely to bleach. This effect of the flagellar geometry should be considered during both livecell imaging and fixed-cell quantitative photobleaching experiments.

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Fig. 2 Comparison of DIC, spinning disk confocal, and TIRF microscopy, and summary of current IFT observations via TIRF. For all kymographs, the X-axis represents distance (base of flagellum on the left, tip on the right), while the Y-axis is time (running from top to bottom). Vertical scale bars: 2 s; horizontal scale bars: 2 µm. (A) DIC kymograph of KAP-GFP fla3. Anterograde and retrograde traces are processive. Acquired at intermediate resolution with an 0.85 NA dry condenser. For higher quality DIC kymographs of IFT (taken with 1.4 NA oil condensers), see Iomini et al. (2001) and Dentler (2005). (B) Spinning disk kymograph of IFT20-GFP Dift20. Compare fidelity with the TIRF kymograph of IFT20GFP Dift20 in (C). Anterograde is processive, but retrograde has broad, slow traces that pause and change speed. (D) TIRF kymograph of IFT27-GFP in a wild-type background. Anterograde and retrograde traces are processive, similar to DIC. (E) TIRF kymograph of KAP-GFP fla3. Anterograde is processive, but there are very few retrograde traces and the flagellar background is higher than that for IFT20-GFP or IFT27-GFP, especially toward the flagellar tip. Compare to the processive retrograde in the DIC kymograph in part A. (F) A hypothetical anterograde particle. The three GFP-labeled IFT proteins are color-coded to match part G. (G) An illustrated kymograph that summarizes the behaviors of all three proteins seen in parts C, D, and E.

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helps link kinesin-2 to IFT complex B, detach from these large particles. While the core of IFT complex B (including IFT27) undergoes processive retrograde transport in a protein complex large enough to be seen by DIC, IFT20 (perhaps bound to neighboring peripheral proteins such as IFT57) appears to undergo slower and less processive transport in complexes that may be too small for DIC visualization. Meanwhile, it is possible that many kinesin-2 proteins do not partake in retrograde transport at all, but rather remain diffuse in the flagellum. This observation potentially conflicts with the model that kinesin2 is transported out of flagella on retrograde IFT particles powered by cytoplasmic dynein (Signor et al., 1999). It should be noted that the divergent retrograde behaviors of

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IFT20-GFP and KAP-GFP could be the result of interference from the GFP tags. For example, another possible explanation for the retrograde behavior seen in IFT20-GFP is that the GFP tag on IFT20 could block the unloading of kinesin at the flagellar tip, yielding slower, stalling retrograde particles that experience a tug-of-war between kinesin and dynein. However, IFT20-GFP and KAP-GFP both fully rescue endogenous protein mutations and both have anterograde transport that is identical to IFT27-GFP and IFT observed by DIC, suggesting that the GFP-labeled proteins function normally. In addition, all three GFP-tagged proteins have reduced retrograde frequencies compared to frequencies that have been reported by DIC (Dentler, 2005; Iomini et al., 2001). This may be due to detection limits of the TIRF technique, or it may present additional evidence that there are several different species of retrograde particles that cannot be distinguished by DIC. Of course, it is of great interest to label other IFT proteins with GFP to compile a complete description of which proteins share different IFT behaviors.

III. Materials and Methods: Technical Considerations of Chlamydomonas TIRF A. Slide Preparation and Cell Immobilization Preparing GFP-labeled Chlamydomonas cells for observation by TIRF microscopy is straightforward. After growing cells in liquid media, simply plate a small volume (~15 µl) on an untreated cover glass (we use #1.5). Though specially conditioned cover glass may also work, Chlamydomonas cells readily adhere to untreated glass. It is necessary to place a spacer between the slide and cover glass (we use a square ring of petroleum jelly) to prevent cells from being crushed. Focusing on the interface between the cover glass and the media should reveal numerous adherent cells whose flagella are perfectly positioned for TIRF microscopy. While the unique gliding motility of Chlamydomonas cells makes TIRF imaging possible, it also poses one of the greatest obstacles to the technique. Although cells often remain in a stationary gliding position for up to a minute, which is ample time to analyze IFT, cells are quite mobile and will either glide out of the field of view or detach from the cover glass altogether. This is further complicated by the phototactic and photophobic behaviors of Chlamydomonas. The mere act of observing the cells with TIRF illumination is enough to either encourage cells to flee the field of view or cluster tightly together on the cover glass, depending on the strain. Mounting cells on an agarose pad is an effective tool for immobilizing Chlamydomonas cells for DIC or confocal microscopy. However, agarose pads rarely immobilize cells in a gliding position. Instead, the autofluorescent cell bodies are pressed against the cover glass (and into the TIRF field), while the flagella do not evenly adhere to the glass. Coating the cover glass with polylysine has also proved problematic for immobilizing gliding cells. At 0.1 mg/ml, polylysine slightly decreases cell motility. At increased concentrations (up to 10 mg/ml), gliding motility nearly stops, but the tips of adherent flagella begin to curl up and cells deflagellate, leaving their flagella behind on the cover glass (Kozminski, 1995b). Additionally, because polylysine has a

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higher refractive index than aqueous media, it can interfere with the propagation of the evanescent wave and cause light scattering and increased background signal (George, 2008). When considering cover glass coatings, it is also important to ensure that the layers are thin so they will not occupy a significant portion of the TIRF field. It is possible that a more specific coating, such as antibodies to the flagellar membrane glycoprotein FMG1 (Bloodgood et al., 1986), might prove more effective. However, even if we could completely immobilize the flagella to a substrate, Chlamydomonas may simply shed their flagella as seen with polylysine. However, it may be possible to prevent deflagellation by utilizing flagellar autonomy ( fa) mutants (Finst et al., 1998; Lewin and Burrascano, 1983). (Finst et al., 1998; Lewin and Burrascano, 1983). We have had greater success by crossing GFP strains to mutants with paralyzed flagella (such as the central-pair mutant pf18). While these cells tend to remain adhered to one spot for significantly longer time periods than wild-type cells, ultimately, they too glide away or detach. There are additional Chlamydomonas mutants that may prove useful for immobilizing cells in the gliding position. ptx mutants are defective in phototaxis, and thus may not react to the excitation light (Horst and Witman, 1993). Concanavalin A has been reported to interfere with gliding motility in the L-23 pf18 mutant (Bloodgood and Salomonsky, 1989). There are also a large number of gli mutants that were isolated by screening for gliding defects (Kozminski, 1995c), though the identities of these genes remain unknown, which complicates potential crosses to GFP strains. It is also important to note that mutant backgrounds may introduce unexpected variables (e.g., pf18 cells regenerate their flagella with slowed kinetics), and thus should always be compared to wild-type cells. Finally, gliding motility is reportedly inhibited by lowering the free calcium concentration with EGTA and adding 100 mM NaCl (Bloodgood, 1995; Kozminski et al., 1993). However, these conditions also induce flagellar resorption through unknown mechanisms (Lefebvre et al., 1978), and consequently are less useful for observing flagella under steady-state length conditions. The immobilization of adherent Chlamydomonas cells during TIRF microscopy remains an unresolved issue, and until a solution is found, confocal and DIC microscopy will maintain an advantage for the prolonged observation of single cells. B. The Angle of Incidence and Depth of the TIRF Field The depth of the evanescent field varies greatly as a function of the excitation laser’s angle of incidence. Greater angles create shallower fields (Mattheyses and Axelrod, 2006), so for most flagellar imaging applications, the incident angle should be reduced to near the critical angle to ensure the deepest possible TIRF field. By adjusting the angle of incidence very close to the critical angle, we have measured TIRF fields on our microscope that are 250–300 nm deep (Engel et al., 2009). Even though this is enough to image completely through Chlamydomonas flagella, it is very important to note that TIRF illumination is not constant, but rather falls off exponentially as a function of distance from the cover glass (Axelrod, 2001). This property of the evanescent field should always be considered when interpreting results. For example, when comparing IFT particles within a flagellum, it is impossible to distinguish whether differences in

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intensity are due to unequal protein content or different positions of the particles on the axoneme (Fig. 1B). However, it is legitimate to compare the average intensity of IFT particles between different flagella (as shown in Engel et al., 2009). The relationship between axoneme geometry and the TIRF field also increases the variability of GFP step sizes during quantitative photobleaching of fixed cells. GFP molecules that are further from the cover glass produce less intensity and are also less likely to bleach (Fig. 1B). Thus, although many GFP step sizes are similar, it is not uncommon to see a few smaller steps toward the end of the bleach. Since even slight variations in the incident angle significantly impact GFP intensity and the depth of the evanescent field, motorized control of the laser trajectory is quite useful, especially when trying to be consistent between imaging sessions. Other parameters, such as laser power and camera settings should also be kept constant when attempting to compare intensities between images. Many new electron multiplying charge-coupled device (EMCCD) cameras are equipped with linear gain, which increases the reliability of comparing intensities between images. However, gain decreases as EMCCD cameras age (Ingley et al., 2008), so imaging sessions should be performed as close together as possible. C. GFP Labeling of Chlamydomonas Flagellar Proteins Fluorescence detection is greatly increased if every copy of a specific flagellar protein is labeled with GFP. Furthermore, complete labeling enables quantification of protein abundance via intensity measurements and quantitative photobleaching (Engel et al., 2009). However, since targeted gene deletion and homologous recombination are not simple feats in Chlamydomonas, the best candidates for GFP labeling are IFT genes that already have identified Chlamydomonas mutations. These mutants can then be rescued with genes encoding GFP-tagged versions of the proteins. Rescuing IFT mutants provides the additional advantage of demonstrating that the exogenous GFP-labeled protein is functioning correctly. Moreover, when tagged proteins are expressed in the presence of the untagged endogenous protein, the latter may outcompete the former for interaction with a limited number of binding partners (Lechtreck et al., 2009b). The existing GFP-tagged IFT20 and BBS4 strains are in null mutant backgrounds, where all endogenous flagellar protein has been replaced with the labeled version (Fig. 3, Lechtreck et al., 2009a). KAP-GFP was expressed in the temperature-sensitive KAP mutant, fla3 (Mueller et al., 2005). However, at all temperatures, the flagellar KAP population is almost completely labeled with GFP (minute levels of the endogenous mutant protein may remain, see Fig. 3). IFT27-GFP, on the other hand, was expressed in wild-type cells. As a result, only half of the flagellar IFT27 protein is GFP-labeled, yielding a much weaker fluorescent signal (compare Fig. 2D with C and E; Qin et al., 2007). Two distinct cloning strategies have been successfully employed for GFP-labeling in Chlamydomonas: (1) insertion of the tag sequence into an exon near the 30 end of the target gene (KAP-GFP) and (2) fusion of fluorescent proteins to the C-terminus of the target proteins (IFT27-GFP, CrPKD2-GFP, IFT20-GFP, IFT20-mCherry, BBS4-GFP). Proteins can be expressed either from vectors containing genomic sequences including the endogenous promoters (KAP-GFP, IFT27-GFP, CrPKD2-GFP) or from vectors using the

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HSP70B/rbcS or FLA14 promoters, the latter of which was used to express cDNAs of IFT20 and BBS4 (Lechtreck et al., 2009a). Because the FLA14 gene does not contain introns, its promoter efficiently expresses intronless cDNA. Furthermore, since FLA14 encodes the flagellar dynein light chain LC8 (Pazour et al., 1998), tagged flagellar proteins

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(such as IFT20-GFP) are expressed by the FLA14 promoter at relatively wild-type levels and have equal flagellar abundance to endogenous protein in wild-type cells (Fig. 3). Chlamydomonas cells can be transformed with linearized plasmids using the glassbead method (Kindle, 1990) or electroporation (Shimogawara et al., 1998). Screening transformants for rescue of a selectable phenotype enables thousands of transformants to be screened and strains expressing fusion proteins to be easily identified. IFT20GFP, for example, was expressed in an ift20 deletion mutant, which is immotile due to the lack of flagella. Transformants were selected for restoration of motility, and expression of the GFP-fusion protein was then verified. If phenotypic selection of cells is not possible, cells expressing the fusion protein may be identified by fluorescence microscopy and western blotting. In our experience, fluorescence-activated cell sorting (FACS) identification of GFP-expressing cells is difficult because the strong autofluorescence of the cell body masks GFP emission, but it may be possible to apply this approach to “white” Chlamydomonas mutants (McCarthy et al., 2004).

IV. Discussion: Future Prospects for Chlamydomonas TIRF A. Simultaneous Multicolor TIRF Simultaneously observing two or more proteins tagged with different color fluorophores holds great promise for understanding the colocalization of multiple IFT proteins and cargos as they transit through the flagellum. Considering that our initial observations appear to have revealed different classes of retrograde behaviors (Fig. 2), it will be of great interest to visualize which proteins traffic together. In principle, there are few limitations to labeling numerous flagellar proteins with different fluorophores. Initially, Chlamydomonas IFT proteins had only been tagged with codon-optimized GFP because of concerns about protein expression. However, IFT20 fused to a synthetic mCherry sequence optimized for human codon bias was successfully expressed (Fig. 4; Lechtreck et al., 2009a), opening the door to twocolor experiments. IFT20-mCherry was observed to be less photostable and half as bright as IFT20-GFP. Initial observations of cells expressing both IFT20-mCherry and BBS4-GFP revealed that BBS4 is carried by only a subset of anterograde particles (Fig. 4) (http//www.elsevierdirect.com/Companions/9780123749734, See Suplementary movie 4). Yellow fluorescent protein (YFP) and cyan fluorescent protein (CFP) variants of codon-optimized GFP have also been expressed in Chlamydomonas (Feldman, 2008), providing additional avenues for multicolor imaging. Simultaneous multicolor TIRF presents several unique hurdles compared to singlechannel imaging. In order to cleanly image two colors simultaneously, several specialized pieces of equipment are required, including a multiline laser launch, a multipass dichroic mirror that directs both excitation wavelengths to the sample, and an emission splitter (such as Photometrics Dual-View or Cairn OptoSplit) that separates each emission wavelength to opposite halves of a single camera or to two different cameras. (The movie in Fig. 4 was acquired with an Olympus I71 Inverted Microscope

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Fig. 4 Simultaneous two-color TIRF of IFT20-mCherry and BBS4-GFP. For all kymographs, the X-axis represents distance (base of flagellum on the left, tip on the right), while the Y-axis is time. Vertical scale bar: 2 s; horizontal scale bar: 2 µm. BBS4-GFP is much lower in abundance than IFT20-mCherry, and is only localized to a subset of the IFT20-mCherry particles. In the merged image, BBS4-GFP is green and IFT20mCherry is red. (See Plate no. 7 in the Color Plate Section.)

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equipped with an Olympus 60/1.4 NA PlanApo objective, a Semrock FF498/581 multipass beam splitter with laser-line filters to clean the two colors of excitation light, a custom-built emission splitter system with a Semrock FF562-Di01 dichroic mirror and Semrock 525/50- and 630/69-nm emission filters that block both excitation wavelengths, and an Andor iXon DV860 back-illuminated EMCCD camera). Most commercially available TIRF systems use a single fiber-optic cable to deliver multiple laser wavelengths to the sample. These systems can only introduce light at one incident angle and at one focus setting, which creates two complications for multicolor imaging. The first issue to consider is the effect of excitation color on laser focusing. Because different wavelengths of light are refracted at different angles when passing through the objective, they do not share the same focal point. Unless each laser is independently focused, the beam will be slightly less collimated, leading to increased light scattering and background signal (George, 2008). The second issue to consider is the effect of laser color on the depth of the evanescent field. The depth of field penetration depends on the refractive indices of the cover glass (n = 1.518) and the aqueous medium (n = 1.33–1.38), in addition to both the angle and wavelength of the incident illumination (Axelrod, 2001; George, 2008). Since the refractive indices of the imaging setup are relatively fixed, the angle of incidence must be adjusted for different excitation wavelengths to maintain the same depth and intensity of TIRF illumination. Even with motorized control of laser trajectory, tuning the incident angle when switching between excitation colors introduces too great a lag for the simultaneous imaging of fast-moving IFT particles. While neither of these wavelength-related effects is severe, simultaneous multicolor TIRF would benefit from an imaging setup with separate light paths for each laser, each with its own independent angle and focus adjustments. B. FRAP, Photoactivation, and Photoconversion Several advanced photobleaching and photoactivation techniques can be combined with TIRF to shed light on additional aspects of IFT. Measuring the kinetics of FRAP (fluorescence recovery after photobleaching) may yield insights into how readily different IFT proteins in the flagella exchange with the cytoplasm. Tagging IFT proteins with photoactivatable or photoconvertable fluorophores would provide a reciprocal approach to this question. Photoactivation in the cytoplasm would facilitate the observation of protein exchange into the flagella. Because Chlamydomonas is biflagellate, photoactivating within one flagellum would also be highly informative. Protein exchange could be judged not only by loss of fluorescent IFT particles within the photoactivated flagellum, but also by the appearance of fluorescence in the dark flagellum. Finally, photoactivation and photoconversion would provide useful tools for understanding the dynamics of IFT turnaround at the flagellar tip. By only activating a small subset of the IFT particles in a flagellum, it would become clear whether particles immediately undergo retrograde transport when they reach the tip, and whether retrograde KAP (Fig. 2E) truly is diffuse or whether it undergoes an active transport mechanism that is undetectable in fully labeled KAP-GFP flagella.

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One potential obstacle to these promising experiments is the expression of photoactivatible and photoconvertible proteins in Chlamydomonas. The design of photoactivatible GFP (PA-GFP) should only require the straightforward task of making a few simple base pair changes to the codon-optimized GFP sequence (following the scheme outlined by Patterson and Lippincott-Schwartz, 2002). The expression of more exotic fluorophores, such as the green-to-red photoconvertable proteins Kaede (Ando et al., 2002), EosFP (Wiedenmann et al., 2004), and Dendra2 (Adam et al., 2009; Labas et al., 2002), and the reversibly photoswitchable Dronpa (Habuchi et al., 2005) may require codon optimization. However, it is possible that, like mCherry, these proteins will express without alterations to their sequence. The greatest challenge to performing FRAP or photoactivation in TIRF will probably be the immobilization of Chlamydomonas cells. As previously discussed, even paralyzed strains glide out of the field of view or detach from the cover glass within minutes. It is likely that Chlamydomonas will have a strong photophobic response to the intense laser energy required for FRAP. Furthermore, the phototropin blue-light receptors localized in Chlamydomonas flagella (Huang et al., 2004) will certainly be stimulated by the 405 nm light used to activate PA-GFP, though the function of these photoreceptors is unknown. It is possible that a combination of the strategies discussed in Section III.A (such as ptx mutants and calcium depletion) will immobilize adherent cells long enough to facilitate these powerful proteintracking techniques. For some questions, such as protein exchange between the flagellar and cytoplasmic pools, confocal microscopy should suffice, allowing immobilization on agarose pads. C. Applications Beyond IFT TIRF should prove equally beneficial to imaging other Chlamydomonas flagellar proteins that are not continuously trafficked by IFT. This includes monitoring the dynamics of membrane-bound signaling proteins, such as CrPKD2-GFP (Huang et al., 2007) and tracking changes in flagellar membrane glycoprotein distribution (Bloodgood et al., 1986). FRAP is not limited to studying the exchange of IFT particles, but can also be used to examine the turnover of membrane proteins and integrated axonemal proteins, such as dynein arms (Watanabe et al., 2004). Based on the incorporation of HA-tubulin in quadraflagellates, it had been concluded that axonemal tubulin does not treadmill (Marshall and Rosenbaum, 2001). However, this hypothesis could be tested more elegantly in live cells by either speckling axonemal tubulin with GFP (Waterman-Storer and Danuser, 2002) or by bleaching a fiducial mark on a fluorescent axoneme and monitoring this mark for movement. In another novel use of the technique, TIRF was recently combined with calcium-responsive dyes to measure transient spikes in flagellar calcium concentration. These transient calcium spikes rapidly increased in frequency prior to deflagellation (Wheeler et al., 2008). Clearly, TIRF microscopy of Chlamydomonas flagella holds great promise for a range of applications beyond the study of IFT.

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V. Summary TIRF microscopy is a versatile new method for observing fluorescence in Chlamydomonas flagella. While there are still a few technical hurdles, such as protein expression and cell immobilization, TIRF provides greatly improved fidelity for imaging flagellar proteins in live cells. It has already provided some glimpses of new IFT behaviors (Fig. 2), but a wealth of untapped potential remains. Substantial inquiries into the mechanisms of IFT, as well as a full gamut of other flagellar biology questions, are waiting to be explored through additional protein labeling and the incorporation of multicolor, FRAP, and photoactivation techniques. Perhaps in the more distant future, it may even be possible to combine TIRF of Chlamydomonas flagella with super resolution technologies such as photoactivated localization microscopy (PALM)/stochastic optical reconstruction microscopy (STORM) (Betzig et al., 2006; Rust et al., 2006) and Structured Illumination (Gustafsson, 2000).

Acknowledgments We thank Kurt Thorn and the Nikon Imaging Center at UC San Francisco for invaluable microscopy resources and assistance. We also thank Joshua Mueller, Mary Porter, Hongmin Qin, and Joel Rosenbaum for generously sharing GFP-labeled strains, as well as Kurt Thorn, Arthur Millius, Sarah Goodwin, Jennifer Blake, and the Marshall Lab for helpful discussions and careful reading of the manuscript. This work was supported by the W. M. Keck Foundation Distinguished Young Scholars Program (W. F. Marshall), the Searle Scholar Program (W. F. Marshall), the Genentech Graduate Fellowship (B. D. Engel), National Institutes of Health Grant GM 030626 (G. B. Witman), and the Robert W. Booth Fund at the Greater Worcester Community Foundation (G. B. Witman).

References Absalon, S., Blisnick, T., Kohl, L., Toutirais, G., Doré, G., Julkowska, D., Tavenet, A., and Bastin, P. (2008). Intraflagellar transport and functional analysis of genes required for flagellum formation in trypanosomes. Mol. Biol. Cell 19, 929–944. Adam, V., Nienhaus, K., Bourgeois, D., and Nienhaus, G.U. (2009). Structural basis of enhanced photoconversion yield in GFP-like protein Dendra2. Biochemistry 48, 4905–4815. Ando, R., Hama, H., Yamamoto-Hino, M., Mizuno, H., and Miyawaki, A. (2002). An optical marker based on the UV-induced green-to-red photoconversion of a fluorescent protein. Proc. Natl. Acad. Sci. USA 99, 12651–12656. Axelrod, D. (2001). Total internal reflection fluorescence microscopy in cell biology. Traffic 2, 764–774. Betzig, E., Patterson, G.H., Sougrat, R., Lindwasser, O.W., Olenych, S., Bonifacino, J.S., Davidson, M.W., Lippincott-Schwartz, J., and Hess, H.F. (2006). Imaging intracellular fluorescent proteins at nanometer resolution. Science 313, 1642–1645. Bisgrove, B.W., and Yost, H.J. (2006). The roles of cilia in developmental disorders and disease. Development 133, 4131–4143. Blacque, O.E., Li, C., Inglis, P.N., Esmail, M.A., Ou, G., Mah, A.K., Baillie, D.L., Scholey, J.M., and Leroux, M.R. (2006). The WD repeat-containing protein IFTA-1 is required for retrograde intraflagellar transport. Mol. Biol. Cell 17, 5053–5062. Bloodgood, R.A. (1977). Motility occurring in association with the surface of the Chlamydomonas flagellum. J. Cell Biol. 75, 983–989. Bloodgood, R.A. (1995). Flagellar surface motility: Gliding and microsphere movements. In “Methods in Cell Biology” (W. Dentler and G.B. Witman, eds.), Vol. 47, pp. 273–279. Elsevier Inc., San Diego.

9. TIRF Microscopy of Chlamydomonas Flagella

175

Bloodgood, R.A., and Salomonsky, N.L. (1989). Use of a novel Chlamydomonas mutant to demonstrate that flagellar glycoprotein movements are necessary for the expression of gliding motility. Cell Motil. Cytoskeleton 13, 1–8. Bloodgood, R.A., Woodward, M.P., and Salomonsky, N.L. (1986). Redistribution and shedding of flagellar membrane glycoproteins visualized using an anti-carbohydrate monoclonal antibody and concanavalin A. J. Cell Biol. 102, 1797–1812. Chung, S.H., and Kennedy, R.A. (1991). Forward-backward non-linear filtering technique for extracting small biological signals from noise. J. Neurosci. Methods 40, 71–86. Cole, D.G., Diener, D.R., Himelblau, A.L., Beech, P.L., Fuster, J.C., and Rosenbaum, J.L. (1998). Chlamydomonas kinesin-II-dependent intraflagellar transport (IFT): IFT particles contain proteins required for ciliary assembly in Caenorhabditis elegans sensory neurons. J. Cell Biol. 141, 993–1008. Dentler, W. (2005). Intraflagellar transport (IFT) during assembly and disassembly of Chlamydomonas flagella. J. Cell Biol. 170, 649–659. Engel, B.D., Ludington, W.B., and Marshall, W.F. (2009). Intraflagellar transport particle size scales inversely with flagellar length: Revisiting the balance-point length control model. J. Cell Biol., 187, 81–89. Feldman, J.L. (2008). Deconstructing cell architecture: Exploring centriole structure, function, and positioning in the green alga Chlamydomonas reinhardtii. Ph.D. thesis, University of California, San Francisco. Finst, R.J., Kim, P.J., and Quarmby, L.M. (1998). Genetics of the deflagellation pathway in Chlamydomonas. Genetics 149, 927–936 Follit, J.A., Tuft, R.A., Fogarty, K.E., and Pazour, G.J. (2006). The intraflagellar transport protein IFT20 is associated with the Golgi complex and is required for cilia assembly. Mol. Biol. Cell 17, 3781–3792. Fuhrmann, M., Oertel, W., and Hegemann, P. (1999). A synthetic gene coding for the green fluorescent protein (GFP) is a versatile reporter in Chlamydomonas reinhardtii. Plant J. 19, 353–361. George, N.B. (2008). Overcoming optical challenges to live-cell TIRF microscopy. Biophotonics 15, 30–33. Gustafsson, M.G. (2000). Surpassing the lateral resolution limit by a factor of two using structured illumination microscopy. J. Microsc. 198, 82–87. Habuchi, S., Ando, R., Dedecker, P., Verheijen, W., Mizuno, H., Miyawaki, A., and Hofkens, J. (2005). Reversible single-molecule photoswitching in the GFP-like fluorescent protein Dronpa. Proc. Natl. Acad. Sci. USA 102, 9511–9516. Heitzer, M., Eckert, A., Fuhrmann, M., and Griesbeck, C. (2007). Influence of codon bias on the expression of foreign genes in microalgae. Adv. Exp. Med. Biol. 616, 46–53. Horst, C.J., and Witman, G.B. (1993). ptx1, a nonphototactic mutant of Chlamydomonas, lacks control of flagellar dominance. J. Cell Biol. 120, 733–741. Hou, Y., Qin, H., Follit, J.A., Pazour, G.J., Rosenbaum, J.L., and Witman, G.B. (2007). Functional analysis of an individual IFT protein: IFT46 is required for transport of outer dynein arms into flagella. J. Cell Biol. 176, 653–665. Huang, K., Diener, D.R., Mitchell, A., Pazour, G.J., Witman, G.B., and Rosenbaum, J.L. (2007). Function and dynamics of PKD2 in Chlamydomonas reinhardtii flagella. J. Cell Biol. 179, 501–514. Huang, K., Kunkel, T., and Beck, C.F. (2004). Localization of the blue-light receptor phototropin to the flagella of the green alga Chlamydomonas reinhardtii. Mol. Biol. Cell 15, 3605–3614. Huang, B., Rifkin, M.R., and Luck, D.J. (1977). Temperature-sensitive mutations affecting flagellar assembly and function in Chlamydomonas reinhardtii. J. Cell Biol. 72, 67–85. Huangfu, D., Liu, A., Rakeman, A.S., Murcia, N.S., Niswander, L., and Anderson, K.V. (2003). Hedgehog signalling in the mouse requires intraflagellar transport proteins. Nature 426, 83–87. Ingley, R., Smith, D.R., and Holland, A.D. (2008). Life testing of EMCCD gain characteristics. Nucl. Instrum. Methods Phys. Res., Sect. A 600, 460–465. Iomini, C., Babaev-Khaimov, V., Sassaroli, M., and Piperno, G. (2001). Protein particles in Chlamydomonas flagella undergo a transport cycle consisting of four phases. J. Cell Biol. 153, 13–24. Kamiya, R. (1982). Extrusion and rotation of the central-pair microtubules in detergent-treated Chlamydomonas flagella. Prog. Clin. Biol. Res. 80, 169–173. Kindle, K.L. (1990). High-frequency nuclear transformation of Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA 87, 1228–1232.

176

Benjamin D. Engel et al. King, S.M. (1995). Large-scale isolation of Chlamydomonas flagella. In “Methods in Cell Biology” (W. Dentler and G.B. Witman, eds.), Vol. 47, pp. 9–12. Elsevier Inc., San Diego. Kozminski, K.G. (1995c). Beat-independent flagellar motilities in Chlamydomonas and an analysis of the function of alpha-tubulin acetylation. Ph.D. Thesis. Yale University. Kozminski, K.G. (1995b). High-resolution imaging of flagella. In “Methods in Cell Biology” (W. Dentler and G.B. Witman, eds.), Vol. 47, pp. 263–271. Elsevier Inc., San Diego. Kozminski, K.G., Beech, P.L., and Rosenbaum, J.L. (1995a). The Chlamydomonas kinesin-like protein FLA10 is involved in motility associated with the flagellar membrane. J. Cell Biol. 131, 1517–1527. Kozminski, K.G., Johnson, K.A., Forscher, P., and Rosenbaum, J.L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523. Labas, Y.A., Gurskaya, N.G., Yanushevich, Y.G., Fradkov, A.F., Lukyanov, K.A., Lukyanov, S.A., and Matz, M.V. (2002). Diversity and evolution of the green fluorescent protein family. Proc. Natl. Acad. Sci. USA 99, 4256–4261. Leake, M.C., Chandler, J.H., Wadhams, G.H., Bai, F., Berry, R.M., and Armitage, J.P. (2006). Stoichiometry and turnover in single, functioning membrane protein complexes. Nature 443, 355–358. Lechtreck, K.F., Delmotte, P., Robinson, M.L., Sanderson, M.J., and Witman, G.B. (2008). Mutations in Hydin impair ciliary motility in mice. J. Cell Biol. 180, 633–643. Lechtreck, K.F., Johnson, E.C., Sakai, T., Cochran, D, Ballif, B.A., Rush, J., Pazour, G.J., Ikebe, M., and Witman, G.B. (2009a). The Chlamydomonas BBSome is an IFT cargo required for export of specific signaling proteins from flagella. (in preparation). Lechtreck, K.F., Luro, S., Awata, J., and Witman, G.B. (2009b). HA-tagging of putative flagellar proteins in Chlamydomonas reinhardtii identifies a novel protein of intraflagellar transport complex B. Cell Motil. Cytoskeleton. 66, 469–482. Lefebvre, P.A. (1995). Flagellar amputation and regeneration in Chlamydomonas. In “Methods in Cell Biology” (W. Dentler and G.B. Witman, eds.), Vol. 47, pp. 3–7. Elsevier Inc., San Diego. Lefebvre, P.A., Nordstrom, S.A., Moulder, J.E., and Rosenbaum, J.L. (1978). Flagellar elongation and shortening in Chlamydomonas. IV. Effects of flagellar detachment, regeneration, and resorption on the induction of flagellar protein synthesis. J. Cell Biol. 78, 8–27. Lewin, R.A., and Burrascano, C. (1983). Another new kind of Chlamydomonas mutant, with impaired flagellar autotomy. Experientia 39, 1397–1398. Marshall, W.F., Qin, H., Rodrigo Brenni, M., and Rosenbaum, J.L. (2005). Flagellar length control system: Testing a simple model based on intraflagellar transport and turnover. Mol. Biol. Cell 16, 270–278. Marshall, W.F., and Rosenbaum, J.L. (2001). Intraflagellar transport balances continuous turnover of outer doublet microtubules: Implications for flagellar length control. J. Cell Biol. 155, 405–414. Mattheyses, A.L., and Axelrod, D. (2006). Direct measurement of the evanescent field profile produced by objective-based total internal reflection fluorescence. J. Biomed. Opt. 11, 014006. McCarthy, S.S., Kobayashi, M.C., and Niyogi, K.K. (2004). White mutants of Chlamydomonas reinhardtii are defective in phytoene synthase. Genetics 168, 1249–1257. Mueller, J., Perrone, C.A., Bower, R., Cole, D.G., and Porter, M.E. (2005). The FLA3 KAP subunit is required for localization of kinesin-2 to the site of flagellar assembly and processive anterograde intraflagellar transport. Mol. Biol. Cell 16, 1341–1354. Mukhopadhyay, S., Lu, Y., Qin, H., Lanjuin, A., Shaham, S., and Sengupta, P. (2007). Distinct IFT mechanisms contribute to the generation of ciliary structural diversity in C. elegans. EMBO J. 26, 2966–2980. Orozco, J.T., Wedaman, K.P., Signor, D., Brown, H., Rose, L., and Scholey, J.M. (1999). Movement of motor and cargo along cilia. Nature 398, 674. Ou, G., Blacque, O.E., Snow, J.J., Leroux, M.R., and Scholey, J.M. (2005). Functional coordination of intraflagellar transport motors. Nature 436, 583–587. Ou, G., Koga, M., Blacque, O.E., Murayama, T., Ohshima, Y., Schafer J.C., Li, C., Yoder, B.K., Leroux, M.R., and Scholey, J.M., (2007). Sensory ciliogenesis in Caenorhabditis elegans: Assignment of IFT components into distinct modules based on transport and phenotypic profiles. Mol. Biol. Cell 18, 1554–1569.

9. TIRF Microscopy of Chlamydomonas Flagella

177

Patterson, G.H., and Lippincott-Schwartz, J. (2002). A photoactivatable GFP for selective photolabeling of proteins and cells. Science 297, 1873–1877. Pazour, G.J., Dickert, B.L., Vucica, Y., Seeley, E.S., Rosenbaum, J.L., Witman, G.B., and Cole, D.G. (2000). Chlamydomonas IFT88 and its mouse homologue, polycystic kidney disease gene tg737, are required for assembly of cilia and flagella. J. Cell Biol. 151, 709–718. Pazour, G.J., and Rosenbaum, J.L. (2002). Intraflagellar transport and cilia-dependent diseases. Trends Cell Biol. 12, 551–555. Pazour, G.J., Wilkerson, C.G., and Witman, G.B., (1998). A dynein light chain is essential for the retrograde particle movement of intraflagellar transport (IFT). J. Cell Biol. 141, 979–992. Pigino, G., Geimer, S., Lanzavecchia, S., Paccagnini, E., Cantele, F., Diener, D.R., Rosenbaum, J.L., and Lupetti, P. (2009). Electron tomographic analysis of intraflagellar transport complexes in situ. J. Cell Biol. 187, 135–148. Piperno, G., and Mead, K. (1997). Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella. Proc. Natl. Acad. Sci. USA 94, 4457–4462. Piperno, G., Siuda, E., Henderson, S., Segil, M., Vaananen, H., and Sassaroli, M. (1998). Distinct mutants of retrograde intraflagellar transport (IFT) share similar morphological and molecular defects. J. Cell Biol. 143, 1591–1601. Qin, H., Diener, D.R., Geimer, S., Cole, D.G., and Rosenbaum, J.L. (2004). Intraflagellar transport (IFT) cargo: IFT transports flagellar precursors to the tip and turnover products to the cell body. J. Cell Biol. 164, 255–266. Qin, H., Wang, Z., Diener, D., and Rosenbaum, J.L. (2007). Intraflagellar transport protein 27 is a small G protein involved in cell-cycle control. Curr. Biol. 17, 193–202. Rosenbaum, J.L., Moulder, J.E., and Ringo, D.L. (1969). Flagellar elongation and shortening in Chlamydomonas. The use of cycloheximide and colchicine to study the synthesis and assembly of flagellar proteins. J. Cell Biol. 41, 600–619. Rust, M.J., Bates, M., and Zhuang, X. (2006). Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat. Methods 3, 793–795. Schneider, L., Clement, C.A., Teilmann, S.C., Pazour, G.J., Hoffmann, E.K., Satir, P., and Christensen, S.T. (2005). PDGFRaa signaling is regulated through the primary cilium in fibroblasts. Curr Biol. 15, 1861–1866. Shimogawara, K., Fujiwara, S., Grossman, A., and Usuda, H. (1998). High-efficiency transformation of Chlamydomonas reinhardtii by electroporation. Genetics 148, 1821–1828. Signor, D., Wedaman, K.P., Orozco, J.T., Dwyer, N.D., Bargmann, C.I., Rose, L.S., and Scholey, J.M. (1999). Role of a class DHC1b dynein in retrograde transport of IFT motors and IFT raft particles along cilia, but not dendrites, in chemosensory neurons of living Caenorhabditis elegans. J. Cell Biol. 147, 519–530. Singla, V., and Reiter, J.F. (2006). The primary cilium as the cell’s antenna: Signaling at a sensory organelle. Science 313, 629–633. Snow, J.J., Ou, G., Gunnarson, A.L., Walker, M.R., Zhou, H.M., Brust-Mascher, I., and Scholey, J.M. (2004). Two anterograde intraflagellar transport motors cooperate to build sensory cilia on C. elegans neurons. Nat. Cell Biol. 6, 1109–1113. Walther, Z., Vashishtha, M., and Hall, J.L. (1994). The Chlamydomonas FLA10 gene encodes a novel kinesin-homologous protein. J. Cell Biol. 126, 175–188. Watanabe, Y., Hayashi, M., Yagi, T., and Kamiya, R. (2004). Turnover of actin in Chlamydomonas flagella detected by fluorescence recovery after photobleaching (FRAP). Cell Struct. Funct. 29, 67–72. Waterman-Storer, C.M., and Danuser, G. (2002). New directions for fluorescent speckle microscopy. Curr. Biol. 12, R633–640. Wheeler, G.L., Joint, I., and Brownlee, C. (2008). Rapid spatiotemporal patterning of cytosolic Ca2þ underlies flagellar excision in Chlamydomonas reinhardtii. Plant J. 53, 401–413. Wiedenmann, J., Ivanchenko, S., Oswald, F., Schmitt, F., Röcker, C., Salih, A., Spindler, K.D., and Nienhaus, G.U. (2004). EosFP, a fluorescent marker protein with UV-inducible green-to-red fluorescence conversion. Proc. Natl. Acad. Sci. USA 101, 15905–15910.

CHAPTER 10

Purification of IFT Particle Proteins and Preparation of Recombinant Proteins for Structural and Functional Analysis Robert H. Behal, Ewelina Betleja, and Douglas G. Cole Department of Microbiology, Molecular Biology and Biochemistry and the Center for Reproductive Biology, LSS142, University of Idaho, Moscow, Idaho 83844-3052

Abstract I. Introduction II. Isolating IFT Particle Proteins from Chlamydomonas Flagella A. Flagellar Isolation B. Extraction of IFT Particle Proteins C. How to Prepare, Run, and Harvest Sucrose Density Gradients D. Purification of IFT Particle Complexes E. Purification of the IFT Complex B Core III. Recombinant IFT Particle Proteins A. Engineering Rationale B. Materials C. Results D. Uses for Recombinant Proteins IV. Summary and Concluding Remarks Acknowledgments References

Abstract Intraflagellar transport (IFT) is characterized by a robust bidirectional movement of large proteinaceous particles along the length of eukaryotic cilia and flagella. Essential for the assembly and function of the organelle, IFT is believed to transport a large array of ciliary components in and out of the organelle. Biochemical analysis of the proteins METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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involved with this transport has been largely dependent on the ability to isolate suitable quantities of intact cilia or flagella. One model organism, Chlamydomonas reinhardtii, has proven to be especially well-suited for such endeavors. Indeed, many of the IFT particle proteins were initially identified through biochemical analysis of green algae. This chapter describes some of the most effective methods for the purification of IFT particle proteins from Chlamydomonas flagella. This chapter also describes complementary approaches where recombinant IFT proteins are generated with affinity tags that allow rapid and specific purification. The recombinant proteins can be used to analyze protein–protein interactions and can be directly delivered to mutant cells to analyze functional domains. Although the techniques described here are focused entirely on Chlamydomonas IFT proteins, the approaches, especially regarding recombinant proteins, should be applicable to the study of IFT machinery in other model organisms.

I. Introduction The phenomenon of intraflagellar transport (IFT) was first documented less than 20 years ago when gliding Chlamydomonas cells were viewed using video-enhanced differential interference contrast microscopy (Kozminski et al., 1993). Amenable to light microscopy, the unicellular biflagellate green algae revealed a robust, bidirectional movement of large (50–300 nm) proteinaceous particles (formerly known as rafts) as they displaced flagellar membrane while moving quickly along the length of the axonemal microtubule doublets (Kozminski et al., 1995). Since its discovery, IFT has been found to be strongly conserved as an essential component in the assembly and function of eukaryotic cilia and flagella [for comprehensive reviews of IFT, see Blacque et al. (2008); Rosenbaum and Witman (2002); Scholey (2003)]. In Chlamydomonas, anterograde IFT particles are moved out to the distal tip of the organelle at ~2 µm/s with approximately 100 particles arriving at the end of every minute. After a distal tip rearrangement, nearly twice as many retrograde particles are returned to the cell body by cytoplasmic dynein 1b/2 at a speed of ~3.5 µm/s (Dentler, 2005; Iomini et al., 2001; Kozminski et al., 1993, 1995). Anterograde movement is driven by kinesin-2; the Chlamydomonas heterotrimeric kinesin-2 is composed of the FLA10 and FLA8 motor subunits along with a third, nonmotor subunit known as FLA3 (KAP) (Cole et al., 1998; Kozminski et al., 1995; Miller et al., 2005; Mueller et al., 2005; Walther et al., 1994). Retrograde movement is driven by cytoplasmic dynein 1b/2; the Chlamydomonas version contains at least four distinct subunits including two copies of the large heavy chain motor subunits (DHC1b), two copies of the intermediate chain (FAP133), two copies of the light intermediate chain (D1bLIC), and the LC8 light chains (Hou et al., 2004; Pazour et al., 1998, 1999; Perrone et al., 2003; Porter et al., 1999; Rompolas et al., 2007). The particles being moved by the IFT motors contain multiple copies of nearly 20 proteins which appear to act as scaffolds responsible for the transport of numerous cargos including axonemal building blocks and membrane proteins [for recent reviews

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Table I Chlamydomonas IFT Particle Proteins Complex A Subunitc

Predicted size

IFT144 IFT140 IFT139 IFT122 IFT121 IFT43

1367 aa/150.8 kDa 1384 aa/154.6 kDa 1355 aa/152.0 kDa 1239 aa/139.2 kDa 1224 aa/136.1 kDa 272 aa/28.8 kDa

Complex B Corea subunits

Complex B Peripheralb subunits

Subunit

Predicted size

Subunit

Predicted size

782 aa/86.3 kDa 683 aa/77.1 kDa 641 aa/71.4 kDa 454 aa/50.4 kDa 343 aa/37.9 kDa 204 aa/22.8 kDa 189 aa/20.4 kDa 192 aa/22.8 kDa

IFT172 IFT80 IFT57 IFT20

1755 aa/197.6 kDa 765 aa/ 85.7 kDa 469 aa/51.3 kDa 135 aa/15.6 kDa

IFT88 IFT81 IFT74/72 IFT52 IFT46 IFT27 IFT25 IFT22

a

The complex B core is the subcomplex that remains after treatment with 300 mM NaCl (Lucker et al., 2005). b The peripheral B subunits are dissociated from complex B in the presence of 300 mM NaCl. c IFT particle subunit names reflect the relative electrophoretic mobility of each protein when run on SDSPAGE as established in Cole et al. (1998).

that focus specifically on Chlamydomonas IFT machinery and mutants, see Cole (2009); Cole and Snell (2009)]. First isolated from Chlamydomonas flagella, the IFT particle proteins represent two biochemically distinct subsets or complexes (Table I; Cole et al., 1998; Piperno and Mead, 1997; Piperno et al., 1998). Complex A contains six distinct protein subunits while complex B contains another dozen which can partially dissociate in response to higher ionic strength to reveal a core set of eight subunits (Brazelton et al., 2001; Deane et al., 2001; Hou et al., 2007; Lechtreck et al., 2009; Lucker et al., 2005; Pazour et al., 2000, 2002; Pedersen et al., 2005; Qin et al., 2001, 2004, 2007; Wang et al., 2009). This chapter focuses on the biochemical purification of these flagellar IFT particle complexes and includes some strategies and methods for working with recombinant versions of specific IFT proteins.

II. Isolating IFT Particle Proteins from Chlamydomonas Flagella A. Flagellar Isolation Flagella were isolated from Chlamydomonas strains using the classic pH shock approach as described by Witman et al. (1972) with modifications as follows: In order to harvest flagella, Chlamydomonas cells are grown in 24–64 l liquid TAP (Gorman and Levine, 1965) or other liquid media in 8 l glass or polycarbonate bottles that have been aerated vigorously using aeration stones and HEPA-filtered pressurized air which can be supplemented with CO2 up to 5%. We typically use TAP medium without CO2 supplementation. Cells can be grown under continuous light or, as we prefer, using a light : dark cycle of 14:10 h. Care must be exercised not to allow the

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cells to achieve a quiescent and largely unflagellated state. Most strains grow well in a range of temperatures from 20–25°C; growth is faster at 25°C but the chance for overgrowth is much greater. We find 22–23°C to work well for routine flagellar preparations. Cells are harvested from large volumes (> 8 l) of liquid medium using a Pellicon tangential flow filtration device (Millipore, Bedford, MA, USA) with a 0.45-µm HVMP-C Pellicon 2 Cassette and/or low-speed centrifugation at 1200–1400 g for 3 min. Four 500-ml conical bottles allow 2 l of cells to be centrifuged at one time; little or no braking will minimize loss of cells due to reduced swirling of the supernatant layer. Cells are immediately resuspended in fresh media at ~1/4–1/5 original volume and vigorously aerated under intense light for 1–8 h at room temperature to ensure maximal flagellation. Tangential flow filtration often helps recently divided cells to lose their mother cell wall and assemble flagella. Care must be taken not to keep cells too concentrated (1/10–1/50 original volume) for too long a period as cells will tend to spontaneously deflagellate and eventually die. Cells are concentrated prior to deflagellation using a combination of tangential flow filtration and centrifugation at 1200–1400 g for 3 min. Cells are resuspended in 200–600 ml 10 mM HEPES, pH 7.2. To increase flagellar load of IFT proteins, sucrose can be added to a final concentration of 5.0% 5–10 min prior to deflagellation. At all stages, cells should be monitored under the light microscope to verify cells are flagellated. When the cells are very dense, the flagella will beat slowly, if at all, and the cells will stop swimming; thus, swimming behavior is not a good indicator of flagella presence when cells become concentrated. Flagella are shed by cells in response to acidification (pH 4.6–4.8) with 0.5 M acetic acid with vigorous mixing. Deflagellation is confirmed using phase microscopy after 25 s; cells are then rapidly neutralized with 0.5 M KOH before 2 min total time has elapsed. If sucrose was not added previously, bring the solution to a final sucrose concentration of 5%; add 1/9 volume 50% sucrose while the cell solution continues to mix. If cells are going to be saved for reflagellation and a second harvesting, do not add protease inhibitors or EGTA and do not underlay with 25–30% sucrose. Instead, proceed directly to 1400 g centrifugation for 5 min in 250-ml conical bottles. If cells are not going to be used for subsequent harvests, add EGTA to a final concentration of 0.1 mM as the solution is rapidly cooled in an ice–H2O bath. Protease inhibitors are added to a final concentration of 2.0 mM phenylmethyl-sulfonyl fluoride (PMSF), 50 µg/ml soybean trypsin inhibitor, 1 µg/ml pepstatin A, 2 µg/ml aprotinin, and 1 µg/ ml leupeptin. The addition of 1 mM DTT is optional and can be avoided if chemical cross-linking or other oxidation-reduction-sensitive experiments are planned. All subsequent steps are carried out on ice or at 4°C. Most of the cell bodies are removed by centrifugation at 1400 g for 5 min at 4°C using 250-ml conical bottles. Flagella will be largely retained in the supernatant which is carefully transferred to 50-ml conical tubes (30 ml/tube) and then underlayed with 7–10 ml cold 25% sucrose in 10 mM HEPES, pH 7.2. The remaining cell bodies are removed from the top layer by centrifugation at 1500 g for 10 min at 4°C. The flagella-containing top layers are carefully removed and pooled.

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Flagella are collected by high-speed centrifugation (SS34 rotor, Sorvall) at 10,000 rpm (7,800g) for 15–20 min. The resulting pellet is thoroughly resuspended in 2–5 ml 5% sucrose in 10 mM HEPES, pH 7.2, with protease inhibitors, and then centrifuged through 2–4 ml 15% sucrose in 5 mM HEPES, pH 7.2, plus half-strength protease inhibitors in 12- or 15-ml conical tubes in an HB-6 rotor or SS34 rotor at 10,000 rpm (7,800g) for 15 min. A 15-ml conical tube will likely not fit in the rotor if the lid is present so it will be necessary to use a rotor that locks onto the centrifuge spindle in the absence of the lid. This step usually produces a stratified pellet consisting of three layers. The bottom layer is small, tight, dark green, and is enriched in cell bodies. The middle layer is white and consists primarily of flagella. The top layer is light green, flocculent, easily removed, and contains flagella and an unidentified amorphous material. Typically only the white layer of flagella is recovered and resuspended in a minimal volume (0.1–1.0 ml) of HMDEK buffer (10 mM HEPES, pH 7.2, 5 mM MgSO4, 1 mM DTT, 0.5 mM EDTA, and 25 mM KCl) containing the same protease inhibitors as described above. Flagella can be extracted immediately or frozen and stored at –80°C for later use.

B. Extraction of IFT Particle Proteins Routine Chlamydomonas IFT particle protein purification consists of two major steps: (1) extraction of solubilized IFT particle proteins and (2) fractionation by sucrose density gradient centrifugation. These methods will be described in detail and will include some discussion on alternative purification utilizing size exclusion chromatography. While immunoprecipitation using specific antibodies bound to an insoluble matrix is a powerful tool to specifically purify IFT complexes (Cole et al., 1998; Lechtreck et al., 2009; Lucker et al., 2005; Qin et al., 2004), subsequent analysis of antibody resins has been largely limited to protein electrophoresis and/or mass spectroscopy and will not be discussed in depth here. 1. Option 1: Detergent Extraction. As described in Cole et al. (1998), fresh or frozen (–80°C) flagella are resuspended in 3- to 10-fold buffer volume to pellet mass using 0.05–0.10% NP-40 (nonidet P-40; nonylphenylpolyethylene glycol; US Biological cat#N3500) in HMDEK plus protease inhibitors to partially dissolve the lipid bilayer. This releases a soluble fraction known as the membrane plus matrix or “M þ M” which contains soluble flagellar proteins and a subset of flagellar membrane-associated proteins. Extract for 10 min on ice before centrifugation. 2. Option 2: Freeze–Thaw Extraction. As slightly modified from Rompolas et al. (2007), freshly isolated flagella are resuspended in 3- to 10-fold HMDEK buffer with protease inhibitors before freezing on dry ice or in liquid N2; if necessary, frozen aliquots can be stored at –80°C before proceeding. Frozen flagellar aliquots are allowed to thaw at room temperature and then refrozen and thawed two additional times before centrifugation. This releases a soluble fraction of matrix proteins. This technique is notable because it allows the extraction of an IFT supracomplex that contains the IFT dynein, kinesin-2, and complex A as

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described by Rompolas et al. (2007). Thus, the freeze–thaw extraction may also preserve other higher order assemblies and should be considered when planning to study matrix components. 3. Additional Extraction Variables: Inclusion of 1–5 mM Mg2þ-ATP during extraction will usually increase the yield of soluble IFT particle proteins. Increasing ionic strength can also increase yield, but it also enhances extraction of axonemal proteins and causes complex B to dissociate. 4. Centrifugation: The solubilized matrix or membrane plus matrix components are largely separated from the insoluble material by 10 min of centrifugation at 16,000 g at 4°C (microcentrifuge). The supernatant can be further clarified by a second centrifugation for 10 min at 16,000 g. If a refrigerated AirfugeTM (Beckman Coulter, Inc, Fullerton, CA, USA) or tabletop ultracentrifuge is available, the second run can be performed at 50,000–100,000 g for 10 min to improve clarification.

