INTERNATIONAL REVIEW OF CYTOLOGY V130, Volume 130

VOLUME 130

SERIES EDITORS Geoffrey H. Bourne James F. Danielli Kwang W. Jeon Martin Friedlander

1949-1 988 1949-1 984 19671984-

ADVl SORY EDITORS Howard A. Bern Dean Bok Stanley Cohen Rene Couteaux Marie A. DiBerardino Donald K. Dougall Charles J. Flickinger Nicholas Gillham M. Nelly Golarz De Bourne Elizabeth D. Hay Mark Hogarth H. R. Kaback Keith E. Mostov Audrey Muggleton-Harris

Andreas Oksche Muriel J. Ord Valdimir R. Pantic M. V. Parthasarathy Lionel I. Rebhun Jean-Paul Revel L. Evans Roth Jozef St. Schell Hiroh Shibaoka Wilfred Stein Ralph M. Steinrnan D. L. Taylor M. Tazawa Alexander L. Yudin

Edited by Kwang W. Jeon Department of Zoology The University of Tennessee Knoxville, Tennessee

Martin Friedlander

Jules Stein Eye Institute UCLA School of Medicine Los Angeles, California

VOLUME 130

W Academic Press, Inc. Harcourt Brace Jovanovich, Publishers San Diego New York Boston London

Sydney Tokyo Toronto

This book is printed on acid-free paper. @ Copyright 0 1991 BY ACADEMIC PRESS, INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.

Academic Press, Inc. San Diego, California 92101 United Kingdom Edition published by ACADEMIC PRESS LIMITED 24-28 Oval Road, London NW1 7DX

Library of Congress Catalog Card Number: 52-5203 ISBN 0-12-364530-1

(alk. paper)

PRINTED IN THE UNITED STATES OFAMERICA 91929394 9 8 7 6 5 4 3 2 1

CONTENTS

Contributors .............................................................................................................

ix

Immunoglobulin Transport in B Cell Development Shiv Pillai Introduction .................................................................................................... Overview of B Lymphocyte Ontogeny .......................................................... lntracellular Retention of Secretory Immunoglobulins .............................. Membrane Immunoglobulin Transport during B Cell Ontogeny ............... Immunoglobulin Secretion in Plasma Cells ................................................

I. II. 111. IV. V. VI. Summary: Choices between RetentionlDegradation and Transport of Immunoglobulins Are Dictated by Function ................................................ References ......................................................................................................

1 2 13 20 29

31 34

The Cytoskeletal System of Nucleated Erythrocytes William D. Cohen Introduction .................................................................................................... Nucleated Erythrocytes: A Phylogenetic and Physiological Portrait ........ The Marginal Band of Mature Erythrocytes ................................................. Marginal Band Biogenesis and Function during Erythrocyte Morphogenesis ............................................................................................... V. The Membrane Skeleton ...............................................................................

1. II. 111. IV.

37 39 43 59 67 V

Vi

CONTENTS

VI . Intermediate Filaments .................................................................................. VII. The Cytoskeletal System of Mammalian Primitive Erythrocytes ................ VIII. Concluding Remarks ..................................................................................... References ......................................................................................................

Structure of the Mouse Egg Extracellular Coat. the Zona Pellucida Paul M. Wassarman and Steven Mortillo I. Introduction .................................................................................................... II. 111. IV . V.

Functions of the Zona Pellucida ................................................................... Characteristics of the Zona Pellucida .......................................................... Ultrastructure of the Zona Pellucida ............................................................ Concluding Remarks ..................................................................................... References ......................................................................................................

75 76 79 80

85 86 89 96 105 108

The Male Germ Cell Protective Barrier along Phylogenesis Mordechai Abraham I. Introduction .................................................................................................... II . Background .................................................................................................... 111 . IV. V. VI .

The Male Germ Cell Protective Barrier along Phylogenesis ...................... Structure of the Male Germ Cell Protective Barrier .................................... Function of the Male Germ Cell Protective Barrier ..................................... Concluding Remarks ..................................................................................... References ......................................................................................................

111 112 119 143 159 173 177

Mitochondria-Rich Cells in the Gill Epithelium of Teleost Fishes: An Ultrastructural Approach M. Pisam and A . Rambourg I. II. 111. IV.

Introduction .................................................................................................... Gill Morphology ............................................................................................. General Ultrastructural Features of Chloride Cells ..................................... Mitochondria-Rich Cells and Modifications of the Environment ..............

191 192 193 204

CONTENTS

V . Mitochondria-Rich Cells and Smoltification in Salmonids ........................ VI . Hormones and Chloride Cells ....................................................................... VII . Concluding Remarks ..................................................................................... References ......................................................................................................

Vii

221 225 227 228

Structure and Function of Plant Cell Walls: Immunological Approaches Takayuki Hoson I. II. Ill. IV . V. VI . VII.

Introduction .................................................................................................... Antibodies as Probes for the Study of Plant Cell Walls .............................. Location and Metabolism of Cell Wall Polymers ........................................ Growth Regulation ......................................................................................... Selective Breakdown of Plant: Cell Walls ..................................................... Other Aspects of Plant Cell Walls ................................................................. Conclusions and Future Prospects .............................................................. References ......................................................................................................

233 234 240 246 254 259 261 263

Biologically Localized Firefly Luciferase: A Tool t o Study Cellular Processes Claude Aflalo Introduction .................................................................................................... Cellular Biogenesis and Organization ......................................................... Firefly Luciferase: An Overview .................................................................... A Local Probe for ATP in Model Systems .................................................... Leading Luciferase into Cellular Compartments ........................................ Light Emission by Luciferase in Biological Systems .................................. Experimental Approaches and Perspectives ............................................... References ......................................................................................................

269 270 280 287 296 300 309 318

Index .........................................................................................................................

325

I. II. 111. IV. V. VI . VII .

This Page Intentionally Left Blank

CONTRIBUTORS

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

Mordechai Abraham (1 1 l ) , Department of Zoology, The Hebrew University of Jerusalem, Jerusalem 91904, Israel Claude Aflalo (269),Department of Biochemistry, The Weizmann Institute of Science, Rehovot 76100, Israel William D. Cohen (37),Department of Biological Sciences, Hunter College of CUNY, New York, New York 10021; and The Marine Biological Laboratory, Woods Hole, Massachusetts 02543 Takayu ki Hoson (233),Department of Biology, Faculty of Science, Osaka City University, Osaka 558, Japan Steven Mortillo (85),Department of Cell and Developmental Biology, Roche lnstitute of Molecular Biology, Roche Research Center, Nutley, New Jersey 07110 Shiv Pillai (l), Massachusetts General Hospital and Harvard Medical School, Boston, Massachusetts 02129 M. Pisam (191),Service de Biologie Cellulaire, Departement de Biologie Cellulaire et Moleculaire, Centre d ’ h d e s Nucleaires de Saclay, 91 191 Gif-sur-Yvette Cedex, France

ix

X

CONTRIBUTORS

A. Rambourg (191), Service de Biologie Cellulaire, Departement de Biologie Cellulaire et Moleculaire, Centre d’Etudes Nucleaires de Saclay, 91 191 Gif-sur-Yvette Cedex, France

Paul M. Wassarman (85), Department ofCell andtDevelopmentalBiology, Roche institute of Molecular Biology, Roche Research Center, Nutley, New Jersey 071 10

INTERNATIONAL REVIEW OF CYTOLOGY. VOL. 130

Immunoglobulin Transport in B Cell Development SHIVPILLAI Massachusetts General Hospital and Harvard Medical School, Boston, Massachusetts 02129

I. Introduction The B lymphocyte differentiation pathway is one of the best-studied models of eukaryotic differentiation. While B cells are of obvious interest to students of immunology, the study of these cells has also yielded numerous insights into cellular processes that are common to many other cell types in higher eukaryotes. Examples of such processes include tissue-specific transcriptional regulation, alternative splicing to yield mRNAs for membrane and secretory proteins, and differential polyadenylation of genes during development. There are a number of extremely interesting cellular and molecular processes unique to immune cells, which make the study of B lymphocytes particularly exciting. In pre-B lymphocytes, unusual molecular events include the rearrangement of immunoglobulin genes and the processes of allelic and isotypic exclusion [which ensure that a given B cell expresses only one rearranged heavy- and one light-chain (H- and L-chain, respectively) allele]. In early B cells, exposure to antigen may lead to clonal deletion, one of the mechanisms by which self-non-self discrimination is achieved. The exposure of slightly more differentiated B cells to antigen leads to the activation of a number of molecular events, including two that are unique to the B lineage: A rearrangement process at the H-chain locus leads to isotype switching, and the process of somatic mutation of rearranged immunoglobulin genes further diversifies the immune repertoire. The transport of immunoglobulin molecules is exquisitely regulated during B lymphocyte differentiation and closely parallels the vastly different functions subserved by membrane and secretory immunoglobulins during B cell ontogeny. The study of the processes by which immunoglobulin transport is regulated, apart from yielding useful insights into protein trafficking in general, also provides a paradigm for the regulation of cellular differentiation at the level of protein transport. In this chapter I begin by providing an overview of the process of B cell differentiation, highlighting features of the pathway which are particularly relevant from the viewpoint of immunoglobulin transport. I then describe the regulation of trans1

Copyright 0 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

2

SHIV PILLAI

port during the process of differentiation in some detail, discussing at each step mechanisms of broad relevance to protein trafficking and cell biology in general. 11. Overview of B Lymphocyte Ontogeny

The goal of B lymphocyte differentiation is to generate a wide range of antigen-responsive cells which will respond to antigen by secreting specific antibodies. Each B cell clone must express a single cell surface antigen receptor in order to maintain clonal specificity; self-reactive B cell clones must be either eliminated or rendered anergic. Immunoglobulin H-chain proteins are synthesized in two forms, the membrane (m) and secretory (s) forms, which are products of alternative splicing at the H-chain locus (reviewed by Wall and Kuehl, 1983). A schematic view of the p H-chain locus is provided in Fig. 1. Many hundred variable region (V) immunoglobulin genes are separated by a large distance from a small group of diversity (D) genes. Downstream of the D cluster is a small set of joining (J) genes separated by the J-C intron from the first exon of the constant region (C) of the p H-chain gene. In the case of the p chain (the other isotypes downstream of p have broadly similar structural features), the C region is made up of four common exons, respectively encoding the c p l to cp4 domains. Two membrane exons, pMl and pM2, encode a total of 42 amino acids, which include 13 amino acids that form an extracellular negatively charged region, 26 amino acids that form a relatively hydrophilic membrane-spanning domain (nine of the 26 amino acids are serines and threonines), and three amino acids that constitute a cytoplasmic tail. Two polyadenylation sites, one immediately downstream of the cp4 exon and the other downstream of the pM2 exon, permit the generation of two mRNAs from the p gene (similar polyadenylation and splice sites characterize every H-chain isotype). A splice donor site in the coding region of the cp4 domain leads to the generation of an alternatively spliced version of the p gene which includes the two membrane exons and encodes the membrane form of the immunoglobulin H chain. The secretory form of the p H chain (ps) includes a tailpiece distal to the splice site in the cp4 exon, but excludes the relatively hydrophobic pMl and pM2 exons. An extremely simplified view of B cell differentiation is presented in Fig. 2. In this scheme the ontogenic process is considered to be made up of three stages, each having a well-defined and -demarcated function; the immunoglobulin molecule at each of these broadly defined stages subserves a distinct function and is transported to a specific subcellular loca-

3

IMMUNOGLOBULIN TRANSPORT

V Genes 100-1000

D Cluster 5 10 genes

-

-

J C lntron

Constant Region

p m Messenger RNA

s Messenger RNA

FIG.1. A schematic view of the p heavy-chain locus and its products after rearrangement and splicing. V, Variable region; D, diversity region; J, joining region; L, leader exon.

tion in order to achieve this function. In very general terms, the pre-B lymphocyte represents the period of differentiation at which the stage is set for the synthesis of immunoglobulins; stepwise rearrangement of immunoglobulin genes leads to the generation of cells which first express intracellular p H chains. Intracellular membrane immunoglobulin at this stage plays a feedback role in differentiation, a role that is expanded on later in this chapter. The B cell stage corresponds to the antigenresponsive portion of ontogeny ; membrane immunoglobulin is expressed on the cell surface, where it functions as the antigen receptor. Antigenactivated B cells differentiate into plasma cells, whose sole function is to

4

SHIV PILLAI

Pre-B lntraceliular Pm lntracellular Ps

B Cell surface

Plasma cell

Prn 2 L2

Secretion of

lntracellular Ps 2 L2 No J chain synthesis No sulfydryl oxidase

pentamers

FIG. 2. A highly simplified view of the process of B cell differentiation.

secrete antibody, thus serving as the effector end cells of the humoral immune system. In the following subsections a more detailed description of all the defined cellular stages in B cell ontogeny is provided. These stages are schematically represented in Fig. 3. A. PRO-BLYMPHOCYTES

The sites of B lymphocyte generation are the spleen and the liver before birth and the bone marrow in adult life. The earliest identifiable precursor of a B cell is a pro-B lymphocyte; a pro-B cell is committed to the B lineage, but has not yet undergone any rearrangements at the H-chain locus. A number of genes of functional significance in the B lineage are known to be expressed at this stage. These include the A5 (Sakaguchi and Melchers, 1986) and uPreB (Kudo and Melchers, 1987) genes, which encode the o and L surrogate immunoglobulin L chains; the RAG-I and RAG-2 genes (Schatz et af.,1989; Oettinger et d.,1990), which encode the V-D-J recombinase; and the rnb-1 (Sakaguchi et al., 1988) and B29 (Her-

IMMUNOGLOBULIN TRANSPORT Pro - B no rearrangement

5

-

Pre B lntracellular

pIwl1

FIG. 3. A detailed view of B lymphocyte ontogeny.

manson et al., 1988) genes, which encode presumed signal transduction/ transport-relatedproteins that associate with membrane immunoglobulin. Rearrangement of immunoglobulin genes in pre-B cells depends on the activity of a site-specific V-D-J recombinase, which is presumably the product of the pre-B and pre-T-specific RAG-I and RAG-2 genes, and other stage-specific DN A-binding factors that bind to specific regulatory

6

SHIV PILLAI

motifs in immunoglobulin genes. These DNA-binding proteins are transacting factors that selectively “open” specific immunoglobulin loci, making these regions accessible to the V-D-J recombinase, which, in turn, recognizes the conserved heptamer-nonamer signals for rearrangement (reviewed by Yancopoulos and Alt, 1986). B. PRE-BLYMPHOCYTES While pro-B lymphocytes are operationally defined as cells committed to the B lineage which have yet to rearrange their immunoglobulin genes, pre-B cells are B cell precursors in the process of rearranging their immunoglobulin genes. Early pre-B cells are generally believed to be large bone marrow pre-B cells actively rearranging their H-chain loci. Late pre-B cells are smaller cells which are either primed to rearrange, or are in the process of rearranging, their L-chain genes. The first rearrangement event that occurs in pre-B cells is a D-J rearrangement involving the H-chain locus. It has been demonstrated that D-J rearrangements may give rise to very unusual D p transcripts; these transcripts initiate upstream of the rearranged D segment and include a conserved ATG codon which is in frame with the coding exons of the p gene (Reth and Alt, 1984). Amazingly, the transcript includes, downstream of the ATG codon, a set of codons derived from the recombinational signal sequences immediately upstream of the D segment, which constitute an upstream hydrophobic leader peptide. Translation of the D p transcript leads to synthesis of the D p protein, which is translocated into the lumen of the endoplasmic reticulum (ER). It has been suggested that high-level expression of the D p protein may serve as a signal for further differentiation of pre-B lymphocytes. Preliminary studies suggest that both Dpm and Dps proteins are synthesized and that Dpm is not transported to the cell surface (S. Pillai, unpublished observations), suggesting that a signal for further differentiation may be generated from an intracellular location. It is unclear whether the generation of a Dp protein-containing cell is an obligate intermediate stage in the process of B cell differentiation, and no direct evidence exists to either confirm or refute any model which invokes the need for synthesis of a Dp protein.’ The next step in pre-B lymphocyte differentiation involves the rearrangement, again at the H-chain locus, of a V segment to the D segment that has undergone a previous rearrangement, to give rise to a D-J structure. There is a one-in-three chance of any V-to-D-J rearrangement giving rise to an in-frame rearranged H-chain immunoglobulin gene. During the joining process the addition of a small number of bases (N regions) or the I

See Note Added in Proof, p. 36.

IMMUNOGLOBULIN TRANSPORT

7

removal of a few bases contributes to junctional diversity and amplifies the creation of the immune repertoire. It is important from the viewpoint of the immune system that any given B cell clone express only a single receptor in order to maintain clonal specificity in the immune system. In any given B cell, there is the theoretical possibility of in-frame rearrangements of the H-chain gene on both maternally and paternally derived chromosomes, giving rise to two nonidentical H-chain proteins. It is now well appreciated that a productive H-chain gene rearrangement that leads to the synthesis of an intracellular p protein initiates a feedback regulation process (reviewed by Alt et al., 1986). This process is capable of shutting off H-chain gene rearrangement on the other allele and also provides a positive signal, permitting further differentiation to the stage of L-chain gene rearrangement. The feedback regulation process is of direct relevance to studies of the transport of immunoglobulins in pre-B cells. It is now clear that the signal for the feedback regulation of differentiation in pre-B cells is dependent on the synthesis of pm, not ps (Nussenzweig et al., 1987; Reth et al., 1987). At this stage of differentiation, pm is associated with the w and L surrogate immunoglobulin L chains, which are the products of the pre-B-specific A5 and vPreB genes, respectively. Important issues related to immunoglobulin transport at this stage include the localization of pm and the site from which a differentiation signal is generated by this protein. C. PRE-B-SPECIFIC SURROGATE IMMUNOGLOBULIN L CHAINS Pre-B cells have long been described as the stage of B cell ontogeny in which cells contain “free” cytoplasmic H chains. It is now well established that in pre-B cells at least a proportion of the intracellular immunoglobulin H chain is not free, but is complexed with the w and L surrogate immunoglobulin L chains. The w surrogate L chain is a 20-kDa protein that is pre-B-specific and forms disulfide-linked tetramers with the p chain (Pillai and Baltimore, 1987a; Cherayil and Pillai, 1991a). The L chain is a 15-kDa protein that is noncovalently associated with the p2-0~2complex and is also pre-B specific (Pillai and Baltimore, 1988). The w protein was not found to be associated with the D p protein in pre-B cell lines, and was not associated with p in a fibroblast cell line expressing a transfected pm gene nor in a T cell line expressing a transfected p gene. In a subset of pre-B cell lines (which we refer to as Type I1 pre-B cell lines, to distinguish them from Type I pre-B cell lines in which pm is only detectable as an intracellular protein), pm is transported to the cell surface (Findley et al., 1982; Gordon etal., 1981; Hardy et al., 1986; Hendershot and Levitt, 1984; Paige et al., 1981). In these lines, cell surface pm was found to be associated with w and L (Pillai and Baltimore, 1987a; 1988) or equivalent human

8

SHIV PILLAI

proteins (Hollis et al., 1989; Kerr et al., 1989). Indeed, in a pre-B cell line, 70213, in which pm can be induced to migrate to the cell surface in the absence of K L chain gene transcription (Paige et al., 1981), it does so in association with w and L ( S . Pillai, unpublished observations). The w chain is the product of the pre-B-specific A5 gene, and the L chain is the product of another pre-B-specific gene, the uPreB gene (Cherayil and Pillai, 1991a; Karasuyama et al., 1990; Kudo and Melchers, 1987; Sakaguchi and Melchers, 1986; Tsubata and Reth, 1990). Similar chains have more recently been described in human pre-B cell lines. The pre-B-specific human homolog of the A5 gene is the 14.1116.1 gene (Chang et al., 1986). The A5 and 14.1/16.1genes are structurally similar to conventional L-chain genes over the 3’ halves of their coding regions. The predicted structure of these genes, which respectively encode the mouse and human w surrogate L-chain proteins, includes a signal peptide followed by an amino-terminal segment which has no homology to any known protein. This is followed by an immunoglobulin J domain-like segment, and finally a domain with a structural resemblance to Ch and CK L-chain domains. A preterminal cysteine residue analogous to the preterminal cysteine in conventional L chains is disulfide linked to the free cysteine in the CHI domain. The sequence of the uPreB gene predicts a V domain-like structure preceded by a signal peptide. A likely but unproven model for the association of the p H chain with surrogate L-chain proteins is depicted in Fig. 4. It is assumed that the dvPreB protein associates with the VH domain of

FIG.4. Presumed structure of the heavy-surrogate light-chain complex.

IMMUNOGLOBULIN TRANSPORT

9

the H chain and that the carboxy-terminal half of the w/h5 protein associates with the CH1 domain, with an interchain disulfide bridge ensuring covalent association. We assume that these proteins associate with p m in pre-B cells in order to participate in the generation of a feedback signal, presumably through the six polypeptides (Pillai and Baltimore, 1988)of the pm activation complex. There are three related models for the function of the o protein in this process. The first model is the solubility/stability model, which assumes that the surrogate L chains serve a purely physical L-chain-like purpose, which is to prevent H chains from aggregating or being in an “inappropriate” conformation; an “appropriate” conformation may be necessary for movement or for engaging the proteins of the pm activation complex in order for a signal transduction event to occur. This model assumes that if, as part of the feedback process, pm needs to be transported to the cell surface or to some intracellular compartment, it cannot attain competence for transport unless ass6ciated with either a conventional L chain or surrogate L chains. The second model is the intracellular ligand/crosslinking mode. In this model, feedback regulation depends on the generation of a signal from an intracellular location, and w cannot be functionally replaced by a conventional L chain. The intracellular site of signal generation is presumed to be the ER itself or the cis-Golgi compartment. In this model, the association of surrogate L-chain proteins with p m in an intracellular location is an absolute prerequisite for both engaging and “turning on” the pm activation complex. The third model for feedback regulation is the cell-surface activation model. This model suggests the existence of a physiological differentiation stage in which, prior to rearrangement of the K locus, the pm-surrogate L-chain complex is transported to the cell surface. In such a model, an extracellular ligand may engage this pm-receptor complex and lead to the generation of a differentiation signal. In theory, such a signal, if it were to activate NFKB(Sen and Baltimore, 1986), could lead to the transcriptional activation of the K locus -and rearrangement of the K gene, leading to the next stage of differentiation. The latter two models are schematically depicted in Fig. 5. The genes for surrogate L-chain proteins are transcribed in pro-B cells prior to the onset of H-chain gene rearrangement. It is unclear whether the w/h5 protein has any function in pro-B cells. It has been suggested that w and L are coassociated in pro-B cells (Misener et al., 1990). In our studies of the w/X5 protein, we have observed a number of protein species coassociated with the w protein in a pro-B cell line (Cherayil and Pillai, 1991b). It is unclear whether any of these proteins are candidate intracellular ligands for the pw receptor which appears later in ontogeny. The major w species

10

SHIV PILLAI

I

A

I

2nd

B

FIG. 5. Models for differentiationsignal generationlfeedback regulation in pre-B lymphocytes. (A) Intracellular ligand crosslinking model. (B) Extracellular ligand activation model.

in pro-B cells is an 02 dimer. This dimer, however, is not secreted (Cherayil and Pillai , 1991b) , It appears likely that the association of p with w and L can occur only as a trimolecular complex (Karasuyama et al., 1990; Tsubata and Reth, 1990; Cherayil and Pillai, 1991a)The D p protein contains the CH1 domain of the p H chain, but lacks a V domain. The presence of the CH1 domain may have been predicted to be sufficient to permit association with the wlh5 protein; however, neither form of Dp protein is coprecipitated with w (Pillai and Baltimore, 1987),presumably because association with vPreB/L (which would require the presence of a V domain) is a necessary prerequisite for p--0 association.* TRANSITIONAL AND IMMATURE D. PRE-BTO B CELLTRANSITION: B CELLSTAGES At least three molecular events related to immunoglobulin genes and proteins characterize the pre-B to B cell transition. At this transition, cells

* See Note Added in Proof, p. 36.

IMMUNOGLOBULIN TRANSPORT

11

acquire the ability to transport membrane immunoglobulin to the cell surface, they constitutively activate transcription of the K L-chain gene, and they shut off transcription of genes encoding surrogate L-chain proteins. The transition of a pre-B cell to the antigen-responsive B stage is an antigen-independent event. The earliest B cell to emerge from the pre-B compartment in the bone marrow is probably a recently defined transitional B cell (Cherayil and Pillai, 1991a). Transitional B cells simultaneously express p--w and p-K cell surface receptors. Cell lines have been derived that represent this transitional B stage which simultaneously express the N and K genes (McGarigle et al., 1991). At the pre-B to B cell transition, the constitutive activation of NFKB and K gene transcription clearly precedes the shut-off of N gene expression. The next stage of B cell differentiation is the immature B cell stage at which cells express p-K or p-A receptors and have permanently shut off surrogate L-chain gene expression. It is presumed that at the immature B stage, cells respond negatively to ligation of their receptors by antigen and die. This is one of the two major mechanisms by which B cells are rendered tolerant to self antigens (reviewed by Goodnow et al., 1990).

E. MATUREIGM/IGD-EXPRESSING B LYMPHOCYTES The next stage of B cell differentiation is the surface immunoglobulin M/immunoglobulin D (IgM/IgD) positive mature B stage. Differentiation until this stage is antigen independent. If cells at the two previous transitional and immature B stages are exposed to antigen, they are presumed to respond negatively and may thus be eliminated. At the mature B IgM/IgD stage, cells respond positively when exposed to antigens by proliferating and undergoing further differentiation. The frequently used term “virgin B cell” covers surface immunoglobulin B cell stages prior to activation by antigen; it includes transitional, immature, and early B cells. There are three aspects to the function of membrane immunoglobulin at the mature B cell stage. Polyvalent antigens such as polysaccharide antigens, which are “T independent” (and can produce biological responses in the absence of T cell help), directly engage membrane immunoglobulin and transduce a signal which leads to B cell activation. Although, in the case of “T-dependent” protein antigens, membrane immunoglobulin may still play a signal transductional role, its major function is to internalize the cognate antigen so that the latter may be processed in an acidic endosomal compartment into peptide fragments; appropriate fragments associate with the cleft of the class I1 histocompatibility molecule and are presented on the cell surface to a cognate helper T lymphocyte. This T cell, in turn, secretes the appropriate lymphokines, such as interleukins 4 and 5, which

12

SHIV PILLAI

drive further differentiation of the concerned B lymphocyte clone. From the standpoint of protein trafficking and immunoglobulin function, membrane immunoglobulin in mature B cells must therefore be part of a signaltransducing protein complex that is transported to the cell surface and must also function as an endocytic receptor. An interesting third aspect to the intracellular transport of membrane immunoglobulin at this stage of differentiation has recently emerged. The exposure of mature B cells to a cognate antigen in the absence of appropriate T cell help can lead to the down-regulation of membrane IgM, but not membrane IgD, on these cells. This is presumed to be one of the mechanisms by which clonal anergy to self antigens may be perpetuated. The actual function of membrane IgD remains unknown. IgD is not expressed at any subsequent stage of B cell ontogeny after the mature B cell stage. F. ACTIVATED AND MEMORY B LYMPHOCYTES

After mature B cells are activated by protein antigens, two unusual events involving rearranged immunoglobulin genes contribute to increased recognitional as well as functional diversity in the B lineage. Somatic mutation is a process by which rearranged immunoglobulin V segments in activated B cells are further diversified by a unique and poorly understood mutational mechanism. Somatic mutation contributes to greatly increased immune diversity at the antigen recognition level. Isotype switching, which involves a somatic deletional recombination event, leads to the generation of activated B cells with unchanged antigen specificities, but the "new" C regions, and contributes to the production of IgG, IgA, and IgE antibodies (reviewed by Rajewsky et al., 1987). Secretion of antibody, from the immunological viewpoint, should be a consequence of exposure to antigen and should not occur in early nonactivated B cells. An additional regulatory event that characterizes the activated B stage is acquisition of the ability to secrete antibody. Until and including the mature B stage, secretory IgM is synthesized but is held back intracellularly . After activation, IgM is no longer sequestered intracellularly and is secreted; for other isotypes, all of which appear in ontogeny only subsequent to activation by antigen, holdback mechanisms are irrelevant. Activated B cells may differentiate to give rise to either long-lived memory B cells or antibody-secreting plasma cells. Memory B cells may be defined as long-lived activated B cells whose life spans are measured in years instead of days. Exposure to antigen leads to proliferation and the secretion of antibody by memory B cells, which may also divide asymmetrically to give rise to plasma cells.

IMMUNOGLOBULIN TRANSPORT

13

G. PLASMACELLS:ENDSTAGEOF B LYMPHOID ONTOGENY Plasma cells are the end-stage cells of B cell ontogeny and are basically designed to be antibody-secreting factories. These cells have numerous ribosomes and extremely abundant reticular ER and Golgi compartments. Largely by posttranscriptional mechanisms, they achieve high secretoryto-membrane immunoglobulin H-chain RNA ratios; the mRNA level of secretory immunoglobulin is also regulated at the level of increased message stability (Mason el al., 1988), possibly because it is efficiently translated and is protected from degradation by virtue of association with ribosomes. These cells express no cell surface class I1 molecules and also do not express membrane immunoglobulin.

111. Intracellular Retention of Secretory Immunoglobulins From a purely teleological viewpoint, immunoglobulins should not be secreted at all stages of B cell ontogeny prior to antigen exposure. The only isotypes that are expressed by B lymphocytes prior to activation by antigen are the p and 6 isotypes. Any mechanism that prevents the secretion of immunoglobulins prior to exposure to antigen should therefore apply to these isotypes. The question of the retention of secretory IgD is a moot one; although rare IgD-producing myelomas do exist, at the mature B stage, when IgD is expressed on the cell surface along with IgM, RNAprocessing mechanisms lead to the generation of pm and ps transcripts, as well as two 6m transcripts, but no 6s transcripts. When mature B cells are activated, further differentiation leads to shut-off of the mechanisms which generate 6 transcripts. Pre-B cells are unusual from a viewpoint which has interesting implications for studying the assembly and transport of multisubunit oligomers. Immune receptors, unlike other receptors made up of more than one subunit, are synthesized in stages; the immunoglobulin H chain is synthesized prior to the L chain, and the T cell receptor p chain is synthesized before the a chain. The process of making coding region joints during immunoglobulin gene rearrangement is designed to be imprecise in order to increase junctional diversity and thus to maximize the immune repertoire. However, the efficiency of this process depends on the sequential rearrangement and expression of H-chain and L-chain genes and the stepwise selection of cells that make in-frame rearrangements. Pre-B cells, as a result, constitute natural developmental models for studying the assembly, transport, and degradation of incomplete oligomeric proteins. In bone marrow pre-B cells, alternative splicing gives rise largely to pm mRNA, and negligible amounts of ps may actually be synthesized by these

14

SHIV PILLAI

cells in uiuo (Thorens et al., 1985). However, most pre-B cell lines synthesize roughly equivalent amounts of pm and ps, at the message as well as at the protein level. ps synthesized by pre-B cell lines is not secreted. There are two putative retention mechanisms that might serve to hold secretory immunoglobulins intracellularly . One potential mechanism applies to any immunoglobulin isotype, is not B lineage specific, and involves the association of free secretory H chains with binding protein (Bip), an abundant ER protein; this mechanism presumably involves the association of Bip with a CH1 domain. The second potential mechanism is specific for the p isotype, is developmentally regulated, and applies equally well to tetramers of ps with conventional or surrogate L chains as it does to free ps. This second mechanism presumably involves retention via a cysteine tailpiece and may not really be distinct in principle from the CH1dependent retention process. A. INTRACELLULAR RETENTION OF H CHAINAND BIP IN PRE-B AND THE SALVAGE PATHWAY CELLS:KDEL SEQUENCES A H-chain binding protein was originally described by Morrison and Scharff (1975) in a nonsecreting myeloma variant which had lost L-chain expression. This protein was rediscovered by Haas and Wabl (1983) in H-chain-only pre-B cell lines and hybridomas derived from pre-B lines. These latter workers described a doublet of proteins, which were 78 and 70 kDa in size and associated with free H chains, and referred to these proteins as the binding protein. Some of the confusion surrounding Bip was resolved when Munro and Pelham (1986) cloned the gene encoding the 78-kDa Bip and discovered that it was identical to a much-studied member of the heat-shock protein family, GRP78, which is induced by glucose starvation. The 70-kDa protein that was originally considered to be part of the Bip complex is probably a cytosolic heat-shock family protein, hsc 70. Bip/GRP78 was found to be coprecipitated with misfolded mutants of influenza hemagglutinin which were incapable of being transported to the cell surface, and the suggestion was made that Bip may play a role in identifying and sequestering misfolded or improperly assembled proteins in the ER (Gething et al., 1986). Bip/GRP78 has been demonstrated to possess a peptide-dependent ATPase activity (Flynn et al., 1989) and can be dissociated from misfolded proteins by the addition of ATP in vitro (Munro and Pelham, 1987). Bip/GRP78 has also been found to be associated transiently with assembly intermediates of some secreted proteins, and it has been suggested that the function of Bip may actually be to catalyze some step in the folding and assembly of proteins in the ER that are destined for export: It may only remain associated with incomplete

IMMUNOGLOBULIN TRANSPORT

15

oligomers or misfolded variants (Kassenbrock et al., 1988). Neither of these two models for the function of Bip has been established, and neither may be correct. Bip constitutes a well-studied example of a “resident” ER luminal protein which has a KDEL retention sequence and is also, like other members of this category of resident ER proteins, a calcium-binding protein. Proteins that possess a KDEL retention signal recycle back and forth between the ER and a pre-Golgi salvage compartment (Lewis et al., 1990). The KDEL receptor binds to proteins such as Bip with a high affinity in the salvage compartment; recycling to the ER occurs rapidly and in this compartment luminal proteins such as Bip have virtually no affinity for the KDEL receptor, which returns “unoccupied” to the salvage compartment. Bip-immunoglobulin H-chain complexes probably cycle back and forth in this manner in pre-B cell lines. The homolog of the Bip gene in Saccharornyces cereuesiae is the KAR2 gene. KAR2 is an essential gene in yeast, is required for nuclear fusion, and is assumed to play some important role, as yet undetermined, in the secretory process (Rose et al., 1989). KAR2 was somewhat unexpectedly found to be required for protein translocation into the ER (Vogel et al., 1990). Such a function would not have been predicted for Bip from studies of mammalian cells. Although it remains clear that Bip/GRP78/KAR2 must play some important role in cell physiology, this function remains to be determined with any certainty. The best-studied, though not necessarily the best-understood, role of Bip may be its putative ability to retain secretory immunoglobulin H chains in an intracellular location in some cell lines. This presumed role in retaining H chains intracellularly is based on the correlation of the association of Bip with nonsecreted H chains. It is equally likely that the association of Bip with H chains is a consequence of a failure of immunoglobulin secretion rather than the cause of retention. The primary basis for suggesting that Bip itself may play a role in the retention of immunoglobulins has come from studies (Bole et al., 1986; Hendershot et al., 1987; Hendershot, 1990) of a number of secreted and nonsecreted immunoglobulin mutants in a range of myeloma lines. In the absence of an L chain, wild-type immunoglobulin H-chain protein of a number of isotypes is retained intracellularly along with Bip. Mutant y immunoglobulins which lack CH 1 domains are secreted in the absence of an L chain. Mutants lacking CH2 or CH3 domains remain associated with Bip and are not secreted. Since L chains associate with the CH1 domain and since L chains displace Bip from nascent H-chain-Bip complexes, it has been proposed that Bip binds to the CH 1 domain of immunoglobulins and serves to retain unassembled H chains which are then degraded intra-

16

SHIV PILLAI

cellularly . It is equally possible that the association of nonassembled immunoglobulin H chains with Bip is the consequence, rather than the cause, of intracellular retention. A second site for Bip association in the cysteine-containing tailpiece of the CH4 domain of ps has been suggested, in part, to explain the retention of p 2 - ~ 2complexes in early B cells. Since this cysteine-dependent mechanism could presumably be involved in the retention of ps in pre-B cells, is there any physiological role for the Bip/CH 1 domain-dependent retention of free ps? This is probably a moot question, since bone marrow pre-B cells may synthesize relatively minuscule amounts of ps mRNA to begin with. However, in a pre-B cell line which has switched to the y isotype (which lacks the cysteine-containing retention tailpiece found in p ) , y s is retained, intracellularly associated with Bip. Association of y s with a K L-chain protein in a transfected derivative of this cell line leads to the “release” of the H chain from Bip (or merely the assumption of an appropriate conformation by the tetrameric complex) and its quantitative secretion as ys2-~2 tetramers (Bachhawat and Pillai, 1991). In the next subsection, a model is suggested that unifies the cysteine tailpiece retention mechanism with the Bip holdback hypothesis. Needless to say, it remains unresolved as to whether Bip binding is a cause or a consequence of retention. B. CYSTEINE TAILPIECE-DEPENDENT RETENTION OF SECRETORY IMMUNOGLOBULINS The retention of secretory IgM is mechanistically and structurally closely linked to the processes involved in the polymerization of this molecule prior to secretion. Both ps and as H chains contain a tailpiece which contains a cysteine residue. This tailpiece plays an important role in the polymerization of these isotypes and their association with the J chain. Issues that relate to the function of this tailpiece in secretion are dealt with in greater detail in the section on immunoglobulin transport at the plasma cell stage (Section V,A). The relevance of this tailpiece to the intracellular retention of ps in the earlier stages of B cell differentiation is considered here. Numerous studies have indicated that in “virgin” splenic B cells as well as in many B cell lines that represent the early antigen-independent stages of B cell differentiation, tetramers of pm and L chain are transported to the cell surface, while ps, which also forms tetramers with an L chain, is retained intracellularly and degraded (Sibley el af., 1980; Vassali et af., 1980; Sidman, 1981; Dulis, 1983; Rubartelli et af., 1983). Three models have been suggested for the failure of IgM secretion by early B cells.

IMMUNOGLOBULIN TRANSPORT

17

1. Early B cells lack an important constituent responsible f o r the assembly of polymerized IgM, and the absence of this assembly step results in failure of secretion. Candidates for a missing or limiting low-level constituent at the B cell stage are (a) the J chain (Koshland, 1985) and (b) the IgM polymerase, a sulfhydryl oxidase (Roth and Koshland, 1981). The role of the J chain and the IgM polymerase is expanded on in the section on transport in plasma cells (Section V,A). However, the J chain is not essential for polymerization of IgM. IgM is polymerized and secreted in the absence of J chain by transfected glioma and pheochromocytoma cells (Cattaneo and Neuberger, 1987). The absence of polymerization and assembly processes in earlier B cell stages may well be tightly linked to the tailpiece-dependent retention process discussed later in this section. However, polymerization is not an absolute prerequisite for secretion; IgA is certainly secreted as monomers as well as dimers, and mutant IgM molecules that cannot form polymers are efficiently secreted (Baker et al., 1986). 2. Early B cells are intrinsically incompetent for imunoglobulin secretion because of a poorly developed ER. Although this argument has frequently been advanced, experimental evidence suggests otherwise. In a B cell line, representative of the immature B stage, transfection of a ps gene with a mutated tailpiece retention signal resulted in immunoglobulin secretion, suggesting that these cells are intrinsically competent for immunoglobulin secretion (Sitia et al., 1990). In a pre-B cell line which has switched to the y isotype (an isotype which lacks a retention signal), introduction of an L-chain gene led to efficient secretion (Bachhawat and Pillai, 1991), further indicating that early B lineage cells do not lack the machinery necessary for the secretion of secretory immunoglobulins. In the case of secretory IgM (as discussed in item l), early B cells may certainly lack some polymerizing activity, and this lack might contribute to the retention process, for tailpiece-containing immunoglobulins. 3 . The ps chain contains structural information that leads to its retention in early B lineage cells. The 19-amino-acidcysteine-containing secretory tailpiece at the carboxy terminus of the ps H chain has been demonstrated to be involved in the retention of ps in a stage-specific manner. Mutation of cys-575 (the penultimate residue) leads to the secretion of secretory IgM by B cell lines which represent early stages of differentiation and which retain and degrade wild-type ps-containing IgM molecules. Engineering this tailpiece onto a y H-chain gene converts IgG to a protein that is now retained by early B lineage cells. The current evidence suggests that in early B cells the cysteine tailpiece serves as a signal for retention of secretory IgM; this mechanism applies to

18

SHIV PILLAI

heavy-light complexes and is therefore independent of the retention mechanism that depends on an available CH1 domain. When a deletion mutant of the ps H-chain gene lacking a CHI domain was transfected into a plasmacytoma cell line, the mutant protein was retained intracellularly and was associated with Bip. When this deletion mutant was further altered, converting the cysteine in the tailpiece at position 575 to an alanine, the resulting protein was efficiently secreted (Sitia et al., 1990). Two interpretations remain for this phenomenon. The first is that this tailpiece-dependent mechanism involves the binding of Bip to the tailpiece and hence the. retention. The alternative mechanism is that the tailpiece plays a role in retention independently of Bip, and the retained intracellular immunoglobulin remains associated with Bip. It is worth noting in this context that, although ps2-L2 tetramers and the CHI deletion mutant of ps are both retained intracellularly by means of the tailpiece mechanism, Bip association is prominent with the deletion mutant and barely detectable in the case of IgM heavy-light complexes (Sitia et al., 1990). This argues for a secondary role for Bip which remains associated with nonassembled and retained secretory immunoglobulins. In the B lineage, this retention mechanism is restricted to pre-B and early B cells. However, it is unlikely that this mechanism is B lineage specific. It appears more likely that the tailpiece-dependent retention mechanism is present in many cell types, but is abrogated in certain differentiated cells. Transfected glioma cells can secrete IgM (Cattaneo and Neuberger, 1987) and transfected fibroblasts can secrete hemimers of ps and surrogate L chains (Karasuyama et al., 1990), suggesting that Bip (which is synthesized by all cells) may not be a major player in such a retention mechanism. However, some nonlymphoid lines, including transfected Chinese hamster ovary cells and HeLa cells, retain secretory IgM (Cattaneo and Neuberger, 1987), suggesting that retention may be the phenotype of cell types which possibly lack some specialized “assembly machinery.” This putative machinery presumably leads to the masking of retention signals and is assumed to be a characteristic feature of certain specialized cells, which include activated B cells, plasma cells, and glioma and pheochromocytoma cells. When mature B cells are activated following exposure to antigen, this retention mechanism is presumably shut off and remains silent in all subsequent stages of B cell differentiation. It is possible that, as B cells differentiate, they acquire the ability to polymerize IgM and thus to mask the tailpiece retention signal. The only other isotype other than the p isotype in which a cysteine-containing tailpiece is present is a. This tailpiece in secreted IgA is believed to play an important role in the process of dimerization and association with the J chain. Since IgA is synthesized

IMMUNOGLOBULIN TRANSPORT

19

only by activated B cells which have undergone class switching (and have presumably shut off the tailpiece-dependent retention mechanism), this mechanism is probably of no relevance to IgA secretion. IgA can be secreted as monomers as well as dimers. In studies of an IgA-expressing B lymphoma, nonactivated lymphoma cells expressed primarily am message and virtually no as message. Activation of these cells with bacterial lipopolysaccharide led to the synthesis of secretory IgA. This IgA was not retained by these cells, arguing that the tailpiece-dependent retention mechanism for IgA is missing in activated B cells (Sitia et al., 1985). Exposure of early B lineage cells to 2-mercaptoethanol leads to immunoglobulin secretion (Alberini et al., 1990), possibly by interference with the cysteine tailpiece-dependent retention mechanism. It is possible to unify the existing knowledge of secretory immunoglobulin retention into a single framework. Two structural features of secretory immunoglobulin H chains may play a role in retention. In cells that lack L chains, an exposed CH1 domain may serve as a retention signal, particularly for the y isotype. Since retention of ps in pre-B cells could occur through the other retention signal, the tailpiece in the CH4 domain, the physiological significance of retention through the CHI domain during B cell differentiation is certainly open to question. Retention through the CHI domain as well as through the CH4 domain may depend on two alternative mechanisms. 1 . The Bip-dependent model. In this model, the CH1 domain of all isotypes as well as the CH4 tailpiece in p and Q is considered to be a binding site for Bip. Bip binding to the CH1 domain can be masked by L-chain association. The binding of Bip to the tailpiece can be masked by the process of IgM polymerization. It is conceivable that the cysteine in the CH1 domain involved in L-chain association may be part of a Bip binding site. It was suggested by Hendershot et al. (1987) that since mutants lacking this cysteine in the Q chain CH1 domain still bound Bip, this residue could not contribute to Bip binding. It is quite likely that in these mutants Bip was bound to the tailpiece and therefore the CHI cysteine might still constitute part of a Bip binding site. 2. The incomplete assembly model. This model assumes that some aspect of the free CHI domain and also of the tailpiece leads to the retention of p H chains in pre-B cells and nonpolymerized IgM in nonactivated B cells. Unpaired cysteines in the CHI domain or in the tailpiece could contribute to misfolding, thus preventing assembly and secretion, or may directly associate with retention “receptors.” Alternatively, some other structural feature of these retention domains may contribute to the holdback process. The association with Bip is considered, in this model, to be a consequence, rather than a cause, of retention.

20

SHIV PILLAI

IV. Membrane Immunoglobulin Transport during B Cell Ontogeny

An important issue for the regulation of pre-B cell differentiation as well as the transport of immunoglobulins is the existence in uiuo of a physiological “Type 11” pre-B stage at which membrane immunoglobulin is transported to the cell surface. The existence of such a stage would suggest that a ligand may exist for the p-o receptor, which could help drive differentiation from the pre-B to the B stage. The B cell stage of lymphoid ontogeny may be subdivided into the transitional, immature, mature, activated, and memory B cell stages. In all these stages, membrane immunoglobulin functions as a signal transduction receptor and must be transported to the cell surface. In the antigenindependent immature stage, signal transduction may lead to cell death. In the later antigen-dependent stages, signal transduction through membrane immunoglobulins leads to proliferation and further differentiation. In these stages, membrane immunoglobulin must also be capable of functioning as an endocytic receptor for the internalization and processing of protein antigens. In these stages, exposure to (self) antigens in the absence of cognate T cell help may also lead to the phenomenon of receptor downregulation, which may be one of the mechanisms of perpetuating clonal anergy . A. ER DEGRADATION VERSUS CELLSURFACE TRANSPORT: FATEOF MEMBRANE IMMUNOGLOBULIN IN PRE-B CELLS

Do pre-B cells express pm on the cell surface? Is the expression of conventional K or X L chains a prerequisite for the cell surface expression of pm? What is the molecular basis for acquisition of the ability to transport pm to the cell surface at the pre-B to B cell transition? Does cell surface expression of pm precede activation of K gene expression? These closely interrelated questions are of interest from the viewpoint of understanding immunoglobulin transport and are central to understanding regulatory mechanisms involved in the pre-B to B cell transition. Most pre-B cells in the bone marrow do not express detectable amounts of pm on the cell surface. Indeed, in pre-B cultures derived from bone marrow or fetal liver, the pre-B population expressing intracellular p is clearly distinguishable from infrequently seen surface p-positive B cells. Anti-p treatment in uiuo eliminates early B cells, but leaves most pre-B cells unaffected, which is consistent with the view that most pre-B cells do not express pm on the cell surface (Burrows et al. 1978). Whether a small subpopulation of pre-B cells which express detectable surface p in associ-

IMMUNOGLOBULIN TRANSPORT

21

ation with surrogate L chains actually exists in the bone marrow or the fetal liver is an important unresolved issue in B cell ontogeny. h e - B cell lines in which the H-chain locus has been productively rearranged fall into two categories. The majority of these lines (which we term Type I) contain only intracellular p. In these lines, p is associated with the o and L surrogate immunoglobulin L chains and is rapidly degraded. In a subset of mouse and human pre-B cell lines (which we term Type I1 pre-B cells), pm is transported to the cell surface. In subclones of a murine lymphoma, 70213, in which most cells do not express pm on the cell surface, exposure to dextran sulfate (a B cell mitogen) leads to surface expression of p in the absence Of K L-chain synthesis (Paige et al., 1981). In Type I1 pre-B cell lines as well as in dextran sulfate-induced 70Z/3 cells, we have demonstrated that pm on the cell surface is associated with the o and L surrogate L chains. From the above studies it is clear that the expression of p on the cell surface does not depend on the synthesis of conventional K L chains, but may, nevertheless, depend on proper assembly with surrogate L-chain proteins as well as some poorly defined molecular event that may be regulated by dextran sulfate in 70213 cells. This event permits pm to evade intracellular degradation and to be transported to the cell surface. One approach to the issue of the cell surface expression of p in the pre-B and B stages of differentiation is to recognize the existence of two competing pathways for membrane immunoglobulin transport in these cells. The rapid degradation of pm in pre-B cells is part of an intracellular editing process that targets incomplete oligomeric receptors for degradation in a pre-Golgi compartment, probably the ER itself. As pre-B cells differentiate, somewhere at the late pre-B stage, they acquire the ability to “rescue” pm complexed with surrogate L chains from the intracellular ERdegradative pathway, and to transport this complex to the cell surface. Even at the B stage, after the synthesis of the K chain, both of these pathways coexist; a portion of the (presumably imcompletely or incorrectly assembled) p m in mature B cells is degraded intracellularly, and the remainder is rescued and transported to the cell surface (Dulis et al., 1982). The ER-degradative pathway for incomplete oligomers of membrane immunoglobulin in pre-B cells can be distinguished from the degradative pathway for incomplete oligomers of secretory immunoglobulin (Bachhawat and Pillai, 1991). This difference may indeed be related to the ability of nonassembled secretory immunoglobulin H chains to bind to Bip (whether or not Bip causes retention or merely associates with molecules that are incapable of movement), while nonassembled membrane immunoglobulin H chains perhaps do not associate with as efficiently with Bip. The Bip-

22

SHIV PILLAI

associated secretory immunoglobulin may shuttle back and forth between the ER and the salvage compartment prior to degradation. Alternatively, transmembrane hydrophilic topogenic sequences may determine the rapid degradation of nonassembled membrane immunoglobulins. The possible role of such a topogenic sequence is described in the next subsection. In pre-B cells, pm that is either free or associated with w and L is rapidly degraded. It is unclear whether p chains that are associated with surrogate L chains are less susceptible to degradation than are free p chains. However, the levels of expression of the genes for surrogate L chains (the A5 and uPreB genes) are similar in both Type I and Type I1 pre-B cell lines. Similarly, the levels of expression of two B lineage genes, the mb-I and B29 genes, whose products play a role in the surface expression of pm in B cells are also comparable to mature B cell levels in both Type I and Type I1 pre-B cell lines. What regulates the ability to transport pm to the cell surface at the pre-B to B cell transition? Issues such as the role of pmassociated proteins, the assembly process, and the hydrophobicity of pm are discussed in detail in Section IV,B. B. TRANSPORT OF MEMBRANE IGM: ASSOCIATED PROTEINS AND THE ROLEOF HYDROPHOBICITY As discussed in Section IV,A, membrane immunoglobulin H chains have two distinct fates after synthesis. One constitutive “pathway” for this protein involves its intracellular degradation; incompletely assembled or misfolded mmebrane immunoglobulins are probably being constantly directed into this degradatory pathway. The site of degradation is a preGolgi compartment which may the ER itself. The second fate of membrane immunoglobulin is to be rescued from the intracellular degradative pathway, presumably by virtue of being properly and completely assembled. What constitutes complete and proper assembly of membrane immunoglobulin? Since intriguing correlations have been made between the acquisition of hydrophobicity and the cell surface transport of IgM, is there an underlying theme, possibly of heuristic value, which connects assembly, hydrophobicity, and cell surface transport? It has long been appreciated that the mere association of pm and K is insufficient for the surface expression of p. Indeed, from studies of Type I1 pre-B cells which express p on the cell surface in association with surrogate L chains, it has been clear that expression of the K L chain was not the key event leading to the surface expression of p. In plasma cells which secrete ps in association with conventional L chains, pm was associated with K , but was not transported to the cell surface (Sitia et al., 1987; Hombach et al., 1988a). Indeed, in a pre-B cell line expressing the y iso-

IMMUNOGLOBULIN TRANSPORT

23

type and a transfected K L-chain gene, ys2-~2tetramers were secreted, while ym2-~2 tetramers remained intracellular (Bachhawat and Pillai, 1991). All these results suggested that an additional posttranslational event or protein may be necessary for the surface transport of membrane immunoglobulin. In studies of the biosynthesis and fate of pm in pre-B cells and in B cells, we demonstrated that, in B cells, three distinct forms of pm may be identified (PiUai and Baltimore, 1987b). The initially synthesized pm protein is relatively hydrophilic (and partitions into the aqueous phase upon fractionation with Triton X-114). This relative hydrophilicity is perhaps not too surprising when one considers that the transmembrane region of pm contains nine serine and threonine residues. The hydrophilic form is referred to as the pml form. After a short while, this protein is converted to a relatively hydrophobic form, referred to as the pm2 form. In B cells, the pm2 form acquires endo H resistance prior to conversion to a terminally glycosylated slower migrating pm3 form, which is transported to the cell surface. Since the acquisition of a relatively long-lived hydrophobic form of pm correlates with the transport of this protein to the cell surface in B cells, it seemed likely that understanding the basis for the acquisition of hydrophobicity might yield information relevant to the mechanism by which pm is transported to the cell surface. The acquisition of relative hydrophobicity by pm postsynthetically is probably relevant to the conformation of the fully assembled membrane IgM molecule that is recognized for transport, and is not in any way a requirement for proper anchoring in the ER membrane (Pillai and Baltimore, 1987b). The relevance of the relative hydrophobicity of the pm molecule to the process of cell surface transport was dramatically highlighted in a study on the transport of wild-type and mutant IgM proteins in nonlymphoid cells (Williams et al., 1990). In non-B cells, transfection of a gene encoding the wild-type membrane p H-chain protein and a A L-chain gene leads to the formation of intracellular membrane IgM tetramers which are not transported to the cell surface. However, when a hydrophilic stretch of five amino acids in the anchor region of the pm gene was replaced by nucleotides encoding hydrophobic amino acids, transfection experiments on fibroblasts revealed that this relatively hydrophobic pm protein could be transported to the cell surface in association with L-chain. From these studies, it seems likely that, whatever the requirements for surface transport of IgM entail, in terms of either associated proteins or posttranslational modifications or both, the net effect of these assembly-related events may well be mimicked by the artificially acquired hydrophobicity of the pm protein. The hydrophilic TTAST stretch may be viewed as a topogenic sequence that targets pm for retention, and posttranslational

24

SHIV PILLAI

processes such as assembly with other hydrophobic proteins may “mask” this region and permit transport to the cell surface. In the past few years, a number of proteins have been described in pre-B and B cells that are associated with membrane immunoglobulin. On the surface of Type I1 pre-B cells, p m is known to be associated with the o and L surrogate L-chain proteins. These surrogate L chains have been described in considerable detail in Section IIC. A functional role of these proteins in permitting the transport of pm to the cell surface has been suggested in transfection experiments. Transfection of the A5 and vPreB genes along with a modified p m gene (incorporating transmembrane and cytoplasmic tail structures from a class I gene) led to the expression of p and surrogate L chains on the surface of a recipient myeloma cell line (Tsubata and Reth, 1990). In addition to w and L , up to six other polypeptides have been reported to associated with p m in Type I1 pre-B cell lines (Pillai and Baltimore, 1988; Takemori et al., 1990). Two of these proteins may correspond to the mb-l/Iga and B29/Igp proteins that have been studied in some detail in B cells and are described here. The other four have been characterized in a limited fashion, and their role in the transport of pm remains to be established. The use of digitonin as a lysis buffer and two-dimensionalnonreducing/ reducing gel systems for analysis has led to the characterization of two proteins associated with membrane IgM in B cells (Hombach et al., 1988b, 1990a).These proteins are believed to form a disulfide-linked heterodimer that is associated with membrane IgM; the exact stoichiometry of this complex and the proportion that is actually disulfide bonded remain to be determined with certainty. The first of these proteins, originally described as B34, is now known as IgM(a). This protein is a 34-kDa protein and has been confirmed to be the product of the mb-Z gene (Hombach et al., 1990a; Sakaguchi et af., 1988). The ability of subclones of a myeloma line to transport p m to the cell surface was correlated directly with the presence of this protein. Introduction of the mb-1 gene into a cell line that initially lacked the ability to transport IgM to the cell surface led to the establishment of a transfectant cell line which expressed IgM(a)protein, and which was now competent to transport pm to the cell surface. A second protein that was found to be associated with membrane IgM in B cells and which is at least in part disulfide linked to IgM(a) is a 39-kDa protein termed Ig(p). This protein, unlike IgM(a) (which is specifically associated with the p isotype), is presumed to be associated with membrane immunoglobulin of all isotypes. Ig(p) has recently been demonstrated to be the product of the B lineage-specific B29 gene (Hombach et al., 1990b; Hermanson et al., 1988).Overexpressionof the B29 gene in a myeloma line led to an increase in the surface expression of IgM. Both mb-1 and B29 are expressed from the earliest pro-B stage of

IMMUNOGLOBULIN TRANSPORT

25

ontogeny and by all surface immunoglobulin-positive lymphocytes. Structurally, the cDNA sequences of both these genes suggest that they encode transmembrane glycoproteins. Both genes encode signal peptides, extracellular domains with N-glycosylation sites, transmembrane anchor regions, and cytoplasmic tails which bear some homology to one another as well as to the cytoplasmic tails of the T cell receptor-related CD3 y chain, suggesting that these proteins may be involved in signal transduction through membrane IgM. Indeed, both these proteins have been demonstrated to be N-glycosylated and to be phosphorylated on tyrosine residues in response to crosslinking of membrane immunoglobulin (Campbell and Cambier, 1990). A separate protein associated with membrane IgD expressed in cell lines which have lost mb-1 and the ability to transport surface IgM has been described. This 35-kDa protein is presumed to be the 6 isotype-specific equivalent of mb-1 (Wienands et al., 1990). This protein is structurally related to the mb-1 product, but must be predicted to be the product of a distinct gene (since it has been identified in clones that lack mb-1 transcripts). The probable existence of a y isotype-specific “mb-1”-like protein has been postulated based on the inability of ym2-~2 tetramers to be transported to the cell surface in a transfected pre-B cell line (Bachhawat and Pillai, 1991). The acquisition of the ability to transport membrane immunoglobulins of various isotypes during development and its correlation with isotype-specific associated proteins remains a poorly explored avenue. Many issues remain to be resolved regarding the role of the products of the mb-1 and B29 genes in transporting membrane IgM to the cell surface. Although it is tempting to draw analogies with the role of the CD3 complex proteins in the transport of the T cell receptor to the cell surface, certain differences need to be addressed. The product of the mb-2 gene (unlike CD3 chains) can apparently be expressed on the surface of some cells in the absence of membrane IgM. Both mb-1 and B29 are expressed at levels comparable to those seen in B cells in Type I pre-B cell lines that express only intracellular p. It is likely that other IgM-associated proteins may exist in the B cell stage that may play a role in transporting membrane IgM to the cell surface; these proteins remain to be identified. A “minimal” model for the structure of the B cell receptor is provided in Fig. 6. An interesting suggestion has been made regarding the possible role of assembly of IgM with mb-1 (this suggestion could apply equally well to B29 or any other IgM-associated protein) in assisting transport to the cell surface. Since, in nonlymphoid cells, replacement of five hydrophilic amino acids in the transmembrane region of the H chain with hydrophoSee Note Added in Proof, p. 36.

26

SHIV PILLAI

gatively charged loop

MEMBRANE

K

K

FIG. 6. A “minimal” model for the B cell receptor.

bic residues is sufficient to make intracellular IgM competent for surface transport, it was suggested that mb-1 may, by associating with the H chain, mask the hydrophilic stretches in the transmembrane region, and the ensuing hydrophobicity is sufficient for ensuring transport to the cell surface (Williams et al., 1990). The possibility that a posttranslational modification such as the myristylation of IgM may play a role in the acquisition of hydrophobicity in the process of assembly and surface transport remains to be excluded (Pillai and Baltimore, 1987b). From the studies of membrane immunoglobulin discussed here as well as from interesting studies of the degradation and transport of the T cell receptor-CD3 complex (Bonifacino et al., 1990), an interesting concept has emerged that may apply generally to the assembly of transmembrane heterooligomers. Hydrophilic stretches in the transmembrane segments of individual chains of multisubunit membrane proteins are probably involved in the recognition of partners and in the assembly process. The process of assembly leads to the masking of the hydrophilic transmembrane stretches and prevents rapid degradation. The process may be regulated by the need to attain a threshold level of hydrophobicity that permits transport to proceed; alternatively, being below this threshold, hydropho-

IMMUNOGLOBULIN TRANSPORT

27

bicity may serve as a signal for ER retention or degradation, thus abrogating the transport of incompletely assembled oligomers. The “packing” process could conceivably be catalyzed by posttranslational processes such as acylation (Pillai and Baltimore, 1987b). There is evidence (B. J. Cherayil and S. Pillai, unpublished observations) suggesting that the ratelimiting step at the pre-B to B transition which drives assembly and transport may indeed be the efficiency of the process of covalent heavy-light assembly, which may be a prerequisite for engaging transmembrane masking proteins such as mb-1 and B29. C. MEMBRANE IMMUNOGLOBULIN RECYCLING, ENDOCYTOSIS, AND DOWN-REGULATION Apart from its ability to be transported to the cell surface in B cells, the membrane immunoglobulin molecule or an associated protein must carry structural information permitting endocytosis, recycling, and receptor down-regulation functions. When the antigen receptor on B cells encounters a cognate protein antigen, the antigen is rapidly internalized into an endocytic compartment. In order for antigen presentation to occur, the contents of this compartment must be accessible to processing proteases and must intersect with vesicles containing class I1 molecules. The internalized protein is then cleaved into peptides, and the appropriate peptide then associates with the antigen-binding groove on the class I1 molecule and is presented on the surface of the cell. Membrane immunoglobulin is then presumably recycled to the cell surface. Ligation of membrane immunoglobulin by a multivalent antigen (as mimicked by antiimmunoglobulin crosslinking) leads to internalization by nonselective endocytosis (Guagliardi et al., 1990). Monovalent protein antigens are internalized into clathrin-coated vesicles (Watts et al., 1989). However, ligands taken up by selective or nonselective endocytosis are known to be delivered to the same endosomes (Tran et al., 1987). The transmembrane region and the cytoplasmic tail of each membrane immunoglobulin H-chain isotype are highly conserved across species. The function of membrane immunoglobulin in terms of signal transduction as well as for the endocytic functions that are essential for antigen presentation is presumed to be mediated in part via the proteins that are associated with pm (including the products of the mb-1 and B29 genes). The cytoplasmic tail of pm contains only three amino acids; the mb-1 and B29 gene products have considerably longer cytoplasmic tails which contain conserved cytoplasmic residues known to be phosphorylated in response to activation. Presumably, ligation of membrane immunoglobulin leads to the appro-

28

SHIV PILLAI

priate conformational changes/phosphorylation events that regulate the generation of a differentiation signal, as well as the internalization and recycling of the receptor itself. Attempts have been made to dissociate the signal transduction and antigen presentation functions of membrane immunoglobulin by making mutations of the transmembrane region and cytoplasmic tail regions of the pm gene, followed by transfection and functional analysis (Webb et al., 1989; Shaw et al., 1990). Replacement of the transmembrane region of pm with a class I1 transmembrane anchor, not surprisingly, abolished the ability of this protein to transduce a signal. Deletion of the KVK cytoplasmic tail severely compromised both signal transduction and antigen presentation functions. The hydroxyl group of the tyrosine residue (position 587) and the adjacent serine residue (position 588) were demonstrated to be of critical importance in signal transduction (Shaw et al., 1990). Interestingly, conversion of Tyr-587 alone to a phenylalanine resulted in a molecule that was able to signal, but which lacks the ability to present antigen. This dissociation of signal transduction from antigen presentation suggests that different protein contacts are required for these two processes and that different pm-associated proteins may help mediate these functions. In Fig. 7, a schematic view of the last 41 amino acids of the pm protein is provided, indicating residues which, based on the analysis of mutations in this region, are known to be of functional importance. Closely related to the functions of membrane immunoglobulin just discussed is the phenomenon of receptor down-regulation, which may be an important mechanism for the generation of clonal anergy. In “double” transgenic mice expressing hen egg white lysozyme as well as the rearranged immunoglobulin genes which recognize lysozyme, a feature of anergic B lymphocytes was the expression of relatively high antigenspecific IgD levels in conjunction with low surface IgM levels (Goodnow et al., 1990). A model for anergy in these cells invokes the specific downregulation of surface IgM, possibly as a result of exposure to monomeric self antigen in the absence of cognate T cell help. Since different proteins have been found to be associated with IgM and IgD, selective posttranslational changes in one isotype-specific protein may be invoked to explain such a specific down-regulation event. It is indeed conceivable that even in memory B cells (which may express other isotypes, including IgG, IgE, and IgA) similar down-regulation mechanisms may play a role in the maintenance of anergy . The role of isot ype-specific membrane immunoglobulinassociated proteins and of specific transmembrane residues on the immunoglobulin molecule and in these processes may be predicted, but remain to be established.

IMMUNOGLOBULIN TRANSPORT

29

E

(-)

E (-) V

N A E

G

E

(-1

(-1 (4

F

EXTRACELLULAR

(-) E N

L W *Fl RetentioniDegradation signal

LFL

L

MEMBRANE

SL

s Y7587) T~

(+I

K (595) (596) (+) K (597)

v

T

(588) INTRACELLULAR

FIG. 7. Membrane exon-encoded residues of the pm protein depicting residues relevant to function and transport. For details, see text.

D. MEMBRANE IMMUNOGLOBULIN RETENTIONIN PLASMA CELLS Plasma cells presumably do not need to respond to antigen and need not express membrane immunoglobulin on the cell surface. This issue has not, however, been established with certainty, and few studies have been performed using actual plasma cells. In many myeloma lines, membrane immunoglobulin is retained intracellularly (Hombach et al., 1988a; Sitia et al., 1987). In some lines, this loss of expression can be explained by the loss of mb-1 expression. In situ hybridization of mb-1 and B29 genes or immunohistochemical studies using antibodies for these proteins must be performed on tissue sections to confirm whether these genes are actually shut off in plasma cells in the course of B cell development. V. Immunoglobulin Secretion in Plasma Cells

Although numerous studies have been published on the secretion of immunoglobulins by plasma cells, many unanswered questions remain regarding the details of this process. In brief, no vital role in the process of

30

SHIV PILLAI

transport itself can be ascribed to posttranslational modifications of immunoglobulin, to the process of assembly into higher oligomers (for IgM and IgA), or to the J chain. An absolute requirement does exist for association with an L chain and for the heavy-light complex to be “conformationally appropriate” for transport. OF A. ROLEOF THE J CHAINAND ASSEMBLY POLYMERIC IMMUNOGLOBULINS

Prior to the establishment of the tailpiece-dependent retention mechanism of secretory IgM in early B cells, a popular model for IgM secretion invoked the need for assembly of J chain-containing polymeric IgM prior to secretion. The immunoglobulin J chain (Koshland, 1985) is a 15-kDa protein associated with the cysteine-containing tailpiece in IgM pentamers and in IgA dimers. The J chain gene is transcriptionally activated in the course of antigen- and lymphokine-induced activation of B lymphocytes. The J chain is essential neither for the oligomerization of IgM monomer units nor for IgM secretion. In a transfected glioma cell line, IgM assembles to form pentamers and hexamers in the absence of J chain and is secreted (Cattaneo and Neuberger, 1987). Indeed, assembly of IgM and IgA into polymeric forms is not a prerequisite for secretion. Monomeric IgA is secreted efficiently and so is a mutant IgM that is incapable of forming pentamers (Baker et al., 1986). The probable role of the J chain lies in the recognition of assembled immunoglobulins by the polyimmunoglobulin receptor prior to transcytosis. The issue of IgM assembly has been reviewed in depth (Davis and Shulman, 1989). AND SECRETION B. POSTTRANSLATIONAL MODIFICATIONS

Numerous posttranslational modifications of immunoglobulins ranging from N-linked and 0-linked glycosylation to tyrosine sulfation have been examined from the viewpoint of secretion (reviewed by Wall and Kuehl, 1983). No known posttranslational modification is essential for the secretory process. C. ROLEOF L CHAINSAND “APPROPRIATE” CONFORMATION FOR SECRETION Immunoglobulin L chains play a crucial role in immunoglobulin secretion. Indeed, it is likely that a small conserved region on the VL domain may serve as a “recognition patch” for the secretory machinery. Studies of the role of L chain in immunoglobulin secretion support, albeit in a

IMMUNOGLOBULIN TRANSPORT

31

limited way, the notion that secretion is not merely a “bulk flow” process for nonretained nonanchored proteins that enter the ER. H-chain proteins in the absence of L chains are retained intracellularly in association with Bip. In an H-chain-only myeloma cell line (Pepe el al., 1986), as well as in a ys-containing pre-B cell line (Bachhawat and Pillai, 1991), introduction of an L-chain gene leads to efficient secretion of fully assembled immunoglobulin. L-chain proteins in the absence of H chains are secreted. A notion that is currently gaining ground is that L chains may possess a steric determinant that is crucial for the secretion of free L chains as well as heavy-light tetramers. L chains with point mutations have been identified that are not secreted (Mosmann and Williamson, 1980; Wu et al., 1983; Nakaki et al., 1989; Dul and Argon, 1990). Nonsecreted L-chain proteins have also been found to be associated intracellularly with Bip. Mutations in both the VL and CL domains have been identified that prevent secretion. Dul and Argon (1990) tested the hypothesis that a group of conserved residues (residues 57-65) on L-chain proteins may form part of a structural patch that is recognized by the intracellular secretory machinery. Conversion of a phenylalanine at position 62 to a serine resulted in a nonsecreted L chain. This mutant L chain bound Bip. In the presence of H-chain protein, however, this L chain was assembled into a functional antigenbinding antibody. This antibody molecule did not bind Bip and was recognized by a panel of monoclonal and polyclonal antibodies against wild-type L chains, indicating that no gross conformational alteration had occurred as a result of the point mutation. The assembled and functional antibodies containing a serine residue at position 62, however, were not secreted. These data suggest that structural recognition of conserved motifs may play a role in the progress of the antibody molecule along the secretory pathway. Figure 8 outlines retention and transport signals that influence immunoglobulin secretion. M. Summary: Choices between RetentiodDegradation and Transport of Immunoglobulins Are Dictated by Function

In pre-B lymphocytes, secretory immunoglobulin has no function. At this stage, ps is retained intracellularly. Two retention signals have been identified, one in the CH1 domain and the other in the CH4 cysteine tailpiece. The retention mechanism may involve association with Bip or possibly with an ER protein or proteins that sequester luminal proteins that contain free cysteines. At this stage, pm is associated with the o and L proteins and is degraded intracellularly. The p 2 - ~ 2 - ~complex 2 (in associ-

32

SHIV PILLAI chain

Recognition patch on L chain for secretion

-

Putative Bip binding sites or retention signals

Tailpiece

Ht

/

FIG. 8. Schematic view of retention and transport signals on secretory immunoglobulins. The domain structure of the heavy chain depicted is typical of hs. L chain, light chain; H chain, heavy chain; V, variable region; VL, variable region of the light chain; CL, constant region of the light chain; SH, sulthydryl.

ation with other proteins) is presumed to be involved in generating a signal for further differentiation. Intracellular degradation of pm in early (Type I) pre-B cells probably involves recognition of a hydrophilic signal in the transmembrane region of pm. Late in the pre-B stage in Type I1 pre-B cells, pm in association with surrogate L chains is transported to the cell surface. Cell surface transport probably depends on efficient covalent pm-o assembly and also the posttranslational acquisition of hydrophobicity. The mb-1/IgMa and B29/Igp proteins may serve to mask the hydrophilic transmembrane residues in pm which target it for ER degradation. In nonactivatedhirgin B cells, secretory ps2-~2tetramers are retained intracellularly by a mechanism involving the cysteine tailpiece on ps and association with Bip. Bip does not associate with tetrameric membrane immunoglobulins (Sitia et al., 1990). Membrane immunoglobulin is transported to the cell surface in association with IgM(a) (which is specific for

IMMUNOGLOBULIN TRANSPORT

33

the p isotype) and Ig(p), which presumably associates with membrane immunoglobulins of all isotypes. At this stage, membrane immunoglobulin functions as the antigen receptor which is involved both in signal transduction and endocytosis (for antigen presentation). Mutations in the transmembrane domain can dissociate signal transduction from endocytosis and suggest that the endocytic pathway can be regulated by pm-associated proteins. Receptor down-regulation may serve to sustain a state of clonal anergy . After B cells are activated by antigen, class switching may occur. Isotype-specific associated proteins may play a role in the surface transport of various membrane immunoglobulin classes. Assembly processes help ps and as to bypass retention mechanisms. The y and E isotypes lack a cysteine tailpiece and are secreted in association with L chains. At this stage, polymerization masks the cysteine tailpiece of ps and a s and retention is abrogated. While considering transport along the secretory pathway, two broad mechanisms have been proposed. One model suggests that, after proteins are translocated into the ER, many categories of retention and transport signals are recognized by receptors which direct vesicles to appropriate locations. Mannose 6-phosphate is a tag for lysosomal transport, and the KDEL signal ensures that ER luminal proteins do not go beyond the salvage compartment. In such a model, proteins destined for the cell surface or for secretion have specific steric features recognized by receptors which guide vesicular transport. The second model is the bulk flow model (Rothman, 1987), which suggests that once a protein enters the ER it will be transported to the cell surface by “bulk flow,” unless retention signals or lysosomal transport signals subvert this process. This model suggests that no receptors are necessary to direct vesicular transport to the plasma membrane. A separate concept particularly relevant to immunoglobulin transport and which is compatible with both models above is the need for oligomerization and assembly as a prerequisite for cell surface transport (Kreis and Lodish, 1986). An “exposed” interface prior to assembly could function as a retention signal which requires masking by the process of assembly. Alternatively, the fully assembled complex may provide the correct stereochemical configuration for further transport. It remains unclear whether transport of membrane and secretory immunoglobulins merely reflects the abrogation of retentioddegradation mechanisms or actually depends in any way on specific steric recognition of domains on immunoglobulin molecules for movement to occur. The recent demonstration of a patch on L-chain molecules which is essential for immunoglobulin movement suggests a possible role for steric recognition events, as opposed to bulk flow, in the absence of retention.

34

SHIV PILLAI

ACKNOWLEDGMENTS I thank David Baltimore for introducing me to pre-B lymphocytes, and Marian Koshland, Bobby Cherayil, Anand Bachhawat, and Sudhir Krishna for many invaluable discussions. I also thank Ravi Iyer for his help with the figures and Michael Reth for communicatingresults prior to publication. This work was supported by National Institutes of Health grant A1 27835 and by the Arthritis Foundation.

REFERENCES Alberini, C. M., Bet, P., Milstein, C., and Sitia, R. (1990). Nature (London)347,485-488. Alt, F. W., Blackwell, T. K., Depinho, R. A., Reth, M., and Yancopoulos, G. D. (1986). Immunol. Rev. 89,5-63. Bachhawat, A-.K., and Pillai, S. (1991). J . Cell Biol. (in press). Baker, M. D., Wu, G. E., Toone, W. M., Murialdo, H., Davis, A. C., and Shulman, M. J. (1986). J . Immunol. 137, 1724-1728. Bole, D. G., Hendershot, L. M., and Kearney, J. F. (1986). J . Cell Biol. 102, 1558-1566. Bonifacino, J. S., Cosson, P., and Klausner, R. D. (1990). Cell 63,503-513. Burrows, P. D., Kearney, J. F., Lawton, A. R., and Cooper, M. D. (1978).J. Immunol. 120, 1526-153 I . Campbell, K. S., and Cambier, J. C. (1990). EMBOJ. 9,441-448. Cattaneo, A., and Neuberger, M. S. (1987). EMBO J . 6,2753-2758. Chang, H., Dmitrovsky, E., Hieter, P. A., Mitchell, K., Leder, P., Turoczi, L., Kirsch, I. R., and Hollis, G. F. (1986). J. Exp. Med. 163,425-435. Cherayil, B. J., and Pillai, S. (1991a). J . Exp. Med. 173, 111-116. Cherayil, B. J., and Pillai, S. (1991b). Manuscript in preparation. Davis, A. C., and Shulman, M. J. (1989). Immunol. Today 10, 118-128. Dul, J. L.,and Argon, Y. (1990). Proc. Natl. Acad. Sci. U.S.A. 87,8135-8139. Dulis, B. H. (1983). J. Biol. Chem. 258,2181-2187. Dulis, B. H., Kloppel, T. M., Grey, H. M., and Kubo, R. T. (1982). J. Biol. Chem. 257 4369-4374. Findley, H. W.,Cooper, M. D., Kim, J. H., Alvarado, C., and Ragab, A. H. (1982). Blood 60, 1305- 1309. Flynn, G. C., Chappell, T. G., and Rothman, J. E. (1989). Science 245,385-390. Gething, M.-J., McCammon, K., and Sambrook, J. (1986). Cell 46,939-950. Goodnow, C. G., Adelstein, S., and Basten, A. (1990). Science 248, 1373-1379. Gordon, J., Hamblin, T. J., Smith, J. L., Stevenson, F. K., and Stevenson, G. T. (1981). Blood 58,552-556. Gu,H., Kitamura, D., and Rajewsky, K. (1991). Cell 65,47-53. Guagliardi, L. E., Koppelman, B., Blum, J. S., Marks, M. S., Cresswell, P., and Brodsky, F. M . (1990). Nature (London) 343, 133-139. Haas, I. G., and Wabl, M. (1983). Nature (London)306,387-389. Hardy, R. R., Dangl, J. R., Hayakawa, K., Jager, G., Herzenberg, L. A., and Herzenberg, L. A. (1986). Proc. Natl. Acad. Sci. U.S.A. 83, 1438-1442. Hendershot, L. M. (1990). J. Cell Biol. 111,829-837. Hendershot, L., and Levitt, D. (1984). J. Immunol. 132,502-509. Hendershot, L. M., Bole, D., Kohler, G., and Kearney, J. F. (1987). J . Cell Biol. 104, I61-767. Hermanson, G., Eisenberg, D., Kincade, P., and Wall, R. (1988). Proc. Natl. Acad. Sci. U.S.A. 85,6890-6894.

IMMUNOGLOBULIN TRANSPORT

35

Hollis, G. F., Evans, R. F., Stafford-Hollis, J. M.,Korsmeyer, S. J., and McKearn, J. P. (1989). Proc. Natl. Acad. Sci. U . S . A . 86,5552-5556. Hombach, J., Sablitzky, F., Rajewsky, K., and Reth, M. (1988a). J. Exp. Med. 167,652-657. Hombach, J., Leclercq, L., Radbruch, A., Rajewsky, K., and Reth, M. (1988b).EMBO J . 7, 345 1-3456. Hombach, J., Tsubata, T., Leclercq, L., Stappert, H., and Reth, M. (1990a). Nature (London) 343,760-762. Hombach, J., Lottspeich, F., and Reth, M. (1990b). Eur. J. Immunol. 20,2795-2799. Karasuyama, H., Kudo, A., and Melchers, F. (1990). J . Exp. Med. 172, %9-972. Kassenbrock, C. K., Garcia, P. D., Walter, P., and Kelley, R. B. (1988). Nature (London) 333,90-93. Kerr, W. G . , Cooper, M. D., Feng, L., Burrows, P. D., and Hendershot, L. M. (1989). Int. Immunol. 4,355-361, Koshland, M. E. (1985). Annu. Rev. Immunol. 3,425-452. Kreis, T. E., and Lodish, H. F. (1986). Cell 46,929-937. Kudo, A., and Melchers, F.(1987). EMBO J . 6,2267-2272. Lewis, M. J., Sweet, D. J., and Pelham, H. R. B. (1990). CeN61, 1359-1363. Mason, J. O., Williams, G., and Neuberger, M. S. (1988). Genes Deu. 2, 1003-1011. McGarigle, M., Krishna, S., and Pillai, S. (1991). Manuscript in preparation. Misener, V., Jongstra-Bilen, J., Young, A. J., Atkinson, M. J., Wu, G. E., and Jongstra, J. (1990). J . Immunol. 145,905-909. Morrison, S. L., and Scharff, M.D. (1975). J . Immunol. 114,655-659. Mosmann, T. R., and Williamson, A. R. (1980). Cell 20,283-292. Munro, S . , and Pelham, H. R. B. (1986). Cell 46,291-300. Munro, S . , and Pelham, H. R. B. (1987). Cell 48,899-907. Nakaki, T.,Deans, R. J., and Lee, A. S. (1989). Mol. Cell Biol. 9,2233-2238. Nussenzweig, M.C., Shaw, A. C., Sinn, E., Danner, D. B., Holmes, K. L., Morse, H. C., and Leder, P. (1987). Science 236,816-819. Oettinger, M.A., Schatz, D. G., Gorka, G., and Baltimore, D. (1990). Science 248, 15171523. Paige, C. J., Kincade, P. W., and Ralph, P. (1981). Nature (London)292,631-633. Pepe, V. H., Sonenshein, G. E., Yoshimura, M. I., and Shulman, M. J. (1986). J. Immunol. l37,2367-2372. Pillai, S., and Baltimore, D. (1987a). Nature (London) 329, 172-174. P h i , S., and Baltimore, D. (1987b). Proc. Natl. Acad. Sci. U.S.A. 84,7654-7658. Pillai, S., and Baltimore, D. (1988). Curr. Top. Microbiol. Immunol. 137, 136-139. Rajewsky, K., Forster, I., and Cumano, A. (1987). Science 238, 1088-1093. Reth, M. G., and Alt, F. W. (1984). Nature (London) 312,418-423. Reth, M. G., Petrac, E., Wiese, P. Lobel, L., and Alt, F. W. (1987). EMBO J . 4,361-366. Reth, M., Hombach, J., Wienands, J., Campbell, K. S., Chien, N., andcambier, J. C. (1991). Immunol. Today 12, 196-201. Rose, M. D., Misra, L. M., and Vogel, J. P. (1989). Cell 57, 1211-1221. Roth, R. A., and Koshland, M. E. (1981a). Biochemistry 20,6594-6599. Rothman, J. E. (1987). Cell 50,521-522. Rubartelli, A., Sitia, R., Zicca, A., Grossi, C. E., and Ferrarini, M. (1983). Blood 62, 495-504. Sakaguchi, N., and Melchers, F. (1986). Nature (London) 324,579-582. Sakaguchi, N., Kawashimura, S., Kimoto, M., Thalmann, P., and Melchers, F. (1988). EMBO J . 7,3457-3464. Schatz, D. G., Oettinger, M. A., and Baltimore, D. (1989). Cell 59, 1035-1048.

36

SHIV PILLAI

Sen, R., and Baltimore, D. (1986). Cell 47,921-928. Shaw, A. C., Mitchell, R. N., Weaver, Y.K., Campos-Torres, J., Abbas, A. K., and Leder, P. (1990). Cell 63,381-392. Sibley, C. H., Ewald, S. J., Kehry, M. R., Douglas, R. H., Raschke, W. C., and Hood, L. E. (1980). J . fmmunol. 125,2097-2105. Sidman, C. (1981). Cell 23,379-389. Sitia, R., Rubartelli, A., Kikutani, H., Hammerling, U., and Stavnezer, J. (1985). J. fmmunol. 135,2859-2864. Sitia, R., Neuberger, M. S., and Milstein, C. (1987). EMBO J . 6, 3969-3977. Sitia, R. Neuberger, M., Alberini, C., Bet, P., Fra, A., Valetti, C., Williams, G., and Milstein, C. (1990). Cell 60,681-690. Takemori, T., Mizuguchi, I. M., Miyazoe, I., Nakanishi, M., Shigemoto, K., Kimoto, H., Shirsawa, T., Maruyama, N., and Taniguchi, M. (1990). EMBO J . 9,2493-2500. Thorens, B., Schulz, M.-F., and Vassali, P. (1985). EMBO J . 4, 361-368. Tran, D., Carpentier, J. L., Sawano, F., Gorden, P., and Orci, L. (1987). Proc. Natl. Acad. Sci. U.S.A. 84,7957-7961. Tsubata, T., and Reth, M. (1990). J . Exp. Med. 172,973-976. Tsubata, T., Tsubata, R., and Reth, M. (1991). Eur. J . fmmunol.21, 1359-1363. Vassali, P., Tartakoff, A., Pink, J. R. L., and Jaton, J. C. (1980). J. Biol. Chem. 255, 11822-11827. Vogel, J. P., Misra, L. M., and Rose, M. D. (1990). J . Cell Biol. 110, 1885-1895. Wall, R., and Kuehl, M. (1983). Annu. Rev. fmmunol. 1,393-422. Watts, C., West, M. A., Reid, P. A., and Davidson, H. W. (1989). Coldspring HarborSymp. Quant. Biol. 54,345-352. Webb, C. F., Nakai, C., and Tucker, P. W. (1989). Proc. Natl. Acad. Sci. U . S . A . 86, 1977- 198 1. Wienands, J., Hombach, J., Radbruch, A., Riesterer, C., and Reth, M. (1990). EMBO J. 9, 449-455. Williams, G. T., Venkitaraman, A. R., Gilmore, D. J., and Neuberger, M. S. (1990). J . Exp. Med. 171,947-952. Wu, G. E., Hozumi, N., and Murialdo, H. (1983). Cell 33,77-83. Yancopoulos, G. D., and Alt, F. W. (1986). Annu. Rev. Immunol. 4,339-368. NOTEADDEDI N PROOF.Since the submission of this review the work of Rajewsky and colleagues (Gu er al., 1991) suggests that the D p protein serves as a negative signal, eliminating early pre-B cells in which a particular reading frame has been generated by D-J rearrangement at the H-chain locus. Some evidence has also been obtained for low levels of D p protein on the surface of pre-B cells in association with surrogate light chains (Tsubata et al., 1991). Evidence has also been obtained indicating that alternatively glycosylated forms of the mb-1 protein associate with IgM and IgD, indicating that only a single Ig(a) protein may exist (unpublished observations cited in Reth et al., 1991).

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 130

The Cytoskeletal System of Nucleated Erythrocytes WILLIAMD. COHEN Department of Biological Sciences, Hunter College of CUNY, New York, New York 10021; and The Marine Biological Laboratory, Woods Hole, Massachusetts 02543

I. Introduction

The blood of all vertebrates other than mammals contains nucleated erythrocytes throughout life, with rare exception. Indeed, this is one of the simple universal characteristics distinguishing mammals from nonmammals. Thus, the erythrocytes of fish, amphibians, reptiles, and birds are typically flattened ellipsoids, with biconvexity produced by a bulging nucleus (Ramon-Cajal, 1933; Andrew, 1965; Rowley and Ratcliffe, 1988) (Fig. 1). In contrast, essentially all erythrocytes in the blood of adult monotremes, marsupials, and placental mammals are anucleate, and they assume the familiar flattened biconcave discoid shape under static (nonflow) conditions (Briggs, 1936; Ralston, 1985). However, during early embryonic development, even mammalian blood is populated by nucleated erythrocytes (Dantschakoff, 1908; Maximow, 1909; Yadav, 1972; Block, 1964). These constitute the “primitive generation” that originates in the so-called “blood islands” of the yolk sac, in common with the first generation of erythrocytes in all other vertebrates. In addition, various invertebrates, including blood clams (Mollusca), certain sea cucumbers (Echinodermata), and marine worms or wormlike species representing several phyla (Annelida, Sipuncula, and Priapulida), also package respiratory proteins within nucleated erythrocytes (Cohen and Nemhauser, 1985). Beyond interest in erythrocytes per se, there are good reasons for studying the cytoskeletal system of nucleated erythrocytes. First, these are cells specialized with respect to the generation and maintenance of a particular shape, and in their mechanical responses to deformation. As such, they are likely to have a highly differentiated cytoskeletal system displaying few extraneous properties. Second, they are readily available in quantity and in pure populations with only minor variation of cell shape or size for a given species, the one major variable being age. Third, their cytoskeletal system is highly compartmentalized compared to other cell types, simplifying interpretation of data and permitting at least some de31

Copyright 6 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

38

WILLIAM D. COHEN

FIG. 1. Erythrocytes of the frog Runa pipiens. (a) Edge and face views; (b) oblique view illustrating nuclear bulge. Scanning electron microscopy of glutaraldehyde-fixed critical point-dried cells.

gree of cytoskeletal fractionation. Fourth, because erythrocytes exist naturally as individual cells in a fluid tissue, their external morphology and internal structure are maintained independently of contact with other cells, and their cytoskeletal system can be considered functionally complete. Fifth, their distinctive universal shape facilitates an experimental approach to questions about the relationship between cellular morphogenesis and biogenesis of the cytoskeleton during cell differentiation. For these reasons, the past decade has seen the exploitation of nucleated erythrocytes as a model system for a variety of studies on cytoskeletal structure, function, and biogenesis. Such work represents a natural step toward analysis of a cytoskeletal system at the next level of complexity beyond that of the mammalian erythrocyte, that is, one with greater resemblance to that of eukaryotic cells in general. In this status report on our understanding of the nucleated erythrocyte cytoskeletal system, coverage is not exhaustive. The focus is on cytoskeleton formation and function, particularly that of the marginal band (MB) of microtubules (MTs). Although much of this chapter is concerned with cytological, ultrastructural, and molecular observations on nonmammalian erythrocytes, recent comparative work on the cytoskeleton of primitive nucleated erythrocytes in developing mammals is also discussed. In pointing out areas of ignorance or disagreement throughout, I have taken the liberty of speculating occasionally in the hope that this will stimulate further investigation.

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

39

11. Nucleated Erythrocytes: A Phylogenetic and Physiological Portrait

In the gallery of cells presented diagrammatically in Fig. 2, nucleated erythrocytes of five nonmammalian vertebrate classes are represented: cartilaginous fish, bony fish, amphibians, reptiles, and birds (Fig. 2, cells a-g). Although cell size is fairly uniform for a given species, among different species there is considerable size variation (Ram6n-Caja1, 1933; Andrew, 1965; Goniakowska-Witalidska and Witalitiski, 1976). The Amphibia typically have relatively large erythrocytes (Fig. 2, cells a-c), those of the giant salamanderAmphiuma tipping the vertebrate scale with a long axis of 70 pm. At the other extreme are erythrocytes of most birds and bony fish (Fig. 2, cells f and g), usually 15 p m or less. In an intermediate range are the erythrocytes of cartilaginous fish (Elasmobranchs)and reptiles, the latter typically somewhat smaller (Fig. 2, cells d versus e). Despite these considerable size differences, the flattened ellipsoidal biconvex shape of nucleated nonmammalian vertebrate erythrocytes is essentially universal. Similar morphology occurs in mammalian primitivegeneration nucleated erythrocytes as well (Fig. 2, cell h), with the exception that many cells have reduced ellipticity or even discoidal profiles, as

-

a b C d e f 9 FIG. 2. Size and morphology of vertebrate erythrocytes. Cells a-g, nonmammals; cells

h-j, mammals. (a) Amphiuma tridactylum (giant salamander); (b) Notophthalmus uiridescens (Eastern newt or salamander); (c) Rana pipiens (leopard frog; face and edge view); (d) Mustelus canis (smooth dopfish); (e) Anolis carolinensis (anole, a lizard); (f) Carassius auratus (goldfish); (9) Callus domesticus (chicken); (h) Monodelphis domestica (gray shorttailed opossum; primitive erythrocyte of neonate); (i) Camelus dromedarius (camel, adult; face and edge view); (i)Homo sapiens (human, adult; face and edge view).

40

WILLIAM D.COHEN

observed in marsupial neonates (Cohen et al., 1990). In mammals of the camel family (e.g., camel, guanaco, and llama), circulating definitive erythrocytes are anucleate, but they are otherwise unusual among mammals, being elliptical even under nonflow conditions and flattened without biconcavity (Cohen, 1976; Cohen and Terwilliger, 1979) (Fig. 2, cell i). In all other mammals, typical mature definitive erythrocytes are biconcave disks when not in flow, as shown in face and edge views of a human erythrocyte (Fig. 2, cellj). The fundamental morphology of nucleated erythrocytes does not depend on the presence of the nucleus itself. In certain species representing several genera of salamanders (Emmel, 1924; Villolobos et al., 1988), >80% of the circulating erythrocytes are anucleate, yet most are flattened and ellipsoidal (Cohen, 1982). In addition, flattened elliptical shape is observed in other types of nucleated blood cells, such as nonmammalian vertebrate thrombocytes (Behnke, 1970a, b; Fawcett and Witebsky, 1964) and invertebrate clotting cells (Cohen and Nemhauser, 1985). Therefore, this shape is correlated with a common environment, rather than a common cargo or function. The real world of mature nucleated erythrocytes is not the microscope slide or the test tube in which we usually examine them, but rather a dynamic flowing tissue in which they are continuously subjected to pressure, deformation, and mechanical and osmotic testing. Their properties as observed under static experimental conditions can be misleading, a point pursued in Sections III,E and IV,A. What, then, is the functional significance of flattened ellipsoidal shape in nucleated erythrocytes? Flattening presumably enhances respiratory gas exchange by reducing diffusion distances, and, perhaps even more importantly, it orients cells in flow in such a way as to reduce overall blood viscosity (Chien, 1975; Fischer, 1978). Ellipticity reduces effective cell diameter for passage through narrow capillaries (Bloch, 1962), and it also appears to be important for cell orientation during blood flow, though its precise contribution is less clear (Cohen, 1978a; Fischer, 1978). Differences between nucleated erythrocytes of nonmammalian vertebrates and the nonnucleated definitive erythrocytes of mammals extend well beyond morphology. Typical adult mammalian erythrocytes are highly adaptable to physiological or experimental flow conditions, under which they rapidly assume shapes such as umbrellas and flattened ellipsoids. Although the nucleated erythrocytes of nonmammals are able to twist and bend as they negotiate turns of small radii, and to regain normal (i.e., equilibrium) shape rapidly, they exhibit much lower deformability than mammalian definitive erythrocytes during capillary flow (Usami et al., 1970; Gaehtgens et al., 1981a,b). In addition, under experimental conditions in uitro, mammalian erythrocytes align and elongate into flat-

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

41

tened ellipsoids in the flow direction without tumbling, whereas nonmammalian vertebrate erythrocytes assume more random positions and flip or tumble (Fischer, 1978; Gaehtgens et al., 1981a). The latter behavior is also exhibited by mammalian erythrocytes that have been made more rigid by glutaraldehyde fixation (Chien, 1975; Fischer, 1978). In mammalian definitive erythrocytes, the membrane exhibits “tank-treading”; that is, any point on the membrane will translocate all the way around the cell under flow conditions (Fischer and Schmidt-Schonbein, 1977; Fischer, 1978; Fischer et al., 1978). The general effect of membrane tank-treading is increased stability of cell orientation in flow and further lowering of blood viscosity, reducing the circulatory work requirement. Neither membrane tank-treading nor its physiological effects have been observed in nucleated erythrocytes of nonmammalian vertebrates, suggesting that tank-treading represents an evolutionary advance in erythrocyte design. Definitive mammalian erythrocytes also readily crenate, forming numerous surface protrusions, whereas those of nonmammals do not. In summary, definitive anucleate mammalian erythrocytes are morphologically much more responsive and behave like “fluid droplets,” while nucleated nonmammalian erythrocytes more closely resemble “solid bodies” (Fischer, 1978; Fischer er al., 1978). The basis for these physiological differences is to be found in large part in differences between the cytoskeletal systems of these two erythrocyte types. That of nucleated erythrocytes consists of the following major components: the MB of MTs; the cell surface-associated cytoskeletal network, or membrane skeleton (MS); and intermediate filaments (IFs) of the vimentin class. These components, interacting as a system, are believed to be responsible for the generation and maintenance of nucleated erythrocyte morphology and for mechanical properties critical to cell function. Of the three, only the MS has a counterpart in typical definitive mammalian erythrocytes. The basic features of the MB/MS system are illustrated in Fig. 3, using the dogfish erythrocyte as a typical example. A somewhat oversimplified diagram of the cytoskeletal system of mature nucleated erythrocytes is presented in Fig. 4. The MB is completely enclosed within the MS and tightly apposed to it as a supporting frame (Fig. 4a). The nucleus is suspended between MS networks lining the inner surface of the plasma membrane on opposite sides of the cell, producing a bulge in edge view (Fig. 4b). For simplicity, the diagram omits IFs, which traverse the region between nucleus and MS. In categorizing erythrocyte behavior as fluid versus solid, it is important to note that the fundamental distinction is not equivalent to primitive versus definitive, mammalian versus nonmammalian, or even nucleated versus nonnucleated. With respect to structure and behavior, primitive

FIG. 3. The cytoskeletal system of dogfish erythrocytes (Mustelus canis), typical of nearly all nonmammalian vertebrates. (a) Erythrocyte cytoskeleton produced by Triton X-100 lysis of cells under microtubule (MT)-stabilizing conditions; uranyl acetate-stained whole-mount, transmission electron microscopy (TEM). The marginal band (MB) and the nucleus (N)are densely stained. The membrane skeleton (MS) appears as a more lightly stained network bordered by the MB. (b) Thin cross section through MB in simultaneously lysed and fixed cell. MTs constituting the MB are enclosed with the MS at the cell extremity. Remnants of the membrane bilayer and hemoglobin (fuzzy clumps) are present due to the preparation method used. (c) MB isolated by detergent-based MS dissolution (whole-mount, TEM). (d) Lane I: SDS-PAGE pattern of whole cytoskeletons, showing spectrin-region (S) and tubulin-region (T) polypeptides, plus some actin (A, faint) and a few other components; lane 2: isolated MBs, at comparable tubulin-region loading, exhibiting the same tubulin pattern as in whole cytoskeleton and only minor amounts of other components. (b) From Cohen et al. (1982a); reproduced from the Journal of Cell Biology, 1982, 93, 828-838, by copyright permission of the Rockefeller University Press. (c and d) From Sanchez et al. (1990); reproduced from the European Journal ofcell Biology, 1990,52,349-358, by permission of Wissenschaftliche Verlasgesellschaft MBH.

