Advances in Botanical Research, Vol. 12

Advances in

BOTANICAL RESEARCH VOLUME 12

Advances in

BOTANICAL RESEARCH Editor-in-Chief J. A. CALLOW

Department of Plant Biology, University of Birmingham, Birminghum, England

Editorial Board H. W. WOOLHOUSE W. D. P. STEWART

E. G. CUTTER W. G. CHALONER

E. A. C. MAcROBBIE

John Innes institute, Norwich, England Department of Biological Sciences, The University, Dundee, Scotland Department of Botany, University of Manchester, Manchester, England Department of Botany, Royal Holloway & Bedford New College, University of London, Egham Hill, Egham, Surrey, England Department of Botany, University of Cambridge, Cambridge, England

Advances in

BOTANICAL RESEARCH Edited by

J. A. CALLOW Depurtmrnt of Plant Biology University of Birmingham Birmingham, Englund

VOLUME 12

I986

ACADEMIC PRESS Harcourl Brace Jovanovich, Publishers

London Orlando San Diego New York Austin Montreal Sydney Tokyo Toronto

PRESS INC. (LONDON) LTD COPYRIGHT 0 1986 BY ACADEMIC ALL RIGHTS RESERVED. NO PART OF THIS PUBLICATION MAY BE REPRODUCED OR TRANSMITTED IN ANY FORM OR BY ANY MEANS, ELECTRONIC OR MECHANICAL, INCLUDING PHOTOCOPY, RECORDING, OR ANY INFORMATION STORAGE AND RETRIEVAL SYSTEM, WITHOUT PERMISSION IN WRITING FROM THE PUBLISHER.

ACADEMIC PRESS INC. (LONDON) LTD. 24-28 Oval Road

LONDON NWI 7DX

Unired Srates Edition published by

ACADEMIC PRESS. INC. Orlando. Florida 32887

LIBRARY OF CONGRESS CATALOG CARD NUMBER:62-21 144 ISBN 0-12-005912-6 PRINTED INTHEUNITED STATESOFAMERICA

86878889

9 8 1 6 5 4 3 2 1

CONTENTS CONTRIBUTORS TO VOLUME 12 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . PREFACE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

vii

ix

Light/Dark Modulation of Enzyme Activity in Plants LOUISE E . ANDERSON I . Occurrence of Light Modulation . . . I I . Proposed Mechanisms .............................. 111. Regulation of Light Modulation ...................... IV . Dark Modulation ................................... V . Changes in the Target Enzyme ....................... VI . The Function of Light Modulation . . . . . . . VII . Effect of Sulfur Dioxide on Light Mo VIII . Osmotic Stress and Light Modulation IX . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . ............................

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

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

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

3 6 24 28 30 36 38 38 39 39

Algal Toxins WAYNE W . CARMICHAEL .. . . . . . . . . . . . . . . . . . I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . .................. I1 . Naming Algal Toxins ............................... 111. Occurrence. Growth. and Toxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV . Isolation Characterization and Toxinology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V . Environmental Role of Algal Toxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ................. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.

.

47

52 52 67 91

93

Plant Transposable Elements PATRICIA NEVERS. NANCY S. SHEPHERD. AND H E I N Z SAEDLER 1. The Phenomenon of Variegation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 . Recognizing Mutations due to Transposable Elements by Classical Genetic

104

Means . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

109

111. Transposable Elements at a Molecular Level ............................. IV . A Model of the Mechanism of Transposition .............................

151

V . Interaction between Transposable Elements .............................. V

170 177

vi

CONTENTS

v1. Induction of Cenomic Instabilities. ...................................... VII. Addendum ............................................................ References ...........................................................

188

193 194

The Dinoflagellate Chromosome D.C. SlGEE I. II.

Introduction ....................................

..................... 208 IV . Dinoflagellate Chromosome Fine Structure . . . . . . . . . . . . . V. . . . . . . . . . . . . . . . 222 VI. Chromosome Changes during the Cell Cycle . . VII. Macromolecular Composition of Dinoflagellate Chromatin . . . . . . . . VIII. IX. References ................. ............................ 260 111. DNA Levels and Chromosome Numbers

AUTHOR I N D E X . . . . SUBJECT INDEX.. ..

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

265 281

CONTRIBUTORS TO VOLUME 12 Numbers in parentheses indicate the pages on which the authors' contributions begin.

LOUISE E. ANDERSON ( I ) , Department of Biological Sciences, University of Illinois at Chicago, Chicago, Illinois 60680, U.S.A. WAYNE W. CARMICHAEL (47), Department of Biological Sciences, Wright State University, Dayton, Ohio 45435, U.S.A. PATRICIA NEVERS (103), Max-Planck-Institut fur Zuchtungsforschung, Egelspfad, D-5000 Koln 30, Federal Republic of Germany HEINZ SAEDLER (103), Max-Planck-Institut fur Zuchtungsforschung, Egelspfad, D-5000 Koln 30, Federal Republic of Germany NANCY S. SHEPHERD' (103), Max-Planck-Institut fur Zuchtungsforschung, Egelspfad, D-5000 Koln 30, Federal Republic of Germany D. C. SIGEE (203, Departments of Botany and Zoology, University of Manchester, Manchester MI3 9PL, England

I Present address: E. I . DuPont de Nemours & Company, Inc., Central Research and Development Department, Wilmington. Delaware 19898, U.S.A.

vii

This Page Intentionally Left Blank

PREFACE The changes in enzyme activity in green plants caused by the transition from light to dark are now regarded as important regulatory processes directing metabolism towards synthesis of sugars and storage compounds in the light, and their breakdown in the dark. Light affects chloroplast enzyme activity in a number of diverse ways, through alteration of stroma1 pH, ion and metabolite levels. However, there are also changes in activity in some enzymes that involve post-translation (probably covalent) modification of the enzyme protein, and these are generally referred to as ‘light modulation’. In her article, Anderson reviews such plant enzyme systems, the biochemical mechanisms involved (probably by reduction of a disulphide bond), their potential molecular basis and the function of modulation in photosynthetic carbon metabolism. One of the most important developments in plant molecular genetics is the rapid improvement of our understanding of the nature and mechanisms of mutation induced by transposable elements. It is interesting to reflect that the origins of this lie in our fascination for variegated plants as horticultural curiosities! Because of our increasing interest in transposable elements for exploring the genetic origins of variation, or as systems for molecular biology and genetic engineering, the review of Plant Transposable Elements by the group at the Max-Planck Institute, Koln (Nevers, Shepherd and Saedler) is particularly welcome. How transposons will be used to isolate genes known only for their phenotypic effects will be seen in the future. The unicellular dinoflagellates are major components of marine and freshwater ecosystems. Apart from their general ecological importance, there are a number of reasons why they are of interest to biologists. In this volume we consider two such aspects. The article by Sigee discusses the very high level of DNA possessed by these organisms, the particular configuration of their chromatin and their nuclear organisation. These are of phylogenetic significance, and to summarise the position as presented in Sigee’s article, it now seems that dinoflagellates are to be regarded as true eukaryotes with some prokaryote features, and that probably they are ‘primitive’ rather than degenerate forms of more advanced ancestors. Algae that can produce toxins effective against animals are found in three of the eight algal divisions, including the dinoflagellates. Carmichael ix

x

PREFACE

reviews various aspects of biology and chemistry of these chemically diverse toxins, some of which can exert potent effects on humans, and considers their potential natural role. Few ideas of such ecological roles appear to have been subject to critical experimentation, and this article should provide a framework for such future work. J. A. Callow

Light/Dark Modulation of Enzyme Activity in Plants' LOUISE E . ANDERSON Department of Biological Sciences University of Illinois at Chicago Chicago. Illinois. U.S.A.

I. Occurrence of Light Modulation .........................................

II. Proposed Mechanisms ..................................................

3 6 8 14 21 21 22 23 24 24 25 26 27 27 27 28 28 29 30 30 30

A . The LEM System ................................................... B. The Ferredoxin-Thioredoxin Reductase. Thioredoxin System ............ C Ferralterin .......................................................... D . Activation of Pyruvic.Pi Dikinase .................................... E . Other Mechanisms .................................................. F Light Modulation of the Activity of Cytosolic Enzymes ................. 111. Regulation of Light Modulation .......................................... A NADP-Linked Malic Dehydrogenase .................................. B. Fructose-Bisphosphatase ............................................. C Sedoheptulose-Bisphosphatase........................................ D. Ribulose-5-Phosphate Kinase ......................................... E Glucose-&Phosphate Dehydrogenase .................................. F. Glyceraldehyde-3-PhosphateDehydrogenase ........................... IV . Dark Modulation ....................................................... A Thioredoxin in Dark Reversal of Light Modulation...................... B . Other Possible Mediators ............................................ C Regulation of Dark Modulation ....................................... V . Changes in the Target Enzyme ........................................... A . Change in the Target Enzyme as a Result of Light Modulation ........... B Change in the Target Enzyme as a Result of Thioredoxin-DTT-Catalyzed Modulation ............................. 32

.

. . . . . .

.

The abbreviations used in this article include the following: CAM. Crassulacean acid metabolism; CMU. 3-(4-chlorophenyl)-l.I-dimethylurea; DCMU. 3-(3.4-dichlorophenyI)I. I-dimethylurea; DCPIP. dichlorophenolindophenol;DTT. dithiothreitol; FCCP. carbonylcyanide p-trifluoromethoxyphenylhydrazone;FeS. iron sulfur; GSH. glutathione; GSSG. oxidized glutathione; LEM. light effect mediator . ADVANCES IN BOTANICAL RESEARCH. VOL. 12

1 Copyright 8 1986 by Academic Press Inc. (London) Ltd . AU rights of reproduction in any form m e N e d .

2

LOUISE E. ANDERSON

VI.

VII. VIII. IX.

C. Change in the Target Enzyme as a Result of Dithiothreitol Treatment.. ... D. Change in Kinetic Parameters of the Target Enzyme.. .................. The Function of Light Modulation ....................................... A. Induction of Photosynthetic Carbon Dioxide Fixation ................... B. CFI-CFo Mg ATPase.. .............................................. C. Stornatal Opening.. ................................................. Effect of Sulfur Dioxide on Light Modulation ............................. Osmotic Stress and Light Modulation. .................................... Conclusions ........................................................... References ............................................................

32 34 36 36 31 31 38 38 39 39

Light activation of the reductive pentose phosphate cycle enzyme NADPlinked glyceraldehyde-3-P dehydrogenase was first observed by Irmgard and Hubert Ziegler while they were assaying for the activity of the enzyme in dehydrated specimens of the resurrection plant Myrothamnus flabellifolia. When they noticed rather large differences in activity levels, they realized that the enzyme was being activated in the plants on the sunflooded lab bench (I. Ziegler, personal communication). Subsequently they demonstrated that the phenomenon occurs in less exotic species such as spinach and maize (Steiger et al., 1971). It is now widely accepted that light/dark modulation of enzyme activity in green plants is an important regulatory process which directs metabolism toward synthesis of sugars and storage products in the light and toward the breakdown of these compounds in the dark. There is reason to suspect that many other processes are also controlled by the chloroplast photomodulation system. Here I will discuss first the extent and distribution of what is thought to be light-dependent posttranslational modification of chloroplastic and cytosolic enzymes, next the various mechanisms which have been proposed for this kind of light modulation, and finally the functions which have been suggested for this process. Because it has not yet been demonstrated that the light-activated and light-inactivated enzymes are actually covalently modified, I will refer to this process by the more general (and more ambiguous)term light modulation. For the purposes of this article light modulation will not be used to refer to those effects which clearly do not involve covalent modification, i.e., pH effects, Mg2+effects, metabolite effects. Light does, of course, also control enzyme activity through the alteration of stromal pH, and ion and metabolite levels. Light modulation of enzyme activity in higher plants has been the subject of several reviews (Anderson, 1979, 1983a,b; Anderson et al., 1980, 1981, 1982, 1983; Buchanan et al., 1979a,b; Buchanan, 1980a,b, 1981,

3

LIGHT/DARK MODULATION

1984a,b; Jacquot, 1984; Losada, 1976) and a workshop after the Sixth International Photosynthesis Congress (Scheibe, 1983a). I. OCCURRENCE OF LIGHT MODULATION

All Orevolving photosynthetic organisms probably have the ability to light activate or inactivate some enzymes. Light modulation has been observed in photosynthetic species ranging from cyanobacteria to C3,C4, and CAM plants (Tables I and 11). It has not yet been observed in any of the photosynthetic bacteria. The enzymes which are light modulated in chloroplasts of pea (Pisum satiuum) are shown in Fig. 1. Four Calvin cycle enzymes and NADP-linked malic dehydrogenase are light activated. Ribulose-bisphosphatecarboxylase is also “light activated” in that as pH and Mg2+levels in the irradiated chloroplast rise, the enzyme is starch

0Z-i

RuBP

6-P-gluconate F 6 P

I

I

4

Pyruvate

I

cytosol

NADP malate-oralacetate

4

chloroplast

Fig. 1. Effect of light on carbon metabolism in the pea leaf. Heavy arrows and sun symbols indicate activation by light; crossing out and sun symbols indicate inactivation by light. The abbreviations used are the following: DHAP, dihydroxyacetone-P; E4P, erythrose-4-P; F6P, fructose-6-P; FBP, fructose-I ,6-P2; GIP, glucose-I-P; G6P, glucose-6-P; G3P, glyceraldehyde-3-P; PGA, P-glyceric acid; R5P, ribose-5-P; RuSP, ribulose-5-P; RuBP, ribulose-l ,5-P2; S7P, sedoheptulose-7-P; SBP, sedoheptulose-l ,7-P2; XuSP, xylulose-5-P. Not shown in figure: transaldolase. From Anderson et a / . (1980) with permission.

4

LOUISE E. ANDERSON

converted to the active form (for a review see Miziorko and Lorimer, 1983). There is new evidence which suggests that an additional activator may be required to convert the enzyme to an activatable form in some plant species (Chastain and Ogren, 1985; Portis, 1985; Salvucci et al., 1985; Seeman and Berry, 1985; Vu et al., 1984). This process is currently not understood and is under intensive investigation. It will not be discussed further here. Two oxidative pentose phosphate pathway enzymes and four enzymes of starch metabolism and glycolysis are light inactivated. In addition, two cytosolic pea leaf enzymes involved in hexose phosphate metabolism are light inactivated. In C4 plants, the chloroplast enzyme pyruvate,Pi dikinase is light activated as is the cytosolic enzyme P-enolpyruvate carboxylase. In Crassulacean acid metabolism plants Penolpyruvate carboxylase is light inactivated (von Willert et al., 1979; Manetas, 1982). Four epidermal enzymes, which function in stomatal carbon metabolism in the pea plant, are also photomodulated. In addition to the enzymes shown in Fig. 1 and Tables I and 11, a lightactivated thylakoid-bound protein kinase controls the distribution of excitation energy between photosystem 1 and photosystem 2 (for a review see Bennett, 1983). There is also evidence that an enzyme in chloroplastic fatty acid biosynthesis, possibly acetyl-CoA carboxylase, is light activated (Roughan et al., 1980; Nakamura and Yamada, 1979). Probably many other enzymes which have not yet been tested are light activated as well, including some of those which are known to be DTT or DTTthioredoxin modulated, such as 6-aminolevulinic acid synthetase (Clement-Metral, 1979; Clement-Metral and Holmgren, 1983) and 6-aminolevulinic acid dehydratase (Balange and Lambert, 1983). To date the photogenes which have been studied appear to be phytochrome controlled (Jenkins et al., 1984), or blue light controlled (Steinmiiller and Zetche, 1985), but it seems possible that some gene expression may be regulated by the same systems involved in photomodulation of carbon enzyme activity. Activatiodinactivation of the enzymes listed in Tables I and I1 varies from about twofold to values approaching infinity. This, of course, reflects the method of calculation of activation (light activity divided by dark activity). Unfortunately there is no measure of absolute dark or light activity at present. For most of these enzymes, at least under optimal assay conditions, dark activity is significant. There is apparently species specificity for modulation. Whether this specificity is at the level of the target enzyme or at the level of the modulating system, or both, is not known. At physiological pH and fructose bisphosphate levels, activity of the dark form of fructose-bisphosphatase is much reduced relative to that of

LIGHT/DARK MODULATION

5

TABLE I Light Acfivafionof Enzyme Activity in Planfs and Cyanobacteria Reductive pentose phosphate cycle enzymes NADP-linked glyceraldehyde-3-P dehydrogenase Ulva, Enferomorpha, Bryopsis, Gracilaria, Polysiphonia (Ziegler and Ziegler, 1967); Ceramium (Ziegler et a/., 1968); Pisum (Anderson and Avron, 1976); Spinaciu (Steiger e f a/., 1971); Vicia, Nicoriana, Brassica, Valerianella, Befa, Myrothamnus, Trificum, Avena, Lemna (Ziegler and Ziegler, 1965);Zea, Saccharum (Steiger et al.. 1971); Tidesfromia (Bjorkman and Badger, 1977); Kalanchoe (Gupta and Anderson, 1978) Ribulose-5-P kinase Anacysfis (Duggan and Anderson, 1975); Pisum (Anderson and Avron, 1976); Spinacia, Zea (Steiger e f a/., 1971); Tidesfromia (Bjorkman and Badger, 1977); Kalanchoe (Gupta and Anderson, 1978) Fructose-bisphosphatase Pisum (Anderson e f a / . , 1979);Spinaciu (Steiger e f a/., 1971); Glycine (Alscher 1982b); Triticum (Leegood and Walker, 1983a) Sedoheptulose-bisphosphatase Pisum (Anderson and Avron, 1976); Spinacia (Champigny and Bismuth, 1976); Trificum (Woodrow and Walker, 1980); Kalanchoe (Gupta and Anderson, 1978) C4 pathway enzymes NADP-linked malic dehydrogenase Pisum chloroplastic (Anderson and Avron, 1976). epidermal (Rao and Anderson, 1983a); Spinacia (Wolosiuk et al., 1977);Zea (Johnson, 1971); Tidesfromia (Bjorkman and Badger, 1977); Kalanchoe (Gupta and Anderson, 1978) Pyruvic,P, dikinase Zea, Amaranfhus (Hatch and Slack, 1969) P-enolpyruvate carboxylase Pisum epidermal (Rao and Anderson, 1983a); Salsolu. Cyperus (Manetas e f al., 1983);Saccharum, Amaranrhus, Sefaria, Cynodon (Karabourniotis ef al., 1985) Other enzymes of carbon metabolism NADP-linked isocitric dehydrogenase Pisum epidermal (Rao and Anderson, 1983a) Glycerate kinase Maize (Kleczkowski and Randall, 1985) Enzymes of nitrate assimilation Nitrite reductase Anucystis (Tischner, 1983) Enzymes of amino acid metabolism Glutamine synthetase Anacystis (Rowell ef al., 1979); Chlorella (Tischner and Huttermann, 1980); Chlumydomonas (Cullimore, 1981) Glutamate synthetase Chlorella (Tischner and Huttermann, 1980) Phenylalanine ammonia lyase Spinacia (Nishizawa e f a/.. 1979) Other enzymes CFI-CFO Spinacia (Vallejos e f a/., 1983) NAD kinase Pisum, Trificum (Muto e f a/., 1981) Pyridine nucleotide-linked ascorbate peroxidase Spinacia, Sedum (Kow e f a/., 1982)

6

LOUISE E. ANDERSON

TABLE I1 Light Inactivation of Enzyme Acriviry in Planrs and Cyanobacteria Oxidative pentose phosphate cycle enzymes Glucose-6-P dehydrogenase Anacysris (Duggan and Anderson, 1975); Pisum chloroplastic and cytosolic (Anderson and Duggan, 1976); Spinacia (Lendzian and Ziegler, 1970) Transaldolase Pisum (Anderson, 1981) Glycolytic enzymes Phosphorylase Pisum chloroplastic (Heuer et a / . , 1982) P-glucomutase Pisum chloroplastic (Heuer et a / . , 1982) P-glucoisomerase Pisum chloroplastic (Heuer et a / . , 1982) P-fructokinase Pisum chloroplastic (Heuer et a / . , 1982). cytosolic (Kachru and Anderson, 1975) C4 pathway enzymes

P-enolpyruvate carboxylase Sedum (Manetas, 1982) Enzymes of amino acid metabolism Aspartate aminotransferase Pisum epidermal (Rao and Anderson, 1983a)

the light form (Charles and Halliwell, 1981b). Likewise greater activation of sedoheptulose-bisphosphatase is observed when physiological levels of sedoheptulose bisphosphate are used (Laing et al., 1981). Real activation of these and the other enzymes shown in Table I may then be considerably greater than is apparent since those values are for optimum assay activity. 11. PROPOSED MECHANISMS

The involvement of the photochemical apparatus in light activation of glyceraldehyde-3-P dehydrogenase was first indicated in experiments in Ziegler’s laboratory (Ziegler et al., 1968). It was shown that the action spectrum for activation of NADP-linked glyceraldehyde-3-P dehydrogenase follows the action spectrum for photosynthesis. Therefore the pigments of the photochemical apparatus, and not phytochrome or the cytoplasmic blue light receptor, are the photoreceptors. Inhibitor experiments and experiments with photosystem I particles demonstrate that the reducing side of photosystem I is involved in light modulation (Fig. 2). DCMU (Latzko et al., 1970; Leegood and Walker, 1980b; Avron and Gibbs, 1974; Anderson and Avron, 1976; Hatch, 1977; Gupta and Anderson, 1980) and CMU (Champigny and Bismuth, 1976),

7

LIGHT/DARK MODULATION

which block electron transport between the two photosystems, inhibit light modulation. This inhibition can be overcome by adding electrons from ascorbate or DCPIP into the system beyond the DCMU block (MvC Akamba and Anderson, 1981). Stimulation of electron transport with antimycin (Champigny and Bismuth, 1976; Leegood and Walker, 1980a,b) or tetramethylethylenediamine (Champigny and Bismuth, 1976) increases light activation, while electron acceptors such as nitrite (Leegood and 2 (Latzko et al., 1970; Leegood and Walker, 1980b), Walker, 1980b) or 0 or compounds which are reduced by photosynthetically generated NADPH, such as oxalacetate and CO2, inhibit or reverse light activation (Leegood and Walker, 1980a,b). Diquat (Avron and Gibbs, 1974; Anderson and Avron, 1976; Gupta and Anderson, 1980) and methylviologen (Wildner, 1975; Hatch, 1977; Mills, 1983), which shunt electrons from photosystem I to 02,are inhibitors, as is pyocyanin (Leegood and Walker, 1980b), which short-circuits electron transport by shunting electrons from the reducing to the oxidizing side of photosystem I. Clearly electron transport to photosystem I is required for light modulation of enzyme activity. In confirmation of the inhibitor experiments, photosystem I particles support light modulation (MvC Akamba and Anderson, 1981). Since FCCP inhibits light modulation of only one enzyme (glyceraldehyde-3-P dehydrogenase) (Anderson and Avron, 1976; Gupta and AnderMV Diquat I I

I

DCMU CMU

Ascorbate D T

I

p/hy

L P S II

\

\

NADP '700

Fig. 2. Effect of inhibitors on photosynthetic electron transport. DCMU and CMU block electron transport from photosystem 11 (Ps 11). Diquat and methylviologen (MV) shunt electrons from the system at the photosystem I level (Ps I). Ascorbate or DCPIP feeds electrons in between the two photosystems.

8

LOUISE E. ANDERSON

son, 1980), proton pumping and photophosphorylation appear not to be necessary for light modulation. Four mechanisms have been advanced to explain the light modulation of the activity of the carbon metabolism enzymes. The light effect mediator, or LEM system, has been developed in this laboratory (Anderson et al., 1981). Buchanan and co-workers have proposed a system which involves ferredoxin and thioredoxin (Buchanan, 1980a). The same group has decribed a ferredoxin-independent factor for light modulation, ferralterin (Buchanan, 1980a). In Australia, Hatch and co-workers (Sugiyama and Hatch, 1981; Ashton et al., 1984) have described an entirely different system involving phosphate and ADP and a single regulatory protein for the activation and inactivation of pyruvate,Pi dikinase in C4 plants. Properties of the proteins thought to be involved in light/dark modulation in the chloroplast are listed in Table 111.

A. THE

LEM SYSTEM

Indications of the involvement of thiol compounds in light modulation first came from experiments of Latzko er al. (1970). They observed that arsenite, which reacts with vicinal dithiols (Fig. 3), and iodoacetamide, which reacts with thiol groups, inhibited light activation of ribulose-5-P kinase in intact chloroplasts, and that in broken chloroplast preparations, dithiothreitol activated the enzyme. They postulated that a vicinal dithiol was involved in light modulation. That the vicinal dithiol was membrane-bound and light generated was shown in experiments with crude broken chloroplast systems (Anderson and Avron, 1976). It was found that if the thylakoid membranes were illuminated and exposed to arsenite, photomodulation was subsequently inhibited in the recombined broken chloroplast system. Arsenite was not effective unless the thylakoids were irradiated during the treatment. The effect of arsenite was reversed by treatment with DTT. These experiments indicated that the vicinal dithiol group was generated when the thylakoids were irradiated. Sulfite, which reacts with disulfide bonds, giving rise to a free thiol group and a thiosulfonic acid (Fig. 3), also inhibited. Sulfite was effective even when the thylakoids were exposed to the inhibitor in the dark. These experiments indicate that a redox-active disulfide bond is reduced by transfer of electrons from the photosynthetic electron transport system giving rise to a vicinal dithiol. The membrane component containing these groups was named the “light effect mediator,” or LEM.

LIGHT/DARK MODULATION

9

Fig. 3. (A) Reaction of arsenite with vicinal dithiol; (B) reaction of sulfite with disulfide.

1. The LEM The LEM is removed from the chloroplast thylakoid membranes when they are treated with Zwittergent, a zwitterionic sulfobetaine detergent (Mohamed and Anderson, 198la). Light modulation is restored when the Zwittergent extract is dialyzed, concentrated, and recombined with the depleted thylakoids. The factor in the detergent extract is sulfite sensitive, but the depleted thylakoid membranes are not (Mohamed and Anderson, 1981b). Arsenite-treated thylakoids which have been stripped of the LEM are as effective as control thylakoids in a reconstituted light activation system. But the LEM activity in the Zwittergent extract of the arsenitetreated thylakoids is increased when the extract is treated with DTT and the DTT is removed by gel filtration (Anderson et al., 1985). Apparently the arsenite- and sulfite-sensitive LEM is the Zwittergent-stripped thylakoid membrane component necessary for light activation. Sonication enhances reconstitution (Anderson et al., 1985). This, and the requirement for a detergent for solubilization, indicate that the LEM is an integral membrane protein. Since it interacts with soluble stromal components, it is likely that the LEM is located on the surface of the thylakoid membrane. The coincidence of activity for activation of NADP-linked malic dehydrogenase, fructose-bisphosphatase, and NADP-linked glyceraldehyde dehydrogenase on different Procion dye-Sepharose columns indicates that there is only one LEM (Anderson et al., 1985). Specificity for the target enzyme must then be a property of the soluble component of the system. After partial purification and filtration through Sephadex G-75 or elution from Red H3BN Sepharose, the LEM has no apparent “thioredoxin activity,” as measured by DTT-dependent activation of NADP-linked malic dehydrogenase (Anderson et al., 1985). Neither thioredoxin nor plastocyanin can replace the LEM preparation in the light activation assay. EPR spectra of the stripped thylakoid membranes indicate the presence of iron sulfur centers A and B. The LEM then does not appear to be identical with any of these factors.

TABLE 111 Factors Proposed to be Involved in LightlDark Modulation Subunits Molecular weight

Number

Molecular weight

Visible spectra

),A(

Remarks

Thylakoid

Light effect mediator (LEM) E, Protein modulase

10,500

Femdoxin, tbioredoxin reductase

38,000

None

39,000

3

Source Pea

Anderson et al. (1985)

Pea

Anderson et al. (1985) de la Torre et al. (1979)

bound Requires ferredoxin None

2 = 12,500 1 = 14,000

408 nm

Spinach 3 Fe/mol

Spinach

Schiinnann (1981) Hutcheson et al. (1983) Yee et al. (1981) Hutcheson and Buchanan (1983a) Schiirrnann et al. (1981) Wolosiuk et al. (1979)

36,000

None

Maize

38,000 31,000

None

Nostoc Kalanchoe

Thioredoxin fa

11,200

Thioredoxin fb

16,000

Spinach.

None Heat labile

Reference

Spinach

16,000

Thioredoxin f

2

Spinach

Soulie e f al. (1981)

3 Isoenzymes

Nostoc Maize

Yee et al. (1981) Hutcheson et al.

Heat stable

Spinach

Wolosiuk et a/.

3 Isoenzymes

Nostoc Maize

Yee et al. (1981) Hutcheson e f al.

Monomeric in high ionic strength solutions

2 = 8,000

16,000 20,000

(1983)

Thioredoxin m

9,000

(1979) 9,000 10,000

(1983)

Thioredoxin mb

11,500

None

Spinach

Schiirmann et al.

Thioredoxin mc

12,300

None

Spinach

Schurmann e f a/.

Ferralterin

28,000

3

28,000

3

(1981)

-

(1981)

e

pYruvate,Pi dikinase activating factor

88,000

2= 1= 2= 1=

12,000 7,000 12,000 7,000

410 nm

3 Fe/mol

Spinach

de la Torre et a/.

410 nm

3 Fe/mol

Nostoc

de la Torre et al.

Maize

Nakamoto and Sugiyama (1982)

(1982) (1982)

12

LOUISE E. ANDERSON

The molecular weight of the LEM is about 10,500 as estimated by gel filtration (Anderson et al., 1985). The LEM is, in size and in having a redox active vicinal dithiol group, very much like thioredoxin (see Section I1,B) or glutaredoxin. But unlike these proteins, the LEM appears to be involved directly or indirectly in a 1 e- transfer reaction. There seems to be no reason to suspect that the LEM is a thioredoxin.

2. Protein Modulase Evidence for a soluble component in this system first came from experiments of Ashton and Anderson (1981) in which a stromal factor was shown to be required for light modulation. This factor was very unstable. Recently it has been resolved into two components (Fig. 4; Anderson et al., 1985). The first component is ferredoxin. The second has been extensively purified by adsorption onto ferredoxin-Sepharose and elution with a salt gradient. Soluble thioredoxin is not required, but involvement of thioredoxin has not been eliminated since some thioredoxin will be bound to the thylakoid membranes (Ashton et al., 1980) used in the light activation assay. It is possible that thioredoxin is a tightly bound modulase subunit. This system is not sensitive to fairly high levels of ferredoxin antibody (Ashton, Mohamed, and Anderson, unpublished). Therefore either this ferredoxin differs from the soluble ferredoxin involved with ferredoxin-NADP reductase in photosynthetic NADP reduction or the antigenic sites are shielded when it is complexed with the modulase protein.

Fig. 4. Separation of protein modulase (PM)activity into two components on Sephacryl S-200. The activity of the first component (0)is enhanced by the addition of the second component (0). which appears to be identical with ferredoxin.

LIGHT/DARK MODULATION

13

Fig. 5 . Hypothetical scheme for light activation catalyzed by the LEM system. In frame 1 all of the components are in the dark, oxidized state. In frame 2 the LEM is reductively activated by transfer of electrons from some component on the reducing side of photosystem 1 before ferredoxin. In frame 3 ferredoxin (Fd) is reduced and passes an electron to protein modulase (PM).In frame 4 protein modulase is reduced and passes its electrons to the target enzyme (TE).In frame 5 the target enzyme is reduced and undergoes a change in conformation to the light form.

If ferredoxin is involved in LEM system-catalyzed light modulation, what then is the role of the LEM? Originally it was thought to be a reductant, first for the target enzyme, then for protein modulase. Since the LEM becomes arsenite sensitive in the light in the absence of the soluble stromal factors ferredoxin and protein modulase, it seemed reasonable to assume that the LEM is reductively activated by the electron transport system. But if electron transport through ferredoxin is required for modulation, and if the LEM is reduced in the absence of ferredoxin (or under conditions which do not allow light modulation of target enzyme activity), then the LEM cannot be either the terminal target enzyme reductant or a 2 e- carrier. It is tempting to speculate that the LEM is a modifier or regulatory protein which must first be activated before light modulation can proceed. Such a possible scheme for light modulation involving these factors is shown in Fig. 5 . Alternatively, the LEM may be a 1 e- carrier which shifts orientation in the thylakoid membrane to expose vicinal dithiols in the light.

14

LOUISE E. ANDERSON

active

MDH<~ MDH2I inactive\

NADP

Fig. 6. Scheme for light modulation involving thioredoxin proposed by Buchanan and coworkers. Electrons are passed from ferredoxin (Fd) to ferredoxin-thioredoxin reductase (Fd,ThR) to thioredoxin (Th) and then to the target enzyme (MDH, NADP-linked malic dehydrogenase). From Anderson (1983a) with permission.

B. THE FERREDOXIN-THIOREDOXIN REDUCTASE, THIOREDOXIN SYSTEM

Buchanan et al. (1967) demonstrated a light- and ferredoxin-dependent activation of fructose-bisphosphatase in broken spinach chloroplast preparations. Light activation, according to the scheme later proposed by Buchanan and collaborators (Buchanan, 1980a), involves electron transport through ferredoxin to the soluble enzyme ferredoxin-thioredoxin reductase which, in turn, reduces thioredoxin. Reduced thioredoxin interacts with the target enzyme converting it to the light form. See Fig. 6. Ferredoxin-thioredoxin reductase may or may not be an iron sulfur protein (see the next section). Several thioredoxins have been implicated in light modulation. These are usually referred to as thioredoxin f (more active with fructose-bisphosphatase)and thioredoxin m (more active with malic dehydrogenase). Variants are designated thioredoxin fa, thioredoxin fb, etc. (see Table 111). 1 . Ferredoxin- Thioredoxin Reductase This enzyme is required for thioredoxin- and ferredoxin-dependent light modulation. It is thought to mediate the transfer of electrons from reduced ferredoxin to oxidized thioredoxin. An analogous enzyme, which is part of the ribonucleotide reductase system in bacterial and mammalian systems, uses NADPH as reductant. The ferredoxin-dependent enzyme does not appear to be structurally related to the NADPH reductases. There is disagreement as to the nature of the ferredoxin-dependent reductase, which was first separated from thioredoxin by Schurmann et al. (1976). Buchanan and co-workers have isolated a colorless protein (M, 38,000) from spinach (de la Torre et al., 1979), maize (Hutcheson et al.,

LIGHT/DARK MODULATION

15

1983), and Anabaena (Yee et al., 1981), which they claim has ferredoxinthioredoxin reductase activity. A smaller (3 1 kDa) reductase has been partially purified from Kulanchoe (Hutcheson and Buchanan, 1983a). This reductase cross-reacted with antibody to maize ferredoxin-thioredoxin reductase, but it was not sufficiently pure for spectral studies. Schiirmann (1981) has isolated a brownish protein (hm,,408nm) from spinach which consists of one 14 kDa and two 12.5 kDa subunits and has reductase activity. The spectrum of this protein is shown in Fig. 7. The highest ratio A408:A279 found is 0.37. The Orsay group (Droux et al., 1984; Jacquot et al., 1983) has isolated a similar brownish reductase from spinach. Antibody against this reductase completely inhibits light activation in broken spinach and pea leaf chloroplast preparations. Schiirmann (1981) has observed that a colorless protein, which is not involved in thioredoxin reduction, copurifies with the reductase. It remains to be determined whether there are two ferredoxin-thioredoxin reductases in spinach or whether one of these is an artifact. A similar protein, which reportedly functions independently of ferredoxin, was isolated earlier by the Berkeley group and named ferralterin (Section 11,C). In view of the similarities between the brownish ferredoxin-thioredoxin reductase and ferralterin, it seems quite possible that they are the same protein. Strangely, although ferralterin has a distinctive EPR spectrum (see Section I1,C) EPR studies of the brownish ferredoxinthioredoxin reductase appear not to have been done. The redox potential for oxidation and reduction of the iron sulfur group of the brownish ferredoxin-thioredoxin reductase has not yet been determined. If it is the same as that of ferralterin (Section I1,C) then some other as yet unidenti-

WAVELENGTH (nm)

Fig. 7. Absorption spectrum of the brownish ferredoxin-thioredoxin reductase. From Schiirrnann (1981) with permission.

16

L O U S E E. ANDERSON

fied group on the protein must be responsible for the electron transport properties of the molecule, since that FeS center gives the protein a very positive redox potential. If either the colorless reductase or the brownish reductase does transfer electrons from ferredoxin to thioredoxin (or to the target enzyme redox active disulfide) then the functional prosthetic group must be capable of a 1 e- to 2 e- transfer. Flavins and quinones usually fulfill this function. Ferredoxin-thioredoxin reductase is not restricted to photosynthetic organisms: An FAD-containing ferredoxin-thioredoxin reductase has been isolated from the obligate anaerobe Clostridium pasteurianum (Hammel et al., 1983). The analogous NADPH-requiring enzyme, a component of the system for ribonucleotide reduction in mammalian and bacterial systems (see above), is a flavoprotein. Ferredoxin-thioredoxin reductase activity has been found in darkgrown barley seedlings (Buchanan et al., 1978), but NADPH thioredoxin reductases have been isolated from wheat flour (Suske et ul., 1979) and dark grown carrot tissue culture cells (Cao et al., 1984). Droux et al. (1984) have shown that their brownish ferredoxin-thioredoxin reductase binds to ferredoxin-Sepharose, which indicates that the two proteins interact in uitro and hence might interact in uiuo. Kalunchoe ferredoxin-thioredoxin reductase and, according to Buchanan (1983), the reductases from Anabaena, maize, and spinach also bind to ferredoxin immobilized on Sepharose. It is not clear whether it is the colorless 38 kDa protein described by the Berkeley group or the iron sulfur protein of Schurmann which is bound in the later experiments. According to Droux et al. (1984) their reductase also binds to thioredoxin-Sepharose. According to Jacquot et al. (1983) the brownish reductase catalyzes the activation of NADP-linked malic dehydrogenase with either spinach or pea thylakoids. In the latter case, the system is thioredoxin dependent only after the thylakoids are treated with Triton X-100in the presence of high levels of salt. SouliC et ul. (1981) have shown that thioredoxin f can be reduced by ferredoxin-NADP reductase with NADPH as reductant. As has been pointed out by Lara et al. (1983), none of the ferredoxinthioredoxin reductases has been shown to reduce thioredoxin, either in catalytic or in substrate amounts. The enzyme may, then, be misnamed and might function in some way other than that proposed (see Fig. 6). For example, thioredoxin, reduced directly by the photosynthetic electron transport system, might interact with the reductase to form an active complex which is able to reduce the target enzyme. In such a complex the electrons for reduction might pass directly from the reductase to the target enzyme. Alternatively, thioredoxin might be a more effective reductant when complexed with the reductase.

LIGHT/DARK MODULATION

17

2. Thioredoxin Thioredoxins are small and rather unusual proteins. Most thioredoxins are heat stable; all contain redox-reactive vicinal dithiol groups. In the oxidized form the disulfide protrudes from the surface of the protein (Holmgren, 1981). Among enzymes studied to date, thioredoxin and the closely related protein glutaredoxin are unique in having protruding active sites. Thioredoxins are thought to be involved in the reduction of nucleotides to deoxynucleotides (see Section II,B,l), in sulfur metabolism, and in other reactions. For their role in dark modulation see Section IV,A. Levels of thioredoxin in chloroplasts are quite high (Scheibe, 1983b). A part of the “thioredoxin-like” activity of the pea leaf chloroplast is membrane bound (Ashton et al., 1980). The chloroplast thioredoxins are classified as thioredoxin f (active in the activation of fructose-bisphosphatase) and thioredoxin m (active in the activation of NADP-linked malic dehydrogenase). Two thioredoxin fs, thioredoxin fa and thioredoxin fb, and several thioredoxin m’s have been isolated (see Table 111). Buchanan’s group found two thioredoxins in spinach chloroplasts (Wolosiuk et al., 1979) and three in barley chloroplasts (Crawford et al., 1979). According to this group, thioredoxin f is heat labile and has a molecular weight of 16,000 when isolated from spinach. This thioredoxin, now named thioredoxin fb, has also been purified to homogeneity by SouliC et al. (1981). Apparently it is a dimer composed of two 8 kDa monomers which dissociate in high ionic strength solutions. The dimer seems to be the active form of the factor (or conditions which favor dimerization favor activation) (Soulik et al., 1981). A second thioredoxin f, now called thioredoxin fa, was first isolated by Schurmann and coworkers (1981) and later by Buc et al. (1983, 1984). The purified (active) protein exists only as a monomer with a pZ of 6.1 (Schurmann el al. 1981). The amino acid compositions of both thioredoxin f s have been determined (Schurmann et al., 1981; SouliC et al., 1981). The thioredoxin m isolated by Wolosiuk and co-workers (1979) from spinach is a 9 kDa heat-stable monomer. Schiirmann and co-workers (1981) find two m-type thioredoxins in spinach. They differ slightly in molecular weight (1 1,500 and 12,300) and in pZ values (5.2 and 5.0). Their amino acid compositions are similar and differ from that of thioredoxin fa. Antibodies to the two m-type thioredoxins cross-react with both of these thioredoxins, but not with thioredoxin fa. Likewise antibody to thioredoxin fa does not cross-react with the thioredoxin m’s. Three thioredoxins (two thioredoxin m’s and one thioredoxin f) have been separated from sorghum and from bean (Phaseolus vulgaris) leaves by Jacquot et al. (1978). Two thioredoxin f s and one thioredoxin m have been isolated from

18

LOUISE E. ANDERSON

Kalunchoe leaves (Hutcheson and Buchanan, 1983a). These appear to be similar in molecular weight and specificity to those from spinach and maize, but considerably less stable. A much larger protein (30 kDa) with thioredoxin activity has been isolated from Scenedesmus by Follmann and co-workers (Berstermann et al., 1983). Two smaller thioredoxins, apparently cytosolic, M, about 12,600, have also been isolated from this green alga. A multiplicity of proteins with thioredoxin activity have been isolated from maize by Jacquot and Gadal (1983). It is suggested that some of these proteins may have other functions in the plant. Buchanan's group (Hutcheson et al., 1983) has separated three thioredoxin f s and three thioredoxin m's from maize. The three forms, in both cases, have different PI'S. Tsugita et al. (1983) have shown that in absence of effectors, such as fructose bisphosphate, only thioredoxin f, and not thioredoxin m, is capable of catalyzing the DTT-dependent activation of fructose-bisphosphatase. Buc et al. (1983) found that both thioredoxin fa and thioredoxin fb, but not thioredoxin m, catalyze the activation of fructose-bisphosphatase, whereas only thioredoxin fb supports the DTT-dependent activation of sedoheptulose-bisphosphatase. NADP-linked malic dehydrogenase, on the other hand, is activated by all of the chloroplast thioredoxins (Tsugita et al., 1983; Buchanan, 1983) and by Escherichia coli thioredoxin (Tsugita et al., 1983). The same specificity for thioredoxins is seen with the ferredoxinthioredoxin reductase system (Schurmann, 1981; Wolosiuk et al., 1979). Spinach thioredoxin f is more efficient than thioredoxin m in catalyzing the activation of phosphoribulokinase and NADP-linked glyceraldehyde3-P dehydrogenase with DTT and in the ferredoxin-thioredoxin system (Wolosiuk et al., 1979). Crawford et al. (1979) report that there are two thioredoxins in spinach leaf cytosol similar to chloroplastic thioredoxin f and thioredoxin m, which they designate thioredoxin cf and cm.The function of these thioredoxins is not known, but it seems possible that they are involved in modulation of enzyme activity in the cytosol, either as light or dark modulators (see below). Berstermann et al. (1983) have purified multiple forms of thioredoxins from soybean and wheat seeds. According to Schmidt (1980, 1981), two thioredoxins which he has isolated from the cyanobacterium Synechococcus 6312 (Anacystis nidulans) are able to catalyze the DTT-dependent activation of fructose-bisphosphatase isolated from that organism, but not the activation of the spinach chloroplast enzyme. Two thioredoxins have been isolated from

LIGHT/DARK MODULATION

19

Anabaena sp. 7119 (Nostoc muscorum) by Crawford et al. (1983a,b). According to these workers (Crawford et al., 1984) both thioredoxins are active in DTT-dependent and ferredoxin-thioredoxin-dependent activation of Anabaena fructose-bisphosphatase, sedoheptulose-bisphosphatase, and ribulose-5-P kinase and also in the DTT-dependent and ferredoxin-thioredoxin light-dependent activation of spinach fructose-bisphosphatase and NADP-linked malic dehydrogenase. Some differences in target enzyme specificity were observed between higher plant and Anabaena thioredoxins. In contrast, Gleason (1985) finds only one thioredoxin in this organism. Reduction, according to Gleason, occurs not by a ferredoxin-thioredoxin reductase, but by a membrane-bound NADPH thioredoxin reductase. Reduced thioredoxin is an efficient reductant for ribonucleotide reduction with the ribonucleotide reductase from this organism. In the presence of DTT this thioredoxin activities spinach NADPlinked malic dehydrogenase and fructose-bisphosphatase, but not Anabaena fructose-bisphosphatase (Whitaker and Gleason, 1984). Fructose-bisphosphatase from the closely related A . cylindrica is activated by the single thioredoxin isolated from A . cylindrica (Ip et al., 1984). Clement-Metral and Holmgren (1983) have purified thioredoxin from Rhodopseudomonas sphaeroides to homogeneity and have partially purified an NADPH-dependent thioredoxin reductase from this photosynthetic bacterium. The thioredoxin is similar to the thioredoxin isolated from Anabaena by Gleason and Holmgren (1981) with respect to amino acid composition. The Rhodopseudomonas system catalyzes the activation of 6-aminolevulinic acid synthetase (Clement-Metral and Holmgren, 1983) and might be responsible for the regulation of bacteriochlorophyll synthesis by 02 in the photosynthetic bacteria. Spinach thioredoxin mc and an active site peptide from thioredoxin f have been sequenced by the Swiss group (Tsugita et al., 1983; Schiirmann et al., 1989, and the sequence of Anabaena 7119 thioredoxin has been determined by Gleason et al. (1985) (see Fig. 8). Like all thioredoxins, these thioredoxins and the thioredoxin isolated from Anabaena cylindrica (Ip et al., 1984)contain the invariant tetrapeptide -Cys-Gly-Pro-Cys-.The sequence around the active site of spinach thioredoxin m shows extensive homology with the same regions of the Anabaena, Escherichia coli (Fig. 8), and Corynebacterium nephridii thioredoxins. In contrast, the active site peptide of thioredoxin f shows little homology with the active site peptides of any of these thioredoxins (Tsugita et al., 1983; Schiirmann, 1983). Spinach fructose-bisphosphatase binds to thioredoxin-Sepharose (Pla and Lopez-Gorge, 1981). Elution can be accomplished by addition of

20

LOUISE E. ANDERSON

Spinach thioredoxin fa Val-Leu-~-Met-Phe-Thr-Gln-Trp-Cys-Gly-Pro-Cys-Lys-Ala-Asn-Gly-Asp-Lys-Glu-Ala-Thr-Gln-His

Spinach thioredoxin mc Met-Val-Asp-Phe-Trp-Ala-Pro-Trp-Cys-Gly-Pro-Cys-Lys-Leu-Ile-Ala-Pro-Val-Ile-Asp-Glu-Leu-~

Anabaena Leu-Val-Asp-Phe-Trp-Ala-Pro-Trp-Cys-Gly-Pro-Cys-Arg-Met-Val-Ala-Pro-Val-Val-Asp-Glu-Ile-~

E . coli

Leu-Val-Asp-Phe-Trp-Ala-Glu-Trp-Cys-Gly-Pro-Cys-Lys-Met-Ile-Ala-Pro-Ile-Leu-Asp-Glu-Ile-~

30

24

40

Fig. 8. Amino acid sequence of spinach thioredoxin fa, thioredoxin mc, and E. coli thioredoxin. Sequences are aligned to show overlap. From Tsugita ef ul. (1983) and Gleason (1985).

MgC12. Binding is not specific for thioredoxin f. Thioredoxin m and the cytosolic thioredoxins are also effective. While the phosphatase does not bind to ferredoxin-Sepharose, it does bind to a number of amino acids and related compounds attached to Sepharose. Hence the binding may be relatively unspecific. Scheibe (1983b) estimates that the thioredoxin m concentration in pea chloroplasts is about 50 p M . Surprisingly, the levels of ferredoxin required by the ferredoxin-thioredoxin system have never been reported. In a broken chloroplast system (66 pg chlorophyll ~ m equivalent - ~ thylakoids and stroma) activity appears to be saturated by 2 p M concentrations (Buchanan et al., 1967). In a very preliminary experiment with purified components of the system (Jacquot et al., 1983) rather high levels of thioredoxin appear to be required for maximal light activation of NADP-linked malic dehydrogenase. On the other hand, the thioredoxin f concentration required for half-maximal activation of fructose-bisphosphatase can be calculated to be approximately 0.5 p M from data of Schurmann et al. (1981). This value is probably only valid for the conditions of the particular assay, but it seems likely that thioredoxin concentrations in the chloroplast (Scheibe, 1983b) could support activation by this system. The major problems with the ferredoxin-thioredoxin system lie in the sequence of events and the nature of the reductase. There is no direct evidence that thioredoxin is reduced by the reductase, nor is there direct evidence that reduced thioredoxin activates enzymes. It is not clear how ferredoxin-thioredoxin reductase, if its iron-sulfur group can only function as an oxidant under physiological conditions, can act as a one electron acceptor and transfer and two electrons to reduce a disulfide bond. Clearly additional biochemical characterization of this system is needed and its relationship to the LEM system defined.

LIGHT/DARK MODULATION

21

C. FERRALTERIN

Ferralterin is an iron-sulfur protein which is extraordinarily similar to Schiirmann’s and Jacquot’s ferredoxin-thioredoxin reductase (see above). This enzyme has been purified from Anabaena and from spinach (de la Torre et al., 1982). Some preparations of this enzyme support light activation of fructose-bisphosphatase without the addition of ferredoxin and thioredoxin. The activity of the enzyme in other preparations is stimulated by addition of ferredoxin and/or thioredoxin (Lara et al., 1983). The authors speculate that this occasional dependence on thioredoxin or ferredoxin could have been related to the state of the protein or of the thylakoid membrane. It may be that only very low levels of thioredoxin and ferredoxin are required for light activation and that these were sometimes present on the thylakoid membranes. The molecular weight of ferralterin is reported to be around 28,000 (de la Torre et al., 1982). Before the final steps of purification the enzyme appears to be somewhat larger, since it emerges from a Sephadex G-100 column ahead of ferredoxin-thioredoxin reductase (M,38,000) (Lara et al., 1980; Crawford et al., 1983b). As isolated, ferralterin consists of three subunits, two apparently identical 7 kDa monomers and a single 12 kDa monomer (de la Torre et al., 1982). It contains 3.3 nonheme irons per mole and a corresponding amount of acid-labile sulfur. Presumably there is one 4Fe4S center or two 2Fe2S centers per molecule. It is brownishyellow in color and has a peak in the visible spectrum at 410 nm. The highest ratio of A410:A280 found for the spinach enzyme is 0.35 and for the Anabaena enzyme, 0.39. The EPR signal from oxidized ferralterin is rhombic with g values of 2.10, 2.01, and 2.00. The midpoint redox potential is +410 mV, which is the most oxidizing potential ever reported for an FeS protein. Therefore it is unlikely that the FeS center of ferralterin is directly involved in electron transport. Exactly how ferralterin accomplishes activation of fructose-bisphosphatase is not known. No reports of activation of enzymes other than fructose-bisphosphatase by ferralterin have been published.

