International Review of Cytology

VOLUME 142

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

1949-1988 1949-1 984 19671984-

ADVISORY EDITORS Airnee Bakken Eve Ida Barak Howard A. Bern Robert A. Bloodgood Dean Bok Stanley Cohen Rene Couteaux Marie A. DiBerardino Donald K. Dougall Charles J. Flickinger Nicholas Gillham Elizabeth D. Hay Mark Hogarth Keith E. Mostov

Audrey Muggleton-Harris Andreas Oksche Muriel J. Ord Vladimir R. Pantic' M. V. Parthasarathy Lionel I.Rebhun Jean-Paul Revel L. Evans Roth Jozef St. Schell Hiroh Shibaoka Wilfred Stein Ralph M. Steinrnan 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 and Department of Physiology UCLA School of Medicine Los Angeles, California

VOLUME 142

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 1992 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. 1250 Sixth Avenue, San Diego, California 92101-431 1 United Kingdom Edition published by Academic Press Limited 24-28 Oval Road, London NW1 7DX Library of Congress Catalog Number: 52-5203 international Standard Book Number: 0-12-364545-X

PRINTED IN THE UNITED STATES OF AMERICA

92 93 94 95 96 97 98

EB

9 8 7 6 5 4 3 2 1

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

ix

Biological Problems of Regenerative Cementogenesis: Synthesis and Attachment of Collagenous Matrices on Growing and Established Root Surfaces Hubert E. Schroeder I. Introduction ......................................... II. Origin, Types, and Function of Root Cementum on Hum 111. Spontaneous Cementogenesis and Matrix Formation on Growing ...... Root Surfaces ....................................... IV. Matrix Formation on Established Root Surfaces in Vitro ...... V. Regenerative Cementogenesis on Established Root Surfaces in Vivo . . . . . . . . . . . . . . VI. Concluding Remarks and Perspectives . . . . . . . . . . .......................... References ............................. .....................

1 2 5 28 43 51 52

lmmunocytochemical Localization of Proteins in Striated Muscle Marvin H. Stromer I. II. 111. IV. V. VI.

Introduction ................................ Localization of Proteins in Skeletal Muscle Cells . . . . . . . . . . . . . . . . . . . Localization of Proteins in Cardiac Muscle Cells ................................. Sarcoplasmic Reticulum, Transverse Tubules, and the Sarcolemma . . . . . . . . . . . . . . . Other Proteins .................................... ..................... Conclusions and Outlook ............................ References . . . . . . . . . . . . . . . . .................................. V

61 62 102 119 127 128 129

vi

CONTENTS

Recent Developments in Vertebrate Cell Culture Technology Satish J. Parulekar. Thomas Hassell. and Satish C . Tripathi I. II. 111. IV. V. VI .

....................... Traditional Cultures . . . . . ........................... Three-Dimensional Cultures . . . . . . . . . . ............................ Commercial Scale Bioreactors . . . . . . . . . . . . . . . . .

................

Design and Optimization Considerations ..................... Concluding Remarks . . . . . . . . . . . . ...........................

.....................

145 147 153 162 192 201 204

Transdifferentiation in Medusae Volker Schmid I II 111 IV

Introduction ....................... ............................ .............. The Concept of Transdifferentiation . . . . . . . . . . . . . . Transdifferentiation in Hydromedusae ...................... Concluding Remarks . . . . . . . ............................ References ............................. ......................

213 214 218 256 258

Symplast as a Functional Unit in Plant Growth Kiyoshi Katou and Hisashi Okamoto ............ I. Introduction ..................................... II. Electrophysiological Structure of the Plant Germ Axis . . . . . . . . . . . 111 . Role of Spatially Separated Proton Pumps in Stem Elongation .................... ..................... IV. Lockhard Equations and Action of Auxin . . . . V . Integration of the Activity of the Symplast in P .................................. VI . ........................................

263 265 273 277 285 299 300

lntracellular Ca2+Messenger System in Plants Shoshi Muto I. Introduction ................................................................... I1. Receptors ..................................................................... Ill. G Proteins ....................................................................

305 306 308

vii

CONTENTS

IV. Regulation of lntracellular Ca2+ Concentration .................................. V . Phosphatidylinositol Turnover . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

................ .......................................................... VIII . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References

....................

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

311 321 328 332 338 339 359

This Page Intentionally Left Blank

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

Thomas Hassell' (145), Celltech Limited, Slough SL1 4EN, Berkshire, England Kiyoshi Katou (263), Biological Laboratory, College of General Education, Nagoya University, Nagoya 464-01, Japan Shoshi Muto (305),Institute of Applied Microbiology, University of Tokyo, Tokyo 113, Japan Hisashi Okamoto (263), Biology Department, Graduate School of Integrated Science, Yokohama City University, Kanazawa 236, Japan Satish J. Parulekar (145), Department of Chemical Engineering, lllinois Institute of Technology, Chicago, Illinois 60616 Volker Schmid (213), Institute of Zoology, Pheinsprung 9, CH-4051 Basel, Switzerland, and Friday Harbor Laboratories, Friday Harbor, Washington 98250 Hubert E. Schroeder ( l ) ,Department of Oral Structural Biology, Dental lnstitute, University of Zurich, CH-8028 Zurich, Switzerland Marvin H. Stromer (61), Department of Animal Science, Muscle Biology Group, Iowa State University, Ames, Iowa 5001 1 Satish C. Tripathi (1 45), Department of Life Sciences, IlT Research Institute, Chicago, lllinois 60616 'Present address: iAF BioVac Inc., Ville de Laval, Quebec, Canada H7N 422.

ix

This Page Intentionally Left Blank

Biological Problems of Regenerative Cementogenesis: Synthesis and Attachment of Collagenous Matrices on Growing and Established Root Surfaces Hubert E. Schroeder Department of Oral Structural Biology, Dental Institute, University of Zurich, 8028 Zurich. Switzerland

1. Introduction

During the past decade, regeneration of periodontal tissues has received increasing and worldwide attention, from both oral biologists and dental clinicians. “Periodontal regeneration” is defined as the restoration of the various components of the periodontium, i.e., alveolar bone, periodontal ligament, root cementum, and gingiva lost due to disease, “in their appropriate locations, amounts, and relationships to each other” (Aukhil, 1991). In contrast to more simple goals such as the reattachment of the periodontal ligament and supraalveolar connective tissue to the torn cell/fiber tissue at the dental root surface, following their short-term separation, or as the spontaneous repair resulting from unguided wound healing, “periodontal regeneration” requires an enormously well-conducted action of various cell populations to appear and function in space and time in order to reconstitude both structural normality and functional integrity. As the development, structure, and function of the human periodontium, being dissimilar in details to that of laboratory animals such as rodents, dogs, and non-human primates, are immensely complex biologically and not entirely understood (Schroeder, 1986), clinical and laboratory experiments in humans and other animals designed to study the regenerative potential of the various tissue components under conditions of different defects and treatment modalities are exceedingly difficult to analyze. In fact such experiments necessarily address the periodontium as a whole rather than being able to examine separately the response of component tissues. In addition, the situation is rendered even more complex by the InlPrnati~lnu/Reuirn, of

Cyro/oas. V o / . 142

1

Copyright Q 1992 by Academic Press. Inc. All rights of reproduction in any form reserved.

2

HUBERT E. SCHROEDER

fact that the creation of artificial defects or the pretreatment of spontaneous lesions caused by bacterial infection and inflammation introduce artificial tissue alterations that interfere with repair and regeneration. For these reasons, clinical and laboratory experiments on periodontal regeneration, reviewed by Egelberg (1987), Nyman et al. (1989), and Minabe (1991), have yielded widely different results that remain insufficient from a clinical, and invalid from a biological, point of view, results that to a varying extent appear as biologically undecodable messages from a blackbox world. Not surprisingly, periodontal regeneration is loaded with problems, clinical, such as reinfection and mechanical disturbance of the wound healing processes, as well as biological. The latter include the physical and chemical denaturation of treated root surfaces; epithelial migration over the exposed root surface sites that should be the substrate for regenerative tissue formation; the different turnover and growth potentials of the four periodontal tissue components; and innumerable problems regarding cells, mediators, growth factors, etc. (Abdallah et al., 1988; Terranova er al., 1989a; Messadi and Bertolami, 1991; Aukhil, 1991). One reason for the unsatisfying situation was and still is the fact that human cementogenesis remained undiscovered for too long. Indeed, at the beginning of the research focusing on periodontal regeneration, our knowledge of dental root cementum was incredibly meager, resting on antique rather than medieval information. Root cementum represents, however, the most cardinal periodontal tissue component that is primary and indispensible for regenerating the tooth-bone connection, i.e., for reconstituting tooth anchorage. This review provides some of the missing data needed to discuss spontaneous and regenerative cementogenesis and unveils some of the biological problems involved.

II. Origin, Types, and Function of Root Cementum on Human Teeth

A new and increasingly accepted classification of root cementum on human teeth was proposed by Jones (1981) and with modifications adopted by Schroeder (1986). It differentiates among four varieties according to the absence or presence of cells and to the source of collagen fibers, i.e., the major matrix component contained within. Consequently, this classification distinguishes between acellular afibrillar cementum (AAC); acellular extrinsic fiber cementum (AEFC); cellular intrinsic fiber cementum (CIFC) that may also occur as an acellular variety (AIFC; Bosshardt and Schroeder, 1990);and cellular, mixed stratified cementum (CMSC). These varieties are summarized in Table I.

3

REGENERATIVE CEMENTOGENESIS TABLE I Types of Human Root Cementum

Terms

Abbreviation

Acellular, afibrillar cementum

AAC

Acellular, extrinsic fiber cementum

AEFC

Cellular, intrinsic fiber cementum

CIFC

Acellular, intrinsic fiber cementum

AIFC

Cellular, mixed, stratified cementum (AEFC + CIFCI AIFC)

CMSC

Organic components Homogeneous matrix, no cells, no collagen fibrils Collagen fibrils as Sharpey’s fibers, no cells Intrinsic collagen fibrils and fibers, cementocytes

Intrinsic collagen fibrils and fibers, no cells Intrinsic collagen fibrils and fibers, collagen fibrils as Sharpey’s fibers, cementocytes

Location At dentinoenamel junction, on enamel Cervical to middle root

Function Unknown

Tooth anchorage

Apical and interradicular root surfaces, resorption lacunae, fractures Apical and interradicular root surfaces

Adaptation, repair

Apical and interradicular root surfaces

Adaptation, root anchorage

Adaptation

Apart from a homogeneous and mineralized ground substance of unknown composition, AAC contains neither cells nor collagen fibrils. In humans, it is found as coronal cementum covering patchwise the cervical enamel, and as an occasional part of cervical AEFC (Schroeder 1986, 1988). Its origin and function are unknown, but it may represent serumderived organic material coprecipitated with mineral (Beertsen and Van Den Bos, 1991). The AEFC lacks cells and is composed entirely of densely packed, well-oriented bundles of collagen fibrils, i.e., the so-called fibers of Sharpey. These fibers continue into the periodontal ligament and connect the root to the alveolar bone. Thus, all AEFC fibers are extrinsic. About 30,000 fibers insert into 1 mm2 of AEFC surface, each fiber being about 4 pm in diameter (Schroeder, 1986). In humans, the rather thin (20 to 250 pm), densely mineralized AEFC shows parallel incremental lines and is found primarily on the cervical and middle root regions, but it may extend further apically. It is formed by fibroblasts of the dental follicle

4

HUBERT E. SCHROEDER

proper, i.e., ectomesenchymal derivative, and later of the periodontal ligament, and serves exclusively for tooth anchorage (Schroeder 1986, 1988). The CIFC contains cells, the cementocytes, but its collagen fibers, being again the major matrix component, are all intrinsic and run a circular or spiral course around the root, i.e., more or less parallel to the root surface. Thus, CIFC lacks Sharpey's fibers. In humans, the less well mineralized CIFC is found mainly in situations of repair, filling resorption lacunae or connecting root fracture fragments. It is also part of CMSC, forming initially on apical root portions and patchwise between layers of AEFC. The CIFC is formed by cementoblasts of the dental follicle proper and later of the periodontal ligament; its function is associated with repair and adaptation. On particular portions of the root, AIFC may form without leaving cementocytes behind. CMSC is a mixture of pure AEFC and CIFC/AIFC; the latter part may contain cementocytes with uneven distribution and density. The CMSC is usually a stratified tissue, with consecutive or alternating layers of AEFC and CIFC/AIFC being unpredictably superimposed on one another. In humans, the inhomogeneously mineralized and, in part, porous CMSC is variably thick, ranging from 100 to 600 p m or more, and occurs primarily in the apical third of the roots and in the furcations. It serves the functions of adaptation, i.e., adynamic reshapeningofthe root surfaceas the tooth shifts and drifts in its socket, and, if superficially covered by AEFC, of root anchorage (Table I; Schroeder, 1986, 1988). Measurements of sequential fluorochrome labeling lines in all deciduous teeth and the permanent molars of one -13-month-old M . fascicularis monkey (Bosshardt er al., 1989) and in the alveolar bone surrounding the first molars (Schroeder er al., 1992) provided data for the formation rate of two cementum varieties, in comparison to that of dentine, and bone TABLE II Rates of Formation of Cementum, Dentine, and Bone

Tissue

Abbreviation

Acellular, extrinsic fiber cementum Cellular, intrinsic fiber cementum Initial layer Appositional layers Crown dentine (first molars) Root dentine (deciduous teeth) Root dentine elongation (deciduous teeth) Alveolar bone crest (first molars) Alveolar bone septum (first molars)

AEFC CIFC

CD RD RDE ABC ABS

Formation-rate (pmlday)" x + s

< 0.10 + 0.02 0.4-3. I 0.1-0.5 3.1 k0.2 2.7-4.6 12.0-36.0 5.0-14.0 I3 .O-22.0

" Measured in one sequentially fluorochrome-labeled M . fuscicularis monkey; from Bosshardt et a / . (1989) and Schroeder et a / . (1992).

REGENERATIVE CEMENTOGENESIS

5

and to root elongation (Table 11). These data demonstrated that AEFC is an extremely slowly forming tissue, initially as well as later in life (Sequeira et al., 1992). In humans, its daily rate of formation, i.e., increase in thickness, is smaller that 0.1 p m , possibly as low as 0.005 to 0.01 p m (Dastmalchi et al., 1990; Sequeira et al., 1992). In contrast, initial CIFC is formed at a fast rate, ranging from 3.1 to 0.4 pm/day. Subsequently, appositional cementum layers, possibly of the AIFC variety, may still form at a faster rate than AEFC, i.e., 0. I to 0.5 pm/day. Comparatively, CIFC may form as rapidly as crown and root dentine and not much slower than alveolar bone (Table 11). 111. Spontaneous Cementogenesis and Matrix Formation on Growing Root Surfaces

In contrast to amelogensis and dentinogenesis, i.e., to rather well-defined developmental systems characterized by particular classes of cells and their morphologically and biochemically defined matrix products, cementogenesis on human teeth was essentially unknown until 1985, although some fragmentary information was available for other mammalian species such as rodents. This information had been derived from studies on mouse incisors and molars (Selvig 1963, 1964, 1967) and on rat molars (Paynter and Pudy, 1958; Diab and Stallard, 1965; Lester, 1969; Formicola et al., 1971 ;Owens, 1980). Although such molars are also covered by both acellular and cellular cementum, on their roots, and albeit most recent investigations of Cho and Garant (1988, 1989) and Yamamoto and Wakita (1990, 1991, 1992) demonstrated some similarity in matrix production and attachment to dentine, there are a number of reasons for the argument that root development and cementogenesis in rodent molars might be unlike that in humans (see below). Therefore, this review focuses primarily on cementogenesis in human teeth and recent observations in rodents will be used only comparatively. A. Acellular Extrinsic Fiber Cementum

In human teeth, AEFC covers the cervical root surfaces and extends from the cemento-enamel junction apically. In single-rooted teeth (i.e., incisors, canines, and most premolars), AEFC coats 60 to 90% of the total root length that varies between 13 (central incisors) and 15.5 mm (canines; Schumacher and Schmidt, 1983; Schroeder, 1988). AEFC is first formed while the roots develop. AEFC formation begins at and along the growing root edge and the AEFC slowly increases in thickness in the coronal direction. As shown in human premolars with incomplete roots developed

6

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

7

50 to 60% of their final length (Bosshardt and Schroeder, 1991a), initiation of AEFC formation, early matrix production, and its attachment to root dentine all take place within a zone of about 300 pm, extending coronally from the advancing root edge (Fig. 1). This zone probably first develops at the outer surface of the initially formed root portion and later shifts in the apical direction as the root grows in length. As a consequence, thin layers of established AEFC are encountered in coronal root regions, while initial AEFC genesis continues apically, i.e., near the advancing root edge. From data defining the time period necessary for root development, it is known that human premolars attain their complete root length within about 6 years following the completion of their crown (Schroeder, 1987). Based on a rough calculation, the time period during which AEFC is formed initially on the growing root spans 43 to 65 months, i.e., 60 to 90% of 6 years. Because AEFC coats 8 to 10 mm (i.e., 60 to 75%) of the final root length in human premolars, AEFC development at and along the growing root may proceed at a rate of 4.6 to 6.9 pm/day (Schroeder, 1987, 1988). These rates, calculated from clinical and morphometric measurements rather than based on direct evidence which is unavailable, are very much lower than the rates of root elongation measured in all deciduous and permanent molar teeth of the M.fuscicularis monkey, ranging between 12 and 36 pmlday (Table 11). As a preliminary estimate, it can be inferred from these data that initiation of AEFC formation within the 300-pm-wide zone proceeds apically with an average speed of 5 to 6 pmlday. In all probability, AEFC is a product of a particular class of fibroblasts (Beertsen and Everts, 1990; Bosshardt and Schroeder, 1991a,b). The advancing root edge includes the leading edge of newly produced predentine and the inorganic edge of mineralized dentine. The latter follows the former within a short distance of up to 50 pm. From this edge, predentine continues both over the pulpal surface along the dentine-odontoblast interface and over the external surface of the newly formed root dentine. At the latter site, it can be followed coronally over a distance of about 250

FIG. 1 Schematic drawing illustrating topographically the initial stages of AEFC genesis on human premolars developed to 5040% of their final root length: 1, fibroblasts contact root/ predentine and become committed: 2, fibroblasts start to form and attach collagen fibrils: 3, inital fiber fringe with maximum fiber density is established: 4, cell fiber fringe meshwork is established and the mineralization front approaches the base of the fringe: 5, mineralization front progresses into initial fiber fringe. AEFC. acellular extrinsic fiber cementum: MD, mineralized dentine: ERM, epithelial rests of Malassez: FPF, fringe-producing fibroblasts: FF, collagenous fiber fringe; PD, predentine;MF, mineralizationfront; NMD, nonmineralized dentine or predentine;CF, committed fibroblasts;ARE, advancing root edge; HRS, Hertwig’s epithelial root sheath;FFB, fiber fringe base. Modified from Bosshardtand Schroeder (1991 a).

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

9

to 300 pm from the advancing edge (Fig. 1). At this edge, the diaphragm, i.e., the most apical part of Hertwig’s epithelial root sheath, touches the predentine but, lateral or external to this edge, the root sheath deviates from the surface of newly formed dentine, continues coronally as a short strand, and eventually breaks up into the discontinuous epithelial rests of Malassez (Fig. 1). In humans, Hertwig’s root sheath, including its diaphragm, consists of the former inner and outer layers of the enamel epithelium, extends by continuous proliferation of the diaphragm (Diab and Stallard, 1965; Kenney and Ramfjord, 1969; Formicola et al., 1971), disintegrates coronally in accordance with its rate of proliferation, and is surrounded by a basal lamina. The latter actually contacts the leading root edge of predentine (Schroeder, 1986). In contrast to previous statements in most current textbooks, Hertwig’s root sheath does not cover much of the external surface of newly formed predentine, at least in human premolars. Rather, that surface at the advancing root edge is almost from its beginning accessible to connective tissue cells of the dental follicle proper. In the triangular region between the laterally deviating root sheath and the surface of newly formed predentine, connective tissue cells with the morphological appearance of fibroblasts can always be encountered. These cells are slender or bulky, are basophilic, and reveal an activated euchromatin-rich nucleus, displaying an -50-nm-thick nuclear fibrous lamina and a cytoplasm with numerous strands of rough endoplasmic reticulum cisternae and a prominent Golgi field. These cells are connected to one another by desmosome-like junctions and project numerous, slender cytoplasmic processes that contact and insert between collagen fibrils of the not yet mineralized outer predentine matrix (Fig.2a). These features are typical for the most apical 30 to 50 pm along the surface of the newly formed root. In about that distance to the advancing root edge, cells of similar appearance begin to produce the first AEFC matrix portions in the form of tiny but discrete bundles of collagen fibrils, slowly increasing in length and density in the apico-coronal direction. Thus, a short fiber fringe is produced, with most of the collagen fibrils arranged in parallel and oriented more or less perpendicular to the root surface. These fibrils

FIG. 2 (a) Electron micrograph depicting the most apical zone of not yet mineralized predentine matrix (PD) contacted and penetrated by cytoplasmic processes (CP) of fibroblast-like cells ( F )that begin to produce cementa1 collagen fibrils (see Fig. I , parts 1 and 2). (b) Electron micrograph of the initial fringe of collagen fibrils and fibril bundles (FF) that insert into the not yet mineralized predentine (PD);the mineralization front (MF) has not reached the future dentino-cementa1 junction (see Fig. 1, parts 3 and 4). Magnification: a,b, x6700. (a) From Bosshardt and Schroeder (1991a). (b) Courtesy Dr. D. D. Bosshardt.

10

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

11

terminate within and intermingle with the collagen fibrils of the dentinal matrix (Fig. 2b). Because of their orientation and parallelity, the fringe fibers are well demarcated from the randomly arranged fibrils of predentine. Within about 200 pm from the advancing root edge, this fringe attains maximum density (i.e., 130-150 fibers per I mm root surface length), which remains constant further coronally (Bosshardt and Schroeder, 1991b; Sequeira et al., 1992). In other words, the base of the fringe at its interface with predentine is fully established within a distance of 200 pm, or, based on the above calculation, within about 30 days of matrix production (Figs. 2b and 3a). Matrix production is not fully understood at present, but it is apparently carried out by the one to three cell layers that form a three-dimensional interwoven cell-fiber fringe meshwork. It is possible that the sequential, intracellular events in the synthesis and secretion of collagen are analogous to those described for human gingival fibroblasts (Yajima et al., 1980), for periodontal ligament fibroblasts in young Balb-C mice (Cho and Garant, 1981)and in 20-day-old rats (Marchi and Leblond, 1983, 1984), and for osteoblasts and odontoblasts in 20-day-old rats (Weinstock and Lebland, 1974;Weinstock, 1975; Leblond, 1989). In these cells, the precursors of Type-I collagen are synthesized in the rough endoplasmic reticulum and processed along the Golgi-secretory granule pathway, the resulting procollagens being released by exocytosis at the cell surface. Extracellularly , these procollagens are then transformed into fibrillar collagen. However, the particular class of fibroblasts forming the initial fiber fringe that represents the first AEFC matrix, in addition to synthesis and release of procollagen, is also engaged in fibril assembly, bundle formation, and fibril bundle orientation. Whereas the assembly and orientation of collagen fibrils is a primary task, bundle condensation and elongation is a subsequent step, increasing the bundle density and allowing AEFC matrix to increase in thickness. Selvig (1964), studying cementogenesis in albino mice, Schroeder (1986), examining early AEFC on human teeth, and Yamamoto and Wakita( 1992),investigating bundle formation during the genesis of acellular cementum in 20-dayold Wistar rats, observed that in tangential and cross sections through the developing and the established fiber fringe, as well as the fiber matrix covering the first, already mineralized layer of AEFC, the fibroblasts

FIG. 3 (a) Electron micrograph depicting the established cell fiber fringe meshwork with the established fringe (FF) of collagen fibril bundles attached to mineralized dentin (MD); the mineralization front (MF) has surpassed the dentino-cementa1 junction (see Fig. I , part 5 ) . (b) Electron micrograph of a tangential section through the cell-fiber fringe meshwork, with cytoplasmic extensions of the fibroblast-like cells (F)encircling compartments for individual collagen fibril bundles (FB). Magnification: a,b x4470. (a,b) Courtesy Dr. D. D. Bosshardt.

12

HUBERT E SCHROEDER

associated with AEFC genesis form circular cytoplasmic recesses and sheet- or plate-like extensions that partially of fully surround the assembly of cross-cut collagen fibrils (Fig. 3b). In fact, cytoplasmic processes of adjacent cells overlap, forming closed compartments (Yamamoto and Wakita, 1992). The formation of collagen fibril bundles has also been studied in both the corneal stroma and the tendon of chick embryos (Birk andTrelstad, 1984, 1985,1986; Fleischmajer er a / . , 1988; Birk et a/., 1990). In both cases, collagen fibrils are produced and bundled into oriented fibers. Bundling occurs because the extracellular space adjacent to fibroblasts is partitioned by the fibroblast surface and its extensions. Individual compartments, probably assuming the form of a cylinder, that house developing fibril bundles are first formed by surface recesses and are later separated from one another by cytoplasmic extensions. Within such compartments, collagen fibril segments of varying lengths can be added to the already existing group of fibrils (Birk et a/., 1990). By means of linear and lateral fusion of fibril segments, it is assumed that collagen fibrils grow in length and fibers in thickness. Birk e t a / . (1990) stated, “it is our hypothesis that synthesis, posttranslational processing, packaging, discharge, and assembly into fibril segments, fibrils, bundles, and tissue-specific macroaggregates are closely regulated by the cell at each stage,” including “the formation of distinct compartments in which the structural elements of the developing matrix would serve to physically position these elements within the developing matrix.” The generation of dense bundles of collagen fibrils that are packed nearly parallel and are oriented nearly perpendicular to the root surface and their elongation are the major events in AEFC matrix production, both initially and subsequently, as AEFC increases in thickness. With increasing density and diameter of the fibril bundles, cytoplasmic extensions may even retract, allowing lateral bundle coalescence that results in particularly thick collagen fibers (Yamamoto and Wakita, 1992). The macroaggregate of oriented fringe fibers, which is very particular for the AEFC matrix, consisting exclusively of Type I ([al(I)]* a2) collagen and noncollagenous components that amount to about 19% of the total matrix, certainly derives from a guiding activity of that particular class of fibroblasts (Christner er a / . , 1977; Smith er al., 1983). In the rat molar, early stages of AEFC matrix formation have been described differently. Hertwig’s epithelial root sheath covers the newly formed predentine externally until the latter starts to mineralize. Instead of deviating from the growing root edge and the newly formed root surface, as occurs in humans, the epithelial sheath has to break up or be penetrated in order to allow connective tissue cells to arrive at the newly formed root surface (Formicola et al., 1971; Yamamoto, 1986; Cho and Garant, 1988, 1989; Yamamoto and Wakita, 1990). As in humans, the latter retains an I-pm-thick, not yet mineralized, layer of predentine, the matrix of which

-

REGENERATIVE CEMENTOGENESIS

13

is loosely knit and reveals only a few collagen fibrils (Owens, 1980; Yamamot0 and Wakita, 1990). The first primitive and still loose collagen fibril bundles that penetrate into this superficial predentine layer appear rapidly. Thereafter, a ruthenium red-positive, phosphotungstic acid-positive, and silver-stain-negative material is deposited into and onto the predentine, giving rise to a layer 1-2 pm thick (Yamamoto, 1986; Yamamoto and Wakita, 1990). Under an electron microscope, this layer shows a reticular structure that includes fine granular and filamentous material. In addition, radiolabeled mannose is deposited on the predentine surface, but not on older portions of the root surface already covered with acellular cementum. As this material first appears within cementoblast-like cells, i.e., cytoplasmic granules, and later in association with collagen fibrils of the cementa1 matrix, both newly formed collagen fibrils and mannose-containing material are believed to be the product of the cementoblast-like cells (Cho and Garant, 1989). Among the known mannose-containing glycoproteins of the connective tissue are fibronectin, structural glycoprotein, and the carboxyl terminal propeptides of procollagen (Cho and Garant, 1989). The origin of the ruthenium red-positive layer is less clear. However, the cementoblast-like cells have been traced by Cho and Garant (1989). They reported that “disruption of the epithelial root sheath appears to be a consequence of directed cell migration by cells of the dental follicle proper which undergo differentiation into precementoblasts.” The latter eventually contact the newly formed outer predentine by means of cytoplasmic processes, become cementoblasts, and temporarily produce part of the acellular cementum matrix. Thereafter, they withdraw from the root surface and assume a fibroblast-like morphotype, referred to as postcementogenic fibroblasts (Cho and Garant, 1989). Later in the process, prior to calcification, further collagen fibril bundles are formed that penetrate only a short distance into the ruthenium red layer and become denser and more bulky; these bundles serve as the major component of the acellular cementum matrix in rat molars. Although there are some similarities of AEFC genesis in rats and humans, it is not known whether a ruthenium red-positive layer and mannosecontaining material also appear in human cementogenesis. Because in the latter all cells, i.e., contacting the predentine and associated with the fiber-fringe meshwork, are apparently engaged in the production of collagen fibrils and bundles thereof, it is likely that all of them are fibroblasts (young and mature), possibly belonging to a particular but still unspecified class. It is possible that the cementoblast-like cells reported on by Cho and Garant (1988, 1989) are also typical fibroblasts, as they were similar ultrastructurally and their profiles were only slightly larger than that of the fibroblasts. Moreover, in humans, none of the connective tissue cells associated with AEFC genesis is comparable in size and structure to

14

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

15

cementoblasts associated with matrix formation of either AIFC or CIFC produced more apically (See 111,B;Bosshardt and Schroeder, 1990, 1991a, 1992). As soon as the first fiber fringe has attained maximum density, the external mineralization front in human premolar roots approaches the zone of interdigitation of the fringe fibers and the dentinal matrix (Bosshardt and Schroeder, 1991a). This occurs at about 0.3 to 1 mm coronal to the advancing root edge (Figs. 1 and 3a). Subsequently, the base of the fiber fringe begins to mineralize (Bosshardt and Schroeder, 1991b). Thus, the fiber fringe becomes identical to the AEFC matrix, although the fringe continues to exist by fiber elongation, whereas the mineralization front involves more and more of the fringe matrix. In humans, early mineralization of the fiber fringe is associated with the appearance of spherical mineralization centers that initially occur as isolated patches, bound to particular bundles of collagen fibrils ahead of the mineralization front, and later fuse with the advancing front, eventually becoming incorporated in mineralized AEFC (Fig. 4a, Schroeder, 1986; Bosshardt and Schroeder, 1991b). In rat molars, early calcification at the dentino-cementa1 junction was shown to proceed in the presence of spherical bodies resembling matrix vesicles (Yamamoto, 1986). Apparently, early and active AEFC mineralization follows a globular pattern, but it is unclear whether in humans this is guided by matrix vesicles. However, it is known that periodontal ligament cells are rich in alkaline phosphatase activity (Chomette et al., 1987; Oshima et al., 1988; Piche et al., 1989) and this enzyme is believed to act in the process of cementum mineralization (Beertsen and Van Den Bos, 1991). Further development of AEFC involves fiber fringe elongation in proportion to the advancement of the mineralization front, which eventually becomes smooth and stabilized (Fig. 4b). The remaining, elongated and not or not yet mineralized fringe of fibril bundles shows still the same, constant density (Bosshardt and Schroeder, 1991b; Sequeira er al., 1992), implying that at the base of fiber implantation, i.e., along the dentinocementa1junction, no additional fibers are added to this base and that the

FIG. 4 (a) Electron micrograph depicting the actively progressing mineralization of the initial fiber fringe (FF), with the first -5-pm-thick portion of acellular extrinsic fiber cementum (AEFC) attached to dentine (D) and the dentino-cementa1 junction, and with mineralization centers ahead of the mineralization front (MF) and within AEFC. (b) Electron micrograph with the of an established first layer, 10 to I5 pm thick, of AEFC attached to dentine (D), former fringe fibers extending into the periodontol ligament (PL); the mineralization front (MF) is smooth and has reached stability. DCJ, dentino-cementa1 junction. Magnification: a,b, x3020 (a) From Bosshardt and Schroeder (1991b). (b) Courtesy Dr. D. D. Bosshardt.

16

HUBERT E. SCHROEDER Apical Root Portion

0 Apposition

of AlFC

<

0 >

Apposition

of ClFC

<

0 Initial Matrix

> of ClFC Matrix Attachment

REGENERATIVE CEMENTOGENESIS

17

initially formed fibers become elongated and possibly thicken in diameter, as outlined previously. Once the first, 15- to 20-pm-thick layer of AEFC has been formed, its fiber fringe becomes connected to the principal fibers of the periodontal ligament that run from the tooth surface to bone and the gingiva. From that particular point in development, the AEFC matrix fiber fringe appears as the terminal ends of periodontal ligament fibers, which insert into AEFC as fibers of Sharpey. The fiber region closely abutting the AEFC surface may serve as cementoid giving rise, through mineralization, to the next and overnext layers of AFEC. The mechanisms of fiber connection and the alteration of fiber portions into cementoid and its mineralization are still unknown, both in humans and in rodents. However, it is known that AEFC, after its initial establishment, thickens with age. In human premolars, this amounts to an increase by about 20 pm between ages 10 and 17 years (Sequeira et a/., 1992). Later in life, AEFC continues to grow in thickness at an approximate rate of 1.5 to 3.0 pm/year, and this is true for erupted and functioning as well as for impacted, nonfunctioning teeth (Zander and Hurzeler, 1958; Azaz et a / . , 1974, 1977; Dastmalchi et al., 1990). B. Cellular/Acellular Intrinsic Fiber Cementum

In human teeth, CIFC/AIFC are a normal part of CMSC that covers the apical and furcational root surfaces and more coronally merges with pure AEFC. In single-rooted teeth (i.e., incisors, canines, and most premolars), CMSC coats the apical 10 to 40% of the total root length, i.e., about 1.0 (mandibular second incisors) to 5.7 mm (mandibular first premolars) of the apical root (Schroeder, 1988). Frequently, CIFC appears as the first layer of CMSC; thus it forms on newly produced dentine when the advancing root edge has reached the respective apical region. In this situation, the

FIG. 5 Schematic drawing illustrating topographically the initial stages of CIFC genesis on human premolars, developed to about 75% of their final root length: I , committed clone of precementoblasts contacts root predentine and produces first matrix fibrils; 2, cementoblasts form initial collagenous matrix attached to predentine; 3, first formed and mineralized CIFC, including cementocytes, grows by apposition; 4. CIFC is covered with a layer of AIFC. AIFC, acellular intrinsic fiber cementum; CB/u, cementoblasts with unipolar matrix production; CIFC, cellular intrinsic fiber cementum; ERM, epithelial rests of Malassez; MD, mineralized dentine; CBlc, cementoblasts with potential to become cementocytes; CC, cementocytes; PD, predentine; MF, mineralization front; CM, initial cementum matrix; CB/m, cementoblasts with multipolar matrix production; NMD, nonmineralized dentine or predentine; CPCB, committed precementoblasts; ARE, advancing root edge; HES, Hertwig’s epithelial root sheath.

18

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

19

initial layer of CIFC is a very fast forming tissue that later increases in thickness and forms still faster than AEFC (Table 11). As shown in human premolars with incomplete roots developed to about 75% of their final length (Bosshardt and Schroeder, 1990, 1992), initiation of CIFC formation, early matrix production, and its attachment to root dentine take place along a very short, i.e., about 100- to 200-pm-long, zone extending coronally from the advancing root edge (Fig. 5). The CIFC is a product of cementoblasts; it resembles bone as a product of osteoblasts (Selvig, 1965; Furseth, 1967, 1969; Schroeder, 1986; Bosshardt and Schroeder, 1990, 1992). The formation of early CIFC follows an apico-coronal gradient that is established over a distance very much shorter than that of AEFC (Fig. 5 ) . In the triangular space between the deviating epithelial root sheath and the newly formed predentine, which may extend coronally up to 100 pm from the advancing root edge, a cluster of large, basophilic, closely abutting cells is situated (Figs. 6a and 6b). These cells, considered to be precementoblasts, have an activated, euchromatin-rich nucleus and abundant cytoplasm rich in rough endoplasmic reticulum cisternae and Golgi fields (Fig. 6b). At the precementoblast-predentine interphase, they form numerous slender, fingerlike, partially branching and randomly oriented cytoplasmic processes extending into the first, still very narrow, newly formed CIFC matrix. The latter consists of loosely and randomly arranged collagen fibrils of varying diameter, which interdigitate with the predentine fibrils (Fig. 9b). Along that interphase, this matrix is not well demarcated from the dentinal matrix, and it gradually becomes thicker in the coronal direction, while precementoblasts withdraw. Immediately coronal to the precementoblast cluster, there is a variably wide and short region of rapid matrix production (Figs. 5 , 6a). Mature cementoblasts, separated from one another, become embedded in CIFC matrix that accumulates between and around them (Fig. 6a, inset). This matrix appears denser than that at the predentine surface and consists mainly of randomly oriented collagen fibrils. However, the cementoblasts

FIG. 6 (a) Light micrograph depicting the root surface zone of rapid CIFC genesis and appositional growth, coronal to the advancing root edge (ARE, see Fig. 5, parts 1-3). (b) Electron micrograph of the cluster of precementoblasts (PC) along the most recently formed root predentin (PD; see Fig. 5, part 1). Coronal to this cluster, cementoblasts (CB) with shallow surface recessions and cytoplasmic processes are surrounded by collagen fibrils of newly produced CIFC matrix (a, inset). Further coronally, cementoblasts (CB) reside along the CIFC surface (a). CC, cementocyte; CIFC, cellular intrinsic fiber cementum; D, dentine; HRS, Hertwig’s epithelial root sheath; MF. mineralization front. Magnifications: a, x570; b, ~ 3 0 2 0 ;inset, X6700 (a,b). From Bosshardt and Schroeder (1992).

20

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

21

enclosed in this matrix have shallow surface recesses bordered by cytoplasmic processes, which begin to compartmentalize the newly formed collagen fibrils (Fig. 6, inset). Thus, in sections cut vertical to the root surface in a corono-apical direction, most of the collagen fibrils, with varying diameters, appear cross-cut in these recessions. As the latter are distributed irregularly in all areas of the cementoblast periphery, these cells are considered to produce CIFC matrix in a multipolar fashion (Bosshardt and Schroeder, 1990). This is supported by the observation of Formicola et al. (1971), that tritiated proline accumulates around such cells in the rat molar. As a consequence, the CIFC matrix rapidly increases in thickness, and some cementoblasts, surrounded by their products, become cementocytes (Figs. 6a, 7b). This area of rapid matrix production continues for a variably short distance along the root surface. It ends approximately level with the mineralized edge of the growing root; i.e., the mineralization front follows an incline up to 45" to the root axis (Figs. 5 and 6a). Cementoblasts that remain at the CIFC surface at this point continue to produce matrix, although more slowly (Bosshardt et al., 1989). This matrix forms a variably narrow layer of cementoid at the mineralized CIFC surface and consists mainly of very densely packed collagen fibrils of rather similar diameter, which to a variable extent are bundled into fibers of varying size. Most of the fibril bundles at the surface of and within mineralized CIFC/AIFC appear cross-cut; i.e., they run parallel to and probably around the root surface rather than perpendicular to it (Figs. 7a and 8b; Schroeder, 1986; Bosshardt and Schroeder, 1990, 1992). Cementoblasts that cover the cementoid of this most recently formed CIFC show typical, deep-seated, partially circular surface recessions and long, slender cytoplasmic processes forming numerous compartments densely filled with cross-cut collagen fibrils (Fig. 7a). These cells also show an activated, euchromatin-rich nucleus with a dense fibrous lamina and a large cytoplasm rich in rough endoplasmic reticulum cisternae and Golgi fields. They also extend finger-like, thin cytoplasmic processes into the bundles of collagen fibrils, which run parallel to and even include such fibrils. Young cementocytes located within already mineralized CIFC appear morphologically identical, and still show slight surface recesses and cytoplasmic processes of varying lengths (Fig. 7b). They are sur-

FIG. 7 (a) Electron micrographs depicting a cementoblast (CB). with deep surface recessions and cytoplasmic processes that compartmentalize the extracellular collagen fibril matrix at the surface of CIFC cementoid (CM: see Fig. 5 . part 3). (b) Electron micrograph of a cernentocyte (CC) within newly formed and mineralized CIFC. surrounded by a matrix halo (MH; see Fig. 5 , part 3). CIFC, cellular intrinsic fiber cementum. Magnifications: a , x 10.000; b, ~ 9 0 0 0 (a) . Courtesy Dr. D. D. Bosshardt. (b) From Bosshardt and Schroeder (1992).

22

HUBERT E. SCHROEDER

rounded by a narrow, halo-like zone of nonmineralized matrix including loosely and randomly arranged collagen fibrils that fill the space between mineralized CIFC and the cementocyte. The mineralization front is indistinct both around cementocytes and along the CIFC surface; focal mineralization centers are absent. The mineralized matrix is very dense and shows a heterogeneous pattern composed of randomly oriented collagen fibrils and numerous bundles of such fibrils cut cross-sectional or in tangential planes. Cementoblasts in the zone of rapidly forming CIFC matrix and that covering the newly formed matrix, i.e., cementoid, at the surface of established and already mineralized CIFC consistently reveal single intracytoplasmic collagen fibrils, either within slender cytoplasmic processes or at the cytoplasm periphery. These fibrils appear to be enclosed in membranebound, electron-lucent compartments. In central portions of the cementoblast-cytoplasm, groups of fibrils in parallel orientation appear in large membrane-bound intracytoplasmic channels (Bosshardt and Schroeder, 1992). It is unclear whether such fibrils are in fact intracytoplasmic or are located in narrow and deep recesses continuous with the extracellular space, as described for tendon and corneal fibroblasts (Birk and Trelstad, 1984, 1986; McBride et al., 1985). Intracytoplasmic, membrane-bound collagen fibrils have been the subject of numerous studies, predominantly concerning gingival and periodontal ligament fibroblasts. Whereas most of these investigations have claimed that such collagen fibrils are undergoing enzymatic degradation (Ten Cate, 1972; Listgarten, 1973; Ten Cate and Deporter, 1974; Eley and Harrison, 1975; Frank et a/., 1976; Garant, 1976; Ten Cate et al., 1976; Rose et al., 1980; Melcher and Chan, 1981; Schellens et al., 1982), a few authors have associated intracytoplasmic collagen fibrils with polymerization and secretion of collagen (Garant and Cho, 1979; Cho and Garant, 1981, 1985). The suggestion that collagen is degraded intracytoplasmatically has been supported by showing either respective enzymatic activity or suggestive ultrastructural details, such as lysosomal bodies. In the case of the periodontal ligament, synthesis and degradation of collagen are part of its specific turnover characteristics (for review, see Schroeder, 1986). Another line of reasoning is that intracytoplasmic collagen fibrils occur “when the rate of procollagen synthesis exceeds the rate of transport to the extracellular compartment,” i.e., in situations of accelerated collagen biosynthesis (Enwemeka, 1991). As no ultrastructural details typical for collagen degradation appear in cementoblasts and because the phenomenon is associated with a process of rapid matrix formation, it has been suggested that intracytoplasmic collagen fibrils of cementoblasts are features of fibril formation and fibril assembly rather than of fibril degradation (Bosshardt and Schroeder, 1992). However, it cannot be ruled out that

REGENERATIVE CEMENTOGENESIS

23

cementoblasts also screen their products and actively participate in removing unsuitable components from the newly formed matrix. In summary, human CIFC genesis involves cementoblasts that may function either in a multipolar, rapid or in a unipolar, slow mode of matrix production (Bosshardt and Schroeder, 1990,1992). In the former case, the rapid matrix development may be one of the reasons for the incorporation of cementocytes, as was suggested by Paynter and Pudy (1958) and Formicola et d. (1971). A dense cementocyte distribution in CIFC may, therefore, signal rapid CIFC genesis. On the other hand, the same cementoblast may be a slow matrix producer, resulting in either CIFC with low cementocyte density or AIFC (Bosshardt and Schroeder, 1990). However, there seems to be a significant difference between the slow-rate matrix production of CIFC and that leading to AIFC. In the former case, cementoblasts form a discontinuous and not strictly unicellular layer at the CIFC surface and the resulting matrix is less well organized. The AIFC is formed by a continuous, unicellular layer of cementoblasts that show typical surface recessions and matrix compartmentalization at the cell-rnatrix interphase, and this matrix shows the highest degree of fibril-bundle organization (Figs. 8a and 8b; Bosshardt and Schroeder, 1990, 1992). Thus, the different modes of matrix production that a cementoblast may be associated with result in dramatic differences in terms of the speed of genesis and the structural pattern of the product. Much less is known about the genesis of cellular cementum in the rat. Apparently, the cellular variety of rat molar cementum is different from human CIFUAIFC in part because cells of the disintegrating epithelial root sheath of Hertwig become trapped between dentine and the newly forming cementum and are thus incorporated (Paynter and Pudy, 1958; Diab and Stallard, 1965; Lester, 1969; Lester and Boyde, 1970; Schroeder, 1986). In addition, Yamamoto and Wakita (1991) have shown that the genesis of cellular cementum in rat molars is associated with the simultaneous formation of Sharpey’s fibers, i.e., compact bundles of collagen fibrils that run perpendicular to and insert into the newly forming cementum matrix of cross-cut or randomly arranged collagen fibrils. In human cementum, AEFC may be formed later on top of already established CIFC/ AIFC, but in the early stages of CIFC genesis, the incorporation of Sharpey’s fibers is a rare event. Cementoblasts are very similar, morphologically, to osteoblasts and may even function analogously, as CIFC resembles bone. In the latter, “protein secretion is generally polarized toward the bone surface, but at regular intervals along the surface of newly forming bone an osteoblast will secrete matrix away from the surface, eventually surrounding itself to become an osteocyte” (Marks and Popoff, 1988). Another way for an osteoblast to become an osteocyte has been proposed by Nefussi er al.

24

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

25

(1991). Osteocyte inclusion may “occur without a matrix synthesis inversion by the future osteocyte and with maintenance of close cell contacts with the replacing cell.” The future osteocyte may cease its matrix-producing activity while an adjoining and replacing preosteoblastic cell turns to actively produce new matrix that covers and embeds the osteocyte. However, Frank and Frank (1969) demonstrated that the osteoblast may also behave as a polarized cell with an alternating secretory activity, i.e., directed either toward the osteoid layer or the opposite side. Similarly, cementoblasts seem to have the option of functioning in either a multipolar or an unipolar fashion. It should be added that the mineralization front may also assume a different progression speed, either rapidly following rapid matrix production or slowly advancing behind slow appositional matrix production (Bosshardt et al., 1989). C. Attachment of Matrices t o Growing Dentine

Attachment of the early, first-formed cementum matrix to root dentine is one of the cardinal aspects of root development. In both humans and rats, cementum matrix attachment occurs on a previously formed dentine surface while the respective tooth erupts. In other words, the dentine surface serving as an attachment site moves in the coronal direction while attachment is secured. The speed of this movement may vary between 10 and 40 pm/day and may be faster during AEFC than CIFC initiation (Schroeder, 1991b; Schroeder et al., 1992). Obviously, this attachment concerns the newly formed cementum matrix rather than the cells producing that matrix. In both AEFC and CIFC the newly formed cementum matrix becomes attached to an outer layer of still nonmineralized dentine. In the case of AEFC, an essential part of this attachment is the implantation and interdigitation of collagen fibril bundles perpendicular to and with the randomly oriented collagen fibrils of the dentinal matrix (Fig. 9a; Bosshardt and Schroeder, 1991a). Whether noncollagenous material, such as fibronectin, other mannose-containing matrix components, or unknown glycoproteins, is necessary to facilitate or enhance matrix attachment is unclear at present, at least for human cementogenesis. However, the cells

FIG. 8 (a) Light and (b) electron micrographs depicting a continuous, unicellular layer of cementoblasts (CB), with surface recessions and fibril compartmentalization against the cementoid layer (CM) covering acellular intrinsic fiber cementum (AIFC), including numerous cross-cut bundles of collagen fibrils (b). Magnifications: a. x 1000; b, ~ 3 0 2 0(a,b) . From Bosshardt and Schroeder (1990).

26

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

27

producing the collagen fibrils and organizing the fiber fringe do not attach to predentine. Rather, by means of cytoplasmic processes, they initially contact the dentinal matrix and this contact may be mandatory in triggering their commitment to function as a particular class of AEFC-forming fibroblasts. In the case of CIFC, precementoblasts appear to attach to the most recently formed part of root dentine at the advancing root edge, but even in this situation contact for cell commitment may be all that is required, as these precementoblasts immediately start to produce the first cemental matrix against that same surface. This early matrix of CIFC is loose and irregular, but its randomly oriented collagen fibrils also intermingle with those of the predentine (Fig. 9b; Bosshardt and Schroeder, 1992). In both cases, AEFC and CIFC, the attachment of first-formed matrix fibrils has to be met and supplemented by two requirements. First, as the attachment site, i.e., the surface of predentine, is moving and newly produced portions of root dentine are continually offered for further attachment, there is a demand for new cementogenic cells to be generated. Whether these new cells move in from the surrounding dental follicle proper, as suggested for rat cementogenesis (Cho and Garant, 1989), or originate by cell division from the cluster of precementoblasts seen in human CIFC genesis is unknown. It is tempting, however, to consider the cluster of human CIFC precementoblasts a pool of dividing cells that do not move but remain located with the deviating epithelial root sheath, furnishing the moving root surface with the population of cementoblasts necessary to continue CIFC genesis. Second, the initial interdigitation and attachment of cementum matrix to predentine must be tightly secured by mineralization. In the case of AEFC, the mineralization front that temporarily remained behind the dentinal root surface starts to advance and gradually approaches and overruns the interphase between the interdigitating cemental matrices and the dentinal matrices, i.e., the future cemento-dentinal junction, as soon as the maximum fiber fringe density and, thus, the maximum collagen fibril interdigitation are established (Bosshardt and Schroeder, 1991a). In other words, the primary interdigitation of the two matrix fibril populations before mineralization and the secondary reenforcement by the mineralizing process allow an intimate fibrous linkage and guarantee a force-resistant cemento-dentinal junction.

FIG. 9 Electron micrographs depicting the interdigitation of cemental with predentinal collagen fibrils at the cernento-dentinal junction of acellular extrinsic fiber cementum (a, AEFC) and cellular intrinsic fiber cementum (b, CIFC). PD, predentine. Magnifications: a, x 14,400; b, x 10,000. (a,b) Courtesy Dr. D. D. Bosshardt.

20

HUBERT E. SCHROEDER

The same is true for CIFC, but in this case, the process of early matrix production and the reenforcement of the rather loosely arranged interdigitation of the cementa1 and dentinal fibril populations by mineralization are much faster. In fact, the mineralization front, being inclined to the root axis, moves in an oblique corono-apical direction rather than parallel to the root surface. Based on such differences in development, structure, and reenforcement of the attachment zone, i.e., the cemento-dentinal junction, it could be assumed that the force resistance between AEFC and dentine is greater than that between dentine and CIFC.

IV. Matrix Formation on Established Root Surfaces in Vitro

Based on a series of experiments (Bellows et al., 1980, 1981, 1982a,b) indicating that “periodontal ligament fibroblasts contract vigorously in vitro, that they attach to exogenous collagen fibers with which they are cultured, and that they orient themselves and the fibers between pairs of mineralized tissue particles,” i.e., dentine, “to which they are attached” (Pitaru and Melcher, 1983),a new interest developed to study the behavior of cells and their products in an in vim-simulated interdental space. In particular, the orientation and attachment of cells and collagen matrices to root surfaces and the formation of a matrix resembling that of natural cementum along such surfaces were the subjects of a series of investigations. A. Culture Material and Experimental Design

The in uirro model used for studying orientation and attachment of cells and endogenously produced collagen fibers to root surfaces in a simulated interdental or periodontal space was first introduced by Pitaru and Melcher (1983) and later modified first by Aukhill and Fernyhough (1986) and then by Bernstein et af. (1988). In principle, it consisted of a cultural system in which cells were grown in the presence of denuded root discs, about 150-300 pm thick, that were cut transversely from middle and apical portions of extracted or otherwise removed, healthy teeth. Following different kinds of pretreatment, the denuded root discs were placed in pairs and in part mechanically secured on the bottoms of plastic culture dishes, with a space of 0.1 to 0.5 mm left between the paired discs. In a modified system (Aukhil and Fernyhough, 1986), 600-pm-thick root discs were placed into slightly wider rings of cortical bone, with a space of 0.1-1 .O mm between tooth and bone. Cells were either added to pretreated

REGENERATIVE CEMENTOGENESIS

29

discs or precultured prior to disc placement. The complete system was cultured under standard conditions supplemented with ascorbic acid for periods up to 90 days. In the original and the modified model, cells and discs were in contact with the bottom of the culture dish, whereas in the model of Bernstein et al. (19881, discs and cells rested on a filter that was suspended in the culture medium (Fig. 10a). The materials that were used to compose the model systems varied widely, at least originally. Cells used were human gingival fibroblasts obtained from the American Type Culture Collection (Rockville, MD; Pitaru and Melcher, 1983,1987; Pitaru et al., 1984a,b; Melcher et al., 1986) or grown from explants of gingival biopsies (Pitaru et al., 1984b; Aukhil and Fernyhough, 1986; Quarnstrom and Page, 1986; Fernyhough et al., 1987),and the periodontal ligament cells grown from the roots of rat molars (Melcher et al., 1986). Root discs originated from porcine mandibular premolars (Pitaru and Melcher, 1983, 1987; Pitaru er al., 1984a,b) or primary molars (Melcher er al., 1986) removed from animal cadavers, and from extracted human teeth of unspecified type (Aukhil and Fernyhough, 1986; Quarnstrom and Page, 1986; Fernyhough er al., 1987). Bone rings were prepared from cortical bovine bone of unspecified origin (Aukhil and Fernyhough, 1986; Fernyhough er al., 1987). It is clear, retrospectively, that a model composed of material, living or dead, derived from such a variety of species (rat, pig, human) and tissues (gingiva, periodontal ligament, teeth, bone) is extremely heterogeneous, inaccurate, and not necessarily valid, biologically. These original model systems were used to answer a series of questions, such as whether mineralized or demineralized root discs are a favored substrate for cell and matrix attachment, whether cell attachment is a prerequisite for fiber attachment, whether a root surface still covered with natural cementum or one with exposed root dentine enhances cell attachment and fiber orientation, whether a matrix analogous or similar to the connective tissues in the interdental and periodontal spaces develops in uitro, and whether some forms of cementum matrix could be generated in uitro (Pitaru and Melcher, 1983, 1987; Pitaru et al., 1984a,b; Aukhil and Fernyhough, 1986; Melcher et al., 1986; Quarnstrom and Page, 1986; Fernyhough et al., 1987). The original model was modified twofold, first by using autologous human material and standardized methods of microscopic evaluation (Preisig and Schroeder, 1988) and second by employing a new culture system in which nourishment is provided from both above and below the cellular matrix (Fig. IOa; Bernstein et al., 1988). In contrast to the original model used primarily to examine the fibro-cellular “tissue” that develops between two opposing root discs, or the root disc and bone, the new culture system was employed to focus primarily on the collagenous matrix that develops at, on, and along the established root

30

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

31

surfaces. This system, using human periodontal ligament cells and alveolar bone cells, both grown from explants of the same subject who also provided the extracted teeth for root disc preparation (Bernstein et d., 1988), had the disadvantage that a particular experiment was based on autologous material from an individual patient and, therefore, restricted in scope.

B. Formation and Attachment of Collagenous Matrices Any in uitro model devised to study aspects of periodontal regeneration obviously must attempt to develop structures similar to those of normal tissues in uiuo, in particular to the tissue most essential for that regeneration, i.e., AEFC. Neither the original nor the lately modified model systems (see Section IV,A) have achieved that goal, although using the system of Bernstein et al. (1988, 1989a,b, 1990) two findings were helpful. First, a collagenous fiber fringe attached perpendicular to and intermingled with the matrix fibrils of established but demineralized root cementum developed occasionally, a fringe similar to that in early AEFC genesis. Second, this autologous and rather standardized system furnished some indications why such model systems are necessarily inefficient, at least for the time being. In the original model of Pitaru and Melcher (1983), employed for 20-30 days and later for short-term culturing, multiple layers of gingival fibroblasts and collagen fibers developed over 10 days in about 60% of the culture sets. These layers formed sheets that radiated perpendicularly from the root disc surfaces and bridged and eventually filled the interdisc space. The fiber component, being phase-contrast refractile and histochemically positive for collagen, assumed parallel orientation and extended across the interdisc space as well as from the root disc periphery not opposed by another disc. Because the refractile material was also seen after 1 to 6 days of culturing, with the discs placed on top of a confluent layer of gingival fibroblasts precultured for 4 to 6 days (Pitaru et al., 1984a,b), the interpretation of this refractile material bridging the interdisc space as though resembling the arrangement and distribution of transeptal and dentogingival fibers in uiuo was slightly premature (Pitaru and

FIG. 10 (a) Schematic drawing of experimental culture model. (b,c) Light and (d) electron micrographs depicting the interdisc space (b) with a matrix forming a dense fringe (FF) of collagen fibrils (c,d) perpendicular to and intermingled with cementa1 fibrils (d) along cementum-lined (C) root surfaces in a 42-day-old culture of human periodontal ligament cells cocultured with autologous root discs (RD). Magnifications: b, X 10; c , X 150; d, x7000. (a-d) From Bernstein et al. (1988).

32

HUBERT E. SCHROEDER

Melcher, 1983). In addition, the idea that precultured fibroblasts migrate toward and attach to the root discs within 24 hr (as seen by time-lapse photography of the disc surfaces in whole-mount culture preparations) because they are attracted by them (Boyko et al., 1980; Pitaru et al., 1984a) gave rise to a series of attachment assays introduced some years later (see Section IV,C). Furthermore, it was suggested that partially demineralized substrates favor cell matrix orientation and attachment, whereas demineralized cementum is more efficient in maintaining that attachment (Pitaru and Melcher, 1983; Pitaru et a / . , 1984a,b). The first light- and electron-microscopic studies on the development and composition of the interdisc matrix synthesized in uitro were reported by Quarnstrom and Page (1986), Fernyhough et al. (1987), and Pitaru and Melcher (1987). Using totally or very briefly decalcified root discs of unspecified human teeth, planed to create a smooth dentine surface, and a preculture of confluent human gingival fibroblasts of the 5th to the 10th passage, Quarnstrom and Page (1986) described the development and composition of the interdisc matrix histologically and biochemically in cultures 5 days to 13 weeks old. Initially, cells attached to and migrated up the vertical root surfaces. Eventually, they filled the entire interdisc space with extracellular matrix. Attachment and mobility of cells to and along the demineralized root surface was explained by the presence of fibronectin and the exposure of collagen fibrils. The first interdisc matrix was shown to contain hyaluronic acid, chondroitin sulfate, dermatan sulfate, and large amounts of fibronectin. Collagen fibrils appeared only after 6 weeks in culture and fibers became denser with time. In late cultures the matrix contained collagen of Types I, 111, and V (Quarnstrom and Page, 1986). Although some fibroblasts of the interdisc matrix assumed a shape indicating a tendency to compartmentalize extracellular collagen fibrils, no fiber fringe structures were observed along the vertical disc-dentine surfaces. Similar results, albeit restricted to microscopic evaluation, were obtained in 20- to 30-day cultures of bovine disc pairs and human gingival fibroblasts (Pitaru and Melcher, 1987). These authors, however, reported that along decalcified cementum surfaces, cells in high density were oriented mainly perpendicular to that surface, whereas along nondecalcified surfaces, less numerous cells lay with their long axis parallel to the disc surface. And the same was true for matrix collagen fibrils. Initial cell attachment at the demineralized cementum surface was established either by cell processes penetrating between cementum collagen fibrils or by the plasma membrane being closely apposed to such fibrils, possibly suggesting a fibronectin-independent or fibronectin-mediated process. Pitaru and Melcher (1987) felt that “the continuous layer of gingival fibroblasts oriented parallel to the non-decalcified cementum and the similarly oriented extracellular fibrillar matrix is reminiscent of a capsular structure,”

REGENERATIVE CEMENTOGENESIS

33

whereas the cell matrix orientation at demineralized surfaces would resemble the in uiuo arrangement. However, the findings of Pitaru and Melcher (1987) as illustrated in their paper were very much unlike the in uiuo situation and the same was true for the observations of Fernyhough et al. (1987), who used the root disdbone model. The modified culture system of Bernstein et al. (1988) yielded more consistent results, in particular with respect to the formation of a dense fiber matrix along established cementum surfaces. In a series of experiments (Preisig and Schroeder, 1988; Bernstein et al., 1988, 1989a,b, 1990) using autologous human periodontal ligament cells and demineralized root discs precoated with fibronectin in 6- to 13-week-old cultures that rested on a Puropor-200 filter supported by a wire-mesh grid (Figs. 1Oa and lob), it was confirmed that dentine-lined root surfaces were often encapsulated by cells and collagen fibrils running parallel to that surface. Along cementum-lined root surfaces, a variably dense fringe of collagen fibrils that interdigitated with those of the cementum matrix developed within 3 to 6 weeks (Figs. 1Oc and 1Od).This fringe with its fibrils oriented perpendicular to the cementum surface extended for a short distance into the interdisc space or the disc surroundings. However, the fringe was not continuous and covered patchwise variably large areas of the root surface. Fibroblasts were positioned irregularly. This fringe being rather short and occasional was not to be equated with the refractile material of cell fiber sheets, radiating from disc surfaces (Pitaru and Melcher, 1983; Pitaru et al., 1984a,b). In older cultures, the interdisc matrix and its cells appeared to deteriorate, whereas new cell matrix layers had formed on top of that space. In other words, the fiber fringe fully developed after 6 weeks did not increase in density with time. When autologous cells derived from human alveolar bone were added to that model system, a more continuous and very much denser fiber fringe developed along most of the cementumlined disc surfaces and this fringe remained relatively unchanged over 13 weeks in culture (Figs. 1Oc and 1Od; 1 la and I lc). Because of a high degree of fibril interdigitation, the interface between the newly formed matrix and the established but demineralized cementum attained structural continuity (Fig. 1 Ic). Thus, the combination of autologous human periodontal ligament cells and alveolar bone cells produced an improved fiber fringe intimately attached to established cementum and indeed somewhat reminiscent of early AEFC matrix and its attachment to dentine (see Section 111,A). The majority of collagen fibrils produced by that combination had a diameter within the 50- to 80-nm range, whereas the fibrils produced by periodontal ligament cells alone were mainly in the 20- to SO-nm range (Table 111). The former distribution was indeed similar to that in cementum (Table 111). In addition, as a less frequent finding, single periodontal ligament cells or single cells in ligament/bone cell mixtures were found to be

34

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

35

surrounded by their collagenous fibril product in a halo fashion (Fig. 1 Ib). This was possibly an indication for some cells assuming a cementoblast phenotype, producing an early form of CIFC matrix (see Section 111,B). Intentionally, the autologous model system of Bernstein et al. (1988, 1989a) was not supplemented with Na-P-glycerophosphate, which had been shown to allow the production and calcification of bone-like matrix nodules (Williams et al., 1980; Ecarot-Charrier et al., 1983, 1988; Sudo et al., 1983; Bhargava et a / . , 1988) and even the assembly, by osteoblasts, of a bone-specific macrostructure in uitro (Gerstenfeld et al., 1988). Bernstein er al. (1989a,b) merely examined the contribution bone cells would make to the development of a “cementoid” fiber fringe matrix attached to the established cementum surface. When Melcher et al. (1986, 1987) cultured fetal rat calvaria cells in the presence of porcine root discs in medium supplemented with ascorbic acid and Na-P-glycerophosphate, they claimed to have found varieties of “cementum” newly formed on the root disc surfaces. However, in no instance did they report a true fibrous attachment with a dense collagen fibril arrangement, and the nodules forming on cementum-lined discs and elsewhere in their cultures were similar to bone-like centers produced in systems without root discs (Bhargava et al., 1988; Maniatopoulos el a / . , 1988). The finding of an improved fiber fringe matrix attached to scaled, cementum-lined but demineralized root surfaces by the addition of alveolar bone cells to autologous periodontal ligament cells (Bernstein et al., 1989a) supported the hypothesis that, in uiuo, cells originating in alveolar bone may migrate to the periodontal ligament and function as cementum-producing cells (McCulloch et al., 1987; Melcher et al., 1987). However, in a subsequent experiment using the same model system with demineralized and nondemineralized discs precoated with normal culture medium, fibronectin, or autologous serum, and with the culture medium in part being supplemented with 10% autologous serum for the first 17 days, Bernstein et a/. (1989b) were unable to further improve the fringe matrix. The collagen fibrils were again in a diameter range of 40-60 nm or beyond (Table 111). The fiber fringe did not become denser or more consistent, and the newly formed matrix again displayed a patchwise distribution. A dense collagenous fiber matrix was also produced along nondemineralized, ce-

FIG. l l (a) Light and (b,c) electron micrographs depicting (a) the interdisc matrix (IDM) with cells surrounded by discrete halos of collagen fibrils (b, arrowheads) or a dense fiber fringe (FF) continuous and intermingled with cementa1 fibrils (c, inset) along cementurn-lined ( C ) root surfaces in a 56-day-old culture of human periodontal ligament cells and alveolar bone cells cocultured with autologous root discs (RD). Magnifications: a, x580; b,c, x2700; inset, ~ 5 8 0 (a,c) . From Bernstein er a / . (1989a).

TABLE Ill Diameter (nm) of Collagen Fibrils of the Fiber Fringe Matrix Grown in Cultures, Compared to Cementum Fibrilsa % Diameter distribution (nm)

Root discs (decalcified) Days in culture 56 56 124 124 Cementum Cementum

F-coated F-coated F-coated F-coated -

-

Cells

N

Diameter (x ? s)

HPLC HPLC + HBC HPLC HPLC + HBC -

80 80 160 160 320 320

42+ 65 ? 472 63 ? 642 71 +

Diameter (x + s)

Range

20-40

40-60

552 9 51 + 10 58 & 10

32- 83 26- 86 26- 86

7 15 6

69 67 59

9 14 9 I1 11

13

Range

20-50

50-80

80- 100

25- 64 38-101 32- 79 39- I05 32- 97 37-109

79 10 61 8 9 4

21 78 39 86 83 72

0 12 0 6 8 24

% Diameter distribution (nm)

Days in culture 35-42 35-42 35-42

Root discs (decalcified)

Cells

N

M-coated F-coated AS-coated

HPLC + HBC HPLC + HBC HPLC + HBC

180 180 180 ~~

" Data from Bernstein

~~

60-80 24 18 35

~

er al., 1988, 1989a,b; N = number of measurements; F, fibronectin; M. medium; AS, autologous serum; HPLC, human

periodontal ligament cells; HBC, human alveolar bone cells.

REGENERATIVE CEMENTOGENESIS

37

mentum-lined, sterile root discs derived from previously diseased and experimentally planed teeth with advanced periodontitis (Bernstein et al., 1990). However, the smooth, hard, and chemically as well as mechanically denatured root surface did not appear to be a suitable substrate for attaching a newly produced matrix. Such surfaces do not provide exposed collagen fibrils with which a new fibril population could intermingle, and this is exactly the situation for clinically treated root surfaces that are needed to serve as attachment sites for regenerating cementum.

C. Factors and Failures Retrospectively, attempts to study regenerative phenomena such as interdental connective tissue matrix development and cementogenesis along established root surfaces in uitro have failed so far. All in uirro models used have focused on an experimental situation in which two factors, i.e., cells and established but denatured and modified root surfaces, were elected to interact. Aspects of normal wound healing processes, including blood clot formation and clearance of traumatized wound edges by phagocytosis and resorption and the several humoral factors that participate in would healing, were intentionally excluded. The most simplified model situation indeed did not permit the observation of anything but the behavior of cells and the arrangement of matrix products in the presence and along denatured root surfaces. Also, both of these model components were heterogeneous, nonspecified biologically, and most often of unknown origin and ontogeny. This is particularly true of the cells used in such model systems, i.e., gingival and periodontal ligament fibroblasts and (alveolar) bone cells freed by enzyme treatment (Melcher et al., 1986). All of these cell populations were heterogeneous mixtures of functionally unknown properties, both when harvested from biopsies and after propagation in culture. Gingival fibroblasts grown from human gingival biopsies represent a mixture of ill-defined groups of cells that include subpopulations of different functional capacity and subpopulations that may be phenotypically stable both in uiuo and in uitro or regulated by extracellular factors (Narayanan and Page, 1983; McCulloch and Bordin, 1991). For example, human gingival fibroblasts grown from normal gingiva include subpopulations with a heterogeneously expressed Clq-receptor (Bordin et al., 1983,1984). Cells with a surface marker for the collagen-like domain of Clq are activated by that molecule and increase their DNA synthesis, rate of proliferation, and total protein production, the latter about threefold, with 40% of this production directed to collagen synthesis, in particular of Types I11 and V. These properties remain stable in culture, at least over 12 doublings

38

HUBERT E. SCHROEDER

(Korotzer et al., 1980; Bordin et a / . , 1983, 1984; Bordin and Page, 1988, 1989). Mass cultures of human gingival fibroblasts derived from one individual biopsy were shown to differ widely in proliferation rate and production of protein, in particular the synthesis of collagens and glycosaminoglycans (Hassel and Stanek, 1983). The types of collagens produced by such mixed populations are I, 111, IV, and V, the latter two as minor species (Hurum et al., 1982; Narayanan and Page et al., 1983), and most cells can produce Types I and I11 collagens simultaneously, whereas some produce only Type I or none (Engel et al., 1980).The two main glycosaminoglycans synthesized by human gingival fibroblasts are hyaluronic acid and heparan sulfate, but chondroitin sulfate and dermatan sulfate (Bartold and Page, 1985) are also synthesized. Clones of human gingival fibroblasts, varying in size, morphology, and growth rates, are rather similar in their ability to synthesize collagen, collagenase, and collagenase inhibitors and to degrade native collagen mats, indicating that such clones can both synthesize and degrade collagen (Hurum et al., 1982). However, any particular fibroblast can either secrete or degrade collagen but cannot do both simultaneously (Yajima, 1988). Furthermore, human gingival fibroblasts can be functionally modified by extracellular factors: prostaglandin E2 inhibits their growth and collagen synthesis (KOet al., 1977); concanavalin A stimulates their synthesis of collagenase about 10-fold (Hurum et al., 1982); guanidine/EDTA extracts of proteins from bovine cementum, dentine, and alveolar bone stimulate their total protein production and collagen synthesis (Somerman et al., 1987a,b). However, it is not known whether these modifications in cellular activity occur or can be induced in all or only in particular subsets of human gingival fibroblasts that, like human dermal fibroblasts, can be made to increase their collagen production, independent of cell growth, by a factor of 2 to 3 in the presence of ascorbic acid (Russell et al., 1981). It is also unclear whether there are stem cells for fibroblast propagation. McCulloch and Knowles (1991) demonstrated that in hamster gingiva and cultures thereof, 40% of the fibroblasts divide in uiuo and in v i m , a further 40% of these cells divide only in uitro, and about 10% do not divide in either situation. That may suggest that in the hamster gingiva, there is a population of actively cycling progenitor fibroblasts, another population that is growth-inhibited in uiuo but capable of growing upon explantation, and a third population of terminally differentiated cells. Truly colony-forming progenitor fibroblasts, i.e., stem cells, may represent a very small population in uivo (i.e., about 0.5% of all isolated cells: McCulloch and Knowles, 1991). For all these reasons, subcultures of human gingival fibroblasts (and that of animal models) in fact are heterogenous mixtures of cell subsets of unknown properties and of unknown proportions that may vary from culture to culture and donor-to-donor tissue. Nevertheless, in continuation

REGENERATIVE CEMENTOGENESIS

39

of the in uitro model work discussed here (Pitaru et al., 1984a,b), such illdefined mixtures of human gingival fibroblasts have been used to study their migration and attachment to untreated and modified porcine root slices and various alloys in short-term cultures (Lowenberg et al., 1985, 1986, 1987) as well as to diseased and nondiseased human roots treated by citric acid- or EDTA-demineralization and collagenase (Fardal and Lowenberg, 1990). Similar attachment studies were performed with mixtures of human gingival fibroblasts added to diseased and nondiseased roots (or slices thereof) freed from cementum and coated with human plasma (Abbas et al., 1987) or fibronectin (Fernyhough and Page, 1983), and to Petri dishes or wells coated with guanidine/EDTA protein extracts of unspecified bovine and human cementum (Somerman et al., 1991). Also, human gingival fibroblasts were cultured for 2 hr on fibronectincoated and/or tetracycline-treated blocks of bovine dentine (Terranova el al., 1986). All authors reporting such experiments usually claimed that cell-attachment assays used to screen conditions of favored or enhanced attachment to established but modified root tissue surfaces such as blank cementum or dentine would contribute directly to the clarification of a principal phenomenon required for regenerative cementogenesis. Similar experiments and arguments were applied to study the behavior of so-called periodontal ligament cells. When cultured, such cells are derived from explants of the residues of natural periodontal ligament tissue that remains at the healthy root surface after tooth extraction. Developmentally, periodontal ligament cells are of ectomesenchymal origins and residents of the dental follicle proper that invests tooth germs. In the mouse, such ectomesenchymal cells of the dental follicle have been shown to give rise to and differentiate into cementogenic, osteogenic, and fibroblastic lineages forming the functionally united root cementum, alveolar bone proper, and periodontal ligament (Ten Cate et al., 1971; Barrett and Reade, 1981; Yoshikawa and Kollar, 1981; Palmer and Lumsden, 1987). However, this totipotency of periodontal ligament cells is restricted to the time period of root development and has so far not been detected in any of the cells residing in the periodontal ligament of fully developed and functioning teeth (McCulloch and Bordin, 1991). For culturing, periodontal ligament cells have been harvested from a variety of different species such as pigs, cattle, dogs, non-human primates, and humans. Following tooth extraction, at least some of such cells can be kept viable at their original site at the root surface for up to 1 year (Litwin et al., 1971) and harvested by trypsinization or scraping (Blomlof and Otteskog, 1981; Ragnarsson et al., 1985; Oikarinen and Seppa, 1987). Morphologically, cultured human periodontal ligament cells are fibroblastlike and spindle-shaped or stellate, with numerous, long cytoplasmic extensions protruding in the direction of their long axis (Ragnarsson et al.,

40

HUBERT E. SCHROEDER

1985; Preisig and Schroeder, 1988).Ultrastructurally, these cells reveal an euchromatin-rich nucleus, and a cytoplasm containing the usual set of organelles and single and compact bands of microfilaments, i.e., vimentin and possibly actin, numerous endocytotic vesicles, and variable amounts of glycogen storage granules (Chomette et al., 1987;Preisig and Schroeder, 1988; Rose et al., 1987). Surprisingly, probably because of the presence of a preosteoblast subset, periodontal ligament cells reveal a high alkaline phosphatase activity similar to that of osteoblasts (Kawase et al., 1986; Piche et al., 1989) and this is also found in periodontal ligament cells of bovine incisors, which also produce increasing amounts of 3’, 5’-cyclic adenosine monophosphate in response to human PTH, and a marker for osteoblasts (i.e., a protein cross-reactive with bovine bone Gla protein; Nojima et al., 1990). Clones of porcine or monkey periodontal ligament cells are capable of synthesizing collagen Types I, 111, and V in varying proportions, i.e., 10 to 30% of the total collagen for Type 111and extremely high or low amounts of Type I, from clone to clone (Limeback et al., 1978, 1983). This is a clear indication that such periodontal ligament cells are a heterogenous mixture of cells producing distinctly different ratios of collagen types. Some clones of human periodontal ligament cells produce a heat-resistant factor that inhibits the PTH-induced resorption process in fetal long bone cultures (Giniger et al., 1991). Further evidence for the heterogeneity of periodontal ligament cells is derived from in uiuo studies. In both wounded and normally functioning periodontal ligament of the mouse, paravascular cells with small nuclei, rather undifferentiated ultrastructurally, appear to have progenitor or stem cell characteristics (Gould et al., 1977, 1980; McCulloch and Melcher, 1983b; Gould, 1983). These paravascular cells constitute 15 to 20% of all ligament cells (McCulloch and Melcher, 1983b),have the capacity for a relatively high rate of proliferation, migrate to the bone and cementum surfaces (McCulloch and Melcher, 1983a), cycle more slowly than cells outside the paravascular region (McCulloch, 1985), and are found in the endosteal spaces of the alveolar bone, from which they migrate via Volkmann’s channels into the periodontal ligament (McCulloch et al., 1987). These cells, being considered a stem cell population of clonal distribution (McCulloch, 1985) may be able to augment fibroblasts, cementoblasts, or osteoblasts because of their totipotency (McCulloch and Bordin, 1991).This is supplemented by the finding that periodontal ligament fibroblasts and preosteoblasts express numerous specific binding sites for epidermal growth factor in uiuo, whereas fibroblasts of other connective tissues have only a few binding sites (Cho et al., 1988). These ligament cells may be of common origin and related to paravascular stem cells. Whether paravascular cells of similar characteristics are present in the human periodontium is, however, unknown.

REGENERATIVE CEMENTOGENESIS

41

Comparative studies with human gingival fibroblasts and human periodontal ligament cells have revealed that both mixed populations are not alike. In vivo studies in the mouse demonstrated that the collagen turnover rate is five times higher in the periodontal ligament than in gingiva (Sodek, 1976, 1977), and in the rat, periodontal ligament cells including preosteoblasts consist of subpopulations that respond with proliferation patterns different from those to traumatic wounding (Gould et al., 1977, 1980), endocrine (Roberts, 1975a,b)and electrical (Davidovitch et al., 1980)stimulation, and orthodontic forces (Roberts et al., 1982; for review see Schroeder, 1986). In vitro, mixed human periodontal ligament cells are significantly more active in their protein and collagen synthesis and have a significantly greater alkaline phosphatase activity than gingival fibroblasts (Somerman er al., 1988; Arceo et al., 1991). Furthermore, mixed human periodontal ligament cells, but not gingival fibroblasts, are “capable of producing mineral-like nodules in vitro” (Arceo et al., 1991). Porcine periodontal ligament cells, either in clones or in mass cultures, are relatively homogeneous in the expression of collagen Type I and fibronectin; i.e., 99% of all cells express both. In contrast, porcine gingival fibroblasts are considerably heterogeneous; only 57 to 78% of the cells express both collagen and fibronectin, 21 to 42% express only collagen, and 0.4 to 0.8% express only fibronectin (Connor et a/., 1983). In attachment assays, human periodontal ligament cells respond preferentially to fibronectin-coated blocks of dentine or wells, whereas gingival fibroblasts are promoted to attach to surfaces coated with bone phosphoprotein (i.e., osteopontin) and with guanidine/EDTA extracts of bone and cementum (Somerman er al., 1987c,d, 1989;Terranova et al., 1987,1989b). However, as cultures of both periodontal ligament cells and gingival fibroblasts are heterogeneous mixtures of different cell subsets, it is unclear whether such differences are typical for all or any particular set of cells. Finally, alveolar bone cells that can be freed from the marrow spaces and bone surfaces are again mixtures of heterogeneous cell populations that differ due to their source and origin. None of the isolation procedures used provides a pure homogeneous population of structurally and functionally identified cells (Nijweide et al., 1986; Marks and Popoff, 1988). Cementoblasts or precementoblastic precursor cells have not been isolated or detected yet at all. In fact, it is entirely unknown where they come from, whether they proliferate, how long and under what conditions they function, and where they go to or reside in reserve. And this is true for the particular class of fibroblasts forming AEFC as well as for the osteoblastlike cementoblasts producing CIFC/AIFC (see Sections III,A and 111,B). The second component of the in vitro model for studying regenerative matrix production, i.e., denuded, planed, and artificially modified root

42

HUBERT E. SCHROEDER

surfaces of human and mostly porcine teeth, is likewise of heterogeneous, often unidentified nature. Not only were cementum- and/or dentine-lined surfaces, denuded or denatured, still mineralized or demineralized, employed side by side, but there was also no differentiation between AEFC and CIFC or CMSC (see Section 11). As these varieties of root cementum differ from one another in terms of their genesis, structure, and composition (Schroeder, 1986), it is possible that their ability to serve as a favored substrate for regenerative cementogenesis is distinctly different as well. For example, if interdigitation of collagen fibril populations of dentinal and cementa1 matrices is required, decalcified AEFC, with its fibrils running perpendicular to the root surface, is a much better candidate than CMSC or CIFC/AIFC, in which most fibrils run parallel to the root surface. The latter is also true for dentine. Apart from such structural differences, the various cementum varieties may, in addition, possess different biochemical characteristics. For example, studies have suggested that extracts of healthy human (and bovine) cementum modify the migration, attachment, and growth of gingival or periodontal ligament fibroblasts (Miki et al., 1987; Somerman et al., 1987a-d, 1989; Nakae et al., 1989; Nishimura et al., 1989; McAllister et al., 1990) and that human and bovine cementum contains two sialoproteins, i.e., bone sialoprotein I1 and osteopontin (Somerman et al., 1990; Olson et al., 1991). However, in none of these studies was root cementum defined as one or the other variety. In conclusion, in vitro models simulating interdental or periodontal relationships have been used under conditions lacking any possibility to define the model component parameters. Under such conditions, it is not possible to study the sequential evolvement of the cells and matrices or the origin and character of a particular matrix or its attachment to established root surfaces. This has already been pointed out by Bernstein er al. (1989b). Furthermore, it is questionable whether the fact that mixed gingival fibroblasts attach to denuded but variably coated dentine has anything in common with biological problems of periodontal regeneration, specifically regenerative cementogenesis. As shown in Section I1 of this review, it is not clear whether cells, acting in wound healing, in the presence of a dense blood clot and during the subsequent development of a granulation tissue, indeed do need to attach to a substrate surface prior to forming and attaching their matrix product. The requirements for future in uitro investigations focusing on periodontal regeneration appear to be of a different order: 1. to characterize the various phenotypically different or modulationdependent cells in the human periodontal ligament; 2. to identify, biologically and functionally, the fibroblast-like class of cells forming AEFC and the cementoblasts forming CIFC/AIFC;

REGENERATIVE CEMENTOGENESIS

43

3. to devise methods for harvesting and culturing these cells and to study their behavior under simplified conditions of an in uitro model; and 4. to focus on matrix attachment rather than cell attachment.

V. Regenerative Cementogenesis on Established Root Surfaces in Vivo Clinical studies focusing on the healing and regenerative response of the periodontal tissues to various treatment modalities, although attractive for many decades, have been increasing in number since about 1975 and respective reports are abundant. Reviews attempting to summarize the essential results and failures of such studies have been provided by Selvig (1983), Polson (1986, 1987), Egelberg (1987), Nyman et al. (1989), and Minabe (1991). In the context of this review, selected and restricted groups of original contributions will be used to discuss the major biological problems involved in spontaneous and guided regenerative cementogenesis. A. Clinical and Experimental Conditions

In addition to numerous clinical studies in humans, various animal models have been developed, employing mostly non-human primates and dogs, to explore the periodontal regeneration potential. Such models included (a) autotransplantation of single-rooted teeth (Butcher and Vidair, 1955; Andreasen and Kristerson, 1981; Proye and Polson, 1982), ( 6 )submucosal implantation of tooth roots at the bone-connective tissue interface (Nyman et al., 1980; Karring et al., 1980), ( c ) the window technique creating artificial submucosal defects in the mid-root periodontium ( Jansen et al., 1955; Nyman et uf., 1982a), ( d )experimental creation of chronic periodontitis lesions around single- or multirooted teeth by means of attaching rubber bands or silk ligatures subgingivally (Caton and Zander, 1975; Caton and Kowalski, 1976; Nyman et af., 1980), and ( e ) the experimental production of furcation defects in multirooted teeth (Ellegaard el al., 1973; Johansson et al., 1978; Crigger et al., 1978). In the diseased human periodontium as well as in all such model systems, the principal obstacle during wound healing and tissue regeneration is the fact that one side of the wound defect is the blank root surface lined by varieties of cementum or exposing dentine, i.e., a hard, smooth tissue wall denuded from organic material and compressed by planing instruments. Such surfaces, which

44

HUBERT E. SCHROEDER

primarily had been exposed to the inflammatory environment of a periodontal pocket and covered by bacteria, have undergone pathological changes of both their organic and their inorganic components, such as hypermineralization, accumulation of rare inorganic elements (e.g., fluoride), and contamination with endotoxins (Selvig, 1983). Periodontal wound healing along such surfaces can possibly and eventually provide only four different types of restoration: ( a ) the exposed and treated root surface is lined by epithelium, i.e., in the form of a long junctional epithelium that grows out from the gingival wound edge and covers and protects the denatured root surface; ( b ) the exposed hard tissue being recognized as an inert foreign body is encapsulated by connective tissue with collagen fibers running parallel to its surface; (c) the exposed root surface undergoes initial resorption followed by either bone contact and ankylosis or the formation of new cementum and a periodontal ligament; and ( d ) the exposed and treated root surface is covered by new cementum of one or the other variety, without undergoing resorption (Selvig, 1983; Nyman et al., 1989;Schroeder, 1991a).These options depend on the proliferational speed with which the tissues bordering the defect can react, i.e., the epithelium and connective tissue of the gingiva, the alveolar bone, and the periodontal ligament. That speed is high in the gingival epithelium and low in the periodontal ligament (see Schroeder, 1986, 1991a).

B. Spontaneous Regeneration Using autotrans- or replantation of healthy teeth as a model, the quality and extent of periodontal regeneration depend on the root surface conditions. In freshly extracted teeth, with vital remnants of the periodontal ligament, i.e., cells and torn collagen fibers, remaining attached to the cementum of their root surface, a scar-like healing results in a reconnection of these fibers with that of the surrounding connective tissue, with normal structure and function of the bone-ligament-cementum complex being restored eventually (Butcher and Vidair, 1955; Polson and Caton, 1982; Houston et al., 1985; Blomlof et al., 1988). The healing result is the same when a healthy root is replanted into an alveolar bone socket of artificially reduced height (Polson and Caton, 1982; Houston et al., 1985). Healing and fiber restoration may be completed in about 3 weeks (Hurst, 1972) and circumscribed, artificial notches placed prior to replantation in the midroot may be lined with new AEFC and fiber attachment (Houston et al., 1985). A periodontitis-affected or a healthy root partially denuded from organic material including some or all old cementum, however, is rapidly covered by a longjunction epithelium (i.e., within 2 weeks; Karring et al., 1984),

REGENERATIVE CEMENTOGENESIS

45

even when that root is autotransplanted into a socket of normal height (Stones, 1934; Polson and Proye, 1982, 1983; Lindhe et a/., 1984; Houston et al., 1985, Bowers et al., 1989~;Blomlof et a / . , 1988). If epithelium is excluded from wound healing, i.e., by crown amputation and mucosal closure, a great portion of the denuded root surface will be lined by connective tissue with its collagen fibers running parallel to that surface, or the latter may undergo resorption with or without ankylosis (Karring et af., 1980, 1984; Nyman et al., 1980; Houston et a/., 1985). In such experiments, an observation made in clinically treated periodontitis patients has been repeated (Skillen and Lundquist, 1937; Listgarten and Rosenberg, 1979; Cole et al., 1980; Bowers et a/., 1985, 1989a,b,c), i.e., that at the apical termination of the denuded root surface, usually marked by an artificial notch, and close to the most apically located healthy or restored part of the periodontal ligament and its fiber attachment to old cementum, a variably short strand of new cementum develops, with or without fiber attachment. This occurs more frequently and to a greater extent if epithelium is excluded (Bowers et a / . , 1989a,b,c). As seen in classical histological sections, this new cementum is variably thin, slightly mineralized (Ogura et a/., 1991), of a cellular variety, and present over nonresorbed old cementum and dentine, but does not, due to tissue shrinkage, stick to the old root tissues (Listgarten and Rosenberg, 1979; Bowers et a/., 1989a,b,c). These experiments suggested that: ( a ) gingival and periodontal connective tissue reattaches to the root surface as long as the latter carries vital remnants of periodontal ligament, ( 6 ) bone regrowth and periodontal ligament regeneration are unrelated phenomena, and (c) cementum regeneration and new fiber attachment are possible provided the denuded root surface is first repopulated by cells derived from periodontal ligament and, possibly, the alveolar bone. In fact, using precultured fibroblasts from that ligament and other tissues such as gingiva, dermis, periosteum, and fascia to envelop replanted roots denuded by mechanical scaling, Andreasen and Kristerson (1981) and Boyko et al. (1981) demonstrated that periodontal ligament cells are unique in regenerating cementum and fiber attachment and cannot be replaced by other cells, except perhaps those of a dental follicle. Such cells, therefore, appear as an indispensible prerequisite for regenerative cementogenesis in uiuo. This conclusion was corroborated by experiments using the window technique, i.e., a surgically created submucosal fenestration in the alveolar wall, combined with a local removal of periodontal ligament and the cementum, covered by a Millipore filter (Nyman et al., 1982a; Pettersson and Aukhil 1986; Aukhil et a/., 1986b; Iglhaut et a/.,1988; Knox and Aukhil, 1988; Selvig er a / . , 1988). In this situation, at least part of the exposed root dentine will be covered within about 3 weeks to 6 months,

46

HUBERT E. SCHROEDER

with thin, patchy new cementum, possibly AEFC (Pettersson and Aukhil, 1986), albeit often clearly not AEFC (Nyman et al., 1982a),and the defect will be filled by bone ingrowth and a new periodontal ligament with partial fiber attachment, but root resorption and ankylosis occur frequently. In non-human primates (M. fuscicularis), cells from both the surrounding, intact periodontal ligament and the cut bone surface, labeled by [3H]thymidine, arrive in the window defect 3 days after wounding, and at 21 days, a thin “cementoid” matrix is deposited along the exposed dentine (Iglhaut et al., 1988). In rats, a 2-week window is filled with a fibroblast-rich connective tissue and after 4 weeks, an electron-dense material lines the denuded dentine, from which dense aggregates of inserted collagen fibrils radiate perpendicularly (Knox and Aukhil, 1988). This initial stage of regenerative cementogenesis somewhat resembles the beginning of normal genesis of acellular cementum in the rat (see Section 111,A).Similar results, although with direct attachment of collagen fibrils to decalcified dentine, were obtained in dogs (Selvig et al., 1988). In these cases, regenerative matrix attachment seemed to follow the principle of interdigitation of collagen fibril populations as seen in human cementogenesis (see Section 111,C).

C. Guided Regeneration

The principle of guided periodontal tissue regeneration was born as an extension of the window technique and first introduced by Nyman et al. (1982b). It is based on the argument that cells proliferating from the periodontal ligament bordering the periodontal defect should preferentially and selectively repopulate the denuded root surface. To achieve that purpose, a topographically dressed piece of a physical barrier, i.e., a nonresorbable membrane such as Millipore or Teflon (Gore-Tex) or a biodegradable membrane such as collagen or Vicryl (Polyglactin 910; Zappa, 1991a,b) is placed over the denuded root surface and covered by mucosal tissue. In that position, the membrane extends from the outer bone surface apical and lateral to the root defect coronally underneath the gingival margin, and allows cells of the adjacent periodontal ligament and of alveolar bone to repopulate the blood clot that fills the space between root and barrier, and later the denuded root surface. At the same time, the barrier prevents gingival epithelium and connective tissue from contacting that surface. Such barrier membranes have been used clinically in humans (Nyman et al., 1982b, 1987; Gottlow et al., 1986; Pontoriero et al., 1987, 1988, 1989; Stahl et al., 1990; Stahl and Froum, 1991a,b) as well as in experimental animals such as dogs (Aukhil et al., 1983, 1986a; Pitaru et al., 1987, 1988, 1989; Caffesse el al., 1988, 1990, 1991; Magnusson et al.,

REGENERATIVE CEMENTOGENESIS

47

1988, Claffey et al., 1989) and non-human primates (Gottlow er al., 1984; Magnusson et al., 1985; Aukhil and Iglhaut, 1988; Ettel et al., 1989; Gottlow et al., 1990) presenting with artificial periodontitis produced by the ligature technique (see Section V,A). In part, such experiments were combined with crown amputation and mucosal closure in order to exclude the epithelium from wound healing (Gottlow et al., 1984; Claffey et al., 1989). Unfortunately, the guided tissue regeneration technique is hampered by clinical problems such as incorrect barrier placement due to topographical complexity, postoperative gingival recession, and shrinkage resulting in the exposure of the coronal edge of the barrier membrane, reinfection along the exposed membrane, and premature loss of the latter. On the other hand, the barrier membrane may help to protect the blood clot from mechanical disturbances (Claffey et al., 1989; Selvig et al., 1990). In the most favorable circumstances, a new cementum matrix is formed along up to 70% of the denuded root surface (Minabe, 1991), extending from the apical or lateral border of the still intact periodontium coronally for distances varying between 1 and 5 mm. This regenerated cementum matrix, presumably calcified in uiuo, is variably thin; tapers coronally ; overlaps the old cementum apically; includes cells (i.e., cementocytes or osteocytes ?); and, unless the denuded root surface has preceedingly undergone superficial resorption by dentoclasts, does not adhere to the established root tissues (Fig. 12), neither old cementum nor dentine (Gottlow er al., 1984, 1990; Ettel er al., 1989; Zappa, 1991b). The outer surface of this new cementum matrix is covered by connective tissue that, in the human, rarely shows dense aggregates of collagen fibers oriented perpendicularly to and inserting into that new cementum, although, on the basis of classical histologic sections, some authors claimed to have demonstrated new fiber attachment after about 3 months of healing (Gottlow et al., 1986). This is different in dogs and non-human primates in which a dense and oriented fiber arrangement in the periodontal ligament and fiber insertion into a new cementum matrix is observed more frequently (Aukhil et al., 1986a; Ettel et al., 1989; Caffesse et al., 1988, 1990, 1991; Pitaru et al., 1988; Gottlow et al., 1990). In contrast, new alveolar bone regenerates more easily and may restore interradicular septi entirely (Caffesse et al., 1990). However, that portion of the denuded root surface not covered by either epithelium or new cementum frequently and unpredictably undergoes resorption and ankylosis. On the other hand, if root resorption is followed by the production of new cementum matrix, the latter is intimately bound and irreversibly attached to that root surface (Aukhil et al., 1983, 1986a; Caffesse et al., 1990, 1991). In fact, Egelberg (1987) stated very clearly that “if initial surface resorption occurs, newly formed collagen fibrils could interdigitate . . . with collagen fibrils from the cementum or the

40

HUBERT E. SCHROEDER

REGENERATIVE CEMENTOGENESIS

49

dentrine matrix, being exposed by the resorptive process. . . . In its absence, the newly formed matrix splits away from the root surface.” Furthermore, along the intact periodontium bordering the denuded root defect, cells of the periodontal ligament react with a proliferative burst only within a very limited zone of 200 to 300 p m during the first 7 days after surgery and there is some evidence, only from experiments with the window technique, that such cells actually migrate toward the denuded root surface (Aukhil and Iglhaut, 1988; Iglhaut et a / . , 1988). There is no evidence, however, to indicate that cells derived from the alveolar bone participate in the regenerative process leading to new cementum matrix formation. But, when the influx of cells from intact portions of bone and the periodontal ligament is blocked by a barrier membrane placed apically around the root, regenerative processes fail to occur (Aukhil et a/., 1987).

D. Biological Problems After several decades of research, it seems clear at present that under both spontaneous and guided conditions, periodontal tissue regeneration, in particular regenerative cementogenesis, along established but formerly diseased and denatured root surfaces is not achieved in the true sense of the term. Even under the best possible, so-called guided conditions, unpredictable repair rather than regeneration (i.e., the true morphological and functional restoration of the periodontium) results from wound healing in that area, at least after short-term observations in the human. The newly formed tissue that in part develops along such root surfaces is a cellular, rapidly generated variety of either cementum or bone, and this tissue, with or without a dense collagen fiber insertion, is not functionally attached to the old, denatured root tissues, be it cementum or dentine. There is no indication that a fiber fringe matrix and AEFC are deposited initially or

FIG. 12 (a-c) Light micrographs depicting newly formed cellular cementum (NC) atop and along a thick layer of old acellular extrinsic fiber cementum (AEFC), with and without root resorption (a). Where resorption (R)took place. NC is firmly attached to the old cementum (a,b); without resorption, NC is simply superimposed and separates (arrows) from the old cementum due to preparatory shrinkage (a.c). The same is true at the apical termination, marked by an artificial notch (N), of the mechanically treated root surface (inset). There is no attachment or insertion of new connective tissue (CT) fibers in NC. [The material originated from human patients undergoing guided regeneration treatment (with a resorbable Vicryl membrane) of lower incisors, after 3 months of regenerative wound healing.] CC, old cellular cementum. Courtesy Dr. Urs Zappa, Bern. Magnifications: a, x 100; b,c, ~ 3 2 0 ; inset. x100.

50

HUBERT E. SCHROEDER

later in time. The reasons for this may be found in basic biological issues regarding cementogenesis. Apart from clinical problems mentioned above, the basic biological issues center around two questions: ( a )the repopulation of the blank root surface and its covering blood clot with cells appropriate for regenerating cementum and the periodontal ligament, and ( b )the functional attachment of the regenerated tissue to the denatured root surface. In his review, Minabe (1991) concluded that “at present, the attainment of a new functional periodontal ligament over extensive portions of treated root surfaces may not be realistic,” because “without a method for selective repopulation of the root surface, the support system of the cementum, periodontal ligament, and bone cannot be reliably renewed.” When teeth develop preeruptively, cells forming root cementum derive directly or indirectly from the dental follicle proper that envelops the growing root. Apparently the appropriate cells receive their differentiation signal in a time-dependent fashion, isochronous with root elongation in the apical direction. Various varieties of cementum are formed along a corono-apical gradient. On the other hand when periodontal defects with denatured root surfaces enter protected wound healing, the respective cells, supposedly deriving from still intact portions of the periodontal ligament and alveolar bone, have to follow an apico-coronal pathway, with the gradient of cementogenesis reversed. In this situation, the questions arise: how fast do these cell populations proliferate, are the appropriate cementogenic cells present or available through precursors, how fast and, how far are they able to migrate, and what are the signals to turn on their matrix-producing activity at a particular wound or defect locality? At present, these questions have no answers. Periodontal ligament cells are developmentally, topographically, and functionally different from gingival fibroblasts, and they represent heterogeneous mixtures of phenotypically or through environmental modulation different subsets of largely unknown ability and proportion. Although osteoblasts precursors might occur, it is unkown whether cementogenic cells or their precursors are part of that mixture or not. But even when present, such cells may not divide frequently, may be unable to respond to differentiation signals or to migrate extensively, or may decrease in number with age. This is particularly valid topographically, i.e., for the remaining, intact ligament tissue around the root apex or in the furcational region. In fact, it is not known whether the cells producing the new, cellular “cementum” derive from the ligament or the alveolar bone. However, the fact that root resorption and ankylosis are frequent and unpredictable side effects, particularly when epithelium is artificially excluded from the wound healing process, indicates that connective tissue and cells other than those with the appropriate cementogenic capacity have arrived first and proliferate along the root surface.

REGENERATIVE CEMENTOGENESIS

51

Whatever origin and composition the new cementum-like tissue may have, its matrix does not attach to the hard and denatured root surface. Clear evidence to the mere deposition of that matrix is the fact that during histologic processing, it splits away from the established root tissues. The reason is lack of matrix fibril interdigitation secured by mineralization as it occurs during natural cementogenesis. Believing that demineralized root surfaces held an induction potential for bone and cementum regeneration, Register (1973) and Register and Burdick (1975, 1976), employing hydrochloric, lactic, citric, phosphoric and other acids, initiated a series of studies showing that superficial demineralization of the denuded root surfaces facilitated and accelerated fiber reattachment (Holden and Smith, 1983). Later on, it was shown in the dog that citric acid demineralization to a depth of 3 to 5 pm indeed favors the interdigitation of the newly formed and acid-exposed collagen fibril populations (Ririe et al., 1980; Selvig, 1983),mimicking the first step of a spontaneous resorption process, i.e., demineralization of the crystals along the resorptive surface (Frank et al., 1974). A similar result, i.e., dense collagen fibril reattachment in superficially demineralized (about 1 to 2 pm deep) dentin, was observed along the apical portion of a natural defect in a 72-year-old woman (Heritier, 1983). Unfortunately, under routine clinical conditions and in further animal experiments root demineralization has met with controversial success.

VI. Concluding Remarks and Perspectives Although some first steps have been made in supplying urgently needed information, the genesis of the various types of cementum on growing human teeth, the conditions under which they increase in thickness, and the functional roles they might fulfill are still unknown. The task of deciphering all this is very complex because ( a ) the various types of cementum developing in selected areas, as a pure or mixed variety, simultaneously or in alternating layers, often behave unpredictably in time and space, ( 6 ) the full spectrum of all possible combinations is not yet known, and ( c )human teeth are not freely available and rarely subject to experimentation. Nevertheless, the need for further information is imperative. One aspect, however, appears secured, i.e., that a force resistant union between cementum and dentine is generated only via matrix fibril interdigitation and subsequent mineralization. The failure in establishing that kind of union is one of the basic problems in regenerative cementogenesis. Another is that the cells producing the various types of cementum are virtually unknown. For that reason, neither in vitro nor in vivo models for

52

HUBERT E. SCHROEDER

studying regenerative cementogenesis are particularly promising, unless much more is known about where these cells come from, when they do what and why, and where they eventually go. Animal models, such as dogs and non-human primates, if still available in the future, may project a response picture too optimistic to be used in analogy to aging humans. Very little is known about the animal's spontaneous cementogenesis, their particular types of root cementum, and their regenerative response potential in general. Future perspectives probably lie along the route of dynamic labeling studies, and the imperative task is to establish knowledge, for the various types of root cementum, about their developmental system characterized by particular classes of cells and their morphologically, biochemically, and functionally defined matrix products.

Acknowledgments The author is indebted to Dr. D. D. Bosshardt, Zurich, for his generosity in permitting the use of part of his unpublished material on developmental cementogenesis; to Dr. U. Zappa, Dental Clinics, University of Bern, for allowing photography and reproduction of part of his experimental material on guided tissue regeneration: to Mr. H. Maag, Department of Orthodontics, University of Zurich, for translating the author's raw sketches into representative drawings; to Mrs. Susi Munzel-Pedrazzoli for competent help in designing and producing the illustrative plates: and to Mrs. Rosmarie Kroni for handling the reference list and for typing and editing the manuscript.

References Abbas, D. K., Albandar, J. M., Helgeland, K., and Johansen, J. R. (1987). Acta Odontol. Scand. 45, 353-360. Abdallah. F., Kon, S., and Ruben, M. P. (1988). J . West. SOC. Periodontol. (Periodont. Abstr.) 36, 53-61. Andreasen, J. O., and Kristerson, L. (1981). fnt. J . Oral. Surg. 10, 189-201. Arceo, N., Sauk, J. J., Moehring, J., Foster, R. A,, and Somerman, M. J. (1991). J . Periodonrol. 62, 499-503. Aukhil. 1. (1991). Dent. Clin. North. Am. 35, 459-467. Aukhil, I . , and Fernyhough, W. S. (1986). J . Periodontol. 57, 405-412. Aukhil, I . . and Iglhaut, J. (1988). 1. Clin. Periodonrol. 15, 374-382. Aukhil, I . , Pettersson, E., and Suggs, C. (1986a). J. Periodontol. 57, 727-734. Aukhil, I . Simpson. D. M., Suggs. C., and Pettersson, E. (l986b). J . Clin. Periodontol. 13, 862-868. Aukhil, I., Pettersson, E., and Suggs, C. (1987). J . Periodonrol. 58, 71-77. Aukhil, I., Simpson, D. M., and Schaberg, T. V. (1983). J . Periodont. Res. 18, 643-654. Azaz, B., Michaeli, Y.,and Nitzan, D. (1977). Oral Surg. Oral Med. Oral Parhol. 43, 572-578. Azaz, B., Ulmansky, M., Moshev, R., and Sela, J. (1974). Oral Surg. 38, 691-694. Barrett, A. P., and Reade, P. C. (1981). J . Periodont. Res. 16, 456-465.

REGENERATIVE CEMENTOGENESIS

53

Bartold. P. M.. and Page. R. C. (1985). J . Periodont. Res. 20, 284-292. Beertsen, W., and Everts, V. (1990). J . Dent. Res. 69, 1669-1673. Beertsen, W., and Van Den BOS,T . (1991). J . Dent. Res. 70, 176-181. Bellows, C. G.. Melcher, A. H., and Aubin. J. E. (1981). J . Cell Sci. 50, 299-314. Bellows, C. G., Melcher, A. H., Bhargava, U., and Aubin. J. E. (l982a). J . Ultrastrucr. Res. 78, 178-192. Bellows, C. G., Melcher, A. H., and Aubin, J . E. (1982b). J . Cell Sci. 58, 125-138. Bellows, C. G., Melcher, A. H., and Brunette, D. M. (1980). J . Cell Sci. 44, 59-73. Bernstein, A. B., Preisig, E., and Schroeder, H. E. (1988). Cell Tissue Res. 254, 659-670. Bernstein. A. B., Preisig, E., and Schroeder, H . E. (1989a). Cell Tissue Res. 255,631-639. Bernstein, A. B., Preisig, E., Pajarola, G., and Schroeder, H. E. (1989b). Cell Tissue Res. 258, 125-135. Bernstein, A. B., Preisig. E., and Schroeder, H. E. (1990). Cell Tissue Res. 259, 603-606. Bhargava, U., Bar-Lev, M . , Bellows, C. G., and Aubin, J. E. (1988). Bone 9, 155-163. Birk, D. E., and Trelstad, R. L. (1984). J . Cell Biol. 99, 2024-2033. Birk, D. E., and Trelstad, R. L. (1985). Ann. N . Y . Acad. Sci. 460, 258-266. Birk, D. E., and Trelstad, R. L . (1986). J . CellBiol. 103, 231-240. Birk, D. E., Zycband, E. I., Winkelmann, D. A,. and Trelstad, R. L. (1990). Ann. N . Y . Acad. Sci. 580, 176-194. Blomlof. L., Lindskog. S. , and Hammarstrom, L. (1988). Swed. Dent. J . 12, 101-112. Blomlof, L., and Otteskog, P. (1981). Scand. J . Dent. Res. 89, 43-47. Bordin, S ., Kolb, W. P., and Page, R. C. (1983). J . fmmunol. 130, 1871-1875. Bordin, S . , and Page, R. C . (1988). In Vitro 24, 719-726. Bordin, S . , and Page, R. C. (1989). Mol. Immunol. 26, 677-685. Bordin. S . , Page, R. C.. and Narayanan, A. S. (1984). Science 223, 171-173. Bosshardt, D. D . . Luder, H. U., and Schroeder, H. E. (1989). J . Biol. Buccale 17, 3-13. Bosshardt, D. D., and Schroeder, H. E. (1990). J . Clin. Periodontol. 17, 663-668. Bosshardt, D. D., and Schroeder, H . E. (1991a). Cell Tissue Res. 263, 311-324. Bosshardt. D. D.. and Schroeder, H. E. (1991b). Cell Tissue Res. 263, 325-336. Bosshardt, D. D.. and Schroeder, H. E. (1992). Cell Tissue Res. 267, 321-335. Bowers, G. M., Chadroff, B., Carnevale, M., Mellonig, J . , Corio, R . , Emerson, J . , Stevens, M., and Romberg, E. (1989a). J . Periodontol. 60,664-674. Bowers, G. M., Chadroff, B., Carnevale, M., Mellonig, J., Corio. R.. Emerson, J., Stevens, M., and Romberg, E. (1989b). J . Periodontol. 60, 675-682. Bowers, G. M., Chadroff, B., Carnevale, M., Mellonig, J . , Corio, R., Emerson, J., Stevens, M., and Romberg, E. (1989~).J . Periodontol. 60, 683-693. Bowers, G. M., Granet, M., Stevens, M.. Emerson, J.. Corio, R., Mellonig, J . , Lewis, S. B.. Peltzman, B., Romberg, E., and Risom, L. (1985). J . Periodontol. 56, 381-395. Boyko, G. A., Brunette. D. M., and Melcher, A. H. (1980).J . Periodont. Res. 15,297-303. Boyko, G. A., Melcher, A. H., and Brunette, D. M. (1981).J . Periodont. Res. 16, 73-88. Butcher. E. O., and Vidair, R. V. (1955). J . Dent. Res. 34, 569-576. Caffesse, R. G., Dominguez, L. E., Nasjleti, C. E., Castelli, W. A., Morrison, E. C., and Smith, B. A. (1990). J . Periodontol. 61, 45-50. Caffesse. R. G . , Nasjleti, C. E., Anderson, G. B., Lopatin, D. E., Smith, B. A,, and Morrison, E. C. (1991). J . Periodontol. 62, 21-29. Caffesse, R. G . , Smith, B. A., Castelli, W. A,. and Nasjleti, C. E. (1988). J . Periodontol. 59,589-594. Caton, J. G.. and Kowalski, G. J . (1976). J . Periodontol. 47, 506-510. Caton, J . G., and Zander, H . A. (1975). J . Periodontol. 46, 71-77. Cho, M.-I., and Garant, P. R. (1981). Anat. Rec. 199, 309-320. Cho, M.-I., and Garant, P. R. (1985). J . Periodont. Res. 20, 185-200.

