International Review of Cytology: A Survey of Cell Biology, Volume 104

INTERNATIONAL

REVIEW OF CYTOLOGY A SURVEYOF CELLBIOLOGY VOLUME104

ADVISORY EDITORS H. W. BEAMS HOWARD A. BERN GARY G. BORISY PIET BORST BHARAT B. CHATTOO STANLEY COHEN RENE COUTEAUX MARIE A. DIBERARDINO CHARLES J. FLICKINGER OLUF GAMBORG M. NELLY GOLARZ DE BOURNE YUKIO HIRAMOTO Y UKINORI HIROTA K. KUROSUMI GIUSEPPE MILLONIG ARNOLD MITTELMAN AUDREY MUGGLETON-HARRIS DONALD G. MURPHY

ROBERT G. E. MURRAY RICHARD NOVICK ANDREAS OKSCHE MURIEL J. ORD VLADIMIR R. PANTIC W. J. PEACOCK DARRYL C. REANNEY LIONEL I. REBHUN JEAN-PAUL REVEL L. EVANS ROTH JOAN SMITH-SONNEBORN WILFRED STEIN HEWSON SWIFT K. TANAKA DENNIS L. TAYLOR TADASHI UTAKOJI ROY WIDDUS ALEXANDER YUDIN

INTERNATIONAL

Review of Cytology A SURVEY OF CELLBIOLOGY

Editor-in-Chief

G. H. BOURNE St. George's University School of Medicine St. George's, Grenada

West Indies

Associate Editors

K. W. JEON

M. FRIEDLANDER

Department of Zoology University of Tennessee Knoxville, Tennessee

The Rockefeller University New York, New York and Hackensack Medical Center Hackensack, New Jersqy

VOLUME104

1986

ACADEMIC PRESS, INC. Harcourt Brace Jovanovich, Publishers

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86878889

9 8 7 6 5 4 3 2 1

Contents

Plasmids of Rhizobium and Their Role in Symbiotic Nitrogen Fixation R . K . PRAKASHAND ALANG . ATHERLY

I. I1. 111. IV .

V.

VI . VII .

VIII .

Introduction . . . . . . . . . ......... ............. ........ Physical Studies on Pla ......................................... Strategies and Genetic Methods for a Functional Analysis of Plasmids . . . . . . . . Functions Controlled by Plasmid Genes . . . . Plasmid-Genome Rearrangements ..................................... Relationship between Rhizobium and Agrobucrerium Plasmids . . . . . . . Restriction Endonuclease Maps ................................ Perspectives ............................................. References . . ..................................................

1 2 4 9 16 16 17 18 19

Mouse Mutants: Model Systems to Study Congenital Cataract AUDREYL . MUGGLETON-HARRIS

I. I1. 111.

IV . V.

VI .

Introduction ........................................................ The Search for Chromosomes Associated with the Mutants Cataract . . . . . . . . . . Lens Crystallins .................................................... Cellular Studies on Lens Epithelial Cells ................................ Manipulations of the Cataractous Phenotype ............................. Possible Areas of Research for the Future ............................... References .........................................................

25 27 27 30 32 33 35

Cell Wall Synthesis in Apical Hyphal Growth J . G . H . WESSELS

I. 11. 111.

IV .

Introduction ........................................................ Observations on Living Hyphae ....................................... The Cytoplasmic Components of Apical Wall Growth ..................... Structure of the Fungal Cell Wall ...................................... V

37 38 46 56

vi

CONTENTS

V . Wall Polymer Assembly at the Hyphal Apex .......................

VI .

Summary .................................................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

65 72 73

Connectin. an Elastic Filamentous Protein of Striated Muscle KOSCAKMARUKAMA

............................. I . Introduction . . I1 . ....................... I11. Iv . Native Connectin ................................................... V . Interaction with Myosin and Actin VI . Location in Myofibrils . . . . . . . . . . VII . Connectin as an Elastic Component .......... VIII . Connectin Transformation during Differentiation .......................... IX . Comparative Biochemistry ............................................ X . Perspectives ............. .... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

81 81 83

86 96 99 103 105

107 110 112

Cell Interactions during the Seminiferous Epithelial Cycle MARTTIPARVINEN. KlMMO K . VIHKO.

AND

JORMATOPPARI

I . Introduction . . . . . . . . ............................................ I1 . Spermatogenic Cell Ty and Their Metabolism ......................... 111. Cycle. Wave. and Transillumination of the Seminiferous Epithelium . . . . . . . . .

IV . Transcriptional Activity during Spermatogenesis and the Function of the Chromatoid Body

V . Cyclic Interaction between Sertoli Cells and Spermatogenic Cells . . . . . . . . . . . .

VI .

VII . VIII . IX .

Localization and Function of Plasminogen Activators in the Seminiferous Epithelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Spermatogenesis in Vitro ......................... . . . . . . . . . . . . . . . . . . . . Interaction between Seminiferous Tubules and the Leydig Cells . . . . . . . . . . . . . Conclusions and Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References .........................................................

115 117 121 125 129 138 142 144

146 147

The Cytoskeleton in Protists: Nature. Structure. and Functions JEANGRAIN

I. I1 . Ill . IV . V.

Introduction ........................................................ Actin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Myosin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Roles of Actin and Myosin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tubulins and Microtubules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

153

154 172 176 I85

VI. VII. VIII. IX.

X. XI.

CONTENTS

vii

Periodic Fibers . . . , . . . , . . . . , . . , . . . . , . . , . . . . . , . , . . , . . . . . . . . . . . , . . . . . . Intermediate Filaments Epiplasm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nonactin Microfilaments Filament Systems of Unkn .................................. . . . . . .. . . Conclusion . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

217 224 228 232 238 24 1 242

The Electrical Dimension of Cells: The Cell as a Miniature Electrophoresis Chamber ARNOLDDE LOOF

I. 11. 111. IV . V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structure of the Plasma Membrane and the Generation of the Transmembrane Potential and Transcellular Currents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Selected Functions of Ionic Currents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Effects of Imposed Electrical Fields on Cellular Activities . . . . . . . . . . . . . . . . . . Closing Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References ............................

INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

25 I 252 294 328 333 335 353

This Page Intentionally Left Blank

rNTERNATIONAL REVIEW OF CYTOLOGY. VOL. 104

Plasmids of Rhizobium and Their Role in Symbiotic Nitrogen Fixation R. K. PRAKASH* AND ALANG. ATHERLYT *Native Plants, lnc. Salt Lake City, Utah 84108 and fDepartment of Genetics, Iowa State University, Ames, Iowa 50011 I

I. Introduction Rhizobia are gram-negative soil bacteria which fix nitrogen in a symbiotic association with plants of the family Leguminosae; however, this classical definition must be extended now to include nodulation and nitrogen fixation on a nonlegume, Purusponiu (Trinick, 1973). Rhizobium purusponiu can infect this tropical tree and fix nitrogen in a manner very similar to that observed in legumes. In both cases, establishment of the symbiosis starts with invasion of plant root or stem by free-living rhizobia followed by a series of steps that result in the formation of a nodule. It is in these nodules that nitrogen fixation takes place. Both the plant and the bacteria undergo differentiation that is regulated by gene expression. The details of recognition and nodule formation are only beginning to be understood. It is the purpose of this review to discuss recent literature and the current state of knowledge concerning these genes, their location, methods of study, and organization. Members of the genus Rhizobium are of great economic importance because of their ability to fix nitrogen. The genus has somewhat informally been separated into those species that are fast growers and those that are slow growers. This separation has recently become more formal (Jordan, 1982) by the creation of a new genus, Brudyrhizobium, which includes all of the slow growing species while the fast-growing species associating with root soybean have now been reclassified as Rhizobium fredii (Scholla and Elkan, 1984). The classification was done mainly because of many clearly distinct differences between slow- and fast-growing species. Some of these differences are summarized in Table I. Recently the clear distinctions between fast- and slow-growing strains of rhizobia have been flurred by the discovery of strains that seemingly have some characteristics of both groups (Stowers and Eaglesham, 1983; see Broughton er ul., 1984, for a listing of numerous references). Thus a third class of rhizobia exist that nodulates many of the same plant species as the slow-growing and fastgrowing strains. An understanding of the genes of Rhizobium involved in plant symbiosis and 1 Copyright D 1986 by Academic Press, Inc. All rights of reproduction in any form reserved.

2

R. K. PRAKASH AND ALAN G. ATHERLY TABLE I SOMEPHENOTYPIC CHARACTERISTICS OF Rhizobium Phenotype

Fast growers

Slow growers

Doubling time Mannitol medium Disaccharide utilization

2-4 hours Acid production

7-20 hours Alkaline production

(+)

(-)

6-Phosphogluconate dehydrogenase Antibiotic sensitivity Free-living acetylene reduction

(+)

(-)

5-50 pg/ml

100-1OOO pg/ml

(-)

(+)

Peritrichous 50.6-63.1%

Subpolar 62.8-65.5%

Deley and Russel (1965)

0.20-0.40 M

0.10 M

Yelton ef a/. (1983)

Flagellation Guanine and cytosine Salt tolerance

Reference

Sadowsky ef a / . (1983); Glenn and Dilworth (1981) Martinez-DeDrets and Arias (1972)

McComb ef a / . (1975) Kurz and LaRue (1975) Pagan ef a/. ( 1975)

nitrogen fixation has moved very rapidly with the fast-growing strains (Rhizobium leguminosarum, Rhizobium trifolii, Rhizobium meliloti, and more recently, R. parasponia and R.fredii). This is largely due to their ease of handling, growth rate and lysis, as well as the ability to apply traditional techniques of genetic analysis. On the other hand progress has been hindered in the genetic analysis of the slow-growing strains [Bradyrhizobium japonicum, Bradyrhizobium sp. (Vigna), Bradyrhizobium sp. (Lupinus), as well as species that nodulate Ornithopus, Cicer, Sesbania, Leucaena, Mimosa, Lablab, Acacia, and Parasponia] due to the difficulty of application of the same techniques.

11. Physical Studies on Plasmids A. DETECTION The techniques initially used to isolate plasmid DNA involved alkaline denaturation, renaturation, and removal of single-stranded chromosomal DNA from the preparation (Currier and Nester, 1976; Nuti et al., 1977; Ledeboer er al., 1976; Casse et a f . , 1979; Prakash et al., 1980;'Hirsch et al., 1982). By analysis on gel electrophoresis, the plasmid profiles of particular strains have been determined. Large plasmids with molecular weights ranging from 90,000,000 to 300,000,000 have been identified in most of the fast-growing strains of rhizobia. For literature before 1980, several recent reviews should be consulted (Denarie er al., 1981; Nuti et al., 1982). Recently, using a modified Eckhardt (1978) procedure, a megaplasmid not observed earlier was detected in several fast-growing rhizobia

PLASMIDS OF RHlZOBlUM

3

(Rosenberg er u l . , 1981; Masterson er al., 1985; Heron and Pueppke, 1984). An R. meliloti megaplasmid has been estimated to be 1000 MDa using electron microscopy (Burkhardt and Burkhardt, 1984). Recent studies indicated the presence of at least two megaplasmids, of the same size in R. meliloti (Banfalvi et al., 1985). Very rarely are plasmids of less than 50 MDa found in rhizobia and usually the number of different plasmids within a strain varies from one to six. When the combined size of plasmids and chromosome are totaled the genome of Rhizobium species becomes very large (5.4-7.6 X lo9 Da; Chakrabarti et al., 1983) which is severalfold larger than the Escherichia coli genome (2.2 X lo9 Da), thus plasmid DNA may represent as much as 25% of the total DNA of some strains. Rhizobium plasmids have been given designations as proposed by Prakash er al. (1980). The indigenous plasmids of Rhizobium are named as pRtr, pRle, pRme, pRph, pRja, etc. (for plasmids of R. trifolii, R. leguminosarum, R . meliloti, Rhizobium phaseoli, B. japonicum, respectively) followed by the number of the strain in which the plasmid was found and serial letter a, b, c, etc. in case of multiple plasmids where a designates the largest plasmid in a series. Plasmids containing symbiotic genes are also sometimes referred to as pSym. Slow-growing strains of Rhizobium have only been recently examined for the presence of plasmids because gentle lysis procedures did not readily adapt to the slow-growing strains. An exception was the analysis of a number of slowgrowing strains isolated from alkaline soils (Gross er al., 1979). More recently, Masterson et (11. (1982) and Cantrell et al. (1982) developed a plasmid isolation procedure that is useful with all Bradyrhizobiurn strains. Using this procedure, several B. japonicum strains (USDA 110, 122DES, 143, 136, 142, 6, and SR) have been shown not to contain any plasmids. Very large plasmids are present in most B. japonicum strains including 61A76, 61A24, USDA strains 31, 71a, 74, 94, and 143 (Haugland and Verma, 1981; Masterson et al., 1985).

B. NUCLEICACIDHYBRIDIZATION DNA hybridization studies have provided information about the relatedness of plasmids derived from the same or different species of bacteria. Jarvis et al. (1980) compared the DNA sequences from total DNA of 35 different fastgrowing strains of Rhizobium by DNA:DNA reassociation kinetics and found 40-80% sequence similarities. On the other hand, Hollis et al. (1981) compared a wide variety of slow-growing species for DNA sequence homology and found that they could be grouped into three homology groups: high homology, moderate homology, and very little sequence homology (10% or less). Haugland and Verma (1981) examined the intraspecific homology of total DNA sequences in three strains of B . japonicum. Strains 61A76 and I10 showed very little sequence homology with eachrother (24%), while a third strain 61A24, showed 50% homology with strain 110. Although strain 61A76 was found to have one large plasmid and 61A24 was found to have two plasmids, no discern-

4

R. K. PRAKASH AND ALAN G. ATHERLY

ible plasmid could be detected in strain 110. Heterologous hybridizations between plasmid and total DNAs from these strains indicated that sequences which were plasmid-borne in one strain were located on the chromosomal DNA in other strains. Jouanin et al. (1981) analyzed the sequence homology of the plasmids of symbiotically effective strains of R. meliloti. chosen because of their wide geographical origin. They used restriction endonuclese patterns and Southern DNA:DNA hybridization to reveal sequence homologies of the purified plasmid DNA. Sequence homology was present between all plasmids tested, irrespective of geographical origin. The homology was concentrated in a few restriction fragments, but because restriction maps are not available they could not provide evidence for any degree of clustering of these sequences. Likewise, they did not indicate any biological function that might be associated with these highly conserved sequences. Prakash et al. (1981) found that plasmids from different fast-growing Rhizobium species share extensive homology. Organization on the restriction endonuclease map indicated that these homologous sequences are clustered within a region that contained sequences for symbiotic nitrogen fixation (Prakash et al., 1982a,b). Masterson et al. (1985) examined the homology of a R. fredii Sym plasmid with the plasmid DNA and total DNA from various other R. fredii strains and 8 . japonicum strains. They found extensive homologous sequences in plasmid DNAs as well as in chromosomal DNA of different R. fredii strains. However, in one R . fredii strain USDA194 and in B. japonicum strains the homology was observed only with total DNA. A possible explanation for highly conserved sequences in plasmid DNA was obtained by Watson and Scholfield (1985). In R. trifolii, they found a repeated sequence that is a reiteration of the nijHDK promoter. This sequence is highly conserved within all geographically distinct isolates and located exclusively in the Sym plasmid. The sequences were specific for R. trijolii. They propose a model in which the expression of symbiotic genes is host-specifically activated via their species-specific sequences. 111. Strategies and Genetic Methods for a Functional Analysis of Plasmids

A. TRANSPONSONS, GENEFUSIONS,AND MARKER EXCHANGE A number of symbiotically important functions are coded by the plasmids of fast-growing strains, but no such functions can be assigned to the plasmids of the slow-growing rhizobia. A wide variety of approaches have been used to identify

PLASMIDS OF RHIZOBIUM

5

and characterize plasmid-borne genes and gene functions. These includes plasmid transfer, plasmid curing, mutant analysis, expression in minicells, fusion of genes with facZ, and restriction endonuclease mapping. Some of these procedures apply to both plasmid genetic analysis as well as to genome analysis. An extremely useful approach for genetic analysis of Rhizobium and other gram-negative bacteria is transposon mutagenesis. Insertion of a transposon results in a relatively stable mutation (insertional inactivation). One disadvantage is that transposon insertion usually results in polar mutations (Berg et a f . , 1980). A variety of suicide vehicles to deliver transposons into Agrobucrerium and Rhizobium are available. The first of these vectors was developed by Van Vliet et al. (1978) and Beringer e t a f .(1978) and has been effectively used to genetically tag plasmids for plasmid transfer (Johnston et a f . , 1978; Hooykaas et a f . , 1981). The vectors were designed to replicate only in E. cofibut not in other nonenteric, gram-negative bacteria. When replication is not possible, the transposon present in the plasmid is rescued by insertion into the host DNA. For unknown reasons the presence of Mu phage DNA sequences in RP4 does not allow replication of RP4 in Rhizobium or Agrobacterium. Unfortunately, the Mu-induced failure of RP4 to replicate can occasionally be overcome, thus complicating interpretation of results (Simon et a f . , 1983). Another complicating event is the frequent simultaneous transposition of both Mu and Tn5 sequences which then makes genetic analysis difficult (Banfalvi et a f . , 1981b; Meade et a f . , 1982; Forrai et al., 1983). To overcome these problems, Simon et al. (1983), Selvaraj and Iyer (1983), and Yakobson and Guiney (1984) have developed broad-host-range mobilization systems for transposon mutagenesis. Simon et uf . (1983) constructed a family of vector plasmids containing the P-type recognition site for mobilization (mob site) which can be mobilized with high frequency from a donor strain carrying the transfer genes (oriT)of Incp-type plasmid RP4 integrated into the chromosome. The mobilization vectors were derived from pACYC184, pACYC177, and pBR325 which are able to replicate in E. cofi and its close relatives but not in Rhizobium and Agrobacterium. Vectors were prepared that include Tn5 (Nm, Km; pSUPlOl1) Tn7 (Sp/Sm; pSUP2017), and Tcr genes integrated into a nonfunctional area of Tn5 (pSUP10141) which then gives Tc‘ and Nmr or Kmr simultaneously. In a similar system developed by Yakobson and Guiney (1984), the 760-base pair oriT region of broad-host-range plasmid RK2 was cloned into Tn5 residing on a derivative of pBR322. The resulting “suicide” plasmid, pEYDGl , can be transferred to a variety of gram-negative bacteria in the presence of RK2 helper plasmid pRK2073. Once transposon-generated mutants are isolated, replicons which have acquired Tn5-oriT can be mobilized to other gram-negative hosts following the introduction of helper plasmid pRK212.1 into the mutants. Selvaraj and Iyer (1983) constructed suicide plasmids containing Tnl , Tn5, or Tn9 in a pl5A-type of replicon with an N-type of bacterial mating system. These

6

R . K . PRAKASH AND ALAN G . ATHERLY

vectors also replicate in E . coli but not in Rhizobium and because of the N-type mating system they are very efficient in matings with E . coli and Rhizobium. These vectors are useful for random mutagenesis as well as for site-specific mutagenesis. Tn5-induced mutant clones of genes can be reinserted into a genome by homologous recombination, selecting for kanamycin. For example, a clone of the recA gene containing Tn5 (pRMB 1002) has been isolated by Better and Helinski (1983) from R . melifoti genome bank. By reinserting this cloned gene into a suicide vector (pSUP205; Simon et al., 1983) the Tn5 in the recA gene can be rescued creating a recA mutant. Such a procedure has been used by Shantharam and Iyer (1986) to construct a double mutant affected in recombination in R. meliloti. Simon et af. (1983) and Hahn and Hennecke (1984) have used this approach to create site-specific mutants of R . melifoti and B . japonicum, respectively. The procedure initially developed for site-directed mutagenesis (Ruvkun and Ausubel, 1981) involved the cloning of restriction fragment containing the mutagenized gene of interest on a P-group broad-host-range vector. This is followed by replacement of the wild-type parental DNA sequence with the mutant sequence by conjugation with a second P-group plasmid, with simultaneous selection for Tn5 and a marker on the second plasmid. R. K. Prakash and A. G . Atherly (unpublished) further simplied the site-specific mutagenesis technique by cloning the Tn5-mutagenized gene in vector pBR322, which can replicate in E . coli but not in Rhizobium. This vector has been found to be mobiliziable with the plasmid DNA that mobilize the P-group broad-host-range vector. Recently lac gene fusions have been used to study a wide variety of genetic functions both at the transcriptional and translational levels (Weinstock et al., 1983). lac fusions are especially useful in analyzing gene functions and regulation where the gene product is difficult, if not impossible, to assay or when the gene in question is turned on/off under undefined conditions. Techniques for fusing genes with lac have been very well developed (Silhavy et al., 1984) and a large number of convenient cloning vectors have been constructed to facilitate genetic analysis (Casadaban et al., 1980; Kahn and Timblin, 1984). These have been applied to the analysis of regulation of nifgene activity in rhizobia. The technique of lac gene fusion has been extremely useful for analyzing trans-acting regulatory functions of the ntrA, n t E , and nifA genes on nijKDH promoters (Sundaresan et al., 1983a,b; Szeto et af., 1984). One criterion for studying lacZ gene fusions to heterologous promoters is for the host cell to have a low level of endogenous P-glactosidase activity. This is true in most fast-growing strains of Rhizobium, and, hence, fur mutants are desirable but not necessary if Rhizobium is used as the host cell (Szeto et al., 1984). A potentially very useful technique of facZ gene fusion makes use of the combination of suicide plasmid and lacZ in a transposable element (Simon et a f . , 1983; Selvaraj and lyer, 1983) and Mu-d lac. Castillo et af. (1984) have constructed mini-Mu transposable elements containing lac operon structural genes.

PLASMIDS OF RHIZOBIUM

7

These mini-Mu elements have selectable genes for either ampicillin or kanamycin resistance and can be used to form transcriptional and translational lac gene fusions. Olsen et al. (1985) developed a procedure for selecting mini-Mu insertions by inserting Mu-d lac (kan) into the suicide vector pGS6 (Selvaraj and Iyer, 1983). They mated this vector to R . fredii strain USDA201. Kanamycinresistant transconjugants arose at a frequency of lop4 and appear to insert randomly into the genome. Several transconjugants exhibited increased lac expression when grown in the presence of Glycine max root exudates and nodule extracts, suggesting fusion to nod (nodulation) andfir (fixation) gene promoters. Another potentially useful promoter probe for Rhizobium has been recently developed for Caulobacter crescentus. Bellofatto et al. (1984) removed the promoter of Tn5 while still retaining its translational start site and inserted it into a suicide plasmid containing Mu phage sequences. Consequently, neomycin phosphotransferase is expressed only when fused to the functional promoter in the genome. B. CLONING VECTORSAND

THE

USE OF PHAGE

A considerable number of DNA cloning vectors have become available over the years; however, the vast majority of them are restricted for use in E . coli. Since 1980, two groups of broad-host-range DNA cloning vectors have been developed for use in specialized functions. The first of these vectors (pRK290) was developed by Ditta et al. (1980) from RK2, a large (56 kb) P-1 incompatibility group plasmid. RK2 and its derivative plasmids have the ability to transfer at a high frequency via conjugation into a wide variety of gram-negative bacteria, including species of Rhizobium, Klebsiella, Serratia, Pseudomonas, Acinetobacter, and Agrobacterium. pRK290 is very stable in R . meliloti, with only a 0.2% loss per generation (Ditta et al., 1980). The 20 kb plasmid confers tetracycline resistance, is not mobilizable (mob ,tra-), and contains a single recognition site for EcoR1 and BglII. However, a second derivative of RK2 (Figureski and tielinski, 1979) contains the tra functions and the neomycin resistance gene ligated to a ColE1 replicon and will mobilize pRK290, respectively. To further extend the use of pRK290, Friedman et al. (1982) and Knauf and Nester (1982) added a cos fragment into the unique Bglll site creating the cosmid pLAFRl , and pVKlOO and pVK102. The plasmid pRK290 has now been reduced in size to 10 kb and has been modified for gene expression studies for a wide variety of gram-negative bacteria (Ditta et al., 1985). A second group of broad-host-range cloning vectors has been developed from the 8.9 kb IncQ/P4 plasmid RSFlOlO (Bagdasarian et al., 1981; Guerry et al., 1974). Simon et al. ( 1983) removed the replication and mobilization functions of pKT210, a chloramphenicol-resistant derivative of RSF1010, and inserted them into pACYC184, pBR325, orpACYC177 which resulted inpSUP104, pSUP204, and pSUP304, respectively. By inserting a 0.40 kb cos fragment into the single +

8

R. K. PRAKASH AND ALAN G . ATHERLY

PsrI site of pSUP104, a cosmid pSUP106 was developed. However, fragments greater than 10 kb cloned into pSUP106 are very unstable and cannot be easily maintained (R. K. Prakash and A. G. Atherly, unpublished). RSFlOlO and its derivatives are nontransmissible plasmids but are efficiently moblized by Inc- 1 plasmids or the mob site of IncP-1 plasmids. David et al. (1983) using derivatives of RSF1010, (pKT248. pKT210; Bagdasarian eral., 1981)demonstratedthat they were transferred at a high rate from E. coli to R. melilori in the presence of RP4 and were stably maintained. A valuable adjunct of plasmid cloning vectors for genetic analysis is the discovery of generalized transducing phage in R . melilori (Casadesus and Olivaries, 1979; Sik e f al., 1980; Martin and Long, 1984; Finan er al., 1984), R . leguminosarum and R. trifolii (Buchanan-Wollasten, 1979), and R. japonicum (Shah et al., 1981, 1983). Not only can phage be used for fine structure mapping and strain construction of Rhizobium but also for transfer of large plasmids from one strain to another. The molecular size of the phage N3 (Martin and Long, 1984) is very large (approximately 195 kb) thus allowing transfer of large plasmids intact. Unfortunately, rhizophage are restricted to only a few species and are usually strain specific. C.

PLASMID

TRANSFER AND R-PRIMES

Plasmids present in Rhizobium species are frequently of very high molecular weight (150,000,000 to more than 500,000,000) and therefore biochemical and genetic analysis is extremely difficult. Procedures that reduce the working size of a DNA segment or plasmid are therefore very helpful. Identification of symbiotic functions can be accomplished by the transfer of whole or partial plasmids from Rhizobium to E. coli or to plasmidless Rhizobium and Agrobacrerium strains. Plasmids coding for symbiotic functions in some strains of R . leguminosarum, R. trifolii, and R. phaseoli are self-transmissible (Higashi, 1967; Johnston er al., 1978; Beynon er al., 1980; Rolfe et al., 1981; Hooykaas er al., 1981, 1982a; Lamb er al., 1982; Scott and Ronson, 1982), but self-transmissibility has not been observed in symbiotic (Sym) plasmids from R. melilori or R. fredii (Kondorosi er al., 1982; Atherly et al., 1985; Appelbaum et al., 1985). In the case of nontransmissible plasmids, several approaches are available to obtain transfer to new hosts. Using R plasmid, Hooykaas et al. (1982a,b) developed a helper plasmid PRL180 to transfer nontransmissible plasmid. Kondorosi er al. (1982) inserted the mob region of the P-1 type plasmid RP4 into the megaplasmid pRme4lb of R. melilori and subsequently obtained pRme4lb transfer at a frequency of 1 X Kanamycin on Tn5 was used as a selective marker but transconjugants were also tested for nod (nodulation) and f i x (fixation). mobRP4 was inserted into the megaplasmid by cloning a fragment of pRme4lb and the mob region into pBR322. This hybrid plasmid was mobilized into R. melilori but will not replicate; however, recombination occurs due to homology between the

PLASMIDS OF RHIZOBIUM

9

resident plasmid and the hybrid pBR322. A similar approach was used by Julliot et al. (1984) for creating R’ plasmids of R. melifoti pSym megaplasmid of strain 201 1. After mobilization of the cointegrate RP4:pSym into E. coli, they obtained a set of RP4-primes that contained large fragments of the pSym megaplasmid. One of them has been extensively characterized (285 kb) and contains the symbiotic genes. Simon (1984) constructed a convenient suicide vector by inserting mob into Tn5 which, in turn, was present in a suicide vector. Thus, by inserting this mob:Tn5 into a Rhizobium plasmid, mobilization could be effected. Appelbaum et al. (1985) used this approach to mobilize (with RP4 in trans) the Syrn plasmid of R. fredii. D. PLASMID DELETION AND CURING P-1 group plasmids will occasionally form cointegrates with resident Rhizobium plasmids if naturally occurring plasmid-plasmid homology sites exist. The P-1 group plasmids apparently can be maintained as integrated units only if a recA or low recA gene activity is present. Ronson and Scott (1983) obtained a cointegrate of pRtr514 and R68.45, which totaled about 770 MDa and transferred it to a variety of different bacterial strains at frequencies as high as Similarly, cointegrates of Sym plasmid from R. fredii strain USDA191 and PRL180 (Hooykaas er al., 1982a,b) were transferred to E. coli and Agrobacreriurn tumefaciens (Engwall et al., 1986). But, cointegrates were very unstable in recA+ E. coli strains yielding only pRLl8O upon conjugation. In recA- E. coli hosts the Sym plasmids suffered random deletions and it was thus possible to create a family of deletions encompassing the entire plasmid. Useful information on plasmid function can also be obtained from plasmidcured strains, such as lost phenotypic characteristics and reintroduction of plasmid fragments to regain only specific plasmid functions. Plasmid curing is most easily accomplished by prolonged exposure to high temperatures (Zurkowski and Lorkiewicz, 1978), but this procedure may also result in internal deletions within plasmids (Banfalvi et al., 1981b). Heat treatment of slow-growing strains of R. japonicum also resulted in deletions with the majority of them occurring in the nif (nitrogen fixation) or nod (nodulation) region (Skogen-Hagenson and Atherly, 1983).

IV. Functions Controlled by Plasmid Genes A. GENETIC EVIDENCE FOR SYMBIOTIC NITROGEN FIXATION GENES The study of symbiotic nitrogen fixation in Rhizobium has been mainly limited to Rhizobium plasmids. A great percentage of DNA in Rhizobium is in the form of large plasmids (up to 25%) and it was of interest to know what genes are

10

R . K . PRAKASH AND ALAN G. ATHERLY

carried on these large plasmids. Some early genetic evidence indicated that in Rhizobium the symbiotic nitrogen fixation genes might be on the plasmid DNA (Higashi, 1967; Dunican and Cannon, 1971). Several genetic and physical studies have now clearly established that, at least in fast-growing Rhizobium species, the genes for symbiotic nitrogen are usually on a large plasmid. Zurkowski and Lorkiewicz (1976, 1978, 1979) and Hooykaas et al. (1981) showed that nonnodulating mutants of R. trifolii, resulting from treatment at high temperature, were due to loss of plasmid DNA. This nod- mutant also lost its ability to attach to the root hair surface and that property was also restored upon the reintroduction of the plasmid. These data strongly suggested the involvement of plasmid DNA in nodulation. Furthermore, Johnston et al. (1978) demonstrated that transfer of the Tn5 marked plasmid into a j r strain of R . leguminosarum restored its normal symbiotic functions, strongly implying that the symbiotic genes were located on plasmids. Brewin et al. (1980a) showed that transfer of R. leguminosarum bacteriocinogenic plasmid pRLl JI into four symbiotic mutants restored them to normal symbiotic function. In R. meliloti, genes controlling the early and late function in symbiosis were reported to be on a megaplasmid (Rosenberg et a f . , 1981). Further Palomares et al. (1978, 1979) showed that in R . meliloti, extrachromosomal DNA was responsible for polygalacturonase, a key enzyme responsible for the early infection process (Ljunggren and Fahraeus, 1961). Thus, it seems clear that early functions in the infection process are plasmid controlled in R . meliloti. Using the genetic strategy described in Sections III,C and D the presence of both nodulation and nitrogen fixation genes on a fastgrowing Rhizobium plasmid has been established by other authors (Banfalvi et al., 1981b, 1983; Kondorosi et al., 1982, 1983; Julliot et al., 1984; Rolfe e t a l . , 1983; Appelbaum et al., 1985; Hombrecher et al., 1981a; Kowalczuk et a / . , 1981; Sadowsky and Bohlool, 1983). In addition to nifand nod genes the Sym plasmid seem to confer host specificity to Rhizobium. Recently Brewin et al. (1981) reviewed the role of Rhizobium plasmids in host specificity. Plasmid controlled host-range specificity in R . leguminosarum was first described by Johnston et al. (1978). In this study a plasmid (pJB5JI) bearing Tn5 was transferred to R . trifolii and R . phaseoli. Transconjugants arose at a frequency of lo-* and all were capable of forming nodules on peas, in addition to retaining their normal host nodulation properties. Later, several other reports provided evidence for the presence of host-range determinants on other Sym plasmid DNA (Brewin et al., 1980c; Hooykaas et al., 1981, 1982a,b; Djordjevic ef al., 1983; Morrison er al., 1983; Lamb et al., 1982; Appelbaum et al., 1985). Besides Rhizobium species, the Sym plasmid has also been transferred to closely related Agrobacterium. Hooykaas et al. (1981, 1982a) transferred the Sym plasmid from R. trifolii and R. leguminosarum to A . tumefaciens LBA288 (a nonvirulent strain cured of its tumorigenic plasmid) and observed that the trans-

11

PLASMIDS OF RHIZOBIUM

conjugants were able to form nodules on clover. Electron microscopy of the nodules showed root hair curling and infection threads filled with bacteria. Furthermore, the bacteria were surrounded by a periplasmic membrane indicating that the agrobacteria had been released from the infection thread into the plant cell. But there were no typical bacteroid-like structures and no nitrogen fixation was observed. Similar findings are obtained with other Rhizobium plasmids when transferred to Agrobacterium (Kondorosi et a / . , 1982, 1983; Broughton et al., 1984; Djordjevic et al., 1983). In most cases the agrobacteria were confined to the infection threads and the nodule was essentially devoid of bacteria (Hirsch e t a / . , 1985; Schofield et al., 1984; Truchet et al., 1984; Wong et a/., 1983).

B. EXPRESSION OF Sym PLASMID

IN

RHIZOBIACEAE

Transfer of Syrn plasmids within Rhizobiaceae often results in variable expression. Hooykaas e f al. (1981, 1982a,b) observed that the Syrn plasmids of R . leguminosarum and R. rrifolii expressed symbiotic nitrogen fixation properties completely when transferred between these two strains. However when these Sym plasmids were transferred to A . tumefaciens or R . meliloti they induced root nodules but did not fix nitrogen. Djordjevic et al. (1983) and Rolfe et al. (1983) found that transfer of plasmid pJB5J1 or pBR IAN, which encodes pea and clover specificity, respectively, to various R . meliloti plasmid-cured strains did not result in the ability of R. mefifoti to nodulate peas and clover. The transfer of pBRl AN to an A . tumefaciens strains conferred clover nodulation to this strain, whereas pJB5JI could not induce this Agrobacrerium strain to nodulate peas. The transfer of pBR 1 AN to various Syrn plasmid-cured or deleted R. leguminosarum or R . rrifolii strains resulted in the ability of these strains to nodulate clover, whereas the reverse was true when pJB5JI was transferred to these strains. When plasmid pJB5JI and pBRlAN were introduced into a fast-growing R. parasponium strain, the resulting transconjugants showed a change in the spectrum of plants that could be nodulated. The plasmids have also been transferred to slowgrowing Rhizobium species, but the plants had an ineffective phenotype. The transfer of a host-range plasmid pJB5J1 from R . leguminosarum to R. fredii elicited only early stages of nodule development on peas (Rviz-Sainz et al., 1984). On the other hand R. fredii Syrn plasmid from strain USDA191 is Fix in ANU265 genetic background (Appelbaum et a/., 1985). ANU265 is a pSymcured derivative of NGR234, a broad-host-range fast-growing strain. This implies that the chromosomal genetic background of NGR234 is very similar to R . fredii. Incompatibility and instability of the Syrn plasmids could also be attributed to variable expression of the Syrn plasmid in different hosts (Djordjevic et al., 1982; Christensen and Schubert, 1983). +

12

R. K. PRAKASH AND ALAN G.ATHERLY

C. PHYSICAL EVIDENCE FOR NITROGEN FIXATION GENES The first physical evidence for the presence of nifgenes on a plasmid was obtained by hybridizing R. leguminosarum plasmid with the cloned nif structural genes KDH of Klebsieflapneumoniae (Nuti et al., 1979). The nifstructural genes of K . pneumoniae are found to be highly conserved in all nitrogen-fixing bacteria (Ruvkun and Ausubel, 1980). With the exception of R. melilori, in most of the fast-growingRhizobium species the nifgenes are located on medium size isolatableplasmids (Prakashetal., 1981; Broughtonetal., 1984;Mastersonet u f . ,1982, 1985).Rhizobium meliloti nifgenes are present on the megaplasmid (Rosenberg et al., 1981; Banfalvi et al., 198lb). Slow-growing strains of Rhizobium. including B. japonicum, have not been shown to carry nif sequences on plasmid DNA (Mastersonet al., 1982; Haugland and Verma, 1981). Likewise in a fast-growing R. fredii strain (USDAl94) homologous sequences for nif structural genes were identified only when total DNA was used (Masterson e t a f . , 1985). DNA hybridization studies indicated that in R. phuseoli (Quinto et al., 1982) and R. fredii (Prakash and Atherly, 1984) the nif structural genes are reiterated. The nif structural genes have been cloned and characterized from several Rhizobium species (Ruvkun and Ausubel, 1980; Schetgens et af., 1984; Hennecke, 1981; Schofield et al., 1983; Quinto et al., 1985). Site-directed mutagenesis and complementation studies verified that the region homologous to K . pneumoniae nif structural genes is involved in nitrogen fixation (Schetgens er al., 1984; Ruvkun and Ausubel, 1981; Ruvkun et al., 1982a). Further, RNA isolated from nitrogen-fixing nodules hybridizes strongly to the nifregion on the plasmid DNA (Prakash et al., 1982a; Krol et al., 1980, 1982; Corbin et af., 1982, 1983). Apart from nifstructuralgene KDH, the nifregulatory gene A of K . pneumoniae, has also been localized on plasmid DNA of R. leguminosarum (Downie et al., 1983a),R . meliloti (Szetoer al., 1984), and in chromosomal DNA of B. japonicum (Fuhrman et af., 1985). In most of the fast-growing Rhizobium species the nifand nod genes have been found to be located on one plasmid except in few strains of R . fredii. In some R . fredii strains the nif and nod genes are not only present on different plasmid but also specified on the chromosome (Masterson et al., 1985; Scholla et al., 1984; Mathis et al., 1985).

D. IDENTIFICATION AND ISOLATION OF NODULATIONGENES In addition to nif structural genes, other genes involved in effective nitrogen fixation and nodulation have been identified. Using both nitrosoguanidine and transposon mutagenesis, Forrai er al. (1983) identified 5 nod- (noduation) and 57 Fix- (fixation) symbiotic mutants of R . meliloti. Of these mutants 1 nodand 11 Fix- mutants were localized on the chromosome while 5 Tn5-induced

PLASMIDS OF RHIZOBIUM

13

Fix- mutants and 1 Tn5 nod- mutants were localized on the megaplasmid. Long et al. (1981) and Meade et al. (1982) also found several nod- and Fixmutants of R. meliloti using Tn5 mutagenesis. Among 19 symbiotic mutants characterized, at least 6 were shown to reside on the megaplasmid (Buikema et al., 1983). Microscopic examination of four nodulation-defective ( n o d - ) mutants (Hirsch et a f . , 1982) showed that some did not induce root hair curling and entered root epidermal cells even though no infection threads were formed. In R. phaseoli, 10 symbiotic Tn5-induced mutants were isolated (Noel et a f . , 1984). Of three that were located on the Sym plasmid, one mutation led to abnormal nodule development and two of the mutations, which eliminated nitrogenase activity, were located outside the immediate region of the nif structural genes. The remaining seven mutants were seemingly on the chromosome. They resulted in slow nodule development and nodule dispersement on the root system. Three of the chromosomally located mutants (Vandenbosch et al., 1985) induced the formation of uninfected root nodule-like swelling on bean. The technique which utilized transponon Tn5 mutagenesis has facilitated the isolation of nodulation genes from several Rhizobium species. In R. trifolii the nodulation region, identified by Tn5 insertion, was cloned and used to isolate the wild-type sequence. Reintroduction of the wild-type sequence, present on a 14 kb Hind111 restriction fragment, into Sym-plasmid-cured R. trifolii strain and A. tumefaciens resulted in the restoration of nodulation on clover (Schofield et a l . , 1984). This 14 kb nod fragment of R. trifolii also formed nodules on clover plants when present in Lignobacter and Pseudomonas strains (Plazinski and Rolfe, 1985). All nodules formed by Lignobacter transconjugants showed bacterial release from the infection threads into host cytoplasm. Pseudomonas transconjugants formed pseudonodule without bacterium, except within the cellular species of the outermost cells. Similarly a 10 kb region of DNA-encoding nodulation functions was cloned from R. leguminosarum (Downie et al., 1983b). Long et al. (1982) cloned the nodulation gene from R. melilori strain 1021 by complementation studies. In this procedure, a clone bank of wild-type R. meliloti sequences was conjugated into each of two nod- R. meliloti mutants. Plants were inoculated with the transconjugants. The bacteria isolated from the nodules contained a plasmid pRmSL26, bearing nodulation genes. Further it was found (Hirsch et al., 1985; Jacobs et al., 1985) that two subclones of pRmSL26, one an 8.7 kb EcoRI fragment and a 5.5 kb PstI fragment elicited the same response in A. tumefaciens on roots of alfalfa as did pRmSL26. In addition, Hirsch et al. (1985) found that 8.7 kb nod fragment alone is sufficient for nodule morphogenesis. In R. meliloti strain 41, the essential nod genes were localized in 8.5 and 6.8 kb EcoRI restriction fragment, respectively. In nod- deletion mutant lacking the 8.5 kb fragment the nodulation ability on alfalfa was restored upon the introduction of Sym plasmid of R. leguminosarum or R. trifolii (Banfalvi et al., 1981a; Djordjevic et al., 1983; Kondorosi er a l . , 1984). Similarly the nod-

14

R. K. PRAKASH AND ALAN G . ATHERLY

mutant of Sym plasmid from R. meliloti, R . trifolii, and broad-host-range Rhizobium strain can be complemented with the 8.7 kb nod fragment of R . meliloti strain 1021 or 14 kb nod fragment of R. trifolii (Fisher et a l . , 1985; Djordjevic et al., 1985). Moreover, the nod region of the 8.5 and 8.7 kb EcoRI nod fragment of R . meliloti strain 1021 and 41, respectively, hybridized with nod genes from other rhizobia (Prakash and Atherly, 1984; Masterson et al., 1985; Broughton et al., 1984). Therefore, these nod gene clusters are called as “common” nod genes. By complementation and sequence analysis, four genes were identified in the common nod region, which were designated as nod A , B, C, and D ,respectively (Torok et a l . , 1984; Fisher et al., 1985; Jacobs et al., 1985). Comparison of nod A, B, C nucleotide and amino acid sequences of R. meliloti and R . leguminosarum (Torok et al., 1984; Rossen et al., 1984) showed that the organization of these three genes is fairly similar and the nucleotide sequence of nod A, B, and C genes shared 72, 69, and 71.4% homology, respectively. Using directed Tn5 mutagenesis, a nod gene cluster of about 3.5 kb was found within the 8.7 kb EcoRI fragment (Jacobs et al., 1985; Kondorosi et al., 1984). The other 6.8 kb region contains two nod gene regions separated by a 1 kb region nonessential for nodulation (Kondorosi et al., 1984). With the use of 3.5 kb nod sequence as a probe, the nod genes from R. fredii were also cloned (Prakash et al., 1986). As in R. meliloti, the nod genes of R. fredii are not clustered, but instead are present at different regions on the plasmid DNA. Further, in R.fredii, the nod genes are reiterated (Prakash and Atherly, 1984).

E. OTHERGENESINVOLVED IN SYMBIOTIC NITROGEN FIXATION 1. Polysaccharide Synthesis In 1974 an attractive theory involving legume seed lectins was proposed for the specific attachment of Rhizobium to their host plant (Bohlool and Schmidt, 1974). This theory has subsequently been pursued and discussed by numerous investigators (Bauer, 1981; Graham, 1981; Dazzo and Truchet, 1983; Dazzo and Gardiol, 1984). The biochemical basis for this theory is that the seed glycoproteins known as “lectins” bind to specific sugar haptens on rhizobial cell-surface polysaccharides, thus aiding in attachment and recognition. Rhizobium cellsurface polysaccharides have been shown to carry the receptor sites of lectin molecules (Bal et al., 1978; Shantharam et al., 1980). Suggestions for the involvement of Rhizobium plasmid genes in the recognition system stem mainly from the properties of plasmid-free symbiotically defective mutants of Rhizobium. A nod- derivative of R. leguminosarum, obtained by culturing at elevated temperatures, lost pea lectin-binding properties because of the loss of its smallest plasmid (Prakash er al., 1980). Also, a strain of R. trijolii cured of a large plasmid failed to nodulate its host. In addition, it lost lectin-binding capaci-

PLASMIDS OF RHIZOBIUM

15

ty and the ability to attach to the root hair surface. When the Sym plasmid was restored from the wild-type strain, the bacteria regained all of the above properties (Zurkowski and Lorkiewicz, 1979; Zurkowski, 1980). Mutants that are incapable of producing exopolysaccharide (EPS) and nitrogen-fixing root nodules have been isolated from Rhizobium (Sanders et a l . , 1978; Chakravorty et al., 1982) suggesting again that polysaccharides may be involved in the recognition process. From a Tn5-derived mutant of R. trifolii the wild-type DNA sequence was cloned which was capable of restoring the mutant strain to synthesize normal levels of EPS and simultaneously restored its ability to nodulate clover (Chakravorty et a l . , 1982). On the other hand, in Azotobacter, independent transfer of R. trifolii lectin-binding property has been shown without the transfer of nodulating ability (Bishop e f a l . , 1977). Evidence that extracellular polysaccharide is involved in the nodulation of R . meliloti comes from the work of Leigh et al. (1985) and Finan et al. (1985b). They obtained mutants in polysaccharide synthesis by screening with the fluorescent strain calcafluor and also by insensitivity to monoclonal antibodies to the rhizobial surface (Johansen et al., 1984). These two independently isolated sets of mutants (Exo-) formed ineffective nodules on alfalfa and fell into six distinct genetic groups as determined by complementation analysis. Apparently, the exopolysaccharide, although not required for nodule formation, is involved in nodule invasion. One of these mutants has been mapped to a second megaplasmid (pRme-SU47b) which is slightly larger than the symbiotic plasmid (pRme-SU47a) (Finan et al., 1985a). The overwhelming evidence supports the suggestion that polysaccharides are involved in the nodulation process; however, the exact mechanism is not known. It is clear that the genes involved in polysaccharide synthesis are located on both the plasmid and the chromosome.

2 . Hydrogen Uptake Genes Large quantities of energy are lost during the nitrogenase-catalyzed reaction as hydrogen gas. Many Rhizobium species possess an active hydrogen uptake system (Hup) permitting hydrogen to be recycled. Thus, Hup+ Rhizobium strains generally are more effective symbionts than those with Hup- phenotypes (Albrecht e f a l . , 1979). Evidence for the presence of hydrogen uptake genes on plasmid DNA comes from the work of Brewin et al. (1980b) who cotransferred the genetic determinants for hydrogenase activity and nodulation ability between R. leguminosarum strains. DeJong et al. (1982) found that transfer of such plasmids to different R . leguminosarum strains improved symbiotic nitrogen fixation. However, the Hup plasmid from R . meliloti expressed low levels of Hup activity in alfalfa (Bedmar et al., 1984; Behki et a l . , 1985). This observation may be related to observed variability in expression of Hup genes in relation to the host plant (Bedmar et al., 1984).

16

R. K. PRAKASH AND ALAN G. ATHERLY

V. Plasmid-Genome Rearrangements The relationship between the chromosome and plasmids present in Rhizobium strains is of interest because of the large size of the plasmids. The DNA-DNA homology studies from different Rhizobium species (see Section I1,B) and nifand nod DNA hybridization (see Sections IV,A and B) indicated that in some strains the DNA sequences are conserved on a plasmid and in other strains it is on the chromosomal DNA. Studies on the organization of nifand nod sequences in R. fredii (Masterson et al., 1985) showed that although the nif and nod hybridization patterns to restriction endonuclease cut DNA were identical within these species, in one strain (USDA194) nifand nod sequences are on the chromosomal DNA. Further, in strain USDA194, the Sym plasmid DNA sequences present in other R. fledii strains are conserved in the chromosomal DNA, suggesting a plasmid-chromosome DNA rearrangement in R. fredii strains. Recently, Cantrell et al. (1982) examined several Hup+ and Hup- strains of B. japonicum for plasmid content. They discovered two plasmids in three spontaneous, nonrevertable hydrogenase mutants (Hup- ) plasmids. The parent strain SR did not contain isolatable plasmids. The authors concluded from these findings that either a large, Hup- -encoding megaplasmid has rearranged to give rise to two smaller plasmids or the two plasmids were generated from the bacterial chromosome. In either event, a structural rearrangement of the Rhizobium genome is very likely. Further evidence that the plasmid and chromosome of B. japonicum strains can undergo rearrangements was provided by Berry and Atherly (1984). They observed that after introduction of a P-group plasmid (RP1) by spheroplast transformation, the RPl DNA was found to be integrated into the chromosome, but, simultaneously an equal-sized piece of chromosomal DNA transposed into a large indigenous plasmid, producing an even larger plasmid. Thus, the very large plasmid of Rhizobium may possess episome-like behavior. This conclusion seems even more likely in light of the finding that B. japonicum possesses at least 18 copies of two insertion-like sequences (Kaluza et al., 1985). Recently, rearrangement of nif genes during heterocyst differentiation in the Cyanobacterium anabaenu has been reported (Golden et al., 1985). But no such DNA rearrangement was observed during transition of Rhizobium to nitrogenfixing bacteriods (Scott et al., 1984; R. K. Prakash and A. G. Atherly, unpublished).

VI. Relationship between Rhizobium and Agrobacterium Plasmids Agrobucterium and Rhizobium both belong to the family Rhizobiaceae. Agrobacteria are closely related to fast-growing rhizobia but not to slow-growing

PLASMIDS OF RHIZOBIUM

17

rhizobia (DeLey, 1968; Gibbins and Gregory, 1972; Heberlein et al., 1967). As in fast-growingRhizobium species, where the Syrn plasmid controls the symbiotic functions, the tumor-inducing ability of A . tumefaciens is controlled by a large plasmid designated as the Ti plasmid (see review by Nester and Kosuge, 1981). The high level of genetic relatedness between Rhizobium and Agrobacterium has been shown in several reports. Hooykaas et al. (1982~)compared chromosomal linkage maps of Rhizobium and Agrobacterium and found a high degree of similarity. Prakash and Schilperoort (1982) demonstrated homologous regions in Rhizobium Syrn plasmids and the Agrobacterium Ti plasmid. Hadley and Szalay ( 1982) showed that T-DNA sequences of Agrobacterium are present in diverse Rhizobium species. The implications of these observations are not clear at present. But, studies with R. fredii plasmid DNA indicate that the observed homology of the Syrn plasmid with 7'-DNA and vir-DNA region of A. tumefaciens is due to the presence of bacterial insertion IS-66-like sequences (Ramakrishnan et al., 1986). Insertion elements have already been described in Rhizobium lupini (Preifer et al., 1981), R . meliloti (Ruvkun et al., 1982b), andB. japonicum (Kaluza et al., 1985).

VII. Restriction Endonuclease Maps Restriction endonuclease maps of plasmids facilitate the identification and genetic characterization of plasmid-borne genes and consequently aids in the genetic manipulation of plasmid genes. Strategies for constructing a physical map of a large plasmid are varied but usually depend upon construction of a plasmid DNA clone bank using a vector that will yield large DNA fragments so as to facilitate construction of overlapping sequences. Prakash et al. (1982b) were the first to establish a restriction fragment map of the symbiotic plasmid (pRle1001a) from R. leguminosarum strain 1001. This 150 MDa plasmid was mapped by hybridization of individual HpaI restriction fragments to blotted SmaI and KpnI digestions of the plasmid. Regions homologous to nifstructural genes, Syrn plasmid DNA of R. trifolii, and Ti plasmid DNA of A . tumefaciens were mapped. A large region around the nifstructural genes was found to be highly conserved in these Rhizobium species. Other regions that are common to both the Syrn plasmid of R. leguminosarum and R . rrifolii were also conserved in octopine and nopaline Ti plasmids of A . tumefaciens. The regions that are transcribed in free-living bacteria and bacteriods were also localized on a restriction endonuclease map of the plasmid pR le lo0 la (Prakash et al., 1982a). One region that is actively transcribed in nitrogen fixation bacteroids includes the regions homologous to nifstructural genes and the region that has homology to the Syrn plasmid of R. trifolii. Similarly Huguet et al. (1983) prepared a physical map of a 150 MDa nonsymbiotic plasmid (pRme4la) from R. meliloti and found extensive regions of homology with both octopine and nopaline Ti plasmids and, surpris-

18

R . K. PRAKASH AND ALAN G. ATHERLY

ingly, less homology with R . meliloti plasmid DNA from strains of various geographical origins. Because of the great difficulty in constructing physical maps of very large plasmids (greater than 150 MDa) no megaplasmid has been totally mapped. However, partial maps have been constructed using R' plasmids derived from the megaplasmid of R . meliloti 41 (pRme4lb) (Kondorosi et a f . , 1983, 1984; Banfalvi et al., 1981b; Rosenberg er al., 1981) and strain 201 (Batut e? al., 1984; David el al., 1984). David et al. (1984) found extensive regions of plasmid gene expression (about 100 kb) from a 285 kb R' fragment and as well two regions (about 20 kb) that are expressed only in vegetatively growing cells. Using restriction endonuclease mapping of segments of plasmid DNA, several laboratories found a close linkage of nod and nif genes in several fast-growing Rhizobium species. In R. meliloti, mutations in nod regions mapped withir. about 25 and 13 kb downstream from the nif KDH operon (Kondorosi et a l . , 1984; Buikema er a f . , 1983). In R. trifolii, nod genes are located some 16 kb from the nifstructural genes KDH (Schofield et al., 1983). In R. leguminosarurn a 45 kb region of DNA has been shown to carry two clusters of genes encoding nifgenes separated by a cluster of nod genes (Downie er al., 1983b).

VIII. Perspectives The amount of genetic information now known about plasmid DNA is incredibly small in relation to the immense information present. An estimated molecular weight of 1 X lo9 for one megaplasmid (Burkhardt and Burkhardt, 1984) and the observation that two such megaplasmids exist in R . meliloti (Kondorosi er af., 1984) indicate that, at least in some Rhizobium strains, the amount of plasmid DNA may be equivalent to the total chromosomal DNA of E. cofi (2.2 X lo9 Da). Very useful genes for the survival of bacteria are frequently present on plasmids; thus it is not surprising that symbiotic functions and nitrogen fixation genes are present on the plasmids of a great many Rhizobium strains. But, by no means have all these genes been identified and their functions determined. Prospects for the future are bright in identifying symbiotic and nitrogen fixation genes due to the advent of many very useful techniques in molecular genetics. In this respect two very useful procedures must include lacZ fusions, including the use of Mu-d lac, and mRNA hybridization to cloned fragments to identify functional genes. Tn5 insertional inactivation of tentatively identified genes, as well as random mutagenesis with Tn5, is extremely helpful in identifying gene functions. At present, the genes involved in competitive ability are of great interest and preliminary evidence suggests they may be on plasmids (DeJong er al., 1982; Olson et al., 1985). Also, the involvement of carbohydrates is strongly implicated in some steps of the symbiotic process, but few genes or mutants have

PLASMIDS OF RHlZOBIUM

19

been identified. Sequences are found on plasmids that are highly conserved (Prakash et al., 1981 ; Masterson et al., 1985; Watson and Scholfield, 1985) and there is little evidence to indicate what functions these sequences play in the symbiotic process, or other necessary functions. The hope for a complete understanding of the symbiotic process is that it will eventually lead to intelligent strain construction. Changes in Rhizobium strains that may be useful include altering the host-specificity genes to broaden the host range, increasing the competitive ability, and optimizing nitrogen fixation efficiency or creating Rhizobium strains specific for particular cultivars, soils, or climatic conditions. It also seems very likely that knowledge of the regulation of genes in the nodule tissue will aid in the construction of strains that export new products to the plant, other than nitrogen. Genes for the synthesis of insect hormones, plant hormones, fungicides, or other chemicals can be engineered into Rhizobium strains with expression regulated during the bacteroid form and the products exported to the plant. In this respect Rhizobium may become more useful to the farmer than just a nitrogen source.

ACKNOWLEDGMENTS We wish to thank Dr. S. Shantharam for his critical reading of this manuscript and Miss Ruth Richman for typing the manuscript.

REFERENCES Albrecht, S. L., Maier, R. J., Hanus, F. J., Russell, S. A,, Emerich, D. W., and Evans, H. J. (1979). Science 203, 1255. Appelbaum, E. R., McLoughlin, T. J . , O’Connell, M., and Chartrain, N. (1985). J. Bacteriol. 163, 385. Atherly, A. G., Prakash, R. K., Masterson, R. V., Du Teau, N. B., and Engwall, K. S. (1985). Proc. World Soybean Conf., 3rd. pp. 229-300. Bagdasarian, M.. Lurz, R.,Ruckert, B., Franklin, F. C. H., Bagdasarian, M. M., Frey, J., and Timmis, K. N. (1981). Gene 16, 237. Bal, A. K., Shantharam, S., and Ratnam, S . (1978). J. Bacteriol. 133, 1393. Banfalvi, Z., Randhawa, G. S., Kondorosi, E., Kiss, A , , and Kondorosi, A. (1981a). Mol. Gen. Genet. 189, 129. Banfalvi, Z., Sakanyan, V., Kohcz, C., Kiss, A,, Dusha, J., and Kondorosi, A. (1981b). Mol. Gen. Genet. 184, 318. Banfalvi, Z . , Kondorosi, E., and Kondorosi, A. (1985). Plasmid 13, 129. Batut, J . , Gherardi, M., Terzaghi. E.. and Huguet, T. (1984). In “Advances in Nitrogen Fixation Research” (C. Veeger and W. E. Newton, eds.), p. 672. Nijhoff, The Hague. Bauer, W. D. (1981). Annu. Rev. Plant Physiol. 32, 407. Bedmar, E. J . , Brewin, N. J., and Phillips, D. A. (1984). Appl. Environ. Microbiol. 47, 876. Behki, R . M., Selvaraj, G., and Iyer, V. N. (1985). Arch. Microbiol. 140, 352.

20

R. K. PRAKASH AND ALAN G. ATHERLY

Bellofatto, V., Shapiro, L., and Hodgeson, D. A. (1984). Proc. Natl. Acad. Sci. U.S.A. 81, 1035. Berg, D. F., Weiss, A., and Crossland, L. (1980). J. Bacteriol. 142, 439. Beringer, I. E., Beynon, I. L., Buchanan-Wollaston,A. V., and Johnston, A. W. B. (1978). Nature (London) 276, 633. Berry, J. 0..and Atherly, A. 0.(1984). J . Bacteriol. 157, 218. Better, M., and Helinski, D. H. (1983). J . Bacteriol. 155, 311. Beynon, J. L., Beringer, J. E.. and Johnston, A. W. B. (1980). J . Gen. Microbiol. 120, 421. Bishop, P. E., D m o , F. B., Appelbaum, E. R., Maier, R. J., and Brill, W. J. (1977). Science 198, 938.

Bohlool, B. B., and Schmidt, E. L. (1974). Science 185, 269. Brewin, N. J., Beringer, J. E., Buchana-Wollaston, A. V., Johnston, A. W. B., and Hirsch, P. R. (1980a). J . Gen. Microbiol. 116, 261. Brewin, N. J., Beringer, J. E., and Johnston, A. W. B. (1980b). J . Gen. Microbiol. 120, 413. Brewin, N. J., DeJong, T. M., Phillips, D. A., and Johnston, A. W. B. (1980~).Nature (London) 288, 71. Brewin, N. J., Beynon, J. L., and Johnston, A. W. B. (1981). In “Genetic Engineering of Symbiotic Nitrogen Fixation” (J. M. Lyon, R. C. Valentine, D. A. Phillips, D. W. Rains, and R. C. Huffaker, eds.), pp. 65-77. Plenum, New York. Broughton, W. J., Heycke, N., Meyer, Z. A. H., and Pankhurst, C. E. (1984). Proc. Nud. Acad. Sci. U.S.A. 81, 3039. Buchanan-Wollaston, V. (1979). J. Gen. Microbiol. 112, 135. Buikema, W. J., Long, S. R., Brown, S . E.,Van Den Bos, R. C., Earl, C., and Ausubel, F. M. (1983). J . Mol. Appl. Genet. 2, 249. Burkhardt, B., and Burkhardt, H. J. (1984). J . Mol. Biol. 175, 213. Cannon, F. C., Reidel, G. E., and Ausubel, F. M. (1979). Mol. Gen. Genet. Cantrell, M. A., Hickok, R. E.. and Evans, H. J. (1982). Arch. Microbiol. 131, 102. Casadaban, M. J., Chou, J., and Cohen, S. N. (1980). J . Bacteriol. 143, 971. Casadeus, J., and Olivaries, J. (1979). J. Bacteriol. 139, 316. Casse, F., Boucher, C., Julliot, J. S . , Michel, M., and Denarie, 3. (1979). J. Gen. Microbiol. 113, 229.

Castillo, B. A., Olfson, P., and Casadaban, M. J. (1984). J. Bacteriol. 158, 488. Chakrabarti, S. K., Mishra, A. K., and Charkrabartty, P. K. (1983). Cum. Sci. 52, 779. Chakravorty, A. K., Zurkowski, W., Shine, J., and Rolfe, B. G. (1982). J. Mol. Appl. Genet. 1, 585. Christensen, A. H., and Schubert, K. R. (1983). J. Bacreriol. 156, 592. Corbin, D., Ditta, G., and Helinski, D. R. (1982). J . Bacteriol. 149, 221. Corbin, D., Barran, L., and Ditta, G. (1983). Proc. Natl. Acud. Sci. U.S.A. 80, 3005. Currier, T. C., and Nester, E. W. (1976). Anal. Biochem. 76, 431, David, M., Vielma, M., and Julliot, J. S. (1983). FEMS Microbiol. Lett. 16, 335. David, M., Domerque, D., and Kahn, D. (1984). In “Advances in Nitrogen Fixation Research” (C. Veeger and W. E. Newton, eds.), p. 676. Nijhoff, The Hague. Dazzo, F. B.,and Gardiol, A. E. (1984). In “Genes Involved in Microbial-Plant Interactions” (D. p. Verma and T. Holm, eds.), pp. 1-23. Springer-Verlag, Berlin and New York. D ~ z o F. , B., and Truchet, G. L. (1983). J. Membr. Biol. 73, 1. DeLey, J. (1968). Annu. Rev. Phyroparhol. 6, 63. DeLey, S., and Russel, A. (1965). J . Gen. Microbiol. 41, 85. DeJong, T. M., Brewin, N. J., Johnston, A. W. B., and Phillips, D. A. (1982). J . Gen. Microbiol. 128, 1829. Denarie, J., Boistard, P., Casse-Delbart, F., Atherly, A. G., Berry, J. O., and Russel, P. (1981). Int. Rev. Cyrol. Suppl. 13, 225.

PLASMIDS OF RHIZOBIUM

21

Ditta, G., Stanfield, S., Corbin, D., andHelinski, D. R. (1980). Proc. Natl. Acad. Sci. U S A . 77, 7347. Ditta, G., Schmidhauser, T., Yakobson, E., Lu, P., Liang, X. W.. Finley, D. R . , Ginney, D., and Helinski, D. R. (1985). Plasmid 13, 149. Djordjevic, M. A., Zurkowski, W., and Rolfe, 8. G. (1982). J . Bacteriol. 151, 560. Djordjevic, M. A., Zurkowski, W., Shine, J., and Rolfe, B. G. (1983). J . Bacreriol. 156, 1035. Djordjevic, M. A., Scholfield, P. R., Ridge, R. W., Morrison, N. A., Bassam, B. J.. Plazinski, J., Watson, J. M., and Rolfe, B. G. (1985). Plant Mol. Biol. 4, 147. Downie, J . A., Ma, Q. S., Knight, C. D., Hombrecher. G., and Johnston, A. W. B. (1983a). EMBO J . 2, 947. Downie, J . A., Hombrecher, G . , Ma, Q. S., Knight, C., Wells, B., and Johnston, A. W. B. (1983b). Mol. Gen. Genet. 190, 359. Dunican, L. K., and Cannon, F. (1971). Plant Soil 7, 73. Eckhardt, T. (1978). Plusmid 1, 584. Engwall, K . S., and Atherly, A. G. (1986). Plant Mol. Biol. 6, 41. Figureski, D., and Helinski, D. R. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 1648. Finan, T. M., Hartwieg, E., Lemieux, K., Bergman, K., Walker, G. C., and Signer, E. (1984). J. Bacteriol. 159, 120. Finan. T. M., deVos, G. F., and Signer, E. (1985a) In “Nitrogen Fixation Research Progress” (H. J. Evans, P. J. Bottornley, and W. E. Newton, eds.). p. 135. Nijhoff, The Hague. Finan, T. M., Hirsch, A. M., Leigh, J. A., Johansen, E., Kuldau, G. A., Deegan, S., Walker, G., and Signer, E. (1985b). Cell 40, 869. Fisher, R. F., Tu, J. K., and Long, S. R. (1985). Appl. Environ. Microbiol. 49, 1432. Forrai, T., Vincze, E., Banfalvi, Z., Kiss, G. B., Randhawa, G. S., and Kondorosi, A. (1983). J. Bacteriol. 153, 635. Friedman, A. M., Long, S. R., Brown, S. E., Buikema, W. J., and Ausubel, F. M. (1982). Gene 18, 289. Fuhrman, M., Fischer, H.M., and Hennecke, H. (1985). M o l . Gen. Genet. 199, 315. Gibbins, A. M., and Gregory, K. F. (1972). J. Bacteriol. 111, 129. Glenn, A. R., and Dilworth, M. J. (1981). Arch. Microbiol. 129, 233. Golden, J. W., Robinson, S. J., and Haselkorn, R. (1985). Nature (London) 314, 419. Graham, T. L. (1981). Int. Rev. Cytol. Suppl. 13, 127. Gross, D. C., Vidaver, A. K., and Klucas, R. V. (1979). J. Gen. Microbiol. 114, 619. Guerry, O., Embden, J., and Falkow, S. (1974). J. Bacreriol. 117, 619. Hadley, R. G., and Szalay, A. A. (1982). Mol. Gen. Gener. 188, 361. Hahn, M., and Hennecke, H. (1984). Mol. Gen. Genet. 193, 46. Haugland, R., and Verma, D. P. S. (1981). J . Mol. Appl. Genet. 1, 205. Heberlein, G. T., Deley, J., and Tijtga, R. (1967). J . Bacteriol. 94, 116. Hennecke, H. (1981). Nature (London) 291, 354. Heron, D. S., and Pueppke, S. G. (1984). J . Bacreriol. 160, 1061. Higashi, S. (1967). J. Gen. Appl. Microbiol. 13, 391. Hirsch, A. M . , Wilson, K. J., Jonee, J. D., Bang, M., Walker, V. V., and Ausubel, F. M. (1981). J . Bacteriol. 158, 1133. Hirsch, A. M., Long, S. R., Bang, M., Haskins, N., and Ausubel, F. M. (1982). J. Bacteriol. 151, 411. Hirsch, A. M., Drake, D., Jocobs, T. N., and Long, S. R. (1985). J. Bacteriol. 161, 223. Hirsch, P. R., Van Montagu, M., Johnston, A. W. B., Brewin, N. J., and Schell, J. (1980). J. Gen. Microbiol. 120, 403. Hollis, A. B., Kloos, W. E., and Elkan, G. H. (1981). J . Gen. Microbiol. 123, 215. Hombrecher, G., Brewin, N. J., and Johnston, A. W. B. (1981a). Mol. Gen. Genet. 182, 133.

22

R. K. PRAKASH AND ALAN G. ATHERLY

Hombrecher, G., Got, R., Dibb, N. J . , Downie, J. A., and Johnston, A. W. B. (1981b). Mol. Gen. Genet. 182, 132. Hooykaas, P. J. J., Van Brussel, A. A. N.,Den dulk-Ras, H., Van Slogteren, G. M. S., and Schilperoort, R. A. (1981). Nature (London) 291, 351. Hooykaas, P. J. J., Den dulk-Ras, H., and Schilperoort, R. A. (1982a). Plasmid 8, 94. Hooykaas, P. J. J . , Snijdewint, F. G. M., and Schilperoort, R. A. (1982b). Plasmid 8, 73. Hooykaas, P. J. J., Peerbolte, R., Renensburg,-Tuink, A. J. G., De Vries, P., and Schilperoort, R. A. (1982~).Mol. Gen. Genet. 188, 12. Huguet, T., Rosenberg, C., Casse-de Bart, F., De Lajudie, P., Jouanin, L.. Batut, J., Boistard, P., Julliot, J. S., and Denarie, J . (1983). In “Molecular Genetics of Bacteria-Plant Interaction” (A. Puhler, ed.), pp. 35-45. Springer-Verlag, Berlin and New York. Jacobs, T. W., Egelhoff, T. T., and Long, S. R. (1985). J . Bacteriol. 162, 461. Jarvis, B. D. W., Dick, A. G., and Greenwood, R. M. (1980). Int. J. Syst. Bucteriol. 30, 42. Johansen, E., Finan, T. M.. Gefter, M. L., and Signer, E. R. (1984). J. Bacreriol. 160, 454. Johnston, A. W. B., Beynon, J. L., Buchanan-Wollaston, A. V., Setchell, S. M., Hirsch, P. R., and Beringer, J. E. (1978). Nature (London) 276, 635. Jordan, D. C. (1982). Int. J . Syst. Bacteriol32, 136. Jouanin, L., Delajudie, P., Bazetoux, S., and Huguet, T. (1981). Mol. Gen. Genet. 182, 189. Julliot, J. S., Dusha, I., Renalier, M. H., Terzaghi, B., Gamerone, A. M., and Boistard, P. (1984). Mol. Gen. Genet. 193, 17. Kahn, M. L., and Timblin, C. R . (1984). J . Bacreriol. 158, 1070. Kaluza, K., Hahn, M., and Hennecke, H. (1985). J. Bacteriol. 162, 5535. Knauf, V., and Nester, E. W. (1982). Plasmid 8, 45. Kondorosi. A., Kondorosi, E., Pankhurst, C. E., Broughton, W. J., and Banfalvi, 2. (1982). Mol. Gen. Genet. 188, 433. Kondorosi, A., Kondorosi, E., Banfolvi, 2.. Broughton, W. J., Pankhurst, C. E., Randhawa, G. S., Wong, C. H., and Schell, J. (1983). In “Molecular Genetics of Bacteria-Plant Interaction” (A. Puhler, ed.), pp. 55-63. Springer-Verlag, Berlin and New York. Kondorosi, E., Banfalvi, 2.. and Kondorosi, A. (1984). Mol. Gen. Genet. 193, 445. Kowalczuk, E., Skorupska, A., and Lorkeiwicz, A. (1981). Mol. Gen. Genet. 183, 388. Krol, A. J. M., Hontelez, J. G. J . , Van Den Bos, R. C . , and Van Kammen, A. (1980).Nucleic Acids Res. 8,4337. Krol, A. J . M., Hontelez, J. G. J., and Van Kammen, A. (1982). J. Gen. Microbiol. 128, 1839. Kun, W. G. W., and La Rue, R. A. (1975). Nature (London) 256,407. Lamb, J. W., Hombrecher, G., and Johnston, A. W. B. (1982). Mol. Gen. Genet. 186, 449. Ledeboer, A. M., Krol, A. J. M., Dons, J. J. M., Spier, R., Schilperoort, R. A., Zaenen, I., Van Larkeke, W., and Schell, J. (1976). Nucleic Acids Res. 3, 419. Leigh, J. A., Signer, E. R., and Walker, G. C. (1985). Proc. Nutl. Acad. Sci. U.S.A. 82, 6231. Ljunggren, H., and Fahraeus, G. (1961). J. Gen. Microbiol. 26, 521. Long, S. R., Meade, H. M.. Brown, S. E., and Ausubel, F. M. (1981).In “Genetic Engineering in the Plant Sciences” (N. J. Panopoulos, ed.), pp. 129-145. Praeger, New York. Long, S. R., Buikema, W. E., and Ausubel, F. M. (1982). Nature (London) 298, 485. McComb, I. A., Elliot, J., and Dilworth, M. J. (1975). Nature (London) 256, 409. Martin, M. O., and Long, S. R. (1984). J . Bacteriol. 159, 125. Marinez-DeDrets, G., and Arias, A. (1972). J . Bacreriol. 109, 467. Masterson, R. V., Russell, R. P., and Atherly, A. G. (1982). J . Bacteriol. 152, 928. Masterson, R. V., Prakash, R. K., and Atherly, A. G. (1985). J . Bacteriol. 163, 21. Mathis, I. N., Barbour, W. H., and Elkan, G. H. (1985). Appl. Environ. Microbiol. 49, 1385. Meade, H. M., Long, S . R., Ruvkun, G. B., Brown, S. E., and Ausubel, F. M. (1982). J . Bacreriol. 149, 114.

PLASMIDS OF RHIZOBIUM

23

Morrison, N. A., Cen, Y. H., Trinick, M. J., Shine, J., and Rolfe, B. G. (1983). J . Bacteriol. 153, 527. Nester, E. W., and Kosuge, T. (1981). Annu. Rev. Microbiol. 35, 531. Noel, K. D., Sanchez, A . , Fernandez, L., Leeman, J., and Cevallos, M. A. (1984). J . Bacteriol. 158, 148. Nuti, M. P., Ledboer, A. M., Lepidi, A. A., and Schilperoort, R. A. (1977). 1. Gen. Microbiol. 100, 241. Nuti, M. P., Lepidi, A. A., Prakash, R. K., Schilperoort, R. A,, and Cannon, F. C. (1979). Nature (London) 282, 533. Nuti, M. P., Lepidi, A. H., Prakash, R. K., Hooykaas, P. J. J . , and Schilperoort, R. A. (1982). In “Molecular Biology of Plant Tumors’’ ( G . Kahl and J. S. Schell, eds.), pp. 561-589. Academic Press, New York. Olson, E., Sadowsky, M., and Verma, D. P. (1985). Biotechnology 3, 143. Pagan, J. D., Child, J . J., Scowcroft, W. R., and Gibson, A. H. (1975). Nature (London) 256,406. Palomares, A., Montoya, E., and Olivares, J. (1978). Microbios 21, 33. Palomares, A., Montoya, E., and Olivares, J . (1979). Microbios 22, 7. Plazinski, J., and Rolfe, B. G. (1985). J. Bacteriol. 162, 1261. Prakash, R. K., and Atherly, A. G. (1984). J. Bacteriol. 160, 785. Prakash, R. K.. and Schilperoort, R. A. (1982). J . Bacferiol. 149, 1129. Prakash, R. K., Hooykaas, P. J. J., Ledeboer, A. M., Kijne, J. W., Schilperoort, R. A,, Nuti, M. P., Lepidi, A. A., Casse, F., Boucher, C., Julliot, J. S . , and Denarie, J. (1980). In “Nitrogen Fixation 11” (W. E. Newton and W. H. Orme-Johnson, eds.), pp. 89-164. Univ. Park Press, Baltimore, Maryland. Prakash, R. K., Schilperoort, R. A,, and Nuti, M. P. (1981). J . Bacteriol. 145, 1129. Prakash, R. K., Van Brussel, A. A. N., Quint, A,, Mennes, A. M., and Schilperoort, R. A. (1982a). Plasmid 7, 28 I. Prakash, R. K.. VanVeen, R. J. H.,and Schilperoort, R. A. (1982b). Plasmid 7 , 271. Prakash, R. K., DuTeau, N. D., and Atherly, A. G. (1986). Submitted. Refer, U. B., Burkardt, H. J., Klipp, W., and Puhler, A. (1981). Cold Spring Harbor Symp. Quant. Biol. 45, 87. Quinto, C., De La Vega, H., Flores, M., Femaudez, L., Ballado, T., Soberon, G., and Palacios, R. (1982). Nature (London) 299, 724. Quinto, C., De La Vega, H., Flores, M., Leemeans, J., Cevallos, M. A,, Pardo, M. A,, Azpiroz, R., Girard, M., Calva, E., and Palacios, R. (1985). Proc. Natl. Acad. Sci. U.S.A. 82, 1170. Ramakrishnan, M. S., Prakash, R. K., and Atherly, A. G. (1986). Submitted. Rolfe, B. G., Djordjevic, M., Scott, K. F., Hughes, J. E., Badenoch-Jones, J., Gresshoff, P. M., Chen, Y., Dudman, W. G., Zurkowski, W., and Shine, J. (1981). In “Current Perspectives in Nitrogen Fixation” (A. Gibson and A. Newton, eds.), pp. 142-145. Australian Academic Press, Canberra. Rolfe, B. G., Djordjevic, M. A., Morrison, N. A,, Plazinski, J., Bender, G. L., Ridge, R., Zurkowski, W., Tellan, J. T., Gresshoff, P. M., and Shine, J. (1983). In “Molecular Genetics of the Bacteria-Plant Interaction” (A. Puhler, ed.), pp. 188-203. Springer-Verlag, Berlin and New York. Ronson, C. W., and Scott, D. B. (1983). In “Molecular Genetics of Bacterial-PlantInteraction” (A. Puhler, ed.), pp. 177-187. Springer-Verlag. Berlin and New York. Rosenberg, C., Boistard, P., Denarie, J., and Casse-Delbart. (1981). Mol. Gen. Genet. 84, 326. Rossen, L., Johnston, A. W. B., and Downie, J. A. (1984). Nucleic Acids Res. 12, 9497. Ruiz-Sainz, J. E., Chandler. M. R.. Jimenez-Diaz, R., and Beringer, J. (1984). J . Appl. Bacteriol. 57, 309. Ruvkun, G. B.. and Ausubel, F. M. (1981). Nature (London) 289, 8 5 .

24

R. K. PRAKASH AND ALAN G. ATHERLY

Ruvkun, G. B., and Ausubel, F. M. (1980). Proc. Narl. Acad. Sci. U.S.A. 11, 191. Ruvkun, G. B., Sundaresan, V., and Ausubel, F. M. (1982a). Cell. Ruvkun, G. B., Long, S. R., Meade, H. M., Van Den Bos, R. C., and Ausubel, F. M. (1982b). J . Mol. Appl. Genef. 1, 405. Sadowsky, H. J., and Bohlool, B. B. (1983). Appl. Environ. Microbiol. 46, 906. Sadowsky, M. J., Keyser, H. H., and Bohlool, B. B. (1983). Inr. J . Sysr. Bacreriol. 33, 716. Sanders, R. E., Carlson, R. W., and Albersheim, P. (1978). Nature (London) 271, 240. Schetgens, T. M. P., Bakkeren, G., Van Dun, C., Hontelez, J. G. J., Van Den Bos, R. C., and Van Kammen, A. (1984). J. Mol. Appl. Genet. 2, 406. Schofield, P. R., Djordjevic, M. A.. Rolfe, B. G., Shine, J., and Watson, J. M. (1983). Mol. Gen. Genet. 192, 459. Schofield, P. R., Ridge, R. W., Rolfe, B. G., Shine, J., and Watson, J. M. (1984). PlantMol. Biol. 3, 3. Scholla, M. H., and Elkan, G. H. (1984). Inr. J. Sysr. Bacreriol. 34, 484. Scholla, M. H., Moorefield, J. H., and Elkan, G. H. (1984). Inr. J . Sysr. Eacreriol. 34, 382. Scott, D. B., and Ronson, C. W. (1982). J . Bacreriol. 151, 36. Scott, D. B., Court, C. B., Ronson, C. W., Scott, K. F., Watson, J. M., Schofield, P. R., and Shine, J. (1984). Arch. Microbiol. 139, 151. Selvaraj, G., and Iyer, V. N. (1983). J. Bacreriol. 156, 1292. Shah, K., Sousa, S., and Modi, V. V. (1981). Arch. Microbiol. 130, 262. Shah, K., Patel, C., and Modi, V. V. (1983). Can. J . Microbiol. 29, 33. Shantharam, S., and Iyer, V. N. (1986). In preparation. Shantharam, S., Gow, J. A., and Bal, A. K. (1980). Can. J . Microbiol. 26, 107. Sik, T., Hovath, J., and Chatterjee, S. (1980). Mol. Gen. Genet. 178, 51 1 . Silhavy, T. J . , Berman, M. L., and Enquist, L. W. (1984). I n “Experiments with Gene Fusions.” Cold Spring Harbor Press, Cold Spring Harbor, New York. Simon, R. (1984). Mol. Gen. Genet. 196, 413. Simon, R., Priefer, U., and Puhler, A. (1983). Biorechnology 1, 784. Skogen-Hagenson, M. J . , and Atherly, A. G. (1983). J. Bacreriol. 156, 937. Stowers, M. D., and Eaglesham, A. R. J. (1983). J. Gen. Microbiol. 129, 3651. Sundaresan, V., Ow, D. W., and Ausubel, F. M. (1983a). Proc. Narl. Acad. Sci. U.S.A. 80,4030. Sundaresan, V., Jones, J. D. G., Ow, D. W., and Ausubel, F. M. (1983b). Nature (London) 301, 728. Szeto, W. W., Zimmerman, J. L., Sundaresan, V., and Ausubel, F. M. (1984). Cell 36, 1035. Torok, I., Kondorosi, E., Stepkowski, T., and Konodorosi, A. (1984). NucleicAcids Res. 12,9509. Trinick, M. J. (1973). Nature (London) 244, 459. Truchet, G., Rosenberg, C., Vasse, J., Julliot, J. S., Camut, S., and Denarie, J. (1984). J . Bacreriol. 157, 134. Vandenbosch, K. A., Noel, K. D., Kaneko, Y.,and Newcomb, E. (1985). J . Bacreriol. 162,950. Van Vliet, F., Silva, B., Van Montagu, M., and Schell, J. (1978). Phsmid 1, 446. Watson, J. M., and Scholfield, P. R. (1985). Mol. Gen. Genet. 199, 279. Weinstock, G. M., Berman, M. L.. and Silhavy, T. J. (1983). I n “Expression of Cloned Genes in Prokaryotic and Eukaryotic Vectors” (T. S . Papas er al., eds.), p. 27. Elsevier, Amsterdam. Wong, C. H., Panhurst, C. E., Kondorosi, A., and Broughton, W. J. (1983). J. CeNEiol. 97,787. Yakobson, E. A., and Guiney, D. G. (1984). J. Bacreriol. 160, 451. Yelton, M. M.. Yang, S. S., Edie, S. A., and Lim, S. T. (1983). J. Gen. Microbiol. 129, 1537. Zurkowski, W. (1980). Microbios 7, 27. Zurkowski, W., and Lorkiewicz, Z. (1976). J. Bacreriol. 128, 481. Zurkowski, W., and Lorkiewicz, Z. (1978). Genet. Res. 32, 311. Zurkowski, W., and Lorkiewicz, Z. (1979). J . Bacferiol. 128, 481.

INTERNATIONAL KEVIEW OF CYTOLOGY. VOL I W

Mouse Mutants: Model Systems to Study Congenital Cataract AUDREYL. MUGGLETON-HARRIS MRC Experiment01 Embryology cind Teratology Unit, Medicul Research Council Laboratories. Carshalton, Surrey SM5 4EF, England

I. Introduction Congenital and early developmental cataracts are common ocular abnormalities and represent an important visual impairment in childhood; 10-38% of all blindness in children is caused by developmental cataracts. One of every 250 newborns (0.4%) has some form of congenital cataract; 26% of those children operated on for congenital cataract are able to attend school, and the majority of those have impaired vision. In the light of the statement that “In view of the inevitable operative risk, in other words, of anesthetic accident, post operative infection and surgical complications, congenital cataract surgery should be undertaken only after the most careful consideration” (Nelson, 1984), it would appear that an analysis of the genetic and phenotypic mechanisms underlying congenital and developmental cataractogenesis is a necessity. In a child with cataracts who is otherwise healthy, between 8.3 and 23% of cataracts are familial, autosomal dominant hereditary being the most frequent mode of inheritance (Nelson, 1984). The limited data on the causative mechanisms of human cataract are related to the fact that man is such a slow breeder, and detailed in vivo morphological classification of inherited cataract and family histories are not readily available. Also the surgical procedure for cataract removal involves fragmentation of the lens, and thus valuable material for histological, cellular, and biochemical analysis is destroyed. Therefore animal models are especially suited for a genetic analysis of cataractogenesis, because the litter size is large and they have a relatively short life-span. Animal Models. There are various animal models with which to study congenital cataract; the advantages of using mouse mutants are obvious: the genetics of the mouse are well researched and the breeding of large litters is comparatively easy. As stated earlier, autosomal dominant hereditary is the most frequent mode of inheritance of congenital cataract, therefore mutants which inherit their cataracts in this manner are specifically interesting. One such mutant is the Cuturuct Fruser (CutFr)mouse. The CatFr mouse develops a cataract prenatally and was first described as a dominant autosomal mutation known as 25 Cupyripht 0 1986 by Academic Prcm Inc. All right? uC reproduction in any t u r n reserved.

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AUDREY L. MUGGLETON-HARRIS

“shriveled” (Fraser and Schabtach, 1962). Mice begin to show lens deterioration between 10 and 14 days of intrauterine life. Initially the cell nuclei in the deep cortex become abnormally pyknotic; degeneration of cytoplasm and destruction of the lenticular nucleus follow. Two weeks postnatally, the anterior epithelia show unusual mitotic activity with the formation of multiple cell layers that infiltrate into the abnormal fibers of the anterior cortex. Cells at the equatorial region retain their ability to differentiate until complete hydration of the lens occurs after 1 year of age. lnhibition of elongated older cells takes place by the breakdown of the nuclear membrane (Gelatt and Das, 1984). The abnormalities associated with this congenital cataract can be appreciated by observing the normal structure of the lens and its development (McAvoy, 1981). The pathogenesis of another genetically malformed lens mutant, Elo, has indicated that the eye lens obsolescence is independent of the extraocular environment and that the y-crystallin synthesis in the lens of the Elo mutant and the normal lens was similar, thereby suggesting that the necrotic process in the lens fibers of the Elo is not due to defects in y-crystallin synthesis. Initial cytological changes in this mutant were the appearance of numerous lysosomal bodies, the destruction of mitochondria at the basal cytoplasm of the lens fibers, and nuclei with elevated perinuclear cistema (Oda et al., 1980; Watanabe et al., 1980). A systematic search for heritable electrophoretic variation among the lens crystallins to date yielded no useful markers for the mammalian crystallin genes. However, isoelectrofocusing of soluble lens proteins from 29 strains of mice revealed 3 electrophoretic phenotypes. Genetic experiments and molecular sieving studies demonstrated that the variation was encoded by three alleles at a locus, designated Len-Z, and that the affected protein was a y-crystallin. Len-/ was determined to be located on mouse chromosome 1. Based upon the nature of the genetic variant, it is plausible to consider Len-1 as the structural locus for one of several mouse y-crystallins. Two mutations affecting the mouse lens, vacuolated lens (VL), and eye lens obsolescence (Elo) have also been mapped to chromosome 1 (Skow, 1982). Certainly in the light of the y-crystallin genes being closely linked to cataractous mutants (Skow, 1982), there is the possibility of some regulation mutation affecting gene expression of the lens crystallins in the C U P mouse. The fact that the present C U P mouse is not on the A I J background as first reported a number of years ago and appears not to be on several alternative albino inbred strains which we have studied could indicate that it is now a “unique” strain of cataractous mouse (Muggleton-Harris et a l . , 1986). A mutation which arose spontaneously in a stock of mice homozygous for the Robertsonian translocation (Rb) Ald is an albino, whose pink eyes are opaque. Examination of the eyes revealed the opacity in the lens, and it was present when the eyes first opened. Breeding showed the character to be inherited as an autosomal dominant, and it was given the name and symbol lens opacity, Lop. Linkage tests of Lop with markers on various chromosomes, to map the locus

MOUSE MUTANTS: MODEL SYSTEMS FOR CATARACTS

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and to exclude allelism with other genes for cataract known to map in other positions, confirm that there is no allelism with several already mapped genes affecting the eye and that there is a locus for a dominant cataract gene of chromosome 10 for Lop. The Lop occupies almost an end position on the chromosome and thus provides a useful marker in linkage studies (Lyon et al., 1981; West and Fisher, 1985).

11. The Search for Chromosomes Associated with the Mutants Cataract

Using available genetic markers, various loci on the chromosomes of the mouse may be eliminated as being associated with the CutFrcongenital cataract. By breeding and cross-breeding the inbred CarFr mouse with the appropriate mice which carry specific genetic markers, the chromosome carrying the C U P gene can be determined by a process of elimination. Once a specific marker is linked to the CatFrmouse, and if the chromosome on which the marker is carried can be identified, then the chromosome with the CatFrgene will also be known. Already a number of chromosomes (specific) are known not to carry the CatFr mutation. Details of the previous work undertaken by Fraser and Schabtach (1962) and the chromosomes mapped by ourselves are given in Table I. The markers that are available for mapping the loci for the CarFrmouse are indicated in Table I. We have crossed inbred and cataractous strains to establish dominance and then backcrossed to establish genetic segregation. The backcross progeny are then typed for all relevant marks (i.e., those at which the parental strains differ). Fifty animals are studied for initial matings, and once provisional evidence indicates a positive, then the numbers will be increased to 200-300 animals. The CutFrmutant has been partially mapped. Previous studies by Fraser and Schabtach (1962) used the genes a, b, c, N, and Sex; we have covered those with an X on the marker list. A map of the chromosomes already mapped at specific loci is now available (Fig. 1). Kosambi’s function was used to convert the frequency of crossing over to physical chromosomal distance (Robinson, 1971). There are some obvious chromosomes yet to cover: chromosome 1 (with which other lens abnormalities are associated) and chromosome 10 (Lop lens defect). 111. Lens Crystallins Specialized proteins called crystallins are associated with the lens structure (Papaconstantinou, 1967, 1967). The synthesis of these proteins in the lens is ‘The CarFr and Lop lens abnormalities are linked on chromosome 10 and are probably allelic. This gene location has been recently confirmed by Muggleton-Hams er al. (1986).

28

AUDREY L. MUGGLETON-HARRIS TABLE I MARKERS FOR MAPPING THE LOCI Locus Idh-1 A0x-1

Marker

I I

Biochemical Biochemical Biochemical Coat color Biochemical Biochemical Biochemical Biochemical Biochemical Coat color Biochemical Biochemical Coat color Biochemical Biochemical Coat color Biochemical Biochemical Biochemical Coat color Biochemical Coat texture Limb mutant Biochemical Biochemical Coat color Biochemical Biochemical Limb mutant

Pep-3

1

2 3 3 3 4 4

Gpd-1 Pgm-2

B Pgm-1

Ldr-1 Miwh

Gpi-I Hbb C

Es-1 Mol-1 Tcf d Es-3 Re Xt

Es-10 Gpr-I

N Ce-2 PgK-2 bm Sex

CUf“ MOUSE

Chromosome

A Adh-3 Cur-2 Sep-1

FOR THE

4 5 6 6 7 7 7 8 9 9

9 I1 II 13 14

15 15 17 17 19 20

Sex

Mapped

X X X

X X X X

well documented for development, differentiation, aging, and cataract (Bloemendal, 1977, 1981; Hoenders and Bloemendal, 1983; Harding and Dilley, 1976; Piatigorsky, 1981). The initiation of crystallin synthesis in the mouse does not require proper development of the lens (Zwaan and Williams, 1968; Zwaan, 1975). Altered patterns of crystallin synthesis and aggregation of lens proteins have been reported for aging and cataractous lenses (Mostafapor and Reddy, 1980; Genis-Galvez et al., 1968; Harding, 1981). The eye lens of the CatFr mouse contains reduced amounts of y-crystallins (Garber et al., 1984), and, in initial studies, the presence of “abnormal” proteins was found in CatFrlenses, but using more sensitive methods, the authors found that traces of these “abnormal” proteins are in fact present in normal lenses (Garber et al., 1985). The

1

2

FIG.I .

3

4

5

6

7

8

9

10

11

12

13

Map of chromosomes covered for specific loci for the Curfr mutant. (---)

cr a / . ( 1986).

14

15

16

17

18

19

X

Fraser and Schabtach (1962): (- - - -) Muggleton-Harris

30

AUDREY L. MUGGLETON-HARRIS

murine a-crystallin-related proteins were not present in cultures of normal or mutant CatFr lenses, nor were they present in the in vitro translation products directed by RNA extracted from such lenses (Mostafapor and Reddy, 1980; Harding, 198 1). Column chromatography of total soluble lens proteins from various cataractous mutants has shown increased amounts of a-crystallin aggregated in the lenses. However, each of these mutations has a distinct etiology, making it likely that the formation of a-crystallin derivatives and high-molecular-weight aggregates is an indirect effect of the mutations. In each case, some primary cataractogenic defect presumably initiates a sequence of events leading to the development of the cataract (Genis-Galvez et al., 1968). One or more of these events could be responsible for the changes noted in crystallin synthesis, structure, or protein aggregation. Therefore the primary genetic mutation which results in the initiation of the cataract is an area of study which needs clarification. It is known that the mRNA for a-crystallin synthesis is present in nearly normal amounts and in translatable form in lenses of CatFrmice at birth (Garbor et al., 1985); and yet, as stated previously, the lens deterioration associated with the CutFr mouse is present between 10 and 14 days of the embryo’s intrauterine life. The observation that the loss of the y-crystallin mRNA correlates in time with the destruction of the lens fiber cells in the CatFr mouse demonstrates the need to study the primary genetic defect.

IV. Cellular Studies on Lens Epithelial Cells The lens epithelial cells from normal and cataractous ( C U P )mice have been cloned and characterized for morphology, replication, and growth patterns. The C U P cells undergo fewer replications than the normal cells and the growth pattern is also different (Muggleton-Harris et al., 1981). One observation made on the cultured lens epithelial cells which was in conflict with a number of previous studies (Eguchi and Kodarna, 1979; Russell et al., 1977; Okada et af., 1971) was that the mouse lens epithelial cells did not undergo differentiation to fibers and form lentoid bodies in vitro. Only if the cells were left undisturbed with cellular or lenticular membrane/capsule present did those alterations take place. Cells in close proximity to one another will form plaques of cells (Muggleton-Harris et al., 198 1). The stages of differentiation in lens epithelial cells in culture has been well documented (Creighton et al., 1981). The cytoskeleton of cultured cells has been linked with elongation (Piatigorsky er al., 1972). Cell volume and nuclear size are altered with elongation in vivo (Hendrix and Zwaan, 1974) and in v i m ; the cytoskeletal structures generating intracellular forces and/or membrane interactions may also be important factors for cell elongation (Piatigorsky, 198 1; Ramaekers and Bloemendal, 1981). Lens epithelial cell elongation has been reported in the absence of microtubules (Beebe et al., 1979),

MOUSE MUTANTS: MODEL SYSTEMS FOR CATARACTS

31

and in rat lens epithelial cell cultures, the presence of neural retina-conditioned medium precipitated cell fiber differentiation and synthesis of y-crystallin (Campbell and McAvoy, 1984). The loss of regulation of cell replication and growth which is a major factor of congenital cataractogenesis may be reflected in a distortion of the microtubulemicrofilament connections to the cell membrane. Transformed fibroblasts show a different distribution of actin-containing material to their normal counterparts (Puck, 1977). Anti-actin serum stains stress fibers in the cultured cells; the morphological appearance of this major component of the cellular microfilament network has been shown to depend upon the cell’s physiological state and upon the presence of pathological changes (Ramaekers and Bloemendal, 1981; Puck, 1977). In cultured bovine MLE the intermediate-sized filaments of the vimentin type are associated with certain regions of the cytoplasm, and the number of bundles of intermediate filaments, with and without microtubules, is arranged almost perpendicular to the microfilament bundles (Ramaekers and Bloemendal, 1981). The epitheloid lens-forming cells grown in vitro as monolayers are connected by gap junctions, and the intermediate filaments occur in close proximity to the intercellular boundary. At the cell-to-cell boundary, terminal anchorage sites of microfilament bundles and a densely stained web of fine intermediate filamentous material appear at the site of membrane anchorage (Ramaekers and Bloemendal, 1981). Actin and tubulin immunofluorescence and y-crystallin synthesis have been reported to be related to stages of differentiation of lens epithelial cells in culture (Creighton et al., 1981). A tabulation of the various observations of this correlation was made with specific stages associated with the cells as they replicate and grow in virro. At stage 1 when the cells are rounded or cuboidal the actin immunofluorescence was diffuse and in globules (Creighton et al., 1976; Hamada and Okada, 1977; Mousa and Trevithick, 1977). When the cells are elongated (fibroblastic polygonal stellate), the actin immunofluorescence is associated with the microfilament fibers (Tamura, 1965; Okada et al., 1971; McDevitt and Yamada, 1969; Piatigorsky and Rothschild, 1972; Creighton et al., 1976; Hamada and Okada, 1977). Observations on cultured lens epithelial cells by Creighton et al. (1977) indicate that the nucleus of the cell moves to one side and the cytoplasm is filmy or lace-like, the actin immunofluorescence appeared fibrous and associated with the microfilaments at this stage (Van der Veen and Heyen, 1959; Tamura, 1965; Friedrich and Glaesser, 1971; Creighton et al., 1976; Hamada and Okada, 1977). At stage 4 the production of y-crystallin has been observed (Papaconstantinou, 1967; McDevitt and Yamada, 1969; Creighton et al., 1976) and there was no alteration in the distribution of actin observed. Stage 5 brought about the disintegration of the nucleus in the cells (Papaconstantinou, 1967; Mamo and Leinfelder, 1958), and at stage 6 the cells became a fibrous mass with globules and/or “lentoid bodies” forming (Mamo and Lein-

32

AUDREY L. MUGGLETON-HARRIS

felder, 1958; Okada et al., 1971; Creighton et al., 1976; Hamada and Okada, 1977; Russell et al., 1977). In cloned mouse lens epithelial cells which are kept isolated from degenerating cells and lens capsule material, cell elongation and the formation of “lentoid bodies” have not been seen (Muggleton-Harris et al., 1981). Conflicting reports on the presence of lens proteins in cultured lens epithelial cells appear to be related to species and culturing conditions. Cloned cells of chicken lens epithelium have formed differentiated colonies when conditioned medium was used, a gelatin layer encouraged clonal growth, and “lentoid bodies” were observed at high cell density (Okada et al., 197 1 , 1973). Growth control of lens epithelium cells by cell substratum and interactions of the cell with the substratum to induce cell shape and alterations in morphology highlight the extracellular influences on the cell’s morphology and behavior (Iwig and Glaesser, 1979a,b). The replication, growth, and morphology of the CatFr mouse lens epithelial cells in vitro have been well characterized (Muggleton-Harris et al., 1981). The distribution of lens proteins and actin microfilaments in these cells is not known, but preliminary studies appear to indicate that the CarFrcells do not synthesize the lens y-crystallins in v i m , whereas the noncataractous mouse lens epithelial cells do (unpublished results).

V. Manipulations of the Cataractous Phenotype Somatic cell hybrids and manipulative studies have been undertaken on the cultured CatFrmouse lens epithelial cells. The micromanipulation of the cells is achieved with the aid of deFonbrane micromanipulators (Lipman and Muggleton-Harris, 1982). The results from these experiments demonstrated that the two parental genotypes which constituted the hybrid had been retained by the clone of cells derived from that hybrid. The parental cells from a noncataractous mouse and the C U P mouse display a finite life-span. The cells of cataractous origin have a decreased number of population doubling levels compared to the capacity of the normal cells when replicated in v i m . Hybrid cells derived from individual cell fusions of these two types have a mode of replication similar to that of normal cells, thus indicating that the cataractous mouse lens epithelial cells have been modified by the addition of noncataractous lens epithelial cell components. Which components of the noncataractous lens epithelial cells had this effect is not known, but the system is available to analyze with the technique of nuclear transfer. This technique was developed and used to identify the components controlling the cellular aspects of replication in vitro (Muggleton-Harris and Hayflick, 1976; Muggleton-Harris and Palumbo, 1979; Muggleton-Harris and DeSimmone, 1980).

MOUSE MUTANTS: MODEL SYSTEMS FOR CATARACTS

33

VI. Possible Areas of Research for the Future If we are to move toward the eventual prevention of cataract, the initiating point in pathogenesis becomes critical. During early morphogenesis, organ or tissue specific “stem” cell lines are established. Initially multipotent, the progeny become committed to the expression of an increasingly restricted number of phenotypes. This process leads to acquisition by cells of specialized structures, e.g., cytoskeletal filaments, altered nuclear-cytoplasmic ratio, and specific functions, e.g., synthesis of lens proteins. Implicit in this process is the idea of a progressive loss of the cell genome, for reprogramming from one type of behavior to another. In vivo the CatF’-differentiated central lens epithelium cells continue to replicate for at least 3 weeks following birth. The noncataractous mouse lens cells cease replicating once the animal is born. This demonstrates a lack of cell regulation during embryogenesis. At which point was this initiated and how can this problem be studied? One approach is chimera studies; chimeric or allophenoic mice (Tarkowski, 1961, 1963; Mintz, 1969, 1971) can readily be made by aggregating 4-8 cell embryos. The embryos are flushed from the oviduct of the mouse, and after removal of the zonae pellucida with acid tyrodes, the embryos are pushed together and held for a few minutes until they adhere. These manipulations are carried out at 37°C. To enable these early embryos to develop they are inserted into the uterus of a pseudopregnant mouse after cultivation to the late morula/early blastocyst stage of development. Differing coat colors, or other genetic variants such as glucose-phosphate isomerase (GPI; EC 5.3. I .9) isozymes will act as genetic markers. The growth and development of the lens can be studied histologically on embryos before birth. Slit lamp observations on a day-to-day basis can monitor the lens in vivo after birth. One area of research would involve the removal of the epithelium and a GPI analysis of the different populations of cells; this will provide an estimate of the contribution of cells from both donor embryos. Strain-specific allelic variants of GPI are analyzed electrophoretically for parental cell lines and the experimental hybrid cells and their resultant clones. The isozyrnes are separated using a Titan I1 ZipZone cellulose acetate plate (Helena Labs, Beaumont, Texas) with 0.025 M Tris base, 0.192 M glycine for 1 hour at 200 V . Following the application of a 2% agar overlay containing 0.3 M Tris-HC1 at pH 8.0,0.3 1 mM NADP, 15 mM fructose 6-phosphate, 20 mM Mgacetate, 2.4 pM dimethylthiazolyldiphenyltetrazolium bromide, 1.3 pM phenazine methosulfate, and 10 IU of glucose-phosphate dehydrogenase, the GPI could be detected. Since such analyses can be accomplished on blood samples from mice, lenses from embryos or chimeras, or manipulated mouse blastocysts, very little material is needed. Thirty cells derived from one hybrid MLE gave

34

AUDREY L. MUGGLETON-HARRIS

clear results that the strain-specific allelic variants of both parental cells were present in the cells (Lipman and Muggleton-Harris, 1982). Rescue of an abnormal developmental defect by embryo aggregation or manipulation of early mouse embryos has been achieved. In a study of inherited photoreceptor cell degeneration (ru/ru), the interaction of mutant and normal pigment epithelium was analyzed in experimental chimeric mice. The eyes of the resulting chimeric mice had particles of normal retina interspersed with particles lacking photoreceptors. The synthesis of rod outer segment disks proceeded normally in photoreceptor cells underlying mutant pigment epithelial cells. The results indicated the site of the mutant gene action to the neural retina (LaVail and Mullen, 1976). Experimental chimerism has also been used as a genetic tool to study the X-linked lethal mutation jimpy up) (Eicher and Hoppe, 1973). Aggregates of the early mouse embryo were made between Tajp/ + x + + / Y crosses and wild-type embryos, the viable X-linked tabby gene serving as a marker to identify chimeras. The experimental chimera was shown to transmit the TajpX chromosome demonstrating that sufficient amounts of the normal cell components of the embryo were present to counteract the lethality of the jp/Y genotype and allow jp/Y cells to differentiate in the testis and form functional jp sperm. When trisomic and diploid (Ts-2n) mouse embryos were aggregated, the resultant chimeras can have Ts and 2n cells in all embryos. This approach has been used for trisomes 15, 16, 17, and 19 and the results indicate that significant numbers of trisomic cells are found in brain, heart, liver, and kidney; the mean proportion of Ts cells was between 40 and 55%. Trisomic cells contribute in a normal manner to development in those chimeras which came to term (Cox et al., 1984). Possible levels of interaction between genotype and environment can be studied at both the cell and embryonic stages. Mouse blastocysts can be injected with genetically distinct embryonic cells, then develop normally and tissues of the resulting offspring are often chimeric consisting of cells from both donor and host origin (Gardner, 1968, 1972). The developmental potential for normal development of embryonal carcinoma cells has been tested in this manner, and in the experimental chimeras, it was found that these “malignant” abnormal cells participated in normal development (Papaioannou et af., 1973). Lack of material with which to study the human congenital cataract makes it necessary to fully utilize available animal models. Mouse mutants enable baseline data to be obtained in sufficient numbers as to be statistically significant. Such data on the embryonic and cellular aspects of congenital cataractogenesis may be used to further our knowledge of this abnormal birth defect. The fields of recombinant DNA and related technologies have revealed much about the structure of the genes associated with lens crystallin synthesis (Piatigorsky, 1981; Garber et af., 1985; Skow, 1982). Both the in vitro and in vivo approaches compliment one another, and eventually advantage will be taken of the rapid

+

MOUSE MUTANTS: MODEL SYSTEMS FOR CATARACTS

35

advances in the gene technology field to challenge the genes directly by incorporating them into the embryonic genome.

REFERENCES Beebe, D. C., Feagans, D. W., Blanchette-Mackie, E. I., and Nau, M. E. (1979). Science 206, 836-838. Bloemendal, H. (1977). Science 197, 127-138. Bloemendal, H. (1981). In “Molecular and Cellular Biology of the Eye Lens” (H. Bloemendal, ed.), pp. 1-47. Wiley, New York. Campbell, M. T., and McAvoy, J. (1984). Exp. Eye Res. 39, 83-94. Cox, R. D., Smith, S. A., Epstein, L. B., and Epstein, C. J. (1984). Dev. Biol. 101, 416-425. Creighton, M. O., Mousa, G. Y., and Trevithick, J. R. (1976). Diflerenfiafion 6, 155-167. Creighton, M. 0.. Mousa, G. Y., and Trevithick, J. R. (1981). Visual Res. 21, 25-35. Eguchi, E., and Kodama. R. (1979). Ophthalmic Res. 11, 308-315. Eicher, E. M., and Hoppe, P. C. (1973). J. Exp. 2001.183, 181-184. Fraser, F. C., and Schabtach. (1962). Gener. Res. 3, 383-387. Friedrich, E., and Glaesser, D. (1971). Acra. Biof. Med. Germ. 27, 41-43. Garber, A. T., Goring, D., and Gold, R. J. M. (1984). J . Biol. Chem. 259, 16, 10376-10379. Garber, A. T., Winkler, C., Shinohara, T.. King, C. R., Inana, G., Piatigorsky. J., and Gold, R. I. M. (1985). Science 227, 74-77. Gardner, R. L. (1968). Nature (London) 220, 596-597. Gardner, R. L. (1972). In “Reproduction of Mammals” (R. V. Short and C. R. Austin, eds.). Cambridge Univ. Press, London and New York. Gellatt, K. M., and Das, N. D. (1984). Curr. Eye Res. 3, 765-778. Genis-Galvez, J.. Maisal, H., and Castro, J. (1968). Exp. Eye Res. 71, 593-605. Hamada, Y., and Okada, T. S. (1977). Dev. Growrh Direr. 19, 265-273. Harding, J. J. (1981). In “Molecular and Cellular Biology of the Eye Lens” (H.Bloemendal, ed.). Wiley, New York. Harding, J. J., and Dilley, K. J. (1976). Exp. Eye Res. 22, 1-73. Hendrix, R. W., and Zwaan, J. (1974). Nature (London) 247, 145-147. Hoenders, H. J., and Bloemendal, H. (1983). J . Geronrol. 38, 278-286. Iwig, M., and Glaesser, D. (1979a). Ophthalmic Res. 11, 293-298. Iwig, M., and Glaesser, D. (1979b). Ophthalmic Res. 11, 298-301. LaVail, M. M., and Mullen, R. M. (1976). Exp. Eye Res. 23, 227-245. Lipman, R. D., and Muggleton-Harris, A. L. (1982). Somatic Cell Genef. 8, 791-800. Lyon, M. F., Jarvis, S. E., Sayers, I., and Holmes, R. S. (1981). Genet. Res. Cambr. 38,337-341. MacAvoy, J. W. (1981). In “Mechanisms of Cataract Formation in the Human Lens” (G. Duncan, ed.). pp. 7-47. Academic Press, New York. McDevitt, D. S., and Yamada, T. (1969). Am. Zool. 9, 1130-1 131. Mamo, J. G., and Leinfelder, P. J. (1958). AMA Arch. Ophthalmol. 59, 417-419. Mintz, B. (1969). Orig. Article Ser. 5, 11-22. Mintz, B. (1971). In “Methods in Mammalian Embryology” (J. C. Daniel, Jr., ed.). Freeman, San Francisco, California. Mostafapor, M. K., and Reddy, V. N. (1980). Invest. Ophthalmol. Visual Sci. 19, 206. Mousa, G. Y., and Trevithick, J. R. (1977). Dev. Growth Difler. 19, 265-273.