D. ACTIVATION OF

PYRUVIC,Pi DIKINASE

The system for activation of pyruvic,Pi dikinase is unique. There is no evidence for thiol involvement (Chapman and Hatch, 1981). Inorganic phosphate and pyruvicpi dikinase-regulatory protein are required for activation of the enzyme, which involves formation of PPi, with the phosphate moieties coming from the required Pi and dikinase threonine-bound phosphate (Ashton and Hatch, 1984; Ashton et al., 1984). Inactivation

LOUISE E. ANDERSON

22

.

E

Y

E

yip (active)

PVRUVATE

4

PPi

Fig. 9. Scheme for the activation and inactivation of pyruvate,Pi dikinase. PDRP is pyruvate,Pi dikinase regulatory protein. PEP is P-enolpyruvate. Phosphorylation of the histidine residue occurs during catalysis. Phosphorylation of the threonine residue occurs during modulation. From Burnell (1984) with permission.

involves the same regulatory protein (Burnell and Hatch, 1983),ADP, and very low levels of ATP (see Fig. 9). During inactivation the P-phosphate of ADP is transferred to the dikinase threonine (Ashton and Hatch,1983a, 1984; Ashton et al., 1984). The regulatory protein is then the only known protein kinase which utilizes ADP rather than ATP as the source of transferable phosphate. The function of the ATP is apparently to phosphorylate a His on the enzyme, rendering it susceptible to ADP-dependent phosphorylative inactivation (Burnell and Hatch, 1984a,b). ADP, AMP, and inorganic pyrophosphate inhibit activation; pyruvate and related compounds inhibit inactivation. It is possible that the light effect is mediated by energy charge and through changes in metabolite levels (Nakamot0 and Edwards, 1983b, 1984a). The enzyme in maize is activated in the dark under anaerobic conditions (Nakamoto and Edwards, 1983a).

E. OTHER MECHANISMS

Experiments of Scheibe and Beck (1979) and Alscher (1982b) suggest that there is a membrane-associated light activation system in spinach and soybean, respectively. This could represent a membrane-bound version of one of the systems described above. Rudd and Anderson (1983) report that light inactivation of a membrane-bound form of P-fructokinase from

LIGHT/DARK MODULATION

23

pea leaf chloroplasts is sensitive to DCMU and to Diquat, which indicates that photosystem I is involved in light modulation of this thylakoid-bound enzyme. A part of the glucose-6-P dehydrogenase, ribulose-5-P kinase, and glyceraldehyde-3-P dehydrogenase activities in the chloroplast are membrane bound and are released to the stroma when the chloroplast is irradiated (Ben-Bassat and Anderson, 1981; Anderson and Ben-Bassat, 1981). Release is DCMU sensitive. It seems possible that the light modulation system is involved in binding and/or release. Fructose-bisphosphatase from Anacystis nidulans is inactivated by glutathione and by DTT (Udvardy et al., 1983). According to these workers there is no detectable glutaredoxin activity in this photosynthetic prokaryote. It is suggested that GSH/GSSG levels may be important in regulation of the activity of this enzyme in uiuo. Halliwell and Foyer (1978) have clearly shown that GSWGSSG levels remain relatively constant during lightldark transitions, which would seem to rule out GSH as a mediator in higher plants. Schriek and Schwenn (1984) found that spinach chloroplast ferredoxinNADP reductase contains a protein capable of catalyzing NADPH-dependent activation of fructose-bisphosphatase. Although thioredoxin stimulates activation, there is substantial (50% of complete system) activation without the dithiol. Ferredoxin is not required. The activating protein has NADPH-ferredoxin and NADPH-DCPIP diaphorase activity. It seems possible that this enzyme is related to protein modulase or ferredoxinthioredoxin reductase. It has been suggested that NADP, NADPH, and ATP might be the light mediators (Lendzian, 1978, 1980; Lendzian and Bassham, 1975; Miiller, 1970; Miiller et al., 1969; Schmidt-Clausen et al., 1969; Ziegler, 1972). Although these compounds will act as effectors in uiuo, there is no indication that this type of modulation involves covalent modification. Activation through NADPH reduction cannot be ruled out (see above). Winter (1980) has suggested that the light inactivation of P-enolpyruvate carboxylase in the CAM plant is actually due to malate inhibition, but this possibility seems to have been eliminated by experiments of Manetas and Gavalas (1983) in which the extracts were desalted by passage through Sephadex G-25 before activity was measured. F. LIGHT MODULATION OF THE ACTIVITY OF CYTOSOLIC ENZYMES

The mechanism of this process has received little attention and is not understood. Since DCMU inhibits light inactivation of pea leaf cytosolic glucose-6-P dehydrogenase (Anderson and Nehrlich, 1977) and activation

24

LOUISE E. ANDERSON

of P-enolpyruvate carboxylase in Salsola soda (Karabourniotis et al., 1983), the photochemical apparatus must be involved.

111. REGULATION OF LIGHT MODULATION Although a number of enzymes are light modulated the only evidence at present for target enzyme specificity is in the difference in activity observed with thioredoxin f and thioredoxin m in the ferredoxin-thioredoxin system. Yet there clearly is differential modulation of target enzyme activity in the intact chloroplast. How is this differential modulation achieved and how is light modulation controlled? A unique solution to these problems seems to have evolved in the green plant. Apparently the target enzymes, rather than the modifying enzymes, interact with modifiers. In some cases these might be acting by blocking the redox-active disulfide bond. In other cases they may be altering the conformation of the target enzyme, either making the redox-active disulfide inaccessible to the modifying enzyme (be it protein modulase, ferralterin, or reduced thioredoxin) or changing the redox potential of the active disulfide so that it can no longer be reduced, or, in dark modulation, oxidized. The effect of modulators on activation has been the subject of a number of investigations.

A.

NADP-LINKED MALIC

DEHYDROGENASE

NADP inhibits the DTT-thioredoxin-dependent activation of NADPlinked malic dehydrogenase and the thioredoxin-dependent inactivation of the enzyme (Ashton and Hatch, 1983~).The effect of NADP is reversed by NADPH, but NADPH itself is without effect. A model has been constructed to show the effects of varying NADPINADPH ratios on the steady state activity levels of the target enzyme (see Fig. 10). Although many assumptions have to be made, it is significant that the modulation system itself must be subject to control.'That NADP does indeed inhibit light activation of this enzyme has been shown by Scheibe and Jacquot (1983). In experiments with intact chloroplasts these workers, as well as Leegood and Walker (1983b), demonstrated that electron acceptors, which would increase the NADP/NADPH ratio, inhibit the activation of this dehydrogenase. The enzyme is activated in the dark in intact maize leaves under anaerobic conditions (Nakamoto and Edwards, 1983a, 1984b).

25

LIGHT/DARK MODULATION

0

0.5

1.0

M D P H (RIM)

Fig. 10. Simulated rates of thioredoxin-dependentactivation and inactivation of NADPlinked malic dehydrogenase activity at different NADPWNADP ratios. From Ashton and Hatch (1983~)with permission.

B. FRUCTOSE-BISPHOSPHATASE

The DTT-dependent activation of fructose-bisphosphatase is enhanced by fructose bisphosphate and Mg2+(Rosa, 1981; Rosa and Whatley, 1984). The DTT-thioredoxin-dependent activation of fructose-bisphosphatase also appears to be enhanced by fructose bisphosphate (Wolosiuk er al., 1980; Nishizawa and Buchanan, 1981; Hutcheson and Buchanan, 1983b). Here it is possible that the substrate protects the enzyme from inactivation by overreduction. Hertig and Wolosiuk (1980) and Hutcheson and Buchanan (1983b) report that Ca2+ and fructose bisphosphate, or Mn2+ and fructose bisphosphate, stimulate thioredoxin f-DTT-dependent activation of fructose-bisphosphatase. The Ao.5 for fructose-bisphosphatase is 0.33 mM, and, for Ca2+,55 p M (Hertig and Wolosiuk, 1980). These concentrations are reasonably close to assumed concentrations in uiuo and it can be concluded that these compounds may modulate activation in the green leaf. Sedoheptulose bisphosphate, glucose I ,Qbisphosphate, or ribulose bisphosphate can replace fructose bisphosphate in the Ca2+-and thioredoxin f-dependent DTT activation (Hertig and Wolosiuk, 1983). Rosa (1981) suggests that Ca2+may be acting by inhibiting the breakdown of the enzyme protector (and substrate) fructose bisphosphate. Whatever the biochemical explanation for the effect, these experiments suggest that both substrate and metal cations might affect modulation in uiuo and that

26

LOUISE E. ANDERSON

the two might act together. Gontero et al. (1984), however, find little or no effect of the carbon substrates and Mg2+on thioredoxin-dependent activation. The strict specificity for thioredoxin fin DTT-thioredoxin activation of fructose-bisphosphatase is lost in the presence of fructose bisphosphate (Schurmann, 1983). Light activation of this enzyme in broken pea leaf chloroplast preparations is inhibited by sedoheptulose bisphosphate, 3-Pglycerate, ribulose bisphosphate, and NAD (Anderson et al., 1981). Fructose bisphosphate is without effect. In spinach chloroplasts light activation of fructose-bisphosphatase is inhibited by anaerobiosis (Leegood et al., 1982). Light activation of NADP-linked malic dehydrogenase was not affected by anaerobiosis in the same experiments. These data indicate that differential regulation of light activation can occur in the intact chloroplast. According to Charles and Halliwell (1981~)addition of dihydroxyacetone-P to the suspension medium enhances activation of fructose-bisphosphatase in spinach chloroplasts. In intact wheat leaves at low light intensities activation of fructose-bisphosphataseis sensitive to atmospheric levels of 0 2 (Leegood and Walker, 1983a). This could be a reflection of the effect of metabolite levels on modulation or a direct effect of O2on the reductant required for activation.

C. SEDOHEPTULOSE-BISPHOSPHATASE

Inorganic phosphate apparently inhibits the DTT-dependent activation of sedoheptulose-bisphosphatase (Woodrow et al., 1983). DTT-thioredoxindependent activation of maize sedoheptulose-bisphosphataseis enhanced in the presence of sedoheptulose biphosphate, but fructose bisphosphate has no effect (Nishizawa and Buchanan, 1981). Ca2+enhances the DTTdependent activation of maize sedoheptulose-bisphosphatase and inhibits the activity of the enzyme (Wolosiuk et al., 1982). Ethylene glycol bis(paminoethy1 ether)N,N ,N’ ,N’-tetraacetate (EGTA), which chelates Ca2 , completely inhibits activation. Mg2+ and sedoheptulose bisphosphate (presumably both are required) enhance the DTT-dependent activation of wheat leaf sedoheptulose-bisphosphatase (Woodrow and Walker, 1982; Woodrow et al., 1983). Clearly the availability of metal cations, as well as substrate, could modulate light activation of both phosphatases in uiuo. Light activation of sedoheptulose-bisphosphatase in wheat chloroplasts is stimulated by the transportable metabolite dihydroxyacetone-P(Woodrow and Walker, 1980). Presumably the dihydroxyacetone-P is con+

LIGHT/DARK MODULATION

27

verted to sedoheptulose bisphosphate in the chloroplast which enhances activation of the enzyme. D . RIBULOSE-5-PHOSPHATE KINASE

Ashton (1983b) has examined the effects of ATP on DTT-dependent activation of maize ribulose-5-P kinase . Since ATP inhibits DTT-dependent activation and also inactivation by GSSG or alkylating reagents, the regulatory disulfide is probably located near the ATP binding site. Activation of this enzyme in broken pea leaf chloroplast preparations is inhibited by AMP, NADPH, NADH, and 6-P-gluconateand stimulated by NADP (Anderson et al., 1981). E. GLUCOSE-6-PHOSPHATE DEHYDROGENASE

The DTT-dependent inactivation of this enzyme in crude extracts of Anabaena uariabifis is reversed by the substrate, glucose-6-P (Cossar et al., 1984). Glucose-6-P, sedoheptulose bisphosphate, fructose bisphosphate, Pi, NAD, NADH, ADP, and NH4Cl inhibit light inactivation of this enzyme in broken pea leaf chloroplast preparations (Anderson et d . , 1981). In intact spinach chloroplasts light inactivation of glucose-6-Pdehydrogenase is inhibited by Pi, which causes transport of metabolites out of the chloroplast (Huber, 1979). Inhibition is reversed by P-glycerate. F. GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE

Activation of the NADP- and NAD-linked enzyme activities in broken chloroplast preparations is inhibited by ATP, ADP, NADPH, NADH, NAD, 6-P-gluconate, 3-P-glycerate, sedoheptulose bisphosphate, and fructose bisphosphate (Anderson et al., 1981). Activation of the NADPlinked activity (but not of the NAD-linked activity) is inhibited by AMP, NADP, fructose bisphosphate, and NH4Cl. Glucose-6-P inhibits activation of the NAD-linked activity, but not activation of the NADP-linked activity. The pH dependency and Mg2+requirements for activation and for inactivation are different for each light modulated enzyme (Anderson et al., 1981). It seems likely that these differences reflect direct influence on, or interaction with, the target enzyme rather than the enzymes of the modulating system. Stepwise and/or differential activation by a single modifying system may be made possible by the sensitivity of the target enzymes to effectors and to pH.

28

LOUISE E. ANDERSON

IV. DARK MODULATION Aerobic conditions appear to be required for dark modulation in uiuo (Nakamoto and Edwards, 1983~).Under anaerobic conditions reversal of dark modulation (mimicking light modulation) has been observed both in intact leaves (Nakamoto and Edwards; 1983c) and in chloroplasts (Leegood and Walker, 1981a). A . THIOREDOXIN IN DARK REVERSAL OF LIGHT MODULATION

Oxidized thioredoxin is required for dark modulation of NADP-linked malic dehydrogenase and glucose-6-P dehydrogenase in the reconstituted pea leaf chloroplast modulation system (Scheibe and Anderson, 1981). Fructose-bisphosphatase is rapidly inactivated in the dark following activation with highly purified components of the ferredoxin-thioredoxin system and illuminated thylakoid membranes (Schurmann, 1983). Oxidized thioredoxin f also has been shown to inactivate DTT-activated fructosebisphosphatase (Soulib et al., 1981) and sedoheptulose-bisphosphatase (Meunier et al., 1983; Gontero et al., 1984a). Antibody to E. coli thioredoxin, which inhibits the activity of purified pea leaf thioredoxin, inhibits dark modulation in broken chloroplast preparations (Scheibe and Anderson, 1981). Clearly thioredoxin is necessary for the reversal of light modulation. Kagawa and Hatch (1977) earlier showed that a small heat stable regulatory protein catalyzed the inactivation of NADP-linked malic dehydrogenase from maize. This factor was probably thioredoxin. The specificity of thioredoxins as dark mediators has not been determined. Scheibe (1981) has shown that levels of oxidized thioredoxin are high in the dark and decrease markedly in the light (Fig. Il), but it is not clear how reduced thioredoxin is oxidized in the chloroplast. The rate of inactivation of fructose-bisphosphatase is decreased if an 02 trap is included in the system. Oxidized ferredoxin or oxidized thioredoxin, added when the light is turned off, increases the rate of inactivation (Schiirmann, 1983; Schurmann and Kobayashi, 1984). Although oxidation of reduced thioredoxin is catalyzed by ferredoxin-NADP oxidoreductase, which transfers electrons from thioredoxin to NADP (SouliC et al., 1981), this reaction seems too slow to account for the rapid oxidation of reduced thioredoxin in intact chloroplasts. A cysteine oxidase which might function in dark modulation has been identified in a variety of photosynthetic organisms ranging from angiosperms to purple nonsulfur photosynthetic bacteria (Schmidt and Kramer, 1983, 1984). According to the scheme postulated by these workers cystine would oxidize either the light form of the target enzyme or thiore-

LIGHT~DARKMODULATION

29

Fig. I I. NADP-linked malic dehydrogenase activity (0)and level of oxidized thioredoxin From Scheibe (1981) with permission.

(m)in chloroplast suspensions upon light/dark/light transitions.

doxin. Since catalase inhibits dark modulation (Brennan and Anderson, 1980) (see also Charles and Halliwell, 1981c), there is also a possibility that peroxides are involved in the oxidation of thioredoxin. Figure 12 depicts a possible scheme for the thioredoxin-dependent dark reversal of light modulation. B. OTHER POSSIBLE MEDIATORS

Various naturally occuring oxidants including dehydroascorbate and oxidized glutathione will reverse reduced thioredoxin-mediatedenzyme activation and it has been suggested that they function as dark modulators

Fig. 12. Scheme for dark modulation. When the light goes out the target enzyme (TE) is in the reduced, light form and thioredoxin (Td) becomes oxidized (frame I). The enzyme passes its electrons to thioredoxin and changes conformation as it changes to the dark form (frame 2). Thioredoxin in turn finds an electron acceptor and returns to its active form as dark modulator (frame 3).

30

LOUISE E. ANDERSON

(Nishizawa and Buchanan, 1981; Schurmann and Wolosiuk, 1978, Wolosiuk and Buchanan, 1979). Since in all of these experiments oxidized thioredoxin would have been generated, it seems likely that the effect of these compounds was mediated by thioredoxin. Udvardy et al. (1984) have shown that various nonprotein Fe3+chelates mimic the effect of dark modulation with partially purified chloroplast and cyanobacterial enzymes. They note that this property is consistent with a role in dark modulation in uiuo. C. REGULATION OF DARK MODULATION

Dark modulation is subject to pH and metabolite control. Fructose bisphosphate, added to the ferredoxin-thioredoxin reductase activation system, accelerates light activation of fructose-bisphosphatase, and slows down or, at high concentrations, totally blocks dark inactivation at a pH approximating to that of the stroma in the illuminated chloroplast (pH 7.9) (Schurmann, 1983). At pH 7.2 (dark stromal pH) fructose bisphosphate has no effect on dark inactivation. The effect of a series of metabolites and of pH and Mg2+concentrations on dark modulation in broken chloroplast preparations has been reported (Anderson et al., 1981). As is the case for light modulation, no clear-cut effect of any single metabolite on dark modulation is evident. It therefore seems likely that the effects are mainly on the target enzyme which becomes more or less susceptible to dark modulation in the presence of the effector.

V. CHANGES IN THE TARGET ENZYME Light modulation probably involves a posttranslational covalent modification of proteins already modified by oxidation of cysteine residues. As yet there is no direct evidence for target enzyme reduction by the light modulation system. It is known that the conformation of the enzyme and its kinetic parameters are altered by light modulation and, except in the case of transaldolase (Anderson, 1981), by DTT-dependent reduction. A. CHANGE IN THE TARGET ENZYME AS A RESULT OF LIGHT MODULATION

1. Oxidation State

There are several reasons to suspect that light modulation is reductive and dark modulation is oxidative. (1) The effects of DTT and light modulation

LIGHT/DARK MODULATION

31

(Anderson, 1979) and of dark modulation and the strong oxidant diamide (Ashton and Anderson, 1981) on enzyme activity are similar. (2) Schiirmann (1983) has shown that reduced ferredoxin is oxidized in the presence of fructose-bisphosphatase, ferredoxin-thioredoxin reductase, and thioredoxin. (3) There is a marked DCMU-sensitive increase in total stroma1 thiol groups when spinach chloroplasts are irradiated (Slovacek and Vaughn, 1982; Slovacek and Monahan, 1984). The increase in the activity of fructose-bisphosphatase and NADP-linked malic dehydrogenase is essentially coincident with the increase in stromal thiol groups. (4) There is a significant increase in both soluble and membrane-bound thiols in an illuminated activation system consisting of thylakoids and partially purified ferredoxin-thioredoxin reductase, fedredoxin, thioredoxin, and NADP-linked malic dehydrogenase (Jacquot et a f . , 1983). An earlier suggestion (Anderson et al., 1978) that light modulation might result in intrapeptide thiol-disulfide exchange rather than in net reduction was based on the assumption, later shown to be incorrect, that no soluble intermediate was involved in light modulation. Nevertheless, modulation of the soluble chloroplast enzyme could involve thiol-disulfide exchange or some other alteration in enzyme structure rather than net disulfide reduction. Activation of the chloroplast coupling factor, CFI, by light and DTT involves reduction of a disulfide on the y subunit (Ketcham et al., 1984; Moroney et al., 1984; Nalin and McCarty, 1984). It is thought that in vivo activation is the result of light-dependent energization which makes the redox-active disulfide accessible to light-dependent reduction by the reducing component of the light modulation system. 2. Conformation There is good evidence that light modulation is accompanied by a change in conformation. Although the light and dark forms of pea leaf chloroplastic glyceraldehyde-3-P dehydrogenase are the same size, they differ in electrophoretic mobility (Anderson and Lim, 1972). This indicates that the exposed charged groups on the two forms differ. Light modulation involves a change in the susceptibility of four enzymes (fructose-bisphosphatase, glucose-6-P dehydrogenase, NADP-glyceraldehyde-3-P dehydrogenase, and ribulose-5-P kinase) to proteolytic degradation in crude extracts (Anderson et af., 1979) and, in the case of glucose-6-P dehydrogenase, in a highly purified system (Srivastava and Anderson, 1983). The exposure of protease-susceptible bonds is apparently altered as a result of the change in conformation associated with modulation. Differences in the thermal denaturation patterns of the light and dark forms of glucosedP dehydrogenase also suggest that light/dark modulation involves confor-

32

LOUISE E. ANDERSON

mational changes in the modulated proteins (Srivastava and Anderson, 1983). Glyceraldehyde-3-P dehydrogenase from some species forms aggregates (Wolosiuk and Buchanan, 1976; Cerff, 1978; Pupillo and Faggiani, 1979; Wara-Aswapati et al., 1980). The relationship of the aggregation state of the enzyme to light modulation is unclear, but it seems likely that it reflects the change in conformation which accompanies modulation. B. CHANGE IN THE TARGET ENZYME AS A RESULT OF THIOREDOXIN-DTT-CATALYZED MODULATION

Jacquot et al. (1984) report that two thiols per 38 kDa subunit appear when maize NADP-linked malic dehydrogenase is thioredoxin-DTT activated, but DTT treatment (48 hr) produces only 1.5 thiols per subunit. According to Scheibe (1984), two thiols per subunit appear when pea leaf NADP-liked malic dehydrogenase is thioredoxin-DTT or DTT activated. In contrast there is, according to Buchanan (1981), no change in the thiol content of fructose-bisphosphatase as a result of thioredoxin-DTT activation, but DTT activation results in an increase in 16 thiol groups. It should be noted that this enzyme reportedly has a very high Cys content (Buchanan et d . ,1971) and that it might be difficult to detect a change in one disulfide in such a protein. Pradel et al. (1981) report an increase of 24 sulfhydryl groups following activation of fructose-bisphosphatase by DTT. Wolosiuk and Buchanan (1976, 1979) have isolated DTT-thioredoxinsensitive and -insensitive forms of ribulose-5-P kinase and glyceraldehyde-3-P dehydrogenase. These apparently differ in molecular weight. The basis for the differences in aggregation and thioredoxin-DTT sensitivity has not been investigated. C. CHANGE IN THE TARGET ENZYME AS A RESULT OF DITHIOTHREITOL TREATMENT

There is an increase of two thiol groups per subunit when pea leaf NADPlinked malic dehydrogenase is reduced with DTT (Scheibe, 1984). Activity lags behind the appearance of the thiol groups (Fig. 13). Analyses of the kinetics of activation and enzyme reduction indicate that all four subunits of the tetramer must be reduced before the enzyme is activated. Activation can, however, also be achieved by treating the enzyme with guanidineaHC1, which would seem to indicate that activation results from a conformational change rather than from reduction per se (Scheibe and Fickenscher, 1985).

LIGHT/DARK MODULATION

I

I

I

0

10

2'0

33

30 6 0

Activation time (Mia)

Fig. 13. Appearance of thiol groups and activity during D'IT-dependent activation of NADP-linked malic dehydrogenase. From Scheibe (1984) with permission.

Fertk et al. (1984) find no free thiols on purified spinach NADP-linked malic dehydrogenase, and three per subunit after reduction. They suggest that at least one Cys per subunit is involved in an interprotomeric disulfide bond. Ashton (1983a) has used Procion Rubine MXB as an affinity label for maize NADP-linked malic dehydrogenase. The dye reacted both with a group in or near the active site and with a reactive thiol, presumably reduced when the enzyme is activated with DTT. Potentially the light and dark forms of the enzyme could be separated and the redox-active dithiol involved in light modulation could be identified with this dye. This is an exciting new approach. It is now important to determine whether the dye interacts with the light-activated form of the enzyme. In a similar set of experiments Ashton (1984) used Procion Red MX2B as an affinity label for the regulatory thiol of maize ribulose-5-P kinase. The DTT-activated form of the enzyme is inhibited by the dye and by sulfhydryl reagents while the nonactivated form is not. Since ADP and ATP inhibit the dye inactivation, the reactive thiol is probably positioned near the active site. Experiments with alkylating reagents give similar results (Omnaas et al., 1985). The ATP-protected thiol appears to be extraordinarily reactive. There is an increase of one thiol per subunit (four per mole enzyme) when maize P-enolpyruvate carboxylase is treated with DTT (Iglesias and Andreo , 1984). The kinetics of thermal denaturation of the DTT-inactivated glucose-6P dehydrogenase are the same as those of the dark form of the enzyme and distinct from those of the light form (Srivastava and Anderson, 1983).

34

LOUISE E. ANDERSON

Likewise proteolytic degradation of the DTT-inactivated enzyme resembles that of the dark form. Clearly the light form of this enzyme is not identical with the DTT-inactivated form. The DTT-activated form of fructosebisphosphatase appears to differ conformationally from the DTTthioredoxin-activated form, since the latter is much more sensitive to trypsin (Schurmann and Wolosiuk, 1978). Apparently the posttranslational modification produced by the light modulation system is not necessarily equivalent to the modification produced by DTT treatment alone or by DTT treatment in the presence of thioredoxin. The light modulation system is probably a considerably more specific reductant than is DTT or DTT- thioredoxin. D. CHANGE IN KINETIC PARAMETERS OF THE TARGET ENZYME

Shifts in pH dependency of pea leaf chloroplastic glucose-6-P dehydrogenase (Ben-Bassat and Anderson, 1981) , of spinach chloroplastic fructose-bisphosphatase (Slovacek and Vaughn, 1982), and of P-enolpyruvate carboxylase from Sedum praealtum (Manetas, 1982) and Mesembtyanthemum crystallinurn (Winter, 1980) as a result of light/dark modulation of activity have been observed. The dark forms of P-enolpyruvic carboxylase from Salsola soda (Karabourniotis et al., 1983) and pea leaf chloroplastic fructose-bisphosphatase (Marques and Anderson, 1985a) exhibit biphasic kinetics, while the kinetics of the light form are linear. This behavior suggests that there are two conformers of the enzyme and that the dark conformer is converted to the light conformer, which has a higher apparent affinity for substrate, when the plant is irradiated. This would be consistent with experiments indicating that light modulation effects conformational change (see above). An alternative explanation for biphasic kinetics, i.e., that there are two different carboxylases and two different phosphatases, cannot be ruled out since crude extracts were used in both sets of experiments. P-enolpyruvatecarboxylase in the CAM plant Sedum praealturn is light inactivated (Manetas, 1982; Manetas and Gavalas, 1983). The dark form of this enzyme has apparent Michaelis-Menten kinetics, while the light form shows positive cooperativity. The differences, also, are indicative of two conformers or of two enzymes. Both forms are inhibited by malate. For most of the enzymes which have been studied, maximal velocity rather than Km changes as a result of light modulation (Table IV). This may simply reflect the fact that most surveys for light-modulatable enzymes have been done with conventional assays containing excess substrate, in which case those enzymes responding to light modulation by a change in apparent affinity for substrate will escape detection.

35

LIGHT/DARK MODULATION

TABLE IV w e c t of Light Modulation on Kinetic Parameters of the Target Enzymes Change Enzyme"

Source

V,,

Km

Glyceraldehyde-3-P dehydrogenase

Pea leaves

Increase

None

Ribulose-5-P kinase

Spinach

Increase

None

Fructosebisphosphatase Glucose-6-P dehydrogenase Glucose-6-P dehydrogenase

Pea leaves

Increase

Decrease

Pea chloroplast Pea cytosol

Decrease

None

Decrease

None

Reference Melandri et al. (1970); Anderson and Duggan (1976) Gardemann er al. (1983) Marques and Anderson (1985a) Anderson and Duggan (1976) Anderson and Duggan (1976)

For changes in P-enolpyruvate carboxylase kinetic parameters, see text.

The effects of inhibitors on the activity of the light and dark forms of these enzymes have received almost no attention, possibly because of the complications that might arise in inhibitor experiments with crude extracts. Winter (1980) has shown that the light (less active) form of Penolpyruvate carboxylase from M.crysralliniurn is more sensitive to inh:bition by malate than is the dark form. Interestingly, the maize enzyme is activated to the same extent by either glucose-6-Por by DTT and glucose6-P (Iglesias and Andreo, 1984), which would seem to indicate that reduction and binding of the modifier glucose-6-Paffect the same group(s) in the active site. Ashton and Hatch (1983b) propose that light/dark conformational changes shift the equilibrium of the enzyme-substrate/product complex away from unity. (Efficient catalysis is thought to require that the equilibrium for the substrate/product complex be close to unity.) Such a change must be reflected in a change in substrate binding since the equilibrium constant for the overall reaction cannot be altered. Consistent with this proposal, marked differences in binding of NADP and NADPH to the DTT-active and -inactive forms of NADP-linked malic dehydrogenase are observed. Whether or not light/dark transitions are entirely identical with the DTT-induced changes in conformation, this is a very neat kinetic explanation for the effect of thiol modulation. Differences in the binding of Mg2+and fructose bisphosphate to the inactive and DTT-activated forms of fructose-bisphosphatase have been

36

LOUISE E. ANDERSON

reported by Meunier et al. (1981). Mixed cooperativity is observed with the inactive form. After DTT activation the substrate and the cation are bound with positive cooperativity. VI. THE FUNCTION OF LIGHT MODULATION A. INDUCTION OF PHOTOSYNTHETIC CARBON DIOXIDE FIXATION

There is considerable controversy about the exact function of light modulation in metabolic regulation in the green plant (Anderson et al., 1981, 1982, 1983; Charles and Halliwell, 1981a; Hiber, 1978, 1980; Leegood and Walker, 1981b; Robinson and Walker, 1980; Wirtz et al., 1982). Photosynthetic carbon dioxide fixation in intact chloroplasts is characterized by an initial lag period. This lag is thought to be related to the time required for the build-up of Calvin cycle intermediates when the chloroplast is illuminated. Arguments have been put forth for (Leegood and Walker, 1981b; Leegood et al., 1982; Anderson et al., 1980, 1981, 1982,1983;Heldt et al., 1981; Huber, 1978)and against (Charles and Halliwell, 1981a-c; Robinson and Walker, 1980; Leegood and Walker, 1980c) a causal relationship between light modulation and induction. Since light activation of the two phosphatases never follows induction, and since inhibition of light modulation prolongs induction (Anderson et al., 1983) and preactivation with DTT shortens induction time (Leegood and Walker, 1981b), it seems clear that the two phenomena are related. Leegood et al. (1982) point out that the great variation in induction times in intact chloroplasts is consistent with an interaction between autocatalysis and light activation. The increase in enzyme activity resulting from light activation must accelerate the build-up of reductive pentose phosphate cycle intermediates in the chloroplast. With each turn of the cycle, metabolite levels, and hence enzyme rates, will be increased. Without this acceleration the lag period will be extended to infinity. This is seen experimentally when chloroplasts are poisoned with arsenite (Gibbs and Calo, 1960; Marques and Anderson, 1985b). On the other hand, if carbon intermediates were added when light activation is blocked, carbon dioxide fixation might be expected to occur, but at a reduced rate, since some rate-limiting enzymes would not be fully active. This has been observed in experiments with arsenite-poisoned chloroplasts (Bamberger and Gibbs, 1965). It would seem then that light activation is involved in the autocatalytic build up of carbon intermediates in the chloroplast. It still remains to be shown experimentally that light activation is required for the build-up of photo-

LIGHT/DARK MODULATION

37

synthetic intermediates in the light and that this build-up is related to induction. A preliminary computer simulation of the reductive pentose phosphate cycle is consistent with this hypothesis (J. B. Anderson and L. E. Anderson, unpublished). The availability of ATP and NADPH, and the light-dependent rise in stromal pH and increase in stromal Mg2+ concentration, will also undoubtedly be involved, along with light modulation, in determining the extent of the lag phase. Experiments with intact maize leaves indicate that light activation is not rate limiting during induction (Usuda et al., 1984). Computer simulation probably represents the only means of estimating the relative contribution of each of these factors (metabolite levels, pH, cation concentrations, light modulation) in the regulation of the activity of the reductive pentose phosphate pathway in the intact chloroplast. Woodrow et al. (1984)have analyzed the effects of reductive activation and of pH and Mg2+ concentrations on the activity of sedoheptulosebisphosphatase. They conclude that in the dark less than 1% of the enzyme molecules will be in the active form. Heldt and co-workers (Gardemann et al., 1983) have measured ribulose-5-P kinase activity levels under conditions which simulate a light/ dark transition. In the presence of metabolites the effect of light activation is dramatically increased. Clearly, modulation of the activity of the light and dark forms of the enzyme is important in chloroplast metabolism in the intact leaf. B.

CFI-CFo Mg ATPASE

The thiol-related light activatioddark inactivation of CFI-CFo is thought to allow rapid responses to illumination and darkness. CFI-CFo is also pH activated, but presumably pH changes in the chloroplast would be slow compared to redox thiol changes (Mills and Mitchell, 1982b). C. STOMATAL OPENING

Three epidermal enzymes which are thought to be involved in metabolism related to stomatal movement, P-enolpyruvate carboxylase, NADPlinked malic dehydrogenase, and NADP-linked isocitric dehydrogenase, are light activated, and one, aspartate amino transferase, is light inactivated. Arsenite and sulfite inhibit light-dependent stomatal opening and light activation of the epidermal enzymes (Rao and Anderson, 1983b). There is an increase in the measureable free thiols when epidermal strips are irradiated which appears to parallel stomatal opening (Rao and Anderson, 1983a). The thiol content of the epidermis is decreased when intact

38

LOUISE E. ANDERSON

plants are exposed to SO2in the light (Rao et al., 1983). Apparently light modulation of the activity of these epidermal enzymes is involved in stomatal opening. VII. EFFECT OF SULFUR DIOXIDE ON LIGHT MODULATION The extreme sensitivity of the LEM to sulfite (Anderson and Avron, 1976) and the fact that plants, and not animals, are SO2 sensitive suggests that light modulation might be the primary process affected by SO2 pollution. In support of this hypothesis it was found that treatment of intact seedlings with SO2 results in inhibition of the system for light inactivation of glucose-6-P dehydrogenase (Anderson and Duggan, 1977). Apparently, inactivation of the light modulation system can and does occur in uiuo when the plant is exposed to SO*. Prolonged exposure to low levels of SO2 almost certainly will disrupt the regulatory system for control of carbon metabolism by light, leading to metabolic chaos and eventually to death or extreme damage to the plant. Recently Alscher (1982a, 1983) has found that light activation of fructose-bisphosphatase in an SO2-sensitivesoybean cultivar is sulfite sensitive, while light activation in an SO2-insensitivecultivar is not affected by sulfite. Muschinek et al. (1985) report that the thylakoids in a more SOZsensitive variety of Pisum is more sensitive to S032-(and to AsOz-) than in a less SO2-sensitivevariety. These experiments suggest that SO2 susceptibility may be related to light modulation. These experiments were done with crude broken chloroplast preparations and with plants fumigated in growth chambers and in the field. They indicate that SO2 susceptibility is related to light modulation. What is not yet known is how the light activation system differs in the two varieties. VIII. OSMOTIC STRESS AND LIGHT MODULATION Even though light modulation was first observed in dehydrated specimens of a resurrection plant (Ziegler and Ziegler, 1965), light activation of fructose-bisphosphatase and sedoheptulose-bisphosphataseis severely inhibited when isolated spinach chloroplasts are subjected to osmotic stress (Boag and Portis, 1984). Metabolite levels indicate that the light-activated enzyme fructose-bisphosphatase does not function normally in stressed chloroplasts (Berkowitz and Gibbs, 1982). The effects of osmotic stress and desiccation on light modulation in resurrection plants have not been studied.

LIGHT/DARK MODULATION

39

IX. CONCLUSIONS Lighddark modulation is an important regulatory process in the green plant. Work in progress in a number of laboratories should soon lead to a resolution of the differences in the proposed mechanisms for modulation, to positive proof that the result of modulation is posttranslational modification and that the modification catalyzed by the LEM system or the ferredoxin-thioredoxin system involves reduction of a disulfide bond, and to a better understanding of the function of modulation in photosynthetic carbon metabolism. There will probably be more surprises for those who, like Irmgard and Hubert Ziegler, have the good luck to do the right experiment and the wit to interpret it correctly.

ACKNOWLEDGMENTS Work in the author’s laboratory was supported by grants from the United States National Science Foundation, Department of Agriculture, and Department of Energy. Myhanh Hoang and Loretta Moore helped with the processing of the manuscript. I thank Drs. Peter Jablonski, Ivan0 Marques, Gyorgi Muschinek, and Duane Ford for critical reading of the manuscript.

REFERENCES Alscher, R. (1983). I n “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 154-163. University of Bayreuth, Bayreuth. Alscher-Herman, R. (1982a). Enuiron. Pollut. 27, 83-96. Alscher-Herman, R. (1982b). Plant Physiol. 70, 728-734. Anderson, L. E. (1974). Biochem. Biophys. Res. Commun. 59,907-913. Anderson, L. E. (1979). I n “Encyclopedia of Plant Physiology, New Series” (M. Gibbs and E. Latzko, Eds.), Vol. 6, pp. 271-281. Springer-Verlag, Berlin. Anderson, L. E. (1981). Biochem. Biophys. Res. Commun. 99, 1199-1202. Anderson, L. E. (1983a). I n “Molecular Models of Photoresponsiveness” (G. Montagnoli and B. F. Erlanger, eds.), pp. 153-173. Plenum, New York. Anderson, L. E. (1983b). I n “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 47-52. University of Bayreuth, Bayreuth. Anderson, L. E., and Avron, M. (1976). Plant Physiol. 57, 209-213. Anderson, L. E., and Ben-Bassat, D. (1981). In “Photosynthesis IV. Regulation of Carbon Metabolism” (G. Akoyunoglou, ed.), pp. 373-378. Balaban International Science Services, Philadelphia. Anderson, L. E., and Duggan, J. X . (1976). Plant Physiol. 58, 135-139. Anderson, L. E., and Duggan, J. X . (1977). Oecologia 28, 147-151. Anderson, L. E.,and Lim, T.-C. (1972). FEES Lett. 27, 189-191. Anderson, L. E., and Nehrlich, S. C. (1977). FEBS Lett. 76,64-66.

40

LOUISE E. ANDERSON

Anderson, L. E., Nehrlich, S. C., and Champigny, M.-L. (1978). Plant Physiol. 61,601-605. Anderson, L. E., Chin, H.-M., and Gupta, V. K. (1979). Plant Physiol. 64, 491-494. Anderson, L. E., Ashton, A. R., Ben-Bassat, D., Mohamed, A. H., and Scheibe, R. (1980). What’s New Plant Physiol. 11, 34-40. Anderson, L. E., Mohamed, A. H., Ashton, A. R., Scheibe, R., Brennan, T., and BenBassat, D. (1981). In “Photosynthesis IV. Regulation of Carbon Metabolism” (G. Akoyunoglou, ed.), pp. 293-302. Balaban International Science Services, Philadelphia. Anderson, L. E., Ashton, A. R., Mohamed, A. H., and Scheibe, R. (1982). BioScience 32, 103-107. Anderson, L. E., Mohamed, A. H., Ashton, A. R., Scheibe, R., and Duraini, M. 0. (1983). In “Thiorbdoxines Structure et Fonctions” (P. Gadal, ed.), pp. 269-276. CNRS, Paris. Anderson, L. E., Jablonski, P., and Mohamed, A. H. (1985). In “Thioredoxin and Glutaredoxin Systems: Structure and Function” (A. Holmgren, C. I. Brandbn, H. Jisrnvall, and B-M. Sjoberg, eds.). Raven Press, New York. In press. Ashton, A. R. (1983a). Biochim. Biophys. Acta 749, 24-31. Ashton, A. R. (1983b). In “Thiorkdoxines Structure et Fonctions” (P. Gadal, ed.), pp. 245250. CNRS, Paris. Ashton, A. R. (1984). Biochem. J . 217, 79-84. Ashton, A. R., and Anderson, L. E. (1981). Biochim. Biophys. Acta 639, 242-249. Ashton, A. R., and Hatch, M. D. (1983a). Biochem. Biophys. Res. Commun. 115, 53-60. Ashton, A. R., and Hatch, M. D. (198313). Arch. Biochem. Biophys. 227,406-415. Ashton, A. R., and Hatch, M. D. (1983~).Arch. Biochem. Biophys. 227,416-424. Ashton, A. R., and Hatch, M. D. (1984). In “Advances in Photosynthesis Research” (C. Sybesma, ed.), VoI. 3, pp. 613-616. Martinus NijhoffIDr. W.Junk Publ., The Hague. Ashton, A. R., Brennan, T., and Anderson, L. E. (1980). Plant Physiol. 66, 605-608. Ashton, A. R., Burnell, J. N., and Hatch, M. D. (1984). Arch. Biochem. Biophys. 230,492503. Avron, M., and Gibbs, M. (1974). Plant Physiol. 53, 136-139. Balange, A. P., and Lambert C. (1983). Plant Sci. Lett. 32, 253-259. Barnberger, E. S., and Gibbs, M. (1965). Plant Physiol. 40, 919-926. Ben-Bassat, D., and Anderson, L. E. (1981). Plant Physiol. 68, 279-283. Bennett, J. (1983). In “Molecular Models of Photoresponsiveness” (G. Montagnoli and B. F. Erlanger, eds.), pp. 175-196. Plenum, New York. Berkowitz, G. A., and Gibbs, M. (1982). Plant Physiol. 70, 1535-1540. Berstermann, A., Vogt, K.,and Follmann, H. (1983). In “Thiorkdoxines Structure et Fonctions” (P. Gadal, ed.), pp. 31-39. CNRS, Paris. Bjisrkman, O., and Badger, M. (1977). Annual Report of the Director. Depf. Plant Biol. Carnegie Inst. 1976-1977 pp. 346-354. Boag, S . , and Portis, A. (1984). Planta 160, 33-40. Brennan, T., and Anderson, L. E. (1980). Plant Physiol. 66, 815-817. Buc, J., Riviere, M., and Meunier, J. C. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 28-37. University of Bayreuth, Bayreuth. Buc, J., Riviere, M., and Meunier, J. C. (1984). In “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 541-544. Martinus Nijhoff/Dr. W. Junk Publ., The Hague. Buchanan, B. B. (1980a). Annu. Rev. Plant Physiol. 31, 341-374. Buchanan, B. B. (1980b). In “The Enzymology of Post-Translational Modification of Proteins” (R. G. Freedman and H. C. Hawkins, eds.), Vol. 1, pp. 345-362. Academic Press, New York. Buchanan, B. B. (1981). In “Photosynthesis IV. Regulation of Carbon Metabolism” (G. Akoyunoglou, ed.), pp. 245-256. Balaban International Science Services, Philadelphia.

LIGHT/DARK MODULATION

41

Buchanan, B. B. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 1-10. University of Bayreuth, Bayreuth. Buchanan, B. B. (1984a). BioScience 34,378-383. Buchanan, B. B. (1984b). In “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 573-580. Martinus Nijhoff/Dr. W. Junk Publ., The Hague. Buchanan, B. B., Kalberer, P. P., and Arnon, D. I. (1967). Biochem. Biophys. Res. Commun. 29, 74-79. Buchanan, B. B., Schiirmann, P., and Kalberer, P. P. (1971). J. Biol. Chem. 246,5952-5959. Buchanan, B. B., Crawford, N. A., and Wolosiuk, R. A. (1978). Plant Sci. Lett. 12, 257264. Buchanan, B. B., Wolosiuk, R. A., and Schiirmann, P. (1979a). What’s New Plant Physiol. 10, 1-4. Buchanan, B. B., Wolosiuk, R. A., and Schiirmann, P. (1979b). Trends Biochem. Sci. 4,9396. Burnell, J. N. (1984). Biochem. Biophys. Res. Commun. 120,559-565. Burnell, J. N., and Hatch, M. D. (1983). Biochem. Biophys. Res. Commun. lll, 288-293. Burnell, J. N., and Hatch, M. D. (1984a). Biochem. Biophys. Res. Commun. 118, 65-72. Burnell, J. N., and Hatch, M. D. (1984b). Arch. Biochem. Biophys. 231, 175-182. Cao, R. Q., Johnson, T. C., Kung, J. E., and Buchanan, B. B. (1984). In “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 4, pp. 877-880. Martinus NijhoW Dr. W.Junk Publ., The Hague. C e s , R. (1978). Eur. J. Biochem. 82,45-53. Charnpigny, M. L., and Bismuth, E. (1976). Physiol. Plant. 36, 95-100. Chapman, K. S.R., and Hatch, M. D. (1981). Arch. Biochem. Biophys. 210, 82-89. Charles, S. A., and Halliwell, B. (1981a). Biochem. J. 200, 357-363. Charles, S. A., and Halliwell, B. (1981b). Planta 151, 242-246. Charles, S. A., and Halliwell, B. (1981~).Cell Calcium 2, 211-224. Chastain, C. J., and Open, W. L. (1985). Plant Physiol. 77,851-856. Clement-Metral, J. D. (1979). FEBS Lett. 101, 116-120. Clement-Metral, J. D., and Holmgren, A. (1983). In “Thiorkdoxines Structure et Fonctions” (P. Gadal, ed.), pp. 59-68. CNRS, Paris. Cossar, J. D., Rowell, P., and Stewart, W. D. P. (1984). J. Gen. Microbiol. WO, 991-998. Crawford, N. A., Yee, B. C., Nishizawa, A. N., and Buchanan, B. B. (1979). FEBS Lett. 104, 141-145. Crawford, N. A., Sutton, C. W., Yee, B. C., Carlson, D. C., andBuchanan, B. B. (1983a). I n “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 38-42. University of Bayreuth, Bayreuth. Crawford, N. A., Yee, B. C., de la Torre, A., Lara, C., Carlson, D. E., and Buchanan, B. B. (1983b). I n “Thioredoxines Structure et Fonctions” (P. Gadal, ed.), pp. 191-198. CNRS, Pans. Crawford, N. A., Sutton, C. W., Yee, B. C., Johnson, T. C., Carlson, D. C., and Buchanan, B. B. (1984). Arch. Microbiol. l39, 124-129. Cullimore, J. V . (1981). Planta 152, 587-591. de la Torre, A., Lara, C., Wolosiuk, R. A., and Buchanan, B. B. (1979). FEBS Lett. 107, 141-145. de la Torre, A., Lara, C., Yee, B. C., Malkin, R., and Buchanan, B. B. (1982). Arch Biochem. Biophys. 2l3,545-550. Duggan, J. X . , and Anderson, L. E. (1975). Planta 122, 293-297. Droux, M., Jacquot, J., Suzuki,A., and Gadal, P. (1984). In “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 533-536. Martinus Nijhoff/Dr. W. JunkPubl., The Hague.

LOUISE E. ANDERSON

42

Fertb, N., Meunier, J.-C., Suave, P., and Ricard, J. (1984). Plant Sci. Lett. 35, 175-179. Gardemann, A., Stitt, M., and Heldt, H. W. (1983). Biochim. Biophys. Acta 722, 51-60. Gibbs, M.,and Calo, N. (1960). Biochim. Biophys. Acta 44, 341-347. Gleason, F. K. (1985). I n “Thioredoxin and Glutaredoxin Systems: Structure and Function” (A. Holmgren, C. I. Banden, H. Jornvall, and B.-M. Sjoberg, eds.). Raven Press, New York. In press. Gleason, F. K., and Holmgren, A. (1981). J. Biol. Chem. 256, 8306-8309. Gleason, F. K., and Holmgren, A. (1983). I n “Thioredoxines Structure et Fonctions” (P. Gladal, ed.), pp. 41-47. CNRS, Paris. Gleason, F. K., Whittaker, M. M., Holmgren, A., and Jornvall, H. (1985). J . Biochem., in press. Gontero, B., Meunier, J.-C., and Buc, J. (1984a). I n “Advances in Photosynthesis Research,” (C. Sybesma, ed.), Vol. 3, pp. 505-508. Martinus Nijhoff/Dr. W. Junk Publishers, The Hague. Gontero, B., Meunier, J.-C., and Ricard, J. (1984b). Plant. Sci. Lett. 36, 195-199. Gupta, V. K., and Anderson, I. E. (1978). Plant Physiol. 61, 469-471. Gupta, V. K., and Anderson, L. E. (1980). J. Exp. Biol. 18, 983-984. Halliwell, B., and Foyer, C. H. (1978). Planta U9, 9-17. Hammel, K. E., Cornwell, K. L., and Buchanan, B. B. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 3681-3685. Hatch, M.D. (1977). Photosynthetic organelles. Plant Cell Physiol., Special Issue 31 1-314. Hatch, M. D., and Slack, C. R. (1969). Biochem. J . 1l2, 549-558. Heldt, H. W., Laing, W., Lorimer, G. H., Stitt, M., and Wirtz, W. (1981). I n “Photosynthesis IV. Regulation of Carbon Metabolism” (G. Akoyunoglou, ed.), pp. 213-226. Balaban International Science Services, Philadelphia. Hertig, C., and Wolosiuk, R. A. (1980). Biochem. Biophys. Res. Commun. W ,325-333. Hertig, C., and Wolosiuk, R. A. (1983). J. Biol Chem. 258,984-989. Heuer, B., Hansen, M. J., and Anderson, L. E. (1982). Plant Physiol. 69, 1404-1406. Holmgren, A. (1981). Curr. ‘fop. Cell. Regul. 19,47-76. Huber, S. C. (1978). FEBS Lett. 92, 12-16. Huber, S. C. (1979). Plant Physiol. 64, 846-851. Huber, S. C. (1980). Plantu 149, 485-492. Hutcheson, S. W., and Buchanan, B. B. (1983a). Plant Physiol. 72,870-876. Hutcheson, S . W., and Buchanan, B. B. (1983b). Plant Physiol. 72,877-885. Hutcheson, S . W., Crawford, N. A., Buchanan, B. B., and Jacquot, J.-P. (1983). I n “Thiorbdoxines Structure et Fonctions” (P. Gadal, ed.), pp. 235-243. CNRS, Paris. Iglesias, A. A., and Andreo, C. S. (1984). Plant Physiol. 75, 983-987. Ip, S.-M., Rowell, P., Aitken, A., and Stewart, W. D. P. (1984). Eur. J . Biochem. 141,497504.