C. How to Prepare, Run, and Harvest Sucrose Density Gradients Because the IFT particle complexes are fairly large, sucrose density gradient centrifugation is an effective technique to separate IFT proteins from other matrix components. 1. Formation of Sucrose Density Gradients: The choice of rotor and ultracentrifuge depends on availability. The most popular swinging bucket rotors hold either 5-ml tubes (SW55Ti, Beckman; AH-650, Sorvall) or 13-ml tubes (SW41Ti, Beckman; TH-641, Sorvall). One convenient approach for gradient formation is to place a solution of 12.5% sucrose in HMDEK or other buffer in a 5-ml ultra tube (Davis and Pearson, 1978). After two cycles of freeze/thaw, the tube will contain a gradient of approximately 5–20% sucrose but does have the caveat that buffers and salts will also form a similar gradient and the Chlamydomonas complex B is sensitive to ionic strength. A more popular approach is to use a standard gradient former. We prefer gradient formers that have pistons in each cylinder so that the level of fluid in each stays even. The outlet tube from the gradient former can be run through a peristaltic pump in order to deliver the fluid at a constant rate; the gradient former shown in Fig. 1 (J17, Jule Inc., Milford, CT, USA) also has the option of bypassing the peristaltic pump by using a feedscrew to control the piston movement and fluid flow. Our preferred technique for pouring the gradient is to place the lighter sucrose solution (e.g., 10% sucrose) in the cylinder containing a magnetic stir bar and the outlet tube, and the heavier solution (e.g., 25% sucrose) in the other cylinder. The end of the outlet tube is fitted with a 10-, 25, or 50-µl capillary tube which is then placed into the centrifuge tube so that the sucrose solution is delivered directly to the bottom of the tube. Care should be taken to avoid pumping air bubbles while pouring the gradient in order to minimize mixing. A little bromophenol blue coloring can be added to the more dense solution to verify the formation of the gradient (Fig. 1, inset).

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Formation of a sucrose density gradient. The J17 (Jule Inc.) gradient former shown is a typical dual cylinder apparatus where the less-dense solution (10% sucrose) is placed in the cylinder with the outlet tube and magnetic stir bar. An equal volume of the more dense solution (25% sucrose; shown with bromophenol blue) is placed in the remaining cylinder. The optional pistons are lowered with a feedscrew to ensure both cylinders contain equal volumes at all times; alternatively the outlet tube can be fed through a peristaltic pump. With constant mixing, the more dense solution is slowly added to the less-dense and the constantly changing mixture is delivered to the bottom of the centrifuge tube via the delivery tubing (inset). The addition of bromophenol blue to the heavier component reveals a linear gradient when completed.

2. Sucrose Density Gradient Calibration: The rate of sedimentation of a given protein complex is described by its sedimentation value (S-value), which has units of Svedbergs or 10–13 s. The S-value of a purified protein can be determined precisely by using analytical ultracentrifugation, but this approach is problematic with complex mixtures such as flagellar extracts. It is possible, however, to indirectly measure the S-value by comparison to sedimentation standards which can be loaded along with the sample protein or loaded onto a second, identical gradient that undergoes the same centrifugation. In our gradients, we routinely use bovine thyroglobulin (19S), porcine catalase (11.3S), BSA (4.6S), ovalbumin (3.5S), and equine cytochrome c (1.86S). Cytochrome c (red) and catalase (greenish-brown) have the added benefit of color so that one can immediately see

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the separation of the proteins; BSA can be incubated with bromophenol blue before loading so that it, too, would be visible. The standard gradient is then fractionated and analyzed by SDS-PAGE; the distance each protein migrated (sedimented) from the top of the gradient is plotted against the S-value which should produce a linear standard curve. 3. Sucrose Density Gradient Fractionation: Gradients are typically fractionated into 18–26 aliquots and subsequently analyzed by SDS-PAGE and Western blotting. In brief, one end of a 10- to 50-µl capillary tube is placed carefully into the bottom of the tube (avoiding any pelleted material) while the other end is connected to tubing that is run through a peristaltic pump; PVC tubing with an inner diameter of 0.76 mm (Rainin; cat#39-624) works well for 5- to 13-ml gradient fractionation. Gradients can also be harvested by syringe needle puncture at or near the bottom of the gradient tube. Fractions are collected in microcentrifuge tubes on ice; aliquots are frozen at –28°C or colder for long-term storage. D. Purification of IFT Particle Complexes Purification of the Chlamydomonas IFT particle proteins has traditionally relied upon physical methods such as sucrose density gradient centrifugation. Initially, all of the known particle proteins cofractionated on such gradients, but subsequent studies have shown that these IFT proteins can be separated into two distinct complexes, A and B. This section describes some of the most popular approaches. The most common method for purification of the IFT complexes utilizes sucrose density gradient centrifugation. In the 1998 experiment shown in Fig. 2, M þ M 20%

Fig. 2

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Sucrose density gradient profile of Chlamydomonas flagellar membrane plus matrix. The membrane plus matrix extract from flagella of wild-type cells (CC-125) was fractionated on an 11-ml (SW41 rotor; Beckman) 5–20% sucrose gradient in 10 mM HEPES, pH 7.2. The Coomassie-stained SDSPAGE gel (7.5% acrylamide) of the gradient profile shows many of the IFT particle polypeptides that cosediment at ~16 S (highlighted by white arrowheads with apparent electrophoretic mobilities listed on the left). Reprinted with labeling modification from Cole et al. (1998) with permission from Rockefeller University Press, New York, NY, USA.

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proteins were fractionated on an 11-ml (SW41, Beckman) sucrose density gradient (in 10 mM HEPES, pH 7.2) that was centrifuged for 12 h at 38,000 rpm (178,000g). It should be noted that all of the IFT proteins cofractionate at ~16S under these conditions. These proteins, however, are segregated into two separable complexes, A and B, which had become apparent early in our studies when using other methods of purification such as immunoprecipitation (antiIFT139 resin) and size exclusion chromatography (Cole et al., 1998). An excellent example of chromatographic separation is shown in Fig. 3, where Rompolas et al. (2007) fractionated matrix components on a Superose-6 HR10/30 column (Amersham Biosciences/GE Lifesciences, Piscataway, NJ, USA); complex A peaks in fraction 6 while complex B peaks a little sooner in fraction 5. This experiment is also notable because the gentle freeze/thaw extraction (no detergent) of the flagella preserved the existence of a large supracomplex containing some of the IFT complex A and IFT motors (not show here) that eluted early on the Superose-6 (see fractions #2–3 in Fig. 3).

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Fig. 3 Gel filtration profile of Chlamydomonas flagellar matrix. The matrix components were extracted from wild-type (CC-124) flagella using three cycles of freeze/thaw and then fractionated using a Superose-6 gel filtration column. Eluted fractions were resolved on 5–15% SDS-PAGE gels and stained with Coomassie Blue (upper panel); similar gels were transferred to nitrocellulose and probed with antibodies (lower panels) against IFT139 (complex A) and IFT81 (complex B). Although the elution profile of both IFT complexes partially overlap, the mean rates of diffusion for A and B differ. Reprinted with modification from Rompolas et al. (2007) with permission from the Company of Biologists, Cambridge, UK.

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While centrifugation is often employed to separate IFT proteins from the majority of other flagellar components, care must be taken. As shown in Fig. 4, the amount of centrifugal force can affect the resolution of IFT components. IFT172 is a peripheral component of IFT complex B: when centrifuged at lower speeds such as 29,000 rpm (80,000 g), it remains associated with complex B. However, when subjected to greater RCF at 55,000 rpm (287,000 g), IFT172 appears to partially dissociate from complex B (compare IFT172 to IFT88 and IFT81 in Fig. 4). A partial dissociation could explain why the sedimentation rates at 55,000 rpm (287,000 g) for all B components, including IFT88 and IFT81 that are shown here, are shifted slightly slower relative to complex A. It should be noted that !2t was the same for these two SW55Ti runs so that most proteins would migrate nearly the same distance in both gradients. While this sort of experiment reveals fragility in complex B, simply increasing the salt concentration leads to even greater dissociation as detailed in the next section.

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Fig. 4 IFT complex B is sensitive to centrifugal shear forces. The membrane plus matrix extract from flagella of fla-2 cells (CC-1390) was fractionated on identical 5-ml (SW55Ti rotor; Beckman) 10–25% sucrose gradients in HMDEK Buffer. (A) After loading 250 µl M þ M, the gradient was centrifuged for 16:00 h at a constant rate of 29,000 rpm (80,000 g). (B) After loading 250 µl M þ M, the gradient was centrifuged for 4:28 h at a constant rate of 55,000 rpm (287,000 g). Each gradient was fractionated into 21 fractions containing 250 µl each; as shown here, only 14 fractions were resolved on 5–20% SDS-PAGE gels and stained with CoomassieBlue. Corresponding transfers were probed with specific complexes A and B antibodies as shown. While most of the IFT proteins cosedimented under either condition, the apparent S-value of IFT172 decreased at the higher speed indicating that complex B is sensitive to the rate of centrifugation.

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Fig. 4 (Continued)

E. Purification of the IFT Complex B Core As isolated from Chlamydomonas, IFT complex B is sensitive to ionic strength. As described previously, the addition of 300 mM NaCl causes several peripheral subunits to dissociate revealing a salt-stable core composed of IFT88, IFT81, IFT74, IFT52, IFT46, IFT27, IFT25, and IFT22 (Lucker et al., 2005). It should be noted that the terms “peripheral”and “core” are biochemical constructs that merely describe the behavior of these subunits in response to increasing salt concentration. For this section we describe the initial separation of the peripheral subunits from the core followed by a chromatographic fractionation of the core. To separate the B core from the peripheral subunits, flagellar matrix components (detergent is optional in the initial extraction) are fractionated (37,000 rpm, 16 h) on a 13-ml (SW41, Beckman) 10–25% sucrose density gradient equilibrated in HMDEK buffer supplemented with 300 mM NaCl (HMDEK-300). SDS-PAGE and Western blot analysis of such a gradient reveals that a subset of complex B subunits (termed the B core) comigrated at ~11S while four of the B subunits (IFT172, IFT80, IFT57, and IFT20) fully separated from the original complex (Fig. 5, panel A). The 16S complex A remains fully intact and is clearly separated from all complex B components. The 11S core fractions are pooled and loaded onto a 1.6 60 cm Sephacryl S-300 column (GE Lifesciences) equilibrated in HMEK-300 buffer. As shown in Fig. 5 (panel B), the

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core components coelute just ahead of the thyroglobulin calibration standard with a diffusion coefficient of 1.8 10–7 cm2/s, consistent with the core having a molecular mass of approximately 500,000 Da. A more rapid, albeit less-reversible, method to isolate the complex B core is to use antibody pulldown in buffer conditions that favor dissociation of the peripheral B subunits (e.g., HMEK-300 or HMDEK-300). Two

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antibodies that are especially effective for complex B pulldowns are the anti-IFT81 monoclonals (combinations of 81.1, 81.3, and 81.4 are more effective than individual use; Cole et al., 1998; Lucker et al., 2005) and the polyclonal anti-IFT74/72 antibody (a-IFT721 and a-IFT722; Qin et al., 2004).

III. Recombinant IFT Particle Proteins A. Engineering Rationale Biochemical analysis of IFT particle proteins has been challenging for multiple reasons. Biochemical analysis has been largely limited to Chlamydomonas flagella and, even so, the quantities available are not large. In addition, the particles readily dissociate into two major complexes, A and B, and complex B itself is not particularly stable. Thus, we have sought to use bacterial and yeast expression of recombinant proteins to study the structure and interactions of IFT particle proteins. Coexpression of multiple proteins can be an especially powerful approach to generate protein complexes. To achieve this, we have adopted the Duet system of plasmids which allow coexpression of up to eight different proteins in one bacterial host; a number of these proteins can be engineered to contain a variety of affinity tags. The Duet vectors incorporate different origins of replication and antibiotic selection markers; therefore, stable cotransformation of a single host bacterium is possible. Furthermore, we have combined select Duet vectors with the pMal vector, which incorporates the maltose-binding protein (MBP), to take advantage of rapid and reversible amylose chromatography. Although we have used this approach to generate complexes containing as many as five different IFT particle proteins we will keep the discussion simple by presenting data on the production and purification of an IFT81/IFT74 subcomplex.

Fig. 5 Purification of the IFT complex B core. The soluble matrix components were extracted from wildtype (CC-124) flagella by trituration and fractionated through a 13-ml (SW41 rotor, Beckman) 10–25% sucrose density gradient in HMEK buffer supplemented with 300 mM NaCl (HMEK-300) to increase ionic strength. (A) Silver-stained 7.5% SDS-PAGE gel and corresponding Western blots probed with specific IFT antibodies as indicated. Complex A proteins (IFT144 and IFT139) cosediment at 16S, whereas complex B dissociates into separate components. The complex B core proteins cosediment at 11S and include IFT88, IFT81, IFT74, IFT72, IFT52, IFT46, and IFT27; IFT25 and IFT22 also cosediment at 11S but are not shown here. IFT172, IFT80, IFT57, and IFT20 fully dissociate from complex B under these conditions. (B) Following fractionation on the 10–25% sucrose density gradient shown in (A), the 11S complex B core was applied to a Sephacryl S-300 16/60 FPLC column equilibrated in HMEK-300. Two-milliliter fractions were collected, fractionated by SDS-PAGE, transferred to nitrocellulose, and probed with antibodies as indicated. The arrows, from left to right, indicate the elution position of calibration standards, thyroglobulin (T), apoferritin (A), alcohol dehydrogenase (ADH), bovine serum albumin (BSA), and ovalbumin (OV), respectively. Note that all of the complex B core subunits coelute (IFT25 and IFT22 are not shown). Reprinted from Lucker et al. (2005) with permission by ASBMB.

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B. Materials • MBP-fusion proteins. We are using the pMalC2X vector (New England Biolabs, Ipswich, MA, USA) for MBP fusions. While this particular plasmid is no longer available through NEB, more recent versions should work as well. The pMalC2X vector contains the pBR322 (ColE1) origin of replication and an ampicillin resistance marker. • Poly-histidine fusion proteins. The pRSFDuet vector (Novagen, San Diego, CA, USA) contains two T7lac promoters and two multiple cloning sites, allowing for the simultaneous expression of two proteins from a single plasmid; proteins encoded in MCS1 contain an N-terminal hexahistidine (His6) tag. The RSF1030 replicon and kanamycin resistance marker allow this vector to be cotransformed with the pMalC2X vector detailed above. Other Duet vectors with unique replicons and selectable markers are available from Novagen. Expression of recombinant proteins from the Duet vectors requires a DE3 bacterial host strain expressing the gene for T7 RNA polymerase. • We routinely transform BL21(DE3) host strains for coexpressions. The first plasmid is transformed into chemically competent cells; subsequent cotransformations utilize electroporation. It is necessary that each of the plasmids for coexpression contain different replicons and different selectable markers. • Amylose resin for MBP affinity chromatography (New England Biolabs, Ipswich, MA, USA, #E8021). • HisPur Cobalt resin for metal chelate chromatography (Pierce, Rockford, IL, USA, #89965). C. Results A powerful aspect of recombinant expression is the ability to generate multiple fusion proteins that contain unique affinity tags. We have relied extensively on MBP as a fusion tag due to its ability to reversibly and specifically bind to amylose resin; elution from amylose requires only maltose. In addition, the presence of the MBP tag tends to stabilize and increase solubility of fusion partner proteins, such as the complex A components. A caveat of MBP use is the fact that it adds an additional 43 kDa of protein to fusion partners. Histidine-tagging (six or more consecutive histidine residues) is also effective and allows rapid purification on metal chelate resins that use Ni2þ or Co2þ as the affinity ligand; elution typically requires 100–250 mM imidazole. A caveat of metal chelate chromatography is that the resin acts as an ion exchange matrix that has nonspecific affinity for some proteins. Complex B components IFT81 and IFT74 have been shown previously to interact in yeast-based two-hybrid analyses while the native proteins are readily cross-linked with DFDNB (Lucker et al., 2005). To illustrate the bacterial coexpression approach, we have simultaneously expressed IFT74 and IFT81, and have demonstrated a specific interaction between the two recombinant proteins by tandem affinity chromatography. In brief, a cDNA fragment encoding the IFT74 protein was cloned into MCS1 of the pRSFDuet vector, creating a His6–IFT74 hybrid protein. Similarly, the protein-encoding sequence of IFT81 was cloned into pMalC2X, resulting in an MBP-IFT81 fusion protein. Following cotransformation, both proteins were simultaneously expressed for 2.0 h at 37°C in

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Fig. 6 Tandem affinity purification of MBP-IFT81 and His6-IFT74. MBP-tagged IFT81 and His6-tagged IFT74 were simultaneously expressed in a single host E. coli. Soluble protein was extracted in ME Buffer (20 mM Tris-HCl and 0.20 M NaCl, pH 7.4) and loaded onto a 1.0-ml amylose affinity column; after washing, MBP-IFT81 was specifically eluted with 10 mM maltose in ACB Buffer and collected in 1.0-ml fractions. Peak amylose fractions were directly loaded onto a 1.0-ml cobalt affinity column; after washing, His6-IFT74 was specifically eluted with 150 mM imidazole into 1.0-ml fractions. Note that the MBP-IFT81 and the His6-IFT74 cofractionated from both columns.

response to 1.0 mM IPTG induction in a volume of 200 mL LB medium. Harvested bacteria were resuspended in 7.5 ml of MBP-Extraction Buffer (ME Buffer: 20 mM TrisHCl, pH 7.4, 0.20 M NaCl plus protease inhibitors described above) and lysed by sonication on ice. After insoluble material was removed by centrifugation (10,000 g, 10 min, 4°C), the clarified protein extract was loaded onto a 1.0-mL amylose column; disposable Poly-Prep (0.2–2.0 ml bed volume, BioRad #731–1550) and Econo-Pac (1– 20 ml bed volume; BioRad #732–1010) columns work well for small volumes of affinity resin. The column was washed extensively with ME Buffer to remove weakly or unbound components while the IFT81–IFT74 complex was specifically eluted with 10 mM maltose in ME Buffer; a majority of the bound protein eluted in the first four fractions (Fig. 6). Peak fractions (1.0 ml) were pooled and immediately applied (without exchanging buffers) to a 1.0-ml HisPur cobalt column and the resin was washed extensively with Equil-Wash Buffer (50 mM Na2HPO4, 0.30 M NaCl, 10 mM imidazole, pH 7.4). Specifically bound IFT81–IFT74 complex was eluted with Elution Buffer (50 mM Na2HPO4, 0.30 M NaCl, 150 mM imidazole, pH 7.4). Stoichiometric coelution of the two proteins is shown in Fig. 6. D. Uses for Recombinant Proteins While it is often preferable to study proteins isolated directly from the model organism, several factors conspire against this approach. Although Chlamydomonas has proven to be an especially powerful model organism for the biochemical study of flagellar machinery, there are limitations to the yield and purity of IFT proteins

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extracted from isolated flagella or whole cells. More importantly, Chlamydomonas has proved resistant to homologous genetic recombination, which has impeded structurefunction studies of individual proteins. To overcome these obstacles, we can utilize recombinant protein expression in heterologous hosts such as yeast and bacteria. Although Escherichia coli lacks much of the post-translational modification machinery found in eukaryotes, the benefits of high protein yield, ease of molecular engineering, and simple and rapid growth conditions make prokaryotic recombinant protein expression an attractive and valuable tool. As indicated above, we routinely coexpress multiple proteins in a single bacterial host. This enables high-yield production of various mixtures of IFT particle proteins, each of which can be uniquely tagged to afford rapid and specific purification. Subsequent analyses of the resulting complexes can include the determination of size, shape, and stoichiometry. Standard deletional analysis can be used to identify specific interaction domains involved in the assembly of the IFT particles. Recombinant flagellar proteins have also been shown to rescue mutant phenotypes by direct protein electroporation into living cells (Hayashi et al., 2002; Watanabe et al., 2004). Lastly, the IFT subcomplexes are also excellent candidates as baits in pull-down assays that search for IFT-interacting proteins in Chlamydomonas extracts.

IV. Summary and Concluding Remarks The important roles of IFT in the assembly and function of cilia and flagella have stimulated considerable research focused on understanding the molecular mechanisms by which IFT operates. Although genetic and cell biological approaches to understanding the function of IFT in key model organisms have proven to be invaluable, biochemical studies exploiting Chlamydomonas have played an early and central role beginning with the isolation and identification of much of the IFT machinery. And it is likely that the biochemical approach will continue to play an essential role as the field focuses on specific problems such as understanding the molecular architecture of the particles, determining how the particles dock onto and release their cargos, and understanding how the different phases of the transport (e.g., anterograde movement or distal tip rearrangement) are regulated. With this in mind, this chapter has been written to provide a detailed description of how to purify IFT particle proteins from Chlamydomonas flagella and a description of how we are using heterologous expression to complement our studies. The successful generation and purification of various subcomplexes including, but not limited to, the IFT81/IFT74 complex described here and an IFT88/IFT52/IFT46 ternary complex (Lucker et al., 2006), demonstrates that it is possible to recreate select portions of the IFT particles. It also gives us reason to think that it may be possible to assemble complete IFT complexes and particles using only recombinant proteins.

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Acknowledgments We gratefully acknowledge members of the Cole laboratory for their contributions with special thanks to Mark Miller (University of Idaho, Moscow) for technical assistance and advice and for critical review of this manuscript, Thomas Esplin (Brigham Young University-Idaho, Rexburg) for technical assistance, and Ben Lucker (Washington State University, Pullman) for insightful and critical discussions.

References Blacque, O.E., Cevik, S., and Kaplan, O.I. (2008). Intraflagellar transport: From molecular characterisation to mechanism. Front. Biosci. 13, 2633–2652. Brazelton, W.J., Amundsen, C.D., Silflow, C.D., and Lefebvre, P.A. (2001). The bld1 mutation identifies the Chlamydomonas osm-6 homolog as a gene required for flagellar assembly. Curr. Biol. 11, 1591–1594. Cole, D.G. (2009). Intraflagellar transport. In “The Chlamydomonas Sourcebook: Cell Motility and Behavior” (G. Witman and E. Harris, eds.), 2nd edn., Vol. 3, Chapter 4, pp. 71–113. Academic Press, San Diego. Cole, D.G., Diener, D.R., Himelblau, A.L., Beech, P.L., Fuster, J.C., and Rosenbaum, J.L. (1998). Chlamydomonas kinesin-II-dependent intraflagellar transport (IFT): IFT particles contain proteins required for ciliary assembly in Caenorhabditis elegans sensory neurons. J. Cell Biol. 141, 993–1008. Cole, D.G., and Snell, W.J. (2009). SnapShot: Intraflagellar transport. Cell 137, 784–784.e1. Davis, P.B., and Pearson, C.K. (1978). Characterization of density gradients prepared by freezing and thawing a sucrose solution. Anal. Biochem. 91, 343–349. Deane, J.A., Cole, D.G., Seeley, E.S., Diener, D.R., and Rosenbaum, J.L. (2001). Localization of intraflagellar transport protein IFT52 identifies basal body transitional fibers as the docking site for IFT particles. Curr. Biol. 11, 1586–1590. Dentler, W. (2005). Intraflagellar transport (IFT) during assembly and disassembly of Chlamydomonas flagella. J. Cell Biol. 170, 649–659. Gorman, D.S., and Levine, R.P. (1965). Cytochrome f and plastocyanin: Their sequence in the photosynthetic electron transport chain of Chlamydomonas reinhardi. Proc. Natl. Acad. Sci. USA 54, 1665–1669. Hayashi, M., Yanagisawa, H.A., Hirono, M., and Kamiya, R. (2002). Rescue of a Chlamydomonas innerarm-dynein-deficient mutant by electroporation-mediated delivery of recombinant p28 light chain. Cell Motil. Cytoskeleton 53, 273–280. Hou, Y., Pazour, G.J., and Witman, G.B. (2004). A dynein light intermediate chain, D1bLIC, is required for retrograde intraflagellar transport. Mol. Biol. Cell 15, 4382–4394. Hou, Y., Qin, H., Follit, J.A., Pazour, G.J., Rosenbaum, J.L., and Witman, G.B. (2007). Functional analysis of an individual IFT protein: IFT46 is required for transport of outer dynein arms into flagella. J. Cell Biol. 176, 653–665. Iomini, C., Babaev-Khaimov, V., Sassaroli, M., and Piperno, G. (2001). Protein particles in Chlamydomonas flagella undergo a transport cycle consisting of four phases. J. Cell Biol. 153, 13–24. Kozminski, K.G., Beech, P.L., and Rosenbaum, J.L. (1995). The Chlamydomonas kinesin-like protein FLA10 is involved in motility associated with the flagellar membrane. J. Cell Biol. 131, 1517–1527. Kozminski, K.G., Johnson, K.A., Forscher, P., and Rosenbaum, J.L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523. Lechtreck, K.F., Luro, S., Awata, J., and Witman, G.B. (2009). HA-tagging of putative flagellar proteins in Chlamydomonas reinhardtii identifies a novel protein of intraflagellar transport complex B. Cell Motil. Cytoskeleton 66, 469–482. Lucker, B.F., Behal, R.H., Qin, H., Siron, L.C., Taggart, W.D., Rosenbaum, J.L., and Cole, D.G. (2005). Characterization of the intraflagellar transport complex B core: Direct interaction of the IFT81 and IFT74/ 72 subunits. J. Biol. Chem. 280, 27688–27696. Lucker, B.F., Miller, M.S., Qin, H., Blackmarr, P., Ferrell, M., Rosenbaum, J.L., and Cole, D.G. (2006). Architectural analysis of the Chlamydomonas intraflagellar transport complex B. Mol. Biol. Cell. 17, Abstract no. 1611 (CD-ROM).

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Robert H. Behal et al. Miller, M.S., Esparza, J.M., Lippa, A.M., Lux, F.G., III, Cole, D.G., and Dutcher, S.K. (2005). Mutant kinesin-2 motor subunits increase chromosome loss. Mol. Biol. Cell 16, 3810–3820. Mueller, J., Perrone, C.A., Bower, R., Cole, D.G., and Porter, M.E. (2005). The FLA3 KAP subunit is required for localization of kinesin-2 to the site of flagellar assembly and processive anterograde intraflagellar transport. Mol. Biol. Cell 16, 1341–1354. Pazour, G.J., Baker, S.A., Deane, J.A., Cole, D.G., Dickert, B.L., Rosenbaum, J.L., Witman, G.B., and Besharse, J.C. (2002). The intraflagellar transport protein, IFT88, is essential for vertebrate photoreceptor assembly and maintenance. J. Cell Biol. 157, 103–113. Pazour, G.J., Dickert, B.L., Vucica, Y., Seeley, E.S., Rosenbaum, J.L., Witman, G.B., and Cole, D.G. (2000). Chlamydomonas IFT88 and its mouse homologue, polycystic kidney disease gene tg737, are required for assembly of cilia and flagella. J. Cell Biol. 151, 709–718. Pazour, G.J., Dickert, B.L., and Witman, G.B. (1999). The DHC1b (DHC2) isoform of cytoplasmic dynein is required for flagellar assembly. J. Cell Biol. 144, 473–481. Pazour, G.J., Wilkerson, C.G., and Witman, G.B. (1998). A dynein light chain is essential for the retrograde particle movement of intraflagellar transport (IFT). J. Cell Biol. 141, 979–992. Pedersen, L.B., Miller, M.S., Geimer, S., Leitch, J.M., Rosenbaum, J.L., and Cole, D.G. (2005). Chlamydomonas IFT172 is encoded by FLA11, interacts with CrEB1, and regulates IFT at the flagellar tip. Curr. Biol. 15, 262–266. Perrone, C.A., Tritschler, D., Taulman, P., Bower, R., Yoder, B.K., and Porter, M.E. (2003). A novel dynein light intermediate chain colocalizes with the retrograde motor for intraflagellar transport at sites of axoneme assembly in Chlamydomonas and mammalian cells. Mol. Biol. Cell 14, 2041–2056. Piperno, G., and Mead, K. (1997). Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella. Proc. Natl. Acad. Sci. USA 94, 4457–4462. Piperno, G., Siuda, E., Henderson, S., Segil, M., Vaananen, H., and Sassaroli, M. (1998). Distinct mutants of retrograde intraflagellar transport (IFT) share similar morphological and molecular defects. J. Cell Biol. 143, 1591–1601. Porter, M.E., Bower, R., Knott, J.A., Byrd, P., and Dentler, W. (1999). Cytoplasmic dynein heavy chain 1b is required for flagellar assembly in Chlamydomonas. Mol. Biol. Cell 10, 693–712. Qin, H., Diener, D.R., Geimer, S., Cole, D.G., and Rosenbaum, J.L. (2004). Intraflagellar transport (IFT) cargo: IFT transports flagellar precursors to the tip and turnover products to the cell body. J. Cell Biol. 164, 255–266. Qin, H., Rosenbaum, J.L., and Barr, M.M. (2001). An autosomal recessive polycystic kidney disease gene homolog is involved in intraflagellar transport in C. elegans ciliated sensory neurons. Curr. Biol. 11, 457–461. Qin, H., Wang, Z., Diener, D., and Rosenbaum, J. (2007). Intraflagellar transport protein 27 is a small G protein involved in cell-cycle control. Curr. Biol. 17, 193–202. Rompolas, P., Pedersen, L.B., Patel-King, R.S., and King, S.M. (2007). Chlamydomonas FAP133 is a dynein intermediate chain associated with the retrograde intraflagellar transport motor. J. Cell Sci. 120, 3653–3665. Rosenbaum, J.L., and Witman, G.B. (2002). Intraflagellar transport. Nat. Rev. Mol. Cell Biol. 3, 813–825. Scholey, J.M. (2003). Intraflagellar transport. Annu. Rev. Cell Dev. Biol. 19, 423–443. Walther, Z., Vashishtha, M., and Hall, J.L. (1994). The Chlamydomonas FLA10 gene encodes a novel kinesin-homologous protein. J. Cell Biol. 126, 175–188. Wang, Z., Fan, Z.C., Williamson, S.M., and Qin, H. (2009). Intraflagellar transport (IFT) protein IFT25 is a phosphoprotein component of IFT complex B and physically interacts with IFT27 in Chlamydomonas. PLoS One 4, e5384. Watanabe, Y., Hayashi, M., Yagi, T., and Kamiya, R. (2004). Turnover of actin in Chlamydomonas flagella detected by fluorescence recovery after photobleaching (FRAP). Cell Struct. Funct. 29, 67–72. Witman, G.B., Carlson, K., Berliner, J., and Rosenbaum, J.L. (1972). Chlamydomonas flagella. I. Isolation and electrophoretic analysis of microtubules, matrix, membranes, and mastigonemes. J. Cell Biol. 54, 507–539.

CHAPTER 11

Studying Cilia in Zebrafish Iain Drummond Departments of Medicine and Genetics, Harvard Medical School and Massachusetts General Hospital, Charlestown, Massachusetts 02129

Abstract I. Introduction A. Molecular Approaches to Ciliogenesis in Zebrafish B. Wholemount Confocal Imaging of Zebrafish Cilia C. An inexpensive System for High-Speed Microvideo of Cilia D. Electron Microscopy Methods for Fish Cilia II. Summary Acknowledgments References

Abstract Zebrafish are ideally suited for analysis of genes required for ciliogenesis and cilia function. Combining genetic manipulation with high-quality in vivo imaging, zebrafish embryos provide a high-throughput system for annotation of the cilia proteome. The specific advantages of the system are the availability of cilia mutants, the ability to target genes of unknown function using antisense methods, the feasibility of observing cilia in living embryos, and the ability to image fixed cilia in wholemount at high resolution. Techniques are described for analysis of mutants, gene knockdown using antisense morpholino oligos, visualizing cilia and cilia orientation in wholemount zebrafish embryos, cilia imaging by high-speed video, and electron microscopy of zebrafish cilia.

METHODS IN CELL BIOLOGY, VOL. 93 Copyright Ó 2009 Elsevier Inc. All rights reserved.

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I. Introduction The convergence of studies of cystic kidney disease, left–right asymmetry, retinal degeneration, and cilia/flagella formation has given rise to the central hypothesis that defects in the formation and function of cilia underlie this wide range of disease pathologies (Pazour, 2004; Veland et al., 2009). A relatively large number of zebrafish mutations associated with cystic kidneys are now known to cause pleiotropic defects which also include photoreceptor cell death, randomization of left–right asymmetry, and hydrocephalus, suggesting that many of these mutants represent ciliogenic genes (Brand et al., 1996; Drummond et al., 1998; Sun et al., 2004; Zhao and Malicki, 2007). Cloning the mutations has confirmed that intraflagellar transport protein genes as well as novel ciliogenic gene mutations are responsible for the observed defects (Omori et al., 2008; Owens et al., 2008; Pathak et al., 2007; Serluca et al., 2009; Sukumaran and Perkins, 2009; Sullivan-Brown et al., 2008; Sun et al., 2004; Tsujikawa and Malicki, 2004; van Rooijen et al., 2008). This work has established the zebrafish as a powerful genetic system for studies of vertebrate cilia. Since zebrafish embryos and larvae contain multiple different ciliated cell types zebrafish embryos can be used to model both sensory and motile cilia functions. Examples of motile cilia can be found in the zebrafish Kupffer’s vesicle epithelium between the 8 and 14 somite stages (http://www.elsevierdirect.com/companions/9780123749734, movie 1) (Essner et al., 2005; Kramer-Zucker et al., 2005), the olfactory placodes at 2–3 days of development (http://www.elsevierdirect.com/ companions/9780123749734, movie 2), the spinal canal from 1 to 3 days of development (http://www.elsevierdirect.com/companions/9780123749734, movie 3), and the pronephros which contains both single and multiciliated cells (http://www. elsevierdirect.com/companions/9780123749734, movie 4) (Liu et al., 2007). Pronephric cilia have a “9 þ 2” microtubule doublet organization and dynein arms characteristic of motile cilia and flagella (Kramer-Zucker et al., 2005; Liu et al., 2007; Pathak et al., 2007). Cilia motility in these organs has been shown to be important for fluid movement (Kramer-Zucker et al., 2005) and proper organ function. Sensory cilia are found in the zebrafish lateral line organs, the olfactory placodes, and the retina where photoreceptors are thought of as modified sensory cilia (Tsujikawa and Malicki, 2004). A. Molecular Approaches to Ciliogenesis in Zebrafish A principal advantage of using zebrafish to study cilia is the availability of cilia mutants and reverse genetic approaches to disrupt the function of candidate ciliogenic genes. Large-scale ENU and retroviral insertional mutagenesis identified many mutants with a spectrum of disorders now known to be linked to cilia defects (Drummond et al., 1998; Sun et al., 2004; Zhao and Malicki, 2007). The genes affecting cilia formation or function in zebrafish as well as candidate cilia genes based on common mutant phenotype are presented in Table I. Since genetic mutants represent not only genes for cilia structural proteins but also genes for cytoplasmic

11. Studying Cilia in Zebrafish

Table I Zebrafish Mutants with Pleiotropic Cilia Associated Phenotypes Name

Alleles

Gene

Description Core intraflagellar transport protein Core intraflagellar transport protein Periferal intraflagellar transport protein Ortholog of human Joubert syndrome gene AAAþ ATPase implicated in chromatinremodeling complexes Cilia paralysis; interaction with planar polarity pathways and convergent extension Subject to IFT; ortholog of C.elegans DYF-3 Ortholog of human autosomal dominant polycystic kidney disease gene

Reference

curly larry moe scorpion pontin

hi3417 hi409 hi2211 hi459 hi1055B

ift57 ift81 ift172 arl13b pontin

Sun et Sun et Sun et Sun et Sun et

al. (2004) al. (2004) al. (2004) al. (2004) al. (2004)

seahorse, zator

hi3308, tg238a, fa20r

lrrc6l

qilin pkd2

hi3959A hi4166

qilin Polycystin2

twister enigma shen yan locke

NC NC NC NC

Sun et al. (2004) Sun et al. (2004) Brand et al. (1996), Zhao and Malicki (2007) Brand et al. (1996) Zhao and Malicki (2007)

garbus elipsa

hi1392 hi2005 tf214a tm138a, tj8, ts277, tl215, to237b tm304 m649, tp49d

oval switch hitter

tz288a hu255H

NC Traf3ip1 MIP- Links IFT particle to rab8. Ortholog of C.elegans gene DYF-11. T3 Ift88 Core intraflagellar transport protein lrrc50 Cilia paralysis. Ortholog of Chlamydomonas Oda7p

Brand et al. (1996), Zhao and Malicki (2007) Drummond et al. (1998), Malicki et al. (1996), Omori et al. (2008) Brand et al. (1996) Tsujikawa and Malicki (2004) Sullivan-Brown et al. (2008), van Rooijen et al. (2008)

Brand et al. (1996), Serluca et al. (2009), Sun et al. (2004), Zhao and Malicki (2007) Ou et al. (2005), Sun et al. (2004) Sun et al. (2004)

(Continued)

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Table I (Continued ) Name

Alleles

Gene

double bubble fleer

m468, m153, m587 m477

NC Fleer

sentinel schmalhans bazooka Joe dizzy junior mr. bubble blowup fusen bubblicious big league chew hubba bubba inflated pao pao tang cyster

w38 tn222 m453, & m527 m605 m304 m305 m348 m478 m620 m696 m194 m786 m629 m329

cc2d2a NC NC NC NC NC NC NC NC NC NC NC NC NC

Description

Reference

Drummond et al. (1998) Ortholog of C.elegans gene DYF-1. Regulator of Drummond et al. (1998), Pathak et al. (2007) tubulin post-translational modification. Ortholog of human Joubert syndrome gene Owens et al. (2008) Cilia paralysis Brand et al. (1996) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998) Drummond et al. (1998)

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transport or cilia assembly proteins, these mutants constitute an important resource for identifying regulatory proteins that may not be represented in the cilia proteome. In addition to genetic mutants, reverse genetic approaches to gene function are feasible in zebrafish. Antisense knockdown approach using morpholino antisense oligos is a relatively high-throughput approach (Bill et al., 2009) but requires appropriate controls to avoid confounding off-target effects of the oligos. Generation of stable genetic mutants in a gene of interest is feasible by selection of ENU mutants (TILLING) (Wienholds et al., 2003) or by gene-targeting with engineered zinc finger nucleases (Foley et al., 2009; Meng et al., 2008). Morpholino knockdown provides the quickest route to gene function and is the most useful first-pass approach to annotating unknown genes and proteins represented in the cilia proteome. Several good reviews of morpholino use in zebrafish have been published (Bill et al., 2009; Eisen and Smith, 2008). We present here methods representing our own approach to the best use of morpholino knockdown, specifically, how to optimize dosage to achieve specific gene knockdown. It is essential to confirm that injection of morpholino antisense oligos (1) prevents expression of targeted mRNA and (2) does not cause off-target effects (artifactual phenotypes). The first consideration requires methods to measure the efficiency of gene knockdown. Morpholino antisense oligos work by steric hindrance (as opposed to RNAaseH activation) and can be used to block mRNA translation (targeting ATG initiation codons and 50 UTR) or to disrupt mRNA processing (splice site-blocking morpholinos). If an antibody to the target protein is available then it is fairly straightforward to quantify the efficiency of knockdown with an ATG morpholino by western blotting or by immunofluorescence. We favor use of splice donor site-blocking morpholinos to characterize the function of unknown proteins since they seem to be more reliable than splice acceptor-blocking morpholinos and because their use allows quantification of morpholino efficiency by RT-PCR when antibodies are not available.

1. Materials morpholino antisense oligos (Gene Tools, LLC) 2 Injection solution 200 mM KCl 10 mM Hepes pH 7.2 0.2% Phenol Red Glass Micropipettes Microinjection apparatus

2. Methods a. Morpholino Design. The first steps in splice site morpholino design are to identify splice sites by alignment of your mRNA target to the genome. Genomic resources for zebrafish are available online (http://zfin.org, http://www.sanger.ac.uk/Projects/

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D_rerio/, http://www.ncbi.nlm.nih.gov/genome/guide/zebrafish/) and alignments can be performed using tools such as Spidey or Splign (NCBI). A frequent outcome of blocking a splice donor site is exon skipping or deletion where the splice acceptor downstream bypasses the blocked splice donor site and finds the next functional splice donor 50 to the blocked site (Obara et al., 2006). This is often the splice donor of the upstream exon (exon skipping) but can also be a cryptic splice donor within the coding sequence of the targeted exon. With these considerations in mind, simple strategies for efficient disruption of protein function emerge. For instance if an mRNA encodes a protein with known catalytic function, targeting the exon encoding the catalytic site is likely to produce loss of function. Often it is possible to produce a full protein null by targeting the splice donor of the first coding exon, thereby deleting the ATG initiation codon. When these strategies are not feasible, targeting an exon that, in the event of exon skipping, would produce an out-of-frame fusion between remaining exons can be a good approach. These approaches are not fool proof; it is not possible to predict the outcome of missplicing using morpholinos. It is always necessary to determine the outcome by RT-PCR. Final design of a morpholino can be done in consultation with the manufacturer, Gene Tools LLC (http://www.gene-tools.com/). For a control morpholino, we also design an “invert sense” oligo. These have the same purine/ pyrimidine ratio as the experimental morpholino but are an irrelevant sequence. For detailed reference on morpholino design and use the reader is referred to excellent reviews on the topic (Bill et al., 2009; Eisen and Smith, 2008). For quantification of morpholino efficacy, design primers in exons flanking the targeted exon for nested RT-PCR. Ideally, the final predicted size of the RT-PCR product should be in the 400- to 600-bp range so as to resolve a smaller deletion product generated by mis-splicing. Deletions can be small (as small as 30 bp in our experience) so final analysis is best performed with 1–2% agarose gels and small products. b. Morpholino Injection and Analysis. This protocol describes how to analyze the effects of injected antisense morpholino on target mRNA splicing and optimize morpholino dosage. 1. Dissolve the morpholino in high-quality distilled water to make a 2-mM stock solution. Store in an evaporation resistant tube at –20° [storage temperature does not seem to be critical (Bill et al., 2009)]. 2. Prepare the injection system. We use a Nanoliter 2000 microinjection device from World Precision instruments (http://www.wpiinc.com). This injector is superior to air pressure-driven devices since it uses an oil column and direct piston displacement to deliver reproducible small volumes independent of micropipette tip size. Pulled glass micropipettes are used for injection; tip size can be controlled by pulling conditions or by breaking the tip under a microscope (Bill et al., 2009). 3. Determine the effective dose of the morpholino. Dilute stock solutions to make 0.5-, 0.25-, and 0.1-mM injection solutions in buffer containing 100 mM KCl, 0.1% Phenol

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Red, and 5 mM Hepes pH 7.2. Some references suggest using Danieau solution for injection solution (Bill et al., 2009), but since this is an extracellular medium with very high Ca2þ relative to cytoplasm, it is best avoided. Fill the injection micropipette with injection solution and morpholino using suction or by backfilling. 4. Inject 4.6 nl of working morpholino solution per embryo into 1- or 2-cell stage embryos. Given the approximate molecular weight of a 25mer morpholino (8000 D), the stock solutions we use generate injection amounts of roughly 18, 9, and 4 ng morpholino per embryo. Methods for immobilizing embryos for injection can be found in The Zebrafish Book online at http://zfin.org/zf_info/zfbook/chapt5/5.1. html. 5. Allow embryos to develop in egg water (http://zfin.org/zf_info/zfbook/chapt1/1.3. html) for 24, 48, and 72 hpf. Prepare RNA at each time point from groups of 10 injected embryos. RNA can be made using standard guanidinium hydrochloride/ phenol/chloroform protocols (MacDonald et al., 1987) or alternatively, RNA can be prepared from individual embryos using Qiagen RNAeasy columns (http://www1. qiagen.com/) to gauge the uniformity of knockdown efficiency. 6. RT-PCR. I. Dissolve RNA in nuclease-free water at 1 µg/ml or 5 µl water per embryo. II. Add 1 µl DNase (RNase-free) to 20 µl RNA/water. This is an essential step if you are analyzing morphant RNA. Genomic DNA will always contaminate guanidinium RNA preps. III. Incubate for 30 min at 37°. Inactivate by heating to 75° for 25 min IV. Reverse transcription: Combine: 1 µl Reverse gene-specific primer (50 µM) 1 µg Total RNA or 2.5 µl (RNA from 1/2 embryo). 1 µl dNTP’s (10 mM) Nuclease free water up to 12 µl Heat at 70° for 10 min. Cool on ice. Add: 4 µl 5 SuperScript buffer 2 µl 0.1 M DTT 1 µl RNase inhibitor RNAase water up to 20 µl 1 µl Superscript III Incubate 50 min at 42° Inactivate by heating to 70° for 15 min V. PCR In a separate tube: 5 µl 10 PCR buffer 1 µl dNTP’s (10 mM) 1 µl forward primer (50 µM) 1 µl reverse primer (50 µM)

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2 µl of primary RT reaction Nuclease-free water up to 50 µl (0.5 µl Taq Polymerase) Use a “Hot Start” PCR: Start denaturation step of PCR. After 2–3 min pause the PCR and pipette 0.5 µl Taq and replace the tube in the PCR machine for the rest of the cycling. PCR cycles: We design RT-PCR primers as 23mers with a melting temperature of 68°. 94° 2 min 94° 30 s 65° 30 s 30 72° 1 min 72° 10 min 1 4° hold VI. Secondary nested PCR: 1 µl of first PCR reaction The first PCR should be diluted to avoid carrying over the outside primers and to dilute any spurious products. The first PCR can be diluted up to as much as 1 ml with water and 1 µl of this used in the secondary PCR. Primers can also be removed using Qiagen gel purification/PCR clean up columns but you should still only use 1/50th to 1/1000th of the primary PCR in the secondary reaction. 5 µl 10 PCR buffer 1 µl dNTP’s (10 mM) 1 µl forward primer (50 µM) 1 µl reverse primer (50 µM) Nuclease-free water up to 50 µl (0.5 µl Taq Polymerase) Use a Hot Start and PCR cycles as above. Analyse products on a 1.5–2% agarose gel. A typical result of morpholino splice site block is shown in Fig. 1. Wild-type mRNA is the predicted size while morpholino-induced mis-splicing results in a smaller RTPCR product. DNA from the gel can be isolated using standard gel purification techniques and sequenced directly using the inside nested reverse and forward RTPCR primers. Titration of the morpholino dose to the minimum necessary to produce complete absence of wild-type mRNA will avoid artifacts due to morpholino toxicity. Morpholino oligos can be expected to disrupt splicing effectively for up to 3 days of development. After that, wild-type mRNA typically reappears due to dilution or breakdown of the morpholino. After optimizing morpholino dosage, analysis of cilia structure and function can proceed using protocols listed below. Final confirmation of the specificity of morpholino knockdown (and validity of cilia phenotypes) ideally requires rescue of phenotypes by coinjection of synthetic wild-type mRNA along with the morpholino. For many genes this is

205

11. Studying Cilia in Zebrafish

spMO

(A) m

(B)

C

wt

24

48

IQ1

72

IQ2

spMO

696

Fig. 1 Disruption of mRNA splicing by splice donor blocking morpholino. (A) Analysis of nested RTPCR products amplifying exons encoding the C-terminus of inversin/NPHP2 (NM_152970) (Otto et al., 2003). In this experiment a morpholino was designed to block the exon 15 splice donor site to disrupt calmodulin binding domains in inversin. Control mRNA (C) shows the size of normal mRNA RT-PCR product and RNA prepared from morpholino-injected embryos (spMO) at 24 h post fertilization (hpf), 48 hpf and 72 hpf reveal a smaller RT-PCR product induced by mis-splicing. By 72 hpf in this case, wild-type mRNA recovers due to dilution of morpholino. (B) Sequencing of morpholino deletion RT-PCR products reveals that blocking the exon 15 splice donor forces use of a cryptic splice donor within the coding sequence of exon 15. This results in an out-of-frame fusion with exon 16 and introduction of a premature stop codon, deleting the C-terminal IQ2 calmodulin binding domain of the Inversin/NPHP2 protein.

not feasible since uniform (ectopic) expression of, for instance, potent morphogens, disrupts development. Fortunately, however, since most if not all cells are ciliated, uniform expression of a ciliogenic gene is not usually detrimental. Injection of 25–200 pg of mRNA can be tolerated by a zebrafish embryo. We use Ambion message machine kits for synthesis of capped mRNA from a plasmid template (http://www.ambion.com/). After in vitro synthesis of capped mRNA, dissolve mRNA in RNase free water and inject in 1– 2 cell stage morpholino embryos as described above.

B. Wholemount Confocal Imaging of Zebrafish Cilia A first-pass analysis of phenotypes caused by cilia gene disruption in zebrafish is to examine cilia structure by immunostaining. The commonly used

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anti-acetylated tubulin monoclonal antibody 6-11-B1 (Sigma T6793) works well to visualize cilia in zebrafish. Other useful commercially available monoclonals that work in zebrafish include anti-gamma tubulin (GTU-88) and anti-polyglutamyl tubulin (B3).

Anti-acetylated tubulin (clone 6-11b-1) Anti-polyglutamylated tubulin (clone B3) Anti-gamma-tubulin (clone GTU-88)

Sigma Sigma

T6793 T9982

Sigma

T6657

The protocol that follows is designed for use of these antibodies but can be adapted for other antibodies and other fixation methods.

1. Materials Phosphate Buffered Saline (PBS) DMSO Tween20 Normal Goat Serum (Sigma G9023) Bovine Serum Albumin (Sigma A8022 ) or Gelatin from cold water fish skin (Sigma G7765) Methanol Benzyl alcohol Glycerol N-propyl gallate Formaldehyde Hydrogen Peroxide SDS

2. Solutions PBST: Blocking solution:

Incubation solution:

Phosphate Buffered Saline þ 0.5% Tween20 Phosphate buffered Saline with 1% DMSO 0.5 % Tween20 1% BSA or 0.3% Gelatin from cold water fish skin 10% Normal Goat Serum Phosphate buffered Saline with 1% DMSO 0.5 % Tween20 2% Normal Goat Serum

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High-salt wash:

Phosphate buffered Saline with 1% DMSO 0.5 % Tween20 2% Normal Goat Serum 0.18% NaCl (final NaCl = 0.27%) Dent’s fixative: 80% Methanol 20% DMSO Formaldehyde (BT) fix 4% Formaldehyde (from paraformaldehyde) 0.1 M Phosphate Buffer pH7.2 3% Sucrose 0.12 mM CaCl2 Rehydration solutions: 75:25 MeOH/PBST 50:50 MeOH/PBST 25:75 MeOH/PBST Antigen retrieval solution: 1% SDS in Phosphate Buffered Saline Mounting medium: 53% Benzyl Alcohol (by weight) 45% Glycerol (by weight) and 2% N-Propyl Gallate

3. Methods One of the main advantages of using zebrafish for immunofluorescence is the transparency of their embryos. To achieve maximum embryo transparency, development of pigmentation can be blocked by raising embryos in 0.003% Phenylthiourea (PTU) egg water. Alternatively embryos can be bleached with hydrogen peroxide after fixation (see below).

4. Fixation 1. Fix embryos in Dent’s fixative for 3 h to overnight at room temperature. If necessary, remove pigment by bleaching fixed embryos overnight in 10% H2O2. After fixation embryos can be stored in 100% methanol at –20° . 2. Rehydrate Dent’s fixed embryos with graded changes of methanol PBT: 75:25 MeOH/PBT 15 min 50:50 MeOH/PBT 15 min 25:75 MeOH/PBT 15 min PBT Dent’s fixative works well for antibodies that preferentially recognize denatured epitopes. If formaldehyde fixation is preferable, antigen retrieval can be performed by denaturing the embryos after formaldehyde fixation. For formaldehyde fixation fix 2 h to overnight at 4° in BT fix and then wash twice in PBS. Permeabilize the embryos in cold acetone for 20 min at –20°. Warm to room temperature and wash with PBS. For antigen retrieval (denaturation), incubate fixed embryos in 1% SDS/PBS for 15 min at room temperature followed by washing with PBS 4 5 min each.