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Muggleton-Harris. A. L., and DeSimmone, D. W. (1980). Somatic Cell Genet. 6, 689-698. Muggleton-Harris. A. L., and Hayflick, L. (1976). Exp. CeNRes. 103, 321-330. Muggleton-Harris. A. L., and Palumbo, M. (1979). Somatic Cell Genet. 5 , 397-407. Muggleton-Harris, A. L., Lipman, R. D., and Kearns, J. (1981). Exp. Eye Res. 32, 563-573. Muggleton-Harris, A. L., Festing, M. F. W., and Hall, M. (1986). Genet. Res., in press. Oda, S., Watanabe, K., Fujisawa, H., and Kamagama, Y. (1980). Exp. Eye Res. 31, 673-681. Okada, T. S., Eguchi, G., and Takeichi, M. (1971). Dev. Growth Difler. 13, 323-335. Okada, T. S., Eguchi, G., and Takeichi, M. (1973). Dev.Biol. 34, 321-333. Nelson, L. B. (1984). Ophthalmol. Surg. 8, 697-688. Papaconstantinou, J. (1965). Biochim. Biophys. Acta 107, 81-90. Papaconstantinou, J. (1967). Science 156, 338-346. Papaioannou, V., BcBurney, M. W., and Gardner, R. L. (1973). Nature (London) 258, 70-73. Piatigorsky, J. (1975). Ann. N.Y. Acad. Sci. 253, 333-347. Piatigorsky, J. (1981). Differentiation 19, 134-153. Piatigorsky, J., and Rothschild, S. S. (1972). Dev. Biol. 28, 382-389. Piatigorsky, J., Webster, H. de F., and Wollberg, M. (1972). J . Cell Biol. 55, 82-92. Puck, T. T. (1977). Proc. Narl. Acad. Sci. U.S.A. 74, 4491-4495. Ramaekers, F. C. S., and Bloemendal, H. (1981). In ‘‘Molecular and Cellular Biology of the Eye Lens” (H. Bloemendal, ed.), pp. 85-135. Wiley, New York. Robinson, R. (1971). “Gene Mapping in Laboratory Animals.” (Part A). Plenum, New York. Russell, P., Fukui, H. N., Tsunemetsu, Y.,Huang, F. L., and Kinoshita, J. H. (1977). Invest. Ophthalmol. Visual Sci. 16, 243-246. Skow, L. C. (1982). Exp. EyeRes. 34, 509-516. Tamura, S. (1965). Jpn. J. Ophthalmol. 9, 1130-1131. Tarkowski, A. K. (1961). Nature (London) 190, 857-860. Tarkowski, A. K. (1963). Narl. CancerInst. Monogr. 11, 51-67. Van der Veen, J., and Heyen, C. F. A. (1959). Nature (London) 183, 1137-1138. Watanabe, K., Fujiosawa, H., Oda, S., and Kameyama, Y. (1980). Jpn. J. Ophthalmol. 31, 683689. West, J. D., and Fisher, G. (1985). Genet. Res. Cambr. 46, 45-56. Zwaan, J. (1975). Dev. Biol. 44, 306-312. Zwaan, J., and Williams, R. M. (1968). J . Exp. Zool. 169, 407-422. Zwaan, J., and Williams, R. M. (1969). Exp. Eye Res. 8, 161-167.

INTERNATIONAL REVIEW OF CYTOLOGY. VOL. I W

Cell Wall Synthesis in Apical Hyphal Growth J. G . H. WESSELS Department of Plant Physiology, Biological Centre, University of Groningen, 9751 N N Hnren, The Netherkinds

I. Introduction Fungi, with the exception of the unicellular yeasts, typically grow by means of hyphae which elongate at their apices. The mycelial colony, consisting of a system of branched hyphae, may thus grow over and through substrates. Consequently, the actively growing part of the organism constantly moves away from its original position while colonizing dead organic substrata (saprotrophs) or other living organisms (biotrophs) as in parasitic and symbiontic associations. Particularly with plants, such associations have developed as manifested by the frequent occurrence of fungi as plant pathogens (Dickinson and Lucas, 1982; Misaghi, 1982) and their association with the roots of nearly all plants known as mycorrhizae (Harley and Smith, 1983). Indeed, there is evidence of a longstanding mutual dependence of plants and fungi. They are the main decomposers of the lignocellulose wall of plants (Crawford, 1981) and they have been observed in a vesicular-arbuscular mycorrhizal association with the roots of primitive Devonian land plants (Nicholson, 1975). While the possession of cell walls makes plants immobile and the loss of cell walls was a factor in the aquisition of mobility in animals, the fungi have evolved as organisms with cell walls which nevertheless have a certain mobility due to apical growth. The typical way cell wall growth occurs in plants, namely, diffuse extension growth, can also be found among fungi but never during the invasive growth of the mycelium. Examples of diffuse wall growth are found in a certain stage of elongation of sporangiophores of zygomycetes such as Phycomyces or in the hyphae of elongating stipes of fruit bodies of the Agaricales (basidiomycetes). In fact, one of the hypotheses to explain microfibril orientation in elongating plant cells (Lloyd, 1984), namely, the multinet-growth hypothesis, was formulated on the basis of observations on fibril orientation during intercalary growth of P hycomyces sporangiphores (Roelofsen, 1959). On the other hand, apical growth of tubular cells also occurs in plants, notably during growth of root hairs and pollen tubes (Sievers and Schnepf, 1981). Cytologically as well as functionally these cells resemble fungal hyphae and therefore reference to research on apical wall synthesis in these systems will be made when appropriate. 31 Copyright 0 1986 by Acadumic Prms. Inc. All rights of rcproduction in any form rc\crvcd.

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The problem of apical hyphal growth does not only concern the morphogenesis of a tubular cell. It is also through the growing hyphal tips that the fungus communicates with its biotic and abiotic environment and translates environmental signals in terms of changes in rate or direction of growth or the formation of adaptive apical structures. The direction of growth is regulated in negative autotropism, i.e., the tendency of the hyphae to grow away from each other (Robinson, 1973) and in chemotropism and positive autotropism as in sexual interactions (Gooday, 1975). Recognition between fungal hyphae and plant-host tissues probably also involves hyphal tips with strong evidence for hyphal wall polymers as elicitors of some of the ensuing reactions (Kosuge, 1981; Albersheim er al., 1983). Acceptance or rejection of the fungus by the host tissue is most probably mediated by mechanisms which either allow or check apical growth of the fungus. In the case of biotrophic fungi, acceptance often leads to morphogenetic changes at hyphal apices resulting in typical intracellular infection structures, such as haustoria in parasitic associations and vesicular arbuscular structures in mycorrhizas. Finally, it should be realized that fungi can only display their invasive growth in solid dead substrata or living organisms by virtue of the production of an array of hydrolytic enzymes which hydrolyze polymers in the milieu. On the one hand, the breakdown products can be taken up and used to sustain growth of the fungus. On the other hand, hydrolysis of the polymers clears the way for penetration of the hyphae into the solid substrata. Although little direct evidence is available, it is likely that these enzymes reach the milieu by secretion at the growing hyphal apices (Sentandreu et al., 1981) where the structure of the wall must allow for the passage of these proteins (Chang and Trevithick, 1974). This brief overview of biological activities associated with apical growth in fungi serves to illustrate the importance of understanding how wall synthesis and expansion at the hyphal apex takes place. This is not an easy problem because the apex represents but a tiny part of a hypha. Also, while in growing hyphae wall synthesis per unit area is maximal at the tip, the total amount of wall material synthesized subapically at the same time is appreciable and may qualitatively differ from wall material synthesized at the apex (Sietsma et al., 1985). Therefore, conventional techniques of biochemistry must be combined with a variety of other approaches, including cytochemical and autoradiographic methods, to begin to understand the process of apical wall growth. 11. Observations on Living Hyphae A. THEPHENOMENON OF APICALGROWTH The phenomenon of apical cellular growth received much attention in the nineteenth century and hypotheses generated at that time to explain polarized growth of tubular cells surrounded by rigid walls survive in modem concepts on

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apical growth. Most notable is a publication of Reinhardt (1892) which reviews concepts prevailing at that time. This work added a wealth of new observations interpreted within the framework of a model which still contains elements worth consideration in the light of modem knowledge. Reinhardt worked with the wide hyphae (16- 18 pm) of Peziza species, particularly Peziza sclerotiorum, which displayed a growth rate of 14-23 p,m min- with a maximum of 34 pm min- I . These values are similar to those recorded for the fast-growing leading hyphae of Neurosporu crussa (Steele and Trici, 1975), another ascomycete which has become the subject of many related observations. The most relevant observations made by Reinhardt (1892) can be summarized as follows. 1. There is a relationship between the shape of the apex and the growth rate of the hypha. Slowly growing hyphae have half-spherical tips; with increasing growth rates the tips assume a more tapered appearance going to half-ellipsoids of revolution. 2. When hyphal growth is temporarily stopped, e.g., by applying water to the cultures, the apices swell. With minor disturbances growth resumes after some time from these apices but the emerging hyphae have a smaller diameter. If growth is checked for a longer time, then the swollen tips flatten and hyphal growth from these tips is irreversibly blocked. Resumption of growth takes place by branches arising just under the modified tip. 3. A positive correlation exists between growth rate of hyphae and their diameters. 4. Flooding of hyphae with water often causes bursting of tips. However, bursting never occurred at the extreme apex but always at the boundary of the apical dome and the cylindrical part of the hypha. 5. The three first observations and observations concerning the the site of Occurrence of curvature in growing hyphae were considered as evidence for growth at the tips. Reinhardt concedes that the observation of subapical bursting of tips could be construed as evidence for growth just under the tip because there the wall is apparently weakest. However, he considers that just in this area the tangential stress in the wall, due to turgor, becomes maximal and turgor pressure could thus cause rupture of the young wall in this area. Direct demonstration of apical growth by observing the displacement of particles applied to the apex was possible only with root hairs of Lepidium sativum.

With regard to the mechanisms involved in apical extension growth Reinhardt ( 1 892) considered the then prevailing theory of leading botanists who regarded enlargement of wall area at the apex as a process in which an elastic wall expands under turgor pressure while new wall material is being added by apposition or intussusception. However, he regarded this theory as inadequate because it would require an increase in mechanical strength of the wall going from the very apex to the base of the extension zone. As he puts it, such an increase in strength could be achieved by a proportional increase in wall thickness or by a change in

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the quality of the molecules that make up the wall. He found no evidence for a change in wall thickness. A change in the quality of the wall molecules-which is precisely the kind of change which recent work in our laboratory suggests (see Section V,B)-was considered unlikely. In both cases one would expect that an increase in hydrostatic pressure as caused by flooding with water would result in swelling and bursting at the extreme tip which he did not observe. Reinhardt (1 892) concluded that the wall must have uniform strength over the whole apex and that the wall grows by intussusception of wall material, maximally at the extreme tip and declining to zero at the base of the extension zone. In his view turgor pressure does not cause plastic expansion of the wall at the apex which is rigid enough to withstand this pressure. The fact that relief of turgor pressure by applying solutions of low osmotic potential stopped growth was interpreted as being due to detachment of cytoplasm from the apical wall, thus interrupting the organized delivery of new wall materials by the cytoplasm to the wall. The basic observations of Reinhardt have been confirmed and extended by later workers. Using artificial markers, direct evidence was obtained that growth of hyphae is confined to the apex (Burgeff, 1915; Smith, 1923; Stadler, 1952) and, moreover, that there exists a gradient in expansion rate of the surface declining from the extreme tip to the base of the extension zone (Castle, 1958). Trinci and Saunders (1977) have confirmed that apices of fast-growing hyphae are not hemispherical but more closely approximate half-ellipsoids of revolution. The many measurements made by Trinci and co-workers (Trinci, 1973; Steele and Trinci, 1975) have also revealed that there is indeed a rather strict positive correlation between the growth rates of individual hyphae, their diameters, and lengths of the extension zones. The uniformity of wall thickness over the apex was confirmed by electron microscopy of sections (Girbardt, 1969; Grove and Bracker, 1970; Trinci and Collinge, 1975). In agreement with Reinhardt’s observations, experiments of Robertson (1958) with Fusarium oxysporum showed that after flooding of hyphae with solutions of high- or low-osmolarity arrestment of growth was accompanied by swelling just under the apex. Continuation of hyphal growth occurred either from the swollen apex, after a brief arrestment, or by subapical branching when arrestment was longer than 40 seconds. The phenomenon that hyphal apices when flooded with solutions of low osmolarity tend to swell and burst in a region just under the very tip has also been noted by others (Robertson, 1958, 1965; Park and Robinson, 1966; Bartnicki-Garcia and Lippman, 1972b; J. H. Sietsma and J . G . H. Wessels, unpublished). Although these independent studies largely confirm the basic observations made by Reinhardt (1892), his model of apical growth has generally been rejected in favor of models featuring expansion of the wall by turgor pressure. It can be surmised that this was partly due to d’Arcey Thompson’s (1918, 1942) considerations on the origin of cell form. Yet, as will be discussed later, modern research on the cytoskeleton in the hyphal apex may support some of the ideas of Reinhardt on the formative capacity of the cytoplasm in apical growth.

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B. TURGOR A N D WALLEXPANSION To what extent the wall at the apex is under turgor pressure is not entirely clear. Relatively few studies have addressed this question. Robertson (1958) flooded colonies of F . oxysporurn, growing on agar, with sucrose solutions and examined the effect on the growth of leading hyphae. When flooded with solutions between 0.125 and 0.25 M sucrose growth of the hyphae was unaffected. Solutions of lower or higher osmotic potential resulted in temporary arrestment of apical extension followed by subterminal branching. Robertson concluded that the internal osmotic potential is equivalent to 0.125-0.25 M sucrose. Park and Robinson (1966) found rapid bursting of apices and extrusion of cytoplasm when hyphae of Aspergillus niger were flooded with 0.5% acetic acid. By combining solutions with varying concentrations of sucrose with 0.5% acetic acid they determined the osmotic equivalent between 0.1 and 0.15 M sucrose. Like Robertson they showed that hyper- or hypotonic solutions caused arrestment of apical growth but the hyphae quickly equilibrated apparently by reestablishing a normal internal hydrostatic pressure after which growth resumed. Using the wide hyphae of N . crassa, Robertson and Rizvi (1968) have made measurements of the water potential by determining swelling and shrinking in diameter of hyphae in various sucrose concentrations, and of the osmotic potential by determining incipient plasmolysis. They arrived at a turgor pressure of 1240 and I750 kPa for apical and basal hyphal compartments, respectively. As pointed out by the authors this pressure difference may be a factor in forcing cytoplasm toward the apex. All these observations and measurements attest to the fact that growing hyphae are turgescent and that internal hydrostatic pressure is a component of the apical growth process. However, to what extent the extending apical wall is subjected to internal hydrostatic pressure is not immediately evident because the underlying cytoplasm represents a highly structured entity (see Section HI). Vacuoles responsible for building up high internal pressures are present only in the basal part of the hypha. In addition, there seem to be no reports establishing a quantitative relationship between the growth rate of hyphae and the magnitude of turgor pressure. It could be argued that bursting of the tip and the violent extrusion of cytoplasm after applying various chemicals represent proof that the growing wall at the apex is under high turgor pressure. Chemicals, such as polyoxin D, acetic acid, chelating substances, salts, alcohols detergents, etc., have all been thought to act by weakening the cell wall at the apical dome (Park and Robinson, 1966; Bartnicki-Garcia and Lipprnan, 1972a,b). Although some of these substances without doubt act in this way, e.g., polyoxin D, some of them may also have acted primarily on the structured cytoplasm underlying the apical wall (see Section 111). However, the strength of the argument that bursting indicates full turgor pressure in the apex is particularly weakened by the fact that swelling and bursting are often localized just under the apex where extension has

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ceased. In fact, often the whole apical dome is blown off under these circumstances (J. H. Sietsma and J. G. H. Wessels, unpublished observation). It thus appears possible that bursting occurs when the apical cytoplasmic organization is destroyed or when a weakened cell wall synthesized at the apex is displaced by growth toward the subapical region where turgor pressure becomes higher. Picton and Steer (1982) have also argued that it seems very unlikely that the thin wall covering the apices of extending pollen tubes has sufficient mechanical strength to contain the osmotic pressure of the cytoplasm. They propose that the pollen tube apex is stabilized by a fibrillar cytoplasmic network similar to that found in amoebae and slime molds. Although there is no proof for the existence of a cytoplasmic component which protects the expanding wall at the hyphal apex against the turgor pressure prevailing elsewhere in the hypha, it seems appropriate to keep this possibility in mind when discussing biophysical models of hyphal growth. In these models the hypha is treated as a tubular wall expanding at one end under uniform turgor pressure, with the role of the cytoplasm reduced to the delivery of wall precursors and enzymes to the growing wall.

C. BIOPHYSICAL MODELSOF APICALGROWTH Inspired by considerations of d'Arcy Thompson (1917) on the origin of cell form, several mathematical models have been put forward to describe hyphal morphogenesis (de Wolff and Houwink, 1954; Da Riva Ricci and Kendrick, 1972; Green, 1974; Trinci and Saunders, 1977; Koch, 1982). These models describe the gradients in expansion rate of wall areas in the apical dome in order to generate a tubular structure. Depending on the shape of the tip, hemispherical or half-ellipsoid, the rate of expansion of any point on the apex is proportional to the cosine or the cotangent of the angle between this point and the longitudinal axis of the hypha (Green, 1974; Trinci and Saunders, 1977; see Fig. 1). In all I

I

I

I

I

I I

I

--+-nsion zone

I,

I

I

FIG. 1. The rate of expansion of any point at the apex is proportional to the cosine of the angle a when the shape of the tip is hemispherical (A) or to the cotangent of the angle a when the shape of the tip is half-ellipsoid (B).

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cases expansion is maximal at the very tip and declines to zero at the base of the apical dome. In these models turgor is considered as the force that drives expansion. For a hemispherical tip, Green has argued that the stress in the wall due to turgor is uniform over the apex and that the decline in expansion must thus be caused by a decreasing tendency of the wall to yield to the internal pressure. Qualitatively this also holds for tips with deviating shapes. Because expansion does not lead to thinning of the wall there must be a gradient in the addition of wall materials to the wall which matches the gradient in expansion of the wall. Indeed, in N. crussu hyphae it was found (Trinci and Saunders, 1977) that the cotangent function describing wall expansion was matched by similar gradients in chitin synthesis, as determined by autoradiography , and in the concentration of cytoplasmic vesicles thought to deliver wall materials to the growing wall. Such correlates, however, do not reveal causal mechanisms and the mathematical models are probably carried too far if they are used to prove or disprove mechanisms as attempted by Saunders and Trinci (1979). Their mathematical model could not have excluded a mechanism of growth as envisaged by Reinhardt (1 892) in the absence of preconceived mechanistic concepts. Nevertheless, growth by expansion of a plastic wall is a much more appealing mechanism than the mechanism proposed by Reinhardt because it allows for the addition of wall materials by apposition and because stretching of the wall is a well-known phenomenon in the nonapical diffuse extension growth of cell walls of plants and some fungal systems, e.g., stage IV sporangiophores of Phycomyces. By different means Roelofsen (1950) and Ahlquist and Gamov (1973) artificially stretched these sporangiophore walls and showed that the growing zone, an area just under the developing sporangium, was the most extensible part of the wall exhibiting permanent nonelastic deformation. Nongrowing parts of the wall displayed only small reversible elastic deformations. In Phycomyces stage IV sporangiophores the extensible wall area clearly arises from a formerly nonextensible wall area and there is little doubt that extension occurs under considerable turgor pressure. The growing apical wall, however, has no history of stiffness, its viscoelastic properties have not been determined, and thus the internal pressure necessary to deform this wall is unknown. Assuming that the wall deposited at the apex has plastic properties, then some mechanism must exist that gradually stiffens this wall so that eventually a tubular wall shape can be maintained even under the highest turgor pressure. Green (1974) has advanced the interesting idea that the wall could automatically acquire this stiffness if the increase in resistance to stretch were a function of previous stretch. Whether this is a realistic proposition is hard to tell on the basis of present knowledge of hyphal wall structure. On the basis of what is known about the structure of the wall, two apparently opposing views have been presented regarding the maintenance of a gradient in

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plasticity of the wall at the growing tip. One view, explicitly formulated by Bartnicki-Garcia (1973), holds that the newly deposited wall is inherently rigid and that lytic enzymes permanently loosen this wall and create gaps for the insertion of new wall material. The wall thus grows by intussusception and the theory is mostly referred to as implying “a delicate balance between lysis and synthesis.” Another view, more recently emanating from work in the reviewer’s laboratory (Wessels et al., 1983; Wessels, 1984), holds that the wall material deposited at the growing apex by apposition is inherently plastic in nature but gradually develops stiffness due to secondary processes occurring in the wall. Both theories will be more fully discussed in Section V.

D. ELECTRICAL CURRENTS AND ION GRADIENTS Elongation of fungal hyphae is a clear example of polarized growth. With the advent of the vibrating electrode (Jaffe and Nuccitelli, 1974) it has been become clear that all tip-growing systems generate endogenous electrical currents such that positive ions enter the tip and leave the system subapically (Jaffe and Nuccitelli, 1977; Quatrano, 1978; Nuccitelli, 1983). The first indications for the occurrence of such currents in fungi stem from Slayman and Slayman (1962) who used intracellular electrodes with the wide hyphae of N. crassa. The first measurements with the vibrating probe with fungi were by Stump et al. (1980) in the water mold Blastocladiella emersonii in which they showed a current of positive ions, presumably protons, entering the tips of the rhizoids and leaving over the surface of the thallus. Since then transhyphal currents have been demonstrated in a variety of filamentous fungi, always showing entry of positive currents at a growing hyphal apex: the ,oomycetes Achlya debaryanum (Armbruster and Weisenseel, 1983) and Achlya bisexualis (Kropf et al., 1983), the ascomycetes N. crassa and A. niger and the basidiomycetes Schizophyllum commune and Coprinus cinereus (Gow, 1984), and the deuteromycete Trichoderma harzianum (Horwitz et al., 1984). Not only do these endogenous currents accompany apical growth, they also precede the emergence of an apical growth center as shown for pollen germination (Weisenseel et al., 1975) and branching in fungal hyphae (Kropf et al., 1983). In addition, the property of applied constant electrical fields to determine the site of organization of an apical growth center, as shown for rhizoid outgrowth in fucoid eggs (Peng and Jaffe, 1976), has also been demonstrated in a fungal system by de Vries and Wessels ( 1982). They showed that at an applied field of 25 mV/cell, 75% polarization of the outgrowth of hyphae from regenerating protoplasts of S. commune occurred toward the anode. It appears that the endogenous currents in fungi are largely carried by protons (Gow et al., 1984), possibly by a spatial difference in the location or activity of an electrogenic proton-translocating ATPase in the plasma membrane (Goffeau

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and Slayman, 1981). Jennings (1979) has suggested a role for a subapically active K + ,Na+ -ATPase for generating an ionic current in the marine fungus Dendryphiella salina. What is the significance, if any, of these self-maintained transhyphal currents? At the moment only speculations are possible.

I . Self-electrophoresis of vesicles toward the growing wall area. This mechanism, originally proposed by Jaffe et al. (1974), has been surmised as operative in fungal hyphae (Bartnicki-Garcia, 1973; Harold, 1977). Alternatively, the vesicles could be carried by water flow caused by electroosmosis (Jennings, 1979). 2 . Polarization of the plasma membrane by lateral movement of charged particles in the lipid bilayer under the influence of the electricalfield (Jaffe, 1977). Once initiated, the electrical current could sustain itself by positioning the pumps generating the current in the proper location along the membrane. Also, lateral diffusion of wall synthetic enzymes in the membrane by the electrical field could determine their proper distribution in the apex. The low strength of the fields sufficient to effect polarization in S. commune protoplasts (10% polarization at 0.7 mV/cell) has suggested such a mechanism to deVries and Wessels (1982) but they were unable to show dislocation of chitin synthesis under influence of the applied electrical field. 3. Direct influence of the membrane potential on the activity of wall synthetic enzymes. Such a mechanism has been suggested on the basis of experiments on cellulose biosynthesis in bacteria (Delmer et a l . , 1982) and plants (Bacic and Delmer, 1981). A disturbing fact in applying any of these theories is that apical hyphal growth is apparently possible without a net positive current traversing the hypha, as shown by Kropf e f al. (1983). In Ac. bisexualis they showed that a maximum inward current developed at a site along the longitudinal wall before the emergence of a branch at that site. During this process, the inward current at the tip of the main hypha decreased and even turned outward before slowly reverting to an inward current. During the period of outward current, lasting about 1 hour, the main hypha continued to elongate at normal speed. However, by using pHsensitive microelectrodes Cow et af. (1984) have shown that during the period of reversed current protons continue to flow into the apex. They concluded that tip growth is always accompanied by an influx of protons but that reversal of the electrical current must have been caused by the flux of other ions. The influx of protons in growing hyphal tips may be related to the reported acidity of the apical cytoplasm as detected with pH indicator dyes (Turian, 1978). The apparent importance of the flux of specific ions makes if attractive to consider a role for cytoplasmic gradients for such ions. 4. Muintenunce qf a polarized cytoskeleton organization in the apex by ion

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gradients. As discussed in Section 111, disturbance of the cytoskeleton with antitubulins and cytochalasins can dramatically influence wall deposition. In addition, evidence is accumulating for a general role of the cytoskeleton in the movement of vesicles (Schroer and Kelly, 1985). The electrical transhyphal currents may be related to the proper distribution in the hyphae of, e.g., H + and Ca2+ necessary for the proper functioning of processes based on this cytoskeleton. Most information relates to the distribution of Ca2+ in apices. X-ray microanalysis by Galpin et al. (1978) showed that in Chaeromium Ca2+ is the only cation with a significant spatial gradient. X-ray microanalysis also showed a Ca2+ gradient in pollen tubes (Reiss et al., 1983). Using chlorotetracycline fluorescence Reiss and Herth (1979a) showed indirectly a tip-to-base gradient in Ca2+ concentration in Achlya hyphae and in various other tip-growing systems. Growing pollen tubes require an optimal external Ca2 concentration and disturbance of the Ca2+ gradient with ionophores and chelaters leads to cessation of growth and a collapse of the ultrastructural polarity of the cytoplasm (Herth, 1978; Reiss and Herth, 1979b, 1980, 1982). Picton and Steer (1982, 1983) have developed interesting ideas about the role of Ca2 in the dynamics of the cytoskeleton in the tips of pollen tubes. Such studies are not yet available for fungal hyphae but Reissig and Kinney (1983) have shown that a Ca2 ionophore induces branching in N . crassa. Also the presence of calmodulin in fungi (Gomes et al., 1979; Grand et al., 1980; Ortega-Perez er al., 1981; Muthukumar et al., 1985), a protein known to regulate many Ca2+-mediated processes (Cheung, 1982), should be considered when searching for possible important roles of Ca2+ in hyphal elongation. +

+

+

111. The Cytoplasmic Components of Apical Wall Growth A. CYTOPLASMIC VESICLES

In an excellent review Grove (1978) has surveyed the apical cytoplasmic organization of the various groups of fungi, as seen after conventional chemical fixation. Emphasis was placed on the occurrence of a large number of vesicles apparently en route for fusion with the plasma membrane. This appears to be an attribute of all tip-growing cells and can be regarded as a highly polarized system of exocytosis. In conventially fixed preparations fusion profiles are frequently seen (Girbardt, 1969; Grove and Bracker, 1970) but they are extremely rare in preparations fixed by freeze substitution (Howard and Aist, 1979; Hoch and Howard, 1980; see Fig. 2). This may be due to the rapid fixation achieved by this procedure and the extreme velocity at which fusion occurs (Hoch and Howard, 1980). Historically, the assemblage of vesicles at the hyphal apex relates to the so-

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called Spitzenkorper (apical body). In phase optics this is a dark area seen in the apex of elongating hyphae of ascomycetes and basidiomycetes but not of zygomycetes and oomycetes (Girbardt, 1957, 1969). Girbardt ( 1969) and McClure et al. (1968) have equated the Spitzenkorper with the whole concentrated assemblage of vesicles while Grove and Bracker (1970) have argued its equivalence with a central vesicle-free zone in the apices of septate fungi (Fig. 2 and see below). Howard (1981) has discussed this controversy and concluded that Girbardt’s view is probably correct. Girbardt (1957, 1969) found that growth is closely associated with the presence of the Spitzenkorper. When growth is arrested the Spitzenkorper vanishes and reappears just before growth resumes and its position is related to the direction of growth. A central position precedes straight growth while an eccentric location is followed by a change in the direction of growth. In the latter case thickening of the wall, and presumably retardation of wall expansion, occurs in the wall area closest to the Spitzenkorper resulting in curvature of the hypha. The vesicles in the hyphal apex vary in size and contrasting contents. For a description of the vesicles after conventional fixation the reader is referred to the review of Grove (1978). Because the details observed after fixation by freeze substitution are superior, the following summary is from the work of Howard and Aist (1979) on the ascomycete Fusarium acuminatum and Hoch and Howard (1980) on the basidiomycete Laetisaria arvafis (Fig. 2). Vesicles of 70-120 nm diameter occur nearly exclusively at the apices and at the sites of branch formation. They are therefore referred to as apical vesicles and two types of these can be distinguished on the basis of staining of the contents with osmium tetroxide. Microvesicles, 20-50 nm diameter and often characterized by an hexagonal outline, were seen throughout the apex (in the ascomycete, also in the central zone free of apical vesicles). In addition, these microvesicles and a special type surrounded by fibrous material (called filasomes) were found along lateral walls and at the sites of septum formation. Apical vesicles and microvesicles were also seen in association with fenestrated sheets of smooth endoplasmic reticulum located subapically in tip cells presumably representing the Golgi equivalent in these septate fungi. These Golgi equivalents occurred in close association with mitochondria and probably are the source of both apical vesicles and microvesicles. No direct information is available on the rate at which vesicles are produced and fuse with the plasma membrane. On the assumption that membrane material for extending the plasma membrane is delivered by fusion of the apical vesicles, Collinge and Trinci (1974) calculated a fusion rate of 38,000 vesicles per minute for fast-growing hyphae of N . crassa. This rate may even be an underestimate because recycling of membrane material was not considered. Using a method in which a short exposure to cytochalasin D only interfered with vesicle transport, Steer and Picton (1984) estimated the rate of production of apical vesicles in

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FIG.2. Apex of Fusarium acuminatum fixed by freeze substitution (A) and conventional technique (B). (A) Apex of a freeze-substituted hyphal tip cell. Apical vesicles of two electron densities partially surround a cluster of ribosome-like particles. Note the presence of a microtubule (MT) within the apical dome and the close association between mitochondria (M) and smooth cistemae (SC). Expansion of the outer cell wall layer can be surmised at the upper right (arrows). Bar, 1.O km. Inset: Enlargement of the boxed area. The apical cell wall is distinctly four layered. Note the apical vesicles (V) and smooth contour of the plasmalemma (P).Bar, 100 nm. (B)Apex of a conventionally fixed hyphal tip cell. The mitochondria (M) have been so grossly distorted that enclaves of cytoplasm (*) appear as though part of the mitochondria. Electron-lucent vesicles and ribosome-like particles

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pollen tubes growing at different rates (28 and 7 pm min-I). There was little difference in the vesicle production rate and the calculations indicated 8 0 4 7 % recycling of membrane material delivered to the plasma membrane at the lower growth rate. What is the role of the vesicles in wall formation at the hyphal apex? In many publications (cf. Grove, 1978) they are loosely regarded as being involved in making new wall. One possibility would be that they carry wall polymers ready for insertion in the growing wall. lntracellular synthesis of wall polymers delivered to the wall by vesicles applies to pectin, hemicellulose, and hydroxyprolinerich glycoprotein in plants (Willison, 1981; Northcote, 1984) and to wail mannoproteins in yeast (Sentandreu et al., 1981). For filamentous fungi there is no good evidence for this. Cytochemical staining does detect polysaccharide material in the apical vesicles (Grove, 1978), but this material may represent glycoprotein enzymes destined for export. Electron microscopic autoradiography of hyphal apices of S . commune, labeled to detect chitin synthesis (van der Valk and Wessels, 1977), and Saprolegnia monocica, labeled to detect glucan synthesis (Fkvre and Rougier, 1982), has failed to detect intracellular synthesis of wall polymers. Girbardt (1969) has expressed doubts that wall constituents could be carried by apical vesicles because they are not seen near septa where active wall synthesis is taking place. However, because lateral walls may contain polymers not present in the septa (Hunsley and Gooday, 1974; van der Valk er al., 1977; Wessels and Sietsma, 1979) it is still possible that the apical vesicles carry such polymers to the growing wall. Apart from their possible role in carrying some wall polymers to the surface, there is little doubt that the vesicles are part of the general secretory pathway studied much better in yeasts (Sentandreu et al., 1981; Scheckman, 1982). Presumably they contain the whole array of hydrolytic enzymes so abundantly excreted by fungi while their membranes contain plasma membrane proteins among which are those functioning in wall synthesis. Again, there is little experimental evidence to substantiate this point. Cytochemistry has revealed the presence of cellulase in apical vesicles of Achlya ambisexualis (Nolan and Bal, 1974). In the same fungus Hill and Mullins (1980) demonstrated the vesicular location of a nucleoside diphosphatase and acid phosphatase while Dargent (1975) detected alkaline phosphatase in apical vesicles and Golgi cisternae of Ac. bisexualis. Various studies have attempted to assign polysaccharide synthase and hydrolase activities to cellular structures by differential centrifugation of homogenates (e.g., see Fkvre, 1979). However, vesiculation of the plasma membrane

(arrows) appear at the cell apex. Bar, 1.0 pm. Inset: Enlargement of the boxed area. The cell wall is seen as two layers (i, ii) and is irregular in outline on the inside. The contents of apical vesicles (V) appear coarsely fibrous. Bar, 100 nm. (With permission from Howard and Aist, 1979.)

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and cross-contamination of membranes make it very difficult to draw conclusions, except for the chitin synthase particles to be discussed below. Microvesicles (Fig. 2) are not confined to hyphal apices but are also abundantly present along lateral walls and developing septa (Girbardt, 1969; Hoch and Howard, 1980; Howard, 1981). Howard (1981) noted a hexagonal outline in sections of these microvesicles but dismissed the possibility that they represented mycovirus on the basis of absence of dsRNA, the usual nucleic acid component of these viruses. Yet, the general Occurrence of mycovirus without any apparent disease symptoms (Lemke, 1979) should be kept in mind. Like Bracker er al. (1976) and Girbardt (1979), Howard (1981) suggested that at least some of the microvesicles could be identical to chitosomes which look similar when sectioned after isolation (Bracker er al., 1976). Chitosomes can be isolated from a variety of chitin-containing fungi (RuizHerrera et al., 1975; Bartnicki-Garcia et al., 1978; review by Bartnicki-Garcia and Bracker, 1984). They measure 40-70 nm, have a membrane-like shell rich in sterols and no glycoprotein, and apparently contain exclusively chitin synthase in an inactive form. By proteolytic digestion chitosomes are activated in virro and then synthesize crystalline chitin from uridinediphospho-N-acetylglucosamine, apparently one microfibril per chitosome. Toluene-permealized germlings of Mucor rouxii after treatment with trypsin-synthesized chitin from added widine diphospho-N-acetylglucosaminewithin the cytoplasm without an apparent preferential localization, as shown by electron microscopic autoradiography (Sentandreu et al., 1984). This may reflect the observation that the microvesicles are not strictly localized at the apex. Bartnicki-Garcia and Bracker (1984) concluded that chitisomes convey a package of chitin synthase zymogen to the cell surface, probably to the plasma membrane. Although there is now good evidence for the operation of chitin synthase in the plasma membrane (see below), they still considered both intracellular synthesis of chitin and synthesis of chitin by chitosomes extruded into the periplasmic space as possibilities. Previous claims as to the presence of inactive chitin synthase in the so-called gamma particles of zoospores of B. emersonii (Mills and Cantino, 1981) have been refuted (Dalley and Sonnebom, 1982; Hohn et al., 1984). B. CYTOSKELETON Microtubules in filamentous fungi have been extensively described in relation to mitosis (Heath, 1978) and nuclear migration (Girbardt, 1968; Raudaskoski and Koltin, 1973) but their abundance in apices only became evident after rapid fixation of hyphae by freeze substitution (Howard and Aist, 1979; Hoch and Howard, 1980; Howard, 1981; see Fig. 2). As in subapical areas the microtubules in the apex run parallel to the hyphal axis and some are seen to end at the apical plasma membrane. The abundance of microtubules at the tiyphal apex has

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now also been demonstrated by immunological techniques in Uromyces phaseofi (Hoch and Staples, 1985). This became possible by using monoclonal antibody against yeast tubulin (Kilmartin and Fogg, 1982) and permeabilization of the fungal wall by lytic enzymes to allow the antibody to gain access to the cytoplasm. Hoch and Staples (1985) even obtained evidence for the presence of microtubule nucleating regions in the hyphal apex. In S . commune microtubules, extending into the apex, have been visualized using antibodies against mammalian a- and p-subunits of tubulin (Runeberg and Raudaskoski, 1986). Fungal microtubules, except those of the oomycetes, are relatively insensitive to standard antimicrotubule agents, such as colchicine, colcimid, and Vinca alkaloids. However, certain systemic fungitoxic benzimidazole derivatives, such as benomyl and its hydrolysis product methyl benzimidazole-Zyl carbamate (MBC), have similar effects on fungal microtubules as colchicine on those of plants and animals (Davidse, 1975; Davidse and Flach, 1977). Morris and coworkers (see Gambino et al., 1984, and lit. cit.) have indicated the p-tubulin subunit as the target for these antitubulins and have examined a number of benomyl resistance mutations of Aspergiffusnidufans with respect to effects on mitosis and nuclear migration. Howard and Aist (1977, 1980) have investigated the effects of MBC on hyphal tip elongation and ultrastructure of F. acuminatum. At a concentration of MBC permitting reduced growth (65-30% of control) light microscopy showed the formation of a wavy hyphal tube, rounding off of the tips, disappearance of the Spitzenkorper, and an increasing distance between the mitochondria-rich area and the tip. The ultrastructural study (Howard and Aist, 1980) showed the extreme susceptibility of apical microtubules to MBC and a dramatic effect on the distribution of the 70-90 nm diameter vesicles. In control cells 50% of all the vesicles in the apical 30 pm of the hypha were found to lie within 2 pm of the apex. While the total number of vesicles did not change much, MBC caused this vesicle distribution to become uniform resulting in a substantial increase in the number of vesicles in subapical regions. Howard and Aist (1980) proposed this to be due to continued fusion of vesicles with the plasma membrane together with a reduction in the rate of transport of the vesicles. Consequently they implied a role for apical microtubules in the transport of apical vesicles. Such a role would seem plausible because of the general implication of microtubules in vesicle transport in animals (Dustin, 1978) and the recent in vitro demonstrations of movement of axoplasmic vesicles along single microtubules in the presence of ATP (reviewed by Schroer and Kelly, 1985). In pollen tubes of Lilium and Clivia, colchicine, at concentrations which led to complete disappearance of subapical microtubules, failed to inhibit growth (Franke et al., 1972). Also in secretory root cells, colchicine apparently did not affect migration of secretory vesicles (Mollenhauer and Morrk, 1976). On the other hand, colchicine has been shown to inhibit apical growth in pollen tubes of

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Nicotiana tabacum (Derksen and Traas, 1985) and in tip-growing moss and algae cells (Schmiedel and Schnepf, 1980; Mizukami and Wada, 1983). In none of these tip-growing plant systems were axially running microtubules observed in the tips (for root hairs, see Traas et a f . , 1985) but, as in fungi, this may reflect the use of conventional fixation procedures. For the moment, the plant systems thus offer no support for the purported role of microtubules in vesicle transport. Microfilaments, 4-7 nm in diameter, have been found in hyphae of many fungi. They are particularly clear in a circumferential filamentous belt in the area of septum formation (Patton and Marchant, 1978; Girbardt, 1979). Girbardt ( 1979) has expressed doubt about contractile properties of this filamentous septal belt because of its position relative to the invaginating plasma membrane and its apparent insensitivity to cytochalasin B. However, cytochalasin B is not usually effective against fungi while cytochalasin A, C, D, and E have been more useful (Betina e t a f . , 1972; Allen e t a f . , 1980; Hoch and Staples, 1983). Again, fixation by freeze substitution has also enabled the visualization of microfilaments in the apical region of fungi, particularly in the central vesicle-free area of septate fungi (Howard and Aist, 1979; Hoch and Howard, 1980; Howard, 1981; Fig. 2), an area which looks finely granular after conventional fixation (Grove and Bracker, 1970). Evidence for equating the microfilaments with actin and myosin is accumulating. Allen and Sussman (1978) have extracted material from N . crassa hyphae which in the presence of ATP precipitates to form actin-like filaments which could be decorated with heavy meromyosin. Cytochalasins which interfere with the polymerization of actin (MacLean-Fletcher and Pollard, 1980; Flanagan and Lin, 1980) inhibit hyphal tip growth and cause irregular deposition of wall materials in N . crassa (Allen et a f . , 1980), Gifbertellapersicaria(Grove and Sweigard, 1980), and U. phaseoli (Herr and Heath, 1982). Rhodamineconjugated phalloin, a derivative of phalloidin which binds to F-actin, has been used to visualize actin in situ in U.phaseoli (Hoch and Staples, 1983, 1985). The actin filaments were present throughout the germ tube but especially in more basipetal regions whereas fluorescent plaques, possibly equivalent to filasomes, occurred near the periphery of the cell’s cytoplasm most abundantly in the hyphal tip region. Using the fluorescent probe 7-nitrobenz-2 oxa- 1,3-diazole-phallacidin (NBD-Ph) or anti-actin antibodies, distinct fluorescence was also noted in apices of S . commune (Runeberg and Raudaskoski, 1986). In addition, intense fluorescence was seen in the septal belt area before the formation of a septum, suggesting the equivalence of the constituting filaments with F-actin. The recent discovery of heavy myosin in Saccharomyces cerevisiae (Watts et a f . , 1985) probably forecasts the demonstration of this cytoskeletal element in the filamentous fungi. Cytochalasins also inhibit tip growth and transport of vesicles in pollen tubes (Herth et a f . , 1972; Franke et a f . , 1972). This inhibition of vesicle transport has been used (Picton and Steer, 1981) to measure the rate of production of vesictes by dictyosomes, although prolonged exposure to cytochalasin also seems to in-

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hibit the latter process (Shannon et a f . , 1984). A possible role for Ca2+ in the proper functioning of the cytoskeleton has already been mentioned (Section 11,D).According to Picton and Steer (1982, 1983) rather high concentration of Ca2+ at the extreme apex of the pollen tube are necessary for the fusion of the vesicles with the plasma membrane while a decreasing Ca2+ concentration in a subapical direction is necessary to maintain the microfilament network at the tip sufficiently relaxed. In summary, the fragmentary information that exists does suggest that the growing fungal hypha contains in its apex a cytoskeleton of microfilaments and longitudinally running microtubules. There are also indications for connections of cytoskeletal elements with the plasma membrane. As in nonwalled systems (animals, amoebae, slime molds) the cytoskeleton could thus have form-determining properties. This is particularly important if it is assumed that the new wall deposited over the apex does not as yet have enough strength to maintain form. Although the bursting of hyphal tips by chemical agents (Park and Robinson, 1966; Bartnicki-Garcia and Lippman, 1972b) or cold (Robinson and Morris, 1984) has been attributed to changes in wall metabolism, some of these influences may actually work via disturbance of the apical cytoskeleton. Most probably the cytoskeleton at the apex is involved in the polar transport of vesicles for fusion with the plasma membrane. In addition, by maintaining connections with plasma membrane proteins the cytoskeleton could be involved in regulating the distribution of certain proteins, e.g., transport proteins or polysaccharide synthases, in the plasma membrane. C. PLASMAMEMBRANE With respect to wall synthesis many autoradiographic studies have shown that wall components, such as chitin and glucans, are most actively synthesized in the apical portion of the growing hypha (e.g., Bartnicki-Garcia and Lipmann, 1969; Gooday, 197 I ; Katz and Rosenberger, 197 1 ; Wessels et al. , 1983). In N . crassa it was shown (Trinci and Saunders, 1977; Gooday and Trinci, 1980) that the gradient in chitin synthesis at the apex closely follows the theoretical gradient to maintain uniform wall thickness, maximal at the extreme tip and declining to a minimal value at the base of the extension zone (Section 11,C). Autoradiography also shows heavy chitin synthesis at the site of forming septa while a low but significant degree of synthesis, particularly of glucan (Gooday, 1971 ; Sietsma el al., 1985), may occur along the whole length of the longitudinal wall. For chitin there is good evidence that its synthesis occurs only on the plasma membrane. Electron microscopic autoradiography of chitin deposition in regenerating protoplasts and hyphal apices of S . commune has shown that chitin is synthesized only at the plasma membrane/wall interface (van der Valk and Wessels, 1977). Similar observations have been made for chitin synthesis along

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longitudinal walls in stipes of C. cinereus fruitbodies (Gooday, 1982). The presence of chitin synthase in the plasma membrane has been ascertained by isolating pure plasma membranes using the concanavalin A method developed by Scarborough (1975). Thus Durin et al. (1975) showed the association of chitin synthase with the plasma membrane of protoplasts from the yeast Sac. cerevisiae. The chitin synthase was largely in an inactive form and could be activated by limited proteolysis. Similar results were reported for protoplasts from yeast and mycelial phases of Candida albicans (Braun and Calderone, 1978) and the mycelial fungus S . commune (Vermeulen et al., 1979) in which the plasma membrane-associated chitin synthase was largely in an active state. Cabib et al. (1983) have provided good evidence that the enzyme accepts N-acetylglucosamine residues from uridinediphospho-N-acetylglucosamine at the cytoplasmic face of the membrane and transfers them vectorially to a growing chain of chitin that is concomitantly extruded at the outside of the membrane. The association of chitin synthase with the plasma membrane is compatible with the presence of chitin synthase in the previously discussed chitosomes if the latter are considered as the transport vehicles for inactive chitin synthase to convey the enzyme to the plasma membrane. A clear distinction between two chitin synthase fractions could be made after subcellular fractionation of concanavalin A-coated protoplasts of S . commune (Vermeulen et al., 1979). About 90% of the active chitin synthase was found associated with the plasma membrane while proteolytic treatment of this fraction resulted in a stimulation of only a factor of 1.7. But these protoplasts also contained a chitin synthase fraction, sedimentable at high centrifugal speed and in an inactive form like chitosomes, which accounted for about 50% of the total chitin synthase activity after proteolytic stimulation. Although this study does not prove that the cytoplasmic inactive form is the precursor for the active plasma membrane-bound chitin synthase, this is a suggestive pathway. How the enzyme becomes active in the plasma membrane is not known. There is no direct evidence for proteolytic activation of chitin synthase in vivo although such a scheme has been proposed for activation of chitin synthase at the site of septum formation in yeast with a zymogenic form of the enzyme uniformly present in the plasma membrane (Cabib et al., 1982, for review). Being an integral membrane protein the influence of the lipid environment can be surmised as important for regulating activity. Arrhenius plots show clear transition points in the activity of the enzyme, delipification inactivates the enzyme, and phospholipids activate or restore the activity of partially delipified enzyme (Durin and Cabib, 1978; Vermeulen and Wessels, 1983; Montgomery and Gooday, 1985). Mutations which alter the lipid composition of membranes have an effect on chitin synthesis (Hanson and Brody, 1979; Pesti et al., 1981). Fungicides which interfere with normal ergosterol biosynthesis cause irregular deposition of chitin as inferred from hyphae stained with the optical brightener diethanol

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(Kerkenaar and Barug, 1984; Marichal et al., 1984). Polyene antibiotics, known to interact with sterols, inhibit chitosomal chitin synthesis (Rast and BartnickiGarcia, 1980). Since the chitosomal lipids differ from those present in total membrane preparations (Hernandez et al., 198I), and thus probably also from the lipids in the plasma membrane, it seems possible that the mere transfer of chitin synthase from the chitosomes to the plasma membrane results in activation of the enzyme. There is a clear need for in vitro reconstitution experiments attempting incorporation of chitosomes into plasma membrane vesicles or liposomes. If it is accepted that chitin synthase acts as a vectorial integral plasma membrane enzyme, how then is the polarized pattern of chitin synthesis in the fungal hypha achieved? On the basis of the available information a few possibilities can be considered. Chitin synthase may be continuously inserted into the plasma membrane at the growing apex while its polarized activity could be regulated by local perturbations of the lipid environment in the membrane or by some other reversible mechanism such as phosphorylation/dephosphorylation.Assuming a long halflife for the enzyme this would imply the persistence of a latent chitin synthase in the subapical plasma membrane. Some evidence for this scheme comes from experiments of Issac et al. (1978): “early” protoplasts, formed preferentially from hyphal tips, had chitin synthase with a much higher specific activity and only slight proteolytic activatability in comparison to “late” protoplasts formed preferentially from older areas of the hyphae. One may ask, however, whether the protoplast membrane faithfully reflects the original hyphal plasma membrane in composition. During protoplasting the membrane may grow and new lipids and proteins may be inserted. This uncertainty even applies to the assumed uniform distribution of chitin synthase in the plasma membrane of intact yeast which was inferred from the distribution of chitin synthase in the protoplast membranes (DurBn et al., 1975; Cabib et al., 1983). There is a clear need for the development of immunological methods to detect chitin synthase in situ. Contrary to a reversible activation/deactivation mechanism, Ruiz-Herrera (1984) has proposed an irreversible mechanism for creating a gradient of activity of chitin synthase at the hyphal apex. The proposed mechanism is based on the property of chitosomes to synthesize in vitro a single chitin microfibril upon proteolytic activation after which activity is irreversibly lost. In his words: ‘‘Chitosomes are directed towards the apical pole, they become activated, synthesize a single chitin microfibril, and are deactivated in a matter of minutes to guarantee the polarized growth of fungi.” This mechanism would imply an extremely high turnover rate of chitin synthase for which no evidence is available at the moment. On the contrary, cycloheximide blocking the synthesis of chitin synthase has little effect on the chitin-synthesizing ability of regenerating protoplasts (de Vries and Wessels, 1975) and the drug causes a displacement rather

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than a decrease of chitin synthesis in growing hyphae (Katz and Rosenberger, 1971). This apparent stability of chitin synthase can be reconciled with a rapid turnover of the enzyme at its site of activity only if there were an enormous reserve of latent chitin synthase molecules in the cytoplasm. The autoradiographic study of Sentandreu et al. (1984) indeed indicates that large amounts of chitin synthase may be present in the cytoplasm. A completely different view of localized chitin synthesis, so far not considered in the literature, could be based on the newly discovered cytoskeletal elements in the apex and septa1 region and the possible connections between these elements and proteins in the plasma membrane. A polarized assemblage of stable chitin synthase molecules in the plasma membrane highest in density at the extreme apex could be just another manifestation of the polarized activity of the apical cytoskeleton. The whole assemblage of chitin synthase molecules, stabilized by connections to the cytoskeleton, could be pushed through the membrane while this membrane extends. A disturbance of the cytoskeleton, e.g., by osmotic shock, could then easily lead to a displacement of chitin synthesis as observed (Katz and Rosenberger, 1971). Preparation of protoplasts would then probably result in a completely new pattern of chitin synthase insertion and stabilization in the plasma membrane. Numerous studies have demonstrated the synthesis from uridinediphosphoglucose of a (1-.3)-P-glucan by particular or vesicular preparations from fungi (ref. cit. in Quigley and Selitrennikoff, 1984; Sonnenberg et al., 1985). Control mechanisms involving nucleotides have been proposed to explain localized activity (Ftvre, 1983; Szaniszlo et al., 1985). However, the evidence that the glucan synthase is another plasma membrane-bound enzyme is scarce. An autoradiographic study (Ftvre and Rougier, 1982) suggests that all wall glucans in Saprolegniu monoica are synthesized at the plasma mernbrane/wall interface while a (1-.3)-P-glucan synthase has been found to occur in the plasma membrane of S. cerevisiae protoplasts (Shematek et al., 1980).

IV. Structure of the Fungal Cell Wall A. WALLPOLYMERS

Cell wall analyses of different taxonomic groups of fungi have revealed a remarkable heterogeneity with respect to polymers or combinations of polymers present, as first pointed out clearly by Bartnicki-Garcia (1968). On the other hand, ultrastructural studies (Bracker, 1967; Troy and Koffler, 1969; Hunsley and Burnett, 1970; van der Valk et al., 1977; Burnett, 1979; Wessels and Sietsma, 1979) have shown a general similarity in construction of the wall, i.e., an inner layer containing chitin or cellulose microfibrils apparently embedded in

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TABLE I VAKIOUS POLYM~K OCCUKRING S I N THE CELLWALLw- FUNGI" Cell wall polymers ~~

Taxonomic groups Basidioniycotina Ascomycotina Zygomycotina Mastigomycotina Chytridiomycetes Hyphochytridiomycetes

Alkali soluble Xy lo-manno-protein ( 1+3)-u-u-glucanh (Galacto)-manno-protein ( 1+3)-a-u-glucanh Glucuronomanno-protein pol yphosphate Glucanc Not determined

Alkali insoluble

(1+3)-P/( I+6)-P-~-Glucan Chitin ( l+3)+3/( l+6)-P-~-Glucan Chitin Polyglucuronic acid Chitosan Chitin Glucan' Chitin Chitin Cellulose (l+3)-P/( 1+6)-P-o-Glucan Cellulose

"From Wessels and Sietsma (1981a) with permission. "In a number of cases the alkali-soluble fraction also contains part of the (l+3)-P/( 1+6)-P-uglucan. Incompletely characterized; probably (1+3)-P and ( 1+6)-P-linked.

other polymers and one or more outer layers. As a rule the outer layers are soluble in dilute alkali leaving the inner layer as an insoluble residue. Table I lists various types of alkali-soluble and alkali-insoluble polymers as they occur among the fungi. It appears that within the fungi convergent evolution has provided for the occurrence of a variety of different polymers fulfilling similar functional requirements of the walls. Chitin is the most characteristic component of fungal walls. It is the common feature of the walls of fungi belonging to the Basidiomycotina, Ascomycotina, and Zygomycotina, which according to Whittaker and Margulis ( 1978) constitute the Kingdom Fungi. Chitin has also been found in two classes of the Mastigomycotina, the Chitridiomycetes and the Hyphochitridiomycetes (BartnickiGarcia, 1968) and recent work even suggests the presence of chitin in oomycetes (Compos-Takaki rt a / ., 1982) which typically have cellulose in their walls. Wall composition in the Mastigomycotina will not be considered further. With regard to the "true fungi," chitin and associated polymers such as P-glucans and glycuronans will be discussed because they are likely to play an important morphogenetic role. (For a more comprehensive treatment of fungal wall polymers, see Bartnicki-Garcia, 1968; Wessels and Sietsma, 198 1 a; Bartnicki-Garcia and Lippnian, 1982.) Chitin is usually considered as a homopolymer of N-acetylglucosamine but

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even in crystalline chitin a number of nonacetylated glucosamine residues may occur. X-ray diffraction of fungal chitin shows a microcrystalline conformation known as cx-chitin (which also occurs after spontaneous crystallization of chitin from solution) in which chains presumably run antiparallel. In addition, two other polymorphs, p and y, occur in nature (cf. Muzzarelli, 1977). Evidence for the Occurrence of chitin in walls is mostly obtained by chemical analysis, infrared spectroscopy, and X-ray diffraction analysis of wall residues obtained after consecutive extractions with alkali and acid. After such treatments electron microscopic observations mostly reveal long interweaving microfibrils of varying width in random orientation (Troy and Koffler, 1969; Hunsley and Burnett, 1970; Burnett, 1979; van der Valk er al., 1977), sometimes short rodlets (Kitazima er al., 1972) or granules (Houwink and Kreger, 1953; Pollack er al., 1983). Usually nontreated walls do not show X-ray diffraction lines of chitin (Houwink and Kreger, 1953; Kreger, 1954; Reid and Bartnicki-Garcia, 1976; Sietsma and Wessels, 1977). For the walls of S. commune it was concluded that in native and in alkali-extracted walls most of the chitin is poorly crystallized. Only after treatments which removed the P-glucan did sharp X-ray reflections of chitin (Sietsma and Wessels, 1977) and abundant microfibrils (van der Valk er al., 1977) appear. On the other hand, microfibrils were seen on the inner surface of the native walls but the relationship between crystallinity and microfibrillar structure is not straightforward (Gow and Gooday, 1983). In contrast, on regenerated protoplasts where initially chitin is deposited without alkali-insoluble pglucans the chitin in the native wall is microfibrillar and highly crystalline (van der Valk and Wessels, 1976). Also when chitin is synthesized in virro with chitosomes (Ruiz-Herrera and Bartnicki-Garcia, 1974) or with membrane preparations (Vermeulen et al., 1979) the chitin is microfibrillar and highly crystalline. In the hyphal wall secondary modifications of chitin and interactions between the chitin chains and P-glucan chains (see below) probably prevent the formation of perfect crystallites of chitin. Chitosan is a homopolymer of glucosamine (deacetylated chitin). As mentioned above even in chitin a number of residues may be deacetylated and thus there may be a continous range of polymers from chitin to chitosan varying in the degree of acetylation. This is in line with the observation that chitosan is synthesized from chitin by a deacetylase (Araki and Ito, 1975; Davis and BartnickiGarcia, 1984). Chitosan and partially deacetylated chitin may occur in the walls of chitin-glucan fungi (e.g., Novaes-Ledieu and Garcia Mendoza, 1981) but these polymers are particularly abundant in the walls of zygomycetes where their cationic nature is balanced by the presence of anionic polymers such as inorganic polyphosphate and glycuronans (Bartnicki-Garcia and Reyes, 1968a,b; Datema er al., 1977a,b). Using nitrous acid which specifically attacks nonacetylated glucosamine residues three fractions could be distinguished in the wall of Mucor mucedo (Datema

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et al., 1977b). One fraction which was solubilized by nitrous acid contained N-acetylglucosamine interspersed with glucosamine. Another fraction became nitrous acid soluble after treatment of the walls with pronase or alkali, indicating a polymer containing (N-acety1)glucosamine to which peptides were linked. The remaining fraction appeared to consist of a homopolymer of N-acetylglucosamine with an X-ray diffraction pattern of a-chitin. In contrast, other zygomycetes, e.g., Phycomyces blakesleeanus (Kreger, 1954) and Mucor rouxii (Bartnicki-Garcia and Nickerson, 1962; Bartnicki-Garcia and Reyes, 1968a), apparently contain the homopolymer chitosan. How this chitosan exists in the native wall is unknown but a significant observation is that the crystalline conformation of chitosan is detected only after extraction of the substance from the wall with dilute acid, a treatment likely to break covalent bonds. With the exception of the hyphal walls of zygomycetes, glucans with (1+3)-p and (1+6)-p linkages occur in the walls of all fungi (Bartnicki-Garcia, 1968; Gorin and Spencer, 1968; Rosenberger, 1976; Fleet and Phaff, 1981; Wessels and Sietsma, 1981a). They may occur as more or less water-soluble glucans forming a gel-like layer around hyphae (Gorin and Spencer, 1968; Buck et al., 1968; Wessels et al., 1972) or as a genuine wall component often existing in an alkali-insoluble complex together with chitin. The extreme insolubility of these glucans makes their analysis extremely difficult. Their extraction often involves the use of strong alkali and acid with the risk of breaking covalent bonds and modifying the glucans. For instance, heating with dilute mineral acid may render the glucans soluble in alkali with simultaneous hydrolysis of ( 1+6)-P-linked residues and crystallization of the remaining ( 1-3)-P-linked chains to form “hydroglucan” (Kreger, 1954). The finding that acid-treated pieces of rhizomorphs of Armillaria mellea showed oriented X-ray reflections of hydroglucan enabled Jelsma and Kreger (1975) to show that hydroglucan has a conformation of three intertwining helices each containing six glucose residues per turn of the helix, in accordance with Marchessault er al. (1977). In native walls or alkaliextracted residues, which only give very diffuse hydroglucan reflections, such ordered structures are probably frequently disrupted by side branches giving the glucan gel-like properties (Sato et al., 1981). The reasons for insolubility of a considerable portion of the (1+3)+- and (1-6)-P-linked glucans in the wall are not completely clear. Since the alkaliinsoluble fraction of the wall always contains (N-acety1)glucosamine it is possible that the glucans are actually part of a glucosaminoglucan. In many fungi a large proportion of the alkali-insoluble glucan can be extracted by dimethyl sulfoxide (Sietsma and Wessels, 1981) but the conclusion that this glucan fraction is apparently unbound to an (N-acety1)glucosamine polymer was not substantiated by showing the absence of amino sugars in the extracted material. In all fungi investigated there was a fraction of glucan which resisted extraction with dimethyl sulfoxide and even extraction with 10 M NaOH at 100°C. In S.

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commune walls this fraction represented nearly all of the glucans insoluble in I M KOH at 60°C. However, after specific hydrolysis of the chitin chains present in these alkali-resistant residues, either with purified chitinase or with nitrous acid after deacetylation of chitin, the glucan became soluble partly in water, the rest in dilute alkali (Sietsma and Wessels, 1979, 1981). The released alkali-soluble glucan very much resembled a glucan also found in the culture medium, a (1+3)-P-glucan with single glucose branches attached by (1+6)-P linkages. The released water-soluble glucan was also (1+3)-@-linked but contained longer (1+6)-P-linked branches. These studies thus indicate that the insolubility of these glucan chains of moderate lengths is caused by linkage to the alkaliinsoluble (acety1)glucosamine-containing polymer. Amino acids, particularly lysine, appear to be involved in linking the two polymers together (Sietsma and Wessels, 1979). Apparently this linkage is stable in alkali but it is very labile to acid (Sietsma and Wessels, 1977). Acidic polysaccharides, although sometimes found in the other groups, are major constituents in the walls of zygomycetes (Bartnicki-Garcia and Reyes, 1968a,b; Miyazaki and Irino, 1970, 1971; Ballesta and Alexander, 1971; Datema et al., 1977a). A heteroglucoronan isolated with alkali from walls of M. rouxii (Bartnicki-Garcia and Reyes, 1968a,b) contained fucose, mannose, and glucuronic acid in a 2:3:5 ratio (mucoran). Another acidic polysaccharide resisted both alkali and acid extraction but could be solubilized by alkali after acid treatment. The water-insoluble polysaccharide extracted in this way (mucoric acid) was microcrystalline and contained mainly glucuronic acid. The X-ray diffraction pattern showed this substance to be identical to a substance previously demonstrated in acid-treated walls of P. blakesleeanus (Kreger, 1954, 1970). Although isolated as two distinct polymers the possibility that mucoran and mucoric acid were derived from a single polymer was not excluded (BartnickiGarcia and Reyes, 1968a). By depolymerizing the glucosamine-containing polymers with nitrous acid, Datema et al. (1 977a) found that this procedure solubilized from the wall of M . mucedo a single heteroglucuronan containing all the neutral and acidic sugars of the wall (fucose, mannose, galactose, and glucuronic acid in a molar ratio of 5: 1: 1:6). This water-soluble heteroglucuronan, apparently held insoluble in the wall by ionic bonds to the glucosamine-containing polymers, could also be extracted quantitatively from the wall by salt solutions of high ionic strength and partially by alkali. By treatment of the isolated watersoluble heteroglucuronan with 1 M HCl at 100°C it was partly converted into a water-insoluble crystalline compound containing only glucuronic acid with the properties of mucoric acid. This suggests that, at least in M. mucedo, mucoric acid, which can be extracted from the wall by alkali after acid treatment, is not a genuine wall component but arises by partial acid hydrolysis of a single heteroglucuronan and subsequent crystallization of acid-resistant homopolymeric stretches of glucoronic acid contained in this heteroglucuronan.

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B. STRUCTURE OF THE HYPHALWALL In order to reveal the molecular architecture of the hyphal wall electron microscopic observations combined with the use of more or less specific enzymic or chemical extractions have been made on the walls of a variety of fungi (lit. cit. in Wessels and Sietsma, 1981a). On the basis of such observations Hunsley and Burnett (1970) have modeled the wall as a coaxially layered structure. For example, going from the outside to the inside, the wall of N . CYUSSU is thought to contain the following layers (also see Burnett, 1979): ( I ) a layer of mixed a-and P-glucans, (2) a glycoprotein reticulum merging into a proteinaceous layer, (3) a distinct layer of protein, and (4)an innermost chitinous region in which chitin microfibrils are embedded in proteinaceous material. As Burnett (1979) has pointed out it should be understood that the coaxially arranged regions are not discrete but grade into each other. As pointed out by Wessels and Sietsma ( 198 la) the techniques used can easily lead to misinterpretations and they considered most published studies in agreement with a model of the wall in which the various wall components are more closely associated with each other forming essentially one layer with some components accumulating at the outside apparently forming extra layers. This simple model applies only to vegetative hyphae and not to the walls of specialized structures, such as spores and aerial hyphae, where genuine outer layers may be present. Figure 3 depicts a model of the hyphal wall of S. commune integrating the results of a number of chemical, enzymatic, and ultrastructural analyses (Wessels et a/., 1972; Sietsma and Wessels, 1977, 1979; van der Valk er al., 1977). In this case a water-soluble gel-like (1+3)-P/(1+6)-P-glucan and an alkali-soluble ( 1+3)-a-glucan (S-glucan) accumulate at the outside of a layer which contains the alkali-insoluble chitin-P-glucan complex. In this complex the glucan chains are ( 1+3)-P-linked with ( 1+6)-P-linked branches attached. In some of the (1+3)-P-linked chains the branches consist of just one glucose residue and these chains thus resemble the gel-like glucan accumulating at the outside of the hyphae. Other (1-3)-P-linked chains carry longer (1+6)-@linked glucan branches. Both types of branched glucans are thought to be attached to “chitin” chains through their reducing ends via amino acids. Probably these substituted chitin chains are hydrogen bonded to microfibrillar chitin. As mentioned earlier this chitin is only weakly crystalline. The presence of hydrogen-bonded triple helices among the glucan chains is inferred from the weak hydroglucan reflections seen in X-ray analyses of the chitin-glucan complex. Treatment of the complex with hot dilute acid breaks the linkages between chitin and glucan and leads to sharper X-ray reflections of hydroglucan and chitin. The model suggests that the excreted gel-like glucan may consist of glucan chains which have either escaped linkage to chitin or have been secondarily split

62

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M EDlUM

L

---G F

I-

plasma membrane2

C Y TO PLASM

FIG.3. Model of the mature hyphal wall of Schizophyllum commune. Partially crystallized chitin chains (A) are hydrogen bonded to chitin chains which carry covalently linked p-glucan chains. The coupling fragment (B) contains amino acids with a high proportion of lysine. The P-glucan chains are (1+3)-/3-linked and carry single (1+6)-P-linked glucose branches (C) or longer (1+6)-@-linked glucan branches (D) or are alternatively (1+3)-p- and (I+6)-P-linked (E). Some unsubstituted or sparsely branched (1+3)-P-glucan segments may form triple helices (H) which add to the strength of the glucan network. Crystalline (1+3)-a-glucan fibrils (alkali soluble) (F) occur throughout the wall and accumulate at the outer surface as a layer. Free water-soluble (1+3)-p-glucan chains with single (1+6)-p-linked glucose branches (G)are also present in the wall and may be excreted into the medium. (Adapted from Wessels and Sietsma, 1981b, with permission.)

off. It was found that the formation of the water-soluble glucan is inversely related to the formation of the alkali-insoluble glucan, depending on genotype and environmental conditions (Sietsma et al., 1977). In addition a small amount of water-insoluble but alkali-soluble p-glucan was found. It should also be noted that in other fungi often a large amount of wall bound p-glucan is found which is alkali insoluble but soluble in dimethyl sulfoxide and therefore possibly not covalently bound to chitin (Sietsma and Wessels, 1981). The most important evidence for postulating linkages between P-glucan chains and chitin is that the glucan chains become soluble in water or alkali after

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63

specific depolymerization of (acety1)glucosamine-containing polymers (Sietsma and Wessels, 1979). Both in regenerating protoplasts and growing hyphae watersoluble and alkali-soluble P-glucan chains were shown to be precursors for the alkali-insoluble glucan which could subsequently be released by depolymerization of chitin (Sonnenberg et af., 1982; Wessels er af., 1983). These precursor glucans are primarily pure ( 1+3)-P-glucans; ( 1-6)-P-linked residues seem to be attached at a later stage of generation of the complex (Sonnenberg ef al., 1982, 1985; Sietsrna et al., 1985). Importantly, in regenerating protoplasts in which the synthesis of chitin can be effectively blocked by polyoxin D this inhibition also leads to complete inhibition of the insolubilization of the glucan chains (Sonnenberg et al., 1982). The model given in Fig. 3 is necessarily speculative and may undergo modifications when other fungal walls are similarly analyzed. As it stands its most characteristic feature is that it predicts a rigid structure based on chitin microfibrils bonded together by P-glucan chains. In essence the model is similar to that proposed for primary walls of plants (Keegstra et al., 1973), although the nature of the polymers and the way they interact are very different. In the zygomycetes which do not contain glucan in their hyphal walls little is known about the location of polymers in the wall. One would expect, however, that the cationic polymers containing glucosamine and the anionic glucuronans and polyphosphates which together with chitin make up most of the wall occur in close association. Possibly in this case chitin microfibrils coated with partly deacetylated chitin chains are fixed into a rigid structure by their ionic interactions with the anionic polymers. In that case an enzymatic apparatus responsible for covalently linking P-glucan chains to chitin as possibly operating in basidiomycetes and ascomycetes could be simply replaced by a deacetylase in the zygomycetes (see Section V,B).

C. WALLSTRUCTURE AT

THE

APEX

It is clear that the tiny amoung of wall material present in growing apices does not significantly contribute to the mass of wall material used in chemical analyses. In fact, such apical wall material may be entirely lacking in broken hyphal wall preparations (Wessels et af., 1983). After autoradiography of pulse-labeled hyphae of S . commune labeled wall material over apices could be detected only before but not after breaking the hyphae for preparing wall preparations. After breaking the hyphae labeled glucan did not precipitate with the wall fragments at low-speed centrifugation while labeled chitin did precipitate but as fragments without recognizable form. However, after a chase, allowing for the labeled wall material to move in subapical direction, the labeled wall material stayed with the broken walls. Now also some tips, apparently arrested in growth at the moment of chasing the radioactivity, were labeled. These observations clearly indicate

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that growing and nongrowing hyphae differ in the type of wall material that covers their apices. The absence of alkali-insoluble P-glucan at the very apex of growing S . commune hyphae has been demonstrated by light microscopic autoradiography (Wessels et al., 1983). This same study showed that the chitin synthesized at the apex is insoluble in alkali. A subsequent study using electron microscopic autoradiography on shadowed preparations (Vermeulen and Wessels, 1984) revealed that the chitin in growing apices, though alkali insoluble, must be in a conformational state quite different from that in nongrowing apices and subapical parts. In contrast to the chitin in these older parts the newly synthesized chitin at apices appeared nonfibrillar, very susceptible to chitinase degradation and partly soluble in hot dilute mineral acid. There have been a few earlier observations indicating discontinuities in the presence of microfibrils at hyphal apices (Marchant, 1966; Strunk 1968) but these have been contradicted by many workers showing a continuous network of chitin microfibrils over the apex after chemical treatments that remove “matrix substance” (cf. Hunsley and Burnett, 1970; Bartnicki-Garcia, 1973; Schneider and Wardrop, 1979; Burnett, 1979). However, the study of Vermeulen and Wessels (1984) suggests that the chemical treatments used to visualize microfibrils could have removed chitin from growing apices so that there is the suspicion that images showing apical microfibrils represent nongrowing apices which abundantly occur in growing mycelia. After chemical treatments to visualize chitin microfibrils these were seen to increase in thickness and density from the very tip toward the base of the apex (Hunsley and Burnett, 1968; Burnett, 1979). This was interpreted as an increase in the number of “elementary fibrils” making up the microfibrils (Burnett, 1979). However, any explanation for this remarkable phenomenon should now also take into account the uncertainty of whether the examined tips were actually growing at the time of fixation. There are a number of light microscopic studies employing fluorescent probes which attest to the fact that the wall covering growing apices is different from that covering nongrowing apices and present in subapical regions. In these studies use was made of fluorescently labeled antibodies (Fultz and Sussman, 1966; Marchant and Smith, 1968; Hunsley and Kay, 1976), fluorescent brighteners such as calcofluor (Gull and Trinci, 1974), and fluorescently labeled wheat germ agglutinin (Galun et al., 1976). The two latter probes probably detect chitin. Differential staining at growing tips may be due to the absence of covering wall materials but also to a difference in the conformation of the polymers that bind these probes. For instance, it can be expected that probes for chitin, such as calcofluor or congo red or wheat germ lectin, would bind much better to noncrystalline chitin possibly present at growing apices than to microfibrillarordered chitin chains. In fact, substances, such as calcofluor and congo red, are

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known to inhibit crystallization of chitin and microfibril formation (Herth, 1980; Elorza et a/., 1983).

V. Wall Polymer Assembly at the Hyphal Apex A. AUTOLYSINS AND APICAL WALLGROWTH Autolysins are considered to be cell envelope-associated wall-lysing enzymes (Farkas, 1979; Rosenberger, 1979). As pointed out by Gooday (1978) a role for such enzymes in keeping the wall at the hyphal apex in a soft and extensible condition has been suggested nearly 100 years ago by Marshall-Ward. In essence, a rigid wall at the hyphal apex would only yield to turgor pressure if locally bonds in the wall polymers are broken. In a wall growth model involving apposition of new wall polymers this would mean that breaks are permanent and that the partially hydrolyzed polymers at the outer surface may even become soluble. Such a model has been of value in explaining wall growth in bacteria (Koch et a/., 1982; Tomasz, 1984) and plant cells (Cleland, 1981). In a wall growth model involving intussusception of new wall materials the breaks in the polymers would be temporary because after extension of the wall the gaps must be filled up with newly synthesized polymer. This is the essence of the unitary model of apical wall growth in fungi as formulated by Bartnicki-Garcia ( 1973). The wall at the hyphal apex is regarded as a rigid entity but growth is possible by the maintenance of a delicate balance of lysis and synthesis of wall polymers. More specifically the model assumes that a high turgor pressure forces broken microfibrils away from each other before synthesizing enzymes can insert new microfibrils or extend the broken ones. Amorphous wall material is thought to be delivered to the wall in vesicles and forced in between the fibrillar network by the turgor pressure. However, concomitant lysis and synthesis of these matrix polymers was not excluded. This concept has gained wide acceptance in the literature, although Bartnicki-Garcia (1973) has indicated that the evidence for participation of wall-lytic enzymes in the apical growth process is circumstantial. Until now no direct evidence has been forwarded and a critical examination of the arguments in favor of a role of lysins in apical growth therefore seems in place. One piece of evidence concerns the presence of autolytic enzymes in many wall preparations. The occurrence of the bulk of these enzymes in older, nongrowing portions of the wall (Polacheck and Rosenberger, 1978; Kritzman et al., 1978; Rosenberger, 1979; Hoch et a/., 1979) would seem to lessen the strength of this argument for a role of these enzymes in apical wall growth. In addition, these wall-bound enzymes [including chitinase, (1+3)-P-glucanase, P-N-acetylglucosaminidase,and P-glucosidase] are subject to catabolite repres-

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sion and their activities rise dramatically during carbon starvation (Polacheck and Rosenberger, 1978; Rosenberger, 1979). This feature is unprobable for enzymes involved in a delicately balanced growth process but more typical for extracellular enzymes involved in degradation of polymers for nutritive purposes (Reese, 1977). The presence of such enzymes tightly bound to older portions of the wall seems strange but could be due to entrapment of export vesicles in the forming wall since the enzymes can be released from the walls by cationic detergents (Polacheck and Rosenberger, 1978). Another piece of evidence forwarded in support of the involvement of lytic enzymes in apical wall growth is the often found correlation between the level of these enzymes and the process of branching or germination. It appears inescapable to infer the action of autolysins whenever a new hyphal apex must arise from a rigid wall area but the increased levels of wall-lytic enzymes detected in the mycelium or in the medium during increased branching can also be regarded as a consequence of the increase in the number of apices. Hydrolytic enzymes are probably excreted at the apices where the vesicles that package these enzymes fuse with the plasma membrane followed by diffusion of the enzymes through the wall into the medium. It has also been pointed out that the increased levels of lytic enzymes found upon branching may result from a lack of nutrients in dense highly branched mycelia leading to relief from catabolite repression (Polacheck and Rosenberger, 1978). It is a generally established fact that some lytic enzymes apparently produced for nutritive purposes are able to attack walls or wall components from the same fungus (Wessels and Sietsma, 1981a). For example, it is possible to liberate protoplasts from T. harziunum by a concentrate of the hydrolytic enzymes excreted into the medium by the organism itself (de Vries and Wessels, 1973). Yet this would not justify implicating these enzymes in normal wall metabolism at the apex. The fact that the first protoplasts are released from apices indicates that the wall is most sensitive in this area. Also by assaying degradation in wall preparations after briefly labeling of wall components it was shown that newly synthesized wall material is most easily attacked by lytic enzymes (Polacheck and Rosenberger, 1975, 1977; Fkvre, 1977; Vermeulen and Wessels, 1984). There thus remains the problem of why potentially lytic enzymes passing the wall at the apex do not destroy the wall in this area. Part of the answer may be that the concentration of the enzymes as they pass the wall may be relatively low and that the susceptible polymers at the apex are quickly integrated in a crosslinked and partially crystalline wall structure where they are much less susceptible to degradation. Moreover, if the wall at the apex were protected against high turgor pressure by a cytoskeleton, as discussed in Section III,B, occasional cuts in the polymers at the apex would also be of little consequence for the maintenance of apical integrity. The bursting tendency of hyphal tips after flooding with water or solutions of

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acids, chelating substances, salts, etc. was construed as a main argument for the concept of a balance between lysis and synthesis during apical wall growth by Bartnicki-Garcia and Lippman (1972b). They noted that the temperature coefficient of bursting in water was 1.3-2.0 and concluded that bursting thus could not be considered as a purely osmotic phenomenon but that the participation of a biochemical reaction, as a rate-limiting step, was indicated. They also argued that the fact that various substances could trigger bursting was difficult to reconcile with a direct effect of these substances on the apical wall but that bursting was more likely to be associated with interference with wall metabolism. Specific interference with chitin synthesis using polyoxin D also induced bursting (Bartnicki-Garciaand Lippman, 1972a). If lysis and synthesis were an integral part of wall formation at the apex any disturbance tipping the balance toward decreased synthesis or increased lysis would lead to disintegration of the wall fabric at the apex. Recent support for the concept of lysis as an integral part of wall growth at the apex has come from studies showing a close association of chitinase with chitin synthase preparations (Lopex-Romero et al., 1982; Humphreys and Gooday, 1984) and an increase in reducing ends in chitin during light-stimulated growth in stage I sporangiospores of Phycornyces (Herrera-Estrella and Ruiz Herrera, 1983). It must be admitted that all these pieces of evidence are very indirect and alternative explanations are possible. As discussed earlier bursting of hyphae is mostly subapical and some of the substances used to induce bursting may have influenced the integrity of the cytoskeleton underlying the apex. Inhibition of synthesis of one of the principal wall polymers could lead to the assembly of a wall of insufficient strength to withstand turgor pressure in subapical parts and would result in bursting of the wall in this region. Studying the antagonism of Ca2+ and acid in bursting of hyphal tips, Dow and Rubery (1975) came to the conclusion that the assumed role of lysins could not explain their results. Also the apparent absence of significant wall turnover in growing walls (Rosenberger, 1979) does not support the activity of lysins in apical wall growth. In addition, no conceptual framework has been advanced to understand how a balance between lysis and synthesis of the wall at the growing apex can be maintained. First, for lysis and concomitant synthesis to account for intussusception of wall material one must assume that the enzymes for both processes operate within the wall fabric. For lytic enzymes this presents no problem. For the synthetic enzymes, however, most of the evidence points to a cytoplasmic localization, e.g., the plasma membrane in the case of chitin synthase (Section 111,C). Second, the activities of both enzyme systems should be precisely balanced with the magnitude of intussusception which should decrease when a wall segment moves from the extreme apex to subapical parts. For synthesizing enzymes located in the plasma membrane mechanisms originating from the cytoplasm can be envisaged to account for their regulation. This is much more difficult to envisage for stable

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lytic enzymes operating within the wall away from the plasma membrane. Humphreys and Gooday (1984) have suggested that a chitinase and a chitin synthase, both present in the plasma membrane, could perform the delicate balance required. However, growth of a rigid wall cannot proceed unless cuts are produced in at least the outer zone of the wall which would require the lysidsynthase complex to move away from the plasma membrane. Control of the activity of wall-lytic enzymes has also presented a problem in explaining intercalary growth of plant cells and bacteria in which extension of the wall probably does involve the continuous breakage of bonds. However, in these cases it is generally thought that new wall material is formed by apposition and it has been suggested, both for plants (Cleland, 1967) and bacteria (Koch et al., 1982), that the lysins act only in the outer zone of the wall on polymers under stress having an extended conformation. This would allow for breakage of bonds in the stretched older portion of the wall while such bonds would not be subject to hydrolysis in the not yet stretched newly deposited wall polymers. But even if apposition of wall material is also assumed at the hyphal apex, it is still difficult to see how the prevailing stress pattern in the apical wall could lead to proper control of lysis according to this hypothetical mechanisms. It is clear that wall-lytic enzymes play important roles in various aspects of the life of fungi. These included net degradation of wall polymers during starvation, degradation of septa for nuclear migration, hyphal fusions, branch initiation, and germination. As discussed by Wessels and Sietsma (1981a) the key enzymes involved need not be identical with the commonly assayed enzymes using artificial substrates. A few specific breaks in the complicated network of the wall structure by special enzymes may be sufficient to weaken the wall sufficiently or to initiate complete degradation by more general hydrolytic enzymes. As may be evident from the foregoing discussion the participation of lytic enzymes in apical wall growth can at least be doubted. Probably the most fruitful route to clarify this issue would be a search for conditional mutants blocked in apical extension. This route has been taken to assess the role of autolysins in bacterial growth (Tomasz, 1984) and has been started for yeasts (Nombela and Santamaria, 1984). IN B. ASSEMBLY

THE

WALL:A STEADY-STATE MODEL

Only if the wall at the hyphal apex is considered inherently rigid is it necessary to invoke the continuous action of lysins to keep the wall plastic and to allow for the insertion of new wall material. However, the new views on fungal wall structure (Section IV) in which the polymers are cross-linked by various bonds make it highly unlikely that this structure is built in one step. At least two steps can be envisaged. First the individual polymers must be deposited outside the plasma membrane, either by synthesis on the plasma membrane or by extrusion

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through vesicles that fuse with the plasma membrane. Second, extracellularly the individual polymers may become modified, undergo partial crystallization, and mutual interactions involving covalent or ionogenic bonds. This would imply that a major part of the assembly of the wall structure occurs in the domain of the wall itself. Because of the macromolecular nature of the assembly system it may take considerable time before the structure has reached its final state and it would seem plausible that during this time the mechanical properties of the wall gradually change from a viscoelastic fluid to a rigid composite. The whole process may be likened to that occurring in man-made composites in which by varying the amount of cross-linking between polymers a wide range of plastic and strength properties can be obtained. Cross-linking reduces crystallinity but the additional stiffening effect of an increasing degree of cross-linking between chains more than counterbalances the increasing flexibility due to loss of crystallinity (Kaufman and Falcetta, 1977). Because fungal walls vary widely in the types and amount of the wall polymers they contain it can be expected that there will also be variation in the types of interactions that generate the final rigid wall structure. Because the view on wall biogenesis as outlined above is based mainly on our studies on the structure of the wall of S. commune (Fig. 3 ) and the processes that lead to its construction, this work will be briefly summarized. For the present discussion we ignore the large amount of crystalline (1+3)-aglucan present in the wall of S. commune because this polymer is probably of little morphogenetic significance. Mutations that block the synthesis of this polymer in A . nidulans (Zonneveld, 1974; Polacheck and Rosenberger, 1977) or 2-deoxyglucose that inhibits its synthesis in S . commune (J. H. Sietsma and J. G. H. Wessels, unpublished) do not have a major effect on hyphal morphology. However, this a-glucan may become the major shape-maintaining wall polymer in established hyphae under conditions that most of the P-glucan and chitin have been removed from the walls by autolytic processes (Wessels and Sietsma, 1979). The chitin-P-glucan complex depicted in Fig. 3 is considered to be of major importance in determining the final shape of the hyphae. Interference with its synthesis by congo red or calcofluor white causes the formation of balloonlike structures from growing apices (C. A. Vermeulen and J. G. H. Wessels, unpublished). The proposed structure of the chitin-P-glucan complex immediately suggests that its synthesis must occur in at least two distinct phases. First the individual chitin and P-glucan chains must be polymerized and deposited outside the plasma membrane. Only then can enzymes in the wall proceed to couple the pglucan chains to chitin probably competing with the tendency of the chitin chains to crystallize. The following observations support to occurrence of the suggested sequence of events (Fig. 4). Not only in regenerating protoplasts (Sonnenberg et al., 1982), where initially

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A

B

I

1'

FIG.4. Highly schematic diagram of possible changes in wall structure in the growing hyphal apex of Schizophyllum commune. Increase in stipling in C indicates degree of cross-linking and crystallizationof p-glucan and chitin chains, paralleled by an increase in rigidity. A shows the mature wall structure in which chitin chains probably assembled in microfibrils are cross-linked to partly crystallized (1+3)-p-glucan chains with (1+6)-p-linked branches attached. B shows individual chitin chains (straight lines) and individual (1+3)-p-glucan chains (wavy lines) as present at the extreme apex. (Adapted from Wessels, 1984, with permission.)

no alkali-insoluble P-glucan is formed, but also in growing hyphae (Wessels et al., 1983; Sonnenberg et al., 1985) pulse-chase experiments with [ *4C]glucose have shown that the precursor for the alkali-insoluble P-glucan is a water-soluble/alkali-soluble ( 1+3)+-glucan. The conversion to the alkali-insoluble form is interpreted as being due to linkage of the glucan chains to chitin because they become soluble again after specific depolymerization of chitin. The conversion to the alkali-insoluble form is a relatively slow process, taking several minutes. By light microscopic autoradiography it could be shown (Wessels ef al., 1983) that both the chitin and glucan chains are synthesized at the apex according to the expected gradient but only label from N-a~etyl-[~H]glucosamine immediately appears in an alkali-insoluble form (chitin). Label from [3H]glucoseis primarily incorporated at the apex into water- and alkali-soluble glucans; at the extreme apex alkali-insolubleglucan is completely missing. While the glucans move in a subapical direction during growth in the chase period, alkali-insoluble glucan appears at the expense of the water-soluble glucan. Like the precursor glucan the alkali-insoluble glucan formed at the apex is mainly (1+3)-P-linked with few, if any, (1-6)-P-linked glucan branches attached (Sietsma et al., 1985; Sonnenberg et al., 1985). But in a subapical direction the number of (1+6)-P-linked glucose residues rises rapidly in the alkali-insoluble glucan. Possibly this is another type of modification which occurs within the domain of the wall. Also the conformational state of the chitin changes while moving from its site of synthesis in subapical direction (Vermeulen and Wessels, 1984). Apparently the

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chitin at the growing apex is not yet crystalline since no microfibrils can be observed and the chitin in this area is extremely sensitive to dissolution by chitinase or hot dilute mineral acid. This contrasts with the conformational state of chitin in subapical parts and indicates a time gap between polymerization of the chitin chains and their final integration into the wall structure. The existence of a gap between polymerization and microfibril formation has been suggested by using congo red or calcofluor white in organisms which form either P-chitin (Herth, 1980) or a-chitin (Elorza et al., 1983). In addition it has been shown that both nascent chitin (Molano et al., 1979; Lopez-Romero et al., 1982) and nascent (1+3)-P-glucan (Pkrez et al., 1984) synthesized in v i m are much more sensitive to enzymic degradation than some time after their formation, again suggesting a conformadonal change with time. When hyphae of S . commune stop growing the alkali-insoluble form of glucan is produced over the whole apex (Wessels er al., 1983) indicating that the process of insolubilization of the glucan (interpreted as linkage of the P-glucan chains to chitin) proceeds with time independent of elongation of the hypha. Also the (1+6)+-linked glucan chains are now formed in the wall at the apex (Sietsma et al., 1985) and chitin microfibrils become visible in this area after treatment with hot dilute mineral acid (Vermeulen and Wessels, 1984). In other words, after elongation stops the wall at the apex assumes the same structural features as observed in the subapical wall. In essence, the scheme suggested above may be applicable to basidiomycetes and ascomycetes in general but details may be different. In particular, the presence of large amounts of alkali-insoluble glucan possibly not linked to chitin in a number of species (Sietsma and Wessels, 1981) has not been accounted for in the above scheme. It is tempting to speculate that in the zygomycetes the role of the chitin-glucan linkages is played by the ionic linkages between partially deacetylated chitin and glucoronans. Similar to the solubilization of P-glucans by depolymerization of chitin (Sietsma and Wessels, 1979), specific depolymerization of the partially deacetylated chitin chains in zygomycetes causes the solubilization of glucuronans (Datema et al., 1977a). Deacetylation of chitin seems to occur by a deacetylase in the wall shortly after synthesis of the chitin chains before crystallization has occurred (Davis and Bartnicki-Garcia, 1984). Assuming hydrogen bonding between partially deacetylated chitin and chitin microfibrils, the generation of the cationic groups by the deacetylase at the apex could lead to cross-linking of the microfibrils by acidic glucuranans which may be hydrogen bonded among themselves. Thus the process of deacetylation of chitin at the apex could also lead to a significant change in the viscoelastic properties of the wall in these fungi. The model of apical wall growth developed above is a steady-state model because the maintenance of elongation depends on the steady delivery to the apex of individual wall components which then interact in the wall to form a rigid

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structure while moving subapically. Any influence stopping elongation would lead to disturbance of the steady-state process and lead to rigidification of the wall over the apex. This would explain why a hyphal apex arrested in growth for a critical time can no longer resume growth (Reinhardt, 1892; Robertson, 1958). Assuming that the time required for rigidification is not dependent on the elongation rate, the model could also easily explain the positive relationship often found between elongation rate and length and width of the extension zone (Reinhardt, 1892; Steele and Trinci, 1975). The observed random orientation of microfibrils in subapical walls and (nongrowing) apices (Burnett, 1979) would present no problem since no realignment of fibrils has to occur. Finally, the deviating wall structure at growing apices may allow for the passage of large-sized proteins intended for export (Chang and Trevithick, 1974).

VI. Summary Although apical growth of hyphae is the most prominent feature of the organisms belonging to the fungal kingdom, we are still largely ignorant of the mechanisms by which it occurs. However, as this review may have shown, many new analyses and observations as well as comparisons with other cellular systems now open new routes for ideas and experimentation. Understanding the mechanisms involved in apical hyphal growth would not only clarify a longstanding morphogenetic problem, namely, the generation of a tubular cell under turgor pressure by elongation at one end. Understanding these mechanisms would also give a better insight in a number of characteristic features which dominate the life of fungi, such as the colonizing ability of the mycelium, the way in which environmental factors influence growth and morphogenesis, the interactions occurring between hyphae, and the interactions between fungi and other organisms in parasitic and mutually beneficial relationships. It appears that the hyphal apex is best viewed as a highly polarized system of exocytosis. Wall materials, extracellular enzymes, and probably other substances are excreted at the growing end of a tubular cell. The most obvious cellular features that accompany this polarized system are the unidirectional flow of vesicles in the cytoplasm fusing with the plasma membrane at the apex, the gradients in wall synthesis at the apex, and the cytoplasmic gradients in ion distribution that are maintained at the apex. New microscopic techniques are also beginning to reveal a cytoskeletal organization of the cytoplasm at the apex which may be crucial to its polarized activity. How these various aspects are causally related can only be a matter of much speculation at the moment. It is clear that once an apical growth center is initiated it is self-maintained. It can be surmised that the distribution of certain proteins in the plasma membrane is of prime importance in this respect. For instance, the suspected spatial distribution

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of proton pumps in the hyphal plasma membrane creates proton currents which in turn may cause the maintenance of the spatial distribution of proton pumps and possibly other proteins in the plasma membrane. The apical gradients in protons and other ions such as Ca2+ could also be instrumental in organizing the cytoskeleton at the apex and allow for the proper interactions of the vesicles with the cytoskeleton to effect their transport to the extreme apex and subsequent fusion with the plasma membrane. Both the polarized distribution of proteins in the plasma membrane and the polarized delivery of vesicles are likely to play a role in the gradient of wall-synthesizing activity generated at the apex. Because of the high turgor pressure the tubular form of the hypha can be maintained only by a rigid wall generated at the apex. It is not certain, however, that the full turgor pressure also acts on the wall at the apex because of the cytoskeletal organization in this region. With regard to the type of wall synthesized at the apex fungi exhibit wide variations. It is clear, however, that a fruitful approach to the study of wall biosynthesis can be expected only after sufficient knowledge is obtained about the details of the structure of a particular wall. In fact, such a structural analysis can reveal clues as to possible mechanisms involved in the synthesis of the wall which then hopefully can be generalized. Growth of the wall at the hyphal apex requires that the wall in this region has plastic properties which contrasts with the requirement of rigidity elsewhere in the hypha. A widely held view involves the participation of wall-lytic enzymes in plasticizing the wall at the apex and in allowing new wall material to be inserted. A critical evaluation of the evidence presented to support this view makes this hypothesis less attractive. As an alternative a steady-state model is discussed based on recent observations in the author’s laboratory. In essence this model holds that the assemblage of polymers synthesized at the apex is inherently plastic. However, this assemblage develops rigidity by interactions, in the wall, between and among the various individual polymers present while the wall segment moves in subapical directions during elongation. This model seems to fit many of the original observations made on living hyphae.

REFERENCES Ahlquist, C. N . . and Gamov, R . 1. (1973). Plarif fhvsio/. 31, 586-587. Albersheim, P., Darvill, A . G . , Davis, K . R . , Lau, J . M., McNeil, M.. Sharp, J . K . , and York, W. S.(1983). In “Structure, Function, and Biosynthesis of Plant Cell Walls” (W. M. Dugger and S. Bartnicki-Garcia, eds.), pp. 19-51. Waverly Press, Baltimore, Maryland. Allen, E. D., and Sussman, A. S. (1978). J . Bacferiol. 135, 713-716. Allen, E. D..Aiuto. R . , and Sussman, A. S. (1980). Protoplasma 102, 63-75. Araki, Y..and Ito. E. (1975). Eur. J . Biochem. 55, 71-78. Armbruster, B . L . . and Weisenseel, M. H. (1983). Proropkusma 115, 65-69. Bacic, A . , and Delmer. D. P. (1981). fluma 152, 346-351.

74

J. G. H. WESSELS

Ballesta, J.-P. G., and Alexander, M. (1971). J. Bacreriol. 106, 938-945. Bartnicki-Garcia, S. (1968). Annu. Rev. Microbiol. 22, 87-108. Bartnicki-Garcia, S. (1973). Symp. SOC. Gen. Microbiol, 23rd., pp. 245-267. Bartnicki-Garcia, S . , and Bracker, C. E. (1984). I n “Microbial Cell Wall Synthesis and Autolysis” (C. Nombela, ed.), pp. 101-1 12. Elsevier, Amsterdam. Bartnicki-Garcia, S . , and Lippman, E. (1969). Science 165, 302-304. Bartnicki-Garcia, S., and Lippman, E. (1972a). J . Gen. Microbiol. 71, 301-309. Bartnicki-Garcia, S . , and Lippman, E. (1972b). J . Gen. Microbiol. 73, 487-500. Bartnicki-Garcia, S . , and Lipprnan, E. (1982). Handb. Microbiol. 4, 229-252. Bartnicki-Garcia, S., and Nickerson, W. J. (1962). Biochim. Biophys. Acta 58, 102-119. Bartnicki-Garcia, S . , and Reyes, E. (1968a). Biochim. Biophys. Acfa 165, 32-42. Bartnicki-Garcia, S . , and Reyes, E. (1968b). Biochim. Biophys. Acra 170, 54-62. Bartnicki-Garcia, S., Bracker, C. E., Reyes, E., and Ruiz-Herrera, J. (1978). Exp. Mycol. 2, 173192. Betina, V., Micekovfi, D., and Nemec, P. (1972). J . Gen. Microbiol. 71, 343-349. Bracker, C. E. (1967). Annu. Rev. Phytoparhol. 5 , 343-374. Bracker, C. E., Ruiz-Herrera, J., and Bartnicki-Garcia, S. (1976). Proc. Narl. Acad. Sci. U.S.A. 73, 4570-4574. Braun, P. C., and Calderone, R. A. (1978). J. Bacreriol. 133, 1471-1477. Buck, K. W., Chen, A. W., Dickerson, A. G., and Chain, E. B. (1968). J. Gen. Microbiol. 51, 337-352. Burgeff, H. (1915). Flora, N . F. 108, 353-488. Burnett, J. H. (1979). I n “Fungal Walls and Hyphal Growth” (J. H. Bumett and A. P. J. Trinci, eds.), pp. 2-25. Cambridge Univ. Press, London and New York. Cabib, E., Roberts, R., and Bowers, B. (1982). Annu. Rev. Biochem. 51, 763-793. Cabib, E., Bowers, B., and Roberts, R. L. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 3318-3321. Campos-Takaki, G. M., Dietrich, S. M. C., and Mascarenhas, Y. (1982). J. Gen. Microbiol. 128, 207-209. Castle, E. S . (1958). J . Gen. Physiol. 41, 913-926. Chang, P. Y. L., and Trevithick, J. R. (1974). Arch. Microbiol. 101, 281-293. Cheung, W. Y. (1982). Fed. Proc. Fed. Am. SOC. Exp. Biol. 41, 2253-2257. Cleland, R. (1967). Planra 77, 182-191. Cleland, R. E. (1981). Encycl. Plant Physiol. New Ser. 136, 255-273. Collinge, A. I., and Trinci, A. P. J. (1974). Arch. Microbiol. 99, 353-368. Crawford. R. L. (1981). “Lignin Biodegradation and Transformation.” Wiley, New York. Dalley, N. E., and Sonneborn, D. R. (1982). Biochim. Biophys. Acra 686, 65-76. Dargent, R. (1975). C. R. Acad. Sci. Ser. D. 282, 1601-1604. Da Riva Ricci, D., and Kendrick, B. (1972). Can. J . Bor. 50, 2455-2462. Daterna, R., Ende, H. van den., and Wessels, J. G. H. (1977a). Eur. J . Biochem. 80, 611-619. Daterna, R., Wessels, J. G. H., and Ende, H. van den. (1977b). Eur. J. Biochem. 80, 621-626. Davidse, L. C. (1975). I n “Microtubules and Microtubule Inhibitors” (M. Borgers and M. de Brabander, eds.), pp. 483-495. North-Holland Publ., Amsterdam. Davidse, L. C., and Flach, W. (1977). J . Cell Biol. 72, 174-193. Davis, L. L., and Bartnicki-Garcia, S. (1984). Biochemisrry 23, 1065-1073. Delmer, D. P., Benziman, M., and Padan, E. (1982). Proc. Narl. Acad. Sci. U.S.A.79,5282-5286. Derksen, I., and Traas, J. A. (1985). In “Sexual Reproduction in Seed Plants, Ferns and Mosses’’ (M. T. M. Willemse and J. H. van Went, eds.), pp. 64-67. Pudoc, Wageningen. De Vries, 0. M. H., and Wessels, J. G. H. (1973). J . Microbiol. Serol. 39, 397-400. De Vries, 0. M. H., and Wessels, J. G. H. (1975). Arch. Microbiol. 102, 209-218. De Vries, S . C., and Wessels, J. G. H. (1982). Exp. Mycol. 6, 95-98.

CELL WALL SYNTHESIS IN APICAL HYPHAL GROWTH

75

De Wolff, P. M., and Houwink, A. L. (1954). Acfa Bor. Neerl. 3, 369-397. Dickinson, C. H., and Lucas, J. A. (1982). “Plant Pathology and Plant Pathogens,” Basic Microbiology, Vol. 6. Blackwell, Oxford. Dow, J. M., and Rubery, P. M. (1975). J. Gen. Microbiol. 91, 425-428. Duran, A., and Cabib, E. (1978). J . Biol. Chem. 253, 4419-4425. Durhn, A., Bowers, B., and Cabib, E. (1975). Proc. Narl. Acud. Sci. U.S.A. 72, 3952-3955. Dustin, P. (1978). “Microtubules.” Springer-Verlag, Berlin and New York. Elorza, M. V., Rico, H., and Sentandreu, R. (1983). J. Gen. Microbiol. 129, 1577-1582. Farkas, V. (1979). Microbiol. Rev. 43, 117-144. Rvre, M. (1977). J. Gen. Microbiol. 103, 287-295. Wvre, M. (1979).In “Fungal Walls and Hyphal Growth” (J. H. Burnett and A. P. J. Trinci, eds.), pp. 225-263. Cambridge Univ. Press, London and New York. Rvre, M. (1983). PIanra 159, 130-155. Fkvre, M., and Rougier, M. (1982). Arch. Microbiol. 131, 212-215. Flanagan, M. D., and Lin, S. (1980). J. Biol. Chem. 255, 835-838. Fleet, G . H., and Phaff, H. J. (1981). Encycl. PIanr Physiol. New Ser. 13, 416-440. Franke, W. W., Herth, W., Van der Woude, J., and M o d , D. J. (1972). Planra 105, 317-341. Fultz, S. A., and Sussman, A. S. (1966). Science 152, 785-786. Galpin, M. F. J., Jennings, D. H., Oates. K., and Hobot, J. (1978). Exp. Mycol. 2, 258-269. Galun, M., Braun, A., Frensdorff, A., and Galun, E. (1976). Arch. Microbiol. 108, 9-16. Garnbino, J . , Bergen, L. C., and Morris, N. R. (1984). J. Cell Biol. 99, 830-838. Girbardt, M. (1957). Planfa 50, 47-59. Girbardt, M. (1968). Symp. SOC.Exp. Biol. 12, 249-259. Girbardt, M. (1969). Proroplasma 67, 413-441. Girbardt, M. (1979). Exp. Mycol. 3, 215-228. Goffeau, A., and Slayman, C. W. (1981). Eiochim. Biophys. Acra 639, 197-223. Comes, S. L., Mennuci, L., and Dac-Maia, J. C. (1979). FEBS Len. 99, 39-42. Gooday, G. W. (1971). J. Gen. Microbiol. 67, 125-133. Gooday, G. W. (1975). In “Primitive Sensory and Communication Systems” (M. G. Carlile, ed.). pp. 155-204. Academic Press, New York. Gooday, G. (1978). In “The Filamentous Fungi” (J. E. Smith and D. R. Berry, eds.), Vol. 3, pp. 51-77. Arnold, London. Gooday, G. W. (1982).In “Basidium and Basidiocarp” (K. Wells and E. K. Wells, eds.), pp. 158173. Springer-Verlag. Berlin and New York. Gooday, G. W., and Trinci, A. P. J. (1980). SOC. Gen. Microbiol. Symp. 30, 207-251. Gorin, P. A. J., and Spencer, J. F. T. (1968). Adv. Carbohydr. Chem. 23, 367-417. Cow, N. A . R. (1984). J. Gen. Microbiol. 130, 3313-3318. Cow, N. A. R., and Gooday, G. W. (1983). Proroplasma 115, 62-58. Cow, N. A. R., Kropf, D. L., and Harold, F. M. (1984). J . Gen. Microbiol. 130, 2967-2974. Grand, R. J. A., Nairu, A. C., and Perry, S. V. (1980). Biochem. J . 185, 755-760. Green, P. B. (1974). Brookhaven Symp. Biol. 25, 166-190. Grove, S . N. (1978).In “The Filamentous Fungi” (J. E. Smith and D. R. Berry, eds.), Vol. 111, pp. 28-50. Arnold, London. Grove, S. N., and Bracker, C. E. (1970). J . Bacreriol. 104, 989-1009. Grove, S . N., and Sweigard, J. A. (1980). Exp. Mycol. 4, 239-250. Gull, K., and Trinci, A. P. J. (1974). Arch. Microbiol. 96, 53-57. Hanson, B., and Brody, S . (1979). J. Bacreriol. 138, 461-466. Harley, J. L., and Smith, S. E., eds. (1983). “Mycorrhizal Symbiosis.” Academic Press, New York. Harold, F. M. (1977). Annu. Rev. Microbiol. 31, 181-203.

76

J. G.H. WESSELS

Heath, 1. B. (1978). “Nuclear Division in the Fungi.” Academic Press, New York. Hernandez, J., Lopez-Romero, E., Cerbon, J., and Ruiz-Herrera, J. (1981). Exp. Mycol. 5, 349356. Herr, F. B., and Heath, M. C. (1982). Exp. Mycol. 6, 15-24. Herrera-Estrella, L., and Ruiz-Herrera, J. (1983). Exp. Mycol. 7, 362-269. Herth, W. (1978). Protoplasma 96, 275-282. Herth, W. (1980). J. Cell Biol. 87, 442-450. Herth, W., Franke, W. W., and Van der Woude, W. J. (1972). Narurwissenschajien 59, 28-39. Hill, J. W.. and Mullins, J. T. (1980). Can. J. Microbiol. 26, 1132-1 140. Hoch, H. C., and Howard, R. J. (1980). Protoplasma 103, 281-297. Hoch, H. C., and Staples, R. C. (1983). Eur. J . CellBiol. 32, 52-58. Hoch, H. C., and Staples, R. C. (1985). Protoplasma 124, 112-122. Hoch, H. C., Hanssler, G.,and Reisener, H.-J. (1979). Exp. Mycol. 3, 164-173. Hohn, T. M . , Lovett, I. S., and Bracker, C. E. (1984). J . Bacteriol. 158, 253-263. Honvitz, B. A., Weisenseel, M. H., Dorn, A , , and Gressel, J. (1984). Planrfhysiol. 74,912-916. Houwink, A. L., and Kreger, D. R. (1953).Antonie van Leeuwenhoek. J . Microbiol. Serol. 19, I 24. Howard, R. J. (1981). J. CellSci. 48, 89-103. Howard, R. J., and Aist, 1. R. (1977). Protoplasma 92, 195-210. Howard, R. J., and Aist, J. R. (1979). J. Ultrasrruct. Res. 66, 224-234. Howard, R . J., and Aist, J. R. (1980). J. Cell B i d . 87, 55-64. Humphreys, A. M., and Gooday, G.W. (1984). J . Gen. Microbiol. 130, 1359-1366. Hunsley, D., and Burnett, J. H. (1968). Nature (London) 218, 462-463. Hunsley, D., and Burnett, J. H. (1970). J. Gen. Microbiol. 62, 203-218. Hunsley, D., and Gooday, G. W. (1974). Protoplusma 82, 125-146. Hunsley, D., and Kay, D. (1976). J. Gen. Microbiol. 95, 233-248. Issac, S., Ryder, N. S., and Peberdy, J. F. (1978). J. Gen. Microbiol. 105, 45-50. Jaffe, L. F. (1977). Narure (London) 265, 600-602. Jaffe, L. F., and Nuccitelli, R. (1974). J. Cell Biol. 63, 614-628. Jaffe, L. F., and Nuccitelli, R. (1977). Annu. Rev. Biophys. Bioeng. 6, 445-476. Jaffe, L. F., Robinson, K. R., and Nuccitelli, R. (1974). Ann. N . Y . Acad. Sci. 238, 372-389. Jelsma, J., and Kreger, D. R. (1975). Carbohydr. Res. 43, 200-203. Jennings, D. H. (1979). In “Fungal Walls and Hyphal Growth” (J. H. Burnett and A. P. J. Trinci, eds.), pp. 279-294. Cambridge Univ. Press, London and New York. Katz, D., and Rosenberger, R. F. (1971). J . Bacteriol. 108, 184-190. Kaufman, H. S., and Falcetta, J. J. (1977). “Polymer Science and Technology.” Wiley, New York. Keegstra, K., Talmadge, K. W., Bauer, W. D., and Albersheim, P. (1973). Planrfhvsiol. 51, 188196. Kerkenaar, A., and Barug, D. (1984). Pestic. Sci. 15, 199-205. Kilmartin, J., and Fogg, J. (1982). In “Microtubules in Micro-organisms” (P. Cappuccinelli and N. R. Moms, eds.), pp. 157-170. Dekker, New York. Kitazima, Y., Banno, Y., Noguchi, T., Nozawa, Y., and Ito, Y. (1972).Arch. Biochem. Biophys. 152, 811-820. Koch, A. L. (1982). J. Gen. Microbiol. 128, 947-951. Koch, A. L., Higgins, M. L., and Doyle, R. I. (1982). J. Gen. Microbiol. 128, 927-945. Kosuge, T. (1981). Encycl. Planr Physiol. New Ser. 13, 584-623. Kreger, D. R. (1954). Biochim. Biophys. Acra 13, 1-9. Kreger, D. R. (1970). Narure (London) 227, 81-82. Kritzman, G.,Chet, I., and Henis, Y. (1978). J . Bacreriol. 134, 470-475. Kropf, D. L., Lupa, M. D. A., Caldwell, J. H., and Harold, F. M. (1983).Science 220, 1385-1387.