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

a

43

b

FIG.4. Model of the nucleated erythrocyte cytoskeletal system, diagrammatically simplified. (a) Face view, showing the marginal band (MB) completely enclosed within the membrane skeleton (MS), and in contact with it. (b) Edge view, showing biconvexity due to the presence of the nucleus. Intermediate filaments, believed to connect the inner cell surface to the nucleus or to the opposing inner cell surface, are omitted for simplicity. Modified from Cohen and Nemhauser (1985), “Marginal bands and the cytoskeleton in blood cells of marine invertebrates,” in Blood Cells of Marine Invertebrates: Experimental Systems in Cell Biology and Comparative Physiology (W. D. Cohen, ed.), pp. 1-49, with permission of WileyLiss Div. of John Wiley & Sons, Inc.

and definitive erythrocytes of nonmammals are not significantly different, and the same can be said for primitive erythrocytes of mammals versus nonmammals, and for the anucleate erythrocytes of certain nonmammalian vertebrates versus nucleated ones, as documented in subsequent sections. Rather, the real distinction is between erythrocytes in which MBs, interacting with other cytoskeletal elements, play a role in the determination of erythrocyte morphology and mechanicalhheological properties (solid body behavior) and erythrocytes in which MBs have no such role (fluid droplets) (Cohen, 1978a; Fischer, 1978). 111. The Marginal Band of Mature Erythrocytes

All of the mature nucleated erythrocytes of nonmammalian vertebrates (Fig. 2, cells a-g) contain MBs. The MB is principally a hooplike continuous MT bundle located close to the plasma membrane in the plane of flattening. Few, if any, MTs are found elsewhere in the cells; thus, the MB is one of the simpler MT systems in terms of organization and spatial

44

WILLIAM D. COHEN

distribution. Therein lies its fundamental attraction as the subject of a variety of experimental studies in recent years. For erythrocytes of nonmammalian vertebrates, MB thickness in the light microscope and MT number per electron microscopic (EM) cross section are positively correlated with cell size (Small and Davies, 1972; Goniakowska-Witalinska and Witalinski, 1976; Cohen, 1978b). Thus, the Amphiuma erythrocyte MB (Fig. 2, cell a) is about 1pm thick and contains hundreds of MTs per cross section (Small and Davies, 1972), whereas the corresponding values for goldfish (Fig. 2, cell f ) are -0.1 pm with eight to 10 MTs (Weinreb and Weinreb, 1965). For diverse species representing most vertebrate classes, the relationship between cell long axis ( L ) and MT number (n) is roughly logarithmic; log n = 0.05L + 0.3, with greater deviations occurring in the smaller cells (derived from the data of Goniakowska-Witalinska and Witalinski, 1976). Species can thus be selected so as to provide MBs of size and thickness appropriate to a particular experimental problem. MBs are also present in mammalian blood platelets (Behnke, 1970b), nonmammalian vertebrate thrombocytes (Fawcett and Witebsky, 1964), and invertebrate nucleated erythrocytes and clotting cells, representing a wide phylogenetic range (Cohen and Nemhauser, 1985). Although these MBs are of considerable interest from a comparative standpoint, the discussion here focuses principally on MBs of vertebrate erythrocytes.

A. STRUCTURE In their original paper on the ultrastructural features of erythrocyte MBs, Fawcett and Witebsky (1964) raised fundamental questions regarding their construction. Remarkably, these questions have not yet been answered unequivocally for a single species. For purposes of discussion, it is helpful to set forth several hypothetical models (Fig. 5). A closed-hoop model (Fig. 5, model I) would seem least likely for any MB, although not totally absurd. MT annealing (Rothwell et al., 1986) allows for the possibility that a +MT end might grow around the cell perimeter and join onto its own -end, but such a rendezvous is certainly a complex scenario. In fact, closed MT hoops have never been seen separating from others during MB isolation experiments, and the model I prediction that MBs will always exhibit the same MT number at opposite sides in cross section is not supported by observation. Moreover, in at least some erythrocyte cytoskeletons, individual MTs appear to traverse the circumference more than one complete turn (Miller and Solomon, 1984; Cohen and Nemhauser, 1985). The overlapping MT segment model (Fig. 5, model 11) is also highly unlikely. Individual MTs, when traceable within erythrocyte cytoskel-

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

I. CLOSED

11.OVERLAPPING SEGMENTS

HOOPS

3

P

V

4-

-4

A

4

111.SINGLE

VERY LONG MICROTUBULE

45

3-

A

4 SMALL NO. LONG MICROTUBULES OF SAME OR MIXED POLARITY

FIG. 5. Possible models of marginal band (MB) construction. Models I and I1 are highly unlikely for any MB; model I11 may be correct for MBs of some species; model IV is probably correct for MBs of most species. Numbers adjacent to arrowheads indicate the numbers of microtubules that would be observed in cross section through the MB at various points.

etons or isolated MBs, appear to be very long, and few MT ends are normally visible. In addition, kinetic data indicate that, for reassembling chicken erythrocyte MBs at least, there are probably only one to three free growing MT ends present (Miller and Solomon, 1984), and chicken erythrocyte MT protein characteristicallyforms very long MTs in vitro (Murphy and Wallis, 1983b).

46

WILLIAM D. COHEN

The literature thus supports models in which MBs are constructed of only one or a few extremely long MTs. For relatively thin MBs of small erythrocytes (e.g., birds and bony fish), the single coiled MT (Fig. 5 , model 111)may well apply. It may also be correct for the mammalian platelet MB, although this has not yet been demonstrated directly (Nachmias et al., 1977; Nachmias, 1980; White et al., 1986). For an elliptical erythrocyte in which L = only 10 pm, a single MT with but 10 windings would be almost 300 pm long! This model predicts a maximum difference of only one MT at opposite sides in a given MB cross section, regardless of the number of windings. Although not yet studied systematically, a difference of 1 is frequently observed in thin MBs such as those of goldfish, in which low MT number makes accurate counting feasible (W. D. Cohen, unpublished observations). However, the single-MT model cannot apply universally because, depending on the species, some (turtle and bullfrog) or all (mudpuppy) MBs have been found to contain MTs of mixed polarity, as determined using the tubulin “hook” method (Euteneuer et al., 1985). For these cells, at least, MBs are believed to consist of a relatively small number of very long MTs of opposite polarities (Fig. 5 , model IV). For two multiply wound MTs, model IV predicts a maximum difference of 2 at opposite MB sides regardless of the number of windings of either MT, with each additional multiply wound MT increasing the maximum difference by 1 . This is as yet untested. The polarity studies by Euteneuer et al. (1985) also revealed (1) a mirror-image relationship between MT polarity patterns in opposite-side MB cross sections, (2) relative constancy of MT number and polarity pattern throughout MB length (i.e., no MT overlap zone), and (3) variation in MT polarity patterns in different MBs of an individual animal and species. Features 1 and 2 are consistent with a model proposed originally for experimentally induced reassembly of the blood clam erythrocyte MB (Nemhauser et al., 1983) (Fig. 9, discussed in Section 111,D). Feature 3 implies a degree of permissible variability in the MB biogenetic mechanism. It is interesting that the percentage of cells having mixed MT polarity was highest in the largest cells studied (100% in the mudpuppy), lowest in the smallest ones (14%in the turtle), and intermediate in those of intermediate size (43% in the bullfrog) (Euteneuer et al., 1985). Such a correlation could be tested using much smaller erythrocytes of birds or bony fish, having very thin MBs. Direct determination of the actual number of MTs composing a given MB is quite difficult in practice. The possibility of artifact due to MT breakage during MB isolation or to poor in situ fixation cannot be eliminated with certainty. In addition, MB curvature precludes complete threedimensional reconstruction by serial sectioning, although up to 60% of MB

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

47

length has been examined using a tilt stage (Euteneuer et al., 1985). The best current approach to this problem may be high-resolution videoenhanced contrast microscopy of relatively thin MBs undergoing experimentally induced disorganization in real time, combined with negative staining to verify that the MTs osbserved by video are truly single. With respect to three-dimensional arrangement, MBs do not consist of MTs packed uniformly into a tight cylindrical shape, but rather more asymmetrical ribbonlike arrays (Behnke, 1970a,b; Small and Davies, 1972; Cohen et al., 1982a). Negatively stained isolated MBs often look ribbonlike when flattened onto the substrate, and in scanning electron micrographs they appear as thick ribbons (Bertolini and Monaco, 1974; Cohen, 1978b; Sanchez et al., 1990). Electron microscopy of MBs in situ or after isolation frequently reveals inter-MT cross-bridges (e.g., Cohen et al., 1982a; Centonze et al., 1984), but their number and spatial distribution are unknown. In very large erythrocytes of amphibians (e.g., Amphiurna) the MB is more highly flattened and ribbonlike at the two ends of the elliptical cell than elsewhere. This was noted by Meves (191l), and can be seen immediately upon cell lysis under MT-stabilizing conditions (Cohen, 1978b). It is possible that such flattening permits the MB to negotiate a smaller radius of curvature at these extremities, thereby allowing large cells with otherwise thick MBs to attain elliptical, as opposed to discoidal. cell shape. Regardless of functional significance, the question remains open as to how this structural differentiation is achieved. B. MECHANICAL PROPERTIES The MB is flexible both in situ and after isolation; that is, it can bend and twist without breaking. This was well documented by Meves (191l), and Fawcett and Witebsky (1964) also noted that MBs could form and sustain loops in situ. Goniakowska-Witalinska (1974) observed figure-eight MB twisting in situ in amphibian erythrocytes swollen by exposure to hypoosmotic media, in which MBs appeared to be separated by a considerable distance from the plasma membrane except at the two ends of the cellular ellipse. MBs in erythrocyte cytoskeletons prepared by detergent lysis also typically twist into figure-eight forms (Bertolini and Monaco, 1974;Cohen, 1978b)(Fig. 6 a +. b), but here, twisting appears to be an accomodation to reduction in MS surface area, with the MB confined within. When the MS is removed from cytoskeletons containing figure-eight MBs, using high salt, proteases, or detergents (Cohen, 1982; Cohen and Ginsburg, 1986; Sanchez et al., 1990), the isolated MBs typically reassume a more planar configuration (Fig. 6c). Therefore, MS shrinkage or contraction probably

48

WILLIAM D. COHEN

-

(el tangential stripping

cell lysis

(a] cytoskeleton in situ

MS

Giz?

-

(b] f i g i r e 8 twisting

(cl isolated MB circularirat ion

FIG. 6. Mechanical behavior of the cytoskeleton and isolated marginal band (MB) in uitro. The nucleus is omitted for simplicity; for a detailed explanation, see text.

imposes forces on the MB sufficient to cause twisting (see Section V,D and Fig. 15). In a figure-eight twist, there are two possible mirror-image configurations: right-handed or left-handed. Counts in several amphibians have shown that the MB twist direction in figure-eight cytoskeletons is nonrandom, strongly favoring right-handedness (Cohen, 1978b). The reason for this is unknown, but MB construction by unequal numbers of MTs of opposite polarity (Euteneuer et al., 1985) could be a factor. Additional information on MB mechanics has been obtained by studying MB structure during isolation and subsequent experimentally induced disorganization (Fig. 6c-e). MBs in intact nucleated erythrocyte cytoskeletons typically retain the ellipticity of living cells after cell lysis (Fig. 6a). However, they become much more circular when released from the cytoskeleton by MS dissolution, regardless of the particular agent or conditions utilized (Bertolini and Monaco, 1974; Cohen, 1982; Cohen and Ginsburg, 1986; Sanchez et al., 1990)(Fig. 6c). Isolated MBs remain intact even during micromanipulation (Cohen, 1978b; Waugh et al., 1986; Waugh and Erwin, 1989),their integrity presumably attributable to the preservation of inter-MT cross-bridges. Isolated MBs transected accidentally, or experimentally by means of microneedles, typically open outward so as to become more linear (Fig. 6d). Continued proteolysis of MBs after proteolytic isolation from cytoskeletons frequently results in the “stripping” of MTs or MT bundles from the MB, with linearization tangential to the curved MB surface (Fig. 6e). Mechanical properties of isolated MBs have been studied by means of micromanipulation on hooks, accompanied by measurements using calibrated glass fibers (Waugh et al., 1986; Waugh and Erwin, 1989). MBs were sufficiently flexible to return to their original shape after extension

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

49

into a highly elongated oval, maintaining essentially constant circumference during deformation and recovery (Fig. 6f). For MBs of the newt Notophthalmus uiridescens isolated using proteases or high KCl, average flexural rigidity (indicating resistance to bending) was -9 x lo-’’ dyn-cm’, and average extensional rigidity (resistance to increase in length produced by axial forces) was 0.017 dyn. Waugh and co-workers concluded that MBs are essentially inextensible compared with the erythrocyte membrane, implying maintenance of constant MB circumference during erythrocyte deformation. In addition, the flexural rigidity of isolated MBs was several thousand times greater than that calculated for the membrane, indicating that MBs stabilize the overlying cell surface against indentation. Cytoskeletal behavior, as summarized in Fig. 6, is consistent with a mechanical model in which the intact MB is stressed due to curvature [originally described as bending strain (Cohen, 1978b; Joseph-Silverstein and Cohen, 1984, 198S)l. In other words, the curved MTs behave as if preferring to be straight. Within the complete system, the MB is assumed to be deformed into an ellipse through forces applied by other cytoskeletal elements (Cohen, 1978b; Waugh and Erwin, 1989) (Fig. 6a). Release from the cytoskeleton during isolation would produce MB circularization due to the spontaneous equalization (i.e., redistribution) of strain (Fig. 6a-c), and loss of continuity by transection or proteolytic stripping would result in linearization as bending strain is relieved (Fig. 6d and e). The mechanical characteristics of the MB in mature nucleated erythrocytes thus indicate that it is a strained flexible structural frame, that is, a relatively passive and stable skeletal structure providing for the attachment and support of other structural elements. This is considered further in Section II1,E. C. MOLECULAR COMPONENTS 1. Tubulin

MBs consist principally of MTs, and thus tubulin is their major molecular component. The properties of nucleated erythrocyte tubulin have been studied by Murphy and co-workers, using chicken erythrocytes (Murphy and Wallis, 1983a,b, 1985; Murphy et al., 1987; Rothwell et al., 1986). Tubulin was found to constitute 1% of total cell protein, with less than one-half in the MB. Erythrocyte tubulin assembled with greater efficiency and lower nucleation rate than did brain tubulin, producing much longer MTs under similar conditions. It was proposed that the assembly of long MTs typical of MBs might be regulated by a tubulin oligomer pool, limiting the nucleation rate and accounting for at least some non-MB tubulin in the cell (Murphy and Wallis, 1985).

50

WILLIAM D.COHEN

The characteristics of chicken erythrocyte tubulin in uitro may derive, in part, from its unusually divergent p subunit (Murphy and Wallis, 1983a; Murphy et al., 1987). This isotype, cp6, is the product of a gene expressed only in developing erythrocytes and thrombocytes in chick bone marrow, i.e., cells assembling MBs (Murphy et al., 1986). It is one of six known chicken p-tubulin isotypes, constituting 95% of the p-tubulin in mature chicken erythrocytes, cp3 making up the remaining 5% (Joshi et al., 1987). Another p-tubulin isotype, MPl , identified in the mouse, is proposed to be involved in MB formation in mammalian primitive erythrocytes and platelets. It differs from cp6 in 18% of amino acid residues, but also shares with it unique amino acid similarities at many positions throughout its structure (Wang et al., 1986; Murphy et al., 1987). Since Mpl was identified using a mammalian bone marrow library, and since mammalian bone marrow erythroblasts do not contain MBs (Repasky and Eckert, 1981, vs. Grasso, 1966), it is possible that Mpl is platelet specific and that there exists another p-tubulin unique to mammalian primitive erythrocyte MBs. It is important to note, however, that the broader question as to whether MB-specific p-tubulins occur universally has not yet been addressed systematically. Data on MB tubulins in other species are more limited. After disassembly of dogfish erythrocyte MBs in living cells at low temperature, four major polypeptides increased in the cytosol (Cohen et al., 1982a),all in the tubulin region of sodium dodecyl sulfate (SDS) gels. Isolated dogfish erythrocyte MBs also yield these four polypeptides, with the same stoichiometry as in the whole cytoskeleton (Sanchez et al., 1990),and preliminary Western blotting indicates that all are tubulins (Sanchez and Cohen, 1990 unpublished observations). MT protein, obtained from dogfish erythrocyte MBs either by low-temperature extraction of whole cytoskeletons or by isolated MB disassembly, reassembles readily into MTs in uitro (Cohen et al., 1982b). 2 . Other MB-Associated Proteins The term “MB-associated proteins” as used here includes those indirectly associated with the MB as well as MT-associated proteins (MAPS) having MT binding sites. Possible functions for the latter include nucleation and stabilization of MTs during MB assembly, cross-bridging of MTs into bundles, and linking of the MB to the MS and the plasma membrane. In erythrocytes of the amphibian Bufo marinus (marine toad), the MB contains a protein antigenically similar to brain MAP 2, and gold-labeled anti-MAP 2 binds to MB cross-bridges in this species (Sloboda and Dickersin, 1980; Centonze et al., 1984, 1985). In addition, a protein of M , 280,000 obtained from the cytoskeleton cross-reacts with MAP 2 antibody and can cause bundling of reassembled brain MTs (Centonze and Sloboda, 1986).

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

51

Recently, Feick et al. (1991; see also the review by Wiche et al., 1991) have isolated syncolin, a high-molecular-weight MT-bundling protein from chicken erythrocytes that is probably the same protein studied in Sloboda’s laboratory. The starting material consisted of taxol-polymerized MTs obtained from extracts of sonicated cells, from which syncolin was released at low temperature. Syncolin comigrated with MAP 2 in SDSpolyacrylamide gel electrophoresis (SDS-PAGE) (M, 280,000) and also showed cross-reactivity with anti-MAP 2. However, it was quite different in structure, exhibiting heat lability and assembling into 13-nm spheroids of 990,000 average molecular weight, rather than being fibrous. Two observations suggested to the authors that syncolin was involved specifically in MT-MT interactions, as opposed to functioning as either a spatial determinant or a MT-MWmembrane link: (1) syncolin remained diffusely associated with the cytoskeleton after low-temperature MB disassembly, but reassociated with the MB after temperature-induced MB reassembly, and (2) in mature erythrocytes, syncolin showed immunofluorescent colocalization only with MBs, even when MBs were highly twisted. These observations do not constitute a compelling argument for MT-MT bridging, however, since the interface between MB and MS would likely be intact even in cytoskeletons containing highly twisted MBs. It would thus be helpful to determine syncolin distribution in intact chicken erythrocyte cytoskeletons versus isolated MBs. Other studies have not demonstrated the presence of high-molecularweight MAP 2- or syncolin-like MAPs, or have yielded equivocal results. Temperature cycling experiments revealed only two putative MAPs of M , -80,000 and 100,000 that coassemble with an invertebrate (blood clam) erythrocyte MB in living cells (Joseph-Silverstein and Cohen, 1986). In Xenopus erythrocytes, cytoskeletal polypeptides of appropriate molecular weight did not cross-react with antibodies to mammalian or avian MAP 2 (Gambino et al., 1985). MBs isolated from amphibian (Triturus cristatus) erythrocytes were reported to contain numerous non-tubulin components, including actin, myosin, M , 90,000 glycoprotein, and other highmolecular-weight proteins (Monaco et al., 1982), but many of these appear to be contaminants resulting from incomplete MS removal (Cohen and Ginsburg, 1986). Neither myosin nor high-molecular-weight MAPs have been found in MBs isolated from dogfish erythrocytes, although minor amounts of spectrin and actin have been observed (Sanchez et al., 1990). Tau protein, but not MAP 2, has been found in MT protein preparations purified from chicken erythrocytes by temperature cycling, and the MB of chicken erythrocyte cytoskeletons binds anti-tau in situ (Murphy and Wallis, 1985). Although a high-molecular-weight component possibly corresponding to syncolin is present in whole dogfish erythrocyte cytoskeletons, MBs isolated from these cytoskeletons by a recently developed

-

52

WILLIAM D.COHEN

detergent-based method have been found to contain tau proteins, but not high-molecular-weight MAPs (Sanchez et al., 1990; Sanchez and Cohen, 1990). The presence of tau in MBs may reflect a role in MT nucleation (Murphy and Wallis, 1985), MT-MT cross-bridging, or both. Considerable information on tau properties is now available, derived in part from studies related to its presence in the paired helical filaments of Alzheimer’s disease (Ihara etal., 1986; Grundke-Iqbal et al., 1986). Like MAP 2, tau proteins have an MT-binding domain consisting of three 19-amino-acid repeats near the carboxy-terminal end (Lee et al., 1988). The long amino-terminal remainder of the molecule, thought to project away from the MT, probably does not have an MT crosslinking function. Rather, the shorter carboxyterminal portion beyond the repeats (Lewis et al., 1989), or possibly including the last repeat (Lewis and Cowan, 1990), appears able to bind to the same region of another tau (or MAP 2) molecule. Thus, binding between two tau molecules on adjacent MTs would create an inter-MT cross-bridge. The ability of tau to induce MT bundling in living cells has been demonstrated elegantly by engineering the expression of tau or tau variants in cultured cells which normally do not produce it (Kanai et al., 1989; Lewis et al., 1989). One appeal of a tau-tau cross-bridge is that MB MTs would require only one type of MAP binding site, rather than a different one for each end of a more specialized bridge protein. In addition, if the same type of bridge were made with a membrane/MS-bound form of tau, MTs could link both to the inner cell surface and to other MTs without having to bind MAPs selectively. Such a mechanism is speculative, of course, and a membrane-bound form of tau has not yet been reported. However, polymerized actin has been localized to the plane of the mature erythrocyte MB (Kim et al., 1987), and binding of tau protein to actin has been demonstrated in uitro (Selden and Pollard, 1983). Studies in two distantly related species suggest that tau is a common component in vertebrate MBs (Murphy and Wallis, 1985; Sanchez et al., 1990). As shown in curved paracrystals, tau has elasticity that decreases with increased phosphorylation [length and stiffness increase (Lichtenberg ef al., 1988; Lee, 1990)l. Such elasticity could be an important factor in MB mechanical properties. MTs bridged by tau carboxy terminal regions would be very closely spaced (Lewis ef al., 1989), as usually observed in situ and in isolated MBs. However, relatively weak tau-tau binding might also permit temporary separation of the MTs making up the MB in regions of extensive cell deformation, with MTs “zipping” back together upon removal of the deforming forces. This could explain why MB MTs in situ are sometimes observed to be separated from each other by a considerable distance, whereas in isolated MBs they are usually tightly bundled.

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

53

Actin and ezrin are two additional proteins that appear to be MB associated. Use of rhodamine-labeled phalloidin on mature chicken erythrocytes has revealed polymerized actin with a distribution similar to that of MB tubulin (Kim et a f . , 1987). It is not yet clear whether any of this actin composes the MB itself, or whether it is principally associated with the MS in the plane of flattening. Another protein localized to the MB plane via monoclonal antibody binding has at least some properties identical to those of ezrin (Birgbauer and Solomon, 1989), a M , -80,000 component isolated previously from intestinal brush border (Bretscher, 1983). Localization of erythrocyte ezrin in the plane of flattening during MB biogenesis follows, rather than precedes, that of the MB MTs, and thus it is not a likely candidate for the spatial control of MB formation in erythroblasts (Birgbauer and Solomon, 1989). However, it might well be involved in the control of MB reassembly after experimentally induced disassembly. INDUCED MB REASSEMBLY IN MATURE CELLS D. EXPERIMENTALLY The basic sequence of morphological stages during the differentiation of nucleated erythrocytes, and the accompanying changes in cytoskeletal properties, are summarized in Fig. 7. The focus in this section is on the mature erythrocyte MB (Fig. 7, cell 4), with earlier stages of the sequence discussed in Section IV. Colchicine or related MT inhibitors applied at physiological temperature appear to have little or no effect on MB MTs or the shape of mature cells, regardless of species (Fig. 7, cell 4-4a) (Behnke, 1970a,b; Barrett and Dawson, 1974; Cohen et al., 1982a; Gambino et af., 1985; Kim et al., 1987). In some species (e.g., frogs), exposure to low temperature (0-4°C) also has no effect, whereas in others (e.g., chicken, dogfish, and blood clam) such exposure induces MB disassembly (cell 4+4b) (Behnke, 1970a,b; Cohen et al., 1982a; Nemhauser et al., 1983). Despite lowtemperature MB disassembly, flattened elliptical cell morphology is retained (Fig. 7, cell 4b). Return of such cells to physiological temperature appropriate to the species results in MB reassembly, usually in 1-2 hours (Fig. 7, cell 4b+4c). Although colchicine and related inhibitors do not induce MB disassembly, they block reassembly at physiological temperature after low-temperature MB disassembly (Fig. 7, cell 4b+4d) (Cohen et al., 1982a; Nemhauser et af., 1983). In the case of mature blood clam erythrocytes, a pair of centrioles is physically associated with the MB in every mature cell (Cohen and Nemhauser, 1980) (Fig. 8). These centrioles form part of an MT organizing center (MTOC) that participates in temperature-induced MB reassembly (Nemhauser et af., 1983). MT growth initiates at this MTOC, and individual MTs or thin MT bundles elongate so as to contact the MS within

54

WILLIAM D. COHEN

KEY: +T=IOWtemp. (0-4Oc) 4 T =physiol. temp COI=colehiclne

CyB= cylochalasin B CaI=calcium ionophore c> =sphere, no MB Q=flattened disc +MB flattened ellipse+ MB

o=

blood clam)

o=

flattened ellipse, noMB

FIG. 7. Properties of the cytoskeleton during morphogenesis and maturationof nucleated erythrocytes. The natural pathway from erythroblast to mature erythrocyte is presented in steps 1-4 (heavy arrows). All other steps (2a-2d; 4a-4e) represent experimental manipulations. See text for a detailed explanation.

-15-20 minutes. Inter-MT cross-bridging occurs after a significant number of MTs are already at the periphery in the plane of flattening. In the sequence illustrated in Fig. 9, it is proposed that MTs grow in opposite directions around the cell periphery (Fig. 9a) and cross each other distally (Fig. 9b), eventually closing the MB proximal to the MTOC (Fig. 9c) and continuing on in their respective directions. This would produce MBs of mixed MT polarity, corresponding to model IV (Fig. 5, Section 111,A). In contrast, centriole-containing MTOCs are not observed during temperature-induced MB reassembly in mature chicken erythrocytes (Miller and Solomon, 1984). Here, MT growth appears to be largely localized to the periphery, and MT numbers as observed in whole mounts are consistent with model I11 (Fig. 5 ) . With the exception of skate erythrocytes (Cohen, 1986), MB-associated centrioles have not been found in vertebrate erythrocytes; however, thin sections of amphibian and reptilian erythrocytes reveal centrioles in regions near the nucleus (Gambino et al., 1984; Euteneuer et a / . , 1985).

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

55

FIG. 8. Blood clam erythrocytes and their cytoskeletal system. (a) Living erythrocytes, seen by phase contrast. Most are ellipsoidal, but occasionally singly pointed cells are observed(arrowhead1. (b-d) Erythrocyte cytoskeletons produced by Triton X-100lysis of cells under microtubule (MT)-stabilizingconditions. (b) A pair of centrioles is associated with the marginal band (MB) in every cell, appearing as adjacent dense dots in phase contrast (arrowhead). N, Nucleus. (c) Uranyl acetate-stained cytoskeleton whole-mount, transmission electron microscopy (TEM). MB-associated centrioles (Ce)are densely stained. The MB is, in this case, twisted into a figure eight, with the membrane skeleton (MS) more readily visible where overlapped near the crossover point. N, Nucleus. (d) A centriole pair in thin-section, TEM. In typical elliptical cells, centrioles are usually located at or near one end of the ellipse. In pointed cells, centrioles are invariably located at the point. (a,b, and d) Noetia ponderosa; (c) Anadara transversa. From Cohen and Nemhauser (1980) and Nemhauser et al. (1983);reproduced from the Journal of CellEiology, 1980,86,286-291; and 1983, %, 979-989, by copyright permission of the Rockefeller University Press.

56

WILLIAM D. COHEN

FIG. 9. Proposed model for experimentally induced marginal band (MB) reassembly in blood clam erythrocytes. (a) Microtubules (MTs) initiate at the centriole-containing organizing center and grow toward the distal end of the cell in diverging directions. (b) MTs pass each other distally and continue growth around the periphery. (c) MTs pass each other proximally, closing the MB. The resulting MB contains MTs of mixed polarity. From Nemhauser et al. (1983); reproduced from the Journal of Cell Biology, 1983,%, 979-989, by copyright permission of the Rockefeller University Press.

E. MB FUNCTION I N MATURE CELLS Retention of mature erythrocyte shape after MB disassembly was initially interpreted as indicating nonparticipation of MBs in cell shape maintenance (Barrett and Dawson, 1974; Behnke, 1970a). This was somewhat illusory, however, because cells were examined under the static conditions of microscopic examination. Challenged with mechanical stress (fluxing through capillary tubes) or hyperosmotic conditions, both dogfish and blood clam erythrocytes pretreated so as to lack MBs behave quite differently from those containing MBs ( Joseph-Silverstein and Cohen, 1984,1985), as summarized in Fig. 10. Here, mature cells with and without MBs at physiological temperature were prepared by MB disassembly at low temperature followed by rewarming with or without colchicine. Mechanical stress produced a significant proportion of deformed cells only in cells lacking MBs (Fig. 10, sequence a-c versus a-d). The same was true for cells without MBs at low temperature, as compared with those containing taxol-stabilized MBs (Fig. 10, a-g versus a-f). These results indicate that MBs can restore and/or stabilize mature erythrocyte shape, regardless of temperature. However, no quantitative information is available on the extent to which MBs resist an imposed force, the yield point or range at which MB deformation begins, or the rapidity of MB recovery and restoration of cell shape. The relationship between MB thickness and the effect of imposed forces is similarly unexplored.

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

57

FIG. 10. Marginal band (MB) function in mature erythrocytes. General effects of mechanical stress (cells with fist) and hyperosmotic conditions ( t 0s) on cells containing or lacking MBs, as observed in dogfish erythrocytes. t T, Physiological temperature (20-22°C); T, 0°C; tax, taxol; col, colchicine. See text for a detailed explanation.

In experiments testing MB influence on cellular responses to hyperosmotic conditions ( Joseph-Silverstein and Cohen, 1984), significantly greater numbers of cells “shriveled” if they lacked MBs (Fig. 10, a-b versus a-e). This indicates that the MB acts as an internal frame supporting the cell surface from within, again resisting distortion of mature cell shape. An interesting feature of the shriveling response was its all-or-none

58

WILLIAM D. COHEN

nature; that is, cells lacking MBs were either relatively normal in morphology or drastically shrunken and distorted, without intermediate stages. One possible explanation is that there is resistance to shrinkage until cellular contents suddenly burst outward through a localized weak spot in the MS and the bilayer. This would parallel the mechanism of hypoosmotic lysis as currently understood for both mammalian and nonmammalian erythrocytes (Lieber and Steck, 1982a,b;Lieber et al., 1987).The capacity of the MB to support and deform the cell surface from within has also been evident in other studies with mature dogfish erythrocytes. These cells acquire pointed ends during long-term incubation in Elasmobranch Ringer’s solution at physiological temperature, a phenomenon useful for testing the causal relationship between cell and MB pointedness. These experiments are included in the discussion of the MS (Fig. 16, Section V). The mechanical properties of MBs (Section II1,B) indicate a functional mechanism based on flexibility, resistance to bending, and the capacity to return to equilibrium shape by the equalization (i.e., redistribution) of strain. These features would all appear to be independent of MT polarity. Therefore, they are compatible with the observed variability of MT polarity within erythrocytes of an individual organism (Euteneuer et d., 1985), and with the possibility that different modes of MB biogenesis (Section IV) or reassembly can achieve the same mechanical end in different species and cell types. TO FRIEDRICH MEVES F. A TRIBUTE

This discussion would be incomplete without reference to the work of Meves, who began study of the “marginal ring” in salamander erythrocytes in 1903, equipped only with a light microscope. Challenged by skeptical contemporaries, Meves showed convincingly that the MB was not a cytological artifact by partially isolating it and by devising special staining procedures that revealed MB substructure. The following admirable observations and conjectures are taken from his classic 1911 paper (translated from the German): One can perceive parallel lines within the ring which become more distinct after the cell body has lost its hemoglobin. The marginal ring now appears to be composed of many ultrafine filaments running in parallel or, what is just as possible, of a single, uninterrupted filament which is wound into a skein along the edge of the blood disk. In the pole areas the filaments often maintain larger intervals between each other; the skein, if one exists, is looser in these areas. . . . In blood cells treated with a gentian violet solution, one frequently observes loop formations at one or both poles of the marginal ring, which are probably caused by torsion of the ring. . . . It is well known that the red blood corpuscle can change its shape passively due to a mechanical influence, be it within the body or without; however, it assumes its original shape as soon as the force ceases. This is made

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

59

possible by the elasticity inherent in the marginal ring, which allows it to return to its natural state.

IV. Marginal Band Biogenesis and Function during Erythrocyte Morphogenesis The morphogenetic sequence for nucleated vertebrate erythrocytes and accompanying cytoskeletal properties are summarized in Fig. 7. The first of three major stages encompassed is jattening, or conversion of the spheroidal erythroblast to a flattened disk concomitant with MB biogenesis (Fig. 7, cell 1-2) (Barrett and Scheinberg, 1972; Barrett and Dawson, 1974; Dorn and Broyles, 1982). The second stage is elliptogenesis, in which the flattened discoidal cell becomes ellipsoidal (Fig. 7, cell 2+3), and the third is maturation, during which cell morphology and properties of the erythrocyte cytoskeletal system change toward greater stability (Fig. 7, cell 3+4) (Barrett and Scheinberg, 1972; Barrett and Dawson, 1974; Dorn and Broyles, 1982, Kim et al., 1987). Details of the sequence and specific timing of major structural and biochemical events are not yet well known. Thus, the separation between elliptogenesis and maturation indicated in Fig. 7 may be somewhat artificial in that at least some properties associated with maturation probably begin to appear during elliptogenesis. Moreover, it is not yet clear whether a discoidal stage precedes the generation of ellipsoids in all species (Twersky et al., 1990). However, such definition and separation of stages are convenient here for purposes of discussion. A. MB BIOGENESIS AND THE MECHANISM OF CELLFLATTENING

MB biogenesis is considered a causal factor in the flattening of nucleated erythrocytes. This was evident in the work by Barrett and Dawson (1974) on chick bone marrow cells, in which newly flattened discoidal cells exposed to low temperature reverted to spheres (Fig. 7, cell 2-2b). These spheres reflattened upon restoration of physiological temperature (here, 37"C), even if cytochalasin B was present (Fig. 7, cell 2b+2c), but such reflattening was blocked by colchicine (Fig. 7, cell 2b+2d). However, exposure to colchicine did not cause MB disassembly in newly flattened discoidal cells maintained at physiological temperature (Fig. 7, cell 2-2a). One fascinating observation was the initiation of flattening while cultured erythroblasts were still physically associated as postmitotic daughter cell pairs (Barrett and Scheinberg, 1972), a phenomenon worthy of further investigation.

60

WILLIAM D. COHEN

Information on properties of immature flattened discoidal cells derives principally from studies of chicken erythrocytes, which have cold-labile MBs even after maturation. Far less is known about discoid cells of other species, such as amphibians (Dorn and Broyles, 1982), in which the MB of mature elliptical cells is stable at low temperature (Fig. 7, cell 444a). An early electron microscopic study of MB biogenesis in chick primitive erythrocytes showed that small MT bundles first appeared near the cell surface in “basophilic erythroblasts” at about 2 days of incubation (Small and Davies, 1972), with complete MBs in “polychromatophilic erythroblasts” at 3 days. More recently, anti-tubulin immunofluorescence has been used to study MB biogenesis in chicken erythroblasts of both the primitive (Kim et al., 1987) and definitive series (Murphy et al., 1986), the latter using bone marrow of chicks made anemic by phenylhydrazine. In this case, MTs were initially organized radially about centrosomes, were subsequently reorganized into cytoplasmic “wreaths” without centrosome association, and finally became tightly bundled MBs at the periphery. In immature primitive erythroblasts, centriole-containing MTOCs were observed at the cell periphery (Kim et al., 1987). Here, however, assembling MTs formed bundles running throughout the cytoplasm, but were not organized into wreaths prior to MB formation. Disagreement on this intermediate stage might be due to a real difference between primitive and definitive MB biogenesis in chicken erythrocytes, but it could also reflect differences in experimental procedures or in specific stages examined. Neither of these studies attempted direct correlation of cytoskeletal structure with cell morphology, so it is unclear whether cells containing the cytoplasmic MT bundles or the MT wreaths had just begun to flatten, or whether they were already flattened (discoidal). When these observations are compared with ones made on amphibian erythroblasts (Ginsburg et al., 1989), the picture becomes more complex. The axolotl larval spleen develops initially as a closed sac containing differentiating nucleated erythrocytes, including spheres, disks, and ellipses. However, it also contains a significant percentage of unusual singly or doubly pointed erythroid cells, with correspondingly singly and doubly pointed MBs. Similar pointed erythrocytes have been noted previously in the circulation of phylogenetically diverse species including the chicken (Lucas and Jamroz, 1961), skate (Cohen, 1986), and slender salamander (Cohen, 1982), but never in such large numbers. Many of the pointed splenic erythroblasts were found to contain a pair of centrioles close to a pointed end, suggesting MTOC function and the hypothetical sequence for MB biogenesis illustrated in Fig. 1 I (Ginsburg et al., 1989). Here, a centrosome nucleates MTs that ultimately form two bundles growing in opposite directions around the periphery (Fig. 1 la). The bundles meet to produce a

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

61

FIG. 1 1 . Proposed model for marginal band (MB) biogenesis and cellular morphogenesis in amphibian erythroblasts. Stages of microtubule (MT) initation and growth from an organizing center (a and b) are similar to those for blood clam MB reassembly (Fig. 9). The cells are initially spheroidal (a), with doubly pointed and singly pointed cells (b and c) following as intermediate stages. MB closure produces a discoidal cell (d). Adapted from Ginsburg et al. (1989), “Cellular morphogenesis and the formation of marginal bands in amphibian splenic erythroblasts,” Cell Motility and the Cytoskelefon U ,159-168, with permission of WileyLiss Div. of John Wiley & Sons, Inc.

doubly pointed cell (Fig. 1lb), followed by closure of the distal (Fig. 1Ic) and then proximal ends (Fig. 1Id). Closure could not involve annealing, because the meeting MT ends would all have the same polarity, but it might be effected by cross-bridging between MTs of the two bundles and their continued peripheral growth in opposite directions [as suggested for blood clam MB reassembly (Fig. 9)]. A critical step in testing this proposal would be the establishment of an in uitro amphibian erythroblast culture system in which the complete differentiation sequence could be followed in individual cells. Although this has not yet been accomplished, the production of singly and doubly pointed cells during amphibian erythrogenesis has been verified in recent time-course studies of Xenopus during recovery from phenylhydrazine-induced anemia (Twersky et al., 1990) (Fig. 12). In considering MB biogenesis, it is important to remember that there may be different modes of MB construction in different species. Thus, intermediate stages in amphibians may prove to be dissimilar to those in chickens, with the end products (i.e., MBs) nevertheless structurally and functionally equivalent. Ultimate formation of a mechanically functional MT bundle is a common feature of MB biogenesis, regardless of the particular structural path taken. The properties of a potential erythrocyte MT-bundling protein such as syncolin are thus of particular interest. In dividing chicken erythroblasts, syncolin distribution differed from that of tubulin. However, it became associated with MTs of the forming MB at later stages, and was localized exclusively to the MB of mature cells (Feick et al., 1991). As noted by Feick e f al., it remains to be determined whether syncolin is an

62

WILLIAM D. COHEN

FIG. 12. Examples of pointed cells appearing in large numbers in the circulation of Xenopus laeuis during recovery from experimentally induced (phenylhydrazine) anemia. (a) Doubly pointed; (b) singly pointed with a long point; (c) singly pointed with a shorter point. Such cells are rarely observed in normal animals. Phase contrast video microscopy of glutaraldehyde-fixed cells. From Twersky et a / . (19%).

initial causal factor in the bundling of erythroblast MTs or whether it functions to further stabilize existing MT bundles. Study of erythrocyte tau distribution during MB biogenesis would similarly shed some light on its possible role as a bundling protein, and clarify its temporal relationship to syncolin binding. In both splenic amphibian erythroblasts and chicken primitive-series erythroblasts (Ginsburg et al., 1989; Kim et al., 1987),centriole-containing MTOCs are located closer to the cell periphery than in many other cell types. These MTOCs presumably function in MB biogenesis both to nucleate MTs and to influence their spatial distribution, as appears to be the case during MB reassembly in blood clam erythrocytes (Nemhauser et al., 1983). MTs nucleated by centrosomes in interphase cells exhibit steadystate “dynamic instability” (e.g., Mitchison and Kirschner, 1984; Schulze and Kirschner, 1986), with some growing at the same time that others rapidly disassemble. A mechanism can thus be envisioned in which peripheral MTOC localization reduces the angle at which some growing MTs approach the cell surface, permitting them to follow the surface contour and to make lateral stabilizing contacts with it. These MTs would continue to elongate at the expense of others that had grown in a different direction (i.e., toward the cell interior) and remained dynamically unstable. With respect to additional molecular mechanisms involved in MB biogenesis, synthesis of MB-specific cp6 tubulin is coincident with that of hemoglobin (Murphy et al., 1986). Nevertheless, the cp6 isotype can copolymerize with other tubulins in uitro and can participate in the formation of interphase MT arrays and functional mitotic spindles after transfection into living cultured cells that normally do not contain it (Joshi et al., 1987; Baker et al., 1990). Conversely, assembly of non-MB (brain) tubulin within MT-depleted chicken erythrocyte cytoskeletons produces MB-like structures containing very long MTs (Swan and Solomon, 1984). Questions thus arise as to how and why the divergent cp6 isotype is produced at the time of MB assembly.

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

63

Assessment of the significance of MB-specific tubulin isotypes demands consideration of the real environment in which the erythrocyte cytoskeleton functions. Assembly of MB-like structures from brain tubulin in cytoskeletons in uitro (Swan and Solomon, 1984), or even of incorrect isotypes into a normal-looking MB in a living cell (if achieved), tells us little about functional properties of the cell. Even mature erythrocytes with no MB have relatively normal morphology (Fig. 7, cells 4b and 4d) when left unperturbed! The significance of cp6 may well lie in the mechanical requirements for long, stable MTs that can effect cell flattening, and ultimately provide critical cytoskeletal responses in the bloodstream (Fig. 10). This is in accord with the suggestion that MTs with reduced cp6 content are less stable; while they may help generate cell flattening, their eventual disassembly would leave more stable cp6-containing MTs to constitute the mature MB. This could account, in part, for reduced MT numbers in the mature cells of some species (Baker et al., 1990). The real question, then, is whether MBs containing incorrect isotypes would be fully functional in the living animal, since even slight alterations of MB mechanical responsiveness might lead to lethal blockage of capillaries. Similarly, incorporation of cp6 into normal-looking, functioning mitotic spindles in cultured cells tells us little about true function. Incorrect mitotic distribution of only an occasional chromosome during early development in the living animal could be lethal. Thus, the consequences of natural selection with respect to both erythroblasts and mitotic cells would be strict developmental programming and the evolutionary conservation of tubulin isotypes. Is this testable? Developments in viral transfection of chicken erythroblast lines, identification and manipulation of tubulin genes, and achievement of terminal erythrocyte differentiation in culture suggest that in vitro production of nucleated erythrocytes containing variant tubulins in their MBs may soon be possible. Initially, experimental comparison could be made between their mechanical/rheological properties and those of normal cells in uitro. The ultimate goal, however, would be functional comparison of cells with normal versus genetically manipulated cytoskeletal systems in the living animal, either by transfusion of in uitro-produced cells into the bloodstream or by engineered production of genetically variant erythrocytes during otherwise normal development. The mechanism by which MB biogenesis causes cell flattening is a separate unresolved problem. As a point of departure, the relatively simple hypothetical model illustrated in Fig. 13 can be considered. Here, the forming MB is depicted as a tight multiply wound coil that gradually expands outward like the spring winding mechanism of a clock. Such expansion does not require MT growth; rather, the release of bending strain by the initial coil drives the expansion. The MT slides along itself as

64

WILLIAM D. COHEN b

a

tight coil expanded coi I FIG. 13. An “expanding coil” mechanism as a possible basis for marginal band (MB) function during erythrocyte flattening. In this structurally oversimplified version, an initially tight microtubule (MT) coil (a) expands outward (b) by the release of bending strain, establishing a plane of flattening. A mechanism of this type would require only mechanical properties already attributed to the MB of mature cells, as discussed in the text.

the diameter of the ring increases, without requiring motile cross-bridges. Reduction in the number of MTs appearing in MB cross sections, observed in some species, is a natural consequence of the process. As presented in pure form in Fig. 13, the model is no doubt unrealistic. MT distribution during MB biogenesis in chick or amphibian erythroblasts is not a neat tight coil, but rather a looser arrangement of curved MT bundles, or a wreath, as noted earlier in this section. However, the same basic mechanism might be at work, with curved MTs or MT bundles pressing outward until meeting MS resistance. Once such pressure began to establish a plane of flattening, it is likely that all additional MTs would find their way into the same plane, since this plane would permit the greatest outward movement and maximal MT straightening. One appeal of such a mechanism is simplicity: It requires only mechanical properties already attributed to the MB of mature cells, with no need to invoke predetermined peripheral “guidetracks.”

B. ELLIPTOGENESIS Little is known about the mechanism by which the discoidal MBcontaining cell becomes elliptical. Isolated MBs tend to assume either a circular or slightly elliptical configuration (Fig. 6c), whereas MBs are usually highly elliptical in situ (Fig. 6a). It is unlikely that isolated MB

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

65

circularization involves loss of MB structure, because it occurs regardless

of the particular isolation medium or technique used (Bertolini and Monaco, 1974; Cohen, 1978b; Waugh and Erwin, 1989; Sanchez et al., 1990). This indicates that the MB alone does not generate ellipticity. In our model (Fig. 4), ellipticity is imposed by the MS through application of force asymmetrically across the flexible MB. Experiments and calculations by Waugh and Erwin (1989) indicate that a force of -12.5 x dyn is needed to deform a circular newt MB into an appropriately elliptical one. They determined that sustained support of this force by the nucleated erythrocyte membrane (presumably, the MS) would be possible if the membrane had greater rigidity than that of the mammalian definitive erythrocyte [shear modulus ( p ) = 0.07 dyn/cm versus -0.01 dyn/cm]. If not, then a mechanical contribution to ellipticity by other cytoskeletal elements, possibly IFs, would be required. Actually, membrane microaspiration experiments yielded a p value of 0.07 dyn/cm or greater for nucleated erythrocytes of representative bony fish, amphibians, reptiles, and birds (Waugh and Evans, 1976),but the extent to which MB presence influenced the data is unknown. Clarifying information might well be obtained by repeating microaspiration studies on experimentally produced erythrocytes lacking MBs versus ones containing MBs ( Joseph-Silverstein and Cohen, 1984). A direct experimental approach to the mechanism of elliptogenesis is inherent in the work of Dorn and Broyles (1982). These workers used density gradients to separate developing erythrocytes during metamorphosis in the bullfrog Rana catesbiana, obtaining essentially pure preparations of disks versus ellipses for hemoglobin analysis. Similar preparations should be valuable for comparing cytoskeletal molecular components before and after elliptogenesis, and possibly for attempts at obtaining synchronous elliptogenesis in uitro. With respect to the latter, it is important to note that the mechanisms involved in both flattening and elliptogenesis are intrinsic; they do not require external forces generated in the bloodstream. This is demonstrated by the terminal differentiation of avian erythroblasts in cultures (Beug ef al., 1982)and of larval amphibian erythroblasts confined within the developing spleen in uiuo (Ginsburg et al., 1989). Thus, nucleated erythrocyte morphogenesis anticipates functional requirements for flow conditions. C. MATURATION The term “maturation” encompasses the emergence of several cytoskeletal properties, one of which is retention of cell shape at low temperature in species that have cold-labile MBs (Fig. 7, cell 444b). It is not

66

WILLIAM D.COHEN

known whether such stabilization follows elliptogenesis or is concomitant with it, but “setting” of the MS to differentiated morphology is apparently involved (Barrett and Scheinberg, 1972; Barrett and Dawson, 1974; see Section V,C). In other species, MBs of mature cells are cold stable. The basis for such differences has not really been explored, but lowtemperature lability of dogfish and blood clam MBs (Fig. 7, cell 4+4a) shows that there is no correlation with homeothermy. Tubulin isotype composition may be a contributingfactor, as indicated in work on chicken erythrocytes (Joshi et al., 1987). Another characteristic of maturity in at least some species is reduction in the number of MTs per MB cross section. In chicken erythrocytes, for example, the number diminishes from -50 (average)in early erythroblasts to 10 in mature cells (Small and Davies, 1972). Such reduction does not occur universally, however; early erythroblasts of the salamander Triturus cristatus contain about the same number (average, 109) as mature erythrocytes (Small and Davies, 1972). A third feature of maturation is that, in appropriate species, essentially all MTs reassemble into a peripheral MB upon rewarming following lowtemperature MB disassembly (Fig. 7, cell 4a+4c). That this occurs in the original plane of flattening can be inferred from the absence of morphological changes in the cells during reassembly (Cohen et al., 1982a). In contrast, at early stages of MB biogenesis in immature cells, not all MTs reassemble in their initial locations (Kim et al., 1987). One explanation adduced for MT reassembly in the original plane of flattening is that factors controlling spatial organization reside in that plane near the cell periphery. This possibility has been examined in vitru by depleting chicken erythrocyte cytoskeletons of native tubulin at low temperature, then reassembling MTs within them using MAP-free brain tubulin (Swan and Solomon, 1984). MTs were found to be localized at or near the cytoskeleton periphery in MB-like fashion. However, such localization in itself does not demonstrate the presence of peripheral determinants unequivocally. MT coils closely resembling MBs can be induced to assemble in synaptosomes (Gray et al., 1982),artificially produced cell fragments that do not normally contain such coils, and in which the presence of preorganized peripheral determinants is highly unlikely. Peripheral MT localization in both cases is more simply explained as a mechanical effect of MT elongation coupled with resistance to bending (see Fig. 13 and Section IV,A). The presence of spatial determinants in nucleated erythrocytes is indicated more convincingly by the finding that the total length of reassembled MTs (i.e., the number of MT windings) was independent of tubulin concentration (Swan and Solomon, 1984).Since not all MB MTs are in contact with the MS, this result implies either that determinants are present both at the MS interface and deeper within the extracted cytoskeleton, or that more localized pe-

-

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

67

ripheral determinants control both MT location and length (or, number of windings). One additional feature of mature erythrocyte morphology is calcium lability. Exposure of mature flattened elliptical cells to the ionophore A23187 rapidly converts them to spheres (Fig. 7, cell 4-4e). This occurs both in Xenopus (Gambino et al., 1985), a species with cold-stable MBs, and in the smooth dogfish (Bartelt, 1982),a species with cold-labile MBs, accompanied by loss of much cytoskeletal structure and, in dogfish erythrocytes, ultimate cell lysis. Experimentally induced lysis of Xenopus erythrocytes in the presence of Ca2+,or exposure of erythrocyte cytoskeletons to a Ca2+-containingcytosolic fraction, causes rapid depolymerization of MB MTs. This effect appears to require calmodulin and other as yet unidentified cytosolic factors (Gambino et al., 1985). It is noteworthy that the presence of calmodulin has been demonstrated in dogfish erythrocyte extracts (Bartelt et al., 1982).In Xenopus erythrocytes, Ca2+induces the partial proteolytic degradation of spectrin, suggesting that selective proteolysis of major cytoskeletal proteins may be the cause of Ca2+ ionophore-induced conversion of mature ellipsoids to spheres (Gambino et al., 1985). Similar conversion of mature nucleated erythrocytes to spheres is induced by prolonged contact with surfaces such as slides or coverslips. It is not known how rapidly such contact initiates cytoskeletal changes. Thus, the interpretation of data reported by many laboratories is complicated by the fact that living erythroid cells have been allowed to attach to surfaces prior to experimental treatments (e.g., Granger and Lazarides, 1982; Gambino et al., 1984; Murphy et al., 1986; Joshi et al., 1987; Kim et al., 1987; Koury et al., 1987). V. The Membrane Skeleton

The MS of nucleated erythrocytes has been less thoroughly studied than that of nonnucleated definitive mammalian erythrocytes. In earlier work (Cohen, 1978b), the noncommittal term “trans-MB material” was applied to a rough network spanning the MB in TEM whole mounts of nucleated erythrocyte cytoskeletons. Although it was suggested that this network might contain actin and spectrin-like molecules, it was initially assumed to contain cytoplasmic elements as well, collapsed onto the planar grid substrate. Subsequently, it became clear that most of this material was associated with the plasma membrane external to the MB, and the terminology was refined to cell “surface-associated cytoskeleton (SAC).” At the same time, “membrane skeleton” came into wider use for the corresponding

68

WILLIAM D. COHEN

structure in mammalian erythrocytes, and thus “MS” is adopted in this review as generic. A. STRUCTURE

The paradigm here is the MS of mammalian definitive erythrocytes (Fig. 14), essentially their entire cytoskeletal system. In stretched-out nega-

tively stained MS preparations, details of the spectrin-actin network are visible (Byers and Branton, 1985), and in deep-etch replicas the mammalian erythrocyte MS appears as a relatively homogeneous interwoven fabric of spectrin, actin, and associated proteins (Coleman et al., 1989). Comparable high-resolution views of the MS in nucleated erythrocytes of nonmammals have not yet been obtained, but fundamental structural equivalence is generally assumed based on similarities in protein composition and function. In intact nucleated erythrocytes, the exterior MS surface is believed to interface with the plasma membrane bilayer throughout the cell, with its interior surface contacting the MB in the plane of cell flattening. The MS is not visualized directly in thin sections of whole nucleated erythrocytes, in which it is obscured by electron-dense hemoglobin, but its presence is indicated by the fact that MB MTs are never found in direct contact with the membrane bilayer. The MS is readily visible, however, in thin sections of cells simultaneously detergent-lysed and fixed, or in TEM whole

--

FIG. 14. General molecular structure of the membrane skeleton (MS), as derived principally from studies of mammalian erythrocytes. The MS is essentially a spectrin ( S ) tetramer network, with foci composed of actin (a) oligomers and band 4.1 protein. The MS is bound to and spectrinthe membrane bilayer through interaction between 4.1 and glycophorin (G), bound ankyrin (ank) and the anion transporter (AT). The general model is thought to apply to the MS of nucleated erythrocytes of nonmammalian vertebrates as well, with variation in spectrin subunits (af-p). as discussed in the text.

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

69

mounts (e.g., Cohen et al., 1982a, 1990).The question arises as to whether the MS might contain a specialized structural/molecular “track” that guides MB reassembly and possibly MB biogenesis during erythrocyte differentiation. Indeed, Granger and Lazarides (1982), using shadowed replicas of sonicated chicken erythrocytes, made the intriguing observation that structural tracks resembling ridges between grooves were continuous with MTs at the inner plasma membrane surface. In itself, however, this is not compelling evidence that tracks exist, since they could be an artifactual imprint resulting from MS pressure against the MB during cell preparation and fixation. Thus, it remains to be determined whether such tracks are real by looking for them in erythrocytes predepleted of MTs at low temperature, or rewarmed to room temperature with MB reassembly blocked by colchicine or other inhibitors. COMPOSITION B. MOLECULAR The molecular components of the erythrocyte MS have been studied extensively in recent years, and details can be found in a number of excellent reviews (e.g., Geiger, 1983; Lazarides, 1987; Lazarides and Woods, 1989; Steck, 1989). The molecular organization of the mammalian erythrocyte MS (Fig. 14) is thought to be similar to that of the nucleated erythrocyte, since nonmammalian equivalents of spectrin (e .g., Jackson, 1975; Cohen et al., 1982a; Bartelt et al., 1982, 1984; Repasky et al., 1982; Glenney et al., 1982), actin, ankyrin (Moon and Lazarides, 1984), band 4.1 (Granger and Lazarides, 1984, 1985; Ngai et al., 1987), and band 3 (anion transporter) (Woods et al., 1986) are present. Terminology regarding the spectrin family proteins has been and continues to be somewhat confusing. Like other representatives, that of the nucleated erythrocyte is a heterodimer, with subunits of Mr -240,000 (a) and -220,000 (p), as observed on SDS-PAGE. The Mr 240,000 a subunit resembles the a subunit of mammalian nonerythroid spectrin (i.e., a-fodrin, or a-“brain spectrin”) (Levine and Willard, 1981) more closely than that of mammalian erythrocytes, and is here designated af.This was demonstrated by Bartelt et al. (1982, 1984) for dogfish erythrocytes on the basis of calmodulin binding and reactivity with antibodies, and by several other laboratories at about the same time (e.g., Repasky et al., 1982; Glenney et al., 1982). Nonmammalian erythrocyte afbinds calmodulin with high affinity, as does a-fodrin, but the a subunit of mammalian erythrocyte spectrin does not. Comparison of nucleotide/amino acid sequences has shown >90% homology between these calmodulin-binding a subunits, as compared with <60% for mammalian erythrocyte a-spectrin (McMahon et al., 1987). The MS of nonmammalian nucleated erythrocytes

70

WILLIAM D. COHEN

thus has more in common with the MS of eukaryotic cells in general than with that of definitive mammalian erythrocytes. Viewed another way, the mammalian erythrocyte MS appears to be a highly modified version of that of nucleated erythrocytes, reflecting the elimination of interactions with other types of cytoskeletal elements, and evolutionary modification for cell fluidity.

C. MS BIOGENESIS Biogenesis of the MS of nucleated erythrocytes has now been studied in some detail, as reviewed by Lazarides (1987) and by Lazarides and Woods (1989), making considerable experimental use of virally transformed chicken erythroblast cell lines (e.g., Beug et af., 1982). In this section, changes in some of the major MS proteins during nucleated erythrocyte differentiation are considered briefly in relation to cellular morphogenesis. Undifferentiated chicken erythroid precursor cells that are still mitotic have been found to produce only the spectrin family protein known specifically as fodrin, designated here as af-& The af and pf subunits are, respectively, M , -240,000 and M , -235,000, with the latter referred to as the y-spectrin subunit by some authors (Lazarides and Woods, 1989). Subsequently, spectrin isoform switching occurs, and erythrocyte &spectrin (M,-220,000) and pf are coexpressed, with the degradation of af-& and af-p heterodimers limiting their accumulation. Induction of terminal differentiation [increased hemoglobin concentration, flattening, and elliptogenesis (Beug et af., 1982)] markedly reduces the rate of pf synthesis and turnover of af-P heterodimers (Lazarides and Woods, 1989). Thus, by the time MB biogenesis begins in postmitotic spherical erythroblasts, spectrin of the af-/?type found in mature cells is already assembling within the MS. The af-p heterodimers are produced in excess, their accumulation in the MS apparently being regulated by stabilization at membrane binding sites (Lazarides and Woods, 1989). Actin in filamentous form is associated with the cortical regions of mitotic erythroblasts. During differentiation, actin filaments become restricted to the region of the MB in the plane of flattening (Kim ef al., 1987), while elsewhere actin associates with the MS as short oligomers. Despite the fact that actin is an important structural component of the MS, little information is currently available regarding the temporal and spatial control of actin assembly in nucleated erythrocytes. Synthesis of ankyrin and protein 4.1 occurs throughout erythrocyte differentiation, whereas synthesis and accumulation of the anion transporter (AT) begin upon induction of terminal differentiation (Koury et af., 1986, using mammalian cells). Since association of spectrin, ankyrin, and

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

71

protein 4.1 to form a transient MS begins earlier, it is believed that the anion transporter serves to stabilize the MS, but not to initiate its assembly. In the final stages of MS assembly in nucleated erythrocytes, synthesis and incorporation of both ankyrin and protein 4.1 continue, while diminishing for other molecular components. These two proteins are substoichiometric to spectrin in immature cells as compared with mature ones, leading Lazarides and Woods (1989) to suggest that final MS “mechanical stability” is achieved by completion of ankyrin and protein 4.1 incorporation. In addition, in early avian erythroid progenitor cells, a protein 4.1 variant lacking the spectrin-actin binding domain predominates, and this may contribute to limited crosslinking and instability of synthesized MS components. During terminal differentiation, synthesis and assembly of protein 4.1 variants containing the spectrin-actin binding domain occur (Lazarides and Woods, 1989), potentially important for stabilization of the MS and for providing its elastic memory during cell maturation (Fig. 7; see also next section). Conversely, earlier presence of the 4.1 variant that limits crosslinking of MS components, together with substoichiometric ankyrin, might facilitate remodeling of the immature MS (and thereby cell shape) on the assembling MB frame. PROPERTIES D. MECHANICAL In the discussion that follows, similarity between MS molecular components in nucleated nonmammalian vertebrate erythrocytes and mammalian erythrocytes is assumed to be paralleled by qualitatively similar mechanical behavior. Mechanical properties studied principally in the mammalian erythrocyte MS are thus treated as applicable to nucleated erythrocytes. Only selected MS properties relevant to cytoskeletal function in nucleated erythrocytes are considered here; further details can be found in other reviews (e.g., Steck, 1989; Mohandas et al., 1983). The MS, particularly its spectrin, is believed to be largely responsible for the elasticity of the erythrocyte membrane. Elastic materials usually consist of filaments crosslinked into networks, and they are reversibly deformable (Steck, 1989). The MS is thought to be held in a partially extended state in the intact membrane by interaction with the bilayer, so that removal of lipids by detergents produces elastic contraction (i.e., shrinkage) of the mammalian MS (Steck, 1989). Although direct observations on MS molecular spacing are lacking for nucleated erythrocytes, the twisted figure-eight geometry of cytoskeletons prepared by detergent lysis indicates similar MS elasticity (see also Fig. 6b and Section 111,B). Figureeight twisting appears to be an accommodation of the MB to elastic contraction of the MS within which it is confined, since MS dissolution

72

WILLIAM D. COHEN

releases planar MBs from twisted cytoskeletons. In figure-eight cytoskeletons, the MS itself assumes a saddle-shaped configuration (hyperbolic paraboloid) in which surface area is reduced versus the more planar initial state (Cohen, 1978b). This shape approaches minimum energy and area, and can be demonstrated in soap films adhering to figure-eight wire models. Ultrastructural evidence of MS elasticity in nucleated erythrocyte cytoskeletons is difficult to obtain because the nucleus distorts and obscures the critical central MS region. However, this problem can be circumvented through use of the exceptional erythrocytes of Batrachoseps salamanders, in which all circulating cells contain MB and MS, but more than 90% are anucleate (Cohen, 1982). In the figure-eight anucleate cytoskeletons, the MS is found to be contracted adjacent to the crossover area (Fig. 15b), as compared with untwisted cytoskeletons (Fig. 15a). Elastic deformation of erythrocyte membranes occurs when applied forces are of relatively low magnitude and short duration (lop6dyn and
CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

73

FIG. 15. The cytoskeletal system of the unusual anucleate erythrocytes of Batrachoseps salamanders. (a) As seen in whole-mount by transmission electron microscopy, it consists essentially of the marginal band (MB) and the membrane skeleton (MS). (b) In figure-eight twisted anucleate cytoskeletons, increased MS density indicates that it is contracted at or near the crossover region (arrows; cf. area further away). (Inset) Living erythrocytes of this species-one nucleated (N), the other anucleate-as observed in phase contrast. Modified from Cohen (1982); adapted from Protoplasma 113, 23-32, with permission of SpringerVerlag.

74

WILLIAM D. COHEN

FIG. 16. Summary of experiments on deformation of the membrane skeleton (MS) and alteration of cell shape, induced by marginal band (MB) discontinuityin dogfish erythrocytes. t T, Physiological temperature (20-22°C); & T, 0°C; col, colchicine. See text for details.

In summary, the MS of the mature nucleated erythrocyte resembles an elastic molecular fabric with a memory for flattened elliptical cell shape. During the morphogenetic sequence (Fig. 71, however, the MS is presumed to be far more dynamic, undergoing changes in molecular composition as well as deformation by the assembling MB.Since disks of at least some species (Fig. 7, cell 2) revert to spheres upon MB disassembly (Fig. 7, cell 2+2b), the disk-stage MS either retains memory for the spherical shape or, more likely, is highly plastic and spontaneously assumes a spheroidal low-energy state. Upon completion of elliptogenesis, the MS becomes “set” to the differentiated shape, presumably by mechanisms

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

75

involving changes in protein assembly and composition as discussed in the previous section (V,C). M. Intermediate Filaments

IFs were first reported in nucleated erythrocytes by Virtanen et al. (19791, who observed them attached to the nucleus in chicken erythrocytes and speculated that they served a nucleus-anchoringfunction. The IFs are of the vimentin class, interlinked by the protein synemin (Granger and Lazarides, 1982). They appear to loop out from the nucleus, attaching along their length to the adjacent MS region on both sides (Granger and Lazarides, 1982; Lazarides, 1987). This fits well with the finding by Georgatos and Blobel (1987a,b) that the nuclear envelope of turkey erythrocytes contains lamin B binding sites for the carboxy-terminal vimentin domain, sites from which IFs are assembled. In addition, the plasma membrane (presumably, the MS) binds the amino-terminal vimentin domain at sites that do not function in IF assembly. However, these data do not account for IFs that appear to bypass the nucleus, attaching to the MS at both ends (Granger and Lazarides, 1982).Candidate MS proteins for the IF-binding function are ankyrin (Georgatos and Marchesi, 1985) and certain variants of protein 4.1 (Lazarides, 1987). It is not known whether vimentin IFs are a universal feature of the nucleated erythrocyte cytoskeleton, but they have been found in erythrocytes of species quite divergent from the chicken, such as amphibians [e.g., Xenopus (Gambino et al., 1984)l. Similarly,it is unclear whether IFs play any critical role in either establishment or maintenance of the flattened elliptical shape. Since erythrocytes of essentially the same morphology, but lacking nuclei, are the normal predominant type in several salamander genera (Emmel, 1924; Cohen, 1982; Villolobos et al., 1988), anchoring of IFs to the nucleus cannot be important for cytoskeletal form or function in at least these species. Anucleate ghosts, produced by fluxing hypoosmotically swollen dogfish erythrocytes through syringe needles, also frequently retain their normal flattened elliptical shape (Cohen et al., 1982a). While there is good evidence that IFs are attached to the nucleus and the MS in species such as the chicken, there appears to be no direct attachment to the MB (Granger and Lazarides, 1982). Assessment of the functional contribution of IFs to the nucleated erythrocyte cytoskeletal system would benefit from the production of living erythrocytes of a given species with and without IFs, or with IFs of altered composition. This cannot be done by simple in uitro treatment with physical or chemical agents because of IF stability, but may become possible through manipulation of vimentin genes (e.g., Ngai et al., 1987).

76

WILLIAM D. COHEN

VII. The Cytoskeletal System of Mammalian Primitive Erythrocytes

As noted in Section I, the first, or primitive-generation, erythrocytes of developing mammals originate in the yolk sac. They are nucleated and significantly larger than the definitives that appear subsequently. In an early brief paper, van Deurs and Behnke (1973) reported the presence of MBs in nucleated erythrocytes observed in thin sections of mouse, rat, and human embryonic liver. However, Koury et al. (1987), using a variety of techniques, concluded that true continuous MBs were not present in primitive erythrocytes obtained from the circulation of mouse embryos at a comparable stage of development. Attempting to clarify this issue, we have taken a simplified approach by utilizing marsupials (Cohen et al., 1990). The young of these mammals are born in an underdeveloped state, with nucleated primitive erythrocytes constituting essentially all of the circulating blood cell population (Block, 1964; Richardson and Russell, 1969; Yadav, 1972;Cutts et al., 1980; Holland et al., 1988).Thus, material is obtained without invasion of the mother, and problems associated with developmental timing are largely avoided. Two marsupial species used extensively in current biomedical research were studied: the tammar wallaby (Macropus eugenii) (Tyndale-Biscoe and Janssens, 1988), a small member of the kangaroo family, and the gray short-tailed opossum (Monodelphis dornestica) (VandeBerg, 1990). In both species, nucleated primitive erythrocytes were found to constitute nearly 100% of the circulating population at birth (i.e., day 0), as anticipated. Cell morphology was similar to that of nonmammalian vertebrate erythrocytes, except for reduced ellipticity (Figs. 2 and 17). Correspondingly, the primitive erythrocyte cytoskeletal system was essentially indistinguishable from that of nonmammalian vertebrate erythrocytes, consisting of MB and MS as observed by anti-tubulin immunofluorescence and in TEM whole mounts and thin sections (Cohen et al., 1990) (Fig. 17). In fetal blood samples from the tammar wallaby, singly pointed nucleated primitive erythrocytes were a significant component, with a few doubly pointed ones also present. Singly pointed cells were observed in smaller number in both species to day 2, but not thereafter. This is consistent with the studies of amphibian erythroblasts discussed in Section IV,A, suggesting that doubly and singly pointed cells are immature stages that parallel stages ofMB biogenesis (Ginsburg et al., 1989;Twersky et al., 1990) (Figs. 11 and 12). The transition from primitive to definitive generation erythrocytes began shortly after birth in both species. By day 2 or 3, -50% of the circulating population consisted of smaller anucleate definitives (lacking MBs), and this increased to over 90% after -1 week. In most blood

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

77

FIG. 17. Presence of the marginal bandhembrane skeleton (MB/MS) system in circulating primitive-generation erythrocytes of mammals during early development. Erythrocytes of marsupial neonates (Monodelphis dornestica, the gray short-tailed opossum). (a) Living erythrocytes in hanging drop, seen by bright-field microscopy. Cells are flattened disks or ellipsoids, with the nuclear bulge (n) visible in edge view. Morphological variants include singly pointed (p), twisted (arrow), and anucleate cells (not shown). (b) MBs of these cells visualized by anti-tubulin immunofluorescence. (c) Uranyl acetate-stained cytoskeleton whole-mount (transmission electron microscopy), with the MB, MS and nucleus (N)visible. From Cohen et al. (1990); reproduced from “The cytoskeletal system of mammalian primitive erythrocytes: Studies in developing marsupials,” Cell Motility and the Cytoskeleton 16, 133-145, by permission of Wiley-Liss Div. of John Wiley & Sons, Inc.

samples taken during this period, a different and intriguing erythrocyte type constituted 1-6% of the cells: anucleate erythrocytes of the same size and general shape as the nucleated primitives. These had been noted previously by several other workers (e.g., Cutts et al., 1980). In TEM whole mounts, their cytoskeletons looked strikingly like those‘ of Batrachoseps (Fig. 15), each consisting of MB and MS, and thus they were identifiable as anucleate primitive erythrocytes. Their percentage in the circulating population was maintained or even increased in the days after birth, while that of nucleated primitives declined. This suggests that anucleate primitive erythrocytes are derived from nucleated primitives, and that loss of erythrocyte nuclei in mammals begins earlier than is generally recognized (i.e., in the primitive generation). It is noteworthy

78

WILLIAM D. COHEN

that cells thought to be nonnucleated primitive erythrocytes (“megalocytes”) were described long ago in human embryos (Knoll, 1927, 1932). By what mechanism(s)might anucleate erythrocytes containing MBs be generated? Here, Batrachoseps and related salamanders can serve as a model. Emmel (1924) reported nonmitotic division of nucleated Batrachoseps erythrocytes in hanging drops and proposed an active division mechanism to account for formation of anucleate “erythroplastids” of various sizes in the blood. Variations of cytoskeletal structure observed in these cells were compatible with this proposal (Cohen, 1982), although division was not confirmed directly. Recently, Villolobos et al. (1988) discovered four other salamander genera with species exhibiting 80% or more anucleate erythrocytes in the circulation. As compared with related species having normal nucleated erythrocytes, all of those with anucleate erythrocytes had either wholly miniaturized bodies or greatly reduced bodies with miniaturized limbs. They also had a relatively high DNA content per haploid genome and free nuclei in blood smears. Villolobos et al. proposed that enucleation occurs in the circulation of Batrachoseps and related salamanders by “random cellular breakage,” as relatively large erythrocytes with large nuclei are forced through capillariek that are unusually narrow due to body miniaturization. Such a passive mechanism might well apply to marsupial neonates if reduction of capillary diameter accompanies production of the smaller definitive erythrocytes in the days after birth. This, as well as possible enucleation by nonmitotic division, remains to be investigated. In our present state of ignorance, a second type of active mechanism might equally well be entertained in which nuclei degenerate intracellularly. However, this would not explain the occurrence of what appear to be free nuclei in blood samples of opossum and wallaby neonates (Block, 1964; Cohen et al., 1990). The presence of the MB/MS cytoskeletal system in mammalian primitive erythrocytes confirms the long-suspected similarity between these cells and the nucleated erythrocytes of all nonmammalian vertebrates, reawakening thoughts of ontogeny recapitulating phylogeny (van Deurs and Behnke, 1973; Palmer, 1980). Whether MB-containing nucleated erythroc-ytes are functionally necessary at these early developmental stages is an open question, however. A proposed sequence for erythrocyte formation in developing marsupials is summarized in Fig. 18. The findings reinforce the original observations by van Deurs and Behnke (1973) on placental mammals and inspire speculation on several issues. True continuous MBs are formed during de3nitiue erythrogenesis in mammals of the camel family only, apparently serving transiently to generate the flattening and ellipticity of bone

CYTOSKELETALSYSTEMOFNUCLEATEDERYTHROCYTES

79

yolk sac i liver spleen bone marrow FIG. 18. Proposed sequence for erythrocyte formation in developing marsupials, summarizing data from several authors. Most circulating cells at birth are nucleated primitive erythrocytes, containing marginal bands (MBs) (a);before birth, pointed ones are frequently observed (b). Within a few days, definitive (anucleate) erythrocytes are produced by the liver and subsequently other organs ( Q). Data suggest that the loss of primitive erythrocyte nuclei produces circulating anucleate primitive erythrocytes containing MBs (O), the fate of which is unknown (?). From Cohen et al. (1990); reproduced from “The cytoskeletal system of mammalian primitive erythrocytes: Studies in developing marsupials, ” Cell Motility and the Cyroskeleton 16,133-145,with permission of Wiley-Liss Div. ofJohn Wiley & Sons, Inc.

marrow erythroblasts, then disappearing prior to or during maturation (Goniakowska-Witalinska and Witalinski, 1977; Cohen and Terwilliger, 1979; Repasky and Eckert, 1981 vs. Grasso, 1966). The mature circulating erythrocytes of these species are thus uniquely elliptical (Fig. 2, cell i). However, production of these MBs may not be a novel Camellidae invention, but rather the continued utilization or reactivation of biogenetic mechanisms originally active in the fetus. Similar considerations might apply to the oval or figure-eight fibrillar structures known as “Cabot’s rings,” found in the circulating nucleated erythrocytes of severely anemic humans (Cabot, 1903; Jordan et al., 1930), should they prove to be constructed of MTs. Finally, the presence of MBs in primitive mammalian erythrocytes encourages the search for erythroid regulatory elements and cytoskeleton-specific genes (e.g., MB-specific tubulins and MAPS) (Murphy et al., 1986; Wang et al., 1986) common to mammalian and nonmammalian genomes.

VIII. Concluding Remarks The past decade has seen increased exploitation of nucleated erythrocytes for basic studies of ubiquitous cytoskeletal elements. Considerable progress has been made in the structural, molecular, and mechanical characterization of the MB, MS, and IFs of these cells, and toward understanding their function. Nevertheless, several long-standing questions regarding cytoskeletal structure and biogenesis remain unanswered, and

80

WILLIAM D. COHEN

new ones have emerged. As is apparent from comparison of Figs. 3 , 8 , and 17, the MB/MS cytoskeletal system exhibits remarkable structural similarity in erythrocytes of highly diverse species. However, molecular species and domains having specific and universal functional significance for the MB/MS system are incompletely identified, and genetic control mechanisms for synthesis of the major cytoskeletalproteins of these cells remain to be explored. In addition, we are largely ignorant of the interactions among major components that ultimately produce a cytoskeletal system with properties greater than the sum of its parts. Their quantitative determination may become possible through genetic manipulation of cytoskeleta1 proteins, coupled with assay of the effects of such manipulation on mechanical and rheological properties of the living cells. With respect to biogenesis of the cytoskeleton, improved methods are needed for studying the morphogenesis of nucleated erythrocytes in vitro, to establish more precisely specific stages in the sequence and to understand the temporal and spatial mechanisms involved. Finally, the degree of molecular similarity between the nucleated erythrocyte cytoskeletal system in nonmammalian vertebrates and that of nucleated primitive erythrocytes in developing mammals is of particular interest, and study of cytoskeletal proteins and their genes with respect to the latter erythrocyte type is in its infancy. In all of these areas, continued experimental utilization of nucleated erythrocytes can be expected to enhance our understanding not only of erythrocytes and other blood cell types, but of cytoskeletal systems in general. ACKNOWLEDGMENTS I am indebted to all of my former and current students and collaborators for their innumerable contributions over the years. For direct input regarding selected aspects of this review, particular thanks are due to Dr. Ursula Euteneuer (University of California, Berkeley), Dr. Thomas Fischer (Aachen, Germany), Ivelisse Sanchez (Hunter College of CUNY), Dr. Laura Twersky (BloomfieldCollege, NJ), Dr. Richard Waugh (University of Rochester), and Dr. Gerhard Wiche (University of Vienna, Austria). Support from the National Science Foundation has made possible much of the author’s work discussed, and is gratefully acknowledged.

REFERENCES Andrew, W. (1965). “Comparative Hematology.” Grune & Stratton, New York. Baker, H. N., Rothwell, S. W., Grasser, W. A., Wallis, K. T., and Murphy, D. B. (1990). J . Cell Biol. 110,97-104. Barrett, L. A., and Dawson, R. B. (1974). Deu. Biol. 36,72-81. Barrett, L. A., and Scheinberg, S. L. (1972). J. Exp. Zool. 182, 1-14. Bartelt, D. C. (1982). Ph.D. dissertation. City University of New York, New York. Bartelt, D. C., Carlin, R. K., Scheele, G. A., and Cohen, W. D. (1982). J. Cell Biol. 95, 278-284,

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

81

Bartelt, D. C., Carlin, R. K., Scheele, G. A., and Cohen, W. D. (1984). Arch. Biochem. Biophys. 230, 13-20. Behnke, 0. (1970a). J. Ultrastruct. Res. 31,61-75. Behnke, 0 . (1970b). Int. Rev. Exp. Parhol. 9, 1-92. Bertolini, J. B., and Monaco, G. (1974). J. Ultrastruct. Res. 54,57-67. Beug, H., Doederlein, G., Freudenstein, C., and Graf, T. (1982). J . Cell. Physiol., Suppl. 1, 195-207. Birgbauer, E., and Solomon, F. (1989). J . Cell Biol. 109, 1609-1620. Bloch, E. H. (1%2).Am. J . Anat. 110, 125-153. Block, M. (1964). Ergeb. Anat. Entwicklungsgesch. 37,237-366. Bretscher, A. (1983). J. Cell Biol. 97,425-432. Briggs, E. A. (1936). Nature (London) W, 762. Byers, T. J., and Branton, D. (1985). Proc. Natl. Acad. Sci. U.S.A. 82,6152-6157. Cabot, R. C. (1903). J. Med. Res. 9, 15-18. Centonze, V. E., and Sloboda, R. D. (1986). Exp. Cell Res. 167,471-483. Centonze, V. E., Ruben, G. C., and Sloboda, R. D. (1984). Cell Motil. Cytoskeleton 6, 376-388. Centonze, V. E., Ruben, G. C., and Sloboda, R. D. (1985). Eur. J. Cell Biol. 39, 190-197. Chien, S. (1975). I n “The Red Blood Cell” (D. M. Surgenor, ed.), Vol. 11. Academic Press, New York. Cohen, W. D. (1976). J. Microsc. (Oxford) 108,221-226. Cohen, W. D. (1978a). Blood Cells 4,449-541. Cohen, W. D. (1978b). J. CeNBiol. 78,260-273. Cohen, W. D. (1982). Protoplasma 1l3,23-32. Cohen, W. D. (1986). Biol. Bull. (Woods Hole, Mass.) 171,338-349. Cohen, W. D., and Ginsburg, M. (1986). I n “Methods in Enzymology” (R. B. Vallee, ed.), Vol. 134, pp. 232-252. Academic Press, Orlando, Florida. Cohen, W. D., and Nemhauser, I. (1980). J. CellBiol. 86,286-291. Cohen, W. D., and Nemhauser, I. (1985). I n “Blood Cells of Marine Invertebrates: Experimental Systems in Cell Biology and Comparative Physiology” (W. D. Cohen, ed.), pp. 1-49. Liss, New York. Cohen, W. D., and Terwilliger, N. B. (1979). J. Cell Sci. 36,97-107. Cohen, W. D., Bartelt, D. C., Jaeger, R., Langford, G., and Nemhauser, I. (1982a). J . Cell Biol. 93,828-838. Cohen, W. D., Langford, G. M., and Sloboda, R. D. (1982b).Biol. Bull. (Woods Hole, Mass.) 164950-61. Cohen, W. D., Cohen, M. F., Tyndale-Biscoe, C. H., VandeBerg, J. L., and Ralston, G. B. (1990). Cell Motil. Cytoskeleton 16, 133-145. Coleman, T. R., Fishkind, D. J., Mooseker, M. S., and Morrow, J. S. (1989). Cell Motil. Cytoskeleton 12,225-247. Cutts, J. H., Krause, W. J., and Leeson, C. R. (1980). Blood Cells 6,55-62. Dantschakoff, W. (1908). Organ. Anat. H e f e 37,471-589. Dorn, A. R., and Broyles, R. H. (1982). Proc. Natl. Acad. Sci. U.S.A. 79,5592-55%. Emmel, V. E. (1924). A m . J. Anat. 33,347-391. Euteneuer, U.,Ris, H. R., and Borisy, G. G. (1985). Eur. J . Cell Biol. 37, 149-155. Fawcett, D. W., and Witebsky, F. (1964). 2.Zellforsch. 62,785-806. Feick, P., Foisner, R., and Wiche, G. ( 1 9 9 1 ) . J. Cell Biol. 112,689-699. Fischer, T. M. (1978). Blood Cells 4,453-461. Fischer, T. M., and Schmidt-Schonbein, H. (1977). Blood Cells 3,351-365. Fischer, T. M., Stohr-Liesen, M., and Schmidt-Schonbein, H. (1978). Science 202,894-896.

82

WILLIAM D. COHEN

Gaehtgens, P., Schmidt, F., and Will, G. (1981a). PJluegers Arch. 390,278-282. Gaehtgens, P., Will, G., and Schmidt, F. (1981b). Pfiuegers Arch. 390,283-287. Gambino, J., Weatherbee, J. A., Gavin, R. H., and Eckhardt, R. A. (1984). J. Cell Sci. 72, 275-294. Gambino, J., Ross, M. J., Weatherbee, J. A., Gavin, R. H., and Eckhardt, R. A. (1985). J . Cell Sci. 79, 199-215. Geiger, B. (1983). Biochim. Biophys. Acta 737,305-341. Georgatos, S . D., and Blobel, G. (1987a). J. Cell Biol. 105, 105-115. Georgatos, S. D., and Blobel, G. (1987b). J. Cell Biol. 105, 117-125. Georgatos, S. D., and Marchesi, V. T. (1985). J. Cell Biol. 100, 1955-1961. Ginsburg, M. F., Twersky, L. H., and Cohen, W. D. (1989). Cell Motil. Cytoskeleton 12, 1 59- 168. Glenney, J. R., Glenney, P., and Weber, K. (1982). Proc. Natl. Acad. Sci. U.S.A. 79, 4002-4005. Goniakowska-Witalidska, L. (1974). Bull. Acad. Pol. Sci., Ser. Sci. Biol. 22,59-65. Goniakowska-Witalidska, L., and Witalidski, W. (1976). J. Cell Sci. 22, 397-401. Goniakowska-Witalihska, L., and Witalidski, W. (1977). J. Zool. 181,309-315. Granger, B. L., and Lazarides, E. (1982). Cell 30,263-275. Granger, B. L., and Lazarides, E. (1984). Cell 37,595-607. Granger, B. L., and Lazarides, E. (1985). Nature (London)313,238-241. Grasso, J. A. (1966). Anat. Rec. 156, 397-414. Gray, E. G., Burgoyne, R. D., Westrum, L. E., Cumming, R.B., and Barron, J. (1982). Proc. R . SOC.London, Ser. B 216,385-396. Grundke-Iqbal, I., Iqbal, K., Quinlan, M., Tung, Y.-C., Zaidi, M. S., and Wisniewski, H. M. (1986). J. Biol. Chem. 261,6084-6089. Holland, R. A. B., Rimes, A. F., Comis, A,, and Tyndale-Biscoe, C. H. (1988). Respir. Physiol. 73,69-86. Ihara, Y., Nukina, N., Miura, R.,and Ogawara, M. (1986). J . Biochem. (Tokyo) 99, 18071810. Jackson, R. C. (1975). J. Biol. Chem. 250,617-622. Jordan, H. E., Kindred, J. E., and Beams, H. W. (1930). Anat. Rec. 46, 139-160. Joseph-Silverstein, J., and Cohen, W. D. (1984). J . Cell Biol. 98,2118-2125. Joseph-Silverstein, J., and Cohen, W. D. (1985). Can. J . Biochem. Cell Biol. 63,621-630. Joseph-Silverstein, J., and Cohen, W. D. (1986). Ann. N . Y. Acad. Sci. 466,444-446. Joshi, H. C., Yen, T. J., and Cleveland, D. W. (1987). J . Cell Biol. 105,2179-2190. Kanai, Y., Takemura, T., Oshima, T., Mori, H., Ihara, Y., Yanagisawa, M., Masaki, T., and Hirokawa, N. (1989). J . Cell Biol. 109, 1173-1184. Kim, S., Magendantz, M., Katz, W., and Solomon, F. (1987). J. Cell Biol. 104,51-59. Knoll, W. (1927). Denskscr. Schweiz. Naturforsch. Ges. 64. Knoll, W. (1932). In “Handbuch der allgemeinen Haematolgie” (H. Hirschfield and A. Hittmair, eds.). Urban and Schwarzenberg, Berlin. Koury, M. J., Bondurant, M. C., and Mueller, T. J. (1986). J. Cell. Physiol. 126,259-265. Koury, S. T., Repasky, E. A,, and Eckert, B. S. (1987). Cell Tissue Res. 249,69-77. Lazarides, E. (1987). Cell 51,345-356. Lazarides, E., and Woods, C. (1989). Annu. Rev. Cell Biol. 5,427-542. Lee, G. (1990). Cell Motil. Cytoskeleton 15, 199-203. Lee, G., Cowan, N., and Kirschner, M. (1988). Science 239,285-288. Levine, J., and Willard, M. (1981). J. Cell Biol. 90,631-643. Lewis, S . A., and Cowan, N. (1990). Nature (London)345,674. Lewis, S . A., Ivanov, 1. E., Lee, G. H., and Cowan, N. J. (1989). Nature (London) 342, 498-505.

CYTOSKELETAL SYSTEM OF NUCLEATED ERYTHROCYTES

83

Lichtenberg, B., Mandelkow, E.-M., Hagestedt, T., and Mandelkow, E. (1988). Nature (London) 334,359-362. Lieber, M. R., and Steck, T. L. (1982a). J . Biol. Chem. 257, 11651-11659. Lieber, M. R., and Steck, T. L. (1982b).J . Biol. Chem. 257, 11660-11666. Lieber, M. R., Hesse, J. E., Nickol, J. M., and Felsenfeld, G. (1987). J . Cell Biol. 105, 1055-1065. Lucas, A. M., and Jamroz, C. (1961). “Atlas of Avian Hematology,” U.S. Dep. Agric. Monogr. 25. U.S. Government Printing Office, Washington, D.C. Lux, S. E., John, K., and Karnovsky, M. (1976). J. Clin. Inuesr. 58,955-963. Maximow, A. (1909). Arch. Mikrosk. Anat. 73,444-561. McMahon, A. P., Giebelhaus, D. H., Champion, J. E., Bailes, J. A., Lacey, S., Camtt, B., Henchman, S. K., and Moon, R. T. (1987). Differentiation 34,68-78. Meves, F. (191I). Arch. Mikrosk. Anat. Enrwicklungsmech. 77,465-540. Miller, M., and Solomon, F. (1984). J. Cell Biol., Suppl. 99,70~-75s. Mitchison, T. J., and Kirschner, M. W. (1984). Nature (London)312,237-242. Mohandas, N., Chasis, J. A., and Shohet, S. B. (1983). Semin. Hernatol. 20,225-233. Monaco, G . , Salustri, A., and Bertolini, B. (1982). J . Cell Sci. 58, 149-163. Moon, R. T., and Lazarides, E. (1984). J. Cell Biol. 98, 1899-1904. Murphy, D. B., and Wallis, K. T. (1983a). J . Biol. Chem. 258,7870-7875. Murphy, D. B., and Wallis, K. T. (1983b). J. Biol. Chem. 258,8357-8364. Murphy, D. B., and Wallis, K. T. (1985). J. Biol. Chem. 260, 12293-12301. Murphy, D. B., Grasser, W. A., and Wallis, K. T. (1986). J. Cell Biol. 102, 628-625. Murphy, D. B., Wallis, K. T., Machlin, P. S., Ratrie, H., and Cleveland, D. W. (1987). J . Biol. Chem. 262, 14305-14312. Nachmias, V. T. (1980). J. Cell Biol. 86,795-802. Nachmias, V. T., Sullender, J., and Asch, A. (1977). Blood 50,39-53. Nemhauser, I., Joseph-Silverstein, J., and Cohen, W. D. (1983).J. Cell Biol. 96,979-989. Ngai, J., Stack, J. H., Moon, R. T., and Lazarides, E. (1987).Proc. Natl. Acad. Sci. U . S . A . 84,4432-4436. Palmer, J. G. (1980). Blood Cells 6, 63-64. Ralston, G. (1985). Ausr. J. Biol. Sci. 38, 121-129. Ram6n-Caja1, S. (1933). “Histology.” Wood, Baltimore, Maryland. Repasky, E. A., and Eckert, B. S. (1981). Anat. Embryol. 162,419-424. Repasky, E. A., Granger, B. L., and Lazarides, E. (1982). Cell 29,821-833. Richardson, B. J., and Russell, E. M. (1969). Aust. J. Exp. Biol. Med. Sci. 47,573-580. Rothwell, S . W., Grasser, W. A., and Murphy, D. B. (1986). J. Cell Biol. 102,619-627. Rowley, A. F., and Ratcliffe, N. A., eds. (1988). “Vertebrate Blood Cells.” Cambridge Univ. Press, Cambridge, England. Sanchez, I., and Cohen, W. D. (1990). J. Cell Biol. 111,239a. Sanchez, I., Twersky, L. H., and Cohen, W. D. (1990). Eur. J . Cell Biol. 52,349-358. Schulze, E., and Kirschner, M. (1986). J . Cell Biol. 102, 1020-1031. Selden, C. S., and Pollard, T. D. (1983). J. Biol. Chem. 258,7064-7071. Sloboda, R. D., and Dickersin, K. (1980). J. Cell Biol. 87, 170-179. Small, J. V., and Davies, H. G . (1972). Tissue Cell 4,341-378. Steck, T. L. (1989). In “Cell Shape: Determinants, Regulation, and Regulatory Role.” Academic Press, San Diego, California. Swan, J. A., and Solomon, F. (1984). J. Cell Biol. 99,2108-2113. Twersky, L. H., Moogan, T., and Cohen, W. D. (1990). J. Cell Biol. 111,27a. Tyndale-Biscoe, C. H., and Janssgns, P. A. (1988). In “The Developing Marsupial: Models H. Tyndale-Biscoe and P. A. Janssens, eds.), pp. 1-7. for Biomedical Research” Springer-Verlag, Berlin.

(c.

84

WILLIAM D. COHEN

Usami, S., Magzinovic, V., Chien, S., and Gregersen, M. I. (1970). Microuasc. Res. 2, 489-499. VandeBerg, J. L. (1990). Ausf. J . Zool. 37,235-255. van Deurs, B., and Behnke, 0.(1973). Z. Anat. Entwicklungsgesch. 143,43-47. Villolobos, M., Leon, P., Sessions, S. K., and Kezer, J. (1988). Herperologica 44,243-250. Virtanen, I . , Kurkinen, M., and Lehto, V.-P. (1979). CellEiol. Int. Rep. 3, 157-162. Wang, D., Villasante, A., Lewis, S. A,, andcowan, N. J. (1986).J . CellEiol. 103,1903-1910. Waugh, R. E., and Erwin, G. (1989). J. Cell Biol. 108, 1711-1716. Waugh, R., and Evans, E. A. (1976). Microuasc. Res. 12,291-304. Waugh, R. E., Erwin, G., and Bouzid, A. (1986). J. Biol. Mech. Eng. 108,201-207. Weinreb, E. L., and Weinreb, S. (1965). Z. Zellforsch. Mikrosk. Anar. 68,830-836. White, J. G., Radha, E., and Krumwiede, M. (1986). Blood 67,873-877. Wiche, G., Oberkanins, C., and Himmler, A. (1991). Znt. Rev. Cytol. W, 217-273. Woods,C. M.,Boyer,B.,Vogt,P. K., andLazarides,E. (1986).J. CellEiol. 103,1789-1798. Yadav, M. (1972). Aust. J . Zool. 20,249-263.