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Antibody staining: 3. Block non-specific binding by incubating fixed embryos for 2 h to overnight in Blocking solution. Incubations are done in eppendorf tubes at 4° on a nutator rocking platform. 4. Incubate with primary antibody (6-11-B1 at 1:1000) in incubation solution overnight at 4°. Monoclonal supernatants can be used at 1:50–1:25 dilution. Be aware that any primary antibody raised in rabbits should be affinity purified on antigen to avoid high background staining of larval fish skin. If necessary to economize on primary antibodies the incubation can be done in 50–100 µl without agitation. 5. After incubation with primary antibody, wash at least 4 30 min with incubation solution. 2% normal goat serum is included in all steps to reduce background. If background staining becomes a problem, the first wash after incubation with primary antibody can employ the “high-salt” wash. Subsequent washes are in the standard incubation solution. 6. Incubate with secondary antibody in incubation solution overnight. Alexa antimouse secondary antibodies (Invitrogen) work well at 1:1000 dilution. Following incubation, wash at least 4 30 min with Incubation solution on a rocking platform. Although background staining with secondary antibodies is not common, a highsalt wash can be used to minimize non-specific staining. Mounting the Sample for Confocal Microscopy To minimize optical distortion caused by mis-match in refractive index of the sample, coverslips, and immersion oil, use a mounting medium that has the same refractive index (1.513) as the immersion oil. We make a mounting medium developed by Gustafsson et al. (1999) that is a mixture of glycerol and benzyl alcohol and contains an antifade compound (N-propyl gallate) that is essential for preventing signal bleaching, especially when using 488 nm fluorophores (FITC, Alexa 488) for large Z image stacks. 7. Embryos can be placed in mounting medium directly after washing. The difference in density of the mounting medium and PBS is significant and causes turbulence but does not damage the sample. It is best to transfer embryos to mounting medium in a depression slide or directly on a microscope slide since the embryos become essentially invisible and can be hard to find in an eppendorf tube. Change to fresh mounting medium and transfer embryos to a standard microscope slide. Using small balls of modeling clay, support the edges of a coverslip to provide space for the embryo and coverslip the sample. Alternatively, make a coverslip bridge with additional coverslips as spacers. The orientation of the embryo can often be shifted by moving the coverslip. This sample configuration is suited for viewing with an upright microscope using a 63 oil immersion objective. Figure 2 shows the results of wholemount immunostaining zebrafish embryo cilia and basal bodies and imaging Z-series stacks by confocal microscopy. These images were generated using two different monoclonal antibodies (see alternate protocol below).

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Fig. 2 Wholemount confocal fluorescence imaging of zebrafish cilia. (A–C) Posterior spinal canal cilia stained with a combination of anti-acetylated tubulin (A; green) and anti-gamma tubulin (B; red). Merged image in (C) reveals orientation of cilia relative to basal bodies. (D–F) Olfactory placode motile multicilia stained with a combination of anti-acetylated tubulin (D; green) and anti-gamma tubulin (E; red). Merged image in (F) highlights the apical basal bodies in placode epithelial cells. These images were made using the double mouse monoclonal antibody modification of wholemount immunostaining. (See Plate no. 8 in the Color Plate Section.)

4. Alternate Protocols a. Visualization of Cilia with HRP/DAB. If visualization of cilia by light microscopy is desired, a horseradish peroxidase-coupled secondary antibody can be used. To visualize HRP after washing out the secondary antibody: 1. Wash once in PBSBT (PBS/0.1% Tween20/0.2% BSA) for 5 min 2. Incubate with 0.3 mg/ml DAB, 0.5% NiCl2 in PBSBT for 20 min. 3. Add H2O2 to 0.03% (1:1000 dilution of 30% stock) and monitor color development (check at 10 min). 4. Stop the reaction by rinsing in PBST and then PBS. Postfix in 2% formaldehyde/ 0.1% glutaraldehyde. b. Histological Sectioning of Wholemount Stained Embryos. Fluorescent signal from secondary antibodies is preserved in glycol methacrylate (JB4)-embedded samples. This allows for histological sectioning of wholemount stained embryos and viewing 3–10 µm sections by standard widefield epifluorescence. Longer wavelength excitation secondary antibodies, for instance Alexa 546 or rhodamine, should be used instead of FITC since fluorescein excitation wavelengths result in significant autofluorescence from the glycol methacrylate/JB4. 1. After the final wash after secondary antibody incubation, dehydrate embryos in sequential changes of 30, 50 75, 85, 95% ethanol. 2. Prepare catalyzed JB4 (Polysciences or Electron Microscopy Sciences) following the manufacturer’s instructions. Dissolve 0.625 g powdered catalyst to 50 ml

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3.

4.

5.

6.

monomer solution A in a 50 ml tube. Wrap the tube in foil to protect it from light and store it at 4°. The catalyzed JB4 solution can be used for 1–2 weeks. Remove final ethanol and add catalyzed JB4 solution to the dehydrated wholemount stained embryos. The embryos will float at the top of the solution and gradually sink as they become infiltrated. Leave at 4° overnight. Draw off the first solution of JB4 being sure to remove traces of ethanol that can inhibit hardening. Add the hardener (solution B) at 40 µl/ml to the embryos. Pour embryos in JB4 into a plastic mold, orient the embryos, and allow the JB4 to harden. Humidity and oxygen will inhibit hardening so this step is best done in a sealed, desiccated chamber. To embed embryos to cut them in cross section, first half fill a mold by pouring a bed of JB4 (0.7 ml) in the mold (15 mm 15 mm) and let it harden. Then add stained embryos in final catalyzed JB4 plus hardener and orient them to be headed to the side of the mold. The “half blocks” of JB4 adhere well even if polymerized separately and this allows for rotating and mounting the final block to have embryos facing the knife for sectioning. JB4 blocks require sectioning with glass knives. Use a Leica RM2255 or equivalent style microtome. JB-4 embedding Kit and embedding molds can be purchased from Polysciences or Electron Microscopy Supplies (EMS).

c. Double Immunofluorescent Staining of Wholemount Zebrafish Using Two Different Mouse-Monoclonal Antibodies. Cilia orientation and its control by planar polarity signaling is an important aspect of ciliogenesis (Park et al., 2008). To detect cilia orientation it is necessary to visualize the basal body and axoneme with separate fluorophores. The best antibody to visualize zebrafish basal bodies is also a mouse monoclonal GTU-88, anti-gamma tubulin, presenting a challenge for double staining with the monoclonal anti-acetylated tubulin. Also, detection of tubulin posttranslational modifications such as polyglutamylation or polyglycylation is best accomplished with mouse monoclonals coupled with anti-acetylated tubulin staining to demonstrate the presence and total length of axonemes. To allow for separate detection of two different mouse monoclonal antibodies in the same tissue, the following modification of the standard wholemount immunofluorescence protocol can be used. Additional Materials Required: Normal mouse serum (Sigma M5905) Goat anti-mouse Fab fragments (Jackson-ImmunoResearch 115-007-003) Methods 1. Process embryos through the first antibody incubation, detection with secondary antibody, and wash steps as described above. 2. Incubate embryos in 10% Normal mouse serum (Sigma M5905) in incubation solution for 2 h at 4°C. 3. Wash with incubation solution 4 30 min at 4°C

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4. Incubate embryos with goat anti-mouse Fab fragments at 1:20 dilution in incubation buffer; (Jackson-ImmunoResearch 115-007-003) overnight at 4°C. 5. Wash with incubation solution 4 30 min at 4°C 6. Block with Blocking solution 2 h at 4°C 7. Incubate with second primary antibody, detect with second fluorescent antibody, and wash as described above. Wash twice with PBS and mount in mounting medium for confocal microscopy. Combining wholemount in situ mRNA detection with immunocytochemistry for cilia Zebrafish embryos are well suited for analysis of cilia gene expression by wholemount in situ hybridization (Liu et al., 2007). mRNAs for ciliogenic genes are enriched in ciliated tissues, particularly in cells with multiple cilia like the olfactory placodes and the pronephric multiciliated cells (Pathak et al., 2007). Reliable protocols for wholemount in situ hybridization have been published elsewhere (Westerfield, 2000). In experiments where it is desirable to assess cilia structure and ciliogenic mRNA expression in the same cells, this can be accomplished by simultaneous incubation with antibodies to Digoxigenin (for in situ mRNA detection) and anti-acetylated tubulin (immunohistochemistry for cilia). Standard in situ hybridization protocols call for mRNA detection using Digoxigenin-labeled antisense mRNA probes and an antiDigoxigenin antibody detection step. Following incubation steps with antisense mRNA probe for your gene of interest and post-hybridization washes (Westerfield, 2000), incubate the embryos in Maleic Acid buffer (0.1 M Maleic Acid, 0.15 M NaCl, pH 7.5) with sheep anti-DIG-Alkaline phosphatase-conjugated antibody FAB fragments (1:5000 dilution; (Westerfield, 2000)) together with anti-acetylated tubulin antibody (6-11-B1, 1:1000 dilution) at 4°C overnight. Following detection of Alkaline phosphatase (Westerfield, 2000), incubate the embryos with HRP-coupled secondary antibody and detect with diaminobenzidine as described above.

C. An inexpensive System for High-Speed Microvideo of Cilia Measuring cilia beat rate and waveform requires high-speed microvideo imaging. For cilia beating at 25 Hz it would be necessary to capture video at 250 frames per second (fps) to have 10 frames for each single beat cycle. Dedicated systems are available to accomplish this task. For example IO industries VideoSavant (http://www. ioindustries.com/products/software/video_savant_4.html) and the Photron fastcam (http://www.photron.com/) have been optimized for high-speed microscopy. Camera systems and software range in price from about $5,000 to $30,000 or more. For labs who want to image cilia on a limited budget, we present a way to do it for about $1,200. We outline the method here but those who take this route are advised to read the manuals. The Dragonfly Express from Point Grey Research (http://www.ptgrey.com) is a Firewire camera that can deliver up to 350 fps at a resolution of 320 240 pixels. Designed primarily for machine vision and industrial imaging, this camera can be

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adapted for videomicroscopy since it can be supplied with a standard C-mount enclosure. The Dragonfly Express requires a standard IEEE-1394b (firewire 800) interface with the imaging computer. We have used this camera with Macintosh Quad G5 and Intel MacPro computers running Mac OS 10.5. It is also expected to work on Windows PCs (with different software) but we have not done this. The cost of the camera is approximately $1,000. We use ASC’s Astro IIDC software (http://www.outcastsoft.com/ASCASTROIIDC. html) to acquire high-speed Quicktime movies. This program costs $110. Astro IIDC is a flexible program used in both astronomy and microscopy and can acquire images at up to 220 fps. We use the “324 242 FMT7” mode to acquire and set the frame rates at 240 fps. Actual acquired frame rates are 220 fps (open the “Inspector” in Quicktime Player to confirm frame rate data). Once maximum frame rates have been achieved with a particular computer configuration, camera settings should be preserved and preferences can be set to acquire a fixed time period of video. For instance, we acquire at 220 fps and set preferences to record 220 frames to make each movie one second long (actual time). Astro IIDC opens Quicktime Player and generates a Quicktime movie and an associated metadata file listing acquisition parameters. Post processing movies is required to slow down the motion and resolve cilia beat rates. Since there does not seem to be inexpensive software packages available to do this on a Macintosh (as far as we know) we adapted Quicktime Pro for this purpose. Upgrading Quicktime Player to Quicktime Pro activates movie editing features and requires purchasing a Pro license (about $30) from Apple Computer. Built-in Quicktime Pro commands that alter movie frame rates are rather limited so to expand the timeline of a high-speed video, we first export it as an image sequence (individual tiff files for each frame) and reassemble it from the separate frames at a new lower frame rate. This gets cumbersome so we have automated the steps required using an Applescript. Below is an example of an Applescript that we use to slow a high-speed (220 fps) video to 15 fps: tell application "Finder" set tmpfilePath to do shell script "cd ~; pwd" do shell script "cd ~/Desktop" if exists folder "tmpFrames" then delete folder "tmpFrames" end if set theFolder to make new folder at desktop with properties {name:"tmpFrames"} end tell tell application "QuickTime Player" export document 1 to tmpfilePath & "/Desktop/tmpFrames/tmpframe" as image sequence using most recent settings open image sequence tmpfilePath & "/Desktop/tmpFrames/tmpframe 001.tif" frames per second 15

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end tell tell application "Finder" delete theFolder end tell tell application "QuickTime Player" set thefile to (choose file name) save self contained document 1 in thefile end tell This script requires that Quicktime Player is open and it will process only the active front Quicktime window. This script makes a temporary folder on the desktop to store movie frames, exports the frames of a movie currently open in Quicktime Player, reassembles the movie at 15 fps, deletes the temporary files, and saves the new movie. The script can be pasted into the Applescript Script editor, compiled, named and saved, and run as a script. To have it appear from a desktop menu, activate “Show Script menu in menu bar” in the Applescript utility and save the script to “/Library/Scripts/” (as administrator). Measuring Cilia Beat Rate and Amplitude Using ImageJ Cilia beating can often be counted visually in slow motion movies but to avoid error it is best to use image processing software. ImageJ is a free image processing program available at http://rsbweb.nih.gov/ij/. We use ImageJ to “reslice” movies to generate a graphical representation of cilia beating. 1. Open a Quicktime movie in ImageJ 2. Select the line tool from the toolbar and draw a line on your movie that crosses images of beating cilia. Drawing a line in the center of a tubule lumen for instance will allow sampling of multiple cilia. 3. From the “Image” menu, choose “Stacks” and “Reslice”. 4. Check the box for “Rotate 90°”; click OK. You will get a new window that represents the single row of pixels under the line you drew set side by side for each frame. This produces a graphical representation of cilia motility that can be correlated with the time scale of the movie to derive beat rate as well as represent beat amplitude. Again, these methods are for high-speed imaging on the smallest budget. Other systems have been built to accomplish these tasks such as the one described by Michael Sanderson at University of Massachusetts Medical School (http://users. umassmed.edu/michael.sanderson/mjslab/Cilia_Page.htm).

D. Electron Microscopy Methods for Fish Cilia Electron microscopy is a standard method required for assessing cilia structural defects. Electron microscopy on zebrafish is performed essentially the same as for other vertebrate tissues but with some alterations that accommodate the lower osmolarity of fish extracellular fluids. Other modifications include use of tannic acid in the

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fixative to enhance contrast and use of partially reduced osmium that produces a less grainy final image. Materials (All available from Electron Microscopy Sciences) Glutaraldehyde Paraformaldehyde Phosphate or Cacodylate buffer Sucrose Tannic Acid Methods 1. Fix overnight at 4° in 1.5% Glutaraldehyde 1% Paraformaldehyde 70 mM NaPO4 pH 7.2 3% Sucrose 0.1% Tannic Acid

7.5 ml 10% stock 3.125 ml 16% stock 3.5 ml 1 M stock 5 ml 30% stock 0.05 g Water to 50 ml

EMS (Electron Microscopy Sciences) premade stocks can be used for the aldehydes; these come in sealed glass vials. Addition of tannic acid enhances the final contrast of the image; particularly of microtubules and other filamentous structures. Add the tannic acid just before fixation; it does not keep well. The buffer is slightly lower osmolarity than a fixative for mammalian tissue. 2. Wash 3 5 min in 0.1 M cacodylate or phosphate buffer, pH 7.4 Avoid the use of mechanical rocking or mixing during the fixation and washes; the overall concern is to be as gentle as possible to the tissue to preserve structure. 3. Post Fix in 1% OsO4 plus potassium ferrocyanide one hour on ice: 1 ml 4% OsO4 3 ml H2O 0.06 g. Potassium Ferrocyanide This is partially reduced osmium; it makes for less grainy images. If desired the potassium ferrocyanide may be omitted. 4. Wash 2 10 min in 0.1 M cacodylate or phosphate buffer, pH 7.4 5. In bloc stain fixed embryos in 1% Uranyl Acetate in 0.1 M cacodylate pH 7.4 for 1 h at Room Temperature. 6. Wash 3 10 min in 0.1 M cacodylate or phosphate buffer, pH 7.4 7. Dehydrate in 25%, 50% 70% 80% 95% 100% EtOH for 20–30 min each 8. Dehydrate with 2 100% EtOH 20–30 min each 9. Infiltrate with Propylene Oxide for 15–20 min (in a fume hood). 10. Mix the Epon 812 (or substitute epoxy embedding medium) according to manufacturers’ instructions.

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25% 300 (25% Epon in propylene oxide) 50% 400 75% Overnight 100% 4h 100% 1h 12. Embed embryos in fresh Epon in flat molds. Position embryo swimming toward the tip of the block for acquiring cross sections. 13. Harden at 60° for 16 h.

11. Infiltrate with

The blocks are now ready for sectioning and staining. We use formvar-coated slot grids (Electron microscopy Sciences; catalog # FF2010-Cu) to maximize visibility of the tissue in the sections. Accurate diagnosis of cilia defects requires direct orthogonal viewing; i.e. the perfect cross section. Tangential sections are not useful unless your electron microscope is equipped with a stage goniometer for tipping the grid. For less able configurations the best approach is to orient the fish embryo to maximize the chance of finding a clear cross section. The easiest way to find a cross section is to image pronephric cilia in cross section. Most pronephric cilia are constrained by the tubule lumen and can be seen in cross sections over the yolk and yolk extension.

II. Summary The methods summarized here provide a foundation for using the zebrafish to characterize the function of genes important in ciliogenesis and cilia function. Clearly this is just a subset of experimental approaches to study ciliogenesis. Although much remains to be done to fully develop the system, the zebrafish has already proven useful for revealing the function of novel ciliogenic genes. Higher resolution analysis of phenotypes including imaging IFT processes, and immunoEM of cilia/basal body components promises to keep the fish at the forefront of cilia research.

Acknowledgments I thank past and present members of my laboratory for their contributions to this work. This work was supported by NIH grants DK053093, and DK070263 to I.A.D and by grants from the PKD foundation.

References Bill, B.R., Petzold, A.M., Clark, K.J., Schimmenti, L.A., and Ekker, S.C. (2009). A primer for morpholino use in zebrafish. Zebrafish 6, 69–77. Brand, M., Heisenberg, C.P., Warga, R.M., Pelegri, F., Karlstrom, R.O., Beuchle, D., Picker, A., Jiang, Y.J., Furutani-Seiki, M., van Eeden, F.J., Granato, M., Haffter, P., et al. (1996). Mutations affecting development of the midline and general body shape during zebrafish embryogenesis. Development 123, 129–142.

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Iain Drummond Drummond, I.A., Majumdar, A., Hentschel, H., Elger, M., Solnica-Krezel, L., Schier, A.F., Neuhauss, S.C., Stemple, D.L., Zwartkruis, F., Rangini, Z., Driever, W., and Fishman, M.C. (1998). Early development of the zebrafish pronephros and analysis of mutations affecting pronephric function. Development 125, 4655– 4667. Eisen, J.S., and Smith, J.C. (2008). Controlling morpholino experiments: don’t stop making antisense. Development 135, 1735–1743. Essner, J.J., Amack, J.D., Nyholm, M.K., Harris, E.B., and Yost, H.J. (2005). Kupffer’s vesicle is a ciliated organ of asymmetry in the zebrafish embryo that initiates left-right development of the brain, heart and gut. Development 132, 1247–1260. Foley, J.E., Yeh, J.R., Maeder, M.L., Reyon, D., Sander, J.D., Peterson, R.T., and Joung, J.K. (2009). Rapid mutation of endogenous zebrafish genes using zinc finger nucleases made by Oligomerized Pool ENgineering (OPEN). PLoS ONE 4, e4348. Gustafsson, M.G., Agard, D.A., and Sedat, J.W. (1999). I5M: 3D widefield light microscopy with better than 100 nm axial resolution. J. Microsc. 195, 10–16. Kramer-Zucker, A.G., Olale, F., Haycraft, C.J., Yoder, B.K., Schier, A.F., and Drummond, I.A. (2005). Ciliadriven fluid flow in the zebrafish pronephros, brain and Kupffer’s vesicle is required for normal organogenesis. Development 132, 1907–1921. Liu, Y., Pathak, N., Kramer-Zucker, A., and Drummond, I.A. (2007). Notch signaling controls the differentiation of transporting epithelia and multiciliated cells in the zebrafish pronephros. Development 134, 1111–1122. MacDonald, R.J., Swift, G.H., Przybyla, A.E., and Chirgwin, J.M. (1987). Isolation of RNA using guanidinium salts. Methods Enzymol. 152, 219–227. Malicki, J., Neuhauss, S.C., Schier, A.F., Solnica-Krezel, L., Stemple, D.L., Stainier, D.Y., Abdelilah, S., Zwartkruis, F., Rangini, Z., and Driever, W. (1996). Mutations affecting development of the zebrafish retina. Development 123, 263–273. Meng, X., Noyes, M.B., Zhu, L.J., Lawson, N.D., and Wolfe, S.A. (2008). Targeted gene inactivation in zebrafish using engineered zinc-finger nucleases. Nat. Biotechnol. 26, 695–701. Obara, T., Mangos, S., Liu, Y., Zhao, J., Wiessner, S., Kramer-Zucker, A.G., Olale, F., Schier, A.F., and Drummond, I.A. (2006). Polycystin-2 immunolocalization and function in zebrafish. J. Am. Soc. Nephrol. 17, 2706–2718. Omori, Y., Zhao, C., Saras, A., Mukhopadhyay, S., Kim, W., Furukawa, T., Sengupta, P., Veraksa, A., and Malicki, J. (2008). Elipsa is an early determinant of ciliogenesis that links the IFT particle to membraneassociated small GTPase Rab8. Nat. Cell Biol. 10, 437–444. Otto, E.A., Schermer, B., Obara, T., O’Toole, J.F., Hiller, K.S., Mueller, A.M., Ruf, R.G., Hoefele, J., Beekmann, F., Landau, D., Foreman, J.W., Goodship, J.A., et al. (2003). Mutations in INVS encoding inversin cause nephronophthisis type 2, linking renal cystic disease to the function of primary cilia and left-right axis determination. Nat. Genet. 34, 413–420. Ou, G., Qin, H., Rosenbaum, J.L., and Scholey, J.M. (2005). The PKD protein qilin undergoes intraflagellar transport. Curr. Biol. 15, R410–R411. Owens, K.N., Santos, F., Roberts, B., Linbo, T., Coffin, A.B., Knisely, A.J., Simon, J.A., Rubel, E.W., and Raible, D.W. (2008). Identification of genetic and chemical modulators of zebrafish mechanosensory hair cell death. PLoS Genet. 4, e1000020. Park, T.J., Mitchell, B.J., Abitua, P.B., Kintner, C., and Wallingford, J.B. (2008). Dishevelled controls apical docking and planar polarization of basal bodies in ciliated epithelial cells. Nat Genet. 40, 871–879. Pathak, N., Obara, T., Mangos, S., Liu, Y., and Drummond, I.A. (2007). The zebrafish fleer gene encodes an essential regulator of cilia tubulin polyglutamylation. Mol. Biol Cell. 18, 4353–4364. Pazour, G.J. (2004). Intraflagellar transport and cilia-dependent renal disease: The ciliary hypothesis of polycystic kidney disease. J. Am. Soc. Nephrol. 15, 2528–2536. Serluca, F.C., Xu, B., Okabe, N., Baker, K., Lin, S.Y., Sullivan-Brown, J., Konieczkowski, D.J., Jaffe, K.M., Bradner, J.M., Fishman, M.C., and Burdine, R.D. (2009). Mutations in zebrafish leucine-rich repeatcontaining six-like affect cilia motility and result in pronephric cysts, but have variable effects on left-right patterning. Development 136, 1621–1631.

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Sukumaran, S., and Perkins, B.D. (2009). Early defects in photoreceptor outer segment morphogenesis in zebrafish ift57, ift88 and ift172 Intraflagellar Transport mutants. Vision Res. 49, 479–489. Sullivan-Brown, J., Schottenfeld, J., Okabe, N., Hostetter, C.L., Serluca, F.C., Thiberge, S.Y., and Burdine, R.D. (2008). Zebrafish mutations affecting cilia motility share similar cystic phenotypes and suggest a mechanism of cyst formation that differs from pkd2 morphants. Dev. Biol. 314, 261–275. Sun, Z., Amsterdam, A., Pazour, G.J., Cole, D.G., Miller, M.S., and Hopkins, N. (2004). A genetic screen in zebrafish identifies cilia genes as a principal cause of cystic kidney. Development 131, 4085–4093. Tsujikawa, M., and Malicki, J. (2004). Intraflagellar transport genes are essential for differentiation and survival of vertebrate sensory neurons. Neuron 42, 703–716. van Rooijen, E., Giles, R.H., Voest, E.E., van Rooijen, C., Schulte-Merker, S., and van Eeden, F.J. (2008). LRRC50, a conserved ciliary protein implicated in polycystic kidney disease. J. Am. Soc. Nephrol. 19, 1128–1138. Veland, I.R., Awan, A., Pedersen, L.B., Yoder, B.K., and Christensen, S.T. (2009). Primary cilia and signaling pathways in mammalian development, health and disease. Nephron. Physiol. 111, p39–p53. Westerfield, M. (2000). The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Danio Rerio).” University of Oregon Press, Eugene. Wienholds, E., van Eeden, F., Kosters, M., Mudde, J., Plasterk, R.H., and Cuppen, E. (2003). Efficient targetselected mutagenesis in zebrafish. Genome Res. 13, 2700–2707. Zhao, C., and Malicki, J. (2007). Genetic defects of pronephric cilia in zebrafish. Mech Dev. 124, 605–616.

CHAPTER 12

Analysis of IFT Kinesins in Developing Zebrafish Cone Photoreceptor Sensory Cilia Christine Insinna*, Katherine Luby-Phelps*,†, Brian A. Link* and Joseph C. Besharse* * Department of Cell Biology, Neurobiology and Anatomy, Medical College of Wisconsin, Milwaukee, Wisconsin 53226-0509 †

Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, Texas 75390-9039

Abstract Introduction Comparison of Photoreceptor Model Systems Zebrafish as a Model for Analysis of Photoreceptor IFT Mutant Zebrafish Antisense Morpholino Oligonucleotides Transient Transgenesis Using Photoreceptor-Specific Promoters Materials and Methods A. Cloning of IFT Constructs Using the Gateway® Technology B. Microinjections C. Screening for GFP Expression D. Fluorescence Analysis VIII. Transmission Electron Microscopy Acknowledgments References I. II. III. IV. V. VI. VII.

Abstract The photoreceptor outer segment (OS), a well-defined sensory cilium, provides an important context for the study of intraflagellar transport (IFT). The early phases of OS development involve successive events that are common to virtually METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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all cilia. Additionally, intense protein trafficking occurs through the cilium and relies on IFT to maintain proper cellular morphology and optimize the photosensitive function. In the past decade, progress has been made in the characterization of photoreceptor OS trafficking in murine and amphibian models. Recently, powerful and cost-effective molecular tools and techniques for zebrafish have opened new opportunities to study photoreceptor IFT. Studies using zebrafish take advantage of its rapid embryogenesis to characterize the early events involved in photoreceptor ciliogenesis and OS assembly. In this overview, we describe phenotypes associated with knockdown strategies or genetic mutations of IFT components in zebrafish and detail a general experimental approach that has enabled us to study the function of the two anterograde IFT motors, KIF17 and kinesin II, in zebrafish cone photoreceptors.

I. Introduction The photoreceptor outer segment (OS) is a modified sensory cilium that requires intraflagellar transport (IFT) for the transport of components essential for its complex architecture and function in phototransduction. The cilium extends from its basal body to form a transition zone, common to all cilia, and its membrane forms an expanded distal domain that accumulates the photopigment, opsin, and other transduction components as a prelude to disc formation (Fig. 1). Although there is a great deal of focus on the transition zone (Besharse and Horst, 1990; Rohlich, 1975), generally referred to as the connecting cilium, the photoreceptor ciliary axoneme extends distally into the OS (Kaplan et al., 1987; Sale et al., 1988) where it terminates as singlet microtubules (Insinna et al., 2008b; Steinberg and Wood, 1975). There is also substantial evidence suggesting that additional singlet microtubules are present in the distal OS (Eckmiller, 1996; Eckmiller, 2000; Roof et al., 1991). During OS formation the ciliary plasmalemma of photoreceptors becomes organized into disc membranes that align perpendicular to the axoneme, which extends distally along one side of the OS (Fig. 1). The discs of rod and cone photoreceptors differ in that those of rods are contained in the cytoplasmic compartment and are separated from the OS plasmalemma, while most discs of cones maintain continuity with the plasma membrane (Young, 1976). This fundamental design difference suggests distinct mechanisms of OS disc formation between rods and cones. However, at this early stage of analysis the same fundamental model for IFT applies equally well for both rods and cones. Historically, tremendous progress has been made in understanding the signaling mechanisms underlying phototransduction within the OS (Burns and Arshavsky, 2005). In contrast, the mechanisms underlying assembly and renewal of the OS have remained obscure. However, it is now clear that IFT plays a critical role in OS assembly because mutations in genes encoding both IFT proteins (Pazour et al., 2002) and IFT molecular motors (Jimeno et al., 2006; Krock et al., 2009; Marszalek et al., 2000) cause severe ablation or disruption of OS assembly and

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Dynein KIF17 Kinesin II IFT particle Cargo

Fig. 1 A current model for photoreceptor IFT. The photoreceptor axoneme extends from a basal body anchored in the IS and extends distally with membrane discs aligned perpendicular to it. The anterograde motors KIF17 and kinesin II carry IFT particles and cargo distally and the cytoplasmic dynein 2 motor returns IFT particles and kinesin motors. Two potential turn around points for switching between anterograde and retrograde IFT are indicated by the curved arrows. The role of KIF17 in the distal OS along singlet microtubules is modeled after C. elegans sensory cilia.

maintenance. Two features of photoreceptors make them a particularly valuable model for the analysis of IFT function. First, a key problem in IFT research is the identification of specific IFT cargo, and it has been suggested that elements of phototransduction machinery may be directly transported by IFT. Since phototransduction proteins are relatively abundant and the biochemical tools for their analysis are well-developed, photoreceptors offer a particularly useful model for the analysis of cargo loading and transport. For example, we have recently reported that

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membrane guanylyl cyclase 1 (Gucy2E), a critical element in the phototransduction cascade, can be isolated in a cargo complex that includes IFT proteins and the kinesin II motor (Bhowmick et al., 2009). Second, the unusual organization of the axoneme in photoreceptors has led to the realization that at least two separate IFT kinesins are required for OS assembly (Insinna and Besharse, 2008). The current model for photoreceptor IFT (Fig. 1) involves IFT protein complexes like those in other types of cilia, but is divergent in that it includes two kinesin 2 family motors, heterotrimeric kinesin II and homodimeric KIF17, for transport of proteins from the site of synthesis in the inner segment (IS) to the OS. As in other ciliary IFT models cytoplasmic dynein 2 is thought to serve as the retrograde motor (Krock et al., 2009). The involvement of two kinesin 2 motors in photoreceptors is analogous to models from Caenorhabditis elegans, where both kinesin II and osm-3, a Kif17 homolog, can support ciliogenesis (Pan et al., 2006). The finding that OSM-3 is required for distal elongation of singlet microtubules in C. elegans (Snow et al., 2004) and the fact that vertebrate photoreceptors have distal singlets, led us to look for a possible role of KIF17 in photoreceptor IFT and to the finding that it is essential for OS elongation (Insinna et al., 2008b). Although we have made some progress in understanding important features of IFT in photoreceptors, much remains to be discovered about the molecular details.

II. Comparison of Photoreceptor Model Systems Current animal models for the analysis of photoreceptor OS trafficking include mice, Xenopus laevis, and zebrafish (Danio rerio), but the three models have different sets of advantages and disadvantages. Current technology, which enables both the production of transgenic mice as well as targeted deletions or “knockins” along with the ability to conditionally mutate genes specifically in photoreceptors, has made mice a model of choice. However, mouse models require extensive development time and are expensive. Both X. laevis and zebrafish offer less expensive alternatives. Transgenic X. laevis using a Xenopus opsin promoter to drive transgene expression has been used to great advantage to study cell biological pathways in rod OS formation (Knox et al., 1998; Moritz et al., 2001; Tam et al., 2004). Such studies have recently included targeting of green fluorescent protein (GFP)-tagged IFT proteins into rod OS (Luby-Phelps et al., 2008). In principle, this approach could be extended to cone OS development in Xenopus (Moritz et al., 2002). However, an ever-widening array of induced gene mutations along with the use of antisense morpholino knockdown techniques and transient transgenesis make zebrafish particularly valuable for analysis OS development. A limitation of both Xenopus and zebrafish has been the inability to target mutations to specific genes. However, a mutational strategy involving zinc-finger nucleases has recently been introduced that shows promise for the widespread introduction of targeted germ line mutations in zebrafish (Meng et al., 2008).

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III. Zebrafish as a Model for Analysis of Photoreceptor IFT From a methodological perspective the rapid embryogenesis of the zebrafish retina and the multiple molecular tools available for its analysis make it a compelling model for the study of IFT in early photoreceptor OS development. For example, loss-offunction studies in zebrafish of ift88, ift57, and ift172 (Sukumaran and Perkins, 2009; Tsujikawa and Malicki, 2004) as well as morpholino knockdowns of ift52, ift57, and kif17 all resulted in failure or disruption of OS formation with little or no effect on the IS (Insinna et al., 2008b; Krock and Perkins, 2008; Tsujikawa and Malicki, 2004). In zebrafish, the expression of rod and cone opsins precedes the formation of the OS at 2.5 days postfertilization (dpf) (Raymond et al., 1995). Although at 3 dpf a small set of rod OS are located ventral to the optic nerve, the majority (~90%) of the retina contains cone cells whose OS develop dramatically between 3 and 5 dpf. By 12 dpf five photoreceptor types (one rod and four cones) become fully differentiated (Kljavin, 1987). An additional, and highly useful, feature of early photoreceptor development in zebrafish is the fact that neurogenesis persists at the retinal periphery where progenitor cells continue to give rise to retinal cell types including new photoreceptors. This means that early developing (older) photoreceptors are located near the retinal center and are developmentally more advanced than the newly emerging photoreceptors at the periphery. The antisense and transient transgenic approaches available for early zebrafish development are particularly valuable for the analysis of the role of IFT in cone OS formation between 3 and 5 dpf. In this chapter, we describe an approach for analysis of IFT in zebrafish that has provided compelling evidence that IFT is a critical mechanism for OS formation. In addition, these approaches have led to the idea that two different kinesin 2 family members, heterotrimeric kinesin II and homodimeric KIF17, play distinct and important roles in photoreceptor IFT. In this work we have made use of antisense morpholino oligonucleotides directed against kinesin heavy chains as well as dominant negative forms of kinesin 2 family members (DNKIF17 and DNKIF3B). These studies have also taken advantage of a late-onset cone transducin a (Ta-CP) promoter originally developed by Breandan Kennedy (Kennedy et al., 2007) to target the expression of dominant negative constructs and GFP-tagged components of the IFT system in cone photoreceptors. The value of this approach is that it specifically targets cone photoreceptors during a critical phase (3–5 dpf) of early OS elongation when cones represent about 90% of the photoreceptors present in the retina.

IV. Mutant Zebrafish Two screens using chemical or insertional mutagenesis have identified a number of mutations in IFT proteins and three of these (Table I) that have been studied in photoreceptors. Two of the mutations (IFT88 and IFT172) cause severe ablation of the OS, while the mutation of IFT57 causes a less severe OS assembly defect (Krock

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Table I Summary of Photoreceptor OS Phenotypes Associated with Components of the IFT Machinery in Zebrafish IFT component

Mutation or treatment

OS phenotype

References

IFT protein IFT88

Complex B

Mutation

No OS

IFT172 IFT57

Complex B Complex B

Mutation Mutation

No OS Reduced OS

IFT20 IFT57 IFT52 IFT140

Complex B Complex B Complex B Complex A

Morpholino Morpholino Morpholino Morpholino

Reduced OS Reduced OS Reduced OS Mild effect; reduced Zop-GFP expression

Kinesins KIF17 KIF3B Kif17 KIF3B

Homodimeric Heterotrimeric Homodimeric Heterotrimeric

Morpholino Morpholino Dominant Negative Dominant Negative

No OS Early lethality Disrupted OS morphogensis Disrupted IS; ribbon synapses not formed

Insinna Insinna Insinna Insinna

Dynein 2 DYNC2H1 DYNC2LI1 DYNC2I1

Dynein 2 heavy chain Dynein 2 light intermediate chain Dynein 2 intermediate chain

Morpholino Morpholino Morpholino

Disorganized, short OS Disorganized, short OS Disorganized, short OS

Krock et al. (2009) Krock et al. (2009) Krock et al. (2009)

Sukumaran and Perkins (2009), Tsujikawa and Malicki (2004) Sukumaran and Perkins (2009) Krock and Perkins, (2008), Sukumaran and Perkins (2009) Tsujikawa and Malicki (2004) Tsujikawa and Malicki (2004) Tsujikawa and Malicki (2004) Tsujikawa and Malicki (2004)

et al. et al. et al. et al.

(2008) (2009) (2009) (2009)

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and Perkins, 2008; Sukumaran and Perkins, 2009; Tsujikawa and Malicki, 2004). In each case the OS defect is phenocopied through injection of antisense morpholino oligonucleotides. These findings along with the earlier work in mice showing that an IFT88 mutation (Pazour et al., 2002) or more recent work showing that conditional deletion of IFT20 (Keady, B.T., Le, Y., and Pazour, G.J., personal communication) in cones impair OS formation, provide a compelling case that IFT is critical for photoreceptor OS formation. It is of some interest that all mutant IFT proteins studied at this level in photoreceptors are components of the anterograde IFT complex B. Although zebrafish mutations in the IFT motors or in the retrograde IFT complex A have not been reported, antisense and dominant negative expression studies show that each plays a critical role (see below).

V. Antisense Morpholino Oligonucleotides Antisense morpholino oligonucleotides directed at either an intron–exon splice junction or the translation start site are widely used in zebrafish for knockdown of gene expression in early embryogenesis. Since the morpholino is generally introduced by microinjection at the one cell stage, the titration of the morpholino during rapid cell division of early embryogenesis or by ongoing target gene expression limits their effect to early embryonic stages. The early differentiation of cone OS by 3–5 dpf makes this approach feasible for analysis of OS IFT. For example, morpholinos directed at IFT88, IFT172, and IFT57 for the most part phenocopy the effects of mutations in those same genes. On the other hand, morpholino oligonucleotides provide the principal evidence for the functions of KIF17, dynein 2, and the IFT complex A protein IFT140 in photoreceptors (see Table I). Although morpholino knockdown of KIF17 causes a strong OS ablation phenotype (Insinna et al., 2008b) comparable to mutations in IFT88 and IFT172 (Sukumaran and Perkins, 2009; Tsujikawa and Malicki, 2004), knockdown of the subunits of the dynein 2 motor and IFT140 reportedly exhibit milder phenotypes (Krock et al., 2009; Tsujikawa and Malicki, 2004). Since these observations pertain to the earliest stages of development in which the cone OS is growing in length, the milder effect of components associated with retrograde IFT is not surprising. Comparison of the effects of antisense morpholino oligonucleotides for the knockdown of KIF17 or the KIF3B subunit of kinesin II illustrate both the power and potential ambiguities associated with this antisense approach from a cell biological perspective. On the positive side, an antisense morpholino directed at kif17 resulted in dramatic knockdown of KIF17 protein and revealed a failure of OS formation in 3 dpf zebrafish embryos. This was a relatively specific effect in that embryos in general were indistinguishable from controls and the retina was well differentiated with normal lamination. More recently, we have found that the KIF17 morpholino effect on photoreceptors is rescued by the coexpression of GFP-tagged KIF17 (unpublished). These combined results indicate that KIF17 is essential for OS formation. Although OS formation was severely ablated, the pronephros was normal and kidney epithelial cilia

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elongated normally (Insinna et al., 2008b). This implies that KIF17 plays a role in OS elongation that is not shared in kidney cilium elongation. A more subtle effect on olfactory cilium development cannot be ruled out. In contrast to the KIF17 results, morpholinos directed at either a splice site or the translation start site of the KIF3B subunit of kinesin II gave more ambiguous results, possibly due to a more widespread requirement for KIF3B in early zebrafish embryogenesis (Insinna et al., 2009). Both KIF3B morpholinos resulted in a high level of embryonic mortality and in surviving embryos caused severe developmental defects in multiple tissues including retina. The retina failed to laminate normally or to form a photoreceptor layer, which precluded the analysis of photoreceptor OS formation for comparison to KIF17 morphants. Attempts to minimize the effects by reducing the concentration of injected morpholino led to an increased survival rate and an absence of a phenotype in the retina. The effects of KIF3B knockdown were much more severe than morpholino knockdowns reported for KIF17 (see above), the dynein 2 retrograde motor complex (Krock et al., 2009), or IFT proteins (Krock and Perkins, 2008; Sukumaran and Perkins, 2009; Tsujikawa and Malicki, 2004), suggesting that KIF3B may play a critical role in early embryonic events distinct from ciliogenesis or IFT. Although consistent with an early, essential function for KIF3B in retinal development, these data could also be explained by potential off-target effects, another possible caveat with morpholino experiments. Standard controls for both efficacy and specificity of knockdown have been recently reviewed (Eisen and Smith, 2008; Robu et al., 2007).

VI. Transient Transgenesis Using Photoreceptor-Specific Promoters Zebrafish transient transgenesis provides a rapid and ideal method for confining the effects of transgene expression specifically to photoreceptors. Typically, plasmid constructs containing a cell-specific promoter upstream of an open reading frame of interest is injected at the one cell stage and expression of the transgene is studied during early embryogenesis. The technique is referred to as “transient transgenesis” because gene expression generally occurs transiently during embryogenesis and often in a mosaic pattern. The zebrafish rod opsin (Zop) (Kennedy et al., 2001) and Ta-CP (Kennedy et al., 2007) promoters provide an opportunity to drive transgene expression in the majority of either rod or cone photoreceptors during early embryogenesis. For example, expression of Zop-IFT52-GFP causes GFP expression specifically in rod cells (Fig. 2). We have used this approach in both X. laevis (Luby-Phelps et al., 2008) and zebrafish (Fig. 2) to study the distribution of GFP-tagged IFT proteins as a prelude to real-time imaging of their translocation. In both species IFT52-GFP is found in cilium-like structures associated with OS elongation. Although the latter finding is consistent with specific OS trafficking of IFT proteins, the strong overexpression driven by these robust promoters causes GFP expression in all cell compartments including the synaptic terminal.

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(A)

(B)

Fig. 2 A cluster of developing rods expressing an IFT52-GFP fusion protein under the control of the Zop promoter. Z-slices taken from an intact, anesthetized 8 dpf larva using a Leica SP2 confocal microscope Leica Microsystems, Inc, Deerfield, IL, USA and a 60W water immersion lens. Arrow points to region of a basal body and arrowheads point to axonemes within the OS. A and B are different z-slices from the same set of cells. (See Plate no. 9 in the Color Plate Section.)

A key question in the current model for photoreceptor IFT relates to the relative roles of KIF17 and kinesin II. Since mutants are not currently available and our comparison using morpholinos was confounded by ubiquitous knockdown of KIF3B and the limited window of efficacy (Insinna et al., 2008b), we used an alternative strategy involving overexpression of dominant negative forms of KIF17 (DNKIF17) and kinesin II (DNKIF3B) proteins in which GFP replaced the motor domain (Insinna et al., 2009). Since 90% of the early developing photoreceptors are cones, we placed these GFP-tagged proteins downstream of the cone Ta-CP promoter that is known to induce the expression of transgenes in the majority of cones at 3 dpf (Fig. 3). At this stage, cones at the central retina have mid- to adult-sized OS and most of the cells at the periphery have a very short OS. We used confocal microscopy to image the localization of transgenes in cone cells expressing high levels of GFP (Fig. 3) and transmission electron microscopy (TEM) to characterize the ultrastructural defects in cones at the central and peripheral retina (Fig. 4). As expected, transgenes were expressed specifically in cones and this approach offered a considerable advantage over morpholinos by restricting the dominant negative phenotypes to cone cells. DNKIF17 had effects that were largely restricted to the OS, causing severe OS vesiculation among older photoreceptors in the retinal center and blockade of OS elongation in peripheral photoreceptors (Fig. 4). Strikingly, IS and synaptic pedicles were like those of wild-type larvae in these experiments. In contrast, DNKIF3B caused extensive cell death among cones throughout the retina, which was preceded by formation of IS vacuoles (Fig. 4). Furthermore, cone synapses were significantly immature and deficient in synaptic ribbons. In general, these results suggest that KIF17 functions principally in OS assembly because the TEM phenotypes were restricted to OS defects. The failure to complete ribbon synapses and IS defects in cells expressing DNKIF3B, however, suggest that KIF3B plays roles in IS membrane trafficking and in formation of the synaptic pedicle in addition to its role in IFT. Since

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(A)

(B)

OS IS ONL OPL

OPL INL GCL

(C) OS

IS

N

S

Fig. 3 Widespread expression of DNKIF3B-GFP under the control of the zebrafish Ta-CP promoter in the eye of a 4 dpf larva. (A) Most cones across the entire retina express the transgene. (B) The transgene is abundant in the IS outer nuclear layer (ONL), and outer plexiform layer (OPL), but does not enter the OS. (C) Diagram illustrating the distribution of the transgene (green) in photoreceptor compartments. Images in (A) and (B) were made using the Leica SP2 confocal microscope. (See Plate no. 10 in the Color Plate Section.)

IS membrane-trafficking events are upstream of the IFT pathway, those defects would be expected to dominate. Although not yet fully realized, the potential for directly studying protein and membrane trafficking in developing photoreceptor cells is great for the zebrafish model. For example, we have recently used the Ta-CP promoter to drive expression of nonphosphorylatable and phosphomimetic KIF17 mutant proteins in zebrafish cones (Insinna et al., 2008a). This work was based on the finding that calciumcalmodulin kinase II specifically phosphorylates KIF17 at a serine residue near its C-terminus (Guillaud et al., 2008). In hippocampal dendrites phosphorylation resulted in release of cargo. However, in the retina we found that KIF17 phosphorylation was required for its association with IFT proteins and entry into the OS. Furthermore, phosphomimetic KIF17 drove increased OS assembly and turnover. These studies benefited from the combination of the transient transgenesis approach, the Ta-CP promoter for cone-specific expression, and the MultiSite Gateway® Technology for cloning of expression constructs.

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Periphery

Center

OS

OS OS IS

IS IS

IS

OS

OS OS

IS

IS

IS

IS

DNKIF17

DNKIF3B

DNKIF17

DNKIF3B

Fig. 4

Images illustrating the structure of cones expressing DNKIF17 or DNKIF3B in either the central or peripheral retina. In the center DNKIF17 causes disruption and vesiculation (asterisks) within the OS, while KIF3B causes disruption of the IS (asterisks). At the periphery DNKIF17 prevents formation of the OS (asterisk), while DNKIF3B causes vesiculation of the IS (asterisk) that does not prevent early OS formation.

Finally, it should be mentioned that the transparency of the organism coupled with ease of transgenesis contribute to the promise of live cell imaging in photoreceptors. For example, by blocking formation of melanin pigment in the eye by treatment with 1-phenyl-2-thiourea we have been able to image developing photoreceptors in living larvae (Fig. 2). This, along with the fact that others have carried out in vivo imaging of subcellular trafficking in other cells of the zebrafish (Baye and Link, 2007; Laughlin et al., 2008), suggests that the tools are now available to study photoreceptor IFT in individual photoreceptors. Application of these techniques using the photoreceptorspecific expression systems described here should enable a much more complete understanding of the role of IFT motors within the OS.

VII. Materials and Methods A. Cloning of IFT Constructs Using the Gateway® Technology The site-specific recombinational cloning system offered by Invitrogen’s MultiSite Gateway® Technology is an ideal system to create the constructs necessary for the study of photoreceptor IFT. We have used PCR fragments containing transgenes of interest (DNKIF17 and DNKIF3B) fused to GFP (Chu et al., 2006; Le Bot et al., 1998;

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Lin-Jones et al., 2003) and site-specific recombination sites in the BP (attB and attP sites) recombination reaction to create the entry clones. We used a similar approach to create the second entry clone containing the zebrafish cone Ta-CP promoter (Kennedy et al., 2007). The Ta-CP and both DNKIF3B and DNKIF17 entry clones were then recombined with pcDNA 6.2/V5-pL-DEST vectors (Invitrogen, Carlsbad, CA, USA) to create the final Ta-CP-DNKIF3B or Ta-CP-DNKIF17 expression clones used in the experiments. Gateway amendable constructs that incorporate transposon-mediated transgenesis for multiple types of epitope tagging experiments and for conditional gene expression have now been generated (Kwan et al., 2007). B. Microinjections Microinjection of one-cell embryos is a routine procedure for the introduction of both morpholinos and expression plasmids. Embryos are collected in a Petri dish containing fish water and transferred to a new dish with fish water to remove any residual debris. One-cell stage embryos are sorted away from older embryos and divided into separate dishes for each construct to be injected. Plasmid constructs are suspended (25 ng/µl) in water plus 0.1% phenol red (Sigma, Saint Louis, MO, USA) in an Eppendorf tube and kept on ice until the injection system is set up. Using a Nanoject II injector (Drummond Scientific, Broomall, PA, USA) a glass needle is filled with a few microliters of the solution which is injected at a volume of ~5.0 nl per cell. Following injection, embryos lacking the traces of phenol red dye are discarded. This first selection step increases the probability of success. Injected and uninjected control embryos are kept at 28°C in an incubator. Rapid formation of melanin pigment in the choroid and retinal pigment epithelium of the eye is a potential impediment for imagining photoreceptors. In experiments where this is an issue, pigment development can be blocked by placing embryos in a dish containing fish water supplemented with 0.003% 1-phenyl-2-thiourea (PTU). Fish water is made by dissolving 0.06 g Instant Ocean sea salts (Spectrum Brands Company, Atlanta, GA, USA) in 1.0 l of deionized water (H2O) and buffering the pH to 7.2 using sodium bicarbonate powder (NaHCO3). Embryos can remain in PTU–fish water until 3–4 dpf. C. Screening for GFP Expression When using GFP-tagged expression constructs such as DNKIF3B (Fig. 3) embryos can be screened for GFP fluorescence in the eyes at 3–4 dpf. Embryos are simply transferred into a Petri dish containing 0.05% Tricaine in embryo medium to anesthetize the fish. The dish is placed on the stage of an epifluorescence microscope (Eclipse TE-300, Nikon, Tokyo, Japan) and eyes are scanned with a 10 objective; observation of uninjected embryos may be necessary to determine the level of background autofluorescence. Embryos selected for GFP expression can be retained for further development or fixed and processed for either fluorescence or electron microscopic analysis.

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D. Fluorescence Analysis We use a fixation protocol that permits direct observation of GFP in frozen sections as well as immunocytochemistry for other cell constituents such as the cone photopigment (opsin) or microtubules. Embryos are fixed in 4% paraformaldehyde in phosphatebuffered saline (PBS) (pH 7) for 1 h at room temperature or overnight at 4°C and washed three times in PBS prior to cryoprotection. For cryoprotection the embryos are incubated in successive solutions of 15 and 30% sucrose in water for 1 h each at 4°C. Sucrose is subsequently replaced with optimal cutting temperature (OCT) medium (Miles Scientific, Elkhart, IN, USA) and incubated overnight at 4°C. The next day wood spatulas are used to transfer the OCT-infiltrated embryos into transparent Tissue Tek cryomolds (EM Sciences, Hatfield, PA, USA) containing OCT medium. A sharp pin is used to align the tip of the embryo heads perpendicularly to one side of the cryomold; multiple embryos are aligned parallel to each other along the same side of the cryomold. They are then frozen in the cryomold at –20°C, and 10- to 12-µm transverse sections are cut in a rostrocaudal direction using a cryostat. Sections are monitored using a dissecting scope to obtain those from the central portion of the eye. Air-dried sections are either first stained with an antibody to an antigen such as cone opsin or directly placed in Fluoromount G media (EM Sciences, Hatfield, PA, USA) and covered with a cover glass (24 60-1 Premium Cover Glass, Fisher Scientific, Pittsburg, PA, USA). The localization of GFP or the antigen of interest is determined in the photoreceptor layer using a confocal laser scanning microscopy with a 63 objective. When using the Ta-CP promoter most cone cells will express the transgene at low levels across the entire photoreceptor layer, while approximately 15–30% of the cones will express GFP at a high level.