Jacquot, J. P. (1984). Physiol. Veg. 22,487-507. Jacquot, J. P., and Gadal, P. (1983). I n “Thioredoxines Structure et Fonctions” (P. Gadal, ed.), pp. 221-233. CNRS, Paris. Jacquot, J.-P., Vidal, J., Gadal, P., and Schiirmann, P. (1978). FEBS Leu. 96, 243-246. Jacquot, J. P., Droux, M., Miginiac-Maslow, M., and Gadal, P. (1983). I n “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 19-27. University of Bayreuth, Bayreuth. Jacquot, J.-P., Gadal, P., Nishizawa, A. N., Yee, B. C., Crawford, N. A., and Buchanan, B. B. (1984). Arch. Biochem. Biophys. 228, 170-178. Jenkins, G. I., Gallagher, T. F., Hartley, M. R., Bennett, J., and Ellis, R. J. (1984). I n “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 4, pp. 863-872. Martinus Nijhoff/Dr. W. Junk Publ., The Hague.

LIGHT~DARKMODULATION

43

Johnson, H. S. (1971). Biochem. Biophys. Res. Commun. 43, 703-709. Kachru, R. B., and Anderson, L. E. (1975). Plant Physiol. 55, 199-202. Kagawa, T., and Hatch, M. D. (1977). Arch. Biochem. Biophys. 184, 290-297. Karabourniotis, G., Manetas, Y.,and Gavalas N. A. (1983). Plant Physiol. 73, 735-739. Karabourniotis, G., Manetas, Y.,and Gavalas, N. A. (1985). Plant Physiol. 77,300-302. Ketcham, S. R., Davenport, J. W., Warncke, K., and McCarty R. E. (1984). J. Biol. Chem. 259,7286-7293. Kleczkowski, L. A., and Randall, D. D. (1985). Plant Physiol. 77, S-496. Kow,Y.W., Smyth, D. A., and Gibbs, M. (1982). Plant Physiol. 69,72-76. Laing, W. A., Stitt, M., and Heldt, H. W. (1981). Biochim. Biophys. Acta 637, 348-359. Lam, C., de la Torre, A., and Buchanan, B. B. (1980). Biochem. Biophys. Res. Commun. 93, 544-551. Lara, C., de la Torre, A., Malkin, R., and Buchanan, B. B. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 53-59. University of Bayreuth, Bayreuth. Latzko, E., v. Garnier, R., and Gibbs, M. (1970). Biochem. Biophys. Res. Commun. 39, 1140-1144. Leegood, R. C., and Walker, D. A. (1980a). FEBS Lett. 116, 21-24. Leegood, R. C., and Walker, D. A. (1980b). Biochim. Biophys. Acta 593,362-370. Leegood, R. C., and Walker, D. A. (1980~).Arch. Biochem. Biophys. 200,575-582. Leegood, R. C., and Walker, D. A. (1981a). Arch. Biochem. Biophys. 212,644-650. Leegood, R. C., and Walker, D. A. (1981b). Plant Cell Environ. 4, 59-66. Leegood, R. C., and Walker, D. A. (1983a). Planta 156,449-456. Leegood, R. C., and Walker, D. A. (1983b). Plant Physiol. 71, 513-518. Leegood, R. C., Kobayashi, Y.,Meimanis, S., Walker, D. A., and Heber, U. (1982). Biochim. Biophys. Acta 682, 168-176. Lendzian, K. J. (1978). Planta 141, 105-110. Lendzian, K. J. (1980). Planta 148, 1-6. Lendzian, K., and Bassham, J. A. (1975). Biochim. Biophys. Acta 396, 260-275. Lendzian, K., and Ziegler, H. (1970). Planta 94, 27-36. Losada, M. (1976). I n “Reflections on Biochemistry” (A. Kornberg, B. L. Horecker, L. Cordudella, and J. Oro, eds.), pp. 73-74. Pergamon, Oxford. Manetas, Y . (1982). Photosyn. Res. 3, 321-333. Manetas, Y.,and Gavalas, N. A. (1983). Plant Physiol. 71, 187-189. Manetas, Y.,Karabourniotis, G., Selinioti, E., and Gavalas, N. A. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 106-114. University of Bayreuth, Bayreuth. Marques, I. A., and Anderson, L. E. (1985a). Plant Physiol. 77,807-810. Marques, I. A., and Anderson, L. E. (1985b). Plant Physiol. 77,S-235. Melandri, B. A., Pupillo, P., and Baccarini-Melandri, A. (1970). Biochim. Biophys. Acta 220, 178-189. Meunier, J. C., Buc, J., Soulib, J.-M., Pradel, J., and Ricard, J. (1981). Eur. J. Biochem. 1l3, 513-520. Meunier, J. C., Gontero, B., Ferte, N., and Buc, J. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 60-68. University of Bayreuth, Bayreuth. Mills, J. D. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 85-88. University of Bayreuth, Bayreuth. Mills, J. D., and Mitchell, P. (1982a). Biochim. Biophys. Acta 679, 75-83. Mills, J. D., and Mitchell, P. (1982b). FEBS Lett. 144, 63-67. Miziorko, H. M., and Lorimer, G. H. (1983). Annu. Rev. Biochern. 52, 507-535.

44

LOUISE E. ANDERSON

Mohamed A. H., and Anderson, L. E. (1981a). Arch. Biochem. Biophys. 209,606-612. Mohamed A. H., and Anderson, L. E. (1981b). In “Photosynthesis IV. Regulation of Carbon Metabolism” (G. Akoyunoglou, ed.), pp. 379-388. Balaban International Science Services, Philadelphia. Moroney, J. V.,Fullmer, C. S., and McCarty, R. E. (1984). J. Biol. Chem. 259,7281-7285. Miiller, B. (1970). Biochim. Biophys. Acta 205, 102-109. Miiller, B., Ziegler, I., and Ziegler, H. (1969). Eur. J . Biochem. 9, 101-106. Muschinek, G., Anderson, L. E., and Alscher, R. (1985). Plant Physiol. 77, S-761. Muto, S., Miyachi, S., Usuda, H., Edwards, G. E., and Bassham, J. A. (1981). Plant Physiol. 68, 324-328. MvC Akamba, L., and Anderson, L. E. (1981). Plant Physiol. 67, 197-200. Nakamoto, H., and Edwards, G. E. (1983a). Plant Sci. Lett. 32, 139-146. Nakamoto, H., and Edwards, G. E. (1983b). Biochem. Biophys. Res. Commun. 115, 673679. Nakamoto, H., and Edwards, G. E. (1983~).Plant. Physiol. 71, 568-573. Nakamoto, H., and Edwards, G. E. (1984a).I n “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 601-604. Martinus Nijhoff/Dr. W. Junk Publ., The Hague. Nakamoto, H., and Edwards, G. E. (1984b). Z. Pflanzenphysiol. 114, S . 315-320. Nakamoto, H., and Sugiyama, T . (1982). Plant Physiol. 69, 749-753. Nakamura, Y.,and Yamada, M. (1979). Plant Sci. Lett. 14, 291-295. Nalin, C. M., and McCarty, R. E. (1984). J . Biol. Chem. 259, 7275-7280. Nishizawa, A. N., and Buchanan, B. B. (1981). J . Biol. Chem. 256, 6119-6126. Nishizawa, A. N., Wolosiuk, R. A., and Buchanan, B. B. (1979). Planta 145, 7-12. Omnaas, J., Porter, M. A., and Hartman, F. C. (1985). Arch. Biochem. Biophys. 236,646653. Pla, A., and Lopez-Gorge, J. (1981). Biochim. Biophys. Acta 636, 113-118. Portis, A. R., Jr. (1985). Plant Physiol. 77,S-594. Pradel, J., SouliC, J.-M., Buc, J., Meunier, J.-C., and Ricard, J. (1981). Eur. J. Biochem. lW,507-511. Pupillo, P., and Faggiani, R. (1979). Arch. Biochem. Biophys. 194, 581-592. Rao, I. M., and Anderson, L. E. (1983a). Plant Physiol. 71, 451-455. Rao, 1. M., and Anderson, L. E. (1983b). Plant Physiol. 71,456-459. Rao, I. M., Amundson, R. G., Alscher-Herman, R., and Anderson, L. E. (1983). Plant Physiol. 72, 573-577. Robinson, S. P., and Walker, D. A. (1980). Arch. Biochem. Biophys. 202, 617-623. Rosa, L. (1981). FEBS Lett. W, 151-154. Rosa, L., and Whatley, F. R. (1984). Plant Physiol. 75, 131-137. Roughan, P. G., Kagawa T., and Beevers, H. (1980). Plant Sci. Lett. 18, 221-228. Rowell, P., Sampaio, M. J. A., Ladha, J. K., and Stewart, W. D. P. (1979). Arch. Microbiol. 120, 195-200. Rudd, T. P., and Anderson, L. E. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 98-105. University of Bayreuth, Bayreuth. Salvucci, M. E., Portis, A. R., and Ogren, W. L. (1985). Plant Physiol. 77, S-591. Scheibe, R. (1981). FEBS Letr. 133, 301-304. Scheibe, R. (Ed.). (1983a). “Light-Dark Modulation of Plant Enzyme Activity.” University of Bayreuth, Bayreuth. Scheibe, R. (1983b). In “ThiorCdoxines Structure et Fonctions” (P. Gadal, ed.), pp. 205211. CNRS, Paris. Scheibe, R. (1984). Biochim. Biophys. Acta 788, 241-247. Scheibe, R., and Anderson, L. E. (1981). Biochim. Biophys. Acta 636, 58-64.

LIGHT/DARK MODULATION

45

Scheibe, R., and Beck, E. (1979). Plant Physiol. 64, 744-748. Scheibe, R., and Fickenscher, K. (1985). FEES Lett. 180, 317-320. Scheibe, R., and Jacquot, J.-P. (1983). PIanta 157, 548-553. Schmidt, A. (1980). Arch. Microbiol. 127, 259-265. Schmidt, A. (1981). I n “Biology of Inorganic Nitrogen and Sulfur” (H. Bothe and A. Trebst, eds), pp. 327-333. Springer-Verlag, Berlin. Schmidt, A., and Kramer, E. (1983). I n “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 115-123. University of Bayreuth, Bayreuth. Schmidt, A., and Kramer, E. (1984). I n “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 525-528. Martinus Nijhoff/Dr. W. Junk Publ., The Hague. Schmidt-Clausen, H. J., Ziegler, I., and Ziegler, H. (1969). Planta 86, 272-285. Schriek. U., and Schwenn, J. (1984). I n “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 705-708. Martinus Nijhoff/Dr. W. Junk Publ., The Hague. Schiirmann, P. (1981). In “Photosynthesis IV. Regulation of Carbon Metabolism” (G. Akoyunoglou, ed.), pp. 273-280. Balaban International Science Services, Philadelphia. Schiirmann, P. (1983). In “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 11-18. University of Bayreuth, Bayreuth. Schiirmann, P., and Kobayashi, Y. (1984). I n “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 629-632. Martinus Nijhoff/Dr. W. Junk Publ., The Hague. Schiirmann, P., and Wolosiuk, R. A. (1978). Biochim. Biophys. Acta 522, 130-138. Schiirmann, P., Wolosiuk, Breazeale, V. D., and Buchanan, B. B. (1976). Nature (London) 263, 257-258. Schiirmann, P., Maeda, K., and Tsugita, A. (1981). Eur. J . Biochem. 116, 37-45. Schiirmann, P., Tsugita, A., Maeda, K., Dalzoppo, D., and Vilbois, F. (1985). “Thioredoxin and Glutaredoxin Systems: Structure and Function” (A. Holmgren, C. 1. Brandtn, H. Jornvall, and B.-M. Sjoberg, eds.). Raven Press, New York, in press. Seeman, J. R., and Berry, J. A. (1985). Plant Physiol. 77, S-624. Slovacek, R., and Monahan, B. (1984). I n “Advances in Photosynthesis Research” (C. Sybesma, ed.), Vol. 3, pp. 713-716. Martinus Nijhoff/Dr. W. Junk Publ., The Hague. Slovacek, R. E., and Vaughn, S. (1982). Plant Physiol. 70,978-981. Soulie, J.-M., Buc, J., Meunier, J.-C., Pradel, J., and Ricard, J. (1981). Eur. J. Biochem. 119,497-502. Srivastava, D. K., and Anderson, L. E. (1983). Biochim. Biophys. Acta 724, 359-369. Steiger, E., Ziegler, I., and Ziegler, H. (1971). Planta 96, 109-118. Steinmiiller, K., and Zetsche, K. (1984). Plant Physiol. 76, 935-939. Sugiyama, T., and Hatch, M. D. (1981). Plant Cell Physiol. 22, 115-126. Suske, G., Wagner, W., and Follmann, H. (1979). Z. Nuturforsch. 34c,214-221. Tischner, R. (1983). I n “Light-Dark Modulation of Plant Enzyme Activity” (R. Scheibe, ed.), pp. 129-138. University of Bayreuth, Bayreuth. Tischner, R. and Huttermann, A. (1980). Plant Physiol. 66, 805-808. Tsugita, A., Maeda, K., and Schiirmann, P. (1983). Biochem. Biophys. Res. Commun. 115, 1-7. Udvardy, J., Godeh, M. M., Cseke, C., and Farkas, G. L. (1983). I n “Thiortdoxines Structure et Fonctions” (P. Gadal, ed.), pp. 199-204. CNRS, Paris. Udvardy, J., Borbely, G., JuhBsz, A., and Farkas, G. L. (1984). FEES Lett. 172, 11-16. Usuda, H., Ku, M.S. B., and Edwards, G. E. (1984). Plant Physiol. 76, 238-243. Vallejos, R. H., Arana, J. L., and Ravizzini, R. A. (1983). J. Biol. Chem. 258, 7317-7321. Von Willert, D. T., Brinckmann, E., Scheitler, B., Thomas, D. A., and Treichel, S. (1979). Planta 147, 31-36. Vu, J. C. V., Allen, L. H., Jr., and Bowes, G. (1984). Plant Physiol. 76, 843-845.

46

LOUISE E. ANDERSON

Wan-Aswapati, O., Kemble, R. J., and Bradbeer, J. W. (1980). Plant Physiol. 66, 34-39. Whittaker, M. M., and Gleason, F. K. (1984). J . Biol. Chem. 259, 14088-14093. Wildner, G. F. (1975). Z . Naturforsch. 3Oc, 756-760. Winter, K. (1980). Plant Physiol. 65, 792-796. Wirtz, W., Stitt, M., and Heldt, H. W. (1982). FEBS Lett. 142, 223-226. Wolosiuk, R., and Buchanan, B. B. (1976). J . Biol. Chem. 251, 6456-6461. Wolosiuk, R. A., and Buchanan, B. B. (1979). Arch. Biochem. Biophys. 189, 97-101. Wolosiuk, R. A., Buchanan, B. B., and Crawford, N. A. (1977). FEBS Lett. 81, 253-258. Wolosiuk, R. A., Crawford, N. A., Yee, B. C., and Buchanan, B. B. (1979). J. Biol. Chem. 254, 1627-1632. Wolosiuk, R. A., Perelmuter, M. E., and Chehebar, C. (1980). FEBS Lett. 109, 289-293. Wolosiuk, R. A., Hertig, C. M., Nishizawa, A. N., and Buchanan, B. B. (1982). FEBS Lett. 140931-35. Woodrow, I. E., and Walker, D. A. (1980). Biochem. J. 191,845-849. Woodrow, I. E., and Walker, D. A. (1982). Arch. Biochem. Biophys. 216,416-422. Woodrow, I. E., Murphy, D. J., and Walker, D. A. (1983). Eur. J . Biochem. 132, 121-123. Woodrow, I. E., Murphy, D. J., and Latzko, E. (1984). J. Biol. Chem. 259, 3791-3795. Yee, B. C., de la Torre, A., Crawford, N. A., Lara, C., Carlson, D. E., and Buchanan, B. B. (1981). Arch. Microbiol. WO, 14-18. Ziegler, H., and Ziegler, I. (1965). Planta 65, 369-380. Ziegler, H., and Ziegler, I. (1967). Planta 72, 162-169. Ziegler, H., Ziegler, I., and Schmidt-Clausen, H.4. (1968). Planta 81, 169-180. Ziegler, I. (1972). Proc. Int. Congr. Photosyn. Res., 2nd pp. 1847-1853.

Algal Toxins

WAYNE W. CARMICHAEL Department of Biological Sciences Wright State University Dayton, Ohio, U.S.A.

I. Introduction.............................................................

. .. . .. .... .. ... . . .... .. .. . . In. Occurrence, Growth, and Toxicity.. . . . . . . . . . . . . . . . . . . . . . . . . . . . , . . . . . . . . . . . A. Chrysophyta ... ...................................................... B. Pyrrophyta. .. . . . . ... .. . .. . . .. . .. . . . . . .. .. .. .. ... . . .. .. .... . .. .. .... . . C. Cyanophyta .......................................................... IV. Isolation, Characterization, and Toxinology . . . . . . . . . . . .. .. . . . .. . . . . . . .. .. . . . A. Prymnesium Toxins.. . .. .. . . . .. . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . ... .. . . . B. Freshwater Cyanophyte Toxins . . . . . , . . . . . . . . . . . . . .. . . . . . . .. . . . . . . . . . . . C. Marine Cyanophyte Toxins ............................................ D. Dinoflagellate Paralytic Shellfish Poisons (PSP) . . . . . . . . . .. . . . . . . . .. . . . , . . E. Dinoflagellate Ciguatera Toxins . . . . . .. . . . . . . . . . . . . ... . . . . . . . . . . . . . . . . . . F. Other Dinoflagellate Toxins . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . .. . . . . . . . . . . . V. Environmental Role of Algal Toxins . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . References . . . . . . . . . .. , . . . . .. . . . . . . . . . .. . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . 11. Naming Algal Toxins. ... .. .. ... .. ... .. .. . .. .. .

47 52 52 52 59 64 67 67 68 78 82 86 89 91 93

I. INTRODUCTION Chemicals of secondary biosynthesis are important to the growth and development of all organisms, plant, animal, and microbial. The term “secondary” derives from the observation that these chemicals are not needed for primary metabolism, i.e., respiration or photosynthesis. The term also serves to suit the fact that for many of the chemicals no obvious functional role exists, hence the term secondary would seem to catalog them well. Secondary chemicals were first suggested by Stahl (1888) as ADVANCES IN BOTANICAL RESEARCH, VOL. 12

47

Copyright 8 1986 by Academic Press Inc. (London) Ltd. All rights of reproduction in any form reserved.

48

WAYNE W . CARMICHAEL

perhaps having evolved for protection against attack by herbivorous animals. Numerous other functions have been suggested for these chemicals, including the following: Regulators of plant growth or biosynthetic activity Storage forms of plant growth regulators Energy reserves Transport facilitators Waste products 6. Detoxification products of environmental poisons (Chew and Rodman, 1979) 1. 2. 3. 4. 5.

While some debate still exists concerning the adaptive significance of the secondary chemicals, evidence continues to accumulate to support an ecological role for these metabolites both in interactions among plants and their associated biota and as protective agents against physical environmental stresses (Rhoades, 1979). Secondary chemicals having a marked biological effect, usually negative, are often called biotoxins. While the role of biotoxins for animals is often clear, i.e., prey gathering or defense, the role of microbial biotoxins is not. Although they may act in some capacity as defense chemicals against herbivores or to improve an organism’s competitive advantage for a food source, it is also possible to find nontoxic forms of a potentially toxic species. Algae responsible for producing toxins are found in three of the eight divisions which make up the algae (Scagel et al., 1982). These are the Chrysophyta (class Prymnesiophyceae), Cyanophyta (cyanobacteria or blue-green algae), and the Pyrrhophyta (class Dinophyceae, also known as the dinoflagellates). In the Chrysophyta the toxic species is the flagellated unicellular Prymnesium paruum. It is a euryhaline chrysophyte widespread in brackish and marine habitats, producing toxins best known for causing mass mortalities of fish (Shilo, 1971, 1981). In the Cyanophyta both marine and freshwater toxic species are found. Marine toxic forms are in the filamentous genera Lyngbya, Schizothrix, and Oscillatoria. Toxins produced are of the type responsible for the contact dermatitis called “swimmers itch” (Moore, 1981b, 1984). Toxic freshwater cyanobacteria include the unicellular colonial Microcystis and the filamentous Anabaena, Aphanizomenon, (Figs. 1-3) Nodularia, and Oscillatoria. These freshwater algae can form thick surface accumulations of cells as the water bloom develops within the water body. They can be found in many eutrophic lakes and ponds at temperate latitudes and are responsible for sporadic but widespread outbreaks of wild and domestic animal illness or death (Gentile, 1971; Carmichael, 1981; Carmichael and Mahmood, 1984; Carmichael and Schwartz, 1984). They are also implicated in

Fig. 1. Scanning electron micrograph of Microcystis aeruginosa. Gelatinous matrix, which holds the cells in a colony form, is embedded with numerous rod-shaped bacteria. Micrograph by M. M. Ecker.

Fig. 2. Scanning electron micrograph of Anabaena circinafis.Numerous rod-shaped bacteria are embedded in the matrix surrounding the individual cells. Anabaena circinafiswas recently identified as the probable cause of animal deaths from a toxic water bloom in Illinois (Beasley et al., 1983). Micrograph by M. M. Ecker.

Fig. 3. Scanning electron micrograph of Aphanizornenon flos-aquae from New Hampshire. Cell in center of filament not covered by the sheath is a heterocyst-primary site of nitrogen fixation. Micrograph taken by M. M. Ecker.

52

WAYNE W . CARMICHAEL

human poisonings in certain municipal and recreational water supplies (Carmichael et al., 1985). The Pyrrhophyta contain most of the algae documented as causing outbreaks of algal disease. They are almost exclusively from the marine environment. Two major disease types are recognized. The first, paralytic shellfish poisoning (PSP), refers mainly to toxic species of Protogonyaulax (formerly Gonyaulax) (Figs. 4 and 5), which are concentrated by filter-feeding molluscs. Recently the tropical dinoflagellate species Gymnodinium catenatum (Morey-Gaines, 1982) (Fig. 6) and Pyrodinium bahamense var. compressa, and the tropical red alga Jania sp. have also been found to produce PSPs (Oshima et al., 1984). The molluscs are in turn eaten by humans, resulting in yearly episodes of illness and death. The second disease type is called ciguatera seafood poisoning. It refers to the concentration by many reef-feeding fish of toxic dinoflagellates recently identified as belonging to the genera Gambierdiscus (Fig. 7), Prorocentrum, Gymnodinium, and Gonyaulax. The fish are in turn eaten by humans (Ragelis, 1984).

11. NAMING ALGAL TOXINS As with most natural toxins, algal toxins have been named from the organisms that produce them or the vectors by which the toxins move through food chains. Only occasionally is a toxin name based on its chemical structure. This seems logical to the investigator doing the research and to his colleagues to whom he must communicate information about the toxin and the organism producing it. Coincidentally, it also reinforces the interest, respect, and image of mystique that the general public accords natural toxins. As more chemical information is known about toxins and when more than one toxic compound is isolated from a given toxic species, the use of this type of nomenclature system has become confusing. However, since most toxins are investigated and the findings published before their structure is entirely understood, the naming of toxins based on the organisms involved will continue. Table I lists the algal toxins and, where known, the origins of their names. 111. OCCURRENCE, GROWTH, AND TOXICITY A . CHRYSOPHYTA

The chrysophyte Prymnesium parvum was first identified as the causative agent of fish deaths by Carter (1938). Most problems have been reported from brackish water environments in England, Holland, Denmark, Bul-

ALGAL TOXINS

53

Fig. 4. Scanning electron micrograph of Protogonyaulax (Gonyaulax) tarnarensis. This species is the primary offender in outbreaks of PSP along the eastern coast of the U.S.A., Western Europe, and the western Pacific. Micrograph by L. Firtz.

Fig. 5. Scanning electron micrograph of Protogonyaulax (Gonyaulax) catenella, a PSP-producing dinoflagellatefrom the west coast of the U.S.A. Strands covering the cells are trichocysts extruded by the organism. Micrograph by G . Gaines.

ALGAL TOXINS

55

Fig. 6. Scanning electron micrograph of Gymnodinium catenatum, a paralytic shellfish poison-producing dinoflagellate from Mazatlan, Mexico (Morey-Gaines, 1982). Micrograph by G . Gaines.

TABLE I Algal Toxins-Terms Used and Main Species Responsible Organism A. Chrysophyta Prymnesium parvum

2

B. Cyanophyta 1. Marine Lyngbya majuscula

Schizorhrix calcicola Oscillatoria nigrouiridis 2. Fresh water Microcystis aeruginosa

Anabaena jlos-aquae (different strains of cyanobacteria)

Toxin

Chemical group

Origin of toxin name

Reference

Ichthyotoxin Cytotoxin Hemolytictoxin

Unknown

Fish toxin Cell toxin Red blood cell toxin

Shilo (1981)

Lyngbyatoxin A Debromoaplysiatoxin Aplysiatoxin Debromoaplysiatoxin Oscillatoxin A

Indole alkaloid Phenolic bislactone Phenolic bislactone Phenolic bislactone Phenolic bislactone

Genus Lyngbya

Moore (1984)

Microcystin Cyanoginosin

Peptide Peptide

Genus Microcystis

Anatoxin-a Anatoxin-b Anatoxin-c Anatoxin-d Anatoxin-b(s) Anatoxin-a(s)

Alkaloid Unknown Peptide Unknown Unknown Unknown

Genus Anabaena

Genus Oscillatoria Carmichael and Mahmood (1984); Botes et al. (1984) Carmichael and Gorham (1978)

Aphanizomenon Jos-aquae

Oscillatoria agardhii rubescens C. Pyrrophyta Gonyaulax (Protogonvaulax) catenella

u -4 l

tamarensis acatenella phoneus Pyrodinium bahamense var. compressa Ptychodiscus brevis (Gymnodinium breve) Dinophysis fortii acuminata Gambierdiscus toxicus

Aphantoxins

Alkaloid

Genus Aphanizomenon

Oscillatoria toxin

Peptide

Genus Oscillatoria

Paralytic shellfish poison (PSP) Saxitoxin

Alkaloid

Sasner et a/.( 1984); Carmichael and Mahmood (1984) Skulberg e f 01. (1984)

Hall and Reichardt (1984) Genus Saxidomas (Alaskan butter clam)

Neosaxitoxin Gonyautoxin 1-4 Cryptic (B1-2.C14)

Genus Gonyaulax

Brevetoxins

Pol ylactone

Genus P . brevis

Baden et al. (1984)

Dinophysistoxin Pectenotoxin

Okadaic acid-like Polyether lactone

Genus Dinophysis

Yasumoto el a/. (1984)

Ciguatoxin

Alkaloid

Cigua-from Cuban name for the marine snail Turbo Livona pica

Ragelis (1984)

Maitotoxin Scaritoxin

58

WAYNE W. CARMICHAEL

Fig. 7. Scanningelectron micrograph of Gumbierdiscus toxicus, collected from the Virgin Islands. Gumbierdiscus toxicus is the primary organism currently identified as being responsible for ciguatera seafood poisoning. Micrograph by G . Gaines.

garia, and especially Israel. The primary concern in Israel comes from the great number of commercial, brackish water fish ponds in which P. paruum was found to grow (Shilo, 1971, 1981). Prymnesium toxin consists of three toxins, hemoly sin, ichthyotoxin, and cytotoxin. Laboratory cultures on modified S50 medium (Shilo, 1967) produced the greatest amount of toxins during the late stages of logarithmic growth and continued at a high level into stationary phase. In contrast to other algal toxins the toxins of P. paruum are secreted into the environment, especially in the late

ALGAL TOXINS

59

stages of growth. Culture conditions can have a significant effect on toxin production. In the absence of light, a marked reduction of toxin production occurs when a carbon-enriched (glycerol) medium is us_ed.Cell multiplication continues during this time (Shilo, 1967). In contrast a 10- to 20fold increase in all the toxins is found in phosphate-starved cells with even more toxin than normal being found in the culture medium. Although the greatest production of the toxins occurs under stress conditions which would imply some protective (ecological)role for these secondary chemicals, Shilo (1981) feels that their formation has no direct role or importance in the life of P . paruum. B. PYRROPHYTA

Most reported PSP outbreaks have occurred in northern latitudes. Protogonyaulax (Gonyaulax) catenella and P . polyedra (Taylor, 1984) are implicated on the west coast of North America while Protogonyaulax tamarensis var. excauata (Schmidt and Loeblich, 1979) is involved in Northern Europe, Great Britain, Japan, and Western Pacific. These marine dinoflagellates exist in three cell types: 1. A resting zygote (cyst, 25-30 X 45-50 pm)(Dale et al., 1978; Turpin et al., 1978; Anderson, 1984) 2. A baagellate motile cell (35-50 pm in diameter) which is the bloomforming stage 3. An asexual temporary cell (cyst, 30-40 p m in diameter) called a pellicle or ecdysal cyst (Anderson and Wall, 1978; Turpin et al., 1978).

Blooms of these dinoflagellates are generally defined as occurring when cell populations are between 104-106 dm-3. Lower cell concentrations of lo3dm-3 can occur at any time during the April-October growing season. When cell concentrations reach about lo6 dm-3 the water can be discolored and referred to as a “red tide.” Red tides are not all toxic, however, since (1) red tides can form from the presence of other, nontoxic dinoflagellates (Yentsch et al., 1978); (2) shellfish can accumulate toxins at levels below those necessary to discolor the water (Hurst and Yentsch, 1981); and (3) not all Protogonyaulax species are toxic when isolated, cultured, and tested (C. M. Yentsch, personal communication). Dense concentrations of toxic marine dinoflagellates can result from several factors (Yentsch and Incze, 1980) in addition to the direct rapid growth of the algal population. These physical factors apparently have a great deal to do with frontal circulation patterns (Seliger et al., 1979). Active phytoplankton growth is situated along or close to these frontal systems that form between well-mixed and season-

60

WAYNE W . CARMICHAEL

ally stratified water masses. Holligan (1979) defines three stages of bloom formation in these frontal areas, as measured by chlorophyll a. The first stage, which depends on upward mixing of nutrient-rich waters, involves growth of phytoplankton at the base of the thermocline. The second stage occurs in the upper part of the thermocline at a depth of about 10 m. The third stage, which often finds the blooms patchy in appearance, may be initiated by upward mixing of the subsurface bloom. While Holligan (1979) does not find evidence to suggest that vertical migration of dinoflagellates occurs, Dugdale (1979) and others find evidence to indicate that migration occurs in response to nutrient gradients, especially nitrates. These studies indicate that dinoflagellates swim downward in the afternoon and evening to take advantage of higher nutrient levels than those near the surface where higher photosynthetic rates have been taking place during the day. There is also some evidence that toxic dinoflagellates exhibit an annual rhythm. Yentsch and Mague (1980) report that Protogonyaulax excuuatu, cultured under constant conditions over a 5-year period, exhibited a circannual rhythm. Growth rates varied from a maximum of two doublings per day for the months of July and August to 0.10 doubling per day in the month of January. Hydrographical mechanisms are also important in the formation of toxic blooms. These mechanisms require no rapid reproduction and provide a means for delivery of an existing population to an area where biological behavior such as phototaxis can result in dense concentrations. These mechanisms can be triggered by meteorological events such as rainfall and wind or the formation of discontinuities between water masses, called frontal zones. These fronts which result from tide or wind-generated convergences and/or density discontinuities are some of the locations most likely to generate red tides (Yentsch and Incze, 1980). Blooms and cultures of toxic dinoflagellatescan vary in toxicity. When cultured in the laboratory, variations are seen in quantity and type of toxin produced. Points to remember when studying dinoflagellate toxicity include the following: 1. The mouse bioassay used to detect toxicity has an inherent variability of about k 20%. 2. Toxicity is a measure of total toxins produced by the population in culture and does not distinguish between the different toxin types.

Growth would seem to be the most obvious regulator of toxicity with optimal growth conditions reponsible for optimum toxin production. However, with some dinoflagellates this may not be so. Hall (1982) studied the effects of nutrient deprivation using modified seawater with Guillard’s “F/2” enrichment (Guillard and Ryther, 1962), on cell growth and

ALGAL TOXINS

61

toxicity of selected Alaskan coast Protogonyaulax clones. The omission of nitrate and phosphate from the culture medium was tested for its effect on growth and toxicity. Despite growth reductions to a level of 3200 and 3000 cells ~ m for - ~minus nitrate and minus phosphate, respectively, compared to 16,300 (over 27 days) for controls, the toxicity per cell was slightly better then the control for minus nitrate and over 5 times as much for the minus phosphate cultures. Growth of the clone over a series of temperatures ranging from 6 to 12°C showed a reduction in growth rate at lower temperatures accompanied by an increase in toxin content per cell. This growth attenuation accompanied by increased toxicity is supported by DuPuy (1968) and Proctor et af. (1975), who concluded that toxin content is inversely related to growth rate. Perhaps this increase in a secondary metabolite under stress conditions can be likened to the production of secondary compounds by higher plants when they are grown under nutrient stress conditions (Janzen, 1974). Hall (1982) also rightfully points out that the conditions that induced higher toxin content, viz., low phosphate and low temperature, are closer to the range of these parameters found under natural conditions. Hall (1982) also investigated toxin composition under the attenuated growth conditions which increased overall toxin composition. The results of his limited experiments in this area showed that toxin composition among strains isolated from different regions of the Alaskan coast had different toxin compositions while those from close neighboring areas were uniform through successive culture runs. He found that the sulfamate toxins (Fig. 11) were dominant across the entire geographical range sampled. It should also be said that toxicity of dinoflagellates has been found to decrease as the cultures approach the end of log growth phase (White and Maranda, 1978). The economic impact of toxic dinoflagellates is felt mainly when the toxins are concentrated by shellfish. Although many different shellfish can accumulate dinoflagellate toxins (Carmichael et a f . , 1985) the complex pattern of toxification and detoxification has been best studied using the blue mussel Mytilus edulis. This is based on the organism’s widespread occurrence, ease of collection, and rapid rate of intoxification and detoxification. In the laboratory M. edulis will toxify in proportion to the filtration rate and the number of cells in the water. Along the coast of Maine (U.S.A.) during the summer when optimal cell densities of about lo6 ~ m are - ~found, M. edulis can toxify at the rate of 10 pg toxin g-I tissue per day. These are the times when Protogonyaulax is in the motile stage (Gilfillan and Hansen, 1975). Under these conditions the lethal levels for humans of 10 mg of toxin are often reached. In a 4-year study (1976-1979) involving shellfish intoxication patterns along the coast of Maine, Hurst and Yentsch (1981) found that the peak toxicity occurred in

62

W A Y N E W. CARMICHAEL

1978. Toxin maxima were found in May and ranged from 25 to 114 pg g-' tissue. These high levels of toxin accumulation are possible only because shellfish are not harmed by the toxins they accumulate (Twarog and Yamaguchi, 1975). Detoxification of the mussel tissue depends on the amount of toxin present and the amount of melanin pigment in the tissue (Price and Lee, 1971). Decay constants can in some cases be determined and used to help determine when to reopen contaminated shellfish beds (Hurst and Gilfillan, 1977). Hurst and Yentsch (1981) concluded from their survey that 1. Considerable variability exists for toxicity between stations a few kilometers apart. However, appearance of toxicity is simultaneous over wide areas. 2. Major occurrences of toxicity are during spring or late summer when water temperatures are changing. During this time toxic cysts and motile cells in the water provide the inoculum for a new bloom. 3. Considerable variation between years is found at all stations. Most stations are characterized by one major outbreak per year. 4. Rates of intoxification and detoxification are rapid, being several hundred micrograms of shellfish toxin per 100 g shellfish tissue per day.

Another important factor influencing the apparent toxicity of PSP-producing dinoflagellates is biotransformation by the shellfish (Shimizu and Yoshioka, 1981; Shimizu et al., 1984). By using homogenates of the east coast scallop Placopecten magellanicus, it was shown that gonyautoxins and neosaxitoxin decreased while saxitoxin content increased. Bioconversion of neosaxitoxin to saxitoxin in Mya arenaria and Mercenaria mercenaria was also demonstrated. Dinoflagellates causing ciguatera disease are found to be most prevalent in the tropics between 35"N and 34"S, especially in the South Pacific islands and the Caribbean. In these areas the disease affects more than 10,000 individuals per year and appears to be increasing in frequency (Halstead, 1967; Ragelis, 1984). Ciguatera has been caused by over 400 species of fish, but especially the herbivorous surgeon fish and parrot fish, and the larger carnivorous reef sharks and red snappers (Banner, 1976). Randall (1958) concluded that all such fish were tied to the coral reef through the food chain. The possibility that ciguatera originates as a result of toxic algae was first mentioned by a British physician, Colin Chisholm, in 1808 (Halstead, 1967). Evidence gathered during the 1960s and 1970s pointed to certain dinoflagellates that grow in close association with macroalgae and/or other bottom structures as progenitors of ciguatera toxins. It was not until a publication by Bagnis et al. (1977) that a causative organism was isolated from the biodetritus layer of waters surrounding

ALGAL TOXINS

63

the Gambier Islands in the South Pacific. It was initially identified as a Diplopsalis sp. by Yasumoto et al. (1977) but later it was decided that this benthic dinoflagellate belonged to a new genus and was named Gambierdiscus toxicus (Adachi and Eukuyo, 1979). Other species of dinoflagellates inhabiting the same microhabitats in the Pacific have been found to produce toxins which may also contribute to the ciguatera syndrome. These include Amphidinium carteri, Amphidinium klebsii, Ostreopsis ovata, Ostreopsis siamensis, Prorocentrum concavum (Nakajima, 198 l), and Prorocentrum lima (Yasumoto et al., 1980; Nakajima et al., 1981; Murakami et al., 1982). Current studies on ciguatera seem to center around those of Tindall and co-workers working in the Caribbean, Yasumoto, Bagnis, and others who are working in the Pacific island area, and Kimura, Hokama, and coworkers in Hawaii. Tindall et al. (1984) surveyed over 70 sites in the Virgin Islands. From these collections 65 strains representing 18 species of dinoflagellates are being cultured at the University of Southern Illinois. These species have all been proved to culture best using the ES medium of Provasoli (1968) to which was added 1-5% soil extract. Initial screening for toxicity showed that all possessed some toxic component and studies to date have concluded that Prorocentrum concavum and Gambierdiscus toxicus are the major contributors of ciguatera toxins to these Caribbean reef and inshore ecosystems. These studies also concluded that no one dinoflagellate or toxin is the major cause of ciguatera. It was also found that the major ciguatera toxin found in the contaminated fish, ciguatoxin, was not the major toxin produced by cultured toxic strains. This implies that ciguatoxin may be formed by biotransformation processes in the fish, that laboratory cultures of toxic dinoflagellatesproduce different toxins in the laboratory than in nature, or that ciguatoxin-producing dinoflagellates have not been successfully isolated and cultured. A recent study by Carlson et al. (1984) examined the effect of extracts of macroalgae on the growth of G . toxicus, P . concauum, and Prorocentrum mexicanum. The macroalgae chosen were those commonly found in association with these toxic dinoflagellates in the Caribbean, viz., Chaetomorpha linum (Chlorophyta), Dictyota dichotoma (Phaeophyta), and Turbinaria turbinata (Phaeophyta). Water-soluble extracts from field collections of these macroalgae were aseptically added to axenic cultures of the toxic dinoflagellates. It was found that soil and macroalgal extracts enhanced the growth of bacterized cultures of G . toxicus and P . concauum but inhibited that of bacterized P . mexicanum. Generation times of axenic cultures of G. toxicus and P. concavum were lengthened. Evidence was found to indicate that bacteria could contribute to variation in toxin production. Variation of toxicity independent of growth due to bac-

64

WAYNE W. CARMICHAEL

teria has also been demonstrated in freshwater toxic cyanobacteria (Carmichael and Gorham, 1977). The dinoflagellate study concluded that enhanced algal growth in the presence of bacteria was not due to increased C02production. It was also speculated that hormones such as cytokinins may play a role in the stimulation of growth by the macroalgal extracts. C. CYANOPHYTA

Cyanobacteria producing toxic substances have been reported since the late 1800s (Carmichael, 1981).The first laboratory cultures of toxic strains were obtained from the colonial species Microcystis aeruginosa by Hughes et al. (1958) using a modified Fitzgerald medium (Fitzgerald et al., 1952). Later, cultures of Microcystis utilized the medium ASM (Artificial Seawater McLachlan), which is a modified ASP (Artificial Seawater Provasoli) (McLachlan and Gorham, 1961). Growth experiments on several single-cell isolates (clones) of Microcystis revealed that toxicity varied between clones and that there were toxic and nontoxic cells within a given colony. A natural consequence of this varying toxicity was that only the most toxic clones were retained in culture for future work (Gorham, 1964). The optimal temperature for growth of the toxic clone M . aeruginosa NRC-1 was not the same as that for optimal toxin production. The same toxic clone was grown in 9 dm3 batch cultures for isolation and characterization of the toxic principle (Bishop et al., 1959). Since the late 1960s, however, NRC-1 has not been grown for toxicity studies and when tested in 1971by this author the toxic characters differed from those when the strain was originally isolated. Recent tests for toxicity in 1980 indicated that overall toxicity was also declining. This has resulted in the use of another toxic strain, M . aeruginosa 7820 (Codd and Carmichael, 1982). It should be noted that over the period of time NRC-1 was cultured, the medium in which it was grown was changed from modified Fitzgerald to ASM to ASM-1 (Gorham et al., 1964), with the current medium being BG11 (Stanier et al., 1971). These media have progressively higher levels of nitrate as NaN03. BG-11 has 18 times the amount of nitrate as ASM. Phosphate concentration as orthophosphate is equal in all three media. These culture media were chosen for their growth-promoting potential and only secondary consideration was given to whether toxin production was affected with extended subculturing. In retrospect it is important to ask whether the decline in toxicity is due to the fact that the cells are now in a nutrient-rich medium and no longer “stressed,” resulting in lowered toxin production. This same pattern is emerging with other genera of toxin-producing cyanobacteria similar to those of Microcystis, including Anabaena (Carmichael, unpublished data) and Oscillatoria (0. M.

ALGAL TOXINS

65

Skulberg, personal communication). Sirenko (personal communication) also finds that culture toxicity declines when Microcystis isolated from

waters in the U.S.S.R. is maintained in the laboratory over a number of months to years. In the early 1960s toxic strains of Anabaena flus-aquae were isolated for laboratory studies by Gorham and co-workers (Gorham et al., 1964). The basic ASM medium was modified by doubling the nitrogen, phosphorus, calcium, and iron contents while increasing by four times the boron, zinc, cobalt, and copper contents to produce a medium termed ASM-1, Most studies on the chemistry and toxicology of Anabaena toxins have been done using the clonal isolate NRC-44-1 (Carmichael, 1981). Toxicity has remained quite stable in this strain over the period of 1961 to 1980. At this time, for reasons of convenience, NRC-44-1 was transferred to BG-11 for maintenance in this author’s culture collection. Growth continued to be good in this strain but a routine check of toxicity in 1982 revealed that the strain had become nontoxic. A check of 25 subcultures of NRC-44-1 maintained as backup cultures in the laboratory revealed that only 1 had a level of toxicity detectable by mouse bioassay. Transfer of this toxic subisolate back to ASM-1 medium has restored its normal level of toxicity, i.e., 40-60 mg kg-1 (intraperitoneal injection in mouse). Subcultures maintained by P. R. Gorham (University of Alberta) on ASM-1 medium and 0. M. Skulberg (Norwegian Institute for Water Research) on Z-8 medium (Skulberg, 1983), have retained their toxicity throughout this time period. Both ASM-1 and Z-8 media contain the same level of nitrate. Light levels, temperature, and pH were maintained the same throughout this time period. At present, investigations are underway to see if nontoxic NRC-44-1 returned to ASM-1 medium will regain its former toxicity levels. Another instance of apparent change in toxicity has been observed in Anabaena flos-aquae S-23-g, a strain isolated by Carmichael from Saskatchewan, Canada in 1975 (Carmichael and Gorham, 1978). This strain, at the time of initial isolation, produced a toxin, anatoxin-d, with neurotoxic properties. Repeated subculturing of this strain on ASM-1 and then changing to BG-11 medium has resulted in the loss of neurotoxin production and expression of a hepatotoxin like that of Microcystis (Jones, 1984). It will be important to examine whether cultures of toxic cyanobacteria now being cultured from Norway (Skulberg, 1983; Skulberg et al., 1984) and South Africa (Eloff el al., 1982; Botes et al., 1982a; Siegelman et al., 1984) will also undergo a decrease in toxin production with time when grown on the standard nutrient-rich culture media. Toxicity of the filamentous c yanobacterium Aphanizomenonflos-aquae

66

WAYNE W. CARMICHAEL

has proved to be the most difficult to study. This is largely due to the difficulty in culturing compared to other toxic cyanobacteria. Phinney and Peek (1961) were among the first to report that blooms of Aph.Jos-aquae could be toxic. Toxicity was not verified, however, by laboratory cultures from the bloom. Gorham (1964) reported some success in culturing Aphanizomenon from sources in Ontario and Saskatchewan but toxicity in the mouse bioassay was negative in all cases. Gentile (1971) reported some success in culturing toxic Aph. Jos-aquae from blooms collected at Klamath Lake, Oregon in 1968. However, no work was done on toxicity or toxicology of the isolates to verify toxicity beyond the standard mouse assay. Toxic isolates were cultured and examined from blooms collected at Kezar Lake, New Hampshire (Gentile and Maloney, 1969). In growth studies toxin production was not increased independently of growth when light or temperature was changed. Toxicity did decrease independently of growth, however, when Aph. flos-aquae was cultured at stress temperatures of 30”C, but not at the more normal environmental temperatures of 15, 20, or 26°C. No experiments were conducted to investigate possible toxin increases by stressing the cells through lowering nitrate or phosphate concentrations. These toxic isolates were subsequently discarded and no toxic Aphanizomenon existed in culture until new isolates from New Hampshire were made by Carmichael in 1980 (Ikawa et al., 1982; Carmichael and Mahmood, 1984). These isolates produce toxins similar to the earlier New Hampshire isolates but as yet no growth studies have been done to investigate possible regulation of toxin production. Toxic marine cyanobacteria have not been studied in laboratory cultures. Instead, all chemical and toxicological studies have used field collections. Toxic marine cyanobacteria are all found in the Oscillatoriaceae family, occupying benthic habitats in warm water areas such as Hawaii and Okinawa (Moore, 1981a,b). The secondary chemicals produced cause a severe “swimmer’s itch” or contact dermatitis among swimmers who come in contact with the filaments. Most reports of these contact irritations come during the summer months and it is not clear whether toxins are produced at other times of the year, although it would seem that they would be. While direct comparisons between marine and freshwater cyanobacteria toxins are difficult to make, at present there is some indication that freshwater cyanobacteria produce contact irritants. This comes from field observations on certain species of Anabaena, Aphanizomenon, Oscillatoria, and Gloeotrichia in the U S A . and Europe over the past 4 years (Skulberg et al., 1984; Carmichael et al., 1985; Codd and Carmichael, unpublished data). When these toxins are characterized and studied it should become clear that the freshwater cyanobacteria are capable of producing secondary chemicals with neurotoxic, hepatotoxic, and dermatotoxic properties.

ALGAL TOXINS

67

Overall there has been little work on the effects of growth on toxin production for any of the toxic algae. Most investigators have been kept busy just maintaining a few strains in culture in order to isolate, purify, and characterize the toxins present. It is possible to outline, however, several studies which could yield information on qualitative and quantitative aspects of toxin production and the role of these secondary chemicals in the ecology of the organism. These include 1. The role of nutrient depletion on toxin production, especially major nutrients such as nitrogen and phosphorus. At least some evidence points to a possible stimulation of toxin production as growth decreases due to a shortage of nitrogen and phosphorus. This should be analyzed over several generations of subculturing. 2. Although the few studies that have been done do not indicate that temperature or light affect toxin production independently of growth, longer term studies could prove useful in examining this possibility. 3. Possible role of zooplankton herbivores through selective feeding of nontoxic over toxic cells. Zooplankton presence could also stimulate production of toxins under herbivore stress, as is now being revealed for higher plants. This possibility does not seem as likely as nutrient stress, however, since it would imply that some chemical communication exists between cells of a bloom. 4. Possible role of extrachromosomal DNA (plasmids) in regulation of toxin production (Hauman, 1981). Expression of the plasmid regulating toxin production (if present) could be part of nutrient or predator stress.

IV. ISOLATION, CHARACTERIZATION, AND TOXINOLOGY A. PRYMNESIUM TOXINS

The toxic principles of P.paruum have been purified and characterized by Ulitzur (1969) and Ulitzur and Shilo (1970). The toxin has a broad spectrum of different biological activities in uiuo and in uitro (Shilo, 1971, 1981). It seems that there is a family of compounds having similar composition but different toxic effects, rather than a single compound with different toxic effects. These effects are generally classed as being cytotoxic, hemolytic, and ichthyotoxic. Since the toxic effects are primarily to fish the main component is often referred to as an ichthyotoxin. Intoxication of fish consists of two stages. Initially there is reversible damage to the gill tissues, resulting in the loss of their selective permeability. The second stage, which leads to death, is the response of the sensitized fish to any of a number of toxicants in the environment including the Prymne-

68

WAYNE W . CARMICHAEL

sium toxin (Shilo, 1981). The gill epithelium of the fish is the primary site of action for the toxin. Extraction of the toxic compounds indicates that they are acidic, polar lipids. Analysis of the toxin revealed 15 amino acids, a number of fatty acids, 0.47% phosphate, and 10 to 12% hexose sugars (Ulitzur and Shilo, 1970). In its composition of fatty acids together with protein and phosphate, the purified toxin resembles proteolipids (Shilo, 1981). More recently isolation and purification of Prymnesium hemolytic toxin and ichthyotoxin have been done by Kim and Padilla (1977) and Kozakai et al. (1982).