54

HUBERT E. SCHROEDER

Cho, M.-I., and Garant, P. R. (1988). J . Periodont. Res. 23, 268-276. Cho, M.-I., and Garant, P. R. (1989). Anat. Rec. 223, 209-222. Cho, M.-I., Garant, P. R., and Lee, Y. L. (1988). J . Periodont. Res. 23, 287-294. Chomette, G., Auriol, M., Armbruster, D., Szpirglas, H., and Vaillant, J. M. (1987). J. Biol. Buccale 15, 217-224. Christner, P., Robinson, P., and Clark, C. C. (1977). Calcif. Tissue Res. 23, 147-150. Claffey, N., Hahn, R., and Egelberg, J. (1989). J. Clin.Periodontol. 16, 371-379. Cole, R. T., Crigger, M., Bogle, G., Egelberg, J., and Selvig, K. A. (1980). J. Periodont. Res. 15, 1-9. Connor, N. S., Aubin, J. E., and Sodek, J. (1983). J . Cell Sci. 63,233-244. Crigger, M., Bogle, G., NilvCus, R., Egelberg, J., and Selvig, K. A. (1978). J. Periodont. Res. 13, 538-549. Dastmalchi, R., Polson, A., Bouwsma, O., and Proskin, H. (1990). J . Clin.Periodontol. 17, 709-7 13. Davidovitch, Z . , Finkelson, M. D., Steigman, S., Shanfeld, J. L., Montgomery, P. C., and Korostoff, E. (1980). Am. J. Orthod. 77, 33-47. Diab, M. A., and Stallard, R. E. (1965). Periodontics 3, 10-14. Ecarot-Charrier, B., Glorieux, F. H., van der Rest, M., and Pereira, G. (1983). J. Cell Biol. 96,639-643. Ecarot-Chanier, B., Shepard, M., Charette, G., Grynpas, M., and Glorieux, F. H. (1988). Bone 9, 147-154. Egelberg, J. (1987). J. Periodont. Res. 22, 233-242. Eley, B. M., and Harrison. J. D. (1975). J. Periodont. Res. 10, 168-170. Ellegaard, B., Karring, T., Listgarten. M., and Loe, H. (1973). J. Periodontol. 44,209-217. Engel, D., Schroeder, H. E., Gay, R., and Clagett, J. (1980). Arch. OralBiol. 25, 283-296. Enwemeka, C. S. (1991). Tissue Cell 23, 173-190. Ettel, R. G., Schaffer, E. M., Holpuch, R. C., and Bandt, C. L. (1989). J . Periodontol. 60, 342-351. Fardal, O., and Lowenberg, B. F. (1990). J . Periodontol. 61, 529-535. Fernyhough, W., Aukhil, J., and Link, T. (1987). J. Periodontol. 58, 762-769. Fernyhough, W., and Page, R. C. (1983). J . Periodontol. 54, 133-140. Fleischmajer, R., Perlich, J. S., Timpl, R., and Olsen, B. R. (1988). J. Histochem. Cytochem. 36, 1425-1432. Formicola, A. J., Krampf, J. I., and Witte, E. T. (1971). J . Periodontol. 42, 766-773. Frank, R. M., Fellinger, E., and Steuer, P. (1976). J . Biol. Buccale 4, 295-313. Frank, R. M., Fiore-Donno, G., Cimasoni, G., and Matter, J. (1974). J. Periodontol. 45, 626-635. Frank, R. M., and Frank, P. (1969). Z . Zellforsch 99, 121-133. Furseth, R. (1967). Acta Odontol. Scand. 25, 613-645. Furseth, R. (1969). Archs Oral. Biol. 14, 1147-1 158. Garant, P. R. (1976). J . Periodontol. 47, 380-390. Garant, P. R., and Cho, M.4. (1979). J. Periodont. Res. 14, 95-106. Gerstenfeld, L. C., Chiprnan, S. D., Kelly, C. M., Hodgens, K. J., Lee, D. D., and Landis, W. J. (1988). J. Cell. Biol. 106, 979-989. Giniger, M. S., Norton, L., Sousa, S . , Lorenzo, J. A., and Bronner, F. (1991). J. Dent. Res. 70, 99-101. Gottlow, J., Karring, T., and Nyman, S. (1990). J. Periodontol. 61, 680-685. Gottlow, J., Nyman, S., Karring, T., and Lindhe, J. (1984). J . Clin. Periodontol. 11,494-503. Gottlow, J., Nyman, S., Lindher, J., Karring, T., and Wennstrom, J. (1986). J. Clin.Periodontol. 13, 604-616. Could, T. R. L. (1983). J. Dent. Res. 62, 873-876.

REGENERATIVE CEMENTOGENESIS

55

Gould, T. R. L., Melcher. A. H., and Brunette, D. M. (1977). Anat. Rec. 188, 133-142. Gould. T. R. L., Melcher, A. H., and Brunette, D. M. (1980).J. Periodont. Res. 15, 20-42. Hassel, T. M., and Stanek. E. J. (1983). Arch. Oral Biol. 28, 617-625. Held, A.-J. (1951). Oral Surg. Oral Med. Oral Pathol. 4, 53-67. Heritier, M. (1983). J. Periodontol. 54, 515-521. Holden, M. J., and Smith, B. A. (1983). J . West. Soc. Periodontol. (Periodont. Abstr.) 31, 45-56. Houston, F., Sarhed, G . , Nyman, S., Lindhe, J., and Karring, T. (1985).J . Clin. Periodontol. U ,716-727. Hurst, R. V. V. (1972). J . Dent. Res. (Suppl.) 51, 1183-1192. Hurum, S., Sodek, J., and Aubin, J. E. (1982). Biochem. Eiophys. Res. Commun. 107, 357-366. Iglhaut, J., Aukhil, I., Simpson, D. M., Johnston, M. C., and Koch, G. (1988). J . Periodont. Res. 23, 107-1 17. Jansen, M. T., Coppes, L., and Verdenius, H. H. W. (1955). J . Periodontol. 26, 292-300. Johansson, 0.. NilvCus, R., and Egelberg, J. (1978). J . Periodont. Res. 13, 525-531. Jones, S.J. (1981). In “Dental Anatomy and Embryology” ( J . W. Osborn, ed.), pp. 193-205, 286, 294. Blackwell Scientific. Oxford. Karring, T., Nyrnan, S., and Lindhe, J. (1980). J. Clin. Periodontol. 7, 96-105. Karring, T., Nyrnan, S.. Lindhe, J., and Siriat, M. (1984). J. Clin. Periodontol. 11, 41-52. Kawase, T., Sato, M., Yarnada, M., Hirayama, A., Miake, K., and Saito, S. (1986). Bone Mineral Res. Suppl. 1,63. Kenney, E. B., and Ramfiord, S. P. (1969). J . Dent. Res. 48, 114-119. Knox, B., and Aukhil, I. (1988). J. Periodont. Res. 23, 60-67. KO, D. S . , Page, R. C., and Narayanan, A. S. (1977). Proc. Natl. Acad. Sci. U . S . A . 8, 3429-3432. Korotzer, T. I., Clagett, J . A., Kolb, W. P., and Page, R. C. (1980). J . Cell Physiol. 105, 503-512. Leblond, C. P. (1989). Anat. Rec. 224, 123-138. Lester, K. S. (1969). J. Ultrastruct. Res. 27, 63-87. Lester, K. S., and Boyde, A. (1970). J . Ultrastruct. Res. 33, 80-94. Limeback, H. F., Sodek, J., and Aubin, J. E. (1983). J. Periodont. Res. 18, 242-248. Limeback, H. F., Sodek, J., and Brunette, D. M. (1978). Eiochem. J . 170, 63-71. Lindhe, J.. Nyman, S., and Karring, T. (1984). J. Clin. Periodontol. 11, 33-40. Listgarten, M. A. (1973). J . Periodonf. Res. 8, 335-345. Listgarten, M. A., and Rosenberg, M. M. (1979). J. Periodontol. 50, 333-344. Litwin, J., Lundquist, G., and Soder, P.O. (1971). Scand. J. Dent. Res. 79, 536-539. Lowenberg, B. F., Aubin, J. E., Deporter, D. A,, Sodek, J., and Melcher, A. H. (1985). J . Dent. Res. 64, 1106-1 110. Lowenberg, B. F., Pilliar, R. M., Aubin, J. E., Fernie, G. R., and Melcher, A. H. (1987). J . Dent. Res. 66, 1000-1005. Lowenberg, 9. F., Thibault, J., Lawrence, C., and Sodek. J. (1986). J . Dent. Res. 65, 1010-10 15.

Magnusson, I., Batich, C.. and Collins, B. R. (1988). J. Periodontol. 59, 1-6. Magnusson, I., Nyman, S., Karring, T., and Egelberg, J. (1985). J . Periodont. Res. 20, 20 I -208. Maniatopoulos, C., Sodek, J., and Melcher, A. H. (1988). Cell Tissue Res. 254, 317-330. Marchi, F., and Leblond, C. P. (1983). Am. J. Anat. 168, 167-197. Marchi, F., and Leblond, C. P. (1984). J. Cell Eiol. 98, 1705-1709. Marks, S. C., and Popoff, S. N. (1988). A m . J. Anat. 183, 1-44. McAllister, B., Narayanan, A. S., Miki, Y., and Page, R. C. (1990). J . Periodont. Res. 25, 99- 105.

56

HUBERT E. SCHROEDER

McBride, D. J., Hahn. R. A,, and Silver, F. H. (1985). I n t . J. Biol. Mncromol. 7, 71-76. McCulloch, C. A. G. (1985). Anat. Rec. 211, 258-262. McCulloch, C. A. G., and Bordin. S. (1991). J . Periodont. Res. 26, 144-154. McCulloch, C. A. G., and Knowles, G. (1991). Cell Tissue Res. 264, 87-94. McCulloch, C. A. G., and Melcher, A. H. (1983a). J . Periodont. Res. 18, 339-352. McCulloch, C. A. G., and Melcher. A. H. (1983b). Am. J . Anat. 167, 43-58. McCulloch, C. A. G . , Nemeth, E., Lowenberg, G., and Melcher, A. H . (1987). Anat. Rec. 219, 233-242. Melcher, A. H., and Chan, J. (1981). J . Ultrastruct. Res. 77, 1-36. Melcher, A. H., Cheong, T., Cox, J., Nemeth, E., and Shiga. A. (1986). J . Periodont. Res. 21, 592-612. Melcher, A. H., McCulloch, C. A. G . , Cheong, T.. Nemeth, E., and Shiga, A. (1987). J . Periodont. Res. 22, 246-247. Messadi, D. V . , and Bertolami, C. N. (1991). Dent. Clin. North. A m . 35, 443-457. Miki, Y., Narayanan, A. S., and Page, R. C. (1987). J. Dent. Res. 66, 1399-1403. Minabe, M. (1991). J . Periodonr. Res. 62, 171-179. Nakae, H. Ikrayanan, A. S . , and Page, R. C. (1989).J . Dent. Res. 68,365, Abstr. No. 1439. Narayanan, A. S . , and Page, R. C. (1983). Collagen Re/. Res. 3, 33-64. Nefussi. J. R., Sautier, J . M.. Nicolas. V., and Forest, N. (1991). J . Biol. Buccale 19, 75-82. Nijweide. P. J., Burger, E. H., and Feyen. J . H. M. (1986). Physiol. Rev. 66, 855-886. Nishimura, K., Hayashi, M., Matsuda, K., Shigeyama, Y.. Yamasaki. A., and Yamaoka, A. (1989). J . Periodonr. Res. 24, 146-148. Nojima, N., Kobayashi, M., Shiomone, M., Takahashi. N.. Suda. T., and Hasegawa, K . (1990). J . Periodont. Res. 25, 179-185. Nyman, S ., Gottlow, J., Lindhe. J., Karring. T., and Wennstrom. J . (1987). J . Periodont. Res. 22, 252-254. Nyman, S . , Karring, T., Lindhe, J., and Planten, S . (1980). J. Clin. Periodontol. 7,394-401. Nyman, S., Lindhe, J., and Karring, T. (1989). I n “Textbook of Clinical Periodontology” ( J . Lindhe, ed.), 2nd edition, pp. 450-476. Munksgaard, Copenhagen. Nyman, S., Gottlow, J., Karring, T., and Lindhe, J. (1982a).J. Clin. Periodontol. 9,257-265. Nyman, S ., Lindhe, J . , Karring, T., and Rylander, H. (1982b). J. Clin. Periodontol. 9, 290-296. Ogura, N. Mera, T., Sato, F., and Ishikawa, I. (1991). J . Periodontol. 62, 284-291. Oikarinen, K. S ., and Seppa, S . T. (1987). Endod. Dent. Traumatol. 3, 95-99. Olson, S., Arzate, H., Narayanan, A. S. , and Page. R. C. (1991). J . Denf. Res. 70, 12721277. Oshima, M., Kuwata, F.. Otsuka, K., and Suzuki, K. (1988). I n “Recent Advances in Clinical Periodontology” ( J . Ishikawa, H. Kawasaki, K. Ikeda. and K. Hasegawa, eds.), pp. 573-577. Excerpta Medica, Amsterdam. Owens, P. D. A. (1980). Arch. Oral B i d . 24, 901-907. Palmer, R. M., and Lumsden, A. G. S . (1987). Arch. Oral B i d . 32, 281-289. Paynter, K. J.. and Pudy, G. (1958). Anat. Rec. 131, 233-251. Pettersson, E. C., and Aukhil, I . (1986). J. Periodont. Res. 21, 543-552. Piche, J. E., Carnes, D. E., and Graves, D. T. (1989). J . Dent. Res. 68, 761-767. Pitaru, S . , and Melcher, A. H. (1983). J . Periodont. Res. 18, 483-500. Pitaru, S . . and Melcher, A. H. (1987). J . Periodont. Res. 22, 6-13. Pitaru. S ., Gray. A., Aubin, J. E., and Melcher, A. H . (1984a). J . Periodont. Res. 19, 408-41 8. Pitaru, S ., Aubin. J. E.. Gray, A., Metzger, Z . , and Melcher, A. (1984b).J . Periodont. Res. 19. 66 1-665.

REGENERATIVE CEMENTOGENESIS

57

Pitaru, S.. Tal, H., Soldinger. M., Azar-Avidan, 0..and Noff, M. (1987). J . Periodont. Res. 22, 331-333. Pitaru, S . , Tal. H., Soldinger, M.. Grosskopf, A.. and Noff, M. (1988). J . Periodontol. 59, 380-386. Pitaru, S.. Tal, H., Soldinger, M., and Noff, M. (1989). J . Periodont. Res. 24, 247-253. Polson. A. M. (1986). J . Clin. Periodonrol. 13, 995-999. Polson, A. M. (1987). Endod. Dent. Traumatol. 3, 45-57. Polson, A. M.. and Caton, J. (1982). J . Periodontol. 53, 617-626. Polson. A. M., and Proye. M. P. (1982). J . Clin. Periodontol. 9, 441-454. Polson, A. M.. and Proye, M. P. (1983). J . Periodontol. 54, 141-147. Pontoriero, R.. Lindhe, J.. Nyman, S. , Karring, T., Rosenberg, E., and Sanavi, F. (1988). J . Clin. Periodonrol. 15, 247-254. Pontoriero, R., Lindhe, J., Nyman. S . , Karring. T., Rosenberg, E.. and Sanavi, F. (1989). J . Clin. Periodontol. 16, 170-174. Pontoriero, R., Nyman, S. , Lindhe, J., Rosenberg, E., and Sanavi. F. (1987). J . Clin. Periodontol. 14, 618-620. Preisig, E., and Schroeder, H. E. (1988). J . Periodont. Res. 23, 211-221. Proye, M., and Polson, A. M., (1982). J . Periodontol. 53, 379-389. Quamstrom, E. E., and Page, R. C. (1986). J . Cell Sci. 84, 183-200. Ragnarsson, B.. Carr, G.. and Daniel, J. C. (1985). J . Dent. Res. 64, 1026-1030. Register. A. A. (1973). J . Periodontol. 44, 49-54. Register, A. A.. and Burdick, F. A. (1975). J . Periodontol. 46, 646-655. Register, A. A., and Burdick, F. A. (1976). J . Periodontol. 47, 497-505. Ririe, C. M., Crigger, M., and Selvig. K. A. (1980). J . Periodont. Res. 15, 314-327. Roberts, W. E. (1975a). Arch. Oral Biol. 20, 465-471. Roberts, W. E. (l975b). Am. J . Anal. 143, 363-370. Roberts, W. E., Mozsary. P. G., and Klingler, E. (1982). A m . J . Anat. 165, 373-384. Rose, G. G., Yajima, T., and Mahan. C. J. (1980). J . Periodont. Res. 15, 53-70. Rose, G. G.. Yamasaki, A,, Pinero, G. J., and Mahan, C. J. (1987). J . Periodont. Res. 22, 20-28. Russel, S. B., Russel, J. D., and Trupin. K. M. (1981). J . Cell. Physiol. 109, 121-131. Schellens, J. P. M.. Everts, F., and Beertsen, W. (1982). J . Periodont. Res. 17, 407-422. Schroeder. H. E. (1986). “The Periodontiurn,” Vol. V/5. “Handbook of Microscopic Anatomy.” Springer-Verlag. Berlin. Schroeder, H. E. (1987). “Orale Strukturbiologie,” pp. 28-33. Georg Thieme Verlag, Stuttgart. Schroeder, H. E . (1988). In “Periodontology Today” (B. Guggenheim, ed.), pp. 32-40. Karger, Basel. Schroeder. H. E. (1991a). “Pathobiologie oraler Strukturen,” 2, “uberarbeitete und erweiterte Aufl,” pp. 205-214. Karger, Basel. Schroeder. H. E. (1991b). Schweiz. Monatsschr. Zahnmed. 101, 279-284. Schroeder, H. E., Luder, H . U., and Bosshardt, D. D. (1992). Schweiz. Monatsschr. Zahnmed. 102, 20-31. Schuhmacher, G. H . , and Schmidt, H. (1983). “Anatomie und Biochemie der Zahne.” 3rd edition, pp. 438. Fischer. Stuttgart, New York. Selvig, K. A. (1963). Acta Odontol. Scand. 21, 175-186. Selvig, K. A. (1964). Acta Odontol Scand. 22, 105-120. Selvig, K . A. (1965). Acta Odontol Scand. 23, 423-441. Selvig, K. A. (1967). “Studies on the Genesis, Composition and Fine Structure of Cementum.” Theses. University of Bergen, Norway. Selvig, K. A., (1983). J . B i d . Buccule 11, 79-94.

58

HUBERT E. SCHROEDER

Selvig, K. A., Bogle, G., and Claffey, N. M. (1988). J . Periodontol. 59, 758-768. Selvig, K. A., N i l v h s , R. E., Fitzmorris, L., Kersten, B., and Khorsandi, S. S. (1990). J . Periodontol. 61, 515-520. Sequeiria, P., Bosshardt, D. D., and Schroeder, H. E. (1992). J . Periodontol. Res. 27, 134- 142. Skillen, W. G., and Lundquist, G. R. (1937). J. Am. Dent. Assoc. 24, 175-185. Smith, A. J., Leaver, A. G., and Smith, G. (1983). Arch. Oral Biol. 28, 1047-1054. Sodek, J. (1976). Biochem. J . 160, 243-246. Sodek, J. (1977). Arch. Oral Biol. 22, 655-665. Somerman, M. J., Archer, S. Y., Hassel, T. M., Shteyer, A., and Foster, R . A. (1987a). Arch. Oral Biol. 32, 879-883. Somerman, M. J., Perez-Mera, M., Merkhofer, R. M., and Foster, R. A. (1987b).J . Periodontol. 58, 349-351. Somerman, M. J., Archer, S. Y., Shteyer, A., and Foster, R. A. (1987~). J . Periodont. Res. 22, 75-77. Somerman, M. J., Prince, C. W., Sauk, J. J., Foster, R. A., and Butler, W. T. (1987d). J . Bone Mineral Res. 2, 259-265. Somerman, M. J., Archer, S. Y., Imm,G. R., and Foster, R . A. (1988). J . Dent. Res. 67, 66-70. Somerman, M. J., Foster, R. A., Imm, G. M., Sauk, J. J., and Archer, S. Y. (1989). J . Periodontol. 60,73-77. Somerman, M. J., Sauk, 1. J., Foster, R. A.. Dickerson, K., Norris, K., and Argraves, W. S. (1991). J . Periodont. Res. 26, 10-16. Somerman, M. J., Shroff, B., Argraves, W. S.. Morrison, G., Craig, A. M., Denhardt, D. T., Foster, R. A., and Sauk, J. J. (1990). J. Biol. Buccale 18, 207-214. Stahl, S. S., and Froum, S. J. (1991a). J . Clin. Periodontol. 18, 69-74. Stahl, S . S., and Froum, S. J. (1991b). J . Clin. Periodontol. 18, 685-689. Stahl, S. S., Froum, S., and Tarnow, D. (1990). J . Clin. Periodontol. 17, 191-198. Stones, H. H. (1934). Proc. R . Soc. Med. 27, 728-744. Sudo, H., Kodama, H.-A., Amagai, Y.. Yamamoto, S . , and Kasai, S. (1983). J . Cell Biol. 96, 191-198. Ten Cate, A. R. (1972). J. Anat. ll2,401-414. Ten Cate, A. R., and Deporter, D. A. (1974). Arch. Oral Biol. 19, 339-340. Ten Cate, A. R., Deporter, D. A., and Freemann, E. (1976). Am. J . Orthod. 69, 155- 168. Ten Cate, A. R., Mills, C., and Solomon, G. (1971). Anat. Rec. 170, 365-380. Terranova, V. P., Franzetti, L. C., Hic. S., DiFlorio, R. M., Lyall, R. M., Wikesjo, U. M. E., Baker, P. J., Christersson, L. A., and Genco, R. J. (1986). J . Periodontol. Res. 21,330-337. Terranova, V. P., Hic, S., Franzetti, L., Lyall, R. M., and Wikesjo, U. M. E. (1987). J . Periodontol. 58, 247-257. Terranova, V. P., Jendresen, M., and Young, F. (1989a). Adu. Dent. Res. 3, 69-79. Terranova, V. P., Odziemiec, C., Tweden, K. S., and Spadone, D. P. (1989b).J . Periodontol. 60,293-301. Weinstock, M . (1975). I n “Extracellular Matrix Influences on Gene Expression” (H. C. Slavkin and R. C. Greulich, eds.) pp. 119-128. Academic Press, New York. Weinstock, M., and Leblond, C. P. (1974). J . Cell Biol. 60,92-127. Williams, D. C., Boder, G. B., Toomey, R. E., Paul, D. C., Hillman, C. C., King, K. L., Van Frank, R. M., and Johnston, C. C.. Jr. (1980). Calcif. Tissue Int. 30, 233-246. Yajima, T. (1988). Histochemistry 90, 245-253. Yajima, T., Rose, G. G., and Mahan, C. J. (1980). J . Periodont. Res. 15, 267-287.

REGENERATIVE CEMENTOGENESIS

Yamamoto, T. (1986). Arch. Hisrol. Jpn. 49, 459-481. Yamamoto, T., and Wakita, M. (1990). J . Periodonr. Res. 25, 113-119. Yamamoto, T., and Wakita, M. (1991). J . Periodonr. Res. 26, 129-137. Yamamoto, T., and Wakita, M. (1992). J . Periodonr. Res. 27, 20-27. Yoshikawa, D. K . , and Kollar, E. J. (1981). Arch. Oral Biol. 26, 303-307. Zahnder, H. A . , and Hurzeler, B. (1958). J . Dent. Res. 37, 1035-1044. Zappa, U . (1991a). Schweiz. Monatsschr. Zahnmed. 101, 1147-1 151. Zappa, U. (1991b). Schweiz. Monatsschr. Zahnmed. 101, 1321-1324.

59

This Page Intentionally Left Blank

lmmunocytochemical Localization of Proteins in Striated Muscle Marvin H. Stromer Muscle Biology Group, Department of Animal Science, Iowa State University, Ames, Iowa 5001 1

1. Introduction

Localization of specific muscle proteins via immunofluorescence and with immunoelectron microscopy are both well-established approaches. The availability of monoclonal antibodies (mAb) now has made it feasible to locate specific epitopes, to probe functional sites on proteins, and to locate structural domains and their arrangement in s h . Although many valuable contributions have been made to our understanding of striated muscle by laboratories around the world, the papers cited in Table I are especially noteworthy for their enhancement of our knowledge about striated muscle. The proteins are listed in the same order in which they appear in this review. This review provides an overview of the results obtained by using antibodies to understand striated muscle better. Emphasis has been on the most recent papers, but when needed for purposes of perspective, a limited number of earlier papers have been included. Results from some studies where antibodies have been used to probe functional sites or domains on proteins have been cited, as have some recent reviews that provide important background information. Reviews by Groschel-Stewart (1980), Pepe (1983), and Pette and Staron (1990) provide valuable information on topics related to this chapter. Because of the breadth of the topic, the very large number of papers that deal with various aspects of this topic, and length limitation, it was impossible to include all papers on this subject. I apologize in advance to anyone who feels that a favorite paper of theirs should have been included. Such omissions were unintentional. /~ll~~r~lulK ~ PoUm i ClM/ 1Jf C?lo/il~?,VlJI. 142

61

Copyright 0 1992 by Academic Press. Inc. All right5 of reproduction in any form reserved.

62

MARVIN H. STROMER

TABLE I Examples of lmmunocytochemical Studies That Have Made Major Contributions to Our Understanding of Striated Muscle

Protein Myosin

Reference

Contribution

Pepe (1967b)

Provided the basis for thick-filament model and understanding how myosin molecules are arranged in thick filaments. Located C-protein isoforms in the A band of C-protein Dennis et a/. (1984) different muscle types. Troponin Ohtsuki (1975) Located the three components of the troponin complex along the thin filament. Described the behavior of nebulin filaments Nebulin Wang and Wright (1988) anchored at the Z line during muscle shortening. Demonstrated that titin filaments extend from Titin Fiirst et a/. (1988) the Z line to near the M line. Described the behavior of titin epitopes in the Maruyama et a/. (1989) A and I bands during muscle shortening and stretch. Desmin Granger and Lazarides (1978) Demonstrated that desmin IFs exist at the periphery of Z lines. Demonstrated that desmin IFs span Z line to Z Richardson et a / . (1981) line between adjacent myofibrils.

11. Localization of Proteins in Skeletal Muscle Cells A. Thick-Filament Proteins and A-Band-Associated Proteins

The early recognition that muscle contraction involved interactions between thick and thin filaments focused attention on the need to know which proteins were present in the A band and how they were arranged. The complexity of this task increased as additional proteins associated with thick filaments were identified. Current emphasis is on the relationship between the specific location of a protein and/or its isoforms and the functional properties of thick filaments. The protein that has received the most attention is myosin.

1. Myosin a . Probing the Structure of the A Band Pepe (1966; 1967a,b; 1968) was among the early workers to use fluorescein-conjugated myosin antibodies for localization with the fluorescence microscope and unlabeled myosin antibodies for electron microscope (EM) localization. Pepe’s success in

PROTEINS IN STRIATED MUSCLE

63

isolating antibodies to specific regions of the myosin molecule by absorbing myosin antibodies with tryptic fragments of myosin permitted him to demonstrate that antigenic sites on thick filaments were accessible to these antibodies only in certain portions of the A band. For example, myosin antibodies absorbed with the heavy meromyosin (HMM) tryptic fragment of the myosin molecule would provide antibodies that would only recognize the light meromyosin (LMM) antigenic sites. Absorption with LMM from a 60-min trypsin digest of myosin would produce an antibody fraction that would recognize both the HMM and the trypsin-sensitive sites. Absorption with LMM from a 15-min trypsin digest would yield an antibody fraction that recognizes only HMM because short-digest LMM contains both the LMM and the trypsin-sensitive sites. Antibodies specific for LMM bound in the outer thirds of each half of the A band (Pepe, 1971). Antibodies specific for the trypsin-sensitive region were localized in seven stripes in the middle third of each half of the A band. HMM-specific antibodies bound to sites in the inner third of each half of the A band only if sarcomere lengths were 2.5 pm or greater so that no actin filaments overlapped with this region of the A band. Reduction or abolition of antiHMM staining at shorter sarcomere lengths indicated that the actin-HMM interaction blocked the epitope required for anti-HMM binding. These myosin antibody staining patterns were an important part of the information used by Pepe (1967a, 1971) to propose a model for how myosin molecules were packed into thick filaments. In a subsequent study, Wachsberger et al. (1983) used a mAb specific for an epitope on the S1 fragment of skeletal myosin that was not related to the ATPase site. When myofibrils in relaxing medium were exposed to this antibody, there was binding along the entire length of the cross-bridge region in the A band. In the absence of relaxing medium, the antibody bound to narrow regions on each side of the bare zone and at the outer edges of the A band. Because this antibody binding pattern was not affected by linkage of myosin crossbridges to actin filaments, it was suggested that differences in molecular packing along the thick filament could cause differences in availability of the antigenic site. A rnAb that recognized an epitope 92 ? 5 nm from the C-terminal end of the LMM or tail fragment of myosin was used by Shimizu et al. (1985a) to map this epitope in the A band and in isolated thick filaments. In contradiction to the earlier results of Pepe (1967b) and of Lowey and Steiner (1972), who observed a nonuniform fluorescent staining of the A band with antibody to LMM, this antibody stained along almost the entire length of the A band. Thin sections of labeled muscle bundles contained 50 stripes spaced 14.3 nm apart in each half of the A band. Isolated thick filaments decorated with this LMM antibody had stripes of variable density that repeated every 14.6 nm along the filament. This pattern was interpre-

64

MARVIN H. STROMER

ted to mean there was an uninterrupted axial spacing of myosin rods in thick filaments and as support for Huxley’s (1963) proposal for how myosin molecules are packed in thick filaments.

b. Mapping Functional Sites on Myosin Molecules The vertebrate skeletal myosin molecule consists of two 223-kDa heavy chains (Strehler et al., 1986) and four light chains (LCs). Each molecule contains two 19-kDa LC2 or 5,5-dithiobis-(2-nitrobenzoate)(DTNB) light chains and combinations of two different alkali light chains, a 20-kDa LCl or A1 chain and/ or a 16-kDa LC3 or A2 chain. Each of the two heads of the myosin molecule contains 1 of LC2 and 1 of either LCl or LC3. The original nomenclature of the alkali light chains (A1 and A2) and the DTNB light chain was based on the method used to isolate them. Now the three types are usually referred to as LC 1, LC2, and LC3 in decreasing order of chain weight. Because of the complexity of both the molecule itself and the functions of myosin, including, among others, ATP hydrolysis, conformation changes, and calcium binding, there has been interest in locating sites on myosin that would affect either the function and/or the structure of the molecule. Shimizu et al. (1985b) found that a mAb that recognized LC2 in fast skeletal muscle would only stain myofibrils after incubation in EDTA or if the myofibrils were denatured by acetone or ethanol. When calcium or magnesium occupies the metal binding site on LC2, there may be a conformational change that inhibits antibody binding. Removal of the metal with EDTA or denaturing LC2 with organic solvents reverses the conformational change and the antibody can bind. Electron microscopy of rotary-shadowed myosin-antibody complexes shows that the antibody binding site and the calcium binding site are located near the head-rod junction of myosin. The question of the spatial location of the N-terminus of myosin heavy chains (MHC) in the head of myosin molecules was investigated by Sutoh et al. (1987) by using an antibody to the N-terminal eight residues. Rotaryshadowed HMM-antibody complexes showed that the antibody was located 12 nm from the head-rod junction and indicated that the N-terminus of the MHC is located in the middle region of the head. Miyanishi et al. (1988) produced antibodies to two of the three tryptic fragments (23, 50, and 20 kDa) of the S1 portion of HMM. Rotary shadowing of the myosin-antibody complexes showed that one-third of the myosin molecules had antibodies to 50-kDa fragments bound at the tip of the head and two-thirds bound 16 nm from the head-rod junction on the side of the head. Antibodies and Fab fragments of antibodies to the 23-kDa fragments both bound 14-18 nm from the head-rod junction at the middle of the head. Dan-Goor et al. (1990) also used trypsin to cleave S1 into 27-, 50-,

PROTEINS IN STRIATED MUSCLE

65

and 20-kDa fragments aligned in order from the N-terminus. One mAb to the 27-kDa fragments reacted with an epitope in the N-terminal23 residues of the 27-kDa N-terminal fragment. A second mAb to the 50-kDafragments reacted with a 3-kDa N-terminal region of the 50-kDa fragment. The 27and the SO-kDa epitope were 14 and 17 nm, respectively, from the head-rod junction of myosin. Binding SI to actin decreased the affinity of antibodies to 50-kDa fragments for S1 but did not affect the affinity of antibodies to 27-kDa fragments for S 1. Antibodies to 50-kDa fragments inhibited the K+ (EDTA) and the actin-activated ATPase activity of SI, but antibodies to 27-kDa fragments had no effect. This suggests that either the 50-kDa epitope is near the actin or ATP binding sites or there is some form of “communication” between these sites and the antibody binding site. The location of the alkali light chains (LCl and LC3) was determined by using antibodies specific for the N-terminal region (Waller and Lowey, 1985). These light chains are located in the “neck” region of the myosin head, very close to the LC2 chain. Tokunaga et al. (1987) used mAbs to determine that the N-terminus of LC2 is located at the head-rod junction and that the N-terminus of LCI is 11 nm from the head-rod junction. This proximity between LC2 and the alkali light chains may explain why LC2 affects interactions between alkali light chains and MHC. Additional detailed information on the location of both the alkali light chains and LC2 was provided by labeling cysteine residues with the thiol-specific reagent 5-(iodoacetamido) fluorescein (Katoh and Lowey, 1989). Each of the two alkali LCs has a single cysteine near the C-terminus, and LC2 was labeled at either cysteine 125 or cysteine 154. Fluorescyl antibodies were then added to the labeled myosin, and the complexes were rotary-shadowed. The cysteine of the alkali light chains was 9 nm closer to the tip of the myosin head than the cysteine of LC2, which was close to the head-rod junction. This indicates that the two types of light chains are not arranged colinearly in the myosin head. The effect of antibodies to the S2 subfragment of myosin on the contractile behavior of myofibrils was examined by Harrington et al. (1990). Antibody-treated myofibrils contracted at an initial rate similar to that of untreated myofibrils. This was followed, in antibody-treated myofibrils, by a much slower contraction that ended with I-pm sarcomere spacing still visible. Untreated myofibrils contracted into nonstriated globular structures. Skinned muscle fibers were able to develop much less isometric force in the presence of antibodies to S2. These results may indicate that the S2 region of the myosin molecule also may be involved in force production in working muscle. Edman er al. (1988) used two MHC mAbs in conjunction with studies on unloaded shortening velocity and myofibrillar

66

MARVIN H. STROMER

ATPase activity and concluded that variability in shortening velocity and myofibrillar ATPase activity that exists among frog twitch fibers is based on differences in MHC composition. c. Location of MHC

mRNA The approach most recently used to locate MHC mRNA is in situ hybridization with a biotinated cDNA probe. An antibody to biotin is then added, followed by a second antibody that is specific for the species in which the biotin antibody was produced and that is complexed with a label (e.g., colloidal gold). Pomeroy et al. (1991)found that MHC mRNA was localized mainly in the peripheral regions of 14-day chick pectoral muscle cells where developing myofibrils are abundant. Some MHC mRNA was also observed near the nucleus. Most MHC mRNA was associated with 4- or 10-nm-diameter filaments believed to be actin and intermediate filaments, respectively. Although MHC mRNA is concentrated near assembling myofibrils, most mRNA is 0.1-0.5 pm from the nearest myofilaments. This suggests that an unknown intervening process such as diffusion or transport associated with cytoskeletal filaments may be involved in transporting protein chains to the assembly sites. Wenderoth and Eisenberg (1991) used a similar approach with cryosections and with sections of LR white-embedded papillary muscle. In both types of preparations, MHC mRNA was in the intermyofibrillar space adjacent to both A and I bands but was not identified as being associated with any structures. Enzymatic detection of MHC mRNA with a light microscope had previously shown that this mRNA is concentrated at the myotendinous junction of stretched fibers, where there is active myofibril assembly (Dix and Eisenberg, 1990), under the sarcolemrna and, possibly, between myofibrils in rabbit leg muscle (Dix and Eisenberg, 1988). Hesketh et al. (1991) used a sulfur-35-labeled riboprobe prepared from slow MHC cDNA to show that, in the rat soleus, about 70% of the myosin mRNA is present in the intermyofibrillar cytoplasm and the remainder is concentrated in a thin subsarcolemmal rim. A second approach to locating MHC mRNA involves a combination of cDNA probes and antibody localization. The rat extraocular (EO) musculature simultaneously expresses at least six different MHC genes at the mRNA level because of a sequential activation of MHC genes without complete repression of any of the genes (Wieczorek et al., 1985).Antibody localization showed that four different isozymes are present and tend to segregate into different cells, but some contain more than one MHC type. Isoform transitions in rat EO muscles seem to be arrested at different stages in different groups of cells; this suggests that tissue-specific or environmental factors may modify the expression of MHC genes. Miller et al. (1989) prepared four mAbs to epitopes on the head fragment of

PROTEINS IN STRIATED MUSCLE

67

fast MHC and used a two-part cDNA expression approach to map their respective epitopes. Each of the epitopes was at or near the ATPase site. One of them (F59) was conserved to varying degrees through vertebrate evolution, as shown by antibody binding to a spectrum of vertebrate muscles. A fifth mAb (F27) recognized an epitope located on the red portion of fast MHC and was also similarly conserved through vertebrate evolution.

d. Use of AntibodiesforFiber Typing Because of the existence of several distinct classification systems for vertebrate twitch-type muscle fibers and because of the evolution of some of these systems, the papers chosen for inclusion in this section will be described by using the fiber-type terminology used by the respective authors. Fiber typing has traditionally been done by using histochemical staining for enzymatic activity, normally myofibrillar ATPase, after preincubation at pH values 4.3, 4.6 or 9-10. The three fiber types identified by this approach are often named slow-twitch-oxidative (type I, or red), fasttwitch-oxidative-glycolytic (type IIA, or intermediate), and fast-twitchglycolytic (type IIB, or white). Since the advent of reliable methods to prepare striated muscle for study in the transmission electron microscope, ultrastructural characteristics such as relative numbers of mitochondria per cell and Z-line width also have been used as indicators of fiber type. Because the speed of contraction of muscle fibers is related to the expression of different MHC isoforms, the immunocytochemical detection of MHC isoforms has been and continues to be emphasized. Production of polyclonal or monoclonal antibodies to specific isoforms of myosin increased not only the sensitivity of detection of fiber types but also the apparent complexity of fiber types. Several early studies compared fiber typing based on histochemical staining or ultrastructural criteria with results from immunocytochemical staining with specific antibodies. Gauthier (1979) found that fast-twitch rat muscle contained red, intermediate, and white fibers on the basis of mitochondria1 content. Some of the red fibers bound antibodies specific for white or fast myosin, whereas others did not. All red fibers had wide Z lines, regardless of their myosin type. This led to the suggestion that rat diaphragm muscle contains four fiber types: red fast, red slow, intermediate, and white. Fiber typing of adult mouse muscle on the basis of Z-line width differed from the type I and I1 classification obtained after using antibodies to fast and slow myosin isozymes (te Kronnie et al., 1980). Human type I fibers have been unambiguously identified by using colloidal gold labeling and a mAb specific for slow MHC (Semper et al., 1988). A combination of enzyme histochemistry and immunohistochemistry for slow and fast myosin demonstrated that human type I fibers had a slow

68

MARVIN H. STROMER

myosin, type IIA and IIB had fast myosin only, and type IIC had various proportions of slow and fast myosin (Billeter et al., 1980). Bormioli et al. (1980) raised antibodies against slow myosin from the chicken anterior latissimus dorsi (ALD) and from the guinea pig soleus (SOL). Fibers from various vertebrate species that stained with anti-ALD myosin corresponded to slow-tonic fibers. Mammalian fibers stained with anti-SOL myosin, however, corresponded to slow-twitch fibers. In mammals, only a minority of EO fibers and nuclear bag fibers were labeled with anti-ALD myosin, Only afew avian slow fibers reacted with anti-SOL myosin. Thus, vertebrate skeletal muscle may have two antigenically distinct, though partly cross-reacting, types of slow myosin. The larger avian posterior adductor profundus (AP) muscle was analyzed with mAbs to myosin and C-protein isoforms and found to contain essentially pure slow-tonic fibers that were similar to those of the ALD (Zhang et al. 1985). This could make the posterior AP muscle a valuable source of slow-tonic fibers. The fast avian adductor superficialis (AS) muscle stained as a type IIA fiber by the histochemical ATPase test, but reacted with both antibodies to slow and fast MHC. The heterogeneity of this muscle is due to the presence of two MHC isoforms and not to the presence of variable amounts of slow myosin in its fibers (Zhang et al., 1989). By a combination of enzyme histochemistry and immunohistochemistry, Schiaffino et al. (1989) and Gorza (1990) have identified a type IIX MHC in rat skeletal muscles. The IIX MHC is localized in a large subset of type I1 fibers, is coexpressed with type IIA or IIB MHC in a small number of fibers, and may be a novel MHC. Muscles with predominantly type IIX MHC have a shortening velocity intermediate between muscles containing type I and type IIB MHC. Antibodies to myosin isoforms also have been used to relate isoform expression to fiber types and function in certain specialized vertebrate striated muscles. A polyclonal antibody to fast MHC from bovine EO muscle did not react with MHC of other myosins (Sartore et al., 1987). Most fibers with a type I1 ATPase stain pattern were labeled with EO antibodies, but some reacted with both anti-EO and either anti-IIA or antifetal myosin isoforms. Sartore et al. (1987) suggested that the EO isoform may exist because these muscles are exceptionally fast contracting. The EO myosin isoform is seemingly unrelated to the superfast myosin in the muscles closing a cat’s jaw. Extraocular antibodies do not stain the muscles closing a cat’s jaw (e.g., posterior temporalis). Similarly, antibodies to rat posterior temporalis myosin do not stain EO muscles. In EO muscle fibers in the rat, both single- and multiple-innervated fibers in the orbital layer labeled along their length with a mAb specific for neonatal/embryonic mammalian myosin, except in the endplate region. All fibers labeled in the endplate region with a mAb specific for fast myosin (Jacoby et al., 1990). These authors hypothesized that twitch or tonic innervation may act locally to suppress either neonatal-like myosin or fast myosin, respectively.

PROTEINS IN STRIATED MUSCLE

69

Myosin isoforms also have been studied in human masticatory muscles and in ear muscles from various mammals. The myosin composition of human masticatory muscles was more heterogeneous than limb muscles and included various proportions of slow and fast myosins, heavy as well as light chains (Thornell et al., 1984a). Many fibers of the human masseter muscle contain more than one myosin isoform (Butler-Browne et al., 1988). This muscle contains, in addition to adult fast and slow myosin, neonatal MHC and embryonic myosin light chains, two components of developing muscle. Myosin isoform-specific antibodies showed that there is no typical fiber-type composition in middle-ear muscles of several species of carnivores and some primates (Mascarello et al., 1983). This suggests that different functional requirements may dictate the myosin isoform composition of middle-ear muscles in different mammals. Staining with mAbs to MHC isoforms demonstrated that different isoforms existed in a single fiber and that there were discrepancies when fiber typing by mAbs was compared with fiber typing by histochemical staining in tensor tympani and stapedius muscles of the rat (van den Berge and Wirtz, 1989a,b). Collectively, the properties of these muscles indicate that they can contract rapidly, are fatigue-resistant, and, thus, are able to protect the inner ear from noise damage.

e. Myosin Isofomis in Muscle Spindles Muscle spindles are specialized receptors in skeletal muscle that respond to stretch of either the entire muscle or of the equatorial region of the spindle as a result of contraction of the intrafusal fibers. Sensory nerve endings exist at the equatorial region, and motor nerve endings are present at the ends of the spindle. Three fiber types are present in spindles: nuclear bag 1 , nuclear bag 2, and nuclear chain fibers. Because the intrafusal fibers express myosin isoforms different from those of the surrounding musculature, there has been interest in the patterns of isoform expression by intrafusal fibers. Species differences exist in the binding of mAbs against embryonic MHC to intrafusal fibers (Maier et al., 1988a). In cat, rat, and rabbit muscle spindles, nuclear bag 1 fibers stained strongly at both polar and juxtaequatorial regions. Nuclear bag 2 fibers stained little or not at all at the poles and moderately at juxtanuclear regions. In rat and rabbit muscle spindles, nuclear chain fibers did not stain but, in the cat, nuclear chain fibers stained as strongly as bag 1 fibers. Intrafusal fibers in EO muscles of the sheep, cow, and pig were studied with antibodies specific for slow-tonic or slowtwitch myosin (Scapolo et al., 1990). In the sheep and cow, bag fibers reacted with both antibodies, but one bag fiber in the pig was labeled with antibodies to slow-tonic myosin but not with antibodies to slow-twitch myosin. In rat and rabbit tibialis anterior muscle spindles, however, nuclear bag 2 fibers and nuclear chain fibers contain two or more myosin isoforms, but nuclear bag 1 fibers contain only one (Maier and Zak, 1989).

70

MARVIN H. STROMER

In adult rat SOL, slow-tonic and neonatal MHC are expressed only in spindle fibers (Pedrosa et al., 1989). Each of the three intrafusal fiber types has a unique and variable composition of MHC isoforms and M-band proteins. Maier and Zak (1990) also found that intrafusal fibers in chicken tibialis anterior muscle spindles and extensor digitorum longus (EDL) muscle spindles contained a nonhomogenous population of MHC. Carpenter et al. (1988) found that all intrafusal fibers in adult rat skeletal muscle reacted with antibodies to chick fast-twitch MHC and with antibodies to chicken slow-tonic MHC. Extrafusal fibers in adult rat skeletal muscle reacted with antibodies to chicken fast-twitch MHC, but not with antibodies to chicken slow-tonic MHC. Carpenter et al. (1988) suggested that extrafusal fibers with similar innervation are immunologically related and that the type of innervation may be an additional way to classify avian and rat fibers. Kucera and Walro (1988) severed either the sensory or the motor nerve supply to neonatal rat SOL muscles. In the neonatal rat, spindles are present but intrafusal fibers are structurally and immunocytochemically immature. Deafferented fibers did not react with antibodies to slow-tonic MHC or with antibodies to fast-twitch MHC. This indicates that intact sensory nerves are required for MHC expression during the postnatal development of rat spindles. Kucera and Walro (1991) used an antibody to a slow-tonic MHC isoform to show that the spread of expression of this isoform from the equator of prenatal rat intrafusal fibers toward the poles of bag 1 and bag 2 fibers may be modulated by sensory and motor neurons. An a-cardiac MHC isoform has been detected with monoclonal antibodies to human a-cardiac MHC in rat nuclear bag spindle fibers (Pedrosa er al., 1990). Neonatal deefferentation showed that motor innervation influences the expression of a-cardiac MHC along the spindle fibers. PedrosaDemellof et al. (1991) have pointed out that some of the discrepancies in the results from different labs with respect to the appearance of MHC isoforms in developing muscle spindles are due to differences in antibody sensitivity and specificity. f . Myosin Isoform Changes during Muscle Development Because of species and muscle differences, comments in this section are grouped by species in the following order: avian, human, rat, mouse, and cat. The avian embryo, especially chicken, has been a popular source of developing skeletal muscle cells. The sequential appearance of three broad categories of fast MHC isoforms at certain developmental stages has led to their designation as embryonic, neonatal, and adult isoforms (Bader et al., 1982; Bandman et al., 1982; Lowey et al., 1983). It is now clear that different isozymes may simultaneously appear in the same fiber. Zhang and Shafiq (1987) found that at Embryonic Day 16 (E16) all ALD fibers reacted strongly with a mAb that recognized slow MHCs and reacted weakly with a second mAb to slow MHC and with a mAb to fast MHC.

PROTEINS IN STRIATED MUSCLE

71

In the early stages after hatching, many fibers reacted with none of these antibodies. The adult pattern of strong reaction with both monoclonals to slow MHC and weak reaction with the antibody to fast MHC was not established until at least 9 weeks after hatching. Until E8, limb bud fibers produce both a ventricular MHC and a fast skeletal MHC (Sweeney et al., 1989). Young myotubes in adult muscle had a similar MHC pattern. At E6, some limb bud fibers that later became slow fibers began to express slow MHC. Sweeney er al. (1989) suggested that muscle progenitor cells commit to a single skeletal muscle lineage and that appearance of a fibertype-specific lineage occurs after localization of cells in premuscle masses. Muscle colonies formed by cloning myoblasts from E4-6 embryo hind limbs were of three types. Most numerous were colonies producing only fast MHC. Less numerous were colonies in which all myotubes produced fast and slow classes of MHC and most rare were colonies producing only slow MHC (Miller and Stockdale, 1986a). Colonies from E10-12 myoblasts contained myotubes that produced only fast MHC. Monoclonal antibodies were used to identify the myotube types. These results, together with data obtained by using monoclonal antibodies to study both chicken and quail embryos (Miller and Stockdale, 1986b), indicated that early myoblasts in the avian embryo are committed to three lineages, independent of innervation, that form primary myotubes differing in their MHC content. In a later phase, secondary myotubes form from myotubes in a single lineage, with maturation and fiber diversity dependent on innervation. Crow and Stockdale (1986) used mAbs to light- and to heavy-chain subunits of myosin to demonstrate that the distribution of embryonic fiber types in chicken thigh muscles was in a spatial pattern that predicted future fiber composition of the muscle. Innervation during the fetal phase is required for maintaining the pattern and controlling the myosin content of the cells. An earlier study, in which postmitotic myoblasts in clones derived from El0 chicken breast muscle were stained with antibodies to M-type creatine kinase and to skeletal MHC, also concluded that myogenic precursor cells were a heterogeneous population (Quinn and Nameroff, 1983). Cerny and Bandman (1986) inhibited spontaneous contraction of cultured chick muscle cells with either potassium (12 mM) or tetrodotoxin and found that neonatal MHC disappeared from virtually all myotubes. Immunocytochemical analysis of the fast fibers of the red strip in the adult chicken pectoralis showed that they contain embryonic fast MHC (Shear et al., 1988). During normal development, these fibers only transiently express neonatal and adult MHC. After denervation, however, the adult isoform is reexpressed, which may indicate that innervation represses certain MHC isoforms in the chicken. A special type of normally quiescent myoblast , the satellite cell, exists between the basal lamina and the sarcolemma of adult muscle cells (Campion, 1984). During muscle repair or regeneration, satellite cells prolifer-

72

MARVIN H. STROMER

PROTEINS IN STRIATED MUSCLE

73

ate, fuse and differentiate into specific muscle fiber types. Feldman and Stockdale (1991) used mAbs specific for avian fast and slow MHC to show that satellite cells from adult chicken and quail muscles are committed to form at least two fiber types. One type formed only fast fibers and the second type formed mixed (fast/slow) fibers. Different proportions of these two types of satellite cells are found in fast and slow muscles. Repeated subculturing of the progeny of these satellite cells resulted in no change in MHC isoform in cells from the chicken pectoralis major, an increase in fast versus mixed fibers in cells from the chicken ALD, and a rapid change from fast to mixed fibers in cells from quail ALD. The intracellular location of myosin isoforms in developing chicken muscle has been studied with both immunofluorescence and immunoelectron microscopy. Gauthier (1990) found that there were populations of myofibrils in pectoralis muscle cells at 7 days after hatching that contained more of a specific isoform than did other myofibrils in the same cell (Fig. 1). Monoclonal antibodies and immunoelectron microscopy have been used to track epitope changes on avian MHC during development (Winkelmann et al., 1983). An epitope located 14 nm from the head-rod junction in the N-terminal fragment of MHC is present in both adult and embryonic pectoralis myosin. Another epitope near the C-terminus of the myosin rod is present in adult fast myosin but not in early developing pectoralis myosin. A third epitope that is present on a MLC near the head-rod junction exists throughout development. Taylor and Bandman (1989) isolated myosin filaments from chicken pectoral muscle at different stages of development. Filaments from muscle at E12, 10 days after hatching, and at 1 year reacted only with mAbs to embryonic, neonatal, and adult myosins, respectively. Three classes of filaments were isolated from El9 muscle. One class reacted only with antibodies to embryonic myosin, a second reacted only with antibodies to neonatal myosin, and a third was recognized by antibodies to both embryonic and neonatal myosins. Three classes of thick filaments also existed in muscle 44 days after hatching, as indicated by reaction with antibodies to neonatal or adult fast-MHC antibodies or to both. Because the neonatal MHC antibody preferentially bound to the center of heterogeneous filaments, Taylor and Bandman

FIG. 1 Chicken pectoralis muscle, 7 days after hatching. Lowicryl section showing the response to gold-labeled anti-embryonic myosin. Specificity is demonstrated by the absence of labeling in the I bands. This mAb discriminates between two types of myofibrils. In the lower right, gold particles are localized in the A bands, except for the central bare zone. In the upper left, only a few particles are present in the A band (X25.000). (Reproduced from the Journal of CcIlBiology, 1990, Vol. 110, p. 697, by copyright permission of the Rockefeller University Press and by courtesy of G . F. Gauthier.)

74

MARVIN H. STROMER

(1989) speculated that neonatal MHC may have a unique role in thickfilament assembly. Monoclonal antibodies specific for adult and for neonatal myosin isoforms have been used to show that avian MHCs form predominantly homodimeric molecules with either two neonatal or two adult MHCs (Lowey et al., 1991). The use of antibodies also has enhanced our understanding of muscle fiber development in human embryonic muscle. Moore et al. (1984) used mAbs to adult fast and slow MHC and could detect adult slow MHC in 14-week-old human embryos, much earlier than could be done with histochemical stains. Zhang et al. (1987) also found that, in human fetal muscle at 19-20 weeks gestation, mAbs could identify three fiber types, those reacting with antibodies to slow MHC, with antibodies to fast MHC, or with neither, but histochemical staining could not discriminate these fibers. Slow MHC was detected with mAbs in all primary myotubes at 9 weeks of gestation but was not present in secondary or tertiary myotubes until 29 weeks of gestation (Draeger et al., 1987). These three types of myotubes appear sequentially and could be detected immunocytochemically but not histochemically . Monoclonal antibodies were also used to show that, in human muscle at 17-20 weeks of gestation, slow MHC continued to be present only in primary myotubes (Ecob-Prince et al., 1989b). Secondary and tertiary myotubes contained neonatal MHC and different levels of fast and embryonic MHC. A fetal MHC that is expressed at a decreasing rate during fetal life was detected by a mAb (Pons et al., 1986). Adult slow MHC was detected by immunostaining in a few fibers in 14- to 16-week fetal muscle and preceded the appearance of adult fast MHC. Antibodies to slow MHC have been used to show that embryonic rat muscle primary fibers initially produce slow myosin (Kelley and Rubinstein, 1980; Harris et al., 1989a). Narusawa et al. (1987) found, however, that in the 16-day rat fetus, all primary myotubes in the anterior tibial, EDL, and SOL muscles expressed both slow and embryonic myosin. Condon et al. (1990a) agreed that primary rat myotubes contain both slow and embryonic myosin and found that the loss of slow myosin is accompanied by the expression of neonatal myosin. As muscle development continues, the expression of slow myosin is inhibited in some fibers, which then express neonatal (Condon et al., 1990a) or fast isoforms (Narusawa et al., 1987). Narusawa et al. (1987) denervated the neonatal SOL and found that the amount of slow MHC declined, but when the neonatal EDL was denervated, the content of slow MHC increased. Narusawa et al. (1987) suggested that the nerve protects and amplifies production of the predominant MHC isoform and inhibits other isoforms. Lyons et al. (1983) stained fibers from the slow SOL and fast EDL at 20 days gestation and found that all fibers reacted strongly with antibodies to adult fast

75 myosin, and some also reacted with antibodies to adult slow myosin. The axial distribution pattern of fibers that recognized the slow myosin antibody resembled the distribution of slow fibers in the adult SOL. Sartore et al. (1982) used an antibody to fetal MHC to show that fetal myosin, which has a heterogeneous fiber distribution in fetal and neonatal rat muscle, is transiently expressed in regenerating muscle. In rat SOL, half the fibers contained embryonic and slow myosin 1 week after birth, and later contained only slow myosin (Butler-Browne and Whalen, 1984). A second group contained embryonic and neonatal myosin at 1 week, and later most contained only fast myosin. A part of this second group began at 4 weeks t o acquire slow myosin. These data were interpreted to suggest that the myosin isozyme sequence is embryonic to neonatal to adult fast myosin. An individual developing fiber may be induced to produce slow myosin as synthesis of other isoforms is repressed. Frozen sections of somites and early limb buds from 10- to 12-day mouse embryos and cultured cells from the same age limb buds stain with antibodies to both slow and embryonic fast myosin (Vivarelli et al., 1988). Thus, it seems that mouse and rat embryonic myoblasts are similar with respect to MHC synthesis, but both differ from avian embryonic myoblasts, which are divided into fast, slow, and mixed. Cells from limbs of 13-day mouse embryos have reduced affinity for the antibodies to slow myosins. By E15, cultured cells from the limb express only embryonic fast myosin, but cryostat sections show that primary fibers contain both slow and embryonic fast myosin and that secondary fibers contain only embryonic fast myosin. Whalen et al. (1984) found that 10% of the fibers in mouse EDL and semimembranosus muscles stained with antibody to adult slow myosin during the first month after birth and that, in the adult, only 0 to 0.8% of the fibers recognized the slow myosin antibody. In the newborn cat, 4.8% of the fast EDL and 26% of the slow SOL primary fibers stained strongly for slow myosin and stained weakly for fetal/embryonic myosin (Hoh et al., 1988a). The secondary fibers all reacted strongly with an antibody specific for fetal/embryonic myosin. With increased maturity, secondary fibers in the fast muscle became fast fibers and those in the slow muscle became slow fibers. In the newborn kitten posterior temporalis muscle, anti-fetal myosin stained almost all fibers uniformly but some fibers stained with antibodies to superfast myosin and lightly with antibodies to slow myosin (Hoh et al., 1988b). By 50 days, slow myosin staining disappeared and superfast myosin replaced fetal myosin, so that the muscle was nearly all superfast myosin, as in the adult. The masseter muscle in the adult cat has a unique myosin isoform composition, with a majority of the fibers expressing superfast myosin and the remainder expressing slow myosin (Hoh and Hughes, 1989). Antibody staining of fetal masseter muscle suggests there are two types of primary PROTEINS IN STRIATED MUSCLE

76

MARVIN H. STROMER

and two types of secondary fibers. Slow primary fibers stain with antibodies to slow myosin and stain transiently with anti-embryonic myosin. Superfast primary fibers stain with embryonidfetal and slow myosin antibodies during the perinatal period but later stain with antibodies to superfast myosin. Superfast secondary fibers express embryonic/fetal myosins in the fetus and replace these with superfast myosins after birth. Slow secondary fibers first express embryonidfetal myosins; shortly after birth, they express slow myosin or slow and superfast myosin, and later they express only slow myosin.

g . Effects of Innervation on Myosin Isoforms The early experiments by Buller et al. (1960), which demonstrated that cross-innervation caused a transformation in contraction speed (i.e., fast muscles became slow), created interest in the effects of the nerve supply on myosin isoforms. Gauthier et al. (1983) cross-innervated the normally fast cat flexor digitorum longus (FDL) with the SOL nerve and similarly cross-innervated the normally slow SOL muscle with the FDL nerve. The FDL muscle, after about 48 weeks, was nearly completely converted to slow fibers. The cross-innervated SOL, however, continued to recognize antibodies against slow myosin but also stained with antibodies to fast myosin light chains. Strips of the superfast cat posterior temporalis muscle were transplanted in the fiber-free bed of either the fast EDL or the slow SOL muscles (Hoh and Hughes, 1988). After 210 days, transplanted muscle strips in the EDL beds reacted only with anti-superfast myosin, but those in the SOL beds reacted mostly with anti-slow myosin, and a few fibers reacted with antisuperfast myosin. The EDL and SOL muscles regenerating in their own beds expressed fetal, slow, and fast myosins only. Because the transplanted superfast strips early in regeneration recognized antibodies to fetal, slow, and superfast MHC, Hoh and Hughes (1988) concluded that the expression of adult myosin isoforms can be modulated by the nerve. Denervation by surgical or chemical methods has also been used to study isoform transformations. Reinnervation of mouse SOL muscles 1 month after sectioning the SOL nerve resulted in the presence of hybrid fibers that stained with mAbs to both type I and type IIA myosin (Desypris and Parry, 1990). Denervation or chronic paralysis of rat calf muscles at El5 or earlier caused most primary myotubes to express only embryonic and neonatal MHC by El9 but not to express slow MHC (Harris et al., 1989b). Denervation or paralysis after El5 did not cause this effect. In these rat muscles, the sequence of myosin isoforms is dependent on active innervation during a particular embryonic period. Normal 1-month-old and adult rat gastrocnemius muscle type IIB (fast glycolytic) fibers can be divided into three subgroups by their graded staining with a mAb to MHC (Leung et al., 1987). Neonatal denervation, however, caused gastrocne-

77 mius fibers to stain uniformly with the antibody. This suggests that the nerve supply is responsible for maintaining the heterogeneity of myosin isoforms in the fibers. Brachial levels of the neural tube were surgically removed at E2 in avian embryos (Phillips et a / . , 1986). At E l I , muscles that had developed aneurally were smaller, but the distribution of fast and slow myotubes was unchanged compared with normal embryos. This finding in the avian embryo seems to differ from the situation in the rat, which has an apparent requirement for innervation to produce different myosin isoforms. Curare treatment of El8 chick ALD, however, causes the expression of a second fiber type that stains with an antibody to fastmyosin LCl (Gauthier et a/., 1984). The authors speculated that the change in the motoneuron pool may have induced the expression of this second myosin isoform. Denervation or tetrodotoxin block of the sciatic nerve in adult rats cause reexpression of embryonic and neonatal MHCs in type IIA fibers (Schiaffino et al., 1988). Immunoreactivity differed along the same fiber, implying that myosin isoform expression within a fiber is coordinated by the nerve. Injection of P-bungarotoxin into rat fetuses before muscle innervation caused no changes in either the fiber types or their intramuscular distribution as detected by antibodies to myosin when compared with controls (Condon et a / . , 1990b). The ultimate fiber type was achieved in some fibers without the normal sequence of isoform changes, and some slow fibers lost their ability to express slow myosin. Six months after adult cats were spinalized at T12-Tl3, the normally slow SOL muscle showed an increase in the percentage of fibers that stained with antibodies to fast MHC or to both fast and slow MHC (Jiang et a/., 1990). Direct electrical stimulation of muscles to affect myosin isozyme transformation also has been a popular approach. Among the early studies, Rubinstein et a/. (1978) continuously stimulated fast rabbit muscles at 10 Hz and found that this caused preexisting fibers to switch from synthesis of fast myosin to synthesis of slow myosin. Stimulating the fast EDL muscle of the rabbit at 10 Hz for 12 hr/day resulted in transformation to a slow-type muscle (Maier et a/., 1986a). Examination of this muscle after 6 to 21 days stimulation showed that fast-twitch fibers were degenerating and that small fibers, often with central nuclei, that reacted with antibodies to embryonic MHC were present. Maier et al. (1986a) concluded that, in addition to the isoform switches in adult fibers, newly formed slow fibers also could contribute to the muscle transformation. Maier et al. (1988b) subsequently found that chronically stimulated fast muscle from adult rabbits contains fibers that react with antibodies to neonatal/embryonic and embryonic MHCs, two isoforms normally not present in adult muscle. Low-frequency stimulation seemingly influences some of the newly formed myotubes to become slow fibers because, as duration of stimulaPROTEINS IN STRIATED MUSCLE

70

MARVIN H. STROMER

tion increased, fewer fibers stained with the neonatallembryonic and embryonic MHC antibodies. Immunogold labeling and examination in the electron microscope were used by Franchi et al. (1990) to study stimulation-induced fiber-type transformation in rabbit muscle. After 4 weeks of stimulation, fibers of the fast tibialis anterior muscle reacted with mAbs to both slow and fast myosins. After 6 weeks, only labeling with antibodies to slow myosin was seen. After 7 weeks of stimulation and 3 weeks of recovery, fibers again reacted with both antibodies. The distribution of gold particles indicated that newly synthesized myosin is incorporated throughout the length and cross section of the A-band. The opposite fibertype transformation was done by Gorza et al. (1988a), who stimulated rat adult slow SOL muscles with fast (100 Hz) stimuli. All stimulated fibers labeled strongly with antibodies to fast myosin, and at least 90% were also weakly labeled with antibodies to slow myosin. The labeling with antibodies to fast myosin could first be detected 7 days after stimulation was begun. No fibers reacted with antibodies to fetal myosin, indicating that the slow-to-fast transformation occurred in preexisting adult fibers in the rat SOL. Coculturing muscle cells and nerves is another approach to studying the effect of nerves on myosin isoform expression. Culturing section of embryonic mouse spinal cord with explanted adult mouse muscle fibers results in the degeneration of the adult fibers and the proliferation of satellite cells to form new myotubes (Ecob et al. 1983). Some of these myotubes become innervated by the embryonic mouse spinal cord neurons and contain, after 3 to 5 weeks, adult fast MHC. Ecob-Prince et al. (1986) subsequently found that fast MHC was absent both in new rat myotubes in cocultures and in older myotubes cultured without spinal cord tissue. The blockage of nerve-induced contractions with a-bungarotoxin did not decrease the number of fibers in cocultures that expressed fast MHC. Thus, it seems that the synthesis of adult fast MHC is dependent on the presence of nerves but not on the nerve-induced contractile activity.

h. OtherExternalInJluences on Myosin Isoforms The effects of a variety of external factors, principally on myosin but on other myofibrillar proteins as well, have been investigated with immunocytochemistry. Stretch hypertrophy of the chicken ALD muscle caused 28-52% of the fibers to stain with antibodies to fast MHC by Day 12 to 19 of stretch compared with <1% in the control ALD (Everett and Sparrow, 1987). Most of these fibers also stained with antibodies to slow MHC. By 50 days of stretch in some birds and by 80 days in all birds, the staining had reverted to normal. Hypertrophy of the ALD induced by weight overload causes nearly complete replacement of one slow-tonic myosin isoform with another slowtonic isoform (Kennedy er al., 1986, 1988). In addition, small fibers, possi-

PROTEINS IN STRIATED MUSCLE

79

bly of satellite cell origin, express a myosin isoform characteristic of embryonic muscle cells. Regeneration of adult chick skeletal muscles after focal freeze injury is accompanied by MHC and C-protein isoform switches that resemble those found in developing muscle, but differ in both temporal and muscle-type-specific patterns (Saad et al., 1987). Thyroxine feeding caused the conversion of slow-tonic chicken fibers to fast-twitch fibers (Carpenter er al., 1987). Antibodies to fast-twitch MHC and slow-tonic MHC showed that newly synthesized MHC was incorporated uniformly across the fiber in cross sections and incorporated homogeneously into each sarcomere. Antibodies to either slow or fast myosin showed that the SOL muscle of 1-day-old rabbits contained 25% slow and 75% fast fibers (Lutz er al., 1978). In the young adult (5-8 months), more than 90% were slow fibers, but in old age (4-7 years), up to 50% of the SOL fibers contained fast myosin. Endurance training of human triceps brachii muscle resulted in an increase in the number of intermediate fibers that were characterized by the presence of both fast and slow isoforms of MHCs and troponin C, I, and T (Schantz and Dhoot, 1987). Bovine slow muscle fibers involved with movement, such as jaw muscles, stained with antibodies to slowmyosin LCl, but slow fibers in deep hip muscles did not, suggesting that postural muscles do not contain this light chain (Young, 1989). 2. C Protein

Early evidence of the existence of slow and fast isoforms of C protein in rabbit muscle (Callaway and Bechtel, 1981) and in adult chicken muscle (Reinach et al., 1982)was verified by the use of isoform-specific antibodies. Myofibrils isolated from neonatal chicken breast muscle reacted with both antibodies, which indicated that both C protein isoforms existed in the same myofibril (Obinata et al., 1984a). In developing chicken skeletal muscle, slow C protein appeared after slow myosin light chains disappeared, but fast C protein appeared after fast myosin light chains accumulated (Obinata et al., 1984b). Bahler et al. (1985~)examined younger chicken embryos and used antibodies to detect cardiac C protein between E3 and El5 in developing skeletal muscles. From El5 on, white skeletal C protein is progressively accumulated and the cardiac isoform is lost. Primary cell cultures continue to express cardiac C protein throughout extended culture times. A complex labeling pattern in rabbit and human skeletal muscle has been described by Dhoot et al. (1985). Anti-slow skeletal C protein stained all type I and a fraction of type I1 rabbit fibers with various intensities. Type I1 rabbit fibers either stained strongly with antibodies to fast C protein or stained with antibodies to both fast and slow C protein. In human skeletal muscle, this antibody to slow C protein

80

MARVIN H. STROMER

stained all fibers, but the antibody to fast C protein stained only type 11 fibers. Fluorescent localization of antibodies to C protein in rabbit psoas, plantaris, and SOL muscles indicated that C protein is present in largest amounts in fast white and fast intermediate fibers and is absent from slow red fibers (Starr et a / . , 1985). With the EM, C protein was localized in stripes 4-1 1 in psoas, in stripes 5-1 1 in psoas and plantaris, in stripes 3 and 5-11 in plantaris, or in no stripes in red fibers (Bennett et al., 1986). Dennis et a / . (1984) observed with the EM that, although there were differences in the number of stripes and positions occupied by antibodies to the slow and fast isoforms, the fact that the same stripes were labeled by the two antibodies in one muscle indicates that two C protein isoforms can coexist in a single stripe.

3. Myomesin and M Protein Myomesin and M protein are both located in the M line at the center of the A band. The nomenclature of these proteins was clarified by Grove el a / . (1984), and any pre-1984 papers included in this section will be presented with the current nomenclature. Myomesin is the name currently used for the 185-kDa M-line protein, and M protein is the 165-kDa M-line protein. An early study with an antibody to M protein extracted from frog muscle demonstrated that the fluorescent label was located in the M line (Dhanarajan and Atkinson, 1980). More recently, Pedrosa et al. (1989) were able to localize myomesin in all three intrafusal fiber types in rat muscle spindles. M-protein antibodies always stained chain fibers, never stained bag 1 fibers, and stained bag 2 fibers with intermediate intensity and regional variability. At the ultrastructural level, antibodies to M protein labeled the entire M line from striation M6 through M6’ in chicken breast muscle (Strehler et a / . , 1983). Low-ionic-strength extraction of chicken pectoralis muscle removes the MI, M4, and M4’ striations and leads to an incomplete labeling of the M line between M6 and M6’ with antibodies to M protein. Changes in the synthesis and location of both myomesin and M protein have been investigated in developing muscle from chickens and rats. In cultured chicken pectoral myogenic cells, myomesin accumulated and was localized in the M line 1.5 days before M protein (Grove et al., 1985). In embryonic chicken pectoralis major muscle, at E7, presumptive type I and I1 fibers both contain M protein and myomesin (Grove et al., 1987). From El0 to E14, M protein is suppressed in presumptive type I and type I1 fibers formed from primary myotubes. All adult type I1 fibers, however, contain M protein as well as myomesin. Developing anterior and posterior latissimus dorsi muscles also show a suppression, beginning at EIO, of M protein, first in primary and then in secondary presumptive slow-tonic

PROTEINS IN STRIATED MUSCLE

81

type I11 fibers (Grove and Thornell, 1988). M protein is also transiently suppressed in presumptive fast-twitch type I1 fibers derived from primary myotubes, but not in those from secondary myotubes. In the rat embryo, incorporation of myomesin into the M line also precedes that of M protein 1990). Fetal and newborn rat and muscle creatine kinase (Carlsson et d., muscles all contained myomesin and M protein. At 4 weeks of age, however, type I fibers in the SOL and EDL muscles no longer contained M protein and had only M6, M4, M4', and M6' striations in the M line. This led to the suggestion that, in the rat, M protein may be located in the central M I striation. Guinea pig type I fibers also lack M protein, but contain all five striations in their M lines (Thornell et al., 1990), which suggests that the location of M protein may be species-specific. The expression and incorporation of myomesin and M protein into fetal rat M lines are nerve-independent, but the suppression of M protein synthesis during postnatal development is nerve-dependent (Carlsson et al., 1990). 4. MM-Creatine Kinase MM-creatine kinase (MM-CK) is the dimeric form of the muscle isozyme of creatine kinase and is an established component of M lines in skeletal muscle. Although much of the MM-CK is in a soluble form, some is bound to the M line. Early experiments showed that labeling chicken skeletal muscle with an excess of anti-M-CK IgG or anti-M-CK Fab fragments caused heavy decoration of the M line (Walliman et al., 1977)and removal of the M line (Wallimann et al., 1978), respectively. It was subsequently shown that labeling with lower concentrations of antibodies to M-CK decorated the M4 and M4' M-line striations (Wallimann et al., 1983; Strehler et al., 1983; Wallimann and Eppenberger, 1985). All three fiber types in muscle spindles label with antibodies to MM-CK, but bag 1 fibers reacted less strongly and did not display clear striations (Pedrosa et al., 1989). Immunogold localization of MM-CK in muscle cryosections after removal of soluble CK showed labeling of both the M line and the sarcoplasmic reticulum (SR) membranes (Rossi et al., 1990). Purified SR vesicles also labeled with antibodies to M-CK. Extraction of SR vesicles with 0.6 M KCl released no MM-CK, but treatment with an EDTA/low-salt buffer released some MM-CK from isolated vesicles. MM-CK seems to be bound fairly tightly to the SR and may be favorably located to regenerate ATP needed by the calcium transport ATPase.

5. 86-kDa Protein The 86-kDa protein has been isolated from a crude C-protein preparation from chicken pectoralis muscle (Bahler et al., 1985a). Antibodies to this

a2

MARVIN H. STROMER

protein do not cross-react with myosin, M-line proteins, titin, or C protein; in immunofluorescence, they label in the A bands of fast-twitch fibers but not slow-tonic or cardiac fibers. With immunoelectron microscopy, antibodies to the 86-kDa protein label 9 stripes spaced 43 nm apart in each half of the A band (Bahler er al., 1985b). Colocalization in strips 5 to 11 occurred when antibodies to the 86-kDa protein and to C protein were added to glycerinated fast-twitch fibers. Stripes 3 and 4, which are closer to the M line and also are spaced with a 43-nm periodicity, are labeled by antibodies to 86-kDa protein but not with antibodies to C protein. 6. F Protein, H Protein, and X Protein Each of these proteins has been localized in the A band. F protein has been identified as phosphofructokinase. Immunofluorescence with antibodies to F protein shows labeling in a zone in each half of the A band (Offer el al., 1988). The location of these two zones within the A band and the increase in zone size as sarcomere length increases suggest that phosphofructokinase is located in the cross-bridge region of thick filaments and that overlapping thin filaments may restrict access of the antibody. Immunofluorescent labeling with antibodies to H protein or antibodies to X protein also produced a labeled zone in each half of the A band that was independent of sarcomere length (Starr er al., 1985). Labeling patterns differed significantly from muscle to muscle. H protein was present in fast white rabbit psoas fibers and was absent in fast and slow red psoas fibers, but all isolated psoas myofibrils seemed to label with antibodies to H protein. In the rabbit plantaris muscle, H protein was absent from fast white fibers, but was present in some slow red fibers. X protein is present in fast and slow red fibers and is absent from fast white fibers. The distance between the two labeled zones in the psoas A band was much less with antibodies to H protein (0.35 pm) than with antibodies to C protein (0.64 pm). The anti-X protein zones were slightly, but significantly, closer (0.52 pm) than the anti-C protein zones. In SOL and plantaris myofibrils, the zones labeled by antibodies to X protein were slightly farther apart (0.67 pm) than those labeled with antibodies to C protein. This heterogeneous labeling pattern with antibodies to H and X proteins also was observed with the EM (Bennett er al., 1986). H protein was present near stripe 3 (14 nm nearer the M line) in psoas fibers, but was absent in SOL and plantaris muscle. X protein was detected in stripes 3-11 in red fibers of all three muscles, in stripe 4 only in psoas and plantaris, in stripes 3 and 4 in psoas and plantaris, or in no stripes. Anti-X protein stripes are 8-9 nm further from the M line than the corresponding stripes labeled by antibodies to C protein and are wider because they consist of a doublet with an internal spacing of 16 nm. The precise locations in the A band of H protein and X

PROTEINS IN STRIATED MUSCLE

a3

protein differ from each other and also differ from that of C protein. Bennett et al. (1986) suggested that the presence and arrangement of these three proteins may define a more intricate system of muscle fiber classification than is currently used. 7. Ribosomes

Antibodies that recognized five major proteins of the large ribosomal subunit from rat liver labeled frozen rat skeletal muscle sections at the inside of the sarcolemma and between myofibrils (Horne and Hesketh, 1990). The intermyofibrillar staining was located between A bands of adjacent myofibrils and decreased, compared with the subsarcolemmal staining, as animal age increased.

8. Skelemins Polyclonal antibodies to the skelemins (200- and 220-kDa) do not crossreact with MHC, talin, fodrin, synemin, or MAPS and, in immunofluorescence, they localize at the periphery of M lines in both skeletal and cardiac muscle (Price, 1987). Skelemins have been detected in human, bovine, and rat muscles, but not in chicken muscles.

6. Thin-Filament Proteins and I-Band-Associated Proteins 1. Actin Because actin is a ubiquitous protein, many attempts to produce actin antibodies have involved SDS treatment of the immunogen, attachment of F-actin to an inert carrier such as Affigel702, or adsorption of actin to alum precipitates. These and other approaches have frequently resulted in antibodies with very different specificities or in very low titers, or in no immunological response. These difficulties in obtaining a reliable antibody for the detection of actin have been circumvented by the use of phalloidin complexed with various fluorochromes or to colloidal gold (Lachapelle and Aldrich, 1988). It should be stressed that the phalloidin approach detects filamentous or F-actin and does not discriminate between actin isoforms. Although the sarcomeric isoform of actin in mature skeletal muscle is skeletal a, the presence of other muscle types (e.g., smooth muscle in blood vessels) in mature muscle and the existence of isoform switches during myogenic differentiation indicate that it would be advantageous to label different isoforms. Antibodies to an amino-terminal peptide that is unique to skeletal a-actin have been used to detect a-actin in skeletal

a4

MARVIN H. STROMER

and cardiac muscle (Bulinski et a/., 1983). Peptides corresponding to the N-terminus of skeletal muscle a-actin and to the N-terminus of nonmuscle y-actin also have been used as antigens (Otey et af., 1988). I bands of isolated rabbit myofibrils stained strongly with antibodies to the a peptide and much less strongly with antibodies to the y peptide. Immunogold labeling of cultured L6 myoblasts indicated that the nonmuscle isoform was found predominantly in the cortical actin filaments and that the skeletal muscle isoform was located predominantly in nascent myofibrils. It should be emphasized, however, that both isoforms were present in all actin-containing structures. For the detection of specific isoforms of actin, the current method of choice seems to be the use of monoclonal antibodies as described by Lessard (1988). This approach has been successfully used by Handel et af. (1989) to localize a- and y-actin isoforms in cultures of rat skeletal muscle. An antibody to N-terminal residues of actin has been used to determine the role of these residues in the actomyosin interaction. DasGupta and Reisler (1991) compared the binding of myosin subfragment 1 (Sl) to actin in the presence or absence of nucleotides when as much as 80% of the actin was saturated with Fab fragments of the antibody to the first seven N-terminal actin residues. In the absence of nucleotide, the antibody decreased the binding of S1 to actin by a relatively small amount. The antibody, however, strongly inhibited the binding of S1 nucleotide complexes to actin, especially at low S1 concentrations. This inhibition was partly released by increasing the S 1 concentration. These results indicate that either the N-terminal residues per se or another part of actin perturbed by the binding of this antibody is more important for the binding of S I to actin in the presence than in the absence of nucleotides. 2. Tropomyosin

Remarkably few immunolocalization studies have been done in intact skeletal muscle with antibodies to tropomyosin. Endo et af. (1966) used antibodies to native tropomyosin, which is a complex of troponin and tropomyosin, to demonstrate that the fluorescent label existed in the I band and, perhaps, along the entire thin filament. I-band fluorescence also was seen when Pepe (1966) used antibodies to tropomyosin to label myofibrils. In the EM, these labeled myofibrils displayed an irregular increase in I-band density. During development of chicken breast muscle, the red strip contains a-fast, a-slow, and p isoforms of tropomyosin (Matsuda et af., 1983). The a-slow isoform is absent from other areas of the breast. P-Tropomyosin is gradually lost in the posterior regions of the developing chicken breast. Lin et a / . (1985) developed several monoclonal antibodies to chicken tropomyosin isoforms. One of these antibodies (CL2)

PROTEINS IN STRIATED MUSCLE

a5

labeled microfilaments with a periodicity of 35-37 nm and was used to isolate skeletal tropomyosin-enriched microfilaments from muscle cells differentiating in vifro (Lin and Lin, 1986). These isolated microfilaments contained muscle and nonmuscle isoforms of both actin and tropomyosin and may have been an intermediate class of microfilaments assembled during thin-filament maturation. More recently, Trombitas et af. (1990a) used polyclonal antibodies to chicken cardiac a-tropomyosin and mAbs to chicken cardiac and to chicken leg tropomyosin to label single, stretched frog semitendinosus fibers. The polyclonal antibody formed 23 stripes spaced 38.4 nm apart along the length of thin filaments in highly stretched sarcomeres. This antibody did not label the 24th stripe adjacent to the Z line, nor were the connecting or gap (titin) filaments labeled in the gap at the A-I junction. Both mAbs stained the I bands periodically, but less intensely than the polyclonal antibodies. The mAbs stained the 24th stripe on either side of the Z line. The mAb to chicken leg tropomyosin had a higher affinity for the region near the Z line. It is unknown at present whether these labeling patterns represent differences in location of tropomyosin isoforms; differences in functional properties, such as extent of phosphorylation; or some other attribute.

3. Troponin Troponin (TN) is a collective term applied to a complex of three regulatory proteins, troponin C (TN-C), which has two calcium-specific binding sites; troponin I (TN-I), which is involved in inhibition of the actin-myosin interaction; and troponin T (TN-T), which is responsible for binding the troponin complex to tropomyosin on thin filaments (Greaser and Gergely, 1973). When chicken breast mucle myofibrils or separated thin filaments were stained with antibodies to TN, a pattern of 24 periods with an approximately 40-nm spacing was observed along the length of thin filaments (Ohtsuki et al., 1967). Antibodies to each of the three TN components were distributed along the thin filament with a spacing of 38 nm (Ohtsuki, 1975). Staining El9 or E20 embryonic chick breast muscle with antibodies to TN produced 25 to 29 striations with 38-nm spacing (Ohtsuki, 1979a) and may indicate the thin filaments in embryonic breast muscle are longer than those in the adult. Antibodies to TN-T, and -T2, two chymotryptic fragments of TN-T, also formed transverse striations at 38-nm intervals along thin filaments (Ohtsuki, 1979b).The striations caused by antibodies to TN-T, began 40 nm from the free ends of thin filaments and the striations caused by antibodies to TN-T, striations began 27 nm from the free ends. A monoclonal antibody to TN-T that stains I bands in adult and embryonic skeletal and cardiac muscle of several vertebrate species has been produced (Lim et d.,1984). An intense but diffuse cytoplasmic staining was

86

MARVIN H. STROMER

seen in chicken gizzard smooth muscle. Although it is well established that smooth muscle contains no TN; this antibody seemingly recognizes a determinant shared by striated and smooth muscle. In adult skeletal muscle, slow and fast isoforms of each of the components of the TN complex are located in type I and type I1 fibers, respectively (Dhoot and Perry, 1979). In developing rat, mouse, and rabbit leg muscles, however, fast and slow isoforms of TN-C, TN-I, and TN-T can exist in the same cell. Fetal rat skeletal muscle, at 12-14 days of gestation, contains five isoforms of embryonic slow skeletal TN-T (Sabry and Dhoot, 1991). Two of these embryonic isoforms are lost by 17 days in utero when adult slow TN-T and small amounts of embryonic cardiac TN-T are present. None of the embryonic isoforms of skeletal TN-T could be detected in postnatal rat skeletal muscle. Two isoforms of slow TN-T are present in adult human skeletal muscle and in fetal human skeletal muscle at 20 weeks of gestation (Sabry and Dhoot, 1991). A third isoform of slow TN-T was found in a subset of adult human skeletal muscles, but was usually not detected in any fetal human skeletal muscles. Matsuda et al. (1981) found two TN-T isoforms in developing chicken breast muscle, but only one in developing leg muscles. The TN-T isoform in leg muscle and the TN-I and TN-C in both breast and leg muscles do not change during development. Adult breast muscle also contains only one TN-T isoform. Cultures of embryonic pectoralis muscle cells stained with antibodies to the three skeletal TN components and with antibodies to cardiac TN-T and TN-C for the duration of the cultures (Toyota and Shimada, 1983). If these cells were cocultured with embryonic motor or sympathetic nerves for 1 week, the cells no longer stained with antibodies to cardiac TN-T and TN-C but stained only with antibodies to skeletal TN components. This suggests a potential role for innervation in TN isoform development. Adult chicken skeletal muscle fibers that were regenerating after cold injury contained new cells that stained like embryonic cells, in other words, stained with antibodies to all skeletal TN components and with antibodies to cardiac TN-T and TN-C (Toyota and Shimada, 1984). As differentiation continued, the adult staining pattern was established. Shimizu and Shimada (1985) prepared mAbs that could distinguish between breast and leg TN-T isoforms. All fibers in all fast muscles of 12-day-old chicken embryos stained with the antibody to leg TN-T. Late in embryonic development, breast TN-T accumulated in leg muscle. By 1 week after hatching, breast muscle fibers that contained breast TN-T increased in number, and leg fibers that contained breast muscle TN-T decreased in number. Hartner et al. (1989) used polyclonal antibodies that reacted only with fast TN-T isoforms or only with slow TN-T isoforms to monitor the fastto-slow TN-T transitions in rabbit hindlimb muscles that received chronic

PROTEINS IN STRIATED MUSCLE

a7

low-frequency stimulation. At least six fast TN-T isoforms and two slow TN-T isoforms were detected. Prolonged stimulation produced a TN-T isoform pattern similar to that found in a normal slow-twitch muscle such as the SOL. 4. a-Actinin

a-Actinin is a 200-kDa protein that consists of two anti-parallel 100-kDa polypeptide chains and is located at the Z line of mature skeletal muscle. Although the first published report of a-actinin localization by immunofluorescence is credited to Masaki et al. (1967), subsequent experiments in several laboratories demonstrated that the procedure for the preparation of that antigen would likely produce a complex protein mixture, one of which was a-actinin. It is generally assumed that the protein mixture used as the antigen is the reason that this antibody also stained the M line and produced background staining that was greater than expected. Development of a method to purify a-actinin led to the production of specific antibodies that labeled only the Z line (Schollmeyer et al., 1972, 1973, 1974). It should be emphasized that, in immunofluorescence, anti-a-actinin staining is uniform across a Z line in either skeletal or cardiac muscle. This is in contrast to the pattern observed after staining with antibodies to desmin, where the label is concentrated at the edge of or between myofibrils, thus producing a punctate fluorescence pattern that indexes at Z lines. The complimentary staining patterns obtained with antibodies to aactinin and desmin were demonstrated, in immunofluorescence, by Lazarides er al. (1982), who showed that isolated Z-disk sheets stained continuously with anti-a-actinin within individual Z lines of the sheet. Antibodies to desmin, however, stained in a netlike pattern that surrounded individual Z lines in the sheet. Although most rat skeletal myoblasts in culture stained homogeneously with antibodies to a-actinin, longitudinal strands that also stained with antibodies to a-actinin were detected in both myoblasts and small and larger immature myotubes (Jockusch and Jockusch, 1980). Spontaneously contracting myotubes in these cultures displayed a Z-line staining pattern with antibodies to a-actinin unless contraction was blocked with 1 p M tetrodotoxin. Administering tetrodotoxin before contractions began resulted in a pattern of chains of well-separated, small stained specks. After contractions began, tetrodotoxin did not affect the a-actinin patterns for 5 days: after 5 days, parallel longitudinal arrays stained with the antibody. Skeletal muscle a-actinin was absent from mononucleated skeletal myoblasts unless the myoblast contained myofilament bundles (Endo and Masaki, 1984). Smooth muscle a-actinin, however, was present in mononucleated myoblasts in the cytoplasm and on sarcolemma-associated structures. After fusion. skeletal a-actinin was

88

MARVIN

H. STROMER

present in myofilament bundles, Z bodies, and Z lines. In immature and mature myotubes, smooth muscle a-actinin was confined to sarcolemmaassociated structures. Chowrashi and Pepe (1982) showed that antibodies to both a-actinin and amorphin [amorphin is an 85-kDa protein that may be identical to phosphorylase b (Maruyama, Kei, et al., 1985)l produced immunofluorescent staining of mature Z lines. Immunofluorescence with antibodies to a-actinin has been used to follow the translocation of sarcomeric a-actinin from Z lines to cortical actin-containing bodies in cultured skeletal muscle cells treated with phorbol myristate acetate (Lin et ul., 1989a). a-Actinin has also been localized in Z lines in several striated muscles in Drosophila (Vigoreaux et al., 1991). a-Actinin has also been localized with the EM. Tokuyasu (1983b) used ferritin or colloidal gold to identify the location of antibodies to a-actinin in ultrathin cryosections of chicken sartorius muscle. An antibody to the rod domain of dystrophin cross-reacts strongly with a 90-kDa protein that has been identified as a fast-twitch glycolytic isoform of myofibrillar aactinin (Hoffman et d . , 1989). Ultrathin cryosections of mouse EDL muscle contain Z lines that label strongly with this antibody, but sections from the slow-twitch SOL muscle do not label. 5. Nebulin

Nebulin is a family of very large sarcomere-associated cytoskeletal proteins (Wang, 1985) whose size varies from 600 to 900 kDa depending on the tissue, the species, and the developmental stage of skeletal muscle (Stedman et al., 1988;Jin and Wang, 1991). Thus far, nebulin has not been found in cardiac muscle. Polyclonal antibodies to nebulin produced an immunofluorescent band on each side of the Z line that coincided with the N, lines in I bands of long sarcomeres P 2 . 5 pm) (Wang and Williamson, 1980). Immunofluorescent staining of myofibrils with 2.8-pm sarcomeres that lacked N, lines because of an improved preparative approach showed a wide, brightly stained band on each side of the Z line and a more lightly stained zone in the thick- and thin-filament overlap region (Wang and Wright, 1988). There was also a narrow stained band at the Z line. The changes in intensity and location of these stained regions at shorter sarcomere lengths were consistent with the movement of nebulin epitopes toward the center of the sarcomere in a manner that paralleled the increased overlap of thick and thin filaments. This was confirmed with immunoelectron microscopy, which showed six antibody-binding stripes or bands along thin filaments in each half of I bands in myofibrils stretched until the A-I gap region was visible. As sarcomere length decreased, these stained bands were sequentially incorporated into the A band and retained

PROTEINS IN STRIATED MUSCLE

89

their same relative positions with respect to the Z line (Fig. 2). This led Wang and Wright (1988) to propose that nebulin exists in a set of inextensible filaments, attached at one end to the Z line, that are at least 1 pm long and that could serve as a template or “ruler” for actin assembly.

FIG. 2 Distribution and stretch response of nebulin epitopes. Rabbit psoas-split fibers were stretched to various sarcornere lengths and labeled with anti-nebulin and protein A-gold conjugates. A preimmune control of nonoverlap sarcorneres in A indicates the absence of prominent I-band striations as opposed to those present in sarcomeres labeled with nebulin antibodies, displayed in decreasing lengths from (B) to (F). T in (A) and (B) indicate titin filaments spanning the gaps of separated A and I segments in these sarcomeres. Antibody stripes in the I band are designated a-f (arrows). The axial position of the distal epitopes that have entered the A band are detectable by gold clusters (brackets) indicated by arrowheads in (E). Gold labeling of the stripe a is indicated by an arrow in (F) (X31,200) (Reproduced from the Journal of Cell Biology, 1988, Vol. 107, p. 2207, by copyright permission of the Rockefeller University Press and by courtesy of K. Wang.)

90

MARVIN H. STROMER

Maruyama et al. (1989) and Pierobon-Bormioli et al. (1989)confirmed that anti-nebulin binding sites in the I band did not change position during either stretch or shortening. Support for the concept that nebulin serves as a template or length regulator for thin filaments was published by Kruger et al. (1991), who found that skeletal muscles that have larger nebulin have proportionally longer thin filaments. The nebulin variants had extra protein segments at either or both ends of the nebulin molecule. Nebulin epitopes in the I band repeat at 40 nm and enhance the intrinsic 40-nm thin-filament repeat (Fig. 3). 6. Filamin

Filamin is a high-molecular-weight (500 kDa) actin-binding protein. Immunofluorescent localization in adult fast chicken pectoral muscle showed that filamin is located at the periphery of Z lines (Gomer and Lazarides, I98 1). In cultured skeletal muscle cells, double immunofluorescence has shown that filamin localizes at the Z line shortly before desmin or vimentin is associated with the Z line. In the slow ALD muscle, however, filamin was localized throughout the I band by immunofluorescence (Gomer and Lazarides, 1983). Although this antibody was prepared by using a smoothmuscle antigen, recognition of skeletal filamin was not impaired. Quantita-

FIG. 3 Epitope profile of nebulin N46 mAb. This antibody labeled a group of 10 stripes spanning from 0.25 to 0.68 pm from the Z line in LD muscle. The repetitive epitopes are spaced at 40 nm and its multiples. Antibody labeling seems to enhance the intrinsic 40-nm thin-filament repeat at the loci (tick marks spacing: 5 repeats). The higher magnification (inset) demonstrates specific labeling of the gold conjugates (x21.000). (Reproduced from Kruger er al., the Journal of Cell Biology, 1991, Vol. 115, p. 103, by copyright permission of the Rockefeller University Press and by courtesy of K. Wang.)

PROTEINS IN STRIATED MUSCLE

91

tion showed that there was approximately 10 times as much filamin per actin in ALD muscle as in pectoral muscles. The functional significance for this difference in amount and location of filamin in avian fast and slow muscles is unknown. 7. Zeugmatin

Monoclonal antibodies prepared with a purified fraction of fascia adherens domains of intercalated disks from chick cardiac muscle included one monoclonal antibody that labeled Z lines in adult skeletal and cardiac muscle and labeled intercalated disks in adult cardiac muscle (Maher et al., 1985). Immunoblots indicated that this mAb stained two bands with molecular weights of more than 500 kDa and several bands with lower molecular weight. Maher et al. (1985) named this protein zeugmatin. In cultures of chick pectoral muscle cells, 18% of the round myoblasts found early in culture stained positively for titin but did not stain with antibodies to zeugmatin or to MHC (Colley et al., 1990). Round myoblasts later expressed zeugmatin, titin, MHC and a-actinin, all of which were localized in a perinuclear punctate distribution. Bipolar myoblasts consistently stained positively with antibodies to all three proteins. This may indicate that zeugmatin has a less important role in myofibrillar protein organization than titin does.

8. Cap ZIPActinin Cap Z is an actin-capping protein that consists of 36- and 32-kDa subunits (Casella et af., 1986). Cap Z is localized by immunofluorescence in the Z lines of both isolated skeletal muscle myofibrils and frozen sections (Casella et al., 1987). Immunogold localization by electron microscopy of frozen sections confirmed this localization. When antibodies to Cap Z were used to label S 1-decorated actin filaments, the protein A-colloidal gold probe was predominantly located at the barbed end (Casella et al., 1987). p-Actinin had previously been described as an actin-capping protein that was located, both by immunofluorescence (Maruyama et a f., 1977) and by immunogold labeling (Funatsu et al., 1988), at the pointed end of actin filaments. A comparison of properties of purified p-actinin with Cap Z (Maruyama et al., 1990) showed that the two proteins had identical subunit molecular weights and, in addition, the amino acid sequences of two peptides from p-actinin were identical to portions of the Cap Z p subunit deduced from cDNA sequences (Caldwell et al., 1989). The finding by Maruyama et al. (1990) that purified p-actinin capped the barbed or Zline end of actin filaments led to the concession that p-actinin and Cap Z

92

MARVIN H. STROMER

are the same protein. A monoclonal antibody to the large a subunit of Cap Z labeled only Z lines of chicken breast muscle (It0 et al., 1991). 9. Spectrin, Vinculin, and Talin

Antibodies against chicken erythrocyte a-spectrin have been localized at or near the sarcolemma in skeletal and cardiac muscle (Repasky et al., 1982). The P subunit of chicken erythrocyte spectrin has been localized in a gridlike network under the sarcolemma of skeletal and cardiac muscle cells (Nelson and Lazarides, 1983).Appleyard et af. (1984)produced mAbs to human spectrin subunits and found that six of seven mAbs to P-spectrin bound at the cytoplasmic surface of the sarcolemma, but none of the mAbs to a-spectrin was bound. In fast-twitch avian pectoralis major muscle fibers, a-spectrin was localized primarily under the sarcolemma (Menold and Repasky, 1984),but slow-tonic ALD muscle fibers showed localization both at the sarcolemma and throughout the sarcoplasm. Another aspect of this study indicated that slow muscle had a threefold greater concentration of a-spectrin relative to other myofibrillar proteins. Dystrophic chicken pectoralis major muscle contained areas in the sarcoplasm that stained intensely with antibodies to a-spectrin (Repasky et af., 1986) and, in addition, staining at the sarcolemma was increased so that the spectrin network usually seen there was obscured. Most sarcoplasmic reticulum vesicles isolated from dystrophic muscle stained with antibodies to aspectrin, but only a small fraction of normal vesicles were labeled. Spectrin, together with y-actin, colocalized with vinculin, a 130-kDa actinbinding protein, in costameres, which are frequently seen as rib-like bands between avian skeletal myofibril I bands and the sarcolemma (Craig and Pardo, 1983; Pardo et al., 1983a). Shear and Bloch (1985) used immunofluorescence to show that vinculin was located at neuromuscular and myotendinous junctions in avian latissimus dorsi muscles. They also used immunogold to localize vinculin in subsarcolemmal electron-dense patches in tonic fibers of the ALD muscle. Vinculin has also been localized at the inner surface of the polysynaptic sarcolemma of neuromuscular junctions in mouse diaphragm and snake muscle (Yorifuji and Hirokawa, 1989). Confocal microscopy has shown that, although vinculin immunoreactivity is most prominent at the periphery of the cell, a reduced amount of vinculin also in present within the Z line of skeletal muscle cells (Terracio et al., 1990). Koteliansky et af. (1985) found vinculin in the fascia adherens of the cardiac intercalated disk and between the external myofibrils and the sarcolemma in cardiac muscle. Antibodies to collagen and collagen adhesion proteins were localized on the exterior surface of cardiac myocytes, directly opposite the intracellular location of both vinculin and talin (Terracio et al., 1989). This relationship suggests a potential linkage

PROTEINS IN STRIATED MUSCLE

93

between the extracellular matrix and the intracellular cardiac cytoskeleton that may involve vinculin and/or talin. Talin is a 225-kDa protein that can bind to vinculin and also is present in the finger-like muscle cell processes that extend into tendons at myotendinous junctions (Tidball et al., 1986).

C. Other Structural Proteins Associated with the Sarcornere

1. Titin Although titin is closely associated with the A band, there is conclusive evidence that titin spans the distance from the Z line to near the M line. For this reason, titin is discussed here rather than with the proteins associated with the A band. Titin is a huge protein, sometimes called connectin (Maruyama et al., 1981), that is usually present as a doublet, T1 and T2, on low-porosity SDS gels. TI is not extractible and has a molecular weight of 28003000 kDa. T2 is believed to be a proteolytic fragment that can be purified under native conditions and has a molecular weight of 2400-2600 kDa. (For recent reviews on titin, see Trinick, 1991; Fulton and Isaacs 1991.) Early immunofluorescence labeling with a mixture of antibodies to T1 and T2 showed heavy labeling at the A-I junction and at the center of the A band and a weaker, more diffuse staining throughout the remainder of the A band (Wang et al., 1979). The labeling at the A-I junctions remained at that location at a variety of sarcomere lengths and suggested that titincontaining structures were associated with the ends of thick filaments. A similar labeling pattern has been observed by others (Ohashi et al., 1981; Ikeya et al., 1983; Maruyama et al., 1984; Itoh et al., 1988). Itoh et al. (1988), however, found that the titin epitope recognized by one of their mAbs remained at the edge of the A band during sarcomere stretching to 3.5 pm. The titin epitope recognized by a second mAb was in the I band near the edge of the A band and did move during stretching. Immunoelectron microscopy after labeling with these same mAbs confirmed that the first antibody labeled three stripes in the edge of the A band that did not move when sarcomeres were stretched to 3.5 pm. The second mAb labeled two stripes in the I band that moved during stretch. This behavior was interpreted to mean that titin filaments in the I band are more extensible than those at both edges of the A band. These same titin mAbs also showed that the A-band epitopes remained constant during stretch, release, and contraction to rest length of frog skinned fibers (Maruyama et al., 1989). The I-band epitopes not only moved during stretch, but also moved closer to each other in the I band during extreme shortening. Horowits et al.

94

MARVIN H. STROMER

(1989) found that movement of thick filaments in activated skeletal muscle was accompanied by movement of a titin epitope in the I band in the same direction. This implies that neither calcium nor cross-bridge activity affected elasticity of titin or binding of titin to thick filaments. K. Wang et al. (1991) were able to correlate the movement of an I band titin epitope with the yield point sarcomere length, thereby suggesting that titin is involved in producing resting tension. Pierobon-Bormioli et a/. (1989) observed that titin epitopes in the edge of the A band and one in the I band near the A-I junction did not move away from the M line at sarcomere lengths <2.8 pm. At longer sarcomere lengths, however, they detected a small movement of titin epitopes from the M line, suggesting some elasticity of titin filaments at the edge of the A band. Immunofluorescence also has demonstrated that titin is present only in fast and slow skeletal muscles and in cardiac muscle and is not present in smooth muscle (Ikeya et al., 1983). Immunoelectron microscopy has advanced our understanding of the location of titin in skeletal muscle. Immunogold labeling showed that titin labeling was concentrated at the A-I junction, extended into the A band, and was also seen associated with superthin filaments at the ends of isolated native mouse thick filaments (Gassner, 1986). Monoclonal antibodies to titin localized titin molecules from near the M line to beyond the ends of the thick filament, a distance of about 1 pm (Whiting et al., 1989). Identification of gap filaments in the space between A and I bands as titincontaining filaments has been done in stretched beef muscle (LaSallle et al., 1983) and rabbit muscle (Wang, 1985). A polyclonal antibody to titin produced five stripes of binding in the I band and Z line (Maruyama, Koscak, et al., 1985). Furst et a / . (1988) used 10 mAbs to titin to map 10 nonrepetitive epitopes in the sarcomere. These studies showed convincingly that antibodies specific for TI bound at or close to the Z line and that the remaining antibodies bound to epitopes in the I band, several near the A-I junction, and in the A band to within 0.2 pm of the M line (Fig. 4). Titin filaments were thus shown to extend from the Z line to near the center of the sarcomere. Sarcomeres fractured at the A-I junction contained thin filaments that retracted to the N1 line, suggesting either that titin filaments do not interact with actin filaments except at the N1 level or that titin has an inelastic domain near the Z line (Trombitas et al., 1990b). Three mAbs have been produced that bind to repetitive titin epitopes that are spaced 42-43 nm apart in the A band (Furst et a / . , 1989a). These epitopes are in a zone between 140 and 440 nm from the M-line center and coincide either closely or exactly with the location of C protein and the 86-kDa protein. A fourth mAb in this set binds to an epitope at both edges of the M line 55 nm from the center of the M line, indicating that titin molecules penetrate the M line. The antibody binding sites for two of these mAbs have been identified on isolated T2 molecules and indicate that the head of T2

PROTEINS IN STRIATED MUSCLE

95

FIG. 4. Relaxed chicken pectoralis muscle labeled first with four monoclonal titin antibodies (a-d) and then with sheep anti-mouse immunoglobulin to amplify the signal. The antibodies used were (a) T20, (b) T21, (c) T22, and (d) T23. The mixture of two other monoclonal antibodies T3 and T12 (e) represents the sum of the two individual staining patterns. A control muscle treated only with the second antibody is shown in (f) ( x 18,600). Bar, 500 nm. (Reproduced from the Journal of Cell Biology, 1988, Vol. 106, p. 1568, by copyright permission of the Rockefeller University Press and by courtesy of D. Fiirst.

is part of the M-line anchoring domain (Nave et al., 1989). A titin epitope located 0.74 pm from the center of the A band was lost when the Cterminal residues of titin were removed with carboxypeptidase Y, but a second epitope 0.69 pm from the center of the A band was not affected (Wang, S.-M., et al., 1991). This also indicates that the C-terminus of titin is located at the Z-line end of the molecule and the N-terminus is located at the M-line. The possible role of titin in sarcomere assembly has been investigated in several laboratories. Hill et al. (1986) found that titin antibodies bound to the A-I junction in nascent, maturing, and mature myofibrils and that titin and MHC synthesis are tightly coupled events in cultured chick embryo breast cells. Titin-positive cells were, however, detected before positive staining for MHC and zeugmatin could be detected in cultured chick embryo cells (Colley et al., 1990). In cultured rat skeletal myoblasts and developing myotubes, a titin mAb produced diffuse and/or fibrillar fluorescence in myoblasts and immature myotubes and a periodic or doublet staining near the A-I junction in more mature myotubes (Handel et al., 1989). Because titin assembly on stress-fiber-like structures (SFLS) appeared before either a or y actin periodicity, Handel et al. (1989) sug-

96

MARVIN H. STROMER

gested that titin has a role in sarcomere organization. In the mouse embryo, desmin is expressed first, then titin, then a-actin and MHC (Furst et al., 1989b). At Gestation Days 11 and 12, immature myofibrils show a periodic distribution of titin epitopes that exist at or close to the Z line, but there is a random distribution of titin epitopes in the A band and at the A-I junction. At Gestation Days 13 and 14, all titin epitopes show the typical myofibrillar banding pattern. Furst et al. (1989b) also proposed that putative elastic titin filaments could act as integrators to unite filaments connected to the Z line with filaments of the A band. Recent studies have suggested that several invertebrates may contain analogs of titin in their skeletal muscles. Fibrillar muscles in both honeybees and Drosophila contain a protein named projectin that has a molecular mass >600 kDa and is localized on either side of the Z line in honeybee myofibrils (Saide, 1981) and between the Z line and the edge of the A band in Drosophila flight muscle (Saide et al., 1989). Nave and Weber (1990) isolated a 600-kDa protein from flight and leg muscles of the honeybee and the locust and the whole body of Drosophila. They named this protein mini-titin and localized it to the I-Z-I region and the edge of the A band: they indicated that, analogous to its vertebrate counterpart, mini-titin was a proteolytic component of a 700-kDa parent molecule. An 800-kDaprotein from lethocerus flight muscle was localized in connecting filaments that link thick filaments and Z lines in flight muscle and in A bands of leg muscle (Lakey et al., 1990). Preliminary DNA sequencing suggested that the insect 800-kDa protein was related to vertebrate titin and to nematode twitchin. Hu et al. (1990) isolated projectin from crayfish claw muscle and found that it had a molecular weight of 1200 kDa and cross-reacted with chicken breast muscle titin. Polyclonal antibodies to crayfish projectin also cross-reacted with honeybee projectin, strongly labeled I bands and H zones of honeybee flight muscles, and weakly labeled beetle flight muscle I bands.

2. Desmin and Vimentin Desmin is a 53-kDa protein that constitutes the 10-nm or intermediate filaments (IFs) in skeletal muscle cells (Stromer, 1990). At the molecular level, a fundamental building block of desmin IFs is the tetrameric protofilament, which consists of two antiparallel coiled coils, as shown by antibody binding at both ends of the protofilament rods (Geisler et al., 1985). The epitope for this mAb was located between residues 324 and 412 in the C-terminal part of the middle domain. Immunofluorescence was used by Lazarides and Hubbard (1976), Campbell et al. (1979), and Thornell et al. (1980) to demonstrate that desmin was located in a punctate pattern at the Z-line level in myofibrils and at the periphery of Z-line profiles in isolated

PROTEINS IN STRIATED MUSCLE

97

Z-line sheets (Granger and Lazarides, 1978). Vimentin, the principal component of 10-nm filaments in fibroblasts and in most cells of mesenchymal origin, is also present in developing muscle cells and has been reported to be colocalized with desmin at the perimeter of mature chicken skeletal muscle Z lines (Granger and Lazarides, 1979). Richardson et al. (1981) were the first to use anti-desmin labeling at the electron microscope level to identify the inter-Z line filaments in avian skeletal muscle as desmin IFs. Tokuyasu et a/. (1983) used immunoferritin labeling of ultrathin frozen sections to demonstrate that desmin is also present in intermyofibrillar spaces. An immunofluorescence study of vimentin and desmin in embryonic, postnatal, and adult chicken skeletal muscle showed that the two proteins were colocalized during embryonic development but, contrary to the results of Granger and Lazarides (1979), that the concentration of vimentin decreased and had virtually disappeared at hatching (Tokuyasu et a / . , 1984). Lateral registry of myofibrils preceded the Z-line localization of both these proteins. In a follow-up immunoelectron microscope study, Tokuyasu et a / . (1985a,b) found that desmin and vimentin coexisted along the entire length of 10-nm filaments and confirmed that some lateral myofibril alignment occurred before 10-nm filaments changed from a longitudinal arrangement to one associated with the Z line. Yagyu et al. (1990) used an immunogold labeling approach with mature porcine muscle and found that, although desmin IFs surrounded Z lines, some longitudinal IFs were also labeled near the periphery of the fibers. Both monoclonal and polyclonal antibodies have localized desmin in the postsynaptic domain of human neuromuscular junctions (Askanas et al., 1990). The question of what specific role of desmin- and/or vimentin-containing IFs have in myofibril assembly is still being actively debated. If desmin, for example, is to have a role in myofibril assembly, it must be present before or at the time assembly begins. Erickson et al. (1987) found that, in anterior portions of 54-hr quail embryos, the myotome was already stained with antibodies to desmin, and the myotome was intensely stained in the 4-day embryo. In the quail embryo, desmin clearly is present well before myofibril organization begins. Chicken myoblasts express desmin before they express skeletal myosin (Yablonka-Reuveni and Nameroff, 1990). The frequency with which these myoblasts expressed desmin was greatest among progeny of satellite cells from adult muscle and was least in very young embryos. Cranial somites in rat embryos at Day 12 were also desmin-positive (Bignami and Dahl, 1984). Rat and mouse embryonic prefusion myoblasts express desmin a few hours earlier than a-actin can be detected (Babai ef al., 1990). Rat embryo somite premyoblasts expressed only vimentin, type 1 myoblasts expressed vimentin and desmin, and type 2 myoblasts expressed desmin and a-actin (Babai et al., 1990). These results again indicate that, in the rat and the mouse, desmin is

98

MARVIN H. STROMER

present very early in development, even before sarcomeric actin is present. Lazarides et al. (1982),however, concluded that vimentin was synthesized throughout chicken skeletal myogenesis in uitro, but that desmin was restricted to postmitotic fusing myoblasts and myotubes. They also hypothesized that the assembly of the Z line was regulated via developmental changes in IFs. Tao and Ip (1991) microinjected quail embryo cells with an antibody that reduced desmin phosphorylation by 75-80%. The injected cells did not fuse into myotubes by 24 hr postinjection, but the uninjected myogenic cells did fuse. This suggests that the desminhimentin cytoskeleton is involved in the capability of myoblasts to fuse and confirms that phosphorylation may affect the structure and/or properties of IFs. This also supports previous reports that indicated that phosphorylation of desmin or vimentin IFs caused disassembly and that phosphorylated subunits were unable to assemble. It is hoped that additional experiments will clarify the role of in uiuo phosphorylation of IFs. Schultheiss et al. (1991) have presented evidence that indicates that desmin IFs are not required for myofibril assembly. Antibodies to desmin and to vimentin showed that expression of a truncated desmin molecule caused a transformation of vimentin or desminhimentin filaments into spherical bodies. Myoblasts and myotubes that contained these spherical bodies retained the ability to assemble laterally aligned myofibrils. Cultured dysgenic mouse myotubes that have no contractile activity lack both organized myofibrils and a mature organization of the desmin network (Tassin et al., 1988). Restoration of contractile activity by coculturing with spinal cord cells produced cross-striated myofibrils, but no reorganization of the desmin I F network. Blocking contraction of normal myotubes with tetrodotoxin caused disorganization of myofibrils and an aberrant pattern of desmin distribution. Tassin et al. (1988) suggested that intermediate filaments are not directly involved in myofibrillar organization, but that contractile activity is required. The structural organization of desmin IFs has been investigated by using an antibody to a synthetic peptide that corresponds to residues 442-450 near the C-terminal tail of desmin (Birkenberger and Ip, 1990). This antibody labeled tetrameric protofilaments but would not label assembled IFs. 3. Paranemin and Synemin Paranemin (280 kDa) and synemin (230 kDa) are two intermediate filamentassociated proteins that are coexpressed and colocalized with desmin and vimentin in embryonic skeletal, cardiac, and smooth muscle cells (Price and Lazarides, 1983). Paranemin is not present in adult avian fast and slow skeletal muscles, and synemin is not present in adult avian cardiac muscle. More recently, synemin has been localized in mammalian muscle with

PROTEINS IN STRIATED MUSCLE

99

immunofluorescence in a punctate patter at Z lines (Bilak et al., 1991). Immunogold labeling indicated that synemin and desmin colocalized in mammalian skeletal and cardiac muscle.