VIII. Transmission Electron Microscopy Since the 1950s photoreceptors from a wide range of species including zebrafish have been the subject of numerous TEM studies and their overall architecture is well defined. The polarized organization of photoreceptors separates their synaptic terminals, nuclear compartment, and IS, and OS into separate regions when viewed longitudinally. Thus, TEM is a method of choice for defining cellular defects that arise from disruption of gene expression through mutations, morpholinos, or dominant negative constructs. The general techniques for conventional TEM have been well defined over the years and vary principally in the details of the fixation protocol and embedment medium. In our recent work on zebrafish (Insinna et al., 2008b, 2009), embryos were fixed overnight at 4°C in a mixture of 2% glutaraldehyde and 2% paraformaldehyde at pH 7.4 in Sorenson’s phosphate buffer at a final concentration of 0.1 M. A volume of 100 ml of 2 Sorenson’s buffer is prepared by mixing 81 ml of 0.2 M Na2HPO4 and 19 ml of NaH2PO4 and adjusting pH to 7.4. After washing out the aldehyde fixative, embryos are postfixed for 1.5–2 h in 1% OsO4 in the same buffer on ice in the ventilated hood of the EM facility. After washing three times for 10 min in Sorenson’s buffer, the tissue is dehydrated in increasing concentrations of methanol up to 100%

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and then transferred to acetonitrile and infiltrated with embedding resin. We currently use a mixture consisting of 25 ml Epon 812, 20 ml Araldyte 502, 60 ml DDSA to which 1.1 ml of DMP-30 is added (1%); these products are all from EM Sciences. The next day embryos are embedded in flat embedding molds (PELCO, Ted Pella, Inc., Redding, CA, USA) and hardened by placing in an oven at 60–70°C overnight or until the resin is polymerized. Sections from these blocks are suitable for both light and electron microcopy. For light microscopy, cut 1-µm sections rostrocaudally with an ultramicrotome and stain sections with toluidine blue. For electron microscopy thin sections (70 nm) from the same block are stained with uranyl acetate and lead citrate and placed on nickel EM grids. These are then imaged in our facility using either the Hitachi 600 (Pleasanton, CA, USA) or JEOL JEM 2100 (Tokyo, Japan) electron microscope. Since the Ta-CP promoter drives the expression of dominant negative IFT motors with a late onset (3 dpf), a wide range of phenotypes can be found in cones at the central retina that have a well-formed OS.

Acknowledgments The authors wish to thank Dr Don Arnold for the DNKIF17 (USC, Los Angeles, CA), Dr Beth Burnside (UC Berkeley, CA) for the DNKIF3B constructs, and Dr Breandan Kennedy (UCD, Ireland) for the Zop and Ta-CP promoters. We also thank Clive Wells for assistance with EM imaging and Monica Humby for technical assistance. This work was supported by NIH grant EY03222 (JCB) and an NIH core grant for Vision Research. C. Insinna was supported by NIH NRSA Training grant T32-EY014537.

References Baye, L.M., and Link, B.A. (2007). Interkinetic nuclear migration and the selection of neurogenic cell divisions during vertebrate retinogenesis. J. Neurosci. 27, 10143–10152. Besharse, J.C., and Horst, C.J. (1990). The photoreceptor connecting cilium. A model for the transition zone. In “Ciliary and Flagellar Membranes” (R.A. Bloodgood, ed.), pp. 389–417. Plenum Publishing Corp, New York. Bhowmick, R., Li, M., Sun, J., Baker, S.A., Insinna, C., and Besharse, J.C. (2009). Photoreceptor IFT complexes containing chaperones, guanylyl cyclase 1, and Rhodopsin. Traffic 10, 648–663. Burns, M.E., and Arshavsky, V.Y. (2005). Beyond counting photons: Trials and trends in vertebrate visual transduction. Neuron 48, 387–401. Chu, P.J., Rivera, J.F., and Arnold, D.B. (2006). A role for Kif17 in transport of Kv4.2. J. Biol Chem. 281, 365–373. Eckmiller, M.S. (1996). Renewal of the ciliary axoneme in cone outer segments of the retina of Xenopus laevis. Cell Tissue Res. 285, 165–169. Eckmiller, M.S. (2000). Microtubules in a rod-specific cytoskeleton associated with outer segment incisures. Vis. Neurosci. 17, 711–722. Eisen, J.S., and Smith, J.C. (2008). Controlling morpholino experiments: Don’t stop making antisense. Development 135, 1735–1743. Guillaud, L., Wong, R., and Hirokawa, N. (2008). Disruption of KIF17-Mint1 interaction by CaMKIIdependent phosphorylation: A molecular model of kinesin-cargo release. Nat. Cell Biol. 10, 19–29. Insinna, C., and Besharse, J.C. (2008). Intraflagellar transport and the sensory outer segment of vertebrate photoreceptors. Dev. Dyn. 237, 1982–1992.

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Insinna, C., Humby, M., Scoma, H., and Besharse, J.C. (2008a). A role for CaMKII-mediated phosphorylation of KIF17 in photoreceptor IFT. In “Annual Meeting of American Society for Cell Biology.” Online Abstracts. 1033. Full length abstracts at: http://www. ascb.org/meetings/. Insinna, C., Humby, M., Sedmak, T., Wolfrum, U., and Besharse, J.C. (2009). Different roles for KIF17 and kinesin II in photoreceptor development and maintenance. Dev. Dyn., 238, 2211–2222. Insinna, C., Pathak, N., Perkins, B., Drummond, I., and Besharse, J.C. (2008b). The homodimeric kinesin, Kif17, is essential for vertebrate photoreceptor sensory outer segment development. Dev. Biol. 316, 160– 170. Jimeno, D., Feiner, L. Lillo, C., Teofilo, K., Goldstein, L.S., Pierce, E.A., and Williams, D.S. (2006). Analysis of kinesin-2 function in photoreceptor cells using synchronous Cre-loxP knockout of Kif3a with RHO-Cre. Invest. Ophthalmol. Vis. Sci. 47, 5039–5046. Kaplan, M.W., Iwata, R.T., and Sears, R.C. (1987). Lengths of immunolabeled ciliary microtubules in frog photoreceptor outer segments. Exp. Eye Res. 44, 623–632. Kennedy, B.N., Alvarez, Y., Brockerhoff, S.E., Stearns, G.W., Sapetto-Rebow, B., Taylor, M.R., and Hurley, J.B. (2007). Identification of a zebrafish cone photoreceptor-specific promoter and genetic rescue of achromatopsia in the nof mutant. Invest. Ophthalmol. Vis. Sci. 48, 522–529. Kennedy, B.N., Vihtelic, T.S., Checkley, L., Vaughan, K.T., and Hyde, D.R. (2001). Isolation of a zebrafish rod opsin promoter to generate a transgenic zebrafish line expressing enhanced green fluorescent protein in rod photoreceptors. J. Biol. Chem. 276, 14037–14043. Kljavin, I.J. (1987). Early development of photoreceptors in the ventral retina of the zebrafish embryo. J. Comp. Neurol. 260, 461–471. Knox, B.E., Schlueter, C., Sanger, B.M. Green, C.B., and Besharse, J.C. (1998). Transgene expression in Xenopus rods. FEBS 423, 117–121. Krock, B.L., Mills-Henry, I., and Perkins, B. (2009). Retrograde intraflagellar transport by cytoplasmic dynein-2 is required for outer segment extension in vertebrate photoreceptors but not arrestin translocation. Invest. Ophthalmol. Vis. Sci. [Epub ahead of print] Krock, B.L., and Perkins, B.D. (2008). The intraflagellar transport protein IFT57 is required for cilia maintenance and regulates IFT-particle-kinesin-II dissociation in vertebrate photoreceptors. J. Cell Sci. 121, 1907–1915. Kwan, K.M., Fujimoto, E., Grabher, C., Mangum, B.D., Hardy, M.E., Campbell, D.S., Parant, H.J.M., Yost, J., Kanki, J.P., and Chien, C.B. (2007). The Tol2kit: A multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev. Dyn. 236, 3088–3099. Laughlin, S.T., Baskin, J.M., Amacher, S.L., and Bertozzi, C.R. (2008). In vivo imaging of membraneassociated glycans in developing zebrafish. Science 320, 664–667. Le Bot, N., Antony, C., White, J., Karsenti, E., and Vernos, I. (1998). Role of xklp3, a subunit of the Xenopus kinesin II heterotrimeric complex, in membrane transport between the endoplasmic reticulum and the Golgi apparatus. J. Cell Biol. 143, 1559–1573. Lin-Jones, J., Parker, E., Wu, M., Knox, B.E., and Burnside, B. (2003). Disruption of Kinesin II function using a dominant negative-acting transgene in Xenopus laevis rods results in photoreceptor degeneration. Invest. Ophthalmol. Vis. Sci. 44, 3614–3621. Luby-Phelps, K., Fogerty, J., Baker, S.A., Pazour, G.J., and Besharse, J.C. (2008). Spatial distribution of intraflagellar transport proteins in vertebrate photoreceptors. Vision Res. 48, 413–423. Marszalek, J.R., Liu, X., Roberts, E.A., Chui, D., Marth, J.D., Williams, D.S., and Goldstein, L.S. (2000). Genetic evidence for selective transport of opsin and arrestin by kinesin-II in mammalian photoreceptors. Cell 102, 175–187. Meng, X., Noyes, M.B., Zhu, L.J., Lawson, N.D., and Wolfe, S.A. (2008). Targeted gene inactivation in zebrafish using engineered zinc-finger nucleases. Nat. Biotechnol. 26, 695–701. Moritz, O.L., Peck, A., and Tam, B.M. (2002). Xenopus laevis red cone opsin and Prph2 promoters allow transgene expression in amphibian cones, or both rods and cones. Gene 298, 173–182. Moritz, O.L., Tam, B.M., Hurd, L.L., Peranen, J., Deretic, D., and Papermaster, D.S. (2001). Mutant rab8 Impairs docking and fusion of rhodopsin-bearing post-Golgi membranes and causes cell death of transgenic Xenopus rods. Mol. Biol. Cell. 12, 2341–2351.

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Christine Insinna et al. Pan, X., Ou, G., Civelekoglu-Scholey, G., Blacque, O.E., Endres, N.F., Tao, L., Mogilner, A., Leroux, M.R., Vale, R.D., and Scholey, J.M. (2006). Mechanism of transport of IFT particles in C. elegans cilia by the concerted action of kinesin-II and OSM-3 motors. J. Cell. Biol. 174, 1035–1045. Pazour, G.J., Baker, S.A., Deane, J.A., Cole, D.G., Dickert, B.L., Rosenbaum, J.L., Witman, G.B., and Besharse, J.C. (2002). The intraflagellar transport protein, IFT88, is essential for vertebrate photoreceptor assembly and maintenance. J. Cell. Biol. 157, 103–113. Raymond, P.A., Barthel, L.K., and Curran, G.A. (1995). Developmental patterning of rod and cone photoreceptors in embryonic zebrafish. J. Comp. Neurol. 359, 537–550. Robu, M.E., Larson, J.D., Nasevicius, A., Beiraghi, S., Brenner, C., Farber, S.A., and Ekker, S.C. (2007). p53 activation by knockdown technologies. PLoS Genet. 3, e78. Rohlich, P. (1975). The sensory cilium of retinal rods is analogous to the transitional zone of motile cilia. Cell Tissue Res. 161, 421–430. Roof, D., Adamian, M., Jacobs, D., and Hayes, A. (1991). Cytoskeletal specializations at the rod photoreceptor distal tip. J. Comp. Neurol. 305, 289–303. Sale, W.S., Besharse, J.C., and Piperno, G. (1988). Distribution of acetylated a-tubulin in retina and in in vitro-assembled microtubules. Cell Motil. Cytoskeleton 9, 243–253. Snow, J.J., Ou, G.,Gunnarson, A.L., Walker, M.R., Zhou, H.M., Brust-Mascher, I., and Scholey, J.M. (2004). Two anterograde intraflagellar transport motors cooperate to build sensory cilia on C. elegans neurons. Nat. Cell Biol. 6, 1109–1113. Steinberg, R.H., and Wood, I. (1975). Clefts and microtubules of photoreceptor outer segments in the retina of the domestic cat. J. Ultrastruct. Res. 51, 307–403. Sukumaran, S., and Perkins, B.D. (2009). Early defects in photoreceptor outer segment morphogenesis in zebrafish ift57, ift88 and ift172 Intraflagellar Transport mutants. Vision Res. 49, 479–489. Tam, B.M., Moritz, O.L., and Papermaster, D.S. (2004). The C terminus of peripherin/rds participates in rod outer segment targeting and alignment of disk incisures. Mol. Biol. Cell 15, 2027–2037. Tsujikawa, M., and Malicki, J. (2004). Intraflagellar transport genes are essential for differentiation and survival of vertebrate sensory neurons. Neuron 42, 703–716. Young, R.W. (1976). Visual cells and the concept of renewal. Invest. Ophthalmol. Vis. Sci. 15, 700–725.

CHAPTER 13

Analysis of Intraflagellar Transport in C. elegans Sensory Cilia Limin Hao*, Seyda Acar*, James Evans*, Guangshuo Ou† and Jonathan M. Scholey* *

Molecular and Cell Biology, University of California, Davis, California 95616

†

Department of Cellular and Molecular Pharmacology, University of California, San Francisco, California 94107

Abstract I. Introduction A. Intraflagellar Transport (IFT) and Cilium Biogenesis on C. elegans Chemosensory Neurons B. Genetic Screens of C. elegans IFT Mutants C. Examination of Ciliary Structure D. In Vivo IFT Motility Assay E. Biochemical Analysis of IFT Motors and Particles II. Rationale A. Genetic Screens of C. elegans IFT Mutants B. Examination of Ciliary Structure C. In Vivo IFT Motility Assay D. Purification of Heterologously Expressed Motor Proteins and In Vitro Motility Assay III. Methods A. Maintenance of WT and dyf Mutant Worms B. Genetic Screens of dyf Mutants and Dyf Assay C. Examination of Ciliary Structure D. Microscopy for In Vivo IFT Motility Assay and Cilia Imaging and Cilia Length Measurement E. Expression and Purification of Kinesin-II and Gliding Motility Assay IV. Materials A. Maintenance of WT and dyf Mutant Worms B. Genetic Screen of dyf Mutants and Dyf Assay C. Examination of Ciliary Structure D. In Vivo IFT Motility Assay and Cilia Imaging E. Purification of Kinesin-II and Gliding Motility Assay METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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V. Discussion A. Genetic Screen of IFT Mutants B. Examination of Ciliary Structure C. In Vivo IFT Motility Assay D. In Vitro Gliding Motility Assay VI. Summary Acknowledgments References

Abstract Cilia are assembled and maintained by intraflagellar transport (IFT), the motordependent, bidirectional movement of multiprotein complexes, called IFT particles, along the axoneme. The sensory cilia of Caenorhabditis elegans represent very useful objects for studying IFT because of the availability of in vivo time-lapse fluorescence microscopy assays of IFT and multiple ciliary mutants. In this system there are 60 sensory neurons, each having sensory cilia on the endings of their dendrites, and most components of the IFT machinery operating in these structures have been identified using forward and reverse genetic approaches. By analyzing the rate of IFT along cilia within living wild-type and mutant animals, two anterograde and one retrograde IFT motors were identified, the functional coordination of the two anterograde kinesin-2 motors was established and the transport properties of all the known IFT particle components have been characterized. The anterograde kinesin motors have been heterologously expressed and purified, and their biochemical properties have been characterized using MT gliding and single molecule motility assays. In this chapter, we summarize how the tools of genetics, cell biology, electron microscopy, and biochemistry are being used to dissect the composition and mechanism of action of IFT motors and IFT particles in C. elegans.

I. Introduction A. Intraflagellar Transport (IFT) and Cilium Biogenesis on C. elegans Chemosensory Neurons Cilia (also termed flagella) consist of microtubule (MT)-based axonemes surrounded by a specialized membrane that project from the surface of most eukaryotic cells, where they play vital roles in cell motility, sensory perception, and signaling. The assembly, maintenance, and functions of cilia depend on IFT, a process during which MT-based kinesin and dynein motors drive the bidirectional movement of multiprotein complexes called IFT particles, together with associated cargo molecules, between the base and the tip of the axoneme (Rosenbaum and Witman, 2002; Scholey, 2003). IFT was originally observed in Chlamydomonas reinhardtii (Kozminski et al., 1993) and

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subsequently the IFT particles were isolated and found to consist of two subparticles, IFT subcomplex A (IFT-A) and IFT subcomplex B (IFT-B). IFT-A was resolved into 6 polypeptides and IFT-B into 11 polypeptides (Cole et al., 1998; Piperno and Mead, 1997). The motors that drive anterograde IFT belong to the kinesin-2 family, the founding member of which was identified in sea urchin eggs and embryos as a heterotrimeric, plus-end directed MT-based motor that is required for ciliogenesis (Cole et al., 1993; Morris and Scholey, 1997). It is now known that heterotrimeric kinesin-2 motors (e.g., kinesin-II and the KIF3 complex) drive IFT in most cilia, but its activity can be augmented by homodimeric kinesin-2 motors (e.g., OSM-3 and Kif17) in cilia that contain singlet axoneme extensions (Cole et al., 1998; Hirokawa et al., 2006; Insinna and Besharse, 2008; Scholey, 2008; Snow et al., 2004). Retrograde IFT is driven by the cytoplasmic dynein class 1b, also known as IFT dynein (Schafer et al., 2003; Scholey, 2008; Signor et al., 1999a). The Caenorhabditis elegans genome encodes orthologs of most of the IFT particle and motor polypeptide components (Hao and Scholey, 2009). C. elegans has 60 ciliated sensory neurons whose ciliated dendritic endings are arranged into distinct sensory organs, two amphids in the head and two phasmids in the tail, which are amenable to observations of IFT and sensory ciliogenesis. The sensory cilia on these neurons are approximately 7-µm long and are differentiated longitudinally into three domains: the transition zone, the middle segment, and the distal segment, within which the axonemes display distinct morphologies. The transition zone harbors the basal body which consists of nine triplet microtubules. The middle segment consists of nine doublet microtubules and the distal segment consists of nine singlet microtubules (Evans et al., 2006; Perkins et al., 1986). The techniques available in C. elegans, most notably forward and reverse genetics, genetic manipulations that permit fluorescent protein labeling and worm transformation, and in vivo IFT motility assays, are allowing researchers to study the composition and mechanism of action of IFT motors, IFT particles, and associated IFT components in considerable detail (Fig. 1).

B. Genetic Screens of C. elegans IFT Mutants Following Sydney Brenner’s pioneering work, which was aimed at understanding the genetic control of animal development and behavior using C. elegans as a model system, a variety of behavioral mutants were isolated by ethyl methane sulfonate (EMS)-mediated mutagenesis and were found to affect behaviors such as chemotaxis (che) (Lewis and Hodgkin, 1977), osmotic avoidance (osm) (Dusenbery et al., 1975), and dauer larva formation (daf) (Riddle, 1977). Electron microscopic reconstruction of the sensory neurons in the head of the animal revealed that some of these mutants exhibited structural defects of sensory cilia located on the dendritic endings of sensory neurons (Perkins et al., 1986). Dye-filling assays demonstrated that the mutants displaying defective cilia structure could not absorb fluorescent dye from the surrounding environment into their amphids and phasmids, whereas the

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(A)

(B)

Fig. 1 The IFT machinery of C. elegans sensory cilia. (A) All the IFT particle components are arranged into three complexes: IFT-A, IFT-B, and BBSome complex. The components in the circles in IFT-A and IFTB correspond to those originally isolated from C. reinhardtii. Other subunits were observed to move together with these subunits undergoing IFT. The arrangement of the BBS components in the BBSome complex is based on Nachury et al. (2007). (B) The subunit compositions of the two anterograde IFT motors, the heterotrimeric kinesin-II and homodimeric OSM-3, and the retrograde motor IFT dynein from C. elegans. IFT dynein probably contains additional light and intermediate subunits.

wild type could do so (Perkins et al., 1986). In an independent dyf (dye-filling defective) mutant screen, 13 dyf mutants representing new genetic loci were isolated, and the molecular lesions responsible for the mutant phenotype associated with most of them have subsequently been elucidated (Table I) (Starich et al., 1995). During the same year that IFT was discovered in C. reinhardtii, and the IFT motor, kinesin-2 was purified from sea urchin embryos, osm-3, was cloned as a ciliary mutant and was deduced to encode a kinesin-like protein (Shakir et al., 1993). Subsequently, the osm-6 mutant was cloned (Collet et al., 1998) and found to encode an IFT52 homologue, a subunit of the IFT-B subcomplex (Cole et al., 1998; Piperno and Mead, 1997). Thus, early C. elegans genetic analysis of animal behavior provided a foundation for the subsequent analysis of ciliary biogenesis and IFT using this organism. To complement forward genetics, transposon insertion mutagenesis, chemical and UV-TMP (trimethylpsoralen) mutagenesis, and PCR-based protocols have been applied to the isolation of deletion alleles of any given genes in C. elegans (Jansen et al., 1997; Liu et al., 1999; Zwaal et al., 1993). This reverse genetic approach is particularly critical for studying sensory ciliogenesis and IFT because the RNAi technique which is so effective in other systems has not been shown to robustly knockdown genes in ciliated sensory neurons, even in sensitized mutant backgrounds. Moreover, the C. elegans gene knockout consortium facility uses UV-TMP mutagenesis and PCR screens to generate mutations in any gene of interest, in response to researchers’ requests (http://celeganskoconsortium.omrf.org/ or http://www.shigen.nig. ac.jp/c.elegans/).

Name che-2 che-3 che-11 che-13 daf-10 daf-19 dyf-1 dyf-2 dyf-3 dyf-4 dyf-5 dyf-6 dyf-7 dyf-9 dyf-10 dyf-11 dyf-12 dyf-13 osm-1 osm-3 osm-5 osm-6 osm-12

Other name

osm-2, che-8, avr-1, caf-2 che-9 osm-4 daf-24

dyf-8

caf-1, klp-2

bbs-7

Gene model/Genetic position (cM)

che

daf

dyf

osm

Function

References

F38G1.1 F18C12.1 C27A7.4 F59C6.7 F23B2.4 F33H1.1a F54C1.5a ZK520.1 C04C3.5a V:4.31 ± 0.29 M04C9.5 F46F6.4 C43C3.3 V:24.22 ± 0.35 I:1.53 ± 0.040 C02H7.1 X:2.18 ± 0.07 C27H5.7a T27B1.1 M02B7.3a Y41G9A.1 R31.3

þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ

þ þ þ þ þ þ / / / / / / þ þ / / þ þ þ þ þ þ

þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ

þ þ þ þ þ þ þ / / / / / þ / / / / / þ þ þ þ

IFT-particle B IFT-dynein heavy chain IFT-particle A IFT-particle B IFT-particle A RFX family transcription factor OSM-3-kinesin activator IFT-particle IFT-particle B associated

Fujiwara et al. (1999) Wicks et al. (2000) Qin et al. (2001) Haycraft et al. (2003) Bell et al. (2006) Swoboda et al. (2000) Ou et al. (2005b) Efimenko et al. (2006) Murayama et al. (2005)

MAP kinase IFT-particle Cell-surface ligand

Burghoorn et al. (2007) Bell et al. (2006) Heiman and Shaham (2009)

IFT-particle B associated

Bacaj et al. (2008)

Y75B8A.12

þ

/

þ

þ

Ciliary distal segment assembly IFT-particle B IFT-kinesin IFT-particle B IFT-particle B Part of ciliary distal segment assembly

Blacque et al. (2005) Bell et al. (2006) Shakir et al. (1993) Haycraft et al. (2001) Collet et al. (1998) Blacque et al. (2004)

13. Analysis of Intraflagellar Transport in C. elegans Sensory Cilia

Table I C. elegans Ciliary Mutants

Keys: che, chemotaxis assay; daf, daufer formation assay; dyf, dye-filling assay; osm, osmotic avoidance assay.

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C. Examination of Ciliary Structure The cilia present on the dendrites of C. elegans chemosensory neurons are fasciculated and localized along the channel formed by two glia-like cells, namely the sheath cell and socket cell. These channel cilia are in direct contact with the environment, which facilitates the sensing of environmental cues (Lewis and Hodgkin, 1977; Ward et al., 1975). When the worms are immersed in a lipophilic fluorescent dye, for example, FITC or DiI, a subset of the channel cilia uptake the dye, producing fluorescently stained sensory neurons. Consequently, sensory neurons that fail to uptake the fluorescent dye have perturbations of their ciliary structure, often due to defects in cilium assembly or maintenance. The ultrastructural analysis of such dyf mutants by transmission electron microscopy (TEM) has indeed revealed all kinds of ciliary structural defects (Perkins et al., 1986). The subsequent molecular analysis of these mutants demonstrated that many of them are caused by mutations in IFT components. Serial section TEM can be employed to confirm that morphology defects of IFT motor mutants observed using light microscopy reflect a change in the ultrastructure of the amphid channel cilia (Chalfie and Thomson, 1982; Evans et al., 2006; Perkins et al., 1986). While the acquired images only represent a snapshot of the cross-sectional view every 50–150 nm along their length, a model of the path of each cilium can be generated and used to help understand the role of specific gene products in IFT and cilium biogenesis. Aligning images of serial sections corresponding to the first 10 µm from the tip of the head of the worm allows the tracking and modeling of individual cilia from their neuronal transition zone to their exposed endings at the amphid pore (Fig. 2A). Additionally, if the micrographs are recorded on film or CCD at a sufficient resolution (~2 nm/pixel) then the architecture of each axoneme can also be determined with the MTs arranged either as singlets, doublets, or triplets (Fig. 2A). Reconstruction of serial transmission sections of the worm’s head allows researchers to observe the morphology and the length of the cilia, the architecture of the axoneme, and other details. In most cases though, mutations in IFT genes cause significant shortening of cilia which can be seen by introducing a fluorescently labeled cilia marker into the mutant allowing observations of cilium length under fluorescence microscopy (Fig. 2B). By correlating results from light and TEM, it has been shown that two kinesin-2 motors, homodimeric OSM-3 kinesin and heterotrimeric kinesin II, function in a partially redundant manner to build full-length amphid channel cilia but are completely redundant for building full-length amphid wing (AWC) cilia. This difference apparently reflects cilia-specific differences in OSM-3 activity, which serves to extend distal singlets in channel cilia but not in AWC cilia which lack such singlets and respond to different stimuli (Chalfie and Thomson, 1982; Evans et al., 2006; Perkins et al., 1986). D. In Vivo IFT Motility Assay Using time-lapse fluorescence microscopy, the movement of specifically labeled IFT proteins along the sensory cilium can be readily visualized in C. elegans. First,

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green fluorescent protein (GFP)-labeled kinesin-II (tagged on the KAP-1 subunit) and OSM-6, an IFT-B component, were observed to move along the cilium (Orozco et al., 1999). The same technique was subsequently applied to other IFT particle components, leading to the identification and characterization of a number of IFT-A and IFTB components such as OSM-1, OSM-5, CHE-2, CHE-11, DAF-10 (Qin et al., 2001). Later, several more IFT particle components, including IFTA-1 (Blacque et al., 2006), DYF-2 (Efimenko et al., 2006), and CHE-13 (Haycraft et al., 2003) were identified and characterized (Fig. 1A). Because the fluorescent signal emitted by GFP-tagged IFT components accumulated in the ciliary tips of che-3 and xbx-1 mutants, these latter two genes were identified as subunits of dynein 1b, the retrograde IFT motor (Schafer et al., 2003; Signor et al., 1999a) (Fig. 1B). With the introduction of spinning disc confocal microscopy and kymography, the IFT assay became a routine method for studying the mechanism of IFT in this system (Snow et al., 2004) (Fig. 3). In addition to kinesin-II, OSM-3 was found to contribute to transport along the cilium by coordinating with kinesin-II to transport IFT particles along the middle segment and to move along and build the distal segment on its own (Evans et al., 2006; Snow et al., 2004). Kinesin-II is a slow motor, moving at 0.5 µm/s in the absence of OSM-3, whereas OSM-3 is a fast motor, moving at 1.3 µm/s in the absence of kinesin-II (Snow et al., 2004). In wild-type animals, kinesin-II and OSM-3 cooperate to move the same IFT particles along the middle segment at a velocity intermediate between those of the individual motors (0.7 µm/s) whereas OSM-3 moves along the distal singlets on its own at its characteristic fast velocity (1.3 µm/s). By analyzing the rates of IFT in bbs (Bardet–Biedl syndrome) mutants, it was found that kinesin-II moves at its characteristic slow rate (0.5 µm/s) and OSM-3 moves at its characteristic fast rate (1.3 µm/s) along the axoneme middle segment, suggesting that BBS complex is responsible for the coordination of the two anterograde IFT motors (Ou et al., 2005b). Several GFP-tagged IFT-A components move at the slow kinesin-II rate (0.5 µm/s) whereas IFT-B components move at the fast OSM-3 rate (1.3 µm/s), indicating that kinesin-II carries the IFT-A subcomplex and OSM-3 carries the IFT-B subcomplex (Ou et al., 2005b; Pan et al., 2006) (Table II). In combination with the analysis of dyf mutants, four novel IFT-B components, DYF-1 (Ou et al., 2005b), DYF-3 (Murayama et al., 2005; Ou et al., 2005a), DYF-11 (Omori et al., 2008), and DYF-13 (Blacque et al., 2005) were identified and characterized; these components had been missed using the powerful approach of C. reinhardtii biochemistry, underscoring the value of using complementary techniques in multiple systems (Cole et al., 1998; Piperno and Mead, 1997). The putative cargo of the IFT machinery was also studied assuming that they may be cotransported with the IFT machinery, for example, OSM-9 and OCR-2 (Qin et al., 2005). This powerful approach also allowed investigators to identify dyf-5, a map kinase, as a modulator of the velocity of OSM-3 (Burghoorn et al., 2007). E. Biochemical Analysis of IFT Motors and Particles Taking advantage of the ease of cell culture and deflagellation, IFT particles have been isolated from C. reinhardtii flagella and their components have been nicely

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(A)

Distal

Middle

Transition

klp-11

osm3/ klp-11

3-D Model

Wild Type

kap-1/ osm-3

kap-1

klp-11

klp-11/ osm-3

kap-1/ klp-11

osm-3

che-3

(B) WT

Fig. 2 (Continued)

osm-3

che-3

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characterized. Antibodies raised against separated IFT components were used for further identification of flagellar mutants and for further investigation of the localization and mechanism of action of IFT components. In addition, by changing the ionic strength, it was determined that isolated IFT particles could dissociate into two subcomplexes, IFT-A and IFT-B, of which IFT-B was found to be formed from a core particle to which a few additional loosely associated polypeptides are bound (Fig. 1A). Recently, the “BBSome,” another subcomplex associated with the IFT machinery, has also been isolated using tandem affinity purification from cultured mammalian cells. This body of work provides useful insights into the native form of the multi-protein complex involved in IFT. However, although most, if not all, the components of the IFT particle have been identified, our understanding of how these IFT components interact with each other and how they interact with motors and cargos is relatively poor (Hao and Scholey, 2009). Unfortunately, C. elegans is not amenable for the biochemical isolation of endogenous IFT particles since it contains few ciliated cells (60 out of ~1000 cells) and the cilia themselves are tiny. Therefore, it is necessary to heterogeneously express IFT proteins in order to investigate the interactions and biochemical properties of these recombinant proteins in vitro. OSM-3, one of the two anterograde IFT motors, has been overexpressed in an Eschericia coli system where both a GFP and a 6 His Tag have been used to facilitate the protein’s purification. The purified recombinant protein was used for sophisticated in vitro gliding motility assays, ATPase assays, single molecule fluorescence, and optical trap experiments (Imanishi et al., 2006; Pan et al., 2006). Heterotrimeric kinesin-2 was initially purified from sea urchin by using pan-kinesin peptide antibodies raised against hyperconserved sequences within the kinesin superfamily motor domain (Cole et al., 1992; Cole et al., 1993). Later purification trials of the orthologous kinesin-II complex from C. elegans resulted only in its partial purification due to the low levels of the native holoenzyme, although it was possible to use coimmunoprecipitations and fractionation procedures to determine its heterotrimeric state, and to reveal that OSM-3 formed a distinct, homodimeric complex (Signor et al., 1999b). Recently we have described the heterologous expression of genes encoding klp-11, klp-20, and kap-1, the three subunits of kinesin-II simultaneously in the baculovirus expression system using Sf9 cell cultures (Pan et al., 2006). The production of heterologously expressed kinesin-II in bulk amounts compared to native kinesin-II has allowed us to isolate the protein in high-purity relatively easily. The

Fig. 2 Examination of ciliary structure. (A) TEM analysis of amphid channel cilia in IFT motor mutants. Representative micrographs of distal and middle segments and transition zones and a side view of the corresponding 3D model that was reconstructed from serial sections from each strain. Three-dimensional reconstructions show trajectories of channel cilia obtained from serial sections taken from the distal tip at the amphid pore down through the basal transition zones to the dendrites (total length ~10 µm). Bars, 250 nm. (B) Amphid channel cilia morphology analyzed by fluorescence microscopy. Amphid channel cilia visualized with IFT particle proteins, OSM-5::GFP or OSM-6::GFP. Arrowheads indicate middle (bottom) and distal (top) segment boundaries. (Reprinted from Evans et al., 2006).

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(A)

M1

M2

M3

M4

OSM-6 GFP

D 140

Middle

Distal

Transport Events

120

D

100 80 60 40 20

M1 M2 M3 M4

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

Velocities (µm/s)

(B) M1

M2

M3

M4

140

OSM-6 GFP in osm-3

Transport Events

120 100 80 60 40 20

M4 M3 M2 M1

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

Velocities (µm/s)

(C) M1

M2

M3

M4

D

OSM-6 GFP in KAP-1

80

D

Transport Events

70 60 50 40 30 20 10

M4 M3 M1 M2

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

Velocities (µm/s)

Fig. 3 Anterograde transport of IFT-particles along the middle and distal segments of C. elegans sensory cilia. Motility of IFT particles (OSM-6::GFP) within sensory cilia of wild-type (A), osm-3 mutant (B) and kap-1 mutant (C). Left column show fluorescence micrographs with corresponding cartoon showing the lines used to generate kymographs along 4 middle segments (M1–M4) and the distal segment (D). Kymographs (middle column) show that motility along the distal segments is faster than along middle segments (no distal segment motility is seen in the osm-3 mutant). Right column, histograms showing IFT velocity profiles along middle and distal segments. Horizontal bars = 2.5 µm. Vertical bar = 5 s. (Reprinted from Snow et al., 2004).

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Table II Velocities of the IFT Motors and Particles in the In Vitro Gliding Assay and In Vivo Motility Assay Pure motor (µm/s)

Motor in cilium (µm/s)

IFT particles (µm/s)

kinesin-II

0.4

0.5

0.5

OSM-3

1.1

1.3

1.2

kinesin-II and OSM-3

0.7

0.7

0.7

Assembly of

Middle segment Full-length cilium/ distal segment Middle segment (redundant)

use of the baculovirus expression system for the production of this complex protein is important in that it allows protein expression in a higher eukaryote system which contains the machinery required for post-translational modifications. This in turn can yield recombinant proteins sharing properties with the native form. Kinesin-II and OSM-3 work together to move an IFT particle along the middle segment of the sensory cilium. The coordination of these two motor proteins was reconstituted by assaying the gliding velocities of mixtures of these two proteins in a series of different ratios. This approach also enabled us to closely correlate the rates of motility of the purified motors with the rates of IFT seen in different domains of wildtype and mutant sensory cilia, allowing us to develop a model for the coordination of the two ciliary motor proteins (Table II) (Pan et al., 2006).

II. Rationale A. Genetic Screens of C. elegans IFT Mutants Using forward genetic approaches in C. elegans, gene inactivation is accomplished in two ways. EMS mutagenesis is the most widely used technique to mutate the C. elegans genome (Brenner, 1974). The ability of EMS to alkylate guanine bases results in base mispairing, primarily producing G/C to A/T transitions, which ultimately causes amino acid changes, deletions, or insertions during subsequent DNA repair. Alternatively, Mos1 transposon-mediated mutagenesis has been used to insert Mos1 DNA fragment into the C. elegans genome to generate mutations (Boulin and Bessereau, 2007). EMS- or Mos1-treated animal can be screened for defects in chemotaxis, osmotic avoidance, dauer larva formation, and dye filling. Among these assays, dye-filling assay is very easy to perform and also provides a direct assessment of the presence of ciliary defects (Perkins et al., 1986). Screens for behavioral mutants are also easy to perform and some of the mutants from these screens are indeed required for IFT or ciliogenesis. However, a significant portion of

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these mutants develop normal cilia and they may therefore play other roles, for example, participating in signal transduction in response to environmental stimulation. Therefore, the dye-filling assay needs to be performed to examine their roles in ciliogenesis (Ou et al., 2007). The identification of EMS-induced mutations in the genome is a time-consuming task. The strategy starts with two-point mapping to determine on which chromosome the mutation localizes by genetic crossing with other phenotypic markers. Then three-point mapping is applied to narrow down the locus of the mutation on the chromosome. The recent application of a snip-SNP (restriction site modified single-nucleotide polymorphism) mapping protocol based on single-nucleotide differences between Bristol (the widely used wild type) and Hawaiian strains of C. elegans significantly speeds up the genetic mapping of ciliary mutants (Wicks et al., 2001). When the mutation is mapped to a small region within several hundred kilobases, a pool of cosmids or fosmids carrying wild-type genes covering this region can be individually transformed into mutant animals by germ-line microinjection, in order to test if they can rescue the mutant phenotype. All the genes in the cosmid or fosmid that rescue the mutant phenotype can then be amplified by PCR to examine their ability to rescue the mutant phenotype, or alternatively, the exons of all the genes on the construct can be amplified by PCR and subjected to sequencing in order to identify the mutation. Since genes containing “X-box motifs” in their promoter regions are likely to be ciliary genes, the identification of candidates for further analysis can be facilitated by bioinformatics searches for putative X-box motifs (see the reason in the next paragraph). Compared with EMS mutations, the identification of Mos1 transposon insertion-induced mutation is relatively straightforward. Mos1 insertions represent molecular tags that allow inverse PCR to amplify the flanking sequences of Mos1 in the C. elegans genome. Moreover, with the advancement of “next-generation” sequencing techniques, whole genome sequencing can be used to identify sought-after mutations (Hillier et al., 2008; Sarin et al., 2008). C. elegans reverse genetics is based on the fact that IFT is a process occurring in the cilium which is a compartmentalized space requiring the transport of cytoplasmically synthesized proteins. Thus, genes contributing to IFT are expressed in ciliated sensory neurons and their products are located in cilia. To identify ciliated sensory neuronexpressed genes, a few strategies have been used: (1) All the currently identified IFT genes and many ciliary genes have an X-box motif in their putative promoter region that can be recognized by DAF-19, an RFX family transcription factor, one isoform of which is specifically expressed in ciliated sensory neurons (Swoboda et al., 2000). On the basis of this, genome-wide searches for genes containing X-boxes in their putative promoter regions predict that hundreds of genes could be regulated by DAF-19 in ciliated sensory neurons (Chen et al., 2006; Efimenko et al., 2005). (2) Serial analysis of gene expression (SAGE) data collected from in vitro cultured sensory neurons provide a list of candidate genes that are expressed in C. elegans sensory neurons (Blacque et al., 2005). (3) Overexpression of a GFP-tagged mRNA poly(A)-binding protein (PAB) in ciliated sensory neurons, followed by the use of a GFP antibody to

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isolate the mRNA species that bind to PAB has been used in the identification of the isolated mRNA which provides information about genes that are specifically expressed in ciliated sensory neurons (Kunitomo et al., 2005). (4) The list of the ciliary genes discovered by comparative genomics (Avidor-Reiss et al., 2004; Li et al., 2004) and proteomics (Liu et al., 2007; Mayer et al., 2008, 2009; Ostrowski et al., 2002; Pazour et al., 2005; Song and Sokolov, 2009) in other organisms can be used to identify likely ciliary genes in C. elegans. Once a set of putative candidate genes are available, these genes can be substantiated as IFT components based on the location of their protein product, IFT assays, and mutant analysis. To isolate C elegans deletion mutants in genes of interest, two steps are involved (Jansen et al., 1997). The first step involves random mutagenesis. A large population of worms is irradiated under UV illumination in the presence of trimethylpsoralen which creates small deletions (presumably in all the genes). Half of the mutated worms are frozen and another half of them are used to extract genomic DNA. The second step is to screen for the mutation of interest by PCR. The primers are designed to flank the desired deletion region in the gene of interest. If a deletion occurs in the region, it brings primers closer together and generates a PCR product smaller than wild type. When a deletion is detected in a DNA pool, one can go back to the corresponding frozen worm stock and recover their siblings which bear the same mutation. Recently, oligonucleotide array comparative genomic hybridization has been used as a new tool to identify the deletion. Using exon-tiled oligonucleotide arrays, DNA samples from WT and mutagenized animals are differentially labeled and hybridized to detect singlecopy single-gene deletions (Flibotte et al., 2009; Maydan et al., 2007). In addition, targeting induced local lesion in genomes (Gilchrist et al., 2006) and PCR product sequencing (Cuppen et al., 2007) have also been adopted to identify mutations of interest.

B. Examination of Ciliary Structure The dye-filling defect phenotype of the dyf mutants typically suggests that cilia structure is abnormal (Hedgecock et al., 1985; Perkins et al., 1986). There are two main ways to examine ciliary structure: TEM and fluorescent microscopy. TEM analysis allows direct observations of axoneme architecture and length, as well as cilia membrane structure (Chalfie and Thomson, 1982; Evans et al., 2006; Perkins et al., 1986). Fluorescence microscopy allows analysis of IFT and cilia length based on the distribution of fluorescently labeled markers. The structure of the basal body, ciliary membrane, axoneme, and matrix has been studied using fluorescent labeling of known components of these domains in the cilium. Ciliary membrane components, OSM-9, OCR-2, TUB-1, etc., have been labeled fluorescently and observed in transgenic worms (Mukhopadhyay et al., 2005; Qin et al., 2005). The alpha, beta, and gamma tubulins are components of the axoneme. The tbb-4 gene, coding for a beta tubulin, has been shown to be specifically expressed in the amphids and phasmids, which makes it a perfect marker to analyze ciliary lengths (Jauregui et al., 2008;

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Portman and Emmons, 2004). The tbg-1 gene encodes a gamma tubulin, which makes it a good marker of the basal body (Tabish, 2007). IFT particle components locate to the matrix of the cilia. So far, almost all the IFT components (Fig. 1A and B) have been labeled with GFP, which are perfect for IFT assays and in most cases are also good markers for ciliary length analysis. In addition, GFP diffuses into cilia by itself when it is expressed in a subset of cilia driven by certain sensory neuron-specific promoters, gcy-5, gcy-7, etc. (Yu et al., 1997). However, the diffusible GFP does not fill the entire full-length cilium.

(A)

(B)

(C)

(D)

Fig. 4 (Continued)

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C. In Vivo IFT Motility Assay C. elegans is transparent and each of its cells can be observed directly under a microscope. Fluorescently tagged IFT proteins can be integrated into the IFT machinery and their motility can be recorded. By analyzing the recorded movie by kymography, the moving velocity can be estimated. In order to perform in vivo motility assays of IFT, transgenic worms that carry a fluorescently tagged IFT protein should be available. An appropriate transgene may be constructed by three commonly used methods (Fig. 4). (1) Worm expression vectors that contain GFP can be used by either traditional cloning (Fire et al., 1990) or gateway recombination (Hope et al., 2004). A gene of interest together with its own putative promoter region is amplified by PCR and cloned into the upstream region of GFP in the vector, either by restriction digestion and ligation in conventional genetic engineering or by recombined arms in the gateway recombination system (Fig. 4A and B). (2) Fusion PCR is used to directly fuse a gene of interest together with its putative promoter to the GFP sequence, and the purified PCR product is directly used as a transgene (Hobert, 2002). This method requires two rounds of PCR. First of all, the gene of interest and GFP are amplified, respectively, using one of the primer pairs that are engineered to recognize each other so that the PCR products can overlap within the engineered sequence. Then the two nonoverlapped primers that were used in the first-round PCR are used for the next-round PCR (Fig. 4C). (3) The GFP encoding sequence is recombined into a gene of interest in a fosmid (Dolphin and Hope, 2006). This method requires two rounds of recombination in a genetically modified bacterial system. In the first round, the PCR product of a gene cassette containing rpsL and tetA(C) genes under control of a strong promoter, ompF, is generated with the primers that are engineered to recognize the sequences flanking the insertion point in the gene of interest. It is transformed into bacteria that contain a single copy of the fosmid, recombination is allowed to occur, and successful recombinants are selected for the TetR marker since the two markers confer streptomycin sensitivity (StrS) and tetracycline resistance (TetR) to the bacterial host, respectively. In

Fig. 4 Schematic cartoon of gene cloning strategies used to make GFP-tagged IFT protein constructs. (A) A gene of interest (e.g., osm-3) together with its promoter is amplified by PCR using two primers engineered into restriction sites (RE1 or RE2), respectively, which recognize the same sites in a vector that carries the GFP sequence. (B) The gene of interest (e.g. osm-3) together with its promoter is amplified by PCR with two primers engineered into a recombination site (attL or attR) which recognizes the same site in a vector that carries the GFP sequence. (C) A gene of interest (e.g., osm-3) together with its promoter is amplified by PCR using two primers. The downstream primer is engineered into a sequence that overlaps the 50 end of the GFP sequence. Another GFP sequence, together with the unc-54 30 UTR, is amplified by PCR. Next, the two PCR products are mixed together and the fusion PCR is performed by using two primers: one upstream of osm-3 and another downstream of the unc-54 30 UTR. (D) Recombineering. The PCR product of a double-selection marker [ompF::rpsL::tetA(C)], amplified by two primers engineered in two 50 mer of sequences that are the same as the sequences flanking the insertion point in a gene of interest (e.g., che-3) in a fosmid, is used to perform first-round recombination. The recombinant is selected by TetR because of the insertion of tetA(C) in the che-3 gene. In the second round of recombination, the PCR product of GFP amplified by two primers engineered in two 50-mer sequences that are the same sequences flanking the insertion point is used. The recombinant is selected by StrR because of the removal of rpsL.

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the next round of recombination, the PCR product of GFP containing the flanking recognition sequences at the insertion point is transformed into the recombinant from the first-round screen for streptomycin resistant recombinant (Fig. 4D). To generate transgenic worms, the construct made by either of the above methods together with an injection marker (e.g., a dominant rol-6 mutant allele in pRF4 that confers a rolling phenotype to transgenic worms) are injected into the worm’s gonad. The transgene forms an extrachromosomal array that contains multiple copies of the transgene (Mello et al., 1991). The bombardment of the transgene can also be used to produce transgenic worms carrying a low copy number of the transgene (Praitis et al., 2001). The successfully created transgenic worms are observed under a fluorescence microscope to see if the transgene is expressed, as judged by the existence of a GFP signal. The transgene is also transformed into the corresponding mutant background to see if it rescues the mutant phenotype, and thus to determine if it is functional. Any transgenic animals carrying a functional transgene can be used to create IFT motility movies under a spinning disc confocal microscope. D. Purification of Heterologously Expressed Motor Proteins and In Vitro Motility Assay Testing purified IFT motor proteins for their motility activity and assaying their biochemical properties is obviously important for improving our understanding of the mechanism of IFT. To this end, motility assays of dyneins and kinesins based on time lapse imaging by video-enhanced fluorescence microscopy of rhodamine-labeled MTs are especially convenient. The assay involves attaching motor proteins to a glass slide and assaying motility using buffer solutions containing ATP and taxol stabilized MTs. Using this approach, the C. elegans IFT motors, heterotrimeric kinesin-II, and OSM-3 have been studied (Scholey, 2008). Motility assays were used to determine the motility properties of each protein individually as well as for a series of mixtures of varying molar ratios of the two proteins. In this study, heterotrimeric kinesin-II was expressed using the baculovirus system and purified using affinity chromatography and gel filtration chromatography, whereas the OSM-3 motor was expressed in E. coli and purified in the Vale Lab. The use of mixtures of the two proteins mimicked the in vivo conditions and led to the generation of a model for IFT in C. elegans that explains how kinesin-II and OSM-3 are functionally coordinated to move the same IFT particle along the middle segment of amphid channel cilia (Table II) (Pan et al., 2006).

III. Methods A. Maintenance of WT and dyf Mutant Worms

1. NGM Plate Preparation and Worm Culturing 1. To a 2-l flask with a stir bar, add 975 ml ddH2O, 2.5 g peptone, 3.0 g NaCl. Stir until dissolved, and then add 19 g of bacto-agar and autoclave at 120°C for 15 min.

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2. Allow NGM agar to cool until the flask may be touched by hand (~55°C), and then add the following sterilized solutions: 1 ml cholesterol (5 mg/ml stock in ethanol), 1 ml 1 M MgSO4, 1 ml 1 M CaCl2, 25 ml 1 M Na-Phosphate, pH 6.0. 3. Pour the plates (35 mm10 mm). When the agar hardens, seed the plates with an overnight culture of OP50 bacterial strain. 4. Pick two adult worms and transfer onto the seeded NGM plates and culture the worms at 20°C for 3–4 days until their progeny grow to adult stage. These worms can be used for the IFT assay and ciliary structure analysis.

2. Freezing Worms 1. Culture six plates of worms at 20°C. When the plates are full of worms, wash off the worms with M9 buffer into a 15-ml plastic tube. 2. Centrifuge at 4000 rpm (2200 g) for 1 min to collect the worms. 3. Aspirate the solution till 1.5 ml is left in the tube and then add 1.5-ml freezing buffer and mix. 4. Dispense them into three cryo-vials, 1 ml per vial, and then put the three vials into a Styrofoam rack and incubate in a –80°C freezer. (The Styrofoam rack allows the temperature to fall gradually). 5. When the worms are frozen (1 h or longer), the worms can be stored in the –80°C freezer or in liquid nitrogen. B. Genetic Screens of dyf Mutants and Dyf Assay

1. Isolation of dyf Mutants 1. Culture wild-type (N2 strain) worms at 20°C to get many L4 larval stage worms or young adults. Wash off the worms into a 15-ml tube. Spin down the worms and resuspend in 3 ml M9. 2. Prepare another 15-ml tube with 1 ml M9 and add 20 µl EMS and mix. 3. Combine the worms and EMS. Incubate on a shaker at 20°C for 4 h. 3. Spin down the worms and wash three times with M9 buffer. 4. Transfer the worms onto fresh NGM plates and allow them to recover for 4 h to overnight. 5. Pick four worms (P0) onto each NGM plate and repeat for 60 plates. 6. Culture the worms for 3 days to obtain F1 progeny, and pick four worms onto each plate and repeat for 200 plates or more. 7. Stain the F2 worms with DiI solution. Observe under a fluorescence dissecting microscope to search for worms that are not able to be stained. 8. The putative dyf worms are singled onto fresh plates and bred true.

2. Dyf Assay 1. Culture the worms till the plate is full of worms of different ages. 2. Make the fluorescent dye solution (DiI for red fluorescence or DiO for green fluorescence) by diluting (1:200) the stock solution with M9 buffer.

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3. For a 3.5-cm plate, add 0.5 ml diluted fluorescent dye solution and swirl the plate to make sure all the worms are immersed in the solution, and incubate for 1 h or longer until the plate surface becomes dry. 4. Observe the stained worms directly under a fluorescence dissecting microscope or pick the worms on a 2% agarose pad with a drop of 10 mM NaN3 and observe under a compound microscope with a 60 objective. C. Examination of Ciliary Structure

1. Preservation of Worms 1. Load worms into specimen holder and insert into high-pressure freezer. 2. Using 2000 bar pressure and liquid nitrogen, freeze specimen and transfer under liquid nitrogen into cryo-vial containing substitution cocktail: 1% OsO4, 0.1% uranyl acetate in 100% acetone. 3. Replace the water in the frozen sample with acetone using a freeze substitution device using the following protocol: a. 3 days at –90°C (Leica AFS) b. 12 h at –25°C c. 3–4 h at 4°C d. 1–2 h at room temperature e. Rinse with 99.99% anhydrous acetone 4. Infiltrate and embed worms in Epon-Araldite resin using the gradient approach. a. 3:1 acetone : resin for 2 h b. 2:2 acetone : resin for 4 h c. 1:3 acetone : resin overnight d. 100% resin overnight 5. Polymerize resin at 70°C for 72 h.

2. Sample Preparation for TEM 1. Align the polymerized resin block to obtain the proper orientation of the worm for obtaining cross-sections. 2. Trim the face of the resin block to a trapezoid just larger than the desired specimen area using a handheld razor blade. 3. Load the trimmed resin block into the ultramicrotome cutting arm and slice 50- to 150-nm thick sections using a glass or diamond knife. 4. Collect the floated serial sections on a formvar coated 3 mm copper TEM slot grid. 5. Stain the grid containing serial sections with 2% uranyl acetate dissolved in methanol for 5 min followed by Reynold’s lead citrate for 2 min.