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 130

Structure of the Mouse Egg Extracellular Coat, the Zona Pellucida PAULM. WASSARMAN AND STEVENMORTILLO Department of Cell and Developmental Biology, Roche Institute of Molecular Biology, Roche Research Center, Nutley, New Jersey 07110

I. Introduction Virtually all unfertilized eggs are surrounded by one or more extracellular coats (Dumont and Brummett, 1985). The zona pellucida (ZP) is a thick extracellular coat that surrounds the plasma membrane of all mammalian eggs. Accordingly, during the process of fertilization in mammals, sperm must bind to and penetrate through the ZP in order to reach and fuse with egg plasma membrane (Gwatkin, 1977; Wassarman, 1987a,b, 1988a, 1991; Yanagimachi, 1988). The ZP is responsible for both species specificity of sperm binding and establishment of a slow block to polyspermy following gamete fusion. In addition, in at least some mammals, the ZP induces bound sperm to undergo a form of exocytosis, the acrosome reaction. Preimplantation development takes place within the ZP, from which expanded blastocysts hatch just prior to implantation in the uterus (McLaren, 1982; Hogan et al., 1986; Perona and Wassarman, 1986). The ZP protects the cleaving embryo as it is transported along the female reproductive tract. Thus, the ZP performs several important functions during its rather brief existence. Over the years, the ZP has received a great deal of attention from biologists. Some of this stems from the fact that, for many experiments, the ZP has proved to be a nuisance. Consequently, various methods have been devised to gently remove the ZP from eggs and embryos (Hogan et al., 1986). In a more positive vein, the ZP also has received a significant amount of attention from investigators interested in its structure and function (Dunbar, 1983; Dunbar and Wolgemuth, 1984; Dietl, 1989). This interest extends back to the turn of the century, when a debate over the cellular origin of the ZP began. Finally, the ZP has received considerable attention as a potential target for development of a contraceptive vaccine (Sacco and Yurewicz, 1989;Dean et al., 1991).Thus, as a research subject, the ZP is covered by a rather extensive and diverse literature. Recently, a monograph (Dietl, 1989) appeared devoted solely to the ZP, suggesting that this unique organelle may have gained the status it deserves. 85

Copyright 0 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

86

PAUL M. WASSARMAN AND STEVEN MORTILLO

As indicated, research on the ZP has been both extensive and diverse, and an attempt is not made to provide encyclopedic coverage of the subject here. Rather, the focus is particularly on experimental approaches that have led to advances in our understanding of the mouse egg ZP structure. Collectively, these approaches, which range from light and electron microscopic to biochemical and molecular genetic studies of the ZP, have provided a much clearer (albeit, low-resolution) picture of the organelle than was available only 10 years ago. As expected, this information has provided insight into the relationship between ZP structure and function. Based on available evidence, it is likely that conclusions drawn from work on the mouse ZP will apply as well to the ZP from other mammalian eggs.

11. Functions of the Zona PeUucida A. FUNCTIONS DURING FERTILIZATION

Free-swimming sperm must bind to and penetrate the ZP in order to fertilize eggs (Phillips, 1991; Drobnis and Katz, 1991)(Fig. 1). Fertilization in mammals is relatively species-specific, and the species specificity is attributable, at least in part, to sperm receptors located in the egg ZP (Gwatkin, 1977; Bedford, 1981, 1982; Wassarman, 1987a,b, 1990; Yanagimachi, 1978, 1984, 1988). When ovulated eggs from one mammal are exposed to sperm from another mammal in uitro,rarely do the sperm bind to and penetrate the ZP. However, in many cases, removal of the ZP from ovulated eggs (i.e., removal of species-specific sperm receptors) eliminates the barrier to fertilization by sperm from heterologous species. Furthermore, in many cases, it is clear that only sperm with an intact plasma membrane overlying their head (“acrosome-intact” sperm) bind to receptors in the ZP (Fig. 2), and such binding induces the sperm to undergo exocytosis (“acrosome reaction”) (Bleil and Wassarman, 1983; Wassarman, 1987a,b, 1989; Kopf and Gerton, 1991)(Fig. 3). Sperm must undergo the acrosome reaction in order to be able to penetrate the ZP and fuse with egg plasma membrane.

B. FUNCTIONS AFTER FERTILIZATION Following fertilization of the egg by a single sperm, the ZP undergoes changes that collectively constitute the so-called “zona reaction” and are responsible for instituting the so-called “slow” block to polyspermy (Gwatkin, 1977;Wassarman, 1987a,b;Yanagimachi, 1988).The zona reaction takes place as a result of the “cortical reaction.” The latter involves

STRUCTURE OF ZONA PELLUCIDA

87

FIG. 1. Light micrograph (Normarski differential interference contrast) of mouse sperm “bound” to an unfertilized mouse egg ZP during culture in uitro. “Attached” sperm were removed from the egg by gentle pipeting with a drawn-out mouth-operatedglass micropipet.

fusion of egg cortical granules with plasma membrane and deposition of cortical granule contents into the perivitelline space (Gulyas, 1980; Ducibella, 1991). The contents, which includes several enzymes, diffuse into the ZP and modify both ZP2 and ZP3 (discussed in Section 111,B). As a result, the ZP becomes a more insoluble (“harder”) structure and the primary and secondary sperm receptor activities of the ZP (attributable to

FIG. 2. Transmission electron micrograph of an acrosome-intact mouse sperm bound to an unfertilized mouse egg ZP during culture in uitro. n, Sperm nucleus; a, sperm acrosome; zp, unfertilized egg zona pellucida; pm, unfertilized egg plasma membrane.

FIG. 3. Transmission electron micrographs of mouse sperm. (a) Acrosome-intact sperm head. (b) Acrosome-reacted sperm head. pm, Plasma membrane; ac, acrosome. (Note: the black dots associated with sperm heads are gold particles added to the sperm preparations.) The acrosome reaction involves multiple fusions between plasma membrane and outer acrosomal membrane at the anterior portion of the sperm head. This results in exposure of the inner acrosomal membrane and associated acrosomal contents. Only acrosome-reacted sperm can penetrate the ZP, and only acrosome-reacted sperm can fuse with egg plasma membrane.

STRUCTURE OF ZONA PELLUCIDA

89

ZP3 and ZP2, respectively) are destroyed. These changes prevent both continued penetration of supernumerary sperm already bound to the ZP and binding of additional free-swimming sperm to the ZP (Fig. 4). C. SUMMARY The ZP regulates the process of fertilization in mammals. The ZP possesses species-specific sperm receptors that enable free-swimming sperm to bind to unfertilized eggs. Binding of sperm to these receptors induces sperm to undergo the acrosome reaction, a prerequisite for fertilization. Once the egg is fertilized, the ZP undergoes changes that help to prevent fertilization by additional sperm (polyspermy). 111. Characteristicsof the Zona Pellucida

A. GENERAL CONSIDERATIONS The mouse egg ZP is a relatively thick (5-7 pm) translucent extracellular coat (Fig. 5 ) that contains -3-4 ng of protein and is extremely porous, permitting relatively large macromolecules (e.g., immunoglobulins) and even small viruses to pass through it (Gwatkin, 1977; Wassarman, 1988b). The ZP first appears when small oocytes (12-15 pm) begin to grow within the ovary, and increases in thickness as oocytes increase in diameter during a 2- to 3-week period (Wassarman, 1988~).During growth of the oocyte, the number and length of its microvilli increase markedly, presumably in order for oocyte surface area to keep pace with oocyte volume. These microvilli, as well as cellular projections that extend from surrounding follicle (cumulus) cells and form junctions with the oocyte (Anderson and Albertini, 1976; Gilula et al., 1978; Larsen et al., 1986),are embedded in the developing ZP. It has been known for some time that the ZP consists of both protein and carbohydrate (Harter, 1948; Braden, 1952; Jacoby, 1962; Guraya, 1974; Nicolson et al., 1975), with a ratio of protein to carbohydrate of -3-5 : 1 (Wassarman, 1988b). The ZP is quite elastic (Green, 1987). A well-known characteristic of the ZP is its ability to be solubilized by any one of a variety of agents, including proteases, mild acid, heat, reducing agents, and certain detergents (Gwatkin, 1977; Dunbar, 1983; Greve and Wassarman, 1985). In some cases, the ZP is solubilized by agents that do not break covalent bonds, suggesting that its structural integrity is maintained by noncovalent interactions. For example, the mouse oocyte ZP is dissolved at 50°C (neutral pH), and the rate of dissolution is significantly affected by the ionic strength of the culture medium (Fig. 6).

90

PAUL M. WASSARMAN AND STEVEN MORTILLO

FIG.4. Light micrograph (dark field) of ouse sperm, eggs, and two-cell embryos cultured in uirro. Sperm are bound to the ZP 0' Jnfertilized eggs, but not to the ZP of two-cell embryos. The latter is due to induction of the secondary block to polyspermy (i.e., "hardening" of the ZP and loss of receptivity to sperm) following fertilization. Sperm can attach, but not bind, to the ZP of artificially activated or fertilized eggs.

STRUCTURE OF ZONA PELLUCIDA

91

FIG. 5 . Micrographs of the mouse egg ZP. Shown are a transmission electron micrograph and a light micrograph (inset; Nomarski differential interference contrast) of the mouse egg ZP. zp, Zona pellucida; pvs, perivitelline space, pm, plasma membrane; pb, first polar body.

Time at 5OoC (hr)

FIG. 6. Dissolution of isolated ZP at 50°C as a function of ionic strength. The percentage of intact ZP (-50 ZP per sample) is plotted against time (hours) at 50°C in the presence of various concentrations of ammonium acetate (10-200 mM) at neutral pH. Dissolution of ZP was monitored by light microscopy (P. M. Wassarman, unpublished observations).

92

PAUL M. WASSARMAN AND STEVEN MORTILLO

B. ZP GLYCOPROTEINS The unfertilized mouse egg ZP is composed of three glycoproteins: ZP1, ZP2, and ZP3 (Bleil and Wassarman, 1980a; Wassarman, 1988b). These glycoproteins appear either as very broad bands following onedimensional sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (Fig. 7), or as a very complex set of spots following highresolution two-dimensional SDS-PAGE. Like many other glycoproteins, their appearance on gels is due to varying degrees of microheterogeneity, which result from differences in glycosylation patterns. Based on their migration on SDS-PAGE, under nonreducing conditions, the average apparent MrS of ZP1, ZP2, and ZP3 are 200,000, 120,000, and 83,000, respectively. All three glycoproteins have isoelectric points below 5 , and, like their broad M , ranges, their relatively acidic nature is due to carbohydrate, not polypeptide. ZP2 and ZP3 are the major components of the ZP and, based on protein estimates, are present in roughly equimolar amounts. ZPl is a relatively minor component of the ZP, apparently present at less than one-quarter the amount of either ZP2 or ZP3. Collectively, ZPl,ZP2, and ZP3 account for more than 90% of the mass of the ZP. Of course, it is possible that additional minor components of the ZP have gone undetected as yet. 1. ZP1 ZP1 ( M , 200,000) consists of two apparently identical 75,000 M , polypeptides, held together by intermolecular disulfides, and an undetermined number of N- and 0-linked oligosaccharides (Wassarman, 1988b; G. S. Salzmann and P. M. Wassarman, unpublished observations). On SDSPAGE, performed under reducing conditions, ZP1 migrates at approximately 120,000 M,.. 2 . ZP2 ZP2 (Mr 120,000)consists of a 77,000 M , polypeptide (679 amino acids), six complex-typeN-linked oligosaccharides,and an undetermined number of 0-linked oligosaccharides(Greve et al., 1982;Wassarman, 1988b;Liang et al., 1990). ZP2 is synthesized as a 713-amino acid polypeptide (Fig. 8), with a 34-amino acid signal-sequence at its amino terminus that is cleaved prior to secretion of the glycoprotein (Liang et al., 1990). The polypeptide has seven potential N-linked glycosylation sites (Am-Xaa-Ser/Thr) and very little a-helix content. ZP2 has 21 cysteine residues, and there is substantial evidence to suggest that some of these are present in intramolecular disulfides (Bleil et al., 1981; Moller and Wassarman, 1989). ZP2 serves as a secondary sperm receptor during fertilization (Bleil et al., 1988) and is partially responsible for the slow block to polyspermy

STRUCTURE OF ZONA PELLUCIDA

93

FIG. 7. High-performance liquid chromatography (HPLC)-purified ZP glycoproteins ZPl,ZP2, and ZP3 from mouse ovaries. (A) Profile of absorption at 280 nm for a preparation of -30,000 ZP. (B) Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDSPAGE) analysis of HPLC fractions followed by staining with silver. Fractions 49-50 contain -120,000). Fractions 61-65 pure ZP1 (M,-200,000). Fractions 55-57 contain pure ZP2 (M, contain pure ZP3 (M, -83,000). For a description of ZP glycoprotein purificationprocedures, see Bleil et al. (1988).

following fertilization (Bleil et al., 1981; Moller and Wassarman, 1989). In the latter case, ZP2 undergoes limited proteolytic cleavage (producing a form of the glycoprotein called ZP2f), catalyzed by a cortical granule protease, following fertilization. Proteolysis is thought to inactivate ZP2 as a secondary sperm receptor and to cause hardening of the ZP.

94

PAUL M. WASSARMAN AND STEVEN MORTILLO

1 -MARWQRKASVSSPCGRSI Y R

FLSLLFTLVTSVNSVSLPQSENPAFPGTLI CDKDEVRIEFSSRFDMEKWNPSWDTLGSE ILNCTYALDLERFVLKFPYETCTIKWGGY

QVNIRVGDTTTDVRYKDDMYHFFCPAIQAE THEISEIWCRRDLISFSFPQLFSRLADEN

QNVSEMGWIVKIGNGTFSHILPLKDAIVQG FNLLIDSQKVTLHVPANATGIVHYVQESSY LYTVQLELLFSTTGQKIVFSSHAICAPDLS VACNATHMTLTIPEFPGKLESVDFGQWSIP EDQWHANGIDKEATNGLRLNFRKSLLKTKP SEKCPFYQFYLSSLKLTFYFQGNMLSTVID PECHCESPVSIDELCAQDGFMDFEVYSHQT KPALNLDTLLVGNSSCQPIFKVQSVGLARF HIPLNGCGTRQKFEGDKVIYENEIHALWEN PPSNIVFRNSEFRMTVRCYYIRDSMLLNAH VKGHPSPEAFVKPGPLVLVLQTYPDQSYQR PYRKDEYPLVRYLRQPIYMEVKVLSRNDPN IKLVLDDCWATSSEDPASAPQWQIVMDGCE YELDNYRTTFHPAGSSAAHSGHYQRFDVKT FAFVSEARGLSSLIYFHCSALICNQVSLDS PLCSVTCPASLRSKREANKEDTMTVSLPGP ILLLSDVSSSKGVDPSSSEITKDIIAKDIA SKTLGAVAALVGSAVILGFICYLYKKRTIR

FNH-713 FIG. 8. Complete amino acid sequence of the ZP2 polypeptide chain. The signal-sequence (residues 1-34) is italicized, and positions of the seven potential N-linked glycosylation sites are underlined. For details, see text, as well as Liang et al. (1990).

3. ZP3 ZP3 (M,83,000) consists of a 44,000 M,polypeptide (402 amino acids), three or four complex-type N-linked oligosaccharides, and an undetermined number of 0-linked oligosaccharides (Salzmann et al., 1983; Wassarman et al., 1985; Wassarman, 1988b). ZP3 is synthesized as a 424amino acid polypeptide (Fig. 9), with a 22-amino acid signal-sequence at its amino terminus that is cleaved prior to secretion of the glycoprotein (Kinloch et al., 1988; Ringuette et al., 1988; Kinloch and Wassarman, 1989). The polypeptide is unusually rich in proline, serine, and threonine residues, has six potential N-linked glycosylation sites (consensus sequence Asn-Xaa-SedThr), and has very little a-helix content. ZP3 has 13 cysteine residues, and, based solely on its migration on SDS-PAGE in the presence and absence of reducing agents, it appears likely that some of

STRUCTURE OF ZONA PELLUCIDA

95

1 -MASS Y F LFLCLL LCGGPELCNSQTLW L LP G

GTPTPVGSSSPVKVECLEAELWTVSRDLF GTGKLVQPGDLTLGSEGCQPRVSVDTDVVR FNAQLHECSSRVQMTKDALVYSTFLLHDPR PVSGLSILRTNRVEVPIECRYPRQGHVSSH PIQPTWVPFRATVSSEEKLAFSLRLMEENW NTEKSAPTFHLGEVAHLQAEVQTGSHLPLQ LFVDHCVATPSPLPDPNSSPYHFIVDFHGC LVDGLSESFSAFQVPRPRPETLQFTVDVFH FANSSRNTLYITCHLKVAPANQIPDKLNKA CSFNKTSQSWLPVEGDADICDESHGNCSN SSSSQFQIHGPRQWSKLVSRNRRHVTDEAD VTVGPLIFLGKANDQTVEGWTASAQTSVAL GLGLATVAFLTLAAIVLAVTRKCHSSSYLV SLPQ-4 24

FIG. 9. Complete amino acid sequence of the ZP3 polypeptide chain. The signalsequence (residues 1-22) is italicized, positions of the six potential N-linked glycosylation sites are underlined, and positions of the 16 cysteine residues (three in the signal-sequence) are underlined. For details, see text, as well as Kinloch et al. (1988), Ringuette et al. (1988),and Kinloch and Wassarman (1989).

these are present in intramolecular disulfides. In this context, the positions of all 13 cysteine residues in mouse ZP3 polypeptide are conserved in both the hamster (Kinloch et al., 1990)and human (Chamberlin and Dean, 1990) ZP3 polypeptides. ZP3 serves a primary sperm receptor and acrosome reaction-inducer during fertilization and is partially responsible for the slow block to polyspermy following fertilization (Bleil and Wassarman, 1980b, 1983; Wassarman, 1987a,b, 1988a, 1989, 1990; Bleil, 1991). In the latter case, ZP3 is inactivated as a sperm receptor following fertilization by limited modification of its O-linked oligosaccharides(producing a form of the glycoprotein called ZP3f) (Florman and Wassarman, 1985; Wassarman, 1989, 1990; J. D. Bleil and P. M. Wassarman, unpublished observations). ZP3f is inactive as both a primary sperm receptor and an acrosome reactioninducer.

C. SOURCE OF ZP GLYCOPROTEINS Mouse ZP glycoproteins are synthesized and secreted by oocytes during their 2-to-3-weekgrowth phase (Bleil and Wassarman, 1980c;Greve et al., 1982; Salzmann et al., 1983; Roller and Wassarman, 1983; Shimizu et al., 1983; Wassarman, 1988b,c). In fact, genes encoding ZP glycoproteins are expressed exclusively by growing oocytes (Philpott et al., 1987; Roller et al., 1989; Kinloch and Wassarman, 1989; Liang et al., 1990; Lira et al.,

96

PAUL M. WASSARMAN AND STEVEN MORTILLO

1990 (Fig. 10). In growing oocytes, as much as 7-8% of total protein synthesis is devoted to ZP glycoproteins, and as much as 3-4% of total polysomal poly(A+) RNA encodes ZP glycoproteins. Thus, laying down the ZP is a major synthetic activity of growing oocytes. In fact, the tremendous expansion of the Golgi apparatus observed during mammalian oocyte growth (Anderson, 1972; Zamboni, 1972; Wassarman and Josefowicz, 1978) undoubtedly reflects, at least in part, processing and secretion of ZP glycoproteins. Numerous secretory vesicles containing nascent ZP glycoproteins can be found throughout the oocyte cytoplasm, as well as fused with oocyte plasma membrane (Wassarman, 1990; S. Mortillo and P. M. Wassarman, unpublished observations (Fig. 11). There is evidence to suggest that N-linked glycosylation of ZP2, but not ZP3, is necessary for efficient secretion of the nascent glycoprotein (Roller and Wassarman, 1983). D. SUMMARY Results of recent biochemical and molecular studies indicate that the mouse ZP is constructed of three glycoproteins-ZPl,ZP2, and ZP3-all of which are synthesized and secreted exclusively by growing oocytes. ZP2 and ZP3 polypeptides are not significantly similar to any other proteins entered into computer data bases. Collectively,these three glycoproteins account for the known functions of the ZP during fertilization.

IV. Ultrastructure of the Zona Pellucida A. GENERAL CONSIDERATIONS

The ZP has been a popular subject for morphological studies. Most investigators agree that the ZP is a fibrillogranular structure (Baranska et al., 1975; Anderson et al., 1978; Wassarman and Josefowicz, 1978; Diet1 and Czuppon, 1984; Greve and Wassarman, 1985) that has a “Swiss cheese” appearance in scanning electron micrographs (Phillipsand Shalgi, 1980a,b; Phillips et al., 1985). However, whether or not the ZP is structurally homogeneous throughout its width (as assessed by dye, antibody, or lectin binding and light and/or transmission electron microscopy) continues to be the subject of some disagreement (Dunbar and Wolgemuth, 1984; Wassarman, 1988b). Certainly, even relatively early morphological studies suggested that, in some species (e.g., rabbit), the ZP appeared to consist of two or more concentric layers (Dickmann, 1965).

FIG.10. I n situ hybridization assay of ZP3 mRNA in mouse ovaries. Shown are light micrographs [(A) bright field; (B) dark field] of the same ovarian section hybridized with a radiolabeled ZP3 probe and subjected to autoradiography. Only the five oocytes, not the surrounding follicle cells, have a concentration of grains above background. FC, Follicle cells; 0, oocyte. For details, see Roller er al. (1989).

FIG. 1 1. Transmission electron micrographs of immunogold-labeled mouse oocytes in ovarian sections. (A) Surface of a growing oocyte showing the ZP labeled with immunogold, plasma membrane (PM), and secretory vesicle (V)containing nascent ZP glycoproteins labeled with immunogold. (B) Golgi apparatus of a growing oocyte showing formation of vesicles (V)containing nascent ZP glycoproteins labeled with immunogold.

STRUCTURE OF ZONA PELLUCIDA

99

B. ELECTRON MICROSCOPY OF THE ZP The fibrous or filamentous nature of the ZP has been revealed in electron micrographs most dramatically by three different procedures. In one procedure, isolated ZP (porcine) were solubilized by lithium-3,5-diiodosalicylate (LIS) (Dietl and Czuppon, 1984), and in another isolated ZP (mouse) were solubilized by either proteases or reducing agents (Greve and Wassarman, 1985). Finally, electron micrographs of small growing oocytes (mouse), in the process of laying down a nascent ZP, have provided additional evidence for the filamentous nature of the ZP (Wassarman and Josefowicz, 1978; S. Mortillo and P. M. Wassarman, unpublished observations).

I . LIS-Solubilized ZP LIS is a chaotropic agent that has a tendency to act like a detergent (Findlay, 1987). When isolated ZP were exposed to 0.2 M LIS and then sedimented at 45,000 g and processed for transmission electron microscopy, individual long fibrils could be seen clearly in such preparations (Dietl and Czuppon, 1984) (Fig. 12). At this LIS concentration, apparently

FIG. 12. Transmission electron micrograph of porcine ZP fibrils. These fibrils were reorientedin aparallel fashion by treatment with 50 mMlithium-3,5-diidosalicylate(LIS). From Dietl and Czuppon (1984). Reproduced from Gamete Research, 9,45-54, by permission of Wiley-Liss Div. of John Wiley & Sons, Inc. 0 1984.

100

PAUL M. WASSARMAN AND STEVEN MORTILLO

a reorientation of fibrils occurred such that they came to lie in parallel to one another. In some cases, the individual fibrils actually protruded from the surface of the ZP or were released from the ZP. It should be noted that the ZP dissolved completely at LIS concentrations higher than 0.3 M. 2. Protease- and Dithiothreitol-Solubilized ZP Protease- and dithiothreitol-solubilized ZP preparations have been sprayed as a fine mist onto a substrate (mica or carbon-coated Formvar) and examined by transmission electron microscopy. Solubilization of ZP by elastase yielded large complexes consisting of interconnected, or branched, filaments (Greve and Wassarman, 1985; Wassarman, 1988b, 1990) (Fig. 13). Heterologous interactions among ZPl,ZP2, and ZP3 were maintained under these experimental conditions. Solubilization of ZP by chymotrypsin, on the other hand, yielded individual, or unbranched, filaments in which heterologous interactions between only ZP2 and ZP3 were maintained. Apparently, ZP1 was selectively degraded under these experimental conditions (presumably the cause of ZP dissolution). Similarly, dithiothreitol-solubilized ZP yielded unbranched filaments, presumably due to reduction of ZPl intermolecular disulfides. The effect of chymotrypsin and dithiothreitolon the branching of ZP filaments suggests that ZPl may serve as a filament crosslinker. Filaments as long as 2-3 pm could be seen in the elastase-, chymotrypsin-, and dithiothreitol-solubilized ZP preparations. The appearance of ZP filaments in micrographs depended very much on the manner in which they were prepared for electron microscopy. For example, ZP filaments have a 140-150 8, structural repeat that was clearly seen in negatively stained preparations adsorbed onto a substrate, as well as in rapidly frozen shadowed preparations (Fig. 14). However, this repeat was obscured in sprayed and shadowed preparations, as well as when glycerol was present in the solubilized samples. In the latter case, the structural repeat “collapsed” into 90 8, “beads” located every 170 8, or so along the ZP filament. Finally, electron micrographs of replicas of freeze-etched and platinum-shadowed ZP preparations clearly show the extensive filamentous lattice that constitutes the egg extracellular coat (S. Mortillo and P. M. Wassarman, unpublished observations) (Fig. 15). 3. ZP of Growing Oocytes

The filamentous nature of the ZP also has been revealed clearly in transmission electron micrographs of small growing oocytes within ovarian follicles (Wassarman and Josefowicz, 1978; S. Mortillo and P. M. Wassarman, unpublished observations) (Fig. 16). As oocytes initiate growth, they secrete nascent ZP nonuniformally into localized “pockets”

STRUCTURE OF ZONA PELLUCIDA

101

FIG. 13. Transmission electron micrographs of mouse ZP filaments. An enzymesolubilized ZP preparation was sprayed onto substrate-coated grids and rotary shadowed. Arrows indicate the number of branches extending from the point of intersection of ZP filaments. For details, see Greve and Wassarman (1985).

between the oocyte surface and surrounding follicle cells. Because the ZP is extremely “thin” at these early stages of oocyte (follicle) growth, its filamentous structure was easily resolved. It is noteworthy that, at these early stages, the ZP filaments appeared to be organized into nearly hexagonal arrays that had a fairly uniform pore size. As growing oocytes continue to increase in diameter, the pockets of ZP filaments coalesce and form a uniform coat around the cell.

102

PAUL M. WASSARMAN AND STEVEN MORTILLO

FIG. 14. Transmission electron micrographs of mouse ZP filaments. (A) Enzymesolubilized ZP preparation adsorbed to substrate-coated grid and negatively stained. (B) Enzyme-solubilized ZP preparation freeze-dried and unidirectionally shadowed. F, Filaments. For details, see Greve and Wassarman (1985) and Wassarman (1988b, 1990). (Panel B: Courtesy of Dr. John E. Heuser.)

STRUCTURE OF ZONA PELLUCIDA

103

FIG. 15. Transmission electron micrographof mouse ZP filaments (arrowheads).Purified oocyte ZP was sprayed onto mica, and freeze-etched platinum-shadowed replicas were prepared for examination by electron microscopy.

FIG. 16. Transmission electron micrograph of a developing mouse follicle. (a) Note the pockets (p; arrowheads) of nascent ZP that form between the growing oocyte and surrounding follicle cells. gv, Germinal vesicle of oocyte. (b) High-magnification view of a pocket of nascent ZP filaments present in the micrograph shown in (a).

STRUCTURE OF ZONA PELLUCIDA

105

C. ARRANGEMENT OF GLYCOPROTEINS I N ZP FILAMENTS The 140-150 A structural repeat described in Section IV,B,2 apparently reflects the periodic arrangement of hundreds of ZP2-ZP3 heterodimers along ZP filaments. That is, ZP filaments are polymers constructed of ZP2-ZP3 heterodimers. Evidence from two types of experiments supports such a tentative model for the structure of ZP filaments (Wassarman, 1988b; J. M. Greve and P. M. Wassarman, unpublished observations). First, quantitative analysis of electron micrographs of ZP filaments, decorated with monoclonal antibodies directed against either ZP2 or ZP3, revealed a periodic arrangement of immunoglobulin G (IgG) molecules that had an -150 A repeat (Fig. 17). Second, SDS-PAGE analysis of chemically crosslinked ZP filaments revealed the presence of a M , “ladder” of oligomers generated from an - 180,000M , species. The latter was composed of both ZP2 and ZP3. In view of these and other observations, as well as the stoichiometry of ZP2 and ZP3 in the ZP, the tentative model proposed above seems a very likely possibility (Fig. 18). To a certain degree, ZP filaments can be thought of as polymers composed of a single dimeric protein consisting of nonidentical subunits. D. SUMMARY Results of ultrastructural studies indicate that the ZP is constructed of long interconnected filaments that possess a structural repeat (periodicity). Additional evidence suggests that ZP filaments are polymers composed of ZP2-ZP3 heterodimers, which presumably account for the observed periodicity, and that ZP filaments are interconnected by ZP1.

V. Concluding Remarks A full appreciation of the molecular basis of ZP functions will require considerable information about ZP structure. Relatively recent studies, especially of the mouse egg ZP, have begun to provide some of the necessary structural information. Perhaps most surprising is the relatively simple composition of the ZP. Either three or four glycoproteins apparently constitute the ZP of eggs from a wide variety of mammals (from mice to humans). Thus far, all evidence suggests that these glycoproteins are unique (i.e., not significantly similar to any entries in computer data bases of protein primary structure) and that they are responsible for all ZP functions. Perhaps most gratifying is the finding that the ZP is not the amorphous structure it was once considered. On the contrary, the ZP is

106

PAUL M. WASSARMAN AND STEVEN MORTILLO I

I

30

1

I

50

60

7b

80

90 100

Antibody Spacing (nm)

FIG. 17. Histograms of distances between adjacent immunoglobulin G (IgG) molecules on ZP filaments decorated with a monclonal antibody directed against ZP3 (Bleil and Wassarman, 1986; Bleil et al., 1988) and rotary shadowed for examination by transmission electron microscopy (J. M. Greve and P. M. Wassarman, unpublished observations). (A) Ninety-six measurements of distances up to 52 nm between IgG molecules. (B) Fifty-seven measurements of distances up to 100 nm between IgG molecules. Distances between IgG molecules on straight portions of ZP filaments were measured on photographic prints by using an ocular micrometer. Note the peaks at approximately 15, 30,45,60,75, and 90 nm.

107

STRUCTURE OF ZONA PELLUCIDA

ZP2

ZP3

ZP2:ZP3

(120,000Mr) (83.000Mr) (180,000Mr)

ZP 1 (200,000Mr)

FIG. 18. Schematic representation of the arrangement of glycoproteins ZP1, ZP2, and ZP3 in mouse ZP filaments. The distance between ZP2-ZP3 dimers is approximately 15 nm (see text). The sites at which filaments are interconnected by ZP1 were chosen arbitrarily. Furthermore, the nature and specificity of ZP1 -filament interactions are unknown. For details, see text, as well as Greve and Wassarman (1985) and Wassarman (1988b).

apparently well organized, with primary and secondary sperm receptors (two different ZP glycoproteins) arranged periodically along ZP filaments. Many aspects of ZP structure need to be explored, since important questions remain unanswered. Some of these questions are concerned with assembly of the ZP during oocyte growth, as well as with the nature of the noncovalent bonds that stabilize ZP-glycoprotein interactions. The latter could be attributable to protein-protein, protein-carbohydrate, and/or carbohydrate-carbohydrate interactions. In view of the fact that the embryo ZP is considerably more insoluble (harder) than the egg ZP, it will be of interest to study these interactions both before and after fertilization. It appears that the extent of filament-filament interactions increases in the ZP following fertilization, thereby accounting for its decreased solubility. Assembly of the ZP during oocyte growth remains an interesting problem in biological construction. Even the simplest questions, for example, whether the ZP is built from the outside in or the inside out, remain to be answered. Perhaps, like the insect chorion (Regier et af., 1982), a glycoprotein framework is laid down and subsequently “filled-in.” It is unclear whether ZP assembly begins within the growing oocyte itself, perhaps in secretory vesicles, or only after secretion of individual nascent glycoproteins from the oocyte. Perhaps studies of ZP assembly will also finally answer the question of whether the ZP is structurally homogenous throughout its width or is subdivided into different structural domains.

108

PAUL M. WASSARMAN AND STEVEN MORTILLO

This brief review of ZP structure may convey some of the recent excitement in this area of research. Many important advances have been made, and our appreciation of ZP structure and function has increased significantly. The rate of recent progress in research on the ZP suggests that we can look forward to many new and unexpected discoveries in the near future. ACKNOWLEDGMENTS We are grateful to the members of our laboratory for their many experimental and conceptual contributions to research described in this review. Our research was supported in part by the National Institute of Child Health and Human Development, the National Science Foundation, the Rockefeller Foundation, and Hoffmann-La Roche, Inc.

REFERENCES Anderson, E. (1972). I n “Oogenesis” (J. D. Biggers and A. W. Schuetz, eds.), pp. 87-118. Univ. Park Press, Baltimore, Maryland. Anderson, E., and Albertini, D. F. (1976). J. Cell Biol. 71,680-686. Anderson, E., Wilkinson, R. F., Lee, G., and Meller, S. (1978). J . Morphol. 156,339-366. Baranska, W., Konwinski, M., and Kujawa, M. (1975). J. Exp. Zool. 192, 193-202. Bedford, J. M. (1981). Nature (London)291,286-288. Bedford, J. M. (1982). I n “Reproduction in Mammals: Germ Cells and Fertilization” (C. R. Austin and R. V. Short, eds.), Vol. 1, pp. 128-163. Cambridge Univ. Press, Cambridge, England. Bleil, J. D. (1981). I n “Elements of Mammalian Fertilization” (P. M. Wassarman, ed.), Vol. 1, pp. 133-152. CRC Press, Boca Raton, Florida. Bleil, J. D., and Wassarman, P. M. (1980a). Deu. Biol. 76, 185-203. Bleil, J. D., and Wassarman, P. M. (1980b). CellU), 873-882. Bleil, J. D., and Wassarman, P. M. (1980~).Proc. Narl. Acad. Sci. U.S.A. 77, 1029-1033. Bleil, J. D., and Wassarman, P. M. (1983). Deu. Biol. 95,317-324. Bleil, J. D., and Wassarman, P. M. (1986). J. CeNBiol. 102, 1363-1371. Bleil, J. D., Beall, C. F., and Wassarman, P. M. (1981). Deu. Biol. 86, 189-197. Bleil, J. D., Greve, J. M., and Wassarman, P. M. (1988). Deu. Biol. 128,376-385. Braden, A. W. H. (1952). Aust. J . Sci. Res. 5,460-471. Chamberlin, M. E., and Dean, J. (1990). Proc. Natl. Acad. Sci. U.S.A. 87,6014-6018. Dean, J., Millar, S. E., Liang, L., Chamberlin, M. E., and Lunsford, R. D. (1991). I n “Elements of Mammalian Fertilization” (P. M. Wassarman, ed.), Vol. 2, pp. 133-146. CRC Press, Boca Raton, Florida. Dickmann, Z. (1%5). I n “Preimplantation Stages of Pregnancy” (G. E. W. Wolstenholme and M. O’Connor, eds.), pp. 169-178. Churchill, London. Dietl, J., ed. (1989). “The Mammalian Egg Coat: Structure and Function.” Springer-Verlag, Berlin. Dietl, J., and Czuppon, A. B. (1984). Gamete Res. 9,45-54. Drobnis, E. Z., and Katz, D. F. (1991). I n “Elements of Mammalian Fertilization” (P.M. Wassarman, ed.), Vol. 1, pp. 269-300. CRC Press, Boca Raton, Florida.

STRUCTURE O F ZONA PELLUCIDA

109

Ducibella, T. (1991). In “Elements of Mammalian Fertilization” (P. M. Wassarman, ed.), Vol. I , pp. 205-232. CRC Press, Boca Raton, Florida. Dumont, J. N., and Brummett, A. R. (1985). In “Developmental Biology-A Comprehensive Synthesis” (L. W. Browder, ed.), Vol. 1, pp. 235-288. Plenum, New York. Dunbar, B. S. (1983). In “Mechanism and Control of Animal Fertilization,” (J. F. Hartmann, ed.), pp. 139-177. Academic Press, New York. Dunbar, B. S., and Wolgemuth, D. J. (1984). In “Modern Cell Biology,” (B. H. Satir, ed.), Vol. 3, pp. 77-11], Liss, New York. Findlay, J. B. C. (1987). in “Biological Membranes-A Practical Approach” (J. B. C. Findlay and W. H. Evans, eds.), pp. 179-218. IRL Press, Oxford. Florman, H. M., and Wassarman, P. M. (1985). Cell41, 313-324. Gilula, N. B., Epstein, M. L., and Beers, W. H. (1978). J . Cell Biol. 78,58-75. Green, D. P. L. (1987). Exp. Cell Res. 169,31-38. Greve, J. M., and Wassarman, P. M. (1985). J. Mol. Biol. 181,253-264. Greve, J. M., Salzmann, G. S., Roller, R. J., and Wassarman, P. M. (1982). Cell31,749-759. Gulyas, B. J. (1980). Int. Reu. Cytol. 63,357-392. Guraya, S. S. (1974). Int. Rev. Cytol. 37, 121-152. Gwatkin, R. B. L. (1977). “Fertilization Mechanisms in Man and Mammals.” Plenum, New York. Harter, B. T. (1948). Anat. Rec. 102,349-367. Hogan, B., Costantini, F., and Lacy, L. (1986). “Manipulating the Mouse Embryo.” Cold Spring Harbor Lab., Cold Spring Harbor, New York. Jacoby, F. (1962). I n “The Ovary” (S. Zuckerman, ed.), Vol. 1, pp. 189-245. Academic Press, New York. Kinloch, R. A., and Wassarman, P. M. (1989). New Biol. 1,232-238. Kinloch, R. A., Roller, R. J., Fimiani, C. M., Wassarman, D. A., and Wassarman, P. M. (1988). Proc. Natl. Acad. Sci. U . S . A . 85, 6409-6413. Kinloch, R. A., Ruiz-Seiler, B., and Wassarman, P. M. (1990). Deu. Biol. 142,414-420. Kopf, G . S., and Gerton, G. L. (1991). In “Elements of Mammalian Fertilization” (P. M. Wassarman, ed.), Vol. I , pp. 153-204. CRC Press, Boca Raton, Florida. Larsen, W. J., Wert, S. E., and Brunner, G. D. (1986). Dev. Biol. 13,517-521. Liang, L.-F., Chamow, S. M., and Dean, J. (1990). Mol. Cell. Biol. 10, 1507-1515. Lira, S. A., Kinloch, R. A., Mortillo, S., and Wassarman, P. M. (1990). Proc. Natl. Acad. Sci. U.S.A. 87,7215-7219. McLaren, A. (1982). I n “Reproduction in Mammals: Embryonic and Fetal Development” (C. R. Austin and R. V. Short, eds.), Vol. 2, pp. 1-25. Cambridge Univ. Press, Cambridge, England. Moller, C. C., a i d Wassarman, P. M. (1989). Deu. Biol. 132, 103-112. Nicolson, G. L., Yanagimachi, R., and Yanagimachi, H. (1975). J . Cell Biol. 66, 263-274. Perona, R. M., and Wassarman, P. M. (1986). Deu. Biol. 114,42-52. Phillips, D. M. (1991). I n “Elements of Mammalian Fertilization” (P. M. Wassarman, ed.), Vol. 1, pp. 249-268. CRC Press, Boca Raton, Florida. Phillips, D. M., and Shalgi, R. (1980a). J. Exp. Zool. 213, 1-8. Phillips, D. M., and Shalgi, R. (1980b). J . Ultrastruct. Res. 72, 172-181. Phillips, D. M . , Shalgi, R., and Dekel, N. (1985). Am. J. Anat. 174, 357-372. Philpott, C. C., Ringuette, M. J., and Dean, J. (1987). Dev. Biol. 121,568-575. Regier, J. C., Mazur, G. D., Kafatos, F. C., and Paul, M. (1982). Deu. Biol. 92, 159-174. Ringuette, M. J., Chamberlin, M. E., Baur, A. W., Sobieski, D. A,, and Dean, J. (1988). Dev. Biol. 127,287-295. Roller, R. J., and Wassarman, P. M. (1983). J. Biol. Chem. 258, 13243-13249.

110

PAUL M. WASSARMAN AND STEVEN MORTILLO

Roller, R. J., Kinloch, R. A., Hiraoka, B. Y., Li, S. S.-L., and Wassarman, P. M. (1989). Development 106,251-261. Sacco, A. G., and Yurewicz, E. C. (1989). I n “The Mammalian Egg Coat: Structure and Function” (J. Dietl, ed.), pp. 128-154. Springer-Verlag, Berlin. Salzmann, G. S., Greve, I. M., Roller, R. J., and Wassarman, P. M. (1983). EMEO J. 2, I451-1456. Shimizu, S.,Tsuji, M., and Dean, J. (1983). J. Eiol. Chem. 258,5858-5863. Wassarman, P. M. (1987a). Science 235,553-560. Wassarman, P. M. (1987b). Annu. Rev. Cell Eiol. 3, 109-142. Wassarman, P. M. (1988a). Sci. A m . 256,78-84. Wassarman, P. M. (1988b).Annu. Rev. Eiochem. 57,415-442. Wassarman, P. M. (1988~).I n “The Physiology ofReproduction” (E.Knobil and J. D. Neill, eds.), Vol. 1, pp. 60-102. Raven, New York. Wassarman, P. M. (1989). I n “Carbohydrate Recognition in Cellular Function” (G. Bock and S. Harnett, eds.), pp. 135-155. Wiley, New York. Wassarman, P. M. (1990). Development 108, 1-17. Wassarman, P. M., ed. (1991). “Elements of Mammalian Fertilization,” Vol. 1 and 2. CRC Press, Boca Raton, Florida. Wassarman, P. M., and Josefowicz, W. J. (1978). J. Morphol. 156,209-236. Wassarman, P. M., Bleil, J. D., Florman, H. M., Greve, J. M., Roller, R. J., Salzmann, G. S., and Samuels, F. G. (1985). Cold Spring Harbor Symp. Quant. Eiol. 50, 11-19. Yanagimachi, R. (1978). Curr. Top Dev. Eiol. l2,83-105. Yanagimachi, R. (1984). Gamete Res. 10, 178-232. Yanagimachi, R. (1988). I n “The Physiology of Reproduction” (E. Knobil and J. D. Neill, eds.), Vol. 1, pp. 135-185. Raven, New York. Zamboni, L. (1972). I n “Oogenesis” (J. D. Biggers and A. W. Schuetz, eds.), pp. 5-46. Univ. Park Press, Baltimore, Maryland.

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 130

The Male Germ Cell Protective Barrier along Phylogenesis MORDECHAI ABRAHAM Department of Zoology, The Hebrew University of Jerusalem, Jerusalem 91904, Israel

I. Introduction

The silver jubilee of the first descriptions of the blood-testis barrier based on modem techniques (Kormano, 1967, 1968; Nicander, 1967; Setchell, 1967, Setchell et al., 1969; Setchell and Waites, 1970) is nearly here; the centenary of the pioneer experiments of intravenous dye injecting (Ribbert, 1904; Bouffard, 1906; Goldman, 1909; Pari, 1910), which led to the discovery of the barrier, is not far away; yet some of the basic tenets of the blood-testis barrier concept are challenged. Germ cell autoantigens, which, until recently, were considered as being hidden behind the barrier (Setchell and Waites, 1975), were discovered in the open compartment of the seminiferous tubules (Yule et al., 1987a,b, 1988) (Section V,C). Dym and Fawcett’s (1970) two-layer barrier formed by Sertoli and myoid cells has been replaced by up to six barrier sites between the circulatory system and the postleptotene germ cells (Section IV,F). The intermediate testicular compartment, with the help of which Russell (1977a) so aptly explained the migration of the spermatocytes, is now doubted (Cavicchia and Sacerdote, 1988) (Section IV,A). But the most controversial aspect of the blood-testis barrier remains its name: “Blood-testis barrier” sounds good and is, moreover, easily remembered; the abbreviationBTB-sounds as good as “BBB” (blood-brain barrier); and it perfectly evokes the image of what it is supposed to represent-its only flaw being that it is not more generally accepted (Section 11,B). The Sertoli cells and the barrier they form have been reviewed by Fawcett (1975), Setchell and Waites (1979, Setchell (1980), Waites and Gladwell (1982), Tindall et al. (1985), Russell and Peterson (1983, and Setchell and Brooks (1988), who have concentrated on the mammalian testis. Pilsworth and Setchell(1981), in their comparative review of invertebrate and vertebrate testes, briefly referred to the BTB. A concise review of the testis barrier and the relationship between germ and somatic cells in the testes of locusts and moths has been published by Szollosi (1982). Since 1975, a steadily increasing number of publications on the 111

Copyright 0 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

112

MORDECHAI ABRAHAM

relationship between somatic cells and germ cells have confirmed the presence of a barrier between them in species belonging to most of the major phyla. In 1975, Marcaillou and Szollosi published their report on the permeability of the testicular follicles in the locust. Two years later, they published a paper about the BTB in locusts (Szollosi and Marcaillou, 1977), the first report of the barrier in invertebrates. In 1979, the barrier was documented in humans by Camatini er al. However, as early as 1973, Koskimies, Kormano, and Alfthan found in human seminiferous tubule fluid several proteins found neither in serum nor in testicular lymph, and concluded that this was due to a BTB. Abraham et al. (1979a) and Marcaillou and Szollosi (1980) described the barrier in fish; Osman et al. (1980), in birds; Franchi et al. (1982), Cavicchia and Moviglia (1983), and Bergmann et al. (1983), in amphibians; and Baccetti et al. (1983), in reptiles. In the invertebrates, other than insects, the barrier was described in a nematode (Marcaillou and Szollosi, 1980), a centipede (Beniouri, 1984), snails (de Jong-Brink et al., 1984; O’Donovan and Abraham, 1987), a flatworm and possibly a leech (O’Donovan and Abraham, 1987), and a coelenterate (O’Donovan and Abraham, 1988a). There are important zoological groups in which no barrier has been observed so far (e.g., Echinodermata, Cephalochordata, and Cyclostomata). In other groups (e.g., Poly- and Oligochaeta, Crustacea, and Urochordata), critical research concerning the barrier is still lacking. The discovery at the beginning of this century (Ribbert, 1904; Bouffard, 1906; Goldman, 1909; Pari, 1910) of a BTB in the mammalian testis was confirmed by numerous investigators in the late 1960s and 1970s (Dym and Fawcett, 1970; Dym, 1973; Gilula et al., 1976; Dym and Cavicchia, 1977; 1978; Cavicchia and Dym, 1978). It has been extended, with a few exceptions (Section 111) into a general principle of a barrier between somatic tissue and male germ cells in the metazoa, from coelenterates to mammals (Marcaillou and Szollosi, 1980; Abraham, 1983; O’Donovan, 1988; O’Donovan and Abraham, 198813).

11. Background

A. DIFFERENT BLOOD-ORGAN BARRIERS

The barrier which protects the male germ cells is not unique in the animal kingdom. Several other tissue-protecting barriers have been described: the BBB, for example (Bodenheimer and Brightman, 1968; Brightman et al., 1970; Shivers, 1979; Betz et al., 1980; Goldstein and Betz, 1986, and many other authors). The BBB is formed by endothelial

MALE GERM CELL PROTECTIVE BARRIER

113

cells lining cerebral capillaries, with tight junctions which prevent the intercellular passage of proteins (Reese and Karnovsky, 1967). Between adjacent endothelial cells, there are closed circumferential belts of tight junctions, with the exception of certain regions in the brain, which are outside the barrier (Rodriguez, 1955). The endothelial cells in endoneural capillaries also have tight junctions, which make them impermeable to peroxidase (Olsson and Reese, 1969). In sharks, the endothelium of brain capillaries is permeable to circulating horseradish peroxidase (HRP). Here, there are no tight junctions between endothelial cells, and injected tracers such as HRP penetrate the subendothelial space. Their spread into the perivascular neuropil is impeded by well-developed tight junctions between lateral membranes of astrocytic processes (Gotow and Hashimoto, 1984). Blood-organ barriers have also been described in invertebrates: for example, the blood-kidney barrier in the land snail Achatina achatina (Skelding, 1973); the blood-eye barrier (Shaw, 1977, 1978) in the compound eye, which seals off the insect retina from the blood and from the lamina synaptic region; and others. B. TERMSFOR THE BLOOD-TESTIS BARRIER The original term “blood-testis barrier” is not universally accepted. Patterned after the “blood-brain barrier,” the term “blood-testis barrier” is, in fact, a misnomer (Abraham et d., 1980). Alternate suggestions include: testicular-tubular barrier (Willson et al., 1973), blood-germ cell barrier (Jones, 1978), blood-seminiferous tubule barrier (Howards et al., 1976), and hemolymph-germ cell barrier (Szollosi and Marcaillou, 1977). Abraham et al. (1980), working on fish, have argued that the barrier is not between blood and the testis as a whole, since, in specimens injected with HRP into blood, the interstitial region of the testis is filled by the marker both inter- and intracellularly. Only the region of the germ cells is protected by inter-Sertoli cell tight junctions. They therefore proposed the term “blood-male gonocyte barrier,” (or “blood-spermatozoa barrier”) (Abraham, 1981). Other terms proposed are: testis permeability barrier (Franchi et al., 1982), blood-gonad barrier (de Jong-Brink et al., 1984), testis-blood barrier (Beniouri, 1984), Sertoli cell barrier (Russell and Peterson, 1985), blood-tubular barrier (Sakiyama et al., 1988), and diapause barrier (Schweich and Leloup, 1988). All things considered, it appeared that a correction of the term was required. The Platyhelminthes and the Coelenterata have no blood, and in order to find a term which is also applicable to animals without a circulatory system, the term “somatic tissue-male germ cell barrier” seemed adequate (O’Donovan and Abraham, 1987, 1988a).

114

MORDECHAI ABRAHAM

In view of the fact that in several recent articles the capillaries in the mammalian testis have been described as having barrier features similar to those of the brain capillaries (Cordon-Cardo et al., 1989; Harik et al., 1989), the original term “blood-testis barrier” appears to be suitable (Section IV,F). However, it would not be applicable to Hydrozoa or Turbellaria, which do not have a circulatory system. An additional problem of terminology is that in mammals, for example, the barrier appears between two groups of germ cells: that is, between the mitotic spermatogonia and the leptotene spermatocytes in the open compartment, and the pachytene spermatocytes and the haploid germ cells in the closed one. In fish, the barrier is situated between spermatids and spermatozoa. It separates between germ cells at different stages of the spermatogenetic process. In the testes of mammals, there is also the problem of a multiplicity of barriers: those of the capillaries, of the immunosuppressive factor in the lymphoid spaces, of the intermyoid cell spaces, of the peritubular basement lamina, and finally that of the inter-Sertoli space (Section IV,F). In insects, too, more than one barrier site has been described (Schweich and Leloup, 1988), as well as in gastropods (O’Donovan and Abraham, 1987),and in fish (Lou and Takahashi, 1989). In order to adapt the term to different barrier sites, the term “male germ cell barrier” or “male germ cell protective barrier” (MGPB) is proposed without reference to its exact site(s) in the testes. Such a term adequately describes the male germ cell barriers in the entire animal kingdom. In this chapter, “male germ cell protective barrier” is generally used, together with the terms used by the individual authors to whose reviewed papers I referred. C. THEDESIGNATION “SERTOLI CELL”FOR SOMATIC CELLSIN TESTES FROM DIFFERENT PHYLA The term “Sertoli cell” originally referred to somatic cells of the male gonad in mammals. Is it proper to use this term in different phyla? In his original account, Sertoli (1865) wrote about cellule ramijcati (branching cells), which are a characteristic trait of the mammalian Sertoli cell, but not of the Sertoli cells in most other phyla. Is the use of the term “Sertoli cell” justified only in those animal groups in which the somatic cells carry out several of the tasks performed by the mammalian Sertoli cell (e.g., surrounding the seminiferous tubules, phagocytosis, and forming a barrier) and are in intimate contact with the germ cells? In the literature the cell is referred to by Soos (1910), who used the name “Sertoli” for the somatic cells in the testes of a snail. Soos considered the auxiliary cell? in the gonads of Helix arbustorum as equivalent to the mammalian Sertoli cells. In many invertebrate species, the somatic cells surrounding the male

MALE GERM CELL PROTECTIVE BARRIER

115

gonocytes resemble the vertebrate Sertoli cells in many aspects. Germ cells develop in close contact with them, somatic cells are rich in lysosomes, they are surrounded by a basal lamina, they are actively phagocytosing, and the nucleus has indented outlines. A further point of analogy is the intercellular occluding junctions, which form a blood-male germ cell barrier. Hinsch and Dehn (1979) considered that the somatic cells of the asteroid Luidiu clathruta bear many similarities to the vertebrate Sertoli cells. They claimed that there is sufficient evidence for these cells to be called Sertoli cells. Chia and Buckland-Nicks (1987) objected to this and contended that the name “Sertoli” should be restricted to those somatic cells which form a BTB, as in vertebrates, insects, or mollusks. However, it must be admitted that the illustrations of the echinoderm somatic cells published by different authors (Section II1,J) are reminiscent of the vertebrate Sertoli cell. The orthodox view that an identical name should not be given to two different biological structures if homology is not established beyond all doubt is no longer universally accepted. In modem biology, the analogy of functions can be determinant for the name of a structure. Roosen-Runge ( 1977) mentioned a pillarlike ramifying supporting cell in the testes of Hydromedusae, of ectodermal origin, resembling closely the mammalian Sertoli cell which develops from mesodermal rudiment. In our view, the use of “Sertoli cell” is admissible for somatic cells of the male gonad from different phyla, even when only part of the functions of the mammalian Sertoli cell are performed. D. METHODSWHICHHAVEHELPEDTO ARRIVEAT AN MGPB

THE

CONCEPTOF

The ability of some mammalian tissues to exclude dyes has been known since early in this century (Pari, 1910). Dyes such as lithionkarmin (Ribbert, 1904), tolidine [sic] + acide H (Bouffard, 1906), and others have been injected intravenously, hypodermically, or intraperitoneally . Histological examination of the different tissues revealed that nervous structures and testes have kept out the dyes. Yet the concept of a barrier between blood and seminiferous tubules arose much later (Kormano, 1967,1968; Setchell, 1967; Setchell et al., 1969; Setchell and Waites, 1970). In fact, Nicander’s (1967) electron microscopic study first showed tight junctions in the inter-Sertoli space. During the early 1970s, transmission electron microscopy provided ultrastructural details of Sertoli-Sertoli junctions. In the later years of the decennium, these were supplemented by intercellular tracer or freezefracture replication techniques, which continue to be the preferred

116

MORDECHAI ABRAHAM

methods. Intercellular tracers injected into the circulatory system, incubated with testis fragments prior to fixation, or mixed with the fixative, represent the variants of this approach which attempt to visualize intercellular relationships in the tissue and to establish the precise site where the intercellular flow of macromolecules is interrupted by the Sertoli cell barrier. The most common tracers are HRP, with a molecular mass of about 40,000 Da and a diameter of 6 nm (Karnovsky, 1967),and lanthanum hydroxide or nitrate. The presence of HRP in the tissue is detected with diaminobenzidine. Parmentier et al. (1985) injected intracardially ,in addition to HRP, mouse IgG (170,000 Da) and IgM (900,000 Da) as macromolecule tracers, suitable to detection by immunofluorescence. Tracer techniques were used in many studies in order to delineate the boundaries between the territory freely irrigated by the blood and regions barred from it, in mammals from minks (Pelletier, 1986, 1988) to humans (Camatini et al., 1979), and in lower vertebrates from fish (Abraham et al., 1980) (Figs. 18-22) to fowl (Osman et al., 1980). Lou and Takahashi (1989) used HRP and bovine serum albumin in the teleost Oreochromis niloticus to locate the sites of the testis barrier. Intercellular tracer techniques were also used to detect the site of the barrier in invertebrates: in Hydra uiridis incubated alive with HRP (O’Donovan and Abraham, 1988a) (Figs. 3 and 4); in a filarial parasite, Dipetalonema dessetae (Marcaillou and Szollosi, 1980), dissected in culture medium containing HRP; in Lithobius forfcatus (Beniouri, 1984) tested with different tracers, such as ferritin, lanthanum nitrate, and HRP incubated with the testes, prior to fixation; in Mamestra brassicae (Schweich and Leloup, 1988) tested with lanthanum nitrate; and in Lymnaea stagnalis (de Jong-Brink et al., 1984) incubated in uitro with lanthanum nitrate, tannic acid, and colloidal gold. O’Donovan and Abraham (1987) injected HRP dissolved in snail physiological solution (Lockwood, 1961) into the hearts of living snails, Leuantinu hierosolyma (Fig. 14). Prior to the injection, the shell region overlying the heart was thinned by a weak solution of H2N03. Further perfection of the tracer techniques is due to Cavicchia and Sacerdote (19881, Ross (19771, and Pelletier and Friend (1983), who administered the tracers both by blood vessels and by perfusion via the seminiferous tubules. Cavicchia and Sacerdote (1988) placed the exposed testes and epididymides of rats on the stage of a Leitz micromanipulator and injected mixtures of fixative and lanthanum, or of peroxidase alone, into the tubules or the rete testis. The marker penetrated the tubuli, from both the tubular lumen and the interstitial capillaries toward the cis and trans parts of the barrier. Ross (1970) monitored the intratubular penetration of

MALE GERM CELL PROTECTIVE BARRIER

117

acriflavine with a fluorescent microscope. Considerable progress in the visualization of intercellular structures was made by introduction of the low-molecular-weight polyene antibiotic filipin, as a cytochemical probe which binds specifically to certain membrane sterols (Kinsky, 1970; Hamilton-Miller, 1973; Robinson and Karnovsky, 1980). Filipin-sterol complexes can be detected at the ultrastructural level after negative staining. In addition, when filipin is used after fixation or simultaneously with fixation, filipin-sterol complexes are readily visualized in freeze-fracture preparations, since they form 20- to 25-nm protuberances inside the split membranes. The precision of the filipin technique was demonstrated by Pelletier and Friend (1983, 1986) (Figs. 25-27), who studied freeze-fracture replicas in order to locate filipin-sterol complexes in the split-membrane surfaces of the adluminal compartment in guinea pig testis. A very detailed analysis of the MGPB in different phyla is due to a combination of freezefracture (or freeze-etch) and tracer techniques. This makes possible en face visualization of the interior of the cell membrane (Figs. 23 and 24) and correlation between the split-membrane structure and tracer penetration [Connell (1978), Suzuki and Nagano (1978), Nagano and Suzuki (19831, Pelletier and Friend (1983, 1986), and Pelletier (1988) in mammals; Pelletier (1990) in birds; Toshimori et al. (1979) and Miranda and Cavicchia (1986, 1988) in insects] (Figs. 25 and 28). A wide range of new methods and perfections of old ones regularly appear in the literature. Russell and Burguet (1977) used osmium:ferrocyanide as a fixative for testes. Junctional particles appeared clearly defined as electron-translucent spots within each bilayer half of the membrane. Bergmann et al. (1984b) used hypertonic fixatives, which produced shrinkage in cysts containing spermatogonia and spermatocytes, whereas cysts possessing spermatids and spermatozoa were not affected. Recently introduced techniques include immunocytochemical methods (LewisJones et al., 1987; Harik et al., 1989) and staining following fixation with tannic acid (Miranda and Cavicchia, 1988), which sharply enhances the contrast of the membranes and their junctional specializations. Morales et al. (1987) analyzed in uiuo, by light microscopy and quantitative autoradiography , the transport of radioactive iron across the seminiferous tubules. Also with the in situ hybridization technique, they studied the anatomical site of transferrin gene expression in the seminiferous epithelium of the rat. A different approach to demonstrate the existence of the BTB uses comparisons of the chemical composition of seminal fluid and blood plasma. The review by Setchell and Waites (1975) presents inter alia various techniques to achieve this objective. Marshall and Bryson (1988)

118

MORDECHAI ABRAHAM

and Marshall et al. (1989) compared the in uitro bidirectional transcytotic activity of the sperm duct epithelium in the brook trout. They quantified the transepithelial transport of 86Rb+,a tracer for K+, concomitantly with Na’ absorption, measured as the short-circuit current. Morisawa et al. (1979), Steyn and Van Vuren (1986), and Vlok and Van Vuren (1988) compared the chemical composition of blood and seminal plasma and reported significant differences in pH and in concentrations of sodium, potassium, calcium, magnesium, chloride, glucose, fructose, total protein, and total lipids, etc. Turner et al. (1981b) compared the concentrations of intravenously infused labeled inulin in blood, seminiferous tubule fluid, and cauda epididymal fluid in order to gauge the tightness of the tubular and epididymal barriers. E. THESERTOLI CELLin Vitro There is a steadily growing number of publications on Sertoli cell in in uitro conditions, sometimes in combination with micromanipulationtechniques. Many facets of the activity of Sertoli cells have been examined in in uitro systems, for example, the regulation of plasminogen activator secretion (Lacroix et al., 1977; Ellison and Jenkins, 1989), the regulation of transferrin secretion (Jenkins and Ellison, 1989), and the formation of the tight junctions (Eddy and Kahri, 1976). Bigliardi and Vegni Talluri (1976) compared the ultrastructure of Sertoli cell junctional complexes in rat testis in uiuo and in uitro. In uitro techniques applied to Sertoli cell activity are of two kinds: (1) tissue culture of pieces of seminiferous tubules and subsequent studies by phase contrast light microscopy and, after fixation, by electron microscopy (Eddy and Kahri, 1976); and (2) cell culture of separated Sertoli cells. Cell- or tissue-cultured Sertoli or epididymal cells were assayed for a large variety of purposes: hormonal stimulation of the production of androgen-binding protein (ABP) (Louis and Fritz, 1977, 1979), inhibin production (LeGac and de Kretser, 1982), age-related differences in response to follicle-stimulatinghormone (FSH) (Lee et al., 1983), ceruloplasmin synthesis (Skinner and Griswold, 1983), and testosterone metabolism in epididymis (Brown et al., 1983). An important step forward in the study of Sertoli cell functions involved the introduction of the “two-chamber assembly” technique (Misfeldt et al., 1976; Byers et al., 1986; Ailenberg et al., 1988). Tung and Fritz (1984) emphasized that morphology, behavior, and activity of Sertoli cells in culture are profoundly influenced by the substratum. Sertoli cells plated onto plastic lose their polarity, flatten, and develop atypical junctional complexes (Tung and Fritz, 1984; Hadley et al., 1985). Sertoli cells grown on a reconstituted basement membrane were columnar, resembling the

MALE GERM CELL PROTECTIVE BARRIER

119

cells in uiuo, with characteristic basally located tight junctions. Hadley et al. (1985) Ailenberg et al. (1988)used as substratum an extracellular matrix (Matrigel, from Collaborative Research, Inc. ; Bedford, MA) prepared from tumor cells. Sertoli cells, or rnyoid cells cultivated separately in uitro, failed to form the basal lamina space (i.e., the basement membrane) which is formed between them in uiuo. Only in coculture, with direct contact between myoid and Sertoli cells, was the basal lamina formed, and at the same time the secretory activity of the Sertoli cells was modulated (Skinner and Fritz, 1985; 1986; Hettle et al., 1986). Another problem which was tackled with the dual-environment chambers method concerns the polarity of the Sertoli cells. Dym et al. (1986) measured the iron transport across the Sertoli cell monolayer by adding to the basal compartment of the culture chamber human transferrin bound to 59Fe. The 59Feentered the Sertoli cells, by dissociation from the human transferrin and became bound to endogenous rat transferrin before being secreted in the adluminal compartment. Thus, it was demonstrated that, in viuo as well as in uitro, 59Feis transcytosed from the base of the Sertoli cells toward the adluminal compartment in a polarized condition. 111. The Male Germ Cell Protective Barrier along Phylogenesis

A. PORIFERA

No MGPB has been described in the phylum Porifera. However, it is interesting to examine the spermatogenetic process described in several species, in view of a unique phenomenon not observed in Metazoa: Spermatozoa are carried to the ova by specialized cells, thus separating them from the somatic tissue. They evoke the image of a miniature spermatophore which delivers a single spermatozoon to the mature oocyte. According to Fell (1974), Roosen-Runge (1977), Reiswig (1983), and others, spermatozoa enter the female inhalant systems. The male germ cell then makes contact with a choanocyte (i.e., the carrier cell), which engulfs it and holds it within a membrane-bound cavity-a spermiocyst. Finally, the male gamete, which has by then lost its flagellum, is transferred to the egg cell to fertilize it. According to Tuzet et al. (1970), in Aplysilla rosea, at the start of spermatogenesis all choanocytes of a flagellated chamber transform simultaneously to spermatogonia, which later undergo meiosis. The germ cells are surrounded by a single layer of pinacocyte-like flattened cells which constitute a spermatic cyst. The degree of isolation provided by the enclosing cells, between the neighboring somatic tissue and the male germ cells, was not examined with the help of intercellular markers. In Suberites massa, Diaz (1973) and Diaz et al. (1973) did not observe a cellular enve-

120

MORDECHAI ABRAHAM

lope in the early stages of spermatogenesis; this cell layer developed later, during maturation divisions, a sequence of stages predictive of future development of the MGPB in higher phyla. B. CNIDARIA The presence of an MGPB in H . viridis was proposed by O’Donovan and Abraham (1988a) (Figs. 1-5). In the hydrozoans, interstitial cells between the epitheliomuscular cells of the epidermis give rise to germ cells (Burnett et al., 1966; Schincariol et al., 1967; Davis, 1973; Tardent, 1974; RoosenRunge, 1977; Miller, 1983), enclosed within compartments of the overall epidermal structure-except for small regions of direct contact between the spermatogonia and the mesoglea (Burnett et al., 1966). Later stages of spermatogenesis were found deeply embedded in the gonadal spaces between the epitheliomuscular cells. This bears a striking resemblance to the manner in which the spermatogonia of the vertebrate amniotes rest against the basal lamina of the seminiferous tubules, whereas the later stages are deeply embedded and surrounded by Sertoli cells (O’Donovan and Abraham, 1988a). Large intracellular vacuoles have been described in both the epidermis and the gastrodermis of Hydra (Lenz and Barmett, 1965; Haynes, 1973; Benos et al., 1977). In Hydra incubated alive with HRP (O’Donovan and Abraham, 1988a), the marker was never found within the intercellular space between the epithelial cells of either epi- or gastrodermis. Septate junctions found between these cells were impermeFIGS.1-5. Hydra uiridis (Coelenterata). From O’Donovan and Abraham (1988a); Reproduced from “Somatic tissue-male germ cell bamer in Hydra uiridis (Hydrozoa, Coelenterata),” Journal ofMorphology, 1988,198,179-188, with permission of Wiley-Liss Div. of John Wiley & Sons, Inc. FIG. 1. Spermary. ep, Epidermis; gd, gastrodermis; gc, germ cells; sz, spermatozoa; mg, mesoglea (thins out at the spermary base, arrows). ~ 2 9 0 . FIG. 2. Septate junction (sj) in the spermary wall. EP, Epidermal cell; vac, vacuole; bf, bifurcation of the intercellular space. ~55,700.(Inset) Same as main figure at lower magnification. gl, Glycocalyx; g, granule. ~ 9 4 0 0 . FIG. 3. Longitudinal section of Hydra after 2-hour exposure to horseradish peroxidase (HRP). G.C.,Gastric cavity; mg, mesoglea; gd, gastrodermis, ep, epidermis. Single arrows, HRP fills the mesoglea in the lower part of the body; double arrows, cells in the basal disk filled with HRP pinosomes; *, HRP in the mesoglea stops diffusing upward. x 100. FIG. 4. Tracer-free intercellular spaces (arrows) in the spermary wall after 12-hour exposure to horseradish peroxidase (HRP) (noncontrasted thin section). HlW fills the glycocalyx (GL). X 15,360. FIG. 5 . Gap junction (arrow) between the epidermis (EP) and the gastrodermis (GD). mg, Mesoglea; tb, transmesogleal bridges between both epithelia. x24,570. (Inset) Gap junction (arrows) between EP and GD, from a different micrograph at high magnification. X73,920.

122

MORDECHAI ABRAHAM

able to HRP. In two parts of the body-the tentacles and the pedal disk-HRP was incorporated into epidermal cells through pinocytosis shortly after the exposure of Hydra to the tracer. The mesoglea in the region of the tentacles and the pedal disk appeared dark with tracer. Spermaries of H. uiridis develop in the upper gastric region of the body stalk. HRP was found to fill the mesoglea of the lower half of the body stalk, but in the upper half, including the spermary region, the mesoglea was always tracer free. Numerous studies have been made of the structure and/or permeability of septate junctions to extracellular tracers (Wood, 1959; Hand, 1971; Hand and Gobel, 1972; Filshie and Flower, 1977; Wood and Kuda, 1980). In H . uiridis, the paracellular flow across the epidermal intercellular spaces of solutes at least the size of HRP was limited by a barrier composed of (1) septate junctions, (2) electron-dense material within the intercellular space, and (3) long and tortuous pathways of intercellular spaces between the epidermal cells in the testis region. The connective tissue of the mesoglea is most similar in structure to the basal Lamina. It is considered the connective tissue of the coelenterates and can serve as a filter and ionic sink for other intercellular materials (Hausman, 1973), a kind of “primeval” connective tissue predating the origin of a circulatory system. In ripe spermaria, the mesoglea is thinner than in other regions of the body, and this might be an additional isolating factor of the germ cells. The gap junctions found between epi- and gastrodermis of H . uiridis provide direct metabolic contact without leakage into the mesoglea and the intercellular spaces (Hufnagel and Kass-Simon, 1976; Larsen, 1983; Verselis et al., 1986).

C. PLATYHELMINTHES In the free-living turbellarian Dugesia biblica, a somatic tissue-male germ cell barrier was described by O’Donovan and Abraham (1987) (Figs. 6-9). The testes of D. biblica consist of follicles surrounded by fusiform parietal cells embedded in the parenchyma (Henley, 1974). Cytoplasmic extensions overlap with those of contiguous parietal cells. The long intercellular spaces between adjacent parietal cells run concentrically with the follical wall. Toward the follicle lumen, these spaces are closed by septate junctions separating the testicular lumina from the general parenchyma. The interseptal region is filled by an electron-dense material. Septate junctions in Platyhelmintheshave been described by Green and Bergquist (1982), Lord and DiBona (1976), and Storch and Welsch (1977), however, not in the testis. The parietal cells have structural features in common with

MALE GERM CELL PROTECTIVE BARRIER

123

the Sertoli cells of vertebrates: They have a lobate nucleus, display phagocytotic activity, contain active Golgi structures, and display occluding junctions in their intercellular space. They achieve their role as molecular barriers by several combined factors: the amount of septa and the possible cation-binding properties of the interseptal material (Wood and Kuda, 1980). An additional factor which may enhance the barrier in D . biblica is the extreme length and orientation of the interparietal cell space. As to the role of the barrier in Platyhelminthes, observations by Gremigni (1974) concerning the differentiation of planarian neoblasts when isolated from the rest of the parenchyma are suggestive of a similar situation for the germ cells, which possibly undergo spermatogenesis when separated from the somatic tissue. Parietal cells bear the morphological characteristics of secretory cells (Franquinet and Lender, 1973), suggesting that the testicular fluid is different in composition from the rest of the tissue fluids. The composition of the testicular fluid may be important for keeping the sperm cells immotile.

D. NEMATODA

The description of a barrier between somatic cells in the testis of D . dessetae is due to the report by Marcaillou and Szollosi (1980). Dipetalonema dessetae, a filarial parasite infesting the rodent Proechimys oris, is 30-40 mm long. The single testis of D . dessetae is a hollow tube containing free germ cells which move from the apical germinal zone through a “growth zone” and then enter a dilated part of the male gonoduct, the seminal vesicle. Along its entire length, the tubular testis is limited by a somatic cell layer, externally covered by a thick sheath of amorphous material. Spermatogonia are located in the apical extremity of the gonad, while spermatocytes in their growth phase occupy the major part of it. When the worm is dissected in culture medium containing HRP, the tracer strongly impregnates the external sheath, penetrates between the somatic cells, and attains the diploid germ cells. In the last segment of the testis, containing spermatids, junctions have been observed between adjacent somatic cells. Here, the tracer impregnates the external sheath and sometimes infiltrates among the peripheral cells of the somatic envelope, but it does not reach the developing germ cells. The haploid spermatids will not differentiate further in the male genital tract (Terry et al., 1961). In nematodes, the tight compartment is very reduced, and the open one is very long; this feature might be related to the long-lasting diploid “growth phase” (Favard, 1961).

124

MORDECHAI ABRAHAM

FIGS.6-9. Dugesia biblica (Platyhelminthes). From O’Donovan and Abraham (1987); reproduced from “Somatic tissue-male germ cell bamer in three hermaphrodite invertebrates: Dugesia biblica (Platyhelminthes), Placobdella cosrafa (Annelida), and Leuanrina hierosolyma (Mollusca),” Journal of Morphology, 1987, 192, 217-227; with permission of Wiley-Liss Div. of John Wiley & Sons, Inc. FIG. 6. Testicular follicles with spermatozoa (Sp) in the lumen (Lum); parietal cells (P)

MALE GERM CELL PROTECTIVE BARRIER

125

E. ANNELIDA

Cytophores in the leech Placobdella costata, as a possible substitute of the barrier forming somatic cells in other phyla, were described by O’Donovan and Abraham (1987) (Fig. 10-12). Placobdella costata (Annelida, Hirudinea) parasitize a turtle, Mauremys caspica riuulata. The walls of the testes are formed by a layer of parietal cells and a thick basement membrane. During early spring, all the spermatogenetic stages are found developing in the lumina of the testes, attached to central cytophores. No specialized intercellular occluding junctions are found between adjacent parietal cells, although septate junctions were found in other tissues of P . costata (O’Donovan and Abraham, 1987). However, a thickening of the plasmalemma and electron-dense material in the intercellular space is found where the thin extensions of one cell overlap the neighboring cell. For the most part, these extensions interdigitate, determining tortuous intercellular pathways between the basement membrane and the testis lumen. Between the cytophore and the germ cells, there is a cytoplasmic continuum. The germ cells and their cytophores have no connection with the parietal cells. According to Roosen-Runge (1977), cytophores are a characteristic feature of annelids. Incomplete cytokinesis during spermatogonial division results in the formation of a central anucleate cytoplasmic mass-a cytophore (Sawada, 1984), in which germ cells undergo spermatogenesis. According to Bondi and Farnesi ( 1976), in Branchiobdella pentadonta, meiosis begins in cytophores containing 16 spermatogonia. The spermatozoa separate from the cytophore only when their differentiation is complete. Chatton and Tuzet (1941), Walsh (1954), and other discussed the origin of the cytophore (reviewed by Roosen-Runge, 1977). According to Anderson et al. (1967), in Lumbricus terrestris, mitochondria and an elaborate endoplasmic reticulum (ER) occur in the cytophore. Stang-Voss (1970) has observed degenerate Golgi structures, annulate lamellae, and ER in the cytophore of Eisenia foetidus in the spermatid stage. The presumed roles of the cytophore are a nutritional

are shown with septate junctions (arrows) between them. Sp.H, Spermatozoon head in a phagocytotic parietal cell. X 17,300. FIG. 7. Extensions of a parietal cell (P. ext) overlappingwith an adjacent parietal cell (P), forming a narrow intercellular space (arrowheads), running concentrically with the follicle lumen (Lum). Nu, Nucleus of parietal cell; Sp, spermatozoa. X 15,000. FIG. 8. Septate junctions between adjacent parietal cells (P) lining the follicular lumen (Lum). Sp, Spermatozoa. ~ 3 6 , 4 0 0 . FIG. 9. Tortuous intercellular space between parietal (P) cells with occluding septate junctions (arrows). e, Extensions of parietal cells; SER, smooth endoplasmic reticulum. X 15,470.

FIGS. 10-12. Placobdella costhta (Annelida). From O’Donovan and Abraham (1987); reproduced from “Somatic tissue-male germ cell barrier in three hermaphrodite invertebrates: Dugesia biblica (Platyhelminthes), Placobdella costata (Annelida), and Leuanrina hierosolyma (Mollusca),” Journal of Morphology, 1987, 192, 217-227; with permission of Wiley-Liss Div. of John Wiley & Sons, Inc.

MALE GERM CELL PROTECTIVE BARRIER

127

center, a center-to-control spermatogenetic synchrony, or a common disposal center of superfluous cytoplasm at the end of spermiogenesis. Ciliated cells in the walls of the testis were also found in other leeches, such as Glossiphonia complanata (Damas, 1968) and Hirudo medicinalis (Patisson, 1965, 1966, 1977). In P . costata, two different cell types were observed in the testicular walls. Structural characteristics in the P. costata testis possibly related to a barrier function are as follows: (1) electrondense material in the interparietal cell space near the testicle lumen, (2) a basement lamina which surrounds the parietal cells, and (3) the connection of the germ cells to the cytophore. It was postulated by O’Donovan and Abraham (1987) that molecules originating in the somatic tissue can only enter the germ cells through the central mass of the cytophore, which thus has itself a barrier role. Eckelbarger (1984) noticed that the testis of a reef-building polychaete, Phragmatopoma lapidosa, is similar to the ovary in that both have germ cell masses attached to the walls of intersegmental blood vessels. It is not mentioned whether the tissue connections between the gametes and the blood vessels are different between males and females. Sawada (1984) distinguishes between two polychaete types, the Arenicola and the Nereis types, with and without a cytophore, respectively. It would be rewarding to compare the organization of the periluminal somatic cells between these two types.

F. CRUSTACEA The junctional relationships between the cellular components of the seminiferous cords of Macrobrachium rosenbergii were studied by Dougherty and Sandifer (1984). The testes of the shrimp are composed of spermatogenic and sustentacular cells. No occluding junctions between the sustentacular cells were found. The authors summarized that “among the spermatogenic cells, preleptotene spermatocytes and encysted spermatozoa were of most frequent occurrence” (p. 115). However, in the

FIG. 10. Testis lumen (Lum). Parietal cells (PI and P2) with thin cytoplasmic extensions (stars) lining the testicle; one of the parietal cells (P2) lost its cilia. In the lumen (Lum), cytophores (Cyt) are shown with germ cells ( G , and GZ)at different stages of spermatogenesis attached to them; the connection of Gzgerm cells with the cytophore is outside the plan of the cut. Sp, Spermatozoa; col, collagen fiber space (Le., basement membrane). x 1880. FIG. 11. Similar to Fig. 10, at higher magnification. Arrows point to the rim of cytoplasm lining the lumen (Lum). G, Germ cells; P, parietal cells; col, collagen fiber space. ~ 4 1 3 0 . FIG. 12. Intercellular space between adjacent parietal cells (P, PI, and P2).Arrows point to the region near the lumen (Lum) with membrane thickenings and intercellular electrondense material. cil, Cilium; vil, villus. X 14,100.

128

MORDECHAI ABRAHAM

results and discussion, the germ cells they described are mainly preleptotene spermatocytes, and only passing mention was made of haploid gonocytes. The sustentacular cells, too, are examined in the region of spermatocytes. The possibility that occluding junctions appear between the sustentacular cells during the later part of meiosis, as in insects, cannot be ruled out. Koulish and Kramer (1986) described in the testis of a barnacle, Balanus eburneus, a thick basal lamina and branching Sertolilike cells which may hinder free diffusion of molecules from the hemolymph to the germ cells and provide some form of BTB. G. MYRIAPODA The MGPB in Myriapoda was studied by Beniouri (1984). Males of the centipede L .forficatus have an unpaired testis situated mediodorsally. The testicular wall is constituted of three cellular layers alternating with three collagen layers (Camatiniand Ceresa-Castellani,1974).Germ cell differentiation is oriented centripetally. Beniouri (1984) tested the permeability of the testicular wall to different tracers, such as ferritin and HRP,which did not penetrate into it. Lanthanum nitrate infiltrated the testicular wall and was stopped by septate junctions in the intercellular spaces of the cells enclosing the germ cells. In specimens treated with 20-hydroxy-ecdysone, the penetration of the marker was enhanced. These results are similar to those obtained by Kambysellis and Williams (1971a,b) in the silkworm, and opposite those obtained by Jones (1978), who found that ecdysterone induces a premature formation of the hemolymph-testis barrier in the testicular follicles of Schistocerca gregaria.

H. INSECTA In insects, a barrier between hemolymph and male germ cells was demonstrated in Orthoptera such as Locusta migratoria (Marcaillou and Szollosi, 1975; Szollosi and Marcaillou, 1977) and S . gregaria (Jones, 1978) and in Lepidoptera such as Bombyx mori (Toshimori et al., 1979), Anagasta kuehniella (Szollosi et al., 1980), Triatoma infestans (Miranda and Cavicchia, 1986, 1988),Heliothis uirescens (Baldwin et a f . , 1987),and M . brassicae (Schweich and Leloup, 1988).The paired testes in insects are composed of follicles which present almost innumerable variations in form and arrangement among different species (Imms, 1957). Each follicle is connected by a short vas efferens with the vas deferens. The vasa deferentia are paired canals which enlarge to form the vesicula seminalis. The

MALE GERM CELL PROTECTIVE BARRIER

129

united vasa deferentia open into the common ejaculatory duct. Inside the follicles are cysts containing clones of germ cells which differentiate from the apex toward the efferent duct. The youngest germ cells, the primary spermatogonia, not yet enclosed in cysts, are at the apices of the follicles; the oldest ones are in the basal region, near the vas efferens. In the midst of the primary spermatogonia is a large apical cell with unknown function (Szollosi and Marcaillou, 1979; Szollosi et al., 1980; Szollosi, 1982). The cells forming the wall of the follicles and the cells forming the germ cell cysts are differently structured in locusts and moths (Lane and Skaer, 1980; Szollosi, 1982). In locusts, the MGPB is in the follicle wall, while in the moths the barrier is in the cyst which envelops the germ cells. The follicle wall in locusts is single layered in the apical half and double layered in the basal half. In the basal half, the inner layer is constituted of cells displaying long overlapping processes which form a complex tortuous network of intercellular spaces along with extensive septate junctions. This inner cell layer of the wall forms the permeability barrier in locusts. Only in the basal half of the follicle is the barrier formed. This is the “closed” compartment with meiotic and postmeiotic germ cells. In the apical “open” compartment, which contains the young germ cells, the tracers permeate the intercellular spaces of the perifollicular cell layer and the cyst envelopes and reach the young germ cells, in S . gregaria (Jones, 1978) as well as in L. migratoria (Marcaillou and Szollosi, 1975; Szollosi and Marcaillou, 1977). The border between the open and closed compartments is located in the region where the germ cells have entered meiotic prophase. The barrier is established in the third larval instar in S . gregaria (Jones, 1978). In L. migratoria, it appears during the fourth instar (Szollosi and Marcaillou, 1977). In moths, the follicle wall is permeable. The cells forming the cyst envelope in moths send long cytoplasmic processes which surround the germ cells, creating a sinuous system of intercellular spaces between adjacent cells (Szollosi, 1982). Gap junctions, septate junctions, tight junction-like structures, and desmosomes are observed between the cyst cells. An impermeable barrier is formed. HRP incubated with the testes, prior to fixation, freely penetrates across the follicular wall at all stages of postembryonic life, but is totally excluded from the interior of the cysts. Only the primary gonia, which are not yet enclosed in cysts, have direct contact with the tracer (Szollosi et al., 1980). Toshimori et al. (1979) distinguished three kinds of cell junctions in the cyst envelope in the testes of B. mori: pleated septate junctions, gap junctions, and tight junctions. In A. kuehniella, tight junctions coexist with septate junctions, and in some areas of the membrane the two oc-

130

MORDECHAI ABRAHAM

cluding systems can be observed side by side (Szollosi, 1982). Miranda and Cavicchia (1988) described between adjacent cells composing the germ cell cysts of T. infestans, gap junctions and septate junctions of two types, smooth and pleated (Green, 1981). In freeze-fracture replicas, these two types of intercellularjunctions appear either as closely packed parallel rows of intramembrane particles on the P-face of the cell membranes, or as curved rows of particles. In the testes of M. brassicae (Lepidoptera, Noctuideae), Schweich and Leloup (1988) distinguished with the aid of lanthanum nitrate, two distinct hemolymph-male germ cell barriers: one formed by pleated septate junctions and tight junctions in the cyst envelope, the other located in the testicular coat in diapausing or prediapausing insects-the diapause barrier. No cytological structure in relation with this second barrier has been identified. A similar case of a barrier without structural basis in the insect testis is that of the apical cysts of locustan testis follicles, which are impermeable to HRP during the last larval instar for 24 hours (Marcaillou et al., 1978; Szollosi, 1982). Baldwin et al. (1987) described a “novel type of occludingjunction” in the testis of H. uirescens with freeze-fracture technique and HRP and Ruthenium Red tracers. This junction is formed by a septum which, according to the authors, is “more impermeable” than the “typical septum” of the septate junctions. However, the Ruthenium Red postfixation tracing, as used by these authors, must be interpreted cautiously. The open gonial compartment of the locust testis is almost always freely penetrated by HRP, at least during larval life. On the sixth or seventh day of the fifth instar, the apical cysts which contain spermatogonia become closed to the marker for 24 hours (Marcaillou and Szollosi, 1975; Marcaillou et al., 1978). At this time, high levels of ecdysteroids can be detected in the hemolymph by radioimmunoassay. Values of 4000 ng/ml and more (Marcaillou et al., 1977) have been recorded. From ultrastructural observations of the testis, it is concluded that the temporary isolation of the gonial compartment is not based on newly formed junctions which could act as a barrier. At the last (ninth) day of the instar, the cysts are again open to HRP. The temporary closing of the open apical compartment in locusts, which corresponds with high ecdysterone titers, has raised much interest. Jones (1978) postulated that the molting hormone, as well as juvenile hormone, might be involved in barrier formation. In uitro experiments performed on follicles of young larvae of S . gregariu led Jones (1978) to conclude that ecdysterone is able to induce a premature formation of the hemolymph-testis barrier in the follicles of second-instargrasshoppers. In order to test Jones’ (1978) hypothesis, Marcaillou and Lauverjat (1986) transplanted premeiotic testicular follicles into the abdominal

MALE GERM CELL PROTECTIVE BARRIER

13 1

cavity of adult L. migratoria, whose hemolymph is known to be free of ecdysteroids (Lagueux et al., 1976). The survival of such transplanted follicles was shown by Lautie (1981). Marcaillou and Lauverjat (1986) found that the cysts of the transplanted follicles achieve spermatogenesis and form normal septate junctions and a tight compartment in an environment lacking ecdysterone, soon after the start of the meiotic process. A different function of ecdysterone was proposed earlier by Kambysellis and Williams (1971a,b), who postulated that the function of ecdysone is to further the penetrability of the testis walls and thereby to facilitate the entry of a macromolecular factor found in the blood of diapausing saturniids. This role of ecdysone is challenged by Friedlander (1989), whose results indicate that 20-hydroxy-ecdysone promotes in uitro spermatogenesis by sustaining functional integrity of the envelope cells. The role of ecdysone as either a barrier former or a barrier stormer has not been substantiated. The role of septate junctions as a transepithelial permeability barrier was reviewed by Noirot-Timothee and Noirot (1980) and by Lane and Skaer (1980). Molecular size is not the only parameter limiting the passage of HRP or lanthanum across septate junctions. Acidic polysaccharides found in the interseptal matrix can, by their anionic groups, bind cations whose presence in the intercellular space can, in turn, restrict the passage of charged molecules of any size (Wood and Kuda, 1980). In the cyst envelopes, the cells are associated by rare and loosely organized septate junctions, and freeze fracture demonstratesfocal tight junctions (Lane and Skaer, 1980) consisting of short rows of fused particles forming ridges on the P-face of the split membranes and corresponding furrows on the Eface. Similar structures have been observed in a variety of tissues (Lane, 1979).

The distribution of tracheolae in the testicular region of locusts and moths is different. In moths, large tracheae are found in the external layer of the follicular wall, while tracheolae penetrate its inner layer and reach the cavity of the follicle, but never penetrate into the interior of the cysts (Szollosi and Marcaillou, 1977; Szollosi et al., 1980; Szollosi, 1982). In locusts, the tracheal network penetrates between the follicles, but never enters them (Szollosi, 1982). It is noteworthy that the penetration of tracheolae into the testicular region parallels the depth of the formation of the barriers. In locusts, in which the barrier is in the inner follicular wall, the tracheolae reach the outer follicular wall, but do not penetrate further. In the moths, in which the barrier lies in the cyst envelope, the tracheolae penetrate into the interior of the follicle, but not into the cyst. In both groups, the tracheolae do not penetrate into the barred region.

FIGS.13-17. Leuantina hierosolyma (Mollusca). From O’Donovan and Abraham (1987); reproduced from “Somatic tissue-male germ cell barrier in three hermaphrodite invertebrates: Dugesia biblica (Platyhelminthes), Placobdella cosrara (Annelida), and Leuantina hierosolyma (Mollusca),” Journal of Morphology, 1987, 192, 217-227; with permission of Wiley-Liss Div. of John Wiley & Sons, Inc.

MALE GERM CELL PROTECTIVE BARRIER

133

I. MOLLUSCA In hermaphrodite gastropods, an MGPB was observed in several species, for example, L. stagnalis (de Jong-Brink et al., 1984; Bergmann et al., 1984a; Anelli et al., 1985) and L . hierosolyma (O’Donovan and Abraham, 1983, 1987) (Figs. 13-17). Luchtel(1972a,b) described a barrier between male and female compartments in three hermaphrodite gastropods: Arion ater rufus, Arion circumscriptus, and Deroceras reticulatum. In gonochoristic gastropods, MGPBs were described by Buckland-Nicks and Chia (1986) in Fusitriton oregonensis, Ceratostoma foliolaturn, and Littorina sitkana and by O’Donovan (1988) in Ampullaria canaliculata and Cerithium adansonii. The ovotestis of the hermaphrodite gastropods consists of several lobular acini with lumina connected to the hermaphrodite duct (Joosse and Reitz, 1969; de Jong-Brink et al., 1981, 1983). Each acinus is surrounded by a thick basement lamina (O’Donovan and Abraham, 1987). In the central part of the acinus is the compartment of male germ cells, separated from the peripheral layer of oocytes by a continuous layer of Sertoli cells which also protrude toward the lumen of the acinus (Luchtel, 1972a,b; de Jong-Brink et al., 1977). The luminal face of Sertoli cells has welldeveloped septate and gap junctions as well as desmosomes at the luminal terminals. The oocytes are surrounded by follicle cells, which are connected to them by desmosomes. The MGPB is formed by the inter-Sertoli septate junctions (de JongBrink et al., 1984; O’Donovan and Abraham, 1987). Bergmann et al. (1984a) observed that lanthanum penetrates the septate junctions, but is stopped by small tight junctions. They supposed that the barrier between the female and male compartments is based on tight, rather than septate, junctions. This view has been challenged by Buckland-Nicks and Chia (1986), who observed only septate and desmosome-like junctions between the Sertoli cells of three gonochoristic prosobranch species, which, according to these authors, form the MGPB. The basal lamina and the underlying connective tissue surrounding the

FIG. 13. Male and female compartments in an acinus of the ovotestis. Fo, Follicle cells; 00,oocyte, S, Sertoli cell; Sp, spermatozoa; Lum, acinar lumen. X8240. FIG. 14. Horseradish peroxidase, retained by the collagen space (CS), slightly penetrates into interfollicular (Fo) cell spaces (arrows). x 10,180. FIG. 15. Adjacent Sertoli (S)cells near the acinarlumen (Lum). Sp, Spermatozoa. x 3780. FIG. 16. Septate junctions (arrows) between adjacent Sertoli (S) cells. ~50,440. FIG. 17. Inter-Sertoli (S) gap junctions (arrows). X81,480.

134

MORDECHAI ABRAHAM

acini contain glycoprotein, collagenous proteins, myofibrocytes, and pigment cells (de Jong-Brink et al., 1984). In electron micrographs, the basal lamina is shown to consist of a lamina densa (0.6-2.0 pm) and a lamina rara (0.5-1.0 pm). Its role as a barrier was examined by several authors. In vitro incubations (de Jong-Brink et al., 1984) with different markers showed that the colloidal gold particles had not passed the lamina densa of the basal lamina; lanthanum nitrate and tannic acid passed the basal lamina and penetrated into the female compartment, up to the first septa of the septatejunctions between the cells lining the male compartment. The total length of the junctions was never crossed. Hill (1977)found that in the slug Agriolimax reticulatus neither ferritin nor HRP could be located within or between any of the cells of the ovotestis. He suggested that the pigment cells and fibroblasts, in conjunction with the basement membrane, formed a barrier to the tracers and possibly to all exogenous protein. In tissue of snails injected in the heart with HRP before being killed, the marker completely outlined the basement membrane, but failed to penetrate the ovotestis (O’Donovan and Abraham, 1987). Only when a very high dose (30 mg) was injected in short pulses over a longer period (45 minutes) did the marker penetrate the ovotestis. O’Donovan and Abraham presumed that the basement membrane forms a primary barrier, whereas the septate junctions between the Sertoli cells constitute a subsequent one, a situation similar to that known in mammalian testis (Dym and Fawcett, 1970). It is interesting that in two terrestrial species, A. reticulatus (Hill, 1977) and L. hierosolyma (O’Donovan and .Abraham, 1987), the basal lamina and the collagen spaces barred HRP from the ovotestis, while in two freshwater species, L. stagnalis (de Jong-Brink et al., 1984) and Viviparus viviparus (Griffond and Gomot, 1979), HRP crossed the basal lamina. In L. stagnalis, the widths of the basal lamina and the collagen spaces vary from 7 to 18 nm, while in L. hierosolyma it is 120 nm, and 100 nm in Helix aspersa (Guyard, 1971), another terrestrial species. The significance of this difference in size and tightness is not evident. The composition of the gonadal fluid versus that of the hemolymph in L. stagnalis was studied by de Jong-Brink et al. (1984). Gonadal fluid was collected by micropuncture from the central collecting ductus efferens. Gonadal fluid has higher osmolarity and a higher concentration of amino acids and proteins, Na+ is partly replaced by K+,and HC03- is almost totally replaced by C1-. The high osmolarity of the gonadal fluid attracts water. This induces a high transmural pressure, necessary to maintain the architecture of the gonadal system and/or to induce a water current, required for transport of sperm and oocytes. The agglutinin assay as described by van der Knaap et al. (1981, 1982) was applied in order to

MALE GERM CELL PROTECTIVE BARRIER

135

study the presence of agglutinins in L. stagnalis. No agglutinins were found in the gonadal fluid (de Jong-Brink et al., 1984). Luchtel(1972a,b) presumed that the barrier is necessary to isolate from each other male and female gonocytes in hermaphrodite snails. It has been shown that in gonochoristic male gastropods differentiation of male germ cells is under control of an androgenic factor. If the function of the barrier were to prevent the developing gonocytes from being influenced by hormones of the opposite sex, it might have been expected that in male gonochoristic snails the barrier would be absent (de Jong-Brink et al., 1984). However, Buckland-Nicks and Chia (1986)and O’Donovan (1988) described an MGPB in several gonochoristic species. An interesting aspect of the MGPB in mollusks has been discussed by O’Donovan (1988), who mentioned the study by von Medem (1945)of Diodora nubecula and the review by Fretter and Graham (1964). Sperm are packed into spermatophores in certain gastropods and cephalopods in which the gametes are surrounded by testicular epithelium and preserved in optimal conditions of immobility. The possible barrier structure of cells lining the spermatophores in other phyla, such as insects, has not yet been studied. J. ECHINODERMATA Sertolilike cells, the so-called interstitial cells, were found in the testes of different species in all classes of echinoderms, but no occluding junctions in the inter-Sertoli cell spaces have been observed. The somatic region surrounding the male germ cells has been studied in Asteroidea, Echinoidea, Ophiuroidea, Holothuroidea, and Crinoidea (Holland, 1967; Longo and Anderson, 1969;Atwood, 1973;Worley et al., 1977;Hinsch and Dehn, 1979;Bickell et al., 1980;Walker, 1980;Buckland-Nicks et al., 1984). More specifically, the presence of a somatic tissue-male germ cell barrier in the testes of Asteroidea and Ophiuroidea was studied by Chia and Buckland-Nicks (1987) by mixing lanthanum nitrate with glutarparaformaldehyde fixative. The somatic cells in the testes of echinoderms are similar to Sertoli cells, because they provide mechanical support for spermatogenesisand phagocytose residual bodies as well as waste sperm. In mammals, the population of Sertoli cells is fixed (Fawcett, 1975). In pulmonate mollusks (de Jong-Brink et al., 1977) and echinoderms (Chia and Buckland-Nicks, 1987), there occurs a cycle of apoptosis and replacement. However, no permeability barrier was observed between the spermatogenic cells and the hemal system or the somatic cells in general. Lanthanum nitrate infiltrated the inter-Sertoli region and penetrated into the testicular lumen. Chia and Buckland-Nicks (1987)concluded that ophi-

136

MORDECHAI ABRAHAM

uroids have no barrier, while asteroids may have an incomplete permeability barrier made by desmosome-like junctions between cells of the spermatogenetic epithelium. They suggested that the blood-testis permeability barrier evolved together with internal fertilization and they would not expect to find a BTB in animals that shed gametes into water. This is an original viewpoint. However, in the majority ot teleost species studied up to now, as well as in the Anura, in which an MGPB was found, external fertilization occurs (Section 111,K).The electron micrographs published by Chia and Buckland-Nicks (1987), which show the penetration of lanthanum between the Sertoli cells and into the testicular lumen, were possibly made before the spawning season, since no spermatozoa can be seen in the illustrations. Nor do the authors mention whether the animals had ripe spermatozoa at the time the experiments were done.