4. Integrin Integrin, a family of membrane glycoproteins, is present in E l 2 avian limbbud muscle in random punctate foci (Bosyczko er al., 1989). By E17- 19, the distribution was uniform with increased density at sites along the sarcolemma. In avian fast-twitch muscle at 3 or more weeks after hatching, integrin was concentrated at acetylcholine receptors and at myotendinous junctions. In cultured cells, Bosyczko er al. (1989) also found integrin concentrated at acetylcholine receptors and at sites where myofibrils terminate at the sarcolemma. No colocalization with talin, vinculin, fibronectin, or lamin was observed. A mAb against the p subunit of chicken integrin also gave a strong fluorescent signal at the myotendinous junction (Swasdison and Mayne, 1989).

5. Clathrin A mAb specific for the clathrin heavy chain localized in a doublet with one element on either side of the Z line soon after multinucleated myotubes were evident (Kaufman et al., 1990). This same pattern was observed in cultures containing myotubes from newborn or adult rat skeletal muscle and from a myogenic cell line. The appearance of clathrin banding in myotubes preceded the appearance of titin, MHC, actin, and desmin and led Kaufman et al. (1990) to suggest that clathrin may have a role in sarcomere assembly. 6. Talin Talin is a 225-kDa protein that has been localized at myotendinous junctions, in other words, at the tapered end of chicken skeletal muscle cells where they form a junction with tendon collagen fibers (Tidball et al., 1986). In addition, very slight periodic bands of fluorescence were seen near the cell surface. Talin is a post synaptic component of rat diaphragm neuromuscular junctions and persists there for at least 6 weeks after denervation (Sealock et a/., 1986). Belkin et a/. (1986) found that talin was localized in costameres of skeletal and cardiac muscle and in the intercalated disc of cardiac muscle. Drenckhahn er a / . (1988) confirmed their previous observations of strong anti-talin staining of the myotendinous junction and a weak periodic stain along the sarcolemma of skeletal and cardiac muscle, but found no labeling of the cardiac intercalated disc.

100

MARVIN H. STROMER

It has been suggested that talin may have a role in cell adhesion to the extracellular matrix, but details of the location of talin at the intercalated disc remain to be established.

D. Use of Antibodies t o Investigate Abnormal or Pathological Aspects of Skeletal Muscle

1. Muscle Dystrophies This section has been included in abbreviated form to present some examples of the use of immunocytochemistry in abnormal muscle. No attempt has been made to review this topic completely. Abnormalities in dystrophin, the product of the Duchenne muscular dystrophy (DMD) gene, have recently been investigated with immunocytochemistry. In many instances, dystrophin is either absent or present in very small amounts and, thus, is labeled only very weakly at the intracellular surface of the sarcolemma. A 7-year-old female with DMD, however, exhibited a mosaic of normal and dystrophin-deficient fibers (Tachi et al., 1990). Four males diagnosed as having Becker muscular dystrophy presented a patchy or discontinuous anti-dystrophin staining pattern at the sarcolemma (Sunohara et al., 1990). Polyclonal antibodies to the Nterminal half of dystrophin produced similar staining patterns in normal and dystrophic human fetal myotubes (Ginjaar et al., 1990). A polyclonal antibody against the C-terminal portion of dystrophin stained only normal human fetal myotubes, suggesting that a truncated dystrophin was present in the dystrophic myotubes. For diagnostic use, Ginjaar er al. (1990) advised the use of both the N-terminal and the C-terminal dystrophin antibodies. Antibodies to dystrophin have also been used successfully on clonal muscle cultures derived from possible DMD carriers to determine the number of clones that are positive or negative for dystrophin (Miranda et al., 1989). Aberrations in other myofibrillar proteins in dystrophic muscle have also been studied with antibodies. Schiaffino et al. (1986) found that antibodies to fetal myosin were a useful marker in identifying regenerating fibers in DMD muscle. Monoclonal antibodies to fetal, adult fast, or adult slow MHC showed that, in DMD, fetal myosin increased with patient age because of muscle regeneration and that a subset of fast fibers (type IIb) was the first to degenerate (Webster et al., 1988).Adult chicken dystrophic muscle continued to express neonatal myosin, which may indicate that muscular dystrophy inhibits MHC gene switching that normally occurs during development (Bandman, 1985). Slow-tonic ALD muscles from normal and dystrophic chickens had nearly identical ratios of the two slow

PROTEINS IN STRIATED MUSCLE

101

isomyosins (Kennedy et al., 1991). A minor fast-twitch fiber population also was present, which doubled in number with overloading in the normal ALD but remained unchanged in the dystrophic ALD. The dystrophic fast-twitch patagialis (PAT) muscle had more fast myosin 3 and less fast myosin I than its normal counterpart. Overloading caused both normal and dystrophic PAT muscles to accumulate fast myosin 3 and the dystrophic PAT to repress slow myosin 1. Dystrophic human muscle has more of a protein that resembles P-spectrin associated with the sarcolemma than normal muscle does (Appleyard et al., 1984). Denervating dystrophic fasttwitch avian posterior latissimus dorsi (PLD) muscle had no effect on aspectrin content, but increased the a-spectrin content of normal PLD twoto threefold (Gregorio et af., 1988). Increased immunofluorescent staining under the sarcolemma was associated with increased content of a-spectrin. Anti-desmin staining has been used to identify regenerating fibers in DMD (Thornell et al., 1980), intermediate filament aggregates in a pelvic girdle-lower limb myopathy (Pellisier et al., 1989), and longitudinal extensions of desmin filaments that occurred in six males suffering from severe postexercise soreness of their thigh muscles (Friden et al., 1984). An antibody to a putative desmin (52 kDa) from normal end dystrophic hamster skeletal o r cardiac muscle stained at the Z line in normal myofibrils, but did not stain at the Z line in some dystrophic hamster myofibrils (Pollock and Atkinson, 1985). Although both titin and nebulin are stained normally in DMD muscle by mAbs (Furst et al., 1987), intermediate-type cells in biopsies from patients with different dystrophies stained with antibodies to both fast and slow TN-I (Dhoot and Pearce, 1984). In normal muscle, the slow and fast forms of TN-I are segregated in type I and type I1 cells, respectively. Myofibrils from chicken dystrophic skeletal muscle showed a much weaker generalized staining with antibodies to TN-C compared with the discrete A- and I-band staining seen in normal myofibrils (Wilson et al., 1978). Normal avian I segments stained with antibodies to TN-C showed a 384 2 7w periodicity when examined in the EM, but dystrophic I segments had no distinct periodicity after staining (Irish et al., 1977). a-Actinin has been shown to be a major component of rod bodies in nemaline myopathy muscle (Schollmeyer et al., 1972, 1973; Jockusch et al., 1980).

2. Rhabdomyosarcoma The quest for the most effective antibody to use for diagnosis of rhabdomyosarcoma (RMS) has led to a diversity of opinions. Antibodies to myosin were preferred to antibodies to myoglobin by Saku et al. (1985). Seidal et al. (1987) found that all well-differentiated tumors stained with antibodies to myoglobin, but that a desmin mAb gave the most consistent results.

102

MARVIN H. STROMER

Antibodies to a-actin (De Jong et al., 1987a; Skalli et al., 1988) were preferable to antibodies to desmin, but Osborn et al. (1986)found that the parallel use of desmin and titin antibodies was the most effective approach. Krenacs et al. (1990) found that antibodies to the sarcoplasmic reticulum calcium transport ATPase were useful on formalin-fixed paraffin sections. In experimental RMS induced in rats by a sarcoma virus, tumor cells contained predominantly embryonic and neonatal MHC and rarely contained adult MHC (Azzarello et al., 1987). If the sarcoma was induced by nickel, RMS cells contained embryonic and adult fast MHC, a minority contained adult slow MHC, and none contained neonatal MHC (Borrione et al., 1988). 3. Other Skeletal Muscle Abnormalities The extent of muscle necrosis in dermatomyositis (De Geeter et al., 1989) and the extent of skeletal muscle damage with fractures in trauma patients (Elgazzar et al., 1989) have been assessed with indium-11 1-labeled antibodies to myosin. Severely atrophic fibers in spinal muscular atrophy (Werdnig-Hoffman disease) contained fetal MHC; intermediate-sized triangulated fibers contained fetal and fast MHC or fast MHC alone; and hypertrophied fibers contained slow MHC or slow and small amounts of fetal MHC (Biral et al., 1989). Biopsies from patients with neurogenic muscle disease revealed the coexpression of multiple myosin isoforms in selected fibers (Sawchak et al., 1989). Muscle cells from a 9-year-old male expressed only the slow isoform of myosin (Biral et al., 1987). Impaired skeletal muscle relaxation that is aggravated by exercise has been associated with a deficiency of the sarcoplasmic reticulum calcium transport ATPase (Karpati et al., 1986; Danon et al., 1988). Cytoplasmic bodies in cytoplasmic myopathies may stain with antibodies to desmin (Osborn and Goebel, 1983), with antibodies to either actin or desmin, or with neither antibody (Schroder et al., 1990). Muscle biopsies from active elderly humans contained a much greater concentration of ring fibers that were identified by labeling with antibodies to desmin (Jakobsson et al., 1990). These ring fibers had increased amounts of actin and spectrin at the periphery. Serum from myasthenia gravis and thymona patients contained autoantibodies against titin (Aarli et al., 1990).

111. Localization of Proteins in Cardiac Muscle Cells

Emphasis in this section will be placed on information that is unique to cardiac muscle and that is not redundant with information from the skeletal muscle system.

PROTEINS IN STRIATED MUSCLE

103

A. Thick-Filament Proteins and A-Band-Associated Proteins

1. Myosin Some of the questions about cardiac myosin that have been addressed with immunocytochemistry include the following: Is cardiac myosin immunologically distinct from skeletal myosin? How many isom yosins or isoforms of myosin exist in cardiac muscle? How are they distributed in the various regions of the heart? Sartore (1981) found that antibodies to ventricular myosin would cross-react with slow skeletal myosin. Two distinct types of atrial MHC were detected by Gorza et al. (1982) in three populations of bovine atrial fibers. One of these atrial MHCs was antigenically related to ventricular myosin. Human atria also contain two distinct types of myosin, and human ventricles contain three types of myosin (Bouvagnet et a/., 1984). Dechesne et al. (1985) determined that the two atrial myosins differed in at least five antigenic determinants and shared at least two. One atrial myosin also differed in at least five determinants and shared two with ventricular myosin, but the other atrial myosin shared at least five and differed in at least two determinants. Monoclonal antibodies demonstrated that human atria contained two distinct P-MHCs, 01 and P2 (Tsuchimochi et a/., 1988), whose expression can be regulated by pressure overload and embryonic development. aMyosin heavy chain was localized in all myofibers of the sinoatrial (SA) node, the atrio-ventricular (AV) node, and in approximately half the myofibers of the ventricular impulse-conducting system in human myocardium (Kuro-o et al., 1986). Only a few ventricular myofibers labeled with antia-MHC. p-Myosin heavy chain was located in all myofibers of the AV node and the ventricular impulse-conducting system, but almost no fibers in the SA node contained p-MHC. Gorza et a / . (1986) found that a large number of SA and AV fibers in the bovine heart stained with antibodies to fetal MHC and that no other myocardial cells reacted with this antibody. Most nodal cells also were positive for a-MHC and a number were positive for P-MHC. This selected group of papers makes the point that cardiac myosin isoforms differ from those found in skeletal muscle and that the specialized regions in the myocardium differ in their myosin composition. Just as skeletal muscles do not contain only one fiber type, the specialized myocardial tissues also do not contain only one type of myosin. There is agreement, however, that there are two principle MHC isoforms, a cardiac-specific a-MHC and a p-MHC that also exists in slow-twitch skeletal muscle. In adult hearts of the larger mammals, including humans, a-MHC predominates in the atria and p-MHC predominates in the ventricles. In the rat heart ventricle, there are three isomyosins, designated V1, V2, and V3, that consist of dimers of a-Q-, a+-, and p-p-MHC, respec-

104

MARVIN H. STROMER

tively (Jin et al., 1990). Adult rat atria contain two isomyosins, A1 and A2.

2. Myosin Isoforms in Developing Heart Myofibers

a. H u m a n Myocardium A mAb to a-MHC stained all atrial myofibers from 14 weeks gestation to an adult, but no ventricular staining occurred from 14 to 18 weeks gestation (Everett, 1986). The majority of infant ventricles either did not stain positively for a-MHC or had a scattering of stained cells, which suggests that the a-MHC appears in ventricles about the time of birth. Antibodies to p-MHC stained all ventricular cells and, after 14 weeks gestation, stained a minority of atrial cells. Bouvagnet et al. (1987) found a similar pattern of MHC expression during embryonic development and that, after birth, many fibers express both a- and pMHC. Wessels et al. (1991b) have provided a detailed analysis of the spatial distribution of a-and p-MHC in developing human hearts from 3 1 to 60 days gestation.

6. Avian Myocardium The earliest myosin detectable in the embryonic chicken heart and the somatic myotome reacted with a mAb to ventricular MHC but not with polyclonal antibodies to fast-twitch or slow-tonic skeletal MHC (Sweeney et al., 1984). The first isomyosin in avian atria and ventricles was subsequently identified by Sweeney et al. (1985) as V3 (pp-MHC). As development advanced, atria began to express a-MHC and to repress p-MHC, but ventricles retained the p-MHC. Sweeney et al. (1987) found that reactivity of embryonic atria with the adult ventricular MHC antibody could not be detected after E6 and suggested that the chick heart, at E6, has a myosin isoform distribution similar to that of the adult. One mAb to chicken pectoralis muscle reacted first with the presumptive ventricle at stage 10 and later with all regions of the developing heart (Gonzales-Sanchez and Bader, 1984). A second mAb that reacted only with atrial cells did not bind until 1 day later (E2) or stage 15. De Jong et al. (1990) could first detect both atrial and ventricular MHC isoforms in stage-8 hearts that consisted of four pairs of somites. Cardiac contraction first occurred at stage 10 in cells that contained both isomyosins. From stage 14, there was a regional loss of one of the isomyosins in the atria and the other in the ventricles so that the isomyosin distribution characteristic of the adult could be formed (De Jong et al., 1987b). The transition from coexpression of both cardiac a-and p-MHC occurred during E12- 13 in the atria and during E12-15 in the ventricles (De Groot et al. 1989), which is considerably later than the transition by E6 observed by Sweeney er al. (1987). Evans et al. (1988) found, however, that at least three MHC isoforms were expressed in embryonic chicken atria and ventricles and

PROTEINS IN STRIATED MUSCLE

105

that, in the adult chicken, atria continued to express at least three MHC isoforms, ventricular impulse-conducting fibers expressed two MHC isoforms, and ventricles expressed one MHC isoform. A MHC specific to the conduction system was first detected at El7 in developing chicken hearts (Gonzales-Sanchez and Bader, 1985). A cytoplasmic myosin isoform was localized in the contractile ring of embryonic chick ventricular cells grown in culture (Conrad et al., 1991).Greaser et al. (1989) described strand-like “wispy” structures that stained positively with antibodies to myosin and that were located alongside SFLS in cultured chick cardiac cells. Schultheiss et al. (1990) observed that both striated and nonstriated myofibrils existed in cultured chick cardiac cells and that there was a variable association between the nonstriated myofibrils and fibril-like structures that stained with antibodies to MHC. Myosin heavy chain antibodies did not stain the nonstriated myofibrils per se.

c. Rat and Mouse Myocardium In 3-week-old rats, all cardiac cells stained with antibodies to V 1 (a-a-MHC) and nearly none with antibodies to V3 (p-p-MHC) (Samuel et al., 1983). In adult controls and in cultured cardiac cells, 50% of the cells reacted with antibodies to V1, 10% with antibodies to V3, and 40% with both antibodies. The myocardium from adult hypophysectomized rats contained cells that reacted only with antibodies to V1. All rat ventricular fibers expressed a-MHCs from 2 days after birth until the second month, when they disappeared in a small endocardia1 fiber population and in a few conductive fibers (Dechesne et al., 1987). P-Myosin heavy chains are also initially present in all cells but then disappear from epicardium to endocardium between the second and fourth weeks, except in conductive fibers. The P-MHCs reappeared during the second month and then were expressed in nearly all fibers. Fetal and adult rat atria expressed a-MHC, but fetal ventricles expressed predominantly P-MHC (Sweeney and Kelley, 1990). The observation by Sweeney and Kelley (1990) that adult rat ventricles expressed exclusively a-MHC differs from that made by Dechesne et a/. (1987) who found P-MHC in nearly all adult rat ventricular fibers. Monoclonal antibodies to embryonic skeletal MHC identified a similar MHC isoform in SA and AV nodes in the rat heart (Gorza et al., 1988b). This isoform is developmentally regulated because it is rarely detected in adult tissue and is also partly controlled by thyroid hormone.

d. Eflect of Thyroid Hormones on Cardiac Myosin Sartore et al. (1981) used an antibody to atrial MHC to follow the influence of thyroid hormone on rabbit ventricular fibers. Control ventricular tissue showed a variable staining pattern, but all ventricular fibers were stained after thyroxine treatment. Hypothyroidism caused by polythiouracil treatment caused the

106

MARVIN

H. STROMER

ventricular fibers to become unreactive or poorly reactive. Lin et af. (1989a) found that the different proportions of a-MHC in adjacent fibers produced a mosaic staining pattern in control adult rabbits with antibodies to a-MHC and that feeding propylthiouracil for 70 days caused the ventricles to express 100%@-MHC.The normal mosaic staining pattern returned when triiodothyronine was administered. Immunogold labeling with antibodies to a-MHC indicated that injecting triiodothyronine caused a 50% increase in labeling at the edges of the A band and a 40% decrease at the center of the A band (Wenderoth and Eisenberg, 1987). Under these conditions, there was a seemingly greater exchange of new myosin molecules at the ends of thick filaments. Rat atria that contained virtually all a-MHC accumulated nearly 5% of their myosin as the @ isoform if thiouracil was administered or if hypophysectomy was done (Samuel et af., 1986).Thyroxine reverses this effect. Dwarf mice develop an adult cardiac myosin phenotype if thyroxine is injected and do so more rapidly than in skeletal muscle (Pruliere et al., 1989). The @- to a-MHC switch was also induced in canine ventricular muscle and in cultured ventricular cells by administering thyroxine (Seko et al., 1990). It is well established that increased concentrations of thryoid hormones are associated with increased amounts of ventricular a-MHC and, conversely, that hypothyroidism is associated with increased amounts of ventricular @-MHC.

e. Effect of Growth in Culture on Cardiac Myosin Newly isolated adult rat ventricular myocytes were rod-shaped cells that all stained with antibodies to a-MHC and 70% stained with antibodies to @-MHC (Eppenberger et al., 1988).After the cells spread out, 100% stained with antibodies to @-MHC and 70% stained with antibodies to a-MHC. This indicates a change from an adult terminally differentiated state to a less-differentiated state in cultures. Zadeh et af. (1986) also observed that cultured chick embryonic atrial and ventricular cells showed a decreased reactivity with tissue-specific MHC antibodies as time in culture increased. The MHC isoform characteristic of the conductive fibers in vivo was never expressed in these cultures. Zadeh er al. (1986) suggested that a unique MHC isoform(s) may be produced in cultured cells and that cultured cardiac cells may not be able to maintain the intrinsic program of tissue-specific MHC expression. Bugaisky and Zak (1989) did not agree with this suggestion and found, instead, that cultured adult rat cardiac cells remain highly differentiated cells that retain many in vivo characteristics, including synthesis of only V1 (a-a-MHC). Klein et al. (1985) labeled cultured neonatal rat heart cells with an antibody to the 26-kDa MLC and observed the incorporation of this light chain during a 72-hr period into sarcomeres in both spontaneously beating and noncontracting cells.

PROTEINS IN STRIATED MUSCLE

107

3. MM-Creatine Kinase and Myomesin A unique feature of adult chicken hearts is the total absence of MM-CK and, because M bridges between adjacent thick filaments are composed of MM-CK, they have no visible M-line structure. This situation occurs because there is no transition from the embryonic isoform, BB-CK, to the muscle-specific MM-CK isoform (Schafer et al., 1985). Myomesin, however, is present in chicken cardiac muscle at the center of the A band. Embryonic chicken hearts transiently expressed MM-CK between E4 and El 1, mainly in areas that participated in formation of the impulseconducting system (Lamers et d., 1989). Microinjecting in uitro-generated MM-CK and M-CK/B-CK mRNA with SP6 RNA polymerase resulted in translation products that could be localized at the M line with immunofluorescence (Schafer and Perriard, 1988). The best reconstitution of M lines was obtained after microinjecting MM-CK mRNA. The hybrid protein made from the M-CK head and the B-CK tail did not bind to A bands, but the protein that contained the M-CK tail and the B-CK head was localized in the M line. This indicates that the carboxyl half of M-CK is responsible for interaction with thick filaments. Although mammalian hearts normally contain both MM-CK and myomesin, the expression and location of these proteins can be influenced by various factors. In differentiating bovine hearts, MM-CK and myomesin were detected by antibodies earlier in the AV bundle and bundle branch cells than in AV node cells (Forsgren et al., 1983). Studies of adult rat hearts and freshly isolated cells showed that MM-CK was located in the M line (Eppenberger ef al., 1988).After these cells spread out in culture dishes, the MM-CK immunostaining was lost, which suggested that there had been a shift to a less-differentiated state. Cryostat sections of canine left ventricle showed that the major myofibrillar sites of MM-CK were the entire A band and the Z line (Otsu et al., 1989). In addition, EM localization showed that MM-CK also was located at the sarcolemma, the SR, and the inner and outer mitochondria1 membranes. This location difference could be caused by the use of whole tissue versus the use by others of isolated myofibrils or cells or could be due to the detection method or to a species difference. 6. Thin-Filament Proteins and I-Band-Associated Proteins

1. Actin

In Syrian hamster embryos, the earliest sign of myofibril organization in embryonic heart cells was the presence of randomly arranged thick filaments and some 6-nm diameter filaments that labeled with antibodies to

108

MARVIN H. STROMER

actin near the sarcolemma (Hill and Lemanski, 1986). Myofibril formation in long-term cultures of adult rat heart cells involves an alignment of myofibrils with bundles of actin stress fibers, suggesting that the actin filaments could serve as an organizing scaffold for myofibril formation (Eppenberger et af., 1987). The a-vascular smooth muscle actin isoform was present in significant amounts in embryonic rat hearts and in somatic myotomal cells between Gestational Day 10 and 17 (Sawtell and Lessard, 1989). At the end of the first postnatal week, this actin isoform was absent from both cardiac and skeletal muscle cells. Adult rat ventricular cells and newly dissociated cells from that source also contained no a smooth muscle actin (Eppenberger-Eberhardt et al., 1990). After these cells had flattened and spread out in culture, a fraction of the cells had a smooth muscle actin in SFLS, but only a little a-actin was in newly formed myofibrils. These adult cardiac cells, therefore, behaved like fetal cells in their transient expression of a smooth muscle actin. In cultured cardiomyocytes, nonmuscle ( y ) actin and tropomyosin were organized with sarcomere-like periodicity in the early stages of sarcomere assembly (Handel et a/., 1991). The SFLS in these cells labeled with antibodies to muscle (a) actin and to tropomyosin. These results indicated that there was no exclusive compartmentalization of muscle and nonmuscle isoforms of several myofibrillar proteins during myofibrillogenesis in cultured avian heart cells.

2. Tropomyosin and Troponin Primitive chick cardiomyocytes in culture contain SFLS that stain positively for muscle tropomyosin, but sometimes stain disproportionately for muscle tropomyosin and a-actin (Wang et af., 1988). This suggests that the synthesis and/or assembly of these two isoforms into SFLS may not be stoichiometric. Purkinje fibers from young rabbits, pigs, and fetal lambs were stained in a cross-banded pattern with antibodies to cardiac tropomyosin and myosin between 3 and 7 days in culture (Canale et af., 1983). During embryonic development, chicken myocardium expresses cardiactype TN-T and -C and a TN-I that is antigenically similar to both cardiac and skeletal TN-I (Toyota and Shimada, 1981).Cultured cardiac myocytes express the same TN isoforms as in the embryo, but express adult isoforms if cultured with motor or sympathetic nerves, nerve extract, or in the nerve-conditioned medium (Toyota and Shimada, 1983).

3. a-Actinin, Vinculin, and Filamin Although vinculin is not considered to be a protein associated with thin filaments, the association of vinculin with a-actinin at the intercalated disk

PROTEINS IN STRIATED MUSCLE

109

and at cardiac cell-substrate contacts makes it logical to present these two proteins in the same section. By using a combination of cell rupturing, immunogold labeling, and freeze-etching of hamster cardiac cells, Isobe et al. (1988) were able to obtain three-dimensional information about the location of a-actinin. In addition to the expected labeling on the Z lines, a-actinin was also located on stellate foci where cytoskeletal filaments converged. a-Actinin was first detected in embryonic hamster hearts as a discontinuous layer under the sarcolemma and in association with filamentous arrays (Hill and Lemanski, 1986). Later in development, a-actinin was located in Z plaques at the sarcolemma and ultimately in both Z lines and intercalated disks. Localization in ultrathin frozen sections of chicken cardiac muscle showed that a-actinin was present in the Z line and near the fascia adherens of the intercalated disk (Tokuyasu et al., 1981). At the fascia adherens, double labeling showed that vinculin was located closer to the membrane than a-actinin. Cultured embryonic chick cardiac cells contained plaques that stained positively for both a-actinin and vinculin (Terai et al., 1989). These plaques are near areas of myofibril assembly at the ventral sarcolemma and overlap cell-to-substrate focal contacts. Rat heart myocytes growing in situ showed the predictable pattern of anti-aactinin staining in the Z line and anti-desmin staining at the Z-line periphery (Samuel et al., 1985). Cultured neonatal rat heart cells retained their antia-actinin staining at the Z line but anti-desmin staining shifted from the Z line to cables and random sarcoplasmic foci. Nine somite-stage chick hearts contained smaller a-actinin-positive dots within larger titin-positive spots (Tokuyasu and Maher, 1987b). Examination with the EM showed that the a-actinin dots were Z bodies that were narrower than the nascent myofibril. This size relationship suggested to Tokuyasu and Maher (1987b) that Z bodies stabilize filaments for the assembly of myofibril. Koteliansky et al. (1985) also have localized vinculin in the intercalated disk and between the peripheral myofibrils and the sarcolemma. Filamin is located in the intercalated disk and at the periphery of the cardiac Z line (Koteliansky et al., 1985, 1986). 4. Caldesmon

This 141-kDa actin-binding and calmodulin-binding protein is present in cardiac muscle in amounts equal to those in skeletal muscle (Ngai and Walsh, 1985). The precise location of caldesmon in cardiac muscle has not been determined. 5. Gelsolin

Rouayrenc et al. (1984) isolated a 92-kDa protein that had all the properties of gelsolin and found that antibodies to this protein localized in the I bands

110

MARVIN H. STROMER

of cardiac myofibrils. They concluded that gelsolin was a myofibrillar protein. Carron et af. (1986) also isolated a 90-kDa protein that had all the properties of gelsolin and prepared antibodies to it. When affinity-purified polyclonal antibodies to gelsolin, to a closely related 92-kDa protein, brevin, and brevin-free IgG and IgA monoclonal antibodies were used, Carron et af. (1986) found no pronounced I-band staining. Instead, they found a diffuse distribution in muscle cells. Because of the very similar properties shared by brevin and gelsolin and because brevin can be a contaminant in antibody preparations, there seems to be obvious potential for uncertainty in the respective localization of these two proteins. 6. Spectrin Antibodies to a spectrin-like protein produced a diffuse staining reaction that was not localized in any particular part of the cell (Goodman et af., 1981). Repasky et af. (1982) observed an intense labeling at or near the sarcolemma in cardiac muscle but no sarcoplasmic staining. Nelson and Lazarides (1983)found that fluorescence with antibodies to P-spectrin was located in a grid-like network at the inner surface of the sarcolemma and that this grid-like pattern was less regular in cardiac than in skeletal cells. Antibodies to mammalian and avian erythroid spectrin and to mammalian brain spectrin recognized a 240-kDa a-spectrin subunit found in hamster hearts and localized in Z lines and intercalated disks in hamster cardiac muscle (Messina and Lemanski, 1989). Immunofluorescent staining of the Z-line region was interpreted as being due to the association of spectrin with T-tubules located near the Z line rather than direct association of spectrin with the Z line. In addition to fluorescent labeling at the Z-line level, at the sarcolemma, and at the intercalated disk, Thornell et af. (1984a)found that antibodies to a 230-kDa spectrin-like protein also faintly labeled longitudinal strands in human ventricular muscle.

C. Other Structural Proteins Associated with the Cardiac Sarcomere

1. Titin Although it is well established that titin is a protein associated with the A band and that it remains attached even to isolated thick filaments (Hill and Weber, 1986), the equal certainty that titin filaments extend from the Z line to near the M line dictates its inclusion in this section. A mAb specific for bovine cardiac titin produced a fluorescent band at each A-I junction (Wang and Greaser, 1985). The distance between successive bands in

PROTEINS IN STRIATED MUSCLE

111

adjacent sarcomeres depended on sarcomere length. Disrupting the M line

or the middle of the A band caused a significant amount of the titin antibody labeling to shift toward the Z line. Shortening the A band by removing myosin molecules from the ends of thick filaments caused the titin antibody staining to move toward the M line. Hill and Weber (1986) used three mAbs to titin to demonstrate that there are both species differences and differences between cardiac and skeletal muscle in the staining with these antibodies. The three antibodies, T I , T3 and T4, were prepared by using a chicken heart residue that contained titin as an antigen, and each recognized a different epitope. TI and T4 stained in the I band in chicken heart myofibrils, but T3 stained at the A-Ijunction. All three antibodies stained at the A-I junction in chicken skeletal muscle. Frozen sections of rat cardiac muscle did not stain with T1 or T4, but T3 did stain rat cardiac sections. Rat skeletal muscle stained with all three antibodies. Several studies have investigated the chronological appearance and location of titin in developing cardiac muscle. Tokuyasu and Maher (1987a) found that titin was present both in a diffuse form and in spots in the 4- to 7-somite premyofibrillar stages. In chick hearts at the 8- to 1 1-somite stage, the number of titin spots decreased while the number of myofibrils with periodically aligned titin spots increased. Double labeling with antibodies to titin and a-actinin showed that, at the 7-somite stage, chick hearts contained titin but no a-actinin; at the 9-somite stage, larger titin spots coincided with smaller a-actinin spots; and at the 10-somite stage, titin lines coincided with a-actinin lines in widened myofibrils (Tokuyasu and Maher, 1987b). Chick cardiac cells at the 10-somite stage contained Factin strands that also stained positively with anti-titin (Tokuyasu, 1989). These actin strands were identified with the phalloidin approach, which labels F-actin but cannot discriminate between actin isoforms. The SFLS in cultured chick cardiac cells stained positively for the a-actin and muscle tropomyosin isoforms (Wang et al., 1988). The SFLS centrally located in myocytes developed sarcomeric titin patterns before or coincidently with the sarcomeric periodicity of either a-actin or muscle tropomyosin. Myosin assembly into sarcomeric repeats was linked with the appearance of titin staining in developing sarcomeres. Komiyama et al. (1990) found a somewhat different sequence of protein appearance in cultured embryonic chick cardiomyocytes from that described by the Tokuyasu group. In premyofibrillar stages, Komiyama et al. (1990) used immunofluorescence to detect the I-Z-I proteins, in other words, a-actinin dots and diffuse actin and TN-C antibody labeling. At a later stage, titin and myosin dots that had nearly identical distributions were detected in these I-Z-I areas. Still later, nonstriated fibrils were present that were positive for a-actinin and TN-C but unreactive or weakly reactive for titin. The a-actinin dots along these fibrils became aggregated to form Z lines at about the same time that

112

MARVIN H. STROMER

titin and myosin exhibited a sarcomeric pattern. It seems likely that the cultured chick cardiac cells studied by Komiyama et al. (1990) synthesized titin at a later developmental stage than the embryonic chick cardiac cells did that were studied by the Tokuyasu group. Schaart et al. (1989) used antibodies to detect titin at 8.25 days after coitus in embryonic mouse heart anlage when no desmin was present. Titin was present in a punctate pattern similar to that observed in embryonic chick hearts by Tokuyasu and Maher (1987a).

2. Desmin and Vimentin Immunofluorescent localization in isolated heart cells showed that desmin was located between Z lines of adjacent myofibrils and at areas of membrane interaction between laterally associated cells (Lazarides and Hubbard, 1976). Cultured chick or rat cardiac cells and frozen sections of cardiac muscle from chicken or rat also showed that desmin was located in a filamentous network and, in rat cardiac muscle, was localized at the intercalated disk and the Z line (Campbell et al., 1979). Isolated adult rat heart cells that retained their rod shape also retained desmin localization at the edge of the Z line and in close association with the sarcolemma (Saetersdal et al., 1989). If these cells contracted into square or round shapes, this organization of desmin was completely absent, suggesting that these desmin contacts were broken. Cultured neonatal rat heart cells lost both lateral myofibrillar registry and anti-desmin staining from the Zline area (Samuel et a!., 1985). This was interpreted to mean that desmin binding to Z lines is correlated with myofibril registry. In immature cultured chick cardiac cells, Dlugosz et al. (1984) saw no anti-desmin staining at the Z line or at the sarcolemma and also failed to find 10-nm filaments between adjacent Z lines with the EM. Instead, numerous longitudinal 10nm filaments were observed. Although Dlugosz et al. (1984)argued against an essential role for desmin in assembling Z lines, the fact that in more mature myofibrils, Z lines were associated with anti-desmin staining leaves open to interpretation their observations on desmin as an intermyofibrillar linker. Desmin filaments in normal or hypertrophied (aortic stenosis or thyroxine injection) rat hearts were first reported to be unchanged (Samuel et al., 1984; Rappaport et al., 1985). Subsequently, Watkins et al. (1987) found that, in hypertrophied rat heart, desmin filaments were also longitudinally oriented to link two out-of-register Z lines and were located along digitations of the intercalated disk connecting two neighboring desmosomes. Kartenbeck et al. (1983) used immunogold labeling to show that desmin IFs were attached either laterally or terminally (end-on) to desmosoma1 plaques in the intercalated disk in cultured rat heart cells. Kartenbeck et al. (1983) also found that rat heart cells contained no cytokeratin

PROTEINS IN STRIATED MUSCLE

113

filaments. Schaart et al. (1989) used immunofluorescence to detect desmin at 8.25 days after coitus in the mouse ectoderm, where it was transiently expressed with vimentin and keratin. Desmin, vimentin, and keratin were expressed by heart muscle cells from 8.5 to 9.5 days after coitus, with desmin present in a striated pattern at 9.5 days after coitus. In the Syrian hamster heart, desmin immunostaining was associated with myofibrils at E9, with Z lines in the newborn, and with intercalated discs 3 days after birth (Osinska and Lemanski, 1989). A mAb to vimentin stained Z lines in single heart cells isolated from 9- to 12-week-old rats (Zernig and Wiche, 1985). The location of desmin in chicken heart cells has also been extensively studied. Tokuyasu (1983a) used ferritin labeling of cryosections to show that longitudinal networks of desmin filaments surrounded chicken cardiac myofibrils and that transverse desmin networks surrounded Z lines. Sugi (1989) used gold labeling with desmin antibodies to identify the 12- to 14nm filaments that formed a loose network around myofibrils in E3 chick hearts. Before aortic-pulmonary (AP) septation in embryonic chicken hearts at E4, anti-desmin staining of the AP septum anlagen was observed (Sumida et al., 1987). During septation on E5, a more intense anti-desmin staining was present and suggested that cells containing desmin may have a function in septation. Two mAbs to desmin equally recognized desmin IFs in adult chicken hearts, but differed in their staining of embryonic chicken heart cells (Danto and Fischman, 1984). One of the mAbs stained all heart cells during development, but the other was unreactive until myofibrils were laterally aligned. It was determined that this differential response was not caused by the presence of a new desmin isoform o r by post-translational modification of an existing protein and, therefore, was assumed to be caused by unmasking of an epitope by a protein associated with intermediate filaments (Fischman and Danto, 1985). Virtanen et af. (1990), however, have produced an antibody that reacts only with desmin in bovine Purkinje fibers and not with desmin from other muscle cells. This result suggests that there are cell-type-specific antigenic determinants in desmin IFs. The quick-freeze deep-etch method of sample preparation combined with immunogold localization has also been used on cultured o r embryonic heart cells. Isobe et al. (1991) found that polyclonal desmin antibodies heavily and continuously stained some individual filaments between myofibrils in cultured hamster heart cells, but left most filaments, myofibrils, and membrane structures unlabeled. Monoclonal antibodies to desmin labeled some individual filaments, with labeled regions interspersed with bare regions along the filament. Monoclonal antibodies to vimentin produced a similar staining pattern, except that bare regions were longer on the intermyofibrillar IFs. The IFs in fibroblasts in the same cultures were

114

MARVIN H. STROMER

continuously stained by the vimentin antibody. Sugi and Hirakow (1991) observed that the 12- to 14-nm filaments that formed a loose network around myofibrils were attached to myofibrils, and some of them converged into Z-line regions in embryonic chicken heart. Immunogold staining identified these filaments as desmin IFs. Human fetal hearts, at mid-gestation, had moderate anti-desmin fluorescence at the Z line in ventricular and atrial myocytes and in proximal parts of the conduction system (Forsgren et al., 1982). Peripheral parts of the conducting system stained intensely, indicating a greater desmin content. Normal and pathological human ventricular myocardium had identical localization of desmin in an intermyofibrillar lattice at the Z line and at the intercalated disk (Thornell et al., 1984b). These results from human hearts support those from other species and indicate that desmin is located transversely between Z lines of adjacent myofibrils, between peripheral myofibrils and the sarcolemma, at the intercalated disk, and longitudinally between myofibrils. As was true with skeletal muscle, there are no data to show that desmin is in the Z line per se. 3. Vinculin and Talin Vinculin was first localized in ultrathin cryosections in the fascia adherens (FA) of chicken cardiac intercalated disks and was closer to the sarcolemma that a-actinin (Tokuyasu et al., 1981). Immunofluorescence observations of cryostat sections of chicken and rat hearts revealed that vinculin was located in the intercalated disk in spots periodically distributed at the sarcolemma, and over the Z-line region of the peripheral myofibrils (Koteliansky and Gneushev, 1983). Glycerinated myofibrils, however, did not react with antibodies to vinculin, which suggested that vinculin was not an integral myofibrillar protein. Pardo ef al. (1983b) called these vinculincontaining rib-like bands between the sarcolemma and the peripheral myofibrils, costameres. They also found vinculin associated with T-tubules in bovine atrial, ventricular, and Purkinje fibers. Rogalski (1987) also found vinculin, together with a 130-kDa sialoglycoprotein (Sgp), in sarcolemma dense plaques at the cardiac Z line. The FA contained vinculin, but the Sgp was absent. Belkin et al. (1986) colocalized vinculin and talin in costameres of chicken cardiac muscle and noted that vinculin staining was greater. Meta-vinculin also was located in the intercalated disc and costameres in human cardiac muscle (Belkin et al., 1988). In addition to finding vinculin in the intercalated disk and in costameres, Thornell et al. (1984b) found abundant vinculin in myotendinous junctions in human muscle. Neonatal rat cardiac cells reorganize their myofibrils and intercalated disks during the first 72 hr in culture. Fascia adherens could not be detected with antibodies to vinculin until after 48 hr in culture (Atherton

PROTEINS IN STRIATED MUSCLE

115

et al., 1986). The sarcomeric organization of myofibrils was stable if the myofibrils were attached to the intercalated disk and unstable if they were attached to non-disk regions. The recent finding, with laser confocal scanning microscopy, that vinculin in isolated cardiac cells from several species is most abundant at the cell periphery and is also present in lesser amounts within the Z line (Terracio et af., 1990) opens the question of where vinculin is located in the Z line. Vinculin and a-actinin were localized in plaques at the ventral sarcolemma in regions of myofibril assembly in cultured chick cardiac cells (Terai e f af., 1989). Increased vinculin labeling was seen where zonula adherens were converted to fascia adherens in 6- to 12-somite chick embryonic hearts (Tokuyasu, 1989).

D. Use of Antibodies t o Investigate Abnormal or Pathological Aspects of Cardiac Muscle

1. Human Transplant Rejection Right ventricular endomyocardial biopsy has been the traditional method of diagnosing rejection. The interest in developing a noninvasive method of detecting the rejection process led to the use of indium-11 l-labeled Fab fragments of antibodies specific for cardiac myosin. Intravenous injection of the labeled Fab fragments is followed, after an appropriate interval, by planar imaging or single-photon emission-computed tomography (SPECT). Scintigrams localize the uptake of the labeled antibody in myocardial areas undergoing transplant rejection or that have been damaged irreversibly by myocarditis or ischemia. For a review on the use of nuclear cardiology techniques, see Zaret and Wackers (1991);for results of clinical trials, see Johnson et af. (1989). Frist et a / . (1987) compared the indium111 FAB approach with the biopsy method on 18 patients and found the indium-11 1 approach to be 80% accurate. Ballester-Rodes e f a / . (1988) quantitated the uptake of indium-1 1 1 by using a heart-to-lung ratio and found that this was useful in monitoring the clinical course of transplant acceptance or rejection. De Nardo et al. (1989) also compared 30 scintigraphic studies after indium-111 Fab injection with biopsy results on 10 heart transplant recipients. Nineteen images were negative, and no false negatives were detected. On the basis of the 11 positive images, De Nardo et a / . (1989) indicated that biopsy was required to prepare a definitive diagnosis. Johnson and Cannon (1991) found a better correlation between indium-11 1 uptake and degree of transplant rejection in animal studies than in human studies. They postulated that this may be due to sampling error in the endomyocardial biopsy. Limitations of the indium-1 1 1 antimyosin method include slow blood clearance, long half-life of indium-11 1, and hepatic uptake (Johnson and Cannon, 1991).

116

MARVIN H. STROMER

2. Animal Models of Transplant Rejection Nishimura et al. (1987)found that uptake of indium-11 1 myosin antibodies was significantly greater in moderate and severe cases of myocardial rejection in the dog and that specific localization was evident in damaged areas. After heterotopic heart transplants in 56 rats, Allen et al. (1989)found that donor heart uptake of indium-11 1 myosin antibodies was significantly greater in both moderate and severe rejection than in lesser degrees of rejection.

3. Human Myocardial Infarct Damage Technetium-99m-labeled Fab fragments of myosin antibodies identified areas of myocardial necrosis in 90% of patients, which was virtually identical with results obtained with pyrophosphate (Khaw et af.,1986). Infarct size determined by pyrophosphate SPECT was 1.7 times larger than infarct size determined by myosin antibody SPECT. Johnson and Seldin (1989) and Antunes et al. (1989) made use of indium-1 ll-labeled Fab fragments of myosin antibodies to obtain diagnostic and/or prognostic information from patients. Correlation between images obtained after administering myosin antibodies labeled with indium-11 1 to a living patient and autopsy evaluation showed that the labeled myosin antibodies are specifically bound to irreversibly damaged cells and that this uptake can be seen even if the label is injected 6 days after infarction (Jain et al., 1990). Matsumori et al. (1990) studied 35 patients with myocardial infarctions (MIS), 5 with myocarditis, and 3 with angina and found that indium-11 1-labeled Fab fragments of myosin antibodies were useful in diagnosing coronary disease and myocarditis.

4. Animal Models of Infarct Damage Experimental MI in the dog has been studied extensively. Khaw et af. (1976) found a greater uptake of iodine-125-labeled (Fab’)2 fragments of myosin antibodies in infarcted areas than if iodine- 125-labeled myosin antibodies were used. Uptake was greater 72 hr after MI than at 24 hr after MI. A better correlation between uptake of iodine-25 (Fab’)2 myosin antibodies and impaired regional blood flow was observed than when technetium-99m-labeled pyrophosphate was used (Beller et al., 1977). Sikorska et al. (1990) found that iodine-1 1 1 (Fab’)2 monoclonal myosin antibodies localized more specifically in infarcted dog myocardium than technetium-99m pyrophosphate did. The location of iodine-131 uptake in damaged cells in heart slices correlated well with scintigrams taken onehalf hour after injection of the label (Khaw et al., 1978). Because the

PROTEINS IN STRIATED MUSCLE

117

energy characteristics of iodine isotopes are not ideal for imaging with a gamma camera, Khaw et al. (1980) developed a method for coupling indium-] I 1 to Fab myosin antibodies. It is, therefore, somewhat surprising that the use of isotopes of iodine continues. Increased membrane damage during the first 45 min of reperfusion of dog hearts was demonstrated with iodine-125 or iodine-131 (Fab’)2 myosin antibodies (Frame et al., 1983). Some of the membrane damage could be prevented by altering the conditions of reflow. Hoberg et al. (1988) found that iodine-123 Fab fragments of myosin antibodies produced better scintigrams in experimental MI dogs because of faster clearance from the blood than iodine-123 myosin antibodies. Uptake of iodine- I23 myosin antibodies measured in infarcted zones in ventricular slices was slightly more than twice as great as the uptake of iodine- 123 Fab fragments of anti-myosin. Polyclonal antibodies to TN-1 were labeled with iodine-131 and were specifically localized in infarcted areas of dog hearts. These antibodies were bound in up to 24 times greater concentrations in necrotic than in normal myocardium (Cummins et al., 1990). Technetium-99m Fab fragments of mAbs to myosin were present in ratios of up to 30 to 1 in infarcted compared with normal myocardium if blood flow was not limited and in ratios up to 10 to I if blood flow was severely restricted (Khaw et al., 1984). The effect of MI or M1-like conditions on the myocardium of rats and rabbits has been studied by various approaches. Nolan et al. (1983)intravenously injected mAbs to myosin after occluding the left coronary artery in rats. Sections of the left ventricle were then stained with fluoresceinlabeled antibodies to mouse IgG. Ischemic zones were labeled, but nonischemic areas were not labeled. Control and anoxic Langendorf perfused rat hearts showed that, after 90 min of anoxia, there was diminished staining with antibodies to vinculin and a-actinin but relatively little change in desmin antibody staining (Ganote and Vander Heide, 1987). Williams et al. (1991) compared the tissue retention fraction of indium-I I 1 myosin antibodies with the amount of creatine kinase released from isolated, perfused interventricular rabbit septum after induction of tissue necrosis. They concluded that indium-1 I 1 myosin antibodies could be used to quantitate the amount of necrosis in this tissue.

5. Effects of Cardiac Hypertrophy and Pressure Overload on Human and Animal Hearts In normal human left atria, fibers staining for P-MHC were rare, but in hypertrophied left atria after mitral stenosis a large proportion of fibers contained 0-MHC (Gorza et al., 1984). Hypertrophied left ventricles in patients with mitral regurgitation showed a MHC pattern similar to that in the normal ventricles, in other words, P-MHC in all fibers and a-MHC in

118

MARVIN H. STROMER

a variable number of fibers. Subsequently, these researchers reported the same result in hypertrophied left atria in human, but found that ventricular hypertrophy in patients with aortic stenosis, systemic hypertension, or tetralogy of Fallot was characterized by an almost complete absence of fibers with a-MHC (Schiaffino et al., 1984). Tsuchimochi et al. (1984) agreed that, in pressure-overloaded human atria, the number of fibers with P-MHC increased as those with a-MHC decreased. Rappaport et al. (1986) found little or no shift in myosin isozymes in humans with chronic compensatory hypertrophy. Rats with aortic stenosis, however, shifted rapidly from a- to P-MHC 2 to 3 days after stenosis. Rappaport et al. (1984) used immunofluorescence to demonstrate that changes in the microtubule pattern occurred soon after induction of hypertrophy in the rat and that desmin distribution was unchanged. Overloaded canine right atria bound more p-MHC antibodies and less a-MHC antibodies compared with normal right atria, but the reactivity of sinoatrial node cells was unchanged (Nomoto et al., 1988). Komura et al. (1988) determined that it is the PlMHC that is highly induced in overloaded canine right atria. Normal right atria contain nearly no 01-MHC but express P2-MHC. The overall pattern in hypertrophied atria is a shift from a- to 0-MHC. Hypertrophied ventricles may respond with nearly no change in MHC isoforms or may lose nearly all a-MHC. 6. Animal Models of Cardiac Abnormality

a . The Mexican Axolotl, Ambystoma mexicanum This salamander possesses a mutant cardiac lethal gene that results in abnormal heart development and function in homozygous animals. Only certain regions of the heart, the conus and the upper ventricle, contract, but there is a beat-type rhythm. Myofibril organization is aberrant, and myosin antibody staining is less in mutants than in normal hearts (Lemanski et al., 1980). Mutant hearts have very little anti-tropomyosin staining (Starr et al., 1989). Antia-actinin staining is significant but is not located in the expected Z-line striations. The quantity of TN-T present in mutant and normal hearts was identical, but instead of the usual I-band location, it was located in amorphous collections at the cell periphery in mutant hearts (Fuldner et al., 1984). The intensity of staining with an antibody to cardiac actin was similar in mutant and normal hearts, but the sarcomeric pattern was not present in mutants (Starr et al., 1989). Immunogold and immunofluorescence labeling with desmin antibodies was diffusely distributed in mutants (Shen and Lemanski, 1989). 6 . Cardiomyopathic UM-X7.1 Syrian Hamster This hamster has been utilized as a model for chronic congestive heart failure. Larger-than-aver-

PROTEINS IN STRIATED MUSCLE

119

age cardiac muscle cells contain randomly oriented myofibrils (Li and Lemanski, 1990). Immunofluorescent staining in individual myofibrils for myosin, actin, and tropomyosin was normal (Lemanski and Tu, 1983). Originally, Lemanski and Tu ( 1983) reported that anti-a-actinin staining in cardiomyopathic cells indicated that Z lines were wide and irregular, but, in a more recent paper, Li and Lemanski (1990)found no abnormalities in individual Z lines. A shift from predominantly a-MHC toward the p isoform was detected on cryostat sections of mid-ventricles from the cardiomyopathic hamster (Jasmin et al., 1988).

c. Murine Viral Myocarditis Experimental myocarditis was induced in BALB/c mice by exposure to encephalomyocarditis virus (Matsumori et al., 1989). Iodine-125 and iodine-131 Fab mAbs to myosin were used to detect damage to the heart. Three days after virus inoculation, uptake of the label increased significantly and reached a maximum on Day 14, when lesions were most extensive and prominent.

IV. Sarcoplasmic Reticulum, Transverse Tubules, and the Sarcolemma

This section will include topics related to what are generally considered to be the more specialized membrane systems in both skeletal and cardiac muscle. The sarcoplasmic reticulum is the intracellular membrane system found in all muscle cells whose function is the regulation of intracellular free-calcium concentrations. Free-calcium concentrations in resting musM . When the stimulus for contraction cle are approximately to arrives at the cell, a cascade of changes is initiated that increases freecalcium concentrations to to lo-’ M , and muscle contraction is initiated. When the stimulus ends, calcium is pumped, against a concentration gradient, into the lumen of the sarcoplasmic reticulum, thereby lowering free-calcium concentrations to levels insufficient to support contraction. The specific location and the extent to which the transverse-tubule system is developed vary among different animals and, to some extent, differ among muscle types. The transverse-tubule system, or T system, is formed by invaginations of the sarcolemma and is seen as tubules perpendicular to the cell surface that protrude radially into the interior of the muscle cell. Physiological evidence has indicated that the T system, through its regularly spaced invaginations, rapidly conducts the stimulus for contraction from the sarcolemma into the interior of the cell. The lumen of the T tubules is located intracellularly but actually is continuous with the

120

MARVIN H. STROMER

extracellular space. The sarcolemma is the cell membrane located at the periphery of the muscle cell.

A. Sarcoplasmic Reticulum

1. Calcium-TransportATPase (Ca2+-ATPase) The calcium-transport ATPase is a 1 10-kDa intrinsic membrane protein of the SR. It is an integral part of the SR membrane and, as the name implies, is responsible for pumping calcium into the lumen of the SR by an ATPdependent process. Immunofluorescence has shown that the Ca2+-ATPase is distributed fairly uniformly in the SR membrane of rat skeletal muscle (Jorgensen et al., 1979) and that fast-twitch glycolytic (FG) fibers have greater amounts of the Ca2+-ATPasethan fast-twitch oxidative-glycolytic (FOG) fibers, which have more than slow-twitch oxidative (SO) fibers (Maier et al., 1986b; Krenacs et al., 1989). Nuclear chain fibers in adult rat, rabbit, and cat muscle spindles, which contract faster than nuclear bag fibers, also have greater concentrations of the Ca2+-ATPase(Maier et al., 1988~).In rat papillary muscle, Ca2+-ATPase was fairly uniformly distributed in the free SR membranes, but was absent from interior and peripheral junctional SR, T tubules, sarcolemma, and mitochondria (Jorgensen et al., 1982). A similar distribution was seen in rat ventricular muscle (Jorgensen and McGuffee, 1987). The CaZt-ATPase has been detected in developing chick hearts by the time the first contraction occurs, and the density and intensity of antibody labeling both increase as rate of embryonic cardiac contraction increases ( Jorgensen and Bashir, 1984). Monoclonal antibodies have been produced to the two Ca2+-ATPaseisoforms, one of which is specific for the enzyme in fast-twitch fibers, and the other recognizes the enzyme in both slow and cardiac fibers (Jorgensen et al., 1988). In developing chick skeletal muscle, all fibers expressed both isoforms, but primary fibers expressed mainly the slow/cardiac isoform, and secondary fibers expressed both isoforms at comparable levels (Kaprielian and Fambrough, 1987). Fiber-type-specific isoforms of Ca2'-ATPase and MHC are not coordinately expressed in chick skeletal muscle. Dulhunty er al. (1987) used a mAb that had the same binding affinity for both isoforms to show quantitatively that rabbit fast-twitch fibers had seven times greater concentrations of Ca2+-ATPasein the terminal cisternae membranes than did slow-twitch fibers. Chronic stimulation of a predominantly fast skeletal muscle caused all fibers to stain with mAbs to the slow fiber Ca2+-ATPaseisoform (Briggs et al., 1990). In control fast muscles, 80% stained with mAbs to the fast isoform and 20% stained with antibodies to the slow isoform. The content of Ca2'-ATPase was

PROTEINS IN STRIATED MUSCLE

121

significantly less in the SR from 68-year-old human subjects than from 28year-old subjects (Klitgaard et ul., 1989). Despite the decrease in Ca2+ATPase, there was no concomitant decrease in calsequestrin or of the ryanodine receptor. It has been reported that T-tubule membranes contain a Mg2+-ATPase that is a 102-kDa concanavalin-A-binding glycoprotein and that has a high degree of structural homology with the SR Ca2+-ATPase (Damiani et al., 1987). Comparative localization with well-characterized mAbs may provide information on the specific arrangement of these two ATPases. It has been proposed that the Ca2+-ATPaseconsists of 10 transmembrane a-helices and, on the cytoplasmic side of the SR membrane, a stalk composed of a-helices and a knob-like structure (Brand1 et ul., 1986; MacLennan, 1990). The smaller part of the knob is called the p-strand sector because it contains all p sheet structure. The other part of the knob has a phosphorylation domain and a nucleotide-binding domain. The larger part of the knob contains a trypsin-sensitive site, T,, between residues 505 and 506. A second trypsin-sensitive site, T,, exists in the p-strand sector between residues 198 and 199. Cleavage at T, produces a 57-kDa fragment called the A tryptic fragment from the N-terminus and a 52-kDa B tryptic fragment from the C-terminus. The A tryptic fragment can be cleaved at T, into a 23-kDa A, fragment and a 34-kDa A, fragment. Epitopes on these and smaller fragments have been utilized to attempt to localize specific regions and functions of Ca2'-ATPase. Matthews et al. (1989) produced antibodies to five segments of the enzyme, including the N- and C-terminal regions and found that, except for antibodies to a peptide from residues 567 through 582, all other antibodies bound strongly to intact isolated SR vesicles. This indicates that the other four segments are located on the cytoplasmic side of the membrane. An antibody to a peptide from residues 877 to 888 bound only to SR vesicles that had been detergent-treated, suggesting that this epitope may be in the lumen of the SR (Matthews et al., 1990). A battery of 28 mAbs recognized epitopes on the A, and B tryptic fragments (Colyer et al., 1989). Of these, 9 antibodies inhibited the catalytic activity of Ca2+-ATPasewhen bound to specific regions of the enzyme. Molnar et al. (1990) also used 14 monoclonal and 5 polyclonal Ca2+-ATPaseantibodies to determine the accessibility of epitopes in native SR vesicles. They found that 2 mAbs to the B-fragment produced nearly 50% inhibition of the rate of ATP-dependent Ca2+transport. 2. Calsequestrin

Calsequestrin is a 55- to 60-kDa SR protein that functions as a calciumbinding protein. It has a large number of medium-affinity calcium-binding sites and is capable of binding 40-60 mol of calcium/mol calsequestrin.

122

MARVIN H. STROMER

The dual function of calsequestrin as a specialized calcium storage site that also could release calcium when needed created a substantial interest in knowing where calsequestrin was located. Immunofluorescence showed that calsequestrin was more discreetly located near the A-I junction in rat skeletal muscle than the more widely disseminated Ca2+-ATPase( Jorgensen et al., 1979). Slow type I fibers contained less calsequestrin than type I1 fibers did (Maier et al., 1986b). Nuclear bag fibers of muscle spindles contained less calsequestrin than the faster-contracting nuclear chain fibers (Maier et al., 1988~).Immunoelectron microscope localization in rat skeletal muscle (Jorgensen et al., 1983), ventricular (Jorgensen and McGuffee, 1987), and atrial cells (Jorgensen et al., 1985) showed that calsequestrin was located in the lumen of both peripheral and interior junctional SR and in corbular SR that is located in the I band. In contrast to the rat, chicken ventricular cells have no T tubules, but the calsequestrin distribution in peripheral junctional and corbular SR in the I band is unaffected (Jorgensen and Campbell, 1984). Antibodies to an avian calsequestrin homolog, first described as a lamin-binding protein named aspartactin, stained slow- and fast-twitch skeletal muscle, cardiac muscle, and the cerebellum (Choi and Clegg, 1990). This homolog has a predicted sequence that is 70-80% identical to mammalian calsequestrin and is present in E5 limb primordia before myoblast fusion begins. From E8 to E18, this protein increases 10-fold, with a time course that precedes myoblast fusion. Although the content of CaZf-ATPase was significantly less in 68-year-old subjects than in 28-year-old subjects, the content of calsequestrin and the ryanodine receptor did not show a concomitant decrease (Klitzgaard et al., 1989). Calsequestrin was localized in frog ventricular cells in peripheral junctional SR and in corbular SR near the sarcolemma, but was absent from the central regions of the cell (McLeod et al., 1991).

3. 53-kDa and 160-kDa Glycoproteins The 53-kDa glycoprotein has the ability to bind ATP with high affinity and is localized at the inner surface of the SR membrane (Leberer et al., 1989). Both polyclonal and monoclonal antibodies to the 53-kDa glycoprotein cross-react with the 160-kDa glycoprotein. The 160-kDa glycoprotein has been named sarcalumenin and is a calcium-binding protein. Comparative immunoelectron microscope labeling with mAbs showed that the 53-kDa glycoprotein and CaZf-ATPasewere codistributed in the lumen of the free SR. Sarcalumenin had a similar distribution but labeling levels were much lower (Leberer et al., 1990).

4. Foot Protein (Ryanodine Receptor) The nomenclature used originates from the observation that there are projecting structures that appear to link terminal cisternal SR membranes

PROTEINS IN STRIATED MUSCLE

123

with T tubules. The structures were originally referred to as feet and subsequently also as bridges, pillars, or spanning protein. This linkage of the two membrane systems forms the well-known triad structure. Isolation of proteins that constitute these feet simultaneously isolated ryanodine receptor activity, hence the dual nomenclature. A wide array of molecular weights have been ascribed to the foot protein, ranging from 160k to 565k. A molecular weight of 565k was calculated from the c-DNA sequence (Takeshima et al., 1989). Kawamoto et al. (1986) suggested that native foot protein had a molecular mass of 630-800 kDa and that a foot was composed of four 300-kDa subunits. Immunogold localization of the 300kDa subunit showed that the antibody bound to the junctional gap of the triad. Trypsin treatment of isolated terminal cisternal vesicles removed the feet and also abolished antibody binding (Kawamoto et al., 1988). Permeabilizing vesicles either before or after trypsin treatment results in antibody binding on the luminal side of the membrane, suggesting that the foot protein spans the width of the membrane. This arrangement would be expected because of the calcium-channel activity associated with the protein. Campbell et al. (1987) found that mAbs to the ryanodine receptor recognized a protein that was enriched in isolated triads but was absent from light SR vesicles and T-tubule vesicles. Zorzato er al. (1989) obtained a 50% inhibition in calcium loading activity and a 25% increase in Ca2+ATPase activity when terminal cisternae were preincubated with antibodies to a putative foot protein. This suggests that calcium release channels are located in the feet and that the putative foot protein is a component of the calcium release channel. Airey et af. (1990) produced antibodies to two isoforms of a 500-kDa protein that bound [3Hl-ryanodine and that localized at the terminal cisternae. These two isoforms were named a and /3 foot proteins. Sutko et al. (1991) used these antibodies to the two foot protein isoforms to demonstrate that, although both isoforms increased markedly at the time of chick hatching, the a isoform was detected in a majority of fibers in E l 0 skeletal muscle, whereas the P isoform was first detected at E15. Sutko et al. (1991) speculated that the two isoforms may have specific roles in rnyofibril assembly or muscle differentiation. 5. Phospholamban

The monomer of phospholarnban has a molecular weight of 6080 and readily associates into pentamers. When cardiac muscle phospholamban is phosphorylated by a CAMP-dependent protein kinase, a process that is enhanced by P-adrenergic agonists, calcium transport by the SR Ca2+ATPase is enhanced. Conversely, when phospholamban is not phosphorylated, calcium transport is inhibited. Phospholamban is uniformly distributed in the network and the corbular SR, but is absent from the junctional SR, T tubules, and sarcolemma of adult canine ventricular cells (Jorgensen

124

MARVIN H. STROMER

and Jones, 1987). Hori et al. (1989) produced a mAb to bovine cardiac phospholamban that blocked phosphorylation of phospholamban and that localized along the cell surface and in a network over the myofibrils in monkey, dog, mouse, and human cardiac cells. This mAb did not stain skeletal or smooth muscle cells. Jorgensen and Jones (1986) found that, although three adult canine skeletal muscles contained much less phospholamban than cardiac muscle, slow (type I ) fibers stained with antibodies to phospholamban in all three muscles, but fast (type 2) fibers did not. It is unclear whether this discrepancy is because of properties of the antibodies or some other factor.

6. High-Affinity Calcium-Binding Protein (Calreticulin) This 55-kDa protein contains a relatively small number of high-affinity calcium-binding sites and has an immunofluorescence staining pattern very similar to that of calsequestrin. In skeletal muscle, localization is predominantly at junctional SR areas, whereas staining in cardiac and smooth muscle is in the SR and, in a variety of nonmuscle cells, is in the ER (Fliegel et al., 1989a). This indicates that the protein is shared in cells that contain SR and ER and that it may be present in a wide variety of tissues. The protein has recently been named calreticulin (Fliegel et al., 1989b).

6. Proteins Associated with the Sarcolemma 1. Dystrophin Dystrophin, the 400- to 430-kDa protein product of the DMD gene, is associated with four glycoproteins (156, 50, 43, and 35 kDa) to form a complex in normal skeletal muscle (Ervasti et al., 1990). Both the 156kDa glycoprotein and dystrophin were present in substantially reduced amounts in DMD patients and in mdx mice. The 156-kDa glycoprotein is a peripheral membrane protein, but the 50-, 43-, and 35-kDa glycoproteins are integral membrane proteins (Ervasti and Campbell, 1991). Each of these glycoproteins contains distinct epitopes, which suggests that they are not fragments of larger proteins or dystrophin. Antibodies to each glycoprotein show a subsarcolemma localization. Dystrophin is also localized in normal adult fibers in a narrow zone of fluorescence at the cytoplasmic side of the sarcolemma. Antibodies to both the N-terminus and the C-terminus of dystrophin show that both ends of the dystrophin molecule are associated with the cytoplasmic surface of human and rabbit sarcolemmal vesicles (Zubrzycka-Gaarn et al., 1991). In embryonic human

PROTEINS IN STRIATED MUSCLE

125

muscle, dystrophin first appears in the sarcoplasm near the myotendinous junction (Wessels et al., 1991a). Dystrophin is distributed throughout the fiber in fetal human muscle, but is restricted to the subsarcolemma site in adult human muscle (Wessels et al., 1991a). This change in distribution suggests that dystrophin accumulates in the sarcoplasm before its association with the sarcolemma. Dystrophin colocalized with talin at focal adhesions in cultured embryonic Xenopus skeletal muscle (Kramarcy and Sealock, 1990). Dystrophin is also present at the neuromuscular junction (Miike et al., 1989; Byers et al., 1991; Huard et al., 1991a)and in the brain (Miike et al., 1989). A mAb whose epitope is between residues 1750 and 2248 of dystrophin recognizes both dystrophin and a-actinin on Western blots (SDS denatured) but only a-actinin in frozen sections of human muscle (Nguyen et al., 1990).This suggests that there are structural homologies between dystrophin and a-actinin. Immunoelectron microscope localization of dystrophin has shown that 5-nm gold particles were almost entirely confined to a 75-nm rim at the sarcoplasmic side of the skeletal muscle sarcolemma (Cullen et al., 1990). Dystrophin also has a subsarcolemma location in cardiac muscle, but it is absent from intercalated disks (Byers et al., 1991). Examination of mouse muscle via the deep-etching replica method with unlabeled or gold-labeled dystrophin antibodies shows that the antibody is associated with rodshaped structures in the cytoskeleton between myofibrils and the sarcolemma (Wakayama and Shibuya, 1991a,b). Antibodies to dystrophin have also been used to isolate dystrophin molecules from dystrophin-enriched sarcolemma preparations (Pons et al., 1990). These molecules appeared as flexible rods 175 ~f:15 nm x 2 nm, some of which had an enlargement at one end, a diameter reduction at one end, or both. Antibodies to dystrophin have been widely used to determine the location of dystrophin in human DMD muscle, either in patients or in carriers, and in two animal models of DMD, the mdx mouse and the xmd dog. In human DMD patients, anti-dystrophin labeling is usually described as absent or markedly decreased (Bonilla et al., 1988a; Uchino et al., 1989; Wakayama et al., 1989). Arahata et al. (1989) found that the presence of higher- or lower-molecular-weight dystrophins resulted in patchy or discontinuous anti-dystrophin staining and that all DMD patient muscle contained no detectable dystrophin. Dystrophin was present in cultures of normal human muscle that were grown with nerve cells, but was absent if the muscle cells were obtained from DMD patients (Ecob-Prince et al., 1989a). Dystrophin was absent in undifferentiated myoblasts from control and DMD patients and from myotubes from one DMD patient (Miranda et al., 1988). Dystrophin was present in mature control myotubes and in reduced amounts in myotubes from a second DMD patient. Antibodies against synthetic peptides and fusion proteins from the amino-terminal

126

MARVIN

H. STROMER

region of human cDNA produced either reduced binding or no binding in DMD fibers (Zubrzycka-Gaarn et a/., 1988). Nicholson et al. (1989), however, found that eight of nine biopsies from DMD patients contained isolated fibers that were dystrophin-positive, indicating that most DMD patients synthesize some dystrophin. Subsequently, Nicholson et a/. (1990) reported that 40% of DMD biopsies contained dystrophin-positive fibers and 20% had very weak labeling on a large number of fibers. DMD carriers have both normal and dystrophin-deficient fibers (Bonilla et a/., 1988b) and present a mosaic pattern of immunostained fibers (Sugita et al., 1990). In the mdx mouse, dystrophin has been described as absent (Uchino et al., 1989) or either absent or markedly deficient (Wakayama et al., 1989). Heterozygous mdx mice show neighboring dystrophin-negative and dystrophin-positive fibers and discontinuous or patchy dystrophin labeling in fibers from 10-day-old mice (Watkins et al., 1989). Older heterozygous mdx mice have very few dystrophin-negative fibers, suggesting that nuclei containing the dystrophin gene can compensate for nuclei without the gene in the same fiber. Fusion of normal rat myoblasts with mdx myoblasts in culture resulted in normal dystrophin distribution in all hybrid myotubes even if the ratio of normal to dystrophic nuclei was low (Huard et al., 1991b). Mouse muscle dystrophin was used to produce a cDNA that expressed, in COS cells, a dystrophin that was indistinguishable from mouse muscle dystrophin and that was localized at the COS cell membrane (Lee et al., 1991).

2. Adherens-Junction-SpecificCell Adhesion Molecule (A-CAM) The A-CAM is a 135-kDa membrane glycoprotein that is located in the FA of chicken cardiac muscle intercalated disks (Volk and Geiger, 1986a). Immunogold labeling indicated that A-CAM is located closer to the FA midline than either vinculin or a-actinin, but is absent from desmosomes. At the junctional membrane, the epitope recognized by the A-CAM antibody is seemingly exposed at the external cell surface. The A-CAM is also present in cultured chick lens cells (Volk and Geiger, 1986b)and may exist in other cells where cell-cell adherens-type junctions are found. 3. T tubules and Other Sarcolemmal Proteins Malouf et al. (1986) used a mAb to a triad-enriched fraction of rabbit skeletal muscle and found that, with immunogold localization, T tubules and the opening of the T tubules at the sarcolemma were labeled. This antibody recognized 65- and 50-kDa bands in immunoblots of surface

PROTEINS IN STRIATED MUSCLE

127

membrane fractions. The presence of dihydropyridine receptors and dihydropyridine-sensitive calcium channels in T tubules was a property exploited by Malouf et af. (1987) to screen other mAbs. One of these increased the rate of single channel opening when interacting with the internal side of the channel protein and immunoprecipitated polypeptide bands with M , values greater than or equal to 175, 90, 55, and 34 kDa. Rosemblatt and Scales (1989) found that antibodies to 37- and 53-kDa proteins from an isolated T-tubule preparation localized to T tubules only. Two other proteins (28 and 100 kDa) from the same preparation were localized by immunofluorescence to both T tubules and the sarcolemma. Jorgensen et al. (1990) demonstrated that mAbs to a 28-kDa protein from T-tubule membranes is localized only in T-tubules and subsarcolemmal vesicles and is more densely distributed in fast (type 11) than in slow (type 1) fibers. A second mAb to a SO-kDa protein from the sarcolemma was localized in the sarcolemma and also was more abundant in fast than in slow fibers. Neither the 28- nor the 50-kDa protein identified by Jorgensen et af. (1990) was classified as a soluble protein and, instead, seemed to be attached to, or an integral part of, the two membrane domains. The 50-kDa protein was detected by immunofluorescence as foci at the sarcolemma in skeletal muscle from 17-day-old rabbit fetuses (Yuan et af., 1990).The 28kDa protein was present in some, but not all, 17-day-old rabbit fetal myotubes as a few foci at the sarcolemma. This indicates that the 50-kDa protein is synthesized before the 28-kDa protein. In older rabbit embryos, the 28-kDa protein was present in finger-like projections that extended from the cell surface toward the interior. In neonatal rabbit muscle, a network of these projections was connected by transient longitudinal structures that also stained with antibodies to the 28-kDa protein. In 10-dayold rabbit muscle, the longitudinal structures were seldom seen, and a transversely oriented network was dominant. Future experiments should clarify the functions of these proteins and their roles in assembly of T tubules and the sarcolemma.

V. Other Proteins

Antibodies to parvalbumin, a sarcoplasmic calcium- and magnesiumbinding protein, have shown that parvalbumin is present in fast-twitch (type 11) rat skeletal muscle fibers and is nearly absent from type I fibers (Celio and Heizmann, 1982). Five subgroups of type I1 fibers are evident, based on distinct staining intensities. In adult mouse muscles, parvalbumin is present in moderate or large amounts in fast-twitch glycolytic (type IIB) fibers, is absent or present in small amounts in fast-twitch oxidative-

128

MARVIN

H. STROMER

glycolytic (type IIA) fibers, and is absent from slow-twitch oxidative (type I) fibers (Ecob-Prince and Leberer, 1989). Culturing adult mouse muscle with embryonic mouse spinal cord produced innervated myotubes that expressed fast MHC and stained histochemically like fast-twitch fibers, but which contained no parvalbumin (Ecob-Prince and Leberer, 1989). Schmitt and Pette (1991) quantitated the amount of parvalbumin present in specific fiber types in rabbit muscle. Type I fibers contained extremely low concentrations of parvalbumin. Type I1 fibers differed in their parvalbumin concentration, depending on subgroup of the fiber. The specific factor(s) that influences the expression of parvalbumin is unknown. An overview of parvalbumin is contained in the review by Pette and Staron ( 1990).

VI. Conclusions and Outlook Information gained about striated muscle by using antibodies is both diverse and impressive. One of the early questions normally asked when a new protein is discovered is where is it located. Antibody localization has been the obvious tool of choice to obtain that information. This, in turn, provides clues about other proteins that may be candidates for interactions because of their location in the same or adjacent domains. The use of mAbs not only has made it possible to locate specific epitopes and to probe functional sites on proteins but also, in many instances, has made us aware of other levels of complexity that will be the subject of future investigations. The many different labels and various detection options currently available increase the importance of carefully characterizing antibodies or their fragments for specificity and cross-reactivity. The exchange of antibodies or other laboratory-to-laboratory interactions are often very useful in clarifying uncertainties about antibodies and should be encouraged. The effects of species differences and epitope accessibility differences, however, should not be overlooked. The recent availability of new fluorochromes that facilitate double labeling and/or that are less photosensitive will continue to ease the problems of recording observations. For EM study of antibody-labeled samples, colloidal gold has become the label of choice. Variability of quality in some commercially available colloidal gold complexes (e.g., colloidal gold complexed with either protein A or protein G) has been documented. Concerns have also been expressed about both resolution limitations and penetrability properties of labels that are suitable for EM studies. A continued search for electron-dense tags that could provide gains in both resolution and accessibility to sites within tissues

PROTEINS IN STRIATED MUSCLE

129

and cells would seem to be very worthwhile. The electron spectroscopic imaging approach with boronated protein A that was pioneered by Bendayan et d.(1989) is a very interesting approach, but requires specialized instrumentation. Another approach has been to use progressively smaller gold particles, with diameters of 5 nm or less. Hainfeld (1987) has covalently linked Fab’ fragments to clusters containing I I gold atoms. Although the smaller-diameter gold particles provide the potential for improved resolution and penetratability, small-diameter probes can sometimes be difficult to observe unless both staining and instrument operation are optimized. Silver enhancement has been used to improve the visability of the very small particles in the EM and has made it possible to visualize gold labels with the light microscope. The prospects for using well-characterized antibodies with detection methods of ever-improving resolutions make this an exciting time to be using antibodies to improve our understanding of striated muscle and other biological systems as well. The greater our emphasis on using quality antibodies and on optimizing all aspects of our detection systems, the more rapidly we will gain information from well-planned experiments. It should, for example, be possible to determine where a-actinin is in the Z line o r to better understand the complex arrangement of proteins that link filaments to membranes. When we have answers to questions such as these, we will have a far better grasp of the significance of these structures.

Acknowledgments I am grateful to Mary Sue Mayes for her very able assistance with the references and to Marylou Weigel for transforming my rough draft into the final copy. Special thanks go to Dr. Marion Greaser for his suggestions and critique of the manuscript. My research has been supported by both the American Heart Association-Iowa Affiliate and Iowa State University. This is Journal Paper No. 5-14770 of the Iowa Agriculture and Home Economics Experiment Station, Ames, Iowa 50011 (Project No. 2921).

References Aarli, J. A,, Stefansson, K., Marton, L. S. , and Wollmann. R. L. (1990). Clin. Exp. Immunol. 82, 284-288. Airey, J. A., Beck, C. F., Murakami, K., Tanksley. S. J., Deerinck, T. J.. Ellisman, M. H., and Sutko, J . L. (1990). J . B i d . Chem. 265, 14,187-14,194. Allen, M. D., Tsuboi, H., Togo, T., Eary, J. F., Gordon, D.. Thomas, R., and Reichenbach, D. D. (1989). Transplantation 48, 923-928. Antunes, M. L., Seldin, D. W., Wall, R. M., and Johnson, L. L. (1989). A m . J . Cardiol. 63, 777-783.

130

MARVIN H. STROMER

Appleyard, S. T., Dunn, M. J., Dubowitz, V., Scott, M. L., Pittman, S. J.. and Shotten, D. M. (1984). Proc. Natl. Acad. Sci. U . S . A . 81, 776-780. Arahata, K., Hoffman, E. P., Kunkel, L. M., Ishiura, S., Tsukahara, T., Ishihara, T., Sunohara, N., Nonaka, I., Ozawa, E., and Sugita, H. (1989). Proc. Natl. Acad. Sci. U.S.A. 86, 7154-7158. Askanas, V., Bornemann, A., and Engel, W. K. (1990). Neurology 40,949-953. Atherton, B. T., Meyer, D. M., and Simpson, D. G. (1986). J . Cell Sci. 86, 233-248. Azzarello, G., Sartore, S., Saggin, L., Gorza, L., D'Andrea, E., Chieco-Bianchi, L., and Schiaffino, S. (1987). J . Cancer Res. Clin. Oncol. 113, 417-429. Babai, F., Musevi-Aghdam, J., Schurch. W., Royal, A,, andGabbiani, G. (1990).Differentiation 44, 132-142. Bader, D., Masaki, T., and Fischman, D. A. (1982). J. Cell Biol. 95, 763-770. Bahler, M., Eppenberger, H. M., and Wallimann, T. (1985a). J . Mol. B i d . 186, 381-391. Bahler, M., Eppenberger, H. M., and Wallimann, T. (1985b). J. Mol. Biol. 186, 393-401. Bahler, M., Moser, H., Eppenberger, H. M., and Wallimann, T. (1985~).Dev. Biol. 112, 345-352. Ballester-Rodes, M., Carrio-Gasset, I., Abadal-Berini, L., Obrador-Mayol, D., BernaRoqueta, L., and Caralps-Riera, J. M. (1988). Am. J . Cardiol. 62, 623-627. Bandman, E. (1985). Science 227, 780-782. Bandman, E., Matsuda, R., and Strohman, R. C. (1982). Dev.Biol. 93, 508-518. Belkin, A. M., Ornatsky, 0. I., Glukhova, M. A., and Koteliansky, V. E. (1988). J. Cell Biol. 107, 545-553. Belkin, A. M., Zhidkova, N. I., and Koteliansky, V. E. (1986). FEES Lett. 200, 32-36. Beller, G. A., Khaw, B. A., Haber, E., and Smith, T. W. (1977). Circulation 55, 74-78. Bendayan, M., Barth, R. F., Gingras, D., Londono, I., Robinson, P. T., Alam, F., Adams, D., and Mattiazzi, L. (1989). J . Hisrochem. Cytochern. 37, 573-580. Bennett, P., Craig, R., Starr, R., and Offer, G. (1986).J. Muscle Res. Cell Moril. 7,550-567. Bignami, A., and Dahl, D. (1984). J. Hisrochem. Cytochem. 32,473-476. Bilak, M. M., Robson, R. M., Bilak, S. R., Bright, M. J., and Stromer, M. H. (1991).J. Cell B i d . 115, 333a. Billeter, R., Weber, H., Lutz, H., Howald, H., Eppenberger, H. M., and Jenny, E. (1980). Histochemistry 65, 249-259. Biral, D., Damiani, E., Scarpini, E., Bet, L., Bresolin, N., Moggio, M., Pellegrini, G., Barbieri, S., and Scarlato, G. (1987). Neurology 37, 1658-1662. Biral, D., Scarpini, E., Angelini, C., Salviati, G., and Margreth, A. (1989). Muscle Nerve l2,43-51. Birkenberger, L., and Ip, W. (1990). J. Cell Biol. 111, 2063-2070. Bonilla, E., Samitt, C. E., Miranda, A. F., Hays, A. P., Salviatti, G., DiMauro, S., Kunkel, L. M., Hoffman, E. P., and Rowland, L. P. (1988a). Cell (Cambridge. Mass.) 54,447-452. Bonilla. E. Schmidt, B., Samitt, C. E., Miranda, A. F., Hays, A. P., de Oliveira, A. B. S., Chang, H. W., Servidei, S., Ricci, E., Younger, D. S.,and DiMauro, S. (1988b). A m . J . Pathol. 133, 440-445. Bormioli, S. P., Sartori, S.. Vitadello, M., and Schiaffino, S. (1980). J. CellBiol. 85,672-681. Borrione, A. C., Zanellato, A. M., Saggin, L., Mazzoli, M., Azzarello, G., and Sartore, S. (1988). Differenfiation 38, 49-59. Bosyczko, D., Decker, C., Muschler. J., and Horowitz, A. F. (1989). Exp. Cell Res. 183, 72-9 I. Bouvagnet, P., Leger, J. O., Pons. F., Dechesne, C., and Leger, J. J. (1984). Circ. Res. 55, 794-804. Bouvagnet, P., Neveu, S., Montoya, M., and Leger, J. J. (1987). Circ. Res. 61, 329-336. Brandl, C. J., Green, N. M., Korcyak, B., and MacLennan, D. H. (1986). Cell (Cambridge, M U S S . )44, 597-607.

PROTEINS IN STRIATED MUSCLE

131

Briggs, F. N., Lee, K. F., Feher, J. J., Wechsler, A. S., Ohlendieck, K., and Campbell, K. (1990). FEES Lerr. 259, 269-272. Bugaisky, L. B., and Zak, R. (1989). Circ. Res. 64, 493-500. Bulinski, J. C., Kumar, S., Titani, K., and Hauschka, S. D. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 1506-1510. Buller, A. J., Eccles, J. C.. and Eccles. R. M. (1960). J. Physiol. (London) 150, 417-439. Butler-Browne, G . S., Eriksson, P. 0.. Laurent, C., and Thornell, L.-E. (1988). Muscle Nerve 11, 610-620. Butler-Browne, G. S., and Whalen, R. G. (1984). Deu. Biol. 102, 324-334. Byers, T. J., Kunkel, L. M.. and Watkins, S. C . (1991). J . CellBiol. 115, 411-421. Caldwell, J. E., Waddle, J. A., Cooper, J. A., Hollands, J. A., Casella, S. J., and Casella, J. F. (1989). J. Biol. Chem. 264, 12.648-12.652. Callaway, J. E., and Bechtel, P. J. (1981). Biochem. J. 195, 463-469. Campbell, G. R., Chamley-Campbell, J., Groschel-Stewart, U., Small, J . V., and Anderson, P. (1979). J. Cell Sci. 37, 303-322. Campbell, K. P., Knudson, C. M., Imagawa, T., Leung, A. T., Sutko, J. L., Kahl, S. D., Raab, C. R., and Madson, L. (1987). J. Biol. Chem. 262, 6460-6463. Campion, D. R. (1984). I n i . Reu. Cyrol. 87, 225-251. Canale, E., Campbell, J. H., and Campbell, G. R. (1983).J . Mol. Cell. Cardiol. 15, 197-206. Carlsson, E., Grove, B. K., Wallimann, T., Eppenberger, H. M., andThornell, L.-E. (1990). Histochemistry 95, 27-35. Carpenter, C. E., Cassens, R. G., and Greaser, M. L. (1988). J . Anim. Sci. 66, 814-818. Carpenter, C. E., Greaser, M. L., and Cassens, R. G. (1987). J. Anim. Sci. 64, 1574-1587. Carron, C. P., Hwo, S., Dingus, J., Benson, D. M., Meza, I.. and Bryan, J. (1986). J. Cell Biol. 102, 237-245. Casella, J. F., Craig, S. W., Maack, D. J., and Brown, A. E. (1987). J . Cell Biol. 105, 371-379. Casella, J. F., Maack, D. J., and Lin, S. (1986). J . Biol. Chem. 261, 10,915-10,921. Celio, M. R., and Heizmann, C. W., (1982). Narure (London) 297, 504-506. Cerny, L. C., and Bandman, E. (1986). J. Cell Biol. 103, 2153-2161. Choi, E. S., and Clegg, D. 0. (1990). Deu. Biol. 142, 169-177. Chowrashi, P. K., and Pepe, F. A. (1982). J . Cell Biol. 94, 565-573. Colley, N. J., Tokuyasu, K. T., and Singer, S. J. (1990). J . Cell Sci. 95, 11-22. Colyer, J . , Mata, A. M., Lee, A., and East. J. M. (1989). Biochem. J . 262, 439-447. Condon, K . , Silberstein, L., Blau, H. M., and Thompson, W. J. (1990a). Deu. Biol. 138, 256-274. Condon, K., Silberstein, L., Blau, H . M., and Thompson, W. J. (1990b). Deu. Biol. 138, 275-295. Conrad, A. H . , Clark, W. A., and Conrad, G. W. (1991). Cell Motil. Cytoskeleton 19, 189-206. Craig, S . W., and Pardo, J. V. (1983). Cell Motil. 3, 449-462. Crow, M. T., and Stockdale, F. E. (1986). Deu. Biol. 113, 238-254. Cullen. M. J., Walsh, J., Nicholson, L. V. B., and Harris, J. B. (1990). Proc. R . Soc. London B 240, 197-210. Cummins, B., Russell, G. J., Chandler, S. T., Pears, D. J., and Cummins, P. (1990). Cardiouasc. R e s . (England) 24, 3 17-327. Damiani, E., Margreth, A., Furlan, A., Dahms, A. S. , A r m , J., and Sabbadini. R. A. (1987). J . Cell Biol. 104, 461-472. Dan-Goor, M., Silberstein, L., Kessel, M., and Muhlrad, A. (1990). J. Muscle Res. Cell Motil. 11, 216-226. Danon, M. J., Karpati, G., Charuk, J., and Holland, P. (1988). Neurology 38, 812-815.

132

MARVIN H. STROMER

Danto, S. I., and Fischman, D. A. (1984). J. Cell B i d . 98, 2179-2191. DasGupta, G., and Reisler, E. (1991). Biochemistry 30, 9961-9966. Dechesne, C. A., Leger, J. 0. C., Bouvagnet, P., Claviez, M., and Leger, J. J . (1985). J . Mol. Cell Cardiol. 17, 753-767. Dechesne, C. A., Leger, J . 0. C., and Leger, J. J. (1987). Deu. Biol. 123, 169-178. De Geeter, F., Deleu, D., Debeuckelaere, S., DeConinck, A., Somers, G., and Bossoyt, A. (1989). Nucl. Med. Commun. 10, 603-607. De Groot, I. J., Lamers, W. H., and Moorman, A. F. M. (1989). Anat. Rec. 224, 365-373. De Jong, A. S., van Kessel-van Vark, M., and van Heerde, P. (1987a). Acta Cytol. 31, 573-577. De Jong, F.. Geerts, W. J. C., Lamers, W. H., Los, J. A., and Moorman, A. F. M. (l987b). Anat. Embryol. (Berl.)177, 81-90. De Jong, F., Geerts, W. J. C., Lamers, W. H., Los, J. A., and Moorman, A. F. M. (1990). Anat. Rec. 226, 213-227. De Nardo, D. Scibilia, G . , Macchiarelli, A. G., Cassisi, A., Tonelli, E., Papalia, U., Gallo, P., Antolini, M., Pitucco. G., Reale, A., Caputo, V., and Marino, B. (1989). J. Heart Transplant 8, 407-412. Dennis, J. E., Shimizu, T., Reinach, F. C., and Fischman, D. A. (1984). J. Cell B i d . 98, 1514-1522. Desypris, G., and Parry, D. J. (1990). A m . J. Physiol. 258, C62-C70. Dhanarajan, Z. C., and Atkinson, B. G. (1980). Can. J. Biochem. 58, 516-526. Dhoot, G. K., Hales, M. C., Grail, B. M., and Perry, S. V. (1985). J. Muscle Res. Cell Motil. 6,487-505. Dhoot. G. K., and Pearce, G. W. (1984). J . Neurol. Sci. 65, 17-28. Dhoot, G. K., and Perry, S. V. (1979). Nature (London) 278, 714-718. Dix, D. J., and Eisenberg, B. R. (1988). J. Histochem. Cytochem. 36, 1519-1526. Dix, D. J . , and Eisenberg, B. R. (1990). J. Cell B i d . 111, 1885-1894. Dlugosz, A. A., Antin, P. B., Nachmias, V. T., and Holtzer, H. (1984). J. Cell Biol. 99, 2268-2278. Draeger, A., Weeds, A. G., and Fitzsimons, R. B. (1987). J. Neurol. Sci. 81, 19-43. Drenckhahn, D., Beckerle, M., Burridge. K., and Otto, J. (1988). Eur. J. Cell Biol. 46, 5 13-522. Dulhunty, A. F., Banyard, M. R. C., and Medveczkey, C. J. (1987). J. Membrane Biol. 99, 79-92. Ecob, M. S., Butler-Browne, G. S., and Whalen, R. G. (1983). Differentiation 25, 84-87. Ecob-Prince, M. S., Jenkison, M., Butler-Browne, G. S., and Whalen, R. G. (1986). J. Cell Biol. 103, 995-1005. Ecob-Prince, M. S., Hill, M. A., and Brown, A. E. (1989a). Muscle Nerve 12, 594-597. Ecob-Prince, M., Hill, M.. and Brown, W. (1989b). J. Neurol Sci. 91, 71-78. Ecob-Prince, M. S . , and Leberer, E. (1989). Differentiation 40, 10-16. Edman, K. A. P., Reggiani, C., Schiaffino. S.. and teKronnie, G. (1988).J. Physiol. (London) 395, 679-694. Elgazzar, A. H., Malki, A. A., Abdel-Dayem, H. M., Owunwanne, A,, Dadah, M., Mahmoud, S., and el-Sayed, M. (1989). Nucl. Med. Commun. 10, 661-667. Endo, M., Nonomura, Y., Masaki, T., Ohtsuki, I., and Ebashi, S. (1966). J. Biochem. 60, 605-608. Endo, T., and Masaki, T. (1984). J. Cell Biol. 99, 2322-2332. Eppenberger-Eberhardt, M.,Flamme, I . , Kurer, V., and Eppenberger, H. M. (1990). Deu. Biol. 139, 269-278. Eppenberger, M. E., Hauser, I., Baechi, T., Schaub, M. C., Brunner, U. T., Dechesne, C. A., and Eppenberger, H. M. (1988). Deu. Biol. 130, 1-15.

PROTEINS IN STRIATED MUSCLE

133

Eppenberger, M., Hauser, I., and Eppenberger, H. M. (1987). Biomed. Biochim. A C ~ U46, S6403645. Erickson. C. A., Tucker, R. P., and Edwards, B. F. (1987). Dgferentiution 34, 88-97. Ervasti, J. M., and Campbell. K. P. (1991). Cell (Cumbridge, Mass.) 66, 1121-1131. Ervasti, J. M., Ohlendieck, K.. Kahl, S. D.. Gaver, M. G., and Campbell, K. P. (1990). Nature (London) 345, 315-319. Evans, D., Miller, J. B., and Stockdale, F. E. (1988). Deu. Biol. 127, 376-383. Everett, A. W. (1986). J. Mol. Cell Cardiol. 18, 607-615. Everett, A. W., and Sparrow, M. P. (1987). J. Muscle Res. Cell Motil. 8, 220-228. Feldman, J. L., and Stockdale, F. E. (1991). Deu. Biol. 143, 320-334. Fischman, D. A., and Danto, S. I. (1985). Ann. N . Y . Acud. Sci. 455, 167-184. Fliegel, L., Burns, K., Opas, M., and Michalak, M. (1989a). Biochim. Biophys. Acta 982, 1-8. Fliegel, L, Burns, K., MacLennan, D. H., Reithmeier. R. A., and Michalak, M. (1989b), J . Biol. Chem. 264, 21.522-21.528. Forsgren, S., Eriksson, A.. Kjorell. U . , and Thornell. L.-E. (1982). Histochemistry 75, 43-52. Forsgren, S., Strehler, E., and Thornell, L.-E. (1983). Histochem. J . 15, 1099-1 I I I . Frame, L. H., Lopez, J. A., Khaw. B. A., Fallon. J. T., Haber, E.. and Powell, W. J., Jr. (1983). J . Clin. Inuest. 72, 535-544. Franchi, L. L., Murdoch, A., Brown, W. E., Mayne, C. N.. Elliott. L., and Salmons. S. (1990). J . Muscle Res. Cell Motil. 11, 227-239. Friden. J., Kjorell. U., and Thornell, L.-E. (1984). In?. J. Sports Med. 5 , 15-18. Frist, W., Yasuda, T., Segall, G., Khaw, B. A., Strauss, H. W., Gold, H., Stinson, E., Oyer, P., Baldwin, J., Billingham, M., McDougall, R., and Haber. E. (1987). Circulation 76, (Suppl. V), V81-V85. Fuldner. R. A.. Lim, S. -S., Greaser, M. L., and Lemanski, L. F. (1984).J . Embryo/. Exp. Morphol. 84, 1-17. Fulton, A. B.. and Isaacs, W. B. (1991). BioEssays 13, 157-161. Funatsu, T., Asami, Y., and Ishiwata, S. (1988). J . Biochem. (Tokyo) 103, 61-71. Furst, D.. Nave, R., Osborn, M., Weber, K., Bardosi, A,. Archidiacono, N., Ferro, M., Romano, V., and Romeo, G. (1987). FEBS Lett. 224,49-53. Furst, D. O., Osborn, M., Nave, R., and Weber, K. (1988). J. Cell Biol. 106, 1563-1572. Furst, D. O., Nave, R., Osborn, M., and Weber, K. (1989a). J. Cell Sci. 94, 119-125. Furst, D. 0.. Osborn, M., and Weber, K. (1989b). J. CellBiol. 109,517-527. Ganote, C. E.. and Vander Heide, R. S. (1987). Am. J . Parhol. 129, 327-344. Gassner, D. (1986). Eur. J . Cell Biol. 40, 176-184. Gauthier, G. F. (1979). J . Cell Biol. 82, 391-400. Gauthier, G. F. (1990). J. Cell B i d . 110, 693-701. Gauthier, G. F., Burke, R. E., Lowey, S., and Hobbs. A. W. (1983).J . CellBiol. 97,756-771. Gauthier, G. F., Ono, R. D., and Hobbs, A. W. (1984). Deu. Biol. 105, 144-154. Geisler. N., Kaufmann, E., and Weber. K. (1985). J. Mol. Biol. 182, 173-177. Ginjaar, 1. B., Bakker, E., denDunnen. J. T., Wessels, A.. van Paassen, M.M. , Kloosterman, M. D., Zubrzycka-Gaarn, E. E., Fischbeck, K. H., Moorman, A. F., and van Ommen, G. J. (1990). Adu. Exp. Med. Biol. 280, 17-23. Gomer, R. H., and Lazarides, E. (1981). Cell (Cambridge. Mass.) 23, 524-532. Gomer, R. H., and Lazarides, E. (1981). J . Cell Biol. 97, 818-823. Gonzalez-Sanchez, A., and Bader, D. (1984). Deu. Biol. 103, 151-158. Gonzalez-Sanchez, A , , and Bader, D. (1985). J . Cell Biol. 100, 270-275. Goodman, S . R., Zagon, I. S., and Kulikowski, R. R. (1981). Proc. Nut/. Acad. Sci. U . S . A . 78. 7570-7574.

134

MARVIN H. STROMER

Gorza, L. (1990). J. Histochem. Cytochem. 38, 257-265. Gorza, L., Sartore, S., and Schiaffino, S. (1982). J . Cell Biol. 95, 838-845. Gorza, L., Mercadier, J. J., Schwartz, K., Thornell, L.-E., Sartore, S., Schiaffino, S. (1984). Circ. Res. 54, 694-702. Gorza, L., Sartore, S., Thornell, L.-E., and Schiaffino, S. (1986).J. CellBiol. 102,1758-1766. Gorza, L., Gundersen, K., Lomo, T., Schiaffino. S., and Westgaard, R . H. (1988a). J . Physiol. (London)402, 627-649. Gorza, L., Saggin, L., Sartore, S., and Ausoni, S. (1988b). J. Mol. Cell Cardiol. 20,931-941. Granger, B. L., and Lazarides, E. (1978). Cell (Cambridge, Mass.) 15, 1253-1268. Granger, B. L., and Lazarides, E. (1979). Cell (Cambridge, Mass.) 18, 1053-1063. Greaser, M. L., and Gergely, J. (1973). J . Biol. Chem. 248, 2125-2133. Greaser, M. L., Handel, S. E., Wang, S. -M., Schultz, E., Bulinski, J. C., Lin, J. -C., and Lessard, J. L. (1989). In “Cellular and Molecular Biology of Muscle Development” (F. Stockdale and L. Kedes, eds.), UCLA Symposia on Molecular and Cellular Biology, Vol. 93, pp. 247-257. A. R. Liss, New York. Gregorio, C. C., Hudecki, M. S., Pollina, C. M., and Repasky, E. A. (1988). Muscle Nerue 11,372-379. Groschel-Stewart, U. (1980). Int. Rev. Cytol. 65, 193-254. Grove, B. K., Cerny, L., Perriard, J. C., and Eppenberger, H. M. (1985). J . Cell Biol. 101, 1413-1421. Grove, B. K., Holmbom, B., and Thornell, L.-E. (1987). Differentiation 34, 106-114. Grove, B. K., Kurer, V., Lehner, C., Doetschman, T. C., Perriard, J. C., and Eppenberger, H. M. (1984). J . Cell Biol. 98, 518-524. Grove, B. K., and Thornell, L.-E. (1988). Muscle Nerve 11, 645-653. Hainfeld, J. F. (1987). Science 236, 450-453. Handel, S. E., Greaser, M. L., Schultz, E., Wang, S. -M., Bulinski, J. C., Lin, J. J. -C., and Lessard, J. L. (1991). Cell Tissue Res. 263, 419-430. Handel, S. E., Wang, S. -M., Greaser, M. L., Schultz, E., Bulinski, J. C., and Lessard, J . L. (1989). Deu. Biol. 132, 35-44. Harrington, W. F., Karr, T., Busa, W. B., and Lovell, S. J. (1990). Proc. Nail. Acad. Sci. U.S.A. 87,7453-7456. Harris, A. J., Duxson, M. J., Fitzsimons, R. B., and Rieger, F. (1989a). Deuelopment 107, 771-784. Harris, A. J., Fitzsimons. R. B., and McEwan, J. C. (398913). Development 107, 751-769. Hartner, K.-T., Kirschbaum, B. J., and Pette, D. (1989). Eur. J. Biochem. 179, 31-38. Hesketh, J., Campbell, G., and Loveridge, N. (1991). Biochem. J. 279, 309-310. Hill, C. S., Duran, S., Lin, Z., Weber, K., and Holtzer, H. (1986). J . Cell Biol. 103, 2185-2196. Hill, C. S., and Lemanski, L. F. (1986). Eur. J . Cell Biol. 39, 300-312. Hill, C., and Weber, K. (1986). J . Cell Biol. 102, 1099-1 108. Hoberg, E., Eisenhut, M., Hofmann, M., Schuler, G., Brachmann, J., Aidonidis, I., Nachtkamp, J., Kubler, W., and Katus, H. A. (1988). Eur. Henrt J . 9, 328-336. Hoffman, E. P., Watkins, S. C., Slayter, H. S., and Kunkel, L. M. (1989). J. CellBiol. 108, 503-5 10. Hoh, J. F., and Hughes, S. (1988). J. Muscle Res. Cell Motil. 9, 59-72. Hoh, J. F., and Hughes, S. (1989). J . Muscle Res. Cell Motil. 10, 312-325. Hoh, J. F., Hughes, S., Hale, P. T., and Fitzsimons, R . B. (1988a). J . Muscle Res. Cell Motil. 9, 30-47. Hoh, J. F., Hughes, S., Chow, C., Hale, P. T., and Fitzsimons, R . B. (1988b). J. Muscle Res. Cell Motil. 9, 48-58. Hori, Y., Takamoto, T., Schichijo, S., Matsuo, Y., and Yokoyama, M. M. (1989). J. Clin. Lab. Immunol. 30, 163-168.

PROTEINS IN STRIATED MUSCLE

135

Home, Z., and Hesketh, J. (1990). Biochem. J . 268, 231-236. Horowits, R., Maruyama, K., and Podolsky, R. J. (1989). J . Cell Biol. 109, 2169-2176. Hu, D. H., Matsuno, A., Terakado, K., Matsuura, T., Kimura, S., and Maruyama, K . (1990). J . Muscle Res. Cell Motil. 11, 497-51 I . Huard, J., Fortier, L. -P., Labrecque, C., Dansereau, G., and Tremblay, J. P. (1991a). Synapse 7 , 135-140. Huard, J., Labrecque, C., Dansereau, G., Robitaille. L., andTremblay, J. P. (1991b). Muscle Nerve 14, 178-182. Huxley, H. E. (1963). J . Mol. Biol. 7 , 281-308. Ikeya, H., Ohashi, K., and Maruyama, K. (1983). Biomed. Res. 4, 1 1 1-1 16. Irish, M. J., Hirabayashi, T., and Wilson, F. J. (1977). Tissue Cell 9, 499-505. Isobe, Y., Hou, G. R., and Lemanski, L. F. (1991). Anat. Rec. 229,415-426. Isobe, Y . , Warner, F. D., and Lemanski, L. F. (1988). Proc. Nail. Acad. Sci. U . S . A . 85, 6758-6762. Ito, M.,Hirono, Y., Watanabe, S., Yamamoto, H., and Maruyama, K. (1991). J. Biochem. 110, 301-305. Itoh, Y., Suzuki, T., Kimura, S., Ohashi, K., Higuchi, H., Sawada, H., Shimizu, T., Shibata, M., and Maruyama, K. (1988). J . Biochem. 104, 504-508. Jacoby, J., KO, K., Weiss, C., and Rushbrook, J. I. (1990). J. Muscle Res. Cell Motil. 11, 25-40. Jain, D., Crawley, J. C., Lahiri, A., and Raftery, E. B. (1990). J. Nucl. Med. 31, 231-233. Jakobsson, F., Borg, K., and Edstrom, L. (1990). Acra Neuropathol. (Berl.)80, 459-468. Jasmin, G., Proschek, L., Dechesne, C., and Leger, J . (1988). Proc. Soc. Exp. Biol. Med. 188, 142-148. Jiang, B., Roy, R. R., and Edgerton, V. R. (1990). Muscle Nerve 13, 1037-1049. Jin, J.-P., Malik, M. L., and Lin, J. J. -C. (1990). Hybridoma 9, 597-608. Jin, J.-P., and Wang, K. (1991). FEES Lett. 281, 93-96. Jockusch, H.. and Jockusch, B. M. (1980). Deu. Biol. 75, 231-238. Jockusch, B. M., Veldman, H., Griffiths, G. W., van Oost, B. A., and Jennekens, F. G. L. (1980). Exp. Cell Res. 127, 409-420. Johnson, L. L., and Cannon, P. J. (1991). Circulation 84,(Suppl. I), 1273-1279. Johnson, L. L., and Seldin, D. W. (1989). Semin. Nucl. Med. 19, 238-246. Johnson, L. L., Seldin, D. W., Becker, L. C., LaFrance. N . D., Liberman, H. A,, James, C., Mattis, J. A., Dean, R. T., Brown, J., Reiter, A., Ameson, V., Cannon, P. J., and Berger, H . J. (1989). J . Am. Coll. Cardiol. 13, 27-35. Jorgensen, A. O., Arnold, W., Pepper, D. R., Kahl, S. D., Mandel, F., and Campbell, K. P. (1988). Cell Motif. Cytoskeleton 9, 164-174. Jorgensen, A. O., Arnold, W., Shen, A. C. -Y., Yuan, S., Gaver, M., and Campbell, K. P. (1990). J . Cell Biol. 110, 1173-1 185. Jorgensen, A. O., and Bashir, R. (1984). Deu. Biol. 106, 156-165. Jorgensen, A. O., and Campbell, K. P., (1984). J. CellBiol. 98, 1597-1602. Jorgensen, A. O., and Jones, L. R. (1986). J. Biol. Chern. 261, 3775-3781. Jorgensen, A. O., and Jones, L. R. (1987). J. CellBiof. 104, 1343-1352. Jorgensen, A. O., Kalnins, V., and MacLennan, D. H. (1979). J . Cell Biol. 80, 372-384. Jorgensen, A. O., and McGuffee, L. J. (1987). J. Histochem. Cytochem. 35, 723-732. Jorgensen, A. O., Shen, A. C., and Campbell, K. P. (1985). J . Cell Biol. 101, 257-268. Jorgensen, A. 0.. Shen, A. C., Campbell, K. P., and MacLennan, D. H . (1983). J. CellBiol. 97, 1573-1581. Jorgensen, A. O., Shen, A. C., Daly, P., and MacLennan, D. H. (1982). J. Cell Biol. 93, 883-892. Kaprielian, Z . , and Fambrough, D. M. (1987). Deu. Biol. 124, 490-503.

136

MARVIN H. STROMER

Karpati, G., Charuk, J . , Carpenter, S., Jablecki, C., and Holland, P. (1986). Ann. Neurol. 20, 38-49. Kartenbeck, J. Franke, W. W., Moser, J . G., and Stoffels, U. (1983). EMBO J.2,735-742. Katoh, T., and Lowey, S. (1989). J. Cell Biol. 109, 1549-1560. Kaufman, S. J . , Bielser, D., and Foster, R. F. (1990). Exp. Cell Res. 191, 227-238. Kawamoto, R. M., Brunschwig, J.-P., and Caswell, A. H. (1988). J. Muscle Res. Cell Motil. 9,334-343. Kawamoto, R. M., Brunschwig, J.-P., Kim, K. C., and Caswell, A. H . (1986). J. Cell Biol. 103, 1405-1414. Kelley, A. M., and Rubinstein, N. A. (1980). Nature (London) 288, 266-269. Kennedy, J. M., Eisenberg, B. R., Reid, S. K., Sweeney, L. J . , and Zak, R. (1988). A m . J . Anat. 181, 203-215. Kennedy, J. M., Kamel, S., Tambone, W. W., Vrbova, G., and Zak, R. (1986). J. CellBiol. 103, 977-983. Kennedy, J. M., Zak, R., and Gao, L. (1991). Muscle Nerve 14, 166-177. Khaw, B. A., Beller, G. A., Haber, E., and Smith, T. W. (1976). J . Clin. Invest. 58,439-446. Khaw, B. A., Fallon, F. T., Strauss, H. W., and Haber, E. (1980). Science 209, 295-297. Khaw, B. A., Gold, H. K., Leinback, R. C., Fallon, J. T., Strauss, W., Pohost, G. M., and Haber, E. (1978). Circulation 58, 1137-1 142. Khaw, B. A., Gold, H. K., Yasuda, T., Leinbach R. C., Kanke, M., Fallon, J. T., BarlaiKovach, M., Strauss, H . W., Sheehan, F., and Haber, E. (1986). Circulation 74,501-508. Khaw, B. A., Mattis, J. A., Melincoff, G., Strauss, H. W.. Gold, H. K., and Haber, E. (1984). Hybridoma 3, 11-23, Klein, I., Daood, M., and Whiteside, T. (1985). J. Cell Physiol. W, 49-53. Klitgaard, H., Ausoni, S., and Damiani, E. (1989). Acta Physiol. Scand. 137, 23-31. Komiyama, M., Maruyama, K., and Shimada, Y. (1990). J . Muscle Res. Cell Moril 11, 419-428. Komuro, I., Nomoto, K., Takaku, F., and Yazaki, Y. (1988). Jpn. Circ. J . 52, 1191-1200. Koteliansky, V. E., Glukhova, M. A., Gneushev, G. N., Samuel, J. L., and Rappaport, L. (1986). Eur. J . Biochem. 156, 619-623. Koteliansky, V. E., and Gneushev, G. N. (1983). FEBS Lett. 159, 158-160. Koteliansky, V . E., Shirinsky, V. P., Gneushev, G. N., and Chernousov, M. A. (1985). Adu. Myocardiol. 5, 2 15-221. Kramarcy, N. R., and Sealock, R. (1990). FEBS Lett. 274, 171-174. Krenacs, T., Molnar, E., Dobo, E., and Dux, L. (1989). Hisrochem. J . 21, 145-155. Krenacs, T., Stiller, D., Krenacs, L., Bahn, H., Molnar, E., and Dux, L. (1990). Acta Hisrochem. (Jena) (Germany) 88, 159-166. Kruger, M., Wright, J., and Wang, K. (1991). J . Cell Biol. 115, 97-107. Kucera. J., and Walro, J. M. (1988). Histochemistry 90, 151-160. Kucera, J . , and Walro, J. M. (1991). Histochemistry 96, 381-390. Kuro-o, M., Tsuchimochi, H., Ueda, S., Takaku, F., and Yazaki, Y. (1986). J. Clin. Inuest. 77, 340-347. Lachapelle, M.. and Aldrich, H. C. (1988). J . Histochem. Cyrochem. 36, 1197-1202. Lakey, A., Ferguson, C., Labeit, S., Reedy, M., Larkins, A., Butcher, G., Leonard, K., and Bullard, B. (1990). EMBO J. 9, 3459-3467. Lamers, W. H., Geerts, W. J . , Moorman, A. F., and Dottin, R. P. (1989). Anat. Embryo/. (Berl.) 179, 387-393. LaSalle, F., Robson, R. M., Lusby, M. L., Parrish, F. C., Stromer, M. H., and Huiatt, T. W. (1983). J. Cell Biol. 97, 258a. Lazarides, E., Granger, B. L., Card, D. L., O’Conner, C. M., Breckler, J., Price. M.. and Danto, S. I. (1982). Cold Spring Harbor Symp. Quant. Biol. 46, 351-378.