3. Imaging and Modeling 1. Load grid with serial sections into TEM sample holder and insert into microscope. 2. After using standard alignment procedures, acquire transmission electron micrographs of the sample area at a magnification sufficient for resolving desired information.

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3. Due to the resulting gap of 50–150 nm between consecutive sections the images should be manually aligned using the manual image deformation and alignment system contained within IMOD (freely available from http://bio3d.colorado.edu/imod). 4. After the sections are aligned, model the positions of the microtubules, cilia, and other relevant organelles as surface renderings using 3DMOD (within the IMOD package) to depict the length, path, and axoneme architecture of each cilium. D. Microscopy for In Vivo IFT Motility Assay and Cilia Imaging and Cilia Length Measurement

1. Immobilization of Worms 1. Prepare agarose pads: Put a drop of melted 2% agarose in M9 buffer on a slide and cross another slide on it and press gently. When it is used, detach the two slides and the agarose layer is formed on one of them. 2. Get a cover slip and transfer 16 µl of 10 mM levamisole on it and then pick four worms into the levamisole solution and incubate 5 min with a cover preventing evaporation. Adult worms are usually used for the IFT assay because their cilia size is large and easy to be observed. 3. Reverse the agarose pad and put it onto the cover slip with anesthetized worms. 4. Seal the slide assembly with Valap: Dip a cotton stick into the melted Valap and brush along the edges of the cover slip. 5. Look at the worms against the light and label the worms with a marker pen on the slide side of the assembly so as to find the worms easily in the following step under higher magnification (100). 6. Put a drop of oil on the cover slip and reversely put onto the platform of a confocal microscope to observe.

2. Image Acquisition 1. Set up the microscope and computer program. An Olympus confocal microscope equipped with a 100 1.35 NA objective and an Ultraview spinning disc confocal head is controlled by the program Ultraview. The exposure time is set to 300 ms. 2. Find a worm based on the labeled mark and focus on the animal’s head for amphids or the tail for the phasmids. 3. For the time lapse, images are collected for 3 min. And for each worm strain, at least three worms are recorded. As for the Z-stack imaging, the upper and the lower limits of an amphid are set. The interval between each section sets to 0.1 µm.

3. Data Analysis for IFT Assay 1. MetaMorph (Molecular Devices Corporation, Sunnyvale, CA, USA) is used to analyze the recorded images: go to File\Open special\Build stack\Quick (or Ctrl þ Q) to open a stack file. 2. Go to Stack\Kymograph, and set line width to 5 in the Kymograph dialog window. Use the single line function in the tool box and draw a line along the cilium in the

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image. Then click “Create” in the Kymograph dialog window to create a kymograph image (representative kymographs can be seen in Fig. 3). 3. Go to Measure\Region Measurements. In the “Region Measurements” window, select “All Regions” in the “Include” down-drag menu. Click “Open Log” and select “Dynamic Data Exchange (DDE),” and then click OK; a Microsoft Excel file is opened. Draw a line along a kymograph track of a particle. By clicking “Log Data,” the values (Distance and Area) will be transferred into a Microsoft Excel file. 4. Go to Measure\Measure Pixel. In the “Measure Pixel” dialog window, click “Config Log” to select “Image name”, “X”, “Y”. And then double click on the lines in the Kymograph drawn in the “Region Measurements”. The values of X, Y will appear in the opened Excel file (The lines can be seen in Fig. 3). 5. Calculate the distance, time, and velocity. Distance = 0.129*(XLine2 – XLine1) *Distance/Area, where 0.129 is the parameter of the 100 objective lens. At 100, 1 pixel = 0.129 µm. Time = 0.3 (YLine2– YLine1), where 0.3 s is the exposure time. Velocity = Distance/Time.

4. Data Analysis for Cilia Image and Length Measurement 1. MetaMorph (Molecular Devices Corporation) is used to analyze the recorded images: go to File\Open special\Build stack\Quick (or Ctrl þ Q) to open a stack file. 2. Go to Process\Stack Arithmetic\Maximum to make a projection image. 3. Go to Measure\Region Measurements. In the “Region Measurements” window, select “All Regions” in the “Include” down-drag menu. Click “Open Log” and select “Dynamic Data Exchange (DDE)”, and then click OK; a Microsoft Excel file is opened. 4. Go to Measure\Measure Pixel. In the “Measure Pixel” dialog window, click “Config Log” to select “Image name,” “X,” “Y.” And then double click on the lines in the Kymograph drawn in the “Region Measurements.” The values of X, Y will appear in the opened Excel file. 5. Calculate the cilium length. Cilium length = 0.129 SQRT(Xline2 – Xline1)2 þ (Yline2-Yline1)2.

E. Expression and Purification of Kinesin-II and Gliding Motility Assay

1. Baculovirus Expression of Kinesin-II 1. Cloning the subunits of kinesin-II into the baculovirus expression vector: cDNAs encoding the kinesin-II subunits, KLP-11, KLP-20, and KAP-1 are amplified by PCR and inserted into Gateway vector pDONR221 (Invitrogen, Carlsblad, CA, USA), and are subsequently cloned into the destination vector, pDEST8 (Invitrogen). The kap-1 gene contains a 6 His Tag which was incorporated by including the 6 His in the 30 end primer. Recombinant bacmids are generated by transforming the pDEST8 vectors containing klp-11, klp-20, and kap-1 into E. coli MAX Efficiency®

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2.

3.

4.

5.

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DH10BacTM cells (Invitrogen). The plasmids in these E. coli cells are modified baculovirus DNA molecules called bacmids. Bacmids are isolated from E. coli DH10Bac and used to transfect Sf9 surface cell cultures to generate baculovirus using Cellfectin® reagent (Invitrogen). After two days of infection the first progeny of viral stocks (P1 viral stocks) are collected. At this stage amplification of the P1 viral stocks is done by following the instructions in the Bac-to-Bac® manual (Invitrogen). The titers of the viral stocks are determined using BacPAKTM Baculovirus rapid titer kit following the manufacturer’s instructions (Clontech Mountain View, CA, USA). The unit for titer of the viral stocks is calculated as number of plaque-forming units per milliliter (pfu/ml). Sf9 cells are cultured using serum-free medium Sf-900TM II (Gibco, Carlsblad, CA, USA) to a concentration of 2106 cells/ml. Mix the three baculovirus stocks by taking equal amount of pfu/ml from each and use this to infect Sf9 cell culture. Culture of Sf9 cells are infected with equal multiplicity of infection [MOI: plaque-forming unit (pfu) of virus per Sf9 cell] from each klp-11, klp-20, and kap-1 viral stocks and are then incubated for three days at 27°C and 130 rpm in a shaking incubator. Harvesting is carried out by centrifugation at 1500 rpm (640 g) at 4°C in a Beckman J6-HC centrifuge JS 4.2 type rotor. The cells are collected and washed once, and the final cell pellet is resuspended in a minimum amount of SFM medium to make a dense slurry and this slurry is used to make small frozen balls by slowly dropping into liquid nitrogenfilled ceramic mortar. The frozen Sf9 cells can be used immediately or stored at –80°C.

2. Purification of Kinesin-II 1. Lysis of Sf9 cells: The frozen Sf9 cells are ground using Micro-Dismembrator S (Sartorius Stedim Biotech, Bohemia, NY, USA) using 20 ml Teflon container and metal balls precooled in liquid nitrogen. The ground powder is suspended in lysis buffer. Lysis of the cells can also be carried out by passing the 5% solution of the cells twice through French press as described in Pan et al. (2006). The subsequent steps of purification are performed at 4°C. 2. The ground powder in the lysis buffer is made into a homogenous mixture by mixing with a glass rod for several minutes. The cell homogenate is then centrifuged at 12,000 rpm (17400 g) for 30 min in a Beckman centrifuge JA-20 type rotor. 3. The supernatant is immediately applied to previously equilibrated Talon® column beads (Clontech Mountain View, CA, USA) and shaken for 1 h with gentle inversions. 4. The Talon® beads and crude extract mixture is then centrifuged at 1500 rpm (640 g) for 5 min in a Beckman J6-HC centrifuge using JS 4.2 type rotor. 5. The supernatant is discarded and the beads are washed twice with lysis buffer. In order to facilitate washing, the mixture is shaken by inversions on a shaker for 10 min. 6. The talon beads slurry is then applied to a disposable 5-ml polypropylene column. 7. The column is washed with lysis buffer containing 10 mM imidazole. The protein is then eluted with lysis buffer containing 150 mM imidazole.

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8. Fractions of 1 ml are collected into Eppendorf tubes and analyzed for protein concentration by Bradford’s assay. 9. The fractions containing proteins are pooled and dialyzed against gel filtration column buffer overnight. 10. The dialysate is passed through a Superose 6 (SuperoseTM 10/300 GL) column. Fractions of 1 ml are collected and they are analyzed by SDS-PAGE for the presence of kinesin-II. 11. The fractions containing the purified kinesin-II protein are collected and concentrated using Amicon Ultra centrifugal filters (Millipore, Billerica, MA, USA) to a concentration of 0.5–1 mg/ml. At the end of the described purification protocol milligram amounts of protein can be obtained with high purity. For illustration SDS-PAGE analysis of samples from different steps of a routine purification, sucrose density gradients, and gel filtration are shown in Fig. 5.

3. Gliding Motility Assay 1. Prepare the flow cell as described in Fig. 6A. Two pieces of the double-sided Scotch tape are attached on two edges of a slide. And then a microscope cover slip with size of 22 22 mm is attached on the tapes to make a flow cell. 2. Use a micropipette to flow in 5 µl of kinesin-II (0.5–1 mg/ml) into the flow cell and incubate for 5 min at room temperature. In the mean time prepare the reaction buffer according to Table III. (The fluorescent microtubules are freshly prepared using Fluorescent Microtubules Biochem Kit following the manufacturer’s instructions (Cytoskeleton, Denver, CO, USA)). Do not allow the flow cell to dry out at any time, allow some extra solution to go out of the flow cell as small droplets. Do not allow air bubbles to be trapped in the flow cell. (Note: When a mixture of heterotrimeric kinesin-II and OSM-3 is used the molarities are used to calculate the ratio of these proteins. The mixture of the proteins should be made before applying to the flow cell by mixing determined moles of two motor proteins.) 3. Flow in 10 µl of blocking solution into the flow cell while touching the opposite side of the flow cell with a piece of filter paper to help flow but not wick off all the solution, and then incubate for 5 min. The set up is shown in Fig. 6B. As the fluid is sucked from one side, the solution pipetted from the other side should fill the flow cell. 4. Flow in 20 µl of reaction buffer and observe under microscope immediately. Seal the flow cell with Valap on both sides. 5. Adjust the focus under microscope (Nikon Eclipse E600 microscope with a digital camera (SenSys from Photometrics, Tucson, AZ, USA) attached and connected to a computer) by aiming the side of the well where you can see the tape edge. Microtubules can be seen when the focus is right. Record time lapse images of the motility for 3 min with 7- to 10-s intervals. 6. Data analysis is performed by using MetaMorph software (Molecular Devices Corporation). This method was used to assay the speed of purified kinesin-II and

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(A)

250 kDa 150 kDa 100 kDa 75 kDa

50 kDa

37 kDa

(B)

97 kDa 66 kDa

(C) 97 kDa 66 kDa

Fig. 5 Preparation and purification of kinesin-II. (A) SDS-PAGE analysis of samples from different steps obtained during the purification of kinesin-II. Lanes: (1) molecular weight marker, (2) high-speed supernatant, (3) supernatant after application to Talon column, (4) rinse with lysis buffer, (5) proteins obtained by washing with 10 mM imidazole containing lysis buffer, (6) Talon column eluate, (7) superose 6 purified kinesin-II. On sucrose density gradients (B) and gel filtration columns (C), the KLP-11, KAP-1, and KLP-20 subunits elute as a monodisperse heterotrimeric complex (S value = 9.8; Rs = 7.1 nm; and native molecular mass = 287 kDa) in a KLP-11/KLP-20/KAP-1 molar stoichiometry of 1.0:1.17:0.89 (protein standard peak positions are also indicated). (Panels B and C are reprinted from Pan et al., 2006.)

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(A)

(B) Cover slip

Glass slide

Double-sided Scotch tape

Flow cell

3 mm

Double-sided Scotch tape

2

Filter paper

1

Pipette tip

Fig. 6 Preparation of the flow cell and the experimental set up for in vitro motility assays. (A) Two pieces of Scotch tape are stuck to the glass slide, leaving a 3-mm gap in between them. Then place a cover slip on top of the tape, gently push down on the cover slip to make it stick, and remove any air bubbles trapped in between the tape, slide and cover slip. Remove the Scotch tape sticking out by peeling off. (B). (1) Flow in the protein solution; the fluid will flow into the cell by capillary action. (2) When adding subsequent solutions, use a filter paper to withdraw the previous solution from one side while replacing it with the next solution by drawing it into the chamber from the other side of the flow cell. The protein sticks to both the cover slip and the glass slide. Focus the microscope on the cover slip in order to see the microtubules being moved by motor protein.

Table III The Reaction Mixture Composition for Gliding Motility Assay Ingredients 2 BRB80 ddH2O 10 Antifade 1 M Phosphocreatine 1000 U/ml Creatine Phosphokinase 2 mM Taxol 50 mM MgCl2 50 mM MgATP Microtubule

Volume (µl) 1.5 12.1 1.0 0.5 0.2 0.2 2.0 2.0 0.5

The volume of the microtubules added can be adjusted depending on the concentration of the MT stock.

OSM-3 individually and also to assay mixtures of OSM-3 and kinesin-II at varying molar ratios ([mol OSM-3] [mol OSM-3 þ mol kinesin-II]) in competitive motility assays, to determine how these two types of anterograde motor could cooperate to move IFT particles along the cilium (Table II; Fig. 7) (Pan et al., 2006).

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1.2

R2=0.99

Gliding velocity (µm/s)

1

0.8

0.6

0.4

0.2

0 0

0.2 0.4 0.6 0.8 Mole fraction of OSM-3 (WT)

1

Fig. 7 Result of competitive MT gliding assays for the two anterograde IFT motors, kinesin-II, and OSM-3. MT gliding rate versus mole fraction (i.e., [mol OSM-3] [mol OSM-3 þ mol kinesin-II], ignoring solvent concentrations) for mixtures of OSM-3 and kinesin-II. Gliding assay rates were plotted versus mole fraction of wild-type (WT) OSM-3. Experimental data (black dots) with standard deviations (error bars) are shown with best fits for one of two proposed models for the mechanism of coordination of the two motors (curved line). For details see (Pan et al., 2006).

IV. Materials A. Maintenance of WT and dyf Mutant Worms M9 buffer: 6 g Na2HPO4, 3 g KH2PO4, 5 g NaCl, 0.25 g MgSO47H2O, ddH2O to 1 l. Freezing solution: 20 ml of 1 N NaCl, 10 ml of 1 M potassium phosphate buffer (pH 6.0), 60 ml of 100% glycerol, add ddH2O to 200 ml. Autoclave and then add 0.6 ml of 0.1 M MgSO4. B. Genetic Screen of dyf Mutants and Dyf Assay EMS. Fluorescent dye stock solution: 2 mg/ml in DMF (dimethyl formamide). DiI, 1,10 dioctadecyl-3,3,30 ,30 -tetramethylindocarbocyanine perchlorate (Molecular Probes, Eugene, OR, USA), DiO, 3,30 -dioctadecyloxacarbocyanine perchlorate (Molecular Probes). C. Examination of Ciliary Structure Chemicals: anhydrous acetone, 1% osmium tetroxide, 0.1% and 2% uranyl acetate, Epon-Araldite resin, Formvar, Reynold’s lead citrate. Apparatus: high pressure freezer,

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freeze substitution device, 70°C incubator, ultramicrotome with glass or diamond knife, 3mm copper TEM slot grid (Ted Pella Inc., Redding, CA, USA) and a Transmission Electron Microscope. Software: IMOD (freely available from http://bio3d.colorado.edu/imod). D. In Vivo IFT Motility Assay and Cilia Imaging Animals: transgenic worms carrying fluorescently tagged genes. Chemicals: agarose (SeaKem® GTG®, Lonza, Rockland, ME, USA), levamisole [(-)-tetramisole hydrochloride, Sigma, St Louis, MO, USA], Valap (equal weight of lanolin, vaseline, and paraffin). Apparatus: dissecting microscope, confocal microscope with spinning disc. Softwares: Ultraview, MetaMorph (Molecular Devices Corporation). E. Purification of Kinesin-II and Gliding Motility Assay Lysis buffer: 50 mM pipes (pH 6.9), 300 mM NaCl, 1 mM MgCl2, 1 mM b-mercaptoethanol, and EDTA-free protease inhibitor tablet (Roche, Indianapolis, IN, USA). Gel filtration buffer (pH 6.9): 80 mM pipes, 200 mM NaCl, 1 mM MgCl2, 1 mM EGTA, 1 mM DTT, and 0.1 mM ATP. Slides: Gold Seal® Micro slides (Gold Seal Products, Porthsmouth, NH, USA). Cover slips: FisherBrand Microscope cover glass, 2222 mm (Fisher Scientific Pittsburgh, PA, USA). Double-sided scotch tape. Blocking solution: 5 mg/ml casein in 1 BRB80 (Sigma). The stock solutions of the ingredients used in the assay are 2 BRB80: 160 mM pipes (pH 6.9), 2 mM MgCl2, 2 mM EGTA. 10 Antifade (Cytoskeleton): 1 M phosphocreatine, 1 kU/ml creatine phosphokinase, 2 mM Taxol in DMSO (Cytoskeleton), 50 mM MgCl2, 50 mM MgATP, 0.5 µl MT; except 2 BRB80 buffer and MT solution, all the solutions for the gliding motility assay can be prepared in advance, snap frozen in liquid nitrogen, and stored at –80°C in aliquots.

V. Discussion A. Genetic Screen of IFT Mutants EMS mutagenesis, transposon insertion, and TMP-UV deletion provide complementary and valuable approaches for the genetic analysis of IFT and ciliogenesis in C. elegans. EMS mutagenesis is characterized by high, unbiased mutagenicity with low mortality. It was the first method used to generate ciliary mutations in C. elegans and was most widely used procedure to create mutants in the past. Although the subsequent cloning of EMS-induced mutants is labor-intensive, it is still irreplaceable in many cases, since it is the only well-established method for reproducibly creating point mutations in a gene of interest. Some point mutations in the target protein have yielded key structural and functional insights into that protein. For example, osm-3 (sa125) bears a G444/E mutation caused by EMS in the hinge region of the stalk of the OSM-3 protein (Snow et al., 2004), and this mutated protein is constitutively active in in vitro single-molecule motility assays whereas the wild-type OSM-3 protein is autoinhibited and requires proper activation for motility (Imanishi et al., 2006).

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Thus, EMS mutagenesis paved the way to discover a de-repression phenomenon of MT-based motor protein action. Mos1 transposon-mediated insertion mutagenesis has two advantages over EMS mutagenesis: (1) it is biologically safe to humans and (2) it is easy to clone the mutated genes by inverse PCR using Mos1 as a molecular tag (Boulin and Bessereau, 2007). Reverse genetics has been widely used in the postgenomic era, which significantly accelerates the process of studying the functions of candidate ciliary genes. It creates ciliary mutants whose phenotypes are often too subtle to be uncovered by forward genetics. For instance, mutations in KAP-1 and KLP-11 subunits of kinesin-II were firstly isolated by reverse genetics. Although it ultimately turned out that both mutations increase the transport speeds of IFT, and lead to complete loss of axonemes when combined with osm-3 mutants, the mutant animals display superficially normal cilia using the criteria of dye-filling assays and animal behavioral assays, making them difficult to detect (Snow et al., 2004).

B. Examination of Ciliary Structure Serial section TEM is a powerful tool for observing cellular ultrastructure. When coupled with light microscopy, the technique adopts the added capability of correlating dynamic, functional observations with static, yet high-resolution structural analysis. One limitation of this technique is the inability to detect the presence of specific and individual proteins unless they can be labeled by antibody decoration. However, when used to detect changes between wild-type and mutant organisms, these techniques provide a convenient framework for deciphering the functional role of individual proteins and how that function relates to ciliary structure.

C. In Vivo IFT Motility Assay The powerful in vivo IFT assay has dramatically advanced our understanding of the roles and mechanisms of action of IFT motors and IFT particle components and it will continue to do so. However, the limiting factor of this technique is that it does not provide information on whether IFT proteins are moving individually or whether they are moving as components of an assembled complex. Although it is assumed that all the IFT particle components move in a single complex based on the coisolation of the two IFT subcomplexes in C. reinhardtii, it is clear that some IFT components move at a different rate from other IFT components. One example is seen in the bbs mutant background, where IFT-A and IFT-B move separately, with IFT-A moving at the slow rate characteristic of kinesin-II and IFT-B moves at the fast rate characteristic of OSM3 (Ou et al., 2005b). However, it is hard to explain why DYF-2, an ortholog of the IFT144 subunit of IFT-A, moves at the intermediate rate (0.7 µm/s) in the bbs mutant background (Efimenko et al., 2006). Another example that is hard to reconcile with existing models for IFT occurs in the mutant nphp-4, a C. elegans ortholog of nephrocystin, where osm-6::GFP moves at the slow rate (0.5 µm/s) while all other IFT components move at the intermediate rate (0.7 µm/s) (Jauregui et al., 2008). These

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exceptions challenge the models proposed so far and suggest that much more work is needed to better understand the mechanism of IFT in this system. D. In Vitro Gliding Motility Assay The low levels of native motor proteins in C. elegans and lack of feasibility for their purification from this system encouraged IFT researchers to use heterologously expressed proteins. In our experience, the baculovirus expression system has proved to be very useful for producing the complex of three different subunits of heterotrimeric kinesin-II in a high yield and an active form. This system should be useful for expressing and purifying other components of the IFT particles. Biochemical data on the IFT particle complex rely substantially on data from C. reinhardtii (Cole et al., 1998; Piperno and Mead, 1997). Conversely, data obtained from C. elegans by the analysis of rates of IFT in IFT mutants nicely complement biochemical data from C. reinhardtii (Ou et al., 2005b). The in vitro reconstitution of IFT particles can, in principle, be achieved by coexpressing IFT particle components in the baculovirus system for protein purification, which may in turn allow the future reconstitution of the entire process of IFT using in vitro motility assays.

VI. Summary We have summarized the methods that have so far been employed to investigate IFT motors and particles in C. elegans sensory cilia. These methods are included in four subjects: (1) forward and reverse genetics; (2) ciliary structure examination by TEM or fluorescence microscopy; (3) in vivo IFT assays; and (4) heterologous expression of motor proteins for in vitro motility assays. We provide detailed protocols for these methods. We also discuss some of the advantages and disadvantages of these methods which make C. elegans such a powerful system for studying IFT and cilium biogenesis. Acknowledgments Work on IFT research in the Scholey laboratory is supported by NIH grant GM50718. G. Ou is currently supported by a postdoctoral fellowship from the Damon Runyon Cancer Research Foundation in the Vale laboratory at UCSF.

References Avidor-Reiss, T., Maer, A.M., Koundakjian, E., Polyanovsky, A., Keil, T., Subramaniam, S., and Zuker, C.S. (2004). Decoding cilia function: Defining specialized genes required for compartmentalized cilia biogenesis. Cell 117, 527–539. Bacaj, T., Lu, Y., and Shaham, S. (2008). The conserved proteins CHE-12 and DYF-11 are required for sensory cilium function in Caenorhabditis elegans. Genetics 178, 989–1002. Bell, L.R., Stone, S., Yochem, J., Shaw, J.E., and Herman, R.K. (2006). The molecular identities of the Caenorhabditis elegans intraflagellar transport genes dyf-6, daf-10 and osm-1. Genetics 173, 1275–1286.

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263

Blacque, O.E., Li, C., Inglis, P.N., Esmail, M.A., Ou, G., Mah, A.K., Baillie, D.L., Scholey, J.M., and Leroux, M.R. (2006). The WD repeat-containing protein IFTA-1 is required for retrograde intraflagellar transport. Mol. Biol. Cell 17, 5053–5062. Blacque, O.E., Perens, E.A., Boroevich, K.A., Inglis, P.N., Li, C., Warner, A., Khattra, J., Holt, R.A., Ou, G., Mah, A.K., McKay, S.J., Huang, P., et al. (2005). Functional genomics of the cilium, a sensory organelle. Curr. Biol. 15, 935–941. Blacque, O.E., Reardon, M.J., Li, C., McCarthy, J., Mahjoub, M.R., Ansley, S.J., Badano, J.L., Mah, A.K., Beales, P.L., Davidson, W.S., Johnsen, R.C., Audeh, M., et al. (2004). Loss of C. elegans BBS-7 and BBS-8 protein function results in cilia defects and compromised intraflagellar transport. Genes Dev. 18, 1630–1642. Boulin, T., and Bessereau, J.L. (2007). Mos1-mediated insertional mutagenesis in Caenorhabditis elegans. Nat. Protoc. 2, 1276–1287. Brenner, S. (1974). The genetics of Caenorhabditis elegans. Genetics 77, 71–94. Burghoorn, J., Dekkers, M.P., Rademakers, S., de Jong, T., Willemsen, R., and Jansen, G. (2007). Mutation of the MAP kinase DYF-5 affects docking and undocking of kinesin-2 motors and reduces their speed in the cilia of Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 104, 7157–7162. Chalfie, M., and Thomson, J.N. (1982). Structural and functional diversity in the neuronal microtubules of Caenorhabditis elegans. J. Cell Biol. 93, 15–23. Chen, N., Mah, A., Blacque, O.E., Chu, J., Phgora, K., Bakhoum, M.W., Newbury, C.R., Khattra, J., Chan, S., Go, A., Efimenko, E., Johnsen, R., et al. (2006). Identification of ciliary and ciliopathy genes in Caenorhabditis elegans through comparative genomics. Genome Biol. 7, R126. Cole, D.G., Cande, W.Z., Baskin, R.J., Skoufias, D.A., Hogan, C.J., and Scholey, J.M. (1992). Isolation of a sea urchin egg kinesin-related protein using peptide antibodies. J. Cell Sci. 101(Pt. 2), 291–301. Cole, D.G., Diener, D.R., Himelblau, A.L., Beech, P.L., Fuster, J.C., and Rosenbaum, J.L. (1998). Chlamydomonas kinesin-II-dependent intraflagellar transport (IFT): IFT particles contain proteins required for ciliary assembly in Caenorhabditis elegans sensory neurons. J. Cell Biol. 141, 993–1008. Cole, D.G., Hall, K., Chinn, S.W., Wedaman, K.P., Skoufias, D., and Scholey, J.M. (1993). A novel heterotrimeric kinesin isolated from sea urchin eggs displays plus-end motility. Mol. Biol. Cell 4, 267A. Collet, J., Spike, C.A., Lundquist, E.A., Shaw, J.E., and Herman, R.K. (1998). Analysis of osm-6, a gene that affects sensory cilium structure and sensory neuron function in Caenorhabditis elegans. Genetics 148, 187–200. Cuppen, E., Gort, E., Hazendonk, E., Mudde, J., van de Belt, J., Nijman, I.J., Guryev, V., and Plasterk, R.H. (2007). Efficient target-selected mutagenesis in Caenorhabditis elegans: Toward a knockout for every gene. Genome Res. 17, 649–658. Dolphin, C.T., and Hope, I.A. (2006). Caenorhabditis elegans reporter fusion genes generated by seamless modification of large genomic DNA clones. Nucleic Acids Res. 34, e72. Dusenbery, D.B., Sheridan, R.E., and Russell, R.L. (1975). Chemotaxis-defective mutants of the nematode Caenorhabditis elegans. Genetics 80, 297–309. Efimenko, E., Blacque, O.E., Ou, G., Haycraft, C.J., Yoder, B.K., Scholey, J.M., Leroux, M.R., and Swoboda, P. (2006). Caenorhabditis elegans DYF-2, an orthologue of human WDR19, is a component of the intraflagellar transport machinery in sensory cilia. Mol. Biol. Cell 17, 4801–4811. Efimenko, E., Bubb, K., Mak, H.Y., Holzman, T., Leroux, M.R., Ruvkun, G., Thomas, J.H., and Swoboda, P. (2005). Analysis of xbx genes in C. elegans. Development 132, 1923–1934. Evans, J.E., Snow, J.J., Gunnarson, A.L., Ou, G., Stahlberg, H., McDonald, K.L., and Scholey, J.M. (2006). Functional modulation of IFT kinesins extends the sensory repertoire of ciliated neurons in Caenorhabditis elegans. J. Cell Biol. 172, 663–669. Fire, A., Harrison, S.W., and Dixon, D. (1990). A modular set of lacZ fusion vectors for studying gene expression in Caenorhabditis elegans. Gene 93, 189–198. Flibotte, S., Edgley, M.L., Maydan, J., Taylor, J., Zapf, R., Waterston, R., and Moerman, D.G. (2009). Rapid high resolution single nucleotide polymorphism-comparative genome hybridization mapping in Caenorhabditis elegans. Genetics 181, 33–37. Fujiwara, M., Ishihara, T., and Katsura, I. (1999). A novel WD40 protein, CHE-2, acts cell-autonomously in the formation of C. elegans sensory cilia. Development 126, 4839–4848.

264

Limin Hao et al. Gilchrist, E.J., O’Neil, N.J., Rose, A.M., Zetka, M.C., and Haughn, G.W. (2006). TILLING is an effective reverse genetics technique for Caenorhabditis elegans. BMC Genomics 7, 262. Hao, L., and Scholey, J.M. (2009). Intraflagellar transport at a glance. J. Cell Sci. 122, 889–892. Haycraft, C.J., Schafer, J.C., Zhang, Q., Taulman, P.D., and Yoder, B.K. (2003). Identification of CHE-13, a novel intraflagellar transport protein required for cilia formation. Exp. Cell Res. 284, 251–263. Haycraft, C.J., Swoboda, P., Taulman, P.D., Thomas, J.H., and Yoder, B.K. (2001). The C. elegans homolog of the murine cystic kidney disease gene Tg737 functions in a ciliogenic pathway and is disrupted in osm-5 mutant worms. Development 128, 1493–1505. Hedgecock, E.M., Culotti, J.G., Thomson, J.N., and Perkins, L.A. (1985). Axonal guidance mutants of Caenorhabditis elegans identified by filling sensory neurons with fluorescein dyes. Dev. Biol. 111, 158– 170. Heiman, M.G., and Shaham, S. (2009). DEX-1 and DYF-7 establish sensory dendrite length by anchoring dendritic tips during cell migration. Cell 137, 344–355. Hillier, L.W., Marth, G.T., Quinlan, A.R., Dooling, D., Fewell, G., Barnett, D., Fox, P., Glasscock, J.I., Hickenbotham, M., Huang, W., Magrini, V.J., Richt, R.J., et al. (2008). Whole-genome sequencing and variant discovery in C. elegans. Nat. Methods 5, 183–188. Hirokawa, N., Tanaka, Y., Okada, Y., and Takeda, S. (2006). Nodal flow and the generation of left-right asymmetry. Cell 125, 33–45. Hobert, O. (2002). PCR fusion-based approach to create reporter gene constructs for expression analysis in transgenic C. elegans. Biotechniques 32, 728–730. Hope, I.A., Stevens, J., Garner, A., Hayes, J., Cheo, D.L., Brasch, M.A., and Vidal, M. (2004). Feasibility of genome-scale construction of promoter::reporter gene fusions for expression in Caenorhabditis elegans using a multisite gateway recombination system. Genome Res. 14, 2070–2075. Imanishi, M., Endres, N.F., Gennerich, A., and Vale, R.D. (2006). Autoinhibition regulates the motility of the C. elegans intraflagellar transport motor OSM-3. J. Cell Biol. 174, 931–937. Insinna, C., and Besharse, J.C. (2008). Intraflagellar transport and the sensory outer segment of vertebrate photoreceptors. Dev. Dyn. 237, 1982–1992. Jansen, G., Hazendonk, E., Thijssen, K.L., and Plasterk, R.H. (1997). Reverse genetics by chemical mutagenesis in Caenorhabditis elegans. Nat. Genet. 17, 119–121. Jauregui, A.R., Nguyen, K.C., Hall, D.H., and Barr, M.M. (2008). The Caenorhabditis elegans nephrocystins act as global modifiers of cilium structure. J. Cell Biol. 180, 973–988. Kozminski, K.G., Johnson, K.A., Forscher, P., and Rosenbaum, J.L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523. Kunitomo, H., Uesugi, H., Kohara, Y., and Iino, Y. (2005). Identification of ciliated sensory neuronexpressed genes in Caenorhabditis elegans using targeted pull-down of poly(A) tails. Genome Biol. 6, R17. Lewis, J.A., and Hodgkin, J.A. (1977). Specific neuroanatomical changes in chemosensory mutants of the nematode Caenorhabditis elegans. J. Comp. Neurol. 172, 489–510. Li, J.B., Gerdes, J.M., Haycraft, C.J., Fan, Y., Teslovich, T.M., May-Simera, H., Li, H., Blacque, O.E., Li, L., Leitch, C.C., Lewis, R.A., Green, J.S., et al. (2004). Comparative genomics identifies a flagellar and basal body proteome that includes the BBS5 human disease gene. Cell 117, 541–552. Liu, L.X., Spoerke, J.M., Mulligan, E.L., Chen, J., Reardon, B., Westlund, B., Sun, L., Abel, K., Armstrong, B., Hardiman, G., King, J., McCague, L., et al. (1999). High-throughput isolation of Caenorhabditis elegans deletion mutants. Genome Res. 9, 859–867. Liu, Q., Tan, G., Levenkova, N., Li, T., Pugh, E.N., Jr., Rux, J.J., Speicher, D.W., and Pierce, E.A. (2007). The proteome of the mouse photoreceptor sensory cilium complex. Mol. Cell Proteomics 6, 1299–1317. Maydan, J.S., Flibotte, S., Edgley, M.L., Lau, J., Selzer, R.R., Richmond, T.A., Pofahl, N.J., Thomas, J.H., and Moerman, D.G. (2007). Efficient high-resolution deletion discovery in Caenorhabditis elegans by array comparative genomic hybridization. Genome Res. 17, 337–347. Mayer, U., Kuller, A., Daiber, P.C., Neudorf, I., Warnken, U., Schnolzer, M., Frings, S., and Mohrlen, F. (2009). The proteome of rat olfactory sensory cilia. Proteomics 9, 322–334. Mayer, U., Ungerer, N., Klimmeck, D., Warnken, U., Schnolzer, M., Frings, S., and Mohrlen, F. (2008). Proteomic analysis of a membrane preparation from rat olfactory sensory cilia. Chem. Senses 33, 145–162.

13. Analysis of Intraflagellar Transport in C. elegans Sensory Cilia

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Mello, C.C., Kramer, J.M., Stinchcomb, D., and Ambros, V. (1991). Efficient gene transfer in C. elegans: Extrachromosomal maintenance and integration of transforming sequences. EMBO J. 10, 3959–3970. Morris, R.L., and Scholey, J.M. (1997). Heterotrimeric kinesin-II is required for the assembly of motile 9 þ 2 ciliary axonemes on sea urchin embryos. J. Cell Biol. 138, 1009–1022. Mukhopadhyay, A., Deplancke, B., Walhout, A.J., and Tissenbaum, H.A. (2005). C. elegans tubby regulates life span and fat storage by two independent mechanisms. Cell Metab. 2, 35–42. Murayama, T., Toh, Y., Ohshima, Y., and Koga, M. (2005). The dyf-3 gene encodes a novel protein required for sensory cilium formation in Caenorhabditis elegans. J. Mol. Biol. 346, 677–687. Nachury, M.V., Loktev, A.V., Zhang, Q., Westlake, C.J., Peranen, J., Merdes, A., Slusarski, D.C., Scheller, R.H., Bazan, J.F., Sheffield, V.C., and Jackson, P.K. (2007). A core complex of BBS proteins cooperates with the GTPase Rab8 to promote ciliary membrane biogenesis. Cell 129, 1201–1213. Omori, Y., Zhao, C., Saras, A., Mukhopadhyay, S., Kim, W., Furukawa, T., Sengupta, P., Veraksa, A., and Malicki, J. (2008). Elipsa is an early determinant of ciliogenesis that links the IFT particle to membraneassociated small GTPase Rab8. Nat. Cell Biol. 10, 437–444. Orozco, J.T., Wedaman, K.P., Signor, D., Brown, S., Rose, L., and Scholey, J.M. (1999). Movement of motor and cargo along cilia. Nature 398, 674–674. Ostrowski, L.E., Blackburn, K., Radde, K.M., Moyer, M.B., Schlatzer, D.M., Moseley, A., and Boucher, R.C. (2002). A proteomic analysis of human cilia: Identification of novel components. Mol. Cell Proteomics 1, 451–465. Ou, G., Koga, M., Blacque, O.E., Murayama, T., Ohshima, Y., Schafer, J.C., Li, C., Yoder, B.K., Leroux, M.R., and Scholey, J.M. (2007). Sensory ciliogenesis in Caenorhabditis elegans: Assignment of IFTcomponents into distinct modules based on transport and phenotypic profiles. Mol. Biol. Cell 18, 1554–1569. Ou, G., Qin, H., Rosenbaum, J.L., and Scholey, J.M. (2005a). The PKD protein qilin undergoes intraflagellar transport. Curr. Biol. 15, R410–R411. Ou, G.S., Blacque, O.E., Snow, J.J., Leroux, M.R., and Scholey, J.M. (2005b). Functional coordination of intraflagellar transport motors. Nature 436, 583–587. Pan, X., Ou, G., Civelekoglu-Scholey, G., Blacque, O.E., Endres, N.F., Tao, L., Mogilner, A., Leroux, M.R., Vale, R.D., and Scholey, J.M. (2006). Mechanism of transport of IFT particles in C. elegans cilia by the concerted action of kinesin-II and OSM-3 motors. J. Cell Biol. 174, 1035–1045. Pazour, G.J., Agrin, N., Leszyk, J., and Witman, G.B. (2005). Proteomic analysis of a eukaryotic cilium. J. Cell Biol. 170, 103–113. Perkins, L.A., Hedgecock, E.M., Thomson, J.N., and Culotti, J.G. (1986). Mutant sensory cilia in the nematode Caenorhabditis elegans. Dev. Biol. 117, 456–487. Piperno, G., and Mead, K. (1997). Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella. Proc. Natl. Acad. Sci. USA 94, 4457–4462. Portman, D.S., and Emmons, S.W. (2004). Identification of C. elegans sensory ray genes using wholegenome expression profiling. Dev. Biol. 270, 499–512. Praitis, V., Casey, E., Collar, D., and Austin, J. (2001). Creation of low-copy integrated transgenic lines in Caenorhabditis elegans. Genetics 157, 1217–1226. Qin, H., Burnette, D.T., Bae, Y.K., Forscher, P., Barr, M.M., and Rosenbaum, J.L. (2005). Intraflagellar transport is required for the vectorial movement of TRPV channels in the ciliary membrane. Curr. Biol. 15, 1695–1699. Qin, H., Rosenbaum, J.L., and Barr, M.M. (2001). An autosomal recessive polycystic kidney disease gene homolog is involved in intraflagellar transport in C. elegans ciliated sensory neurons. Curr. Biol. 11, 457–461. Riddle, D.L. (1977). A genetic pathway for dauer larva formation in C. elegans. Stadler Genet. Symp. 9, 101– 120. Rosenbaum, J.L., and Witman, G.B. (2002). Intraflagellar transport. Nat. Rev. Mol. Cell Biol. 3, 813–825. Sarin, S., Prabhu, S., O’Meara, M.M., Pe’er, I., and Hobert, O. (2008). Caenorhabditis elegans mutant allele identification by whole-genome sequencing. Nat. Methods 5, 865–867. Schafer, J.C., Haycraft, C.J., Thomas, J.H., Yoder, B.K., and Swoboda, P. (2003). XBX-1 encodes a dynein light intermediate chain required for retrograde intraflagellar transport and cilia assembly in Caenorhabditis elegans. Mol. Biol. Cell 14, 2057–2070.

266

Limin Hao et al. Scholey, J.M. (2003). Intraflagellar transport. Annu. Rev. Cell Dev. Biol. 19, 423–443. Scholey, J.M. (2008). Intraflagellar transport motors in cilia: Moving along the cell’s antenna. J. Cell Biol. 180, 23–29. Shakir, M.A., Fukushige, T., Yasuda, H., Miwa, J., and Siddiqui, S.S. (1993). C. elegans osm-3 gene mediating osmotic avoidance behaviour encodes a kinesin-like protein. Neuroreport 4, 891–894. Signor, D., Wedaman, K.P., Orozco, J.T., Dwyer, N.D., Bargmann, C.I., Rose, L.S., and Scholey, J.M. (1999a). Role of a class DHC1b dynein in retrograde transport of IFT motors and IFT raft particles along cilia, but not dendrites, in chemosensory neurons of living Caenorhabditis elegans. J. Cell Biol. 147, 519– 530. Signor, D., Wedaman, K.P., Rose, L.S., and Scholey, J.M. (1999b). Two heteromeric kinesin complexes in chemosensory neurons and sensory cilia of Caenorhabditis elegans. Mol. Biol. Cell 10, 345–360. Snow, J.J., Ou, G., Gunnarson, A.L., Walker, M.R., Zhou, H.M., Brust-Mascher, I., and Scholey, J.M. (2004). Two anterograde intraflagellar transport motors cooperate to build sensory cilia on C. elegans neurons. Nat. Cell Biol. 6, 1109–1113. Song, H., and Sokolov, M. (2009). Analysis of protein expression and compartmentalization in retinal neurons using serial tangential sectioning of the retina. J. Proteome Res. 8, 346–351. Starich, T.A., Herman, R.K., Kari, C.K., Yeh, W.H., Schackwitz, W.S., Schuyler, M.W., Collet, J., Thomas, J.H., and Riddle, D.L. (1995). Mutations affecting the chemosensory neurons of Caenorhabditis elegans. Genetics 139, 171–188. Swoboda, P., Adler, H.T., and Thomas, J.H. (2000). The RFX-type transcription factor DAF-19 regulates sensory neuron cilium formation in C. elegans. Mol. Cell 5, 411–421. Tabish, M. (2007). Expression of gamma-tubulin during the development of nematode Caenorhabditis elegans. Mol. Biol. Rep. 34, 233–240. Ward, S., Thomson, N., White, J.G., and Brenner, S. (1975). Electron microscopical reconstruction of the anterior sensory anatomy of the nematode Caenorhabditis elegans?2UU. J. Comp. Neurol. 160, 313–337. Wicks, S.R., de Vries, C.J., van Luenen, H.G., and Plasterk, R.H. (2000). CHE-3, a cytosolic dynein heavy chain, is required for sensory cilia structure and function in Caenorhabditis elegans. Dev. Biol. 221, 295– 307. Wicks, S.R., Yeh, R.T., Gish, W.R., Waterston, R.H., and Plasterk, R.H. (2001). Rapid gene mapping in Caenorhabditis elegans using a high density polymorphism map. Nat. Genet. 28, 160–164. Yu, S., Avery, L., Baude, E., and Garbers, D.L. (1997). Guanylyl cyclase expression in specific sensory neurons: A new family of chemosensory receptors. Proc. Natl. Acad. Sci. USA 94, 3384–3387. Zwaal, R.R., Broeks, A., van Meurs, J., Groenen, J.T., and Plasterk, R.H. (1993). Target-selected gene inactivation in Caenorhabditis elegans by using a frozen transposon insertion mutant bank. Proc. Natl. Acad. Sci. USA 90, 7431–7435.

CHAPTER 14

Functional Genomics of Intraflagellar Transport-Associated Proteins in C. elegans Peter N. Inglis*, Oliver E. Blacque†, and Michel R. Leroux* * Department of Molecular Biology and Biochemistry, Simon Fraser University, 8888 University Drive, Burnaby, BC V5A 1S6, Canada † School of Biomolecular and Biomedical Science, UCD Conway Institute, University College Dublin, Belfield, Dublin 4, Ireland

Abstract I. Introduction A. Caenorhabditis elegans Sensory Cilia B. Intraflagellar Transport in C. elegans II. Identification of Novel IFT-Associated and Ciliary Proteins III. Characterization of Candidate and Known IFT-Associated Proteins A. Expression Analysis of Candidate IFT-Associated Genes Using Promoter–GFP Fusion Constructs B. Subcellular Localization of Candidate IFT-Associated Proteins Using GFP-Tagged Fusion Proteins C. Measuring the IFT Velocities of GFP-Tagged IFT-Associated Proteins In Vivo D. Functional Characterization of IFT-Associated Proteins and Assignment to Distinct Modules E. Roles of IFT-Associated Proteins in C. elegans Sensory-Dependent Behaviors and Signaling IV. Conclusions Acknowledgments References

Abstract The nematode Caenorhabditis elegans presents numerous advantages for the identification and molecular analysis of intraflagellar transport (IFT)-associated proteins, which play a critical role in the formation of cilia. Many proteins were first described METHODS IN CELL BIOLOGY, VOL. 93 Copyright 2009 Elsevier Inc. All rights reserved.

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as participating in IFT in this organism, including IFTA-1 (IFT121), DYF-1 (fleer/ IFT70), DYF-2 (IFT144), DYF-3 (Qilin), DYF-11 (MIP-T3/IFT54), DYF-13, XBX-1 (dynein light intermediate chain), XBX-2 (dynein light chain), CHE-13 (IFT57/HIPPI), orthologs of Bardet–Biedl syndrome proteins, and potential regulatory protein, IFTA-2 (RABL5/IFT22). Transgenic animals bearing green fluorescent protein (GFP)-tagged proteins can be generated with ease, and in vivo imaging of IFT in both wild-type and cilia mutant strains can be performed quickly. The analyses permit detailed information on the localization and dynamic properties (velocities along the ciliary axoneme) of the relevant proteins, providing insights into their potential functions in processes such as anterograde and retrograde transport and cilium formation, as well as association with distinct modules of the IFT machinery (e.g., IFT subcomplexes A or B). Behavioral studies of the corresponding IFT-associated gene mutants further enable an understanding of the ciliary role of the proteins—e.g., in chemosensation, lipid homeostasis, lifespan control, and signaling—in a multicellular animal. In this chapter, we discuss how C. elegans can be used for the identification and characterization of IFT-associated proteins, focusing on methods for the generation of GFP-tagged IFT reporter strains, time-lapse microscopy, and IFT rate measurements.

I. Introduction A. Caenorhabditis elegans Sensory Cilia Cilia are motile and/or sensory organelles found on the surfaces of most unicellular eukaryotes and cell types in metazoans (Bae and Barr, 2008; Ginger et al., 2008; Inglis et al., 2007; Pedersen et al., 2008; Satir and Christensen, 2007, 2008). In humans, they are linked to a growing number of so-called ciliopathies, such as Bardet–Biedl syndrome (BBS), Joubert syndrome, and Meckel syndrome (MKS), that are altogether characterized by obesity, retinal degeneration, kidney disease, brain malformations, skeletal defects, sensory anomalies, and other ailments (Badano et al., 2006; Fliegauf et al., 2007; Hildebrandt et al., 2009; Quinlan et al., 2008; Sharma et al., 2008; Veland et al., 2009; Zaghloul and Katsanis, 2009). Cilium biogenesis almost universally depends on intraflagellar transport (IFT), a microtubule-dependent trafficking system used to deliver ciliary components such as structural precursors and signal transduction components (discussed in detail in Section I.B below). The nematode Caenorhabditis elegans represents an excellent model system for the study of nonmotile (sensory) cilia, as worms represent one of the few metazoans that possess solely sessile axonemal structures. They are found exclusively at the tips of sensory neuron dendritic processes, where they are responsible for detecting and transducing a myriad of environmental cues (Bae and Barr, 2008; Bargmann, 2006; de Bono and Maricq, 2005; Inglis et al., 2007). In the hermaphrodite, 60 of its 302 neurons are ciliated; in the more complex male neuronal network, additional cilia largely involved in mechanosensory and chemosensory processes required for mating are present, mostly in the tail of the animal (Bae and Barr, 2008; Barr and Garcia, 2006). These ciliated neurons are

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Fig. 1 C. elegans contains 60 ciliated neurons in the adult hermaphrodite and additional ciliated neurons in the male (most in the tail). The expression pattern of a gene expressed in all ciliated neurons is revealed by fluorescence microscopy of a transgenic animal carrying a bbs-8 gene promoter (bbs-8p)-GFP transcriptional fusion. 1, inner/outer labial ciliated neuron cell bodies; 2, amphid ciliated neuron cell bodies; PHA, PHB, phasmid ciliated neurons; PDE, ciliated neuron; den, dendrites; arrows, dendritic endings.

revealed by the expression pattern of a ciliary gene, bbs-8 (Fig. 1). For the most part, analysis of IFT in the worm has been restricted to the subset of rod-shaped cilia found in amphid and phasmid sensory neurons, which are found in the head and tail of the animal, respectively (Fig. 2) (Blacque et al., 2008; Inglis et al., 2007; Scholey, 2008; Scholey et al., 2004). Although several review articles (Bae and Barr, 2008; Bargmann, 2006; Inglis et al., 2007) provide detailed descriptions of C. elegans ciliated cell types, as well as the structures and sensory functions of the various cilia, we present the most useful information pertinent for analysis of IFT below. The amphid cell bodies are located near the pharynx, and extend dendrites that ultimately terminate in ciliary endings, close to the mouth of the animal (Figs. 1 and 2). Transmission electron microscopy (TEM) performed on C. elegans amphid cilia over 30 years ago revealed that their axonemal structures are essentially identical to those of vertebrate primary (nonmotile) cilia (Ward et al., 1975). Specifically, the amphid channel axonemes are divided into three principle regions: an approximately 1.0-µmlong transition zone (TZ) and 4.0-µm-long middle segment (MS), both consisting of canonical 9 2 þ 0 microtubule doublets (i.e., 9 microtubule doublets without a central pair), and an approximately 2.5-µm-long distal segment (DS) composed of microtubule singlets (9 1 þ 0). Such cilia are found at the anterior-most portions of

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Fig. 2 Whole nematode with close-up views (fluorescent images and schematics) of GFP-tagged CHE-2 (IFT80) protein localizing to amphid cilia in the head of the animal and to phasmid cilia in the tail. BB, basal body; TZ, transition zone.

the ASE, ASG, ASH, ASI, ASJ, and ASK neurons, where they occur as single rods, and in the ADF and ADL neurons, where they terminate in pairs (Bae and Barr, 2008; Inglis et al., 2007; Perkins et al., 1986; Ward et al., 1975). Collectively, the ciliary axonemes of these eight neuron types form a ciliary bundle that protrude into the amphid channel, which is created by the ciliary axonemes penetrating through a sheath cell and becoming enveloped at the distal end by a socket cell and surrounding cuticle. Belt desmosomes between the various cell types seal off this environmentally exposed channel from the interior milieu of the worm. The pair of neuron types known as the phasmids (PHA and PHB) are found in the posterior of the animal (Figs. 1 and 2), near the anus, and possess rod-shaped cilia that also form part of a sensory channel cilia

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similar to the amphid channel. Four additional amphid neurons (AWA, AWB, AWC, and AFD) possess morphologically distinct cilia that terminate within the cuticle and are therefore not environmentally exposed (Bae and Barr, 2008; Inglis et al., 2007; Perkins et al., 1986; Ward et al., 1975). Although IFT rate analyses are often done on amphid cilia because more particles can be detected simultaneously for kymograph analyses (see Section III.C below), in some cases, visualization of IFT particles or IFTassociated phenotypes in the phasmid cilia is less convoluted, as they are only found in bundles of two. A more recent study has also examined the mechanism of IFT in the wing-shaped AWB cilium (Mukhopadhyay et al., 2007). The foundation required for building cilia, namely, a basal body derived from a modified centriolar structure, is conserved across organisms (Marshall, 2008). Interestingly, however, Ward and colleagues (Ward et al., 1975) discovered in their TEM analyses that terminally differentiated C. elegans cilia do not emanate from a canonical triplet microtubule-based basal body (see schematic in Fig. 3). Instead, the base of the cilia displays an apparently degenerate basal body that contains the canonical basal body marker g-tubulin (Bobinnec et al., 2000) and by TEM appears to consist almost exclusively of transitional fibers, the ubiquitous pinwheel-like structures that emanate from the very distal end of basal bodies and connect to the ciliary membrane (Perkins et al., 1986). Notably, the tips of the transitional fibers are thought to be the principal assembly/docking sites for IFT particles (Deane et al., 2001). Following this region is a more extensive, approximately 1-µm-long canonical TZ region which exhibits Y-shaped links that connect the doublet microtubules of the ciliary axoneme to the ciliary membrane (Fig. 3). Because a typical basal body structure is not observed, the bases of C. elegans cilia are often referred to exclusively as the TZ. However, we now favor referring to the (degenerate) basal body and TZ separately as they likely will display distinct protein compositions and serve different functions. For example, the C. elegans retinitis pigmentosa 2 (RP2) protein likely localizes specifically to the transitional fibers and not the TZ (Blacque et al., 2005; Stephan et al., 2007); this distal basal body region is likely also the site of IFT particle assembly/docking (Deane et al., 2001). In contrast, several proteins, most associated with ciliopathies, have been observed to localize within the basal body–TZ region and may be specifically associated with TZs, namely, NPHP-1, NPHP-4, and three proteins containing B9 domains (XBX-7/MKS-1, TZA-1/MKSR-2, and TZA-2/MKSR-1) (Bialas et al., 2009; Jauregui and Barr, 2005; Williams et al., 2008; Winkelbauer et al., 2005). B. Intraflagellar Transport in C. elegans The movement of IFT particles was first observed in Chlamydomonas reinhardtii by differential interference contrast (DIC) videomicroscopy (Kozminski et al., 1993), with the bidirectional movement of specific IFT components along ciliary axonemes subsequently confirmed using GFP-tagged proteins in transgenic C. elegans (Orozco et al., 1999). Research in Chlamydomonas has provided key insights into the nature of the canonical IFT machinery, which consists of an anterograde heterotrimeric kinesin-2 motor (Kinesin II), two biochemically separable subcomplexes A and B

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Fig. 3 Model for intraflagellar transport in C. elegans. Six distinct steps are indicated (1–6) for the canonical IFT pathway in amphid and phasmid rod-type cilia, with the individual IFT modules and ciliary structural elements shown in the legend. BB, basal body; TZ, transition zone. See text for details.

containing 15 polypeptides, and a retrograde motor (dynein 1b) (Blacque et al., 2008; Hao and Scholey, 2009; Pedersen and Rosenbaum, 2008; Pedersen et al., 2008; Scholey, 2008). While all of these originally described components of IFT particles appear to participate in nematode IFT (Table I), recent studies in C. elegans have revealed a greater complexity to the makeup and mechanism of IFT. For example,

14. Functional Genomics of Intraflagellar Transport

Table I Intraflagellar transport (IFT)-associated genes found in C. elegans IFT modules/ components

Descriptions

C. elegans sequences

C. elegans proteins

Selected alleles

C. elegans references

Kinesin-II

95 kDa motor 85 kDa motor Accessory Motor

F20C5.2 Y50D7A.6 F08F8.3 M02B7.3

KLP-11 KLP-20 KAP-1 OSM-3

tm324a ok2942a ok676 p802

Snow et al. (2004)

Heavy chain Light int. chain Int. chain Light chain

F18C12.1 F02D8.3 C17H12.1 D1009.5

CHE-3 XBX-1 DYCI-1 XBX-2

e1124 ok279 tm3700a tm2097

IFT144 IFT140 IFT139 IFT122 IFT121 IFT43

ZK520.3 C27A7.4 ZK328.7 F23B2.4 C54G7.4 N.D.