B. FRESHWATER CYANOPHYTE TOXINS

1 . Neurotoxins Known chemical groups of toxins from freshwater cyanobacteria include alkaloids and peptides. The alkaloid toxins are the most rapidly acting. They function as neurotoxins paralyzing peripheral skeletal muscles, then respiratory muscles, with death due to respiratory arrest occurring between a few minutes to a few hours. These toxins are produced by various strains and species of Anabaena and are generally referred to as anatoxins (Table I). Anatoxin-a (antx-a) is the only alkaloid toxin from this group which has been chemically characterized (Fig. 8). It is a potent depolarizing neuromuscular blocking agent (Carmichael et al., 1975, 1979; Spivak et al., 1980). Chemically the compound is the secondary amine, 2-acetyl9-azabicyclo[4.2.l]non-2-ene,and is isolated from the filamentous strain Anabaena flos-aquae NRC-44-1 (Huber, 1972; Devlin et al., 1977). Synthesis of antx-a has been done through a ring expansion of cocaine (Campbell et al., 1977) from 1,5-cyclooctadiene (Campbell et al., 1979), and by intramolecular cyclization between an iminium salt and a nucleophilic carbon atom (Bates and Rapoport, 1979).

4

Fig. 8. Structure of anatoxin-a hydrochloride, a depolarizing neuromuscular blocking agent produced by the filamentous cyanobacteriumAnabaena flos-aquae clone NRC-44-1.

ALGAL TOXINS

69

Poisoning of domestic and wild animals by anatoxins comes from ingestion of the toxic cells and extracellular toxin from a water bloom. Given the LD5o for intraperitoneal bioassay in mouse for purified anatoxin-a as 200 pg kg-I body weight and the approximate oral lethal dose for certain animals (Carmichael et af., 1977; Carmichael, 1981), it is estimated that a lethal bolus of water bloom ranges from a few cubic centimeters to a few cubic decameters, depending upon animal species, toxicity of the bloom, and amount of food material in the animal’s gut. The presence of more than one toxin in the bloom can also result in different signs of poisoning and survival times. Known toxins of Aphanizomenon Jos-aquae are also neurotoxic alkaloids, generally referred to as aphantoxins. Sasner and his colleagues presented evidence that aphantoxin from water blooms of Aph. Josaquae was similar to the paralytic shellfish poisons saxitoxin and neosaxitoxin (Sawyer et al., 1968; Jackim and Gentile, 1968; Alam et a f . , 1973, 1978; Alam and Euler, 1981; Sasner et af., 1981; Adelman et al., 1982). Current research with aphantoxin is concerned with a toxic strain isolated from a small pond near Durham, New Hampshire in 1980 by Carmichael, referred to as NH-1 and NH-5 (Carmichael, 1982; Ikawa et a f . ,1982). The toxins of NH-5 are primarily neosaxitoxin (80%) and saxitoxin (15%). Depending on extraction procedure it appears that aphantoxin also contains some precursors to neosaxitoxin and saxitoxin (Carmichael and Mahmood, 1984; Sasner et al., 1984). At present the structures of the aphantoxins have not been determined but assuming that they are the same as PSPs, and all evidence to date supports this, they will have structures like those in Fig. 11. Detection methods for these toxins in water supplies have not been adequately developed but high-performance liquid chromatography (HPLC) is being used in some cases. Both Astrachan and Archer (1981) and Wong and Hindin (1982) have used HPLC for the detection of anatoxin-a. In our laboratory we routinely use HPLC to purify other neurotoxins of Anabaena and Aphanizornenon (Carmichael and Mahmood, 1984). These methods could be modified to detect low levels of the toxins in water reservoirs or recreational waters. 2 . Hepatotoxins The peptide toxins of the freshwater cyanobacteria are primarily found in various strains of Microcystis aeruginosa. These toxins are responsible for most poisonings by cyanobacteria since M. aeruginosa is more common in its worldwide distribution than the other toxigenic cyanobacteria. Although other cyanobacteria, including Anabaena, Aphanizomenon, and Oscilfatoria (Carmichael, 1981, 1982; Skulberg et af., 1984) are

70

WAYNE W . CARMICHAEL

thought to produce peptide toxins, Microcystis peptides have been studied most. Toxic blooms have been found in ponds and lakes thoughout the world including the U.S.A., Canada, the U.S.S.R., Europe, South Africa, India, Japan, the Middle East, and Australia. In temperate zones most toxic blooms occur in mid to late summer (Carmichael et al., 1985). Rats and mice injected either intravenously or intraperitoneally with acutely toxic doses of the cells or toxin extract (LDso 25-100 pg/kg, intraperitoneally in mouse) die within 1 to 3 hr. Death is preceded by pallor and prostration, with episodes of unprovoked leaping and twitching, Upon necropsy, the animals show grossly enlarged livers engorged with blood, with the remainder of the carcass being exsanguinated. Liver weight is increased, and at death composes about 8 to 10% of body weight of mice as compared to about 5% in controls (Slatkin et al., 1983; Falconer et al., 1981; Theiss, 1984). The blood content of livers of mice poisoned by Microcystis increases from 66 mm3g-' liver in controls to 534 mm3g-l for mice killed 45 min after toxin injection (Falconer et al., 1981). Histological examination of the liver reveals extensive centrilobular hemorrhagic necrosis with loss of characteristic architecture of the hepatic cords. Transmission electron microscopic examination indicates that both hepatocytes and hepatic endothelial cells are destroyed. The only alterations noted prior to cell rupture are slight mitochondria1 and cell swelling. Damaged cells have extensive fragmentation and vesiculation of the membrane (Runnegar et al., 1981). Gross and histological examination of intestine, heart, spleen, kidneys, and stomach show no consistent abnormalities; lungs are mildly congested with occasional patches of debris. Thrombi thought to contain platelets are found in the lungs of affected animals (Slatkin et al., 1983; Falconer et al., 1981). It is suggested that these thrombi may be a direct effect of the toxin and may secondarily cause the liver effects by creating sufficient pulmonary congestion to cause right heart failure which, in turn, could cause blood pooling and congestion in the liver (Slatkin et al., 1983). However, in time course studies, Falconer et al. (1981) reported that the pulmonary thrombi did not appear in histological preparations taken at 15 and 30 min after toxin injection and were present only in later preparations (while liver damage was noted as early as 15 min). This evidence, along with other evidence of effects on isolated hepatocytes and the rapid onset of the liver effects in uiuo, have led other researchers to believe that the liver damage is a direct effect of the toxin on the hepatocyte membrane and that the immediate cause of death in acutely dosed animals is hemorrhagic shock (Falconer et al., 1981; Runnegar and Falconer, 1981; Theiss, 1984). Occasionally, hemorrhages are noted in organs other than the liver (Ostensvik et al., 1981). This could possibly be due to coagulation problems associated with the liver damage.

ALGAL TOXINS

71

The action of Microcystis toxin on isolated rat hepatocytes was investigated by Runnegar et al. (1981), Runnegar and Falconer (1981), and Foxall and Sasner (1981). In cell suspension, normal hepatocytes viewed under the scanning electron microscope show a rounded appearance with a surface covered with microvilli. Within 5 min of incubation with the toxin, obvious lumps can be seen on the cell surface, which increase with time to present a distorted lumpy cell without microvilli. These changes occur only in live cells and are found to be dose dependent, with the minimum concentration of toxin causing deformation being directly related to the LDso for the toxin. There is no difference in trypan blue exclusion by hepatocytes incubated with or without the toxin, the toxin causes no cell lysis, and there is no release of aspartate amino transferase (ASAT) into the medium. The toxin appears to be transported into the cell via the bile acid transporters in the cell membrane. Deformation of the cells by Microcystis toxin is blocked by addition of sodium deoxycholate to the medium. The blocking effect also shows a dose response, with increasing concentration of toxin requiring a higher bile acid concentration. In another study by Grabow et al. (1982) the effect of Microcystis toxin was tested on isolated liver, lung, cervix, ovary, and kidney cell cultures. Cells of all cultures were damaged or disintegrated after overnight incubation in the presence of Microcystis toxin. While no mechanism of action was proposed it was suggested that the toxin may act on cell membranes. The effect on isolated hepatocytes of the presence and absence of calcium in the medium was also investigated (Runnegar and Falconer, 1982). It has been suggested by Farber and his co-workers that calcium entry into the cell is the final common step in the death of cells injured by membrane toxins and that cells can be protected by excluding calcium from the medium (Farber, 1981). Conversely, it has been reported by other researchers that toxic injury to hepatocytes is not dependent on calcium and in some cases calcium in the medium actually protects cells from injury by various toxins (Smith et al., 1981). Tests with Microcystis toxin and isolated hepatocytes revealed that the presence or absence of calcium in the medium makes no difference to the toxic effect (Runnegar and Falconer, 1982). It was reported that toxic extracts from algal blooms would agglutinate red blood cells (Carmichael and Bent, 1981). However, Runnegar and Falconer reported no red blood cell agglutination with extracts isolated in their laboratory (Runnegar and Falconer, 1982). The effects of the toxin on mouse liver slices, isolated mitochondria, and microsomes were investigated and no specific effects were noted (Runnegar and Falconer, 1981). The toxin did not significantly affect the incorporation of labeled leucine, uridine, or methylthymidine into trichlo-

72

WAYNE W. CARMICHAEL

roacetic acid-insoluble precipitates from incubated mouse liver slices, indicating that it exerts no major effect on protein, RNA, or DNA synthesis. The toxin did not affect oxygen consumption of liver slices and only slightly increased that of isolated mitochondria. Measurements of glycogen degradation in liver slices incubated with toxin showed a variable increase in glycogen loss and glucose appearance in the incubation medium (Runnegar and Falconer, 1982). Runnegar and Falconer (1981) noted similarities in both structure and gross pathological effects of Microcystis toxin and phalloidin, a bicyclic peptide from the poisonous mushroom Amanita phalloides. Phalloidin produces a hepatic hemorrhagic necrosis similar to that produced by Microcystis, but these similarities do not extend to the cellular level. Phalloidin produces its toxic effects by inhibiting the depolymerization of F actin to actin; Microcystis does not. The molecular mechanism of action of Microcystis toxin is, at this time, unknown. Time course studies of dosed rats and mice have been done by several researchers (Slatkin et al., 1983; Ostensvik et al., 1981; Runnegar and Falconer, 1981). Liver damage can be noted in histological sections as early as 15 min after dosing, with the centrilobular hemorrhagic necrosis steadily progressing outward with time. Platelet counts taken during the course of the toxic reaction show a steady decrease in the circulating platelet count that is inversely proportional to the increase in liver weight (Slatkin et al., 1983; Jones, 1984). Runnegar and Falconer (1981) noted changes in aspartate amino transferase (ASAT) and lactic dehydrogenase at 15 min, and by 30 min had increased to 50 times control levels. Ostensvik et al. (1981) found changes in ASAT commencing at 30 min, but no changes in alanine amino transferase (ALAT) nor bilirubin during the course of toxicity. Ostensvik et al. (1981) monitored the effect of an extract of a predominantly Microcystis water bloom on blood pressure of rats. The blood pressure decreased markedly immediately after intravenous administration, but began increasing at 1.5 min and was back to normal levels within 10 min of injection. During the next 40 min, the blood pressure decreased slowly to very low levels consistent with hemorrhagic shock and remained constant at these low levels until the rats died at about 90 min. Theiss (1984), using purified toxic peptide of M. aeruginosa strain 7820, also found that blood pressure responses, both arterial and venous, were indicative of a direct hepatotoxin which caused death by hemorrhagic shock. Jackson et al. (1984) inoculated sheep intraruminally with M. aeruginosa water bloom suspension. The majority of the lethally poisoned sheep died within 18 to 23 hr. The carcasses of the sheep showed that the

ALGAL TOXINS

73

primary site of toxicity was the liver, which had centrilobular to near massive hepatocyte necrosis. Small hemorrhages were also noted in many other areas of the body. Lungs were mildly edematous, but other organs were normal except for the hemorrhages. It was noted that neutrophils were attracted into the lumen of bile ductules without a cholangitis. This has been noted in other animals poisoned with Microcystis and may have diagnostic significance in differentiating algal poisoning from other plant hepatotoxicities. These sheep experiments revealed a sharp dose-response curve in that up to 90% of the lethal dose of bloom could be ingested in a single administration without measurable effect. From the animal response it was estimated that the 27-kg sheep used would have to ingest about 650 cm3of the thick algal bloom as it occurred on the pond to provide the approximately 30 g of dried cells necessary to cause acute lethal toxicity. Foxall and Sasner (1981) reported that cladocerans, amphibians, crustacea, and teleosts were not affected by the toxin, but all mammals and birds tested to date are sensitive to it. Some effects have been reported on cardiac muscle and blood hemostasis (Kirpenko and Kirpenko, 1980). No effect has been reported on skeletal muscle or isolated nerve preparations. Foxall and Sasner (1981) reported that young hepatocytes in uitro are not affected and neither are young mice nor rats. These mice were not affected by lethal doses of the toxin until they had reached the age of approximately 20 days. This would suggest a possible activation of the toxin by the liver enzyme systems, but to date no one has investigated this possibility. Female mice were slightly more sensitive to the toxin than male mice. Foxall and Sasner (1981) reported no antibiotic activity against green algae, yeast, or bacteria and no toxicity to certain zooplankton, crayfish, amphibians, and teleosts. However, Grigor’yeva et al. (1977) reported a wide spectrum of antimicrobial activity against Escherichia coli, Shigella jlexneri, Salmonella typhimurium, Staphylococcus aureus, Enterococcus, and Candida. The mechanism involved was a decreased thiamine content and inhibition of dehydrogenase activity. Kirpenko et al. (1982) found that Microcystis toxin extracts inhibited the development of saprophytic microflora in artificial ponds. Kirpenko et al. (1981) found that toxic Microcystis caused embryolethal, teratogenic, and gonadotoxic effects in the rat. Mutagenesis involving anomalies of chromosome and chromatid apparatus was also reported. However, Runnegar and Falconer (1982) reported no mutagenicity when a purified extract was tested by the Ames Salmonella assay.

74

WAYNE W . CARMICHAEL

Reports on Microcystis poisoning are often conflicting. The only consistent pathological findings in Microcystis toxicity are the swollen, blood-engorged liver with hemorrhagic necrosis and the mildly edematous lungs. These conflicting reports may be attributed to possible differences in the effects of different strains of toxic Microcystis, the use of extracts from mixed blooms that contain other algal or bacterial species, or differences in research technique. Microcystis has been linked to an outbreak in 1979 of hepatoenteritis among an aboriginal population on Palm Island (Queensland, Australia) (Bourke et al., 1983). An epidemiological investigation revealed that 139 children and 10 adults were affected shortly after a copper sulfate treatment of a dense algal bloom in Solomon Dam, the source of the island’s reticulated water supply. All of the affected individuals were using water from Solomon Dam and no incidence of disease was reported among persons not receiving water from this supply. The disease had three welldefined phases which presented in the following order: hepatitis phase-2 days; lethargic phase with severe electrolyte derangement-1 to 2 days; diarrheal phase-5 days. Repeated sanitary assessments of the environment of Palm Island failed to uncover any other possible cause for the outbreak. Falconer et al. (1983a) examined the results of routine assays for hepatic enzymes in plasma of persons who obtained drinking water from a reservoir (Malpas Dam, Armidale, New England, Australia) containing a heavy bloom of toxic M . aeruginosa during periods before, during, and after the algal bloom. These results were compared with corresponding assays from an adjacent population which did not use water from this source. The residents supplied with water from the bloom-infested Malpas Dam reservoir showed a significant rise in y-glutamyltransferase (GGT) during the bloom period, while no such increase occurred in residents not receiving their water from the Malpas Dam. ALAT also showed an increase during this period, but it was not statistically significant (p I 0.10). ASAT and alkaline phosphatase showed no significant increases. GGT is characteristically released after alcohol or toxin damage to liver cell membranes and is a more sensitive indicator of liver damage than alkaline phosphatase or ASAT. The toxin of M . aeruginosa is normally contained within the algal cells and is released only when the cell is damaged, either by poisoning the algae with copper sulfate, by mechanical rupture of the cell, by breakdown in the stomach, or age-related death of the cell. Damage could occur to the cell when algae in a reservoir are transported through a municipal water system; therefore presence of the blooms in a water supply reser-

ALGAL TOXINS

75

voir might be expected to result in the presence of the toxin in drinking water. Some toxins and odor and flavor organics associated with algal blooms can be removed by filtration of reservoir water through sand topped by a layer of granular activated carbon (Falconer et al., 1983b). The exact composition and structure of the Microcystis toxin have remained elusive, despite the efforts of many scientists since the attempts of Louw (1950) to identify the toxic principle from a Microcystis bloom in South Africa. Considerable variation exists in the toxins from different strains of the algae. The toxin also appears to have an unusual structure which does not lend itself to classical techniques. Working with different strains and utilizing different procedures for purification, most workers now agree that the toxin is a pentapeptide, with some common and some variable amino acids. On hydrolysis the toxin is found to contain five amino acids and methylamine in approximately equimolar amounts. Amino acids isolated from all various toxin preparations invariably include aspartic acid (or P-methylaspartic acid), glutamic acid, and alanine. Glutamic acid and alanine, the invariant amino acids of Runnegar and Falconer (1981) and Elleman et al. (1978), and methionine, have no free carboxyl groups while the a-carboxyls of glutamic and P-methylaspartic acid were shown to be free. The toxin does not react with ninhydrin, nor were any amino groups dansylated in the intact toxin, indicating the absence of any free amino groups (Bishop et al., 1959; Murthy and Capindale, 1970; Rabin and Darbre, 1975; Toerien et al., 1976; Elleman et al., 1978; Botes et al., 1982a; Eloff et al., 1982; Santikarn et al., 1983). From amino acid analysis, a minimum molecular weight of 654 was derived by Runnegar and Falconer (1981). Other proposed molecular weights, derived by different researchers from various strains and utilizing different techniques, have also been reported (Runnegar and Falconer, 1981). The existence of the toxin as dimers, trimers, or even larger groups of identical or similar subunits could explain the wide variability of proposed molecular weights of the toxin (Eloff et al., 1982). The lack of free amino groups has led to the speculation that the toxin is cyclic (Bishop et al., 1959; Santikarn et al., 1983; Williams, 1983; Botes et al., 1984) and/or has a blocking group on the terminal amide group (Botes et al., 1982a,b). The extreme hydrophobicity of Microcystis toxins, as exemplified by their chromatographic behavior on paper, cannot be accounted for in terms of their peptide composition and could logically reside in the properties of such a blocking group. The UV spectrum of the toxin shows an absorption maximum at 240 nm; again, this cannot be accounted for by the peptide portion of the molecule, since Botes et al. (1982a) found no

76

WAYNE W. CARMICHAEL

aromatic amino acid present. They do suggest that this absorbance peak could be due to the presence of a conjugated diene chromophore in the blocking group. Eloff et al. (1982) found that the toxins present in different strains were very similar, but variations in toxin composition were found. In some organisms, as many as six different toxins were obtained, but generally one or two toxins accounted for >90% of the toxin in a single isolate. In all cases the toxin contained P-methylaspartic acid, glutamic acid, alanine, methylamine, and two other amino acids in equimolar ratios. The additional amino acids present in major toxins were as follows:-leucine and arginine, leucine and alanine, tyrosine and arginine, methionine and arginine, leucine and tyrosine, alanine and tyrosine, or arginine and arginine. The peptide containing leucine and arginine was present in 9 of the 13 toxic isolates. Botes et al. (1982a) isolated four toxin variants (BE-2 to BE-5) from a laboratory strain of Microcystis aeruginosa (WR-70). The strain was cultured in a modified Volk and Phinney medium (1968) with the trace element mix of Stanier et al. (1971). All variants were composed of five amino acid residues with one residue each of P-methylaspartic acid, glutamic acid, and alanine common to all. For the remaining two residues combinations of leucine and arginine, tyrosine and arginine, leucine and alanine, and tyrosine and alanine were found. Methylamine was detected in acid hydrolysates in all cases. Configuration assignments (Botes et al., 1982b) of the a-carbon atom of the amino acid residues have been made by stereospecific enzymatic transformations, showing that the constant residues are in the D-form, whereas the L-configuration could be assigned to all the variant residues. The relative configuration of the @carbon atom of P-methylaspartic acid could be made by comparison of the electrophoretic mobility of the toxin-derived residue with literature values for the authentic compound. The presence of N-methyldehydroalanine, which gives rise to methylamine upon acid hydrolysis, has been confirmed by identifying N-methylalanine in the acid hydrolysates of the toxin variants, after reduction of the toxin with sodium borohydride. The use of 400 MHz proton NMR spectroscopy showed the toxins to be more complex than suggested by amino acid analysis alone. Apart from the unambiguous assignment of the amino acid residues, an apolar side chain of 20 carbon atoms was demonstrated. The toxin is thought to exist in dimers, trimers, and other larger aggregates with each subunit consisting of a pentapeptide plus an apolar side chain. In this study the molecular weight of each subunit was estimated at 909. Based on mass spectrometry, using the BE-4 toxin, Santikarn et al. (1983) and Williams (1983) proposed that the blocking group is a highly

77

ALGAL TOXINS

unsaturated hydrocarbon with a molecular weight of approximately 3 13. This would be in addition to the 909-Da subunit of BE-4 toxin. The structure of this side group was reported to be a novel p-amino acid. There is some new evidence that the peptide may be cyclic. This would explain the resistance of the intact toxin to Edman degradation and degradation by proteolytic enzymes. Neither fast atom bombardment mass spectrometry nor the electron impact mass spectrum of methylated Microcystis toxin BE-4 shows evidence of the sequence of ions normally observed in the spectra of linear peptides (Santikarn et al., 1983; Williams, 1983). Botes et al. (1984) showed that the p-amino acid residue was a part of the linear amino acid sequence. The molecular weight of the entire toxin was concluded to be 909. They also proposed the term cyanoginosin for the monocyclic heptapeptides. To designate the variable Lamino acids they propose a two-letter suffix after cyanoginosin. Thus the BE-4 toxin becomes cyanoginosin-LA (Fig. 9). A low-toxicity strain of M. aeruginosa collected by Watanabe and Oishi (1980, 1982) and Watanabe et al. (1981) from Lake Suwa in Japan, in contrast to reports of toxins obtained from other world-wide sources, was ninhydrin positive, inactivated by proteases, and not toxic when administered orally to mice. The main amino acids composing this toxin were glutamic acid, aspartic acid, alanine, glycine, arginine, and leucine. The molecular weight was reported to be approximately 2950, with a minimum of 770, as determined by HPLC. There have been no reported livestock deaths from Microcystis in Japan. Siegelman et al. (1984) presented a method for microdetection of Microcystis toxins. This involves extraction of 20 mg of lyophilized cells with 1 cm3of 38% ethanol, 5% n-butanol, 50 mM ammonium acetate for 1 hr, followed by centrifugation for 5 min at 12,000 g . One cm3 of n-butanol followed by 1 cm3 of water are added, with vortexing, to the supernatant. The sample is then centrifuged (10 min at 500 g), and the upper phase of nbutanol is collected and dried by evaporation. The residue is extracted with I cm3 of 26% acetonitrile, 500 mM ammonium acetate, stored at -10°C for 16 hr, and then centrifuged for 10 min at 12,000 g . These partially purified extracts are stable for several months when stored at - 10°C.

r

o-Ala-m-Masp-B-Adda

- o-Glu-

Mdha

1

Fig. 9. Proposed general structure for cyclic heptapeptide toxins of Microcystis aeru1984). X and Y, variable L-amino acids which can differ between strains. Masp, P-methylaspartic acid; Adda, p-amino acid residue of 3-amino-9-methoxy2,6,8-trirnethyl-10-phenyldeca-4,6-dienoic acid; Mdha, methyldehydroalanine. ginosa (Botes er a/.,

78

W A Y N E W . CARMICHAEL

Aliquots of 5 to 25 mm3 of these extracts are examined by HPLC as follows : Precolumn-pellicular octadecyl (C-18) beads (SynChropak, RSC) (0.4 x 5.0 cm) Column-3-pm octadecyl (C-18) silica (Hypersil, Shandon) (0.46 x 5.0 cm) Solvent-26% acetonitrile, 500 mM ammonium acetate, pH 6.0 Flow- 1.O cm3 min-I Detection-238 nm at 0.04 AUFS (absorbance units full scale)

In our laboratory we are using a modification of this procedure to purify and compare toxic peptides of Microcystis, Anabaena, and Oscillatoria. These modifications employ C- 18 semipreparative columns, Sephadex G-25 gel filtration columns, and Bond Elut C-18 preparative cartridges (Theiss, 1984; Carmichael, unpublished data). Kirpenko et al. (1976) have reported on the use of an enzyme photometric assay to detect Microcystis toxins. The method uses cholinesterase and acetylcholine in the presence of the toxin to yield a product which can be measured with the color indicator bromothymol blue.

C. MARINE CYANOPHYTE TOXINS

Marine cyanobacteria have not presented as many problems as the freshwater cyanobacteria, but several species of the family Oscillatoriaceae (unbranched filamentous, lacking heterocysts) produce toxic compounds that cause dermatitis in man and animals and promote tumors in laboratory animals. These toxins may also have toxic affects on some forms of marine life (Moore, 1981b). The most common and best documented toxic reaction to marine cyanobacteria is a severe contact dermatitis known as “swimmer’s itch.” This is a cutaneous inflammation characterized by erythema, followed by blisters and deep desquamation within 12 hr of exposure to the algae. The dermatitis is induced by exposure to two compounds, debromoaplysiatoxin and lyngbyatoxin A (Fig. lo), which have been isolated from Lyngbya majuscula (Moikehai and Chu, 1971; Moikehai et al., 1971; Kato and Scheuer, 1975; Hashimoto et al., 1976; Mynderse et al., 1977; Cardelh a et al., 1979). Debromoaplysiatoxin has been isolated from deep water strains of L . majuscula; while lyngbyatoxin A is found in a Hawaiian shallow water variety of L . majuscula. The dermatitis produced by lyngbyatoxin A is the same as that from debromoaplysiatoxin. However, the shallow water variety of Lyngbya majuscula has never been impli-

ALGAL TOXINS

79

cated in an outbreak of swimmer’s itch, mainly because it grows on the leeward side of Oahu and swimmers rarely come into contact with the broken filaments of the alga in the water, since the normal tradewinds blow them out to sea rather than toward the shore. Debromoaplysiatoxin, along with several closely related compounds, has also been isolated from other Oscillatoriaceae (Mynderse and Moore, 1978; Moore, 1981b). Debromoaplysiatoxin and oscillatoxin A, for example, are major toxic constituents of a mixture of cyanobacteria from Enewetak identified as Schizothrix calcicola and Oscillatoria nigroviridis. Recent work (Moore et al., 1984) shows that oscillatoxin A has a structure similar to the aplysiatoxins. Solomon and Stroughton (1978) reported reactions to debromoaplysiatoxin occurring in humans at concentrations of 5.6 mg ~ m and - ~in making it one of the most potent dermatotoxins animals at 0.5 pg known. Since swimmers who come in direct contact with toxic strains of the algae develop erythema, followed by blisters and deep desquamation, within 12 hr of exposure, the active agent is classed as a primary irritant rather than an allergen. During animal testing, a second application of the debromoaplysiatoxin produced a similar reaction to the first, which is not suggestive of sensitization. Debromoaplysiatoxin, a phenolic substance, has been isolated from the midgut gland of the sea hare (Stylocheilus Eongicauda) along with a bromine-containing analog bromoaplysiatoxin (Kato and Scheuer, 1976). Lyngbya majuscula is a favorite food of the sea hare, and the digestive tract of this animal appears to be unaffected by the toxin. The sea hare, in fact, utilizes an unknown metabolite in the algae for metamorphosis (Switzer-Dunlap and Hadfield, 1977). Hashimoto et al. (1976) observed rabbitfish (Siganus fuscescens) feeding on sea grass entangled with L . majuscula, but observed no harm to the digestive tract of the fish. However, he suggested that ingestion of L . majuscula might be related to an epidemic of rabbitfish poisoning among the natives in the Ryukyus Islands in 1969, but no further study was made of this. Other marine life infesting toxic Lyngbya may not be so fortunate as the sea hare and the rabbitfish. Lightner (1978) suggested that Lyngbya spp. might be responsible for mass mortalities of raceway-reared blue shrimp (Penaeus stylirostris) in the large shrimp culture facility in Penasco, Mexico. The affected shrimp had severe necrosis in the epithelia lining the midgut, dorsal cecum and hindgut gland, along with subsequent hemocytic enteritis. However, Moore (1981a) could detect no mouse toxicity or activity against P-388 lymphocytic mouse leukemia in lipophilic extracts of a sample of the toxic algae obtained from Lightner’s laboratory in 1979. The appearance of the necrosis in the shrimp was slow and Moore sug-

80

WAYNE W . CARMICHAEL

gests that perhaps a very small contaminant in the algae that normally grow in the rearing tanks may have been responsible. Debromoaplysiatoxin and lyngbyatoxin A, isolated from the lipophilic extract of Lyngbya majuscula, show activity against P-388 lymphocytic mouse leukemia at sublethal doses (Mynderse et al., 1977; Cardellina et al., 1979), as do the aqueous extracts of some marine Oscillatoriaceae (Kashiwagi et al., 1980). Water-soluble toxins are also present in several other cyanobacteria belonging to the Oscillatoriaceae, but none of these has been isolated and characterized (Moore, 1981b). Cooper (1964) reported that the natives in Marakei atoll associate toxic fish with an alga they called “tan-tan.” This cyanobacteria was identified as Schizothrix calcicola, which contains both lipid- and water-soluble toxins which had a small amount of anticholinesterase activity (Banner, 1967). Lyngbya majuscula also possess water-soluble toxins (Kashiwagi et al., 1980)but little is known of these toxins at the present time. The gross structure of lyngbyatoxin A, as determined by chemical and spectral data, is the same as that of teleocidin A (Fig. lo), a poisonous substance associated with several strains of Streptomyces (Cardellina et al., 1979; Sakabe et al., 1966). Teleocidin was first isolated from the mycelia of Streptomyces mediocidicus (Takashima and Sakai, 1960). It is now known that teleocidin is composed of two compounds, teleocidin A ( M , = 437) and teleocidin B ( M , = 451) (Fujiki et al., 1982a, 1984). Teleocidin A produces dermatotoxic effects that are the same as those of lyngbyatoxin A and both are highly toxic to certain types of fish. Lyngbyatoxin A, bromoaplysiatoxin, debromoaplysiatoxin, and teleocidin are also reported to be tumor promoters in mice (Fujiki et al., 1981, 1983, 1984). The in uitro effects of these substances are identical to tetradecanoylphorbal 13-acetate (TPA), the tumor-promoting constituent of croton oil. To screen for the tumor-promoting activity of these compounds a three-test procedure was used. The first test is on irritation of mouse ear, the second on induction of ornithine decarboxylase (ODC) in the skin of the back of mice, and the third on adhesion of human promyelocytic leukemia cells (HL-60) (Fujiki et al., 1984). Using these tests it was shown that lyngbyatoxin A, bromoaplysiatoxin, and teleocidin were all potent promoters, but debromoaplysiatoxin was much weaker. In addition to the above, lyngbyatoxin A and teleocidin show a number of effects on mammalian cells in culture that are similar to those of TPA. These include induction of differentiation of human promyelocytic leukemiacells (HL-60) (Nakayasu et al., 1981; Fujiki et al., 1982b), aggregation of human lymphoblastoid cells (Hoshino et al., 1980), and stimulation of prostaglandin production and choline turnover in HeLa cells. All of these effects are similar to those produced by the structurally dissimilar phorbol ester tumor promoter TPA (Sakamoto et al., 1981).

81

ALGAL TOXINS

OH

bh

B

Fig. 10. (A) Debromoaplysiatoxin and bromoaplysiatoxin (R = Br), a dermatotoxic phenolic bislactone produced by several genera of marine cyanobacteria. (B) Lyngbyatoxin A, a dermatotoxic indole alkaloid, is produced by Lyngbya majuscula. This toxin has the same structure as teleocidin A , which is produced by strains of the filamentous bacterium Streprornyces.

The discover of these tumor-promoting properties suggests that, in addition to the problem posed by the dermatotoxicity of L y n g b y a majuscula, the toxins produced by these cyanobacteria could conceivably be involved in the development of human cancer, even though there is no direct evidence that L y n g b y a majuscula has caused human cancer. Freshwater cyanobacteria have not been investigated for the production of these dermatitis toxins with tumor-promoting activity. Some evidence exists that they are being produced however. This includes reports of contact irritation from freshwater recreational waters (Billings, 1981; Carmichael et al., 1985; Carmichael and Codd, unpublished data). It appears that freshwater cyanobacteria are capable of producing three

82

WAYNE W . CARMICHAEL

groups of toxins: neurotoxic alkaloids, hepatotoxic peptides, and the as yet unidentified dermatitis toxins which may have some similarity to marine cyanobacteria toxins. These three groups are in addition to the presence of lipopolysaccharide endotoxins, which are a part of the cyanobacterial cell wall and do not apparently contribute to their environmental toxicity (Raziuddin et al., 1983). D . DINOFLAGELLATE PARALYTIC SHELLFISH POISONS

(PSP)

PSP has been known as mussel or clam poison depending on the source of extracts. Occasionally it has been called plankton poison and mytilotoxin. The poison was later named saxitoxin (STX), a name derived from Saxidomus gigenteus (Alaskan butter clam), which was then thought to be the only toxic material responsible for all PSP along the Pacific coasts (Schuett and Rapoport, 1962). From the early work of Meyer et al. (1928) and Muller (1939), it was observed that the poison was water soluble, acid stable, and alkaline labile. Using these observed properties, Schantz et al. (1957) was able to purify saxitoxin using carboxylate cation exchange resins. Further analysis of the purified poison as the dihydrochloride.salt (white hygroscopic powder) indicated a molecular weight of 300 to 400 (diffusion coefficient of 4.8 X cm2 sec-I; pK, of 8.2 and 11.6; specific optical rotation of 130 2"). Saxitoxin was also found to be insoluble in lipid solvents, could be detoxified by hydrogen reduction, and showed no absorption above 220 nm (Schantz et al., 1957, 1966; Schantz, 1960). The correct structure of saxitoxin (Fig. 11) was shown to be an alkaloid plus a purine (Schantz et al., 1975). Schantz (1960) and hvans (1970) found out that a large component of toxins isolated from North Atlantic mussels was not being retained by the carboxylate cation exchange resins. With modifications in the purification method involving gel filtration on polyacrylamide gel (Shimizu et al., 1975) and TLC (Buckley et al., 1976), they were able to identify the unretained toxins as derivatives of saxitoxin. It was obvious then that saxitoxin was not the sole cause of toxicity. Ghazarossian et al. (1974) suggested that the saxitoxin derivatives were n-oxides. Subsequently Shimizu et al. (1978) and Boyer et al. (1978) demonstrated structurally that the derivatives are n-1-hydroxysaxitoxin and 1 1-hydroxysaxitoxin sulfates, which are commonly known as neosaxitoxin and gonyautoxins. Hall et al. (1980) and Koehn et al. (1982) isolated and identified other toxins including saxitoxin from Pacific isolates of cultured Protogonyaulax spp. This array of 12 toxins can be divided into 2 groups (Fig. 11):

*

ALGAL TOXINS Carbamate

- R4

83

(H)

&l& H H H

OH OH OH

H H OS03H H OSO,

-

H OSO, H H OSO, H

N Sulfocarbamoyl

STX GTXz GTX 3 neoSTX GTXl GTX4

H

- Rg (SO3-)

R1R2b H

H H OH OH OH

H H OS03H

H OSO,

H OSO, H H OSO, H

B 1(GTX5) C1 C2(GTXe) BP(GTX6) C3 c4

Fig. 1 1 . Structure of paralytic shellfish poisons produced by several genera of marine dinoflagellates. At least some of these toxins have been found in species of Protogonyuulux, Pyrodinium, the red alga Juniu, and the freshwater cyanobacterium Aphanizomenon (Carmichael er al., 1985).

1 . The carbamate toxins-H is the substituted group at &. This group includes saxitoxin, neosaxitoxin, gonyautoxins 1 , 2, 3, and 4. 2. The N-sulfocarbamoyl toxins-sulfite (SO3-) is the substituted group at R4 and includes B1 (gonyautoxin 5 ) , B2 (gonyautoxin 6), C1, C2 (gonyautoxin 8), C3, and C4.

Another naturally occurring toxin that had been reported by Hsu et a f . (1979) and Wichmann et al. (1981) (as an unknown) is gonyautoxin 7. A similar array of toxins has been reported from sources found in subtropical and tropical waters (Yasumoto et a f . , 1983). Oshima et a f . (1984) demonstrated that the tropical marine dinoflagellate Pyrodinium bahamense var. compressa can produce STX, neoSTX, GTXS, GTX6, and decarbamoylsaxitoxin (dcSTX). In addition they showed that the red algal genus Jania produced gonyautoxins which were ingested by coral reef crabs and gastropods. In preliminary tests they found that these gonyautoxins could be bioconverted in the digestive organs of these crabs to STX. In addition, Harada et al. (1982) reported the presence of unidentified new toxins from turban shells (actually top shells) which were designated as TST (turban shell toxin). The relative potencies of these toxins were found to be different. Sax-

84

WAYNE W . CARMICHAEL

itoxin at an LDsoof 10 pg kg-' or 5000 MU mg-' (1 MU is defined as the amount of toxin needed to kill a 20 g mouse in 15 min) is the most potent (Schantz et al., 1966). Hall (1982) and Proctor et al. (1975) showed that the potencies of some of these toxins change upon hydrolysis (0.2 M HCl, 100°C for 5 min) indicating transformations. Toxins B1, C1, C2, B2, C3, and C4 after hydrolysis were found to convert to STX, GTX2, GTX3, neoSTX, GTXI, and GTX4, respectively, indicating hydrolysis of the R4 group (SO3-) (Hall et al., 1980). Neosaxitoxin is as potent as saxitoxin (Shimizu et al., 1978; Hsu et al., 1979; Hall, 1982) even though Genenah and Shimizu (1981) reported a much lower value (2363 ? 101 MU mg-I). Potencies, according to Hall (1982), are STX and neoSTX > GTX3 > C2 > B2 > B1 > C1 (no data given for GTX,, C3, GTX4, and C4; while according to Genenah GTX3 was > STX > GTXl > neoSTX > GTX2 > GTX4 > B1 (GTX5). As mentioned earlier, characterization studies of STX indicated two dissociable groups with pK, values of 8.3 and 11.6 -+ 0.1. Rogers and Rapoport (1980) reported values of 8.22 and 11.28. A similar titration of neoSTX by Shimizu et al. (1978) revealed a third dissociable group with a pK, of 6.75. These pK, values correspond to the dissociation of N-lhydroxyl (6.75), C-8 guanidium (8.5), and C-2 guanidium (1 1.6). The observed differences in the binding to carboxylate cation exchange resins are due to the presence of charges under the acidic condition. STX and neoSTX have a net charge of + 2 whereas GTXs (1 through 4) have a net charge of + 1 (Boyer et al., 1978). Hence it was once postulated that the net charge was the sole determinant of potency. However, structural variation studies on potency of synthetically derived material indicated otherwise. Manipulations of C-I2 OH groups (Fig. 11) made the molecule completely devoid of activity (Kao et al., 1981; Kao and Walker, 1982). Removal of the carbamoyl group (-CONHz) reduces potency to less than 70% of that of STX (Kao and Walker, 1982). Also effective in reducing potency (in uiuo by 15 times that of STX) is the substitution of the carbamate group with a sulfo group (SO3-) (Hall, 1982). Symptoms of paralytic shellfish poisoning are primarily neurologic and appear rather quickly, often within 30 min after ingestion of contaminated shellfish. Most deaths occur from respiratory paralysis within 12 hr of illness. The prognosis is considered good if one survives after 24 hr (Schantz, 1971). The amounts of toxins ingested to cause illness vary with the individual but ranges from 3000 to 20,000 MU. The illness is characterized by paresthesia of lips, mouth, face, and extremities, often followed by nausea, vomiting, weakness, and incoherence. As the illness progresses, respiratory distress and muscular paraly-

ALGAL TOXINS

85

sis become more severe and death results from respiratory paralysis. In a large outbreak of PSP in England in 1968, the common symptoms reported were paresthesia, weakness of extremities, lightheadedness, and floating sensation (McCollum et al., 1968). Since the toxins leave the system quickly, resuscitation is recommended for treatment. But in severe cases, treatment should consist of a cathartic or enema to remove the unabsorbed toxins from the intestinal tract. Gastric lavage should be considered if vomiting has not occurred (Hughes and Merson, 1976). Artificial respiration should be supplied if respiratory difficulties appear (Schantz, 1971). Evans (1965) and Kao (1975) reported some hypotensive effects of the toxins with high doses. Potency varies widely depending on the recipient and routes of administration. From experiments with mice, rats, and rabbits, intraperitoneal (ip) or intravenous (iv) administration of STX is far more potent than oral administration, ranging from 10 to 50 times in potency (Evans, 1972). Thus substances that increase intestinal absorption or peripheral circulation will definitely intensify the symptoms of illness. The mechanism of action for saxitoxin is interference of the transmission of electrical impulses along the nerve and muscle membranes. The toxin causes a reversible blockage of the permeability to sodium ions without affecting permeability to potassium ions. Thus the propagation of nerve-muscle action potentials is blocked (Evans, 1964, 1970; Kao and Nishiyama, 1965). Kao and Walker (1982), using analogs of STX in their studies of Na+ channel blocking, showed that active groups of STX were the C-12 OH groups, the 7,8,9 guanidinium and carbamyl group, but not the N-1 of 1,2,3 guanidinium. They proposed that STX binds the receptor located in the outside of the membrane very close to the opening of the Na+ channels. The guanidinium group is then thought to be electrostatically attracted by the fixed anionic charges around the opening of the Na+ channel and the bulk of the molecule (STX) covers up the channel. The idea of having a toxin-binding site distinct from the Na+ channel was recognized by Twarog and Yamaguchi (1975). Later Spaulding (1980) showed that frog muscle fibers subjected to trimethyloxonium ion (which derivates carboxylate groups) became less susceptible to STX blockade. Cohen and Barchi (1981) noted the binding sites to have glycoprotein characteristics. Detection of paralytic shellfish poisons has been a concern for public health and fisheries people for many years. The officially employed method of detection, worldwide, is the mouse bioassay (Horwitz, 1980; Houser, 1965). Because of the variable nature of this assay method and the inability to distinguish types of toxins present other methods are being developed. Carlson et al. (1984) have developed a radioimmunoassay for

86

WAYNE W . CARMICHAEL

saxitoxin but it does not bind to neosaxitoxin. Other immunoassay methods are also being developed including a rabbit antiserum which has successfully been used as a therapeutic agent in mice to prevent death from saxitoxin (Davio and Pickering, 1983). HPLC is also proving a powerful tool in the assay of PSPs. This method uses the fluorescence property of the PSP derivatives when oxidized under alkaline conditions. Initially problems existed in the detection of neosaxitoxin and gonyautoxin 1 and 4 due to its poor fluorescence (Sullivan and Iwaoka, 1983; Sullivan et al., 1983). These problems have been corrected, making it possible to detect by fluorescence all of the known carbamate and N-sulfocarbamoyl toxins of PSP (Sullivan and Wekell, 1984). E. DINOFLAGELLATE CIGUATERA TOXINS

Ciguatoxin (CTX) is the name given to the first active component isolated which was felt to be responsible for ciguatera poisoning. It is a lipidsoluble neurotoxin. It imparts no different taste or color to the fish and is not affected by cooking or freezing. It was isolated from contaminated red snapper (Lutjanus),shark (Carcharhinus), and the moray eel (Gymnothorux) by a series of organic solvents, liquid-liquid partition, and chromatography. The isolated toxin is a transparent, yellow oil. This final product had an MLD (minimum lethal dosage) of 0.5 mg kg-' and was identified as having the molecular formula CuH65NOs,with the nitrogen being quaternary (Scheuer et al., 1967). Since then it has been further purified and found to have a polyether structure similar to okadaic acid or brevetoxin C, with a molecular weight of about 1111 (Nukina et al., 1983), a crude extract LD5o of 28.1 mg kg-', and a purified toxin LDw of 0.25 pg kg-' intraperitoneally in mouse (Hokama et al., 1984). Research on ciguatera toxin at the University of Southern Illinois found that there is more than one ciguatoxin-like compound. From cultures of Gambierdiscus toxicus grown in the laboratory, three distinct lipid toxins, GT-1, GT-2, and GT-3, were isolated, which all act as competitive antagonists against acetylcholine and histamine sites on guinea pig ileal smooth muscle, and whose effects are not offset by calcium (Miller et al., 1984). It may be that these toxins are individual cyclic polyether residues similar to okadaic acid isolated by Tachibana et al. (1981). Another toxin thought to contribute to ciguatera poisoning is the watersoluble substance known as maitotoxin (MTX), which is found in the liver and gut of affected fish (Yasumoto et al., 1971) and from laboratory cultures of G . toxicus (Dickey et al., 1984). It has been suggested that MTX is a precursor of CTX (E. P. Ragelis, personal communication). On the isolated guinea pig atrium, MTX shows positive inotropic effects similar

ALGAL TOXINS

87

to CTX, but only at low concentrations; and a decrease in contactibility leading to complete suppression with higher concentrations (Miyahara et al., 1979). In anesthetized cats, MTX induced hyperventilation, then respiratory depression, slight bradycardia, with hypertension at low doses and cardiac arrest prior to respiratory arrest. MTX seemed to be more potent in these experiments than CTX (Legrand et al., 1982). MTX has been reported as having a lethal dose ranging from an MLD of 0.2 pg kg-l (Takahashi, 1982) to 1.1 mg kg-' (Dickey et al., 1984). It has also been demonstrated to cause an increase in calcium ion influx into cells, a release of norepinephrine (Takahashi et al., 1982), and to inhibit Na+- and K+-ATPase from microsomes of cat and human kidneys (Bergmann and Nechay, 1982). MTX has been isolated from cultures of G. toxicus and following a latent period it produces irreversible contraction of the guinea pig ileum (Dickey et al., 1984). A third, and less well-studied, toxin thought to contribute to ciguatera poisoning was isolated from the parrotfish (Chungue ef al., 1977) and termed scaritoxin (Bagnis et al., 1974). Individuals having eaten this toxic fish showed initial symptoms similar to ciguatera poisoning but then after 5 to 10 days progressed into a second phase which persisted for more than a month, consisting of equilibrium disorders and kinetic tremors. Like CTX, it is lipid soluble and distinguished from it by its chromatographic properties. Likewise, many of its physiological properties are similar and it has been suggested that it might be a metabolite of CTX. Several attempts have been made to find more reliable methods of detecting the presence of ciguatoxin than the sometimes used cat, mongoose, or mouse bioassays (Halstead, 1967). A radioimmunoassay (RIA) using CTX-human serum albumin to produce anti-CTX in sheep was developed by Hokama et al. (1977). Testing has shown that this method is relatively accurate but economically feasible only in fish weighing greater than 9 kg (Kimura et al., 1982a). This RIA method shows close correlation with mouse bioassay and the in uitro guinea pig atrium assay (Miyahara et al., 1979), and cross-reacts with okadaic acid, a toxic component of sponge (Kimura et al., 1982b). The mouse bioassay involves injecting a fish extract intraperitoneally into mice and watching symptoms for a 48-hr period. Symptoms have been shown to be dose dependent and include lowered rectal temperature, reduced pain reflexes, reduced grip and tightrope responses, reduced locomotor activity, and death (Hoffman et al., 1983). An enzyme immunoassay has been developed and is being used in parts of Hawaii to survey fish (Hokama et al., 1983). This enzyme immunoassay is reported to be as sensitive as the radioimmunoassay but is easier to run, economically feasible for screening all sizes and varieties of fish, and could be used for testing liver in addition to flesh samples. The

88

W A Y N E W . CARMICHAEL

enzyme assay uses the sheep anti-ciguatoxin used for the radioimmunoassay but instead of iodine employs horseradish peroxidase to couple with the antibody (Hokama et al., 1984). The symptoms of ciguatera disease begin within minutes of ingestion to 30 hr afterward. As there are no definitive tests used to diagnose ciguatera poisoning, it is made solely from symptomatology and history. It is often differentiated from other types of poisoning by the presence of paresthesia of the extremities. The first symptoms present are usually gastrointestinal, lasting a few hours; followed by neurological disturbances which last up to several weeks or even months (Halstead, 1967). Additional symptoms can be dermatological and neuromuscular (Morris et al., 1982; Lawrence el al., 1980). Ten to 12% of the intoxications lead to death by respiratory paralysis (Rayner, 1972). With severe poisoning, bradycardia was observed and an electrocardiogram showed inverted T waves in some patients, indicating myocardial ischemia, which was reversible upon recovery (Hanno, 1981). Symptoms have been known to vary in different locations and with ingestion of different fish (Chretien et al., 1981). An unexplained sensitization has often been observed after ciguatera poisoning and patients often experience the same symptoms of poisoning after the ingestion of nontoxic fish or after alcohol ingestion (Barr, 1982). Banner (1967), however, has suggested that this may be psychosomatic. Treatment mostly involves supportive care, but may include gastric lavage, cathartics, atropine, and protopam chloride (Dembert and Pearn, 1982; Bagnis, 1967). Alkaloids from the plant Duboisia myoporoides have been used by natives of New Caledonia as an antidote, and are thought to contain atropine, nicotine, hyoscyamine, and scopolamine (Dufva et al., 1976). Ciguatoxin has been shown to have anticholinesterase activity in uitro (Li, 1965) but not in uiuo (Rayner, 1968). Kosaki and Anderson (1968) observed depressed respiration, a fall in arterial pressure, bradycardia, arrhythmias, and neuromuscular responses in rats after administration of lethal doses of ciguatoxin extracted from various fish, and suggested that the toxicity may be due to demyelination of peripheral and central nervous tissue. In anesthetized cats, ciguatoxin caused respiratory depression, leading to respiratory arrest with bradycardia. An ECG indicated ventricular extrasystole and idioventricular rhythm (Legrand et al., 1982). Setliff et al. (1971) demonstrated that ciguatoxin increases the passive permeability of the cell membrane of frog skin, thus increasing the influx of sodium ions by competitively inhibiting calcium binding at receptor sites which regulate steady state sodium ion permeability. This suggested that possibly ciguatoxin inhibited Na+,K+-ATPase activity. However, when highly purified extrakts of ciguatoxin were used, no inhibition of

ALGAL TOXINS

89

Na+,K+-ATPase was seen, indicating that impurities may have had an effect on some previous results (Rayner and Szekerczes, 1973). A biphasic response is observed on isolated atria from rats and rabbits, with an initial inhibition followed by stimulation of atrial contraction, indicating both cholinergic and adrenergic action and possibly the release of catecholamines (Ohshika, 1971). CTX was later shown to exhibit both positive inotropic and chronotropic effects on the isolated atria of the guinea pig (Miyahara et al., 1979). CTX has also been shown to induce contraction of the guinea pig vas deferens, by releasing stores of norepinephrin, which is probably due to the increased permeability to sodium ions (Shizumi et al., 1981). Possible effects of the ciguatera disease on an unborn infant were observed by Pearn et al. (1982). A cesarean section was performed 2 days after a full-term gravid woman became poisoned with ciguateric fish. The woman experienced fetal movements which began simultaneously with her own symptoms. The infant was born with left-sided facial palsy, but with no heart abnormalities. This suggest that the toxin can cross the placenta, which would not be surprising due to its small molecular size. F . OTHER DINOFLAGELLATE TOXINS

While several other dinoflagellate genera have some species producing various toxic factors, the main genera other than Protogonyaulax (Gonyaulax) involved in poisonings are Ptychodiscus (formerly in the genus Gymnodinium) and Dinophysis (Taylor, 1984). The best known toxin producer in Gymnodinium is the fish killer G . breve. Recently it was transferred to the genus Ptychodiscus by Steidinger (1979). Baden et al. (1984) was able to isolate and purify two toxins from laboratory cultures of P . breuis termed T34 and T17. Both polyether toxins are neurotoxic and ichthyotoxic. One of the toxins also caused bronchioconstriction in anesthetized guinea pigs, which suggests it is responsible for the airborne respiratory irritant in humans associated with these toxic blooms along the Florida coast. The structure of these two “brevetoxins” is given in Fig. 12. The Dinophysis toxins are collectively referred to as diarrhetic shellfish poisoning (DSP). While the difficulty in culturing these dinoflagellates has slowed definitive studies it is thought that D . fortii and D . acuminata are the main toxic species (Taylor, 1984; Yasumoto et al., 1984). DSP produces gastrointestinal disturbances but not fatalities in humans ingesting various shellfish, i.e., Mytilus, which have concentrated the dinoflagellate. DSP toxins isolated from shellfish in Japan are all polyether-based compounds referred to as okadaic acid, dinophysistoxin, and pecteno-

90

W A Y N E W . CARMICHAEL

a

b

Key: a=T34

b=T17

Fig. 12. Structure of brevetoxins, T34 and T17, produced by the Florida red tide dinoflagellate Ptychodiscus brevis (Baden et al., 1984).

okadaic acid dinophysistoxin- 1 dinophysistoxin-3

: :

:

R 1 = H , R,=H R, = H, R, = CH, R, = acyl, R, = CH,

47

R 40

pectenotoxin-1 pectenotoxin-2

: :

R =OH R=H

Fig. 13. Structure of polyether lactone toxins involved in cases of diarrhetic shellfish poisoning (DSP). The toxins are produced by the dinoflagellate Dinophysis acuminata found in the shellfish areas of northern Japan (Yasumoto et a / . , 1984).