PROTEINS IN STRIATED MUSCLE

137

Lazarides, E., and Hubbard, B. D. (1976). Proc. Natl. Acad. Sci. U . S . A . 73,4344-4348. Leberer, E., Charuk, J. H., Clarke, D. M., Green, N. M., Zubrzycka-Gaarn, E., and MacLennan. D. H. (1989). J. Biol. Chem. 264, 3484-3493. Leberer, E., Timms, B. G., Campbell, K. P., and MacLennan, D. H. (1990). J . Biol. Chem. 265, 10,118-10,124. Lee, C. C., Pearlman, J. A.. Chamberlain, J. S . , and Caskey, C. T. (1991). Nature (London) 349, 334-336. Lemanski, L. F., Fuldner, R. A., and Paulson, D. J . (1980). J . Embryol. Exp. Morphol. 55, 1-15.

Lemanski, L. F., and Tu, Z. H. (1983). Deu. Biol. 97, 338-348. Lessard, J. L. (1988). Cell Motil. Cytoskeleton 10, 349-362. Leung, B., Kula, R. W., and Shafig, S. A. (1987). Exp. Neurol. 97, 429-440. Li, J . , and Lernanski, L. F. (1990). Anat. Rec. 228, 46-52. Lim, S.-S., Tu, Z.-H., and Lemanski, L. F. (1984). J . Muscle Res. Cell Motil. 5, 515-526. Lin, J . J.-C., Chou, C. -S., and Lin. J. L.-C. (1985). Hybridoma 4, 223-242. Lin, J. J.-C., and Lin, J. L.-C. (1986). J . Cell Biol. 103, 2173-2183. Lin, Z. Y., Eshleman, J., Grund, C., Fischman, D. A,, Masaki, T., Franke, W. W., and Holtzer, H. (1989a). J. Cell Biol. 108, 1079-1091. Lin, Z., Wenderoth, M. P., and Eisenberg, B. R. (1989b). Am. J . Anat. 185, 455-461. Lowey, S . , Benfield, P. A., LeBlanc, D. D., and Waller, G. S. (1983). J . Muscle Res. Cell Motil. 4, 695-716. Lowey, S., and Steiner, L. A. (1972). J . Mol. Biol. 65, 111-126. Lowey, S., Waller, G., and Bandman, E . (1991). J. Cell Biol. 113, 303-310. Lutz, H., Ermini, M., Jenny, E., and Joris, F. (1978). Aktuel Gerontol. 8, 667-673. Lyons, G. E., Haselgrove, J., Kelley, A. M., and Rubinstein, N. A. (1983). Differentiation 25, 168-175. MacLennan, D. H. (1990). Biophys. J . 58, 1355-1365. Maher, P. A., Cox, G. F., and Singer. S. J. (1985). J . Cell Biol. 101, 1871-1883. Maier, A., Gambke, B., and Pette, D. (1986a). Cell Tissue Res. 244, 635-643. Maier, A., Leberer, E., and Pette, D. (1986b). Histochemistry 86, 63-69. Maier, A., Gambke, B., and Pette, D. (1988a). Histochemistry 88, 267-271. Maier, A., Gorza. L., Schiaffino, S., and Pette, D. (1988b). Cell Tissue Res. 254, 59-68. Maier, A., Leberer, E., and Pette, D. (1988~).Histochemistry 88, 273-276. Maier, A., and Zak, R . (1989). Anat. Rec. 225, 197-202. Maier, A., and Zak, R . (1990). Am. J . Anat. 187, 338-346. Malouf, N. N., Coronado, R., McMahon, D., Meissner, G., and Gillespie, G. Y. (1987). Proc. Natl. Acad. Sci. U.S.A. 84, 5019-5023. Malouf, N . N., Taylor, S. , Gillespie, G. Y., Bynum, J. M., Wilson, P. E., and Meissner, G. (1986). J . Histochem. Cytochem. 34, 347-355. Maruyama, Kei, Kuroda. M., and Nonomura, Y. (1985). Biochim. Biophys. Acta 829, 229-237. Maruyama, Kei, Kurokawa, H., Oosawa, M., Shimaoka, S., Yamamoto, H., Ito, M., and Maruyama, Koscak (1990). J. Biol. Chem. 265, 8712-8715. Maruyama, Koscak, Kirnura, S. , Ishi, T., Kuroda, M., and Ohashi, K. (1977). J. Biochem. (Tokyo) 81, 215-232. Maruyama, Koscak, Kimura, S., Ohasi, K., and Kuwano. Y. (1981). J . Biochem. (Tokyo) 89, 701-709. Maruyama, Koscak, Matsuno, A., Higuchi, H., Shimaoka, S . , Kimura. S., and Shimizu, T. (1989). J . Muscle Res. Cell Motil. 10, 350-359. Maruyama, Koscak, Sawada, H., Kimura, S., Ohashi. K., Higuchi, H., and Umazume, Y. (1984). J . Cell Biol. 99, 1391-1397.

138

MARVIN H. STROMER

Maruyama, Koscak, Yoshioka, T., Higuchi, H., Ohashi, K., Kimura, S., and Natori, R. (1985). J . Cell Eiol. 101, 2167-2172. Masaki, T., Endo, M., and Ebashi, S. (1967). J. Eiochem. 62,630-632. Mascarello, F., Veggetti, A., Cerpene, E., and Rowlerson, A. (1983). J . Anat. 137,95-108. Matsuda, R., Bandman, E., and Strohman, R. C. (1983). Deu. Eiol. 95,484-491. Matsuda, R., Obinata, T., and Shimada, Y. (1981). Dev. Eiol. 82, 11-19. Matsumori, A., Ohkusa, T., Matoba, Y., Okada, I., Yamada, T., Kawai, C., Tamaki, N., Watanabe, Y., Yonekura, Y., Endo, K., Konishi, J., Khaw, B. A,, and Harbor, E. (1989). Circulation 79, 400-405. Matsumori, A., Yamada, T., Tamaki, N., Kawai, C.. Watanabe, Y ., Yonekura, Y., Endo, K., Konishi, J., Yoshida, A., and Tamaki, S. (1990) J p n . Circ. J . 54, 333-338. Matthews, I . , Colyer, J., Mata, A. M., Green, N. M., Sharma, R. P., Lee, A. G.. and East, J. M. (1989). Eiochem. Eiophys. Res. Commun. 161, 683-688. Matthews, I., Sharma, R. P., Lee, A. G., and East, J. M. (1990). J . Eiol. Chem. 265, 18,737- 18,740. McLeod, A. G., Shen, A. C.-Y., Campbell, K. P., Michalak, M.. and Jorgensen, A. 0. (1991). Circ. Res. 69, 344-359. Menold, M. M., and Repasky, E. A. (1984). Muscle Nerue, 7, 408-414. Messina, D. A., and Lemanski, L. F. (1989). Cell Motil. Cytoskeleton 12, 139-149. Miike, T., Miyatake, M., Zhao, J., Yoshioka, K., and Uchino, M. (1989). Brain Dev. 11, 344-346. Miller, J. B., and Stockdale, F. E. (1986a). Proc. Natl. Acad. Sci. U.S.A. 83, 3860-3864. Miller, J. B., and Stockdale, F. E. (1986b). J . Cell Eiol. 103, 2197-2208. Miller, J. B., Teal, S. B., and Stockdale, F. E. (1989). J . Eiol. Chem. 264, 13,122-13,130. Miranda, A. F., Bonilla, E., Martucci, G., Moraes, C. T., Hayes, A. P., and Dimauro, S. (1988). Am. J . Pathol. 132, 410-416. Miranda, A. F., Francke, U., Bonilla, E., Martucci, G., Schmidt, B., Salviati. G., and Rubin, M. (1989). A m . J. Med. Genet. 32, 268-273. Miyanishi, T., Toyoshima, C., Wakabayashi, T., and Matsuda, G. (1988). J. Eiochem. (Tokyo) 103,458-462. Molnar, E., Seidler, N. W., Jona, I., and Martonosi, A. N. (1990). Eiochirn. Eiophys. Acta 1023, 147-167. Moore, A., Hurko, O., and Walsh, F. S. (1984). J . Neuroimmunol. 7, 137-149. Narusawa, M., Fitzsimons, R. B., Izumo, S.. Nadal-Ginard, B., Rubinstein, N. A,, and Kelly, A. M. (1987). J . Cell Biol. 104, 447-459. Nave, R., Fiirst, D. 0.. and Weber, K. (1989). J. Cell Eiol. 109, 2177-2187. Nave, R., and Weber, K. (1990). J . Cell Sci. 95, 535-544. Nelson, W. J., and Lazarides, E. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 363-367. Ngai, P. K . , and Walsh, M. P. (1985). Eiochem. Eiophys. Res. Commun. 127, 533-539. Nguyen, T. M., Ellis, J. M., Ginjaar, I. B., van Paassen, M. M., van Ommen, G.-J. B., Moorman, A. F. M., Cartwright, A. J., and Morris, G. E. (1990). FEES Lett. 272,109-1 12. Nicholson, L. V., Davison, K., Johnson, M. A., Slayter, C. R., Young, C., Bhattacharya, S., Gardner-Medwin, D.. and Harris, J. B. (1989). J . Neurol. Sci. 94, 137-146. Nicholson, L. V . , Johnson, M. A., Gardner-Medwin, D., Bhattacharya, S., and Harris, J. B. (1990). Acta Neuropathol. ( Bed.) 80, 239-250. Nishimura, T., Sada, M., Sasaki, H., Yutani, C., Hayashi, M., Amemiya, H., Fujita, T., Akutsu, T., and Manabe, H. (1987). Eur. J. Nucl. Med. 13, 343-347. Nolan, A. C., Clark, W. A., Jr., Karwoski, T., and Zak, R. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 6046-6050. Nomoto, K., Komuro, I., Kuro-o, M., Tsuchimochi, H., Takaku, F., Machii, K., and Yazaki, Y. (1988). Circ. Res. 62, 1088-1092.

PROTEINS IN STRIATED MUSCLE

139

Obinata, T., Kitani, S., Masaki, T., and Fischman, D. A. (1984a). Deu. Biol. 105,253-256. Obinata, T., Reinach, F. C., Bader. D. M., Masaki, T., Kitani, S., and Fischman, D. A. (1984b). Dev. Biol. 101, 116-124. Offer, G., Starr, R., and Trinick, J. (1988). Adu. Exp. Med. Biol. 226, 61-73. Ohashi, K., Fischman, D. A., Obinata, T., and Maruyama, K. (1981). Biomed. Res. 2, 330-333.

Ohtsuki, I. (1975). J. Biochem. (Tokyo) 77, 633-639. Ohtsuki, I. (1979a). J. Biochem. 85, 1377-1378. Ohtsuki, I. (1979b). J. Biochem. 86, 491-497. Ohtsuki, I., Masaki, T., Nonomura, Y.. and Ebashi, S. (1967). J. Biochem. 61, 817-819. Osborn, M., and Goebel, H. H. (1983). Acta Neuropathol. (Berl.)62, 149-152. Osborn, M., Hill. C., Altmannsberger, M., and Weber, K. (1986). Lab. Invest. 55, 101-108. Osinska, H. E., and Lemanski, L. F. (1989). Anat. Rec. 223, 406-413. Otey, C. A., Kalnoski, M. H., and Bulinski, J. C. (1988). Cell Motil. Cytoskeleton9,337-348. Otsu, N. Hirata, M., Tuboi, S., and Miyazawa, K. (1989). J. Hisrochem. Cytochem. 37, 1465-1470.

Pardo, J. V., Siliciano, J. D.. and Craig, S. W. (1983a). Proc. Narl. Acad. Sci. U . S . A . 80, 1008- I012. Pardo. J. V., Siliciano, J. D., and Craig, S. W. (1983b). J. Cell Biol. 97, 1081-1088. Pedrosa-Domellof, F., Soukup, T., and Thornell, L.-E. (1991). Histochemistry 96,327-338. Pedrosa, F., Butler-Browne, G. S., Dhoot, G. K., Fischman, D. A., and Thornell, L.-E. (1989). Histochemistry 92, 185-194. Pedrosa, F.. Soukup, T., and Thornell, L.-E. (1990). Hisrochemistry 95, 105-113. Pellissier, J. F., Pouget, J., Charpin, C., and Figarella, D. (1989). J. Neurol. Sci. 89, 49-61. Pepe, F. A. (1966). J. CellBiol. 28, 505-525. Pepe, F. A. (1967a). J. Mol. Biol. 27, 203-225. Pepe, F. A. (3967b). J. Mol. Biol. 27, 227-236. Pepe, F. A. (1968). Int. Rev. Cyrol. 24, 193-231. Pepe, F. A. (1971). In “Progress in Biophysics and Molecular Biology” ( J . A. V. Butler and D. Noble, eds.), Vol. 22, pp. 75-96. Pergamon, New York. Pepe, F. A. (1983). In “Handbook of Physiology” (L. D. Peachey, R. H. Adrian, and S. R. Geiger, eds.), Chap. 10, pp. 113-141. American Physiological Society, Bethesda, MD. Pette, D., and Staron, R. S. (1990). Rev. Physiol. Biochem. Pharmacol. 116, 1-76. Phillips, W. D., Everett, A. W., and Bennett, M. R. (1986). J. Neurocytol. 15, 397-405. Pierobon-Bormioli, S., Betto, R., and Salviati, G. (1989). J. Muscle Res. Cell Motil. 10, 446-456.

Pollock, M., and Atkinson, B. G. (1985). Can J. Biochem. Cell Biol. 63, 430-438. Pomeroy, M. E., Lawrence, J. B., Singer, R. H., and Billings-Gagliardi,S. (1991). Dev. Biol. 143, 58-67.

Pons, F., Augier, N., Heilig, R.,Leger, J., Mornet, D., and Leger, J. J. (1990). Proc. Narl. Acad. Sci. U . S . A . 87, 7851-7855. Pons, F., Leger, J. O., Chevallay, M., Tome, F. M., Fardeau, M., and Leger, J. J. (1986). J. Neurol. Sci. 76, 151-163. Price, M. G. (1987). J. Cell Biol. 104, 1325-1336. Price, M. G., and Lazarides, E. (1983). J. Cell Biol. 97, 1860-1874. Pruliere, G., Butler-Browne, G. S., Cambon, N., Toutant, M., and Whalen, R. G. (1989). Eur. J . Biochem. 185, 555-561. Quinn, L. S., and Nameroff, M. (1983). Differentiation 24, 111-123. Rappaport, L., Samuel, J. L., Bertier, B., Bugaisky, L., Marotte, F., Mercadier, A., and Schwartz, K. (1984). Eur. Heart J. 5(Suppl. F), 243-250. Rappaport, L., Samuel, J. L., Bertier-Savelle, B., Marotte, F., and Schwartz, K. (1985). Basic Res. Cardiol. 8O(Suppl. I), 129-132.

140

MARVIN H. STROMER

Rappaport, L., Swynghedauw, B., Mercadier, J. J., Lompre, A. M., de la Bastie, D., Samuel, .I. L., and Schwartz, K. (1986). Fed. Proc. 45, 2573-2579. Reinach, F. C., Masaki, T.. Shafiq, S., Obinata, T . , and Fischman, D. A. (1982). J . Cell Biol. 95, 78-84. Repasky, E. A., Granger, B. L., and Lazarides, E. (1982). Cell (Cambridge, Mass.) 29, 821-833. Repasky, E. A., Pollina, C. M., Menold, M. M., and Hudecki, M. S. (1986). Proc. Natl. Acad. Sci. U . S . A . 83, 802-806. Richardson, F. L., Stromer. M. H., Huiatt, T. W., and Robson, R. M. (1981). Eur. J . Cell Biol. 26, 91-101. Rogalski, A. A. (1987). J. Cell Biol. 105, 819-831. Rosemblatt, M. S., and Scales, D. J. (1989). Mol. Cell. Biochem. 87, 57-69. Rossi, A. M., Eppenberger, H. M., Volpe, P., Cotrufo. R., and Wallimann, T. (1990). J . Biol. Chem. 265, 5258-5266. Rouayrenc, J. F., Fattoum, A., Gabrion, J., Audemard, E., and Kassab, R. (1984). FEBS Lett. 167, 52-58. Rubinstein, N., Mabuchi, K., Pepe, F., Salmons, S., Gergely, J., and Sreter, F. (1978). J . Cell Biol. 79, 252-261. Saad, A. D., Obinata, A., and Fischman, D. A. (1987). Deu. Biol. 119, 336-349. Sabry, M. A., and Dhoot, G. K. (1991). J . Muscle Res. Cell Motil. 12, 262-270. Saetersdal, T., Dalen, H., and Roli, J. (1989). Histochemistry 92, 467-473. Saide, J. D. (1981). J . Mol. Biol. 153, 661-679. Saide, J. D., Chin-Bow, S., Hogan-Sheldon, J., Busquets-Turner. L., Vigoreaux, J. 0.. Valgeirsdottir, K., and Pardue, M. L. (1989). J. Cell. B i d . 109, 2157-2167. Saku, T., Tsuda, N., Anami, M., and Okabe, H. (1985). Acta Paihol. Jpn. 35, 125-136. Samuel, J. L., Bertier, B., Bugaisky, L., Marotte, F., Swynghedauw, B., Schwartz, K., and Rappaport, L. (1984). Eur. J. Cell Biol. 34, 300-306. Samuel, J. L., Jockusch, B., Bertier-Savalle, B., Escoubet, B., Marotte, F., Swynghedauw, B., and Rappaport, L. (1985). Basic Res. Cardiol. 8O(Suppl. 2), 119-122. Samuel, J. L., Rappaport, L., Mercadier, J. J., Lompre, A. M., Sartore, S., Triban, C., Schiaffino, S., and Schwartz, K. (1983). Circ. Res. 52, 200-209. Samuel, J. L., Rappaport, L., Syrovy, I . , Wisnewsky, C., Marotte, F., Whalen. R. G., and Schwartz, K. (1986). Am. J. Physiol. 250, 333-341. Sartore, S. (1981). Biochim. Biophys. Acta 667, 143-156. Sartore, S., Gorza, L., Pierobon Bormioli, S., Dalla Libera, L., and Schiaffino, S. (1981). J . Cell Biol. 88, 226-233. Sartore, S., Gorza, L., and Schiaffino, S. (1982). Nature (London) 298, 294-296. Sartore, S., Mascarello, F., Rowlerson. A., Gorza, L., Ausoni, S., Vianello, M., and Schiaffino, S. (1987). J. Muscle Res. Cell Motil. 8, 161-172. Sawchak, J. A., Lewis, S., and Shafiq, S. A. (1989). Muscle Nerve 12,679-689. Sawtell, N. M., and Lessard, J. L. (1989). J . Cell Biol. 109, 2929-2937. Scapolo, P. A., Costerbosa, G. L., Barazzoni, A. M., Lucchi, M. L., and Bortolami. R. (1990). Anat. Rec. 227, 314-320. Schaart, G., Viebahn, C., Langmann, W.. and Ramaekers, F. (1989). Deuelopment 107, 585-596. Schger, B. W., and Perriard, J. C. (1988). J . Cell Biol. 106, 1161-1170. Schafer, B., Perriard. J. C., and Eppenberger. H . M. (1985). Basic Res. Cardiol. 8O(Suppl. 2), 129-133. Schantz, P. G., and Dhoot, G. K. (1987). Acra Physiol. Scund. 131, 147-154. Schiaffino, S., Gorza, L., Dones, I., Cornelio, F., and Sartore, S. (1986). Muscle Nerve 9, 51-58.

PROTEINS IN STRIATED MUSCLE

141

Schiaffino, S ., Gorza, L., Pitton, G., Saggin, L., Ausoni. S., Sartore, S., and Lomo, T. (1988). Deu. Biol. U7, 1-1 I . Schiaffino, S.. Gorza. L.. Saggin. L.. Valfre, C.. and Sartore, S. (1984). Eur. Henrt 1. ~ ( S U P P IF), . 95-102. Schiaffino, S . , Gorza, L., Sartore, S . , Saggin. L., Ausoni. S . , Vianello. M., Gunderson, K., and Lomo, T . (1989). J. Muscle Res. Cell Motil. 10, 197-205. Schmitt, T. L., and Pette, D. (1991). Histochemistry 96, 459-465. Schollmeyer, J. E., Goll, D. E., Robson, R. M., and Stromer, M. H. (1973). J. CellBiol. 59, 306a. Schollmeyer, J. E., Stromer, M. H., and Goll, D. E. (1972). Biophys. J. 12, 280a. Schollmeyer, J. V., Goll. D. E., Stromer, M. H., Dayton, W.. Singh, I . , and Robson, R. M. (1974). J. Cell Bid. 63, 303a. Schroder, J. M., Sommer, C., and Schmidt, B. (1990).Acta Neuropathol. (Berl.)80,406-414. Schultheiss, T., Lin, Z., Lu, M.-H., Murray, J., Fischman, D. A., Weber, K.. Masaki, T., Imamura, M., and Holtzer. H. (1990). J. Cell Bid. 110, 1159-1 172. Schultheiss, T., Zhongxiang, L., Ishikawa, H., Zarnir. I.. Stoeckert, C. J., and Holtzer, H. (1991). J . Cell Bid. 114, 953-966. Sealock, R., Paschal, B., Beckerle, M., and Burridge, K. (1986).Exp. CellRes. 163,143-150. Seidal, T., Kindblom, L. G., and Angervall, L. (1987). Appl. Pathol. 5, 201-219. Seko, Y., Naito, S., Imataka. K., Fujii, J., Nakane, P. K.. Takaku, F.. and Yasaki. Y. (1990). Cell Biochem. Funct. 8, 117-130. Semper, A. E., Fitzsimons, R. B., and Shotten, D. M. (1988). J . Neurol. Sci. 83, 93-108. Shear, C. R., Bandman, E., and Rosser, B. W. (1988). Deu. Biol. U7, 326-337. Shear, C. R., and Bloch, R. J. (1985). J. Cell Bid. 101, 240-256. Shen, P. S., and Lemanski, L. F. (1990). J . Morphol. 201, 243-252. Shimizu, T., Dennis, J. E., Masaki, T., and Fischman. D. A. (1985a). J. Cell Biol. 101, 1115-1 123. Shimizu, T., Reinach, F. C., Masaki, T., and Fischman, D. A. (1985b). J. Mol. Biol. 183, 271-282. Shimizu, N., and Shimada, Y. (1985). Deu. Biol. 111, 324-334. Sikorska, H., Rousseau. J.. Desputeau, C., Gervais. A., Savoie, S., Ghaffari, M. A., Bisson, L., and van Lier, J. E. (1990). In?. J . Rad. Appl. Instrum. (B) (England) 17, 567-584. Skalli, O., Gabbiani, G., Babai, F., Seemayer. T. A.. Pizzolato, G., and Schurch, W. (1988). Am. J. Pathol. 130, 515-531. Starr, R., Almond, R., and Offer, G. (1985). J. Muscle Res. Cell Moril. 6, 227-256. Starr, C. M.. Diaz, J. G., and Lemanski, L. F. (1989). J . Morphol. 201, 1-10. Stedman, H., Browning, K.. Oliver, N., Oronzi-Scott, M., Fischbeck, K., Sarkar, S.. Sylvester, J., Schmickel, R., and Wang, K. (1988). Genomics 2, 1-7. Strehler, E. E., Carlsson, E., Eppenberger, H . M., and Thornell, L.-E. (1983). J. Mol. Bid. 166, 141-158. Strehler, E. E., Strehler-Page, M.-A., Perriard, J.-C., Periasamy, M., and Nadal-Ginard, B. (1986). J. Mol. Biol. 190, 291-317. Stromer, M. H. (1990). I n “Cellular and Molecular Biology of Intermediate Filaments” (R. D. Goldman and P. M. Steinert, eds.), pp. 19-36. Plenum, New York. Sugi, Y. (1989). J. Electron Microsc. (Tokyo) 38, 126-131. Sugi, Y., and Hirakow, R. (1991). Cell Tissue Res. 263, 459-469. Sugita, H., Arahata, K., Tsukahara, T., and Ishiura, S. (1990). In “Pathogenesis and Therapy of Duchenne and Becker Muscular Dystrophy” (B. A. Kakulas and F. L. Mastaglia, eds.). pp. 33-44. Raven Press, New York. Sumida, H., Nakamura. H., Akimoto, N., Okamoto, N., and Satow, Y. (1987). Arch. Histol. Jpn. 50, 525-53 1.

142

MARVIN H. STROMER

Sunohara, N., Arahata, K., Hoffman, E. P., Yamada, H., Nishimiya, J., Arikawa, E., Kaido, M. Nonaka, I., and Sugita, H. (1990). Ann. Neurol. ( U . S . A . )28,634-639. Sutko, J. L., Airey, J. A., Murakami, K., Takeda, M., Beck, C., Deerinck, T., and Ellisman, M. H. (1991). J . Cell B i d . 113, 793-803. Sutoh, K., Tokunaga, M., and Wakabayashi, T. (1987). J. Mol. Biol. 195, 953-956. Swasdison, S., and Mayne, R. (1989). Cell Tissue Res. 257, 537-543. Sweeney, L. J., Clark, W. A., Jr. Umeda, P. K., Zak, R., and Manasek, F. J. (1984). Proc. Natl. Acad. Sci. U.S.A. 81, 797-800. Sweeney, L. J., and Kelley, S. W. (1990). J . Mol. Cell. Cardiol. 22, 361-370. Sweeney, L. J., Kennedy, J. M., Zak, R., Kokjohn, K., and Kelley, S. W. (1989). Deu. Biol. 133, 361-374. Sweeney, L. J., Nag, A. C., Eisenberg, B., Manasek, F. J., and Zak, R. (1985). Basic Res. Cardiol. 8O(Suppl. 2). 123-127. Sweeney, L. J., Zak, R., and Manasek, F. J. (1987). Circ. Res. 61, 287-295. Tachi, N., Sasaki, K., Yamada, T., Imamura, S., and Mike, T. (1990). Pediatr. Neurol. 6 , 54-56. Takeshima, H., Nishimura, S., Matsumoto, T., Ishida, H., Kangawa, K., Minamino, N., Matsuo, H., Ueda, M., Hanaoka, M., Hirose, T., and Numa, S. (1989). Nature (London) 339, 439-445. Tassin, A. M., Pincon-Raymond, M., Paulin, D., and Rieger, F. (1988). Deu. B i d . 129, 37-47. Tao, J. -X.,and Ip, W. (1991). Cell Motil. Cytoskeleton 19, 109-120. Taylor, L. D., and Bandman, E. (1989). J. Cell Biol. 108, 533-542. te Kronnie, G., Pool, C. W. Scholten, G . , and van Raamsdonk, W. (1980). Eur. J. Cell Biol. 22,772-779. Terai, M., Komiyama, M., and Shimada, Y. (1989). Cell Motil. Cytoskeleton 12, 185-194. Terracio, L., Gullberg, D., Rubin, K., Craig, S., and Borg, T. K. (1989). Anat. Rec. 223, 62-7 1. Terracio, L., Simpson, D. G., Hilenski, L., Carver, W., Decker, R. S., Vinson, N., and Borg, T. K. (1990). J. Cell Physiol. 145, 78-87. Thornell, L.-E., Edstrom, L., Eriksson, A., Hendriksson, K.-G., and Angqvist, K.-A. (1980). J. Neurol. Sci. 47, 153-170. Thornell, L.-E., Billeter, R., Eriksson, P. O., and Ringqvist, M. (1984a). Arch. Oral B i d . 29, 1-5. Thornell, L.-E., Johansson, B., Eriksson, A., Lehto, V.-P., and Virtanen, I. (1984b). Eur. Heart J. S(Suppl. F), 231-241. Thornell, L.-E., Carlsson, E., and Pedrosa, F. (1990). In “The Dynamic State of Muscle Fibers” (D.Pette, ed.), pp. 369-383. de Gruyte, New York. Tidball, J. G., O’Halloran, T., and Burridge, K. (1986). J. Cell Biol. 103, 1465-1472. Tokunaga, M., Suzuki, M., Saeki, K., and Wakabayashi, T. (1987). J. Mol. Biol. 194, 245-255. Tokuyasu, K. T. (1983a). J. Cell Biol. 97, 562-565. Tokuyasu, K. T. (1983b). J . Histochem. Cytochem. 31, 164-167. Tokuyasu, K. T. (1989). J. Cell B i d . 108, 43-53. Tokuyasu, K. T., Dutton, A. H., Geiger, B., and Singer, S. J. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 7619-7623. Tokuyasu, K. T.. Dutton, A. H., and Singer, S. J. (1983). J. Cell Biol. 96, 1727-1735. Tokuyasu, K. T., and Maher, P. A. (1987a). J. Cell Biol. 105, 2781-2793. Tokuyasu, K. T., and Maher, P. A. (1987b). J. Cell Biol. 105, 2795-2801. Tokuyasu, K. T., Maher, P. A., and Singer, S. J. (1984). J. Cell B i d . 98, 1961-1972. Tokuyasu, K. T., Maher, P. A., and Singer, S. J. (1985a). J. Cell Biol. 100, 1157-1 166.

PROTEINS IN STRIATED MUSCLE

143

Tokuyasu, K. T., Maher, P. A., Dutton, A. H., and Singer, S. J . (1985b). Ann. N . Y . Acad. Sci. 455, 200-212. Toyota, N., and Shimada. Y. (1981). J . Cell Biol. 91, 497-504. Toyota, N., and Shimada, Y. (1983). Cell (Cambridge, Mass.) 33, 297-304. Toyota, N., and Shimada, Y. (1984). Cell Tissue Res. 236, 549-554. Trinick, J. (1991). Curr Opinion Cell B i d . 3, 112-1 19. Trombitas, K., Baatsen, P. H. W. W., Lin, J. J.-C., Lemanski, L. F., and Pollack, G. H. (1990a). J. Muscle Res. Cell Motil. 11, 445-452. Trombitas, K., Pollack, G. H., Wright, J., and Wang, K. (1990b). Proc. l2th Int. Cong. Electron Microsc. 3, 478-479. Tsuchimochi, H., Kuro-o, M., Koyama, H., Kurabayashi, M., Sugi, M., Takaku, F., Furuta, S., and Yasaki, Y. (1988). J . Clin.Inuest. 81, 110-1 18. Tsuchimochi, H., Sugi. M., Kuro-o, M., Ueda, S.. Takaku, F., Furuta, S., Shirai, T., and Yazaki, Y. (1984). J . Clin. Inuest. 74, 662-665. Uchino, M., Araki, S., Miike, T., Teramoto, H., Nakamura, T., and Yasutaki, T. (1989). Muscle Nerve 12, 1009-1016. van den Berge, H., and Wirtz, P. (1989a). J. Anat. 164, 215-228. van den Berge, H., and Wirtz, P. (1989b). J . Anat. 166, 157-169. Vigoreaux, J. O., Saide, J . D., and Pardue, M. L. (1991). J . Muscle Res. Cell Motil. 12, 340-354. Virtanen, I., Narvanen, O., and Thornell, L.-E. (1990). FEBS Lett. 267, 176-178. Vivarelli, E., Brown, W. E., Whalen, R. G., and Cossu. G. (1988). J . Cell B i d . 107, 2 191-2 197. Volk, T., and Geiger, B. (1986a). J. CellBiol. 103, 1441-1450. Volk, T., and Geiger, B. (1986b). J. Cell B i d . 103, 1451-1464. Wachsberger, P., Lampson, L., and Pepe, F. A. (1983). Tissue Cell 15, 341-349. Wakayama, Y., Jimi, T., Misugi, N., Kumagai, T., Miyake, S.. Shibuya, S.. and Miike, T. (1989). J. Neurol. Sci. 91, 191-205. Wakayama, Y.. and Shibuya, S. (1991a). Acta Neuropathol. 82, 178-184. Wakayama, Y., and Shibuya, S. (1991b). J. Electron Microsc. 40, 143-145. Waller, G. S.. and Lowey, S. (1985). J. Biol. Chem. 260, 14.368-14,373. Wallimann, T. Doetschman, T. C., and Eppenberger, H. M. (1983). J. Cell Biol. 96, 1771- 1779. Wallimann, T., and Eppenberger, H . M. (1985). Cell Muscle Motil. 6, 239-285. Wallimann, T.. Pelloni, G. W., Turner, D. C., and Eppenberger, H. M. (1978). Proc. Natl. Acad. Sci. U.S.A. 75,4296-4300. Wallimann, T., Turner, D. C., and Eppenberger, H. M. (1977). J. Cell B i d . 75, 297-317. Wang, K. (1985). I n “Cell and Muscle Motility” ( J . W. Shay, ed.), Vol. 6. pp. 315-369. Plenum, New York. Wang, K., McCarter, R., Wright, J., Beverly, J., and Ramerez-Mitchell, R. (1991). Proc. Natl. Acad. Sci. U . S . A . 88, 7101-7105. Wang, K., McClure, J., and Tu, A. (1979). Proc. Nut/. Acad. Sci. U . S . A . 76, 3698-3702. Wang, K., and Williamson, C. (1980). Proc. Natl. Acad. Sci. U . S . A . 77, 3254-3258. Wang, K., and Wright, J. (1988). J. Cell B i d . 107, 2199-2212. Wang, S.-M., and Greaser, M. L. (1985). J. Muscle Res. Cell Motil. 6, 293-312. Wang, S.-M., Greaser, M. L., Schultz, E., Bulinski. J . C.. Lin. J. J.-C., and Lessard, J. L . (1988). J. Cell B i d . 107, 1075-1083. Wang, S.-M., Sun, M.-C., and Jeng, C.-J., (1991). Biochem. Biophys. Res. Commun. 176, 189-193. Watkins, S. C., Hoffman, E. P., Slayter, H. S., and Kunkel, L. M. (1989). Muscle Nerve 12. 861-868.

144

MARVIN H. STROMER

Watkins, S. C., Samuel, J. L., Marotte. F., Bertier-Savalle, B.. and Rappaport, L. (1987). Circ. Res. 60, 327-336. Webster, C., Silberstein, L.. Hays, A . P., and Blau, H. M. (1988). Cell (Cambridge, M a s s . ) 52,503-513. Wenderoth, M. P., and Eisenberg, B. R. (1987). J . Cell Biol. 105, 2771-2780. Wenderoth, M. P.. and Eisenberg. B. R . (1991). J . Hisrochem. Cytochem. 39, 1025-1033. Wessels. A . , Ginjaar, I. B.. Moorman, A. F. M., and van Ommen, (3.4.B. (1991a). Muscle Nerve 14, 1-7. Wessels, A. Vermeulen, J. L. M., Viragh, F.. Kalman, V . F.. Lamers, W. H., and Moorman, A. F. M. (1991b). Anar. Rec. 229, 355-368. Whalen, R. G . , Johnstone, D., Bryers, P. S . , Butler-Browne. G. S . , Ecob, M. S., and Jaros, E. (1984). FEBS Lett. 177, 51-56. Whiting, A . , Wardale, J., and Trinick. J . (1989). J . Mol. Biol. 205, 263-268. Wieczorek, D. F., Periasamy, M., Butler-Browne. G. S. , Whalen, R. G., and Nadal-Ginard, B. (1985). J . Cell B i d . 101, 618-629. Williams, B. A., Huang, S.-C., Krivokapich, J.. Buxton, D. B.. and Schelbert, H. R. (1991). Am. J . Physiol. 261, H319-H326. Wilson, F. J., Camiscoli, D.. Irish, M. J., and Hirabayashi, T. (1978). J . Histochem. Cytochem. 26, 258-266. Winkelmann, D. A., Lowey. S. , and Press. J. L. (1983).Cell(Cambridge,Mass.)34,295-306. Yablonka-Reuveni, Z., and Nameroff, M. (1990).Diffrrentiation 45, 21-28. Yagyu, M., Robson, R. M., and Stromer, M. H. (1990). J . Cell Biol. 111, 42a. Yorifuji, H., and Hirokawa, N. (1989). J . Electron Microsc. Tech. 12, 160-171. Young, 0. A. (1989). J . Muscle Res. Cell Motil. 10, 403-412. Yuan, S . , Arnold, W., and Jorgensen, A . 0. (1990). J . Cell Biol. 110, 1187-1 198. Zadeh, B. J., Gonzalez-Sanchez, A., Fischman, D. A . , and Bader, D. M. (1986). Deu. B i d . 115, 204-214. Zaret, B. L., and Wackers, F. J. Th. (1991). Mod. Concepts Cardiouasc. Dis. 60, 37-42. Zernig, G . , and Wiche, G. (1985). Eur. J . Cell Biol. 38, 113-122. Zhang. Y., Lewis, S. G . , and Shafiq, S. A. (1989). Histochemistry 91, 143-149. Zhang, Y., Rushbrook, J. I., and Shafiq, S. A. (1985).J . Muscle Res. Cell Moril. 6,333-345. Zhang, Y., Sher, J. H., Leung, B., and Shafiq, S. A . (1987). J . Neurol. Sci. 80, 1-12. Zhang, Y. T., and Shafiq, S. A. (1987). J . Exp. Zoo/. 243, 51-62. Zorzato, F., Chu, A., and Volpe, P. (1989). Biochem. J . 261, 863-870. Zubrzycka-Gaarn, E. E., Bulman, D. E . , Karpati, G., Burghes, A . H. M., Belfall, B.. Klamut, H. J., Talbot, J., Hodges, R. S. , Ray, P. N., and Worton, R. G. (1988). Nature (London) 333,466-469. Zubrzycka-Gaarn, E. E., Hutter, 0. F., Karpati, G., Klamut. H. J., Bulman, D. E.. Hodges, R. S., Worton, R. G., and Ray, P. N. (1991). Exp. Cell Res. 192, 278-288.

Recent Developments in Vertebrate Cell Culture Technology Satish J. Parulekar,' Thomas Hassell,t and Satish C. Tripathit ' Department of Chemical Engineering, Illinois Institute of Technology, Chicago, Illinois 60616 t Celltech Limited, Slough, Berkshire, United Kingdom Department of Life Sciences, IIT Research Institute, Chicago, Illinois 60616

*

1. Introduction Structural and functional relationships are of utmost importance in the study of living beings, including vertebrates. The studies involving these relationships require a detailed and in-depth investigation of various organs and the tissues and cell types that constitute them. One way to carry out such studies would be to opt for in vivo systems. In vivo studies present us with the real three-dimensional geometry coupled with the advantage of intercellular matrix, functional association of various cell types, tissues, and organs and highly coordinated physiological and biochemical characteristics. However, there are some inherent problems associated with in vivo systems. The other option is to look for in virro model systems. In vitro techniques are easier to perform than in vivo techniques, can yield homogeneous population of cells, are highly reproducible, and are relatively inexpensive. Three types of model systems have been employed in prior literature, viz., individual cell cultures (two-dimensional or traditional cultures), group cell cultures (three-dimensional cultures), and organ cultures (three-dimensional, in vivo-like, cultures). Just as the discovery of anesthesia was responsible for modern advances in surgery, the development of cell culture techniques has provided a new dimension to the thinking, understanding, and interpretation of today's biology. Advances in biotechnology in the past two decades have made possible the production of diagnostic and therapeutic biochemicals that promise the potential to control or eradicate some of mankind's most devastating diseases. Biochemicals manufactured from animal cells in culture have lnrrrnurionul Reuiriv oJ (pfoloyy. V d . 142

145

Copyright Q 1992 by Academic Press. Inc. All rights of reproduction in any form reserved.

146

SATISH J. PARULEKAR ET AL.

importance in medical and clinical fields (Spier and Horaud, 1985). These products include a wide variety of proteins, such as vaccines, blood chemistry compounds, hormones and endocrines, monoclonal antibodies (mAbs), anti-cancer agents, and other therapeutic biomolecules (Feder and Tolbert, 1983; Glacken et al., 1983a,b; Hu and Dodge, 1985; Katinger, 1988; Pollard and Khosrovi, 1978; Runstadler and Cernek, 1988; Spier and Horaud, 1985). A wide array of potential production cell strains and cell lines has been derived either by direct isolation of cells from normal or transformed tissues or by the application of the various techniques of cell strain improvement in uitro (Katinger, 1988).A new and highly competitive biotechnology industry has emerged to exploit the commercialization of these products. Much of the activity in the biotechnology industry during the past decade has focused on the early stages of product development-genetic engineering, the production of small amounts of diagnostic and therapeutic products, and preclinical or early clinical trials to test the efficacy and safety of these biochemicals. Now the biotechnology industry is beginning to focus upon the large-scale bioprocessing-the manufacture of finished biochemical products made by living cells-that is necessary for the successful commercial introduction of natural and recombinant proteins at reasonable cost. Proteins with complex tertiary or quaternary structure or proteins that require post-translational modification may not be properly expressed in bacteria and hence must be produced by genetically engineered animal cells (Colbere-Garapin and Garapin, 1985). Some of the proteins produced by recombinant mammalian cells are hepatitis B virus surface antigen (Michel et al., 1985), glycoprotein D of herpes simplex virus (Laskey et al., 1984), and tissue-type plasminogen activator (tPA). Consequently, hybridoma cells and genetically altered animal cells are increasingly being used as host organisms for the production of these health proteins. As a consequence of the generally low specific productivity per cell by animal cells for these proteins, and the large volume of therapeutic-grade material that will often be required, the biotechnology industry is now beginning to address the need for cost-efficient, animal-cell culture technologies. Vaccines constitute the largest market outlet of any biological produced by animal cells, with foot-and-mouth disease (FMD) vaccine being the principal product. Human vaccines produced from mammalian cells include polio, rubella, rabies, measles, and encephalitis (Glacken et al., 1983a,b; Jacobs, 1979). Most of these products have thus far been low tonnage, high-cost items manufactured in small, batch systems. However, as the markets expand, the incentives for development of production methods more suitable for large-scale operations will increase (Pollard and Khosrovi, 1978). For producing genetically engineered proteins by the use of mammalian cells, it is desirable to use cells that are immortal. Using a continuous and

VERTEBRATE CELL CULTURE TECHNOLOGY

147

transformed cell line, Chinese hamster ovary (CHO) cells as the host, human immune interferon has been produced in an in uitro mammalian cell culture system. Another example using immortalized cells is the antibodysecreting hybridomas. Lymphocytes, which produce antibodies of interest, are often mortal. These can be fused with a cancerous type, myeloma, which is immortal but does not produce antibodies of practical use. By fusing the two types of cells and properly selecting the hybrids (hybridomas), cell lines that produce antibodies of interest and are immortal can be obtained (Hu and Dodge, 1985). In addition to diagnostic applications, mAbs have already been used on a laboratory scale for the single-step purification of many proteins (Staeheheln et al., 1981) and have significant potential in large-scale biological separations (Secher and Burke, 1980). Additionally, monoclonal antibodies have significant therapeutic value in the treatment of tumors and of various immunological diseases (Olsson and Mathe, 1982; Ritz and Schlossman, 1982; Vitetta et al., 1983). The complexity of animal cell culture is further illustrated by the fact that the supplementation of the medium with serum is usually necessary for cell growth. The most commonly used sera are of horse or bovine origin. The elucidation of the role of serum has been facilitated by the development of “defined” or serum-free media for animal cell cultivation. Most of these media are supplemented with a number of hormones or other growth factors (Bettger et al., 1981; Murakami et al., 1982). The supplemented hormones or growth factors are involved in nutrient transport, maintenance of cellular energy balance, control of macromolecular synthesis, and stimulation of product formation (Mather et al., 1981). Since many of these supplemented factors are purified from serum, the application of serum-free medium to large-scale cell culture may not be economical, except when the downstream product isolation is made much less complicated by the use of serum-free medium. The status of various vertebrate cell culture-related technologies developed so far is reviewed here. Considerations involved in commercial or industrial realization of the concepts developed in a biologist’s laboratory are reviewed in detail. While going over the history of this technology, some of the major technical breakthroughs in this century-old technology, future commercial expectations from this technology, and societal implications of this technology are also discussed.

II. Traditional Cultures A. Background

Animal cells can generally be categorized according to their anchoragedependence. Some cell types can grow in suspension, whereas some cell

140

SATISH J. PARULEKAR ET AL.

types must attach to a compatible surface and spread out in order to grow. Others can grow either in suspension or anchored to a surface. Most cells in this third category, such as HeLa cells, are transformed cells derived from anchorage-dependent cell types (Hu and Dodge, 1985). The development of vertebrate cell culture can be traced back to some of the earliest observations on cultivation of chick embryo cells in warm saline (Roux, 1885). The cultivation of animal tissue can be traced back to a study by Harrison (1907), who devised a method for direct observation of a growing nerve. Nerve tissue was dissected from frog embryos and successfully grown in a “hanging drop” of clotted frog lymph. The technique enabled observation of the rate and nature of growth of nerve fibers over periods ranging from 1 to 4 weeks. Burrows (1910) improved the technique by substitution of frog lymph by chicken plasma clot and later, with Carrel (1912), applied the technique to the study of warm-blooded animals. Carrel pioneered further development of vertebrate-cell cultures by demonstrating the repeated subculturing of a strain of chick embryo cells in the Carrel flask in 1923 (Witkowski, 1979). The development of continuous cell lines of human tumors, i.e., HeLa (Gey et al., 1952), generated significant interest in human tissue culture. Classical studies by Hayflick and Moorhead (1961) with cells of finite life span further aided the establishment of cell lines with the true number of human chromosomes. The post- 1950 period represents rapid expansion in the development of vertebrate-cell culture technology. A series of key innovations led to accelerated development of this technology and applications of the techniques of vertebrate-cell culture on a wide basis. The development of media (Eagle, 1955, 1959; Earle et al., 1951; Fischer et al., 1948; Ham, 1965; McCoy et al., 1959) for cultivation of animal cells was instrumental in this rapid expansion. The applicability of animal-cell culture techniques was broadened further by other major developments including the use of antibiotics to eliminate or minimize contamination, the development of techniques for trypsin-aided passage of cells (Moscona and Moscona, 1952), and technical improvements in the supply of media and serum. The increased availability and applicability of these techniques led to a tremendous increase in both scale and number of applications of vertebrate-cell cultures. The standardization of culture conditions for cultivation of vertebrate-cell systems and cell lines for these has facilitated assay and production of viruses and cells in large numbers, which makes their biochemical analysis more convenient. Research in cell biology, molecular biology, and virology and cancer-related research have depended heavily on cell culture techniques. The introduction of cell fusion techniques (Harris and Watkins, 1965; Littlefield, 1964; Soreiul and Ephrussi, 1961) established somatic cell genetics as a major component in the genetic

VERTEBRATE CELL CULTURE TECHNOLOGY

149

analysis of higher animals, and contributed greatly, via the monoclonal antibody technique (Kohler and Milstein, 1975), to the study of immunology.

6 . Anchorage-Dependent Cultures Using a trypsin solution, Rous and Jones (1916) were able to disperse cells from tissue and grow them in a clot. They subsequently redispersed them from the clot and grew them again. Normal diploid cells exhibit a variety of social behaviors. Both cell movement and cell multiplication are inhibited at high cell density when cell-cell contact is established (Abercrombie and Heaysman, 1954; Stoker and Rubin, 1967). Thus, a “monolayer” of cells is formed when the growth surface is completely covered with cells. At this stage, cells need to be detached, usually by treatment with a proteolytic agent like trypsin, and transferred to a larger surface area for further multiplication (Hu and Dodge, 1985). For the cultivation of anchorage-dependent cells, scale-up often depends on the increase in total surface area. To cultivate anchorage-dependent cells, a compatible wettable surface such as glass or plastic is required for attachment to occur (McKeehan et al., 1981).The most commonly used devices for anchoragedependent cell growth are Petri dishes, tissue culture flasks, and roller bottles. A 1-liter roller bottle provides approximately 500 cm2 of surface area on the inner surface of the bottle wall for cell attachment and growth. The bottles are laid on a roller apparatus in an incubator and rotated at a low speed. Typically, each bottle contains approximately 100 ml of medium. Thus, when bottles rotate, cells growing on the wall are alternately exposed to medium and oxygen in gas phase. These devices are for laboratory and small-scale production use, as they provide only a small surface area (for cell growth) per unit vessel volume. The number of roller bottles required for large-scale production of cells is staggering. In addition, the variation in the cell cultures from a series of roller bottles makes the monitoring of cellular kinetics and control of the growth environment to obtain optimal production rates practically impossible (Glacken et al., 1983a,b). Surface-to-volume ratios for other culture systems employed for cultivation of anchorage-dependent vertebrate cells are provided in Table I. Anchorage-dependent cells are frequently chosen for vertebrate cell cultures instead of the more easily grown freely suspended cells because of their genetic stability and normal diploid nature. These factors have been considered to be particularly important in the production of humaninjectable products, for which all aspects of product safety are under constant review. Diploid and primary cells from solid tissues have evolved

150

SATISH J. PARULEKAR ET AL.

TABLE I Surface-to-Volume Ratios (Si V, cm-’) of Various Methods Used to Culture Anchorage-Dependent Cells

Material

SI v

Reference ~

Inclined roller bottles Gyrogen with tubes Multiple propagator Spiral film Plastic bags Jensen’s IL410 tubular spiral film Glass-bead propagator Artificial capillaries Microcarriers (20 g/liter) (25 g/liter)

0.2-0.7 I .2 1.7 4.0 5.0 9.4 10.0 30.7 I22 153

Bailey and Ollis (1986) Girard et al. (1980) Weiss and Schleicher (1968) House et a/. (1972) Munder et a / . (1971) Jensen (1981); Jensen et a / . (1974) Wohler et a/. (1972) Knazek et a / . (1972) Bailey and Ollis (1986) van Wezel (1967)

Note: The multiple propagator (stacked-plate propagator with airlift tube), the spiral film (coiled sheet with spacers to maintain sheet separation), plastic bags, and Gyrogen with tubes have limited use for large-scale applications with their relatively low S/V.

an intricate cell-cell and cell-substrate interdependence, whereas cells of the circulatory system such as lymphocytes have of necessity developed substrate-independent growth characteristics. Transformed cells, on the other hand, clearly display a different pattern of growth in uitro. They are not contact-inhibited and usually they grow past the confluent monolayer stage and, by growing in multilayer, eventually reach a saturation density several times higher than that of a single monolayer. Theoretically, transformed cells could grow indefinitely in the same culture, the limiting factors being unfavorable environmental conditions, either in the medium or in the gas phase. Other strains of transformed cells are in an intermediate state; i.e., they are able to grow either attached to a surface or in suspension. Baby hamster kidney (BHK) 21-C13 cells are an example of a cell line in which there are strains growing in monolayer or in suspension or in both conditions.

C. Suspension Cultures

The establishment of permanent cell lines from anchorage-dependent primary cell isolates often confers on these cells the ability to grow either in an anchorage-dependent mode or, after a suitable adaptation period, in suspension. The CHO cell line is well known for this dual capability, and for this and other reasons this cell line is a favorite host for the expression of heterologous genes introduced by recombinant-DNA techniques. There is much experimental evidence that suspension cell systems, which differ

VERTEBRATE CELL CULTURE TECHNOLOGY

151

with cell type and cultivation technique, also need a matrix that is macromolecular in nature and serves as a cell surface protector. Serum, for example, contributes more to the medium than a well-balanced mixture of nutrients; its macromolecules serve as important protectants in submerged systems. This macromolecular function is evident from experiments with media in which serum has been replaced. The addition of macromolecules such as Methocel (Keay, 1975; Radlett et al., 1971; Taylor et al., 1971) was essential for cell growth when submerged deep culture techniques were employed (Birch, 1980). Both the type of matrix and the technique of deep culture have to be adapted to the particular requirements of the cell line. In some cases it was shown that aggregates of anchoragedependent cells can substitute for a matrix (Tolbert er al., 1980). Normal human fibroblasts, considered to be entirely anchorage-dependent for proliferation, have been grown in methylcellulose medium without solid matrices (Peehl and Stanbridge, 1981). Certain similarities in growth behavior of microorganisms and anchorage-independent animal cells and the tremendous wealth of knowledge on microbial suspension cultures have benefited cultivation of anchorage-independent cells. Large-scale cultivations of vertebrate cells such as BHK cells in suspension culture are routine (Clarke and Spier, 1980).

0. Considerations in Selection of Growth Environment 1. Surface-Attached Growth versus Growth in Suspension

For the production of viral vaccines or interferon, cells are often propagated for a number of population doublings to increase cell number and are then subjected to infection or induction for product formation. Several other considerations have influenced researchers to set up monolayer systems despite the advantages of suspension cultures. These include (a) flexibility for cultivating either primary cells or diploid or heteroploid cell lines in monolayer systems, (b) easy removal of spent growth medium before infection of cells, and (c) high product (e.g., FMD virus) concentrations (Panina, 1985). The product concentration is related to the number of cells in the culture and to the volume of the medium used in the production phase. It is therefore advantageous to have in the culture system the highest number of cells possible and the flexibility of operating with as little medium as possible. The number of cells in a culture system is related, within limits, to the surface area available for growth, and therefore, operating systems with a large surface area in a relatively small volume of medium are of interest. Cell attachment on a surface, cell spreading, and cell growth are all dependent on the relationships between cells and their supporting surface.

152

SATISH J. PARULEKAR ET AL.

Considering the nature of an artificial substratum, both chemical and physical factors have been identified as having a controlling influence. The macromolecular organization, the charge structure, the wettability, and the physical form are considered to be of importance (Panina, 1985). Cells attached to a variety of surfaces spread in the direction of least curvature of the substratum (Curtis and Varde, 1964; Harrison, 1914; Ivanova and Margolis, 1973; Rosenberg, 1963; Rovensky ef al., 1971; Weiss, 1961, 1962). The extent of the substratum is also of importance. The circumference of glass bead substrata or the length of glass fiber substrata must be more than the normal length of the cells, in order to permit cell spreading and growth (Maroudas, 1972, 1973a,b). For an extracellular product, surface-attached growth may be advantageous because it avoids the need for a cell separation stage. In the extreme case, deliberate immobilization may be desirable, provided that adequate activity of cells can be maintained. If an intracellular product or the cells themselves are required, problems with detachment of cells from surfaces may be circumvented in suspension culture (Pollard and Khosrovi, 1978). Continuous operation offers the advantages of better equipment utilization and lower manpower requirements. This mode may be preferable to batch operation if culture stability can be demonstrated in the pilot plant. The decisive economic factors are directly related to the scale of operation, which is dictated by the projected market potential of the product; continuous processes are often chosen for large-scale production. With many bioreactors for surface-attached growth, e.g., roller bottles, the surfaceto-volume ratio decreases as the scale increases and this may make capital costs for large systems prohibitive. 2. Medium Design

Vertebrate cell culture technology has now given rise to a multi-million dollar industry (Ratafia, 1987). Common to all processes involving growth of cells is the need for culture medium. The cost of the culture medium forms a major part of the production process (Griffiths, 1986). Since the major innovative work of the late 1950s and early 1960s (Eagle, 1955, 1959; Levintow and Eagle, 1961) in which culture media were designed on the basis of cellular requirements, further advances have been made; the potential of systematic quantitative studies in optimizing media was demonstrated by several researchers (Birch and Pirt, 1970; Griffiths, 1970; Ham, 1974; Lambert and Pirt, 1975). However, most recent studies on culture media have concentrated on serum and its replacement by defined medium components. Detailed quantitative studies will become increasingly important as vertebrate cells become more widely used for production processes. Via

VERTEBRATE CELL CULTURE TECHNOLOGY

153

medium analysis and cell growth results, conditions for large-scale culture for production of FMD vaccine have been found to be suboptimal (Spier and Griffiths, 1985). Similarly, a comparison of the currently used techniques of hybridoma cultivation in uiuo (ascites) with those in uitro shows disappointing yields for the latter-being only 1% of those for the former. Such processes would benefit from detailed analysis to produce an improved medium design. An alternative approach to medium design lies in investigations into the composition of the cell itself. Analysis of the constituents of the cell can provide an indication of the components necessary for the production of cells. Herbert (1976) employed this approach to describe elementary analyses of the composition of yeast and bacteria in terms of their C, H , 0, and N content. The information provided by the elemental balances (Parulekar and Lim, 1985)provides a molecular formula for a given microorganism based on the proportion of gram-atoms of each element present in the cell. Such formulae may then be used to generate theoretical elemental balance equations for cell growth. This information is commonly used to give a theoretical indication of the components necessary in any culture medium designed to support the cell growth, as each of these components must be present in some form in the medium so that new cellular material can be synthesized.

111. Three-Dimensional Cultures A. Spider Web Cultures

Ross Harrison's innovative spider web culture system (1914) was probably the first successful attempt toward growing animal-cell cultures in three dimensions (3-D). The 3-D cultures are those in which cells are exposed to growth medium from all sides, a situation that most closely approximates the in uiuo organization of cells. In Harrison's technique of growing vertebrate cells, a single spider was introduced in a clean glass jar that contained a number of glass rings. The spider, within a period of 1-2 days, spun a web covering the jar bottom. The rings were then lifted out with forceps and the web was cut off around them. The rings containing web were sterilized by dry heat at 150°C. The rings were then fastened to a slide with Vaseline and a drop of growth medium was placed upon the web. The explants were seeded onto the drop of growth medium. The growth medium containing explants was mounted with a glass coverslip whose undersurface was coated with spider web. This enabled the tissue

154

SATISH J. PARULEKAR ET AL.

to stay between the two layers of the web. Two examples of cells in Harrison’s (1914) cultures are shown in Fig. 1. Using spider web cultures, Harrison was able to observe some of the features of locomotory behavior of cells. In particular, he demonstrated that cell shape was determined by the nature of the substratum on which they anchored and grew. Thus, cells on spider web fibers would be bi- or tripolar, whereas those on glass coverslips would be flat. He also noted that cell movement is directed by the type of substratum. Harrison (1914) stressed that the presence of a substratum was a prerequisite for the cell growth under either normal or chemotactic conditions. He related the arrangement of cells on web fibers to the sheath cells of an embryonic nerve. He attributed this relationship to stereotropism (now known as chemotaxis) and further postulated that these relationships were analogous to the close application of mesenchymal cells to blood vessels and muscles. As such, Harrison’s pioneering work opened the doors for further studies in the area of 3-D culture systems. However, his own spider web cultures could not be used further for the following reasons. Fibers from spider webs were not strong enough and thus, the actual number of cells that could grow on them was limited. Spiders, particularly the best suited species such as Tiginaria and Agalena, were seasonal. This restricted the use of this technology to certain period of each year. The amount of web fiber available per spider was limited. Efficacy of Harrison’s culture system was neither fully understood nor exploited. Indeed, over 60 years passed before the life scientists revisited Harrison’s idea. These later systems (Curtis and Seehar, 1978; Tripathi and Curtis, 1982; Tripathi, 1984, 1989a; Tripathi and Kerr, 1989; Tripathi and McGrath, 1989) are described in the following paragraphs.

6. Cultures on Sponge Leighton (195 1) developed a short-term cell culture method in which cells and cell aggregates growing into a sponge may produce a basic tissue pattern having similarity to the pattern observed in uiuo. In Leighton’s method, commercial cellulose sponge was cut into pieces measuring 8 x 5 x 1 mm and sterilized. The pieces of sponge were placed in 1.5 x 15-cm Pyrex culture tubes, about 2.0 cm from the bottom. One to four explants of tissue (size I x 1 x I mm to 1.5 x 1.3 x 2.0 mm) were placed on the exposed surface of the sponge. A single drop of chicken plasma and one drop of diluted chick-embryo extract (20-25%) were placed on the tissue explant and sponge. After the explants firmly adhered on to the sponge, six drops of growth medium were added (Figs. 2 and 3). The tubes were incubated at 37.5”C in a roller drum at 0.25 rpm. The

VERTEBRATE CELL CULTURE TECHNOLOGY

155

FIG. 1 (a) Cell from medullary cord with two processes attached to crossed web fibers ( x 300). (b) Bipolar and tripolar cells from medullary cord attached to spider web fibers 2 days after explantation ( x 300). (c) Cells similar to those in (b) 8 days after explantation ( x 300). (d) Two types of pigment cells, 6 days old ( x 300). Reprinted by permission of Wiley-Liss, a division of John Wiley and Sons, Inc., Journal of Experimenral Zoology, Vol. 17, p. 533, R. G . Harrison, copyright 0 1914.

156

SATISH J. PARULEKAR ET AL.

a

e of sponge

b

FIG.2 Sponge-tube cultures. (a) culture tube showing the luminal surface of the sponge from above, through the opposite side of the tube. (b) Culture tube showing the lateral surface of the sponge. (After Leighton, 1951.)

growth medium contained 50% chick-embryo extract, horse serum, and Earl's balanced salt solution in the ratios 1:2:2 and 1:2:4. Leighton's cultures were an attempt toward the development of cell culture techniques at a level where the cells could form distinctive tissue patterns. Thus, it might be possible to develop Leighton's system further to accommodate a variety of studies such as those involving cell accumulation, cell differentiation, cell modulation, tissue formation, and the relationship of structure and function. Leighton's approach might indeed provide valuable adjuncts to the methods and criteria used in pathologic diagnosis, and in the evaluation of the effects of therapeutic agents. Leighton's system, however, lacked certain basic necessities required of a culture system. For example, it was impossible to follow-up the progress of individual cells in the growing culture and a large number of cultures were required for carrying out structural studies due to the fact that it was necessary to sacrifice each culture prior to such studies.

C. Silica-Fiber Cultures Curtis and Varde (1964) grew fibroblasts on silica fibers (8-40 pm in diameter). In these cultures, silica fibers were arranged to bridge explants. The growth medium consisted of cock serum, saline, and 9-day chickembryo extract. The cultures were grown for 19 hr at 37.5"C as standing drops and thereafter as hanging drops. In these cultures, sheets of cells that slung between two or more fibers were formed (Fig. 4). The sheets were suspended in the medium and the cells were without any solid substratum except their lateral adhesion to neighbors and the adhesion of cells at the edge of the sheets to the fibers. In addition, sheets formed between a fiber and the explant where the fiber entered the explant, but these sheets did not extend far into the outgrowth (Fig. 4). Curtis and Varde (1964) used their silica-fiber cultures for studying the effect of topological factors

VERTEBRATE CELL CULTURE TECHNOLOGY

157

FIG. 3 (a) Mixed chick-embryo tissues in sponge. Multinucleated striated muscle cells interspersed with endothelial-lined channels ( X 400). (b) Mixed chick-embryo tissues in sponge showing anastomizing endothelial-lined channels probably derived from chick-embryo heart ( X 400). (After Leighton, 1951.)

158

SATISH J. PARULEKAR ET AL.

FIG. 4 Silica-fiber cultures. Fibroblasts, when cultured between silica fibers (diameter 30 p n ) , form sheets that require two bounding edges as supports. (After Curtis and Varde,

1964.)

VERTEBRATE CELL CULTURE TECHNOLOGY

159

on cell behavior. However, this culture system was not used subsequently due to the lack of good optics.

D. Cultures on Hydrated Collagen Lattices Elsdale and Bard (1972) used hydrated collagen lattices (HCLs) containing 0.1% (w/w) of collagen in the form of native bundles varying in diameter from 500 to 590 nm and possessing a 64-nm lateral period. These characteristics approximate to the concentration and form of collagen in extracellular connective tissue matrices. An example of this culture system is shown in Fig. 5 . From their work on HCL cultures, Elsdale and Bard (1972) postulated that most tissues are permanent residents of the tissues to which they belong by virtue of their attachment to the extracellular connective tissue matrix. They studied the behavior of simian virus-transformed rat (SVTR) fibroblasts in and on HCLs and suggested the possibility that cells may undergo changes that lead to their uncoupling from matrix. They

FIG. 5 Phase-contrast photomicrograph of human embryonic lung fibroblasts within an hydrated collagen lattice (HCL). Bipolar spindle form predominates. Pseudopodia are not expanded into the fanlike ruffling membranes characteristic of cells plated onto plastic at subconfluent densities. (After Elsdale and Bard, 1972.)

160

SATISH J. PARULEKAR E l AL.

further suggested that uncoupling might affect their resident status and may allow cells to be passively transported via the circulation. However, the study by Elsdale and Bard (1972), if pursued further, might help in understanding certain aspects of metastasis and possibility of a link between observed clustering and focal growth of SVTR cells on HCLs and the focal growth often observed at the advancing fronts of tumors. However, this culture system has become redundant for the following reasons. Only single cells can be grown and, therefore, this culture is not related to in uiuo situation. In these cultures, cells do not form definitive aggregates or patterns.

E. Sail-Sheet Cultures Curtis and Seehar (1978) used sail-sheet cultures (SSCs) for the study of the effect of mechanical stretching of cells on their cell cycle. In these 3D cultures, cells are attached to one another only at the edges like the sails of a yacht and they form sheets (Fig. 6). Sail-sheet cultures were later used to study the effect of mechanical stress on cellular morphology (Tripathi and Kerr, 1989);the locomotory behavior of fibroblasts (Tripathi and Curtis, 1982; Tripathi, 1984, 1989a); and the effect of tension on cytoskeleton (Tripathi, 1989b), cellular proliferation (Tripathi and Fygensen, 1992), and protooncogene expression (Tripathi and Vosseller, 1991).Tripathi and McGrath (1989) were able to form SSCs from dispersed cells, a technology that may be significant in developmental biology and healing of wounds. Recently, Marrow-Tech., Inc., has commercialized the SSC technology and has developed a dermal model kit (ZKI 100) that can be used to determine the extent of cytotoxicity induced by compounds/ formulations such as alcohols, antimicrobial preservatives, colognes, conditioners, detergents, metal chlorides, moisturizers, perfumes, pesticides, petrochemicals, shampoos, and solvents. F. Bergenholtz’s Culture System

The method of organ culture developed by Bergenholtz et al. (1977) and use of SEM generated significant information on the 3-D visualization of cellular morphology. Epithelial surfaces (1 x 3 mm) taken from the palatal mucosa of adult exsanguinated cats of both sexes were maintained in an organ culture system. The culture chambers were made of plexiglass with one inlet and one outlet for gas exchange. A sterile disposable plastic Petri dish (diameter 6 cm) was placed in each chamber. Each dish was equipped with a platform (2.5 x 2.5 cm) made from titanium expanded metal and

FIG. 6 Sail-sheet cultures. (a) A 7-day-old culture of chicken heart fibroblasts grown within the meshes of Nitex fabric under phase-contrast optics. Mesh size 200 X 200 pm? (Bar = 49 pm); (b) Scanning electron micrograph of chicken heart fibroblasts (7 days old) growing on a gold grid and beginning to fill the corners (Bar = 10 pm).

162

SATISH J. PARULEKAR ET AL.

FIG. 7 Schematic diagram of the platform illustrating the position and size of the explant. Reprinted by permission of Wiley-Liss, a division of John Wiley and Sons, Inc., Anafomical Record Vol. 189, pp. 433-442, A. Bergenholtz, G. Halimans, and L. Hanstrom, copyright 0 1977.

supported by 3-mm-long legs (Fig. 7). Each Petri dish was provided with 4.5 ml of growth medium. Four pieces of 27 X 5-mm Millipore filters (0.22pm-pore diameter) were placed on the top of each platform (grid). The cultures were maintained in 50% 02,45% NZ,and 5% CO, and incubated at 37°C. Bergenholtz’s system is suitable for studying the behavior of mesenchyma1 cells growing on a biological substratum, i.e., their own connective tissue matrix. Using this system, it is possible to study the proliferation and morphology of epithelial and connective tissue cells and epithelial-connective tissue interactions. This system may also be suitable for studying the effect of substances that might influence such intercellular interactions. A drawback of this system is its inability to allow visualization of and measurements on various aspects of behavior of living cells within their cultured environment. A wide variety of organ types cultured in v i m have served as excellent model systems for understanding the functional and structural relationships.

IV. Commercial Scale Bioreactors A. Considerations in Selection of Production System

There have been many approaches to the problem of providing an animalcell culture system that can not only be scaled to meet the needs of the research, but also be employed at the pilot plant and production plant scales (Spier and Griffiths, 1985). The most important point for the selection of a production system is the question of whether surface cultivation is necessary, or whether the cells can be grown in a suspension system

163

VERTEBRATE CELL CULTURE TECHNOLOGY

(Griffiths, 1988). The techniques for operation of dynamic depth cultures can be distinguished into three types (Fig. 8), viz., ( a ) surface (matrix) moved, liquid static (e.g., roller bottles); (b) surface (matrix) static, liquid moved (e.g., packed beds or columns, hollow-fiber reactors); and (c) matrix (and cells) moved, liquid moved (e.g., microcarrier culture). Several attempts have been made to develop new systems for large-scale operation. A major focus of the research effort was on increasing the surface area (for cell growth) per unit vessel volume and on implementing convenient and appropriate environmental control (Keary and Burton, 1979). Examples of these systems include vessels packed with glass beads (Wohler et al., 1972), stacked-plate columns or roller bottles (Schleicher, 1973; Weiss and Schleicher, 1968), rotating multiple tubes (Girard ef al., 1980), and roller bottles with spiral film inside (House et al., 1972; House, 1973). One limitation that is envisaged for large-scale operation of these systems is the supply of nutrients to the central regions of the bioreactors. Two of the most important advances in the field of vertebrate-cell culture technology have been the use of microcarriers, originally described by van Wezel (1967), and hollow fibers (Knazek ef al., 1972; Schonherr and van Gelder, 1988; Tyo ef al., 1988).

MATRIX TYPE

sdid single troys multipletrays discs plates tubes

DEEP CULTURE TECHNIOUE

BIOREACTOR

static

D

/

matrix

G

m

etC.

soft membranes (hdlowlfibers

polymer beads

submerged

..*:

0 .

microcarrier microcapsules

i

macromoleculor

submerged

FIG.8 Techniques for operation ofdeep cultures of animal cells. (After Katinger and Scheirer, 1985.)

164

SATISH J. PARULEKAR ET AL.

1. Unit and Multiple Process Systems Multiple process systems can be defined as those systems that can be scaled-up by increasing the number of units in operation in roughly the same proportion as the increase in the scale. Such a procedure has a number of disadvantages including higher capital and operating costs, less controllability, and poor operability (Spier, 1985). Although most of the products that are derived from cultured animal cells are high-value, small volume materials and the production cost can be readily taken up in the selling price, there is still much to be gained by operating with greater consistency, higher yields, and maximum product quality. In contrast to the multiple process systems, the scaleup of the unit process system is achieved by an increase in the size of the culture equipment without a substantial increase in the number of vessels. An important advantage inherent in the use of a unit process system is that it is practicable to monitor and control process parameters such as temperature, pH, redox potential, and levels of dissolved oxygen, glucose, and lactic acid dehydrogenase (Griffiths, 1988). 2. Bioreactor Operation Mode Animal cells can be produced using three types of cultures: batch, fedbatch, and continuous/perfusion. In a batch system, no nutrients except oxygen are replenished. The only parameters that can be controlled are temperature, aeration, and pH. Thus, the cells in a batch culture are subject to constantly changing environment, where nutrients are depleted while desired products and waste products accumulate. Cell growth and/ or product formation can be prematurely inhibited due to nutrient limitation or toxic buildup of waste products. The products of animal-cell metabolism include lactic acid, ammonia, and carbon dioxide. If lactic acid and ammonia are left to accumulate to high enough concentrations, cell metabolism and even cell growth can be inhibited. This problem can be partially alleviated by replacing the “spent” medium with “fresh” medium, the so-called repeated-batch culture. The periodic replacement of a constant fraction of the culture by fresh medium, which is the most common means of subcultivating animal cells and often termed solera culture (Pirt and Callow, 1964), is a succession of batch cultures in which part of the old culture is used as inoculum. Solera culture has been used to produce Namalwa cells on a semi-industrial scale for the production of human interferon (Reuveny ef ul., 1980). The cells are still subjected to a constantly changing environment in a repeated-batch culture unless the medium is replaced frequently, which can be very expensive if the medium consists of relatively expensive components such as serum. In addition, when the medium is replaced, some components that are not exhausted

VERTEBRATE CELL CULTURE TECHNOLOGY

165

at the time of the medium change are discarded needlessly. Conditions that are optimal for the maximal cell multiplication may not be optimal for the maximal formation of a particular cell product. Furthermore, conditions optimal for product formation may be attained only transiently or perhaps not at all in conventional batch culture. In addition, a cell product may be inhibited by further production of that product or by production of another cell product or metabolite. A cell product may also inhibit further cell multiplication. A better mode of operation would be a fed-batch system that feeds vital components (only as needed) to the culture with a corresponding increase in culture volume, enabling maintenance of a constant nutrient concentration. The fed-batch process requires that methods exist to monitor and control the nutrients in question. More environmental parameters are controllable in this system and thus one can better optimize cell growth and product generation. Although cellular waste products are still allowed to accumulate in this system, it may be possible to limit the accumulation of these waste products by adjusting appropriately the nutrient flow rate. Significant reductions in generation of lactic acid and ammonia in mammalian cell cultures have been reported by controlled feeding of glucose and glutamine, respectively, to the cultures (Glacken et al., 1983b).If a portion of a fed-batch culture is withdrawn at intervals, the culture can be maintained more or less continuously. Fed-batch culture is particularly attractive for production of those biologicals, the synthesis of which is maximal after cells have reached a stationary phase (Parulekar and Lim, 1985). The chemostat culture (steady-state continuous culture) enables large quantities of animal cells to be produced under precise physiological conditions. Cells can be cultivated in a chemostat in a number of unique environments, under a variety of growth-limiting conditions, and at a wide range of cell growth rates. Other advantages afforded by chemostat culture include the ability to control culture conditions and to automate the process, reduction in size of the culture vessel required to produce a given quantity of cells, reduction in downtime (the time required for sterilization, inoculation, harvesting, etc.), and production of cells in the precise physiological state required (Tovey, 1985). A whole range of different environments can be established and maintained indefinitely in a chemostat. Cell products are continuously removed from the bioreactor, preventing inhibition of product formation or inhibition of cell multiplication. The continuous culture more closely approaches the situation in uiuo. In the living animal, an efficient circulatory system delivers nutrients to cells and removes their waste products and thus provides the cells with a stable environment. The industry standards for large-scale continuous animal-cell cultivations remain the stirred tank reactor (Acton and Lynn, 1977; Pullen et af., 1985; Radlett et af., 1985; Scattegood et al., 1983) and the airlift reactor

166

SATISH J. PARULEKAR ET AL.

(Birch et al., 1987; Margaritis and Wallace, 1984). These reactors provide a homogeneous environment and are amenable to scaleup and process control, and there is a large body of engineering knowledge about these reactors from traditional chemical processing and microbial fermentation literature. With the advent of microcarriers (Levine et al., 1979; van Wezel, 1967) and surface dissociation techniques (Hu et al., 1985), anchorage-dependent cells may be cultured in homogeneous fermentors, and reactor-to-reactor propagation of cultures is simple, so that virtually any cell type may be employed in such a system. 3. Shear Sensitivity The extreme fragility of most animal cells to mechanical and hydraulic shear forces is a particular problem when using cultivation vessels on a large scale, because of the exponential increase in peak energy within turbulent systems (Katinger and Scheirer, 1985). In some cases, the use of low-shear mixing systems like airlift or draft-tube reactors might solve the problem. The other way of protecting cells from shear force is shielding. This is done with systems in which some kind of “encapsulation’’ is used. The capsules may be made of semipermeable membranes like polylysine, or of porous, spongy materials (Scheirer, 1988). Cell lines such as HeLa cells and lymphoblastoids, which do not require a surface on which to grow, can be grown in equipment similar to that employed for bacterial fermentations. However, since mammalian cells do not have a protective cell wall, they are much more sensitive to shear than microbial cells. Thus, whereas increased surface-to-volume ratio is a goal that catalyzed the development of new techniques for cultivating anchorage-dependent cells, shear reduction is the goal that catalyzed the development of novel techniques for cultivating suspension cells (Glacken et al., 1983b). Many attempts have been made to use surface-dependent cells within suspension systems. The best known of these is the microcarrier approach (Butler, 1988; van Wezel, 1985), but other possibilities, such as fiber culture (Peehl and Stanbridge, 1981) and aggregate culture (Tolbert et al., 1980), have also been used successfully. However, if there is a preference for a classical surface-type cultivation, it should be noted that there are some fermentor types that have been reported as successful units (Panina, 1985; Spier, 1985). These include some of the filled-column-type (Merk, 1982; Whiteside et al., 1979) and others of the membrane type, such as those employing hollow fibers or flat membranes (Klement et al., 1987; McAleer et al., 1987).The cells are protected from hydraulic and mechanical shear within the membrane devices and the membranes often enable retention of cells and the desired product(s) and exclude distinct substances, e.g., immunoglobulins, from the medium. Oxygenation of the

VERTEBRATE CELL CULTURE TECHNOLOGY

167

cells through membranes may also be possible (Lehmann et al., 1988; Tolbert et al., 1985). These devices duplicate in principle the environment of cells within organs and may be used for suspension-type cells as well as for surface-dependent ones. In cultivation systems where cells are retained by filters (or molecular sieves) and the culture is continuously perfused with fresh nutrients and waste products are removed, the effect on culture development is further improved and cells at very high densities can be kept viable (Feder and Tolbert, 1983; Tolbert et al., 1981). B. Perfusion-Based Reactors

The various forms of perfusion culture, in which suspended cells are retained by filters or immobilized on beads, provide an in uiuo-like environment. They are suitable for cultivation of all types of cells and very good growth results have been reported with various types of cells at various scales of operation (Feder and Tolbert, 1983; Nahapetian et al., 1986; Whiteside and Spier, 1981; Whiteside et al., 1979). Perfusion systems are particularly advantageous for nonpropagating (stationary phase) cells, when cells of limited life span are used and when the kinetics of product formation are typically non-growth-related, such as with urokinase and monoclonal antibodies. Additionally, the combination of perfusion and external recycling of media offers the application of “quasi-homogeneous” cultivation as well as use of packed-bed and fluidized-bed reactors. Both nutrient and waste product concentrations can be controlled by varying the dilution rate for the perfusion bioreactor. Increased dilution rates result in increased nutrient and decreased waste-product concentrations. Thus, a high degree of control can be exhibited over the bioreactor environment. However, the loss of nutrients in the bioreactor effluent makes the operation less cost effective. In order for perfusion systems to be economical, these must support higher cell densities than those achieved using other techniques, or must employ dialysis or filtration to recover expensive macromolecular serum components and to permit exchange of spent medium with fresh medium outside the culture (Graff and McCarthy, 1957; Himmelfarb et al., 1969; Kruse et al., 1963; Thayer, 1973; Tovey, 1985). Thus, the choice between a fed-batch system and a perfusion system depends on factors such as (a) the optimal medium turnover rate of the perfusion system that balances high final cell densities with low medium usage; (b) the extent of reduction in waste-product accumulation that can be achieved in a controlled, fed-batch system; (c) the number of medium changes required to keep the concentration of waste products below the maximum permissible level in the fed-batch system, if the generation of waste products cannot be sufficiently reduced

168

SATISH J. PARULEKAR ET AL.

through adjustments in environmental parameters alone; (d) the cost of medium dialysis in the perfusion system (Schonherr and van Gelder, 1988); and (e) the cost of equipment peculiar to perfusion systems. The excretion rate of the desired extracellular product is usually related to the growth rate of the culture. In the case of a growth-dependent type of secretion, there is no way of using perfusion-operated cell encapsulation systems because there is almost no growth after confluence within the capsules. In such cases, any system with logarithmic growth, like a continuous culture or a batch culture, must be used. If there is a good production rate during the stationary culture phase, one should avoid logarithmic growth systems and prefer either retention systems or long-lasting batch cultivations. For an optimized system, it may be necessary to use a combination of continuous and perfusion systems to enable flexible choice of growth rates between maximum and almost zero. The spin filter culture described by Himmelfarb et al. (1969), Tolbert et al. (1981, 1988),and Tolbert and Feder (1983)is a special perfusion reactor (similar in principle to the rotofermentor described by Margaritis and Wilke, 1978), where cells are kept in the reactor and only the medium is removed from the suspension by filtration through a rotating filter cartridge placed inside or outside the bioreactor (Fig. 9). The rotation minimizes clogging of the filter cartridge by cell mass, which occurs during separation of animal cells from liquids by filters. The filtration unit separates the cells from the product-containing cell-conditioned medium and returns the cells to the growth vessel as fresh medium is added. The rotating filter cartridge

reservoir 4 4 %

I

I

Gmwlh vessel

37%

FIG. 9 Perfusion culture system for suspension cultures. (After Tolbert et a / . , 1988.)

VERTEBRATE CELL CULTURE TECHNOLOGY

169

generates a laminar liquid layer near the filter surface. The centrifugal force generated by the rotation forces the cells to move away from the filter and keeps the fluid on the filter surface free from cells. Besides high cell densities, it should also be possible to get better product generation. Cells are not wasted with the effluent but stay in the production process. Spin filter cultures (with growth vessels of 100-liter capacity) can operate for prolonged periods (typically 90 days) with cell densities 10 to 30 times those attainable in a conventional spinner vessel. The medium consumption and serum requirement on a per cell basis are about one-third of those in conventional reactors, thereby simplifying the subsequent purification required to obtain the desired product (Katinger and Scheirer, 1985). C. Two-Stage Reactors

Whenever a production system requires continuous renewal of biomass and the product formation requires a separate production step (e.g., virus production, interferon production), the application of two-step or multistep continuous culture systems should be considered, since it combines the optimum biomass production in the first reactor and the possibility of optimizing and conditioning the production phase in the following reactor(s) (by addition of inducers, viruses, etc.). The heterogeneous continuous culture will probably become the system of choice as inducible promoters are cloned in animal-recipient cell strains by recombinant-DNA techniques (Collins, 1982; Hofschneider, 1982). The production of animal-cell products in chemostat culture has to date been largely confined to studies on the production of viruses and interferon (Gori, 1965; Kilburn and van Wezel, 1970; Mogensen et al., 1972; Tovey er al., 1973; van Hernert et al., 1969). A persistent virus that establishes a persistent infection without causing a cytopathic effect and without drastically reducing cell growth rate can be propagated in cells cultivated in a single-stage chemostat. However, a virus that causes a lytic infection would have to be produced in the second stage (where conditions are optimal for maximal virus formation) of a two-stage chemostat, the host cells being produced in the first stage where conditions are optimal for the maximal output of cells. Tovey et al. (1973) showed that repeated induction of interferon could be obtained in chemostat cultures of mouse LS cells without the development of a refractory state (cessation of interferon production).

0. Airlift Reactors In contrast to the conventional stirred tank fermentor systems, airlift fermentor systems have no moving parts, and the only power requirement

170

SATISH J. PARULEKAR ET AL.

comes from the air compressors, which deliver the air through a sparging system. There is no mechanical agitation and the air bubbles forced through the sparger system are responsible for the induced liquid mixing and accompanying mass transfer. The presence of a draft tube inside the airlift fermentor stabilizes the liquid circulation patterns and increases the interfacial area between the gas and liquid phases. The main advantages of airlift fermentors are low shear rates, low energy requirement, and relative simplicity of construction, as aseptic seals are not required around the rotating shafts in these reactors. Airlift bioreactors have been successfully scaled-up to commercial size units (production capacities of 60,000 tondyear) for production of single cell protein by ICI, England (Hines, 1978; Zanetti, 1984), and in a Dainippon process (Zanetti, 1984) in Curtea de Arges, Romania. When air sparging is acceptable, an airlift reactor can be used as an alternative to a mechanically stirred reactor (Katinger el al., 1979; Katinger and Scheirer, 1982). Although mixing efficiency is somewhat lower than that in a stirred tank reactor and there is increased foam formation (both of which problems can be overcome by extending the draft tube to just below the surface and just above the bottom of the bioreactor), there is practically no technical limit to the scaling-up of airlift reactors (Katinger and Scheirer, 1985). A bioreactor based on a bubble column and with extremely low power input and shear forces has been developed by Katinger e? af. (1979). Katinger and Scheirer (1985) tested these bioreactors with net volumes up to 700 liters and found good results with some lymphoblastoid cell lines and BHK21. This airlift-type bioreactor needs, unlike the more conventional types, less energy per volume in larger vessels than smaller ones. This is the reason that larger reactors work more effectively than smaller ones. Scaledown is restricted to sizes larger than 5 liters. The maximum local power dissipation density within the airlift reactor is similar, irrespective of the reactor size, whereas in conventional reactors the local power dissipation density increases with reactor size (Katinger and Scheirer, 1985). E. Cell Propagators

Flat or surface-developed plates have been used in a number of configurations in cell propagators. The materials used for the plates have varied. Many of the systems described have been based on glass plates (Litwin, 1976; Mann, 1972; Parisius et af., 1974; Weiss and Schleicher, 1968), whereas others have been predominantly stainless steel (Spier, 1985). Other materials commonly used include treated polystyrene (Pakos and Johansson, 1983; Skoda and Pakos, 19771, titanium (Molin and Heden, 1969), polycarbonate, polyethylene terephthalate, and methyl pentene

VERTEBRATE CELL CULTURE TECHNOLOGY

171

(Izuka, 1979). The surface area of the plates has been augmented by specially shaped corrugations or by using fused metal spheres of titanium or aluminium (McAleer et al., 1978). Although the techniques of handling banks of static multitray units have been developed to a considerable extent (Pakos and Johansson, 1983), the cumbersome manipulations and the complexity of the automated equipment that handles the units militate against the facile spread of this technology. To overcome some of the disadvantages of the static-plate propagators, multiplate propagators were developed (Fig. 8). The primary advance evidenced by the dynamic systems was the generation of a more thoroughly mixed system. Concentration gradients of metabolites, excretory products, or hydrogen ions were virtually eliminated. In addition, the vertical plate systems offer a higher surface area-to-medium volume ratio, as only half of the plates need be submerged at any time. Inevitably, the mechanical complexity of the system is increased by the addition of motors, bearings, supports, and shafts. It is logical progression from the plate systems to consider the infinitely extended unit plate convoluted in a way that enables it to fit into a conveniently sized chamber or vessel. A unit based on a spiral of treated polystyrene film or sheet has been widely investigated (House et al., 1972; Nicklin and House, 1976; Taylor and Evans, 1975) and is available as a presterilized packaged apparatus. Other spiral systems have also been developed and used, at least on the small scale (less than 2 liters), with some success; melinex polyester (ICI, UK) has been used by Spier (1985), and the application of a polycarbonate spiral system has also been reported (Fontages et al., 1974). One approach to improving such spiral systems requires the control of medium flow and gas exchange throughout the bulk of the apparatus. This can be achieved by growing the cells in plastic (Teflon) bags that are permeable to gases like O,, N,, and COz, but impermeable to fluids. The system consists of a long Teflon tube (typically 5.7 cm wide and about 900 cm long) that is wound in a spiral with a layer of corrugated aluminium between successive turns. Tubes are attached to each end of this long bag for the movement of fluids and the assembly is mounted on a reel that is automatically rotated 360" every hour, reversing direction after each revolution; this enables the top and bottom inner surfaces of the bag to be used for cell growth. The cells can be detached from the substratum by mechanically stretching the unreeled bag. Permeability to air permits bag reuse and facilitates bag sterilization. F. Microcarrier-Based Reactors

Most animal cells require for their growth in uitro a suitable substratum to which they can attach and spread. This phenomenon has delayed the

172

SATISH J. PARULEKAR ET AL.

application of homogeneous culture techniques for animal cell cultivation, as generally used in standard microbiological cultivation processes. It has led to the development of various types of more or less nonhomogeneous stationary culture systems for large-scale production. The disadvantages of these systems regarding control and scaling-up of the cultivation process are obvious and have been extensively described in the literature (Levine et af.,1979; Reuveney, 1983; Spier, 1982; van Hemert et af., 1969). The homogeneous cultivation of anchorage-dependent cells became possible through the development of the microcarrier culture system (van Wezel, 1967, 1972, 1973). Through the development of better microcarriers as substrates, it has grown to be the most preferred system for the large-scale cultivation of animal cells for production of cellular components and viral vaccines (Butler and Spier, 1984; Butler et af.,1986; Glacken et af., 1983b; Hirtenstein and Clark, 1983; Jo et af.,1991; Reuveney, 1983; Spier, 1980, 1982; van Wezel, 1972, 1973, 1985). The concept of microcarrier culture is rather simple. It comprises the cultivation of cells on small solid particles, the so-called microcarriers, suspended in the growth medium by stirring. Cells attach to and spread on the microcarriers and gradually grow out to a confluent monolayer (Levine et af.,1977a,b; van Wezel, 1967). By using small particles of about 200 pm at a concentration of roughly lo4 particles/ml of culture, a quasihomogeneous system that resembles the traditional microbial submerged culture with all of its advantages is achieved. In microcarrier culture, the features of both suspension culture and monolayer culture are brought together in one system. A high transparency of the bead material is required to allow microscopic observation of cells. In some cases, the transparency might be improved by transferring the beads, after staining, to a liquid with high refractive index (such as glycerol). Microcarrier cultures may be started by inoculating well-agitated bioreactors (containing microcarriers) with cells of interest. For better cell attachment and cell growth, it is essential that the agitation be as slow as possible and just enough to keep the microcarrier beads in suspension and the inoculum size be kept low (Pharmacia, 198 1 1. One of the major advantages of the homogeneous microcarrier culture system is that at any time during the cultivation process, samples may be withdrawn for monitoring cell growth, for microscopic control of cells on the carriers, or for a more extensive control (in the absence of extraneous agents) after subcultivation in standard monolayer cultures. Although monitoring of cell growth may be performed by detaching the cells from the microcarriers with a standard trypsinization procedure and subsequent cell counting, more reliable results are obtained by counting released nuclei (Sanford et af., 1951). Cells on the microcarriers may be directly analyzed microscopically or after staining with hematoxylin (van Wezel,

VERTEBRATE CELL CULTURE TECHNOLOGY

173

1985). Simple microscopic techniques enable process operators to monitor the performance of the system by assessing the “state” of the cellsattached, rounded, flattened out, granular, or exhibiting cytopathology, syncytia, overgrowth, confluency, or nodule formation. All the cells throughout the system experience an identical environment in terms of the concentrations of metabolites such as oxygen, lactic acid, ammonia, and hydrogen ions. Once the most favorable conditions for growth and productivity have been defined, all the cells in the system can be made to function at the limits of their capability. The scaleup of microcarrier-based reactors also presents fewer problems because there is no need to mimic the inevitable gradients that are inherent in the inhomogeneous systems such as the packed-bed reactors (Spier, 1985). A suspension of microcarriers has the highest surface-to-volume ratio of any technique developed to date and therefore is particularly attractive for production of anchoragedependent cells on a large scale. Thus, all other factors being equal, less labor, fewer materials, and less medium are required to produce a given quantity of cells in microcarrier cultures compared to other systems. Adjustments to the culture environment can be easily made, to continually maximize cell growth and product formation, while at the same time keeping waste-product formation to a minimum. In addition, since the environmental parameters can be controlled, greater consistency from culture to culture can be expected. Microcarrier suspension cultures have a few unattractive characteristics. First, not every cell line can be easily removed from the microcarriers while still maintaining high cell viability. Since smaller microcarrier cultures cannot be used to inoculate larger microcarrier cultures, a largescale production facility utilizing microcarriers still must depend on cells produced from roller bottles for inocula. Many investigators have experienced considerable difficulties when using particular cell lines at the larger scales of operation (greater than 10 liters). Although in some systems, such as the BHK monolayer cell system, the cells attach to the beads when the latter are in motion (Butler and Spier, 1984; Butler et al., 1986; Spier, 1976), other cells require that in the initial stages of the culture the microcarriers be stationary (Pharmacia, 198 1). As this initial attachment and “flattening out” phase is critical to the subsequent development of the cells, the conditions for optimal performance are often sharply defined. Small changes in the cell seed, serum composition, or hydrodynamic environment can therefore lead to relatively large changes in system performance. Additional reasons favoring the use of alternative systems are (a) the cost of the substratum is considerable; (b) the substratum afforded by most microcarrier systems is different from that to which the cells have been accustomed; (c) the microcarrier systems have proved to be difficult to aerate without generating a foam in which the microcarriers collect; and

174

SATISH J. PARULEKAR ET AL.

(d) the removal of spent fluids or chemicals that control gene expression is difficult, as the void volume of the sedimented microcarriers is significant and in some cases the bioreactors containing microcarriers may have to be diluted out several times to remove traces of unwanted materials (Spier, 1985). The growth of anchorage-dependent cells on microcarrier beads depends directly on the surface available for growth. This is true up to the point where the microcarrier particles reach sufficient concentration to poison the cells and thus reduce cell yield. The toxicity of the support causes long lag phases, death of the cells in the early stages of development, and limited cell yields. Further development of new microcarrier supports will improve the applicability and reduce the toxic effects (Crespi et al., 1981; Feder and Tolbert, 1983; Fohring et al., 1980; Giard et al., 1979; Levine et al., 1979; Pharmacia, 1981; Santero, 1972; van Wezel, 1967; Weiss and Schleicher, 1968). Since anchorage-dependent cells can detach from a microcarrier (Thilly and Levine, 1979) and reattach to new beads, it should thus be possible to supply new beads to the culture and obtain some form of continuous flow culture. Such systems are projected to prove useful for the production of interferon+ or vaccine strains of certain viruses from human diploid fibroblasts. The cell yield in a microcarrier culture may be increased considerably through replenishment of culture medium. The simplest way to replenish the medium is the batchwise procedure. This is achieved by stopping the stirrer, allowing the microcarrier beads to settle, withdrawing the supernatant culture fluid, and adding fresh medium. A disadvantage of this procedure is that the cells are suddenly exposed to totally nonadapted medium. Even if the medium is conditioned at the same temperature and pH, it will always introduce a short lag phase. Also, detachment of cells from the microcarriers is observed during this procedure. These problems may be overcome by partial replenishment. Good results are obtained if only up to 75% of the medium is replaced. More ideal systems for medium replenishment are the continuous flow systems with either recycling or perfusion of medium (Fig. 10) (Kluft ef al., 1983; Tolbert et al., 1988). With these systems the replenishment should be gradually increased in relation to the cell concentration in the bioreactor (growth vessel in Fig. 10). In this way, sudden changes in composition of the culture fluid and culture conditions are avoided. Retention of cell-laden microcarriers in the bioreactor may be achieved by a sieve mounted on the outlet tube or with the help of a settler (Fig. 10) so that the microcarriers and culture fluid are separated by gravity. The first system has the disadvantage that the sieve may be blocked during the process, whereas in the second system microcarriers and culture fluid may not be totally separated at high flow rates. A more reliable system is the application of a cylindrical sieve

175

VERTEBRATE CELL CULTURE TECHNOLOGY

FIG.10 Perfusion culture systemforcellsattached tomicrocarriers. (AfterTolbert e t a / . , 1988.)

mounted on the stirrer shaft (Fig. 11). Even at high recycling or perfusion rates, microcarriers and cells will be dislodged from the filter by shear and centrifugal forces (van Wezel, 1985). In addition to the supply of nutrients, the recycling and perfusion system may also assist in control of pH and PO, (dissolved oxygen level) and the perfusion system may aid in removal of metabolic inhibitors of cell growth. The perfusion system appears to be particularly useful for large-scale production of biologicals excreted by cells, such as the production of tissue plasminogen activator (tPA) by

-El-

@:... :.:c::.+..!? Z Y ..... ..?: .2:.:ii.i.y

Filter

:..._.. . . , :...:.. .... ._ .... ..:. .: ......

...

Air

Medium

+

C02

Culture

Culture fluid

FIG. 11 Continuous perfusion system for replenishment of medium or continuous production of cellular components. (After van Wezel, 1985.)

176

SATISH J. PARULEKAR ET AL.

Bowes melanoma cells (Kluft et al., 1983). At the 40-liter scale and a perfusion rate of one culture volume per day, 750 to 1000 liters of TPAcontaining culture fluid could be generated within 25 days. Particularly now that genes of various viral and cellular components are being cloned in mammalian cells by recombinant DNA techniques, the application of microcarrier-based reactors in large-scale production of cellular components may increase significantly. Microcarrier culture has been employed for the cultivation of recombinant Chinese hamster ovary cells for consistent production of human immune interferon for prolonged periods (up to 36 days) (Hu and Dodge, 1985). However, thus far, the large-scale (1000 liters or more) application of microcarrier culture is mainly limited to the production of viral vaccines such as those for polio and rabies (Montagnon et al., 1981, 1983; van Wezel et al., 1978a), and FMD virus (Meignier et al., 1981). Based on the initial work by van Wezel, improved DEAE-Sephadex beads with a controlled lower charge density were developed (Levine et al., 1979; Hu and Wong, 1985). Using microcarriers with a controlled charge density (in the vicinity of 2.0 meq of DEAE/g of dry Sephadex dextran), a higher microcarrier concentration could be used and still support normal cell growth. A microcarrier concentration of 15 g/l was used by Hu et al. (1985) for cultivation of human fibroblasts. The surface area provided by microcarriers is, of course, dependent on the size of microcarriers used (Hu and Wang, 1987). In the study mentioned above using 15 g/l of microcarriers, a surface area of approximately 68,000 cm2/1was available for cell growth and a maximum cell concentration of 3.5 x lo6 cells/ml was achieved as opposed to 2 x lo5 cells/ ml, which is typically obtained in roller bottles. Besides microcarriers with controlled charge density, other types of microcarriers have been developed including dextran-based beads coated with gelatin, polyacrylamide beads, polystyrene beads, hollow glass beads, cylindrical cellulose beads, and fluorocarbon droplets stabilized with polylysine. At present, dextran-based microcarriers are probably the most widely used. One example of the application of this type of microcarrier is the production of chick-embryo fibroblasts in a 150-liter agitated vessel (Tolbert et al., 1981). DEAE-polyacrylamide beads have also been used in large scale cultivation of cells (Tobert and Feder, 1983). Cylindrical cellulose derivatized with DEAE-charged groups has also been reported to support the growth of chick-embryo fibroblasts in the form of cell-microcarrier aggregates, but does not support the growth of normal diploid cells. Further studies are necessary to test the wide variety of microcarrier surfaces for their ability to support the growth of the wide range of cell types currently being investigated for potential large-scale applications.

VERTEBRATE CELL CULTURE TECHNOLOGY

177

G. Microencapsulation-Based Reactors

In a microencapsulation system, cells are encased within spheres composed of alginate, agarose, or lysine (Lim and Moss, 1981a,b; Nilsson et al., 1983). Cells grow to a high concentration inside the capsule and are used for product formation. The microcontainers or microcapsules minimize the mechanical stress experienced by the cell surface. The polymer resins in which the cells are encapsulated have diameters of 50-500 pm, allowing the embedded cells to build up their own microenvironment, which is not destroyed by convection in the dynamic environment within the bioreactor (Katinger and Scheirer, 1985). The encapsulated cells are cultivated in standard suspension culture vessels. The advantages of this system are that (a) higher cell densities are obtained compared to standard suspension systems; (b) increased product generation per cell can be obtained; (c) cells can be separated from the medium simply by means of gravity settling, as opposed to centrifugation; and (d) product can be partitioned into either the microcapsule or the medium, separate from the cells, whichever is desired. A disadvantage of this system is, of course, the cost of encapsulation and removal of cells and products from the capsule (Katinger and Scheirer, 1985). A more serious limitation to the use of this technique on a large scale is that of transfer of oxygen and nutrients. The cell mass in the microcapsule may limit transport of nutrients and oxygen to cells in the center of the capsule. This can have adverse effects on cell metabolism and product formation, as well as complicating the implementation of any process control scheme. The doubling time for cell growth in microcapsules can be considerably larger than that in a normal suspension culture (Hu and Dodge, 1985). Hybridoma cells seem to be the most promising candidates for mass production in the near future. Hybridomas in many cases do not grow and do not produce product at expected levels in submerged deep culture. Microencapsulation has been used for the production of monoclonal antibodies (Hu and Dodge, 1985; Katinger and Scheirer, 1985; Lim and Moss, 1981a,b; Nilsson et al., 1983). A stainless-steel screen mesh can be employed to retain the beads while allowing draining or filling of fluids. Cell numbers per unit volume in microcapsules can be 50 times greater than those attainable in a continuous suspension culture. The ratio of microcapsule volume to culture volume is normally between 0.2 and 0.5 (Hu and Dodge, 1985). In a study on production of high-molecular-weight globulins by hybridomas encapsulated in polylysine beads, antibody concentrations comparable to those of mouse ascites cultures have been reported after freeing the content of the capsules, resulting in globulin titers 100 times higher than those attainable in conventional in uitro cultures (Lim and

178

SATISH J. PARULEKAR

ET AL.

Moss, 1981a,b). This excellent production was obviously a result of both improved microenvironment and improved nutrient supply achieved by perfusion. The techniques of microencapsulation of living cells, which are currently further developed and applied on a broad basis to plant cells and microorganisms, will stimulate research and promote their application to cultivation of the more fragile and sensitive animal cells. H. Fluidized-Bed Reactors

In the last 15 years or so, there has been a significant increase in the number of applications of fluidization in biotechnology. Some of the advantages of the fluidized-bed bioreactor systems are superior mass and heat transfer characteristics, very good mixing between the multiple phases, and relatively low energy requirements. Additionally, the low shear rates attained make the fluidized-bed bioreactor systems suitable for shear-sensitive cells such as mammalian cells. In most cases reported in the literature, fluidizedbed bioreactors have been employed in conjunction with immobilized cells. One such study (Runstadler and Cernek, 1988)deals with large-scale economic production of medical proteins from hybridomas and genetically engineered mammalian cells. Attachment-independent, attachment-dependent, or attachment-preferred animal cells can be cultured in a fluidized-bed bioreactor by first immobilizing them in porous microspheres. The microspheres are made of materials such as collagen and weighted with a noncytotoxic material to achieve a high specific gravity (typically 1.6 and higher), so that they will remain suspended in the high velocity (of the order of 70 cm min-'), upward-flowing, fluidizing culture liquor. The sponge-like microsphere (with a typical diameter of 500 pm) contains interconnected pores and channels of the order of 20-40 pm wide, allowing cells to enter easily and populate the largely empty interior of the sphere. Several thousand cells will typically populate a single microsphere and the size of the microsphere relative to the cell size is what enables the cells to have the high oxygen and nutrient transfer rates required by the cells. The cubic relationship between the diameter and the volume of the microsphere means that seven-eighths of the cell population inhabits the outer half of each sphere. The high porosity (85%), coupled with the biocompatibility and biological attractiveness to the cells of the collagen, enables cells to grow to high concentrations. Typical cell densities per milliliter of microsphere volume for hybridomas and animal cells are of the order of lo8 to lo9 cells/ml (Runstadler and Cernek, 1988). The porous space in the microspheres permits inoculation with cells after the microspheres are placed in the bioreactor. This means that the microspheres do not have to be manufac-

VERTEBRATE CELL CULTURE TECHNOLOGY

179

tured with cells in them and can be stored for long periods of time prior to their use in the bioreactor. Fluidized-bed technology is used principally to achieve uniformity throughout the bioreactor vessel and to provide mass transfer of oxygen and carbon dioxide at rates sufficient to support the high cell densities that are achieved in the microspheres. The microspheres, which contain over 95% of the cells in the bioreactor vessel, never enter the recycle loop, but are retained in the suspended slurry in the bioreactor (Fig. 12) (Runstadler and Cernek, 1988). The membrane gas exchanger in the recycle loop is a tube-and-shell design using permeable silicone rubber membranes to separate the culture liquor and the gas side of the exchanger. Oxygen is transferred by permeation through the silicone membrane from the gas (shell) side of the exchanger to the culture liquor which flows in tubes. Carbon dioxide permeates in the opposite direction from the culture liquor to the shell side (Fig. 12). The gas exchanger must be sized to supply oxygen in amounts sufficient to meet the oxygen requirements of cells at the high densities maintained in the bioreactor. Typical oxygen supply rates per liter of reactor volume approach 10 mmol hr-I. The all-liquid system (oxygen and carbon dioxide being only present in the culture liquor in the dissolved state) eliminates all gas-liquid interfaces and the resultant foaming found in many bioreactor systems that causes considerable damage to cells and some proteins. In addition, the fluid dynamics of the bioreactor and recycle loop flow must minimize shear forces so that cells and sensitive proteins manufactured by the cells are not damaged. For the same cell line with the same cell-specific productivity, the fluidized-bed/microsphere culture produces cell products at a rate

FIG. 12 Fluidized-bed bioreactor system with recycle flow. (After Runstadler and Cernek, 1988.)

180

SATISH J. PARULEKAR ET AL.

15-60 times greater than that produced by conventional chemostat or microcarrier/stirred-batch modes. This means, simply, that 1-liter fluidized-bed/microsphere continuous culture bioreactor has the same production capability for attachment-dependent cells as a 60-liter microcarrier/ stirred-batch system using the same cell line. Unlike other continuous culture techniques, however, the fluidized bedimicrosphere system can be scaled-up indefinitely. Microsphereifluidized-bed and chemostat hybridoma cultures were run side-by-side after having been started with inoculum from the same frozen cell seed bank (Runstadler and Cernek, 1988). After 30-40 generations, the chemostat culture lost most or all of its MAb productivity, but MAb productivity in the fluidized bed system remained undiminished for over several hundred cell generations.

I. Hollow-Fiber Reactors

There is a need for animal-cell culture systems in which the cell is made to perform as though it were still part of the organ from which it has been derived. Such “pseudoorgan” cultures (Markus and McAleer, 1981) may be constructed about a bed of capillary tubes (hollow-fiber system) made from a variety of organic polymers such as polysulfone/polyacrylonitrile (Monsanto, 1978). The cells may be grown on the “inside” or within the lumen of the tubes or on the “outside” of the tubes. In either case, the cells will grow across the gaps between the tubes if on the outer surface or they will completely fill the tube lacuna if on the inside. By choosing an appropriate material for capillary construction, it is possible to provide cultured cells with an environment similar to that which pertains in uiuo. The semipermeable nature of the artificial capillaries mimics the performance of the body capillaries in that selective exchange of molecules can take place, and with suitable control of the cell bathing fluids, toxic products of metabolism as well as potentially valuable cell excretion products may be removed from the system, thus improving the longevity of the cells and enabling product recovery (Fig. 13). The hollow-fiber bioreactor shows some potential advantages, which may be summarized as follows: simultaneous separation of the cells from their extracellular products, high inherent productivities per unit bioreactor volume due to high cell densities, elimination of “washout” because the cells are trapped, and possible reduction in the overall capital and operating costs of the bioreactor unit. Scaleup may be accomplished by the modular addition of several units in parallel. A high cell concentration can be achieved because of the ability of the capillary network to allow for diffusion of nutrients to and metabolic wastes away from the cells without washing out cells.

181

VERTEBRATE CELL CULTURE TECHNOLOGY

Medium container

Continuous reDlenirhrnent

pH, PO?, T controlled

Dialysis

(a)

LINEAR FLOW

Nutrients

- __. -

- - Waster - - - --Doyr Medium container pH, P 0 2 , T controlled

(b) CLOSEDLOOP

Intermittent replenishment

--. -.

-

Medium container

pH, PO2. T

control chamber

Continuous replenishment

FIG. 13 Three medium-supply systems for hollow-fiber cultures. (a) Linear flow: temperature-, pH-, and pOz-controlled fresh medium is pumped through the fibers of the cartridge; the concentrations of nutrients and waste products in the medium are at a constant level during culture. (b) Closed loop: the controlled medium is pumped through the cartridge and recirculated to the medium container; nutrients are consumed and waste products accumulate; the medium is replenished batch-wise at predefined levels of nutrient or wasteproduct concentrations. (c) Open loop: a combination of the systems described above in which a small volume of medium is controlled and recirculated. (After Schonherr and van Gelder, 1988.)