K. CHORDATA 1 . Cephalochordata

In the lancelet, Branchiostoma floridue, there are several dozen metamerically arranged separate testes, each roughly l mm in diameter (Holland and Holland, 1989). Between the visceral peritoneum and the germ cells, there is a hemal layer, bounded by a 40-nm-thick basal lamina, but not by cellular endothelia. Spermatogonia are in contact with this basal lamina, but no pinocytosis between them was observed. Between the germ cells, supporting cells are found, with probably phagocytotic and possibly steroidogenic function. The supporting cells are widely scattered without cell-to-cell contact. The germ cells up to the spermatid stage are interconnected by intercellular bridges. According to the presented evidence, the lancelet testis has no MGPB. Whether the basement lamina surrounding the hemal vessels or the closely structured and interconnected germ cells themselves may restrain the flow of macromolecules could be ascertained only by experimentation with intercellular tracers. 2 . Cyclostomata In Myxine glutinosa, during the spermatogenetic process, testicular follicles are surrounded by a layer of Sertoli cells interconnected through numerous indentations of the peripheral cytoplasm (Alvestad-Graebner and Adam, 1977). No occluding junctions were described; however, the cell apices of neighboring cells have desmosome-likestructures. Enclosing the Sertoli cell layer is a basal lamina and a layer of connective tissue. The authors did not apply tracer techniques to determine whether the flow of

MALE GERM CELL PROTECTIVE BARRIER

137

macromolecules inside the testis is impeded, nor were the possible changes in the inter-Sertoli space followed during the whole spermatogenetic process. 3. Teleostei The structure of the MGPB in teleosts was described in several species: Aphanius dispar (Abraham et al., 1979a, 1980) (Figs. 18-22), Poecilia reticulata (Marcaillou and Szollosi, 1980; Bergmann et al., 1984b), Abudefduf marginatus (Mattei et al., 1982), Oryzias latipes (Shibata and Hamaguchi, 1986), Ameca splendens (Bergmann et al., 1984b), Cyprinus carpio (Parmentier et al., 1985;Timmermans et al., 1985), and 0 .niloticus (Lou and Takahashi, 1989). The structural pattern of the teleost testes was studied by Grier (1975, 1976, 1981), who distinguished between the restricted spermatogonial (cyprinodont) testis type, in which the spermatogonia are confined to the terminus of the testis tubule, and the unrestricted spermatogonial (cyprinid) testis type, in which the spermatogonia are along the entire length of the tubule. The restricted type corresponds to the tubular type of Billard (1986), while the unrestricted corresponds to Billard’s lobular type. In both types, the Sertoli cells are at the periphery of the cysts in contact with the basal lamina and form a complete layer around the gonocytes. Germ cells have no contact in teleosts with the basal lamina (Mattei et al., 1982). The morphological basis for the barrier in vertebrates consists of junctional complexes between the Sertoli cells, which constitute the wall of the testicular tubules, and between the myoid cells which surround them (Dym, 1973; Gilula et al., 1976). Myoid cells with densely packed microfilaments are located externally to the Sertoli cells. These cells form, in teleosts, a discontinuous layer (Gresik et al., 1973). Shibata and Hamaguchi (1986) distinguished, in the testes of 0. latipes, between type A spermatogonia, which are separated from each other by the cytoplasm of the Sertoli cells, and type B spermatogonia, which are organized in cysts. The germ cells from cysts are connected to each other by intercellular bridges and are at a synchronous stage of spermato- or spermiogenesis. Tight junctions between the Sertoli cells in testes of several freshwater teleost species were observed in 1972 by Billard et al. However, the concept of a barrier in fish testis between blood or somatic cells and gonocytgs was not pronounced at that time. The flow of macromolecules in the testicular interstitium was studied with the help of HRP, lanthanum chloride, or bovine serum albumin as tracers, injected in uiuo (Abraham et al., 1979a, 1980; Parmentier et al., 1985; Shibata and Hamaguchi, 1986; Lou and Takahashi, 1989) or incubated in uitro (Marcaillou and Szollosi, 1980; Bergmann et al., 1984b;

FIGS.18-22. Aphanius dispar (Teleostei). The figures are from specimens injected with horseradish peroxidase. From Abraham er al. (1980); reproduced from Cell Tissue Research, 1980,211,207-214 by permission of the publisher, Springer, Heidelberg.

MALE GERM CELL PROTECTIVE BARRIER

139

Shibata and Hamaguchi, 1986). In cysts of A. dispar testis, containing ripe spermatozoa, adjoining Sertoli cells display elaborate junctional complexes consisting of tight junctions with fused outer leaflets of the adjoining membranes, and alternating with these structures there are desmosomes. Along the junctional complexes, the adjunctional cytoplasm contains bundles of filaments. HRP injected in uiuo does not penetrate the occluding junctions. In younger cysts, including cysts with early spermatids, the marker penetrates freely beyond the Sertoli cell layer to surround the gonocytes (Abraham et al., 1979a, 1980). The endothelium of the capillaries and the interstitial cells are filled with numerous labeled pinocytotic vesicles, also displayed by both myoid and Sertoli cells. In 0. latipes, the tracers are visible inside young cysts, among type B spermatogonia, spermatocytes, or early spermatids (Shibata and Hamaguchi, 1986). There is no barrier in different teleost species throughout the meiotic process, the barrier appears toward the end of spermiogenesis (Abraham et al., 1979a, 1980; Marcaillou and Szollosi, 1980; Parmentier et al., 1985;Timmermans et al., 1985; Shibata and Hamaguchi, 1986). According to Bergmann et al. (1984b), the Sertoli tight junctions have appeared already at the time meiosis is complete. Parmentier et al. (1985) injected HRP and carp immunoglobulin G (IgG) monoclonal antibodies intracardially into the common carp, and found 24 hours later that both markers penetrated into testicular tissue and traversed the limiting cell layers of cysts containing spermatogonia, spermatocytes, or spermatids. The markers were concentrated around the immature germ cells, but did not penetrate in the lumina of the cysts containing mature spermatozoa. Injected IgM left the capillaries in only small amounts and were restricted to the interstitial region of the testes. In the testes of normal 0. niloticus (Lou and Takahashi, 1989), HRP was arrested by the inter-Sertoli junctions, while bovine serum albumin

FIG. 18: Capillary with erythrocyte (ery) from the testicular stroma; endothelium with intense pinocytotic activity. The intercellular spaces send “peg protrusions” into the Sertoli (Sert) cells (arrows). ~ 5 4 0 0 . FIG. 19. Myoid cell (MY) with pinocytotic vesicles; between myoid and Sertoli cells (SERT) is the enlarged intercellular space. NU, Nucleus; peg, protrusion of the intercellular space. ~ 2 7 , 0 0 0 . FIG. 20. Cell from testicular stroma with numerous pinocytotic vesicles in proximity of a stromal capillary. X 14,400. FIG. 21. The junctional complex between adjacent Sertoli (SERT) cells (arrows). MY, Myoid cell; peg, protrusions of the intercellular space into the Sertoli cells. x 16,200. FIG. 22. The junctional complex of adjacent Sertoli (SERT) cells near the cyst lumen (Lum) contains bundles of filaments in the subjacent cytoplasm. X 18,450. (Inset) The junctional complex at higher magnification. X28,800.

140

MORDECHAI ABRAHAM

was prevented from passing by the basement membrane which surrounds the spermatogenetic cysts. The injected HRP, in A . dispar marks the route of the extravascular circulation. The extravascular spaces originating in the pericapillary spaces form an extensive network penetrating the entire testis (Figs. 18-22). The most conspicuous element of this network is the 100- to 200-nm-wide basement membrane space (Abraham et al., 1980) between the myoid and the Sertoli cells, which possibly forms and additional biochemical BTB (Gravis et al., 1977). In P . reticulata (Marcaillou and Szollosi, 1980), testicular cysts containing gonia are limited by a single layer of flat cells, while the wall of cysts containing spermatids has developed into a thick envelope. Actually, the envelope consists of Sertoli cells whose apical membranes are deeply indented by the heads of the elongating spermatids, similar to the Sertoli cell-germ cell contact in mammals. In testicular fragments incubated with HRP, the tracer traverses the limiting layer of the young cysts, penetrates into the intercellular spaces, and bathes all the gonia of the cyst. It is captured in small amounts by the germ cells by means of micropinocytotic vesicles. In older cysts, the tracer is blocked between the inter-Sertoli cell spaces. The cytoarchitecture of the testis in Goodeidae resembles that of Poeciliidae, probably connected to the mechanism by which naked unencapsulated sperm bundles or spermatozeugmata are formed (Grier et al., 1978). Differences in chemical composition between blood and seminal plasma in different teleost species were reported by several authors. According to Steyn and Van Vuren (1986), in Clarias gariepinus, pH and potassium values of seminal plasma are significantly higher compared to that of blood plasma, while calcium, magnesium, and chloride are significantly lower. Fructose is totally absent in the seminal plasma, while glucose, total protein, total lipids, and alkaline phosphatase are significantly lower. Higher values of potassium in seminal plasma compared to blood plasma values were also observed by Morisawa et al. (1979) for Carassius auratus, Cyprinus carpio, and Salmo gairdneri. In the study by Vlok and Van Vuren (1988), in Barbus aeneus, significant differences between seminal plasma and blood plasma were evident in the glucose, total protein, total lipids, sodium, potassium, pH, magnesium, chloride, and calcium concentrations. 4 . Amphibia An MGPB was described by Franchi et al. (1982)in Triturus carnifex, by Bergmann et al. (1983) in Rana esculenta and Salamandra salamandra, by Cavicchia and Moviglia (1983) in Bufo arenarum, by Bergmann et al.

MALE GERM CELL PROTECTIVE BARRIER

141

(1984b) in Xenopus laeuis and Bufo bufo, and by O’Donovan (1988) in Rana ridibunda. The organization of the testes in Anura and Urodela is similar; the germ cells are arranged in cysts resembling their grouping in the teleost testes. Here, as in teleosts, the tight junctions are regularly associated with desmosomes (Bergmann et al., 1984b) or form a complex together with septate-like junctions, both of which are associated with subsurface cisternae of granular ER. In X . laeuis, smooth muscle cells form an incomplete sheath around seminiferous tubules, while Rana temporaria and B . bufo lack such a peritubular contractile layer (Unsicker, 1975). Bergmann et al. (1983) observed punctate tight junctions between Sertoli cells which surround male germ cell cysts, in both frogs and salamanders. Working with lanthanum nitrate tracer, they concluded that the Sertoli barrier appears at the end of meiosis, as in teleosts. Spermatogonia and spermatocysts are in open compartments, and in S . salamandra the cysts containing early spermatids are still penetrated by the tracer. Only late spermatids and spermatozoa are shielded by the barrier. In later work, using hypertonic fixatives together with lanthanum, Bergmann et al. (1984b) observed that young cysts are open and show shrinkage artifacts, as the effect of the hypertonic fixative which they use. In Anura, Bergmann et al. (1984b) also describe desmosomes between the Sertoli cells, which are nonexistent in urodeles, according to their observations. Apparition of the barrier after meiosis was also observed by Franchi et al. (1982) in Triturus cristatus carnifex. The only discordance to this general conclusion about the timing of the barrier formation is that of Cavicchia and Moviglia (1983), who observed in B. arenarum an earlier apparition of the barrier, similar to that occurring in mammals.

5 . Reptilia The MGPB in reptiles was demonstrated by Baccetti et al. (1983) in Lacerta muralis and Lacerta sicula, and by Bergmann et al. (1984b) in Anolis carolinensis. In reptiles, as in other Amniota, the barrier is established shortly after the onset of meiosis. Focal inter-Sertoli tight junctions prevent the penetration of lanthanum to germ cells in zygotene or later stages. These tight junctions are associated with subsurface cisternae of granular ER. In L. muralis and L . sicula, the Sertoli-Sertoli junctional complexes contain tight-, gap-, and septate-like junctions. The Sertoli occluding junctions are near the base of the seminiferous epithelium. There are also desmosome-like junctions surrounded by 7-nm filaments, which appear after the mating period. According to Baccetti et al. (19831, the cytoskeleton of the Sertoli cells in lizards contains vimentin, abundant

142

MORDECHAI ABRAHAM

actin, and microtubules in the cell periphery; however, no actin layer was observed between the plasma membrane and the subsurface cistern, as is known in mammalian Sertoli cells (Section IV,A). 6 . Aves An MGPB in birds was reported in Gallus domesticus by Cooksey and Rothwell (1973), Osman et al. (1980), Bergmann and Schindelmeiser (1987), and Pelletier (1990); in Taeniopygia guttata and Lonchura striata by Bergmann et al. (1984b); in Coturnix coturnix by O’Donovan (1988); and in Anasplatyrhynchos by Pelletier (1990). In the avian testes, between adjacent Sertoli cells, focal tight junctions associated with subsurface cisternae of ER are present. In addition, septate-like junctions basal to the tight junctions (Bergmann et al., 1984b) were found. These septate junctions appear as lucent bridges on an electron-dense background of intercellular lanthanum. They are also associated with subsurface cisternae. The formation of the barrier in the testes of G . domesticus was correlated with the occurrence of early spermatids (Bergmann and Schindelmeiser, 1987). In T. guttata and L . striata, lanthanum reaches spermatogonia, but is prevented from surrounding primary leptotene spermatocytes (Bergmann et al., 1984b). Pelletier (1990) studied the Sertoli occluding junctional complex in a seasonal breeding bird, A . platyrhynchos, and a nonseasonal bird, G . domesticus. The junctional complex, along the long axis of the Sertoli cells, is formed by occluding, gap, and adhering junctions. No septate junctions were observed. The occluding junctions were made up of continuous junctional strands forming an occluding zonule and discontinuous strands forming focal occluding junctions. 7 . Mammalia Structural and functional aspects of the MGPB were examined in several mammalian species (Figs. 23-28): in humans by Koskimies et al. (1973), de Kretser et al. (1975), Camatini et al. (1979), and Bergmann et al. (1989); in species of monkey by Dym (1973) and Camatini et al. (1982b); in the dog by Connell(l978,1980); in the mink by Pelletier (1986,1988);in the bull by Wrobel and Schimmel(l989); in the sheep by Setchell(l967); in the swine by Toyama (1976); in the squirrel by Vogl and Soucy (1985); and there has been a considerable number of papers on the barrier in rats, mice, guinea pigs, and hamsters (reviewed by Setchell and Waites, 1975; Nagano and Suzuki, 1983). The BTB in boars was reported by Yazama et al. (1988). Investigations of the structure and function of the Sertoli cell barrier

MALE GERM CELL PROTECTIVE BARRIER

143

based on the mammalian testis are presented in different sections of this chapter (Sections IV and V). A classical study of the Sertoli cell occluding and gap junctions in mature and developing mammalian testis has been done by Gilula, Fawcett, and Aoki (1976), who examined with freezefracture techniques the BTB in Sprague-Dawley rats. The results are summarized as follows: (1) The inter-Sertoli occluding junctions differ from “common” tight junctions, appearing near the base of the seminiferous epithelim; (2) there are up to 40 parallel circumferentially oriented rows of intramembrane particles; (3) there is no anastomosis between rows; (4) the rows of particles are preferentially associated with the Efracture face (B-fracture face of the former terminology); (5) adjacent to the occluding junctions are typical gap junctions in the immature and atypical gap junctions in the mature testis; and (6) the occluding junctions are resistent to dissociation by hypertonic solution. Sertoli cell junctions in different mammalian species are similar to those in rats or mice (Nagano and Suzuki, 1976). A dissent to this uniformity is presented in the results of Connell(l978) in the dog and of Camatini et al. (1979,1981) in the human male, where septate junctions were found between adjacent Sertoli cells.

IV. Structure of the Male Germ Cell Protective Barrier A. STRUCTURES OF THE SERTOLI CELLAND THE OCCLUDING JUNCTIONS

The structure of the Sertoli cell has been studied by many authors (reviewed by Russell and Peterson, 1985). The cells are irregularly columnar in shape and are anchored to the basal lamina by hemidesmosomes (Connell, 1974; Russell, 1977b). Hypertonic fluids cannot separate Sertoli cells from the basal lamina through cell shrinkage (Gilula et al., 1976; Russell and Peterson, 1985). Three-dimensional stage V Sertoli cell model reconstructions have been made by Russell and Wong (1981), Wong and Russell (1983), and Weber et al. (1983). The cell dimensions, as determined from the model, are about 90 pm in the axis from the periphery of the seminiferous tubule toward its center, 41 pm in the longitudinal axis of the tubule, and 30 pm in the width of the Sertoli cell. The Sertoli cell architecture is stabilized by an intricately organized cytoskeleton composed of microtubules and microfilaments (Christensen, 1965; Pollard, 1980; Griffith, 1980). Sano et al. (1987) found a very high level of tubulin in mouse

MALE GERM CELL PROTECTIVE BARRIER

145

Sertoli cells. The effects of treatment with microtubule disrupting drugs, such as colchicine, colcimide, or vinblastine, on Sertoli cell function were studied by several authors in different mammalian species (Rattner, 1970; Handel, 1979; Aoki, 1980; Russell et al., 1981). Such treatment destroyed the microtubular framework of the Sertoli cell. In the cytoplasm, opposite the tight junctions, there are subsurface cisternae of ER, and bundles of microfilaments, the so-called ectoplasmic specializations (ESs). ESs are hexagonally packed bundles of actin filaments (Toyama, 1976) situated between the plasmalemma and the ER cistern (Franke et al., 1978) at sites adjacent to tight junctions (Fawcett, 1975)and zones of contact with spermatogenic cells. Actin can be detected immunocytochemically on fixed and embedded tissue (Bussolati et al., 1980). The function of Sertoli cell actin is a hotly debated subject. It was first thought that the ESs are contractile and contain ATPase activity (Gravis et al., 1976; Toyama, 1976). Vogl and Soucy (1985) were able to prove that ES filaments in the squirrel contain actin, but not myosin, that glycerinated models did not contract in the presence of exogenous ATP and Ca2+,and that ES filaments are linked to the general cell infrastructure (Russell, 1977b). Their role is to support the plasmalemma adjacent to occluding junctions (Grove and Vogl, 1989) and to regulate the movement of Sertoli cell surface during migration of germ cells from the basal to adluminal compartment (Russell and Peterson, 1985). Russell (1977b) has suggested that ESs may contribute to maintaining the shape of Sertoli cell apical crypts in which spermatids mature. The disappearance of ESs, during spermatogenesis or by experimental treatment, is correlated with changes in the function of the occluding junctions (Russell, 1977b). Weber

FIGS. 23-24. Musfela uison (mink). From Pelletier (1988). Reproduced from "Cyclic modulation of Sertoli cell junctional complexes in a seasonal breeder: The mink (Mustela vison)," The American Journal of Anatomy, 1990, 183, 68-102, with permission of WileyLiss Div. of John Wiley & Sons, Inc. Fig. 25. Guinea pig (Mammalia). From Pelletier and Friend (1986). Reproduced from Journal of Andrology, 1986,7, 127-139, with permission of J. B. Lippincott Co. FIG. 23. Freeze-fracture replica of a discontinuous zonule in an adult mink during the active spermatogenic phase with meandering strands of particles (open arrows) and intercalated gap junctions (arrows). x46,920. FIG. 24. Freeze-fracture replica of a continuous zonule in an adult mink during the active spermatogenic phase. Junctional particles are ubiquitous (arrows) and are associated preferentially with grooves on the E-face. Most rows of particles are continuous and juxtaposed. ~82,800. FIG. 25. Freeze-fracture replica of adult testis perfused with a filipin-fixative solution. In the occluding zonule, composed of uninterrupted strands, no filipin penetrated. Clusters of filipin perturbations (arrows) appear in membrane domains free of junctional strands. x45.080.

FIGS. 26-28. Guinea pig (Mammalia). Figs. 26 and 27 are from specimens treated with gossypol. From Pelletier and Friend (1986). Reproduced from Journal of Andrology, 1986,7, 127-139, with permission of J. B. Lippincott Co. Fig. 28 is reproduced from "The Sertoli cell junctional complex: Structure and permeability to filipin in the neonatal and adult guinea pig," The American Journal of Anatomy, 1983,168,213-228.

MALE GERM CELL PROTECTIVE BARRIER

147

et al. (1988) intratesticularly injected cytochalasin D (CD), a microfilament

inhibitor which disrupted the ES microfilaments. The effect of CD on the structure of the junctions was followed by freeze-fracture technique, while the leakiness of the barrier was shown by tracers such as lanthanum hydroxide, antiserum against haploid germ cells, and radiolabeled inulin, which entered the closed compartment in higher amounts than in control animals. Proposed first by Russell (1977a, 1978), it became a generally held idea (Dym and Cavicchia, 1977; Bergmann, 1987) that, during the migration of spermatocytes from the open to the closed compartment, the junctional complexes, prior to breaking down above migrating germ cells, are reformed below them, thereby maintaining the integrity of the BTB. For a short period, the spermatocytes are thus enclosed from both above and below, in the so-called intermediate compartment. In birds, too, (Bergmann and Schindelmeiser, 1987), the existence of an intermediate tight compartment for leptotene primary spermatocytes has been suggested. This view has been challenged by Cavicchia and Sacerdote (1988), who injected electron-dense tracers together with fixatives directly into the seminiferous tubules, or the rete testis of rats, and at the same time perfused tracer through the spermatic artery. The marker thus penetrated the tubuli from both the tubular lumen side and the interstitium side, toward the barrier. The result was that all the germ cells were surrounded by the tracer. No leptotene spermatocytes devoid of tracer have been observed, indicating therefore that only two distinct compartments are present; there is no intermediate compartment. However, Cavicchia and Sacerdote (1988) could not rule out the possibility of a momentary intermediate compartment-a transit chamber (Russell and Peterson, 1985). The absence of an intermediary compartment was also shown by Ross (1977) and by Pelletier and Friend (1983, 1986).

FIG. 26. Freeze-fracture replica of a Sertoli cell junctional membrane from a 41-day-old gossypol-fed guinea pig. The discontinuous strands, far apart even at the base of the epithelium, permitted the passage of filipin. ~30,940. FIG. 27. Freeze-fracture replica of an apical Sertoli cell junctional area from a gossypoltreated adult guinea pig. Distensions separate strands that offered no resistance to the filipin tracer introduced to both the vascular and luminal poles. X21,840. FIG.28. Close to the spermatogonia, the intercellular cleft between adjacent Sertoli cell membranes is occasionally filled with amorphous material (*). This portion of a Sertoli cell junctional complex from a normal sexually mature guinea pig shows no cellular contacts. Filipin-induced membrane perturbations appear as corrugations (arrows) on the adjacent plasma membranes. ~ 7 5 , 5 3 0 .

148

MORDECHAI ABRAHAM

B. SERTOLICELLJUNCTIONS DURING GROWTH, ANNUAL REPRODUCTIVE CYCLE,AND SPERMATOGENETIC PROCESS In the rat, occluding junctions between Sertoli cells take shape between 16 and 19 days of age (Vitale et al., 1973; Dym and Fawcett, 1970). At this time, the tubular lumen also appears. The start of secretion of ABP by rat Sertoli cells into the lumen of the seminiferous tubules correlates well with the formation of the barrier and the tubule lumen. ABP appears in the tubular lumen 14 days after birth, and at 18-20 days of age, it is found in the caput epididymidis (Tindall et al., 1975). The functional barrier to watersoluble markers develops later and more gradually than the barrier to electron-opaque markers (Setchell et al., 1988). The correlation between the formation of the barrier and that of the lumina in the seminiferous tubuli has been observed in several species (Flickinger, 1%7). In the mink (Mustela vison), a seasonal breeder, the Sertoli cell barrier and the tubular lumen are formed at the same time, by days 220-240 after birth (Pelletier, 1986). Pelletier divided the spermatogenetic cycle into 12 stages, similar to those described in rodents by Leblond and Clermont (1952a,b) and by Clermont (1960; reviewed by Russell, 1980). At the peak months of the active spermatogenetic phase, the BTB separates spermatogonia and young spermatocytes from the older generation of germ cells. In the hamster (Mesocricetus auratus), at the age of 25 days, the Sertoli cell barrier is established. However, the testicular lumen is formed a few days earlier (Vignon et al., 1985).In 31- and 41-day-old guinea pigs, meiosis was complete, and the tubuli had lumina (Pelletier and Friend, 1983,1986). The Sertoli cell junctional complex was fully developed and showed the characteristics of an adult testis, including intermediate, gap, and occluding junctions. In Holstein bulls (Curtis and Amann, 1981), the lumen in the seminiferous tubules appears from 4 to 5 months of age, and the Sertoli cell barrier, 1 month later. In humans, lanthanum tracer is blocked by the Sertolijunctions at puberty (Furuya et al., 1980),when primary spermatocytes also appear. In the domestic fowl too, an effective barrier in the testes appears at puberty (Bergmann and Schindelmeiser, 1987). Several authors studied the formation of the Sertoli occluding junctions with freeze-fracture techniques in different mammalian species (Vitale et al., 1973; Nagano, 1980; Nagano and Suzuki, 1976, 1978, 1983; Gilula et al., 1976; Meyer et al., 1977; Pelletier and Friend, 1983). In embryonic mouse testes (days 14-20 of gestation), junctional particles are located in the center of the grooves in the E-face of Sertoli cells. In addition, there are ridges and grooves devoid of particles, presumably an initial stage in the formation of occluding junctions of the Sertoli cells (Nagano and Suzuki,

MALE GERM CELL PROTECTIVE BARRIER

149

1978). In the peritoneal mesothelium of the mouse embryo, the initial sign of tight junction formation is the close apposition of adjacent cell membranes in the junctional domain without tight junctional particles (Suzuki and Nagano, 1979). In the dog testis, until prepuberty, the intramembranous particles on the Sertoli cell E-face are disorganized, and the interSertoli cell space is permeable to tracers (Connell, 1980). As the interior of the seminiferous epithelium becomes impermeable to tracers, the junctional rows display a parallel alignment and become circumferentially arranged around the cell. Ross (1970) demonstrated that adjacent Sertoli cells form a continuous contact zone. The junctional specializations in this zone were investigated by Flickinger and Fawcett (1967). Coincidently with the development of a tubular lumen and the establishment of a competent BTB, particulate elements associated with the E-face (not the P-face, as in other epithelia) become organized into junctional strands. In the basal region of the inter-Sertoli space of different mammalian species, Pelletier and Friend (1983) and Pelletier (1986, 1988) distinguished, during spermatogenesis, between “continuous zonules” of the junctional strands, which block the passage of protein tracers, and “discontinuous zonules,” which lie near the tubular lumen and are incompetent in blocking entry of tracers into the seminiferous epithelium. The development of continuous zonules coincides with the initiation of spermatogenesis. Over 100 rows of junctional strands were found in the junctional complexes of two adjacent Sertoli cells (Russell and Peterson, 1985). Parvinen (1982) presumed that there is a functional cycle of the Sertoli cells. Cyclic modulation of Sertoli cell junctional complexes in seasonal breeders was studied by Pelletier (1986, 1988, 1990) with freeze-fracturetracer techniques. In the newborn mink, split-membrane particles are associated with both P- and E-faces. Coincidently with the development of a tubular lumen and the association of the particles of the junctional strands with the E-face, the barrier becomes functional (Figs. 23 and 24). In the adult mink, cell junctions occur on the entire lateral Sertoli cell plasma membrane from the base to the apex, during the active spermatogenetic phase. Pelletier (1986, 1988) distinguished three structurally and functionally different regions of these cell junctions. In the basal one-third of the Sertoli cell, near the basal lamina, tight, gap, and adhering junctions were present, which proved permeable to tracers. In the middle one-third, parallel uninterrupted rows ofjunctional particles, the impermeable continuous occluding zonules and adherens junctions were observed. In the luminal one-third, the zonules were discontinuous, being made up of meandering and broken rows of particles; they were permeable to tracers. Pelletier (1988) proposed that the formation of the Sertoli cell junctional

150

MORDECHAI ABRAHAM

complexes begins at the base of the cell and is dismantled at its apex, follows the direction of germ cell migration, and opposes the apicobasal direction of junction formation reported for most epithelia. During the inactive spermatogenetic phase, the junctional strands resembled those of the newborn mink and were incompetent in blocking the entry of HRP into the seminiferous tubules. The timing of occludingjunction formation during pubertal development and in adult animals during testicular recrudescence is similar. In a seasonal breeding duck, A. platyrhynchos, the Sertoli junctional complex during spermatogenesis resembles that of the mink and is formed by occluding, gap, and adhering junctions (Pelletier, 1990). However, in the duck, 7-nm filaments were associated with the occluding junctions only during testicular regression. The junctional complex was composed basally by forming focaljunctions, apicolaterally by an occluding zonule, and toward the apical side by dismantling focal junctions. During the breeding season, the apical region of the Sertoli cells expands to accommodate a large number of germ cells, and this creates the illusion that the barrier is more basally situated than it is. This becomes apparent at the end of the breeding season, when the Sertoli apical projections disappear. The degradation process of the occluding junctions was investigated by Russell (1979a,b), who described tubular evaginations of Sertoli cells 24 pm in length, with bulbous distal dilations which penetrate into an adjacent Sertoli cell. These so-called “tubulobular complexes” may be structures connected with the dissolution process of old junctional contacts (Russell and Clermont, 1976; Russell, 1979a). The formation of the BTB in the mammalian testis relative to spermatocyte maturation was studied by Russell (1978), Ross (1977), Dym and Cavicchia (1977, 1978), Connell (1980), Bergmann and Dierichs (1983), Sun and Gondos (1986), and Pelletier (1986, 1988). In the rat and the rabbit, the spermatocytes still found in the open compartments are at early leptotene prophase; in the monkey, at late leptotene; and in the dog and the mink, at early zigotene prophase. C. DESMOSOMES AND GAPAND SEFTATEJUNCTIONS BETWEEN BARRIER-FORMING CELLS 1. Desmosomes

The functional complex of most epithelia includes a desmosome and/or a zonula adherens (Farquhar and Palade, 1963). Nagano and Suzuki (1983) have not observed desmosomes between adjacent Sertoli cells of the adult mammalian testis, nor have they been observed in a reconstructed Sertoli cell (Russell and Wong, 1981; Wong and Russell, 1983). However, Wong

MALE GERM CELL PROTECTIVE BARRIER

151

and Russell (1983) found in type B configuration of Sertoli cells apically positioned desmosomes which secure the delicate luminal extensions that grasp the spermatid heads. Desmosome-like junctions between Sertoli cells and germ cells were also described by Gravis (1979). Inter-Sertoli cell desmosomes were described in the teleostean and amphibian testes by Abraham et al. (1980) and by Bergmann et al. (1984b) and in the rooster by Cooksey and Rothwell(l973). In desmosomes, there is a dense accumulation of 10-nm subsurface filaments. Some desmosomes show a faint intermediate line (Pelletier and Friend, 1983). In zonulae adherentes, subsurface densities and 10-nm filaments are periodically spaced along the interface of two Sertoli cells. Both types of desmosomes are most frequently seen at the adluminal extent of the rows of tight junctions (Pelletier and Friend, 1983).

2 . Gap Junctions Adjacent Sertoli cells develop gap junctions during fetal life (Nagano and Suzuki, 1978), prepubertal life (Camatini et al., 1982b), and puberty as spermatogenesis is initiated (Gilula et al., 1976; Nagano and Suzuki, 1976; Meyer et al., 1977; Camatini et al., 1982b). Gap junctions are found between tight junctional strands, mainly in the developing animal. McGinley et al. (1979) described gap junctions between Sertoli cells and spermatogonia and spermatocytes. Gap junctions are the sites for metabolic and ionic coupling (Gilula et al., 1972)and also for the coordination of activities between Sertoli and germ cells in rats (McGinley et al., 1979; Palombi et al., 1980). In several lepidopteran species, it was shown that numerous gap junctions appear between the male germ cells and the somatic cells of the cyst envelopes in A. kuehniella (Szollosi and Marcaillou, 1980) and in two other moths, Platysamia Cynthia and Hyalophora cecropia (Szollosi, 1982). Lawrence et al. (1978) demonstrated that hormonal stimulation can be transmitted between two cells belonging to different cell types by means of CAMPdelivered from one cell to another through gapjunctions. Both cyst cells in insects and Sertoli cells in mammals form the MGPB and at the same time presumably transmit molecular messages related to the function of the germ cells. Gap junctions, together with septate junctions and desmosomes, were described by O’Donovan (1988) between adjacent Sertoli cells, as well as gap junctions between Sertoli cells and germ cells in a gonochoristic mollusk, C. adansonii. 3 . Septate Junctions It is generally admitted that the barrier-forming junctions in invertebrates are of the septate type, while in vertebrates, these are the tight junctions (Gilula, 1973; Staehelin, 1973, 1974; Noirot-Timothee and Noirot,

152

MORDECHAI ABRAHAM

1980). Connell (1978) argued that paired parallel rows of junctions in canine testes are true septate junctions distinctly different from the nearby tight junctions. They develop prior to tight junctions and appear in thinsection micrographs as septa crossing the intercellular space, but do not exclude lanthanum, as do tight junctions. Camatini et al. (1979, 1981) demonstrated septate junctions in human testis. Nagano and Suzuki (1983) were skeptical that the paired junctional rows are septate and found it difficult to compare the mammalian Sertoli septate junctions with those found in invertebrates. Bergmann et al. (1984b) described in the avian testis both septate-like and tight junctions. In a recent comprehensive freeze-fracture study of guinea pig testis Sertoli junctions (Pelletier and Friend, 1983), septate junctions were not mentioned. Concomitance between tight and septate junctions between the somatic cells in insect testes was noted by Toshimori et al. (1979), Szollosi (1982), and Schweich and Leloup (1988). In an insect, T. infestans, Miranda and Cavicchia (1988) described pleated and smooth septate junctions (Section 111,H). Coexistence of gap and septate junctions in the hepatic caecum of Duphnia was noted by Hudspeth and Revel (1971), while simultaneity of smooth and pleated septate junctions in malpighian tubule epithelium was reported by Meyran (1982).

D. MGPB IN THE EXCURRENT DUCTEPITHELIUM OF THE TESTES

Is there a barrier between the lumen and the external region of the excurrent ducts, and how strong is it? This is a fundamental question, since the proximity of the efferent ductules to the seminiferous tubules might reduce the effect of the tubular barrier if the ductules were permeable. The efferent ducts, from the testes to the outside, are differently structured in the various animal groups. In some groups, these are short ducts lined by a unique cell type, while in others, such as mammals, the ducts are formed by separate units, which have different structures and functions. A review of the structure and function of the epithelium lining the different ducts which connect the testes of the rat to the exterior can be found in the report by Hamilton (1975). Cell junctions in the mammalian seminiferous tubules and different sections of the excurrent ducts were studied by Suzuki and Nagano (1978) and by Nagano and Suzuki (1983) in freeze-fracture replicas and in lanthanum-perfused thin sections. There is a notable difference between the inter-Sertoli space in the seminiferous tubules and the interepithelial space along the efferent ducts. The inter-Sertoli barrier has parallel strands ofjunctional particles associated more with the E-face, while in the efferent ducts from the rete testis to the ductus deferens the particle strands are

MALE GERM CELL PROTECTIVE BARRIER

153

anastomosing and appear more on the P-face. The highest number of junctional strands was found between the epididymal cells, and the lowest number, between the cells of the ductuli efferentes. The number of junctional strands between the rete testis cells was near that of the interepididymal space. The results with lanthanum perfusion suggested that the barrier of the ductuli efferentes is weak. Freeze-fracture studies by Greenberg and Forssmann (1983) revealed a continuous decrease in the number of tight junctional strands from the proximal toward the distal region of the guinea pig epididymal duct. Junctional anastomosing strands appear on the P-face, and intermingled with them are desmosomal figures. Ultrastructural, cytochemical, and immunocytochemical studies of the normal efferent ducts in mammals were done by several authors: Ramos and Dym (1977) in monkeys; Hermo and Morales (1984), Morales et al. (1984), and Pelliniemi et al. (1981) in rats; Flickinger et al. (1978) in the hamster; Yokoyama and Chang (1971) in the Chinese hamster; Jones et al. (1979) in the rabbit; and Greenberg and Forssmann (1983) in the guinea pig. The efferent ducts in experimental conditions were studied by Hoffer (1982) in the rat and by Goyal and Hrudka (1980,1981) in the bull. Efferent ducts are lined by ciliated and nonciliated (i.e., principal) cells. Endocytotic activity of the principal cells has been shown by Yokoyama and Chang (1971), Ramos and Dym (1977), Flickinger etal. (1978), Jones et al. (1979), Goyal and Hrudka (1980, 1981), and Pelliniemi et al. (1981). According to Hermo and Morales (1984) and Morales et al. (1984), the principal cells in the rete testis and the ductuli efferentes of the rat are capable of both fluid-phase and adsorptive endocytosis. Markers of adsorptive endocytosis-cationic ferritin and concanavalin A-ferritin-and markers of fluid-phase endocytosis-HRP and albumin bound to colloidal goldwere internalized and directed toward the lysosomal apparatus of the cell. The fate of the cells after having been filled by lysosomes is not known. It can be presumed that they become apoptotic and are replaced by new cells. In rat epididymis in uiuo, Na+ absorption and K+ secretion are inhibited by castration and are restored by testosterone replacement therapy (Wong and Yeung, 1978). The absorption of fluids and materials from the lumen along the efferent ducts and the epididymis was stressed by Flickinger et al. (1978) and by Greenberg and Forssmann (1983). The presence of microvilli in the ductuli efferentes is indicative of resorptive processes (Tormey, 1966). Tightjunctions situated at the luminal side of the epithelial cells suggest that the molecular flow which these junctions are barring is orientated in an apicobasal direction (Diamond and Bossert, 1967; Brightman and Reese, 1969; Bennett and Trinkaus, 1970). In the Sertoli epithelium, occludingjunctions are at the basolateral site, a possible exception being duck Sertoli cells

154

MORDECHAI ABRAHAM

(Pelletier, 1990), which have apically situated junctions. The absorptive activity probably determines a slow continuous flow of the fluid contained in the lumen extending along the whole efferent system, from the seminiferous tubules to the urethra. The flow of the luminal fluid starts in the seminiferous tubules of the testes (Fawcett, 1975). Such movement of the lumen content toward the ductus deferens possibly contributes to the effectiveness of the barrier in the testis itself. de Jong-Brink et al. (1984) noted that the high osmolarity of the gonadal fluid in the snail L. stagnalis attracts water that induces a high transmural pressure and a water current toward the hermaphroditic duct (Section 111, I). The importance of the epididymal environment for sperm survival was stressed by Whalen and Luttge (1969), Rastogi (1979), and Rastogi et al. (1979), who showed that reversible sterility can be produced by the use of the androgen antagonist cyproterone acetate. The transepithelial transport of 86Rb+,a tracer for K + , was examined in uitro in the sperm duct of the brook trout (Saluelinusfontinalis) (Marshall and Bryson, 1988; Marshall et al., 1989). Concomitantly, Na+ absorption was measured as the short-circuit current. Cells of the sperm duct actively secrete Rb' ions toward the lumen, after stimulation with dibutyrylCAMP,and take up Na+. Purified chum salmon carbohydrate-rich gonadotropic hormone added to either the apical or basolateral side of the epithelial cells produced a rapid sustained rise in both secretion and uptake. The stimulated Rb' transport was reduced by bilateral replacement of C1- with gluconate, indicating that Rb+ secretion is dependent on C1-. Injection of the antiandrogen cyproterone acetate (in uiuo) significantly reduced Rb+ secretion (in uitro), suggesting that androgens may maintain the active transport characteristics of the BTB. The active K + secretion by the sperm duct accounts for the high concentration of K + in seminal plasma, which is important in maintaining the spermatozoa in a nonmotile situation. Marshall and colleagues did not present information concerning the ultrastructure of the sperm duct epithelium in S. fontinalis. The ultrastructure and cytochemistry of the vasa efferentes and the vas deferens of the teleostean male gonad were studied by van den Hurk and Barends (1974), van den Hurk et al. (1974a,b, 1978), and van den Hurk and Testerink (1975) in the black molly (Poecilia latipinnu) and the rainbow trout (S. gairdneri). The testicular efferent duct system of the black molly consists of short efferent ducts with stratified columnar epithelium, connecting the testis tubules with a wide longitudinal main duct of simple cuboidal epithelium in each half of the testis. The cell apices in the main duct have numerous long microvilli and zonulae occludentes between neighboring cells, while the lateral cell membranes have a whorled configuration. Zonulae occludentes can also be found between the cells of the

MALE GERM CELL PROTECTIVE BARRIER

155

efferent ducts. The presence of malate and lactate dehydrogenases and NADHD activities in the epithelium of the trout vas deferens suggests that this epithelium is involved in regulating the composition of the seminal fluid, while the presence of acid phosphatase points to a phagocytotic function. Periodic acid-Schiff staining demonstrates a strong mucoprotein content in the seminal fluid, creating a suitable physiological environment presumably effective in the immobilization of the sperm cells (van den Hurk and Barends, 1974). The immobilization of spermatozoa by mucopolysaccharides was also suggested by Mann (1967) in the spermatophoric plasma of cephalopods. According to Bergmann et al. (1983), in the efferent ductules of R. esculenta there are no tight junctions. O’Donovan (1988) found the vas deferens in R. ridibunda permeable to HRP, in spite of the tortuous interdigitations of the epithelial cells. Convoluted intercellular spaces and extracellular material between the cells of an epithelium can reduce the movement of substances and participate in the formation of blood-organ barriers (Staehelin, 1974; Lane et al., 1977; Szollosi and Marcaillou, 1977). Nemetallah et al. (1985) found the blood-epididymal barrier in guinea pigs to be more restrictive to molecular transfer than the BTB. Turner et al. (1980) came to a similar conclusion. Wong and Uchendu (1990) found that captopril, an inhibitor of the angiotensin-converting enzyme, was prevented from entering the epididymal lumen of rats by a blood -epididymis barrier. According to Barratt et al. (1989), lead acetate administered to male rats by gavage affected the spermatozoa in the epididymis, while in the testis the germ cells were protected by the BTB. Permeability of septate junctions from excurrent duct epithelia to HRP was reported by O’Donovan (1988) in the hermaphrodite duct of the snail L. hierosolyma.

E. TESTICULAR VASCULAR SYSTEM While the Sertoli cell constituted the main interest in a large number of investigations concerned with the BTB, its opposite pole, the capillary, was less studied. Sakiyama et al. (1988) considered the testis capillaries similar to those of the liver (Goresky, 1982), where plasma proteins are as diffusible through the capillary wall as are smaller molecules. In skeletal and cardiac muscles of mice, when HRP is injected systemically, it crosses the walls of the endothelium within minutes, either in micropinocytotic vesicles or through open slits between neighboring endothelial cells (Karnovsky, 1965, 1967). Kormano (1968) found that intravenously injected Trypan Blue into rats extravasated from the testis capillaries very slowly. After injection of CdC12, the extravasation was increased. An increase in capillary permeability in the testes and changes in the BTB

156

MORDECHAI ABRAHAM

after CdCl2 treatment were found by Waites and Setchell (1966; Setchell and Waites, 1970). Increased vascular permeability in the testes of guinea pigs after CdCl2 administration has been shown by Johnson (1969). Nemetallah and Ellis (1985) found that intratesticular injection of a histamine releaser, “48/80,” or of CdC12 resulted in intense capillary engorgement with blood and ablation of the BTB in rats and guinea pigs (Section V,E). O’Donovan-Lockard et al. (1990) injected CdCl2 into the dorsal blood space and intraperitoneally into Tilapia hybrids (Teleostei). They found general internal hemorrhage, but no effect on testicular capillaries or the Sertoli cell barrier, confirming the view (Chiquoine, 1964) that the cadmium effect on the testes can be observed only in animals with scrota1 testes. Evidence was produced (Cordon-Cardo et al., 1989; Hank et al., 1989; Gallatin et al., 1986; Jalkanen et al., 1987) that testicular endothelium is selective for the molecules released into the perivascular space (Section IV,F). The formation of the blood vessels during mammalian ontogenesis was studied by Mayerhofer and Bartke (1989). In the testes of newborn mammals, blood vessels are poorly developed and highly permeable to dyes. Soon, angiogenic processes appear, and endothelial cell migration starts. Parallel to the formation of the Sertoli cell barrier, mature testicular capillaries are formed. However, blood and lymph vessels are not the only components of the circulatory system. The basement membranes, interposed between the myoid and the Sertoli cell layers which surround the seminiferoustubules, are part of a channel system connected to the pericapillary spaces. The extravascular spaces consist of two basal laminae of the apposing cells and a deposit of collagen fibers which may serve as a resilient filling preventing collapse of the spaces (Abraham et al., 1979b, 1982). Abrah-amet al. (1980) referred to the intercellular space between the myoid and Sertoli cells as the “connective tissue space,” 100-200 nm wide, which sends numerous indentations into the Sertoli cells (Figs. 18, 19, and 21). The connective tissue or collagen spaces surrounding the seminiferous tubules form “peg protrusions” (Vilar, 1973), noted by several authors (Burgos et al., 1970; de Kretser er al., 1975). Peg protrusions penetrate deep into the Sertoli cells, sometimes extending as far as threequarters of the width of the cells, stopping short of the adluminal border of the cell. In cases of human hydrocele, Chakraborty et al. (1976) found branching protrusions of the basement membrane that extended 10 pm or deeper into the Sertoli cells. The functional significance of these protrusions seems to be in conveying metabolites to the Sertoli cells on which germ cells are dependent. Kuopio and Pelliniemi (1989) described ultrastructurally identifiable continuous basement membranes found around seminiferous tubules and the interstitial capillaries. Small patches of base-

MALE GERM CELL PROTECTIVE BARRIER

157

ment membranes were, additionally, found on the free surfaces of Leydig cells, between two Leydig cells, and in macrophage-Leydig cell contact sites. These findings were confirmed by immunocytochemicallocalization of laminin and collagen type IV in the same area. Possibly, there is a basement membrane-mediated interaction of Leydig cells with other testicular structures. In fetal and newborn rats, the basement membranes cover large surfaces of the fetal-type Leydig cells (Kuopio et al., 1989).

F. SITESOF DIFFERENT MGPBS IN THE TESTES The original concept of the BTB refers to an obstacle interposed between capillaries and seminiferous tubules which hinders the free passage of molecules between them (reviewed by Setchell and Waites, 1975). Kormano (1968) studied the distribution of intravenously injected Trypan Blue with fluorescence microscopy in the rat and found a low extravasation of the stain from the testis vessels and a much higher outflow from the capillaries of the caput epididymidis. The capillaries of the testes have, therefore, a barrier for this stain. It was soon established that the key elements of the barrier are not the capillaries, but the intercellular spaces of Sertoli cells, where an impressive battery of tight junctional ridges stops the free transit of certain molecules (Connell, 1978; Nagano and Suzuki, 1978). Dym and Fawcett (1970) and Fawcett et al. (1970) proposed that the peritubular layer of myoid cells which lay external to the Sertoli cells form the first screen of the partition which separates blood-borne molecules from the germ cells. Junctional complexes between adjacent myoid cells stop the free flow of molecules of large size; this is the semipermeablelayer of Posinovec (1989). The Sertoli cell occludingjunctions form the second screen. The term “BTB” was formerly considered incorrect (Section II,B), since the site of the barrier was not the capillary system, as in the brain. Several publications in recent years cause us to reconsider this statement. In the carp, Parmentier et al. (1985) found that autologous immunoglobulins (IgM) of 700,000 Da are restricted to blood capillaries. Harik et al. (1989) have shown with ligand binding that brain microvessels have an extremely high density of the glucose transporter. These authors emphasize the brain’s isolation behind the BBB, which is impermeable to watersoluble compounds. Endothelial cells, when tested with immunocytochemical methods using monoclonal antibodies to human erythrocyte glucose transporter, showed a positive reaction only in vessels from peripheral nerves, retina, brain, and testes. No other endothelium reacted positively, suggesting that endothelial cells from capillaries of nervous tissue and testes have blood-tissue barrier characteristics. It was also

158

MORDECHAI ABRAHAM

found by Cordon-Cardo et al. (1989) that endothelial cells from the capillaries of the testes and the central nervous system contain a cell membrane glycoprotein which binds cytotoxic nonpolar molecules (Section V,E). Similarity between brain and testis structure was found by Campbell et al. (199O), who identified distinct isoenzymes of glutathione S-transferase (GST) in human brain and testes not found elsewhere in the organism. The authors supposed that these GSTs may be involved in blood barrier functions common to both organs. The endothelium of the testis capillaries as an additional site of the BTB also appears from the studies by Gallatin et al. (1986) and by Jalkanen et al. (1987). According to these authors, it may be that the first barrier against immunological aggression in the testis is at the level of the capillary endothelium that controls extravasation of lymphocytes. In order to enter the testicular interstitium, lymphocytes may be required to express specific ligands that would allow them to adhere to receptors on endothelial cells (Section V,C). Cytochemical studies (Gravis et al., 1977; Gravis, 1978) have shown that the peritubular basal lamina, interposed between the myoid and Sertoli cells, is rich in alkaline phosphatase, secreted by the myoid cells. It was postulated by Gravis et al. (1977) that the alkaline phosphatase-rich sheath that completely surrounds the seminiferous tubules constitutes a biochemical BTB through which substances passing from the blood to the germinal epithelium must penetrate and traverse. Rothwell and Tingari (1973) examined the barrier role of the boundary tissue of the seminiferous tubule in the testis of the domestic fowl. The connective tissue space (Abraham et al., .1980) forms in teleosts a selective filter (Lou and Takahashi, 1989) allowing the passage of cationic, but not anionic, proteins (Hadley and Dym, 1987). In the interstitial region there are additional barriers. Pollanen et al. (1988) describe a high-molecular-mass (130,000 Da) immunosuppressive protein factor in the testicular lymph vessels of the rat which inhibits lymphocyte proliferation. The seminiferous tubules in the rat testis are surrounded by lymph sinusoids (Fawcett et al., 19731, which therefore form an effective immunological barrier around the tubules. Besides this immunosuppressive protein (Pollanen et al., 1988), it is probable that an additional barrier exists against immunological aggression: Both Leydig cells and gonocytes were proposed as inhibitors of lymph cell proliferation in uitro (Section V,C). In conclusion, the list of components of the MGPB in the mammalian testis contains various endothelial factors, possibly the Leydig cells, the lymph sinusoids with the immunosuppressive protein, the myoid cell barrier, the alkaline phosphatase-rich connective tissue space, and the Sertoli-Sertoli occluding celljunctions. In no other animal group was such an impressive number of testicular barriers described. In the insects,

MALE GERM CELL PROTECTIVE BARRIER

159

however, a second barrier was mentioned by Schweich and Leloup (1988). The same is true in L. hierosolyma (O’Donovan and Abraham, 1987), in which it is presumed that the basal lamina space forms a primary barrier (Fig. 14), whereas the septatejunctions between the Sertoli cells constitute a subsequent one (Fig. 16). In the freshwater snail L. stagnalis (de JongBrink et al., 1984), only large colloidal gold particles do not pass the lamina. In teleost testis, too, the basal lamina space and the inter-Sertoli junctions form the two barriers (Lou and Takahashi, 1989). The efficiency of the teleost myoid cells as an additional barrier has not yet been confirmed.

V. Function of the Male Germ Cell Protective Barrier

A. SECRETORY ACTIVITY OF THE SERTOLI CELL Different periods in the lifetime of mammalian Sertoli cells are distinctively marked by the differences in their secretory activity. In the pig, during fetal (Tran et al., 1977, 1981; Blanchard and Josso, 1974) and perinatal (Price, 1979) life, Sertoli cells secrete a glycoprotein,,the antimullerian hormone (Picard et al., 1978). This activity declines after birth (Picon, 1969, 1970). Sertoli cell maturation involves acquisition of the characteristic shape and nuclear features of the adult cell, formation of occluding junctions, and blockage of mitotic activity, as well as initiation of the secretion of ABP (Hansson et al., 1973), shown to be correlated with formation of the BTB (Tindall et al., 1975; Tran et al., 1981). Many of the macromolecules required for somatic cell maintenance and development which are present in serum are not available to the developing germinal cells within the tubules. The protein content of the tubules, required for germinal cell viability, is determined largely by the secretion of Sertoli cells (Waites and Neaves, 1977). Sertoli cells supply many of the macromolecules for the adluminal compartment that hepatocytes provide for the systemic circulation (Cheng et al., 1990). Among the molecules secreted by mammalian Sertoli cells (Tindall et al., 1975, 1985) are ABP and y-glutamyl transpeptidase, a membrane-bound enzyme which mediates the translocation of amino acids across the plasma membrane. Skinner and Griswold (1980) have demonstrated that Sertoli cells also secrete transferrin, a 70,000- to 80,000-Da iron transport glycoprotein. The secretion of testicular transfenin was shown to be regulated by FSH, insulin, testosterone, and vitamin A (Skinner and Griswold, 1982). In addition to this, Jenkins and Ellison (1989) also found that epidermal growth factor and retinol regulate transferrin secretion. Sertoli cells are

160

MORDECHAI ABRAHAM

targets for both FSH (Hansson et al., 1973; Lipshultz et al., 1982) and testosterone (Louis and Fritz, 1979). Inhibin, a glycoprotein hormone which selectively suppresses FSH synthesis and/or release and diminishes FSH response to exogenous luteinizing hormone-releasing hormone (Franchimont et al., 1975, 1979; Scott et al., 1980), is also secreted by Sertoli cells. Inhibin may control spermatogenesis as well as the estrus cycle. Miyamoto et a / . (1989) have shown that testes in immature bulls produce high levels of inhibin, which is released into the circulation. After the onset of puberty, the release of inhibin into the circulation decreases due to formation of the BTB. Inhibin activity in neonatal rat testes has also been reported (Au et al., 1986). A handy method for inhibin bioassay in uitro was developed by Scott et al. (1980). Sertoli cells also synthesize ceruloplasmin, a 130,000-Da protein similar immunologically to serum ceruloplasmin, which can function in copper transport and act as a ferroxidase (Skinner and Griswold, 1983). Other physiologically possibly important molecules secreted by mammalian Sertoli cells are the plasminogen activators (Lacroix et al., 1977), which facilitate the transient breakdown of the Sertoli occludingjunctions to permit the passage of spermatocytes. Plasminogen activators catalyze the conversion of plasminogen to the active protease plasmin. In the teleostean testis and probably also in the amphibian testis, continuous destruction and regeneration of the barrier are not necessary. Here, all gonocytes from a given cyst ripen synchronously (Abraham et al., 1980; Marcaillou and Szollosi, 1980;Parmentier et al., 1985; Lou and Takahashi, 19891, and there is no need for separation inside the cyst between germ cells that should, and those that should not, be barred from the circulatory system. Besides proteases, Sertoli cells also synthesize protease inhibitors. According to Hirschhauser and Baudner (1972), there is a trypsinlike enzyme in spermatozoa which is inhibited by a seminal plasma protease inhibitor. Cheng et al. (1990) demonstrated that Sertoli cells are the source of the nonspecific protease inhibitor az-macroglobulin, secreted into the adluminal compartment, where it could limit the damage of proteases released by both degenerating germ cells and Sertoli cells. The peptide inhibitor of acrosin (Setchell, 1980) is also secreted by Sertoli cells.

OF THE BARRIER IN PHYSIOLOGICAL, PATHOLOGICAL, B. EFFICACY AND EXPERIMENTAL CONDITIONS

The exchange of substances between the fluid outside the seminiferous tubules and the fluid inside can vary over a wide range, from almost instantaneous entry (e.g., tritiated water and ethanol) to almost complete exclusion (e.g., albumin and inulin) (Setchell and Waites, 1975; Setchell

MALE GERM CELL PROTECTIVE BARRIER

161

and Main, 1978). The possibility of a slower entry of tritiated water through the BTB than into the general blood plasma was considered by Tanaka and Ujeno (1989). Salonen and Eriksson (1989) studied the pharmacokinetics of ethanol in the rat testis and various efferent ducts after intraperitoneal injections of this substance. In orbital capillary blood, the highest ethanol concentrations were measured 10 minutes after the injection; in the seminiferous tubules, after 20 minutes; and in the rete testis, after 30-60 minutes. Results indicate a barrier between blood and rete testis. Anderson et al. (1985) examined the levels of acetaldehyde in mice testes subsequent to acute ethanol administration and found a lower amount of aldehyde in testes than in blood, suggesting the existence of a BTB to acetaldehyde. High aldehyde dehydrogenase activity was present in mitochondrial and cytosolic fractions of the testes. Biomembranes are readily permeable to acetaldehyde. The molecule is oxidized after having passed the Sertoli cell barrier. Freeman et al. (1989) found that there is a BTB for casodex, a nonsteroidal antiandrogen, since the tissue : serum ratio was less than unity 1 hour after injecting the drug. The rate of entry into the tubules does not dependjust on molecular size; substances with high lipid solubility enter more rapidly than hydrophilic compounds. The transfer rate of drugs across the BTB is in positive correlation with their lipophilicity and their molecular size (Ouyang and Lien, 1984). Glucose enters the tubules by a process of facilitated diffusion (Middleton, 1973) by a carrier mechanism operating in the Sertoli cells. Testosterone enters relatively quickly, but the closely related compound 5a-dihydrotestosterone enters much more slowly (Cooper and Waites, 1975), although it is slightly more lipid soluble. According to Setchell and Main (1975), only free steroids enter the tubules. Turner et al. (1981a) have found that circulating androgens have limited direct access to the intralumind compartment of the male hamster reproductive tract. Pardridge (1988) compared the “free steroid hormone hypothesis” to the “bound steroid hormone hypothesis.” According to Vermeulen (1977), gonadal steroids are bound by albumin and sex hormone-binding globulin. The possibility that such steroid-protein complexes traverse the BTB conjointly was stressed by Bordin and Petra (1980) and by Siiteri et al. (1982). Studies performed by Pardridge (1988) and by Sakiyama et al. (1988) presented evidence that protein-bound sex steroids are taken up by Sertoli cells and are transcytosed into the seminiferous tubules, presumably by a receptor-mediated endocytosis mechanism. Attempts to test the efficiency of the MGPB under experimental conditions are numerous. Pelletier and Friend (1986) examined thin sections and freeze-fracture replicas of testes in prepubertal and adult guinea pig specimens force-fed gossypol acetate, a male antifertility agent which induces azoospermia (Shepu et al., 1980), causes structural changes in the testes of

162

MORDECHAI ABRAHAM

mammals (Wang et al., 1988),and diffuses through the BTB only at a high concentration gradient (Wang et al., 1989). Freeze-fracture replicas of continuous zonules at the bases of Sertolicells from adult gossypol-treated animals showed seemingly intact uninterrupted junctional strands which excluded filipin introduced by vascular perfusion. In prepubertal treated animals, no continuous zonules were formed, and the junctional complexes were entirely permeated by filipin (Pelletier and Friend, 1986) (Fig. 26). Sertoli cells were atypically structured and smaller than in controls, with large vacuoles in the cytoplasm (Hoffer, 1982, 1983) and a significant decrease in the secretion of ABP (Zhuang et al., 1983). Spermatogenesis was arrested at the pachytene spermatocyte stage. In human males exposed to 1,2-dibromo-3-chloropropane(DBCP), another male antifertility drug, seminiferous tubules devoid of germ cells can be found (Biava et al., 1978; Potashnik et al., 1979; Shemi et al., 1982, 1988, 1989).However, the Sertoli cells and their junctions appear to be unaffected. DBCP was also used in fish as a spermatogenesis-suppressingagent (O’Donovan-Lockard et al., 1990). Sullivan et al. (1979) studied the effect on rat testis of fluoroacetate in the drinking water and found degenerative changes of the germ cells, which they attributed to an impairment of Sertoli cell function. Harada et al. (1987, 1988) have shown that l-methy1-5-thiotetrazole, known as a side-chain molecule of cephalosporin antibiotics when injected subcutaneously into juvenile rats, inhibits formation of the BTB, blocks spermatid formation, and induces both reduction in the number of young germ cells and decreases of transferrin and ABP in the testicular cytosol. The treatment also causes apparition of IgG in the seminiferous tubules. The loosening of the Sertoli cell barrier, penetration of lanthanum, and histopathological changes in the tubuli 24 hours after oral administration of 1,3-dinitrobenzene to rats were shown by Shinoda et al. (1989). The effect of a vitamin A-deficient diet on spermatogenesis was investigated by Huang and Hembree (1979), Huang and Marshall (1983), and Huang et al. (1983, 1988). During chronic vitamin A deficiency (i.e., over 100 days), both serum and testicular retinol were reduced to a minimum, and there was severe germ cell loss due to disruption of the Sertoli cell barrier, documented by the penetration of lanthanum into the adluminal compartment. Mehan et al. (1989) noted degenerative changes in germ cells and alterations in Sertoli cell cytology after 5 weeks of a vitamin A-deficient diet. The probably important role of retinol in normal testicular function was demonstrated by Morales and Griswold (1987), who were able to restore spermatogenesis by injecting retinol acetate into rats with disturbed testicular activity due to retinol deficiency. The effect of microwave irradiation on the germinal cells and on the BTB was examined by

MALE GERM CELL PROTECTIVE BARRIER

163

Zhiyuan et al. (1985), who found that the Sertoli cell junctions remained unchanged after the irradiation. C. AUTOIMMUNE AGGRESSION IN MAMMALS AND THE MGPB The role of the occluding junctions between the Sertoli cells in mammals, as a bulwark barring autoimmune aggression, has often been stressed in the literature (Dym and Fawcett, 1970; Fawcett et al., 1970; Vitale et al., 1973; Castro and Seiguer, 1974; reviewed by Setchell and Waites, 1975). Late-appearing antigens of pachytene spermatocytes, spermatids, and spermatozoa, not recognized by the animal’s immune system, are isolated behind this protective barrier. Cell membrane-specific antigens which are absent in the spermatogonia, appear during spermatogenesis (O’Rand, 1980; O’Rand and Romrell, 1977; Tung and Fritz, 1978; Tung et al., 1979). The distribution of the antigenic sites on spermatozoa was studied by O’Rand (1980) and by Schmell et al. (1982). Tracer studies with lanthanum and HRP have demonstrated that the adluminal compartment of the tubules is not accessible to these tracers (Dym and Fawcett, 1970; Dym, 1973; Connell, 1978) and probably is also not permeable to immunoglobulins or lymphocytes, under normal circumstances. However, in animals immunized experimentally against their own mature germ cells, either by injections or by vasectomy (Castro and Seiguer, 1974), an inflammatory process appears-called allergic orchitis or autoimmune aspermatogenic orchitis-which raises tubular permeability. Antibodies to homologous spermatozoa were also observed in normal serum (Johnson, 1968). Both humoral antibodies and immune cells find their way into the seminiferous tubules (Brown et al., 1967; Brown and Glynn, 1969; Johnson, 1970b,1972; Jones et al., 1971), leading to desquamation of the germinal cells, while the inter-Sertoli junctions appear to be unharmed (Kierszenbaum and Mancini, 1973). The crossing of immunoglobulin into the seminiferous tubules is interpreted differently by various authors. The first barrier of the semipermeable myoid cell layer (Posinovec, 1989), acceding to the basal lamina, is apparently easily crossed (Castro and Seiguer, 1974). It has been shown by Mancini et al. (1966) and by Brown et al. (1967) that kininlike substances appear during allergic orchitis which might stimulate the contraction of the myoid cells, thus opening the intercellular passage to the humoral antibodies into the basal compartment. The damage to the spermatogonia and early spermatocytes which may result from this first crossing is presumably minimal. The Sertoli cell barrier, which defends the adluminal compartment, remains unchanged after the induction of allergic orchitis and prevents the entrance of intercellular markers such as HRP or lanthanum

164

MORDECHAI ABRAHAM

nitrate even into those seminiferous tubules in which the germ cells were severely damaged (Willson et al., 1973; Castro and Seiguer, 1974). In human males suffering from azoospermia, the Sertoli cell barrier appears to be intact. The whole population of early spermatids degenerates and is phagocytosed by Sertoli cells which have numerous phagolysosomes with a high content of sodium P-glycerophosphatase (Viehberger, 1985). Ross (1977) reported, after ligation of the efferent ductules in mice, that lanthanum failed to traverse the Sertoli-Sertoli junctions of the seminiferous tubules. In similar experiments, Neaves (1973) noted that the inter-Sertoli junctions became permeable. An intact Sertoli cell barrier in both cryptorchid and contralateral scrota1testis of rats was also reported by Stewart et al. (1990), after 150 days of experimental unilateral cryptorchidism. However, the germinal epithelium of the cryptorchid testis was degenerate. Russell and Peterson (1985) presented in table form many pathological and experimental situations in which spermatogenesis was severely disrupted, while the Sertoli occludingjunctions remained unchanged. Pathological changes in germ cells with apparently intact Sertoli junctions were also shown in studies dealing with varicocele, an abnormal dilatation of the spermatic vein in human males, or experimentally induced in rats and dogs, complicated with infertility, abnormal testicular histology, and changes in sperm motility (MacLeod, 1965; Howards et al., 1976; McFadden and Mehan, 1978; Cameron et al., 1980; Saypol et al., 1981; Turner et al., 1982, 1987; Rajfer et al., 1987). In human males with varicocele, the Sertoli cell barrier appears intact in the electron microscope (Cameron et al., 1980). Turner et al. (1981b) analyzed isotope concentration of [3H]inulin in carotid artery blood, seminiferous tubule fluid, and cauda epididymal fluid after having infused labeled inulin intravenously. Experimental unilateral varicocele did not alter the intratubularconcentrations of inulin either in the testis or in the epididymis; however, it altered intratesticular testosterone concentrations (Rajfer et al., 1987). In germinal aplasia, in infertile males, lanthanum tracer penetrated deeper into the inter-Sertoli spaces than in controls, although in transmission electron microscopy and in freeze-etching replicas no structural modifications were observed (Camatini et al., 1981). According to Johnson (1970a,b) and Willson et al. (1973), the efficiency of the Sertolijunctions diminishes in allergic orchitis, permitting access of the immunoglobulins to the adluminal compartment. Kierszenbaum and Mancini (1973) suggested that the passage of antibodies into the tubular lumen is via the Sertoli cells themselves. Johnson (1972), who observed the presence of fluorescein-conjugatedalbumin or globulin in 50% of the testicular tubules 9 hours after intravenous injection of the tracers, had an alternative explanation, according to which macromoleculespenetrate the

MALE GERM CELL PROTECTIVE BARRIER

165

rete testis and then secondarily reflux into the seminiferous tubules. The rete testis has been found to be permeable to immunoglobulins by other authors as well (Tung et al., 1971; Koskimies et al., 1971), even though tight junctional complexes exist between epithelial cells in this region (Dym, 1976; Nagano and Suzuki, 1983). Suzuki and Nagano (1978) considered the ductuli efferentes, not the rete, as the most permeable section in the duct system. High serum titer of antisperm immunoglobulins, acting as a sperm-immobilizingand -agglutinating or antifertilizingfactor (Haas et al., 1980; Huang et al., 1981; Menge and Peegel, 1980; Brannen-Brock and Hall, 1985), diminishes the fertility of spermatozoain different mammalian species, including humans. The path of penetration of these immunoglobulins into the compartments containing spermatozoa is not known. Slavis et al. (1990) proposed a common immune etiology for infertility in two experimentally traumatized and immunized groups of rats, through possible disruption of the BTB. Wentworth and Mellen (1964) reported that experimentally induced sperm antibodies can enter the male reproductive tract in the Japanese quail. Induced autoimmune sterilization of the testes in freshwater and seawater species of teleosts was proposed by Laird et al. (1978,1980) Secombes etal. (1982,1985,1986), Lou andTakahashi (1987), Timmermans (1987), O’Donovan (1988), and O’Donovan-Lockard et al. (1990).

Studies by Yule et al. (1987a,b, 1988), although not completely negating the immunoprotective role of Sertoli cells, had a more cautious approach to the immunofunction of the barrier. Yule et al. (1988) showed that germ cells located in the basal compartment, outside the Sertoli cell barrier, are also autoantigenic and accessible to circulating activated T cells. In mice immunized with syngeneic testis homogenate, they found deposits of IgG in the basal portion (the open compartment) of 30-40% of seminiferous tubules and developed experimental allergic orchitis. IgG deposits are preferentially formed in seminiferous tubules at stages 7- 12 of the spermatogenetic cycle. The initial targets in experimental allergic orchitis induction are autoantigens on germ cells (preleptotene spermatocytes) outside the BTB. IgG bound to preleptotene spermatocytesmay be carried passively into the adluminal compartment of the seminiferous tubules as these cells differentiate and move behind the barrier. This IgG is absorbed from the circulation by the testis and therefore is found only in the serum of mice orchiectomized before immunization (Mahi-Brown et al., 1988). The presence of immunogenic autoantigensoutside the BTB contradicts earlier views which emphasized that antisera to isologous or autologous testes, produced in the rabbit or the rat, react exclusively with antigens of germ cells located behind the barrier (O’Rand and Romrell, 1977; Tung and Fritz, 1978). A testis-specific autoantigenic isoenzyme lactate dehy-

166

MORDECHAI ABRAHAM

drogenase was detectable only in pachytene spermatocytes and haploid cells (Hintz and Goldberg, 1977). A sperm surface fertilization antigen was isolated from murine testes by Naz and Bhargava (1990). The appearance of cell membrane-specific antigens during spermatogenesis was also studied in teleosts, a vertebrate group with a welldefined Sertoli cell barrier (Marcaillou and Szollosi, 1980; Abraham et al., 1980). In bony fish, germ cell antigens were demonstrated by Muller and Wolf (1979) and by Pechan et al. (1979). Parmentier et al. (1984) and Parmentier and Timmermans (1985) prepared hybridoma clones of IgG and IgM monoclonal antibodies against carp sperm. The antigens were distributed over the sperm heads and midpieces, but were absent from the tails. Immunofluorescence was also observed on the surface membranes of spermatogonia, spermatocytes, and spermatids, but not in somatic cells. Some of the antibodies reacted with surface membranes of oogonia and early prophase oocytes. The immunofluorescent reaction also occurred with male and female germ cells of another cyprinid (Barbus conchonius). Similar results were published by Timmermans and Taverne (1983). Immune reactivity of carp primordial germ cells to monoclonal antibodies against spermatozoa has been shown by Timmermans et al. (1985). Evidence for local membrane differentiation on sperm heads was obtained by Davies et al. (1983) by studying lectin binding on the heads of ripe spermatozoa of Xiphophorus helleri. Normal seminiferous cysts of teleosts contain no immunoglobulin. However, following autoimmune response to spermatozoa, autologous immunoglobulins have been detected inside the cysts in S . gairdneri (Secombes et al., 1987), while Lou and Takahashi (1989) found IgM in the milt of Tilapia specimens 17 days after injection of allogeneic spermatozoa. The latter authors also found that after autoimmunization, both the basement membrane barrier and the inter-Sertoli cell barrier become deficient, and the markers bovine serum albumin and HRP break in. The breakdown of the barriers was evident 1 month after immunization, prior to the appearance of immunopathological changes in the testis (Lou and Takahashi, 1988). Immune cells invaded the barrier sites 45 days after immunization. Leukocytes sensitized by germ cell antigen and adjuvant can activate macrophages to phagocytize spermatozoa (Secombes, 1986). In the testes of mammals, the afferent and the efferent limbs of the immune system do not function with the same efficiency (Lewis-Jones et al., 1987). Once effector cells are activated, the protective barriers of the testis are no longer sufficient. T cells from mice immunized with testis can be transferred to naive syngeneic mice, in which they infiltrate the testis and cause orchitis (Tung, 1978). Protection operates only on the afferent limb of the immune response. The ease in activation of the efferent arm of

MALE GERM CELL PROTECTIVE BARRIER

167

the immune system appears also in cases of accidentally or experimentally induced ischemia in one of the testes of humans, rats, or rabbits, which results in histological damage to the contralateral testis, called sympathetic orchiopathia (Wallace et al., 1982; Nagler and White, 1982; Cerasaro et al., 1984; York and Drago, 1985). Sympathetic orchiopathia of the contralateral testis can be induced by vasectomy or by torsion or ligation of the ipsolateral testicular vessels. Vasoconstriction and ischemia as a result of intraperitoneally injected epinephrine (Gravis et al., 1977) caused testicular degeneration in the adult Syrian hamster, but the Sertoli-Sertoli junctions remained intact. Lewis-Jones et al. (1987) have shown with immunocytochemical methods that in rats with ipsolateral ischemic testis, 7 days after induced ischemia, the contralateral testis is penetrated by IgM antibodies which break the BTB, and 3 weeks later by IgG antibodies. These authors suggested that ischemic necrosis in the ipsolateral testis adversely affects immunoregulatory cell function and thus stimulates autoantibody production. This study was done with a light microscope, and the published illustrations do not permit a good interpretation of what happens with the Sertoli cells and in the inter-Sertoli space. The effect of vasoligation or vasectomy on spermagglutinin formation was also studied by Rumke and Titus (1970). Induced activity of the efferent immune limb was also demonstrated by Pelletier et al. (1981), after immunizing guinea pigs with isologous spermatozoa combined with Freund’s complete adjuvant and pertussis vaccine. They found distended gaps between adjacent Sertoli cells, followed by a massive destruction of the germinal cells, as a possible result of immunoglobulins penetrating into the adluminal compartment. Adjuvant-induced penetration of HRP into the seminiferous epithelium of guinea pigs was also studied by Willson et al. (1973). A possible interpretation of these experiments is that the adjuvant sensitizes the afferent limb of the immune reaction. The studies by Pollanen et al. (1988) and by Yule et al. (1988) emphasized that the testis of rats and probably of mammals in general is an immunologically privileged site which harbors few lymphocytes. This is in spite of large amounts of interleukin 1-like activity secreted by the Sertoli cells which has the potential to promote lymphocyte growth (Khan et al., 1987). Lymphocytes are confined to regions of the tubuli recti and the rete testis (Dym and Romrell, 1975). This is probably the reason that foreign tissue and organ grafts can survive in testicular tissue (Head et al., 1983; Maddocks et al., 1984; Head and Billingham, 1985; Pollanen and Maddocks, 1988). According to El-Demiry e? al. (1985), T suppressor or cytotoxic lymphocytes are found in the normal human testis and epididymis. If the host is sensitized to the graft by previously transplanting tissue from the same donor to the skin, a rapid cell-mediated rejection of the grafts

168

MORDECHAI ABRAHAM

occurs (Head et al., 1983). The rat testis contains large numbers of class I1 MHC antigen-positive macrophages, and it can be assumed that it is at this site that antigen presentation to T inducer lymphocytes occurs (Pollanen and Maddocks, 1988). Pollanen et al. (1988) described a high-molecularmass (130,000 Da) immunosuppressive protein factor in lymph from the rat testicular interstitial tissue with inhibits lymphocyte proliferation. Head and Billingham (1985) reported that immune privilege in the testis is significantly stimulated by steroid hormones. In several organ systems, the immune response does not conform to the general known pattern (e.g., Raju and Grogan, 1969; Franklin and Prendergast, 1970; Kaplan and Streilein, 1977a,b; Barker and Billingham, 1977; Streilein, 1987; Caspi et al., 1987; Billingham and Silvers, 1968). The brain, the eyes, the testes, and the hamster cheek pouch belong to those organs to which the somewhat pretentious title of immunological sanctuary, or the more humble title of immunologically privileged site, has been awarded (Barker and Billingham, 1977; Streilein, 1987; Caspi et al., 1987). It was thought that the antigens are secluded in these sanctuaries, or that there is a scarcity of lymphocytes, which reduces the activity along the afferent limb of the immune reaction and, as a consequence, the immune mechanism does not become aware of the antigens. One interesting possibility for a mechanism of local immune privilege is direct interference with antigen presentation and T cell activation by some cell type resident in the respective organ. The eye is exempt from certain immune reactions. In the case of experimental autoimmune uveoretinitis, retinal glial cells-the Muller cells which accompany the photoreceptor cells-act as a local suppression mechanism of immune reactions (Caspi et al., 1987; Streilein, 1987). The inhibition requires direct contact between the Muller cell and the T cell. In the testis, both Leydig cells and gonocytes were described as inhibitors of lymph cell proliferation in uitro (Hurtenbach et al., 1980; Born and Wekerle, 1982). Local immunosuppressive mechanisms which limit lymphocyte functions at the afferent arm of the antihaploid antigen’s immune response may be determinant in the prevention of testis autoimmune disease. In the mink, M.uison, a seasonal breeder, the BTB decays at the end of the breeding season, leaving the autoantigenic germ cells in the seminiferous tubules defenseless. It is conceivable that immunoreactants (i.e., sensitized lymphocytes) could have access to the autoimmunogens, thus inducing the formation of antisperm antibodies (Pelletier, 1986, 1988). However, infertility has been observed in only one of the strains (Tung et al., 1981, 1984). Presumably, immunosuppressivefactors prevent destruction of the seminiferous epithelium. It may be that the first barrier against immunological aggression in the testis is at the level of the capillary endothelium, which controls extrava-

MALE GERM CELL PROTECTIVE BARRIER

169

sation of lymphocytes (Gallatin et af., 1986; Jalkanen et af., 1987). There are several immunoregulatory mechanisms which probably operate on the afferent limb of the immune response to either prevent a response or direct it toward suppression (Mahi-Brown et al., 1988), such as T cell-mediated suppression, T cell activation by a resident testicular cell, localized general suppression, control or inhibition of antigen presentation, or control of lymphocyte extravasation and circulation in the testis. However, after formation of testis-specific activated T cells, these mechanisms are overcome, and orchitis occurs.

D. ROLEOF THE MGPB IN MAINTAINING SPERMATOZOAN IMMOTILITY Ripe spermatozoa of species from many phyla are known to survive for weeks while inside the testes, but immediately upon release, they begin to move and lose their fertility within a very short time. It might be presumed that a favorable microenvironment inhibits metabolism and motility of the spermatozoa while inside the testes. In many animal groups, sperm transferred during copulation survive in the female organism for many weeks and fertilize the female gamete a long time after mating. A long motionless survival after ripening also characterizes the sperm inside the spermatophores in many invertebrate species (Fretter and Graham, 1964; Adiyodi and Adiyodi, 1983). The literature mentions (Setchell and Waites, 1975) the role of the blood-spermatozoa barrier in preserving ripe spermatozoa from molecules that may trigger their motility and thus reduce their energy level. The effective functioning time of an extruded spermatozoon is very short (Nikolsky, 1963). The movement of ripe sea urchin spermatozoa is inhibited by some amino acids (Tyler and Rothschild, 1951). In mammals, testicular spermatozoa are not motile (Setchell et al.. 1969) although they apparently possess the means required for movement. In the course of their passage through the epididymis, the spermatozoa undergo morphological and physiological changes associated with the development of motility potential (Voglmayr, 1975; Nalbandov, 1976). Spermatozoa from the efferent ducts of rabbits or from the caput epididymidis were infertile, while those from the cauda epididymidis were fertile (Lambiase and Amann, 1973). The movement of spermatozoa while still inside the testis or the spermarium has been observed by Zihler (1972), O’Donovan and Abraham (1988a), and de Jong-Brink et af. (1984). However, this might happen a short time before spermiation. Fish spermatozoa are similar in structure to sea urchin sperm; their 9 + 2 axonemal structure is not surrounded by accessory fibers. They are, however, dissimilar in motility time once released into water. The flagellar beating of sea urchin sperm in sea water might last for hours. There is an

I70

MORDECHAI ABRAHAM

equilibrium between the synthesis of ATP by the mitochondria and its hydrolysis by the dynein ATPase (Christen et al., 1987). Trout sperm are motile for less than 30 seconds after spawning in fresh water, as well as in isotonic solution. Sperm motility was not initiated when the dilution medium contained an elevated concentration of potassium (20-40 mM), but dilution in an isotonic solution of NaCl triggered an immediate activation of motility and sperm swam vigorously. After 20 seconds, sperm became abruptly immotile. When millimolar amounts of Ca2+were also present in the dilution medium, flagella kept beating 1-2 minutes. The progressive decrease of the flagellar beat frequency been observed during the period of trout sperm movement might be related to the rapid exhaustion of intraflagellar ATP. In the rainbow trout, the release of suppression by potassium induces the increase of intracellular CAMP, which converts the immotile axoneme (9 + 2) to a motile one; CAMPis the internal factor to trigger sperm motility (Christen et al., 1987; Morisawa and Suzuki, 1980; Morisawa and Okuno, 1982). While in the seminiferous tubules, mammalian spermatozoa are bathed in a fluid with an ionic composition differing from that of blood plasma (Schlenk and Kahmann, 1938; Setchell and Waites, 1975). The tubular fluid has a much higher potassium content than the blood plasma, less Ca2+ and Mg2+, and practically no glucose or fructose (Setchell and Waites, 1975). According to Baynes et al. (1981) and Vlok and Van Vuren (1988), the in uiuo inhibition of sperm motility in teleosts can be attributed to potassium, which is present in high concentrations in the seminal plasma. This tendency to high potassium and low calcium and magnesium in the seminal plasma can be seen clearly in Clarias gariepinus and Oncorhynchus keta (Morisawa et al., 1979). Sperm motility in chum salmon is also inhibited by potassium in high concentration in the seminal plasma (Morisawa and Suzuki, 1980). A reduction of potassium concentration in the environment surrounding the spawned spermatozoa induces the initiation of sperm motility. The active K+ secretion by the sperm duct accounts for the high concentration of K+ in seminal plasma (Marshall et al., 1989) (Section 111,K).In rat epididymis, Na+ absorption and K+ secretion are inhibited by castration and are restored by testosterone replacement therapy (Wong and Yeung, 1978). The factors inhibiting motility are K+ in salmonids (10-40 mM), and osmotic pressure in cyprinids (200 mosm). The pH of the external medium also plays a determinant role in the motility of trout spermatozoa. At pH 7.0 or less, the spermatozoaare immotile, and at pH 9.0, motility intensity is maximal. There is also an effect of temperature. The beating frequency of the flagella is low at 5°C and increases rapidly with temperature rise. The duration of spermatozoan mobility changes in the different phases of the reproductive season. Benau and

MALE GERM CELL PROTECTIVE BARRIER

171

Terner (1980) found that salmonid spermatozoa are motile for 30-55 seconds at the peak of the spawning season, and only for 15 seconds at its end, caused presumably by the decrease in sperm CAMP concentration. The relationship between medium salinity and fertility of teleost spermatozoa was studied by Billard (1978). In a snail, L. stagnalis (de Jong-Brink ef al., 1984), gonadal fluid has higher osmolarity and a higher concentration of amino acids proteins, K+, and C1- than does the hemolymph. Storage of mature spermatozoa in uitro in physiological solution identical to the seminal fluid without having recourse to cryopreservative techniques was proposed by Abraham (1981). This would permit the use of spermatozoa for medical, aquacultural, or cattle-breeding purposes without having recourse to cryopreservative techniques. The influence of the seminal fluid and various saline diluents on the fertilizing ability of spermatozoa, as well as the techniques of cryoconservation of ripe sperm of different fish species, has been studied by Billard (1981,1986, 1987) and by Legendre and Billard (1980). Steyn et al. (1985) have successfully preserved C. gariepinus semen by cryopreservation at - 196°C. Legendre and Billard (1980) provided different technical hints to improve the cryopreservation procedures for the trout sperm. According to Chao et al. (1975) and Billard (1987), the fertilization rate of cryogenically preserved sperm is low. A short-term preservation of sperm under oxygen atmosphere in rainbow trout ( S . gairdneri) was proposed by Billard (1981). The role of oxygen in sperm preservation is still uncertain. In nature, in the insects (an animal group in which oxygen transport is separated from metabolite transport) the oxygen transport system of the tracheae does not penetrate into the barred region of haploid male germ cells (Section 111,H). It might be that a high K+ medium and anaerobic conditions are nearest to the natural conditions.

E. MEDICAL ASPECTSOF THE MGPB The tightness of the Sertoli cell barrier poses a problem in ailments affecting the testes, since the barrier is impenetrable to drugs. The opening of the barrier should make treatment possible. Osmotic opening of the BBB (Rapoport and Thompson, 1973; Pickard et al., 1977) was attempted with the help of hypertonic solutions in order to shrink the cells. The opening of the Sertoli cell barrier seems to be more difficult. Gilula el al. (1976) found the occluding junctions in the mammalian testes to be resistent to dissociation by hypertonic solution. Bergmann et al. (1984b), using hypertonic fixatives together with lanthanum, observed in amphibian gonads the opening of young cysts together with shrinkage artifacts, but postmeiotic cysts remained closed. Sertoli tight junctions were

172

MORDECHAI ABRAHAM

considered the “tightest” in the mammalian body (Claude and Goodenough, 1973; Fawcett, 1973), a distinction which is disputed today (discussed by Pelletier, 1990). Assaykeen and Thomas (1965) and Nemetallah et al. (1985) noted in the dark mink (M. vison) with spontaneous autoimmune allergic orchitis aspermatogenesis associated with elevated testicular histamine content. The effect of histamine on the barrier was tested by Nemetallah and Ellis (1985). They succeeded in ablating the BTB in rats and guinea pigs through intratesticular injection of a histamine releaser, 48/80, while the blood-epididymal barrier remained intact. An interesting physiological problem connected to the existence of the MGPB arises from the study of lymphocytic leukemia in adults and children. It has been proposed that relapse of acute lymphocytic leukemia in the testes or the meninges of adults may be the result of malignant cells surviving because of the failure of the drugs to penetrate through the Sertoli cell barrier or through the brain capillaries (Cordon-Cardo et al., 1989). Cordon-Cardo and co-workersalso found that endothelial cells from blood-tissue barrier sites such as the central nervous system, the testes, and the papillary dermis contain a cell membrane glycoprotein which is an operative component of the barrier and can bind cytotoxic nonpolar molecules. No other capillaries in the organism contain this cell surface protein which impedes drug penetration into these pharmacological “sanctuaries.” It appears, therefore, that both the testicular endothelium and the inter-Sertoli junctions inhibit the efficient treatment of leukemia. Childhood leukemia occurs before the Sertoli cell junctions are formed (Gondos and Sun, 1985); its treatment is probably less difficult. The topic of Sertoli cell junction and chemotherapy of childhood leukemia was also investigated by Camatini et al. (1982a). In the human male, spermatogoniaare normally in contact with the basal lamina. Dislocation of spermatogonia in human seminiferous tubules was studied by Holstein et al. (1984), Schulze (1988), and Bergmann et al. (1989) in elderly and/or infertile human males, in cases of carcinoma in situ, and after long-term estrogen treatment. Bergmann et al. (1989) considered the pathological dislocation of spermatogonia to be the result of Sertoli cell disfunction. Dislocation of spermatogonia has also been observed in the terminal segment of the human seminiferous epithelium, where Sertoli tight junctions were absent (Lindner, 1982). The correlation of ultrastructural changes in human Sertoli cells, germ cell disorders, and the level of serum FSH has been studied by Chemes et al. (1977) and by de Kretser et al. (1981). Setchell and Main (1978) discussed a yet unsubstantiated problem, that a nontoxic substance could penetrate the BTB and be transformed inside the barrier into a toxic metabolite which at the same time, due to size or

MALE GERM CELL PROTECTIVE BARRIER

173

structure becomes nonpermeant, thereby effectively concentrating the toxic compound inside the tubuli.

VI. Concluding Remarks The main element of the MGPB in the testes of metazoa is formed by occluding junctions between adjacent somatic cells which block the free passage of molecules and cells toward the germ cell region. Such a barrier was found in a variety of species of invertebrates and vertebrates from coelenterates to mammals: Hydra, a flatworm, a nematode, a centipede, insects, mollusks, teleosts, amphibians, reptiles, birds, and mammals. In annelids and crustaceans, the existence of a barrier has not yet been confirmed. In echinoderms, the lancelets, and hagfish, no barrier was found. In the testes of Chondrichthyes, the intensively secreting Sertoli cells form cysts which surround the male germ cells during spermatogenesis (Collenot and Damas, 1980; Callard et al., 1989; Dubois and Callard, 1990). However, the existence of an MGPB has not been experimentally tested. In the shark Heterodontus portusjacksoni, Jones et al. (1984) have demonstrated a barrier to the movement of Naf and K+between the blood and the epididymal lumen. No data are available for minor phyla. In several phyla, more than one barrier regulates the flow of metabolites between somatic and germ cells. In mammals, five, or possibly six, different barrier sites have been reported. The capillary endothelia, lymph sinusoids, peritubular basal laminae, myoid cells, Sertoli cells, and possibly the Leydig cells as well serve as barriers. They are endowed with mechanisms that determine which cells or metabolites can penetrate into the testicular interstitium and the seminiferous tubules (Section IV,F). The discovery that in the testicular capillaries, metabolites and lymphocytes do not extravasate freely (Gallatin et al., 1986; Jalkanen et al., 1987; CordonCardo et al., 1989; Harik et al., 1989) contradicts the previously accepted view (Setchell and Waites, 1975) that the capillaries in the vertebrate testes are open, and serum freely permeates the interstitial region. Due to the considerable number of animal groups in which an MGPB was reported, it is surprising that in three groups of deuterostomata-the echinoderms (Chia and Buckland-Nicks, 1987), the lancelets (Holland and Holland, 1989), and the hagfish (Alvestad-Graebner and Adam, 1977)-no MGPB was found (Section III,J and K). It would seem important to corroborate these results with the help of tracers and for all stages of spermatogenesis. If confirmed, the next task would be to determine which peculiarities of the male germ cells of these deuterostomata make the barrier redundant. The flagellar beating of sea urchin sperm may last for hours, while that of trout

174

MORDECHAI ABRAHAM

may last for less than 30 seconds (Christen et al., 1987).Are the absence of MGPB and the long duration of flagellar activity causally related? In female gonads, no germ cell protecting barrier has been observed in any species. Circulating metabolites in the interspace of somatic cells have free access to the female germ cells at all stages of oogenesis (Dumont and Wallace, 1978; Wallace and Selman, 1981; Abraham et al., 1984; Parmentier et al., 1985). Whereas in male gonads, the volume of the germ cell decreases during gametogenesis, the opposite is true for the female germ cell. Its volume increases during vitellogenesis, and a barrier between the circulatory system and the oocytes would be counterproductive. Proteins from outside the ovaries moving toward the oocytes are endocytosed by the oolemma. The difference between male and female gonads, regarding the barrier, can best be understood by comparing the relationship between germ cells and the circulatory system in hermaphrodite species. In H. uiridis, ovaries develop in the lower part of the body, where the mesoglea [a “primeval” circulatory system (Hausman, 1973)l is well developed and the penetration of exogenous HRP into it is highest and quickest (O’Donovan and Abraham, 1988a). In the spermarium region, the mesoglea is thinner than in other regions, and is always HRP free (Section 111,B). For D . biblica, only data for the male gonad were available (Section 111,C). In the hermaphrodite land snails (Section III,I), the female and male gonads are organized into two concentric belts. The ovaries form the external belt, in close contact with the circulatory system, and the male gonad forms the internal belt, which is barred from it. In the hermaphrodite teleost Sparus aurata, male and female gonads are separated by the zona trabeculata, which consists of dense masses of collagen fibers (M. Abraham and Y.Zohar, unpublished observations). Here, the ovaries are close to the central gonadal cavity. Occasionally, somatic cells with occludingjunctions in the intercellular spaces can be observed at the boundary between male and female gonads. Of the different MGPBs, the best studied are those formed between Sertoli cells (reviewed by Setchell and Waites, 1975; Tindall et al., 1985; Russell and Peterson, 1985; and others). Most cases examined revealed a similar structural pattern of the barrier: septate junctions in invertebrates and tight junctions in vertebrates (although in some cases vertebrate testes with septate-likejunctions were found, and invertebrate testes with tight junctions) (Section III,H and K and Section IV,C). Additional structural elements of the barrier were electron-dense material in the intercellular space, tortuous intercellular pathway, basement membrane with dense collagen space, and secretory cell features of the Sertoli cells surrounding the male germ cells. The secretory activity of the Sertoli cells determines the molecular content of the tubular fluid (Section V,A). The ontogenesis of the Sertoli cell barrier was thoroughly investigated in mammals (Section

MALE GERM CELL PROTECTIVE BARRIER

175

IV,B). Cyclic formation and disappearance of the barrier have been described in mammals and birds (Pelletier, 1986, 1988, 1990). This finding emphasizes that the protection of male germ cells becomes necessary only at certain stages of spermatogenesis. The phenology of the formation and dissipation of the occluding junctions proved to be related to the stages of spermatogenesis and may serve as a basis for the interpretation of the role of the barrier. Since germ cells in some mammals enter the closed compartment early in leptotene, a meiosis-promoting role of the barrier (Setchell and Waites, 1975), through storage of a meiosis-inducing substance (Parvinen, 1982), was postulated. In other amniotes, too, (Section III,K), and in insects (Section III,H), the closed compartments around male germ cells are formed early, in the meiotic prophase. By contrast, in several mammalian species, the closed compartment appears at the end of leptotene. The final rejection of the meiosis-promoting role of the barrier coincides with the study of the Sertoli barrier in anamniotes (Section III,K), in which the barrier forms at a late stage (i-e., when the germ cells are already haploid). Various hypotheses concerning the role of the blood-spermatozoa barrier have been discussed by Setchell and Waites (1975). One of the suggestions concerning the role of the Sertoli cell barrier centers on the protection of the newly formed haploid cells from immunogenic contact and autoallergic destruction. Several authors have observed that the barrier is efficient against the formation of antibodies along the afferent limb of the immune reaction, but if antibodies are introduced into the organism from the outside, the barrier becomes ineffective (Section V,C). The Sertoli cell barrier is not the only agent responsible for the prevention of immunogenic contact of the haploid cells. The presence of autoimmunogenic germ cells outside the Sertoli cell barrier (Tung, 1980; Yule er al., 1987a,b) indicates that additional immunosuppressive mechanisms must be active in preventing autoimmune disease of the testis. No data are available about the immunoregulatory activity of the barrier in invertebrates. The immunological role of the barrier may have arisen late in evolution. Coelenterates exhibit allogeneic incompatibility (Theodor, 1970), yet information available about immune reactions in these or higher invertebrates (Ratcliffe et al., 1985) fails to support the view that they possess physiological mechanisms which enable them to distinguish between the antigens of their own diploid and haploid cells. As for the female gonads, haploid germ cells-which are supposedly the main autoimmunogenic factor-do not appear in the ovary. Female germ cells do not become haploid until immediately before fertilization. In mammalian ovaries (Shulman, 1975), ovarian antigens are localized in the zona pellucida, the theca interna, and the atretic follicles. It is not known whether these antigens are autoimmunogenic. Yule et al. (1988) considered that autoim-

176

MORDECHAI ABRAHAM

mune defense is not one of the factors in the evolution of the BTB. The protective role of the Sertoli cell barrier against experimentally formed or introduced immunoglobulins seems unimportant, although its significance in preventing immunogenic contact is well founded (Section V,C). The reason for the segregation between male germ cells and somatic tissue is not obvious. It has been emphasized that the bamer might have appeared early during evolution, in order to protect the medium in which the spermatozoa are kept immotile until spermiation (Setchell and Waites, 1975; Abraham, 1983). An alternative or complimentary hypothesis concerning the role of the barrier is related to the fact that in primitive multicellular animals (e.g., Hydra and Dugesia) there is no early segregation between somatic and germ lines. In Coelenterata and Platyhelminthes, the transformation of somatic cells into germ cells is an epigenetic phenomenon. At the embryonic stage, germ cells are neither differentiated nor determined, and over a long period of development the future germ cells share the fate of the somatic cells. Eventually, they segregate from the somatic cells under the influence of external inductive actions by delayed regional chemodifferentiation. O’Donovan and Abraham (1988a) postulated, as one of the roles of the barrier in Coelenterata or Platyhelminthes, triggering of the differentiation of multipotential cells into male germ cells. The same may apply for more evolved phyla in which no continuous germ line has been observed (Nieuwkoop and Sutasurya, 1981). Such ontogenetically late differentiation has also been considered for Annelida. However, in a number of polychaete species, primordial germ cells have been traced from embryonic cells (Shroeder and Hermans, 1975). According to Loomis (1964), in Hydra littoralis, sexual differentiation is controlled by “microenvironmental” metabolites which leak from the ectodermal cells and induce gamete differentiation. In the diecious Hydra oligactis, Campbell (1985) found germ cells which determine the phenotypic sex of Hydra. In diecious Hydra species, Bosch and David (1986) assumed the existence of a suppressive factor, produced by male stem cells, which represses the differentiation of female stem cells. The MGPB might isolate this suppressive factor, thus controlling the formation of female gametes. The MGPB appeared at the dawn of metazoan evolution, together with the first organization of cells into tissues (O’Donovan and Abraham, 1988a). The segregation of totipotential embryonic cells from somatic cells, in order to give rise to male germ cells, might be the primal raison d’&treof the barrier. At the same time, it presumably served as a protective mechanism, permitting the spermatozoa to remain immotile for longer periods, in a state of “subripeness” prior to spermiation (Abraham, 1983). In the female gamete, subripeness is achieved by a mechanism which

MALE GERM CELL PROTECTIVE BARRIER

177

discontinues meiosis until ovulation. The necessity to keep the spermatozoa in an immotile state in a controlled and osmotically stable environment, might have maintained the barrier throughout phylogenesis from coelenterates to vertebrates. In mammals, and possibly in other amniotes, the barrier has come to fulfill additional functions, such as inhibition of immunogenic contact.