DYF-2 CHE-11

Efimenko et al. (2006) Qin et al. (2001)

DAF-10 IFTA-1

m160 e1810 gk508b, gk477b e1387 nx61

IFT172 IFT88 IFT81 IFT80 IFT74/72 IFT70 IFT57/55 IFT54

T27B1.1 Y41G9A.1 F32A6.2 F38G1.1 C18H9.8 F54C1.5 F59C6.7 C02H7.1

OSM-1 OSM-5 IFT-81 CHE-2 IFT-74 DYF-1 CHE-13 DYF-11

p808 p813 tm2355, tm2356 e1033 tm2393, tm2397 mn335 e1805 mn392

Bell et al. (2006) Haycraft et al. (2001) Kobayashi et al. (2007) Fujiwara et al. (1999) Kobayashi et al. (2007) Ou et al. (2005) Haycraft et al. (2003) Bacaj et al. (2008); Kunitomo and Iino (2008); Li et al. (2008); Omori et al. (2008)

OSM-3 Kinesin IFT-Dynein

IFT-A

IFT-B

Snow et al. (2004) Shakir et al. (1993) Signor et al. (1999) Schafer et al. (2003) Efimenko et al. (2005)

Bell et al. (2006) Blacque et al. (2006)

(Continued )

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Table I (Continued ) IFT modules/ components

BBSome

Cargo

Descriptions

C. elegans sequences

C. elegans proteins

Selected alleles

C. elegans references

IFT52 IFT46 IFT27 IFT25 IFT22 IFT20 Qilin

R31.3 F46F6.4 N.D. N.D. T28F3.6 Y110A7A.20 C04C3.5 C27H5.7

OSM-6 DYF-6

p811 m175, mn346

Collet et al. (1998) Bell et al. (2006)

IFTA-2

tm1724 gk548a, tm2935a m185 mn396

Schafer et al. (2006) Ou et al. (2007) Murayama et al. (2005) Blacque et al. (2005)

BBS1 BBS2

Y105E8A.5 F20D12.3

BBS-1 BBS-2

Blacque et al. (2004) Blacque et al. (2004)

BBS3 BBS4 BBS5 BBS7 BBS8 BBS9

C38D4.8 F58A4.14 R01H10.6 Y75B8A.12 T25F10.5 C48B6.8

ARL-6 BBS-4 BBS-5 OSM-12 BBS-8 BBS-9

ok1111 tm3231a, tm3044a, ok2053a ok3472a tm3038b gk507a n1606 nx77 gk471a

TRPV channel

B0212.5

OSM-9

TRPV channel

T09A12.3

OCR-2

n1516, ok1677, ky10 ak47, ok1711

Colbert et al. (1997); Qin et al. (2005) Tobin et al. (2002)

DYF-3 DYF-13

Fan et al. (2004) Ou et al. (2007) Blacque et al. (2004) Blacque et al. (2004)

Peter N. Inglis et al.

Each protein is grouped into one of the IFT modules described in the text. For some components first characterized in C. elegans, namely, DYF-2, IFTA-1, and IFTA-2, association with the noted IFT subcomplexes A (IFT-A) or B (IFT-B) has only been recently confirmed by Cole and Snell (2009). The association of DYF-1, DYF-3, DYF-11, and DYF-13 with IFT-B has been shown biochemically by Omori et al. (2008) and Follit et al. (2009). N.D., not detected in C. elegans. a Uncharacterized allele. b Likely not a null mutant, based on preliminary phenotyping and cDNA sequencing (unpublished observations).

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worms have an additional kinesin-2 motor, homodimeric OSM-3 (KIF17 in vertebrates), that contributes to anterograde IFT and is widely conserved across metazoans, and is additionally found in some protists (Shakir et al., 1993; Silverman and Leroux, 2009; Snow et al., 2004). Other proteins were first shown to participate in IFT in C. elegans, namely, six different BBS proteins (BBS-1, BBS-2, BBS-3, BBS-5, BBS-7, and BBS-8), dynein motor subunits (XBX-1 and XBX-2), CHE-13, IFTA-1, DYF-1, DYF-2, DYF-3, DYF-11, and DYF-13 (Bacaj et al., 2008; Blacque et al., 2004, 2005; Efimenko et al., 2005, 2006; Fan et al., 2004; Haycraft et al., 2003; Kunitomo and Iino, 2008; Li et al., 2008; Murayama et al., 2005; Omori et al., 2008; Ou et al., 2005a, b, 2007). A complete list of proteins known to be IFT-associated (i.e., visualized to undergo IFT) is presented in Table I, and fluorescent microscopy images of a GFPtagged IFT protein (CHE-2) localizing to amphid head cilia and phasmid tail cilia (Fujiwara et al., 1999), is shown in Fig. 2. Together, the studies carried out principally in Chlamydomonas and C. elegans strongly support the notion that the IFT machinery consists of separate functional modules, namely, IFT subcomplexes A (IFT-A) and B (IFT-B), kinesin motors (Kinesin-II and OSM-3), dynein motor, BBS, and ciliary cargo modules (Blacque et al., 2008; Hao and Scholey, 2009; Ou et al., 2007). In C. elegans, module classification has relied on detailed phenotypic analyses, incorporating IFT rate measurements and protein localization descriptions, as well as phenotypic assessment of mutant alleles, detailing cilia structure/morphologies, as well as cilia-related behaviors such as chemo- and osmo-sensation. In general, the phenotypic profiles of individual module components tend to be very similar. A revised view of the different modules and mechanism of action of the IFT machinery, incorporating the latest C. elegans findings, is described in Fig. 3. According to the emerging model of IFT in C. elegans amphid and phasmid channel cilia, the two kinesin-2 anterograde motors (Kinesin-II and OSM-3) act cooperatively in the doublet microtubule-containing Middle Segments (MSs), together trafficking at a rate of 0.7 µm/s. This velocity likely arises from the averaged faster speed of OSM-3 (1.1–1.2 µm/s) and slower motility of Kinesin-II (0.5 µm/s) (Pan et al., 2006; Snow et al., 2004). Kinesin-II does not exit the MSs, and OSM-3 is solely responsible for transport of the remaining IFT machinery (IFT-A, IFT-B, BBS proteins, dynein, and presumably cargo) in the singlet microtubule-containing Distal Segments (DSs), traveling at 1.2 µm/s (Fig. 3). Consistent with a partially redundant role for the motors, in the absence of Kinesin-II (klp-11 or kap-1 mutants), OSM-3 builds full-length cilia by transporting the IFT machinery in MSs and DSs at 1.2 µm/s (Fig. 4). Conversely, in osm-3 mutant animals, Kinesin-II travels at 0.5 µm/s and only generates shortened cilia consisting only of MSs (Fig. 4). Dynein is transported in an inactive form by anterograde IFT to the ciliary tip, and following a remodeling of the IFT machinery, it is then used to recycle the IFT machinery (including the inactive OSM-3 kinesin) and other ciliary components back to the basal body, at a rate of 1.10 µm/s (Signor et al., 1999; Snow et al., 2004); at or near the basal body/transitional fibers, remodeling of IFT components and assembly of new IFT particles take place (Fig. 3). Notably, disruption of IFT-A components typically results in slightly shortened cilia with IFT protein accumulations at the tip (Fig. 5), whereas mutations affecting IFT-B components are associated with

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Fig. 4 Defects in IFT and ciliary structures observed in kinesin or dynein motor mutants. Schematics for wild-type, kinesin-2 (klp-11 or kap-1), osm-3, or IFTdynein (che-3 or xbx-1) mutants use the same conventions depicted in Fig. 3, and are shown alongside representative fluorescence images of GFP-tagged IFT-A, IFT-B, or BBS components. In the schematics, the four distinct regions of amphid/phasmid cilia are indicated (BB, basal body/transitional fiber; TZ, transition zone; middle and DSs) are indicated along with variations (e.g., absent DS in osm-3 and IFT protein accumulations in the IFT-dynein mutants). Fluorescent images: cil, cilia; *, accumulations.

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Fig. 5 Defects in IFT and ciliary structures observed when core IFT and BBS proteins are disrupted. The schematics and fluorescent images for the indicated wildtype and mutant strains follow the same convention as that shown in Fig. 4. An “X” shows that a specific module is disrupted.

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severe shortening of the ciliary axoneme (Fig. 5), suggesting—as observed in Chlamydomonas—seemingly distinct roles for IFT-A and IFT-B in retrograde and anterograde transport, respectively (Blacque et al., 2008; Hao and Scholey, 2009; Pedersen and Rosenbaum, 2008; Pedersen et al., 2008; Scholey, 2008). Such distinct activities for IFTA and IFT-B subcomplex proteins are also suggested from IFT studies in Trypanosoma brucei (Absalon et al., 2008). Variations on the “canonical” IFT model exist for at least one other type of C. elegans cilium in which IFT has been characterized. In AWB cilia, the Kinesin-II and OSM-3 motors are at least partially decoupled, and either motor can build an apparently complete ciliary structure (Mukhopadhyay et al., 2007). Components that regulate the movement of the motors but may not be part of the IFT machinery (DYF-5) have also been uncovered in C. elegans, pointing to distinct mechanisms that regulate IFT (Burghoorn et al., 2007; Chen et al., 2006). Adding to the complexity of the IFT mechanism, C. elegans BBS protein orthologs (Table I) have been shown to traffic in cilia via the IFT machinery, and are required to maintain the interaction between the two anterograde motor complexes (Blacque et al., 2004; Ou et al., 2005, 2007; Pan et al., 2006). Specifically, disruption of individual BBS proteins (BBS-1, BBS-7, or BBS-8) causes the two anterograde motors to separate in MSs, concomitant with the separation of IFT subcomplexes A and B (Fig. 5). The working model arising from these findings is that in MSs, the two kinesin-2 motors exist in a tug-of-war state (because of their different intrinsic translocation motilities) and that through intermolecular interactions, proteins such as BBS components prevent the motors from dissociating (Ou et al., 2005; Pan et al., 2006). Although the ciliary structure is only slightly compromised (truncated) in bbs mutants, an interesting additional discovery from these mutants has been that Kinesin-II remains associated with all known IFT-A components, moving at a unified slow rate of 0.5 µm/s, whereas OSM-3 remains associated with known IFT-B components and collectively travels at 1.2 µm/s (Fig. 5) (Ou et al., 2005, 2007). On this basis, several novel C. elegans IFT-associated proteins were proposed to associate with IFT-A (IFTA-1) or IFT-B (CHE-13, IFTA-2, DYF-1, DYF-2, DYF-3, DYF11, and DYF-13) (Table I) (Efimenko et al., 2006; Li et al., 2008; Ou et al., 2007). Four of these, namely, DYF-1, DYF-3/Qilin, DYF-11, and DYF-13, have now been confirmed as associating with the proposed IFT-B module (Follit et al., 2009; Omori et al., 2008); others have been recently validated in a short IFT review by Cole and Snell (2009). Furthermore, in the context of how the large IFT macroassemblies are structurally organized, the data from the bbs mutants suggest that IFT-A proteins are more closely/tightly associated to the Kinesin-II motor, whereas IFT-B proteins are more closely/tightly associated with OSM-3. Lastly, disrupting either motor in a bbs mutant apparently prevents the tug-of-war between the motors and restores the association of the IFT-A and IFT-B subcomplexes (not shown), such that they move together at the speed of the residual motor (Pan et al., 2006); this suggests that the two discrete IFT subcomplexes are likely to be physically associated within the IFT machinery.

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In addition to specific molecular defects in IFT rates and cilium morphologies, defects in the functions of the majority of IFT-associated proteins causes a variety of sensory and/or signaling anomalies that can be studied in the context of animal nematode behavior and physiology. For example, most mutants have chemosensory (e.g., osmoavoidance, chemotaxis) and lifespan defects; a nonexhaustive list of additional phenotypes includes small body sizes, lipid homeostasis anomalies, altered foraging/roaming behaviors, and thermosensory defects (Apfeld and Kenyon, 1999; Bae and Barr, 2008; Bargmann, 2006; Fujiwara et al., 2002; Inglis et al., 2007; Kobayashi et al., 2007; Li et al., 2008; Perkins et al., 1986). The sensory and behavioral phenotypes likely reflect changes in a number of signaling pathways, many of which have yet to be fully resolved. For example, disruption of IFT-associated genes results in lifespan alterations that may be largely attributable to defects in insulin/ IGF1-based signaling (Apfeld and Kenyon, 1998; Schafer et al., 2006). The ciliary phenotypes associated with the disruption of IFT-associated proteins are detailed in the relevant sections below. The dual-kinesin IFT pathway described for amphid and phasmid cilia, and variations thereof found in other cilia (e.g., AWB neuron cilia), may be very similar to that found in more complex animals—e.g., as suggested for vertebrate photoreceptors, which have a bipartite doublet–singlet microtubule ciliary structure and an OSM-3 ortholog (KIF17) (Silverman and Leroux, 2009; Insinna and Besharse, 2008). This, combined with the extensive GFP-tagged IFT strains and ciliary/IFT mutant toolkit available, makes C. elegans an appealing model system for the molecular analysis of IFT and ciliopathy-associated proteins. In this chapter, we will discuss approaches that have been used to uncover C. elegans IFT-associated proteins and summarize the various methods that can be used for their characterization. Many of the techniques presented are also generally applicable to the study of basal body/TZ-associated or ciliary proteins, including some which may play a role in the IFT process.

II. Identification of Novel IFT-Associated and Ciliary Proteins C. elegans researchers have invested considerable effort exploiting various genetic, genomic, and bioinformatic tools to identify the “ciliome” of the organism, that is, the complement of proteins required for proper cilium structure and function. Notably, many of the candidates uncovered have been associated with IFT. A landmark study by Swoboda et al. (2000) revealed that the restricted expression of most ciliary/IFT genes in the worm ciliated neurons is controlled by its sole RFX-type transcription factor, DAF-19, which recognizes a specific 14-bp regulatory element known as an X box. Subsequent indepth bioinformatic analyses (Blacque et al., 2005; Efimenko et al., 2005) have revealed that virtually all known C. elegans genes encoding IFT/BBS proteins possess X boxes and are regulated by DAF-19, and led to the identification of many novel IFT-associated

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proteins (Table I) that were not previously identified in the original Chlamydomonas biochemical purification of IFT subcomplexes A and B (Cole and Snell, 2009; Cole et al., 1998; Piperno and Mead, 1997). In parallel, a number of genome-wide transcriptional profiling studies have been employed to identify ciliated cell-specific genes, including comparative SAGE analysis of ciliated versus nonciliated cell types (Blacque et al., 2005; Colosimo et al., 2004; Etchberger et al., 2007), comparative microarray studies of daf-19(–) and daf19(þ) strains (Chen et al., 2006), and the isolation of mRNAs specifically expressed in ciliated neurons (Kunitomo et al., 2005). Additionally, putative IFT/ciliary genes can be revealed if their expression pattern in C. elegans is restricted to ciliated sensory neurons (e.g., see Fig. 1), in experiments performed either on promising candidates (Blacque et al., 2005; Chen et al., 2006; Efimenko et al., 2005; Kunitomo et al., 2005) or in genome-wide projects (Hunt-Newbury et al., 2007). Together, these bioinformatic and functional genomic studies in C. elegans have helped identify hundreds of known and candidate ciliary genes which represents an excellent resource, accessible in a webaccessible database at http://www.ciliome.com, for discovering novel proteins either directly or indirectly associated with IFT (Inglis et al., 2006). The resource is most useful when combined with other bioinformatic, genomic, and proteomic studies aimed at uncovering the ciliomes of other species with motile cilia (Gherman et al., 2006; Inglis et al., 2006) and, for example, the mouse ciliary photoreceptor proteome (Liu et al., 2007), which is nonmotile like all C. elegans sensory cilia. Another useful resource for identifying novel IFT proteins consists of the various genetic screens directed at uncovering C. elegans ciliogenic genes. Methods for conducting de novo genetic screens in C. elegans are elegantly summarized by Jorgensen and Mango (Jorgensen and Mango, 2002). In principle, however, any existing mutant possessing Dyf, Osm, Che, Odr, Ttx, Daf, Mev, and/or Avr phenotypes could result from the disruption of an IFT/ciliary gene (Bae et al., 2008; Fujii et al., 2004; Hedgecock et al., 1985; Ou et al., 2007; Perkins et al., 1986; Starich et al., 1995). Most genetic lesions have been mapped to a specific genomic region (typically a chromosome arm), and their cloning could reveal novel ciliary/IFT genes. There exists a great variety of techniques used to resolve the genetic position of specific mutations, which include standard twoand three-point mapping strategies (Fay, 2006a,b), snip-SNP mapping (Wicks et al., 2001), whole-genome sequencing (Sarin et al., 2008; Shen et al., 2008), and sensitive comparative genomic hybridization studies capable of detecting small deletions or individual SNPs (Maydan et al., 2007, 2009). Typically, several of the above approaches can be used in a complementary manner in order to greatly expedite mutant identification. A list of candidate genes can also be generated by cross-referencing mapped regions with “ciliomic” data described above. Direct confirmation of the mutation can be done in several ways, most notably rescue by microinjection of the corresponding wild-type gene (e.g., with a promoter-cDNA/genomic DNA construct, or an available Fosmid) (Evans, 2006).

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III. Characterization of Candidate and Known IFT-Associated Proteins A. Expression Analysis of Candidate IFT-Associated Genes Using Promoter–GFP Fusion Constructs A typical first step in characterizing a putative IFT or ciliary gene is to observe its expression pattern in C. elegans and assess if it is restricted or enriched for in ciliated neurons. This may be an important step in prioritizing a list of candidates, especially since this experimentation can be easily carried out in a high-throughput fashion. For some genes of interest, expression patterns may already have been determined via large-scale expression projects (Dupuy et al., 2007; Hunt-Newbury et al., 2007), whose results are freely available in Wormbase (http://www.wormbase.org). For all other genes, one must create one or more transgenic strain(s) that harbor an expression construct consisting of upstream promoter sequences fused to GFP, and subsequently assess the cell/tissue distribution using fluorescence microscopy. This procedure is described below. Expression constructs are typically created either by subcloning into a GFP (or other fluorescent protein) expression vector (e.g., pPD95.75) (Boulin et al., 2006), using a PCR fusion-based method termed “stitching” (Hobert, 2002; Hunt-Newbury et al., 2007) or by introduction into a Gateway vector (Reboul et al., 2001); notably, the latter two approaches are particularly amenable to high-throughput expression studies. Whichever method is utilized, a typical construct should have at least 1.5 kb of upstream 50 sequence (or the full intergenic 50 UTR), several codons from the ORF of the gene (including the ATG start codon) fused to in-frame to GFP, and a 30 UTR terminator sequence (typically from the unc-54 or let-858 gene). Furthermore, a 50 nuclear localization signal sequence is normally incorporated so that fluorescence signals are brightest in the nucleus, which aids in cell identification. The reader is directed to the article by Boulin and colleagues (2006) for additional details on the generation of reporter gene constructs. Strain(s) carrying the transcriptional fusion construct are then generated by introducing the transgene using standard microinjection techniques and injection markers; useful markers include rol-6(su1006), which marks transgenic strains with a dominant, distinctive rolling phenotype, or dpy-5(þ), which rescues the dpy-5(e907) dumpy phenotype (see, e.g., Li et al., 2008). The generation of transgenic strains via microinjection or microparticle bombardment is described in detail in Evans (2006). The extrachromosomal arrays obtained are normally adequate to determine expression patterns. Visualization of expression patterns can be achieved using conventional epifluorescence or confocal microscopy. Subsequent assignment of specific cells expressing GFP can then be performed using the known anatomical data available for the worm and individual cells, focusing on the 60 ciliated neuron bodies that cluster mainly in the head of the animal and to a lesser extent, its tail (Bae and Barr, 2008; Bargmann, 2006; Inglis et al., 2007) (see also WormAtlas; http:// www.wormatlas.org). Expression of the candidate IFT/ciliary gene in some, most,

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or all of the 60 ciliated sensory neurons, but potentially not in other nonciliated cell types, is a good indication that a gene may have cilia-specific functions (Blacque et al., 2005; Chen et al., 2006; Efimenko et al., 2005; Kunitomo et al., 2005). Typical expression patterns observed in adult hermaphrodites (amphid and phasmid neurons) and males (tail), as well as in two embryonic stages, are shown in Fig. 1 for a representative gene (bbs-8) expressed in all ciliated neurons. For these expression studies, there are several potential caveats to bear in mind. For a proportion of promoter-GFP fusion constructs, no expression pattern may be observed. Another problem may be that the observed expression pattern does not reflect the endogenous expression characteristics of the gene. Both problems likely occur because the gene fusion is not expressed in its natural genomic context. In such cases, creating different promoter–GFP fusions (e.g., with a more extensive 50 UTR or by incorporating one or more introns/exons before fusing to GFP) may be helpful. Alternatively, a recombineering approach wherein GFP is introduced in a large genomic fragment (usually a Fosmid) may create a construct with the correct expression pattern (Sarov et al., 2006; Zhang et al., 2008). Another caveat may be that the transgene is not expressed in all of the relevant cells at once, creating mosaic expression patterns, or a situation in which transmission from animal to animal is inefficient. In this case, one should opt to integrate the extrachromosomal arrays into the genome using irradiation (Evans, 2006). B. Subcellular Localization of Candidate IFT-Associated Proteins Using GFP-Tagged Fusion Proteins Although restricted expression in ciliated sensory neurons is a hallmark of C. elegans ciliary/IFT genes, determining the subcellular localization of a protein of interest is necessary for its assignment and characterization as a basal body/ ciliary, or more specifically, IFT-associated protein. Because IFT is a dynamic cellular process, it is necessary to construct a translational protein fusion with GFP (or a suitable variant) for live time-lapse microscopy. Generating such protein constructs shares the same overall strategy as making transcriptional reporters (Section III.A), with the exception that, typically, the entire coding region of the gene is fused to GFP at the C-terminus and no NLS is included (see, e.g., Blacque et al., 2004). One can use cDNA, although the expression characteristics of the entire genomic sequence (introns and exons) is usually preferable. Also, although the endogenous promoter of the gene can be used to make the translational fusion constructs, one may also substitute a generic ciliated cell-specific promoter that is deemed suitable, such as a bbs-1 promoter (Ansley et al., 2003; Blacque et al., 2005)—the advantage being that it may be more easily amenable to highthroughput studies (e.g., using Gateway vectors). Additional details regarding the generation of translational fusions are provided in Boulin et al. (2006). Visualization of the GFP-tagged protein should be performed in live animals immobilized with 10–15 mM of the muscle relaxant levamisole or 10 mM muscimol.

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Capturing still images of the GFP signal may reveal specific localization to basal body/ TZ regions and/or to ciliary axonemes (see Fig. 2). Real-time observation of the signal may also indicate a clear, bidirectional movement of the GFP-tagged protein along cilia—in particular those of amphid or phasmid neurons (see Kymographs in Fig. 6). Irrespectively, one should record time-lapse images of relevant amphid and phasmid cilia to assess the possibility of IFT (see the detailed methods presented in Section III.C, below). Representative time-lapse movies of GFP-tagged IFT-associated proteins can be seen in numerous publications (Blacque et al., 2004, 2005, 2006; Efimenko et al., 2006; Jauregui et al., 2008; Kunitomo and Iino, 2008; Mukhopadhyay et al., 2007). In generating and observing GFP-tagged proteins, at least four caveats deserve mentioning. First, making a suitable construct may be difficult due to the large

Fig. 6 Kymograph analyses of a GFP-tagged IFT-associated protein in C. elegans. Shown on the left are fluorescent still-images captured from time-lapse recordings of CHE-2::GFP (IFT80) in amphid cilia (left panels), along with the respective kymographs obtained from MSs and DSs. Also shown are schematics of individual particle movements, illustrating both anterograde and retrograde traces. The top panels are from a wild-type (N2) strain and bottom panels are from a bbs-8 mutant. Note the different particle velocities in the N2 middle and DSs (0.7 and 1.1 µm/s, respectively) and same velocity in the bbs-8 mutant (CHE-2 associates with OSM-3 and moves at the constant rate of 1.2 µm/s in both MS and DS). BB/TZ, basal body/ transition zone region.

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size of the genomic open reading frame, or size/availability of the cDNA. In such a situation, a recombineering approach, capable of dealing with extremely large coding regions, may be warranted (Sarov et al., 2006; Zhang et al., 2008). Second, it is typically very important to obtain transgenic strains that do not express the translational fusion construct excessively, creating GFP signals that are too bright and thus can obscure details of the protein localization (e.g., specifically to the basal body/TZ region, or the ciliary axoneme), and may prevent visualization of IFT (see below). Such overexpression can also increase the possibility that the protein will be seen in potentially nonphysiologically relevant regions, for example, in the cell body, and along dendrites, and might in principle create overexpression phenotypes that could affect the structures or functions of cilia. It is therefore important to create several strains that express different amounts of the transgene by titrating the amount of construct injected in the animal; as a general guideline, using decreasing amounts of constructs (typically 1–10 ng/µl) has proven to be useful. As with the transcriptional fusion constructs, integration of the extrachromosomal arrays may also be desirable for consistent (and low-level) expression across cell types. The use of a microparticle bombardment method rather than microinjection for introducing transgenes may be of assistance, as it is reported to favor the creation of integrated transgenic animals with low copy numbers of the DNA construct (Praitis, 2006). As a side note, unc119(þ) is commonly used as a bombardment marker (Ferguson and Fisher, 2009) but should probably not be used given that UNC-119 has basal body/ciliary functions in C. elegans (Ou et al., 2007) and elsewhere. The third important consideration is that visualization of motility for a bona fide IFT-associated protein may be obscured. One possibility is that the GFP-tagged protein is too highly concentrated (bright) in cilia, and the other possibility is that a significant fraction of the protein is immobile; the latter behavior could reflect a protein that requires IFT for its trafficking but is deposited within the ciliary axoneme, matrix, or membrane. In such situations, it may be useful to perform a fluorescence recovery after photobleaching (FRAP) study and observe whether the recovered signal, previously obscured, displays IFT-like motility. Finally, the overproduced tagged protein may not be functional; for example, addition of GFP to small GTPases may have negative consequences. To assess functionality, one should assess if the transgene encoding the tagged gene of interest rescues known phenotypes of available mutant alleles for the gene (see Section III.D below). In addition, if antibodies are available, one can investigate if the endogenous protein displays the same localization pattern as the GFP-tagged protein. If the construct is not functional, one possibility is that the GFP tag interferes with activity or protein–protein interactions; making an N-terminally tagged GFP fusion protein construct may solve this problem. After confirmation that a GFP-tagged protein undergoes IFT-like movement in cilia, it is of interest to determine whether the protein displays dynamic properties consistent with a specific association with the IFT machinery, by measuring velocities, testing for

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dependence on the IFT machinery for its movement, and potential role in ciliogenesis and C. elegans sensory/behavioral biology, as described below.

C. Measuring the IFT Velocities of GFP-Tagged IFT-Associated Proteins In Vivo

1. Key Considerations for Visualizing the Motilities of IFT-Tagged Proteins It is most important that the expression level of the GFP-tagged protein construct is not too high, as strong fluorescence signals may interfere with IFT visualization and rate analysis, or too low, where the long exposure times required can lead to significant photobleaching effects. For well-established GFP-tagged IFT/BBS proteins, a typical time-lapse videomicroscopy recording on a standard epifluorescence compound microscope (100 lens; numerical aperture or N.A. of 1.4), equipped with a standard charge-coupled device (CCD) camera (see below), is 3 frames/s, for at least 30–60 s. Lower exposures of 100 or 200 ms may be possible with more sensitive cameras and relatively bright signals. These guidelines are usually adequate for robust measurement of IFT rates. However, the precise parameters employed for a specific GFP-tagged protein depends on its abundance (level) in cilia, the nature of the fluorophore (e.g., EGFP vs dsRed), and whether IFT visualization is in amphid, phasmid, or other ciliary types. In addition, parameters may differ markedly between different microscopy hardware setups (e.g., compound vs spinning disc systems and lowvs high-sensitivity cameras).

2. Minimum Microscopy Equipment Required Some microscope systems are optimized for performing time-lapse microscopy whilst reducing sample damage and photobleaching—e.g., spinning disc confocal microscopes—and these are clearly suitable for quantitative analyses of IFT in C. elegans. However, visualization and rate measurements of IFT do not require expensive or specialized equipment except for a standard widefield microscope setup that includes a good-quality CCD camera. From our collective experience with several different microscope systems, we suggest, as a general guide, the following minimum hardware requirement: a. Upright or inverted epifluorescence microscope; the stage need not be motorized. b. Appropriate filterset(s) for visualizing EGFP or other GFP derivatives. c. DIC or phase-contrast optics to visualize specific features within the amphid and phasmid regions of the worm. d. Light source (e.g., Xenon or Mercury lamp) that should ideally be attenuable. e. Low-powered objective lens for localizing the nematodes on the slide (e.g., 10 or 20). f. 63 or 100 oil objective, preferably with a N.A. of 1.3 or higher.

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g. Although a temperature-controlled stage may be optimal, we find that this is not necessary under the observation times/conditions. h. Sensitive CCD or EMCCD camera (e.g., CoolSNAP HQ from Photometrics, Orca AG from Hamamatsu, iXon DV885 camera from Andor technologies, or equivalent). Since the moving IFT particles are very small (<1 µm), cameras with high resolutions are preferred (e.g., pixel well areas of <10 µm). i. Suitable microscopy software and computer system for acquisition of time-lapse movies; the system should permit sufficiently fast acquisition and storage of data so as to record movies of 100–400 ms exposure per frame, and at least 2–3 frames/s.

3. Preparation of Nematodes and Slides for Time-Lapse Microscopy Protocols for the maintenance of C. elegans are described in detail by Theresa Stiernagle (Stiernagle, 2006). To prepare slides, wash well-fed worms off of standard agar plates with 1 ml M9 buffer into a 1.5-ml collection tube. Following a brief centrifugation (400g for 30 s), decant the supernatant and wash 2–3 using M9 buffer. Following the last wash, remove as much supernatant as possible, and add 100–200 µl of 10–15 mM levamisole (or 10 mM muscimol) to the worm pellet. Let sit for several minutes. Meanwhile, melt a solution of 2–4% agarose (prepared in H2O) in a microwave; this will be used to provide a sticky pad onto which the worms can rest on the slide. Prepare the pads by placing a drop of the agarose solution onto a standard glass slide and immediately stack a second slide on top of the drop, in an orientation perpendicular to the first slide. Do not pull the slides apart until you are ready to add worms to the pad. If using a nonintegrated strain with poor transmission of the GFP marker, it may be prudent at this point to transfer the worms from the 1.5-ml tubes back onto an unseeded NGM plate. Individual worms possessing the GFP-tagged protein marker can then be selected and transferred onto the agarose pad, being careful not to scratch the delicate surface. This transfer must be relatively rapid, as the agarose pad and the nematodes can desiccate quickly. After finishing the transfer, dispense onto the pad 1–2 µl of 10 mM levamisole or muscimol to immobilize the animals, and immediately place a glass coverslip on top. Alternatively, for integrated or high-transmission lines, 1–2 µl of the worm pellet (in a small amount of M9) can be applied directly onto the agarose pad using a 20-µl pipettor, where 1/3 of the pipette tip is cut off to prevent worms from sticking to the plastic tips. Add a further 1–2 µl of 10 mM levamisole or muscimol to the worm pad and then position the glass coverslip on top. Cilia and IFT can be observed with either a 63 or 100 oil objective using the aforementioned microscopy system requirements. IFT measurements are most easily made on the bundle of amphid channel cilia, which are found close to the head of the animal and are generally oriented along its main body axis (Fig. 2). It is also possible to visualize and measure IFT within the two pairs of phasmid cilia, found near the tail (Fig. 2), or in other ciliated sensory neurons (e.g., AWB)

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as reported (Mukhopadhyay et al., 2007). Normally, visualization of anterograde movement is significantly easier than retrograde movement (1.10 µm/s), probably because kinesin-mobilized IFT particle “trains” are comparatively larger in size (Iomini et al., 2001).

4. Time-Lapse Microscopy To properly visualize IFT, it is best to restrict the area of investigation to a suitable amphid or phasmid cilia bundle. Cropping this specific region using the acquisition software focuses on the region of interest and minimizes the computational processing lag times. Narrowing the observation region to the channel cilia will also reveal subtle movements of the worm, which can impair kymograph analyses (see below). Indeed, keeping the worms motionless is a major difficulty facing IFT analyses. Although anesthetics such as sodium azide are highly effective at restricting muscle activity, they inhibit ATP-dependent processes such as IFT. Levamisole and muscimol are therefore the most commonly used paralysis reagents for IFT measurements, but they are not always completely effective at inhibiting motion in every worm, particularly in the nose of the animal where the amphid cilia reside. Immobilizing worms by gluing (Richmond, 2006) has not been reported in the literature for IFT measurements, likely because it would be more laborious, and cilium-sensed volatile/soluble odorants in the glue could conceivably interfere with ciliary transport. On the other hand, microfluidic devices that facilitate immobilization of the worm and microscopic visualization of GFP signals (Dittman, 2009) might prove useful for IFT measurements. Cilia should ideally be entirely within the same focal plane so that the length of the axoneme in view is sufficient to allow kymographic detection of moving particles. If there is difficulty capturing sufficient axonemal structure in focus, capturing a movie with merged Z-stacks at each time point may be useful if the microscope system has a sufficiently fast motorized Z-stage and the camera is sensitive enough to permit brief exposures. However, by adjusting the focal plane, a sufficient in-focus length of axoneme can be visualized in almost every worm. The length of the recordings depends mostly on the strength of the fluorescence signal and photobleaching effects. These factors can often be limiting, restricting the recordings to short recording times (15–20 s or less) and thus a small overall number of frames. In such situations, it is important to optimize the recording parameters of exposure time, exposure gain, and intensity of fluorescence excitation. In addition, the use of highly sensitive CCD camera technology such as EMCCD is beneficial as it dramatically shortens the exposure times required, albeit often at the expense of image resolution. By considering the above-mentioned parameters, it is relatively easy to optimize IFT motility assays. Another consideration is that statistically relevant numbers of recordings are made for each IFT marker. Typically, a total of 100 or more particles are measured for each of the middle and DSs, from at least several (e.g., 10) different worms. When dealing with mutant strains where IFT may be impaired, and rates for

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any given IFT marker may be altered in the middle and/or DSs, it is important to perform appropriate statistical analyses for differences compared to wild-type—e.g., a Student’s t-test, to query potential differences in mean values, and a different statistical method to assay for variance.

5. Kymograph Analyses Because of the highly convoluted arrangement of the amphid channel cilia (or even those of the phasmid channel), automated or manual particle tracking applications that are found in many analysis software packages are largely ineffective. For this reason, we highly recommend the use of kymograph analyses. Figure 6 depicts such analyses performed in amphid cilia middle and DSs, both in wild-type and bbs-8 mutant animals. Several proprietary software packages, including Metamorph® (Molecular Devices, Downingtown, PA, USA), possess kymograph functions. However the free ImageJ analysis software (http://rsb.info. nih.gov/ij/) has an available kymograph plugin (MultipleKymograph; http://www. embl.de/eamnet/html/body_kymograph.html), which comes packaged with a rate analysis macro (Read velocities from tsp). It is important to note that the rates calculated by this macro are, in fact, incorrect; the x and y axes of the kymograph generated by MultipleKymograph need to be converted into practical values. The following is a brief, step-by-step description of IFT rate analysis using the ImageJ MultipleKymograph plugin: a. Obtain time-lapse recording of IFT, as described above. If opening the file in ImageJ, it is best to save your movie as a stacked TIFF file, making sure to include the calibration bar in the images. If the acquisition software encodes files compatible with the Bioformats importer ImageJ plugin (http://www.loci.wisc.edu/ ome/formats-imagej.html), it may be possible to open the files directly in ImageJ with various metadata (e.g., one can import Openlab LIFF files directly into ImageJ). b. Open file in ImageJ. c. Adjust the brightness/contrast for ideal resolution of individual particles. d. Convert the file to 8-bit, if it is not already. e. Generate tracks as outlined in the MultipleKymograph plugin instruction manual (http://www.embl.de/eamnet/downloads/kymograph/kymograph_description.pdf), or, alternatively, simply draw a line that traces the movement of particles, from base to tip. f. Measure the length of the kymograph line (in µm) by clicking on “Analyze,” then “Measure.” If imported with the Bioformats plugin, the output value should be the actual length. Otherwise, it will become necessary to convert the measured length by first measuring the length of the calibration bar in order to obtain the µm/ pixel ratio. g. Run the MultipleKymograph plugin (Click on “Plugins,” then “MultipleKymograph”). You will then be prompted to enter the line width. This must be an odd value and

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generally does not affect the output significantly. The kymograph will appear in a new window. It might need to be magnified several times before the lines become visible. h. Trace over an individual moving particle with the straight line tool. i. Measure the rate of particle movement by clicking on “Plugins,” then “Macros,” then “read velocities from tsp.” This macro needs to be installed every time ImageJ is opened. The output window will appear, showing values for “dx” (the number of pixels on the x axis covered by the trace line), “dy” (the number of pixels on the y-axis covered by the trace line), and “speed” (dx divided by dy). j. Convert the values on the kymograph into actual velocities. The x-axis conversion ratio is the actual length (in µm) of the kymograph track obtained in (f) divided by the number of pixels along the x axis of the kymograph. The y-axis conversion ratio is the actual time of the movie (in seconds) divided by the number of pixels along the y-axis of the kymograph. k. Multiply the uncalibrated speed obtained in (i) with the x-axis conversion and then divide this value by the y-axis conversion to obtain the actual velocity in µm/s. Note that all of the calculations described in this section can be expedited by producing a worksheet in a spreadsheet application, such as Microsoft Excel. D. Functional Characterization of IFT-Associated Proteins and Assignment to Distinct Modules As described in the Introduction, moving IFT assemblies can be subdivided into distinct protein modules or subcomplexes, namely, subcomplex A (IFT-A), subcomplex B (IFT-B), kinesin (Kinesin-II and OSM-3), dynein, BBS proteins, and IFT cargo. Proteins associated with each module generally show the same behavior and trafficking profiles in various mutant backgrounds (e.g., see Ou et al., 2007), and loss-of-function mutants of individual components within the same module typically exhibit comparable cilium structure, function and/or IFT proteintrafficking phenotypes. These principles appear to apply to all IFT-associated components, which include the “core” proteins required for the function of the IFT machinery (e.g., motors, IFT particle, BBS proteins) and the more “peripheral” cargo proteins that dock onto the core machinery but are not necessary for its operation. Indeed, the cargo “module” are of particular interest since very few bona fide IFT-associated cargos have been uncovered and the manner with which they dock onto the IFT machinery—e.g., direct binding to motor protein or IFT-A/ B modules—is virtually unknown (Blacque et al., 2008; Silverman and Leroux, 2009). In this section, we present experimental methods to functionally characterize IFT-associated proteins uncovered through methodologies discussed above in Sections III.A–C. In essence, two complementary experimental approaches are taken: (i) in Section III.D.1 below, we describe how to employ fluorescencetagged fusions of the protein of interest and analyze cilium localization and IFT behavior in various IFT mutant backgrounds; (ii) in Sections III.D.2–4, we discuss methods to analyze cilium structure/function and IFT motility in strains carrying a

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mutation in the novel ciliary/IFT gene of interest. These primarily genetic complementary approaches serve to generate IFT behavior and phenotypic profiles for novel IFT components, which can inform the assignment of the respective proteins to specific (or novel) IFT modules.

1. Analysis of GFP-Tagged IFT Proteins in IFT Gene Mutants Using standard C. elegans genetic crossing techniques, transgenes containing gfptagged fusions to the IFT gene of interest (Section III.B) are crossed into appropriate IFT gene mutants (described below). Transgenic animals are then assessed for the ciliary localization and IFT motility of the GFP-tagged protein of interest using methods outlined in Sections III.B and III.C. Below follows a description of the most useful mutant backgrounds for teasing out phenotypic profiles for IFT components, both known and novel. a. Wild-type background. Whilst most IFT module components display identical ciliary localization patterns and IFT motility behaviors in wild-type cilia (Fig. 2), one exception is the heterotrimeric Kinesin-II motor module. Consisting of two motor subunits (KLP-11, KLP-20) and an accessory protein (KAP-1), Kinesin-II ciliary localization is restricted to the MSs of amphid and phasmid channel cilia (Snow et al., 2004). Although not yet known, it is possible that “peripheral” IFT-associated proteins (e.g., cargoes or motor regulators) may be exclusively associated with the Kinesin-II motor module. If so, a key observation would be the exclusion of the novel IFT-associated protein from DSs of wild-type amphid/phasmid channel cilia. b. Bardet–Biedl syndrome mutants. The bbs gene mutants are probably the most informative for distinguishing components of different IFT modules. Since published work shows identical IFT phenotypes in all examined bbs mutants, we suggest to initially examine novel GFP-tagged IFT proteins in a single available bbs mutant (e.g., bbs-1, bbs-7/osm-12, or bbs-8) that has already been characterized insofar as its effect on the IFT process (Blacque et al., 2004; Ou et al., 2005, 2007; Pan et al., 2006). As described in the Introduction, IFT assemblies are destabilized in the MSs of bbs mutant amphid channel cilia, with Kinesin-II and OSM-3-kinesin motor modules uncoupled and moving at respective speeds of 0.5 and 1.2 µm/s (Fig. 5). IFT-A subcomplex components travel with the slow-moving Kinesin-II module (0.5 µm/s) and fail to enter DSs, whereas IFT-B components are associated with the fast moving OSM-3 motor module (1.2 µm/s) that translocates along both middle and DSs (Fig. 5). In addition, IFT-B proteins tend to accumulate at the tips of the middle and DSs of bbs mutants, whereas this does not appear to be the case for IFT-A proteins (Fig. 5) (Blacque et al., 2004, 2006; Ou et al., 2005, 2007). C. elegans IFTA-2, a putative cargo protein based on the finding that its disruption causes signaling anomalies but not IFT or ciliary structure defects (Schafer et al., 2006), has been observed to travel with the fast-moving OSM-3/IFT-B assemblies in the bbs-8 mutant (Ou et al., 2007). However, given the recent finding that disruption of the Trypanosoma RABL5

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ortholog causes ciliogenesis defects (Adhiambo et al., 2009), it is possible that this protein is not a cargo component per se but rather represents a “core” IFT protein whose function is less critical in the nematode. The only bona fide C. elegans IFTassociated cargoes, OCR-2 and OSM-9 (Colbert et al., 1997; Qin et al., 2005; Tobin et al., 2002), have not been assigned to an IFT module—a deficiency that will be of interest to address. For components of the BBS protein module or “BBSome” (Nachury et al., 2007), individual BBS subunits accumulate at the ciliary base and fail to enter the ciliary axonemes and undergo IFT when present in a mutant background affecting a different BBS component—e.g., BBS-1::GFP undergoes IFT in a wild-type background but not in a bbs-5 mutant, and the same holds true for other BBS::GFP and bbs mutant combinations (Fig. 5) (Ou et al., 2007). This suggests that disruption of one of the BBSome subunits may destabilize the entire BBSome assembly, which is perhaps not surprising since many of the mammalian BBS proteins (BBS1, 2, 4, 5, 7, 8, and 9) form a hetero-oligomeric protein complex (termed “BBSome”) that can be isolated biochemically (Nachury et al., 2007). Together, these phenotypic profiles for IFT module components in bbs mutants provide evidence that a novel IFT protein is (i) a BBS module component; (ii) a Kinesin-II or IFT-A module component; since these modules have different localization patterns in wild-type cilia (see Section III.D.1.a above), we can distinguish between these two modules; or (iii) an OSM-3-kinesin or IFT-B module component (cannot distinguish between the two). c. IFT subcomplex B mutants. In IFT-B mutants (e.g., che-2, che-13, or osm-5), most IFT proteins accumulate within the basal body/TZ region, fail to enter the severely truncated axonemes, and have IFT motility defects, consistent with the notion that the IFT-B module serve functions closely related to anterograde IFT (Blacque et al., 2008; Cole, 2003; Pedersen and Rosenbaum, 2008; Scholey, 2008). Such observations for IFT-B gene mutants are important as they can help to demonstrate that the ciliary translocation of a novel IFT protein is dependent on an intact IFT process, thereby strengthening the notion that the protein is an integral component of this process. Although IFT-B mutants are less useful than bbs mutants in assigning novel IFT-associated components to distinct modules, they may assist in providing evidence that the novel IFT protein associates with the IFT-B module. In certain IFT-B mutants, the ciliary mislocalization defects for some IFT-B components are more severe than other IFT-B components; specifically, in che-13 and osm-6 mutants (Collet et al., 1998; Haycraft et al., 2003), OSM-5::GFP is fully delocalized, failing even to accumulate at the basal body/TZ region, whereas OSM-6::YFP is found at the basal body/TZ of che-13 mutants (Haycraft et al., 2001, 2003). Also, in the reverse experiment, CHE-13::GFP accumulates at the TZs of osm-5 mutants (Haycraft et al., 2003). The differential effect of che-13 and osm-5 mutations on the ciliary localization of IFTB module components has been suggested to reflect a possible hierarchy in the assembly of the IFT-B complex, where incorporation of OSM-5 into the IFT-B module is dependent on the prior incorporation of CHE-13 (Haycraft et al., 2003). The inference is that in various IFT-B mutants, the IFT-B module still assembles partially,

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lacking specific components. In the case of che-13 mutants, the IFT-B module is severely disrupted, which in turn disrupts the incorporation of OSM-5. Since these observations appear specific to components of the IFT-B module, che-13 or osm-6 mutant alleles may in some cases serve as useful tools for providing evidence that a novel IFT protein associates with the IFT-B module. In Section III.D.1.d below, we further discuss how analysis of IFT in a mutant allele of a known or novel IFT protein could serve to place components of a particular module within a possible hierarchical topology. d. IFT subcomplex A mutants. In IFT-A mutants (e.g., che-11 or daf-10), most IFT proteins accumulate within the truncated bulbous cilia (particularly at the distal tips) and fail to visibly undergo IFT, consistent with the notion that IFT-A modules serve functions closely related to retrograde IFT (Bell et al., 2006; Blacque et al., 2008; Cole, 2003; Pedersen and Rosenbaum, 2008; Qin et al., 2001; Scholey, 2008). Like investigations using IFT-B mutants, similar observations allow us to conclude that novel proteins are integral components of the IFT process. As proposed above for the IFT-B module, the assembly of IFT-A may also be hierarchical. Unlike daf-10 mutants where a light intermediate chain component of the IFT-Dynein module (XBX-1) accumulates within ciliary axonemes, XBX-1::YFP fails to enter che-11 mutant cilia (Schafer et al., 2003). Similarly, a putative IFT-A module component (IFTA-1) also failed to enter the cilium of a che-11 mutant, whereas in the reverse experiment, CHE-11::GFP accumulated in ifta-1 mutant cilia (Blacque et al., 2006). These findings hint at a model where CHE-11 incorporates into the IFT-A assembly before IFTA-1 and DAF-10. According to this model, the IFT-dynein module cannot dock with the poorly formed IFT-A modules in che-11 mutants but can dock with the less severely affected IFT-A modules in daf-10 mutants. As all of these observations appear specific to IFT-A and IFT-dynein module components, the che-11 mutant may be useful for providing evidence that a novel IFT protein is a component of the IFTdynein or IFT-A modules. e. Kinesin-II motor mutants. In the amphid channel cilia of Kinesin-II mutants (e.g., klp-11), all IFT proteins examined translocate with the OSM-3-kinesin motor at 1.2 µm/s along middle and DSs (Fig. 4). In the osm-3 kinesin mutant, IFT proteins are restricted to the MSs (the DSs are not built), where they associate with the Kinesin-II motor and move at 0.5 µm/s (Fig. 4). Observing such motility behaviors would serve to further demonstrate that a novel IFT protein is an integral part of the IFT system. In addition, if a novel IFT component associates exclusively with one of the Kinesin-II motor modules (e.g., cargo proteins or motor regulators), we would predict that this component would not be trafficked at all in the respective kinesin-2 mutant. However, to date, such additional tightly associated Kinesin-II moduleassociated components have not been uncovered; even DYF-1, which regulates the function of OSM-3 by allowing it to dock onto the IFT particle, still undergoes IFT in the MS of osm-3 mutant animals by way of its presumably indirect association with the Kinesin-II motor (Ou et al., 2005).

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2. Disrupting the Gene of Interest Just as the behavior of a known or novel IFT protein in various existing IFT gene mutants is highly informative, much information can be gleaned from the observation of GFP-tagged proteins in a strain carrying a mutant allele of the known or novel gene of interest. Luckily, cilia are not essential for C. elegans viability, facilitating the obtention and characterization of ciliary mutants. Such mutants may be uncovered, as discussed in Section II, in genetic screens for C. elegans animals with ciliaassociated phenotypes. Alternatively, using a reverse genetic approach, mutations in candidate genes that are shown or suspected to encode ciliary/IFT-associated proteins can be obtained. If a mutant strain is not already available, requests for knockouts can be made to the C. elegans Gene Knockout Consortium (http://celeganskoconsortium. omrf.org) or the National BioResource Project (http://www.shigen.nig.ac.jp/c.elegans/ index.jsp). Alternatively, deletion mutants can be generated by in-house mutagenesis and PCR screening. One possibility is to screen for imprecise Mos1 transposon excision if an appropriate parental Mos1 strain is available (Robert and Bessereau, 2009). Trimethyl psoralen (TMP)-UV mutagenesis can also be used to generate null deletion alleles (Ahringer, 2006). Although RNA interference (RNAi) is widely known to be generally ineffective in sensory (ciliated) neurons, even with the use of RNAisensitive strains such as eri-1, rrf-3 and eri-1;lin-15B, a promising approach is to express double-stranded RNA directly in the ciliated neurons of interest (Esposito et al., 2007). Once a suitable mutant is obtained, care must be taken to determine if a fortuitous gene duplication did not take place (by PCR screening for the wild-type open-reading frame), and the mutant must be outcrossed to the wild-type (N2) strain at least five times to eliminate possible secondary mutations in other genes. A critical first step in analyzing the mutant is to determine if introduction of the wild-type gene rescues any observable phenotype uncovered (see below).