Most hollow-fiber systems for cell culture are based on a cylindrical geometry (Fig. 13). The hollow-fiber capillaries are mounted inside a cylinder and potted at both ends. Cells are inoculated into the shell side, or the extracapillary space, through a port. Medium is generally introduced from one end of the cylinder into either the lumen or the extracapillary space and flows longitudinally toward the exit end of the reactor (Ku et al., 1981). Spent medium exits from both the shell side and the tube side. It is possible to select the molecular weight cutoff of the fiber to control the flux of molecules of different molecular weights across the fiber. Most of the hollow fibers used today to immobilize different cell systems are made from cellulose, modified cellulose, cellulose acetate, polypropylene, polysulfone, acrylic copolymers, and other polymers (Fohring e t al., 1980; Hopkinson, 1983; Schonherr and van Gelder, 1988). These hollow fibers have a highly porous surface wall about 70 pm thick to which the

182

SATISH J. PARULEKAR ET AL.

cells grow by attachment, and a cylindrical lumen of about 200 pm in diameter. The surface of the lumen is covered with a thin ultrafiltration layer that separates the immobilized cells and the lumen. Commercial hollow-fiber bioreactor units are available with luminal surface capacity ranging between 100 and 10,000 cm2. Units having as high as 100,000-cm2 luminal surface area can be obtained by adding a number of modules in a parallel flow mode. Hollow-fiber bioreactors have given encouraging results, and a wide variety of products have been produced. The attempt to cultivate mammalian cells in a hollow-fiber reactor dates back to the study by Knazek et al. (1972). Anchorage-dependent human choriocarcinoma cells were used for the production of human chorionic gonadotrophin (hCG). The concentration of hCG increased approximately twofold every 30 hr for 18 days. Hollow-fiber systems are also used for the cultivation of suspension-grown cells (Wiemann et al., 1983) and for production of MAbs. Notwithstanding the obvious advantages of pseudoorgan culture, problems arise because of the difficulties in obtaining a homogeneous environment within the hollow-fiber reactors. This often results from cell growth occluding the space in which the medium flows. Excessive cell growth may distort or even rupture the membrane, and thus destroy the fiber. The pores of the membranes in the hollow fibers may be plugged if the feed stream is not adequately prefiltered. Precipitates in serum-containing supply medium can obstruct the lumen of the fibers. The pores of the membranes can be clogged by the supply medium or by cell lysates. Depending on the kinetics of cell growth and product formation, the diffusion of nutrients and/or products through the ultrafiltration membrane might be a rate-limiting step. Due to possible diffusion limitations, the accumulation of toxic products might inhibit the metabolic activity of the cells entrapped in the hollow fiber. Thus, although these phenomena are not serious in small reactors, such as the ones commercially available (e.g., capafusion system) (CELLCO, 1975), when the system is scaled-up, yields do not increase proportionately. The countermeasure to this is to increase the complexity of the unit by adding thin fiber pads with special diffuser plates (micrometallic filters), which not only ensure even distribution of liquid but also allow retention of the cells within the apparatus (Monsanto, 1978). Larger scale units would have to be built of batteries of the largest successful stand-alone unit. Mass transfer in diffusion processes is less sensitive to clogging effects than in filtration processes. Schonherr and van Gelder (1988) found that in dialysis processes (Fig. 13), which are based on diffusion, hollow-fiber membranes retain high mass transfer rates even during prolonged cell cultivations. Because of the high cell concentrations, gradients of nutrient and metabolite concentrations occur along the axial direction of the reactor (Ku et

183

VERTEBRATE CELL CULTURE TECHNOLOGY

al., 1981). At times such concentration gradients can be growth inhibitory. As a result, cell concentrations are usually highest in the reactor near the medium inlet and decrease along the length of the reactor. Solutions to this problem include recirculation of the supply medium at substantial rates (Fig. 13) and periodic reversal of the direction of medium flow across the hollow fibers (Fig. 14) (Tyo et af., 1988). J. Membrane-Based Reactors

The Membroferm (Scheirer, 1988) is a modular fermentation system in which the cells can be grown between flat sheets of membranes. As the membranes are separated by a fluorocarbon matrix, cells can be grown either on the surface of the matrix fibers or, for suspension cells, in the openings of the fabrics. The goals for this functional design have been (Scheirer, 1988) ( a ) free choice of any commercially available flat membrane, ( 6 ) good geometry for diffusion of nutrients and circulation of medium, (c) excellent scaleup potential, ( d ) in situ sterilization, ( e )accumulation of product within the system, (f)harvesting of product independently from cells and medium, and ( g ) flexibility in operation mode and process control. By different combinations of the spacers, several different operation modes become possible. The simplest is the alternating stacking of medium and cell chambers to give a two-chamber system (Fig. 15). Any microfiltration membrane that retains the cells can be used. In principle, there is no difference from a hollow-fiber cartridge, but there is a wider choice of membranes, a better geometry for nutrient diffusion, and a larger volume (up to 4 liters) of cell mass (Scheirer, 1988). For retaining the product, an appropriate ultrafiltration membrane can also be used. In the three-chamber setup, the cells are immobilized

I

Cycle No. 1

I

Cycle No. 2

FIG. 14 Schematic diagram of the fluid cycling process used in ACUSYST hollow-fiber technology. This cyclingensures a consistent environment for up to IORcellsml-' and greater, depending upon the cell line. (After Tyo et a/.,1988.)

SATISH J. PARULEKAR ET AL.

184

b

a A

I I I I

4

Product

4

Medium

4

Product

Cells

Cells Medium

FIG. 15 Membroferm reactors with (a) two-chamber system and (b) three-chamber system. (After Scheirer, 1988.)

between two different flat membranes (Figs. 15 and 16). The membrane between the nutrient medium and the cells is an ultrafilter with a molecular weight cutoff appropriate for the particular product. The membrane must be permeable enough to provide the cells with nutrients and oxygen. Products desired to be retained, such as antibodies, must not pass this barrier and should thus remain within the cell compartment. The other side of the cell chamber is a microfiltration membrane of a pore size suitable for retaining the cells from the product chamber, e.g., a 0.2-pm sterile filtration membrane. The product traverses this membrane by diffusion into the product chamber (Figs. 15 and 16). With this configuration, it is possible to feed the cells continuously and wait for a suitable product concentration. This closely resembles the situation for mouse ascites; i.e., product harvest can be operated repeatedly at suitable time intervals by replacing the contents of the product chamber with fresh medium without removing cells from the reactor. There may be an advantage over mouse ascites, because of the exclusion of large serum proteins from the product by the feeding membrane barrier. As mentioned above, there is the possibility of using the third chamber for other purposes, e.g., oxygenation or induction. Stacking of the system is possible (Fig. 15) (Scheirer, 1988).

VERTEBRATE CELL CULTURE TECHNOLOGY

185

A @-

U FIG. 16 Periphery installation of a Membroferm with conditioning vessel (A), and reservoirs of growth medium ( B) , spent medium (C), product-collecting medium (D), and product (E). Probes are indicated as follows: TI, temperature indicator; FI, Flow-rate indicator; pOzl. oxygen tension indicator: PHI. pH indicator; C, automated control circuit. (After Scheirer. 1988.)

K. Fiber-Bed Bioreactors Glass has been a popular choice among the numerous materials used for cell immobilization. One of the common glass supports is in the form of beads. External medium circulation loops driven by a pump (Thornton et af., 1985) or airlift (Whiteside and Spier, 1981) have been used to provide adequate liquid flux to the glass-bead beds. Since particles such as glass beads tend to be densely packed, small glass beads often result in high hydraulic resistance. Usually glass beads of 1- to 3-mm diameter are used in order to avoid high pressure drops. However, the larger the bead diameter, the lower is the surface-to-volume ratio for the beads (a key parameter as far as mass transfer is concerned). The low surface-to-volume

186

SATISH J. PARULEKAR ET AL.

ratio for cell attachment makes high cell densities difficult to achieve and potentially leads to low volumetric productivities (Crouch et af., 1985). Packed beds of small-diameter glass fibers have been considered a good candidate as an alternative support for cell attachment and growth due to their high surface-to-volume ratios. In addition, at a constant surface-tovolume ratio, packed beds of glass fibers provide controllable packing density and result in both lower pressure drops and lower mass transfer resistance than packed beds of glass beads. Recombinant y-CHO cells that have been engineered to produce y-interferon have been grown on glass fibers of diameters of 80 p m (Perry, 1987), and 30 and 24 pm (Chiou et al., 1991). An important barrier for scaleup is the shear sensitivity of animal cells. Due to the lack of cell wall and relatively large size, animal cells are easily damaged by fluid motion. Cells attached to a fixed support are not able to reduce fluid forces and torques by rotation, making anchorage-dependent cells more susceptible to shear forces resulting from agitation and aeration. This leads to limitations for the existing oxygenation methods for largescale animal-cell cultures (Spier and Griffiths, 1982). Among the oxygenation methods, gas sparging has been shown to be a simple, efficient, and scaleable method. However, it has been reported that sparging leads to cell damage and decreased cell growth (Handa et af., 1987). These detrimental effects could be the result of either gas-liquid interfacial forces or hydrodynamic effects. If these adversities can be solved, sparging would be a preferable in situ method of providing oxygenation for anchoragedependent cells (Chiou et af., 1991). The fiber-bed bioreactor consists of an inner draft tube and an annular packed glass fiber bed (Fig. 17). Cells are immobilized on the glass fibers in the annular region and medium fills both the draft tube and the annular region. Air is introduced from the base of the draft tube and oxygenates the medium in the draft tube, and bubbles disengage at the upper fluid surtace. The density difference between the aerated fluid in the draft tube and the bubble-free fluid in the annular fiber bed leads to the global circulation of the medium in the bioreactor. This bioreactor design provides the following unique characteristics (Chiou et d., 1991; Murakami et af., 1991): (a) the reactor can provide a very high surface-to-volume ratio in the annular section; (b) in situ oxygenation with no external loop can be achieved; (c) the cells are not exposed to gas-liquid interfaces since no gas bubble is entrained by the oxygenated medium in the annular region; ( d ) nutrients are convectively supplied to the immobilized cells when the fluid is flowing through such a system; ( e ) very little hydrodynamic cell damage can result since low turbulence and low shear stress occur due to the low liquid velocity in the annular fiber bed; (f) no mechanical agitation or external pump is required (this avoidance of moving parts prevents

VERTEBRATE CELL CULTURE TECHNOLOGY

107

FIG. 17 A cross-sectional view of the fiber-bed bioreactor. Arrows indicate the direction of Row of medium in the reactor. Reprinted by permission of Wiley-Liss, a division of John Wiley and Sons, Inc., Biotechnology and Bioengineering, Vol. 37, p. 756, T-W. Chiou, S. Murakami, D. I . C. Wang, and W-T. Tu, copyright 0 1991.

cell damage by local high shear regions and also decreases the risk of contamination); (8)the reactor can be operated in a batch or continuous/ perfusion mode; and (h) high scaleup potential is expected. The cell concentration was found not to be significantly different across the radial direction of the fiber bed (Chiou er al., 1991), which indicates that channeling was not a serious problem for cell cultivation in the fiberbed reactor. The shear stress on the cells was very low and only a small fraction of the cells was detached from the fibers. High cell densities (typically 7 X 10" cells/liter) and high oxygen uptake rates (6-8 mmol/ liter-hr) were reached without encountering nutrient transport limitations (Chiou et al., 1991).

L. Packed-Bed Reactors

Anchorage-dependent cells are ideally suited for immobilization within appropriately designed bioreactors (Nilsson and Mosbach, 1980). Cell attachment is mediated through specialized structures, called adhesion plaques, on the underside of the cell (Brown et al., 1988). Furthermore, unlike the majority of microorganisms that have been incorporated into

188

SATISH J. PARULEKAR ET AL.

immobilized cell systems, anchorage-dependent mammalian cells secrete at least a portion of the factors required for adherence to appropriate surfaces. For example, collagen and fibronectin facilitate cell attachment and are also produced by many anchorage-dependent cell lines (Alitalo er al., 1981; Brown er al., 1988;Castellini et al., 1986).Additional attachment factors are normally provided by serum supplements added to the basal growth medium. The primary advantages of immobilized cell systems are cell-free spent media (which simplifies downstream processing), extended production periods, high cell densities, and rapid media turnover rates within the bioreactor (van Brunt, 1986). The maximum cell densities reached by immobilization in any reactor are limited by its mixing parameters and also by the cell type, morphology, cell size, etc. The practical limits may be assumed to be somewhere between lo7 and lo8 cells/ml of bioreactor volume. Packed-bed reactors used for vertebrate cell growth are normally of the perfusion type. The cells are firmly attached to the solid substratum in the bioreactor while the bulk of the medium is held in another vessel which is attached to the bioreactor via a circulation system. Calcium alginate gels have been employed for immobilization of MAb-secreting cells (Nilsson and Mosbach, 1980). Besides this, two fused element systems have been used for vertebrate cell immobilization; one is based on the generation of a glass matrix, and the other is based upon special-shaped stainless-steel elements combined in such a fashion as to promote maximum mixing (Huber, 1974; Huber and Schutz, 1975a,b). The packed-bed perfusion system for growing industrial quantities of anchorage-dependent cells is simple to use, robust, and reliable. The apparatus can be built in-house and it is easy to monitor and control, as the medium-holding vessel can provide adequate accommodation for probes and the aeration of this vessel does not interfere with the cells growing on the bed elements. A significant advantage of this type of system is the ability to remove spent growth medium or chemicals that control the cellular biochemistry rapidly and efficiently and to change the culture fluid volume-to-cell volume ratio. This latter feature enables cell growth to be achieved under one set of conditions, whereas product generation may be achieved under conditions of higher cell concentrations. Methods have been developed to trypsinize the cells from such culture vessels (Spier, 1982) and the cells so produced have been shown to be useful seed materials for the next culture. It has also been shown that the cells grow throughout the matrix of the bed. Although packed-bed bioreactors suffer from the disadvantages inherent in the inhomogeneous nature of the system, they can be scaled-up by a large factor (up to 15,000liter medium volume) and therefore they are expected to serve a useful function for the manufacture of materials derived from cultured anchoragedependent animal cells (Whiteside, 1983).

189

VERTEBRATE CELL CULTURE TECHNOLOGY

1. Glass-Bead Reactors

For their growth in large-scale cultures, anchorage-dependent cells must be provided with a compatible matrix possessing high surface-to-volume ratios and a structure that allows for the unrestricted delivery of oxygen and nonvolatile components of growth medium. The packed glass-bead system satisfies this requirement (Varani et al., 1983). The components of the glass-bead system are united through a circulating fluid loop of constant volume to which fresh medium is added and from which conditioned medium is removed (Fig. 18). The primary function of continuous medium circulation is to shuttle oxygen from the oxygenator to the cells within the packed bed and to remove excess CO, produced through cellular respiration. A secondary and less-well-recognized function of active medium circulation is to prevent the overgrowth of cells and the resulting channeling of fluids within the packed bed (Brown et al., 1988). Regardless of the cell line chosen or the medium in which it is grown, cells do not divide as rapidly in a mature glass-bead system as they would in a monolayer culture. In this sense, the glass-bead cultures assume the characteristics of an extended stationary phase seen in refed static cultures. The major problems to be overcome with scaleup include provision of adequate pumping and high-capacity oxygenators, sterilizability , and inoculation strategies (Brown et al., 1988). The cell lines grown in glassbead bioreactors have included (target product indicated in parentheses) mouse mammary fibroblast and Chinese hamster ovary (heterologous proteins), human kidney carcinoma (constitutive product), human melanoma (tissue plasminogen activator), and hepatocyte/hepatoma hybridoma (acetyl cholinesterase) (Brown et d.,1988). Samplo Port n

Product Collectlon

Glass Beads

-

Dlrectlon of Flow

FIG. 18 Schematic diagram of glass-bead bioreactor system. (After Brown rr a / . , 1988.)

190

SATISH J. PARULEKAR ET AL.

2. Ceramic Reactors One of the newer technologies employs nonporous and porous ceramic matrices (Fig. 19) to immobilize cells, suspension or attachment-dependent, within a continuously recycled perfusion culture loop (Berg, 1985; Berg and Bodeker, 1988). The ceramic matrix provides a large surface area in a small volume; it minimizes shear forces on the cells; and serumfree medium can easily be employed. The smooth-surfaced (nonporous) ceramic is generally employed in processes where cell harvesting, be it by trypsinization, cold-shock, or changes in pH, is necessary. The porous ceramic contains pores of diameters of ca. 50 pm and has porosity in the range of 40%. This type of core is generally used for the production of secreted biomolecules in low-serum or serum-free medium. Because of the porous nature of this ceramic, it offers a larger surface area per unit matrix volume than the nonporous ceramic, but does not allow for complete cell harvesting (Berg and Bodeker, 1988). The schematics of the ceramic bioreactor are shown in Fig. 19. The main loop provides for constant recirculation of the medium through the

..

Harvest

Medium

Feed

reservoir

reservoir

reservoir

FIG. 19 Schematic of the ceramic bioreactor. Closed-loop process for medium circulation. pH and p 0 2 are monitored before and after the medium has perfused through the culture. Gas permeators maintain controlled levels of O2and COz. Auxiliary pump is for the addition of basic solution for pH control. The two secondary flow paths are for the continuous feed and harvest of medium. (After Berg and Bodeker, 1988.)

VERTEBRATE CELL CULTURE TECHNOLOGY

191

culture, while the two secondary loops provide for the continuous replenishment with fresh medium and continuous removal of the exhausted medium containing products and/or cellular toxins (continuous feed and harvest). The PO, and the pH of the medium are measured upstream of the immobilized culture and these values are utilized for the constant feedback control of these parameters. This is accomplished by the continuous mixing of the gases (N,, o,, CO,) in the correct percentages within the gas permeators so that the desired values or set-points for the pH and p 0 2 are maintained. The pH electrode activates the auxiliary pump for the addition of a base solution to raise the pH value, when required. To obtain uniform and consistent growth throughout the length of the core, the flow rate of the medium must increase proportionally in response to the increase in number of cells immobilized within the ceramic during the growth phase, and plateau just as the cell growth plateaus. It has been found that p 0 , value of the medium is the best parameter to determine medium flow rates in a perfusion system as utilization of oxygen is much more rapid than that of any other nutrient. Because of this, the downstream PO, value is used to determine the flow rate (see Fig. 19). Because the cells are immobilized in the closed ceramic core, it is impossible to visualize the cells or to determine cell numbers by counting. The oxygen consumption rate (OCR) provides a reliable, continuous, on-line method for determination of cell numbers in the ceramic core as well as the metabolic state of these cells (Lydersen et al., 1985). Once cells become confluent, the rate of oxygen utilization per cell is decreased and the direct linearity between cell number and OCR is not as reliable for prediction of total cell numbers (Balin et al., 1976). A variety of biomolecules (including vaccines, urokinase, pro-urokinase, tissue-type plasminogen activator, murine and human MAbs, factor VIII, interferons, interleukin-2, human growth hormone, and colonystimulating factor) have been produced from mammalian cells grown on the ceramic matrix. These were produced from genetically engineered cell lines, from cells that inherently secrete the product, or from primary cells that required an induction step for production (Berg and Bodeker, 1988; Marcipar et al., 1983). For example, the tPA production rate was 2.5-fold higher in the Opticell ceramic culture system than in shake flasks (Berg and Bodeker, 1988), which was indicative of the better cell growth, a higher metabolic activity, and increased productivity of the ceramic matrix culture. Since ceramic matrix reactors are operated in a plug-flow fashion, nutrient and metabolite concentration gradients may have a significant effect on cell growth. Like the hollow-fiber systems, the scaleup will probably be more successful using units in parallel or using units with other configuration. The true ability of the ceramic matrix to be scaled has

192

SATISH J. PARULEKAR ET AL.

yet to be convincingly shown in terms of very large production units, i.e., those comparable with stirred tank fermentors with volumes in excess of 1000 liters. Research must be carried out not only on multiple ceramic matrices, but also on ceramics with larger surface areas. In addition, for a true production facility employing the ceramic matrices for cell immobilization, one must be able to include, on-line, all aspects of production, from medium preparation/storage, sterilization, adaptable scaling, to final concentrated product recovery (Berg and Bodeker, 1988).

V. Design and Optimization Considerations

The optimization of an in uitro cell culture system is possible on two general levels of manipulation, first by improvement of the genotypes of the cell strains and second by process design and control, which are aimed at bringing about the optimum phenotypic expression of the genetic potential of any production strain. The makeup of a culture system is a composite of three basic elements: ( a ) cultivation techniques, which include such aspects as reactor configuration and how to suspend, mix, or immobilize a cell population; ( 6 )the choice of a proper cultivation process (batch, fed-batch, or continuous culture; i.e., flow of phases within or around a reactor); and (c) sophisticated parameter estimation and control. In designing the entire cultivation system and the engineering of a bioreactor, the techniques of suspending cells or supporting cells with solid or soft matrices, the methodology of feeding cells with nutrients, etc., must be adapted to the particular cellular requirements and characteristics of cell growth and product formation. In static deep culture systems, generally used at the small scale for all types of animal cells, the interfacial mass transfer is predetermined by the geometric configuration and the liquid level. Their scaleup is easily achieved by keeping the relative interfacial area constant, a fact that leads to multiples of geometrically identical units (Katinger and Scheirer, 1985). The new demands for animal-cell culture will necessitate economization of fermentations, a large part of which may be improved reactor productivity through better engineering design. Chief among the engineering problems of animal-cell culture is the provision of adequate gas-liquid mass transfer without affecting cell viability, product activity, or downstream processing (Aunins et al., 1986). A cell culture will display special characteristics (cell growth and/or product release) in response to a certain set of chemical and/or physicochemical factors. The physical environment of a cell culture is largely affected by the configuration of a bioreactor and the power supplied, and the chemical input to a large extent is determined by the composition of

VERTEBRATE CELL CULTURE TECHNOLOGY

193

the nutrient medium. The resulting cell activity is fed back upon both levels of input, physically and chemically (e.g., by cell growth or cell aggregation and by secretion of substances). The overall result of these biological, chemical, and physical interrelations defines the status of the cell population (Katinger and Scheirer, 1985). Optimization of cell growth and/or product formation is achieved by interaction at different levels, by cell strain improvement (selection, mutation, and genetic recombination), improved parameter control, formulation of suitable nutrient media and stimuli (Spier, 1980), choice of proper techniques of cultivation (bioreactor configuration and bioreactor performance), and application of a suitable cultivation method (open or closed, homogeneous or heterogeneous, filtration or dialysis). Compared to bacterial cell cultures, animal-cell cultures are 1.5-2 orders of magnitude less productive of biomass due to lower cell densities and lower rates of propagation. Submerged deep culture reactors for animal-cell cultures must therefore be designed for effective liquid mixing combined with low stress factors (mechanical and hydrodynamic) and for adequate interfacial mass transfer capacity. An important part of the bioreactor design is to assess its implication on the downstream processing steps (Rosevear and Lambe, 1988). These may include cell separation as well as product separation and purification steps. One approach that has important economic benefits is to have a bioreactor design that achieves simultaneous fermentation with cell and/ or product separation (Crespi et a l. , 1981; Cysewski and Wilke, 1977; Glacken et af., 1983a,b; Margaritis and Wallace, 1984). Usually a membrane is placed within, or outside, the bioreactor to separate the cells from their metabolic products during the course of fermentation. Another means of achieving the same objective is to immobilize the cell system in different ways as discussed in the previous section. This approach results in high cell densities per unit bioreactor volume, which increases manyfold the volumetric productivity of the bioreactor system. Increased volumetric productivities may result in reduced capital and operating costs for the bioreactor system. Last but not the least, in terms of economic significance and commercialization of a process, is the potential of scaling up the novel bioreactor design from the laboratory to the industrial level. Some bioreactor systems may work extremely well in the laboratory, but become impossible or very inefficient to operate when they are scaled-up. A. Cellular Metabolism

An important criterion in optimization of vertebrate-cell cultures, or for that matter any cell culture, is a basic understanding of the microbiology of the particular cell system. This includes, among other things, the mor-

194

SATISH J. PARULEKAR ET AL.

phology and the nutritional requirements for optimal cell growth. One would like to know as much about the biochemical pathways responsible for cell growth and the production of primary and secondary metabolites as possible. This information is needed to control the bioreactor macroenvironment, which in turn affects the microenvironment within the cell, and thus helps to optimize the formation of the product(s) of interest. To reduce the costs of producing animal-cell products on a large scale, it is desirable to maximize the cell density. The higher the cell density, the higher the product titer will be. As the product titer is increased, the costs of purification of the biological product decreases. Since serum components are thought to catalyze cellular reactions and are not depleted to a great extent in the medium, no additional serum components are needed at higher cell densities. Therefore, at higher cell densities, the unit yield of product per unit of serum used in the medium is higher. Since serum is the predominant component of animal cell culture media, high cell densities greatly reduce the cost of producing animal biologicals. Research in the field of cultivation procedures should be centered upon control of nutrient requirements in order to achieve higher cell yields per unit culture volume. The development of serum-free medium is of great interest, as this will facilitate culture manipulation. For instance, it will be unnecessary to change to a serum-free medium when virus is added to a culture, and the downstream processing of cellular components will be greatly facilitated. This is of major interest in view of the increased application of mammalian cells as hosts for recombinant-DNA genes for the production of cellular biologicals. The kinetics of cell growth and product formation is indispensable in determining residence times in a bioreactor system. Whether the product is growth or non-growth associated will also in part determine the sequence and type of bioreactor system. As the number of genetically modified cell systems used in the production of important metabolites is increasing, we need to develop useful kinetic models to predict their behavior in a bioreactor system. The possible inhibitory effects of the substrate and/or products on the metabolism of the cell type being used are also part of the kinetic requirements. If a genetically modified cell system is to be used successfully in a bioreactor system, it is extremely important that we know its genetic stability under actual bioreactor operating conditions. When dealing with a cell line whose genetic stability is limited, there is a risk of loss of productivity upon scaleup. Due consideration must be given to this possibility when selecting a culture system. Examples of such cell lines may be found among hybridomas, particularly interspecies hybridomas, and genetically engineered cell lines. First of all, one should try to stabilize the cell line by cell-biological means. Then one might try to slow down the loss of genes by reducing the overall number of cell

VERTEBRATE CELL CULTURE TECHNOLOGY

195

duplications and by decreasing the population growth rate. Both can be achieved by appropriate cultivation systems and methods. Total cell duplications are minimized when the loss of cell mass is prevented with the help of cell retention systems. Such systems also enable the control of cell growth rate. It has been shown that prolonged exposure to ammonia concentrations in excess of 4 mM can inhibit cell growth. Ammonia generation can be a serious problem in animal cell cultures. For example, it is estimated that a culture of HeLa cells growing from a cell density of 3.4 x 10’ cells/ml to a cell density of 1 X lo7 cells/ml could generate approximately 30 mM ammonia (Glacken et al., 1983b). On a small scale, frequent replenishment with fresh medium would keep the concentration of these toxic compounds from reaching inhibitory levels. On a large scale, this is not desirable, since serum-supplemented medium is very expensive, and thus, it would be very wasteful to discard serum components (which account for the great majority of the medium cost) to remove lactate and ammonia (Glacken et al., 1983b). One strategy is to modify the culture medium so that the cell’s metabolism is altered to generate less waste products. Eagle et al. (1958) substituted galactose for glucose as the carbohydrate source in the medium, resulting in a 67-fold decrease in lactic acid generation. Fleischaker (1982) reported the same effect by feeding glucose to FS-4 cells in a fed-batch manner, such that the glucose concentration in the medium was relatively low at all times [OS mM (fed-batch) vs 20 mM]. Glacken et al. (1983b) demonstrated that ammonia generation can be reduced by over 60% by continually feeding glutamine to the culture to keep the glutamine concentration at a consistently low level [0.2 mM (fedbatch) vs 4 mM]. 8. Mixing in Bioreactors

Mixing phenomena, although not necessarily detrimental to the cells, have to be carefully considered in the design of any submerged deep culture system and whenever free cell surfaces are exposed to fluid or mechanical stress. It is obviously very complicated to achieve optimum mixing in a submerged reactor in order to achieve reasonable mass transfer rates, with an appropriate degree of homogeneity in the bulk liquid, and minimize detrimental effects on the microenvironment of the cell surfaces (Katinger and Scheirer, 1985). The minimum mechanical power input per unit mass of fluid needed to achieve a homogeneous cell distribution is expected to depend on the system geometry (circulation patterns, dead spaces, shear rates, foaming, etc.), cell properties (density, size, and shape), solids holdup, and the physical properties of the fluid. Local fluid shear rates

196

SATISH J. PARULEKAR ET AL.

may be high enough to break cells even at the minimum mechanical power input. The net specific growth rate of cells may be reduced as a result and the system may fail totally. With scaleup based on power input per unit mass of fluid, these effects are expected to be magnified in large-scale equipment. The conditions needed to prevent cell lysis are in direct conflict with those needed to maximize mass transfer rates and to ensure an even distribution of cells and reactants. The hydrodynamic characteristics of the bioreactor are of paramount importance because they affect both the mass and the heat transfer capability of the bioreactor. The geometric configuration of the bioreactor vessel and the mixing characteristics of a particular impeller (or other mixing device) are important. Effective mixing to minimize pH, temperature, and concentration gradients within a stirred tank bioreactor are especially important during scaleup to industrial size units. The viscosity of the fermentation broth and whether it behaves as a Newtonian or non-Newtonian fluid will have a profound influence on mixing as well as mass and heat transfer (Atkinson and Mavituna, 1983). The shearing effects of a specific impeller system often dictate whether shear-sensitive and fragile mammalian cells can be cultivated in the particular bioreactor system. In order to accommodate shear-sensitive cell systems, novel impeller designs or other mixing systems that give gentle mixing at very low shear rates have been devised (Feder and Tolbert, 1983; Margaritis and Wallace, 1982; 1984). Most of the mechanically agitated vessels for animal cells grown on microcarriers or in suspension use marine-type impellers or pitched-blade paddles for agitation. However, there have been some attempts to modify the agitation mechanism. In the studies by Tolbert and Feder (1983), a sail-type agitator was used for both suspension and microcarrier cultures. With this agitator, the agitation rate required (as low as 10 rpm) to provide the same pumping capacity is much lower than that of other types of agitators, such as paddles. Such an agitation mechanism provides a low shear force (Tolbert er al., 1988). Another nonconventional mixing mechanism involves the use of the “scull” type agitator (Barteling, 1984). The scull is anchored on the head plate of the fermentor while the shaft rotates at an angle to the fermentor axis. An indigenously fabricated impeller assembly, with all parts made from Teflon, has recently been employed for propagation of recombinant vaccinia virus in HeLa cultures (Chillakuru et al., 1991). Impellers that can force vertical circulation currents over the entire vessel are considered to be most effective for liquid mixing (lowest power for specific circulation time). Most designs aim to recirculate material through the high-shear region close to the impeller to ensure effective mixing. The vibromixer is composed of a horizontal disk with conical apertures attached to a vertical shaft that rapidly oscillates up and down

VERTEBRATE CELL CULTURE TECHNOLOGY

197

(Girard, 1977). The vibromixer circulates the medium in vertical, rather than horizontal, direction, resulting in less shear force to obtain an adequate dispersion of cells. Careful design of mechanically agitated vessels may be able to overcome the problem of cell fragility for some cell lines but it is inevitable that applicability of such designs will be limited; there will undoubtedly be a scale of operation or a cell line for which the local shear rates at the impeller give unacceptable levels of cell lysis (Pollard and Khosrovi, 1978). Zwerner et al. (1975) reported results which indicate that the choice of the proper agitation system has to be carefully evaluated for any particular case. With different culture vessels (equipped with spinner, vibro, and marine impellers) of sizes up to 14 liters, they found a marked influence of agitator system on the expression of surface antigens by mouse cells. Besides the agitated vessels, bubble column reactors have also been employed for the cultivation of some suspension grown cells (Katinger et al., 1979). For large-scale cultivation with microcarriers, standard commercially available biorectors may be applied as long as they are equipped with a mixing device by which the microcarriers can be kept in suspension at slow agitation and they do not have slack points where microcarriers may accumulate (van Wezel and van der Velden-de Groot, 1978). In this regard, it is advisable to use bioreactors with a rounded bottom and smooth walls. C. Oxygen Transport

The most critical feeding substance, particularly with high cell density animal cell cultures, is oxygen, because of its low solubility in aqueous solutions. At oxygen concentrations below approximately 1 mg/liter, the metabolism is altered drastically (Glacken et al., 1983b);one has to ensure that no portion of the cell culture is exposed to such low concentrations of oxygen. This is particularly important with encapsulation systems in which the oxygen supply is governed by diffusion (Scheirer, 1988). Traditionally, large-scale animal-cell cultures have employed surface aeration and in some cases low levels of sparging to accomplish aeration. Surface aeration suffers from the fact that surface area per unit volume in a bioreactor (such as a stirred tank) decreases with the inverse of the reactor diameter for geometrically similar systems. Sparged aeration may be carried to the extreme in airlift operation, providing large mass transfer coefficients under gentle agitation. However, vigorously sparging a serumcontaining culture can cause protein foaming, cell death due to hydrodynamic or interfacial effects, and product or nutrient denaturation at the gas-liquid interface (Aunins et al., 1986). The creation and suppression

198

SATISH J. PARULEKAR ET AL.

of foam raise several problems: microcarriers and suspension cells may be entrained in the foam phase; some products, nutrients, and proteins carried over in the foam phase may denature at the gas-liquid interface; and antifoams that remedy these problems may be toxic to cells or difficult to separate from the product. Direct sparging of an oxygen-carrying gas through the medium can cause cell lysis above certain gas flow rates (Kilburn and Webb, 1968). Some cells appear to be more sensitive to the presence of bubbles than others but the reasons for this are not well understood. The influence of scaleup on gas-cell interactions cannot be predicted until cell fracture mechanisms are more clearly understood. Scaleup efforts should thus be focused on techniques that provide increased interfacial area and increased driving force. The latter can be achieved by overpressuring the reactor and feeding pure oxygen; this may have limits with oxygen toxicity (Kilburn and Webb, 1968). The interfacial area may be increased by installing more tubing, or by installing a surface impeller that disrupts the surface film but does not significantly contribute to overall power input. Although these techniques will greatly increase the absolute level of oxygen transfer, they are unlikely to affect the scaleup dependency. The ability to provide an adequate oxygen supply to mammalian cells at increasing culture volumes is the most critical barrier to the scaleup of mammalian cell cultures, especially suspension systems. Unless oxygen is constantly supplied to the medium, oxygen limitation will result. Oxygen may be supplied indirectly across tubular polymeric membranes. The oxygen transfer rate could be controlled by adjustment of the oxygen partial pressure within the tubes without risk of bubble formation within the medium (Fig. 20) (Aunins et al., 1986; Lehmann er al., 1988). The membranes would also act as a sink for carbon dioxide produced in conjunction with cell metabolism. Silicone rubber, often used as a coating for glassware to prevent cell adhesion, has a suitably high oxygen permeability. The aeration rates from both the free surface and a coiled silicon tube in a membrane-oxygenated stirred tank were investigated by Aunins et al. (1986). The major effect of scaleup at constant power unit volume on the mass transfer rate achievable in the bioreactor is the loss of interfacial area per unit volume. A Chemap fermentor, equipped with two impellers, the lower one a four-blade pitched turbine for off-bottom suspension of solids and the upper one a four-blade flat turbine, was modified by wrapping a silicone tubing (0.15-cm i.d., 0.2 cm 0.d.) in a helix around the four fermentor baffles (Fig. 21). The tubing vented to the reactor headspace. Sparged aeration of cultures was shown to be a promising technique for those cultures that can withstand the hydrodynamic and/or interfacial forces generated by bubbles in culture (Aunins et al., 1986). If interfacial adsorption is not a problem and hydrodynamic damage of cells may be

VERTEBRATE CELL CULTURE TECHNOLOGY

199

Aualmn

I

FIG. 20 Membrane stirrer reactor for bubble-free aeration with a porous membrane. The head-space is closed but connected to the gas input to achieve pressure equilibrium between the head-space and membrane. Bubbling cannot occur. (After Lehmann et al., 1988.)

avoided, sparged reactors represent the most economical homogeneous reactor type for suspension and possibly microcarrier cultures. D. Control of Environment in Bioreactor

The basic function of the bioreactor is to control the macroenvironment that will influence the metabolic activity of the cells which grow in that

SATISH J. PARULEKAR ET AL.

200 A i r in

Air out

Baffles

.4-FBT Silicone tubing

- 4-PBT

1

10 LITE R FERMENTOR

FIG. 21 Ten-liter fermentor with silicone tubing. Impellers are four-bladedflat-blade (top) and pitched-blade (bottom) turbines. Reprinted by permission of Wiley-Liss, a division of John Wiley and Sons, Inc., Biotechnology and Bioengineering Symposium. No. 17, p. 703, J. G. Aunins, M. S. Croughan, D. I. C. Wang, and J. M . Goldstein, copyright 0 1986.

environment. The regulation of important parameters, such as temperature, pH, oxygen concentration, C 0 2 concentration, and other metabolic parameters requires the design of appropriate sensors (Mattiasson et al., 1983), which need to be interfaced with microprocessors for efficient computer control of the process. The bioreactors for large-scale cultivation processes (generally above 5-liter working volume) are generally equipped sufficiently to control temperature, pH,pO,, and stirrer speed. It is preferable that they are equipped with a supply of nitrogen and oxygen for PO, control and a CO, supply for pH control. The most common procedure for pH control above the set-point is the addition of more or less C 0 2 to the gas mixture passed over the culture surface. Below the set-point, the pH can be controlled by addition of a solution of bicarbonate or sodium hydroxide. As the main cause of changes in the pH is the conversion of glucose to lactate, minimizing accumulation of lactate will facilitate the pH control.

VERTEBRATE CELL CULTURE TECHNOLOGY

201

Oxygen control is generally the major problem in large-scale cultivations employing microcarriers. The main reasons for this are (a) transfer of oxygen from the gas headspace to the culture fluid is low, as only slow agitation is used, since otherwise cells may be detached from the microcarriers; and (b) sparging of air or oxygen should be avoided, as cells may be damaged by air bubbles and, in addition, foam formation may result in the flotation of microcarriers. Therefore, oxygen is normally controlled by increasing the 0, tension in the gas mixture passed over the culture surface (van Wezel, 1982). Other possibilities are aeration of the culture fluid outside the fermentor in a recycling system, or even more directly by sparging air through the culture fluid inside a cylindrical filter mounted on the stirrer shaft (van Wezel, 1985).

VI. Concluding Remarks Recent advances in vertebrate cell culture technology have resulted in better understanding of structural and functional properties of cells and their organelles and mechanisms of various cellular processes, the latter being of immense value in the development of cell-based products. The vertebrate-cell-based products (Table 11) are being developed for health care in both human and veterinary areas, biologicals, growth mediators, hormones, enzymes, and monoclonals. This field of research is growing rapidly and will continue to witness growth for next several years and well into the twenty-first century. In view of this, optimization of the animalcell-based production processes is of great importance. Evidence to support this contention is provided by the projected financial markets for sales of biotechnology-based products, which are predicted to reach about 11.4 billion U.S. dollars (basis: year 1989) by the year 2000 (Shame1 and Chow, 1989) (Table 111). The health care market is a natural target for the vertebrate cell culture technology since it has a great potential for high-valueadded products. Human and animal health care products will continue to account for the majority of sales. The contribution of products that are made exclusively by animal-cell culture will therefore be significant. The understanding of the areas such as cellular growth requirements and the development of new media is expanding massively though much is still unknown. Work on the identification of key components of the serum has progressed considerably and is continuing to do so; yet many fundamentals of vertebrate cell cultures are still poorly understood. As a result, cultivation of vertebrate cells continues to remain an art rather than a science. Nonetheless, there is increased awareness of the urgency of transforming vertebrate-cell culturing from an art to an exact science and

TABLE II Products from Vertebrate Cells Vaccines Human Veterinary Disease

Cell culture

Foot-and-mouth disease Newcastle disease Marek’s disease Cattle plague Rabies Canine distemper Canine hepatitis Feline rhinopneumonitis Bovine viral diarrhea Rift Valley fever African horse sickness Equine encephalitis Equine rhinopneumonitis Swine fever Fowl pox Mink enteritis

BHK calf and pig kidney Pig kidney. chick embryo Chick embryo Kidney CEF, BHK Dog kidney Pigiferrel kidney CEC Embryonic kidney CEF Monkey kidney CEC Pig kidney Kidney CEF Mink kidney

Principal vaccines Polio (Salk) Polio (Sabin) Measles Mumps Rubella Rabies Yellow fever Influenza

Limited application vaccines

Hormones Hormone

Herpes simplex Growth hormone Cytomegalovirus Prolactin Varicella-Zoster Insulin Respiratory syncytial virus ACTH Adenovirus Thyroid Tickborne encephalitis Parathyroid Adrenal Luteinizing Folliclestimulating Thyrocalcitonin Erythropaetin TSH Thyrotropin (TSH) Gonadal

Cell source

Whole cells and other products

Assays Bone marrow lymphocytes and skin Fetal rat pancreas grafts Hybrid cells Pituitary cells. HeLa V3 Cell constituents Chromosomes. DNA, mRNA, Organelles Parathyroid adenoma Tumor antigens Adrenal cortex tumors Attachment factors Anterior pituitary (human. Fibronectin rat. sheep) Laminin Medullary carcinoma Transferrin Monoclonal antibodies Antibodies Fibroblast and Hemopoietic cells Interferons Sheep pituitary l ymphoblastoid Fibrinolytic Placental and follicle cells Enzymes lmmunoregulator lnterleukins Plateletderived growth Growth factors factor. epidermal growth factor. and nerve growth factor Other products Heparin Tumor angiogenesis factor Albumin Blood clotting factors Chondroitin sulfate Rat pituitary tumor

Whole cells

203

VERTEBRATE CELL CULTURE TECHNOLOGY TABLE 111 United States Biotechnology Market 1990-2000 (in U S $ millions)a

Human diagnostics/ Analytical reagents/ Instruments Human biotherapeutics EnvironmentaVmarine Agricultural/industrial Total

1990

1995

5660

9980 5000 2186 1498 18,664

1100

1287 623 8670

2000

AAGR~ 1990-2000

18,310 10,380 4,695 4,490 37,875

12.5% 25.0% 14.0% 21.8% 15.9%

“ Source: “Biotech Forum Europe” Vol. 9, p. 100. Business Comunication Co., Heidelberg, Germany (1992). AAGR: Average annual growth rate. as a result, the “black box” approach of viewing the vertebrate cells is gradually being abandoned. The development of culture media specifically designed to meet the specific requirements of various cell types is becoming increasingly important in order to increase the productivity of individual metabolites. This may be particularly important in reducing costs of scaleup. Culture media tailor-made to meet the nutritional requirements of individual cell lines minimize their requirements of essential and expensive nutrients and hence, result in the elimination of costly and nonessential nutrients. The advent of improved culture processes is leading to increases in both the quality and the quantity of MAbs. Work on the use of MAbs for therapeutic purposes is making breakthroughs; the earliest likely applications may very well be in the areas such as treatment of specific human cancers. The development of targeted drug delivery systems (“magic bullets”) based on monoclonal antibodies would revolutionize cancer therapy by allowing physicians to destroy cancerous cells selectively while minimizing the damage to normal cells. Current and future advancements in the technologies leading to the large-scale production of needed monoclonal antibodies will have ever increasing diagnostic and therapeutic applications. Further developments in vertebrate-cell culture technology will result in improved processes for enhanced generation of a vast number of products with diagnostic and therapeutic application, and improvement in methodology and process equipment for producing these products will act in tandem to create a massive increase in both quantity and range of applications of vertebrate cells. An improved understanding of cellular metabolism will be crucial in the optimization of vertebrate cell culture

204

SATISH J. PARULEKAR ET AL.

systems and will afford means to better understanding of issues such as the interactive mechanisms of energy production in vertebrate-cell cultures; interrelationships among various dynamic processes such as cell growth, glucose consumption, and lactate production (Zielke ef al., 1984); and roles of glutamine and glutamate in glycolysis. The majority of the molecular biology laboratory techniques work well at the level of micro- or milligrams; however, their scaleup to the level of production quantities becomes a challenging task. On the other hand, many traditional separation techniques used for chemicals need not necessarily be applicable to biological products because of the lower chemical, mechanical, and thermal stabilities of latter. Thus, improved and novel separation techniques will continually be in demand in the near future as new products are generated from vertebrate-cell cultures. Also, the scaleup of biological processes requires and will continue to require a large investment in both time and manpower. Work on the optimization of the design and operation of bioreactors used for cultivation of vertebrate cells is progressing at a rapid pace and will continue to do so due to the fact that biological processes result in rather dilute and often impure product streams. This imposes need for increasing the volumetric productivity of vertebrate-cell cultures and reducing the number of processing steps to make production of the desired metabolites more economical. Better bioreactor designs will be necessary for improving heat and mass transfer in these cultures. Also, work on the development of improved control strategies will continue to expand to facilitate effective operation of vertebrate-cell culture systems.

References Abercrombie, M., and Heaysman, J. E. M. (1954). Exp. Cell Res. 6 , 293. Acton, R. T., and Lynn, J. D. (1977). Adv. Biochem. Eng. 7 , 85. Alitalo, K., Keski-Cja, L., and Vakeri, A (1981). Int. J . Cancer 27, 755-761. Atkinson, B . , and Mavituna, F. (1983). “Biochemical Engineering and Biotechnology Handbook,” pp. 581-669. The Nature Press, New York. Aunins, J. G., Croughan, M. S. , Wang, D. 1. C., and Goldstein, J. M. (1986). Biotechnol. Bioeng. Symp. No. 17, 699-723. Bailey, J . E . , and Ollis, D. F. (1986). In “Biochemical Engineering Fundamentals,” 2nd ed., Chap. 9. McGraw-Hill, New York. Balin, A. K., Goodman, D. B . , Rasmussen, H., and Cristofalo, V. (1976). J . Cell Physiol. 89, 235-249. Barteling, S. J. (1984). Dev. Biol. Stand. 55, 143-147. Berg, G . J. (1985). Dev. B i d . Stand. 60,297-303. Berg, G. J., and Bodeker, B . G . D. (1988). In “Animal Cell Biotechnology,” R. E. Spier, and J. B. Griffiths (eds.), Vol. 3, pp. 321-335. Academic Press, New York/London. Bergenholtz, A . , Halimans. G., and Hanstrom, L. (1977). Anar. Rec. 189, 433-442.

VERTEBRATE CELL CULTURE TECHNOLOGY

205

Bettger, W. J., Boyce, S. T., Wathall, B. J., and Ham, R. G. (1981).Proc. Natl. Acad. Sci. U . S . A . 78, 5588. Birch, J. R. (1980). Deu. Biol. Stand. 46, 21-27. Birch, J. R., and Pirt, S. J. (1970). J . Cell Sci. 7, 661-670. Birch, J . R., Thompson, P. W., Boraston, R.. Oliver, S., and Lambert, K. (1987). In “Plant and Animal Cells: Process Possibilities” (C. Webb, and F. Mavituna, eds.), pp. 162-171. Ellis Horwood, Chichester. Brown, P. C., Figueroa, C., Costello, M. A. C., Oakley, R., and Maciukas, S. M. (1988). In “Animal Cell Biotechnology” (R. E . Spier, J. B. Griffiths, eds.), Vol. 3, pp. 251-262. Academic Press, New York/London. Burrows, M. T. (1910). J . A . M. A . 55, 2057. Butler, M. (1988). In “Animal Cell Biotechnology” (R. E. Spier, and J. B. Griffiths, eds.), Vol. 3, pp. 283-303. Academic Press, New York/London. Butler, M., Hassell, T., and Rowley, A. (1986). In “Process Possibilitiesfor Plant and Animal Cell Culture” (F. Mavituna, and C. Webb, eds.), pp. 64-74. Ellis Horwood, Chichester. Biofechnol. 34, 187-196. Butler, M., and Spier, R. E . (1984). .I. Carrel, A. (1912). J . Exp. Med. 15, 516-528. Castellani, P., Siri, A., Rossellini, C., Infusini, E.. Borsi, L., and Zardi, L. (1986). .I. Cell B i d . 103, I67 I - 1677. CELLCO (1975). British Patent 1,395,291. Chillakuru, R. A., Ryu, D. D. Y., and Yilma, T. (1991). Biotechnol. Prog. 7, 85-92. Chiou, T.-W., Murakami, S., Wang, D. I. C., and Tu, W.-T. (1991). Biofechnol. Bioeng. 37, 755-761. Clarke, J. B., and Spier, R. E. (1980). Arch. Virol. 63, 1-9. Colbere-Garapin, F., and Garapin, A. C. (1985). In “Animal Cell Biotechnology” (R. E. Spier, and J. B. Griffiths, eds.), Vol. 3, pp. 405-429. Academic Press, New York/London. Collins, J. (1982). “Symp. Biotechnol.” Arzneim-Forsch, Bad-Schliersee, Federal Republic of Germany. Crespi, C. L., Imamura, T., Leong, P., Fleischaker, R. J., Brunengraber, H., Thilly, W. G., and Giard, D. J. (1981). Biotechnol. Bioeng. 23, 2673-2689. Crouch, C. F., Fowler, H . W., and Spier, R. E. (1985). J. Chem. Techno/. Biotechnol. B 35, 273-28 I . Curtis, A. S. G., and Seehar, G. M. (1978). Nature (London) 274, 52-53. Curtis, A. S. G., and Varde, M. (1964). J. Natl. Cancer Insf. 33, 15-23. Cysewski, G. R., and Wilke, C. R. (1977). Biotechnol. Bioeng. 19, 1125-1143. Eagle, H. (1955). J. Biol. Chem. 214, 839-853. Eagle, H. (1959). Science 130, 432-437. Eagle, H., Barban, S., Levy, M., and Schulze, H. 0. (1958). J. B i d . Chem. 233, 551-558. Earle, W. R., Schilling, E. L., and Shannon, J. E. (1951). .I Natl. . Cancer Znst. 12, 179-193. Elsdale, T., and Bard, J. (1972). J. Cell Biol. 54, 626-637. Feder, J., and Tolbert, W. R. (1983). Sci. Amer. 248, 36-43. Fischer, A., Astrup, T., Ehreusvard, G., and Oehlenschlager, V. (1948). Proc. Soc. Exp. Biol. Med. 67, 40-46. Fleischaker, R. J. (1982). In “An Experimental Study in the Use of Instrumentation to Analyze Metabolism and Product Formation in Cell Culture.” Ph.D. thesis, Massachusetts Institute of Technology, Cambridge, MA. Fohring, B., Tjia, S. T., and Zenke, W. M. (1980). Proc. Soc. Exp. Biol. Med. 164, 222. Fontages, R., Beaudry, Y., and Matot, G. (1974). Biotechnol. Bioeng. Symp. 4, 859-860. Gey, G. O., Coffman, W. D., and Kubicek, M. T. (1952). Cancer Res. 12, 364-365. Giard, D. J . , Loeb, D. H., Thilly, W. G., Wang, D. 1. C., and Levine, D. W. (1979). Biotechnol. Bioeng. 21, 433-442.

206

SATISH J. PARULEKAR ET AL.

Girard, H. C. (1977). In “Cell Culture and Its Applications,” (R. T. Acton, and J. D. Lynn, eds.), pp. 111-127. Academic Press, New York. Girard, H. L., Sutcu, M., Erdern, H., and Gurhan, I. (1980).Biotechnol. Bioeng. 22,477-493. Glacken, M. W., Fleischaker, R. J., and Sinskey, A. J. (1983a). Tibtech. 1, 4. Glacken, M. W., Fleischaker, R. J., and Sinskey, A. J. (1983b). Ann. N.Y. Acad. Sci. 413, 355-372. Gori, G. B. (1965). Appl. Microbiol. 13, 909-917. Graff, S., and McCarthy, K. S. (1957). Exp. Cell Res. 13, 348-357. Griffiths, J. B. (1970). J . Cell Sci. 6, 739-749. Griffiths, J. B. (1986). Tibtech. 4, 268-272. Griffiths, J. B. (1988). In “Animal Cell Biotechnology,” (R. E. Spier, J. B. Griffiths, eds.), Vol. 3, pp. 179-220. Academic Press, New York/London. Ham, R. G. (1965). Proc. Natl. Acad. Sci. U.S.A. 53, 288-293. Ham, R. G. (1974). I n Vitro 10, 119-129. Handa, A., Emery, A. N., and Spier, R. E. (1987). Deu. B i d . Stand. 66, 241-253. Harris, H., and Watkins, J. F. (1965). Nature (London) 205, 640-646. Harrison, R. G. (1907). Proc. SOC.Exp. Biol. 4, 140-143. Harrison, R. G . (1914). J. Exp. Zool. 17, 521-544. Hayflick, L., and Moorhead, P. S. (1961). Exp. Cell Res. 25, 585-621. Herbert, D. (1976).In “Continuous Culture: Applications and New Fields.” (A. C. R. Dean, ed.), pp. 1-30. SOC.Chem. Ind., London. Himmelfarb, P., Thayer, P. S., and Martin, H. E. (1969). Science 164,555-557. Hines, D. A. (1978). In “Biotechnology,” p. 55. Verlag Chemie, Weinheim. Hirtenstein, M., and Clark, J. (1983).In “Immobilized Cells and Organelles,” (B. Mattiasson, ed.), Vol. 1, pp. 57, 58. CRC Press, Boca Raton, FL. Hofschneider, P. H. (1982). “Symp. Biotechnol.” Arzneim-Forsch, Bad-Schliersee, Federal Republic of Germany. Hopkinson, J. (1983). In “Immobilized Cells and Organelles” (B. Mattiasson, ed.), Vol. 1. CRC Press, Boca Raton, FL. House, W. (1973). In “Tissue Culture: Methods and Applications” (R. F. Kruse, and M. K. Patterson, eds.), p. 338. Academic Press, New York. House, W., Schierer, M., and Maroudas, N. G. (1972). Exp. Cell. Res. 71, 293-296. Hu, W.-S., and Dodge, T. C. (1985). Biotechnol. Prog. 1, 209-215. Hu, W.-S., Meier, J., and Wang, D. I. C. (1985). Biotechnol. Bioeng. 27, 585. Hu, W.-S., and Wang, D. I. C. (1985). Biotechnol. Bioeng. 27, 1466. Hu, W. S., and Wang, D. I. C. (1987). Biotechnol. Bioeng. 30, 548-557. Huber, M. (1974). U.S. Patent 3,785,620. Huber, M., and Schutz, G . (1975a). U.S. Patent 3,871,624. Huber, M., and Schutz, G. (1975b). U.S. Patent 3,871,625. Ivanova, 0. Y., and Margolis, L. B. (1973). Nature (London)242, 200-201. Izuka, M. (1979). British Patent 1,539,263. Jacobs, J. B. (1979). I n “Developments in Biological Standardization” (R. H. Regamey, R. Spier, and F. Horodniceanu, eds.), Vol. 42, pp. 13-18. Karger, Basel. Jensen, M. D. (1981). Biotechnol. Bioeng. 23, 2703-2716. Jensen, M. D., Wallach, D. F. H., and Lin, P. S. (1974). Exp. Cell Res. 84, 271-281. Jo, E.-C., Gu, M. B., Kim, D.-I., and Paik, S. B. (1991). Biotechnol. Bioeng. 38, 247-253. Katinger, H. (1988). In “Animal Cell Biotechnology” (R. E. Spier, and J. B. Griffiths, eds.), Vol. 3, pp. 239-250. Academic Press, New York/London. Katinger, H., and Scheirer, W. (1985). In “Animal Cell Biotechnology” (R. E. Spier, J. B. Griffiths, eds.) Vol. I , pp. 167-194. Academic Press, New York/London. Katinger, H . W. D., and Scheirer, W. (1982). Acta Biotechnol. 2, 3-41.

VERTEBRATE CELL CULTURE TECHNOLOGY

207

Katinger, H. W. D., Scheirer, W., and Kromer. E. (1979). Ger. Chem. Eng. (Eng. Transl.) 2, 3 1-38, Keary, L, and Burton, C. W. (1979). Proc. Biochem. 14, 29. Keay, L. (1975). Biotechnol. Bioeng. 17, 745-764. Kilburn, D. G., and van Wezel, A. L. (1970). J . Gen. Virol. 9, 1-7. Kilburn, D. G., and Webb, F. C. (1968). Biotechnol. Bioeng. 10, 801. Klement, G., Scheirer, W., and Katinger, H. W. D. (1987). Deu. Biol. Stand. 66, 221-226. Kluft, C., van Wezel, A. L., van der Welden, C. A. M., Emeis. J. J., Verheijen, J. H., and Wijngaards, G. (1983). In “Advances in Biotechnological Processes” (A. Mizrahe and A. L. van Wezel, eds.), Vol. 2, pp. 97-110. A. R. Liss, New York. Knazek, R. A.. Gullino, P., Kohler, P. O., and Dedrick, R. (1972). Science 178, 65-67. Kohler, G., and Milstein, C. (1975). Nature (London). 256, 495-497. Kruse, P. F., Jr., Myhr, B. C., Johnson, J. E., and White, P. B. (1963). J. Natl. Cancer Instir. 31, 109-123. Ku, K., Muo, M. J., Delente, J., Wildi, B. S., and Feder, J. (1981). Biotechnol. Bioeng. 23, 79. Lambert, K. J., and Pirt, S. J. (1975). J . Cell Sci. 17, 397-41 I. Lasky, L. A., Dowbenko, D., Simonson, C. C., and Berman, P. W. (1984). BiolTechnology 2, 527. Lehmann, J., Vorlop, J., and Buntemeyer, H. (1988). In “Animal Cell Biotechnology” (R. E. Spier. and J . B. Griffiths, eds.), Vol. 3, pp. 221-237. Academic Press, New Yorkl London. Leighton, J. (1951). J. Natl. Cancer Inst. 12, 545-560. Levine, D. W., Wang, D. 1. C., and Thilly, W. G. (1977a). In “Cell Culture and Its Applications” (R. T. Acton, and J. D. Lynn, eds.), pp. 191-216. Academic Press, New York. Levine, D. W., Wong, J . S., Wang, D. I. C., and Thilly, W. G. (1977b). Somatic Cell. Genet. 3, 149-155. Levine, D. W., Wang, D. 1. C., and Thilly, W. G. (1979). Biotechnol. Bioeng. 21, 821-845. Levintow, L.. and Eagle, H. (1961). Annu. Rev. Biochem. 30, 605-640. Lim, F., and Moss, R. D. (1981a). In “Microencapsulation: Selected Papers from the 47th International Symposium on Microencapsulation” (M. H. Ferguson, ed.), pp. 1-4. American Pharmaceutical Society, Washington, D.C. Lim, F., and Moss, R. D. (1981b). J. Therm. Sci. 70, 351-354. Littlefield, J. W. (1964). Science 145, 709-710. Litwin, J. (1976). In “Proceedings, Gen. Meet. Eur. Soc. Animal Cell Technol., Ist, 1976 Amsterdam” (R. E. Spier, and A. L . van Wezel, eds.), pp. 49-54. Rijks Inst. Volksgezondheid, Bilthoven, The Netherlands. Lydersen, B. K., Pugh, G. G., Paris, M. S., Sharma, B. P., and NOH, L. A. (1985). Biol Technology 3, 63-67. Mann, G. F. (1972). Bol. of. Sanit. Panam. (Engl. ed.) 6, 33-36. Marcipar, A., Henno, P., Lentowojt, E., Roseto, A,, and Broun, G. (1983). Ann. N . Y. Acad. Sci. 413, 416. Margaritis, A,, and Wallace, J. B. (1982). Biotechnol. Bioeng. Symp. No. 12, 147-159. Margaritis, A., and Wallace, J. B. (1984). BiolTechnolog.v May, 447-453. Margaritis, A.. and Wilke, C. R. (1978). Biotechnol. Bioeng. 20, 709-726. Markus, H. Z., and McAleer, W. J. (1981). U.S. Patent 4,301,249. Maroudas, N. G . , (1972). Exp. Cell Res. 74, 337-342. Maroudas, N. G . (1973a). Exp. Cell Res. 81, 104-110. Maroudas, N. G . (1973b). Nature (London) 244, 353-354. Mather, J., Wu, R., and Sato, G. (1981).In “The Growth Requirement of Vertebrate Cells, in Vitro” (C. Waymouth, B. G. Ham, and P. G. Chaple, eds.), pp. 244-257. Cambridge Univ. Press.

208

SATISH J. PARULEKAR ET AL

Mattiasson, B., Maudenius. C. F.. Axelsson. J . P.. Danielsson, B.. and Hagander. P. (1983). Ann. N . Y . Acad. Sci. 413, 193-196. McAleer, W. J., Markus, H. Z., Bailey, F. J., Herman, A. C.. Harder, B. J., Wampler, D. E., Miller, W. J . , Keller, P. M., Buynak, E. B., and Hilleman, M. R. (1987). J . Virol. Method. 7, 263-271. McAleer, W. J . , Schlabach, A. J., and Spier. R. E. (1978). British Patent 1.509.826. McCoy, T. A., Maxwell. M.. and Kruse, P. F. (1959). Proc. Soc. Exp. Biol. Med. 100, 115-1 18. McKeehan, W. L., McKeehan, K. A., and Han, R. G. (1981).I n “The Growth Requirement of Vertebrate Cells in Vitro” (C. Waymouth, B. G. Ham, and P. G. Chaple, eds.) pp. 118-130. Cambridge University Press. Meignier, B., Mangeot, A., and Favre. H. (1981). Deu. B i d . Stund. 46, 246-249. Merk, W. A. M. (1982). Deu. Biol. Stand. 50, 137-141. Michel, M.-L., Sobczuk, E., Malpiece, Y., Tiollais, P., and Streeck, R. (1985). BiolTechnology 3, 561. Mogensen, K. E., Tovey, M. G., Pirt, S. J.. and Mathison, G. E. (1972). J . Gen. Virol. 16, 111-1 14. Molin, O., and Heden, C. G. (1969). Prog. Iwzmirnobiol. Stand. 3, 106-110. Monsanto Co. (1978). British Patent 1,514,906. Montagnon, B. J . , Fanget, B., and Nicolas, B. J . (1981). Deu. B i d . Stand. 47, 55-64. Montagnon, B. J., Vincent-Falquet, J. C., and Fanget, B. (1984). Deu. B i d . Stand. 55, 37. Moscona, A. A., and Moscona. M. H. (1952). J . Anat. 86, 287. Munder, P. G., Modolell, M., and Wallach. D. F. H. (1971). FEBS Lett. 15, 191-195. Murakami, H., Masui. H., Sato, G. H., Sueoka, N.. Chow, T. P., and Kano-Sueoka. T. (1982). Proc. Natl. Acad. Sci. U . S . A . 79, 1158-1 162. Murakami, S., Chiou, T.-W., and Wang, D. I. C. (1991). Biotechnol. Biorng. 37, 762-769. Nahapetian, A. T., Thomas, J. N., and Thilly, W. G. (1986). J. Cell Sti. 81, 65-103. Nicklin, P. M., and House, W. (1976). Biotechnol. Bioeng. 18, 723-727. Nilsson, K., and Mosbach, K. (1980). FEBS Lett. 118, 145-150. Nilsson, K., Scheirer, W.. Merten, 0.-W.. Osberg, L., Liehl. E., Katinger. H. W. D., and Mosbach, K. (1983). Nature (London)302, 629-630. Olsson, L., and Mathe. G. (1982). I n “Recent Results in Cancer Research” (G. Mathe, G. Bonadonna, and S . Salmon, eds.), Vol. 80, pp. 334-337. Springer-Verlag, Berlin. Pakos, V., and Johansson, A. (1983). “Abstr., Proc. Gen. Meet. Eur. SOC.Animal Cell Technol., 6th. 1983, Copenhagen.” Panina, G. F. (1985). I n “Animal Cell Biotechnology” (R. E. Spier, J . B. Griffiths, eds.), Vol. I , pp. 211-242. Academic Press, New York/London. Parisius, J. L. E. W., Cucakovich, N. B., and Maccorine, G. M. G. (1974). British Patent 1,358,321. Parulekar, S. J., and Lim, H. C. (1985). Adu. Biochem. Bioeng.lBiotechnol. 32, 207-258. Peehl, D. M., and Stanbridge, E. J. (1981). Proc. Natl. Acad. Sci. U . S . A . 78, 3053-3057. Perry, S. D., (1987). M.S. thesis, Massachusetts Institute of Technology, Cambridge, MA. Pharmacia Fine Chemicals (1981). “Microcarrier Cell Culture: Principles and Methods.” Pharmacia Fine Chemicals, AB, Upsalla, Sweden. Pirt, S. J., and Callow, D. S. (1964). Exp. Cell Res. 33, 413-421. Pollard, R., and Khosrovi, B. (1978). Proc. Biochem. (3,31-37. Pullen, K. F., Johnson, M. D., Phillips, A. W., Ball. G. D., and Finter, N . B. (1985). Deu. B i d . Stand. 60, 175. Radlett, P. J., Pay, T. W. F., and Garland. A. J . M. (1985). Deu. B i d . Stand. 60, 163. Radlett, P. J . , Telling, R. C., Stone, C. J., and Whiteside, J. P. (1971). Appl. Microbiol. 22, 534-537.

VERTEBRATE CELL CULTURE TECHNOLOGY

209

Ratafia, M. (1987). BiolTechnology 5, 692-694. Reuveny, S. (1983). Adv. Biotechno/. Processes 2, 1-32. Reuveny, S . , Bino. T., Rosenberg. H.. Traub, A.. and Mizrahi, A. (1980).Deu. Biol. Stund. 46,281-288. Ritz, J., and Schlossman. S. F. (1982). Blood 59, 1 - 1 1 . Rosenberg, M. D. (1963). Science 139, 41 1-412. Rosevear, A., and Lambe, C. (1988) In “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths, eds.), Vol. 3, pp. 397-440. Academic Press, New YorklLondon. Rous, P., and Jones, F. S. (1916). J . Exp. Med. 23, 549. Roux, W. (1885). 2. Biol. 21, 411. Rovensky, Y. A,, Slavnaja. I. L., and Vasiliev, J. M. (1971). Exp. Cell Res. 65, 193-201. Runstadler, P. W . , Jr., and Cernek, S. R. (1988). In “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths. eds.), Vol. 3, pp. 305-320. Academic Press, New York/London. Sanford, K. K., Earle, W. R., and Evans, V. J. (1951). J . Nntl. Cuncer Inst. 11, 773-795. Santero, G. G. (1972). Biotechnol. Bioeng. 14, 753-775. Scattegood, E. M., Schlabach, A. J., McAleer, W. J., and Hilleman, M. R. (1983). Ann. N . Y . Acud. Sci. 413, 332. Scheirer, W. (1988). In “Animal Cell Biotechnology” (R. E. Spier and J . B. Griffiths, eds.), Vol. 3, pp. 263-281. Academic Press, New York/London. Schleicher, J. B. (1973). I n ‘‘Tissue Culture: Methods and Applications.” (R. F. Kruse and M. K. Patterson, eds.), p. 333. Academic Press. New York. Schonherr, 0. T., and van Gelder. P. T. J. A. (1988). In “Animal Cell Biotechnology” (R. E . Spier and J. B. Griffiths, eds.). Vol. 3, pp. 337-355. Academic Press, New Yorki London. Secher, D., and Burke, D. C. (1980). Nuture (London) 285, 446. Shamel, R. E., and Chow, J. J. (1989). Chem. Eng. Progr. 85(12), 33-37. Skoda, R., and Pakos, W. (1977). Belgian Patent 854,898. Sorieul, S., and Ephrussi, B. (1961). Nature (London) 190,653-654. Spier, R. E . (1976). Biotechnol. Bioeng. 18, 659-667. Spier, R. E. (1980). Adu. Biochem. Eng. 14, 119-162. Spier, R. E. (1982). .IChem. . Techno/. Biotechno/. 23, 304. Spier, R. E. (1985). In “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths, eds.), Vol. I , pp. 243-265. Academic Press, New YorklLondon. Spier, R. E., and Griffiths, J. B. (1982). Deu. Biol. Stund. 55, 81-92. Spier, R. E., and Griffiths, J. B. (1985). In “Animal Cell Biotechnology” (R. E. Spier and J . B. Griffiths, eds.), Vol. I , pp. 3-1 1. Academic Press, New York/London. Spier, R. E., and Horaud, F. (1985). In “Animal Cell Biotechnology” (R. E. Spier and J . B. Griffiths, eds.), Vol. 2, pp. 431-458. Academic Press, New York/London. Staeheheln, T.. Hobbs, D. S., Kung, H. F., and Pestka, S. (1981). In “Methods in Enzymology” (S. Pestka, ed.), Vol. 78, pp. 505-512. Academic Press, New York. Stoker, M., and Rubin, H. (1967). Nuture (London), 215, 171-172. Taylor, G. W., Kondig, J. P., Nagle, S. C., Jr., and Higuchi, K. (1971). Appl. Microbiol. 21, 928-933. Taylor, W. G., and Evans, V. J . (1975). Biotechnol. Bioeng. 17, 1847-1851. Thayer, P. S. (1973). In ‘‘Tissue Cultures: Methods and Applications” (P. F. Kruse and M. K. Patterson, Jr., eds.), pp. 345-351. Academic Press, New York. Thilly, W. G., and Levine, D. W. (1979). In “Methods in Enzymology” (W. B. Jakoby and Z. H . Pastan, eds.), Vol. 58, pp. 184-194. Academic Press. New York. Thornton, B., McEntee, I. D., and Griffiths, B. (1985). Deu. Biol. Stund. 60, 475. Tolbert, W. R., and Feder, J. (1983). Annu. Rep. Ferm. Proc. 6 , 35. Tolbert, W. R., Feder, J., and Kimes, R. C. (1981). In Vitro 17, 885-890.

210

SATISH J. PARULEKAR ET AL.

Tolbert, W. R., Hitt, M. M., and Feder, J. (1980). In Vitro 16, 486-490. Tolbert, W. R., Lewis, C., White, P. J., and Feder, J. (1985). I n “Large Scale Mammalian Cell Culture” ( J . Feder and W. P. Tolbert, eds.), pp. 97-123. Academic Press, Orlando. Tolbert, W. R., Srigley, W. R., and Prior, C. P. (1988). I n “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths, eds.), Vol. 3, pp. 373-393. Academic Press, New Yorkl London. Tovey, M. G. (1985). 1n“Animal Cell Biotechnology” (R. E., Spier and J. B. Griffiths, eds.), Vol. 1 , pp. 195-210. Academic Press, New York/London. Tovey. M. G., Mathison, G. E., and Pirt, S. J. (1973). J . Gen. Virol. 20, 29-35. Tripathi, S. C. (1984). 1n“Symposium on Regulation of Cell Proliferation. Proceedings of Eighth All-India Cell Biology Conference,” S4. M. K. University, Madurai, India. Tripathi, S. C. (1989a). I n Vitro 25, 980-986. Tripathi, S. C. (1989b). B i d . Cell 67, 351-353. Tripathi, S. C., and Curtis, A. S. G. (1982). 1n“Proceedings of Michael Abercrombie Memorial International Meeting of British Society of Cell Biology,” p. 22. University College, London, UK. Tripathi, S. C., and Fygensen, M. (1992). In Vitro Cell Dev. Biol. Submitted. Tripathi, S . C., and Kerr, J. (1989). Tissue Cell 21, 747-752. Tripathi, S. C., and McGrath, M. F. (1989). Biotechniques 7, 454-455. Tripathi, S. C., and Vosseller, K. A. (1991). Co(d Spring Harbor Symp. Quant. Biol. 56, 175. Tyo, M. A., Bulbulian, B. J., Meken, B. Z., and Murphy, T. J. (1988). In “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths, eds.), Vol. 3, pp. 357-371. Academic Press, New York/London. van Brunt, J. (1986). BiolTechnology 4, 505-510. van Hemert, P., Kilburn, D. G., and van Wezel, A. L. (1969). Biotechnol. Bioeng. 11, 875-885. van Wezel, A. L. (1967). Nature (London) 216, 64-66. van Wezel, A. L . (1972). Prog. Immunobiol. Stand. 5 , 187. van Wezel, A. L. (1973). I n “Tissue Culture: Methods and Applications” (R. F. Kruse and M. K. Patterson, eds.), p. 372. Academic Press, New York. van Wezel, A. L. (1982). J . C h e m . Techno/. Biotechnol. 32, 318-323. van Wezel, A. L. (1985). In “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths, eds.), Vol. 1, pp. 265-282. Academic Press, New York/London. van Wezel, A. L., van Steenis, G., Hannik, C. A., and Cohen, H. (1978). Deu. Biol. Stand. 41, 159-168. van Wezel, A. L., and van der Velden-de Groot, C. A. M. (1978). Process Biochem. 13,6-8. Varani, J., Dame, M., Beal, T. F., and Wass, J. A. (1983).Biorechnol. Bioeng.25,1359-1372. Vitetta, E. S ., Krolick, K. A., Miyama-Inaba, M., Cushley, W., and Uhr, J. W. (1983). Science 219, 644. Weiss, P. A. (1961). Exp. Cell R e s . (Suppl) 8, 260-281. Weiss, P. A. (1962). In “Biological Interactions on Normal and Neoplastic Growth” (M. J. Brennan and W. L. Simpson, eds.), pp. 3-20. Little, Brown, Boston. Weiss, R. E., and Schleicher, J. B. (1968). Biotechnol. Bioeng. 10, 601-616. Whiteside, J. P. (1983). “Investigations on Anchorage-Dependent Animal Cells Grown in Packed Bed Propagators.” Ph.D. thesis, University of Surrey. Whiteside, J. P., and Spier, R. E. (1981). Biorechnol. Bioeng. 23, 551-565. Whiteside, J. P.. Whiting, B. R., and Spier, R. E. (1979). Deu. B i d . Stand. 42, 113-119. Wiemann, M. C., Ball, E. D., Fanger, M. W., McIntyre, 0. R., Bernier, G., and Calabresi, P. (1983). Clin. R e s . 31, 511A. Witkowski, J. A. (1979). Med. Hist. 23, 279-296.

VERTEBRATE CELL CULTURE TECHNOLOGY

211

Wohler, W., Rudiger, H . W., and Passargem, E. (1972). Exp. Cell Res. 74, 571-573. Zanetti, R. (1984). Chem. Eng. 91, 18-21. Zielke, H . R., Zielke, C. L., and Ozand, P. T. (1984). Fed. Proc. 43, 121-125. Zwerner, R. K . , Runyan, C., Cox, R . M.,Lynn, D., and Acton, R. T. (1975). Biorechnol. Bioeng. 17,629-657.

This Page Intentionally Left Blank

Volker Schmid Institute of Zoology, Rheinsprung 9, CH-405 I Basel, Switzerland, and Friday Harbor Laboratories, Friday Harbor, Washington 98250

1. Introduction

The question of how stable differential gene expression is achieved in ontogeny has long been a field of interest in developmental biology. Is stability a consequence of loss of genomic information (Tobler, 1986) or of irreversible alterations in the genome (Caplan and Ordahl, 1978)? Stability in gene expression can be regarded as a consequence of a variety of ontogenic processes to which the genome, cytoplasmic factors, cell cycle events (Freeman, 1981; Jeffery, 1985; Edger and Schubiger, 1986), components of the cytoskeleton (Ben-Ze’ev, 1980; Zanetti and Salursh, 1984; Franke, 1987), and the cellular (Moscona et al., 1983) and extracellular environment (Trelstad, 1984; Acheson et al., 1986; Ekblom, 1989; Timpl, 1989; Ingber and Folkman, 1989b;Anderson, 1990) contribute. In the lattercase the influence is exerted by molecules interacting with transmembrane glycoproteins which, directly or indirectly (Ingber and Folkman, 1989a),can transfer the information from the extracellular environment to the cytoplasm and the nucleus (Watson, 1991). Therefore, cellular commitment and the stability of a functionally specialized cell can be viewed as being the consequence of its individual ontogenic history. Despite great progress achieved in understanding the control of gene expression it is still unclear how the different extracellular and cellular elements cooperate to establish and maintain functional stability of cells. Investigations on this problem have mostly concentrated on the question whether functional stability and cellular commitment can be changed, and if so, to what extent. In plants there is clear evidence that fully specialized adult cells can be totipotent (Vasil and Vasil, 1972; Meins, 1986, 1989; Osborne and McManus, 1986; Okada, 1991); cellular commitment is therefore fully reversible and can extend even to the full morphogenetic potential necessary to shape the organism. In animals, however, gene expression in committed and differentiated cells seems in general to be stable. This statement, howInrrmofkmul Rruies, of Ci,roloay. V d . 142

213

Copyright Q 1992 b y Academic Press. Inc. A l l rights of reproduction in any form reserved.

214

VOLKER SCHMID

ever, does not fully refer to the nuclei of somatic cells. Nuclear transplantation studies have indeed clearly demonstrated that the nuclei of adult somatic cells are still able to express pluripotency when transplanted into oocyte cytoplasm (DiBerardino et al., 1984; DiBerardino, 1988). Success in these studies is, however, rare and it is assumed that this could be the consequence of changes in the chromatin structure of the implanted nucleus which prevent normal cooperation between the nucleus and the recipient cytoplasm (DiBerardino, 1980). Results are greatly improved when the implanted nuclei are conditioned prior to implantation by serial cloning. One has to conclude that the full repertoire of genes is still present, but reactivation in the foreign oocyte cytoplasm becomes increasingly difficult the further advanced in development the implanted nucleus is. The low number of transplantation successes with nuclei of advanced donor stages and observations on in uitro cultures has led to the hypothesis that under “normal” conditions genes are progressively and finally irreversibly repressed (Caplan and Ordahl, 1978; Bennett, 1983). There are, however, an increasing number of observations that are clearly incompatible with this view. Activation of silent genes through DNA-binding proteins (Topscott et al., 1988; Lassar et al., 1989), experiments with heterocaryons (DiBerardino et al., 1984; Okada, 1991), and studies on transdetermination and transdifferentiation, has proven that stability in most cases is not aconsequence of loss or irreversible inactivation of genes. In the case of transdetermination and transdifferentiation nuclei of somatic, specialized cells in uiuoand in uitro can change their commitment without the necessity ofbeing exposed to embryonic cytoplasm. This change in commitment is called transdetermitation in embryonic cells and transdifferentiation in specialized somatic cells. In transdifferentiation processes the initial events for gene activation are therefore of extracellular origin whereas in nuclear transplants these events seem to reside in the recipient cytoplasm. In recent years several reviews on transdifferentiation have been published (Pritchard, 1986; Yamada, 1989). The most comprehensive review, however, covering the historical background as well as the different systems or the molecular mechanisms was published by Okada( 1991). Therefore the emphasis of this review is based on an in uitro transdifferentiation system that was established using isolated medusa tissues of differentiated cells.

II. The Concept of Transdifferentiation A. Terminology

Transdifferentiation can be defined as a switch (or reprogramming or rechanneling) of cells that have already expressed specific differentiated

TRANSDIFFERENTIATION IN MEDUSAE

215

traits into another cell type distinguished from the original cells by a set of phenotypic characters (Okada, 1991). In sensu strict0 this term can be used only when committed cells expressing in part or fully the differentiation characters undergo a change in commitment and gene expression. In all known cases of true transdifferentiation the newly aquired state is stable and a switch to the original cell type has never been reported. Transdifferentiation seems therefore to be an unidirectional process (Okada, 1991). The term transdifferentiation is actually the most widely used, however, for historical reasons, the following terms are still present in the literature and can give rise for confusion. Metaplasia, cellular metaplasia, cell type conversion, cell transformation, or modulation are terms that occasionally are used to denote the change from one differentiated cell type (state) into another. Occasionally these terms are used synonymously with transdifferentiation; often, however, they are used simply to describe a change in the phenotype of cells or tissues. For a better insight into this terminology problem the reader should refer to the review of Okada (1991). The term transdetermination basically refers to a change of commitment in undifferentiated cells. Transdetermination therefore most likely occurs in ontogeny and has been convincingly documented in imaginal discs of Drosophila (Hadorn, 1966; Gehring, 1967). Since transdetermination and transdifferentiation have the same roots and only differ in the expression of the committed state it is often difficult to distinguish these two terms.

B. Transdifferentiation in Plants and Animals

1. Plants Totipotency of plant cells cultured in uitro is a well-established fact. Regeneration of an entire new plant from fragments of carrot phloem was reported by Steward e f al. (1964). Later it was demonstrated that even isolated and cultured single cells have this potential (Vasil and Vasil, 1972). Because determination and differentiation on the molecular basis seem to be different in plants than in animals, this totipotency does not necessarily involve transdifferentiation. Contrary to many animal tissues, determination in plants could arise primarily at the tissue and organ level (Henshaw et al., 1982). In plants, only a small percentage of cells seem to progress to an advanced state of differentiation so that the living tissue retains a greater capability for future developmental changes. Within an individual explant, only a very low proportion of the cells, approximately exhibit totipotency (Meins, 1982). Liberation of these cells from the constraints that operate in uiuo allows them to express their residual

216

VOLKER SCHMID

flexibility (Osborne and McManus, 1986). The question therefore is whether stably specialized cells can also exibit totipotency. There are evidently examples that stable states arise during development in plants and that some of these states persist in populations of dividing cells (Meins, 1986). Detailed studies of habituation for auxin and cytokinin have shown that some determined states can be inherited by individual cells and that these cells undergo a process similar to transdetermination (Meins, 1989). Furthermore, isolated cells from the mesophyll were reported to transdifferentiate without mitosis into tracheary elements (Fukuda and Komamine, 1985). 2. Animals

The following will briefly summarize the most used in uivo and in v i m systems in animals. For a more detailed review the reader should refer to the book on transdifferentiation by Okada (1991). All well-known cases of transdifferentiation in uiuo or in uitro have strong ties to regeneration processes. The ability to use diffentiated cells to bring about regeneration seems to be advantageous for the organism.

In Vivo Systems In most of the well-regenerating invertebrates like sponges, coelenterates, planaria, and annelida, transdifferentiation most likely occurs frequently, but only in a few cases has sufficient evidence been published to support the existence of this event. In sponges multifunctionality of cell types in both the gastral and the dermal layer is well known and in some species transdifferentiation-like processes are a part of normal life (Bergquist, 1978). Apart from the medusa system described in this article, transdifferentiation was investigated in the freshwater polyp Hydra. Successful attempts to regenerate an entire polyp from isolated tissue layers have been reported for the endoderm (Normandin, 1960; Haynes and Burnett, 1963; Davis et al., 1966) and ectoderm of a different species by Lowell and Burnett (1969). Similar experiments in my own laboratory (not published) and by others (Smid, 1984) with the same or different species were not successful. Experiments with the marine hydroid polyp Podocoryne carnea have readily shown polyp regeneration from isolated ectoderm, but not from the endoderm (Achermann, 1980). Changes in differentiation phenotypes, however, occur constantly in epitheliomuscle cells and some cells of the nervous system in the Hydra polyp (Bode, 1992). The changes in nerve cell phenotype occur when cells or tissues are displaced along the body column.

a.

TRANSDIFFERENTIATION IN MEDUSAE

21 7

Planarians are another classical example used for regeneration experiments and the question whether regeneration blastema stems from neoblasts, dedifferentiated specialized cells, or both has not been answered convincingly (Okada, 1991). Using a karyological marker Gremigni and his collaborators (Gremigni, 1988) showed that premeiotic germ cells can contribute to the blastema and are capable of differentiating into some foreign cell types. With regard to the original cell type the term transdifferentiation, however, is not appropriate for characterizing this interesting event. Although the differentiation potential of neoblasts has been investigated in a series of elegant experiments (Bagufia and Romero, 1981; Baguna er al., 1989) the transdifferentiation question is still not answered convincingly for planarians. In arthropods the cells responsible for silk production in the labial gland of the silkworm undergo a transdifferentiational change during metamorphosis and become specialized for the active transport of salts in the moth (Kafatos and Williams, 1964; Kafatos, 1972). In vertebrates, transdifferentiation problems in the amphibian lens and limb regeneration systems have long been the subject of intensive investigation. Regeneration of the lens by transdifferentiation of melanocytes from the dorsal iris is a fact and the different steps have been investigated in great detail (Colucci, 1891; Wolff, 1895; Yamada, 1989). The contribution of transdifferention to limb regeneration is still controversial. The use of kariotypic markers (Steen, 1968; Namewirth, 1974) and monoclonal antibodies (mAbs) (Kinter and Brockes, 1984) clearly demonstrates that the blastema cells are heterogenous. The transdifferentiation potential of vertebrate cells participating in regeneration is limited; e.g., dermal connective tissue can form all mesodermal tissue except muscle or muscle can transform into cartilage (Okada, 1991). In very detailed studies it was shown that in the pancreas of mammals destruction of the acinar tissues by different drugs and initiation of regeneration by various diets induced ectopic transdifferentiation of both the ductal and the periductal interstitial cells into pancreatic hepatocytes (Rao et al., 1990). In uiuo studies in general have one main problem: How can the cell type under investigation be labeled reliably and its developmental fate followed through the possible transdifferentiation process? To avoid this problem, tissue and cell cultures based on only one cell type have been established.

b. In Vifro Systems Due to difficulties in culturing invertebrate cells most in vitro transdifferentiation systems use vertebrate cells. The first clonal transdifferentiation system was established by Eguchi and Okada (1973) using retinal pigment epithelium (RPE) of chicken embryos. The RPE gradually transdifferentiate to lentoid cells, expressing typical lens crystallins. Recently, similar observations were reported from cultured

218

VOLKER SCHMID

RPE of adult human eyes (Eguchi, 1988). The culture conditions have been elegantly worked out and today several transdifferentiation steps can be monitored at will by changing the culture conditions (Itoh and Eguchi, 1986). In this system, molecular analysis of transdifferentiation is most advanced; the lens-specific genes are isolated and characterized and the expression patterns have been analyzed both in uitro (Agata and Eguchi, 1984; Eguchi, 1988; Mochii et al., 1988a,b)and in uiuo (Agata et al., 1983; Okada, 1983; Kondoh et al., 1987; Inoue et al., 1989). Transdifferentiation from pigment epithelial cells from other sources into neuronal cells (Okada, 1991) and among different subclasses of chromatophores (Ide, 1986) emphasize the great flexibility of this cell type. Another example of transdifferentiation among derivatives of neural crest cells is the transformation of chromaffin cells, an endocrine cell type in the adrenal gland, into cholinergic neuronal cells (Unsicker et al., 1978; Anderson, 1989). Finally, an elegant system for testing the transdifferentiation/transdetermination potential of minced muscle and other tissue should be mentioned. Demineralized bone matrix seems to contain a factor that is identical to human transforming growth factor (TGF-P) (Okada, 1991). When tissues are cultured on this matrix, production of cartilage is induced (Nathanson, 1986). A detailed analysis of the responding cell types in the heterogenous explant and in pure populations of cloned muscle cells revealed that the bone matrix factor induces committed myoblasts and authentic fibroblasts to transform in cartilage-producing chondrocytes (Nathanson et al., 1978).

111. Transdifferentiation in Hydromedusae A. The Animal and Its Tissue Organization

Medusae (jellyfish) belong to the Cnidaria, a group of animals that lack a true mesoderm and are essentially classified as bilayered animals (Fig. 1). However, they have developed real organs and highly specialized tissues, nerve nets (Mackie, 1980), and a variety of sense organs, e.g., a visual system (Weber, 1981) or mechanical receptors (Tardent and Schmid, 1972). Additionally, they have two properties that have been of crucial importance for the establishment of an experimental transdifferentiation system; first, many species are able to regenerate most of their body parts (Schmid, 1974) and second, the swimming organ (the umbrella or bell) is composed of three tissue layers, of which two consist of only one welldifferentiated cell type (Figs. 1-7; Schmid, 1972). Both cell types, the striated muscle and the endoderm of the subumbrella, were used for our transdifferentiation studies.

isolation of tissues medusa

i

e ex

e

ex

collagenase

'I

I

distilled watei

Dt

4

6 , . , ~ , n

f

ld

regenerated manubrium 8d

nc 1

ectoderm: smooth muscle interstitial cells gland cells nematoblasts-cytes gametes.

endoderm : digestive cells interstitial cells gland cells nematoblasts gametes. secretory cells

FIG. 1 Schematic drawing of the in uitro system used to regenerate manubria or tentacles. am, outer mesogloea; d, days; dam. degenerating exumbrella and outer mesogloea; e, endoderm of the subumbrellar plate; ex, exumbrella; f, flagella; im, inner mesogloea; ma, manubrium; nc, nematocytes; st, striated muscle; t, tentacles; v, velum; y cells correspond to (FM)RF-amide-positive cells; * gametes observed in the grafting experiments (after Schmid and Alder, 1986).

220

VOLKER SCHMID

1. The Striated Muscle Tissue

In situ striated muscle of all the different medusa species investigated consists of mononucleated cells that are out of cell cycle (Figs. 2-7; Schmid, 1972, 1988), presumably in Go (resting stage). The cells form a thin, monolayered, epithelial tissue that lines the subumbrellar cavity and adheres to the inner mesogloea, an extracellular matrix (ECM) that is part of the bell. Although the gross morphology is the same in all the species investigated, the bundles of myofibers vary in thickness, probably in direct relation to the swimming behavior of the animals.

2. The Endoderm of the Subumbrella The subumbrellar plate endoderm forms a continuous monolayer beneath the straited muscle of the medusae (Figs. 1, 2-4). The cells are extremely flattened and of hexagonal shape and spread between inner and outer mesogloea. No DNA replication was observed in this type of endoderm in hatched medusae (Fig. 6; Schmid, 1972). Arrays of septate junctions are rarely observed in the endodermal layer. However, they are readily apparent in lanthanum-impregnated material (Weber and Schmid, 1984). Gap junctions are common in the endodermal lamella (Weber and Schmid, 1984). The endoderrn cells are ultrastructurally poorly differentiated and their function is unclear. 3. The Extracellular Matrix (Mesogloea) Within the last two decades it has become clear that the ECM not only serves as a mechanical support to maintain tissue shape and integrity but also carries information for the proper function of organ systems for cellular determination, differentiation, and morphogenesis. In our search for the mechanisms responsible for maintenance of the differentiated state of straited muscle tissue of jellyfish, the structural organization of the ECM (mesogloea) to which this tissue adheres is of special interest. All methods that successfully alter the state of cellular differentiation (transdifferentiation) in the tissue, either directly (collagenase, pronase, hyaluronidase) or indirectly (tumor promoters, diacylglycerol, lectins, mAbs), seem to affect the structural integrity of the ECM-ligand-receptor complex. The great bulk of the bell of hydromedusae consists of transparent extracellular matrix material which is separated by the intervening subumbrellar plate endoderm into an outer and inner portion (Figs. 1, 2-4). The mesogloea does not contain cells. In contrast to other organisms, the ECM of medusae is their most prominent structure. It essentially shapes the body of the medusa and serves different functions, e.g., as a substratum

FIG. 2 Medusa of Podocoryne carnea after being treated with collagenase for 4 hr. The mononucleated striated muscle (st) has separated from the subumbrellar plate endoderm (sp). ma, manubrium; te, tentacles, Bar, 100 p m (from Schmid and Alder, 1984). FIG. 3 Enlarged view of a medusa treated with collagenase and stained with DAPI. The striated muscle (st)has detached from the subumbrellar plate endoderm (sp). ex, exumbrella. Darkfield micrograph. Bar, 50 pm. (from Schmid and Alder, 1984). FIG.4 Mirror image of Fig. 3. Nuclei (n) of exumbrella (ex) and subumbrellar plate endoderm (sp) stain brightly with DAPI, and nuclei of striated muscle (st) do not stain. Fluorescence micrograph. FIG. 5 Fragment of isolated striated muscle stained with DAPI after isolation. Nuclei and cytoplasm of contaminating cells of the subumbrellar plate endoderm stain brightly (two are encircled). Fluorescence micrograph. Bar, 100 p m (from Schmid and Alder, 1984). FIG. 6 Distribution of S-phase nuclei in a freshly liberated medusa of P . cornea. Replicating nuclei are made visible with the thymidine analog bromodeoxyuridine (BrdU, I-hr pulse). The bell of the medusa is completely free of DNA-replicating nuclei. Arrows indicate sites of DNA replication. Bar, 100 pm. (courtesy of Ch. Baader). FIG. 7 (a) En face micrograph showing the striated muscle cells of the subumbrellar (bell) surface of P . carnea after silver nitrate incubation. Bar, 100 pm. (courtesy of Ch. Weber). (b) Striated muscle cells in a whole-mount preparation of the medusa P . carnea. The mAb 6AIO stains contractile elements of striated and smooth muscle cells. Bar, 10 pm. (courtesy of B. Rupp). (c) Like (b), demonstrating the smooth muscle (sm) over the radial canal (arrowheads) Bar, 50 pm. (courtesy of B. Rupp).

222

VOLKER SCHMID

for cell movement (Schmid et al., 1984; Schmid and Bally, 1988), the control of morphogenetic processes (Schmid et al., 1976;Schmid, 1978), and the regulation of buoyancy (Denton, 1963;Chapman, 1966;Mackay, 1969).The outer mesogloea of hydrozoan jellyfish was found to contain a species-specific meshwork of striated fibers of different diameters ( Weber and Schmid, 1985). In the mesogloea, molecules that exhibit several features of well-known vertebrate ECM were identified: a laminin-like molecule that appears cross-shaped on electron micrographs (Beck et al., 1989),a fibronectin-like molecule (Schlage, 1988)(both detectable by their immunoreactivity at the exumbrella side), and a species-specific collagen consisting of three different a-chains, of which at least two can be decorated with Con A (Schmid et a f . ,1991).The a-chains are linked by disulfide bridges. Acetic acid extraction of the mesogloea and subsequent salt precipitation yields fibrils that appear banded in the electron microscope and support species-specific adhesion and spreading of isolated tissue. These precipitated fibrils are mainly composed of disulfide-linked collagen (Schmid and Bally, 1988).

6. The Isolation of Tissues Consisting of Only One Differentiated Cell Type

Because contamination of transdifferentiating tissues with other cell types is a problem that often raises controversial arguments, the isolation procedure for the striated muscle and the subumbrellar plate endoderm is outlined in detail. A precise isolation protocol is given in Schmid et al. (1982).

1. Mononucleated Striated Muscle Freshly hatched medusae of Podocoryne carnea that were reared in the laboratory (Schmid, 1979), or medusae of Polyorchis sp., Sarsia sp., or Stomotoca atra collected in plankton (Schmid, 1978) were stained with (4’-6-diamidino-2-phenylindole.2 HCI, (DAPI; Serva, Heidelberg, Germany), 0.05 pg/ml, for 60 min. The DAPI binds to AT-rich regions of DNA and when applied in uiuo for 1 hr brightly stains the nuclei of all cell types except those of striated muscle (Figs. 3-5). The stained medusae were then washed and incubated in collagenase (Millipore Corp.,No. 5275, 150 units/ml culture medium) for 6-8 hr at 26°C. This treatment digests the inner mesogloea and separates the striated muscle from the subumbrellar plate endoderm (Fig. 1; Schmid et al., 1982). Interradial fragments were excised with microscissors and watchmaker’s forceps, the peripheral part with the velum and the circular canal was removed, and the loose striated muscle was gently pulled off. Immediately after isolation the

TRANSDIFFERENTIATION IN MEDUSAE

223

striated muscle was examined by fluorescence microscopy for contamination by DAPI-stained cells of the subumbrellar plate endoderm (Figs. 3-5). Alternatively, striated muscle can be isolated mechanically without collagenase treatment by the same dissection procedure; this, however, is more difficult to do and many more isolated fragments are contaminated by endodermal cells.

2. Subumbrellar Plate Endoderm After the striated muscle is excised, the remainder of the extirpated interradial medusa fragment consists of the subumbrellar plate endoderm, the bulky outer mesogloea (extracellular matrix), and the exumbrellar tissue (Fig. 1). If cultured, this fragment rounds up within the next 12 hr, and the subumbrellar plate endoderm becomes concentrated in the center and is covered by the outer mesogloea, over which the exumbrella cells form a continuous tissue layer. These rounded-up fragments are rinsed for about 10 sec with distilled water and transferred back to culture medium (Fig. 1). The osmotic shock kills all the peripherally arranged exumbrella cells, whereas most of the endoderm, protected by the outer mesogloea, survives (Schmid et al., 1982). The remnants of the dead exumbrella cells are rinsed off by sucking the fragment repeatedly through a pipet, and the endoderm, together with remaining portions of the outer mesogloea, is transferred to culture medium at 12°C for further observation.

C. The Transdifferentiation Potential of Isolated Tissues

1. Striated Muscle Tissue a . Stability of the Differentiated State Striated muscle that has been mechanically isolated contains variable amounts of inner mesogloea. This can be observed directly in the large medusa species (Polyorchis, Stornotoca) where the inner mesogloea is sufficiently thick. In the small species Podocoryne carnea the adhering ECM can be stained by ECMspecific monoclonal antibodies (see later in Figs. 32,33). When such fragments are cultured, the muscle tissue covers the mesogloea within a few hours. In the course of the next 2-3 days the volume of the isolated fragments increases, resulting in typical swollen isolates. DNA synthesis, flagellum formation, or the disappearance of the cross-striated myofibers is rarely observed and cellular commitment remains fairly stable (Schmid, 1978; Schmid and Alder, 1986). Since these isolates demonstrate the contractions characteristic for striated muscle tissue, the cells must be functional in the original sense. Transdifferentiation to new cell types

224

VOLKER SCHMID

rarely occurs and no medusa parts are regenerated. The isolates can live for 1-2 months until they finally fall apart.

b. Destabilization Striated muscle tissue isolated with collagenase and post-treated with activating agents is in general not able to maintain the functional state of a mononucleated cross-striated muscle cell. In this case, the isolated muscle tissue shrinks shortly after activation treatment and forms an aggregate-like structure (Schmid, 1978; Schmid and Alder, 1984). By this process the flat spindle-shaped muscle cells, originally arranged as a single-layered epithelium, become cuboidal. Two to 3 days after isolation, the bundles of striated myofibers may eventually disappear completely, although there is considerable variation between different fragments and even within cells of the same fragment in this respect (Figs. 8, 9). The presence of bundles of striated myofibers seems to have no influence on the ability of the cells to transdifferentiate and/or to participate in regeneration. Earliest DNA synthesis starts 24 hr postisolation followed by mitoses (after 3 days). De nouo formation of flagella (Fig. 8) in all the peripheral cells of the isolate is first observed at 48 hr (Schmid, 1974). In the following sections, isolated striated muscle cells exhibiting these features will be referred to as destabilized.

c. Transdifferentiation to New Cell Types The following observations refer exclusively to tissues isolated from P. carnea. After mitosis is completed (between 2 and 4 days after isolation) the destabilized isolates consist entirely of two new cell types (Schmid and Alder, 1984) present in about equivalent proportions. One cell type was identified as a smooth muscle cell, the other as a sensory nerve type of cell that shows strong positive immunoreactivity to FMRF-amide/or RF-amide (Figs. 8-12). The regional distribution of the nerve cells stained with the antibody against RF-amide is the same as that defined by the antibody against FMRFamide. Most data presented below were obtained with the antibody against RF-amide. a . Formation of Smooth Muscle-like Cell Type Smooth muscle cells (Fig. 8) are characterized by vacuoles filled with electron-dense material that can be found beneath the surfaces of these large cells, which are located at the periphery of the isolate (Fig. 8). These cells are flagellated, whereas striated muscle cells do not have flagella, neither in uiuo nor in uirro (Schmid, 1972; Schmid er al., 1982). In addition, this cell type has a few very large and “empty” vacuoles in the mid-part of the cell body (Fig. 8). With increasing age they progressively contain fewer fragments of striated myofilaments but instead develop at their base an arrangement of contractile filaments characteristic of the smooth muscle cells of the animal (Figs. 8 (insert), 9).

TRANSDIFFERENTIATION IN MEDUSAE

225

FIG. 8 Muscle fragment 10 days after isolation: smooth muscle cells and y cells (nerve cells) are well differentiated. f, flagella, ny, nuclei of y cells: s, smooth muscle-like arrangement of myofilaments (see insert): sn, nucleus of smooth muscle cells; sv, small vesicles of y cells; solid arrow, bundles of striated myofilaments; open arrows, peripheral vacuoles typical for smooth muscle. Bar, 5 pm; insert, 2 pm (from Schmid and Alder, 1984). FIG. 9 Muscle fragment 3 days after isolation. Smooth muscle-like arrangement of myofilaments (s) are continuous with bundles of striated myofilaments (arrow). Bar, 2 pm. FIG. 10 y cell (nerve cell); 10-day-old isolate. ny, nucleus of y cell; sv. small vesicles. Bar, 2 pm (from Schmid and Alder, 1986). FIG. 11 Five-day-old isolate spread on ECM (mesogloea) of the polyp stage of the jellyfish. Nerve cells are specifically stained with antiserum against RF-amide without any background on nonnerve cells. Whole mount demonstrating the network of nerve cell processes (from Schmid and Alder, 1986). FIG. 12 Enlarged part of a RF-amide-positive nerve cell demonstrating the presence of undegraded myofibers in its cytoplasm (arrow); ny, nuclei of nerve cell. Bar, 10 pm.

226

VOLKER SCHMID

To trace the change in the F-actin pattern during the transdifferentiation process NBD-phallacidin was used for fluorescence labeling of F-actin structures (Figs. 13-16; Weber et al., 1987a). The initial arrangement of well-organized bundles of striated myofilaments remained unchanged up to 24 hr (Fig. 13). Then the pattern of myofilaments changed strikingly, and the striated bundles of myofilaments became less organized (Figs. 14,

FIGS. 13-16 Whole-mount preparations of collagenase-treated isolates of different ages. The isolates were labeled with NBD-phallacidin to trace the change of the F-actin pattern of the contractile complex during transdifferentiation process (from Weber et a!., 1987a). FIG. 13 Fluorescence micrograph of a destabilized isolate, 8 hr after isolation. The excised monolayer of striated muscle cells rearrange to form a multicellular spheroid, which shows a compact appearance. At this moment the initial pattern of well-organized bundles of striated myofibrils remains unchanged. FIG. 14 Actin pattern of a destabilized isolate 52 hr after isolation. There is selective dissassembly of striated myofibrils. FIG. 15 Six-day-old isolate, destabilized after collagenase treatment. These isolates are characterized by a progressive loss of striated myofilaments on one hand, and the formation of a fine network of continuous strands of nonstriated filaments on the other. FIG. 16 In contrast to the latter, in 6-day-old isolates not treated with collagenase, the striated F-actin pattern of the muscle cells remains unchanged. Bar (same in Figs. 13-16), 20 pm.

TRANSDIFFERENTIATION IN MEDUSAE

227

15). Between the second and third day the striated myofilaments of the contractile complex either broke up at the Z-bands into short fragments or bifurcated laterally along their longitudinal axis. The newly arranged fibrils branched and formed a fine pattern of successively arranged fragments of myofilaments. At that point the myofilament pattern already showed most of the characteristics of myofilament arrangement of smooth muscle cells. During the following days, a progressive loss of striated myofilaments on one hand and development of densely packed continuous strands of contractile filaments at the base of each smooth muscle cell on the other were observed. Occasionally smooth muscle-like filaments form a continuum with bundles of striated myofibers (Fig. 9). With the exception of the remnants of striated myofibrils this cell type is indistinguishable at the ultrastructural level and in contractility from smooth muscle cells seen in some medusa organs (e.g., tentacle and manubrium; Schmid and Alder, 1984). All isolates of striated muscle showed this transitional change after activation treatment, whereas none of the untreated isolates, which were used as controls, exhibited DNA synthesis or flagellum formation. Instead, they remained in the differentiated state, and the original striated arrangement of the F-actin pattern of the contractile complex also remained unchanged (Fig. 16) and functional (spontaneous contractions) for weeks (Weber el al., 1987a). Recently, the mAb sm-1, specific for smooth muscle, was characterized (Ch. Weber, personal communication). It reacts with a 147-kDa protein, which is present in smooth muscle cells but absent in all the other cell types. During the transdifferentiation process mAb sm-1 detects a de nouosynthesized smooth muscle-specific molecule that becomes visible within 16 to 24 hr after the activation of the striated muscle fragments (Figs. 17-20). The 147-kDa protein from Podocoryne is a tissue-specific protein but it does not seem to display a distinct species specificity. The antibody cross-reacts even with protein from vertebrates. In the mouse, the mAb sm-1 reacts with a 147-kDa protein on blots from both smooth muscle and striated muscle. This finding invites speculations about evolutionary aspects of the relationship between striated and smooth muscle systems in both coelenterates and vertebrates (Weber et al., in preparation). With the use of actinomycin D, the role of transcription in the transdifferentiation process of striated to smooth muscle cells was examined (Weber et al., 1987b). Whole-mount preparations of muscle fragments labeled with NBD-phallacidin demonstrated that the overall pattern of striated myofilaments remains unchanged in isolates treated with actinomycin D prior to 48 hr after activation. The ultrastructural organization of the thick myosin and the thin actin filaments in the striated muscle cells of these isolates corresponds to the striated contractile complex of

228

VOLKER SCHMID

FIGS. I7 and 18 Immunostaining with the monoclonal antibody sm-1 on whole-mount preparations (courtesy of Dr. Weber). FIG. 17 Phase-contrast micrograph of freshly isolated not destabilized fragment of striated muscle spread on extracellular matrix for 4 hr. The contractile complex of the muscle cells consists of well-organized bundles of striated myofibrils (arrow) nu, nucleus. FIG. 18 Fluorescence micrograph of same area as shown in Fig. 17 labeled with mAb sm-I. Single smooth muscle cells showing anti-sm- I-positive immunoreaction. Bar, 40pm. FIGS. 19 and 20 Immunostaining with the mAb sm-1 on whole-mount preparations of isolated destabilized muscle fragments of different age (from Weber er nl., in preparation). FIG. 19 Fluorescent micrograph of a 4-day-old isolate spread on an ECM for 3 hr before fixation. The cells show sm-I-positive immunoreaction. Bar, 95 p m . FIG. 20 Nine-day-old aggregate. Only half of the isolate shows positive immunoreaction with sm-I, whereas the other half (dotted line) is not destabilized and therefore nonreactive. Bar, 100 wm.

TRANSDIFFERENTIATION IN MEDUSAE

229

isolates that remain in the differentiated state (Schmid, 1978). Furthermore, they share characteristics such as absence of flagella and lack of DNA synthesis with the differentiated state of striated muscle cells in untreated isolates. In destabilized isolates exposed to actinomycin D 3 to 4 days after isolation, the drug no longer has a visual impact on the transdifferentiation potential of the isolates. When 6 days old, the isolates formed de novo flagella and produced both smooth muscle cells and RF-amide-positive cells. These two cell types are indistinguishable in ultrastructure from corresponding cell types of destabilized isolates of the same age not treated with actinomycin D (Weber ef al., 1987a). When translation was inhibited by cycloheximide for up to 48 hr formation of the smooth muscle did not occur. Therefore formation of this cell type requires transcription and translation, but not DNA synthesis (see also Section E). b. Formation of RF-Amide-Positive Nerve Cells The RF-amidepositive cell type is distinctly smaller in size in comparison to smooth muscle cells (Figs. 8,lO). The cytoplasm bearing the base of the flagellum always forms a narrow bottle-shaped outgrowth at the periphery of the isolate. It contains more ribosomes than smooth muscle cells, and toward the base, numerous small vesicles filled with electron-dense material are seen (Figs. 8,lO). The vesicles strongly resemble the dense-cored vesicles found in nerve cells of coelenterates (Westfall and Kinnamon, 1978). To date, however, no synaptic structures have been identified. Vacuoles or filament arrangements like those found in smooth muscle cells have never been observed in this cell type. Occasionally cytoplasmic bridges connect developing cells of this type (unpublished). Although dense-cored vesicles are observed after mitosis (third day), the earliest immunoreactivity to RF-amide is seen from Day 4 on (Figs. 1 1,12). Fragmentary bundles of striated myofilaments are often seen in both cell types when young, but they gradually disappear with increasing age. Occasionally, however, they can be seen even in mature nerve cells expressing RF-amide immunoreactivity (Fig. 12). Both cell types are found in all destabilized isolates tested and remain present until the isolates fall apart after 3-12 weeks. Because of the stem-cell-like division pattern of the smooth muscle cells, the number of nerve cells increases in older isolates (see Section E,2). The fact that all destabilized isolates always expess these two cell types have raised speculations on their functional and developmental connections (Weber, 1989). The functional roles of smooth muscles with respect to locomotion and protective behavior have been investigated in other hydromedusae, mainly on the basis of electrophysiological data (Gladfelter, 1972; Mackie, 1975; Mackie and Singla, 1975). Immunocytochemical investigations with a RF-amide antiserum demonstrate the distribution

230

VOLKER SCHMID

of RF-amide-like immunoreactivity in the nervous system of many different coelenterates (Grimmelikhuijzen, 1983; Coe, 1987) but also in other phyla (Williams and Dockray, 1983; Marder et al., 1987). Immunoreactivity with (FM)RF-amide antisera in other hydrozoans is restricted to a subset of nerves associated with smooth muscle cells and certain sensory structures (Grimmelikhuijzen and Spencer, 1984; Mackie et al., 1985; Coe, 1987). In double-labeling experiments of Podocoryne medusae, the distribution of neurons showing RF-amide-like immunoreactivity also corresponds most strikingly to the distribution of the smooth muscle system (Weber, 1989). In addition to suggesting a functional relationship between the immunoreactive neuronal system and the smooth muscle system (Mackie et al., 1985), this tight association also invites conclusions about developmental aspects of this neuromuscular pattern. During transdifferentiation, the nerves that are immunoreactive with RFamide are always a division product of the smooth muscle cells (Alder and Schmid, 1987), demonstrating again a close developmental relationship. Whether similar developmental mechanisms are also valid during ontogeny of the medusa, and whether these mechanisms result in the observed proximity of the smooth muscle pattern and the distribution of neurons showing RF-amide-like immunoreactivity, is still highly speculative and is the subject of further investigation.

d. Formation of the Regenerate When the destabilization and culture conditions are appropriate (Schmid and Alder, 1984), 20-30% of the activated muscle fragments form by the fifth day an inner endodermal layer separated from the outer, ectodermal layer by a basal lamina (Schmid and Alder, 1984). In some rare cases cells from both layers still contain portions of striated myofilaments (Fig. 21). Occasionally bundles of striated myo-

FIG. 21 Muscle fragment 6 days after isolation. The fragment has formed an outer, ectodermal layer (ec) and an inner, endodermal layer (en) separated by a basal lamina (m).Cells of both tissues contain bundles of striated myofilaments (solid arrows). Bar, 5 p m (Figs. 21-26 from Schmid and Alder, 1984). FIG. 22 Muscle fragment 6 days after isolation (same specimen as in Fig. 21). The first nematoblasts differentiate in the endodermal layer; the thread (t) of the nematocyst is still outside the capsule (c). The same cell contains nonvacuolated bundles of striated myofilaments (solid arrow). n, nucleus of nematoblast; sv, small vesicles of y cell. Bar, 5 pm. FIGS. 23 and 24 The electron micrographs demonstrate the presence of bundles of striated myofilaments (arrows) in the cytoplasm of nematocytes. c, capsule of nematocyte; n, nucleus of nematocyte. Bars, 2 pm. FIG. 25 Striated myofilaments in a gland cell. Bar, 2 p m . FIG. 26 Striated myofilaments (arrow) in a digestive cell. nd nucleus of digestive cell. Bar, 2 pm.