CELL WALL SYNTHESIS IN APICAL HYPHAL GROWTH

77

Lemke, P. A . (1979). “Viruses and Plasmids in Fungi.” Dekker, New York. Lloyd, C. W. (1984). In!. Rev. Cvtol. 86, 1-51. Lopez-Romero, E. Ruiz-Herrera, J., and Bartnicki-Garcia, S. (1982).Biochim. Biophys. Acta 702, 233-236. McClure, W. K.,Park, D., and Robinson. P. M. (1968). J. Gen. Microbiol. 50, 177-182. Maclean-Fletcher, S.,and Pollard, T. D. (1980). Cell 20, 329-341. Marchant, R. (1966). Ann. Bot. 30, 821-830. Marchant, R., and Smith. D. G. (1968). Arch. Mikrobiol. 63, 85-94. Marchessault, R. H.,Deslandes, Y., Ogawa, K., and Sundararajan, P. R. (1977). Can. J. Chem. 55, 300-303. Marichal, P., Gorrens, J., and Van den Bossche, H. (1984). J. Med. Vet. Mycol. 22, 13-21. Mills, G. L., and Cantino, E. C. (1981). Arch. Microbid. 130, 72-77. Misaghi. 1. J. (1982). “Physiology and Biochemistry of Plant-Pathogen Interactions.” Plenum, New York. Miyazaki, T., and Irino, T. (1970). Chem. Pharm. Bull. IS, 1930-1934. Miyazaki, T., and Irino, T. (1971). Chem. Pharm. Bull. 19, 2545-2549. Mizukami, M., and Wada, S. (1983). Protoplasma 114, 151-162. Molano, J., Polacheck, 1.. DurBn, A , , and Cabib, E. (1979). J. Biol. Chem. 254, 4901-4907. Mollenhauer, H. H., and Morrk, D. I. (1976). Protoplasma 87, 39-48. Montgomery, G. W. G., and Gooday, G. W. (1985). FEMS Microbiol. Lett. 27, 29-33. Muthukumar, G., Kulkarni, R. K., and Nickerson, K. W. ( I 985). J. Bacteriol. 162, 47-49. Muzzarelli, R . R. A. (1977). “Chitin.” Pergamon. Oxford. Nicholson, T. H. (1975). In “Endomycorrhizas” (F. E. Sanders, B. Mosse, and P. B. Tinker, eds.), pp. 25-34. Academic Press, New York. Nolan, R. A,. and Bal, A. K. (1974). J. Bacteriol. 117, 840-843. Nombela, C., and Satamaria, C. (1984). In “Microbial Cell Wall Synthesis and Autolysis” (C. Nombela, ed. 1, pp. 249-259. Elsevier. Amsterdam. Northcote, D. H. (1984). In “Structure, Function, and Biosynthesis of Plant Cell Walls” (W. M. Dugger and S. Bartnicki-Garcia, eds.), pp. 222-234. Waverly Press, Baltimore, Maryland. Novaes-Ledieu, M.. and Garcia Mendoza. C. (1981). Can. J. Microbiol. 27, 779-787. Nuccitelli, R. (1983). Mod. Cell Biol. 2, 451-481. Ortega-Perez, R., Van Tuinen, D., MarmC, D., Cox, J. A,, and Turian, G. (1981). FEBS Lett. 133, 205-208. Park, D., and Robinson, P. M. (1966). Ann. But. 30, 425-439. Patton, A. M., and Marchant, R. (1978). Arch. Microbid. 118, 271-277. Peng, H. B., and Jaffe, L. F. (1976). Dev. B i d . 53, 277-284. Pdrkz, P., Garcia-Acha, I . , and DurBn, A. (1984). FEMS Microbiol. Lett. 23, 233-238. Pesti, M.,Campbell, J. M., and Peberdy, J. F. (1981). Curr. Microbiol. 5, 187-190. Picton, 1. M.. and Steer, M. W. (1981). J. Cell Sci. 43, 263-272. Picton, I . M., and Steer, M. W. (1982). J. Theor. B i d . 98, 15-20. Picton, J. M.. and Steer, M. W. (1983). Protoplasma 115, 11-17. Polacheck, I., and Rosenberger. R . F. (1975). J. Bacteriol. 121, 332-337. Polacheck, I., and Rosenberger, R. F. (1977). J. Bacteriol. 132,650-656. Polacheck, I., and Rosenberger, R. F. (1978). J. Bacreriol. 135, 741-747. Pollack, J. H., Lange, C. F., and Hashimoto, T. (1983). J. Bacteriol. 154, 965-975. Quatrano, R . S. (1978).Annu. Rev. Plant Phvsiol. 29, 487-510. Quigley, D.R.. and Selitrennikoff, C. P. (1984). Erp. Mvcol. 8, 202-214. Rast. D.R., and Bartnicki-Garcia, S. (1980). Proc. Nut/. Acud. Sci. U.S.A. 78, 1233-1236. Raudaskoski. M., and Koltin. Y. (1973). J. Bacteriol. 116,981-988. Reese. E. T. (1977). Recent Adv. Phytochein. 11, 31 1-367.

78

J. G. H. WESSELS

Reid, I. D., and Bartnicki-Garcia, S. (1976). J . Gen. Microbiol. 96, 35-50. Reinhardt, M. 0. (1892). Juhrb. Wiss. Eor. 23, 479-566. Reiss, H.-D., and Herth, W. (1979a). PIunru 146, 615-621. Reiss, H.-D., and Herth, W. (1979b). Plunru 145, 225-232. Reiss, H.-D., and Herth, W. (1980). Plunru 147, 295-301. Reiss, H.-D., and Herth, W. (1982). Plunru 156, 218-225. Reiss. H.-D., Herth, W., and Schnepf, E. (1983). Protoplusmu 115, 153-159. Reissig, J. L., and Kinney, S. G. (1983). J . Eacreriol. 154, 1397-1402. Robertson, N. F. (1958). Ann. Eor. 22, 159-173. Robertson, N. F. (1965). Trans. Er. Mycol. SOC. 48, 1-8. Robertson, N. F., and Rizvi, S. R. H. (1968). Ann. Eor. 32, 279-291. Robinson, P. M. (1973). Eor. Rev. 39, 367-384. Robinson, P. M., and Moms, G. M. (1984). Trans. Er. Mycol. SOC. 83, 569-573. Roelofsen. P. A. (1950). Rec. Truv. Eor. Neerl. 42, 73-110. Rwlofsen, P. A. (1959). “The Plant Cell Wall.” Encyclopedia of Plant Anatomy, 3 Part 4. Gebriider Bomtrager, Berlin-Nikolassee. Rosenberger, R. F. (1976). In “The Filamentous Fungi” (J. E. Smith and D. R. Berry, eds.), Vol. 11, pp. 328-344. Arnold, London. Rosenberger, R. F. (1979). In “Fungal Walls and Hyphal Growth” (J. H. Bumett and A. P. J. Trinci, eds.), pp. 266-277. Cambridge Univ. Press, London and New York. Ruiz-Herrera, J. (1984). In “Microbial Cell Wall Synthesis and Autolysis” (C. Nombela, ed.), pp. 113-120. Elsevier, Amsterdam. Ruiz-Herrera, J., and Bartnicki-Garcia, S. (1974). Science 186, 357-359. Ruiz-Herrera, J., Sing, V. O., Van der Woude, W. J., and Bartnicki-Garcia, S . (1975). Proc. Nurl. A d . Sci. U.S.A. 72, 2706-2710. Runeberg. P., and Raudaskoski, M. (1986). Eur. J. Cell Eiol.,in press. Sato, T., Norisuye, T., and Fujita, H. (1981). Carbon Res. 95, 195-204. Saunders, P. T., and Trinci, A. P. J. (1979). J . Gen. Microbiol. 110, 469-473. Scarborough, G. A. (1975). J. Eiol. Chem. 250, 1106-1111. Schekman, R. (1982). Trends Eiochem Res. July, 243-246. Schmiedel, G., and Schnepf, E. (1981). Plunru 147, 405-413. Schneider, E. F., and Wardrop, A. B. (1979). Can. J . Microbiol. 25, 75-85. Schroer, T. A., and Kelly, R. B. (1985). Cell 40, 729-730. Sentandreu, R., Larriba, G., and Elorza, M. V. (1981). Encycl. PIunr Physiol. New Ser. 13, 487512. Sentandreu, R., Martinez-Ramon, A., and Ruiz-Herrera, J. (1984). J. Gen. Microbiol. 130, 11931199. Shannon, T. M., Picton, J. M., and Steer, M. W. (1984). Eur. J. Cell Eiol. 33, 144-147. Shematek, E. M., Braatz, J. A., and Cabib, E. (1980). J. Eiol. Chem. 255, 888-894. Sietsma, J. H., and Wessels, J. G. H. (1977). Eiochim. Eiophys. Acru 496, 225-239. Sietsma, J. H., and Wessels, J. G. H. (1979). J . Gen. Microbiol. 114, 99-108. Sietsma, J. H., and Wessels, J. G. H. (1981). J . Gen. Microbiol. 125, 209-212. Sietsma, J . H., Rast, D., and Wessels, J. G. H. (1977). J . Gen. Microbiol. 102, 385-389. Sietsma, J. H., Sonnenberg, A. S. M., and Wessels, J. G. H. (1985). J. Gen. Microbiol. 131, 13311337. Sievers, A., and Schnepf, E. (1981). Cell Eiol. Monogr. 8, 265-299. Slayman, C. L., and Slayman, C. W. (1962). Science 136, 876-877. Smith, J. H. (1923). Ann. Eor. 37, 341-343. Sonnenberg, A. S. M., Sietsma, J. H., and Wessels, J. G. H. (1982). J . Gen. Microbiol. 128,26672674.

CELL WALL SYNTHESIS IN APICAL HYPHAL GROWTH

79

Sonnenberg, A. S. M., Sietsma, J. H., and Wessels, J. G.H. (1985). Exp. Mycol. 9, 141-148. Stadler, D. R. (1952). J. Cell. Comp. Physiol. 39, 449-474. Steele, G.C., and Trinci, A. P. J. (1975). New Phyrol. 75, 583-587. Steer, M. W., and Picton, J. M. (1984). I n “Structure, Function and Biosynthesis of Plant Cell Walls” (W. M. Dugger and S. Bartnicki-Garcia, eds.), pp. 483-494. Waverly Press, Baltimore, Maryland. Strunk, C. (1968). Arch. Mikrobiol. 60,255-261. Stump, R. F., Robinson, K. R., Harold, R. L., and Harold, F. M. (1980). Proc. Nurl. Acad. Sci. U.S.A. 77, 6673-6677.

Szaniszlo, P. J., Kang, M. S., and Cabib, E. (1985). J. Bucteriol. 161, 188-194. Thompson, D’Arcy, W. (1917, 1942). “On Growth and Form.” Cambridge Univ. Press, London and New York. Tomasz, A. (1984). In “Microbial Cell Wall Synthesis and Autolysis” (C. Nombela, ed.), pp. 312. Elsevier, Amsterdam. Traas, J. A., Braat, P.,Emons, A.-M. C., Meekes, H., and Derksen, J. (1985). J. Cell Sci. 76,303320.

Trinci, A. P. J. (1973). Arch. Mikrobiol. 91, 113-126. Trinci, A. P. J., and Collinge, A. J. (1975). J. Gen. Microbiol. 91, 355-361. Trinci, A. P. J., and Saunders, P. T. (1977). J . Gen. Microbiol. 103, 243-248. Troy, F. A., and Koffler. H. (1969). J. Biol. Chem. 244, 5563-5576. Turian, G.(1978). Experienriu 34, 1277-1279. Van der Valk, P., and Wessels, J. G.H. (1976). Proroplustnu 90, 65-87. Van der Valk, P., and Wessels, J. G.H. (1977). Acra Bot. Neerl. 26, 43-52. Van der Valk, P., Marchant, R., and Wessels, J. G.H. (1977). Exp. Mycol. 1, 69-82. Vermeulen, C. A., and Wessels, J. G.H. (1983). Curr. Microbiol. 8, 67-71. Vermeulen, C. A., and Wessels, J. G.H. (1984). Proroplustnu 120, 123-131. Vermeulen, C. A., Raeven, M. B. J. M., and Wessels, J. G.H. (1979). J . Gen. Microbiol. 114,8797.

Watts, F. Z., Miller, D. M., and On,E. (1985). Nature (London) 316, 83-86. Weisenseel, M. M., Nuccitelli, R., and Jaffe, L. F. (1975). J . Cell Biol. 66, 556-567. Wessels, J. G.H. (1984). I n “Microbial Cell Wall Synthesis and Autolysis” (C. Nombela, ed.). Elsevier, Amsterdam. Wessels, I. G.H., and Sietsma, J. H. (1979). In “Fungal Walls and Hyphal Growth” (J. H. Burnett and A. P. J. Trinci, eds.), pp. 28-48. Cambridge Univ. Press, London and New York. Wessels, J. G. H., and Sietsma, J. H. (1981a). Encycl. Plan? Physiol. New Ser. 13, 352-394. Wessels, J. G. H., and Sietsma, J. H. (1981b). In “Cell Walls ’81” (D.G. Robinson and H. Quader, eds.), pp. 135- 142. Wissenschaftliche Verlagsgesellschaft, Stuttgart. Wessels, I. G.H., Kreger, D. R., Marchant, R.,Regensburg, B. A., and de Vries, 0. M. H. (1972). Biochim. Biophys. Acra 273, 346-358. Wessels, J. G.H., Sietsma, J. H., and Sonnenberg, A. S. M. (1983). J. Gen. Microbiol. 129, 16071616.

Whittaker, R. H., and Margulis, L. (1978). Biosystems 10, 3-18. Willison, J. H. M. (1981). Encycl. Plan? Physiol. New Ser. 13, 513-541. Zonneveld, B. J. M. (1974). J. Gen. Microbiol. 81, 445-451.

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INTERNATIONAL REVIEW OF CYTOLOGY. VOL. 104

Connectin, an Elastic Filamentous Protein of Striated Muscle KOSCAK MARUYAMA Department of Biology, Faculty of Science, Chiba University, Chibn 260, Japan

I. Introduction Connectin, also called titin, is a very long, flexible filamentous protein of striated muscle that links thick myosin filaments to Z disks in a sarcomere. Despite its presence in rather high amount (about 10%) in myofibrils next to myosin (43%) and actin (22%), as listed in Table I, connectin has only recently been characterized. This was because physicochemical investigations have been very difficult due to its huge molecular weight of a few million. At present, a proteolytic product of the mother molecule (a-connectin, T,), p-connectin or T, has been purified as native form, but a-connectin has not yet been isolated. Connectin superthin filaments are shown to serve as an elastic component of striated muscle. Therefore, it is regarded as the fourth type of cytoskeletal structure following microtubules, intermediate filaments, and actin filaments. The problem is whether connectin-like filaments are present in nonmuscle cells or not. The present review deals with this problem after a full description of muscle connectin (cf. Wang, 1984, 1985; Locker, 1984a; Maruyama and Kimura, 1985; Ohtsuki et af., 1986).

11. The Third Filament? Over a century ago, a great German physiologist Johannes Miiller indicated in his famous textbook on human physiology that skeletal muscle could be considered to be elastic bodies (Miiller, 1840). Since then, it has long been assumed that muscle consists of two components, contractile and elastic. The elastic property of muscle was ascribed to the function of extracellular collagen fibers attached to muscle cell membranes (sarcolemma). In 1954, Natori first demonstrated that an elastic component exists in muscle cells using his famous demembraned fibers of the Natori type (Natori, 1954). He postulated the presence of an “internal elastic structure.” In the same year, H. E. Huxley and Hanson, together with A. F. Huxley and Niedergerke (1954), proposed the sliding theory based on the movement of the thin (actin) filaments relative to the thick (myosin) 81 Copyrighl 0 1986 by Academic Press. Inc. All rights or reprduclion in any form reserved.

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KOSCAK MARUYAMA TABLE I MYOFIBRILLAR PROTEIN OF VERTEBRATE SKELETAL MUSCLE“

Protein Contractile Myosin Actin Regulatory Troponin Tropomyosin M-Protein C-F’rotein a-Actinin Cytoskeletal Connectin Nebulin

Molecular weight (x 103)

Content wt%

Localization (band)

520 42

43 22

A I I I M line A Z line

70

33 x 2 165 135 95 x 2 2800 750

10 5

A-I I

OMinor structural proteins less than 1% of the total proteins are omitted. For a complete list, see Ohtsuki er al. (1986).

filaments in a sarcomere (H. E. Huxley and Hanson, 1954). In order to explain the continuity of myosin-removed myofibrils, they assumed the presence of an elastic filament called an S-filament linking the free ends of actin filaments in a sarcomere. However, this elastic filament model was not subsequently mentioned in the development of the sliding theory. In 1962, Sjostrand observed very thin filaments at the gap region between myosin and actin filaments when muscle was extremely stretched beyond the overlap of the two sets of filaments. Sjostrand (1962) called these filaments that were thinner than actin filaments “gap filaments,” and assumed that they were continuous with the tapered ends of myosin filaments. A year earlier, A. F. Huxley and Peachy (1961) mentioned the possible presence of “fine filaments” connecting the ends of both myosin and actin filaments from their observations of highly stretched muscle fibers. Graham Hoyle and his associates examined fine structures of a variety of striated muscles both in invertebrates and vertebrates and reached the conclusion that there were “superthin” or T-filaments (3 nm in width) connecting adjacent Z lines in a sarcomere (McNeill and Hoyle, 1967; Hoyle et af., 1968). Guba et al. (1968) reported that there were residual filaments after the extraction of myosin and actin. They claimed that those superthin filaments consisted of a protein designated fibrillen. dos Remedios (1969) observed that there was a third filament in addition to the myosin and actin filaments in a sarcomere that was resistant to salt extraction (dos Remedios and Gilmour, 1978). Revival of Sjostrand’s gap filament was brought about by Locker in 1975.

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TABLE II CHRONOLOGICAL LIST OF THE CONCEFT OF SUPERTHIN FILAMENT IN STRIATED MUSCLE Reference Inner elastic structure S filament connecting actin filaments Fine filament connecting myosin and actin filaments Gap filaments connecting myosin and actin filaments Connecting filaments connecting myosin and Z disk in insect muscle Superthin T filament running from Z to Z disks Fibrillen Third thin filament running from Z to Z disks G-filament connecting myosin and Z disks Connectin 10-nm filament connecting Z to Z disks Titin Projectin in insect muscle Cross-linked matrix

Natori (1954) H. E. Huxley and Hanson (1954) A. F. Huxley and Peachy (1961) Sjostrand (1962) Auber and Couteaux (1963) McNeill and Hoyle (1967) Guba er al. (1968) dos Remedios (1969); dos Remedios and Gilmour (1978) Locker and Leet (1975) Maruyama et al. (1976, 1977a) Price and Sanger (1977) Wang et al. (1979) Saide (1981) Lowey et al. (1983)

Locker and Leet (1975) first observed that bovine neck muscle (sternomandibularis) can be stretched to five times its rest length, up to 12 pm between the neighboring Z disks, spanned by very thin gap filaments, continuous with the thick filaments (Locker and h a t , 1976a,b). Locker called them G-filaments and assumed that they formed cores of myosin filaments, emerging at one end only, and arriving at the Z disk (cf. Locker, 1984a). Thus, as listed in Table 11, morphological and physiological observations strongly suggested the presence of the third superthin filament in addition to the thick and thin filaments in a sarcomere of vertebrate striated muscles. However, this was not widely accepted. One reason was, and still is, the lack of a clear-cut image of the compatibility of the third filament model toward the sliding theory of muscle contraction. Finally, it is to be noted that there are third filaments connecting the edges of myosin filaments to Z disks in fibrillar flight muscles of some insects, e.g., bee, fly, waterbug, etc. that rhythmically and quickly repeat contraction and relaxation (Auber and Couteaux, 1963; reviewed by Pringle, 1978). Projectin, the connecting filament protein, was isolated from honeybee thoracic muscle (Saide, 1981). 111. Connectin versus Titin

Stimulated by Natori’s pioneering work (Natori, 1954), the present writer began to identify the chemical entity of the elastic filament connecting Z disks in myofibrils of vertebrate skeletal muscle after removal of myosin and actin (rab-

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bit, chicken, and frog) in 1975. The residue was completely insoluble in salt solution, e.g., 1 M K1, LiBr, KSCN, etc. Dilute acid (1 N acetic acid, 0.1 N HCl, etc.) or alkali (0.01 and 0.1 N NaOH) also failed to solubilize. Most of the residue was resistant to even 0.1% SDS or 8 M urea. Alkali-treated ghost skinned fibers of frog muscle behaved just like rubber (Maruyama et al., 1976). When the insoluble residue, after removal of myosin, actin, and regulatory proteins and also connective tissue, was washed with water and then solubilized in I % SDS solution, the SDS-gel electrophoresis pattern showed that the main components were a high-molecular-weight (HMW) component which hardly moved and a 42kDa band (Maruyama et al., 1977a). The HMW component was cut out of the gel and used as an antigen. The FITC-labeled antiserum stained the filamentous material in the muscle residues (Maruyama et al. 1977a). Immunofluorescence observation showed that the A-I junction area of a sarcomere was most intensely stained (Maruyama et al., 1980). The 42-kDa component was nothing but denatured actin (Maruyama et al., 1983). King and Kurth (1980) isolated HMW connectin by chromatography on DEAE-Sepharose CL-6B in the presence of guanidine-HC1 and urea. Locker and Daines (1980) separated maleylated protein by DEAE-cellulose chromatography. The relationship between ‘‘gap filaments” and salt-insoluble connectin was discussed by Locker and Daines (1980). It is worth mentioning the peculiar solubility behavior of muscle structural proteins. Myosin can be easily extracted with a large volume of HasselbachSchneider solution: the remaining myosin would be very small in amount, if extraction is repeated several times. Actin extraction with 0.6 M KI is always incomplete. Some actin is left behind. If myofibrils are treated with 0.6 M KI from the beginning, a large amount of actin remains unextracted together with myosin. The proteins aggregate around deteriorated Z disks as first pointed out by Granger and Lazarides (1978). The most likely situation is that actin, myosin, and other proteins are bound to free connectin filaments extending from the Z disk and the whole mass aggregates near the Z disk. These become completely insoluble in salt solutions and some of them are not soluble even in a SDS solution. This explains the formation of the “elastic matrix” mentioned by Lowey et al. (1983). Quite independent of the connectin work aimed at the elastic filament of skeletal muscle, Kuan Wang accidentally discovered a giant protein by SDS-gel electrophoresis of total SDS e.xtract of whole muscle (Wang et al., 1979). Originally, he intended to find smooth muscle actin-binding protein (filamin according to Wang’) in skeletal muscle without success. Instead, he found at least three

’

Actin-binding protein (ABP) was first purified from rabbit lung macrophages (Hartwig and Stossel, 1975; Stossel and Hartwig, 1975). Chicken gizzard ABP was isolated by Wang (1977; cf. Wang er al., 1975) and named filamin.

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1

2

3

4

a

b

c

d

FIG. 1. SDS-gel electrophoresis patterns of a direct SDS extract of chicken breast muscle myofibrils. (a) 2.8% polydcrylamide gels, (b) 2.5%. (c) 2.3%,(d) 2.0%.( I ) a-Connectin (titin I ) , (2) p-connectin (titin 2), (3) nebulin, (4) myosin heavy chain. D. H. Hu (unpublished).

high-molecular-weight protein bands (bands 1, 2, and 3). Figure 1 shows SDSgel electrophoresis pakerns of a SDS extract of chicken breast muscle. By gel filtration in the presence of 0.1% SDS, Wang was able to isolate bands 1 and 2 in a denatured state and to raise antibodies against them. Bands 1 and 2 were named titin because of their huge molecular weight of one million. In myofibrils, they were located in the A - I junction area and also in Z disks as revealed by an immunofluorescence study. The content of titin was as large as 10% of the total myofibrillar proteins, and therefore Wang called it the third major structural protein of muscle. We immediately confirmed that Wang’s titin was identical to connectin (Maruyama et al., 1981a). Although King and Kurth (1980) separated connectin by gel filtration from guanidine- and HCI-solubilized muscle residues, Wang’s procedure was simpler and more reproducible. Then, our efforts were concentrated on the isolation of connectin in a native form. We noticed that some amount of connectin is soluble in a salt solution, coextractable with myosin in Guba-Straub solution (Maruyama et al., 1981a). It turned out that this is p-connectin (T,), derived from a-connectin (T2) by endog-

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enous protease. a-Connectin is not soluble. We have selected the conditions in which myosin is not solubilized and isolated native p-connectin (Kimura et al., 1982; Kimura and Maruyama, 1983a; Kimura et al., 1984b). Meanwhile, Wang’s group in the United States and Trinick’s group in the United Kingdom were successful in isolating connectin by different means (Wang et al., 1984; Trinick et al., 1984).

IV. Native Connectin A. PREPARATION With our observations that some connectin is solubilized from chicken breast muscle together with myosin in Guba-Straub or Hasselbach-Schneider solution (Maruyama et al. 1981a), we sought the conditions in which connectin alone is soluble without myosin. First, it was ascertained that connectin is soluble in the presence of 0.2 M NaCl (pH 7.0) or 0.075 M phosphate buffer (pH 6.5). With 0.1 M sodium phosphate buffer, connectin was solubilized above pH 6.5. At pH 7.0, both connectin and myosin were soluble, but the latter was not soluble at pH 6.5. However, a-actinin, actin, and other proteins were also extracted by 0.1 M phosphate buffer (pH 6.5). Therefore, myofibrils were first washed well with 5 mM NaHCO, followed by extraction with 0.1 M phosphate buffer at pH 5.6. These procedures removed a-actinin and other proteins. The precipitate was briefly washed with water and then extracted with 0.1 M phosphate buffer, pH 6.6. The filtrate consisted largely of connectin (Kimura and Maruyama, 1983a). For further purification, hydroxyapatite chromatography is highly recommended. However, it was noticed that fresh myofibrillar preparations resulted in a low yield of connectin. Therefore, myofibrils were prepared from muscle strips stored overnight at 0°C (Kimura et al., 1984b). This was due to proteolysis of aconnectin to p-connectin (see Section IV). The native connectin we obtained was p-connectin. By this procedure the yield was as high as 400 mg starting from 100 g of muscle. This is approximately 40% of the original content of connectin. Trinick and his associates (1984) have separated connectin from myosin in a salt extract of rabbit psoas myofibrils by sedimenting myosin in 0.2 M KCI. Contaminated C-protein and other proteins can be removed by DEAE-cellulose column chromatography. The eluted connectin is precipitated by (NH,),SO, at 35% saturation and it was dissolved in 0.5 M KCI containing 50 mM Tris-HC1 buffer, pH 7.9. This procedure is very convenient for concentrating connectin. Pure connectin can be obtained by gel filtration. The yield is approximately 200 mg from 100 g of muscle. Wang’s procedure (Wang er al., 1984) was similar to the procedures of C-protein purification (Reinach et al., 1982) and the yield was low: 100 mg from 100 g of muscle. Crude myosin preparations were subjected to

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DEAE-Sephadex chromatography to remove myosin. The unabsorbed material containing C-protein and connectin was then subjected to hydroxyapatite chromatography. Connectin tends to degrade during this preparation procedure, unless the procedure is performed quickly in the cold. B. SIZEAND SHAPE The mobility of connectin in SDS-gel electrophoresis is very slow: in a 510% polyacrylamide gel, connectin hardly moves and remains at the top of the gel. In 2-3% polyacrylamide gels it moves slowly (cf. Fig. 1). Since the apparent molecular weight (MW) of connectin is very large, appropriate markers for MW determination are not available. Therefore, artificial markers were prepared by cross-linking myosin heavy chains (MHC) with a maleimide derivative (Knight and Offer, 1978). Myosin heavy chains were rapidly dimerized by the cross-linking reaction of their SH groups with the maleimide reagent and within a few minutes of incubation monomers completely disappeared. Dimers were further cross-linked with each other forming 2 X dimers, 4 X dimers, etc. A linear relationship between logarithms of MWs and electrophoretic mobilities was observed up to 3200 kDa (8 X dimers) using 2.0% polyacrylamide gels, as seen in Fig. 2. From the mobility of isolated native connectin, its MW was roughly estimated to be 2100 kDa between 5 X dimers (2000 kDa) and 6 X dimers (2400 kDa) of MHC. This native connectin corresponds to the lower band (p-connectin) of direct SDS extract of intact myofibrils. The upper band (a-connectin) had an apparent MW of 2800 kDa. Wang (1982) claimed the MWs of 1800 and 1200 kDa for T , and T, (a-and p-connectins) by a similiar procedure, but he assumed that dimers, trimers, etc. had been formed by cross-linking reaction of MHC. Our sedimentation equilibrium measurements of MW of p-connectin in 0.5 M KCl and 0.1 M phosphate buffer (pH 7.0) also showed a value of 2700 kDa (Maruyama et al., 1984a). The sedimentation pattern was a single hypersharp peak having a sedimentation coefficient of 17 S in 0.1 M phosphate buffer, pH 7.0, in agreement with the value of 13.4 S in 0.5 A4 KCI and 0.05 M phosphate buffer, pH 7.5 (Trinick et al.. 1984). The shape of the sedimentation pattern showed that the connectin molecule is highly asymmetric. At the same time, the presence of a smaller peak of approximately 30-40 S suggested formation of side-by-side aggregates. An interesting observation was made in the presence of 1% SDS: the sedimentation coefficient of 15 S in the native state changed to 11 S in the denatured state. Since the peak became more hypersharp, it is likely that the change in the sedimentation coefficient was not due to subunit dissociation, but to a shape change in the presence of SDS. King (1984) estimated the MW of connectin in SDS to be lo6 by lightscattering technique. Electron microscopic images by a low angle rotary shadowing procedure showed that there were more straight rods in the presence of SDS

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KOSCAK MARUYAMA

a

C

e

rno bil it y FIG. 2. Molecular weight determinations of a- and p-connectins by SDS-gel electrophoresis: 2% polyacrylamide gels. The two arrows indicate a- and p-connectins. (a) Cross-linked myosin heavy chains, (b) isolated native connectin, (c) a + b, (d) direct SDS extract of chicken breast muscle. Myosin oligomers, (0)albumin oligomers. Modified (e) cross-linked bovine serum albumin. (0) from Maruyama ei al. (1984a).

than in its absence (H. Sawada and S. Kimura, unpublished). From the asymmetric nature of connectin, it was expected that the viscosity value was large. In a conventional Ostwald type viscometer, the value of intrinsic viscosity was less than 2 (g/dl) suggesting an axial ratio of about 50. However, it turned out that the viscosity of connectin greatly depended on the velocity gradient in the measurements. Thus at a very low velocity gradient of 0.0007 second-', a connectin solution of 0.3 mg/ml had a viscosity value of as high as 17,000 CPand the value dropped to 230 CP at 0.08 second-'. This is a thyxotropic nature owing to an entanglement of very thin filaments that can be easily disentangled by weak force. Electron microscopic observations have revealed that connectin is a very long,

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

89

flexible filament (Maruyama et al., 1984a; Wang et al., 1984; Trinick ef al., 1984). A low angle rotary shadowing method gave various kinds of images: from straight filament to entangled knitting wool (Fig. 3). The length distribution was heterogeneous ranging from 0.2 to 1 pm. Flow birefringence measurements

FIG. 3. Low angle shadowing images of connectin filaments. Note that myosin molecule (160 nm long) is included. Bar, 0.2 Fm. From H . Sawada and S. Kimura (unpublished; cf. Maruyarna er a / . , 1984a).

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KOSCAK MARUYAMA

suggested an approximate length of 0.4 p n in solution. Electron microscopic pictures showed that there were some beaded structures in the filaments. Trinick et al. (1984) observed that the width of the connectin filament was 4-5 nm in negatively stained samples. C. OTHERPROPERTIES Connectin is soluble as filaments in KCl concentrations higher than 0.2 M at pH 7-8 and aggregates in lower concentrations of KCl. At 0.05 M KCl it precipitates. Even in 0.2 M KCl, shaking or agitation results in fiber-like aggregate formation. On concentration by a rotary evaporator, lateral association occurs leading to formation of an elastic rubber-like bundle. UV absorption spectra are of a protein nature with a maximum at 280 nm. The value of A,, at 1 mg/ml was approximately 1.2 (light path, 1 cm). A slight shoulder around 290 nm is always seen, but its origin is unknown. Amino acid composition shows that connectin is an acidic protein (Table 111). On the whole, connectin is similar to actin in amino acid composition: proline, TABLE 111 AMINOACIDCOMPOSITION OF CONNECTIN FROM CHICKEN BREASTMUSCLP

Native connectin Asx Thr Ser Glx Pro GlY Ala Cysl2

Val Met Ile Leu TYr Phe LYS

His Arg

96 76 60 111 74 74 65 2 87 12 60

66 31 26 86 15 59

Denatured connectinb 1

2

95 75 69 116 74 71 62 II 85 10 59 67 30 27 82 15 55

93 66 67 I I8 67 76 75 6 78 16 56 76 30 29 79

uNumber of residues per lo00 residues. bPrepared according to Wang et al. (1979).

18

50

91

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN TABLE IV A M I N OACID COMFQSITIONS OF CONNECTIN, C-PROTEIN, AND PROJECT IN",^

Asx Thr Ser Glx Pro GlY Ala Cysl2 Val Met Ile Leu TYr Phe LYS His Arg

T

co

CI

c2

c3

P

95 75 69 I16 74 71 62 I1 85 10 59 67 30 27 82 15 55

93 66 67 118 67 76 75 6 78 16 56 76 30 29 79 18 50

96 59 57 I I7 71 71 66 14 I 04 17 51 69 29 37 86 14 44

99 58 62 I24 69 85 74 ND 85 13 43 68 27 35 89 17 52

I07 69 58 I I9 70 70 80 ND 68 18 62

I09 66 74 I25 76 93 68 8 53 12 36 69 27 32 87 23 51

64 35 32 93 15 48

UNumber of residues per 1000 residues. "T, Denatured connectin (chicken breast) (Maruyama c/ a/.. 1981); CO, native connectin (chicken breast) (Maruyama CI a / ., 1981 a); C I, C-protein (rabbit skeletal) (Offer cr a/.. 1973); C2, C-protein (rabbit white) (Callaway and Bechtel, 1981); C3, C-protein (rabbit red) (Callaway and Bechtel, 1981); P, projectin (Honeybee flight) (Saide, 1981); ND, not determined.

valine, and lysine are abundant and alanine, methionine, isoleucine, and tyrosine occur less in connectin than in actin. Methylhistidine is not present in connectin. A rather high content of proline (-9%) is in parallel with the lack in the a-helix portion (see below). The fact that connectin is rich in nonpolar amino acids may be related to its tendency to form salt-insoluble aggregates. A striking fact in the amino acid composition is that connectin is almost identical with C-protein (Table IV; Fig. 4). Although immunological crossreactivity was not detected using both antisera against connectin and C-protein, this fact is of some interest in view of their localizations in myofibrils (see Section VI). The amino acid composition of connectin is also very similar to projectin, an elastic protein of insect flight muscle (Table 1V; Fig. 4), whose MW is 360 kDa (Saide, 1981). This is of special interest in that both proteins serve as the elastic component of muscle. It is generally thought that a polypeptide of such a huge MW as connectin does not consist of a single peptide. It is reasonable to assume that several peptides are cross-linked as in collagen or elastin. The presence of hydroxylysinonorleucine,

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KOSCAK MARUYAMA

FIG. 4. Star diagrams of the amino acid compositions of connectin, C-protein, and projectin. Relative contents of the amino acids listed in Table IV are presented in star diagrams. ( I ) Denatured connectin (a + p). (2) native p-connectin, (3) projectin, (4) C-protein (rabbit skeletal), (5) C-protein (rabbit white), and (6) C-protein (rabbit red).

a cross-linker in collagen and elastin, was suggested by tritium incorporation experiments (Fujii and Maruyama, 1982). However, the amino acid analyses showed that the presence of the diamino acid was negligible (Maruyama et al., 1983). Gruen et al. (1982) also denied its presence. They could not detect glutamyllysine either. The carbohydrate content is very small and less than 1% by weight (Table V). Recently, Gassner et al. (1985) have emphasized the possibility of connectin as a glycoprotein. It is not clear whether the small amount of carbohydrates is covalently bonded to connectin or not (cf. Gassner, 1986). Circular dichroism measurements led Trinick and his associate (1984) to the conclusion that connectin completely consists of a random coil. This was an important finding. However, when we tried to measure circular dichroism in

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TABLE V CARBOHYDRATE CONTENTSI N CONNECTIN FROM CHICKEN BREASTMUSCLE"

Fucose Mannose Galactose Glucose GluNAc NeuNAc

Native connectin

Denatured connectinb

0.38 0.48 0.81 5.77 0.34 0

0.68 0.95 0.91 9.41

0.12 0

<'Dataare given as nrnol/rng dry weight. bPrepared according to Wang el al. (1979) (S Handa and S. Kimura, unpublished).

1983, it was not possible due to a strong orientation effect. Recently, we have obtained data by decreasing the protein concentration to 0.1 mg/ml in 0.5 M KCl. Trinick et al. (1984) used a connectin solution of 1 mg/ml for their study. The curve we obtained is quite different from that of Trinick et al. (1984), although the presence of a-helix is very small, if any (Fig. 5 ) . Although an accurate analysis is difficult, our curve suggests abundance of p-structure in addition to a smaller content of random coil (Maruyama et al., 1986). D. PROTEOLYSIS 1. Spontaneous Breakdown

Connectin is very easily degraded into smaller but still large fragments both in vivo and in vitro. As was first demonstrated by Wang et al. (1979), direct SDS extract of fresh whole muscle contains a- and p-connectins in an approximate ratio of 5: 1. In addition, a faint band just below p-connectin is always seen. This is tentatively called p'-connectin (Yoshidomi et a l . , 1985), but it has not yet been characterized (cf. Fig. 13). Isolated native p-connectin is mainly derived from a-connectin hydrolyzed in situ. Therefore, the question arises of whether p-connectin present in a SDS extract of whole muscle fibers is a real component of myofibrillar structural proteins or not. a-connectin might be partly degraded by intrinsic protease before its inactivation with SDS. This possibility cannot be ruled out, although we cannot readily agree with this view, because the ratio of p-connectin to aconnectin did not change by procedures of SDS extraction: muscle was first homogenized in water followed by addition of a SDS solution or directly homogenized in a SDS solution. Connectin was degraded into a form corresponding to

94

KOSCAK MARUYAMA

FIG. 5 . Circular dichroism spectra of native connectin solution: 0.5 M KCI, 5 mM phosphate buffer, pH 7.5. From H. Yoshidomi (unpublished; cf. Maruyama et a / . . 1986).

the smear without the two distinct bands in the SDS-gel electrophoresis pattern, when stored in a SDS solution for more than a few hours at room temperature. It must be kept in the semisolid condition at 0°C. When chicken breast muscle was placed in the ice, degradation of a-connectin to p-connectin completed within a day, and only p-connectin was present after 8

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

95

hours at 30°C (D. H. Hu, unpublished). This observation led us to prepare native (3-connectin effectively using muscle strips kept overnight at 0°C (Kimura et al., 1984b). On the other hand, it took 4-5 days at 0°C in well-washed myofibrils before a-connectin disappeared. Protease inhibitors, phenylmethylsulfonate (0.5 mM), p-chloromercuribenzoate (2 mM), leupeptin (10 pg/ml), and pepstatin (10 kg/ml) did not prevent the degradation at all. On the other hand, 1 mM CaCI, promoted it, although 10 mM EGTA only slightly delayed hydrolysis. Since calcium-activated neutral protease did not effectively hydrolyze a-connectin, it is still unknown what type of protease is responsible for in vivo hydrolysis of connectin. Seki and Watanabe (1984) observed that in carp muscle stored at 0°C aconnectin changed to (3-connectin within 10 hours, whereas it took 72 hours in rabbit skeletal muscle. a-Connectin was detected in a small amount in myofibrils prepared from bovine skeletal (longissimus) muscle stored for 3 days at 2°C and only a trace of a-connectin was present in myofibrils prepared from the muscle kept for 24 hours at 25°C (Lusby et al., 1983). Both a- and (3-connectins were largely broken down by heating bovine longissimus dorsi muscle for 40 minutes at 60 or 80°C (King et al., 1981; King, 1984; Locker and Wild, 1984). Isolated native connectin in solution is slowly degraded even at 0°C. It took 2 weeks at 0°C for (3-connectin to be split into 1900- and 400-kDa fragments that were similar to tryptic or chymotryptic products (see Section IV,D,2). 2. Effects of Proteases Various kinds of proteolytic enzymes, trypsin, chymotrypsin, papain, serine protease, and nagarse rapidly digested connectin in myofibrils. Interestingly, calcium-activated protease was not effective (Maruyama et al., 198Ib). Here, effects of trypsin and chymotrypsin will be described in detail. Both are very effective even in the weight ratio of 1:lOOO to chicken skeletal myofibrils. Trypsin ( I : 1000) digested a-connectin to (3-connectin within 1 minute at 25"C, and slowly split (3-connectin to 1900- and 400-kDa fragments. Chymotrypsin also acted in a similar way. In the weight ratio of 1:250, tryptic action produced I900-, 1700-, I400-, I 300-, I050-, 800-, 600-, and 400-kDa fragments, respectively. In the chymotryptic peptides, 1700- and 1300-kDa fragments were not present, and 1200-kDa fragments were produced. In both cases, the 1900- and 400-kDa fragments remained after prolonged hydrolysis, but this tendency was more evident in the chymotryptic action. Isolated native connectin was also very sensitive to trypsin and chymotrypsin. As seen in Fig. 6 , trypsin, in the weight ratio of 1:lOO or 1500, was more effective in splitting connectin than chymotrypsin. However, chymotrypsin in these weight ratios did not hydrolyze the 400-kDa fragment. Therefore, chymotryptic digestion has been used for the preparation of this fragment (Kimura et al., 1984b).

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KOSCAK MARUYAMA

FIG.6. Digestion of native p-connectin by trypsin and chymotrypsin. Connectin, 0.5 mg/ml, was incubated in 0.1 M sodium phosphate buffer, pH 7.0 at 25°C. The reaction was stopped by the addition of trypsin inhibitor, I mg/ml or 1 mM PMSF; 2% polyacrylamide gels were used. (a) Trypsin, 1/100 or 11500 by weight ratio; (b) chymotrypsin, 1/100 or 1/500 by weight ratio. Incubation time (minutes) is given under each lane. Lane C, intact sample. From H. Yoshidomi (unpublished).

V. Interaction with Myosin and Actin A. BINDING TO MYOSIN It was observed that connectin causes aggregation of myosin filaments around 50-120 mM KCI at pH 7.0. Turbidity increased and then gradually percipitation of aggregates occurred as seen in Fig. 7 (Kimura and Maruyama, 1983b; Kimura

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

97

et al., 1984a). The turbidity measurements revealed that the action of connectin is effective up to 0.15 M KCl, and the KCl dependence is reversible: upon dilution to 75 mM KCl aggregation took place, and on elevation to 0.15 M KCl the suspension became clear again. The aggregate formation of myosin filaments was ascertained by flow birefringence measurements and directly visualized under an electron microscope (Fig. 7). Myosin filaments were randomly aggregated in the presence of connectin. The myosin aggregates are easily sedimented in the presence of connectin under conditions in which myosin filaments or connectin alone remained in the supernatant. Thus the extent of binding of connectin to myosin filaments was

Fic. 7 . Effects of native connectin on aggregation of myosin and actin filaments at 80 mM KCI. (a) 1, Myosin; 2, connectin: 3, 4, myosin and connectin; (b) I , actin; 2, connectin; 3, 4, actin and connectin; (c) myosin + connectin; (d) actin + connectin. Modified from Kimura and Maruyama (1983b) and Kimura e r a / . (1984a).

98

KOSCAK MARUYAMA

studied quantitatively. There was a saturation around 0.12 mg connectin per 0. I mg myosin. This observation indicates a stoichiometric ratio of 0.3 mol of connectin per mol of myosin (Yoshidomi and Maruyama, 1985). Assuming that a myosin filament consists of 300 myosin monomers, as much as 90 connectin filaments can bind to one myosin filament. This value is much greater than the content of connectin relative to myosin in myofibrils (12 connectin filaments per myosin filament). Thick filaments in sarcomere are associated with several regulatory proteins of which C-protein is the most abundant (Offer et a l . , 1973). It is of interest whether C-protein affects the binding of connectin to a myosin filament. So far, there was no difference in the binding of connectin to C-protein-free and Cprotein-bound myosin filaments. Although binding of C-protein to connectin was reported (Koretz and Wang, 1984), we could not confirm this (Yoshidomi and Maruyama, 1985). The interaction of connectin with myosin appeared to be electrostatic. Using various neutral salt ions, such as K + , Na+, Li+, C1-, Br-, I - , SO:-, and citrate it was observed that the interaction depends on ionic strength but not on molar concentrations (Y. P. Huang, unpublished). It is well known that a myosin molecule consists of head S , , neck S , , and rods. We have examined the binding of connectin to these subfragments (Maruyama et al., 1985a). The head portion did not interact with connectin at all, whereas L-meromyosin and the rod portion effectively caused aggregation of myosin filaments. The neck S , fragments formed some complex with connectin at 0.04-0.06 M KCl. Heavy meromyosin formed a complex with connectin in the same range. This was likely due to contaminated light meromyosin that interacted with connectin in the presence of KCI lower than 0.15 M. Thus connectin binds to the tail portion of myosin that forms a filamentous structure. So far p-connectin (2100 kDa) is used in experiments for the interaction with myosin. Does a smaller fragment of connectin filament bind to myosin filament? The answer is yes. The 400-kDa fragment formed by chymotryptic action has been isolated by gel filtration (Kimura et al., 1984b). The one-fifth fragments reacted with myosin filaments to form sediment. However, smaller fragments of chain weights of 100 kDa or less did not cause aggregation of myosin. Finally, we should mention that a crude myosin preparation is easily precipitable upon dilution to an ionic strength of less than 0.05, whereas purified myosin is not. These familiar phenomena may be explained by myosin-aggregating action of connectin.

B. BINDINGTO ACTIN Actin filaments are also affected by connectin filaments in the presence of KCI up to 0.15 M (Kimura and Maruyama, 1983b; Kimura et al., 1984a). The turbidity increased and flocculent precipitates were formed when kept standing

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

99

-0-0-

I

5

10

15

20

Time(min)

FIG. 8. Effects of native connectin on the degree of flow birefringence of an F-actin solution. Factin. 0.25 mglml, 40 mM KCI, 10 mM phosphate buffer. At the first arrow connectin, 0.1 mglml, was added. At the second arrow KCI was added to give a final concentration of 150 mM. Velocity gradient, 100 seconds - I . From K . Maruyama (unpublished).

for a few hours. As shown in Fig. 8, degree of flow birefringence of an F-actin solution decreased by addition of connectin at a low velocity gradient just as by a-actinin (Maruyama and Ebashi, 1965). In fact, bundles of actin filaments are formed (Fig. 7). The effects of connectin on F-actin were also dependent on KCl concentrations as on myosin. The bundling action of connectin disappeared by raising the KCl concentration to 0.15 M (Fig. 8). Distinct from the action of aactinin (Drabikowski et al., 1968), the bundle formation by connectin was not affected by tropomyosin at all (H. Yoshidomi, unpublished). On the other hand, 400-kDa fragments of connectin did not exert any action on actin filaments at 50-120 mM KCl (Kimura er al., 1984b). Therefore, it is believed that flexible connectin filaments enhance an intrinsic tendency of bundle formation of actin filaments. The effect of connectin filaments on actin filaments may be attributed to topological restrictions. It may not be physiologically significant. Kimura et al. (1984a) reported that connectin enhances the onset of superprecipitation of actomyosin by MgATP resulting in an increase in the ATPase activity. The extent of this elevation was to a much smaller extent than a-actinin (Maruyama, 1966). Therefore, this effect of connectin appears to be an artifact in vitro.

VI. Location in Myofibrils A. IMMUNOFLUORESCENCE

STUDY

A simple method to locate a given protein in situ is an immunofluorescence observation using fluorescein isothiocyanate (F1TC)-labeled antibodies against

100

KOSCAK MARUYAMA

the protein. Anti-connectin was first prepared using cut top gels of salt-extracted muscle residues after electrophoresis in the presence of SDS (Maruyama et al., 1977a, 1980). Wang et al. (1979) used denatured connectin isolated by gel filtration in the presence of SDS. The two groups showed that connectin is most concentrated at the A-I junstion area in a sarcomere. It is also present in the A band except for the center region. The 1 band is only faintly fluorescent. We have observed that antiserum against chicken skeletal muscle connectin reacted with frog skeletal muscle connectin. When skinned muscle fibers of the frog were highly stretched so that gaps were formed between the A and I bands (sarcomere length, 3.5 pm), treatment with antibodies resulted in an elongation of an A band from 1.6 to 2.8 pm, as shown in Fig. 9 (Maruyama et al., 1984b). This clearly indicates that connectin filaments are present in the gap region. In fact, as presented in Fig. 9, very thin filaments are seen in this area in confirmation with Sjostrand ( 1962). More direct evidence that anti-connectin antibodies were deposited on the gap filament was reported (La Salle et al., 1983; Wang, 1985).

FIG. 9. “Gap filaments” in stretched myofibrils of frog skeletal muscle and immunofluorescence location of connectin in stretched myofibrils. ( I ) Thin section. Bar, 0.5 pm. (2) a, phase contrast image of a myofibril; b, phase contrast image of anti-connectin-treated myofibril; c , fluorescent image ofanti-connectin-treated myofibril. Bar, 10 pm. Modified from Maruyama ei a/. (1984b).

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

101

When myosin and actin are thoroughly extracted away from myofibrils, the strongest fluorescence is observed at both sides of the Z lines (Maruyama et af., 1977a, 1984b; Wang et af.,1979). This turned out to be due to accumulation of residual myosin, actin, and other proteins together with connectin (cf. Granger and Lazarides, 1978; Hattori and Takahashi, 1979). Fluorescent fiber-like structures are seen between Z lines. On the contrary, when myosin was removed, connectin was shown to be translocated to the M band region (Wang, 1985).

B. ELECTRONMICROSCOPY Trinick ( I 98 1) described “end-filament” attached to isolated thick filaments, 5 nm wide and 85 nm long. Later, Trinick and his associates (1984) identified the end-filaments as connectin filaments by comparison with isolated connectin under an electron microscope using negatively stained samples. Three to four connectin filaments are extended from each end of a thick filament of rabbit psoas myofibrils. These observations are in good agreement with our observations on thin sections of stretched fibers of frog muscle (Maruyama et af., 1984b). The question every investigator in this field has raised is where connectin filaments starting from thick filaments are terminated. Locker (cf. 1984a) and Maruyama and Kimura (1985) assumed that connectin filaments are directly linked to Z disks. On the other hand, Wang (1984, 1985) has claimed that connectin is attached to nebulin meshwork and eventually linked to the Z disk. Moreover, Wang ( I 985) assumes that a connectin filament spans from an N, line to other N, lines in a sarcomere. The difficulty is that connectin filaments cannot be identified in intact 1 bands because of the presence of many actin filaments. Also its exact location on a myosin filament remains obscure. We have prepared thin sections of anti-connectin-treated muscle fibers of frog skeletal muscle. First there was no sign of antibody staining in a sarcomere (Maruyama et al., 1984b). However, it was noticed that antibody deposits were only restricted to a few layers of peripheral myofibrils (Maruyama et af., 1985b), as seen in Fig. 10. Evidently, penetration of antibodies into myofibrillar bundles was very slow. Several distinct stripes were symmetrically present in each half of the A band of anti-connectin-treatedmuscle fiber at rest length (Fig. 10).The first stripe was seen approximately 0.15 pm away from the center of the A band. Thus the two stripes are at the edges of a pseudo H zone. The positions of other stripes from the center of the A band were approximately 0.3, 0.6, 0.7, 0.8, 0.85, and 0.9 pm, respectively. The latter two were outside of the A band. These bands were more markedly recognized in antiserum-treated myofibrils swollen at a low ionic strength (Fig. lo), where diffusion of antibodies was more rapid than in intact fibers.

FIG. 10. Immunoelectron micrographs of anti-connectin-treated myofibrils of frog skeletal muscle. (a) Rest length myofibrils; (b) stretched myofibrils; (c) low ionic strength extracted myofibrils. Bar, 1 pm. From T. Yoshioka (unpublished; cf. Maruyama et al., 1985b).

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

103

More important were the deposits of antibodies in I bands and Z lines (Fig. lo), clearly indicating that antigenic sites distribute all the way through Z lines. Stretched fibers did not show regular stripes at all. Antibody labeling was randomized because myosin filaments were pulled randomly to either side of Z disks (Fig. 10). These observations with polyclonal antibodies against connectin show that connectin filaments directly link each end of a myosin filament to its neighboring Z disk as an elastic component.

VII. Connectin as an Elastic Component As Natori (1954) first showed, there is an elastic component in myofibrils that results in passive tension generation upon extreme stretch and also in the return of separated thick and thin filaments to the original state on release (cf. Rb. Natori et al., 1980; Yoshioka et a/., 1981). First we postulated that an elastic filament like connectin directly links neighboring Z disks in a sarcomere (Maruyama et al., 1976). Electron microscopic observations suggest the presence of relatively few filaments running through Z disks in myosin- and actin-extracted myofibrils (cf. Wang and Ramirez-Mitchell, 1983). Wang and his associate (1983) regarded the remaining filaments as intermediate filaments (cf. Price and Sanger, 1979). However, intermediate filaments are known to connect adjacent Z disks at the periphery of a myofibril in cardiac muscles (Tokuyasu, 1983), but not in skeletal muscle except in early stages of embryonic development (Tokuyasu er d., 1984). Therefore, Wang’s view is not valid. Although the span of a few connectin filaments between adjacent Z disks is not definitely denied, the remaining filaments are likely to be associated connectin filaments freed from thick filaments. Some myosin or actin might be trapped in them. It is true that those remaining filaments connect KI-extracted Z disks in ghost myofibrils (Wang and Ramirez-Mitchell, 1983; Maruyama et a/., 1984b). Our work using polyclonal antibodies against connectin has definitely shown that connectin filaments directly link myosin filaments to Z lines starting from 0. I5 k m away from the center of the A band in a symmetrical way (Maruyama et al., 1985b). The presence of distinct stripes in each half of a sarcomere indicates that connectin filaments are in register along myosin filaments at rest length (see Fig. 10). A tentative model of connectin filaments in a sarcomere is depicted in Fig. 1 1 . However, we cannot exclude the possibility that a connectin filament

c-FIG I I

I A model of connectin filaments

in

a sarcomere

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KOSCAK MARUYAMA

spans a whole sarcomere by way of its association with thick filament between adjacent Z disks. If so, a connectin filament must consist of an end-to-end (or head-to-head) dimer as judged by symmetrical distributions of epitopes (Wang and Ramirez-Mitchell, 1984; Maruyama et al., 1985b). Locker (1984a) and Magid et al. (1984) have postulated that connectin is present in the core of a thick filament. This possibility is not excluded, but the presence of connectin filaments has been observed alongside the end region of a thick filament (Trinick et al., 1984; Maruyama et al., 1984b). Therefore, at least several connectin filaments attach to the surface of a thick filament starting from the edge of the center bare zone (Maruyama et al., 1985b). Taking the MW values of connectin and myosin as 3 X lo6 and 5 X lo5,their contents of 10 and 44% in the total myofibrillar proteins, and 300 myosin monomers to form a thick filament, the presence of 12 connectin filaments per thick filaments is calculated. It is expected that six connectin filaments are present in each half of the thick filament. In the above calculation, it is assumed that a connectin filament consists of a single chain of molecule. Recent work using monoclonal antibodies has showed the specific location of each species as a pair of symmetrical stripes at the I band, A-I junction, and A band (Wang and Ramirez-Mitchell, 1984). This stimulating observation strongly suggests that a long peptide of connectin with one epitope for one species of monoclonal antibody extends from the myosin filament to the Z disk. Since a connectin molecule does not contain any a-helical structure but consists of random coil (Trinick et al., 1984) and p structure (Maruyama et a l . , 1986), a single peptide of 3 X lo6 MW could have a length of up to 7 pm. At rest length, half sarcomere length is about 1.2 pm and at extreme stretch it is 3.5 pm. Therefore, connectin filaments must be compactly folded in a sarcomere. Trinick (1981) pointed out that “end-filament” has beaded structure, and this has been confirmed in isolated connectin filaments (Trinick et al., 1984; Wang er al., 1984). In this connection, it is of special interest to determine whether the pspiral structure in elastin fibers is present in connectin filament or not (cf. uny, 1984). Higuchi and Umazume (1985) made an important observation on the passive tension generation by stretch of skinned muscle fibers of the bullfrog. A brief extraction of myosin with a salt solution resulted in a decrease in tension development in proportion to the amount of myosin extracted. Only 10% of tension generation remained after almost complete removal of myosin. The decrease in tension generation can easily be explained by the release of connectin filaments at the attachment site on the myosin filament. It has already been indicated that some connectin is solubilized together with myosin. Another significant aspect of passive tension generation of skinned muscle fibers is the effect of a brief trypsin digestion. Mild tryptic treatment (0.2 pg/ml) led to the decrease in passive tension generation (Fig. 12). At the same time, a-connectin was de-

105

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

0

1.o

1

3

5

7

(min)

I

C

.-0 v)

C

al I-

0-l

I

1

1

r

5 Trypsin Treatment

I

10 ( m i d

FIG.12. Decrease in generation of passive tension and hydrolysis of connectin during trypsin treatment of skinned fibers of frog skeletal muscle. SDS-gel electrophoresis patterns show progressive breakdown of a- and p-connectins. From H. Higuchi and S . Kimura (unpublished).

graded to p-connectin. Under this mild tryptic digestion, any other proteins including nebulin were not hydrolyzed. This fact strongly suggests that a-connectin filaments serve as an elastic component in a sarcomere. It is to be noted that a string of glycerinated myofibril from rabbit psoas muscle contracted into a small mass on addition of MgATP, whereas after a mild treatment with trypsin it fell into a number of small pieces in the presence of MgATP (Maruyama and Yamamoto, 1979). This observation again suggests that a trypsin-sensitive structure is involved in the elastic continuity of a myofibril. The change in length of connectin filament during contraction and relaxation has been shown in cardiac myofibrils using a fluorescent stripe stained with monoclonal antibodies as marker (S. M. Wang and Greaser, 1986).

VIII. Connectin Transformation during Differentiation A. CHANGES DURING DEVELOPMENT OF THE CHICK It has been well established that a number of muscle structural proteins undergo characteristic changes in isoform expression during embryonic and neonatal development (cf. Obinata et al., 1984). We have investigated changes in connectin isoforms during embryonic and

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KOSCAK MARUYAMA

a

b

C

Fic. 13. Connectin isoforms of embryonic, neonatal, and adult skeletal muscles. (a) Just hatched chick. Note that the uppermost band is embryonic cx-connectin and the band just below is neonatal aconnectin. Two faint bands, embryonic P- and P'-connectins are seen. (b) a + c. ( c ) Adult connectin. Note that there is adult P'-connectin just below p-connectin. Modified from Yoshidomi c'r a/. ( 1985).

neonatal development of the chick using SDS-gel electrophoresis checked by an immunoblot technique (Yoshidomi et al., 1985). Although the band was very faint, connectin-like high-molecular-weight protein was detected in breast muscles of 7-day incubated chick embryo. Cruen e?al. (1982) reported that connectin is expressed in 7.5-week-old sheep fetus much later than the appearance of myosin. The 13-day chick embryo has bands which reacted with antiserum against adult breast muscle connectin. The embryonic a-connectin showed slower mobility in SDS-gel electrophoresis than the adult one, as clearly seen in Fig. 13. Its apparent MW was estimated to be as high as 3.4 million (Table VI). This embryonic a-connectin is present up to 3 days posthatch. It is of some interest to note that the high-MW band of embryonic a-connectin corresponds to that of slow muscle (Section IX,A). In developing chick fast muscle, the first cardiac TABLE VI APPARENT MOLECULAR WEIGHTSOF CONNECTIN ISOFORMSI X J R I N G DEVELOPMENT OF THE CHICK"

Embryonic Neonatal Adult

a

P

P'

3.4 3.1 2.8

2.4 2.4 2. I

2. I 2.1 2.0

G i v e n as million (lo6).

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

107

type of C-protein is expressed, followed by the slow muscle type; finally the fast muscle type is expressed (Obinata et a/., 1984). Unfortunately, our polyclonal antibodies are not tissue and species specific and, therefore, tissue specificity cannot be distinguished. Neonatal a-connectin appears just after hatch and is present up to 7 days posthatch (Fig. 13). Adult type a-connectin, its MW being slightly smaller than the neonatal one, is expressed at 5 days posthatch. Changes in p-connectin are more complicated. First, on close examination, the connectin bands are not doublet bands as claimed before (Wang et al., 1979). There is another faint band just below the p band (Fig. 13). This p’ band appears not to be a proteolytic product of a-connectin, because the amount does not change when a-connectin is degraded (H. Yoshidomi, unpublished). Adult pconnectin appears at 5 days posthatch. In 17-day embryo, there are two p bands: the mobility of the lower band is the same as that of adult p-connectin. Although this band is not distinguishable from adult p, it is tentatively called embryonic p’, and the upper embryo-specific p band is termed embryonic p, which is present up to 3 days posthatch. The changes in the types of connectin during development of the chick are summarized in Table VI. The relative amount of connectin markedly increased after hatch reaching a value of adult connectin (10% of the total myofibrillar protein) at 7-10 days after hatch.

B . CHANGES DURING MYOFIBRILLOGENESIS Recently, Greaser and his associates (S. M. Wang et al., 1984) have studied the fate of connectin during myofibrillar differentiation in cultured rat leg muscle cells, using fluorescent monoclonal antibodies against bovine cardiac connectin (S. M. Wang and Greaser, 1985). Newly synthesized connectin filaments are scattered randomly in the cytoplasm, and then these filaments gradually associate with actin and myosin to form longitudinal bundles. As myofibrils are formed, the banded patterns of connectin at the A-I junction area are observed. Sometimes, the appearance of connectin periodicity preceded the band formations of myosin and actin. This suggests an important role of connectin in sarcomere assembly. Immunoelectron microscopic examinations will be of great help in understanding the situation.

IX. Comparative Biochemistry A. GENERAL SURVEY There are two ways to detect connectin in various types of tissues or organisms: detection of high-molecular-weight protein bands in 2-3% polyacrylamide gels in the presence of SDS preferably with confirmation by an

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KOSCAK MARUYAMA

immunoblot and also by immunofluorescenceexaminations of fixed cells. Antibodies raised against chicken breast muscle connectin cross-reacted with frog leg muscle connectin (Maruyama et al., 1984b). Ikeya et al. (1983) investigated cryostat sections of chicken tissues using an immunofluorescence technique: connectin was detected only in striated muscles, breast, anterior latissimus dorsi (ALD), and cardiac muscles. There was no sign of connectin in tendon, blood vessels, gizzard, nerves, and liver. A SDS-gel electrophoresis survey confirmed these observations (Hu et al., 1986a). It is rather surprising that connectin is specific for striated muscle just like troponin. S. M. Wang et al. (1984) also noticed that connectin is not found in cultured nonmuscle cells. It is noteworthy to mention “high-molecular-weight (HMW)” proteins in chicken gizzard. As already reported by Wang et al. (1979), total SDS extract of fresh gizzard does not contain any HMW bands corresponding to a MW of 1 million in SDS-gel electrophoresis. However, when gizzard was thoroughly extracted with Hasselbach-Schneider solution and 0.6 M KI to extract myosin and actin followed by 1 N acetic acid (to remove desmin), connectin-like HMW band appeared at the top of the gel. The fraction was separated by gel filtration in the presence of SDS. Amino acid analyses revealed that it contained hydroxyproline by as much as lo%, suggesting that the HMW protein in question was crosslinked collagen (Y. Kuwano and K. Maruyama, unpublished). Our earlier observation that salt- and acid-extracted residue of gizzard contained connectin (Maruyama et al., 1977b) is no longer valid. SDS-electrophoresis examinations showed that slow muscle (ALD) a-connectin has a larger MW than fast breast muscle a-connectin. Chicken cardiac aand p-connectins have the same molecular weights as breast muscle ones, but there is no p’-connectin. In all the vertebrate skeletal muscles examined, connectin has been detected in SDS-gel electrophoresis: rabbit (Wang et al., 1979; Trinick et al., 1984), bovine (Locker and Daines, 1980; Lusby et al., 1983; King, 1984), rat (Gruen et al., 1982), sheep (King and Kurth, 1980; King etal., 1981), chicken (Maruyama et al., 1981a, 1984a), snake (Hu et al., 1986a), frog (Maruyama et al., 1984b), and carp (Seki and Watanabe, 1984). In invertebrates, detailed studies have not yet been made except for a brief note: insect (Wang et al., 1979; Locker and Wild, 1986; Hu et al., 1986a); crayfish (Hu et al., 1986a); C . elegans (Hu et al., 1986a), and chordate, Amphioxus (Hu et al., 1986a).

B. NATIVECONNECTIN FROM CARDIAC MUSCLE Native connectin has been purified from pig heart (Ito et al., 1986). The method used was the same as that for the preparation from chicken breast muscle (Kimura et al., 1984b) except that it was not necessary to keep excised muscle overnight at 0°C. It appears that a-connectin is rapidly degraded to p-connectin

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

109

in cardiac muscle. Myofibrils were prepared from pig heart immediately after delivery from a local slaughter house. The yield was approximately 80 mg starting from 100 g of muscle. The SDS-gel electrophoresis pattern showed that cardiac native connectin moved at the same mobility as chicken breast muscle p-connectin. A negatively stained sample showed under an electron microscope that there were entanglements of very thin filaments as elegantly presented by Trinick et a f . (1984). Surprisingly, the amino acid composition was almost identical with that of chicken native connectin (Table IV). Cardiac connectin also interacted with both myosin and actin filaments in the presence of 50-150 mM KCl at pH 7.0 and 25°C.

C. Physarum PLASMODIAL PROTEIN Ozaki and Maruyama ( 1980) first indicated that salt-extracted residues of plasmodium of the slime mold Physarum polycephalum contained a SDS-insoluble material from which acidic polysaccharides could be separated by centrifugation as transparent gels. These SDS-insoluble residues could be partly dispersed in 6 M guanidine-HCI. Under an electron microscope, entanglements of very thin filaments were observed. Its amino acid composition was very similar to that of muscle connectin (Table VII). Therefore, it is assumed that there are connectin-like filaments in Physarum plasmodium. Recently, Wohlfarth-Bottermann’s school has been able to isolate connectinlike protein from cytoplasmic droplets of plasmodium in the presence of SDS (Gassner et a f . , 1985; Gassner, 1986). The HMW protein isolated from a SDS extract of Physarum plasmodium freed of extracellular slime formed huge, elastic, gel-like aggregate after removal of SDS by dialysis. Electron microscopy by negative staining and by rotary shadowing revealed the presence of superthin filaments 2-3 nm wide. Gassner et a f . (1985) emphasized that the isolated superthin filaments closely resemble those observed in plasmodia1 endoplasmic droplets and also in their SDS-treated ghost cytoskeletal matrix. It is suggested that Physarum connectin-like filament is a major component in the elastic cytoskeletal matrix. We have confirmed the above result (Hu et a f . , 1986b) using whole plasmodium. A homogenate in 1 mM NaHCO, was briefly centrifuged and the precipitate was used for the starting material. To the precipitate SDS was added, and the solution was centrifuged for 2 hours at 200,000 g to sediment the polysaccharides. The supernatant was subjected to gel filtration to isolate connectin-like protein. The amino acid composition was similar to that reported by Gassner et a f . ( 1985) and also to that by Ozaki and Maruyama (1980) (Table VII). Negatively stained sample showed a thin filamentous structure under an electron microscope. Native HMW protein has been obtained from direct gel filtration of a bicarbo-

110

KOSCAK MARUYAMA

AMINOACIDCOMPoSlTlON

Asx Thr Ser Glx Pro GIY

Ala Val Met Ile

Leu TYr Phe LYS His A%

TABLE VII Physarum

OF

SDS insoluble('

Isolatedh

100 64 67 116 62 70 84 60 18 49 88 29 42 74 24 49

loo 55 68 126 63 73 84 58 17 48 86 29 39 77 26 50

CONNECTIN-LIKE PROTEIN

SDS insoluble matrices" 101

53 70 I28 61 80 96 52 14 37 83 25 34 71 34 60

Isolated'' 102 58 69 I28 60 72 90 58 20 45 86 27 38 71 38 60

OOzaki and Maruyama (1980). bD.H. Hu and S . Kimuras (unpublished). G a m e r et al. (1985).

nate extract of plasmodium (Hu et al., 1986b). This fraction caused aggregation of rabbit skeletal muscle myosin as muscle connectin. Thus, it is very likely that connectin-like protein is present in the slime mold. However, judging from the mobility in SDS-gel electrophoresis and the presence of an SDS-insoluble form, Physarum connectin-like proteins are covalently cross-linked.

X. Perspectives Connectin (also called titin) has a long history of doubt, neglect, and revival. Some of the pioneering researchers include Natori (Japan), Sjostrand (United States and Sweden), Hoyle (United States), Guba (Hungary), dos Remedios (Australia), and Locker (New Zealand). Even after the present writer took up this elastic protein in 1976, followed by Wang from a different point of view in 1979, most scientists were and still are reluctant to accept it. The present writer feels that this reluctance is rather natural because the protein molecule in question is very large and quite outside the reasonable range of sizes of hitherto known proteins and, in addition, its width (5 nm) is near the limit of resolution under an

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

111

electron microscope. It may be added that all the workers who followed our and Wang’s research so far have belonged to meat science laboratories: Takahashi (Japan), King (Australia), Locker (New Zealand), Greaser (United States), Robson (United States), and Trinick (United Kingdom). Is there really a single peptide of three million MW consisting of more than 20,000 amino acids? Wang’s elegant work showing the epitope distribution corresponding to each of monoclonal antibodies (Wang and Ramirez-Mitchell, 1984) strongly suggests so. From a protein chemical view it is extremely difficult to test this problem simply because of the size and tendency of degradation. Despite difficulty, gene cloning will give a final answer. At present we have not yet been successful in the isolation of the mother molecule, a-connectin. What has been isolated independently in Japan, the United States, and the United Kingdom is its proteolytic product, p-connectin (Kimura and Maruyama, 1983a; Wang et al., 1984; Trinick et al., 1984). aConnectin is not easily soluble in salt solution, but a small amount can be solubilized together with its degradation peptides (S. Kimura, unpublished). Therefore, isolation of a-connectin does not seem impossible. Returning to the connectin filaments in a sarcomere of striated muscle, details of the structure have remained obscure. First, are they present as a singlestranded filament consisting of random coil and p-structure? In the rest state, the length may be in the order of 1 pm and it may be extended to longer than 3 pm when stretched. To generate tension, an elastic structure should be present in connectin filaments. One of these candidates is P-spiral structure in elastin fibers (Urry, 1984), since connectin is rich in P-sheet structure (Maruyama et al., 1986). Connectin filaments directly link myosin filaments to Z disks in a sarcomere. How does this attach to the Z disk? Is there any cementing substance? Connectin filaments bind to the rod portion of myosin filaments, but details of the manner of binding are not known, It is also possible that a few connectin filaments exist in a core of myosin filament (Locker, 1984a; Magid et al., 1984). It is known that there are high-density areas called N , and N, lines in the 1 band (cf. Locker, 1984b; Cooke, 1985). Nebulin constitutes an N, line (Wang and Willaimson, 1980). Wang (1984, 1985) assumes that connectin filaments are linked to nebulin meshwork. At present the nature of native nebulin is completely unknown. Furthermore, as pointed out by Locker (1984b), there is no nebulin in cardiac myofibrils. We have also observed that the content of nebulin in cardiac myofibrils is very small, if any (Hu et a/., 1986a). It is possible that connectin attaches to both N , and N, lines in skeletal muscle and this is a problem that needs clarification. The mechanism of myofibrillar formation during differentiation of muscle cells is of vital importance from a biological point of view. Greaser’s school has presented very suggestive evidence that connectin filaments are deeply con-

112

KOSCAK MARUYAMA

cerned in the sarcomeric assembly of myosin and actin filaments (S. M. Wang el al., 1984). More detailed work is due along this line. Finally, from the comparative view of cell motility, the presence of connectinlike superthin filaments in the slime mold is of special interest. A thorough investigation is desirable. Are there similar type of filaments in the amoeba and other motile cells?

ACKNQWLE~GMENTS

I express my cordial thanks to Dr. S. Kimura for painful joint work and also to other members of our laboratory for their cooperation. I am much indebted to Professors R. Natori of Jikei Medical School, S. Ebashi of the National Institute for Physiological Sciences, and Sir Andrew Huxley of Trinity College, Cambridge, for their constant encouragement. I am deeply indebted to Dr. H. Sawada of the University of Tokyo for his constant help in the present work. Finally, acknowledgments are due to Professor H. Noda of the University of the Air for his critical reading of the manuscript.

REFERENCES

Auber, J., and Couteaux, R. (1963). J . Microsc. 2, 309. Callaway, J. E., and Bechtel, P. J. (1981). Biochem. J . 195, 463. Cooke, P. (1985). In “Cell and Muscle Motility” (J. W. Shay ed.), Vol. 6, pp. 287-313. Plenum, New York. dos Remedios, C. G. (1969). Ph.D. thesis, University of Sydney. dos Remedios, C. G., and Gilmour, D. (1978). J . Biochem. 84, 235. Drabikowski, W., Nonomura, Y., and Maruyama, K. (1968). J . Biochem. 63, 761. Fujii, K., and Maruyama, K. (1982). Biochem. Biophys. Res. Commun. 104, 633. Gassner, D. (1986). In “The Molecular Biology of Physarurn polycephalum” (W. F. Dove, ed.), pp. 225-236. Plenum, New York. Gassner, D., Shraideh, Z., and Wohlfarth-Bottermann, K. E. (1985). Eur. J . Cell Biol. 37, 44. Granger, B. L., and Lazarides, E. (1978). Cell 22, 1253. Gruen, L. C., King, N. L., Kurth, L., and McKenzie, L. J. (1982). Inr. J . Pepride Res. 20, 401. Guba, F., Harsanyi, V., and Vajda, E. (1968). Acta Biochim. Biophys. Acad. Sci. Hung. 3, 433. Hartwig, J. H., and Stossel, T. P. (1975). J . Biol. Chem. 250, 5699. Hattori, A., and Takahashi, K. (1979). J . Biochem. 85, 47. Higuchi, H., and Umazume, Y. (1985). Biophys. J . 48, 137. Hoyle, G., McNeill, A., Wallcott, B., and Selverston, A. (1968). Symp. Biol. Hung. 8, 34. Hu, D. H., Kimura, S., and Maruyama, K. (1986a). J . Biochem. 99, 1485. Hu, D. H., Kimura, S., Suzuki, T., and Maruyama, K. (1986b). I n “The Molecular Biology of Physarum polycephalum” (W. F. Dove, ed.), pp. 237-242. Plenum, New York. Huxley, A. F., and Niedergerke, R. (1954). Narure (London) 173, 971. Huxley, A. F., and Peachy, L. D. (1961). J. Physiol. (London) 157, 150. Huxley, H. E., and Hanson, J. (1954). Narure (London) 173, 973. Ikeya, H., Ohashi, K., and Maruyama, K. (1983). Biomed. Res. 4, I 1 I . Ito. Y.,Kimura, S., and Maruyama, K. (1986). J. Biochem., in press. Kimura, S., and Maruyama, K. (1983a).J. Biochem. 94, 2083.

CONNECTIN, AN ELASTIC FILAMENTOUS PROTEIN

113

Kimura, S . , and Maruyama, K. (1983b). Biomed. Res. 4, 607. Kimura, S., Sawada, H., and Maruyama, K. (1982). Biophysics Jpn. 22, 1 4 3 . Kimura, S., Maruyama, K., and Huang, Y. P. (1984a). J. Biochem. 96, 494. Kimura, S . , Yoshidomi, H., and Maruyama, K. (1984b). J. Biochem. 96, 1947. King, N. L. (1984). Meur Sci. 11, 27. King, N. L., and Kurth, L. (1980). In “Fibrous Proteins: Scientific, Industrial, and Medical Aspects” (D. A. D. Parry and L. K. Creamer, eds.), Vol. 2, pp. 57-66. Academic Press, New York. King, N. L., Kurth, L., and Shorthose, W. R. (1981). Meat Sci. 5, 389. Knight, P., and Offer, G. (1978). Biochem. J. 175, 1023. Koretz, J. F., and Wang, K. (1984). Biophys. J. 45, 104a. La Salle, F., Robson, R. M., Lusby, M. L., Parrish, F. C., Stromer, M. H., and Huiatt, T. W. (1983). J. Cell B i d . 97, 258a. Locker, R. H. (1984a). Food Microsrrucr. 3, 17. Locker, R. H. (1984b). J . Ultrustrucr. Res. 88, 207. Locker, R. H., and Daines, G. J. (1980). In “Fibrous Proteins: Scientific, Industrial, and Medical Aspects” (D. A. D. Parry and L. K. Creamer, eds.), Vol. 2, pp. 43-55. Academic Press, New York. Locker, R. H., and Leet, N. G. (1975). J. Ultrusrrucr. Res. 52, 64. Locker, R. H., and Leet, N. G. (1976a). J. Ultrustrucr. Res. 55, 157. Locker, R. H., and Leet, N. G. (1976b). J. Ultrustrucr. Res. 56, 31. Locker, R. H., and Wild, D. J. C. (1984). Meat Sci. 11, 89. Locker, R. H., and Wild, D. J. C. (1986). J. Biochem. 99, 1473. Lowey, A. G., Wilson, F. J . , Taggart, N. M., Grene, E. A., Frasca, P., Kaufman, H. S., and Sorrell, M. J. (1983). Cell Motil. 3, 463. Lusby, M. L., Radpath, J . F., Parrish, F. C., Jr., and Robson, R. M. (1983). J. FoodSci. 48, 1787. McNeill, P. A., and Hoyle, G. (1967). Am. Zool. 7, 483. Magid, A,, Ting-Beall, H. P., Carvell, M., Kontis, T., and Lucaveche, C. (1984). In “Contractile Mechanisms in Muscle” ( G . H. Pollack and H. Sugi, eds.), pp. 307-327. Plenum, New York. Maruyama, K. (1966). J. Biochem. 59, 422. Maruyama, K. (1976). J. Biochem. 80, 405. Maruyama, K., and Ebashi, S. (1965). J. Biochem. 58, 13. Maruyama, K., and Kimura, S. (1985). In “Cell Motility: Regulation and Function” (H. Ishikawa, S. Hatano, and H. Sato, eds.), pp. 561-569. Univ. of Tokyo Press, Tokyo. Maruyama, K., and Yamamoto, K. (1979). In “Cross-Bridge Mechanism in Muscle Contraction” (H.Sugi and G. H.Pollack, eds.), pp. 319-328. Univ. of Tokyo Press, Tokyo. Maruyama, K., Nonomura, Y., and Natori, R. (1976). Nature (London) 262, 58. Maruyama, K., Matsubara, S . , Nonomura, Y.,Kimura, S . , Ohashi, K., Murakami, F., Handa, S . , and Eguchi, G. (1977a). J. Biochern. 82, 317. Maruyama, K., Murakami, F., and Ohashi, K. (1977b). J. Biochem. 82, 339. Maruyama, K., Kimura, S., Toyota, N., and Ohaski, K. (1980). In “Fibrous Proteins: Scientific, Industrial, and Medical Aspects” (D. A. D. Parry and L. K. Creamer, eds.), Vol. 2, pp. 33-41, Academic Press, New York. Maruyama, K., Kimura, S., Ohashi, K., and Kuwano, Y. (1981a). J. Biochem. 89, 701. Maruyama, K., Kimura, M., Kimura, S . , Ohashi, K., Suzuki, K., and Katunuma, N. (1981b). J. Biochem. 89, 7 I I . Maruyama, K., Yamada, N., Ikeya, H., and Kimura, S. (1983). In “Muscular Dystrophy: Biomedical Aspects” (S. Ebashi and E. Ozawa, eds.), pp. 201-208. Jap. Sci. SOC. Press, Tokyoldpringer-Verlag, Berlin and New York. Maruyama, K., Kimura, S . , Yoshidomi, H., Sawada, H., and Kikuchi, M. (1984a). J . Biochem. 95, 1423.

114

KOSCAK MARUYAMA

Maruyama, K., Sawada, H., Kimura, S.. Ohashi, K., Higuchi, H., and Umazume, Y. (1984b). J. CeN Biol. 99, 1391. Maruyama, K., Kimura, S.. Yamamoto, K., Wakabayashi, T., and Suzuki, T. (1985a). Biomed. Res. 6, 423. Maruyama. K., Yoshioka, T., Higuchi, H.,Ohashi, K., Kimura, S . , and Natori, R. (1985b). J . Cell Biol. 101, 2167. Maruyama, K., Ito, Y.,and Arisaka, F. (1986). J . Biochem., in press. Miiller, J. (1840). In “Handbuch der Physiologie des Menschen,” Vol. 11, pp. 59-62. J. Holscher, Coblenz. Natori, R. (1954). Jikeikui Med. J . 1, 119. Natori, Rb., Umazume, Y.,and Natori, R. (1980). Jikeikui Med. J . 27, 83. Obinata, T., Reinach, F. C., Bader, D. M., Masaki, T., Kitani, S., and Fischman. D. A. (1984). Dev. Biol. 101, 116. Offer, G., Moos, C., and Starr, R. (1973). J . Mol. Biol. 74, 653. Ohtsuki, K., Maruyama, K., and Ebashi, S. (1986). Adv. Protein Chem., in press. Ozaki, K., and Maruyama, K. (1980). J. Biochem. 88, 883. Price, M., and Sanger, I. W. (1979). J . Exp. Zool. 208, 263. Pringle, J. W. S. (1978). Proc. R . SOC. London Ser. B M1, 107. Reinach, F. C., Masaki, T., Shafig, S. A., Obinata, T., and Fischhan, D. A. (1982). J . CellBiol. 95, 78.

Saide, J. D. (1981). J . Mol. Biol. 153, 661. Seki, N., and Watanabe, T. (1984). J . Biochem. 95, 1161. Sjostrand, F. S. (1962). J . Ulrrusfrucf.Res. 7, 225. Stossel, T. P., and Hartwig, J. H. (1975). J . Biol. Chem. 250, 5706. Tokuyasu, K. T. (1983). J . CeNBiol. 97, 562. Tokuyasu, K. T., Maher, P. A., and Singer, S. J. (1984). J . Cell Biol. 98, 1961. Trinick, J. A. (1981). J . Mol. Biol. 151, 309. Trinick, J. A,, Knight, P., and Whiting, A. (1984). J . Mol. Biol. 180, 311. Urry, D. W. (1984). J . Protein Chem. 3, 403. Wang, K. (1977). Biochemisrry 16, 1857. Wang, K. (1982). In “Methods in Enzymology” (D.W. Frederiksen and L. W. Cunningham, eds.), Vol. 85, pp. 264-274. Academic Press, New York. Wang, K. (1984). In “Contractile Mechanisms in Muscle” (G. H. Pollack and H. Sugi, eds.), pp. 439-452. Plenum, New York. Wang, K. (1985). In “Cell and Muscle Motility” (J. W. Shay, ed.), Vol. 6, pp. 315-369. Plenum, New York. Wang, K., and Ramirez-Mitchell, R. (1983). J. Cell Biol. 96, 562. Wang, K., and Ramirez-Mitchell, R. (1984). Biophys. J. 45, 392a. Wang, K., and Williamson, C. L. (1980). Proc. Null. Acud. Sci. U.S.A. 77, 3254. Wang, K., Ash, J. F., and Singer, S . J. (1975). Proc. Nurl. Acud. Sci. U.S.A. 72, 4483. Wang, K., McClure, J., and Tu, A. (1979). Proc. Nutl. Acud. Sci. U.S.A. 76, 3698. Wang, K., Ramirez-Mitchell, R.,and Palter, D. (1984). Proc. N d . Acud. Sci. U.S.A. 81, 3685. Wang, S. M., and Greaser, M. L. (1985). J . Muscle Res. Cell Motil. 6 , 293. Wang, S. M., and Greaser, M. L. (1986). In preparation. Wang, S. M., Schultz, E., and Greaser, M. L. (1984). J. Cell Biol. 99, 436a. Yoshidomi, H., and Maruyama, K. (1985). Zool. Sci. 2, 925. Yoshidomi, H., Ohashi, K., and Maruyama, K. (1985). Biomed. Res. 4, 207. Yoshioka, T., Natori, R., and Umazume. Y. (1981). Jikeikui Med. 28, 153.

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 104

Cell Interactions during the Seminiferous Epithelial Cycle MARTTIPARVINEN, KIMMOK. VIHKO,AND JORMATOPPARI Institute of Biomedicine, Department of Ancitomy, University of Turku, SF-20520 Turku, Finland

I. Introduction Spermatozoa belong to the most differentiated cells of the body. Their special features are a haploid number of chromosomes, a tightly packed inactive form of the chromatin, a very small amount of cytoplasm, and an ability of independent movement by a flagellum. The development of spermatozoa in the seminiferous epithelium includes three main phases: spermatogonial multiplication, meiosis, and spermiogenesis (Fig. IA). Cells in these phases are called spermatogonia, spermatocytes, and spermatids, respectively. The seminiferous epithelium also contains somatic Sertoli cells that completely surround spermatocytes and spermatids; all nutrients and hormones influencing spermatogenic cell development must pass these cells after the onset of meiosis. Pituitary follicle-stimulating hormone (FSH) and androgens secreted by the Leydig cells are the main hormones that regulate spermatogenesis (Steinberger, 1971). Sertoli cells contain receptors for these hormones and are generally considered their only targets in the seminiferous epithelium (Fritz, 1978; RitzCn et al., 1981). Spermatogenic cells perhaps are not directly influenced by FSH and androgens, but are dependent on factors that are produced by Sertoli cells under stimulation by these hormones. Today, the local control mechanisms are poorly understood, but there is an increasing amount of evidence about their existence. The developing spermatogenic cells are not randomly arranged in the seminiferous epithelium but form associations with constant composition (Fig. 1B ) . At the bottom, Sertoli cells and spermatogonia have direct contact with the basal lamina. Spermatocytes form the next layer and haploid spermatids are found at the top of the epithelium. Given cell types are always found together in the epithelium. This gives rise to the stages of the cycle of the seminiferous epithelium, defined most accurately by the morphology of the developing acrosomes and of the nuclei of the young spermatids (Leblond and Clermont, 1952; Fig. 1B).The stages follow each other along the seminiferous tubules in a wave1961). Experiments with seminiferous tubule preparalike fashion (Perey et d., tions isolated by a transillumination-assisted microdissection procedure (Fig. 2; I I5 Copyrig,h( 0 IYK6 by Academic Press. Inc. All rights oI reprluclion in any form reserved.

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FIG. 1 . (A) Phase contrast photomicrographs of living spermatogenic cells of the rat in different stages of their development. (From Parvinen, 1982, reproduced by copyright permission from The

REGULATION OF THE SEMINIFEROUS EPITHELIUM

I17

Parvinen and Vanha-Perttula, 1972; Parvinen, 1982) have revealed several biochemical parameters that have a cyclic variation suggesting that the function of the Sertoli cells is influenced by the associated spermatogenic cells. The current knowledge about the local regulation of the seminiferous epithelium at different stages of the epithelial cycle is summarized in this review.

11. Spermatogenic Cell Types and Their Metabolism

Spermatogonia with 2C amount of DNA (Fig. IA) multiply mitotically and form a stem cell population for spermatogenesis. After six successive mitoses, meiosis starts with a replication of DNA in preleptotene spermatocytes. During the prophase of meiosis, characterized by 4C amount of DNA, the homologous chromosomes pair, crossing-over occurs, and finally the cells divide into two successive meiotic divisions without DNA replication between, to form spermatids with a haploid (IC) amount of DNA. The nucleus of the spermatids becomes attached by a flagellum, transforms, and the chromatin eventually condenses. Finally the spermatozoon is released from the seminiferous epithelium at sperniiation. A. SPERMATOGONIAL DEVELOPMENT Spermatogonia are located in the basal compartment of the seminiferous epithelium, separated from the adluminal compartment by tight junctions between

Endocrine Society.) Spermatogonial development includes type A (A), intermediate (In), and type B (B) spermatogonia. characterized by a diminishing size of the nuclei and an increasing amount of heterochromatin attached to the nuclear envelope. Meiosis starts at preleptotene (PI) and includes leptotene (L), zygotcnc (Z), and pachytene (P) stages. During early pachytene (eP), the size of the nuclei is similar to zygotene, but at mid-pachytene (mP), an enlargement of the nuclei with increasing diffuseness of the chromosomes takes place. Late pachytene (IP) is characterized by large nuclei that eventually undergo first (div) and second meiotic divisions, with a short interphase (secondary spermatocyte) between to produce haploid spermatids. These differentiate through steps (1- 19) that are recognized by the morphology of the acrosomic system (arrow in 3). and of the nuclei. The step 17 spermatids are depicted at a lower magnification ( X 3 5 0 ) in order to demonstrate their typical arrangement in bundles; all other cells are magnified X1500. Residual bodies (rb) that become separated from the testicular spermatozoa (sz) at spermiation are phagocytosed by the Sertoli cells. (B) The "road map" of spermatogenesis. (From Dym and Clermont, 1970, reproduced by copyright permission from Alan R. Liss, Inc.) The 14 cell associations or stages of the cycle of the seminiferous epithelium are defined according to criteria of Leblond and Clermont (1952, for details, see Table I). The letters and figures designating the different cell types are the same as in A, except that Di indicates diakinesis-stage primary spermatocytes, I1 indicates secondary spermatocytes, and m indicates the mitotic peaks of spermatogonia at given stages of the cycle.

DARK-

PALE-

WEAK

SPOT-

STRONG SPOT-

DARK ZONE

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neighboring Sertoli cells (Dym and Fawcett, 1970). These junctions form perhaps the most effective part of the blood-testis barrier that restricts a free passage of a number of compounds (Setchell, 1978). Spermatogonia are divided in two main types, noncycling (A,) and the ones that differentiate into spermatocytes (Clermont and Bustos-Obregon, 1975; Dym and Clermont, 1970; Clermont and Hermo, 1975). Type A, spermatogonia are resistant to ionizing radiation and repopulate the seminiferous epithelium after a dose of X-irradiation that selectively destroys cycling spermatogonia (Dym and Clermont, 1970). It has been proposed that a chalone secreted by the spermatogonia normally prevents the A, spermatogonia from entering the cell cycle (Clermont and Mauger, 1974). The cells that multiply mitotically and provide the stem cell population for meiosis and spermiogenesis are called type A,-,, intermediate, and type B spermatogonia. Type A spermatogonia have a pale dustlike chromatin, while type B spermatogonia contain accumulations of heterochromatin attached to their nuclear envelope. Intermediate-type spermatogonia, found in rodents, but not in man, have an amount of heterochromatin that is between the type A and B spermatogonia (for review, see Clermont, 1972). B. MEIOSIS The DNA replication of preleptotene spermatocytes in late stage VII and during stage VIlI of the cycle (Monesi, 1962; Hilscher, 1967; Clermont, 1972) is considered the onset of meiosis. During preleptotene, the spermatocytes start to lose their contact with the basal lamina through a penetration of Sertoli cell processes that meet each other and form tight junctional complexes at stage IX Fiti. 2. A freshly isolated unstained rat seminiferous tubulus in transmitted light under a stereomicroscope at a magnification of X 3 5 (top panel, from Parvinen and Soderstrom, 1977, reproduced by permission from Elsevier/North Holland Biomedical Press). The middle panel shows four main transillumination patterns (magnification X 100) with corresponding histology ( X 4 0 0 ) from stages XI. I, 111, and VII of the cycle (middle, from Parvinen and Ruokonen, 1982, reproduced by permission from J. B. Lippincott Company). In the bottom panel, a scheme of the transillumination-assisted microdissection of rat seminiferous tubules is presented (from Parvinen, 1982, reproduced by copyright permission from The Endocrine Society). The point of spermiation at stage VIII of the cycle is recognized by an abrupt stop of the dark absorbing center of the tubule. A pale absorbing zone covers stages IX-XII; the elongating nuclei of the spermatids have not condensed (XI). The weak spot zone covers stages XIII-I; the steps 13-15 spermatids with condensed nuclei are arranged in bundles, These bundles increase in density and plunge deep into the Sertoli cells in stages 11-V and give rise to a strong spot absorption zone (111). At stage VI the bundle arrangement is released and the maturing step 19 spermatids become located at the top of the seminiferous epithelium inducing a dark homogeneous central absorption (VII). The scheme at the bottom indicates the pools of stages that have been collected for biochemical studies of the cycle. For further details, see Table I.

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(Russell, 1977a). After this step, spermatogenic cells become completely surrounded by the Sertoli cells for the rest of spermatogenesis. The leptotene stage of meiosis is characterized by a finely beaded filament configuration of the chromosomes. Tight junctions of Sertoli cells are found both below and above the leptotene spermatocytes that are located in the intermediate Compartment of the seminiferous epithelium (Russell, 1977a). In stage XII, the junctions above the spermatocytes that now have reached the zygotene stage of meiosis disappear and the cells become located in the adluminal compartment of the seminiferous epithelium. During zygotene the homologous chromosomes pair and become connected by a synaptonemal complex (Rasmussen and Holm, 1980). A rotatory movement of the chromosomes (Parvinen and Soderstrom, 1976a) that is regulated by a colcemid-sensitive mechanism in the nuclear envelope (Salonen et al., 1982) has been suggested to be involved in the pairing. Pachytene starts when the pairing of the chromosomes is completed. Crossing-over takes place probably during early pachytene (Hotta et al., 1977). In mid-pachytene spermatocytes at stages VI and VII of the cycle the chromosomes become diffuse in their structure. At the end of meiosis, the chromosomes partly split during a short diplotene step (Di), and this is followed by two reduction divisions in short succession without an interphase DNA synthesis. The first meiotic division is characterized by large division figures that result in formation of secondary spermatocytes (11). They resemble young spermatids morphologically but are larger. After the second meiotic division, one primary spermatocyte gives rise to four haploid spermatids. C. SPERMIOGENESIS

Spermiogenesis is a successive series of events in which the haploid cells undergo nuclear transformation and condensation of the chromatin and develop special structures such as acrosome on the anterior head of the nucleus and a flagellum attached on its posterior surface. According to morphological criteria, spermiogenesis can be divided in Golgi (steps l-3), cap (4-7), acrosome (8-121, and maturation (13-19) phases (Leblond and Clermont, 1952; Fig. 1A and B). Steps 1-7 spermatids are transcriptionally active as detected by [3H]uridine autoradiography (Monesi, 1964; Soderstrom and Parvinen, 1976a). During the same steps, acrosome is formed to the spermatids. It is a cap-like structure rich in glycoproteins, and it is thought to have a function in the penetration of the spermatozoon into the ovum. The Golgi complex participates in the formation of the acrosome. Shortly after the meiotic divisions, a defined Golgi complex is assembled in the spermatids (Soderstrom and Parvinen, 1978). Simultaneously, a characteristic cytoplasmic organelle, the chromatoid body, obtains its typical lobulated shape. Toward the end of step 1, the acrosomic vesicle is formed. The morphology of the spermatid nuclei and of

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the developing acrosomes are used as “markers” of the cell associations or stages of the cycle of the seminiferous epithelium (Leblond and Clermont, 1952; Table I). Several other classifications of the seminiferous epithelium are based on histological arrangement of the maturing spermatids (reviewed by Courot et al., 1970), but they are not practical for cytological analyses. During elongation of the spermatid nuclei, their protein composition changes. Histones are present in steps 1-8 spermatids, but not after step 12. They are replaced by nuclear proteins designated TP and TP2 characteristic for steps 1315. These become replaced during steps 16- I9 by basic nuclear protein TP3 and by the sperm basic nuclear protein S I (Grimes et al., 1977; Meistrich et al., 1978a). These changes correlate with the ultrastructure of the chromatin (Kierszenbaum and Tres, 1975). During the first change of the nucleoproteins, the beaded structure of thc chromatin is replaced by a smooth type. Later, changes in structure and chemical composition lead to side-by-side alignment of the chromatin fibers that facilitates the packaging of the male gamete genome (Kierszenbaum and Tres, 1978). During step 15 of rat spermiogenesis, a reduction in phosphotungstic acid stainability probably also reflects the second nucleoprotein transition (Courtens and Loir, 1981). The chromatin of the spermatozoon further condenses during the epididymal transit where an increasing number of the disulfide bonds has been observed (Calvin and Bedford, 1971). The flagellum develops from the centriolus early during spermiogenesis. In living cells, its movements have been observed during late step I . It becomes attached to the nucleus around steps 6 and 7. A characteristic feature of late acrosome and early maturation phase spermatids is their arrangement in bundles containing 6- 1 1 (average 8) cells (Wing and Christensen, 1982). This arrangement starts at stage XI1 and is released at stage VI of the cycle. During stages IV and V the penetration into the Sertoli cells is deepest. The reason for this arrangement of the spermatids is not known, but it may be associated with the final maturation of the nuclei and of the acrosomes.

111. Cycle, Wave, and Transillumination of the Seminiferous Epithelium

In the seminiferous epithelium, one or two generations of spermatogonia along the basement membrane, one or two generations of spermatocytes, and one or two generations of spermatids bordering the lumen of the seminiferous tubule evolve synchronously through the whole spermatogenesis. The various generations form cellular associations, also called stages, appear, and follow each other in a given area of the seminiferous tubule in a fixed sequence. In different animal species the number of typical cell associations may vary depending on the arbi-

122

MARTTI PARVINEN ET AL. TABLE 1 CRITERIA FOR IDENTIFICATION OF DIFFERENT STAGESOF THE CYCLEOF RAT SEMINIFEROUS EPITHELIUM('

Stage I

11

111

IV

V

VI

VII

VIII

Duration (days)"

Average length (mm)"

Trdnsihnination and microdissection"

I .7

2.6 (0.1-7.2)

Definite weak spots in tubular center

0.8

I .2 (0.2-4.5)

Onset of strong spot zone, development of peripheral stripes

0.4

0.6 (0.1-5.0)

11 and 11 together form the first half of

Flattening of the acrosomic granule against nucleus; maximal penetration of step 17 spermatid bundles into Sertoli cells Head cap formation in young spermatids

0.5

0-9

I .3

I .4 (0.2-5.7)

Head cap covers onequarter to one-third of nuclear circumference; step 18 spermatids lose their bundle arrangement and move centripetally Maximal size of the head cap; step 19 spermatids at the top of the epithelium Polarization of nuclei and peripheral orientation of acrosomes of step 8 and spermiation of step 19 spermatids

I .o

I .5 (0.1-6.2)

Criteriab Newly formed spermatids develop idiosomes; step 15 spermatids in loose bundles Appearance of proacrosomic granules in young spermatids; movement of late spermatid bundles toward periphery Formation of a single acrosomic granule

strong spot zone Strong spots

(0.1-3.9)

2.3

3.L

(0.3-9. I )

1.2

1.4 (0.2-6.2)

Stages IV and V form the second half of strong spot zone Fusion of strong spots to homogeneous dark central absorption

Homogeneously dark tubular center

Abrupt cessation of dark absorption at spermiation

REGULATION OF THE SEMINIFEROUS EPITHELIUM TABLE I

Stage

IX X XI

XI1

XI11

XIV

Criteriah

(Conrinued)

Duration (days)“

Initiation of spermatid elongation

0.3

Slightly elongated spermatid nuclei Much elongated spermatid nuclei Straight, narrow, and darkening spermatid nuclei with onset of bundle arrangement Curved apex of the spermatids; definite bundle arrangement Dividing primary and secondary spermatocytes

0.3

Average length (mm)” 0.8 (0.1-3. I )

0.3 1.3

I23

0.4 (0. I - I .9) 0.6 (0.1-6.0) I .s (0.1-6.8)

0.8

0.8 (0.1-6. I )

0.7

0.8 (0.1-3.3)

Transillumination and rnicrodissectione Pale absorption, peripheral lipid droplets Pale absorption Pale absorption Late half of the pale zone

Appearance of weak spots in tubular center XI11 and XIV comprise the first half of weak spot zone

“In histological sections stained with PAS-hematoxylin-stained sections, living cell preparations, and in freshly isolated unstained seminiferous tubules. bAs defined by Leblond and Clermont (1952). “According to Clermont and Harvey (1965). ”According to Perey C t d.(1961). ‘Parvinen and Vanha-Perttula ( 1 9 7 2 ~Parvinen and Ruokonen (1982). and Parvinen (1982); Fig. 2.

trary criteria used for their identification (for references, see Courot et a / ., 1970; Clermont, 1972). A clear distinction should be made between the “cycle” and “wave” of the seminiferous epithelium. The cycle is a dynamic histological phenomenon in time at any one area of the seminiferous tubule, whereas the wave refers to the succession of the different cellular associations along the seminiferous tubules at any time. Each stage of the cycle has a constant duration. A complete cycle, i.e., the time required for development of a cell to a next more advanced type in the same cell association, lasts 12.0 days in the Sherman rat (Clermont e t a / ., 1959) and 12.9 days in the Sprague-Dawley rat (Clermont and Harvey, 1965). The length of the wave of rat seminiferous epithelium (the distance of two identical cell associations with all other stages between) has been shown to vary from 0.55 to 4.25 cm (average 2.33 cm) and 80% of all waves contain modulations (Perey e t a / . , 1961). When traced from the rete testis, the cell associations follow each other in descending order without jumps (continuity of the segmental order).

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MARTTI PARVINEN ET AL.

Modulations are local reversals to an ascending order. Instead of a next lower number, a segment may be followed by one with a next higher number so that two or more segments are arranged in an ascending order in a distal direction. Thereafter, the descending pattern is resumed without breach in the continuity of the segmental order (for details, see Perey et al., 1961). Biochemical and endocrinological studies of the wave of the seminiferous epithelium have become possible along with the transillumination technique in which variations in light absorption allow the recognition of different segments of the epithelial wave in freshly isolated seminiferous tubules (Parvinen and Vanha-Perttula, 1972; Fig. 2). An increased light absorption is associated with the condensation of the chromatin of the spermatid nuclei at step 12 of spermiogenesis. Therefore, stages IX-XI1 have a pale absorption, and around the border of stages XI1 and XIII, weak spots in transilluminated seminiferous tubules indicate a condensation of step 13 spermatid nuclei and their concomitant arrangement in bundles with increasing density. Stages XIII-I of the cycle are characterized by weak spot absorption type, and a marked increase of the density of the spots occurs at stage 11, concomitantly with a deep penetration of the step 16 spermatid bundles into the Sertoli cells and an increase of their absorption probably due to the development of the outer dense fibers into their flagella. Stages 11-V of the cycle are characteiized by a strong spot absorption pattern due to a deep penetration of steps 16-17 spermatid bundles into the seminiferous epithelium. At stage V1, the bundle arrangement of the step 18 spermatids is released and the cells move centripetally in the seminiferous epithelium to be located at its top at stages VII and VIII. This is reflected by a transition of the dark absorbing spots to a homogeneously dark center of the transilluminated seminiferous tubules. At the site of spermiation in stage VIII of the cycle, the dark absorption abruptly stops and the pale absorbing zone of the seminiferous tubules reappear. Taking the relative average lengths of the various segments of the wave of the seminiferous epithelium in account, a transillumination-assisted microdissection method has been developed (Parvinen and Ruokonen, 1982). Seminiferous tubule segments at defined stages of the cycle can be pooled and collected in amounts that are sufficient for a variety of biochemical analyses (5-10 cm, equivalent to -5-10 mg wet weight of tissue) during a 1- to 3-hour dissection time. The pools represent stages I, 11-111, IV-V, V1, V1Ia+, VIIc-d, VIII, IXXI, XII, and XIII-XIV (Fig. 2). Transillumination is not accurate for identification of all stages of the cycle. Stages VI, VII, VIII, and IX, owing to their characteristic absorption pattern, can be isolated with great accuracy, but errors up to -+3 stages may occur in the region of stage XI1 (Table 11). A reason for this may be that stages IX, X, and XI are very short (average lengths 0.8, 0.4, and 0.6 mm, respectively). If a precise identification is needed, the “marker cells” can be identified during the separation process by phase contrast microscopy.

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TABLE 11 ACCURACYOF T H t TRANSILLUMINATION ASSISTHI MICRODISSECTION M ~ T H O DA" S CONTROLLtD BY H~STOLOGICAL EXAMINATION* Correct

Transillumination

I 11-111

IV-v VI Vll,-h VII, -* VIII IX-XI XI1 XIII-XIV

I1

38 23 26 in 23 24 18

31 35 21

Histology XIII-11 XIV-I11 111-v v-VI vI-vIId VIIa-V1Id VIId-VIII IX-XI1 IX-XI11 XII-I

(%)

50 65 96 I8 74 61 72 90 51 51

UParvinen and Ruokonen (1982). bLeblond and Clermonl (1952).

Short segments (<0.5 mm) of seminiferous tubules are carefully squashed between glass slides and the excess fluid is blotted by a piece of lens paper. The spermatogenic cells float out from the tubule segment and eventually form a monoiayer of slightly flattened cells that allow an accurate recognition (Soderstrom and Parvinen, 1976a, 1978). This isolation procedure is practical only for experiments that can be carried out with 1- to 2-mm samples of seminiferous tubules. The details of the transillumination-assisted microdissection method with correlations to the morphological criteria of the seminiferous epithelium are summarized in Table 1.

IV. Transcriptional Activity during Spermatogenesis and the Function of the Chromatoid Body The first observations about cellular distribution of RNA during spermatogenesis were made by Daoust and Clermont (1955) by histochemical pyronin staining technique. Spermatogonia showed a clear positive reaction, whereas the affinity of the cytoplasm for the stain gradually decreased and was low at the beginning of pachytene. In mid-pachytene the reaction increased to be again gradually decreased during late pachytene, diplotene, in secondary spermatocytes and spermatids. In the young spermatids the chromatoid body was intensively stained with pyronin. Direct evidence about the cellular synthesis of RNA was obtained with [ 'Hluridine labeling and autoradiography . Monesi

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MARTTI PARVINEN ET AL.

(1964, 1971) found a high rate of RNA synthesis in type A spermatogonia of the mouse. It decreased in intermediate and type B spermatogonia and no labeling was found in leptotene, zygotene, and early pachytene primary spermatocytes. In mid-pachytene, the RNA synthesis rapidly increased but decreased again toward late pachytene and diakinesis. Cells at meiotic divisions did not incorporate radioactivity from [3H]uridine. The secondary spermatocytes and early spermatids (steps 1-7) were slightly labeled but after condensation of the chromatin, no labeling could be observed. The labeling pattern is similar in the golden hamster (Utakoji, 1966) and rat (Soderstrom and Parvinen, 1976a), but in the ram maximal RNA synthesis appears later during diplotene (Loir, 1972). In contrast to the autosomes, the XY bivalent remains unlabeled during meiosis (Monesi 1964, 1971; Utakoji, 1966). It was originally believed that the nucleolus also remains unlabeled during the prophase of meiosis. This was interpreted as an indication of an inhibition of the ribosomal RNA synthesis (Henderson, 1964; Das et al., 1965; Utakoji, 1966). It was later found that the lack of [3H]uridinelabeling of the nucleoli associated with the XY bivalent is due to the location of rRNA genes in autosomes (Henderson et al., 1972). During the zygotene and early pachytene stages of the meiotic prophase, autosomes synthesize rRNA (Kierszenbaumand Tres, 1974a), and [3H]uridinelabel can be seen in pachytene XY -bivalents if the label has been introduced during zygotene-early pachytene (Kierszenbaum and Tres, 1974a). High resolution autoradiographic studies have confirmed and extended the light microscopic observations (Kierszenbaum and Tres, 1974a,b, 1975). It was also found that the pachytene chromosomes with typical lampbrush loops are sites of nonribosomal RNA synthesis (Kierszenbaum and Tres, 1974b). Biochemical studies of RNA synthesis during meiosis and spermatogenesis are few. Muramatsu et al. (1968) found that a large proportion of the RNA synthesized in whole-testis preparations had a high molecular weight and a rapid turnover. The first attempt to study testicular RNA synthesis in specific cell types was made by Galdieri and Monesi (1974) who selectively destroyed spermatogonia by 300 rads of X rays and studied the ribosomal RNA synthesis in spermatocyte-enriched testis. The low rate of rRNA synthesis was also observed in this study. Pachytene spermatocytes and round spermatids isolated after labeling with radioactive RNA precursors contained both ribosomal and poly(A) RNA (mRNA) (Geremia ef al., 1978) that was considered as evidence of gene expression in haploid spermatids. Recently, a-tubulin gene has been shown to be specifically expressed during the haploid phase in the mouse spermatogenesis using a cDNA probe (Distel et al., 1984). Therefore, genes responsible for the morphogenesis and structural components of the spermatozoon are at least partly expressed during the haploid phase. It is likely, however, that many mRNAs coding the proteins of the spermatozoon are synthesized before meiotic divisions in the spermatocytes and stored in long-lived form for translation during late spermiogenesis where the genome is inactive. +

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A storing organelle for the long-lived mRNA may be the chromatoid body that is a lobulated cytoplasmic structure with 1-2 pm in diameter unique for spermatogenic cells (for review, see Sud, 1961a). It was first described by Benda (1891) and suggested to be derived from nucleus or nucleolus. This origin was also suggested by electron microscopic observations (Comings and Okada, 1972). On the other hand, a cytoplasmic origin of the chromatoid body has been proposed (Sud, 1961a; Fawcett et al., 1970; Fawcett, 1972), since during its first appearance in pachytene spermatocytes at stage ViII of the rat seminiferous epithelial cycle (Russell and Frank, 1978), the chromatoid material is intimately associated with mitochondria. During pachytene and diplotene stages of the meiotic prophase it grows and specific intranuclear membrane modifications may be involved in the transport of material from the nucleus to the chromatoid body (Russell, 1977b). During meiotic divisions, the chromatoid body has been found to be dispersed in the cytoplasm as small 30-nm granules (Russell and Frank, 1978) and condenses to a prominent lobulated form at the end of step 1 of spermiogenesis (Soderstrom and Parvinen, 1978). During early spermiogenesis the chromatoid body is located on the surface of the nucleus close to the Golgi complex and the developing acrosome (Susi and Clermont, 1970). In living cells, it has been shown to move rapidly and to have frequent contact with the Golgi complex (Parvinen and Jokelainen, 1974). The chromatoid body is apparently dependent on the function of the haploid genome during early spermiogenesis (L. M. Parvinen et al., 1978; Soderstrom, 1977). Radioactivity derived from [ 3H]uridine incorporates to the chromatoid body clearly after nuclear labeling (Fig. 3B) both in spermatocytes (Soderstrom and Parvinen, 1976b; Soderstrom, 198 1) and in spermatids (Soderstrom and Parvinen, 1976b) suggesting an incorporation of RNA into this organelle. This is in accordance with observations about an increased amount of nuclear pore complexes in the vicinity of the chromatoid body (Fawcett et al., 1970) and material continuities observed through nuclear pores between chromatoid body and intranuclear material (Soderstrom and Parvinen, 1976c; Parvinen and Parvinen, 1979; Fig. 3A). High-resolution immunohistochemical studies have indicated the presence of actin and RNA in the chromatoid body (Walt and Armbruster, 1984). Later in spermiogenesis, the chromatoid body moves to the area of the attachment of the flagellum to the spermatid nucleus (at step 7), where it forms a ringlike structure (Fawcett et al., 1970). Toward the end of spermiogenesis, the chromatoid body diminishes in size and is found in the region of the developing flagellum. It becomes removed from the spermatozoon at spermiation in the residual body. The chromatoid body obviously has several functions during spermiogenesis. The interaction with the Golgi complex (Soderstrom and Parvinen, 1976c) suggests that it may be involved in the formation of the acrosomic system. Perhaps the most interesting of the proposed functions of the chromatoid body is its

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Fic. 3. (A) Electron micrograph of the chromatoid body (arrow) in rat step 3 spermatid. It is located at the proximity of an outpocketing in the nuclear envelope. Time-lapse cinemicrographic analysis of living spermatids in the same step revealed that this location is very transient. Material continuity is seen through a nuclear pore complex to the chromatoid body. (From Parvinen and Parvinen, 1979. Reproduced from The Journal of Cell Biology, 1979, 80, 621-623 by copyright ; I pm. (B)Electron permission of The Rockefeller University Press.) Magnification, ~ 2 6 , 0 0 0 bar, microscopic autoradiogram of a rat step 1 spermatid labeled for 2 hours with ["Iuridine and chased for 14 hours in the presence of nonradioactive nucleotide precursors. The chromatoid body is labeled and some grains are seen in the nucleus (n). No grains are seen in the cytoplasm outside the chromatoid body, except in the space between the nucleus and the chromatoid body (arrow). (From SMerstrom and Parvinen, 1976b. reproduced by copyright permission from The Rockefeller University Press.) Both of these observations suggest that the chromatoid body incorporates RNA in a storage that may be necessary for coding the protein synthesis during late spermiogenesis when the genome of the spermatid becomes inactive. Magnification, X9000; bar, I pm.