3. Analyzing the Structural Integrity of Cilia in the Mutant of Interest One may be able to quickly assess the severity of a mutation in a ciliogenic gene with a simple dye-filling (Dyf) assay (an example of which is shown in Fig. 7). Briefly, worms are exposed for approximately 1 h to a lipophilic dye, such as DiI, DiO, or FITC; typically, one uses a 2-mg/ml stock diluted from 1:200 to 1:1000 in M9 buffer. Wild-type (N2) worms take up the dye through exposed amphid and phasmid ciliary membranes, causing the dendrites and cell bodies of the sensory neurons to fluoresce (Hedgecock et al., 1985; Starich et al., 1995). While all IFT/BBS gene mutants characterized to date display severe Dyf phenotypes (that is, a complete or nearcomplete loss of dye uptake), certain mutations, including those altering genes associated with nephronophthisis (nphp), result in more subtle Dyf defects that are observed at lower concentrations of dye (e.g., 2 mg/ml DiI diluted 1:1000 in M9 buffer) (Jauregui et al., 2008). In some cases, expression of GFP-tagged IFT reporters can interfere with ciliogenesis in certain sensitized mutant backgrounds (e.g., OSM-6:: GFP in nphp-4 strain), raising the possibility of synthetic Dyf phenotypes (SynDyf)

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Fig. 7 Dye-filling assays can reveal potential structural defects (i.e., shorter axonemes) in the environmentally exposed cilia. Shown are fluorescent images of animals following incubation with DiI. Note the near absence of dye entry into the amphid and phasmid ciliated neurons of the IFT gene mutant che-11 compared to the wild-type animal.

(Jauregui et al., 2008); it may be prudent to perform Dyf assays to test for such dominant transgene effects. For the most part, Dyf phenotypes are reliably associated with abrogation of amphid/phasmid ciliary structures. However, they may also result from defects in dendritic extension (Heiman and Shaham, 2009) or in the organization of the sheath and socket cells that surround Dyf cilia (Perkins et al., 1986). Accordingly, a Dyf phenotype is indicative, but not proof, that cilium integrity/structure is compromised. To directly examine cilium structure, it is useful to introduce one or more ciliary GFP markers in the mutant of interest. One possibility is to examine the localization pattern of GFP-tagged IFT/BBS proteins introduced into the mutant by genetic crosses;

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using this approach, the lengths and overall shapes/orientation can be measured. Another possibility is to employ ciliated cell-specific transcriptional GFP markers, which diffusely localize throughout the cell, including the ciliary compartment. Since GFP accumulates at the basal body/TZ as a bulge of fluorescence, thereby labeling the base of the organelle, these markers are very useful for assessing cilium length and morphology. Particularly useful cell-specific markers include srb-6p::gfp, which can be used to label phasmids, and gcy-5p::gfp, which labels a sole amphid cell, ASER (Fig. 8) (Blacque et al., 2006). Alternatively, ciliary axonemes can be visualized in animals expressing a construct encoding a tagged version of b-tubulin (tbb-4) (Jauregui et al., 2008). In all cases, ciliary lengths and overall shapes can be measured and compared to wild-type animals. The nature of the ciliary defect(s) present in the mutant can also be resolved in much greater detail using TEM, which permits the direct visualization of the individual ciliary transitional fibers, TZs, as well as middle and DSs (Perkins et al., 1986; Ward et al., 1975). This technique, although technically challenging, has proven to be particularly useful in identifying subtle ciliary defects, for example, those found when the TZ-localized nephrocystin proteins are disrupted (Jauregui et al., 2008).

Fig. 8 Cilium length measurements using GFP reporters expressed specifically in ciliated neurons of transgenic animals. Shown are fluorescent images showing the labeling of the ciliated neuron ASER using a gcy-5p::GFP reporter (left panel), and PHA/PHB neurons using the srb-6p::GFP reporter (right panel). BB/TZ, basal body/transition zone region; CBs, cell bodies.

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Probing the ciliary structure defects present in the mutant of interest will help to shed light on its potential ciliogenic role. For example, severely truncated cilia may be indicative of a gene encoding an IFT-B-associated protein—a possibility that may be strengthened by assigning it to this module, as discussed in Section III.D.1) above. Somewhat shorter cilia containing IFT protein accumulations at the tip are indicative of an IFT-A mutant, an argument that again could be strengthened by assigning the encoded protein to this module (Section III.D.1); alternatively, the mutant may affect the function of IFT-dynein directly and cause prominent IFT protein accumulations within cilia. The presence of a slightly truncated cilium, with small IFT protein accumulations near the midpoint (between the MSs and DSs), and the tip of cilia, suggests that the mutation is in a bbs gene.

4. Assessing Possible IFT Defects in the Mutant of Interest To examine possible IFT defects in the mutant of interest, it is advantageous to introduce several GFP-tagged components of the IFT machinery and observe their localization and dynamic behavior. This is simply done by genetically crossing a wild-type strain containing a GFP-tagged IFT protein marker into your mutant background. For a thorough analysis, we recommend using IFT reporters for IFT subcomplex A (e.g., CHE-11::GFP), IFT subcomplex B (e.g., CHE-2::GFP, OSM-5:: GFP, OSM-6::GFP, or CHE-13::GFP), OSM-3::GFP, KAP-1::GFP (a subunit of Kinesin-II), XBX-1::YFP (dynein light intermediate chain subunit of IFT dynein; the CHE-3 dynein heavy chain protein has yet to be tagged), and any one of the many BBS proteins examined to date (BBS-1, 2, 3, 5, 7, and 8). Care should then be taken to compare the localization and motility (including IFT rate determinations; Section III.C) of each IFT-associated protein marker in the mutant and wild-type strains. These analyses should shed light on the nature of the IFT defects, if any. As discussed in detail in Section III.D.1, some of the problems may be severe and readily apparent (e.g., a particular IFT/BBS marker does not penetrate the cilia or undergo IFT), or may be more subtle, e.g., their velocities may be altered. Together, these observations, combined with assignment into putative IFT modules (Section III.D.1), will help to shed significant insights into the role of the known or novel IFT proteins in the IFT process. It should be noted that one may also design a screen for any such specific ciliogenesis defects using a mutant library and one or more GFP-IFT marker(s). For example, Ou and colleagues identified a number of different genetic loci from a Dyf mutant library that affect the cilia, based on observation of an OSM-6::GFP marker in the mutants (Ou et al., 2007). Similarly, a forward genetic screen for genes that affect the trafficking of a GFP-tagged ciliary protein can be conducted; for example, Bae et al. (2008) screened for mutants exhibiting mislocalization of the ciliary protein PKD-2 and uncovered a strain with a disruption in the xbx-1 gene, which encodes the IFT-associated dynein light intermediate chain protein.

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E. Roles of IFT-Associated Proteins in C. elegans Sensory-Dependent Behaviors and Signaling The ciliated neurons of C. elegans are responsible for conferring most, if not all, of the ability of the organism to transduce environmental stimuli and enable complex behaviors. The majority of its sensory capabilities and associated behaviors can be tested relatively easily and collectively represent an excellent means of assessing cilium-based sensory and signaling processes that may require the proper function of IFT-associated proteins. Below, we briefly summarize the various ciliary phenotypes observed in C. elegans. It should be noted that although the phenotypes are often associated with ciliary defects, they need not arise specifically from cilia dysfunction. Detailed methodologies for phenotypic assays are beyond the scope of this paper, but excellent protocols are readily available; for example, refer to the outstanding, all-inclusive article by Anne Hart (Hart, 2006). Many of the mutants isolated in the initial C. elegans mutagenesis screens performed over 30 years ago were specifically defective in chemotaxis (Che) (Dusenbery et al., 1975) and osmo-avoidance (Osm) behaviors (Culotti and Russell, 1978), and later were determined to result in ultrastructural defects to sensory cilia (Perkins et al., 1986). Indeed, specific cilia in both the head and the tail are responsible for recognizing and initiating responses to a large number of attractive and repulsive water-soluble and volatile chemical signals (Bargmann, 2006; de Bono and Maricq, 2005; Inglis et al., 2007; Melkman and Sengupta, 2004). Recognition of thermal signals, either physiological (15–25°C) or noxious (>30°C), also appears to rely on normal ciliary structure/function. For example, bbs mutants show defects in thermotaxis (Ttx) at normal temperatures and avoidance of high temperatures (Tan et al., 2007). Furthermore, IFT gene mutants are known to be impaired in sensing noxious temperatures (Wittenburg and Baumeister, 1999). Abrogating the function of the cilium-localized cyclic nucleotide-gated channels TAX-2 and TAX-4 also results in thermotactic phenotypes (Coburn and Bargmann, 1996; Komatsu et al., 1996). Ciliary genes are also implicated in the regulation of C. elegans roaming (food-searching) behavior and body size. Mutations in the genes encoding core IFT components IFT-74, IFT-81, and CHE2 (IFT-80) result in decreased body size and roaming characteristics (Fujiwara et al., 2002; Kobayashi et al., 2007); the roaming defect, albeit less pronounced, is also observed in other ciliary mutant strains, including nphp-4, tza-1/mksr-2, and tza-2/ mksr-1 (Williams et al., 2008). Cilia also appear to be involved in the regulation of additional physiological processes in C. elegans. Most IFT gene mutants live significantly longer than wildtype, hinting at an association between environmental perception and lifespan that depends on the DAF-2/DAF-16 IGF1/insulin signaling pathway (Apfeld and Kenyon, 1999). This notion has been supported by recent studies examining nematodes deficient in one or more of the following cilia-associated genes, namely, ifta-2, nphp-1, nphp-4, xbx-7/mks-1, tza-1/mksr-1, and tza-2/mksr-2 (Bialas et al., 2009; Schafer et al., 2006; Winkelbauer et al., 2005). The formation of the alternative, stress-resistant dauer

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lifestage can be impaired in some ciliary mutants, leading to a Daf phenotype (Fielenbach and Antebi, 2008). Finally, IFT-associated genes have also been implicated in alterations to lipid homeostasis in the worm. An allele of bbs-1(mg409) was identified in an enhancer screen for a mild lipid accumulation phenotype observed in the kat-1 mutant, which affects the gene encoding a 3-ketoacyl-coA thiolase involved in b-oxidation of fatty acids (Mak et al., 2006). Such an increased lipid accumulation phenotype is also observed when the IFT gene mutations che-2 and osm-5 are combined with kat-1. Similarly, disruption of the IFT-B component DYF-11/MIP-T3 causes a marked increase in lipid accumulation when compared to wild-type (Li et al., 2008).

IV. Conclusions IFT represents a motility system that involves a remarkably large number of different components—kinesin and dynein molecular motors, the multiprotein IFT subcomplexes A and B, BBS proteins, regulatory factors, and ciliary cargo. Notwithstanding its manifest complexity, it is likely that yet additional IFT-associated components will be uncovered. The principles and techniques outlined in this chapter are intended not only to facilitate the characterization of such novel players in C. elegans, but also to assist in the continued functional dissection of known IFT-associated proteins. It should be noted that many components not directly associated with the IFT machinery within the cilium may be of critical importance to the ciliary trafficking process; various cell body proteins may help to target IFT/BBS proteins to their docking site at the basal body, and other factors could participate in their assembly. For example, the basal body/TZ-localized nephrocystin proteins NPHP-1 and NPHP-4 were recently studied in the context of an assembly role for the IFT-BBS machinery (Jauregui et al., 2008). The remodeling of the IFT machinery that occurs during the transition of anterograde-to-retrograde transport is likely also facilitated by a number of proteins, one of which is EB1, a microtubule plus-end tracking protein EB1 that interacts with IFT172 (Pedersen et al., 2005) at the ciliary tip. In brief, C. elegans, in conjunction with other excellent model system (e.g., Chlamydomonas, Tetrahymena, Trypanosoma), is well poised to dissect in great detail the molecular mechanisms of IFT, ostensibly one of the most complex and biomedically important intracellular transport system of eukaryotes.

Acknowledgments We thank Maureen Barr for providing helpful comments on the manuscript. This work was funded by the Canadian Institutes of Health Research (CIHR grant MOP-82870 to M.R.L.) and a Science Foundation Ireland President of Ireland Young Researcher Award (to O.E.B.). P.N.I. acknowledges doctoral scholarships from the Natural Sciences and Engineering Research Council of Canada (NSERC) and the Michael Smith Foundation for Health Research (MSFHR), and M.R.L. is the recipient of a MSFHR senior scholar award.

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299

References Absalon, S., Blisnick, T., Kohl, L., Toutirais, G., Dore, G., Julkowska, D., Tavenet, A., and Bastin, P. (2008). Intraflagellar transport and functional analysis of genes required for flagellum formation in trypanosomes. Mol. Biol. Cell 19, 929–944. Adhiambo, C., Blisnick, T., Toutirais, G., Delannoy, E., and Bastin, P. (2009). A novel function for the atypical small G protein Rab-like 5 in the assembly of the trypanosome flagellum. J. Cell Sci. 122, 834–841. Ahringer, J., ed. (2006). Reverse genetics. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–43. doi/10.1895/wormbook.1.47.1, http://www.wormbook.org. Ansley, S.J., Badano, J.L., Blacque, O.E., Hill, J., Hoskins, B.E., Leitch, C.C., Kim, J.C., Ross, A.J., Eichers, E.R., Teslovich, T.M., Mah, A.K., Johnsen, R.C., et al. (2003). Basal body dysfunction is a likely cause of pleiotropic Bardet-Biedl syndrome. Nature 425, 628–633. Apfeld, J., and Kenyon, C. (1998). Cell nonautonomy of C. elegans daf-2 function in the regulation of diapause and life span. Cell 95, 199–210. Apfeld, J., and Kenyon, C. (1999). Regulation of lifespan by sensory perception in Caenorhabditis elegans. Nature 402, 804–809. Bacaj, T., Lu, Y., and Shaham, S. (2008). The conserved proteins CHE-12 and DYF-11 are required for sensory cilium function in Caenorhabditis elegans. Genetics 178, 989–1002. Badano, J.L., Mitsuma, N., Beales, P.L., and Katsanis, N. (2006). The ciliopathies: An emerging class of human genetic disorders. Annu. Rev. Genomics Hum. Genet. 7, 125–148. Bae, Y.K., and Barr, M.M. (2008). Sensory roles of neuronal cilia: Cilia development, morphogenesis, and function in C. elegans. Front. Biosci. 13, 5959–5974. Bae, Y.K., Lyman-Gingerich, J., Barr, M.M., and Knobel, K.M. (2008). Identification of genes involved in the ciliary trafficking of C. elegans PKD-2. Dev. Dyn. 237, 2021–2029. Bargmann, C. I. (2006). Chemosensation in C. elegans. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–29. doi/10.1895/wormbook.1.123.1, http://www.wormbook.org. Barr, M.M., and Garcia, L.R. (2006). Male mating behavior. WormBook 1–11. Bell, L.R., Stone, S., Yochem, J., Shaw, J.E., and Herman, R.K. (2006). The molecular identities of the Caenorhabditis elegans intraflagellar transport genes dyf-6, daf-10 and osm-1. Genetics 173, 1275–1286. Bialas, N.J., Inglis, P.N., Li, C., Robinson, J.F., Parker, J.D., Healey, M.P., Davis, E.E., Inglis, C.D., Toivonen, T., Cottell, D.C., Blacque, O.E., Quarmby, L.M., et al. (2009). Functional interactions between the ciliopathy-associated Meckel syndrome 1 (MKS1) protein and two novel MKS1-related (MKSR) proteins. J. Cell Sci. 122, 611–624. Blacque, O.E., Cevik, S., and Kaplan, O.I. (2008). Intraflagellar transport: From molecular characterisation to mechanism. Front. Biosci. 13, 2633–2652. Blacque, O.E., Li, C., Inglis, P.N., Esmail, M.A., Ou, G., Mah, A.K., Baillie, D.L., Scholey, J.M., and Leroux, M.R. (2006). The WD repeat-containing protein IFTA-1 is required for retrograde intraflagellar transport. Mol. Biol. Cell 17, 5053–5062. Blacque, O.E., Perens, E.A., Boroevich, K.A., Inglis, P.N., Li, C., Warner, A., Khattra, J., Holt, R.A., Ou, G., Mah, A.K., McKay, S.J., Huang, P., et al. (2005). Functional genomics of the cilium, a sensory organelle. Curr. Biol. 15, 935–941. Blacque, O.E., Reardon, M.J., Li, C., McCarthy, J., Mahjoub, M.R., Ansley, S.J., Badano, J.L., Mah, A.K., Beales, P.L., Davidson, W.S., Johnsen, R.C., Audeh, M., et al. (2004). Loss of C. elegans BBS-7 and BBS-8 protein function results in cilia defects and compromised intraflagellar transport. Genes Dev. 18, 1630–1642. Bobinnec, Y., Fukuda, M., and Nishida, E. (2000). Identification and characterization of Caenorhabditis elegans gamma-tubulin in dividing cells and differentiated tissues. J. Cell Sci. 113, 3747–3759. Boulin, T., Etchberger, J. F., and Hobert, O. (2006). Reporter gene fusions. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–23. doi/10.1895/wormbook.1.106.1, http://www.wormbook.org. Burghoorn, J., Dekkers, M.P., Rademakers, S., de Jong, T., Willemsen, R., and Jansen, G. (2007). Mutation of the MAP kinase DYF-5 affects docking and undocking of kinesin-2 motors and reduces their speed in the cilia of Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 104, 7157–7162.

300

Peter N. Inglis et al. Chen, N., Mah, A., Blacque, O.E., Chu, J., Phgora, K., Bakhoum, M.W., Newbury, C.R., Khattra, J., Chan, S., Go, A., Efimenko, E., Johnsen, R., et al. (2006). Identification of ciliary and ciliopathy genes in Caenorhabditis elegans through comparative genomics. Genome Biol. 7, R126. Coburn, C.M., and Bargmann, C.I. (1996). A putative cyclic nucleotide-gated channel is required for sensory development and function in C. elegans. Neuron 17, 695–706. Colbert, H.A., Smith, T.L., and Bargmann, C.I. (1997). OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans. J. Neurosci. 17, 8259–8269. Cole, D.G. (2003). The intraflagellar transport machinery of Chlamydomonas reinhardtii. Traffic 4, 435–442. Cole, D.G., Diener, D.R., Himelblau, A.L., Beech, P.L., Fuster, J.C., and Rosenbaum, J.L. (1998). Chlamydomonas kinesin-II-dependent intraflagellar transport (IFT): IFT particles contain proteins required for ciliary assembly in Caenorhabditis elegans sensory neurons. J. Cell Biol. 141, 993–1008. Cole, D.G., and Snell, W.J. (2009). SnapShot: Intraflagellar transport. Cell 137, 784–784.e1. Collet, J., Spike, C.A., Lundquist, E.A., Shaw, J.E., and Herman, R.K. (1998). Analysis of osm-6, a gene that affects sensory cilium structure and sensory neuron function in Caenorhabditis elegans. Genetics 148, 187–200. Colosimo, M.E., Brown, A., Mukhopadhyay, S., Gabel, C., Lanjuin, A.E., Samuel, A.D., and Sengupta, P. (2004). Identification of thermosensory and olfactory neuron-specific genes via expression profiling of single neuron types. Curr. Biol. 14, 2245–2251. Culotti, J.G., and Russell, R.L. (1978). Osmotic avoidance defective mutants of the nematode Caenorhabditis elegans. Genetics 90, 243–256. de Bono, M., and Maricq, A.V. (2005). Neuronal substrates of complex behaviors in C. elegans. Annu. Rev. Neurosci. 28, 451–501. Deane, J.A., Cole, D.G., Seeley, E.S., Diener, D.R., and Rosenbaum, J.L. (2001). Localization of intraflagellar transport protein IFT52 identifies basal body transitional fibers as the docking site for IFT particles. Curr. Biol. 11, 1586–1590. Dittman, J. (2009). Worm watching: Imaging nervous system structure and function in Caenorhabditis elegans. Adv. Genet. 65, 39–78. Dupuy, D., Bertin, N., Hidalgo, C.A., Venkatesan, K., Tu, D., Lee, D., Rosenberg, J., Svrzikapa, N., Blanc, A., Carnec, A., Carvunis, A.R., Pulak, R., et al. (2007). Genome-scale analysis of in vivo spatiotemporal promoter activity in Caenorhabditis elegans. Nat. Biotechnol. 25, 663–668. Dusenbery, D.B., Sheridan, R.E., and Russell, R.L. (1975). Chemotaxis-defective mutants of the nematode Caenorhabditis elegans. Genetics 80, 297–309. Efimenko, E., Blacque, O.E., Ou, G., Haycraft, C.J., Yoder, B.K., Scholey, J.M., Leroux, M.R., and Swoboda, P. (2006). Caenorhabditis elegans DYF-2, an orthologue of human WDR19, is a component of the intraflagellar transport machinery in sensory cilia. Mol. Biol. Cell 17, 4801–4811. Efimenko, E., Bubb, K., Mak, H.Y., Holzman, T., Leroux, M.R., Ruvkun, G., Thomas, J.H., and Swoboda, P. (2005). Analysis of xbx genes in C. elegans. Development 132, 1923–1934. Esposito, G., Di Schiavi, E., Bergamasco, C., and Bazzicalupo, P. (2007). Efficient and cell specific knockdown of gene function in targeted C. elegans neurons. Gene 395, 170–176. Etchberger, J.F., Lorch, A., Sleumer, M.C., Zapf, R., Jones, S.J., Marra, M.A., Holt, R.A., Moerman, D.G., and Hobert, O. (2007). The molecular signature and cis-regulatory architecture of a C. elegans gustatory neuron. Genes Dev. 21, 1653–1674. Evans, T. C., ed. (2006). Transformation and microinjection. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–15. doi/10.1895/wormbook.1.108.1, http://www.wormbook.org. Fan, Y., Esmail, M.A., Ansley, S.J., Blacque, O.E., Boroevich, K., Ross, A.J., Moore, S.J., Badano, J.L., May-Simera, H., Compton, D.S., Green, J.S., Lewis, R.A., et al. (2004). Mutations in a member of the Ras superfamily of small GTP-binding proteins causes Bardet-Biedl syndrome. Nat. Genet. 36, 989–993. Fay, D. (2006a). Genetic mapping and manipulation: Chapter 2—Two-point mapping with genetic markers. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–6. doi/10.1895/wormbook.1.91.1, http://www.wormbook.org.

14. Functional Genomics of Intraflagellar Transport

301

Fay, D. (2006b). Genetic mapping and manipulation: Chapter 3—Three-point mapping with genetic markers. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–7. doi/10.1895/wormbook.1.91.1, http://www.wormbook.org. Ferguson, A.A., and Fisher, A.L. (2009). Retrofitting ampicillin resistant vectors by recombination for use in generating C. elegans transgenic animals by bombardment. Plasmid 62, 140–145. Fielenbach, N., and Antebi, A. (2008). C. elegans dauer formation and the molecular basis of plasticity. Genes Dev. 22, 2149–2165. Fliegauf, M., Benzing, T., and Omran, H. (2007). When cilia go bad: Cilia defects and ciliopathies. Nat. Rev. Mol. Cell Biol. 8, 880–893. Follit, J.A., Xu, F., Keady, B.T., and Pazour, G.J. (2009). Characterization of mouse IFT complex B. Cell Motil. Cytoskeleton 66, 457–468. Fujii, M., Matsumoto, Y., Tanaka, N., Miki, K., Suzuki, T., Ishii, N., and Ayusawa, D. (2004). Mutations in chemosensory cilia cause resistance to paraquat in nematode Caenorhabditis elegans. J. Biol. Chem. 279, 20277–20282. Fujiwara, M., Ishihara, T., and Katsura, I. (1999). A novel WD40 protein, CHE-2, acts cell-autonomously in the formation of C. elegans sensory cilia. Development 126, 4839–4848. Fujiwara, M., Sengupta, P., and McIntire, S.L. (2002). Regulation of body size and behavioral state of C. elegans by sensory perception and the EGL-4 cGMP-dependent protein kinase. Neuron 36, 1091– 1102. Gherman, A., Davis, E.E., and Katsanis, N. (2006). The ciliary proteome database: An integrated community resource for the genetic and functional dissection of cilia. Nat. Genet. 38, 961–962. Ginger, M.L., Portman, N., and McKean, P.G. (2008). Swimming with protists: Perception, motility and flagellum assembly. Nat Rev Microbiol 6, 838–850. Hao, L., and Scholey, J.M. (2009). Intraflagellar transport at a glance. J. Cell Sci. 122, 889–892. Hart, A. C., ed. (2006). Behavior. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–67. doi/10.1895/wormbook.1.87.1, http://www.wormbook.org. Haycraft, C.J., Schafer, J.C., Zhang, Q., Taulman, P.D., and Yoder, B.K. (2003). Identification of CHE-13, a novel intraflagellar transport protein required for cilia formation. Exp. Cell Res. 284, 251–263. Haycraft, C.J., Swoboda, P., Taulman, P.D., Thomas, J.H., and Yoder, B.K. (2001). The C. elegans homolog of the murine cystic kidney disease gene Tg737 functions in a ciliogenic pathway and is disrupted in osm-5 mutant worms. Development 128, 1493–1505. Hedgecock, E.M., Culotti, J.G., Thomson, J.N., and Perkins, L.A. (1985). Axonal guidance mutants of Caenorhabditis elegans identified by filling sensory neurons with fluorescein dyes. Dev. Biol. 111, 158–170. Heiman, M.G., and Shaham, S. (2009). DEX-1 and DYF-7 establish sensory dendrite length by anchoring dendritic tips during cell migration. Cell 137, 344–355. Hildebrandt, F., Attanasio, M., and Otto, E. (2009). Nephronophthisis: Disease mechanisms of a ciliopathy. J. Am. Soc. Nephrol. 20, 23–35. Hobert, O. (2002). PCR fusion-based approach to create reporter gene constructs for expression analysis in transgenic C. elegans. BioTechniques 32, 728–730. Hunt-Newbury, R., Viveiros, R., Johnsen, R., Mah, A., Anastas, D., Fang, L., Halfnight, E., Lee, D., Lin, J., Lorch, A., McKay, S., Okada, H.M., et al. (2007). High-throughput in vivo analysis of gene expression in Caenorhabditis elegans. PLoS Biol. 5, e237. Inglis, P.N., Boroevich, K.A., and Leroux, M.R. (2006). Piecing together a ciliome. Trends Genet. 22, 491–500. Inglis, P. N., Ou, G., Leroux, M. R., and Scholey, J. M. (2007). The sensory cilia of Caenorhabditis elegans. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–22. doi/10.1895/wormbook.1.126.2, http://www.wormbook.org. Insinna, C., and Besharse, J.C. (2008). Intraflagellar transport and the sensory outer segment of vertebrate photoreceptors. Dev. Dyn. 237, 1982–1992. Iomini, C., Babaev-Khaimov, V., Sassaroli, M., and Piperno, G. (2001). Protein particles in Chlamydomonas flagella undergo a transport cycle consisting of four phases. J. Cell Biol. 153, 13–24.

302

Peter N. Inglis et al. Jauregui, A.R., and Barr, M.M. (2005). Functional characterization of the C. elegans nephrocystins NPHP-1 and NPHP-4 and their role in cilia and male sensory behaviors. Exp. Cell Res. 305, 333–342. Jauregui, A.R., Nguyen, K.C., Hall, D.H., and Barr, M.M. (2008). The Caenorhabditis elegans nephrocystins act as global modifiers of cilium structure. J. Cell Biol. 180, 973–988. Jorgensen, E.M., and Mango, S.E. (2002). The art and design of genetic screens: Caenorhabditis elegans. Nat. Rev. Genet. 3, 356–369. Kobayashi, T., Gengyo-Ando, K., Ishihara, T., Katsura, I., and Mitani, S. (2007). IFT-81 and IFT-74 are required for intraflagellar transport in C. elegans. Genes Cells 12, 593–602. Komatsu, H., Mori, I., Rhee, J.S., Akaike, N., and Ohshima, Y. (1996). Mutations in a cyclic nucleotide-gated channel lead to abnormal thermosensation and chemosensation in C. elegans. Neuron 17, 707–718. Kozminski, K.G., Johnson, K.A., Forscher, P., and Rosenbaum, J.L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523. Kunitomo, H., and Iino, Y. (2008). Caenorhabditis elegans DYF-11, an orthologue of mammalian Traf3ip1/ MIP-T3, is required for sensory cilia formation. Genes Cells 13, 13–25. Kunitomo, H., Uesugi, H., Kohara, Y., and Iino, Y. (2005). Identification of ciliated sensory neuronexpressed genes in Caenorhabditis elegans using targeted pull-down of poly(A) tails. Genome Biol. 6, R17. Li, C., Inglis, P.N., Leitch, C.C., Efimenko, E., Zaghloul, N.A., Mok, C.A., Davis, E.E., Bialas, N.J., Healey, M.P., Heon, E., Zhen, M., Swoboda, P., et al. (2008). An essential role for DYF-11/MIP-T3 in assembling functional intraflagellar transport complexes. PLoS Genet. 4, e1000044. Liu, Q., Tan, G., Levenkova, N., Li, T., Pugh, E.N.J., Rux, J.J., Speicher, D.W., and Pierce, E.A. (2007). The proteome of the mouse photoreceptor sensory cilium complex. Mol. Cell. Proteomics 6, 1299–1317. Mak, H.Y., Nelson, L.S., Basson, M., Johnson, C.D., and Ruvkun, G. (2006). Polygenic control of Caenorhabditis elegans fat storage. Nat. Genet. 38, 363–368. Marshall, W.F. (2008). Basal bodies platforms for building cilia. Curr. Top. Dev. Biol. 85, 1–22. Maydan, J.S., Flibotte, S., Edgley, M.L., Lau, J., Selzer, R.R., Richmond, T.A., Pofahl, N.J., Thomas, J.H., and Moerman, D.G. (2007). Efficient high-resolution deletion discovery in Caenorhabditis elegans by array comparative genomic hybridization. Genome Res. 17, 337–347. Maydan, J.S., Okada, M., Flibotte, S., Edgley, M., and Moerman, D.G. (2009). De novo Identification of Single Nucleotide Mutations in Caenorhabditis elegans Using Array Comparative Genomic Hybridization. Genetics 181, 1673–1677. Melkman, T., and Sengupta, P. (2004). The worm’s sense of smell. Development of functional diversity in the chemosensory system of Caenorhabditis elegans. Dev. Biol. 265, 302–319. Mukhopadhyay, S., Lu, Y., Qin, H., Lanjuin, A., Shaham, S., and Sengupta, P. (2007). Distinct IFT mechanisms contribute to the generation of ciliary structural diversity in C. elegans. EMBO J. 26, 2966–2980. Murayama, T., Toh, Y., Ohshima, Y., and Koga, M. (2005). The dyf-3 gene encodes a novel protein required for sensory cilium formation in Caenorhabditis elegans. J. Mol. Biol. 346, 677–687. Nachury, M.V., Loktev, A.V., Zhang, Q., Westlake, C.J., Peranen, J., Merdes, A., Slusarski, D.C., Scheller, R.H., Bazan, J.F., Sheffield, V.C., and Jackson, P.K. (2007). A core complex of BBS proteins cooperates with the GTPase Rab8 to promote ciliary membrane biogenesis. Cell 129, 1201–1213. Omori, Y., Zhao, C., Saras, A., Mukhopadhyay, S., Kim, W., Furukawa, T., Sengupta, P., Veraksa, A., and Malicki, J. (2008). Elipsa is an early determinant of ciliogenesis that links the IFT particle to membraneassociated small GTPase Rab8. Nat. Cell Biol. 10, 437–444. Orozco, J.T., Wedaman, K.P., Signor, D., Brown, H., Rose, L., and Scholey, J.M. (1999). Movement of motor and cargo along cilia. Nature 398, 674. Ou, G., Blacque, O.E., Snow, J.J., Leroux, M.R., and Scholey, J.M. (2005a). Functional coordination of intraflagellar transport motors. Nature 436, 583–587. Ou, G., Koga, M., Blacque, O.E., Murayama, T., Ohshima, Y., Schafer, J.C., Li, C., Yoder, B.K., Leroux, M.R., and Scholey, J.M. (2007). Sensory ciliogenesis in Caenorhabditis elegans: Assignment of IFT components into distinct modules based on transport and phenotypic profiles. Mol. Biol. Cell 18, 1554–1569.

14. Functional Genomics of Intraflagellar Transport

303

Ou, G., Qin, H., Rosenbaum, J.L., and Scholey, J.M. (2005b). The PKD protein qilin undergoes intraflagellar transport. Curr. Biol. 15, R410–R411. Pan, X., Ou, G., Civelekoglu-Scholey, G., Blacque, O.E., Endres, N.F., Tao, L., Mogilner, A., Leroux, M.R., Vale, R.D., and Scholey, J.M. (2006). Mechanism of transport of IFT particles in C. elegans cilia by the concerted action of kinesin-II and OSM-3 motors. J. Cell Biol. 174, 1035–1045. Pedersen, L.B., Miller, M.S., Geimer, S., Leitch, J.M., Rosenbaum, J.L., and Cole, D.G. (2005). Chlamydomonas IFT172 is encoded by FLA11, interacts with CrEB1, and regulates IFT at the flagellar tip. Curr. Biol. 15, 262–266. Pedersen, L.B., and Rosenbaum, J.L. (2008). Intraflagellar Transport (IFT) Role in Ciliary Assembly, Resorption and Signalling. Curr. Top. Dev. Biol. 85, 23–61. Pedersen, L.B., Veland, I.R., Schroder, J.M., and Christensen, S.T. (2008). Assembly of primary cilia. Dev. Dyn. 237, 1993–2006. Perkins, L.A., Hedgecock, E.M., Thomson, J.N., and Culotti, J.G. (1986). Mutant sensory cilia in the nematode Caenorhabditis elegans. Dev. Biol. 117, 456–487. Piperno, G., and Mead, K. (1997). Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella. Proc. Natl. Acad. Sci. USA 94, 4457–4462. Praitis, V. (2006). Creation of transgenic lines using microparticle bombardment methods. Methods Mol. Biol. 351, 93–107. Qin, H., Burnette, D.T., Bae, Y.K., Forscher, P., Barr, M.M., and Rosenbaum, J.L. (2005). Intraflagellar transport is required for the vectorial movement of TRPV channels in the ciliary membrane. Curr. Biol. 15, 1695–1699. Qin, H., Rosenbaum, J.L., and Barr, M.M. (2001). An autosomal recessive polycystic kidney disease gene homolog is involved in intraflagellar transport in C. elegans ciliated sensory neurons. Curr. Biol. 11, 457–461. Quinlan, R.J., Tobin, J.L., and Beales, P.L. (2008). Modeling ciliopathies: Primary cilia in development and disease. Curr. Top. Dev. Biol. 84, 249–310. Reboul, J., Vaglio, P., Tzellas, N., Thierry-Mieg, N., Moore, T., Jackson, C., Shin-i, T., Kohara, Y., ThierryMieg, D., Thierry-Mieg, J., Lee, H., Hitti, J., et al. (2001). Open-reading-frame sequence tags (OSTs) support the existence of at least 17,300 genes in C. elegans. Nat. Genet. 27, 332–336. Richmond, J.E. (2006). Electrophysiological recordings from the neuromuscular junction of C. elegans. In “WormBook” (The C. elegans Research Community, ed.), pp. 1–8. doi/10.1895/wormbook.1.112.1, http://www.wormbook.org. Robert, V.J., and Bessereau, J.L. (2009). Manipulating the Caenorhabditis elegans genome using mariner transposons. Genetica [Epub ahead of print]. Sarin, S., Prabhu, S., O’Meara, M.M., Pe’er, I., and Hobert, O. (2008). Caenorhabditis elegans mutant allele identification by whole-genome sequencing. Nat. Methods 5, 865–867. Sarov, M., Schneider, S., Pozniakovski, A., Roguev, A., Ernst, S., Zhang, Y., Hyman, A.A., and Stewart, A.F. (2006). A recombineering pipeline for functional genomics applied to Caenorhabditis elegans. Nat. Methods 3, 839–844. Satir, P., and Christensen, S.T. (2007). Overview of structure and function of mammalian cilia. Annu. Rev. Physiol. 69, 377–400. Satir, P., and Christensen, S.T. (2008). Structure and function of mammalian cilia. Histochem. Cell Biol 129, 687–693. Schafer, J.C., Haycraft, C.J., Thomas, J.H., Yoder, B.K., and Swoboda, P. (2003). XBX-1 encodes a dynein light intermediate chain required for retrograde intraflagellar transport and cilia assembly in Caenorhabditis elegans. Mol. Biol. Cell 14, 2057–2070. Schafer, J.C., Winkelbauer, M.E., Williams, C.L., Haycraft, C.J., Desmond, R.A., and Yoder, B.K. (2006). IFTA-2 is a conserved cilia protein involved in pathways regulating longevity and dauer formation in Caenorhabditis elegans. J. Cell Sci. 119, 4088–4100. Scholey, J.M. (2008). Intraflagellar transport motors in cilia: Moving along the cell’s antenna. J. Cell Biol. 180, 23–29.

304

Peter N. Inglis et al. Scholey, J.M., Ou, G., Snow, J., and Gunnarson, A. (2004). Intraflagellar transport motors in Caenorhabditis elegans neurons. Biochem. Soc. Trans. 32(Pt 5), 682–684. Shakir, M.A., Fukushige, T., Yasuda, H., Miwa, J., and Siddiqui, S.S. (1993). C. elegans osm-3 gene mediating osmotic avoidance behaviour encodes a kinesin-like protein. Neuroreport 4, 891–894. Sharma, N., Berbari, N.F., and Yoder, B.K. (2008). Ciliary dysfunction in developmental abnormalities and diseases. Curr. Top. Dev. Biol. 85, 371–427. Shen, Y., Sarin, S., Liu, Y., Hobert, O., and Pe’er, I. (2008). Comparing platforms for C. elegans mutant identification using high-throughput whole-genome sequencing. PLoS ONE 3, e4012. Signor, D., Wedaman, K.P., Rose, L.S., and Scholey, J.M. (1999). Two heteromeric kinesin complexes in chemosensory neurons and sensory cilia of Caenorhabditis elegans. Mol. Biol. Cell 10, 345–360. Silverman, M.A., and Leroux, M.R. (2009). Intraflagellar transport and the generation of dynamic, structurally and functionally diverse cilia. Trends Cell Biol 19, 306–316. Snow, J.J., Ou, G., Gunnarson, A.L., Walker, M.R., Zhou, H.M., Brust-Mascher, I., and Scholey, J.M. (2004). Two anterograde intraflagellar transport motors cooperate to build sensory cilia on C. elegans neurons. Nat. Cell Biol. 6, 1109–1113. Starich, T.A., Herman, R.K., Kari, C.K., Yeh, W.H., Schackwitz, W.S., Schuyler, M.W., Collet, J., Thomas, J.H., and Riddle, D.L. (1995). Mutations affecting the chemosensory neurons of Caenorhabditis elegans. Genetics 139, 171–188. Stephan, A., Vaughan, S., Shaw, M.K., Gull, K., and McKean, P.G. (2007). An essential quality control mechanism at the eukaryotic basal body prior to intraflagellar transport. Traffic 8, 1323–1330. Stiernagle, T. (2006). Maintenance of C. elegans. WormBook 1–11. Swoboda, P., Adler, H.T., and Thomas, J.H. (2000). The RFX-type transcription factor DAF-19 regulates sensory neuron cilium formation in C. elegans. Mol. Cell 5, 411–421. Tan, P.L., Barr, T., Inglis, P.N., Mitsuma, N., Huang, S.M., Garcia-Gonzalez, M.A., Bradley, B.A., Coforio, S., Albrecht, P.J., Watnick, T., Germino, G.G., Beales, P.L., et al. . (2007). Loss of Bardet Biedl syndrome proteins causes defects in peripheral sensory innervation and function. Proc. Natl. Acad. Sci. USA 104, 17524–17529. Tobin, D., Madsen, D., Kahn-Kirby, A., Peckol, E., Moulder, G., Barstead, R., Maricq, A., and Bargmann, C. (2002). Combinatorial expression of TRPV channel proteins defines their sensory functions and subcellular localization in C. elegans neurons. Neuron 35, 307–318. Veland, I.R., Awan, A., Pedersen, L.B., Yoder, B.K., and Christensen, S.T. (2009). Primary cilia and signaling pathways in mammalian development, health and disease. Nephron Physiol 111, p39–53. Ward, S., Thomson, N., White, J.G., and Brenner, S. (1975). Electron microscopical reconstruction of the anterior sensory anatomy of the nematode Caenorhabditis elegans. J. Comp. Neurol. 160, 313–337. Wicks, S.R., Yeh, R.T., Gish, W.R., Waterston, R.H., and Plasterk, R.H. (2001). Rapid gene mapping in Caenorhabditis elegans using a high density polymorphism map. Nat. Genet. 28, 160–164. Williams, C.L., Winkelbauer, M.E., Schafer, J.C., Michaud, E.J., and Yoder, B.K. (2008). Functional Redundancy of the B9 Proteins and Nephrocystins in C. elegans Ciliogenesis. Mol. Biol. Cell 19, 2154–2168. Winkelbauer, M.E., Schafer, J.C., Haycraft, C.J., Swoboda, P., and Yoder, B.K. (2005). The C. elegans homologs of nephrocystin-1 and nephrocystin-4 are cilia transition zone proteins involved in chemosensory perception. J. Cell Sci. 118, 5575–5587. Wittenburg, N., and Baumeister, R. (1999). Thermal avoidance in Caenorhabditis elegans: An approach to the study of nociception. Proc. Natl. Acad. Sci. USA 96, 10477–10482. Zaghloul, N.A., and Katsanis, N. (2009). Mechanistic insights into Bardet-Biedl syndrome, a model ciliopathy. J. Clin. Invest. 119, 428–437. Zhang, Y., Nash, L., and Fisher, A.L. (2008). A simplified, robust, and streamlined procedure for the production of C. elegans transgenes via recombineering. BMC Dev. Biol. 8, 119.

CHAPTER 15

Generating Conditional Mutants to Analyze Ciliary Functions: The Use of Cre–Lox Technology to Disrupt Cilia in Specific Organs Amber K. O’Connor*, Robert A. Kesterson†, and Bradley K. Yoder* *

Department of Cell Biology, University of Alabama at Birmingham, Birmingham, Alabama 35294

†

Department of Genetics, University of Alabama at Birmingham, Birmingham, Alabama 35294

Abstract I. Introduction II. The Design of a Conditional Targeted Allele A. Construct Generation B. Recombineering III. ES Clone Selection and Manipulation IV. The Cre Line V. The Reporter VI. Practical Notes VII. Conditional Alleles in Cilia Biology VIII. Methods A. Tissue Culture Media and Solutions B. Isolation of Targeting Vector DNA C. Preparation of Mouse Embryonic Fibroblasts D. ES Cell Growth and Electroporation E. Picking Drug-Resistant ES Clones and Creating Replica Plates F. Southern Analysis (adapted from Ramirez-Solis et al., 1992) G. Expansion and Verification of Targeted ES clones H. Preparing ES Cells for Microinjection Acknowledgments References

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Abstract The list of human disordered associated with cilia dysfunction, the ciliopathies, continues to highlight the importance of understanding the many roles of the long overlooked primary cilium. Much of the insights into the clinical importance of the cilium have come from analyses in model organisms, especially the mouse. However, the early embryonic lethality and severe developmental defects associated with cilia disruption has hindered progress in exploring cilia functions in late development or in adult tissues. This hurdle is being surmounted through the use of conditional alleles of genes encoding ciliary proteins and Cre deletor lines with inducible Cre activity or with lines expressing Cre in a cell-type-specific manner. Results from these approaches are providing important insights into the diverse array of cellular and tissue activities regulated by the cilium. Here we provide a recent account of the Cre/lox strategy. The generation and use of well-designed conditional alleles, as well as careful manipulation of embryonic stem cells are discussed. We also provide specific examples to illustrate the use of Cre/lox approaches to evaluate ciliary function in several tissues. With the recent characterization of multiple cilia proteomes along with efforts of several consortia to generate conditional alleles of all genes in the mouse, further use of conditional mutation approaches promise to yield many advances and surprises as we explore the functions of this increasingly complex organelle.

I. Introduction The advent of gene targeting has made the mouse a major research instrument for analyzing gene function and has established the mouse as an important model system for research into the molecular and physiological basis of human disease. Mouse models of the ciliopathies, the collective disease set caused by alterations in cilia structure or function, have been indispensable for determining the basic roles of primary cilia. However, classical gene-targeting approaches have limitations with regards to analyzing ciliogenic proteins since many of these mutations result in gestational lethality preventing analyses at later developmental stages, in specific tissues or cell types, or in adults. This limitation has been overcome using “conditional” alleles and Cre/lox technology that allow for spatial and temporal control of gene inactivation. Here we provide an overview of the technologies, strategies, and pitfalls that can be encountered when evaluating the role of cilia proteins via the Cre/lox system. First we will discuss the generation and use of alleles that are modified or “floxed” with flanking regulatory elements, resources to determine whether targeted embryonic stem (ES) cells might already be available, or if one should be engineered to more precisely fit research objectives. Of almost equal importance is selection of the ideal line of mice expressing Cre recombinase in the desired target tissue. It is through Cre expression that one gains control of when and

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where recombination occurs. We will also discuss the main options and features of some common reporter mouse lines as well as approaches to determine recombination efficiency. Lastly, we will briefly discuss examples of where these techniques have been applied in the past as a means of advancing cilia research.

II. The Design of a Conditional Targeted Allele Several conditional or floxed alleles of ciliogenic genes [i.e., ift88tm1Bky known here as ift88 fl (Haycraft et al., 2007); ift20tm1.1Gjp (Jonassen et al., 2008); and kif3atm2Gsn (Marszalek et al., 2000)] have been described. These alleles have been invaluable for determining the general importance of cilia and are readily available; however, the complete loss of cilia that results from these alleles is generally lethal. This helped to establish the cilium as an essential organelle, yet prevents critical analysis of ciliary function at later time points and in specific contexts. Thus, to explore more elaborate roles of the cilium, it has become necessary to employ more refined techniques to allow for temporally and spatially controlled disruption of ciliogenesis or cilia signaling activities. Candidate genes for these types of analyses are being identified through mutagenesis screens in simpler model organisms such as Caenorhabditis elegans. In addition, cilia or flagella proteomes have been reported for several species (Chlamydomonas reinhardtii and C. elegans) and mammalian cell types (photoreceptors, tracheal epithelium; Table I). Browsing these databases reveals a wealth of information regarding genes and potential pathways regulated by cilia. Once a gene of interest has been identified, the next step for ciliary analysis using mice as the model system is to search the databases of the targeting consortiums, such as KOMP and EUCOMM (Table II), to determine whether an allele is currently available. EUCOMM and KOMP construct conditional and classical gene knockout alleles and have been generating targeted ES cells at a rapid pace. If your gene is currently not on the targeting list of the consortium, justified requests are considered.

A. Construct Generation The generation of a novel floxed allele is completed through homologous recombination of a targeting vector in ES cells. When choosing the best way to make a conditional allele, it is often desirable to delete the entire gene of interest (or majority thereof), to ensure a true null mutation. This is done by inserting loxP sites (see below) on either side of the gene. These loxP sites have polarity and dictate how the DNA will be rearranged. Two loxP sites in the same orientation will cause the irreversible excision of the DNA between, while loxP sites that are in the opposite orientation (facing each other) cause inversion of the intervening DNA (Feil, 2007) leaving the LoxP sites available for subsequent inversion events possibly re-producing the wildtype allele. If deleting the whole gene is not feasible, a region of a gene may be selected

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Table I Cilia Proteomes Investigator

Organism

Katsanis Pazour Leroux Avidor-Reiss

Compilation Chlamydomonas reinhardtii Compilation Caenorhabditis elegans and Drosophila melanogaster

Blacque

C. elegans

Efimenko Li

C. elegans Homo saipan

Smith

Tetrahymena thermophila

Stolc

C. reinhardtii

Ostrowski

H. saipan

Pierce

Mus musculus

URL or reference http://www.ciliaproteome.org/ http://labs.umassmed.edu/chlamyfp/index.php http://www.ciliome.com Decoding cilia function: defining specialized genes required for compartmentalized cilia biogenesis (Avidor-Reiss et al., 2004) Functional genomics of the cilium, a sensory organelle (Blacque et al., 2005) Analysis of xbx genes in C. elegans (Efimenko et al., 2005) Comparative genomics identifies a flagellar and basal body proteome that includes the BBS5 human disease gene (Li et al., 2004) Robust method for proteome analysis by MS/MS using an entire translated genome: demonstration on the ciliome of T. thermophila (Smith et al., 2005) Genome-wide transcriptional analysis of flagellar regeneration in C. reinhardtii identifies orthologs of ciliary disease genes (Stolc et al., 2005) A proteomic analysis of human cilia: Identification of novel components (Ostrowski et al., 2002) The proteome of the mouse photoreceptor sensory cilium complex (Liu et al., 2007)

Table II Gene Trap/Conditional Allele Resources Consortium European Conditional Mouse Mutagenesis Program Knockout Mouse Project

International Gene Trap Consortium The Jackson Laboratory Nagy Lab Cre Database

Web address

Services

http://www.eucomm.org/

Generation of mutated genes in C57BL/6N ES cells using conditional gene trapping and gene targeting approaches Generation of mouse ES cells repository for alleles containing a null mutations

http://www.komp.org/ or http://www. knockoutmouse.org/ http://www.genetrap.org/ http://jaxmice.jax.org http://www.mshri.on.ca/ nagy/

A repository for publicly available gene trap cell lines Provide mouse breeding, repository, and research services Cre transgenic mouse line database

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for deletion with certain considerations. The first is that the region being deleted should be essential for normal function of the gene; this is generally accomplished by targeting the first coding exon. Second, it is advisable that deletion of the target region result in a change in the reading frame, unless the goal is to make in-frame deletions in your gene of interest (i.e., remove a potential protein interaction domain). Since the “upstream” loxP site is often placed into the promoter region containing key regulatory elements, care must be taken to not disrupt those domains when known. Lastly, it can be beneficial to engineer a unique enzyme site into your vector for linearization for the purpose of targeting and future screening (see below). Once these considerations are made, a targeting vector is engineered containing a large piece of the genome and conditional genetic elements that allow for controlled excision of genetic information; it is this vector that is linearized and used for homologous recombination in ES cells. Because the recombination elements described below are inserted into noncoding regions, they have a reduced likelihood of altering the expression of your and other genes; however, caution is necessary when analyzing phenotypes of both heterozygous and homozygous conditional alleles. Choosing where to insert the conditional elements is important as introns can contain regulatory elements. Aligning the genomic sequence with homologs from other species can help identify conserved intronic regulatory elements to be avoided. Detailed phenotyping is required to ensure that the conditional allele does not inhibit or alter gene function. As a first step for cilia mutants, crossing the conditional allele with a null, if available, and ensuring that the phenotype is the same as the heterozygous null should be done (e.g., demonstrate that the conditional allele is functionally wild type). A targeting vector is engineered with several kilobases (kb) of homology that flank the region being altered, typically a minimum of 2 kb on either side, with as much a 6 kb being recommended for increased efficiency (Fig. 1A). A lone loxP site is engineered into an intron which will be the 50 Cre recombination boundary; the 30 loxP site/Cre boundary is inserted in a downstream intron along with a neomycin resistance gene (neo, see below) for positive selection. The neo gene is flanked on both sides by Frt sites which are recognized by Flp recombinase. FLP is a recombinase (see below) similar to Cre that recognizes genetic sites referred to as Frt (Flp recombinase target) sites (Buchholz et al., 1998). The Neo gene allows for the selection of ES cells that have incorporated the targeting vector (thus killing off all other nontransfected cells) and subsequent enrichment of recombinant ES cells. The neo cassette can be excised by in vivo or in vitro Flp activity leaving two loxP sites flanking the region to be deleted by Cre. Although the inclusion of Frt sites makes vector design more complex, it has the benefit that it will facilitate subsequent removal of the selection marker after successful targeting. Use caution when leaving selection markers in place in vivo as they can interfere with the expression of nearby genes and possibly create hypomorphic mutations with reduced gene function; however, these may prove to be informative, recall that the hypomorphic mutation in the IFT88 gene in the ORPK (ift88Tg737Rpw) (Moyer et al., 1994; Pazour et al., 2000; Taulman et al., 2001) mouse was a revelation in cilia biology. It is also recommended that the second loxP site (the

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(A) Schematic overview of conditional allele generation. A targeting vector is generated with several kilobases of homology to the targeted genomic locus (thin black line). A lone 50 LoxP site (triangle) is inserted in an upstream intron. In a downstream intron a Neo cassette flanked by FRT sites (hexagons) is inserted, along with a 30 LoxP site. Together these sites delineate the boundary of the genomic region being deleted, represented in grey, boxes 3 and 4. After homologous recombination (dashed X-shaped lines) occurs in ES cells and selection against the vector backbone/TK gene (thick grey lines) is complete colonies containing the targeted allele are expanded, preserved, and characterized (Fig. 2). The generation of the conditional allele occurs when Flp recombinase initiates recombination at the FRTsites (dashed semicircle at hexagons) of the targeted allele removing the selection cassette, either in vitro or in vivo, leaving a FRTsite and the two LoxP sites flanking one or more exons (grey boxes). The final recombination step removing a segment of the gene (grey region boxes 3 and 4) to produce a null allele, occurs when Cre activity initiates and causes recombination at the LoxP sites (dashed semicircle at triangles), leaving one loxP site and removing the intervening DNA in vivo. (B) Schematic overview of conditional allele generation via recombineering. A BAC is chosen that contains the gene of interest (black region), and a targeting vector is generated with two homology arms in the same orientation as the sequence of the BAC (denoted ABC, DEF). The region of interest including the homology arms must contain the gene of interest and enough homology with the genome (i.e., several kilobases on both sides) to facilitate homologous recombination in ES cells. The gene of interest is captured from the BAC via recombination (dashed X-shaped lines, upper panel). Once the gene is in the targeting vector the first cassette, containing a kanamycin resistance gene (Kan) flanked by LoxP sites, is added in an upstream intron of the construct using recombineering (looped-arrow) resulting in the first targeting recombinant. A subsequent Cre recombination event is carried out in bacteria removing the Kan cassette (dashed semicircle at triangles) resulting in a derivative of the initial targeting recombinant containing only the 50 LoxP site. The second selection cassette, Neo flanked by FRT sites and a single LoxP site, is added in a 30 intron in the construct using recombineering resulting in the second targeting recombinant molecule, the final targeting vector. After homologous recombination occurs in ES cells and selection against the vector backbone is complete colonies containing the targeted allele are expanded, preserved, and characterized (Fig. 2). The generation of the conditional allele occurs when Flp recombinase initiates recombination at the FRT sites of the targeted allele removing the selection cassette, either in vitro or in vivo, leaving a FRTsite and the two LoxP sites flanking one or more exons. The final recombination step removing a segment of the gene to produce a null allele occurs when Cre activity initiates and causes recombinaiton at the LoxP sites, leaving one LoxP site and removing the intervening DNA in vivo.