TRANSDIFFERENTIATION IN MEDUSAE

231

232

VOLKER SCHMID

filaments are even found in differentiating nematoblasts (Fig. 22). Between the fifth and sixth day directional swimming of the isolates indicates the expression of the proximodistal axis of the future regenerate. In the following days the normal inventory of at least seven to eight new cell types differentiate, perfectly arranged to form a tentacle or a manubrium (Figs. 23-26), the sexual and feeding organs of the medusa. In some cases both manubrium and tentacle were expressed in the same regenerate (Schmid, 1988). The origin of most of the newly formed cell types from striated muscle cells is visually documented by the few rare cases in which the striated myofibers were not completely degraded and remained visible in the cytoplasm of the new cell types (Figs. 22-26). Usually the regenerates are very small and so far no attempts have been made to feed the regenerated manubrium and thus initiate gamete formation (see later).

2. The Endoderm of the Umbrella Due to the isolation procedure, freshly isolated endoderm is covered by the outer mesogloea but might also contain remnants of the inner mesogloea (Fig. 1; Schmid et al., 1982). In the following 12-24 hr the endoderm forms a sphere and reduces the gap junctions (visualized by lanthanium; Weber and Schmid, 1984), and the ECM material completely disappears from the surface of the isolates. The removal of the adhering ECM follows a characteristic pattern of selective degradation by the endodermal cells (Weber et al., 1987b). The process involves three distinct steps: an initial extracellular condensation within the ECM fibrillar network, followed by intercellular internalization of the fibrillar elements and subsequent endocytosis of ECM material. The first step immediately follows the removal procedure of the exumbrellar cells and is completed within hours. This process cannot be interrupted by dihydrocytochalasin B (H,CB), a drug that destroys the actin microfilaments. The second step lasts 24-28 hr, is mediated by cell mechanisms, and can be stopped by H,CB. The third step is a slow process (of up to 14 days). It involves intercellular degradation of fibrillar material, endocytosis, and completion of digestion within lyosomes (Weber et al., 1987b). The endoderm is extremely sensitive to temperature in the isolated state. When cultured like muscle isolates at 22"C, it dies within 2-4 days; however, when cultured at 10-12°C it will survive for weeks and form functional flagella (Schmid er al., 1982). Gap junctions can be fully restored, at least in the first days, if the endoderm is allowed to spread into monolayer on isolated and permanently stretched ECM (Weber and Schmid, 1984). Contrary to the situation in the isolated muscle, DNA synthesis and mitosis were never observed in the isolated endoderm, even after collagen-

TRANSDIFFERENTIATION IN MEDUSAE

233

ase treatment (Schmid er al., 1982). Endoderm seems unable to form new cell types or a regenerate when subjected to the same collagenase treatments that are successful with muscle. However, the potential to transdifferentiate is present in endoderm, as demonstrated by the following experiments.

3. Combinations of Striated Muscle and Endoderm Endoderm can be activated to synthesize DNA and to transdifferentiate to new cell types when combined with isolated striated muscle (Figs. 27-29; Schmid er al., 1982). In this case regeneration leads to a fully functional manubrium, able to produce gametes (Fig. 29), and occassionally to tentacles. The ability to activate endoderm is present only in destabilized striated muscle (i.e., treated with collagenase prior to grafting; Table I). Muscle isolated mechanically, without activating treatment, and then grafted to endoderm, stably maintains its own differentiated state, and neither DNA synthesis nor formation of new cell types is observed in the endoderm (Table I; Schmid et al., 1982); no regenerate is formed. Direct desmosomal contact between the muscle and the endoderm does not suffice for the activation of the endoderm. The critical condition for activation is that the number of activated muscle cells must be large enough (20-40) to cover the endoderm (Schmid er al., 1984).

4. The Transdifferentiation Potential of the Endoderm As the isolated striated muscle alone can give rise to all the cell types found in complete regenerates (see above), the grafting experiment described in the previous section cannot conclusively demonstrate the transdifferentia-

FIG. 27 Isolated destabilized striated muscle (st) grafted to endoderm (en), 4 hr after grafting. Bar, 100 p m (Figs. 27-29 from Schmid et a / . , 1982). FIG. 28 Regeneration 4 days after grafting; the endodermal cavity contains a ball-shaped fragment of endoderm (fen) not incorporated into the endodermal wall. Bar, I 0 0 pm. FIG. 29 Regenerated and fed manubrium (4 weeks): nematocytes (nc) are well developed at the manubrial lips and eggs are maturing in the gonads (go). SEM. Bar, 50 pm.

TABLE I The Effect of the Time of Collagenase Treatment on DNA Synthesis in the Regeneratesa

Treated with collagenase for Not treated with collagenease Days of culture 1

2 3 7-8 Regeneration

4 hr

24 hr

48 hr

72 hr

n

sm(r)

en(r)

n

sm(r)

en(r)

n

sm(r)

en(r)

n

sm(r)

en(r)

4 4 4 3

01369 01301 11372 121335 0

01388 0131 01205 01182

4 4 3 2

01483 01227 01219 31151 0

01222 01138 01122 0197

4 4 4 4

01230 151185 541228 1701183

01335 11263 221230 1891190

3 4 4 2

21302 621160 17/37 26/41

01131 01172 31131 221123

+

+

sm(r)

en(,-)

4 181152 3 361233 4 301263 -

41213 91222 451194

n

+

-

The striated muscle (sm) was treated prior to grafting upon endoderm (en). Grafting occurred immediately after collagenase treatment. n, number of experiments; r, labeling index (number of labeled cellsltotal number of cells) (From Schmid and Alder, 1986). I'

235

TRANSDIFFERENTIATION IN MEDUSAE

tion of the endoderm. More evidence is given by experiments in which the destabilized striated muscle was treated with mitomycin C, a drug that irreversibly inhibits DNA replication (Schmid et al., 1982). With the hypothesis that cell cycles are a prerequisite for cellular commitment, and therefore also for transdifferentiation (see below), one would predict that destabilized muscle treated with mitomycin C cannot contribute any further new cell types to the regenerate, other than those already present or initiated at the time of drug treatment. This treatment was applied between Days 3 and 4 after isolation, when only smooth muscle and RF-amidepositive cells could have been present. After grafting to endoderm small but complete functional regenerates were formed, and the histodynamics and DNA synthesis pattern in both tissue layers clearly demonstrates transdifferentiation of the endoderm comparable to that of the muscle (Table 11; Schmid et al., 1982). The reverse experiment, in which mitomyocin-C-treated endoderm is grafted onto destabilized muscle, confirmed the great transdifferentiation potential of the muscle (Table 11). In both types of mitomycin-C experiments, the regenerated manubria occasionally contained gametes (Fig. 29; oocytes; Schmid et al., 1982). This indicates that both cell types ultimately have the ability to transdifferentiate to gametes.

TABLE I1 Formation of New Cell Types in Grafts Consisting of Striated Muscle and Endoderma Control

Treated with mitomycin C

Age

(days)

n

ec

ten

1

4

2

5

a (342) a; b (390) a-c (893) a-d (623)

a (219) a (280) a: b; d (844) a-e: g (654)

3

II

4

7

5

3

b-d

b-g

(346)

(366)

n

mec

en

n

2

a (144)

a (219)

2

3 2

a; b

I

(105) a: b

(75)

3

7

a:b;e

(34) 8-1 1

5

b-e:g

(385)

b-g (516)

a; b (352) a; b; d; f (259) a-g (654)

4 5

3

ec

men

a; b

(389) a-c (571) a-c (861) b-d ( 1272) b-e

(709) 3

b-e; g

(253)

ec, striated muscle; en, endoderm. The tissues were alternately treated with mitomycin C before grafting. mec, striated muscle treated with mitomycin C; men, endoderm treated with mitomycin C; a, original cell types of the ectoderm or the endoderm; b, interstitial cells; c , gland cells; d, nematoblasts; e , nematocytes; f, secretory cells; g, gametes; n, total of examined specimens; number in parentheses is the total number of examined nuclei (From Schmid et ul., 1982).

236

VOLKER SCHMID

It seems that the activation potential of destabilized muscle fragments resides in its ability to degrade ECM by proteolytic activity. This activity was observed highest 30-50 hr after destabilization treatment (Schmid et al., 1992a).

D. Initiation of Transdifferentiation

1. The Role of the Extracellular Matrix a . Tissue-ECM Interactions in Hydromedusae Molecular and biochemical studies have demonstrated the existence of glycoproteins in the extracellular matrix that are connected through transmembrane molecules to the cytoskeleton. Cell adhesion, migration, differentiation,and intercellular communication may largely depend on these molecules. These ECM components seem to occur throughout the animal kingdom and to be conservative in their chemical composition (Miura and Kimura, 1985; Beck et al., 1989; TilletBarrett et al., 1992). Various observations in our laboratory indicate that the initiation of the transdifferentiation process is influenced by components of the ECM to which the muscle tissue strongly adheres (Schmid, 1978; Schmid et al., 1992a; Reber-Muller and Schmid, 1992). To reveal this muscle-ECM relationship, we showed that the ability of the isolated muscle cells to adhere and spread on a substratum is not restricted to their own ECM (tissue specificity) but that other ECMs of the medusa or the polyp stage can serve the same purpose. The results clearly demonstrate that the ECMs carry no tissue-specific components for cell adhesion and spreading. This is true for all species tested (Schmid and Bally, 1988). However, a strong species specificity exists throughout the whole portion of the ECM (Schmid and Bally, 1988). The adhesion factor can be largely removed by extracting the ECM with 5 M guanidine-HCI. Normal cell adhesion and spreading properties are restored when the extracted ECM is incubated with its extract. The factor is also extractable with 0.5 M acetic acid and can be precipitated with 1 M sodium chloride. The precipitate is almost insoluble in water, is of fibrillar structure, and supports specific adhesion and spreading of striated muscle tissue. It consists mainly of a cysteine-containing collagen (Schmid and Bally, 1988).

b. Transdifferentiation by Enzyme Treatment Transdifferentiation can be initiated in this system by enzymes that seem to degrade primarily components of the ECM. Hyaluronidase is the least effective of the activating enzymes. In Podocoryne (not published) and in Stomotoca atra (Schmid, 1978), DNA replication is initiated in 10-20% of the cells when

TRANSDIFFERENTIATION IN MEDUSAE

237

tissues were treated for 1 to 2 days at RT. Collagenase was used as a semipurified fraction (type 111, fraction A; Sigma Chemical Co.) for both isolation of the tissues and postisolation treatment. Tissue fragments isolated with collagenase but not treated with this enzyme after isolation destabilize at low rates (20-40%, unpublished). When treated again with this enzyme after isolation at 26-28°C for 4-8 hr, 60-80% (average) of all cells will undergo DNA replication and transdifferentiation (Alder and Schmid, 1987).The most potent enzyme is Pronase (1.25 mg/ml seawater, Streptomyces griseus, Boehringer Mannheim, Germany). Treatment for 2 to 4 min of mechanically isolated fragments at 22°C is sufficient to induce 100% DNA replication and transdifferentiation (Figs. 31, 40; Schmid et al., 1992a).

c. Inhibition and Induction of Transdifferentiation by ECM Grafts In ontogeny and regeneration, migration of tissues and cells along an ECM is an indispensible process. Cell and tissue mobility requires that the ECM ligand-cell receptor complex, which mediates cell substrate adhesion, be altered. There is good evidence that the molecular composition of the substrate on which migration takes place can influence cell fate (Kornblihtt and Gutman, 1988; Chiquet-Ehrismann, 1991). However, recent reports suggest that mechanical forces transmitted by the substrate may also act as biological regulators (Ingber and Folkman, 1989a,b; Streuli & Bissell, 1990; Ingber, 1990;Mochitate etal., 1991).These forces change the mechanochemical interplay (Ingber and Folkman, 1989a) between the ECM and the attached cells, and cell shape, distribution of integral membrane molecules, and the cytoplasm are all affected. One of these, pertubation of cell shape, is reported to change nuclear functions such as the synthesis of DNA and RNA and the expression of differentiation-specific genes (Bissell and Barcellos-Hoff, 1987; Ingber and Folkman, 1989b; Ben-ZC ev, 1991). In wounded jellyfish large areas of exposed ECM are recovered with migrating tissue before DNA replication and regeneration are initiated (Schmid et al., 1976). Cell migration in uiuo, however, exposes the cells with new substrate, which can vary in both structure and molecular composition. We have addressed the question whether the migration process or cell shape can alter gene expression in differentiated cells. To mimic the in uiuo situation and promote tissue/cell migration we combined isolated striated muscle tissue of medusae with differently sized and variously structured cell-free ECMs (Fig. 30; Schmid er al., 1992a). The influence of the ECM grafts on cell cycle activity and the transdifferentiation process was then investigated. The isolates were either grafted on small pieces of floating ECMs (Fig. 30g) or on large permanently stretched ECMs (Fig. 30h). In both cases adhesion and cell migration onto the grafted ECMs occurred.

238

VOLKER SCHMID

FIG. 30 Schematic drawing summarizing isolation of ECMs and combination of ECMs with isolated striated muscle tissue of medusae. Polyps and medusae (a) were incubated in CaMgfree seawater (CaMg RW).The obtained cell-free ECMs (b) were air-dried onto coverslips (c) and afterward transferred to a culture dish (d). Isolated striated muscle tissue was grafted onto the ECMs (e) with the help of a piece of coverslip. For floating cultures the ECMs together with the adhering and spreading tissue were removed from the coverslip (f). Cultures with permanently stretched ECMs remained unchanged (h). Strippled areas, cell-free ECMs from medusa or polyp: cross-hatched areas, inner mesogloea to which the muscle tissue adhere; gm, grafted mesogloea; im, inner mesogloea (ECM); pcs, piece of broken coverslip; st, striated muscle tissue (from Schmid er a / . , 1992a).

a. D N A Replication and Transdifferentiation Induced in Floating Grafts Consisting of Isolated Striated Muscle Tissue and Small Pieces of ECM of Different Origins Striated muscles were isolated without enzymes and combined with pieces of ECM from different locations of the medusa or its polyp (Fig. 30). Their fate was compared with those of uncombined isolates (control 1) and of isolates activated by pronase treatment (control 2; Fig. 31). In the combinations most of the muscle cells leave the native ECM, overgrow the grafted ECM, and completely cover it within 12 to 24 hr depending on the size of the ECM. Both the grafted and the native ECM are gradually degraded and the isolate that was initially of swollen appearance, shrinks. From the third day onward both combinations of muscle and ECM and pronase-activated isolates (control 2) start to swim with newly formed flagella. DNA replication and transdifferentiation to RF-amide-positive nerve cells occur much more frequently in the combined fragments than in untreated controls. The extent and rate of these processes in the experimental combinations are similar to those seen in isolates artificially activated with pronase. The source of the ECM, polyp or medusa, has no effect on the outcome (Fig. 3 I , Schmid et a f . , 1992a).

239

TRANSDIFFERENTIATION IN MEDUSAE %

120

W

contr. 1

100

80 60 40

20 0

labeled nuclei to total nuclei

RF-amide positive nerve cells

FIG. 31 DNA replication and formation of RF-amide-positive nerve cells in combinations of isolated striated muscle with ECMs from the medusa or the polyp. The combinations were cultured under floating conditions. Contr. I , untreated, uncombined specimens: contr. 2, uncombined specimens activated with pronase; im, inner mesogloea of medusa; om. outer mesogloea of medusa: pm, mesogloea of polyp (from Schmid rf a / . , 1992a).

6 . DNA Replication Induced by Grafting Isolated Striated Muscle on Large, Permanently Stretched, Polyp ECM Large pieces of polyp ECM were kept permanently stretched on coverslips (Fig. 30h). Mechanically isolated striated muscle tissue adheres quickly to these stretched ECMs, and the cells start to spread into a monolayer. Staining with a mAb of the ECM adhering to the muscle tissue before grafting reveals that the migrating cells leave their own ECM behind (Figs. 32, 33). The grafted ECM, however, is too large to be covered completely by the cells. Depending on the size of the isolates, cell migration ceases after 22-24 hr. At this point 60 to 80% of the cells have migrated from their own ECM onto the grafted ECM. DNA replication is much increased compared with untreated controls, but never reaches the same level as that in floating ECM combinations (Fig. 38). Radiolabeled nuclei and RF-amide positive cells (putative neurons) are mostly to be found at the contact zone between the muscle and the grafted ECM and seldom in the peripheral, most highly spread portions of the muscle tissue (Fig. 39). As shown in pulse-labeling experiments, the cells that initiate DNA replication are those that extend from their own to the grafted ECM (gap cells, Figs. 34, 35, 39; Schmid et al., 1992a). When cell membranes were silver-stained, it could be shown that gap cells adhere to both the native and the grafted ECM (Figs. 36,37; Schmid et al., 1992a). In this region proteolytic activity degrades both types of ECM underneath the cells (zones 1-3 in Fig. 39). After removal of the cells with

240

VOLKER SCHMID

FIGS. 32 and 33 Combination of muscle cells with permanently stretched polyp ECM. Same objects: the native ECM was stained with mAb 19 prior to grafting; the tissue was allowed to spread 12 hr before fixation (from Schmid et a / . , 1992a). FIG. 32 The distribution of the DAPI-stained nuclei shows the spreading pattern; arrowheads delimit the junctional area where cells move from the native ECM to the grafted ECM. Bar, 100 p m . FIG. 33 The native Ab-labeled ECM is left behind when cells spread on the grafted ECM. FIGS. 34 and 35 Combinations of muscle cells with permanently stretched polyp ECM. Same specimen, autoradiography of a spread specimen fixed after 40 hr (Fig. 34). DNA replication starts at the junctional area between the native ECM and the grafted ECM; the native ECM was stained with mAb 44 prior to grafting (arrowheads point to the stained thread-like structures of the native ECM). Bars, 100 pm (Fig. 34) and 50 pm (Fig. 35) (from Schmid ef al., 1992a). FIG. 36 Combinations of muscle cells with permanently stretched polyp ECM. Specimen, silver-stained after the spreading process had ended (26 hr); cell membranes are clearly visible; arrow delineates the junctional area beneath the native and the grafted ECM. Bar, 100 p m (from Schmid et a / . , 1992a). FIG. 37 Combinations of muscle cells with permanently stetched polyp ECM; Enlarged portion of Fig 36; the course of the cell membranes demonstrates that cells use both the native and the grafted ECM as substrates. n, nucleus. Bar, 100 p m (from Schmid et a / . , 1992a).

241

TRANSDIFFERENTIATION IN MEDUSAE

. . .

7

1

I

-

+ & -A -.-

2

4

6

A

A

8

days

FIG. 38 DNA replication in combinations of muscle cells and permanently stretched polyp

ECMs. Solid circles, muscle tissue activated with pronase and not grafted onto ECMs; solid triangles, muscle tissue not treated with the pronase and not grafted onto ECMs; open triangles, muscle tissue not treated with pronase but grafted onto ECMs. Ordinate, percentage of labeled nuclei (from Schmid et a / . , 1992a).

CaMg-free seawater, degraded areas can first be detected in the grafted ECM 6 hr after the migration process has come to an end (28-30 hr). Occasionally in older grafts, a large hole in the underlying stretched polyp ECM can be observed (Schmid et al., 1992a). c . Inhibition of DNA Replication by ECM Grafts in Activated Isolates The results presented above raise the question whether DNA replication and transdifferentiation are inhibited when the cells stretch into a monolayer. To test this hypothesis isolated muscle tissue was activated by pronase treatment and grafted on permanently stretched polyp ECM, either immediately after pronase treatment or after termination of the first cell cycle. In ungrafted controls all cells undergo DNA replication, whereas in fragments grafted immediately after activation treatment, DNA replication is reduced and restricted to the site of grafting. If those cells that still adhere to their own ECM are removed after 8-15 hr, the remaining cells, being those that have spread out on the grafted ECM, rarely show any DNA replication (Fig. 40). Since pronase treatment reliably initiates cell cycle activity, we conclude that the spreading and flattening process on the stretched polyp ECM suppresses DNA replication and transdifferentiation. If this is correct, actively cycling cells should be prevented from conducting further cell cycles when allowed to spread on permanently stretched ECM. This is actually the case. When pronase-activated isolates

242

VOLKER SCHMID

a

4

3

2

1

Pm

b

4

3

2

1

C %

6.0

4.0

2.0

0.0 4

3

2

1

FIG. 39 Dynamics of DNA replication in striated muscle tissue cultured on permanently stretched polyp ECM. Specimens were continuously cultured in [3H]thymidineuntil time of fixation. (a) Schematic drawing ofthe specimen at the time of fixation (sideview); (b) distribution of labeled nuclei at 43 hr in the zones indicated in (a); ( c ) like (b), time of fixation was 48 hr. im, native ECM of the muscle tissue; pm, stretched ECM of the polyp (from Schmid et a / . , 1992a).

TRANSDIFFERENTIATION IN MEDUSAE

loo

243

. . . .

1 I

50-

I

j

. A A

‘

4

A

2

4

1 A

,

a

6

days

FIG. 40 DNA replication in pronase-activated muscle tissue cultured on stretched polyp ECM. Specimens were permanently cultured in the presence of [3H]thymidine. Solid circles, pronase activated tissue not grafted to ECM; open triangles, specimens were grafted to stretched polyp ECM; solid triangles, specimens where all tissue that had not spread on the grafted ECM were removed 15 hr after grafting. (from Schmid et al., 1992a).

are allowed to spread on polyp ECM between the first and the second cell cycle, subsequent cell cycles are reduced after 5 to 12 hr (Table 111). Interestingly, those cells that have already completed the first cell cycle before grafting are able to differentiate into nerve cells even while being TABLE 111 The Effect of Spreading on DNA Replication and Formation of RF-Amide-Positive Nerve Cellsa ~~

Specimens spread on polyp ECM

DNA replication Formation of RF-amide-posi tive nerve cells

Unspread controls

n

tot

a

S

n

tot

a

S

9 9

441559

7.31

8.0

5 5

5381726

75.48

16.12

651559

14.85

22.11

3771726

54.50

19.28

Pronase-activated fragments of isolated striated muscle spread on permanently streched polyp ECM were compared with unspread controls. Grafting of the tissues onto the ECM and spreading occurred after completion of the first cell cycle (50-54 hr). Labeling with [3H]thymidine was performed from 64 hr until fixation at 148 hr. n . number of specimens; tot, labeled nuclei to the total of nuclei examined; a, average; s, standard deviation (After Schmid e t a / . , 1992a).

244

VOLKER SCHMID

well spread. These observations demonstrate that the inhibition affects only DNA replication, and not the differentiation process to new cell types. d . The Effect of Art$ciul Stretching of Activated Isolates under Fluorocarbon An experiment was designed (Fig. 41) to distinguish between the following hypotheses: (a) DNA replication is inhibited by a factor associated with the stretched polyp ECM; (b) DNA replication is rendered impossible because of changes in cell shape and cytoarchitecture. Because pronase-activated muscle cells are loosely arranged and dissociate easily, they are not well suited to sustain applied stretching forces. The isolated striated muscle tissue was therefore activated with collagenase instead. This enzyme is a less potent activator of DNA replication and transdifferentiation than pronase (see Section III,D,l ,bJ; however, the number of activated muscle cells in preparations treated with collagenase is still significantly higher than that in controls not treated with any enzyme. The results clearly demonstrate that both initiation of DNA replication and formation of flagella were delayed for the time the isolated muscle tissue was kept in a stretched configuration under fluorocarbon (Fig. 42). The control c2 show that this is not an artifact due to fluorocarbon. Together these experiments indicate than inhibition of DNA replication control c1

isolated striated muscle

I

I d

I I

I

control c 2

unstret ched

s, and sa

stretched

FIG. 41 Schematic drawing of the depression slide (d, side view) in which the collagenaseactivated isolated striated muscle (ist) with a very small amount of seawater is cultured unstretched (control c,) or stretched (sI and s2) under a layer of fluorocarbon (Fc) (from Schmid ef a / . , 1992a).

TRANSDIFFERENTIATION IN MEDUSAE

245

does not depend on the presence or quality of a grafted ECM, but is more likely correlated with the ability of the cells to maintain or reexpress a stretched monolayer, a morphology that the muscle tissue also possesses in uiuo. Whereas striated muscle cells are inhibited from DNA replication when they firmly adhere and spread out on a substrate, it has to be pointed out that in other vertebrate systems cells are triggered to proliferate under comparable conditions (Ingber and Folkman, 1989a). In our system we demonstrated that the first activated cells are the gap cells. The gap cells are confronted with two substrates that are heterogeneous in their physical properties (the malleable native ECM and the rigid grafted ECM). Because cell migration from the native ECM to the grafted ECM stops 24 hr after grafting, the gap cells are prevented from moving into a more homogeneous environment and appear unable to maintain their differentiated state in this position. DNA replication is induced and proteolytic enzymes destroy both types of ECM underneath the cells at the site of grafting. This possibly represents the first step in reestablishing a more adequate substrate. Since the zone of activated cells enlarges with time we assume that proteoloytic degradation of ECM components activates the additional cells. In the case of the permanently stretched polyp ECM the activation zone is restricted to the adjacent cells of the gap zone. The gap cells, unable to move to a more homogeneous substrate, seem to detect their altered configuration and do not maintain their differentiated state in this position. There are many ways cells could perceive inhomogeneities of the substrates. Mechanotransducers like stretch-activated and stretch-inactivated ion channels could be activated in gap cells. A strong correlation between shape-dependent alkalinization of capillary endothelial cells and an increase in DNA systhesis, has been reported. Growth stimulation in many cells is accompanied by an increase in intracellular pH (Watson, 1991), and cytoplasmic alkalinization results from the activation of the Na+/H+exchangers. Change in cell shape can also result in acceleration of CAMP synthesis by adenylate cyclase. The activity of this enzyme seems to require an intact microfilament system (Watson, 1991).It becomes evident that cell shape and the factors that influence cell shape delimit the cells' responsiveness. The interplay between the adhesion substrate and the gap cell seems to be disturbed and mechanotransducers such as the ones mentioned or others provide the gap cells the information to change this situation. It seems very likely that under conditions of in uiuo regeneration, the differentiated tissues that participate in wound healing are activated for DNA replication and possibly for transdifferentiation by similar mechanisms.

A

l00

80

60

40

20

Yo

100

-

80

-

60

-

40

-

20

-

B

TRANSDIFFERENTIATION IN MEDUSAE

247

2. The Role of Protein Kinases

Studies with tumor-promoting phorbol esters have shown that a wide range of biological phenomena can be induced by these agents (Castagna, 1987; Blumberg, 1988). Phorbol esters like TPA are known to act as mitogens and to stimulate DNA synthesis (Glimelius and Weston, 1981; Rozengurt, 1986); however, there are also several reports for inhibition of growth (Doctrow and Folkman, 1987; Huang and Ives, 1987). In this case an induction of differentiation can be observed (Montesano and Orci, 1985). TPA, the most potent tumor promoter, is a lipophilic substance that intercalates into the cell membrane, where it acts as a substitute for diacylglycerol (Nishizuka, 1984). It directly activates protein kinase C (PKC) by increasing the affinity of this enzyme to Ca2+(Niedel et al., 1983). Protein kinase C is thought to play a crucial role in the transduction of extracellular signals into the cell (Nishizuka, 1988). Therefore, most, if not all, of the biological effects observed in cell cultures treated with phorbol esters are thought to be mediated by this serinehhreonine, Ca2'-activated phospholipid-dependent protein kinase. Transdifferentiation can be initiated by treating striated muscle tissue immediately after nonenzymatic isolation for 3 hr with TPA (Kurz and Schmid, 1991). The number of cells having replicated their DNA and transdifferentiated to RF-amide-positive cells by TPA treatment is presented in Table IV. Within the first 5 days only 5% of all nuclei in control cultures become labeled (continuous incubation). The same is true for isolates treated with the inactive phorbol ester TPA-methylether (Table IV). The role of protein kinase C in the initiation of DNA replication is furthermore stressed by experiments in which its activity was inhibited. K-252a has been reported to be a potent inhibitor of the protein kinase C in uitro (Yamada et al., 1987). In isolates treated simultaneously with TPA and K-252a the rate of transdifferentiation was strongly reduced (Table IV). Mezerein has been reported to be a weak tumor promoter in mouse skin. This nonphorbol diterpene can activate protein kinase C in uiuo and in uitro (Couturier et al., 1984). Incubation of isolated striated muscles for 10-24 hr with mezerein initiates transdifferentiation, however, at a reduced rate when compared to TPA (Table v). Diacylglycerol (DiC,) can

FIG. 42 The effect of stretching under fluorocarbon on flagellum formation (a) and DNA replication (B) in isolated fragments of striated muscle. Abscissa: days after isolation, ordinate: percentage flagellated of total isolates (A) or percentage labeled to the total of examined nuclei (B). c , , control, cultured without further treatment; c2,control, cultured permantently under fluorocarbon; s,. isolates stretched for I day; s2. isolates stretched for 2 days under fluorocarbon (from Schmid et a / . , 1992b).

248

VOLKER SCHMID

TABLE IV DNA Synthesis and Formation of RF-Amide-PositiveNerve Cells in TPA., Diacylglycerol (DiC8)-,and Mezerein-Treated Isolates of Striated Muscle Tissue'

Treatment

u

n

f

Control

8 8 8 8 8 8 4 4 10 4 4 10

18 18 17 12 10 48 7 8 5 9 9

1325 608 581 359 769 1727 388 470 363 836 433 643

TPA Control DiC, Control

Mezerein

10

Time of labeling (hr)

% Labeled

0-120 0-24 24-48 48-72 0-72 0-72 0-72 0-120 0-72 0-120

5 t 13.3 6 f 6.3 81 f 24.7 10 t 6.1 3 f 3.2 95 f 13.7 2 2 1.4 1 ? 1.2

-

nuclei (t SD)

-

16 ? 10.6 51 f 19.5 -

% RF-amide-positive

cells

(2

SD)

2 f 7.5 38 k 9.9 39 f 10.6 38 t 7.8 1 t 1.8 23 f 14.6 1 f 1.6 23 f 7.1

' u , age in days of the specimens when processed for autoradiography and immunocytochemistry; n , number of fragments examined; f, total number of nuclei counted (DAPI stained) (From Kurz and Schmid, 1991).

mimic the effect of TPA on isolated striated muscle cells (Table V). As already observed after TPA treatment, DiC,-treated isolates underwent compaction and displayed transient dissociation of cells. Compared to TPA cultures, the appearance of flagellated aggregates was delayed: The majority of the isolates acquired autonomous ability at the fourth day.

TABLE V Effect of TPA-Methylether and K.252a on DNA Synthesis and the Formation of RF-Amide-Positive Nerve Cellsa

Treatment

a

n

r

Control TPA TPA-methylether Control TPA K-252a + TPA

8 8 8 7 7 7

23 32 25 18 26 29

1905 784 1791 1465 613 1564

Time of labeling (hr)

% Labeled

0-120 0-120 0-120 0-72 0-72 0-72

3 t 6.2 81 t 14.7 5 k 9.1 3 t 2.4 94 f 6.5 16 f 32.7

nuclei ( f SD)

% RF-amide-positive

cells ( t SD) 1 28 1 1

41 5

2.5 12.0 t 3.3 ? 0.9 f 8.9 2 22.4 f

f

'a, age in days of the specimens when processed for autoradiography and immunocytochemistry; n , number of fragments examined; t . total number of nuclei counted (DAPI stained) (From Kurz and Schmid, 1991).

TRANSDIFFERENTIATION IN MEDUSAE

249

Among the various effects reported for TPA, the initiation of transdifferentiation to new cell types is one that has not been observed before. Diacylglycerol, the physiological activator of PKC in uiuo, can mimic the effect of TPA in our system. Furthermore, transdifferentiation induced by TPA can be counteracted by simultaneous treatment with K-252a. The same inhibitory effect can be observed when muscle tissue is incubated simultaneously with H7, another inhibitor of protein kinase C (data not published). These results suggest that PKC may be implied in the activation of striated muscle cells for transdifferentiation. Since K-252a also has effects on protein kinase A (Kase et al., 1987), it cannot be ruled out that other transduction pathways may be involved. In contrast to the effects of TPA on muscle cells of rat and chicken, the activation for transdifferentiation in our system cannot be reversed by removal of the drug. A short time exposure (3 hr) to TPA causes irreversible changes. Degraded myofibrils, for example, are not restituted and transdifferentiation proceeds.

3. Induction of Transdifferentiation by Dissassembly of the Microfilament System Observations in vertebrate systems indicate that cell-shape and cell-volume changes can have strong effects on gene expression (Watson, 1991). Initiation of transdifferentiation in this system with enzymes or drugs (see previous and following sections) is often correlated with dissociative effects. The activation process evidently leads to changes in cell shape. Changes in cell shape, however, require alterations in the structural organization of the cytoskeleton. We have followed the question whether along the signal transduction pathway, initiation of transdifferentation could be achieved by destructive processes at the level of cytoskeleton, namely the microfilaments. This was done by incubating the isolated striated muscle tissue in dihydrocytochalasin-B, a drug that leads to dissassembly of the actin filaments. Observations on isolated muscle tissue of Gonionemus (not published) and experiments with Podocoryne demonstrate that DNA replication and transdifferentiation are effectively induced by this drug (Kurz and Schmid, 1993). Again the activation process correlates with dissociative effects and therefore cell-shape changes. 4. Induction of Transdifferentiation by Molecular Interactions with Carbohydrates As was demonstrated in Section D, 1 ,b, transdifferentiation of striated muscle can be initiated in the mechanochemical interplay between cells and the ECM are altered (Schmid er al., 1992a). To identify ligands involved in those interactions two monoclonal antibodies specific for differ-

250

VOLKER SCHMID

ent ECM ligands and different lectins were investigated on their ability (a) to inhibit cell adhesion and spreading of the muscle tissue on grafted ECM and (b) to induce DNA replication and transdifferentiation (ReberMiiller and Schmid, 1992). Experiments with mAbs 19 and 44 can be summarized as follows: MAb 19 strongly effects cell adhesion and spreading in a time-dependent manner. When the grafted permanently stretched ECM (Fig. 30) was pretreated with this antibody for 3 hr, adhesion and spreading were reduced. If, however, pretreatment was extended to 12 hr cell adhesion was strongly reduced and no spreading occurred. DNA replication and transdifferentiation could be induced in striated muscle tissue isolated without enzymes when treated directly with this antibody for short periods (15-60 min; Table VI). In this case the inner mesogloea to which the muscle tissue adheres was intensively stained by mAb 19. As usually observed by activating treatments with enzymes or drugs, treatment of the tissue with this mAb produced dissociative effects that increased with the time of treatment. In whole mounts of medusae ECMs are unspecifically and very intensively stained by mAb 19 (Fig. 43). This antibody is directed against carbohydrate residues of different ECM ligants (Fig. 45). In whole mounts mAb 44 stains fibrillar structures in medusa and polyp ECMs (Fig. 44). The antigen resists deglycosylation (Fig. 45). Monoclonal antibody 44 has no inhibitory effect in cell adhesion and spreading and does not promote DNA replication and transdifferentiation (Table VI). Lectins are a group of proteins that specifically recognize mono and disaccharides (Liz and Sharon, 1991). The results observed with mAb 19 suggest that carbohydrates are involved in the ECM-ligand-receptor complex which communicates cell adhesion, spreading, and stability of TABLE VI Initiation of DNA Synthesis by Treatment of Isolated Striated Muscle with rnAbsa

Treatment mAb 19 mAb 44 mAb 93 Control

Time of treatment (min) 60-70 60-70 60

-

n

tot

% Labeled nuclei ( f s)

18 14 8 9

2281484 221710 41128 351881

47.1 37.1 3.1 2 9.9 3.1 4.4 4.0 f 4.8

* *

“Labeling with [)H]thymidine was performed from 24 hr until fixation at 168 hr. The muscle was treated by incubation in mAb-containing hybridoma-conditioned supernatant. mAb 19 is specific for carbohydrates, mAb 44 is specific for an ECM protein, and mAb 93 specifically stains a component of the contractile system of striated muscle cells. n , number of fragments; tot, labeled nuclei to the total of nuclei examined: s, standard deviation (After Reber-Muller and Schmid, 1992).

TRANSDIFFERENTIATION IN MEDUSAE

251

FIGS.43and44 Whole-mount preparations of the hydromedusa Podocoryne carnea. The ECMspecific staining pattern with mAb 19 is presented in Fig. 43, and with mAb 44 presented in Fig. 44. Bars, 20 pm (from Reber-Muller and Schmid, 1992). FIG. 45 Immunoblotting of Podocoryne carnea proteins. Lane I , staining pattern with carbohydrate-specific rnAb 19. Lane 2, staining pattern of mAb 44. The mAb stains bands with calculated molecular weights of 220 and 105 kDa, respectively. Lanes 3 and 4, staining with carbohydrate-specific mAb 19 after chemical deglycosylation of proteins on Western blot, prior to antibody labeling. Lane 3, experimental strip: Lane 4. control. Lanes 5 and 6, deglycosylated proteins on Western blots probed with mAb 44, lane 5, experimental strip; lane 6, control. Note that mAb 19 does not stain deglycosylated blots whereas the staining pattern of mAb 44 remains unchanged indicating that mAb 44 recognizes a protein epitope (from Reber-Muller and Schrnid, 1992).

the differentiated state in striated muscle cells. Therefore, we have tested various lectins for their effects on cell adhesion, spreading, and transdifferentiation. Whereas Phaseolus- and BS-I-lectin had no effect, Con A and WGA inhibited cell adhesion and spreading on pretreated ECMs and led to initiation of DNA replication when directly applied on isolated muscle tissue (Reber-Miiller and Schmid, 1992). The effective concentrations again were correlated with transient dissociation of cells from the treated muscle fragments.

252

VOLKER SCHMID

E. DNA Replication and Transdifferentiation

Besides the proliferative aspects of cell division, it is becoming more and more apparent that specific phases of the cell division cycle, such as DNA replication, are important control elements in cell differentiation (Holtzer er al., 1979; Satoh, 1982; Brown, 1984; Newport and Kirschner, 1984). Furthermore, commitment and expression of the differentiated state are somehow related to the number of DNA replication cycles (Satoh, 1982). In specialized cell types the acquisition of stability of the differentiated state is frequently accompanied by a withdrawal from the cell division cycle (DiBerardino et al., 1984; Baserga, 1985; Blau et al., 1985; Brachet, 1985). This is also true for the striated muscle tissue of the medusa Podocoryne carnea (Schmid, 1972). Transdifferentiation studies in other systems and nuclear cloning experiments indicate that new programming of the genome in many cases requires cell cycle activity, specifically DNA replication (Gurdon er al., 1975;Okada, 1991). The role of cell activity in the medusa transdifferentiation system was investigated by using drugs that specifically inhibit DNA replication.

1. Transdifferentiation without DNA Replication In pulse-labeling experiments with [3H]thymidineit was demonstrated that destabilized muscle isolates go through several cycles of DNA replication (Fig. 46). The question was put forward whether there is a correlation between the number of cell cycles and the cell types formed by transdifferentiation. Cells can be arrested reversibly at the G,-S transition by aphidicolin, a nontoxic, reversible inhibitor of DNA polymerase a (Ikegami er al., 1979; Spadari et al., 1982). Such cells are prevented from entering the S phase, and when continuously incubated in aphidicolin and [3H]thymidine, no incorporation of label and no metaphase chromosomes were observed (Alder, 1982; Alder and Schmid, 1987). Isolated and collagenase-treated striated muscle, which has been incubated continuously in aphidicolin, demonstrates the normal pattern of transdifferentiation up to the third day, e.g., formation of flagellated smooth muscle cells. However, no other cell types (e.g., nerve cells) are formed before the isolates disintegrate after 2-3 weeks (Alder and Schmid, 1987). Formation of smooth muscle cells is therefore completely independent of DNA replication and mitosis. This confirms reports of de nouo formation of flagella and smooth muscle-like arrangement of myofilament in destabilized isolates when mitosis was inhibited with mitomycin C (Schmid, 1975). Furthermore, the first important step in this transdifferentiation system, the formation of smooth

253

TRANSDIFFERENTIATION IN MEDUSAE

lYDl 80

% labeled nuclei

40

t *O! 0

L L 2

I

I

star1 of collagenase treatrnenl

1

i3

f

5

4

Y

5

6

7

8

days without separation of layers

Y;s

wilh separaton of layers

d ;g ;se; nb

FIG. 46 DNA synthesis in collagenase-treated isolates of striated muscle cells. At various time isolates were incubated in [3H]thymidine for 2 hr. The curve represents the total of five independent experiments. Each point with error bars represents the average of at least four isolates. Bottom: first appearance of cell types (Schmid and Alder, 1984). d, digestive cells; f, flagella of smooth muscle cells; g, gland cells; nb, nematoblasts; s, smooth muscle cells; se, secretory cells, y. nerve cells (after Alder and Schmid, 1987).

muscle cells, is evidently not disturbed by these drugs. Recent observations with the smooth muscle-specific mAb sm-1 demonstrated that the antigen is already synthesized before DNA replication has started (Ch. Weber, personal communication, see Section III,C, 1 ,c). By the fifth day of continuous aphidicolin treatment, 9 of 33 isolates displayed an elongated shape with a cavity. In these cases, an outer, epidermal layer was separated from an inner, gastrodermal layer by formation of basal lamella. The same events occurred in controls that were not treated with aphidicolin (see above; Schmid and Alder, 1984). But in contrast to the controls, directional swimming was not developed in the aphidicolin-treated isolates. Other cell types, such as digestive cells, gland cells, secretory cells, and nematoblasts, did not appear in the gastrodermal layer of these aphidicolin-treated regenerates. Another cell cycle inhibitor, cytosine arabinoside, confirmed the results that have been obtained with aphidicolin (unpublished). The formation of nerve cells, the second new cell type, is completely prevented by continuous aphidicolin treatment, suggesting a dependence on events of the first cell cycle. Thus, the temporal succession of transdifferentiation events beyond the formation of smooth muscle cells is broken down. The fact that a basal lamella that separates an epidermal from a

254

VOLKER SCHMID

gastrodermal layer can be formed during continuous aphidicolin treatment (see above) demonstrates that this process does not depend on DNA replication (Alder and Schmid, 1987). Consequently the process of delamination (cell layer formation based on mitosis) can be excluded as a causal mechanism leading to separation of layers. The real mechanism is not yet known (Schmid and Alder, 1984).The number of isolated muscle fragments in which separation of layers occurred is about the same as that in isolates not treated with aphidicolin. Furthermore, these observations indicate that in this system degradation of myofibers is uncoupled from DNA replication and transdifferentiation to new cell types (Schmid and Alder, 1984, 1986). Transdifferentiation-like events without DNA replication in other systems has been recently reviewed (Beresford, 1990). It appears that this occurs mostly between cell types that are closely related to each other in ontogeny. In nontransdifferentiation systems (e.g., heterocaryons, myoblast-myotube transition) similar observations have been made (Nadal-Ginard, 1978; Bennett, 1983; Chiu and Blau, 1984).

2. Transdifferentiation with DNA Replication To investigate the effect of the first cell cycle on transdifferentiation events, isolated striated muscle fragments were incubated in aphidicolin after collagenase treatment. The drug was removed at the expected initiation time of the first S phase (Fig. 47; Alder and Schmid, 1987). This is the time when about 50% of the cells in collagenase-activated controls would have started with DNA replication. The aphidicolin-treated isolates proceeded with DNA replication (after removal of the drug) just as the controls do. If aphidicolin is then added again after the first S phase (76 hr), the cells will complete the first cell cycle and accumulate at the GI-S transition (Ikegami et al., 1979; Spadari et al., 1982). In all cases, nerve cells appeared on the fourth day. The ratio of smooth muscle cells to nerve cells was about 1:l (51:49%) as in the controls not treated with aphidicolin. However, no directional swimming (polarity) and no other cell types, typical of regenerate formation, were expressed in those isolates [8 of 32 (21%)] where formation of an epidermal and a gastrodermal layer was observed (Alder and Schmid, 1987). When aphidicolin was used to allow one cell cycle, only nerve cells and smooth muscle cells were formed (Alder and Schmid, 1987). The time of addition of the drug is not very critical. We can add it in G,, M, or GI phase to be sure that we do not interrupt the S phase and that the cells accumulate again at the next GI-S transition and complete one cell cycle. Regeneration formation was incomplete after one cell cycle despite the fact that separation of the layers sometimes occurred. The lack of polarity

255

TRANSDIFFERENTIATION IN MEDUSAE 100 1

80 60

%labeled nuclei 40 ..

20

0

1 1

2

3

4

f

Y

s

S

5

6 days

FIG. 47 Transdifferentiation with one cell cycle. Isolated striated muscle was incubated in aphidicolin. At the expected initiation time of the first S phase, the drug was removed (56 hr). After completion of the first S phase, aphidicolin was added again (76 hr). [3H]Thymidine was continuously present. Flagellated smooth muscle cells (s) appear during the first round of DNA replication (data points with error bars). Nerve cells (y) are first seen on the fourth day. The bar (bottom) indicates the time when aphidicolin was removed (from Alder and Schmid, 1987).

(directional movement) suggests the need for additional mechanisms that are bound to an additional cell cycle. The question whether two cell cycles are sufficient for complete regenerate formation was examined by again using aphidicolin. Aphidicolin was applied after collagenase treatment and then removed from the isolates at the expected initiation time of the first S phase. Provided that the drug is added after completion of the second S phase, the cells will complete two cell cycles. The labeling rate of nuclei is comparable to the labeling rates of controls not treated with aphidicolin. In all cases, nerve cells appeared on Day 4 (Alder and Schmid, 1987). The ratio of smooth muscle cells to nerve cells was about !:I (53:47%), as that in untreated controls. By the fifth day, all isolates [5 of 25 (20%)] that had formed a gastrodermal layer expressed directional swimming (polarity). In the gastrodermal layer, the whole inventory of cell types necessary to form a functional regenerate appeared (Alder and Schmid, 1987). Two cell cycles without separation of layers produced no other cell types than smooth muscle and RF-amidepositive nerve cells. This occurs only when separated layers are found. In this case polarity is expressed; digestive cells, gland cells, secretory cells, and nematoblasts are differentiated, and a regenerate is formed. Aphidi-

256

VOLKER SCHMID

Colin treatment in this experiment apparently had no effect on the formation of regenerates that are indistinguishable from controls. With regard to cell types and anatomical arrangement, the regenerates seem to be normal. These observations demonstrate strict correlations between cell cycle events and differential gene expression. The same seems to be true for transdetermination of Drosophila imaginal disks (Hadorn, 1966; Haynie and Bryant, 1976; Szabad et al., 1979). In Wolffian lens regeneration (see II,B,2) four complete cycles are required before the descendants of an iris epithelial cell become definitely dedifferentiated and it is estimated that an additional two cell cycles are needed before they become irreversibly committed to lens cells (Pritchard, 1986; Yamada, 1989). Preceeding cell divisions and/or DNA replication are not essential in all instances of transdifferentiation. However, when changes occur between distinct cell types with ontogenically remote origins, cell cycles seem to play an important role (Okada, 1991).

IV. Concluding Remarks

The transdifferentiation processes observed in medusae clearly demonstrate that well-differentiated animal cells can retain the ability to change cellular commitment and to form various new cell types that can be organized into a functional regenerate. The only possible alternative explanation of our findings would be that the original material was contaminated by cells of other types. This hypothesis can be excluded on the following grounds: 1 . Previous authors have concluded that tissue layers as prepared for these experiments are free of contamination cell types (Kuhn, 1914; Weiler-Stolt, 1960; Frey, 1968; Bolsterli, 1977). 2. Contaminating cells can be reliably identified and controlled for through the use of the DAPI method (Schmid et al., 1982). This procedure is used throughout our experiments. 3. When contaminating cells were known to be present in the isolated muscle there was no effect on the rate of formation of endoderm or a regenerate (Schmid and Alder, 1984). 4. In those cases in which degradation of striated myofibers was delayed, these can be found freely in the cytoplasm of the newly formed cell types (see Figs. 12; 22-26). Since the striated myofibers are not contained in vacuoles, and therefore not phagocytosed into the cell, the new cell

TRANSDIFFERENTIATION IN MEDUSAE

251

types must originate from striate muscle. The contamination argument thus becomes irrelevant. Initiation of transdifferentiation follows the signal transduction pathway in which the extracellular environment, the cell membrane, and the cytoplasm are involved. Changes in the ECM-ligant-cell receptor complex disturb the interplay between the cell and its substrate (experiments with ECM grafts, enzyme treatments). This leads to changes in cell shape and, as known from vertebrate systems, activates DNA replication and transdifferentiation in medusa cells. Destabilization, however, also occurs if elements involved in signal transduction on the membrane/cytoplasms level are affected. Protein kinase C seems to play a key role, as is indicated by experiments with TPA, DiC8, mezerein, and the corresponding inhibitors of PKC. Zn uiuo, however, the first activating signal seems to originate from tissue that are involved in covering the wound area and at places where the stabilizing interplay between the cells and their substrate is disturbed. With the exception of the formation of smooth muscle, cell cycles are necessary for this in uitro transdifferentiation to take place. The demonstrated transdifferentiation of striated muscle is unparalled in animals and shows that a tissue layer composed of fully differentiated cells can undergo pluripotent transdifferentiation and retain the capacity for morphogenesis. This implies that the striated muscle can express, at least in part, the developmental potential of the fertilized egg. In many animals ontogenesis is understood as a time-dependent cascade of events, in which genome, cytoplasmatic determinants, and extracellular information finally shape a functional organism. It is difficult to understand how a functionally specialized somatic cell, the final product of such a complicated line of events, can maintain its developmental potential, but the experiments described here show that in medusae this is indeed the case. In interpreting our observations the evolutionary background of the animal has to be considered. Although medusa belong to the “primitive” phylum Coelenterata, they are well-differentiated animals. They have developed a multitude of specialized cell types, in which gene products are stably expressed. Yet, these animals have a great ability to regenerate, very likely based on the transdifferentiation potential demonstrated here.

Acknowledgments I thank Barbara Rupp for generous assistance in preparation of the manuscript. Furthermore, I thank Dr. P. Schuchert and Susanne Reber-Mutter for correcting the manuscript. The work discussed in this chapter was supported by different grants from Swiss Science Foundation.

258

VOLKER SCHMID

References Achermann, J. (1980). In “Developmental and Cellular Biology of Coelenterates” (P. Tardent and R. Tardent, eds.), pp. 273-279. Elsevier, Amsterdam. Acheson, A., Edgar, D.. Timpl, R., and Thoenen. H. (1986). J. Cell B i d . 102, 151-159. Agata K . , and Eguchi, G. (1984). Deu. Growth Dgf. 26, 385. Agata, K . , Yasuda, K., and Okada, T. S. (1983). Deu. B i d . 100, 222-226. Alder, H. (1982). Thesis, University of Zurich, unpublished. Alder, H., and Schmid, V. (1987). Deu. B i d . 124, 358-369. Anderson, D. J. (1989). Trends Genet. 5 , 174-179. Anderson, H. (1990). Experientia 46, 2-13. Baguiih, J., and Romero, R. (1981). Hydrobiologia 84, 181-194. Baguiia, J., Sa16, E., and Auladell, C. (1989). Deuelopment 107, 77-86. Baserga, R. (1985). “The Biology of Cell Reproduction.” Harvard Univ. Press, Cambridge, London. Beck, K., McCarthy, R. A., Chiquet, M., Masuda-Nakagawa, L., and Schlage, W. K. (1989). I n “Cytoskeletal and Extracellular Proteins: Structure, Interactions and Assembly,” 2nd EBSA Symposium, Springer Series in Biophysics, Vol. 3, pp. 102-105. Springer-Verlag, New York. Bennett, D. C. (1983). Cell (Cambridge, Mass.) 34, 445-453. Ben-Ze’ev, A. (1980). Cell (Cambridge, Mass.) 21, 365-377. Ben-Ze’ev, A. (1991). Bio Essays 13, 207-212. Beresford, W. A. (1990). Cell Dgf. Deu. 29, 81-94. Bergquist, P. R. (1978). “Sponges.” Hutchinson University Library, London. Bissell, M. J., and Barcellos-Hoff, M. H. (1987). J . Cell Sci. Suppl. 8, 327-343. Blau, H. M., Pavlath, G. K., Hardeman, E. C., Chiu, C. P., Silberstein, L., Webster, S. G., Miller, S. C., and Webster, C. (1985). Science 230, 758-766. Blumberg, P. M. (1988). Cancer Res. 48, 1-8. Bode, H. R. (1992). Trends Genet., in press. Bolsterli, U. (1977). J . Morphol. 154, 259-290. Brachet, J. (1985). “Molecular Cytology,” Vol. I , “The Cell Cycle.” Academic Press, Orlando. Brown, D. D. (1984). Cell (Cambridge, Mass.) 37, 359-365. Caplan, A. I., and Ordahl, C. P. (1978). Science 201, 120-130. Castagna, M. (1987). Biol. Cell. 59, 3-14. Chapman, G. (1966). In “The Cnidaria and Their Evolution” ( J . W. Rees, ed.), pp. 147-172. Academic Press, New York/London. Chiquet-Ehrismann, R. (1991). Curr. Opinion Cell Biol. 3, 800-804. Chiu, C. P., and Blau, H. M. (1984). Cell (Cambridge. Mass.) 37, 879-887. Coe, I. R. (1987). Thesis. Univ. of Victoria, B.C. Canada, pp. 1-103. Colucci (1891). R . Accad. Bologna, Sez. Sci. Nar. (Ser. 5 ) 1, 167-203. Couturier, A., Bazgar, S., and Castagna, M. (1984). Biochem. Biophys. Res. Commun. U1, 448-455. Davis, L. E., Burnett, A. L., Haynes, J. F., and Muman, V. R. (1966). Deu. Biol. 14, 307-329. Denton, E. J. (1963). Endeavour 22, 85-87. DiBerardino, M. A., (1980). Differentiation 17, 17-30. DiBerardino, M. A., (1988). Cell DifJ Deu. 25(Suppl.), 129-136. DiBerardino, M. A., Hoffner, J. N., and Etkin, L. D. (1984). Science 224, 946-952. Doctrow, S. R., and Folkman, J. (1987). J . Cell B i d . 104, 679-687. Edgar, B. A., and Schubiger, G. (1986). Cell (Cambridge, Mass.) 44, 265-372, 871-877.

TRANSDIFFERENTIATION IN MEDUSAE

259

Eguchi, G. (1988). Cell D$f. Dev. 25(Suppl.), 147-158. Eguchi, G., and Okada, T. S. (1973). Proc. Natl. Acad. Sci. U . S . A . 70, 1495-1499. Ekblom, P. (1989). FASEB J . 3, 2141-2150. Franke, W. W. (1987). Cell (Cambridge, Mass.) 48, 3, 4. Freeman, G. (1981). Wilhelm Roux’ Arch. 190, 123-125. Frey, J. (1968). Wilhelm Roux’ Arch. Entw. Mech. Org. 160, 428-464. Fukuda, H., and Komamine, A. (1985). I n “Cell Culture and Somatic Cell Genetics of Plants” (I. K. Vasil, ed.), Vol. 2, pp. 149-212. Academic Press, New York/London. Gehring, W. (1967). Dev. Biol. 16, 438-456. Gladfelter, W. B. (1972). Helgolander Wiss. Meeresunters. 23, 38-79. Glimelius, G., and Weston, J. (1981). Dev. Biol. 82, 95-101. Gremigni, V. (1988). Zoolog. Sci. 5, 1153-1163. Grimmelikhuijzen, C. J. P. (1983). Histochemistry 78, 361-381. Grimmelikhuijzen, C. J. P., and Spencer, A. N. (1984). J . Comp. Neurol. 230, 361-371. Gurdon, J. B., Laskey, R. A., and Reeves, 0. R. (1975). J. Embryol. Exp. Morphol. 34, 93-1 12.

Hadorn, E. (1966). I n “Mayor Problems in Developmental Biology,” 25th Symp. SOC.Dev. Biol. (M. Locke, ed.), pp. 85-104. Academic Press, New York. Haynes, J. F., and Burnett, A. L. (1963). Science 142, 1481-1483. Haynie, J. L., and Bryant, P. J. (1976). Nature (London) 259, 659-662. Henshaw, G. G., O’Hara, J. F., and Webb, K. J. (1982). Symp. Br. SOC.CellBiol. 4,23 1-251. Holtzer, H., Rubinstein, N., Fellini, S., Yeoh, G., Chi, J. Birnbaum, J., and Okayama, M. (1979). Q. Rev. Biophys. 8, 523-557. Huang, C. L., and Ives, H. E. (1987). Nature (London) 329, 849, 850. Ide, H. (1986). I n “Current Topics in Developmental Biology,” Vol. 20, “Commitment and Instability in Cell Differentiation” (T. S. Okada and H. Kondoh, eds.), pp. 79-87. Academic Press, Orlando/London/Tokyo. Ikegami, S., Shonan, A., Oguro, M., Nagano, H., and Mano. Y. (1979). J . Cell Physiol. 100, 439-444.

Ingber, D. E. (1990). Proc. Natl. Acad. Sci. U . S . A . 87, 3579-3583. Ingber, D. E., and Folkman, J. (1989a). J. Cell Biol. 109, 317-330. Ingber, D. E., and Folkman, J. (1989b). Cell (Cambridge, Mass.) 58, 803-805. Inoue, K., Ozato, K, Kondoh, H., Iwamatsu, T.. Wakamatsu, Y.,Fujita, T., and Okada, T. S. (1989). Cell D$J Dev. 27, 57-68. Itoh, Y., and Eguchi, G. (1986). Cell D.$J 18, 173-182. Jeffery, W. R. (1985). Cell (Cambridge, Mass.) 41, 11, 12. Kafatos, F. C. (1972). I n “Current Topics in Developmental Biology” (A. A. Moscona and A. Monroy, eds.), Vol. 7, pp. 125-191. Academic Press, New York/London. Kafatos, F. C., and Williams, C. M. (1964). Science 146, 528-540. Kase, H.,Iwahashi, K., Matsuda Y., Yamada, K., Takahashi, M., Murakata, C., Sato, A., and Kaneka, M. (1987). A . Rev. Cell Biol. 108, 1079-1091. Kinter, C. R., and Brockes, J. P. (1984). Nature (London) 308, 67-69. Kondoh, H., Katoh, K., Takahashi, Y., Fujisawa, H., Yokoyama, M, Kimura, K, Katsuki, M, Saito, M, Nomura, T., Hiramoto, Y, and Okada, T. S. (1987). Deu. Biol.120, 177-185. Kornblihtt, A. R., and Gutman, A. (1988). Biol. Rev. 63, 465-507. Kiihn, A. (1914). Ergebn. Fortschr. 2001.87. Kurz, E., and Schmid, V. (1991). In: “Coelenterate Biology: Recent Research on Cnidaria and Ctenophora,” Symp. 5th Int. Conf. Coelenterate Biol. Hydrobiologia (R. B. Williams, P. F. S. Cornelius, R. G. Hughes, and E. A. Robson, eds.), pp. 11-17. Kluwer Academic. Kurz, E., and Schmid, V. (1993). In preparation Lassar, A. B., Buskin, J. N., Lockshon, D., Davis, R. L., Apone, St., Hauschka, St. D., and Weintraub, H. (1989). Cell (Cambridge, Mass.) 58, 823-831.

260

VOLKER SCHMID

Lis, H., and Sharon, N. (1991).Curr. Opinion Struct. Biol. 1, 741-749. Lowell, R. D., and Burnett, A. L. (1969).Biol. Bull. 137, 312-320. Mackay, N. C. (1969).C o m p . Biochem. Physiol. 30,481-488. Mackie, G . 0.(1975).J . Neurobiol. 6, 357-378. Mackie, G. 0.(1980).Trends Neurosci. January, 13-16. Mackie, G. O.,and Singla, C. L. (1975).J . Neurobiol. 6, 339-356. Mackie, G. O.,Stell, W. K., and Singla, C. L. (1985).Acta Zool. 66, 199-210. Marder, E.,Calabrese, R. L., Nusbaum, M. P., and Trimmer, B. (1987).J . C o m p . Neurol. 259, 150-163. Miura, S., and Kimura, S. (1985).J. Biol. Chem. 260, 15,352-15,356. Meins, F. (1982).In “Molecular Biology of Plant Tumors” ( J . Schell and E. Kahl, eds.), pp. 233-264. Academic Press, New York. Meins, F. (1986).In “Current Topics in Developmental Biology” (T. S. Okada and H. Kondoh, eds.), pp. 383-396. Academic Press, Orlando/London/Tokyo. Meins, F. (1989).Annu. Reu. Genet. 23, 395-408. Mochii, M., Takeuchi, T., Kodama, R., Agata, K., and Eguchi, G. (1988a).Cell. Diff. 23, 133-42. Mochii, M., Agata, K., Kobayashi, H., Yamamoto, T. S., and Eguchi, G. (1988b).Cell Difj 24,67-74. Mochitate, K., Pawelek, P., and Grinnel, F. (1991).Exp. Cell Res. 193, 198-207. Montesano, R., and Orci, L. (1985).Cell (Cambridge, Mass.) 42, 469-477. Moscona, A., Brown, M., Degenstein, L., Fox, L., and Soh, B. M. (1983).Proc. Natl. Acad. Sci. U.S.A. 80,7239-7243. Nadal-Ginard, B. (1978). Cell (Cambridge, Mass.) 15, 858-864. Namewirth, M. (1974).Deu. Biol. 41, 42-56. Nathanson, M. A. (1986).I n “Current Topics in Developmental Biology,” Vol. 20,“Commitment and Instability in Cell Differentiation” (T. S. Okada and H. Kondoh, eds.), pp. 39-62. Academic Press, Orlando/London/Tokyo. Nathanson, M. A., Hilfer, S . R., and Searls, R. L. (1978).Deu. Biol. 64,99-117. Newport, J. W.,and Kirschner, M. W. (1984).Cell (Cambridge, Mass.) 37, 731-742. Neidel, J. E.,Kuhn, L. J., and Vandenbark, G. R. (1983).Proc. Natl. Acad. Sci. U.S.A. 80, 36-40. Nishizuka, Y. (1984).Nature (London) 308, 693-698. Nishizuka, Y. (1988).Nature (London) 334, 661-665. Normandin, K. K. (1960).Science 132, 678. Okada, T. S. (1983).Cell D$f. 13, 177-183. Okada, T. S. (1991).“Transdifferentiation.” Clarendon Press, Oxford. Osborne, D. J., and McManus, M. T. (1986).In “Ethylene: Biochemical, Physiological and Applied Aspects” (Y. Fuchs and E. Chalutz, eds.), pp. 221-230. Nijhoff/Dr. W. Junk, The Hague. Pritchard, D. J. (1986).“Foundations of Developmental Genetics.” Taylor & Nancis, London/Philadelphia. Rao, M. S., Yeldandi, A. V., and Reddy, J. K. (1990).Cell D$J 29, 155-164. Reber-Miiller, S., and Schmid, V. (1992).Submitted. Rozengurt, E. (1986).Science 234, 161-166. Satoh, N. (1982).Differentiation 22, 156-163. Schlage, W.K. (1988).Biol. Chem. Hoppe-Seyler369,359-363. Eur. J . CellBiol.47,395-403. Schmid, V. (1972).Wilhelm Roux’ Arch. 169, 281-307. Schmid, V. (1974).Am. Zool. 14, 773-781. Schmid, V. (1975).Exp. Cell Res. 94, 401-408. Schmid, V. (1978).Deu. Biol. 64,48-59.

TRANSDIFFERENTIATION IN MEDUSAE

261

Schmid, V. (1979). Ann. SOC.Fr. Biol. Deu. pp. 35-38. Schmid, V. (1988). Cell D i f f .22, 173-182. Schmid, V., and Alder, H. (1984). Cell (Cambridge, Mass.) 38, 801-809. Schmid, V., and Alder, H. (1986). I n “Current Topics in Developmental Biology” (A. A. Moscona, A. Monroy, eds.), Vol. 7, pp. 117-135. Academic Press, San Diego. Schmid, V., Baader, C., Bucciarelli, A., and Reger-Muller, S. (1992a). Submitted. Schmid, V., Schuchert, P., Piraino, S., and Boero, F. (1992b).Scienria Marina,56, 131-136. Schmid, V., and Bally, A. (1988). Deu. B i d . 129, 573-581. Schmid, V., Bally, A., Beck, K., Haller, M., Schlage, W. K., and Weber, Ch. (1991). In “coelenterate Biology: Recent Research on Cnidaria and Ctenophora,” Symp. 5th Int. Conf. Coelenterate Biol. (R. B. Williams, P. F. S. Cornelius, R. G. Hughes, and E. A. Robson, eds.), pp. 3-10. Kluwer Academic. Schmid, V., Schmid, B., Schneider, B., Stidwill, R., and Baker, G. (1976). Wilhelm Roux’ Arch. 179, 41-56. Schmid, V., Weber, Ch., and Keller, D. (1984). Wilhelm Roux’ Arch. 193, 36-41. Schmid, V., Wydler, M., and Alder, H. (1982). Deu. Biol. 92, 476-488. Smid, I. (1984). Thesis. University of Zurich, Rieker & Partner AG. Spadari, S., Sala, F., and Pedrali-Noy, G. (1982). TIBS, January, 29-32. Steen, T. P. (1968). J. Exp. Zool. 167, 49-71. Steward, F. C., Maples, M. O., Kent, A. E., and Holsten, R. D. (1964). Science, 143,20-27. Streuli, C. H., and Bissell, M. J. (1990). J . CellBiol. 110, 1405-1415. Szabad J., Simpson P., and Nothiger, R. (1979). J . Embryol. Exp. Morphol. 49, 229-241. Tardent, P., and Schmid, V. (1972). Exp. Cell Res. 72, 265-275. Tillet-Barret, E., Franc, J-M., Franc, S., and Garronne, R. (1992). FEBS, Eur. J . Biochem., in press. Timpl, R. (1989). Eur. J. Biochem. 180, 487-502. Tobler, H. (1986). In “Results and Problems in Cell Differentiation,” Vol. 13, “Germ Line-Soma Differentiation” (W. Henning, ed.), pp. 1-69. Springer-Verlag, Berlin/Heidelberg. Topscott, S. J., Davis, R. L., Thayer, M. J . , Cheng, R. F., Weintraub, H, and Lassar, A. B. (1988). Science 242,405-411. Trelstad, R. L. ed. (1984). “The Role of the Extracellular Matrix in Development.” A. R. Riss, New York. Unsicker, K., Krisch, B., Otten, J., and Thoenen, H. (1978). Proc. Narl. Acad. Sci. U . S . A . 75, 3498-3502. Vasil, J. K., and Vasil, V. (1972). In Vifro8, 117-127. Watson, P. A. (1991). FASEB J . 5, 2013-2019. Weber, Ch. (1981). J . Morphol. 167, 313-331. Weber, Ch. (1989). Cell Tissue Res. 255, 275-282 Weber, Ch., and Schmid, V. (1984). Exp. Cell Res. 155, 149-158. Weber, Ch., and Schmid, V. (1985). Tissue Cell 17, 81 1-822. Weber, Ch., Alder, H., and Schmid V. (1987a). Cell D$f. 20, 103-115. Weber, Ch., Kurz, E., and Schmid, V. (1987b). Tissue Cell 19, 757-771. Weiler-Stolt, B. (1960). Wilhelm Roux’ Arch. 152, 389-454. Westfall, J. A., and Kinnarnon, J. C . (1978). J . Neurophysiol. 7 , 365-379. Williams, R. G., and Dockray, G. J . (1983). Brain. Res. 276, 213-229. Wolff, G. (1895). Archivfiir Mikroskoische Anatomie und Entwicklunsmechanik der Organismen 1, 380-390. Yamada, T. (1989). Int. Rev. Cyfol. 117, 215-255. Yamada, T., Iwahashi, K., and Kase, H. (1987). Biochem. Biophys. Res. Commun. 144, 35-40. Zanetti, N. C., and Salursh, M. (1984). J. Cell Biol. 99, 115-123.

This Page Intentionally Left Blank

Symplast as a Functional Unit in Plant Growth Kiyoshi Katou' and Hisashi Okarnotot * Biological Laboratory, College of General Education, Nagoya University, Nagoya 464-0 1, Japan

t Biology Department, Graduate School of Integrated Science, Yokohama City University, Kanazawa, Yokohama 236, Japan

I. Introduction

The cell-wall capsule and well-developed vacuole are characteristic of plant cells. These must have played important roles in expanding the habitat of plants from the paleozoic sea toward the dry land. The solutes accumulated in the vacuole keep cellular water potential low by lowering osmotic potential ( -RTCi) and enabling osmotic water uptake. The uptake of water by plant cells stresses the wall capsule. This generates intracellular hydrostatic pressure (Pi) and keeps the cell turgid. Increase in Pi elevates cellular water potential (qi= Pi- RTC') and opposes water entry. Solute accumulation, of course, primarily contributes to the adjustment of water potential of the cell by generating the potential ability of cellular water uptake. However, Pisimultaneously plays an indispensable role in the adjustment of cellular water potential. This adjustment occurs very rapidly because Pi intimately depends on the mechanical properties of the cell wall. Cooperative interaction of these two opposing forces (Piand - R T C ) adequately adjusts the cellular water potential against perturbation of the extracellular water potential. Thus, plant cells with the cell-wall-vacuole system can overcome cellular burst and/or wilting even in fresh and/or brackish water environment where the water potential is apt to vary rapidly. It must have been very difficult, however, for plant cells to migrate from the aquatic environment to the dry land even though they had acquired the cell-wall-vacuole system. Atmospheric water potential is too low compared with the typical cellular water potential of -0.2 MPa of higher plant cells because the water potential of very humid air of 90% relative humidity is - 14.5 MPa at 25°C. Rainfall may immediInrernarional Review of Cylology, Vol. 142

263

Copyright 0 1992 by Academic Press. Inc. All rights of reproduction in any form reserved.

264

KlYOSHl KATOU AND HISASHI OKAMOTO

ately elevate the extracellular water potential though. The atmospheric water potential is apt to vary greatly and rapidly. It could be said, therefore, that the water environment of the dry land is extraordinarily severe on the life of plant cells. The dry land, however, is undoubtedly advantageous for plants to perform photosynthesis and gas exchange efficiently. This could be the reason why plants expanded their habitat from the acquatic environment to the dry land. The habitat of terrestrial plants is exactly the boundary between the earth and the atmosphere. Roots absorb water and minerals from the soil, and leaves in the air engage in gas exchange and photosynthesis. The aerial parts of the plants, however, are always exposed to water deficits whereas the subterranean parts are subjected to oxygen shortage. Axial organs that consist of the vascular bundle, cortex, and epidermis are basic and common features in the structural organization of higher plants that live on dry land. The vascular bundles conduct fluid, and the gas-filled intercellular or apoplastic spaces well developed in the cortex serve as the pathway for gas conduction. In addition, the epidermis covered with cuticle, having low permeability to water, enables the plant to lose water into the air in a well-regulated manner through stomata. Thus, the plants could avoid both water deficits in the aerial parts and oxygen shortage in the subterranean parts. The extinct ancient plants, presumably the ancestors of present higher plants, might have not succeeded in migrating from the aquatic environment to dry land until they acquired an axial structure such as the stem. After the migration, it became possible for the plants to develop an organ that sustained high photosynthetic productivity in the air. In this connection, it seems very suggestive that primitive higher plants such as Rhynia, a fossil fern (psilophytes), consisted of no roots and leaves but only stems. The cell-wall-vacuole system of the cells and the axial structure of the organs must be specific structural characteristics of higher plants. These are entirely different from those of animals whose cells are always bathed in an isoosmotic body fluid within bursae. These distinct characteristics of higher plants caused their pattern of growth to be quite unique. The embryonic tissue called the apical meristem where cell division always takes place is located at the top of an axial organ and is quite active throughout the life of a plant. The elongating region neighbors basipetally on the apical meristem. Newly developed cells, by sequential cell division from the apical meristem, never move but are added to the apex. From then on, they begin to elongate along the organ axis keeping their original positions that are determined during the sequential cell division. These morphological features of plant growth (Hara, 1972) are quite different from those of animals. The elongation of plant cells is characterized both by vigorous absorption of solutes and water and by synthesis of cell-wall materials. During

SYMPLAST IN PLANT GROWTH

265

elongation, the cells increase in volume intensively due to vacuolation, after which the morphological development of the organ ceases. The mature cell is usually occupied by a large vacuole more than 90% of its volume. Cell elongation is always preceded by cell division. This is the typical process of growth in higher plants. As a result, there is a gradient in the age of the constituent cells along the organ axis. The hypocotyl of cowpea (Vigna unguiculata, previously named V . sesquipedalis) grows in the early stage of the germination period followed by epicotyl growth in the later stage; because of this, cowpea is known as an epigeal plant. The meristematic region, the elongating region, and the matured region are arranged, in order of cell age, along the axis from the cotyledonary node toward the base. In the later stage of germination when the growth of the hypocotyl ceases, this characteristic pattern of growth moves into the epicotyl. During the germination period of cowpea, the germ as an embryonic organ develops under the supply of nutrient materials from the cotyledon, which is already in a physiologically aged state, and ceases growth within the germination period (Oota, 1964). The cells in the elongating region actively attract nutrient materials from the cotyledon and may simultaneously receive regulatory signals. Such a region has been called an attraction center (Mothes, 1960; Oota, 1964), a kind of sink, on which hormonal and environmental signals of growth regulation must have great influence. In this chapter, we present our biophysical studies on the attraction center of the elongating stem based on the integrated activities of constituent cells. First, the physiological anatomy of the cowpea hypocotyl, based mostly on the results obtained through electrophysiological methods, is discussed. Then, the regulatory mechanism of the elongation growth of the hypocotyl is examined on the basis of such anatomical knowledge. II. Electrophysiological Structure of the Plant Germ Axis

We define the term “electrophysiological structure” as the semimacroscopic structure of an organ that cannot exactly be recognized without electrophysiological analysis.

A. Elongation Growth of the Stem and Surface Electrical Potential For a long time, intimate relationships have been recognized among plant elongation growth, bioelectric potentials, and respiration (Lund et al.,

266

KlYOSHl KATOU AND HISASHI OKAMOTO

1947). Biinning (1953) explained the relationship in terms of a parallelism between respiratory activity and positive surface electric potential without showing causal relationships between them. The relationship between the surface potential and growth was carefully reexamined in a germinating epigeal plant of Vigna sesquipedalis ( V . unguiculata, cowpea) by the method of surface induction (Okamoto, 1955). In this study, it was observed that near linear realtionship between positive surface potential and local growth rate appeared only in the earliest stage of germination. In the later stages, a valley of the surface potential appeared in the most rapidly elongating region of the hypocotyl (Fig. 1). Later, the valley could be observed to move acropetally with organ growth into the epicotyl. The situation was the same as that in the hypogeal plant Phaseolus angularis ( V . angularis, azuki bean) (Ichimura and Okamoto, 1958). Stem elongation requires a lot of water and nutrient solutes. Thus, in the elongation zone, the activity of cellular transport must be inevitably P D

mV

I

0

-20

-4 0

0

20

40

mm

E

FIG. 1 Distribution of electric potential on the surface of the hypocotyl of Vigna unguiculata (3-day-old etiolated seedling at 28" C). Electric potentials are referred to as the surface potential of the base of the hypocotyl. C, cotyledon; E, elongation zone of the hypocotyl.

267

SYMPLAST IN PLANT GROWTH

high. The valley of the distribution of surface potential may indicate the activity of cellular transport, i.e., absorption of solutes and water from the xylem fluid. A certain mechanism of generation of surface electric potential on the axial organ was proposed based on the idea of electric polarity of the constituent cells stemming from their oxidation-reduction activity (Lund er al., 1947). It gained, however, no additional proof experimentally. To understand the generation mechanism of the surface potential, electrophysiological examinations on the fine structure of axial organs are essential. B. Electrophysiological Anatomy of the Cowpea Hypocotyl

1. H+ as the Active Ion in Electrogenesis The distribution of the activity of the ion that participates predominantly in the generation of bioelectric potential must be parallel or antiparallel to that of the surface potential if the differences in electric potential on the surface of the germ axis were caused by diffusion of such ion along the organ axis (Okamoto, 1955). Further study (Katou and Okamoto, 1970) showed that the H + in the xylem sap is the only ion whose activity distribution along the germ axis (curve A in Fig. 2) has a close relationship c

PH

0

20

40

mm

FIG. 2 Distribution of pH in bleeding sap and tissue homogenate along the axis of the hypocotyl of Vigna unguiculata. (A) xylem sap; (B)phloem sap; (C) tissue homogenate. (Modified from Katou and Okamoto, 1970.)

268

KlYOSHl KATOU AND HISASHI OKAMOTO

with the potential distribution. The region where the xylem sap was the most acidic (pH 4.0) coincided with the most rapidly elongating region. In contrast to the pH of the xylem sap, the pH of the cell sap showed no regional difference but was constant (pH 6.0) throughout the hypocotyl (curve C in Fig. 2). Negative surface potential and acidic sap pH (pH 4.6) were also observed in the elongating region of bean roots of Phaseolus mungo, whereas the homogenate pH was 6.2 (Ishikawa et al., 1984). The distribution of K+ concentration in the xylem sap of V. unguiculata was decreased from the cotyledonary node toward the base. No valley was observed (Ichino et al., 1973). The distribution of surface electric potential, therefore, might be caused by diffusion of H', not in the bulk tissue of the hypocotyl, but in the specific channels of the xylem vessles. The electric potential difference, however, was kept between both cut surfaces even when the intact hypocotyl was cut into segments. Its polarity reflected the distribution pattern of the surface electric potential in the hypocotyl (Katou and Okamoto, 1970). In addition, this electric polarity immediately disappeared under anoxia (Ichino et al., 1973) and was very sensitive to both HCN and CO (Mizuno, 1978). Therefore, it is concluded that the surface electric potential is generated by the electrogenic activity of the constituent cells. The pH difference between the xylem sap and the parenchyma cells is greatest in the elongation zone; i.e., the xylem sap is more acidic by 2 pH units than the vacuolar sap (Fig. 2), and presumably by 3 pH units than the cytoplasm (Kurkdjian and Guern, 1989). This sharp pH difference must be generated by active H+ secretion from the parenchyma cells into the xylem vessels. This suggests that the electrogenic transport of H+ that determines the pH distribution within the xylem apoplast, represented as curve A in Fig. 2, plays an important role in the generation of the distribution of the surface potential along the hypocotyl axis.

2. Radial Potential Difference in the Hypocotyl Symbols used: Vp Electric potential of the parenchyma cell with reference to the rooting medium V , Electric potential of the hypocotyl surface with reference to the rooting medium V , Electric potential of the xylem vessel or apoplast with reference to the rooting medium Vps Intracellular potential with reference to the hypocotyl surface V,, Intracellular potential with reference to the xylem apoplast V,, Radial potential difference (electric potential difference between the surface and the xylem apoplast)

269

SYMPLAST IN PLANT GROWTH mV

120 0)

mE Ym

2 0

-

n M

1

I

E

n

40

mn

H

80

m

.r

c, c, m

a .r

40

0

CL

0 0

I

20

Hypocotyl l e n g t h

FIG. 3 Histogram of the radial electric potential difference along the hypocotyl axis. Open bar: respiration-dependent component; solid bar: passive component; M, mature zone; E, elongation zone; H, hook zone. (Modified from Ichino, 1978.)

Ichino (1978) found that the electric potentials of the freshly cut stumps of a hypocotyl with respect to the rooting medium (V,) showed no regional differences and that the electric potential difference between the surface (15 to 20 mm beneath the stump) and the cut stump (V,,) was always positive. He showed distinct regional differences in V,,, the pattern of which was parallel to the pattern of the surface potential distribution. The value of V,, was minimum in the elongation zone. These results indicate that the potential difference along the xylem vessels and its contribution to the distribution of V , are very small, as suggested by Oda (1955). The characteristic pattern of V,, depended on aerobic metabolism. It became flat under anoxia. The respiration-dependent component of V,, was minimum in the elongation zone but maximum in the matured zone (Fig. 3). He concluded that the radial electrogenic activity contributed to the generation of the distribution of surface electric potential and that the elongation zone was in a depolarized state. The cellular basis of this radial electrogenic activity, however, did not become evident until the electrogenic activity of the constituent cells in situ was examined using the intracellular microelectrode.

3. Cell Membrane Potential in Hypocotyl Segments Usually, microelectrode experiments are carried out under the condition that the excised plant tissue be bathed in an appropriate medium (Higin-

270

KlYOSHl KATOU AND HISASHI OKAMOTO

botham, 1973; Okamoto, 1976), the condition of which is essentially the same as that of animal and/or aquatic plant cells. The natural environment of the aerial parts of terrestrial higher plants, however, is the air. Therefore, it was difficult to understand a priori what kind of information could be obtained through microelectrode experiments on the membrane potential of the cells in situ. The membrane potential of the cells of hypocotyl segments with reference to the hypocotyl surface ( V,,) was measured with a microelectrode (Katou, 1978). There were no regional differences either in the values of V,, or in its respiration-dependent component between the hook, the elongation zone, and the mature zone. Thus, the distribution of V , cannot be explained by the axial distribution of V,, itself. In addition, no radial differences in the values of V p sand its respiration-dependent component were observed. A cluster of V,, in a certain cross-sectional area of the hypocotyl segment showed synchronized changes upon respiratory inhibition or light irradiation. Those cells seemed to behave like a giant single cell. Thus, it may be said that the parenchyma cells between the surface and the xylem vessels form a symplast connected to each other with plasmodesmata, as observed in roots and leaves (Dunlop and Bowling, 1971a; Liittge, 1971; Robards and Clarkson, 1976). The fact that the cellular electrical resistance of the cortical cells of Vigna hypocotyl (Liu et al., 1991) was only one-tenth that reported in cultured cells of single-cell aggregates (Ohkawa et al., 1981) electrophysiologically supports the symplast idea (Spanswick, 1972). Electron microscopic observations on plasmodesmata interconnecting the cells in cowpea hypocotyl (Maeda, cited in Oakmoto 1991) also supported this view. However, distinct regional differences in the sensitivity of V,, to light or CO were observed along the axis. It seems appropriate, therefore, to say that the hypocotyl cylinder consists of multiple layers of disk-like symplast penetrated by the vascular bundles (the disk symplast model) (Katou, 1978).

C. Two-Pump Hypothesis The absence of regional differences in V,,, despite the existence of regional differences in V,,, is an apparent contradiction. In addition, it seems curious that the electrogenic activity of the radial potential ( V s x )is minimum in the maximally elongating zone where transport activity should be maximum. These situations become understandable if two electrogenic ion pumps back to back are assumed. That is, one is located at the surface membrane of the disk-like symplast (the surface pump) and the other at the boundary membrane between the symplast and the xylem apoplast (the xylem pump) (Okamoto et al., 1978).

271

SYMPLAST IN PLANT GROWTH

According to the two-pump hypothesis, V,, consists of two membrane potentials of Vpsand Vpx and is represented by vsx

=

v,,

- vps

(1)

The regional differences in V,, must be caused by the regional differences in the hidden membrane potential of Vp, (Fig. 4). Of course, V,, partially depends on respiration, and its value and respiration dependency should be maximum in the elongation zone because V,, is minimum in this zone. There is no potential difference along the xylem vessels. Thus, the potential difference between two given points of the hypocotyl surface is determined by the difference in V,, at those points. The estimated polarity of the respiration-dependent component of V,, supports the idea that the xylem pump is the electrogenic proton pump that extrudes H + from the symplast into the xylem apoplast. The xylem pump must, therefore, contribute to bringing out the regional differences in the pH of the xylem sap. The three electric potentials, V,, Vps, and V,, of intact hypocotyl of V. unguiculata were measured simultaneously using the microelectrode combined with surface induction. Their electrogenic components (AV) were estimated from their reversible changes upon anoxia (i.e., AV = Vanoxia- Vair);AV,, and AVsx could be approximated by AV, and AVs, respectively, because of no regional difference in V, both in air and under anoxia (Ichino, 1978). The value and the time course of the changes

W 0

c a3

L

W

* r .W

.-0 .I-

c W .I-

0

LL

Hypocotyl a x i s FIG. 4 Scheme of the dual structure of radial electric potential difference and its relation to the surface potential distribution along the hypocotyl.

272

KlYOSHl KATOU AND HISASHI OKAMOTO mV I-10

min

0

0

-

-I

Elongating zone

\

al

L

E

L

2 Ic

-40

-40

,__.. _....--.-----,

-

.r

-a

+

m

.r

c, E

-80

-80

-1 20

-120

c,

a

, FIG. 5 Typical examples of the simultaneous recording of the anoxia responses of V,, Vp,and Vps.(Okamoto et al., 1979.)

in V, caused by anoxia were distinctly different from those in Vps (Fig. 5). Regional differences in AV, were observed despite no regional difference in AVps along the axis; AV, was maximum in the elongation zone (Table I). The fact that the response of Vpxwas different from that of Vps suggests a certain insulating structure existing somewhere in the apoplast between the cortex and the stele. The starch sheath was later reported to prevent apoplastic transport (Pearce and Penny, 1986).The most depolarized radial potential and its smallest electrogenic activity in the elongation zone

TABLE I Dual Structure of the Electrogenic Activity Responsible for the Radial Potential Difference of Vigna unguiculata

Zone Hook

Electric potential (mV) VS

VP VPS

Elongation

VS

VP VPS

Mature

VS VP V",

*

-5.8 14.0 -110.7f 11.7 -105.2 f 6.0 -17.3 6.8 -127.2 f 11.5 -109.6 f 9.2 11.4 15.4 - 117.2 20.0 -128.6 f 10.8

*

*

*

Respiration-dependent component AV (mV)

*

6.4 -26.8 36.6 f 4.1 63.6 f 7.3 -10.1 f 9.5 47.6 f 9.5 57.4 f 7.3 -44.6 f 8.1 27.3 f 4.6 72.1 9.7

*

Nore: The values given are the mean f standard deviation for each nine experiments. (Summarized from Okamoto et a / . , 1979.)

SYMPLAST IN PLANT GROWTH

273

(Ichino, 1978) are due to the counterbalance between V,, and Vps. The value of AV,/AV,, is 0.8 in the elongation zone and 0.4 in the mature zone (Okamoto et al., 1979). A transient change in V, induced by anoxia in the elongation zone is due to a large and delayed change in Vp compared with that in VPs. The smallest electrogenic activity of the radial potential in the elongation zone is only the apparent one. It indicates rather the maximum electrogenic activity, which means the maximum transport activity of the plasmalemma at the xylem-symplast boundary in the elongation zone. The two-component structure of the radial electric potential as represented by Eq. (1) (i.e., the membrane potentials back to back) had been proposed in roots (Dunlop and Bowling, 1971b; Dunlop, 1974; Bowling, 1976). It explained the back potential observed in the responses of the trans-root potential against an increase in the concentration of KCl in the rooting medium. However, the active nature of the inner membrane potential that should be attributed to the activity of the stelar cells was quite ambiguously discussed in that model. The two-pump hypothesis adequately explains the observed contradiction between the surface potential and the membrane potential of the cells in situ. The two-pump model for the hypocotyl, together with the disk symplast model, suggests that the electrogenic activity of the symplast membrane of either the surface or the xylem-symplast interface would play crucial roles in the biophysical regulation of the elongation growth of the hypocotyl. The two-pump idea postulating the stelar or xylem pump was discussed to explain radial ion transport across roots (Hanson, 1978; Liittge and Higinbotham, 1979)but with no direct experimental proof. It was not until 1983 that the xylem pump in roots was clearly recognized by electrophysiological methods (De Boer et al., 1983). The electrophysiological structural model of the hypocotyl of V. unguicufata based on both the two-pump model and the disk symplast model is diagrammatically presented in Fig. 6. A similar structure has been found in roots (De Boer et al., 1983; Ishikawa et al., 1984).

111. Role of Spatially Separated Proton Pumps in Stem Elongation

A. The Surface Pump The stimulating effect of acidic pH on plant elongation growth was reported for the first time in Pharbitis roots (HerCik, 1925). Later, it was reconfirmed as “acid growth” of Avena coleoptile (Bonner, 1934). Bonner

274

KlYOSHl KATOU AND HISASHI OKAMOTO

I

Xylem

I

Starch sheath

Epidermis

FIG. 6 Schematic illustration of the electrophysiological structure of the hypocotyl of Vignu unguiculuta.

observed that the acids increased the plasticity of coleoptiles, which resembled the effect of auxin on the plasticity of the cell wall (Heyn, 1940). The acid effect on plant elongation growth was rediscovered around the beginning of 1970s and the following hypothesis, termed the acid-growth theory, was proposed. The regulatory factor of wall extensibility in auxininduced promotion of elongation growth is H+ (Rayle and Cleland, 1970, 1972; Hager et al., 1971) that is extruded by proton pumps at the plasmalemma and driven by ATP (Hager et al., 1971). Auxin induces membrane hyperpolarization in the cells of higher plants (Etherton, 1970; Mark et af., 1974; Cleland et al., 1977; Nelles, 1977; Mizuno et al., 1980; Bates and Goldsmith, 1983; Felle et al., 1986; Ikoma and Okamoto, 1988). Elongation of the stem is mainly restricted by the peripheral cell-wall layers (Tanimoto and Masuda, 1971 ; Masuda and Yamamoto, 1972; Firn and Digby, 1977; Brummell and Hall, 1980; Kutschera and Briggs, 1987). The role of the surface pump, therefore, has been studied in detail in connection with the theory of acid growth for auxin action (Mizuno et al., 1980; Mizuno and Okamoto, 1982,1983;Katou and Ichino, 1982). Indole-3-acetic acid (IAA) aerosol, acetic acid aerosol, or CO, induces hyperpolarization of the surface membrane of the symplast, followed by a marked enhancement of elongation of the hypocotyl segments. Hyperpolarization strictly depends on respiration, and the enhanced elongation

SYMPLAST

IN PLANT GROWTH

275

significantly correlates to the degree of hyperpolarization (Mizuno et al., 1980; Katou and Ichino, 1982). Weak acids are known to induce membrane hyperpolarization and decrease cytoplasmic pH (Boron and De Weer, 1976; Thomas, 1976; Kurkdjian et al., 1978; Brummer et al., 1984; Hager and Moser, 1985; Felle and Bertl, 1986; Felle, 1987). Growth burst following anaerobiosis (Krauss and Hager, 1976; Parrish and Davies, 1977) was explained by the enhanced H+ export stemming from the lowering of cytoplasmic pH during anaerobiosis due to the formation of lactic acid (Hager, 1980). Thus, the electrogenic surface pump must be the IAAactivated proton pump that participates in the yielding of surface wall layers. In other words, the surface pump should be the proton pump suggested in the acid-growth theory of auxin action (Hager et al., 1971; Rayle and Cleland, 1977). Auxin might stimulate H' export by acidifying the cytosol (Brummer and Parish, 1983; Felle et al., 1986)or by increasing the number of the electrogenic proton pump at the plasmalemma (Hager et al., 1991).

6. The Xylem Pump It has been a matter of controversy whether the xylem pump or stelar pump really exists in the roots (Anderson, 1975; Pitman, 1977; De Boer et al., 1983; Clarkson, 1988), since Crafts and Broyer (1938) proposed the model of leaky stele (no xylem pump hypothesis) for solute transport across roots. Pitman's analyses (1977) of active solute fluxes for the loading of symplast and vessels in roots provided evidence in favor of the xylem pump. The first direct experimental proof of the xylem pump, however, was given not in roots but in hypocotyls (Okamoto et al., 1979). The role of the xylem pump in radial solute transport across roots was discussed by Hanson (1978) based on the chemiosmotic concept. The xylem proton pump was thought to provide a driving force for solute secretion into the xylem vessels. In contrast to that in roots, the xylem pump in hypocotyls must play a role in the absorption of solutes from the xylem fluid. It was very difficult, however, to access the xylem-symplast boundary experimentally until the method of xylem perfusion (Van Be1 and Reinhold, 1975) was well established (Clarkson et al., 1984; Okamoto et al., 1984; De Boer and Prins, 1985). Direct demonstration of the xylem pump as an electrogenic proton pump was performed by xylem perfusion (Mizuno et al., 1985). Simultaneous measurements of the membrane potential of the xylemsymplast boundary (Vpx)and the pH of the xylem perfusate (Fig. 7) showed that anoxia caused immediate depolarization of V,, followed by alkalization of the xylem perfusate several minutes later. Reacidification took

276

KlYOSHl KATOU AND HISASHI OKAMOTO

PH

5.5

5.4

0

20

m in

FIG. 7 Effect of anoxia on the membrane potential of the xylem-symplast boundary ( V J and the pH of the xylem perfusate of a hypocotyl segment of Vigna unguiculata. (Mizuno et a / . , 1985.)

place after the recovery of V,, due to reaeration. Anoxia-induced changes in the pH of the xylem perfusate, however, were not as great as expected from the distribution of pH in the xylem sap of intact seedlings. The values of the sap pH were shown to be 4.0 and 5.0 in the maximally elongating zone and mature zone, respectively. This is caused by the high pH buffering capacity of the cell-wall materials in the xylem apoplast (Mizuno and Katou, 1991). Thus, the xylem pump is the respiration-dependent electrogenic proton pump that extrudes H + from the symplast into the xylem apoplast. The pH distribution in the xylem saps as shown in Fig. 2 must be generated by continued H+-pumping by the xylem pump during the germination period. Perfusion experiments with excised internodes of the stem of tomato showed that the uptake of solutes from the xylem vessels was accompanied by coupled proton transport (Van Be1 and Reinhold, 1975; Van Be1 and Van Erven, 1979). In intact seedlings, the xylem pump generates the membrane potential enough to move K' from the xylem vessels into the symplast along the gradient of its electrochemical potential (Okamoto et al., 1978). Simultaneous measurements of Vpx and the Kt concentration of the xylem perfusate in excised hypocotyl segments clearly showed the participation of the xylem pump in solute absorption from the xylem fluids (De Boer et al., 1985). Anoxia caused simultaneous membrane depolarization and inhibition of K+uptake from the xylem vessels. Indole3-acetic acid or fusicoccin induced respiration-dependent membrane hyperpolarization in the xylem-symplast boundary followed by the stimulation of K+ uptake (Fig. 8). There is little doubt, therefore, that the maximum electrogenic activity of the xylem pump and the lowest pH of

277

SYMPLAST IN PLANT GROWTH

L,.._.'

u

m m," 1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

I

l

l

1

time in min. FIG. 8 Typical example of the effect of addition of indole-3-acetic acid (IAA)( lo-' mol m-)) to the perfusion solution upon Vpxand the potassium concentration of the xylem perfusate (K+out).(De Boer e r a / . , 1985.)

the xylem sap are in favor of the absorption of nutrient solutes from the xylem fluids necessary for elongation growth. Auxin applied by xylem perfusion has been shown to stimulate both the surface pump and the xylem pump simultaneously (Fig. 9) (Ikoma and Okamoto, 1988). This simultaneous enhancement of both proton pumps provides key information to the understanding of the biophysical regulation of stem elongation growth as discussed below.

IV. Lockhart Equations and Action of Auxin A. Lockhart Equations for Axial Organs

Plant cell elongation is regulated by two physically different processes. One is yielding of the cell wall and the other is water absorption. Simultane-

vm/ h 80

0

20

40

60

80 min

FIG.9 Typical example of responses of Vpxp Vpsrand growth rate to IAA ( l o - ' mol m-3). The perfusion solution was changed from the standard solution (0.25 mol rn-) CaSO,, 30 rnol rn-) sucrose, 1 mol rn-) KCI, pH 5 . 5 ) to that containing IAA at time 0. The rate of perfusion was kept at 120 pI hr-l throughout the experiment. (Ikoma and Okamoto, 1988.)

278

KlYOSHl KATOU AND HISASHI OKAMOTO

ous involvement of these physically different processes is a marked characteristic of the elongation growth. Lockhart (1965) analyzed these processes theoretically. When a plant cell elongating along its major axis is considered, the relative rate of steady elongation due to irreversible yielding of the cell wall is described by the following mechanical but empirical equation: (I/V)(dVldt) = cp(AP

-

Y)

for AP > Y.

(2)

Here, V is the cell volume, cp is the wall extensibility, AP is the turgor pressure of the cell, and Y is the yield threshold of the cell wall; (AP - Y ) can be termed as the effective turgor pressure for wall yielding. Water absorption by the cell is essentially osmosis through the cell membrane (the plasmalemma). Therefore, the relative rate of volume increase due to water absorption can be expressed by the thermodynamic equation (l/V)(dV/dt) = L(RTAC - A P ) ,

(3)

where L is the relative hydraulic conductivity; AC is the difference in osmotic concentration between the cell and the outer medium. Equations (2) and (3), often called the Lockhart or growth equations, must be simultaneous (Lockhart, 1965; Ray et al., 1972). Turgor pressure (AP)has been difficult to measure directly. Thus, the following equation, derived from the Lockhart equations [Eqs. (2) and (3)] by eliminating AP, has often been used for studying the biophysical control of elongation growth (Ray et al., 1972; Penny and Penny, 1978; Boyer, 1985; Tomos, 1985): (l/V)(dVldt) = {Lcp/(L + cp)} (RTAC - Y ) .

(4)

Much attention has been paid to how the parameters on the right-handside of Eq. (4) play their role in the regulation of elongation growth. This equation may indeed be a useful tool, but the determination of all the parameters under various growth conditions has not been so easy (Penny and Penny, 1978; Tomos, 1985). In addition, the parameters that describe physically different processes, i.e., a mechanical process and a transport process, have been mixed up in one equation. Therefore, for the analyses of plant elongation growth, Eq. (4) may not be as useful as was considered before. In addition, most of the studies have been done not on single cells but on axial organs or tissues that have definite multicellular organization. Thus, modification of the Lockhart equations should be done to make these equations suitable for describing elongation of axial organs in higher plants. Some working equation should be derived instead of Eq. (4). Elongation of axial organs in higher plants is governed by essentially the same biophysical processes as those in cell elongation. It should be

279

SYMPLAST IN PLANT GROWTH

considered, however, that the functional unit in the elongation of axial organs is not a constituent cell but a disk-like symplast. Its immediate apoplastic environment is divided by the insulating structure into two parts: one on the surface side and the other on the xylem side. Yielding of the cell wall that mechanically governs stem elongation takes place in the epidermal wall layer, and water absorption is restricted in the xylem parenchyma. Therefore, the Lockhart equations have to be slightly modified as follows:

(1/V)(dV/dt) = @.(Pi- y S ) (l/V)(dV/dt) = LS{URT(C’ - CX)

for Pi > yS -

(Pi

-

P”)}.

(5)

(6)

Here, @ is the physiological extensibility of the surface wall layer of the stem; P’ and Px are hydrostatic pressure of the cell and that of the xylem vessels with reference to the atmosphere, respectively; Y s is the yield threshold of the surface wall layer; Ls is the apparent hydraulic conductivity; u is the reflection coefficient, and Ci and Cxare the osmotic concentrations of the symplast cell and the xylem vessels, respectively. The expression “simultaneous,” as in the case of Eqs. (2) and (3) or Eqs. (5) and (6), means that the elongation growth should simultaneously fulfil the biophysical requirements for both yielding of the wall layer and absorption of water. The intracellular hydrostatic pressure (Pi),however, plays conflicting roles in both processes; i.e., it promotes wall yielding but at the same time suppresses water absorption. Therefore, it must be high enough to force the cell-wall layer to yield but low enough to keep the cellular water potential low, which is necessary for water uptake. These conflicting demands on Pi may be important specific characteristics of the elongation growth to be understood. Therefore, elimination of such an important key parameter as Pi from Eqs. ( 5 ) and (6), which has been a usual procedure in solving growth equations, may not be appropriate. Direct determination of Pi is rather necessary for understanding the state of stem elongation growth.

6. Effects of Auxin on the Mechanical Properties of Surface Wall Layers of the Stem

A great interest has been taken in auxin-induced changes in the cell-wall properties that cause plant cell elongation (Heyn, 1940; Burstrom, 1957; Cleland, 197 1 ; Rayle and Cleland, 1977; Cosgrove, 1986; Masuda, 1990). Different kinds of rheological analyses on the properties of the cell wall indicate that auxin increases the extensibility of the cell wall (Le., cp or a). The yield threshold of the cell wall ( Y s ) , however, is hardly affected by

280

KlYOSHl KATOU AND HISASHI OKAMOTO

auxin (Green and Cummins, 1974); Cleland, 1977; Cosgrove et al., 1984; Cosgrove, 1985). The yield threshold ( Y s ) is one of the in uiuo properties of the cell wall, measurements of which are carried out by reducing Pi to stop the elongation growth. Shifts in Pi were carried out by controlling the extracellular osmotic concentration (Cleland, 1977; Green and Cummins, 1974), water depletion (Meyer and Boyer, 1972; Boyer et al., 1985), or stress relaxation under a limited supply of water (Cosgrove, 1985). Either method requires a long time to determine Y . Green and Cummins (1974) altered Pi abruptly by perfusing with solutions of varying osmotic concentration but the shift of Piwas quite large (0.1 to 0.2 MPa). Living plant cells or tissues, however, may adaptively respond to the changes in Pi. It should be necessary to use a small andlor short step change in Pi for the measurement of Y to avoid unnecessarily huge perturbations that evoke artifacts. Thus, hydrostatical shifts in Piusing a pressure probe as carried out in the cells of Phycomyces (Ortega et al., 1989) would be preferable, but direct application of this method to the multicellular organs of higher plants was impossible. Ortega (1985) augmented the equation of wall yielding to describe the elastic extension of plant cells in response to the changes in Pi as

(lIV)(dVldt) = p(AP

-

Y ) + (l/&)(dP'/dt),

(7)

where E is the elastic modulus of the cell wall. Okamoto et al. (1989) and Ortega et al. (1989) independently reported mathematical treatments of Eq. (7) that expressed the physiological extensibility of the cell wall or cell-wall layer to be A{(llV)(dVldt)}lAPi. Thus, could be determined if the Pi and the relative rate of elongation were simultaneously measured before and after ajump in Pi.The Piof hypocotyl segments can be changed abruptly by changing the xylem pressure of P x either positively (Okamoto et al., 1989) or negatively (Okamoto et al., 1990) using a slightly modified system for xylem perfusion of hypocotyl segments. With the method of positive pressure jump, the yield threshold ( Y s ) could be estimated by linear extrapolation of enhanced growth. The method of positive pressure jump is favorable to successive measurements of the growth parameters of the segments under xylem perfusion, and the method of negative pressure jump is favorable to directly determine Y s . Both methods gave the same value of Y s . In the case of the hypocotyl segments of cowpea (V. unguiculata), the value of Y s determined by the method of pressure jump was found to be smaller than Pi only by several ten kilopascals (Okamoto et al., 1990; Nakahori et al., 1991), whereas previously reported values were smaller than P i by several hundred kilopascals (Meyer and Boyer, 1972; Green and Cummins, 1974; Cleland, 1977; Cosgrove, 1985). In other words, the elongation of the segments of

28 1

SYMPLAST IN PLANT GROWTH

cowpea hypocotyls is quite sensitive to perturbations of P i as can be readily understood from Eq. (5). Indole-3-acetic acid increased @ of V. unguiculata as has been repeatedly reported. However, IAA simultaneously decreased Y s significantly (ca. by 0.05 MPa) (Fig. 10) though it had no effect on P i (Okamoto et al., 1990; Nakahori et al., 1991). The Y s once decreased by IAA was not affected by anoxia. Thus, the IAA-enhanced growth is under the control of both the wall extensibility and yield threshold. Time courses of the changes of the growth parameters just before and after IAA-induced enhancement, which were measured in hypocotyl segments of V. unguiculata by the method of positive pressure jump, are shown in Fig. 11. An adjustment of Y s was also observed in the recovery growth of the excised segments of cowpea hypocotyls against osmotic stress (Liu et al., 1989). Thus the yield threshold of the surface wall layers ( Y s ) of the hypocotyls of cowpea is never constant but adjustable as observed in single cells of Nitella (Green et al., 1971) or Phycomyces (Ortega et al., 1989). An adjustable wall yield threshold has been reported in pea stems whose elongation was retarded by an inhibitor of gibberellin synthesis (Cosgrove and Sovonick-Dunford, 1989)or in growing bean leaves (Van Volkenburgh and Cleland, 1986). The auxin-insensitive yield threshold previously reported for coleoptiles (Green and Cummins, 1974; Cleland, 1977) or stems (Cosgrove, 1985)must be the limit value of Y s below which any adjustment is impossible. In the light of recent knowledge, it is concluded that auxin-induced enhancement of wall yielding of the stem is driven by both increase in umlh 400

+ a, m L

r

c,

200

z

a

0

-83

-60

-40

-20

0

20

Xylem pressure ( P x ) in kPa

FIG. 10 Typical examples of the linear relationships between the rate of elongation growth of

hypocotyl segments and the xylem pressure, summarized from the successive pressure jump experiments in either the absence or the presence of IAA. The intersections of the lines with the abscissa indicate the relative positions of adjustable yield threshold. The origin corresponds to the intracellular pressure of 0.6 MPa. The affixed times (h) indicate the times elapsed after application of IAA. (Okamoto et al., 1990.)

282

KlYOSHl KATOU AND HISASHI OKAMOTO

kPa

100

-

“s, I

’”&

50-f

I

-

if

3Omin

FIG. 11 Eflect of IAA on the growth parameters. The effective turgor is represented under the standard perfusion pressure (10 kPa). Using the pressure jump method, we cannot accurately determine the values of @ and effective turgor while the growth rate is rapidly changing. Therefore, the calculated value of @ was overestimated and that of the effective turgor was underestimated. These estimates are represented by open symbols. Vertical bars show SD of 10 to 12 experiments. (Nakahori et a / . , 1991.)

physiological extensibility and decrease in yield threshold. Now, we can draw one of Lockhart’s equations of wall yielding on the ( I / V ) ( d V / d t )- P i plane (see Fig. 12, lines 1 and 2) based on experimental measurements.

C. Water Relations during Rapid Enhancement of Water Absorption by Auxin in the Stem

The mechanism of auxin-induced enhancement of water absorption has greatly attracted the attention of plant-growth physiologists (Galston and Purves, 1960; Penny and Penny, 1978; Boyer and Wu, 1978; Cosgrove and Cleland, 1983b), since the auxin-induced enhancement of water uptake and its dependency on aerobic metabolism were found in excised discs of potato tuber tissues (Reinders, 1938; Hackett and Thimann, 1952). However, it has been a matter of controversy. Possible effects of auxin on the growth parameters on the right-hand-side of Eq. (4)have repeatedly been examined. The equation shows that increase in the wall extensibility

SYMPLAST IN PLANT GROWTH

283

and/or decrease in the wall yield threshold induce the enhancement of water uptake. It is possible that increase in hydraulic conductivity of the cell membrane (L,) and/or solute concentration of the cell sap (C') also induce the enhancement of water uptake. Increase in @ and decrease in Y s induced by auxin were shown to take place as discussed in the previous section. An auxin-induced rapid increase in C' was also reported by Yoda and Ashida (1961) but was not further confirmed (Penny er al., 1972; Cosgrove and Cleland, 1983b). It has been shown that C' rather decreases during auxin-induced growth (Ordin et al., 1956; Kholdebarin and Oertli, 1977; Boyer and Wu, 1978; Stevenson and Cleland, 1981; Kutschera and Schopfer, 1986). The cellular hydraulic conductivity (L,) has been shown, by various methods, to remain unchanged upon application of auxin (Galston and Purves, 1960; Penny and Penny; 1978; Cosgrove and Cleland, 1983b). Boyer and Wu's conclusion (1978),using a psychrometer, that auxin increases the hydraulic conductivity of auxin-sensitive hypocotyl tissue may be misleading because of their disregard of the physiological fine structure of the hypocotyl. If auxin causes an increase in wall extensibility (a)and a decrease in the yield threshold of the wall ( Y S ) without increasing C'and/or Ls, P i should decrease upon the application of auxin [Eq. (6)]. Thus, the behavior of Pi would provide key information on what kind of regulation of water transport is active in elongating plant cells or symplasts.

D. Diagrammatic Representation of Lockhart Equations and the Action of Auxin The effects of auxin on various growth parameters become rather understandable if the Lockhart equations are graphically solved by plotting (l/V)(dV/dt)againt AP or Pi (Katou and Furumoto, 1986a,b). The elongation of the stem can be expressed by the diagrammatic representations of Eqs. ( 5 ) and (6) on the (l/V)(dVldt)-Piplane (Fig. 12). The process of wall yielding described by Eq. ( 5 ) is represented by straight lines marked with Arabic numerals. The process of water uptake described by Eq. (6) is represented by broken lines marked with Roman numerals. Each parameter in the equations is assumed not to depend on Pi in this case. The gradients of the lines of Eqs. ( 5 ) and (6) correspond to the apparent wall extensibility (@) and the apparent hydraulic conductivity ( L s ) , respectively. The x-axis intercepts of the lines of wall yielding and water absorption represent the yield threshold ( Y s ) and RT(C' - Cx) + Px,respectively. The point of intersection of both lines represents a certain steady state of the elongation growth.

204

KlYOSHl KATOU AND HISASHI OKAMOTO

/

/ Pi Internal Cell Pressure (Pa) FIG. 12 Diagrammatic representation of the growth equations of growing axial organ of higher plants. Lines with Arabic and Roman figures are defined by Eqs. (5) and (6). Points a and b represent the states of stationary growth in the presence and absence of IAA, respectively. Point c represents the state under anoxia. Point d represents the state of IAA-enhanced growth without active promotion of water uptake.