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participation in the RNA metabolism of the spermatogenic cells. Although this is far from clearly understood, there is evidence that a considerable proportion of the RNA synthesized in pachytene spermatocytes is preserved through spermatid development until late spermiogenesis (Geremia et al., 1978). The chromatoid body may be a storing organelle for this long-lived mRNA which may direct the protein synthesis during late spermiogenesis when the genome of the spermatid is inactive. It is also possible that presynthesized proteins are stored in the chromatoid body, to be assembled in spermatid structures at appropriate times. Studies on these problems await techniques such as labeling with specific antibodies and cDNA probes.

V. Cyclic Interaction between Sertoli Cells and Spermatogenic Cells Several morphological observations suggest that Sertoli cells might have a cyclic function depending on the stage of the cycle of the seminiferous epithelium. Leblond and Clermont (1952) found the nuclei of the Sertoli cells in stages IX-XIV flattened parallel to the basement membrane, whereas some of them during stages I-VIII became triangular or oblong with their long axis perpendicular to the basal lamina. Shortly after phagocytosis of the residual bodies, the lipid content of the Sertoli cells rapidly increases (Lacy, 1960; Niemi and Kormano, 1965; Posalaki er al., 1968; Kerr and de Kretser, 1975) to stay high during stages IX-XIV and to decline in stages I-VIII (Kerr et al., 1984). The distribution of some other organelles also supports the concept of the cyclic function of the Sertoli cells. The volume density of the flattened type of the endoplasmic reticulum was greater in stages VII and VIII of the cycle than elsewhere (Assaf, 1980) coinciding with the minimum volume density of the Sertoli cell vesicles (Ulvik and Dahl, 1981). Several enzyme activities of the Sertoli cells have been shown to vary during the cycle of the seminiferous epithelium. The maximal activity of acid phosphatase appeared at stages VII and VIII around the tubular lumen, but little activity was found in stages IX-11. In stages III-V, during the maximal penetration of maturing spermatids into the Sertoli cells, a temporary increase in the acid phosphatase activity could be observed (Niemi and Kormano, 1965). Hydrolytic enzymes of the Sertoli cells show high activity in stages I-VIII, coinciding with their low lipid content (Posalaki et al., 1968). A similar distribution with acid phosphatase has been observed in thiamine pyrophosphatase in the Sertoli cells (Hilscher et al., 1979). Isoenzymes of acid phosphatase do not show similar cyclic distributions. While acid phosphatase IV typical for mature seminiferous tubules in the rat has a clear peak activity in stage VII of the cycle, acid phosphatase 111 shows an even

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distribution throughout different stages of the cycle (Parvinen and Vanha-Perttula, 1972). Aminopeptidase I11 shows a cyclic distribution in its activity with a maximum around stages IV-VIII, although this cycle is not as prominent as that of acid phosphatase IV (Parvinen and Vanha-Perttula, 1972). Acid phosphatase IV has recently been found to be located mainly in round spermatids (T. VanhaPerttula and A. R. Bellvt, personal communication). A. LOCALACTIONOF FOLLICLE-STIMULATING HORMONE Several observations suggest that the responsiveness of the Sertoli cells to circulating hormones is modified during the appearance of advanced spermatogenic cells at puberty. In spite of an increase in the number of the FSH receptors, there is a marked fall in the FSH-stimulated cAMP production (Steinberger et al., 1978). Simultaneously, the activity of aromatase, an enzyme that converts testosterone to estradiol, falls rapidly while the production rate of androgen-binding protein (ABP) rises (Ritzkn el al., 1981). An explanation for these observations is that the spermatogenic cells directly modify the activity and hormone responsiveness of their neighboring Sertoli cells. Although the adult seminiferous tubules have a small response to FSH stimulation (Fritz, 1978; RitzCn er al., 1981), a significant variation occurs both in the binding of FSH and in the FSH-stimulated cyclic AMP production when tubules at different stages of the cycle are compared (Parvinen et al., 1980). FSH is maximally bound at stage I and minimally in stages VI and VII (Parvinen er al., 1980; Purvis er al., 1984; Fig. 4). The adenylate cyclase of the Sertoli cells shows a maximal response to FSH in stages 11-111 (Gordeladze er al., 1982). Similar distribution is found in the FSH-stimulated secretion rate of CAMP. Stages I-V have a clear response while stages VII and VIII are virtually insensitive (Parvinen er al., 1980; Purvis er al., 1984; Fig. 4). The pattern of the FSHstimulated cAMP secretion is almost identical with the distribution of Mn2 dependent adenylate cyclase (Gordeladze et al., 1982; Fig. 4), a soluble enzyme localized specifically in spermatids (Gordeladze and Hansson, 198 1). Cyclic nucleotides together with protein carboxylmethylation have been suggested to have a role in regulating sperm motility (de Turner et al., 1978; Gagnon er al., 1979). Protein carboxylmethylase is specifically located in haploid cells and in the sperm tails (Braun and Dods, 1975; Bouchard et al., 1980). The distribution of this enzyme throughout different stages of the cycle of the seminiferous epithelium is almost identical with that of the Mn2 -dependent adenylate cyclase (Cusan et al., 1981; Fig. 5). A phenomenon that has a very good correlation with FSH-stimulated cAMP production and with adenylate cyclase activities during the cycle of the seminiferous epithelium is the location of the maturation phase (steps 15-17) sperrnatid bundles deep in the Sertoli cell cytoplasm close to their nuclei. This may +

+

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2 100 V

D C

i

1

IV-VVllla-b

Vlll

XI1

FIG. 4. Summary of the data supporting the concept that stages XIII-V of the cycle of rat seminiferous epithelium are preferentially regulated by FSH. (From Parvinen cf a / . . 1985, reproduced by copyright permission from INSERM.) Maximal binding of FSH occurs in stages XIII-I (solid circles) and in stages X U - V I it is significantly (p < 0.01) higher than in stages VII-XI1 (Parvinen et a / . , 1980). In the presence of phosphodiesterase inhibitor (MIX), FSH stimulates the production of cyclic AMP significantly more in stages 11-VI than in other stages (open circles). A significant stimulation over the basal level was also found in stages IX-I, but not in stages VII and VIII (Parvinen el a / . . 1980). This distribution is almost identical with that of the activity of Mn2+dependent adenylyl cyclase of the spermatids (open squares, Gordeladze el al., 1982).

be regulated by a cytoskeletal activity of the Sertoli cells, which is known to be regulated by cAMP and FSH (Means et al., 1978). Sertoli cells in vitro have been shown to contract when FSH or cAMP is added into the medium (Spruill et al., 1981). Furthermore, an isoelectric variant of vimentin-type intermediate filament protein, probably involved in Sertoli cell shape changes, has been shown to become phosphorylated by FSH and a cAMP analog both in cultured Sertoli (De Philip and Kierszenbaum, 1982) and in intact seminiferous tubules (Kierszenbaum et al.. 1985). In stage VI, a rapid decrease in FSH responsiveness is concomitant with the centripetal movement of the step 18 spermatids in the seminiferous epithelium and simultaneous release of their bundle arrangement. The levels of the FSH response stay low during stages VII and VIII but start to rise again in stages XII-XIII, concomitantly with the onset of the bundle arrangement of a new generation of steps 12-13 spermatids (Fig. 4). Still to be clarified is the role of the cytoskeletal components in the normal maturation and

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.-C

E LO

-zF 30 1

a

*

20

u1 0,

E

10

Q

I

11-111 VI Vllc-d IX-XI XIII-XIV IV-V Vlla-b Vlll XI1

FIG. 5 . The activity distributions of two enzymes specific for haploid spermatogenic cells, protein carboxylmethylase (open circles) and Mn2+ -dependent adenylate cyclase (solid circles), are almost identical during the cycle of rat seminiferous epithelium (Cusan er a / . , 1981). It has been suggested that both enzymes play a role in sperm motility and maturation.

release of the spermatids. Furthermore, elevated activity of adenylate cyclase may be associated with the movements of the chromatoid body and of the newly formed flagellum in the steps 2-5 spermatids.

B. ANDROGENS IN THE SEMINIFEROUS EPITHELIUM The most obvious primary targets of androgens in the seminiferous epithelium are the Sertoli cells. They contain both nuclear and cytosol androgen receptors (Mulder et al., 1975; Tindall et al., 1977; Sanborn et al., 1977) that bind to chromatin with high affinity (Tsai et al., 1980) resulting in activation of RNA polymerase activity (Lamb el al., 1981). There is evidence of androgen receptors in the nuclei of spermatogenic cells (Tsai et al., 1980), particularly in the sonication-resistant late spermatids (Wright and Frankel, 1980). Their binding activity even exceeds the mean value observed in seminiferous tubule nuclei (Frankel and Chapman, 1984). In some steps a direct androgen action may therefore occur in the spermatogenic cell themselves, but since transcription in late spermatids is minimal or absent (Monesi, 1964; Kierszenbaum and Tres, 1975), it is difficult to establish any major role for androgens in these cells on the basis of what is presently known about the mechanism of action of steroid hormones in other model systems (Chan and O’Malley, 1976). Therefore, the current concept is that androgens primarily act on Sertoli cells that, in turn,

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I33

regulate the development of spermatogenic cells (Fritz, 1978; RitzCn et a / . , I98 1). The molecular mechanisms of androgen action in the adult full-functioning seminiferous epithelium have remained poorly understood. There are several indirect observations about local androgen action in the seminiferous epithelium. Shortly after hypophysectomy, all spermatogenic cells do not degenerate simultaneously; some stages become affected first. Midpachytene primary spermatocytes and steps 7 and 19 spermatids at stages VII and VIII of the cycle are first to degenerate after hypophysectomy. This is largely prevented by substitution with leutinizing hormone (LH) and completely brought to normal low levels by administration of both LH and FSH, while FSH alone has no effect (Russell and Clermont, 1977). The same cells show specific early degeneration when the rats are treated with monospecific LH antiserum (Dym et a / . , 1977) or with agents that disrupt the hormonal stimulation of spermatogenesis (Russell et al., 1981). That cells at very different steps in the same cell association show simultaneous degeneration strongly suggests that the local conditions, mainly provided and controlled by Sertoli cells, are particularly sensitive to androgen stimulation in stages VII and VIII of the cycle of the seminiferous epithelium. This view is supported by biochemical observations. Maximal concentration of endogenous testosterone was found in stage VIII of the cycle (23.0 5.0 pg/cm), and high levels in the immediate neighborhood at stages VIIc-d and IX-XI (17.8 3.6 pg/cm) (Parvinen and Ruokonen, 1982; Fig. 6). The value at stage VIII was significantly (p < 0.001) higher than at any other stage. Lowest testosterone concentrations were found in stages XIII-V (13.3-15.0 pg/cm) that are significantly lower (p < 0.05) than the values measured in stages VI1 and IX-XI.Therefore, areas with high (stages VII-IX) and low (stages XIII-V) androgen levels can be distinguished in the seminiferous epithelium. The distribution is almost opposite with the distribution of the FSH responsiveness (compare Figs. 9 and 11). The observations suggest that the preferential androgen action occurs in stages VII-XI while stages XIII-V are preferentially controlled by FSH. A more direct indicator of androgen action in the seminiferous epithelium may be the concentration of the nuclear androgen receptor. Significantly higher concentrations were found in the nuclei isolated from stages IX-XI1 and XIII-I of the epithelial cycle than from stages 11-VI and VII-VIII (Isomaa et a / . , 1985; Fig. 6). Measurements of the androgen receptor concentration in the cytosol of the seminiferous tubule preparation were obscured by a component that bound large amounts of the 'H-labeled ligand. This binder is of interest, since it most obviously is not androgen-binding protein (ABP) that does not bind [3H]methyltrienolone (Bardin et a / . , 1981), and since it is not present in measurable amounts in primary Sertoli cell cultures or in cell lines that are derived from Sertoli or myoid cells (Nakhla et al., 1984). It may therefore be located in spermatogenic cells.

+

*

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20

11-111 I

VI IV-V

Vlk-d IX-XI XIII-XIV V l b b Vlll XI1

FIG. 6. Summary of the main data supporting the concept that stages VII-XI1 of the cycle of rat seminiferous epithelium are more influenced by androgens than other stages. (From Parvinen er al., 1985, reproduced by copyright permission from INSERM.) In stage VIII, the concentration of endogenous testosterone (solid circles) was significantly (p < 0.001) higher than in any other stage (Parvinen and Ruokonen, 1982). This distribution is similar to the secretion rate of androgen-binding protein (open circles): in stages VIII-XI it was significantly (p < 0.01) higher than in all other stages except XI1 (Ritztn et al., 1982). The concentration of nuclear androgen receptors (squares) in the seminiferous tubules was maximal in stages IX-XII, significantly (p < 0.01) higher than in stages 11-VIII (Isomaa et al., 1985).

The maximal nuclear androgen receptor concentration in stages IX-I of the cycle immediately follows the maximal endogenous testosterone concentration in stage VIII. The high androgen receptor concentration around stage XIV coincides with meiotic divisions that are critically dependent on androgen stimulation (Steinberger, 1971; Stevens and Steinberger, 1983). The distribution of endogenous testosterone rather than that of androgen receptor concentration is similar with the total RNA synthesis throughout the wave of the seminiferous epithelium (Parvinen and Soderstrom, 1976b). The mechanism that regulates the meiotic RNA transcription and its rapid onset in mid-pachytene spermatocytes remains to be investigated; a direct androgen action on pachytene spermatocytes seems unlikely. C. CYCLICALLY SECRETED PROTEINS

The cycle of the seminiferous epithelium influences the synthesis and secretion of proteins by the seminiferous tubules. Two distinct peaks were found in

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total amount of secreted radioactive proteins in stages VI and XI1 (Fig. 7A) with remarkably different electrophoretic patterns (Fig. 7B and C). At least I5 cyclically secreted proteins were found by two-dimensional polyacrylamide gel electrophoresis after [3sS]methionine labeling (Wright et al., 1983). One of

Stage-related incorporation of radioactivity derived from [3sS]methionine(cpm x 10 - s, FIG. 7. into total proteins secreted by rat seminiferous tubules showed peaks in stages VI and XI1 of the cycle. ( A . Wright ~t a/.. 1983; Parvinen. 1982. reproduced by copyright permision from The Endocrine Society.) The patterns of secreted proteins in stages VI and XI1 were almost completely different when analyzed by two-dimensional polyacrylamide gel electrophoresis. ( B and C , Wright vr ul.. 19x3, reproduced by permission from The Society for the Study of Reproduction.) This is considered an indication of changes induced by sperrnatogenic cells on the gene expression of Sertoli cells that are the obvious wurces of most of the secrctcd proteins.

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them, designated cyclic protein-2 (CP-2), exhibited a pronounced cycle of secretion. Its peak at stage VI was 30 times greater than the nadir at stages XII-XIV. This glycoprotein or group of glycoproteins exhibits a substantial heterogeneity in its carbohydrate content, resulting in MW 32,000-38,000 and pZ 4.9-6.3. During postnatal development, the secretion of CP-2 starts at day 16, is increased 3-fold at day 36, and an additional 3-fold at day 45 with concomitant changes in the heterogeneity of the glycosylation (Wright and Luzarraga, 1984). Cyclic protein-2 is most likely a product of the Sertoli cells (Wright et af., 1983). Its function is not known, but an involvement in Sertoli cell-mid-pachytene spermatocyte interaction is possible. The developing spermatocytes reach midpachytene around an age of 16 days (Clermont and Perey, 1957), and RNA transcription of these cells is activated at stage VI of the adult seminiferous epithelial cycle (Soderstrom and Parvinen, 1976a). In contrast to secreted products, the synthesis of most cellular proteins by tubular segments is relatively constant throughout the cycle of the seminiferous epithelium (Wright et af., 1983), although some polypeptides can be regarded as markers for stages IV, VIII, and X-XIV (De Philip er af., 1982). An indication of the cyclic function of the Sertoli cells depending on the stage of the cycle of the seminiferous epithelium is the varying secretion of a specific Sertoli cell product, ABP. It was highest in stages VII-XII, as determined by steady-state polyacrylamide gel electrophoresis. A more sensitive radioimmunoassay revealed maximal ABP secretion at stages VIII-XI and minimal ABP secretion at stages IV-V of the cycle (RitzCn el al., 1982; Fig. 6). Although the exact function of ABP is not known, it seems to be secreted by the seminiferous tubules in stages in which the proposed androgen action is maximal. It would not be surprising if the maximal endogenous testosterone concentration of the seminiferous tubules is at least partly maintained by high local concentration of ABP. Immunohistochemical localization of ABP (Pelliniemi el al., 1981; Attramadal et af.,1981), however, does not support the findings on secreted ABP, but is in agreement with measurements of endogenous ABP concentration that is low and roughly equal in all stages of the cycle (RitzCn et al., 1982). D. OTHERCYCLICALLY SECRETED FACTORS It seems unlikely that sex hormones directly influence the control of meiosis (Haffen, 1977). Rather, a meiosis-inducing substance (MIS) that is not species or sex specific has been shown to induce meiosis in fetal mouse testes, where meiosis-preventing substance (MPS) normally prevents spermatogenic cells from entering meiosis prematurely (Byskov and SaxCn, 1976; 0 and Baker, 1976; Byskov, 1978). Also in the adult testis, MIS and MPS have been shown to interact in the regulation of meiosis (Grinsted et af., 1979). Meiosis in the seminiferous epithelium starts at a defined stage of the cycle, i.e., preleptotene

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FIG. 8. Semiquantitative estimations of the secretions of meiosis-inducing substance (MIS, solid circles) and meiosis-preventing substance (MPS, open circles) by rat seminiferous tubule segments at different stages of the cycle. (From Parvinen, 1982, reproduced by copyright permission from The Endocrine Society.) High MIS secretion is concentrated in stages VII and VIII, where preleptotene spermatocytes replicate their DNA at the onset of meiosis. Some secretion of MPS is observed in all stages of the cycle that may have a function in preventing the spermatogonia from entering meiosis prematurely.

spermatocytes at stages VII and VIII. Indications of high secretion rates of MIS were found more often in these stages than in others (Fig. 8). The secretion of MPS was not cyclic, and it may prevent the stem cells from entering meiosis at stages other than VII and VIII, where a pulse of MIS triggers some of these cells to start meiosis (Parvinen et al., 1982). Stages VII and VIIl of the cycle have been shown to secrete a macromolecule that has an inhibitory action on FSH-stimulated aromatase activity of cultured immature Sertoli cells by a mechanism that does not interfere with FSH-receptor interaction or inhibit adenylate cyclase (Boitani et al., 1981). This finding is in accordance with the concept of the preferential dependency of stages VII and VlIl on androgen stimulation. One of the major proteins secreted by rat Sertoli cells in culture is a transferrinlike protein (Skinner and Griswold, 1980). It promotes the growth of established testicular cell lines in serum-free medium (Mather, 1980). Sertoli cell-secreted transferrin probably mediates the transportation of iron from serum to the developing sperm. Immunoreactive transferrin has been localized in the developing acrosomes and over the nuclei of steps 1-17 spermatids in the rat, but the transferrin receptor protein was located in spermatocytes, early round spermatids, and dividing spermatogenic cells (Sylvester and Griswold, 1984). Secretion of transferrin by isolated seminiferous tubule segments was maximal in stages 1X-XI1 of the cycle (Mather et al., 1983), showing a correlation with total [ 3H]thymidine incorporation in the seminiferous tubule segments (Lahdetie et al., 1983).

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E. TESTICULAR GROWTHFACTORS Seminiferous epithelium of the mouse testis contains a potent mitogen that induces DNA synthesis and mitoses in 3T3 cells (Feig et al., 1980). The growth factor is particularly enriched in Sertoli cells, but some activity is also found in isolated primitive type A spermatogonia and in preleptotene spermatocytes. This growth factor with a molecular weight of 15,700 and a pt between 4.8 and 5.8 has been found in the seminiferous epithelium of several mammalian species (Feig et al., 1983). A polypeptide with a molecular weight of 45,000 that stimulates DNA synthesis in cultured fibroblasts has been found in ram rete testis fluid (Brown et al., 1982). These growth factors may have separate roles, the former primarily acting on spermatogenic cells in the seminiferous epithelium, whereas the latter may have an effect on epididymal epithelium. Seminiferous tubules and Sertoli cell-enriched monolayers secrete a compound that is immunologically similar to somatomedin. Maximal secretion of this compound was found in stage VIII of the cycle (Johnsonbaugh et al., 1982). Somatomedin C immunoreactivity has been demonstrated in cultured Sertoli cells and in pachytene spermatocytes cocultured with Sertoli cells, but not in spermatogonia or leptotene-zygotene spermatocytes (Tres et a/., 1986). Since both Sertoli cells and pachytene spermatocytes also displayed binding sites for exogenously added somatomedin C , this growth factor possibly has a role in the meiotic process (Tres et al., 1986).

VI. Localization and Function of Plasminogen Activators in the Seminiferous Epithelium Plasminogen activators (PAS) are highly specific serine proteinases that are involved in tissue remodeling and cell migration processes under normal and pathological conditions (for review, see Dana et al., 1985). There are two known types of PAS, called urokinase (u-PA) and tissue-type PA (t-PA), with approximate molecular weights of 50,000 and 70,000, respectively (Dan@and Reich, 1978). PAS proteolytically activate plasminogen into plasmin that degrades most proteins. Isolated immature rat Sertoli cells in culture secrete PA under stimulation of FSH and dibutyryl CAMP(Lacroix et al., 1977; Lacroix and Fritz, 1982). During postnatal development, the testicular PA level increases concomitantly with the appearance of the most mature spermatogenic cells (Lacroix et al., 1982). Several functions have been proposed for PA in the seminiferous epithelium, such as the opening up of the Sertoli cell junctions when spermatocytes migrate into the adluminal compartment, spermiation analogously with ovulation (Strickland and Beers, 1975), and phagocytosis of the residual bodies (Lacroix et al., 1982).

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A. CELLULAR AND HORMONAL REGULATION OF PA SECRETION

In the adult rat seminiferous epithelium, PA is secreted in stages V11 and VlII more than 100-fold compared with other stages of the cycle (Lacroix er al., 1981). These stages contain tissue-restructuring processes such as the release of the preleptotene spermatocytes from the basement membrane at the onset of meiosis (Russell, 1977a) and spermiation (Leblond and Clermont, 1952). To find out which of these processes is dependent on PA, two approaches were used. PA secretion was measured in 1-mm segments isolated sequentially through stages VI-IX of the cycle and in the same stages at defined time intervals after local irradiation of the testes when desired spermatogenic cell generations were low in number (Dym and Clermont, 1970). Maximal secretion of PA occurred in substages VII, and VII, of the cycle (Fig. 9A), and it was almost abolished 8 days after irradiation when preleptotene spermatocytes were selectively reduced in number (Vihko et al., 1984). A normal PA secretion pattern resumed concomitantly with the reappearance of preleptotene spermatocytes 37 days after irradiation. These observations suggest that preleptotene spermatocytes upon their release from the basal lamina at the onset of meiosis are important regulators of PA secretion by the seminiferous tubules. The temporal sequence of the release is in agreement with morphological analyses after fixation with hypertonic solutions (Russell, 1977a). Preleptotene spermatocytes became detached from the basement membrane at stages VI, VII, and VIII of the cycle while spermatogonia remained flat against that structure. Simultaneously, Sertoli cell processes penetrate the space between basal lamina and preleptotene spermatocytes (Fig. 9D and E). These processes meet and form a tight junctional complex at stage IX of the cycle. During stages IX-XI the leptotene spermatocytes are located in the intermediate compartment of the seminiferous epithelium, i.e., tight junctions of the Sertoli cells are located both above and beneath these cells. At stage XI1 of the cycle, the tight junctions above are released and zygotene spermatocytes become located in the adluminal compartment of the seminiferous epithelium (Russell, 1977a). Since PA secretion is low at this stage, the opening up of the Sertoli cell junctions most obviously is not dependent on PA secretion. Normal amounts of PA were secreted on day 51 after irradiation when spermiating spermatids and the residual bodies were low in number. Therefore, these phenomena are obviously not dependent on proteolysis triggered by secreted PA.

B. PAS DURING POSTNATAL AND in Vitro DIFFERENTIATION The role of preleptotene spermatocytes in the regulation of endogenous and secreted PA levels in the seminiferous epithelium was further studied in immature postnatal rats, where the temporal sequence in the appearance of different

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65

FIG.9. Localization and function of plasminogen activators in the seminiferous epithelium. Endogenous PA shows some cyclic distribution (dotted line in A), but PA secretion is clearly limited in stages VII and VIII (solid line in A, Lacroix etal., 1981; Parvinen, 1982, reproduced by copyright

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spermatogenic cell types is known (Clermont and Perey, 1957). The testicular uPA level increased at 9 days of age, concomitantly with the appearance of the preleptotene spermatocytes (Vihko et al., 1986b). Other increases were observed at the time of the first meiotic reduction divisions and during the final maturation of the testis. The cyclic secretion of PA by seminiferous tubule segments started at an age of 28 days, and from 40 days onward it was found associated with the stages in which preleptotene spermatocytes were found. Since in vivo, the onset of PA secretion starts at stage VII and ceases at stage IX of the cycle, it became of interest to study how these phenomena are regulated in vitro (Toppari et al., 1986b). Stage VI tubules were insensitive to testosterone and FSH, but a combination of these hormones together with insulin and retinoic acid (4F) stimulated PA secretion. Stage VIII tubules were insensitive to testosterone, but FSH and 4F had a pronounced effect. After 3 days in culture, stage VI tubules become sensitive to FSH stimulation whereas this ability diminished in stage VIII suggesting a biochemical differentiation of these stages in vivo. Retinoic acid in the 4F combination proved to be the component that stimulates PA secretion at stages I-VII,-, of the cycle. The action of FSH is clearly distinct; it specifically stimulated the PA secretion of stages VII,-,-XI of the cycle (Vihko et al., 1986a). These observations suggest that the spermatogenic cells modulate the actions of these factors in a stage-dependent fashion. The molecular forms of PAS were analyzed using polyacrylamide gel electrophoresis and zymography (Saksela, 1984). Most of the secreted PA was found to be of urokinase type, but a small amount of t-PA was occasionally found in the culture medium when stage VIII tubules were stimulated by FSH or 4F. The endogenous PA of the seminiferous tubules always consisted of both types of PA. U-PA was the predominant form of PA particularly in stages VII and VIII of the cycle (Fig. 9F) and in the seminiferous tubules during postnatal development. The t-PA fraction increased in activity at stage VIII and, during stages IX-XI1 of the cycle, u-PA and t-PA fractions were virtually equal in activity (Fig. 9F). The activity of the t-PA stayed high during stages XII-XIV of the cycle, but was permission from The Endocrine Society). In a detailed analysis, maximal secretion of PA was found in substages VII, and VIId (B, Vihko er al.. 1984, reproduced by permission from The Society for the Study of Reproduction), coincident with the release of the preleptotene spennatocytes from the basal lamina (L. D. Russell, 1977a, reproduced by copyright permission from Alan R. Liss, Inc.), In stage IX, PA secretion decreased to a low level when leptotene spermatocytes became completely separated from the basal lamina. In a selective absence of preleptotene spermatocytes 8 days after local Xirradiation, PA secretion in stages VII and VIII almost totally ceased (C). These observations support the concept that plasminogen activator has a role in the release of preleptotene spermatocytes from the basement membrane at the onset of meiosis. Zymographic analysis revealed that the endogenous PA was mainly of urokinase type (MW 43,000) in stages VII and VIII (F, lane c), and of tissue-type (MW 65,000) in stages IX-XI1 (F, lane b). Equal amounts of both types were found in other stages (lanes a and d). The secreted PA was always of urokinase type, independent of the age of the animals (e, 28 days; f, 40 days; g, 48 days; and h, 80 days).

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again low during stages 11-VII of the cycle. Immunohistochemical analysis with specific antibodies against both types of PA showed an u-PA reaction in Sertoli cells at stages VII and VIII whereas t-PA was located in pachytene and diakinetic primary spermatocytes at stages VII-XI11 of the cycle (Vihko et al., 1985b). The observations suggest that the two molecular forms of PA have different functions during the cycle of the seminiferous epithelium. The role of Sertoli cell-secreted u-PA in the release of the preleptotene spermatocytes from the basal lamina at the onset of meiosis is suggested by several observations, but the function of t-PA in spermatogenic cells remains to be established.

VII. Spermatogenesis in Vitro Virtually all DNA found in the spermatozoa is synthesized at the onset of meiosis by preleptotene spermatocytes. When traced by [3H]thymidineand autoradiography, these cells developed to late pachytene in organ culture from testes of mature rats (Steinberger and Steinberger, 1965), mice, guinea pigs, rabbits and monkeys (Steinberger and Steinberger, 1967), and men (Steinberger, 1967). In coculture with Sertoli cells in chemically defined medium, meiotic prophase spermatocytes have been shown to differentiate to secondary spermatocytes (Tres and Kierszenbaum, 1983), but thus far no convincing reports have been published about the formation of spermatids in v i m from spermatocytes in mammals. A. CULTURES OF DEFINED SEGMENTS OF SEMINIFEROUS TUBULES Seminiferous tubule segments from stages XI1 and XI11 of the cycle were cultured in order to see if the late pachytene and diakinetic primary spermatocytes are able to complete meiosis in v i m . After 2 days in chemically defined medium without added hormones or other factors, numerous meiotic division figures were seen, and after 6 days, several young spermatids with capshaped acrosomic systems had developed from late pachytene primary spermatocytes (Parvinen et al., 1983). Surprisingly, additions of insulin, ceruloplasmin, transferrin, epidermal growth factor, testosterone, hydrocortisone, FSH, and LH did not influence the survival of the tubular segments significantly, as evaluated by semiquantitative visual estimation. In turn, the physiological contacts between Sertoli cells and spermatogenic cells were an essential prerequisite for differentiation. Pachytene spermatocytes are able to develop through meiotic divisions around stage VII of the cycle, and if started from stage XII, they are able to differentiate up to step 6 spermatids (Fig. 10A) and to express some antigens characteristic of early spermiogenesis (Toppari et al., 1985a; Toppari and Parvinen, 1985).

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FIG. 10. (A) Electron micrograph of a step 6 spermatid that has differentiated from diakinetic primary spermatocytc (stage XIII) during 7 days in v i m in chemically defined medium without added hormones or growth factors. No obvious alterations of normal morphology is seen; the acrosomic system (between the arrowheads) covers an area between one-third and one-quarter of the nuclear circumference, and the chromatoid body (cb) is located at the caudal pole of the nucleus. ; I p m , (B) Light micrograph of a stage VIIl tubulus cultured for 3 days in Magnification, ~ 5 0 0 0bar, chemically defined medium without added hormones or growth factors. Instead of differentiating beyond stage XI (compare XI in Fig. 2), the round spermatids did not elongate and degenerated (asterisk). The maturing spermatids were retained in the seminiferous epithelium, instead of spermiation (arrow). Magnification. X800; bar, 10 F m .

B . FLOWCYTOMETRIC EVALUATION OF THE CULTURED SEMINIFEROUS TUBULE SEGMENTS Flow cytometry is a useful method in evaluation of the proportions of testicular cells containing IC (haploid), 2C, and 4C amounts of DNA (Clausen et a / . , 1977; Meistrich et a / . , 1978b; Pfitzer et al., 1982). Analysis of pooled samples from seminiferous tubule segments revealed a stage-dependent variation with two haploid peaks in some stages of the cycle (Clausen et al., 1982). With a new propidium iodide fluorochrome technique, analyses of single 1.5-rnm segments of rat seminiferous tubules with identified stages of the cycle became possible (Toppari et a / . , 1985b, 1986a). All stages have three distinct peaks at locations corresponding to I , 2, and 4C amounts of DNA. A fourth peak with hypohaploid fluorescence intensity appears during stage I . The fluorescence intensity of this cell population further decreases during stages 1I-11I and reaches a constant value of 0.3C during stages IV-VII. It further decreases to 0.24C at stage Vlll and to as low as 0.07C in the cauda epididymidis. The reduction of fluorescence intensity in haploid cells is concomitant with the second nucleoprotein transition

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that starts at step 15 of spermiogenesis (Grimes et al., 1977; Meistrich et al., 1978a) and a reduction of phosphotungstic acid stainability (Courtens and Loir, 1981). The chromatin of the spermatozoon further condenses during epididymal transit, where an increase of the disulfide bonds has been demonstrated (Calvin and Bedford, 1971). The most profound alterations in the flow cytometric diagrams occur in stages VIII (spermiation) and XIV (meiotic divisions) of the cycle. The relative proportions of the nuclei in different DNA classes correlate well with morphometric estimations at different stages of the cycle of the seminiferous epithelium (Wing and Christensen, 1982). All types of spermatogenic cells do not have an equal ability to differentiate in vitro. The step 8 spermatid nuclei start to elongate, but, during the first day in culture, severe malformations are observed in their ultrastructure followed by rapid degeneration. Another feature of cultured stage VIII tubules is that spermiation does not occur although pachytene and preleptotene spermatocytes in the same cell association differentiate normally (Fig. 10B; Toppari er al., 1985b). Failures of chromatin condensation in step 15 seen in flow cytometry and of the release of the bundle arrangement of the step 18 spermatids at stage VI further suggest an unsuccessful late spermatid differentiation in vitro. The findings resemble the ones seen in the seminiferous epithelium after hypophysectomy . The spermatids did not differentiate beyond the elongation step (Clermont and Morgentaler, 1955). Although flow cytometry is a precise method for quantitation of the seminiferous epithelium, almost no effects of hormones on relative cell numbers could be demonstrated. To support the whole spermatogenesis and especially spermiogenesis in virro, further development of the culture conditions is needed.

VIII. Interaction between Seminiferous Tubules and the Leydig Cells The function of seminiferous epithelium is dependent on testosterone, a product of Leydig cells in the testicular interstitial tissue. Leydig cell function is regulated by pituitary LH, but an increasing number of observations suggest that these cells are also influenced by neighboring seminiferous tubules. If spermatogenesis is suppressed by antispermatogenic chemicals applied locally to the testis, Leydig cells adjacent to atrophic tubules showed morphological evidence of stimulation, whereas in the same testis unaffected seminiferous tubules were flanked by normal Leydig cells (Aoki and Fawcett, 1978). The local effect of seminiferous tubules on Leydig cells has been suggested to be mediated by Sertoli cells: their function impaired after treatment with hydroxyurea, vitamin A deficiency, fetal irradiation, cryptorchidism, or efferent duct ligation leads to a subsequent alterations in the Leydig cell function (de Kretser, 1982). Morphometric observations suggest that the local interaction between Leydig

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cells and seminiferous tubules is dependent on the stage of the cycle of the seminiferous epithelium. Leydig cells were significantly larger in triangles limited by seminiferous tubules at stages VII and VIII of the cycle (Bergh, 1982), and peritubular Leydig cells showed a similar dependency on the stage of the seminiferous epithelial cycle (Bergh, 1983). These observations are in agreement with the maximal testosterone concentration and ABP secretion rate in the same stages. When seminiferous tubules from defined stages of the cycle were coincubated with isolated Leydig cells, an influence on both basal and hCG-stimulated testosterone production was observed, and the effect was independent of FSH or GnRH action. All stages of the cycle had a stimulatory effect on Percoll-purified Leydig cells, but the effect was inhibitory on crude Leydig cell preparations (Parvinen et al., 1984).The stimulatory effect by stages VII and VIII of the cycle was significantly higher (p < 0.05) than in stages 11-VI. This is considered as evidence of a stage-dependent interaction between seminiferous tubules and Leydig cells, in agreement with measurements of their size distribution (Bergh, 1983; Fig. 11). The mechanism of the paracrine interaction between Leydig cells and semi-

I

IV-V Vlla-b

Vlll

XI1

FIG. 11. Comparison of the profile area of peritubular Leydig cells (open circles, Bergh, 1983) and of the relative augmentation of hCG-stimulated testosterone production of isolated Leydig cells by seminiferous tubules (solid circles, Parvinen er a / . . 1984) at different stages of the cycle of the seminiferous epithelium. Peritubular Leydig cells are significantly (p < 0.025) larger than the perivascular ones in the proximity of seminiferous tubules at stages 111-VIII of the cycle, and the stimulation of testosterone production by stage VII and Vlll tubules was significantly 0, < 0.05) greater than by stage 11-VI ones. Both observations suggest that a paracrine stimulation of Leydig cells by seminiferous tubules is particularly active in stages VII and Vlll of the cycle.

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niferous tubules is not known, but endogenous peptides, “GnRH-like substances” (Sharpe er al., 1981), and a high-molecular-weight factor (Grotjan and Heindel, 1982) from seminiferous tubules have been suggested. It has been shown that a GnRH agonist, LHRH-A, has a dose-dependent effect on testicular capillary permeability and on the testosterone concentration in the testicular interstitial fluid (Sharpe, 1984). Therefore, an increasing amount of evidence suggests that GnRH-like substances may play a role in the paracrine control of the testicular function. The activity of a proopiomelanocortin-like gene in the Leydig cells suggests that this compound also may have a function in the local regulation of the testicular function (Chen er al., 1984), perhaps specifically in some stages of the epithelial cycle (Gizang-Ginsberg and Wolgemuth, 1985). For further references in this rapidly growing area, see the reviews of Sharpe (1983, 1984).

IX. Conclusions and Future Prospects It has been well established for some time that the normal sperm-producing function of the testis is dependent on stimulation by pituitary gonadotropins, LH and FSH, and that targets for these hormones are Leydig cells in the interstitial tissue and Sertoli cells in the seminiferous epithelium, respectively. The effect of LH on the seminiferous epithelium is relayed by testosterone produced by the Leydig cells. Therefore, the two main hormones directly influencing the function of the seminiferous epithelium are FSH and testosterone. Sertoli cells in the seminiferous epithelium are the primary targets of FSH and testosterone, and most investigators agree that they are the only targets of these hormones. However, there are a few observations suggesting a direct androgen action on some spermatogenic cell types. Both FSH and androgens seem to have a cyclic action depending on the stage of the cycle of the seminiferous epithelium. The preferential action of FSH is associated with stages containing meiotic reduction divisions, early spermiogenesis, and a very close association of maturation phase spermatids with Sertoli cells ( X U - V ) . The parameters related to androgen action predominate at stages in which the final maturation of the spermatids, spermiation, onset of meiosis, onset and highest activity of meiotic RNA transcription, and the onset of nucleoprotein transitions in the spermatids occur (VII-XI). The high concentration of androgen receptors suggests that the region of the meiotic reduction divisions (XIV) might be influenced by both FSH and androgens. The stage-dependent variation of the hormone responses in the seminiferous epithelium strongly suggests an existence of local paracrine regulation and cell interaction mechanisms in the seminiferous epithelium, that are dependent on spermatogenic cells associated with the Sertoli cells at each stage of the cycle of

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the seminiferous epithelium. The nature of this interaction is obscure, but some advances have been made. The secretion of a proteolytic enzyme, urokinase-type plasminogen activator, seems to be dependent on both cellular and hormonal regulation in the seminiferous epithelium. Testicular GnRH-like factors and proopiomelanocortin-derived peptides may play a role in seminiferous tubuleLeydig cell interaction. In the seminiferous epithelium, a Sertoli cell-derived growth factor has been suggested to have a role in local regulation together with other factors, such as meiosis-inducing and -preventing substances, a somatomedin-like compound, and the spermatogonial chalone. The seminiferous epithelium has long been a difficult area for biochemical studies, owing to its highly complex structure. However, it has proved to be an excellent model system for studies of cell interaction in general. A highly organized sequence of cell differentiation events is in intimate association with cyclically functioning Sertoli cells and available for biochemical and cytochemical studies.

ACKNOWLEDGMENTS This project has been supported by grants from The Academy of Finland, The Population Council and from the University of Turku.

Aoki, A , , and Fawcett, D. W. (1978). Eiol. Reprod. 19, 144-158. Assaf, A. A. (1980). M.Sc. thesis, McGill University, Montreal. Attramadal, A.. Bardin, C. W . , Gunsalus, G . L., Musto, N. A., and Hansson, V. (1981). Eiol. Reprod. 25, 983-988. Bardin, C. W., Musto, N., Gunsalus, G., Kotite, N., Cheng, S.-L., Larrea, F., and Becker, R. (1981). Annu. Rev. Phvsiol. 43, 189-198. Benda, C. (1891). Arch. Anat. Phvsiol. 549-552. Bergh, A. (1982). In/. J. Androl. 5, 325-330. Bergh. A. (1983). Int. J. Androl. 6 , 57-65. Boitani, C., RirzCn, E. M., and Parvinen, M. (1981). Mu/.Cell. Endocrinol. 23, 11-22, Bouchard, P., Gagnon, C., Phillips. D. M . , and Bardin, C. W. (1980). J . Cell B i d . 86, 417-423. Braun, T., and Dods, R. F. (1975). Proc. Natl. Acud. Sci. U.S.A. 72, 1097-1 100. Brown, K. D., Blakeley, D. M., Henville, A , , and Setchell, B. P. (1982). Eiochem. Biopphys. Res. Curnmun. 105, 391-397. Byskov, A. G . (1978). Int. J. Androl. Suppl. 2, 29-38. Byskov, A. G . , and Saxen. L. (1976). Dev. B i d . 52, 193-200. Calvin, H. I . , and Bcdford. J. M. (1971). J. Reprod. Fertil. Suppl. 13, 65-75, Chan, L.. and O'Malley, B. W. (1976). N. EngI. J. Med. 294, 1322-1328, 1372-1381, 14301437.

148

MAR’ITI PARVINEN ET AL.

Chen, C.-L. C., Mather, J. P., Moms, P. C., and Bardin, C. W. (1984). Proc. Narl. Acad. Sci. U.S.A. 81, 5672-5675. Clausen, 0. P. F., Purvis, K., and Hansson, V. (1977). Biol. Reprod. 17, 555-560. Clausen, 0. P. F., Parvinen, M., and Kirkhus, B. (1982). Cyromerry 2, 421-425. Clermont, Y. (1972). Physiol. Rev. 52, 198-236. Clermont, Y., and Bustos-Obregon, E. (1975). Am. J . Anar. 123, 237-248. Clennont, Y., and Harvey, S. C. (1965). Endocrinology 76, 80-89. Clermont, Y., and Hermo, L. (1975). Am. J . Anar. 142, 159-176. Clermont, Y.,and Mauger, A. (1974). Cell Tissue Kinet. 7, 171-178. Clermont, Y., and Morgentaler, H. (1955). Endocrinology 57, 369-382. Clermont, Y., and Perey, 9. (1957). Am. J . Anur. 100, 241-268. Clermont, Y., Leblond, C. P., and Messier, B. (1959). Arch. Anar. Microsc. Morphol. Exp. Suppl.

48, 37-46. Comings, D. E., and Okada, T. A. (1972). J. Ulrrusrrucr. Res. 39, 15-23. Courot, M., Hochereau-de Reviers, M.-T., and Ortavant, R. (1970). In “The Testis” (A. D. Johnson, W. R. Gomes, and N. L. Vandemark, eds.), Vol. I, pp. 339-432. Academic Press, New York. Courtens, J.-L., and Loir, M. (1981). J . Ulrrasrrucr. Res. 74, 327-340. Cusan, L., Gordeladze, J. 0.. Parvinen, M., Clausen, 0. P. F., and Hansson, V. (1981). Biol. Reprod. 25, 915-919. Dan#, K., and Reich, E. (1978). J . Exp. Med. 147, 745-757. Dan#, K., Andreasen, P. A., Gr@ndahl-Hansen,J., Kristensen, P., Nielsen, L. S.,and Skriver, L. (1985). Adv. Cancer Res. 44, 139-266. Daoust, R., and Clermont, Y . (1955). Am. J . Anat. 96, 255-283. Das, N. K., Siegel, E. P., and Alfert, M. (1965). J. Cell Biol. 25, 387-395. De Kretser. D. M. (1982). Inr. J . Androl. Suppl. 5 , 11-17. De Philip, R. M., and Kierszenbaum, A. L. (1982). Proc. Nut/. Acad. Sci. U.S.A. 79,6551-6555. De Philip, R. M., Tres. L. L., and Kierszenbaum, A. L. (1982). Exp. Cell Res. 142, 489-494. de Turner, E. A., Aparicio, N. J., Turner, D., and Schwarzstein, L. (1978). Fertil. Sreril. 29, 328331.

Distel, R. J., Kleene, K. C., and Hecht, N. B. (1984). Science 224, 68-70. Dym, M., and Clermont, Y. (1970). Am. J . Anur. 128, 265-282. Dym, M., and Fawcett, D. W. (1970). Biol. Reprod. 3, 308-326. Dym, M., Raj, H. G. M., and Chemes, H. E. (1977). In “The Testis in Normal and Infertile Men” (P. Troen and H. R. Nankin, eds.), pp. 97-101. Fawcett, D. W. (1972). In “The Genetics of Spermatozoon” (R. A. Beatty and S. GluecksohnWaelsch, eds.), pp. 37-67. Edinburgh. Fawcett, D. W., Eddy, E. M., and Phillips, D. M. (1970). Biol. Reprod. 2, 129-153. Feig, L. A., BellvC. A. R., Horbach-Erickson, N., and Klagsbrun, M. (1980). Proc. Narl. Acad. Sci. U.S.A. 77, 4774-4778. Feig, L. A., Klagsbrun, M., and Bellvk, A. R. (1983). J. Cell Biol. 97, 1435- 1443. Frankel, A. I., and Chapman, J. C. (1984). J . Sreroid Biochem. 20, 1301-1311. Fritz, 1. B. (1978). In “Biochemical Actions of Hormones” ( G . Litwack, ed.), Vol. V, pp. 249281. Academic Press, New York. Gagnon, C., Axelrod, J., Musto, N., Dym, M., and Bardin, C. W. (1979). Endocrinology 105, 1440-144s.

Galdieri, M., and Monesi, V. (1974). Exp. Cell Res. 85, 287-295. Geremia, R., D’Agostino, A., and Monesi, V. (1978). Exp. Cell Res. 111, 23-30. Gizang-Ginsberg, E., and Wolgemuth, D. J. (1985). Dev. Biol. 111, 293-305. Gordeladze, J . O., and Hansson, V. (1981). Mol. Cell. Endocrinol. 23, 125-136.

REGULATION OF THE SEMINIFEROUS EPITHELIUM

149

Gordeladze, J. O., Parvinen, M., Clausen, 0. P. F., and Hansson, V. (1982). Arch. Androl. 8,4351.

Grimes, S . R., Jr., Meistrich. M. L., Platz, R. D., and Hnilica, L. S . (1977). Exp. CellRes. 110, 31-39. Grinsted, J., Byskov, A. G.,and Andreasen, M. P. (1979). J. Reprod. Fertil. 56, 653-656. Grotjan, H. E., Jr., and Heindel, J. 1. (1982). Ann. N.Y. Acud. Sci. 383, 465-457. Haffen, K. (1977). In "The Ovary" (S. Zuckerman and B. J. Weir, eds.), Vol. I, pp. 69-112. Academic Press, New York. Henderson, S. A. (1964). Chromosomu 15, 345-366. Henderson, S. A. (1972). Proc. Nutl. Acud. Sci. U.S.A. 69, 3394-3398. Hilscher, B., Passia, D., and Hilscher, W. (1979). Andrologiu 11, 169-181. Hilscher, W. (1967). Arch. Anut. Microsc. Morphol. Exp. 56 (Suppl.), 75-84. Hotta, Y.. Chandley, A. C., and Stem, H. (1977). Nature (London) 269, 240-242. Isomaa, V., Parvinen, M., Jinne, 0. A., and Bardin, C. W. (1985). Endocrinology 116, 132-137. Johnsonbaugh, R . E., Ritzkn, E. M., Hall, K., Parvinen, M., and Wright, W. W. (1982). Proc. Eur. Testis Workshop. 2nd, Abstr. D 12. Kerr, J. B., and de Kretser, D. M. (1975). J . Reprod. Fertil. 43, 1-8. Kerr, J. B., Mayberry, R. A,, and Irby, D. C. (1984). Cell Tissue Res. 236, 699-709. Kierszenbaum, A. L., and Tres, L. L. (1974a). J . Cell Biol. 60,39-53. Kierszenbaum, A. L., and Tres, L. L. (1974b). J . Cell Biol. 63, 923-925. Kierszenbaum, A. L., and Tres, L. L. (1975). J . Cell Biol. 65, 258-270. Kierszenbaum, A. L., and Tres, L. L. (1978).Fed. Proc., Fed. Am. Soc. Exp. Biol. 37,2512-2516. Kierszenbaum, A. L., Spruill, W. A,, White, M. G., Tres, L. L., and Perkins, J. P. (1985). Proc. Nutl. Acud. Sci. U.S.A. 82, 2049-2053. Lacroix, M., and Fritz, I. B. (1982). Mol. Cell. Endocrinol. 26, 247-258. Lacroix, M., Smith, F. E., and Fritz, I. B. (1977). Mol. Cell. Endocrinol. 9, 227-236. Lacroix, M., Parvinen, M., and Fritz, I. B. (1981). Biol. Reprod. 25, 143-146. Lacroix, M., Smith, F. E., and Fritz, I. B. (1982). Mol. Cell. Endocrinol. 26, 259-267. Lacy, D. (1960). J. R. Microsc. SOC. 79, 209-225. Lahdetie, J . , Kaukopuro, S., and Parvinen, M. (1983). Hereditus 99, 269-278. Lamb, D. J . , Tsai, Y.-H., Steinberger, A., and Sanbom, B. M. (1981). Endocrinology 108, 10201026. Leblond, C. P., and Clermont, Y. (1952). Ann. N.Y. Acad. Sci. 55, 548-573. Loir, M. (1972). Ann. Biol. Anim. Biochim. Biophys. 12, 203-219. Mather, J. P. (1980). Biol. Reprod. 23, 243-252. Mather, J. P., Gunsalus, G. L., Musto, N. A., Cheng, C. Y.,Parvinen, M., Wright, W., PirezInfante, V., Margioris, A,, Liotta, A., Becker, R., Krieger, D. T., and Bardin, C. W. (1983). J . Steroid Biochem. 19, 41-51. Means, A. R., Dedman, J. R., Tindall, D. J., and Welsh, M. J. (1978). Int. J . Androl. Suppl. 2, 403-423. Meistrich, M. L., Brock, W. A., Grimes, S. R.,Jr., Platz, R. D., and Hnilica, L. S. (1978a). Fed. Proc., Fed. Am. Soc. Exp. Biol. 37, 2522-2525. Meistrich. M. L., Lake, S . , Steinmetz, L. L., and Gledhill, B. L. (1978b). Mutut. Res. 49, 383396. Monesi, V. (1962). J . Cell Biol. 14, 1-18. Monesi, V. (1964). J. Cell Biol. 22, 521-532. Monesi, V. (1971). J. Reprod. Fertil. Suppl. 13, 1-14. Mulder, E., Peters, M. J., devries, J., and van der Molen, H. J. (1975). Mol. Cell. Endocrinol. 2, 171-182. Muramatsu, M., Utakoji, T., and Sugano, H. (1968). Exp. Cell Res. 53, 278-283.

150

MARIT1 PARVINEN ET AL.

Nakhla, A. M., Mather, I. P., Janne, 0. A., and Bardin, C. W. (1984). Endocrinology 115, 121128. Niemi, M., and Kormano, M. (1965). Anat. Rec. 151, 159-170. 0 , W. S., and Baker, T. G. (1976). J. Reprod. Fertil. 48, 399-401. Parvinen, L.-M., Jokelainen, P. T., and Parvinen, M. (1978). Hereditas 88, 75-82. Parvinen, M. (1982). Endocr. Rev. 3, 404-417. Parvinen, M., and Jokelainen, P. T. (1974). Biol. Reprod. 11, 85-92. Parvinen, M., and Parvinen, L.-M. (1979). J. Cell. Biol. 80, 621-623. Parvinen, M., and Ruokonen, A. (1982). J. Androl. 3, 211-220. Parvinen, M., and Soderstrom, K.-0. (1976a). Nature (London) 260, 534-535. Parvinen, M., and Soderstrom, K.-0. (1976b). J . Steroid Eiochem. 7, 1021-1023. Parvinen, M., and Soderstrom. K.-0. (1977). I n “Techniques of Human Andrology” (E. S . E. Hafez, ed.), pp. 113-123. Elsevier, Amsterdam. Parvinen, M., and Vanha-Perttula, T. (1972). Anat. Rec. 174, 435-450. Parvinen, M., Marana, R., Robertson, D. M., Hansson, V., and Ritztn, E. M. (1980). I n “Testicular Development, Structure and Function” (A. Steinberger and E. Steinberger, eds.), pp. 425432. Raven, New York. Parvinen, M., Byskov, A. G., Yding Andersen, C., and Grinsted, J. (1982). Ann. N.Y. Acad. Sci. 383, 483-484. Parvinen, M., Wright, W. W., Phillips, D. M., Mather, J. P . , Musto, N. A , , and Bardin, C. W. (1983). Endocrinology 112, 1150-1152. Parvinen, M., Nikula, H., and Huhtanierni, I. (1984). Mol. Cell. Endocrinol. 37, 331-336. Parvinen, M., Vihko, K. K., and Toppari, J. (1985). In “Recent Progress in Cellular Endocrinology of the Testis’’ (J. M. Saez, M. G. Forest, A. Dazord, and J. Bertrand, eds.), pp. 387-406. INSERM, Paris. Pelliniemi, L. J., Dym, M.. Gunsalus, G. L., Musto, N. A., and Bardin, C. W. (1981). Endocrinology 108, 925-93 I . Perey, B., Clermont, Y., and Leblond, C . P. (1961). Am. J. Anat. 108, 47-77. Mtzer, P., Gilbert, P., Rolz, G., and Vyska, K. (1982). Cytometry 3, 116-122. Posalaki, Z., Szabo, D., Bacsi, E., and Okros, I. (1968). J. Hisrochem. Cytochem. 16, 249-262. Purvis, K., Parvinen, M., Gautvik, K., and Hansson, V. (1984). In “Regulation of Target Cell Responsiveness” (K. W. McKerns, A. Aakvaag, and V. Hansson, eds.), Vol. I, pp. 21-35. Plenum, New York. Rasmussen, S . W., and Holm, P. B. (1980). Hereditas 93, 187-216. Ritztn, E. M., Hansson, V., and French, F. S. (1981). In “The Testis. Comprehensive Endocrinology” (H. Burger and D. de Kretser, eds.), pp. 171-194. Raven, New York. Ritzkn, E. M., Boitani, C., Parvinen, M.,French, F. S., and Feldman, M. (1982). Mol. Cell. Endocrinol. 25, 25-34. Russell, L. D. (1977a). Am. J. Anat. 148, 313-328. Russell, L. D. (1977b). Biol. Reprod. 17, 184-191. Russell, L. D., and Clermont, Y. (1977). Anat. Rec. 187, 347-365. Russell, L. D., and Frank, B. (1978). Anat. Rec. 190, 79-98. Russell, L. D., Malone, J. P., and Karpas, S. L. (1981). Tissue Cell 13, 369-380. Saksela, 0.. Vaheri, A., Schleuning, W.-D., Mignatti, P., and Barlati, S. (1984). Int. J . Cancer 33, 609-616. Salonen, K., Paranko, J., and Parvinen, M. (1982). Chromosoma 85, 611-618. Sanborn, B. M., Steinberger, A., Tcholakian, R. K., and Steinberger, E. (1977). Steroids 29,493502. Setchell, B. P. (1978). In “The Mammalian Testis,’’ pp. 263-273. Paul Elek, London. Shape, R. M. (1983). Q. J. Exp. Physiol. 68, 265-287.

REGULATION O F THE SEMINIFEROUS EPITHELIUM

15 1

Sharpe, R. M. (1984). Biol. Reprod. 30, 29-49. Sharpe, R. M., Fraser, H. M.. Cooper, I., and Rommerts, F. F. G. (1981). Nature (London) 290, 785-787. Skinner, M. K., and Griswold, M. D. (1980). J. Biol. Chem. 255, 9523-9525. Soderstrom, K.-0. (1977). Cell Tissue Res. 184, 411-421. Soderstrom, K . - 0 . (1981). Exp. Cell Res. 131, 488-491. Soderstrom, K.-0.. and Parvinen, M. (1976a). Mol. Cell. Endocrinol. 5, 181-199. Sderstrom, K.-O., and Parvinen, M. (1976b). J . Cell Biol. 70, 239-246. Soderstrom, K.-0.. and Parvinen, M. (1976~).Cell Tissue Res. 168, 335-342. Sijderstrom, K.-O., and Parvinen, M. (1978). Acta Anar. 100, 557-572. Spruill, W. A., White, M. G., Steiner, A. L., Tres, L. L., and Kierszenbaum, A. L. (1981). Exp. Cell Res. 131, 131-148. Steinberger, A., and Steinberger, E. (1965). J . Reprod. Fertil. 9, 243-248. Steinberger, A., and Steinberger, E. (1967). J. Reprod. Ferril. Suppl. 2, 117-124. Steinberger, A., Hintz, M., and Heindel, J. J. (1978). Biol. Reprod. 19, 566-572. Steinberger, E. (1967). Anat. Rec. 157, 327-328. Steinberger, E. (1971). Physiol. Rev. 51, 1-22. Steinberger, E. (1977). Steroids 29, 493-502. Stevens, R. W., 111, and Steinberger, E. (1983). J. Androl. 4, 57. Strickland, S . , and Beers, W. H. (1976). J . Biol. Chem. 251, 5694-5702. Sud, B. N. (1961a). Q. J . Microsc. Sci. 102, 273-292. Sud, B. N. (1961b). Q. J . Microsc. Sci. 102, 495-505. Susi, F. R., and Clermont, Y. (1970). Am. J. Anar. 129, 177-192. Sylvester, S. R., and Griswold. M. D. (1984). Biol. Reprod. 31, 195-203. Tindall, D. J . , Miller, D. A., and Means, A. R. (1977). Endocrinology 101, 13-23. Toppari, J . , and Parvinen, M. (1985). J. Androl. 6, 334-343. Toppari, J., Brown, W. R. A., and Parvinen, M. (1985a). Ann. N.Y.Acad. Sci. 438, 515-518. Toppari, J., Eerola, E., and Parvinen, M. (1985b). J. Androl. 6 , 325-333. Toppari, J . , Mali, P., and Eerola, E. (1986a). J . Hisrochem. Cyrochem. (in press). Toppari, J., Vihko, K. K., Rasanen, K. G. E., Eerola, E., and Parvinen, M. (1986b). J. Endocrinol. 108 (in press). Tres, L. L., and Kierszenbaum. A. L. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 3377-3381. Tres, L. L., Smith, E. P., van Wyk, J. J., and Kierszenbaum, A. L. (1986).Exp. Cell Res. 162,3350.

Tsai, Y . H., Sanbom, B. M., Steinberger, A,, and Steinberger, E. (1980). J. Steroid Biochem. 13, 711-718. Ulvik, N. M., and Dahl, E. (1981). Cell Tissue Res. 221, 31 1-320. Utakoji, T. (1966). Exp. Cell Res. 42, 585-596. Vihko, K. K., Suominen, J. J. O., and Parvinen, M. (1984). Biol. Reprod. 31, 383-389. Vihko, K. K., Kristensen, P., Toppari, J., Saksela, O., Dan@,K.,and Parvinen, M. (1985). J. Cell Biol. 101, 367a. Vihko, K. K., Toppari, J., and Parvinen, M. (1986a). Submitted for publication. Vihko, K. K., Toppari, J., Saksela, O., Suominen, J. J. O., and Parvinen, M. (1986b). Acra Endocrinol. (in press). Walt, H., and Armbruster, B. L. (1984). Cell Tissue Res. 236, 487-490. Wing, T.-Y., and Christensen, A. K. (1982). Am. J . Anat. 165, 13-25. Wright, W. W., and Frankel, A. I. (1980). Endocrinology 107, 314-318. Wright, W. W., and Luzarraga, M. (1984). Biol. Reprod. 30 (Suppl.), 1, 48. Wright, W. W., Parvinen, M., Musto, N. A., Gunsalus, G. L., Phillips, D. M., Mather, J. P., and Bardin, C. W. (1983). Biol. Reprod. 29, 257-270.

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INTERNATIONAL REVIEW OF CYTOLOGY. VOL. 104

The Cytoskeleton in Protists: Nature, Structure, and Functions JEAN GRAIN UA CNRS 138, Groupe de Zoologie et Protistologie, Universiti Clermont 11, Complexe Scientifique des Cizeawr, 631 70 AubiZre, France

I. Introduction The term cytoskeleton is at present commonly used to mean those cell structures which are involved in either or both of two phenomena: ( 1 ) maintenance of cell shape, i.e., preservation of morphological type, implying a degree of rigidity, or in deformable cells a measure of elasticity, and (2) active cell deformation, or active displacement of various cell organelles (cell locomotion, contractility, cytoplasmic currents, migration of chromosomes, vesicles, organites, displacement of membrane receptors, etc.). Both of these phenomena involve proteins used for the construction of temporary or permanent structures. Some of these proteins, actin for example, are involved in both rigidity (gelation) and contractility. Links between the various structures made up from these proteins, and between these structures and others such as the plasma membrane, the nucleus, etc. can provide the means for active deformation and movement to occur, while at the same time constituting networks and layers or sheaths which maintain cell shape. Thus, the term cytoskeleton has a broad meaning which goes beyond that of “cell skeleton,” which is a static notion, and includes a dynamic aspect. In short, the cytoskeleton comprises the set of protein structures involved in maintenance of cell shape and in movement, along with proteins not organized into visible structures but which are associated with these and control their function in either role. Studying the cytoskeleton is thus a study of structures, of their chemical components and those of associated substances, of the interactions occurring among them, and of the function of each of these various elements. Any attempt at reviewing the cytoskeleton of protists must face the wide diversity of types of organization and behavior encountered. Certain cytoskeletal elements known in higher eukaryotes have been found in certain protists, which suggests that they have been conserved in the course of evolution. On the other hand, knowledge is scant with regard to other elements, which seem specific to protists. Here, we shall first deal with those ubiquitous elements also found 153 Copyright 0 19116 by Academic Press. Inc All rights of reproduction in any form reserved.

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JEAN GRAIN

outside the protists, and then with those specific to them. We shall deliberately omit structures constituting the nucleoskeleton and those involved in nucleus division, and the various alveolar and vesicular systems which by virtue of the hydrostatic pressure inside them may be regarded as constituting a hydroskeleton. The generally accepted structural elements of the cytoskeleton are actin microfilaments, myosin filaments, intermediate or 10-nm filaments, and microtubules. To these can be added elements often found in protists, such as periodic fibers, submembranar epiplasm, and other filaments which differ from the classical ones.

11. Actin

Actin has been shown to be present in some protists, along with or without myosin. The properties of this actin, the structures it may constitute, and its interrelations with other proteins have been thoroughly studied, first in protists which move by ameboid movements and more recently in other protists which sometimes appear lacking in myosin (ciliates, flagellates, sporozoa). OF PROTIST ACTINS A. PROPERTIES

Different types of actin have been found in vertebrate cells: two nonmuscular actins (p and y), two in smooth muscle (a-like and y-like), one (a)in skeletal muscle, and one in cardiac muscle (Vandekerckhove and Weber, 1981). 1. Myxomycetes and Amoeba Various proteins similar to muscle actin have been isolated from Physarum (Nachmias et al., 1970), Dictyostelium (Wooley, 1972),Acunthamoeba (Pollard et a f . , 1970), and Entamoeba histofytica (Gadasi, 1982). As with actin from different metazoan cells, they represent 14 to 20% of the total proteins of the cell. Their molecular weight (MW) is of the order of 42-43K except for Entamoeba (48K) (Gadasi, 1982). They are all able to activate Mg2+-ATPase activity of muscle myosin, but less strongly than muscle actin [3 times less for actin of Acanrhumoeba or of Dicryosteliurn (Table I) (Korn, 1978)l. In their Gform, they bind to DNase I and inhibit its activity, except for actin of Enfamoeba (Gadasi, 1982). Their amino acid composition is only slightly different from that of other actins; they all possess one particular amino acid, N-methylhistidine. The actin of Acanrhamoeba, which alone contains an N-methyllysine, possesses, within an overall amino acid sequence identical to that of others actins, punctual differences in 6% of its residues; the position of certain amino acids enables the characteristics of muscle actin and nonmuscle actin and specific characteristics to

TABLE I COMPARISON OF T H E PROPERTIES OF NONMUSCLEA N D MUSCLEA C r l N S " ~ '

Critical concentration (mg/ml)

2 mM MgClz

0.1 M KCI

Source of actin

25°C

5°C

25°C

Rabbit skeletal muscle Human platelet Rat liver Chick embryo brain Acanthamorba castellanii

0.03 0.03 0.02 0.02 0.06

0.03 0.03 0.02 0.02 0.06

0.03 0.09 0.08 0.07 0.09

5°C 0.1

0.51 0.48 0.5 0.45

0.5 M KCI/ ImM MgClz

0.5 M KCI/ 1 mMCaClz

25°C

25°C

5°C

KaPP(pV)

0.02 0.09

0.15 0.32

0.94 3.2

0.01 0.04

0.22 0.39

7. I 9.6 9.6 9.6 21.7

OKom (1978). the concentration of F-actin required for half-maximal activation of the Mg2 -ATPase of rabbit skeletal muscle heavy meromyosin in 2.5 mM MgCI2/2.0 mM ATPl2.4 mM imidazole chloride, pH 7.0, at 24°C. bKapp is

+

TABLE I1 PARTIAL AMINVACIDSEQUENCE DATAFOR MUSCLEA N D NONMUSCLE ACT INS^,^ Amino acid residues

Sources Rabbit skeletal muscle Human heart Bovine brain Human platelet Acanthamoebu

16

17

Cys

Leu

Val

Val/Ile

Met

Cys

10

Met<‘

106 Thr

Val“

129

176

228

Val

Met

Ala

Val Thr Thr

Leu

Ala

Thrc

Leu<

His”

266 Ile

Leu”

271

278

Ala

Tyr

Ala Cys Cys

Tyr Phe

Alac

286 Ile

294

296

298

Val“

316

357

364

Ala

Asn

Met

Tyr

Ile

Thr

Ala

Ala

Asn

Leu

Tyr Tyr Tyr

Ile Ile Ile

Ser Ser

Ala Ser

Gly“

Vald

Leu‘

Leu”

Serf

Sere

Phe

Tyrc

305

Phed

“From Korn (1978). hThe table shows only those residues where one or more of the four other actins for which partial sequence data are available differs from the totally sequenced rabbit skeletal muscle actin.
157

THE CYTOSKELETON IN PROTISTS TABLE 111 DIFFERENT FORMSOF ACTINWITH REGARD TO THEIR ACIDITY

OH 6

Acanthamoeba

Nonmuscle cells Embryonic muscle Nonmuscle cells Embryonic muscle Embryonic muscle Striated muscle t

l

5.44

p

5.42 5.40

a

Entamoeba Dictyostelium

X X

7

H+

be identified (Table 11) (Korn, 1978; Mackenzie, 1980); the isoelectric point (PI) of this 6-actin of Acanthamoeba is more basic than the pl of other actins, due to histidine in position 224 (Korn, 1978), whereas the actin pl of Entamoeba is more acidic than that of muscle a-actin (Table 111) (Gadasi, 1982), though its peptide map may not be very different from that of muscular actin. As regards muscular a-actin, the actin of Dictyostefium is more basic (Uyemura et al., 1978) and that of Physarum is more acidic. The latter differs from Dictyostefium and Acanthamoeba actins respectively by 4 (Vandekerckhove and Weber, 1980) and 5 amino acids (Vandkerckhove and Weber, 1978); it presents 25 substitutions of amino acids compared with the skeletal muscle actin of mammals, and 17 compared with nonmuscular actin of mammals (Vandekerckhove and Weber, 1978); some of these substitutions may present a functional importance, for example, in the degree of activation of Mg2 -myosin ATPase by actin. Evidently then, in protists, the actins have different compositions as is the case for actins of metazoan (Table IV), where NH,-terminal peptide sequences are variable, though common sequences are extensive (7-9; 11- 15) (Table V) (Vandekerckhove and Weber, 1981). +

TABLE IV OF DIFFERENT AMINO ACIDSBETWEEN VARIOUS ACTINS NUMBER

0

Nonmuscle actins Smooth muscle actins Skeletal muscle actin

{

0 17

y-like a-like

0 3

0

25

a

8

0 a

actins

actins

Skeletal muscle actin

158

JEAN GRAIN

TABLE V OF THE N H 2 - T TRYPTIC ~ ~ ~PEPTIDES ~ ~ ~OF VARIOUS ~ ACTINS" AMINOACIDSEQUENCES Mammalian sarcomeric actins .I I Acetyl&pklu-

BI&p&p&lu-

t

I

Mammalian smooth muscle actins

BI- Glu- Glu- Glu- Asp- Ser- T

t t

h

r

~

~

~

C

y

s

-

~

~

u

-

C

y

BI- Glu- Glu- Clu-Thr- Thr

t

Mammalian cytoplasmic actins

I

BI- Asp- Asp- Asp- Ile- Ala-

Val-]Asp- Asn- Gly-Ser- Gly] Met-Cys. Lys

BI- Glu- Clu- Glu- Ile- Ala

Ile-LAsp-Asn- Gly-Ser-Glyt Met-Cys. Lys

5:

Actin from Ph.vsarum polvcephalum

BI- Glu- Gly- Glu- Asp- Val- G l n - v i Ile-[Asp- Asn- Gly-Ser- Gly] Met-Cys. Lys Actin from Dictvostrlium discoidrum

I

8 Acetyl- Asp- Gly- Glu- Asp- Val- G l n - v j Ile-[Asp- Asn- Gly- Ser- Gly] Met-Cys, Lys Actin from yeast Succhuromvcr.s cerrvisiuc

9

BI(?)- Asp- Ser- Glu-Val- A l a - F l a - ]

He-/Asp-Asn-Gly-Ser- Glyj Met-Cys .Lys

Actin from Drosophilu melunoguster Schneider L-2 cells (predicted) 10

BI( Asp, Glu) Glu- N

f

- N

L

N -]Asp- Asn-Gly-Ser-GlyiMet-Cys Lyr

OFor details, see the original article. From Vandekerckhove and Weber (1981).

In the vegetative form of Dictyosteliurn, 17 genes which are potentially capable of coding for actin have been identified (Firtel et al., 1979); the analysis of the nucleotide sequence of 7 of these genes shows that only 4 of them are able to code for a normal (major) actin; the 3 others are possibly involved in the synthesis of minor actins (differing from the normal sequence by a few amino acids).

s

l

L

?

THE CYTOSKELETON IN PROTISTS

159

The rate of synthesis of these minor actins would be very weak or this synthesis would take place at precise times of differentiation (Vandekerckhove and Weber, 1980). In the ciliate Oxytricha, a macronuclear gene of actin, when cloned, presents an homology of 65.7% with a yeast actin gene, its sequence foreshadowing the synthesis of an actin shorter than the others (356 amino acids in place of 374-373, and with an important difference in structure which could be related to a high functional specialization (Kaine and Spear, 1982); nevertheless, the presence of actin in Oxytricha has not been detected; furthermore the actins of ciliates are not well known at present. Thus, the overall arrangement of the actin has been conserved from unicellular organisms up to mammalians suggesting that, from a common origin, evolution has gradually separated different types of actin, coded for by different genes which constitute gene families; thus, only one gene of actin exists in yeast, one (or maximum 2) in the alga Dunaliefla (Marano et al., 1982) and in Oxytricha, 17 in Dictyostelium, 6 in mammals and birds (Vandekerckhove and Weber, 1981), some of which have already been cloned (Schwartz et al., 1980), and approximately 20 in humans (Humphries et al., 1981); moreover, it seems that introns are absent within the actin genes of Dictyostelium and Oxytricha, whereas there is one intron in those of yeast and Dunaliella. From these observations, it might be concluded that like Dunaliefla, hypotrich ciliates were separated very early from the principal trunk of eukaryotes (Humphries et al., 1981). G-Actin polymerizes to F-actin to give microfilaments (Mf); the structure will be seen later. This transformation, studied in vitro in the presence of salts, begins with a change of conformation from monomeric G-actin to monomeric F-actin, then continues with the process of nucleation [forming a tetramer (Tobacman and Korn, 1983)], and with the actual polymerization. During this phenomena, the lengthening of Mf is realized by adding monomers, each being associated with a molecule of ATP; this one is not immediately submitted to hydrolysis, but will be after the Mf is built (Mockrin and Korn, 1983; Pantaloni et al., 1983). Nucleation requires a critical G-actin concentration which varies according to actin types, the nature and concentration of ions present in the middle, and temperature; it is higher for the actin of Acanthamoeba and nonmuscular actins in general than for muscle actin (Table I) (Korn, 1978); the F-actin polymer of Acanthamoeba seems more labile than its muscle homolog; it is more sensitive to temperature changes (Pollard, 1975). The purified actin of Dicryosrelium polymerizes with a rate, an intensity, an optimal KCl concentration (0.1 M),and a critical concentration (1 pM) similar to those of skeletal muscle actin (Clarke and Spudich, 1977). Yet, F-actin of Physarum can depolymerize easily in the presence of Mg2+ (Totsuka and Hatano, 1970), and large pools of G-actin are found in the cell (Hinssen, 1972); apparently then, two pools of actin occur in the cytoplasm of Physarum, one of polymerizable actin and the other of nonpolymerizable actin, as found in blood platelets (Clarke and Spudich, 1977); certain factors may prevent polymerization, for example, proteins of the profilin

160

JEAN GRAIN

type able to associate with G-actin. In Amoeba, Ca2+ at low concentration (
B. ACTIN-BINDING PROTEINS(ABP) EXCEPTMYOSIN: THEIRROLES(TABLEVI) We shall come back later to the presence of myosin in protists, to its association with actin, and to the possible existence of proteins which regulate interactions between actin and myosin (troponin-tropomyosin) and are normally linked with actin in muscle cells. 1. The 95K protein of Dicryostelium (Glenney et al., 1980) has a Ca2+sensitive action; like the villin of intestinal villi (MW 95K), the gelsolin of rabbit lung macrophages (MW 91K or 270K?) and the actinogelin of Ehrlich tumor cells, it causes, in the presence of Ca2+, a breakdown of the microfilaments of F-actin and, by binding to G-actin, it prevents polymerization. In the absence of free Ca2+, its configuration changes and it then serves as a link between microfilaments, producing an F-actin gel. 2. The severin of Dicryostelium (MW 40K) (Brown et al., 1982; Yamamoto et al., 1982) binds to F-actin in the presence of Ca2 and causes the fragmentation of the microfilaments and then their partial depolymerization, which frees small oligomers and subunits of G-actin, producing a pool of mobile actin which can be moved toward any other site where it may be required (50 to 70% of the actin is free in the cell); the same pool can be used for exchanges of subunits with the microfilaments. 3. The profilin of Acanthamoeba (MW 11.7K) (Reichstein and Kom, 1979; Tseng et al., 1984) is distributed uniformly in the cell; it inhibits nucleation much more strongly than elongation of F-actin under physiological conditions; there is a similar protein in Physarum (Ozaki et al., 1983). 4. The fragmin of Physarum (MW 42K) (Brown et al., 1982; Sugino and +

TABLE VI ACTIN-BINDING PROTEINS FROM PROTISTS (EXCEPTMYOSIN) ~

Protist

Name

Acanthamoeba

Profilin

I 1,700

Inhibition of nucleation

Dictyostelium

Protein

95,000

Polymerization (G + F) Linkage - of Mf + gelBreaking of Mf -+ short fragments Prevention of polymerization

Dictyosrelium

Severin

40,000

Physarum

Fragmin

42,000

Acanthamoeba

Gelactin Gelactin Gelactin Gelactin

I I1 I11 IV

23,000 55,000

Proteins with analogous functions in other nonmuscle cells

Function

MW

Profilin, 16,000, spleen

Villin, 95,000, intestinal villi Gelsolin, 91 ,000 (or 218,000?), macrophages of rabbit lung

}

Breaking of Mf + short fragments with Ca2+ Partial depolymerization Breaking of Mf + short fragments } with Ca2+ Linkage of Mf

+ gel

64000 78,000

Actinogelin, ascites cells

Protein, 200,000, sea urchin ovocytes Gelsolin (without Ca2+) Filamin (200-250,000) (linking form) embryonic cells a-Actinin (55,000). sea urchin sperm and eggs Band 4-1 (78,000), erythrocytes

Physarum

p-Actinin

86,000

Polymerization G -+ F

Gelsolin (without Ca2+) Spectrin (900,000), erythrocytes (with one chain phosphorylated)

Dictyostelium

Membrane Protein

30,000 32,000

Linkage Mf-plasma membrane

a-Actinin, intestinal villi Protein 105-1 10,000, cross-bridges intestinal villi Protein 105,000, platelets (without Ca2+) Spectrin (without Ca2+)

Pseudomicrothorax

a-Actinin

?

Linkage Mf-Mt Linkage Mf-epiplasm (?)

a-Actinin ? Spectrin ?

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JEAN GRAIN

Matsumura, 1983), in the presence of Ca2+, cleaves the microfilaments giving short fragments thus lowering the tension of the actomyosin threads. After removing the Ca2+, this effect subsists for a long time; that means the fragmin remains bound to the actin. As Ca2 usually induces contractility in Physarum, it is unlikely that the fragmin is involved in relaxation in the presence of Ca2 ; it may be that it serves to disorganize the peripheral F-actin gel thus permitting contraction, or that just after contraction it recycles actin in the ectoplasm after breaking it up (Sugino and Matsumura, 1983). 5. The gelactins I, 11, 111, IV of Acanthamoeba (MW 28K to 78K) (Maruta and Korn, 1977; Korn, 1978) favor the formation of actin gels by linking the microfilaments together; they have the same role as the 200K protein of sea urchin ovocytes and coelomocytes (De Rosier and Censullo, 198 I), gelsolin, filamin, and the 95K protein of Dictyostelium (in the absence of Ca2+), although having a different MW; they are more similar to an a-actinin (?) (MW 55K), extracted from sea urchins gametes, which causes the gelation of actin. Gelactin IV has the same MW as the 4-1 band of erythrocytes which favors the maintenance of the F-actin gel generated by spectrin (Cohen and Korsgren, 1980). 6. The p-actinin of Physarum (MW 86K) (Korn, 1978) is made up of two subunits of MW and pl identical to these of actin; in the presence of MgCI,, it brings about the polymerization of the G-actin, forming a Mg2+-actinpolymer, and inhibiting the depolymerization of F-actin. 7. The a-actinin of Pseudomicrothorax has been detected by immunofluorescence in the cytopharyngeal apparatus, where microfilaments of actin linked to microtubules occur, and in the epiplasm, in the epiplasmic ridges, and all along the ciliary furrows. It may be involved in anchoring actin microfilaments on microtubules in its first location, and in anchoring kinetosomes within the epiplasm in its second site (de Haller, 1977), and perhaps even in the transient fixation of temporary microfilaments to the epiplasm. Its chemical characterization remains to be performed. 8. The 30K and 32K membrane proteins of Dictyostelium (Jacobson, 1980; Luna et al., 1981) are involved in the fixation of microfilaments to the plasma membrane, either by lateral or by terminal linking. They have the same role as aactinin of muscle cells, fibroblasts (Hoessli et al., 1980), and intestinal microvilli, the 105K protein constituting the cross-bridges in these villi (Glenney and Weber, 1980; Glenney et al., 1980), the 105K protein of platelets in the absence of Ca2+ (Rosenberg et al., 1981), and the spectrin (MW 900K) of erythrocytes (Fowler et a f . , 1981). Thus, the proteins able to bind to the actin of protists have various actions, the same protein being able, according to the conditions (with regard to Ca2+), to play different roles. They can prevent the polymerization of G-actin, promote this polymerization, cut the microfilaments giving short fragments, associate the microfilaments in a gelated network, depolymerize the F-actin, and link the +

+

THE CYTOSKELETON IN PROTISTS

163

microfilaments to plasma membranes. Other ABPs, not yet identified, certainly exist, for instance, those which enable the microfilaments to associate in tracti or bundles, and which in protists would have the same role as the filamin of mammalia, the fascin of sea urchins, the villin of intestinal villi, and the ABP of 260K of platelets. Clearly then, the state of the actin is conditioned not only by various physicochemical factors, but also by these binding proteins, which may be either temporarily or permanently bound.

C. ACTIN-CONTAINING STRUCTURES They consist of microfilaments (Mfs) of F-actin which are directly visible or visualizable by various procedures such as antibody labeling, decoration with HMM or the S1 fragment of myosin. The Mfs are generally 6 nm in diameter and 1 or more km in length. They may be either isolated, or collected in networks (gels), or associated in parallel in tracti and bundles. They may be located at the periphery of the cell (cortical zone, junctions, microvilli), or in intracellular tracti, or in the contractile ring of division, or in regions of active phagocytosis. In protists, these Mfs have been detected in different forms and with different localization (cortical region, interior of the cell); in some particular cases, the presence of Mfs has not been detected, though that of actin involved in contractility has been demonstrated. 1. Arrangement of Mfs in the Cortical Regions In Amoeba proteus, the Mfs localized in the cortical layer (ectoplasm) form a network (gel) involved in the cortical contraction; this plasmagel sheath is separated from the membrane in the forepart of pseudopods (frontal zone) (Grebecka and Hrebenda, 1979); elsewhere, the Mfs are attached to the plasma membrane at points disposed in a polygonal arrangement (Korohoda and Stockem, 1975) (Fig. la). However, methods using fluorescence of HMM or of anti-actin for light microscopic observation show an uniform distribution of actin, without distinctly visible structures (Opas, 1980). The isometric contraction is accompanied by an arrangement of the Mfs parallel to the membrane (Grebecka and Hrebenda, 1979; Wehland et a l . , 1979) (Fig. lb). In Aranthamoeba, the arrangement of the Mfs is very similar to that of Amoeba; the network may form nodes that become closer in places in glycerinated models (Pollard, 1975). In Dictyostelium, the Mfs of the cortical network are also attached to the protoplasmic face of the plasma membrane (Spudich, 1974), on proteic sites, by lateral or terminal strong links (Jacobson, 1980; Luna er a l . , 198 1). In Physarum (see review in Vanden Driessche, 1979), upon contraction, Gactin polymerizes to F-actin and Mfs arranged in a network are produced; they are associated with myosin. Relaxation is accompanied by a partial depolymer-

164

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a

FIG I . Cortical organization in Amoeba proreus. (a) Ectoplasmic microfilaments form a threedimensional network; they are attached to the plasma membrane at points disposed in a polygonal arrangement (inset). (b) Anterior part during isometric contraction; the microfilaments are roughly parallel to the membrane. (a) Redrawn from Korohoda and Stockem (1975); (b) drawn from a micrography after Grebecka and Hrebenda (1979).

ization of the Mfs. Here again, the actin Mfs are attached to the membrane at least at the level of the frontal zone of the plasmodium. The Mg*+-actin polymer, obtained experimentally, has the same helical structure as muscle F-actin, but possesses interruptions, which is quite specific to Myxomycetes (Tanaka and Hatano, 1976). In the heliozoan Acrinophrys, the leading margin of the large pseudopodium which surrounds the prey after capture contains an irregular network of Mfs, the nature of which remains unknown; some of these Mfs could be linked to the membrane of the growing pseudopodium (Hausmann and Patterson, 1982). In the alga Acetabularia, longitudinal filaments, which are involved in the cytoplasmic current, contain an actin-like protein (Dazy er al., 1981). In the terminal knob of tentacles of the suctorian Heliophrya, single Mfs are associated with the epiplasm, a proteic layer situated just under the plasma membrane (Hauser and Van Eys, 1976) (Fig. 2). In Paramecium, some cortical tracti, attached to the epiplasm, are composed FIG. 2. Diagram of a longitudinally sectioned tentacle of Heliophrya erhardi in the inactive state. The haptocysts (ha) in the knob region are held in a vertical position by the thin epiplasmic filament layer (efl). At their anchorage site (as) the microtubules of the outer circle (ot) reunite with those of the inner ribbons (it) from which they are separated at the level of the sleeve region (slr). In the cytoplasm, at the tentacle base, microfilaments are arranged parallel to the microtubules (mf) within and outside the microtubule cylinder, and as a ring around the microtubule skeleton (mfr). From Hauser and van Eys (1976).

166

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of filaments which could contain actin (Tiggeman and Plattner, 1981; Tiggeman er al., 1981); they can be considered as the “infraciliary lattice”; but this is not decorated with HMM, and it is doubtful whether actin is present in these tracti (Cohen et al., 1984). The microfilaments occurring in the collar and other parts of cells of some choanoflagellates (Fig. 7c-e) consist of actin; their arrangement is comparable with those found in the collar tentacles of sponge choanocytes and in the microvilli of intestinal epithelial cells (Leadbeater, 1983).

2 . Endoplasmic Mfs These are associated with various organelles, particularly sets of microtubules. In Devescovinidae (termite flagellates), a thin layer of Mfs surrounds the middle and posterior parts of the microtubular axostyle, without any visible links with these microtubules (Tamm, 1976) (Fig. 3). In the tentacles of Heliophrya (Suctoria), a transversal ring of Mfs surrounds the base of the microtubular axoneme, other Mfs lying along the microtubule (Hauser and Van Eys, 1976) (Fig. 2). In the cytopharyngeal apparatus of the ciliate’Pseudomicrothorax, Mfs make links between neighboring nemadesmas, while others constitute a circular sheet attached to be outside of the nemadesmal basket. and some are associated with

FIG. 3. Transverse section through the posterior part of the axostyle complex of the Cryptotermes cuvifrons devescovinid; a microfilamentous sheet (mf) surrounds the axostyle. x 35,000. [FromTamm (1978). Reproduced from The Journal of Cell Biology. 1978, 78,76-92 by copyright permission of The Rockefeller University Press.]

THE CYTOSKELETON IN PROTISTS

167

FIG. 4. Pseudornicrorhorar dubius. (a) Thin section through the cytopharyngeal basket at the level of the cytostome. nd, Nemadesmata; ndl, nemadesmal lamellae. Bar, 1 pm. (b) A higher magnification of a nemadesmal lamellae reveals two arm-like structures projecting from each microtubule (arrowheads). Filamentous material (mf) is seen associated with the arm-like projections. Bar, 10 nm. (c) Thin section of a nemadesmata (nd). Bar, 0.5 pm. (d) Detail of a: filamentous material (mf) is associated with the nemadesmata (nd). Bar, 0.5 pm. (e) Thin section of an isolated cytopharyngeal basket, taken through the decorated region (see Fig. 5d) as indicated in the inset. Filamentous material (mf) is seen connecting the nemadesmata (nd). Bar, 0.25 pm. From Hauser et nl. (1980).

the arms born by microtubules of the nemadesmal lamellae (Fig. 4); for the latter, it is likely that links exist with the membrane of the growing digestive vacuole (Hauser et al., 1980); the constituent actin of these Mfs has been detected by immunofluorescence (Fig. 5).

THE CYTOSKELETON IN PROTISTS

169

In the buccal apparatus of Tetrahymena, a thin layer of Mfs (the superficial part of which can be decorated with HMM) is situated under the oral ribs of the right dorsal buccal wall (Fig. 6); some elements of this layer could be directly linked to the plasma membrane of the buccal cavity (MCtCnier, 1981); the Mfs, parallel to the ribs (Williams and Bakowska, 1982), are thickened at regular intervals, and are linked to the microtubules of the ribs (Sattler and Staehelin, 1979). In Paramecium, the detection of actin by HMM (with fluorescence, or with electron microscopy), shows the presence of actin Mfs at the level of digestive vacuoles, either in formation (pouch) or in transfer along the postoral fibers, and also all around the new digestive vacuoles which become free in the cytoplasm (Cohen et al., 1984). In the chrysomonad Synura, short Mfs, decorated with HMM, are stretched between the membrane of the scale-forming vesicles and that of the plast along which the vesicles are displaced (Mignot and Brugerolle, 1982) (Fig. 7). 3 . Presence of Actin without Evident Mfs The gliding of the sporozoites of Eimeria (Sporozoa) could involve a contractile system of Mfs since it is inhibited by cytochalasin B, as is the capping of surface ligands at the back of the cell. Nevertheless, Mfs have never been detected (Russell and Sinden, 1981). Actin is present in the cytoplasm of the euglenian Distigma, where it seems implicated in the euglenoid movement (a mode of nonflagellar locomotion), but Mfs are not described (Gallo et al., 1982). It is the same situation for the volvocale Dunaliella where actin could be involved in the migration of the nucleus toward the flagellar pole, just before mitosis (Marano et al., 1982). Diffuse actin has been detected in isolated pellicles in the ciliate Pseudomicrothorax (Hauser et a l . , 1980) (Fig. 5 ) . Doubts continue about the presence of actin at the level of somatic kinetosomes, as has been described in Paramecium (Tiggeman and Plattner, 1981; Tiggeman et a l . , 1981); in this ciliate, cyclosis could be due to an actomyosin complex, but Mfs have not been found (Kuznicki and Sikora, 1971, 1973; Sikora, 1981). In the axopods of Echinosphaerium (Heliozoa), the presence of actin Mfs has only been revealed in isolated fractions, by decoration with HMM (Edds, 1975). It is therefore conceivable that the rarity of Mfs or the transience of the

Fic. 5. Pseudomicrothorax dubius. Cell ghosts are stained with anti-actin (aA) prepared on a sucrose gradient (a) or on a metrizamide gradient (b) and with anti-a-actinin (aaA) (c). (d-g) Isolated cytopharyngeal baskets: (d) phase contrast micrograph of an isolated organelle showing the decorated region (dr) of the nemadesmal bundles (e) and (0 fluorescence pictures after staining with anti-actin (g) isolated organelle stained with anti-a-actinin (aaA). Pellicle structures stained with anti-tubulin (h, aT), anti-actin ( i , aA), and anti-a-actinin Q, aaA). cp, Cytoprocte; epr, epiplasmic ridges; mn, macronucleus. Bars, 20 pm. From Hauser er al. (1980).

P

UMN

FIG.6. Schemes of the oral apparatus of Te/rahyrnena. (a) General organization. (b) Transverse section. UM, Paroral or undulating membrane (two ciliary rows); Ml-3, adoral membranelles (three ciliary rows each). Several microtubular formations are issued from these organelles: the connectives (C), the rib microtubules (R Mt), and the deep-fiber bundle (DFB). Actin microfilaments (A) lie between the rib microtubules and the fine filamentous network (FFR). The nature of FFR, of the undulating membrane network (UMN), and of filaments (Fb) on each side of the UM is still unknown. FIG. 7. Actin microfilaments in flagellates. (a) Synura: the microfilaments (Mf), stretched between the scale-forming vesicle (V) and the plast (P), are seen in cross sections. (b) Synura: oblique section perpendicular to a at the level of the Mf. A. Axial component of the scale-forming system; Mt, microtubules; S, scale in formation. (c-e) Codosiga botryris: longitudinal (c) and transverse sections (d. e) of the collar tentacles showing actin Mf (arrows). (a) X60.000; (b) X32.500; (c) X33.600; (d) X60,OOO; (e) X60,OOO. (a, b) From Mignot and Brugerolle (1982); (c-e) from G. Brugerolle, unpublished.

THE CYTOSKELETON IN PROTISTS

171

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JEAN GRAIN

arrangement of the actin as F-actin is responsible for the nondetection of Mfs in those cases where actin, proved to be present, seems involved in cell motility. 111. Myosin

Most of the nonmuscle cells containing actin also possess a protein which is analogous to muscle myosin; this protein is a minor one in this case, while it is major in muscle cells. In protists, myosin has first been detected in organisms with ameboid movements (Myxomycetes and Amoeba). A. CHARACTERISTICS OF PROTIST MYOSINS 1. Molecular Constitution

Except for the myosins I of Acantharnoeba which have only one heavy chain and one pair of light chains, and form a globular tailless molecule (Pollard, 1981), the myosins all have the same monomeric structure as muscle myosin, i.e., 2 heavy parallel chains which form the tail, and 2, 3, or 4 light chains which are connected to the ends of the heavy chains to form the two globular parts of the head; the light chains have different compositions and functions, and are coded by different genes. The variability of the myosins lies in their size and their MW which vanes from 170K (Acanthamoeba) to 450-500K (Table VII); this variability does not exclude affinities with actins of other origins: the myosin of Physarum can be linked with muscle actin and acquires an ATPase activity (Vanden Driessche, 1979). Their common characteristic is their ATPase activity (Table VII) located in one of the light chains (d’Haese, 1980; Morita and Matsumoto, 1980); it is generally Mg2+ dependent and stimulated by actin, it varies according to the origin of the myosin and the cations present in the medium, and it is stimulated by Ca2+ in Physarum, Dictyostelium, Amoeba, and Chaos, and in Acanthamoeba for myosin 11; EDTA stimulates the ATPase activity of the myosin I of Acanthamoeba but not that of the myosins of Dictyostelium, Physarum, and Amoeba, which is different from muscle myosins (Clarke and Spudich, 1977). The ATPase activity of pure myosin is also activated by actin in Dictyostelium and Physarum, while a cofactor is required for the myosin I of Acanthamoeba (Clarke and Spudich, 1977); this cofactor is a kinase which phosphorylates the heavy chain of myosin I (Kom, 1978), which is different from the phosphorylation necessary for the light chains of the nonmuscle cells of vertebrates, and muscle cells. The phosphorylation sites (serins) of the myosin I1 of Acanthamoeba are near the free end of the tail (Collins et al., 1982). 2 . Polymerization Under physiological conditions, the nonmuscle myosins assemble in slender bipolar filaments, shorter than those of the muscle myosin. This polymerization

TABLE VII NONMUSCLE CELLS

COMPARISON OF MYOSINS OF

ATPase activity ( p n o l rnin - 1 rng - 1 ) Source Human platelet

Rabbit macrophage

Bovine medulla

Glial cells

Physarum

K+

Subunits 200,000 x 19,000 X 16,000 X 200,000 20,000 15,000 200,000 x 20,000 x 17,000 x 200,000 19,000 17,000 225,000 x 21,000 x 17,000 x

2 2} 2

+ EDTA

0.9 l o

2 2 2

+

Mg2

Mg2+

+

0.002 0.006

2

0.41

0.57

0.045

0.35

0.64

0.01

0.0002

0.56

2 2 2

Ca2

0

0.38

=

2.0

+ F-actin

0.029 0.170

Not phosphorylated

=

0.045 0.40

Without cofactor protein With cofactor protein

0

2

0.017

0.001

I

o.oO04

0

7

0.44

iT

Phosphorylated

(continued)

TABLE VII

(Continued)

ATPase activity (pmol minSource

Subunits

Dicryosteliurn

Amoeba proteus

Chaos carolinensis Acanrhamoebn castelhii

IA

IB

IC

I1

210,000 18,000 16,000 225,000 225,000 130,000 X 1 17,000 X 1 14,000 X 1 125,000 X 1 27,000 x 1 14,000 X 1 130,000 x 1 20,000 x I 17,000 X 1 14,000 X 1 170,000 X 2 17,500 X 2 17,000 X 2

K+

+ EDTA

I

mg - I )

Mg2

0.08

0.005

7

0.2

0.01

0.14

0.1

0.02 0.03

f f

0.12

0.01

3.2

0.38

0.04

7

0.08

f

0.08 1.23

Not phosphorylated Phosphorylated with cofactor

0.12

0.83

0.036

f

0.060

With or without cofactor protein

0.02

+

{

Mg2+

+ F-actin

Ca*

+

0.18

175

THE CYTOSKELETON IN PROTISTS

a

FIG. 8 . Drawings of models of Acunrhamoeba myosin I1 thin filaments. (a) Two-dimensional scale drawing with approximate dimensions. (b) Exposed view showing the proposed longitudinal positions of the 16 myosin 11 molecules. [After Pollard (1982). Reproduced from The Journal of Cell Biology. 1982, 95, 816-825 by copyright permission of The Rockefeller University Press.]

has been carried out in v i m in various protists; thin filaments 600 to 800 nm in length are formed from the myosin of Dictyostelium, in the presence of MgCI,, while in its absence loose aggregates are formed; the purified myosin of Physarum gives either aggregates or thin filaments and that of Amoeba assembles in filaments 1500 nm in length (in Clarke and Spudich, 1977). The myosin 11 of Acanrhamoeba can form two types of bipolar filaments during polymerization: thin filaments of diameter 6 to 7 nm and length 200 nm, composed of 16 molecules arranged opposite in twos, the heads of which are separated, from one pair to the next, by 15 nm (Fig. 8), and thick filaments of diameter 14 to 19 nm and length 300 nm, composed of 40 molecules, apparently formed by lateral association of 3 to 6 thin filaments in the presence of divalent cations (Mg2 , 67 mM; Ca2 , 10 mM) (Pollard, 1982). The myosin I of Acanthamorbu cannot form filaments on its own; it can, however, link up to actin microfilaments (Pollard and Korn, 1973). +

+

B. STRUCTURES While in skeletal muscles, the myosin filaments are permanently visible (thick filaments), they are practically undetectable in nonmuscle cells; condensations of myosin may be seen on the stress fibers (actin) of HeLa cells (Hermann and Pollard, 1981; Sanger et al., 1983), or in the terminal web of intestinal microvilli (Herman and Pollard, 1981), but they do not have the form of filaments. It might be that the myosin polymerizes little and for only a short time; at the moment of

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JEAN GRAIN

contraction, its normal form would then either be rare and very thin filaments, or monomers (Herman and Pollard, 1981). This hypothesis is supported in the amoebae Chaos where the in vivo assembly of myosin into supramolecular structures is initiated by conditions favoring the contraction, but physiologically reversible, hence temporary, scarcely visible filaments are formed (in Clarke and Spudich, 1977). In a glycerinated model of Amoeba discoides (which is capable of contraction), we can observe, in the absence of ATP, a loose network of fine filaments of myosin (2 to 3 nm in diameter) within which are dispersed tracti 15 to 25 nm in diameter (Fig. 9a); the presence of ATP induces contraction, and the fine filaments assemble into numerous thick tracti which order like the myosin fibers of muscles, giving an overall periodic appearance; an important physiological role is evident (Fig. 9b) (Holberton and Preston, 1970). In Physarum, contraction causes myosin bundles 12 nm in diameter to appear transiently; they disappear upon relaxation (Vanden Driessche, 1979). In Acanthamoeba, myosin I1 seems to always be polymerized in thin filaments, rather difficult to detect (Pollard, 1982). In moving Amoeba proteus, thick filaments are detected mainly in the uroid, tightly associated with actin microfilaments (Stockem et al., 1982). In the anterior pole of the coccidian Toxopfasma gondii, myosin has been detected by immunofluorescent localization; it may be associated with the conoid, and not be resolved by electron microscopy; it would be involved, probably with the help of actin, in the movement of the conoid and in the invasion of host cells (Schwartzman and Pfefferkorn, 1983).

IV. Roles of Actin and Myosin The simultaneous presence of these two proteins in nonmuscle cells results in a contractility phenomena. However, there are cases in which myosin has not been detected, and where actin is abundant, playing a role in several cellular mechanisms (transport, division) in the protists. A. ACTIN-MYOSININTERACTIONS It is necessary to bear in mind that in muscle cells the attachment of myosin heads ( S 1) to actin stimulates the myosin Mg2 -ATPase activity, hence ATP is hydrolyzed, liberating the energy used to produce a rotation of the myosin heads and the gliding of the filaments (contraction); the attachment of myosin to actin is itself Ca2+-dependent, and this Ca2+ regulation of the contractile activity of the actomyosin complex can occur in two modes. In the actin-linked mode, which alone exists in skeletal muscles, the site of fixation of the myosin on actin is usually hidden by a protein linked to actin, tropomyosin, in the absence of free +

THE CYTOSKELETON IN PROTISTS

177

FIG.9. Amoeba model cells. (a) Representative area from the cytoplasm of a nonactivated model showing a network of 20- to 30-A microfilaments (arrows); the few thick filaments are not noticeably aligned. (b) A filament tract in an activated cell sectioned longitudinally; linear repeat arises from the lateral alignment of thick filaments (double-headed arrows). (a) X56,800; (b) X52,OOO. From Holberton and Preston (1970).

Ca2+ (conc. M); as a result of stimulus, Ca2+ is liberated in the hyaloplasm; when free Ca2+ concentration is M, it becomes attached to another protein, troponin, also linked to actin; troponin is then deformed, dis-

FIG.9B.

See legend on p. 177.

THE CYTOSKELETON IN PROTISTS

I79

placing the tropomyosin to which it is linked, and the actin site of myosin fixation is revealed. This interpretation is contestable, because it would mean that in vivo, myosin heads, even when charged with ATP or ADP + Pi, always remain attached to actin even during the relaxation of the muscle fiber (in the absence of Ca2 ), where the troponin-tropomyosin complex inhibits myosinATPase activity by blocking the liberation of Pi from the S I -ADP-Pi-actin complex (Chalovich et a f . , 1981). In the myosin-linked mode, which occurs in several smooth muscles, one light myosin chain is capable of blocking (due to its particular state it may be not phosphorylated) the attachment of myosin to actin in the absence of Ca2 , without the intervention of a troponin-tropomyosin system. Furthermore, it is to be noted that actin and myosin filaments are always polymerized, and have a regular distribution. In nonmuscle vertebrate cells in cultures, tropomyosin and troponin have been detected in many cases (reviews of Lazarides, 1975; Korn, 1978; Schloss and Goldman, 1980; Sanger et al., 1983), which evokes an actin-linked mode of regulation of the contraction. In other cases, as in contraction of the terminal web of intestinal epithelium cells (Keller and Mooseker, 1982) or clustering of lymphocyte membrane receptors (Bourguignon et a f . , 1982), contraction of the actomyosin complex is regulated by Ca2 , which stimulates the phosphorylation of a myosin light chain by means of a kinase, the activity of which Ca2+- and calmodulin-dependent (myosin-linked mode). Homologies of structure and function between calmodulin and troponin C (a subunit linked to Ca2+) have been suggested to relate the two modes of regulation of contraction by Ca2+ (Korn, 1978). It should be remembered that the formation of visible filaments of myosin is temporary, limited to the contraction phase, and that a cofactor is required to obtain ATPase activity in the actin-myosin complex; this cofactor could be a kinase that phosphorylates the light myosin chains. In protists, the actin-myosin interactions are variable. The ATPase activity of Physarum myosin is enhanced by association with muscle actin, and reciprocally, the actin of Physarum stimulates the ATPase activity of muscle myosin (Vanden Driessche, 1979). The contractility of the actomyosin system is regulated by Ca2+; in Amoeba and Dictyosfelium, a concentration of about l o p 6 M of Ca2+ is required for contractions. The regulation by Ca2+ is based on the actin-linked mode in Dictyosfelium (Korn, 1978), where the response to a chemotactic stimulus results in an immediate incorporation of 32P in the 210K heavy myosin chain (Malchow et al., 1981). In Physarum, the two modes, actin-linked and myosin-linked, occur (Korn, 1978); since an extract of Physarum can restore the Ca2+ sensitivity of a muscle actomyosin which was desensibilized during its purification (Vanden Driessche, 1979), it certainly contains a homolog of troponin; furthermore, in the presence of Ca2+, addition of muscle tropomyosin to the Mg2 -actin polymer causes a change of conformation of the latter (Tanaka and Hatano, 1976); in fact, a Ca2+-sensitive factor has been isolated from +

+

+

+

180

JEAN GRAIN

Physarum; this factor must be linked to the actin; it is composed of 3 proteins of MW 38K (not very different from troponin T), 24K (not very different from troponin I), and 35K (not very different from tropomyosin) (Clarke and Spudich, 1977). Such a factor also seems to occur in Dictyostelium (Clarke and Spudich, 1977). The role of fragmin (which, in the presence of Ca2 , cuts the microfilaments of actin and decreases the tension of the actomyosin tracti, leading to relaxation in Physarum) seems to contradict the fact that contraction is stimulated by Ca2+; thus it is not exactly known whether Ca2+ acts in Physarum as a stimulator of the contraction or as an inhibitor of cytoplasmic streaming (Sugino and Matsumura, 1983). In the particular case of Acanrhamoeba, Ca2+ does not stimulate contraction (Marx, 1975); the actin of Acanthamoeba alone stimulates in vitro the ATPase activity of muscle myosin; if muscle tropomyosin is added, this stimulation is enhanced, whereas addition of muscle tropomyosin to muscle F-actin reduces the stimulation of the ATPase activity of the muscle myosin (Kom, 1978). The actin and myosin filaments which interact in contractile phenomena are not permanently constituted in protists which have ameboid movements. In myxomycetes, the contraction is accompanied by temporary polymerization of actin and myosin; the myosin involved is not markedly polymerized, slipping doubtlessly occurring between microfilaments of actin and oligomers of myosin (Vanden Driessche, 1979). In amoebae the solidification of ectoplasm in posterior parts of the cell, and the gelation of endoplasm in the frontal part are related to depolymerization and repolymerization of actin (reviewed in Grebecki, 1982); the actin microfilaments are visible only in the moving amoeba. In Physarum, a connectin-like protein has been detected, which forms, beneath the plasmodium cell membrane, an elastic network that may act as a cytoskeletal structure (Ozaki and Maruyama, 1980). An actomyosin complex seems to govern the contraction of the axopods of Echinosphaerium (Heliozoa), where movements take place independently of the microtubules; in the isolated cytoplasm, where microfilaments and thicker filaments (perhaps of myosin) have been detected, contraction is initiated by calcium ions at concentrations above 2.4 x lo-’ M ,and relaxaM ATP (Edds, 1975). In immobilized tion occurs on the addition of Paramecium aurelia, endoplasmic cyclosis would imply interactions between actin and myosin, but there is no direct probe for this (Kuznicki and Sikora, 1971, 1973; Sikora, 1981). The contractility of the sphincter which surrounds the adhesive disk in Giardia is probably due to actomyosin (actin, myosin, and aactinin have been detected) (Erlandsen et d . , 1978). The overall rotary movement of the axostyle, flagella, nucleus, and Golgi apparatus of devescovinid flagellates is not linked to a dynein microtubule system, since specific inhibitors of the dynein-ATPase do not prevent the ATP-mediated reactivation of the glycerinated model, contrary to myosin-ATPase inhibitors (Yamin and Tamm, +

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1982); the motive force for rotation is probably generated by interaction between visible periaxostyle microfilaments and myosin filaments situated in the peripheral ectoplasm (Tamm, 1976, 1978); however, the presence of such myosin filaments is not yet firmly established. Evidently then, though we can be certain of the presence of actomyosin involved in contraction and locomotion in amoebae and myxomycetes, elsewhere, whenever there is cell movement with the proven presence of actively involved actin, that of myosin is not definitely established, but only suggested by indirect indications. Indeed, numerous cases are known in which the presence of myosin is envisaged, though in the absence of any proof, even indirect, of its presence: in Actinophrys (Heliozoa), the capture of prey is performed by small pseudopodia, which contain filamentous material and degraded microtubule axonemes; ingestion is due to a large pseudopod the filaments of which are 6.3 nm in diameter; the presence of myosin has not been proven, so the hypothesis of a traction force generated by an actomyosin complex, with attachment of the microfilaments to the plasma membrane (Hausmann and Patterson, 1982), is without actual support. In addition, the hypothesis of the action of actomyosin in ingestion by the tentacles of the suctorian Heliophrya (Hauser and Van Eys, 1976) has not received any confirmation, even if microfilaments 5 to 7 nm in diameter have been detected in the tentacles; this example seems to be closer to the mechanism that occurs in the normal oral apparatus of ciliates (Section IV,B).

B . ROLEOF ACTINI N

THE

ABSENCE OF MYOSIN

The microfilaments of actin can be linked to microtubules, to membranes, or both. They are thus elements of a mechanism which directs intracellular displacement of organelles, or localized contractile systems. Actin may also play a role in locomotion without any microfilaments being detectable. Here are some examples.