ALGAL TOXINS

91

toxin (Yasumoto er al., 1984). Figure 13 illustrates the structure of these DSP toxins. V. ENVIRONMENTAL ROLE OF ALGAL TOXINS Since so little is known about the ecological or metabolic roles of algal toxins it is important to consider the possible roles in the light of what is known about plant toxins. Since the 1960s evidence has been accumulating to show that vascular plant secondary chemicals have distinct ecological roles (Fraenkel, 1959, 1969). These plant secondary chemicals provide predation protection throughout the growing and dormant seasons. Evidence is now accumulating which shows that chemical protection can be regulated and their production quite rapid. For example, leaves of red oak trees defoliated by gypsy moth larvae show higher secondary chemical production in subsequent years (Schultz and Baldwin, 1982). Responses have also been observed in which the length of time for chemical response has been days to months. Time response can be even more rapid, however, as in the studies of Carroll and Hoffman (1980). They found that damaged leaves of Cucurbita moschata mobilize substances to the damaged region within 40 min. There is even some evidence that damaged plants can chemically communicate the need for increased chemical defense to undamaged plants within the nearby area (Baldwin and Schultz, 1983). This indicates the presence of two broad classes for chemical defenses of plants: those present before herbivore attack and those that change in response to herbivore attack (Levin, 1976). In these cases of secondary chemical production it has been in response to environmental stress. This stress can come from predation pressures as just presented, or from growth pressures such as would be provided by secondary chemical production in response to low soil nutrients (Janzen, 1974; McKey, 1979; Chew and Rodman, 1979). Central to the concept of an ecological role for secondary chemicals is the concept of cost and benefit for the defense of the organisms. If these secondary chemicals are serving an ecological rather than a metabolic (i.e., nutrient storage) role, then the energy cost involved in producing them should be less than the protective value obtained from their presence. While hypotheses abound supporting this idea within secondary plant chemicals, there are no careful quantitative models of costs and benefits (McKey, 1979). With all of these concepts in mind, some points worth considering if algal toxins are to be considered as having an ecological role include the following:

92

W A Y N E W . CARMICHAEL

1, Evidence is accumulating which shows that zooplankton herbivores of algae do respond to the toxins (Snell, 1980; Lambert, 1981; Ransom, 1978; Porter and Orcutt, 1980). What is not clear is just how much, if any, avoidance of toxic cells or filaments occurs relative to nontoxic cells or filaments. If there is a selective advantage to being toxic then it should be possible to show this in laboratory experiments. An important need in the testing of this idea will be the development and use of sensitive assays for toxicity of cells. This would also make it possible to check for increases or changes in toxin production due to predation pressures. Since these algae exist largely as single cells, or in some cases as filaments, there is a need to show chemical communication between the cells if production of toxins as opposed to simple selection of toxic cells is to be demonstrated. For the present it is perhaps too much to envisage that toxin production could be influenced by such mechanisms. At present it is hard to see why many water blooms of cyanobacteria and even dinoflagellates are apparently nontoxic if predation pressures are to have a significant influence on toxin production. 2. What is more tempting to envisage is the possible role of growth conditions on toxin production. This includes the possibility that nutrient composition, pH, temperature, or light may independently or in combination influence toxin production. The algae may in turn produce higher levels or different kinds of toxins in order to minimize predation pressure or to store the toxins for some metabolic need. This seems to be supported by the preliminary work of Hall (1982), who demonstrated higher specific toxin levels per cell in dinoflagellates grown on decreased nitrogen or phosphorus. It also could explain decreases observed in toxin producticin in our laboratory for almost all toxic cyanobacteria when they are grown in nutrient-rich media. This reduction in toxin production is most common among the peptide toxin-producing strains and almost never observed among the alkaloid toxin-producing strains. The decrease in toxin production is also slow, requiring months or more of repeated subculturing before significant reductions are observed. This implies that if toxin production is to be influenced by growth conditions it will take more than a few subcultures. Just as important, however, could be the qualitative changes in toxin production that growth changes could influence. 3. Finally, it has been pointed out that the costs of storing some kinds of secondary compounds are higher than those of others (McKey, 1979). Substances such as tannins and saponins exert their toxic effect by reducing digestion of proteins in the gut of the herbivore. Other toxins such as alkaloids are absorbed from the gut and are active against specific metabolic processes. Rhoades and Cates (1976) argue that the digestion inhibi-

ALGAL TOXINS

93

tors impose higher costs on the plants since they can influence plant processes and thus must be segregated from other cellular components. The more specific metabolic toxins such as alkaloids are effective against herbivores in lower concentrations and are less likely to influence plant metabolism. This perhaps is part of the reason why algae produce the more toxic, more metabolically specific alkaloids, peptides, and related toxins. Whatever the role for algal toxins, it should be evident that (1) they are diverse groups of physiologically potent secondary chemicals that are more chemically related to higher plant toxins than they are to bacterial or animal toxins; (2) their production should not be considered fortuitous or accidental but rather part of evolutionary processes by which the organisms have adapted to particular needs and requirements of their environment; and (3) studies on production of the toxins including possible environmental and genetic influences will serve to aid an overall understanding of secondary chemical production.

REFERENCES Adachi, R., and Fukuyo, T. (1979). Bull. Jpn. SOC.Sci. Fish. 45, 67-71. Adelman, W. J., Jr., Fohlmeister, J. F., Sasner, J. J., Jr., and Ikawa, M. (1982). Toxicon 20, 513-516. Alam, M., and Euler, K. L. (1981). I n “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 405-414. Plenum, New York. Alam, M., Ikawa, M., Sasner, J. J., Jr., and Sawyer, P. J. (1973). Toxicon 11, 65-72. Alam, M., Shimizu, Y., Ikawa, M., and Sasner, J. J., Jr. (1978). J . Enuiron. Sci. Health AW, 493-499. Anderson, D. M. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 125-138. American Chemical Society, Washington, D.C. Anderson, D. M., and Wall, D. (1978). J . Phycol. 14, 224-235. Astrachan, N. B., and Archer, B. G. (1981). I n “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 437-446. Plenum, New York. Baden, D. G., Mende, T. J., Poli, M. A., and Block, R. E. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 359-368. American Chemical Society, Washington, D.C. Bagnis, R. (1967). Bull. SOC.Parhol. Exp. 60,580-592. Bagnis, R., Loussan, E., and Thevenin, S. (1974). Med. Trop. 34, 523-527. Bagnis, R., Chanteau, S., and Yasumoto, T. (1977). Bull. Soc. Parho[. Exp. 3 , 320. Baldwin, I. T., and Schultz, J. C. (1983). Science 221, 277-279. Banner, A. H. (1967). I n “Animal Toxins” (F. E. Russel and P. R. Saunders, eds.), pp. 157165. Pergamon, Oxford. Banner, A. H. (1976). In “Biology and Geology of Coral Reefs” (A. 0. Jones and R. Endean, eds.), Vol. 111, pp. 177-211. Academic Press, New York. Barr, M. (1982). Med. J. Aust. 1, 14-15. Bates, H. A., and Rapoport, H. (1979). J . Am. Chem. SOC. 101, 1259-1265.

94

WAYNE W . CARMICHAEL

Beasley, V. R., Coppock, R. W., Simm, J., Ely, R., Buck, W., Corley, R. A., Carlson, D. M., and Gorham, P. R. (1983). J . A m . V e t . Med. Assoc. 182, 413-414. Bergmann, J. S., and Nechay, B. R. (1982). Fed. Proc., Fed. A m . Soc. Exp. Biol. 41,7547. Billings, W. H. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 243-255. Plenum, New York. Bishop, C. T., Anet, E. F. L. J., and Gorham, P. R. (19.59). Can J . Biochem. Physiol. 37, 453-47 1. Botes, D. P., Kruger, H., and Viljoen, C. C. (1982a). Toxicon 20, 945-954. Botes, D. P., Viljoen, C. C., Kruger, H., Wessels, P. L., and Williams, D. H. (1982b). Toxicon 20, 1037-1042. Botes, D. P., Tuinman, A. A., Wessels, P. L., Viljoen, C. C., Kruger, H., Williams, D. H., Santikarn, S., Smith, R. J., and Hammond, S. J. (1984). J . Chem. SOC.Perkin Trans. I, 23 11-23 18. Bourke, A. T. C., Hawes, R. B., Neilson, A., and Stallman, N. D. (1983). Toxicon Suppl. 3, 45. Boyer, G. L. Schantz, E. J., and Schnoes, H. (1978). J. Chem. Soc. Chem. Commun. 889890. Buckley, L. J . , Ikawa, M., and Sasner, J. J., Jr. (1976). J . Agric. Food Chem. 24, 107-1 I I . Campbell, H. F., Edwards, 0. E., and Kolt, R. J. (1977). Can. J . Chrm. 55, 1372-1379. Campbell, H. F., Edwards, 0. E., Elder, J. W., and Kolt, R. J. (1979). Pol. J . Chem. 53,2736. Cardellina, J. H., 11, Marner, F. J., and Moore, R. E. (1979). Science 204, 193-195. Carlson, R. D., Morey-Gaines, G., Tindall, D. R., and Dickey, R. W. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 271-288. American Chemical Society, Washington, D.C. Carlson, R. E., Lever, M. L., Lee, B. W., and Guire, P. E. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 181-192. American Chemical Society, Washington, D.C. Carmichael, W. W. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 1-13. Plenum, New York. Carmichael, W. W. (1982). S. Afr. J . Sci. 78, 367-372. Carmichael, W. W., and Bent, P. E. (1981). Appl. Enuiron. Microbiol. 41, 1383-1388. Carmichael, W. W., and Gorham, P. R. (1977). J . Phycol. 13, 97-101. Carmichael, W. W., and Gorham, P. R. (1978). Mitt. I n t . Verein. Limnol. 21, 285-295. Carmichael, W. W., and Mahmood, N. A. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 377-389. American Chemical Society, Washington, D.C. Carmichael, W. W., and Schwartz, L . D. (1984). Farmers Bulletin No. 2275. U.S. Dept. of Agriculture. Carmichael, W. W., Biggs, D. F., and Gorham, P. R. (1975). Science 187, 542-544. Carmichael, W. W., Gorham, P. R., and Biggs, D. F. (1977). Can. Vet. J . 18,71-75. Carmichael, W. W., Biggs, D. F., and Peterson, M. A. (1979). Toxicon 17, 229-236. Carmichael, W. W., Jones, C. L. A . , Mahmood, N . A,, and Theiss, W. C. (1985). In “Critical Reviews in Environmental Control” (C. P. Straub, ed.), pp. 275-313. CRC Press, Boca Raton, Florida. Carroll, C. R., and Hoffman, C. A. (1980). Science 209, 414-416. Carter, N. (1938). Arch. Protistenkd. 90, 1. Center for Disease Control, Foodborne Disease Surveillance (1983). Annual Summary 1981. U.S. Dept. of Health and Human Services, Public Health Service. Chew, F. S . , and Rodman, J. E. (1979). In “Herbivores-Their Interactions with Secondary Plant Metabolites” (G. A. Rosenthal and D. H. Jansen, eds.), pp. 271-307. Academic Press, New York. Chretien, J. H., Fermaglich, J., and Garagusi, V. F. (1981). Arch. Neurol. 38, 783.

ALGAL TOXINS

95

Chungue, E., Bagnis, R., Fusetani, N., and Yasumoto, T. (1977). Toxicon 15, 89-93. Codd, G. A,, and Carmichael, W. W. (1982). FEMS Microbiol. Lett. 13, 409-411. Cohen, S. A., and Barchi, R. L. (1981). Biochim. Biophy.5. Acta 645, 253-261. Cooper, M. J. (1964). Pac. Sci. 18, 411-440. Dale, B., Yentsch, C. M., and Hurst, J. W. (1978). Science 20, 1223-1225. Davio, S. R., and Pickering, R. P. (1983). Toxicologist 4, 13 (Abstract No. 52). Dembert, M. L., and Pearn, J. H. (1982). A m . J . Public Health 72, 1298. Devlin, J. P., Edwards, 0. E., Gorham, P. R . , Hunter, N. R., Pike, R. K., and Stavric, B. (1977). Can. J . Chem. 55, 1367-1371. Dickey, R. W., Miller, D. M., and Tindall, D. R. (1984). In “Seafood Toxins’’ (E. P. Ragelis, ed.), pp. 257-269. American Chemical Society, Washington, D. C. Dufva, E., Loison, G . , and Holmstedt, B. (1976). Toxicon 14, 55-64. Dugdale, R. C. (1979). I n “Toxic Dinoflagellate Blooms” (D. L. Taylor and H. H . Seliger, eds.), pp. 257-262. Elsevier, Amsterdam. DuPuy, J. L. (1968). PhD thesis, University of Washington, Seattle. Elleman, T. C . , Falconer, I. R., Jackson, A. R. B., and Runnegar, M. T. (1978). A u s f . J Biol. Sci. 31, 209-218. Eloff, J. N., Siegelman. H. W., and Kycia, H. (1982). S. A,/). J . Sci. 78, 377 (Abstract). Evans, M. H. (1964). Br. J . Pharmacol. Chemother. 22, 478-485. Evans, M. H. (1965). Br. J . Exp. Pathol. 46, 245-253. Evans, M. H . (1970). Br. J . Pharmacol. 40, 847-865. Evans, M. H. (1972). I n t . Rev. Neurobiol. 15, 83-166. Falconer, I. R., Jackson, A. R. B., Langley, J., and Runnegar, M. T. (1981). A u ~ t J. . Bio. Sci. 34, 179-187. Falconer, I. R., Beresford, A. M., and Runnegar, M . T . C. (1983a). Med. J . Aust. 1, 511514. Falconer, I. R., Runnegar, M. T. C., and Huynh, V . L. (1983b). Tenth Fed. Convention Aust. Water Wastewater Assoc., loth, Sydney, April 26-1. Farber, J. L. (1981). Life Sci. 29, 1289. Fitzgerald, G. P., Gerloff, G. C . , and Skoog, F. (1952). Sewage Ind. Wastes pp. 888-896. Foxall, T. L., and Sasner, J. J., Jr. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 365-387. Plenum, New York. Fraenkel, G. S . (1959). Science 129, 1466-1470. Fraenkel, G. S . (1969). Enfomol. Exp. Appl. 12, 473-486. Fujiki, H., Mori, M.,Nakayasu, M.. Terada. M., Sugimura. T., and Moore, R. E. (1981). Proc. Nail. Acad. Sci. U . S . A . 78, 3872-3876. Fujiki, H., Suganuma, M., Matsukura, N., Sugimura, T . , andTakayama, S . (1982a). Carcinogenesis 3, 895-898. Fujiki, H., Suganuma, M., Nakayasu, M., Hoshino, H., and Moore, R. E. (1982b). Gann 73, 495-497. Fujiki, H., Suganuma, M., Sugimura, T . , and Moore, R. E. (1983). In “Human Carcinogenesis” (C. C. Harris, ed.), pp. 303-324. Academic Press, New York. Fujiki, H., Suganuma, M., Hakii, H., Bartolini, G . , Moore, R. E., Takayama, S., and Sugimura, T . (1984). J . Cancer Res. Oncol. 108, 174-176. Genenah, A. A., and Shimizu, Y. (1981). J . Agric. Food Chem. 29, 1289-1291. Gentile, J. H. (1971). In “Microbial Toxins: Algal and Fungal Toxins’’ (S. Kadis, A. Ciegler, and S . J. Ajl, eds.), Vol. VII, pp. 27-66. Academic Press, New York. Gentile, J. H., and Maloney, T . E. (1969). Can. J . Microbiol. 15, 165-173. Ghazarossian, V. E., Schantz, E. J., Schnoes, H. K . , and Strong, F. M. (1974). Biochem. Biophys. Res. Commun. 59, 1219-1225. Gilfillan, E. S . , and Hansen, S . A. (1975). I n “Toxic Dinoflagellate Blooms, 1st International

96

W A Y N E W. CARMICHAEL

Conference” (V. R. LoCicero, ed.), pp. 367-375. Mass. Sci. and Techno]. Found., Wakefield. Gorham, P. R. (1964). In “Algae and Man” (D. F. Jackson, ed.), pp. 307-336. Plenum, New York. Gorham, P. R., McLachlan, J., Hammer, U. T., and Kim, W. K. (1964). Verh. In?. Verein. Limnol. XV, 796-804. Grabow, W. 0. K., DuRandt, W. C., Prozesky, 0. W., and Scott, W. E. (1982). Appl. Environ. Microbiol. 43, 1425-1433. Grigor’yeva, L. V., Kirpenko, Yu. A., Orlovskiy, V. M., and Stankevich, V. V. (1977). Gidrobiol. Zh. 13, 57-62 (Ukranian). Guillard, R. R. L., and Ryther, J. H. (1962). Can. J . Microbiol. 8, 229-239. Hall, S . (1982). PhD thesis, Universit. of Alaska, Fairbanks. Hall, S., and Reichardt, P. B. (1984). In “SeafoodToxins” (E. P. Ragelis, ed.), pp. 113-123. American Chemical Society, Washington, D.C. Hall, S., Reichardt, P. B., and Neve, R. A. (1980). Biochem. Biophys. Res. Commun. 97, 649-653. Halstead, B. W. (1967). “Poisonous and Venemous Marine Animals of the World,” Vol. 11. U.S. Govt. Printing Office, Washington, D.C. Hanno, H. A. (1981). J . Am. Med. Assoc. 245, 464. Harada, T., Oshima, Y., and Kamiya, H. (1982). Bull. Jpn. Soc. Sci. Fish. 48, 821-825. Hashimoto, Y., Kamiya, H., Yamazato, K., and Nozawa, K. (1976). I n “Animal, Plant and Microbial Toxins” (A. Ohsaka, K. Hayashi, and Y. Sawai, eds.), Vol. I , pp. 333-338. Plenum, New York. Hauman, J. H. (1981). I n “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 97-102. Plenum, New York. Hoffman, P. A., Granade, H. R., and McMillan, J. P. (1983). Toxicon 21, 363-370. Hokama, Y., Banner, A. H., and Boylan, D. B. (1977). Toxicon 15, 317-325. Hokama, Y., Abad, M. A., and Kimura, L. H. (1983). Toxicon 21, 817-824. Hokama, Y., Kimura, L. H., Abad, M. A., Yokochi, L., Scheuer, P. J . , Nukina, M., Yasumoto, T., Baden, D. G., and Shimizu, Y. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 307-320. American Chemical Society, Washington, D.C. Holligan, P. M. (1979). In “Toxic Dinoflagellate Blooms” (D. L. Taylor and H. H . Seliger, eds.), pp. 249-256. Elsevier, Amsterdam. Horwitz, W. (1980). “Official Methods of Analyses of the Association of Official Analytical Chemists,” p. 298. Washington, D.C. Hoshino, H., Miwa, M., Fujiki, H., and Sugimura, T. (1980). Biochem. Biophys. Res. Commun. 95, 842. Homer, L. S., ed. (1965). “National Shellfish Sanitation Program Manual of OperationsPart 1” U.S. Dept. of Health, Education and Welfare, Public Health Service Publication No. 33. Hsu, C.-P., Marchand, A., Shimizu, Y., and Sims, G. G. (1979). J. Fish. Res. Board Can. 36, 32-36. Huber, C. S. (1972). Acru Ctystollogr. B28, 2577-2582. Hughes, E. O . , Gorham, P. R., and Zehnder, A. (1958). Can. J. Microbiol. 4, 225-236. Hughes, J. M., and Merson, M. H. (1976). New Engl. J . Med. 295, 1117-1120. Hurst, J. W., and Gilfillan, E . S. (1977). In “Proceedings of the Tenth National Shellfish Sanitation Workshop” (D. S. Wilt, ed.), pp. 152-161. United States Dept. of Health, Education and Welfare, Shellfish Sanitation Branch, Washington, D.C. Hurst, J. W., and Yentsch, C. M. (1981). Can. J . Fish. Aquut. Sci. 38, 152-156. Ikawa, M., Wegener, K., Foxall, T. L., and Sasner, J. J., Jr. (1982). Toxicon 20, 747-752.

ALGAL TOXINS

91

Jackim, E., and Gentile, J. H. (1968). Science 162,915-916. Jackson, A. R. B., McInnes, A., Falconer, I. R., and Runnegar, M. T. C. (1984). Vet. Pathol. 21, 102-113. Janzen, D. H. (1974). Biotropica 6,69-103. Jones, C. L. A. (1984). MSc thesis, Wright State University, Dayton, Ohio. Kao, C. Y. (1975). In “Toxic Dinoflagellate Blooms, 1st International Conference” (V. R. LoCicero, ed.), pp. 347-353. Mass. Sci. Technol. Found., Wakefield. Kao, C. Y., and Nishiyama, A. (1965). J . Physiol. (London) 180, 50-66. Kao, C. Y., and Walker, S. E. (1982). J . Physiol. (London) 323, 619-637. Kao, C. Y., James-Kracke, M. R., Kao, P. N., Koghn, F., and Schnoes, H. K. (1981). Fed. Proc., Fed. A m . SOC.Exp. Biol. 40, 240. Kashiwagi, M., Mynderse, J. S.,Moore, R. E., and Norton, T. R. (1980). J . Pharm. Sci. 69, 735-738. Kato, Y., and Scheuer, P. J. (1975). Pure Appl. Chem. 41, 1-14. Kato, Y., and Scheuer, P. J. (1976). Pure Appl. Chem. 48, 29-33. Kim, V. S., and Padilla, G. M. (1977). Life Sci. 21, 1287-1292. Kimura, L. H., Abad, M. A., and Hokama, Y. (1982a). J . Fish Biol. 21, 671-680. Kimura, L. H., Hokama, Y., Abad, M. A., Oyama, M., and Miyahara, J. T. (1982b). Toxicon 20,907-912. Kirpenko, Yu. A., and Kirpenko, N. I. (1980). Gidrobiol. Zh. 16, 53-57. (Ukranian) Kirpenko, Yu. A., Lukina, L. F., Orlovskiy, V. M., and Peskov, V. A. (1976). Byull. Zzobreteniia 16, 125-126 (Ukranian). Kirpenko, Yu. A., Sirenko, L. A., and Kirpenko, N. I. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 257-269. Plenum, New York. Kirpenko, Yu. A., Stankevich, V. V., and Kirpenko, N. I. (1982). Gidrobiol. Zh. 18, 34-36 (Ukranian). Koehn, F. E., Hall, S., Wichmann, C. F., Schnoes, H. K., and Reichardt, P. B. (1982). Tetrahedron Lett. 23, 2247-2248. Kosaki, T., and Anderson, H. (1968). Toxicon 6, 55-58. Kozakai, H., Oshima, Y., and Yasumoto, T. (1982). Agric. Biol. Chem. 46, 233-236. Lambert, W. (1981). Verh. In?. Verein. Limnol. 21, 1436-1440. Lawrence, D. N., Enriquez, M. B., Lumish, R. M., and Maceo, A. (1980). J. A m . Med. ASSOC. 244, 254-258. Legrand, A. M.,Galonnier, M., and Bagnis, R. (1982). Toxicon 20, 311-315. Levin, D. A. (1976). Annu. Rev. Ecol. Syst. 7 , 121-159. Li, K. (1965). Science 147, 1580-1581. Lightner, D. V. (1978). J . Invert. Pathol. 32, 139. Louw, P. G. J. (1950). S.Afr. Ind. Chem. 4, 62-66. McCollum, J. P. K., Pearson, R. C. M., Ingham, H. R., Wood, P. C., and Dewar, H. A. (1968). Lancer 2, 767. McKey, D. (1979). In “Herbivores-Their Interactions with Secondary Plant Metabolites” (G. A. Rosenthal and D. H. Jansen, eds.), pp. 56-133. Academic Press, New York. McLachlan, J., and Gorham, P. R. (1961). Can. J. Microbiol. 7, 869-882. Meyer, K. F., Sornmer, H., and Schoenholz, P. (1928). J. Prevent. Med. 2, 365-394. Miller, D. M., Dickey, R. W., and Tindall, D. R. (1984). In “Seafood Toxins” (E. P Ragelis, ed.), pp. 241-255. American Chemical Society, Washington, D. C. Miyahara, J. T., Akau, C. K., and Yasumoto, T. (1979). Res. Commun. Chem. Pathol. Pharmacol. 25, 177-180. Moikehai, S.N., and Chu, G. W. (1971). J. Phycol. 7,8-13. Moikehai, S. N., Chu, G. W., and Berger, L. R. (1971). J . Phycol. 7, 4-8.

98

WAYNE W. CARMICHAEL

Moore, R. E. (1981a). In “Marine Natural Products: Chemical and Biological Perspectives” (P. J. Scheuer, ed.), Vol. IV, pp. 1-52. Academic Press, New York. Moore, R. E. (1981b). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 15-23. Plenum, New York. Moore, R. E. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 369-376. American Chemical Society, Washington, D.C. Moore, R. E., Blackman, A. J., Cheuk, C. E., Mynderse, J. S., Matsumoto, G. K., Clardy, J., Woodard, R. W., and Craig, J. C. (1984). J. Org. Chem. 49, 2484-2489. Morey-Gaines, G. (1982). Phycologia 21, 154-163. Morns, J. G., Jr., Lewin, P., Smith, C. W., Blake, P. A., and Schneider, R. (1982). A m . J. Trop. Med. Hyg. 31, 574-578. Muller, H. (1939). J. Pharmacol. Exp. Ther. 53, 67-89. Murakami, Y., Oshima, Y., and Yasumoto, T. (1982). Bull. Jpn. SOC.Sci. Fish. 48, 69-72. Murthy, J . R . , and Capindale, J. B. (1970). Con. J . Biocbem. 48, 508-510. Mynderse, J. S., and Moore, R. E. (1978). J . Org. Chem. 43, 2301-2303. Mynderse, J. S., Moore, R. E., Kashiwagi, M., and Norton, T. R. (1977). Science 196,538540. Nakajima, I., Oshima, Y., and Yasumoto, T. (1981). B14ll. Jpn. Soc. Sci. Fish. 47, 10291033. Nakayasu, M., Fujiki, H., Mori, M., Sugimura, T., and Moore, R. E. (1981). Cancer Lett. U ,271-277. Nukina, M., Tachibana, K., Koyanagi, L. M., and Scheuer, P. J. (1983). A m . Chem. Soc. Div. Agric. Food Chem. Natl. Meet., 186th, Washington, D.C. Abstract No. 135. Ohshika, H. (1971). Toxicon 9, 337-343. Oshima, Y., Kotaki, Y., Harada, T., and Yasumoto, T. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 161-170. American Chemical Society, Washington, D.C. Ostensvik, O., Skulberg, 0. M., and Soh, N. E. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 315-324. Plenum, New York. Pearn, J., Harvey, P., DeAmbrosis, W., Lewis, R., and McKay, R. (1982). Med. J . Aust. 1, 57-58. Phinney, H. K., and Peek, C. A. (1961). In “Algae and Metropolitan Wastes,” pp. 22-27. U.S. Dept. of Health, Education and Welfare, Cincinnati, Ohio. Porter, K. G., and Orcutt, J. D. (1980). In “The Evolution and Ecology of Zooplankton Communities” (W. C. Kerfoot, ed.), pp. 268-281. Univ. Press of New England, Hanover, New Hampshire. Price, R. J., and Lee, J. S. (1971). J . Fish. Res. Board. Can. 28, 1789-1792. Proctor, N. H., Chan, L. S., and Trevor, A. J. (1975). Toxicon 13, 1-9. Provasoli, L. (1968). In “Culture and Collection of Algae” (H. Watanabe and A. Hattori, eds.), pp. 63-75. Jpn. SOC.Plant Physiol., Hakone. Rabin, R., and Darbre, A. (1975). Biochem. SOC. Trans. 3, 428-430. Ragelis, E. P. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 25-36. American Chemical Society, Washington, D. C. Randall, J. (1958). J. Bull. Mar. Sci. GulfCaribb. 8, 236-267. Ransom, R. E., Nerad, T. A., and Meier, P. G. (1978). J . Phycol. 14, 114-116. Rayner, M. D. (1968). Science 160, 70-71. Rayner, M. D. (1972). Fed. Proc., Fed. A m . SOC.Exp. B i d . 31, 1139-1145. Rayner, M. D., and Szekerczes, J. (1973). Toxico/. Appl. Pharmacol. 24, 489-496. Raziuddin, S., Siegelman, H. W., and Tornabene, T. G. (1983). Eur. J . Biochem. 137, 333336.

ALGAL TOXINS

99

Rhoades, D. F. (1979). I n “Herbivores-Their Interactions with Secondary Plant Metabolites” (G. A. Rosenthal and D. H. Jansen, eds.). pp. 4-54. Academic Press, New York. Rhoades, D. F., and Cates, R. G. (1976). Recent Adv. Phytochem. 10, 168-213. Rogers, R. S., and Rapoport, H. (1980). J. Am. Chem. Soc. 102, 7335-7339. Runnegar, M. T. C., and Falconer, I. R. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 325-342. Plenum, New York. Runnegar, M. T . C., and Falconer, 1. R. (1982). S . Afr. J . Sci. 78, 363-366. Runnegar, M. T., Falconer, I. R., and Silver, J. (1981). Arch. Pharmacol. 317, 268-272. Sakabe, N., Harada, H., Hirata, Y.,Tomiie, Y., and Nitta, I. (1966). Tetrahedron Lett. 23, 2523-2525. Sakamoto, H., Terada, M., Fujiki, H., Mori, M., Nakayasu, M., Sugimura, T., and Weinstein, I. B. (1981). Biochem. Biophys. Res. Commun. 102, 100-107. Santikarn, S., Williams, D. H., Smith, R. J., Hammond, S. J., Botes, D. P., Tuinman, A., Wessels, P. L., Viljoen, C. C., and Kruger, H. (1983). J. Chem. Soc. Chem. Commun. 12, 652-654. Sasner, J. J., Jr., Ikawa, M., Foxall, T. L., and Watson, W. H. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 389-402. Plenum, New York. Sasner, J. J., Jr., Ikawa, M., and Foxall, T. L. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 391-406. American Chemical Society, Washington, D.C. Sawyer, P. J., Gentile, J. H., and Sasner, J. J., Jr. (1968). Can. J . Microbiol. 14, 1199-1204. Scagel, R. F., Bandoni, R. J., Maze, J . R., Rouse, G. E., Schofield, W. B., and Stein, J. R. (1982). “Nonvascular Plants-An Evolutionary Survey.” Wadsworth, Belmont, California. Schantz, E. J. (1960). Annals N . Y . Acad. Sci. 90, 843-855. Schantz, E. J. (1971). I n “Neuropoisons: Their Pathophysiological Actions” (L. L. Simpson, ed.) Vol. I , pp. 159-168. Plenum, New York. Schantz, E. J., Mold, J. D., Stranger, D. W., Shavel, J., Riel, P., Bowden, J. P., Lynch, J. M., Wyler, R. S., Riegel, B., and Sommer, H. (1957). J. Am. Chem. Soc. 79, 52305235. Schantz, E. J., Lynch, J. M., Vayvada, G., Matsumoto, K., and Rapoport, H. (1966). Biochemistry 5 , 1191-1 195. Schantz, E. J., Ghazarossian, V. E., Schnoes, H. K., Strong, F. M., Springer, J. P., Pezzanite, J. O., and Clardy, J. (1975). J. Am. Chem. Soc. 97, 1238-1239. Scheuer, P. J., Takahashi, W., Tsutsumi, J . , and Yoshida, T. (1967). Science 155, 12671268. Schmidt, R. J., and Loeblich, A. R., 111. (1979). I n “Toxic Dinoflagellate Blooms” (D. L. Taylor and H. H. Seliger, eds.), pp. 83-88. Elsevier, Amsterdam. Schuett, W., and Rapoport, H. (1962). J. A m . Chem. Soc. 84, 2266-2267. Schultz, J. C., and Baldwin, I. T. (1982). Science 217, 149-151. Seliger, H. H., Tyler, M. A,, and McKinley, K. R. (1979). I n “Toxic Dinoflagellate Blooms” (D. L. Taylor and H. H. Seliger, eds.), pp. 239-248. Elsevier, Amsterdam. Setliff, J. A., Rayner, M. D., and Hong, S. K. (1971). Toxicol. Appl. Pharmacol. 18, 676684. Shilo, M. (1967). Bacteriol. Rev. 31, 180-193. Shilo, M. (1971). I n “Microbial Toxins; Algal and Fungal Toxins” (S. Kadis, A. Ciegler, and S. J. Ajl, eds.), Vol. VII, pp. 67-103. Academic Press, New York. Shilo, M. (1981). In “The Water Environment: Algal Toxins and Health” (W. W. Carmichael, ed.), pp. 37-48. Plenum, New York.

100

WAYNE W . CARMICHAEL

Shimizu, Y., and Yoshioka, M. (1981). Science 212, 546-547. Shimizu, Y . , Alam, M., Oshima, Y., and Fallon, W. E. (1975). Biochem. Biophys. Res. Commun. 66,731-737. Shimizu, Y., Hsu, C.-P., Fallon, W. E., Oshima, Y.,Miura, I., and Nakanishi, K. (1978). J. Am. Chem. SOC.100, 6791-6793. Shimizu, Y., Kobayashi, M., Genenah, A., and Ichihara, N. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 151-160. American Chemical Society, Washington, D.C. Shizumi, Y., Shibata, S., and Tachibana, K. (1981). J. Pharrnacol. Exp. Ther. 217,475. Siegelman, H. W., Adams, W. H., Stoner, R. H., and Slatkin, D. N. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 407-413. American Chemical Society, Washington, D.C. Skulberg, 0. M. (1983). “Culture Collection of Algae at Norwegian Institute for Water Research.” Oslo. Skulberg, 0. M., Codd, G. A., and Carmichael, W. W. (1984). Ambio 13, 244-247. Slatkin, D. N . , Stoner, R. D., Adams, W. H., Kycia, J. H., and Siegelman, H. W. (1983). Science 220, 1383-1385. Smith, M. T., Thor, H., and Orrenius, S. (1981). Science 213, 1257. Snell, T. W. (1980). Oecologia (Berlin) 46, 343-346. Solomon, A. E., and Stroughton, R. B. (1978). Arch. Dermatol. 114, 1333-1335. Spaulding, B. C. (1980). J. Physiol. (London) 305, 485-500. Spivak, C. E., Witkop, B., and Albuquerque, E. X. (1980). Mol. Pharrnacol. 18, 384-394. Stahl, E. (1888). Jena. Z. Med. Natunviss. 22, 557-684. Stanier, R. Y., Kunisawa, R., Mandel, M., and Cohen-Bazire, (1971). Bacteriol. Rev. 35, 171-205. Steidinger, K. (1979). I n “Toxic Dinoflagellate Blooms” (D. L. Taylor and H. H. Seliger, eds.), pp. 435-442. Elsevier, Amsterdam. Sullivan, J. J., and Iwaoka, W. T. (1983). J . Assoc. Off. Anal. Chem. 66, 297-303. Sullivan, J. J., and Wekell, M. M. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 197205. American Chemical Society, Washington, D.C. Sullivan, J. J . , Iwaoka, W. T., and Liston, J. (1983). Biochem. Biophys. Res. Commun. 114, 465-472. Switzer-Dunlap, M., and Hadfield, M. (1977). J . Exp. Mar. Biol. Ecol. 29, 245. Tachibana, K. (1980). PhD thesis, University of Hawaii, Honolulu. Tachibana, K., Scheuer, P. J., Tsukitani, Y., Kikuchi, H., Van Engen, D., Clardy, J., Gopichand, Y., and Schmitz, F. J. (1981). J . Am. Chem. SOC. 103, 2469-2471. Takashima, M., and Sakai, H. (1960). Bull. Agric. Chem. SOC.Jpn. 24, 652-655. Takahashi, M., Ohizumi, Y., and Yasumoto, T. (1982). J . Biol. Chem. 257, 7287-7289. Taylor, F. J. R. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 77-97. American Chemical Society, Washington, D.C. Theiss, W. C. (1984). PhD thesis, Wright State University, Dayton, Ohio. Tindall, D. R., Dickey, R. W., Carlson, R. D., and Morey-Gaines, G. (1984). I n “Seafood Toxins” (E. P. Ragelis, ed.), pp. 225-240. American Chemical Society, Washington, D.C. Toerien, D. F., Scott, W. E., and Pitout, M. J. (1976). Water S . Afr. (Pretoria) 2, 160-162. Turpin, D. H., Dabell, P. E. R., and Taylor, F. J. R. (1978). J. Phycol14, 235-238. Twarog, B. M., and Yamaguchi, H. (1975). I n “Toxic Dinoflagellate Blooms, 1st International Conference” (V. R. LoCicero, ed.), pp. 381-393. Mass. Sci. Technol. Found., Wakefield. Ulitzur, S. (1969). Verh. Inr. Verein. Limnol. 17, 771-777. Ulitzur, S., and Shilo, M. (1970). Biochim. Biophys. Acta 201, 350-363.

ALGAL TOXINS

101

Volk, S. L., and Phinney, H . K. (1968). Cun. J. Bot. 46, 619-630. Watanabe, M. F . , and Oishi, S. (1980). J p n . J . Limnol. 41, 5-9. Watanabe, M. F., and Oishi, S. (1982). Appl. Enuiron. Microbiol. 43, 819-822. Watanabe, M. F., Oishi, S., and Makao, T . (1981). Verh. I n t . Verein. Limnol. 21, 14411443. White, A. W., and Maranda, L. (1978). J. Fish. Res. Board Can. 35, 397-402. Wichmann, C. R., Boyer, G., Divan, C. L., Schantz, E. J., and Schnoes, H. K. (1981). Tetrahedron Lett. 22, 1941-1944. Williams, D. H. (1983). In?. J. Muss Spectrosc. Zon Phys. 53, 37-44. Wong, S. H., and Hindin, E. (1982). A m . Water Works Assoc. J . 74, 528-529. Yasumoto, T., Hashimoto, Y., Bagnis, R., Randall, J. E., and Banner, A. H. (1971). Bull. Jpn. SOC. Sci Fish. 37, 724. Yasumoto, T., Nakajima, I., Chungue, E., and Bagnis, R . (1977).Bull. Jpn. Soc. Sci. Fish. 43, 69-74. Yasumoto, T., Oshima, Y., Murakami, Y., Nakajima, I., Bagnis, R.and Rukuyo, Y. (1980). Bull. Jpn. Soc. Sci. Fish. 46, 327-331. Yasumoto, T., Oshima, Y., Tajiri, M., and Kotaki, Y. (1983). Bull. Jpn. Soc. Sci. Fish. 49, 633-636. Yasumoto, T., Marata, M., Oshima, Y., Matsumoto, G. K . , and Clardy, J. (1984). In “Seafood Toxins” (E. P. Ragelis, ed.), pp. 207-216. American Chemical Society, Washington, D.C. Yentsch, C. M., and Incze, L. (1980). In “Mussel Culture in North America” (R. Lutz, ed.), pp. 223-246. Elservier, Amsterdam. Yentsch, C. M., and Mague, F. C. (1980). Int. J. Chronobiol. 7, 77-84. Yentsch, C. M., Dale, B., and Hurst, J. W. (1978). J . Phycol. 14, 330-332.

This Page Intentionally Left Blank

Plant Transposable Elements

PATRICIA NEVERS, NANCY S. SHEPHERD* AND HEINZ SAEDLER Mux-Plrnc.k-In.stitiltjlir Ziic,htiln~.sforsc.hunC:Egelspfud Kiiln. Federril RC.pilhlic of Gcrtnany

I. The Phenomenon of Variegation.. ......................................... A. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Examples of Variegation Excluded from Detailed Consideration . . 11. Recognizing Mutations due to Transposable Elements by ... Classical Genetic Means.. .... A. Reports of Mutable Alleles ... B. Segregation Ratios and Wh ................ C. Reversion Events . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Reverse Variegation . . . . . . . . . . . . . . ............. E. Variegation Associated with Chromosomal Anomalies .................... F. Stable and Unstable Allelic Variants of Mutable Alleles G. Transposition: The Most Characteristic Pr 111. Transposable Elements at a Molecular Level. A. Cloning Strategy. ...................... B. Structural Features of Plant Transposable Elements ...................... C. Multiple Copies of Plant Transposable Elem D. Detection of Transposition at the Molecular E. How a Transposable Element Influences Ge F. Proteins Encoded by Transposable Elements . . . . . . G . Reversion Events at the Molecular Level.. . H. The Molecular Basis for Allelic Derivatives IV. A Model of the Mechanism of Transposition.. . A. Observations on Which the Model is Based. B. Excision of a Transposable Element in Plants C. Integration of a Plant Transposable Element. . . . . . . D. Predictions.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

104 104 107 109 109 109 137 139 141

153

172

* Present address: E.I. Dupont de Nemours & Company, Inc., Central Research & Development Department, Wilmington, Delaware 19898, U.S. A. 103 ADVANCES IN BOTANICAL RESEARCH, VOL. 12

Copyright 0 1986 by Academic Press Inc. (London) Ltd. All rights of reproduction in any form reserved.

104

PATRICIA NEVERS ET A L .

E. Large-Scale Rearrangements Induced by Transposable Elements in Plants F. Specificity of Integration .............................................. V. Interaction between Transposable Elements A. General Remarks ..................... B. Dosage Effects ..... C. Interaction with Alte D. Other Modifiers .... ................................ E. Possible Interaction between Two Transposable Elements of Antirrhinurn

..

176 177

183

majus ............................................................... 185 VI. Induction of Genomic Instabilities .................. . . . . . . . . . . . . . 188 V11. Addendum ..................................... ........ 193 References .............................................................. 194

I. THE PHENOMENON OF VARIEGATION A. INTRODUCTION

Where uniformity is the rule, irregularities like a flaw in a woven pattern or a spot on a plain-colored surface immediately strike the eye. Thus among living organisms the phenomenon of variegation, the unusually spotted, striped, or otherwise mosaic appearance of some individuals as opposed to their normally uniform counterparts, has long been a source of fascination. In plants variegation is most easily recognized as irregularities in pigment patterns on leaves, flowers, and seeds, although other characteristics such as leaf form, flower form, or starch content (in maize kernels) may also be subject to variegation. As early as 1588 Jacob Theodor von Bergzabern described with great enthusiasm the multicolored and variegated kernels of maize from the New World. And variegation in tulips, illustrated in the still-life paintings of many Dutch masters, became almost a craze in the seventeenth century in the Netherlands. Indeed, variegated tulips were so coveted that a single bulb was worth as much as several pigs, sheep, tons of grain, or even 1000 lb of cheese (Dubos, 1958, cited in Pollard, 1959). Although present scientific interest in variegation is certainly more rational, it is no less animated. Researchers hope that variegation may contribute to our understanding of an organism’s capacity for change and provide a starting point for unraveling the complex mechanisms underlying its development. As discussed below, variegation can be caused by several different factors. The focal point of this article will be variegation in plants due to genetic alterations implemented by transposable elements. The distinguishing feature of this kind of variegation, as opposed to other examples

PLANT TRANSPOSABLE ELEMENTS

105

discussed below, is that the two or more different kinds of tissue in the variegated plant differ genotypically, that is, with respect to the DNA of their nuclear genomes. In the simplest case one kind of tissue contains a transposable element at a given locus and the other does not. The terminology used in connection with transposable elements reflects historical developments in this field of research. Prior to the discovery of transposable elements as discrete physical entities, their existence was predicted on the basis of classical genetic evidence. In this period a premolecular terminology arose, some terms of which are still relevant today while others may require redefinition. The term “mutable allele” was used to refer to a special mutational condition of a locus characterized by frequent reversion to the wild type or to a new, intermediate condition in somatic and/or germinal cells. Likewise the term “mutability” most commonly was used to designate the capacity of an allele to revert to the wildtype condition. Since the phenotypes of plants with mutable alleles are usually variegated, these alleles were given such names as variegata, mutabilis, marmorata, maculata, variabilis, and recurrens. McClintock (1950a) proposed that discrete, transposable genetic elements are responsible for some cases of variegation in maize. She called them “controlling elements” (McClintock 1956a,b) since they seemed capable of influencing the expression of a given locus when integrated in or next to it. Later on, when elements of the same nature were discovered in bacteria, they were classified in two categories according to their size and complexity. The term “insertion sequence” (insertion element, IS element) was assigned to short elements up to about 2000 base pairs (bp) in length while larger, more complex elements containing additional genes unrelated to insertion function were termed “transposons” (Campbell et al., 1977). We have chosen to use the term “transposable element” rather than “controlling element” to avoid confusion with the terminology of other fields of genetic research. Briefly we shall define a transposable element as a discrete segment of DNA that is capable of changing its position in the genome. However, it must be pointed out that the criteria for deciding whether a particular segment of DNA is transposable vary depending upon the experimental methods used. As discussed in Section 111, data from molecular research have provided new criteria beyond those obtained by classical genetic methods. Thus DNA sequence analyses have shown that transposable elements may share certain structural features that might be useful for classification. Furthermore, transposition can now be demonstrated by DNA hybridization techniques that were unknown 20 years ago (see Section 111,D). In general, a transposable element is characterized by its ability to

106

PATRICIA NEVERS E T A L .

integrate into the DNA at some position in the genome and thereby alter the expression of the gene(s) in and around itself. In some instances integration of the element is manifested phenotypically as a mutation. If, for instance, a gene involved in anthocyanin synthesis is affected, integration of an element may result in flowers that lack red/purple anthocyanin pigmentation and are basically yellow or colorless. However, mutations induced by transposable elements are often unstable. In some cells of the mutant plant, for example, the element may be excised from its former position at a given locus. In these cells expression of the locus is then restored. Thus in the case of the anthocyanin gene mentioned above, groups of cells with normal pigmentation appear against a colorless background, resulting in a typical variegated pattern. But although frequent excision events may be the basis for the “mutability” of some alleles, transposable elements can generate genetic instability in other ways. Rearrangements of DNA sequences including deletions, duplications, inversions, and translocations may occur within the element itself or in its vicinity and thus elicit alterations in gene expression and phenotype (McClintock, 1951a, 1956b, 1965a). In some cases chromosome breaks or alterations in heterochromatin structure may also occur adjacent to transposable elements and lead to phenotypic changes (McClintock, 1946, 1947). Therefore we shall extend the term “mutability” to include all genetic alterations induced by transposable elements. Before delving into the intricacies of variegation and transposable elements it is worthwhile briefly considering why this subject has gained such attention in recent years. First of all there is a growing feeling that transposable elements may be important in evolution. The innumerable different levels of expression and control that a transposable element can evoke at a locus reflect an enormous potential of these elements for inducing variability. The rearrangements that occur in connection with transposable elements may be instrumental in restructuring the genome in the course of evolution. Besides evolutionary significance, a possible role in development has also been postulated for transposable elements. Here it is the sensitivity of some elements to changes in their physical and physiological environment, manifested as altered variegation patterns (McClintock, 1965a, 1971; Peterson, 1965b, 1966; Fowler and Peterson, 1978), that suggest that these elements may normally be involved in developmental control. Finally some investigators foresee eventually being able to exploit transposable elements for genetic engineering purposes, particularly in monocotyledons. Two very thorough reviews of maize transposable elements have already been published (Fincham and Sastry, 1974; Fedoroff, 1983). In this

PLANT TRANSPOSABLE ELEMENTS

107

article we shall present a more basic approach to plant transposable elements and include information on mutable alleles of other plants. In the sections that follow we shall outline the classical and molecular genetic evidence available on transposable elements in plants and try to relate the two. We shall also advance speculations concerning mechanisms underlying the activities of transposable elements in hopes of stimulating a general discussion.