Steady state elongation before application of auxin is represented by the intersection b (Fig. 12). The effects of auxin on the yielding properties of the surface wall layer are diagrammatically represented by a downward shift of the x-axis intercept of line 2 together with increasing gradient; i.e., line 2 moves to line 1 upon the application of auxin. No change in the parameters for water transport cause line I1 to remain unchanged. Thus, a new steady state of auxin-promoted elongation can be represented by intersection d. These changes should result in a decrease in Pi (Burstrom, 1971; Ray et al., 1972; Cosgrove, 1981; Boyer et al., 1985; Tomos, 1985). Direct measurements of Piusing a micro-pressure probe, however, showed that Pi did not decrease during auxin-enhanced rapid elongation of pea stems (Cosgrove and Cleland, 1983b). This finding was confirmed later in segments of cowpea stems by continuous measurements of Piduring auxininduced promotion of stem elongation (Nakahori et al., 1991). The values of Piof the cells in the elongation zone of wheat roots were nearly constant, showing no causal correlation with the rates of elongation growth (Pritchard et al., 1987). In addition, no changes in Pi of the cells of mustard hypocotyls were observed upon initiation of phototropic curvature in both the shaded and the illuminated sides (Rich and Tomos, 1988). Contrary to the above expectation, a decrease in Pi was observed when the growth was retarded by respiratory inhibition (Cosgrove and Cleland, 1983b; Nakahori et al., 1990, 1991). No radial differences in Pi or Ci were shown in stems (Cosgrove and Cleland, 1983b; Rich and Tomos, 1988; Nakahori

SYMPLAST IN PLANT GROWTH

205

et al., 1991). These indicate that the water potential of the symplast remains unchanged during the rapid enhancement of water uptake induced by auxin but decreases when the growth is retarded by respiratory inhibition. Being confronted with these contradictions, Cosgrove and Cleland (1983b) mentioned the possible contribution of apoplastic solutes that are in high concentration in the elongating zone to the water relations in the elongating cells of pea stems. Cosgrove (1987) suggested, however, a wellknown mechanism that the motive force for water transport into expanding cells arises from a slight reduction in Pi due to wall yielding. He might have considered that a decrease in Pi enough to sustain the water influx necessary for elongation growth was within a permissible error of the micro pressure probe method (0.3 kPa). Growth enhancement without decreasing PI, however, is possible as illustrated in Fig. 12. If there exists a certain auxin-sensitive mechanism that increases the x-axis intercept of line 11, the new steady-state growth promoted by auxin can be represented by the intersection between lines I and 1 (point a in Fig. 12), because auxin is known to have no effect on membrane hydraulic conductivity ( L J . The auxin-induced increase in the x-axis intercept of line I1 can be caused by an increase in (C' - CX)RT+ Px or the addition of a respiration-dependent nonosmotic term to Eq. (6). In conclusion of sections I11 and IV, auxin controls stem elongation through regulating both the surface proton pump and the xylem proton pump. The surface proton pump participates in the regulation of yielding of the surface wall layers as described by Eq. (3,and the xylem proton pump engages in the active transport of solutes from the xylem fluid into the symplast. An apparent active or nonosmotic transport of water from the xylem fluid, which is induced by auxin, must be intimately linked to the active transport of solutes driven by the xylem proton pump (Fig. 13).

V. Integration of the Activity of the Symplast in Plant Growth

A. Involvement of Respiration-Dependent Water Uptake during Stem Elongation Growth In stems of growing higher plants, water necessary for elongation growth is supplied via the xylem vessels. Water absorbed from the xylem vessels is transported radially toward the epidermis. The radial transport would not be fully apoplastic but symplastic because the apoplastic pathway is interrupted at the starch sheath by a certain barrier (Pearce and Penny, 1986) (Figs. 6, 15). The main barrier to this radial water movement, there-

286

KlYOSHl KATOU AND HISASHI OKAMOTO Aux I n

Synthet i c metabol ism

The xylem H'-punw

The s u r f a c e H+-pump

Secondary a c t i v e s o l u t e transport

Wall

Resp i r a t i on-dependent

Ad j u s t ab Ie y i e Id threshold

osmos I s

(I Act I v e water uptake

extensiblllty

Active s o l u t e uptake

f

/

J.

Yielding of the surface wall

Stem e l o n g a t i o n

FIG. 13 Scheme of the action of auxin in the biophysical regulation of stem elongation growth.

fore, is the boundary membrane between the xylem apoplast and the symplast where the electrogenic xylem proton pump is working very actively (Van Be1 and Reinhold, 1975; Okamoto et al., 1978; Mizuno et al., 1985). Growth promotion of an abraded pea hypocotyl segment whose lateral surface was exposed to air was induced by acid aerosol, but ceased by anoxia even under a sufficient supply of acid-aerosol (Mizuno and Okamoto, 1982). The peripheral wall layer that restricts stem growth (Tanimot0 and Masuda, 1971; Masuda and Yamamoto, 1972; Firn and Digby, 1977; Brummell and Hall, 1980; Kutschera and Briggs, 1987) must be loosened by acid aerosol. The growth retardation under anoxia indicates that respiration-dependent water absorption behaves as a limiting process in the elongation of the segment. The situation is also true as in the case of anoxia-caused inhibition of endogenous growth of hypocotyl segments, because recently it was found that the extensibility of the cell-wall layers remained unchanged or only slightly decreased, even under anoxia (Okamot0 et al., 1989, 1990). These phenomena clearly indicate that elongation

SYMPLAST IN PLANT GROWTH

287

of the aerial organs requires not only loosening of the peripheral wall layers but also respiration-dependent water uptake from the xylem vessels. It is rather likely that the xylem proton pump participates in water uptake from the xylem apoplast. It has already been shown that the xylem pump is stimulated by auxin (De Boer et al., 1985; Ikoma and Okamoto, 1988) and plays a role in solute absorption from the xylem (Van Be1 and Van Erven, 1979; De Boer et al., 1985). When a segment of the hypocotyl of V . unguiculata was subjected to osmotic stress imposed by xylem perfusion, it showed a transient shrinkage followed by adaptive reelongation. Elongation thus resumed under osmotic stress, however, began to immediately shrink upon anoxia. Reaeration caused the segment to elongate again. The change in the volume of the hypocotyl segment was always in proportion to that of the length because only a slight change was observed in the diameter (Okamoto and Katou, 1988). Thus, stress-induced reversion in the elongation of the hypocotyl segment should be caused by the reversion of water transport across the boundary membrane between the symplast and the xylem apoplast. The changes in the activity of the xylem proton pump always preceeded those in the activity of water transport (Okamoto and Katou, 1988). It is unlikely that changes in the hydraulic conductivity or in the reflection coefficient of the cell membrane are responsible for this reversion of water transport, but the driving force for water transport must reverse. In lentil roots, Kuzmanoff and Evans (1981) had reported rapid growth adaptation to osmotic stress similar to that in Vigna hypocotyls. It seems quite likely that respiration-dependent water uptake takes part in the osmotic adaptation of the axial organs. All this suggests that respiration-dependent water uptake efficiently regulates elongation growth of the axial organs in higher plants under various conditions. Active transport of water similar to that of solutes, however, is energetically impossible because of the membrane hydraulic m sec-' Pa-'). conductivity (L,) having a quite large value ( to The process of respiration-dependent water uptake itself should follow the thermodynamic equation [Eq. (6)]. B. Models on Respiration-Dependent Water Transport

Respiration-dependent transport of water in higher plants has been well known as the isoosmotic or nonosmotic exudation from cut stumps of excised roots (Van Overbeek, 1942; Arisz et al., 1951; House and Findlay, 1966; Oertli, 1966; Ginsburg and Ginzburg, 1971). A theoretical basis for the respiration-dependent flow of water was given by Kedem and Katchalsky (1958). The theory reviewed by Dainty (1963) was based on

288

KlYOSHl KATOU AND HISASHI OKAMOTO

the thermodynamic coupling of water and solute fluxes across a nonideal semipermeable membrane (u < 1 ; the KK membrane). According to the theory, water and solute fluxes across the membrane are described by the following well-known simultaneous equations: JT

=

L,(AP

-

uRTAC)

+ (1

V)GF (9) is the water flux across the membrane; q ,the solute flux across JY = wRTAC

-

Here, the membrane; AP is the difference of hydrostatic pressure across the membrane; AC is the difference of solute concentration across the membrane; L, is the hydraulic conductivity of the membrane; u is the reflection coefficient of the membrane to solute; w is the passive mobility of solute in the membrane; c i s the mean concentration of solute in the membrane. These equations suggest that a frictional drag between water and solute molecules takes place during their passage across the KK membrane. It is believed that frictional drag takes place if solute and water molecules simultaneously permeate through water-filled pores penetrating the cell membrane. The shrinking or swelling of a fully turgid plant cell was explained by a water-filled pore model of the cell membrane. Curran (1960) proposed a model of two membranes of different u values in series for isoosmotic fluid transport across the intestinal epithelium of rat. Active solute transport out of the outer compartment across the cell membrane that is thin and semipermeable (i.e., u = 1) increases the concentration of solute in the middle narrow compartment followed by osmotic water influx. This results in an increase in the hydrostatic pressure in the middle compartment. The inner membrane, however, is much thicker with large water-filled pores (i.e., (+ 4 1 ) through which water permeates, dragging solute molecules. Thus, both water and solute lead from the middle compartment into the inner compartment through the inner membrane. Curran and MacIntosh (1962) experimentally confirmed such flow using a model system consisting of two artificial membranes with different u values. Curran’s model greatly influenced studies on the mechanism of respirationdependent water transport across roots (Ginsburg, 1971). In a membrane penetrated by many water-filled pores, osmotic water flow necessarily involves a bulk flow of water through the pores. The osmotic permeability of the membrane for water ( P f ) ,therefore, must always be larger than the diffusional permeability for water (Pd). The reported large values of PfIP, in both biological and artificial phospholipid membranes were in favor of the pore theory. Cass and Finkelstein (1967), however, found that Pd of artifical bilayer membrane made of biological lipids was not so small but nearly the same as P, when the effects of unstirred layer (Dainty, 1963) were carefully eliminated. The actual values

SYMPLAST IN PLANT GROWTH

289

of Pd (or P f ) of artificial lipid bilayer membranes ( - l ~ l O - ~m sec-I) indicate that the permeation of water across most cell membranes takes place through the lipid bilayer. It has been shown that Pd attributable to ion channels that accompany the bulk flow of water (i.e.. Pf/Pd of ion channels > 1) was only 1% of the actual P f value of the cell membrane of animal cells except erythrocyte, having larger P f (-2x m sec-’) (Finkelstein, 1984). In plant cells, determination of the true value of P , is very difficult because the cell wall layer builds a great unstirred layer for water movement just outside the cell membrane. Gutknecht (1967), however, succeeded in estimating the Pd of Valonia cell avoiding the effect of the unstirred layer and found P f / P , = I for the plasmalemma. This strongly suggests that there are no interactions between net flux of water across the plasmalemma and the fluxes of solutes. The main pathway of water transport across the plasmalemma is also the lipid bilayer except the plasmalemma of Characean cells, whose P, is much larger, as in erythrocyte cells. From present knowledge, the values of the reflection coefficient (a)of the plasmalemma for physiological solutes have been determined to be close to 1.0 (Hastings and Gutknecht, 1978; Palta and Stadelmann, 1980; Tyerman and Steudle, 1982; Steudle, 1989). The plasmalemma that behaves as the KK membrane assumed as the second membrane in Curran’s model has not yet been found in higher plants. Therefore, it is quite likely that the movement of water across the plasmalemma is caused not by frictional drag but by osmosis. Respiration-dependent and apparent nonosmotic transport of water is possible by pure osmosis if the structural and/or anatomical characteristics of biological system are taken into consideration. Arisz et al. (1951) proposed a particular part of the vessel element called the “active volume” into which active solute secretion and the following osmotic water flow take place across a root. If not the solute but the water absorbing zone is extended beyond the active volume toward the base of the root, the sap originally absorbed into the active volume is progressively diluted to be isoosmotic with the rooting medium during the upward flow along the vessel elements (Fig. 14). The underlying process of water movement in their model is purely osmotic in nature, but the respiration-dependent and apparent loose coupling between net fluxes of water and solute takes place within a semi-macroscopic system. The model of Arisz er al., (1951) would really be the first clear proposal of the concept of local osmosis and/or standing osmotic gradient that has become well known as the mechanism of secretion of isoosmotic fluid in animal epithelia (Ballantine, 1959, cited in Oschman et al., 1974; Diamond and Bossert, 1967). The standing gradient model supposes a standing gradient of solute concentration within channels of narrow intercellular spaces or folded surfaces. Apparent loose coupling of water and solute movements originates from the transport of

290

KlYOSHl KATOU AND HISASHI OKAMOTO

FIG. 14 The active volume model for root exudation. Om,osmotic value of outer solution; Ob. osmotic value of xylem sap; S, salt secretion; b, bleeding, rate of water absorption; V, active volume of xylem where salt secretion and water transport take place. (Arisz e r a / . , 1951.)

solutes into or out of the channels where volume flow along the channel and solute diffusion occur simultaneously. The channel system as a whole, so to speak, behaves like a coupling membrane with low IT, but no direct coupling between water and solute fluxes occurs anywhere in the system. In Curran’s model, frictional coupling takes place during the movements of water and solute through the second membrane. Governing equations of motion within the channel system are different from those of Curran’s model of double membranes (i.e., div J s = 0 and div v = 0 in Curran’s model but not in the standing gradient model). Therefore, Friedman’s consideration (1986) to regard the standing gradient model as an approximate mapping of Curran’s model of double membranes seems to be inappropriate.

SYMPLAST IN PLANT GROWTH

291

Respiration-dependent water uptake would easily take place in plant stems if transfer cells with well-developed wall ingrowths (Pate and Gunning, 1972) are located around the xylem vessels, because the function of the wall ingrowths could be considered to be the same as that of the infoldings in animal epithelia (Diamond and Bossert, 1967; Oschman et al., 1974). However, no wall ingrowths were found in the cells of elongating Vigna hypocotyl. Therefore, we have reexamined the anatomical characteristics of the stem that may enable respiration-dependent uphill water transport (Katou and Furumoto, 1986a,b). C. The Apoplast Canal Concept for Respiration-Dependent Water Uptake in the Stem

The xylem of the hypocotyl of V . unguiculata is tightly packed with parenchyma cells and walls. The cells of the xylem parenchyma are forming a symplast together with the cells of the cortex. A certain insulating structure at the radial wall of the starch sheath (see Section I1 and Fig. 6) prevents the apoplastic radial transport between the stele and the cortex across the starch sheath. The stele, and therefore the xylem parenchyma, is apoplastically isolated from the cortex and the epidermis. Therefore, the models of radial water transport in which the radial flux of water along a continuous gradient of the water potential in the apoplast is assumed (Cosgrove, 1987; Boyer, 1988) are unlikely. All the water and solutes absorbed in the xylem parenchyma from the xylem apoplast should be transported toward the epidermis via symplastic pathway (Fig. 15). The immediate environment of the cells of the xylem parenchyma, however, is not the vessels but the canals of the thin apoplastic cell wall between the cells. The xylem proton pump generates a proton-motive force between the apoplastic wall and the symplast across the plasmalemma (Mizuno et al., 1985). Active solute uptake takes place across the plasmalemrna of the symplast driven by this proton-motive force (Van Be1 and Reinhold, 1975; Van Be1 and Van Erven, 1979; De Boer et al., 1985). The apoplast canal of the cell wall behaves as an unstirred zone with smaller diffusion coefficients for solutes, where solute movements are not directly disturbed by the flow of the xylem sap through the vessels. Therefore, one may suppose that the active net uptake of solutes by the symplast creates a respiration-dependent depletion of solutes within the narrow canal of the cell wall. This solute depletion provides larger driving force for water uptake than expected from the difference of water potential between the symplast and the xylem vessels. Enough solute depletion in the canal enables water to be transported from the xylem vessels into the symplast even against an overall uphill gradient of the water potential.

292

KlYOSHl MTOU AND HISASHI OKAMOTO

epidermis cortex starch sheath vessel xylem parenchyma

xylem vessel

1 1 ,,

x=O

,,

J

,,

1

,,

1

,,

J xylem

vessel

x= h

FIG. 15 (Top) Sketch of a cross-sectional area of a segment of the hypocotyl of Vigna unguiculata. Plasmodesmata between the cells are not shown in this sketch. (Bottom)Scheme ofthe canal system in the xylem of the hypocotyl of Vigna unguiculara. (Katou and Enomoto, 1991.)

Despite the apparent active transport of water, the movement of water underlying this process is purely passive, i.e., osmotic in nature, which can be described by Eq. (6). Water transport, however, is greatly influenced by the rate of net solute transport. Solute transport is powered by the protonmotive force generated by the xylem proton pump as discussed earlier. Thus, the apoplast canal system as a whole can couple net fluxes of water and solute. The wall ingrowths or invaginations, which have been shown to play a role in fluid secretion in the Limonium salt gland as a coupling space (Hill and Hill, 1973), are not always necessary constituents for the coupling of water and solute fluxes. The simplest physical model of the apoplast canal system for the simultaneous transport of water and solute can be considered in the xylem parenchyma between the two main vessels (Katou and Furumoto, 1986a,b)(Fig. 15). The apoplast canal is a thin film of the cell wall sandwiched between the cells of the xylem parenchyma. The canal ends are both open to a pair of the xylem vessels.

293

SYMPLAST IN PLANT GROWTH

The symbols employed are: Apparent concentration of the solution absorbed through the canal system into the symplast (=J,(t,O)/u(r,O) = J , ( t , h ) / u(t,A))(osmol m-3) Concentration of solute in the symplast (osmol m-3) Concentration of solute in the apoplast canal (osmol mP3) Concentration of solute in the xylem fluid (osmol m-)) The solute diffusion coefficient in the apoplast canal (m2sec- I ) The thickness of the apoplast cell wall (m) The net flux of solute uptake across the plasmalemma into the symplast cells that are in contact with the apoplast canal (osmol rnp2 sec-') Osmotic water flux across the plasmalemma of the symplast that is in contact with the canal (m sec-I) Solute flux along the canal (osmol rn-' sec-I) The hydraulic conductivity of the symplast cell membrane (m sec-l Pa-') The length of the apoplast canal (pm) Intracellular hydrostatic pressure of the symplast with reference to atmospheric pressure (Pa) Hydrostatic pressure of the xylem vessels with reference to atmospheric pressure (Pa) The rate of volume flow along the canal (linear velocity) (m sec-9 t and x are denoted in minutes and micrometers, respectively. Consider the movements of solute and water in the canal system quantitatively. It is convenient that the x axis is designated along the canal, and that one and the other ends of the canal are set as x = 0 and x = A, respectively. Solute flow along the canal is driven by both diffusion and volume flow:

J,

=

-D grad C,+ C,v.

(10)

Water transport from the canal into the symplast cell is driven by osmosis:

I"

=

L,(uRT(C' - C,)

-

(Pi - P X ) } ,

( 1 1)

where the reflection coefficient u = 1 is assumed for most physiological solutes, as discussed earlier. For an infinitesimal segment of canal (Fig. 16), solute flow and water flow should be conserved throughout as div J, = - dC,/dt - 21,/6 and div v = - 2 I J 6 . Therefore the canal equation becomes the following set of nonlinear differential equations (Katou and Furumoto, 1986a,b, 1988; Katou and Enomoto, 1991):

294

KlYOSHl KATOU AND HISASHI OKAMOTO

J,

+ dJ,

v + dv

FIG. 16 Solute flow (J,) and water flow (v) through an infinitesimal segment of apoplast canal. I , and I , are the rate of net solute uptake and that of water uptake across the plasmalemma of the symplast, respectively. S is the thickness of the cell wall adoplast. (Modified from Katou and Furumoto, 1988).

dC,ldt = D div grad C , - div (C,v) - 2416

div v =

-

2I,/6.

(12) (13)

These equations are termed the “canal equations.” For a one-dimensional model canal, these could be solved numerically using a computer (Katou and Furumoto, 1986a,b; Katou and Enomoto, 1991), although a singleended canal extending from a vessel to the barrier of the starch sheath may also be active. The boundary conditions of the canal equations for the one-dimensional model canal are that the water flux along the canal must be null at the midpoint of the canal and that the solute concentration in the xylem vessels is equal to that at the canal end. Thus, u(t, X12) = 0 and C,(t,O) = C,(t,h) = C”(t)if all the solutes are assumed to be absorbable.

D. Auxin-Enhanced Steady-State Water Uptake Calculated by the Apoplast Canal Model

Steady-state water uptake in the absence or presence of auxin was examined applying the apoplast canal model on a model pea hypocotyl segment (Katou and Furumoto, 1986b). In the case of steady-state elongation, it should be dC,ldt = 0 in Eq. (12). The canal equations have been solved numerically for a model pea stem in steady-state elongation by the Runge-Kutta-Verner procedure where Piand Ci were kept constant. Several pairs of standing profiles of the solute concentration (C,) and the water flux along the canal ( u ) have been obtained with varying I,. The profiles of I,-dependent standing-solute

SYMPLAST IN PLANT GROWTH

0

295

100

200

300

Canal length in pm FIG. 17 Standing profiles of solute concentration (C,) and volume flow velocity ( u ) within the apoplast canal of growing model hypocotyl of Vigna unguiculata. Parameters adopted for the canal equations are as follows (Katou and Enomoto, 1991): A = 340 pm, 6 = 2 pm, D = 3 X lo-" mz sec-l, L, = 1.0 x m sec-' Pa-I, C' = 290 mol m-3, P,= 0.62 MPa, P x = 0; because hypocotyl segments were considered, C,(O) and C,(h) was chosen as 55.9 mol m-3 taking free space solutes into consideration (Cosgrove and Cleland, 1983a). I , was vaned from 105 (1) to 285 (7) nosmol m-2 sec-I in 30 nosmol m-? sec-' step. In each condition, a stationary growth was assumed.

depletion within the canal are presented in Fig. 17 (Curve Cs).The distributions of the linear velocity along the canal were also calculated (Curve v in Fig. 17). Net solute uptake was indeed able to generate distinct solute depletion in the canal under each condition imposed. The range of I , values adopted in the calculation seemed to be adequate as shown in a number of reports on the transport activity of plant cell membranes (Raven, 1976).

296

KlYOSHl KATOU AND HISASHI OKAMOTO

Solute depletion increases as the rate of net solute uptake increases. The positive and negative values of u indicate the flow along and against the x axis, respectively. The profiles of u in Fig. 17 indicate that the xylem fluid enters the canal through both open ends, and that the flowing fluid is thoroughly absorbed through the symplast membranes of both lateral sides of the canal until it reaches the midpoint of the canal where the direction of fluid flow reverses. The rate of water absorption by this canal from the xylem vessels is equal to u(0) - u(h). Under the conditions in Fig. 17, net water uptake always occurs though the water potential of the symplast is higher than that of the vessels. It increases as net solute uptake increases. The solute concentration of the fluid absorbed by the canal system (Cap,) depends on I,. Calculated Cap,that is equal to J,(O)lu(O) or J,(h)lu(h)was found to decrease with increase in u(0) or I , (Katou and Furumoto 1986b). Calculated results indicate that net solute uptake from the apoplast canal generates solute depletion within the canal which increases the water potential of the canal and enables symplast cells to absorb water from the vessels against gradient of water potential. Thus, no changes in Pi,C',and L, are necessary for the parenchyma symplast to absorb water from the xylem fluid. Water transport through the apoplast canal system is powered by net transport of solutes that is known to be driven by proton-motive force across the plasmalemma (Van Be1 and Reinhold, 1975; De Boer et al., 1985). Therefore, water transport through the apoplast canal system depends on respiration. It seems to be Loosely coupled with the transport of solutes, but this coupling is an apparent one. Auxin is known to rapidly hyperpolarize the cell membrane (Etherton, 1970; Marre et al., 1974; Cleland et al., 1977; Mizuno et al., 1980; Bates and Goldsmith, 1983; Ikoma and Okamoto, 1988) followed by activation of solute uptake from the xylem vessels (De Boer et al., 1983, whereas C', Pi,and L, are hardly affected by auxin as previously discussed (see Section IV, C and D). The parameter C"in Eq. (6)should not be considered as the solute concentration in the vessels but the average solute concentration in the apoplast canal. Auxin-induced increase in the x-axis intercept of Eq. (6)(Fig. 12) takes place without adding a new term of nonosmotic water movement to the right-hand side of Eq. (6) but with decreasing C" that is caused by solute depletion within the canal (line I1 shifts to line I in Fig. 12). By this shift, together with the changes in the mechanical properties, the rate of elongation increases as represented by point a in Fig. 12 (i.e., b +. a). Respiratory inhibition causes the x-axis intercept to decrease because collapse in the standing-solute depletion in the canal inevitably increases Cx (line I1 shifts to line I11 in Fig. 12), which results in decrease in P'. Thus, the apoplast canal model could give a comprehensive explanation for most contradictions reported on the mechanism of auxin-enhanced water uptake in the elongating cells of higher plants in situ.

SYMPLAST IN PLANT GROWTH

297

Average increase in solute depletion induced by auxin, which enhances water uptake, was calculated to be 0.03 MPa (in terms of osmotic pressure). The hydraulic conductivity of the cell membrane is indeed high enough that the regulation of water transport requires only a small change in solute concentration in the apoplast canal. The fact that the volume of the canal is quite small compared with that of the symplast indicates that the apoplast canal system as the mechanism of respiration-dependent water transport is very efficient and economical. The apparent free space situated outside the xylem vessels observed by the method of xylem perfusion through the tomato stem (Van Bel, 1978) favors the apoplast canal model. The apoplast canal model itself might be regarded as an approximate mapping of the standing-osmotic gradient model for animal epithelia (Diamond and Bossert, 1967). The canal equations [Eqs. (12) and (13)] are mathematically equivalent to the equations for the standing gradient model when d CJdt = 0. However, it would rather be appropriate to say that the apoplast canal model is a reevaluation of the idea of the active volume model for isoosmotic exudation in roots proposed by Arisz et al. (1951) (Fig. 14), taking the fine anatomical and electrophysiological knowledge of the stem into consideration. In addition, the great contribution of hydrostatic pressure to the process of water transport is one of the specific problems of plant physiology. We could not draw any advances in the study of water transport in higher plants from the apoplast canal idea if a pressure probe for directly measuring Pi had not been invented by German scientists (Steudle and Zimmerman, 1971; Husken et al., 1978).

E. Numerical Calculation for the Dynamic Regulation of Plant Elongation Growth The elongation growth of axial organs rapidly recovers just after exposure to osmotic stress. The Pi decreases exponentially upon osmotic stress and remains at a reduced and stable level from then on (Itoh et al., 1987; Nakahori et al., 1990). The response seems to be pure osmosis of passive nature, but the reduced Pi is not fully passive, still partially maintained by respiration (Nakahori et al., 1990) (Fig. 18). Rapid adaptive increase in C' does not take place except the initial rapid increase in sap concentration due to shrinkage of the symplasts (Kuzmanoff and Evans, 1981). The recovery of elongation growth takes place even though Pi and C' do not change adaptively. Thus, the water potential of the symplast did not change at the time of growth recovery. Some experiments suggest that aerobic metabolism and the active uptake of solute may be involved in this rapid recovery of elongation growth (Kuzmanoff and Evans, 1981; Nakahori et al., 1987; Liu et al., 1989). The apoplast canal model must

298

KlYOSHl KATOU AND HISASHI OKAMOTO al

MPa

c

5 0.6 2n

Mannitol

v)

-L

m

0.4

3

?

E

s"

0.2

.

C #-I

0

120 Time i n min

60

180

FIG. 18 Responses of the intracellular pressure (Pi)and elongation growth (G) to osmotic stress. The osmotic concentration of the perfusion solution was increased at the first arrow (Mannitol) from 150 to 200 osmol m-3 with mannitol. At the second arrow (dw), the perfusion solution was replaced by distilled water. (Nakahori et a l . , 1990.)

adequately explain this contradictory situation similar to that observed in auxin-enhanced elongation. The original equations for the standing gradient model (Diamond and Bossert, 1967) and for the earlier canal model (Katou and Furumoto, 1986a,b;Taura et al., 1988) are only applicable to a steady-state transport of water and solutes, i.e., a C,lat = 0. These are ordinary differential simultaneous equations. For time-dependent plant responses, therefore, we adopted the apoplast canal model assuming a series of quasi-steady states. In the adaptive growth recovery against osmotic stress, the assumption of a C,ldt = 0 is no longer valid because some parameters of the canal equations change dramatically. Equations (12) and (13), the simultaneous nonlinear partial differential equations, are the canal equations extended to describe transient or time-dependent rapid movements of solutes and water (Katou and Furumoto, 1988; Katou and Enomoto, 1991). For a one-dimensional model canal, we have successfully numerically solved the extended canal equations applying the finite difference method of time-forward and centered space using a computer (Katou and Enomoto, 1991). Figure 19 shows calculated growth patterns of the segments of cowpea hypocotyl upon additional osmotic stress applied by xylem perfusion. Curves 1 , 2, 3, and 4 represent the patterns of growth with increments in 0,43,67, and 91% of initial I , after the stress, respectively. Osmotic stress stimulates the electrogenic xylem pump that participates in solute uptake from the vessels (Okamoto et af., 1984; Nakahori et al., 1987; Liu et af., 1989). The results clearly show that activated solute uptake plays an indispensable role in the adaptive recovery of growth against osmotic stress. Of course, such activation is a result of the activa-

299

SYMPLAST IN PLANT GROWTH

-

40-

-

0

15

30

45

60

Time (rnin)

FIG. 19 Calculated patterns of growth upon osmotic stress for different degree of promotion of the net uptake of solute. The length represented on the ordinate is given in meters because it is calculated by the integration of Iu(r.0) - u ( t , A)} from t = 0 to t min. Therefore, this change is only proportional to the real change in the length of a hypocotyl segment during elongation. The solute concentration of the xylem fluid (Cx)was increased from 55.9 to 135.9 osmol m-3 at t = 0 min. Curves, I , 2, 3, and 4 represent the patterns of growth with increments in I, of 0, 45, 70, and 95 nosmol m-2 sec-I, respectively, where the steady-state value of I, prior to the stress of 105 nosmol m-’ sec-I. (Katou and Enomoto, 1991.)

tion of the xylem proton pump at the boundary membrane between the symplast and the xylem apoplast (Liu et al., 1989). In addition, the apoplast canal model has been shown to explain well a pure passive response of stem elongation to osmotic stress under respiratory inhibition (Katou and Enomoto, 1991).

VI. Conclusion The elongation growth of the stem of higher plants is undoubtedly supported by cellular activities of the constituent cells. The stem, however, is characteristically organized by constituent cells, which can be realized as an anatomical structure. Most cells in any given cross section of the stem form a disk-like symplast. The stem may be regarded as multiple layers of disk-like symplasts penetrated by vascular bundles (the disk symplast model). There are spatially separated electrogenic proton pumps at the plasmalemma of the symplast: the surface pump and the xylem pump (the two-pump model). The symplasts are arranged chronologically along the germ axis; i.e., the age gradient of the symplast appears along

300

KlYOSHl KATOU AND HISASHI OKAMOTO

the stem axis. This structural organization would be the specific characteristic of plant axial organs. Simultaneous involvement of two physically different processes could be stressed as a typical characteristic of plant elongation growth. One is a rheological process of yielding of epidermal wall layer. Another is a thermodynamic process of membrane transport in the xylem. These processes are joined up by intracellular hydrostatic pressure. It is of great interest that each electrogenic proton pump at the plasmalemma simultaneously participates in the regulation of each elemental process for plant elongation growth. The surface proton pump would modify the parameters of wall yielding by affecting the properties of cell-wall macromolecules just outside the surface membrane of the symplast, the molecular mechanisms of which are expected to be investigated. The xylem proton pump provides driving forces for secondary active transport of nutrient solutes by symplast cells in the xylem parenchyma. Simultaneously, it can provide a driving force for net water transport when organized as a constituent of the apoplast canal system, the mechanism of which has been shown by computer simulation of a semi-macroscopic biophysical model (i.e., the apoplast canal model). Thus, solute and water uptake in the xylem are under full regulation by the xylem proton pump. Auxin enhances both processes via simultaneous activation of the surface and the xylem proton pumps. These suggest that a symplast in a given cross-sectional area, rather than a constituent cell, should be regarded as the functional unit of stem elongation growth. The situation in coleoptiles andlor roots would be the same as that in stems. The elongation growth of the stem can be regarded as one of the highly integrated activities of higher plants. Thus, it could be said that the symplast situated in a semi-macroscopic stratum plays an indispensable role in the integration of molecular, subcellular, and cellular activities into stem elongation.

Acknowledgment The authors thank Mr. T . Taura for assistance in drawing the figures.

References Anderson, W. P. (1975). In “Ion Transport in Plant Cells and Tissues” (D. A. Baker and J. L. Hall, eds.), pp. 231-266. North-Holland, Amsterdam. Arisz, W. H . , Helder, R . I., and Van Nie, R. (1951). J . Exp. Bor. 2, 257-297. Bates, G . W., and Goldsmith, M. H. (1983). Planra 159, 231-237.

SYMPLAST IN PLANT GROWTH

301

Bonner, J. (1934). Protoplasma 21, 406-423. Boron, W. F., and De Weer, P. (1976). J . Gen. Physiol. 67 91-112. Bowling, D. J. F. (1976). “Uptake of Ions by Plant Roots.” Chapman and Hall, London. Boyer, J. S. (1985). Annu. Rev. Plant Physiol. 36, 473-516. Boyer, J. S. (1988). Physiol. Plant. 73, 311-316. Boyer. J. S .. Cavalieri, A. J., and Schulze, E.-D. (1985). Planta 163, 527-543. Boyer, J. S ., and Wu, G. (1978). Planta 139,227-237. Brummell, D. A., and Hall, J. L. (1980). Planta 150, 371-379. Brummer, B., and Parish, R. W. (1983). FEBS Lett. 161, 9-13. Brummer, B., Felle, H., and Parish, R. W. (1984). FEBS Lett. 174, 223-227. Biinning, E. (1953). “Entwicklungs- u. Bewegungsphysiologie der Pflanze.” SpringerVerlag, Berlin. Burstrom, H. G. (1957). Symp. Soc. Exp. Biol. 11 44-62. Burstrom, H. G. (1971). Nature (London) 234, 488. Cass, A.,and Finkelstein, A. (1967). J. Gen Physiol. 50, 1765-1784. Clarkson, D. T. (1988). In “Solute Transport in Plant Cells and Tissues” (D. A. Baker, and J. L. Hall, eds.), pp. 251-304. Longman Scientific & Technical, Essex. Clarkson, D. T., Williams, L., and Hanson, J. B. (1984). Planta 162, 361-369. Cleland, R. (1971). Annu. Rev. Plant Physiol. 22, 197-222. Cleland, R. E. (1977). Symp. Soc. Exp. Biol. 31, 101-1 15. Cleland, R. E., Prins, H. B. A , , Harper, J. R., and Higinbotham, N. (1977). Plant Physiol. 59, 395-397. Cosgrove, D. J. (1981). Plant Physiol. 68, 1439-1446. Cosgrove, D. J. (1985). Plant Physiol. 78, 347-356. Cosgrove, D. J. (1986). Annu. Rev. Plant Physiol. 37, 377-405. Cosgrove, D. J. (1987). Plant Physiol. 84, 561-564. Cosgrove, D. J . , and Cleland, R. E. (1983a). Plant Physiol. 72, 326-331. Cosgrove, D. J . , and Cleland, R. E. (1983b). Plant Physiol. 72, 332-338. Cosgrove, D. J . , and Sovonick-Dunford, S. A. (1989). Plant Physiol. 89, 184-191. Cosgrove, D. J., Van Volkenburgh, E., and Cleland, R. E. (1984). Planta 162, 46-54. Crafts, A. S ., and Broyer, T. C. (1938). A m . J . Bot. 25, 529-535. Curran, P. F. (1960). J. Gen. Physiol. 43, 1137-1148. Curran, P. F., and MacIntosh, J. R. (1962). Nature (London) 193, 347-348. Dainty, J. (1963). Adv. Bot. Res. 1, 279-326. De Boer, A. H., Katou, K., Mizuno, A., Kojimd, H., and Okamoto, H. (1985). Plant Cell Environ. 8, 579-586. De Boer, A. H., and Prins, H. B. A. (1985). Plant Cell Environ. 8, 587-594. De Boer, A. H., Prins, H. B. A., and Zanstra, P. E. (1983). Planta 157, 259-266. Diamond, J. M., and Bossert, W. H. (1967). J. Gen. Physiol. 50, 2061-2083. Dunlop, J. (1974). J. Exp. Bot. 25, 1-10. Dunlop, J., and Bowling, D. J. F. (1971a). J. Exp. Bot. 22, 434-444. Dunlop, J., and Bowling, D. J. F. (1971b). J. Exp. Bot. 22, 453-464. Etherton, B. (1970). Plant Physiol. 45, 527-528. Felle, H. (1987). J . Exp. Bot. 38, 340-354. Felle, H., and Bertl, A. (1986). J. Exp. Bot. 37, 1416-1428. Felle, H., Brummer, B., Bertl, A., and Parish, R. W. (1986). Proc. Natl. Acad. Sci. U.S.A. 83, 8992-8995. Finkelstein, A. (1984). Curr. Top. Membr. Trans. 21, 295-308. Firn, R., and Digby, J., (1977). Aust. J . Plant Physiol. 4, 337-347. Friedman, M. H. (1986). “Principles and Models of Biological Transport.” Springer-Verlag, Berlin.

302

KlYOSHl KATOU AND HISASHI OKAMOTO

Galston, A. W., and Purves, W. K. (1960). Annu. Rev. Plant Physiol. 11, 239-276. Ginsburg, H. (1971). J. Theor. B i d . 32, 147-158. Ginsburg, H., and Ginzburg, B. Z. (1971). J . Membr. B i d . 4, 29-41. Green, P. B., and Cummins, W. R. (1974). Plant Physiol. 54, 863-869. Green, P. B., Erickson, R. 0.. and Buggy, J. (1971). Plant Physiol. 47, 423-430. Gutknecht, J. (1967). Science 158, 787-788. Hackett, D.P., and Thimann, K. V. (1952). Plant Physiol. 25, 648-652. Hager, A. (1980). Z. Naturforsch. C 35, 794-804. Hager, A., Debus, G., Edel, H . G . , Stransky, H.. and Serrano, R. (1991). Planta 185, 527-537.

Hager, A., and Moser, I. (1985). Planta 163, 391-400. Hager, A., Menzel, H., and Krauss, A. (1971). Plantu 100, 47-75. Hanson, J. B. (1978). Plant Physiol. 62, 402-405. Hara, N. (1972). “Morphology of Plants.” Shokabo, Tokyo. [in Japanese] Hastings, D. F., and Gutknecht, J. (1978). J . Theor. Biol. 73, 363-366. HerEk, F. (1925). Pub. Far. Sci. Univ.Masaryk (Brno) 49, 3-21. Heyn, A. N. J. (1940). Bot. Rev. 6, 515-574. Higinbotham, N. (1973). Annu. Rev. Plant Physiol. 24, 25-46. Hill, A. E., and Hill, B. S. (1973). J . Membr. Biol. 12, 129-144. House, C.R., and Findlay, N. (1966). Nutitre (London) 211, 649-650. Husken, D.,Steudle, E., and Zimmermann. U. (1978). Plant Physiol. 61, 158-163. Ichimura, K., and Okamoto, H. (1958). Bot. Mag. Tokyo 71, 201-207. Ichino, K . (1978). Plant Cell Physiol. 19, 253-262. Ichino, K., Katou, K., and Okamoto, H. (1973). Plant Cell Physiol. 14, 127-137. Ikoma, S.,and Okamoto, H. (1988). Plant Cell Physiol. 29, 261-267. Ishikawa, H., Yamamura, K., Furukoshi, M., Ohta, M., and Sakata, M. (1984). Plant Cell Physiol. 25, 1045-1051. Itoh, K., Nakamura, Y., Kawata, H., Yamada, T., Ohta. E., and Sakata, M. (1987). Plant Cell Physiol. 28, 987-994. Katou, K. (1978). Plant Cell Physiol. 19, 523-535. Katou, K., and Enomoto, M. (1991). Plant Cell Physiol. 32, 343-351. Katou, K., and Furumoto, M. (1986a). Protoplasma 130, 80-82. Katou, K., and Furumoto, M. (1986b). Protoplasma 133, 174-185. Katou, K., and Furumoto, M. (1988). Protoplasma 144, 62-63. Katou, K., and Ichino, K. (1982). Planta 155, 486-492. Katou, K., and Okamoto, H. (1970). Plant Cell Physiol. 11, 385-402. Kedem, O., and Katchalsky, A. (1958). Biochim. Biophys. Acta 27, 229-246. Kholdebarin, B.,and Oertli, J. J. (1977). Z. Pjanzenphysiol. 83, 393-402. Krauss, A., and Hager, A. (1976). Z. Naturforsch. C 31, 312-316. Kurkdjian, A., and Guern, J. (1989). Annu. Rev. Plant Physiol. Plant Mol. Biol. 40,271-303. Kurkdjian, A., Leguay, J.-J., and Guern, J. (1978). Resp. Physiol. 33, 75-89. Kutschera, U.,and Briggs, W. R. (1987). Plant Physiol. 84, 1361-1366. Kutschera, U.,and Schopfer, P. (1986). Planta 167, 527-535. Kuzmanoff, K. M., and Evans, M. L. (1981). Plant Physiol. 68, 244-247. Liu, Q.. Katou, K., and Okamoto, H. (1989). Plant Cell Physiol. 30, 1039-1046. Liu, Q.,Katou, K., and Okamoto, H. (1991). Plant Cell Physiol. 32, 1021-1029. Lockhart, J. A. (1965). J. Theor. B i d . 8, 264-275. Lund, E. J., et al. (1947). “Bioelectric Fields and Growth.” Univ. of Texas Press, Austin. Luttge, U. (1971). Annu. Rev. Plant Physiol. 22, 23-44. Liittge, U.,and Higinbotham, N. (1979). “Transport in Plants.” Springer-Verlag. New York. Marre, E., Lado, P., Ferroni, A., and Ballarin Denti, A. (1974). Plant Sci. Lett. 2, 257-265.

SYMPLAST IN PLANT GROWTH

303

Masuda, Y. (1990). Bot. M a g . Tokyo 103, 345-370. Masuda, Y . , and Yamamoto, R. (1972). Physiol. Plant. 27, 109-115. Meyer, R. F . , and Boyer, J. S. (1972). Planta 108, 77-87. Mizuno, A. (1978). Plant Cell Physiol. 19, 1315-1326. Mizuno, A., and Katou, K. (1991). Plant Cell Physiol. 32, 403-408. Mizuno, A., Katou, K.. and Okamoto, H. (1980). Plant Cell Physiol. 21, 395-403. Mizuno, A., Kojima, H., Katou, K., and Okamoto, H. (1985).Plant Cell Enuiron. 8,525-529. Mizuno, A., and Okamoto, H. (1982). Plant Cell Enuiron. 5, 131-135. Mizuno, A., and Okamoto, H. (1983). Plant Cell Enuiron. 6, 243-246. Mothes, K. (1960). Naturwissenshaften 47, 337-351. Nakahori, K., Katou, K., and Okamoto, H. (1987). Plant Cell Physiol. 28, 901-910. Nakahori, K.. Katou, K., and Okamoto, H. (1991). Plant Cell Physiol. 32, 121-129. Nakahori, K., Koizumi, K., Muramatsu, S., Ohtani, H., Masuko, M., Nakase, S., Katou, K.. and Okamoto, H. (1990). Plant Cell Physiol. 31, 859-864. Nelles, A. (1977). Planta 137, 293-298. Oda, K. (1955). Sci. R e p . Tohoku Uniu. 4th Ser. (Biol.) 21, 109-121. Oertli. J . J. (1966). Physiol. Plant. 19, 809-817. Ohkawa, T., Kohler, K., and Bentrup, F.-W. (1981). Planta 151, 88-94. Okamoto, H. (1955). Bot. M a g . Tokyo 68, 1-13. Okamoto, H. (1976). Plant Cell Physiol. 17, 1273-1280. Okamoto, H . (1991). “Unknown Vital Activity of Plants.” Otsuki-Shoten, Tokyo. [in Japanese] Okamoto. H., Ichino, K., and Katou, K. (1978). Plant Cell Enuiron. 1, 279-284. Okamoto, H., and Katou, K. (1988). Plant Cell Physiol. 29, 509-515. Okamoto, H., Katou, K., and Ichino, K. (1979). Plant Cell Physiol. 20, 103-114. Okamoto, H., Liu, Q.. Nakahori, K., and Katou, K. (1989). Plant Cell Physiol. 30,979-985. Okamoto, H., Miwa, C., Masuda. T., Nakahori, K., and Katou, K. (1990).Plant Cell Physiol. 31,783-788. Okamoto, H., Mizuno, A., Katou, K., Ono, Y.,Matsumura, Y.. and Kojima, H. (1984). Plant Cell Enuiron. 7, 139-147. Oota, Y. (1964). Kagaku (Science) 34, 196-199. [in Japanese] Ordin, L., Applewhite, T. H., and Bonner, J. (1956). Plant Physiol. 31, 44-53. Ortega, J. K. E. (1985). Plant Physiol. 79, 318-320. Ortega, J. K. E., Zehr, E. G., and Keanini, R. G. (1989). Biophys. J . 56, 465-475. Oschman, J . L., Wall, B. J., and Gupta, B. L. (1974). Symp. SOC.Exp. Biol. 28, 305-350. Palta, J . P., and Stadelmann, E. J. (1980). Physiol. Plant. 50, 83-90. Parrish, D. J., and Davies, P. J. (1977). Plant Physiol. 59, 745-749. Pate, J . S . , and Gunning, B. E. S. (1972). Annu. Reu. Plant Physiol. 23, 173-196. Pearce, D., and Penny, D. (1986). Physiol. Plant. 67, 61-66. Penny, P., and Penny, D. (1978). I n “Phytohormones and Related Compounds-A Comprehensive Treatise” (D. S. Letham, P. B. Goodwin. and T. J. V. Higgins, eds.), Vol. 11, pp. 537-597. ElseviedNorth-Holland Biochemical Press, Amsterdam. Penny, P., Penny, D., and Marshall, D., and Heyes, J. K. (1972). J . Exp. Bot. 23, 23-36. Pitman, M. G. (1977). Annu. Reu. Plant Physiol. 28, 71-88. Pritchard, J . , Tomos, A. D., and Wyn Jones, R. G. (1987). J. Exp. Bot. 38, 948-959. Raven, J. A. (1976). I n “Encyclopedia of Plant Physiology,” New Series, Vol. 2, “Transport in plant 11, Part A, Cells” (U. Luttge. M. G. Pitman, eds.), pp. 129-188. Springer-Verlag, Berlin. Ray, P. M., Green, P. B., and Cleland, R. (1972). Nature (London) 239, 163-164. Rayle, D. L., and Cleland, R. (1970). Plant Physiol. 46, 250-253. Rayle, D. L., and Cleland, R. (1972). Planta 104, 283-296.

304

KlYOSHl KATOU AND HISASHI OKAMOTO

Rayle, D. L., and Cleland, R. (1977). Curr. Top. Dev. Biol. 11, 187-214. Reinders, D. (1938). Proc. Kon. N e d . Akad. v. Wetensch., Amsterdam 41, 820-831. Rich, T. C. G., and Tomos, A. D. (1988). J. Exp. Bot. 39, 291-299. Robards, A. W., and Clarkson, D. T. (1976). In “Intracellular Communication in Plants: Studies on Plasmodesmata” (B. E. S . Gunning and A. W. Robards, eds.), pp. 181-201. Springer-Verlag, Berlin. Spanswick, R. (1972). Planta 102, 215-227. Steudle, E. (1989). I n “Plant Membrane Transport: The Current Position” (J. Dainty, M. I. de Michelis, E. Marrk, and F. Rasi-Caldogno, eds.), pp. 551-554. Elsevier Science Publishers B. V. (Biomedical Division), Amsterdam. Steudle, E., and Zimmermann, U. (1971). 2. Narurforsch. 26, 1302-1311. Stevenson, T. T., and Cleland, R. E. (1981). Plant Physiol. 67, 749-753. Tanimoto, E., and Masuda, Y. (1971). Plant Cell Physiol. 12, 663-673. Taura, T., Iwaikawa, Y., Furumoto, M., and Katou, K. (1988). Profoplasma 144, 170-179. Thomas, R. C. (1976). J. Physiol. 255, 715-735. Tomos, A.D.(1985). In “Control of Leaf Growth” ( N . R. Baker, W. J., Davis, and C.K. Ong, eds.), pp. 1-33. Cambridge Univ. Press., Cambridge. Tyerman, S. D., and Steudle, E. (1982). Aust. J. Plant Physiol. 9, 461-479. Van Bel, A. J. E. (1978). J. Exp. Bor. 29, 295-303. Van Bel, A. J. E., and Reinhold, L. (1975). 2. Pjanzenphysiol. 76, 224-228. Van Bel, A. J. E., and Van Erven, A. J . (1979). Plant Sci. Lett. 15, 285-291. Van Overbeek, J . (1942). Am. J. Bot. 29, 677-683. Van Volkenburgh, E., and Cleland, R. E . (1986). Planra 167, 37-43. Yoda, S., Ashida, J. (1961). Nature (London) 192, 577-578.

lntracellular Ca2+ Messenger System in Plants Shoshi Muto Institute of Applied Microbiology, University of Tokyo, Tokyo, Japan

1. Introduction Plants sense continually fluctuating environmental conditions and transduce these changes into physiological responses. However, the mechanism of signal transduction in plants is little known. Nevertheless, the pathway of signal transduction in plant cells may be outlined following the scheme established in animals (Fig. 1). Environmental signals, e.g., light, temperature, atmospheric gases, nutrients, are perceived as primary signals by membrane-localized receptors. These are transduced to intracellular messengers (second messengers) by transducers neighboring the receptor. The second messengers modify proteins and enzymes and bring about changes in biochemical reactions. These changes subsequently lead to physiological responses. In long-term responses, protein synthesis uia gene activation is included, but short-term responses are carried out without protein synthesis. In the late 1960s, cAMP was established as a second messenger of intracellular signaling in animal cells. Since then, many attempts have been made to prove that cAMP works as a second messenger in plant cells as well. However, no convincing evidence has been obtained. Plant calmodulin was discovered in the late 1970s (Muto and Miyachi, 1977; Anderson and Cormier, 1978) when the concept of Ca2+ as the second messenger in animal cells was being accepted. This discovery triggered the study of a role for Ca2+other than as a macronutrient in plant growth and development. Previous works have shown that the change in cytoplasmic free Ca2+ concentration modifies many physiological processes in plants (Hepler and Wayne, 1985; Kauss, 1987; Poovaiah and Reddy, 1987). Various components and machineries of the intracellular Ca2+messenger system have also been found and, currently, the role of

Internarional Review of Cytology, V d . 142

305

Copyright 0 1992 by Academic Press, Inc. All rights of reproduction in any form reserved.

306

SHOSHI MUTO ENVIRONMENTALSIGNALS -RECEPTORS Phytochrome E l i c i t o r binding p r o t e i n s

1

TRANSDUCERS

t

PROTEIN

1

MOD I F ICATION Phosphorylation/ Dephosphorylation

PHYS’I OLOG ICAL RESPONSES

FIG. 1 Signal transduction in plant cells.

Ca2+as the second messenger in plant cells is being established. There is increasing evidence to this concept. The aim of this chapter is to critically review the present status of knowledge on Ca2+ signaling in plant cells and to point out the unique features of plants in signal transduction.

II. Receptors

The initial step of signal transduction is perception or recognition of the signal by a specific receptor on the membrane. Auxin-binding proteins have been purified and sequenced as putative auxin receptors (Napier and

INTRACELLULAR Ca2+ MESSENGER SYSTEM

307

Venis, 1991). However, there has been no evidence supporting involvement of Ca2+signaling after binding auxins to the binding proteins. Induced phytoalexin synthesis is one of the most extensively studied of the defense responses of plants against pathogens (Dixon and Lamb, 1990). Elicitors derived from pathogens and plant cell wall induce the same response in plants (Scheel and Parker, 1990; Ryan and Farmer, 1991). The first step of elicitor-induced phytoalexin synthesis is most likely some type of recognition of the elicitor at the plant cell surface. Elicitation of isoflavanoid phytoalexin synthesis in soybean by Phytophthora megasperma f. sp. glycinea has been well studied. The elicitors of this fungus have been identified as 1,3-1,6-P-glucans (Ayers et al., 1976a,b). In soybean, the membranes contain high affinity binding sites for 1,3-1,6-P-glucan elicitors (Schmidt and Ebel, 1987; Cosio et al., 1988). The binding of glucans to membranes was saturable, reversible, and associated with the plasma membrane. Binding constants (K,) were determined to be 10-30 nM using '2SI-labeled-tyramineglucan conjugates. The binding activity was solubilized from soybean microsomal membranes (Cosio et al., 1990). Among P-glucan fragments having various degree of polymerization, the branched 1,3-1,6-hepta- P-glucoside had the highest affinity to the binding site (Kd = 0.75-3 nM) (Cheong and Hahn, 1991; Cosio et al., 1991), and has been found to provide a minimum structural requirements for elicitor activity (Sharp et al., 1984a,b). The binding site was assumed to be a protein or glycoprotein, as it was inactivated by proteolysis and by heat treatment (Cheong and Hahn, 1991). Conrath et al. (1991) reported that a rapid and transient Ca2+uptake, K+ release, and external alkalinization of suspension cultured parsley cells was induced by an elicitor preparation from P . megasperma f. sp. glycinea cell wall. This suggests that the perception of elicitor by its receptor couples to intracellular Ca2+ signaling. In wheat, it was reported that a 67-kDa glycoprotein from germ tube walls of Puccinia graminisi f. sp. tritici elicits the cellular hypersensitive lignification response (Kogel et al., 1988). Specific binding sites for this elicitor were detected in highly purified plasma membrane vesicles of wheat leaf cells by immunoblot analysis (Kogel et al., 1991).Furthermore, analyses showed that the binding sites with molecular masses of 30 and 33 kDa have a K , (app) of 2 p M . In cultured soybean cells, phytoalexin glyceollin production is induced by chitosan, fragments of deacetylated chitin. This induction accompanies Ca2+-mediatedstimulation of callose (1,3-linked glucan) synthesis in the cell wall (Kauss, 1987). Although the receptor for chitosan is not known, this finding suggests the involvement of Ca2+ in internalization of the phytoalexin elicitor signal. Receptors of phytotoxins have been identified. One of these is for fusi-

308

SHOSHI MUTO

coccin, which stimulates the plasma membrane H+-ATPase in higher plants. The first step in the mode of action of fusicoccin is its recognition of binding sites with high affinity and specificity. The fusicoccin-binding proteins were solubilized from maize tissues and functionally reconstituted into liposomes together with H+-ATPase (Aducci et al., 1988). The fusicoccin-binding proteins from several plants have high affinity to fusicoccin with K , ranging from 1.7 to 5.2 nM. They are composed of two polypeptides of 29-3 1 kDa and 30-33 kDa, which are located in the plasma membrane (Feyerabend and Weiler, 1989; Meyer and Weiler, 1989; De Boer et al., 1989; Oecking and Weiler, 1991). Thuleau et al., (1988) reported that zinniol, a toxin produced by the Alternaria group, binds to carrot protoplast and isolated membranes in a saturable and reversible manner. Binding of zinniol to the receptor stimulated Ca2+entry to protoplasts and partially reversed the inhibitory effect of calcium channel blockers. Phytochrome is the extensively studied photoreceptor of many divergences of red-light-triggered physiological and developmental events in plants. Chloroplast rotation in Mougeotia (Hepler and Wayne, 1985), regulation of rotational streaming of the cytoplasm in Vallisneria gigantea (Takagi and Nagai, 1985, 1988; Takagi et al., 1990), and spore germination in Onoclea (Hepler and Wayne, 1985) are some of the physiological and developmental processes in plants and in which Ca2+ participation has been well characterized. Red light perception of phytochrome often modulates intracellular CaZt level and thus initiates the Ca2+signaling pathway. Red-light-induced Ca2+flux is considered to be caused by activation of Ca2+channel or inhibition of Ca2+pump in the plasma membrane or in the membranes of intracellular Ca2+storage organelles. However, the process by which phytochrome modulates Ca2+transport remains unclear. A recent report on Auena sariua suggests involvement of G protein in phytochrome action (Romero et al., 1991; see Section 111).

111. G proteins

GTP-binding proteins (G proteins) are central to the transduction of many receptor signals in animal cells (Birnbaumer et al., 1990). They are a highly conserved family of heterotrimeric membrane-associated proteins composed of a,p, and y subunits (G,, G,, and G,). The G protein cycles between inactive GDP-bound and active GTP-bound forms. The G,, which is unique for each G protein, binds GDP or GTP. Upon binding agonists, receptors catalyze the exchange of GDP for GTP binding to G,. GTPbound G, dissociates from the G, G, complex and GTP-bound G, regulates

INTRACELLULAR Ca2' MESSENGER SYSTEM

309

appropriate effector enzymes such as adenylate cyclase or the cyclic GMP phosphodiesterase. GTP bound to G, is hydrolyzed to GDP by GTPase activity of G , and the G, returns to the GDP bound form. GDP-bound G, recombines with the G, G, complex. On the other hand, the G, G, complex of G proteins is required for efficient receptor-catalyzed guanine nucleotide exchange on G,. It also functions as an attenuator of G, activation of effector enzymes. Early works focusing on the function of G proteins have been reported. Dillenschneider et al. (1986) showed that GTP stimulated the release of inositol phosphates from membranes of suspension cultured Acer pseudoplatanus cells. Recently, studies on phytochrome-stimulated GTP binding to protein have been reported (Romero et al., 1991). The binding of GTP to proteins in extract of Auena seedlings is stimulated by red light but this stimulation was negated by far red light. Warpeha et al. (1991) reported that blue light activated G protein in the plasma membranes of etiolated pea seedlings. The GTPase activity was induced by low fluence of blue light administered to plasma membrane-enriched fraction. In this study, it was observed that anti-G,, (transducin a subunit) antisera immunoreacted to a 40-kDa peptide that was ADP-ribosylated by cholera and pertussis toxins. The ability of the 40-kDa peptide to serve as substrate for the toxinmediated ribosylation was mediated by blue-light irradication. It was also observed that the 40-kDa peptide bound a nonhydrolyzable photoaffinitylabeling analong of GTP only after blue-light irradiation. So far, a heterotrimeric G protein has not been isolated or purified in plants. However, the occurrence of subunits of G proteins in plant membranes has been shown using different methodical approaches. One possible approach is the use of radiolabeling. Binding of [3SS]GTFyS, a nonhydrolyzable GTP analog, to crude plant membranes of Lemna paucicostata (Hasunuma and Funadera, 1987),Pisum satiuum (Hasunuma et al., 1987), and Funaria hygrometrica (Zbell et al., 1989) and to the thylakoid membranes of spinach chloroplasts (Millner, 1987)were reported. In the Cucurbita pep0 microsomal membrane (Jacobs et al., 1988) and in the plasma membranes of several other plants (Blum et al., 1988), G proteins were detected by antisera raised against the highly conserved amino acid sequence (G,-common peptide) of animal G,, ( a subunit of stimulatory G protein), Gi, ( a subunit of inhibitory G protein), and G,,. Western blot analysis conducted in several plant species showed sets of proteins with molecular masses of 33 and 50 kDa ( C . pepo), 37 and 31 kDa (Vicia faba), 36 and 34 kDa (Arabidopsis thaliana), and 38 and 34 kDa (Commelina communis). In the eyespots of a green alga Chlamydomonas reinhardtii (Korolkov et al., 1990),a 24-kDa protein cross-reacted with anti-G, antiserum, and 21- and 29-kDa proteins cross-reacted with anti-Gp antiserum were detected. On the other hand, in Dunaliella tertiorecta, 26- and

310

SHOSHI MUTO

36-kDa proteins were detected on immunoblots using anti-G, and anti-G, antiserum, respectively (Takahashi and Muto, unpublished data). Recently, a G protein was purified from the plasma membranes of maize seedling roots (Bilushi et a f . , 1991). The protein had a molecular mass of 61 kDa and consisted of at least two subunits of 34 and 27 kDa. However, a peptide corresponding to the y subunit was not detected. More recently, G protein was identified through ADP-ribosylation of G, catalyzed by pertussis and cholera toxins. Romero et al. (1991) reported that a 24-kDa protein from etiolated A. satiua seedlings was ADPribosylated by cholera toxin. The ADP-ribosylation was found to be sensitive to the presence of GDP or GTP. Furthermore, an antibody to G,cOmmDn showed a high specificity for the 24-kDa ADP-ribosylated protein. A gene (GPAl) coding for a G, from A. thaliana was isolated using a DNA probe generated by a polymerase chain reaction based on a protein sequence from mammalian and yeast G, (Ma et al., 1990). The deduced amino acid sequence showed that the GPAl gene product (GPal) has 383 amino acid residues (44,582 Da). The GPal protein exhibits similarity to all known G, and has all of the consensus regions for a G protein. The GPAl-encoded mRNA is most abundant in vegetative plant tissues. A gene encoding a G,-like protein (Cblp) was isolated from C. reinhardtii by a method similar to that used for isolation of GPAl (Schloss, 1990). The predicted amino acid sequence of the Cblp is similar to that of mammalian, fruit fly, and yeast G,. There is no evidence indicating that G proteins that were detected immunologically, by ADP-ribos ylation or deduced from the gene sequence, consist of heterotrimeric G proteins. The molecular masses of most of immuno detected and ADP-ribosylated proteins were smaller than those of animal G, (39-50 kDa), whereas the molecular masses of GPal deduced from gene sequencing and some immunodetected and ADPribosylated proteins were similar to those of animal G,. Purification and characterization of native heterotrimeric G proteins are necessary to resolve whether the small molecular masses of plant G,’s are native polypeptides or are artificially produced during extraction from cells. After SDS-polyacrylamide gel electrophoresis and electroblotting to nitrocellulose membrane, [a-32P]GTP-and [y-’*P]GTP-binding proteins with molecular masses of 23.4-28.5 kDa were detected in the C. p e p microsomal fraction (Drgibak et a f . , 1988) and in the outer envelope membrane of pea chloroplasts (Sasaki et al., 1991). Similarly, [35S]GTPySbinding proteins with molecular masses of 28 and 24 kDa were detected in soybean plasma membrane (Zbell et a f . , 1990). All these proteins resemble animal proteins with molecular masses of 29-30 kDa, which possess conserved GTP-binding and hydrolysis domains. This family of small GTPbinding proteins include the ras protooncogenes and about 20 other gene

INTRACELLULAR Ca2+ MESSENGER SYSTEM

311

products (Burgoyne, 1989). They may have important roles, possibly in signal transduction (Downward, 1990) and in mediating a variety of intracellular interactions including aspects of organelle traffic in cells (Balch, 1990). The small GTP-binding proteins, unlike the GTP-binding heterotrimeric G,, retain their ability to bind [cx-~~PIGTP or [-Y-~~S]GTPYS after SDS-polyacrylamide gel electrophoresis and transfer to nitrocellulose membranes (Bhullar and Haslam, 1987). The small GTP-binding proteins have also been detected as substrates for ADP-ribosylation by certain botulinum toxins (Bokoch et al., 1988). In Zea mays, DNA sequences homologous to the oncogenes myb, ras, and src have been detected (Zabulionis et al., 1988). For A . thaliana, a putative gene, ara, homologous to the ras-related gene family has been cloned (Matsui et al., 1989). A ras-related rgpl gene was isolated from 5-azacytidine-induced dwarf rice (Oryza sativa) seedlings (Sano and Youssefian, 1991). cDNA of rgpl encodes 226 amino acids with a molecular mass of 24,859 Da. This protein was expressed in transformed Escherichia coli and showed GTP-binding activity. The fundamental role of G protein in Ca2+signaling may be the activation of the Ca2+channel and phospholipase C on the plasma membrane, which subsequently leads to an increase in cytoplasmic Ca2+concentration. GTP stimulation of Ca2+ release from plant microsomes has been reported (Allan et al., 1989). Recently, Fairley-Grenot and Assmann (1991) demonstrated that GDPpS increased inward K+ current through the K+ channel in the plasma membrane of V . f a v a guard cells, in contrast to GTPyS, which has the opposite effect. Reduction of inward current by GTPyS was eliminated by BAPTA, a Ca2+ chelator. Intracellularly applied cholera and pertussis toxins decreased K+ current. These results suggest the involvement of a G protein in the inhibition of inward K+ current, which may have resulted from an associated increase in cytosolic Ca2+concentration. The increase in cytoplasmic concetration may be caused by Ca2+ influx across the plasma membrane and/or by inositol 1,4,5-trisphophate (Imp,)-induced release from intracellular stores via the Ca2+channel (see Section V,E).

IV. Regulation of lntracellular Ca2+ Concentration

Ca2+plays a vital role as a second messenger in intracellular signaling. The cytoplasmic Ca2+concentration of plant cells must be kept as low as lO-'M (Hepler and Wayne, 1985; Bush and Jones, 1990; Felle, 1989) because a small quantitative change causes a 10- to 100-fold increase in the concentration. The cytoplasmic Ca2+concentration is maintained at a

312

SHOSHI MUTO

low level by active extracellular extrusion and sequestration to intracellular organelles, When cells are stimulated, influx of extracellular Ca2+and/ or efflux of Ca2+sequestrated in organelles occur leading to an increase in cytoplasmic Ca2+concentration. These Ca2+fluxes are considered to be mediated by ion channels. Figure 2 summarizes Ca2+transport in plant cells.

A. Ca2+ Pump and H+/Ca2+Antiporter

1. Plasma Membrane A number of reports provided evidence for the presence of an ATPdependent Ca2+transport system in microsomal fractions or partially purified plasma membrane fractions from several higher plants (Butcher and Evans, 1987; Cocucci, 1986; Giannini et af., 1987; Gross and Marme, 1978; Kubowicz el al., 1982; Malatialy et al., 1988; Paliyath and Thompson, 1988; Rasi-Caldogno et a / . , 1987,1989; Williams et a / . , 1990). Most evidence was based on the measurement of ATP-driven 45Ca2+transport into plasma membrane vesicles. 45Ca2+transport was Mg2+-and ATPdependent. Although ATP was the preferred substrate for driving Ca2+ uptake, GTP could drive transport at about 50% of the total level served for ATP. On the other hand, a protonophore, carbonyl cyanide rn-chlorophenylhydrazone (CCCP), was found to have no effect on Ca2+transport. This suggests the absence of a secondary Ca2+transport system driven by

'f

+- H' H+/Ca%ntiporter

ATP

Cflchannel \vacuole

, l

Cd'

ATP

INTRACELLULAR Ca2+ MESSENGER SYSTEM

313

a proton gradient such as the vacuolar H+/Ca2+antiport system (Section IV,A,3). The Ca2+ accumulated in the membrane was released by an ionophore, A23187. The Ca2+transport had K , values for Ca2+ of 1527 p M and optimum pH values of about 7. Furthermore, Ca2+uptake was inhibited by vanadate and erythrosin B. In these studies, the degree of sealing of the plasma membrane vesicles and relative contamination by sealed vesicles from other membrane components were uncertain. Highly pure plasma membrane vesicles can be prepared from a variety of plant tissues by aqueous two-phase partitioning (Larsson, 1983). However, most of plasma membrane vesicles prepared by this method had outside-out sidedness, and thus were not suitable for Ca2+uptake studies. Graf and Weilar (1989) were able to prepare inside-out plasma membrane vesicles from C. communis by aqueous two-phase partitioning. 4sCa2+ uptake into the vesicles had a pH optimum of 7.2 and K , (app) for Ca2+ of 4.4 pM and for Mg-ATP of 30 pM. The CCCP did not inhibit ATPdependent Ca2+transport. Ca2+uptake was inhibited by erythrosin (Is0, 0.1 p M ) , ruthenium red (Is0, 30 pM),La3+ (Is0, 10 pM),vanadate (I,,, 500 p M ) , and calmodulin inhibitors trifluoperazine (I,,, 70 p M )and W-7 (I,,, 100 P M ) . Recently, the presence of both Ca2+-ATPaseand a tonoplast-type H + / Ca2+antiporter was reported in highly purified (essentially tonoplast-free) corn plasma membrane vesicles (Kasai and Muto, 1990) in which outsideout sidedness was reversed by Triton X-100 treatment (Grouzis et al., 1987). This result was supported by the following observations: (a) Ca2+ transport into the membrane vesicles was Mg2+-ATP-dependentand stimulated by C1- of NO,-. These ions also greatly stimulated H+ transport into the vesicles as reported by Churchill and Sze (1983). (b) The Ca2+ uptake stimulated by Cl- was dependent on the activity of H+ transport into the vesicles. (c) The CCCP and vanadate canceled the Cl--stimulated CaZf uptake. (d) Artificially imposed acid inside pH gradient caused Ca2+ uptake into the vesicles. The K , (app) for Ca2+ in this Ca2+transport system was 0.4 p M . A similar H+/Ca2+antiporter was reported in pinto bean leaf plasma membrane (Castillo and Heath, 1990)and well characterized in Neurospora plasma membrane (Stroobant and Scarborough, 1979). Rasi-Caldogno et al. (1987) also observed H+/Ca2+antiport in radish plasma membrane vesicles, but the plasma membrane Ca2+-ATPaseitself was considered to act as a direct ATP-fueled nH+/Ca*+antiporter. Although the plasma membrane Ca2+-ATPasewas identified as a Ca2+ pump based on the similarities between kinetics of the Ca2+-ATPaseand Ca2+uptake into the membrane vesicles (Rasi-Caldongo et al., 1989),there has been no direct evidence to support this notion strongly. Recently, Kasai and Muto (1991) reported solubilization and functional reconstitution of Ca2+pump from corn leaf plasma membrane. The purified plasma

314

SHOSHI MUTO

membrane by aqueous two-phase partitioning was solubilized with a nonionic detergent C,2Es. Ca2+-ATPase was separated from vanadatesensitive Mg2+-ATPase (H+ transporting ATPase) by DEAE anionexchange high-performance liquid chromatography. These ATPases were separately reconstituted into liposomes. The liposomes reconstituted with Ca2'-ATPase showed ATP-dependent Ca2+ uptake but those with the vandadate-sensitive Mg2+-ATPasedid not. In addition, the nigericin-induced acid inside pH gradient caused essentially no Ca2+uptake into the liposomes reconstituted with Ca2+-ATPase. This indicates that the H + / Ca2+antiporter, which coexists in the plasma membrane (Kasai and Muto, 1990), is not present in the reconstituted liposomes. These results direclty indicate that Ca2'-ATPase functions as a Ca2+pump on the plasma membrane. In a related study, an ATP-driven Ca2+-transport system of C. communis plasma membrane was solubilized with CHAPS and functionally reconstituted into liposomes (Graf and Weiler, 1990). The ATP-driven Ca2+ transport system reconstituted into liposomes had characteristics similar to those of plasma membrane including K , (app)for Ca2+,inhibition by vanadate, and optimum pH. The CCCP did not affect Ca2+transport in the reconstituted system. This Ca2+transport in the reconstituted system was primarily active through a Ca2+-transportingATPase. Calmodulin activation of Ca2+uptake into the plasma membrane vesicles (Dieter and Marme, 1980, 1983; Hsieh et al., 1991; Stosic et al., 1983; Zocchi and Hanson, 1983; Zocchi, 1988) has been reported. However, the activation effect is contradictory. ATP-dependent Ca2+uptake into the plasma membrane vesicles from C. communis (Graf and Weiler, 1989) and radish seedling (Rasi-Caldogno et al., 1989) was not activated by calmodulin. Kasai and Muto (1990) reported that Ca2+ uptake into corn leaf plasma membrane was inhibited by calmodulin inhibitors but this inhibition was not overcome by the addition of calmodulin. Calmodulin activation of Ca2+-ATPaseactivity in the plasma membrane (Dieter and MarmC, 1981; Robinson et al., 1988) has been reported. Calmodulin-dependent Ca2+-ATPasewas purified from corn coleoptile by a calmodulin-affinity column (Dieter and Marme, 1981; Briars et al., 1988). On the other hand, exogenously added calmodulin had only a slight activation effect on the corn leaf plasma membrane CaZf-ATPaseeven after the endogenous calmodulin was removed from the enzyme preparation by DEAE anion-exchange chromatography (Kasai and Muto, 1990). This experimental evidence may indicate that calmodulin-dependent and -independent Ca2+-ATPases are present in corn plasma membrane and that distribution of these enzymes is different among tissues. Stosic et al. (1983) reported that ATP-dependent Ca2' transport capacity and the effect of calmodulin on its activity were different in different tissues of spinach.

INTRACELLULAR Ca2+ MESSENGER SYSTEM

315

In addition, calmodulin was also observed in highly purified pea leaf plasma membrane (Collinge and Trewavas, 1989). 2. Endoplasmic Reticulum

ATP-dependent 45Ca2+uptake was also demonstrated in endoplasmic reticulum (ER) membrane isolated from roots of garden cress (Lepidium satiuum) (Buckout, 1983, 1984). The Ca2+transport was directly coupled to ATP hydrolysis, as CCCP did not inhibit the transport. Oxalate greatly stimulated Ca2+ uptake. The K , (app) values for ATP and free Ca2+ were 2.5 and 73 p M , respectively. The pH optimum was 7.5. Calmodulin inhibitors, phenothiazines, inhibited Ca2+-transportactivity (Ki, 35 p M ) , but bovine brain calmodulin did not show alteration in the activity. The inhibition was thought to be related to nonspecific interaction of drugs with the membrane. Using chlortetracycline as a probe, Ca2+uptake into ER was measured (Lew et al., 1986). The Ca2+uptake had a pH optimum of 7.5, a K , for ATP of 0. I mM, and a K , for Ca2+of about 70 nM. Calmodulin stimulated the Ca2+uptake twofold, but this stimulation was due to contamination by a calmodulin-stimulated system derived from the plasma membrane. Vanadate is a strong inhibitor for Ca2+uptake, with an Is,, of 5 mM. Bush and Sze (1986) reported that Ca2+transport in ER vesicles isolated from cultured carrot cells was also inhibited by vanadate with an ZS0 of 12 p M . It is also insensitive to inhibitors of the tonoplast Ca2+ transport system [NO,-, CCCP, N , N'-dicyclohexylcarbodiimide(DIDS), and 4, 4'-diisothiocyano-2, 2 ' 4 l b e n e disulfonate (DCCD)]. The Ca2+transport exhibited cooperative Mg-ATP dependence. Ca2+-dependentkinetics of the transport were complex with K , (app) of 0.7-2 p M . The ER was also isolated and purified from a fluorescent Ca2+indicator indo- 1-loaded protoplasts prepared from barley (Hordeum uulgare) aleuron layers. The steady-state levels of Ca2+within ER was measured (Bush et al., 1989). The Ca2+level in the lumen of ER was at least 3 p M . This concentration is at least 10-15 times higher than cytosolic Ca2+.Ca2+ transport into barley ER was inhibited by vanadate, but insensitive to NO,-, FCCP, DCCD, and nigericin. The K , (app) values for ATP and Ca2+in Ca2+transport were 67 and 0.5 p M , respectively. The Ca2+transport activity into aleuron ER was stimulated several-fold with gibberellic acid (GA,) treatment. Lew et al. (1986) reported GA,, indol acetic acid (IAA) and abscisic acid (ABA) at 10 p M had no effect on Ca2+uptake into zucchini ER when added to the uptake medium. As Ca2+ transport of the plasma membrane and ER has fairly high affinity for Ca2+( K , < p M ) (Bush and Sze, 1986; Bush et al., 1989; Kasai

316

SHOSHI MUTO

and Muto, 1990; Rasi-Caldogno et al., 1989), it is considered that they are responsible for keeping the basal cytoplasmic Ca2+. 3. Vacuole Ca2+transport across vacuolar membrane (tonoplast) has been well established. Rasi-Caldogno et al. (1982a,b) reported two nonmitochondrial systems involved in ATP-dependent Ca2+accumulation in membrane vesicles from pea. In the lighter membrane fraction obtained from a density gradient centrifugation accumulated Ca2+by an ATP-dependent H+/Ca2+antiport system. This system was inhibited by carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) and nigericin, but stimulated by KCI. It was saturated by 0.8-1 mM Mg-ATP and strictly required ATP, but was severely inhibited by free Mg2+or Mn2+.This was the first indication of a H+/Ca2+antiport system in plants. Schumaker and Sze (1985) studied CaZ+accumulation into right-sideout tonoplast vesicles isolated from oat roots. The Ca2+accumulation was insensitive to vanadate and stimulated by 20 mM C1- but inhibited by 10 mM DIDS or 50 mM DCCD. This Ca2+transport system had a K , (app) for Mg-ATP of 0.24 mM. These properties are similar to those of tonoplast ATPase. The Ca2+transport was abolished by compounds such as ionophores and uncouplers, which dissipated the pH gradient. These results suggest that a pH gradient generated by the H+-ATPase drives Ca2+accumulation into the tonoplast vesicles via a H+ICa2+antiport. The properties of the H+/Ca2+antiporter was studied directly by using artificially imposed pH gradients across tonoplast vesicles isolated from oat roots (Schumaker and Sze, 1986).The Ca2+uptake was tightly coupled to Ht loss as increasing CaZ+levels progressively dissipated the steadystate pH gradient. The Ca2+uptake displayed saturation kinetics with a K, (app) for Ca2+of 10 p M . La3+and Mn2+inhibited Ca2+uptake. Ruthenium red and DCCD specifically inhibited the H+/Ca2+exchange. The H+/ Ca2+exchange in tonoplast vesicles is electrogenic, generating an interior positive membrane potential. An artificially imposed interior negative A J, (transmembrane electrical potential) stimulated Ca2+uptake. These results support a simple model of one Ca2+taken up per H+ lost. Blumwald and Poole (1986) studied the kinetics of H+/Ca2+antiport in tonoplast vesicles isolated from red beet storage tissue using artificial pH gradients. The K, (app) for Ca2+was 42 p M . They concluded that the H+/Ca2+exchange is electroneutral (stoichiometric radio, 2 H+/I Ca2+),as K + and valinomycin had virtually no effect on the exchange. Recently, Blackford et af. (1990) showed that the inherently high protonic conductance of tonoplast vesicles can lead to rapid redistrubtion of H+ in response to manipulation of A I+, using red beet tonoplast. To overcome this high H+ permeability, a strategy

INTRACELLULAR Ca2+ MESSENGER SYSTEM

317

that independently controlled ApH (transmembrane pH difference) and A JI was adopted, and evidence that Ca2+uptake by tonoplast vesicles is enhanced by inside-positive A JI was provided. The simplest interpretation of this is that the transport system translocates positive charge in the outward direction, i.e., that the stoichiometric H+/Ca2+ratio for transport is greater than 2 (if n can be taken as an integer, its value must be at least 3). The antiporter is thermodynamically competent to account for Ca2+ accumulation in vacuoles and that its reversal occurs in uiuo is unlikely. The pH gradient and potential difference across the vacuolar membrane are also generated by tonoplast pyrophosphatase (Rea and Sanders, 1987). Recently, Chanson (1991) demonstrated a Ca2+/H+antiport driven by the pyrophosphate-dependent H+ pump in vaculor membrane from maize roots. To identify the protein responsible for H+/Ca2+exchange activity, proteins in tonoplast-enriched vesicles were solubilized by octylglucoside. The H+/Ca2+exchange activity was reconstituted into phospholipid vesicles via detergent dilution (Shumaker and Sze, 1990). Sealed phospholipid vesicles that generate an imposed pH gradient were formed. Reconstituted vesicles took up Ca2+in response to an imposed pH gradient (acid inside). Inclusion of 50 mM oxalate into the reconstitution mixture was necessary to trap Ca2+ inside the liposome. Ca2+ uptake could be prevented by dissipation of pH gradient with gramicidin or Triton X- 100. Reconstituted Ca2+uptake varied as a function of extraliposomal Ca2+concentration. The K , (app) for Ca2+of 10 pM was the same as that obtained for the antiporter in membrane vesicles. Ruthenium red, DCCD, and La3+inhibited Ca2+uptake but have no effect on the formation of the pH gradient. The vacuole occupies 90% of the mature plant cell volume but the affinity for Ca2+ of vacuolar Ca2+uptake is very low ( K , > 10 p M ) . It contains the majority of intracellular Ca2+.This suggests that the vacuole has a high capacity for Ca2+uptake. Internal volume of tonoplast vesicles from barley roots was estimated using an electron-spin resonance technique and intervesicular Ca2+concentrations were estimated to be as high as 5 mM (DuPont et al., 1990). ATP-driven uptake of Ca2+created 800- to 2000-fold concentration gradients within minutes, indicating the importance of vacuoles in intracellular Ca2+sequestration. 4. Mitochondrion

Ca2+uptake by plant mitochondria is dependent on respiratory substrates and/or ATP. It absolutely requires inorganic phosphate. Ruthenium red inhibits Ca2+ uptake. It has been shown that Ca2+ transport by plant

318

SHOSHI MUTO

mitochondria varies in many aspects between different plant species (Chen and Lehninger, 1973; Dieter and Marme, 1989; Day and Wiskich, 1984) and tissues, and ages in the same species (Dieter and Marme, 1989). This may be caused by variable degrees of mitochondrial intactness. Coupled mitochondria isolated from corn coleoptiles accumulated Ca2+in a phosphate-dependent manner (Martins et al., 1986). The Ca2+ uptake was accompanied by a decrease in A $, H + extrusion, and an increase in the for Ca2+influx was about 31 p M . This value rate of respiration. The is at least 10 times higher than cytoplasmic Ca2+concentration. Thus the in viuo contribution of mitochondrial maintenance of cytoplasmic Ca2+ concentration is questioned. 5. Chloroplast

Light-dependent Ca2+uptake into intact chloroplasts isolated from wheat and spinach has been shown (Muto et al., 1982). This Ca2+ influx was stimulated by uncouplers FCCP, CCCP, and nigericin, but inhibited by 3(3,Cdichlorophenyl)-l, 1-dimetylurea (DCMU) and ruthenium red. It was also insensitive to vanadate (Greimer et al., 1985a,b). Based on these results, the light-dependent Ca2+influx into chloroplast is thought to be electrogenic and mediated by a uniport-type carrier. The importance of this Ca2+transport in the regulation of cytoplasmic Ca2+concentration is obscure because the affinity for Ca2+ is very low (K,(app) = 189 pM) (Muto et al., 1982; Kreimer et al., 1985a). 6, Ca2+ Channels

Elevation of the cytosolic Ca2+concentration from resting levels to excitatory levels by stimulus is performed via activation of Ca2+ channels located on the plasma membrane and the membranes of Ca2+sequestrating organelles. Evidence for the existence of a Ca2+channel in plants was first demonstrated in algae. Channels in the plasma membrane that initiate the charophyte action potential in response to various environmental stimuli allow the movement of Ca2+ into the cytoplasm (Hayama et al., 1978; Lunevsky et al., 1983; Kikuyama et al., 1984; Shiina and Tazawa, 1987; Tsutsui, 1990). The Ca2+channel of charophyte is sensitive to 1,Cdihydropyridine (DHP) (Shiina and Tazawa, 1987), an antagonist of one class of voltage-dependent plasma membrane Ca2+channels of animal cells. In animal systems, InsP, acts to release Ca2+from intracellular stores by binding to specific receptors in the ER and may also contribute to the activation of plasma membrane Ca2+channels (Berridge and Irvine, 1989). An InsP,-binding protein that had properties consistent with the receptor

INTRACELLULAR Ca2+ MESSENGER SYSTEM

319

has been isolated from bovine and rat cerebella and functionally reconstituted into liposomes (Supattapone et al., 1988; Ferris et al., 1989). Cloning the InsP,-binding protein cDNA and its functional expression revealed that the same molecule mediates both InsP, binding and the release of Ca2+from the ER (Furuichi et al., 1989; Ferris et al., 1989). In plants, InsP,-induced Ca2+release from the microsomes (Drgbak and Ferguson, 1985; Reddy and Poovaiah, 1987) and vacuoles (Schumaker and Sze, 1987; Ranjeva et al., 1988; Canut et al., 1989) has been observed in various tissues. Lew et al. (1986) reported that InsP, (10 p M ) did not cause the release of sequestered Ca2+from zucchini ER. These observations suggest that InsP, acts on the vacuole but not on ER to release Ca2+ in plant cells. The InsP,-mediated Ca2+eflux from red beet microsomes was inhibited by heparin (Brosnan and Sanders, 1990), which binds to the InsP, receptor in animal cells. This suggests that the InsP, receptors of animals and plants possibly share a common Ca2+ release mechanism despite differences in cellular locations. Alexandre et al. (1990) have reported that InsP, activates Ca2+channel in red beet root vacuoles by patch clamp studies. The Ca2+current was specific to InsP, and dependent on InsP, concentration. They measured an InsP,-activated single Ca2+channel with an outside-out patch. This Ca2+channel has an unusually high conductance of 30 pS when compared to the Ca2+channels of animal cells, which generally have low conductance (Bean, 1989). Contrary to the finding of Alexandre et al. (1990), InsP,dependent Ca2+channel in outside-out patches of vacuolar membranes from tobacco suspension cultured cells and red beet root cells could not be recorded using the same composition of pipet and bath solutions as reported by them (Zhang, Yabe, and Muto, in press). Instead, in the absence of InsP, channels with 38-pS conductance in positive voltage range and with 23-pS and 47-pS in negative voltage range were frequently observed in red beet vacuoles. These channels were obviously not Ca2+ channels because their reversal potentials were 0 in unsymmetrical Ca2+. It has been reported that channel current in positive range (Hedrich and Neher, 1987) and in negative range (Hedrich and Kurkdjian, 1988) could be blocked by high concentrations of Ca2+( > M ) and Zn2+ (> M ) , respectively, in sugar beet vacuole. However, the presence of 1 mM Ca2+plus 0.1 mMZn2+is insufficient to block all vacuolar currents in positive and negative voltage ranges at least in outside-out mode recording in red beet and tobacco vacuoles (Zhang, Yabe, and Muto, in press). Furthermore, K+ and C1- channels in tobacco vacuole were identified under this condition. Recently, a voltage-gated InsP,-insensitive outward rectified Ca2+channel with a 12-pS conductance was found in vacuoles of sugar beet roots ( Johannes et al., 1991). We found another type of Ca2+channel in vacuoles

320

SHOSHI MUTO

of tobacco, which is inward rectified and has a 19-pS conductance (Zhang, Yabe, and Muto, in press). This channel was efficiently inhibited by Cd2+ but was insensitive to La3+,Gd2+,Ni2+,verapamil, and nifedipine. A similar Ca2+channel was also found in red beet vacuoles. They exist in a high density with high activity but their physiological significance is still unknown. Simultaneous photometric measurements of cytosolic Caf2 and patch clamp recording of ion currents across the plasma membrane of single V . faba guard cells were conducted to distinguish signal-dependent activation of Ca2+ channels in the plasma membrane from release of Ca2+from intracellular organelles (Schroeder and Hagiwara, 1990). It was demonstrated that ABA induced transient repetitive increases in cytosolic Ca2+, which were accompanied by concomitantly occurring increases in an inward-directed ion current. Since depolarization terminated both increases of cytosolic Ca2+and inward ion current, ABA-mediated Ca2+transients were considered to be produced by passive influx of Ca2+from the extracelMar space through Ca2+-permeablebut nonselective channels. The InsP, sensitive Ca2+channel will be further described in Section V,E. Binding of Ca2+channel blocker to plant membrane fraction has been reported (Hetherington and Trewavas, 1984; Andrejauskas ef al., 1985, 1986; Dolle and Nultsh, 1988). Graziana ef al. (1988) examined inhibitory effects of various Ca2+channel blockers on Ca2+entry into carrot protoplasts. Phenylalkylamines, diphenylbutylpiperidines, and bepridil were the most efficient inhibitors, whereas DHPs or diltiazem had no effect. High-affinity membrane receptors for these compounds were present on the plasma membrane. A phenylalkylamine verapamil-binding protein was partially purified from maize coleoptile membranes (Harvey et al., 1989).The major component of verapamil-binding protein was a 169-kDa glycoprotein, which has a mass similar to that of the verapamil-binding subunit of the L-type Ca2+ channel in animal tissues. A 75-kDa polypeptice was identified as a phenylalkylamine calcium channel blocker (LU 49888)-binding protein which is located mainly at the plasma membrane (Thuleau et al., 1990). This protein differs significantly from its animal counterpart in size. To verify whether the verapamil-binding proteins are Ca2+channels, reconstitution of these proteins in artificial membranes and measurements of Ca2+current are needed. Patch clamp recording of Ca2+current in reconstituted verapamil-binding proteins appeared recently in a meeting abstract (Thuleau et al., 1991). Ion channels that are activated by stretching of the plasma membrane have been demonstrated in several plants (Hedrich and Schroeder, 1989; Tester, 1990). Recently, three distinct stretch-activated channels, permeable to C1-, K', and Ca2+,were recorded by outside-out patches of plasma

INTRACELLULAR Ca2+ MESSENGER SYSTEM

321

membranes of guard cells of V.faba (Cosgrove and Hedrich, 1991). They were distinguishable from spontaneous channels for these ions on the basis of conductance, kinetics, voltage-dependence, and sensitivity to membrane stretch. The stretch-activated Ca2+channels had a low conductance (3 pS) and high selectivity over K + . The stretch-activated channels may play important roles in osmoregulation and sensing mechanical perturbation of plant cells.

V. Phosphatidylinositol Turnover In animal cells, phosphatidylinositol (PI) turnover occurs following stimulation by certain agonists. Agonists bind to specific receptors. Cleavage of phosphatidylinositol4,5-bisphosphate[PI(4,5)P2]by a specific phospholipase C coupled to a G protein produces the second messengers InsP, and diacyl glycerol (DG). DG stimulates protein kinase C and/or is metabolized via other metabolic pathways. The InsP, mobilizes Ca2+from intracellular stores and brings about increase in cytoplasmic free Ca2+concentration. Thus the PI turnover is tightly linked to Ca2+signaling. In plant cells, evidence suggesting the importance of PI turnover in signal transduction is accumulating. If PI turnover functions during transduction after certain stimuli in plant cells, then the following must occur: (a) localization of inositol phospholipids and enzymes involved in their metabolism [PI and phosphatidylinositol 4-phosphate (PIP) kinases and phospholipase C] in the plasma membrane; (b) transient changes in levels of polyphosphoinositides and their hydrolysis products (especially InsP, and DG) by stimuli; (c) increases of free Ca2+in cytosol; (d) activation of protein kinase C by Ca2+and DG, and activation of other Ca2+-dependent enzymes. A. Presence of lnositol Phospholipids and lnositol Phosphates

The presence of inositol phospholipids and inositol phosphates in plant tissues has been established (Boss and Massel, 1985; Heim and Wagner, 1986; Morse et al., 1987;Drgbak et al., 1988; Irvine et al., 1989). Phosphatidylinositol is the most common phosphoinositide in plants but quantities of PIP and PIP, in most cells are very low. They were identified and quantified based on radiolabeling with [3H]in~~itol or [32P]followed by cochromatography with authentic standards using one or several thinlayer chromatography systems. Even using this method some investigators

322

SHOSHI MUTO

have failed to identify PIP, in certain cultured cells. Inositol phosphates in extracts from [3H]inositol-labeledcells were separated and identified by anion-exchange high-performance liquid chromatography. Identification and quantification of InsP, require great care because the cellular level of InsP, is extremely low. In addition, inositol metabolites such as glucuronic acid (Cot6 et al., 1987)interfere with the separation of inositol phosphates. Problems in extraction, separation, and characterization of the inositol phospholipids and inositol phosphates were recently reviewed by Cote et al. (1990). B. Phosphatidylinositol and Phosphatidylinositol 4-phosphate Kinases

Sandelius and Sommarin (1986)have established the presence of phosphoinositide kinases in plant membranes. The microsomes isolated from wheat phosphorylated PI to form PIP and PIP to PIP, when supplied with [y-32P]ATP.Enrichment of the phosphorylation activity in the plasma membrane was demonstrated by aqueous two-phase partitioning. Phosphatidylinositol and PIP kinases of wheat required 5-15 m M Mg2+. The Mg2+requirement was partially substituted by Mn2+but not by Ca2+, which partially inhibited at 100 p M (Sommarin and Sandelius, 1988). Recently, Kamada and Muto (1991) demonstrated that submicromolar concentrations of Ca2+inhibited PI and PIP kinases in the plasma membrane from tobacco cultured cells. The presence of DG kinase in addition to PI and PIP kinases in plants has been reported in suspension cultured cells of Catharanthus roseus (Heim et al., 1987) and Nicotiana tabacum (Heim et al., 1987; Kamada and Muto, 1991). C. Phospholipase C

Soluble and membrane-bound forms of phospholipase C have been demonstrated in plants (Pfaffmann et al., 1987). The membrane-bound enzyme of celery hydrolyzes PIP, to release InsP, (McMurray and Irvine, 1988). Melin et al. (1987) have reported the presence of polyphosphoinositidespecific phospholipase C in highly purified plasma membranes from wheat seedlings. The enzyme preferentially hydrolyzed PI(4)P and PI(4,5)P, and had a phosphoinositide substrate profile different from that of soluble phospholipase C. Plasma membrane location of phosphoinositide-specific phospholipase C has also been reported in soybean and bushbean hypocotyls (Pfaffman et al., 1987), oat root (Tate et al., 1989), tobacco suspension cultured cells (Kamada and Muto, 1991), and a green alga, Dunaliella

INTRACELLULAR Ca*+ MESSENGER SYSTEM

323

salina (Einspahr et al., 1989).The enzyme was sensitive to Ca2+;however, the optimum concentrations were different among plant species. Phospholipase C of celery was markedly activated at a 0.1-1 pM Ca2+range but full activation was produced at a millimolar range (McMurray and Irvine, 1988). Soybean and bushbean enzymes were maximally active at 0.5 mM Ca2+(Pfaffman er al., 1987), whereas Dunaliella and wheat enzymes were active at 100 and 10 pM Ca2+,respectively. Phospholipase C activity of tobacco was markedly activated at a submicromolar Ca2+range (Kamada and Muto, 1991). Activation of phospholipase C and inhibition of PI and PIP kinases of tobacco plasma membrane by submicromolar Ca2+concentrations suggest the regulation of PI turnover in the plasma membrane by Ca2+.Coupled to a signal, phospholipase C hydrolyzes PIP, and produces InsP,, which induces Ca2+ mobilization from the intracellular pool raising the Ca2+ concentration in the cytoplasm. Ca2+ in turn activates Ca2+-dependent enzymes, especially phospholipase C. Consequently more InsP, is synthesized and causes further increase of cytoplasmic Ca2+.Whereas, PI and PIP kinases are inhibited by Ca2+,less PIP, is synthesized. Thus activated phospholipase C is limited in its substrate and produces less InsP,. In this way Ca2+works as a feedback inhibitor for the regulation of cytoplasmic Ca2+concentration via PI turnover. Dillenschneider et al. (1986) reported that release of inositol phosphates from membrane-isolated Sycamore suspension cultured cells that had been labeled with [3H]inositol was stimulated by guanine nucleotides in a dosedependent manner. Einspahr et al. (1989)demonstrated PIP,-specific phospholipase C of Dunaliella plasma membrane to be activated by GTPyS over a wide range of Ca2+ concentrations. These reports suggest the involvement of G protein in activation of phospholipase C. However, many investigators have failed to observe guanine nucleotide stimulation of inositol phospholipid breakdown in their assay systems (Melin et al., 1987; McMurray and Irvine, 1988; Biffen and Hanke, 1990; Kamada and Muto, 1991). D. Hydrolysis of lnositol 1,4,5-trisphosphate

In mammalian systems, the InsP, response is attenuated by the rapid metabolism of InsP,. Ins( 1,4)P,, Ins( 1,3,4)P,, and Ins( 1,3,4,5)P, are frequently found as the prevalent inositol phosphate isomers shortly after agonist stimulation (Shears, 1989). Memon et al. (1989a) examined the metabolism of exogenously added [,H]Ins( 1,4,5)P, in microsomal and soluble fractions of carrot suspension cultured cells. The Ins (1 ,4,5)P, was metabolized to InsP, (83%) and InsP

324

SHOSHI MUTO

(6%) in the microsomal fraction. When InsP, was added to the soluble fraction, approximately equal amounts of InsP, and InsP were recovered. The soluble fraction of suspension cultured tobacco cells hydrolyzed InsP, InsP,, and InsP, (Joseph et al., 1989).The major products of Ins (I ,4,5)P3 hydrolysis were Ins( 1,4)P2and Ins(4,5)P2.The Ins( 1,4)P2was hydrolyzed extensively to Ins(4)P. A micromolar Ca2+range markedly stimulated the hydrolysis of all the inositol phosphates. Half-maximal stimulation of Ins( 1 ,4)P2hydrolysis was obtained at a micromolar concentrations of Ca2+. Li+, which is an inhibitor of mammalian inositol monophosphatase (Hallcher and Shermann, 1980), had no effect on the tobacco enzyme. The insensitivity to Li+ has also been reported with a monophosphatase from Lillium pollen (Gumber et al., 1984). 1,4,5)P3in a Drobak et al. (1991) examined the metabolism of [32P]Ins( pea root soluble fractions. The Ins( 1 ,4,5)P, was rapidly converted into lower and higher inositol phosphates. The major dephosphorylation product was Ins(4,5)P2,whereas Ins( 1,4)P2was minor product. Small amounts of nine other metabolites including inositol and Ins( 1,4,5,X)P4were produced. Conversion of Ins( 1 ,4,5)P3to Ins(4,5)P2and Ins( 1,4,5,X)P4 was inhibited by 55 p M Ca2+and stimulated by lowering the Ca2+concentration to 40 nM. These reports demonstrate that the enzymes that rapidly convert Ins(1,4,5)P, are present in plants and that pathways of inositol phosphate metabolism in plants may be different from that in mammals.

E. lnositol 1,4,5-trisphosphate-lnduced Ca2+ Release from lntracellular Stores

Drobak and Ferguson (1985) demonstrated that up to 30% of Ca2+that had been taken up by microsomes isolated from zucchini hypocotyls in the presence of ATP was released by 19 p M InsP,. This release occurred within 0.5 min and was followed by a slower reuptake of Ca2+.Similar results were obtained with corn coleoptile microsomes (Reddy and Poovaiah, 1987). An InsP,-induced release of Ca2+from tonoplast-enriched vesicles of oat roots was reported (Schumaker and Sze, 1987). Ca2+accumulated in the vesicles via a H+/Ca2+antiporter system that is driven by the tonoplast H+-pumpingATPase. It was released by InsP, but not Ins(1,4)P2or Ins( l)P. The release was dependent on InsP, concentration with a K,,,(app) of 0.6 p M . An InsP,-dependent Ca2+release has also been observed with intact vacuoles isolated from A. pseudoplatanus suspension cultured cells (Ranjeva et al., 1988).These results suggest that at least the vacuole is the target organelle for InsP,.

INTRACELLULAR Ca2 + MESSENGER SYSTEM

325

In several animal systems, InsP, provides a link between membrane receptors and the release of Ca2+from intracellular stores, particularly the endoplasmic reticulum (Berridge and Irvine, 1989). Although the plant InsP, receptor has not yet been investigated, comparison of plant and animal receptors is of interest because of their different intracellular locations. A dose-dependent InsP,-induced increase of inward Ca2+current was observed in perfused tonoplast-free cells of Nitella syncurpa (Zherelova, 1989). This Ca2+influx was accompanied by the appearance of an inward CI- current. Theil et al. (1990) introduced InsP, into the cytoplasm of characean algae Chara and Nitella; InsP, induced depolarization of the membrane potential. The effect of InsP, was abolished when EGTA was added. An increase in cytoplasmic Ca2+ induced action potential in a manner similar that of InsP,. These results suggest that characean cells have an InsP,-sensitive system that is able to affect membrane transport uia an increase in cytoplasmic CaZ+concentration. An InsP3-induced Ca2+ efflux from fusogenic carrot protoplasts has been reported (Rincon and Boss, 1987). Release of Ca2+from internal stores was postulated, but it is unlikely that InsP, permeates across the plasma membrane to act on the storage organelles. F. Physiological Responses Involving Phosphatidylinositol Turnover

Rapid changes in phosphoinositide metabolism have been reported during the initiation of the cell cycle. Cytokinin-induced PI synthesis was observed in cytokinin-starved soybean suspension cultured cells that respond to kinetin treatment by reinitiating cell division (Connett and Hanke, 1987). However, PIP, was not detected in the cells. Ettlinger and Lehle (1988) reported that 2,4-dichlorophenoxyaceticacid (2,4-D) stimulated rapid and transient (within minutes) changes in [3H]inositol-labeled InsP, and InsP, in C . ruseus suspension cultured cells that were arrested in the G, phase. However, their results showed that radioactivities recovered in InsP, were much greater than those in PIP before stimulation, and increased to about 270% 1 min after stimulation. These results may indicate that conversion of PI to PIP and hydrolysis of PIP to InsP, are very rapid, and hydrolysis of InsP, is limited. Heim and Wagner (1989) demonstrated that growing cells of C . roseus incorporated [3H]inositol in PIP, and InsP, but growtharrested cells did not. Recently, Grabowski et al. (1991) investigated [,HIinositol incorporation in auxin-starved C. ruseuscells under partially modified experimental conditions of Ettlinger and Lehle (1988), and observed that 2,4-D caused rapid and transient changes in PIP and InsP, only. The

326

SHOSHI MUTO

amount of labeled InsP, and PIP, was too small to be quantified. The amount of inositol phosphates did not exceed that of labeled phosphoinositides. They also demonstrated that 2,4-D induced a rapid increase of PI kinase activity but the magnitude was only 20%. These results may indicate that 2,4-D-induced PI turnover is an essential process for biogenesis of membranes, which is required for cell division. MorrC et al. (1984a) also reported that PI degradation in soybean membrane is enhanced by 2,4-D but not by 2,3-D. An increase in cytoplasmic Ca2+levels should be shown to prove the second messenger role of InsP, in signal transduction of 2,4D-induced cell division. Auxin-stimulated phosphatidylinositol turnover was reported with microsomes from etiolated soybean hypocotyls (Sandelius and MorrC, 1987). This turnover activity is distinct from the usual de nouo biosynthetic pathway, but closely resembles phosphatidylinositol-myo-inositol exchange activities of ER (Sexton and Moore, 1981). Release of inositol phosphates from phospholipids prelabeled with [ y -32P]ATPin carrot microsomal membranes was stimulated by IAA (Zbell and Walter-Back 1988). Gibberellic acid stimulation of PI metabolism has been reported in barley aleurone layers (Murthy et al., 1989). Dunaliella salina grows in saline environments ranging from 0.5 to 5 M NaCl and successfully accommodates rapid and drastic changes in extracellular osmolarity (Ginzburg, 1987). Einspahr et al. (1988b) have investigated osmotic shock-induced transient polyphosphoinositide turnover in Dunaliella. Within 2 min after a sudden hypoosmotic shock, the levels of [32P]-prelabeledPIP, and PIP dropped to 65 and 79%, respectively. This was paralleled by a rapid elevation in the level of phosphatidic acid. Increase of phosphatidic acid level can be explained by either activation of phospholipase D or activation of phospholipase C followed by utilization of the resulting DG for phosphatidic acid synthesis by DG kinase. Ha and Thompson (1991) showed that the plasma membrane DG content of Dunaliella increased by 40% during the first 30 sec following hypoosmotic shock. Molecular species of DG that increased in amount after hypoosmotic shock was characteristic of the plasma membrane PI, PIP, and PIP,. It is tempting to postulate the involvement of PI turnover in transduction of hypoosmotic shock since Dunaliella plasma membrane has phospholipase C which is activated by micromolar range of Ca2+and further by GTPyS (Einspahr et al., 1989). On the other hand, hyperosmotic shock induced a rise in PIP2levels to 131% of control values, whereas the level of phosphatidic acid dropped to 56% of controls (Einspahr et al., 1989). Hyperosmotic shock caused marked increases of lysophophatides and lysophosphatidic acid, suggesting activation of phospholipase A (Einspahr et al., 1988a).

INTRACELLUUR Ca2+ MESSENGER SYSTEM

327

Involvement of InsP, in stornatal movement was demonstrated using an inactive photolabile precursor caged InsP, (P5-1 -(2-nitrophenyl)ethyl ester of InsP,). Blatt et al. (1990) showed that InsP, released from caged InsP, by photolysis in guard cells of V. faba reversibly inactivated K+ channels that were thought to mediate K+ uptake, while simultaneously activating an inward current to depolarize the membrane potential and promote K + efflux through a second class of K+ channel (Blatt, 1988). Using the fluorescent calcium indicator fura-2, McAinsh et al. (1990) have reported that ABA induces a rapid increase in cytosolic Ca2+in guard cells of C. communis and that this increase precedes stornatal closure. Gilroy et al. (1990) measured InsP,-induced changes in cytosolic Ca2+of C. communis guard cells using the fluorescent Ca2+indicator fluo-3, monitoring stornatal aperture. Increasing cytosolic Ca2+ concentration greater than 600 nM induced stornatal closure. Similarly, release of InsP, from the caged form initiated an increase in cytosolic Ca2+concentration followed by stomatal closure. An increase of cytosolic Ca2+ preceded stornatal closure even when guard cells were pretreated with La3+,which inhibits Ca2+influx at the guard cell plasma membrane. This observation suggests that Ca2+ was mobilized from internal stores. On the other hand, Schroeder and Hagiwara ( 1990) suggested that ABA-induced transient repetitive increases in cytosolic Ca2+of V. faba guard cells are produced by passive influx of Ca2+from the extracellular space through Ca2+-permeablechannels. However, the possibility that release of Ca2+from intracellular organelles contributes to the observed rise in cytosolic Ca2+cannot be excluded. Gilroy et al. (1991) observed in C. communis guard cells that ABA induced an increase in cytosolic Ca2+ concentration in a minority of cells, even though stornatal closure always occurred. This suggests the presence of both Ca2+-dependent and -independent transduction pathways linking ABA perception to stornatal closure. Enhancement of light-induced stornatal opening and induction of stomatal opening under darkness by DG was reported in C. communis (Lee and Assmann, 1991). An inhibitor of protein kinase C, H-7, inhibited the lightinduced stornatal opening and enhanced dark-induced stomatal closure. DGs also activated an ATP-dependent voltage-independent current of V. faba guard cell protoplast. Leaflets of Samanea saman open and close rhythmically, a process that is driven by an endogenous circadian clock. Light has a rapid direct effect on the movements and this response is under the control of phytochrome. Morse et al. (1987) demonstrated that a brief (5-30 sec) irradiation with white light decreased PIP and PIP, and increased InsP, and InsP, in Samanea motor organ (pulvini) labeled with [3H]inositol. A transient increase of DG level after 30 sec of white-light illumination was also reported in Samanea (Morse et al., 1989). Light-induced rapid decreases in PIP and

328

SHOSHI MUTO

PIP, accompanied with an increase in InsP, and InsP, were also reported in etiolated Brassica oleracea seedlings (Acharya et al., 1991). Elicitor treatment of cultured cells leads to increases in the amount and activity of mRNAs encoding enzymes of phytoalexin biosynthesis (Ryan and Farmer, 1991). The accumulation of these mRNAs was shown to result from transient increases in their transcriptional rates. Strasser et al. (1986) reported no significant influence of elicitor treatment on hydrolysis of polyphosphoinositides in soybean and parsley cells. On the contrary, Kurosaki et al. (1987) reported that elicitor treatment of carrot cultured cells increased phosphatidylinositol-degrading phospholipase activity without a notable lag. Memon et al. (1989b) found that PIP and PIP, stimulated the activity of the vanadate-sensitive ATPase associated with the plasma membranes from sunflower hypocotyls and carrot suspension cultured cells. They further demonstrated a light-induced in uiuo decrease in plasma membrane PIP kinase activity in etiolated hypocotyls (Memon and Boss, 1990). This decrease was not due to an increase in phospholipase C, since PIP2 level was lowered without increases in InsP, and InsP,. The decrease in PIP kinase accompanied a decrease in the vanadate-sensitive H+-ATPase. These results suggest an alternative mechanism that involves inositol phospholipids as direct membrane effectors rather than as a source of InsP, and DG.