I . Actin and Ingestion Microfilaments of diameter 4 to 6 nm establish links between microtubules and membranes in the cytopharyngeal apparatus of the ciliate Pseudomicrothorax; they are attached to the arms of the microtubules of the nemadesmal lamellae which support the cytopharynx, and they are fixed at their other end to the membrane of the digestive vacuoles (Hauser et a l . , 1980); the presence at this level of an ATPase activity (Hauser and Hausmann, 1982) indicates that the microtubule-microfilament interactions provide the motive force for the displacement of the digestive vacuoles into the cytopharynx. Microfilaments of actin appear at the site of formation and around the di-

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gestive vacuoles during phagocytosis in Paramecium; they play a role either in the formation of the vacuoles, or more likely in their course along the microtubules of the postoral fibers (Cohen et al., 1984). In Tetrahymena, the layer of actin microfilaments situated under the oral ribs (Fig. 6), which establishes a link with the overlying microtubules and with the membrane of the buccal cavity, probably plays an important role in the formation of the digestive vacuole; its elements are contractile (MCtCnier, 1981); their contraction causes closure of the ingestion pouch (Nilsson, 1976; Sherman et al., 1981). Ca2 at low concentration stabilizes the cytoskeleton which supports the walls of the pouch, while at high concentration it induces contractility and closure of the pouch (Sherman et al., 1982) by activating an ATPase. The level of free Ca2+ ions may be regulated by a calmodulin (Williams and Bakowska, 1982) present at the level of the buccal apparatus; a calmodulin inhibitor, trifluorperazin, prevents the formation of digestive vacuoles. The exact mechanism of the relationships between microfilaments, membrane, and microtubules has not yet been fully elucidated however. The existence of calmodulin in Tetrahymena has been reported several times (Jamieson et d.,1979; Suzuki et al., 1981) in ciliary fractions and in the supernatant (Nozawa, 1982); it is a protein of MW 15K which increases the activity of the guanylate cyclase of Tetrahymena in the presence of Ca2 . It has no influence on the activities of the adenyl cyclase and phosphodiesterase of the ciliate, but it activates the phosphodiesterase of the rat brain and has no effect on the guanylate cyclase of different mammal cells. The calmodulins extracted from the cells of rats, sea anemones, Chlamydomonas, and Dicfyostelium do not activate the guanylate cyclase of Tetrahymena, while those of Paramecium do; the calmodulins of these two ciliates seem evolutionarily close; the amino acid sequence of the calmodulin of Tetrahymena differs from that of the rat brain by I I substitutions and one deletion, which is linked to the specificity for different substrates. In the tentacles of the suctorian Heliophrya (Fig. 2), the microfilaments situated under the epiplasm of the end knob must be passively attached to the apex of the microtubule ribbons; ATP induces their contraction (and that of the epiplasm); while food is being taken, these microfilaments serve to push apart the microtubule ribbons, which increases the diameter of the ingestion passage; moreover, by attaching to the internal static arms of the microtubules of the axonemal ribbons on one part, and to the peritrophic membrane on the other part, the microfilaments must generate the force which draws the membrane inward (Hauser and Van Eys, 1976). The retraction (or contraction) of the tentacles in Discophrya is stimulated by Ca2+ (Al-Khazzar et a / . , 1983); it occurs without disorganization of the microtubules in Trichophrya (Mogensen and Butler, 1983). The microfilaments of the sheath surrounding the nemadesmas and those situated between these nemadesmas of the cytopharyngeal basket of Pseudomic+

+

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rothorax must, by their link with the microtubules. play a role in the contraction and distension of the basket (Hauser et al., 1980);.ATPase activity is evident at the level of the sheath (Hauser and Hausmann, 1982). 2. Actin and Positioning In Heliophrya, the microfilaments of the base of the tentacle knob may be involved in positioning the haptocyts (extrusive organelles) (Fig. 2) (Hauser an Van Eys, 1976). In the axopods of centrohelidian Heliozoa, the displacement and the positioning of the kinetocysts and their attached membrane domains are explained by the supposition that relations exist between microfilaments and stable plasma membrane domains, or between microfilaments and the axonemal microtubules; actin has not been detected at this level, but this hypothesis is supported by the existence of microfilaments in the food-cup tentacles (pseudopods without microtubuls) (Fig. 10) (Bardele, 1976). Some filaments, detected in the cortex of Paramecium aurefia, are linked to the epiplasm; they form a network (fasciae) with meshes in the center of which the trichocysts insert their tip, without any direct contact with the fasciae; these filaments would play only a passive role by simply determining the site of insertion of the trichocysts (Plattner et a f . , 1982); nevertheless, it is not certain that these filaments contain actin. The attachment of trichocysts to the cell membrane is a necessary condition for a correct positioning of the nuclei during the division in P . aurelia; this attachment would induce changes into the membrane and the cortex yielding a precise cortical state necessary for the control of this positioning; this latter would involve cytoskeletal links (microfilaments or microtubules) between surface and nuclei (Beisson and Rossignol, 1975; Ruiz et a l . , 1976; Cohen and Beisson, 1980). In Synura (Chrysophyceae) the actin microfilaments are, without doubt, responsible for the anchorage of the scale-forming vesicles along the plasts; they form ribbons of superimposed filaments (Fig. 7), which disappear after the mineralization of the scale is achieved (Mignot and Brugerolle, 1982). 3 . Actin and Division The assembly of the buccal filament complex of Tetrahymena is necessary for the completion of stomatogenesis (Gavin, 1976); cytochalasin B, applied before its assembly (stage 4), stops morphogenesis; after this stage, cytochalasin B has no effect, and the division is completely achieved. Consequently, cytochalasin B would have also no effect on the filaments of the division ring which is situated at the level of the division furrow. This observation introduces the problem of the presence or absence of actin in the division “contractile” ring in ciliates; it is known that cytochalasin B (and cytochalasin D)inhibits cell division in mouse

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ff

MT

c

1

FIG. 10. In centrohelidian axopod kinetocysts (K),attached to highly ordered membrane domains as well as bacteria (B), perform bidirectional movements. The motive force for membrane domain displacement is envisioned in an actomyosin-like system located somewhere in the narrow space between plasma membrane and axopodial microtubules (MT).C , Cilium; FP, food-cupforming pseudopod. From Bardele (1976).

eggs and embryos, by producing a hypercontraction and a displacement of the microfilaments, with their detachment from the cell membrane (Siracusa et al., 1980);the absence of action of cytochalasin B on the division ring in Tetrahymena is a proof for the absence of actin.

4. Actin and Locomotion As seen earlier, sporozoites of Eimeria (Coccidia) are able to move by gliding on the substratum; this gliding, and the capping of surface ligands at the posterior part of the cell, are both inhibited by cytochalasin B; it is likely that this mode of locomotion involves contractile microfilaments which displace surface ligands, but these filaments have not been detected (Russell and Sinden, 1981).

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Actin is present in the cytoplasm of the euglenian Distigmu, but not in the cortex; in the propulsive phase of displacement by euglenoid movement, actin, concentrated in the cell dilatation which moves from the rear frontward, must enable the sol to gelate, thus providing a solid structure which acts as a mechanical support for the interactions between the plasma membrane and subpellicular microtubules which generate the motor force; in the recovery phase, actin is uniformly distributed in the cell, the gel is solated, and a current from the frontal part is produced; actin is never seen as microfilaments (Gallo and Schrevel, 1982; Gallo et al., 1982). Given all these examples in which actin seems to have a function in various manifestations of cell or intracellular movement (except for Synura), one may wonder if the presence of myosin is an absolute requirement for the action of actin (in which case its lack of detection would be due to technical difficulties), or if, on the other hand, as suggested for locomotor extension of cells in culture, mobility is different from the muscular type, and “une croissance et un desassemblage contrdes des fibres et des faisceaux d’actine commandent la formation et la rktraction des extensions mobiles de la cellule” (de Brabander, 1983); in that case, the microfilaments would not be contractile and this mechanism might explain the examples of transport and positioning of organelles at the very least. It would also be necessary to look for actin-binding proteins which could favor, for example, the formation of microfilament networks in the heliozoan pseudopods, as it occurs during the extension of pseudopods in rabbit macrophages in culture (Hartwig et al., 1980), or during the growth of the acrosomal process of sperm (Tilney et a / ., 1973). However, the hasty interpretations made concerning the passive role of actin cables (or stress fibers) thought to be devoid of myosin and only transmitters of intracellular tensions should be viewed with circumspection; recently, myosin, arranged in wide strips, a-actinin in narrow strips, and tropomyosin have been shown to occur in stress fibers of epithelial cells and fibroblasts in culture (Sanger et al., 1983), which makes these structures functionally comparable to the sarcomeres of skeletal muscle. The presence of myosin must therefore be sought, by sophisticated techniques, in all the systems described above in protists, in which actin is evidently present.

V. Tubulins and Microtubules We cannot deal in detail here with the work carried out on tubulins, microtubule-associated proteins (MAPS), construction of microtubules, and their organizing centers (MTOCs). Numerous reviews have examined these matters in depth (Grain, 1969, 1984; Bardele, 1973; Sleigh, 1973; Warner, 1974; Dustin, 1981, 1984; Cachon and Cachon, 1981a; Gibbons, 1981; Haimo and Rosenbaum, 1981). Microtubules are regarded as highly important elements of the

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cytoskeleton, involved in various cell functions: flagellar and ciliary locomotion, axopod and filopod formation, shape maintenance, cytokinesis, nuclear division, migrations, and positioning of intracellular organelles. We shall restrict ourselves to a rapid review of the arrangements and location of the microtubular structures present in protists, and then go on to look at the variability of tubulins, before finishing with the MAPS for which an attempt will be made to pinpoint their possible roles in the functions performed by the overall microtubular systems. Fusorial and nuclear microtubules will not be dealt with. A. MICROTUBULE ARRANGEMENTS AND MTOCs 1. Kinetosomes, Flagella, and Cilia Those of protists do not present any marked peculiarities in the arrangement of their microtubules. However, there are some anomalies in the number of microtubules in the flagellum of male gametes of gregarines (type 6 + 0 in Lecudina, type 3 0 in Diplauxis) (Premier et al., 1980) and in different mutants in Chlamydomonas. The axoneme of so-called sensory cilia is partially disorganized (Grain, 1984); in Dileptus, during regeneration, such an axoneme can be reorganized into an axoneme of locomotor cilia without any total dedifferentiation or resorption (Golinska, 1982); the reverse situation, i.e., conversion of a locomotor ciliary shaft into a sensory one, requires resorption of the locomotor cilium, its subsequent replacement, and the formation of a new sensory ciliary unit anterior to the transformed one (Golinska, 1983). The kinetosome can appear de novo (neogenesis) in Naegleria and Tetramitus (amoeboflagellates) and in the ciliate Oxytricha upon excystment (Grimes, 1973);in that case, the MTOC is probably a mass of astructural protein material, and would resemble MTOCs of some actinopodianaxopods. The kinetosome can appear in the immediate environment of another already constituted kinetosome (autogenesis), e.g., during the duplication of locomotor units which precedes the division; the MTOC is then a fibrogranular mass in Paramecium (Dippell, 1968) and in Tetrahymena (Allen, 1969); taxol, which inhibits the formation of a second centrosphere (MTOC for the mitotic spindle) in the amoebae of Physarum, does not prevent the building of new kinetosomes near the old ones in the same organism (Wright et al., 1980b, 1982). The kinetosome triplets can probably be used as germs for the polarized growth of the peripheral doublets of the axoneme, by adding tubular subunits at the tip of the A and then the B tubules (Warner, 1971) as has been shown in vitro for the A tubules in Chlamydomonas (Borisy et al., 1974). The central tubules are extended from their apical end in vitro (Borisy et al., 1974), but it cannot be ruled out that, in vivo, in Chlamydomonas, growth would also occur by the proximal addition of tubulin subunits, since their distal end is strongly attached

+

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to the flagellar membrane by a cap material (Dentler and Rosenbaum, 1977); their MTOC would be either the axosome or the distal cap.

2 . Kinetosome-Associated lntracellular Microtubules These generally join the MTOCs next to the base of the kinetosome; in some cases, they can be further away, but are then linked to the kinetosome by fibers of variable structure. The microtubules form sets directed either inward (deep fibers) or along the surface in the cortex (tangential fibers). The MTOC is most often a dense mass surrounding the proximal (deep) part of the kinetosome. Its presence is more or less widespread in the ciliates (in Grain, 1984), where it can form a complete or incomplete muff, or a cork, and can be prolonged in sheets or strings (Fig. 11); sometimes it is well developed, and links

A

a

FIG. 1 1 . Ciliate nemadesmata. (a) Transverse section. (b-g) Longitudinal sections showing the diversity of the relationships between nemadesmata and kinetosomes in AIIoiozona (b), Nassula (c), Cycloposrhium (d), Fronronia (e), Coleps (f), and Lacrymaria (g). The matrix plate is associated, sometimes through filaments (Mf), with the dense muff (M) surrounding the proximal part of the kinetosome. The matrix plate consists of several layers: deep (LP). middle (LM), apical (LA), and distal (LD). In Fronronia. a large periodic structure (arrowhead) supports the matrix plate. From Grain (1984).

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b C -

FIG. 12.

A three-dimensional reconstruction of the flagellar rootlet system and transition zone in

Codosiga borrvris. a, A-tubule; ax, axosome: b, B-tubule: c , C-tubule; ca, composite arc: cf. central

filament; i , interstitial material; mt, microtubule; tp, transverse plate. From Hibberd (1975).

several kinetosomes as in the flagellate fyrsonympha (Bloodgood et a / . , 1974). It can be composite, heterogeneous, with dense bands separating clear zones in the choanoflagellate Codosiga (Fig. 12) (Hibberd, 1975) and in the amoeboflagellate forms of fhysarum (parakinetosome) (Wright et a / ., 1979, 1980a). It can be a paracrystalline structure (preaxostyle) spread between two kinetosomes as in the flagellate Monocercornonoides (Brugerolle and Joyon, 1973), or a set of fine criss-crossed fibrils as in Saccinobaculus (McIntosh et a f . ,

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Fiti. 13. Diagram depicting the anterior end of the axostyle of Saccinohacrtlus. The primary row of microtubules (PR) has a centriole-like structure (C) at each end. These centrioles are surrounded by dense material and the rows of microtubules are linked each other by criss-crossed fibrils (F). [Redrawn from Mclntosh e/ a / . (1973). Reproduced from The Journal of Cell Biology, 1973, 56, 304-323 by copyright permission of The Rockefeller University Press.]

1973) (Fig. 13); this last example resembles to some extent particular cases in which sets of microtubules originate at an MTOC distant from the kinetosome, but which is connected to it by other fibers which can be fine filaments in Frontoniu, Prorodon, Brooklynellu (in Grain, 1984), and in the dorsal part of Srylunvchia (Gortz, 1982), a fibrillar reticulum with condensation nodes in Frontoniu (Didier, 1970), microtubules for the MTOC of the ribbon mt apl in Physurum amoebae (Wright et ul., 1979), or a set of microtubules or microtubules plus periodic fiber in Polytomellu (Steams and Brown, 1981). The simplest deep microtubular systems (microtubular kinetosome rootlets) are composed of only one or two microtubules; they are found in some ciliates (Fig. 14) (Grain, 1984). Many of the others are strips or ribbons of microtubules joined to each other in a single plane; in this category are the 5 to 9 microtubule ribbons of the amoeba of Physarum (Wright et a/., 1979), the ribbon of Chilomonus which remains associated with a dense lamella or with periodic lamellae over all its length (Fig. 15)(Mignot et ul., 1968), and those of the subkinetid arrays of the ciliates Phyllopharyngidea (Fig. 14) (Grain, 1984). (The cytopharyngeal ribbons of the ciliates are particular differentiations of tangential ribbons and do not belong to this category: see below.) The most developed are the bundles of

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d FIG. 14. Microtubular kinetosome rootlets in ciliates. They are simple in Dileprus or Monodinium (a) and in Bulantidium coli (b), and more developed in Ophryoglena (c); they form subkinetal arrays in the Phyllopharyngidea (d and e). From Grain (1984).

microtubules which make up the axostyles of the endosymbiont flagellates of insects (Fig. 16) and the nematodesmas (or nemadesmata) of certain flagellates and ciliates (Fig. 11). In the axostyle of Monocercomonoides, the microtubules are set out in parallel rows, with long links (desmoses) within a row, and shorter ones between neighboring rows; the arrangement is a paracrystalline square array (seen on transverse sections) (Brugerolle and Joyon, 1973). In Saccinobaculus, there is an irregular stagger between contiguous rows and there are temporary and permanent links

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FIG. 15. Schematic three-dimensional reconstitution of the anterior part of Chilornorios showing the different fibrillar systems. B , C, and E, Microtubular structures; RH, rhizostyle; BD, dorsal edge; V . vestibuluni; VP. pulsatile vacuole; F , and F2, flagella. From Mignot e t a / . (1968).

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Monocercomonoides

Pyrsonympha

Saccinobaculus Devescovinidae

---%-S. ambloaxostylus

---%as. lata

Tritrichomonas

FIG. 16. Various types of axostyles in flagellates. Note the presence of permanent bridges (dark) inside a row (dotted line) and of temporary bridges (light) between adjacent rows in two species of Saccinobaculus, S. ambloaxostylus, and S . lata. Dotted line, a row.

between microtubules, the latter being distributed along the microtubules with a periodicity which varies with the species (Mclntosh et al., 1973; Bloodgood and Miller, 1974). In Pyrsonympha, the axostyle can include 2000 to 4000 microtubules; only the primary row is linked to the MTOC; the other rows are first distant from the primary row, and then connect to it and to each other more closely (Bloodgood et al., 1974). The axostyle of Devescovinidae is of a different type; it consists of a wide longitudinal ribbon of microtubules which winds around on itself in a spiral; there are desmoses between neighboring microtubules within the ribbon but not between the microtubules of different turns of the spiral (Tamm, 1978). Finally, the axostyle of Trichomonas is only a simple cylinder of microtubules surrounding the nucleus (Hollande and Valentin, 1968). The nematodesmas are supported on a sometimes composite dense lamella,

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the matrix plate. They are constituted, in ciliates (Fig. I I ) , by bundles of parallel microtubules, arranged in a quincux array and connected to each other by desmoses staggered throughout their length; the aspect of the transverse sections is paracrystalline of a hexagonal type. The nematodesmas are usually localized around the cytopharynx, and collectively they form a more or less organized “basket” which may be consolidated by microfilaments and/or by microtubules, giving a “nasse” (Hauser et al., 1980; in Grain, 1984); they are rarely associated with somatic kinetosomes. In certain flagellates, they are also associated with an ingestion apparatus; their arrangement is of the square array type in Entosiphon (Mignot, 1963); their origin has not been detected in Kofoidinium (Cachon and Cachon, 1974a). Tangential microtubule systems are generally ribbons, or radiating sets of microtubules, which leave the base of the kinetosome laterally and run under the cell pellicle; they exist in Chilomonas (Fig. 15) (Mignot et al., 1968), in the choanoflagellate Salpingoeca where the number of microtubules regularly increases with the distance from the perikinetosomal MTOC (Laval, 1971), in the diplozoa Trepomonas where certain ribbons border the wall of the flagellar depression (Brugerolle et al., 1973), in Giardia where particular elements (microribbons) lie on each microtubule (Holberton, 1981; Holberton and Ward, 1981), and in Monocercomonoides where, in addition to the ribbon of microtubules running along the pellicle situated under the recurrent flagellum, there is a vast fan-shaped array of microtubules, the pelta (Brugerolle and Joyon, 1973); they are always present in euglenians, kinetoplastids, and dinophycea. In ciliates (Grain, 1984), the ribbons of tangential ectoplasmic microtubules are localized in the ectoplasm and constitute transverse fibers (T) and postciliary fibers (Pc) (Fig. 17). Transverse fibers are in the form of a 2 to 10 microtubule ribbon, which occurs laterally to the proximal part of the kinetosome at its left anterior side, close to triplets 3, 4, or 5 , then extends upward toward the pellicle and continues across to the left, parallel to the nearby pellicle and at right angles to the kinety; in the oral area of kinetofragminophorea, extensions of T fibers give up the cytopharyngeal ribbons. The postciliary fibers form a ribbon of 2 to 20 microtubules, which originates in the proximal part of the kinetosome, at its right-posterior side, very close to triplet 9; it extends upward into the interkinetal crest, toward the pellicle, and then continues posteriorly; in the somatic areas (Fig. 18), its plane can be orthogonal or parallel to the surface, or it can be folded, forming a prism; if well developed, it can associate with other ribbons and form extensive arrays like the km fibers of heterotrichida (Fig. 27a); in the oral area of oligohymenophorea, the Pc fibers extend into the oral ribs of the naked buccal wall. 3. Microtubules Unrelated to Kinetosomes Certain microtubules are of unknown origin; no MTOC has been found for them. Into this category fall the subpellicular permanent microtubules found in

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FIG. 17. Three-dimensional concept of the structures found in a small segment of the somatic cortex of Tefruhymcnapyriformis. The top of the picture is toward the anterior end of the cell; the right side of the picture is toward the left of a kinety. At the left and right are segments of two kineties with basal portions of their cilia (C). It, Longitudinal microtubules below the epiplasmic layer (e); bt, two basal microtubules running along the left side of each kinety. From each kinetosome, three fibrillar systems are starting: the kinetodesmal fiber (Kd) from the anterior right part, the postciliary fibers (Pc) from the posterior right side, and the transverse fibers (T) from the anterior left side. From Allen (1967).

the somatic regions in some ciliates and situated either under the epiplasm or between the epiplasm and the pellicular alveoli, or inside the epiplasm (Grain, 1984). Certain cortical microtubules appear transiently upon division; they are parallel to the division furrow in Stentor (Diener et al., 1983) and Chlumydomonas (Johnson and Porter, 1968), while they are perpendicular to this furrow (i.e., longitudinally oriented) in the ciliates Nassula (Tucker, 197 la), and ParaFic. 18. Ectoplasmic fibrillar systems associated with kinetosomes in ciliates. (a) General case. (b) In Paruisotricha, note the kinetodesmal fibers (Kd) are perpendicular to the kinety (Ci). A lateral right view (c) and a transverse section (d) of the somatic cortex in Ealuntidiurn show the postciliary fibers (Pc), the ribbons of which are superimposed. In Chilodochona (e and 0 , the postciliary fibers each forms groups of three microtubules. M,Dense muff; T, transverse fiber; LT, transverse blade; R, right side of the kinety; L, left side of the cell; A, anterior pole of the kinety. a and b are seen from the inside of the cell. From Grain (1984).

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R C

a

I

M

Pc

e

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FIG. 19. Semischematic representation of the sporozoite apex of Stylocephalus. A, Superimposed apical rings; mp, the peripheral muff; cm, membranal complex; mt, microtubules alternating with intertubular zones (t); me, external membrane. From Desportes (1969).

mecium where they make up the cytospindle, the elements of which are extended in the interkinetal crests along the whole length of the cell (Sundararaman and Hanson, 1976; Cohen et al., 1982); in this case they could be formed by the transfer, toward the cortex, of tubulin molecules from systems which disappear during division (as postoral fibers in Paramecium) (Cohen et al., 1982). The cytospindle also appears in the exconjugants of Paramecium, just after they separate (in Cohen et al., 1982). Helically arranged microtubules in the conoid of sporozoa (Fig. 19) (Desportes, 1969; Porchet-Hennerk and Vivier, 1971; Vivier and Petitprez, 1972) and the microtubules of the ribbons in suctorian tentacles are also of unknown origin. In the reticulopods of Foraminifera, the microtubules have no known MTOC; they are arranged in bundles of varying sizes and can exhibit lateral and axial translocations (Travis et al., 1983). Others are of known origin. The subpellicular microtubules of infective forms of sporozoa originate on a polar ring (MTOC) (Fig. 19) at the base of the conoid (Desportes, 1969; Porchet-Hennerk and Vivier, 1971; Vivier and Petitprez, 1972; Scholtyseck et af.,1972; Russell and Bums, 1983); they probably form the microtubules which are found in gregarine of the Selenidium type (Schrevel, 1971). The microtubule complexes, called axonemes, of the axopods of actinopods have MTOCs which are either grouped together in a single element with a granular texture, the axoplast (central in the centroaxoplastidian Radiolaria, in the centrohelidian Heliozoa, and in the Acantharia; situated in a nuclear depression in the periaxoplastidian Radiolaria), or are smaller and specific to each

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axoneme [each MTOC is a discrete granular mass, lying against the nuclear membrane in the heliozoan Actinophrys (Fig. 20) (Ockleford and Tucker, 1973) and Ciliophry (Davidson, 1982), or a dense mass articulated on a layer of similar material lying against the nuclear membrane in Sticholonche (Fig. 37) (Cachon et al., 1976)]. In these axonemes, the number, length, and position of the links between microtubules condition the actual overall architecture (Fig. 2 1) (Cachon and Cachon, 1974b; Roth et al., 1970; Tilney, 1971). 4. Remarks Low temperatures (Cachon and Cachon, 19801, mechanical shocks, and ultrasounds (Cachon and Cachon, 1981b; Cachon et a l . , 1981) are able to induce the disorganization of microtubules in Sticholonche, that is followed by a reorganization of tubulin, either in twisted filaments or in chevrons (half-disk piled up) (Cachon and Cachon, 1980, 1981b; Cachon et al.. 1981) or in macrotubules or paracrystals (Cachon and Cachon, 1980). Growth of microtubules in vitro from germs occurs at both ends; on the other hand, in vivo, growth occurs at the end opposite the MTOC in the axopods of actinopods (Ockleford and Tucker, 1973; Cachon and Cachon, 1974b) in the nematodesmas of ciliates (Tucker, 197 Ib), in Polytomella (Stearns and Brown, 1981) and in Eimeria (Russell and Burns, 1983); the microtubule disorganization in the axopods upon retraction is also apical. Induction of the growth can be stimulated by taxol in the axopods of Actinophrys (Hausmann et a / ., 1983) and in the kinetosomes of the amoeba of Physarum (Wright et a l . , 1982). One might wonder whether all the structures considered as MTOCs are in fact true organizing centers of microtubule complexes. It is possible that certain MTOCs would only be sites of nucleation of microtubules, as in Polytomella where the MTOCs contain an initiation protein (Steams and Brown, 1981), or in fyrsonympha where the overall arrangement of the axostyle cannot be due to the MTOC, since this arrangement is subsequently established by specific links between microtubules (Bloodgood et al., 1974). Other MTOCs, as that of Polytomella (Steams and Brown, 1981), of the sporozoite of Eimeria (Russell and Bums, 1983), of cells of mammalia in culture (Brinkley et a l . , 19811, and the matrix plates of the nematodesmas in ciliates, seem to contain sites which specify the arrangement of the microtubules; in addition, the MTOC of Pyrsonympha (Bloodgood and Fitzharris, 1979) may well play a role in the motricity of the axostyle. B. PROTISTTUBULINS

Both a- and P-tubulins which unite to form the heterodimers constituting all the microtubules are among the most evolutionarily conserved proteins. However, Dustin wrote, in 1981: “il ne fait pas de doute qu’il existe des variations

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interspbcifiques des tubulines; leur Ctude commence a peine. ” Works on protists already make their contribution to this study, and more reveal an intraspecific microheterogeneity of the tubulins. 1. a- and P-Tubulins Show Some Differences This is the rule in protists as in superior eukaryotes. Their MWs and their electrophoretic mobilities are different, although the electrophoretic separation is not always sufficient to attribute proper MWs to the two subunits (Bordier et al., 1982). lsoelectric focusing reveals that the pl of a-tubulin is more basic than that of P-tubulin. Their peptide maps are different. The variability is higher, from all points of view, for a-tubulin, although P-tubulin, in spite of an apparent uniformity, can present a certain microheterogeneity (Gallo et al., 1983; Crossley and Holberton, 1983). The a- and P-tubulin mRNAs of Tetrahymena have been isolated and purified; they sediment in different ways in the presence of 70% formamide (25 S for a, 16 S for P) (Marcaud et a!., 1979). During the division, or the regeneration of the flagella in Chlamydomonas, the synthesis of 2 atubulin mRNAs and 2 P-tubulin mRNAs proves the existence of 4 distinct genes which work in coordination (Brunke et al., 1982). In Trypanosoma brucei, the majority of the a- and P-tubulin genes are physically linked in pairs, and clustered in long tandem repeats of alteming a- and P-tubulin sequences; this arrangement may facilitate coordinate expression of the subunits comprising functional tubulin heterodirners (Thomashow et al., 1983). 2. Intraspecific Microheterogeneity of the Tubulins in Protists The tubulins in different cell constituants exhibit differences which are identifiable by several methods. a. By Biochemical Analysis. Five varieties of tubulin have been isolated from Chlamydomonas; they differ by their rate of phosphorylation (Dustin, 1981); the amino acid sequences are not exactly the same for the tubulins of the flagellar, subpellicular, and nuclear spindle microtubules (Pipemo and Luck, 1977). Two-dimensional electrophoresis and peptide mapping reveal, in the trypanosomid Crithidia fasciculata, some differences between the aP heterodirners of the tubulins extracted from the flagellum, the subpellicular microtubules, and the unpolymerized cytoplasmic pool (Russell, 1983). Separation by isoelectric focusing reveals several isoforms for the P-tubulin [4 in Trypanosoma brucei (Gallo et al., 1983), 2 in Giardia, and 5 in Polytomella (Crossley and Holberton, 1983)] and for the a-tubulin [4in T. brucei (Gallo et a l . , 1983) and 3 FIG. 20. Axopodia in Actinophrys sol. (a) A large axoneme in cross section. Long links (arrows) extend between microtubules in adjacent rows. (b) With a fixation procedure which does not preserve the links, several C-microtubules (arrows) are present. (c) The base of a large axoneme contacting the nuclear envelope (E).(a) x 110,OOO;(b) ~ 5 2 , 0 0 0(c) ; X 8 1 ,OOO. From Ockleford and Tucker (1973).

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FIG. 21. Microtubular patterns of the axopodial axonemes in (a) Actinophrys (Heliozoa), (b) Acantharia, (c) Sricholonche zuncleu (Radiolaria), and (d) polycystines sphaerellaires centroaxoplastidiata. (a-c) From Cachon era/. (1973); (d) from Cachon and Cachon (1972).

THE CYTOSKELETON IN PROTISTS

20 I

FIG. 21d.

in Giardia (Crossley and Holberton, 1983)]. However, this intraspecific microheterogeneity is more restricted for P-tubulins; in Polytomella, that of axonemes and that of cytoplasmic microtubules have similar electrophoretic mobilities (McKeithan and Rosenbaun, I98 1). In Physarum, the myxamoeba contains microtubule structures during the whole cycle, and tubulins al,a3,and P I are present; on the other hand, the plasmodium forms microtubules only during mitosis, just after the late synthesis (end of G 2 )of tubulins a I ,a 2 ,P I , and P2; the tubulins a3,a2,and P2 are specific to different stages of the cycle; they differ from a ,and p I , respectively, by their PI, but not by their peptide maps (Burland et al., 1983). It is necessary to note that a glycosylated tubulin, different from the microtubule tubulins, would seem to be associated with the ciliary membrane in various eukaryotes (Stephens, I983), and notably in Chlamydomonas and Tetrahymena (Dentler, 1980); however, its existence and its originality are contested for the ciliary membrane of Paramecium (Adoutte et al., 1980). b. By Immunocytochemical Techniques. By preparing monoclonal antibodies against cattle brain tubulin, it has been possible to determine two clones of hybridomes with specific reactions on Trypanosomu brucei, one with flagellum tubulins and the other with subpellicular microtubule tubulins; these

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two types of tubulin therefore differ by at least one antigenic determinant which is implicated in their specialization (Gallo and Anderton, 1983). It is necessary to investigate the causes of this intraspecific microheterogeneity. While the a-tubulin of vertebrates may be more or less modified, after in vivo translation, by a natural amino acylation that gives the heterogeneity of the tubulins in the cell, no tyrosylation has ever been noted in invertebrates and protists (in Dustin, 1981). Moreover, the number of repeats of the a-p pairs of genes in T. brucei is consistent with the high level of tubulin gene expression and with the presence of distinct classes of microtubules (Thomashow et af., 1983). However, in Pofytomeffa,two tubulins have been separated: the al-tubulin predominant in the cytoplasmic microtubules, which is comparable to brain atubulin, and the a,-tubulin predominant in the flagella (McKeithan and Rosenbaum, 1981); it would be conceivable that these tubulins are coded for by different genes, but, when their corresponding poly(A) mRNAs are isolated and then translated in vitro, only a,-tubulin is obtained (McKeithan et al., 1983). In addition, the two genes of a-tubulin found in Chfamydomonas(Brunke et al., 1982) both code for the same a,-tubulin (identical to that of Pofyromella); apparently then, the a,-tubulin is formed by posttranslational conversion of a,tubulin; furthermore, this conversion would be reversible. Its nature remains unclear; it probably does not involve phosphorylation (McKeithan et al., 1983). It is the same problem for the a,-tubulin of Physarum which seems specific to flagella and for which it is impossible to assert that it results from the modification of the al-tubulin (Burland et al., 1983); nevertheless, during the cell cycle, a2-and P,-tubulins, present only in the plasmodium, are probably coded for by specific genes, which involves the existence of 4 genes in Physarum ( a Ia,, , PI, P,) (Burland et al., 1983); however, it is impossible to rule out the hypothesis that only 2 genes exist (one a and one p), and that a variable evolution of the corresponding mRNAs can lead to a differential expression (aI, a2or a3,and p, or P2) of these 2 genes. Finally, one might wonder about the meaning of this intraspecific microheterogeneity. In Physarum, the myxamoeba which has mitotic spindle, cytoplasmic, centriolar, and flagellar microtubules, possesses 3 tubulins- a,,a3, and PI- while the plasmodium, which forms only mitotic spindle microtubules during late G , possesses 4 tubulins- a,,a2, P I , and p2. This would seem to indicate that it is not necessary to have as many different tubulins as there are different functional types of microtubules in the amoebae, and that several types of tubulins can constitute a single type of microtubule in the plasmodium. It is also important to note that a single electrophoretic species may consist of several polypeptide species. Therefore, it is difficult at present to ascertain “whether multiple a- and P-tubulin reflect functional specializations, or whether individual tubulin subunits are involved in multiple functions” (Burland et al., +

1983).

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203

3 . Interspecific Variability of the Tubulins This variability occurs in the protist phylum as well as between protist and metazoan species. a. In Protists. P-Tubulins shown few or no interspecific variations; those of Physarum (Chang et al., 1983), Tetrahymena (Roobol et al., 1980), Leishmania (Bordier et al., 1982), and Chlamydomonas (Fliss and Suyama, 1979; Little et al., 1981) have the same electrophoretic mobility as those of vertebrates; this similarity also occurs in the peptide maps, those of the P-tubulins of Physarum (Clayton et al., 1980; Little et al., 1982; Chang et al., 1983), Chlamydomonas (Little et al., 1981), and Leishmania (Bordier et a l . , 1982) (Fig. 16) being very similar to those of mammal brains or of sperm flagellae of various invertebrates. If their MWs sometimes appear different, their estimation is doubtful when these differences are not shown on the same gel, during simultaneous electrophoresis. On the other hand, a-tubulins present a certain diversity. While the electrophoretic mobility of the a-tubulins of Physarum (Clayton et al., 1980; Roobol etal., 1980; Chang et al., 1983; Adoutte et al., 1984), Paramecium (Adoutte et al., 1982), Tetrahymena (Adoutte et al., 1984), and Trypanosoma brucei (Gallo et al., 1983) is higher than that of the P-tubulins of vertebrates, creating an a to P inversion of these protists as regards vertebrates, that of the a-tubulins of Chlamydomonas (Fliss and Suyama, 1979; Little et al., 1981) and Leishmania (Bordier et al., 1982) remains lower than that of P-tubulins of vertebrates, giving no inversion. However, it is necessary to consider the a to p inversion with prudence, since, in Chlamydomonas, it may or may not occur, according to the origin of SDS used in electrophoresis (Remillard and Witman, 1982). Valid comparisons between species must be made in simultaneous electrophoresis, on the same gel. The peptide maps of the a-tubulins of Physarum (Clayton et al., 1980; Little et al., 1982; Chang et al., 1983), Leishmania (Bordier et al., 1982) (Fig. 22), Chlamydomonas (Little et al., 1981, 1982), and Terrahymena (Little et al., 1982) are very different from those of metazoans. Although comparisons between protist a-tubulins are difficult to establish from data dispersed in the literature, differences between species of protists would likely to be as high as those found between species of metazoans (Little et al., 1981); nevertheless, a certain similarity exists between the a-tubulins of the cilia of Tetrahymena, the flagella of Chlamydomonas and Pteridium (Little et al., 1981), and the cytoplasmic microtubules of Physarum (Little et al., 1982). Antisera against tubulins of Paramecium and Tetrahymena show identical behavior when tested against various species of protists; they positively and strongly react with some species of protists (not exactly the same for the two antisera), but never with the tubulins of vertebrates; when applied to the peptide maps of a-and P-tubulins of Paramecium, they react against a single a-peptide and a single P-peptide, i.e., against only one antigenic determinant in each of the

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a

FIG. 22. Peptide maps of Leishmania tropica (L) and pig (P) tubulin subunits (aand p). Drawn after Bordier er a / . (1982).

two proteins; the absence of reaction of the antitubulin of Paramecium against certain groups of protists (trypanosomids, myxomycetes) indicates a loss of the reactive determinants, and gives a rough indication of the degree of phylogenetic divergence (Adoutte et a/., 1984). Thus, the application of such narrow-spectrum antibodies becomes a tool for investigating evolutionary relationships. Protist tubulins, contrary to those of vertebrates, are good phylogenetic markers. Furthermore, it is possible to establish relationships between species by other methods; the translation of poly(A) mRNAs of Tetrahymena gives, among other things, a tubulin that is similar to the a-tubulin of Chlamydomonas (Myhal +

THE CYTOSKELETON IN PROTISTS

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and Hufnagel, 1979), which perhaps indicates a common origin for the two genus, and then a (posttranscriptional?) differentiation of the two tubulins. b. Between Protists andMetazoans. The tubulins of Physarum and Tetruhymenu have a weak affinity for colchicine, unlike that of brain tubulin which fixes colchicine 50 times more than that of Physarum and 20 times more than that of Tetrahymena (Roobol et al., 1980). The tubulin of Giardia contains less serine, proline, threonine, and more leucine that that of rat brain (Crossley and Holberton, 1983). While the electrophoretic mobility of P-tubulin is identical in protists and in vertebrates (Little et a l . , 198 1 ; Adoutte et a l . , 1982; Bordier et al., 1982), that of a-tubulin of various protists is higher than that of the two tubulins of vertebrates, creating an a to P inversion with regard to vertebrates; it is difficult at present to ascertain whether it is a characteristic specific to protists (with some exceptions) (Adoutte et a l . , 1984). It should be remembered that posttranslational transformations of tubulins in protists are unknown, while some have been identified in vertebrates (in Dustin, 1981). The numbers of isoforms are different for vertebrate tubulins [ 10 for a,4 or 5 for P-tubulin of pig brain (Gallo et al., 1983)j and for protists (see Section V,B,2). Peptide maps of the P-tubulins of Physarum (Chang et al., 1983) and Leishmania (Bordier et a f . , 1982) show some similarities with that of pig brain atubulin (Fig. 16);that of P-tubulin of Chlamydomonus is very close to that of atubulins of mammalians, molluscans, and echinids (Little et al., 1981). On the other hand, for the a-tubulins, peptide maps are very different between Physarum (Chang et al., 1983), Leishmania (Bordier et al., 1982), and pig brain (Fig. 22); that of a-tubulin of the axonemes of Chlamydomonas differs from that of invertebrates and mammals, but closely resembles that of the sperm flagellum of the fern Pteridium (Little et a f . , 1981), which would indicate a possible phylogenetic relationship. Antibodies against the tubulins of Paramecium and Tetrahymena, which react with the tubulins of some protists, never react with vertebrate tubulins; on the other hand, antibodies against tubulins of turkey or pig react with the tubulins of all the tested protists and vertebrates (Adoutte et a l . , 1982, 1984); behavioral differences between these antibodies evidently result from differences between the tubulins of vertebrates and of protists. According to Adoutte et al. (1984), since vertebrate tubulins have a high degree of conservation of their amino acid sequence, those of a given vertebrate species are not recognized as foreign proteins by rabbit or sheep, and antibodies, only obtained with difficulty by hyperimmunization, can be considered as autoantibodies not directed against stranger determinants specific of the given species, but rather directed against determinants which are present in the tubulins of many species, hence a broad spectrum reaction of these antibodies; on the other hand, the tubulins of protists, which have sufficient differences with those of vertebrates (MW, electrophoretic mobilities), are recognized as “foreign” proteins by rabbit and sheep, hence the

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production of specific antibodies which react only against the tubulins that possess the same determinants as the tubulins injected in the mammal, and the narrowness of the reaction spectrum. Peptide maps of the P-tubulins of the axonemes of Chfamydomonas and Terrahymena and of the cytoplasmic microtubules of Physarum and Leishmania (Fig. 22) are similar to those of cytoplasmic tubulins of vertebrates (Little et al., 1981; Bordier et al., 1982; Chang et al., 1983), and to those of axonemal tubulins of invertebrates and plants (Pteridium) (Little et al., 1981, 1982). On the other hand, peptide maps of a-tubulins of the protists are very different from those of vertebrates (Little er al., 1981; Bordier et af., 1982; Chang et af., 1983) and invertebrates (Adoutte et al., 1982) (Fig. 22), while they are very similar to those of the sperm flagellum of the fern Preridium (Little et af., 1981, 1982). 4. Remarks All these similarities and differences have led Little et af. (1981, 1982) to consider the evolution of tubulins in the following way: from an ancestral single gene, a first phylogenetic divergence separated the 2 tubulins subunits. During the evolution of the lower eukaryotes, only one type of tubulin was used to build cytoplasmic and axonemal microtubules, hence the resemblances between the tubulins of algae, plants, ciliates, and myxomycetes. Then, during the evolution of metazoans, the duplication and the modifications of the tubulin gene led to the use of one type of tubulin for the axonemes (with few further mutations or changes), and of another type for cytoplasmic microtubules (with fast mutations and a large diversification). Although this hypothesis is tempting, it seems premature to construct such evolutionary models for the tubulins, since there are not enough data at present, and there is still uncertainty about the diversity of tubulins (are they the results of the expression of different genes, or of postranslational modifications?). Furthermore, crossing immunological reactions performed by Adoutte er al. (1984) show that, even in the phylum protists, the unity of tubulins is not the rule; some antigenic determinants, recognized by the antibodies obtained against tubulins of ciliates, are also present in dinoflagellates, so they are very old; those determinants would be progressively lost upon phylogenetic divergences leading to euglenians and chloromonadina on the one hand, and trypanosomids on the other; in the latter case, differences already exist between cytoplasmic and axonemal tubulins (Gallo and Anderton, 1983; Russell, 1983). c . MICROTUBULE-ASSOCIATED PROTEINS (MAPS) AND THE ROLES OF MICROTUBULE STRUCTURES As yet, little is known about the MAPS in protists, except for those localized in cilia and flagella.

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1. MAPs of Structures Other Than the Axonemes of Cilia and Flagella

The MAPs associated with the cytoskeletal microtubules of the myxamoebae of Physarum are of low MW: 49K (main one), 59K, and 57K; their presence induces the formation of microtubules, but they are not in large amounts (Roobol et al., 1980); no MAPs of high MW of the neuro-MAP type have been found, as in other different nonneuronal structures which also have a 49K MAP (Roobol et al., 1980). In the adhesive disk of Giurdiu, the microtubules bear lateral lamellae (Fig. 23a) composed of two layers of globular tubulin subunits (Fig. 23b), giving parallel protofilaments that are perpendicular to the longitudinal axis of the microtubule (Holberton, 1981; Holberton and Ward, 1981); between these two layers is a substance composed of a protein, giardin, which is in fact a doublet of 3 1 and 30K; its strong interaction with tubulin enables it to link and maintain the alignment of the tubulin dimers of the two outer layers (Crossley and Holberton, 1983). Other proteins, present in the isolated disk (one with a MW > 120K), have not been localized in the structures. The lamellae and microtubules of the disk constitute a noncontractile whole which probably plays a role in the maintenance of the form. Certain flagellate axostyles are animated by often violent bending movements; these are formed from several rows of microtubules and possess interrow linkages; these latter must play some role in the movement of the axostyle; in Saccinobaculus (McIntosh et a l . , 1973) the bending of the axostyle, which progresses anteroposteriorly, takes place through sliding of rows of microtubules over each other (McIntosh, 1973) caused by rearrangement of the temporary interrow linkages; in the resting stage, the link is attached to two microtubules from neighboring rows and bonded to a Mg2+-ATP complex (a difference compared with skeletal muscles). The hydrolysis of ATP provokes the rotation of the link, hence a displacement of the two microtubules over each other, and immediately after that the detachment of one extremity, with Mg2 , ADP, and Pi being liberated. The new fixation of Mg2 -ATP leads to the relaxation of the link and its refixation on the following microtubule site (Bloodgood, 1975). Given that one protein with ATPase activity of the dynein-type (with only one enzyme subunit of MW 500K, but without any structural subunit) and another protein of MW 80K similiar to nexin have been found (Mooseker and Tilney, 1973), it might well be that MAPs are involved in the mechanism of movement. We should no longer speak of contractile microtubules, as has often been done in the past. +

+

2 . MAPs of Cilia and Flagella Figure 24 summarizes recent data concerning the identification of these MAPs, collected from works on Chlamydomonas and Tetrahymena (Witman et al., 1978; Gibbons, 1981; Huang et al., 1981; Piperno el al., 1981; Dutcher e f

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al., 1984). In the flagellum, the elastic nexin links normally limit the displacement of the doublets in relation to each other, and so moderate the action of the dynein arms; the radial spokes, which contain 17 proteins, 5 of which are phosphorylated [only 4 structure genes and 1 regulating gene have been detected (Huang et al., 198111, act, with the central complex (microtubules + sheath or projections), to transform the sliding of the doublets into a bending of the flagellum; the heads are detached from the sheath in the straight parts, and attached to it in the curved parts of the flagellum (Witman et al., 1978). The outer dynein arms are composed of five parts, the relative motion of which enables either a rigid state or a relaxed loose state to be had, which can explain their role in the sliding of neighboring doublets (Goodenough and Hauser, 1982). A tubulin of MW 53K has been identified in the ciliary membrane of Tetrahymena (Williams et al., 1980). We can include in the MAPs, albeit artificially, the proteins which compose the paraflagellar rod of the euglenians and trypanosomids (Fig. 25); in trypanosomids, it has a paracrystalline aspect, while, in Euglena, it is composed of a layer of 22-nm-diameter filaments, which is twisted in a helix (Hyams, 1982), and it is linked to at least one of the axoneme doublets. In Crithidia, the rod, which is small, contains a single major protein of MW 71K; in Herpetomonus, which has a larger rod, two proteins are found (MW 71 and 65K) (Walkosz, 1982); this is also the case for Euglena (MW 80 and 69K) (Hyams, 1982). A certain degree of ATPase activity has been detected at the level of the rod (Piccini et al., 1975), suggesting that it participates in the movement of the flagellum. Furthermore, the rod should control the frequency and the importance of the beating wave (Hyams, 1982).

3 . MAPs in Other Microtubular Systems In all the other microtubular systems, the MAPs are not characterized, though they must certainly be present. Some of them must, in particular, constitute, in whole or in part, the links between microtubules, between microtubules and microfilaments, between microtubules and membranes, and perhaps between kinetosomes in the polykinetid groups, like pairs, membranelles, cirri, etc., in ciliates. 4. A Number of Roles May be Ascribed to the Microtubular Systems 1. A role in the maintenance of cell shape: a priori, it might be assumed that all the subpellicular microtubules have this role. This is confirmed by certain FIG. 23. Sucking disk of Giurdia. (a) Vertical section through a trophozoite showing microtubules (mt) and microribbons (mr). The disk is strongly domed and microribbons are parallel. (b) Perspective drawing of the subunit arrangement in microribbon faces, showing the discontinuous seam with a disk microtubule. Ribbon core structures are drawn crudely from their appearance in transverse sections. (a) X71,200. (a) From Holberton and Ward (1981); (b) from Holberton (1981).

3 protofilaments ( A d ) 4 proteins(UW
+

1 actin-like protein

Radialspoke N I 0

8:

56 K

84 K

84 K 106 K 128 K 142 K Z50 K

:

14 K

:

K

:

25 50 45 62 97

K K

K K

20 K 66 K 97 K

:

5 7 K Str 110 K 128 K

32 K

:

220 K

:

270 K 360 K

:

in .n CTZ

CTl+CT2 in CT1 Central tubules sheet complex

FIG. 24. h t e i n s found in cilia and flagella.

FIG. 25. Paraflagellar rods (PR) in the euglenian Anisonema costarum (a, b) and in the kinetoplastidia Herperomonas muscarum (c, d). (a, c) Longitudinal sections; (b, d) transverse or 21 1

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observations: in Colpodu (ciliate) subjected to high pressure, the ribbons of the cortical arrays separate from each other (each ribbon remaining intact), bringing about a change in cell shape. At the level of the somatic furrow, this disorganization also reaches the interior of the ribbons and the deep microtubules (nemadesmata); hence, the furrow disappears (Lynn and Zimmerman, 1981). Normally, the rigidity of the microtubule systems is ensured by links between the microtubules; these links are stronger within a ribbon than between two neighboring ribbons. 2. A role of an internal supporting structure: the deep microtubules originating at the kinetosomes and spreading over or around the nucleus in various flagellates must help keep these structures properly positioned in relation to each other; this kinetosome-nucleus junction allows the maintenance of the cohesion of the rotary axis in Devescovinidae (Tamm, 1978), the axostyle rigidity being ensured by links between the microtubules (a single twisted row). The ribbons of microtubules of the sucking tentacles of the suctorians are, during ingestion, shifted away from each other, liberating a large central space (Fig. 26); this movement seems to be provoked only by microfilaments (Hauser and Van Eys, 1976); the microtubule ribbons play, in fact, the role of a solid axis in the tentacle; they are also passive during the displacement of the trophic membrane toward the base of the tentacle; they are not involved with the contractility which results in the tentacle retraction (Hackney and Butler, 1981). In the radiolarian axopods, arms (MAPS?) which are linked to microtubules have been detected; some of them link microtubules together, others have a free end; they were attributed the role of cytoplasm and various organite carriers (Cachon and Cachon, 1975). In the light of experiments and observations on the axopods of Heliozoa, it appears that the provoked absence of microtubules does not prevent cyclosis (Cachon and Cachon, 1974b); the microtubules should act only as a more or less rigid axis for the axopod, and might possibly support actin microfilaments involved in displacement of cytoplasm or organites (Bardele, 1976); the whole structure should be slightly elastic (Ockleford and Tucker, 1973). The apical destruction or the apical extension of the microtubules would be the consequence, but not the cause, of the shortening or lengthening of the axopod (Ockleford and Tucker, 1973). In the oral-ribs of the mouth of Tetruhymenu (Fig. 6), the microtubules originating at the paroral (right ciliary buccal row) establish, with the overlying membrane, links which show up as alignments of intramembrane particles (Sattler and Staehelin, 1979); they strengthen the naked wall of the buccal cavity and give rigidity to the crests which guide the food particles toward the cytostome. In oblique sections. M , Mastigonemes;Ax, axoneme; A, anterior flagellum; R , recurrent flagellum. (a) X5O,ooO, (b) X 40,000; (c) X57,MM; (d) X57,OOO. (a, b) From Mignot; (c, d) from Brugerolle er al. (1973).

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FIG. 26. Heliophryu (Suctoria). Cross sections through the mid-region of a contracted (a) and of an expanded (b) inactive tentacle shaft. (c) Base of a contracted tentacle in the cytoplasm. (d) Base of an expanded tentacle in the cytoplasm. cb. Cross-bridges in the outer circle; cob, connective bridges; fa, free arms; irb, interrow bridges; mtc, microtubule circle; mtr, microtubule rows; og, osmiophilic granules. x65.000. From Hauser and van Eys (1976).

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Colpoda, the destruction of the subepiplasmic microtubules in the right vestibule lip induces the eversion of the lip (Lynn and Zimmerman, 1981). The microtubules of the nematodesmal lamellae of P seudomicrothorax act as a rigid support, bearing arms on which are fixed microfilaments (though the arms may themselves be microfilaments); these microfilaments are connected to be membrane of the digestive vacuoles and displace them (Hauser et al., 1980); this is very probably also the case for most of the cytopharyngeal ribbons in ciliates. The nematodesmas of the "nasse" of Nassula (Tucker, 1968) and of Chifodonella (Soltynska, 1971) are spread widely at the top during ingestion, and afterward, the microfilaments cause them to become more closely packed, so that they form a rigid support structure, with passive movements. 3. A double role: both shape maintenance and cell motility. The destruction of the subpellicular microtubules in Selenidium (Gregarinia) by colchicine not only leads to the deformation of the epicytic folds (support role), but also to the termination of locomotion (Stebbings et al. 1974); it is possible that these microtubules are involved in the bending of the folds, by gliding over each other (Mellor and Stebbings, 1980). Is their role active or passive? The absence of microtubules in the folds of Gregarina, which are concerned with ondulatory movements (Schrevel, 19711, favor the second hypothesis; the origin of contractility would be elsewhere. In the sporozoite of Eimeria, longitudinal microtubules are linked to the plasma membrane at the site of double rows of intramembranal particles, which could help the gliding of microtubules during the extension and retraction of the anterior part of the cell when it enters the host (Dubremetz and Torpier, 1978). In the foraminiferida Allogromia, the reticulopodial microtubules are assembled in bundles that can exhibit axial and lateral translocations, the latter being accompanied by lateral association or dissociation between adjacent bundles; these movements determine the shape, extension, or shortening of the filopodia (or lamellipodia); in addition, the microtubules are actively involved in the bidirectional cytoplasmic streamings in the reticulopodia and filopodia, i .e., in the transport process of cytoplasmic particles, of large hyaline bulks of cytoplasm (Travis et al., 1983), and of extracellular particles adhering to the external side of the plasma membrane (Bowser and McGee-Russell, 1982). Links have been observed between the microtubules and the intracytoplasmic particles (Travis and Allen, 1981) and between the microtubules and the plasma membrane (Bowser et al., 1983). Finally, a dynein-ATPase inhibitor causes the transport of intracellular and extracellular particles to cease, without any destruction of the microtubules; thus, a dynein-like ATPase is implied in bidirectional transport (Bowser et a f . , 1983). The intractyoplasmic transport of a whole cytoplasmic domain involves motility-organizing vesicles (MOVs) which are closely associated with microtubular bundles, and possess cross-connections with the organelles of the cytoplasmic domain, and with the plasma membrane (McGee-Russell et al., 1982).

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In Distigma, displacement by changes in cell shape (euglenoid movement) is probably due to the links between the subpellicular microtubules and either the plasma membrane or the submembranal dense layer (epiplasm?); these links might work like the dynein arms of the cilia, giving a gliding of the microtubules with respect to the pellicle, but their nature is unknown (Gallo and Schrevel, 1982). In Astasia, the body contraction would involve an ATPase activity which is located at the level of the microtubules and beneath the cell membrane and regulated by Ca2+ (Murray, 1981). In Stentor (Fig. 27a), the contraction of the myonemes (see later), which results in the shortening of the cell, causes the passive sliding of the postciliary microtubule ribbons over each other, with rupture of the desmoses between contiguous ribbons; on relaxation, the myonemes are inactive; the desmoses between neighboring ribbons are reestablished and, by a mechanism which may be identical to that of the dynein arms, cause active sliding of the ribbons in the opposite direction, leading to an elongation of the cell (Huang and Pitelka, 1973). Furthermore, in Spirostomum, in which the structural elements are identical to that of Stentor, the relaxation is inhibited by ATPase inhibitors and by colchicine, which supports the hypothesis of an active role of the microtubules and the possible ATPase activity of the desmoses in the sliding of the ribbons (Holberton and Ogle, 1975). 4. Role in the guiding and positioning of organelles: the involvement of microtubules in the positioning of kinetosomes during morphogenesis, either on the whole cell surface (Allen, 1969), or in the buccal primordium of Tetrahymenu (Williams and Frankel, 1973) or in the somatic ventral primordia in Paraurostyla (Jerka-Dziadosz, 1980), has been envisaged. They may also be involved in the positioning of nuclei in Paramecium (Beisson and Rossignol, 1975; Cohen et al., 1982), and of cortical mitochondria in Tetrahymena thermophila (Aufderheide, 1983). In Paramecium, microtubules are concerned with the transport and guiding of trichocysts toward the cortex (Beisson and Rossignol , 1975; Plattner et al., 1982); for example, in P . tetraurelia, the contact between the free deep end of one (or two) microtubule, the other end of which is attached to a kinetosome, and an endoplasmic trichocyst would be a signal for the exiting of the trichocyst from the endoplasmic cyclotic streaming; then the microtubule would guide the trichocyst onto its attachment site on the plasma membrane (Plattner et al., 1982). 5 . Role in cell division: the cortical longitudinal microtubules which appear during the division of Paramecium may play several roles: (1) in the elongation of the cell (by transporting and guiding membranal material), (2) in the maintenance of the shape (true cytoskeleton) for the whole cortex of the cell [which has become more fragile, during the elongation, when the successive kinetodesmal fibers do not overlap (Cohen et al., 1982)], and for the region of the division furrow (where the infraciliary lattice (see below) disappears (Sundararaman and Hanson, 19i6), and (3) during cytokinesis (again by guiding new membranal

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material) (Sundararaman and Hanson, 1976). A correct elongation of the macronucleus in dividing Paramecium occurs only if the intranuclear microtubules are sufficiently numerous and have a normal peripheral position (Cohen et al., 1980). In Chlamydomonas, the microtubules which appear in the future division plane could, together, help the membrane to invaginate, and maintain the rigidity of the edges of the forming furrow (Johnson and Porter, 1968). 6. Finally, the frequent presence of subpellicular microtubules in protists could suggest relationships between microtubules and membrane receptors, either direct or via other proteins such as those of the epiplasm, for instance. [Such relationships between microtubules and (hormonal) receptors have been shown in Rana erythrocytes (Cherksey ef al., 1980).]

VI. Periodic Fibers These fibers are abundant in protists, where they are essentially linked to kinetosomes. Three types can be distinguished, according to their position. A. KINETOSOMALPERIODIC ROOTLETS They run into the endoplasm and they have variable structures, not always well elucidated. 1. Some are composed of tightly arranged fine longitudinal filaments, bearing regularly arranged thickened zones which results in a periodic aspect of the whole; major transversal bands give the major period; the number of subperiods is variable. In general, periodic rootlets of this type have, in protists, a low major period (10 to 50 nm) unlike those of metazoans which have a larger period (50 to 100 nm) (Dingle and Larson, 1981). They are fairly rare in ciliates; they help consolidate the adhesive basal disk of trichodines (Fig. 28) (Favard et al., 1963; Maslin-Liny, 1983) or cross over the body in Loxophyllurn (Fig. 29) (Rodrigues de Santa Rosa, 1974). They are more widespread in flagellates; they are either roughly cylindrical or blade-shaped [Chilomonas (Fig. 15) (Mignot et al., 1968), Trepomonas (Brugerolle et al., 1973)l; they form the rhizoplasts, or the parabasal fibers. In Naegleria, the periodic rootlet which appears at the same time as the kinetosome (Larson and ~

FIG. 27. Cortical structures in heterotrichs. In Slenror (a), a transverse section of the cortex shows a somatic pair of kinetosomes (Ks), the postciliary ribbons or km-fiber (Pc), and a myoneme (M) surrounded by a vacuolar reticulum (V). In Condylostoma. a tangential section (b) shows the relative positions of km-fibers (Pc) and myonemes (M).(c) Detail of a myoneme of Condylosloma (longitudinal view). (a) X28.500; (b) X8.100; (c) X20,oOO. (a) From Bernard (1980); (b,c) from Bohatier (1978).

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FIG. 28. Periodic ciliary rootlet in Trichodina nigra. (a) General view. (b) Detail of the rootlet. (a) X18,oOO. (b) X35,OOO. (a) From Maslin-Uny (1983); (b) from Maslin-Uny and Bohatier ( 1984).

Dingle, 1981b) is a juxtaposition of filaments 5 nm in diameter; it contains a major protein of MW 170K (Larson and Dingle, 1981a), unlike those of the branchial cilia of Aequipecten which have two main proteins (ankyrin: 230 and 250K). The rhizoplasrs of Tetruselmis (Chlorophyceae) which are attached at

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FIG. 29. Loxophyllum meleagris. (a) A large periodic fiber (CF)runs along the kinetosomal periodic rootlet (RC). (b) Detail of these two fibers. (a) X42.000; (b) X56.000. From Grain (1984).

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their deep end, one to the nuclear membrane, the other to the plasma membrane, are contractile (induction by Ca2+ and other cations) (Robenek and Melkonian, 1979); but their structure is peculiar, as the major period is anomalously long (400 to 600 nm); this type of contractile rootlet is mainly found in unicellular algae (Dingle and Larson, 1981). 2. The costa of trichomonadina is composed of stacked disks each bearing parallel lamellae which cross them (Fig. 30b). It is located in the endoplasm, beneath the recurrent flagellum and the “undulating membrane” (Fig. 30a). It undergoes an undulatory movement which is propagated from front to rear, which causes a deformation and a change of the direction of cell displacement (Mattern and Honigberg, 1971); the curving is due to a modification of the angle of the lamellae in relation to the disk carrying them, giving a “zig-zag” appearance to neighboring disks; in the straight parts, the lamellae stay perpen-

Q

rnernbrane-

a

UfiWood

particles

w

FIG. 30. (a) Diagram of a trichomonad flagellate (Trichomonas termopsidis). (b) Longitudinal section (but not median) of the costa of Trichomonas gigantea showing the transverse plates (P)and their lamellae (L). Bar, 250 nm. From Amos et al. (1979).

FIG. 30B.

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dicular to the disk (Amos et al., 1979). In Trichomonas thermopsidis the costa contains a major protein (MW 90K) and of protein of higher MW (300K) in smaller amount; it contains no actin, nor any myosin, spastin, tubulin, or ankyrin; it shows an ATPase activity in the presence of divalent cations; however, ATP does not reactivate isolated costa, though a thermostable ATPase is present (Amos et al., 1979); so, ATP is doubtlessly not the natural source of energy. B. CONNECTIONS BETWEEN KINETOSOMES These are structures connecting the proximal ends of the kinetosomes; they are generally made of fine filaments, bearing irregularly spaced transverse dense striations. In Chlamydomonas, the striated link between the two kinetosomes, normally at 90" to each other, contract in the presence of Ca2+ , reducing this angle (Hyams and Borisy, 1975); mutants devoid of this link have an unusual orientation of their kinetosomes and of the beating wave of the flagella, and show anomalous repartition of the kinetosomes on division; unlike the anterior idea, it seems that this connection has no role in the anchoring of the kinetosomes, nor in the absorption of mechanical stress due to the beating of the flagella, in the realization of the conjugation, in the maintenance of the flagellar length equality (Wright et al., 1983). In ciliates, similar connections have been found; in a few cases they are well developed, as in Lacrymaria (Fig. 31) (Bohatier, 1972), but their role is unknown.

FIG.31. Periodic connections between kinetosomes in Lacrymaria. (1972).

X35.000. From Bohatier

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223

FIG.32. Schematic drawing of a kinetid in Tetrahyrnena. Identification of the cortical proteins from data obtained by Vaudaux (1976) and by Vaudaux et al. (1977).

C. THEKINETODESMAL FIBEROF CILIATES This fiber arises from the dense material of the muff which surrounds the proximal part (base) of the kinetosome, on its anterior right side near triplets 58; generally it extends anteriad toward .the pellicular surface, in the ectoplasm, and on the right side of the kinety (Figs. 17 and 32). Its main period varies between 16 to 50 nm. It is apparently made up of longitudinal filaments (Hufnagel, 1969). It can extend to a varying degree, can sometimes have an anomalous orientation, and can participate in the construction of skeletal rods in Asromaru and Hysterscinetidae (Grain, 1984). In the isolated pellicles of Terrahyrnenu, it often remains tightly fixed to the kinetosome, but in some cases, it is dissociated from the kinetosome even where kinetosome distribution is maintained in the ghost; that means it does not have a role in the maintenance of the shape of the cortex in this ciliate (Collins et af., 1980); however, such a role cannot be denied when the kinetodesmal fiber is involved in skeletal constructions.

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It may have some degree of contractility (in Collins et al., 1980), as well as being able to stabilize the kinetosome during beating of the cilium. The constitutive protein has a high MW (250K) in various species and strains of Tetrahymena (Vaudaux, 1976; Vaudaux er al., 1977) (Fig. 33), comparable to that of the ankyrin of mollusks. It would be interesting to further investigate this comparison, and to determine whether there is any interspecific variability within the ciliates. It should be added that, compared with the microtubules and microfilaments, the periodic fibers of protists have been somewhat neglected in biochemical studies, as have the other categories of skeletal elements that will now be examined.

VII. Intermediate Filaments These are well known and have been thoroughly studied in Metazoa. They have an average diameter of 10 nm (7 to 1 1 nm), and are found in various cellular types. The constituent proteins differ according to the origin of the cell, and their MW varies between 50K and 70K, except for the neurofilin, the subunits of which have high MW (160 to 210K); these proteins also present an immunological diversity (Lieska et al., 1980; Yen et al., 1980; Cabral et al., 198 1; Milstone and McGuire, 1981); they comprise five classes. Nevertheless, all intermediate filaments have common properties: size, morphology, identical proportions of proteins containing a-helical segments and giving similar X-ray diffraction pictures, homologous amino acid sequences, and molecules comprised of two parts, one constant (a-helical, with similar amino acid sequences) and the other variable (with various amino acid sequences) which is responsible for the differences in solubility, and for the immunological and functional diversity (Milstone and McGuire, 1981). In neurons, the intermediate filaments (or neurofilaments) are involved in the axonal transport due to their interactions with the microtubules (Leterrier et al., 1981; Runge et al., 1981). In the other types of cells, the intermediate filaments make up a tridimensional reticulum, often associated with actin microfilaments or with microtubules, and they can also constitute a peripheral nuclear mesh (Henderson and Weber, 1980). Their role is not clear; they could control the positioning of the nucleus (Blose, 1979), but they do not seem to play a role in the maintenance of the cell shape or in motility (de Brabander, 1983). The intermediate filaments are, in contrast to microtubules and actin microfilaments, extremely stable; they are not dissociated by nonionic detergents (Triton X-100) or by high ionic strength solutions. In protists, intermediate filaments are rare, or not well known; there is a great number of filamentous structures, specially in ciliates, the nature, size, and

T.austraZis AU50.1;11(11) T.pyfifo&s GL-C;A T.vorax Tur

T.boreaZis lJM665;3(III) T.caMdensis SMI I;7(III) T.pigmentosa IL3;8(I) T. tropicazis TC105;9(I) T. furgasoni GL-ATCC;C

T.mericrmis Hom3;2(IX) T.eZZiotti GL.Eichel;B T-boreaZis SLA;5(I) T.vorax VzS T.pigmentosa UM1091;6(II) T . h y p e m n g u M s EN1 12;

T . t h e m o p h i k WH6; I

T.thenophila WH14; I T.thenophiZa B ; I T.themophila D ;I

IO(1)

T ' . C U ~ ~ ~ Aul15-3; CO~~S 12(IV) T.patula L-FF

FIG. 33. Diagram of the five different patterns (I-V) of high-molecular-weight structural cortical proteins found among the strains and species of Tetrahyrnena examined (listed under each type with species name, strain designation, syngen/phenoset). After Vaudaux et al. (1977), modified.

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FIG.34. Cell surface of Gregarina rhyparobiae. Note the presence of apical filaments (0 at the top of the epicyte folds. p.m, Plasma membrane; c.m, the two inner cytomembranes. Bar, 0.3 pm. From Philippe er al. (1982).

stability characteristics of which are not given, while their arrangement is perfectly described (see Sections IX and X). Some cases have however been reported. In Gregarina blaberae. the internal filaments, situated beneath the internal membrane of the epicytic folds (Fig. 34), are intermediate filaments of diameter 12 nm (Schrevel et al., 1983); though their diameter is estimated at 7 nm in Gregarina garnhami (Walker et al., 1979), the absence of actin in the pellicle of various species (Mackenzie, 1980; Philippe et al., 1982) proves that they are not actin microfilaments. A protein common to three species (MW 52K) may be that of the internal filaments (Philippe and Schrevel, 1982; Philippe et al., 1982). These may be involved in locomotion by sliding; they may maintain the folds by imposing a rectilinear direction on the sliding; the motive force for this would originate in the relationships established between the mid and outer (plasmic) membrane by the rippled-dense structures [sometimes called “outer filaments” (Walker et al., 1979)l; the internal filaments are, moreover, attached in some sites to the internal membrane of the fold, which allows the sliding of the “internal plus mid membranes” complex along the internal filaments only out-

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side of these anchorage points; this sliding, which in transmitted toward the surface by means of the “external filaments,” causes swellings (varicosities) at short distances from each other; these swellings press on the substrate, then interact with the excreted mucus to provoke displacement (Mackenzie and Walker, 1983). The intermediate filaments might also be involved in morphogenesis, their number increasing with the growth of the epicytic folds (Schrevel et a/., 1983). In the euglenian flagellate Cyclidiopsis acus, filaments with a central densification join the membranes of neighboring vesicles which contain large paramylon rods; their aspect is similar to that of tonofilaments (Mignot, 1975). In Trypanosoma and Crithidia (Fig. 3 9 , the flagellum, fixed to the cuticle of the

FIG. 35. Flagellar attachment of Crirhidiafascicularu to the cuticular lining (cl) of the rectum. The gap between the flagellar membrane and the cuticular lining is occupied by a fibrous coat (fc) which lines the entire hindgut. The thickened inner leaflet of the flagellar membrane and the mass of filaments (fil) emerging from a dense band are features common to all adhesions. ax, Axoneme. X57,OOO. Drawn from a micrography of Brooker (1971).

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digestive tract of the insect host, first contains a submembranal thickening from which filaments leave which run toward the axoneme; the overall appearance is reminiscent of a hemidesmosome (Brooker, 1971; Evans et al., 1979; SoutoPadron and de Souza, 1979). In these two cases, and if the filaments are truly intermediate filaments, they would play the role of a link between membranes or between organelles and membrane. According to Numata and Watanabe (1982), the contractile ring involved in the division of ciliates should be considered as being composed of intermediate filaments. However, the diameter measured when they are in a relaxed state suggests that they should be classed as nonactin microfilaments. Intermediate filaments would also seem to be present at the level of the buccal apparatus (Numata and Watanabe, 1982) (see Section IX).

VIII. Epiplasm This is a layer of proteinaceous material of variable thickness joined to the inner alveolar membrane of the cortex in ciliates. It is especially well developed in somatic regions (Fig. 32). A. STRUCTURE Generally, the epiplasm seems to be made up of dense nonstructured relatively homogeneous matter (Hausmann and Mulisch, 1981); however, in the chonotrich Chifodochona, where it is developed in the unciliated regions, it is composed of several fibrogranular layers of different textures (Grain and Batisse, 1974); it is also thick in Suctoria and Entodiniomorpha. After extraction of pellicles of Tetrahymena, the epiplasm resolves into filaments between 7 and 10 nm in diameter (Collins et al., 1980), which are, in the dividing cells, linked to the overlying alveolar membrane (Yasuda et al., 1980). In Polypfastron, the epiplasm is seen, after extraction, to be composed of filaments 4 nm in diameter, isoluble in KI and PTA (Viguks et al., 1983a) (Fig. 36). In other protists, layers of subalveolar material may be considered as homologous of the epiplasm of ciliates; this is the case for the layer covering the internal filaments of the epicytic folds in Gregarina (Walker er al., 1979; Schrevel et a l . , 1983), for the whole of superposed lamellae in the peridinia Kofoidiniurn FIG. 36. Cortical cytoskeletal structures in Polyplastron. An oblique section of the cortex (a) shows the epiplasm (Ep), the cortical microtubules (Mt). the nodular reticulum (R). the internal filamentous layer (FL),and the barren kinetosomes (Ks).b shows another plane of sections with the same elements. (c) After extraction by Triton X-100 and PTA, only the plasma membrane (M) and the filamentous epiplasm (Ep) subsist. (a) X24,OOO; (b) X50,OOO; (c) X50,000. From Viguts er al. (1983).

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(Cachon and Cachon, 1974a), and for the dense cortical layer of euglenians. In ciliates, the epiplasm, in the ciliated parts, is in continuous relation with the “rosette” (or terminal plate) of the kinetosome (Vaudaux, 1976) (Fig. 32); the external pad of the rosette is probably directly linked to the plasma membrane, because the rosette remains attached to this membrane during obtention of isolated pellicles (Williams et al., 1979).

B. NATURE Analysis of isolated pellicles show that the MW of the major protein of the epiplasm in different species and strains of Tetrahymena varies between 122 and 145K (Vaudaux, 1976; Vaudaux et al., 1977) (Fig. 33); similarly, the MW of the main protein of the rosette varies between 145 and 174K (Vaudaux et al., 1977). In the genus Pseudomicrothorax, isolated epiplasms of both species Pseudomicrothorax dubius and Pseudomicrothorax agilis were found to be highly heterogeneous with regard to their protein composition (R. Peck, personal communication). However, in P . dubius, the epiplasm contains a major component migrating as a single band of 78K in SDS-PAGE, but showing several isoelectric variants when examined by bidimensional analysis. Such a polymorphism is also observed for the main components of the epiplasm in P . agilis, but, in this cell, the major polypeptides show slightly distinct MW ranging from 75 to 90K. Isolated epiplasms from both cells also contain polypeptides with very low molecular weights comprised between 1 1 and 13K. In Polyplastron, the major protein has a MW of 96K and consists in a single molecular species (Vigubs et MCttnier, 1983). However, as judged by immunoelectron microscopy, additional proteins of MW 145 and 175K also might enter in the constitution of the epiplasmic layer of this ciliate (B. Vigds, personal communication).

C. ROLE The main role that the epiplasm seems to play is the strengthening of the cellshape maintenance system; thus, in the absence of any membrane in isolated pellicles of Tetrahymena, it maintains the stability of the pellicle and remains attached to the kinetodesmal, postciliary, and transverse fibers by solid links (Vaudaux, 1976). It also keeps in place the rosettes of the kinetosomes, or at least their external pads, by establishing solid relationships between the rosettes within a kinety, or between neighboring kineties (Collins et al., 1980). After glycerination, the epiplasm remains compact and plays the role of a true cytoskeletal structure in Pseudomicrothorax (Hausmann and Mulisch, 198 1). In

THE CYTOSKELETON IN PROTISTS

23 1

the division furrow of Tefrahymena, the epiplasm is first linked to the alveolar membrane and to the filamentous contractile ring; when this ring undergoes contraction, the epiplasm separates from the alveoles, becomes disorganized, and falls in with the ring, making the cortex more plastic and thus allowing cytodierese to take place (Yasuda et al., 1980). The major protein of the epiplasm is probably the functional equivalent of the spectrin, although the latter has a MW above 200K; a small amount of actin (MW 42K) seems to be associated with it in Tetrahymena (Williams et al., 1979); after solubilizing with KCI, it precipitates forming a filamentous network identical to that formed by spectrin under the same conditions. a-Actinin is present in the epiplasmic ridges and in the epiplasm along the kinetal furrows in Pseudomicrothorax (Hauser et al., 1980), and might be involved in anchorage of actin. Like spectrin (and associated actin) which is linked to the plasma membrane proteins and regulates their lateral mobility, the epiplasm might play some role in the control of the migration and positioning of the cortical structures and in their anchoring, in ciliates (Williams ef al., 1979). In Tefrahymena fhermophila, the epiplasm is involved in the anchorage of cortical mitochondria, establishing links with these organelles when they arrive at their cortical site (Aufderheide, 1983), probably guided by cortical microtubules.

D. REMARKS The epiplasm presents, in all ciliates (and some other protists), a situational analogy and perhaps a functional one; but its structure and chemical composition are variable. This chemical variability in the various Tefrahymena, for instance, might be the mark of evolution by diversification from a common starting strain; intraspecific variation being nonexistent in T. thermophifa (Fig. 33), this would mean that, in the other cases, differences between strains belonging to one species are already differences between actual newborn species; so, the two strains of T. vorax represent, in fact, two different species within the complex T . v o r a (Vaudaux er al., 1977). However, immunological relationships among epiplasmic components of ciliates and other protozoa could be demonstrated using antibodies raised against Pseudomicrothorax dubius isolated epiplasm. Since numerous immunoreactive proteins were detected by immunoblots in ciliates, flagellates, sporozoa (including Plasmodium), and Amoeba (Peck et al., 1986), their selective localization in the epiplasm or similar structures could be confirmed as yet by immunoelectron microscopy in the ciliate Paramecium fetraurefiaand in the phytoflagellate Euglena acus (B. Vigubs, G. Brugerolle, and G . Bricheux, personal communication). Further investigations would permit to assess whether in ciliates, and in a more general point of view in protozoa, the

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epiplasm, or homologous subsurface cytoskeletal structures, are concerned with a unique protein, (or with a little number of proteins), which has been modified during evolution and speciation.

IX. Nonactin Microfilaments The discovery, in protists, of numerous filamentous systems, the constitutive elements of which are filaments of diameter 5 6 nm, and which do not contain actin, even though they are sometimes an integral part of contractile structures, leads us to distinguish a new category of cytoskeletal elements, which has not yet been identified in metazoans, the “nonactin microfilaments.” Among the best-known examples, some are elements of contractile systems, others are not. A. CONTRACTILE NONACTINMICROFILAMENTS These occurs in systems the contraction of which is not of the actomyosin interaction type; generally, the contraction is Ca2 -dependent and ATP-independent. Their chemical nature and their MW remain unknown, even if it has been proven that they are not decorated with HMM. +

1. Well-Known Cases (de Haller, 1977)

1. At the base of the axopods of Sdcholonche, bundles of nonactin microfilaments connect the axopodial microtubules to the nuclear membrane on which the axopod leans (Fig. 37); these filaments are 2.5 nm in diameter; they are wound together in twos. The contraction of the bundles is Ca*+-dependent and ATPindependent, and it causes the movement of the axopod (Cachon er al., 1976). 2. Myonemes. These are tracti of filaments which, in ciliates Geleidae and heterotrichs and in some other protists, are laid out lengthwise in the cortex, enabling the contraction of the body or part of the body to occur. They are not invaded by endoplasmic reticulum dependences, but are surrounded by perimyonemal vesicles which must regulate the amount of free Ca2+ ions. In heterotrichs (Fig. 27), the myonemes are situated in the somatic cortex, under the microtubular arrays formed by the postciliary fibers; they are surrounded by an important vacuolar reticulum. In Srenfor, in the relaxed state, the filaments are 3 to 5 nm in diameter, and are well ordered, in parallel with the longitudinal axis of the myoneme; in the contracted state, they are less well ordered and form short helical twisted tubular-like filaments 8 to 12 nm in diameter. Contraction is initiated by Ca2 , relaxation by EGTA; the contraction-relaxation cycle can be experimentally repeated a few times, with responses which become weaker and weaker. The filaments are not decorated with HMM. +

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FIG. 37. Schematic drawing illustrating the basal region of an axopod of Sricholonche zanclea (Radiolaria). Note the presence of 2.5-nm filaments (0 which move the axopod like an oar by alternative contraction. Mt, Microtubules; H, microtubule-organizing center; D, dense material lying against the nuclear membrane (ne); nc, nuclear capsule; N, nucleus; ef, extracellular fibrils; E, extracellar environment; m. plasmalemma. From Cachon and Cachon (1978).

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The contractile system is thus different from the actomyosin plus ATP one. The contraction seems to be due simply to a change in conformation of the filaments (Huang and Pitelka, 1973; Kristensen et al., 1974). In digitonin-extracted models of Spirostomum, an other heterotrich, the contraction of the myonemes, produced after adjunction of Ca2+, is followed by spontaneous relaxation (even in the presence of Ca2+) which can be inhibited by ATPase inhibitors and by colchicine (Holberton and Ogle, 1975); relaxation seems to be due to motion of the postciliary microtubular ribbons which, by the ATPase activity of their desmoses, slide over each other until they return to their initial relative position. Contraction is due to the nonactin microfilaments of the myonemes; relaxation is passive for the myonemes and is due to the links between adjacent ribbons of microtubules. In Lacrymaria, the neck myonemes exhibit a periodic structure, the thickening of which may be due to the twisting of the filaments (Cachon and Cachon, 1981~).In Condylostoma, such a periodic aspect can be seen (Fig. 27b and c) (Bohatier, 1978). In vorticellids the subpellicular myonemes are linked to “motrice plates” which assume the junctions between the perimyonemal vesicles, the myonerne, and the subpellicular alveoli (Allen, 1973), which enables the transmission of external information and reaction by contraction. 3. Spasmonemes. These are tracti of filaments situated in the stalk of vorticellids. They are penetrated by vesicles of the endoplasmic reticulum which can release (during contraction) or capture (during relaxation) the Ca2 ions. Contraction does not involve thickening of the filaments which keep their diameter of 2 to 3 nm (Amos, 1975; Cachon and Cachon, 1981~); it is induced by Ca2+ , is ATP-independent, and its intensity depends on the Ca2 concentration (Ochiai et al., 1979); it is fast (5 to 20 msec), and not purely mechanical, the energy used being that of the bonds between Ca2+ and the calcium-binding proteins (CBP) (Ochiai et al., 1979); at least 3 proteins, extracted by urea, are CBP; they have a MW under 20K (18, 18, and 16K); they constitute the “spastin-B complex” which is calmodulin-like (Yamada and Asai, 1982); however they differ from calmodulin, since they do not bind to troponin C. Spastins are not structural proteins; these therefore remain to be identified (Yamada and Asai, 1982). Relaxation, induced by EGTA, is stimulated by ATP and requires Mg2+ (de Haller, 1977); it is assisted by the outer sheet of the spasmoneme which, being elastic, is a passive antagonist of the spasmoneme (Ochiai et al., 1979). The contraction-relaxation cycle can be experimentally induced up to 140 times consecutively. 4. The contractile division ring. While the division furrow of various cells of metazoans contains a ring of actin microfilaments, with myosin, a-actinin (Pol+

+

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lard, 1981), and filamin (Nunnally et a f . , 1980), in ciliates the division ring does not seem to contain actin, but this remains to be demonstrated with certainty; it is made up of filaments parallel to the surface, whose diameter varies according to the state of contraction; it can be 2.5, 5, or 10 to 15 nm in Tetrahymena (Yasuda et al., 1980); it is 3 to 5 nm in the relaxed state, and 10 to 15 nm in the contracted state in Stentor, where the contractile division ring shows relationships with the myonemes, that could be proof of its myonemal origin (Diener et al., 1983); it is 4 to 10 nm in Nassufu (Tucker, 1971a ); in Thuricofu (Eperon, 1982), the contractile ring presents fine filaments and thicker filaments which call to mind the two possible states (relaxed and contracted). In Tetrahymena the contractile ring may contain (Yasuda et a f . , 1980) or may not contain (Jerka-Dziadosz, 1981) transverse strips or granules, and it is linked to the epiplasm at the beginning of the division; then it goes deeper and drags the epiplasm which becomes disorganized (Yasuda et a f . , 1980); it might, after division, partially subsist at the apex of the opisthe daughter cell in the form of a ring of filaments linked to the base of kinetosomes, but not linked to the epiplasm (Jerka-Dziadosz, I98 I ) . The division ring of Stentor is accompanied, outside, by microtubules parallel to it, the role of which is unknown (Diener et a l . , 1983), while, in Nassula, the microtubules are longitudinal and reduce the action of constriction (Tucker, I97 la). In Terrahymena, there are no microtubules under the division furrow. From whole cells of Tetruhymena a protein with a MW of 38K (FFP 38) has been extracted by KCI (Numata et a f . , 1980a). The story of this protein is curious; it was, at first, compared with actin; its MW and its p1 (6.7) are different from those of actin; it does not stimulate the Mg2+-ATPase activity of muscular myosin, and it does not make up the F-actin-G-actin conversion; an antiserum prepared against this protein does not react with actin, myosin, or tropomyosin of skeletal muscle, but reacts with the isolated buccal apparatus and with the division furrow, the isolated pellicle and the mitochondria of Tetrahymena (Numata et a f . , 1980a); since polymerization in vitro produces filaments 13 to 15 nm in diameter (Numata et al., 1980b; Yasuda et ul., 1980), it might be that some filaments of the contractile division ring are composed of this protein (Numata et al., 1980a), and that polymerization in vitro produces filaments in the contracted state, with a diameter similar to that of contracted filaments seen in vivo. But the same authors (Numata and Watanabe, 1982) later established that the protein which effectively gives birth, in vitro, to filaments of diameter 14 nm has, in fact, a MW of 49K, and that the FFP 38 was not a component of these filaments. Consequently, they now consider that these filaments are intermediate filaments (Numata and Watanabe, 1982), they localize the 49K protein at the level of the buccal apparatus, the mitochondria, and the mucocysts, and they no longer speak about the division furrow (Numata et al., 1983); they attribute to this protein a crucial role in the morphogenesis of the oral

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primordium and in the rearrangement of the old buccal apparatus, owing to the fact that it is associated with the microtubules and that it is a protein belonging to intermediate filaments (Numata er al., 1983).

2. Doubrjiul Cases Only a few will be mentioned here. In the mucron of the gregarine Lecudina, filaments 6 to 7 nm in diameter, regrouped in a whole which evokes the myonemes of ciliates, seem to play a role in the contraction and the functioning of the sucker. The loss of the epimerite in the gregarina Sycia is due to the contraction of a basal sphincter which is formed by filaments 12.5 nm in diameter (Schrevel and Vivier, 1966) which could be classed as contracted nonactin filaments. But the nature of the filaments is unknown in both cases. In the same way, filaments of unknown nature 5 nm in diameter constitute spicule-retracting myonemes in Acantharia (Febvre, 1974). It may be wondered whether a certain number of so-called periodic fibers, as those which bind the two kinetosomes of Chilomonas, for instance (Mignot et al., 1968), which are made up of longitudinal filaments with transverse strips, are not simply nonactin microfilamentous tracti; measurements are lacking in most of the descriptions, as are chemical identifications. There is also a doubt about the nature of the internal filaments of the epicytic folds of Gregarina which are, according to some authors, 7 nm in diameter (Walker et al., 1979), and according to others 12 nm in diameter (Schrevel er al., 1983) (Chap. VII).

B. NONCONTRACTILENONACTINMICROFILAMENTS 1. Well-Known Cases The ecto-endoplasmicboundary of the ciliate Isotricha is made up of a double layer of filaments 4 nm in diameter (Fig. 38) which seems noncontractile; it is devoid of actin, but contains 2 major proteins of MW 23 and 22K (pt between 4.6 and 5.4) (Vigubs etal., 1983a,b, 1984). The interkinetosomal reticulum of tsotricha, located between the plasma membrane and the ecto-endoplasmic boundary, is composed of filaments linking the proximal parts of neighboring kinetosomes (Fig. 38). These filaments are 6 nm in diameter; they are devoid of actin, insoluble in KCI, and contain 2 major FIG.38. Nonactin microfilamentous structures in lsotricha prostoma. The entire cortex (a) is divided in two parts, the ectoplasm (Ec) and the endoplasm (En),by a microfilamentous bilayer, the ecto-endoplasmic boundary (EEB). A detail of the EEB is given in b. The fine interkinetosomal reticulum (FR) is present between the proximal parts of the kinetosomes (c). (a) x25,OOO: (b) X40,OOO; (c) X30,OOO. (a) From B. Viguks (unpublished); (b) from Viguks er al. (1984); (c) from Viguks et a/. (1985).

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proteins (MW 58 and 63K; pf 5.3 and 5 . 8 ) (Viguks et al., 1983a,b, 1985). This reticulum could be considered as the equivalent of the infraciliary lattice found in the cortex of other ciliates (Allen, 1971). The internal cortical filamentous layer of Polyplastron (ciliate) is made up of filaments 6 nm in diameter (Fig. 36a and b), which contain major proteins of MW 230, 170, 145, and 22K (Vigds and MttCnier, 1983). In the same genus, a reticulum composed of nodules linked by filaments is located between the cortical microtubules, under the epiplasm (Fig. 36a and b); these filaments have a diameter of 3 nm and a major protein of MW 60K (pf 5.6) (Viguks and MCtCnier, 1983). It might be thought that such arrays of nonactin microfilaments play a role in the elasticity and maintenance of the shape of the body; indeed, in fsotricha, the ecto-endoplasmic boundary sends out tracti surrounding the nuclei, which are parts of the “nucleosuspensor. ” The precise role of the ecto-endoplasmic boundaries, which are frequent in ciliates, and the exact nature and ubiquity of the constitutive proteins remain to be investigated. 2. Doubtful Cases The filaments of the “hemidesmosomes” of the trypanosomids are certainly noncontractile. In Crithidia (Fig. 3 3 , they are the size of microfilaments (3 nm in diameter) (Brooker, 1971), while their position calls to mind intermediate filaments of the tonofilament type (Evans et al., 1979).

X. Filament Systems of Unknown Nature There remain, in protists, after all that has been mentioned above, a quantity of filament systems for which we have a very precise idea of the spatial arrangement, but for which biochemical data and indications regarding dimensions and possible function are entirely lacking. To cite examples only among ciliates, filament networks not linked to kinetosomes are found in peniculina and chonotrichida (in Grain, 1984). Many other systems are linked to the kinetosomes (Fig. 39), and form more or less organized networks, with or without condensation nodes, at the level of the buccal apparatus in certain hymenostomata; the most highly organized and regular system is found at the base of the peniculus of Paramecium aurelia (Schneider, 1964), with superimposed condensation nodes. The stacking of such nodes gives the whole fiber its overall periodic appearance, at the level of the somatic kinetosomes of Sicuophora for example (Fig. 40) (de Puytorac and Grain, 1968). Some of these filament systems can establish distant relationships between kinetosomes and other fibrillar structures, or between various kinetosomes.

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C

b

e

FIG. 39. Filamentous systems of unknown nature in ciliates: (a) Fronronia depressa (peniculus PI). (b) Disemarosromu (peniculus). (c) Urocentrurn turbo (parorale). (d) U.rurbo (peniculus). (e) F . depressa (parorale). (f) Campanella: infundibulum fiber, linked to the kinetosomes (C) of the polykineties, with a regular arrangement in zone 2, and packing of the nodes at the periphery (T). (8) Detail o f f : filaments (M) and nodes (N). From Grain (1984).

The role which can, for the moment, be ascribed to these systems is the constitution of resistant supporting structures, giving, for instance, the buccal apparatus of Terruhymena (Wolfe, 1970) or Turuniellu (Iftode and Grain, 1975) its structural integrity, assisted in this by the microtubules. The only biochemical indication presently available concerns a system of

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FIG.40. Filaments of unknown nature in Sicuophora xenopi. Note link (arrow) with a somatic kinetosome (Ks) and the periodic aspect. X68.000.

filaments (UMN),found at the level of the buccal apparatus of Tefrahyrnena (Nilsson, 1976) (Fig. 6), which is linked to the paroral and runs along this throughout its length; it is a nonorganized network, and it seems to be constituted of proteins of MW 82 and 86K (Williams and Bakowska, 1982).

THE CYTOSKELETON IN PROTISTS

24 1

XI. Conclusion In this overview of the cytoskeleton of protists, we have seen various types of elements some of which have been thoroughly studied (structure, chemical nature, and function) and others which still require investigation. 1. We have seen, in protists, that so-called ubiquitous elements such as actin (and myosin) are present, but it seems that they are not present in all the welldeveloped cytoskeletal structures in all the protists. Only microtubules seem to occur in all of them. On the other hand, other elements, such as intermediate filaments, are very rare in protists, perhaps because they await identification. Some, like the epiplasm, could be the equivalent of submembranal elements of the cells of metazoans. Many protists possess a category of cytoskeletal filaments unknown in metazoans, namely nonactin microfilaments, which may or may not be contractile, and which are equal in size, or smaller than actin microfilaments: these elements are specific to protists, at present. 2. This review has brought to the fore a certain number of problems that need to be addressed, and areas that requires research. Problems still arise regarding the mechanism of contraction of the actomyosin system in amoebae: regulation by Ca2+ or something else? The proteins which enable the association of actin microfilaments in bundles to take place in amoebae and myxomycetes need to be found and characterized. It seems necessary to continue to clone the actin genes of protists, to analyze their nucleotidic sequences, and compare them with those of metazoans in order to assess evolutionary distances between the different phyla. In the same way, it is necessary to find out precisely to what extent the heterogeneity of a-tubulin for different species corresponds, or does not correspond, to a heterogeneity in the composition of the gene, some doubts subsisting after the translation of identified mRNA (Fliss and Suyama, 1979; Myhal and Hufnagel, 1979). It would be useful to find out, with sophisticated methods, if actin and myosin are really absent from the contractile division ring of ciliates, and if the periodic fibers have or do not have an ATPase activity. 3. With respect to the elements specific to protists, we need to find out if the variability of the epiplasmic proteins is the mark of a diversifying evolution, or if we are dealing with converging structures, without any phyletic relation at all. The new category of cytoskeletal elements, the nonactin microfilaments, must be studied in detail; it is necessary to know their major constituent proteins, their associated proteins, and their functions; it seems useful to find out if such proteins and such filaments are present in various species of protists, and in cells

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of metazoans. Finally, the numerous systems still imperfectly studied in ciliates (Section X) must be identified. 4. When the different elements of the cytoskeleton in some protists are known, it will be necessary to define the functional relationships between them, for example, to determine the exact mechanism of the interrelation between microfilaments and microtubules in contractility or in transport (what enzymes and what source of energy are involved?). Though protists have been extensively used in the studies of actomyosin relations, and of the cytoskeletal elements of the cilium and flagellum, they evidently possess a quantity of systems the variety of which is much greater than that of cells of metazoans. Accordingly, further research on these systems should help in our understanding of new types of cell functioning.

ACKNOWLELXMENTS I thank Dr. J. Beisson and Dr. A. Adoutte, Centre de Gtnttique Moltculaire du CNRS, Gif-surYvette (France), for their critical reviews of the manuscript. Collaborations and discussions with G. Mtttnier, B. Vigues, R. Peck, J. P. Mignot, C. A. Grolikre, J. Senaud, P. de Puytorac, and G. Brugerolle are appreciated. I also thank all the scientists who have kindly provided illustrations. I am indebted for technical assistance to J. Roudeix, C. Vincenot, and J. L. Vincenot. Financial support was given in part by the CNRS, under the grant to UA 040 138.

REFERENCES Adoutte, A., Rarnanathan, R., Lewis, R. M., Dute, R. R., Ling, K. Y., Kung, C., and Nelson, D. L. (1980). J . Cell Biol. 84, 717-738. Adoutte, A., Cohen, J., Hill, A. M., Pantaloni, D., and Beisson, J. (1982). Biol. Cell. 45, 265a. Adoutte, A., Claisse, M., and Cance, J. (1984). Origins Life 13, 177-182. Al-Khazzar, A. R., Butler, R. D., Eamshaw, M. J., Emes, M. J., and Sigee, D. C. (1983). J . Prorozool. 30, 35A. Allen, R. D. (1967). J. Protorool. 14, 553-565. Allen, R. D. (1969). J. Cell Biol. 40,716-733. Allen, R. D. (1971). J. Cell Biol. 49, 1-20. Allen, R. D. (1973). J. Cell Biol. 56, 559-579. Amos, W. B. (1975). In “Molecules and Cell Movement” (S. Inou6 and R. E. Stephens, eds.), pp. 41 1-436. Raven, New York. Amos, W. B., Grimstone, A. V., Rothschild, L. J., and Allen, R. D. (1979). J. Cell Sci. 35, 139164.

Aufderheide, K. J. (1983). J . Prorozool. 30, 457-460. Bardele, C. F. (1973). Cytobiologie 7, 442-488. Bardele, C. F. (1976). Z. Nururforsch. 31c, 190-194. Beisson, J., and Rossignol, M. (1975). Mol. Biol. Nucleocyroplusmic Relut. 291-294. Bernard, F. (1980). These Doct. 3e cycle, Univ. Clermont 11. Bloodgood, R. A. (1975). Cyrobios 14, 101-120. Bloodgood, R. A., and Fitzharris, T. P. (1979). Cyrobios 23, 109-117.

THE CYTOSKELETON IN PROTISTS

243

Bloodgood, R. A., and Miller, K. R. (1974). J . Cell Biol. 62, 660-671. Bloodgood, R. A,, Miller, K. R., Fitzhanis, T. P., and McIntosh, J. R. (1974). J. Morphol. 143, 77-106.

Blose, S. H. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 3372-3376. Bohatier, J. (1972). These Doct. 3e cycle, Univ. Clermont 11. Bohatier, J. (1978). Protistologica 14, 433-450. Bordier, C., Garavito, R. M., and Armbruster, B. (1982). J. Prorozool. 29, 560-565. Borisy, G. G., Olmsted, J. B., Marcum, J. M., and Allen C. (1974). Fed. Proc., Fed. Am. SOC. Exp. Biol. 33, 167-174. Bourguignon, L. Y.W., Nagpal, M. L., Balazovich, K., Guerriero, V., and Means, A. R. (1982). J . Cell Biol. 95, 793-197. Bowser, S. S., and McGee-Russell, S. M. (1982). J. Cell Biol. 95, 329a. Bowser, S. S., McCee-Russell, S. M., and Rieder, C. (1983). J. Prorozool. 30, 12A. Brinkley, B. R., Cox, S. M., Pepper, D. A., Wibble, L., Brenner, S. L., and Pardue, R. L. (1981). J . Cell Biol. 90, 554-562. Brooker, B. E. (1971). Proroplasma 73, 191-202. Brown, S. S., Yamamoto, K., and Spudich, J. A. (1982). J . Cell Biol. 93, 205-210. Brugerolle, G., and Joyon, L. (1973). Profisfologica9, 71-80. Brugerolle, C . , Joyon, L., and Oktem, N. (1973). Prorisrologica 9, 339-348. Brunke, K. J., Young, E. E., Buchbinder, B. U., and Weeks, D. P. (1982). Nucleic Acids Res. 10, 1295- 1310. Burland, T. G., Gull, K., Schedl, T., Boston, R. S., and Dove, W. F. (1983). J. Cell Biol. 97, 1852- 1859.

Cabral, F., Gottesman, M. M., Zimmerman, S. B., and Steinert, P. M. (1981). J. Biol. Chem. 256, 1428-1431.

Cachon, J., and Cachon, M. (1971). Arch. Prorisrenkd. 113, 80-97. Cachon, J., and Cachon, M. (1972). Arch. Proristenkd. 114, 51-64. Cachon, J., and Cachon, M. (1974a). Prorisrologica 10, 217-222. Cachon, J., and Cachon, M. (1974b). Ann. Biol. 13, 523-560. Cachon, J., and Cachon, M. (1975). C. R. Acad. Sci. 280, 2341-2343. Cachon, J., and Cachon, M. (1978). Arch. Proristenkd. 120, 148-168. Cachon, J., and Cachon, M. (1980). Biol. Cell. 37, 23-34. Cachon, J., and Cachon, M. (1981a). Ann. Biol. 20, 131-139. Cachon, J., and Cachon, M. (1981b). Biol. Cell. 40, 23-32. Cachon, J., and Cachon, M. (1981~).BioSystems 14, 313-326. Cachon, J., Cachon, M., Febvre-Chevalier, C., and Febvre, J. (1973). Arch. Proristenkd. 115, 137153.

Cachon, J., Cachon, M., and Tilney, L. G. (1976). J. Microsc. Biol. Cell. 27. Cachon, J., Cachon, M., and Bruneton, J. N. (1981). Biol. Cell. 40, 69-72. Chalovich, J. M., Chock, P. B., and Eisenberg, E. (1981). J . Biol. Chem. 256, 575-578. Chang, M. T., Dove, W. F., and Laffler, T. G. (1983). J. Biol. Chem. 258, 1352-1356. Cherksey, B. D., Zadunaisky, J. A,, and Murphy, R. B. (1980). Proc. Nafl. Acad. Sci. U.S.A. 77, 6401-6405. Clarke, M., and Spudich, J. A. (1977). Annu. Rev. Biochem. 46, 797-822. Clayton, L., Quinlan, R. A , , Roobol, A,, Pogson, C. I., and Gull, K. (1980). FEBSLerr. 115,301305.

Cohen. C. M.. and Korsgren, C. (1980). Eiochem. Eiophys. Res. Commun. 97, 1429-1435. Cohen, J., and Beisson. J. (1980). Generics 95, 797-818. Cohen, J., Beisson. J., and Tucker, J . B. (1980). J. Cell Sci. 44, 153-167. Cohen, J., Adoutte, A,, Grandchamp, S . , Houdebine, L. M., and Beisson, I. (1982). Biol. Cell. 44, 35-44.

244

JEAN GRAIN

Cohen, J . , Garreau de Loubresse, N., and Beisson, J. (1984). J. Submicrosc. Cyrol. 16, 103-104. Collins, 1. H., Cote, J. P., and Korn, E. D. (1982). J. Biol. Chem. 257, 4529-4534. Collins, T., Baker, R. L., Wilheim, J. M., and Olmsted, J. (1980). J . Ultrustruct. Res. 70, 92-103. Crossley , R., and Holberton, D. V. ( 1983). J . Cell Sci. 59, 8 I- 103. Davidson, L. A. (1982). J. Protozool. 29, 19-29. Dazy, A. C., Hoursiangou-Neubmn, D., and Sauron, M. E. (1981). Biol. Cell. 41, 235-238. De Brabander, M. J . (1983). Recherche 145, 810-820. De Haller, G. (1977). Ann Biol. 16, 241-258. de Puytorac, P., and Grain, J . (1968). Protistologicu 4, 121-130. Dentler, W. L. (1980). J . Cell Biol. 84, 364-380. Dentler, W. L., and Rosenbaum, J. L. (1977). J . Cell Biol. 74, 747-759. De Rosier, D. J., and Censullo, R. (1981). J . Mol. Biol. 146, 77-99. Desportes, I. (1969). Ann. Sci. Nat. 2001.(Paris) 11, 31-96. D'Haese, J. (1980). FEBS Lett. 121, 243-245. Didier, P. (1970). Ann. St. Biol. Besse-en-Chundesse 5, 1-274. Diener, D. R., Burchill, B. R., and Burton, P. R. (1983). J. Prorozool. 30, 83-90. Dingle, A. D., and Larson, D. E. (1981). BioSystems 14, 345-358. Dippell, R. V. (1968). Proc. Natl. Acud. Sci. U.S.A. 61, 461-468. Dubremetz, J. F., and Torpier, G . (1978). J . Ulrrasrrucr. Res. 62, 94-109. Dustin, P. (1981). Ann. Biol. 20, 141-160. Dustin, P. (1984). "Microtubules," 2nd Ed. Springer-Verlag. Berlin and New York. Dutcher, S. K., Huang, B., and Luck, D. J. L. (1984). J . Cell Biol. 98, 229-236. Edds, K. T. (1975). J. Cell Biol. 66, 156-164. Eperon. S. (1982). Thtse Doct.-es-Sci. Lausanne. Eperon, S., and De Haller, G. (1982). Protistologica 18, 85-102. Erlandsen, S. L.. Schollmeyer, 3. V., Feely, D. E., and Chase, D. G. (1978). J . Cell Biol. 79,264a. Evans, D. A., Ellis, D. S . , and Stamford, S. (1979). J. Prorozool. 26, 557-563. Favard. P., Carasso, N., and Faur6-Fremiet, E. (1963). J. Microsc. 2, 337-368. Febvre, J . (1974). Protistologica 10, 141-158. Firtel, R. A . , Timm, R., Kimmel, A. R., and Mc Keown, M. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 6206-6210.

Fliss, E. R., and Suyama, Y. (1979). J. Prorozool. 26, 505-509. Fowler, V. M., Luna, E. J . , Hargreaves, W. R., Taylor, D. L., and Branton, D. (1981). J. Cell Biol. 88, 388-395. Gadasi, H. (1982). Biochem. Biophys. Res. Commun. 104, 158-164. Gallo, J. M., and Anderton. B. H. (1983). EMBO J . 2, 479-483. Gallo, I. M., and Schrevel, J. (1982). Biol. Cell. 44, 139-148. Gallo, J. M., Karsenti, E., Bornens, M., Delacourte, A., and Schrevel, J . (1982). Biol. Cell. 44, 149-156.

Gallo, J. M., Pineau, N . , and Schrevel, J. (1983). J. Prorozool. 30,68A. Gavin, R. H. (1976). J . Exp. Zool. 197, 59-64. Gavin, R. H. (1980). J. Cell Sci. 44, 317-333. Gibbons, 1. R. (1981). J. Cell Biol. 91, 107s-124s. Glenney, J. R., Jr., and Weber, K. (1980). J . Biol. Chem. 255, 10551-10554. Glenney, J. R.,Jr., Bretscher, A . , and Weber, K. (1980).Proc. Nurl. Acad. Sci. U.S.A. 77,64586462.

Golinska, K. (1982). J . Embryo/. Exp. Morphol. 68, 99-114. Golinska, K. (1983). J. Cell Sci. 62, 459-475. Goodenough, U. W., and Heuser, J. E. (1982). J . Cell Biol. 95, 798-815. Gortz, H. D. (1982). J. Protozool. 29, 353-359.

THE CYTOSKELETON IN PROTISTS

245

Grain, J. (1969). Ann. B i d . 8, 53-97. Grain, J. (1984). In “Trait6 de Zoologie” (P. P. Grass& ed.), Vol. 2, pp. 35-179. Masson, Paris. Grain, J., and Batisse, A. (1974). J . Protozool. 21, 95-1 I I . Grebecka, L., and Hrebenda, B. (1979). Acru Protozool. 18, 493-502. Grebecki, A. (1982). Ann. Biol. 21, 275-306. Grimes, G. W. (1973). J. Cell Sci. 13, 43-53. Hackney, C. M., and Butler, R. D. (1981). J. Prorozool. 28, 151-157. Haimo, L. T., and Rosenbaum, 1. L. (1981). J. Cell B i d . 91, 125s-130s. Hartwig, J. H., Tyler, J., and Stossel, T. P. (1980). J. Cell Biol. 87, 841-848. Hauser, M., and Hausmann, K. (1982). Di&w‘erentiation 22, 67-72. Hauser, M., and Van Eys, H. (1976). J . Cell Sci. 20, 589-617. Hauser, M., Hausmann, K., and Jockusch, B. M. (1980). Exp. Cell Res. 125, 265-274. Hausmann, K., and Mulisch, M. (1981). Arch. Prorisrenkd. 124, 410-416. Hausmann, K., and Patterson, D. J. (1982). Cell Motil. 2, 9-24. Hausmann, K., Linnenbach, M., and Patterson, D. J. (1983). J . Ultrustrucr. Res. 82, 212-220. Henderson, D., and Weber, K. (1980). Exp. CeNRes. 129, 441-453. Herman, 1. M., and Pollard, T. D. (1981). J. Cell Biol. 88, 346-351. Hibberd, D. J. (1975). J. CeNSci. 17, 191-219. Hinssen, H. (1972). Cytobiologie 2. Exp. Zellforsch. 5, 146-164. Hoessli, D., Rungger-Brande, E., Jockusch, B. M., and Gabbiani, G. (1980). J . CellBiol. 84,305314.

Holberton, D. V. (1981). J. CellSci. 47, 167-185. Holberton, D. V., and Ogle, W. S. (1975). J . Protozool. 22, 39A. Holberton, D. V., and Preston, T. M. (1970). Exp. Cell Res. 62, 473-477. Holberton, D. V., and Ward, A. P. (1981). J . Cell Sci. 47, 139-166. Hollande, A,, and Valentin, I. (1968). Protistologicu 4, 127-140. Huang, B., and Pitelka, D. R. (1973). J. Cell Biol. 57, 704-728. Huang, B., Piperno, G.,Ramanis, Z., and Luck, D. J. L. (1981). J. Cell Biol. 88, 80-88. Hufnagel, L. A. (1969). J. Cell Biol. 40, 779-801. Humphries, S. E., Whittal, R., Minty, A., Buckingham, M. E., and Williamson, R. (1981). Nucleic Acids Res. 9, 4895-4908. Hyams, J. S. (1982). J . CellSci. 55, 199-210. Hyams, J. S., and Borisy, G. G. (1975). Science 189, 891-893. Iftode, F., and Grain, J. (1975). J. Protozool. 22, 88-96. Jacobson, B. S. (1980). Biochem. Biophys. Res. Commun. 97, 1493-1498. Jamieson, G. A., Vanaman, T. C., and Blum, J. J. (1979). Proc. Nutl. Acud. Sci. U.S.A. 76,64176475.

Jerka-Dziadosz, M. (1980). Protisrologicu 16, 571-589. Jerka-Dziadosz, M. (1981). J. Cell Sci. 51, 241-253. Johnson, U. G., and Porter, K. R. (1968). J . Cell Biol. 38, 403-425. Kaine, B. P., and Spear, B. B. (1982). Nature (London) 295, 430-432. Keith, C. H., Feramisco, J. R.,and Shelanski, M. (1981). J . CellBiol. 88, 234-240. Keller, T. C. S., 111, and Mooseker, M. S. (1982). J. Cell B i d . 95, 943-959. Korn, E. D. (1978). Proc. Nut/. Acud. Sci. U.S.A. 75, 588-599. Korohoda, W., and Stockem, W. (1975). Microsc. Actu 77, 129-141. Kristensen, B. I., Nielsen, L. E., and Rostgaard, J. (1974). Exp. Cell Res. 85, 127-135. Kuznicki, L., and Sikora, J. (1971). Acta Protozool. 8, 439-446. Kuznicki, L., and Sikora, J. (1973). Acru Protozool. 12, 143-150. Larson, D. E., and Dingle, A. D. (1981a). J. Cell Bid. 89, 424-432. Larson, D. E., and Dingle, A. D. (1981b). Dev. Biol. 86, 227-235.