B . EXAMPLES OF VARIEGATION EXCLUDED FROM DETAILED CONSIDERATION

Some of the most common cases of variegation involve no heritable changes in the different tissues of the organism but are due to the direct effects of an external factor operating at random on the organism. The most common example of this kind is variegation caused by virus infection, which often leads to yellow or white blotches on leaves infected by the virus. In these cases the external agent directly destroys tissue in certain parts of the plant and the variegation pattern persists only as long as the virus is active in the organism. They must be clearly distinguished from cases in which viral infection induces mutations secondarily, sometimes even unstable mutations, that persist after the viral agent has disappeared, as discussed in Section VI. Another example of nonhereditary variegation is that caused by mineral deficiencies. Normal pigmentation can be restored when the deficiency is corrected. More interesting than mosaic patterns due to random effects of an environmental factor are variegation patterns that are heritable. Since plants contain three different genomes, in the chloroplast, mitochondrion, and in the nucleus, mutations can in principle occur in any one and lead to a variegated phenotype. Since we are limiting our article to mutations of the nuclear genome, let us first consider how chloroplast or mitochondria1 mutations can be distinguished from nuclear ones. Kirk and Tilney-Bassett (1978) have compiled and classified many examples of greedwhite variegation patterns that all arise from chloroplast defects in some plant cells. Some of these are the consequence of nuclear mutations while others are due to mutations of the chloroplast genome itself. Plastid mutations can be identified on the basis of the following criteria (Kirk and Tilney-Basset, 1978): (1) Segregation of the trait in crosses is non-Mendelian. (2) Inheritance is maternal, as demonstrated by nonreciprocity in crosses. (3) Cells containing both defective and nonde-

108

PATRICIA NEVERS ET A L .

fective chloroplasts (mixed cells) can be observed. (4) The distribution of colorless cells corresponds to what is expected by random sorting out of chloroplasts during cell division. Essentially the same criteria apply to mitochondrial mutations, although mixed cells and sorting out cannot be observed directly as with chloroplasts. Our decision to exclude cases of variegation due to chloroplast and mitochondrial mutations is a purely economical one, and we are aware that mutable alleles due to transposable elements very likely also occur in the genomes of these organelles. Indeed, some cases have been reported that resemble nuclear activation of an unstable chloroplast allele (Potrykus, 1970; Sears, 1984; other cases are summarized in Kirk and TilneyBassett, 1978). Plasmid-like DNAs with structures resembling transposable elements (see Section 111) have been found in mitochondria of S cytoplasmic male-sterile maize (Pring et al., 1977; Levings and Pring, 1980; Kim et al., 1982; Weissinger et al., 1982; Timothy et al., 1983). Reversion of male sterility in maize has been attributed to instability of mitochondrial DNA, possibly associated with transposition of such plasmids (Levings et al., 1980; Kemble et al., 1983). The most obvious indication that a mutation of the nuclear genome is involved in inducing a variegated phenotype is Mendelian segregation of this trait. However, not all mutations induced by transposable elements exhibit strict Mendelian segregation ratios (see Section II,C), and not every mutation that causes a variegated phenotype and segregates in Mendelian fashion is due to a transposable element. As indicated by Kirk and Tilney-Bassett (1978), there are many cases of greedwhite variegated mutants that exhibit Mendelian segregation ratios but cannot be unequivocally attributed to the presence of a transposable element. In these cases the mutated gene is apparently one required for normal chloroplast development, but its protein product may be labile and become inactivated in response to some physiological or environmental factor. Alternatively the mutation may be a non-transposon-induced one that causes expression of the gene to be inhibited in certain parts of the plant. Two aspects distinguish these cases from most mutants caused by transposable elements. First, the phenotypic switch observed here is from the wild-type state (green) to the mutant condition (white). Second, these mutants differ in breeding behavior compared to those harboring transposable elements (see Section 11,C-F); they breed true and rarely or never throw off genetically stable white or green progeny or other unusual offspring. This suggests that the tissue of the variegated plants is uniform in genotype, as has been verified in a number of cases summarized in Kirk and Tilney-Bassett ( 1978).

PLANT TRANSPOSABLE ELEMENTS

109

11. RECOGNIZING MUTATIONS DUE TO TRANSPOSABLE ELEMENTS BY CLASSICAL GENETIC MEANS A . REPORTS OF MUTABLE ALLELES

Mutations attributable to transposable elements were analyzed as early as the middle of the nineteenth century (Darwin, 1868). In Tables I, 11, and I11 we have compiled detailed lists of such mutations. Many examples may have been overlooked, especially those cited in languages other than French, German, or English or published in journals with limited circulation. We have included only examples of mutable alleles in higher plants, omitting a few cases known in algae, ferns, and fungi (Anderson, 1923; Andersson-Kotto, 1930; Bgrresen and Fjeld, 1977; Clutterbuck, 1970). As mentioned earlier, only mutable alleles of the nucleus are considered. We have included only examples for which some genetic data are available, although there are certainly many more candidates including several under investigation at the John Innes Institute in Norwich (B. J. Harrison and R. Carpenter, personal communication). Since it is tempting to anticipate a transposable element behind many different kinds of genetic anomaly, we have tried to establish a set of criteria for identifying transposable element-induced mutations in the nucleus. These will be elucidated in Section I1,B. B. SEGREGATION RATIOS A N D WHAT CAN BE DEDUCED FROM THEM

In order to demonstrate that a variegated phenotype is due to a mutation of the nuclear genome, segregation of the variegated trait must be analyzed in crosses. In the most common case, integration of a transposable element at a locus causes a mutation that is recessive to wild type and exhibits monoallelic segregation, as described below. A hybrid carrying a dominant wild-type gene involved in anthocyanin synthesis in one homolog and a transposable element inserted at the same locus in the chromosome homolog will have fully colored flowers. The progeny of this hybrid will include wild-type and variegated plants in a ratio of 3 : 1 . When the same mutable allele is combined with a stable recessive allele in a heterozygote, the variegated phenotype of the mutable allele will prevail since it is formally dominant to the stable recessive one. In the following generation, after selfing, the hybrids segregate three variegated plants to each stable recessive one. In backcrosses of the hybrid with a homozygous stable recessive tester strain, 50% of the progeny will be variegated and 50% stable recessive. An example of F2 progeny showing monoallelic segregation of a mutable allele is outlined in Fig. 1 .

TABLE I Examples of Mutable Alleles in Plants“

Plant Aquilegia vulgaris L. (common columbine) Avena sativa L. (common oat) Bullota nigru L. (black horehound) Brassica campestris L. (field cabbage)

Z

Mutable alleleb variegata yellow-striped variegata purple variegated War)

Callistephus chinensis (China aster, Queen Marguerite)

Capsicum annum (red pepper)

variegata

Celosia cristata L. (cock’s comb, woolflower)

yellow mutable (a) apm

Chrysanthemum sinense (chrysanthemum)

mutable magenta

Phenotype Yellow-green leaves with a few large green spots Leaves have yellow and green stripes Yellow-white leaves with many green areas of variable size Foliage, stem, and seed pods contain patches of purple or pinkish-white which may bleach to white White petals among yellow ones; pink petals among salmon-colored ones ; bright blue petals among light blue ones; some flowers also spotted in same colors Green leaves with whitish or white patches of variable size; green “ticks” on white patches also observed Flowers have yellow background with red sectors Flowers have a light pink-dark pink variegation Flowers variegated with white and intense magenta

Germinal reversions

+ + + + +

New alleles (S,I,V)c

Modifier‘

References‘ Baur (1911, 1912) Christie (1922)

(S)V

S

SJ,V

S

+

Kirk and Tilney-Bassett (1978) Orakwue and Crowder (1983)

Hondelmann (1959)

Imai (1936, 1938)

Terasawa (1922)*; Kihara (1979) Kihara (1979)

S

Miyake and Imai (1934-1935)

Dahlia variabilis Desf. (dahlia) Delphinium ajacis L. (rocket larkspur)

rose-alpha lavender-alpha uariegated pink (P*)

mottling (m)

--

Elymus junceus (Russian wild ryegrass)

U

Epilobium paruijorum

marmorata

Fagopyrum sagittatum (buckwheat) Glycine max (soybean)

C"

Hosta minor

dotted (dr)

Impatiens balsamina L (impatiens, garden balsam)

pm

Y ,grn

Stems and ray florets have a mosaic distribution of anthocyanin pigmentation Flower has dark purple spots on a rose background Flower has small purple spots on a lavender background Flowers have blue spots, streaks, and sectors on a pink background Flowers have pink sepals with rose markings which may be distinct streaks and sectors White leaves have green stripes when the independently segregating factor, G , is present White leaves with many green areas of variable size

+

Yellow sectors of variable size on a green background Yellow sectors of variable size on a green background (pale green sectors also observed)

+

Leaves have green spots on a white background Flowers are stable white; if the independent controlling element, M, is present, pale and dark sectors of anthocyanin color appear

+

+

Lawrence (1931); R. Carpenter (personal communication) Demerec (1931)

+

Demerec (1931)

+

Dawson (1955)

-

Dawson (1955)

+

+

Michaelis (1957)*; Kirk and Tilney-Bassett ( 1978) Ah-Khan (1971)

s.v

(+)

+

Lawrence (1974)

s,v

+

Peterson and Weber (1969); Sheridan and Palmer (1977); Kirk and Tilney-Bassett (1978) Vaughn and Wilson (1980) Sastry et a / . (1980)

(continued)

TABLE I (Continued)

Plant Lathyrus odoratus L. (sweetpea) Malva parviJora

Mirabilis jalapa L. (four-o'clock flower, marvel of Peru)

Mutable alleleb

Ci laciniata

chlorina (chf)

L

t4

P

RU

Nigella damascena L. (love in a mist, devil in a bush)

striata L

Phenotype White flower with purple flakes of variable size Leaf form changes from laciniata (narrow leaf) to ''hypernormat' ' Homogeneous yellowish-green leaves unless the unlinked V, regulatory element is present. If V is present, green flecks and sectors of variable size occur. Homogeneous white flower becomes variegated with yellow sectors, stripes, and spots in presence of the unlinked regulator, V (also assuming rr) Homogeneous yellow flowers (if dominant Y allele is present) become variegated with red sectors, stripes, and spots when the unlinked regulator V is present Striped flowers Dark blue flowers with large, light blue sectors; flowers with varying shades of blue from pale to dark blue on one plant

Germinal reversions

New alleles (S,I,V)c

Modifier'

References'

+

Punnett (1922, 1932, 1935) Lilienfeld (1929)*; Kihara (1979)

+

Correns (1909, 1910); Delool and Tilney-Bassett (1984)

+

+

Engels et al. (1975); Spitters et al. (1975)

+

+

Engels et al. (1975); Spitters et al. (1975)

(+I

+

+ +

V

I

Correns (1908, 1909) ToxopCus (1928)

Nicotiana tabacum L. (tobacco)

v(s1)

v(Ds1)

Oryza sativa L. (rice)

white striped

Pelargonium zonale (horse-shoe geranium)

variegata

Petunia hybrida (garden petunia)

anlsii

anl+‘*

“Speckled” flower [v(sl) + homozygous]: scattered, small clusters of anthocyanin pigmentation on a uniformly colorless background “Sectorial” flower [ v ( S l ) / v ( s l ) ] : frequent, clearly defined sectors of speckled tissue on a solidly pigmented background “DsLsS” flower [v(Dsl)lv(sl)]: similar to “sectorial” except that frequent sectors of light speckling are on a dark speckled background White plant with large green areas when the modifier is present

+

V

+

Smith and Sand (1957); Sand (1969, 1976)

+

V

+

Smith and Sand (1957); Sand (1969, 1976)

-

V

+

Smith and Sand (1957); Sand (1969, 1976)

+

S

+

Mitra (1932)*;Mitra and Ganguli (1934)*; Kirk and Tilney-Bassett (1978) Imai (1936)

+

SJ,V

+

White-dark red spots on pale colored flowers

+

S,LV

Red and pink spots on white flowers

t

S,I

Bianchi et a!. (1978); Mulder et al. (1981); Gerats et al. (1982, 1983) Gerats et al. (1984a); A. G . M. Gerats (personal communication) Bianchi et al. (1978); Doodeman et al. (1984a)

Green leaves with white patches; green “ticks” on white patches also observed White flowers with red spots

(+)

+

(continued)

TABLE I (Continued)

Plant Petunia hybrida (cont.)

Mutable alleleb anll"+

White flowers with red spots

+

ad-1

White, intermediate, and red spots on pale-colored flowers Red spots on white flowers, some white sectors Red spots on white flowers, some white sectors White, intermediate, and red spots on white background Red spots on pale background, some white sectors Yellow-green leaves with green spots Branches with normal leaves on mutant plants Pale flowers with red-gray spots

+ + + + + +

an2-14 an2-16 rt-3 rt-38 Yg3' alp ad"

dw5 p1-m

Pharbitis nil (Japanese morning glory)

Phenotype

Germinal reversions

willow leaf ( m ' )

Dwarf plants White flowers with purple stripes, leaves striped and slightly corrugated Two kinds of variegation depending on modifier: green leaves with white edges or white leaves with green spots Willow leaf is a slender, three-lobed leaf that rarely but consistently gives rise to a maple leaf bud sport

New alleles (S,I,V)'

Referencesc Doodeman et al. (1984b) Cornu (1977); Farcy and Cornu (1979) Cornu (1977)

SJ

Cornu (1977)

SJ

Cornu (1977)

s.1

Cornu (1977)

+

V

+

SJ s.1

+

Modifier'

Doodeman et al. (1984a) Doodeman et al. (1984b) A. G . M. Gerats (personal communication) Bianchi ef al. (1974)* Maizonnier and Cornu (1971)

+

Potrykus (1970)

Imai (1926)

Yellow-green leaves with a few large green areas

+

yellow-inconstant (Y i )

Yellow leaves with a few green areas of variable size

+

xanthic (x)

Yellow leaves with a few green areas

Pecked (p)

White flower with anthocyanin flecks of variable size Flowers have normal colored flecks or parts on a "duskish" or dilute colored background Flowers have anthocyanin spots on a yellowish or white background Yellow leaves with green areas of variable size

duskish (dk)

speckled (sp) Phaseolus vulgaris L. (French bean, green bean) Pisum sativum L. (garden pea) Portulaca grundiJlora (rose-moss, sun plant)

yellow mutable (us)

(S)V

+

S

+

+

A2

White flowers with purple spots

+

inconstant creumish

Creamish colored flower with yellow stripes; may contain orange stripes superimposed on the yellow

+

s,v

Imai (1934)*; Miyake and Imai (1934)*; Kirk and Tilney-Bassett (1978) Imai (1926,* 1927, 1930, 1934)*; Miyazawa (1929*, 1932)*; Miyake and Imai (1934)*; Kirk and Tilney-Bassett (1978) Miyake and Imai (1934)*; Kirk and Tilney-Bassett (1978) Imai (1931)

+

Imai (1931, 1934*, 1934- 1935)

+

Imai (1921, 1931)

+

Coyne (1966, 1967, 1969); Kirk and They-Bassett (1978) De Haan (1930) Ikeno (1929)*; Imai and Kanna (1935)

(continued)

TABLE I (Continued)

Plant

Mutable alleleb

Phenotype

recessive j a k e (e)

Magenta-colored spots and streaks of various sizes on the petals, stems, and leaves Dwarf plants with some normal branches White flower with magenta flakes

Portulaca grandifora (cont.)

Primula sinensis

--

Sorghum bicolor (chocolate corn) Verbena hybrida (common garden verbena)

Variegated seed coat of fine red stripes on a white background White flower with purple spots

Germinal reversions

New alleles (S,I,Vp

Modifield

References‘

+

Ikeno (1929)*; Fabergt and Beale (1942)

+ +

Blakeslee (1920) De Winton and Haldane (1935); Harrison and Fincham (1964) McWhirter (1973)

+ -

I,V

Eyster (1928)

a We have assembled a list of plants where unstable nuclear genes are thought to be involved in producing a variegated phenotype. Mutable alleles of Antirrhinum majus and Zea mays are not included, but are given in Tables I1 and 111, respectively. Many examples listed are discussed in Kirk and They-Bassett (1978). Other examples of plant variegation are mentioned in the literature, but there are insufficient published data to determine the nature of the instability (see for instance the 1930 review by De Haan). These are not included in the table. Imai (1936) lists many plants having a chlorophyll variegation pattern-some of which may be due to unstable nuclear mutations. However, due to the difficulty of analyzing greedwhite variegation patterns (see text and Kirk and Tilney-Bassett, 1978) only a few of these plants are listed in this table. Most forms of variegation listed involve a recessive to dominant phenotypic change. Only four cases of green + white (yellow) variegation are listed. These cases of “reverse variegation” are given as examples and are more dimcult to explain in terms of transposable elements (see Section 11,D). Note that examples of “rogues” and “eversporting” lines have not been included in thk table. Cases where variegation seems to be associated with chromosomal anomalies (see text) have also been excluded. Although only flower color variegation may be mentioned, other plant parts may also show a variegated pigmentation. Derivatives of the original unstable mutable allele are denoted by S (stable recessive allele), I (stable allele showing an intermediate phenotype), and V (new variegated forms). The presence of a modifier is indicated by +. The modifier may be necessary for variegation or may only alter the pattern of variegation (see text). References with a * indicate that the reference was not available to us.

TABLE I1 Mutable Alleles of Anthirrhinum majusa Reversionb Mutable allele albostriata (stri)

bicolor (bic)

--

Phenotype

S

G

Allelic derivatives

Green spots on white leaves; some normal green leaves Large green sectors on yellow-white leaves

+

+

Forms with different degrees of spotting

+

+

Interaction with other mutable alleles and/or modifiersc. Codominance of alleles observed in homozygotes

Yellow-green leaves

-

+

cupuliformis (cup)

Plants with cup-shaped leaves and some shoots with normal leaves Symmetrical, radial

+

+

-

+

Forms with flowers with varying degrees of radial symmetry

-

+

Forms with varying degrees of altered petals and anthers

4

dejiciens-globifera (def-gli) delilah-recurrens (del-rec)

flowers Flower petals reduced to sepals; male tissue missing Tube of corolla basically colorless with purple-red streaks

+

Baur (1910, 1924); Stubbe (1933, 1966) Kuckuck and Schick (1930); Schick (1933); Stubbe (1933, 1966) Schick and Stubbe (1932); Schick (1933, 1935); Stubbe (1933; 1966); Baur (1932) Stubbe (1933; 1966)

chamaeleon (cham)

cycloidea-radialis recurrens (cyc-rad-rec)

References

No interaction with niu-rec and pal-rec

Isolated by Baur and Stubbe, cited in Kuckuck (1936); Stubbe (1966) Baur (1918, 1924); Hertwig (1926); Stubbe (1966) Harrison (1965, 1971)

(continued)

TABLE I1 (Continued) Reversionb Mutable allele depauperatamutabilis (dep-mut) discolor (dis)

-m

eluta-recurrens (el-rec)

flavostriata Mast)

galbana (ga) gilvostriata (gi)

graminifoliamutabilis (gram-mut)

Phenotype

S

G

Small, pointed undulated leaves; fewer smaller flowers Leaves with green spots on white background; blisters on some leaves due to unequal growth; plants with some normal leaves el'lel+ flowers redd; el-lel- flowers ivory with pale red flush; el-rec flowers red with ivory spots Leaves with yellow background and green spots

-

+

+

+

+

AUelic derivatives

+

Yellow-green leaves with green sectors Cream-colored leaves with light green spots

+

+

+

+

Very narrow leaves, sepals, and petals

-

+

References Stubbe (1974)

Schick and Stubbe (1932); Kutscher (1935); Schick (1935); Stubbe (1936, 1966) Derivatives with different numbers of spots

+

Interaction with other mutable alleles a n d o r modifiers'

No dosage effects; effects of different el-rec alleles additive

Doring (1941); Stubbe (1966)

Kuckuck and Schick (1930); Stubbe (1933, 1966); Bergfeld (1958) Stubbe (1974) Derivatives with varying degrees of variegation Derivatives that exhibit different degrees of germinal reversion

Scherz (1927); Stubbe (1933, 1966) Mechelke and Stubbe (1954); Stubbe (1966)

illustris (ill)

incolorata 2recurrens (inc2)

-

incolorata recurrens (inc)

\o

lucida (luc)

microfolia (mic)

mutabilisrecurrens (mut-rec)

Young plants with narrow, upright leaves; less branching, yellow spot on lower lip enlarged and surrounded by red Mottled leaves, red seedlings

Flowers with red spots and streaks on ivory background Young plants with lighter colored leaves; smaller flowers; less intensive pigmentation on lower lip Undulated cotyledons; smaller leaves; fewer flowers per inflorescence; growth of main stem inhibited White flowers with red stripes and streaks

Stubbe (1941, 1966)

+

t

+

+

-

+

-

t

+

+

N o dosage effect of allele in homozygous state; codominance observed

Linnert (1972, 1978)

Isolated by Stubbe, cited in Kuckuck (1936); Stubbe (1966) Schick and Stubbe (1932); Stubbe (1966)

Stubbe (1941, 1966)

Derivatives with different frequencies of germinal reversion

Stubbe (1933, 1966); Kuckuck (1936)

(continued )

TABLE I1 (Continued) Reversionb Mutable allele nitens-mutabilis (nit-mut) nivea-recurrens (niv-rec)

Phenotype

S

G

Allelic derivatives

Longer and lighter colored hypocotyl; lighter colored leaves Flowers with red spots and stripes on white background

-

+

+

+

Derivatives with different hypocotyl lengths Forms with stable white and stable intermediate flowers; others with altered patterns of variegation Forms with stable white and stable intermediate flowers; others with altered variegation patterns

pallida-recurrens (pal-rec)

Flowers with magenta flakes and stripes on ivory background

+

+

perlutea-recurrens (per1-rec)

Pale yellow flowers with red spots

+

+

Interaction with other mutable alleles andor modifiersc

References Stubbe (1966)

“Stabilizer” of mutability

“Stabilizer” J, K eosinea

Allele shows no dosage effects in homozygous state

Isolated by Stubbe, cited in Kuckuck (1936); Hamson and Carpenter (1973); Harrison (unpublished results) Baur (191 1); Mather (1947); Harrison and Fincham (1964); Fincham and Hamson (1967); Harrison and Fincham (1968); Harrison and Carpenter (1973); Jeffries and Sastry (1981, 1982); Sastry et al. (1980) Stubbe (1940, 1966); Doring (1941)

perlutea-mutabilis (perl-mut) perluteavariabilis (perl-uar)

punctata (punc)

-!2

retusa (rea)

spectabilis (spec)

+ Yellow flowers with red spots; flowers with narrower lips; leaves rolled inward; reduced growth and viability Some leaves needle-like, others normal Pale-colored flowers with pale red spots on the lower lip Leaves lighter colored and wavy; upper lip extended with pink flush; major yellow and red spots on lower lip with reduced pigmentation Young plants with lighter colored leaves; width and length of main stem reduced; length of branches reduced

+ (+Y

+

Forms with flowers with varying degrees of spotting and deformation and with leaves with different degrees of rolling

Stabilizer that prevents germinal reversion; factor that enhances germinal reversion

Stubbe (1966); Doring (1941) Stubbe et a / . (1981); Meyer ef al. (1981, 1982)

Baur (1926)

+

+

Stubbe (1941, 1%)

-

+

Stubbe (1%6)

-

+

Stubbe (1966)

(continued)

TABLE I1 (Continued) Reversionb Mutable allele striata (str)

tonsa (ton)

-

uaria (va)

N

variicolor (vac) victrir (vic)

Phenotype

S

Smaller plants; smaller flowers with colorless sectors on lower lip Smaller plants; leaves dented small flowers with extremely short lips Light yellow to yellow-green leaves with pale spots Pink flowers with colorless sectors Upper and lower lips of flowers red

+

Stubbe (1966)

-

Stubbe (1933, 1966)

+

Stubbe (1941, 1966)

+

Stubbe (1966)

-

G

Interaction with other mutable alleles and/or modifiers‘

ALlelic derivatives

References

Stubbe (1933, 1941, 1966); Doring (1937)

a Information on many of the mutable alleles described here is derived from Stubbe (1966). and more complete descriptions of the phenotypes can be found in this source. The capital letter “S” stands for somatic reversion and “G” for germinal reversion. Where “codominance” or “no dosage effects” are indicated, the effect of the mutable allele has been shown to be additive when two copies of it are present in homozygotes. Note that in a number of cases no somatic variegation but frequent reversion events in germinal tissue have been observed. There is some discrepancy concerning the designation of mutable and wild-type alleles of this locus. The terminology used here is according to Stubbe (1966). Parentheses indicate that germinal reversion occurs only in the presence of an appropriate modifier.

TABLE 111 Mutable Alleles in Maize“ Chromosome 1 P-RR

Pericarp and cob color are red, brown, or orange Somatic instability is usually expressed as colored pencarp sectors on a colorless background Mutable alleleb p-VV = p-rr-Mp (“variegated’ ’)

-

N

Elementc

Emerson (1914, 1917, 1922, 1929); Eyster (1924, 1925, 1926); Anderson (1924); Brink and Nilan (1952); Barclay and Brink (1954); Brink (1954, 1958); Brink and Barclay (1954); Fradkin and Brink (1956); McClintock (1956a); Wood and Brink (1956); Kedhamath and Brink (1958); Van Schaik and Brink (1959); Greenblatt and Brink (1962, 1963); Orton and Brink (1966); Greenblatt (1%6, 1968, 1974, 1984) Hayes (1917); Anderson (1924); Barclay and Brink (1954) Barclay and Brink (1954)

MP

p-MM (“mosaic”) Q36 (“striped”)

bz2

W

Bronze-2: Bronze aleurone and plant Somatic instability expressed as bronze aleurone or plant exhibiting purple or red sectors

Mutable allele

Chromosome 3 a

Referencesd

Element

bd-m

Ds

bd-S

Ds

References McClintock (1956a); Osterman and Schwarz (1981); Rhoades and Dempsey (1982, 1983) Bianchi et a / . (1969)

Anthocyaninless: absence of anthocyanin pigments in the plant and aleurone, and brown pericarp with P-rr Somatic instability is usually expressed as colored sectors on a pale or colorless background

Mutable allele

Element

a-ml

SPm

a-m2

SPm

References McClintock (1951a,b, 1952, 1953b, 1954, 1955, 1956a-c, 1957, 1958, 1967a, 1968); Peterson (1965, 1966, 1968); Neuffer (1966) McClintock (1951b, 1952, 1953b, 1954, 1956a, 1961b, 1963, 1964, 1965b, 1967a, 1968); Gorman and Peterson (1978) (continued)

TABLE I11 (Continued) Element‘

Mutable alleleb a-m3 a-m4 a-m5 a-standard = a-dt

Ds Ds SPm rDt

a-m = am-1:Cache a-m(dense) = a l - m

rDt En

a-rug a-var a-mrh ap-m a-px Two unstable a alleles

rug En. rMrh (Nonautonomous) (Autonomous)

References” McClintock (1951b, 1952, 1956b, 1964, 1965b, 1978a); Neuffer (1%5) McClintock (1952, 1953a); Neuffer (1965) McClintock (1961b, 1962, 1964) Emerson (1932); Rhoades (1936, 1938, 1941, 1945); McClintock (1951a, 1956b); Peterson (1965, 1966); Doerschug (1976) Nuffer (1955, 1961, 1965, 1966); McClintock (1951a); Doerschug (1976) Peterson (1957, 1961, 1965, 1966, 1970b, 1978b, 1980); Reddy and Peterson (1977); Nowick and Peterson (1981) Friedemam and Peterson (1980, 1982); Peterson and Friedemam (1983) Peterson (1980) Rhoades and Dempsey (1982, 1983) Nuffer (1956, 1962) Nuffer (1962) Langhnan and Rhoades, cited in McClintock (1951a)

Shrunken-2: Kernels are large, sweet, and watery at milk stage and collapse to a shrunken form upon drying. Shrunken-2 may be the structural gene for ADP-glucose pyrophosphorylase (Tuschall and Hannah, 1982) Somatic instability is expressed as nonshrunken patches on an otherwise shmnken kernel Mutable allele sh.2-ml Chromosome 4 Tu c2

Element

References Hannah and Nelson (1976); Hannah et a / . (1980); Tuschall and Hannah (1982)

Ds

Tunicate ear: Long glumes on ear and tassel Instability at the Tu locus was observed by Mangelsdorf in 1948 (cited in McClintock, 1951a) Colorless aleurone: Little or no anthocyanin color in aleurone, reduced plant color Somatic instability is usually expressed as colored sectors on a pale or colorless background Mutable allele c2-ml c2-m2

Element SPm rSpm

c2-m3

c-2

MP

References McClintock (1964, 1967b) McClintock (1964, 1967b, 1978a) B. McClintock (personal communication); Wienand et al. (1982) I. M. Greenblatt (personal communication)

CL-ldf

c2-m4490 c2-rn826019 c2-m82602I c2-m826040 c2-rn826134 c2-m826204 Chromosome 5 a2

Anthocyaninless: Absence of anthocyanin pigments in the plant and aleurone, and red pericarp with P-rr Somatic instability is usually expressed as colored sectors on a pale or colorless background

Mutable allele'

bt

Element

a2-mI

rSpm

a2-rn2 a2-m3 a2-m4 a2-ms a2-mI I511

(Not Ac-Ds) (Not Ac-Ds) Ds rSpm En

a2-m4 1596 a2-rn4 1629 a2-m6 8140 a2-m6 8144 a2-rn7 8018

En En En En En

References McClintock (1948, 1951a, 1952, 1957, 1958, 1971); Reddy and Peterson (1977, 1978) McClintock (1952, 1953a) McClintock (1952, 1953a) McClintock (1952, 1953a) McClintock (1964) Peterson (1963, 1968, 1974, 1978a); Fowler and Peterson (1974, 1978); Reddy and Peterson (1976, 1977, 1978) Peterson (1978a) Peterson (1974, 1976a,b, 1978a) Reddy and Peterson (1976); Peterson (1978a) Reddy and Peterson (1976); Peterson (1978a) Peterson (1976a, 1978a);Reddy and Peterson (1976)

Brittle endosperm: The mature kernel is collapsed and often translucent and brittle Instability is expressed as kernels mosaic for bt and Bt (blistered appearance)

Mutable allele bt-m Unstable bt Pr

Greenblatt (1965, 1975) J. L. Kermicle (personal communication) Peterson (1983) Peterson (1983) Peterson (1983) Peterson (1983) Peterson (1983)

Idj Ds

Element rSpm-P

References R. L. Phillips (personal communication) Rhoades, cited in McClintock (1951a)

Red aleurone: The mutant allele (pr) changes purple aleurone color to red and may also change purple anther color to red Instability is expressed as various grades of anthocyanin intensity varying from red to purple (continued)

TABLE I11 (Continued) Mutable allele* pr-ml pr-m2 pr-m3

EIementc rSpm rSpm

Referencesd McClintock (1951b) McClintock (1965a) McClintock (1965a)

Chromosome 6

Y

Yellow endosperm: Carotinoid pigments are present in the endosperm Instability is expressed as white endosperm with yellow sectors

Mutable allele

Element

Robertson, cited in Neuffer er a / . (1968) McClintock (195la,b)

white mutable (w-m) y mutable

5 m

Chromosome 7 02

References

Opaque-2: In 02 kernels the endosperm is soft, floury, and dull in reflected light and opaque to transmitted light Instability is expressed as sectors of flint tissue on an opaque background in seed endosperm

Mutable allele 02-m(r) 02-h 02-col (=02-COL UMBIAN) 02-AGROCERES 02-ch (= 02-CHARENTES) 02-261 02-mh

Element

References

rBg rB2h rSd

Salamini (1980, 1981); Salamini et a / . (1982); Montanelli et a / . (1984) Salamini (1981) Salamini (1981); Salamini et a / . (1982); Montanelli et al. (1984)

rSd rBg

Salamini (1981); Montanelli er a / . (1984) Salamini et a / . (1982) Montanelli et a/. (1984) Montanelli et a/. (1984)

Glossy: The wild-type allele ( C / )sustains the synthesis of epicuticular waxes in the maize seedling (Bianchi et a/., 1977). The g / seedling is bright green with water spray adhering in fine drops Instability of 12 g / alleles was noted due to a high reversion frequency (0.17 x to GI (Salamini and Borghi, 1%). The reason for this instability is unknown

Chromosome 9 c

Aleurone color: The wild-type allele ( C ) produces anthocyanin color in the aleurone while c is colorless. C' is a dominant colorless allele Instability is expressed as colored aleurone sectors on a pale or colorless background Mutable allele<

c-ml c-m2 C-ml

c-m4 c-m5

C'-mutable c-standard c-ml I702 c-m4 1963 c-m5 5292 ~-m5 5301 c-m5 5320 C-m.5 5351 c-m5 5453 c-m6 8613 c-m6 8655 sh

Element

Ds Ds (Not Ac-Ds) Ds SPm En En

En En En

En En

En En

References McClintock (1948, 1949, 1950a. 1951a.b, 1952, 1953a) McClintock (1948, 1950a, 1951a, 1952, 1953a) McClintock (1952, 1953a) McClintock (1952, 1953a, 1964) McClintock (1963, 1978a) McClintock (1951a.b) McClintock (1951a) Peterson (1963, 1978a); Gorman and Peterson (1978) Peterson (1976, 1978a) Peterson (1976, 1978a); Reddy and Peterson (1983) Peterson (1978a); Reddy and Peterson (1983) Peterson (1976, 1978a) Peterson (1978a); Reddy and Peterson (1983) Peterson (1978a); Reddy and Peterson (1983) Peterson (1978a); Reddy and Peterson (1983) Peterson (1978a); Reddy and Peterson (1983)

Shrunken endosperm: Structural gene for the enzyme sucrose synthase. In s h kernel the endosperm collapses to form an indented crown Instability is expressed as germinal reversions to sh' Mutable allelef

sh-m(5933)

Element

Ds

References McClintock (1952, 1953b); Burr and Burr (1980, 1981a,b, 1982, 1983); Doring et al. (1981, 1983, 1984); Geiser et al. (1982); Fedoroff et a / . (1983a,b); Courage-Tebbe et a/. (1983)

(continued)

TABLE I11 (Continued) Mutable alleleb

sh-m(6233)

Ds

sh-m(6258)

Ds

sh-m(6795)

Ds

sh-5582 sh-5584 sh-5586 sh-5596 m

bz

Elementc

Referencesd McClintock (1952, 1953b); Burr and Burr (1980, 1981a,b, 1983); Doring et a / . (1981, 1984); Geiser et al. (1982); Fedoroff et a / . (1983~);Weck e f al. (1984) McClintock (1952, 19853b); Burr and Burr (1980, 1981a, 1982, 1983); Fedoroff et a / . (1983b) McClintock (1952, 1953); Burr and Burr (1981a,b, 1982); Geiser et a / . (1982); Fedoroff et a / . (1983a) Mottinger et a / . (1984b) Mottinger et al. (1984b) Mottinger et al. (l984b); S. Dellaporta (personal communication) Mottinger et al. (1984b)

Tz86

Bronze: A bz mutation changes a purple aleurone to brownish pale, the plant becomes reddish brown, and the tassel anther will fluoresce under UV light Somatic instability is recognized by a phenotypic bz + Bz change in aleurone and plant tissue Mutable allele

Element

bz-ml bz-m2 (=Acbz-mZ)

Ds Ac

bz-mil bz-m4

(Not Ac-Ds) Ds

bz-mut (=bz-m-rh) bz-ml3 bz-m805137 bz-m826301 bz-m826302

rMut rSpm = Rs

References McClintock (1951a,b, 1952); Dooner and Nelson (1977) McClintock (1951a,b, 1952, 1955, 1956a-c, 1962); Dooner and Nelson (1977); Fedoroff et a / . (1984) McClintock (1952, 1953a) McClintock (1953a, 1956~); Dooner and Nelson (1977); Dooner (1980); Gerats et a / . (1983) Rhoades and Dempsey (1982, 1983) Klein and Nelson (1983) Peterson (1983) Peterson (1983) Peterson (1983)

wx

Waxy endosperm: The wild-type allele wx+ determines the amylase content of endosperm tissue and pollen (Sprague et al., ( 1943) Somatic instability is observed by 12-KI treatment of endosperm tissue

Mutable allele wx-ml

Ds

wx-m2 wx-m3

wx-m5 wx-m6

(Not Ac-Ds) (Autonomous) (Not Ac-Ds) (Autonomous) (Not Ac-Ds) Ds Ds

wx-m7

AC

wx-m8

rSpm = Spm-I8

Acwx-m9

AC

WX-BJ WX-B~

AC Ds

wx-m

(Autonomous)

wx-m4 d

W N

Element

References McClintock, 1948; 1950a, 1951a,b, 1952, 1953a); Nelson (1968); Echt and Schwartz (1981); Rhoades and Dempsey (1983) McClintock (1948, 1951a, 1952, 1953a) McClintock (1951a,b, 1952, 1953) McCIintock (1951a. 1952, 1953) McClintock (1952, 1953a, 1964, 1965b) McClintock (1952, 1953a, 1964); Nelson (1968); Echt and Schwartz (1981); Fedoroff et al. (1983b); Shure et al. (1983) McClintock (1953a, 1964, 1965b, 1978a); Peterson (1974); Schwartz-Sommer e f a / . (1984); Behrens et al. (1984) McClintock (1961b, 1978a); Nelson (1968); Echt and Schwartz (1981); Shure et al. (1983); Schwartz-Sommer et al. (1984) McClintock (1963, 1964); Fedoroff et al. (1983b); Shure et al. (1983); Pohlman et al. (1984) Nelson (1968); Echt and Schwartz (1981) Nelson (1976) cited in Echt and Schwartz (1981); Echt and Schwartz (1981) Sager (1951)

TABLE I11 (Continued) Chromosome 10 R’

Aleurone andplant color: Controls the expression of anthocyanin pigment in aleurone and certain plant parts Somatic instability is usually expressed as sectors of anthocyanin pigment on a colorless background

Mutable allele8

Element

R-st (“stippled”)

I-R

mR”J(“R-Navajo”) R-mb (“marbled”)

MP

R-r#2

rFcu and rSpf

r#IO

rFcu and rSpf rFcu

r-cu

R% r-ml r-m3 r-m9

Ds DS

Ds

References Fogel (1950)*;Ashman (1960, 1965); Kermicle (1970, 1973, 1984); Dooner and Kermicle (1971); Williams e r a / . (1984) Williams and Brink (1972); Brink and Williams (1973) Brink (1956); Brink and Mikula (1957); Brink and Weyers (1957); Weyers (1961) Sastry and Kurmi (1970) cited in Gonella and Peterson (1978); Gonella and Peterson (1978) Peterson (1981) Gonella and Peterson (1977, 1978) Dooner and Kermicle (1974) Kermicle (1980) Kermicle ( 1980) Kermicle (1980)

Chromosome location unknown Luteus: Gene associated with chlorophyll production lu Somatic instability is expressed as a yellow seedling variegated for normal green tissue Mutable allele lu-m (“mutable luteus”)

Element Spm? (autonomous)

References McClintock (1946, 1951a, 1961b); Smith (1960)*

Yellow-green chlorophyll: The recessive y g locus mutates to form a darker color chlorophyll

Mutable allele

Element

References McClintock (1948, 1951a) McClintock (1948, 1951a)

Pmp-1817

-W

A dominant mutable allele affected chlorophyll development in the mature plant Instability is expressed as virescent older leaves with dark green stripes. Element at the locus is unknown, but it is not of the E n 4 system (Peterson, 1963, 1964) Variegated white seedlings: White seedling with sectors of pale yellow or of normal green tissue. Element unknown (McClintock , 1946) Variegated light green: Light green seedling with streaks of normal green tissue. Element unknown (McClintock, 1946)

The nomenclature and phenotype given for a particular gene were taken from Neuffer er a / . (1968). The nomenclature of the various mutable alleles may be different from that given in a particular publication-for often even one author uses various designations for the same allele. For example, the mutant a,m-’ (B. McClintock’s nomenclature) has been referred to as a l- m - / or as u-ml in the literature. We chose the latter nomenclature for simplicity. Many of these mutable alleles are discussed in recent review articles on maize transposable elements (Fedoroff, 1983; Doring and Starlinger, 1984). Only the original mutable allele is listed, although many alleles gave rise to both stable and unstable derivatives. Many new unstable alleles are being generated, especially by the mutator system (Mu)described by Robertson (1978). This system induces mutations at a frequency approximately 30 times greater than the usual spontaneous mutation rate for maize, with approximately 35% of the new mutants being unstable (Robertson, 1978). These unstable loci include a, a2, bz, and hundreds of mutable nuclear chloroplast pigment mutants (D. S . Robertson, personal communication). To date, there is not enough information concerning most of these unstable alleles to include them in the table. Orton and Brink (1966) describe 146 independent unstable mutations at the P locus, most of which are due to Mp at the locus. Element refers to the element responsible for the mutable allele in the original unstable mutant. If the regulatory element is listed, the allele is considered to be autonomously controlled. The element listed is designated according to the name given in the publication(s) although several such elements are thought to be functionally equivalent (e.g., Spm = En, Ac = M p ) . The designation of an element at a nonautonomous locus is given by a small r as the first letter. This r stands for “receptor” (e.g., rDr means the receptor element that requires the regulator Dt to be present in the genome in order to produce the instability). Most of the elements are discussed in the text or listed in Table IV. However, a few comments are given as follows: rSpm = I : since Spm and En are functionally equivalent (Peterson, 1965a) then the rSpm (receptor for S p m ) is functionally equivalent to I, the receptor element for the en regulatory element; M u l : An element thought to be associated with a high frequency of mutation (Strommer et al., 1982); E d : barley stripe 1, an element present in a line previously infected with barley stripe mosaic virus; I d j Defective I, where I is the receptor element for the E n 4 two-element system; rSpm-P: A receptor element that responds to the Spm element isolated by R. Phillips; rE2h, rSd, and rBg may be the same receptor element for the two-element Bergamo system ( B g ) (Salamini, 1981); Tz86: Transposon Zea 86. The reference list for some mutable alleles may not be complete but should suffice to initiate an inquiry into the details of that allele. Most references to articles in the Maize Genetics Cooperation News Letter have not been included. Peterson (1978a) lists 72 independently derived En-controlled mutables of the a2 and c loci. Only a few of the better characterized alleles are included in this table. f sh-ml and sh-m2 mutable alleles (both controlled by the Ac-Ds system) are only mentioned in Table 1 of McClintock (1964). It was unclear if these are the same as alleles already listed. mRnj consists of 26 mutable alleles which all carry Mp (Brink and Williams, 1973). Reference was not available to us.

132

PATRICIA NEVERS E T A L .

Plant: Antirrhinum majus Alleles: pullida-recurrens pallida-tubocolorata

(red-and-white variegated flowers) (white flowers with ring of red color at base)

FI Genotype: pal-reclpal-tub Phenotype: Variegated flowers

F2 Expected phenotypes: pal-rec Gametes I pal-rec pal-tub

variegated variegated

pal-tub

variegated white

3 variegated : 1 white

Temperature at which F,grown 25°C 15°C

Observed phenotypes: Total numbers of plants scored Variegated 2017 3199

1493 I589

Phenotypes White

Revertant

Others

499 843

55 672

0 96

Fig. 1. Monoallelic segregation of the pallida-recurrens allele of Antirrhinum mujus. The pallida locus is involved in a late step in anthocyanin synthesis (Harrison and Stickland, 1974). The pallida-recurrens allele is thought to be caused by integration of a transposable element at the pallida locus. Frequent somatic reversion of this allele leads to variegated pigmentation in flowers. The pallida-tubocolorata allele is a stable, recessive allele characterized by white flowers with only very little pigmentation at the base of the corolla. Note that in the Ff, revertants and other new phenotypes are observed that are not expected by normal Mendelian genetics. The incidence of such individuals (and the degree of somatic variegation) is higher at 15°C than at 25°C. The number of variegated progeny is accordingly somewhat lower at 15°C. (Data taken from Harrison and Fincham, 1964.)

Sometimes the segregation ratios indicate that not one nuclear gene but two or more are involved in producing the variegated phenotype. One contributing factor is always a mutable allele that frequently reverts to wild type, presumably due to the presence of a transposable element. Regarding the second locus, two different cases can be distinguished: those where the second factor is required for mutability of the first component, and those where the second factor only modulates mutability of the first. In some instances the second factor is known to be a transposable element itself, while in others this is not clear (see Section V). The most prominent examples where a second independently segregating locus is required for mutability of another one are those of the two-

PLANT TRANSPOSABLE ELEMENTS

133

component systems in Zea mays. Here the second factor is a transposable element capable of autonomous integration, excision, transposition, and other types of activities typical of such elements. This component has been termed the “regulator” (McClintock, 1961a) or autonomous component (Fedoroff, 1983). The element it regulates is defective with respect to all of these activities and can perform none of them of its own accord. This defective element can inhibit expression of a locus simply by virtue of insertion, but it is stable unless a suitable regulatory element is present somewhere in the genome. The regulator can activate in trans the defective component, probably by means of some diffusible protein or RNA “signals.” Accordingly the defective component has been termed the “receptor” (Fincham and Sastry, 1974; Peterson, 1965a) or nonautonomous component (Fedoroff, 1983). Data from a cross exhibiting a segregation ratio typical for two-component transposable element systems are shown in Fig. 2. Information concerning the “signals” produced by a regulatory element is derived primarily from classical genetic experiments. One function that seems to be common to all regulators is a “mutator” function required for excision, transposition, and other activities of both the regulator and related receptor components (McClintock, 1954, 1957, 1961b, 1965a; Fedoroff, 1983). The complex regulatory component of the Spm(En) system is believed to encode one or possibly two other activities in addition to the mutator function: a suppressor, required for complete inhibition of expression at a locus under the control of an element from this system (McClintock, 1954), and possibly an activator for maintaining transcription of the element (McClintock, 1971). A model of how these three functions interact was presented by Nevers and Saedler (1977). The interaction between the two components in a two-component system is highly specific, that is, a given receptor component responds to signals of certain regulatory components but not to others (Peterson, 1978a; Peterson, 1980; Friedemann and Peterson, 1982). This specificity has been exploited by Peterson and his co-workers (Peterson, 1980; Friedemann and Peterson, 1982) to classify several two-component systems in maize, summarized in Table IV together with very recent data on two new systems (Rhoades and Dempsey, 1982, 1983). On the basis of specificity of interaction, the independently isolated regulatory elements En and S p m are considered to be functionally equivalent (Peterson, 1965a), likewise Ac and M p (Barclay and Brink, 1954). It should also be noted that regulatory elements that show the same basic specificity of interaction may not be functionally identical. Thus Ac2 can readily transactivate some Ds elements but not others, and Spfvaries in its ability to mobilize certain receptor components (see Table IV).

A. A two-component system Plant: Mirubilis julapu L. Alleles: Y (dominant allele; red flowers) Y“ (white flowers when V is not present in the genome; white flowers with red spots when V is present in the genome) V (regulatory component whose activity is required for mutability of Y “ ) y (regulatory component mutated, inactive, or missing)

F, Genotype: Phenotype:

Gametes

I

YP vu Red flowers

F2 Expected phenotypes: Y, v Y, u P,v red red red red

red red red red

P>u

red red variegated variegated

red red variegated white

12 red : 3 variegated : I white

Observed phenotypes: 38 red : 7 variegated : 3 white (0.3 < p < 0.5)

B. A mutable allele plus an independently segregating modifier Plant: Alleles:

Antirrhinum majus pallida-recurrens (red-and-white variegated flowers) pallida-tubocolorata (white flowers with ring of red color at base) “Stabilizer” (factor that reduces mutability of pal-rec) StlSr (low mutability of pal-rec) Stlst (medium mutability of pal-rec) stlst (high mutability of pal-rec)

FI Genotype: Phenotype:

pal-reclpal-tub, Stlsr

Medium variegated flowers

F2 Expected phenotypes: pal-rec

Gametes pal-rec, St pal-rec, sr pal-tub,

pal-rec

pal-tub St

si

pal-tub st

low variegated

medium variegated

low variegated

medium variegated

medium variegated

highly variegated

medium variegated

highly variegated

low variegated

medium variegated

white

white

medium variegated

highly variegated

white

white

st

pal-tub, st

6 medium variegated : 3 highly variegated : 3 low variegated : 4 white

Observed phenotypes: 99 medium variegated : 54 highly variegated : 73 low Variegated : 65 white 214 medium variegated : 152 highly variegated :90 low variegated : 136 white

PLANT TRANSPOSABLE ELEMENTS

135

Evidence for two-component systems in which the second component seems to be a regulatory element comparable to those of maize has also been obtained for Oryza sutiva, Mirabilis jalapa, Elymus junceus, and Imputiens balsaminu L. (see Table I for references). Other cases in which segregation ratios indicate that two loci contribute to the variegated phenotype are those in which a modifier of variegation is involved. In these cases one locus apparently carries an autonomous transposable element capable of executing all the activities required to produce a variegated phenotype. But a second locus can influence the activities of the transposable element so that a new kind of variegation pattern occurs. However, this modifier is not absolutely required for variegation. Sometimes it has a stabilizing effect on the transposable element in question (Demerec, 1931; Imai, 1931; Rhoades, 1938; Harrison and Fincham, 1968; Harrison and Carpenter, 1973, 1980; Gerats et al., 1982), so that fewer reversion events occur. Sometimes the modifier enhances the instability of a transposable element so that the degree of variegation and/or germinal reversion is increased (McClintock, 1956c, 1957, 1958, 1965a; Punnett, 1935; Ashman, 1960, 1965; Mulder etal., 1981). Still other modifiers can alter the time in development at which reversion events occur (Punnett, 1935; Sand, 1976; Reddy and Peterson, 1983). For example, in the presence of a modifier, fine spotting, indicative of delayed events, may occur instead of the coarse spotting characteristic of earlier reversion events (Reddy and Peterson, 1983; Sand, 1976). In all cases involving modifiers, plants that are heterozygous for both loci segregate progeny with two different kinds of variegation pattern, as shown in Fig.

Fig. 2. Examples of diallelic segregation ratios obtained with mutable alleles. (A) Segregation of the factors of a two-component system in Mirabilis jalapa. The Y locus is involved in pigment synthesis in this plant. The Y" allele is a mutable allele that may harbor an element comparable to a receptor component of Zea mays. The V allele is an independently segregating factor required for mutability of Y" and is thus comparable to a maize regulatory component. In all plants a wild-type R allele is present in the genome, which permits red pigments to be synthesized. Note that P is stable when the V component is missing or inactive. (Data taken from Spitters er a / . , 1975.) (B) Segregation of a mutable allele and an independently segregating modifier of this allele in Antirrhinum majus. The pallida-recurrens allele is an unstable allele of the pallida locus, which is involved in anthocyanin synthesis. The pallidatubocolorara allele is a stable recessive mutation causing pigmentation in the flower to be restricted to the base of the corolla. "Stabilizer" is a semidominant locus, perhaps a transposable element (see text), that reduces instability of the pal-rec locus so that less somatic variegation is observed. The observed values differ slightly from the expected ones, possibly due to difficulties in scoring. Scoring may be complicated by the fact that pal-reclpal-rec plants have twice as many flakes as pal-reclpal-rub ones (Harrison and Fincham, 1968). Note that the pal-rec allele is by itself capable of inducing variegation, regardless of whether or not the stabilizer is present in the genome. (Data taken from Harrison and Fincham, 1968.)