ACKNOWLEDGMENT Professor Jacob Lorch has valiantly struggled to extirpate from the manuscript stylesediments of different languages and streamline the text into English prose. My sincere thanks are due to him.

REFERENCES Abraham, M. (1981). Eur. Maricult. Soc., Spec. Publ. 6,207-219. Abraham, M. (1983). Abstr. Conf. Eur. Soc. Comp. Endocrinol., 12th p. 53. Abraham, M., Rahamim, E., and Tibika, H. (1979a). Abstr. Conf. Eur. SOC.Comp. Endmrinol., 10th p. I . Abraham, M., Sandri, C., and Akert, K. (1979b). Cell Tissue Res. 199,397-407. Abraham, M., Rahamim, E., Tibika, H., Golenser, E., and Kieselstein, M. (1980). Cell Tissue Res. 211,207-214. Abraham, M., Kieselstein, M., Hilge, V., and Lison, S. (1982). Cell Tissues Res. 225, 567-579.

Abraham, M., Hilge, V., Lison,. S., and Tibika, H. (1984). Cell Tissue Res. 235,403-410. Adiyodi, K. G., and Adiyodi, R. G., eds. (1983). “Reproductive Biology of Invertebrates,” Vol. 11. Wiley, New York. Ailenberg, M., Tung, P. S., Pelletier, M., and Fritz, I. B. (1988). Endocrinology (Baltimore) 12292604-2612.

Alvestad-Graebner, I., and Adam, H. (1977). Zool. Scr. 6, 113-126. Anderson, R. A., Jr., Quigg, J. M., Oswald, C., and Zaneveld, L. J. D. (1985). Biochem. Pharmacol. 34,685-695. Anderson, W. A., Weisman, A., and Ellis, R. A. (1967). J . Cell Biol. 32, 11-26. Anelli, G., Franchi, F., and Camatini, M. (1985). J . Submicrosc. Cytol. 17,213-222. Aoki, A. (1980). Eur. J . Cell Biol. 22,467. Assaykeen, T. A., and Thomas, J. A. (1965). Endocrinology (Baltimore) 76,839-843. Atwood, D. G. (1973). Z. ZeNforsch. 141,319-330. Au, C. L., Robertson, D. M., and de Kretser, D. M. (1986). Biol. Reprod. 35,37-43. Baccetti, B., Bigliardi, E., Talluri, M. V., and Bumni, A. G. (1983). J . Ultrastruct. Res. 85, 11-23.

Baldwin, K. M., Loeb, M. J . , and Riemann, J . G. (1987). Tissue Cell 19,413-421. Barker, C. F., and Billingham, R. E. (1977). Adv. Zmmunol. 2 5 , 1 4 4 . Barratt, C. L. R., Davies, A. G., Bansal, M. R., and Williams, M. E. (1989). Andrologia 21, 161-166.

Baynes, S. M., Scott, A. P., and Dawson, A. P. (1981). J. Fish Biol. 19,259-267. Benau, D., and Terner, C. (1980). Gamete Res. 3,247-257. Beniouri, R. (1984). Cytobios 40,159-170. Bennett, M. V. L., and Trinkaus, J. P. (1970). J . Cell Biol. 44,592-610.

178

MORDECHAI ABRAHAM

Benos, J. D., Kirk, R. G., Barba, W. D., and Goldner, M. M. (1977). Tissue Cell 9, 11-22. Bergmann, M. (1987). Adu. Anat., Embryol. Cell Biol. 105, 1-76. Bergmann, M., and Dierichs, R. (1983). Anat. Embryol. 168,269-275. Bergmann, M., and Schindelmeiser, J. (1987). Inr. J. Androl. 10,481-488. Bergmann, M., Greven, H., and Schindelmeiser, J. (1983). Cell Tissue Res. 232, 189-200. Bergmann, M., Greven, H., and Schindelmeiser, J. (1984a).I n t . J . Inuertebr. Reprod. Deu. 7, 291-296. Bergmann, M., Schindelmeiser, J., and Greven, H. (1984b). Cell Tissue Res. 238, 145-150. Bergmann, M., Nashan, D., and Nieschlag, E. (1989). Cell Tissue Res. 256, 183-190. Betz, A. L., Firth, J. A., and Goldstein, G. W. (1980). Brain Res. 192, 17-28. Biava, C. G., Smuckler, E. A., and Whorton, D. (1978). Exp. Mol. Parhol. 29,448-458. Bickell, L. R., Chia, F. S., and Crawford, B. J. (1980). J . Morphol. 166, 109-126. Bigliardi, E., and Vegni Talluri, M. (1976). Cell Tissue Res. 172,29-38. Billard, R. (1978). Aquaculture 14, 187-198. Billard, R. (1981). Aquaculture 23,287-293. Billard, R. (1986). Reprod. Nutr. Deu. 26,877-920. Billard, R. (1987). Fish. Sci. 1, 1-25. Billard, R., Jalabert, B., and Breton, B. (1972). Ann. Biol. Anim., Biochim., Biophys. 12, 19-32. Billingham, R. E., and Silvers, W. K. (1968). Proc. R . SOC.London, Ser. B 161, 168-190. Blanchard, M. G., and Josso, N. (1974). Pediatr. Res. 8,968-971. Bodenheimer, T. S., and Brightman, M. W. (1968). Am. J. Anat. l22,249-268. Bondi, C. and Farnesi, R. M. (1976). J. Morphol. 148,65-88. Bordin, S . , and Petra, P. H. (1980). Proc. Nail. Acad. Sci. U.S.A. 77,5678-5682. Born, W., and Wekerle, H. (1982). Am. J. Reprod. Immunol. 2,291-295. Bosch, T. C. G., and David, C. N. (1986). Proc. Nail. Acad. Sci. U.S.A. 83,9478-9482. Bouffard, G . (1906). Ann. Inst. Pasteur, Paris 20,539-546. Brannen-Brock, L. R., and Hall, J. L. (1985). Arch. Androl. 15, 15-19. Brightman, M. W., and Reese, T. S. (1969). J. Cell Biol. 40,648-677. Brightman, M. W., Klatzo, I., Olsson, Y.,and Reese, T. S. (1970). J. Neurol. Sci. 10, 215-239. Brown, P. C., and Glynn, L. E. (1969). J. Parhol. 98,277-282. Brown, D. V., Amann, R. P., and Wagley, L. M. (1983). Biol. Reprod. 28, 1257-1268. Brown, P. C., Glynn, L. E., and Holborow, E. J. (1967). Immunology W, 307-314. Buckland-Nicks, J., and Chia, F. S. (1986). Cell Tissue Res. 245, 305-313. Buckland-Nicks, J., Walker, C. W., and Chia, F. S. (1984). J . Morphol. 179,243-262. Burgos, M. H., Vitale-Calpe, R., and Aoki, A. (1970). I n “The Testis” (A. D. Johnson, W. S. Gomes, and N. L. Vandemark, eds.), Vol. I , pp. 551-649. Academic Press, New York. Burnett, A. L., Davis, L. E., and Ruffine, F. E. (1966). J. Morphol. 120, 1-8. Bussolati G., Alfani, V., Weber, K., and Osborn, M. (1980). J . Histochem. Cytochem. 28, 169-173. Byers, S. W., Hadley, M. A., Djakiew, D., and Dym, M. (1986). J. Androl. 7,59-68. Callard, G . , Mak, P., DuBois, W., and Cuevas, M. E. (1989). J . Exp. Zool. Suppl. 2,23-34. Camatini, M., and Ceresa-Castellani, L. (1974). J. Subrnicrosc. Cytol. 6,353-365. Camatini, M., Franchi, E., and deCurtis, I. (1979). J. Submicrosc. Cyrol. 11,511-516. Camatini, M., Franchi, E., and deCurtis, I. (1981). Anat. Rec. 200, 293-297. Camatini, M., Franchi, E., deCurtis, I., Anelli, G., and Masera, G. (1982a).Anar. Rec. 203, 353-363. Camatini, M., deCurtis, I., and Franchi, E. (1982b).J . Ulrrasrruct. Res. 79, 314-326. Cameron, D. F., Snydle, F. E., Ross, M. H., and Drylie, D. M. (1980). Ferril. Steril. 33, 157-160.

MALE GERM CELL PROTECTIVE BARRIER

179

Campbell, R. D. (1985). J. Exp. Zool. 234,451-458. Campbell, E., Takahashi, Y., Abramovitz, M., Peretz, M., and Listowsky, I. (1990).J . Biol. Chem. 265,9188-9193. Caspi, R. R., Roberge, F. G., and Nussenblatt, R. B. (1987). Science 237, 1029-1032. Castro, A. E., and Seiguer, A. C. (1974). Virchows Arch. B 16,297-309. Cavicchia, J. C., and Dym, M. (1978). Biol. Reprod. 18,219-228. Cavicchia, J. C., and Moviglia, G. A. (1983). Anat. Rec. 205, 387-396. Cavicchia, J. C., and Sacerdote, F. L. (1988). Tissue Cell 20, 577-586. Cerasaro, T. S . , Nachtsheim, D. A., Otero, F., and Parsons, C. L. (1984). J. Urol. 132, 577-579. Chakraborty, J., Nelson, L., Ihunjhunwala, J., Young, M., and Kropp, K. (1976). Cell Tissue Res. 174,261-271. Chao, N.-H., Chen, H.-P., and Liao, I.-C. (1975). Aquaculture 5,389-406. Chatton, E., and Tuzet, 0. (1941). C. R . Hebd. Seances Acad. Sci. 213,373-376. Chemes, H. E., Dym, M., Fawcett, D. W., Javadpour, N., and Sherins, R. J. (1977). Biol. Reprod. 17, 108-123. Cheng, C. Y., Grima, J., Stahler, M. S . , Guglielmotti, A., Silvestrini, B., and Bardin, C. W. (1990).Biochemistry 29, 1063-1068. Chia, F. S., and Buckland-Nicks, J. (1987). Znt. 1.Znuertebr. Reprod. Deu. 12, 173-184. Chiquoine, A. D. (1964). Anat. Rec. 149,23-36. Christen, R., Gatti, J. L., and Billard, R. (1987). Eur. J . Biochem. 166,667-671. Christensen, A. K . (1965). Anat. Rec. 151,335. Claude, P., and Goodenough, D. A. (1973). J. Cell Biol. 58,390-400. Clermont, Y. (1960). Ferril. Steril. 11, 563-573. Collenot, G., and Damas, D. (1980). Cah. Biol. Mar. 21,209-219. Connell, C. J. (1974). Anat. Rec. 178,333. Connell, C. J. (1978). J. Cell Biol. 76,57-75. Connell, C. J. (1980). In “Testicular Development, Structure, and Function” (A. Steinberger and E. Steinberger, eds.), pp. 71-78. Raven, New York. Cooksey, E. J., and Rothwell, B. (1973). J. Anat. 114,329-345. Cooper, T. G., and Waites, G . M. H. (1975). J . Endocrinol. 65, 195-205. Cordon-Cardo, C . , Brien, J. P. O., Casals, D., Rittman-Grauer, L., Biedler, J. L., Melamed, M. R., and Bertino, J. R. (1989). Proc. Natl. Acad. Sci. U.S.A. 86,695-698. Curtis, S. K., and Amann, R. P. (1981). J. Anim. Sci. 53, 1645-1657. Damas, D. (1968). Bull. SOC.Zool. Fr. 93,375-385. Davies, J. A. C., Winfrey, V., and Olson, G . E. (1983). Gamete Res. 4, 309-324. Davis, L. E. (1973). In “Biology of Hydra” (A. L. Burnett, ed.), pp. 171-219. Academic Press, New York. de Jong-Brink, M., Boer, H. H., Hommes, T. G . , and Kodde, A. (1977). Cell Tissue Res. 181, 37-58. de Jong-Brink, M., Schot, L. P. C., Schoenmakers, H. J. N., and Bergamin-Sassen, M. J. M. (1981). Gen. Comp. Endocrinol. 45,30-38. de Jong-Brink, M., Boer, H. H., and Joose, J. (1983). In “Reproductive Biology of Invertebrates” (K. G . Adiyodi and R. G. Adiyodi, eds.), Vol. I, pp. 297-355. Wiley, New York. de Jong-Brink, M., de With, N. D., Hurkmans, P. J. M., and Bergamin Sassen, M. J. M. (1984). Cell Tissue Res. 235,593-600. de Kretser, D. M., Kerr, J. B., and Paulsen, C. A. (1975). Biol. Reprod. 12, 317-324. de Kretser, D. M., Kerr, J. B., and Paulsen, C. A. (1981). Znt. J. Androl. 4, 129-144. Diamond, J. M., and Bossert, W. H. (1967). J . Gen. Physiol. 50,2061-2083. Diaz, J. P. (1973). Bull. SOC.Zool. Fr. 98, 145-156. Diaz, J. P., Connes, R., and Paris, J. (1973). C.R . Hebd. Seances Acad. Sci. 277,661-664.

180

MORDECHAI ABRAHAM

Dougherty, W. J., and Sandifer, D. A. (1984). Tissue Cell 16, 115-124. DuBois, W., and Callard, G. (1990). J. Exp. Zool. Suppl. 4, 142-144. Dumont, J. N., and Wallace, R. A. (1978). J. Exp. Zool. 204, 193-218. Dym, M. (1973). Anat. Rec. 175,639-656. Dym, M. (1976). Anar. Rec. 186,493-524. Dym, M., and Cavicchia, J. (1977). Biol. Reprod. 17,390-403. Dym, M., and Cavicchia, J. (1978). Biol. Reprod. 18, 1-15. Dym, M., and Fawcett, D. W. (1970). Biol. Reprod. 3,308-326. Dym, M., and Romrell, L. J. (1975). J . Reprod. Fertil. 42, 1-7. Dym, M., Djakiew, D., Hadley, M. A,, and Byers, S. W. (1986). Anat. Rec. 214,35A. Eckelbarger, K. J. (1984). J. Ultrastruct. Res. 89, 146-164. Eddy, E. M., and Kahri, A. I. (1976). Anat. Rec. 185,333-358. El-Demiry, M. I. M., Hargreave, T. B., Busuttil, A., James, K., Ritchie, A. W. S., and Chisholm, G. D. (1985). Br. J . Urol. 57,769-774. Ellison, J. D., and Jenkins, N. (1989). Anim. Reprod. Sci. 20, 1-10. Farquhar, M. G., and Palade, G. E. (1963). J. Cell Biol. 17, 375-412. Favard, P. (1961). Ann. Sci. Nut., Zool. 3,53-152. Fawcett, D. W. (1973). In “The Regulation of Mammalian Reproduction” (S. Segal, R. Crozier, P. Corfman, and P. Condiliffe, eds.), pp. 116-135. Thomas, Springfield, Illinois. Fawcett, D. W. (1975). In “Handbook of Physiology” (R. Greep, E. Astwood, D. Hamilton, and S. Geiger, eds.), Vol. 5, pp. 21-55. Am. Physiol. SOC.,Washington, D.C. Fawcett, D. W., Leak, L. V., and Heidger, P. M. (1970). J . Reprod. Fertil., Suppl. 10, 105- 122. Fawcett, D. W., Neaves, W. B., and Flores, M. N. (1973). Biol. Reprod. 9,500-532. Fell, P. E. (1974). In “Reproduction of Marine Invertebrates” (A. C. Giese and J. S. Pearse, eds.), Vol. 1, pp. 51-132. Academic Press, New York. Filshie, B. K., and Flower, N. E. (1977). J . Cell Sci. 23, 151-172. Flickinger, C. J. (1967). Z . Zel(forsch.Mikrosk. Anat. 78,92-113. Flickinger, C. J., and Fawcett, D. W. (1967). Anat. Rec. 158,207-222. Flickinger, C. J., Howards, S. S.,and English, H. F. (1978). A m . J . Anat. 152,557-586. Franchi, E., Camatini, M., and deCurtis, I. (1982). J. Ultrastruct. Res. 80, 253-263. Franchimont, P., Chari, S.,Hagelstein, M. T., and Duraiswami, S.(1975). Nature (London) 257,402-404. Franchimont, P., Verstraelen-Proyard, J., Hazee-Hagelstein, M. T., Renard, C. H., Demoulin, A., Bourguignon, J. P., and Hustin, J. (1979). Vitam. Horm. ( N . Y . ) 37, 243-302. Franke, W. W., Grund, C., Fink, A., Weber, K., Jockusch, B. M., Zentgraf, H., and Osborn, M. (1978). Biol. Cell. 31,7-14. Franklin, R. M., and Prendergast, R. A. (1970). J. Immunol. 104,463-469. Franquinet, R., and Lender, T. (1973). Z. Mikrosk.-Anat. Forsch. 87,4-22. Freeman, S.N., Mainwaring, W. I. P., and Furr, B. J. A. (1989). Br. J. Cancer60,664-668. Fretter, V., and Graham, A. (1964). In “Physiology of Mollusca” (K. M. Wilbur and C. M. Yonge, eds.), Vol. 1, pp. 127-164. Academic Press, New York. Friedlander, M. (1989). J. Insect Physiol. 35,29-39. Furuya, S.,Kumamoto, Y., Mori, M., and Sugiyama, S.(1980).In “Normal and Cryptorchid Testis” (E. S. E. Hafez, ed.), pp. 73-93. Nijhoff, The Hague, The Netherlands. Gallatin, M., St. John, T. P., Siegelman, M., Reichert, R., Butcher, E. C., and Weissman, I. L. (1986). Cell 44,673-680. Gilula, N. B. (1973).Am. Zool. W, 1109-1117. Gilula, N. B., Reeves, 0. R., and Steinbach, A. (1972). Nature (London)235,262-265.

MALE GERM CELL PROTECTIVE BARRIER

181

Gilula, N. B., Fawcett, D. W., and Aoki, A. (1976). Deu. Biol. 50, 142-168. Goldman, E. (1909). Bruns’ Beitr. Klin. Chir. 64, 192-265. Goldstein, G. W., and Betz, A. L. (1986). Sci. A m . 255,70-79. Condos, B . , and Sun, E. L . (1985). Ann. Clin. Lab. Sci. 15,320-321. Goresky, C. A. (1982). Fed. Proc., Fed. Am. SOC.Exp. Zool. 41,3033-3039. Gotow, T., and Hashimoto, P. H. (1984). J. Neurocytol. 13,727-742. Goyal, H. 0..and Hrudka, F. (1980). Andrologia 12,401-414. Goyal, H. O., and Hrudka, F. (1981). Andrologia 13,292-306. Gravis, C. J. (1978). A m . J . Anat. 151, 213-226. Gravis, C. J. (1979). Z. Mikrosk.-Anat. Forsch. 93, 321-342. Gravis, C. J., Yates, R. D., and Chen, I.-L. (1976). A m . J. Anat. 147,419-432. Gravis, C. J . , Chen, I.-L., and Yates, R. D. (1977). I n “Male Reproductive System: Fine Structure Analysis by Scanning and Transmission Electron Microscopy” (R. D. Yates and M. Gordon, eds.), pp. 35-55. Masson, New York. Green, C. R. (1981). Tissue Cell l3, 173-188. Green, C. R., and Bergquist, P. R. (1982). J. Cell Sci. 53,279-305. Greenberg, J., and Forssmann, W. G . (1983). Anat. Embryol. 168, 195-209. Gremigni, V. (1974). Bull. Zool. 41, 359-377. Gresik, E. W., Quirk, J. G., and Hamilton, J. B. (1973). Gen. Comp. Endocrinol. 21,341-352. Grier, H . J. (1975). J. Morphol. 146, 229-250. Grier, H. J. (1976). Cell Tissue Res. 168,419-432. Grier, H. J. (1981). Am. Zool. 21,345-357. Grier, H. J., Fitzsimons, J. M., and Linton, J. R. (1978). J. Morphol. 156,419-438. Griffith, L. M. (1980). Eur. J. Cell Biol. 22,290. Griffond, B., and Gomot, L. (1979). Cell Tissue Res. 202, 25-32. Grove, B. D., and Vogl, A. W. (1989). J. Cell Sci. 93, 309-323. Guyard, A. (1971). Ph.D. thesis. University of BesanGon, BesanGon, France. Haas, G. G., Cines, D. B., and Schreiber, A. D. (1980). N. Engl. J. Med. 13,722-727. Hadley, M., and Dym, M. (1987). Biol. Reprod. 37, 1283-1289. Hadley, M. A., Byers, S. W., Suarez-Quian, C. A., Kleinman, H. K., and Dym, M. (1985). J . Cell Biol. 101, 1511-1522. Hamilton, D. W. (1975). I n “Handbook of Physiology” (R. Creep, E. Astwood, D. Hamilton, and S. Geiger, eds.), Vol. 5, pp. 259-302. Am. Physiol. SOC.,Washington, D.C. Hamilton-Miller, J. M. T. (1973). Bacteriol. Rev. 37, 166-196. Hand, A. R. (1971). Anat. Rec. 169, 333A. Hand, A. R., and Gobel, S. (1972). J. Cell Biol. 52,397-408. Handel, M. (1979). J. Embryol. Exp. Morphol. 51,73-83. Hansson, V., Reusch, E., Trygstad, O., Torgersen, O., Ritzen, E. M., and French, F. S. (1973). Nature (London),New Biol. 246,56-58. Harada, Y., Abe, K., Murakami, Y., Ogasawara, H., Misawa, N., Inomata, N., and Yasuda, M. (1987). Teratology 36,467. Harada, Y., Takagi, H., Yamazaki, Y., Inomata, N., and Furukawa, Y. (1988). J. Toxicol. Sci. 13, 314. Hank, S. I., Whitney, P., Perry, G . , Kalaria, R. N., Anderson, L., Lundahl, P., and Cleveland, 0. H. (1989). Neurology 39 Suppl. I , 371. Hausman, R. H. (1973). I n “Biology of Hydra” (A. L. Burnett, ed.), pp. 393-453. Academic Press, New York. Haynes, J. F. (1973). I n “Biology of Hydra” (A. L. Burnett, ed.), pp. 233-237. Academic Press, New York.

182

MORDECHAI ABRAHAM

Head, J. R., and Billingham, R. E. (1985). Transplantation 40,269-275. Head, J. R., Neaves, W. B., and Billingham, R. E. (1983). Transplantation 36,423-431. Henley, C . (1974). I n “Reproduction of Marine Invertebrates” (A. C. Giese and J. S. Pearse, eds.), pp. 267-343. Academic Press, New York. Hermo, L., and Morales, C. (1984). Am. J . Anat. 171,59-74. Hettle, J. A., Waller, E. K., and Fritz, I. B. (1986). Biol. Reprod. 34,895-904. Hill, R. S. (1977). Cell Tissue Res. 183, 131-141. Hinsch, G. W., and Dehn, P. F. (1979). Int. J . Inuertebr. Reprod. 1, 179-185. Hintz, M., and Goldberg, E. (1977). Deu. Biol. 57,375-384. Hirschhauser, C., and Baudner, S. (1972). Fertil. Sferil. 23, 393-398. Hoffer, A. P. (1982). Arch. Androl. 8,233-246. Hoffer, A. P. (1983). Biol. Reprod. 28, 1007-1020. Holland, N. D. (1967). Biol. Bull. (Woods Hole, Mass.) l33, 578-590. Holland, N. D., and Holland, L. Z. (1989). Acta Zool. (Stockholm)70,211-219. Holstein, A. F., Bustos-Obregon, E., and Hartmann, M. (1984). Cell Tissue Res. 236, 35-40. Howards, S. S., Jessee, S. J., and Johnson, A. L. (1976). Biol. Reprod. 14, 264-269. Huang, H., and Hembree, W. (1979). Biol. Reprod. 21,891-904. Huang, H., and Marshall, G. (1983). Biol. Reprod. 28, 1163-1 172. Huang, H., Dyrenfurth, I., and Hembree, W. (1983). Endocrinology (Baltimore) 112, 11631171. Huang, H. F. S., Yang, C. S., Meyenhofer, M., Gould, S., and Boccabella, A. V. (1988). Acta Anat. 133, 10-15. Huang, T. T. F., Tung, K. S. K., and Yanagimachi, R. (1981). Science 213, 1267-1269. Hudspeth, A. J., and Revel, J. P. (1971). J. Cell B i d . 50,92-101. Hufnagel, L., and Kass-Simon, G. (1976). I n “Coelenterate Ecology and Behavior” (G. D. Mackie, ed.), pp. 695-704. Plenum, New York. Hurtenbach, U., Morgenstern, F., and Bennett, D. (1980). J . Exp. Med. 151,827-838. Imms, A. D. (1957). “A general Textbook of Entomology,” 9th Ed. Methuen, London. Jalkanen, S., Bargatze, R. F., de 10s Toyos, J., and Butcher, E. C. (1987). J. Cell Biol. 105, 983-990. Jenkins, N., and Ellison, J. D. (1989). Anim. Reprod. Sci. 20, 11-20. Johnson, M. H. (1968). J. Reprod. Fertil. 16,503-506. Johnson, M. H. (1969). J . Reprod. Fertil. 19,551-553. Johnson, M. H. (1970a). J . Pathol. 101, 129-139. Johnson, M. H. (1970b). J . Reprod. Fertil. 22, 119-127. Johnson, M. H. (1972). Fertil. Steril. 23, 383-392. Johnson, M. H. (1973). Adu. Reprod. Physiol. 6,279-324. Jones, R. (1978). J . Cell Sci. 31, 145-164. Jones, N. A., Willson, J. T., and Katsh, S. (1971). Anat. Rec. 169, 350. Jones, R., Hamilton, D. W., and Fawcett, D. W. (1979). Am. J. Anat. 156,373-400. Jones, R. C., Jones, N., and Djakiew, D. (1984). J. Exp. Zool. 230,417-426. Joosse, J., and Reitz, D. (1969). Malacologia 9, 101-109. Kambysellis, M. P., and Williams, C. M. (1971a). Biol. Bull. Mar. Biol. Lab., Woods Hole, MUSS.141,527-540, Kambysellis, M. P., and Williams, C. M. (1971b). Biol. Bull. Mar. B i d . Lab., Woods Hole, Mass. 141,541-552. Kaplan, H. J., and Streilein, J. W. (1977a). J . Immunol. 118,809-814. Kaplan, H. J . , and Streilein, J. W. (1977b). J. Immunol. 120,689-693. Karnovsky, M. (1965). J. Cell Biol. 27,49a. Karnovsky, M. (1967). J . Cell B i d . 35,213-236.

MALE GERM CELL PROTECTIVE BARRIER

183

Khan, S. A., Soder, O., Syed, V., Gustafsson, K., Lindh, M., and Ritzen, E. M. (1987). I n t . J . Androl. 10,495-503. Kierszenbaum, A. L., and Mancini, R. E. (1973). J. Reprod. Ferril. 33, 119-122. Kinsky, S. L. (1970). Annu. Rev. Pharmacol. 10, 119-142. Kormano, M. (1967). Histochemie 9,327-338. Kormano, M. (1968). Acta Histochem. 30, 133-136. Koskimies, A. I., Kormano, M., and Lahti, A. (1971). J. Reprod. Fertil. 27,463-465. Koskimies, A. I . , Kormano, M., and Alfthan, 0. (1973). J. Reprod. Fertil. 32,79-86. Koulish, S . , and Kramer, C. R. (1986). Tissue Cell 18, 383-393. Kuopio, T., and Pelliniemi, L. J. (1989). Cell Tissue Res. 256,45-51. Kuopio, T., Paranko, J., and Pelliniemi, L. J. (1989). Differentiation (Berlin)40,198-206. Lacroix, M., Smith, F. E., and Fritz, I. B. (1977). Mol. Cell. Endocrinol. 9,227-236. Lagueux, M., Him, M., de Reggi, M., and Hoffmann, J. (1976). C. R . Hebd. Seances Acad. Sci., Ser. D282,1187-1190. Laird, M. L., Ellis, A. E., Wilson, A. R., and Holliday, F. G. T. (1978). Ann. Biol. Anim., Eiochim., Eiophys. 18, 1101-1106. Laird, L. M., Wilson, A. R., and Holliday, F. G. T. (1980). Reprod. Nurr. Deu. 20, 17811788. Lambiase, J . T., and Amann, R. P. (1973). Fertil. Steril. 24,65-67. Lane, N. J. (1979). Tissue Cell 11, 1-18. Lane, N . J., and Skaer, H. (1980). Adu. Insect Physiol. 15,35-213. Lane, N . , Skaer, H., and Swales, L. (1977). J . Cell Sci. 26, 175-199. Larsen, W. J. (1983). Tissue Cell 15,645-671. Lautie, N. (1981). Arch. Zool. Exp. Gen. 122, 199-210. Lawrence, T. S . , Beers, W. H., and Gilula, N. B. (1978). Nature (London) 272,501-505. Leblond, C. P., and Clermont, Y. (1952a). Ann. N . Y . Acad. Sci. 55,548-573. Leblond, C . P., and Clermont, Y. (1952b). A m . J . Anat. 90, 167-214. Lee, B . C., Pineda, J. L., Spiliotis, B. E., Brown, T. J., and Bercu, B. B. (1983). Biol. Reprod. 28, 1207-1215. LeGac, F., and de Kretser, D. M. (1982). Mol. Cell. Endocrinol. 28,487-498. Legendre, M., and Billard, R. (1980). Bull. Fr. Piscic. 278, 11-33. Lenz, T. L., and Barrnett, R. J. (1965). J. Ultrastruct. Res. 13, 192-211. Lewis-Jones, D. I., Richards, R. C., Lynch, R. V., and Joughin, E. C. (1987). Br. J . Urol. 59, 452-457. Lindner, S.G. (1982). Andrologia 14, 352-362. Lipshultz, L. I., Murthy, L., and Tindall, D. J. (1982). J . Clin. Endocrinol. Metab. 55, 228-237. Lockwood, A. P. M. (1961). Comp. Biochem. Physiol. 2,241-289. Longo, F. J . , and Anderson, E. (1969). J. Ultrastruct. Res. 27,486-509. Loomis, W. F. (1964). J. Exp. Zool. 156,289-306. Lord, B. A. P., and DiBona, D. R. (1976). J . Cell Biol. 71,967-972. Lou, Y . H., and Takahashi, H. (1987). Hokkaido Daigaku Suisangakubu Kenkyu Iho 38, 14-26. Lou, Y. H., and Takahashi, H. (1988). Hokkaido Daigaku Suisangakubu Kenkyu Iho 39, 201-209. Lou, Y. H., and Takahashi, H. (1989). Cell Tissue Res. 258,491-498. Louis, B. G . , and Fritz, I. B. (1977). Mol. Cell. Endocrinol. 7,9-16. Louis, B. G., and Fritz, I. B. (1979). Endocrinology (Baltimore ) 104,454-461. Luchtel, D. (1972a). Z. Zellforsch. Mikrosk. Anat. 130,279-301. Luchtel, D. (1972b). Z. Zellforsch. Mikrosk. Anat. 130,302-311. MacLeod, J. (1 965). Fertil. Steril. 16, 735-757.

184

MORDECHAI ABRAHAM

Maddocks, S., Oliver, J. R., and Setchell, B. P. (1984). Co1loq.-Inst. Natl. Sante Rech. Med. U3,497-502. Mahi-Brown, C. A., Yule, T. D., and Tung, K. S. K. (1988). Am. J . Reprod. Immunol. Microbiol. 16, 165-170. Mancini, R. E., Huidobro, J., Fernandez, E., Collazo, E., and Monastirsky, R. (1966). Proc. SOC.Exp. Biol. Med. 123,227-230. Mann, T. (1967). Colloq. Endocrinol. 16,233-248. Marcaillou, C., and Lauvejat, S. (1986). Can. J . Zool. 64,2053-2061. Marcaillou, C., and Szollosi, A. (1975). C. R . Hebd. Seances Acad. Sci. 281,2001-2004. Marcaillou, C., and Szollosi, A. (1980). J. Ultrastruct. Res. 70, 128-136. Marcaillou, C., Szollosi, A., and Porcheron, P. (1977). Bull. SOC.Zool. Fr. 102,313. Marcaillou, C., Szollosi, A., Porcheron, P., and Dray, F. (1978). Cell Tissue Res. 188,63-74. Marshall, W. S., and Bryson, S. E. (1988). Can. J. Zool. 66, 1603-1609. Marshall, W. S., Bryson, S. E., and Idler, D. E. (1989). Gen. Comp. Endocrinol. 75,118-128. Mattei, X . , Mattei, C., Marchand, B., and Leung Tack Kit, D. (1982).J. Ultrastruct. Res. 81, 333-340. Mayerhofer, A., and Bartke, A. (1989). Anat. Rec. 223,77A. McFadden, M. R., and Mehan, D. J. (1978). J. Urol. 119,372-374. McGinley, D. M., Posalaky, Z. Porvaznik, M., and Russell, L. (1979). Tissue Cell 11, 741-754. Mehan, S. P., Asnani, M. V., and Ramachandran, A. V. (1989). Nutr. Res. 9, 1381-13%. Menge, A. C., and Peegel, H. (1980). Arch. Androl. 4, 171-178. Meyer, R., Posalaky, Z., and McGinley, D. (1977). J. Ulrrastrucr. Res. 61,271-283. Meyran, J. C. (1982). J . Ultrastruct. Res. 79, 31-46. Middleton, A. (1973). Ph.D. dissertation. University of Cambridge, Cambridge, England. Miller, R. L. (1983). I n “Reproductive Biology of Invertebrates” (K. G. Adiyodi and R. G. Adiyodi, eds.), Vol. 11, pp. 23-73. Wiley, New York. Miranda, J. C., and Cavicchia, J. C. (1986). Tissue Cell 18,461-468. Miranda, J. C., and Cavicchia, J. C. (1988). Tissue Cell 20, 61 1-619. Misfeldt, D. S., Hamamoto, S. T., and Pitelka, D. R. (1976). Proc. Nafl.Acad. Sci. U.S.A. 73, 1212-1216. Miyamoto, A,, Umezu, M., Ishii, S., Furusawa, T., Masaki, J., Hasegawa, Y., and Ohta, M. (1989). Anim. Reprod. Sci. 20, 165-178. Morales, C., and Griswold, M. D. (1987). Endocrinology (Baltimore) 121,432-434. Morales, C., Hermo, L., and Clermont, Y. (1984). Anat. Rec. 209, 185-195. Morales, C., Sylvester, S., and Griswold, M. D. (1987). Anal. Rec. 218,93A. Morisawa, M., and Okuno, M. (1982). Nature (London)295,703-704. Morisawa, M., and Suzuki, K. (1980). Science 210, 1145-1147. Morisawa, M., Hirano, T., and Suzuki, K. (1979). Comp. Biochem. Physiol. 64,325-329. Muller, U., and Wolf, U. (1979). Dlfferentiation (Berlin) 14, 185-187. Nagano, T. (1980). J. Electron Microsc. 29,250-255. Nagano, T., and Suzuki, F. (1976). Anat. Rec. 185,403-418. Nagano, T . , and Suzuki, F. (1978). Cell Tissue Res. 189,389-401. Nagano, T., and Suzuki, F. (1983). Inr. Rev. Cyrol. 81, 163-190. Nagler, H. M., and White, R. (1982). J . Urol. Us,1343-1348. Nalbandov, A. V. (1976). “Reproductive Physiology of Mammals and Birds.’’ Freeman, San Francisco, California. Naz, R. K.,and Bhargava, K. K. (1990). Mol. Reprod. Deu. 26, 175-183. Neaves, W. B. (1973). J. Cell Biol. 59,559-572. Nemetallah, B. R., and Ellis, L. C. (1985). Arch. Androl. 15,41-48. Nemetallah, B. R., Howell, R. E., and Ellis, L. C. (1985). Arch. Androl. 15,79-82.

MALE GERM CELL PROTECTIVE BARRIER

185

Nicander, L. (1967). Z . Zellforsch. 83,375-397. Nieuwkoop, P. D., and Sutasurya, L. A. (1981). “Primordial Germ Cells in the Invertebrates: From Epigenesis to Preformation.” Cambridge Univ. Press, London. Nikolsky, G. V. (1963). “The Ecology of Fishes.” Academic Press, New York. Noirot-Timothee, C., and Noirot, C. (1980). Int. Rev. Cyrol. 63,97-140. O’Donovan, P. (1988). Ph.D. thesis. The Hebrew University, Jerusalem, Israel. O’Donovan, P., and Abraham, M. (1983). Absrr., Conf. Eur. SOC.Comp. Endocrinol., 12th p. 58. O’Donovan, P., and Abraham, M. (1987). J. Morphol. 192,217-227. O’Donovan, P., and Abraham, M. (1988a). J . Morphol. 198,179-188. O’Donovan, P., and Abraham, M. (1988b). Abstr., Conf.Eur. SOC.Comp. Endocrinol., 14th p. 45. O’Donovan-Lockard, P., Sagi, G., Villwock, W., Hilge, V., and Abraham, M. (1990). Eur. Aquacult. SOC.,Spec. Publ. 11, 199-214. Olsson, Y., and Reese, T. S. (1969). Anat. Rec. 163,318-319. O’Rand, M. G. (1980). Dev. Biol. 75,431-441. O’Rand, M. G., and Romrell, L. J. (1977). Dev. Biol. 55,347-358. Osman, D. I., Eckwall, H., and Ploen, L. (1980). I n t . J. Androl. 3,553-562. Ouyang, Y., and Lien, E. J. (1984). Acta Pharm. Jugosl. 34,201-205. Palombi, F., Ziparo, E., Galdieri, M., Russo, M. A., and Stefanini, M. (1980). Biol. Cell. 39, 249-252.

Pardridge, W.M. (1988). I n “Steroid-Protein Interactions” (R. Frairia, H. L. Bradlow, and G. Gaidano, eds.), Vol. 538, pp. 173-192. Ann. N.Y. Acad. Sci., New York. Pan, G. A. (1910). Frankf. Z . Pathol. 4, 1-28. Parmentier, H. K., andTirnmermans, L. P. M. (1985). J. Embryol. Exp. Morphol. 90,13-32. Parmentier, H. K., Timmermans, L. P. M., and Egberts, E. (1984). Cell Tissue Res. 236, 99-105.

Parmentier, H. K., van den Boogaart, J. G. M., and Timmermans, L. P. M. (1985). Cell Tissue Res. 242,75-81. Parvinen, M. (1982). Endocr. Rev. 3,404-417. Patisson, C . (1%5). C . R . Hebd. Seances Acad. Sci. 261,2950-2953. Patisson, C. (1966). Ann. Univ. ARERS 4,67-75. Patisson, C. (1977). Ann. Sci. Nut., 2001.19,315-347. Pechan, P., Wachtel, S.S., and Reinboth, R. (1979). Differentiation (Berlin) 14, 189-192. Pelletier, R. M. (1986). A m . J. Anat. 175,91-117. Pelletier, R. M. (1988). A m . J. Anar. 183,68-102. Pelletier, R. M. (1990). A m . J. Anat. 188, 87-108. Pelletier, R. M., and Friend, D. S. (1983). A m . J . Anat. 168,213-228. Pelletier, R. M., and Friend, D. S. (1986). J. Androl. 7, 127-139. Pelletier, R. M., Nemirovsky, M. S., Calvert, R., and Hugon, J. S. (1981). Anat. Rec. 199, 197-2 11.

Pelliniemi, L. J., Dym, M., Gunsalus, G. L., Musto, N. A., Bardin, C. W., and Fawcett, D. W . (1981). Endocrinology (Baltimore) 108,925-931. Picard, J. Y., Tran, D., and Josso, N. (1978). Mol. Cell. Endocrinol. 12, 17-30. Pickard, J. D., Durity, F., Welsh, F. A., Langfitt, T. W., Harper, A. M., and MacKenzie, E. T. (1977). Brain Res. 122, 170-176. Picon, R. (1969). Arch. Anat. Microsc. Morphol. Exp. 58, 1-19. Picon, R. (1970). C . R . Hebd. Seances Acad. Sci. 271,2370-2372. Pilsworth, L. M., and Setchell, B. P. (1981). In “The Testis” (H. Burger and D. de Kretser, eds.), pp. 9-38. Raven, New York. Pollanen, P., and Maddocks, S. (1988). J. Reprod. Fertil. 82,437-445.

186

MORDECHAI ABRAHAM

Pollanen, P., Soder, O., and Uksila, J. (1988). J. Reprod. Immunol. 14, 125-138. Pollard, T. D. (1980). Eur. J. Cell Biol. 22,289. Posinovec, J. (1989). Diabetol. Croar. 18,91-95. Potashnik, G . , Yanai-Inbar, I., Sacks, I., and Israeli, R. (1979). Isr. J . Med. Sci.15,438-442. Price, J. M. (1979). Am. J. Anar. 156, 147-158. Rajfer, J., Turner, T. T., Rivera, F., Howards, S. S., and Sikka, S. C. (1987). Biol. Reprod. 36,933-937. Raju, S., and Grogan, J. B. (1969). Transplantation 7,475-483. Ramos, A. S., and Dym, M. (1977). Biol. Reprod. 17, 339-349. Rapoport, S. I., and Thompson, H. K. (1973). Science 180,971. Rastogi, R. K. (1979). J. Steroid Biochem. 11, 101-105. Rastogi, R. K., Milone, M., Di Meglio, M., Caliendo, M. F., and Chieffi, G. (1979). J . Reprod. Fertil. 57,73-77. Ratclfie, N. A., Rowley, A. F., Fitzgerald, S. W., and Rhodes, C. P. (1985). Int. Rev. Cytol. 97, 183-350. Rattner, J. B. (1970). J . Cell Biol. 47, 168a. Reese, T. S., and Karnovsky, M. J. (1967). J. Cell Biol. 34,207-217. Reiswig, H. M. (1983). In “Reproductive Biology of Invertebrates” (K. G. Adiyodi and R. G. Adiyodi, eds.), Vol. 11, pp. 1-21. Wiley, New York. Ribbert, H. (1904). Z. Allg. Physiol. 4,201-214. Robinson, J. M., and Karnovsky, M. J. (1980). J . Hisrochem. Cytochem. 28, 161-168. Rodriguez, L. A. (1955). J. Comp. Neurol. 102,27-45. Roosen-Runge, E. (1977). In “Developmental and Cell Biology Series” (M. Abercrombie, D. R. Newth, and J. G. Torrey, eds.). Cambridge Univ. Press, London. Ross, M. H. (1970). Adu. Androl. 1,83-86. Ross, M. H. (1977). Am. J. Anat. 148,49-56. Rothwell, B., and Tingari, M. D. (1973). J. Anat. 114,321-328. Rumke, P., and Titus, M. (1970). J. Reprod. Fertil. 21,69-79. Russell, L. D. (1977a). Am. J . Anat. 148,313-328. Russell, L. D. (1977b). Tissue Cell 9,475-498. Russell, L. D. (1978). Anat. Rec. 190,99-I 12. Russell, L. D. (1979a). Am. J. Anat. 155,259-280. Russell, L. D. (1979b). Anar. Rec. 194,213-232. Russell, L. D. (1980). Gamete Res. 3, 179-202. Russell, L. D., and Burguet, S. (1977). Tissue Cell9,751-766. Russell, L. D., and Clermont, Y. (1976). Anat. Rec. 185,259-278. Russell, L. D., and Peterson, R. N. (1985). Int. Reu. Cyrol. 94, 177-211. Russell, L. D., and Wong, V. (1981). J. Androl. 2,24-25. Russell, L. D., Malone, J. P., and MacCurdy, D. S. (1981). Tissue Cell 13,349-367. Sakiyama, R., Pardridge, W. M., and Musto, N. A. (1988). J. Clin. Endocrinol. Metab. 67, 98-103. Salonen, I., and Eriksson, C. J. P. (1989). Alcohol Clin.Exp. Res. W, 746-751. Sano, M., Ohshima, A., Kawamura, N., Kitajima, S., and Mizutani, A. (1987).J . Exp. Zool. 241951-59. Sawada, N. (1984). Fortschr. Zool. 29,W-114. Saypol, D. C., Turner, T. T., Howards, S . S., and Miller, E. D. (1981). J. Clin. Invest. 68, 39-45. Schincariol, A. L., Haborsky, J., and Winner, G. (1967). Can. J. 2001.45,590-593. Schlenk, W., and Kahmann, H. (1938). Biochem. Z. 295,283-301. Schmell, E. D., Yuan, L. C., Gulyas, B. J., and Thomas August, I. (1982). Ferril. Steril. 37, 249-257.

MALE GERM CELL PROTECTIVE BARRIER

187

Schulze, C. (1988). Cell Tissue Res. 251,31-43. Schweich, P., and Leloup, A. M. (1988). Ann. SOC.R . Zool. Belg. 118,29-40. Scott, R. S., Burger, H., and Quigg, H. (1980). Endocrinology (Baltimore) 107, 1536-1542. Secombes, C. J. (1986). Vet. Immunol. Immunopathol. 12, 193-201. Secombes, C. J., Laird, L. M., and Priede, I. G. (1982). In “Reproductive Physiology of Fish” (C. J. J. Richter and H. I. T. Goos, eds.), p. 81. Pudoc, Wageningen, The Netherlands. Secombes, C. J., Lewis, A. E., Needham, E. A., Laird, L. M., and Priede, I. G. (1985). J . EXP.ZOO/.233,425-431. Secombes, C. J . , van Winkoop, A., van den Boogaart, J. G. M., Timmermans, L. P. M., and Priede, I. G. (1986). Aquaculture 52, 125-135. Secombes, C. J., Laird, L. M., and Priede, I. G. (1987). Aquaculture 60,287-302. Sertoli, E. (1865). Morgagni 7,31-39. Setchell, B. P. (1967). J. Physiol. (London) 189, 63-65. Setchell, B. P. (1980). J. Androl. 1,3-10. Setchell, B. P., and Brooks, D. E. (1988). In “The Physiology of Reproduction” (E. Knobil and J. D. Neill, eds.), Vol. I, pp. 753-836. Raven, New York. Setchell, B. P., and Main, S. J. (1975). Curr. Top. Mol. Endocrinol. 2,223-233. Setchell, B. P., and Main, S. J. (1978). EHP, Environ. Health Perspect. 24, 61-64. Setchell, B. P., and Waites, G. M. H. (1970). J. Endocrinol. 47, 81-86. Setchell, B. P., and Waites, G. M. H. (1975). I n “Handbook of Physiology” (R. Greep, E. Astwood, D. Hamilton, and S. Geiger, eds.), Vol. 5, pp. 143-172. Am. Physiol. SOC., Washington, D.C. Setchell, B. P., Voglmayr, J. K., and Waites, G. M. H. (1969). J. Physiol. (London) 200, 73-85. Setchell, B. P., Pollanen, P., and Zupp, J. L. (1988). Int. J. Androl. 11,225-233. Shaw, S . R. (1977). J. Comp. Physiol. 113,257-282. Shaw, S . R. (1978). Cell Tissue Res. 188,35-61. Shemi, D.,Sod-Moriah, U. A., Kaplanski, J., Potashnik, G., and Yanai-Inbar, I. (1982). Andrologia 14, 191-199. Shemi, D., Marx, Z., Kaplanski, J., Potashnik, G., and Sod-Moriah, U. A. (1988). Andrologia 20,331-337. Shemi, D., Sod-Moriah, U. A., Abraham, M., Friedlander, M., Potashnik, G., and Kaplanski, J. (1989). Andrologia 21,229-236. Shepu, X., Shudong, Z., Shuyan, S. Yanwan, W., Yi, L., Zenghua, Z., and Xixin, M. (1980). Sci. Sin. 23,642-659. Shibata, N., and Hamaguchi, S. (1986). Zool. Sci. 3,331-338. Shinoda, K., Okarniya, H., Imazawa, T., Furukawa, F., Toyoda, K., Sato, H., and Takahashi, M. (1989). Eisei Shikensho Hokoku 107,63-67. Shivers, R. R. (1979). Brain Res. 169,221-230. Shroeder, P. C., and Hermans, C. 0. (1975). I n “Reproduction of Marine Invertebrates” (A. C. Giese and J. S. Pearse, eds.), vo1.3, “Annelids and Echiurans” pp. 1-213. Academic Press, New York. Shulman, S. (1975). “Reproduction and Antibody Response.” CRC Press, Cleveland, Ohio. Siiteri, P. K., Murai, J. T., Hammond, G. L., Nisker, J. A., Raymoure, W. J., and Kuhn, R. W. (1982). Recent Prog. Horm. Res. 38,457-510. Skelding, J. M. (1973). Z . Zellforsch. 147, 1-29. Skinner, M. K., and Fritz, I. B. (1985). Proc. Natl. Acad. Sci. U.S.A. 82, 114-118. Skinner, M. K., and Fritz, I. B. (1986). Mol. Cell. Endocrinol. 44,85-97. Skinner, M. K., and Griswold, M. D. (1980). J. Biol. Chem. 225, 1923-1925. Skinner, M. K., and Griswold, M. D. (1982). Biol. Reprod. 27,211-221.

188

MORDECHAI ABRAHAM

Skinner, M. K., and Griswold, M. D. (1983). Biol. Reprod. 28, 1225-1229. Slavis, S. A., Scholz, J. N., Hewitt, C. W., Black, K. S., Campbell, R. S., Patel, M., Zimmerman, J., Peake, M. L., and Martin, D. C. (1990). J. Urol. 143,638-641. Soos, L. (1910). Ann. Mus. Natl. Hung.8,304-343. Staehelin, L. A. (1973). J. Cell Sci. 13,763-786. Staehelin, L. A. (1974). Int. Rev. Cytol. 39, 191-284. Stang-Voss, C. (1970). Z . Zeluorsch. Mikrosk. Anat. 109,287-296. Stewart, R. J., Boyd, S., Brown, S., and Toner, P. G. (1990). J . Pathol. 160,51-55. Steyn, G . J., and Van Vuren, J. H. J. (1986). Comp. Biochem. Physiol. A 83A,421-425. Steyn, G. J., Van Vuren, J , H. J., Schoonbee, H. J., and Chao, N.-H. (1985). Water SA 11, 15-18.

Storch, V., and Welsch, U. (1977). Cell Tissue Res. 184,423-425. Streilein, J. W. (1987). FASEB J. 1, 199-208. Sullivan, J., Smith, F. A., and Garman, R. H. (1979). J. Reprod. Fertil. 5,201-207. Sun, E. L., and Gondos, B. (1986). Cell Tissue Res. 243,575-578. Suzuki, F., and Nagano, T. (1978). Anat. Rec. 191,503-520. Suzuki, F., and Nagano, T. (1979). Cell Tissue Res. 198,247-260. Szollosi, A. (1982). In “Insect Ultrastructure” (R. C. King and H. Akai, eds.), Vol. I, pp. 31-60. Plenum, New York. Szollosi, A., and Marcaillou, C. (1977). J. Ultrastruct. Res. 59, 158-172. Szollosi, A., and Marcaillou, C. (1979). J . Ultrastruct. Res. 69, 331-342. Szollosi, A., and Marcaillou, C. (1980). Cell Tissue Res. 213, 137-147. Szollosi, A., Riemann, J., and Marcaillou, C. (1980). J. Ultrastruct. Res. 72, 189-199. Tanaka, M., and Ujeno, Y. (1989). Annu. Rep. Res. React. Inst., Kyoto Uniu. 22, 118-121. Tardent, P. (1974). Am. Zool. 14,447-456. Terry, A., Terry, R. J., and Worms, M. J. (1961). J. Parasitol. 47,703-71 1. Theodor, J. L. (1970). Nature (London) 227,690-692. Timmermans, L. P. M. (1987). Sarsia 72,331-339. . Timmermans, L. P. M.,and Taverne, N. (1983). Acta Morphol. N e e r l d c a n d . 21, 182. Timmermans, L. P. M., Parmentier, H. K., and van den Boogaart, J. G . M. (1985). Cell Biol. Int. Rep. 9,518. Tindall, D. J., Vitale, R., and Means, A. R. (1975). Endocrinology (Baltimore) 97,636-648. Tindall, D. J., Rowley, D. R., Murthy, L., Lipshultz, L. I., and Chang, H. C. (1985). Int. Rev. Cytol. 94, 127-149. Tormey, J. M. (1966). Nature (London) 210,820-822. Toshimori, K., Iwashita, T., and Oura, C. (1979). Cell Tissue Res. 202,63-73. Toyama, Y. (1976). Anat. Rec. 186,477-492. Tran, D., Meusy-Dessolle, N., and Josso, N. (1977). Nature (London) 269,411-412. Tran, D., Meusy-Dessolle, N., and Josso, N. (1981). Biol. Reprod. 24,923-931. Tung, K. S. K. (1978). Science 201,833-835. Tung, K. S. K. (1980). In “Immunological Aspects of Infertility and Fertility Regulation” (D. S. Dhindsa and G. F. B. Schumacher, eds.), pp. 33-91. ElsevierlNorth-Holland, New York. Tung, K. S. K., Unanue, E. R., and Dixon, F. J. (1971). J . Immunol. 106, 1463-1472. Tung, K. S. K., Han, L. B., and Evan, A. P. (1979). Deu. Biol. 68,224-238. Tung, K. S. K., Ellis, L. C., Teuscher, C., Meng, A., Blaustein, J. C., Kohno, S., and Howell, R. (1981). J. Exp. Med. 154, 1016-1032. Tung, K. S. K., Ellis, L. C., Childs, G. V., and Dufau, M. (1984). Endocrinology (Baltimore) 114,922-929.

Tung, P. S . , and Fritz, I. B. (1978). Deu. Biol. 64,297-315.

MALE GERM CELL PROTECTIVE BARRIER

189

Tung, P. S., and Fritz, 1. B. (1984). Biol. Reprod. 30,213-229. Turner, T. T., D’Addario, D. A., and Howards, S. S. (1980). J . Reprod. Fertil. 60,285-289. Turner, T. T., Cochran, R. C., and Howards, S. S. (1981a). Biol. Reprod. 25,342-348. Turner, T. T., Giles, R. D., and Howards, S. S. (1981b). J . Reprod. Fertil. 63,355-358. Turner, T. T., Saypol, D. C., and Howards, S. S. (1982). Biol. Reprod. 26 (Suppl. I), 74. Turner, T. T., Jones, C. E., and Roddy, M. S. (1987). Biol.Reprod. 36,926-932. Tuzet, O., Garrone, R., and Pavans de Ceccatty, M. (1970). Ann. Sci. Nut., Zool. l2,27-50. Tyler, A., and Lord Rothschild (1951). Proc. SOC.Exp. Biol.Med. 76,52-62. Unsicker, K. (1975). Cell Tissue Res. 158,215-240. van den Hurk, R., and Barends, P. (1974). Proc. K. Ned. Akad. Wet., Ser. C C77,205-214. van den Hurk, R., and Testerink, G. J. (1975). Proc. K. Ned. Akad. Wet., Ser. C C78, 1-10. van den Hurk, R., Meek, J., and Peute, J. (1974a). Proc. K. Ned. Akad. Wet., Ser. C C77, 460-469. van den Hurk, R., Meek, J., and Peute, J. (1974b). Proc. K. Ned. Akad. Wet., Ser. C C77, 470-476. van den Hurk, R., Peute, J., and Vermeij, J. A. J. (1978). Cell Tissue Res. 186,309-325. van der Knaap, W. P. W., Sminia, T., and Edelenbosch, P. (1981). Dev. Comp. Immunol. 3, 37-44. van der Knaap, W. P. W., Doderer, A., Boemgter-Barendsen, L. H., and Sminia, T. (1982). Biol. Bull. (Woods Hole, Mass.) 162, 404-412. Vermeulen, A. (1977). I n “Androgens and Antiandrogens” (L. Martini and M. Motta, eds.), pp. 53-65. Raven, New York. Verselis, V., White, R. L., Sprag, D. C., and Bennett, M. V. (1986). Science 234,461-464. Viehberger, G. (1985). Fortschr. Zool. 30,453-455. Vignon, X., Terquem, A., and Dadoune, J. P. (1985). Biol. Cell. 53,48A. Vilar, 0 . (1973). I n “Human Reproduction” (E. S. E. Hafez and T. N. Evans, eds.), pp. 12-37. Harper & Row, New York. Vitale, R., Fawcett, D. W., and Dym, M. (1973). Anat. Rec. 176, 333-344. Vlok, W., and Van Vuren, J. H. J. (1988). Comp. Biochem. Physiol. A WA,49-51. Vogl, A. W., and Soucy, L. J. (1985). J . CellBiol. 100,814-825. Voglmayr, J. K. (1975). I n “Handbook of Physiology” (R.Greep, E. Astwood, D. Hamilton, and S. Geiger, eds.), Vol. V, pp. 437-451. Am. Physiol. SOC.,Washington, D.C. von Medem, F. (1945). Zool. Jahrb., Abt. A&. Zool. Physiol. Tiere 61, 1-44. Waites, G. M. H.,and Gladwell, R. T. (1982). Physiol. Rev. 62,624-671. Waites, G . M. H., and Neaves, W. B. (1977). I n “The Testis” (A. D. Johnson and W. R. Gomes, eds.), Vol. 5, pp. 91-157. Academic Press, New York. Waites, G. M. H., and Setchell, B. P. (1966). J . Endocrinol. 34,329-342. Walker, C. W. (1980). J . Morphol. 166,81-107. Wallace, R. A., and Selman, K. (1981). Am. Zool. 21,325-343. Wallace, D. M. A., Gunter, P. A., Landon, G. V., Pugh, R. C. B., and Hendry, W. F. (1982). Br. J . Urol. 54,765-768. Walsh, M. P. (1954). Trans. Am. Microsc. SOC.73,59-65. Wang, J. M., Qiu, J. P., Wu, X. L., Zhang, Z. R., Shao, Y. U., and Cao, R. Q. (1988). J . Androl. 9,397-402. Wang, J. M., Wen, G. Y.,Zhang, Z. R., Wu, X. L., Jiang, D. H., Tao, L., Cao, R. Q., and Zhou, Q. (1989). Contraception 39,569-575. Weber, J. E., Russell, L. D., Wong, V., and Peterson, R. N. (1983). A m . J . Anat. 167, 163-179. Weber, J. E., Turner, T. T., Tung, K. S. K., and Russell, L. D. (1988). A m . J . Anat. 182, 130- 147.

190

MORDECHAI ABRAHAM

Wentworth, B. C., and Mellen, W. J. (1964). J. Reprod. Fertil. 8,215-223. Whalen, R. E., and Luttge, W. G. (1969). Nature (London) 223,633-634. Willson, J. T., Jones, N. A., Katsh, S., and Smith, S. W. (1973). Anat. Rec. 176,85-100. Wong, V., and Russell, L. D. (1983). Am. J . Anar. 167, 143-161. Wong, P. Y. D., and Uchendu, C. N. (1990). J . Endocrinol. 125,457-465. Wong, P. Y. D., and Yeung, C. H. (1978). J . Physiol. (London) 275, 13-26. Wood, R. L. (1959). J. Eiophys. Biochem. Cytol. 6,343-352. Wood, R. L., and Kuda, A. M. (1980). J . Ultrastruct. Res. 70, 104-117. Worley, E. K., Franz, D. R., and Hendler, G. (1977). Eiol. Bull. (Woods Hole, Mass.) 153, 237-253. Wrobel, K. H., and Schimmel, M. (1989). Cell Tissue Res. 257,93-104. Yazama, F., Nishida, T., Kurohmaru, M., and Hayashi, Y. (1988). Jpn. J . Vet. Sci. 50, 1259-1261. Yokoyama, M., and Chang, J. F. (1971). J . Histochem. Cytochem. 19,766-774. York, J. P., and Drago, J. R. (1985). J . Urol. 133,294-297. Yule, T. D., Montoya, G. D., Russell, L. D., andTung, K. S. K. (1987a).J. CellEiol. 105, 168a. Yule, T., Shepherd, K., Russell, L. D., and Tung, K. S. K. (1987b). Ann. N . Y. Acad. Sci. 513,523-525. Yule, T . D., Montoya, G. D., Russell, L. D., Williams, T. M., and Tung, K. S. K. (1988). J . Immunol. 141, 1161-1 167. Zhiyuan, Z., Xiufen, W., Hanmin, L., Jie, Y.,Zhenshan, L., Guangrong, L., and Huaxun, R. (1985). Acta Acad. Med. Sichuan 16, 195-198. Zhuang, L. Z., Phillips, D. M., Gunsalus, G. L., Bardin, C. W., and Mather, J. P. (1983). J . Androl. 4,336-344. Zihler, J. (1972). Wilhelm Roux’s Arch. Dev. Eiol. 169,239-267.

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 130

Mitochondria-Rich Cells in the Gill Epithelium of Teleost Fishes: An Ultrastructural Approach M. PISAMAND A. RAMBOURG Service de Biologie Cellulaire, Dkpartment de Biologie Cellulaire et Molkculaire, Centre d’Etudes Nucleaires de Saclay, 91 191 Gif-sur-Yvette Cedex, France

I. Introduction Teleostean fishes maintain a constant concentration gradient between electrolytes of their internal medium and those of their external medium. Thus, in sea water, the teleost has to eliminate ions which are absorbed by ingestion (Smith, 1930; Krogh, 1939); in contrast, in fresh water, the fish must retain ingested ions and compensate for passive loss by active uptake (Maetz, 1974; Evans, 1984). Some teleosts may live exclusively in sea water (seawater stenohaline fish) or in fresh water (freshwater stenohaline fish), whereas others are able to adapt to both environments by keeping their internal medium fairly constant in spite of a wide variation in the osmolarity of the external environment (euryhaline fish). The ionic exchanges required for teleost osmoregulation are thought to be mainly located in the gill (Smith, 1932; Garcia-Romeu and Maetz, 1964; Maetz and Garcia-Romeu, 1964; Maetz, 1971; de Renzis and Maetz, 1973; Payan, 1978) and, more specifically, in the gill epithelium (Maetz, 1976; Evans, 1980; Girard and Payan, 1980). About 50 years ago, Keys and Willmer (1932) suggested that a certain type of gill epithelial cells might be responsible for chloride excretion in seawater eel and called it the “secreting chloride cell.” They described it as a large ovoid eosinophilic cell (average diameter of 15 pm) displaying access to both internal and external media. Then, Copeland (1948), presumably for the first time, gave it the name “chloride cells” and showed that, in fresh water- and sea water-adapted Fundulus hereroclitus, it was replete with mitochondria1 and osmiophilic materials (Copeland, 1950). The involvement of chloride cells in ionic exchanges when some species of fish were placed in a hyperosmotic environment has been supported by the following observations. 1 . Transfer from fresh water to sea water induced increases in the number (Olivereau, 1970; Shirai and Utida, 1970; Utida et al., 1971; 191

Copyright Q 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

192

M. PISAM AND A. RAMBOURG

Thomson and Sargent, 1977; Foskett et af., 1981) and size (Shirai and Utida, 1970; Coleman et af., 1977, Pisam, 1981; Pisam et af., 1987) of the chloride cells in the branchial epithelium. 2. In the opercular epithelium, which, in contrast to the branchial epithelium, may be easily dissected out and studied in vitro in an Ussing chamber, chloride cells, with the same ultrastructural features as those of branchial chloride cells, have been described in a few species by Karnaky and Kinter (1977), Marshall (1977), Marshall and Nishioka (1980), Foskett et af. (1981), and Zadunaisky (1984). When the short-circuit current technique was used in conjunction with the staining of chloride cells by DASPMI (dimethylaminostyrylmethylpyridiniumiodine,a specific fluorescent stain for mitochondria, which are numerous in these cells), there was a linear correlation between chloride cell density and the rate of chloride transport (Karnaky et al., 1979, 1984; Marshall and Nishioka, 1980; Foskett et af., 1981). In the opercular epithelium of the sea wateradapted fish, Tifapia,examined by the vibrating probe technique, Foskett and Scheffey (1982) observed that the current generated by the shortcircuited epithelium was directly over the chloride cells, demonstrating that chloride cells were the sites of active chloride secretion and high ionic permeability in seawater teleosts. In contrast, in a freshwater environment, the role of chloride cells in active branchial ionic absorption (Krogh, 1938, Evans, 1984) has not been as convincingly demonstrated. Yet, correlation between chloride cell number and the Ca2+or Na+ uptakes (Perry and Wood, 1985; Avella et al., 1987; Perry and Laurent, 1989) suggested that they are likely sites for such ion transport. The purpose of this chapter is to describe the ultrastructure of chloride cells in the gill epithelium of teleostean fishes and to show to what extent changes in water salinity may affect this structure in various species.

11. Gill Morphology

Gill morphology has been carefully described by Laurent and Dune1 (1980), Laurent (1984), and Hughes (1984). In teleosts, the gills consist of several branchial archs, beautifully illustrated in scanning electron micrographs (Olson and Fromm, 1973; Kendall and Dale, 1979; Jacobs et al., 1981). Each branchial arch bears several primary lamellae (or filaments), which, in turn, give rise to rows of secondary lamellae (Figs. 1 and 2). Each primary lamella is vascularized by arterioles and a central venous sinus. Two types of vascular anastomoses are observed: (1) arterioarterial

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

193

anastomoses (Figs. 2 and 3), in which an afferent and an efferent arteriole are connected by means of arterial pillar capillaries located within the core of the secondary lamellae, and (2) arteriovenous anastomoses (Fig. 3), in which the efferent arteriole is directly connected with the central venous sinus located in the core of the primary lamella (Laurent and Dunel, 1976; King and Hossler, 1986). Primary and secondary lamellae are covered by an epithelial sheet, which, as proposed by Laurent and Dunel (1978), is subdivided into two regions: a primary epithelium covering the primary lamellae and a secondary epithelium covering the secondary lamellae. In a section running parallel to the long axis of the primary lamella and thus transecting the secondary lamellae (Fig. 2), the primary epithelium fills the interlamellar space separating the bases of two consecutive secondary lamellae. According to fish species or size, this epithelium may be pseudostratified (Pisam et al., 1989)(e.g., Fig. 1) or pluristratified (Morgan and Tovell, 1973). Among the more superficial cells in contact with the outside medium, two types have been regularly identified: the flattened, sometimes piriform, pavement cells and the ovoid chloride cells. The latter are voluminous, mainly encountered in the interlamellar regions of the primary epithelium (Laurent and Dunel, 1980; Pisam et al., 1987; Wendelaar Bonga et al., 1990) and on the side of the primary lamella supplied with afferent blood (Copeland, 1948; Karnaky et al., 1976a; Laurent, 1984; Hossler et al., 1986). In the secondary epithelium, the presence of chloride cells is more erratic and may vary according to the fish species; thus, the chloride cells, which are absent from the secondary lamellae in the guppy (Pisam et al., 1987) are frequently observed in the goldfish (Kikuchi, 1977). It may also depend on the salinity of the external medium, as these cells are seen to proliferate when fishes from several species are adapted to demineralized water (Mattheij and Stroband, 1971; Laurent and Dunel, 1980; Laurent et al., 1985),whereas when the fish is transferred to sea water, they disappear (Pisam et al., 1988, 1989) or are scanty (Maina, 1990).

III. General Ultrastructural Features of Chloride Cells

A. THECELLSURFACE: APICALCAVITY AND BASOLATERAL TUBULAR INVAGINATIONS 1 . Apical Cavity As recently as 1948, Copeland, studying seawater-adapted fishes with the light microscope, observed at the apex of gill chloride cells a structure referred to as an “excretory vesicle.” Since such a vesicle was rarely

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

195

encountered in fresh water-adapted fishes, he postulated that its appearance correlated with the adaptation of the fish to high-salinity living conditions (Copeland, 1950). Electron microscopic studies revealed that the so-called excretory vesicle could be equated to an invagination of the apical membrane of the chloride cell (Figs. 28 and 30). Indeed, while, in fresh water-adapted fishes, the apical plasma membrane of chloride cells was only slightly depressed (Getman, 1950; Straus and Doyle, 1961; Straus, 1963; Coleman et al., 1977; Kikuchi, 1977; Lacy, 1983; Lubin et al., 1989; Maina, 1990), it was deeply invaginated in seawater-adapted fishes to form a deep narrow apical cavity which obviously corresponded to the excretory vesicle (Kessel and Beams, 1962; Philpott, 1962; Philpott and Copeland, 1963; Foskett et al., 1981). Moreover, in keeping with Copeland’s assumption, the depth of the apical cavities (Hossler et al., 1979; Lubin et al., 1989) and their frequency (Getman, 1950; Coleman et al., 1977; Hossler et al., 1985) increased with the salinity of the outside medium. Incidentally, a similar effect was also obtained in the yellow perch by Leino et al. (1987) under conditions of osmoregulatory stress induced by acidified soft water. A Na+, K+-activated ATPase has been cytochemically detected on the cytoplasmic side of the apical plasma membrane (Hootman and Philpott, 1979). An extraneous coat, sometimes filling the whole depth of the apical cavity, has also been shown to be loosely attached to its outer aspect (Figs. 11, 28, and 30). As demonstrated by histochemical (Philpott, 1968; Karnaky and Philpott, 1969; Petrik and Bucher, 1969; Pisam et al., 1980) and autoradiographic studies with [3H]glucosamine (Pisam et al., 1980) or [3H]fucose (Pisam et al., 1983), this extraneous coat, which consists of a finely granulated material, is rich in polysaccharides (Pisam et al., 1983) and traps chloride ions in the apical cavity (Copeland, 1948; Philpott, 1965). It might thus modify the composition of the immediate environment of the cell and thereby be involved in ion-exchange processes (Pisam et al., 1980).

FIG. 1. Cross section of a primary lamella (i.e., filament) of teleostean gill stained with Toluidine Blue (as seen in the light microscope). The afferent and efferent regions of the primary lamella (PL) contain the afferent (aa)and efferent (ea) arteries, respectively. The midregion bears secondary lamellae (SL). CVS, Central venous sinus; Ca, cartilage; C, chloride cell. x365. FIG. 2. Electron microscopic view of a longitudinal section through the midregion of a primary lamella stained by reduced osmium. The primary lamella (PL) is made up of a longitudinally oriented central venous sinus (CVS) covered by the primary epithelium containing “pavement” (p) and chloride (c) cells. The secondary lamellae (SL) consist of arterial pillar capillaries (PC) covered by the secondary epithelium. x 2200.

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

197

2. Tubular System From the outset of electron microscopic studies, it was realized that the cytoplasm of chloride cells contained a highly developed membranous system made up of anastomosed tubules. Yet, despite the fact that a continuity had sometimes been observed between some membranous tubules and the plasma membrane (Doyle and Gorecki, 1961; Straus and Doyle, 1961; Straus, 1963), the exact nature of this membranous system was far from being elucidated. Some authors equated it to the agranular reticulum (Kessel and Beams, 1962; Philpott, 1962; Philpott and Copeland, 1963). Others, more numerous, using extracellular space markers such as horseradish peroxidase (Philpott, 1966), lanthanum salts (Philpott, 1967; Ritch and Philpott, 1969), or Ruthenium Red (Komuro and Yamamoto, 1970), estimated that it was an extension of the basolateral plasma membrane (Philpott, 1964, 1968; Karnaky et al., 1976a; Kikuchi, 1977; Sargent et al., 1977; Sardet et al., 1979). Finally, the specific staining of intracytoplasmic membranes by the potassium ferrocyanidereduced osmium technique (Karnovsky, 1971) clearly demonstrated that the smooth tubular system filling most of the cytoplasm of the chloride cells actually consisted of two distinct membranous systems present side by side: a faintly stained endoplasmic reticulum continuous with the nuclear envelope and a densely stained tubular system in continuity with the laterobasal plasma membrane (Pisam, 1981) (Fig. 5). The same conclusion was drawn by Fujita and Yamamoto (1984) when they observed that a tannic acid-glutaraldehyde fixative heavily delineated the tubular system, while the endoplasmic reticulum remained utterly unlabeled. The tubular system extends throughout the whole cytoplasm, except for the Golgi area and a narrow band located just beneath the apical surface (Fig. 12). As a rule, it consists of tubules of constant diameter anastomosed in a network, the meshes of which enclose mitochondria (Figs. 8 and 12). Ritch and Philpott (1969) gave the first evidence that the membrane of the tubules contained repeating units. Adding lanthanum to the fixative, they

FIG. 3. Schematic representation of gill lamellae vasculature in teleost fish. From Pisam er al. (1987).

FIG. 4. Base of a secondary lamella (SL) and interlamellar region (IR) in the gill epithelium of a fresh water-adapted guppy; manganese-lead (Mn-Pb) staining. An a-chloride cell (pale cell), located at the base of the secondary lamella, is in close contact with the pillar capillary (PC). A /%chloridecell (dark cell) is observed in the interlamellar region; flattened sheets of its endoplasmic reticulum (ER) are found between the nucleus (n) and the basal surface of the cell. x7500. From Pisam et al. (1987).

ULTRASTRUCTURE O F MITOCHONDRIA-RICH GILL CELLS

199

filled the lumen of the tubules with opaque material and thus revealed, on the outer leaflet of the membrane enclosing the lumen, rows of 15-16 units regularly arranged around the periphery of the tubule. Along the tubular axis, these units were separated by a center-to-center 60 %, spacing. When the membrane of the tubules was examined in freeze-fracture by Sardet et al. (1979), the P (cytoplasmic) fracture face displayed a corncob appearance, as particles 70-75 %, in diameter repeated themselves along the tube length with a period of 80-100 A. On the E face, repeating lines with a period of 70-80 %, were not exactly perpendicular to the longitudinal axis of the tubules. Their tilting along the tubes, observed by several techniques, suggested instead that the arrangement of the repeating units along the tube was helicoidal. Kamiya (1972), followed by Sargent et al. (1975), and Hootman and Philpott (1978), reported that samples of isolated chloride cells were endowed with a high Na+, K+-ATPase activity. Thereafter, histochemical techniques (Hootman and Philpott, 1979) and autoradiographic studies investigating the fixation sites of [3H]ouabain(Karnaky et al., 1976b; Silva et al., 1977) have clearly localized the Na+, K+-ATPase activity on the tubular system. It was therefore postulated by Sardet et al. (1979) and by Philpott (1980) that the repeating units of the membrane of the tubules might represent pumping units in which the transport-associated ATPase, supplied with ATP by the numerous mitochondria located in the neighborhood, would be directly involved in fish osmoregulation.

FIGS.5-8. Portions of chloride cells in the guppy, showing the membranous tubules of the tubular system (T) densely stained by the reduced osmium technique m, Mitochondria; ER, endoplasmic reticulum. ~35,000.From Pisam er al. (1987). FIG. 5 . a-Chloride cell in a fresh water-adapted guppy. The tubular system forms a network with relatively wide polygonal meshes; it is in continuity with the laterobasal plasma membrane (LBM), which is in close contact with the basement membrane (BM) of a pillar capillary. At the arrowhead, an endoplasmic reticulum cisterna is seen to cross a mesh of the tubular system. FIGS. 6 and 7. P-Chloride cell in a fresh water-adapted guppy. In Fig. 6, the tubular system is unevenly distributed throughout the cytoplasm. It may be anastomosed to form small polygonal meshes (arrowheads). In Fig. 7, near the laterobasal plasma membrane (LBM), slightly wider membranous tubules form compact parallel arrays (white arrow). FIG. 8. Fully developed seawater (SW) a-chloride cell. The tubular system consists of an extensive network made up of numerous small and regular polygonal meshes (arrowheads). BM, Basement membrane of the pillar capillary.

200

M. PISAM AND A. RAMBOURG

B. THEGOLGIAPPARATUS AND VESICULOTUBULAR SYSTEM As shown in many cells (Rambourg and Clermont, 1990), the Golgi apparatus is a continuous ribbonlike structure which, in chloride cells, forms a supranuclear mass. Due to its numerous twists and anastomoses, this Golgi ribbon is sectioned at various angles, and thus, in thin sections (Fig. 9), it is usually seen as a series of independent stacks of saccules distributed throughout the supra- and juxtanuclear areas of the cytoplasm. In sections perpendicular to the surface of the ribbon, each stack of saccules is structurally polarized. One face of the stack consists of a series of circular, ovoid, or elongated profiles connected by lightly stained bridges; this is the cis element of the Golgi ribbon, which is exclusively composed of interconnected membranous tubules (Rambourg and Clermont, 1990) (Fig. 10). Subjacent to the cis element, two or three closely superposed and strictly parallel membranous elements are flattened poorly fenestrated saccules, making up the mid compartment. On the trans aspect of the stack and forming the trans compartment, several elements do not exactly follow the longitudinal axis of the Golgi ribbon; all of them show a central flattened saccular part and a peripheral network of anastomotic tubules that curve away from the overlying elements (Fig. 10); thus, they display a “peeling off” configuration. Intermingled with the trans elements of the Golgi apparatus and probably originating from them, numerous tubules and vesicles are rich in polysaccharides and are parts of the so-called vesiculotubular system. Indeed, although vesicles pertaining to this system had been observed in the apical cytoplasm of chloride cells by Doyle and Gorecki (1961), Straus and Doyle (1961), Threadgold and Houston (1961, 1964), Shirai and Utida (1970), and Kikuchi (1977), the vesiculotubular system was only recognized as a specific entity when the potassium ferrocyanide-reduced 0smium technique of Karnovsky (1971) was applied to fish gills (Pisam, 1981). It comprises vesicles and tubules with a content of intermediate

FIGS.9 and 10. Golgi region of a chloride cell from a sea water-adapted guppy. Reduced osmium staining. FIG.9. The perinuclear area shows cross and oblique sections through the stacks of saccules forming the Golgi apparatus (arrowheads). n, Nucleus; DB, dense bodies. X 16,000. FIG.10. A higher magnification of a stack of Golgi saccules as seen in cross section. Starting from cis face, two elements (1 and 2) are highly fenestrated and consist mainly of anastomosed tubules. The third saccular element (3) is perforated by a few pores. On the trans face, elements 4 and 5 show a central saccular portion and a tubular periphery. Element 5 is “peeling off” from the overlying stack (curved arrow). ~60,000.

202

M. PISAM AND A. RAMBOURG

density between that of the tubular system and that of the endosplasmic reticulum. Tubules and vesicles show a tendency to accumulate at the apex of the cell just beneath the cell surface (Fig. 12). It was thus proposed that apical vesicles could pinch off the extremities of the tubules of the tubular system (Bradley, 1981) and thereby be involved in ion transfer from the tubular system to the apical surface (Philpott and Copeland, 1963; Threadgold and Houston, 1964; Sardet, 1980). While this hypothesis cannot be ruled out altogether, it should be stressed that similar structures are encountered in the trans region of the Golgi apparatus as well as in the region between the Golgi area and the apical surface (Fig. 12). They contain polysaccharides (Pisam, 1981; Pisam et al., 1980,1983), which are also found at the cell surface and on the trans aspect of the Golgi apparatus. They are thus likely, as indicated by autoradiographic studies, to be involved in the transport of glycoproteins from the Golgi apparatus to the apical surface (Pisam et al., 1983).

c .

ENDOPLASMIC RETICULUM, MITOCHONDRIA,AND CYTOSKELETON

The endoplasmic reticulum is a continuous organelle distributed homogeneously throughout the whole cytoplasm of the chloride cell. It frequently intercrosses the tubular system (Figs. 5 and 6) and consists of flattened cisternae interconnected by membranous tubules (Pisam, 1981). The abundance of mitochondria was recognized fairly soon as one of the conspicuousfeatures of chloride cells. As first described by Karnaky et al. (1976a), they are large rod-shaped organelles about 0.35 pm in diameter and approximately 1.6 p m in length. They are closely associated with the tubular system and, as the latter, are found in most of the cytoplasm, except for the Golgi area and a thin apical zone (Fig. 12).

FIGS. 11 and 12. Portions of chloride cells from a sea water-adapted guppy. Reduced osmium staining. FIG. 11. The apical region of the cell contains two kinds of microfilaments: (1) fine filaments (arrowheads) forming a web located just beneath the apical plasma membrane (am) and (2) thicker filaments (mf) distributed throughout the cytoplasm or anchored on desmosomes (d). MT, Microtubules. ~35,000. FIG. 12. Numerous vesicles and tubules of the vesiculotubular system whose staining is intermediate between that of the tubular system (T)and the endoplasmic reticulum (ER) are present near the apical surface (as; short arrows), in the trans region of the Golgi apparatus (9; long arrows), and in the area located between the Golgi region and the apical cell surface (white arrows). X20,OOO. From Pisam (1981).

204

M. PISAM AND A. RAMBOURG

In chloride cells, microfilaments are mainly encountered in the apical region (Bradley, 1981). As pointed out by Pic and Lahitette (1981), they are of two kinds according to their diameter: Fine filaments 25-40 8,in diameter form a filamentous network located just beneath the apical plasma membrane (Fig. 11); this apical web, which may be of variable density, is prominent in the vicinity of apical junctions. Thicker filaments 70-100 A in diameter are either isolated or gathered in bundles distributed throughout the cytoplasm (Fig. 11); some bundles of these microfilaments are found to be anchored on desmosomes binding chloride cells with adjacent cells (Pic and Lahitette 1981) (Fig. 11). Microtubules about 125 8, in diameter are mainly oriented along a basoapical axis (King et al., 1989) (Fig. 11); they might play a role in maintaining cell polarity and promoting cell motility (Maetz and Pic, 1977). IV. Mitochondria-RichCells and Modifications of the Environment A.

(Y-AND @CHLORIDE CELLSI N

FRESH WATER-ADAPTED FISHES

It was long assumed, until recently, that only one type of chloride cells, the mitochondria-rich celljust described, was present in the gill epithelium of most fishes. It was this cell type that increased in size and underwent ultrastructural modifications when fishes were transferred from fresh water to sea water. As recently as 1961, Doyle and Gorecki noted that in the euryhaline killifish, Fundulus, electron-dense and light cells coexisted in the gill epithelium of all specimens regardless of the milieu to which they were adapted. They also reported that in freshwater specimens, the base of

FIGS. 13 and 14. Apical region of chloride cells (a and p) from a fresh water-adapted guppy. Manganese-lead staining. ~ 2 3 , 0 0 0 From . Pisam et al. (1987). FIG. 13. The a-cell spreads along the pillar capillary (PC) visible at the bottom. Its laterobasal plasma membrane (LBM) follows the outline of the basement membrane of the capillary; its flattened apical plasma membrane (am), endowed with a fuzzy coat (fc), is in contact with the external medium. The tubular system (T) consists of a network with relatively wide meshes. The endoplasmic reticulum (ER) is made up of interconnected dilated cisternae. A few elements of the vesiculotubular system (VT) are seen under the apical membrane. m, Mitochondria. FIG. 14. The apical membrane (am), covered with a fuzzy coat (fc), is relatively wavy. Some components of the vesiculotubular system (VT) and numerous darker membranebound bodies (MB) are located beneath the apical membrane and in regions between the Golgi apparatus (9) and the apex of the cell. The endoplasmic reticulum (ER) consists of flattened cisternae surrounding the nucleus (n) or forms a network that intertwines with tubular system (T). m, Mitochondria.

206

M. PlSAM AND A. RAMBOURG

the light cell was in close contact with the basement membrane of the adjacent blood vessel. In the guppy, Straus (1963) also observed light and dark cells, but indicated that in this species most cells were intermediate between the two extremes of cytoplasmic densities. Finally, according to Bierther (1970), two types of chloride cells could be distinguished according to their shape and location in the primary and secondary epithelia of the stickleback gill. In these studies (see also Kikuchi, 1981), characterization of cell types was impaired by the lack of selectivity of techniques contrasting thin sections prior to their examination with the electron microscope. The use of selective metallic impregnations such as the aforementioned reduced osmium technique of Karnovsky (1971) or the manganese-lead method introduced by Pisam et al. (1987) allowed a better preservation and staining of intracytoplasmic membranes and thus led to the clear identification of two types of chloride cells in the gill epithelium of a fresh water-adapted euryhaline fish, the guppy Lebistes reticulatus. One type of cell, the a-chloride cell (a-cell) was located at the base of the secondary lamellae of the gill, in close contact with the basement membrane of the pillar capillary and, thus, mainly related to the arterial blood. This pale elongated cell displayed a flat hardly invaginated apical surface. The nucleus was at the base of the cell. The mitochondria, the elements of the tubular system, and those of the endoplasmic reticulum were evenly distributed throughout the cytoplasm (Fig. 4), except for a narrow zone located just beneath the apical plasma membrane. The tubular system consisted of membranous tubules with a fairly constant diameter of 30 nm; they were anastomosed to form a regular network with relatively wide roughly polygonal meshes (Fig. 5 ) . The endoplasmic reticulum comprised interconnected flattened or dilated cisternae which frequently intertwined with the network made up by the tubular system (Figs. 5 and 13). The vesiculotubular system was poorly developed and restricted to a few tubules and vesicles mainly located below the apical plasma membrane (Fig. 13). The other cell, the p-chloride cell (P-cell), was darker, ovoid or cuboidal, and located in the interlamellar region of the primary epithelium facing the central venous sinus (Fig. 4). Its apical surface was more invaginated and wavy than that of the a-cell (Fig. 14). The nucleus was located at the base of the cell, and numerous mitochondria accumulated along its basolateral surface (Fig. 4).The endoplasmic reticulum and the tubular system differed from the equivalent systems of the a-cell. The tubular system was unevenly distributed throughout the cytoplasm. Its tubular elements, the caliber of which was more irregular than in the a-cell, showed various tridimensional configurations. In some regions, tubules with a diameter of

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

207

30-40 nm formed a tight network, whereas in other regions, wider tubules were more loosely anastomosed (Fig. 6). At the periphery of the cell, wider less regular tubular elements (between 50 and 70 nm in diameter) were continuous with the basolateral plasma membrane and formed parallel arrays (Fig. 7). The endoplasmic reticulum was more conspicuous than in the a-cell. Parallel flattened cisternae of rough endoplasmic reticulum were located in the perinuclear region (Figs. 4 and 14). They were in continuity with a network of rough- and smooth-surfaced cisternae that intertwined with the tubular system in regions where the tubular elements were tightly anastomosed. The vesiculotubular system consisted, as in the a-cell, of tubular and vesicular elements located in the apical zone. In the same area, there were, in addition, numerous membrane-bound bodies of various sizes and shapes that showed an electron-dense content after manganese-lead staining (Fig. 14). The nature of these bodies is thought to differ from that of the tubules and vesicles of the vesiculotubular system. A relationship to a physiological function, however, has not yet been elucidated. Two types of chloride cells strikingly similar to the a-and p-cells of the guppy were recently found in the gill epithelium of two freshwater stenohaline fishes: the loach Cobitis taenia and the gudgeon Gobio gobio (Pisam et al., 1990). In both species, a pale elongated cell resembling the a-cell was located at the base of the secondary lamellae in close contact with the arterioarterial pillar capillaries (Fig. 15). Darker ovoid cells were usually observed in the interlamellar region of the primary epithelium facing the central venous sinus. They were either isolated (Fig. 16) or grouped in multicellular complexes (Fig. 17) in which all cells were alike (Fig. 17) and linked together by deep narrow apical junctions (Fig. 19), similar to those encountered between chloride and pavement cells (Fig. 20). As in p-cells, the apical cytoplasm of dark cells was filled with numerous spheroidal membrane-bound bodies (Fig. 18), and elements of their tubular system were frequently arranged in parallel arrays (Fig. 21).

B. ULTRASTRUCTURAL FEATURES OF MITOCHONDRIA-RICH CELLS TRANSFER OF EURYHALINE FISHES FROM FRESH WATER TO SEAWATER

DURING

The transfer of euryhaline fishes from fresh water induces some important modifications of the chloride cells, which, as mentioned in Section I, were shown to increase in size and sometimes in number. As a result of the clear distinction between light a-cells and dark p-cells, it soon appeared

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

209

that the transformation of the gill epithelium in response to an increase in salinity was more elaborate than initially reported. Thus, as shown in the guppy by Pisam et al. (1987), three types of mitochondria-rich cells were involved in this process. Indeed, while the p-cells degenerated, there was a hypertrophy of the a-cells and the appearance of a new type of mitochondria-rich cells: the acessory cells closely related to the a-cells. 1. Degeneration of p-Cells

After 2 days in full-strength sea water, the p-cells increased in volume without a concomitant development of the various intracytoplasmic membranous systems. As a result, their cytoplasm was more electron lucent than that of the a-cells, and they appeared as large pale cells located in the interlamellar region of the primary epithelium (Fig. 22); the diameter of the meshes formed by the tubular system became larger, and the parallel elements observed at the periphery of the cell tended to be more irregular in caliber; they frequently formed masses of interwined, irregular, and dilated tubules (Figs. 22 and 23). After 4 days in sea water, the percentage of p-cells, which in fresh water approximated 33.3% of the total number of chloride cells, was reduced to 4.6%; they displayed a denser cytoplasm in which elements of the tubular system formed dense clusters (Fig. 23).

FIGS.15-21. Chloride cells in the gill epithelium of the loach. From Pisam et al. (1990). FIG. 15. An ovoid elongated light chloride cell (Ic) is located at the base of a secondary lamella in close contact with the pillar capillary (PC). n, Nucleus. ~ 2 8 0 0 . FIG. 16. An isolated square dark chloride cell (dc) may be seen between two pavement cells (p) in the primary epithelium of an intermellar region. n, Nucleus. ~ 2 8 0 0 . FIG. 17. Two dark chloride cells (dc) are adjacent to each other in an interlamellar space. Flattened cisternae of endoplasmic reticulum (ER) are seen in the perinuclear region. n, Nucleus. ~ 2 8 0 0 . FIG. 18. At left, the apical region of alight chloride cell (Ic) contains a few elements of the vesiculotubular system (VT) located below the wavy apical plasma membrane (am). At right, the apex of the dark chloride cell (dc) is filled with numerous large irregular membrane-bound bodies (MB) and displays a slightly indented apical membrane (am). T, Tubular system; m, mitochondria. x 10,000, FIGS. 19 and 20. In Fig. 19, the apical junction 6 , between parallel arrows) binding the apical portions of two adjacent chloride cells (c) is as narrow and deep as the junction between a pavement cell (p) and a chloride cell (c) shown in (Fig. 20). ~ 3 1 , 0 0 0 . FIG. 21. In the dark chloride cell (dc), the relatively wide elements of the tubular system (T) frequently form compact parallel arrays. m, Mitochondria. x9800.

210

M. PISAM AND A. RAMBOURG

After 8 days, p-cells were rarely found; they showed, then, characteristic signs of degeneration; their denser cytoplasm contained altered mitochondria as well as densely packed hardly identifiable membranous structures (Fig. 24). Most probably, the degeneration of p-cells had already been observed with the light microscope by Shirai and Utida (1970) in the gill epithelium of freshwater eels progressively adapted to sea water. Indeed, using an acid fuchsin stain for demonstrating the mitochondria of chloride cells with the light microscope, these authors reported the presence of two types of chloride cells in fresh water-adapted eels. One cell type strongly stained with fuchsin is likely to correspond to the p-cell. Another cell type was paler and larger and seems to be the equivalent of the a-cell. During the first 3 days of seawater adaptation, they noticed an increase in the number of the large pale cells, which then decreased to a number smaller than the initial number of pale cells in fresh water-adapted eels. This apparent increase in the number of large pale cells that they attributed to a transient increase of the number of a-cells was, instead, probably due to the initial transformation of the darker p-cells, which increased in volume and became paler than the a-cells. Wendelaar Bonga and van der Meij (1989) also reported a degeneration of chloride cells in the gills of the teleost Oreochromis mossambicus. Degenerating chloride cells, which were rarely encountered in fresh wateradapted fishes, were constantly observed 3 days after the transfer of these fishes into sea water. Under these conditions, the cytoplasm of the cells undergoing degeneration was electron transparent, while their tubular system became dilated. Later, there was a condensation of the cytoplasm which contained a pycnotic nucleus, swollen mitochondria, and occasion-

FIGS.22-24. Portions of chloride cells in a sea water-adapted guppy. Reduced osmium staining. From Pisam et al. (1987). FIG. 22. After 2 days in sea water, the P-chloride cell (p) appears to be paler than the typical seawater a-chloride cell (a) located near the pillar capillary (PC). The intensely stained tubular system (T) of the P-cell is irregularly distributed throughout the cytoplasm, and may contain masses of irregular dilated tubules (arrowheads). X8.500. FIG. 23. After 4 days in sea water, a portion of a p-chloride cell (p)only shows irregularly dilated elements of the tubular system (arrow) surrounding altered mitochondria (m). PC, Pillar capillary; a, a-chloride cell. x 17,000. FIG. 24. After 8 days in sea water, a P-chloride cell @) shows signs of degeneration; it contains altered mitochondria (m) and densely packed membranous componenfs (arrowheads). PC, Pillar capillary; a , a-chloride cells. ~ 6 5 0 0 .