246

JEAN GRAIN

Laval, M. (1971). Prorisrologica 7, 325-336. Lazaridts, E. (1975). J. Cell Biol. 65, 549-561. Leadbeater, B. S. C. (1983). Prorisrologica 19, 157-166. Leterrier, J. F., Liem, R. H. K., and Shelanski, M. L. (1981). J. Cell Biol. 90, 755-760. Liebes, L. F., Fleit, H., Zucker-Franklin, D., and Silber, R. (1980). Biochim. Biophys. Acra 633, 245-257. Lieska, N.,Chen, J . , Maisel, H . , and Romero-Herrera, A. E. (1980). Biochim. Biophys. Acra 626, 136- 153. Little, M., Luduena, R. F., Langford, G . M., Asnes, C. F., and Farrell, K. (1981). J. Mol. Biol. 149, 95-107. Little, M., Luduena, R. F., Keenan, R., and Asnes, C. F. (1982). J. Mol. Evol. 19, 80-86. Luna, E. J., Fowler, V. M., Swanson, J., Branton, D., and Taylor, D. L. (1981). J . Cell Biol. 88, 396-409. Lynn, D. H., and Zimmerman, A. M. (1981). Protoplasma 108, 29-38. McGee-Russell, S . M., Bowser, S. S., and Koury, S. T. (1982). J. Cell Biol. 95, 329a. Mackenzie, C. (1980). Cell Biol. Int. Rep. 4, 769. Mackenzie, C., and Walker, M. H. (1983). J . Protozool. 30, 3-8. McIntosh, J. R. (1973). J . Cell Biol. 56, 324-339. McIntosh, J . R., Ogata, E. S . , and Landis, S. C. (1973). J. Cell Biol. 56, 304-323. McKeithan, T. W., and Rosenbaum, J. L. (1981). J . Cell Biol. 91, 352-360. McKeithan, T. W., Lefebvre, P. A., Silflow, C. D., and Rosenbaum, J . L. (1983). J. CellBiol. 96, 1056- 1063. Malchow, D., Boehme, R., and Rahmsdod, H. J. (1981). J . Biochem. 117, 213-218. Marano, F., Galleron, C., Minty, A. J., Montarras, D., and Bornens, M. (1982). Cell Biol. Inr. Rep. 6, 1085-1092. Marcaud, L., Millet, M.,Portier, M. M., and Hayes, D. H. (1979). Colloq. Microbiol. CNRS p. 214. Maruta, H., and Korn, E. D. (1977). J . Biol. Chem. 252, 399-402. Marx, J. L. (1975). Science 18, 34-37. Maslin-Leny, Y. (1983). These Doct. cycle, Univ. Claude Bernard, Lyon. Mattern, C. F. T., and Honigberg, B. M. (1971). Trans. Am. Microsc. SOC.90, 309-313. Mellor, J. S., and Stebbings, H. (1980). J . Exp. Biol. 87, 149-161. Mbtknier, G . (1981). J. Prorozool. 29, 308. Mktknier, G . (1984). J. Prorozool. 31, 205-215. Mignot, J . P. (1963). C.R. Acad. Sci. (Paris) 257, 2530-2533. Mignot, J. P . (1975). Protisrologica 11, 177-185. Mignot, J. P., and Brugerolle, G. (1982). J. Ultrastrucr. Res. 81, 13-26. Mignot, J. P.,Joyon. L., and Pringsheim. E. G . (1968). Protisrologica 4, 493-506. Milstone, L. M., and McGuire, J. (1981). J. Cell Biol. 88, 312-316. Mockrin, S . C., and Korn, E. D. (1983). J. Biol. Chem. 258, 3215-3221. Mogensen, M. M., and Butler, R. D. (1983). J . Protozool. 30, 39A. Mooseker, M. S., and Tilney, L. G. (1973). J. Cell Biol. 56, 13-26. Morita, F., and Matsumoto, A. (1980). J . Biochem. 88, 1883-1886. Murray, J . M. (1981). J . Cell Sci. 49, 99-117. Myhal. M. L., and Hufnagel, L. A. (1979). J. Protozool. 26, 672-675. Nachmias, V. T., Huxley, H. E., and Kessler, D. (1970). J . Mol. Biol. 50, 83. Nilsson, J. R. (1976). C. R. Trav. Lab. Carlsberg 40,215-355. Nozawa, Y. (1982). J. Protozool. 29, 473-474. Numata, O., and Watanabe, Y. (1982). J . Biochem. (Tokyo) 91, 1563-1573. Yasuda, T., Hirabayashi, T., and Watanabe Y.(1980a). Exp. CellRes. 129,233-230. Numata, 0..

THE CYTOSKELETON IN PROTISTS

247

Numata, 0.. Yasuda, T., Hirabayashi, T., and Watanabe, Y. (1980b). J. Biochem. 88, 1499-1504. Numata. O., Hirono, M., and Watanabe, Y. (1983). Exp. Cell Res. 148, 207-220. Nunnally, M. H., D’Angelo, J. M., and Craig, S. W. (1980). J. Cell Biol. 87, 219-226. Ochiai, T., Asai, H., and Fukui, K. (1979). J. Protozool. 26, 420-425. Ockleford, C. D., and Tucker, J. B. (1973). J . Ulfrastrucr.Res. 44, 369-387. Opas, M. (1980). Bull. Acad. Pol. Sci., Ser. Sci. Biol. 28, 51 1-514. Ozaki, K., and Maruyama, K. (1980). J. Biochem. 88, 883-888. Ozaki, K., Sugino, H., Hasegawa, T., Takahashi, S., and Hatano, S. (1983). J. Biochem. (Tokyo) 93, 295-298. Pantaloni, D., Carlier, M. F., and Hill, T. L. (1983). Club Francais du Cytosquelette, Banyuls-surMer, Juin. Peck, R.,Viguts, B., andde Haller, G . (1986). 18th Annual Meeting of the USGEBNSSBE, Basel, Switzerland. Philippe, M., and Schrevel, I. (1982). Biochem. J. 201, 455-464. Philippe, M., Vinckier, D., Dubremetz, J., and Schrevel, J. (1982). J. Protozool. 29, 424-430. Piccinni, E., Albergoni, V., and Coppellotti, 0. (1975). J. Protozool. 22, 331-335. Piperno, G., and Luck, D. J. L. (1977). J. Biol. Chem. 252, 383-391. Piperno. G., Huang, B., Ramanis, Z., and Luck, D. J. L. (1981). J . Cell Biol. 88, 73-79. Plattner, H., Westphal, C., and Tiggemann, R. (1982). J. Cell Biol. 92, 368-377. Pollard, T. D. (1975). In “Molecules and Cell Movement” (S. InouC and R. E. Stephens, eds.), pp. 259-286. Raven, New York. Pollard, T. D. (1981). J. Cell Biol. 91, 156s-165s. Pollard, T. D. (1982). J. Cell Biol. 95, 816-825. Pollard, T. D., and Kom, E. D. (1973). J. Biol. Chem. 248, 4691-4697. Pollard, T. D., Shelton, E., Weihing, R. R., and Korn, E. D. (1970). J. Mol. Biol. 50, 91-97. Porchet-Hennere, E., and Vivier, E. (1971). Ann. Biol. 10, 77-113. Premier, G . , Vivier, E., Goldstein, S . , and Schrevel, J. (1980). Science 207, 1493-1494. Reichstein, E., and Kom, E. D. (1979). J. Biol. Chem. 254, 6174-6179. Remillard, S. P., and Witman, G. B. (1982). J. CellBiol. 93, 615-631. Robenek, H., and Melkonian, M. (1979). Arch. Protistenkd. 122, 340-351. Rodrigues de Santa Rosa, M. (1974). Thtse Doct. 3‘ cycle, Univ. Clermont. Roobol, A,, Pogson, C. I . , and Gull, K. (1980). Exp. Cell Res. 130, 203-215. Rosenberg, S., Stracher, A,, and Lucas, R. C. (1981). J. CellBiol. 91, 201-211. Roth, L. E., Pihalaja, D. J., and Shigenaka, Y. (1970). J. Ulfrustruct. Res. 30, 7-37. Ruiz, F., Adoutte, A,, Rossignol, M., and Beisson, J. (1976). Genet. Res. Cumbr. 27, 109-122. Runge, M. S., El-Maghrabi, M. R., Claus, T. H., Pilkis, S. J., and Williams, R. C., Jr. (1981). Biochemistry 20, 175- 180. Russell, D. G . (1983). J. Protozool. 30,41A. Russell, D. G., and Burns, R. G . (1983). J. Protozool. 30, 41A. Russell, D. G., and Sinden, R. E. (1981). J. Cell Sci. 50, 345-359. Sanger, J. W., Sanger, J. M., and Jockusch, B. M. (1983). J . Cell Biol. 96, 961-969. Sattler, C. A,. and Staehelin, L. A. (1979). J. Ultrastruct. Res. 66, 132-150. Schloss, I. A,, and Goldman. R. D. (1980). J. Cell Biol. 87, 633-642. Scholtyseck, E., Mehlhorn, H., and Senaud, J. (1972). Z. Parasirenkd. 40, 281-294. Schrevel, J. (1971). Protisfologica 7, 101-130. Schrevel, J . , and Vivier, E. (1966). Protistologicu 2, 17-28. Schrevel, J., Caigneaux, E.. Gros, D., and Philippe, M. (1983). J. Cell Sci. 61, 151-174. Schwartz, R. J., Haron, J. A., Rothblum, K. N., and Dugaiczyk, A. (1980). Biochemistry 19,58835890. Schwartzman, J. D.. and Pfefferkorn, E. R. (1983). J. Protozool. 30, 657-661.

248

JEAN GRAIN

Sherman, G. B., Buhse, H. E., Jr., and Smith, H. E. (1981). Trans. Am. Microsc. SOC. 100, 366372. Sherman, G. B., Buhse, H. E., Jr., and Smith, H. E. (1982). J. Prorozool. 29, 360-365. Sikora, J. (1981). Proroplasma 109, 57-77. Siracusa, G . , Whittingham, D. G.,and Felici, De M. (1980). J. Embryol. Exp. Morphol. 60,7 1-82, Sleigh, M. (1973). “The Biology of Protozoa. Contemporary Biology” (E. J. W. Barrington and A. J. Willis, eds.). Arnold, London. Soltynska, M. (1971). Acra Prorozool. 9, 49-82. Souto-Padron, T., and De Sousa, W. (1979). J. Prorozool. 26, 551-557. Spudich, J. A. (1974). J. Biol. Chem. 249, 6013-6020. Steams, M. E., and Brown, D. L. (1981). J. Ulrrasrrucr. Res. 77, 366-378. Stebbings, H., Boe, G. S., and Garlick, P. R. (1974). Cell Tissue Res. 148, 331-345. Stephens, R. E. (1983). J . Cell Biol. 96, 68-75. Stockem, W. (1977). Proc. Inr. Congr. Prorozool., Srh, 131-139. Stockem, W., Hoffmann, H. U.,and Gawlitta, W. (1982). Cell Tissue Res. 221, 505-509. Sugino, H., and Matsumura, F. (1983). J . CellBiol. 96, 199-203. Sundararaman, V., and Hanson, E. D. (1976). Genet. Res. Cumbr. 27, 205-211. Suzuki, Y., Nagao, S., Hirabayashi, K., and Watanabe, T. (1981). J. Biochem. 89, 333-336. Tamm, S. L. (1976). Cell Motil. 3, 949-967. Tamm, S. L. (1978). J. Cell Biol. 78, 76-92. Tanaka, H., and Hatano, S. (1976). J . Mechunochem. CellMotil. 3, 195-200. Taylor, D. L. (1976). In “Cell Motility” (R. Godman, T. Pollard, and J. Rosenbaum, eds.), pp. 797-823. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Taylor, L., Condeelis, J. S . , Moore, P. L., and Allen, R. D. (1973). J . Cell Biol. 59, 378-394. Thomashow, L. S., Milhausen, M., Rutter, W. J., and Agabian, N. (1983). Cell 32, 35-43. Tiggeman, R., and Plattner, H. (1981). Eur. J. CeLlBiol. 24, 184-190. Tiggeman, R., Plattner, H., Rasched, I., Baeuerle, P., and Wachter, E. (1981). J. Histochem. Cyrochem. 29, 1387-1396. Tilney, L. G. (1971). J. Cell Biol. 51, 837-854. Tilney, L. G., Hatano, S . , Ishikawa, H., and Mooseker, M. S . (1973). J. Cell Biol.59, 109-126. Tobacman, L. S., and Korn, E. D. (1983). J . Biol. Chem. 258, 3207-3214. Totsuka, T., and Hatano, S. (1970). Biochim. Biophys. Acra 223, 189-197. Travis, J. L., and Allen, R. D. (1981). J. Cell BioL. 90, 21 1-221. Travis, J. L., Kenealy, J. F. X.,and Allen, R. D. (1983). J . Cell Biol. 97, 1668-1676. Tseng, P. C. H., Runge, M. S., Cooper, J. A., Williams, R. C., Jr., and Pollard, T . D. (1984). J . Cell Biol. 98, 214-221. Tucker, J. B . (1968). J. CellSci. 3, 493-514. Tucker, J. B. (1971a). J. Cell Sci. 8, 557-571. Tucker, J. B. (1971b). Narure (London) 232, 387-389. Uyemura, D. G., Brown, S. S., and Spudich, J. A. (1978). J . Biol. Chem. 253, 9088-9096. Vandekerckhove, J., and Weber, K. (1978). Nature (London) 276, 720-721. Vandekerckhove, J., and Weber, K. (1980). Narure (London) 284, 475-477. Vandekerckhove, J., and Weber, K. (1981). Eur. J. Biochern. 113, 595-603. Vanden Driessche, Th. (1979). Ann. Biol. 18, 417-445. Vaudaux, P. (1976). J. Prorozool. 23, 458-464. Vaudaux, P. E., Williams, N. E., Frankel, J., and Vaudaux, C. (1977). J. Prorozool. 24,453-458. Viguts, B., and Mtttnier, G. (1983). Eur. Conf. Ciliare Biol., Srh, GenPve. Viguts, B., Grolitre, C. A., Sbnaud, J., and Grain, J. (1983a). Eur. Con$ Ciliare Biol.. 5th. GenPve. Viguts, B., Mtttnier, G., and Grolitre, C. (1984). Biol. Cell. 51,67-78.

THE CYTOSKELETON IN PROTISTS

249

Vigubs, B., MCtCnier, G., Grolibre, C., Grain, J., and SCnaud, J. (l983b). J. Protozool. 30,74A. Viguts, B., MCttnier, G., Grolitre, C., Grain, J., and SCnaud, J. (1985). J. Protozool. 32, 38-44. Vivier, E., and Petitprez, A. (1972). Protisrologicu 8, 199-221. Walker, M. H., Mackenzie, C., Bainbridge, S. P., and Orme, C. (1979). J. Protozool. 26, 566574. Walkosz, R. A. (1982). J. Protozool. 29, 472. Warner, F. D. (1971). J. Ultrustruct. Res. 35, 210-232. Warner, F. D. (1974). In “Cilia and Flagella” (M. A. Sleigh, ed.), pp. 11-37. Academic Press, London. Wehland, J., Weber. K., Gawlitta, W., and Stockem, W. (1979). Cell Tissue Res. 199, 353-372. Weihing, R. R., and Kom, E. D. (1972). Biochemistry 11, 1538-1543. Williams, N. E., and Bakowska, J. (1982). J. Protozool. 29, 382-389. Williams, N. E., and Frankel, J. (1973). J . Cell B i d . 56, 441-457. Williams, N. E., Vaudaux, P. E., and Skriver, L. (1979). Exp. Cell Res. 123, 31 1-320. Williams, N. E., Van Bell, C., and Newlon, M. (1980). J . Protozool. 27, 345-350. Witman, G. B., Plummer, J., and Sander, G. (1978). J. Cell Biol. 76, 729-747. Wolfe, J. (1970). J. Cell Sci. 6, 679-700. Woolley, D. E. (1972). Arch. Biochern. Biophys. 150, 519-530. Wright, M., Moisand, A., and Mir, L. (1979). Protoplusmu 100, 231-350. Wright, M., Mir, L., and Moisand, A. (1980a). Protoplusma 103, 69-81. Wright, M., Moisand, A., and Mir. L. (1980b).Protoplusma 105, 149-160. Wright, M., Moisand, A., and Oustrin, M. L. (1982). Protoplusmu 113, 44-56. Wright, R. L., Chojnacki, B., and Jarvik, 1. W. (1983). 1. CeNBiol. 96, 1697-1707. Yamada, K., and Asai, H. (1982). J. Biochem. 91, 1187- 1195. Yamamoto, K., Pardee, J. D., Reidler, J., Stryer, L., and Spudich, J. A. (1982). J . Cell Biol. 95, 71 1-719. Yamin, M. A,, and Tamm, S. L. (1982). J. Cell B i d . 95, 589-597. Yasuda. T., Numata, O., Ohnishi, K., and Watanabe, Y. (1980). Exp. Cell Res. 128, 407-417. Yen, S. H., Liem, R. K., Jenq, L. T., and Shelanski, M. L. (1980). Exp. CeNRes. 129,313-320. Zimmer, B., and Werz, G. (1981). Exp. Cell Res. 131, 105-113.

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INTERNATIONAL REVIEW OF CYTOLOGY.VOL. 104

The Electrical Dimension of Cells: The Cell as a Miniature Electrophoresis Chamber ARNOLDDE LOOF Zoological Institute of the University, B-3000 Leuven, Belgium

I. Introduction It was in 1893 that the famous Belgian chemist Ernest Solvay donated a new Institute of Physiology to the University of Brussels. This institute was partly dedicated to the study of bioelectricity. During the inauguration ceremony he gave a remarkable lecture entitled “Observations gtnkrales sur le r d e de 1’ClectricitC dans les phtnomhnes de la vie animale.” He stated that “the phenomena of life can and should be explained by the action of only physical forces which govern the Universe, and that, among these forces, electricity plays a predominant role” (author’s translation). At this moment, it can indeed be stated that all active animal, plant, and prokaryotic cells-thus not just excitable cells-have an “electrical dimension,” the importance of which may be much greater than is generally realized. It has also become clear, especially as the result of the formulation of the ionic theory on resting and action potentials in excitable cells by Hodgkin-Huxley and Katz, that in biological systems electric current is carried by ions and not by electrons. Membrane potentials, the rather static aspect of the electric dimension of cells, can be accurately measured thanks to the intracellular glass microelectrode technique. The more dynamic aspect of bioelectricity, namely the ability of at least some cells to drive self-generated electric currents through themselves, also became much easier to investigate thanks to the development of the very sensitive vibrating probe technique by Jaffe and Nuccitelli (1974). In this review, which cannot possibly cover all aspects of electrophysiology, the concept that a cell in which the ion pumps-ion channels are asymmetrically distributed over the plasma membrane can be a powerful miniature electrophoresis chamber with all the properties inherent to this will be focused on. For readers not familiar with electrophysiology, some preliminary data on the ionic theory on resting membrane potential are induced. 25 I Copyright 8 1986 by Academic Press, Inc. All rights of reproduclion in any form reserved.

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11. Structure of the Plasma Membrane and the Generation of the Transmembrane Potential and Transcellular Currents

A. CONSTITUENTS OF THE PLASMAMEMBRANE The plasma membrane, also known as the cell surface membrane, is a barrier of highly selective permeability which regulates the relationship between the cell’s internal activities and the environment. It is a complex lipid-proteinaceous-carbohydrate structure. Its selective permeability to ions makes it possible for a potential difference to be built up between its cytoplasmic and external sides. Under certain circumstances, the cell can drive an electrical current, carried by ions, through itself. The available literature on membrane structure is so vast that it is impossible to summarize it here. Only a few data are given. Excellent introductions to membrane structure and function and to the bioelectrochemistry of cell surfaces have been written by Finean ef al. (1979) and Dolowy (1984). The widely accepted model for the structure of the cell membrane, formulated by Singer and Nicholson (1972), assumes it to be a lipid bilayer (Danielli and Davson, 1935), in which fully active protein molecules are embedded by their hydrophobic parts (Fig. 1). Calorimetric and nuclear magnetic resonance studies have detected a “bound” water component of the order of 30% of the dry weight of the membrane. In addition, X-ray diffraction methods have established the water content to be essential for the integrity of membrane lipoprotein structure. Lipids of the cell membrane are amphiphilic molecules. They feature both a hydrophobic hydrocarbon region and a polar head group. They can be grouped into three categories: sterols (such as cholesterol), glycerophospholipids (e.g., phosphatidic acid, phosphatidylserine, etc.), and neutral lipids (sphingosines, cerebrosides, etc.). Not only does the lipid composition of membranes differ from cell type to cell type, but within the same cell the plasma membrane, nuclear envelope, endoplasmic reticulum, Golgi system, mitochondria, and other membrane structures all have different lipid compositions. Membrane lipids of mitochondria and chloroplasts resemble those of bacteria and blue-green algae more than those of eukaryotes. The membrane dry mass comprises 20-80% protein, depending on the cell type. Most membranes, myelin and liver membranes being exceptions, usually contain more protein than lipids (Singer, 1975). There are two classes of cell membrane proteins. The intrinsic proteins are embedded in the lipid layer while the extrinsic are merely adsorbed at the cell surface. Amino acid sequence analysis of several intrinsic proteins showed that they all have segments of polypeptide chains with a preponderance of nonpolar amino acid residues. It is assumed that, in the membranes, these domains are normally accommodated in the nonpolar environment provided by the hydrophobic portions of lipids. Some

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FIG. I . An artist’s impression of the structure of the plasma membrane with emphasis given to the localization of ion pumps (P)and channels (Ch). These are protein structures which are thought to span the lipid bilayer. A Na+ ,K+-ATPase and an ion channel which opens when a hormone (H) binds to its receptor (Re) are represented. Ion pumps or channels might perhaps be anchored to elements of the cytoskeleton, here schematically represented as actin (A). Inset: the membrane behaves electrically as a resistance (Rbcapacitance (C) network, shunted in parallel. E is the potential difference over the membrane. Some ideas of Singer and Nicholson (1972) and Chiabrera et a/. (1984) were used to make this drawing.

intrinsic proteins, such as the very hydrophobic phosphorylase of Staphylococcus aureus, may be almost entirely immersed in the membrane interior. Most intrinsic proteins, however, are in contact with the aqueous environment of the cell’s interior and/or exterior. These contact sites are predominantly polar. Some protein structures such as ion pumps and channels most probably span the membrane. The functions of membrane proteins are numerous: they include the maintenance of the shape and motility of the cell by means of cytoskeletonforming proteins, transport of various substances across the cell membrane, participation in the cell’s metabolism via proteins possessing enzymatic activity, regulation of the cell metabolism and function by hormone receptor proteins, participation in immunological processes by proteins acting as antigens, etc. (Dolowy, 1984).

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Mammalian cell membranes contain 1-8% carbohydrate, this figure being about 25% in amoebae (Finean et al., 1979). Carbohydrates may be covalently linked to lipid as well as to intrinsic protein components, not only those of the plasma membrane but also those of the subcellular organelles. Differences in carbohydrate composition between different areas of the plasma membrane of rat hepatocytes have been demonstrated by binding studies with plant lectins (Kawakami and Hirano, 1984), while agglutinin binding led to the definition of two different endomembrane compartments in several mammalian tissues (Torrisi and Pinto da Silva, 1984). The major carbohydrate residues in the plasma membrane of mammalian cells are sialic acid, galactose, mannose, sometimes glucose, L-fucose, N-acetylgalactosamine, and N-acetylglucosamine. Sialic acid has been identified as one of the principal contributors to the negative charge on cell surfaces. Many heterosaccharides are essential to the functioning of some recognition sites on cell surfaces including receptors, antigenic determinants, and binding sites for lectins and agglutinins and enveloped viruses. In the plasma membrane sugars are present exclusively on the outer cell surface. They usually are strongly negatively charged, while it may be assumed, as an approximation, that intrinsic membrane proteins have no high net electric charge. During the recent decade a substantial amount of research has been done on the orientation and distribution of proteins and lipids in the membrane. Etemadi (1980a) reviewed the techniques employed for the assessment of the asymmetric distribution and orientation of membrane proteins and lipids (Etemadi, 1980b). Such methods involve immunological techniques, lectins, functions of proteins and their perturbations, chemical reagents, enzymatic isotopic labeling, enzymatic cleavage of membrane proteins, and physical techniques. Asymmetry of orientation is explicitly or implicitly admitted for a variety of membrane proteins. For lipids there seems to be no rule for the unequal distribution between the two membrane leaflets of different membranes. In some cases, depending on the methods used, contradictory results have been reported (Etemadi, 1980b). From biochemical data, it is well established that the cell surface carries a net negative charge (Dolowy, 1984). Fixed surface charges result in the formation of a diffuse electrical double layer in which the charge at the surface is balanced by charges of opposite sign in the medium and by the dipole moment of water molecules immediately adjacent to the surface. In the context of this article, no further emphasis will be laid on membrane structure and surface phenomena. The fluidity of the plasma membrane, the asymmetrical distribution of ion pumps and channels, and the consequences of this are believed to be of prior importance and will therefore be more fully discussed. First we will summarize the ways by which cells manage to build up a potential difference of the order of 10-200 mV over their plasma membrane. These, at first sight, low values do in fact correspond, taken into account that the plasma membrane is very thin, to a potential difference of the order of 100,OOO

THE ELECTRICAL DIMENSION OF CELLS

255

V/cm, a field strength greater than that usually encountered by engineers and sufficient to cause dielectric breakdown in many materials (Moore, 1983). AND PIEZOELECTRICITY B. OHM’SLAW,CAPACITANCE,

Before discussing the electrical properties of biological membranes, it is perhaps useful to remind the reader of the relations which are used to describe electricity in physical systems. Ohm’s law states that V = IR where V is the potential difference (in volts), I is the current (in amperes), and R is the resistance (in ohms). The reciprocal of the resistance is called conductance. Ohm’s law is also valid in biological systems although here in general current is carried by ions and not by electrons as in conductors. In the membrane, R is regulated via ion channels while ion pumps help to build up the potential difference over the membrane. A capacitor consists of two plates of conducting material separated by an insulator. If a potential V is applied across the capacitor, a quantity of charge Q,proportional to the potential difference, builds up on the plates of the capacitor. This is given by the formula Q = VC, where C is the capacitance. When the voltage changes, charge flows away from one plate into the other with a current I = C(dv/dt)where dvldt is the rate of change of voltage with time. The plasma membrane, mainly because of its lipid bilayer structure which acts as the insulator, behaves as a capacitor with a capacitance of the order of 1 kF/cm2. A simple model to represent the electrical properties of the resting plasma membrane is given in Fig. 1: the membrane behaves electrically as a resistance and capacitance in parallel. Upon mechanical deformation, some materials become electrically polarized or they undergo a deformation when placed in an electric field. When the effect is linearly proportional to the cause, the phenomenon is called piezoelectricity. The material is said to have piezoelectric properties. Piezoelectricity has been found not only in many inorganic crystals but also in various biological materials, including bone, tendon, dentin, ivory, trachea, aorta, intestine, elastin, nucleid acids, and wood. Fibrous proteins are the principal source of biological piezoelectricity (review Zimmerman, 1982). This form of electricity may be especially important for the proper functioning of bone. ON C. THEORIES

THE

ORIGIN OF THE RESTING MEMBRANE POTENTIAL Em

1. The Ionic Theory a. Intracellular Perjlusion Experiments on Squid Axons. Baker et al. (1962a,b) squeezed the axoplasm out of the giant squid axon, so that a flattened tube was left consisting mainly of the axon membrane, the Schwann cells, and the endoneural sheath. This tube was reinflated by filling it with a perfusion fluid

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ARNOLD DE LOOF

isotonic to sea water. When the perfusion solution was the same as that outside the membrane, the membrane potential, measured with a glass microelectrode in the tube and the ground electrode in the external medium, was within 1 mV of zero. When the external solution was sea water and the perfusion solution was an isotonic potassium chloride solution, the potential difference was about 55 mV, the inside of the tube being at negative potential (indicated as -55 mV). Isotonic solutions of other potassium salts, K2S0, or K isethionate, produced (resting) potentials of the order of -68 and -59 mV, respectively. When the isotonic KCl perfusion fluid was replaced by an isotonic NaCl solution, the potential fell to near zero. When the perfusion fluid was isotonic NaCl solution and the external solution isotonic KCI, the potential difference was now +40 to +60 mV (inside of tube positive). The transplasma membrane potential (Em)of the untreated axon is close to the value obtained with isotonic KCl as the perfusion fluid. The concentrations of the major ions in squid axoplasm are K + (400 mM), Na+ (50 mM), C1- (40-150 mM), Ca2+ (0.4 d), Mg2+ (10 mM), isethionate (250 mM), and other organic ions (about 110 mM). In hemolymph they are K + (20 mM), Na+ (440 mM), C1- (560 d), Ca2+ (10 mM), and Mg2+ (54 mM) (Aidley, 1981; simplified after Hodgkin, 1958). In other cell types or organisms, these values may be different but it is a general feature that the K concentration inside the cell is usually much higher than in the external medium. The Na+ concentration is usually much lower in the cytoplasm than in the blood. Some insects, however, have very high K concentrations in their hemolymph (Florkin and Jeuniaux, 1974) which imposes special adaptations, e.g., for impulse conduction in neurons. Part of the ions in the cytoplasm may be bound to macromolecules and therefore ion activities are more relevant than total ion concentrations. The activities of some ions can be measured with ion selective electrodes. The usual situation is that the inside of a cell is electronegative to the outside by some 20-100 mV in animal cells, and to 200 mV and over in plant cells and in some bacteria. When the measuring electrode is slowly retracted from the cell, the potential difference does not usually change greatly as long as the tip of the electrode is still in the cytoplasm. The potential suddenly falls to zero at the moment that the electrode leaves the plasma membrane: this clearly shows that it is the plasma membrane which is the site of this potential difference (resting potential). Although theoretically much higher membrane potentials might be possible in nature, the values are restricted to a maximum of about 250 mV for “safety” reasons. At the application of a potential equal or higher than 500 mV, the membrane is either fractured or disturbed (Martirosov, 1983). b. The Principle of the Concentration Cell. Let us assume that compartment 1 of the system shown in Fig. 2 contains a higher concentration of an electrolyte XY in aqueous solution than compartment 2. 1 is separated from 2 by a differen+

+

AwlBF THE ELECTRICAL DIMENSION OF CELLS

X+

25 7

A'I

1 2 1 2 FIG. 2. (A) The principle of a concentration cell. Compartment I , which is separated from compartment 2 by a differentially permeable membrane, contains a higher concentration of diffusible ions X . As more X diffuse from 1 to 2, a potential difference between the two compartments builds up. (B) Donnan equilibrium system. In the cytoplasm, organic ions (A-), which cannot pass through the membrane by simple diffusion, are also present. The Donnan rule states that the product of the concentrations of the diffusible ions in one compartment is equal to the product of the diffusible ions in the other compartment, at equilibrium. This is given by the formula [ X + ] , [ Y - ] , = [X+],[Y-],. Redrawn after Aidley (1981). +

+

tially permeable membrane, through which X + can pass but Y- cannot. As a result, a number of cations X + of compartment 1 will diffuse down their gradient to compartment 2. Thus net positive charges are taken away from 1 and concentrated in 2: a potential difference between 1 and 2 is set up. As more X + move from 1 to 2, the potential difference becomes higher, or, in other words, the electrical gradient increases. But, the higher the potential difference, the more difficult it becomes for X + to move against the electrical gradient. After some time an equilibrium position is reached at which the electrical gradient (which tends to move X + from 2 to 1) just balances the concentration or chemical gradient (which tends to move X + from compartment 1 to 2). The term concentration cell is derived from the difference in concentration of the diffusible ion in the two compartments. The potential difference over the membrane is given by the Nernst equation. It is measured as the potential of compartment 2 with respect to compartment 1.

where E = potential difference in volts, R = the gas constant (8.314 J degree Kelvin- I mol- I , T = absolute temperature, z = the charge of the ion (e.g., K = + 1, C1- = - 1 , Ca2+ = +2, SO:- = -2), F = Faraday's constant (96,500 C/mol), and [XI,and [XI, = the molar concentrations, or, more strictly, the activities of X + in compartments 1 and 2, respectively. More simply, the Nernst equation at 18°C can be given, now in millivolts, as +

For the mathematical deduction of this formula, see Richter (1979), Petersen (l980), or Aidley (1981).

258

ARNOLD DE LOOF

For the reader not familiar with electrophysiology a few examples follow, with concentrations of K+ and Na+ as found in squid hemolymph (X,) and axoplasm 58 [4403 = +55 mV = sodium equilibrium or reversal potential ENa+ = - log,, 1 POI 58 EK+ = -

[2ol

1 loglo [400]

58

4,-= -1 log,,

= -75 mV = potassium equilibrium potential

[5601 = -43 mV [1ool

= chloride equilibrium potential

c. Application of the Principle to Living Cells. In these examples, only one ionic species was considered at a time, and it has been assumed that the membrane was equally permeable to the three ions. However, in the cell several ions are simultaneously present. Furthermore, the plasma membrane of living cells is a differentially permeable membrane which is much more selective to K than to Na+ or C1- ,as was shown by diffusion experiments with radiolabeled anions and cations. This can be taken into account by introducing permeability coefficients P, measured in pm/second. If PK+is taken as 1, then PNa+is of the order of about 0.01 to 0.05 and so is P,,-. If only the major ions involved are taken into account, if several assumptions formulated in the “constant field theory” of Goldman (1943;see also Hodgkin and Katz, 1949) are also taken into account, and when there is no current flowing through the membrane, a fairly good approximation of the resting potential (Em) over the plasma membrane of a given cell is obtained by the following equation, which is known as the Hodgkin-Katz-Goldman equation: +

In our example

+

1 [20] 0.01 [440] 1 [400] 0.01 [50] ~ 5 1= -70 mV = 58 log,, [4061

Em = 58 log

lo

+

+ 0.01 [loo]

+ 0.01 [560]

Due to the small values of PN,+ and P,, - , Na and C1- do not significantly contribute to the resting potential. In this case, a cell behaves almost like a K + electrode. However, when the permeability of the plasma membrane for Na+ increases, Na ions will rapidly enter the cytoplasm and cause depolarization of the membrane (Le., make the cytoplasmic side less negative). Increased influx of C1- will cause a hyperpolarization (i.e., make the cytoplasmic side more negative), etc. Any change in flux of a particular ion through the membrane will be +

+

259

THE ELECTRICAL DIMENSION OF CELLS

associated with a change in transmembrane potential. Similarly, chemicals or physical factors which change the transmembrane potential cause changes in ratio(s) of the ionic concentration(s) in the cytoplasm versus the external medium. 2. The Association-tnduction Hypothesis In the theory of resting potentials of Hodgkin-Katz, it is essential that at least a large portion of the cellular K and water exist in the free state. If all the K in the cytoplasm is not in the free state, the measured potential difference over the membrane cannot be due to the concentration gradient of K + and has to be dependent upon another principle. This is the essence of the association-induction (AI) hypothesis of Ling (1982). The basic assumptions of this alternative model are the following. +

+

1. The smaller hydrated K is adsorbed preferentially over hydrated Na due to the (electrostatic) adsorption energy which favors K + adsorption to certain fixed anionic sites such as the f3 and y carboxyl residues of aspartic and glutamic acid. This assumption is well documented for nonliving fixed charge systems such as soils, permutites, sulfonate, and other anion exchange resins and also for living material. In this respect, Edelmann (1977, 1978, 1980, 1981) demonstrated that in striated muscle the cell K + is preferentially localized at the A bands and therefore probably not free. Myosin, the distribution of which in voluntary muscle is restricted to A bands, accounts for 60% of the cell’s f3 and y carboxyl residues. 2. The cell’s resting potential is a surface adsorption phenomenon. In the cell surface-in essence a two-dimensional version of a fixed charge system-the density and nature of the anionic sites on and very close to the cell’s surface on one hand and the concentration of counterions (K ) that are absorbed on these fixed charges at the cell surface on the other hand determine the cell’s resting potential. Elaboration of this model for resting potentials has led to the following equation in which KNaand K, are the adsorption constants for Na+ and K + at the surface, respectively (Ling, 1960): +

+

+

Although only a portion of the relations predicted by the Hodgkin-Katz-Goldman equation is included in this equation, most experimental evidence in favor as well as evidence which is in conflict with the Hodgkin-Katz theory may be explained (Ling, 1982). 3. The major route of entry of cations is the adsorption-desorption route which involves three steps: (1) adsorption onto a fixed anionic site, (2) libration around those fixed ionic sites, and (3) desorption of the first cation. The saltatory

260

ARNOLDDELOOF

route of penetration through the interstices of nonpolarized water is a second way of entry. 4. A large part of the cell water exists as polarized multilayers around a matrix of extended protein chains and therefore has a reduced solvency for Na+ , sugars, and amino acids. This has been demonstrated for aqueous solutions of isolated proteins and for synthetic proteins. In this regard, polarized water at the cell surface may be regarded as the seat of cell surface selective permeability. 5 . Physiologically active compounds such as hormones, drugs, ATP, Ca2+, etc., collectively called cardinal adsorbents, interact with key cardinal sites on proteins. In this way cardinal adsorbents control (1) permeability through modifying the polarized state of cell water and (2) the selective adsorption of K + . Both effects of cardinal adsorbents have been explained in terms of (anti)cooperativity, inductive, and direct electrostatic effects (Ling, 1969).

In our opinion, the A1 hypothesis draws attention to phenomena which have thus far been neglected by membrane physiologists, e.g., the existence of adsorbed ions stresses the dilemma of ionic activities versus ionic concentrations and the physiological state of cellular water. However, on the other hand, the A1 hypothesis undervalues the evidence in favor of the essential role of ion pumps for generating Em according to the widely accepted Hodgkin-Katz theories on the resting potential and on the theories of Hodgkin and Huxley (1939, 1952a-d) on action potentials. Habib and Bockris (1982, 1984) who, like Ling, point to certain experimental discrepancies within the ionic theory on resting membrane potentials, presented an electrodic model to explain charge transfer across biological membranes. The evidence they obtained suggests that in some biological situations the transfer of electrons across interfaces influences the potential difference across the cell membrane. D. MECHANISMS FOR MAINTAINING INEQUALITY OF THE DISTRIBUTION OF DIFFUSIBLE IONS 1. Donnan Equilibrium System

In the cytoplasm there are also indiffusible organic ions present, which leads to an inequality of the distribution of the diffusible ions according to the Donnan rule which is in fact also based on the principle of the concentration cell (Fig. 2). This rule states that, at equilibrium, the product of the diffusible ions in one compartment is equal to the product of the concentrations of the diffusible ions in the other compartment. Boyle and Conway ( 1 941) measured the intracellular potassium and chloride concentrations in frog muscle after equilibration with various solutions for 24 hours at 2-3°C and they found that their data well fitted the Donnan equilibrium hypothesis [K+Ii[C1-Ii = [K +],[Cl-], at all external potassium concentrations above about 10 mM. For details on the deduction of

26 1

THE ELECTRICAL DIMENSION OF CELLS

this formula, see Aidley (1981). A second mechanism by which inequalities in ionic distribution arise is active transport.

2 . Active Transport a. ton Pumps. Active transport systems capable of accumulating or extruding ions against a gradient of electrochemical potential are widespread in the membranes of plant, animal, and bacterial cells. Two different types of active transport may be distinguished. Primary active transport systems (or “ion pumps”) utilize redox energy, ATP hydrolysis, or light as energy sources. Secondary active transport systems (review West, 1980), better known as co- or countertransport systems, are driven by electrochemical gradients generated by primary active transport. Cotransport or symport is the flow of two solutes in the same direction; countertransport or antiport is the flow in opposite directions. For example, there exists in most excitable membranes a coupled exchange of Na+ for Ca2+ (Barzilai er al., 1984; Hale et al., 1984). In heart cells this system is normally used for outward transport of Ca2+ but it is reversible (Bridge and Bassingthwaighte, 1983). The H+/Ca2+ antiport in membrane vesicles of peas is another example (Rasi-Caldogno et al., 1982). Kinetic models of cotransport systems have been reviewed by Turner (1983). Ion pumps are supposed to be large transmembrane proteins. When an appropriate energy source is available the protein goes through a cycle of transitions, and thereby translocates one or several ions through the membrane. ATP is used by two different classes of ion pumps, which differ from each other in several aspects. Class I contains the proton translocating ATPases of mitochondria, chloroplasts, and bacteria. To Class I1 belong the Na+ ,K+-ATPase of animal cells and the Ca2+-ATPase of sarcoplasmatic reticulum. Recently, Specht and Sweadner ( 1984) showed that two molecular forms of Na ,K -ATPase could be isolated from the central nervous system. The proton translocating ATPase of Fungi and plants has structures and reaction mechanisms typical of Class I1 but it is sensitive to dicyclotrexylcarbodiimide as is Class I ATPase. Their stoichiometry is 1 H /ATP as compared to 2-4 H /ATP for Class I (Goffeau and Slayman, 1981; Maloney, 1982; Serrano, 1983, 1984; Lauger, 1984). Ion pumps could, in principle, be electroneutral as, e.g., the in siru gastric proton pump (Malinowska et af., 1981), but most of those so far studied are electrogenic. When, e.g., a Na+,K+ pump should take up one K + from the external medium, transport it through the membrane to the cytoplasm and simultaneously transports one Na from the cytoplasm to the external medium, there is no net transfer of charge across the membrane. Such an ion pump is electroneutral: it has no effect on the membrane potential since for each positive charge which enters the cell, another charge is extruded. The Na+ ,K+-ATPase, first described by Skou (1960), proved not to be an electroneutral pump. The enzyme seems to be composed of two different subunits: the larger catalytic and +

+

+

+

+

262

ARNOLD DE LOOF

ouabain binding (review Anner, 1985) a unit and a smaller p subunit of unknown function. The data so far available suggest that the enzyme spans the plasma membrane, the site of ouabain binding facing the cellular environment and the site of ATP hydrolysis facing the cytosol (Fig. 1). The vectorial exchange of Na+ and K + across the membrane is supposed to include the following steps. Binding of three Na+ ions to high affinity sites on the inner aspect of the membrane supports phosphorylation of the enzyme by ATP. This results in a subsequent change in the conformation of the enzyme which exposes the ion binding sites to the cellular environment. Here, the affinity of these sites for ions is then altered so that the three Na ions are displaced to the external medium by two external K ions. Dephosphorylation of the enzyme results in the return of its conformation to the original condition, bringing two bound K + ions into the cell. Per molecule ATP hydrolyzed there is thus a net extrusion of one positive charge out of the cell. The Na+ ,K+-ATPase makes the inside of the cell more negative than can be expected from the Nernst equation: it is an electrogenic ion pump (Robinson and Flashner, 1979; Cantley, 1981; Jgrgensen, 1982). In the squid axon, the pump contributes 1-2 mV to the resting membrane potential and generates a net outward electric current of a few FA cm-2. It can be forced to run backward (De Weer and Rakowski, 1984). It is not completely clear whether the 3 Na+ -2 K + stoichiometxy is fixed or variable or if it is dependent on conditions such as membrane potential, available free energy, and steepness of the ion gradients. A typical feature of insect Na+ ,K+ -ATPase is its apparent inaccessibility to ouabain (Towle, 1984). In cultured chick myotubes, up to 400 Na+,K+-ATPase related particles/ p,m2 of plasma membrane were counted (Pumplin and Fambrough, 1983). Bacteriorhodopsin, a retinal-containing protein in the plasma membrane of Halobacterium hulobium, acts as a light-driven, electrogenic ion pump (Stoeckenius et al., 1979). Schobert and Lanyi (1982) found that another retinal-containing protein of the same bacterium utilizes light for uphill chloride transport. The Cl- pump in the plasma membrane of the giant unicellular marine alga Acetabularia is electrogenic (Wendler et al., 1983) as is the Mgz+/KClATPase of plant plasma membranes (Sze and Churchill, 1981). The H+ translocating ATPase of Fungi and plants is also electrogenic (Serrano, 1984). Electrogenic ion pumps convert chemical into electrical energy. Pumping activity may be modulated by an electric field which is present in the membrane. These pumps create a transmembrane electric field which may modulate some metabolic activities of the cell or other transport processes. The thermodynamic and kinetic properties of electrogenic ion pumps were recently reviewed by Lauger (1984). b. Ion Channels: Types and Gating Mechanisms. There is much evidence that ions cross the plasma membrane through selective channels (ion selective, valence selective, nonselective; Latorre and Miller, 1983) which are most proba+

+

263

THE ELECTRICAL DIMENSION OF CELLS

bly proteinaceous in nature. The development of the extracellular patch clamp method by Neher et a f . (1978) has substantially contributed to the characterization of the different channel types as this method allows measurement of currents through single membrane channels. Most, if not all, of the channels seem to have at least two conformations: one that allows the passage of ions through the channel and one that does not. Several recent studies indicate that sodium and calcium channels contain substrate proteins which are phosphorylated, and that phosphorylation of those channels may be responsible for regulating ion flux. The available evidence suggests that these proteins are phosphorylated by cyclic AMP-dependent kinases (Osterrieder et al., 1982; Costa et al., 1982; Bittar et a f . , 1982). The selectivity filter is a region of the channel which limits the ions that pass through the open channel on the basis of charge, the ability to form hydrogen bonds, and size. A list of the radius of the ion and its primary hydration shell for the most common inorganic ions is given by Edwards (1982). Whether ion selectivity is high, moderate, or low is largely, but not exclusively, determined by the hydration of the ion involved: anions are relatively unhydrated and cations are usually more hydrated. Edwards (1982), who reviewed the selectivity of ion channels in nerve and muscle, estimates that the number of channels in nature is not too large; probably of the order of a dozen. As still more refined methods are developed, new channels may be discovered, or those already described may prove more complex than hitherto supposed. It is also possible that channels which show similar selectivities are not identical in structure. Chloride channels. In order to explain the many selectivity data obtained for anions, a minimum of three channels is required: a small selective one to fit the frog muscle data, a larger one to fit the results obtained with cat spinal motor neuron and probably a number of other cells, and a still larger one to fit the data from the cortical neuron. A few additional anion channel types may be present in other cell types. The CI- channel, as do the large cation channels, admits all ions up to a certain size, with no cut-off on the small side (Edwards, 1982). Sodium channels. Most data so far obtained suggest that sodium channels in nerve and muscle cells are functionally similar across wide phylogenetic boundaries. They were usually thought to represent a single, homogeneous population that initiates the action potential at the threshold and unerringly transmits it along the surface membrane (review Edwards, 1982; Agnew, 1984). However, there are several indications that distinguishable populations of Na channels exist, the most recent described being Na+ threshold channels (Gilly and Armstrong, 1984). In chick heart and brain, two sodium channel subtypes have been proposed (Rogart et al., 1983). Data from vertebrates and molluscs suggest a highly conserved molecular structure during evolution. Tetrodotoxin, saxitoxin, and derivatives are selective blockers (Agnew, 1984). They bind to a site accessible from the outside of the membrane, causing a blocking of sodium currents. These +

264

ARNOLD DE LOOF

toxins have been used to follow the isolation of sodium channel protein from different tissues. The principal constituent seems to be a large glycoprotein of 250,000 Da. Four properties are essential to Na+ channel function: permeation selectivity, activation and inactivation mechanisms, and high rates of ion transport. At resting potentials the channel is closed but on rapid depolarization it is activated to permit a very fast ion flux (at least lo6 ions per second per single channel at normal membrane potentials). The high single channel conductance allows a channel to efficiently depolarize large areas of membrane. The generalization that invertebrate muscles lack Na+ channels no longer holds (Schwarz and Stiihmer, 1984). Models, mimics, and modifiers of sodium channel gating have also been recently reviewed by French and Horn (1983). Potassium channels. For this channel type, which is also quite selective, the data for starfish egg, muscle, and nerve are more or less consistent with the view that only a single selectivity filter exists (Edwards, 1982). However, additional filters may be required to explain some discrepancies. So far, specific high affinity blocking agents have not been found. Intracellular ATP directly blocks K + channels in pancreatic B cells (Cook and Hales, 1984). According to Thompson and Aldrich (1980) there are four major types of potassium channels in nervous tissue: (1) the outward or delayed rectifier, (2) the inward or anomalous rectifier, (3) the fast transient potassium channels, and (4) the Ca2+-activated potassium channels. Since then, maxi-K+ channels have also been described (review Latorre and Miller, 1983) and also a novel class of ionic channel, namely the Na+-activated K + channel (Kameyama et al., 1984). When the resistance of a conductor is not independent of the voltage across it, the conductor is said to show rectification (Katz, 1949; Aidley, 1981). The effect that depolarization of electrically excitable membranes results in a large increase in the permeability of the membrane to K+ ions is known as delayed rectification. Inward or anomalous rectification means that the conductance of the membrane for outward current is much less than the conductance for inward current (for details see Latorre and Miller, 1983). Hille and Schwarz (1978) calculated that at least one K ion can pass through an open K + channel per microsecond. There are variations in the relative amounts of the “classical” 4 types among similar cell types in different organisms, among the different cell types of the same organism, and there may even be regional differences in the distribution of potassium permeability units within a single cell, e.g., in a neuron. The relative importance of the different types of potassium channels among different cells is a major factor shaping the excitability of the cell. Calcium channels. Acetylcholine may open channels through which different ion types, calcium included, can pass. These are not considered as Ca2+ channels. In a wide variety of cells, the membrane’s permeability to Ca2+ increases upon depolarization of the membrane potential. The “Ca2+ channel,” first found in invertebrate muscle, may be more universal than the Na+ channel, but

265

THE ELECTRICAL DIMENSION OF CELLS

is much more difficult to characterize, mainly because no ideal preparation has so far been found for its study. According to Edwards (1982) there appear to be no fewer than three Ca2+-selective filters. Hagiwara and Byerly (1981) state that the selectivity of the Ca2+ channel cannot be measured in terms of relative permeation, as is done in other channel types, because the reversal potentials for currents through the Ca2+ channel cannot be measured. Instead, the Ca2+ permeation mechanism is characterized in terms of a model in which polyvalent cations bind to a membrane site from which they can then pass through the membrane. Although there is in fact little evidence to establish its channel nature, this voltage-dependent permeation mechanism for Ca2 is referred to as the Ca2 channel (Hagiwara and Byerly, 198 I). Voltage-dependent Ca2 currents can be modulated by certain external chemical messengers such as adrenalin, noradrenalin, serotonin, GABA, somatostatin, and enkephalins (Edwards, +

+

+

1982).

Larger cation channels. At least two large cation selective channels seem to exist. Similar selectivities are found for the acetylcholine (ACh) channel at the frog neuromuscular junction and for the glutamate channel at the crayfish neuromuscular junction. The ACh channel of the eel electroplaque is larger (Edwards, 1982; Latorre and Miller, 1983). The outer membrane of mitochondria contains voltage-dependent anion-selective channels (Mannella ef al., 1983). Gating mechanisms. In general, gating of ion channels presumably involves the creation, or completion, of an aqueous pathway communicating between the aqueous solutions on the two sides of the membrane. This probably involves rapid changes in the spatial configuration of the channel molecules between conducting or “open” and nonconducting or “closed” states (French and Horn, 1983; Horn, 1984). Some channels may be opened by a number of transmitters for postsynaptic channels, while some hormones or other compounds may also be active (chemically operated channels). Some channels, like the sodium and calcium channels, may respond to changes in membrane voltage (voltage gated channels) (Hagiwara, 1983). Specific phosphorylation is also a possible gating mechanism (Latorre and Miller, 1983). The attachment of a sperm to the egg of Urechis causes the opening of sodium channels (Could-Somero, 1981). The opening of some channels is controlled by changes in intracellular ion concentrations, such as, e.g., three kinds of Ca2+-activated K + channels which play a crucial role in the regulation of membrane potential and secretion (Petersen and Maruyama, 1984). Theoretically a very large number of channel selectivity control mechanisms is possible, as was first suggested by Swann and Carpenter (1975) for transmittercontrolled channels in Aplysia neurons. How many combinations are actually used in nature remains to be determined. Some physical factors and nonaqueous solvents can also modify channel gating kinetics. A variety of different experimental approaches such as the introduction of purified channels into recon-

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stituted membranes (e.g., Ca*+-dependent K+ channel, Latorre et al., 1982; Na+ channel, Weigele and Barchi, 1982; Rosenberg et al., 1984) and the monoclonal antibody technique (Na+ channel, Casadei et al., 1984) is now being used to better understand the complexity of mechanisms involved in the gating of ion channels (for references on this topic as far as the sodium channel is concerned, see French and Horn, 1983). The study of ion channels has substantially contributed to the finding that different cell types of the same organism usually have different membrane properties, e.g., with respect to ion channels. c. TransmembraneDifference of Electrochemical Potential for Protons. In most eukaryotes, the electrochemical gradient for Na+ , generated by Na+ ,K+ATPase, is the intermediate form of energy between ATP and cotransport of nutrients with Na+ . In bacteria, mitochondria, and yeast, it is not a Na+ gradient but a H+ gradient which is used for this purpose (Boyer et d . , 1977). According to the generalized chemiosmotic hypothesis of Mitchell (1961, 1966, 1968, 1973, 1977), initially formulated to explain oxidative phosphorylation, energy derived from hydrolysis of ATP via the proton ATPase (Racker et al., 1979; Senior and Wise, 1983) or via the transport of solutes down a concentration gradient (i.e., the reverse of active transport) or via photochemical reactions is transformed into a transmembrane difference of electrochemical potential for protons, ApH . ApH depends partially on ApH, which is the pH difference across the membrane, and partially on A$, which is the actual membrane potential, whose contribution to ApH+ is a function of the external pH. The general formula is AGH+ = A$ - 2.3 (RT/F)ApH 2.3(RT/F), the factor to convert ApH units into electrical units, is 58.8 at room temperature. The “common currency of energy exchange,” particularly in the bacterial cell, is not ATP but ApH+. It is the immediate driving force for a wide range of processes in energy transducing cells and organelles (Kaback, 1983) with regard to active transport. The chemiosmotic hypothesis predicts that transport is driven by A$ (interior negative) for cationic substrates, by ApH (interior alkaline) for anionic substrates, and by A&H+ for neutral substrates (Mates et al., 1982). +

+

E. UNIFACIAL, BIFACIAL,AND MULTIFACIAL CELLS 1. The Fluidity of the Plasma Membrane The viscosity of the lipid moiety in membranes is temperature dependent. At physiologically ambient or body temperature, most of the lipid in the bilayer is supposed to be rather fluid (Peters, 1981). As a result membrane proteins will be subjected to lateral mobility and tend to distribute themselves randomly over the

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267

plasma membrane. Singer and Nicholson (1972), who proposed this ‘fluid mosaic model’ of biological membranes, based their hypothesis mainly on their data of an apparently random distribution of integral membrane proteins over the erythrocyte membrane of mammals. The model was supported by the finding that upon fusion of different tumor cells the antigens bound to the cell membrane are redistributed over the fused cells (Frye and Edidin, 1970). Another evident case is that of the collection of cell surface receptors driven into a “cap” followed by pinocytosis when lymphocytes are exposed to multivalent ligands such as lectins or antigens. Another example is the diffusion-mediated aggregation of some peptide hormone receptors into a microcluster as a prelude to endocytosis. Rhodopsin can also diffuse laterally even in unmodified membranes. There is also evidence of considerable freedom of movement of lipid molecules in membranes (Finean et al., 1979). It thus appears that a number of molecules which are intrinsic components of membranes can be redistributed within the plane of the membrane. This can be achieved either passively by diffusion, or in a directed way which involves coupling to some energy-dependent mechanism, e.g., lateral electrophoresis or by the movement of contractile filaments which are connected in some way to the cytoplasmic surface of the plasma membrane. Protein mobility in cell membranes has been extensively reviewed, most recently by Axelrod (1983) and McCloskey and Po0 (1984). A cell with a fluid membrane, in which all proteins would be able to diffuse laterally and randomly, would have a rather uniform plasma membrane. Such a cell could be called unifacial. It would be at equipotential, wherever a measuring microelectrode would be inserted. In our search through the literature, we have not as yet found an account of a cell type in which all membrane proteins can diffuse freely. 2. Restricted Lateral Diffusion of Some Integral Membrane Proteins In general, the above mentioned examples of lateral diffusion are much better known to nonspecialists than the mass of data indicating that, in many different cell types, at least some membrane proteins are restricted in their lateral mobility and are not randomly distributed. Edidin (1972) stated “that a consideration of tissue architecture and cytochemistry quickly suggests that there must be mechanisms acting to restrict mobility of membrane molecules when cells are organised in tissues.” We could add that recent data show that this view is also valid for at least some single cells, as will be discussed further. An already classical example of a nonunifacial cell is that of the concentration of nicotinic acetylcholine receptors on skeletal muscle fibers beneath the innervating nerve terminal as was already shown by Miledi in 1960 and later confirmed by Axelrod et al. (1976). Since then it has become evident that receptors for other neurotransmitters are, in general, located in a rather small area of

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postsynaptic membranes (see, e.g., Gulley and Reese, 1981). Another example is that of rhodopsin in visual rods as will be described further. Evidence has also been presented to show that lipid may be organized into domains and that such organizational heterogeneity may not only have structural but also functional significance (Karnovsky et al., 1982). In the context of this article, we will go into some detail on the distribution of ion pumps and ion channels over the plasma membrane. 3. Segregation of Ion Pumps and ton Channels in Different Cell Types a. In Epithelial Cells of Diferenriated Organisms. A major function of epithelia is the transport of ions, water, nutrients, and excretion products. Since the direction of transport is evidently very important, it is not surprising that the intrinsic membrane molecules, which carry out the transport function, have a well-defined and constant location (Sabatini et al., 1983). Epithelial cells typicallymhave (at least) two distinct plasma membrane domains, one apical and one basolateral, each with specialized functions. Such cells are (at least) bifacial. Almers and Stirling (1984) reviewed the data which have been published since 1967 on the distribution of the Na ,K -ATPase, the amiloride-sensitive Na channel, the furosernide-sensitive C1- pump, and the phlorizin-sensitive glucose pump in a variety of epithelia of vertebrates. The location of the Na+ ,K+ATPase has been visualized by autoradiography, after application of [3H]ouabain to tissue sections and by immunocytochemistry. For example, in rabbit small intestinal epithelial cells very little Na+ ,K -ATPase activity is present in the apical membrane but it is abundant in the zonula occludens and in the basolateral membrane (Almers and Stirling, 1984). This pump is present in all salt-transporting epithelia, regardless of whether they secrete or absorb NaCI. In most epithelia, the pump is located only in the basolateral membrane. This is the case for intestine, frog skin, gall bladder, urinary bladder, renal tubule, teleost gill, salivary gland, pancreas, shark and bird salt glands, sweat gland, cornea, and trachea. In epithelia of invertebrates, the bulk of the evidence also supports a basolateral location for Na+ ,K+-ATPase (Towle, 1984). In choroid plexus, retinal pigmented epithelium, and ciliary of the vertebrate eye, the Na ,K -ATPase is located only in the apical membrane. Depending on the type of epithelium, the transport direction of Na+ can be either from the apical to the basolateral membrane of vice versa. In most cases, the amiloridesensitive Na channel, the furosemide-sensitive C1- pump, and the phlorizinsensitive glucose pump are located in the apical membrane of the different epithelia. For original references to the numerous data, see the review by Almers and Stirling (1984). Epithelia contain two major diffusional pathways: a transcellular route in which ions penetrate the apical and basal membranes, and a paracellular pathway consisting mainly of zonulae occludentes (tight junctions) in series with the +

+

+

+

+

+

+

THE ELECTRICAL DIMENSION OF CELLS

269

lateral spaces between epithelial cells. There is no general rule for the permeability to ions of the zonula occludens: some epithelia are rather leaky, while in other tissues, this region may be rather impermeable to ions (Fromter and Diamond, 1972; Petersen, 1980; Hanrahan, 1984). Intermediate situations are possible. In leaky epithelia, most transepithelial diffusion of ions occurs through the paracellular route through the zonula occludens and lateral intercellular spaces. Such epithelia have typically low transepithelial potentials (V, < 11 mV), low resistances (R, < 133 Cl cm2), and maintain only small concentration gradients. In tight epithelia, most of the transepithelial diffusion takes place through the cells themselves because the combined resistance of apical and basal membranes is lower than that of the zonulae occludentes and intercellular spaces. They can build up characteristically high transepithelial potential differences (up to 100 mV) and high chemical gradients in vivo (up to 106-fold). Their transepithelial resistance is of the order of 300-10,OOO Cl cm2 (Hanrahan, 1984). Some examples of organs possessing leaky epithelia are the gallbladder (Fromter, 1972), kidney proximal tubule of Necturus (Guggino et al., 1982), rabbit intestine (Frizzell and Schultz, 1972), prawn intestine (Ahearn, 1980), and Aplysia intestine (Gerencser, 1982). Tight epithelia are present in toad urinary bladder (Reuss and Finn, 1974), toad stomach (Spenny et al., 1974), and rabbit urinary bladder (Lewis et a l . , 1976). Insect epithelia usually have properties consistent with leakiness (low resistance) and tightness (large transepithelial potential, ionic and osmotic gradients) (Hanrahan, 1984). For some time, it has been assumed that true tight junctions were absent in invertebrates and that their permeability bamer restricting the paracellular entry of ions and molecules was the result of septate junctions. Lane and Chandler (1980), however, showed that tight junctions do occur within the central nervous system of the house spider Tegenaria. The distribution of tight junctions in insects is limited (Hakim and Baldwin, 1984). b. In Epithelial Cells in the Embryo and Regenerating Tissue. When the blastoderm of an animal embryo behaves as a tight epithelium and when there is transepithelial ion transport toward the blastocoel resulting in a concentration or voltage gradient being built up (see for examples, Jaffe and Nuccitelli, 1977), an efflux of ions from the blastocoel will take place at those places where the tight epithelium becomes leaky. This seems to happen during gastrulation and in neurulae of Xenopus (Robinson and Stump, 1984). Yolk sac, amnion, and allantois compartments of the domestic fowl are neither in ionic nor in osmotic equilibrium with the blood (Simkiss, 1980). This suggested that their epithelia actively transport ions. The chick epiblast is indeed capable of unidirectional apical to basal Na transport, toward the underlying intraembryonic space (Stern and MacKenzie, 1983). The transepithelial potential difference and the short circuit current in the isolated yolk sac of the rat reach peak values of about 4 +

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mV (fetal side positive) and 20 pA cmP2, respectively (Chan and Wong, 1978). Active ion transport from the external medium through the skin of amphibians generates a potential difference over the skin, where the inside potential becomes positive with respect to the external medium. A fresh wound in the skin acts as a leak for ions and this seems to be the origin of wound currents (Borgens, 1982, 1983; Person, 1983; Section 111,F). c. In Neurons. Ion pumps and channels seem to be unevenly distributed over the plasma membrane, especially in myelinated nerve fibers. Voltage-dependent channels are unevenly distributed over soma and axon (review Poo, 1985; Waxman and Ritchie, 1985). Ca2+ channels are concentrated in soma and presynaptic terminals. They function by linking electric excitation to the release of a synaptic transmitter (exocytosis) and they may be involved in controlling some aspects of metabolism in the soma of the neuron. Na+ channels are rather sparsely distributed over the nerve cell body and dendrites. According to Dodge and Cooley (1973) the Na+ channel density in the nonmyelinated initial segment (axon hillock) of vertebrate motoneurons is as high as that of the node of Ranvier. In vertebrate motoneurons, at least, the nerve impulse originates at the axon hillock, and not in the soma (Coombs et al., 1957; Fuortes et al., 1957). In addition, Catterall(l981) also found a 7-fold higher Na+ channel density over the neurites than over the cell body of cultured spinal cord neurons. In myelinated nerve axons, Na+ channels are highly concentrated, perhaps even solely located, within the nodal zone of the node of Ranvier, which is the site of electrical excitation (Ellisman and Levinson, 1982). In the rabbit, Ritchie and Rogart (1977) estimated the number of Na+ channels in the nodes to be 12,000/pm2which is six times higher than in the frog node (Conti et al., 1976). Freeze fracture studies also showed much higher numbers of large particles-supposed to be Na+ channelsin the nodal than in the paranodal region in several animal species (Rosenbluth, 1976; Kristol et al., 1978; Tao-Cheng and Rosenbluth, 1980). The concentration gradients for membrane proteins at the transition from the node of Ranvier to the paranodal region, as shown by morphological and electrophysiological measurements, is steep. K channels are scarce or completely absent at the nodes but localized at the paranodal and internodal regions of the axon membrane (Chiu and Ritchie, 1981, 1982; review by Poo, 1985). The physiological function of K channels in the internodal and paranodal region is, as yet, not fully understood. Na+ ,K+-ATPase is relatively uniformly distributed over the entire nerve (and muscle) membranes (Poo, 1985; Pumplin and Fambrough, 1982). Almers and Stirling (1984) mention a higher concentration of Na ,K -ATPase in the nodal rather than in the paranodal region. d. In Skeletal Muscle Cells. Intrinsic membrane proteins are unevenly distributed over the sarcolemma. Almers et al. (1983) found that in frog skeletal muscle fibers, the Na+ channels seem to be concentrated in strips or oblong patches running along a fiber while the distribution of K + channels was found to +

+

+

+

27 1

THE ELECTRICAL DIMENSION OF CELLS

be irregular. A correlation between the areas of increased Na+ channel density and other cellular structures has not yet been found. Since in skeletal muscle fibers, contrary to the situation in smooth and cardiac muscle, the oblong nuclei are located just underneath the sarcolemma, a causal relationship between the Na+ channel patches and nuclear activity might well become apparent in the near future (De Loof, 1985b). This item will be dealt with further. Acetylcholine receptors are strictly confined to the “motor endplate” sarcolemma (Fertuck and Salpeter, 1976). By means of the vibrating probe technique Betz et ul. (1980) showed that the endplate seems to act as a current source to the rest of the muscle (Fig. 3). The concentration of Na+ channels is probably less at the endplate than elsewhere, while voltage-dependent Ca2 channels may be more concentrated here. The transverse tubular system of frog skeletal muscle is also an example of the specialization of distinct plasma membrane areas. Over 90% of the Ca2+ channels, about 50% of the Na+ channels and the delayed K + channels, and 75-80% of the inward rectifier channels are located here while about 80% of the Na+ ,K+-ATPase is found in the sarcolemma. The picture which has emerged shows that the ionic channels involved in the maintenance of the resting potential, namely the K + and CI- channels, are evenly distributed over the sarcolemma and the transverse tubular systems. The sarcolernmal membranes also contain channels which are involved in excitation (voltage-dependent K ,Na+ channels) (review by Almers and Stirling, 1984). Neurons and skeletal muscle cells are clear examples of cells which are multifacial with respect to the localization of intrinsic membrane proteins (review Poo, 1985). e. In Vertebrate Photoreceptors. Vertebrate photoreceptors, rods, and cones consist of an outer segment, which is connected by a narrow stalk, the ciliary junction, to the inner segment (Fig. 6). The latter contains the nucleus and makes synapses with other neurons of the retina. Na+ ,-K+ -ATPase is exclusively located here while the visual pigment rhodopsin is absent. The outer segment contains abundant rhodopsin in both the plasma membrane and the numerous disks. It also contains a Na+ -permeable, light-sensitive channel. Although rhodopsin can freely move in the membrane it does not diffuse into the inner segment. Ionic currents involved in transforming light into electrical signals will be described in Section III,D (Fein and Szuts, 1982; Almers and Stirling, 1984; Poo, 1985). f. In Heputocytes. The hepatocyte is a polarized epithelial cell the plasma membrane of which is differentiated into three morphologically distinct domains. The exchange of metabolites with the blood takes place at the sinusoidal surface, characterized by irregular microvilli. The lateral surface, marked by junctional elements, functions in cell-cell adhesion and communication. Secretion of bile constituents takes place at the bile canalicular domain, formed between two adjacent cell membranes and separated from the lateral surface by tight junctions (Evans, 1980; Kawakami and Hirano, 1984). Na+,K+-ATPase is mainly lo+

+

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cated at the sinusoidal and contiguous regions (Blitzer and Boyer, 1978; Poupon and Evans, 1979). To our knowledge, the location of other ion pumps or of ion channels has not, as yet, been documented for hepatocytes. The research group of Bartles used polyclonal and monoclonal antibodies in combination with morphological and biochemical methods to localize rat hepatocyte integral plasma membrane proteins. They identified a number of such proteins that exhibit a polarized distribution. So far, they concentrated primarily on specific enzymes and some receptors (Bartles et al., 1985a,b; Hubbard et al., 1985). Their techniques should be very useful to find out whether ion pumps and channels are also asymmetrically distributed over the plasma membrane.

4. Segregation of Sites of Inward and Outward Current as Revealed by the Vibrating Probe Technique When ion pump and channel activity are sufficiently separated from one another, e.g., in the basolateral and apical parts of the plasma membrane, and provided that a cell drives an electrogenic ion flux (electric current) through itself, ionic current will enter the cell at one side and leave at another. By convention the direction of current is that in which positive charges move. Extrusion of net positive charge and influx of net negative charge are both depicted as outgoing current. Extracellular ionic currents can be measured by a very sensitive measuring device, the so-called vibrating probe, originally developed by Jaffe and Nuccitelli (1974) and since then gradually improved (Dom and Weisenseel, 1982; Workshop on Vibrating Probe Technique, Nuccitelli organizer, 1985). The principle of the vibrating probe is as follows. A large electrode, located far from the site of the sensing electrode, serves as the reference. The sensing electrode is a glass micropipet, which is not as in classical electrophysiological techniques filled with, e.g., 3 M KCl solution, but with low resistance solder (or a gold-copper wire in a simplified version, Nawata, 1984) to which a small, platinum black ball of about 25 pm diameter is attached. This probe is vibrated laterally (or in three dimensions in the newer versions), usually at around 200 Hz, between two extracellular points about 30 pm apart. Any voltage difference between these points is converted by this vibration into a sinusoidal output, the amplitude of which is measured with a lock-in amplifier (in newer versions with a two phase lock-in amplifier). Therefore, the measured signal is proportional to the voltage difference and thus to the density of current flowing between these points. The novel idea in this system is to use the low impedance and, hence, low noise, of platinum black electrodes at relatively high frequencies (such as 200-300 Hz) to measure very low frequency, that is, nearly steady currents. The probe measures potential differences. The resistivity (e) of the medium in which the measurements are made is determined with a conductometer. The current density measured at the site of the sensing electrode is then

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given by J = 2.83V cos aled. V is the potential displayed by the lock-in amplifier, a is the angular deviation from the direction of the current flow, d is the distance between the two end points of the vibration, and 2.83 converts the signal amplitude from root mean square to peak to peak (Dom and Weisenseel, 1982). As tne diameter of the measuring electrode tip is of the order of 25 pm, extracellular current patterns, if existing, can be measured only in cells or systems which are not too small. If ion pumps and channels are randomly distributed, the current loops will be so small that they cannot be picked up by this method, even though it is 100-1000 times more sensitive than the classical

PELVETIA

C H A O SC H A O S (amoeba )

BLASTOCLADIELLA

LlLLY POLLEN TUBE

z

ACHLYA

XENOP~JS BLASTOMERE C O T T O N BUG OOCYTE STAGE OF XENOPUS OVARIOLE

BARLEY R O O T

MUSCLE FIBER

FIG. 3. Some examples of extracellular ion current patterns as measured by the vibrating probe technique. Figures redrawn with permission of the authors and the publishers. Egg of the brown alga Pelvetia (Nuccitelli and Jaffe, 1974; Jaffe et a/., 1974); sporulating water mold Blastocladiella (Stump et a / . , 1980); growing lily pollen (Weisenseel er al. Reproduced from The Journal of Cell Biology. 1975, 66, 556-567 by copyright permission of The Rockefeller University Press); growing hyphae of water mold Achl.ya (Kropf er a/. Reproduced from The Journal of Cell Biology, 1984,99, 486-496 by copyright permission of The Rockefeller University Press); roots of Hordeum sarivum (barley) (Weisenseel er a/.. 1979; Weisenseel, 1982); amoeba Chaos chaos (Nuccitelli er a/. Reproduced from The Journal of General Physiology, 1977, 69, 743-763 by copyright permission of The Rockefeller University Press); Xenopus luevis oocyte (Robinson, 1979); two-blastomere stage of Xenopus (Kline er a/. Reproduced from The Journal of Cell Biology, 1983, 97, 1753-1761 by copyright permission of The Rockefeller University Press); ovariole of the cotton bug Dysdercus (Dittmann ct a / . , 1981); skeletal muscle fiber of rat hind foot (Betz er a/.. 1980. Reprinted by permission from Nature, 287, 235-237. Copyright 0 1980 Macmillan Journals Limited.).

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electrophysiological techniques. Although this method does not give direct indications as to the nature of the ion pumps and channels involved, it does show that segregation of electrical membrane properties is probably the rule in all developing systems. Measurements have also been made in a large variety of systems, e.g., in the amoeba Chaos chaos (Nuccitelli et a f . , 1977), growing pollen tubes (Weisenseel et a f . , 1975; Weisenseel, 1982), in the water molds Bfastocfadieffa (Stump et a f . , 1980) and Achfya (Kropf et a f . , 1983, 1984), and plant roots (Weisenseel et a l . , 1979), etc. In Fig. 3 a schematic review is given of the major results so far obtained with this technique. In all these systems, unicellular and multicellular, current enters and leaves at well-segregated sites (Nuccitelli, 1983). For recent data, see Nuccitelli (1986).

5 . Mechanisms to Establish and Maintain Membrane Protein Segregation There must be continuously operating mechanisms to maintain the steep concentration gradients of some intrinsic membrane proteins in differentiated cells. Diffusion-mediated trapping or contact with extracellular structures or components, e.g., the basement membrane, may be one of the mechanisms involved (review by Poo, 1985). Another mechanism, perhaps of importance to segregate membrane proteins that require free mobility to be physiologically active, e.g., some hormone receptors, is electrophoresis in the plane of the membrane (lateral electrophoresis). Photoreceptors, salt-transporting epithelia, melanophores, developing eggs, meroistic ovarian follicles of insects, and probably many other cell types maintain a steady potential difference across themselves. The cytoskeleton seems to be, more and more definitely, a very essential element in regulating the mobility of some intrinsic membrane proteins. According to Almers and Stirling (1984), anchoring of membrane proteins may be fairly common. These authors discuss the fact that anchorage to parts of the cytoskeleton which are in contact with the plasma membrane is, at this moment, the most likely mechanism for reduced mobility of transport proteins in the membrane (Fig. I ) . Recently, Reggio et a f . (1984) proposed that the NH,-terminal part of the Ca2 -ATPase molecule could be responsible for a common function, namely, the bridging of actin filaments to membranes. Spectrin, postsynaptic densities, actin, myosin, and ferrocyanide binding material are usually found in high concentration subjacent to the cell membrane regions that contain highly localized or integral membrane proteins with restricted mobility. The exact mechanisms whereby transport proteins in the membrane might be directly or perhaps indirectly (e.g., through the protein ankyrin-Bennett and Stenbuck, 1979a,b) attached to cytoskeletal components are as yet not elucidated. The zonula occludens (tight junction) has for some time been thought to be essential for maintaining the segregation of proteins in the apical and basolateral parts of the plasma membrane (Pisam and Ripoche, 1976; Ziomek et a f . , 1980; Sabatini et al., 1983). More recent evidence shows that, at least in some epi+

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thelial membranes, proteins are not able to freely move, even after disruption of tight junctions. In myelinated nerve, the myelin sheath is not essential for maintaining the concentration of the Na channels in the node of Ranvier (review by Almers and Stirling, 1984). Cells, which are bifacial with respect to the location of ion pumps and ion channels over two well-segregated regions of the plasma membrane, have the possibility of driving an ionic flux through themselves. This flux does not necessarily imply electrogenic transport. In the event that there is no net transport of charges through the membrane, e.g., when one Na+ and one C1--thus an ion pair-are transported simultaneously, the transcellular ionic flux is electroneutral but present. Such coupled electroneutral 1:1 transport of Na and C1has been observed across the luminal membrane of the epithelial cells in the gall bladder and some other epithelia (for references, see Garcia-Diaz and Armstrong, 1980; Baerentsen er al., 1982). A more complex electroneutral ion flux is at work in the skin of Rana esculenta where there is an active electrogenic ion pump. However, the excretion of protons provides a favorable electrical driving force making passive apical sodium entry possible. Since the exchange ratio is 1:1, the system is also electroneutral. When there is net transport of charge, the ionic flux is electrogenic. An example is the H+-ATPase in the apical membranes of isolated turtle bladder and other tight urinary epithelia which translocates protons without coupling them to the movement of other ions (Steinmetz and Andersen, 1982). When, as the result of the segregation of ion pumps and channels, some cells drive an electrogenic ion flux through themselves, self electrophoresis will occur. This will be described further. From the examples mentioned above, it can be concluded that all cells are probably multifacial or bifacial for one or more components of their plasma membrane. +

+

F.

IONIC AND

ELECTRICAL COMPARTMENTALIZATION WITHIN

THE

CELL

Numerous membrane-limited organelles occur in the cytoplasm of eukaryotic cells. The question arises whether ion pumps and channels are found only in the plasmalemma or whether there are indications that some types of these transport proteins might perhaps also be structural elements of cytoplasmic membranes. 1. Electrophysiological Survey Gulian and Diacumakos ( 1977) made electrophysiological maps of the nucleus, Golgi region, and mitochondria in HeLa cells. The characteristics of their glass microelectrodes were tip resistances between 22 and 55 MC! and tip potentials less than 3 mV. The mean potential differences obtained by direct micropipette electrode impalement of intracellular regions from randomly selected interphase HeLa cells were cytoplasm, - 10.0 ? 0.5 mV; mitochondrion, -21 .O

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f 1.0 mV; Golgi region, -23.0 ? 1.5 mV; nucleus, -43.5 ? 1.5 mV. The positioning of the micropipette tip was confirmed by electron microscopic examination of marker solutions that had been microinjected into specific intracellular regions. They observed only single transients upon penetration of the nuclear envelope despite microelectrode movement throughout the nucleoplasm. However, multiple sharp transients were observed as microelectrodes advanced through the Golgi region which contains layers of membrane cisternae. Recent evidence has indicated that the Golgi apparatus has at least three functionally distinct regions (cis, medial, and trans) (Griffiths et al., 1982, 1983; Schachter, 1984). A study of the distribution of agglutinin binding sites in intracellular membranes of different rat tissues and human leukocytes led, in all cases, to the definition of two different endomembrane compartments. Mitochondria, peroxisomes, endoplasmic reticulum, and the nuclear envelope do not have binding sites for wheat germ agglutinin while the plasma membrane and the membrane of lysosomes, phagocytic vacuoles, and secretory granules have many. The Golgi apparatus reacts weakly (Torrisi and Pinto da Silva, 1984).

2. Vacuoles and Lysosomes Gutknecht (1967) demonstrated in the green sea weed Valonia that the vacuole is electropositive to the external sea water, due to a large positive potential across the tonoplast. In Kalanchoe leaf cells, the tonoplast is about 25 mV positive to the cytoplasm, which in its turn is about 180 mV negative to the external medium. It seems that metabolically regulated K transport out of vacuoles concentrates K + in the cytoplasm (Rona er al., 1980). Liittge and Ball (1979) postulated that Kalanchoe leaf cells have proton pumps at the plasmalemma and tonoplast which respectively extrude protons out of the cytoplasm to the external medium and into the vacuole. In parenchymal cells of oat coleoptiles the vacuole is also positive (>40 mV) to the cytoplasm but still negative to the external solution (Bates el al., 1982). It is now well established that ATPases, which perhaps pump protons, exist at the tonoplast and other vacuolar membranes (Matile, 1978; Doll, 1979; Guy er al., 1979; Marin, 1980). The proton pumps of the plasmalemma and the tonoplast of higher plants have recently been reviewed by Marrk and Ballarin-Denti (1985). Most lysosomal enzymes, when isolated, exhibit a pronounced pH optimum. Coffey and de Duve (1968), therefore, inferred that the intralysosomal pH would probably be low, in general lower than the average cytoplasmic pH. Okhuma and Poole (1978) and Poole and Okhuma (1981), using a pH-dependent fluorescent probe, reported that the intralysosomal pH in living cells is maintained in the range 4.7-4.8. According to Szego and Pietras (1984) two mechanisms for regulating the pH difference across lysosomal membranes have been postulated. The first one is a Donnan-type equilibrium system due to the presence of nondiffusible negatively charged groups within the lysosomes (i.e., acidic glyco+

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lipids, anionic lipoproteins, and glycoproteins with relatively low isoelectric points). According to Hollemans et al. (1980) this mechanism is used in rat liver lysosomes. The second one, more and more supported by strong experimental evidence, states that a lysosomal proton pump, driven by Mg2+ and ATP, is involved (Schneider et al., 1978; Schneider, 1981; Okhuma et al., 1982). Whether the lysosomal H -ATPase may be related to the well-characterized H -ATPase found in secretion granules of adrenal chromaffin cells (Johnson and Scarpa, 1976; Pollard et al., 1979; Geisow, 1982) remains to be confirmed. Recently, Reggio et al. (1984) reported that antibodies against lysosomal membranes of rat liver revealed a 100,000 Da protein that cross-reacts with purified H+,K+-ATPase from gastric mucosa. The antigen was also found in those compartments that have recently been demonstrated to be acidified by an ATP driven pump (some compartments of an endocytic pathway in macrophages, small amounts in the plasma membrane of liver, but large amounts in coated vesicles, some parts of the Golgi complex, etc.). For the role played by the low pH in lysosomes, we refer to the review of Szego and Pietras (1984). Golgi vesicles of liver (Zhang and Schneider, 1983) and those from lactating rat mammary glands (Virk et al., 1985) possess a ApH of about 1 pH unit. Golgi system membranes of adipocytes seem to have a glucose transport activity similar to the plasma membrane (Smith et al., 1984). Lutoids, which are abundant in some plants, e.g., Hevea brasiliensis, are single-membrane microvacuoles with lysosomal characteristics. They have a lower pH (about 5.5) than that of their cytoplasmic environment (about 7) and accumulate numerous mineral and organic ions. Critin (1982) presented data strongly supporting the presence of an inward electrogenic proton-translocating ATPase on the membrane of intact lutoids. Newly formed and especially aged neurohypophyseal neurosecretory granules are more internally acidic than the cytoplasm, but this seems to originate from a Donnan equilibrium (Scherman and Nordmann, 1982). Clathrin-coated vesicles contain an ATP-dependent proton pump that may play a role in the acidification events that are essential in receptor-mediated endocytosis (Forgac et al., 1983). +

+

3. Mitochondria The question about the “right” membrane potential of mitochondria is also as yet not settled (Tedeschi, 1980). The chemiosmotic hypothesis of the coupling mechanism states that oxidative phosphorylation requires that there should be a total promotive force of 210-270 mV across the osmotic barrier of the cristae membrane of respiring mitochondria (electrically negative inside the mitochondrion, Mitchell, 1961, 1966, 1967; Mitchell and Moyle, 1969). Gulian and Diacumakos (1977) measured in HeLa cells in situ only -2 I mV for the impaled mitochondria. Experiments with isolated giant mitochondria of Drosophila yielded values of 10-20 mV, positive inside (Tupper and Tedeschi, 1969).

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Similar positive values were found in the metabolically viable giant mitochondria isolated from the liver of cuprizone-fed mice (Maloff er al., 1978a). The reason why the values are positive might be that the ionic composition of the incubation medium is quite different from that of the in vivo cytoplasm. If impalement had caused a substantial leak, which is of course possible with such small organelles, then the metabolic viability, according to Mitchell’s hypothesis, would have been lost. It, however, remained intact as revealed by biochemical analyses (Maloff er al., 1978b). During the recent decade, indirect methods have been used for estimating the membrane potential of mitochondria and, depending upon the conditions, values closer to the ones predicted by Mitchell have been reported (see, e.g., Rottenberg, 1984). According to Ling (1981) the discrepancy between calculated and actually measured values is due to the fact that several premises of the chemiosmotic theory are incorrect: mitochondria1 functioning could be explained by his association-induction hypothesis. Chloroplasts also seem to be able to create their own ionic environment (Robinson and Downton, 1984). 4. Sarcoplasmic Reticulum The sarcoplasmic reticulum of skeletal muscle cells is very rich in Ca2+ATPase and is a storage site for Ca2 , the essential ion controlling muscle contraction. Merocyanin 540 is a potential sensitive dye. Shifts in fluorescence have been observed during Ca transport in reconstituted vesicles of sarcoplasmic reticulum (Haeyaert et al., 1980). This suggests that there is a charge deficit, despite the simultaneous transport of Mg2 , K ,and, perhaps, H and organic ions (Somlyo et al., 1981). +

+

+

+

5 . Nucleus The question of whether there is also a potential difference across the nuclear envelope has as yet received little attention as it is intuitively assumed that because of the presence of the numerous “pores,” such a potential difference cannot be built up. The scarce experimental data suggest that the nuclei of some somatic cells, at least, may be electrically and ionically compartmentalizedfrom the cytoplasm. Loewenstein and Kanno (1963) examined the electrical potential and resistance in situ and in isolated giant nuclei of salivary glands of Drosophila flavorepleta, with microelectrodes of 10-35 M a , and tip potentials lower than 2.5 mV. The measured resistance of the nuclear envelope was of the order of 1 fl cm2. This is smaller than that of the cell membrane but still large enough to represent a formidable barrier to ion diffusion. The nucleoplasm was, on the average, 15 mV negative with respect to the cytoplasm. The potential declined to zero and the resistance to a fraction of its original value when the nuclear membrane was experimentally perforated by repeatedly driving an empty micro-

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pipette through it. From these experiments Loewenstein and Kanno concluded that it was unlikely that the sites where the double nuclear envelope fuses together (later called the pore complexes) are bridged by nucleoplasm or cytoplasm. They speculated that these “pores” might be additional barriers which confer its high electrical resistance upon the nuclear envelope. In salivary gland cells of another insect, Chironomus thummi, the nucleoplasm was found to be 2-5 mV negative to the cytoplasm (Ito and Loewenstein, 1965). We will refer to nuclei which are able to maintain a potential difference over the nuclear envelope as “closed” nuclei. When Kanno and Loewenstein (1963) performed similar experiments on the germinal vesicle of an amphibian oocyte, they found that this type of nucleus was a rather permeable structure, its resistance was indistinguishable from that of the cytoplasm and nucleoplasm, and no nucleuspotential could be measured. Such a type of nucleus will be referred to as “open.” If there is indeed a potential difference between nucleoplasm and cytoplasm, one would expect to find differences in ionic concentrations-or better activities-in both compartments. Such differences in ionic concentrations between nucleus and cytoplasm of frog oocytes were already reported in 1962 by Naora et al. who dissected out clean and whole nuclei from frozen oocytes. The Na+ and K + content (microequivalents per gram water) of the nucleus were respectively 3.2 and 2.4 times higher than that of the cytoplasm (excluding yolk platelets). The Na+ :K ratio of the nucleus was 1.1 and that of the cytoplasm (excluding yolk platelets) was 0.72. Autoradiographic experiments after incorporation using [22Na, 42K, 14C]leucine and [t4C]alanine showed, in each case, a marked accumulation within the nucleus. Only a very small fraction of the amino acids which accumulated in the nucleus could be precipitated by trichloroacetic acid, which indicates that most amino acids were not incorporated into proteins. These data suggested that, at the level of the nuclear envelope, active mechanisms might be present for uphill movement of ions and amino acids and that the germinal vesicle is of the “closed” type in the developmental stage used by Naora et a f . (1962). However, these data are not supported by more recent work on concentrations and activities of K and Na in oocytes of R a m pipiens. In small oocytes (<40 pg), the ooplasm resembled the cytoplasm of somatic cells and the nucleus and cytoplasm little differed in their K + and Na+ concentrations (Jones et al., 1979; Pieri et al., 1977; Rick et al., 1981; Tluczek et al., 1984). Although there are marked changes in H20, K + , and Na+ content between the germinal vesicle and the ooplasm, they seem to be due to the presence of yolk. If yolk is not taken into account, rather similar values for ooplasm and nucleoplasm are found (Tluczek et a/., 1984). Paine et al. (1981) also found higher K + and lower Na concentrations in the germinal vesicle than in the cytoplasm of the salamander Desmognathus. They showed that nucleus/cytoplasm cation concentration dif+

+

+

+

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ferences are due to (1) partial exclusion of diffusive Na+ and K + by the cytoplasm and (2) differential Na and K binding by the nucleus and cytoplasm. These authors injected gelatin at 34°C to perform their measurements but this temperature is high enough to cause heat shock effects and changes in germinal vesicle envelope properties. Using energy-dispersive X-ray microanalysis, Labadie et al. (1983) also conclude that in Xenopus the concentration of the major ions per volume of free water in the nucleus is probably rather similar to that in the cytoplasm. All these newer data confirm the findings of Kanno and Loewenstein (1963) that there is no potential difference between the germinal vesicle and cytoplasm/yolk of the amphibian oocyte. Data on the ionic concentrations of nuclei and cytoplasm in somatic cells are much scarcer. Langendorf et al. (1961, 1966) reported differences in ionic concentrations between nucleo- and cytoplasm of rat liver cells. Andrews et al. (1983) reported a marked accumulation of P and K + in the nucleus (388 and 190 mmol/kg wet weight, respectively) relative to the cytoplasm (67 and 85 mmollkg wet weight) in domestic duckling erythrocytes, but they did not observe such pronounced differences in the epithelial cells of the salt gland. We have not found enough recent data to either confirm or reject these older results or the electrophysiological data of Gulian and Diacumakos (1977). The question of whether the nuclei of somatic cells are indeed “closed” remains to be answered. If the measured potential differences over the nuclear envelope are not artifacts, then one might expect that active mechanisms are needed for maintaining the ionic/voltage gradients. No experimental data are so far available. The ubiquitous presence of a nuclear envelope in all eukaryotic cells suggests that this envelope is essential to proper nuclear functioning and not just a double membrane with numerous open pores. In our opinion it is not illogical to assume that the nuclear envelope is essential for generating the right ionic conditions for transcription, which may be different from those needed for translation in the cytoplasm. The nuclear pore complexes might turn out to be structures which, under the right conditions, can easily open and close as is the case for gap junctions. The discovery of Goldfine et al. (1982) that receptors for insulin are present in the nuclear envelope of rat liver cells also suggests that this membrane might play a very important role in cell function. Although the amount of hard data is still very limited, the picture emerges that it may very well be possible that membrane bound cell organelles may create their own ionic/electrical environment, either by a Donnan-type equilibrium system orland active pumping. Changes in the ionic environment or the electrical conditions at the outside of these organelles may provoke changes in their interior. Compartmentalization might be imposed by physiological needs. Many enzymes are localized in specific organelles, e.g., lysosomes, nucleus, mitochondria, Golgi systems, and their optimal activity might require a specific ionic (including H + ) environment. +

+

THE ELECTRICAL DIMENSION OF CELLS

G.

28 1

GRADIENTS WITHIN CELLS: TRANSCYTOPLASMIC CURRENTS

VOLTAGE A N D IONIC

In the preceding section we discussed the fact that segregation of ion transport proteins in the plasmalemma occurs in many cell types. This is especially clear in salt-transporting epithelia and in those systems in which extracellular currents can be measured by the vibrating probe technique. It is thus conceivable that such cells could generate intracellular potential and/or concentration gradients along which solutes are driven. Theoretically, intracellular potential gradients could be significant (a few millivolts) on condition that the resistivity of the cytoplasm would be high enough (Petersen, 1980). Even with the very sensitive equipment available today, it is not easy to make accurate measurements of intracellular potential differences in small cells. In classical electrophysiological research it is often not ascertained at which precise part of the cytoplasm the measurement is made, as it is generally assumed that the cell’s interior is at isopotential. There is always the problem of possible leak artifacts due to the penetration of the membrane with microelectrodes. Another problem is the possible leakage of electrolytes from the interior of the microelectrode into the cell with a subsequent alteration of the intracellular electrolyte concentration and/or cell volume. Differences in En, over the apical membrane as compared to that of the basolateral membrane have been reported in a number of epithelia. Zeuthen and Monge (1975) reported that in epithelial cells of the rabbit ileum the En, at the luminal end was -6 mV, while at the serosal end it was -36 mV. In the salivary gland cells of the adult fleshfly, Calliphora, the luminal site of the plasmalemma, at rest, is about 4 mV more negative than the apical site. This difference increases to 20 mV after stimulation with CAMP(Berridge and Prince, 1972). In rabbit corneal epitheleum, which is considered as a “tight” Cl--secreting epithelium, the resting potentials across apical and basal barriers are about 49 and 75 mV, respectively (Marshall and Klyce, 1984). Zeuthen (1977, 1978) also studied the epithelial cells of Necturus gallbladder. The electrode tip was manually advanced through the cytoplasm and intracellular gradients of electrical potential of 0.6 m V / p n were measured. The mean potential difference across the mucosal membrane was recorded as -29 mV while in all cases the potential across the basal membrane was noted as -52 mV. Zeuthen found the same potential profile and membrane potentials on penetrating the epithelium from the serosal site and measurements with multibarreled electrodes gave roughly the same internal potential gradient. The resistivity of the cytoplasm could be determined by a triple-barreled microelectrode. It varied from 10 times that of Necturus saline at the mucosal end of the cell to 4 times in the middle and 6 times at the serosal end. The resistance of the single-barreled electrodes used by Zeuthen was 5-60 MR. Ion activities were measured with

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ion-selective electrodes (Zeuthen, 1978). The Na+ activity varied from about 39 mM at the mucosal end to between 8 and 19 mM at the serosal (basal) end. In cells from animals stored at 5"C, the CI- activity varied from about 55 mM in the mucosal end to 28 mM in the serosal end, and the K + activity from 50 mM to between 95 and 131 mM. Zeuthen (1978) states that within the cell both the recorded electrical and chemical gradients cause Na+ to move toward the serosal end. Suzuki and Fromter (1977) also undertook a study on the potential and resistance profile of the Necturus gallbladder cell but they were able to produce data similar to those of Zeuthen only when using low resistance electrodes ( 1 1 2 Mil). Using high resistance electrodes (230 M a ) the intracellular electrical potential was found to be independent of the depth of penetration of the electrode into the cell. These authors consider Zeuthen's data as largely due to artifacts and as of the order of lo00 times greater than can be inferred from calculated values. They conclude that although it is very conceivable that epithelial cells which transport solutes and water from one compartment into another might generate significant intracellular concentration or potential gradients along which the solutes might be driven across the cytoplasmic space, the Necturus gallbladder epithelium does not possess the necessary electrical properties to permit this. According to Petersen (1980) the question of intracellular potential gradients in this cell type cannot as yet be considered as definitely settled. Regional distinctions-apical versus basal-in the distributions of Na+ ,K and CI - were measured in the salt gland cells of 15-day-old domestic ducklings, with particular emphasis being placed on those from individuals adapted to salt water (Andrews et al., 1983). Slayman and Slayman (1962) used microelectrodes (20-35 M a ) to make intracellular measurements in the filamentous fungus Neurospora crassu. At a distance of 8 mm behind the growing hyphal tips, the average membrane potential was - 127 10 mV. There was a slight gradual decline in average potential toward the center of the colony but, toward the periphery, a more abrupt drop to about 25 mV at the growing hyphal tips was measured. By means of the vibrating probe, Gow (1984) showed that not only this species but also several other fungi generate steady electrical currents around their hyphal tips: it may be a universal characteristic of hyphal growth. Growing hyphae of Achlyu bisexualis generate a longitudinal pH gradient in the surrounding medium (Gow ef al., 1984) and its amino acid proton symport seems to be the mechanism of current entry (Kropf et al., 1984). Bingley (1966) also used microelectrodes for the study of membrane potentials of Amoeba proteus. The advancing part was about 40 mV more positive than other parts; this corresponds to 1 V/cm. In vertebrate photoreceptors a steady current-outward across the cell membrane of the proximal and inward across the membrane of the outer segmentflows continuously between the 2 segments (Almers and Stirling, 1984, Section +

*

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283

IILD). In Limulus neutral photoreceptors a cytoplasmic Ca2+ gradient has been found (Harary and Brown, 1984). Another example of a voltage gradient between different parts of the plasmalemma of a cell and the resulting electrophoretic movement of cell organelles is that of the melanophores of the teleost Oryzias. During the dispersion phase of the melanosomes, which is promoted by the hormone MSH, the Em of the cell body region is about -37 mV while the Em at the tips of the branches is about -20 mV. Kinosita (1963), who made these measurements, suggested that the melanosomes, which carry a negative charge, are displaced as the result of self electrophoresis in the cytoplasm: in the dispersion phase they move toward the tips of the branches. In the aggregation phase, the potential gradient is reversed, again under the influence of hormones (adrenaline), and the granules aggregate in the cell body region. MSH promotes the entry of Na+ into the cell. Jaffe (1966) was able to measure potential differences between the poles of germinating fucoid eggs. Since the signal produced by one such egg is just at the lower limit of detection, the signal was multiplied by lining up many eggs in a long loose fitting capillary. Parallel development of all eggs was induced with light applied at one end and the voltage difference in the media at both ends of the tube was then measured. Voltage differences became measurable when the eggs began to elongate. These data have been of crucial importance for the formulation of the self electrophoresis principle and the development of the vibrating probe technique (Jaffe er al., 1974; Jaffe and Nuccitelli, 1974). For most cells, especially somatic epithelial cells, their small size is the limiting factor for accurately establishing whether or not there is a continuous voltage gradient in their cytoplasm. Large ovarian follicles have proved to be good material in this context. As early as 1905 Hyde brought two Zn electrodes in contact with membranes of eggs of different vertebrate species and, depending upon the sites of contact, potential differences, which changed in polarity in the course of embryonic development, could be measured. This was not an easy technique but it was almost as sensitive as the most sophisticated modem equipment. In insects there are panoistic and meroistic types of ovarioles. In a panoistic ovariole, the ovarian follicle consists of an oocyte which is surrounded by a layer of follicle cells, as is the case in vertebrates. In ovarian follicles of meroistic ovarioles, three major cell types occur, namely one oocyte, a species-specific number of trophocytes, all being surrounded by a layer of follicle cells. There are two types of meroistic ovarioles. In the polytrophic type, the trophocytes or nurse cells are connected to the oocyte by short open cytoplasmic bridges (ring canals). In the telotrophic type, the nurse cells lie together at the apex of each ovariole in a tropharium which is connected to the young oocytes by means of a long tenuous cytoplasmic cord which is stabilized by a microtubular cytoskeleton. As the electrophysiology of insect ovarian follicles is best documented for

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polytrophic types (review De Loof, 1983), it is these latter which will be discussed here. Oocyte and trophocyte are sibling cells which originate from the same cystoblast. When the cystoblast divides cytokinesis remains incomplete so that the division products remain interconnected by a cytoplasmic bridge or ring canal. Depending upon the insect species, there may be several more divisions, always with incomplete cytokinesis. Thus all descendants of one cystoblast remain interconnected by bridges and form a cytoplasmic continuum. In Drosophila, there are 15 trophocytes and one oocyte (Koch et al., 1967). A similar situation prevails in the fleshfly Sarcophaga buflata (Cardoen et al., personal communication) and many other Diptera. In the moth Hyafophora cecropia, there are 7 trophocytes and 1 oocyte. The different cytoplasmic bridges all fuse together to form a central fusome (Fig. 4). This property of the ovarian follicles of this insect species makes them very suitable for electrophysiological experiments, as shown by Woodruff and Telfer (1973, 1974, 1980). Meroistic ovarian follicles are very interesting systems for the study of differentiation and polarized transport. Indeed, although forming a cytoplasmic continuum, being genetically identical and being exposed to the same hormonal environment in the hemolymph, only the trophocytes become polyploid and very active in RNA synthesis. RNA is unidirectionally transported from the trophocytes to the oocyte. It was during their research into how this polarized transport took place that Woodruff and Telfer tested the hypothesis of the existence of a voltage gradient between the trophocyte and oocyte compartment. By means of classical electrophysiological methods, an oocyte potential of -40 mV relative to the hemolymph and a trophocyte potential of -45 mV were recorded (Fig. 4). Thus, over the central fusome (width about 50 km, path length less than 50 pm, with a very low resistance in the cytoplasm of about 10 kfl according to Jaffe and Woodruff, 1977, 1979) there is an average potential difference of about 5 mV, the oocyte being at positive potential (Telfer et al., 1981). This is quite remarkable. The potential gradient in the bridge is greater than 1 V/cm and the bridge current of the order of 0.5 pA (Woodruff and Telfer, 1974; Jaffe and Woodruff, 1979; Telfer er al.. 1981). In order to verify whether the polarized transport of the (supposedly negatively charged) RNA particles could be due to the voltage gradient, Woodruff and Telfer microinjected fluorescently labeled lysosyme (FLY), which is positively charged at a physiological pH, or its negatively charged form obtained by carboxymethylation (FMcLy) into either the trophocytes or the oocyte. They observed that within the follicle these injected proteins migrated just as they would be expected to do in an electrophoresis chamber: relative to the trophocyte Compartment the oocyte is at positive potential (Fig. 4). For details see review by De Loof (1983). Similar experiments have been carried out in the telotrophic ovariole in the hemipteran Rhodnius profixus. Here the equilibrium potential of the tropharium is about 3 mV negative to that of the oocyte, when measured in Rhodnius Ringer’s solution. A clear electrical polarity

THE ELECTRICAL DIMENSION OF CELLS FI uorescein

Basic (+)FLv

e

Acidic ( - 1 FMcLv

0Space penetrable by fluorescent protein \

285

f

Gap junctional membranes Direction of ion currents

FIG. 4. Self-electrophoresis in the ovarian follicle of the moth Hyulophoru cecropia as demonstrated by Woodruff and Telfer (1973, 1974, 1980). The oocyte (Ooc) is interconnected to seven nurse cells (NC) by means of a cytoplasmic bridge. Fluorescein, positively charged fluorescent lysosyme (FLY) and negatively charged fluorescent methylcarboxylated lysosyme (FMcLy) were injected either into the nurse cell compartment or into the oocyte. The lower row of drawings of a-c gives the localization of fluorescent molecules about 60 minutes after injection. Fluorescein, injected either into the oocyte or into a nurse cell, rapidly spreads not only over all cells of the follicle into which it was injected but also to more anteriorly and posteriorly located follicles, showing that they all are electrically coupled. Microinjected FLYand FMcLy do not diffuse into the follicle cells or to adjacent follicles. FLy and FMcLy migrate in opposite directions, illustrating the self-electrophoresis principle. The oocyte (-40 mV) and nurse cells (-45 mV) behave as anode and cathode, respectively. Noncharged molecules diffuse in both directions. By means of the vibrating probe technique. Jaffe and Woodruff (1979) showed that current enters the nurse cell compartment and leaves the oocyte (d). A more detailed analysis of the system revealed that, although a net influx of current is found over the nurse cell compartment (0,there is a current flow out of the nurse cells into the spaces between these cells and the overlying follicle cells (FC) (e, Woodruff, 1984). a-d, Redrawn after De Loof (1983) with copyright permission; e and f, kindly provided by R. Woodruff (unpublished).

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was observed in the tropharium itself it seems as if the more apically situated parts are more negative than the basal part (Telfer et al., 1981). In another hemipteran, Dysdercus, a similar situation seems to prevail although it has as yet not been investigated with microinjection experiments (Dittmann et al., 1981; De Loof, 1983). The vibrating probe technique has been applied to panoistic and meroistic ovarian follicles and the data so far obtained provide additional evidence that these structures are indeed miniature electrophoresis chambers: in H. cecropia (Jaffe and Woodruff, 1979; Woodruff, 1982), Dysdercus (Dittmann et al., 1981), Rhodnius (Huebner, 1984), Drosophila (Overall and Jaffe, 1985), and Sarcophaga (Verachtert and De Loof, 1984, 1986), current enters the tropharium and leaves the oocyte compartment. In the beetle Ips, current enters solely at the tip of the tropharium and leaves at the more posterior region (Huebner, 1984). Panoistic ovarian follicles also show regions in which current enters or leaves although there is no apparent morphological difference between the anterior and the posterior parts (Periplanera, Huebner, 1984). Heinrich et al. (1983) studied the Na+ , K + , and Ca2+ distribution in developing follicles of Drosophila melanogaster with a laser microprobe mass analyzer after precipitation of the cations with potassium antimonate. Ionic asymmetries were observed in vitellogenic ovaries. The concentration of the three ions increases along the anteroposterior gradient. To our knowledge, no data on ion activity, obtained with ion selective electrodes, have as yet been published for insect ovarian follicles. A cytoplasmic Ca2 gradient has been autoradiographically visualized in eggs of a brown sea weed Pelvetiu by Gilkey (for figure, see Nuccitelli, 1983). +

IONIC SPECIES:AN EXTREMELY VERSATILE SYSTEM H. FEWINORGANIC 1. In Differentiated Organisms In the preceding pages, we have seen that if inorganic trace elements are not considered, only a few inorganic species are used by the different cell types, be it plant, animal, or prokaryote; the major ones are K , Na , Ca2 , Mg2 , H , CI- , and HCO, Up to now about a dozen ion channels and about half a dozen ion pumps have been described. Gating and closing of channels can be achieved in different ways, depending on the channel type. There are voltage-gated channels, receptor-operated,and chemically mediated ones. Physical factors may also be important and combinations are possible.Ion pump activity may be influenced by a variety of factors, both physical and chemical. Data have also been given on the asymmetrical distribution of ion pumps and channels, on electrical/ionic compartmentalization, and gradients within the cell. Not only inorganic, but also organic ions contribute to the ionic environment of the cytoplasm. The transport +

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of organic ions is often coupled to that of an inorganic ion (cotransport, countertransport). For example, the uptake of several amino acids is Na+ dependent. Ca2+ plays an important role in many physiological processes. The intracellular Ca2+ concentration, which is of an order of magnitude of lo-’ M versus about l o w 3M in the extracellular fluids, has to be kept under rigorous control in order not to disturb normal cellular functions (McLean and Urist, 1968). Therefore it is not surprising that for this particular ion there is some evidence for compartmentalization of the Ca2+ that has to be transported through the cell, especially in cells which handle large amounts (Terepka et al., 1976). In this section we will try to give an idea of the endless versatility in “ionic environments” which can be generated in the cytoplasm or in cell organelles, although only a limited number of inorganic ionic species are involved. The ‘‘ionic environments” comprise ionic concentrations, potential and ionic gradients, ionic currents, and secondary chemical gradients. This ionic environment at a given location in the cell, e.g., in the nucleus, is dependent upon successive levels of control (De Loof and Geysen, 1983; De Loof, 1985a,b). Many factors operate at the level of the plasma membrane (first level of control). The way neighboring cells make contact is a second control level. It may make a difference whether neighboring cells are interconnected by gap junctions or not. Gap junctions are structures which allow cell to cell communication for ions and small (up to a molecular mass of the order of 2000 Da) molecules (review Loewenstein, 1981). The cytoplasmic Ca2+ concentration is a major determinant for the opening and closing of gap junctions. They have been found in many differentiated tissues but they are already operating in young embryos (Fundulus and axolotl, Bennett et al., 1978; mouse, Lo and Gilula, 1979; Patella, Dorresteijn er al., 1983; Xenopus blastomeres, Guthrie, 1984; Warner et al., 1984; and embryonic Xenopus muscle cells in vitro, Chow and Poo, 1984). Gap junctions make interconnected cells into a functional unit. Tight junctions between neighboring cells allow the generation of large transepithelial potential differences. The route followed by ion fluxes may be different in tight versus leaky epithelia. The properties of the limiting membrane of the cell organelle, e.g., of a closed nuclear envelope, may be a third control level. The site at which the given cell organelle is located is important, e.g., in or out of the main cytoplasmic ion flux (if present) or in a location where, as the result of self electrophoresis, higher or lower concentrations of charged macromolecules occur (secondary chemical gradients) (fourth possible control level). The exact location of the chromosomes in the nucleus may be a fifth control level to be taken into account in order to understand the influence of ions on gene expression. The only organism in which the spatial organization of interphase chromosomes has been mapped is Drosophila. In its salivary gland cells, each

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polytene chromosome arm folds up in a characteristic way, contacts the inner aspect of the nuclear envelope at specific sites, and is topologically isolated from all other arms (Agard and Sedat, 1983; Mathog et al., 1984). The ovalbumin gene is associated with the nuclear matrix of chicken oviduct cells (Robinson et al., 1982). As in bacteria (Anderson and Roth, 1981), the chromosomal position of an artificially introduced mammalian gene in a mammalian genome, can determine the amplification frequency and mechanism, and the size and stability of the resulting amplified genes (Wahl et al., 1984). The inactivated second Xchromosome of female mammals is attached to the inner aspect of the nuclear envelope (Barr body). In neutrophils, the small lobe extruding from the nuclear envelope is thought by some to contain the inactivated X-chromosome (Brown, 1980). De Loof et al. (1982) suggested that the inactivated X-chromosome is located at a site where it minimally experiences ionic fluxes. All this is schematically represented in Fig. 5. Taking into account the various control levels, it may not be excluded that the different cell types of an organism all have different “ionic environments” in their cytoplasm. It seems possible that there may be different ways to realize a given ionic environment. 2. Generation of Asymmetry as a Key to Differentiation: The Double Asymmetry Principle As we have briefly shown there exists a lot of evidence to indicate that the plasma membrane properties of the different cell types and tissues of a differentiated organism may differ from each other with respect to the types of ionic channels and pumps which occur, the relative abundance of these, their distribution over the plasma membrane, and also, perhaps, their association with other intrinsic membrane proteins, e.g., hormone receptors. A very basic but difficult question now arises. What mechanism(s) underlies the formation of these differences? If this question could be answered, then at least part of the basic principles of differentiation would be clarified. Although differentiation is usually considered as a very complex process, it may not, a priori, be excluded that, after all, the basic principle(s) involved might be much simpler than is often thought. Four major principles have to be taken into account (De Loof and Geysen, 1983; De Loof, 1985a-c). 1. It is not the DNA which is the unidimensional description of the whole future organism, but it is the fertilized egg with itsfive dimensions (four dimensions of spacetime plus electrical dimension) which is the unicellular description of it. 2. “The fertilized egg contains it all; all the rest follows of necessity” (Raven, 1959). This means that only mechanisms already present in the egg will govern differentiation.