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Targeting vectors have also been generated through PCR-based cloning. However, if PCR is used at any point in the generation of the targeting vector, it is essential that the final vector be sequenced. Any mutations within coding regions as well as in introns should be corrected with site-directed mutagenesis or a similar technique. It is recommended that isogenic DNA, isolated from the ES cells being targeted, be used for generating the homologous regions in the targeting vector as this is thought to increase recombination frequency (te Riele et al., 1992). B. Recombineering Recombineering is emerging as a new method for the generation of both transgenic and targeted mouse alleles without the use of traditional cut-and-paste cloning (Bouvier and Cheng, 2009; Sharan et al., 2009). The major advantage of the recombineering strategy is that once established, it can greatly increase the rate of vector construction and larger regions of homology can be accommodated that tend to increase the efficacy of homologous recombination in ES cells. Recombineering makes use of lambda Red phage recombinase enzymes and large pieces of the mouse genome, often in bacterial artificial chromosome (BAC) vectors. BACs can be identified and obtained using Ensembl or similar genetic browser. Using engineered strains of Escherichia coli that contain the three lambda recombinase genes exo, bet, and gam, recombination events between DNA molecules can be controlled. The expression of these genes is regulated by a temperature-sensitive repressor whose activity is alleviated at 42°C allowing their rapid transcription. The exo gene encodes an exonuclease that creates 30 single-stranded overhangs that are subsequently bound and protected by the Beta (bet gene) protein. To ensure that the endogenous recombination system (RecBCD) does not degrade the single-stranded genetic elements, gam acts as to inhibit that system. Together these proteins facilitate the recombination between two pieces of DNA with as little as 50 nucleotides of homology. The recombineering strategy employed for generating a final targeting construct varies based on application (knockout vs knockin vs conditional), see http://recombineering.ncifcrf.gov/ for general details, protocols, and links for ordering reagents or recent reviews by Sharan et al. (2009) and Bouvier et al. (2009) for comprehensive details about recombineering. There are multiple ways to generate conditional alleles via recombination, the most basic of which results in a vector that is similar to the one described above, although other strategies exist. The general protocol requires that the genomic segment of interest is retrieved from the appropriate BAC clone, via recombination, into a targeting vector (Fig. 1B), from which positive and negative selection is carried out using recombination to yield the final targeting construct. Briefly, a prokaryotic promoter-driven kanamycin (Kan) cassette flanked by loxP sites is inserted in an intron at the 50 conditional boundary. This cassette allows for the positive selection of recombinant bacteria through Kan resistance, subsequent Cre-mediated recombination of the Kan gene leaves a lone 50 loxP site for the conditional allele (this Cre recombination step is completed in the bacteria). The second recombination event inserts a eukaryotic promoter-driven neo cassette, flanked by Frt sites, and a second

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loxP site at the 30 conditional boundary as above (Fig. 1B). This neo cassette allows for the selection of recombinant ES cells and the subsequent removal of the cassette by Flp leaves the 30 loxP site intact. The ES cell targeting and mouse generation is the same with recombineered constructs as with the traditional constructs, the major advantage of the former is the rapid generation of the targeting vector. An alternative vector construction strategy used in the recent past was the three loxP strategy; however, it is inefficient, and the establishment of Flp has largely replaced this strategy. In general, animals were generated with a targeted allele maintaining the neo cassette which itself is flanked by loxP sites; these animals are crossed to a Cre line known to express in germ cells (see below) allowing natural generation of recombinants. This cross can produce both mice with conditional and mutant alleles depending on the efficiency of the Cre. The advantages of this method include the possibility of partially disrupting gene function in vivo with the neo cassette, and the chance of obtaining mutant animals in parallel with true conditional animals in one cross.

III. ES Clone Selection and Manipulation After the targeting vector is generated, a single wild-type or “normal” allele is converted to the targeted allele using homologous recombination in ES cells (protocol below). Briefly, ES cells are maintained on feeder cells in the presence of factors to inhibit differentiation [i.e., leukemia inhibitory factor (LIF)]. Your vector is cut (with your unique enzyme) and electroporated into the ES cells. After recovery, selection is applied to the ES cells, colonies grow and are isolated, expanded, preserved, and characterized. It is important to consider the genetic background of the ES cells you are using; historically most ES cells for this type of work are from the 129 inbred background, which are hardy, amenable to in vitro manipulation, and frequently contribute to the germline, though today B6 cells are also frequently used. Again, the use of isogenic DNA has been shown to increase the frequency of homologous recombination and BAC libraries are available for both the 129 line and B6 line. ES cells are derived from male embryos and generate male mice once injected into blastocytes, this permits targeting genes on the Y chromosome and facilitates the mating schemes used to establish a line of conditional mice (see below). Positive and negative selection schemes are used to enrich for correctly targeted ES clones. Positive selection is generally carried out with the neo gene; and negative selection, against random integration events, is normally done with the thymadine kinase (TK) gene. Neo imparts resistance to G418 which blocks peptide synthesis and kills nontransfected cells within 7 days helping to identify cells that have undergone recombination integrating the cassette. Negative selection is used to ensure that only the region of interest (within the targeting vector) was integrated to the genome. The TK gene is generally included in the vector backbone outside of the homology region and as such should be excluded from integration. TK is lethal in the presence of gancyclovir or 1-[2-deoxy]2-fluoro-b-D-arabinofuranosyl (FIAU), nucleotide analogs that require TK for their incorporation into the DNA. An alternative to TK is the use of

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the diphtheria toxin A fragment gene whose presence is toxic and requires no drug treatment to eliminate cells carrying the gene. Keep in mind that the feeder cells (mouse embryonic fibroblasts) used during ES cell selection need to be drug resistant as well. In most cases, using both negative and positive selection has been shown to substantially increase the frequency of obtaining correctly targeted ES cells. However, using positive and negative selection only increases the odds of obtaining properly targeted clones, and by no means should one assume that because a clone survives both selection processes that it is properly targeted. Once isolated ES clones have grown under selection, they need to be expanded and preserved while being characterized. Freezing down the cells at an early passage ensures the use of robust, young, ES cells for injection after they have been evaluated. Proper targeting can be confirmed through a variety of methods, with Southern blotting (Fig. 2) being the most definitive and reliable. It is important that the probes used be located outside of the homology arms such that one can distinguish between homologous versus nonhomologous recombination events. It is also essential that these probes be free of repetitive elements and tested on blots with nontargeted ES cell DNA prior to doing the ES cell screening to ensure that the restriction digest patterns agree with those predicted by sequence analysis (or previously published), and that the

(A)

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x 10 kb

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Fig. 2 Schematic representation of allele characterization via Southern blotting. DNA probes for Southern blot analysis are selected which are external to the targeting vector homology arms (grey lines, 50 and 30 below the thick black line representing the DNA) and will result in varied fragment sizes after digestion (enzyme sites denoted by “x” here only one enzyme is used). The wild-type allele (A) will yield a 10-kb fragment with both probes after digestion. The targeted allele (B) will give rise to two products due to the enzyme site within the selection cassette; 6.5 and 5 kb for the respective 50 and 30 probes. The conditional allele (C) will give rise to a band slightly larger than wild type, a 10.5-kb band with either probe, due to the insertion of FRT, LoxP, and intervening DNA. The null allele (D) yields one band with either probe that is smaller than the wild type due to the removal of the exons. Adapted with permission from “Animal models for disease: knockout, knockin, and conditional mutant mice” by LePage and Conlon (LePage and Conlon, 2006).

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probes are able to effectively detect a single copy of the gene. It may be necessary to engineer restriction enzyme sites into the targeting construct to facilitate fragment generation for blotting. Once a correctly targeted ES clone is established, it is also recommended that you confirm your targeting strategy by expressing Cre in the ES cells to ensure that the deleted allele can be generated and detected as well. After confirming successful integration of the conditional allele, the early passage ES clones are thawed, expanded, and injected into the blastocyst of a host that is genotypically distinct from the generator line, and chimeric mice are generated (unless this is going to be a routine procedure in a lab, this step is best performed by university core facilities). Normally, the host has a notable phenotypic variation, such as coat color, that will allow the easy identification of germline transmission. A common technique is to use an albino blastocyst host such that the offspring have a mixture of the wild-type (albino) and ES-derived (pigmented) cells; the male mice obtained (F0 generation) are chimeric as indicated by their marbled coat color. Subsequently, mating the chimeric males with albino females allows for the determination of whether the mutant cells (pigmented) contributed to the chimeric animal’s germ cells. If the F1 mice are pigmented, they have inherited genetic information from your ES cells. The repetitive generation of pigmented offspring in the F1 generation denotes hardy germline contribution of the ES cells; however, it does not automatically denote transmission of the mutant allele. After genotyping to confirm the passage of the targeted allele (which is expected to occur only 50% of the time in pigmented offspring), the heterozygous F1 offspring are then intercrossed to generate homozygous conditional animals.

IV. The Cre Line It is through Cre expression that one gains control of where and when gene deletion occurs once a conditional line is established. Cre recombinase is a tyrosine recombinase isolated from the bacteriophage P1 that catalyzes the cleavage and ligation of DNA at specific nucleotide recognition sites, loxP sites, via phosphoryl transfer (Hoess et al., 1990). As mentioned above these loxP sites have polarity and can cause excision or inversion of the intervening DNA. The distance between loxP sites can vary and this approach has been used to generate small and very large deletions, although the efficiency of deletion is decreased with longer distance. When choosing a Cre mouse line, the primary question is of your target, either a specific cell type, organ, or the whole animal. Many tissue- or cell-type-specific Cre lines have been generated; a quick literature search will inform you of the effectiveness, availability, leakiness, and reliability of a given Cre line. A partial list of available Cre mice can be found on the website for Dr Nagy’s Laboratory or via The Jackson Laboratory (Table II). If there does not appear to be a Cre line available to meet your needs, you may have to generate one; however, this can be a daunting task if expression patterns and promoter sequences have not been well characterized. One also needs to determine whether an inducible or constitutively active Cre line will be used.

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Constitutively active Cre lines are generally driven by an endogenous promoter that regulates the timing and pattern of Cre expression, with deletion of the targeted allele occurring once sufficient Cre is expressed. The timing of expression depends on the promoter and as such Cre can be expressed early in, and throughout development. This allows for analysis only at the onset of Cre expression. In the case of ciliogenic alleles, many constitutively active Cre lines prove fatal; however, such experiments can be utilized as proof of principle and to test the integrity of the conditional allele as they should mimic a null allele if deletion occurs widely and early enough. The use of inducible Cre lines allows for temporal control of Cre expression. CreERT2 (improved tamxoifen sensitivity from the earlier CreER versions) is one widely used variant where the recombinase enzyme is linked to a modified human estrogen receptor (ER) (Feil et al., 1996; Indra et al., 1999). This is an improvement over other technologies such as tetracycline-inducible Cres as they tend to be leakier and are more complicated requiring three components. CreER and CreERT2 have mutations engineered in the ER portion of the protein making them insensitive to endogenous estrogen compounds; a synthetic estrogen analog (tamoxifen) is used as an activating ligand. When unbound by ligand, the ER portion of the fusion inhibits nuclear import of Cre; however, when tamoxifen binds to the ER domain a conformational shift occurs revealing a nuclear import signal and the whole fusion protein can enter the nucleus. Once in the nucleus the Cre enzyme initiates recombination at loxP sites. Route of administration and dosage of tamoxifen should be optimized for each inducible Cre line. Efficient deletion often requires multiple daily injections. Further, tamoxifen remains in the system for about 48 h, which should be considered when wanting to generate deletions during a specific stage of development, especially with promoters that are expressed transiently. It is also worth noting that tamoxifen is not the active form of the ligand and must be processed by the liver to produce 4hydroxytamoxifen (Feil, 2007). In addition, tamoxifen itself may have estrogenic effects on a wide variety of tissues that can complicate the analyses of resulting phenotypes.

V. The Reporter After the conditional allele and Cre deletor lines are established, it is necessary to evaluate temporal and spatial activity of the Cre gene. Several reporter mice have been designed to assist in analysis of Cre activity and expression patterns. These reporter mice have their own characteristics that make them beneficial for specific analyses, some of the more widely used ones are described here. Mao et al. (1999) modified the Rosa26 gene trap originally generated by the Soriano laboratory (Friedrich and Soriano, 1991; Soriano, 1999) to express bgeo (b-galactosidase neo fusion, Gt(ROSA)26Sortm1Sho) after Cre-mediated deletion of “stopper” sequences (transcriptional and translational stop signals) upstream of the bgal gene, known here as Rosa26. When Cre activity occurs, the selection cassette and stop sequences are removed and bgal is expressed (Fig. 3B and E). Z/AP (Tg(CAG-geo/ALPP)1Lbe) is a

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dual Cre reporter designed by Lobe et al. (1999) that expresses bgal before Cre activity and the human alkaline phosphatase gene subsequent to Cre activity (Fig. 3F). Z/EG (Tg(CAGgeo/GFP)21Lbe) is a second dual Cre reporter engineered by Novak et al. (2000), which is a derivation of the Z/AP but after recombination the reporter gene expressed is the enhanced green fluorescent protein (EGFP) (Fig. 3G). Finally, the mT/mG (Gt(ROSA) 26Sortm4(ACTB-tdTomato,-EGFP)Luo) reporter developed by Muzumdar et al. (2007) expresses red fluorescent protein before Cre recombinase activity and EGFP after, allowing for single-cell reporter resolution that can be evaluated in vivo, or in unfixed samples.

(A) IFT88 Δ/wt; PrxCre +/–

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Mutant

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Fig. 3 Detection of Cre expression with Reporters. (A) Mating schematic wherein the male carries the Cre transgene and a delta allele and the female has two conditional alleles and the reporter gene. Four possible offspring are generated two of which are experimentally relevant. (B) Cre activity in the skin of a Prx1-Cre; R26R mouse present only in dermal cells, as revealed by the lacZ reporter gene activity (blue). In some tissues the b-galactosidase activity is detected in small puncta (as shown here), while in other tissues it is seen throughout the cytoplasm. (C) In wild-type control mice, primary cilia are present in the epidermal (e) and dermal (DC, dermal condensate; DF, dermal fibroblast) compartments of the skin as shown by colocalization of antibodies raised against acetylated a-tubulin (green) and IFT88 (red). (D) In Prx1-Cre; ift88 fl/D conditional mutant mice, primary cilia are ablated from most DC and DF cells, but cilia are unaffected on epidermal (e) cells. Rare dermal cilia are detected, likely due to incomplete Cre activity (arrow). Scale bars are 10 µm. Schematic representation of the Rosa26 (E), Z/EG (F), and Z/AP (G) reporter alleles. Upper alleles are expressed before Cre activity, and the lower alleles are present after Cre activity. Where the boxes represent coding sequences, triangles are LoxP sites, STOP represents stoper sequences unique to the Rosa26 line, AAA represents a polyA tail downstream of the translational stop within b-geo. (See Plate no. 11 in the Color Plate Section.)

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(E)

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Fig. 3 (Continued)

These reporters have distinct advantages that can be exploited for specific types of experiments. For example, the Z/EG and mT/mG reporters have advantages in that tissues do not need to be fixed or stained to assess Cre activity, which is very useful for whole organism and live imaging studies. These reporters are also used for isolating mutant cells by sorting for ones that are GFP positive after Cre deletion using fluorescent-activated cell sorting. In contrast, the Rosa26 and Z/AP lines are commonly used in fixed samples to analyze Cre activity. Importantly, with all of these reporter lines, the cells in which Cre is active become indelibly marked. This allows the progeny of the original cell to be followed over time and have been useful for studies such as those in which mutant cell contribution in renal cysts (Patel et al., 2008) was evaluated, and for cell fate studies during development (Ahn and Joyner, 2004). CreER is especially useful for lineage analysis studies because low doses of tamoxifen can be used to affect only a small number of cells that can then be traced in vivo. While reporter alleles make it easy to detect spatial and temporal patterns of Cre activity, they do not indicate whether or not the gene of interest underwent recombination. Deletion of your target gene can be evaluated by several standard techniques such as Southern and western blot, PCR as well as reverse transcription PCR to evaluate alternative transcripts, and immunofluorescence. One needs to consider the efficiency of allele recombination, the half-life of the targeted protein, as well as the stability of the cilium as it may remain long after the gene has been disrupted. This is seen in the case of ciliogenic gene ift88, where cilia have been detected in vivo for more than 2 days after Cre induction and for more than 5 days in vitro (personal unpublished observations). Cilia resorption correlates with cell division, thus cilia loss occurs faster

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in cells that are rapidly dividing (e.g., during embryonic stages). It has also been estimated that the penetrance of cilia loss is about 80% with the most robust Cre lines (see below and Fig. 3C). These are important factors to account for when assessing the potential phenotypes of mutants.

VI. Practical Notes We provide a practical example for many of the aforementioned points using Fig. 3 (panels B–D adapted from Lehman et al., 2009). An ideal mating between a conditional mouse that also contains a mesenchymal-specific Prx1-Cre transgene (Logan et al., 2002) [Tg(Prrx1-cre)1Cjt known here as Prx1-Cre] and a conditional mouse that is also homozygous for the Rosa26 reporter is shown (Fig. 3A). Note that one of the animals has already lost one of the ift88 alleles (e.g., ift88D/wt), which is done to reduce the number of recombination events that must occur to produce a mutant. Once you ensure that there are no phenotypes associated with homozygous floxed mice, this mating scheme provides littermate controls in animals that have the genotype ift88wt/fl while also expressing Cre. This is a better control than an ift88fl/fl Cre-negative animal as it will control for any off target effects of the Cre recombinase. It is important that you evaluate the possibility that Cre could contribute to the phenotype as this has been shown to occur in some tissues (Lee et al., 2006). This mating scheme will also allow you to evaluate Cre activity in the context of a wild-type background with the Cre reporter lines. Here the Rosa26 reporter is bred onto the conditional line and maintained as homozygous such that ift88fl/fl;Rosa26þ/þ animals can be crossed and all offspring can be used experimentally (e.g., are conditional and Rosa26 positive). It is also of note that in our hands we have detected Cre expression from the Prx1-Cre transgene in female germ cells; thus we maintain the Cre allele only in male mice. This “leaky” germ line expression can cause recombination in the germ cells producing a null allele that can be passed to offspring. Potential “leakiness” should be evaluated for each of Cre line; however, in some instances it may be advantageous. Mentioned above was the use of an ift88D/wt animal that can be generated through a mating with a leaky Cre like the Prx1-Cre. Additionally, the EIIa Cre (Lakso et al., 1996) line is often used for this purpose due to early mosaic deletion. The use of a Rosa26 reporter is shown in panel B where the Cre expression domain was determined using a b-galactosidase assay on a skin section. Prx1-Cre is expressed specifically in the dermal layers of the skin as indicated by blackish staining. Remember that this analysis does not infer cilia loss. Panel D addresses cilia loss via immunofluorescence, the wild-type animal shown in C contains cilia on nearly all cells in the dermis and epidermis, whereas the mutant animal in D shows substantial loss of dermal cilia. Note the examples of dermal cells still presenting cilia, speaking to the incomplete loss of the ift88 allele. Evaluation of the deletion by western blotting in this example is complicated due to the mosaic nature of Cre expression (e.g., in the dermis but not in the epidermis). Furthermore, genotyping of these animals to evaluate allele deletion could produce results that would indicate both the floxed, mutant, and wild-type alleles in the

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same animal for the same reason. This happens when the Cre line used is expressed in a cell type that contributes to the tissue used for genotyping (tail or ear biopsy) or in tissues that cannot be dissected apart (dermis from epidermis). The use of well-designed conditional alleles, thoroughly characterized Cre lines, as well as reporter analyses are all necessary before evaluation of the consequences and roles of cilia mutations.

VII. Conditional Alleles in Cilia Biology The recent generation of floxed alleles of ciliogenic genes, the increasing availability of tissue-specific and inducible Cre deletor lines, along with the characterization of several ciliary proteomes is providing great opportunities to explore ciliary function. Since congenic loss of cilia results in early embryonic lethality, the use of conditional mutation approaches permits analyses at later developmental stages and in postnatal life. This has had a major impact on our understanding of cilia function and contributed greatly to the emergence of the cilium as a critically important organelle both in homeostasis as well as development. There are now several examples where the Cre/loxP approach has been used to define the role of the cilium in diverse tissues and organs to address the molecular and cellular basis of the ciliopathies. For example, polydactyly, a common phenotype among cilia mutant mice, which is also seen in several human ciliopathy disorders (Bardet–Biedel syndrome, Meckel syndrome) (Mykytyn et al., 2004) (OMIM#209900, OMIM#249000), was evaluated with the ORPK (ift88Tg737Rpw) (Moyer et al., 1994) and kif3atm2Gsn mice (Haycraft et al., 2007). Using a series of Cre deletor lines in 2007 Haycraft et al. (2007) demonstrated that polydactyly arises through the loss of cilia specifically on the limb mesenchyme cells. Further, Davenport et al. (2007) used the tamxoifen-inducible approach with a systemically expressed Cre (Hayashi and McMahon, 2002) (CAG Cre: Tg(CAGcre/Esr1)5Amc) to induce cilia loss in adult mice. These animals developed hyperphagia and became obese; a phenotype shared with human Bardet–Biedl and Alström’s syndrome patients. Using additional Cre lines with sequentially more refined spatial expression patterns, the obesity phenotype associated with cilia loss was shown to result from cilia dysfunction specifically on the pro-opiomelanocortin neurons in the hypothalamus (see Kesterson et al., 2009 and Davenport et al., 2007). Similar approaches have been used to explore the importance of cilia in different populations of the mammalian skin, as well as during bone development (Haycraft et al., 2007; Lehman et al., 2009). In summary, conditional alleles are powerful tools in today’s research laboratories as they allow precise and discrete analysis of gene function. Their use in cilia biology will better allow us to delineate the exact role of cilia in multiple cell types and at varying times throughout the lifespan of an animal. With the recent description of several cilia proteomes, we now have an intriguing list of ciliary components that need to be further analyzed. Thus, the genetic resources are available to continue the remarkable progress we have made recently in our understanding of how cilia function to regulate homeostasis, development and signaling activities.

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VIII. Methods A. Tissue Culture Media and Solutions

1. Mouse Embryonic Fibroblast (MEF) Media To Dulbecco’s modified Eagle’s medium (DMEM) add: ES-tested fetal calf serum to 10% 200 mM L-glutamine to 2 mM b-Mercaptoethanol to 0.1 mM 10 mg/ml Pen-Strep to 0.1 mg/ml

2. ES Media To DMEM containing high glucose add: ES-tested FCS to 20% 600,000 U LIF per liter 200 mM L-glutamine to 2 mM 10 mg/ml Pen-Strep to 0.1 mg/ml b-Mercaptoethanol to 0.1mM Optional: 100 mM sodium pyruvate to 1 mM 10 mM nonessential amino acids to 0.1 mM

3. Blastocyst Injection Medium (BIM) To DMEM add: ES-tested fetal calf serum to 10% 1 M HEPES buffer to 0.02 M

4. 2 ES Freezing Medium Supplement ES medium with DMSO to 20%

5. Trypsin–EDTA 0.25% trypsin is used to disassociate ES cell clusters while 0.05% is generally used for MEF maintenance.

6. PBS–0.1% gelatin Add gelatin to PBS to 0.1% and sterilize by autoclaving (stored at room temperature).

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7. Southern Lysis Buffer 10 mM Tris pH 7.5 10 mM EDTA pH 8.0 10 mM NaCl 0.5% sarcosyl 1 mg/ml fresh proteinase K

B. Isolation of Targeting Vector DNA 1. Perform standard Midiprep or Maxiprep on bacterial cultures. 2. Digest 25–40 µg of targeting vector DNA with an appropriate restriction enzyme to linearize the construct. 3. Perform phenol–chloroform extraction to remove the enzyme: Add equal volume of phenol–chloroform to the digest, vortex, spin in benchtop centrifuge on max >5 min. Save upper layer containing the DNA. 4. Precipitate linearized DNA with NaCl and ethanol. Add 1/20th volume of 5 M NaCl to the supernatant then add 2.5 volumes of EtOH. Incubate on ice for 10 min prior to centrifugation at room temperature in benchtop centrifuge on maximum speed. Discard supernatant and wash pellet once with 70% ethanol. Aspirate off the EtOH and resuspend the DNA pellet at 1 µg/µl in sterile water. Store the targeting vector at –20°C.

C. Preparation of Mouse Embryonic Fibroblasts Transgenic mouse strains that carry drug-resistance genes are commercially available for the production of drug-resistant MEF cells to withstand the selection applied to the ES cells. MEF cells are prepared from E13.5 mouse embryos and are always seeded onto gelatinized culture surfaces. Gelatin-coated dishes are prepared as follows: Flood the plate with the 0.1% gelatin (~1 ml in a 10 cm plate), swirl the plate to insure complete coverage of the bottom surface, aspirate off the solution and allow the plates to dry in the hood. Many plates can be gelatinized and stored for later use. 1. Set up timed pregnancies with drug-resistant mice. 2. Sacrifice pregnant female mice at 13.5 days postcoitum dissect out the mouse embryos, decapitate, eviscerate, and rinse in PBS. 3. Disassociate carcasses by passing embryos through an 18G needle a. Some researchers utilize trypsin–EDTA to help disassociate the cells: transfer embryos into 10 ml of trypsin–EDTA in a 50-ml tube. Incubate the tubes in a 37°C water bath for 10 min vigorously shaking the tubes at 2-min intervals. 4. Add 10 ml of MEF medium and pellet gently (e.g., 1000 g for 1.5 min). 5. Resuspend pellet in 1 ml MEF medium per embryo, plate in one 10-cm plate per embryo (e.g., for seven embryos, resuspend in 7 ml medium and distribute 1 ml to

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seven plates containing 9 ml of MEF medium per plate). Incubate at 37°C and leave the dishes undisturbed overnight. The next day, change the medium to remove dead cells and debris. When plates become highly confluent (2–4 days after initial seeding), expand each 10-cm plate to a 15-cm plate. When these plates are highly confluent expand the cultures again going from one 15-cm plate to two 15-cm plates, continue to expand in this manor until many plates are confluent (~30 separate 15-cm plates). When the cultures again are highly confluent cells must be treated to stop proliferation. Two techniques are commonly used mitomycin C treatment or g-irradiation. a. Mitomycin C treatment: treat cells with 10 µg/ml of mitomycin C supplemented into regular MEF medium for 2–3 h at 37°C. Wash at least twice with PBS (be sure to remove all of the drug!) lift the cells, pellet, and gently resuspend cells. b. g-irradiation treatment: lift cells, pool, expose to 3000–10,000 rads of g-irradiation, pellet gently, resuspend cells. Dilute cells by mixing MEF medium 1:2 with the 2 freezing medium the final volume of cells will vary but a good starting point is 5 ml per 15-cm plate. Count cells and aliquot ~3.5 106 each into cryogenic vials, place each vial in an isopropanol-jacketed cryocontainer and store at –80°C. The next day, transfer vials in liquid nitrogen for long-term storage. Thaw a vial of cells to determine the surface area that the feeder cells will cover, this will vary with each batch of MEFs. Use the following guidelines: Plate

MEF cells

10 cm 6 cm 12 well 24 well

3.5 106 1 106 5 105 2 105

D. ES Cell Growth and Electroporation Because the formation of large ES cell clusters may trigger differentiation (Fig. 4C), it is advisable to passage the ES cells when 70–80% confluent but no less than every 3 days. Split the cells 1:2 for rapid culture expansion, be careful not to seed too sparsely or the cells may quiesce. 1. Two days prior to the scheduled seeding of ES cells, thaw the appropriate number of vials of MEFs to seed a six-well plate (one well per vial of ES cells) and five 10-cm dishes with feeder monolayers. 2. Thaw a vial of ES cells at 37°C, transfer to a 15-ml conical flask, and add 5 ml ES medium. Pellet the ES cells gently.

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(B)

(A)

100 µm

(C)

50 µm

50 µm

Fig. 4 Morphology of ES cells at different stages of the procedure. (A–C) ES cells on feeder cells. ES cells form oval, shiny colonies with clear boundaries (A, B). Good, pluripotent ES cell cultures reach the same density and morphology again after 2 days. (C) Spontaneous differentiation of ES cell colonies on feeders. The round and shiny shape disappeared and distinct cells can be seen. A good ES cell culture should not show such colonies. In addition, most other ES cell colonies in this picture do not show clear boundaries, are of different sizes, some are flat and not shiny. Used with permission from “Generation of a defined and uniform population of CNS progenitors and neurons from mouse embryonic stem cells” by Bibel et al. (2007).

3. Gently resuspend the thawed ES cells in ES medium, and transfer to one well of a six-well plate with feeders. Incubate at 37°C. 4. When the ES cells are 70–80% confluent, expand them to a 10-cm plate with feeders. 5. The next day (roughly), the ES cells (20 million) will cover 70–80% of the surface area and are ready for electroporation. Change the media 2–3 h before starting the protocol. 6. Lift cells with trypsin and triturate to generate a single-cell suspension. Add ES medium, transfer the suspension to a 15-ml tube, and pellet the ES cells gently. 7. Wash cells twice in PBS by gently resuspending and pelleting again. 8. After the last wash, resuspend the cells in 800 µl of PBS and add 25 µl of the stored DNA solution (1 µg/µl linearized targeting vector in water). Transfer the suspension to an electroporation cuvette with a 0.4-cm gap size. Remove potential air bubbles. 9. For the Bio-Rad Gene Pulser system (Bio-Rad Hercules, CA, USA): Place the cuvette in the Bio-Rad Gene Pulser and electroporate at 230 V, 500 µF. The time constant should be between 6 and 8 ms (all conditions will need to be optimized for each apparatus). Let the cells rest in the cuvette for 5 min. 10. Transfer cells to 40 ml ES media and split into four 10-cm plates of feeders. 11. 24–36 h after the electroporation, replace the ES medium on the plates with drugcontaining ES medium. Use the following final drug concentrations as guidelines, it is recommended that a kill curve be performed to determine the lowest dose necessary to kill wild-type cells: 350 µg/ml G418, 140 µg/ml hygromycin B, 2.75 µg/ml puromycin, 0.2 µM FIAU, 2 µM ganciclovir, use based on the selection of your vector/construct. 12. Refresh the selective medium daily. Generally the ES colonies will be ready for picking 8–10 days after electroporation but that can vary with the ES line being used. Cells without the allele will start

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dying at about day 3. By day 6 the whole plate will look dead and everything will be floating. Between days 7 and 10 colonies will come up and be ready to pick. Individual colonies should be clearly visible through the bottom of the plates and have a uniform appearance with sharp edges (Fig. 4A and B). The center of an ES colony has usually a higher cell density and is somewhat darker in color. ES colonies with a flat pancake-like appearance should not be picked due to likely differentiation (Fig. 4C).

E. Picking Drug-Resistant ES Clones and Creating Replica Plates 1. Two days prior to the scheduled picking of ES cells, thaw the appropriate number of vials of MEFs to seed four 96-well flat-bottomed plates to pick 384 colonies, adjust as necessary. 2. Pick colonies one plate at a time. Screen the colonies in the 10-cm plates before picking; it is helpful to mark the best looking colonies on the bottom of each plate. Cross out differentiated colonies, or ones that may have come from two cells, so they will not be picked. Use of more expensive “gridded” 15-cm dishes can aid in colony localization and increase the speed of the process; alternatively, divide the plate into quadrants with a marker. 3. Place 30 µl of room temperature trypsin (0.25%) in each well of a 96-well roundbottomed plate (without feeders). 4. Remove the ES media from the cells, and replace with serum-free DMEM, set a Pipetman to 5–10 µl use the tip of the pipette to detach the colonies and transfer each colony to a well of the round-bottomed 96-well plate with the trypsin solution. Triturate several times to make sure the colony comes out of the tip. Use a new tip for each colony. 5. After picking a plate of colonies, put the plate at 37°C for 5 min. 6. Using a multichannel pipettor, add 150 µl of ES media to inhibit trypsin (180 µl total volume) and triturate each well to disassociate the cells. Transfer 60 µl from each well to the corresponding wells of two 96-well plates to be grown for DNA isolation. Transfer the remaining 60 µl from each well to the corresponding well of a 96-well plate with MEFs to be grown and frozen. Be sure to change tips and keep track of the wells during this process. Add 200 µl of ES media to all wells of these duplicate “DNA plates” and “master” plate for freezing; allow the cells to grow until confluent. 7. Remove the plates to be frozen from the incubator and remove the medium with a multichannel pipette (use clean tips for each row). Wash ES cells by pipetting 100 µl PBS down and up in the wells using a multichannel pipette. After washing all wells, add 30 µl trypsin–EDTA to each of the wells and incubate the plate for 5 min at 37°C. 8. Take 70 µl ES medium and triturate the trypsinized ES cells using a multichannel pipette. Then, mix the contents of each well with 100 µl ice-cold 2 freezing medium. (Pipette vigorously up and down at least 5 times to dilute the 2 freezing medium.)

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Quickly seal the plate with parafilm, wrap it in several layers of paper towel, and transfer it to a styrofoam box with a lid. Place the box in a –80°C freezer. 9. Proceed to characterizing the targeting reaction with the other duplicate “DNA plates” of cells. F. Southern Analysis (adapted from Ramirez-Solis et al., 1992) Be sure to test this protocol with wild-type ES cells before trying to characterize targeted clones. It is necessary to ensure your probes work and that the DNA digests as expected. 1. When the cells are confluent, wash the plates twice with PBS and aspirate off. Using the multichannel pipette, add 50 µl of lysis buffer into each well. 2. Incubate the plates overnight at 60°C in a humidified chamber (such as a plastic container with a wet sponge inside). 3. The next day, prepare a fresh solution of 75 mM NaCl in ethanol (add 150 µl of 5 M NaCl per 10 ml of cold absolute ethanol and mix well; it is okay if some salt precipitates). 4. Using the multichannel pipette, add 100 µl of the NaCl/ethanol solution per well. Allow the plate to rest on the bench at room temperature for 15–30 min, or until the precipitated DNA is clearly visible under low-power magnification. The DNA adheres to the plastic, so look at the perimeter of each well to see the precipitated “cobweb” DNA. 5. Invert the plate to discard the solution (the DNA will remain adhered to the plate), remove as much of the solution as possible by inverting onto paper towels. Using the multichannel pipette, gently add 150 µl of 70% cold ethanol to sidewalls to wash each well. Invert the plate to discard the 70% ethanol, drain, and repeat the wash two to three times. If available, the use of a tabletop centrifuge equipped with microtiter plate buckets can be used. 6. After the final wash, invert the plate to discard the 70% ethanol, and allow the plate to air-dry for ~20 min but do not over dry. 7. While the plate is drying, prepare the restriction enzyme cocktail (1 restriction buffer specified for the enzyme being used, 1 mM spermidine, 100 µg/ml bovine serum albumin, and 10–15 units of enzyme). 8. Using the multichannel pipette, add 30 µl of restriction enzyme cocktail to each well and mix by pipetting up and down. Change tips between rows. 9. Once the cocktail has been added to all the wells, incubate the plates overnight at 37°C in a sealed humidified chamber. 10. The next day, prepare agarose gel(s) for electrophoresis. 11. Remove the 96-well plate from the incubator and add 4–5 µl of loading buffer to each digest. Load the gel and run at 80 V for 4 h depending on the fragment(s) being distinguished. 12. After the electrophoresis is complete, image gel with fluorescent ruler and then transfer the DNA to a membrane (i.e., Zeta-probe by Bio-Rad) using the manufacturer’s protocol. Label the bottom of blot using pencil (samples, date, etc.) prior to screening with labeled external probes.

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G. Expansion and Verification of Targeted ES clones Correctly targeted ES clones must be thawed and expanded for: (1) Southern blot confirmation of the targeted mutation (again after thawing), (2) karyotyping, and (3) preparation of multiple frozen stocks. 1. Two days prior to the scheduled expansion of the targeted ES cell clones, thaw enough MEFs to prepare multiple 24- and 6-well plates with feeder monolayers as below. 2. Remove one 96-well plate with targeted ES clones from the –80°C freezer and thaw at 37°C 3. Careful mark “positive” wells to be thawed and label corresponding well of 24-well dish with each clone ID accordingly. 4. Once thawed, transfer 96-well “positive” clones containing targeted ES cells to be expanded to one well of a 24-well dish with MEFs. Resuspend the 200 µl of cells by vigorously pipetting up and down several times, and transfer the cell suspension to the 24-well dish containing 1 ml of ES medium. Incubate at 37°C. 5. The next day, change the medium on the 24-well plate to remove any traces of freezing medium or debris. 6. Monitor the growth of the ES cells closely and split clones that have reached 70– 80% confluency, expand and transfer to a six-well plate, and finally to a 10-cm plate. 7. When ES clones are 70–80% confluent lift the cells and pellet gently. 8. Aspirate the medium from the tube and suspend the cell pellet in 6 ml ES medium. Pipette 0.5 ml of cells into several cryogenic vials and add 0.5 ml 2 Freezing medium to each cell suspension, mix well, and transfer to an isopropanol-jacketed cryocontainer and store at –80°C. Freezing can also be done at earlier stages during expansion (e.g., six-well dishes). 9. Divide the remaining 0.5 ml cells over two wells of a 24-well plate with MEF cells. Grow to superconfluency, use one well for DNA extraction and confirmation of targeted mutagenesis via Southern blot, and the other for karyotyping see Nagy for details (2003).

H. Preparing ES Cells for Microinjection Once characterized for correct targeting thaw a vial of ES cells, serially dilute, and seed into three wells of a 24-well dish. Cells can be further expanded and grown to confluence on 12- or 6-well plates for injecting. 1. Make BIM Blastocyst Injection Medium. 2. Remove media from plates, wash two times with PBS. 3. Trypsinize at 37°C for 5 min. Add ES media and transfer to a nongelatinized 6-cm dish and triturate thoroughly. 4. Incubate for 30 min for differential attachment. Fibroblasts should lay down on plate.

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5. Aspirate media carefully and rinse ES cells off the plate with 200–500 µl of BIM (depending on the number of cells) 6. Collect cells and place on ice. Use them for microinjection within 1–2 h following trypsinization. Typically, 10–15 cells are injected per blast and 10–12 blasts/ embryos are injected per targeted clone. Optimally, three to four independently targeted ES clones will be injected.

Acknowledgments We would like to thank N. F. Berbari and C. A. Clement, for critical review of the manuscript and helpful discussions and T. M. Kesterson for technical assistance. Figure 3A was crafted by C. A. Clement. This work was supported by grant RO1 HD056030 (Dr B. Yoder) to B.K.Y., P30 DK074038 (Dr L. Guay-Woodford) to R.A.K., and by NIH T32 training grant AR047512-07 (Dr J.M. McDonald) to A.K.O.

References Ahn, S. and Joyner, A.L. (2004). Dynamic changes in the response of cells to positive hedgehog signaling during mouse limb patterning. Cell 118(4), 505–516. Avidor-Reiss, T., Maer, A.M., Koundakjian, E., Polyanovsky, A., Keil, T., Subramaniam, S. and Zuker, C.S. (2004). Decoding cilia function: defining specialized genes required for compartmentalized cilia biogenesis. Cell 117(4), 527–539. Bibel, M., Richter, J., Lacroix, E. and Barde, Y.A. (2007). Generation of a defined and uniform population of CNS progenitors and neurons from mouse embryonic stem cells. Nat. Protoc. 2(5), 1034–1043. Blacque, O.E., Perens, E.A., Boroevich, K.A., Inglis, P.N., Li, C., Warner, A., Khattra, J., Holt, R.A., Ou, G., Mah, A.K., McKay, S.J., Huang, P., et al. (2005). Functional genomics of the cilium, a sensory organelle. Curr. Biol. 15(10), 935–941. Bouvier, J. and Cheng, J.G. (2009). Recombineering-based procedure for creating Cre/loxP conditional knockouts in the mouse. Curr. Protoc. Mol. Biol. Chapter 23: Unit 23 13. Buchholz, F., Angrand, P.O., and Stewart, A. F. (1998). Improved properties of FLP recombinase evolved by cycling mutagenesis. Nat. Biotechnol. 16(7), 657–662. Davenport, J.R., Watts, A.J., Roper, V.C., Croyle, M.J., van Groen, T., Wyss, J.M., Nagy, T.R., Kesterson, R.A. and Yoder, B .K. (2007). Disruption of intraflagellar transport in adult mice leads to obesity and slow-onset cystic kidney disease. Curr. Biol. 17(18), 1586–1594. Efimenko, E., Bubb, K., Mak, H.Y., Holzman, T., Leroux, M.R., Ruvkun, G., Thomas, J.H. and Swoboda, P., (2005). Analysis of xbx genes in C. elegans. Development 132(8), 1923–1934. Feil, R. (2007). Conditional somatic mutagenesis in the mouse using site-specific recombinases. Handb. Exp. Pharmacol. 178, 3–28. Feil, R., Brocard, J., Mascrez, B., LeMeru, M., Metzger, D. and Chambon, P. (1996). Ligand-activated sitespecific recombination in mice. Proc. Natl. Acad. Sci. USA 93(20), 10887–10890. Friedrich, G. and Soriano, P. (1991). Promoter traps in embryonic stem cells: A genetic screen to identify and mutate developmental genes in mice. Genes Dev. 5(9), 1513–1523. Hayashi, S. and McMahon, A.P. (2002). Efficient recombination in diverse tissues by a tamoxifen-inducible form of Cre: A tool for temporally regulated gene activation/inactivation in the mouse. Dev. Biol. 244(2), 305–318. Haycraft, C.J., Zhang, Q., Song, B., Jackson, W.S., Detloff, P.J., Serra, R. and Yoder, B.K. (2007). Intraflagellar transport is essential for endochondral bone formation. Development 134(2), 307–316. Hoess, R., Abremski, K., Irwin, S., Kendall, M. and Mack, A. (1990). DNA specificity of the Cre recombinase resides in the 25 kDa carboxyl domain of the protein. J. Mol. Biol. 216(4), 873–882.

15. Generating Conditional Mutants to Analyze Ciliary Functions

329

Indra, A.K., Warot, X., Brocard, J., Bornet, J.M., Xiao, J.H., Chambon, P. and Metzger, D. (1999). Temporally-controlled site-specific mutagenesis in the basal layer of the epidermis: Comparison of the recombinase activity of the tamoxifen-inducible Cre-ER(T) and Cre-ER(T2) recombinases. Nucleic Acids Res. 27(22), 4324–4327. Jonassen, J.A., San Agustin, J., Follit, J.A. and Pazour, G.J. (2008). Deletion of IFT20 in the mouse kidney causes misorientation of the mitotic spindle and cystic kidney disease. J. Cell Biol. 183(3), 377–384. Kesterson, R.A., Berbari, N.F., Pasek, R.C. and Yoder, B.K. (2009). Utilization of conditional allels to study the role of the primary cilium in obesity. Methods in Cell Biology. Lakso, M., Pichel, J.G., Gorman, J.R., Sauer, B., Okamoto, Y., Lee, E., Alt, F.W., Westphal, H. (1996). Efficient in vivo manipulation of mouse genomic sequences at the zygote stage. Proc. Natl. Acad. Sci. USA 93(12), 5860–5865. Lee, J.Y., Ristow, M., Lin, X., White, M.F., Magnuson, M.A. and Hennighausen, L. (2006). RIP-Cre revisited, evidence for impairments of pancreatic beta-cell function. J. Biol. Chem. 281(5), 2649–2653. Lehman, J.M., Laag, E., Michaud, J. and Yoder, B.K. (2009). An essential role for dermal primary cilia in hair follicle morphogenesis. J. Invest. Dermatol. 129(2), 438–448. LePage, D.F. and Conlon, R.A. (2006). Animal models for disease: Knockout, knock-in, and conditional mutant mice. Methods Mol. Med. 129, 41–67. Li, J.B., Gerdes, J.M., Haycraft, C.J., Fan, Y., Teslovich, T.M., May-Simera, H., Li, H., Blacque, O. E., Li, L., Leitch, C.C., Lewis, R.A., Green, J.S., et al. (2004). Comparative genomics identifies a flagellar and basal body proteome that includes the BBS5 human disease gene. Cell 117(4), 541–552. Liu, Q., Tan, G., Levenkova, N., Li, T., Pugh, E.N., Jr., Rux, J.J., Speicher, D.W. and Peirce, E.A. (2007). The proteome of the mouse photoreceptor sensory cilium complex. Mol. Cell Proteomics 6(8), 1299–1317. Lobe, C.G., Koop, K.E., Kreppner, W., Lomeli, H., Gertsenstein, M. and Nagy, A. (1999). Z/AP, a double reporter for cre-mediated recombination. Dev. Biol. 208(2), 281–292. Logan, M., Martin, J.F, Nagy, A., Lobe, C., Olson, E.N and Tabin, C.J. (2002). Expression of Cre Recombinase in the developing mouse limb bud driven by a Prxl enhancer. Genesis 33(2), 77–80. Mao, X., Fujiwara, Y., and Orkin, S.H. (1999). Improved reporter strain for monitoring Cre recombinasemediated DNA excisions in mice. Proc. Natl. Acad. Sci. USA 96(9), 5037–5042. Marszalek, J.R., Liu, X., Roberts, E.A., Chui, D., Marth, J.D., Williams, D.S. and Goldstein, L.S. (2000). Genetic evidence for selective transport of opsin and arrestin by kinesin-II in mammalian photoreceptors. Cell 102(2), 175–187. Moyer, J.H., Lee-Tischler, M.J., Kwon, H.Y., Schrick, J.J., Avner, E.D., Sweeney, W.E., Godfrey, V.L., Cacheiro, N.L., Wilkinson, J.E. and Woychik, R.P. (1994). Candidate gene associated with a mutation causing recessive polycystic kidney disease in mice. Science 264(5163), 1329–1333. Muzumdar, M.D., Tasic, B., Miyamichi, K., Li, L. and Luo, L. (2007). A global double-fluorescent Cre reporter mouse. Genesis 45(9), 593–605. Mykytyn, K., Mullins, R.F., Andrews, M., Chiang, A.P., Swiderski, R.E., Yang, B., Braun, T., Casavant, T., Stone, E.M. and Sheffield, V.C. (2004). Bardet-Biedl syndrome type 4 (BBS4)-null mice implicate Bbs4 in flagella formation but not global cilia assembly. Proc. Natl. Acad. Sci. USA 101(23), 8664–8669. Nagy, A. (2003). “Manipulating the Mouse Embryo: A Laboratory Manual.” 3rd edn., 764 pp. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Novak, A., Guo, C., Yang, W., Nagy, A. and Lobe, C.G. (2000). Z/EG, a double reporter mouse line that expresses enhanced green fluorescent protein upon Cre-mediated excision. Genesis 28(3–4), 147–155. Ostrowski, L.E., Blackburn, K., Radde, K.M., Moyer, M.B., Schlatzer, D.M., Moseley, A. and Boucher, R.C. (2002). A proteomic analysis of human cilia: Identification of novel components. Mol. Cell Proteomics 1(6), 451–465. Patel, V., Li, L., Cobo-Stark, P., Shao, X., Somlo, S., Lin, F. and Igarashi, P. (2008). Acute kidney injury and aberrant planar cell polarity induce cyst formation in mice lacking renal cilia. Hum. Mol. Genet. 17(11), 1578–1590. Pazour, G.J., Dickert, B.L., Vucica, Y., Seeley, E.S., Rosenbaum, J.L., Witman, G.B. and Cole, D.G. (2000). Chlamydomonas IFT88 and its mouse homologue, polycystic kidney disease gene Tg737, are required for assembly of cilia and flagella. J. Cell Biol. 151(3), 709–718.

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Amber K. O’Connor et al. Ramirez-Solis, R., Rivera-Perez, J., Wallace, J.D., Wims, M., Zheng, H. and Bradley, A. (1992). Genomic DNA microextraction: A method to screen numerous samples. Anal. Biochem. 201(2), 331–335. Sharan, S.K., Thomason, L.C., Kuznetsov, S.G., Court, D.L. (2009). Recombineering: A homologous recombination-based method of genetic engineering. Nat. Protoc. 4(2), 206–223. Smith, J.C., Northey, J.G., Garg, J., Pearlman, R.E. and Siu, K.W. (2005). Robust method for proteome analysis by MS/MS using an entire translated genome: Demonstration on the ciliome of Tetrahymena thermophila. J. Proteome Res. 4(3), 909–919. Soriano, P. (1999). Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat. Genet. 21(1), 70–71. Stolc, V., Samanta, M. P., Tongprasit, W. and Marshall, W.F. (2005). Genome-wide transcriptional analysis of flagellar regeneration in Chlamydomonas reinhardtii identifies orthologs of ciliary disease genes. Proc. Natl. Acad. Sci. USA 102(10), 3703–3707. Taulman, P.D., Haycraft, C.J., Balkovetz, D.F. and Yoder, B.K. (2001). Polaris, a protein involved in leftright axis patterning, localizes to basal bodies and cilia. Mol. Biol. Cell 12(3), 589–599. te Riele, H., Maandag, E.R., and Berns, A. (1992). Highly efficient gene targeting in embryonic stem cells through homologous recombination with isogenic DNA constructs. Proc. Natl. Acad. Sci. USA 89(11), 5128–5132.