VI. Calmodulin

When intracellular Ca2+concentration is increased by a stimulus, Ca2+ binds to receptor proteins and acts as the second messenger. The Ca2+ receptor proteins have high affinity for Ca2+.Upon binding Ca2+ they change in their ability to interact with other proteins and in their conformation. Calmodulin is the most ubiquitous Ca2+receptor protein with four Ca2+binding sites and mediates Ca2+messages in most eukaryotic cells (Cheung, 1980). Calmodulin is a highly conserved protein with a molecular mass of 17 kDa and is composed of 148 amino acid residues (Roberts et a f . , 1986). Amino acid sequences of spinach (Lukas et a / . , 1984) and wheat (Toda et al., 1985) calmodulin have been reported; they have minor differences to vertebrate calmodulin sequences. The cDNAs encoding calmodulin have been cloned from cDNA libraries of potato (Jena et al., 1989) and barley (Ling and Zielinski, 1989). Two cDNA clones encoding calmodulin isoforms were isolated from Arabidopsis (Ling er al., 1991). Deduced amino acid sequences of the two isoforms share 97% identity. The nucleotide sequences encoding the two

INTRACELLULAR Ca2+ MESSENGER SYSTEM

329

isoforms share 87% sequence identity. Braam and Davis (1990) reported that at least four touch-induced ( T C H ) genes were expressed in Arabidopsis in response to water spray, subirrigation, wind, touch, wounding, or darkness. TCHl cDNA encoded the putative Arabidopsis calmodulin and TCH2 and TCH3 were calmodulin-related genes. The calmodulin-Ca2+ complex activates several plant enzymes directly or indirectly via protein phosphorylation (Dieter, 1984; Poovaiah and Reddy, 1987). However, little is known about calmodulin-dependent physiological processes in plants. In this review the role of calmodulindependent NAD kinase in the regulation of nicotinamide coenzyme levels in green plant cells is described. Plant calmodulin was found during the course of studies on light-induced conversion of NAD' to NADP' in green plant cells. The light-induced conversion of NAD' to NADP+ was first discovered in Chlorella cells (Oh-hama and Miyachi, 1959) and a similar conversion was reported with higher plant leaves (Ogren and Krogman, 1965). The conversion reaction in Chlorella cells was extensively studied in relation to photosynthesis and the association of the reaction with photosynthetic electron transport and photophosphorylation was demonstrated (Matsumura-Kadota e f al., 1982). Since NAD kinase is the only enzyme that phosphorylates NAD+ to NADP', it was proposed that this enzyme catalyzed the conversion reaction using photochemically produced ATP. To elucidate the controlling mechanism of NAD kinase activity, purification and characterization of the enzyme were carried out (Muto and Miyachi, 1977). NAD kinase from pea seedlings lost its activity on a DEAE-cellulose column due to dissociation of an activator protein from the enzyme. The purified enzyme, which had essentially no activity, regained the activity by the addition of activator. Ca2+-dependency of NAD kinase activation by the activator protein was shown by Anderson and Cormier (1978). They also indicated that the NAD kinase activator stimulated the activity of the modulator protein (ca1modulin)-dependent cyclic nucleotide phosphodiesterase. Subsequently, the NAD kinase activator was identified as calmodulin because of the striking similarities of its properties to those of animal calmodulin (Anderson e f al., 1980). The light-induced conversion of NAD' to NADP' was further investigated with higher plant leaves (Muto et al., 1981). The conversion reaction was observed in intact leaves, mesophyll protoplasts, and intact chloroplasts from wheat seedlings. Subcellular fractionation of mesophyll protoplasts from several plants showed that most NAD kinase was localized in the chloroplasts (Muto e f al., 1981). Therefore, the conversion of NAD' to NADP+ was considered to occur in the chloroplast. Subcellular distribution of calmodulin within wheat leaf cells was analyzed by activation of calmodulin-free pea NAD kinase and radioimmuno-

330

SHOSHI MUTO

assay (Muto, 1982). Most of the cellular calmodulin was localized in the cytosol (ca. 90%), and the rest in mitochondria1 (5-9%), chloroplast (1-2%), and microsomal (< 1%) fractions. Calmodulin concentration in the chloroplast, which was calculated assuming a stromal space of 30 pl/ mg chlorophyll and that all chloroplast calmodulin is present in the stroma, was 1-4 pg/ml and sufficient to saturate the chloroplast NAD kinase = 22 ng/ml) (Muto, 1983). Based on these findings and an observation that the intact chloroplasts from wheat and spinach leaves actively took up Ca2+when illuminated (Muto et al., 1982; Kreimer et al., 1985a,b), it was postulated that the lightinduced conversion of NAD' to NADP' was catalyzed by calmodulindependent NAD kinase and was regulated by Ca2+flux into the stroma in the light. Jarrett et al. (1982) reported that a calmodulin-like protein was present in the stroma of pea chloroplasts and that trifluoperazine, a calmodulin antagonist, inhibited the light-induced conversion of NAD+ to NADP+ at a concentration that was also required for calmodulin inhibition. These findings were consistent with the above hypothesis. However, this hypothesis was questioned by the finding of calmodulin-independent NAD kinase in the chloroplasts. Simon et al. (1982) reported that NAD kinase was located in the cytoplasm and chloroplasts of spinach leaves and that the cytoplasmic enzyme was calmodulin-dependent, whereas the chloroplast enzyme was calmodulin-independent and located exclusively in the stroma. Simon et al. (1984) further showed that the calmodulin-dependent and -independent NAD kinases were present in the envelope and the stroma of pea chloroplasts, respectively. The intracellular location of calmodulin-dependent and -independent NAD kinases differs from plant species to species (Muto and Miyachi, 1986). In pea leaf mesophyll cells, 64% of the total NAD kinase located in the chloroplasts and the rest was present in the cytoplasm. The calmodulindependent activity of chloroplasts constituted 40% of the total NAD kinase activity and this activity was located exclusively in the outer and inner membranes of envelope and was completely inactivated when intact chloroplast was treated with trypsin. The stromal NAD kinase was calmodulinindependent and insensitive to the trypsin treatment of chloroplasts. Spinach showed similar intracellular distribution of the total NAD kinase, but the chloroplast NAD kinase was mostly calmodulin-independent and was present in the stroma. With spinach, which was obtained from a local market, NAD kinase was sometimes located exclusively in the chloroplast and most of the activity was calmodulin-dependent. No calmodulin was detected in the chloroplast envelope. The outer envelope membrane was permeable to [ '251]-labeledcalmodulin but the inner membrane was not. This suggests that NAD kinase in the envelope is accessible to calmodulin. The stroma prepared from the purified intact chloroplasts contained only a negligible amount of calmodulin. Therefore,

331

INTRACELLULAR Ca2+ MESSENGER SYSTEM

NAD kinase in the stroma may not be accessible to calmodulin. The small amount of calmodulin detected in the intact chloroplasts (Muto, 1982; Jarrett e? al., 1982) may be due to the cytoplasmic calmodulin that is present between the outer and inner envelope membranes and the surface of the outer membrane. The light-induced conversion of NAD+ to NADP+ occurred irrespective of calmodulin dependency of NAD kinase in the chloroplasts (Muto and Miyachi, 1986). The conversion was inhibited by trifluoperazine in a dosedependent manner (Muto and Miyachi, 1986; Jarrett e? d.,1982). This suggests that the effect of calmodulin inhibitor is not mediated by calmodulin but caused by the inhibition of electron transport through photosystem I1 (Burr ef af.,1982). The stromal NAD kinase was activated when intact chloroplasts were illuminated (Muto et al., 1981; Muto and Miyachi, 1986) or the stromal extract was incubated with dithiothreitol (S. Muto, unpublished results). These findings suggest that the activation of stromal NAD kinase is mediated by the ferredoxin/thioredoxin system (Buchanann, 1980). Taken together the conversion of NAD+ to NADP' is considered to be a reaction catalyzed by photoactivated stromal calmodulin-independent NAD kinase using ATP produced by photophosphorylation. The NADP' produced is reduced by photosystem I and used for the operation of the CO, fixation cycle (Fig. 3). The topology in the envelope of NAD kinase

CaM-dependt

co2

1

PGA

N

T

'I

NADP+ GAP

4

FIG.3 Roles of calmodulin-dependent and -independent NAD kinases in regulation of chloroplast and cytoplasmic nicotinamide coenzyme levels. Calmodulin-dependent NAD kinase is present in the envelope and supplies NADP+ to the cytoplasm. Calmodulin-independent NAD kinase is present in the stroma and supplies NADP+ to the chloroplast. For details see text. DHAP, dihydroxyacetone phosphate; diPGA, diphosphoglyceric acid; GAP, glyceraldehyde 3-phosphate; PGA, 3-phosphoglyceric acid; RuBP, ribulose 1.5-bisphosphate

332

SHOSHI MUTO

is not clear, but the enzyme must be active only at the cytoplasmic side because it was inactivated by the trypsin treatment of the intact chloroplasts (Muto and Miyachi, 1986) and can access calmodulin only at the cytoplasmic side. Thus NAD kinase located in the envelope may function to supply NADP' to the cytoplasm. Light-induced Ca2+influx into spinach protoplasts, which is dependent on photosynthetic electron transport, has been reported (Heimann et al., 1986), suggesting a transient increase of cytoplasmic free Ca2+upon illumination. This coyld cause the activation of calmodulin-dependent NAD kinase in the envelope leading an increase of cytoplasmic NADP+ concentration. NADP+ may be used for the operation of a triose phosphate/3phosphoglycerate shuttle system facilitated by the phosphate translocator at the inner envelope membrane (Heldt, 1976). NADP' is reduced by irreversible NADP+-dependent glyceraldehyde-3-phosphate dehydrogenase leading an indirect transport of reducing power from the chloroplast to the cytoplasm (Fig. 3). The reducing power may be used for biosynthetic pathways in the cytoplasm.

VII. Protein Kinases A. Ca2+-DependentProtein Kinases

1. Protein Kinase C

In animals there are at least two kinds of Ca2+-dependentprotein kinases. These are Ca*'-dependent protein kinases, which are activated by Ca2+-calmodulin complex, and protein kinase C, which is a Ca2+-and phospholipid-dependent enzyme and is activated by DG (Nishizuka, 1984). A small amount of DG markedly increases the apparent affinity of protein kinase C for Ca2+. The enzyme exists in both soluble and membrane fractions of cell extracts. When cells are stimulated, protein kinase C is transiently activated by DG, which is produced in the membrane during signal-induced turnover of phosphoinositides (Section V). The activated enzyme translocates from the cytoplasm to the plasma membrane. In plants, a protein kinase identical to protein kinase C has not yet been found but the presence of a protein kinase C-like enzyme has been demonstrated. Phosphorylation of endogenous substrate of a partially purified protein kinase of wheat suspension cultured cells was activated fourfold by phosphatidylserine plus phorbol ester, which mimics DG in the presence of Ca2+ (Olah and Kiss, 1986). In uiuo phosphorylation of 27- and 32-kDa proteins in protoplasts prepared from etiolated oat was

INTRACELLULAR Ca2+ MESSENGER SYSTEM

333

stimulated by red-light irradiation (Park and Chae, 1990). Phosphorylation was inhibited by a protein kinase C inhibitor, H-7, but was enhanced by DG or phorbol ester. Elliott and Kokke (1987b) reported that a protein kinase purified from Amaranthus tricolor extracts was activated by a micromolar concentration of Ca2+.Phospholipid and diolein further activated the enzyme depending on their concentrations and phospholipid species. The K , for Ca2+ of the enzyme decreased in the presence of phospholipid and further decreased in the presence of phospholipid plus diolein. The enzyme associated reversibly with inside-out erythrocyte membrane vesicles (Elliott and Kokke, 1987b) and plant membranes (Elliott et a f . , 1988) in a Caz+-dependentmanner. Antiserum raised against the regulatory domain of bovine brain protein kinase C cross-reacted with three major bands on a nitrocellulose blot of partially purified enzyme (Elliott and Kokke, 1987a; Elliott et a f . , 1988). The molecular cloning of genes encoding protein kinases from plants provides an important tool to elucidate their expression pattern and biological functions. Oligonucleotides corresponding to conserved regions of animal protein-serinehhreonine kinases were used to isolate cDNAs encoding plant homologs. The cDNA clones encoding putative kinase have been obtained from bean and rice (Lawton er al., 1989) and maize (Biermann et al., 1990). The encoded proteins have sequence similarity to the catalytic domains of protein kinase A and protein kinase C. However, the regulatory domains have no similarity to any known protein kinase. The polymerase chain reaction has been applied to amplify cDNA synthesized from poly(A)+ RNA from pea seedlings using degenerate oligonucleotides corresponding to conserved regions within the catalytic domain of known protein-serinehhreonine kinases (Lin er a f . , 1991). The deduced amino acid sequences from obtained cDNA are equally similar to those of protein kinase A and protein kinase C.

2. Ca2+-and Calmodulin-Dependent Protein Kinases CaZ+-and calmodulin-dependent protein kinases have been demonstrated in several plants (Blowers et a f . ,1985; Blowers and Trewavas, 1987; Polya and Davies, 1982; Polya et a f . , 1983; Polya and Micucci, 1984; Salinath and Marme, 1983), but have not been purified to homogeneity and characterized from plant sources. At least seven Caz+/calmodulin-dependent protein kinases have been reported in animals (Fujisawa, 1990). Among them Ca2+/calmodulin-dependentprotein kinase I1 has been found in a wide range of organisms and tissues and utilizes numerous substrates. This multifunctional kinase is considered to be involved in regulation of various cellular functions (Nairn et al., 1985; Stull et a f . , 1986). Some

334

SHOSHI MUTO

instances of enzyme regulation by Ca2+-and calmodulin-dependent protein kinase in plants will be described in Section VII,C.

3. Ca2+-and Lipid-Stimulated Protein Kinases Ca2+-and lipid-stimulated protein phosphorylation and protein kinases have been reported in several plants. Phosphorylation of soybean membrane protein by the membrane-associated protein kinase was stimulated by DG plus Ca2+(MorrC el al., 1984b). Phosphorylation of polypeptides in spinach chloroplast envelope was inhibited by EGTA, by treatment of the membrane with phospholipase C, or by extraction of the membrane with acetone. The acetone-treated membrane partially recovered its protein kinase activity by the addition of lipid fraction extracted from the membrane (Muto and Shimogawara, 1985). A Ca2+-and phospholipid-activated protein kinase was partially purified from zucchini hypocotyl (Schafer et al., 1985). This kinase had minimal activity in the absence of added Ca2+or phospholipid and was markedly stimulated by 1 pM Ca2+ and further enhanced by phosphatidylserine, phosphatidylethanolamine, and phosphatidic acid. No activation was observed by DG. Lipid-activated plant protein kinases, including kinases that do not require Ca2+,were recently reviewed by Harmon (1990).

4. Ca2+-DependentProtein Kinases Ca2+-dependentprotein kinase (CDPK), which is activated by the direct binding of Ca2+ and is independent of calmodulin or phospholipid, was reported in suspension cultured soybean cells (Harmon et al., 1987). The kinase was renatured after SDS-polyacrylamide gel electrophoresis and autophosphorylated and phosphorylated histone H 1 in a Ca2+-dependent manner. The concentration of Ca2+required for half-maximal (&) histone HI phosphorylation and autophosphorylation was ca. 2 p M . The mobility of kinase in SDS-polyacrylamide gel electrophoresis was changed in the presence of Ca2+.The kinase electroblotted onto nitrocellulose membrane bound 45Ca2+in the presence of NaCl and MgCI,. These results indicate that the enzyme itself is a Ca2+-bindingprotein. A CDPK was purified to homogeneity from soybean (Putnam-Evans et al., 1990). The enzyme is a monomeric protein with a molecular weight of 52,200. Histone type 111-S (lysine-rich histone H1) was the best substrate found for CDPK so far, and the phosphorylated amino acid was serine. The CDPK autophosphorylated both serine and threonine residues. Colocalization of CDPK with F-actin in onion and soybean root cells has been shown by immunocytochemical studies, suggesting a possible role in the

INTRACELLULAR Ca2 MESSENGER SYSTEM +

335

regulation of the cytoskeleton for this enzyme (Putnam-Evans et al., 1989). Recently, a cDNA clone encoding soybean CDPK was identified and sequenced (Harper et al., 1991). The predicted protein from cDNA had a molecular mass of 57,175 Da. This protein contained a catalytic domain similar to that of calmodulin-dependent protein kinases I1 from rat brain. Adjacent to the kinase domain is a calmodulin-like sequence with 39% amino acid identity to spinach calmodulin. The calmodulin-like region contains four putative EF-hand Ca2+-bindingmotifs. The presence of a calmodulin-like domain in CDPK may be responsible for inhibition of the enzyme by calmodulin inhibitors, which has been reported with the calmodulin-free enzyme (Harmon er al., 1987). The CDPKs, which are activated by IO-’M Ca2+,were partially purified and characterized from two species of green algae, D . salina (Guo and Roux, 1990) and D. tertiolecta (Yuasa and Muto, 1992). Both CDPKs were inhibited by calmodulin inhibitors but not regulated by calmodulin. The CDPK from D . tertiolecta bound to the microsomes when activated by Ca2+,like mammalian protein kinase C, and phosphorylated a number of polypeptides in the membranes isolated from the algal cells. Essentially no polypeptide in soluble fraction was phosphorylated. These results suggest a possibility that D . tertiolecta CDPK plays a role in regulation of membrane functions. Recently, CDPK was purified 80,000-fold from chromatin of etiolated pea nuclei (Li et al., 1991). Half-maximal activation occurred at 3 X lO-’M Ca2+.Histone type 111-S, ribosomal S6 protein, and casein were efficiently phosphorylated. In intact isolated pea nuclei, the enzyme preferentially phosphorylated several chromatin-associated proteins. Interestingly, the most phosphorylated protein had the same molecular weight as the nuclear protein substrate whose phosphorylation has been reported to be stimulated by phytochrome in a Ca2+-dependentmanner (Datta et al., 1985). Similar CDPKs have been reported in a number of plants including silver beet leaf (Klucis and Polya, 1987, 1988), oat leaf (Minichiello et al., 1989), and alfalfa (Bogre et al., 1988). In some instances, the Ca2+requirement for plant CDPKs can be overcome to varying degrees by certain fatty acids (Klucis and Polya, 1987, 1988; Minichiello et al., 1989).

B. Protein Kinase A

Protein kinase A plays central roles in signal transduction in mammalian cells. The mammalian CAMP-dependent protein kinase (protein kinase A)

336

SHOSHI MUTO

is a tetramer composed of two catalytic and two regulatory subunits (Rosen et a f . , 1977; Edelman et al., 1987). The regulatory subunit tightly binds cAMP and dissociates from the tetramer. Upon dissociation the catalytic subunits exert catalytic activity. Though protein kinase A has not been found in plants, a CAMP-stimulated protein kinase has been reported in Lemna (Kato et a f . , 1983, 1984). The enzyme bound cAMP but its molecular weight was not changed. Recently, Polya et a f . (1991) demonstrated that a partially purified protein kinase from petunia petals phosphorylated Kemptide, a synthetic peptide substrate for CAMPdependent protein kinase. This protein kinase is a basic protein with a molecular weight of 30,000 and is inhibited by the regulatory subunit of bovine CAMP-dependent protein kinase in a CAMP-reversible manner and by inhibitor proteins of mammalian CAMP-dependent protein kinase. The occurrence of cAMP has been reported in a variety of higher plant tissues; however, there is a large diversity among reported levels of cellular cAMP (Brown and Nelson, 1981). Recently, determination of CAMP levels in higher plants by radioimmunoassay has been carried out with great care. This is to ensure the elimination of interfering compounds and microbial origin of cAMP and the absence of artificial formation of cAMP from endogenous ATP during the assay, together with maximal recovery of endogenous cAMP (Spiteri et a f . , 1989). The cAMP level was below the detection limit of assay (0.5 pmollg fresh wt) in all tested plant tissues. This result is not in favor of CAMP-dependent regulation of protein phosphorylation in plants. In algae, cAMP likely functions as the intracellular signal (Goodenough, 1989; Kooijman et a f . , 1990). The CAMP-dependent protein kinases have been isolated from a marine diatom, Cylindrotheca fusiformis (Lin and Volcani, 1989). They are activated by micromolar concentrations of CAMP. Their molecular weight of 75,000-78,000 remained unchanged upon preincubation with CAMP, suggesting that the enzymes are monomeric and possess the CAMP-binding and catalytic domains on the same polypeptide. The CAMP-stimulated protein phosphorylation has been reported in zoospores of brown algae, Undaria pinnatifida and Laminaria angustata (Ohmori et a f . , 1991).

C. Substrate Proteins for Protein Kinases

Stimulus-induced increase of cytoplasmic Ca2+concentration may activate Ca2+-dependentprotein kinases and bring about phosphorylation of proteins in integral components of signal transduction. The phosphorylated

INTRACELLULAR Ca2+ MESSENGER SYSTEM

337

proteins may change their biochemical activities related to cellular response. Thus the identification of substrate proteins for the kinases is a key to elucidating the role of protein kinases in the cellular response. However, a few proteins have been identified as substrates for Ca2+dependent protein kinases in plants. Among the number of proteins phosphorylated via the Ca2+- and phospholipid-stimulated manner in the envelope membrane of spinach chloroplasts, the most prominent phosphorylation was observed on the small subunit of ribulose 1,S-bisphosphate carboxylase/oxygenase (Muto and Shimogawara, 1985). The phosphorylated amino acid residues were serine and threonine (S. Muto, unpublished data). However, its physiological significance was unclear. Calmodulin-dependent phosphorylation of carrot quinate:NAD+ oxidoreductase, which catalyzes reversible oxidation of quinate (Refeno et al., 1982; Ranjeva et al., 1983), and maize phosphoenolpyruvate carboxylase (Echevarria et al., 1988) has been reported. Quinate:NAD+ oxidoreductase is activated by phosphorylation and inactivated by dephosphorylation. The phosphorylated phosphoenolpyruvate carboxylase is both more active and less sensitive to feedback inhibition by malate (Nimmo et al., 1987). The microsomal H+-ATPase of corn root has been reported to be regulated by reversible Ca2+-and calmodulindependent phosphorylation of membrane proteins (Zocchi, 1985). The H+-ATPase activity was decreased with increasing phosphorylation of the membranes. Schaller and Sussman (1988) reported phosphorylation of the plasma membrane H+-ATPase of oat root by a Ca2+-dependent protein kinase, though the effect of phosphorylation on the enzyme activity was not demonstrated. Plasma membrane-bound Ca2+- and calmodulin-dependent protein kinase (Blowers and Trewavas, 1987) and Ca2+-dependentprotein kinase (Harmon et al., 1987) were autophosphorylated and regulated their own activities. A set of basic polypeptides that are encoded by ABA responsible gene RAB-17 (Vilardell et al., 1990) were rapidly synthesized during embryogenesis induced by ABA in maize (Sanchez-Martinez et al., 1986). These polypeptides have been found to be highly phosphorylated in uiuo (Vilardell et al., 1990). Recently, they were shown to be real substrates for casein kinase 2 isolated from rat liver cytosol and from maize embryos (Plano et al., 1991). Little is known about the role of Ca2+in the regulation of gene activation. Possible regulatory roles of Ca2+are transcriptional control of specific genes by Ca2+-dependent phosphorylation/dephosphorylationof trans-acting factors and control of translation by Ca2+-dependent phosphorylation/dephosphorylationof initiation factors.

338

SHOSHI MUTO

Specificity to artificial substrate for plant Ca2+-dependentprotein kinase has been reviewed by Polya and Chandra (1990).

VIII. Concluding Remarks

It has been revealed that plant plasma membranes have receptors for extracellular signals and G proteins, which may act on the Ca2+channel or phospholipase C to transduce signals percepted by the receptors to intracellular second messenger Ca2+or InsP,. However, many problems remain to be resolved. Various kinds of receptors for agonists such as hormones and nuerotransmitters are known in animals. For example, human epidermal growth factor receptor has protein tyrosine kinase activity that phosphorylates and activates phospholipase C-y in the plasma membrane (Ullrich and Schlessinger, 1990). Such a receptor or protein tyrosine kinase has not been reported in plants. Much effort is required for identification and characterization of receptors. The presence of heterotrimeric G proteins has been supported by various experimental evidences, though they have not been purified from plants and characterized. There is no direct indication that G proteins stimulate Ca2+channel or phospholipase C. Plasma membrane Ca2+channels have not yet been characterized. With respect to PI turnover in signal transduction, protein kinase C has not been found in plants and thus the role of DG as a signal molecule is unclear. InsP,-induced release of Ca2+from tonoplast vesicles or intact vacuoles indicates the presence of InsP, receptor on the vacuolar membrane, but the receptor protein has not been identified. In plant cells, Ca2+seems to be central in signal transduction, whereas the cAMP messenger system seems to be lacking. This is the most remarkable difference in signal transduction pathways in plants and animals. However, the cAMP messenger system in plants should not be totally negated because CAMP-stimulated protein kinases have been reported. In addition, cAMP obviously works as a messenger in algae. A possible involvement of cAMP in special processes of growth and development of higher plants may be considered. As well, the presence of other intracellular messenger systems has been suggested, e.g., pathogen infection- or elicitor-induced rapid changes in membrane potential and proton transport, H202and/or superoxide production, and lipid peroxidation (Dixon and Lamb, 1990; Ryan and Farmer, 1991);regulation of enzymes related to photosynthesis by the ferredoxin/thioredoxin redox system (Buchanann, 1980); and auxin-induced phospholipid breakdown by phospholipase A (Morre, 1990).

INTRACELLULAR Ca2 MESSENGER SYSTEM +

339

References Acharya, M. K., Dureja-Munjal, I., and Guha-Mukherjee, S. (1991). Phytochem. 30, 2895-2897. Aducci, P., Ballio, A., Blein, J.-P., Fullone, M. R., Rossignol, M., and Scalla, R. (1988). Proc. Natl. Acad. Sci. U.S.A. 85, 7849-7851. Alexandre, J., Lassalles, J. P., and Kado, R. T. (1990). Nature (London) 343, 567-570. Allan, E., Dawson, A., Drobak, B., and Roberts, K. (1989). Cellular Signalling 1, 23-29. Anderson, J. M., Charbonneau, H., Jones, H. P., McCann, R. 0.. andcormier, M. J. (1980). Biochemistry 19, 31 13-3120. Anderson, J. M., and Cormier, M. J. (1978). Biochem. Biophys. Res. Commun. 84,595-601. Andrejauskas, E., Hertel, R., and Marmt, D. (1985). J. Biol. Chem. 260, 5411-5414. Andrejauskas, E . , Hertel, R., and MarmC, D. (1986). Biochem. Biophys. Res. Commun. 138, 1269- 1275. Ayers, A. R., Ebel, J., Finelli, F., Berger, N., and Albersheim, P. (1976a). Plant Physiol. 57,751-759. Ayers, A. R., Ebel, J., Valent, B., and Albersheim, A. R. (l976b).Plant Physiol. 57,560-565. Balch, W. E. (1990). Trends Biochem. Sci. 15,473-477. Bean, B. P. (1989). Annu. Rev. Physiol. 51, 367-384. Berridge, M. J., and Irvine, R. F. (1989). Nature (London) 41, 197-205. Bhullar. R. P.. and Haslam, R. J. (1987). Biochem. J. 245, 617-620. Biermann, B., Johnson, E. M., and Feldman, L. J. (1990). Plant Physiol. 94, 1609-1615. Biffen, M., and Hanke, D. E. (1990). Plant Sci. 69, 147-155. Bilushi, S . V., Shebunin, A. G., and Babakov, A. V. (1991). FEES Lett. 291, 219-221. Birnbaumer, L., Abramowitz, J., and Brown, A. M. (1990). Biochim. Biophys. Acra 1013, 163-224. Blackford, S . , Rea, P. A., and Sanders, D. (1990). J . Biol. Chem. 265, 9617-9620. Blatt, M. R. (1988). J. Membrane Biol. 102, 235-246. Blatt, M. R., Thiel, G., and Trentham, D. R. (1990). Nature (London) 246, 766-769. Blowers, D. P., Hetherington, A., and Trewavas, A. (1985). Planta 166, 208-215. Blowers, D. P., andTrewavas, A. J. (1987). Biochem. Biophys. Res. Commun. 143,691-696. Blum, W., Hinsch, K.-D., Schultz, G., and Weiler, E. W. (1988). Biochem. Biophys. Res. Commun. 156, 954-959. Blumwald, E., and Poole, R. J. (1986). Plant Physiol. 80, 727-731. Bogre, L., Oliih, Z., and Dudits, D. (1988). Plant Sci. 58, 135-144. Bokoch, G. M., Parkos, C. A., andMumby, S. M. (1988). J. Biol.Chem. 263,16,744-16,749. Boss. W. F., and Massel, M. 0. (1985). Biochem. Biophys. Res. Commun. 132, 1018-1023. Braam, J., and Davis, R. W. (1990). Cell (Cambridge, Mass.) 60,357-380. Briars, S. A., Kessler, F . , and Evans, D. E. (1988). Planta 176, 283-285. Brosnan, J. M., and Sanders, D. (1990). FEES Lett. 260, 70-72. Brown, E. G., and Nelson, R. P. (1981). Phytochemistry 20, 2453-2463. Buchanann, B. B. (1980). Annu. Rev. Plant Physiol. 31, 341-374. Buckhout, T. J. (1983). Planta 159, 84-90. Buckhout, T.J. (1984). Plant Physiol. 76, 962-967. Burgoyne, R. D. (1989). Trends Biochem. Sci. 14, 394-396. Burr, R., Troxel, K. S. , and Crane, F. L. (1982). Biochem. Biophys. Res. Commun. 104, 1182-1188. Bush, D. R., Biswas, A. K., and Jones, R. L. (1989). Planta 178,411-420. Bush, D. R., and Sze, H. (1986). Plant Physiol. 80, 549-555. Bush, D. S., and Jones, R. L . (1990). Plant Physiol. 93, 841-845. Butcher, R. D., and Evans, D. E. (1987). Planfa 172, 265-272.

340

SHOSHI MUTO

Canut, H., Graziana, A., Boudet, A. M., and Ranjeva, R. (1989). FEES Lett. 253, 173-177. Castillo, F. J . , and Heath, R. L. (1990). Plant Physiol. 94, 788-795. Chanson, A. (1991). J. Plant Physiol. 137, 471-476. Chen, C. H., and Lehninger, A. L. (1973). Arch. Biochem. Biophys. 157, 183-186. Cheong, J.-J., and Hahn, M. G. (1991). Plant Gel( 3, 137-147. Cheung, W. Y. (1980). Science 207, 19-27. Churchill, K. A., and Sze, H. (1983). Plant Physiol. 71, 610-617. Cocucci, M. C. (1986). Plant Sci. 47, 21-27. Collinge, E., and Trewavas, A. J. (1989). J . Biol. Chem. 264, 8865-8872. Connet, R. J. A., and Hanke, D. E. (1987). Planta 170, 161-167. Conrath, U . , Jeblick, W., and Kauss, H. (1991). FEES Lett. 279, 141-144. Cosgrove, D. J., and Hedrich, R. (1991). Planta 186, 143-153. Cosio, E. G., Frey, T., and Ebel, J. (1990). FEES Lett. 264, 235-238. Cosio, E. G., Frey, T., Verduyn, R., van Boom, J., and Ebel, J. (1991). FEES Left. 271, 223-226. Cosio, E. G., Poepperl, H., Schmidt, W., and Ebel, J. (1988). Eur. J . Biochem. 175,309-315. Cott, G . G., Morse, M. J., Crain, R. C., and Satter, R. L. (1987). Plant Cell Rep. 6,352-355. Cote, G . G . , Quarmby, L. M., Satter, R. L., Morse, M. J., and Crain, R. C. (1990). In “Inositol Metabolism in Plants” (D. J. M o d and W. F. Boss, eds.), pp. 113-137. Wiley-Liss, New York. Datta, N., Chen, Y.-R., and Roux, S. J. (1985). Biochem. Biophys. Res. Commun. US, 1403- 1408. Day, D. A., and Wiskich, J. T. (1984). Physiol. V k g . 22, 241-261. DeBoer, A. H., Watson, B. A., and Cleland, R. E. (1989). Plant Physiol. 89,250-259. Dieter, M. (1984). Plant Cell Enuiron. 7, 371-380. Dieter, M . , and Marmt, D. (1980). Proc. Natl. Acad. Sci. U . S . A . 77, 7311-7314. Dieter, M., and Marmt, D. (1981). FEES Lett. 125, 245-248. Dieter, M., and Marm6, D. (1983). Planta 159, 277-281. Dieter, M., and Marmt, D. (1989). Planta 150, 1-8. Dillenschneider, M., Hetherington, A., Graziana, A., Alibert, G., Berta, P. Haiech, J., and Ranjeva, R. (1986). FEES Letr. 208,413-417. Dixon, R. A., and Lamb, C. J. (1990). Annu. Rev. Plant Physiol. 41, 339-367. Dolle, R., and Nultsch, W. (1988). Arch. Microbiol. 149, 451-458. Downward, J . (1990). Trends Biochem. Sci. 15, 469-472. Drprbak, B. K., Allan, E. F. Comerford,, J. G., Roberts, K., and Dawson, A. P. (1988). Biochem. Biophys. Res. Commun. 150, 899-903. Drprbak, B. K., and Ferguson, 1. B. (1985). Biochem. Biophys. Res. Commun. 130, 1241- 1246. Dr@bak,B. K., Ferguson, I. B., Dawson, A. P., and Irvine, R. F. (1988). Plant Physiol. 87, 2 17-222. Drprbak, B. K., Watkins, P. A. C., Chattaway, J. A., Roberts, K., and Dawson, A. P. (1991). Plant Physiol. 95, 412-419. DuPont, F. M., Bush, D. S., Windle, J. J.. and Jones, R. L . (1990). Plant Physiol. 94, 179- 188. Echevarria, C., Yidal, J., Le Marechal, P., Brulfert, J., Ranjeva, R., and Gadal, P. (1988). Biochem. Biophys. Res. Commun. 155, 835-840. Edelman, A. M., Blumenthal, D. K., and Krebs, E. G. (1987). Annu. Reu. Biochem. 56, 567-613. Einspahr, K. J . , Maeda, M., and Thompson, G. A., Jr. (1988a). J . CellBiol. 107, 529-538. Einspahr, K. J., Peeler, T. C., and Thompson, G. A., Jr. (1988b). J. Biol. Chem. 263, 5775-5779.

INTRACELLULAR Ca2+ MESSENGER SYSTEM

341

Einspahr, K. J., Peeler, T. C., andThompson, G. A.. Jr. (1989). Plant Physiol. 90, I 115-1 120. Elliott, D. C.. Founier, A., and Kokke, Y. S. (1988). Phytochemistry 27, 3725-3730. Elliott, D. C., and Kokke, Y. S. (1987a). Biochem. Biophys. Res. Commun. 145,1043-1047. Elliott, D. C., and Kokke, Y. S. (1987b). Phytochemistry 26, 2929-2935. Ettlinger, C., and Lehle, L. (1988). Nature (London) 331, 176-178. Fairley-Grenot, K., and Assmann, S. M. (1991). Plant Cell 3, 1037-1044. Felle, H. (1989). Plant Physiol. 91, 1239-1242. Ferris, C. D., Huganir, R. L., Supattapone, S., and Snyder, S. H. (1989). Nature (London) 342, 87-89. Feyerabend, M., and Weiler, E. W. (1989). Planta 178, 282-290. Fujisawa, H. (1990). BioEssays 12, 27-29. Furuichi, T., Yoshikawa, S., Miyawaki. A . , Wada, K., Maeda, N., and Mikoshiba, K. (1989). Nature (London) 342, 32-38. Giannini, J. L., Ruiz-Cristin, J., and Briskin, D. P. (1987). Planr Physiol. 85, 1137-1 142. Gilroy, S., Fricker, M. D., Read, N. D., and Trewavas, A. J. (1991). Planr Cell3,333-344. Gilroy, S . , Read, N. D., and Trewavas, A. J. (1990). Nature (London) 346,769-771. Ginzburg, M. (1987). Adu. Bot. Res. 14, 95-181. Goodenough. U. W. (1989). J . Cell Biol. 109, 247-252. Grabowski, L., Heim, S., and Wagner, K. G. (1991). Plant Sci. 75, 33-38. Graf, P., and Weiler, E. W. (1989). Physiol. Plant. 75, 469-478. Graf, P.. and Weiler, E. W. (1990). Plant Physiol. 94, 634-640. Graziana, A., Fosset, M., Ranjeva, R., Hetherington, A. M., and Lazdunski, M. (1988). Biochemistry 27, 764-768. Gross, J., and Marme, D. (1978). Proc. Natl. Acad. Sci. U . S . A . 75, 1232-1236. Grouzis, T.-P., Gibrat, R., Rigaud, J., and Grignon, C. (1987). Biochim. Biophys. Acta 903, 449-464. Gumber, S. C., Loewus, M. W., and Loewus, F. A. (1984). Plant Physiol. 76, 40-44. Guo, Y.-L., and Roux, S. J . (1990). Plant Physiol. 94, 143-150. Ha, K. S . , and Thompson, G. A., Jr. (1991). Plant Physiol. 97, 921-927. Hallcher, L. M., and Schermann, W. R. (1980). J. Biol. Chem. 255, 10.896-10,901. Harmon, A. C. (1990). In “Inositol Metabolism in Plants” (D. J. MorrC, W. F. Boss, and F. A. Loewus. eds.), pp. 319-334. Wiley-Liss, New York. Harmon, A. C., Putnam-Evans, C., and Cormier, M. J. (1987). Plant Physiol. 83, 830-837. Harper, J. F., Sussman, M. R., Schaller, G. E., Putman-Evans, C., Charbonneau, H . , and Harmon, A. C. (1991). Science 252, 951-954. Harvey, H. J., Venis. M. A., and Trewavas. A. J. (1989). Biochem. J . 257, 95-100. Hasunuma, K., and Funadera, K. (1987). Biochem. Biophys. Res. Commun. 143, 908-912. Hasunuma, K., Furukawa, K., Tomita, K., Mukai, C.. and Nakamura, T. (1987). Biochem. Biophys. Res. Commun. 148, 133-139. Hayama, T . , Shimmen, T., and Tazawa, M. (1979). Protoplasma 99, 305-321. Hedrich, R., and Kurkdjian, A. (1988). EMBO J. 7, 3661-3666. Hedrich, R., and Neher, E. (1987). Nature (London) 329, 833-836. Hedrich, R., and Schroeder, J . I. (1989). Annu. Rev. Plant Physiol. Plant Mol. Biol. 49, 539-569. Heim, S . , Bauleke, A,, Wylegalla, C., and Wagner. K. G. (1987). Plant Sci. 49, 159-165. Heim, S.. and Wagner, K. G. (1986). Biochem. Biophys. Res. Commun. 134, 1175-1181. Heim, S., and Wagner, K. G. (1989). Plant Sci. 63, 159-165. Heimann, K.. Kreimer. G., Melkonian. M., and Latzko, E. (1986). 2. Nuturforsch. C 42, 283-287. Heldt, H. W. (1976). In “Intact Chloroplast” ( J . Barber, ed.). pp. 215-234. ElsevierlNorthHolland Biomedical Press, Amsterdam.

342

SHOSHI MUTO

Hepler, P. K., and Wayne, R. 0. (1985). Annu. Rev. Plant Physiol. 36, 397-439. Hetherington, A. M., and Trewavas, A. J. (1984). Plant Sci. Lett. 35, 109-113. Hsieh, W.-L., Pierce, W. S., and Sze, H. (1991). Plant Physiol. 97, 1535-1544. Irvine, R. F., Letcher, A. J., Lander, D. J., Drebak, B. K., Dawson, A. P., and Musgrave, A. (1989). Plant Physiol. 89, 888-892. Jacobs, M., Thelen, M. P., Famdale, R. W., Astle, M. C., and Rubery, P. H. (1988).Biochem. Biophys. Res. Commun. 155, 1478-1486. Jarrett, H. W., Brown, C. J., Black, C. C., and Cormier, M. J. (1982). J . Biol. Chem. 257, 13,795- 13,804. Jena, P. K., Reddy, A. S. N., and Poovaiah, B. W. (1989). Proc. Natl. Acad. Sci. U.S.A. 86,3644-3648. Johannes, E., Brosnan, J. M., and Sanders, D. (1991). BioEssays 13, 331-336. Joseph, S. K., Esch, T., and Bonner, W. D., Jr. (1989). Biochem. J . 264, 851-856. Kamada, Y., and Muto, S. (1991). Biochim. Biophys. Acra 1093, 72-79. Kasai, M.,and Muto, S. (1990). J. Membrane Biol. 114, 133-142. Kasai, M., and Muto, S. (1991). Plant Physiol. 96,565-570. Kato, R., Uno, I., Ishikawa, T., and Fujii, T. (1983). Plant Cell Physiol. 24, 841-848. Kato, R., Uno, I., Ishikawa, T., and Fujii, T. (1984). Plant Cell Physiol. 25, 691-696. Kauss, H. (1987). Annu. Rev. Plant Physiol. 38, 47-72. Kikuyama, M., Oda, K., Shimmen, T., Hayama, T., and Tazawa, M. (1984). Plant Cell Physiol. 25, 965-974. Klucis, E., and Polya, G. M. (1987). Biochem. Biophys. Res. Commun. 147, 1041-1047. Klucis, E., and Polya, G. M. (1988). Plant Physiol. 88, 164-171. Kogel, G., Beissmann, B., Reisener, H. J., and Kogel, K. H. (1988). Physiol. Mol. Plant Pathol. 33, 173-185. Kogel, G., Beissmann, B., Reisener, H. J., and Kogel, K. (1991). PIanta 183, 164-169. Kooijman, R., d e Wild, P., van den Briel, W., Tan, S., Musgrave, A., and van den Ende, H. (1990). PIanta 181, 529-537. Korolkov., S. N., Garnovskaya, M. N. Basov, A. S. Chunaev, A. S., and Dumler, 1. L. (1990). FEBS Lett. 270, 132-134. Kreimer, G., Melkonian, M., and Latzko, E. (1985a). FEES Lett. 180, 253-258. Kreimer, G., Melkonian, M.,and Latzko, E. (1985b). Planta 166, 515-523. Kubowicz, B. D., Vanderhoef, L. N., and Hanson, J. B. (1982). Plant Physiol. 69,187-191. Kurosaki, F., Tsurusawa, Y., and Nishi, A. (1987). Planr Physiol. 85, 601-604. Larsson, C. (1983). In “Isolation of Membranes and Organelles from Plant Cells” ( J . L. Hall and A. L. Morr6, eds.), pp. 277-309. Academic Press, New York. Lawton, M. A., Yamamoto, R. T., Hanks, S. K., and Lamb, C. J. (1989). Proc. Natl. Acad. Sci. U . S . A . 86, 3140-3144. Lee, Y., and Assmann, S. M. (1991). Proc. Natl. Acad. Sci. U . S . A . 88, 2127-2131. Lew, R. R., Briskin, D. P., and Wyse, R. E. (1986). Plant Physiol. 82, 46-53. Li, H., Dauwalder, M., and Roux, S . J. (1991). PIant Physiol. 96, 720-727. Lin, P. P.-C., and Volcani, B. E. (1989). Biochemistry 28, 6624-6631. Lin, X., Feng,X.-H., and Watson, J. C. (1991). Proc. Natl. Acad. Sci. U.S.A.88,6951-6955. Ling, V., Perera, I., and Sielinski, R. E. (1991). Plant Physiol. 96, 1196-1201. Ling, V., Zielinski, R. E. (1989). Plant Physiol. 90, 714-719. Lukas, T. J., Iverson, D. B., Schleicher, M., and Watterson, D. M. (1984). Plant Physiol. 75,788-795. Lunevsky, V. Z., Zherelova, 0. M., Vostrikov, I. Y.,and Berestovsky, G . N. (1983). J . Membrane Biol. 72, 43-58. Ma, H . , Yanofsky, M. F., and Meyerowitz, E. M. (1990). Proc. Narl. Acad. Sci. U.S.A. 87, 3821 -3825.

INTRACELLULAR Ca2+ MESSENGER SYSTEM

343

Malatialy, L., Greppin, H., and Penel, C. (1988). FEBS Lett. 233, 196-200. Martins, I. S., Carnieri, E. G. S., and Vercesi, A. E. (1986). Biochim. Biophys. Acra 850, 49-56. Matsui, M., Sasamoto, S., Kunieda, T., Nomura, N., and Ishizaki, R. (1989). Gene 76, 313-3 19. Matsumura-Kadota, H., Muto, S., and Miyachi, S. (1982). Biochim. Biophys. Acra 679, 300-307. McAinsh, M. R., Brownlee, C., and Hetherington, A. M. (1990). Nature (London) 343, 186- 188. McMurray, W. C . , and Irvine, R. F. (1988). Biochem. J. 249, 877-881. Melin, P.-M., Sommarin, M., Sandelius, A. S., and Jergil, B. (1987). FEES Lett. 223,87-91. Memon, A. R., and Boss, W. F. (1990). J. Biol. Chem. 265, 14,817-14,821. Memon, A., Rincon, M., and Boss, W. F. (1989a). Plant Physiol. 91, 477-480. Memon, A. R., Chen, Q.. and Boss, W. F. (1989b). Biochem. Biophys. Res. Commun. 162, 1295- 1301. Meyer, C., and Weiler, E. W. (1989). Plant Physiol. 89,692-699. Millner, P. A. (1987). FEBS Lett. 226, 155-160. Minichiello, J., Polya, G. M., and Keane, P. J. (1989). Plant Sci. 65, 143-152. Morrk, D. J. (1990). In “Inositol Metabolism in Plants“ (D. J. Morrk, W. F. Boss, and F. A. Loewus, eds.), pp. 227-257. Wiley-Liss, New York. MorrC, D. J., Gripshover, B., Monroe, A., and Morrk, J. T. (1984a). J. Biol. Chem. 259, 15,364-15,368. MorrC, D. J., MorrC, J. T., and Varnold, R. L. (1984b). Plant Physiol. 75, 265-268. Morse, M. J., Crain, R. C., Cote, G. G., and Satter, R. L. (1989). Plant Physiol. 89,724-727. Morse, M. J., Crain, R. C., and Satter, R. L. (1987). Proc. Natl. Acad. Sci. U . S . A . 84, 7075-7078. Murthy, P. P. N., Renders, J. M., and Keranen, L. M. (1989). Plant Physiol. 91, 1266-1269. Muto, S. (1982). FEES Lett. 147, 161-164. Muto, S. (1983). Z. PJlanzenphysiol. 109, 383-393. Muto, S., Izawa, S., and Miyachi, S. (1982). FEES Lett. 139, 250-254. Muto, S., and Miyachi, S. (1977). Plant Physiol. 59, 55-60. Muto, S., and Miyachi, S. (1986). I n “Molecular and Cellular Aspects of Calcium in Plant Development” (A. J. Trewavas, ed.), pp. 197-1 14. Plenum, New York/London. Muto, S., Miyachi, S., Usuda, H., Edwards, G. E., andBassham, J. A. (1981). PlantPhysiol. 68, 324-328. Muto, S., and Shimogawara, K. (1985). FEBS Lett. 193, 88-92. Naim, A. C., Hemmings, H. C., Jr., and Greengard, P. (1985). Annu. Rev. Biochem. 54, 93 1-976. Napier, R. M., and Venis, M. A. (1991). Trends Biochem. Sci. 16, 72-75. Nimmo, G. A., McNaughton, G. A. L., Fewson, C. A., Wilkins, M. B., and Nimmo, H. G. (1987). FEES Letr. 213, 18-22. Nishizuka, Y.(1984). Nature (London) 308, 693-698. Oecking, C., and Weiler, E. W. (1991). Eur. J . Biochern. 199,685-689. Ogren, W. L., and Krogman, D. W. (1965). J . Biol. Chem. 240, 4603-4608. Oh-hama, T., and Miyachi, S. (1959). Biochim. Biophys. Acta 30, 202-210. Ohmori, M., Muto, S., and Miyachi, S . (1991). J. Phycol. 27, 540-543. OIBh, Z., and Kiss, Z. (1986). FEES Lett. 195, 33-37. Paliyath, G., and Thompson, J. E. (1988). Plant Physiol. 88, 295-302. Park, M.-H., and Chae, Q. (1990). Biochem. Biophys. Res. Commun. 169, 1185-1 190. F‘fathann, H., Hartmann, E., Brightman, A. O., and MorrC, D. J. (1987). Plant Physiol. 85, 1151-1155.

344

SHOSHI MUTO

Plana, M., Itarte, E., Eritja. R., Goday, A., Pagts, M., and Martinez, M. C. (1991). J . Biol. Chem. 266, 22,510-22,514. Polya, G. M., and Chandra,. (1990). Curr. Top. Plant Biochem. Phvsiol. 9, 164-180. Polya, G. M., Chung, R., and Menting, J . (1991). Plant Sci. 79, 37-45. Polya, G. M., and Davies, J. R. (1982). FEBS Lett. 150, 167-171. Polya, G. M., Davies. J. R., and Micucci. V. (1983). Biochim.Biophys. Acta 761, 1-12. Polya, G. M., and Micucci, V. (1984). Biochim. Biophys. Acta 785, 68-74. Poovaiah, B. W., and Reddy, A. S. N. (1987). CRC Crit. Rev. Plant Sci. 6, 47-103. Putman-Evans, C. L., Harmon, A. C., and Cormier. M. J. (1990). Biochemistry 29, 2488-2495. Putman-Evans, C., Harmon, A. C., Palevitz, B. A., Fechheimer. M.. and Cormier, M. J. (1989). Cell Motil. Cvtoskel. 12, 12-22. Ranjeva, R., Carrasco, A., and Boudet, A. M. (1988). FEBS Lett. 239, 137-141. Ranjeva, R., Refeno, G., Boudet. A. M.. and Marme, D. (1983). Proc. Nail. Acad. Sci. U.S.A. 80, 5222-5224. Rasi-Caldogno, F., De Michelis, M. I.. and Pugliarello. M. C. (1982a). Biochim. Biophys. Acta 693, 287-295. Rasi-Caldogno, F., De Michelis, M. I.. and Pugliarello. M. C. (1982b). In “Plasmalemma and Tonoplast: Their Functions in the Plant Cell” (D.Marmt. E. Marrt. and R. Hertel, eds.), pp. 361-367. Elsevier Biomedical Press, Amsterdam. Rasi-Caldogno, F.. Pugliarello, M. C., and De Michelis, M. 1. (1987). Plant Physiol. 83, 994- 1000. Rasi-Caldogno, F., Pugliarello, M. C.. Olivari, C.. and De Michelis, M. I. (1989). Planr Physiol. 90, 1429- 1434. Rea, P. A., and Sanders, D. (1987). Physiol. Plant. 71, 131-141. Reddy, A. S. N., and Poovaiah, B. W. (1987). J. Biochem. (Tokyo) 101, 569-573. Refeno, G., Ranjeva, R.. and Boudet, A. M. (1982). Plunta 154, 193-198. Rincon, M., and Boss, W. F. (1987). Plant Physiol. 83, 395-398. Roberts, D. M., Lukas, T. I.. and Watterson. D. M. (1986). CRC Crit. Rev. Plant Sci. 4, 31 1-339. Robinson, C., Larsson, C., and Buckhout, T. J. (1988). Physiol. Plant. 72, 177-184. Romero, L. C., Sommer, D., Gotor, C., and Song, P.-S. (1991). FEBS Lett. 282, 341-346. Rosen, 0. M., Rangel-Caldogno, R., and Erlichman, J. (1977). Citrr. Top. Cell Regul. 12, 39-74. Ryan, C. A., and Farmer, E. E. (1991). Annu. Rev. Plant Physiol. Mol. Biol. 42, 651-674. Salimath, B. P.. and Marmt, D. (1983). Planta 158, 560-568. SBnchez-Martinez, D.,Puigdomtnech, P., and Pages, M. (1986). Plant Physiol. 82,543-549. Sandelius, A. S., and Morrt, D. J. (1987). Plant Physiol. 84, 1022-1027. Sandelius, A. S., and Sommarin, M. (1986). FEBS Lett. 201, 282-286. Sano, H., and Youssefian, S. (1991). Mol. Gen. Genet. 228, 227-232. Sasaki, Y., Sekiguchi, K., Nagano, Y., and Matsuno, R. (1991). FEBS Left. 293, 124-126. Schafer, A., Bygrave, F., Matzenauer. S., and Marme, D. (1985). FEES Lett. 187, 25-28. Schaller, G. E., and Sussman, M. R. (1988). Plunta 173, 509-518. Scheel, D., and Parker, J. E. (1990). Z. Naturforsch. C 45, 569-575. Schloss, J. A. (1990). Mol. Gen. Genet. 221, 443-452. Schmidt, W. E., and Ebel, J. (1987). Proc. Nail. Acad. Sci. U.S.A. 84, 4117-4121. Schroeder, J. I., and Hagiwara, S. (1990). Proc. Nail. Acad. Sci. U . S . A . 87, 9305-9309. Schumaker, K. S., and Sze, H. (1985). Plant Physiol. 79, 1 I 11-1 117. Schumaker, K. S., and Sze, H. (1986). J. B i d . Chem. 261, 12,172-12,178. B. i d . Chem. 262, 3944-3946. Schumaker, K. S., and Sze, H. (1987). .I Schumaker, K. S., and Sze, H. (1990). Plant Physiol. 92, 340-345.

INTRACELLULAR CaZ+ MESSENGER SYSTEM

345

Sexton. J. C., and Moore, T. S., Jr. Plant Physiol. 68, 18-22. Sharp, J. K., McNeil, M., and Albersheim, P. (1984a). J . Biol. Chem. 259, 11,321-11,336. Sharp, J. K., Valent, B., and Albersheim, P. (1984b). J . Biol. Chem. 259, 11,312-11,320. Shears, S . B. (1989). Biochem. J. 260, 313-324. Shiina, T., and Tazawa, M. (1987). J. Membrane Biol. 96, 263-276. Simon, P., Bonzon, M., Greppin, H., and Marmk, D. (1984). FEBS Lett. 167, 332-338. Simon, P., Dieter, P., Bonzon, M., Greppin, H., and Marme, D. (1982). Plant Cell Rep. 1, 119-122. Sommarin, M.,and Sandelius, A. S. (1988). Biochim. Biophys. Acra 958, 268-278. Spiteri, A., Viratelle, 0. M., Raymond, P., Rancillac, M., Labouesse, J., and Pradet, A. (1989). Plant Physiol. 91, 624-628. Stosic, V., Penel, C., Marme, D., and Greppin, H. (1983). Plant Physiol. 72, 1136-1138. Strasser, H., Hoffmann, C., Grisebach, H., Matern, U. (1986). Z. Naturforsch. C 41, 7 17-724. Stroobant, P., and Scarborough, G. A. (1979). Proc. Natl. Acad. Sci. U.S.A. 76,3102-3106. Stull, J . T., Nunnally, M. H., and Michinoff, C. H. (1986). Enzymes 17, 114-159. Supattapone, S., Worley, P. F., Baraban, J. M., and Snyder, S. H. (1988). J . Biol. Chem. 263, 1530-1534. Takagi, S., and Nagai, R. (1985). Plant Cell Physiol. 26, 941-951. Takagi, S., and Nagai, R. (1988). Plant Physiol. 88, 228-232. Takagi, S . , Yamamoto, K. T., Furuya, M., and Nagai, R. (1990). Plant Physiol. 94, 1702- 1708. Tate, B. F., Schaller, G. E., Sussman, M. R., and Crain, R. C. (1989). Plant Physiol. 91, 1275- 1279. Tester, M . (1990). New Phytol. 114, 305-340. Theil, G . , MacRobbie, E. A. C., and Hanke, D. E. (1990). EMBO J . 9, 1737-1741. Thuleau, P., Graziana, A., Canut, H., and Ranjeva, R. (1990). Proc. Natl. Acad. Sci. U . S . A . 87, 10,000-10,004. Thuleau, P., Graziana, A., Ranjeva, R., and Schroeder, J. 1. (1991). “Program and Abstracts, Third Int. Congress Plant Mol. Biol., October, 1991, Tuscon AZ.” Thuleau, P., Graziana, A., Rossignol, M., Kauss, H., Auriol, P., and Ranjeva, R. (1988). Proc. Narl. Acad. Sci. U . S . A . 85, 5932-5935. Toda, H., Yazawa, M., Sakiyama, F., and Yagi, K. (1985). J. Biochem. (Tokyo)98,833-842. Tsutsui, I. (1990). I n “The role of Calcium in Biological Systems” (L. J. Anghileri, ed.), Vol. V, pp. 97-1 18. CRC Press, Boca Raton. FL. Ullrich, A., and Schlessinger, J. (1990). Cell (Cambridge, Mass.) 61, 203-212. Vilardell, J., Goday, A., Freire, M. A., Torrent, M., Martinez, M. C., TornC, J. M., and Pages, M. (1990). Plant Mol. Biol. 14, 423-432. Warpeha, K. M. F., Hamm, H. E., Rasenick, M. R., and Kaufman, L. S. (1991). Proc. Natl. Acad. Sci. U . S . A . 88, 8925-8929. Williams, L., Schueler, S. B., and Briskin, D. P. (1990). Plant Physiol. 92, 747-754. Yuasa, T., and Muto, S. (1992). Arch. Biochem. Biophys. 296, 175-182. Zabulionis, R. B., Atkinson, B. G., Procunier, J. D., and Walden, D. B. (1988). Genome 30, 820-824. Zbell, B., Paulik, M., and Morre, D. L. (1990). Protoplasma 154, 74-79. Zbell, B., Schwendemann, I., and Bopp, M. (1989). J . Plant Physiol. 134, 639-641. Zbell, B., and Walter-Back, C. (1988). J. Plant Physiol. 133, 353-360. Zhang, P., Yabe, I., and Muto, S. (1992). Protoplasma, in press. Zherelova, 0. M. (1989). FEBS L e f t . 249, 205-207. Zocchi, G. (1985). Plant Sci. 40, 153-159. Zocchi. G. (1988). Plant Sci. 54, 103-107. Zocchi, G., and Hanson, J. B. (1983). Plant Cell Enuiron. 6, 203-209.

This Page Intentionally Left Blank

Index A

in striated muscle cardiac muscle, 107-109, I11, 118 sarcomere, 94-95, 99 thick-filament proteins, 65 thin-filament proteins, 83-84, 88-91 transdifferentiation in medusae and, 227,

A-band-associated proteins in striated muscle antibody use, 101 cardiac muscle, 106 sarcomere, 93-94, 96 thick-filament proteins, 62-80, 82 Abscisic acid, intracellular calcium messenger system and, 315, 320, 327,

232

a-Actin, proteins in striated muscle and, 83-84, 95-97, 102

a-Actinin, proteins in striated muscle and, I29

337

Acellular afibrillar cementum (AAC), regenerative cementogenesis and, 2-3 Acellular extrinsic fiber cementum (AEFC), regenerative cementogenesis and established root surfaces in uitro, 31, 33-34, 42

cardiac muscle, 108-109, I II , 114-1 15, 117-118

sarcolemma, 125-126 skeletal muscle, 87-88, 91, 101 P-Actinin, proteins in striated muscle and, 9 1-92

Actinomycin D, transdifferentiation in medusae and, 227, 229 Adenylate cyclase, transdifferentiation in medusae and, 245 Adherens-junction-specific cell adhesion molecule (A-CAM), proteins in striated muscle and, 126 Adhesion plaques, vertebrate cell culture technology and, 187 ADP-ribosylation, intracellular calcium messenger system and, 309-31 I Age, regenerative cementogenesis and, 17 Agitation, vertebrate cell culture technology and, 196-197 Airlift reactors, vertebrate cell culture technology and, 169-170 Algae, intracellular calcium messenger system and, 335-336, 338 Alkaline phosphatase, regenerative cementogenesis and, 15, 40-41 A1kalinization intracellular calcium messenger system and, 307

established root surfaces in uiuo, 46, 49 function of cementum, 2-5 growing root surfaces, 5-17, 19, 23, 25. 27-28

Acellular intrinsic fiber cementum (AIFC), regenerative cementogenesis and established root surfaces in uitro, 41-42

function of cementum, 2-5 growing root surfaces, 15-25 Acer pseudoplatanus, intracellular calcium messenger system and, 309, 324 Acetone, intracellular calcium messenger system and, 334 Acetylcholine, proteins in striated muscle and, 99 Acid aerosol, symplast and, 286 Acid exposure, regenerative cementogenesis and, 51 Acid growth, symplast and, 273-275 Actin regenerative cementogenesis and, 40

347

348 symplast and, 275 transdifferentiation in medusae and, 245 Alternaria, intracellular calcium messenger system and, 308 Alveolar bone, regenerative cementogenesis and, I established root surfaces in uitro, 33, 36, 40-4 I established root surfaces in uiuo,44, 46-47, 49-50 function of cementum, 3-5 Arnaranthus tricolor, intracellular calcium messenger system and, 333 Amino acids intracellular calcium messenger system and, 309-31 I , 328, 333 proteins in striated muscle and, 91 Ammonia, vertebrate cell culture technology and, 164-165, 173, 195 Anchorage-dependent cultures, technology and bioreactors, 166, 172-173, 186-189 traditional cultures, 147-151 Ankylosis, regenerative cementogenesis and, 44-47, 50 Anoxia, symplast and, 271-273, 275-276, 286-287 Anterior latissimus dorsi (ALD), proteins in antibody use, 100-101 thick-filament proteins, 68, 70, 77-78 thin-filament proteins, 90-92 Antibiotics, vertebrate cell culture technology and, 148 Antibodies, see also Monoclonal antibodies proteins in striated muscle and, 61, 127-129 cardiac muscle, 103-1 1 I , 113-1 19 detection of abnormalities, 100-102 myosin, 62-79 sarcolemma, 124-125, 127 sarcomere, 93-98 sarcoplasmic reticulum, 120- 124 thick-filament proteins, 79-83 thin-filament proteins, 83-88, 90-92 transdifferentiation in medusae and, 224. 227, 250 vertebrate cell culture technology and, 147, 184 Antigens proteins in striated muscle and cardiac muscle, 103, 111, 113

INDEX

skeletal muscle, 63, 68, 84, 87, 90 vertebrate cell culture technology and, 197 Aortic-pulmonary septation, proteins in striated muscle and, 113 Aphidicolin, transdifferentiation in medusae and, 252-253, 255-256 Apoplast, symplast and, 264, 276, 285, 300 electrophysiological structure, 268, 27 1-272 integration of activity, 285-287, 291-299 Arabidopsis, intracellular calcium messenger system and, 328-329 Arabidopsis thaliana. intracellular calcium messenger system and, 309-31 1 Ascorbic acid, regenerative cementogenesis and, 38 ATP intracellular calcium messenger system and calmodulin, 329, 33 I concentration, 312, 314-317 phosphatidylinositol turnover, 324, 326-327 protein kinases. 337 proteins in striated muscle and, 64-65,81, 120, 122 symplast and, 274 ATPase intracellular calcium messenger system and, 328, 337 concentration, 313-314, 316 receptors, 307-308 proteins in striated muscle and sarcoplasmic reticulum, 120-123 skeletal muscle, 63, 65-68, 81, 102 Atrio-ventricular (AV) node, proteins in striated muscle and, 103, 107 Autophosphorylation, intracellular calcium messenger system and, 334, 337 Auxin intracellular calcium messenger system and, 306-307, 325-326, 338 symplast and integration of activity, 286-287, 294-298 Lockhart equations, 277-279, 283-285 mechanical properties, 279-282 proton pumps, 274-275 water relations, 282-283 transdifferentiation in medusae and, 216

349

INDEX

Avena intracellular calcium messenger system and, 308-310 symplast and, 273 Axial organs, symplast and, 277-279, 287, 297. 300

B

Baby hamster kidney cells, vertebrate cell culture technology and, 150-151, 170, 173 Bacteria regenerative cementogenesis and, 2, 44 vertebrate cell culture technology and, 146, 153, 166, 193 Bergenholtz’s system, vertebrate cell culture technology and, 160, 162 Bioreactors, vertebrate cell culture technology and, 204 commercial scale, 167-187 optimization, 195- 197, 199-201 packed-bed reactors, 187-192 production system, 162-167 Bone, regenerative cementogenesis and, 1-2,4 established root surfaces in uitro, 28-29, 33-34, 36, 40-42 established root surfaces in uiuo, 44, 46, 49-50 growing root surfaces, 17, 19, 23 Brassicu olerucea. intracellular calcium messenger system and, 328 Brevin, proteins in striated muscle and. I10

C C protein, in striated muscle, 62, 79-83, 94 Calcification, regenerative cementogenesis and, 35, 47 Calcium proteins in striated muscle and, 127 sarcoplasmic reticulum, 119-124 skeletal muscle, 64,81, 85, 94, 102 transdifferentiation in medusae and, 247 Calcium-dependent protein kinases, intracellular calcium messenger system and, 332-335

Calcium messenger system, intracellular, in plants, 305-306, 338 calmodulin. 328-332 concentration. 31 1-312 channels, 3 18-321 chloroplasts. 3 I8 endoplasmic retictilum, 3 15-3 16 mitochondria. 317-318 plasma membrane, 312-315 vacuoles. 316-317 G proteins, 308-31 1 phosphatidylinositol turnover, 32 1-328 protein kinases, 332-337 receptors, 306-308 Caldesmon, proteins in striated muscle and, I09 Calmodulin intracellular calcium messenger system and. 305. 328-332 concentration, 313-315 protein kinases. 333-334. 337 proteins in striated muscle and, 109 Calreticulin, proteins in striated muscle and, I24 Calsequestrin, proteins in striated muscle and, 121-122 Cancer therapy, vertebrate cell culture technology and, 203 CapZ, proteins in striated muscleand, 91-92 Carbohydrate. transdifferentiation in medusae and, 249-251 Carbon dioxide, vertebrate cell culture technology and, 164, 167, 169, 189, 198, 200 Cardiac muscle, proteins in antibody use. 101, 115-1 19 sarcolemma. 125-126 sarcomere, 98-99 sarcoplasmic reticulum, 122, 124 structural proteins, 110-1 15 thick-filament proteins, 79, 83, 102-107 thin-filament proteins, 84-88, 91-92, 107-1 10 Carrot, intracellular calcium messenger system and, 308, 315, 320,323,328,337 Cntharurifhus roseus, intracellular calcium messenger system and, 322, 325 CCCP. intracellular calcium messenger system and, 312-315 cDNA intracellular calcium messenger system and, 319, 328-329, 333, 335

350 proteins in striated muscle and, 66-67,91, 123, 126 Cell cycle, transdifferentiation in medusae and, 254-257 Cellular intrinsic fiber cementum (CIFC), regenerative cementogenesis and established root surfaces in uitro, 35, 4 1-42 function of cementum, 2-5 growing root surfaces, 15-25, 27-28 Cellular, mixed stratified cementum (CMSC), regenerative cementogenesis and, 2-4, 17.42 Cementoblasts, regenerative cementogenesis and established root surfaces in uitro, 35, 40-42 growing root surfaces, 13, 15, 19, 21-23, 2.5 Cementocytes, regenerative cementogenesis and, 3-4, 21-22 Cementogenesis, regenerative, 1-2, 51-52 established root surfaces in uitro, 28 collagenous matrices, 3 1-37 culture material, 28-31 factors, 37-43 established root surfaces in uiuo, 43 biological problems, 49-5 1 clinical conditions, 43-44 guided regeneration, 46-49 spontaneous regeneration, 44-46 function of cementum, 2-5 growing root surfaces, 5 acellular extrinsic fiber cementum (AEFC), 5-17 cellular/acellular intrinsic fiber cementum, 17-25 dentine, 25-28 Ceramic reactors, vertebrate cell culture technology and, 190-192 Chemotaxis, vertebrate cell culture technology and, 154 Chinese hamster ovary cells, vertebrate cell culture technology and, 147, 150, 176, 186, 189 Chlamydomonas reinhardtii, intracellular calcium messenger system and, 309-310 Chlorella, intracellular calcium messenger system and, 329 Chloroplasts, intracellular calcium messenger system and, 308, 318, 334

INDEX

calmodulin, 329-332 G proteins, 309-310 Chlortetracycline, intracellular calcium messenger system and, 315 Chromatin, intracellular calcium messenger system and, 335 Chromatography, intracellular calcium messenger system and, 314, 321-322 Chromosomes transdifferentiation in medusae and, 252 vertebrate cell culture technology and, 148 Citric acid, regenerative cementogenesis and, SI Clones intracellular calcium messenger system and, 319, 328-329, 333, 335 regenerative cementogenesis and, 38, 40-4 1 transdifferentiation in medusae and, 214 Cnidaria, transdifferentiation in medusae and, 218 Coleoptile intracellular calcium messenger system and, 314, 318, 320, 324 symplast and, 273-274, 281, 300 Collagen regenerative cementogenesis and established root surfaces in uitro, 28-29, 31-38, 40-42 established root surfaces in uiuo, 44-47, 49, 51 function of cementum, 2-4 growing root surfaces, 9, 11-13, 15, 19, 21-23, 25, 27 transdifferentiation in medusae and, 222, 236 vertebrate cell culture technology and, 159-160, 178, 188 Collagenase regenerative cementogenesis and, 38 transdifferentiation in medusae and DNA replication, 252, 254-255 initiation, 237, 244 isolation of tissues, 222-223 potential, 224, 232-234 Commelina commmunis, intracellular calcium messenger system and, 309, 313-314, 327 Concanavalin A, transdifferentiation in medusae and, 222, 251

INDEX

Connective tissue regenerative cementogenesis and, 13, 29, 37,44-47 vertebrate cell culture technology and, 159, 162 Corn, intracellular calcium messenger system and, 313-314, 318, 324 Cortex, symplast and, 291 COS cells, proteins in striated muscle and, 126 Costameres, proteins in striated muscle and, 114 Cowpea, symplast and, 265-266, 270, 280-28 I , 298 Creatine kinase in striated muscle, 81, 117 Cucurbira pepo, intracellular calcium messenger system and, 309 Culture technology, vertebrate cell, see Vertebrate cell culture technology Cyclic AMP intracellular calcium messenger system and, 305, 335, 338 proteins in striated muscle and, 123 regenerative cementogenesis and, 40 transdifferentiation in medusae and, 245 Cylindrotheca jitsiformis, intracellular calcium messenger system and, 336 Cysteine proteins in striated muscle and, 65 transdifferentiation in medusae and, 236 Cytokeratin, proteins in striated muscle and, I12 Cytokinin, intracellular calcium messenger system and, 325 Cytoplasm intracellular calcium messenger system and, 308, 31 1 calmodulin, 330-332 concentration, 31 1-312, 316, 318 phosphatidylinositol turnover, 321,323, 325-326 protein kinases, 332, 336 proteins in striated muscle and cardiac muscle, 105 sarcolemma, 124 sarcoplasmic reticulum, 121 skeletal muscle, 66, 85, 87, 92, 102 regenerative cementogenesis and established root surfaces in uitro, 39-40 growing root surfaces, 9, 12-13, 19, 21-22, 27

351 symplast and, 275 transdifferentiation in medusae and, 213-214, 245, 256-257 potential, 229-230 Cytoskeleton intracellular calcium messenger system and, 335 proteins in striated muscle and cardiac muscle, 109 sarcolemma, 125 skeletal muscle, 66, 88, 93, 98 transdifferentiation in medusae and, 213, 236, 249 vertebrate cell culture technology and, 160

0 Decalcification, regenerative cementogenesis and, 32, 36, 42, 46 Demineralization, regenerative cementogenesis and, 32-33, 35, 39, 42, 51 Dentine, regenerative cementogenesis and, 4-5, 51 established root surfaces in uirro, 32-33, 38-39, 41-42 established root surfaces in uiuo, 45-47, 49 growingroot surfaces, 5,7,9,19,23,25-28 Dentoclasts, regenerative cementogenesis and, 41 Dephosphorylation, intracellular calcium messenger system and, 324, 337 Depolarization intracellular calcium messenger system and, 320, 325, 327 symplast and, 269, 272, 275-276 Desmin in striated muscle, 62 cardiac muscle, 112-1 14, I18 skeletal muscle, 87, 90, 96-99, 101-102 Desmosomes proteins in striated muscle and, 112, 126 regenerative cementogenesis and, 9 Destabilization, transdifferentiation in medusae and, 224, 236 DG, intracellular calcium messenger system and, 338 phosphatidylinositol turnover, 321-322, 326-328 protein kinases, 332, 334

352 Diacylglycerol, transdifferentiation in medusae and, 247-249, 257 2,4-Dichlorophenoxyaceticacid (2.4-D), intracellular calcium messenger system and, 325-326 Dihydrocytochalasin B, transdifferentiation in medusae and, 232, 249 Dihydropyridine, proteins in striated muscle and. 127 I ,4-Dihydropyridine, intracellular calcium messenger system and, 318, 320 Diploid cells, vertebrate cell culture technology and, 149, 151, 174, 176 Disease, regenerative cementogenesis and, 43 DNA intracellular calcium messenger system and, 310-311 proteins in striated muscle and. 96 regenerative cementogenesis and, 37 transdifferentiation in medusae and. 214 initiation, 237, 247-248, 250 isolation of tissues, 222 potential, 223-224. 227, 229, 232-235 vertebrate cell culture technology and. 150, 169, 176, 194 DNA polymerase, transdifferentiation in medusae and, 252 DNA replication, transdifferentiation in medusae and, 220, 235, 257 initiation, 236-246, 249-250 Drosophiln

proteins in striated muscle and, 88, 96 transdifferentiation in medusae and, 215, 256 Drugs intracellular calcium messenger system and, 315 transdifferentiation in medusae and, 235. 249-250, 252-253 vertebrate cell culture technology and, 203 Duchenne muscular dystrophy, proteins in striated muscle and, 100, 124-126 Dunnliella, intracellular calcium messenger system and, 309, 322-323, 326, 335 Dystrophin, proteins in striated muscle and, 88, 100, 124-126

INDEX E

EDL muscle, proteins in. 70. 74-75, 77, 81, 88 Electric potential, symplast and. 265-269, 271-273 Electrogenesis intracellular calcium messenger system and. 316, 318 symplast and, 299 electrophysiological structure, 267-273 integration of activity. 286, 298 proton pumps, 275-276 Electron microscopy proteins in striated muscle and, 128-129 antibody use, 101 cardiac muscle, 107, I12 sarcomere, 97 thick-filament proteins. 62. 64,67, 78. 80, 82 thin-filament proteins, 84, 88, 91 regenerative cementogenesis and, 13, 32 symplast and, 270 transdifferentiation in medusae and, 222 Electron transport, intracellular calcium messenger system and, 329, 331-332 Electrophoresis, intracellular calcium messenger system and, 310-311 Electrophysiological structure, symplast and. 265-274. 297 Elongation regenerative cementogenesis and, 11-12 symplast and, 265, 299-300 auxin, 277-280, 283-285 electrophysiological structure, 265-273 integration of activity, 286-287, 294, 296-299 proton pumps, 273-277 Encapsulation, vertebrate cell culture technology and, 177-178, 197 Endocytosis regenerative cementogenesis and, 40 transdifferentiation in medusae and, 232 Endoderm, transdifferentiation in medusae and, 256 isolation of tissues, 222-223 potential, 232-236 tissue organization. 218. 220 Endoplasmic reticulum intracellular calcium messenger system and. 315-316, 318-319, 325-326

INDEX

proteins in striated muscle and, I24 Enzymes intracellular calcium messenger system and, 305. 314. 338 calmodulin, 329-330, 332 G proteins, 308-309 phosphatidylinositol turnover, 321-324, 328 protein kinases, 332-336 proteins in striated muscle and, 66-68, 120-121 regenerative cementogenesis and. 15. 22, 37 transdifferentiation in medusae and, 236-238, 244-245, 247, 249-250, 257 Epidermal growth factor, regenerative cementogenesis and, 40 Epithelium regenerative cementogenesis and, 2 established root surfaces in uiuo, 44-45, 47, 50 growing root surfaces, 9, 12, 19, 27 symplast and, 288-289, 291, 297 transdifferentiation in medusae and, 217-218, 220, 224, 256 vertebrate cell culture technology and, 160, 162 Epitopes, proteins in striated muscle and, 61, 128 cardiac muscle, 1 1 1, 113 myosin, 63, 65-67, 73 sarcolemma, 124-126 sarcomere, 93-96 sarcoplasmic reticulum, 121 thick-filament proteins, 79-83 thin-filament proteins, 88, 90 Erythrocytes intracellular calcium messenger system and, 333 symplast and, 289 Escherichia coli, intracellular calcium messenger system and, 31 I Euchromatin, regenerative cementogenesis and, 9, 2 1, 40 Extracellular matrix proteins in striated muscle and, 93, 100 regenerative cementogenesis and, 32 transdifferentiation in medusae and, 257 isolation of tissues, 223

353 potential, 232-236 tissue organization. 220-222 vertebrate cell culture technology and, 159 Extraocular (EO) muscles, proteins in, 66, 68-69

F

F-actin, transdifferentiation in medusae and, 226-227 Fascia adherens (FA), proteins in striated muscle and, 114, 126 Fast-twitch glycolytic fibers (FG), proteins in striated muscle and, 120, 127 Fast-twitch oxidative-glycolytic (FOG) fibers, proteins in striated muscle and, 120, 127-128 Fermentation, vertebrate cell culture technology and bioreactors, 166, 169-170, 179 optimization, 193, 196, 198, 201 Fiber-bed reactors, vertebrate cell culture technology and, 185-187 Fiber typing, proteins in striated muscle and, 67-69, 71 Fibrils, regenerative cementogenesis and, 5 I established root surfaces in uirro, 31-33, 35-37 established root surfaces in uiuo, 46-47, 51 growing root surfaces, 9, 11-13. 15, 19, 2 1-23, 27-28 Fibroblasts proteins in striated muscle and, 97, I13 regenerative cementogenesis and. 4 established root surfaces in uifro,28-29, 31-33, 35, 37-42 established root surfaces in uiuo, 45-46, 50 growing root surfaces, 7, 9, 11-13, 22. 27 transdifferentiation in medusae and, 218 vertebrate cell culture technology and, 151 bioreactors. 174, 176, 189 three-dimensional cultures, 156, 158-160 Fibronectin proteins in striated muscle and. 99

354

INDEX

regenerative cementogenesis and established root surfaces in uifro,32-33, 36, 39, 41 growing root surfaces, 13, 25 transdifferentiation in medusae and, 222 vertebrate cell culture technology and, 188 Filamin in striated muscle, 90-91, 108-109 Flagella, transdifferentiation in medusae and, 252 initiation, 238, 244 potential, 223-224, 227, 229, 232 Fluidized-bed reactors, vertebrate cell culture technology and, 178-180 Fluorescein, proteins in striated muscle and, 65, 117 Fluorescence intracellular calcium messenger system and, 315, 327 proteins in striated muscle and cardiac muscle, 110, 114 sarcolemma, 124 skeletal muscle, 63, 80, 87, 95, 99 transdifferentiation in medusae and, 226 Fluorescence microscopy, proteins in striated muscle and, 62 Fluorocarbon, transdifferentiation in medusae and, 244-245 FMRF-amide, transdifferentiation in medusae and, 224, 230 Foot-and-mouth disease, vertebrate cell culture technology and, 146, 151, 153, I76 Foot protein in striated muscle, 122-123 Funaria hygrometrica, intracellular calcium messenger system and, 309 Fusicoccin, intracellular calcium messenger system and, 308

G G proteins, intracellular calcium messenger system and, 308-31 1, 338 Galactose, vertebrate cell culture technology and, 195 GDP, intracellular calcium messenger system and, 308-31 1 Gelsolin, proteins in striated muscle and, 109-1 10

Gene expression transdifferentiation in medusae and, 213, 215, 237, 249, 256 vertebrate cell culture technology and, 173 Genetic engineering, vertebrate cell culture technology and, 146, 178, 191, 194 Genotype, vertebrate cell culture technology and, 192 Gibberellic acid, intracellular calcium messenger system and, 315, 326 Gingiva, regenerative cementogenesis and established root surfaces in uifro, 29, 31-32, 37-39, 41-42 established root surfaces in uiuo, 44-47, 50 growing root surfaces, I I , 17, 22 Glass-bead reactors, vertebrate cell culture technology and, 189 Glucose. vertebrate cell culture technology and, 164-165, 195, 200, 204 Glutamate, vertebrate cell culture technology and, 204 Glutamine. vertebrate cell culture technology and, 165, 195, 204 Glycogen, regenerative cementogenesis and, 40 Glycolysis proteins in striated muscle and, 67,76,88, 120, 127-128 vertebrate cell culture technology and, 204 GI ycoprotein intracellular calcium messenger system and, 307, 320 proteins in striated muscle and, 99, 121-122, 124, 126 regenerative cementogenesis and, 13 transdifferentiation in medusae and, 213, 236 Glycosaminoglycans, regenerative cementogenesis and, 38 Golgi, regenerative cementogenesis and, 9, 11, 19, 21 Conionemus, transdifferentiation in medusae and, 249 GTP, intracellular calcium messenger system and, 308-312, 323, 326 GTPase, intracellular calcium messenger system and, 309 Guided regeneration, cementogenesis and, 46-49

INDEX

355 H

H protein in striated muscle, 82 Heavy meromyosin (HMM) in striated muscle, 63-64 HeLa cells, vertebrate cell culture technology and, 148, 166, 195-196 Hematoxylin, vertebrate cell culture technology and, 172 Hepatocytes, transdifferentiation in medusae and, 217 High-performance liquid chromatography, intracellular calcium messenger system and, 314. 322 Histones, intracellular calcium messenger system and, 334 Hollow-fiber reactors, vertebrate cell culture technology and, 180-183, 191 HomoIogy intracellular calcium messenger system and, 3 11, 333 proteins in striated muscle and, 121-122, 125 Hordeum uulgare, intracellular calcium messenger system and, 315 Hormones, vertebrate cell culture technology and, 147, 201-202 Hybridization proteins in striated muscle and, 66,76,215 vertebrate cell culture technology and, 147 Hybridoma cells, vertebrate cell culture technology and, 146-147, 153, 194 bioreactors, 177-178, 189 Hydra, transdifferentiation in medusae and, 216 Hydrated collagen lattices, vertebrate cell culture technology and, 159-160 Hydraulic conductivity, symplast and auxin, 278-279, 283 integration of activity, 287-288, 293, 297 Hydrogen intracellular calcium messenger system and, 337 concentration, 312-3 18 phosphatidylinositol turnover, 324, 328 receptors, 307-308 symplast and, 267, 274-276 vertebrate cell culture technology and, 153, 171, 173

Hydromedusae, transdifferentiation in DNA replication, 252-256 initiation, 236-251 isolation of tissues, 222-223 potential, 223-236 tissue organization, 218-222 Hydrostatic pressure, symplast and, 263, 293, 297, 300 Hyperpolarization, symplast and, 274-276. 296 Hypertrophy, proteins in striated muscle and, 118 Hypocotyl intracellular calcium messenger system and, 322, 324, 326, 328, 334 symplast and, 265 auxin, 280-281, 283 electrophysiological structure, 266-271, 273 integration of activity, 286-287, 291-292, 294-295,298-299 proton pumps, 274-276

I-band associated proteins in striated muscle, 122 cardiac muscle, 109, I18 skeletal muscle, 84-85, 93-94, 101 lmmunocytochernical localization of proteins in striated muscle, see Proteins in striated muscle Immunoelectron microscopy, proteins in striated muscle and, 61 sarcolemma, 125 sarcoplasmic reticulum, 122 skeletal muscle, 73, 82, 88, 93-94, 97 Immunofluorescence, proteins in striated muscle and, 61 antibody use, 101 cardiac muscle, 107, 110-1 13, 118-1 19 sarcolemma, 127 sarcomere, 94, 96-97, 99 sarcoplasmic reticulum, 120, 122, 124 thick-filament proteins, 73, 82-83 thin-filament proteins, 87-88, 90-92 Immunoglobulin proteinsinstriatedmuscleand.81, 110,117 vertebrate cell culture technology and, 166

356 Immunology, vertebrate cell culture technology and, 147, 149 Immunoreactivity, transdifferentiation in medusae and, 224, 229-230 in situ hybridization, proteins in striated muscle and, 66 Indole-3-acetic acid, symplast and, 274-277, 280, 282 Infarct damage, proteins in striated muscle and, 116 Inflammation, regenerative cementogenesis and, 2, 44 Inhibitors intracellular calcium messenger system and, 308 calmodulin, 330-331 concentration, 313, 315-317, 319-320 G proteins, 309, 3 I I phosphatidylinositol turnover, 322-324, 327 protein kinases, 333, 335-337 proteins in striated muscle and sarcoplasmic reticulum, 121, 123 skeletal muscle, 64-65, 74, 84, 100 regenerative cementogenesis and, 38, 40 symplast and, 270, 276 auxin, 28 1, 284-285 integration of activity, 286, 296, 299 transdifferentiation in medusae and, 235, 257 DNA replication, 252-253 initiation, 241, 243-245. 247, 249, 251 vertebrate cell culture technology and, 150 bioreactors, 164-165, 175, 182-183 optimization, 194-195 Inositol phosphates, intracellular calcium messenger system and, 309, 321-324 Inositol phospholipids, intracellular calcium messenger system and, 321-323, 328 Inositol 1,4,5-trisphosphate (Imp,), intracellular calcium messenger system and, 31 I , 338 concentration, 3 18-320 turnover, 321-328 Interferon, vertebrate cell culture technology and, 147, 151, 169, 174, 176, 186 Intermediate filaments, proteins in striated muscle and, 96-98, 113-1 14 Intracellular calcium messenger system, see Calcium messenger system, intracellular, in plants

INDEX

Iodine, proteins in striated muscle and, 116-117, 119 Ischemia, proteins in striated muscle and, 115, 117 Isomyosins, proteins in striated muscle and, 101. 103-104 Isozymes, proteins in striated muscle and, 66-67, 70, 75, 77

J

Jellyfish, see Transdifferentiation in medusae

L Lactate, vertebrate cell culture technology and, 195, 200, 204 Lactic acid symplast and, 275 vertebrate cell culture technology and, 164-165, 173, 195 Lamin, proteins in striated muscle and, 122 Laminaria angustata, intracellular calcium messenger system and, 336 Laminin, transdifferentiation in medusae and, 222 Lectins, transdifferentiation in medusae and, 250-25 I Lemnu, intracellular calcium messenger system and, 309, 336 Lens, transdifferentiation in medusae and, 2 17-218 Lepidium satiuum, intracellular calcium messenger system and, 315 Ligands, transdifferentiation in medusae and, 220, 249-250 Light intracellular calcium messenger system and. 305, 318, 327, 329-331, 333 symplast and, 270 Light chains (LCs), proteins in striated muscle and cardiac muscle, 106 myosin, 64-65, 69, 71, 76-77, 79 Light meromyosin (LMM) in striated muscle, 63 Light microscopy proteins in striated muscle and, 66, 129 regenerative cementogenesis and, 32

INDEX

357

Lilliurn, intracellular calcium messenger system and, 324 Limonium, symplast and, 292 Lipids intracellular calcium messenger system and, 334. 338 symplast and, 289 Liposomes, intracellular calcium messenger system and, 310, 314, 317 Lockhart equations, symplast and, 277-285 Lymphoblastoids, vertebrate cell culture technology and, 166, 170 Lymphocytes, vertebrate cell culture technology and, 147, 150 Lysosomes, transdifferentiation in medusae and, 232

M M-band proteins in striated muscle, 70 M protein in striated muscle, 80-81 Magnesium intracellular calcium messenger system and, 312-316, 322 proteins in striated muscle and, 121, 127 Maize, intracellular calcium messenger system and. 308. 320, 333, 337 Manganese, intracellular calcium messenger system and, 316, 322 Mannose, regenerative cementogenesis and, 13, 25 Medusae, transdifferentiation in, see Transdifferentiation in medusae Melanoma cells, vertebrate cell culture technology and, 175, 189 Membrane-based reactors, vertebrate cell culture technology and, 183-185 Membrane potential intracellular calcium messenger system and, 316, 327 symplast and, 269-271 Mesogloea, transdifferentiation in medusae and, 220-223, 232, 250 Messenger RNA intracellular calcium messenger system and, 310. 328 proteins in striated muscle and, 66, 107 Mezerein, transdifferentiation in medusae and, 247-248, 257

Microcarriers, vertebrate cell culture technology and bioreactors, 166, 171-177, 180 optimization, 196-1 99 Microencapsulated-based reactors, vertebrate cell culture technology and, 177- I78 M icrofilamen t s regenerative cementogenesis and, 40 transdifferentiation in medusae and, 232 Microsomes, intracellular calcium messenger system and, 307, 330 concentration, 312, 319 G proteins, 309, 3 I 1 phosphatidylinositol turnover, 322-324, 326 Microtubules, proteins in striated muscle and, 118 Mineralization, regenerative cementogenesis and, 4,45, 51 established root surfaces in uitro, 28-29, 42 growing root surfaces, 11-12, IS, 17. 21-22, 25, 27-28 Mitochondria intracellular calcium messenger system and, 317-318, 330 proteins in striated muscle and, 67, 107, I20 Mitomyocin C, transdifferentiation in medusae and, 235, 252 Mitosis, transdifferentiation in medusae and, 216, 252 MM-creatine kinase, in striated muscle, 81, I07 Monoclonal antibodies proteins in striated muscle and, 61, 128 antibody use, 100-101 cardiac muscle, 103-105, 110-1 11, 113, 116-1 17, I I9 myosin, 64-71, 73-74, 76, 78 sarcolemma, 125- I26 sarcomere, 93-97, 99 sarcoplasmic reticulum, 120-124 thin-filament proteins, 84-86, 91-92 transdifferentiation in medusae and. 217, 223, 227, 239, 249, 253 vertebrate cell culture technology and, 146-147, 203 bioreactors, 167, 177, 180, 188 traditional cultures, 148- 149

358 Mougeotia, intracellular calcium messenger system and, 308 mRNA, see Messenger RNA Muscle, see Cardiac muscle; Skeletal muscle; Striated muscle Muscle creatine kinase, in striated muscle, 81 Muscle dystrophies, proteins in striated muscle and, 100-101 Mutation, proteins in striated muscle and, 1 I8 Myoblasts, proteins in striated muscle and myosin, 71, 75 sarcolemma. 125 sarcomere, 95, 97-98 sarcoplasmic reticulum, 122 thin-filament proteins, 84, 87, 91 Myocarditis, proteins in striated muscle and, 115-116, 119 Myocardium, proteins in, 103-105, 108, 114, 116-117 Myocytes, proteins in striated muscle and, 106, 109, 111, 114 Myofibers proteins in, 103-106 transdifferentiation in medusae and, 220, 224, 254, 256 Myofibrils proteins in antibody use, 101 cardiac muscle, 105, 107-115, 118-1 19 myosin, 65-67, 73, 78 sarcomere, 95-98 sarcoplasmic reticulum, 123-124 thick-filament proteins, 79, 82-83 thin-filament proteins, 84-85, 87-88, 91-92 Myofilaments, transdifferentiation in medusae and, 224, 226, 229-230, 252 Myomesin in striated muscle, 80-81, 107 Myosin in striated muscle, 62-79 antibody use, 100-102 cardiac muscle, 103-106, 115-1 17, 119 sarcomere, 97 transdifferentiation in medusae and, 227 Myosin heavy chains (MHC) in striated muscle, 64,66-71, 73-75, 78, 83, 128 antibody use, 102 cardiac muscle, 103-106, 117-1 19

INDEX

sarcomere, 95-96, 99 sarcoplasmic reticulum, 120 thin-filament proteins, 91 Myotubes, proteins in striated muscle and, 107, 125-128 antibody use, 100 sarcomere, 95, 98-99 thick-filament proteins, 71, 74, 76-78, 80-8 I thin-filament proteins, 87-88

N

NAD, intracellular calcium messenger system and, 329-331, 337 NAD kinase, intracellular calcium messenger system and, 329-332 NADP, intracellular calcium messenger system and, 329-332 Nebulin, in striated muscle, 62, 88-90, 101 Nematoblasts, transdifferentiation in medusae and, 230, 253 Neoblasts, transdifferentiation in medusae and, 217 Neural crest cells, transdifferentiation in medusae and, 218 Neuronal cells, transdifferentiation in medusae and, 218 Nicotiana tabarum, intracellular calcium messenger system and, 322 Nitella intracellular calcium messenger system and. 325 symplast and, 280-281 Nitrogen, vertebrate cell culture technology and, 153, 171, 200 Nucleotides intracellular calcium messenger system and. 309, 323, 328-329 proteins in striated muscle and, 121

0 Odontoblasts, regenerative cementogenesis and, 1 I , 23 Oligonucleotides, intracellular calcium messenger system and, 333 Oncogenes, intracellular calcium messenger system and, 3 I 1

INDEX

359

Onoclea, intracellular calcium messenger system and, 308 Ontogeny, transdifferentiation in medusae and, 213, 230, 256-257 Osmosis, symplast and, 263 auxin, 278-281 integration of activity. 287-290, 292, 297-299 Osmotic shock, intracellular calcium messenger system and, 326 Osteoblasts. regenerative cementogenesis and, 11, 25, 35, 40 Osteocytes, regenerative cementogenesis and, 23-25 Oxygen symplast and, 264 vertebrate cell culture technology and bioreactors, 164.171,173,175,177-179, 184 optimization, 197-201 packed-bed reactors, 189, 191 traditional cultures, 149, 153 Oxygen consumption rate, vertebrate cell culture technology and, 191 Oxygenation, vertebrate cell culture technology and, 166, 184, 186, 189

P

Packed-bed reactors, vertebrate cell culture technology and, 187-192 Paranemin in striated muscle, 98-99 Parenchyma cells, symplast and, 268, 270, 279, 291, 296, 300 Parvalbumin, proteins in striated muscle and, 127-128 Patagialis muscle, proteins in, 101 Pathogens, intracellular calcium messenger system and, 307, 338 Pea intracellular calcium messenger system and, 309-310, 315, 329-330, 333 symplast and, 281, 284, 286, 294 Peptides intracellular calcium messenger system and, 309 proteins in striated muscle and, 83-84,89, 121, 125 Perfusion symplast and, 275-276, 280

vertebrate cell culture technology and, 188, 190 Perfusion-based reactors, vertebrate cell culture technology and, 167-169, 175- 176 Periodontal ligament, regenerative cementogenesis and established root surfaces in uirro, 28-29, 3 I , 36-37, 39-42 established root surfaces in uiuo, 44-46, 49-50 growing root surfaces, 1 I , 15, 17, 22 Periodontal regeneration, 1-2, 42-43 Periodontitis, regenerative cementogenesis and, 37, 45, 47 PH intracellular calcium messenger system and, 313-317 symplast and, 268, 273, 275-276 transdifferentiation in medusae and, 237 Phagocytosis, regenerative cementogenesis and, 37 Phalloidin. proteins in striated muscle and, 83, I l l Pharbitis, symplast and, 273 Phaseolus, symplast and, 266, 268 Phenotype proteins in striated muscle and, 106 regenerative cementogenesis and, 35, 37-38, 40-41 transdifferentiation in medusae and, 215-2 I6 vertebrate cell culture technology and, 192 Phenylalkylamines. intracellular calcium messenger system and, 320 Phorbol esters intracellular calcium messenger system and, 332 transdifferentiation in medusae and, 247 Phosphatidylinositol 4.5-bisphosphate (PIP?),intracellular calcium messenger system and, 321-322, 325-327 Phosphatidylinositol 4-phosphate (PIP), intracellular calcium messenger system and, 321, 323, 327 Phosphatidylinositol 4-phosphate (PIP) kinases, intracellular calcium messenger system and, 32 1-323, 328 Phosphatidylinositol turnover, intracellular calcium messenger system and, 321-328, 338

360 Phospholamban, proteins in striated muscle and, 123-124 Phospholipase A, intracellular calcium messenger system and, 326, 338 Phospholipase C, intracellular calcium messenger system and, 334, 338 phosphatidylinositol turnover, 32 1-323, 326, 328 receptors, 3 1 1 Phospholipase D, intracellular calcium messenger system and, 326 Phospholipids intracellular calcium messenger system and, 317, 338 phosphatidylinositol turnover, 322, 326 protein kinases, 332-334 symplast and, 288 transdifferentiation in medusae and, 247 Phosphorylation intracellular calcium messenger system and, 322, 329, 332-337 proteins in striated muscle and, 85,98, 121, 123- 124 Photoreceptors, intracellular calcium messenger system and, 308 Photosynthesis intracellular calcium messenger system and, 332 syrnplast and, 264 Phycomyces, symplast and, 280 Phytoalexin, intracellular calcium messenger system and, 307, 328 Phytochromes, intracellular calcium messenger system and, 308-309, 327 Phytophtora megasperma, intracellular calcium messenger system and, 307 Phytotoxins, intracellular calcium messenger system and, 307 Pisum satiuum, intracellular calcium messenger system and, 309 Planarian, transdifferentiation in medusae and, 217-218 Plants growth in symplast, see Symplast, plant growth in intracellular calcium messenger system in, see Calcium messenger system, intracellular, in plants transdifferentiation in medusae and, 215-216 Plasma membrane, intracellular calcium messenger system and, 338

INDEX

concentration, 312-315, 318, 320 G proteins, 309-31 1 phosphatidylinositol turnover, 321-322, 325, 327-328 protein kinases, 332, 337 receptors, 307-308 Plasmalemma, symplast and, 278, 299-300 integration of activity, 289, 291, 293-294, 296 proton pumps. 274-275 Plasmodesmata, symplast and, 270, 292 Podocoryne, transdifferentiation in medusae and, 227, 230, 236, 249 Podocoryne carnea, transdifferentiation in medusae and, 216, 222-224, 252 Polymerase chain reaction, intracellular calcium messenger system and, 310 Polymerization intracellular calcium messenger system and, 307 regenerative cementogenesis and, 22 vertebrate cell culture technology and, 180-181, 198 Polyorchis, transdifferentiation in medusae and, 222-223 Polypeptides intracellular calcium messenger system and, 308, 310, 336-337 proteins in striated muscle and, 87, 127 Potassium intracellular calcium messenger system and, 307, 311, 320-321, 327 symplast and, 276-277 Potato intracellular calcium messenger system and, 328 symplast and, 282 Precementoblasts, regenerative cementogenesis and, 13, 27, 41 Predentine, regenerative cementogenesis and, 9, 11-13, 19, 27 Preosteoblastic cells, regenerative cementogenesis and, 25.40-41 Procollagens, regenerative cementogenesis and, I I , 13 Pronase, transdifferentiation in medusae and, 237-238, 241, 244 Propagators, vertebrate cell culture technology and, 170-171 Prostaglandins, regenerative cementogenesis and, 38

INDEX

Protein intracellular calcium messenger system and, 305 calmodulin, 328-330 concentration, 317-3 19 G proteins, 308-31 I protein kinases, 334-337 receptors, 306-308 regenerative cementogenesis and. 23, 37-38. 40-41 transdifferentiation in medusae and, 214, 227, 250 vertebrate cell culture technology and, 146-147, 179, 184, 197-198 Protein in striated muscle, 61-62, 127-129 cardiac muscle, 102 antibody use, 115-1 19 structural proteins, 110-1 15 thick-filament proteins, 103-107 thin-filament proteins, 107-1 10 sarcolemma, 119-120, 124-127 sarcoplasmic reticulum, 119-124 skeletal muscle, 62 antibody use, 100-102 C protein, 79-80 F protein. 82-83 H protein, 82-83 86-kDa protein, 81-82, 94 M protein, 80-81 MM-creatine kinase, 81 myomesin, 80-81 myosin, 62-79 ribosomes, 83 sarcomere, 93-100 skelemins, 83 thin-filament proteins, 83-93 X protein, 82-83 Protein kinase intracellular calcium messenger system and, 332-327. 338 proteins in striated muscle and, 123 transdifferentiation in medusae and, 247-249 Protein kinase A intracellular calcium messenger system and, 333, 335-336 transdifferentiation in medusae and, 249 Protein kinase C intracellular calcium messenger system and, 321, 327, 332-333, 335 transdifferentiation in medusae and, 247, 249. 257

361 Proteolysis intracellular calcium messenger system and, 307 transdifferentiation in medusae and, 236, 239, 245 vertebrate cell culture technology and, 149 Protofilament, proteins in striated muscle and, 96, 98 Proton-motor force, symplast and, 291, 296 Proton pumps. symplast and, 271, 273-277, 285, 299-300 integration of activity, 286-287, 291-292, 299 Protooncogenes intracellular calcium messenger system and, 310 vertebrate cell culture technology and, 160 Protoplasts, intracellular calcium messenger system and, 308, 332 calmodulin, 329, 332 concentration, 315, 320 phosphatidylinositol turnover, 325, 327 protein kinases, 332 Pircciniu grarninisi, intracellular calcium messenger system and, 307

R Radial potential, symplast and, 272-273 rus, intracellular calcium messenger system and, 31 I Reacidification, symplast and, 275 Recombination, vertebrate cell culture technology and, 169, 176, 194, 196 Regeneration, transdifferentiation in medusae and, 217 Regenerative cementogenesis, see Cementogenesis, regenerative Reinfection, cementogenesis and, 47 Replication, DNA, see DNA replication Resorption, regenerative cementogenesis and, 44,46-47, 49-51 Respiration intracellular calcium messenger system and, 318 symplast and auxin, 285 electrophysiological structure, 270-272 integration of activity, 285-296, 299 proton pumps, 274, 276

362

INDEX

Retinal pigment epithelium (RPE), transdifferentiation in medusae and, 217-218 RF-amide, transdifferentiation in medusae and DNA replication, 255 initiation, 238-239, 243, 247-248 potential, 224, 229-230, 235 Rhabdomyosarcoma, proteins in striated muscle and, 101-102 Rhynia, symplast and, 264 Ribosomes, transdifferentiation in medusae and, 229 RNA intracellular calcium messenger system and, 333 transdifferentiation in medusae and, 237 RNA polymerase, proteins in striated muscle and, 107 Root cementum, regenerative cementogenesis and, see Cementogenesis, regenerative Root sheath, regenerative cementogenesis and, 9, 13 Root surfaces, regenerative cementogenesis and, 12 biological problems, 49-5 1 clinical conditions, 43-44 collagen, 31-37 culture material, 28-3 1 factors, 37-43 guided regeneration, 46-49 spontaneous regeneration, 44-46 Rough endoplasmic reticulum, regenerative cementogenesis and, 9, 11, 19, 21 Ryanodine receptor, proteins in striated muscle and, 122-123

S Sail-sheet cultures, vertebrate cell culture technology and, 160-161 Samanea saman, intracellular calcium messenger system and, 327 Sarcolemma, proteins in striated muscle and, 119-120, 124-127 cardiac muscle, 107-1 10, 112, 114-1 15 skeletal muscle, 66, 71, 87, 92, 99-101 Sarcomere, proteins in striated muscle and, 93-100

cardiac muscle, 106, 110-115, 118 thick-filament proteins, 65, 79, 82-83 thin-filament proteins, 85, 88 Sarcoplasmic reticulum, proteins in striated muscle and, 81, 92, 102, 107, 119-124 Sarsia, transdifferentiation in medusae and, 222 Satellite cells, proteins in striated muscle and, 71, 73, 79, 97 SDS-polyacrylamide gel electrophoresis, intracellular calcium messenger system and, 310-311 Second messengers, intracellular calcium messenger system and, 305-306, 31 I , 321, 328, 338 Sequences intracellular calcium messenger system and, 306 calmodulin, 328-329 G proteins, 309-3 I 1 protein kinases, 333, 335 proteins in striated muscle and, 75.91.96, I23 Sharpey’s fibers, regenerative cementogenesis and, 3-4, 17 Shear sensitivity, vertebrate cell culture technology and, 166-167, 178, 196 Signal transduction intracellular calcium messenger system and, 305-306, 338 G proteins, 308, 31 1 phosphatidylinositol turnover, 326 protein kinases, 335-336 transdifferentiation in medusae and, 249, 257 Silica-fiber cultures, vertebrate cell culture technology and, 156, 158-159 Simian virus-transformed rat fibroblasts, vertebrate cell culture technology and, 159- I60 Single-photon emission-computed tomography (SPECT), proteins in striated muscle and, 115-1 16 Skelemins in striated muscle, 83 Skeletal muscle, proteins in antibody use, 100-102 cardiac muscle, 102-103, 108-1 1I sarcolemma, 124-125 sarcomere, 93-100 sarcoplasmic reticulum, 120, 122, 124

INDEX

thick-filament proteins, 62-83 thin-filament proteins, 83-93 Slow-twitch oxidative fibers, proteins in striated muscle and, 120 Smooth muscle proteins in, 94, 98, 124 transdifferentiation in medusae and, 257 DNA replication, 252-255 potential, 224-230, 235 Soleus muscle, proteins in myosin, 68. 70, 74-79 thick-filament proteins, 80-82, 87-88 Soybean, intracellular calcium messenger system and G proteins, 310 phosphatidylinositol turnover, 322, 325-326. 328 protein kinases, 334 receptors, 307 Spectrin in striated muscle, 92-93, 101-102, I10 Spider web cultures, vertebrate cell culture technology and, 153-155 Spinach, intracellular calcium messenger system and, 328, 330, 332, 334-335 Sponge transdifferentiation in medusae and, 216 vertebrate cell culture technology and, 154, 156-157 Spontaneous regeneration, cementogenesis and, 44-46, 49 Stem cells regenerative cementogenesis and, 38, 40 transdifferentiation in medusae and, 229 Stenosis, proteins in striated muscle and, 118 Stereotropism, vertebrate cell culture technology and, 154 Stomotoca, transdifferentiation in medusae and, 222-223. 236 Streptomyces qriseus, transdifferentiation in medusae and, 237 Stress-fiber-like structures, proteins in, 95, 105, 108 Striated muscle protein in, see Protein in striated muscle transdifferentiation in medusae and, 252, 256 initiation, 238-241, 245, 247, 249-251 isolation of tissues, 222-223 potential, 223-231, 233-235, 238 tissue organization, 218, 220

363 Subumbrella, transdifferentiation in medusae and, 218, 220, 222-223 Surface-attached growth, vertebrate cell culture technology and. 151-152 Surface potential, symplast and, 267. 269 Surface pump, symplast and, 273-275, 299 Sycamore, intracellular calcium messenger system and, 323 Symplast, plant growth in, 263-265,299-300 auxin Lockhart equations, 277-279, 283-285 mechanical properties, 279-282 water relations, 282-283 electrophysiological structure, 265 cowpea hypocotyl, 267-270 elongation growth, 265-267 two-pump hypothesis, 270-274 integration of activity apoplast canal, 291-294 auxin-enhanced uptake, 294-297 dynamic regulation, 297-299 models, 287-291 respiration, 285-287 proton pumps surface pump, 273-275 xylem pump, 275-277 Synemin in striated muscle, 98-99

T

Talin in striated muscle, 92-93, 99-100, 114- 1 I5 Teeth, regenerative cementogenesis and, see Cementogenesis, regenerative Temperature intracellular calcium messenger system and, 305 vertebrate cell culture technology and, 164, 174-175, 191, 196, 200 Tetrodotoxin, proteins in striated muscle and, 87, 98 Thick-filament proteins in striated muscle cardiac muscle, 103-107 skeletal muscle, 62-83 Thin-filament proteins in striated muscle, 83-93, 107-1 10 cardiac muscle, 107-1 10 skeletal muscle, 83-93 Thymidine, transdifferentiation in medusae and, 252

364

INDEX

Thyroid hormones, proteins in striated muscle and, 105-106 Thyroxine, proteins in striated muscle and, 105-106

Tissue specificity proteins in striated muscle and, 66, 106 transdifferentiation in medusae and, 236 Titin in striated muscle, 62 cardiac muscle, 109-1 12 skeletal muscle, 91, 93-96, 99, 101-102 Tobacco, intracellular calcium messenger system and, 319-320, 322-324 Tomato, symplast and, 297 Tonoplasts, intracellular calcium messenger system and, 313, 315-317, 324-325 Tooth, regenerative cementogenesis and, see Cementogenesis, regenerative Toxicity, vertebrate cell culture technology and, 174, 198 Toxins intracellular calcium messenger system and, 309-31 1 vertebrate cell culture technology and, 191 TPA, transdifferentiation in medusae and, 247-249, 257 Transcription intracellular calcium messenger system and, 328, 337 transdifferentiation in medusae and, 227, 229 Transdifferentiation in medusae, 2 13-214, 256-257 animals, 216-218 hydromedusae DNA replication, 252-256 initiation, 236-25 I isolation of tissues, 222-223 potential, 223-236 tissue organization, 218-222 plants, 215-216 terminology, 214-215 Translation proteins in striated muscle and, 107 transdifferentiation in medusae and, 229 Translocation, intracellular calcium messenger system and, 317, 332 Transplant rejection, proteins in striated muscle and, 115-1 16 Transverse tubules, proteins in striated muscle and, 119-122, 126-127 Tropomysoin, proteins in striated muscle and, 84-85, 108, 111, 119

Troponin in striated muscle, 62 cardiac muscle, 108, 1 I I , 117-1 18 skeletal muscle, 84-87. 101 Trypsin intracellular calcium messenger system and, 330, 332 vertebrate cell culture technology and, 149, 172 Tumors proteins in striated muscle and, 101-102 transdifferentiation in medusae and, 247 vertebrate cell culture technology and, 147- I48 Turgor pressure, symplast and, 278

U

Umbrella, transdifferentiation in medusae and. 232-233 Undaria pinnatifida, intracellular calcium messenger system and, 336

v Vaccines, vertebrate cell culture technology and, 146, 202 bioreactors, 172, 174, 176 traditional cultures, 151, 153 Vacuoles intracellular calcium messenger system and, 316-317, 319-320, 324 symplast and, 263-265, 268 Valinomycin, intracellular calcium messenger system and, 316 Vallisneriu gigunteu, intracellular calcium messenger system and, 308 Valonia, symplast and, 289 Vanadate, intracellular calcium messenger system and, 313-316, 318, 328 Vascular bundles, symplast and, 264, 270, 299 Verapamil, intracellular calcium messenger system and. 320 Vertebrate cell culture technology, 145-147, 20 1-204 bioreac tors airlift reactors, 169-170 cell propagators, 170-171 fiber-bed reactors, 185-187

INDEX

fluidized-bed reactors, 178-180 hollow-fiber reactors, 180-183 membrane-based reactors, 183-185 microcarriers, 171-177 microencapsulation, 177-178 packed-bed reactors, 187-192 perfusion, 167-169 production system, 162-167 two-stage reactors, 169 optimization. 192-193 bioreactor environment, 199-201 cellular metabolism, 193-195 mixing in bioreactors, 195-197 oxygen transport, 197-200 three-dimensional cultures Bergenholtz's system, 160, 162 hydrated collagen lattices, 159-160 sail-sheet cultures, 160- 16 I silica-fiber cultures, 156, 158-159 spider web cultures, 153-155 sponge, 154, 156-157 traditional cultures anchorage-dependent cultures, 149- 150 background, 147- I49 growth environment, 151-153 suspension cultures, 150-151 Vicia fuba, intracellular calcium messenger system and, 309, 311, 320-321, 327 Vigna, symplast and, 265 auxin, 280-28 I electrophysiological structure, 266, 268, 270-273 integration of activity, 287, 291-292, 295 proton pumps, 274, 276 Vimentin regenerative cementogenesis and, 40 in striated muscle, 90, 96-98, 112-1 14 Vinculin in striated muscle, 126 cardiac muscle, 108-109, 114-1 15, I17 skeletal muscle, 92-93, 99 Viruses, vertebrate cell culture technology and, 148. 169, 172, 174, 176, 196

365

w Water, symplast and absorption auxin. 277-279, 282-283 integration of activity, 285-286. 289-290, 296 depletion, 280 potential, 264, 285 transport, 287-292, 296-297 uptake, 285-287, 291-297, 300 Wheat intracellular calcium messenger system and, 307, 322-323. 329-330, 332 symplast and, 284 Wound healing regenerative cementogenesis and, 2, 37 established root surfaces in uiuo, 43-45, 47, 49-50 transdifferentiation in medusae and, 245 X

X protein in striated muscle, 82 Xenopus, proteins in striated muscle and. 125 Xylem, symplast and, 299-300 auxin, 279-280, 285 electrophysiological structure, 267-27 1, 273 integration of activity, 285-287, 290-292, 294, 296-299 proton pumps, 275-277

2 Zeu mays. intracellular calcium messenger system and, 31 I Zeugmatin in striated muscle, 91, 95 Zinc, intracellular calcium messenger system and, 319 Zinniol, intracellular calcium messenger system and, 308

This Page Intentionally Left Blank