THE ELECTRICAL DIMENSION OF CELLS

'

DIFFERENT TYPES

OF ION

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PUMPS

1

H

W l U

2

3

4

FIG. 5 . Schematic representation of the major control levels which contribute to the establishment of a given ionic environment (ionic concentrations, potential and ionic gradients, ionic currents, secondary chemical gradients) in the cytoplasm or in cell organelles (adapted from De Loof, 1984, with copyright permission). 1. Factors operating at the plasmalemma: Single cells (C, H). A given cell type may contain different types of ion channels (A) and pumps (B). As the exact structure of no single pump or channel is as yet elucidated, the representation in A and B is just schematic. Some cell types are excitable, others are not. If the plasma membrane would be perfectly fluid, the transport proteins for ions could be rather symmetrically distributed over the plasmalemma (C,, H3). Ionic current loops which might occur between neighboring pumps and corresponding channels would be so small as not to be measurable with currently available equipment. Such cells would be unifacial with respect to their membrane properties and they would be at equipotential. Asymmetrical distribution of ion pump/channel activity (Cz, Cq. Hq) seems to be the rule, not the exception (bifacial and multifacial cells). An infinite number of combinations is theoretically possible. If cells drive ionic fluxes through

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3. Tissue-specific protein synthesis is not the primary cause of differentiation, but its result. 4. By definition, differentiation is the generation of asymmetry.

In somatic cells of developing animals and plants, the asymmetry does not seem to have been introduced in the DNA itself. Whole plants can develop from a single differentiated cell or a piece of a plant. Gurdon (1962, 1974) and Hoffner and Diberardino (1980) showed that nuclei of differentiated amphibian cells still contain a complete genome able to generate a complete new organism when introduced into an enucleated egg. In a study of five developmentally regulated regions of genomic DNA of Drosophila, no detectable changes in organization, modification, methylation, or selective amplification were found during development (Levine et al., 1981). It is therefore logical to assume that asymmetry is introduced in the operative key(s) to the genetic information. Control of gene expression is very complex: in addition to mechanisms operating at the level of the DNA/RNA (Davidson, 1976; Lewin, 1980; Brown, 1981; Raff and Kaufman, 1983) there are also epigenetic factors (Gurdon, 1974; Lprvtrup, themselves, these may be electroneutral (C2, C3) or electrogenic (C4). Ionic gradients and voltage gradients may exist in the cytoplasm (D4). Cell groups. In multicellular organisms, cells may be grouped into epithelia, muscle, liver, etc. Different types of contact between neighboring cells may exist. Epithelia may be tight (Dz) or more or less leaky (D3) whereby the transepithelial voltage difference which may be generated may vary from type to type. Large cytoplasmic bridges are known to exist, e.g., in meroistic insect ovarian follicles and in Volvox. Gap functions may be present (D,, lower half) or absent (D,, upper half) between neighboring cells. A given cell (group) may release a substance(s), which may influence the activity of specific ion pumpslchannels of other cells (inductors for embryonic cells, neurotransmittors for excitable cells, hormones for neighboring or distant cells). 2. Factors operating in the cytoplasm. A proportion of the ions in the cytoplasm are free (ion activity) and part may be bound to macromolecules, e.g., proteins. In case of transcytoplasmic ion fluxes, the route followed by the ions will evidently depend upon the localization of the ion pumpslchannels in the plasmalemma (D4). but also upon local differences in conductivity in the cytoplasm, if these exist (C4. homogeneous conductivity in cytoplasm; Cs. conductivity highest just underneath the plasma membrane). Secondary messengers, CAMP, Ca* , and apparently inositol triphosphate also (Berridge and Prince, 1984) may be used to coordinate ion pumplchannel activity located in different parts of the plasma membrane. In the case of ion pumplchannel activity being asymmetricallydistributed and the transcytoplasmic ion flux being electrogenic, the cell will behave as a miniature electrophoresis chamber with all the properties inherent to this structure (C4, HI). Gradients of inorganic ions and of free charged macromolecules may be found (HI). It is not inconceivable that these, in turn, affect the permeability of the plasma membrane and of some ionically compartmentalized cell organelles. 3. The behavior ofmembrane bound cell organelles. Little is known about the possible mechanisms used to maintain the ionic compartmentalization of cell organelles. If a Donnan equilibrium system is used, the internal ionic composition of the organelles will change in accordance to changes in the cytoplasm where they are found. In the case of active mechanisms being involved, the same reasoning applicable to the factors operating at the plasmalemma can be used. Not only the type of ion pumpslchannels present in the membranes of the cell organelles but perhaps also their location +

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1983; De Loof and Geysen, 1983) and these latter factors are especially important for differentiation and morphogenesis. Some of the epigenetic factors seem to be located in the oolemma and underlying cortex as has been shown by many classical experiments on fate maps. This has also been recently illustrated by an experiment in which eggs of the brown alga Pelvetiu were grown in a gradient of the Ca2 ionophore A23 187: these eggs tended to form their rhizoid outgrowth on the sides that were exposed to the higher concentration of the ionophore, thus at the site where the membrane was probably most permeable to Ca2 (Robinson and Cone, 1980). Similarly, A23 187 also polarizes ooplasmic segregation in eggs of the ascidian Boltenia villosa (Jeffery, 1982; more details in Section 111,A). Asymmetry can only be introduced in an almost perfectly reproducible way in those molecular mechanisms which have a well defined and constant location in the cell and which are practically free from the effects of diffusion or cytoplasmic streaming as occurs in Drosophila (Gutzeit and Koppa, 1982). From the data which have recently become available, the oolemma is a good candidate but it can only be of importance for differentiation if two conditions are fulfilled. The first condition is that it would not be completely fluid but quite well fixed and that it could be divided in an asymmetrical way. The second condition +

+

(symmetrically or asymmetrically) distributed in the membrane of the cell organelle) and their susceptibility to chemical and physical factors or changes in voltage gradients within the cytoplasm have to be taken into account. Some cell organelles have a well-defined location in the cytoplasm. This is well documented for the nucleus (N) which, in most differentiated tissues, is always located in a cell type-specific site (F1,2,3,4). This may be important as shown in G4,5,6; it may make a difference whether a given cell organelle is located in a transcytoplasmic electrogenic ion flux or not. 4. The ionic environment of the chromosomes (ionic concentraiions, aciiviiies, gradients, fluxes). This depends on more factors than generally realized. The electrical resistance of the nuclear envelope may be similar to that in cytoplasm as is the case for what we have called an “open” nuclear envelope, and the ionic activity of the nucleoplasm will then be similar to that of the cytoplasm (El). If the resistance is significantly higher (closed envelope), the ionic composition of the nucleoplasm may be different from that of the cytoplasm. If ion pumplchannels would be present, they might be symmetrically (E2) or asymmetrically (E3) distributed over the nuclear envelope. If they would be asymmetrically distributed, transnuclear ion fluxes would be possible. In this respect, the location of the chromosomes (G: g l, g2, g3) in the nucleus is important, e.g., attached to the nuclear envelope (g2 and g,) or more in the center (gl) of the nucleus, in ( g f , g,) or out (g2) the flux of ions. The germinal vesicle in oocytes of the insect Heieropyza pulsates rhythmically (Went, 1977). It is not known whether contractility of the nucleus (E4.5 and E6) is rare or occurs rather frequently in other cell types. It may not be excluded that ion fluxes through the nuclear envelope are generated during contraction and relaxation. It is as yet unknown whether the more or less deep invaginations, so often observed in electron micrographs of nuclei, e.g., in macrophages, are due to regular pulsations or to other mechanisms. These data taken together indicate that the term ionic environment of the chromosomes includes not only concentrations of inorganic and organic ions but also, in certain circumstances, ionic and voltage gradients. 5 . Although it is known that some prokaryotes display high Em values (Section 1II.H) very little is known about the symmetry of distribution of ion pump/channel activity. Theoretically this distribution could be symmetrical (H,)or asymmetrical (H4). A I mV potential difference over both ends of a I-pm-long rod-shaped bacterium corresponds to a gradient of 10 V/cm.

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maintains that there should be a causal relationship between the properties of the plasma membrane and nuclear activity. In recent years, the vibrating probe technique and other experiments yielded evidence that the plasma membrane of egg cells is indeed asymmetrical with respect to the distribution of transport proteins (first element of asymmetry). This was shown by the fact that, in animal eggs, electric current, as measured by the vibrating probe, enters the egg at the animal pole and leaves at the vegetative pole (Jaffe and Nuccitelli, 1974; Jaffe et al., 1974; Jaffe and Woodruff, 1979; Jaffe and Guerrier, 1981) while the opposite situation prevails in the eggs of the algae Fucus and Pelveria (Jaffe, 1966, 1975, 1982) (Fig. 3). Kline et al. (1983) found an asymmetrical distribution of ion pump/channel activity in the Xenopus 2-cell stage embryo, which persists to the 32-cell stage (Nuccitelli, 1983), while Dictus er al. (1984) and Tetteroo et al. (1984) studied the lateral mobility of plasma membrane lipids in the Xenopus egg, by means of photobleaching experiments. Their measurements showed that regional differences related to animal-vegetal polarity are more than 100 times enhanced upon fertilization which lead to the formation of two distinct macrodomains within the plasma membrane. The new furrow membranes have properties different from the preexisting animal and vegetal plasma membranes. Preliminary work has shown that the lipid immobility of the animal half persists after the first cleavage and up to at least the 32-cell stage. The mechanisms by which the membrane lipids and ion pumpskhannels are immobilized in the oolemma have not as yet been studied. Gadenne er al. (1984) reported that in Xenopus from early blastula until neurilation, the lateral mobility of proteins and lipids increases significantly. The second element of asymmetry is asymmetrical mitosis (cleavage). A common feature of early cleavage is that mitosis very soon becomes asymmetrical. In the human embryo, the first cleavage already yields a larger and a smaller cell (Langman, 1976), the membrane properties of which have as yet not been analyzed. In Xenopus, as in other amphibians, the first cleavage takes place in the plane of bilateral symmetry, yielding two identical blastomeres. The second cleavage is again in a vertical plane, perpendicular to that of the first cleavage. The result is four cells, of which only two contain half of the grey crescent. Thus two different pairs of cells are formed. The asymmetry of the third cleavage, which takes place in a horizontal plane, is very pronounced: four small cells and four large ones are now formed. Other examples of asymmetrical cleavage are found in Dentaliurn (mollusc), Ilyunassa (mollusc), fish, and Branchiostoma, etc. (Balinsky, 1975; Grant, 1978). Because the properties of the plasma membrane are asymmetrically distributed and, as is the case in Xenopus, fixed up to at least the 32-cell stage, blastomeres are formed with identical nuclei (Spemann, 1938; Gurdon, 1962, 1974) but with different plasma membrane properties (ion pumps, ion channels, lipids, and perhaps other membrane proteins, such as receptors, etc.). For plants, the topic of polarity and asymmetrical mitosis was already re-

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viewed in 1958 by Biinning. More recently, Sugimoto and Yasuda (1983) emphasized the importance of asymmetrical mitosis for differentiation of thymic lymphocytes. Asymmetrical cell division may be primarily involved in the maintenance of immature “stem cells” while concomitantly producing differentiated cells of smaller sizes. The most extreme example of asymmetrical cytokinesis is the pinching off of the polar bodies from the oocyte during meiosis. Asymmetrical mitosis may as well result in a simultaneous asymmetrical distribution of some cytoplasmic or yolk components. The question, then, is what is the relative importance of the asymmetry generated in the plasma membrane and in the cytoplasm/yolk? Yolk itself, although it contains the germ cell determinant, does not seem to be a major factor governing differentiation (some centrifugation experiments, normal development of vitellogenin-free eggs obtained in male Bombyx, Yamashita and Irie, 1980; cytoplasmic streaming in Drosophda, Gutzeit and Koppa. 1982). In some organisms, some molecules as, e.g., specific proteins or maternal mRNAs (Jeffery, 1984a, review 1984b; Cox et a / . , 1984), might perhaps have become anchored at precisely located cytoplasmic regions, which are not subject to cytoplasmic streaming. They would also become asymmetrically distributed over the blastomeres as the result of the double asymmetry principle and perhaps be a causal element of differentiation (Berry, 1985). The question then is how have these mRNAs or proteins reached their specific location after being released from their site of synthesis? One possibility might be that they do so as the result of self-electrophoresis which seems to occur in all developing oocytes but in this case the asymmetry of the oolemma would still be the primary cause of differentiation. Another possibility is the phenomenon of high-affinity binding to specific cytoplasmic structures. Attempts are being currently undertaken to find out whether or not maternal mRNAs are randomly distributed in the egg, but this problem will not be easy to solve. The existence of gradients is well documented in embryology. Horstadius (1952) introduced data on gradients in sea urchin eggs. Since then, a lot of work has been done on the search for chemical (macromolecular) gradients. Only a limited number of naturally occurring inductors have been identified, but mostly riot in a completely pure form (e.g., Tiedemann, 1981; Yamada, 1981). The question can now be raised whether embryologists have not overlooked the existence and importance of electrical/ionic gradients, which may, in their turn, cause secondary macromolecular gradients. In our opinion the basic idea underlying differentiation in general is the formation of daughter cells with identical nuclei but with different plasma membrane properties (De Loof, 1985a,b). Asymmetrical distribution of maternally derived cytoplasmic factors over the blastomeres may be an additional element in some organisms. A number of data, suggesting a causal relationship between nuclear activity, plasma membrane properties, ions, and externally applied electric fields, will be discussed in Sections 111 and IV.

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111. Selected Functions of Ionic Currents In the next sections we will review the functions which have been attributed to ionic currents. The reader will see that often complete proof for a given function is, as yet, not available. This is mainly due to the very nature of ionic/electrical phenomena themselves: since they are membrane bound, disturbance or disruption of the membrane destroys them. This means that for most purposes intact cells have to be studied, but as most somatic cells are so small, analyses on such intact cells are very difficult. Furthermore, changes in one ionic species are usually linked to simultaneous changes in other ionic species. Indirect evidence is, so far, often the only way to suggest possible functions.

A. TRANSCELLULAR IONICCURRENTS: SIGNALS AND EFFECTORS OF POLARITY 1. Polarized Transport in the Cytoplasm Morphological polarity, which is characteristic of many cell types, e.g., neurons, epithelial cells, etc., is very well documented from histological research. Scanning electron microscopy also revealed such polarity in cells which showed no sign of this condition when viewed under the conventional microscope. For example, neutrophils are highly polarized, pseudopods spread out from one pole of the cell, while the plasma membrane overlying the nucleus has a markedly ruffled appearance (Klebanoff et al., 1978). “Active polarity” of cells may be visualized in a variety of different ways. The vibrating probe technique demonstrated that many cell types are electrically polarized. This has recently been reviewed by Nuccitelli (1983). Polarized transport of ions from one fluid compartment into another is also very well documented. Migration of pigment granules in melanophores and release of neurotransmitter by the presynapse are other examples. Many cell types synthesize proteins which are secreted into the external medium. The release site is usually very well defined. After packaging in the Golgi region, the secretory granules must somehow be transported to the site of release. The exact mechanism is as yet unknown. For some time, it has been thought that the cytoskeletal elements would actually cause the transport by the process of treadmilling. Although the cytoskeleton is a very flexible and permanent structure, to our knowledge, no evidence has so far been obtained, in vivo. in favor of “treadmilling” by any method, including the new very sensitive immunogold techniques combined with videocamera analysis (De Brabander et al., 1985). It seems plausible that self-electrophoresis may be the major force which makes polarized transport of secretory granules, and charged macromolecules in general, possible. For self electrophoresis to be possible, ion pump/ion channel ac-

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tivity should be asymmetrically distributed over the plasma membrane. This is clearly the case in many cell types as has already been discussed (Section 11,E). The basic questions are as follows: Are these transcellular electrical ionic currents just epiphenomena, are they the result of polarity, are they causally related to the generation of polarity, or are there combinations of these possibilities? At what moment in development is polarity established in a given system (Jaffe and Nuccitelli, 1977; Jaffe. 1982)? It is not probable that a universally valid answer can be given: the time at which an asymmetrical distribution of ion pumps/channels is established may vary from cell type to cell type. In developing insect meroistic ovarian follicles, transcellular currents can be measured from the very early beginning of yolk deposition. Full grown eggs of the algae Fucus and Pelvetia, on the other hand, are essentially apolar (Jaffe, 1966). As the result of various stimuli, e.g., illumination of no matter which side, in the course of a day or less, growth is initiated at one pole (the illuminated one), the cell visibly polarizes and divides into two quite different cells: an attachment cell or rhizoid at the growth pole and a thallus cell at its antipode. The result of the induction, e.g., caused by unilateral illumination, is that part of the egg cell membrane becomes more permeable to certain ions that are normally at a much higher electrochemical potential outside or inside the cell. In fucoid eggs, the inward current which is measured during germination is carried by Ca2 (influx) and Cl - (efflux) while K seems to be the probable outward current carrier (Jaffe, 1966, 1968, 1975; Nuccitelli, 1978, 1983; Jaffe and Nuccitelli, 1974). Electrogenic ionic flux through the resistance of the cytoplasm under the leaky membrane region will necessarily generate a cytoplasmic field that is relatively positive under that leaky portion of the membrane. A result of this field is the generation of movement: vesicles and other cytoplasmic constituents with a negative surface charge will be pulled toward the leaky membrane region. When these vesicles, which might be themselves relatively leaky to particular ions, are inserted into the membrane, this latter will again become more permeable resulting in further ion fluxes. Such a positive feedback loop would serve to establish and maintain localized growth expansion (Jaffe e? al., 1975; Jaffe and Nuccitelli, 1977). The release of synaptosomal vesicles, containing neurotransmitter(s), into the synaptic cleft, may also be, at least partially, due to self-electrophoresis. Upon depolarization there is an inward current of Na and Ca2 in the presynapse and the field created in this way may be essential in making the vesicles move toward and make contact with the membrane. Another example of polarized transport is that of rhodopsin which is synthesized in the inner segment of the rod cell but which is concentrated in the outer segment. In the isolated rat retina it has been shown that in darkness a steady current of the order of 70 pA flows inward through the plasma membrane of the rod outer segments which is balanced by an equal outward current distributed +

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along the remainder of each rod. Flashes of light produce a photocurrent which is generated primarily in the zones of the outer segment being illuminated, and which transiently reduces the ionic current. The unilluminated zones of the outer segments, as does the rest of each rod cell, act as sinks for the photocurrent. It has been estimated that there is a 2 mV cytoplasmic voltage drop across the neck region (Hagins et al., 1970). All elements for transport by self-electrophoresis are thus present (Section III,D, Fig. 6). Szego and Pietras (1984) describe what they call the “truly extraordinary” phenomenon of perinuclear polar accumulation of large numbers of lysosomes within seconds to minutes of plasma membrane perturbation by ligand (steroids, peptides, etc.) binding: the lysosomes form a cap at one pole of the nucleus. The nature of the force for this translocation is so far unknown but since membrane perturbation may result in changes in transcellular ion fluxes, it might perhaps be electrical. There are two well-documented examples of induction of polarity in egg cells by inducing changes in permeability to ions of the plasma membrane. When the eggs of the brown alga Pelvetia were grown in a gradient of the Ca2 ionophore A23187, they tended to form their rhizoidal outgrowths on the sites that were exposed to the higher concentration of ionophore (Robinson and Cone, 1980). This is in agreement with the earlier findings of Jaffe and his students on this system. The occurrence of crescents (the grey crescent) is well known in amphibian and ascidian eggs. Eggs of the ascidian Boltenia villosa normally form one orange crescent. When these eggs were exposed to a gradient of the ionophore, the formation of the orange crescent was induced in the region corresponding to the highest concentration of A23 187. Eggs lying between two glass fibers releasing the ionophore often showed the formation of 2 orange crescents half the size of normal, and this within 30 minutes after exposure. The artificially induced crescents contained both pigment granules and mitochondria as do normal crescents. Treatment with A23 187 results in abnormal development (Jeffery, 1982). It is also interesting to note that calcium accumulation was also observed within the growing tips of pollen tubes (Jaffe et al., 1975; Reiss and Herth, 1978) and that A23 187 treatment induces many lateral branches in Neurospora (Reissig, 1977). These results point to the importance of the differential permeability to ions of the oolemma and the possible role of Ca2+ gradients in the polarization process. +

2. Electrophoresis in the Plane of the Membrane (Lateral Electrophoresis) According to the fluid mosaic model of Singer and Nicholson (1972) proteins would be free to migrate laterally unless there are means present to restrict this mobility. When a cell with a fluid membrane is brought into a steady electric field, charged membrane proteins should be redistributed within the plane of the

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plasma membrane by lateral electrophoresis. Jaffe (1977) introduced the idea that rather small extracellular fields could polarize the distribution of freely mobile proteins. According to his calculations, a population of 10 pm membrane particles would become one-tenth to one-half polarized in 3 hours by a steady voltage drop of only 0.8-4.0 mV across the cell. Such a drop is present over several epithelia. Since it usually is very hard or even impossible to do the experiments on lateral electrophoresis in intact organisms, experiments have been designed which try to mimic the supposed in vivo conditions as closely as possible. The first confirmation of Jaffe’s theoretical prediction came from Po0 and Robinson (1977), who showed that in Xenopus muscle cells in culture, the concanavalin A receptor could be grossly polarized within 4 hours in a 400 mV/mm field (I .2 mV/cell), independent of cell metabolism. The Con A receptor is a negatively charged glycoprotein, which moved toward the negative pole, thus just ‘opposite to the expected direction. When sialic acid residues were removed by neuraminidase, or when the cell’s surface charge was changed by means of 3,3’-dioctadecylindocarbocyanine,a positively charged lipid, the direction of migration of the Con A receptor in an imposed field was toward the positive pole. The cause of this “anomaly” seems to be due to electroosmosis (McLaughin and Poo, 1981). In the same cell type acetylcholine receptors are also partially accumulated toward the cathodal pole in a uniform field of 10 V/cm but in contrast to the Con A receptor, the ACh receptors accumulated by the electric field form stable localized receptor aggregates (Orida and Poo, 1978, 1980; Po0 et al., 1979). Lateral electrophoresis has also been observed in sea urchin eggs (Robinson, 1977), mouse fibroblasts (Zagyansky and Jard, 1979), frog neurons (Pate1 and Poo, 1984), and mouse macrophages (Orida and Feldmann, 1982). Lectin receptors of Phaseolus vulgaris (Orida and Feldmann, 1982), ricin receptors (Zagyansky and Jard, 1979), freeze-fracture membrane particles in Micrasterias (Brower and Giddings, 1980), and Fc receptors of rat basophilic leukemia cells (McCloskey er al., 1984) are also redistributed by an imposed field. Lateral electrophoresis might be important in bringing hormone neurotransmitter and neuromodulator receptors to the site where they can be reached by their respective ligands. If lateral electrophoresis would be operating during the very early stages of embryonic development, it might cause an asymmetrical distribution of membrane receptors over the different blastomeres. Protein diffusion in cell membranes, its biological implications, and the effects of lateral electrophoresis have been extensively reviewed in the last decade. Recent reviews are those of Andersson (1979), Edidin (1981), Po0 (1981), Jacobson and Wojcieszyn (1981), Peters (1981), Webb er al. (1982), Axelrod (1983), McCloskey and Po0 (1984), and, especially for excitable membranes, Po0 (1985). Several intrinsic proteins are not subject to lateral diffusion, probably because they are anchored to the cytoskeleton (Almers and Stirling, 1984; Su et al., 1984).

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B. IONICCURRENTS: ARETHESECAUSALLY RELATED TO THE GENESIS OF BODYFORM? The typical form of a plant or an animal is evidently coded for in the DNA. How the information in this linear molecule is translated into mechanisms that inform the developing embryo where to form the forelimbs or the tail, for example, and what forces guide the differentiating cells to their final position within the organism remain to be determined. There are some indications that electrical currents produced by the cells of the developing embryo itself may play a major role. Jaffe and Stem (1979) explored the electrical fields above chick embryos with the vibrating probe technique. Steady currents with exit densities of the order of 100 pA/cm2 leave the whole primitive streak (which is in fact an elongated blastopore), the epicenter lying near Hensen’s node, and return elsewhere through the epiblast. They are probably pumped into the intraembryonic space by the epiblast and then leak out of the primitive streak zone because this latter is a zone of continuous junctional disruption. More recently, Robinson and Stump (1984) reported that the epicenter of the outgoing current in Xenopus neurulae (stages 14-22) is the blastopore, which apparently provides a low-resistance (roughly 1 M a ) leakage pathway for ions pumped inward across the skin. They found, with the vibrating probe, that inward current exists at all other regions of undamaged embryos. A minimum of inward current near the middle of the neural groove was shown, this being consistent with the dissolution of tight junctions which occurs as neurulation proceeds (Decker and Friend, 1974; Decker, 1981). Thus, in both chick and Xenopus embryos, the region of outward current is a site of tissue involution (Keller, 1976), with the cells migrating in a direction opposite to that of the electric field vector. Na+ is essential for the current in Xenopus and at neurulation the Xenopus embryos possess a Na+ transport system similar to that found in adult frog skin. The leak at the blastopore allows the epithelium of the neurula to arrive at a near short-circuited condition (Robinson and Stump, 1984). A role which has been suggested for these large currents is neurite and myoblast guidance but the question as to whether the currents are large enough to be effective in this respect is still unanswered. Several reports have shown that in vitro weak externally applied electric fields of the order of 1 mV per cell diameter can influence cell behavior and redistribute membrane proteins (lateral electrophoresis, Jaffe and Nuccitelli, 1977). Marsh and Beams (1946) and Jaffe and Po0 (1979) observed that the neurites of explanted chick ganglia grow preferentially toward the cathode. Goldfish retinal neurites also turn and grow toward the cathode (Freeman el al., 198 1). By means of a novel circulatory vibrating probe, Freeman et al. (1985) measured the steady currents generated by goldfish retinal ganglion cell growth cones. Current, predominantly carried by Ca2+, enters the tip of the filopodium, flows

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axially down it and back outward near the base, completing the loop in the extracellular medium. This could at least partially explain the behavior of growth cones in an electric field. Hinkle et ul. (1981) found that polarity and differentiation of explanted Xenopus myoblasts and neuroblasts are influenced by the presence of a small electric field. The field causes myoblast elongation perpendicular to the field vector (threshold roughly 36 mV/mm), while neurites of cultured neuroblasts grow toward the direction of the cathode (threshold about 7 mV/mm). These authors also found that a larger fraction of the explanted neural tube cells differentiate into neurons in the presence of an electric field. Stump and Robinson ( 1983) observed that Xenopus neural crest cells migrated toward the cathode in an applied electrical field of 10 mV/mm or greater. Such a field corresponds to a voltage drop of < I mV across the diameter of the cell. If the culture medium and conditions could be further optimalized the strength of the field might perhaps be substantially reduced. Stump and Robinson (1983) and Robinson and Stump (1984) discuss the possibility that the current pathway (produced by the skin battery) in the Xenopus neurulae runs from the ectoderm through the mesoderm and endoderm, into the fluid-filled regions of the archenteron and then out of the blastopore. The pathway followed by the currents depends on the resistivities of the internal tissues and on the location of the ‘‘leaks’’ that allow current to flow into the archenteron and then out of the blastopore. The direction of the electrical field is at least crudely consistent with the direction of neurite and myoblast orientation. They calculate a lower limit value for the electrical field within the embryo to be around 0.18 mVlmm, which is about one or two orders of magnitude below the threshold for orientation found in vitro (Hinkle et al., 1981). However, the in vitro threshold conditions may be an overestimate since in vivo conditions such as substrate adhesion cannot be reproduced. Furthermore, the effect of constraining the cells to a two-dimensional surface may raise the threshold for orientation. Perhaps some of the target structures for the neural crest act as current sinks, thereby maximizing current flow toward themselves. Stump and Robinson (1983) think that this could, for instance, explain neural crest aggregation to form plexuses in the aorta or colon. They also suggest that this does not need to be an active process but could result from something as simple as an absence of tight junctions in some regions. Quail somite fibroblasts and neural crest cells exhibit a two-stage response to an imposed steady electric field. Within 10 minutes they begin to align with their long axes perpendicular to the field lines (threshold 150 mV/mm, only 3 mV/cell width); they then begin migrating toward the cathode (Nuccitelli, 1983; Nuccitelli and Erickson, 1983). The responses of cells to electrical fields have been recently reviewed by Robinson (1985). At the sites were the limbs will appear in embryos of Xenopus (Robinson, 1983) and axolotl’s (Borgens er al., 1983) large outward currents were measured with the vibrating probe, and this before there was any visible sign of limb

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formation (Robinson, 1983; Borgens et af., 1983). Large currents are also present during wound healing and during regeneration of amputated limbs (Section 111,F). In wild carrot, Daucus carota, endogenous currents traverse the embryo. By means of the vibrating probe, it has been shown that current enters the apical pole and leaves the region near the presumptive radicle. This polarity precedes differentiation of vascular tissue and cotyledon development. In subsequent stages of embryogenesis, inward current is found at the cotyledons and outward current at the radicle/root (Brawley et a f . , 1984). In a vertical, freely growing root of Lepidum sativum, current enters the root cap, the meristem, and the beginning of the elongation zone. It leaves the root along most of the elongation zone and in the root hair zone (see Fig. 3 for barley). After the root is tilted to a horizontal position, within about 30 seconds in most cases, current was found to flow acropetally at the upper site of the root cap and basopetally at the lower side. In the following minutes the current pattern undergoes further changes (Behrens et af., 1982). Local irradiation of Vaucheria tubes with blue light results in the establishment of a new outgrowth at the irradiated site. Here again, in most cases large currents precede the outgrowth. This and other examples have been described by Weisenseel and Kicherer (1982). The idea has emerged that electrical currents produced by the developing embryo itself-although very weak-might be the major determinant in the establishment of the body form typical for each species (Mackenzie, 1982). According to Borgens (1984), limb development and limb regeneration might both be initiated by an integumentary wounding.

C. IONS AND NUCLEAR ACTIVITY Electrophysiological studies usually focus exclusively on effects at the level of the plasma membrane. Studies on gene expression mainly focus on events in the nucleus but only very rarely is any attention paid to possible effects mediated by the plasma membrane or the nuclear envelope.This dichotomy in approach, the very different methodologies, and specialized terminologies (languages) used by electrophysiologists and biochemists make integrated research projects, in which the interrelation plasma membrane-nuclear activity is studied, almost nonexistent. That nuclear activity is somehow-at least partially-controlled by the plasma membrane is self-evident. It is at the level of this membrane that changes in the cell’s environment are first experienced. It is here that stimuli have to be translated into a message that can be understood by the nucleus so that, at this site, commands can be given as to which proteins have to be synthesized in order to cope with the changes in the environment. The question then is which messenger system is used between the plasma membrane and the nucleus? From a theoretical viewpoint the following considerations are important (De Loof and

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Geysen, 1983; De Loof, 1985a-c). Factors localized in the plasma membrane which have the capacity to influence nuclear activity must do so either by moving out of the membrane to the nuclear compartment as primary messengers, without making use of secondary messengers, or by using secondary messengers without being displaced from the membrane. Taking into account the large numbers of genes present in the genome of all organisms, the versatility of the messenger system must be very high. This could be realized either by a large number of different primary messengers which would act on specific genes or by variable balances of a rather limited number of secondary messengers or by a combination of both. That proteins could provide a ubiquitous system of secondary messengers is not very likely. Indeed, proteins themselves are gene products and their synthesis must also be controlled by other regulators. In the case of these again being proteins, one enters a system which could go on ad infinitum but which most probably would not function properly. Cyclic AMP, inositol triphosphate, and Ca2 do function as secondary (Ca2 perhaps tertiary) messengers; they occur in all cells (Berridge and Irvine, 1984) and they are messengers for a variety of different hormones, peptides, as well as steroids. It is not possible, however, to generate a highly versatile system with only three factors. Already, more than 20 years ago, Kroeger (1963a,b), who studied the mechanism of puff induction in giant chromosomes of the dipteran Chironomus, found that replacement of Na+ by K + in the explant medium in which the salivary glands were cultured led to a progression of the puffing patterns closely resembling that which in normal animals was supposed to be caused by the rising titer of ecdysone, the steroid molting prohormone of insects. Puff 111 d , , which normally appears upon ecdysone treatment, could also be induced by explantation of the glands into an ecdysone-deficient medium containing a relatively high potassium concentration. Kroeger also found that there were rather specific Na+ , Ca2+, and Mg2+ puffs. Kroeger’s basic idea was that changing balances of “simple” ions such as Na , K , Ca2 , Mg2 , H , C1- , etc. would act as an extranuclear (epigenetic) control mechanism of gene expression: in other words, ions might be a universal messenger system between the plasma membrane and the nucleus. This hypothesis, formulated shortly after the Jacob and Monod (196la,b) theory on the regulation of the lac operon, has quite often been misunderstood and has been subject to lively controversy (Ashburner and Cherbas, 1976; Kroeger, 1977). Kroeger’s hypothesis deals with epigenetic mechanisms and does not exclude that there are also additional mechanisms operating at the level of the DNA/RNA. The major objection to Kroeger’s-in our opinion very valuable-hypothesis was that it was very difficult to understand how a few universal ions, present in all cells, could harbor the selectivity to activate specific genes. Kroeger’s obser+

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vations were not entirely unprecedented. Barth et al. (1960) had already reported that pigment cells could be easily induced from presumptive epidermis from Ram pipiens by a culture medium in which 50% of the NaCl was replaced by KCl, herewith bringing into doubt former data which suggested that induction was due to organic molecules (Barth, 1964). Later, Barth and Barth (1969, 1972, 1974a,b) reported that small aggregates of cells prepared from explants of ventral ectoderm of R. pipiens gastrula could be induced to differentiate into a variety of cell types just by altering the ionic composition of the medium in which they were treated or cultured. One of their most remarkable results they obtained was that increasing the K + concentration in the culture medium caused lithiuminduced ectoderm cells to differentiate into notochord (Barth and Barth, 1974b). More recently, Rosoff and Cantley (1983) indirectly showed that Kroeger’s concept might be valid. They found that by pharmacologically increasing the cellular Na+ content by monensin, a Na+ -ionophore, or by ouabain, a specific inhibitor of Na+ ,K+-ATPase, surface IgM expression could be induced in a pre-B lymphocyte tumor cell line. If only ionic concentrations were involved and if each gene were to be regulated independently, the objection against Kroeger’s concept would have been partially justified. It has taken nearly two more decades before it has become clear that steroid hormones, often considered as inducers of specific genes which act directly at the level of the DNA, exert their first effects at the level of the plasma membrane, where they change the permeability to certain ions (for list of references, see Szego and Pietras, 1984). It has also become clear that the action of steroids is usually not on individual genes but on sets of genes. This was clearly shown for the steroid hormone 20-OH ecdysone. Poeting et al. (1982) detected 26 20-OH ecdysone-induciblegenes in the salivary glands of third instar larvae of Drosophilia melanogaster and 11 in the larval fat body in vitro. Only one protein was found to have similar properties in both tissues. Furthermore, the development of the vibrating probe by Jaffe and Nuccitelli (1974) emphasized that cells are dynamic units, and that besides ionic concentrations, one also has to take into account ionic and potential gradients, ionic currents, and secondary chemical gradients. Neither should organic ions be overlooked. The factors which are involved in controlling the ionic environment of the genes are schematically summarized in Fig. 5 . From this figure, it is clear that although few ionic species are involved, the number of possible combinations is extremely high. The versatility of the system is at least as high as that of the genetic code itself, an essential prerequisite for any system of epigenetic control of gene expression. A system of epigenetic control of gene expression by ions would have several advantages. It is universal and can be used by all organisms. It is based on a physical principle and does not-apart from ion pumps and channels-depend upon gene products. It allows the cell to cope simultaneously with a multitude of stimuli, both physical and chemical, in the external environment. In other words,

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it is a system which can handle a balance of factors and that is exactly what happens in all cells. Each cell has to continuously adjust itself to its changing environment. From the analysis of the versatility of the system shown in Fig. 5, it seems unlikely that in an organism there are two tissue or cell types in which all chromosomes experience an identical ionic environment. According to De Loof (1985b) this is a major reason why the different cell types of an organism, although all having the same genome, do activate different sets of genes; this may also be the major explanation for tissue specificity of hormone action. The effects of ions/electrical currents on nuclear activity may be situated at different levels, e.g., the interaction with DNAIhistones, nonhistone proteins, the nuclear skeleton, and the activity of nuclear enzymes. Active genes have, in general, a more open configuration than inactive genes. Usually, but not alwyas, a specific chromatin structure, detected as a site hypersensitive to cleavage by DNase I, appears to be necessary, but not sufficient, for gene activation. In a gene in the process of being transcribed, the interaction between DNA and histones has been altered and it has been suggested that many of the nonhistone chromosomal proteins play a key role in determining the general chromatin structure of such sets of genes related to their active state (Lewin, 1980; Cartwright et al., 1982; Nicolini, 1983). A gene will be transcribed when the initiation sequence and all control sequences in the upstream and/or perhaps in the gene itself are accessible so that the RNA polymerase can align itself in good order and start transcription. There is, as yet, not enough experimental evidence and no consensus as to the question of whether it is the spatial conformation of the controlling upstream (or in the structural gene itself?) regions or the specificity of some base sequences or both which are the controlling elements. At this moment, the suggestion that spatial conformation-which evidently will partially depend upon the base sequence-is the key factor is at least as valuable as any other hypothesis. The question, then, is how is the right conformation realized? Is it either by the interaction between specific parts of DNA and certain compounds such as, e.g., hormone-receptor complexes, or by changes in the ionic environment, or a combination of both? Ions may play a role in the transition between the different higher order structures of chromatin. Ions may affect the chemical interactions between DNA, histones, and nonhistone proteins. This was the explanation suggested by Lezzi and Gilbert (1970) and Lezzi (1970) for the selectivity of puff induction in isolated salivary gland chromosomes of Chironomus by certain ions. Although today the exact mechanism of puffing is not yet completely understood, these experiments clearly showed that different regions of the chromosomes do react in different ways, to certain ions. Two regions were observed, I-19-A and region I-18-C. If isolated chromosomes were exposed to 150 mM NaCl (plus 12 mM MgCI,), certain bands in the I-19-A region became diffuse, whereas the I-18-C region remained unchanged. Upon exposure to 150 mM KCI (plus 12 mM

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MgCl,), it was the band in the I-18-C region that became diffuse while the 1-19A region remained unchanged. The specific effects of Na+ and K + on the structure of the specific bands were fully dependent on the presence of Mg2 in the incubation medium. The specificity of Na+ and K + was further enhanced by addition of Ca2+ (1-9 mM). Actual RNA synthesis was not required for the induction by Na+ or K + of specific structural changes as this took place in single salt solutions in the absence of the four ribonucleoside triphosphates and exogenous RNA polymerase. Lezzi (1970) suggested that the swelling of a band, caused by a medium of a moderately high ionic strength, is a prerequisite for the initiation of RNA synthesis at the respective site. This view is supported by the fact that chromosomes that are expanded in all their bands by being placed in a medium of high ionic strength synthesize RNA over their entire length. Specific inducers such as hormones or receptor complexes are thus not essential under these conditions. Lezzi (1969) also performed experiments on isolated rat liver chromatin and was able to show differential effects of Na+ and K + on template activity. These indirect effects on chromatin structure may be due to the effects of ions on nonhistone proteins. About 40% of the nonhistone chromosomal protein mass seems to consist of myosin, actin, troponin, and tubulin (Douva et al., 1975; Pederson, 1977). In addition, calmodulin has been found in manually isolated nuclei of Xenopus (Cartaud and Ozon, 1980). In the event that changes in Ca2+ concentration in the nucleus are generated, it may well be that contractionrelaxation of contractile proteins occurs. It may not be excluded that this is one of the factors involved in the higher order transitions of chromatin. Isolated nuclei of Chironornus salivary gland cells quickly swell when brought into a hypertonic solution and shrink in a hypotonic solution: this system is reversible (H. Kroeger and M. Lezzi, personal communication). Whether this is due to a reaction of the nuclear skeletal proteins (the contractile chromosomal proteins) or to changes in hydration is as yet not documented. Changing the volume of the nucleus may be a mechanical means of changing the structure of some chromosome regions. Nuclei transplanted into unactivated amphibian eggs are known to condense into metaphase chromosomes whereas those transplanted into activated eggs decondense and enlarge. An essential element of activation is the increase in intercellular Ca2 (Jaffe, 1983). Chromatin of sperm nuclei injected into unactivated eggs is induced to form metaphase chromosomes apparently because of low Ca2+ concentrations. When Ca2+ concentration increases, as is the case after fertilization, the chromatin deconden'ses (Lokha and Masui, 1984). Ionic fluxes, perhaps, may also be causally related to pulsations and invaginations of the nucleus. In the germinal vesicle of the oocyte of the insect Heteropyza, constrictions of the nucleus occur periodically, through which the nuclear content is shifted to and fro. The duration of the pulses varied between 3 and 10 +

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minutes, depending upon the developmental stage (Went, 1977). Pulsation of these nuclei is reversibly inhibited by cytocholasin B (Kaiser et al., 1982). Nuclear invaginations are also regularly present during oogenesis of ferns and mosses (Bell, 1975) and the so-called accessory nuclei observed in developing oocytes of Hymenoptera and other insect orders are supposed to have originated by budding from the oocyte nucleus (King and Fordy, 1970). Yamada (1962, I98 1 ) observed that invaginations of regular shape and small magnitude occur in the nuclei of untreated ectodermal cells and that upon treatment with bone marrow factor (an inductor) these invaginations become irregular and much more extended. As they also occur in cryofixed material, they are probably not fixation artifacts. Changes in ionic concentrations and sometimes also of fluxes have been found in mitotic, cancer, and developing cells. A large number of experimental observations which suggested that a significant correlation may exist between the level of the Em value in somatic cells and their mitotic activity allowed Cone (1970, 1971a,b) to propose a unified theory of the basic mechanisms of normal mitotic control and oncogenesis. Herein the intracellular ionic conditions associated with various levels of E,, value are considered to regulate the preparation of DNA synthesis and other essential mitotic functions. The theory links the activity of the mechanisms which generate Em and which are evidently plasma membrane bound, with cellular metabolism and with external environmental influences through an explicit system of interacting feedback circuits (Cone, 1970). Low Em values and high [Na+Ii have a mitogenic effect. Em values in cultured cells varied as a consequence of neoplastic transformation and as a function of cell density (e.g., contact inhibition) in culture (Cone, 1970, 1971a,b; Cone and Tongier, 1973). In support of this theory, 2s-Nagy er af. (1981) found that in three types of invasive urogenital cancer cells, the average intracellular [Na J increased more than three fold, [ K + J decreased between 13 and 32%, and [CI- J also increased but less than "a+ 1. The intranuclear Na+ / K + ratios were about 5-fold higher in the cancer cells, all values relative to controls in normal cells. Moyer et a / . (1982) also found that in a large number of transformed or tumor cells, Na+ was most frequently, but not always, increased while [ K + ] was not consistently higher or lower. Walsh-Reitz er a/. (1984) reported that increased availability of Na+ in the culture medium stimulates cell growth in kidney epithelial cell cultures (BSC-I line). A cellular protein, which inhibits cell Na+ accumulation, blocks proliferation. Rat pheochromocytoma cells respond to nerve growth factor by the acquirement of a phenotype resembling neuronal cells. This factor causes an increase in Na+,K+ pump activity and it is this increased Na influx which stimulates growth (Boonstra et al., 1983). Na flux and its effect on intracellular K + and pH have a major role in the complex system that regulates proliferation of 3T3 mouse fibroblasts as shown by Schuldiner and Rozengurt (1982), Lopez-Rivas et ul. (1982), and Burns and +

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Rozengurt (1984). Transformation of these cells by polyoma virus made Em 13 mV more negative and the relative permeability coefficient of some ions was also changed (Killion, 1984). Voltage-gated K+ channels may play a role in mitogenesis in human T-lymphocytes (De Coursey et al., 1984). A rise in intracellular pH of about 0.3 pH units was observed in mitogen-stimulated murine lymphocytes and of about 0.5 pH during exponential growth in three virus-transformed lymphocyte cell lines (Gerson and Kiefer, 1982). An increase in the concentration of free Ca2+ during metaphase appears to stimulate the onset of anaphase. Such an increase, regulated by the cell itself, may contribute to the initiation of chromosome separation in mammalian cells (Izant, 1983). An efflux of Ca2+ from membrane compartments into the spindle before the onset of anaphase was found in endosperm cells of Haemanthus by Wolniak et al. (1983). Ionic concentrations and Emvalues not only change upon transformation of a cell into a cancer cell but can also change as a function of development as clearly shown in liver cells of mice. Maternal liver cells have a mean Em of -41 k 1 mV while the intracellular K + concentration is about 95 ? 7 mM. The respective values in fetal liver are -23 ? 1 mV and 62 4 mM. Both values in fetal liver are increased to values comparable to those of maternal liver during the first 8 days of neonatal life. The activity of a membrane Na+ ,K+ exchange pump thus seems to increase with development (Chapman and Wondergem, 1984). Na+ and K + accumulation also takes place during development of dorsal root ganglia from embryonic chick (Skaper and Varon, 1983). A large increase in the number of voltage-dependent K and Na+ channels was observed in embryonal carcinoma cells (PCC4) that were induced to undergo neuronal differentiation (Ebihara and Speers, 1984). In a recent paper, La Gamma et al. (1985) described that depolarization inhibits increase of preprometenkephalin mRNA in rat adrenal medullae grown in explant culture. It may be concluded that the intracellular ionic environment does influence nuclear activity. Until present, only direct data about ionic concentrations are available. Because it is usually very difficult to measure ionic and electrical gradients within the cell, their effect upon nuclear activity is still to be elucidated. Some indirect data on effects of currents are available from imposed fields (Section IV).

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D. EXCITABLE CELLS An excitable cell is a cell which readily and rapidly responds to suitable stimuli, and in which the response includes a fairly rapid electrical change at the plasma membrane. Nerve, muscle, sensory cells, and the electric organ of electric fishes are the best studied objects. Our present knowledge of the electrical

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dimension of cells is largely based on the brilliant pioneering research of Hodgkin, Huxley, Katz, and their collaborators. Squid giant axons have been extensively used by Hodgkin and his colleagues to unravel the mechanism of impulse conduction (for summary of earlier work, see Hodgkin, 1964; Aidley, 1981). The basis for the understanding of muscle contraction was especially laid by H. E. Huxley (for summary see H. E. Huxley, 1972) and A. F. Huxley (for summary see A. F. Huxley, 1974) and their co-workers. Katz (1966) and his associates studied the physiology of the neuromuscular junction. As there are excellent textbooks (e.g., Richter, 1979; Aidley, 1981; Petersen, 1980) on the physiology of excitable cells, it is unnecessary and indeed impossible to give here, in a few pages, a sufficiently documented summary of the very large number of data which are now available. Suffice it to recall, as most students of biology probably know, that nerve cells are electrically excitable because of the presence in their plasma membranes of voltage-sensitive ion channels that are selective for Na+, K + , or Ca2+ (Catterall, 1984). Influx of Na+ leads to depolarization of the plasma membrane. The increased intracellular Na concentration activates the Na ,K pump and the membrane can again repolarize. Different types of neurotransmitters are essential in chemically transmitting synapses, major ones being acetylcholine, serotonin, norepinephrine, dopamine, and y-aminobutyric acid (GABA). Ca2 influx in the presynapse is essential for the release of acetylcholine and other neurotransmitters. In contrast to what had long been assumed, acetylcholine is now believed to be released, at least partially, directly from the cytoplasm into the synaptic cleft and not from the synaptic vesicles. The synaptic vesicles would function primarily during the extrusion of Ca2+ (Dunant and Israel, 1985). There also exist electrically transmitting synapses (Aidley, 1981). Frog and rat muscle fibers have been extensively used to unravel the contraction mechanism of skeletal muscle. Resting Em values of frog and rat muscle in plasma are -99 and -90 mV, respectively, both values being close to E,. In skeletal muscle fibers both K + and C1- are close to an electrochemical equilibrium, whereas a strong Na+ pump is required to maintain a steady state for Na . Under some conditions a K pump is required to maintain a steady state for K + (Sjodin, 1982). Release of acetylcholine at the motor endplate causes influx of Na and Ca2 , Following a depolarization of the muscle fiber membrane, a transient rise in sarcoplasmic Ca2 concentration occurs which precedes a rise in muscle tension (Ridgway and Ashley, 1967). This Ca2+ is released from the terminal cisternae of the sarcoplasmic reticulum but the exact mechanism involved is still not fully elucidated. Possible mechanisms cited include depolarization, charge movements, and Ca2 -induced Ca2 release (Miyamoto and Racker, 1982). Ca2+ makes interaction between the actintropomysin-troponin system and myosin possible. Thereafter Ca2 is again taken up by the sarcoplasmic reticulum and recycled to the terminal cisternae +

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(Winegrad, 1968). The movement of internal Ca2+ and its role in excitationcontraction coupling has been extensively studied and recently reviewed by Grinell and Brazier (1981). Transport of electrolytes in muscle in general was reviewed by Sjodin (1982). Betz et al. (1980), Betz and Caldwell (1984), and Caldwell and Betz (1984) applied the vibrating probe technique to whole muscle and isolated muscle fibers of rat and frog. In whole muscle, outward current was detected near the middle of the muscle, and the maximum outward current was invariably located at the endplate zone. In the flanking regions membrane current flowed inward. In isolated muscle fibers the peak outward current also occurred at the endplate. This current is dependent on the activity of the sodium pump. According to Betz and Caldwell (1 984), the electric field created by this current may be important for long-term interactions between muscle and nerve. Another aspect of the physiology of excitable cells is the control of their nuclear activity (Thoenen and Edgar, 1982; Sutcliffe et al., 1984). For muscle this is also an unsolved problem. It is known that insulin, adrenaline, noradrenaline, and thyroid hormones influence muscular activity. Insulin causes membrane hyperpolarization probably by a direct effect on the Na ,K -ATPase and by creating new active pump sites (Erlij and Grinstein, 1976). Catecholamines also have a hyperpolarizing effect but by means of mechanisms other than those used by insulin. Thyroid hormone increases the rate of Na+ and K + exchange in rat skeletal muscle via an increase in membrane permeability to Na+ and K + . For details and references see the review of Sjodin (1982). Almers et al. ( 1983) found that in frog skeletal muscle the Na channels occur in oblong patches longitudinally spread over the fiber. In skeletal muscle the nuclei are located immediately underneath the plasma membrane. It cannot be excluded, but is as yet unproven, that the Na+ channel patches might be just above the nuclei. If this is the case, the nuclei would then experience a Na+ current during each depolarization. This might help to explain how “doing work” stimulates muscle development (De Loof, 1985b). If the same mechanism would be used by the heart, which is continuously at work, the heart muscle would hypertrophy. Remarkably, be it a coincidence or not, the nuclei in the heart muscle fibers (as is also the case in neurons) are not located close to the plasma membrane but far away from it and nearly in the middle of the contractile filaments where they probably will only minimally experience the ion fluxes taking place at the plasmalemma. The electric organs of the different types of electric fish are composed of columns of cells called electroplaques. In all species except Gymnotes carupo (Bennett and Grundfest, 1959), these electroplaques are modified muscle cells which have lost their contractile function and which are individually innervated by an excitor nerve. In the main electric organ of the electric eel, Electrophorus electricus, up to 6000 rows of electroplaques are arranged in series with each other. In Torpedo marmorata, the two electric organs are each made up of some +

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400 prisms, each of which is a stack of electroplaques. The electroplaques of the electric eel can be stimulated electrically which results in an all or nothing action potential appearing across the innervated face. As there is no potential change across the noninnervated face, the voltage produced by the stimulated electroplaque is due to the asymmetry of the response of its two faces. In marine electric fish, the electroplaques cannot be stimulated electrically, but they can by acetylcholine which is simultaneously emitted from all synapses, the number of these being estimated at more than 500 billion in Torpedo (Dunant and Israel, 1985). The ventral side of the Torpedo electroplaques has a high electrical resistance while the dorsal membrane has a low resistance. The neurotransmitter acts on receptor molecules of the electroplaques by opening Na channels on the ventral side only. The open Na+ channels allow Na+ to enter rapidly through each ventral membrane and it is this movement of Na which constitutes an electric current. As many electroplaques are arranged in series, a substantial current is propagated into the surrounding sea water where the ions derived from the salts present are induced to move by the electric field. The average amount of current produced by both electric organs of Torpedo can exceed 7 A. It is this system which gave rise to the new ideas concerning the release of acetylcholine from the presynapse (Dunant and Israel, 1985). The electric eel can produce electric discharges of 600-850 V (Aidley, 1981). Action potentials have been measured in a number of animal species at fertilization. This is described in Section 111,E.There are also a number of gland cells that possess voltage-sensitive ion channels. The examples known at present are mammalian pancreatic B cells. cells from the adrenal cortex, chromaffin cells from the adrenal medulla, cells in the anterior pituitary, and various gland cells from invertebrates. The number of gland cells yet to be explored in detail with microelectrodes is still very large (Petersen, 1980). The research which has already been done on gland cells, on the stimulus-secretion coupling, the pancreatic acinar cell, the salivary and lacrimal glands, and the forementioned electrically excitable cells is too extensive to be reviewed here. I refer to the excellent book of Petersen (1980) for details and references especially related to effects of hormones on the electrical properties of gland cells. Photoreceptors are also excitable cells in which rather complex ionic currents are involved in impulse transmission (Fig. 6). They also are an example of a cell which almost continuously drives an ionic current through itself and probably behaves as a miniature electrophoresis chamber. In vertebrates, illumination causes hyperpolarization; this in contrast to most other sensory cells, including the photoreceptors of invertebrates, where the appropriate stimulus causes depolarization. Light also causes reduction of the conductance of the cell membrane of the outer segment containing visual pigment to some ion or ions. It also alters the ionic current which flows continuously between the outer and inner segments. Very schematically, the system is supposed to function as follows. +

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disc

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IUTER EGMENT

ILIARY-

rnitochondi

\INER EGMENT‘

nvclws-

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FIG.6 . Representation of a rod cell and the changes in ionic current following exposure to light. Visual pigment is concentrated in the outer segment (a). In the dark (b) the permeability of the plasma membrane for Na+ is high. Light (c) induces a decrease in Na+ permeability and Ca*+ flows from the interior of the outer segment to the interstitial space. In d a purely hypothetical representation of initial charge configurations in the rhodopsin molecules in a dark-adapted state is shown: the comma represents the vectorial orientation of the molecules. Light causes simultaneous displacement of fixed charges (e) so that positive charges move perpendicularly to the plane of the membrane and toward the cytoplasmic interface and this without any change in the vectorial orientation of rhodopsin. The incremental change in charge density of each membrane surface sets a flow of ions in the extracellular and cytoplasmic spaces (f) into motion. As the fixed charges, now displaced, become neutralized by the diffusible ions, the currents diminish to zero. The result of exposure to light is hyperpolarization. Redrawn and partially modified with copyright permission mainly after Fein and Szuts in “Photoreceptors: Their Role in Vision.” Cambridge University Press, 1982. Some ideas of O’Brien (1982) were also used to draw this figure.

There seems to be a metabolically driven Na+ extrusion pump in the plasma membrane of the inner segment, where large numbers of mitochondria are located, and perhaps also in the nuclear region. Na+ channels are present in the outer segment and probably also in the synaptic region. In the dark the Na+ channels in the outer segment are open and a rod is kept depolarized. The pump

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serves to maintain the ionic gradient of Na+ which provides the driving force for the flow of current into the outer segment and from there through the interior of this segment through the ciliary segment back into the inner segment. In the ciliary segment, the diameter of which is about 0.3 pm, the current density must be of the order of about 500 pA/pm2, which is 20 times higher than in an axon during an action potential (Aidley, 1981). Light seems to reduce the influx of Na+ ions into the outer segment which results in hyperpolarization of this segment. The absorption of one photon prevents about one million ions flowing through the rod membrane. An explanation for this phenomenon may be that light, through the mediation of rhodopsin, causes Na+ channels to close, which could account for the decreased conductance (Aidley, 1981). Ca2 also seems to be involved (Hagins, 1972). Rods contain large amounts of Ca2+ concentrated in their outer segments. Illumination causes a graded efflux of rod Ca2+ content (O'Brien, 1982), amounting to lo4 ions per rhodopsin molecule bleached in dim light (Schroder and Fain, 1984). Ca2+ has a strong effect on the number of open light-sensitive channels (Hodgkin et al., 1985j. While the view was long held that light simply releases internal Ca2+ to bind to and block the light-sensitive conductance, Yau and Nakatani (1984b) showed that there is an electrogenic 3 Na+ : 1 Ca2+ exchange in outer segments. Rods also have a light-sensitive K + conductance (Yau and Nakatani, 1984a). However, there is also the effect of change in distribution of positive (about 30) and negative (about 50) charges within the rhodopsin molecule caused by light. The fixed charges on the molecule contribute to the charge density carried on each membrane surface. They are neutralized by diffusible counterions from the surrounding aqueous solutions. The counterions are mobile charges which are set in motion by a voltage gradient, thereby creating a current. A brief flash of light causes a simultaneous displacement of positive charges toward the cytoplasmic surface of the membrane. The result is that the cytoplasmic side picks up a partial positive charge and the extracellular side becomes commensurately more negative: an incremental voltage appears across the region in which the rhodopsin is embedded. These charge displacements occur without any molecular reorientation and without any ionic current flow across the membrane. However, ionic current flow will be initiated in the surrounding solutions by the resulting incremental voltage. The diffusible anions and cations counterflow until the excess and debit charges are once again neutralized at the membrane-water interface (Fein and Szuts, 1982). The electrical signal produced by closing the Na+ current in the outer segment spreads to the inner segment and through gap junctions into neighboring rods. As the signal spreads, it is modified by additional currents that flow through each inner segment. At the synaptic base, the modified signal causes closure of channels through which Ca2+ enters. This results in a fall in intracellular Ca2+ concentration which interrupts the continuous exocytosis of transmitter-laden vesicles. The decrease +

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in the rate at which transmitter molecules are liberated acts as a message that is communicated to adjacent neurons (Schwarz, 198 1). E. ELECTRICAL PROPERTIES OF OOCYTES DURING GROWTH AND MATURATION AND OF EGGSAT FERTILIZATION Parts of this topic, which attracted much interest in recent years, have been reviewed by Masui and Clarke (1979), Hagiwara and Jaffe (1979), Shapiro (1981), Whitaker and Steinhardt (1982), Hagiwara (1983), Shen (1983), L. F. Jaffe (1983), Jaffe and Cross (1984), Nuccitelli and Grey (1984), and in the Ciba Foundation Symposium Vol. 98 (1983). Oocytes (before maturation) and egg cells (after maturation) possess ionselective channels activated by membrane voltage (Hagiwara and Jaffe, 1979; Hagiwara, 1983; Barish and Baud, 1984) which show similarities as well as differences to those found in cells of differentiated organisms (e.g., the Ca2+ channel in ascidians: Hirano and Takahashi, 1984). At this moment numerous indications for a role of inorganic ions in vitellogenesis, maturation, fertilization, and activation are known. A single, uniform mechanism for their action cannot yet be proposed. Ca2+ seems to be important for activation in all species so far studied. 1. Excitability of the Oocyte Membranes

The Em of animal oocytes is in the range of that of most other animal cell types: between about -20 and -80 mV. In meroistic vitellogenic ovarian follicles, ionic current enters the trophocyte compartment and leaves the oocyte as already described in Section II,G. In vitellogenic panoistic ovarioles, extracellular current patterns have also been measured (Huebner, 1984; Overall and Jaffe, 1985; Verachtert and De Loof, 1986). This indicates that within a follicle the distribution of ion pump/ion channel activity is clearly asymmetrical during yolk deposition. In insects, there have not so far been any reports on spontaneous action potentials during vitellogenesis (continuous records of up to 12 hours). After Miyazaki er al. ( 1 972) discovered electrical excitability in tunicate eggs, action potentials have been found in oocytes and eggs of several classes of animals (review by Hagiwara and Jaffe, 1979). Xenopus oocytes sometimes show such membrane potential fluctuations, which seem to be caused by the spontaneous opening of the acetylcholine receptor channels present in the egg cell membrane but whose function is unknown. Application of acetylcholine to the Xenopus oocyte usually causes depolarization of the membrane, through interaction with a muscarinic receptor by which chloride channels are opened. When acetylcholine is applied to one of the poles, the chloride current is stronger at the animal pole (Kusano et al., 1982). Electrically excitable, Na+-selective channels can be induced in the plasma membrane of Xenopus oocyte when this is

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submitted, by voltage clamp, to prolonged positive potentials (Kado et al., 1979; Baud et a/., 1982; Baud, 1983; Baud and Kado, 1984). The electrical properties of mouse and human oocyte membranes are sensitive to acetylcholine only before fertilization and not afterward (Eusebi eta/., 1979, 1983). Jaffe and Cross ( 1984) speculate that acetylcholine could play a role in the communication between the follicle cells and the oocyte. The mouse oocyte can be stimulated to generate action potentials before ovulation when the oocyte diameter is 40-60 p m (Yoshida, 1983). Bland eta/. (1983) reported that the action potential in the mouse egg is Ca2+ dependent. Mature oocytes can also produce action potentials (function unknown) due to the presence of a voltage-sensitive calcium channel (Okamoto et a l . , 1977). In metaphase 1 oocytes of Rana pipiens, the Em oscillates continuously between -30 and +60 mV without the addition of a neurotransmitter. Two voltage-sensitive ion channels, a Na+ channel and a C1channel, are involved. Opening of the C1- channels causes the fall, while opening the Na+ channels causes the rise (Schlichter, 1983a). Schlichter (1983b) proposes that the action potentials load the oocyte with Na+ which may regulate the rate of maturation either directly or indirectly; during the maturation period the Na+ activity of the frog oocyte cytoplasm will increase 4-5 times. The C1- channels disappear 20 minutes after fertilization (Jaffe and Cross, 1984). Variations in the C1- permeability of amphibian oocyte membrane, which may be Ca2+ dependent, have also been associated with responses to acetylcholine, to adenosine, to fertilization, and to activation (Barish, 1983). Ca2 +-dependent action potentials of long duration (3- 10 minutes for the plateau phase) can be induced in the fertilized egg of the ctenophore, Mnemiopsis feidgi (Barish, 1984). 2. Maturation Oocytes are generally blocked in the prophase of the first meiotic division and have an (absolute?) requirement for a hormonal stimulus to release this block and to induce progression to the second meiotic metaphase as an unfertilized egg. This is called maturation. Maturation involves germinal vesicle breakdown, expulsion of the first polar body, and formation of metaphase I1 chromosomes. In echinoderms, it is I-methyl adenine which is the inductor of maturation. I Methyl adenine causes an increase in free intracellular Ca2+ in the starfish Marastherias glacialis but not in Asterias forbesi (Eisen and Reynolds, 1984). In vertebrates progesterone is the major hormone involved but other steroids, e.g., testosterone, are also active (Maller, 1983; Le Goascogne et al., 1985), as is insulin (Hirai rt a/., 1983; Lessmann and Marshall, 1984; Morrill eta/., 1984a). Increased progesterone synthesis is triggered by luteinizing hormone (LH). The addition of progesterone to the incubation medium of Rana pipiens follicular oocytes does not change the resting potential of the oocyte plasma membrane (about -60 mV) within 1 hour, although Ca2+ efflux already starts. By 10-15 minutes a fall in intracellular CAMP is observed, followed by a transient rise in

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intracellular cGMP 1-2 hours later. The pH of the cytoplasm increases slightly. [3H]Uridine incorporation increases at 3-5 hours. Plasma membrane potential reaches a value of about -25 mV about 6-10 hours after exposure to progesterone (1-2 hours before nuclear breakdown), when Ca2 efflux begins to decline (O’Connor et al., 1977; Weinstein et al., 1982; Morrill et al., 1984b). By the end of the first meiotic division (20-24 hours) the oocyte cytoplasm becomes essentially at isopotential with the medium. Depolarization coincides with the disappearance of Na+ ,K+ -ATPase activity, as shown by a decrease in ouabain-binding sites (Weinstein et al., 1982), and with a simultaneous decrease in both K + and CI- conductances of the oocyte membrane (Weinstein et al., 1982; Morrill et al., 1984b; Taglietti er al., 1984). Na+,K+-ATPase activity reappears following the second meiotic division (Weinstein et al., 1982). By means of the vibrating probe technique, Robinson (1979) found that progesterone treatment brings the transcellular electrical currents in Xenogus to a complete stop. Within 1 hour, luteinizing hormone alone causes a hyperpolarization to about -72 m V perhaps by increasing the K + conductance of the oolemma, or by increasing the oocyte-follicle cell ionic coupling, or by increasing the conductance of the follicle cell plasma membrane. Pituitary hormones may thus play a direct role in processes which accompany or follow maturation (Weinstein er al., 1983). Maturation is inhibited by cycloheximide, not through direct interference with protein synthesis but with Ca2 release (Morrill et al., 1984b). The role, if any, of the increase in pH by about 0.4 pH units and the reduction in ion conductances during amphibian oocyte maturation is still unclear (Lee and Steinhardt, 1981; Ciccirelli et al., 1983; Baud and Barish, 1984). The increase of about 0.05 pH units during maturation of starfish oocytes induced by 1-methyl adenine, does not seem to play an essential role in activation (Johnson and Epel, 1982). In protostomians, the chemical nature of the hormonal trigger(s) for maturation remains to be elucidated. For insects the steroid ecdysone secreted by the follicle cells has been suggested as a maturation inducing factor (De Loof et al., 1981). Progesterone-induced breakdown of the germinal vesicle is a clear example of the fact that at least this steroid hormone may act through a plasma membrane receptor and that CAMPis not exclusively used as a second messenger for peptide hormones (Baulieu, 1983; Blondeau and Baulieu, 1984; Baulieu et al., 1985). It also illustrates that a steroid hormone may be responsible for inducing changes in ionic concentrations between cytoplasm and nucleus, and in physicochemical interactions between various macromolecules, specific ions, and water (Cameron et al., 1983; Tluczek et al., 1984). 7-

+

3. Fertilization In mammals, at least, Na+ ,K+-ATPase activity and the resulting K + influx are important for the acrosome reaction (Mrsny and Meizel, 1981). The attachment of a single spermatozoon to the egg causes, in most but not all species so far studied, a very fast (within a few seconds) transient electrical excitation of the

315

THE ELECTRICAL DIMENSION OF CELLS

oolemma, the fertilization potential. In most cases this is a depolarization which is generally dependent upon an increased sodium permeability. In the marine worm Urechis caupo the increased Na influx takes place through special Na + channels which are not gated by voltage but by sperm, a sperm opening only a fraction of the available channels. The open Na+ channels seem to be localized in a “patch” near the fertilizing sperm. During the first 15 seconds a voltagegated Ca2 action potential acts as an enhancing mechanism (Gould-Somero, 1981). In the annelid Chaetopterus, the potential of the egg membrane shifts upon fertilization from -58 to +40 mV and the potential remains positive for 65 & 33 minutes. Two records of Saccoglossus, a hemichordate, showed a response of similar magnitude, from resting potential of -70 to about +20 mV within seconds. Four minutes after the rise began, the potential had returned to -50 mV (L. A. Jaffe, 1983). In the sea urchin interaction of the sperm with the egg (Em of the unfertilized egg is about -70 to -80 mV) results in a depolarizing response. An action potential is superimposed on the initial phase, which results in a rapid Na+dependent shift of the Em to a peak of +20 to +30 mV within 2-3 seconds after sperm attachment. The repolarization of the membrane, which is accompanied by an increase in K conductance and by intracellular changes in Ca2 , attains -20 to -30 mV by 3 minutes and the original Em of about -70 mV by 7-12 minutes (Hulser and Schatten, 1982; Lynn and Chambers, 1984). Preventing the depolarization normally associated with fertilization by means of a voltage clamp suppresses sperm entry in the sea urchin egg (L. A. Jaffe, 1976; Lynn and Chambers, 1984). In Paracentrotus, the plasma membrane of unfertilized eggs contains a preexisting Na+ , K + pump which is obligatorily stimulated at fertilization (Ciapa et a f . , 1984a). This stimulation is an absolute necessity for the Na+-H exchange. The alkalinization of eggs resulting from the acid efflux is a prerequisite for the enhancement of the Na+ ,K pump. Early events of fertilization, such as exocytosis and calcium release, which may be involved in the stimulation of the Na+ ,K pump, must necessarily be coupled to the rise in intracellular pH (Ciapa et al., 1984b). In the ascidian oocyte, one of the primary events at fertilization is also a rapid depolarization of the plasma membrane accompanied by a decrease in resistance and in voltage noise, The large specialized “fertilization channels” have a unitary conductance of about 40 pS, which is about the largest hitherto observed in biological membranes (Dale and de Felice, 1984). In Xenopus the depolarization which is the direct result of sperm penetration is of the order of 10 mV and this lasts for about 10-20 minutes. It seems to result from an increase in the chloride conductance leading to a net efflux of C1- (Grey et a / . , 1982; Webb, 1984; Webb and Nuccitelli, 1985a,b). In Rana esculenta and Rana ridibunda, the time of rise of the fast depolarization was shorter than in Xenopus and the total duration of the positive phase of the fertilization potential lasted 20-30 +

+

+

+

+

+

+

3 16

ARNOLD DE LOOF

minutes (Charbonneau et al., 1983). In Rana pipiens, the shift to the positive side lasts 10-20 minutes: in this species increased C1- and K + conductance is induced by fertilization, while a voltage-sensitive Na conductance, present in unfertilized eggs, disappears after fertilization (Jaffe and Schlichter, 1985; see also Webb and Nuccitelli, 1985b). In the urodeles Pleurodeles waltli and Ambystoma mexicanum, which both exhibit physiological polyspermy, Em in most eggs did not change in any consistent pattern during 45 minutes after fertilization (Charbonneau et al., 1983). In Xenopus, a transient acidification of the order of 0.03 2 0.02 pH units starts 2 to 6 minutes after fertilization and lasts for about 1 1 minutes. An association with the rise in Ca2 activity, believed to occur at the time of the cortical reaction, has been suggested. Later there is a permanent intracellular pH increase of the order of 0.3 pH units. So far there has been no evidence for a Na -H and CI --HCO, exchange in this species (De Laat et a / . , 1974; Nuccitelli et al., 1982; Webb and Nuccitelli, 1982). The intracellular pH increase requires extracellular Na+ , but the way in which this ion is used is still unclear (Epel et al., 1974; Chambers, 1975, 1976; See Epel, 1974 for review). Miyazaki and Igusa (1981a) reported that golden hamster eggs showed recurring, transient hyperpolarizations in response to fertilization (by several spermatozoa?). The first three to four hyperpolarizations, which occur at intervals of 40-50 seconds, raise the Em from a resting potential of about -30 to -75 to -80 mV. The resting potential gradually rises to -40 mV. The hyperpolarizations are caused by an increase in the membrane permeability to K + ions exclusively, which is based on a Ca2+-activated K+-current (Miyazaki and Igusa, 1981b, 1982; Georgiou et al., 1983). The Em of unfertilized mouse eggs is of the order of -4 I 2 4 mV. During the 60 minutes following insemination there are no dramatic changes in potential. Only small oscillations of 4 1 mV, accompanied by drops in egg membrane resistance from 96 34 to 44 +- 15 starting 7 ? 5 minutes after insemination and lasting about 1 minute, have been observed (Jaffe et al., 1983). According to Georgiou et al. (1983), some of the published values of Em and resistance of mammalian eggs may be underestimates of the real values. Prevention of the entry of more than one sperm (fast block to polyspermy) or at least decreased receptivity of the egg to sperm is one of the major functions which has been attributed to the fertilization potential in many species (Jaffe, 1976; Hagiwara and Jaffe, 1979; Nuccitelli and Grey, 1984). Eggs of fish (Nuccitelli, 1980), urodeles (Charbonneau eral., 1983), ascidians (Dale e t a / ., 1983), hamster (Miyazaki and Igusa, 1981a), mouse, and perhaps also rabbit (Jaffe et al., 1983; Jaffe and Cross, 1984) do not seem to have a (pronounced) electrical polyspermy block although some authors do not rule out the possibility that such a system, perhaps in an attenuated form, might be involved. Some data indicate that depolarization alone is not always a sufficient means for preventing polyspermy . +

+

+

*

+

*

a,

317

THE ELECTRICAL DIMENSION OF CELLS

4. Activation The egg awakens-is activated-at fertilization. Although the entire protocol of biochemical events involved is not yet fully elucidated, it is clear that Ca2+ is a key factor. Eggs of the medaka fish (Gilkey et al., 1978), sea urchins, and probably most deuterostomians seem to be activated by a Ca2 explosion which is propagated at about 10 Fm/second in a wave sustained by the Ca2+-stimulated release of Ca’ from internal sources, probably from specialized regions of the egg’s endoplasmic reticulum, analogous to those of muscle cells (Gilkey et al., 1978). Gardiner and Grey (1983) discovered in ripe Xenopus eggs subsurface cisternae which may be a source of activation calcium. They are 2-3 times more concentrated in the cortex of the animal hemisphere than in the cortex of the vegetal one. This might be related to the observed greater wave speed and excitability of the animal hemisphere. It may not be excluded that the tiny ‘‘agranules” around the large cortical vesicles of ripe medaka eggs, reported by Yamamoto (1962), are another storage site of Ca2 . L. F. Jaffe (1983) points to the similarity between exocytosis which is caused by fertilization and the general pattern of exocytosis (e.g., in the presynapses) induced in natural conditions in other systems: both are shortly preceded by a large increase in free Ca2 (Rubin, 1982). Nuclei transplanted into unactivated amphibian eggs condense into metaphase chromosomes whereas those transplanted into activated eggs decondense and enlarge. Ca2+ concentration is the essential factor involved (Lokha and Masui , 1984). Protostomian eggs are also primarily activated by Ca2 which, unlike the situation in deuterostomian eggs, enters the cytosol from the medium and does so in response to depolarization of the oolemma (L. F. Jaffe, 1983). All over the surface the calcium pulses are synchronous (not propagated in a wave) and prolonged (order of magnitude 10 minutes or more). The exocytosis process in protostomian eggs is far slower than that found in those of deuterostomians. If Chlamydomonas gametic cells of one mating type are mated with gametes of the opposite mating type, a rapid increase in Ca2+ efflux, which lasts about 6 minutes, is observed (Bloodgood and Levin, 1983). We may draw the conclusion that the ionic/electrical phenomena which are inherent both to fertilization and activation are rather similar to those which occur in other excitable cells. However, the functions of the phenomena in the different cell types may be quite different. +

+

+

+

+

F. ELECTRICAL PHENOMENA AND

IN VERTEBRATE REGENERATIVE GROWTH WOUNDHEALING

Until now, we have been discussing electrical phenomena mainly at the cellular level. There is at least one instance in which they have also been examined at the tissue level, viz. regeneration and wound healing. These topics have recently been reviewed in an authorative way by Borgens

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(1982, 1983), Becker (1982, 1984), and Person (1983). When the plasma membrane of a cell, which is internally negative with respect to its extracellular environment, is injured, there will be an instantaneous and steady influx of positive charge through the hole which will be maintained by the ionic pumps in the plasma membrane until the lesion has healed. Since the concentration of Ca2+ and Na+ is often higher outside the cell, these ions will quickly enter the lesion. This sudden rise may locally affect some biochemical processes and structures. Carafoli and Crompton (1976) reported that small increases in internal Na+ cause a release of internally stored Ca2 from the mitochondria, and as the endoplasmic reticulum i; also a storage site for Ca2+, perhaps also from this. In this respect Schlaepfer (1974) suggested that the rise of Ca2+ in the cut end of axons may cause the disassembly of contractile elements and microtubules there. Charge influx may induce or enhance self-electrophoresis which may result in the local accumulation of charged macromolecules or cell organelles, e.g., mitochondria, rapidly accumulate at the cut end of nerves but not as a result of multiplication (Borgens et al., 1980). The same principle can be applied to animal organisms as a whole. Most animals so far studied maintain large (about 30-80 mV) transcutaneous potential differences so that the skin is usually internally positive with respect to the outside (Ussing, 1964; Kirschner, 1973). In the cavy and in man (Barker et af., 1982), the transcutaneous potential difference varies substantially (30 to 80 mV) over the different parts of the body (hairy versus naked regions, foot versus thigh, etc.). For data about man, see the drawing in Borgens (1982). If a transcutaneous wound is made charge will flow out of the wound. Such wound currents were already observed by Dubois-Reymond who, as early as 1843, determined that small wounds in human skin were externally positive with respect to the adjacent uninjured areas. Only in the last two decades has the study of electrical phenomena as related to regeneration attracted renewed interest and lead to practical clinical applications. With the vibrating probe technique it has been shown that the densest current (of the order of 50-100 kA/cm2) leaves the core tissues of regenerating amputated limbs of newts. Small currents (of the order of 1-3 pA/cm2) uniformly spread over the entire limb and enter the skin (Borgens et al., 1977a). Frogs lose the ability to regenerate limbs at that moment during larval development when subdermal lymph sacs appear. The low electrical resistance of these sacs is most probably the cause of the skin-driven current in adult frogs being shunted around and not through the central core of the stump (Schwan, 1963; Borgens et al., 1979a). A variety of means, e.g., amiloride treatment, keeping the animals in a Na+-deficient medium, etc., have been used to inhibit or reverse the current flow in regenerating amphibians. The usual result is inhibition, retardation, or abnormal limb regeneration. In a Na+-free medium, there was no regeneration for weeks, but it at once became very rapid compared to that found in the +

THE ELECTRICAL DIMENSION OF CELLS

319

controls. In these animals, currents were not Na+ dependent and amiloride insensitive, in contrast to the controls in a Na+-rich medium (Borgens et al., 1977a, 1979b; Borgens, 1982). Lasalle (1980) was unable to detect proximodistal potential differences in the skin surface of amputated stumps of axolotls. Since this species regenerates exquisitely, he questioned the relevance of wound currents in regenerative growth. Vanable er al. (1983) found, by means of the vibrating probe technique, that axolotl limb stumps do produce currents that are similar to those now already measured in a variety of newts and salamanders from different habitats: in most species the current ranges from about 10-100 pA/cm2 several hours after amputation and declines with time (Borgens et al., 1984). Insulin has a stimulatory effect on macromolecular events occurring in cultured regeneration blastemata (Vethamany-Globus et al., 1984) perhaps by its effects on Na+ ,K+-ATPase or ion transport (Section 111,G). Wound currents leaving the accidentally cut end of childrens’ fingers, amputated at the level of the terminal phalange, have been measured by an adapted vibrating probe (Illingworth and Barker, 1980). If in these children no skin flap is sewn over the cut end and if no attempt is made to reattach the severed finger or toe phalange, but if the wound is simply cleansed and covered with a nonadherent dressing, regeneration will occur in patients (up to the age of puberty). In 11-12 weeks this regeneration is complete as it includes regrowth of the missing part, fingernail, and dermatoglyphic pattern (Douglas, 1972; Illingworth, 1974). This treatment is now fairly common practice in Australian hospitals. When in salamanders or children a skin flap is sewn over the wound (Polezhaez and Favorine, 1935; Illingworth, 1974; Mesher, 1976), no regeneration will occur. Darwin (1861) reported the ability of children to regenerate fingers. In mammals the skin battery seems to be similar to that of amphibians. Sewing a skin flap over the wound disturbs the normal wound currents but in humans it has not yet been proven that this is the (only?) cause of nonregeneration. When the density of current traversing the core tissues in the stump of a frog limb was artificially enhanced by electric current delivered from a battery, limb regeneration was initiated on condition that the cathode was located at the stump’s end (Smith, 1974; Borgens et al., 1977b). If the anode as placed there, which means that current was now driven in an opposite direction as compared to the natural wound current in salamanders, gross degeneration of the limb was observed. Since metal electrodes were used in the latter experiments, degeneration due to metal ion contamination (a problem in all experiments where current is driven in tissues through implanted metal electrodes instead of wick electrodes) cannot be ruled out (Borgens et al., 1977b). It has been observed that wound currents leaving the stump even increase after denervation of the latter (Borgens et al., 1977a). Hence, the nerves are not the site of wound current generation. This is in agreement with other experiments which strongly suggest that it is the skin which is essential for current generation.

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This does not mean that nerves are not necessary for regeneration. Upon applying an imposed current, a gross increase in the amount of nervous tissue-up to 20% of new tissue in adult frogs-is seen (Borgens et al., 1977b). This hyperinnervation may be essential for speeding up the development of other tissues. We refer to Section III,D in which nuclear activity in excitable cells was discussed. When holes are punched through the ears of a rabbit, regeneration will occur. The bioelectric activity in this regeneration process has been described by Chang and Snellen (1982). The ability to regenerate is also a typical feature of bone tissue. It is often overlooked that bone is a complex, dynamic, living tissue. Electrical signals may be essential to its growth, remodeling, and repair (review Weiss er al., 1980; Borgens, 1982). 1. Steady potentials, the source of which is unknown, have been measured in growing bone. The epiphyseal plate and the cortical bone of the diaphysis are positive with respect to the areas of the long bone just below the epiphyseal plate. If the bone is not viable, these potentials disappear (Friedenberg and Brighton, 1966). 2. Steady injury potentials, the source of which is unknown and which are negative in the area of the lesion with respect to undamaged portions, have been described. They do not occur in dead bone (Friedenberg and Brighton, 1966; Weiss et al., 1980). 3. The collagen component of dry bone-but not its crystalline hydroxyapatite structure-is responsible for the fact that upon mechanical stress short lived potential differences (of the order of millivolts for a few tenths of a second) are induced. Thus bone, dead or living, can act as a piezoelectric transducer. The unloaded portion of a long bone is electropositive with respect to the areas of compressive stress (Fukuda and Yasuda, 1957; Cochran et al., 1968; Bassett, 1971; Yasuda, 1974; Williams and Breger, 1975; Steinberg et al., 1977; Korostoff, 1977). 4.During the intermittent stress and relaxation of normal movement, streaming potentials are generated as the result of movement of body fluids through the small channels in bone tissue. The channel walls preferentially bind ions of the same sign as fluid is forced through the bone from the compression side to the tension side (Borgens, 1982). It is becoming more and more apparent that the signals that mediate repair, remodeling, and even possibly the growth of bone may be electrical in nature (Borgens, 1982). The healing of difficult bone fractures in man can also be speeded up by driving a weak dc current through implanted electrodes, the cathode(s) being in the lesion (reviews of Brighton, 1981; Person, 1983). External electromagnetic fields are presently used to enhance bone fracture healing, but the results so far obtained are under debate (Section IV).

THE ELECTRICAL DIMENS!ON OF CELLS

32 1

Another remarkable example of regeneration is that of the severed spinal cord of lamprey larvae. Immediately after transection current densities of about 0.50.8 mA/cm2 (compared to about 20 p.A/cm* in core tissues of limb stumps in frogs) enter the cut face of the spinal cord as measured by the vibrating probe. It is assumed that the incurrent actually enters the cut ends of the (inside negative) axons themselves. The current declines to a stable level of about 4 pA/cm2 after 2 days posttransection. Eventually the transected spinal cord of the lamprey larva can regenerate across the complete transection, this coinciding with a return of behavioral function (Borgens et al., 1977a,b). The mechanisms of action by which natural and applied electrical fields influence regenerative growth are only partially understood. These fields may have effects on the gross morphology, the character of the cytoskeleton, the rate (Borgens et al., 1977; Borgens, 1982) and direction (Hinkle et al., 1981) of growth, elongation, nuclear activity, self electrophoresis in the cytoplasm, and lateral electrophoresis in the plasma membrane (Jaffe, 1977) of the cells in the vicinity of the wound. The observations in this limited field indicate that electrical phenomena probably are important also for pattern formation.

G . ELECTROPHYSIOLOGY, ENDOCRINOLOGY, AND OSMOREGULATION In addition to communication between adjacent cells by means of gap junctions (Loewenstein, 1981), the different cells and tissues which make up an organism communicate by two major pathways: the nervous system, with its neurotransmitters for rapid communication, and the endocrine system, with its hormones for slower but longer lasting effects. As more and more hormones, which also may act as a neurotransmitter or neuromodulator, are found and as it becomes gradually clearer that many steroids also exert effects at the plasma membrane level, the strict distinction between nervous and endocrine systems gradually becomes more attenuated. There still are two pronounced dualities in endocrinology. The first one is that peptide and steroid hormones are still often thought to act in completely different ways. Because of their molecular mass, it was not thought possible that peptide hormones could enter the cell and would be obliged to make use of secondary messengers: for example, CAMP and Ca2+. Recently polyphosphoinositides have been recognized as new secondary messengers in signal transmission for a wide variety of neurotransmitters, hormones, and growth factors (Berridge and Irvine, 1984). It has also been suggested that Ca2+ might not be a secondary but a tertiary messenger, as its concentration increase seems to be caused by inositol triphosphate (Marx, 1984). Steroid hormones, most of them being lipid soluble, are often thought to diffuse freely into all cells, to be retained here only by “target cells,” and to make no use of secondary messengers. Many peptide hormones, after binding to their plasma membrane receptor, are internalized by endocytosis as documented

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for ACTH, EGF, glucagon, insulin, human chorionic gonadotropin, LH, LHRH, prolactin, and TSH (review Szego and Pietras, 1984). It may be possible that peptide hormones also exert functions inside the cell, perhaps after being degraded to smaller peptides. Specific high-affinity binding sites for insulin have already been found in the nuclear envelope (Goldfine et al., 1982). The second duality is that steroid hormones are quite often supposed to exert their effects only at the level of transcription after binding to a nuclear receptor. Nevertheless, it is well known that some steroids, such as aldosterone (Hamlyn and Duffy, 1978; Moura and Worcel, 1982) and cortisone (Massa et al.. 1975), affect Na+ ,K -ATPase activity or/and plasma membrane permeability to ions. Within the limited space available here, I will only briefly summarize data on the effect of the peptide hormone insulin on ion transport (review Moore, 1983) and nongenomic effects of steroids (De Loof el al., 1982; De Loof and Geysen, 1983; review Duval et al., 1983). It is known that insulin affects transport of glucose, amino acids, and ions across the plasma membrane. Less well known is the fact that insulin stimulates Na+ ,K -ATPase activity, not by increasing the number of pump sites but by allosteric effects. These effects are caused by the binding of insulin to a membrane receptor and not to Na+ ,K+-ATPase itself. Moore (1983) estimates that one insulin receptor controls 15-80 Naf , K + pumps. Insulin causes hyperpolarization in frog skeletal muscle, mammalian skeletal and cardiac muscle, some epithelia, and probably also in fat cells. This hyperpolarization most likely affects the electrophysiological properties of action potential generation in skeletal and cardiac muscle. The hormone, probably by activating Na :H exchange, increases the cytoplasmic pH in frog skeletal muscle. This does not occur in rat heart. It stimulates Na -dependent amino acid uptake in a wide variety of biological tissues. Insulin also changes Ca2 fluxes by two different mechanisms: inhibition of the high affinity Ca2 -ATPase and also through the Na+ :Ca2+ exchange system. For some processes, the action of insulin is intimately linked to the Na+ electrochemical gradient across the plasma membrane. The intracellular rise in pH which is elicited by insulin is the probable trigger for several functions of this hormone, for example, glycolysis, DNA and/or protein synthesis, protein phosphorylation and dephosphorylation (Moore, 1983). Insulin also affects renal reabsorption of Na+ and also plays an important role in K + homeostasis (Knochel, 1977; Cox er ul., 1978). It stimulates nucleoside triphosphatase activity, the enzyme that regulates mRNA efflux, in the nuclear envelope (Purello et al., 1982). An interesting finding was that anti-insulin-receptor antiserum and plant lectins mimic the direct effects of insulin on nuclear envelope phosphorylation (Purello et al., 1983). Insulin is evidently not the major hormone involved in osmoregulation. Several hormones, steroids (e.g., aldosterone, cortisone), peptides (e.g., vasopressin, Handler and Orloff, 1973; Helman et al., 1983), and prostaglandins (Hall et al., 1976; Els and Helman, 1981) are involved in transport of water and solutes in the +

+

+

+

+

+

+

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323

different salt-transporting epithelia of the body, such as the kidneys, salivary glands, Malpighian tubules of insects, etc. Water passively follows the movement of ions, although, at this moment, the question of whether in salt/watertransporting epithelia water passes through the cells themselves or through a paracellular (transjunctional) pathway is as yet not definitely answered (Hill, 1980; Durbin, 1981; Spring and Ericson, 1982). The vibrating probe technique has been successfully used for the definite identification of the mitochondria-rich “chloride cells” in gills of a Tilapia species as being the only site of electrogenic ion transport in this heterogeneous epithelium (Scheffey er al., 1983). Osmoregulation is often considered a system for creating optimal conditions for the other physiological processes: it keeps the ionic composition of the body fluids more or less constant. In fact it does much more than that. It makes it possible for all cells of the body to realize their electric dimension, as this depends completely upon ions. The cells/tissues engaged in osmoregulation are specialists in transporting large amounts of ions through themselves for the benefit of the other cells of the organism. The regulation of protein synthesis in target tissues is a major, but not the unique, action of steroid hormones. The two-step hypothesis of Jensen et al. (1968) on the transport mechanism of the steroid to the nucleus and its intranuclear mode of action is not sufficient to account for all known effects of steroids. It is becoming increasingly criticized (Martin and Sheridan, 1982; Rousseau, 1984; Szego and Pietras, 1984; De Loof, 1985~);as it offers no explanation of the tissue-specific activity or developmental stage specificity of steroids, neither can it account for the fact that nonsteroid molecules can sometimes mimic the effect of a steroid (e.g., Baulieu and Schorderet-Slatkine, 1983). Furthermore, the proposal that cytoplasmic estrogen receptor is translocated to the nucleus after binding the steroid does not seem to be correct (King and Greene, 1984; Welshons and Lieberman, 1984; Schrader, 1984). There are other conflicting data (De Loof et al., 1982). Furthermore, it does not take into account the fact that steroids do not activate single genes but sets of genes and that these sets are brought to expression only when the balance of all control factors (different hormones, nutrients, physical factors) is right. The finding that progesterone brings the extracellular currents around the frog oocyte to a stop (Robinson, 1979) and induces maturation by a plasma membrane-mediated effect (Baulieu et a l . , 1978; Baulieu, 1983) strongly suggests that, contrary to what had long been assumed, the first effects of steroids may occur at the plasma membrane. A plasma membrane receptor for progesterone has recently been characterized in Xenopus oocytes by Blondeau and Baulieu (1984). However, Tso et al. (1982) observed that microinjection of progesterone in paraffin oil also induces maturation in Xenopus. At present, the list of the interactions of steroids with membrane structures and functions is already a long one (review Duval et al., 1983; Szego

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and Pietras, 1984). A whole range of receptor-mediated effects of steroids on membrane composition and properties, effects on cell excitability, interaction with the cyclic nucleotide system (Maller, 1983), ion, glucose, and nucleoside transport, lysosomal and mitochondria1 functioning, and a wide variety of enzyme activities (Tomkins and Maxwell, 1963; Duval et d., 1983), have been reported. Cortisone very rapidly counteracts the effect of MSH on melanosome dispersal (explained by self-electrophoresis, Kinosita, 1963). In the electric fish, Srernopygus, 5a-dihydrotestosterone lowers the discharge frequencies of electroreceptors in accord with that of their associated electric organ (Meyer and Zak, 1982). Androgens affect the electric organ itself in Brienomyrus brachyistius (long biphasic) while estradiol has only a weak effect (Bass and Hopkins, 1983). They also provoke changes of ultrastructure in rat ventral prostate and seminal vesicle (Arnold ef al., 1983). A list of effects of some steroids on concentrations of cellular Na + , K , and CaZ , on the activity of Na+ ,K ATPase and Ca2 -ATPase, on membrane potential and other electric parameters, is given by Szego and Pietras (1984, pp. 90-93). The natural maturation hormone of echinoderms, 1 methyl adenine, induces changes in the electrical properties of starfish oocytes (Moody and Lansman, 1983). Juvenile hormone of insects, a sesquiterpenoid, causes the potential difference existing between the tropharium and oocytes in the ovarian follicle of the insect Rhodnius to rise from 3 to 9-10 mV (Telfer ef al., 1981). The steroid molting hormone of arthropods, 20-hydroxyecdysone, induces, within 15 minutes, an increase in volume of a cultured lepidopteran cell line (Englisch et al., 1984). Tissue and developmental stage specificity is not based on a difference in the base sequences of the genes in the different tissues (Levine et al., 1981; De Loof, 1985b); neither are there tissue-specific receptors for steroids. This means that epigenetic factors are responsible. In our opinion, some steroid hormones activate different gene sets in different tissues, mainly because all these tissues have different plasma membrane properties and, hence, most probably have a different ionic environment (ionic and potential gradients, ionic currents, secondary chemical gradients) in their nuclei, which probably results in differences in higher order structure of chromatin in the various cell types and tissues (De Loof, 1985b,c). The classical approach in endocrinological research, which is to study the effect of one hormone at a time and, even then, to focus on only one of its effects does not sufficiently take into account the fact that so many processes are coupled in the cell. For example, changes in concentration of one ionic species induced by a hormone will probably have effects on other ions (H for example) and perhaps on Em and transcellular currents. These in turn may induce secondary effects. Recent data on the mode of action of polypeptide growth factors like epidermal growth factor (EGF) and platelet-derived growth factor (PDGF) illustrate this. These mitogens stimulate proliferation of fibroblasts and epithelial +

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cells. Among other effects, they induce a Na influx due to the activation of an electroneutral Na -H + antiport. One of the predicted consequences of such an antiport system, namely, unchanged plasma membrane potential and cytoplasmic alkalinization, has been experimentally demonstrated (Glaser et al., 1985). We have suggested that the concept that many cells are miniature electrophoresis chambers and that the targets of steroid hormones are often salt/watertransporting epithelia may yield a more coherent understanding of hormone action in general. By its very nature, such a system could handle a balance of factors, very much in the same way as an interneuron can simultaneously handle all the information coming from the numerous synapses: “all” the interneuron does is to continuously make the sum of all ionic fluxes passing through its plasma membrane. In fact, this is what any active cell continuously does. The difference is that in neurons the ion fluxes are roughly restricted to the plasma membrane, while in other nonexcitable cells, as the result of the asymmetrical distribution of the ion pumpslchannels, the ion fluxes may also traverse the cytoplasm (or a part of it) (De Loof, 1985b,c). +

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H. ELECTRICAL PHENOMENA I N BACTERIA, YEAST, AND MITOCHONDRIA In Section II,H it was briefly mentioned that the membrane potential of bacteria is usually high. The chemiosmotic theory predicts that the pH of the medium in which the bacteria grow will influence the membrane potential. Because bacteria, yeasts, and mitochondria are usually very small, direct measurements with microelectrodes are very difficult or, in most cases, even impossible to perform and therefore indirect methods, which also have their limitations, are often used. By measuring the distribution of thallous ions, Bakker (1978) calculated that respiring cells of E . coli develop a membrane potential of -160 mV with Dlactate and - 180 mV with glucose as respective substrates. For Streptococcus faecalis he reported values of up to - 180 mV in glycolysing cells. In giant cells of E . coli, induced by growth in the presence of 6-amidinopenicillanic acid, membrane potentials have been directly measured with intracellular microelectrodes and indirectly from the steady-state distribution of [’Hltetraphenylphosphonium. Both techniques yielded similar results: Em approximates were -85 mV at pH 5.0 and -142 mV at pH 8.0, with an average slope of -22 mV/pH unit over the pH range 5.0-7.0 (Felle et a l . , 1980). By use of an electrode sensitive to tetraphenylphosphonium, Muratsugu et al. ( 1982) evaluated the membrane potential of E . coli at - 176 I+_ 5 mV. Huttunen and h e r m a n (1980) used safranine, a nontoxic positively charged dye, as a probe for membrane potential determination in membrane vesicles of two strains of E . coli. With formate as a substrate, values of the order of -160 to -170 mV were found: these are in the same range as those obtained with whole bacteria by

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Bakker (1978). Cyanide dyes have also been used, but some of these have been reported to be toxic to both bacteria and mitochondria and the response of cyanines is nonlinear above 80-100 mV (Laris and Persadingh, 1974; Renthal and Lanyi, 1976). Singh and Bragg (1979) reported membrane potentials in the range of 100-150 mV in membrane vesicles of E. coli. Zaritsky et a f . (I98 I), using [3H]triphenylmethylphosphonium,estimated the membrane potential in Bacillus subtifis to be 120 mV, inside negative. They warn that if no sufficient corrections are made, the values obtained on membrane vesicles may give an overestimation of the real values. In Staphylococcus aureus A$ is -85 to -90 mV and ApH is about -100 mV at pH 5.0. At pH 7.5, A$ is about - 130 mV and ApH is 0 (Mates et al., 1982). Finean et a f . (1979) mention AjiH values of about 180-240 mV (negative inside) across the membranes of several respiring bacteria. A$ is usually dominant but, especially in species which flourish in substantially acidic or alkaline environments, ApH can be appreciable. Lower values are usually reported but these are nevertheless higher than those normally found in eukaryotic cells. To our knowledge, there are, as yet, no data available on the symmetry of distribution of ion pumps or channels in the cell wall of bacteria. It is more or less intuitively assumed that this distribution is symmetrical. However, in many-if not all-eukaryotic cells, the distribution is asymmetrical. It should not be ruled out a priori that this could also be the case in prokaryotes. If there would be a potential difference as small as I mV from one end to the other of a I-pm-long rod-shaped bacterium (e.g., - 139 to - 140 mV) this would already correspond to a voltage gradient of 10 V/cm. A 10 mV difference between both ends, e.g., -135 to -145 mV, corresponds to a voltage gradient of 100 V/cm. If bacteria would be able to maintain such voltage gradients, they could also be miniature electrophoresis chambers, perhaps even of the high-voltage type (De Loof et al., 1982). The majority of transport processes of solutes across the membranes of bacteria such as E. coli seem to utilize energy obtained directly from AGH+. Ions are handled in two different ways. Electroneutral H symports (for anions such as glutamate, glucose 6-phosphate, gluconate, glucuronate, and phosphate) or H antiports (for cations) are driven by energy withdrawn from ApH while electrogenic uniports are driven in appropriate directions by A$. In E. coli, the internal Na concentration is kept low by a Na -H antiport. K and several basic amino acids, e.g., lysine, are moved by electrogenic uniport, while Martirosov and Trchounian (1982) and Trchounian (1984) also propose a K -H antiport system as another mechanism for the regulation of the intracellular K concentration in E. coli. Electrogenic symport of the permeant and a proton, with energy withdrawn from both ApH and A$, is the preferred mechanism for molecules that are either un-ionized or electrically neutral, such as, e.g., pgalactoside, galactose, arabinose, leucine, cystine, proline, serine, and glycine (Kaback, 1977, 1983; Harold, 1977, 1978; Finean et al., 1979; Skulachev and +

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Hinckle, 1981).Jacob and Monod (1961a,b), whose hypothesis strongly influenced thinking on the mode of action of steroid hormones, reported that when a sugar-p-galactose or even one of its structural analogs-was added to E . coli growing in a glucose-containing medium, the entire set of genes of the lac operon was switched on. After being taken up in the cytoplasm, the sugar is thought to bind to the repressor and inactivate it by allosteric interaction so that it is released from the operator site. However, because galactose uptake depends upon APH , the ionic environment in the bacterial cytoplasm and perhaps also transcellular ion fluxes will also change as the sugar enters. This may perhaps also be a (major?) factor of importance for release of the repressor. In this respect, recent data reported by Hudson and Schultz (1984)for Necturus small intestine are interesting. Addition of galactose to the mucosal bathing solution causes, within 2 minutes, an increase of the intracellular Na activity from 12 to 20 mmol/liter. This soon declines again but the final steady state in the presence of galactose was characterized by a 3 - to 4-fold increase in the rate of the transcellular Nu+ transport (from 13 to 50 pA/cm2) in the absence of a significant increase of the intracellular Na activity. This clearly illustrates that ionic concentrations alone, without taking currents into account, may give an incomplete view of the “ionic envir0nment”within the cytoplasm. Pena et al. (1984)used 3,3’-dipropylthiocarbocyanideas a membrane potential probe in the yeast Saccharomyces cerevisiae by following both its fluorescence changes and its uptake by the cells under different conditions. Noncorrected values ranged between -152 and -223 mV, but the corrected values (derived after consideration of various factors, e.g., binding of the dye to internal compartments) are substantially lower and closer to the values reported by De La Pena et al. (1982).These authors found that S . cerevisiae cells are able to maintain a AFH whose values vary from - 150 mV at pH 4.5to -90 mV at pH 7.0.As shown for bacterial cells (Zilberstein et a l . , 1979;Michel and Oesterholt, 1980)and bacterial membrane vesicles (Ramos et a l . , 1976),ApH and A+ vary independently as functions of external pH. ApH decreases from 1.8 (- 105 mV) to 0.2(- IOmV) while A+ increases from -45 to -80 mV as the external pH increases from pH 4.5 to 7.0.Yeast cells exchange K + for H+ (Conway et a l . , 1950;De La Pena et a l . , 1982) and accumulate K + against a concentration gradient (Rothstein, 1976)by making use of the electrical potential created by a proton ATPase (Pena, 1975). The data on the membrane potential of mitochondria were cited in Section 11. The mechanism by which mitochondria transfer the chemical energy of a substrate into ATP is electrical in nature. Azzi (1984)clearly described the way in which mitochondria use 0, for cell life as follows. This complex mechanism to generate electricity, or better “proticity,” is essentially similar to that which is found in a hydrogen-burning fuel cell. Such a cell consists of two elements: the the other, one in contact with 0, becomes positively charged and produces 02-; +

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in contact with H,, becomes negatively charged and produces H . The connection between the 2 elements with a H+ conductor (H,O) results in a source of “proticity.” Mitochondria function essentially in this second mode. They have an electron transport chain which allows separation of OH- in one compartment from H in another compartment. In this way proticity is produced and this form of electrical energy can be utilized for ATP synthesis. Cytochrome c oxidase represents an element of the respiratory chain capable of electric conduction through its metal centers and separation of H+ from OH- through a proton pump. The complexicity of this enzyme has been reviewed by Azzi (1984). The chemiosmotic concept is not restricted to the processes we mentioned in bacteria and mitochondria. It also accounts for a wide range of bioenergetic phenomena such as bacterial mobility, the uptake and storage of neurotransmitters in presynaptic nerve termini, active transport, transhydrogenation of NADP by NADH, nitrogen fixation, transfer of genetic information, sensitivity of resistance to certain antibiotics, cellulose synthesis, and processing of secreted proteins (for references, review Kaback, 1983). In vivo studies have shown that active nutrient uptake in plasma membranes of fungi and higher plants also occurs by proton cotransport (Serrano, 1977, 1983; Eddy, 1978; Poole, 1978) and that the electrical membrane potentials (Poole, 1978; Spanswick, 1981) observed in fungi (Serrano, 1980) and plants (Marrk, 1979) can be explained by an electrogenic proton-pumping ATPase located in the plasma membrane. The characteristics of the plasma membrane ATPases of fungi were reviewed by Goffeau and Slayman (1981) and those of plants by Hodges ( 1976). +

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IV. Effects of Imposed Electrical Fields on Cellular Activities In recent years increasing interest has been directed to the effects of imposed dc current and pulsed electromagnetic fields (PEMFs) on cellular activity; this interest was mostly shown in the regeneration of limbs, nerve, and bone tissue. Smith (1974) and Borgens ef al. (1977a) observed that when fields are imposed on frog forelimb stumps by implanted battery-electrode assemblies-with the negative pole to the distal end-the stumps produce a growth consisting of an extension of the radio-ulna, some muscle, islands of cartilage, and a large quantity of nerve. When no current is delivered to the implanted electrode or when the field is imposed in the reversed direction no such growth occurs. These effects are apparently not due to substances produced by the metal electrodes, as they can also be obtained when current is delivered to the distal end of the stump by a “liquid wire” salt bridge (Borgens et af., 1977a). In axolotls, which easily regenerate amputated limbs, it is likely, but not yet completely proven, that a sufficient augmentation of the natural stump current by an imposed field through

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a “liquid wire” electrode also stimulates a more rapid regeneration than normal (Vanable et al., 1983). The technique of driving a weak dc current via implanted electrodes through bone fractures which do not unite spontaneously is used in specialized orthopedic clinics. Regeneration requires de novo protein synthesis, cell multiplication, and directed growth. Whether and how all these activities are causally influenced by an imposed field is not as yet clear. Cheng et al. (1982) found that direct currents (50-500 bA) driven through skin flaps of Wistar rats incubated in vitro caused an up to 3-fold higher incorporation of labeled isoleucine or glycine as compared to controls. At 500 FA ATP concentrations were up to 4 times higher. According to the authors, this suggests that the biochemical effects of electric current can be adequately explained by a modification in proton movements, this on the basis of the chemiosmotic theory. Rubinacci and De Loecker (personal communication, 1985) showed that there may be a cause-effect relationship between endogenous currents and protein synthesis. They fixed skin flaps removed from the back of rats between the two halves of a Ussing-type chamber so that the skin formed a barrier between the two buffer compartments of the chamber. The two fluid compartments were connected by a salt bridge. Since the natural skin battery is not disturbed by this treatment, ions are pumped from one compartment to the other. Without the short circuit through the salt bridge, a transcutaneous potential difference will build up to an equilibrium situation. The salt bridge short circuits both compartments and ionic fluxes can continue to flow as long as the tissue is viable and enough energy is present. When the skin was short circuited by a low resistance salt bridge, after an initial increase, de novo protein synthesis, as measured by 14C-labeledamino acid incorporation, became constantly reduced by up to 40%. When an imposed current was driven through the skin in a direction opposite to the one produced by the skin itself, de novo protein synthesis was increased by up to 20%. Bloom et al. (1982) removed the appropriate segment of the neural tube of chick embryos at day 2 in ovo, thus prior to the outgrowth of nerves. The muscles were then chronically stimulated via surgically implanted electrodes. They found that growth was enhanced significantly beyond the level characteristic of unstimulated aneurogenic muscles. These results implicate activity per se as an important factor necessary for the proper growth and survival of at least brachial muscles during early embryogenesis. Myosin ATPase profiles were not altered by the stimulation regime. Galvanotaxis has been observed in a wide variety of organisms and also more recently in cell cultures (Jaffe and Nuccitelli, 1977). Orida and Feldman (1982) observed that the pseudopodial protrusive activity of the mouse peritoneal macrophage occurred toward the positive pole of the electric field, when this was greater than 400 mV/mm. Galvanotaxis of fibroblast and neural crest cells ap-

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peared at voltage gradients of around 150 mV/mm (Nuccitelli and Erickson, 1983; Erickson and Nuccitelli, 1984; Cooper and Keller, 1984; Nuccitelli, 1983), but was not observed in chick mesoderm cells (Stem, 1981). Fish epidermal cells migrate directionally toward the cathode in electric fields of 0.5- 15 V/cm, but in the upper range of this field strength, cell clusters and cell sheets break apart into single migratory cells (Cooper and Schliwa, 1984, 1985). Cells coupled by gap junctions will react as a single unit to an applied electric field. Cooper (1984) calculated that in a given field, the hyperpolarization and depolarization of cell membranes at the ends of an electrotonically coupled tissue are increased by a factor of 10-100 over single, uncoupled cells. He suggests that gap junctional coupling therefore increases the likelihood that cells are influenced by the weak electric fields which are commonly found in developing and regenerating tissues, as described in other parts of this review. At a field strength of 5 V/cm, Xenopus epithelial cells elongate perpendicularly with respect to the field (Luther ef al., 1983). Another effect of imposed field is the separation of proteins within the cells themselves by electrophoresis as was clearly shown by Dan (1972), who succeeded in the separation of hemoglobins contained in nucleated erythrocytes of fetal mouse. Pulsed electromagnetic fields (PEMFs) have also been used to induce time variable ionic currents in tissues, thereby modifying a number of cellular functions. Reviews of the literature up to 1976 on the biological effects of electric and magnetic fields of extremely low frequency have been published by Sheppard and Eisenbud (1977) and Marino and Becker (1977). Considerable interest was shown by surgeons because of the possible practical applications in regeneration, especially in bone repair and treatment of pseudarthroses (Bassett et a/., 1974; Bassett, 1982; Spadaro, 1982; Smith and Nagel, 1983; Marino, 1984; Papatheofanis et a/., 1984; Wittbjer et al., 1984; Fontanesi et a/., 1984; Becker, 1984) and treatment of tumors in mice (Smith and Feola, 1982; Akamine et al., 1985). Selective changes in cellular calcium (Bassett et a / . , 1979), in cyclic AMP levels (Fitton-Jackson and Bassett, 1980), and in the synthesis of collagen and proteoglycans (Fitton-Jackson and Farndale, 1981) have been reported. ATP hydrolysis may be induced by passage of a direct current through a solution of Ca2+-ATP (Leonard and Wade, 1982). Shteyer er a/. (1980) described electromagnetically induced DNA synthesis in calvaria. Liboff et a/. (1984) reported that human fibroblasts exhibited enhanced DNA synthesis when exposed to sinusoidally varying magnetic fields for a wide range of frequencies and amplitudes. They suggest that their results bring into question the allegedly specific magnetic wave shapes now used in therapeutic devices for helping the fusion of difficult bone fractures. They also suggest the possibility that mutagenic interactions arise directly from short-term changes in the earth’s field. Pulsing electromagnetic fields induce cellular transcription in isolated salivary glands of the dipteran Sciara coprophila, as monitored with [3H]uridine transcription auto-

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radiography, cytological peak translation, and analysis of isolated RNA fractions. In the single pulse mode, a 4-fold increase in total RNA was seen after 15 and 45 minutes exposure of the glands. The mRNA class exceeded control values 1 I-fold after 15 minutes and 13-fold after 45 minutes. The pulse train mode induced qualitative and quantitative responses in the patterns of RNA synthesis. The responses were different from those induced by ecdysone or heat shock (Goodman et af., 1983). These results suggest that some pulse modes may influence chromatin structure. Initiation of uncoiling of DNA by other types of pulses was reported by Hinsenkamp et af. (1978) and Chiabrera et af. (1979). Pulse trains are used clinically to elevate cellular calcium and to trigger calcification of fibrocartilage in disunited fractures. Single pulses lower cellular calcium in chondrocytes and stimulate bone accretion in patients with osteoporosis and avascular necrosis (Goodman et af., 1983). Archer and Ratcliffe ( 1983) exposed cultured 7-day embryonic chick tibiae to small alternating currents induced by pulsed magnetic fields. The main conclusion was that normal chondrogenesis was impaired by experimental treatment. Ca2+ uptake in cultured chick tibiae was observed by Colacicco and Pilla (1983). Low-energy, low-frequency fields inhibit the responses of bone cells in v i m to parathyroid hormone (Luben et al., 1982). The effects occurred with induced extracellular fields as small as 1 mV/cm or less, even though transmembrane potential gradients are typically 105V/cm. Luben et al. (1982) argue that the effectiveness of such weak stimuli in generating cellular processes must depend on a series of amplification mechanisms either before or during the transmembrane coupling of the stimuli. The field effects might perhaps be mediated by interference with hormone-receptor interaction or by blocking receptorcyclase in the membrane. Externally applied electric fields have been used to stimulate neurite outgrowth in culture (Sisken and Smith, 1975; Sisken et af., 1984; Patel and Poo, 1984) and nerve regeneration and function (Ito and Bassett, 1983; Murray et af., 1984; Cullen and Spadaro, 1983; Orgel et af., 1984; O’Brien et al., 1984, this list being not at all exhaustive). Delport et af.(1984) applied various types of PEMFs to isolated rat skin and examined the uptake of two labeled amino acids and [6-3H]thymidine. Stimulation of DNA synthesis was never observed. The uptake of labeled amino acids was stimulated only when PEMFs characterized by specific combinations of different electromagnetic parameters (maximum 5 kHz, maximum pulse width under 20 psec) were applied vertically upon the skin tissue. The uptake pattern of the two amino acids was different. No effects were observed with the electromagnetic fields running parallel to the skin flaps. An increased amino acid transport across the plasma membranes is supposed to be the immediate result of the effective PEMFs. Exposure to a 2-psec, 3-kV/cm pulse elicited a 50% loss of intracellular K in mouse spleen lymphocytes in vitro. No significant effects +

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on the response to various mitogens were observed (Smith and Cleary, 1983). The aggregation between lectins and lymphocyte surface receptors could be affected strongly by a low level electric field induced in the cell suspension by a time-varying magnetic field (Chiabrera et al., 1984). Concanavalin A receptors on the surface of cultured Xenopus myoblasts redistributed in response to monopolar, pulsed electric fields (Lin-Liu and Adey, 1984). Em values of MadinDarby canine kidney cells in vitro were reduced by approximately 20 mV in 4 hours by a pulsed electric field (660 psec, 150 mV/cm, 15 Hz; Strope er al., 1984). High-strength electric currents and fields can alter plant physiology by the production of heat within the plant tissue and by ionization of air molecules at the plant tips. An applied weak electric field of 5 kV/cm, 60 Hz resulted in a decrease of about 5% in the germination rate of sunflower seeds (Marino et a / ., 1983). ATP synthesis in chloroplasts of lettuce can to some extent be stimulated by an external electric field (Vinkler and Korenstein, 1982). Pulsed magnetic fields stimulated the number of roots that formed on plant cuttings of Forsythia hardwood and Begonia, but not the length. Other species did not respond (Smith and Mays, 1984). Continuous and intermittent exposure of the slime mold Physarum polycephalum to low-frequency electromagnetic fields both lengthen the mitotic cycle but have opposite effects on the respiration rate (Goodman et a / ., 1984). Electric field-induced cell-to-cell fusion seems to offer a lot of new possibilities in basic and applied biological research (Zimmerman and Vienken, 1982; Berg et al., 1984). Delgado et al. (1982) reported that very weak, lowfrequency electromagnetic fields induced a very significant effect on embryological development of chicken eggs exposed for 48 hours. However, Maffeo et al. (1985) duplicated Delgado’s experiments as carefully as possible, but they could not observe differences between exposed, sham exposed, and control eggs. The possibility of hazardous effects resulting from electric fields around 60-Hz high-voltage overhead transmission lines on intact animals and plants has been repeatedly suggested. Sometimes 60-Hz bioeffects were observed, sometimes not (for references, see Hart and Marino, 1982). In this respect, it has already been established that many organisms are surrounded by a dipolar continuous current field. In all fish, the mouth region acts as the positive and the gill regions as the negative pole of the field. The field, which is modulated by mechanical breathing, can be used for communication as well as for the location of matter. Shoals of migrating fish appear to be held together by their common electric field, They probably use certain potential levels in the sea for navigatory purposes. An act of perception can be initiated by an astonishingly low isolated amount of energy. Although already repeatedly emphasized, it is still largely overlooked that electrical fields are also produced by terrestrial animals. Various types of insects (e.g., bees), birds, and furred animals reach relatively high amplitudes. The mechanisms by which this is achieved, the influence of atmospheric ions, and the biological significance have been excellently reviewed by

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Warncke (1979). Chronic exposure of rats from conception to about 120 days of age to an unperturbed vertical 60 Hz, 80 kV/m electric field did not alter food or water intake but resulted in a slight initial developmental delay (Seto et a / . , 1983). The problem with intact animals is that it is difficult to differentiate whether the observed effects are due to direct interaction of the 60-Hz electric field with the general cellular activities or because the animal has sensed the field via a nervous system mediation which then causes changes in metabolism. To overcome this problem Lymangrover et al. (1983) tested whether PEMFs could affect corticosterone production (a hormone involved in stress) in slices of adrenal cortex of rats. They found that exposure to a 60-Hz electric field at 10 kV/m caused a highly significant, 3-fold elevation in the steroidogenic response of this tissue after administration of adrenocorticotropic hormone under in vitro superfusion conditions: 5 , 100, and 1000 kV/m had no effect. Eisenbach et al. (1983a) studied the effect of an electromagnetically induced electric field on macroscopic assays of chemotaxis and mobility in Escherichia coli. Under the conditions used, mobility was doubled and chemotaxis was inhibited by 70%. The authors suggest that an electrical process, but not fluctuations in the membrane potential (Margolin and Eisenbach, 1984), may be involved in the chemotaxis machinery. An imposed field might perturb the hypothetical electric signaling, modulate one or more proteins involved in chemotaxis, or modulate the ion currents involved in chemotaxis (Eisenbach, 1982; Eisenbach et af., 1983b, 1984). The interpretation of the results obtained on different objects with different methodologies is very difficult because it is as yet unclear which molecules or biochemical reactions are influenced by a given field. Strong fields may cause heating of the object if no measures are taken to prevent this. Different waveforms produce different biological responses (Bassett et d., 1982; Goodman et al., 1983). According to Ito and Bassett (1983) it may be possible that the same type of PEMF may stimulate bone or neural repair by different mechanisms of action.