TABLE IV Two-Component Systems of Transposable Elements in Maize"

Responsive allele:

I a-m(r)

rSpm a-ml

Ds a-Ds

Ds *b

Ds bz2-m

rdt a-dt

rFcu r-cu

rFcu r#IO

ruq a-ruq

rBg o2-m(r)

rMrh a-mrh

rMut bz-mut

+

-

-

+

+

-

+

Data are taken from Friedemann and Peterson (1982), and from Rhoades and Dempsey (1983). * indicates Ds in standard position on chromosome 9, which generates variegation by inciting chromosome breakage. A + symbol indicates that a receptor element is capable of responding to signals from the regulator, usually manifested by a variegated phenotype. A - symbol means that there is no response or the response has not been tested. Where neither nor - is present, no test has been performed. (-) means that the response requires unusually high doses of the regulator element (see text).

+

PLANT TRANSPOSABLE ELEMENTS

137

2. However, it should be emphasized that definite conclusions concerning the presence of a modifier can be drawn only if a clear-cut diallelic segregation ratio can be demonstrated. In other cases where progeny with different degrees of variegation are observed but no Mendelian segregation ratios, other explanations are possible. Here one could imagine that multiple modifiers are responsible (see Section V) or that alterations have occurred in or around the transposable element causing mutability. Origin of Receptor Elements There is considerable evidence that some receptor elements originate from regulatory components. Thus in a number of cases it was found that when the autonomous regulatory element En(Spm) transposes away from a locus, part of it seems to be left behind. This residual element behaves like a receptor element in that it responds to transactivating signals from a regulatory element elsewhere in the genome (McClintock, 1962; Peterson, 1961, 1968; Reddy and Peterson, 1976). In other cases alterations within an autonomous element inserted at a particular locus seem to give rise to nonautonomous derivatives without transposition of the element away from the locus. For example, derivatives of the bz-m2 mutant, which harbors A c at the bz locus, have been found to behave as if a Ds element were present at the bz locus (McClintock, 1955, 1956b, 1962). The Ds element at the wx locus in the mutant wx-m9 also arose in the same manner (McClintock, 1963). Thus in these cases one can expect that the structure of the receptor component resembles that of the corresponding regulators. Support for this idea on the basis of molecular data will be presented in Section 111. C. REVERSION EVENTS

Since most of the mutable alleles analyzed so far are alleles of the nuclear genome, we began our considerations on how to identify transposable element-induced mutations be examining segregation patterns typical for nuclear mutations. We shall now direct our attention to a prominent feature of most mutable alleles, their reversion to a phenotypically wild-type condition in somatic and/or germinal tissue. As early as 1910 Correns interpreted the greedwhite variegation of some mutants of Mirabifisjulapa as being due to a transition from the homozygous recessive state to a dominant heterozygous state in the colored parts of the plant. Depending on whether this transition occurs early or later in development, larger or smaller sectors of colored tissue arise. Although this hypothesis has not yet been verified by molecular data for Mimbilis jalupa, supporting evidence is available for other unstable mu-

138

PATRICIA NEVERS E T A L .

tants discussed in Section III,G. In these cases the two types of tissue in the variegated plant definitely differ genotypically. However, when molecular data are not available, the strongest indication that variegation involves mutational change is when variegated plants produce some wildtype or nearly wild-type offspring that later breed true. This can be attributed to reversion events occurring in germinal tissue, which results in wild-type gametes. If reversions to wild type occur frequently in germinal tissue, distorted Mendelian ratios will be found in crosses involving a mutable allele. Wildtype or nearly wild-type plants may occur where they are not at all expected, or an excess of wild-type individuals may be observed (see Fig. 1). Somatic and germinal reversion may not always involve excision of the element responsible for mutability at a given locus (McClintock, 1953, 1956c; Doodeman et al., 1984a). Other possibilities will be discussed in Section 111. One question for which there is no universal answer is whether the reversion events that occur in somatic tissue are equivalent to those that take place in germinal tissue. In a few cases that have been analyzed at a molecular level, reversion was shown to involve excision of the transposable element, and this could be demonstrated in both somatic tissue and germinal revertants (see Section 111,G). The fact that the frequencies of somatic and germinal reversion events are often strongly correlated also lends support to the concept of equivalence. The stabilizing effect of higher temperatures (25°C) on both somatic and germinal reversion of the mutable pul-rec mutation of Antirrhinum majus (Harrison and Fincham, 1964; see Fig. 2) provides further evidence for a positive correlation of this kind. There are, however, some striking exceptions such as the perlutea-uuriubilis allele in Antirrhinum (Stubbe et al., 1981) and the mutable alleles of Callistephus chinensis (Hondelman, 1959), where no positive correlation can be observed. In still other cases either somatic or germinal reversion may be missing completely. For example, some mutants of Antirrhinum have been reported in which frequent reversion in germinal tissue obviously occurs, leading to a large number of revertants among the progeny of the mutant, but there is no incidence of somatic reversion (see Table 11). Reversion in germinal tissue of the A . mujus mutant perlutea-variabilis occurs only in the presence of a specific modifier (Stubbe et al., 1981). The modifier of the mutable u locus in Nicotiana has the same effect (Sand, 1976). Germinal reversion but no somatic reversion is also observed in innumerable inbred lines with a tendency toward “sporting” or toward producing socalled “rogues.” In these lines wild-type or off-type individuals appear

PLANT TRANSPOSABLE ELEMENTS

139

(Burr and Burr, 1981a). very frequently, sometimes at the rate of 1 x This subject has been reviewed by other authors (Dermen, 1960; Jensen, 1965; Pearson, 1969) and interpreted by Burr and Burr (1981a) as follows: “Rogue” or “sporting” lines are often commercial lines containing mutations that have been maintained for many generations by strong artificial selective pressure. It is conceivable that a large number of these mutations originated by insertions of transposable elements and are thus basically unstable. They revert frequently to wild type, and the entire population would eventually revert to wild type in the absence of counterselective pressure. Cases in which somatic variegation but no germinal reversion is observed have already been discussed (Section 1,B). Since reversion does not take place in germinal tissue, it is questionable whether the somatic events observed are due to genetic alterations. Thus transposable elements may not be involved in these cases. D. REVERSE VARIEGATION

In considering reversion events we have so far discussed mainly those examples in which reversion occurs from a homozygous recessive state to a heterozygous one, manifested, for example, by pigmented spots on a colorless or nearly colorless background. The reverse phenotype, that is, colorless spots on a pigmented background, has also been observed (McClintock, 1968; Peterson and Weber, 1969; Peterson, 1980; Reddy and Peterson, 1984; Cornu, 1977; Harrison and Carpenter, 1977, 1979, 1980; Doodeman et al., 1984b; Gerats et al., 1984b). This phenomenon can be interpreted in terms of transposable elements as well. Several different theoretical explanations for the reverse variegated phenotype are conceivable. In the maize mutants a-m2 (McClintock, 1968) and a-m(Au) (Peterson, 1980) elements belonging to the Sprn(En) system are known to be integrated at the a locus, yet expression of the locus is apparently unimpaired, and kernels of these mutants are basically pigmented. Here one could imagine that the element is located in the control region of the locus in such a way that transcription originates at a promoter within the element and continues into the locus (see Nevers and Saedler, 1977). Expression of the locus would therefore depend on active transcription of the element. If for some reason transcription should no longer be initiated at the promoter in the eIement, as during an inactive phase, then expression of the a locus would also cease. Thus occasional inactivation of transcription might explain the white sectors sometimes observed with such mutants. Another case involves revertants of the transposable element-induced mutants anl-sl+ and a d - s l p - + in Petunia hybrida (Bianchi et al., 1978;

140

PATRICIA NEVERS ET A L .

Doodeman et al., 1984b). Plants heterozygous for the original mutable allele and an anZ-+l+ revertant allele have red flowers with many white spots. Such spots are not observed in heterozygotes carrying the mutable allelle and a true wild-type allele. Moreover, the white spots on the flowers of the hybrid revertant contain colored dots. As discussed by Doodeman et al., (1984a), this suggests that the element is retained at the locus in phenotypic revertants and continues to be unstable. Rearrangements in the sequence of either the element or the target site may be responsible for switching from the mutant to the revertant condition and back again, although other explanations are conceivable. A revertant line derived from the unstable pal-rec mutant of A. majus has been isolated that also has red flowers with many pale-colored and some white spots. In this case a completely different interpretation has been advanced (Harrison and Carpenter, 1979). It is based on the assumption that the element responsible for the pal-rec mutation is excised in the process of reversion and that this element and/or others continue to transpose in the revertant. The pale spots are thought to arise by insertion of a transposable element at a locus involved in anthocyanin synthesis, generating a phenotypically distinguishable heterozygous condition. To explain the occurrence of white spots either a one-step or a two-step process can be postulated. In the first instance one could imagine that an element integrates in a single chromosome homologue and generates a dominant mutation leading to acyanic tissue. This is plausible since dominant mutations of this kind are known, for example, the c-Z and c2-Zdfmutations in Zea mays and the Eluta mutation in A. majus, which are thought to result in accumulation of an inhibitor of pigmentation. On the other hand, if insertion of a transposable element leads to a recessive mutation in one chromosome homologue, then a two-step process is required to generate a white spot. In this case some form of gene conversion or mitotic crossing over would be necessary to reproduce the recessive mutation in the second chromosome homologue. Both processes would require a high frequency of transposition or possibly a preference for integration at loci involved in anthocyanin synthesis. The novel aspect of the interpretation of reverse variegation in A. majus presented by Harrison and Carpenter (1979) is that spotting is due to transposition and integration of transposable elements. Accordingly this phenomenon was termed “resurgence of instability.” That reverse variegation in A. majus arises by the same mechanism that generates red spots on white flowers in a pal-rec line, that is, by transposition, is supported by the fact that both kinds of spotting are affected by the same factors (Harrison and Carpenter, 1979). In both cases the frequency of spotting is reduced at higher temperatures (25°C) and under the influence of a stabi-

PLANT TRANSPOSABLE ELEMENTS

141

lizing gene elsewhere in the genome. It is possible that “resurgence of instability” is due to specific mobilization of elements related to the one responsible for the pal-rec mutation. Molecular analyses of the elements responsible for the original pal-rec mutation and for a new niu-rec allele derived from the unstable pal-rec revertant are being conducted at present (H. Sommer, personal communication) and should help to clarify the matter.

E. VARIEGATION ASSOCIATED WITH CHROMOSOMAL ANOMALIES

So far we have discussed two characteristic aspects of transposable element-induced mutations in the nuclear genome: nearly Mendelian segregation of the variegated trait and reversion of the mutable allele in somatic and germinal tissue. An unusual set of variegated mutants is known which are definitely of nuclear origin and exhibit germinal reversion but nonMendelian segregation ratios. These are mutants in which defined chromosomal alterations such as breaks and translocations occur that can be observed cytologically. In maize, for example, the transposable element Ds (Dissociation) induces chromosomal breaks that can be observed directly (McClintock, 1945, 1946, 1947, 1948, 1951a). These breaks only occur when an Ac regulatory element is present in the genome (McClintock, 1947; Barklay and Brink, 1954). In a variegated plant of Nicotiana, mutant tissue exhibits a conspicuous block of heterochromatin at a defined position in the genome that is missing in the genome of revertant tissue (Burns and Gerstel, 1967). Many other examples of chromosomal anomalies associated with a variegated phenotype are summarized by Kirk and Tilney-Bassett (1978). It is not certain whether transposable elements are involved in all of these instances, but the case of Ds suggests that this may be true in some instances. F. STABLE A N D UNSTABLE ALLELIC VARIANTS OF MUTABLE ALLELES

None of the criteria discussed so far for recognizing transposable elements is absolutely foolproof. If segregation ratios and evidence for somatic and germinal reversion are ambivalent, a third characteristic of transposable elements can be considered: their ability to generate new derivatives with heritably altered phenotypes. Some of these derivatives may have a stable recessive or intermediate phenotype while others may exhibit an altered pattern of variegation with respect to frequency and timing of somatic reversion. Cases in which new alleles have been derived from a mutable one are shown in Tables I and I1 (Table 111 does not

142

PATRICIA NEVERS ET AL.

include this information although many mutable alleles of Zea mays have produced allelic variants). When a derivative with an altered phenotype is isolated, the first question to be answered is whether or not this is due to a mutation. It is a wellestablished fact, for example, that environmental factors such as temperature and light can strongly influence the degree of variegation observed, irrespective of any mutational change in the organism (Winton and Haldane, 1935; Rhoades, 1941; FabergC and Beale, 1942; Sand, 1957; Harrison and Fincham, 1964; Harrison and Carpenter, 1973, 1979; Bianchi et al.. 1978). The obvious indication that a hereditary alteration is involved is when a true-breeding line exhibiting the altered phenotype can be established. This, of course, depends in turn upon the stability of the allele responsible for the altered phenotype. If the new allele is itself highly unstable, it may be difficult if not impossible to establish a truebreeding line. Assuming that only one or two factors are responsible for the altered phenotype, another question to be answered is where the mutation has taken place, at the locus controlled by a transposable element or at an independent modifying locus? This can be determined by examining segregation ratios in appropriate crosses. In the two-element systems of Zea mays, for example, mutations within or adjacent to either the receptor or the regulator component can lead to an altered phenotype. Mutational alterations of independently segregating Ac and Spm regulator elements are known which cause a reduction in mutability, manifested by kernels with fewer andor smaller spots of revertant tissue (McClintock, 1956c, l957,1961a, 1963,1964, 1965b). Such weak “states” of the regulator may alternate with stronger “states” of the element (McClintock, 1959; 1961a,b, 1963,1965b). Still other allelic derivatives of En(Spm) and A c are known that exhibit cyclic changes in activity (McClintock, 1957, 1958, 1959, 1961b, 1962, 1964, 1971; Peterson, 1966). In addition to mutations in or around an independently segregating locus, alterations can occur at the locus immediately controlled by a transposable element. For example, colorless, intermediate, and various different variegated alleles of loci apparently controlled by autonomous elements have been reported for Callistephus chinensis (Hondelman, 1959), Anfirrhinurn majus (Fincham and Herrison, 1967), and Petunia hybrida (Bianchi et al., 1978; Gerats et al., 1984a,b; Farcy and Cornu, 1979). Likewise, many new derivatives of En-controlled a and a2 loci in Zea mays have been found (Peterson, 1960, 1961, 1968, 1970b, 1976a,b, 1980; Fowler and Peterson, 1974, 1978; Reddy and Peterson, 1976). Many of the latter seem to have arisen when En(Spm) transposed away from the locus.

PLANT TRANSPOSABLE ELEMENTS

143

Under the influence of an appropriate regulator component (McClintock, 1955) mutations can also occur in or around a receptor element integrated at a given locus (McClintock, 1955, 1957, 1958, 1965a, 1968; Peterson, 1961, 1970a; Reddy and Peterson, 1984). McClintock (1955) referred to these variants as “states of a locus.” The best documented examples of this kind involve derivatives of mutable alleles of the a locus in maize. The receptor elements in some of these derivatives no longer respond to mutator signals of the regulator, and kernels of these derivatives have a stable, nonvariegated phenotype even when an active regulator is present in the genome. Pigmentation in such stable derivatives is homogeneous and can vary from null to an almost wild-type level. Other derivatives exhibit altered patterns of somatic variegation. A unique allelic variant of an Spm-I-controlled locus is a2-ml. It contains a receptor element of the Spm system that responds to the suppressor of Spm but not to the mutator function (McClintock, 1957, 1958, 1961a,b). Kernels of this derivative are almost fully pigmented in the absence of Spm and colorless in its presence. Still other derivatives of the a locus contain elements that are unstable only in particular parts of the plant. For example, in the Enbearing mutant a-m(crown) variegation is observed only in the crown of the kernels, while instability is restricted to the base of the kernel in am(flow) (Peterson, 1965b, 1966). In these derivatives En-(Spm) seems to be programmed to respond to tissue-specific signals. It is also conceivable that mutations in the immediate vicinity of a transposable element lead to an altered phenotype. Deletions of a flanking sequence, for example, might affect excision of the transposable element or its influence on gene expression. One such case involving the maize element MuZ has been verified by DNA sequence analysis (L. Taylor and V. Walbot, personal communication; see Section 1,H). Similarly, mutation of an external promoter might affect transcription across the element and alter transcription from sequences within the element itself. It has been suggested that a deletion may have fused the bz locus to the control region of the sh locus in the maize mutant bz-m4, and that this is responsible for the altered timing and tissue specificity of bz expression observed in the mutant (Gerats et a f . ,1983b). However, it is now known that the bzm4 mutant contains a deletion that extends to a position 500 bp upstream from the translational start signal of the s h locus (U. Courage, personal communication; Weck et al., 1984).To accommodate this finding with the fusion hypothesis one would have to assume that the sh locus is subject to control by a DNA sequence that is further upstream than the promoter region of the sh locus. Finally the exact position of a transposable element in the genome may have a decisive effect on the kind of variegation pattern produced. Peter-

144

PATRICIA NEVERS ET A L .

son (1976a, 1977) has proposed that varying insertional sites of a transposable element at a given locus may be responsible for various derivatives of mutable alleles in maize. Likewise Brink and Williams (1973) concluded that different modes of integration of Mp(Ac) at the r locus in maize are responsible for the wide array of derivatives with different patterns of somatic variegation observed. The frequency of germinal reversion of the maize od-m(r) allele controlled by the receptor element of the Bg system also seems to be dependent on the exact position of the regulatory component on the same chromosome: the farther away the regulator is located, the lower the frequency of reversion (F. Salamini, personal communication). Molecular analysis of derivatives of mutable systems will be required to distinguish between the alternative explanations for “states of a locus” listed above. Examples of such investigations are presented in Section II1,H. G. TRANSPOSITION: THE MOST CHARACTERISTIC PROPERTY OF

TRANSPOSABLE ELEMENTS

1 . General Remarks The property of transposable elements which is unique is at the same time the one most difficult to demonstrate unequivocally, and that is their ability to transpose. Several lines of classical genetic evidence outlined below suggest that transposition in plants occurs by excising an element from one position and reinserting it at another. Based on the assumption that somatic reversion also occurs by means of excision and that excising is inevitably accompanied by transposition, some authors even go so far as to equate each spot or stripe on a variegated plant with a transposition event. While this may hold true in some cases, caution is warranted in making any generalizations about transposition in plants. First, the means available for studying transposition are relatively crude, involving populations of elements, cells and plants in in uiuo studies and populations of molecules in in uitro experiments. Ideally one would like to be able to follow the movements of one uniquely marked element in a single cell. Second, there is some evidence that transposition may occur by more than one mechanism in a single organism. In prokaryotes, for example, one mode of transposition seems to be a process in which a copy of the element moves (Grindley and Sherratt, 1978; Shapiro, 1979; Arthur and Sherratt, 1979), a mode sometimes referred to as replicative transposition. In this case excision and transposition seem to be independent events. However, preliminary results indicate that transposition in prokaryotes may sometimes proceed via an alternate route, namely via excision and reintegration (Berg, 1983; Bresler et al., 1983; Harayama et al., 1984;

145

PLANT TRANSPOSABLE ELEMENTS

N. Kleckner, personal communication). By the same token it is possible that more than one mechanism of transposition also operates in plants. Finally, as discussed in Section III,G, phenomena such as colored spots on white tissue that appear phenotypically to have arisen by excision of a transposable element may in fact have some other molecular basis. It follows that reversion events of this kind may have nothing to do with transposition. Since transposition as detected by genetic means is a rare event, one must develop a strategy for picking out the few organisms in which transposition has occurred among perhaps thousands of others. It is in this aspect of selection that most of the cases of transposition reported so far differ. Several examples will be outlined below. 2 . Transposition of a Receptor Element To follow the movements of a transposable element it must possess some property by which it can be identified and mapped wherever it is in the genome. In the case of a receptor element this is difficult since these elements usually cannot elicit a recognizable phenotype at each and every site in the genome. Nevertheless, the first and historically most important case of transposition reported is that of the receptor element Ds in Zea mays described by McClintock (1945, 1946, 1950a,b, 1951a,b). In this instance she was able to exploit a unique property of some forms of this element, namely their ability to provoke chromosomal breaks wherever they are integrated provided Ac is present in the genome. A heterozygous, Ac-containing line was used that is heterozygous for a number of different markers. In this line Ds is at its standard position proximal to the wx locus on the short arm of one chromosome 9 homologue (Fig. 3). A

*

9 h

I

vI

I

C ShBz

I

wx

I v

?

11

centromere

Fig. 3. Standard position of Ds on chromosome 9 of Zea mays according to McClintock (1951a). Symbols below the line indicate wild-type alleles of genetic loci. Yg: yellow-green; mutants have yellow-green leaves rather than green ones. C : aleurone color; mutants have colorless kernels rather than purple ones. Sh: shrunken endosperm; kernels of mutants are collapsed; locus encodes the enzyme sucrose synthase. Bz: bronze; mutants have kernels with reddish-brown aleurone rather than purple. Wx: waxy; mutants have kernels with low amylose content. The arrow indicates transposition of Ds to another position on the chromosome.

146

PATRICIA NEVERS ET A L .

series of dominant genetic markers (c-Z, bz+,and wx+) are situated distal to the Ds element. On the other homologue of chromosome 9, the corresponding recessive markers (c+, bz-, and wx-)are present and Ds is missing. It should be noted that the c-Z mutation is dominant to wild type and causes kernels to be colorless, while c+ kernels are normally fully colored. When a break occurs at Ds in some cells, the whole set of dominant markers can be lost and the respective recessive ones unmasked in these cells and their progeny. This results in a variegated kernel with clones of tissue exhibiting the recessive phenotypes. The Dsinduced chromosomal breaks can also be observed by cytological means, and the position of these breaks was found to correspond well with the position of Ds determined by genetic means. Transposition of Ds to a more distal position on chromosome 9 was signaled when kernels appeared with new patterns of marker loss or other altered phenotypes. 3. Transposition of a Regulatory Element The position of an active regulatory element in the genome can be determined more readily than that of a receptor element because a regulatory element can theoretically produce transactive signals at almost any integration site in the genome. In combination with a standard tester allele containing an appropriate receptor element, these signals in turn lead to a characteristic phenotype that can be followed in classical genetic mapping experiments, provided the element does not transpose too frequently (Rhoades and Dempsey, 1983). When the element is stable enough for genetic mapping, the main problem in detecting transposition of a regulatory element is to identify suitable candidates in which mapping seems merited. In some instances transposition was found simply by chance. Starting with a maize line with the En(Spm)-induced mutable allele pg-m, for example, the element was subsequently found at the a locus (Peterson, 1961, 1970b). Similarly the elements Ds and Ac also appeared spontaneously at other loci such as u, a2, c, bz, and wx (McClintock, 1948, 1949, l950,1951a,b, 1952, 1953,1962,1963, 1964, 1965a). The appearance of the transposable element at a new locus was often concomitant with its loss at the former locus in these cases. Mutable alleles of other loci have also been found in pal-rec and pal-rec revertant lines of A. majus (B. J. Harrison, 1965, personal communication), in the perlutea-variabilis line of A. majus (Meyer et al., 1981) and in a revertant of an unstable mutant of P. hybrida (Doodeman et al., 1984b). However, on the basis of classical genetic data it is not possible to determine whether the same DNA entity is responsible for the new mutations as that which was present at the original mutable locus.

147

PLANT TRANSPOSABLE ELEMENTS

In other experiments a concerted effort was made to detect transposition of a particular element to specific loci. For example (Peterson, 1968), a maize line carrying En(Spm) at the a locus with the genetic constitution a2+a2+,c+c+,r+r+,a+a-m was crossed to a tester strain carrying alleles of all these loci in the homozygous recessive form. Most of the F1 hybrids were therefore a2+a2-,c+c - ,r+ r- ,a+a- , or a2+a2-,c+c- ,r+ r- ,a-ma - . Among these some individuals were found exhibiting instability at the a2 locus and having the constitution a2-mla2-. The instability could be attributed to the presence of En(Spm). Similar experiments were used to detect transposition of Ac to a and a2 (McClintock, 1953). McClintock (1956b) reported that transposition of Ac could be detected by selecting kernels with apparently recombinant phenotypes. One possible cross of this kind is shown in Fig. 4. In a few exceptional kernels with a recombinant phenotype, A c was found to have left its position on chromosome 9 and transposed to a new site. Transposition of Spm(En) in maize has been successfully detected by using lines with a variegated phenotype due to an autonomous element at a particular locus and examining derivatives with a stable, nonvariegated phenotype. In these derivatives the element can be shown to be missing at its original position but present at another (McClintock, 1954, 1955; Peterson, 1961, 1970b; Fowler and Peterson, 1974; Reddy and Peterson, 1976;

Parental lines

Genotype

c+ A c wx+ c+ - wx-

Phenotype

C+,W.x+ nonvariegated

Genotype

c+ Ac wx+ -

Phenotype

C-I - W X - DS C-I, W.x+ variegated

X

.1 Majority of P~ogenY

Recombinant phenotypes Note: c+: c-I: wx+:

wx-:

+

C-I - W X - DS C-I - W X - DS C-I,W x - nonvariegated c+ - wx- -

C-1 - W X - DS C-I,Wx - nonvariegated

Some C-I,Wx', nonvariegated and some C-I. W x - variegated kernels Colored kernels; Colorless kernels; dominant to c+ Full starch content in kernels Reduced starch content in kernels

Fig. 4. Diagrammatic representation of a cross in maize in which transposition of Ac was detected in apparently recombinant progeny (McClintock, 1956b). In the parental line heterozygous for A c and the wx locus, classical crossing over can be expected to occur between these two loci, generating recombinant C-I, Wx+ individuals with no Ac and C-I,Wx- plants with A c still linked to the wx locus. Closer examination of the latter class of recombinant progeny revealed that in 50% of them Ac was no longer at its original position in chromosome 9 but had moved to a new one.

148

PATRICIA NEVERS ET A L .

Nowick and Peterson, 1981). Very detailed studies of En(Spm) transposition were conducted in this manner (Peterson, 1970b; Nowick and Peterson, 1981) using lines with mutable En(Spm)-induced alleles of the a locus as starting material. In 25% of the nonvariegated derivatives En was recovered on the same chromosome as the a locus, indicating preference for transposition from one position to another on the same chromosome (Peterson, 1970a). Qualitatively similar results have been obtained for Mp(Ac) (Van Schaik and Brink, 1959; Orton, 1966; Greenblatt, 1984). In still other experiments dosage effects have been exploited to detect transposition (see Section V for detailed discussion of dosage effects). For example, using a line of maize carrying the receptor component of the Dt (Dotted) system at the a locus and a single regulator element elsewhere in the genome, exceptional kernels were isolated with a phenotype characteristic for two doses of the regulatory element, i.e., kernels with a high degree of spotting (Doerschug, 1976). In plants from these kernels two copies of the regulatory component were indeed found, one at the original position and the other at a second site. This was interpreted as evidence for transposition. Similar results were also obtained with Ac (McClintock, 1956b). 4. Transposition of Mp(Ac) in Detail At first glance the finding of increased numbers of a transposable element would appear to contradict the concept of transposition via excision and reintegration. But detailed studies by Brink, Greenblatt and their coworkers (Brink and Nilan, 1952; Greenblatt and Brink, 1962, 1963; Greenblatt, 1966, 1968, 1974, 1984; van Schaik and Brink, 1959) have shown that increased copy numbers of a transposable element can also be explained by a loss/gain mechanism of transposition. In these experiments transposition of the autonomous element Mp(Ac) (Modulator) was examined. A maize line was used that is heterozygous for the p locus, a locus involved in pericarp pigmentation. One homologue carries a stable recessive allele of p while the other bears the allele p-uu (Emerson, 1917) consisting of Mp integrated at the p locus. Kernels of this heterozygous line have characteristic sectors of fully red pericarp against a white background. The degree of variegation is termed “medium.” On some ears of such heterozygotes twinned sectors can be observed of approximately equal size, one with red kernels and the other with lightly variegated kernels. Twinned sectors are thought to be the products of two different daughter cells of a common progenitor cell. Fully red kernels arise by loss of Mp from the p-vv locus (Brink, 1958) while lightly variegated ones are characteristic of lines containing two copies of Mp (McClintock, 1948; Emerson, 1929; Brink and Nilan, 1952).

PLANT TRANSPOSABLE ELEMENTS

149

Upon closer examination it was found that plants from the lightly variegated kernels did indeed contain two copies of M p , one at the original position at the p locus and another elsewhere in the genome (Greenblatt and Brink, 1962, 1963). In 61 to 67% of the cases the second copy was present on the same chromosome as the first, implying a “homing” tendency in transposition (Greenblatt and Brink, 1962, 1963; van Schaik and Brink, 1959; Greenblatt, 1984). Plants grown from fully red kernels were found to have lost M p from the p-uu locus, restoring wild-type expression of the locus. The simplest interpretation of these results is that M p is excised from one chromatid and reinserted in another prior to division of the progenitor cell of the twinned sectors, whereby it frequently selects a site on the sister chromatid. More detailed analysis of the revertant kernels from twinned sectors led to a refinement of the concept of M p transposition outlined above. It was found that although some of the revertant plants were completely devoid of M p , others contained a copy of the element at another position in the genome (Greenblatt and Brink, 1962,1963; Greenblatt, 1966,1984). Moreover, in 12 out of 13 such revertants that harbored a cryptic copy of Mp the position of the element was identical to the position of the second copy in the lightly variegated sector (Greenblatt and Brink, 1962, 1963). This has been explained by assuming that Mp is excised from a replicated portion of the chromosome and reinserted in an unreplicated segment of the same chromsome or another (Greenblatt, 1968, 1984; Fincham and Sastry, 1974). Inspection of the model presented in Fig. 4 shows that this explanation is plausible for twinned sectors that contain M p in the revertant kernels. A series of other types of twinned and untwinned sectors have been observed. These are summarized in Fig. 5 together with the putative structures of their respective p-uu chromosomes. It can be seen that all of these cases except for untwinned, light-variegated sectors can be explained by excision of Mp from one chromatid followed by loss or reinsertion of the element at another position. It is also conceivable that some of these constellations, in particular that of an untwinned, light-variegated sector, arise from others by a gene conversion step. Alternatively one might invoke a replicative mode of transposition to explain the latter (Fedoroff, 1983). Greenblatt (1984) has presented an interpretation of M p transposition that contradicts some aspects of the model dicussed above. His main premise is that transposition of M p occurs during replication of the p-uu chromosome and that M p can only integrate into unreplicated portions of a chromosome. The latter hypothesis is based on the fact that he finds no insertions of M p in an interval about 4 map units long immediately proximal to the p locus, while the interval immediately distal to p is a preferred

150

PATRICIA NEVERS E T A L .

t

Mpst P locus

Medium-Variegated

LightVariegated

Red

LightVariegated

Red

MediumVariegated

Red

MediumVariegated

Red

TWINNED SECTORS

TWINNED SECTORS

UNTWINNED R E D SECTOR

UNTWINNED R E D SECTOR

IMp present in red sector1

IMp missing in red sectorl

IMp missing in red sectorl

IMp present in red sector)

MediumLlghtVariegated Variegated

UNTWINNED LIGHT VARIEGATED SECTOR

Fig. 5 . A model of transposition of the M p ( = A c ) element in Zea mays. In the figure only one homologue of chromosome 1 is shown. This chromosome contains the transposable element M p (designated by a diamond-shaped symbol) integrated at the p locus, which is involved in pericarp pigmentation. The second homologue of chromosome 1 is not shown, but is assumed to carry a stable, recessive allele of the p locus. When the Mp-bearing homologue of chromosome 1 replicates, two chromatids are formed that enter two different daughter cells after cell division during development of the cob. These daughter cells may give rise to sectors of kernels that differ genotypically and phenotypically from the rest of the plant, as shown in the bottom line of the figure. Transposition of M p seems to occur after the replication fork has passed over the p-uu locus. Theoretically M p can be envisioned as transposing any time between replication of p-uu and the next round of cell division. It may move to an unreplicated site on the same chromosome, resulting in the first kind of twinned sector shown on the left. Or it may select a site on a replicated portion of the same chromatid or on the sister chromatid. Finally it may transpose to another chromosome or be lost entirely. Further theoretical considerations are required to explain an untwinned, lightvariegated sector. Here a different mode of transposition may be involved, or gene conversion or mitotic crossing over may have taken place subsequent to an initial transposition event.

region for integration. H e has interpreted this observation in terms of a replication fork moving in a proximal to distal direction with respect t o p . In keeping with this hypothesis Greenblatt (1968, 1974, 1984) has proposed other explanations for the origin of untwinned sectors and for twinned sectors in which Mp is missing in revertant kernels. He maintains that untwinned sectors arise when one of the original two daughter cells is lost

PLANT TRANSPOSABLE ELEMENTS

151

or forms cob tissue rather than kernels. To explain twinned sectors apparently lacking Mp in revertant kernels he has suggested that the element is present but in an inactive state.

5 . Transposition in Plants Other than Zea mays Evidence for transposition in other plants is minimal. Results from classical genetic experiments that support the notion of transposition in A . majus have already been described in Section II,D and molecular evidence will be presented in Section II1,D.

111. TRANSPOSABLE ELEMENTS AT A MOLECULAR LEVEL A. CLONING STRATEGY

With the dawn of gene technology in the 1970s it became possible to isolate as physical entities the elements McClintock, Peterson, and others had predicted as the source of instability of mutable alleles. The initial strategy employed is a straightforward one. What is required is a locus which encodes a known protein product and is thought to contain a transposable element in a given mutant line. With the help of antibodies against the gene products of the locus and appropriate cDNA probes, the genomic sequence of the locus can be cloned from wild-type plants. This in turn can be used to isolate the corresponding sequence from the mutant. Most of the elements which have been cloned were isolated in this manner and are summarized in Table V. Once a particular transposable element has been cloned, it can be used to isolate other elements of the same kind at different positions in the genome by virtue of sequence homology. This procedure allows isolation of loci that might otherwise be inaccessible by the straightforward cloning techniques described above. A further refinement can be achieved by establishing schemes to directly mutagenize a particular locus with a specific transposable element for which a molecular probe exists. This concept of “tagging” a locus with a transposable element has been practiced in prokaryotes for some time and similar attempts are being made in plant molecular genetics. Thus isolation of the Ac element Ac9 at the Acwz-m9 locus in maize was exploited to isolate the bz (bronze) locus via a mutant (bz-m2)harboring Ac at this second position (Fedoroff et al., 1984). There are hopes that other elements can be applied in the same way. The efficiency of transposon tagging obviously depends on a number of factors. If mutants bearing a known transposable element at the locus in question do not exist, then crosses must be designed to isolate new mu-

TABLE V Cloned Plant Transposable Elements and Other Insertion Elements Plant

Locus

Symbol

Protein

Allele

Element

Reference

Antirrhinum majus

nivea

niv

Chalcone synthase

niv-53

Tam1

Bonas et a/. (1984a)

Glycine max Zea mays

lectin I alcohol dehydrogenase-1

lel adhl

Seed lectin Alcohol dehydrogenase

niv-43 /el adhl-Frn335

Tam2 Tgml Dsl

K. Upadhyaya (personal communication) Goldberg et a / . (1983) Sutton et a/. (1984)

adhl-2Fl1 adhl -S3034 bz-m2

Ds M uI Ac

A. Merckelbach (personal communication) Bennetzen et al. (1984) Fedoroff et al. (1984)

sh-m5933 sh-m6233 sh-5586 wx-m6

Ds5933 Ds Tz86 Ds6

Geiser er al. (1982) Weck et al. (1984) S . Dellaporta (personal communication) Fedoroff et al. (1983a)

wx-m7 wx-in8 wx-m9 Acwx-m9

Ac Spm-I8 Ds9 Ac9 Cinl-001

Behrens et a / . (1984) Schwarz-Sommer et a /. (1984) Fedoroff et ul. (1983a) Fedoroff er al. (1983a) Shepherd et a/. (1982)

bronze

bz

shrunken

sh

waxy

wx

line CI

Icl

UDP glucose :flavonoid Glucosyl transferase Sucrose synthase

UDP-glucose-starch glucosy ltransferase

Unknown

nfl

PLANT TRANSPOSABLE ELEMENTS

153

tants containing appropriate transposable elements. Here it is convenient to work with an element that can be readily identified by classical genetic means such as one that belongs to a two-component system. This element must, of course, be able to transpose to the locus of interest at a reasonably high frequency. Alternatively, one may be able to select an element that is preferentially mobilized under certain inducing conditions. The Mu1 element of Zea mays may be of this type (Freeling, 1984). Another important factor is the number of copies of the marker element present in the genome from which the tagged locus is to be isolated (see Section 11,C): the higher the copy number, the more sorting out is required to identify the right locus. The isolation of transposable elements in plants as real, physical units has opened up a wealth of new perspectives on transposition and provided a new set of criteria for defining transposable elements, some of which will be discussed in the following sections. B . STRUCTURAL FEATURES OF PLANT TRANSPOSABLE ELEMENTS

Most plant transposable elements exhibit two structural features which have been shown to be typical for transposable elements in prokaryotes and animals. That is, the element possesses inverted repeat sequences at its termini, and generates a duplication at the target site upon integration (Fig. 6 and Table VI). In prokaryotes the lengths of both the terminal inverted repeat sequences and the duplication are characteristic for a given family of elements (Calos and Miller, 1980; Kleckner, 1981). Although sequence data on plant transposable elements are still too limited to make the same generalization with certainty, families of elements having similar physical characteristics do seem to exist. The elements are grouped in Table VI according to the size of the target site duplication. It is immediately apparent that all members of the Ac-Ds system make an 8bp duplication of the target site and have a terminal inverted repeat of 11 bp. Four of the Ds elements for which sequence data are available contain the same 11-bp sequence in inverse orientation at their very ends, while Ds9 isolated from the wx-m9 allele has an altered base that was also found in Ac-9 in the Acwx-m9 progenitor allele (see Fig. 6). Both Ac elements present at the wx locus in the mutants wx-m7 and Acwx-m9 have a terminal G nucleotide instead of the terminal A found in most Ds elements. The significance of the termini of a transposable element for excision, transposition, and other activities is a well-established fact for prokaryotic elements (for review see Kleckner, 1981). Therefore it is not surprising that related elements like Ac and Ds from the same plant species

@ A A A A C A A C A T C A

c,

A - 1 A - 1

- 1 - 1 - 1 - 1 - G - T - T - G - 1 - A - G - 1 -

A - 1 A - 1

A A A C A A C A

G

0 . .

...

+c

4

- 1 - 1 - 1 - G - 1 - 1 - G - 1 1 -A C - G A - 1 -G+

0 . .

A A A A G A A C

A - 1

A - 1 G G A - 1

1 -A 1 -A A - 1

1 -A C - G A - 1 0 . .

0 . .

1 1 1 1

-C - 1 - 1 - G

A - T 1 - A

...

- G+

--&

-

C --G A - 1

...

-c

--G0 . .

c3

(3 A - 1

A - T

A - 1 A - 1 G -C 1 - A A - T G - C G - C G - C A - 1

A - 1 A - 1

G -C 1 - A A - 1 G - C G - C G - C A - 1

1 - A A - 1

G-C G 1 1 G

-

C A A C

A - 1

1 - A A - 1 G - C A - 1 +G C

........ ........ ........ ........ ..... ..... ......... - ......

->l

>pG

-l-A-

+ l - A

D

0 . .

Fig. 6. The termini of plant DNA elements. A single DNA strand of each element is shown (5'+3'), and the terminal inverted repeat sequences are paired. Dots below the figure indicate the number of bases present in the flanking target site duplication. The sequence data shown here were derived from references listed in Table VI. Ds9 (a derivative of Ac9) in the wx-m9 allele of maize retains the sequence shown here for an A c element.

155

PLANT TRANSPOSABLE ELEMENTS

TABLE VI Physical Characteristics of rhe Cloned Elements

Elements

Allele

Insert size (kb)

Target site duplication (bp)

Terminal inverted Repeat (bp)”

Tam1

niv-53

17

3

13*

Tam2

niv-43

5

3

14*

Tgml Spm-I8

/el wx-m8

3.4 2

3 3

35 13*

Cinl-001

nfl

0.691

56

A c9 Ac

A c wx-m9

wx-m7

4.563‘ 4.5

8 8

11 11

Ac Dsl Ds

bz-m2 adhl -Fm335 adhl-2F1I

4.3 0.405 1.3

8 8

lI* Il*

Ds5933 Ds Ds6 Ds9

sh-d933 sh-m6233 wx-mi5 wx-m9

8 8

11*

Mu1

adhl-S3034

1.376

Tz86

s h-5586

3.6

30 (Ds=2 kb)“ 4 (Ds=2 kb)d 2 4.369

8’

6*

II* I I C

Reference Saedler ef al. (1983) Bonas et al. (1984b) K . Upadhyaya (personal communication) Vodkin et al. (1983) Schwarz-Sommer er a/. ( 1984) Shepherd e t a / . (1984) Pohlman et a / . (1984) M. Muller-Neumann (personal communication) Fedoroff et a/. (1984) Sachs et al. (1983) Doring et a/. (1984) A. Merckelbach (personal communication) Doring ef a/. (1984) Weck et a/. (1984) Fedoroff et a / . (1983a) Pohlman et al. (1984)

9

215

Bennetzen er al. (1984) J . L. Bennetzen (personal communication)

10

None

S. Dellaporta (personal communication)

A * indicates a “perfect” inverted repeat. The Cinl-001 element is not immediately Ranked by a direct repeat; however, two other Cinl elements were found to have a 5-bp direct repeat immediately Ranking the element. It has not been shown that the direct repeats are a result of element integration. The exact size of the Ac9 element was determined by DNA sequence analysis. The size of the other two cloned Ac elements is thought to be the same (or very similar) to that of Ac9; however, DNA sequence analysis has not been completed. The size of a single Ds unit is thought to be approximately 2 kb although the size of the DNA insert is larger (see text). In Pohlman er al. (1984) the DNA sequence of only one end of the Ds9 element was determined. The DNA sequence of this end was found to be identical to that of the progenitor element, Ac9 (i.e., differing from all other Ds element termini: see Fig. 6).

156

PATRICIA NEVERS ET A L .

share terminal sequence homology. However, a very unexpected observation is that elements from several different plant species all share a common sequence at their termini. The Tgml element from soybean, the Taml and Tam2 elements from A . majus, and the Spm-I8 receptor element from maize all end in the sequence CACTA. Furthermore all four elements generate a 3-bp duplication of the target site. One explanation for this phenomenon might be convergent evolution of these elements in all three species. Alternatively it may mean that these elements all derived from a common progenitor early in evolution or that horizontal spread of a progenitor element occurred at a later stage in evolution. If and when interspecific transformation systems are available, it may be possible to determine whether the elements from different species can interact functionally. The termini of many plant transposable elements exhibit even more extensive inverted repeat homology, as demonstrated for the elements Taml, Tam2, and Spm-I8 in Fig. 7 . By pairing the regions of homology in the ends of one DNA strand of the element, elaborate stem-and-loop structures can be formed that may be of significance for the mechanism of transposition (see Section IV). The maize element Ac also exhibits a structure of this kind (Courage et al., 1984). Although terminal inverted repeats and generation of a target duplication seem to be characteristic of transposable elements, this observation does not necessarily mean that every DNA sequence with these structural idiosyncracies can transpose. Two elements have been isolated that are structurally similar to transposons but cause stable mutations and thus may be defective mobile elements. These are the Tgml insertion in the Lel mutant of soybean (Goldberg e f al., 1983; Vodkin et al., 1983) and the Tam2 element of Antirrhinum majus (K. Upadhyaya, personal communication). There is no classical genetic evidence for transposition of these elements, but some molecular evidence in favor of transposition of Tam2like elements is presented in Section III,D. Another insertion sequence with structural characteristics of transposable elements is the Cinl elemenl of Zea mays (Shepherd et al., 1982, Fig. 7. Structural characteristics of the termini of the elements Taml and Tam2 from A. mujus and Spm-Z8 from Z . mays. The stem-and-loop structures shown are formed by pairing stretches of inverted repeat sequence homology within the termini of one strand of each element. Arrows on the right indicate regions of homology shared by Tam1 and Tam2; those on the left indicate homologous sequences common to all three elements. The two different kinds of arrows represent two different stretches of sequence homology. The target site duplication is illustrated by the boxed-in sequences at the base of each element. Data from Bonas er al. (1984b), Schwarz-Sommer ef al. (1984), and K . Upadhyaya (personal communication).

Spm - 18 of Z,mays

\

/

\

/

C

G

T;

c

G A

C

A

A

C T C T

G A A T / I5 \ \

/

T

A\ \ 51

1 /

T

If C T

C

/ c

A

G A

G T

G T

Tam2 of A. maim

C

G\

\

2

G

/ \

'A T T

/

A

T

I

p

G A G T

G

Ti.

T \

/

:23

14

G A

G

G A A

A

G

I I '

C

I

A

'

T

A

; c

C C

,

T T A T

T

C

C

r:

T T

A C

T T T 1 C T T G T A

G I G

/

lGTTJ

C

G \

/ A

41

I \

A

A I I

A A

' '

A C

2 : c I

A

I

C

I

A

I r

G

T T

T A A T

G\

r \

:35 \

31;

I

\

/

\

' A A

r; T

A T

I \

C

C

G

I A

I

\

\ 51

A A G A

63)

180 \

Ji

/

G

/ \

A

G'

I 15 \ .;\'A I I I

I

I /

\

C

/c

I \ I

G

A

\

/

C A T T

C C

9

4; C

A A

T'

/

A A

A

/

\

\

\

101

~

/

\

\

\

1

\

\

\

/

1 \

Tam 1 of A. majus

G

C

C T \

\

75) /

158

PATRICIA NEVERS ET A L .

1984). Here the terminal 5 bp are identical to those of the copia transposable element of Drosophila melanogaster (Lewis et al., 1980). It also contains the dinucleotides TG-CA at its very ends (Shepherd et al., 1983),which resemble those of such elements as copia, the Tyl element of yeast, the majority of retrovirus proviruses (for reviews see Shapiro, 1983), and mouse IAP genes (Cuff et al., 1983). The size and internal structure of Cinl are similar to the flanking LTR (long terminal repeat) structures of the elements listed above (Shepherd et al., 1984). However, for Cinl there is no evidence for actual transposition except perhaps on an evolutionary scale (Shepherd et al., 1982, 1984). The structural features discussed above may not be characteristic for all plant transposable elements. A notable exception is the maize Tz86 element isolated from the unstable sh mutant sh-5586. This element apparently has no terminal inverted repeat sequences (S. Dellaporta, personal communication). In contrast to the apparently conserved terminal sequences of transposable elements, the internal sequences seem to be subject to much more variation. Several different categories of DNA insertions, ranging in size from 0.4 to 30 kb, have been described, each of which behaves genetically as a Ds receptor element capable of being excised when functions are provided in trans by an active Ac regulatory component (see Table VI). Although all of these elements share a terminal 1 l-bp repeat that is quite similar, DNA sequence analysis of Dsl (Sutton et al., 1984) and Ds5933 (Doring et al., 1984a) revealed that they share very little sequence homology and do not hybridize to each other. However, hybridization experiments and/or restriction analysis have shown homology between Ds5933 and the other cloned Ds elements (Doring et al., 1984a; Weck et al., 1984; Fedoroff et al., 1983b; Pohlman et al., 1984). Thus it is possible that at least two different classes of Ds elements exist that structurally have only the 1 1-bp terminal repeats in common. It follows that the 1l-bp sequence may be all that is required as a substrate for excision and transposition in response to the mutator function of A c . For the expression of all the complex functions of a regulatory component such as Ac, the internal sequences are more critical. For example, a small, internal 200-bp deletion within the 4.5-kb Ac element of Acwx-m9 mutants can eliminate its regulatory capacity and convert it to an element that behaves genetically like a passive receptor element, namely Ds9 (Fedoroff et al., 1983a; Pohlman et al., 1984). On the basis of the sequence homology outlined above it would seem that Ds6, Ds9, Ds5933, and the Ds elements in sh-m6233 and adhl-2F11 may all represent deletion derivatives of Ac and therefore may be considered defective regulatory elements. This is in keeping with the origin of two-component sys-

PLANT TRANSPOSABLE ELEMENTS

159

tems dicussed in Section I1,B. The 405-bp element D s l , on the other hand, does not follow suit. Its origin is presently unclear. It is possible that some plant transposable elements include DNA sequences that play no part in transposition and other element-induced activities. One such candidate may be the unusually large 30-kb insertion at the maize sh locus in the mutant sh-m.5933 that is flanked by compound Ds elements (Geisir et al., 1982; Fedoroff et al., 1983b,c; Courage-Tebbe et a f . , 1983; Doring et a f . , 1984a). Other possible candidates are the large Ds insertions described by Burr and Burr (1981b, 1982). C. MULTIPLE COPIES OF PLANT TRANSPOSABLE ELEMENTS

Southern hybridization analysis has revealed that in plants multiple copies of various different transposable elements are present as cryptic components of the genome (Geiser et al., 1982; Fedoroff et al., 1983b; Peacock et a f . , 1983; Sutton et al., 1984; Schwarz-Sommer et a f., 1984; Bonas, 1984). The banding patterns on which copy number estimates are based vary depending upon the stringency of the hybridization experiments, the particular line being examined, and which part of the element is used to probe the genome. The choice of probe in turn depends upon the complexity of the element being investigated. To look for functionally, relatively simple receptor elements, where only the termini may be of importance, as discussed in Section III,B, a terminal fragment of a larger element or the entire sequence of a smaller element may be appropriate as a probe. However, since the distinguishing feature of an autonomous element is its ability to produce transactive signals, and since these are most likely encoded in more internal sequences of the element, a fragment from this region lacking the terminal sequences is normally used as a probe for autonomous elements. In a commonly used assay procedure the genomic DNA is cut with an enzyme that generates one fragment homologous to the probe per copy of the element. This genomic fragment must also contain sequences adjacent to the element. Thus each fragment that hybridizes to the probe represents one copy of the element at a unique position in the genome. Alternatively the genomic DNA can be cut with an enzyme that releases an internal fragment of the element in question and then probed with the same internal fragment. The copy number is then estimated on the basis of the intensity of the hybridization signals (Bennetzen, 1984). By these means 30 or more copies of both Ds and Spm-Z8 were found in the maize genome (Geiser et a f . , 1982; Peacock et a f., 1983; Schwarz-Sommer et al., 1984). Similarly 10 to 50 copies of the 1.4-kb Mu1 element were found in a maize mutator line (Bennetzen, 1984; see Section VI for details on mutators). In contrast, only 4-10 sequences

160

PATRICIA NEVERS ET AL.

homologous to internal sequences of the regulator element Ac responsible for the wx-m9 mutation of maize occur elsewhere in the Acwx-m9 genome (Fedoroff et al., 1983b). Likewise, sequences homologous to internal parts of the Antirrhinum elements Taml and Tam2 are found, respectively, four to seven times in the genome (Bonas, 1984; K. Upadhyaya, personal communication). As discussed in Section III,B, these multiple copies may represent a whole array of deleted or otherwise altered derivatives of a standard fulllength element. This is supported by evidence from Fedoroff et al. (1983b), who find that the 4 to 10 elements homologous to an internal portion of Ac and present as cryptic elements in the Acwx-m9 genome are of varying lengths. Similar results have been found for the Antirrhinum element Taml (Bonas, 1984) and for Mu1 from maize (Bennetzen, 1984). The Cinl DNA insertion element of Zea mays has structural characteristics of transposable elements (Shepherd et al., 1984) and belongs to the middle repetitive class of DNA from this organism (Gupta et al., 1984). Full-length Cinl elements as well as altered forms were found dispersed in the maize genome (Gupta et al., 1983). This supports the notion that dispersed repetitive sequences in general consist of elements that once were or still are transposable, as has been proposed by several authors (Britten and Davidson, 1976; d’Eustachio and Ruddle, 1983; Flavell et al., 1981; Freeling, 1984). D . DETECTION OF TRANSPOSITION AT THE MOLECULAR LEVEL

Since more sophisticated in uitro assays for transposition in plants have not yet been developed, the only molecular evidence for transposition (as opposed to excision) is derived from Southern blot analysis. In Section II1,C the procedure for determining copy numbers of elements was outlined. If the pattern of hybridization signals obtained in this manner differs between different plants, several interpretations are possible. The most obvious explanation is that sequence polymorphism has led to divergent restriction sites (although this is unlikely between sibling plants). Alternatively sequence alterations within a transposable element or in its immediate vicinity may result in altered hybridization signals. Finally these differences in banding pattern may also mean that some copies of the element in question have changed position. On the basis of altered Taml hybridization signals between sibling plants of the niu-53 mutant of A . majus, Taml-like elements are considered to be transposable (Bonas, 1984). Similarly altered banding patterns in different lines of A . majus obtained with a Tam2 probe suggest that

PLANT TRANSPOSABLE ELEMENTS

161

Tam2-like elements are also transposable (E. Krebbers and K. Upadhyaya, personal communication). It is not certain, however, whether the Tam2 copy originally cloned at the nivea locus is itself mobile. Recent results (E. Krebbers, personal communication) indicate that this particular copy of Tam2 can be excised under certain circumstances, so it is possible that is also transposes. For elements of a higher copy number (e.g., Cinl) or elements containing middle to highly repetitive DNA sequences (e.g., Tgml) the use of Southern hybridization experiments to detect transposition is more difficult. It should be noted that while changes in hybridization signals in appropriate Southern blots may provide positive evidence for instability associated with transposable elements, the opposite observation, namely constant hybridization patterns, cannot be so readily interpreted. Here the elements may be potentially transposable but stabilized due to some kind of epistatic control (e.g., high levels of repressor or low levels of activator). E. HOW A TRANSPOSABLE ELEMENT INFLUENCES GENE EXPRESSION

Integration of a piece of foreign DNA in or at a locus can theoretically influence the expression of the locus at a number of different levels. The simplest possibility is that the element itself contains no sequences of significance for transcription or translation and influences expression only by virtue of its length and position. The importance of the position of integration has been emphasized by Peterson (1976a, 1977), and varying insertional sites have already been discussed as a possible source of allelic variants of mutable alleles (Section 11,F). However, as documented by the few elements that have been sequenced so far, most transposable elements probably contain splicing sequences as well as transcriptional and translational start and stop signals. These additional parameters increase the complexity of the means by which a transposable element can influence the expression of a locus at which it resides. Finally protein products of transposable elements may directly regulate expression of loci controlled by related elements. A number of the plant transposable elements that have been isolated have been found to be integrated within the 5' control region of a locus. These include Tam1 and Tam3 at the nivea locus in Antirrhinum majus (Bonas et al., 1984a; H. Sommer, personal communication), Ds in the adhl-Fm335 mutant of Zea mays (Sachs et al., 1983; Peacock et al., 1983), and Ac in the wx-m7 mutant of maize (Behrens et al., 1984;

162

PATRICIA NEVERS ET AL.