212

M.PISAM AND A. RAMBOURG

ally disrupted nuclear and cellular membranes. Finally, the degenerating cells showed a tendency to be globular and were possibly engulfed by macrophages. It would thus seem that while most chloride cells are able to transform themselves into seawater chloride cells, other mitochondriarich cells, presumably p-cells, degenerate during the transfer of fresh water-adapted fish into sea water. 2. Hypertrophy of a-Cells In the guppy, the a-cells increased markedly in size during adaptation of the fishes to sea water. Their surface area, which averaged 75 pm2 in fresh water-adapted fishes, increased to -180 pm2 in sea water. They always remained in contact with the pillar capillaries of the secondary lamella (Figs. 8,25, and 26), but occasionally some enlarged a-cells were seen to occupy the whole interlamellar space (Fig. 27), thus facing the central venous sinus by their enlarged basal surface. As they increased in size, the a-cells became progressively darker, and in fully adapted seawater fishes, the a-cells appeared to be as dark as the p-cells in fresh water-adapted fishes. In the gill epithelium of eels transferred to sea water, the number of dark chloride cells increased during the first week of adaptation to reach a maximum during the second week; since the increase in the number of the dark cells paralleled the decrease in the number of paler cells, Shirai and Utida (1970) postulated that the increase in the number of the former might

FIG. 25. An a-chloride cell in a fresh water (FW)-adapted guppy (reduced osmium staining). The cell is located at the base of the secondary lamella in close contact with the pillar capillary (PC).n, Nucleus. X4500. From Pisam et al. (1987). FIGS. 26-28. a-Chloride cells and accessory cells in sea water (SW)-adapted guppies (reduced osmium staining). FIG. 26. The a-chloride cell located next to the pillar capillary (PC) has greatly increased in volume to become a seawater a-chloride cell. Its apical membrane (am) is deeply invaginated. A, Process of an accessory cell; n, nucleus. X4500. From Pisam et al. (1987). FIG. 27. After days in full-strength sea water, a-chloride cells are frequently seen to occupy the whole interlamellar space between two pillar capillaries (PC). A, Process of an accessory cell. X4000. From Pisam et al. (1987). FIG..28. An accessory cell (A) is seen next to a chloride cell (C). In the accessory cell, the stained tubular system (T) appears less developed and more loosely anastomosed than in the chloride cell, where it forms a tight meshwork, in the meshes of which are found numerous mitochondria (m). Numerous elements of the vesiculotubular system (VT)are present near the apical surface of the chloride cell; they are rare at the apex of the accessory cell. The extraneous coat (ec) in the apical cavity of the chloride cell appears as a dense orderly feltwork. n, Nucleus; ER, endoplasmic reticulum: g, Golgi apparatus; as, apical surface. x 15,000. From Pisam (1981).

214

M. PISAM AND A. RAMBOURG

be the result of the transformation of the latter. Similarly, Coleman et al. (1977) reported that dark chloride cells were more common in sea wateradapted than in fresh water-adapted Tilapia aurea. They did not, however, relate this increase in number to the transformation of a paler cell. The cytoplasmic features of seawater chloride cells have been described repeatedly in various species (Philpott and Copeland, 1963; Bierther, 1970; Shirai and Utida, 1970; Doyle and Epstein, 1972; Karnaky et al., 1976a; Sardet et al., 1979; Philpott, 1980; Karnaky, 1980; Pisam, 1981; Laurent, 1984; Hwang, 1987; King et al., 1989); they are, thus, illustrated briefly here as exemplified in the guppy (Pisam et al., 1987). In this species, the seawater chloride cells contained numerous mitochondria. The extensive tubular system formed a tight network with numerous small polygonal meshes (Figs. 8,22, and 28), but, as also noted by Karnaky et al. (1976a) in Cyprinodon variegatus, the diameter of the tubules was not modified. This proliferation of the tubular system has usually been correlated with the increase in Na+, K+-ATPase activity detected in gill filaments of sea water-adapted fishes by Utida et al. (1971), Karnaky et al. (1976a), and Hossler (1980), and with the proliferation of mitochondria (Getman, 1950; Shirai and Utida, 1970; Coleman et al., 1977; Hwang, 1987; King et al., 1989) which are assumed to provide energy for the Na+ pump. The endoplasmic reticulum developed in a network of anastomosed smooth flattened sheetlike cisternae tightly interdigitated with the tubular system (Figs. 12 and 28). The vesiculotubular system also became conspicuous and consisted of an increased number of tubules and vesicles, the diameter of which was greater than that of similar elements in fresh water-adapted a-cells (Figs. 12 and 28). 3. Appearance of Accessory Cells As already stressed by Dunel and Laurent (1973) and as shown by Sardet et al. (1979), Dunel-Erb and Laurent (1980), Hootman and Philpott (1980), Laurent and Dunel (1980), Pisam et al. (1980), Lacy (1983), and Chretien and Pisam (1986), one most intriguing ultrastructural change induced by the transfer of a fresh water-adapted fish into sea water was the development of the so-called accessory cells next to, and in close association with, the apical portion of seawater chloride cells. In the guppy, these cells, sometimes called adjacent cells (Sardet et al., 1979), were pear shaped or semilunar and exhibited, although to a lesser extent, the ultrastructural features of chloride cells. Thus, the mitochondria1crests were smaller and less numerous, the tubular system more loosely anastomosed, and the vesiculotubular system less conspicuous, with fewer tubules and vesicles

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

215

(Fig. 28). The apical surface (straight or slightly convex) was covered with a thin layer of filamentous material, which, in contrast to the material filling up the apical cavity of the chloride cell, seemed to be tightly bound to the apical plasma membrane (Pisam et al., 1980). A striking ultrastructural feature of accessory cells was the presence of lateral cytoplasmic processes that penetrated the apical portion of the chloride cell, formed with the latter numerous plasma membrane interdigitations, and finally reached the outside medium filling the apical cavity (Fig. 26). As a result, the apices of the seawater chloride cells formed with their associated accessory cells a mosaic of cell apices, making up the wall of the apical cavity. This mosaic of cells was linked to surrounding pavement cells by deep apical junctions which appeared multistranded when examined by freeze-fracture microscopy (Sardet et al., 1979). In contrast, chloride cells and accessory cells were linked by shallow apical junctions which were permeable to lanthanum (Sardet et al., 1979); when freezefractured they only displayed one (Sardet et al., 1979) or a few strands (Kawahara et al., 1982). Similarjunctions were observed in the low-resistance chloride-secreting opercular epithelium of the sea water-adapted killifish (Ernst et al., 1980), in the electrolyte-secreting epithelium of the avian salt gland (Ellis et al., 1977; Riddle and Ernst, 1979), and in the salt-secreting epithelium of elasmobranch rectal glands (Ernst et al., 1979). In all these cases, as in the gill epithelium, the apical surfaces of adjacent electrolyte-secreting cells were narrow and highly interdigitated. As in the gill epithelium, the shallow and presumably leaky apical junctions exhibited a marked tortuosity. It was thus proposed that they could provide a paracellular pathway for ions secreted in sea water (Sardet et al., 1979; Ernst et al., 1980; Forrest et al., 1982; Foskett et al., 1983). It had been postulated by Sardet et al. (1979), Hootman and Philpott (1980), and Wendelaar Bonga et al. (1990) that accessory cells, with their small size and less developed intracytoplasmic membranous systems, might, in fact, correspond to young chloride cells. This view, however, was not supported by Dunel and Laurent (1973) or by Laurent and Dunel (19801, who claimed that the accessory cell was a new cell type characteristic of seawater or sea water-adapted fishes. Consequently, cell renewal and differentiation in the gill epithelium were studied by autoradiography at various time intervals after [3H]thymidine injection into fresh water- or sea water-adapted guppies (Chretien and Pisam, 1986); it was then found that chloride cells and accessory cells had different origins and modes of differentiation. In agreement with some observations by Conte and Lin (1967) and by Shirai and Utida (1970), chloride cells, in both fresh water-

216

M. PISAM AND A. RAMBOURG

and sea water-adapted fishes, originated from undifferentiated cells in contact with the basement membrane; they never left this contact as they increased in volume to reach the free surface of the primary epithelium. In contrast, accessory cells in sea water-adapted fishes originated from undifferentiated cells located in the intermediate layers of the primary epithelium in contact with mature chloride cells; they were in close relationship with the apical portion of the chloride cell and never reached the basement membrane. The first sign of their differentiation was the appearance of a rudimentary tubular system, which always arose from the lateral surface adjacent to the chloride cell. It was thus concluded that the accessory cell was indeed a new cell type, the differentiation of which was seemingly induced by mature chloride cells (Chretien and Pisam, 1986). CELLSIN STENOHALINE C. CHLORIDE CELLSAND ACCESSORY SEAWATER FISHES

As expected from what has just been described in the sea water-adapted guppy, the gill epithelium of stenohaline seawater fishes contains only one type of chloride cell: the sea water-type chloride cell. This chloride cell, however, as just mentioned, is always associated with accessory cells (Dunel-Erb and Laurent, 1980). In the turbot (Scophthalmus maximus) (Pisam et al., 1990),the chloride cells were exclusively encountered in the primary epithelium of the interlamellar space. Like the a-cells of freshwater or fresh water-adapted fishes, they were located at the base of the secondary lamellae and thus related to the arterial blood of the pillar capillary. Like the seawater cells of sea water-adapted euryhaline fishes, they were voluminous cuboidal cells that sometimes filled most of the interlamellar space and thus were also related to the venous sinus (Fig. 29). Their tubular system consisted of slightly distended membranous tubules tightly anastomosed to form an extensive network, with small regular polygonal meshes frequently penetrated by cisternae of the endoplasmic reticulum. Numerous mitochondria were uniformly distributed throughout the cytoplasm, except for a narrow apical zone filled with the numerous tubules and vesicles of the vesiculotubular system (Figs. 30 and 31). The apical surface was deeply invaginated in an apical cavity containing a filamentous feltwork (Fig. 30). The accessory cells were small mitochondria-rich cells appended to the apical portion of the chloride cells (Figs 29 and 30) and linked to them by a shallow apicaljunction (Fig. 3 1). As first observed in the sole by Dunel-Erb and Laurent (1980),their cytoplasm resembled that of the chloride cell and showed a similar density. It contained an extensive tubular network, with

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

2 17

small regular polygonal meshes frequently intercrossed by cisternae of the endoplasmic reticulum. A conspicuous difference between the two cytoplasms, however, was the apical zone, which in accessory cells was almost devoid of vesiculotubular elements (Figs. 30 and 31). The accessory cells sent numerous cytoplasmic processes that penetrated and interdigitated with the apical portion of the chloride cell before reaching the outer medium at the bottom of its apical cavity (Fig. 30). The numerous interdigitations between the two cytoplasms increased the length of the intercellular space, which, in front views, displayed a marked tortuosity. There was a concomitant amplificationof the length of the shallowjunction at the apical part of the intercellular space, thereby providing the fish with an expanded paracellular pathway to excrete ions into a hyperosmotic medium.

D. CHLORIDE CELLSAND ACCESSORY CELLSDURING TRANSFER OF SEAWATER-ADAPTED FISHES TO FRESHWATER Relatively few studies have been dedicated to the morphological changes induced in chloride cells by the transfer of sea water-adapted fishes into fresh water. Indeed, Hossler et al. (1985) pointed out that, under these conditions and in contrast to what is commonly observed when fishes are adapted to sea water, morphological and even biochemical modifications might lag far behind physiological responses. Yet a striking ultrastructural alteration, occurring within hours after transfer into diluted sea water (Newstead, 1971), is the disappearance of the apical cavity (Philpott and Copeland, 1963; Shirai and Utida, 1970; Doyle and Epstein, 1972; Hossler, 1980; Maina, 1990; Pisam et al., 1990), usually accompanied by a loosening of the tubular system and occasionally by a proliferation of the rough endoplasmic reticulum (Shirai and Utida, 1970; Maina, 1990). Thus, when the intertidal sculpin Oligocottus maculosus was briefly exposed to sea water diluted 1/100 with glass-distilled water (Newstead, 1971), mitochondria and membranous tubules were reorganized in parallel arrays preferentially located at the periphery of the chloride cells; the closely packed mitochondria in these arrays were separated from each other by a single layer of straight unbranched tubules parallel to the long axis of the mitochondria. The hyperosmotic adapted teleost Oreochromis a!caficus studied by Maina (1990) did not seem to tolerate adaptation to diluted sea water; there was an initial increase in the diameter and the degree of interdigitation of the membranous tubules, followed by a gradual decrease and the appearance of signs of intracellular degeneration.

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

2 19

In the stenohaline seawater turbot S. maximus, which, at a juvenile stage, may be encountered in the diluted sea water of estuaries, the situation was more complex (Pisam er al., 1990). In fishes adapted to 5%0 salt water, the overall appearance of the cytoplasm of the chloride cells remained unaltered, and the only modification was the disappearance of the apical cavity. The apical surface was no longer depressed, but sent instead numerous protrusions into the external medium (Fig. 32). The accessory cells were still present and bound to the apical portions of chloride cells by shallow junctions. However, the number of cytoplasmic interdigitations between the two cell types was reduced. As a result, the length of the junction sealing off the apical portion of a poorly convoluted intercellular space was significantly decreased, thereby preventing undue ionic excretion in the gills of a fish placed in a hypoosmotic medium. Transfer of the fish into fresh water induced further morphological changes. Both chloride and accessory cells increased in size but decreased in number. In chloride cells, the flattened apical surface was partially or completely separated from the outside medium by accessory cells (Fig. 33). The membranous elements of the tubular system were narrower and more loosely interconnected. The mitochondria had a tendency to accumulate in the perinuclear region, while some regions of the cytoplasm were completely devoid of organelles. The accessory cells sent only rare interdigitations into chloride cells. These interdigitations never reached the apical surface, and apical junctions were difficult to find. According to Pisam et al. (1990), these ultrastructural modifications reflected the difficulty, or perhaps the impossibility, for this seawater stenohaline fish to face such unphysiological

FIGS. 29-31. Chloride and accessory cells in a seawater turbot (reduced osmium staining) From Pisam et al. (1990). FIG. 29. Two voluminous chloride cells (C) are located on either side and at the base of a secondary lamella (SL). Their basolateral membrane is in contact with the basal lamina (bl) of the primary epithelium. PC, Pillar capillary; P, pavement cell; A, accessory cell. x3300. FIG. 30. An accessory cell (A) sends an interdigitation (arrow) into the apical portion of the chloride cell (C). It reaches the outside medium and thereby forms a part of the wall of the apical cavity, which is filled with a filamentous feltwork (f). In contrast to the accessory cell, the chloride cell displays in its apical region numerous elements of the vesiculotubular system (VT). In both cells, the membranous tubules of the tubular system (T) are intermingled with cisternae of the endoplasmic reticulum (ER). ~ 2 1 , 0 0 0 . FIG. 31. In seawater, the apical junction between the accessory cell (A) and the chloride cell (C) (arrowhead) is shallower than the one between pavement (P) and accessory cells (arrow). Note the numerous vesiculotubular elements (VT) in the chloride cell. X41,OOO.

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

22 1

conditions. Particularly, the loss of any contact between chloride cells and the outside medium and the concomitant disappearance of apical junctions were interpreted by these authors as indicating the shut-off of all excretory pathways to prevent any further ionic leakage in such a hypoosmotic environment. V. Mitochondria-Rich Cells and Smoltification in Salmonids

So far, it has been shown that the ultrastructural modifications of the gill epithelium in teleosts were mainly dependent on the salinity of the outside medium. Thus, while two types of chloride cells were commonly encountered in fresh water-adapted fishes, there was only one cell type, the so-called a-cell, in seawater fishes. Furthermore, accessory cells appended to the apical portion of transformed a-cells were exclusively observed in the latter animals. This generally accepted rule, however, did not seem to apply to all cases and, particularly, did not apply to salmonids which did or did not undergo smoltification. Thus, the rainbow trout does not undergo smoltification, and fishes of small size may hardly be adapted to sea water (Johnson and Cheverie, 1985). Yet, when the gill epithelia of these fishes were examined by electron microscopy, chloride cells with basolateral membrane invaginations partly consisting of flattened unfenestrated cisternae were always associated with smaller denser mitochondria-rich cells with the following ultrastructural characteristics: they were endowed with an extensive tubular system uniformly distributed throughout the cell (Fig. 36), and they were always adjacent to the apical portion of the chloride cells to which they were bound by junctions significantly shallower (Fig. 35) than those bind-

FIGS. 32 and 33. Apical portions of chloride and accessory cells in the gill epithelium of turbots adapted to diluted salt water and fresh water (reduced osmium staining). X22,OOO. From Pisam et al. (1990). FIG. 32. In 5%0 salt water (SW), the apex of the chloride cell (C) protrudes into the outside medium and contains elements of the vesiculotubular system (VT). The apex of the accessory cell (A), at left, is deprived of vesiculotubular elements and contains flattened cisternae of endoplasmic reticulum (ER). In both cell types, the tubular system (T) i s still prominent and consists of anastomosed membranous tubules. FIG. 33. In fresh water (FW), an accessory cell (A) covers a chloride cell (C) and separates it from the outside medium; its cytoplasm is denser than that of the chloride cell. In the chloride cell, the loosely anastomosed and narrow membranous tubules of the tubular system (T) stand out sharply against the pale background.

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

223

ing the apex of chloride cells to adjacent pavement cells (Fig. 34); they were therefore tentatively equated with accessory cells. Indeed, when the trout was progressively adapted to artificial sea water (Pisam et al., 1989), these small mitochondria-rich cells were found to behave like the accessory cells in sea water-adapted euryhaline fishes. They sent numerous cytoplasmic interdigitations within the apical portion of the chloride cell, while there was a further shallowing of the junction, sealing off the thereby expanded intercellular space (Fig. 37). The Atlantic salmon (Salrno salar) during its early life as a parr lives in fresh water. Then, in contrast to the rainbow trout, it undergoes a series of morphological, physiological, and biochemical transformations known as smoltification to reach a stage referred to as “freshwater smolt,” in which the fish is ready to pass into sea water to become a “seawater smolt.” When the gill epithelium of the Atlantic salmon was examined at all these stages, the gill epithelium of the parr contained chloride cells (Fig. 38) with a tubular system made up of loosely anastomosed tubules, as is the case in fresh water-adapted fishes (Fig. 40); these chloride cells, however, were frequently associated with smaller cells that resembled those observed in the rainbow trout and were thus also equated with accessory cells (Pisam et al., 1991). In freshwater smolts, the chloride cells almost doubled in size (Fig. 39), while their tubular system developed extensively to form a tight network, with regular meshes distinctly smaller than those encountered in parr chloride cells (Fig. 41). Half of the accessory cells appended to these enlarged chloride cells did not reach the outside medium, yet they were

FIGS. 34 and 35. Apical portions of the intercellular spaces in freshwater trout gill epithelium, stained by the reduced osmium technique. ~30,000.From Pisam et al. (1989). The staining of the intercellular space (is)disappears at the level of the apical junction (j, located between two parallel arrows), binding the apical portion of the chloride cell (C) to a pavement (P) cell (Fig. 34). A significantly shallower junction may be observed between an accessory cell (A) and the chloride cell (C) (Fig. 35). am, Apical membrane. FIGS. 36 and 37. Apex of chloride and accessory cells in gill epithelium of trout. x 18,000. From Pisam et al. (1989). FIG. 36. In freshwater trout, the tubular system (T) of the chloride cell (C) consists of sheets and tubules loosely anastomosed. The accessory cell (A) displays a tubular system made up of tightly anastomosed tubules. am, Apical membrane; FW, fresh water. FIG. 37. In seawater trout, the tubular system (T) of the chloride cell (C) appears as a dense network of anastomosed tubules around numerous small polygonal meshes. The accessory cell (A) sends numerous interdigitations (curved black arrows) into the apex of the chloride cell, am, Apical membrane. (Inset) The short apical junctions (arrows) located between chloride and accessory cells. SW, Salt water. x40,000.

ULTRASTRUCTURE O F MITOCHONDRIA-RICH GILL CELLS

225

bound to the latter by shallowjunctions. Transfer of freshwater smolts into sea water increased the number of accessory cells, all of them now being in contact with the outside medium; extensive plasma membrane interdigitations appeared between them and the chloride cells (Fig. 42), thereby producing an amplification of the length of the shallow junction at the apical parts of the intercellular space. It was thus clear that, in contrast to most euryhaline fishes, the Atlantic salmon (Pisam et al., 1988) and, to a lesser extent, the rainbow trout may, in fresh water, promote most of the ultrastructural transformations required for survival in a hyperosmotic environment. Indeed, the appearance of accessory cells in both species as well as the enlargement of chloride cells with an extensive tubular system in freshwater smolts of the Atlantic salmon were not triggered by sea water, but might instead result from a complex synergy between external (e.g., temperature, photoperiod, and trophic capacity of the external medium) and internal (e.g., hormonal and nervous) factors (Saunders and Henderson, 1978; Fontaine, 1975; Hoar, 1976; Komourdjian et af., 1976b; Wedeweyer et af., 1980; Folmar and Dickhoff, 1980; McCormick and Saunders, 1987).

M. Hormones and Chloride Cells In the previous section we stated that the nature of the stimuli which might induce morphological changes in chloride cells is still largely unknown. Nevertheless, among these, slowly acting hormones have been repeatedly suspected to play a major role in fish adapatation to sea water or fresh water (Lahlou, 1980;Foskett et al., 1983;Mayer-Gostan et al., 1987).

FIGS.38 and 39. Chloride cells (C) located at the base of secondary lamellae (SL) in Atlantic salmon (reduced osmium staining). x2500. From Pisam er al. (1988). In freshwater smolt (Fig. 39) the chloride cell is more voluminous than in parr (Fig. 38) and is accompanied by an accessory cell (A). FIGS.40 and 41. Portions of chloride cells in which the tubular system is selectively impregnated by reduced osmium. ~22,500.From Pisam er al. (1988). In the parr (Fig. 40). the tubular system (arrows) forms a loose network, with wide polygonal irregular meshes. In smolt kept in fresh water (Fig. 41), the tubular system (arrows) is more tightly anastomosed. ER, Endoplasmic reticulum; m, mitochondria. FIG. 42. In the seawater smolt, the accessory cell (A) sends apical interdigitations in the apex of the chloride cell ( C ) .The reduced osmium staining of the intercellular space separating the chloride cell from accessory cell may be followed almost up to the apical surface of the gill epithelium (arrows); it is absent at the level of a deep narrow junction located between the accessory cell and an adjacent pavement cell (P). T, Tubular system. x 19,000. From Pisam et al. (1988).

226

M. PISAM AND A. RAMBOURG

When freshwater eels were injected for 2 weeks with a high dose of cortisol, their gill epithelia contained an increased number of chloride cells, with hyperdevelopment of free ribosomes, enlarged mitochondria, and exceptional branching of the tubular reticulum; these modifications were associated with high levels of gill Na+, K+-ATPase (Doyle and Epstein, 1972), an enzyme known to be located in membranes of the tubular system. Thus, since cortisol induced changes specific for seawater adaptation, it was usually considered the specific hormonal mediator for the latter. Indeed, a long-term treatment of freshwater eels with cortisol (Forrest et al., 1973) markedly accelerated their seawater adaptation. Yet, there was, on the one hand, an initial lag in sodium outflux at the time of transfer, despite a high level of gill Na+, K+-ATPase; on the other hand, eels receiving cortisol for only 2 days had a significant increase in sodium outflux, although there was no detectable increase in gill Na+, K+-ATPase activity (Forrest et al., 1973). In the first case, the delayed salt excretion might be attributed to a lack of exposure of chloride cells to sea water (Doyle and Epstein, 1972; Foskett et al., 1981) and, consequently, to the lack of development of the shallow apical junctions; these observations, nevertheless, clearly indicate that other factors besides cortisol are involved in the complete differentiation of functional seawater chloride cells (Foskett et al., 1983). Cortisol has also been reported to be a sea water-adapting hormone during smoltification and seawater adaptation of anadromous salmonids, but in these species cortisol failed to restore Na', K+-ATPase activity and to promote ion regulation in hypophysectomized fishes adapted to diluted sea water (Bjornsson et al., 1987; Richman et al., 1987). Recent studies have thus focused on growth-independent effects of growth hormone on seawater adaptation. The results indicate that, indeed, growth hormone might play a role of prime importance in parr-smolt transformation and seawater dapatation (Komourdjian et al., 1976a; Prunet et al., 1989; Boeuf et al., 1989, 1990). In fact, when freshwater yearlings of the sea trout Salmo trutta were injected with growth hormone, cortisol, or both hormones, gill Na+, K+-ATPase, gill interlamellar chloride cell density, and chloride cell apical-to-basal length increased by all hormone treatments, but most significantly by combined growth hormone-cortisol treatment. Furthermore, the growth hormone-cortisol-treated fishes showed only insignificant changes in ion osmotic homeostasis when transferred into sea water, suggesting a synergistic effect of the two hormones (Madsen, 1990). Curiously enough, cortisol, a presumed sea water-adapting hormone, seemed to also play a role in fish adaptation to diluted environments

ULTRASTRUCTURE OF MITOCHONDRIA-RICHGILL CELLS

227

(Laurent and Perry, 1990). A daily injection of cortisol for 10 days in the freshwater rainbow trout Salmo gairdneri increased whole-body influxes of sodium and chloride ions and, concomitantly, induced a significant increase in the number and individual apical area of gill chloride cells per square millimeter of filament epithelium. When examined with the electron microscope, the gill chloride cells displayed the same ultrastructural features as those of a trout transferred into ion-deficient water; they developed at their apical surface numerous microridges (microplicae),thereby increasing the surface area of their apical membrane, which, in control freshwater fishes, formed only irregular short microvilli. The linear increase of sodium or chloride influxes as a function of chloride cell apical surface area strongly suggests that they are dependent on the availability of apical membrane transport sites. Thus, according to Laurent and Perry (1990), an increased need of transport sites in cortisol-injected trout or fishes living in ion-deficient water is compensated by the addition to the apical plasma membrane of an enlarged number of coated vesicles, presumably originating from the Golgi complex. Finally, it has been shown that a major hormonal modification following seawater transfer of teleosts was a drastic reduction in prolactin secretion (Dharmamba and Nishioka, 1968; Nicoll et al., 1981; Prunet et al., 1985; Hasegawa et a f . ,1987). Further electrophysiologicalstudies demonstrated that several prolactin injections into sea water-adapted fishes induced a significant inhibition of the current generated by the isolated opercular membrane (Foskett et a f . , 1982). According to Foskett et a f . (1983), this effect might be operated by blocking differentiation of new chloride cells or by causing dedifferentiation of the existing cells. Yet, at the present time, ultrastructural proof supporting such an interpretation is still to be provided.

VII. Concluding Remarks During the last 10 years, ultrastructural studies have emphasized the heterogeneity of the mitochondria-rich cell population in the gill epithelium of teleosts. Studies of sea water-adapted fishes have stressed the importance of a close association between seawater chloride cells and accessory cells for a good adaptation to seawater life. As proposed by Sardet et al. (1979), the increase in length of the intercellular space between these two cell types and the concomitant amplification of its apical shallow junction would provide the fish with an expanded paracellular pathway to excrete ions into a hyperosmotic medium. The enlarging of

228

M. PISAM AND A. RAMBOURG

chloride cells, the appearance of accessory cells, and the amplification of the space between the two cell types might result from a complex poorly understood synergy between environmental and internal factors. During smoltification,only the last modifications,namely, a further increase in the length of the shallow apical junction, seem to be triggered by an external factor: the contact of the apical surface of the cells with sea water. Among internal factors, cortisol and growth hormone are usually considered sea water-adapting hormones, but mechanisms by which they may influence an ultrastructural remodeling of chloride cells remain to be established. In contrast to what occurs in seawater fishes, the analysis of the ultrastructural features of mitochondria-rich cells in fresh water-adapted fishes may be somewhat confusing, since, as pointed out by Dejours (1988), freshwater environments are characterized by highly variable chemical properties, including pH as well as ionic or gaseous composition. Yet, studies of the gill epithelium of freshwater teleosts have revealed the existence of at least two types of chloride cells. The a-cells, located at the base of the secondary lamellae, are closely related to the arterioarterial pillar capillaries; they give rise to seawater chloride cells when euryhaline fishes are transferred into sea water. The p-cells, facing the central venous sinus, usually degenerate in a seawater environment; their function in fresh water is unknown at the present time. ACKNOWLEDGMENTS We are grateful to Mrs. Claudine Juillard for her secretarial work and Miss Corinne Le Moal for her skillful technical assistance. We also wish to acknowledge the useful advice of Drs. M. Chretien, N. Mayer-Gostan, P. Pic, and P. Prunet, who read the manuscript.

REFERENCES Avella, M., Masoni, A., Bornancin, M., and Mayer-Gostan, N. (1987). J . Exp. Zool. 241, 159-169.

Bierther, M. (1970). Z. ZeNforsch. 107,421-446. Bjornsson, B., Yamauchi, K., Nishioka, R. S., Deftos, L. J., and Bern, H. A. (1987). Gen Comp. Endocrinol. 68,421-430. Boeuf, G., Le Bail, P. Y., and Prunet, P. (1989). Aquaculture 82,257-268. Boeuf,G., Prunet, P.,andLeBail, P. Y. (1990). C. R. Hebd. SeancesAcad. Sci., Ser. 111310, 75-80.

Bradley, T.J. (1981). J . Exp. Zool. 217, 185-198. Chretien, M., and Pisam, M. (1986). Biol. Cell. 56, 137-150. Coleman, R., Yaron, Z., and Ilan, Z. (1977). J. Fish Biol. 11,589-594. Conte, F. P., and Lin, D. H. Y. (1967). Comp. Biochem. Physiol. 23,945-951.

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

229

Copeland, D. E. (1948).J . Morphol. 82,201-228. Copeland, D. E. (1950). J. Morphol. 87,369-379. Dejours, P. (1988). “Respiration in Water and Air: Adaptation-Regulation-Evolution.” Elsevier, Amsterdam. de Renzis, G., and Maetz, J. (1973). J. Exp. Biol.59,339-358. Dharmamba, M.,and Nishioka, R. S. (1968). Gen. Comp. Endocrinol. 10,409-420. Doyle, W . L., and Epstein, F. H. (1972). Cyrobiologie 6,58-73. Doyle, W. L., and Gorecki, D. (1961). Physiol. Zool. 34,81-85. Dunel, S., and Laurent, P. (1973). J. Microsc. (Oxford) 16,53-57. Dunel-Erb, S . , and Laurent, P. (1980). J. Morphol. 165, 175-186. Ellis, R. A., Goertemiller, C. C., Jr., and Stetson, D. L. (1977). Nature (London) 268, 555-556. Ernst, S. A., Hootman, S. R., Schreiber, J. H., and Riddle, C. V. (1979). J. CellBiol. 83,83a. Ernst, S . A., Dodson, W. C., and Karnaky, K. J. (1980). J. Cell Biol. 87,488-497. Evans, D. H. (1980). A m . J. Physiol. 238, R224-R230. Evans, D. H. (1984). In “Fish Physiology” (W. S. Hoar and D. J. Randall, eds.), Vol. 10, 239-283. Academic Press, Orlando, Florida. Folmar, L. C., and Dickhoff, W. W. (1980). Aquaculture 21, 1-37. Fontaine, M.(1975). Adv. Mar. Biol. W, 241-355. Forrest, J. N., Jr., Cohen, A. D., Schon, D. A., and Epstein, F. H. (1973). Am. J. Physiol. 224,709-713. Forrest, J. N., Jr., Boyer, J. L., Ardito, T. L., Murdaugh, H. V., and Wade, J. B. (1982). Am. J . Physiol. 242, C388-C392. Foskett, J. K., and Scheffey, C. (1982). Science 215, 164-166. Foskett, J. K., Logsdon, G. D., Turner, T., Machen, T. E., and Bern, H. A. (1981). J . Exp. Biol. 93,209-224. Foskett, J. K., Machen, T. E., and Bern, H. A. (1982).Am. J . Physiol. 242, R380-R389. Foskett, J. K., Bern, H. A., Machen, T. E., and Conner, M . (1983). J . Exp. Biol. 106, 255-281.

Fujita, M., and Yamamoto, T. (1984). Arch. Histol. Jpn. 47, 113-118. Garcia-Romeu, F., and Maetz, J. (1964).J . Gen. Physiol. 47, 1195-1207. Getman, H. C. (1950). Biol. Bull. (Woods Hole, Mass.) 99,439-445. Girard, J. P., and Payan, P. (1980). Am. J. Physiol. 238, R260-R268. Hasegawa, S., Hirano, T., Osagawa, T., Iwata, M., Akiyama, T., and Arai, S. (1987). Fish Physiol. Biochem. 4, 101-110. Hoar, W. S. (1976). J . Fish. Res. Board Can. 33, 1234-1252. Hootman, S. R., and Philpott, C. W. (1978). Anat. Rec. 190,687-702. Hootman, S . R., and Philpott, C. W. (1979). Anat. Rec. 193,99-130. Hootman, S . R., and Philpott, C. W. (1980). A m . J. Physiol. 238, R199-R206. Hossler, F. E. (1980). A m . J . Physiol. 7, R160-Rl64. Hossler, F. E., Ruby, J. R., and McIlvain, T. D. (1979). J. Exp. Zool. 208,399-406. Hossler, F. E., M u d , G., Karnaky, K. J., Jr., and Epstein, F. H. (1985). J . Morphol. 185, 377-386. Hossler, F. E., Harpole, J. H., and King, J. A. (1986). J . Submicrosc. Cyrol. 18,519-528. Hughes, G . M. (1984). In “Fish Physiology” (W. S. Hoar and D. J. Randall, eds.), Vol. 10, pp. 1-72. Academic Press, Orlando, Florida. Hwang, P. P. (1987). Mar. Biol. 94,643-649. Jacobs, D., Esmond, E. F., Melisky, E. L., and Hocutt, C. H. (1981). Can. J. Fish Aquar. Sci. 38, 16-22.

230

M. PISAM AND A. RAMBOURG

Johnson, C. E., and Cheverie, J. C. (1985). Can. J. Fish. Aquat. Sci. 42,1994-2003. Kamiya, M. (1972). Comp. Eiochem. Physiol. E 43B, 61 1-617. Karnaky, K. J., Jr. (1980). Am. J. Physiol. 238,R185-Rl98. Karnaky, K. J., Jr., and Kinter, W. B. (1977). J . Exp. Zool. 199,355-364. Karnaky, K. J., Jr., and Philpott, C. W. (1969). J . Cell Eiol. 43,64a. Karnaky, K. J., Jr., Ernst, S. A,, and Philpott, C. W. (1976a). J. Cell Eiol. 70, 144-156. Karnaky, K. J., Jr., Kinter, L. B., Kinter, W. B., and Stirling, C. E. (1976b). J . CellEiol. 70, 157-177. Karnaky, K. J., Jr., Degnan, K. J., Garreston, L. T., and Zadunaisky, J. A. (1979). Bull. Mt. Desert I d . Biol. Lab. 19, 109-111. Karnaky, K. J., Jr., Degnan, K. J., and Zadunaisky, J. A. (1984). Am. J . Physiol. 246, R770-R775. Karnovsky, M. J. (1971). Proc. Am. SOC.CellEiol. 284, 146a. Kawahara, T., Sasaki, T., and Higashi, S . (1982). J . Electron Microsc. 31, 162-170. Kendall, M., and Dale, J. (1979). J. Fish. Res. Board Can. 36, 1072-1079. Kessel, R. G., and Beams, H. W. (1962). J. Ultrastruct. Res. 6,7747. Keys, A., and Willmer, E. N. (1932). J. Physiol. (London)76,368-378. Kikuchi, S . (1977). Cell Tissue Res. 180,87-98. Kikuchi, S . (1981). Annu. Rep. Iwate Med. Univ. Sch. Lib. Art. Sci. 16,33-45. King, J. A. C., and Hossler, F. E. (1986). Scanning Electron Microsc. 4,1477-1488. King, 1. A. C., Abel, D. C., and DiBona, D. R. (1989). Cell Tissue Res. 257,367-377. Komourdjian, M. P., Saunders, R. L., and Fenwick, J. C. (1976a). Can. J. Zool. 54, 531535. Komourdjian, M. P., Saunders, R. L., andFenwick, J. C. (1976b). C0n.J. Zool. 54,544-551. Komuro, T., and Yarnamoto, T. (1970). J . Electron Microsc. 19, 194a. Krogh, A. (1938). Z. Vergl. Physiol. 25,335-350. Krogh, A. (1939). In “Osmotic Regulation in Aquatic Animals,” pp. 130-153. Cambridge Univ. Press, LondodDover, New York. Lacy, E. R. (1983). Am. J. Anat. 166, 19-39. Lahlou, B. (1980). In “Environmental Physiology of Fishes” (M. A. Ah, ed.), NATO Ser., pp. 201-240.) Plenum, New York. Laurent, P. (1984). In “Fish Physiology” (W. S.Hoar and D. J. Randall, eds.), pp. 73-183. Academic Press, Orlando, Florida. Laurent, P., and Dunel, S . (1976). Acta Zool. (Stockholm) 57, 189-209. Laurent, P., and Dunel, S . (1978). C. R . Hebd. Seances Acad. Sci. 286, 1447-1450. Laurent, P., and Dunel, S . (1980). Am. J . Physiol. 238, R147-RI59. Laurent, P., and Perry, S. F. (1990). Cell Tissue Res. 259,429-442. Laurent, P., Hobe, H., and Dunel-Erb, S . (1985). Cell Tissue Res. 240,675-692. Leino, R. L., McCormick, J. H., and Jensen, K. M. (1987). Cell Tissue Res. 250,389-399. and Bradley, T. M. (1989). J. Fish Eiol. 34,259-272. Lubin, R. T., Rourke, A. W., Madsen, S . S . (1990). Gen. Comp. Endocrinol. 79,l-11. Maetz J. (1971). Philos. Trans. R . SOC.London, Ser. E 262,209-251. Maetz, J. (1974). Eiochem. Biophys. Perspect. Mar. Biol. 1, 1-167. Maetz, J. (1976). Ciba Found. Symp. 38, 133-159. Maetz, J., and Garcia-Romeu, F. (1964). J . Gen. Physiol. 47, 1209-1227. Maetz, J., and Pic, P. (1977). J. Exp. Zool. 199, 325-338. Maina, J. N. (1990). Anat. Embryol. 181,83-98. Marshall, W. S. (1977). J. Comp. Physiol. E 114, 157-165. Marshall, W. S.,and Nishioka, R. S . (1980). J. Exp. Zool. 214, 147-188. Mattheij, J. A. M., and Stroband, H. W. J. (1971). Z . Zel(forsch. l21,93-101.

ULTRASTRUCTURE OF MITOCHONDRIA-RICH GILL CELLS

23 1

Mayer-Gostan, N., Wendelaar-Bonga, S. E., and Balm, P. H. M. (1987). I n “Vertebrate Endocrinology: Fundamentals and Biomedical Implications” (P. K. T. Pang and M. P. Schreibman, eds.), Vol. 2, pp. 21 1-238. Academic Press, Orlando, Florida. McCormick, S. D., and Saunders, R. L. (1987). Am. Fish. SOC.Symp. 1,211-229. Morgan, M., and Tovall, P. (1973). Z . Zellforsch. 142, 147-162. Newstead, J. D. (1971). 2.Zellforsch. 116, 1-6. Nicoll, C. S., Wilson, S. W., Nishioka, R., and Bern, H. A. (1981). Gen. Comp. Endocrinol. 44,365-373. Olivereau, M. (1970). C. R . Seances SOC.Biol. Ses Fil164, 1951-1955. Olson, K., and Fromm, P. (1973). Z . Zellforsch. 143,439-449. Payan, P. (1978). J . Comp. Physiol. W, 181-188. Perry, S. F., and Laurent, P. (1989). J. Exp. Biol. 147, 147-168. Perry, S. F., and Wood, C. M. (1985). J . Exp. Biol. 116,411-434. Petrik, P., and Bucher, 0. (1969). Z . Zellforsch. %,66-74. Philpott, C. W. (1962). Anat. Rec. 142,267-268. Philpott, C. W. (1964). J. CellBiol. 23,74a. Philpott, C. W. (1965). Protoplasma 60,7-23. Philpott, C. W. (1966). J. CellBiol. 31, 86a. Philpott, C. W. (1967). J. CellBiol. 35, 104a. Philpott, C. W. (1968). I n “Cystic Fibrosis” (R. Portes and M. O’Connor, eds.), pp. 109-122. Ciba Found. LondonKhurchill, London. Philpott, C. W. (1980). A m . J. Physiol. 238,R171-Rl84. Philpott, C. W., and Copeland, D. E. (1963). J . CellEiol. 18,389-404. Pic, P., and Lahitette, P. (1981). J . Comp. Physiol. 141,523-529. Pisam, M. (1981). Anat. Rec. 200,401-414. Pisam, M., Sardet, C., and Maetz, J. (1980). Am. J . Physiol. 238, R213-R218. Pisam, M., Chretien, M., Rambourg, A., and Clermont, Y. (1983). Anat. Rec. 207,385-397. Pisam, M., Caroff, A., and Rambourg, A. (1987). Am. J . Anat. 179,40-50. Pisam, M., Prunet, P., Boeuf, G., and Rambourg, A. (1988). Am. J. Anar. 183,235-244. Pisam, M., Prunet, P., and Rambourg, A. (1989). Am. J . Anat. 184,311-320. Pisam, M., Boeuf, G., Prunet, P., and Rambourg, A. (1990). Am. J . Anat. 187,21-31. Pisam, M., Prunet, P., and Rambourg, A. (1991). Manuscript in preparation. Prunet, P., Boeuf, G., and Houdebine, L. M. (1985). J. Exp. Zool. 235, 187-196. Prunet, P., Boeuf, G., Bolton, J. P., and Young, G. (1989). Gen. Comp. Endocrinol. 74, 355-364. Rambourg, A., and Clermont, Y. (1990).Eur. J . Cell Biol. 51, 189-200. Richman, N. H., Nishioka, R. S., Young, G., and Bern, H. A. (1987). Gen. Comp. Endocrinol. 67, 194-201. Riddle, C. V., and Ernst, S. A. (1979). J . Membr. Biol. 45,21-35. Ritch, R . , and Philpott, C. W. (1969). Exp. Cell Res. 55, 17-24. Sardet, C. (1980). Am. J . Physiol. 238,R207-R212. Sardet, C., Pisam, M., and Maetz, J. (1979). J. Cell Biol. 80,96-117. Sargent, J. R., Thomson, A. J., and Bornancin, M. (1975). Comp. Biochem. Physiol. B 51B, 75-79. Sargent, J. R., Pirie, B. J. S., Thomson, A. J., and George, S. G. (1977). I n “Physiology and Behavior in Marine Organisms” (D. S. McLusky and A. J. Berry, eds.), pp. 123-134. Pergamon, Oxford, England. Saunders, R. L., and Henderson, E. B. (1978). J . Fish. Res. Board Can. 35, 1542-1546. Shirai, N., and Utida, S. (1970). Z . Zellforsch. 103,247-264. Silva, P., Solomon, R., Spokes, K., and Epstein, F. H. (1977). J . Exp. Zool. 199,419-426.

232

M. PISAM AND A. RAMBOURG

Smith, H. W. (1930). Am. J. Physiol. 93,480-505. Smith, H. W. (1932). Q. Rev. Biol. 7 , 1-26. Straus, L. P. (1%3). Physiol. Zool. 36, 183-198. Straus, L. P., and Doyle, W. L. (1961). Am. Zool. 1,392a. Thomson, A. J., and Sargent, J. R. (1977). J . Exp. Zool. u)o,33-40. Threadgold, L. T.,and Houston, A. H. (1961). Nature (London) 190,612-614. Threadgold, L. T., and Houston, A. H. (1964). Exp. Cell Res. 34, 1-23. Utida, S., Kamiya, M., and Shirai, N. (1971). Comp. Biochem. Physiol. A 38A, 443-447. Wedeweyer, G., Saunders, R. L., and Clarke, W. L. (1980). Mar. Fish. Rev. 42, 1-14. Wendelaar Bonga, S. E., and van der Meij, C. J. M. (1989). Cell Tissue Res. 255,235-243. Wendelaar Bonga, S. E., Flik, G., Balm, P. H. M., and van der Meij, J. C. A. (1990). Cell Tissue Res. 259,575-585. Zadunaisky, J. A. (1984). I n “Fish Physiology” (W. S. Hoar and D. J. Randall, eds.), Vol. 10, pp. 129-176. Academic Press, Orlando, Florida.

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 130

Structure and Function of Plant Cell Walls: Immunological Approaches TAKAYUKI HOSON Department of Biology, Faculty of Science, Osaka City University, Osaka 558, Japan

I. Introduction Plant cells are surrounded by a cell wall which gives them characteristics different from animal cells. The cell wall used to be regarded as merely a dead structureless envelope, but it is actually a delicately balanced complex entity having a diverse range of roles in the support of plant life. The important functions of plant cells depend greatly on the presence of cell walls and their characteristics. A plant cell cannot exist for long without its cell wall. Thus, the cell wall is an indispensable organelle and a fascinating object for study. The life cycle of a flowering plant begins with germination, which is followed by a complex series of processes of growth and differentiation. Reproductive growth then starts, and the plant reaches a stage of senescence. Throughout its life, the cell wall provides the plant with the mechanical strength with which to construct and protect itself against the environment. At the same time, the cell wall must allow the plant to grow. This is accomplished by qualitatively and quantitatively controlled weakening of the cell wall at appropriate times. The extent and location of the cell wall weakening determine the rate and direction of cell expansion, and therefore control plant growth and morphogenesis. When plant cells differentiate, dramatic changes occur in their cell walls. Each differentiated cell possesses its characteristic cell wall. During senescence, dissociation of certain types of cells is observed (e.g., cell abscission and fruit softening). Such dissociation is brought about by selective breakdown of the cell wall. Furthermore, the cell wall has important roles in plant resistance to pathogenic organisms, in the perception of environmental signals, and in cell-to-cell recognition. The diversity of cell wall functions arises from the extraordinary complexity of its structure. The cell wall can be divided into primary and secondary cell walls, the primary one being the wall of growing cells, and the secondary one being that which is formed after growth has stopped and differentiation has started. Both walls consist of a microfibrillar phase and a matrix phase. The microfibrillar phase, cellulose, is a polymer of 233

Copyright 0 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

234

TAKAYUKI HOSON

(1 + 4)-P-linked D-glucosyl residues. Cellulose constitutes 20-50% of the primary cell wall. The matrix, consisting of a variety of polysaccharides, proteins, and phenolic compounds, is very complex and varies with cell type, position, condition, and age. Its composition also differs between Gramineae and other monocotyledons or dicotyledons. The general structure of the plant cell wall has been reviewed by Darvill et al. (19801, McNeil et al. (1984), Dey and Brinson (1984), Fry (1988), and Brett and Waldron (1990). The structural characterization of the plant cell wall has been attempted by many researchers. Developments in gas chromotography-mass spectrometry and nuclear magnetic resonance spectroscopy have greatly advanced our understanding of the primary structure of cell wall polymers, especially polysaccharides (McNeil et al., 1984). However, little is known about the secondary, tertiary, and quaternary structures. X-Ray diffraction analysis can be used to determine the conformation of some crystallized polysaccharides (Preston, 1979). Circular dichrometry provides information about the secondary structure. However, these methods are applicable to only some cell wall polymers. What seems best are immunological methods which offer potent probes for clarifying the location, metabolism, and function, as well as the structure of cell wall polymers. The molecular biological studies of today would not have been possible without immunological methods for identifying gene products. Study of the plant cell wall using immunological approaches has rapidly advanced in recent years. It is timely to summarize the advances in this field and to discuss some future prospects. 11. Antibodies as Probes for the Study of Plant Cell Walls

In order to understand the structure and function of plant cell walls, potent probes are needed to specifically distinguish details of cell wall components. Antibodies, lectins, and enzymes could be used for this purpose. Antibodies are the most suitable, as they are more precise than other candidates. Antibodies can now be generated against different types of target cell wall componets. A. ANTIBODIES RAISEDAGAINST CELLWALLCOMPONENTS The primary cell wall contains 5-10% protein (Lamport, 1970; Darvill et al., 1980; Cassab and Varner, 1988). Various enzymes have been detected in the cell wall, and they may be involved in metabolic activities associated with cell wall functions. As shown in Table I, antibodies have been raised against diverse wall enzymes and utilized for different purposes since the

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

235

TABLE I ANTIBODIES RAISEDAGAINST PLANT CELL WALL ENZYMES Enzyme

Material

Cellulase

Pea epicotyl Bean leaf abscission zone Avocado fruit Kidney bean leaf abscission zone (1+3),( 1+4)-P-Glucanase Endoglucanase Barley aleurone layer Maize coleoptile Exoglucanase

(1-+3)-P-Glucanase Polygalacturonase a-Galactosidase P-Fructosidase Chitinase L ysozyme Peroxidase

(1+3)-P-Glucan synthase

Maize coleoptile Cultured tobacco cells Bean leaf Soybean seedling Tomato fruit Mung bean seed Tomato fruit Cultured carrot cells Cultured tobacco cells Bean leaf Wheat germ Cultured peanut cells Potato tuber Maize seedling Peach seed Cultured soybean cells

Reference Byrne et al. (1975) Sexton et al. (1980) Awad and Lewis (1980) Koehler et al. (1981) Woodward and Fincher (1982) Hoson and Nevins (1989~);Inouhe and Nevins (1991b) Hoson and Nevins (1989~); Labrador and Nevins (1989) Felix and Meins (1985) Vogeli et al. (1988) Takeuchi et al. (1990) Tucker et al. (1980); Ali and Brady (1982);DellaPenna et al. (1986) Hankins er al. (1979) Iki et al. (1978) Laurihre et al. (1989) Shinshi et al. (1987) Vogeli et al. (1988) Audy et al. (1988) Cairns et al. (1980); Stephan and van Huystee (1980); Hu et al. (1987) Espelie and Kolattukudy (1985) Kim et al. (1988) Quesada et al. (1990) Fink et al. (1990)

1970s. Cellulase and peroxidase have been purified from various sources to generate antibodies. Cell wall protein, especially that from dicotyledons, is exceptionally rich in hydroxyproline. Much of this amino acid is a component of a highly basic glycoprotein named extensin (see Lamport, 1970; Lamport and Catt, 1981; Cooper et af., 1984; Cassab and Varner, 1988). Extensin appears to have a role in growth regulation and resistance to pathogens. Antibodies have been raised against extensin from soybean seed coat (Cassab and Varner, 1987), a melon callus (Mazau et af., 1988), and carrot roots (Stafstrom and Staehelin, 1988). At first, extensin was thought to be an insoluble polymer. Later, the existence of salt-extractable extensin was reported

236

TAKAYUKI HOSON

(Chrispeels, 1969; Stuart and Varner, 1980). Antibodies were also raised against this putative precursor of extensin (Kieliszewski and Lamport, 1986; Conrad et al., 1987). Differences in the amount and structure of extensin were found between dicotyledons and Gramineae (see Darvill et al., 1980;Cassab and Varner, 1988).Less extensin is present in Gramineae cell walls. Antibodies have been generated against maize extensin (Stiefel et al., 1988; Kieliszewski et al., 1990). Extensin is the major component of Chlamydomonas cell walls, and immunological characterization of this extensin has been carried out (Smith et al., 1984; Roberts et al., 1985; Woessner and Goodenough, 1989). Arabinogalactan proteins are another class of hydroxyproline-rich glycoproteins. They differ in structure from extensin (see Clarke et al., 1979; Fincher et al., 1983), and their protein content is usually between 2 and 10%. Galactose and arabinose are the major monosaccharides of their carbohydrate moieties; uronic acids are also common components. Arabinogalactan proteins are present at the plasma membrane-cell wall interface and may be involved in signal perception and resistance to environmental stress. They interact with Yariv antigens (Jermyn and Yeow, 1975), and this has been utilized to purify and characterize them. Also, antibodies have been raised against them (Anderson et al., 1984; Tsumuraya et al., 1984). The generation of antibodies specific for cell wall carboyhdrates is a recent achievement. In general, carbohydrates are also effective antigens. Actually, antibodies directed at sugar residues are obtained when cell wall glycoproteins are used as antigens (Anderson et al., 1984; Smith et al., 1984; McManus et al., 1988; Lauribre et al., 1989). Antibodies have been raised against pectic polysaccharides, rhamnogalacturonan I (Moore et al., 1986)and polygalacturonic acid (Liners et al., 1989;Knox et al., 1990), and against hemicelldosic ones, such as xyloglucan (Moore et al., 1986) and (1 --f 3),(1 + 4)-/3-~-glucan (Hoson and Nevins, 1989a). Pure (1 + 3),( 1 + 4)-/3-~-glucantriggered the production of antibodies, while other polysaccharides were coupled to, or mixed with, a protein carrier to raise antibodies. Misaki and co-workers established a method for generating antibodies specific for a-L-aribinose(Kaku et al., 1986;Misaki et al., 1988) and xyloglucan oligosaccharides (Sone et al., 1989a,b) coupled to bovine serum albumin. The method was extended to obtain antibodies that can recognize different mono- and oligosaccharides (Northcote et at., 1989). It should be noted that antibodies are also raised against Darabinose residues, when Freund’s complete adjuvant, which contains bacterial polysaccharides, is used to emulsify antigens (Kaku et al., 1986). Antibodies have been raised against unidentified antigens. Extracts of whole tissues (Evans et al., 1988), cell walls (Huber and Nevins, 1981;

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

237

Morrow and Jones, 1986; Hatfield and Nevins, 1988; Melan and Cosgrove, 1988; Cohn et al., 1989), or protoplasts (Norman et al., 1986; Lynes et al., 1987) and proteins secreted from cultured cells (Satoh and Fujii, 1988; Esaka et al., 1990) have been used as antigens. The antibodies generated are useful for characterizing compounds in antigen fractions. In most of the studies just mentioned, polyclonal antibodies were raised in rabbits or mice. Monoclonal antibodies have been produced against cell wall enzymes (Hu et al., 1987; Kim et al., 1988), glycoproteins (Smith et al., 1984; Anderson et al., 1984), and polysaccharides (Liners et al., 1989; Knox et al., 1990). Because monoclonal antibodies recognize only a specific epitope, they may be more advantageous for some of our purposes. In some studies, Fab fragments, the immunologically working ends of immunoglobulins, are used instead of whole antibody molecules (Melan and Cosgrove, 1988; Ward et al., 1988). Immobilized papain, available commercially, can be used to prepare Fab fragments. In addition to antibodies, enzymes (Rue1 and Joseleau, 1984; Stone, 1984; Bonfante-Fasolo et al., 1990) and lectins (Baldo et al., 1982, 1984; Hayashi and Maclachlan, 1984; Hoson and Masuda, 1987, 1991; Benhamou et d . , 1990a) have been used as probes to study the structure and function of the plant cell wall. Enzymes appear to have sufficient specificity, but their catalytic activity can decrease their usefulness as probes. Lectins are proteins of nonimmune origin which are capable of specific recognition and binding to sugar moieties of different carbohydrates, without altering the covalent structure of any of the recognized sugar ligands (see Etzler, 1985; Lis and Sharon, 1986). Lectins with different sugar specificities and with potential utility in the study of the plant cell wall have been purified (Table 11). Because lectins are easy to obtain and handle, they are especially suitable for the screening of different cell wall components or preliminary studies. The limitations to their use are that their specificities are not always high and that lectins specific for arabinose or xylose residues have not been found in nature. This is why Kaku et al. (1986) tried to generate antibodies which recognize a-L-arabinofuranose residues.

B. PURIFICATION AND SPECIFICITY Antiserum raised against different cell wall components has been directly used for the study of the plant cell wall, but usually the serum is purified to obtain specific probes. Immunoglobulins are sometimes prepared via ammonium sulfate precipitation. A more convenient and reliable way of purifying immunoglobulinG (IgG) is affinity chromatography using protein A or G, surface proteins from Staphylococcus species, which can

238

TAKAYUKI HOSON TABLE I1 LECTINS IMPORTANTTO THE STUDYOF PLANTCELLWALLS

Sugar specificity a-D-Mannose, a-D-glucose &D-GalaCtOSe a-L-Fucose N-Acetylglucosamine N -Acet ylgalactosamine D-Ghcuronic acid D-Galacturonic acid

Cell wall component

Source of lectin

Enzyme or structural glycoprotein Galactan, xyloglucan, arabinoxylan Xyloglucan

Canavalia ensiformis," Lens culinaris, Pisum satiuum Abrus precatorius,b Ricinus communis (120),bViscum albumb Tetragonolobus purpureas, Ulex europaeus (I) Bandeiraea simplicifolia (II), Triticum uulgaris,' Ulex europaeus (11) Dolichos biporus, Glycine max, Helix pomatia Limulus polyphemus Aplysia depilans

Enzyme or structural glycoprotein Enzyne or structural glycoprotein Arabinoxylan Polygalacturonic acid

Concanavalin A. Extremely hazardous. Wheat germ agglutinin.

bind with the Fc fragment of IgG. Immobilized proteins A and G are commercially available. If sufficiently purified antigens are used to generate antibodies, no further purification is needed. In some cases, especially when carbohydrates coupled to a protein carrier are used as antigens, antibodies raised against contaminating compounds should be removed. By incubating crude antibodies with contaminating compounds, undesired antibodies can be precipitated. The complete removal of antibodies to contaminating compounds has successfully been carried out by affinity chromatography, using these compounds as ligands (Bennett and Christoffersen, 1986; Kaku et al., 1986; Moore et al., 1986; Northcote et al., 1989; Fink et al., 1990). Monoclonal antibodies specific for target cell wall components can be obtained by screening the hybridomas. The specificity of antibodies thus purified is examined by immunodiffusion assays, such as double-diffusion (Ouchterlony, 1949),the quantitative precipitation reaction, the complement fixation assay, the hapten competition test, immunoelectrophoresis, and immunoblot analysis. For protein antigens, Western blot analysis has been used routinely. Polyclonal antibodies were raised against oat caryopsis (1 + 3), (1 + 4)-P-~-glucanswith an average molecular weight of 1.5 x lo4 (Hoson and Nevins, 1989a). The specificity of these antibodies was examined by precipitation reaction (Fig. 1) and dot blot analysis (Fig. 2). The latter has been carried out on nitrocellulose membrane. Because the binding of polysaccharides to this membrane is not always strong, a cellulosecoated thin-layer plate may be more useful for the analysis of polysac-

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

" 0

239

100

200 300 Polysaccharide (pg) FIG.1. Quantitative precipitation of (1 + 3),(1 + 4)-p-~-glucanantibodies with various P-D-glucans. Antibodies (70 pg of protein) were incubated with different amounts of glucans for 72 hours. G, Oat caryopsis (1 + 3),(1 + 4)-p-~-glucan;Li, lichenan; La, laminarin; P, pachyman; C, carboxymethylcellulose.

FIG.2. Dot blot analysis of the specificity of (1 -+ 3),(1 + 4)-P-~-glucanantibodies to different glucans. Each glucan ( I , 2, and 5 pg) was dotted onto a cellulose-coated glass plate, and made to react with the antibodies. The plate was stained with Coomassie Brillant Blue solution. G, Oat caryopsis (1 + 3),(1 -+ 4)-P-~-glucan;Li, lichenan; La, laminarin; P, pachyman; C, carboxymethylcellulose.

240

TAKAYUKI HOSON

charides with a strong affinity for cellulose. In both tests, antibodies reacted with the oat glucans and lichenan. However, they did not exhibit any reaction with laminarin, pachyman, carboxymethylcellulose,or xyloglucan. Thus, these antibodies are specific for (1 + 3),(1 + 4)-p-Dglucans. C. APPLICATIONS The most popular applicationsof antibodies are for studying the location of various components of the plant cell wall. By fluorescence microscopy, using the secondary antibodies conjugated with fluorescencedyes, we can easily find the location and distribution of certain types of cell wall components within a tissue, organ, or plant body. The occurrence of particular components among different species can also be examined. Electron microscopy by the immunogold method can further clarify the subcellular localization of cell wall components. This method provides much information on the synthesis, processing, and secretion pathways of cell wall polysaccharides and proteins. The content of certain types of cell wall components can be determined histochemically by measuring the strength of fluorescence or density of gold particles. The specificity of antibodies is very precise, and antibodies can recognize the conformation of cell wall molecules. Thus, minute differences in the structure of cell wall polysaccharides or proteins among different sources or isomers can be detected wit.h antibodies. Cell wall components can be purified with an immunoaffinity column having antibodies as ligands. Immunoassays, particularly the enzyme-linked immunosorbent assay, have been widely used to determine the concentrations of various cell wall components. Moreover, the immunoprecipitation of in uitro translation products is indispensable for molecular biological studies of the plant cell wall. Antibodies serve as specific inhibitors of biochemical reactions (e.g., enzymatic activity) and physiological events (e.g., growth and leaf abscission). Conventional inhibitors often lack specificity and show different side effects. Because specific inhibitors are needed to understand the mechanism of the regulation of cell wall functions, antibodies may also greatly contribute to their study. 111. Location and Metabolism of Cell Wall Polymers

Cell wall polysaccharides as well as structural glycoproteins have been localized at various levels. The occurrence and metabolism of several enzymes have also been reported. The blotting of fresh tissue onto nitro-

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

24 1

cellulose membrane, called tissue printing, is a useful technique for localizing cell wall proteins (Cassab and Varner, 1987; Spruce et al., 1987). Indirect staining with fluorescein isothiocyanate (FITC) or gold conjugates of anti-IgG antibodies is the standard procedure. Details and characteristics of the procedure have been described by Knox (1982), Lloyd (1987), and Herman (1988). A. POLYSACCHARIDES 1 . Xyloglucan

Xyloglucan is the principal hemicellulosic polysaccharide of the primary cell wall of higher plants, especially dicotyledons. It consists of a backbone of (1 + 4)-P-linked D-glucosyl residues and thus is hydrogen-bonded to cellulose (see McNeil et al., 1984; Fry, 1989; Hayashi, 1989). Fluorescence microscopy using FITC-labeled fucose-binding lectins has shown that xyloglucan is localized both on and between cellulose microfibrils in pea stems (Hayashi and Maclachlan, 1984). Thus, the presence of such a binding between xyloglucan and cellulose is supported histochemically . The location of xyloglucan has also been studied by an indirect fluorescence method, using antibodies raised against xyloglucan oligosaccharides by Sone et al. (1989a,b). The cell wall of etiolated azuki bean epicotyls exhibits only weak autofluorescence when exposed to blue light. Administration of preimmune serum does not influence the fluorescence (Fig. 3A). However, when epicotyl sections are treated with antibodies against xyloglucan heptasaccharide (Xyl3Glcd, the cell wall fluorescence increases, indicating binding of the antibodies to the walls of the epidermal, parenchymatous, and vascular bundle cells (Fig. 3B). The fluorescence regions are distributed almost evenly along the cell wall. FITClabeled fucose-binding lectin, Ulex europaeus agglutinin (UEA) I, is also bound to the cell wall of azuki bean epicotyls (Fig. 3C). Distinct fluorescence regions are observed along the epidermis and outer cell layers of the parenchyma. The binding of UEA I to the cell wall is inhibited in the presence of fucose (Fig, 3D), suggesting the specificity of binding of the lectin to fucose residues of the cell wall. Thus, fucose-containing xyloglucans are localized at the cell wall of the outer part of the stems, although xyloglucans are present in the wall of all types of cells in azuki epicotyls. A study using the same antibodies indicated that xyloglucan is restricted to the primary cell wall and is absent from the middle lamella in soybean cotyledons (Sone et al., 1990). Antibodies raised against xyloglucan polysaccharides were also exclusively bound to the primary cell wall in root tip and leaf tissues of red clover (Moore and Staehelin, 1988), while the

242

TAKAYUKI HOSON

FIG. 3. Fluorescence micrographs of sections from subapical regions of azuki bean epicotyls and maize coleoptiles treated with antibodies. (A and B) Azuki thin sections were treated with (A) preimmune serum (PIS) or (B) xyloglucan heptasaccharide antibodies, and then incubated with fluorescein isothiocyanate (F1TC)-labeled secondary antibodies. (C and D) Azuki sections were treated with FITC-labeled Ulex europaeus agglutinin (UEA) I in the (D) presence or (C) absence of fucose. (E and F) Maize thin sections were treated with (E)PIS or (F) (1 + 3),( 1 + 4)-p-~-glucanantibodies, and then incubated with FITC-labeled secondary antibodies. Bar, 100 pm.

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

243

antibodies were bound to the entire cell wall, including the middle lamella, in suspension-cultured sycamore cells (Moore et al., 1986). Suspensioncultured cells excrete much xyloglucan into the medium, causing the cells to produce copious amounts of compensate for the loss. The presence of xyloglucan in the middle lamella may be a special event occurring only in cultured cells (Moore and Staehelin, 1988). Electron microscopy using antibodies against xyloglucan polysaccharides supported the view that the Golgi apparatus is the site of synthesis of this polysaccharide (Moore and Staehelin, 1988). Xyloglucan is present within the Golgi cisternae and vesicles. The concentration of xyloglucan in the appressed central regions of the Golgi cisternae is very low, and it is not detected in the endoplasmic reticulum. The antibodies also showed that xyloglucan is present in the forming cell plate of dividing cells and in adjacent Golgi stacks and vesicles (Moore and Staehelin, 1988). Thus, the Golgi apparatus is clearly the source of materials for the cell plate. 2 . ( I + 3),(I + 4)-P-~-G/ucan O-D-Glucan containing both 1,3 and 1,4-linkages is a characteristic hemicellulosic polysaccharide in the primary cell wall of Gramineae (Nevins et al., 1978; McNeil et al., 1984). A study using FITC-conjugated Bacillus subtilis (1 + 3),(1 + 4)-p-~-glucanaseas a probe indicated that this polysaccharide is present in the inner layer of the aleurone cell wall (Stone, 1984). The location of (1 + 3),(1 + 4)-P-~-glucan was further studies with antibodies specific for this polysaccharide. These antibodies become bound to the cell walls of the epidermis, parenchyma, and vascular bundles of maize coleoptiles (Fig. 3F), while the preimmune serum does not (Fig. 3E). Distinct antibody binding is observed along the inner layers of the cell wall, indicating that the newly synthesized P-D-glucans are deposited in the inner wall (Hoson and Nevins, 1989b). This glucan is absent from the cell wall of embryos, but increases rapidly with coleoptile growth, and then declines after elongation has been completed (Carpita, 1984; Lutteneger and Nevins, 1985; Hoson and Nevins, 1989b). There was a good correlation between the glucan content and the amount of antibodies bound to the cell walls (Hoson and Nevins, 1989b). Antibodies became bound to the cell wall of coleoptiles, mesocotyls, and roots of different Gramineae species, but not to the cell wall of some dicotyledons tested. The results clarified the occurrence and distribution of (1 + 3),(1 + 4)-p-~-glucan. 3. Polygalacturonic Acids Pectic polysaccharides have been localized using different types of antibodies. Rhamnogalacturonan I is restricted to the middle lamella and is

244

TAKAYUKI HOSON

especially evident in the junctions between cells in suspension-cultured sycamore (Moore et al., 1986) and in root tip or leaf tissues of red clover (Moore and Staehelin, 1988). Rhamnogalacturonan I, like xyloglucan, is present within the Golgi cisternae and vesicles. This polysaccharide is not detected in the endoplasmic reticulum. While xyloglucan is present in the forming cell plate of dividing cells, rhamnogalacturonan I is practically absent from the cell plate (Moore and Staehelin, 1988). These observations partially support, but partially contradict, the supposed location of this polysaccharide. Rhamnogalacturonan I antibodies recognize the long repeating regions of polygalacturonic acid (Moore and Staehelin, 1988). Monoclonal antibodies specific for unesterified and methyl-esterified polygalacturonic acids were used to locate these polysaccharides (Knox et al., 1990). In carrot root apex, esterified polygalacturonic acid is located evenly throughout the cell wall, whereas unesterified polygalacturonic acid is localized at the inner surface of the primary cell wall and the middle lamella and abundantly on the outer surface at intercellular spaces. The cell walls of Gramineae contain less pectic polysaccharides than do those of dicotyledons and other monocotyledonous families (Darvill et al., 1980; Jarvis, 1984; Masuda and Yamamoto, 1985). The location of polygalacturonic acid is also characteristic in Gramineae (Knox et al., 1990). In the root apex of oat, unesterified polygalacturonic acid is localized at the region of intercellular spaces of the cortical cells. Methyl-esterified polygalacturonic acid is present in the cell wall of the cortex and the stele, but is absent from the wall of the epidermal or root cap cells. In oat coleoptiles or leaves, the cell walls of all tissues contain the unesterified polygalacturonic acid, but not the esterified one. 4 . Other Polysaccharides Antibodies specific for a-L-arabinofuranoseresidues have been used to locate arabinogalactans in dicotyledons and arabinoxylans in Gramineae. These antibodies were found to be bound to the primary cell wall and middle lamella of soybean cotyledons by an indirect immunofluorescence method (Misaki et al., 1988). Support for this came from work on bean roots with the immunogold method, using the same type of antibodies (Northcote et al., 1989). Arabinogalactans are also detected in the cell plate of dividing bean cells, although they are absent from the secondary walls of bean cells as well as suspension-culturedZinnia cells. The specific organelle is detected within the cytoplasm and the vacuole with the antibodies (Northcote et al., 1989). However, this organelle has not been characterized. Arabinoxylans in rice endosperm are present in the primary cell walls (Misaki et al., 1988). In pollen tubes of Nicotiana alata, a - ~ -

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

245

arabinofuranose-containingpolysaccharides are localized at the outer fibrillar wall layer (Anderson ef al., 1987). Using xylanase-gold conjugates, Rue1 and Joseleau (1984) observed that xylans occurred in the primary cell wall of parenchyma cells. Antibodies raised aginst 1,4-p-linked xylan oligosaccharides also showed the presence of xylans in growing cell walls of bean and Zinnia cells (Northcote er al., 1989). The heaviest staining is observed in the secondary cell wall of xylem cells. Xylans are not present in the cell plate of dividing root cells. Northcote et al. (1989) studied the location of callose using antibodies against 1,3-~-oligoglucans.This polysaccharide is detected in the primary cell wall and the cell plate in bean roots. In growing cell walls, callose is located specifically in the plasmodesmata. During cell plate formation, callose is dispersed over the whole plate area. These results confirm the observations by conventional histochemistry. B. STRUCTURAL GLYCOPROTEINS The location of extensin was examined in carrot roots, using polyclonal antibodies raised against glycosylated extensin 1 , the most abundant hydroxyproline-rich glycoprotein in this material (Stafstrom and Staehelin, 1988). Extensin is present quite uniformly throughout the primary cell wall, but is absent from the expanded middle lamella at the junction of three or more cells. The glycoprotein is also reduced in the narrow middle lamella at the center of the cell wall between two cells. The finding that entensin cannot enter the pectin-rich middle lamella indicates that the wall surrounding a cell is synthesized by that cell alone (Stafstrom and Staehelin, 1988). In soybean seeds, extensin is mainly localized in the seed coat, hilum, and vascular tissues (Cassab and Varner, 1987). The seed coat of soybean consists of two layers: the palisade epidermis and hourglass cells. Extensin is not detected at the early developmental stages, but starts accumulating first in the cell wall of the palisade epidermis. In mature seeds, much extensin is present in the walls of both seed coat layers. Extensin was also immunolocalized in the cell wall of maize roots (Ludevid et al., 1990), although no detailed subcellular localization has been reported. The synthesis and secretion pathways of extensin in higher plants have not been fully studied by immunological methods. In Chlamydomonas, immunohistochemical observation demonstrated that extensin is synthesized in the endoplasmic reticulum, glycosylated in the Golgi stack, and transported to the cell surface via transitional vesicles (Grief and Shaw, 1987).

246

TAKAYUKI HOSON

C. ENZYMES Cell wall enzymes have been localized immunohistochemically. Of the wall enzymes, (1 + 3),(1 + 4)-p-~-glucanase(Stuart et al., 1986; Hoson and Nevins, 1989c), (1 + 3)-p-~-glucanase(Mauch and Staehelin, 1989; Benhamou et al., 1989; Keefe et al., 1990), chitinase (Mauch and Staehelin, 1989;Keefe etal., 1990;Behamou etal., 1990b),and peroxidase (van Huystee and Lobarzewski, 1982; Griffing and Fowke, 1985; Chibbar and van Huystee, 1986;Espelie et al., 1986;Kim et al., 1988)have been the main objects of study. Because the location and metabolism of wall enzymes are directly related to the particular functions of the cell wall, details of these studies are described in the following sections.

IV. Growth Regulation

Growth of plant cells requires extension of their cell walls. The rate and direction of plant cell growth are determined by how much their cell walls are extensible and in which direction. When plant genes, hormones, or environmental factors alter the growth rate or direction, they must influence directly or indirectly the mechanical properties of the cell wall. The plant hormone auxin, in particular, induces plant cell elongation growth by increasing the mechanical extensibility or by decreasing the yield threshold of the cell wall (see Cleland, 1971; Masuda, ,1978, 1990; Taiz, 1984). Changes in the mechanical properties of the cell wall are brought about by biochemical modifications of certain components. Table I11 summarizes the metabolic turnover of cell wall polysaccharides due to auxin in dicotyledons and in Gramineae. As can be seen, several types of matrix polysaccharides undergo auxin-induced modifications, and the types differ between dicotyledons and Gramineae, probably because of differences in matrix composition. Thus, we first need to determine the metabolic modification crit.ica1 for auxin-induced changes in the mechanical properties of the cell wall (i.e., cell wall loosening). The cause-effect relationship between auxin-induced cell elongation via wall loosening and changes in the cell wall polysaccharides has been examined by imposing conditions which modify growth rate or interfere with the metabolism of wall polysaccharides. If a certain change in polysaccharides is observed in segments in the presence of a 0.2-0.3 M mannitol solution in which auxin-induced elongation is suppressed osmotically, such a change is not a consequence of auxin-induced elongation. However, it is not clear whether the change is really a cause of elongation. Inhibitors of enzymic or metabolic reactions have been utilized

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

247

TABLE I11 AUXIN-INDUCED METABOLIC TURNOVER OF CELL WALL POLYSACCHARIDES Polysaccharide Dkot y ledons Xyloglucan

Turnover Degradation

Pea internode

Solubilization

Pea internode

Decrease in molecular weight Autolysis Polygalacturonic Solubilization acid Autolysis (Arabin0)galactan Degradation Decrease in molecular weight Autolysis Gramineae (1+3),( h 4 ) p -D-GIu can

Material

Degradation

Azuki epicotyl Azuki epicotyl Pea internode Azuki epicotyl Pea internode Azuki epicotyl Azuki epicotyl Chick-pea epicotyl Oat coleoptile Barley coleoptile

Autolysis Arabinoxylan

Degradation Autolysis

Xy loglucan

Solubilization Decrease in molecular weight

Rice coleoptile Maize coleoptile Maize coleoptile Oat coleoptile Maize coleoptile Oat coleoptile Oat coleoptile

Reference Labavitch and Ray ( 1974a,b) Terry and Bonner (1980); Terry et al. (1981) Nishitani and Masuda (1981, 1983) Hoson (1990) Terry et al. (1981) Hoson (1990) Gilkes and Hall (1977) Nishitani and Masuda (1980, 1981) Nishitani and Masuda ( 1980) Seara et al. (1988) Loescher and Nevins (1972); Sakurai and Masuda (1977) Sakurai and Masuda (1978) Zarra and Masuda (1979) Inouhe and Nevins (1991a) Darvill et al. (1978) Heyn (1986) Nock and Smith (1987) Inouhe et al. (1984) Inouhe ef al. (1984)

to suppress changes in polysaccharides and examine the influence on auxin-induced growth. Unfortunately, conventional inhibitors lack specificity, yielding confusing results. This can be overcome by using antibodies raised against wall polysaccharides or wall enzymes which are involved in the metabolism of the polysaccharides to offer high specificity. If we can clarify the cause-effect relationship between auxin-induced cell elongation and changes in the cell wall polysaccharides, the next steps toward understanding the mechanisms of regulation of plant cell growth would be (1) to precisely identify chemical changes in cell wall components

248

TAKAYUKI HOSON

due to auxin, (2) to clarify the characteristics of wall enzymes involved in the modifications in the wall components, and (3) to find how auxin regulates the enzyme activity. For these purposes, antibodies specific for wall components serve as sensitive probes as well as inhibitors. A. PRELIMINARY STUDY APPROACHES Two preliminary approaches have been taken to understand the role of the cell wall in growth regulation in higher plants. Huber and Nevins (1981) generated antiserum against the wall protein fraction extracted from maize coleoptile cell wall with 3 M LiC1. The antiserum inhibited indole-3-acetic acid (1AA)-induced elongation of coleoptile segments. The cuticle had to be removed by abrasion of the coleoptile surface to observe significant inhibition of elongation. Preimmune serum or the serum preincubated with the cell wall preparation did not exhibit growth inhibition, suggesting the specificity of the effect. The cell wall extracts were not purified further in this study. However, the antiserum clearly inhibited autohydrolysis of the isolated wall fraction from maize coleoptiles (Huber and Nevins, 1981). Because (1 + 3),(1 + 4)-P-~-glucanis mainly hydrolyzed during this autolysis of the wall of Gramineae coleoptiles (Lee et al., 1967; Huber and Nevins, 1979), the degradation of this glucan may be closely associated with auxin-induced cell elongation. Subsequently, Nevins et al. (1987) and Hatfield and Nevins (1988) established a procedure for evaluating cell wall protein fractions for their ability to reverse growth inhibition caused by antibodies against whole wall proteins. The same approach has been used to study auxin-induced growth in dicotyledons. Antiserum was generated agaisnt extracellular proteins obtained from pea internode segments by centrifugation (Morrow and Jones, 1986). Antibodies thus prepared did not inhibit auxin-induced elongation of pea segments. Antibodies were also raised against 3 M LiCl extracts of pea epicotyl cell wall (Melan and Cosgrove, 1988). The antibodies did not influence in uitro extension of pea segments. Thus, the above approach was not successful in dicotyledons. However, it was found that 3 M LiC1-extracted wall proteins from cucumber hypocotyls had activity for inducing in vitro extension of inactivated segments (McQueen-Mason et al., 1990). Further studies are required with dicotyledons. Another type of approach was carried out using lectins. Hoson and Masuda (1987) examined the effect of 12 kinds of lectins, representing six groups of sugar specificities, on auxin-induced elongation of segments of azuki bean epicotyl (dicotyledon)and oat coleoptile (Gramineae). Concanavalin A, Tetragonolobus purpureas agglutinin, and Ulex europaeus agglutinin I suppressed elongation of azuki segments. Elongation of oat

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

249

segments was suppressed by concanavalin A, wheat germ agglutinin, and agglutinins from Dolichos bgorus, Glycine max, or Limulus polyphemus. These lectins were bound to the cell wall of the segments, as examined by fluorescence microscopy, and inhibited auxin-induced cell wall loosening (decrease in the minimum stress-relaxation time of the cell wall), as measured by the stress-relaxation method (Yamamoto et al., 1970). These results demonstrate that the wall components containing a-D-mannose (a-D-glucose) or a-fucose in dicotyledons and those containing a-Dmannose (a-D-glucose), N-acetyl-D-glucosamine, N-acetyl-mgalactosamine, or D-glucuronic acid in Gramineae play important roles in auxininduced cell elongation via cell wall loosening (Hoson and Masuda, 1987). Because xyloglucan present in the cell wall of dicotyledons contains Lfucose on the side chains (Kato and Matsuda, 1976; Kato et al., 1981), xyloglucan may be one of the components critical for the growth of dicotyledonous cells. The results obtained in these studies suggest that the breakdown of xyloglucan in dicotyledons and ( 1 + 3),( 1 + 4)-/3-~-glucanin Gramineae is at least one of the causes of auxin-induced cell elongation. The significance of the breakdown of these wall polysaccharides in growth regulation has been studied further. B. DICOTYLEDONS 1 . Xyloglucan

Mechanisms by which fucose-binding lectins inhibit auxin-induced elongation of segments of dicotyledonous organs have been studied (Hoson and Masuda, 1989, 1991). Polysaccharides extracted from the cell wall of etiolated azuki bean epicotyls with 24% KOH were separated with a gel filtration column. Xyloglucan determined by the iodine method shows a single peak with an average molecular weight of 1.5 x lo6. IAA treatment of the segments causes a shift of distribution of xyloglucan from higher to lower molecular weight. When epicotyl segments were incubated with IAA in the presence of fucose-binding lectins, the decrease in molecular weight of xyloglucan due to IAA was completely inhibited. The cell wall preparation, from which pectic polysaccharides had been removed by enzyme digestion, autolyzed xyloglucan (Hoson, 1990). Fucose-binding lectins inhibited such a hydrolysis of xyloglucan. The inhibitory effect of the lectins on xyloglucan metabolism was not observed when the lectins were preincubated with fucose. Thus, a fucose-binding lectin is a specific inhibitor of the breakdown of xyloglucan. These findings, as well as the inhibitory effects on auxin-induced elongation and wall loosening, support

250

TAKAYUKI HOSON

the view that the breakdown of xyloglucan is associated with the cell wall loosening responsible for auxin-induced stem elongation in dicotyledons. Antibodies raised against xyloglucan oligosaccharides (Sone et al., 1989a,b) were further used to clarify the relationship between auxininduced xyloglucan breakdown and elongation (Hoson et al., 1991a). Of the antibodies tested, those against hepta- and octasaccharide inhibit the IAAinduced decrease in the molecular weight of xyloglucans in a 24% KOH fraction of azuki epicotyls (Table IV). The autolytic release of xylosecontaining products from the pectin-removed cell wall is also suppressed by hepta- and octasaccharide antibodies, which were found to inhibit IAA-induced cell elongation and cell wall loosening in azuki epicotyl segments (Table IV). Preimmune serum or antibodies against xyloglucan disaccharide (isoprimeverose) exhibit little or no effect on IAA-induced xyloglucan breakdown, elongation, or cell wall lossening. Thus, the view that the breakdown of xyloglucan is the cause of cell wall loosening responsible for auxin-induced stem elongation in dicotyledons was further confirmed by the antibody experiments. Some differences were found in the effects of fucose-binding lectins and xyloglucan antibodies on xyloglucan breakdown in azuki epicotyls. Although both inhibited the auxin-induced decrease in the molecular weight of xyloglucans in the 24% KOH fraction, only the lectins exhibited an inhibitory effect on the breakdown of xyloglucans present in the fraction extracted with KOH after pectinase treatment (unpublished observations). Xyloglucan breakdown in the latter fraction is associated with the removal of terminal L-fucosyl residues. Because both cell wall loosening and cell elongation due to auxin are similarly suppressed by the lectins and the antibodies, removal of the terminal fucose may trigger further degradation of xyloglucans, which may result in the cell wall loosening responsible for cell elongation. Plant stems are composed of inextensible epidermis cells and extensible parenchyma cells. Auxin induces elongation of the stem segments by loosening the thick growth-limiting outer epidermal cell wall (see Kutschera, 1989; Masuda, 1990). When segments from growing regions of plant stems are split longitudinally and floated on the buffer solution, the segments bend outward due to “tissue tension.” Auxin inhibits this outward bending. Antibodies specific for xyloglucan heptasaccharides reverse the inhibitory effect of auxin on the bending (unpublished observations). The results indicate that the breakdown of xyloglucan in the outer epidermal cell wall plays an essential role in the auxin-induced elongation of dicotyledons. Thus, only xyloglucans present in the outer epidermal wall are involved in the regulation of auxin-induced elongation, although xyloglucans exist in the cell wall of any type of cells in azuki epicotyls (Fig.

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

25 1

TABLE IV EFFECTOF XYLOGLUCAN ANTIBODIES ON ELONGATION, CELLWALL AND XYLOGLUCAN BREAKDOWN I N AZUKIAND LOOSENING, OATSEGMENTS~ Treatment Azuki epicotyl Initial Buffer IAA' IAA + P I 9 IAA + AbXG2g IAA + AbXG78 IAA + AbXG88 Oat coleoptile Initial Buffer IAA' IAA + P I 9 IAA + AbXG2g IAA + AbXG7R IAA + AbXG8R

Elongation (76)

3.5 t 0.2" 9.7 f 0.5 8.9 f 0.3 8.7 t 0.3 6.1 f 0.4d 6.3 2 0.3"

2.8 t 0.3" 22.2 -C 0.9 22.6 t 1.4 19.9 f 0.8 21.1 f 0.6 21.7 t 0.7

(msec)

Molecular massC (MDa)

13.9 t 0.3d 13.2 f 0.6" 10.0 t 0.5 10.5 t 0.6 9.8 f 0.9 12.3 t 0.7d 12.5 f 0.6d

1.56 t 0.04" 1.50 t 0.07d 1.25 t 0.09 1.23 f 0.04 1.30 f 0.10 1.55 f 0.08" 1.53 f 0.03"

15.0 f 0.7d 15.5 f 1.0" 10.5 2 0.5 10.6 t 0.4 10.7 t 0.5 10.3 t 0.5 10.4 t 0.6

0.72 f 0.02" 0.66 5 0.00" 0.55 ? 0.02 0.59 2 0.00 0.66 t 0.02" 0.64 f 0.01" 0.64 f 0.01"

TOb

Segments were grown for 4 hours. Values represent means ? standard errors. Minimum stress-relaxation time. Average molecular mass of xyloglucans, as determined with a Sepharose CL-4B column. Mean value significantly different from the IAA treatment at the 5% level. ' Indole-3-acetic acid at IO-'M. Preimmune serum at 400 pg/ml. Antibodies raised against xyloglucan disaccharide (Xyl,Glc,), heptasacchande (Xyl$31c4), and octasaccharide (GallXyl&31c4)at 400 pg/ml. a

3B). Fucose-containing xyloglucans are localized at the cell wall of the outer part of the stems (Fig. 3C). The presence of terminal fucosyl residues in xyloglucan molecules appears to be closely associated with the regulation of auxin-induced elongation in dicotyledons. 2 . Other Polysaccharides Antibodies have been raised against pectic polysaccharides and utilized for the study of their location (Moore e? al., 1986; Moore and Staehelin, 1988; Knox et al., 1990) or of changes in their conformation induced by calcium (Liners et al., 1989). The role of polygalacturonase, a pectindegrading enzyme, in the regulation of fruit softening has been studied, using antibodies specific for this enzyme (Ali and Brady, 1982; DellaPenna e? al., 1986; Tieman and Handa, 1989). However, these antibodies have

252

TAKAYUKI HOSON

not been used to clarify the cause-effect relationship between the auxininduced breakdown of pectic polysaccharides and elongation via cell wall loosening, or to understand the mechanism by which auxin induces breakdown of the polysaccharides. Antibodies raised against a-L-arabinofuranose residues (Kaku et al., 1986) were found to inhibit auxin-induced elongation of azuki bean epicotyl segments (Hoson and Masuda, 1986). However, they exhibited an inhibitory effect only at high concentration, when nonspecific effects appear. Lectins which recognize both D-galactose and N-acetylgalactosamine residues did not inhibit auxin-induced elongation of azuki segments (Hoson and Masuda, 1987). These data suggest that the breakdown of (arabin0)galactans is not always associated with auxin-induced cell elongation. In autolysis of the cell wall of dicotyledons, (arabin0)galactans are mainly degraded (Labrador and Nicolas, 1985; Seara et al., 1988). Cosgrove (1989) reported that autolysis of these polysaccharides was not related to the in uitro extension of segments from cucumber hypocotyls. Thus, the breakdown of (arabino)galactans appears to be unnecessary for auxin-induced elongation in dicotyledons, although these polysaccharides are actively metabolized in the cell wall of growing plant tissues. C. GRAMINEAE 1 . (1 + 3),(1 + 4)-P-~-Glucan

The significance of the breakdown of (1 + 3),( 1 + 4)-P-~-glucanin auxin-induced cell elongation in Gramineae has been studied, using antibodies against this polysaccharide as well as enzymes which are involved in its breakdown. The results indicate that breakdown of the glucan plays an essential role in auxin-induced elongation. Antibodies specific for ( 1 + 3),(1 + 4)-P-~-glucanwere generated against the oat caryopsis glucan fraction (Hoson and Nevins, 1989a). Auxin decreases the noncellulosic glucose content of the maize coleoptile cell wall (Table V), as reported in coleoptiles of other species (Loescher and Nevins, 1972; Sakurai and Masuda, 1977, 1978; Zarra and Masuda, 1979). The antibodies inhibit an auxin-induced decrease in noncellulosic glucose content (Table V). The isolated cell wall preparation of maize coleoptiles autolyzes the glucan, which is suppressed by the antibodies. Preimmune serum or glucan antibodies pretreated with the glucan do not inhibit the glucan degradation. Thus, the antibodies are specific inhibitors of both in viuo and in uitro degradation of (1 + 3),( 1 + 4)-P-~-glucan.The glucan antibodies also specifically inhibit auxin-induced elongation and cell wall loosening (decrease in the minimum stress-relaxation time) in maize coleoptile segments (Table V). These results support the idea that

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

253

TABLE V EFFECTOF (1-*3),(1-.4)-P-D-GLUCAN ANTIBODIES ON ELONGATION, CELLWALLLOOSENING, AND GLUCAN BREAKDOWN I N MAIZE COLEOPTILE SEGMENTS" Treatment Initial Buffer IAA' IAA + IAA + IAA + IAA +

Elongation

TOb

(%I

(msec)

Glucan contentC (pg/segment)

-

17.1 f O S d 17.4 2 0.5d 14.0 f 0.8 15.0 f 0.7 14.4 f 0.6 17.0 f 0.8d 15.4 f 0.6

74.0 2 0.3d 71.7 f O S d 62.1 f 0.5 64.5 2 0.9 61.6 f 1.9 69.1 f 0.5d 61.5 f 0.1

5.0 t 0.4d

PIY Glucang Abh Ab(g1ucan)'

11.0 f 0.7 10.6 f 0.7 10.2 f 0.5 6.7 f O S d

9.8 f 0.6

Segments were grown for 4 hours. Values represent means standard errors. Minimum stress-relaxation time. ' Glucan content was determined by gas chromatography. Mean value significantly different from the IAA treatment at the 5% level. Indole-3-acetic acid at 10-5M. f Preimmune serum at 200 pglml. R (1+3),(1+4)-p-~-Glucan at 200 pglml. Antibodies raised against (1+3),(1+4)-p-~-glucan at 200 pglml. Antibodies pretreated with (1+3),(1+4)-p-~-glucan.

the degradation of (1 + 3),( 1 + 4)-P-~-glucanby cell wall enzymes is the cause of the cell wall loosening responsible for auxin-induced cell elongation in Gramineae. Using maize coleoptiles, Hatfield and Nevins (1988) observed that only one wall protein fraction, which has no hydrolase activity, was capable of precipitating the growth-inhibiting factor of antibodies raised against whole wall proteins. Also, both endoglucanase and exoglucanase fractions reacted with the antibodies and could effectively reverse the growthinhibiting activity (Hoson and Nevins, 1989~).Actually, antibodies raised against these glucanase fractions are capable of inhibiting the auxininduced elongation of maize coleoptile segments, indicating that they are involved in cell elongation. Antibodies against two glucanase fractions also inhibit auxin-induced cell wall loosening, as measured by the stressrelaxation method (Hoson and Nevins, 1989~).Furthermore, these antibodies inhibit the auxin-induced decrease in noncellulosicglucose content and the autolytic reactions of isolated cell walls (Hoson and Nevins, 1989c; Labrador and Nevins, 1989; Inouhe and Nevins, 1991b). Thus, auxininduced degradation of (1 + 3),( 1 + 4)-P-~-glucan,cell wall loosening, and cell elongation are inhibited when either the substrate (glucan) or the enzyme (glucanases)is conjugated by antibodies. These data clearly con-

254

TAKAYUKI HOSON

firm the significance of degradation of the glucan in auxin-induced elongation in Gramineae. When auxin induces growth of plant organs, the thick outer epidermal cell wall is a target of auxin even in coleoptiles, which are hollow cylinders having both an outer and an inner epidermis (Kutschera et al., 1987; Masuda et al., 1989). Split segments of maize coleoptiles bend outward, and auxin inhibits such a bending. Antibodies specific for (1 + 31, (1 + 4)-P-~-glucanreverse the inhibitory effect of auxin on the bending (Hoson et al., 1991b). Moreover, the antibodies are effective only when they are administered through the outer epidermis (Hoson et al., 1991b). The results indicate that the degradation of (1 + 3), (1 + 4)-P-~-glucan in the outer epidermal cell wall is required for auxin-induced elongation of Gramineae tissues. 2 . Other Polysaccharides Of the lectins with different sugar specificities, those that recognize D-glucuronic acid inhibit auxin-induced cell elongation and cell wall loosening in oat coleoptile segments (Hoson and Masuda, 1987). Antibodies specific for the a-L-arabinofuranose residues exhibit similar physiological effects in oat segments (Hoson and Masuda, 1986). These results suggest that the breakdown of (g1ucurono)arabinoxylan is associated with the cell wall loosening responsible for auxin-induced cell elongation. The effects of these lectins and antibodies on the breakdown of this polysaccharide should be analyzed. Antibodies raised against xyloglucan oligosaccharides were also used to clarify the relationship between auxin-induced xyloglucan breakdown and elongation in oat coleoptiles (Hoson et al., 1991a). All of the antibodies tested inhibited the IAA-induced decrease in the molecular weight of the water-insoluble hemicellulose, but none influenced IAA-induced cell elongation or cell wall loosening (Table IV). These results contradict the view that the breakdown of xyloglucan is the cause of the cell wall loosening responsible for auxin-induced cell elongation also in Gramineae. This may be the first time that the expected relationship between the breakdown of a certain type of wall polysaccharide and auxin-induced elongation was clearly denied by antibodies raised against cell wall components. V. Selective Breakdown of Plant Cell Walls

Dissociation of certain types of cells occurs during the life cycle of plants due to selective breakdown of their cell walls. Immunological approaches have been used to understand the mechanism of this cell wall breakdown.

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

255

A. GERMINATION During germination of Gramineae caryopses, different hydrolases, especially a-amylase, are synthesized in the aleurone layers and the scutellum in response to gibberellic acid, a plant hormone, produced in the embryo. These enzymes are released into the endosperm to hydrolyze storage materials. The cell walls of the aleurone layers and the scutellum, as well as those of endosperm cells, should be hydrolyzed for these enzymes to reach the endosperm. The cell wall of aleurone cells consists of two layers. The inner thin cell wall is enriched with (1 + 3),( 1 + 4)-P-~-glucan,while the thicker outer wall consists mainly of arabinoxylan (Fincher, 1989). The cell walls of the scutellum and the endosperm also contain (1 + 3),( 1 + 4)-P-~-glucanand arabinoxylan. Germinating barley caryopses have at least two types of ( 1 -+ 3), (1 + 4)-P-~-glucanaseisoenzymes. Polyclonal antibodies raised against these isoenzymes are mutually cross reactive (Woodward and Fincher, 1982). These antibodies also showed that isoenzyme I, with a molecular weight of 28,000 and an isolectric point (PI) of 8.5, was synthesized predominantly in the scutellum, while isoenzyme 11, with a molecular weight of 30,000 and a PI higher than 10.0, was synthesized exclusively in the aleurone (Stuart et al., 1986). When in uitro translation products of mRNA from aleurone cells are immunoprecipitated with the antibodies, a polypeptide with a molecular weight of 33,000 is detected (Mundy and Fincher, 1986). Gibberellic acid greatly increases the translatable mRNA encoding the glucanases (Mundy and Fincher, 1986). No such mRNA is detected in ungerminated barley caryopses by hybrization histochemistry study using the cDNA probe (McFadden e? al., 1988). Expression of the glucanase genes is first detected in the scutellum 1 day after the initation of germination and is confined to the epithelial layer. After 2 days, the levels of mRNA decrease in the scutellar epithelium, but increase in the aleurone. Induction of glucanase gene expression in the aleurone layer progresses from the proximal to the distal end of the caryopsis (McFadden et al., 1988). Enzymes capable of degrading arabinoxylan are detected in germinating caryopses. The activity of xylanase, one such enzyme, is very low in barley aleurone at the early stage of germination and increases in response to gibberellic acid (Taiz and Honigman, 1976; Dashek and Chrispeels, 1977). Barley aleurone xylanase has been purified (Benjavongkulchai and Spencer, 1986), but has not been characterized immunologically. Little is known about the breakdown processes of the cell wall in germinating seeds of species other than Gramineae. The cell wall of the endosperm in some species, such as date palm, contains (ga1acto)mannans as

256

TAKAYUKI HOSON

storage carbohydrates. The characteristics of a-galactosidase, one of the enzymes capable of hydrolyzing this polysaccharide, were studied with antibodies specific .for this enzyme. Immunogold localization of a-galactosidase indicated that the enzyme was transferred to and through the Golgi apparatus, and finally deposited within the protein body in developing soybean cotyledons (Herman and Shannon, 1985). During germination of date palm seeds, a-galactosidase present in the protein body decreases, while the enzyme appears in a thin region of the inner cell wall of the endosperm (Chandra Sekhar and DeMason, 1990). In endosperm undergoing digestion, a-galactosidase is localized in the flocculent contents of vacuoles, which results from storage protein breakdown, then accumulates in the inner cell wall of cell cavities formed by the dissolution of the cytoplasm. Finally, the enzyme diffuses throughout the endosperm cell wall. Thus, a-galactosidase is stored in the protein body of endosperm cells and released to the cell wall following cell disintegration.