V. Closing Remarks It seems as if in biology two closely linked cellular properties, namely the asymmetrical distribution of ion pump/channel activity in the plasma membrane and the ability of cells to drive ioniclelectric currents through themselves, have, hitherto, been largely underestimated. Cells which are symmetrical with respect to all their plasma membrane properties are probably rare, if they exist at all. Asymmetrical mitosis, where it occurs, may also be more important than generally thought. When the numerous data, collected by investigators in quite different disciplines, are brought together in a conceptual framework, the idea emerges

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that all cells are probably miniature electrophoresis chambers, at least during part of their life cycle. This electrical dimension of cells has already provided keys to a better understanding of polarity, differentiation during embryonic development, pattern formation, and the mode of action of hormones. Practical applications have already been realized: natural regeneration of amputated last joints of fingers or toes of children below the age of puberty (Illingworth, 1974) and the healing of difficult bone fractures with the help of weak currents (Section IV). The electrical dimension of cells may also contribute to a better definition of the living state: in our opinion a cell is dead when it has irreversibly lost its ability to realize its electrical dimension. Nature apparently uses ions, and the electric current which can be carried by them, as a kind of universal language for coordinating intracellular activities and certain relations between adjacent cells. Nature makes use of only a limited number of basic principles, as suggested by Solvay (1893), but shows herself a master by developing these few principles to a very high degree of versatility. This is of crucial importance if these principles are to be of use under a great variety of different conditions, which are furthermore often subject to continuous change. The urkaryote, which is supposed to be the common ancestor of all eukaryotes, lived in the “primordial sea.” During that period hormones or other macroregulatory substances were probably not as yet formed or were scarce. Changes in the cell’s external environment, which somehow required induction or change in the synthetic activity of the nucleus, had to be translated at the level of the plasma membrane into messages which could be understood by the nucleus, permitting enzymes to start transcription at the right site or by making other parts of the genome inaccessible to these enzymes. In our opinion, changes in ion permeability of the plasma membrane, perhaps accompanied by changes in ionic/electrical current, were the most appropriate and easiest means of delivering the message. Or as Szego and Pietras (1984) state: “If ever there was a universal indicator of cellular activation (or subduction), it is surface membrane destabilization (or stabilization). All else follows from this primary event. Changes in the ionic constitution of the nucleus may have resulted in changes in chromatin conformation. If ion fluxes were indeed the original key to the epigenetic use of the genomic information of the urkaryote, it is reasonable to assume that this basic principle has been preserved in the course of evolution, just as the basic principles of the genetic code have been preserved. It also follows that, besides changes in the genetic code, changes in the distribution of ion pumps/channels over the plasma membrane, changes in the functioning of ion pumplchannel activity, and in the symmetry of mitosis may have been important, but hitherto overlooked, mechanisms in evolution. Indeed, just as happens during differentiation in embryonic development when many cell types are formed with identical genetic information in their nuclei but which are all in ”

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different ways restricted by epigenetic mechanisms in the use they make of this information, it is conceivable that, in the course of evolution, variability could have been generated by introducing restrictions in the operative key to the genetic code without introducing changes in the genes themselves (De Loof, 1985a; Peferoen and De Loof, 1986). Oppenheimer (in Willier and Oppenheimer, 1974) predicted that with respect to the understanding of embryonic development, the most likely postulate to appear in the near future would involve factors, ranging from the ionic to the macromolecular, relating to the structure and function of membranes (“These are the cell organelles whose time is now ripe”). This seems at present to have become true. She also wrote that “scientific pendula have long swings”: it, indeed, took nearly 3 centuries (Wu, 1984) from the early experiments with electric fish to the formulation of the self electrophoresis concept and its implications. Although this concept offers exciting new perspectives for several disciplines of biology, research in this area will not be as easy as it is in molecular biology. While it is common practice to homogenize cells in order to study biochemical processes, the effects of ionslelectric current on cells or organisms should evidently be carried out on intact structures as homogenization destroys all electrical properties. We are confident that in the near future more and more adequate techniques will be developed allowing greater experimental refinement.

ACKNOWLEDGMENTS My special thanks to Mr. H. Van den bergh for his very valued bibliographic support and help with text correction. I am very grateful to my colleagues and collaborators Dr. F. Ollevier, Dr. J. Vereecke, Dr. M. Peferoen, J. Evans, J. Geysen, and B. Verachtert for their helpful comments. Thanks to Mrs. M. Van der Eeken for typing the several drafts of the manuscript and to Mrs. J . Puttemans for making the illustrations.

REFERENCES Agard, D. A,, and Sedat, J. W. (1983). Nature (London) 302,676-681. Agnew, W . S. (1984). Annu. Rev. Physiol. 46, 517-530. Ahearn, G. A. (1980). Am. J . Physiol. 239, CI-CIO. Aidley, D. J. (1981). “The Physiology of Excitable Cells.” Cambridge Univ. Press, London and New York. Akamine, T., Muramatsu, H., Hamada, H.,and Sakou, T. (1985). J . Cell. Physiol. 124, 247-254. Almers, W . , and Stirling, C. (1984). J. Membr. Eiol. 77, 169-186. Almers, W., Stanfield, P. R., and Stiihmer, W. (1983). J . Physiol. (London) 336, 261-284. Andersson, G. (1979). J. Theor. Eiol. 77, 1-18. Andersson, R. P., and Roth, J. R. (1981). Proc. Nut/. Acad. Sci. U.S.A. 78, 3113-3117.

336

ARNOLDDELOOF

Andrews, S. B., Mazurkiewicz, J. E., and Kirk, R. G. (1983). J . Cell Biol. 96, 1389-1399. Anner, B. M. (1985). Biochem. J . 227, 1-11. Archer, C. W., and Ratcliffe, N. A. (1983). J. Exp. Zool. 225, 243-256. Arnold, C., Gulbenkian, S., Carmo-Fonseca, M., and David-Ferreira, J. F. (1983). B i d . Cell. 47, 161- 170. Ashburner, M., and Cherbas; P. (19,776). Mol. Cell. Endocrinol. 5 , 89-107. Axelrod, D. (1983). J . Membr. Biol. 75, 1-10. Axelrod, D., Ravdin, R., Koppel, D. E., Schlessinger, J., Webb, W. W., Elson, E. L., and Podleski, T. R. (1976). Pioc. Nutl. Acud. Sci. U.S.A. 73, 4594-4598. Azzi, A. (1984). Experienriu 40,901-906. Baerentsen, H. J., Christensen, O., Thomsen, P. G., and Zeuthen, T. (1982). J . Membr. Biol. 68, 215-225. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1962a). J. Physiol. (London) 164, 330-354. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1962b). J. Physiol. (London) 164, 355-374. Bakker, E. P. (1978). Biochemistry 17, 2899-2904. Balinsky, B. I. (1975). “An Introduction to Embryology.” Saunders, Philadelphia, Pennsylvania. Barish, M. E. (1983). J. Physiol. (London) 342, 309-325. Barish, M. E. (1984). Dev. Biol. 105,29-40. Barish, M. E., and Baud, C. (1984). J. Physiol. (London) 352, 243-263. Barker, A. T., Jaffe, L. F., and Vanable, J. W., Jr. (1982). Am. J . Physiol. 242, R358-R366. Barth, L. G., and Barth, L. J. (1969). Dev. Biol. 20, 236-262. Barth, L. G., and Barth, L. J. (1972). Dev. Biol. 28, 18-34. Barth, L. G., and Barth, L. J. (1974a). Dev. Biol. 39, 1-22. Barth, L. G., Barth, L. J., and Nelson, I. (1960). Anut. Rec. 137, 337-338. Barth, L. J. (1964). In “Physiology of the Amphibia” (J. A. Moore, ed.), pp. 469-544. Academic Press, New York. Barth, L. J., and Barth, L. G. (1974b). Biol. Bull. 146, 313-325. Bartles, J. R., Braiterman, L. T., and Hubbard, A. L. (1985a). J. Cell Biol. 100, 1126-1138. Bartles, J. R., Braiterman, L. T., and Hubbard, A. L. (1985b). J . Biol. Chem. 260, 12792-12802. Barzilai, A., Spanier, R., and Raharnimoff, H. (1984). Proc. Nutl. Acud. Sci. U.S.A. 81, 65216525. Bass, A. H., and Hopkins, C. D. (1983). Science 220, 971-973. Bassett, C. A. L. (1971). In “Biochemistry and Physiology of Bone” ( G . H. Bourne, ed.), 2nd Ed., pp. 1-76. Academic Press, New York. Bassett, C. A. L. (1982). Culcif. Tissuelnr. 34, 1-8. Bassett, C. A. L., Pawluk, R. J., and Pilla, A. A. (1974). Science 184, 575-577. Bassett, C. A. L., Chokski, H. R., Hernandez, E., Pawluk, R. J., and Strop, M. (1979). In “Electrical Properties of Bone and Cartilage: Experimental Effects and Clinial Applications” (C. T. Brighton, J. Black, and S. R. Pollack, eds.), pp. 427-453. Grune & Stratton, Orlando, Florida. Bassett, C. A. L., Valdes, M. G., and Hernandez, E. (1982). J. Bone Jt. Surg. Am. Vol. 64, 888895. Bates, G. W., Goldsmith, M. H., and T. M. Goldsmith (1982). J. Membr. Biol. 66, 15-23. Baud, C. (1983). Dev. Biol. 99, 524-528. Baud, C., and Barish, M. E. (1984). Dev. Biol. 105, 423-434. Baud, C., and Kado, R. T. (1984). J. Physiol. (London) 356, 275-289. Baud, C., Kado, R. T., and Marcher, K. (1982). Proc. Nutl. Acud. Sci. U.S.A. 79, 3188-3192. Baulieu, E. E. (1983). Exp. Clin. Endocrinol. 81, 3-16. Baulieu, E. E., and Schorderet-Slatkine, M. (1983). J. Steroid Biochem. 19, 139-145. Baulieu, E. E., Godeau, J. F., and Schorderet-Slatkine, M. (1978). Nature (London) 275,593-598. Baulieu, E. E., Schorderet-Slatkine, S., Le Goascogne, C., and Blondeau, J.-P. (1985). Dev. Growth Direr. 27, 223-231.

THE ELECTRICAL DIMENSION OF CELLS

337

Becker, R. 0. (1982). J. Bioelectr. 1, 239-264. Becker, R. 0. (1984). J. Bioelectr. 3, 105-1 18. Behrens, H: M., Weisenseel, M. H., and Sieves, A. (1982). Pluntfhysiol. 70, 1079-1083. Bell, P. R. (1975). Endeuvour 34, 19-22. Bennett, B., and Stenbuck, P. J. (1979a). J. Biol. Chem. 254, 2533-2541. Bennett, B., and Stenbuck, P. J. (1979b). Nature (London) 280, 468-473. Bennett, M. V. L., and Grundfest, H. (1959). J. Gen. Physiol. 42, 1067-1104. Bennett, M. V. L., Spira, M. E., and Spray, D. C. (1978). Dev. Biol. 65, 114-125. Berg, H., Augsten, K.,Bauer, E., Forster, W., Jacob, H. E., Miinlig, P., and Weber, H. (1984). Bioelectrochem. Bioenerg. 12, 119-134. Bemdge, M. J., and Irvine, R. F. (1984). Nature (London) 312, 315-321. Bemdge, M. J., and Prince, W. T. i1972). In “Advances in Insect Physiology” (J. E. Treherne, M. J. Berridge, and V. B. Wigglesworth, eds.), Vol. 11, pp. 1-49. Academic Press, New York. Berry, S. J. (1985). In “Developmental Biology” (L. W. Browder, ed.), Vol. 1, pp. 351-384. Plenum, New York. Betz, W. J., and Caldwell, J. H. (1984). J. Gen. Physiol. 83, 143-156. Betz, W. J., Caldwell, J. H., Ribchester, R.R., Robinson, K. R., and Stump, R. F. (1980). Nature (London) 287, 235-237. Bingley, M. S. (1966). Exp. Cell Res. 43, 1-12. Bittar, E. E., Chambers; G., and Fisher, E. H. (1982). J. Physiol. (London) 333, 39-52. Bland, K. P., Bountra, C., and House, R. C. (1983). J . Physiol. (London) 343, 103P. Blitzer, B. L., and Boyer, J. L. (1978). J. Clin. Invest. 62, 1104-1 108. Blondeau, J.-P., and Baulieu, E. E. (1984). Biochem. J . 219, 785-792. Bloodgood, R. A., and Levin, E. N. (1983). J . Cell Biol. 97, 397-404. Bloom, J. W., Butler, J., Brierly, J., and Cosmos, E. (1982). J. Neurosci. 5, 414-420. Boonstra, J., Moolenaar, W. H., Harrison, PH. H., Moed, P., van der Saag, P. T., and De Laat, S. W. (1983). J. Cell Biol. 97, 92-98. Borgens, R. B. (1982). Int. Rev. Cyrol. 76, 245-298. Borgens, R. B. (1983). frog. Clin.Biol. Res. llO(A), 597-608. Borgens, R. B. (1984). Dizerentiution (Berlin) 28, 87-93. Borgens, R. B., Vanable, J. W., Jr., and Jaffe, L. F. (1977a). Proc. Narl. Acud. Sci. U.S.A. 74, 4528-4532. Borgens, R. B., Vanable, J. W., Jr., and Jaffe, L. F. (1977b). J. Exp. Zool. 200, 403-416. Borgens, R. B., Vanable, J. W., Jr., and Jaffe, L. F. (1979a). J. Exp. Zool. 209, 49-55. Borgens, R. B., Vanable, J. W., Jr., and Jaffe, L. F. (1979b). J. Exp. Zoo/. 209, 377-386. Borgens, R. B., Jaffe, L. F., and Cohen, M. J. (1980). Proc. Nutl. Acud. Sci. U.S.A. 77, 12091213. Borgens, R. B., Rouleau, M. F., and DeLanney, L. E. (1983). J. Exp. Zool. 228, 491-503. Borgens, R. B., McGinnis, M. E., Vanable, J. W., Jr., and Miles, E. S. (1984). J. Exp. Zool. 231, 249-256. Boyer, P. D., Chance, B., Ernster, L., Mitchell, P., Racker, E., and Slater, E. C. (1977). Annu. Rev. Biochem. 46, 955-1026. Boyle, P. J., and Conway, E. J. (1941). J. Physiol. (London) 100, 1-63. Brawley, S. H., Wetherell, D. F., and Robinson, K. R. (1984). Proc. Nurl. Acud. Sci. U.S.A. 81, 6064-6067. Bridge, J. H. B., and Bassingthwaighte, J. B. (1983). Science 219, 178-180. Brighton, C. T. (1981). J . BoneJr. Surg. Am. Vol. 63, 847-851. Brower, D. L., and Giddings, T. H. (1980). J. Cell Sci. 42, 279-290. Brown, B. A. (1980). “Hematology, Principles and Procedures.” Lea & Febiger, Philadelphia, Pennsylvania. Brown, D. D. (1981). Science 211, 667-674.

338

ARNOLD DE LOOF

Bunning, E. (1958). Proroplasmatologia 8, 2-86. Bums, C. P., and Rozengurt, E. (1984). J. Cell Biol. 98, 1082-1089. Caldwell, J . H., and Betz, W. J. (1984). J . Gen. Physiol. 83, 157-173. Cameron, I. L., LaBadie, D. R. L., Hunter, K. E., and Hazlewood, C. F. (1983). J . Cell. Physiol. 116, 87-92. Cantley, L. C. (1981).In “Current Topics in Bioenergetics” (D. R. Sanadi, ed.), Vol. I I , pp. 201237. Academic Press, New York. Carafoli, E., and Crompton, M. (1976). Symp. SOC. Exp. Biol. 30, 89-1 15. Cartaud, A,, and Ozon, R. (1980). J . Biol. Chem. 225, 9404-9408. Cartwright, I. L., Abmayr, S. M., Fleischmann, G., Lowenhaupt, K., Elgin, S . , Keene, G., and Howard, G. C. (1982). CRC Crir. Rev. Biochem. 13, 1-86. Casadei, J. M., Gordon, R. D., Lampson, L. A., Schotland, D. L., and Barchi, R. L. (1984). Proc. Narl. Acad. Sci. U.S.A. 81, 6227-6231. Catterall, W. A. (1981). J. Neurosci. 1, 777-783. Catterall, W. A. (1984). Science 223, 653-661. Chambers, E. L. (1975). J . Cell Biol. 67, 60a. Chambers, E. L. (1976). J. Exp. 2001.197, 149-154. Chan, S. T. H., and Wong, P. Y. D. (1978). J . Physiol. (London) 279, 385-394. Chang, K. S., and Snellen, J. W. (1982). J. Exp. Zool. 221, 193-203. Chapman, L. M., and Wondergem, R. (1984). J. Cell. Physiol. 121, 7-12. Charbonneau, M., Moreau, M., Picheral, B., Vilain, J. P., and Guerrier, P. (1983). Dev. Biol. 98, 304-3 18. Cheng, N., Hoogmartens, M. J., Mulier, J. C., Sansen, W. M., and De Loecker, W. (1982). Biochem. SOC. Trans. 10, 468-469. Chiabrera, A., Hinsenkamp, M., Pilla, A. A., Ryaby, J., Ponta, O., Belmont, A,, Beltrame, F., Grattarola, M., and Nicolini, C. (1979). J . Hisrochem. Cyrochem. 27, 375-381. Chiabrera, A., Grattarola, M., and Viviani, R. (1984). Bioelecrromagnetics 5, 173-191. Chiu, S . Y.,and Ritchie, J. M. (1981). J. Physiol. (London) 313, 415-437. Chiu, S. Y., and Ritchie, J. M. (1982). J . Physiol. (London) 322, 485-501. Chow, I., and Poo, M.-M. (1984). J. Physiol. (London) 346, 181-194. Ciapa, B., De Renzis, C., Girard, J. P., and Payan, P. (1984a). J . Cell. Physiol. 121, 235-242. Ciapa, B., Allemand, D., Payan, P., and Girard, J. P. (1984b). J . Cell. Physiol. 121, 243-250. Ciba Foundation (1983). “Molecular Biology of Egg Maturation” (Ciba Found. Symp. No. 98). Pitman, London. Cicirelli, M. F., Robinson, K. R.,and Smith, L. D. (1983). Dev. Biol. 100, 133-146. Cochran, G. V. B., Pawluck, R. J., and Bassett, C. A. L. (1968). Clin. Orrhop. 58, 249-270. Coffey, J. W., and de Duve, C. (1968). J . Biol. Chem. 243, 3255-3263. Colacicco, G., and Pilla, A. A. (1983). Bioelecrrochem. Bioenerg. 10, 119-131. Cone, C. D., Jr. (1970). Oncology 24, 438-470. Cone, C. D., Jr. (1971a). J . Theor. Biol. 30, 151-181. Cone, C. D., Jr. (1971b). J . Theor. Biol. 30, 183-194. Cone, C. D., Jr., and Tongier, M. (1973). J . Cell. Physiol. 82, 373-386. Conti, F., Hille, B., Neumcke, B., Nonner, W., and Stampfli, R. (1976).J . Physiol. (London)262, 699-727. Conway, E. J., Brady, R. G., and Carton, E. (1950). Biochem. J . 47, 369-374. Cook, D. L., and Hales, N. (1984). Nature (London) 311, 271-273. Coombs. J. S . , Curtis, D. R., and Eccles, J. C. (1957). J . Physiol. (London) 139, 232-249. Cooper, M. S. (1984). J. Theor. Biol. 111, 123-130. Cooper, M. S., and Keller, R. E. (1984). Proc. Narl. Acad. Sci. U . S . A . 81, 160-164. Cooper, M. S . , and Schliwa, M. (1984). Biophys. 1.45, 98a.

THE ELECTRICAL DIMENSION OF CELLS

339

Cooper. M. S., and Schliwa, M. (1985). J. Neurosci. Res. 13, 223-244. Costa, M. R.. Casnellie. J. E., and Catterall, W. A. (1982). J . Biol. Chem. 257, 7918-7921. Cox, K.H., Deleon, D. V., Angerer, L. M., and Angerer, R. C. (1984). Dev. B i d . 101,485-502. Cox, M.. Sterns. R. H., and Singer, 1. (1978). N . Engl. 1.Med. 299, 525-532. Cretin, H. (1982). J . Memhr. Biol. 65, 175-184. Cullen, J. M., and Spadaro, J. A. (1983). J. Bioelectr. 2, 57-75. Dale, B., and de Felice, L. (1984). Dev. B i d . 101, 235-239. Dale, B., De Santis, A., and Ortolani, G. (1983). Dev. B i d . 99, 188-193. Dan, M. S . (1972). Exp. Cell Res. 72, 582-584. Danielli, J. F., and Davson, H. (1935), J . Cell. Comp. Physiol. 5 , 495-508. Darwin. C. (1861 ). ”The Variation of Animals and Plants under Domestication.” Murray, London. Davidson, E. N. (1976). “Gene Activity in Early Development.” Academic Press, New York. De Brabander, M., Gevens, G . , Muydens, R., Moeremans, M . , and De Mey, J. (1985). Cytohios 43, 273-283. Decker, R. S . (1981). Dev. B i d . 81, 12-22. Decker, R. S . , and Friend, D. S. (1974). J. Cell Biol. 62, 32-47. De Coursey, T. E., Chandy, K. G., Gupta, S., and Cahalan, M. D. (1984). Nature (London) 307, 465-468. De Laat, S. W., Buwalda, R. J. A,, and Habets, A. M. M. C. (1974). Exp. C e l l R e s . 89, 1-14. Delgado, J . M. R., Leal, J., Monteagudo, J. L., and Gracia, M. G. (1982). J. Anur. 134, 533-551. De La Pena, P., Barros. F., Gascon, S., Ramos, S . , and Lazo, P. S. (1982). Eur. J . Biochem. 123, 447-453. De Loof, A. (1983). Comp. Biochem. Phvsiol. A 74, 3-9. De Loof, A. ( 1985a). Comp. Biochem. Physiol. A 80,453-459. De Loof. A. (1985b). Ann. Soc. R . Zool. Belg. 115, 121-136. De Loof, A. (1985~)./nsec/ Biochem. 16, 169-173. De Loof, A,, and Geysen, J. (1983). Bioelectrochem. Bioenerg. 11, 383-388. De Loof, A.. Huybrechts, R., and Briers, T. (1981). Ann. Soc. R . Zool. Belg. 110, 179-184. De Loof, A., Briers, T., Huybrechts, R.. Ollevier, F., Peferoen, M., Stoppie, P., and Stynen, D. (1982). Ann. Soc. R . Zool. Belg. 112, 3-22. Delport, P. H., Cheng, N . , Mulier, J. C . , Sansen, W. M., and De Loecker. W . (1984). Biochem. Soc. Trans. 12, 437-438. De Weer, P., and Rakowski, R. F. (1984). Natrtre (London) 309, 450-452. Dictus, W.. Van Zoelen. E., Tetterloo, P., Tertoolen, L.. De Laat, S . , and Bluemink, J. (1984). Dev. B i d . 101, 201-211. Dittmann, F., Ehni. R., and Engels, W. (1981). Wilhelm Rou.r’sArch. Dev. B i d . 190, 221-225. Dodge, F. A., Jr., and Cooley, J. W. (1973). IBM J . Res. Dev. 17, 219-229. Doll. S., Rodier. F., and Willenbrink, J. (1979). Planta 144, 407-41 I . Dolowy, K. (1984). Prog. Sut$ Sci. 15, 245-368. Dorn, A,. and Weisenseel, H. (1982). Protoplasma 113, 89-96. Dorresteijn, A. W. C., Wagemaker, H. A . , De Laat, S. W., and van den Biggelaar. J. A. M. ( 1983). Wilhelm Rori.r’s Arch. Dev. B i d . 192, 262-269. Douglas. B. S. (1972). Airst. Paediatr. J . 8, 86-91. Douva, S . , Harrington, C. A., and Bonner, J . (1975). Proc. Nut/. Acad. Sci. U.S.A. 72,3902-3906. Dubois-Reymond. E. (1843). Ann. Phvs. Chem. (Leipzig) 58, 1-30. Dunant, Y..and Israel, M. (1985). Sci. Am. 252,40-48. Durbin. R . P. (19x1). J. Memhr. B i d . 61, 141. Duval. D.. Durant. S.. and Homo-Delarche, F. (1983). Biochim. Biophvs. AC/U 737, 409-422. Ebihara, L.. and Speen, W. C. ( 1984). Biophvs. J . 46, 827-830. Eddy. A. A . (197X). Curr. Top, Memhr. Transp. 10, 271-360.

340

ARNOLDDELOOF

Edelmann, L. (1977). J. Microsc. (OxfbrdJ 112, 243-248. Edelmann, L. (1978). Microsc. Acru Suppl. 2, 166-174. Edelmann, L. (1980). Histochemistry 67, 233-242. Edelmann, L. (1981). In “International Cell Biology 1980-1981” (H. G. Schweiger, ed.), pp. 941948. Springer-Verlag, Berlin and New York. Edidin, M. (1972). I n “Membrane Research” (C. F. Fox, ed.), pp. 15-25. Academic Press, New York. Edidin, M. (1981). I n “Membrane Structure” (M. Finean, ed.), pp. 37-82. Elsevier, New York. Edwards, C. (1982). Neuroscience 7, 1335-1366. Eisen, A,, and Reynolds, G. T. (1984). J . CellEiol. 99, 1878-1882. Eisenbach, M. (1982). Biochemistry 21, 6818-6825. Eisenbach, M., Zimmerman, J. R., Ciobotariu, A., Fischler. H., and Korenstein, R. (1983a). Eioelecrrochem. Eioenerg. 10, 499-5 10. Eisenbach, M., Raz, T., and Ciobotariu, A. (1983b). Biochemistry 22, 3293-3298. Eisenbach, M., Margolin, Y., Ciobotariu, A., and Rottenberg, H. (1984). Eiophys. J . 45,463-467. Ellisman, M. H., and Levinson, S. R. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 6707-671 1. Els, W. J., and Helman, S. I. (1981). Am. J . Physiol. 241, F279-F288. English, L. H., Magelky, B. K., and Marks, E. P. (1984). In Vitro 20, 71-78. Epel, D., Steinhardt, R., Humphreys, T., and Mazia, D. (1974). Dev. B i d . 40, 245-255. Erickson, C. A., and Nuccitelli, R. (1984). J. CeN Eiol. 98, 296-307. Erlij, D., and Grinstein, S. (1976). J . Physiol. (London) 259, 13-31. Etemadi, A. H. (1980a). Eiochim. Biophys. Acra 604, 347-422. Etemadi, A. H. (1980b). Eiochim. Eiophys. Actu 604,423-475. Eusebi, F., Mangia, F., and Alfei, L. (1979). Nature (London) 277, 651-653. Eusebi, F., Pasetto, N., and Siracusa, G. (1983). Biol. Reprod. 28, 86. Evans, W. H. (1980). Eiochim. Eiophys. Acra 604, 27-64. Fein, A., and Szuts, E. Z. (1982). “Photoreceptors: Their Role in Vision.” (IUPAB Biophysics series). Cambridge Univ. Press, London and New York. Felle, H., Porter, J. S . , Slayman, C. L., and Kaback, H. R. (1980).Biochemistry 19, 3585-3590. Fertuck, H. C., and Salpeter, M. M. (1976). J. CeNEiol. 69, 144-158. Finean, J. B., Coleman, R., and Michell, R. H. (1979). “Membranes and Their Cellular Functions.” Blackwell, Oxford. Fitton-Jackson, S. F., and Bassett, C. A. L. (1980). In “Tissue Culture in Medical Research (11)” (R. J. Richards and K. T. Rajan, eds.), pp. 21-28. Pergamon, Oxford. Fitton-Jackson, S., and Farndale, R. (1981). Trans. Orrhop. Res. SOC. 6, 300. Florkin, M., and Jeuniaux, C. (1974). In “The Physiology of Insecta” (M. Rockstein. ed.), 2nd Ed. Vol. 5 , pp. 255-307. Academic Press, New York. Fontanesi, G., Dal Monte, A., Rinaldi, E., Traina, G. C., Mammi, G. I., Giancecchi, F., Rotini, R., Poli, G., Negri, V., Virgili, B., and Caldossi, R. (1984). J . Bioelecrr. 3, 155-175. Forgac, M., Cantley, L., Wiedenmann, B., Altstiel, L., and Branton. D. (1983). Proc. Nut/. Acud. Sci. U.S.A. 80, 1300-1303. Freeman, J . A., Weiss, I. M., Snipes, G. I., Mayes, B., and Norden, I. J. (1981). SOC.Neurosci. Symp. Absrr. 7, 550. Freeman, J . A., Manis, P. B., Snipes, G. J., Maynes, B. N., Samson, P. C., Wikswo, J . P., and Freeman, D. B. (1985). J. Neurosci. Res. 13, 257-283. French, R. J., and Horn, R. (1983). Annu. Rev. Biophys. Eioeng. 12, 319-356. Friedenberg, Z. B., and Brighton, C. T. (1966). J . Bone Jt. Surg. Am. Vol. 48, 915-923. Frizzell, R. A., and Schultz, S. C. (1972). J. Gen. Physiol. 59, 318-346. Fromter, E. (1972). J . Membr. B i d . 8, 259-301. Fromter, E., and Diamond, I. M. (1972). Nurure (LondonJ New Eiol. 235, 9-13.

THE ELECTRICAL DIMENSION OF CELLS

34 1

Frye, L. D., and Edidin, M. (1970). J. Cell Sci. 7, 319-335. Fukuda, E., and Yasuda, I. (1957). 1. Phys. SOC. Jpn. 12, 1158-1162. Fuortes, M. G. F., Frank, K., and Becker, M. C. (1957). J. Gen. Physiol. 40, 735-752. Gadenne, M., Van Zoelen, E. J. J., Tencer, R., and De Laat, S. W. (1984). Dev. Biol. 104,461468.

Garcia-Diaz, J. F., and Armstrong, W. Mc. D. (1980). J. Membr. Biol. 55, 213-222. Gardiner, D. M., and Grey, R. D. (1983). J . Cell Biol. 96, 1159-1 163. Geisow, M. (1982). Nature (London) 298, 515-516. Georgiou, P., Bountra, C., Bland, K. P., and House, C. R. (1983). Q. J . Exp. Physiol. Cogn. Med. Sci. 68, 687-700. Gerencser, G. A. (1982). Comp. Biochem. Physiol. A 72, 721-725. Gerson, D. F., and Kiefer, H. (1982). 1. Cell. Physiol. 112, 1-4. Gilkey, J. C., Jaffe, L. F., Ridgway, E. B., and Reynolds, G. T. (1978). J . Cell Biol. 76,448-466. Gilly, W. F., and Armstrong, C. M. (1984). Nature (London) 309, 448-450. Glaser, L., Whiteley, B., Rothenberg, P., and Cassel, D. (1985). Bioessuys 1, 16-20. Goffeau, A., and Slayman, C. W. (1981). Biochim. Biophys. Acra 639, 197-223. Goldfine, I. D., Purello, F., Clawson, G. A., and Vigneri, R. (1982). J . Cell. Biochem. 20, 29-39. Goldman, D. E. (1943). J . Gen. Physiol. 27, 37-60. Goodman, E. M.,Greenebaum, B., Marron, M. T., and Canick, K. (1984). J . Bioelecrr. 3,57-66. Goodman, R., Bassett, C. A. L., and Henderson, A. S. (1983). Science 220, 1283-1285. Could-Somero, M. (1981). Nature (London) 291, 254-256. Cow, N. A. R. (1984). J . Gen. Microbiol. 130, 3313-3318. Cow, N. A. R., Kropf, D. L., and Harold, F. M. (1984). J. Gen. Microbiol. 130, 2967-2974. Grant, Ph. (1978). “Biology of Developing Systems.” Holt, New York. Grey, R. D., Bastiani, M. J., Webb, D. J . , and Schertel, E. R. (1982). Dev. Biol. 89, 475-484. Griffiths, G., Brands, R., Burke, B., Louvard, D., and Warren, G. (1982). J. Cell Biol. 95, 781792.

Griffiths, G., Quinn, P., and Warren, G. (1983). J. Cell Biol. 96, 835-850. Grinell, A. D., and Brazier, M. A. B., eds. (1981). “The Regulation of Muscle Contraction Excitation-Contraction Coupling.” Academic Press, New York. Guggino, W. B., Windhager, E. E., Boulpaep, E. L., and Giebisch, G. (1982). J . Membr. Biol. 67, 143-154.

Gulian, D., and Diacumakos, E. G. (1977). J. Cell Biol. 72, 86-103. Gulley, R. L., and Reese, T. S. (1981). J. Cell B i d . 91, 298-302. Gurdon, J. B. (1962). Dev. Biol. 4, 256-273. Gurdon, J. B. (1974). “The Control of Gene Expression in Animal Development.” Oxford Univ. Press (Clarendon), London and New York. Guthrie, S. C. (1984). Narure (London) 311, 149-151. Gutknecht, J. (1967). J. Gen. Physiol. 50, 1818-1821. Gutzeit, H., and Koppa, R. (1982). J . Embryol. Exp. Morphol. 67, 101-111. Guy, M., Reinhold, L., and Michaeli, D. (1979). Planr Physiol. 64, 61-64. Habib, M. A,, and Bockris, J. O’M.(1982). J. Bioelecrr. 1, 289-294. Habib, M. A., and Bockris, J. O’M. (1984). J. Bioelectr. 3, 247-280. Haeyaert, P., Verdonck, F., and Wuytack, F. (1980). Arch. Int. Pharmacodyn. Ther. 244,333-335. Hagins, W. A. (1972). Annu. Rev. Biophys. Bioeng. 1, 131-158. Hagins, W. A., Penn, R. D., and Yoshikami, S. (1970). Biophys. J . 10, 380-412. Hagiwara, S. (1983). “Membrane Potentials Dependent Ion Channels in Cell Membrane.” (Society of General Physiologists Distinguished Lecture Series, Vol. 3). Raven, New York. Hagiwara, S., and Byerly, L. (1981). Annu. Rev. Neurosci. 4, 69-125. Hagiwara, S., and Jaffe, L. F. (1979). Annu. Rev. Biophys. Bioeng. 8, 385-416.

342

ARNOLDDELOOF

Hakim, R. S., and Baldwin, K. M. (1984). Am. Zoo/. 24, 169-175. Hale, C. C., Slaughter, R. S., Ahrens, D. C., and Reeves, J. P. (1984). Proc. Null. Acud. Sci. U.S.A. 81, 6569-6573. Hall, W . J., O’Donoghue, J. P., O’Regan, M. G., and Penny, W. I. (1976). J . Phvsiol. (London) 258, 731-753. Hamlyn, J. M., and Duffy, T. (1978). Biochem. Biophys. Res. Commun. 84, 458-464. Handler, J. S., and Orloff, J. (1973). In “Handbook of Physiology,” Section 8: Renal Physiology (J. Orloff and R. W. Berlina, eds.), pp. 791-814. Am. Physiol. SOC., Bethesda, Maryland. Hanrahan, J . W. (1984). Am. Zoo/. 24, 229-240. Harary, H. H., and Brown, J. E. (1984). Science 224, 292-294. Harold, F. M. (1977). Curr. Top. Bioenerg. 6 , 83-149. Harold, F. M. (1978). In “The Bacteria” (I. C. Gunsalus, L. N. Ornston, and T. R. Socatch, eds.), Vol. 6, pp. 463-521. Academic Press, New York. Harold, F. M., Kropf, D. L., and Caldwell, J. H. (1985). Exp. Mvcol. 9, 183-186. Hart, F. X.,and Marino, A. A. (1982). J. Bioelecrr. 1, 129- 154. Heinrich, V., Kaufmann, R., and Gutzeit, H. (1983). Diflerentiution 25, 10-15. Helman, S. I . , Cox, T. C., and Van Driessche, W. (1983). J. Gen. Physiol. 82, 201-220. Hill, A. (1980). J . Membr. Biol. 56, 177-182. Hille, B., and Schwarz, W. (1978). J. Gen. Physiol. 72, 409-442. Hinkle, L., McCaig, C. D., and Robinson, K. R. (1981). J. Physiol. (London) 314, 121-135. Hinsenkamp, M., Chiabrera, A., Ryaby, J., Pilla, A. A., and Bassett, C. A. L. (1978). Arm Orrhop. Belg. 44, 636-65 I . Hirai, S., Le Goascogne, C., and Baulieu, E. E. (1983). Dev. B i d . 100, 214-221. Hirano, T., and Takahashi, K. (1984). J. Physiol. (London) 347, 327-344. Hodges, T. K. (1976). Encycl. Plant Physiol. 2, 260-283. Hodgkin, A. L. (1958). Proc. R . Soc. London Ser. B . 148, 1-37. Hodgkin, A. L. (1964). “The Conduction of the Nervous Impulse.” Univ. Press, Liverpool. Hodgkin, A. L., and Huxley, A. F. (1939). Nature (London) 140, 710. Hodgkin, A. L., and Huxley, A. F. (1952a). J. Physiol. (London) 116, 449-472. Hodgkin, A. L., and Huxley. A. F. (1952b). J. Physiol. (London) 116, 473-496. Hodgkin, A. L., and Huxley, A. F. (1952~).J. Physiol. (London) 116, 497-506. Hodgkin, A. L., and Huxley, A. F. (1952d). J. Physiol. (London) 117, 500-544. Hodgkin, A. L., and Katz, B. (1949). J. Physiol. (London) 108, 37-77. Hodgkin, A. L., McNaughton, P. A., and Nunn, B. J. (1985). J . Physiol. (London) 358,447-468. Hoffner, N. J., and Diberardino, M. J. (1980). Science 209, 517-519. Hollemans, M., Donker-Koopman, W., and Tager, J. M. (1980). Biochim. Biophys. Acru 603, 171177. Horn, R. (1984). In “Ion channels: Molecular and Physiological Aspects” (W.D. Stein, ed.), pp. 53-97. Academic Press, New York. Horstadius, S. (1952). J. Exp. Zoo/. 120, 421-436. Hubbard, A. L., Bartles, J. R., and Braiterman, L. T. (1985). J. Cell Biol. 100, I 1 15-1 125. Hudson, R. L., and Schultz, S. G. (1984). Science 224, 1237-1239. Huebner, E. (1984). In “Advances in Invertebrate Reproduction” (W. Engels, ed.), pp. 97-105. Elsevier, Amsterdam. Hiilser, D., and Schatten, G. (1982). Gamete Res. 5, 363-377. Huttunen, M. T., and Akerman, K. E. 0. (1980). Biochim. Biophys. Acru 597, 274-284. Huxley, A. F. (1974). J. Physiol. (London) 243, 1-43. Huxley. H. E. (1972). In “The Structure and Function of Muscle” ( G . H. Bourne, ed.), 2nd Ed., Vol. I , pp. 301-387. Academic Press, New York. Hyde. 1. (1905). Am. J . Physiol. 12, 241-275.

THE ELECTRICAL DIMENSION OF CELLS

343

Illingworth, C. M. (1974). J. Pediufr. Surg. 9, 853-858. Illingworth, C. M., and Barker, A. T. (1980). Clin. Phvs. Phvsiol. Meus. I, 87-89. Ito, H., and Bassett, C. A. L. (1983). Clin. Orihop. Relur. Res. 181, 283-290. Ito, S . , and Loewenstein, W. R. (1965). Science 150, 909-910. Izant, J. G. (1983). Chromosoma 88, 1-10. Jacob, F., and Monod, J. (1961a). J . Mol. Biol. 3, 3 18-356. Jacob, F., and Monod, J. (1961b). ColdSpring Harbor Symp. Quanr. Biol. 26, 193-211. Jacobson, K., and Wojcieszyn, 1. (1981). Commenis Mol. Cell. Biophys. 1, 189-199. Jaffe, L. A. (1976). Nature (London) 261, 68-71. Jaffe, L. A. (1983). In “The Physiology of Excitable Cells’’ (A. Grinnell and W. Moody, eds.), pp. 211-218. Liss, New York. Jaffe, L. A., and Cross, N. L. (1984). Biol. Reprod. 30, 50-54. Jaffe. L. A., and Schlichter, L. C. (1985). J. Physiol. (London) 358, 299-319. Jaffe, L. A., Weisenseel, M. H., and Jaffe, L. F. (1975). J. Cell Biol. 67, 488-492. Jaffe, L. A., Sharp, A. P., and Wolf, D. P. (1983). Dev. Biol. 96, 317-323. Jaffe, L. F. (1966). Proc. Nut/. Acud. Sci. U.S.A. 56, I 102-1 109. Jaffe, L. F. (1968). Adv. Morphog. 7, 295-328. Jaffe, L. F. (1975). Science 187, 70-72. Jaffe, L. F. (1977). Nature (London) 265, 600-602. Jaffe, L. F. (1982). Symp. Soc. Dev. Biol. 40, 183-215. Jaffe, L. F. (1983). Dev. Biol. 99, 265-276. Jaffe, L. F., and Guerrier, P. (1981). Dev. Biol. 83, 370-373. Jaffe, L. F., and Nuccitelli, R. (1974). J . Cell Biol. 63, 614-628. Jaffe, L. F.. and Nuccitelli, R. (1977). Annu. Rev. Biophys. Bioeng. 6, 445-476. Jaffe, L. F., and Poo, M. M. (1979). J. Exp. Zool. 209, 115-128. Jaffe, L. F., and Stem, C. D. (1979). Science 206, 569-571. Jaffe, L. F., and Woodruff, R. I. (1977). J. Cell Biol. 75, 23A. Jaffe, L. F.. and Woodruff, R. I. (1979). Proc. Nut/. Acud. Sci. U.S.A. 76, 1328-1332. Jaffe, L. F., Robinson, K. R., and Nuccitelli, R. (1974). Ann. N.Y. Acud. Sci. 238, 372-389. Jeffery, W. R. (1982). Science 216, 545-547. Jeffery, W. R. (1984a). Dev. Biol. 103, 482-492. Jeffery, W. R. (1984b). Bioessays I, 196-199. Jensen, E. V., Suzuki, T., and Kawashima, T. (1968). Proc. Nail. Acud. Sci. U.S.A. 59,632-638. Johnson, C. H., and Epel, D. (1982). Dev. Biol. 92, 461-469. Johnson, R. G., and Scarpa. A. (1976). J . Biol. Chem. 251, 2189-2191. Jones, R. T., Johnson, R. T., Gupta, B. L., and Hall, T. A. (1979). J. Cell Sci. 35, 67-85. Jergensen, J. P. (1982). Biochim. Biophys. Acru 694, 27-68. Kaback, H. R. (1977). J. Cell. Physiol. 89, 575-593. Kaback, H. R. (1983). J . Membr. Biol. 76, 95- 112. Kado, R. T., Marcher, K., and Ozon, R. (1979). C. R. Hebd. Acud. Sci. 288, I 187- 1 189. Kaiser, J., Lang, A. B., and Went, D. F. (1982). Exp. Cell Res. 139, 460-463. Kameyama, M., Kakei, M., Sato, T., Shibasaki, T., Matsuda, H., and Irisawa. H. (1984). Nuture (London) 309, 354-356. Kanno, Y.,and Loewenstein, W. R. (1963). Exp. Cell Res. 31, 149-159. Karnovsky, M. J., Kleinfeld, A. M., Hoover, R. L., and Klausner, R. D. (1982). J . Cell Biol. 94, 1-16.

Katz, B. (1949). Arch. Sci. Physiol. 3, 285. Katz. B. (1966). “Nerve, Muscle and Synapse.” McGraw-Hill, New York. Kawakami, H., and Hirano, H. (1984). Hisrochemisrry 80, 415-420. Keller, R. E. (1976). Dev. Biol. 51, 118-137.

344

ARNOLD DE LOOF

Killion, J. J. (1984). Biophys. J. 45, 523-528. King, P. E., and Fordy, M. R. (1970). Z. Zellforsch. Mikrosk. Anat. 109, 158- 170. King, W. J., and Greene, G. L. (1984). Nature (London) 307, 745-747. Kinosita, H. (1963). Ann. N.Y. Acud. Sci. 100, 992-1004. Kirschner, L. B. (1973). In “Transport Mechanisms in Epithelia” (H. H. Ussing and N. A. Thorn, eds.), pp. 447-460. Munksgaard, Copenhagen. Klebanoff, S. J., and Clarck, R. A. (1978). “The Neutrophil: Function and Clinical Disorder.” North-Holland Pub]., Amsterdam. Kline, D., Robinson, K. R., and Nuccitelli, R. (1983). J . Cell Biol. 97, 1753-1761. Knochel, J. P. (1977). Kidney fnr. 11, 443-452. Koch, E. A., Smith, P. A., and King, R. C. (1967). J. Morphol. 121, 55-70. Korostoff, E. (1977). J . Biomech. Eng. 10, 41-44. Kristol, C., Sandri, C., and Akert, K. (1978). Brain Res. 142, 391-400. Kroeger, H. (1963a). J . Cell Comp. Physiol. 62 (Suppl. I), 45-49. Kroeger, H. (1963b). Nature (London) 200, 1234-1235. Kroeger, H. (1977). Mol. Cell. Endocrinol. 7, 105-110. Kropf, D. L., Lupa, M. D. A., Caldwell, J. H., and Harold, F. M. (1983). Science 220, 1385-1387. Kropf, D. L., Caldwell, J. H., Gow, N. A. R., and Harold, F. M. (1984). J . Cell Biol. 99, 486496.

Kusano, K., Miledi, R., and Stinnakre, J. (1982). J . Physiol. (London) 328, 143-170. Labadie, D. R. L., Hazlewood, C. F., Forster, J., and Cameron, I. L. (1983). Physiol. Chem. Phys. Med. NMR 15, 201-208. LaGamma, E. F., White, J. D., Adler, J. E., Krause, J. E., McKelvy, J. F., and Black, I. B. (1985). Proc. Narl. Acad. Sci. U.S.A. 82, 8252-8255. Lane, N. J., and Chandler, H. J. (1980). J . Cell Biol. 86, 765-774. Langendorf, H., Siebert, G., Lorenz, I., Hannover, R., and Beyer, R. (1961). Biochem. 2. 335, 273-284.

Langendorf, H., Siebert, G., Kesselring, K., and Hannover, R. (1966). Nature (London) 209, 11301131.

Langman, J. (1976). “Medical Embryology” (Dutch version). Bohn, Scheltema & Holkema, Utrecht. Laris, P. C., and Persadingh, H. A. (1974). Biochem. Biophys. Res. Commun. 57, 620-626. Lasalle, B. (1980). Dev. Biol. 75, 460-466. Latorre, R., and Miller, C. (1983). J . Membr. Biol. 71, 11-30, Latorre, R., Vergara, C., and Hildago, C. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 805-809. Lauger, P. (1984). Biochim. Biophys. Acta 779, 307-341. Lee, S. C., and Steinhardt, R. A. (1981). Dev. Biol. 85, 358-369. Le Goascogne, C., Sananbs, N., GouCzou, M., and Baulieu, E.-E. (1985). Dev. Biol. 109, 9-14. Leonard, F., and Wade, C. W. R. (1982). J. Bioelecrr. 1, 231-237. Lessmann, C. A., and Marshall, W. S. (1984). J. Exp. Zool. 231, 257-266. Levine, M., Garen, A., Lepesant, J. A., and Lepesant-Kejzlarova, J. (1981). Proc. Narl. Acad. Sci. U.S.A. 78, 2417-2421. Lewin, B. (1980). “Gene Expression: 2. Eukaryotic Chromosomes.” Wiley, New York. Lewis, S. A., Eaton. D. C., and Diamond, J. M. (1976). J . Membr. Biol. 28, 41-70. Lezzi, M. (1969). Physiol. Chem. Phys. 1, 447-461. Lezzi, M. (1970). Inr. Rev. Cyrol. 29, 127-168. Lezzi. M., and Gilbert, L. (1970). J. Cell Sci. 6, 615-628. Liboff, A. R., Williams, T., Jr., Strong, D. M., and Wistar, R., Jr. (1984). Science 223, 818-820. Ling, G. N. (1960). J. Gen. Physiol. 43, 149-174. Ling, G. N. (1969). Inr. Rev. Cyrol. 26, 1-61. Ling, G. N. (1981). Physiol. Chem. Phys. 13, 29-96.

THE ELECTRICAL DIMENSION OF CELLS

345

Ling, G. N. (1982). Physiol. Chem. Phys. 14, 47-96. Lin-Liu, S., and Adey, W. R. (1984). Biophys. J . 45, 121 1-1218. Lo, C. W., and Gilula, N. B. (1979). Cell 18, 411-422. Loewenstein, W. R. (1981). Physiol. Rev. 61, 829-913. Loewenstein, W. R., and Kanno, Y. (1963). J. Cen. Physiol. 46, 1123-1140. Lokha, M. J . , and Masui, Y. (1984). Dev. Biol. 103, 434-442. Lopez-Rivas, A., Adelberg, E. A., and Rozengurt, E. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 6275-6279. Lgvtrup, S. (1983). Biol. Rev. Cambridge Philos. SOC. 58, 91-130. Luben, R. A., Cain, C. D., Chen, M., Rosen, D. M., and Aden, W. R. (1982). Proc. Narl. Acad. Sci. U.S.A. 19, 4180-4184. Luther, P. W., Peng, H. B., and Jung-Ching Lin, J. (1983). Nature (London) 303, 61-64. Liittge, U., and Ball, E. (1979). J. Membr. Biol. 47, 401-422. Lymangrover, J. R., Keku, E., and Seto, Y. J. (1983). Life Sci. 32, 691-696. Lynn, J. W., and Chambers, E. L. (1984). Dev. Biol. 102, 98-109. McCloskey, M., and Poo, M.-M. (1984). Int. Rev. Cyrol. 87, 19-81. McCloskey, M., Liu, Z. Y . , and Poo, M.-M. (1984). J . Cell Biol. 99, 778-787. Mackenzie, D. (1982). New Sci. 93, 217-220. McLaughlin, S., and Poo, M.-M. (1981). Biophys. J . 34, 85-93. McLean, F. C., and Urist, M. R. (1968). “Bone: An Introduction to the Physiology of Skeletal Tissue,” 3rd Ed. Univ. of Chicago Press, Chicago. Maffeo, S., Miller, M. W., and Carstensen, E. L. (1984). J . Anar. 139, 613-618. Malinowska, D. H . , Koelz, H. R., Hersey, S . J., and Sachs, G. (1981). Proc. Narl. Acad. Sci. U.S.A. 78, 5908-5912. Maller, J. L. (1983). Adv. Cyclic Nucleotide Res. 15, 295-336. Maloff, B. L., Scordilis, S. S., and Reynolds, C. (1978a). J. Cell Biol. 78, 199-213. Maloff, B. L.. Scordilis, S. P., and Tedeschi, H. (1978b). J. Cell Biol. 78, 214-226. Maloney, P. C. (1982). J . Membr. Biol. 67, 1-12. Mannella, C A,, Colombini, M., and Frank, J. (1983). Proc. Nut/. Acad. Sci. U.S.A. 80, 22432247. Margolin, Y., and Eisenbach, M. (1984). J. Bacreriol. 159, 605-610. Marin, B. (1980). In “Plant Membrane Transport: Current Conceptual Issues” (R. M. Spanswick, W. J. Lucas, and J. Dainty, eds.), pp. 435-436. Elsevier, Amsterdam. Marino, A. A. (1984). J. Bioelecrr. 3, 235-244. Marino, A. A,, and Becker, R. 0. (1977). Physiol. Chem. Phys. 9, 131-147. Marino, A. A , , Hart,F. X.,and Reichmanis, M. (1983).IEEE Trans. Biomed. Eng. BME-30,833834. Marri3, E. (1979).In “Recent Advances in the Biochemistry of Cereals” (D. L. Laidman and R. G. WijnJones, eds.). pp. 3-25, Academic Press, New York. M a d , E., and Ballarin-Denti, A. (1985). J. Bioenerg. Biomembr. 17, 1-21. Marsh, G., and Beams, H. W. (1946). J. Cell. Comp. Physiol. 27, 139-157. Marshall, W. S., and Klyce, S. D. (1984). Biochim. Biophys. Acra 778, 139-143. Martin, P. M., and Sheridan, P. J. (1982). J. Steroid Biochem. 16, 215-229. Martirosov, S. M. (1983). Bioelecrrochem. Bioenerg. 10, 335-344. Martirosov, S. M . , and Trchounian, A. A. (1982). Bioelecrrochem. Bioenerg. 9, 459-467. Marx. J . L. (1984). Science 224, 271-274. Massa, E. M., Morero, R. D., Bloj, R., and Farias, R. N. (1975). Biochem. Biophys. Res. Commun. 66, 115-122. Masui, Y., and Clarke, H. J. (1979). Inr. Rev. Cyrol. 57, 185-282. Mates, S. M., Eisenberg, E. S., Mandel, L. J., Patel, L., Kaback, H. R., and Miller, M. H. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 6693-6697.

346

ARNOLDDELOOF

Mathog, D., Hochstrasser, M., Gruenbaum, Y., Saumweber, H., and Sedat, I. (1984). Nature (London) 308, 4 14-42 1. Matile, Ph. (1978). Annu. Rev. Planr Physiol. 29, 193-213. Mescher, A. L. (1976). J . Exp. Zoo/. 195, 117-128. Meyer, J. H., and Zak, H. H. (1982). Science 217, 635-637. Michel, H., and Oesterhelt, D. (1980). Biochemistry 19, 4607-4614. Miledi, R. (1960). J. Physiol. (London) 151, 1-23. Mitchell, P. (1961). Nature (London) 191, 144-148. Mitchell, P. (1966). Biol. Rev. Cambridge Philos. SOC. 41, 445-502. Mitchell, P. (1967). Fed. Proc., Fed. Am. SOC. Exp. Biol. 26, 1370-1379. Mitchell, P. (1968). “Chemiosmotic Coupling in Oxidative and Photosynthetic Phosphorylation.” Glynn Research, Bodmin, England. Mitchell, P. (1973). J. Bioenerg. 4, 63-91. Mitchell, P. (1977). Symp. SOC. Gen. Microbiol. 27, 483-523. Mitchell, P., and Moyle, J . (1969). Eur. J. Biochem. 7, 471-484. Miyamoto, H., and Racker, E. (1982). J. Membr. Biol. 66, 193-201. Miyazaki, S., and Igusa, Y. (1981a). Nature (London)290, 702-704. Miyazaki, S., and Igusa, Y. (1981b). In “The Mechanisms of Gated Calcium Transport Across Biological Membranes” (S. T. Onishi and M. Endo, eds.), pp. 305-31 1. Academic Press, New York. Miyazaki, S . , and Igusa, Y . (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 931-935. Miyazaki, S., Takahashi, K., and Tsuda, K. (1972). Science 176, 1441-1443. Moody, W. J., and Lansman, J. B. (1983). Proc. Narl. Acad. Sci. U.S.A. 80, 3096-3100. Moore, R. D. (1983). Biochim. Biophys. Acra 737, 1-49. Momll, G. A., Kostellow, A. B., and Weinstein, S. P. (1984a). Biochim. Biophys. Acra 803, 7177. Momll, G. A., Ziegler, D. H., Kunar, J., Weinstein, S. P., and Kostellow, A. B. (1984b). J . Membr. Biol. 77, 201-212. Moura, A. M., and Worcel, M. (1982). Clin. Sci. 63, 350-365. Moyer, M. P., Moyer, R. C., and Waite, M. R. F. (1982). J . Cell. Physiol. 113, 129-133. Mrsny, R. J., and Meizel, S. (1981). J. CellBiol. 91, 77-82. Muratsugu, M., Kamo, N., and Kobatake, Y. (1982). Bioelectrochem. Bioenerg. 9, 325-331. Murray. H. M., O’Brien, W. J., and Orgel. M. G. (1984). J. Bioelecrr. 3, 19-32. Naora, H.,Naora, H.,Izawa, M., Allfrey, V. G., and Mirsky, A. E. (1962). Proc. Natl. Acad. Sci. U.S.A. 48, 853-859. Nawata, T. (1984). Plant Cell Physiol. 25, 1089-1094. Neher, E., Sakmann, B., and Steinbach, J. H. (1978). Pfluegers Arch. 375, 219-228. Nicolini, C. (1983). Anticancer Res. 3, 63-86. Nuccitelli, R. (1978). Dev. Biol. 62, 13-33. Nuccitelli, R. (1980). Dev. Biol. 76, 499-504. Nuccitelli, R. (1983). Mod. Cell Biol. 2, 451-481. Nuccitelli, R. (1986). Prog. Clin. Biol. Res. 210. Nuccitelli, R., and Erickson, C. A. (1983). Exp. Cell Res. 147, 195-201. Nuccitelli, R., and Grey, R. D. (1984). Dev. Biol. 103, 1-17. Nuccitelli, R., and Jaffe, L. F. (1974). Proc. Natl. Acad. Sci. U.S.A. 71, 4855-4859. Nuccitelli, R., Poo, M.-M., and Jaffe, L. F. (1977). J. Gen. Physiol. 69, 743-763. Nuccitelli, R., Webb, D. J., Lagier, S. T., andMatson, G.B. (1982). Proc. Narl. Acad. Sci. U.S.A. 78,4421-4425. O’Brien, D. F. (1982). Science 218, 961-966. O’Brien, W. J.. Murray, H. M., and Orgel. M. G. (1984). J. Bioelecrr. 3, 33-40. O’Connor, C. M., Robinson, K. R., and Smith, L. D. (1977). Dev. Biol. 61, 28-40.

THE ELECTRICAL DIMENSION OF CELLS

347

Okamoto. H., Takahashi, K., and Yamashita, N. (1977). J. Physiol. (London) 267, 465-495. Okhuma, S., and Poole, B. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 3327-3331. Okhuma, S., Moriyama, Y., and Takano, T. (1982). Proc. Natl. Acad. Sci. U . S . A . 79,2758-2762. Orgel. M. G., O’Brien, W. J., and Murray, H. M. (1984). Plast. Reconsrr. Surg. 73, 173-182. Orida, N., and Feldmann, J. D. (1982). Cell Motil. 2, 243-255. Orida, N . , and Poo, M.-M. (1978). Nature (London) 275, 31-36. Orida, N., and Poo. M.-M. (1980). Exp. Cell Res. 130, 28 1-290. Ostenieder, W., Brum, G., Hescheler, J., and Trautwein, W. (1982). Nature (London) 298, 576578. Overall, R., and Jaffe, L. F. (1985). Dev. Biol. 108, 102-119. Paine, P. L., Pearson, T. W., Tluczek, L. J., and Horowitz, S. B. (1981). Nature (London) 291, 258-261. Papatheofanis, F. J., Papatheofanis, B. J . , and Day, R. D. (1984). J. Bioelectr. 3, 223-233. Patel, N.,and Poo, M.-M. (1979). J. Neurosci. 2, 483-496. Patel, N. B., and Poo, M.-M. (1984). J . Neurosci. 4, 2939-2947. Pederson, T. (1977). Biochemistry 16, 2771-2777. Peferoen, M., and De Loof, A. (1986). In preparation. Pena, A. (1975). Arch. Biochem. Biophys. 167, 397-409. Pena, A., Uribe, S., Pardo, J. P., and Borbolla, M. (1984). Arch. Biochem. Biophys. 231, 217225. Person, P. (1983). In “Nerve, Organ and Tissue Regeneration Research Perspectives” (F. Seil, ed.), pp. 407-429. Academic Press, New York. Peters, R. (1981). Cell Biol. Int. Rep. 5, 733-760. Petersen, 0. H. (1980). “The Electrophysiology of Gland Cells.” Academic Press, London. Petersen, 0. H., and Maruyama, Y.(1984). Nature (London) 307, 693-696. Pieri, C., Zs-Nagy, I., Zs-Nagy, V . , Giuli, C., and Bertoni-Freddari, C. (1977). J. Ulrrasrrucr. Res. 59, 320-331. Pisam, M., and Ripoche, P. (1976). J . Cell Biol. 71, 907-920. Poeting, G. A,, Koerwer, W., and Pongs, 0. (1982). Chromosoma 87, 89- 102. Polezhaez, L. V.. and Favorine, W. N. (1935). Wilhelm R o w ’ Arch. Entwicklungsmech. Org. 133, 701-727. Pollard, H. B., Shindo, N., Creutz, C. E., Pazoles, C. J., and Cohen, J. S. (1979). J. B i d . Chem. 254, 1170-1177. Poo, M.-M. (1981). Annu. Rev. Biophys. Bioeng. 10, 245-276. Poo, M.-M. (1985). Annu. Rev. Neurosci. 8, 369-406. Poo, M.-M., and Robinson, K. R. (1977). Narure (London) 265, 602-605. Poo, M.-M., Lam, J . W., Orida, N., and Chao, A. W; (1979). Biophys. J. 26, 1-21. Poole, B., and Okhuma, S. (1981). J. Cell B i d . 90, 665-669. Poole, R. J . (1978). Annu. Rev. PIanf Physiol. 29, 437-460. Poupon, R., and Evans, W. H. (1979). FEBS Letr. 108, 374-378. Pumplin, D. W., and Fambrough, D. M. (1983). J. Cell B i d . 97, 1214-1225. Purello, F., Vigneri, R., Clawson, G. A., and Goldfine, I. 1. (1982). Science 216, 100-1006. Purello, F., Burnham, D. B., and Goldfine, 1. D. (1983). Science 221, 463-464. Racker, E., Violand, B., O’Neal, S . , Alfonso, M., and Telford, J. (1979). Arch. Biochem. Biophvs. 198, 470-477. Raff, R. A , , and Kaufman, T. C. (1983). “Embryos, Genes and Evolution.” Macmillan, New York. Ramos, S., Schuldiner, S., and Kaback, H. R. (1976). Proc. Natl. Acad. Sci. U . S . A . 73, 18921896. Rasi-Caldogno, F.. De Michelis, M. I., and Pugliarello, M. C. (1982).Biochim. Biophys. Acta 693, 287-295.

348

ARNOLD DE LOOF

Raven, C. P. (1959). “An Outline of Developmental Physiology.” Pergamon, New York. Reggio, H., Bainton, D., Harms, E., Coudrier, E., and Louvard, D. (1984). J. CellEiol. 99, 151 11526. Reiss, H. D., and Herth, W. (1978). Protoplusmu 97, 373-377. Reissig, J. L. (1977). J . Cell Eiol. 75, 30a. Renthal, R., and Lanyi, J. K. (1976). Eiochemisrry 15, 2136-2143. Reuss, L., and Finn, A. L. (1974). J. Gen. Physiol. 64, 1-25. Richter, K. (1979). “Allgemeine Elektrophysiologie.” Fischer, Jena. Rick, R., Dorge, A,, Vonamim, E., Weigel, M., and Thurau, K. (1981). In “Epithelial Ion and Water Transport” (A. D. C. Macknqght and J. P. Leader, eds.), pp. 117-125. Raven, New York. Ridgway, E. B., and Ashley, C. C. (1967). Eiochem. Eiophys. Res. Commun. 29, 229-234. Ritchie, J. M., and Rogart, R. B. (1977). Proc. Natl. Acad. Sci. U.S.A. 74, 21 1-215. Robinson, J. D., and Flashner, M. S. (1979). Eiochim. Eiophys. Acta 549, 145-176. Robinson, K. R. (1977). J. Cell Eiol. 75, 25a. Robinson, K. R. (1979). Proc. Nurl. Acud. Sci. U.S.A. 76, 837-841. Robinson, K. R. (1983). Dev. Biol. 97, 203-211. Robinson, K. R. (1985). J. Cell Eiol. 101, 2023-2027. Robinson, K. R., and Cone, R. (1980). Science 207, 77-78. Robinson, K. R., and Stump, R. F. (1984). J . Physiol. (London) 352, 339-352. Robinson, S. I., Nelkin, B. D., and Vogelstein, B. (1982). Cell 28, 99-106. Robinson, S. P., and Downton, W. J. S. (1984). Arch. Eiochem. Eiophys. 228, 197-206. Rogart, R. B., Regan, L. J., Dziekan, L. C., and Galper, J. B. (1983). Proc. Nurl. Acud. Sci. U.S.A. 80, 1106-1 110. Rona, J.-P., Pitman, M. G., Liittge, U.,and Ball, E. (1980). J. Membr. Eiol. 57, 25-35. Rosenberg, R. L.. Tomiko, S. A., and Agnew, W. S. (1984). Proc. N d . Acud. Sci. U.S.A. 81, 5594-5598. Rosenbluth, J. (1976). J. Neurocytol. 5, 731-745. Rosoff, Ph. M., and Cantley, L. C. (1983). Proc. Nurl. Acad. Sci. U.S.A. 80, 7547-7550. Rothstein, A. (1976). In “Ion Transport and Metabolism” (A. Kotijk and A. Kleinzeller, eds.), pp. 270-284. Academic Press, New York. Rottenberg, H. (1984). J. Membr. Eiol. 81, 127-138. Rousseau, G. G. (1984). Eiochem. J . 224, 1-12. Rubin, R. P. (1982). “Calcium and Cellular Secretion.” Plenum, New York. Sabatini, D. D., Griepp, E. B., Rodriguez-Boulan, E. J., Dolan, W. I., Robbins, E. S., Papadopoulos, s., Ivanov, I. E., and Rindler, M. J. (1983). Mod. Cell Eiol. 2, 419-450. Schachter, H. (1984). Eiol. Cell. 51, 133-146. Schackmann, R. W., Christen, R., and Shapiro, B. M. (1984). J. Eiol. Chem. 259, 13914-13922. Scheffey, C., Foskett, J. K., and Machen, T. E. (1983). J. Membr. Eiol. 75, 193-203. Scherman, D., and Nordmann, J. J. (1982). Proc. Natl. Acud. Sci. U.S.A. 79, 476-479. Schlaepfer, W. W. (1974). Bruin Res. 69, 203-215. Schlichter, L. C. (1983a). Dev. Eiol. 98, 47-59. Schlichter, L. C. (l983b). Dev.Eiol. 98, 60-69. Schneider, D. L. (1981). J . Eiol. Chem. 256, 3858-3864. Schneider, D. L., Burnside, J., Gorga, F. R., and Nettleton, C. J. (1978).Eiochem. J . 176, 75-82. Schobert, B., and Lanyi, J. K. (1982). J . Eiol. Chem. 257, 10306-10310. Schrader. W. T. (1984). Narure (London) 308, 17-18. Schrijder, W. H., and Fain, G. L. (1984). Nature (London) 309,268-270. Schuldiner, S., and Rozengurt, E. (1982). Proc. Narl. Acud. Sci. U.S.A. 79, 7778-7782. Schwann, H. P. (1963). Eiophysik (Berlin) 1, 198-208. Schwartz, L. M., and Stiihmer, W. (1984). Science 225, 523-525.

THE ELECTRICAL DIMENSION OF CELLS

349

Schwarz. E. A. (1981). J. CellBiol. 90,271-278. Senior, A. E., and Wise, J. G. (1983). J. Membr. Biol. 73, 105-124. Serrano, R. (1977). Eur. J . Biochem. 80, 97-102. Serrano, R. (1980). Eur. J. Biochem. 105, 419-424. Serrano, R. (1983). Arch. Biochem. Biophys. 227, 1-8. Serrano. R . (1984). Curr. Top. Cell. Regul. 23, 87-126. Seto, Y. 1.. Hsieh, S. T., Majeau-Chargois, D., Dunlap, W. P., and Lymangrover, J. R. (1983). J . Bioelecrr. 2, 197-205. Shapiro, M. B. (1981). In “Fertilisation and Embryonic Development in Vitro” (L. Mastroianni, Jr. and I. D. Biggers, eds.), pp. 233-260. Plenum, New York. Shen, S. S. (1983). “Mechanisms of Animal Fertilisation,” pp. 213-267. Academic Press, New York. Sheppard, A. R., and Eisenbud, M. (1977). “Review of the Literature on Biological Effects of Electric and Magnetic Fields of Extremely Low Frequency.” New York Univ. Press, New York. Shteyer. A., Norton, L. A., and Rodan, G. A. (1980). J. Dent. Res. 59, 362 (Abstr. 380). Simkiss, K. (1980). Am. Zool. 20, 385-393. Singer, S. J. (1975). In “Cell Membrane Biochemistry, Cell Biology and Pathology” ( G . Weissmann, ed.), pp. 35-75. H. P. Publ. Co, New York. Singer, S. J., and Nicholson, G. L. (1972). Science 175, 720-731. Singh, A. P., and Bragg, P. D. (1979). Arch. Biochem. Biophys. 195, 74-80. Sisken. B. F., and Smith, S. D. (1975). J . Embryol. Exp. Morphol. 33, 29-41. Sisken, B. F., McLeod, B., and Pilla, A. A. (1984). J. Bioelecrr. 3, 81-101. Sjodin, R. A . (1982). J. Membr. Biol. 68, 161-178. Skaper, S. D., and Varon, S . ( I 983). Dev. Biol. 98, 257-264. Skou, J. C. (1960). Biochim. Biophvs. Acta 42, 6-23. Skulachev. V. P.. and Hinckle, P. C., eds. (1981). “Chemiosmotic Proton Circuits in Biological Membranes.” Addison-Wesley, Reading, Massachusetts. Slayman, C. L., and Slayman, C. W. (1962). Science 136, 876-877. Smith, G . K., and Cleary, S. F. (1983). Biochim. Biophys. Acra 763, 325-331. Smith. M. M.. Robinson, F. W., Watanabe, T., and Kono, T. (1984). Biochim. Biophys. Acra 775, 121- 128. Smith, R. L., and Nagel, D. (1983). Clin. Orrhop. Relaf. Res. 181, 277-282. Smith, S . D. (1974). Ann. N.Y. Acad. Sci. 238, 500-507. Smith, S . D., and Feola, J. M. (1982). J. Bioelecrr. 1, 207-229. Smith, S. D., and Mays. R. (1984). Bioelectrochem. Bioenerg. 12, 567-573. Solvay, E. (1894). “Du Rble de I’EIectricitt dans les Phtnomenes de la Vie Animale.” Hayez, Brussels. Somlyo, A. V., Gonzales-Serrates, S. H., Shumann, H., McClellan, G., and Somlyo, A. P. (1981). J. Cell Biol. 90, 577-594. Spadaro, J. A. (1982). J. Bioelecrr. 1, 99-128. Spanswick, R. M. (1981). Annu. Rev. Plant Physiol. 32, 267-284. Specht, S . C., and Sweadner, K. J. (1984). Proc. Nurl. Acad. Sci. U.S.A. 81, 1234-1238. Spemann, H. (1938). “Embryonic Development and Induction.” Yale Univ. Press, New Haven, Connecticut. Spenny, J. G.. Shoemaker, R. L., and Sachs, G. (1974). J. Membr. Biol. 19, 105-128. Spring. K. R., and Ericson. A. C. (1982). J. Membr. Biol. 69, 167-176. Steinberg, M. E., Labosky, D. A,, Jimenez, S., Lane, J. M., Korostoff, E., and Pollack, S. R. (1977). Trans. Orrhop. Res. Soc. 2, 285. Steinmetz, P. R., and Anderson, 0. S. (1982). J. Membr. Biol. 65, 155-174. Stem, C. D. (1981). Exp. Cell Res. 136, 343-350. Stem, C. D., and MacKenzie, D. 0. (1983). J. Embryol. Exp. Morphol. 77, 73-98.

350

ARNOLD DE LOOF

Stoeckenius, W., Lozier, R. M., and Bogomolni, R. A. (1979). Eiochim. Eiophys. Acfu 505, 215278. Strope, E. R., Findl, E., Conti, J. C., and Acuff, V. (1984). J. Eioelecfr. 3, 329-346. Stump, R. F., and Robinson, K. R. (1983). J. Cell Eiol. 97, 1226-1233. Stump, R. F., Robinson, K. R., Harold, R. L., and Harold, F. M. (1980). Proc. Null. Acud. Sci. U.S.A. 77, 6673-6677. Su, Y. X., Lim, S., and Edidin, M. (1984). Eiochim. Eiophys. Acfu 776, 92-96. Sugimoto, N., and Yasuda, T. (1983). Thymus 5, 297-310. Sutcliffe, J. G., Milner, R. J., Gottesfield. J. M., and Reynolds, W. (1984). Science 225, 13081315. Suzuki, K., and Fromter, E. (1977). Pfuegers Arch. 371, 109-1 17. Swann, J. W., and Carpenter, D. 0. (1975). Nature (London) 258, 751-754. Sze, H., and Churchill, K. A. (1981). Proc. Nut/. Acad. Sci. U.S.A. 78, 5578-5582. Szego, C. M., and Pietras, R. J. (1984). Int. Rev. Cytol. 88, 1-302. Taglietti, V., Tanzi, F., Romero, R., and Simoncini, L. (1984). J. CeN. Physiol. 121, 576-588. Tao-Cheng, J. H., and Rosenbluth, J. (1980). Bruin Res. 199, 249-265. Tedeschi, H. (1980). Eiol. Rev. Cambridge Philos. Suc. 55, 171-206. Telfer, W. H., Woodruff, R. I . , and Huebner, E. (1981). Am. Zoo/. 21, 675-686. Terepka, A. R., Coleman, J. R., Armbrecht, H. J., and Gunter, T. E. (1976). Symp. Soc. Exp. Eiol. 30, 117-140. Tetteroo, P. A. T., Bluemink, J. G . , Dictus, W. J. A. G., Van Zoelen, E. J. J., and De Laat, S. W. (1984). Dev. Eiol. 104, 210-218. Thoenen, H., and Edgar, D. (1982). Trends NeuroSci. (Pers. Ed.) Sept., 311-313. Thomson, S. H., and Aldrich, R. W. (1980). In “The Cell Surface and Neuronal Function” (C. W. Cotman, G. Poste, and G. L. Nicholson, eds.), pp. 49-85. Elsevier, Amsterdam. Tiedeman, H. (1981). Forfschr. Zoo/. 26, 121-131. Tluczek, L., Lau, Y. T., and Horowitz, S. B. (1984). Dev. Eiol. 104, 97-105. Tomkins, G., and Maxwell, E. S. (1963). Annu. Rev. Eiochem. 12, 677-708. Torrisi, M. R., and Pinto Da Silva, P. (1984). J. Cell Eiol. 98, 29-34. Towle, D. W. (1984). Am. Zuol. 24, 177-185. Trchounian, A. A. (1984). Bioelecfrochem. Eiuenerg. 13, 231-232. Tso, J., Thibier, C., Mulner, O., and Ozon, R. (1982). Proc. Nurl. Acud. Sci. U.S.A. 79, 55525556. Tupper, I., and Tedeschi, H. (1969). Science 166, 1539-1540. Turner, R. J . (1983). J. Membr. B i d . 76, 1-15. Ussing, H. H. (1964). Harvey Lecr. 59, 1-30. Vanable, J. W., Hearson, L. L., and McGinnis, M. E. (1983). frog. Clin. Biol. Res. llO(A), 587596. Verachtert, B., and De Loof, A. (1984). Inf. Congr. Enfumol., 17fh. Hamburg Absfr. Vol. p. 161. Verachtert, B., and De Loof, A. (1986). In “Ionic Currents in Development” (R. Muccitelli, ed.), pp. 173-179. Liss, New York. Vethamany-Globus, S . , Globus, M., Darch, A., Milton, G., and Tomlinson, B. L. (1984). J. Exp. 2001.231, 63-74. Vinkler, C., and Korenstein, R. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 3183-3187. Virk, S. S., Kirk, C. J., and Shears, S. B. (1985). Eiochem. J . 226, 741-748. Wahl, G. M., de Saint Vincent, B., and De Rose, M. L. (1984). Nufure (London) 307, 516-520. Walsh-Reitz, M. M., Toback, F. G . , and Holley, R. W. (1984). Proc. Narl. Acad. Sci. U.S.A. 81, 793-796. Warncke, U. (1979). I n “Electromagnetic Bio-Information” (F. Popp, G. Becker, H. Konig, and W. Peschka, eds.), pp. 55-79. Urban and Schwarzenberg, Munchen.

THE ELECTRICAL DIMENSION OF CELLS

35 1

Warner, A. E., Guthrie, S. C., and Gilula, N. B. (1984). Narure (London) 311, 127-131. Waxman, S. G., and Ritchie, I. M . (1985). Science 228, 1502-1507. Webb, D. J. (1984). Bioelecfrochem. Bioenerg. 13, 429-438. Webb, D. J., and Nuccitelli, R. (1982). Kroc. Found. Ser. 15, 293-324. Webb, D. J., and Nuccitelli, R. (1985a). Dev. Biol. 107, 395-406. Webb, D. J., and Nuccitelli, R. (1985b). Comp. Biochem. Physiol. 82A, 35-42. Webb, W. W., Barak, L. S., Tank, D. W., and Wu, E. S. (1982). Biochem. Soc. Symp. 46, 191205. Weigele, J. B., and Barchi, R. L. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 3651-3655. Weinstein, S . P., Morrill, G. A., and Kostellow, A. B. (1982). Biochem. Biophys. Rex Commun. 108, 876-880. Weinstein, S. P., Morrill, G. A., and Kostellow, A. B. (1983). Dev. Growth Difler. 25, 11-21. Weisenseel, M. H. (1982). In “Biophysik” (W. Hoppe, W. Lohmann, H. Markl, and H. Ziegler, eds.), 2nd Ed., pp. 475-480. Springer-Verlag, Berlin and New York. Weisenseel, M. H., and Kicherer, R. M. (1982). Cell Biol. Monogr. 8, 379-400. Weisenseel, M. H., Nuccitelli, R., and Jaffe, L. F. (1975). J. Cell Biol. 66, 556-567. Weisenseel, M. H.. Dom, A,, and Jaffe, L. F. (1979). Plant Physiol64, 512-518. Weiss, A. B., Parsons, J. R., and Alexander, H. (1980). J. Med. Soc. N.J. 77, 523-526. Welshons, W. V., Lieberman, M. E., and Gorski, J. (1984). Nature (London) 307, 747-749. Wendler, S., Zimmermann, U., and Bentrup, F.-W. (1983). J . Membr. Biol. 72, 75-84. Went, D. F. (1977). Dev. Biol. 55, 392-396. West, 1. C. (1980). Biochim. Biophys. Acfa 604, 91-126. Whitaker, M. J., and Steinhardt, R. A. (1982). Q. Rev. Biophys. 15, 593-666. Williams, W. S., and Breger, L. (1975). J . Biomech. 8, 407-413. Willier, B. H., and Oppenheimer, J. M.(1974). “Foundations of Experimental Embryology,” 2nd Ed. Hafner, New York. Winegrad, S . (1968). J. Gen. Physiol. 51, 65-83. Wittbjer, J., Glantz, P. O., Rohlin, M., and Thomgren, K. G. (1984). Acra Odontol. Scund. 42, 141-151. Wolniak, S . M., Helper, P. K., and Jackson, W. T. (1983). J. Cell Biol. 96, 598-605. Woodruff, R. I., and Telfer, W. H. (1973). J. Cell Biol. 58, 172-188. Woodruff, R. I . , and Telfer, W. H. (1974). Ann. N.Y. Acad. Sci. 238, 408-419. Woodruff, R. I., and Telfer, W. H. (1980). Nature (London) 286, 84-86. Woodruff, R. I . , Huebner, E., and Telfer, W. H. (1984). In “Advances in Invertebrate Reproduction” (W. Engels, ed.), p. 652. Elsevier, Amsterdam. Workshop on Vibrating Probe Technique, R. Nuccitelli, organizer (1985). See Nuccitelli, R. (1986). Wu, C . H. (1984). Am. Sci. 72, 598-607. Yamada, T. (1962). J. Cell Comp. Physiol. 60 (Suppl. I ) , 49-64. Yamada, T. (1981). Nerh. J. Zool. 31, 78-98. Yamamoto, T. (1962). Embryologia 7, 228-251. Yamashita, 0.. and hie, K. (1980). Nature (LondonJ 283, 385-386. Yasuda, I. (1974). Ann. N.Y. Acad. Sci. 238, 457-465. Yau, K. W., and Nakatani, K. (1984a). Narure (London) 309, 352-354. Yau, K. W., and Nakatani, K. (1984b). Nature (London) 311, 661-663. Yoshida, S. (1983). J. Physiol. (London) 339, 631-642. Zagyanski, Y. A,, and Jard, S. (1979). Nature (London) 280, 591-593. Zaritsky, A.. Kinara, M.. and Macnab, R. M. (1981). J. Membr. Biol. 63, 215-231. Zeuthen, T. (1977). J . Membr. Biol. 33, 281-309. Zeuthen, T.(1978). J . Membr. Biol. 39, 185-218. Zeuthen, T., and Monge, C. (1975). Philos. Trans. R. Soc. London Ser. B . 271, 277-281.

352

ARNOLD DE LOOF

Zhang, F., and Schneider, D. L. (1983). Biochem. Biophys. Res. Commun. 114, 620-626. Zilberstein, D.,Schuldiner. S., and Padan, E. (1979). Biochemisrry 18, 669-673. Zimmerman, R. L. (1982). J . Bioelectr. 1, 265-287. Zimmerman, U.,and Vienken, J . (1982). J. Membr. Biol. 67, 165-182. Ziomek, C. A , , Schulman, S., and Edidin, M. (1980). J . Cell B i d . 86, 849-857. Zs-Nagy, I . , Lustyik, G . , Zs-Nagy, V., Zarandi, B . , and Bertoni-Freddari, C. (1981). J . CellBiol. 90, 769-777.

Index

A Acanthamoeba. actin-binding proteins of,

160-

I62 Acetylene reductase, of Rhizobium species, 2 Actin binding of connectin, 98-99 extraction of, 84 hyphal microfilaments and, 52 lens cell development and, 3I of protists. actin-binding proteins, 160- 163 actin-containing structures, 163-172 properties of, 154-160 role in absence of myosin division, 183-I84 ingestion, 181-183 locomotion, 184- I85 positioning, 183 structures containing arrangement of microfilaments in cortical regions, 163- 166 endoplasmic microfilaments, 166- 169 presence of actin without evident microfilaments, 169- 172 Actin-binding proteins, roles of, 160- 163 a-Actinin protistan, 161, 162 solubilization of, 86 P-Actinin, protistan, 161,162 Actin-myosin interaction, in protists, 176-181 Active transport, maintaining inequality of distribution of diffusible ions ion channels: types and gating mechanisms,

262-266 ion pumps, 261-262 transmembrane difference of electrical potential for protons. 266

Adenosine triphosphatase, microtubule-associated protein and, 207,209,214,215 Adenylate cyclase, FSH and, 130- I32 Agrobacreriurn, plasmids, relationship to Rhizobium plasmids, 16-I7 Amoebas, actin, properties of. 154-160 Amino acid composition, of connectin, 90-91 plasmodia1, 1I0 Amino acid sequences of actins, 154-159 of protistan tubulins, 199-201 Androgens, in seminiferous epithelium, 132-

134 Antibiotic sensitivity, of Rhizobium species, 2 Apex, hyphal, cell wall structure at, 63-65 Apical growth cytoplasmic components of cytoskeleton, 50-53 plasma membrane, 53-56 vesicles, 46-50 of hyphae, biophysical models of, 42-44 phenomenon of, 38-40 Ascomycotina, cell wall polymers of, 57 Association-induction hypothesis, of resting membrane potential, 259-260 Autolysins, apical wall growth and, 65-68

B Bacteria, electrical phenomena in, 325-327 Basidiomycotina, cell wall polymers of, 57 Benzimidazole-2-yl carbamate, fungal microtubules and, 51 Bioelectricity, historical background, 25I Body form, genesis, ionic currents and, 298-

301

353

354

INDEX

Bone, regeneration of, 320 Bursting, of hyphal tips, 66-67 C

Calcium channels, 264-265 Calcium ions actin-myosin interaction in protists and, 179-180 egg activation and, 317 excitable cells and, 308 gradients, apical growth and, 46 Calmodulin, in protists. 182 Cancer, intracellular ionic conditions and, 306-307 Capacitance, plasma membrane and, 255 Carbohydrates, of plasma membrane, 254 Cardiac muscle, native connectin from, 108109 Cataract animal models and, 25-27 congenital, prevalence of, 25 Cataractous phenotype, manipulations of, 32 Cell(s) excitable, ionic currents and, 307-312 ionic and electrical compartmentalization within electrophysiological survey, 275-276 mitochondria, 277-278 nucleus, 278-280 sarcoplasmic reticulum, 278 vacuoles and lysosomes, 276-277 unifacial, bifacial, and multifacial fluidity of plasma membrane, 266-267 mechanism to establish and maintain membrane protein segregation, 274275 restricted lateral motion of some integral membrane proteins, 267-268 segregation of ion pumps and ion channels in different cells, 268-270 segregation of sites of inward and outward current, 273-274 voltage and ionic gradients within, 281286 Cell division actin and, 183-184 role of microtubular systems in, 215, 217 Cell shape, 238 epiplasm and, 230-231 microtubular systems and, 209, 212

Cellular activities, effects of imposed electric fields on, 328-333 Cell wall fungal, structure of apical structure, 63-65 hyphal wall, 61-63 wall polymers, 56-60 Chitin in fungal cell walls, 57-58 in hyphal apices, 64, 70-71 in hyphal walls, 63 sites of synthesis, 53-56 Chitin synthase. activation of, 54-55 Chitosan, in fungal cell walls, 58-59 Chitosomes, composition and localization, 50, 54 Chloride channels, 263 Chromatoid body, function of, 127-129 Chromosomes associated with mutant cataracts, 27 location, effect of ions on gene expression and, 287-288 Chymotrypsin, effect on connectin, 95 Cilia microtubules and, 186- I87 microtubule-associated proteins and, 207209 Circular dichroism, of connectin, 92-93 Cloning vectors, functional analysis of Rhizobium plasmids and, 7-8 Cointegrates, of Rhizobium plasmids, 9 Concentration cell, principle of application to living cells, 258-259 origin of resting membrane potential and, 256-258 Connectin, 180 comparative biochemistry general survey, 107- 108 native connectin from cardiac muscle, 108- 109 Physarum plasmodia1 protein, 109- I10 as elastic components, 103-105 interaction with myosin and actin, 96-99 location in myofibrils electron microscopy, I0 1- I03 immunofluorescence studies, 99- 101 native other properties, 90-93 preparation, 86-87 proteolysis, 93-96 size and shape, 87-90

355

INDEX occurrence of. 81 transformation during differentiation during chick development, 105- 107 during myofibrillogenesis, 107 versus titin, 83-86 C-protein, connectin and, 98 Curing, of Rhizobium plasmids, 9 Current, inward and outward, segregation of sites of, 273-274 Cycloheximide, chitin synthase and, 55-56 Cytochalasins, fungal microfilaments and, 5253 Cytoplasm, polarized transport in, 294-296 Cytoskeletal organization, polarized, maintenance in apex by ion gradients, 45-46 Cytoskeleton apical wall growth and, 50-53 functions of, 153

D Deletion, of plasmids in Rhizobium. 9 Dictyosteliurn, actin-binding proteins of, 160, 161, 162 Differentiation connectin transformation and, 105- 107 generation of asymmetry as key to, 288-293 of spermatocytes, plasminogen activators during, 139-142 Digestive vacuoles, actin and, 181-182 Division ring, of protists, 234-236 Donnan equilibrium system, maintaining inequality of distribution of diffusible ions, 260-26 1

E Eggs, developing, voltage differences within, 283 Electrical currents, apical growth and, 44-46 Electrical potential. gradients within cells, 281-286 Electric organs, ionic currents and, 309-310 Electron microscopy of connectin, 87-90 in muscle, 101-103 Electrophoresis, in plane of membrane, 296298 Electrophysiological survey, of compartmentalization within cells, 275-276

Endocrinology, electrophysiology and, 321325 Endoplasm, of protists, microfilaments in, 166-169 Enzymes apical vesicles and, 49-50 autolytic localization in hyphae, 66 role of, 68 of Sertoli cells, cyclic activity of, 129I30 Epiplasm, of protists, 23 1-232 nature, 230 role, 230-231 structure, 228-230 Epithelia, intracellular potential gradients in, 28 1 Epithelial cells, segregation of ion pumps and ion channels in in differentiated organisms, 268-269 in embryos and regenerating tissues, 269270 EvoIu tion of tubulins, 206

F Fertilization, ionic currents and, 3 14-3 16 Filaments, intermediate, of protists, 224-228 Flagella microtubules and, 186- 187 microtubule-associated proteins and, 207209 Flagellation, of Rhizobium species, 2 Follicle-stimulating hormone local action on Sertoli cells, 130-132 secretion of plasminogen activators and, 141 Fragmin, 180 properties of, 160, 162 Fungi cell wall growth in, 37-38 cell wall structure apical, 63-65 hyphal, 61-63 wall polymers, 56-60

G Galvanotaxis, electric current and, 329-330 Gap junctions, ion fluxes and, 287

356

INDEX

Gating mechanisms, for diffusible ions, 265266 Gelactins, properties of, 162 Gel electrophoresis, of connectin, 87 Gene(s), active, configuration of, 304-305 Gene fusion, functional analysis of Rhizobium plasmids and, 6-7 Genome, of Rhizobium, size of, 3 P-Glucans, in fungal cell walls, 57, 58 ( I+3)-P-Glucans in fungal cell walls, 57, 59, 60 in hyphal walls, 61-63, 70 synthesis in hyphae, 56 ( I +6)-P-Glucans in fungal cell walls, 57, 59-60 in hyphal walls, 61-63, 70, 71 Golgi region, compartmentalization in, 276, 277 Growth factors, testicular, I38

H Hepatocytes, segregation of ion pumps and ion channels in, 272 Host range, specificity in Rhizobium, 10 Hydrogan uptake, genes, in Rhizobium, 15 Hyphae observations in vivo biophysical models of apical growth, 4244 electrical currents and ion gradients, 4446 phenomenon of apical growth, 38-40 turgor and wall expansion, 4 1-42 potential gradients within, 282 Hyphal apex, wall polymer assembly at assembly in wall: a steady-state model, 6872 autolysins and apical wall growth, 65-68 Hyphal walls, structure of, 61-63

I Immunochemistry, of protistan tubulins, 201202. 203-204, 205-206 lmmunofluorescence study, of connectin, 99101

Ingestion, role of actin in, 181-183 Insulin, ion transport and, 322

Ions diffusible, mechanisms for maintaining inequality of distribution of active transport, 261 -266 Donnan equilibrium systems, 260-261 Ion channels, types, 262-263 calcium, 264-265 chloride, 263 gating mechanisms, 265-266 larger cations, 265 potassium, 264 sodium, 263 Ion gradients, apical growth and, 44-46 Ionic currents selected functions of causal relationship to genesis of body forms, 298-301 electrical phenomena in bacteria, yeasts and mitochondria, 325-328 electrical phenomena in vertebrate regenerative growth and wound healing, 317-321 electrical properties of oocytes during growth and eggs at fertilization, 312-31 7 electrophysiology, endocrinology and osmoregulation, 321-325 excitable cells, 307-312 ions and nuclear activity, 301-307 transcellular ionic currents: signals and effectors of polarity, 294-298 Ionic theory, of origin of resting membrane potential application of principle to living cells, 258259 intracellular perfusion experiments on squid axons, 255-256 principle of the concentration cell, 256-258 Ion pumps, maintaining inequality of distribution of diffusible ions, 261-262 Ion pumps and ion channels, segregation in different cell types in epithelial cells of differentiated organisms, 268-269 in epithelial cells in embryos and regenerating tissue, 269-270 in hepatwytes, 272 in neurons, 270 in skeletal muscle cells, 270-272 in vertebrate photoreceptors. 272

357

INDEX

K Kinetodesmal fibers, of ciliates, 223-224 Kinetosomes connections between, 222 microtubules and, 186 periodic rootlets of, 2 17-222

L Large cation channels, 265 Lens crystallin electrophoretic phenotypes, 26 synthesis of, 27-30 Lens epithelial cells, cellular studies on, 3031 Leydig cells, interactions with seminiferous tubules, 1 4 4 1 4 6 Lipid chitin synthase and, 54-55 of plasma membranes, 252, 254 Locomotion, actin and, 184- I85 Lysosomes, compartmentalization and, 276277

M Mastigomycotina, cell wall polymers of, 57 Meiosis, in spermatocytes, 119- 120 Meiosis-inducing factor, cyclic secretion of, 136-137 Melanosomes, intracellular potential gradients and, 283 Membrane potential, influence on activity of wall synthetic enzymes, 45 Membrane proteins, integral mechanisms to establish and maintain segregation of, 274 restricted lateral diffusion of, 267-268 Microfibrils, of chitin in fungal cell walls, 58 in hyphal apices, 64, 71 Microfilaments arrangement in cortical regions of protists, 163-166 formation by protist actin, 159-160 in fungal apical hyphae. 52 Microtubules arrangements and MTOCs in protists, I97 kinetosome-associated intracellular microtubules, 187- 193

kinetosomes, flagella and cilia, 186- 187 microtubules unrelated to kinetosomes, 193-197 in hyphal apex, 50-52 kinetosome-associated, 187- 193 unrelated to kinetosomes, 193-197 Microtubule-associated proteins, 206 of cilia and flagella, 207-209 of other microtubular systems, 209 roles ascribed to microtubular systems, 209217 of structures other than axonemes of cilia and flagella, 207 Microtubule-microfilament connections, lens cell development and, 31 Mitochondria compartmentalization and, 277-278 electrical phenomena in, 327-328 Mitogens, ionic effects of, 324-325 Mitosis asymmetrical, differentiation and, 292-293 intracellular ionic conditions and, 306-307 Molecular constitution, of protistan myosin, 172 Motility, role of microtubular systems and, 2 14-2 15 Mouse. congenital cataract mutants and, 2527 Muscle ionic currents in, 308-309 microfibrillar proteins of, 82 third filament in. 81-83 Myofibrils, location of connectin in electron microscopy, 101-103 immunofluorescence study, 99- 101 Myonemes, nature of, 232, 234 Myosin binding of connectin, 96-98, 104 extraction of, 84 polymers, as molecular weight markers, 87 protistan characteristics of, 172- 175 structures, 175- 176 Myxomycetes, actin, properties of, 154- 160

N Na , K +-Adenosine triphosphatase localization of, 268, 272 membrane potential and, 261-262 +

358

INDEX

Neurons, segregation of ion pumps and ion channels in, 270 Nitrogen fixation genes, in Rhizobium genetic evidence for, 9- I 1 physical evidence for, 12 Nodulation, plasmid genes and, 10 Nodulation genes, identification and isolation of, 12-14 Nonactin microfilaments, of protists contractile, 232-236 noncontractile, 236-238 Nuclear activity, ions and, 301-307, 309 Nucleic acids of Rhizobium species, guanine and cytosine content, 2 of Rhizobium plasmids, hybridization of, 34 Nucleus, compartmentalization and, 278-280

0 Ohm’s law, in biological systems, 255 Oocytes, electrical properties activation, 317 excitability of membranes, 312-313 fertilization, 3 14-3 I6 maturation, 313-314 Organelles, guiding and positioning of, role of microtubular systems in, 215 Osmoregulation, hormones and, 322-323 Ovarian follicle, electrophysiological measurements on. 282-286

P Peptide maps, of tubulins, 199-206 Phage, functional analysis of Rhizobium plasmids and, 8 6-Phosphogluconate dehydrogenase, of Rhizobium species, 2 Photoreceptors ionic currents and, 310-312 vertebrate, segregation of ion pumps and ion channels in, 272 Physarum, actin-binding protein of, 160- 162 Physarum polycephalum, native connectin from, 109-110 Piezoelectricity, in biological materials, 255 Plasma membrane apical wall growth and, 53-56

constituents of, 252-255 fluidity of, 266-267 polarization, apical growth and, 45 Plasmids of Rhizobium detection, 2-3 nucleic acid hybridization, 3-4 relationship to Agrobacterium plasmids, 16-17 restriction endonuclease maps of, 17- 18 strategies and genetic methods for functional analysis in Rhizobium cloning vectors and use of phage, 7-8 plasmid deletion and curing, 9 plasmid transfer and R-primes, 8-9 transposons, gene fusions and marker exchange, 4-7 Plasmid RK2, derivatives, as cloning vectors, 7 Plasminogen activators, localization and function in seminiferous epithelium, 138 cellular and hormonal regulation of secretion, 139 during postnatal and in virro differentiation, 139- I42 Polarity, transcellular ionic currents as signals and effectors of electrophoresis in plane of membrane, 296298 polarized tranport in cytoplasm, 294-296 Polygalacturonase, in Rhizobium, 10 Polymer(s), of fungal cell walls, 56-60 Polymerization, of protistan myosin, 172, 175 Polysaccharides, acidic, in fungal cell walls,

60 Polysaccharide synthesis, genes, in Rhizobium. 14-15 Positioning, actin and, 183 Potassium channels, 264 Profilin, properties of, 160, 161 Projectin, 83 amino acid composition, 91 Proteins cyclicly secreted by seminiferous epithelium, 134-136 of plasma membranes, 252-253, 254 of spermatid nuclei, 121 Protein carboxymethylase, sperm and, 130 Proteolysis, of connectin effects of proteases, 95-96 spontaneous breakdown, 93-95

359

INDEX Protists actin of actin-binding proteins, 160-163 actin-containing structures, 163- 172 properties of, 154-160 actin-myosin interaction in, 176- 18 I epiplasm of, 231-232 nature, 230 role, 230-23 I structure, 228-230 filament systems of unknown nature in, 238-240 myosin of characteristics, 172- I75 structures, 175- 176 nonactin microfilaments of contractile division ring, 234-236 doubtful cases, 236 myonemes, 232-234 spasmonemes, 234 well-known cases, 232 intermediate filaments of, 224-228 nonactin microfilaments, noncontractile doubtful cases, 238 well-known cases, 236-238 periodic fibers of connections between kinetosomes, 222 kinetodesmal fibers of ciliates, 222-224 kinetosomal periodic rootlets, 217-222 tubulins and microtubules of, 185-186 arrangements and MTOCs, 186- 197 associated proteins and roles of structures, 206-2 17 tubulins, 197-206 Protons influx, growing hyphal tips and, 45 transmembrane difference of electrical potential for, 266 Pseudomicrothorax, actin-binding protein of, 161, 162

R Receptors, microtubular systems and, 217 Regenerative growth, electrical phenomena and, 317-321, 328-329 Resting membrane potential, theories of origin of association-induction hypothesis, 259-260 ionic theory, 255-259

Restriction endonuclease maps. of Rhizobium plasmids, 17-18 Retinoic acid, plasminogen activator secretion and, 144 Rhizobium functions controlled by plasmid genes expression of Sym plasmid in Rhizobiaceae, 11 genetic evidence for nitrogen fixation genes, 9-11 identification and isolation of nodulation genes, 12-14 other genes involved in symbiotic nitrogen fixation, 14-15 physical evidence for nitrogen fixation genes, 12 phenotypic characteristics of, 2 physical studies on plasmids detection, 2-3 nucleic acid hybridization, 3-4 plasmid-genome rearrangements in, 16 relationship of plasmids to Agrobucrerium plasmids, 16-17 restriction endonuclease maps of plasmids, 17-18 strategies and genetic methods for functional analysis of plasmids cloning vectors and use of phage, 7-8 plasmid deletion and curing, 9 plasmid transfer and R-primes, 8-9 transposons, gene fusions and marker exchange, 4-7 Ribonucleic acid, synthesis during spermatogenesis, 125- 126 R-primes, Rhizobium plasmids and, 9

S Succhuromyces commune, hyphal wall assembly in, 69-72 Salt tolerance, of Rhizobium species, 2 Sarcoplasmic reticulum, compartmentalization and, 278 Sedimentation coefficient, of connectin, 87 Seminiferous epithelium cycle, wave and transillumination of, 121125 localization and function of plasminogen activators in, 138 cellular and hormonal regulation of secretion, 139

360

INDEX

during postnatal and in vitro differentiation, 139-142 Seminiferous tubules, interactions with Leydig cells, 144-146 Sertoli cells cyclic interaction with spermatogenic cells, 129-130 androgens in seminiferous epithelium, 132-134 cyclicly secreted proteins, 134-136 local action of follicle-stimulating hormone, 130-132 other cyclicly secreted factors, 136- 137 testicular growth factors, 138 spermatocyte meiosis and, I 19-120 spermatogonia and, 119 Severin, properties of, 160, 161 Skeletal muscle cells, segregation of ion pumps and ion channels in, 270-272 Sodium channels, 263-264 Somatomedin C, in testis, 138 Spasmonemes, nature of, 234 Spectrin, epiplasmic protein and, 231 Spermiogenesis, stages of, 120-121 Spermatogenesis transcriptional activity during, 125-126 in vitro cultures of defined segments of seminiferous tubules, 142- 143 flow cytometric evaluation of cultured seminiferous tubule segments, 143144 Spermatogenic cells cyclic interaction with Sertoli cells, 129I30 androgens in seminiferous epithelium, 132- 134 cyclicly secreted proteins, 134- 136 local action of follicle-stimulating hormone, 130-132 other cyclicly secreted factors, 136- 137 testicular growth factors, 138 metabolism of meiosis, 119-120 spermatogonial development, 117-1 19 spermiogenesis, 120- 12 1 Spermatogonia, development of, 117-1 19 Spitzenkorper. nature of, 47 Squid axons, intracellular perfusion experiments, 255-256

Steady-state model, of hyphal wall assembly, 68-72 Steriod hormones, ionic effects and, 302, 303, 321-322, 323-324 Substratum, lens cell development and, 32 Supporting structure, role of microtubular systems and, 212-214 Symbiotic functions, plasmids coding for in Rhizobiurn, transfer of, 8-9 Sym plasmid, expression in Rhizobiaceae, 11

T Testosterone, Leydig cells and, 144-145 Titin, identification of, 85 Transfemn-like protein secretion by Sertoli cells, 137 Transmembrane potential and transcellular currents generation of few inorganic ion species: an extremely versatile system, 286-293 ionic and electrical compartmentalization within cells, 275-280 mechanism for maintenance of inequality of distribution of diffusible ions, 260-266 Ohm’s law, capacitance and piezo electricity, 255 plasma membrane constituents, 252-255 theories on origin of resting membrane potential, 255-260 unifacial, bifacial, and multifacial cells, 266-275 voltage and ionic gradients within cells: transcytoplasmic currents, 281-286 Transposons, functional analysis of Rhizobium plasmids and, 5-6 Trypsin, effect on connectin, 95, 105 Tubulins interspecific variability among protists, 203-205 between protists and metazoa, 205-206 of protists, 197-199, 206 differences between a- and P-tubulins, I99 interspecific variability, 203-206 intraspecific microheterogeneity, 199-202 Turgor, hyphal wall expansion and, 41-42

361

INDEX

V Vacuoles, compartmentalization and, 276 Vesicles cytoplasmic, apical wall growth and, 46-50 self-electrophoresis toward growing hyphal wall area, 45 Vibrating probe technique, extracellular ionic currents and, 273 Viscosity, of connectin, 88

Wound healing, electrical phenomena and, 3 17-321

x X-irradiation, spermatogonia resistant to, I 19

Y Yeast, electrical phenomena in, 327

W Wall materials addition, hyphal growth and, 43, 44 apical vesicles and, 49

Z Zygomycotina, cell wall polymers of, 57, 60

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