B. LEAFABSCISSION Abscission is the process by which plants shed organs, such as leaves, buds, stems, petals, stigmas, stamens, fruits, and seeds. In abscission, a highly coordinated sequence of biochemical events leads to cell wall breakdown in one or two rows of cells present in the particular region called the abscission zone (Sexton and Roberts, 1982). Many studies have focused on leaf abscission, especially that in the bean. Ethylene, a gaseous plant hormone, induces it. Horton and Osborne (1967) first reported that cellulase activity increased dramatically during bean leaf abscission. There are two types of cellulase isoenzymes in bean leaves, one form having an acidic PI of 4.5 and the other, a basic PI of 9.5 (Lewis et al., 1972). Antibodies were raised against the purified 9.5 cellulase (Sexton et al., 1980; Koehler et al., 1981). The antibodies do not cross-react with 4.5 cellulase. By assaying cellulase activity before and after immunoprecipitation with the antibodies, 9.5 cellulase activity was found to be predominantly confined to the abscission zone, while 4.5 cellulase activity was more widely spread throughout the plant (Durbin et al., 1981; del Campillo et al., 1988). The 9.5 cellulase activity is not detectable prior to the induction of abscission by ethylene. After the induction, 9.5 cellulase activity increases rapidly by de n o w synthesis. The 4.5 cellulase activity is highest in young tissues, with its level falling after induction. Furthermore, the antibodies injected into the bean abscission zone inactivate the enzyme and inhibit abscission (Sexton et al., 1980). These results indicate that 9.5 cellulase is involved in the cell wall breakdown processes responsible for leaf abscission.

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

257

There are two abscission zones in the primary leaves of bean; the distal abscission zone is at the base of the leaf blade in the pulvinus-petiole junction, and the proximal abscission zone is in the petiole-stem junction. The studies mentioned above focused on the proximal zone. The occurrence and distribution of 9.5 cellulase in both abscission zones were studied, using antibodies specific for 9.5 cellulase to detect the enzyme on nitrocellulose tissue prints (Reid er al., 1990; del Campillo ef al., 1990). In the distal zone, the cellulase appears first in the cells of the separation layer adjacent to vascular traces and extends toward the periphery. In the proximal zone, the enzyme accumulates first in the cortical cells in the adaxial side and then extends to the abaxial side. Thus, there are some differences between the two abscission zones in the 9.5 cellulase distribution in the separation layer (del Campillo et al., 1990). The 9.5 cellulase antibodies immunoprecipitate in virro translation products of abscission zone poly(A)+ RNA with a molecular weight of 51,000 (Tucker et al., 1988). The antibodies specific for bean leaf 9.5 cellulase reacted with two forms of cellulase present in anthers of Lathyrus odoratus (Sexton et al., 1990). These enzymes may be associated with anther dehiscence. Another cell wall enzyme that appears to be implicated in leaf abscission is polygalacturonase. Morr6 ( 1968) reported an increase in polygalacturonase-like activity in the bean abscission zone. In the abscission zones of tomato and elder leaves, both cellulase and polygalacturonase activities increased after the induction of abscission (Roberts er al., 1989). Polygalacturonase activity is primarily restricted to the abscission zone, while cellulase is not. Thus, polygalacturonase instead of cellulase may be involved in leaf abscission in these species. A study using polygalacturonase cDNA as a probe could not detect any increase in the level of polygalacturonase mRNA during abscission (Roberts et al., 1989). Immunological characterization of the roles of polygalacturonase in leaf abscission remains to be carried out. C. FRUITSOFTENING Ripening is the final phase of fruit development, involving a series of coordinated biochemical events which result in dramatic changes in the texture, color, and flavor of fruits. Softening, a common and characteristic feature associated with ripening, is brought about by a selective and extensive breakdown of the cell wall (Labavitch, 1981; Huber, 1983; Brady, 1987). Various changes occur in the structure of different cell wall components during fruit ripening. Therefore, we should determine which of the many changes play critical roles in the wall softening, as in the case of growth regulation. The mechanisms by which certain types of changes

258

TAKAYUKI HOSON

in the cell wall are regulated must be clarified as well. Immunological approaches have contributed to these purposes. Polygalacturonase in tomato fruit has been intensively studied. In tomato fruit, there is a good correlation between the rate of softening and the degree of pectin degradation (see Labavitch, 1981; Brady, 1987). Pectic polysaccharides in tomato fruits are hydrolyzed by a successive action of pectin methylesterase and polygalacturonase. The activity of the former does not always change with fruit softening, and polygalacturonase appears to be the limiting enzyme of pectin degradation (Labavitch, 1981). Tomato polygalacturonasecomes in three forms: PG 1, PG 2A, and PG 2B. PG 1 is probably a dimer of the PG 2s (Tucker et al., 1980). The PG 2s have similar molecular size and structure (Ali and Brady, 1982). There is disagreement as to which is the endogenous form of the enzyme (Pressey, 1988; Bruinsma et al., 1989). Antibodies raised against these polygalacturonase isoenzymes are mutually cross reactive (Tucker et d., 1980; Ali and Brady, 1982). Radioimmunoassay using antibodies against the PG 2s showed that polygalacturonase was not detectable in green tomatoes and rapidly increased during ripening (Tucker and Grierson, 1982). PG 1 appears first and the PG 2s appear later, but eventually the PG 2s become predominant, according to immunoelectrophoresis analysis (Brady et al., 1982). There are several tomato mutants that show little or no ripening. The level of polygalacturonase measured with the antibodies is low in such mutant fruits (Tucker and Grierson, 1982). Immunoassay of polygalacturonaseand the determination of mRNA encoding this enzyme by immunoprecipitation indicate that polygalacturonaseis synthesized de nouo during ripening (Tucker and Grierson, 1982; Grierson et al., 1985). Translation of polygalacturonase mRNA in uitro results in the synthesis of a single polypeptide with an apparent molecular weight of 54,000 (Sato et al., 1984; DellaPenna and Bennett, 1988). Immunocytolocalization of polygalacturonase in tomato fruits shows that the enzyme first appears in the columella region followed by sequential appearance in the exopericarp and endopericarp (Tieman and Handa, 1989). Peach fruits contain polygalacturonase which crossreacts with antibodies raised against tomato polygalacturonase(Lee et al., 1990). The results mentioned above support the view that the breakdown of pectin by polygalacturonase is associated with cell wall softening during the ripening of tomato fruits. However, the data obtained in two lines of studies contradict this view. Polygalacturonase antisense RNA was introduced into tomato plants, causing a substantial reduction in the level of polygalacturonaseproteins in the transformed fruit and an enzyme activity of almost 10% of the normal level. However, the fruit still underwent

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

259

softening, as judged by the compressibility test (Smith et al., 1988; Grierson et al., 1989). The polygalacturonase gene was ligated to a promoter and inserted into the fruit genome of one of the ripening-impaired mutants, and then plants were generated. Expression of this gene resulted in the accumulation of active polygalacturonase and the degeneration of pectin in the transgenic plant. However, no significant effect on fruit softening was detected. Thus, polygalacturonase is the primary determinant of pectin degradation, but this degradation alone is not sufficient for the induction of cell wall softening (Bennett et al., 1989; Giovannoni et al., 1989). That is, polygalacturonase is not the sole determinant of fruit softening in the tomato. In avocado fruit, both polygalacturonase and cellulase activities increase during the climacteric stage. Because cellulase activity started to increase 3 days before polygalacturonase was detected (Awad and Young, 1979), cellulase may contribute to the wall softening in this species. Antibodies were raised against avocado cellulase (Awad and Lewis, 1980) and have been utilized to study the mechanism by which cellulase production is induced during ripening. Immunoprecipitation by the antibodies indicated an increase in cellulase mRNA at the onset of ripening (Christoffersen et al., 1984; Tucker and Laties, 1984). The increase in cellulase activity may be caused by de nouo synthesis of this enzyme. The in uitro translation products of cellulase mRNA have a molecular weight of 54,000 and may undergo cleavage of the signal peptide, glycosylation, and carbohydrate trimming before they become mature enzymes (Bennett and Christoffersen, 1986). Although the above results suggest that cellulase is involved in cell wall softening in the avocado, how it degrades the cell wall is unknown. There is a possibility that cellulose is not a native substrate of this enzyme (Hatfield and Nevins, 1986). The activity of enzymes capable of hydrolyzing hemicellulosic polysaccharides also increases during the ripening of fruits in some species (Labavitch, 1981;Huber, 1983; Brady, 1987). Immunological characterization of these hydrolases remains to be done. Fruit softening may also be caused by the synthesis of highly methylated polygalacturonic acid or the ionic displacement of apoplastic calcium (Brady, 1987; Knee, 1989). There is a need to evaluate the contribution of these mechanisms to fruit softening.

VI. Other Aspects of Plant Cell Walls The hydroxyproline-rich cell wall glycoprotein, extensin, is thought to have a role in the cessation of cell elongation, formation of cell wall architecture, and maintenance of mechanical strength (Lamport, 1970;

260

TAKAYUKI HOSON

Lamport and Catt, 1981). Histochemical studies using antibodies or nucleic acid probes indicate that extensin accumulates in the cell wall of plants infected with fungi or bacteria (Mazau and Esquerre-Tugaye, 1986; Corbin et al., 1987; Sauer et al., 1990). Its accumulation is especially obvious in the walls of uninfected cells close to sites where the growth of fungi and bacteria is restricted. Extensin is also induced by wounding (Corbin et al., 1987; Ludevid et al., 1990; Sauer et al., 1990) or during cold acclimation (Weiser et al., 1990). Extensin and its mRNA are localized at actively dividing cells (Ludevid et al., 1990) or in the region initiating vascular elements (Keller et al., 1989; Stiefel et al., 1990). Thus, extensin may be involved in the defense against pathogenic organisms, resistance to various types of stresses, and cell differentiation. Cell wall glycoproteins obtained from maize are found to crossreact with antibodies raised against dicotyledonous extensins (Kieliszewski and Lamport, 1987; Kieliszewski et al., 1990). Immunological methods can also be utilized for such the structural analysis of extensins and their genes (Cassab and Varner, 1988). Peroxidase, widely distributed in the cell wall of plants, is responsible for the formation of lignin, suberin, ferulic acid bridges, or extensin bridges, and may be associated with growth regulation, formation of cell wall structure, and resistance to pathogenic organisms or stresses (Fry, 1986; Cassab and Varner, 1988; Lewis and Yamamoto, 1990). The molecular structure and function of peroxidase have been studied with immunological probes (van Huystee, 1987). Many isoenzymes of peroxidase have been found and distinguished into two major groups: anionic and cationic. Immunological characterization of these isoenzymes indicates that the structure of anionic peroxidase is distinctly different from that of the cationic type (van Huystee and Maldonado, 1982), and that peroxidases of the same group obtained from different species sometimes differ in structure (Lobarzewski and van Huystee, 1982; van Huystee and Maldonado, 1982). Even within the same group of peroxidases from one species, immunologically different types are found (Quesada et al., 1990). Thus, the molecular structure of peroxidase is very diverse. Most of the major peroxidases are glycosylated and can be detected in Golgi vesicles under immunocytochemical examination (Griffing and Fowke, 1985; Chibbar and van Huystee, 1986). In wound-healing potato tuber tissue, peroxidase is localized in the inside cell wall of the periderm, where it may play a role in the deposition of the aromatic polymeric domain of suberin (Espelie et al., 1986). The results of immunosassay using monoclonal antibodies specific for maize peroxidase suggest that the enzyme present in the wall participates in the stimulation of coleoptile and the inhibition of mesocotyl growth by red light exposure (Kim et al., 1989).

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

26 1

Chitinase and /3-1,3-glucanase are other enzymes believed to be involved in the resistance of plants to pathogenic organisms. They may directly act as lytic enzymes of the fungal cell wall or they may indirectly induce phytoalexin elicitors (Dixon and Lamb, 1990). Chitinase and p-1,3glucanase activities are increased by pathogen infection or by exogenously applied ethylene, a putative mediator of defense reactions (Abeles et al., 1970;Boller et al., 1983;Mauch et al., 1984). Immunoprecipitation of these enzymes confirmed the above observations and showed that they were synthesized de nouo (Vogeli et al., 1988). The synthesized enzymes are processed in the Golgi apparatus and transported to the cell wall (Mauch and Staehelin, 1989; Benhamou et al., 1989). Because the occurrence of chitinase in the cell wall is either preceded by or coincides with that of /3-1,3-glucanase,the glucan fragments released by their action may act as elicitors of chitinase production (Benhamou er al., 1990b).Tobacco mesophyll protoplasts synthesize characteristic proteins which cross-react with antibodies specific for chitinase and p-1,3-glucanase (Grosset et al., 1990). Thus, the production of these enzymes appears to be widely induced when plants are exposed to different kinds of stresses. Interestingly, chitinase and /3-1,3-glucanase are synthesized during flower formation in tobacco explants (Neale et al., 1990). Cell wall components play a principal role in cell-to-cell recognition (e.g., pollen-stigma interaction and legume-rhizobia symbiosis). Little information about these interactions has been obtained by immunological approaches, and further studies are needed. VII. Conclusions and Future Prospects Immunological approaches have been used to understand the structure and function of the plant cell wall. The occurrence and location of polysaccharides, structural glycoproteins, and enzymes constituting the cell wall have been examined with antibodies specific for these compounds. The antibodies are also useful probes for clarifying the conformational structure and the regulation of metabolism or gene expression of different wall components in relation to their roles in diverse functions of the cell wall. The main advantage of using antibodies as probes is that they yield much information about the cell wall in siru. The results of biochemical analyses and morphological studies carried out independently can be combined with immunological findings. The immunological approach should offer an answer to the cause-effect relationship between the metabolism of a particular wall component and a

262

TAKAYUKI HOSON

specific function of the cell wall. Antibodies have been utilized as specific and potent inhibitors of in uiuo breakdown of cell wall components during auxin-induced cell elongation (Huber and Nevins, 1981; Hoson and Nevins, 1989a,c; Hoson e? al., 1991a; Inouhe and Nevins, 1991b) and leaf abscission (Sexton et al., 1980). However, other uses of antibodies are limited to the study of in v i m activities of cell wall enzymes, such as chitinase, p- 1,3-glucanase, xylanase, and nitrate reductase (Schlumbaum et al., 1986; Vogeli et al., 1988; Ward e? al., 1988; Fuchs et al., 1989). In applying antibodies to plant tissues as in uiuo inhibitors, some steps must be observed; for example, the cuticle must be removed by abrasion to inhibit the breakdown of xyloglucan or (1 + 3),(1 + 4)-P-~-glucanby antibodies, and the antibodies should be injected into the abscission zone of leaves to suppress cellulase activity. Nevertheless, antibodies are useful as inhibitors of events occurring in the cell wall, because the cell wall constitutes the apoplast and therefore antibodies need not be taken up into the cytoplasm. Another topic not yet addressed by immunological approaches is the occurrence of physiologically active oligosaccharides in the cell wall and their quantitation. Certain types of oligosaccharides released from the plant cell wall exhibit diverse activities, such as elicitation of phytoalexin synthesis; stimulation of the synthesis of protease inhibitors, lignin, chitinase, or ethylene; induction of hypersensitive death of the cells; induction of flowering; and inhibition of auxin-induced elongation. These active oligosaccharides are called oligosaccharins (Albersheim and Darvill, 1985). Because they are effective at nanomolar concentrations, the immunoassay is the best way to determine their individual endogenous concentrations. Antibodies should help to confirm the significance of oligosaccharins in regulating different processes during the life cycle of plants. Characterizationof the structure and function of the cell wall by molecular biological methods is gaining prevalence, with nucleic acid probes being used instead of antibodies. However, molecular biological methods cannot completely replace the immunological methods for the following reasons. First, the translation products can be detected only by antibodies. Second, the translation products undergo further successive processing before they become functional proteins. Third, a probe must detect not only enzymes, but also wall substrates, most of which are polysaccharides. Finally, plants are strongly influenced by environmental factors, which the study of genes alone cannot cover. Thus, immunological approaches are needed for study of the plant cell wall. At present, immunological approaches are being used in almost 10% of cell wall studies. During the next decade, this percentage should increase,

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

263

and with it, our understanding about the structure and function of the plant cell wall. ACKNOWLEDGMENTS I thank Professors Y. Masuda and S. Kamisaka (Department of Biology), Professor A. Misaki and Dr. Y. Sone (Department of Food and Nutrition, Osaka City University), and Professor D. J. Nevins (University of California, Davis) for their cooperation and advice during the course of this study. Thanks are also due to Mrs. J. Noguchi for advice in preparing the manuscript.

REFERENCES Abeles, F. B., Bosshart, R. P., Forrence, L. E., and Habig, W. H. (1970). Plant Physiol. 47, 129-134. Albersheiin, P., and Darvill, A. G. (1985). Sci. Am. 253,58-64. Ali, Z. M., and Brady, C. J. (1982). Ausr. J . Plant Physiol. 9, 171-178. Anderson, M. A., Sandnn, M. S., and Clarke, A. E. (1984). Plant Physiol. 75, 1013-1016. Anderson, M. A., Hams, P. J., Bonig, I., and Clarke, A. E. (1987). Planra 171,438-442. Audy, P., Benhamou, N., Trudel, J., and Asselin, A. (1988). Plant Physiol. 88, 1317-1322. Awad, M., and Lewis, L. N. (1980). J . Food Sci. 45, 1625-1628. Awad, M., and Young, R. E. (1979). Plant Physiol. 64,306-308. Baldo, B. A., Boniface, P. A., and Simmonds, D. H. (1982). Aust. J . Planr Physiol. 9, 647-661. Baldo, B. A., Barnett, D., and Lee, J. W. (1984). Aust. J . Planr Physiol. 11, 179-190. Benhamou, N., Grenier, J., Asselin, A., and Legrand, M. (1989). Plant Cell 1, 1209-1221. Benhamou, N., Chamberland, H., and Pauze, F. J. (1990a). Plant Physiol. 92,995-1003. Benharnou, N., Joosten, M. H. A. J., and De Wit, P. J. G. M. (1990b). Plant Physiol. 92, 1108-1 120. Benjavongkulchai, E., and Spencer, M. S. (1986). Planra 169,415-419. Bennett, A. B., and Christoffersen, R. E. (1986). Plant Physiol. 81,830-835. Bennett, A. B., DellaPenna, D., Fischer, R. L., Giovannoni, J., and Lincoln, J. (1989). In “Cell Separation in Plants” (D. J. Osborne and M. B. Jackson, eds.), pp. 11-19. Springer-Verlag, Berlin. Boiler, T., Gehri, A., Mauch, F., and Vogeli, U. (1983). Planra l57,22-31. Bonfante-Fasolo, P., Vian, B., Perotto, S., Faccio, A., and Knox, J. P. (1990). Planta 180, 537-547. Brady, C. J. (1987). Annu. Rev. Plant Physiol. 38, 155-178. Brady ,C. J., MacAlpine, G., McGlasson, W. B., and Ueda, Y . (1982). Ausr. J . Plant Physiol. 9, 171-178. Brett, C., and Waldron, K. (1990). “Physiology and Biochemistry of Plant Cell Walls.” Unwin Hyman, London. Bruinsma, J., Knegt, E., and Vermeer, E. (1989). In “Cell Separation in Plants” (D. J. Osborne and M. B. Jackson, eds.), pp. 127-138. Springer-Verlag, Berlin. Byrne, H., Christou, N. V., Verma, D. P. S., and Maclachlan, G. A. (1975). J. Biol. Chem. 250, 1012-1018. Cairns, E., van Huystee, R. B., and Cairns, W. L. (1980). Physiol. Plant. 49,78-82. Carpita, N. C. (1984). Planr Physiol. 76,205-212. Cassab, G . I., and Varner, J. E. (1987). J. Cell Biol. 105,2581-2588.

264

TAKAYUKI HOSON

Cassab, G. I., and Varner, J. E. (1988). Annu. Rev. Plant Physiol. Plant Mol. Biol. 39, 321-353.

Chandra Sekhar, K. N., and DeMason, D. A. (1990). Planta 181,53-61. Chibbar, R. N., and van Huystee, R. B. (1986). J. Plant Physiol. 125,477-486. Chrispeels, M. J. (1%9). Plant Physiol. 44, 1187-1193. Christoffersen, R. E., Tucker, M. L., and Laties, G. G. (1984). Plant Mol. Biol. 3,385-391. Clarke, A. E., Anderson, R. L., and Stone, B. A. (1979). Phytochernistry 18,521-540. Cleland, R. (1971). Annu. Rev. Plant Physiol. 22,197-222. Cohn, N. S., Mitchell, J. P., and Zhou, Z. (1989). Plant Sci. 60, 137-143. Conrad, T. A., Lamport, D. T. A., and Hammarschmidt, R. (1987). Plant Physiol. 83, 1-3. Cooper, J. B., Chen, J. A., and Varner, J. E. (1984). I n “Structure, Function, and Biosynthesis of Plant Cell Walls” (W. M. Dugger and S. Bartnicki-Garcia, eds.), pp. 75-88. Am. SOC.Plant Physiol., Rockville, Maryland. Corbin, D. R., Sauer, N., and Lamb, C. J. (1987). Mol. Cel. Biol. 7,4337-4344. Cosgrove, D. J. (1989). Absrr. Cell Wall Meet. 5th abstr. 1. Darvill, A., Smith, C. J., and Hall, M. A. (1978). New Phyrol. 80,503-516. Darvill, A., McNeil, M., Albersheim, P., and Delmer, D. P. (1980). I n “The Biochemistry of Plants” (N. E. Tolbert, ed.), Vol. 1, pp. 91-162. Academic Press, New York. Dashek, W. V., and Chrispeels, M. J. (1977). Planta W, 251-256. del Campillo, E., Durbin, M., and Lewis, L. N. (1988). Plant Physiol. 88,904-909. del Campillo, E., Reid, P. D., Sexton, R., and Lewis, L. N. (1990). Plant Cell 2,245-254. DellaPenna, D., and Bennett, A. B. (1988). Plunt Physiol. 86, 1057-1063. DellaPenna, D., Alexander, D. G.,and Bennett, A. B. (1986). Proc. Natl. Acad. Sci. U.S.A. 83,6420-6424.

Dey, P. M., and Brinson, K. (1984). Adv. Carbohydr. Chem. Biochem. 42,265-382. Dixon, R. A., and Lamb, C. J. (1990). Annu. Rev. Plant Physiol. Plant Mol. Biol. 41, 339-367.

Durbin, M. L., Sexton, R., and Lewis, L. N. (1981). Plant Cell Enuiron. 4,67-73. Esaka, M., Enoki, K., Kouchi, B., and Sasaki, T. (1990). Plant Physiol. 93, 1037-1041. Espelie, K. E., and Kolattukudy, P. E. (1985). Arch. Biochem. Biophys. 240,539-545. Espelie, K. E., Franceschi, V. R., and Kolattukudy, P. E. (1986). Plant Physiol. 81,487-492. Etzler, M. E. (1985). Annu. Rev. Plant Physiol. 36,209-234. Evans, P. T., Holaway, B. L., and Malmberg, R. L. (1988). Planta 175,259-269. Felix, G . , and Meins, F., Jr. (1985). Plantu 164,423-428. Fincher, G . B. (1989). Annu. Rev. Plant Physiol. Plant Mol. Biol. 40,305-346. Fincher, G . B., Stone, B. A., and Clarke, A. E. (1983). Annu. Rev. Plant Physiol. 34,47-70. Fink, J., Jeblick, W., and Kauss, H. (1990). Planta 181,343-348. Fry, S . C. (1986). Annu. Rev. Plant Physiol. 37, 165-186. Fry, S. C. (1988). “The Growing Plant Cell Wall: Chemical and Metabolic Analysis.’’ Longman, Essex, England. Fry, S. C. (1989). J. Exp. Bot. 40, 1-11. Fuchs, Y., Saxena, A., Gamble, H. R., and Anderson, J. D. (1989). Plant Physiol. 89, 138- 143.

Gilkes, N. R., and Hall, M. A. (1977). New Phyrol. 78, 1-15. Giovannoni, J. J., DellaPenna, D., Bennett, A. B., and Fischer, R. L. (1989). Plant Cell 1, 53-63.

Grief, C., and Shaw, P. J. (1987). Planta 171,302-312. Grierson, D., Slater, A., Speirs, J., and Tucker, G. A. (1985). Planta 163,263-271. Grierson, D., Smith, C. J. S., Moms, P. C., Watson, C., Bird, C. R., Ray, J., Schuch, W., Knapp, J. E., Davies, K., and Picton, S. J. (1989). I n “Cell Separation in Plants” (D. J. Osborne and M. B. Jackson, eds.), pp. 1-9. Springer-Verlag. Berlin.

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

265

Griffing, L. R., and Fowke, L. C. (1985). Protoplasma 128,22-30. Grosset, J., Meyer, Y., Chartier, Y., Kauffman, S., Legrand, M., and Fritig, B. (1990). Planr Physiol. 92, 520-527. Hankins, C. N., Kindinger, J. I., and Shannon, L. M. (1979). Planr Physiol. 64, 104-107. Hatfield, R., and Nevins, D. J. (1986). Plant Cell Physiol. 27,541-552. Hatfield, R., and Nevins, D. J. (1988). Plant Cell Physiol. 29,713-720. Hayashi, T. (1989). Annu. Rev. Plant Physiol. Plant Mol. Biol. 40, 139-168. Hayashi, T., and Maclachlan, G. (1984). Plant Physiol. 75,596-604. Herman, E. M. (1988). Annu. Rev. Plant Physiol. Plant Mol. Biol. 39, 139-155. Herman, E. M., and Shannon, L. M. (1985). Plant Physiol. 77,886-890. Heyn, A. N. J. (1986). Plant Sci. 45,77-82. Horton, R., and Osborne, D. J. (1967). Nature (London) 214, 1086-1088. Hoson, T. (1990). Planr Cell Physiol. 31,281-287. Hoson, T . , and Masuda, Y. (1986). In “Cell Wall ’86” (B. Vian, D. Reis, and R. Goldberg, eds.), pp. 242-243. Groupe Parois, Paris.. Hoson, T., and Masuda, Y. (1987). Physiol. Plant. 71, 1-8. Hoson, T., and Masuda, Y. (1989). Abstr. Cell Wall Meet., 5th abstr. 132. Hoson, T., and Masuda, Y. (1991). Physiol. Plant. 82,41-47. Hoson, T . , and Nevins, D. J. (1989a). Planr Physiol. 90, 1353-1358. Hoson, T., and Nevins, D. J. (1989b). Physiol. Plant. 75,452-457. Hoson, T., and Nevins, D. J. (1989~).Physiol. Plant. 77,208-215. Hoson, T., Masuda, Y., Sone, Y., and Misaki, A. (1991a). Planr Physiol. 96,551-557. Hoson, T., Masuda, Y., and Nevins, D. J. (1991b). Plant Physiol. (in press). Hu, C., Carbonera, D., and van Huystee, R. (1987). Plant Physiol. 85,299-303. Huber, D. J. (1983). Hortic. Rev. 5, 169-219. Huber, D. J., and Nevins, D. J. (1979). Plant Cell Physiol. 20,201-212. Huber, D. J., and Nevins, D. J. (1981). Physiol. Plant 53,533-539. Iki, K., Sekiguchi, K., Kurata, K., Tada, T., Nakagawa, H., Ogura, N., and Takehana, H. (1978). Phytochemistry 17,311-312. Inouhe, M., and Nevins, D. J. (1991a). Plant Physiol. 96,285-290. Inouhe, M., and Nevins, D. J. (1991b). Planr Physiol. 96,426-431. Inouhe, M., Yamamoto, R., and Masuda, Y. (1984). Plant Cell Physiol. 25, 1341-1351. Jarvis, M. C. (1984). Plant Cell Environ. 7, 153-164. Jermyn, M. A., and Yeow, Y. M. (1975). Ausr. J . Plant Physiol. 2,501-531. Kaku, H., Shibata, S., Satsuma, Y., Sone, Y., and Misaki, A. (1986). Phytochemistry 25, 2041-2047.

Kato, Y., and Matsuda, K. (1976). Plant Cell Physiol. 17, 1185-1198. Kato, Y., Iki, K., and Matsuda, K. (1981). Agric. Biol. Chem. 45,2745-2753. Keefe, D., Hinz, U., and Meins, F., Jr. (1990). Planra 182,43-51. Keller, B., Templeton, M. D., and Lamb, C. J. (1989). Proc. Natl. Acad. Sci. U.S.A. 86, 1529-1533.

Kieliszewski, M., and Lamport, D. T. A. (1986). Phytochemistry 25,673-677. Kieliszewski, M., and Lamport, D. T. A. (1987). Plant Physiol. 85,823-827. Kieliszewski, M., Leykam, J. F., and Lamport, D. T. A. (1990). Plant Physiol. 92, 316326.

Kim, S.-H., Terry, M. E., Hoops, P., Dauwalder, M., and Roux, S. J. (1988). Plant Physiol. 88, 1446-1453. Kim, S.-H., Shinkle, J. R., and Roux, S. J. (1989). Proc. Natl. Acad. Sci. U.S.A. 86, 9866-9870.

Knee, M. (1989). In “Cell Separation in Plants” (D. J. Osborne and M. B. Jackson, eds.), pp. 145-154. Springer-Verlag, Berlin.

266

TAKAYUKI HOSON

Knox, R. B. (1982). I n “Techniques in Immunocytochemistry” (G. R. Bullock, ed.), Vol. 1, pp. 205-238. Academic Press, New York. Knox, J. P., Linstead, P. J., King, J., Cooper, C., and Roberts, K. (1990). Planta 181, 5 12-52 1.

Koehler, D. E., Lewis, L. N., Shannon, L. M., and Durbin, M. L. (1981). Phytochernistry 20, 409-412.

Kutschera, U. (1989). Physiol. Plant. 77, 157-163. Kutschera, U . , Bergfeld, R., and Schopfer, P. (1987). Planta 170, 168-180. Labavitch, J. M. (1981). Annu. Rev. Plant Physiol. 32,385-406. Labavitch, J. M., and Ray, P. M. (1974a). Plant Physiol. 53,669-673. Labavitch, J. M., and Ray, P. M. (1974b). Plant Physiol. 54,499-502. Labrador, E., and Nevins, D. J. (1989). Physiol. Plant. 77,487-492. Labrador, E., and NicolBs, G . (1985). Physiol. Plant. 64,541-546. Lamport, D. T. A. (1970). Annu. Rev. Plant Physiol. 21,235-270. Lamport, D. T. A., and Catt, J. W. (1981). I n “Plant Carbohydrates 11. Extracellular Carbohydrates” (W.Tanner and F. A. Loewus, eds.), pp. 133-165. Springer-Verlag, Berlin. Lauribre, M., Lauritre, C., Chrispeels, M. J., Johson, K. D., and Sturm, A. (1989). Plant Physiol. 90, 1182-1188. Lee, E., Speirs, J., Gray, J., and Brady, C. J. (1990). Plant Cell Environ. W, 513-521. Lee, S.-H., Kivilaan, A,, and Bandurski, R. S. (1967). Plant Physiol. 42,968-972. Lewis, N. G., and Yamamoto, E. (1990). Annu. Rev. Plant Physiol. Plant Mol. Biol. 41, 455-4%.

Lewis, L. N., Lew, F. T., Reid, P. D., and Barnes, J. E. (1972). In “Plant Growth Substances 1970” (D. J. Carr, ed.), pp. 234-239. Springer-Verlag, Berlin. Liners, F., Letesson, J.-J., Didembourg, C., and Van Cutsem, P. (1989). Plant Physiol. 91, 1419-1424.

Lis, H., and Sharon, N. (1986). Annu. Rev. Biochem. 55,35-67. Lloyd, C. W. (1987). Annu. Rev. Plant Physiol. 38, 119-139. Lobarzewski, J., and van Huystee, R. B. (1982). Plant Sci. Lett. 26,39-46. Loescher, W., and Nevins, D. J. (1972). Plant Physiol. 50,556-563. Ludevid, M. D., Ruiz-Avila, L., Vallks, M. P., Stiefel, V., Torrent, M., Torn& J. M., and Puigdomtnech, P. (1990). Planta 180,524-529. Luttenegger, D. G., and Nevins, D. J. (1985). Plant Physiol. 77, 175-178. Lynes, M., Lamb, C. A., Napolitano, L. A., and Stout, R. G. (1987). Plant Sci. 50,225-232. Masuda, Y. (1978). Bot. Mag., Spec. Issue 1, 103-123. Masuda, Y. (1990). Bot. Mag. 103,345-370. Masuda, Y., and Yamamoto, R. (1985). In “Biochemistry of Plant Cell Walls” (C. T. Brett and J. R. Hillman, eds.), pp. 269-300. Cambridge Univ. Press, Cambridge, England. Masuda, Y., Inouhe, M., and Yamamoto, R. (1989). I n “Plant Water Relations and Growth under Stress” (M. Tazawa, M. Katsumi, Y. Masuda, and H. Okamoto, eds.), pp. 291-297. Yamada Sci. Found., Osaka/Myu, Tokyo. Mauch, F., and Staehelin, L. A. (1989). Plant Cell 1,447-457. Mauch, F., Hadwiger, L. A., and Boller, T. (1984). Plant Physiol. 76,607-611. Mazau, D., and Esquerre-Tugaye, M.-T. (1986). Physiol. Mol. Plant Palhol. 29, 147-157. Mazau, D., Rumeau, D., and Esquerre-Tugaye, M.-T. (1988). Plant Physiol. 86,540-546. McFadden, G . I., Ahluwalia, B., Clarke, A. E., and Fincher, G. B. (1988). Planta 173, 500-508.

McManus, M. T.,McKeating, J., Secher, D. S., Osborne, D. J., Ashford, D., Dwek, R. A., and Rademacher, T. W. (1988). Planta 175,506-512.

IMMUNOLOGICAL APPROACHES TO PLANT CELL WALLS

267

McNeil, M., Darvill, A. G., Fry, S. C., and Albersheim, P. (1984). Annu. Reu. Biochem. 53, 625-663. McQueen-Mason, S . , Durachko, D. M., and Cosgrove, D. J. (1990). Plant Physiol., Suppl. 93,75. Melan, M. A., and Cosgrove, D. J. (1988). Plant Physiol. 86,469-474. Misaki, A., Kaku, H., and Sone, Y. (1988). Carbohydr. Res. 173, 133-144. Moore, P. J., and Staehelin, L. A. (1985). Planta 174,433-445. Moore, P. J., Darvill, A. G., Albersheim, P., and Staehelin, L. A. (1986). Plant Physiol. 82, 787-794. Morre, D. J. (1968). Plant Physiol. 43, 1545-1559. Morrow, D. L., and Jones, R. L. (1986). Physiol. Plant. 67,397-407. Mundy, J., and Fincher, G. B. (1986). FEBS Lett. 198, 349-352. Neale, A. D., Wahleithner, J. A., Lund, M., Bonnett, H. T., Kelly, A., Meeks-Wagner, D. R., Peacock, W. J., and Dennis, E. S. (1990). Plant Cell 2,673-684. Nevins, D. J., Yamamoto, R., and Huber, D. J. (1978). Phytochemistry 17, 1503-1505. Nevins, D. J., Hatfield, R., Hoson, T., and Inouhe, M. (1987). In “Physiology of Cell Expansion during Plant Growth” (D. J. Cosgrove and D. P. Knievel, eds.), pp. 122-132. Am. SOC.Plant Physiol., Rockville, Maryland. Nishitani, K., and Masuda, Y. (1980). Plant Cell Physiol. 21, 169-181. Nishitani, K., and Masuda, Y. (1981). Physiol. Plant 52,482-494. Nishitani, K., and Masuda, Y. (1983). Plant Cell Physiol. 24,345-355. Nock, L. P., and Smith, C. J. (1987). Plant Physiol. 84, 1044-1050. Norman, P. M., Wingate, V. P. M., Fitter, M. S., and Lamb, C. J. (1986). Planta 167, 452-459. Northcote, D. H., Davey, R., and Lay, J. (1989). Planta 178,353-366. Ouchterlony, 0 . (1949). Acta Pathol. Microbiol. Scand. 26,507-519. Pressey, R. (1988). Planta 174,39-43. Preston, R. D. (1979). Annu. Rev. Plant Physiol. 30,55-78. Quesada, M. A., Tigier, H. A., Bukovac, M. J., and Valpuesta, V. (1990). Physiol. Plant. 79, 623-628. Reid, P. D., del Campillo, E., and Lewis, L. N. (1990). Plant Physiol. 93, 160-165. Roberts, J. A., Taylor, J. E., Lasslett, Y.V., and Tucker, G. A. (1989). In “Cell Separation in Plants” (D. J. Osborne and M. B. Jackson, eds.), pp. 61-68. Springer-Verlag, Berlin. Roberts, K., Phillips, J., Shaw, P., Grief, C., and Smith, E. (1985). In “Biochemistry of Plant Cell Walls” (C. T. Brett and J. R. Hillman, eds.), pp. 125-154. Cambridge Univ. Press, Cambridge, England. Ruel, K., and Joseleau, J.-P. (1984). Histochemistry 81,573-580. Sakurai, N., and Masuda, Y. (1977). Plant Cell Physiol. 18,587-594. Sakurai, N., and Masuda, Y. (1978). Plant Cell Physiol. 19, 1217-1223. Sato, T., Kusaba, S., Nakagawa, H., and Ogura, N. (1984). Plant Cell Physiol. 25, 10691071. Satoh, S., and Fujii, T. (1988). Planta 175,364-373. Sauer, N., Corbin, D. R., Keller, B., and Lamn, C. J. (1990). Plant Cell Environ. W, 257-266. Schlumbaum, A., Mauch, F., Vogeli, U., and Boller, T. (1986). Nature (London) 324, 365-367. Seara, J . , NicolAs, G., and Labrador, E. (1988). Physiol. Plant. 72,769-774. Sexton, R., and Roberts, J. A. (1982). Annu. Rev. Plant Physiol. 33, 133-162. Sexton, R., Durbin, M. L., Lewis, L. N., and Thomson, W. W. (1980). Nature (London)283, 873-874.

268

TAKAYUKI HOSON

Sexton, R., del Campillo, E., Duncan, D., and Lewis, L. N. (1990). Plant Sci. 67,169-176. Shinshi, H., Mohnen, D., and Meins, F., Jr. (1987).Proc. Natl. Acad. Sci. U.S.A. 84,89-93. Smith, C. J. S., Watson, C. F., Ray, J., Bird, C. R., Moms, P. C., Schuch, W., and Grierson, D. (1988). Nature (London)334,724-726. Smith, E., Roberts, K., Hutchings, A., and Galfre, G. (1984). Planta 161,330-338. Sone, Y., Misaki, A., and Shibata, S. (1989a). Carbohydr. Res. 191,79-89. Sone, Y., Kuramae, J., Shibata, S., and Misaki, A. (1989b). Agric. Biol. Chem. 53, 28212823. Sone, Y., Shibata, S., and Misaki, A. (1990). Abstr. Inr. Workshop Front. Plant Microb. Glycans p. 25. Spruce, J., Mayer, A. M., and Osborne, D. J. (1987). Phytochemistry 26,2901-2903. Stafstrom, J. P., and Staehelin, L. A. (1988). Planta 174,321-332. Stephan, D., and van Huystee, R. B. (1980). Can. J. Biochem. 58,715-719. Stiefel, V., Perez-Grau, L. L., Albericio, F., Giralt, E., Ruiz-Avila, L., Ludevid, M. D., and Puigdomhech, P. (1988). Plant Mol. Biol. 11,483-493. Stiefel, V., Ruiz-Avila, L., Raz, R., Vallts, M. P., Gomez, J., Pages, M., MartinezIzquierdo, J. A., Ludevid, M. D., Landale, J. A., Nelson, T., and Puigdomhech, P. (1990). Plant Cell 2,785-793. Stone, B. A. (1984). In “Structure, Function, and Biosynthesis of Plant Cell Walls” (W. M. Dugger and S. Bartnicki-Garcia, eds.), pp. 52-74. Am. SOC.Plant Physiol., Rockville, Maryland. Stuart, D. A., and Varner, J. E. (1980). Plant Physiol. 66,787-792. Stuart, I. M., Loi, L., and Fincher, G. B. (1986). Plant Physiol. 80,310-314. Taiz, L. (1984). Annu. Rev. Plant Physiol. 35,585-657. Taiz, L., and Honigman, W. A. (1976). Plant Physiol. 58,380-386. Takeuchi, Y., Yoshikawa, M., Takeba, G., Tanaka, K., Shibata, D., and Horino, 0. (1990). Plant Physiol. 93,673-682. Terry, M. E., and Bonner, B. A. (1980). Plant Physiol. 66,321-325. Terry, M. E., Jones, R. L., and Bonner, B. A. (1981). Plant Physiol. 68,531-537. Tieman, D. M., and Handa, A. K. (1989). Plant Physiol. 90, 17-20. Tsumuraya, Y., Nakamura, K., Hashimoto, Y., and Yamamoto, S. (1984). Agric. Biol. Chem. 48,2915-2917. Tucker, G. A., and Grierson, D. (1982). Planta 155,64-67. Tucker, M. L., and Laties, G. G. (1984). Plant Physiol. 74,307-315. Tucker, G . A,, Robertson, N. G., and Grierson, D. (1980). Eur. J . Biochem. 112,119-124. Tucker, M. L., Sexton, R., del Campillo, E. and Lewis, L. N. (1988). Plant Physiol. 88, 1257- 1262. van Huystee, R. B. (1987). Annu. Rev. Plant Physiol. 38,205-219. van Huystee, R. B., and Lobarzewski, J. (1982). Plant Sci. Lett. 27,59-67. van Huystee, R. B., and Maldonado, B. (1982). Physiol. Plant. 54,88-92. Vogeli, U., Meins, F., Jr., and Boller, T. (1988). Planta 174, 364-372. Ward, M. R., Tischner, R., and Huffaker, R. C. (1988). Plant Physiol. 88, 1141-1145. Weiser, R. L., Wallner, S. J., and Waddell, J. W. (1990). Plant Physiol. 93, 1021-1026. Woessner, J. P., and Goodenough, U. W. (1989). Plant Cell 1,901-911. Woodward, J. R., and Fincher, G . B. (1982). Eur. J. Biochem. 121,663-669. Yamamoto, R., Shinozaki, K., and Masuda, Y. (1970). Plant Cell Physiol. 11,947-956. Zarra, I . , and Masuda, Y. (1979). Plant Cell Physiol. 20, 1125-1133.

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 130

Biologically Localized Firefly Luciferase: A Tool to Study Cellular Processes CLAUDEAFLALO Department of Biochemistry, The Weizmann Institute of Science, Rehovot 76100, Israel

I. Introduction Metabolic processes in 'situmust be coordinated and respond efficiently to global changes in the environment. They are often studied using average intracellular concentrations of enzymes, reactants, and intermediates measured by destructive methodologies, or in cell-free systems in which the original cellular organization has been artificially randomized. In both approaches, a homogeneous distribution of freely diffusible soluble components (enzymes and metabolites) is implicitly assumed. Consequently, the involvement of physical processes and heterogeneous distribution of enzymes in cellular metabolism has been considerably overlooked. It is increasingly realized that the spatial organization of cellular systems must play a major role in their macroscopic behavior (reflected in conventional studies). Although considerable structural information is accumulating in support of a high degree of organization, it has not yet been integrated and translated into comprehensive mechanistic representations at the functional level. This is hindered by the experimental difficulties in measuring the concentrations of metabolites nonhomogeneously distributed in the crowded intracellular milieu, and localized processes can only be inferred by indirect experimental evidence. Thus, the design of localized probes should contribute exclusive information and help to better understand the complex machineries involved in cellular functions. The purpose of this chapter is to introduce a new approach to studying metabolic processes in cellular environments by using firefly,luciferase (FL) as a general probe for protein assembly and metabolism, on one hand, and to monitor local bioenergetic metabolite concentrations, on the other. Both these topics are relevant to the broader context of cellular structurefunction relationships. The working hypothesis is that neither proteins nor metabolites behave in the cell as in isolated systems (near-ideal behavior in solution), but are rather channeled between their respective sites of production and operation (or utilization), which are localized in the cell. Firefly luciferase catalyzes the emission of light in the presence of ATP, firefly luciferin (LHz), and molecular oxygen with a very high quantum 269

Copyright 0 1991 by Academic m s s , Inc. All rights of reproduction in any form RSeNed.

270

CLAUDE AFLALO

yield. Initial studies have demonstrated the applicability of local [ATPI probing in artificial systems. The next step involves the introduction of the enzyme into cells so it will be directed to specific compartments, using genetic engineering methodology. The first system investigated is yeast mitochondria, with FL targeted to the cytoplasmic face of the outer membrane. In this chapter, different concepts and approaches used in the multidisciplinary study of cell metabolism are presented, in addition to an updated review on various aspects of FL structure and mechanisms relevant to its use as a probe. The high level of organization in the cell is achieved by complex mechanisms for the recognition and assembly of functional structures. This may result in the tightly controlled and coordinated metabolic fluxes which characterize the process of life itself. Effort has been made to introduce and develop each topic separately and show the potential of its use in coordination. 11. Cellular Biogenesis and Organization

A detailed description of these fields is beyond the scope of this chapter. It is nevertheless necessary to define the subject in its proper context.

A. PROTEIN TRAFFIC AND ASSEMBLY Most organelle-specific proteins in eukaryotic cells are encoded in the nucleus and synthesized in the cytoplasm as precursors directed to their target compartment. The precursors must reach a new compartment distinct from the site of their synthesis before being functional. This process involves the energy-dependent translocation of polypeptide chains through one or more biological membranes, and probably requires unfolding of the chains. During or after translocation, the precursors are sometimes processed and further assembled into functional proteins. The correct localization is attained through the specific recognition of signal sequences on the precursor (Blobel, 1983) by a complex molecular machinery, under current identification and characterization (Pain et ul., 1988; Vestweber et al., 1989; Rassow et al., 1989; Hinekgt ql., 1990). In many instances, it has been possible to identify discrete sequences which determine the addressing of the various polypeptides. Targeting sequences do not seem to present extensive homology in their primary sequence, but are characterized by specific residue composition and their potential for folding into defined secondary structures (von Heijne, 1986; Roise and Schatz, 1988). Such signal sequences are often necessary and sufficient to

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

27 1

target foreign soluble proteins to the correct location in cells (Hurt et al., 1984, 1985; Vijaya et al., 1988). The localization of genetically engineered chimeric proteins in which signal peptides have been fused to soluble polypeptides, extensively used to characterize the signal sequences, also provides essential information on the molecular machinery for ultrastructure assembly. While most of the targeting information is contained in relatively short sequences, it is expected that structural restrictions should apply to the rest of the protein for its correct import and assembly. Again, the use of fusion genes and their chimeric products represents a versatile tool for studying the interaction of the precursor with the import machinery and the structural requirements for assembly in the correct topology (Hurt et al., 1985; Froshauer et al., 1988). The need for an import-competent conformation of the precursors is also illustrated by the dependence of translocation on phosphodiester bond (ATP or GTP) hydrolysis in the cytosolic phase. It has been shown that the tight folding of proteins through binding of ligands (Eilers and Schatz, 1986; Chen and Douglas, 1987) or disulfide bridges (Vestweber and Schatz, 1988) strongly impairs the translocation of precursors after the binding (and internalization)of the “leader peptide.” Nevertheless, the targeting and import machinery is surprisingly tolerant toward the molecular species fused to leader peptides. In a limited number of cases, correct targeting of such chimeric polypeptides has been achieved in highly purified cell-free systems (On0 and Tuboi, 1988). It seems, however, that this “self-assembly” does not generally occur at high efficiency and with strict specificity for the correct target (Pfanner et al., 1988). In many cases, the assistance of extrinsic factors (i.e., other than the protein and isolated organelles) is needed. Molecular “chaperones” (Deshaies et al., 1988; Ellis and Hemmingsen, 1989) play a central role in maintaining the precursors in a proper conformation until their “delivery” to the specific receptors on the target. They represent a range of cytoplasmic and organelle-specific proteins, also identified as heat-shock or stress proteins, whose role is to prevent and/or reverse faulty interactions between cytoplasmic precursors (or partially unfolded mature proteins under stress) and exposed surfaces in the cell and assist in their processing during or after translation. At least two main routes for permanent protein topogenesis can be described, based on the relationship in time between translation in the cytoplasm and the early events of targeting. In the secretory pathway (Walter and Lingappa, 1986), proteins are generally cotranslationally translocated into the lumen of endoplasmic reticulum (ER), in which they are further processed (proteolysis and/or covalent modification) and

272

CLAUDE AFLALO

sorted on the way to their final assembly in the ER, Golgi apparatus, lysosomes (Johnson et al., 1987; or vacuoles in yeast, Rothman et al., 1989), plasma membrane (Vijaya et al., 1988), or secretory vesicles. In the second mode, precursors are fully translated on free polysomes and then directed to the nucleus (Kalderon et al., 1984), mitochondria (Douglas et al., 1986; Grivell, 1986; Hart1 and Neupert, 1989), chloroplasts (Smeekens et al., 1986; Keegstra, 1989), and peroxisomes (Gould et al., l989,1990a,b; Lazarow and Fujiki, 1985). The direction of precursors to the correct (sub) compartment is done in uiuo at extremely high fidelity, in spite of the apparent similarity among the various signal sequences. Moreover, it seems that the mechanism for direction and import has been conserved along the evolutionary scale, so that yeast mitochondria1 precursors, translated in uitro using rabbit cytoplasmic factors, are readily imported into rat liver mitochondria (On0 and Tuboi, 1988). Some reports cited above present evidence for the redirection of natural or engineered precursors to different targets in uitro, raising the question of hierarchy between signal sequences (Colman and Robinson, 1986). The pathways just described lead to permanent localization of proteins in the various compartments in the cell. However, a number of proteins lacking obvious targeting or trans-membrane sequences show tight detergent-sensitive binding to membranes. This attachment is dependent on covalent posttranslational modification of the polypeptides with lipidcontaining moieties (Magee et al., 1989). This would enable a transient association of the protein with the membrane subject to control. Another type of transient localization of proteins may occur through proteinprotein interactions solely, when one of the components is itself permanently localized. €3. STRUCTURAL BASISFOR THE ORGANIZATION OF METABOLISM

Stable assembly of distinct catalytic units is recognized at different levels in the cell. Some examples are membrane-enclosed organelles and multienzyme complexes (e.g., pyruvate or succinate dehydrogenase), in which metabolic processes take place in relative isolation from other cellular compartments. Thus, the general relationship between such cellular structures and their metabolic function is well established. However, even at the level of single subcellular units as familiar as the mitochondrion, a wide gap exists between their actual structure and that assumed in the early mechanisms proposed for their function (e.g., the chemiosmotic coupling concept). The present information available on oxidative phosphorylation and other processes in energy-transducing membranes reveals heterogeneity in function, which may reflect their complex structure. A

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

273

considerable amount of experimental evidence does not support a simple delocalized proton-coupling mechanism (Westerhoff et al., 1984), and various modifications to the simple model involving thermodynamic (slips) or kinetic (allosteric regulation) effects have been postulated. However, most discrepancies may be explained by an effective microcompartmentation of protons and catalytic units in the absence of physical barriers (Westerhoff et al., 1988). This view may be extended to the wider context of cellular metabolism. The cytoplasm was traditionally considered as a concentrated solution (Fig. 1A) harboring delocalized processes. However, it contains a large amount of surfaces (membranes and cytoskeletal elements) to which enzymes or larger structures present at high concentrations could reversibly adsorb and thus form functional units, or metabolons (Srere, 1987; Kaprelyants, 1988). The occurrence of such units has been inferred for various catalytic pathways from a nonrandom distribution of the participating enzymes (Masters, 1981; Clegg, 1984), and some peculiarities of the kinetic behavior in some metabolic pathways (discussed in Section 11,C). Macromolecularassociation, while greatly enhanced by molecular crowding (Minton, 1983) as a general cellular phenomenon (Ottaway and Mowbray, 1977), often occurs selectively between functionally related soluble species (e.g., consecutive enzymes in a pathway). The specificity of the attachment may be achieved through recognition sites located at the periphery of proteins, as described for chiral-specific dehydrogenases (Srivastava et al., 1985). In addition, proximity can be achieved by adsorption to a common (restricted) surface, as illustrated in Fig. 1B. The association between the interacting catalytic and/or structural species is often modulated by the occupancy of specific binding sites by lowmolecular-mass substrates, products, or effectors. These facts have led to the proposal that the sequential reactions in a metabolic pathway may occur as a “relay at the surface” in which an adsorbed enzyme successively binds its substrate, dissociates, transfers its product to the next vicinally adsorbed enzyme, and reassociates with the surface (Ryazanov and Spirin, 1989). A direct consequence of the colocalization of catalytic systems in a limited space is the parallel occlusion of their intermediates at these loci. This is supported by the relatively high enzyme : intermediate concentration ratio, determined experimentallyfor a wide variety of metabolic pathways (Srere, 1987; Srivastava and Bernhard, 1986a,b),and by considering that most cellular water might be adsorbed to cellular surfaces or under their influence (see Clegg, 1984). The concept of heterogeneous distribution for macromolecules described above may be extended to a higher level and involve subcellular structures. Physical association between chloroplasts, mitochondria, and

274

CLAUDE AFLALO

B

A

Dialysis-bag model

Surface adsorption model

FIG. 1. Two views of cellular organization. The cellular components schematically represented are various organelles, proteins, and metabolites (A) and (0).(A) The dotted background indicates aqueous solution, in which organelles and molecules are homogeneously distributed as traditionally assumed in mechanistic studies. (B) In the modem view, the components are adsorbed on the membranal surfaces and the cytoskeleton is represented by a lattice.

peroxisomes has led to the proposal of their cooperation in the oxidative photosynthetic carbon cycle of photorespiration in leaves, while mitochondria and glyoxysomes apparently directly exchange metabolites of fatty acid p-oxidation in endosperm (Tolbert, 1981). A current view is that organelles can bind through microtubule-associated proteins to the cytoskeleton (Hirokawa, 1982). The latter consists of microtubuli (tubulin filaments) interconnected by the F-actin microtrabecular lattice (Porter, 1984; Schliwa et al., 1981). The formation and dissociation of cytoskeletal filaments are vectorial energy-dependent processes (Carlier, 1989), and thus are subject to metabolic control. The influence of the metabolic status on the cytoskeleton has been postulated from the possible participation of several glycolytic enzymes in the formation of actin filaments (Morton et al., 1982; Tillmann and Bereiter-Hahn, 1986). It has been shown that cytoplasmic factors effect the unidirectional motion of organelles upon the cytoskeletalfibers (Vale, 1987). This translocation activity has been reconstituted in uitro with various purified filaments, organelles (or latex beads), and molecular motors. The direction of the motion is determined by two factors: the intrinsic polarity of the cytoskeletalfilaments and the nature of

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

275

the motor bound to specific receptors at the surface of the organelle. While such a cytoskeleton-effected motility of ultrastructures within large cells is perceived as an obvious need (e.g., transport of neurotransmitter vesicles in neurons and separation of chromosomes), it represents a more subtle metabolic advantage in other systems by enabling a dynamic association or segregation between functionally related systems, according to metabolic needs.

C. INVOLVEMENT OF PHYSICAL PROCESSES AND LOCAL CATALYSIS IN CELLULAR METABOLISM Enzymatic catalysis requires the binding of substrates to the active site on the enzyme, their chemical transformation, and the release of products. The actual rates of binding and release of reactants are determined by their local concentrations, or rather chemical activities, in the environment of the active site. For catalysis in well-mixed dilute aqueous solutions, there is no difference between the local (effective) concentration of reactants and that in the bulk medium which is used in the determination of the kinetic parameters of the enzymatic reaction. However, in the more realistic cases mentioned above, diffusion or mass transfer of reactants to or from the microenvironment of enzymes may kinetically limit (at steady state) the overall rate of catalysis, while heterogeneous partition of reactants (at equilibrium) may affect the observed dependence on their concentration in the bulk medium. The influence of diffusion and other physical phenomena on heterogeneous catalytic systems was first recognized and evaluated in the field of chemical engineering (see Engasser and Horvath, 1976). Using immobilized enzymes as a model (see Mosbach, 1978), Katchalski and co-workers (Goldman and Katchalski, 1971) developed a theoretical approach to heterogeneous catalysis in biological systems (see Goldstein, 1976). This pioneering work has uncovered the biological implications of diffusion control in cellular metabolism. The extent of diffusion control is determined essentially by the ratio of the rate of diffusion (diffusivity and length of path) to that of catalysis. The lower this ratio, the further the observed kinetic behavior diverges from that in well-stirred solution in terms of apparent affinities for reactant and also transient kinetics (Easterby, 1981). With diffusion limitation, it is expected that substrates and products will be depleted and accumulated, respectively, in the microenvironment of the immobilized enzyme, relative to the bulk medium at steady state (see Boag, 1969, for comprehensive treatment for oxygen in tissues). On the other hand, the relative substrate depletion [i.e. (SlOcd- Sbulk)/Sbulkl will determine the extent of departure of the macroscopic kinetic behavior

276

CLAUDE AFLALO

from that in solution. The occurrence of such concentration gradients has been associated with an increased efficiency of coimmobilized coupled enzymes, compared to that observed with the same system in solution (Mattiasson and Mosbach, 1971; Goldman, 1973). In a concentrated enzymatic phase, the accumulated product of a first reaction is more accessible to the second enzyme than in solution, leading to the concept of tunneling or channeling of intermediates and its quantitative evaluation (Kurganov, 1986; Keleti and Ovadi, 1988; Welch et al., 1988). These features represent the basis for modeling (Kernevez, 1980) a compartmentalized metabolism in uiuo (see Ottaway and Mowbray, 1977), in view of the various possibilities for microcompartmentation or physical association between “soluble” enzymes as mentioned above. These considerations are very relevant to the study of the mechanism of membrane-associated enzymes in isolated systems, as illustrated in the following example. In the photophosphorylation of ADP by isolated thylakoids, ATP is found tightly bound to ATP synthase in the presence of excess hexokinase added to the medium. This has been interpreted as entirely due to its occupancy of the active site as a catalytic intermediateof phosphorylation (Rosen er al., 1979). However, there is strong experimental evidence suggesting that the newly released ATP accumulates at steady state near the thylakoid membranes during photophosphorylation and can rebind to the ATP synthase before it becomes accessible to exogeneous hexokinase or equilibrates with exogeneous ATP added to the medium (Aflalo and Shavit, 1982). The involvement of diffusion limitations was also inferred from the increased apparent affinity of ATP synthase to ADP following hypotonic shock of thylakoids, with no effect on the total activity (Aflalo and Shavit, 1983, 1984). The limitation was relieved upon artificial reduction of the catalytic rate, as expected in a diffusion control situation. Therefore, the physical-as opposed to catalytic-steps may represent alternative limiting factors in the macroscopic behavior of enzymes sequestered in a restricted environment. The analysis of macroscopic data, while accurately describing some effective properties of the “mechanistic black box” investigated, yields kinetic parameters inherent to the heterogeneous system, but not the microscopic ones intrinsic to the catalytic components operating in situ. These aspects of research suffer from the lack of more direct ways to demonstrate and quantitate the effects of physical steps on the observed kinetic behavior in nonhomogeneous catalysis. Considering the high enzyme : intermediate concentration ratio in situ, and the respective dissociation constants, it is plausible that most metabolites in the cell occur as bound species (Srere, 1987). There is now strong evidence that the transfer of intermediates (I) between two soluble en-

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

277

zymes (El and E2) carrying out coupled reactions might occur directly between the enzymes in a transient complex (EI-I-E~)rather than through a release-diffusion-binding sequence. Bernhard and co-workers demonstrated experimentally the direct transfer of NAD(H) between selected cytoplasmic dehydrogenases in uitro under proper conditions (Srivastava and Bernhard, 1986a). This was done by showing that the complex El-I represents a preferred substrate for E2 as compared to free I in solution, provided that the binding of I by El and E2 is of opposite chirality (Srivastava et al., 1985;Srivastava and Bernhard, 1986b).It was also shown that I in the preformed El-I complex is transferred faster to E2 than released in solution (Srivastava and Bernhard, 1987). These conclusions were extended to mitochondrial dehydrogenases (Fukushima et al., 1989) and proposed as a general cellular phenomenon (Ryazanov and Spirin, 1989). Such a direct and selective transfer mechanism may insure channeling of intermediates from their site of production to that of utilization, so they will not be scavenged by concurrent systems during their diffusion as free species. At the cellular level, the occurrence of asymmetry in the distribution of structures and its relation with asymmetry in function were first postulated with the discovery of distinct pools for glycine in kidney cells (Garfinkel, 1963; Garfinkel and Lajtha, 1963). Since then, a considerable amount of evidence has been presented in favor of this principle for various metabolic pathways catalyzed by cytoplasmic components previously assumed to be homogeneously distributed. Functional compartmentation of various pathways for energy transformation has been demonstrated in different biological systems as transient association between cytosolic ATPproducing reactions and endergonic processes known to be localized in the cell (Lynch and Paul, 1987).The best-known example is the formation of a glycolytic complex (Masters, 1981) and its association with the plasma membrane or cytoskeletal elements (Knull, 1990).The occurrence of metabolic units spanning multiple compartments separated by biological membranes has also been proposed. For example, cooperation between mitochondrial [matrix and (inter)membranal spaces] and cytosolic enzymes has been described for the urea cycle (Cheung et al., 1989)and the creatine phosphate shuttle between mitochondria and myofibrils (Bessman and Carpenter, 1985). Although these associations often occur at relatively stable infrastructures [e.g., contact sites between mitochondrial membranes (Brdiczka et al., 1990)], they may be transient and responsive to metabolic status. The reversible dissociation of these complexes is often modulated by substrate or effector binding. On the other hand, the association of an enzyme to another structure may effect its catalytic behavior through allosteric cooperative effects between the association site and the

278

CLAUDE AFLALO

catalytic site. This “ambiquity” of proteins is expected to have a reciprocal effect on cellular function by forcing metabolic fluxes in selected directions (Wilson, 1988). The selectivity may be achieved by reducing diffusional limitation in the sequence catalyzed by the enzymes in a complex, as compared to that occurring between enzymes delocalized in solution, or physically segregated into separate microcompartments. Channeling of intermediates in metabolic pathways has been assessed by their unavailability to isotopic dilution by exogenous metabolites (Kurganov, 1986; Keleti and Ovadi, 1988). In isolated systems, radiolabeled tracers have been used to demonstrate intermediates channeling. For example, medium [‘4C]ADPis converted to [I4C]ATPat a much higher rate than that of its uptake (i.e., equilibration with the matrix pool) into energized mitochondria (Murthy and Pande, 1985). This occurs as if the phosphorylation flux is “short circuited” in a functional complex around the adenine nucleotide translocase of the inner membrane. D. USEOF ENDOGENOUS ENZYMESAS REPORTERS FOR LOCAL OF REACTANTS CONCENTRATIONS Average metabolite concentrations are commonly assessed in deproteinated crude extracts of cells or tissues using exogenous specific enzymes. A similar principle may also be applied to estimate local concentrations using endogenous enzymes naturally localized in the cell. Their activities can be measured in intact cells under experimental conditions such that the rate of catalysis will reflect the local concentration of a single metabolite (see Jones and Aw, 1990). The ideal conditions for this are defined as follows: 1. The enzyme should be far from its equilibrium, as in the case of an exergonic reaction or when a product is efficiently removed by a coupled reaction. 2. The rate of reaction should be essentially limited by the concentration of the metabolite to be determined. This implies that the concentration range for the metabolite should be substantially below the intrinsic K,,, of the enzyme. 3. The measurement should be made by noninteracting procedures such as determination of the accumulation of a nonmetabolized product, the disappearance of a saturated cosubstrate, or, most conveniently, a spectroscopic change occurring with catalysis. 4. The correlation between enzyme activity and the local concentration of its limiting substrate (or effector) should be verified independently. This can be done on the basis of the enzyme’s response in a solu-

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

279

bilized system as a first approximation, and critically reassessed in siru, considering the possible influence of structural and regulatory factors. Alternatively, one may also use enzymes at equilibrium and derive the concentration of one reactant from the knowledge of that of the others and the equilibrium constant for the catalyzed reaction. This requires that the enzyme activity is high relative to associated reactions in its vicinity. The fulfillment of all the above conditions is rather rare for any single enzyme. Nevertheless, this approach was generalized from the pioneering studies by Chance and co-workers in the noninvasive measurements of H202 and 0 2 in situ. (Seis and Chance, 1970; Chance et al., 1973; Tamura et al., 1989). A very important aspect of these approaches is that they permit estimation of metabolite concentrations at specific subcellular sites. Thus, in principle, heterogeneities in metabolites can be detected using specific distributions of enzymes among various cellular compartments or microenvironments, not necessarily separated by physical barriers (membranes). Their actual demonstration relies on considerable experimental manipulation of the investigated system. Intracellular diffusion gradients of O2and ATP have been demonstrated in intact hepatocytes (Jones, 1986) using endogenous enzymes located at different cellular sites. The average [ATP] was modulated by affecting oxidative phosphorylation and/or glycolysis. While its concentration in the cytoplasm probed by ATP sulfurylase is well correlated with the macroscopic measurements, it is substantially higher than that probed with the plasma membrane Na+,K+-ATPase, distal to mitochondria (Aw and Jones, 1985). The difference increases as the cellular ATP content is lowered by various means, all leading to the complete arrest of K+ uptake at the plasma membrane when only 60% of bulk cell ATP is depleted, in agreement with the cytoplasmic probe. This suggests complete depletion of ATP in a restricted space near the membrane (distal to mitochondria as the major ATP producer in the conditions tested) which does not contribute significantly to the average ATP content. This conclusion is further supported by modeling the system assuming diffusionalrestrictions for the transfer of ATP between inhibited mitochondria and the basal plasma membrane through an ATP-consuming volume. However, a direct effect of the factors used to lower ATP content on Na+,K+-ATPase, while improbable, cannot yet be ruled out completely. Another limitation is the relatively long time needed to determine the enzymatic activities as opposed to the quasiinstantaneous spectroscopic measurements feasible in other systems (Tamura et al., 1989). The latter are independent of the

280

CLAUDE AFLALO

diffusional restrictions which may apply in the macroscopic measurements of bulk chemical reactants used to assess local activities. In summary, the cell represents a highly structured biological unit endowed with a multitude of functions. The latter must be efficiently coordinated in order for the cell to fit in a constantly changing environment. This adaptability cannot be satisfactorily explained assuming independent functionality of the various cellular systems. Molecular mechanisms have evolved for the correct localization and assembly of complex structures which are able to mutually interact, resulting in a harmonious and efficient operation of all the physicochemical processes occurring whereby. Wide structural evidence supports the view of an organized cellular metabolism; theoretical treatments start to emerge and describe heterogeneous distribution of catalytic systems and modulation of physical factors as efficient means to control metabolism. However, a direct demonstration of localized processes is still very difficult to obtain and relies on the development of novel approaches involving sophisticated experimental and mathematical tools for the analysis of metabolism in situ. The use of FL as a biologically localized probe may represent a versatile analytical tool to implement the current research in cellular metabolism.

111. Firefly Luciferase: An Overview

A. HISTORY AND APPLICATIONS Luminescent phenomena have fascinated mankind ever since its selfconsciousness. The first known report of bioluminescence is attributed (Harvey, 1957) to Aristotle (384-322 BC). Robert Boyle (1668) contributed to its demystification by discovering that light is emitted with no perceptible heat in dependence on air (oxygen). Applying emerging biochemical approaches, Raphael Dubois (1885 and 1887) first demonstrated the involvement of organic compounds in light-emitting reactions in the firefly and the clam. He was able to restore light production by mixing two crude extracts from the same organism, and concluded that a heat-stable component (luciferine) served as a substrate to a heat-labile catalytic component (luciferuse) in the luminescent reactions occurring in each organism. The concept of specific luciferin-luciferase systems was then extended to other bioluminescent reactions from other species, and it was realized that the source of light was the low-molecular-mass compound, as suggested by the subsequent discoveries of chemiluminescent reactions in solution. The next major step in the study of the firefly bioluminescence reaction was the demonstration of the participation of Mg2+and ATP by

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

28 1

William McElroy (1947). With the purification of luciferase from the North American firefly Photinus pyralis (Green and McElroy, 1956), the study of its reaction by McElroy, joined later by Marlene DeLuca, entered the modem biochemistry and biophysics phase (DeLuca and McElroy, 1978). The partial elucidation of the enzyme’s mechanism (see Section II1,B) has served as a guideline and a source of inspiration for the understanding of bioluminescent processes in other organisms. Of the latter, bacterial luciferase (Ziegler and Baldwin, 198 I) and aequorin from coelenterate (Cormier, 1978), using flavin mononucleotide (FMNH2) and Ca2+ as cofactors, respectively, are among the best-known (and -used) luciferases. In addition to the understanding of basic aspects of energy transduction, the efforts invested in the study of bioluminescent reactions have been stimulated by the enormous potential of their use in a wide spectrum of applications (Kricka, 1988). The extremely high sensitivity of modern light-measuring devices, combined with the specificity inherent to their enzymic nature, confers to these reactions the characteristics of invaluable analytical tools (Whitehead et al., 1979). In the following section, I concentrate on FL, but similar considerations apply to other bioluminescent systems. In addition to the detection of minute amounts of ATP or oxygen by FL, the luminescent reaction can be coupled to different enzymes in order to determine the concentrations of many metabolites in biological extracts under proper experimental conditions. This versatility may be further extended to the determination of the coupled enzymes’ activity by continuous monitoring of the ATP produced or consumed by these enzymes. A major challenge of firefly luminescence is to replace radioactive or enzymatic detection systems in immunoassays. This has been promoted by the recent synthesis of luciferin derivatized at the carboxyl or phenolic residue with various substituents (Miska and Geiger, 1987). While the derivatives have no bioluminescent activity, free luciferin may be released by their enzymatic hydrolysis with an appropriate enzyme and elicit light production by added luciferase. With the advent of molecular biology, the gene of FL was first isolated and cloned in bacteria (de Wet et al., 1985);since then, it has been cloned and expressed in a variety of organisms (see Section V,A). After the determination of its sequence (de Wet et al., 1987), the applicability of the clone as a reporter of genetic events associated with transcription, translation, and assembly was soon realized (Gould and Subramani, 1988). The recent cloning and expression of new genes from the Jamaican click beetle Pyrophorus plagiophthalamus (Wood et al., 1989b) have opened the way to detailed studies of the structure-function relationships in beetle luciferases. While genetically distinct, they emit light at different colors using the same substrates.

282

CLAUDE AFLALO

B. CHARACTERIZATION OF THE ENZYME 1 . Molecular Mechanism of Light Emission

FL catalyzes a complex reaction which uses ATP, molecular oxygen, and luciferin (LH3 as substrates to produce AMP, pyrophosphate (PPi), C02, oxyluciferin (LO), and light. Firefly luciferin is a unique heterocyclic acidic chromophore found among luminous beetles only. The sequence of reactions (see Fig. 2) has been partially elucidated through the study of the enzyme response to luciferjn analogs (Seliger and McElroy, 1964), their spectroscopic properties in solution (White et al., 1980), and fast kinetic analysis of light emission (DeLuca and McElroy, 1974). In a first reversible stage, luciferin is activated by ATP in the presence of Mg2+to yield luciferyl adenylate (LH2-AMP)tightly bound to a hydrophobic pocket and free PPi. In addition, luciferase can also catalyze the reversible activation of dehydroluciferin (L) to form a tightly bound dehydroluciferyl adenylate (L-AMP) (see Fig. 2A). The latter represents a dead-end product, since it cannot participate in the subsequent oxidative step. These steps are accompanied by a large conformational change in the enzyme (DeLuca and Marsh, 1967). Light emission can be elicited in the absence of Mg2+ by the addition of synthetic LH2-AMP instead of the natural substrates (Rhodes and McElroy, 1958). The oxidative decarboxylation of the LH2-AMP occurs through a complex sequence of reactions. First, a proton is abstracted from the a-carbon, generating a carbanion to which molecular oxygen adds and transiently forms a peroxylactone with the carbonyl residue from which AMP is displaced. Next, C02 is released through electronic rearrangement and LO is generated (still tightly bound to the active site) in an electronically excited state which may exist in various tautomeric forms (Morton et al., 1969). The excited intermediate spontaneously decays to the ground state, concomitantly with the emission of a photon, whose wavelength depends on the configuration of LO as shown in Fig. 2B (enol, deprotonated: yellow-green, 560 nm; keto: red, 610 nm). In turn, physicochemical factors (e.g., low pH, high temperature, bivalent cations, and sulfhydryl reagents) may alter the conformation of the active site and lead to a shift in the color of the light output from yellow-green, under optimal conditions, to red (Seliger and McElroy, 1964).

This early postulation of the relationship between the active site and the color of the light emitted has been confirmed by the recent studies on P. plagiophthalamus luciferases, indicating that discrete and independent changes in the color emitted by naturally occurring isozymes are due to

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

283

A

L

L-AMP

B

enol: yellow-green emission

keto: red emission

LO FIG. 2. Molecular mechanism of luciferase reactions. (A) Reversible activation of dehydroluciferin by ATP. (B) Mechanism of light emission with luciferin. L, Dehydroluciferin; L-AMP, dehydroluciferyl adenylate; LO, oxyluciferin.

variation of a few amino acid residues (Wood et al., 1989b). With the sequence of luciferase at hand, the way to refine the structure-function relationship studies of the molecular mechanism is open.

284

CLAUDE AFLALO

2. Structure

FL isolated from an acetone powder of light organs has been purified to homogeneity and can be repeatedly crystallized (Green and McElroy , 1956). Unfortunately, the crystals (long needles) are unsuitable for X-ray crystallography, and attempts to get different crystals were unsuccessful. The early physical characterization of the enzyme in nondenaturating conditions indicates an apparent molecular mass of 50 kDa for the minimal catalytically active unit (Green and McElroy, 1956). In spite of its hydrophobic nature, the purified enzyme is soluble in aqueous media, but tends to aggregate at low ionic strength (DeLuca, 1969). It was commonly accepted (see Alter and DeLuca, 1986) that the active form of the enzyme occurs as a dimer (100 kDa). The basis for this comes from the apparent stoichiometries for binding of reactants (Lee et al., 1970; Lee and McElroy, 1971a), indicating “half-site reactivity” for Mg.ATP (1 : 100 kDa), while free ATP, AMP, and luciferin bind at the expected ratio (1 :50 kDa). In addition, active-site titration using L suggests that two ATPs and one L are needed to yield one L-AMP per 100 kDa of enzyme (DeLuca and McElroy, 1984). However, the calculated molecular mass of the active recombinant enzyme (550 residues) encoded by a single gene (de Wet et al., 1987) is 61 kDa. Moreover, the native enzyme and that cloned and expressed in various systems migrate at the same rate on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), in accordance with that expected for the translated sequence (de Wet et al., 1985,1987;Ow et al., 1986). Finally, an active enzyme can be synthesized in uitro from a single mRNA species (Wood et al., 1984)and the recombinant luciferase is kinetically and catalytically undistinguishable from that isolated from the firefly (Wood 1989). It appears then that the active enzyme contains only one type of subunit, which is rather tightly packed compared to typical globular proteins in solution. Also, the mature species is not specifically modified (proteolytic cleavage or posttranslational covalent modification) as a mature species, except for the alkylation of the amino terminus, which impairs sequencing of the firefly enzyme by Edman’s degradation. The possibility of its occurrence as a homodimer during catalysis is discussed further, along with the kinetic properties of the enzyme. A moderate homology (45-48% identity) is found between the protein sequences of luciferases from different species (Wood et d., 1989b; Masuda et al., 1989). A search of the protein data base indicates a relatively high (34% identity) sequence homology with 4-coumarate : CoA ligase (Schroder, 1989).This plant enzyme catalyzes the reversible activation of coumarate by ATP, and the resultant adenyl residue is further

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

285

displaced by coenzyme A (CoA) to yield an activated thioester of coumarate at the carboxyl residue. The functional similarity between the enzymes is extended by the formation of L-CoA catalyzed by luciferase in the presence of the nonoxydable analog L, ATP, and CoA (McElroy and Seliger, 1961). It is significant that the regions of homology between the plant enzyme and beetle luciferases are restricted to conserved domains along the sequence, as indicated by hydropathy analysis. These results are indicative of a more conserved tertiary structure and suggest that these enzymes have evolved from a remote common ancestor, and may be useful to map the ATP binding site of luciferases. 3. Initial and Steady-State Kinetics of Light Production The kinetics of the reaction can be followed by monitoring light emission. The measured light intensity represents, in fact, the instantaneous rate of photon emission which is stoichiometrically related to that of substrate consumption. The dependence of light production on substrate concentration (varied in a wide range) strongly diverges from ideal Michaelis-Menten behavior. The concentrations of ATP, LH2, and 02, giving apparent half-saturation, are about 100, 10, and 0.1 p M , respectively. The odd pre-steady-state kinetics of light production hinder a straightforward analysis. At saturating substrate concentration, the addition of the last substrate results (after a short lag) in the emission of a flash (lo) followed by a steady emission of lower light intensity (Iss). This is equivalent to an initial burst of activity and indicates that the release of bound product, subsequent to photon emission, is rate limiting at steady state. Under nonsaturating conditions (low [ATP] or [LH*]),no flash is observed and the constant light output is proportional to the limiting substrate concentration (Lundin et al., 1976; Miska and Geiger, 1987) (see Fig. 8). The difference in the dependence on [ATP] of the peak height and the steady-state light output has led to the proposal that two distinct catalytically active sites exist for ATP, the first one (lower affinity)being responsible for the initial flash and the second producing the lower steady light intensity (DeLuca and McElroy, 1984). This is also based on the apparent stoichiometry of two ATPs for one L to saturate the active site (1 : 100 kDa) (Lee et al., 1970; DeLuca and McElroy, 1984). Cooperative regulation through two allosteric binding sites for nucleotides has also been considered from the effect of various nucleotides on light production elicited by ATP (Ugarova, 1989). However, these early conclusions contradict the actual molecular mass of the enzyme (see Section III,B,2) and need to be critically reassessed. The following arguments may be considered against the involvement of multiple binding sites or a dimeric form of the enzyme: (1) The dependence of light production on [LH2] indicate

286

CLAUDE AFLALO

that a flash is emitted at high concentrations only, as is the case with ATP. High concentrations of each substrate inhibit the steady-state light output more than the peak height (C. Aflalo, unpublished observations). Moreover, luciferase luminescence is subject to noncompetitive inhibition with respect to both ATP and luciferin (Lemasters and Hackenbrock, 1977), suggesting that inhibition is due to a slow release of bound product (Gates and DeLuca, 1975), rather than the existence of additional regulatory sites for ATP. (2) The determination of a stoichiometry of 2 for the reactive ATP: L ratio might have been biased by the relatively low [L] used in active-site titration with ATP, as compared with the symmetric titration with L done in the presence of a large excess of ATP (DeLuca and McElroy, 1984). The former condition is dictated by the experimental set-up for the fluorimetric determination of the tightly bound product L-AMP in the presence of fluorescent dehydroluciferin. (3) The complex kinetic behavior, including the pre-steady-state kinetics and the differential dependence of the peak height and the steady-state light output, may be simulated by numerical integration (C. Aflalo, unpublished observations), assuming single binding sites for ATP and LH2 and positive modulation of the inhibitory oxyluciferin release by PPi (see below). The following discussion is based on experimental data reported in two major communications (DeLuca and McElroy ,1974; McElroy and Seliger, 1961). The kinetically relevant steps are summarized below. FL + LH2 + MgATP FL + LH2-AMP FL t L + MgATP FL LH2-AMP + 0 FL LO* AMP FLLOAMP

2

tf t)

-

+ +

FL LH2-AMP + MgPPi FL LH2-AMP FL L-AMP + MgPPi

(1)

(la) (lb)

FL LO* AMP + COz FL LO AMP + hv FL+LO+AMP

Step 1 may be considered a fast equilibrium reaction since the time required to reach peak light emission (-0.3 seconds) is largely independent of enzyme or substrate concentration, and light emission with synthetic LH2-AMP (starting from step la) is not elicited faster. The reversible activation of L (step lb) to yield tightly bound L-AMP represents a dead-end reaction. Step 2 is believed to include two relatively slow processes (tentatively, proton abstraction from the a-carbon and a conformational change in the enzyme) responsible for the lag and rise of luminescence observed at short times (see also Lemasters and Hackenbrock, 1979; Aflalo and DeLuca, 1987). After the fast photon emission (step 3), the active-site occupancy by product causes a progressive depletion of free enzyme and results in the decrease of light output until a steady state is reached between the fast catalysis and the slow release of product. AMP

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

287

should be bound during light emission, since AMP analogs affect the color of the light emitted (DeLuca et af., 1973). It is uncertain whether it is released concomitant with (i.e., rapidly, step 3), or subsequent to, light emission (step 4, rate limiting). It may be calculated that, at saturated substrate concentrations, the light output at steady state (10% of peak height), represents about 4% of the theoretical maximum rate if no limiting step occurred subsequent to photon emission. The rate of decay to steady state and the ratio ZO : Z,, (see Fig. 8) strongly depend on the nature and concentration of the reactants present in the assay. The decay is accelerated at high substrate concentration in parallel with an increase in the peak height, indicating a faster accumulation of the bound product(s), relative to its (constant) release. I,, is markedly lowered in the presence of pyrophosphatase or when synthetic LH2-AMP is used as a substrate instead of LH2 + Mg ATP, so that the catalytic cycle is almost completely inhibited after the initial flash. It is likely that the release of product(s) is positively modulated by PPi (see also Schram et al., 1981). Both the intensities of the flash and the steady light output may be enhanced by several factors which tend to decrease the effective polarity of the solvent (e.g., glycerol and polyethylene glycol). Similar effects are obtained at high protein (albumin) concentration, or in the presence of neutral detergents (Kricka and DeLuca, 1982). It seems that these factors also stimulate the release of product from the (hydrophobic) catalytic site. The presence of lipids tightly bound to the enzyme [isolated with no delipidation step (Ugarova and Dukhovitch, 1987)] may indicate their possible role in the modulation of light emission in uiuo. One can conclude that a high substrate concentrations the reaction is governed by bound product inhibition and that, paradoxically, increasing the initial catalytic rate (i.e., increase in peak height) results in a relative reduction of that at steady state. At limiting substrate concentrations, the overall rate is much lower and the amount of enzyme-intermediate complex is likely to represent the limiting factor. This important corollary has been used extensively to design improved conditions for the determination of ATP with FL (Lundin et af., 1976; Lemasters and Hackenbrock, 1979).

IV. A Local Probe for ATP in Model Systems The usefulness of soluble FL to specifically measure [ATP] at high sensitivity is well established (Lundin et af., 1976; DeLuca and McElroy, 1978; Whitehead et al., 1979). The ATP assay has been coupled to ATPconverting reactions in soluble systems (Lundin, 1982), isolated mitochondria (Lemasters and Hackenbrock, 1979), and chloroplast thylakoids

288

CLAUDE AFLALO

(Beard and Dilley, 1986) for continuous monitoring purposes. While a soluble enzyme monitors ATP in bulk solution, a localized enzyme should respond to [ATP] in its immediate vicinity. The localization of FL and bacterial luciferase in a restricted environment has been achieved by their covalent attachment to solid supports. The chemical immobilization confers to the enzymes a better stability, with no apparent loss of light production (Wienhausen et al., 1982). DeLuca and co-workers first introduced the immobilized probe concept to develop a versatile analytical tool for the determination of metabolite concentrations in liquid samples. The system consists of bacterial luciferase (and oxidoreductase to regenerate FMNH2 from NADH) coimmobilized with dehydrogenases and other coupled enzymes (DeLuca, 1984; Wienhausen and DeLuca, 1986). In this system, the light output from luciferase is empirically correlated with the level of NADH produced locally by the coimmobilized couple enzymes in response to added cosubstrates.

A. CONTINUOUS MONITORINGUSINGIMMOBILIZED LUCIFERASE We further developed this approach using FL immobilized on Sepharose (Pharmacia, Sweden) beads to monitor [ATP] directly in the microenvironment of colocalized enzymes which produce or consume ATP (Aflalo and DeLuca, 1987,1988). As a control system, the enzymes were immobilized under identical conditions, except that luciferase and the ATPconverting enzymes were attached to separate beads. The global catalytic power of the first system (coimmobilized FL-kinase, or, in short, Im : FLkinase) can be reconstituted by mixing the beads of the separate batches (i.e., Im : FL + Im :kinase) in the right proportion. Controls have shown that immobilized luciferase responds linearly to low [ATP] added to the medium and as the enzyme in solution, does not significantly consume it under the test conditions. However, upon rapid addition of ATP to the beads suspended in a luciferin solution, light output evolves slower than with the enzyme in solution, due to a lower diffusibility of ATP to the immobilized luciferase. Light output from immobilized luciferase was monitored during ATP production by pyruvate kinase (PK) immobilized on the same beads (Im : FL-PK) (Fig. 3a) or on separate beads (Im : FL + Im : PK) (Fig. 3b). Upon addition of phosphoenol pyruvate and ADP to the stirred suspension, a burst of light is observed in the coimmobilized Im : FL-PK system; this is followed by a steady increase. With the Im : FL + Im : PK system, the same slow increase is observed after a lag corresponding to the burst observed with Im : FL-PK. None of these patterns is observed with the enzymes in solution. This result suggests that the ATP generated by the concentrated immobilized PK accumulates rapidly inside the beads before

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

289

FIG. 3. Local and macroscopic production of ATP by immobilized pyruvate kinase (PK). The reaction mix contained 0.1 mM LH2and 10 mMglucose in buffer and equivalent amounts of luciferase and PK (20 U/ml) coimmobilized(a) on the same Sepharose beads or (b) on separate beads. The final concentrations of the added reactants were: ADP, 1 p M ; phosphoeno1 pyruvate (PEP), 1 mM; hexokinase (HK), 2 U/ml. The stirring protocol is indicated by the bar at the bottom. Reproduced with permission from Aflalo, C., and DeLuca, M. (1987), Biochemistry 26,3913-3919. Copyright 0 1987 American Chemical Society.

it diffuses out to the bulk medium. A steady state is reached in which the rate of appearance of ATP in the medium equates that of its formation in the beads. Thus, with Im : FL-PK, the slow increase in the local [ATP] at steady state reflects its accumulation in the medium. When the stirring is stopped, a biphasic increase of light output is observed with both systems. In the fast phase, the light output from Im: FL-PK increases, while no change is observed with Im :FL. This indicates the establishment of additional diffusional restrictions (at-and beyond-the surface of the bead) for ATP. The slower increase with both systems represents the settling of the beads to the bottom of the cuvette, bringing them together into a larger compartment, from which ATP diffuses out slower. The accumulation of ATP inside the beads is clearly demonstrated by its inaccessibility to a large excess of soluble hexokinase added to the bulk medium.

290

CLAUDE AFLALO

Similar behavior can be observed with ATP consumption by immobilized hexokinase (Fig. 4). In the absence of glucose, addition of ATP to luciferase coimmobilized with hexokinase (Im : FL-HK) (Fig. 4a.B) or attached to separate beads (Im : FL + Im: HK) (Fig. 4b.B) elicits light production at the same rate. When a saturating concentration of glucose is added, the light output from Im : FL-HK decreases rapidly in a first phase to a steady state, which further decays exponentially to zero. The decrease in light output from the Im : FL + Im : HK system occurs as a monotonic slow decay in parallel with the second phase of the coimmobilized system (Fig. 4B). When glucose is added before ATP, the addition of the latter results in a fast increase of the light output to the steady-state value, followed by the slow decay as previously observed (Fig. 4C). This experimental set-up enables the kinetic resolution of the variation in ATP directly in the environment of HK inside the beads (local [ATPI), using the coimmobilized luciferase, or alternatively in the medium (bulk [ATPI), using the separately immobilized enzymes. One can derive kinetic parameters from the data in Fig. 4a.B and separate them into two groups according to their dependence on the beads concentration. While kdif and ~ H K , describing properties inherent to the heterogeneous system, are constant, the effective parameter kobs is directly proportional to the bead concentration. Similarly, the observed values for I,, and I0 depend on the amount of beads, but their ratio (representing a microscopic property of the beads) remains constant (Aflalo and DeLuca, 1987). In fact, this ratio represents an effectiveness factor (see derivation in the mathematical modeling section) which can be used to evaluate directly intrinsic properties of HK in its restricted environment from macroscopic measurements. This is illustrated in the following example. If the experiment described in Fig. 4a.C is repeated at varying glucose concentrations, one can derive the dependence of kobs on the latter. This yields an apparent K0.5 for glucose lower by about one order of magnitude than that expected for the enzyme in solution. This value reflects the strong limitation imposed by ATP diffusion, since the beads are preequilibrated with glucose added in excess over ATP. However, the macroscopic values of kobs can be corrected by simply dividing them by the experimentally determined ratio I,, :Io, yielding back the true dependence of HK on glucose concentration in the microenvironment of the bead (Aflalo and DeLuca, 1987) (see Section V,B). This approach just described is the first report on the continuous monitoring of local [ATPI in a heterogeneous system. The use of localized FL enables the kinetic resolution of the variation of ATP in the microenvironment of the enzyme from that in the bulk phase. This, in turn, permits a direct assessment of intrinsic parameters of enzymes sequestered at high concentration in a definite compartment, as illustrated in the following.

29 1

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

A

B

C

a

hv

b

ATP Glu Glu ATP FIG. 4. Local and macroscopic consumption of ATP by immobilized hexokinase (HK). The reaction conditions were as described in Fig. 3, with constant stirring. The total [HK] was 50 Ulml. ATP and glucose were added at final concentrations of 100 nM and 0.2 mM, respectively. Top panel (a), coimmobilized FL-HK; bottom panel (b), separately immobilized FL and HK. (A) Schematic representation of the immobilized systems. (B) Transient depletion of ATP in the microenvironment of HK and in the bulk medium. (C) Steady-state measurements after preequilibration with glucose. kdS. kHK,and kobsrepresent the pseudo-first-order rate constants for the ATP diffusion-limited light evolution and the fast and slow decays of light output in (B) respectively. Reproduced with permission from Aflalo, C., and DeLuca, M. (1987). Biochemistry%, 3913-3919. Copyright 0 1987 American Chemical Society.

B. MATHEMATICAL MODELINGWITH SOLUBLE AND LOCALIZED LUCIFERASE'

I. Kinetic Analysis of the Firefly Luciferase Reaction The light output from luciferase can be represented as the rate of photon emission, which is related to the reaction velocity as follows:

z=

'

(,+)

=

c(vol)(7)d [ATP]

Adapted with permission from Aflalo, C., and DeLuca, M. (1987), Biochemistry 26, 3913-3919. Copyright 0 1987 American Chemical Society.

292

CLAUDE AFLALO

where c is a constant depending on the geometry and electronic properties of the photometer, and vol is the total volume. For the sake of simplicity, the velocity at steady state will be considered to obey a Michaelis-Menten behavior. Thus, at all [ATP],

and at low [ATPI,

I=

C( V O ~kFL[ ) FL][ATP]

(la)

It must be stressed, however, that kFL has a different value than k,atlKm calculated at relatively high [ATP] (see Section III,B,3). More elaborate models have been proposed for the complex time dependence (Aflalo and DeLuca, 1987; Slooten and Vandenbranden, 1989; see also Ugarova, 1989).

2 . Effect ofDiffusion on the Kinetic Behavior of Immobilized Enzymes a. External Diffusion. Due to the action of an enzyme immobilized on a solid support, the local concentration of substrate S should be lower than that in the bulk medium S m . At steady state, the rate of substrate disappearance from the medium by diffusion (Fick’s first law) is equal to that of catalysis at the interface. Therefore,

where Dm,A, x , and kt represent the diffusion coefficient of substrate in the bulk medium, the area of the support exposed to the medium, the distance from the support, and a mass transfer coefficient for the substrate, respectively. The latter is an empirical proportion factor which depends on Dm, A, and also on the hydrodynamic properties of the system [geometry, dimensions, and mixing (see Engasser and Horvath, 1976)l. At low substrate concentrations, we may derive S from a simplified Eq. (2) as

The extent of diffusion control (i.e., the ratio S: S m ) may be substantially reduced by efficiently mixing the bulk solution, resulting in a fast mass transfer compared to the rate of catalysis. b. Internal Diffusion. When an enzyme is embedded in a porous matrix, the substrate is also, in this case, subject to two concurrent processes:

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

293

diffusion in the support, which is largely independent of external mixing, and its consumption by the enzyme. Introducing Fick’s second law, the phenomenological equation for the system, assuming a spherical symmetry of radius R , is

at any t > 0 and 0
Assuming that a steady state is reached rapidly (no significant variation in S,,,)? that external diffusion is negligible, and that the enzyme concentration [El is constant throughout the whole support, one may set the following boundary conditions S = S, dS _ -0

at r = R at r = O

dr

The analytic solution of Eqs. (5) is R

s = s, sinh(crR)

so = s,

sinh(cYr) r

ffR sinh(cuR)

where

The dimensionless parameter CUR describes the physico-chemical properties inherent to the heterogeneous catalytic system, and will determine the substrate concentration profile in the support, as well as the catalytic efficiency. The higher the ratio of the rate for enzymatic catalysis to diffusion and/or the size of the support, the steeper the concentration gradient in the support as shown in Fig. 5a. The catalytic efjciency may be defined as the ratio between the effective activity of the immobilized enzyme to that expected if it was in solution. The macroscopic activity is the rate of disappearance of substrate (at the interface) from the medium.

294

CLAUDE AFLALO

a

b

r/R WR FIG.5 . Diffusion control in a spherical heterogeneous catalytic system. (a) Concentration profile of substrate consumed in the support at steady state. (b) Variation of the effectiveness factor with the characteristic properties of the system. The curves were generated using Eqs. (6) and (9), respectively. Reproduced with permission from Aflalo, C., and DeLuca, M. (1987), Biochemistry 26,3913-3919. Copyright 0 1987 American Chemical Society.

For a single bead, it is given by Fick’s first law

where V , represents the volume of the bulk medium. For the same amount of enzyme in solution, the velocity would be

The effectiveness factor p is found by dividing Eq. (7) by Eq. (8) and rearrangement p =

3

-[aR coth(CUR) - 11 ff2R2

(9)

This quantity is independent of the number of beads in suspension. The behavior of this function is such that it is approximately equal to one at low CUR (no diffusion control), and decreases in a near exponential fashion as CUR increases (Fig. 5b).

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

295

3. Quantitation of Light Output from (Co)immobilized Luciferase At low [ATP], the measured light output is the sum of the emissions from the individual FL molecules homogeneously distributed in the support. However, since gradients of ATP may occur, one should calculate the total output by integrating the differential form of Eq. (la) over the volume of the spherical support. Thus for a single bead

Z = c~FJFL]loR[ATP],4?r?dr

(10)

where [FLI represents the local concentration of the enzyme in the support. At steady state, we may substitute [ATP] using Eq. (6). After integration and rearrangement, we get for light output at steady state (see Fig. 4,B and C) 4

3 a2R2

Is, = C-~TR~~FL[FL][ATP],,-, -[aR coth(aR) - 13 3

(1 1)

where a is a function of all the processes consuming ATP. Thus, for singly immobilized FL, assuming a very low rate of ATP consumption ( ~ F L+ 0 and /.LFL + 1; see Fig. 5 ) , we get no concentration gradient inside the support (i.e., [ATP], = [ATP],), and the light output is identical to that produced with the enzyme in solution [Zss = Zo in Fig. 4; compare with Eq. (la)]. On the other hand, for FL coimmobilized with an enzyme consuming ATP, Eq. (1 1) reduces to the simple expression

Is, = lop

(12) where l o represents here the light output elicited by the addition of a known concentration of ATP in the medium with no ATP consumption by the coimmobilized enzyme in a single bead. The values of l o and Z,, are experimentally measurable as the total light output from all beads in suspension in the absence or presence of glucose, respectively, as shown in Fig. 4B. These considerations are the basis for calibration of localized luciferase. In summary, this approach demonstrates the feasibility of continuous monitoring of ATP in definite microenvironments by artificially localized luciferase. Local measurements for biological compounds in easily manipulatable artificial systems represents a powerful means to develop and design physicochemical systems approximating the cellular ones. The parallel development of mathematical models enables the generation of predictions which, in turn, may be challenged in artificial systems of increased complexity. Furthermore, the simulated behavior in the models may be directly confronted with the macroscopic behavior of cellular

296

CLAUDE AFLALO

systems, using independently assessed physicochemical parameters of the cell (e.g., dimensions; enzymes and metabolites concentrations). Thus, local [ATP] monitoring by luciferase may, in principle, be extended to more realistic biochemical systems, provided that the enzyme is accurately targeted to defined cellular compartments, while retaining its activity, and that it can be properly calibrated.

V. Leading Luciferase into Cellular Compartments

A. CLONING, EXPRESSION, AND LOCALIZATION OF NATIVE LUCIFERASE The FL gene (luc)has been successfully cloned and expressed in viruses (Rodriguezet al., 1989),bacteria (de Wet et al., 1985),yeast (Aflalo, 1990; Gould et al., 1990), plants (Ow et al., 1986; Schneider et al., 1990), cultured mammalian cells (de Wet et al., 1987), and transgenic mice (DiLella et al., 1988; Lira et al., 1990).Polyadenosyl mRNA isolated from P . pyralis (Wood et al., 1984)or Luciola mingrelica (Kutuzova et al., 1989) fireflies’ light organs has been translated in cell-free systems to yield a bioluminescent product. The amount of active enzyme produced depends on the in uitro system used and the concentration of RNA in the reaction. In addition, mRNA transcribed in uitro has been introduced in plant cells by electroporation (Gallie et al., 1989), in mouse cells through cationic liposomes (Malone et al., 1989) or injected into Xenopus laeuis oocytes (Kutuzova et al., 1989), respectively. In all cases, in uiuo translation resulting in bioluminescence activity was observed. Similar results were obtained with luciferase from Luciola cruciata cloned in bacteria (Masuda et al., 1989).This suggests that the correct folding of the protein chain does not require specific posttranslationalmodification. However, comparative studies of the fate of the translated and mature enzyme (purified from the firefly) in cells or cell-free extracts indicate that some specific processing may occur in the firefly, resulting in improved stability of the enzyme (Kutuzova et al.. 1989). The de nouo synthesis of luciferase endowed with luminescence activity opens wide possibilities for use of the enzyme as a genetic probe (Gould and Subramani, 1988; Wood et al., 1989a). However, light emission in intact cells has so far little value by itself for quantitative or kinetic analysis. The straightforward interpretation of in situ data is hindered by the uncertainty of the environmental conditions under which the enzyme is operating. In order to develop accurate calibration procedures, the localization of the enzyme and other physicochemical parameters (e.g., cosub-

BIOLOGICALLY LOCALIZED FIREFLY LUCIFEFUSE

297

strate concentration and pH) should be determined. Native luciferase expressed in transformed yeast, plant, or animal cells is spontaneously directed to peroxisomes (Gould et al., 1990a), as is the case in the firefly light-producing cells (Keller er al., 1987). The exact location has been assessed by indirect immunofluorescence or immunogold labeling microscopy of whole cell preparation. The direct subcellular localization of luminescent activity in single cells has not yet been achieved at reasonable resolution. The localization of luciferase to peroxisomes occurs posttranslationally due to targeting sequence located at the carboxy terminus of the protein (Gould et al., 1987). The minimal peroxisome targeting sequence is a conserved carboxy-terminal tripeptide (Ser-Lys-Leu) which is necessary to direct many natural proteins to peroxisomes, and sufficient to redirect soluble proteins to these organelles. Moreover, antibodies raised against the tripeptide recognize many peroxisomal proteins (Gould et al., 1990b). It is notable that the Japanese firefly lacks such a sequence (Masuda er al., 1989).Mutagenetic analysis indicated that only alimited number of conservative changes in this sequence are allowed which do not result in the delocalization of the protein to the cytoplasm (Gould et al., 1989). In order to specifically localize the enzyme in biological systems, one can use the cellular machinery responsible for the correct targeting and assembly of endogenous proteins in defined compartments.

B. RETARGETING LUCIFERASE TO OTHERCELLULAR ORGANELLES Fusion genes between the 5‘-coding region of a Saccharomyces cerevisiae mitochondria1 protein (70-kDa outer membrane protein) and luciferase cDNA (fuc)have been constructed (Aflalo, 1990). The “mitochondrial” moiety of the chimeric products includes an amino-terminal sequence of 30-40 residues (Hase er al., 1983) which targets the parent protein to mitochondria and anchors it to the outer membrane through a transmembrane a-helix (Hase er al., 1984). The construction is schematically presented in Fig. 6. The final fusion gene has been transferred to a yeast-bacteria shuttle plasmid under the natural promoter of the yeast gene and used to transform yeast cells. The native luciferase gene (under a galactose promoter) was also cloned in yeast as a control. The fusion and native genes are both expressed, and the products are endowed with bioluminescence activity in intact yeast and in cell-free extracts. The fusion gene is expressed in parallel with mitochondria proliferation, and its active product is highly enriched in isolated mitochondria. When the native gene is overexpressed in the presence of galactose, the resulting bioluminescence activity is present mostly in the postmitochondrial supernatant, and may further be enriched in a microparticulate fraction.

298

CLAUDE AFLALO pKW 114 4.lKbp

B BIN

b

f

I

Iuc H

1.7kbp

co‘ 27kbp

d

,

PT7 B I E E

H-B

co 2 lkbp

co 27kbp

FIG.6. Construction of 70K-luc fusion genes. Selected genes and open reading frames are emphasized. (a-c) Steps in the construction of the 70K-luc fusion gene encoding for 292 residues from the yeast protein. The letters in brackets indicate the origin of the corresponding segments. The dashed segment (c) corresponds to a deleted fragment. (d) and (e) represent the shortened (92 residues) fusion gene in the bacterial and yeast plasmids, respectively. (f and g) Schemes for pKWl14 (bacteria, native luciferase) and pFL1-ca (yeast, fusion gene). 70K, Original 70-kDa outer membrane protein gene from yeast; ca’ and ca, fusion genes; 2pD, 2pm yeast EcoRI-D fragment; amp, ampicillin (penicillin)resistance gene; B, BamHI; BI, Bgn; BII, BgnI; E, EcoRI; H, HindIII; N,NsiI; ori, E. coli origin of replication; p l 7 , l7 phage promoter (@lo); S, SmaI; URA3, yeast orotidine-5’-phosphate decarboxylase gene. Reproduced with permission from Aflalo, C. (1990),Biochemisrry 29,4758-4766. Copyright 0 1990 American Chemical Society.

However, a small fraction remains associated with highly purified mitochondria. Since the inducible proliferation of peroxisomes is very low under these conditions (Distel et al., 1987), it is possible that the native enzyme has been misrouted at low efficiency. The fusion protein is specifically directed in uiuo to the cytoplasmic face of the outer membrane of yeast mitochondria (Aflalo, 1990) as the parent 70-kDa protein (Reizman et al., 1983). This has been assessed by the effect of detergent or limited proteolysis on luciferase activity in isolated mitochondria containing the chimeric product. Figure 7 shows the time course of proteolysis (A) and the distribution of residual bioluminescence (B) following trypsin treatment of mitochondria-associated and soluble luciferase (in the presence of mitochondria). While the purified luciferase does not associate with wild-type mitochondria and is relatively insensitive to trypsin, the mitochondria-bound fusion product is first clipped off, releas-

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

299

ing all the active luciferase moiety to the medium (not shown), and further completely destroyed by the protease, indicating that no active luciferase is present in internal compartments. Finally, the fraction of native enzyme associated with intact mitochondria remains insensitive to trypsin (before solubilization with detergent), indicating that it must be located beyond the outer membrane. Luciferase has also been targeted to chloroplasts in tobacco plants (Schneider et al., 1990). A fusion gene between a 5' fragment of the ribulose bisphosphate carboxylase (RUBISCO) small-subunit gene and luc at the 3' end has been constructed. The plant-derived fragment encodes for 57 amino-terminal residues, including the transit peptide responsible for targeting the small subunit of RUBISCO to the stroma of chloroplasts. As with the native luc, the chimeric gene is expressed in whole plant tissues, and bioluminescence activity is associated with the expression. In this case also, the fusion product is correctly targeted, processed, and assembled in the stroma of chloroplasts, while the product of the native gene is directed to an internal (foreign) subcompartments still retains its catalytic ability. Thus, the polypeptide chain can unfold in the cytoplasm, cross a biological membrane, and correctly refold in a different environment.

B

A

-E

12

El suspension 0 supernatant

pellet

E 100 0 0

.-.-E>

e

L

60

a

40

;

20

.-.-S>

0

a

8 E 4 d0

a,

a

80

s

4

s

0

A

Time (min)

o pFLl

+ FL

pMD45

pFL1-ca

FIG. 7. Localization of purified and cloned native or targeted luciferase. Mitochondria ( I mg/ml) isolated from wild type (pFLI) or yeast cells transformed with the native (pMD45) or fusion gene (pFL1-ca) were incubated on ice with trypsin (10 pg/ml). Wild-type mitochondria were supplemented with soluble luciferase (FL, 10 ng/ml). At the indicated times, trypsin inhibitor (0.2 mg/ml) was added and aliquots from the suspension were analyzed for luciferase activity at saturating substrate concentrations, or separated into soluble and particulate fractions and then analyzed. (A) Time course of proteolysis. arb., arbitrary units. (B) Residual activity in untreated (con) or treated (15 minutes, try) samples.

300

CLAUDE AFLALO

VI. Light Emission by Luciferase in Biological Systems Light emission from cloned luciferase in situ depends on a variety of factors which may affect the amount of product present in the cell, its intrinsic catalytic ability, and the local conditions under which the enzyme is operating. IN WHOLECELLS A. LIGHTEMISSION

The total amount of luciferase produced in the cell depends on the efficiency of transcription and translation, determined mainly by the type of promoter used with the cloned gene (see Gould and Subramani, 1988; Wood, 1989; Aflalo, 1990; Schneider et al., 1990), and other regulatory elements added upstream (e.g., enhancers) or downstream [e.g., poly(A) sequences] to the gene (Wood, 1989; de Wet et al., 1987)or mRNA (Gallie et al., 1989). The efficiency of bulk expression is best assessed by quantitative determination of mRNA, or that of the product peptide in immunoblots from solubilized extracts of whole cells. A reasonably good correlation is generally found between these two parameters. The level of bioluminescence in transgenic plant tissues closely reflects that of transcription of the luciferase gene (Scheider et al., 1990). Moreover, the overexpressed luciferase appearing in the microparticulate fraction of yeast cells is active and represents most of the total activity (Aflalo, 1990). Finally, studies’of the translation and stabilization of luciferase in X.laeuis oocytes (Kutuzova et al., 1989) indicate that the enzyme is correctly assembled in a conformation compatible with eukaryotic cellular environment. Early characterization studies of the cloned enzyme in bacterial extracts indicate that it is undistinguishable from mature luciferase. Indeed, the kinetic pattern of light emission, its spectral distribution, and sensitivity to inhibitors are similar for both enzymes in the presence of bacterial extracts (Wood, 1989). These conclusions may be somewhat extended to the chimeric luciferase targeted to mitochondria, as shown in Fig. 8. The dependence of soluble luciferase activity on [LH2] is significantly affected by the presence of mitochondria (see Fig. 8 insets). Moreover, in the presence of neutral detergents, the differences in the kinetic behavior of both systems are further reduced (unpublished observations). It seems, then, that the intrinsic catalytic properties of the enzyme are rather insensitive to moderate variations in the primary structure at either end of the sequence. Additional experimental evidence is needed to confirm this preliminary proposal, which is relevant to the design of genetically engineered luciferase.

301

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

16 12 o8 '' OO

400 800 1200 1600 2000

IE-l

I

100

10

1000

10 I IE-l

16 12 0' 8 ..

I€-2

'0

20

40

60

00

100

I E-3 IE-3 IE-2 IE-l

4 0 I

10

f

100

FIG. 8. Dependence of light production of soluble and mitochondria-bound luciferase on ATP and luciferyl adenylate (LH2). Isolated mitochondria containing 20 pg of protein and/or 0.2 ng of purified luciferase were assayed for light production activity. The reaction was initiated by injection of LH2. The peak height (I, closed symbols) and steady light emission (I,,, open symbols) are reported. The samples assayed were soluble firefly luciferase (FL) alone (circles), FL supplemented with wild-type mitochondria (triangles), and FL bound to mitochondria isolated from cells transformed with pFL1-ca (squares). (A and B) Dependence on [ATP]. [LH2] = 0.1 mM. (C and D) Dependence on [LH2]. [ATP] = 2 mM. (Insets) Dependence of the ratio Z, :I,, on substrate concentration. Reproduced with permission from Atlalo, C. (1990), Biochemistry 29,4758-4766. Copyright 0 1990 American Chemical Society.

The activity of cloned luciferase in situ may, in addition, be significantly affected by the accessibility of substrates and other environmental conditions occurring locally. Preliminary light measurements with intact cells containing cloned luciferase indicate that the most likely limiting factor is the intracellular concentration of luciferin diffusing in from the medium (Gould and Subramani, 1988; Aflalo, 1990). The bioluminescent activity of transformed cells with exogenous luciferin increases at low pH, due to a better permeability of protonated LH2 through the cell membrane. In animal cells, this is further improved by the addition of dimethylsulfoxide, or nigericin in the presence of potassium (Gould and Subramani, 1988). It is expected that the acidic LH2 molecule will accumulate at equilibrium in

302

CLAUDE AFLALO

the more basic of two aqueous phases separated by a membrane. Some features of light emission by transformed yeast cells are presented in Fig. 9. Upon addition of LH2, a slow increase in light output is observed, representing the build-up of intracellular [LH2] (not shown). At saturation, the light emission shows the characteristic flash pattern indicating bound product inhibition. However, this occurs at a much slower rate than with soluble luciferase, due to the kinetic limitation imposed by the slow diffusion of luciferin. Oxygen is present at saturating concentrations (-250 p M at 25°C) in solutions equilibrated with air. However, actively respiring cells rapidly consume all the oxygen in standing suspensionsand reduce the light output according (Fig. 9B-D). After additional incubation in anaerobic conditions, a fast flash is generated upon reoxygenation. The peak height is dependent on the length of the anaerobic incubation (Fig. 9A,3), indicating that the inhibitory product is slowly released from the inactive enzyme. The flash emission does not occur with cells with a low respiratory capacity (see Fig. 9A,2, for yeast grown in the presence of glucose). The results indicate that even under severe anaerobiosis, the local [ATP] is not a limiting factor for light production. Thus, the average [ATP] in normal cells (in the millimolar range) apparently saturates the light production by luciferase in intact cells. However, these data illustrate the use of luciferase as a localized oxygen sensor. Bioluminescence of intact transformed cells is currently used as a qualitative tool to detect positive transformation, at relatively low sensitivity to cell-free extracts. At this stage, one can only speculate on its quantitative aspects. However, further studies on the catalytic behavior of the native enzyme and its localized fusion products in various environments (e.g., cell-free extracts) will help to better define the factors affecting the light output in situ. This will provide additional insights on localized processes in the cell. At present, straightforward measurements of ATP are possible at low concentrations only, and the use of the probe is limited by its calibration. A strict control of the conditions for the calibration and operation of the localized probe is possible only in isolated systems, in which luciferase is exposed to the medium, as illustrated in the following section.

B. MONITORING LOCAL[ATP] AT THE MITOCHONDRIAL SURFACE 1 . Local versus Bulk [ATP] Measurements

Low [ATP] emerging from isolated mitochondria can be determined by directly monitoring the light output from the outer membrane-bound enzyme or soluble luciferase added to wild-type mitochondria as a control. The light output by luciferase in these conditions is linearly related to

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE Time after LH2 addition (rnin)

A 20

25

30 35 -40 45

50

1 1 -

Time after LH2 additon (min)

55

10

20

15

TI

\

c

Mixing:

Time after LH2 addition(min)

off

on

303

D

\ 2.5~ \ \

Time after LH;! addition (min)

Mixing'off on

FIG. 9. Light production by cloned luciferase and oxygen consumption in intact yeast cells. Yeast cells transformed with pFL1-ca (fusion gene) or pMD45 (native gene inducible by galactose) were grown in the presence of glucose (repressor of mitochondria proliferation) or a nonfermentable carbon source. The cells were harvested (log phase), washed in water, and resuspended in Na-MES (Na-morpholino ethane sulfonate), pH 5.5. The luciferase reaction was initiated by addition of 50 p M luciferyl adenylate (LH2) and vortexing to saturate the suspension with oxygen. (A) Light output from cells transformed with the native or fusion gene of luciferase and grown as indicated: I , fusion, lactate; 2, fusion, glucose; 3, native, galactose. Arrows indicate reoxygenation. (B) Native, galactose. Effect of cell concentration (in multiples of that in A3) on the light output and oxygen consumption, simultaneously recorded using a microelectrode. (C) Fusion, lactate. (D) Native, galactose; Same as (B), with continuous magnetic stimng, as indicated at the bottom. Reproduced with permission from Aflalo, C. (1990), 0 Biochemistry 29, 4758-4766. Copyright 1990 American Chemical Society.

[ATPI up to 1-2 pM (see Fig. 8B). The steady-state [ATP] generated by the mitochondria and consumed by exogenous soluble HK has been assessed in both systems (Fig. 10). In the absence of HK, no significant difference could be detected in the kinetics of ATP formation in the rapidly stirred mitochondria1suspensions, nor in the calibration of both the localized and soluble luciferase using exogenous ATP. However, while in the control system the steady-state [ATP] can be reduced to undectable concentrations at high enough levels of HK (Fig. lob), the light output from mitochondria-boundluciferase is reduced to a finite value (Fig. 10a) under identical conditions. This value is further lowered in the presence of atractyloside, which blocks the transport of nucleotides through the mito-

304

CLAUDE AFLALO Time lminl 20

10

I

I

30

40

I

I

a

b

HK

HK

1.

ti

mit0 ATP

t ADP

t

ATP

I

atrac

mitt! ltTP

f

ATP

t

ADP

FIG. 10. Light emission from localized or soluble luciferase during oxidative phosphorylation coupled to hexokinase. Mitochondria (40 pg/ml) isolated from transformed (pFL1-ca) (a) or wild-type (b) yeast cells were suspended in a stirred medium containing 0.6 M sorbitol, 20 mM tricine (pH 7.8), 20 mM KCI, 3 mM Mg-tricine, 5 mM Pi, 10 m M K-succinate, 0.1 mM luciferin, and 1 mglml bovine serum albumin; (b) 10 ng/ml purified luciferase was added. The indicated additions (in small volume) were 0.1 p M ATP, 1 p M ADP, and 10 pg/ml atractyloside, and at the upper arrows, yeast hexokinase was added from serial dilutions so that its concentration doubled each time (2-64 U/ml).

chondrial inner membrane and thus inhibits oxidative phosphorylation. The residual activity reflects the contribution of adenylate kinase to ATP formation in the intermembrane space. These results indicate that when ATP is efficiently depleted in the medium by HK (as detected by soluble luciferase),the ATP emerging from the mitochondria is still available to the localized probe. Thus, at least for high catalytic rates of ATP consumption in solution, the coupled heterogeneous system may be limited by diffusion of nucleotides to and from the surface of mitochondria.

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

305

2 . Dependence on the Rate of ATP Formation

The extent of the actual limitation by diffusion is directly related to the catalytic ability of the concurrent enzymic steps. Thus, upon increasing the specific (i.e., local) activity of phosphorylation, more ATP should accumulate at steady state near the outer membrane, as suggested by the effect of atractyloside (Fig. 10a) and other inhibitors of oxidative phosphorylation. On the other hand, increasing the overall (i.e., bulk) rate of phosphorylation in the system by adding more mitochondria should not affect significantly the flux of ATP emerging from the membrane, and thus leave the local accumulation of ATP unaffected. This rationale is illustrated in the experiment presented in Fig. 1 1. The inital rate of phosphorylation in the absence of HK (see Fig. 10) was found to depend on the concentrations of added ADP and mitochondria. The experiments in Fig. 10 were repeated in the presence of tripled amounts of ADP or mitochondria, resulting in either case in a proportional increase in the observed initial rate. The steady-state concentrations of ATP at variable HK efficiencies were replotted as a function of l/[HK] in

B

A

-T

1 .o

1.o

0.8

0.8

0.6

a 0.6

0.4

2

-

-

I P

m

r I-

5

n

0.01

0.0

"

0.2

"

0.4

'

'

"

0.6

llHK ImllUi

0.8

0.4

. '

1.0

llHK Iml/U~

FIG. 11. Titration of steady-state ATP at the surface of mitochondria or in the bulk medium. Experimental conditions were as in Fig. 10. (A) Mitochondria bearing the luciferase fusion product. Circles, [mito] = 40 pg/ml, [ADP] = 1 p M ; triangles, [mito] = 120 pg/ml, [ADP] = 1 p M ; squares, [mito] = 40 pg/ml, [ADP] = 3 p M . (B) Wild-type mitochondria supplemented with purified luciferase. Symbols and concentrations are as in (A).

306

CLAUDE AFLALO

order to derive their extrapolated limit at infinitely high ATP consumption in the medium. The values obtained with the local probe are finite and depend on added [ADP], but not on the mitochondria concentration ([mito]) (Fig. 11A). The control experiments with wild-type mitochondria and soluble luciferase yielded zero extrapolation values (Fig. 11B). Therefore, with excess HK the nucleotide in the medium is all in the form of ADP, and the extrapolated values with the local probe represent the net accumulation of ATP near the membrane when its concentration in the bulk medium is zero. The results strongly suggest that the accumulation of ATP near the mitochondria depends on its flux through the membrane (i.e., per unit area) rather than on the total rate of its appearance in the bulk medium, indicating that a true local event is being monitored by the probe.

3. Modeling A similar behavior can be simulated by reducing the reactions to a simple model, including two pseudo-first-order catalytic reactions coupled through the diffusion of intermediates. In the first one, the sequences of steps-including the transfer of ADP to internal mitochondrial compartments, its phosphorylation, and the export of ATP-is assumed to occur as a single catalytic step located at the surface of spherical particles in suspension. Control experiments have shown that the initial rate of ATP formation is linearly dependent on both [ADP] and [mito] in the ranges used in this work. The second reaction of dephosphorylation of ATP in the bulk medium by HK is similarly related to [,4TP] and [HK]. The diffusion of nucleotides between the bulk medium and the surface can be described as a mass transfer whose rate is assumed to be proportional to the difference in the concentrations of the transferred nucleotide in both compartments and the concentration of surfaces (i.e., [mito]). For the sake of simplicity, it is assumed that the nucleotides diffuse in a near-planar unstirred layer sticking to the mitochondria surface, and that the volumes of both mitochondria and the diffuse layer are negligible compared to that of the bulk medium. The model is schematically outlined in Fig. 12a, and the pertinent equations describing the system are formulated in Fig. 12b, assuming an identical mass transfer coefficient of ADP and ATP. One can solve the system of differential equations for steady state, and by introducing [AXP] as the sum of endogenous and added nucleotides (ADP + ATP, represented as 100% in the ordinate of Fig. 10). A simple expression for the local [ATP] at infinite HK concentration may be reached [Eq. (21) in Fig. 121.

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

307

a

kmit kdit kHK

=

=

kox A [mito] D A [mito]/6 k"[HK]

b

d[ADP]b

=

-

d[ATP]b

kmlt

[ATPIs,axtrap.= k d l f + k ml t

=

kox D16 + ko x

. [AXP]

(21)

FIG. 12. Model for coupling two catalytic steps in a heterogeneous system. The indices s and b represent surface and bulk, respectively. R and 6 represent the radius of the (spherical) mitochondria and the width of a hypothetical unstirred layer, respectively. A is the specific area of mitochondria (cm2/mgof protein), k,, and k" are second-order rate constants, and D is the diffusion constant. The second-order mass transfer coefficient is D/6. [Eqs. (13)-(16)] Rate laws for each nucleotide species. [Eq. (17)] Mass conservation in the bulk medium. [Eqs. (18)-(19)] Steady-state solutions for Eqs. (13)-(16). [Eq. (20)] Steady-state rate of ATP appearance in the medium in the absence of hexokinase (HK) [Eq. (21)]

308

CLAUDE AFLALO

Since both kmit and kdjf are proportional to the concentration of mitochondria, it follows that the fraction of ATP accumulated at the surface is proportional to [AXP] and independent of [mito], in accordance with the results in Fig. 11. The ratio kmit/kmit + kdif can thus be determined experimentally, as the slope of the extrapolated values versus the total nucleotide concentration. In the absence of HK, the phosphorylation of added ADP may be represented by a single process with a composite pseudo-first-order rate constant (mass transfer and surface catalysis occurring in sequence) (Easterby, 1981), as shown in Fig. 12 Eq. (20). This value, proportional to the concentration of mitochondria, can be determined experimentally as the slope of the observed initial velocity of phosphorylation versus added [ADP]. Next, the values of kmit and kdif can be derived by solving the system of equations [Eqs. (20)-(21)] as formulated in Fig. 12b. It turns out that the ratio kdiflkmit is in the range 15-20 so that diffusion does not contribute significantly as a rate-limiting step with stirred suspensions in the absence of HK. However, as the rate constant for HK increases (kmit -e kdif 5 ~ H K ) ,diffusion becomes more limiting and the relative accumulation of ATP at the surface increases. While this relatively simple model does not reflect the physical reality, it is useful to describe the effective properties of the system. A more exact (and elaborated) mathematical model has been developed (Aflaloand Segel, 1991),which describes the experimental data with less “unrealistic” assumptions. The steady-state value of local [ATP] in the presence of excess HK is 5-10% of the total nucleotide concentration added to isolated mitochondria. This value represents a lower limit for the true concentration gradient, since HK may selectively associate with the outer membrane of yeast mitochondria (Krause et al., 1986; but see Kovac et al., 1986). Another aspect which was not considered is the microdistribution of luciferase on the outer membrane. Indeed, the membrane-bound enzyme would detect only the fraction of “vectorial” ATP (emerging from the porin molecules) which has diffused laterally along the membrane. In contrast, “scalar” ATP added to the medium would be simultaneously and equally accessible to the membrane probe, in the absence of a net flux for its formation or utilization. The resolution of this requires a finer resolution for luciferase localization, using, for example, immunogold labeling of mitochondria preparations (Douma et al., 1985; Hines et al., 1990). Finally, a substantially higher diffusion limitation should also be expected in the more intricate (and unstirred) cellular environment. The commonly observed aggregation of mitochondria around ATP-consuming sites in some cells represents a trivial solution for the cell to overcome such a limitation.

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

309

MI. Experimental Approaches and Perspectives A. LIGHTMEASUREMENT Various light-measuring instruments may be used for the detection and quantitation of luminescence (for a review of commercially available radiometers, see Stanley, 1986). They include photon counters, which evaluate the number of quanta emitted in a predetermined lapse of time, and luminometers, which measure light energy flux as an electrical current. Many modern radiometers can operate in either mode. Light detection itself is achieved using two main kinds of devices. The photomultiplier is based on a photocathode which acts as an electron emitter when illuminated. The photoelectrons are captured and multiplied by a succession of anodes under increasing voltage. The response (pulse or current), proportional to the incident light over a wide dynamic range, may be amplified up to 108-fold.The second device is the photodiode, or charge-coupled device (CCD), on which each incident photon creates a single electron-hole pair on a siliconjunction. The resulting signal can be amplified to alesser extent than in the photomultiplier, resulting in less sensitivity. On the other hand, photodiodes are sturdier and more compact and may be miniaturized. Recent advances in CCD technology have led to the development of array detectors consisting of two-dimensional matrices of light-sensitive elements (pixels), which enable quantitative imaging at high resolution (500 x 500 pixels/cm*) and real-time measurement capability when coupled to image intensifiers (Hooper et al., 1990). Relatively high sensitivity can be reached using digital image processing of stored signals, including background subtraction, integration, and averaging of the collected data. These techniques have been used to assess the expression of luciferase in single mammalian cells infected with a recombinant vaccinia virus (Hooper et al., 1990), or plant protoplasts after the insertion of (luciferase) mRNA by electroporation (Gallie et al., 1989). Further improvement of the sensitivity and microscopic video imaging is needed to enable real-time analysis of light production at subcellular spatial resolution, which is the ultimate challenge for this rather expensive and sophisticated technology. The use of spectroluminometers has also been described for analysis of the color of light emitted upon varying environmental factors or with different luciferases isolated or cloned from P. plagiophthalamus (Seliger et al., 1964; Wood et al., 1989b). Another approach for semiquantitative light detection involves direct exposure of two-dimensional objects (Petri dishes or microtiter plates) to photographic films (Kricka and Thorpe, 1986; Wood and DeLuca, 1987). This is particularly useful for rapid screening procedures of recombinant colonies or plaques expressing luciferase. Moreover, the film darkening

310

CLAUDE AFLALO

may be quantified by scanning densitometry. This method has also been used to show the spatial distribution of luciferase luminescence from whole recombinant tobacco plants (Ow et al., 1986; Schneider et al., 1990). Liquid scintillation counters may also be used for routine light measurements. In these widely available instruments, the light is collected by two photomultipliers, which can be operated independently, giving a total count from both detectors (chemiluminescence mode), or in conjunction through a coincidence circuit, yielding counts representing luminescent events recorded simultaneously by the detectors (scintillation mode). The rate recorded in the coincidence mode is proportional to the square of the light emitted (Kimmich et al., 1975), in contrast with the first mode which directly measures light intensity, as with common radiometers (Nguyen et al., 1988). While increased sensitivity (i.e., signal-to-background ratio) may be attained by measuring steady light emission over a long time, this method is not suitable for monitoring sudden changes of light output in response to the addition of reagents, since the time required to insert the vial in the counting chamber is relatively long. Presently, the time evolution of luminescence can be assessed at the highest sensitivity as a space-average phenomenon with the light-emitting units in stirred solutions or suspensions. The configuration of the light detection chamber is important for the optimization of light collection from the luminescent object. The light-sensitive (planar) surface should be as close as possible to the object, since the light flux decreases drastically with the distance. Thus, when a single detector is used, light emitted from the side of the object opposite the detector is lost unless it is reflected back to the detector by a surface of appropriate geometry. Some commercially available luminometers are equipped with mixing devices which insure a homogeneous distribution of the light emitters and a fast dilution of added reagents, and help to sustain an adequate oxygen concentration in respiring biological suspensions (see Figs. 9C and D).

B, PRACTICAL CONSIDERATIONS 1. Use of FL as a Probe for Metabolites Concentrations

The quantum yield of luciferase bioluminescence [i.e., photon : molecule of luciferin consumed = 0.88 (McElroy and Seliger, 1961)l is one of the highest known in photochemistry. The turnover rate for catalysis is rather low compared to other enzymatic reactions (0.01-0.03 sec-', saturated). The combination of these features represents the basis for using luciferase as a probe, since the light signal can be measured at

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

31 1

high sensitivity with a relatively slow consumption of the substrate being monitored. In addition, the reaction is very specific toward ATP, and light production is only marginally affected by other nucleoside di- or triphosphates (Lee ef al., 1970). Both the peak and steady-state light intensities are proportional to the amount of luciferase in the reaction. The fast-evolving peak value is routinely used to determine the native or recombinant enzyme concentration in biological samples at high sensitivity and with relatively low interference from concurrent reactions (e.g., ATP or 02 consumption) likely to occur in the extracts. Of course, the most attractive feature of this analytic system is the linear relationship between light output and the concentration of the limiting substrate, sustained in a wide range (10-'2-10-6 M for ATP). The linear range can be somewhat expanded by lowering the catalytic rate at the expense of sensivity. This may be done by lowering the cosubstrate concentration or by adding competitive inhibitors (respectively to the limiting substrate being monitored). Another approach is to relieve the bound product inhibition through the addition of neutral surfactants or polar additives (see Kricka and DeLuca, 1982). Under these conditions, in addition to ATP, a wide range of metabolites can be determined when luciferase is coupled to enzymes which use them as cosubstrates in ATPgenerating or -consuming reactions. Moreover, the activity of such enzymes in biological extracts can be similarly monitored in the continuous mode by following the time course of ATP formation or consumption. Light evolution as a response to variation in ATP concentration occurs generally fast enough (-0.5 seconds) to permit such an application. The reader is referred to a comprehensive review of these aspects by Lundin (1982). A major drawback of this method is that a linear response is elicited only by relatively low (i.e., submicromolar) [ATP]. This does not permit the study of enzymes or more complex biological systems under physiologically relevant conditions with respect to ATP whose average concentration in cells is in the millimolar range. Another limitation with fast-kinetics systems may be the time dependence of light production by luciferase. Lemasters and Hackenbrock (1979) have developed calibration and data manipulation procedures to deal with the dependence on [ATP] in the continuous monitoring of oxidative phosphorylation in isolated mitochondria. Moreover, pre-steady-state kinetic data for luminescence can be converted to the real variation of ATP in fast reactions by calculation using kinetic models based on the study of purified enzyme (Aflalo and DeLuca, 1987, 1988; Slooten and Vandenbranden, 1989; C. Aflalo, unpublished observations). Nevertheless, the straightforward use of luciferase as a

312

CLAUDE AFLALO

probe for ATP (or other substrates) concentration is limited to controled conditions in which the latter is kept in the micromolar range. This may be achieved only with isolated or permeabilized systems, or alternatively, if luciferase were to be directed to cellular compartments with either intrinsically low ATP content or a high ATP consumption ability.

2. Genetic and Chemical Engineering of Luciferase In principle, with appropriate structural and mechanistic information from luciferase, it may be possible to modify the enzyme so it will be more suitable for probing ATP in a cellular environment. The aim would be to decrease the apparent affinity for ATP in order to maintain a linear relationship between the steady light output and [ATP] in the physiological range. However, it may be anticipated that a modification of the active site by covalent derivatization or mutagenesis of specific residues could also affect other characteristics of the enzyme relevant to its use as a probe. Ideally, the modification should exclusively shift the luciferin activation equilibrium (step 1 in Section 111,B,3) to the left, with no effect on the specificity toward ATP (or luciferin), or the low turnover rate which enables noninteracting continuous measurements. This prospective approach is hindered by the uncertain validity of active site-directed chemical modification data acquired before the primary structure of the enzyme was resolved. Three partial sequences of peptides derived from luciferase chemically derivatized with reactive substrate analogs have been reported (Travis and McElroy, 1966; Lee and McElroy, 1971b; Lee et al., 1981).None of these putative active site-relatedpeptides could be recognized in the translated sequence for cloned luciferase (Wood, 1989). So far, a first attempt for mutagenesis of nonessential but catalytically related (Alter and DeLuca, 1986) cysteine residues to alanine did not affect the activity of luciferase expressed in bacteria(D. C. Vellom, personal communication). Thus, it seems that some aspects of the early characterization of the enzyme involving active site-directed chemical modification need reinvestigation in conjunction with the genetic approaches currently used to study the color of light emitted by different beetle luciferases (Wood, 1990; Wood et al., 1989b). The latter are oriented to map the luciferin binding site. A parallel study is needed for the pyrophosphate (in ATP) binding site, which is more relevant to the proposed modification. The process of the cloning and direction of luciferase to foreign cells and compartments involves genetic manipulation of the coding sequence, and may, a priori, result in a change in the enzyme properties. There is no available experimental evidence for that in eukaryotic cells. However, with bacteria transformed with luc, a large amount of antiluciferase reac-

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

313

tive protein may be detected in an insoluble fraction which contains no bioluminescent activity. The effect of short deletions from either the 3' or 5' end of the coding region in luc cloned in bacteria was studied systematically. The total light production in extracts is drastically reduced when as few as eight residues are deleted from either the amine or carboxy terminus (Wood, 1989). The decrease in activity with longer deletions is correlated with the amount of luciferase in soluble extracts, determined by Western blot analysis. These results suggest that the deletions affected the stability of luciferase in the bacterial cell environment rather than the intrinsic catalytic ability of the truncated products. The terminal fragments may contribute to the solubilization of the enzyme or prevent interactions between hydrophobic domains on different molecules normally located in the core of the protein.

C. PERSPECTIVES 1 . Luciferase as a Probe for Protein Metabolism

A complete picture for gene expression should include information on the various processes leading to the establishment of the final phenotype. These include the transcription and translation of genetic material, targeting (processing) and assembly of the resultant polypeptide chain(s), and the catabolic degradation of the mature protein. This pathway is generally studied through the chemical determination of translated protein in extracts by radioactive (pulse-chase labeling) or immunochemical methods, which provide sensitive means to determine the amount of protein. However, this approach does not provide information on the functionality of the polypeptides investigated. Catalytic ability is the best criterion for correct folding and assembly of translated enzymes, but it is often quantitated with much lower sensitivity compared with the chemical methods. Therefore, correlation between structural and functional expression is difficult to test with most systems, especially in the case of low-level gene expression, which is often required for viable cells. Numerous applications of cloned luciferases as reporter genes have been reviewed by Gould and Subramani (1988), Wood et al. (1989a), and Wood (1990). All take advantage of the unique light emission ability of the enzyme from firefly (or other organisms) in the biological systems investigated. The ease and sensitivity for determination of light emission confer to luciferase a significant advantage over different probes for protein metabolism. Indeed, bioluminescence from cloned luciferase may be quantitated at noninvasive levels of expression. This insures a low interference of the enzyme with other cellular systems. In genetic applications, as opposed to the use of the enzyme as a chemical sensor, the limiting

314

CLAUDE AFLALO

factor for light emission is the concentration of the enzyme itselfin biological samples supplemented with saturating substrates. The focus of these applications is the regulation of active enzyme synthesis, and the posttranslational or degradation aspects have been only marginally considered. A few studies of the stability of the enzyme in foreign environments have emerged which use the enzyme activity (Kutuzova et al., 1989). However, the information gathered by activity measurements alone may be incomplete and must be confronted with chemical determinations as mentioned above. An example of discrepancy between both approaches in an overexpression bacterial system was given in Section VI,B,2. Studies of the stability of cloned luciferase to heat or related stress assessed by both the methods indicate that the enzyme is not degraded, but becomes insoluble as the activity is lost (Nguyen et al., 1989). This illustrates the use of the enzyme as a promising tool to study the molecular mechanisms of thermolability and resistance to stress. This can be extended to wider studies of the interaction of the enzyme with the cellular machineries involved in protein traffic and processing. The possibility of formation of a tightly bound product (e.g., L-AMP in Fig. 2) enables the possible isolation of a translocation intermediate, since the tightly folded polypeptide chain will not cross the membrane (see Section 11,A). Finally, the analysis of experimental cases in which the fusion protein failed to be localized in an active form may provide important information on the cellular import machinery. 2 . Targeting Luciferase to Cellular Compartments New fusion genes may be designed to direct luciferase to various cellular (micro)compartments.The fusion genes should be constructed by isolating appropriate restriction fragments encoding for leader sequences directing natural proteins to specific cellular compartments in cells (see Section 11,A). These can be ligated to the FL gene, and the resulting fusion genes can be inserted into shuttle plasmids behind selected promoters. These plasmids may be propagated in bacteria. The selection of positive transformants is easily performed by screening colonies for bioluminescence using luminometric or photographic standardized assays (Wood and DeLuca, 1987). After reisolation of the plasmids, they can be used to transform eukaryotic cells. Care should be taken to choose appropriate lengths of the fused sequences to minimize their possible interaction with the luciferase moiety until the final assembly of the protein. Also, the peroxisomal targeting sequence at the carboxy terminus of luciferase should be deleted or modified to avoid ambiguity in the final localization of fusion proteins. The use of fusion genes has been proposed (Froshauer et al., 1988) to determine membrane protein topology, normally assessed by the interac-

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

315

tion of proteases or antibodies with hydrophilic domains exposed to the medium. The approach is based on the creation of a chimeric gene consisting of the coding sequence of a soluble enzyme fused in frame with fragments of varying length encoding for the membrane protein. Thus, the soluble enzyme domain should appear on the same side of the membrane as the point to which it was fused in the original membrane protein. Walter and associates (Green et al., 1989) have applied this approach with soluble galactokinase fused to various fragments of yeast arginine permease, a plasma membrane protein including multiple trans-membrane segments. They used the glycosylation of galactokinase in the lumen of microsomal vesicles to determine its location relative to the membrane. One can take advantage of the characteristics of luciferase in a similar system to assess the topology, since the membrane-bound enzyme responds differently to substrate addition according to its relative location. While a flash emission of high intensity is expected with the enzyme exposed to the medium (saturating [ATP] and [LHz], pH 7.8), a very low light output should develop slowly with the enzyme inside the vesicles, due to the low accessibility of substrates (see Section V1,A). Of course, this “activity-oriented” approach may (and should) be complemented by the physical demonstration of exclusive localization of the protein in the correct compartment with the exclusive topology. This may be conveniently done by immunogold labeling (Keller et al., 1987; Douma et al., 1985) of whole-cell preparations for electron microscopy. It remains to be demonstrated that the luciferase activity will be retained in the various fusion proteins during or after their translocation through biological membranes and compartment-specific processing. Experimental evidence to date has shown that the enzyme is active in peroxisomes (de Wet et al., 1987), cytoplasm (Gould et al., 1987), mitochondria (Aflalo, 1990), and chloroplasts (Schneider et al., 1990). An intriguing and highly speculative prospect would be the design of chimeric proteins consisting of FL fused to peptide domains known to interact specifically with proteins or structures localized in the cell. For example, mellitin-luciferase or troponin-luciferase chimerae would bind to calmodulin or actin filaments, respectively. Such an approach may provide significant refinements to the biological localization of the probe. However, the probability of adverse interaction with the target is expected to rise due to steric hindrance from the large luciferase domain. 3. Prospects for Local [ATP] Measurements Most isolated cells have a stable adenylate concentration which may be modulated experimentally to some extent. The steady-state average concentration of cellular ATP is determined by the ratio of its rate of formation

3 16

CLAUDE AFLALO

(mainly by oxidative phosphorylation and glycolysis) to that of consumption by various energy-requiring systems. Thus, modulation of the activity of these systems by addition of exogenous substrates or inhibitors may be used to control the total ATP content of cells. Morever, it is possible to modulate [ATP] in selected compartments by specifically affecting localized processes (see Section 11,D). Although a most significant reduction of [ATP] in compartments sustaining a high rate of its consumption is conceivable, it remains doubtful that one could reach ATP levels suitable for linear analysis by luciferase in most compartments of viable cells. Indeed, the calculated [ATP] in cells is higher than the latter by three orders of magnitude. As a noninteracting probe, it is expected that the enzyme will detect only freely diffusible ATP in its microenvironment as opposed to the possibly channeled ATP which is committed in the respective reactions of its production and utilization. An interesting prospect for localized luciferase is the assessment of free ATP in concentrated cellular phases as the mitochondria1 matrix, or muscular filaments which have a potentially high nucleotide binding capacity. The use of luciferase as a local probe for ATP seems more adequate in isolated and/or reconstituted systems artificially depleted from adenine nucleotides. Channeling of ATP may be demonstrated when ATP is preferentially utilized in a selected system from a specific source, and the operation of the selected system is relatively unaffected by ATP generated or consumed by other systems. This rationale was illustrated in Sections IV,A and VI,B in an artificial and a biological system, respectively. In both cases, tunneling of ATP between the site of its formation and the proximal luciferase was due to its slow transfer by diffusion to a remote compartment, in which it is efficiently consumed. In intact hepatocytes, a similar reasoning has been used to propose the establishment of [ATP] gradients (Aw and Jones, 1985). In erythrocytes (Mercer and Dunham, 1981) and smooth muscle (Lynch and Paul, 1987), ion transport by Na+, K+-ATPase at the plasma membrane appears to be tightly coupled to glycolysis, and relatively independent of the bulk (cellular) [ATP]. The reduction of ion transport rate under aerobic conditions in the absence of glycolysis cannot be explained only by diffusional restrictions for ATP transfer between mitochondria and the plasma membrane, in view of the high concentration of bulk cellular ATP, and its expected diffusivity, although significant gradients of ADP could be predicted by the same models (Lynch and Paul, 1988).Thus, the mode of interaction between the coupled systems must be elucidated in more detail. To date, a single attempt was made to demonstrate the direct transfer of ATP between kinases (Dillon and Clark, 1990), as for NADH with dehydrogenases (Srivastava and Bernhard, 1986a,b). Such a mechanism may

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

317

provide an alternative explanation for the experimentally inferred channeling of nucleotides in cells. Some systems have been reconstituted in vitro from isolated catalytic components to better assess the coupling mechanisms and the kinetic features resulting from physical association (see Masters, 1981; Keleti and Ovadi, 1988; Wilson, 1988). The use of luciferase colocalized with ATP-consuming systems, in conjunction with soluble (delocalized) luciferase, represents an independent means to assess the fate of ATP in such a reconstituted coupled system. Under the experimental conditions used to test for direct transfer (Srivastava and Bernhard, 1986a,b), the concentration of free ATP would be extremely low. 4 . Monitoring Local Events in Cryptic Compartments As mentioned in Section VI,A, the most probable limiting factor for light production in normal cells is the local luciferin concentration. A relatively simple model system for light emission in situ is the soluble enzyme expressed in bacteria. The study of light production in permeabilized cells should help to define the contribution of some environmental factors. This situation is essentially analogous to the model system described in Section IV (effect of colocalized enzyme activity). The next step would be the determination of internal ATP and luciferin concentration (diffusing in from the medium) in intact cells under different metabolic states expected to affect the limiting conditions. This can be done in extracts from cells rapidly separated from the suspension medium. The light output from the localized enzyme in these conditions may be confronted with these measurements in order to better define the limitation on light production imposed by these factors. For example, the addition of glucose to cells incubated at low pH in the presence of citrate causes a marked transient reduction in the steady light output (see also the effect of citrate itself, shown in Fig. 9,C and D). On the other hand, the addition of uncouplers results in a large increase in light production, followed by an irreversible inhibition. These preliminary data indicate that the luciferin-limited variation of light output may reflect repartition of luciferin associated with proton movements. At saturating luciferin concentrations, local changes in pH or divalent cation concentration may also affect the intensity and the color of light emitted by the enzyme. Thus, light emission by luciferase restricted to defined compartments may indicate specifically a wide range of events occurring locally, which are not detectable by conventional means. A systematic study of the dependence of luminescence on various factors with the enzyme in solution may help to design calibration procedures for the localized enzyme.

318

CLAUDE AFLALO

With luciferase localized in various defined cellular compartments, a survey of metabolism in situ may be initated through assessment of the local concentration of reactants using the different probes. This long-range perspective would be applicable only after adequate calibration procedures are developed for the probe in cryptic compartments. So far, our present experimental and theoretical modeling abilities fall short of this ideal use of luciferase. However, the unique properties of bioluminescent enzymes and the possibility to localize them specifically at various cellular sites enable novel approaches to integrative studies of the structure and function of biological systems. These may provide valuable information on assembly processes and local events previously inferred indirectly by conventional methodologies. The systematic analysis of the latter should promote the development of more focused theoretical treatments for metabolic function in situ, which could, in turn, be tested directly in model systems. Finally, these models may be useful to elucidate complex problems in cellular metabolism in which the use of local probes is not presently applicable. ACKNOWLEDGMENT This chapter is dedicated to the memory of Marlene DeLuca, whose spirit, it is hoped, is reflected here. The support of the Israel Ministry of Integration to the author is gratefully acknowledged.

REFERENCES Aflalo, C. (1990). Biochemistry 29,4758-4766. Aflalo, C., and Segel, L. A. (1991). Manuscript in preparation. Aflalo, C., and DeLuca, M. (1987). Biochemistry 26,3913-3919. Aflalo, C., and DeLuca, M. (1988).In “Ion Pumps: Structure, Function and Regulation” (W. Stein, ed.), pp 337-342. Liss, New York. Aflalo, C., and Shavit, N. (1982). Eur. J . Biochem. U6,61-68. Aflalo, C., and Shavit, N. (1983). FEBS L e f t .154, 175-179. Aflalo, C., and Shavit, N. (1984). Curr. Top. Cell. Regul. 24,435-445. Alter, S . C., and DeLuca, M. (1986). Biochemistry 25, 1599-1605. Aw, T. Y., and Jones, D. P. (1985). Am. J . Physiol. 249, C385-C392. Beard, W. A., and Dilley, R. A. (1986). FEBS Left.201,57-62. Bessman, S. P., and Carpenter, C. L. (1985). Annu. Rev. Biochem. 54,831-862. Blobel, G . (1983). In “Methods in Enzymology” ( S . Fleischer and B. Fleischer, eds.), Vol. 96, pp. 663-691. Academic Press, New York. Boag, J. W. (1969). Curr. Top. Radiaf. Res. 5 , 141-195. Brdiczka, D., Bucheler, K., Kottke, M., Adams, V., and Nalam, V. K. (1990). Biochim. Biophys. A c f a 1018,234-238. Carlier, M.-F. (1989). Inr. Reu. Cytol. 115, 139-170. Chance, B., Oshino, N., Sugano[T., and Mayersky, A. (1973). Ado. Exp. Med. Biol. 37A, 277-292.

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

319

Chen, W. J., and Douglas, M. G. (1987). J. Biol. Chem. 262, 15605-15609. Cheung, C.-W., Cohen, N. S., and Raijman, L. (1989). J . Biol. Chem. 264,4038-4044. Clegg, J. S. (1984). Am. J . Physiol. 246, R133-R151. Colman, A., and Robinson, C. (1986). Cell 46,321-323. Cormier, M. J. (1978). I n “Bioluminescence in Action” (P. J. Hemng, ed.), pp. 75-108. Academic Press, New York. DeLuca, M. (1969). Biochemistry 8, 160-166. DeLuca, M. (1984). Curr. Top. Cell. Regul. 24, 189-195. DeLuca, M., and Marsh, M. (1%7). Arch. Biochem. Biophys. 121,233-240. DeLuca, M., and McElroy, W. D. (1974). Biochemistry W , 921-925. DeLuca, M., and McElroy, W. D. (1978). I n “Methods in Enzymology” (M. A. DeLuca, ed.), Vol. 57, pp. 3-15. Academic Press, New York. DeLuca, M., and McElroy, W. D. (1984). Biochem. Biophys. Res. Commun. 123,764-768. DeLuca, M., Leonard, N. J., Gates, B. J., and McElroy, W. D. (1973). Proc. Natl. Acad. Sci. U.S.A. 70, 1664-1669. Deshaies, R. J., Koch, B. D., and Scheckman, R. (1988). Trends Biochem. Sci. W , 384-388. de Wet, J. R., Wood, K. V., Helinski, D. R., and DeLuca, M. (1985). Proc. Natl. Acad. Sci. U.S.A. 82,7870-7873, de Wet, J. R., Wood, K. V., DeLuca, M., Helinski, D. R., and Subramani, S. (1987). Mol. Cell. Biol. 7,725-737. DiLella, A. G., Hope, D. A., Chen, H., Tmmbauer, M., Schwartz, J., and Smith, R. G. (1988). Nucleic Acids Res. 16,4519. Dillon, P. F., and Clark, J. F. (1990). J . Theor. Biol. 143,275-284. Distel, B., Veenhuis, M., and Tabak, H. F. (1987). EMBO J . 6 , 3 1 1 1-3 116. Douglas, M. G., McCammon, M. T., and Vassarotti, A. (1986).Microbiol. Rev. 50,166-178. Douma, A. C., Veenhuis, M., de Koning, W., Evers, M., and Harder, W. (1985). Arch. Microbiol. 143,237-243. Easterby, J. S. (1981). Biochem. J . 199, 155-161. Eilers, M., and Schatz, G. (1986). Nature (London) 322,228-232. Ellis, R. J., and Hemmingsen, S. M. (1989). Trends Biochem. Sci. 14,339-342. Engassser, J.-M., and Hovath, C. (1976). Appl. Biochem. Bioeng. 1,127-220. Froshauer, S . , Green, G., Boyd, D., McGovern, K., and Beckwith, J. (1988). J. Mol. Biol. u)0,501-511. Fukushima, T., Decker, R. V., Anderson, W. M., and Spivey, H. 0. (1989). J. Biol. Chem. 264, 16483-16488. Gallie, D. R., Lucas, W. J., and Walbot, V. (1989). Plan? Cell 1,301-31 1. Garfinkel, D. A. (1963). J. Biol. Chem. 238,2435-2439. Garfinkel, D. A., and Lajtha, A. (1963). J. Biol. Chem. 238,2429-2434. Gates, B. J., and DeLuca, M. (1975). Arch. Biochem. Biophys. 169,618-621. Goldman, R. (1973). Biochimie 55,953-966. Goldman, R., and Katchalski, E. (1971). J. Theor. Biol. 32, 243-257. Goldstein, L. (1976). In “Methods in Enzymology” (K. Mosbach, ed.), Vol. 44, pp. 397-443. Academic Press, New York. Gould, S. J., and Subramani, S. (1988). Anal. Biochem. 175,5-13. Gould, S . J., Keller, G.-A., and Subramani, S. (1987). J. Cell Biol. 105, 2923-2931. Gould, S. J., Keller, G.-A., Hosken, N., Wilkinson, J., and Subramani, S. (1989). J. Cell Biol. lOS, 1657-1665. Gould, S. J., Keller, G.-A., Schneider, M., Howell, S. H., Garrard, L. J., Goodman, J. M., Distel, B., Tabak, H., and Subramani, S . (1990a). EMBO J . 9,85-90. Gould, S. J., Krisans, S., Keller, G.-A., and Subramani, S . (1990b). J. Cell Biol. 110,27-34.

320

CLAUDE AFLALO

Green, A. A., and McElroy, W. D. (1956). Biochim. Biophys. Acta 20, 170-178. Green, G. N., Hansen, W., and Walter, P. (1989). J . Cell Sci.,Suppl. 11, 109-113. Grivell, L. A. (1986). I n t . Rev. Cytol. 3, 107-141. Hartl, F.-U., and Neupert, W. (1989). J. Cell Sci.,Suppl. 11, 187-198. Harvey, E. N. (1957). “A History of Luminescence from the Earlier Ages until 1900.” Am. Philos. SOC.,Philadelphia, Pennsylvania. Hase, T., Reizman, H., Suda, K., and Schatz, G. (1983). E M B O J . 2,2169-2172. Hase, T., Muller, U., Reizman, H., and Schatz, G. (1984). E M B O J . 3,3157-3164. Hines, V.,Brandt, A., Griffiths, G . , Horstmann, H., Brutsch, H., and Schatz, G. (1990). EMBO J . 9, 3191-3200. Hirokawa, N. (1982). J. Cell B i d . 94, 129-142. Hooper, C. E., Ansorge, R. E., Browne, H. M., and Tomkins, P. (1990). J . Biolumin. Chemilumin. 5 , 123-130. Hurt, E. C., Pesold-Hurt, B., and Schatz, G. (1984). EMBOJ. 3,3149-3156. Hurt, E. C., Pesold-Hurt, B., Suda, K., Oppliger, W., and Schatz, G . (1985). EMBO J . 4, 2061-2068.

Johnson, L. M., Bankaitis, V. A,, and Emr, S. D. (1987). Cell 48,875-885. Jones, D. P. (1986). Am. J . Physiol. 250, C663-C675. Jones, D. P., and Aw, T. Y. (1990). I n “Structural and Organizational Aspects of Metabolic Regulation” (P. Srere, M. E. Jones, and C. Mathews, eds.), pp. 345-361. Liss, New York. Kalderon, D., Roberts, B. L., Richardson, W. D., and Smith, A. E. (1984). Cell 39,499-509. Kaprelyants, A. S. (1988). Trends Biochem. Sci. W, 43-46. Keegstra, K. (1989). Cell 56,247-253. Keleti, T., and Ovadi, J. (1988). Curr. Top. Cell. Regul. 29, 1-33. Keller, G.-A., Could, S., DeLuca, M., and Subramani, S. (1987). Proc. Natl. Acad. Sci. U.S.A. 84,3264-3268. Kernevez, J.-P. (1980). “Enzyme Mathematics.” North-Holland Publ., Amsterdam. Kimmich, G. A., Randles, J., and Brand, J. S. (1975). Anal. Biochem. 69, 187-206. Knull, H. R. (1990). I n “Structural and Organizational Aspects of Metabolic Regulation” (P.Srere, M. E. Jones, and C. Mathews, eds.), pp. 215-228. Liss, New York. Kovac, L., Nelson, B. D., and Emster, L. (1986). Biochem. Biophys. Res. Commun. W, 285-29 1.

Krause, J., Hay, R., Kowollik, C., and Brdiczka, D. (1986). Biochim. Biophys. Acta 860, 690-698.

Kricka, L. J. (1988). Anal. Biochem. 175, 14-21. Kricka, L. J., and DeLuca, M. (1982). Arch. Biochem. Biophys. 217,674-681. Kricka, L. J., and Thorpe, G. H. G. (1986). I n “Methods in Enzymology” (M. A. DeLuca and W. D. McElroy, eds.), Vol. 133, pp. 404-420. Academic Press, Orlando, Florida. Kurganov, B. I. (1986). J. Theor. B i d . 119,445-455. Kutuzova, G . D., Skripkin, E. A., Tarasova, N. I., Ugarova, N. N., and Bogdanov, A. A. (1989). Biochimie 71,579-583. Lazarow, P. B., and Fujiki, Y. (1985). Annu. Rev. Cell. B i d . 1,489-530. Lee, R. T., and McElroy, W. D. (1971a). Arch Biochem. Biophys. 145,78-84. Lee, R. T., and McElroy, W. D. (1971b). Arch. Biochem. Biophys. 146,551-556. Lee, R. T., Denburg, J. L., and McElroy, W. D. (1970). Arch. Biochem. Biophys. 141,38-52. Lee, Y . , Esch, F. S., and DeLuca, M. (1981). Biochemistry 20, 1253-1256. Lemasters, J. J., and Hackenbrock, C. R. (1977). Biochemistry 16,445-447. Lemasters, J. J., and Hackenbrock, C. R. (1979). I n “Methods in Enzymology” (S.Fleischer and L. Packer, eds.), Vol. 56, pp. 530-544. Academic Press, New York.

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

321

Lira, S. A., Kinloch, R. A., Mortillo, S., and Wassarman, P. M. (1990). Proc. Natl. Acad. Sci. U . S . A . 87,7215-7219. Lundin, A. (1982). In “Clinical and Biochemical Luminescence” (L. J. Kricka and T. J. N. Carter, eds.), pp. 43-74. Dekker, New York. Lundin, A., Rickardsson, A., and Thore, A. (1976). Anal. Biochem. 75,611-620. Lynch, R. M., and Paul, R. J. (1987). Am. J. Physiol. 252, C328-C339. Lynch, R. M., and Paul, R. J. (1988). In “Microcompartmentation” (D. P. Jones, ed.), pp. 17-35. CRC Press, Boca Raton, Florida. Magee, A. I., Gutierrez, L., Marshall, C. J., and Hancock, J. F. (1989). J. Cell Sci., Suppl. 11, 149-160. Malone, R. W., Felner, P. L., and Verma, I. M. (1989). Proc. Narl. Acad. Sci. U.S.A. 86, 6077-6078.

Masters, C . J. (1981). CRC Crit. Rev. Biochem. 11, 105-144. Masuda, T., Tatsumi, H., and Nakano, E. (1989). Gene 77,265-270. Mattiasson, B., and Mosbach, K. (1971). Biochim. Biophys. Acra 235,253-257. McElroy, W. D., and Seliger, H. H. (1961). In “Symposium on Light and Life” (W. D. McElroy and H. B. Glass, eds.), pp. 219-257. Johns Hopkins Univ. Press, Baltimore, Maryland. Mercer, R. W., and Dunham, P. B. (1981). J . Gen. Physiol. 78,547-568. Minton, A. P. (1983). Mol. Cell. Biochem. 55, 119-140. Miska, W., and Geiger, R. (1987). J. Clin. Chem. Clin. Biochem. 25,23-30. Morton, D. J., Wiedemann, J. F., Clarke, F. M., Stephan, R., and Stewart, M., (1982). Micron W,377-379. Morton, R. A., Hopkins, T. A., and Seliger, H. H. (1969). Biochemistry 8, 1598-1607. Mosbach, K. (1978). Curr. Top. CeN. Regul. 14, 197-241. Murthy, M. S. R., and Pande, S. V. (1985). Biochem. J. 230,657-663. Nguyen, V. T., Morange, M., and Bensaude, 0. (1988). Anal. Biochem. 171,404-408. Nguyen, V. T., Morange, M., and Bensaude, 0. (1989). J . Biol. Chem. 264, 10487-10492. Ono, H., and Tuboi, S. (1988). J . Biol. Chem. 263,3188-3193. Ottaway, J. H., and Mowbray, J. (1977). Curr. Top. Cell. Regul. 12, 107-208. Ow, D. W., Wood, K. V.,DeLuca, M., de Wet, J. R., Helinski, D. R., and Howell, S. H. (1986). Science 234, 856-859. Pain, D., Kanwar, Y. S., and Blobel, G. (1988). Nature (London) 331,232-237. Pfanner, N., Pfaller, R., and Neupert, W. (1988). Trends Biochem. Sci. W, 165-167. Porter, K. R. (1984). J . Cell. Biol. 99,3~-12s. Rassow, J., Guiard, B., Wienhues, U., Herzog, V., Hartl, F.-U., and Neupert, W. (1989). J . Cell Biol. 109, 1421-1428. Reizman, H., Hase, T., van Loon, A. P. G. M., Grivell, L. A., Suda, K., and Schatz, G. (1983). EMBO J . 2,2161-2168. Rhodes, W. C., and McElroy, W. D. (1958). J . B i d . Chem. 233, 1528-1537. Rodriguez, D., Rodriguez, J.-R., Rodriguez, J. F., Trauber, D., and Esteban, M. (1989). Proc. Narl. Acad. Sci. U.S.A. 86, 1287-1291. Roise, D., and Schatz, G. (1988). J. Biol. Chem. 283, 4509-4511. Rosen, G., Gresser, M., Vinkler, C., and Boyer, P. D. (1979). J. Biol. Chem. 254, 1065410661.

Rothman, J. H., Yamashiro, C. T., Kane, P. M., and Stevens, T. H. (1989). Trends Biochem. Sci. 14,347-350.

Ryazanov, A. G . , and Spirin, A. S. (1989). Biochemistry (Engl. Transl.) 54,558-563. Schliwa, M., van Berkom, J., and Porter, K. R. (1981). Proc. Narl. Acad. Sci. U.S.A. 78, 4329-4333.

322

CLAUDE AFLALO

Schneider, M., Ow, D. W., and Howell, S. H. (1990). Plant Mol. Biol. 14,935-947. Schram, E., Ahmad, M., and Moreels, E. (1981). I n “Bioluminescence and Chemluminescence” (M. DeLuca and W. D. McElroy, eds.), pp 491-496. Academic Press, New York. Schroder, J. (1989). Nucleic Acids Res. 17,460. Seis, H., and Chance, B. (1970). FEBS Lett. 11, 172-176. Seliger, H. H., and McElroy, W. D. (1964). Proc. Natl. Acad. Sci. U.S.A. 52,75-81. Seliger, H. H., Buck, J. B., Fastie, W. G., and McElroy, W. D. (1964). J. Gen. Physiol. 48, 95-104. Slooten, L., and Vandenbranden, S. (1989). Biochim. Biophys. Acra 976, 150-160. Smeekens, S., Bauerle, C., Hageman, J., Keegstra, K., and Weisbeek, P. (1986). Cell 46, 365-375. Srere, P. A. (1987). Annu. Rev. Biochem. 56,89-124. Srivastava, D. K., and Bernhard, S. A. (1986a). Curr. Top. Cell. Regul. 28, 1-68. Srivastava, D. K., and Bernhard, S. A. (1986b). Science 234, 1081-1086. Srivastava, D. K., and Bernhard, S. A. (1987). Biochemistry 26, 1240-1246. Srivastava, D. K., Bernhard, S. A., Landridge, R., and McClarin, J. A. (1985). Biochemistry 24,629-635. Stanley, P. E. (1986). I n “Methods in Enzymology” (M. A. DeLuca and W. D. McElroy, eds.), Vol. 133, pp. 587-603. Academic Press, Orlando, Florida. Tamura, M., Hazeki, O., Nioka, S., andchance, B. (1989).Annu. Rev. Physiol. 51,813-837. Tillmann, U., and Bereiter-Hahn, J. (1986). Cell Tissue Res. 243,579-585. Tolbert, N. E. (1981). Annu. Rev. Biochem. SO, 133-157. Travis, J., and McElroy, W. D. (1966). Biochemistry 5,2170-2175. Ugarova, N. N. (1989). Biochemistry (Engl. Transl.) 54,580-584. Ugarova, N. N., and Dukhovitch, A. F. (1987). I n “Bioluminescence and Chemiluminescence: New Perspectives” (J. Scholmerich, R. Andreesen, A. Kapp, M. Emst, and W. G. Woods, eds.), pp. 409-412. Wiley, Chichester, England. Vale, R. D. (1987). Annu. Rev. Cell. Biol. 3,347-378. Vestweber, D., and Schatz, G. (1988). J. Cell Biol. 107,2037-2043. Vestweber, D., Brunner, J., Baker, A., and Schatz, G. (1989). Nature (London) 341, 205-209. Vijaya, S., Narayanaswamy, E., Zavala, F., and Moss, B. (1988). Mol. Cell. Biol. 8, 17091714. von Heijne, G. (1986). EMBO J . 5, 1335-1342. Walter, P., and Lingappa, V. R. (1986). Annu. Rev. Cell. Biol. 2,499-516. Welch, G. R., Keleti, T., and Vertessy, B. (1988).J . Theor. Biol. WO, 407-422. Westerhoff, H.V., Melandri, B. A., Venturoli, G., Azzone, G. F., and Kell, D. B. (1984). Biochim. Biophys. Acta 768,257-292. Westerhoff, H. V., Kell, D. B., Kamp, F., and van Dam, K. (1988). I n “Microcompartmentation” (D. P. Jones, ed.), pp. 115-154. CRC Press, Boca Raton, Florida. White, E. H., Steinmeitz, M. G., Miano, J. D., Wildes, P. D., and Morland, R. (1980). J. Am. Chem. SOC. 102,3199-3208. Whitehead, T. P., Kricka, L. J., Carter, T. J. N., and Thorpe, G. H. G. (1979). Clin Chem. 25, 1531- 1546. Wienhausen, G., and DeLuca, M. (1986). I n “Methods in Enzymology” (M. A. DeLuca and W. D. McElroy, eds.), Vol. 133, pp. 198-208. Academic Press, Orlando, Florida. Wienhausen, G., Kricka, L. J., Hinkley, J. E., and DeLuca, M. (1982). Appl. Biochem. Biotechnol. 7,463-473.

BIOLOGICALLY LOCALIZED FIREFLY LUCIFERASE

323

Wilson, J. E. (1988). I n “Microcompartmentation” (D. P. Jones, ed.), pp. 171-190. CRC Press, Boca Raton, Florida. Wood, K. V. (1990). J . Biolumin. Chemilumin. 5, 107-114. Wood, K. V. (1989). Ph.D. thesis. University of California, San Diego, California. Wood, K. V., and DeLuca, M. (1987). Anal. Biochem. 161,501-506. Wood, K. V., Lam, Y. A., and McElroy, W. D. (1989a). J . Biohmin. Chemilumin. 4, 289-301. Wood, K. V., Lam, Y. A,, Seliger, H. H., and McElroy, W. D. (1989b). Science 244, 700-702. Wood, K. V., de Wet, J. R., Dewji, N., and DeLuca, M. (1984). Biochem. Biophys. Res. Commun. W, 592-596. Ziegler, M. M., and Baldwin, T. 0. (1981). Curr. Top. Bioenerg. l2,65-113.

This Page Intentionally Left Blank

INDEX

A

Abudefdf marginatus, 137 Accessory cells, appearance, 214-216 Activated B lymphocytes, 12 Agriolimax reticulatus, 134 a- and P-chloride cells in freshwater-adapted fishes, 204-207 a cells, hypertrophy, 212-214 Ameca splendens, 137 Amphibia, presence of MGPB, 140-141 Amphiuma, 39,47 Ampullaria canaliculata, 133 Anadara transversa, 55 Anagasta kuehniella, 128, 129, 151 Anas platyrhynchos, 142, 150 Annelida, presence of MGPB, 125-127 Anolis carolineus, 141 Antibodies as probes for study of plant cell walls, 234-240 applications, 240 purification and specificity, 237-240 raised against cell wall components, 234-237 Aphanius dispar, 137, 138-138, 140 Apical cavity of chloride cells, 193-196 Arion circumscriptus, 133 Arion ater rufus, 133 ATP, probe for ATP in model systems, 287-2% [ATPI at mitochondria1 surface, monitoring local, 302-308 ATP formation in FL, dependence on rate, 305-306 [ATP] measurements of FL

local versus bulk, 302-304 local, prospects, 315-317 Autoimmune aggression in mammals and the MGPB, 163-169 Auxin-induced metabolic turnover of cell wall polysaccharides, 246,247 Aves, presence of MGPB, 142

B

B cell differentiation pathway and immunoglobulin transport immunoglobulin secretion in plasma cells, 29-30 J chain, role of, and assembly of polymeric immunoglobulin, 30 L chains, role of, and appropriate conformation for secretion, 30-31 posttranslational modifications and secretion, 30 intracellular retention of secretory immunoglobulin, 13-19 cysteine tailpiece-dependant retention, 16-19 KDEL sequences and salvage pathway, 14-16 membrane immunoglobulin transport during B cell ontogeny, 20-27 ER degradation versus cell surface transport, 20-22 membrane immunoglobulin recycling, endocytosis and down-regulation, 27-28 325

326

INDEX

membrane immunoglobulin retention in plasma cells, 29 transport of membrane IgM, 22-27 overview, 2-13 activated and memory B lymphocytes, 12 mature IgMIIgD-expressingB lymphocytes, 11-12 plasma cells and end stage of B lymphoid ontogeny, 13 pre-B lymphocytes, 6-7 pre-B-specific surrogate immunoglobulin L chains, 7-10 pre-B to B-cell transition, 10-11 pro-B lymphocytes, 4-6 studies, 1-2 summary, 31-33 8-cells, degeneration, 209-212 (1+3),(1~4)-8-~-Glucan, 243,252-254 effect of in maize coleoptile segments, 253 Balanus eburneous, 128 Barbus aeneus, 140 Barbus conchonius, 166 Basolateral tubular invaginations, 196- 199 Batrachoseps, 72,78 Biologically localized firefly luciferase, see Firefly luciferase Bip gene, in pre-B cells, and intracellular retention, 14-16 Blood-organ bamers, different, 112-1 13 Blood-testis barrier, I 1 1-1 12, see also Male germ cell protective barrier concept of, 157-158 efficacy of in physiological, pathological and experimental conditions, 160-163 terms, 113-1 14 Bombyx mori, 128 Bound steroid hormone hypothesis, 161 Branchiobdella pentadonta, 125 Branchiostomafloridae, 136 BTB, see Blood-testis barrier Byfo arenarum, 140, 141 Bufo byfo, 141 C

Carassius auratus, 140 Carassium carpio, 140

Cell flattening, MB biogenesis and mechanism, 59-64 Cell surface, 193-199 apical cavity, 193-196 tubular system, 196-199 Cell surface transport versus ER degradation, 20-22 Cell wall components, antibodies raised against, 234-237 Cell wall functions, diversity, 233-234 Cell wall polymers, location and metabolism, 240-246 enzymes, 246 polysaccharides, 241-245 (1+3),( 1+4)-P-D-Glucan, 243 other polysaccharides, 244-245 polygalacturonic acids, 243-244 xyloglucan, 241-243 structural glycoproteins, 245 Cellule ramificati, 114 Cellular biogenesis and organization endogenous enzymes, use of as reporters for local concentrations of reactants, 278-280 involvement of physical processes and local catalysis in cellular metabolism, 275-278 protein traffic and assembly, 270-272 structural basis for organization of metabolism, 272-275 Cellular compartments of FL, leading cloning, expression and localization of native luciferase, 2%-297 retargeting to other cellular organelles, 297-299 targeting luciferase to, 314-315 Cellular organelles of FL, retargeting to other, 297-299 Cephalochordata, presence of MGPB, 136 Ceratostoma foliolaturn, 133 Cerithium adansonii, 133, 151 Chlamydomonas cell walls, 236 Chloride cells and accessory cell during transfer of sea-water adapted fishes to fresh water, 217-221 and accessory cells in stenohaline seawater fishes, 216-217 Chloride cells, general ultrastructural features

327

INDEX cell surface, 193-199 apical cavity, 193-196 tubular system, 196-199 endoplasmic reticulum, mitochondria, and cytoskeleton, 202-204 golgi apparatus and vesiculotubular system, 200-202 Chordata, presence of MGPB, 136-143 amphibia, 140-141 aves, 142 cephalochordata, 136 c yclostomata, 136- 137 mammalia, 142-143 reptilia, I4 1-142 teleostei, 137-140 Clarias gariepinus, 140, 170 Cloning, expression and localization of native luciferase, 2%-297 Cnidaria, presence of MGPB, 120-122 Cobitis taenia, 207 Coturnix coturnix, 142 Crustacea, presence of MGPB, 127-128 Cyclostomata, presence of MGPB, 136- I37 Cyprinodon variegatus, 214 Cyprinus carpio, 137 Cysteine tailpiece-dependant retention of secretory immunoglobulin, 16- 19 Bip-dependent model, 19 incomplete assembly model, 19

D Deroceras reticulatum, 133 Desmosomes and gap and septate junctions between barrier-forming cells, 150- 152 desmosomes, 150-151 gap junctions, 151 septate junctions, 151-152 Diapause barrier, 130 Dicotyledons, 249-252 other polysaccharides, 25 1-252 xyloglucan, 249-25 1 Digitonin, use of as lysis buffer, 24 Diodora nubecula, 135 Dipetalonema dessetae, 116, 123 Dithiothreitol- and protease-solubilized ZP, 100 Dolichos biflorus. 249

Down-regulation, and membrane IgM, 27-28 Dugesia biblica, 122-123, 124, 174, 176 E

Echinodermata, presence of MGPB, 135-136 Eisenia foetidus, 125 Electron microscopy and ZP, 99-104 Elliptogenesis of MB cells, 64-65 End stage of B cell ontogeny, 13 Endocytosis, and membrane IgM, 27-28 Endogenous enzymes, use of as reporters for local concentrations of reactants, 278-280 Endoplasmic reticulum, mitochondria, and cytoskeleton, 202-204 Environment, modifications of and mitochondria-rich cells a-and p-chloride cells in fresh water-adapted fishes, 204-207 chloride cells and accessory cell during transfer of sea-water adapted fishes to fresh water, 217-221 chloride cells and accessory cells in stenohaline seawater fishes, 216-217 ultrastructural features of cells during transfer of euryhaline fishes from fresh water to sea water, 207-217 accessory cells, appearance, 214-216 degeneration of p-cells, 209-212 hypertrophy of a cells, 212-214 Enzyme characteristic of FL, 282-287 initial and steady-state kinetics of light production, 285-287 molecular mechanism of light emission, 282-283 structure, 284-285 Enzymes, 246 antibodies raised against plant cell wall, 234, 235 ER degradation versus cell surface transport, 20-22 Erythrocytes, nucleated, cytoskeletal system, see also Nucleated erythrocytes mammalian primitive, 76-79

328

INDEX

mature, marginal band, 43-59 morphogenesis, 69-67 Euryhaline fishes, ultrastructural features of cells during transfer of from fresh water to sea water, 207-217 Excurrent duct epithelium of testes, MGPB in, 152-155 F

Fertilization, ZP, functions after, 86-89 during, 86 summary, 89 Firefly luciferase (FL), biologically localized cellular biogenesis and organization endogenous enzymes, use of as reporters for local concentrations of reactants, 278-280 involvement of physical processes and local catalysis in cellular metabolism, 275-278 protein traffic and assembly, 270-272 structural basis for organization of metabolism, 272-275 experimental approaches and perspectives light measurement, 309-310 perspectives, 313-3 18 luciferase as probe for protein metabolism, 313-314 monitoring local events in cryptic compartments, 317-318 prospects for local [ATP] measurements, 3 15-3 17 targeting luciferase to cellular compartments, 3 14-3 15 practical considerations, 310-313 use of FL as probe for metabolites concentrations, 3 10-3 I2 genetic and chemical engineering of luciferase, 312-313 leading into cellular compartments cloning, expression and localization of native luciferase, 2%-297 retargeting to other cellular organelles, 297-299

light emissions by in biological systems, 300-308 in whole cells, 300-302 monitoring local [ATP] at mitochondria1 surface, 302-308 dependence on rate of ATP formation, 305-306 local versus bulk [ATP] measurements, 302-304 modeling, 306-308 overview enzyme characteristic, 282-287 initial and steady-state kinetics of light production, 285-287 molecular mechanism of light emission, 282-283 structure, 284-285 history and applications, 280-281 probe for ATP in model systems, 287-2% continuous monitoring using immobilized luciferase, 288-291 mathematical modeling with soluble and localized luciferase, 291-296 kinetic analysis of FL reaction, 291-292 kinetic behavior of immobilized enzymes, effect of diffusion, 292-294 quantitation of light output from (Co) immobilized luciferase, 295-296 Fishes, fresh water-adapted, a-and P-cells, 204-207 FL, see Firefly luciferase Free steroid hormone hypothesis, 161 Fruit softening, 257-259 Fundulus heteroclitus, 191,204

Gallus domesticus, 142 Gap junctions, desmosomes, 151 Germination of plant cell walls, 255-256 Gill cells of teleost fishes, see also Teleost fishes gill morphology, 192-193 Gloosphonia complanata, 127

329

INDEX Glycine m a , 249 Glycoproteins, arrangement of in ZP filaments, 105 Glycoproteins, structural, 245 Glycoproteins, ZP, 92-95 ZP1,92 ZP2,92-94 ZP3,94-95 source, 95-96 Gobio gobio, 207 Golgi apparatus and vesiculotubular system, 200-202 Gramineae, 252-254 (1 + 3),(1 + 4)-P-D-GlUCan, 252-254 other polysaccharides, 254 Growth regulation of plant cell walls, 246-254 dicotyledons, 249-252 other polysaccharides, 251-252 xyloglucan, 249-25 1 gramineae, 252-254 (1 + 3),(1 4)-P-~-Glucan,252-254 other polysaccharides, 254 preliminary study approaches, 248-249

-

Immunoglobulin transport in B cell development, see B cell lymphocyte differentiation pathway and immunoglobulintransport Immunological approaches to plant cell walls, see Plant cell walls, immunological approaches In vitro Sertoli cells, 118-1 19 Initial kinetics of light production, 285-287 Insecta, presence of MGPB, 128-132 Intermediate filaments (IFs) in nucleated erythrocytes, 75 Inter-Sertoli septate junctions, MGPB formed by, 133 Intracellular retention of secretory immunoglobulin, 13-18

J J chain, role of, and assembly of polymeric immunoglobulin, 30

K H

H chain and Bip in pre-B cells, intracellular retention, 14-16 Heliothis virescens, 128 Helix aspersa, 134 Hirudo medicinalis, 127 Hormones and chloride cells, 225-227 Hyalophora cecropia, 15 1 Hydra littoralis, 176 Hydra oligactis, 176 Hydra viridis, 120-122, 173 Hydrophobicity, role of, and associated proteins, 22-27 Hypertrophy of a-cells, 212-214 I

IgM/IgD-expressing B lymphocytes, mature, 11-12 Immunoglobulin secretion in plasma cells, 29-30

KDEL sequences and salvage pathway, 14-16 Kinetic analysis of FL reaction, 291-292 Kinetic behavior of immobilized FL enzymes, effect of diffusion, 292-294 Kinetics of light production, initial and steady-state, 285-287

L L chains, role of and appropriate conformation for secretion, 30-31 pre-B-specific surrogate immunoglobulin, 7-10 Lacerta muralis, 141 Lacerta sicula, 141 Leaf abscission of plant cell walls, 256-257 Lebistes reticulus, 206 Lectins important to study of plant cell walls, 237,238

330

INDEX

Levantina hierosolyma, 116, 132, 133, 134, 159 Life cycle of plant cell walls, 233 Light emission, molecular mechanism, 282-283 Light emissions by in biological systems, 300-308 in whole cells, 300-302 monitoring local [ATPI at mitochondria1 surface, 302-308 dependence on rate of ATP formation, 305-306 local versus bulk [ATPI measurements, 302-304 modeling, 306-308 Light measurement of FL, experimental approaches and perspectives, 309-310 Light production, initial and steady-state kinetics, 285-287 , Limulus polyphemas, 249 LIS-solubilized ZP, 99-100 Lithobius forjicatus, 116, 128 Locusta migratoria, 128, 129, 131 Lonchuria striata, 142 Luciferase genetic and chemical engineering, 3 12-3 13 immobilized, continuous monitoring using, 288-291 localized, mathematical modeling, 291-2% kinetic analysis of FL reaction, 291-292 kinetic behavior of immobilized enzymes, effect of diffusion, 292-294 quantitation of light output from (Co) immobilized luciferase, 295-2% Luciferase as probe for protein metabolism, 313-31.4 monitoring local events in cryptic compartments, 317-318 prospects for local [ATP] measurements, 315-317 targeting luciferase to cellular compartments, 3 14-315 Luciola cruciata, 2% Luciola mingrelica, 296 Luidia clathrata, 115

Lumbricus terrestris, 125 Lymnaera stagnalis, 116, 133, 134-135, 154, 159, 171

M Macrobrachium rosenbergii, 127 Macropus eugenii, 76 Male germ cell protective banier (MGPB) background blood-organ barriers, different, 112-113 MGPB, methods helping to arrive at concept, 115-1 18 Sertoli cell for somatic cells in testis in different phyla, 114-1 15 Sertoli cell in vitro, 118-1 19 terms, 113-1 14 conclusions, 173- 177 function autoimmune aggression in mammals and the MGPB, 163-169 efficacy of barrier in physiological, pathological and experimental conditions, 160-163 MGPB, medical aspects, 171-173 MGPB, role of in maintaining spermatozoan immortality, 169- 171 secretory activity of sertoli cell, 159-160 along phylogenesis annelida, 125-127 chordata, 136-143 cnidaria, 120-122 crustacea, 127-128 echinodermata, 135-136 insecta, 128-132 mollusca, 133-135 myriafoda, 128 nematoda, 123 platyhelminthes, 122- 123 porifera, 19-120 structure desmosomes and gap and septate junctions between barrier-forming cells, 150-152 desmosomes, 150- 151

33 1

INDEX gap junctions, 151 septate junctions, 151-152 MGPB in excurrent duct epithelium of testes, 152-155 Sertoli cell and occluding junctions, 143-147 Sertoli cell junctions during growth, annual reproductive cycle, and spermatogenic process, 148-150 sites of different MGPB in testes, 157-159 testicular vascular system, 155-157 Mamestra brassicae, 116, 128, 130 Mammalia, presence of MGPB, 142-143 Mammalian primitive erythrocytes, 76-79 Mammals, autoimmune aggression and MGPB, 163-169 Marginal band (MB) biogenesis and function during erythrocyte morphogenesis, 59-67 elliptogenesis, 64-65 maturation, 65-67 MB biogenesis and mechanism of cell flattening, 59-64 Marginal band reassembly, experimentally induced, 53-56 Mastela vison, 145 Mature erythrocytes, marginal band, 43-59 experimentally induced MB reassembly, 53-56 MB function in, 56-58 molecular components, 49-53 other MB-associated proteins, 50-53 tubulin, 49-50 mechanical properties, 47-49 structure, 44-47 tribute to Friedrich Meves, 58-59 Mature IgMIIgD-expressing B lymphocytes, 11-12 Mauremys caspica rivulata, 125 MB, see Marginal band Membrane immunoglobulin ER degradation versus cell surface transport, 20-22 recycling, endocytosis and down-regulation, 27-28 retention in plasma cells, 29 transport, 22-27 transport during B cell ontogeny, 20-27

Membrane skeleton (MS) of nucleated erythrocytes, 67-75 biogenesis, 70-71 mechanical properties, 71-75 molecular composition, 69-70 structure, 68-69 Memory B lymphocytes, 12 Mesocricetus auratus, 148 Metabolites concentrations, use of FL as probe, 310-312 Metabolism of FL, structural basis for organization, 272-275 Meves, Friedrich, tribute, 58-59 MGPB, see Male germ cell protective barrier Mitochondria-rich gill cells of teleost fishes, see Teleost fishes Model system for coupling catalytic steps in a heterogenous system, 306-308 Molecular components, of mature erythrocytes, 49-53 proteins, other MB-associated, 50-53 tubulin, 49-50 Molecular mechanism of light emission, 282-283 Mollusca, presence of MGPB, 133-135 Monodelphis domestica, 39,76,77 Mouse egg extracellular coat, structure, see Zona pellucida MS, see Membrane skeleton Mustelus canus, 42 Myriafoda, presence of MGPB, 128 Myxine glutanosa, 136 N

Nematoda, presence of MGPB, 123 Noetia ponderosa, 55 Notophthalmus viridescens, 49 Nucleated erythrocytes, cytoskeletal system conclusions, 79-80 intermediate filaments, 75 mammalian primitive erythrocytes, 76-79 marginal band biogenesis and function during erythrocyte morphogenesis, 59-67

332

INDEX

elliptogenesis, 64-65 maturation, 65-67 MB biogenesis and mechanism of cell flattening, 59-64 mature cells, marginal band of, 43-59 experimentally induced MB reassembly, 53-56 MB function in, 56-58 molecular components, 49-53 other MB-associated proteins, 50-53 tubulin, 49-50 mechanical properties, 47-49 structure, 44-47 tribute to Friedrich Meves, 58-59 membrane skeleton, 67-75 mechanical properties, 71-75 molecular composition, 69-70 MS biogenesis, 70-71 structure, 68-69 phylogenic and physiologic portrait, 39-43 study, 37-38

0 Oligocottus maculosis, 217 Onchorhynchus keta, 170 Oocyte, growing, ZP of, 100-104 Oreochromis mossambicus, 210,217 Oreochromis niloticus, 116, 139 Oryzias latipes, 137, 139 Oryzias niloticus, 137

P Photinus pyralis, 28 1, 296 Phragmatopoma lapidosa, 127 Phylogenesis, MGPB annelida, 125- 127 chordata, 136-143 amphibia, 140- 141 aves, 142 cephalochordata, 136 cyclostomata, 136-137 mammalia, 142-143 reptilia, 141-142 teleostei, 137-140 cnidaria, 120-122 crustacea, 127-128

echinodermata, 135- 136 insecta, 128-132 mollusca, 133-135 myriafoda, 128 nematoda, 123 platyhelminthes, 122- 123 porifera, 19-120 Pylogenic portrait of nucleated erythrocytes, 39-43 Physical processes and local catalysis in cellular metabolism, involvement, 275-278 Physiologic portrait of nucleated erythrocytes, 39-43 Placobdella costata, 125, 127 Plant cell walls, immunological approaches antibodies as probes for study, 234-240 applications, 240 purification and specificity, 237-240 raised against cell wall components, 234-237 conclusions, 261-263 diversity of cell wall functions, 233-234 future prospects, 261-263 growth regulation, 246-254 dicotyledons, 249-252 other polysaccharides, 251-252 xyloglucan, 249-251 gramineae, 252-254 (1 + 3),(1 + 4)-P-D-GlUCan, 252-254 other polysaccharides, 254 preliminary study approaches, 248-249 life cycle, 233 location and metabolism of cell wall polymers, 240-246 enzymes, 246 polysaccharides, 241-245 (1 + 3),(1 + 4)-P-D-GlUCan, 243 other polysaccharides, 244-245 polygalacturonic acids, 243-244 xyloglucan, 241-243 structural glycoproteins, 245 other aspects, 259-261 selective breakdown, 254-259 fruit softening, 257-259 germination, 255-256 leaf abscission, 256-257 structural characterization of plant cell wall, 234

333

INDEX Plasma cells and end stage of B lymphoid ontogeny, 13 Plasma cells, membrane IgM retention, 29 Plasma cells, immunoglobulin secretion, 29-30 J chain, role of, and assembly of polymeric immunoglobulin, 30 L chains, role of, and appropriate conformation for secretion, 30-31 posttranslational modifications and secretion, 30 Platyhelminthes, presence of MGPB, 122- 123 Plafysamia Cynthia, 151 Poecilia latipinna, 154 Poecilia reticulata, 137, 140 Polygalacturonic acids, 243-244 Polymers, cell wall, location and metabolism, 240-246 enzymes, 246 polysaccharides, 24 1-245 structural glycoproteins, 245 Polysaccharides, 241-245,251-252,254 (1 + 3),(1 --* 4)-/3-~-Glucan,243 other polysaccharides, 244-245 polygalacturonic acids, 243-244 xyloglucan, 241-243 Porifera, presence of MGPB, 119-120 Posttranslational modifications and secretion, 30 Pre-B cells, fate of membrane immunoglobulin, 20-22 Pre-B to B-cell transition, 10-1 1 Pre-B lymphocytes, 6-7 Pre-B-specilic surrogate immunoglobulin L chains, 7-10 Pro-B lymphocytes, 4-6 Protease- and dithiothreitol-solubilized ZP, 100

Protein traffic and assembly of FL, 270-272 Proteins, other MB-associated, 50-53 Pyrophorus plagiophtalarnus, 281,282, 309 Q

Quantitation of light output from (Co) immobilized luciferase, 295-296

R Rana catesbiana, 65 Rana espculenta, 140, 155 Rana ridibunda, 141, 155 Rana temporaria, 141 Recycling, membrane immunoglobulin, 27-28 Reptilia, presence of MGPB, 141-142 Retentioddegradation and transport of immunoglobulin, choices between, 31-33 Rhamnogalacturonan 1 antibodies, 244 Ribulose biphosphate carbox ylase (RUBISCO), 299

S Saccharornyces cerevesiae, Bip gene in, 15 Saccharomyces cerevisae, 297 Salamandra salamandra, 140 Salmo gairdneri, 154, 166,227 Salmo salar, 223 Salrno trutta, 226 Salmonids, smoltilication and mitochondria-rich cells, 221-225 Salvelinus fontinalis, 154 Schistocerca gregaria, 129, 130 Scophthalmus maximus, 216,219 Secretion in plasma cells, immunoglobulin, 29-3 1 Secretory immunoglobulin, intracellular retention, 13-18 cysteine tailpiece-dependant retention, 16-18 KDEL sequences and salvage pathway, 14-16 Septate junctions, desmosomes, 151-152 Sertoli cell in vitro, 118-1 19 junctions during growth, annual reproductive cycle, and spermatogenic process, 148-150 occluding junctions, 143-147 secretory activity, 159-160 for somatic cells in testis in different phyla, 114-115 Smoltification in salmonids, and mitochondria-rich gill cells, 221-225

334

INDEX

Somatic cells, sertoli cells in testis in different phyla, 114-1 15 Sparus aurata, 174 Spermatozoan immortality, role of MGPB in maintaining, 169-171 Staphylococcus species, 237-238 Steady-state kinetics of light production, 285-287

Stenohaline seawater fishes, chloride cells and accessory cells, 216-217 Structural characterization of plant cell wall, 234

Testes, sites of different MGPB in, 157- 159

Testicular vascular system, 155-157 Tetragonolobus purpureas, 248 Tilapia aurea, 214 Tilapia specimens, 166 Transitional and immature B cell stages, 10-11 Triatoma infestans, 128, 130, 152 Triturus carnifex, 140 Triturus cristatus, 66 Tubulin, 49-50

T

U

Taeniopygia guttata, 142 Teleost fishes, mitochondria-rich gill cells, ultrastructure chloride cells, general features cell surface, 193-199 apical cavity, 193-1% tubular system, 196-199 endoplasmic reticulum, mitochondria, and cytoskeleton, 202-204 Golgi apparatus and vesiculotubular system, 200-202 conclusions, 227-228 environment, modifications a-and p-chloride cells in fresh water-adapted fishes, 204-207 chloride cells and accessory cell during transfer of sea-water adapted fishes to fresh water,

Ulex europaeus, 248 Ultrastructural features of cells during transfer of euryhaline fishes from fresh water to sea water, 207-217 accessory cells, appearance, 214-216 degeneration of p-cells, 209-212 hypertrophy pf a cells, 212-214 Ultrastructure of zona pellucida, 96-105 arrangement of glycoproteins in ZP filaments, 105 electron microscopy, 99-104 LIS-solubilized ZP,99-100 oocyte, growing, ZP of, 100-104 protease- and dithiothreitol-solubilized ZP, 100 general considerations, 96-98 summary, 105

217-221

chloride cells and accessory cells in stenohaline seawater fishes, 2 16-2 17

ultrastructural features of cells during transfer of euryhaline fishes from fresh water to sea water, 207-217 accessory cells, appearance, 2 14-2 16

degeneration of p-cells, 209-2 12 hypertrophy of a cells, 212-214 gill morphology, 192-193 hormones and chloride cells, 225-227 and smoltification in salmonids, 221-225 Teleostei, presence of MGPB, 137-140

v Vesiculotubular system and golgi apparatus, 200-202 Virgin splenic B cells, 16-17 Vitamin-A deficient diet, effect on spermatogenesis, 162-163 Viviparus uiuiparus, 134 W

Whole cells, light emissions in biological systems, 300-302

INDEX X Xenopus laevis, 61,62,67, 141, 296 Xiphophorus helleri, 166 Xylanase-gold conjugates, use, 245 Xyloglucan, 241-243,249-251 effect of antibodies in azuki and oat segments, 250,251 Xyloglucan oligosaccharide, 250

z Zona pellucida (ZP), structure characteristics, 89-96 general considerations, 89-91 source of ZP glycoproteins, 95-96 summary, % ZP glycoproteins, 92-95 ZP1.92

335

Zp2.92-94 ZP3,94-95 conclusions, 105-108 description, 85-86 functions, 86-89 after fertilization, 86-89 during fertilization, 86 summary, 89 ultrastructure, 96-105 arrangement of glycoproteins in ZP filaments, 105 electron microscopy, 99-104 LIS-solubilized ZP, 99-100 oocyte, growing, ZP of, 100-104 protease- and dithiothreitol-solubilized ZP, 100 general considerations, 96-98 summary, 105 Zonula adherens, 150-151 ZP, see Zona pellicuda

This Page Intentionally Left Blank