Schwarz-Sommer et al., 1984). Considering the complexity of eukaryotic control regions, including TATA, CAAT, or AGGA sequences as well as possible binding sites for regulatory proteins (Proudfoot, 1979; Benoist et al., 1980; Messing et al., 1983), the mere presence of a piece of foreign DNA can be envisioned as having serious effects on expression. For the Dsl element in adhl-Fm335, located between the promoter and the first exon of the adhl locus, there is some indication that the element interferes with expression more directly. Here the decisive piece of evidence is that the ADH protein of the mutant is more thermolabile than its wildtype counterpart but of normal size (Osterman and Schwartz, 1981). The following explanation for this observation has been proposed (Sachs et al., 1983; Peacock et al., 1983): Sequence analysis revealed that the 405bp element contains six ATG translational start signals, each of which is quite closely followed by a translational stop codon. Thus it can be ruled out that translation simply starts within Ds and continues into the locus, leading to an enlarged fusion product. This complies with the observation that both mRNA and protein are of approximately normal size in the mutant (Osterman and Schwartz, 1981; Sachs et al., 1983; Peacock et al., 1983). However, at 10 and at 15 codons beyond the first ATG in the element, suitable intron donor sequences can be found. If translation starts at the first ATG in Ds, then sequences between such an intron donor site and the distal end of the first intervening sequence may be removed by splicing. This would result in a slightly shorter message, which has indeed been found (Sachs et al., 1983; Peacock et al., 1983). A protein would be synthesized that contains an altered N-terminus, which in turn might account for its thermolability. Altered proteins have been reported for a number of other mutants harboring transposable elements at the locus in question. These include the Ds-controlled alleles sh2-ml (Hannah and Nelson, 1976), sh-m6258, and sh-m6795 (Fedoroff et al., 1983b,c), wx-B3 (Echt and Schwartz, 1981) and bz-wm, a derivative of bz-m2 (Dooner and Nelson, 1977); bz-m13 (Klein and Nelson, 1983) harboring the Spm receptor component (Z,rSpm), and A c wx-m9 (Schwartz and Echt, 1982; Shure et al., 1983). Since none of the mutants contains an element in the promoter region of the locus, explanations other than the one described above are required. More detailed protein and DNA sequence analyses are necessary before any final conclusions can be drawn. In addition to control regions, both exons and introns may be sites for integration of transposable elements. Integration within an exon would be expected to have direct and drastic effects on expression of the locus. In this case the inserted element could cause a shift in reading frame or, if it

PLANT TRANSPOSABLE ELEMENTS

163

remains in frame, lead to a protein with additional amino acids. The Ds element responsible for the adhl-2F11 allele is located in the fourth exon of the adh gene (Doring et al., 1984b; Dennis et al., 1984; A. Merckelbach, personal communication). Likewise the Spm-I8 receptor element responsible for the wx-m8 mutation is also located within an exon (Zs. SchwarzSommer, personal communication). Several other cases have been reported in which integration within an intron could be demonstrated. These include Tam2 at the niuea locus in A . majus (K. Upadhyaya, personal communication), Ds5933 at the maize sh locus (Courage-Tebbe et al., 1983), the Ds responsible for the maize allele sh-m6233 (Weck et al., 1984), the Ds elements in the mutants sh-m6258 and sh-m6795 (Sheldon et al., 1984), and Mu1 in the maize adhl-S3034 mutant and its derivatives (Strommer el al., 1982; Bennetzen et al., 1984). In adhl-S3034 and two derivatives of this allele, normal protein is produced but at lower levels, namely 40, 13, and 0% (Stommer et al., 1982). Tam2 is integrated within the consensus splicing site between the first exon and intron (H. Sommer, personal communication), which could explain its mutational effect. Mere insertion of foreign DNA within an intron should not be expected to interfere with expression of the locus. The fact that the elements listed above do indeed affect expression suggests that they either contain disruptive signal sequences or have caused sequence disorders in their immediate vicinity. This is further supported by results obtained with the mutants sh-m6258 and sh-m6795 (Fedoroff et al., 1983b), where shorter than wild-type messages were found, as if transcription terminated in or at the Ds elements responsible for these alleles. In some instances insertion of a transposable element at a locus may have no phenotypically visible effect on expression. Examples of this kind include derivatives of the maize mutant a-ml (McClintock, 1956a, 1965a, 1968) and the mutant bz-m13 (Klein and Nelson, 1983), each of which bears a receptor element of the Spm(En) system at the mutated locus, as well as the maize mutant R-r#2, which contains the receptor element of the Spfsystem (Gonella and Peterson, 1978). Provided that a suitable regulatory element is not present elsewhere in the genome, expression of the controlled loci is normal in these mutants, and they produce almost fully colored kernels. Only under the influence of the appropriate regulatory component can effects of the receptor elements on expression be seen. The suppressor function then reduces basal expression of the loci to zero while the mutator function mediates reversion of the mutant loci to the wild-type condition in somatic and germinal tissue. More subtle effects on expression can be expected as the result of interplay between a given transposable element and “modifying factors,”

164

PATRICIA NEVERS ET A L .

which may or may not be related elements. Examples of such interactions are discussed in Section V. F. PROTEINS ENCODED BY TRANSPOSABLE ELEMENTS

In prokaryotes many transposons and insertion elements encode one or more proteins required for the various activities of the element, and the corresponding DNA sequences in the element are actively transcribed. Proteins known to be encoded by prokaryotic transposons include a “transposase,” involved in transposition, and a repressor of the transposase gene. In the case of the transposon Tn3 the repressor also functions as a “resolvase,” an enzyme required for a special recombinational step characteristic of the transposition pathway of certain transposons (for reviews see Heffron, 1983; Iida et al., 1983; Kleckner, 1983). In plants evidence for proteins encoded by transposable elements is mostly of classical genetic nature. As discussed in Section II,B, at least three different functions of the maize Spm regulator element have been hypothesized: a mutator, a suppressor, and an activator function. Although the mutator and suppressor activities can mutate independently (McClintock, 1965a, 1968), it is unclear whether these represent activities of separate proteins or of a single protein. Molecular data concerning proteins encoded by transposable elements in plants are scanty at present, but will certainly be available in a very short time. Contributions can be expected from sequence data analyzed for open reading frames. Indeed, an open reading frame corresponding to 203 amino acids has been found in Ds5933 (Doring et al., 1984a). The Ac element in the mutant Ac wx-m9 is reported to contain two open reading frames encoding polypeptides of 839 and 210 amino acids (Pohlmann et al., 1984). The larger open reading frame is thought to correspond to the structural gene of a function required for transposition since a mutation in this sequence converts Ac to a Ds element (Fedoroff et al., 1983a; Pohlmann et al., 1984). Conflicting results have been found for the Ac’s in Ac wx-m7 and Ac wx-m9 (M. Muller-Neumann and P. Starlinger, personal communication). These results indicate that each Ac contains two smaller open reading frames separated by approximately 150 bp instead of a single larger one. Four different open reading frames, two on each strand, have been found in the element Mu1 (J. Bennetzen, personal communication). A.nalysis of transcripts from transposable elements using Northern hybridization techniques and in uitro and in uiuo expression systems will provide further information. Preliminary results suggest that transcripts from En(Spm) (Zs. Schwarz-Sommer, personal communication) and Taml (R. Piotrowiak, personal communication) have been identified.

PLANT TRANSPOSABLE ELEMENTS

165

Fig. 8. Southern hybridization analysis of somatic reversion events in the niu-53 mutant of A. mujus. The niu-53 mutant harbors the element Tam1 at the niuea locus. Genomic DNA was digested with EcoRI, which cleaves twice within the sequence of Taml. A 32P-labeled fragment which contains most of the sequence of the chalcone synthase gene was used as a probe. Lane 1: Niu+ DNA; lane 2: DNA from niu-53 flowers; lane 3: DNA from niu-5.3 leaves. Note that the 5.7-kb fragment characteristic of Niu+ DNA can also be generated with DNA from the mutant, indicating that the mutant contains Niu+ tissue in which Tam1 has been excised. Data taken from Bonas ef al. (1984a). G . REVERSION EVENTS AT THE MOLECULAR LEVEL

When a specific probe is available, somatic reversion events can be analyzed with the help of Southern hybridization procedures (Wienand et al., 1982; Bonas et al., 1984a; Schwarz-Sommer et al., 1984). For example, when wild-type DNA of Antirrhinum majus is cut with EcoRI, a 5.7-kb fragment hybridizes to a probe containing most of the niuea locus. In contrast, when DNA from the niu-53 mutant harboring Taml at the niuea locus is cut with EcoRI and hybridized to the same probe, two new bands, 4.4 and 4.7 kb in size, are seen. This is due to the fact that Taml contains two EcoRI cleavage sites so that digestion with this enzyme generates two unique junction fragments, each containing part of the niuea locus and one end of T a m l . However, in the same blot the 5.7-kb fragment characteristic for the wild-type locus can also be observed, as shown in Fig. 8. This 5.7-kb band derives from tissue in which Taml is no longer present at the niuea locus. Thus it appears that in this case somatic reversion occurs by means of excision of T a m l .

166

PATRICIA NEVERS ET A L .

The molecular events underlying germinal reversion can be examined by DNA sequence analysis of the restored locus in revertant plants. The DNA sequence in and around the target site of the transposable element in question is compared in the progenitor line, in the mutant and in revertants. The major stumbling point here may be to locate the real progenitor line of mutants that were isolated some time ago. In the cases examined so far, germinal reversion has been shown to arise by exicision of the element from the locus at which it resided in the mutant. Strangely enough, however, the excision events do not result in exact restoration of the original sequence of the locus. This would require removal of the entire element plus one copy of the target site duplication generated during integration. Instead, in the cases reported so far it seems that the duplication or part of it is retained after excision (Sachs et al., 1983; Bonas et al., 1984b; Pohlman et al., 1984; Weck et al., 1984). These examples are summarized in Table VII. Sometimes one or two base pairs adjacent to the duplication junction are missing. In other instances inversion of one or two base pairs has occurred, also in the region immediately adjacent to the duplication junction. The possible significance of these findings will be discussed in Section IV. TABLE VII DNA Sequence Analysis of Revertants" Allele

Reference

Wild type Mutant Revertant

niu+ niu-53 ::Tam1 niu-531

-ATA-ATA -ATAiTA-

Bonas et al. (1984)

Wild type Mutant Revertant Revertant Revertant Revertant

adhl+ adhl-Fm335 adhl-RVI adhl-RV2 adhl-RV3 adhl-RV4

-GGGACTGA-GGGACTGA GGGACTGA-GGGACTG~GGACTGA-GGGACTGZCGACTGA- G G G A C T G m ACTGA-GGGACTG~GGACTGA-

Sachs et al. (1983) Sutton et al. (1984)

Wild type Mutant Revertant

sh+ sh-m6233 sh-m6233 r1

-CTTGTCCC-CTTGTCCC CTTGTCCC-CTTGTC:CTTGTCCC-

Weck et al. (1984)

Wild type Mutant Revertant

wx+ Ac wx-m9 wx9-rl

-CATGGAGA-CATGGAGA CATGGAGA-CATGGAGA LTGGAGA-

W T n m ]ATAl

a

Pohlman et al. (1984)

The positions which are underlined in the DNA sequence of the revertants indicate that these positions differ from those present in the direct repeat flanking the element in the mutant. A dot indicates a deleted base pair.

PLANT TRANSPOSABLE ELEMENTS

167

This mode of excision has not been observed in either prokaryotes or animals and may be unique to plants. The fact that a slightly altered sequence remains after excision means that altered proteins may also be formed, providing the sequence is present within the transcriptional unit of the locus. Indeed, altered proteins have been observed in revertants of the following maize mutants: wx-ml (Echt and Schwartz, 1981), bz-m2 (Dooner and Nelson, 1979), sh2-ml (Tuschall and Hannah, 1982), and wxm6 (Shure et al., 1983). This opens up an entirely new perspective on how transposable elements may contribute toward genome variability and fitness of plants in evolution. While excision has been shown to be responsible to reversion in a number of instances, it cannot be concluded that this is the only means by which reversion occurs. It is conceivable, although not yet extensively documented, that offspring with a revertant phenotype can arise without excision of the element. Burr and Burr (1982) have reported that Ds is retained in a revertant of a mutable sh allele although in rearranged form. Reversion without excision has also been proposed for revertants of mutable alleles in Petunia hybrida (Bianchi et d . , 1978; Doodeman et d . , 1984a). Reversion events of this kind could result from mutations within the transcriptional and translational signal sequences of the element that are otherwise responsible for its mutational effect or within adjacent sequences of the target site. Alternatively, mutations at other loci might contribute toward wild-type or nearly wild-type expression of a locus harboring a transposable element. Suppressor mutations at secondary loci, as described for the gypsy element in Drosophila (Modolell et al., 1983) and the Tyl element in yeast (Fink et al., 1980), for example, may eliminate the inhibitory effects of the elements. H . T H E MOLECULAR BASIS FOR ALLELIC DERIVATIVES OF MUTABLE ALLELES

The possible origins of derivatives of mutable alleles with new, genetically heritable phenotypes was discussed in Section II,F. At the moment three approaches seem to be promising for examination of such phenomena at a molecular level. First, one can look for structural alterations in and around the transposable element at the locus in question. Second, alterations in expression of genes encoded by transposable elements can be investigated by analyzing the corresponding mRNAs or proteins. Finally, changes in copy number, size, and positions of related transposable elements can be examined by Southern hybridization. Molecular evidence of this kind is still sparse. Fedoroff et al. (1983a) and Pohlman et al. (1984) report that a 0.2-kb deletion within an open

168

PATRICIA NEVERS E T A L .

reading frame of A c results in the loss of its regulatory capacities and converts it to an element that behaves like Ds.This convincingly demonstrates how an autonomous element can become inactivated, but a deletion mutation cannot explain the cyclic changes in activity referred to in Section I1,F. Molecular information on the unstable maize mutant adhlS3034 and a derivative, adhl -S3034a, suggests how stable recessive alleles can arise from a mutable one. While integration of Mu1 at the adhl locus in adhl43034 still allows expression of mRNA and protein at a level 40% that of wild type, a small 74-bp deletion immediately adjacent to the element in adhl-S3034a eliminates expression completely by destroying the first exodintron junction (Strommer et al., 1982; L. Taylor and V. Walbot, personal communication). This is reminiscent of deletions induced by transposable elements in bacteria (Reif and Saedler, 1975). Further molecular evidence for changes in state concerns the ability of Ds to induce chromosomal breaks in the presence of Ac. The pertinent results derive from studies on the mutant sh-m5933 in Zea mays (Courage-Tebbe et al., 1983; Doring et al., 1984a). This mutant contains a complex 30-kb insertion within the sh locus that responds to Ac signals and thus behaves in its entirety as a receptor element. The sequence of the 3' end of the insertion relative to the sh locus consists of a 4-kb structure made up of one basic 2-kb Ds unit inserted in another such 2-kb sequence in inverse order. This is illustrated schematically in Fig. 9. Adjacent to the sh locus, at an unknown distance upstream from its 5' end, is a duplication of the 5' end of the locus including one double Ds structure. When the sh-m5933 mutant reverts to wild type, the 30-kb insertion is lost, but the duplication remains. Ds-induced chromsome breakage also persists in these revertants so that kernels of revertant plants have variegated pigmentation, as discussed in Section II,G. In one revertant, sh-r5, Ds-induced breakage in the endosperm is delayed, manifested phenotypically by fewer and smaller spots of color. In this revertant a 2.0-kb deletion was detected in the double Ds structure still present, suggesting that this is responsible for the altered timing of somatic variegation. Two different hypotheses have been advanced concerning the molecular basis of Ds-induced breakage (Weck et al., 1984; Doring and Starlinger, 1984). One notion is that Ds-induced breakage is due to pairing between inverted regions of double Ds structures on sister chromatids followed by crossing over within this paired section. This would result in a dicentric chromosome lacking the Ds-distal segment and an acentric chromsome containing a duplication of the Ds-distal segment. The latter would be lost during cell divison. How an A c function might mediate these events is unclear, but one possibility is that it enhances mitotic

c c

4 .

Sh revertants with continued

Ds breakage

*-c

Sh-r5 w i t h de\ayed Ds breakage

Fig. 9. Structure of the Ds-induced maize allele sh-m.5933 and revertant derivatives according to Courage-Tebbe et al. (1983) and Doring et al. (1984a). The drawing is merely a schematic representation of relevant structural features and is not drawn to scale. The sequence between the black box on the far right of the upper line and the middle black box represents the 30-kb insertion integrated within the sh locus of sh-m.5933. The 3’ end of the locus is to the right of this structure. To the left of the 30-kb structure is a duplication of the 5‘ end of the sh locus and part of the 30-kb insertion. The orientation of this duplication has not yet been determined. Duplicated sequences are indicated by bars above the line. The bracket designates the only segment of DNA for which complete sequence data are currently available. The hatched triangle indicates a basic 2-kb Ds sequence, which is inserted within another such sequence at the 3‘ end of the 30-kb insert. Part of this double Ds structure seems to be missing at the 5’ end of the insert. In most revertants of sh-m5933 (see the middle figure), the 30-kb insert is removed, but the duplication of the 5’ end is retained together with a partial double Ds structure at which breakage presumably continues to occur. In one revertant with an altered pattern of Ds-induced chromosomal breakage (see the lower figure), 2.0 kb of the truncated double Ds structure has been deleted, although it is not certain which part of this structure is missing. The small arrows below each double Ds structure indicate terminal inverted repeat sequences.

170

PATRICIA NEVERS ET AL.

crossing over. The second idea about how Ds breakage might occur also involves a double Ds structure. Since each 2-kb Ds in the double Ds structure contains terminal inverted repeat sequences, the composite structure includes two sets of direct repeats of the same sequences, as shown in Fig. 9. These may be recognized by an Ac function and cleaved as in excision, resulting in a chromosome break. In this case breakage and excision would be two facets of the same process, as has been proposed by McClintock (1949). Both mechanisms described here are based on a double Ds structure. It follows that simple Ds structures like Dsl (Sachs et al., 1983), the 1.3-kb Ds in adhl-2Fl1 (Doring et al., 1984b), and the 2kb Ds6 element (Weck et al., 1984) could not generate breakage in this manner. Furthermore, it seems that no chromosomal breaks or only very few occur at A c itself (McClintock, 1963). Therefore the Ds elements generated by deletions or other alterations in the A c sequence, as disucssed in Section III,B, should also fail to produce breaks, unless the alteration affects a sequence that otherwise prevents breakage at Ac. Elucidation of the ability of the various cloned Ds elements to produce breaks should help to clarify this question.

IV. A MODEL OF THE MECHANISM OF TRANSPOSITION A . OBSERVATIONS O N WHICH THE MODEL IS BASED

As discussed in Section II,G, there is evidence that transposition in plants occurs by means of excision of a transposable element and reinsertion of the same element at another position in the genome. This is contradictory to the mode of transposition thought to be the most prevalent mechanism in prokaryotes. Here transposition is believed to occur primarily by replication of a transposable element and integration of a copy of this element at a new site in the genome (for model see Shapiro, 1979; Grindley and Sherratt, 1978). Any model of transposition in plants must take the “cutand-paste” pathway into account. Structural features of transposable elements and their integration sites provided clues to the mechanism of transposition in prokaryotes, and, as mentioned in Section 111, plant transposable elements resemble those of other organisms in a number of structural properties. The terminal sequences of the elements of a given class of elements are almost completely identical. Furthermore, insertion of a given plant transposable element at a locus is accompanied by a duplication of the target site sequence that is of characteristic length (see Table VI). A striking difference between prokaryotic and plant elements is ob-

PLANT TRANSPOSABLE ELEME,NTS

171

served with respect to the product of excision. In plants, as opposed to prokaryotes, it appears that reversion to wild type is not accompanied by exact restoration of the wild-type DNA sequence. In all seven cases investigated to date the target site duplication is retained in an altered form, as discussed in Section II1,G and summarized in Table VII. The unusual thing about all of these anomalous revertant sequences is that the alterations are all found at the junction between the duplicated segments. Since this is the point at which the element was formerly attached to the sequence, it is plausible that the anomalies occurred during the process of excision. This peculiarity of reversion events in plants provided the basis for the model of excision and transposition discussed below.

B. EXCISION OF A TRANSPOSABLE ELEMENT IN PLANTS

By analogy with models of prokaryotic transposition (Grindley and Sherratt, 1978; Shapiro, 1979; Arthur and Sherratt, 1979; Galas and Chandler, 1981; Harshey and Bukhari, 1981) the key enzyme in excision of a plant transposable element is thought to be a specific transposase capable of identifying the ends of a particular class of transposable elements. This enzyme can be envisioned as recognizing the inverted repeat sequences of an element and drawing them together, which in turn would bring the duplicated target site sequences into close approximation. This is shown in Fig. 10a. Then the transposase sets staggered nicks at the 5’ ends of the duplicated target ‘sequence, which are a specific distance apart. By this process the element will be severed from the target site DNA and contain complementary, single-stranded copies of the duplication on each 5’ end. The same overlapping single-stranded sequences will be present in the DNA adjacent to the element. Let us now assume that the element is not released immediately but remains interjected between the single-stranded segments of the chromosome in a DNNprotein complex. While the free 3‘ ends of the element may not be accessible to polymerase at this point since they are buried in the complex, it is possible that the free 3‘ ends of the target sequence are. Starting at these ends polymerase begins to fill up the gaps in the duplicated sequence. At the same time a 5’ exonuclease repair enzyme starts nibbling off loose ends. Assuming that the 5’ fringes attached to the element protrude from the complex, they would be vulnerable and may be degraded quite extensively. Depending upon how efficient the exonuclease is, it may produce a “clean” element free of sticky ends, or one with one or two additional, complementary nucleotides at one or both termini of the element.

172

PATRICIA NEVERS ET A L .

As shown in Fig. 10a the free 5’ ends present in the chromosome may also be subject to exonuclease degradation. Removal of one or two bases at one of these ends would result in a shortened template for repair synthesis. After completion of repair synthesis and ligation of the chromosomal DNA, an imperfect duplication lacking one or two base pairs on either side of the ligation junction would arise. This would explain the sequences found for revertants of the niu-53 mutation of A . majus (Bonas e?al., 1984b), the sh-m6233 mutation of Zea mays (Weck et al., 1984), the maize Ac wx-m9 mutation (Pohlman et al., 1984), and for some of the revertants of the Ds-induced mutation adhl-Fm335 (Sachs e? al., 1983). An extension of the model is required to explain the inversion found at the duplication junction in some revertants of the adhl-Fm335 mutant. Here template switching may occur during repair synthesis in the transposase complex, as shown in Fig. 10a. As polymerase approaches the interjected element, it may switch from the chromosomal template sequence to the protruding single-stranded ends of the transposable element. Following correction of the original template sequence in this area or some other form of mismatch repair, an inversion one or more base pairs long would be formed at the duplication junction. C. INTEGRATION OF A PLANT TRANSPOSABLE ELEMENT

For simplification the first case considered is that of integration of a “clean” linear transposable element with no sticky 5’ ends. Once again it is assumed that a specific transposase forms a complex between the termini of the element and the target site sequence. Then this enzyme makes staggered nicks separated by a characteristic number of nucleotides (see Table VI) in the target sequence, generating complementary singlestranded fringes, as proposed in models of prokaryotic transposition. The element is inserted between these single-stranded recipient DNA sequences and ligated where possible. The gaps opposite the singlestranded sequence stretches are filled by repair synthesis followed by ligation. The result is an integrated element flanked by a duplication of the target site, as shown in Fig. lob. D . PREDICTIONS

The model as it has been presented suffices to explain the data on plant transposable elements that are currently available. More interesting are a number of predictions by which the validity of the model could be tested. Let us first consider other kinds of excision events that could be expected to arise by the process outlined above. For example, repair syn-

PLANT TRANSPOSABLE ELEMENTS

173

thesis of the target site sequence might be completed without interference by 5‘ exonuclease degradation of the template. A perfect duplication of the target site would then remain in the revertant after excision of the element. On the other hand, if exonuclease degradation of the singlestranded 5‘ chromosomal ends proceeds extremely rapidly and efficiently, one end and part or all of the other may be removed, resulting in a deletion at the original target site sequence. Removal of only one singlestranded chromosomal end by exonuclease degradation would allow precise restoration of the wild-type sequence. Whether or not a protein with wild-type function can be formed following such excision events depends upon the position of the integration site within the locus, possible changes in reading frame due to sequence alterations, and the consequences of loss or addition of amino acids at that point in the peptide chain. With respect to inverted base pairs at the duplication junction the model clearly predicts that when more than one base pair is involved the individual base pairs are not simply inverted in situ, but rather the entire segment. A real inversion should be observed instead of individual transversions. The one relevant case reported so far (Sachs et al., 1983) does not permit distinction since the two inverted base pairs are identical. Other predictions associated with the model concern integration events. Here it is a question of the extent to which the single-stranded 5’ sequences attached to the ends of the excised element are degraded prior to transposition. If they are not attacked at all, these “sticky ends” might possibly rehybridize, allowing the excised element to possibly circularize and reintegrate in a manner similar to the bacteriophage A (see Campbell, 1983, for review). Conversely, partial degradation of the 5’ sticky ends protruding from the transposase complex during excision would decrease the probability of circularization. Furthermore, integration of an element carrying additional nucleotides at one or both ends may present other difficulties, as shown in Fig. lob. When such an element is inserted between the single-stranded sequences of a new integration site, the additional nucleotides of the element will cause it to be slightly displaced from the target sequence at this end. Ligation between the 3‘ strand of the element at this end and the target sequence may be impaired. Polymerase begins to fill the gap, and ligation to the 5’ ends of the element is accomplished. Now it is a question of whether or not the 3‘ ends of the element become ligated to the target sequence before the next round of replication occurs. If ligation is accomplished in time, then mismatches will be generated at both ends of the integrated element. The missing bases may then be filled in by a repair mechanism generating an integrated element displaced from the target site duplication by one or more base pairs on each end. Alternatively, if ligation is delayed and the entire structure is repli-

174

PATRICIA NEVERS ET A L . Transposase

a 5’ 3’

Transposase

I

I

GAT I C G G T A + - C 3’ C C A W 5’

GATTCGGT-

+c TAAGCCT-AAG t-Transposase

1-

Tranrposase

)--(....

I-

H G A T T C G G T -CTAAGCCA-

I

- I I C G G T A ~ A A W C A T ~

I--

+

NGATTCG ~ c r t c c c i ~ -CIAAGCCAA~AGAAGCCA+--O

+

p

Fig. 10. A model of the mechanism of transposition in plants. (a) Excision of a transposable element. The wavy line represents a transposable element that is flanked on each end by a duplication of the target site sequence. In the upper line the element is still integrated at the target site, but a transposase enzyme has set staggered nicks at the 5’ ends of the flanking duplication sequences. The figures below this show possible events that occur during excision leading to the revertant structures and excised elements shown in the lower part of the drawing. The hatched circle represents a protein complex, dashed lines indicate 5’ exonuclease activity, and the dotted line symbolizes polymerase activity. On the left-hand side of the drawing the 5‘ fringes attached to the transposable element are removed completely by the 5‘ exonuclease generating a “clean” element. The same 5’ exonuclease also removes some nucleotides of the flanking, single-stranded target site duplication, generating abbreviated templates for repair synthesis. After removal of the element and ligation of the target sequence, an altered duplication lacking one or two base pairs on each side of the former junction is left behind. On the right-hand side of the drawing an excision event is shown where the 5‘ fringes attached to the element are not completely degraded, resulting in an excised element with one or two additional base pairs on each end. During repair synthesis of the target site sequence, polymerase switches templates, as shown by the arrows extending from the dotted lines. When excision has been completed, a duplication remains with inversions one or more base pairs long on each side of the junction. (b) Integration of a transposable element. Symbolism as in (a). The upper line shows the sequence of the target

175

PLANT TRANSPOSABLE ELEMENTS Transposase,

b

c t

YTransposase

.

U.. .-8 ..

~

+

G

C

G

C

A

T

-M

T

A

A

T

C

G

T

A

G

C

r

~

~

~

-****.*‘Polymerase

~

c CTATGAC GGATAC TG

cc TATGAC ~

H G

C A G G A T A C T G ~ G ~ GGATACTG~ J ~ T ~

A

~

Of

o-+

C C T A T G A C~ ~~ G A ~ C C T A T G A C GGATAC TG -crlccntnctT;i 0

+

H C C T A T G A CI C GGATAC T G G I V-

W

site that has been nicked by a transposase enzyme. On the left, an element with no additional terminal base pairs is inserted between the overlapping single-stranded fringes of the target site sequence. Repair synthesis and ligation are completed with no difficulty. Insertion of an element with additional nucleotides at its 5’ ends is illustrated on the right. If repair synthesis is completed before the next round of general replication, and if the gaps opposite the additional, intervening nucleotides are filled, the integrated element will be displaced from the target site duplication by one or more base pairs at each end. If replication occurs before the gaps opposite the intervening nucleotides have been filled, then two different structures will be generated, each consisting of an integrated element that is displaced from the target site duplication at only one end by additional base pairs.

c

~

176

PATRICIA NEVERS E T A L .

cated without any previous mismatch repair, then the two cases shown at the lower right Fig. 10b should result. Here the element is displaced from the target site duplication at only one end. Another question concerns the possible participation of a 5' exonuclease activity such as the one postulated in connection with excision. If degradation of the overlapping 5' ends of the chromosomal target sequence occurs, then the integrated element will be flanked by a duplication of less then characteristic length. It should be noted that while imprecise excision events have been well documented in plants (see Table VII), there is no definitive evidence for aberrant integration events at the moment. It is possible that integration occurs by a slightly different, more precise process than excision.

E. LARGE-SCALE REARRANGEMENTS INDUCED BY TRANSPOSABLE ELEMENTS IN PLANTS

The currently favored model for transposition in prokaryotes proposes that large-scale rearrangements such as deletions and inversions of sequences adjacent to transposable elements are different molecular consequences of the same transposition event. If one assumes, as we have, that transposition in plants proceeds by a process different than the replicative one proposed for prokaryotes, then another explanation must be found for events of this kind observed in association with transposable elements in plants (McClintock, 1951a, 1956b, 1965a). Here rearrangements might occur as the result of homologous recombination between copies of an element scattered throughout the genome, as shown in Fig. 11. Pairing of this kind might serve to stabilize the DNA/protein complex thought to be formed during these processes. The soybean insertion element Tgml also exhibits sequence homology between its termini and flanking sequences of the lectin gene (L. Vodkin, personal communication). This element has 30-bp, nearly perfect inverted repeat sequences at its termini. Five 5Cbp direct tandem repeats are found in the left arm of the element, each of which contains inverted repeats that can be drawn as hairpin structures. A 7-bp sequence, ACATCGG, is found in the stems of these repeats as well as in the terminal inverted repeat sequences and the target site sequence (L. Vodkin, personal communication). More degenerate forms of the basic 54 repeating unit structure can also be found, and these too contain the 7-bp ACATCGG sequence or a close derivative of it. All together there are 15 hairpin structures and 33 occurrences of the 7-bp sequence or its derivatives within T g m l . The significance of this structure is unknown.

PLANT TRANSPOSABLE ELEMENTS

177

F . SPECIFICITY OF INTEGRATION

Doring and Starlinger (1984) have analyzed the target site sequences surrounding a number of cloned transposable elements. They have found a 6to 10-bp perfect or nearly perfect direct duplication in the target site sequence next to the integrated element for five insertion s,ites of Ds elements, for the Mu1 insert in adhl-S3034 and for Spm-I8 and Taml. Two tandem repeats of 58 bp each are also found about 800 bp upstream from the TgmZ insertion site in the lectin gene (L. Vodkin, personal communication). Doring and Starlinger suggest that transposable elements in plants may have some preference for integrating next to a direct duplication of this kind. Alternatively one could imagine that such duplications represent “fossil sequences” from previously integrated elements. Another possibility has been suggested for the integration specificity of Tam1 (Bonas et al., 1984b). Stretches of sequence homology in direct and inverse orientation can be found within each terminus of the element. If the endmost inverted repeats in the termini of one strand of Taml are paired, and if further basematching occurs within each terminus, a stemand-loop structure reminiscent of a tRNA can be formed, as illustrated by the middle-sized line in Fig. 12. The single-stranded loops exposed in this structure contain extensive homology to sequences of the Taml target site in the niuea locus. One could imagine that the exposed sequences in Tam1 interact with homologous sequences of the target site during integration and/or excision, as shown in Fig. 12.

V. INTERACTION BETWEEN TRANSPOSABLE ELEMENTS A . GENERAL REMARKS

Several lines of evidence suggest that expression of a locus harboring a transposable element may not be governed solely by the resident element but ultimately by the interaction of this element with related elements elsewhere in the genome. The most obvious and well-documented example is that of the two-component systems in maize, where a regulatory element provides functions that mediate its own activities as well as those of related receptor components. In view of the evidence discussed in Section I11 that some receptor elements are simply altered versions of a regulatory element, partially or totally defective in autonomous expression, and that multiple copies of such elements are commonly present in the genome of an organism, the picture becomes exceedingly complex. In

178

PATRICIA NEVERS ET A L . A

Transposition to the homologous chromosome

A

S

-

0

C

D

w

- 1

transposition

B

A 1

,

4,.Au--

L

C

D

4

I

recombinalion

A

___

+

C

deletion

D

acentric ring

2

=

0

A,,

D c d -

- 1

acentric deletion

ring

3

A

_. -.

A

-.

B,

7 -.

- 0 -

L

A

... .

- 3

.

I a -

C D

C

D

paricentric inversion

pericentric inversion

Fig. 1 I . Schematic representation of events possibly leading to gross chromosomal rearrangements in plants. Wavy line indicates a transposable element while the arrow beneath it shows its orientation. Each line designates a chromosome, whereby A , B, C,and D are genetic loci, and the open circle represents the centromere.

PLANT TRANSPOSABLE ELEMENTS B

Transposition withln a chromosome

A

0

=

"

D

C

ryuvI*M

transposition

- 1

A

=

1

B

"

D

C

Y E E e

r between homologous elements

A

--

0 - C D

\L CD deletion

2

acentric deletion 4

A

B

A =

3

B

C

0

C D

U

dicentric duplication

n

c

A

1

. f B

B

"

a

c

A

.

A

0

V _

C D

n

t

I a

acentric deletion duplication dicent ric deletion duplication

acentric deletion

__._. 8 3a ~

C -

Fig. 11 (Continued)

dicentric duplication

179

180

PATRICIA NEVERS E T A L . C

Transposition t o another chromosome

A ,

x

-

Y

A -

x

Y

X

-

I

Y-

_y

“IU

-

C

D

Z

2

-

A

-

B

1

B

-

-

B

C D

z

reciprocal translocation acentric dicenlric

reciprocal translocation

C D

Fig. 11 (Continued)

Fig. 12. Possible basis for integration specificity of Tam2 in A. mujus. The thin line represents one strand of the target site sequence in the niveu locus of A. mujus. The middlesized line shows the terminal sequences of one strand of an integrated Tam2 element. Starting at the ATA duplication flanking the integrated element, a stem structure is formed by pairing the inverted repeat sequences at each end of Turnl. Further pairing of inverted repeats within each individual terminus generates an elaborate stem-and-loop structure. Sequences exposed in such loop structures are available for pairing with homologous sequences of the target site, as shown by the thick lines.

PLANT TRANSPOSABLE ELEMENTS

181

plants we may be dealing with a system such as that found for P elements in Drosophila, where copies of different sizes are scattered throughout the genome, but only the larger ones provide in trans all the functions required for transposition (Spradling and Rubin, 1982; Rubin and Spradling, 1982; Rubin, 1983). B. DOSAGE EFFECTS

That additional copies of completely functional regulatory elements in plants can alter the expression of a locus under control of a single, related element is illustrated by several well-known examples of “dosage effects.” Here a number of different kinds of effects have been observed depending upon the kinds of proteins involved. In the presence of two copies of Ac, for example, there is a delay in timing and an apparent reduction in frequency of Ds-induced chromosome breakage and somatic reversion (McClintock, 1948; Brink and Nilan, 1952; Wood and Brink, 1956). Kernels of the Ds-controlled mutant p-uu with two copies of Mp(Ac) in the genome have fewer and smaller sectors of pericarp pigmentation. This effect may be due to accumulation of a repressor of the element-encoded transposase gene in the cell. The same interpretation may also apply to Robertson’s mutator maize lines. When a nonmutator line is crossed to a mutator line, 30 to 50 times more mutations are found than in control crosses, many of which are unstable and may thus be due to insertions of transposable elements (Robertson, 1978). In particular the element Mu1 is present in multiple copies in mutator lines but lacking in nonmutator ones (Bennetzen, 1984). As the copy number of the purported elements increases in appropriate crosses, the mutation frequency (= transposition?) levels off and then even decreases (Robertson, 1983). Thus in the two cases described above increased copy number of particular transposable elements is associated with decreased mobility of these elements, which may be due to increased levels of an inhibitor. The opposite effect is observed with the dt (Dotted) system in maize. Here the dosage effect seems to be simply an additive one. With two copies of the regulatory element, as opposed to one, the mutability of a corresponding receptor component is higher, as demonstrated by increased variegation (Rhoades, 1936, 1938; Nuffer, 1955, 1961). The same kind of dosage effect was observed for the Fcu regulatory element of the Fcu two-element system in maize (Gonella and Peterson, 1977). Likewise Friedemann and Peterson (1982) found that two doses of the Uq regulatory element, operating on a receptor element at the a locus in maize, lead to increased spotting, but a third copy has no further effect, as if the system were saturated. Thus there may be copy number and accordingly

182

PATRICIA NEVERS E T A L .

also protein threshold levels above which no further dosage effect is observed. The dosage effects one sees below this threshold value may correspond to titration of transposase activity. A curious example of a transposable element with additive dosage effects is Ac2, a new maize element isolated and characterized by Rhoades and Dempsey (1982, 1983). That this element can be legitimately considered an Ac is based on its ability to transactivate certain Ds elements. However, its behavior is anomalous in a number of respects. While one dose of a standard Ac is capable of inducing somatic reversion of the Dscontrolled allele bz2-m, two doses of Ac2 are required to achieve the same effect. Moreover, whereas two or three doses of a standard Ac lead to reduced mutability of b d - m , three doses of Ac2 have the opposite effect, namely enhanced mutability. Ac2 also differs from a standard Ac in its ability to stimulate chromosome breakage at a Ds in standard position on chromosome 9 (Fig. 3). Four doses of Ac2 as opposed to one dose of a standard Ac are required for chromosome breakage to occur. Four doses of Ac2 also induce infrequent reversion of the Ds-controlled allele wx-ml, but have no effect on Ds wx-m9. Both alleles are highly mutable in the presence of a single dose of a standard Ac. Thus the regulation and possibly also the structure of Ac2 must differ from those of a standard Ac. Another intriguing feature of Ac2 is its ability to distinguish between the Ds elements in b d - m , wx-ml, and wx-m9, which suggests that these Ds elements also differ in structure. Along this line of thought it is interesting to note that both the exceptional 0.4-kb element Dsl and the Ds responsible for the adhl-2Fll allele were derived from the bz2-m mutant (Osterman and Schwartz, 1981; Doring et al., 1984b).Thus it is possible that one or both of these Ds elements are structurally related to the Ds present at the bz locus in bz2-m. Therefore it would be interesting to see whether Ac2 exhibits the same dosage effects with these two Ds elements as with the Ds in bz2-m. Dosage effects of the regulator element Spm can only be observed in conjunction with a unique allele of a2, the allele 122-ml. The receptor component at the a2 locus in the a2-ml mutant allows expression of the locus, and a2-ml kernels are pigmented in the absence of an Spm regulatory element. This receptor element is unusual in that it responds to the suppressor function of Spm but not to the mutator function (McClintock, 1957, 1958, 1961a,b).Thus under the influence of an active Spm regulator, a2-ml kernels are uniformly colorless and show no variegation. However, if the standard regulator component is replaced by one that frequently alternates between active and inactive states, variegation can indeed be seen. In combination with such an Spm, a2-ml kernels are basically colorless due to Spm suppressor activity, but have spots of pigmented tissue

PLANT TRANSPOSABLE ELEMENTS

183

in which Spm has become inactive. When two such regulator elements are present in the genome, pigment is produced only when both are inactive. Since two inactivation events are less probable than one, a2-ml kernels with two doses of the cyclically alternating Spm have fewer pigmented spots than those with one dose (McClintock, 1957, 1958, 1959, 1961a). C. INTERACTION WITH ALTERED ELEMENTS

In the examples of dosage effects with two component systems summarized above it is clear that the interacting factors are transposable elements and that those contributing to the dosage effect are completely functional regulatory components. More subtle effects on expression can be obtained with mutationally altered regulatory elements. One such case is the En-Malt (= Restrainer) (Peterson, 1976a) element reported by Reddy and Peterson (1983). When En-Malt is combined with certain alleles of the c locus, each containing a functional, autonomous En, ajine pattern of spotting is observed instead of the usual coarse pattern, indicating that the timing of somatic reversion events is delayed. A standard En does not have this effect. En-Malt apparently produces this delay only in combination with an autonomous En, but not with a standard receptor component at the c locus. In the latter case the normal coarse pattern of variegation is observed. Another example is the “Modifier” of Spm(En) reported by McClintock (1956a, 1965a). The evidence available suggests that this is a defective regulatory component. It can transpose, but not autonomously. By itself it cannot induce mutability of an appropriate receptor component, and it also seems to lack the suppressor function of a normal regulatory element described elsewhere in this article (Section 11,B). However, it can somehow interact with so-called ‘‘weak” Spm regulatory components (McClintock, 1958). “Weak” Spm regulatory components normally induce low mutability of a receptor element. In the presence of “Modifier,” however, normal mutability is restored. Perhaps “Modifier” can complement the “weak” regulatory component by providing an efficient transposase subunit, or perhaps it enhances transcription of the weak Spm element. D. OTHER MODIFIERS

In the examples cited above there is reasonable evidence that the modifying factors are themselves transposable elements related to the elements whose activity they control. In other instances the data are less conclusive. The modifier of the unstable r-st allele in maize, m-st (Ashman, 1960), is a case in point. The r locus is involved in anthocyanin synthesis

184

PATRICIA NEVERS ET A L .

and r-st is an unstable allele of this locus which exhibits spotted aleurone pigmentation. In the presence of m-st a high degree of somatic reversion is observed, while loss of m-st is accompanied by a reduction in spotting. M-st also appears to be transposable (Ashman, 1969). The modifying factor “Stabilizer” (Harrison and Fincham, 1968; Harrison and Carpenter, 1973a,b) in A . majus reduces the instability of the pal-rec allele (Harrison and Fincham, 1968) and seems to be specific for this allele since it has no effect on the niuea-recurrens allele (Harrison and Carpenter, 1973b). Furthermore, “stabilizer” itself seems to be subject to activity changes (Harrison and Fincham, 1968; Harrison and Carpenter, 1977, 1980). The specificity and instability of “stabilizer” support the notion that a related transposable element is involved. The reported “stabilizer” of the niu-rec allele may represent another such case (Harrison and Carpenter, 1973b). In still other cases there is evidence for neither specificity nor instability of the modifying factor. In a number of these instances, however, it has been shown by clear-cut classical genetics (see Section I1,B) that a single, independently segregating modifying factor influences the mutability of a particular allele. These include the “speckled-reduced” factor in Pharbitis nil (Imai, 1931); a modifying gene in Delphinium ajacis (Demerec, 1931); the 0 3 modifier responsible for increased somatic instability of the g l pigment locus in Lathyrus (Punnett, 1932); the In1 allele in Petunia, which decreases color intensity in flowers as well as the somatic reversion frequency of unstable alleles of a1 (Gerats et al., 1982); the fleck-timer Flt(3) in Nicotiana, which delays somatic reversion of unstable u alleles and promotes reversion in germinal tissue (Sand, 1976); and the factor S in the “Sippe 50” line of A . majus, which promotes germinal reversion of the mutable perlutea-uariabilis allele but has no effect on somatic reversion (Meyer et al., 1982). Finally there are several examples of genetically less well-defined modifying factors affecting the mutability of a particular allele. In these cases no readily interpretable segregation ratios have been obtained. Modifier(s) of the regulatory element Dt (dotted) in maize cause a decrease in somatic reversion events associated with the receptor element present at the a locus (Rhoades, 1938). Jeffries and Sastry (1981) report that there are at least three different loci in addition to the previously mentioned “stabilizer” that influence the instability of a pal-rec mutation in A . majus. Further evidence is provided by examination of the frequence of germinal reversion of the mutable inc2-recurrens allele in successive generations in A . majus (Linnert, 1978). The average frequency of reversion in heterozygous inc-recline- plants was found to vary significantly from generation to generation. Moreover the frequency of germinal reversion in inc-

PLANT TRANSPOSABLE ELEMENTS

185

reclinc-rec homozygotes derived from crosses with another line differed significantly from that of homozygotes established by inbreeding the increc parent plant. This was attributed to multiple modifying factors varying in different genomic backgrounds and segregating independently of the inc2 locus. Another case concerns a mutable allele of Mirabilis jalapa. By statistical analysis Delool and Tilney-Basset (1984) found that the different spotting frequencies of 12 individuals of the unstable uariegata mutant of Mirubilis julupu are heritable. they suggest that polygenic control of mutability is responsible for the pattern of inheritance found. A similar case may be that of the perlutea-variabilis allele of A . majus (Stubbe et al., 1981), where a given population of plants exhibits a continuous spectrum of altered flower forms, leaf rolling, and anthocyanin spotting. Here too the various different phenotypes seem to be heritable. It should be noted that in most of the instances cited above it is impossible to say conclusively whether segregation of multiple modifying factors is responsible for the varying phenotypes. It is equally possible that frequent mutational alterations at the mutable locus generate the different variants. However, in view of the evidence for multiple copies of related transposable elements (see Section I1,C) in the genomes of different plant species, it is tempting to speculate that in at least some of the cases listed above the putative modifying factors consist of structurally and functionally related elements.

E. POSSIBLE INTERACTION BETWEEN TWO TRANSPOSABLE ELEMENTS OF

Antirrhinum majus

In plants several different cases have been reported in which two different alleles of a given locus interact in a fashion that is not compatible with what is predicted by classical Mendelian genetics. When two such alleles are brought together in a heterozygote, a heritable change of one allele occurs with high frequency, and the corresponding parental allele can no longer be recovered in subsequent generations. This phenomenon has been termed paramutation, as opposed to classical mutation since it occurs regularly in appropriate heterozygotes, and since the change that occurs is always in one particular direction (for reviews see Brink, 1973; Kermicle, 1973; Hagemann and Berg, 1977; Gavazzi, 1977). A similar case has also been reported for two alleles of Antirrhinum rnajus and attributed to paramutation (Harrison and Carpenter, 1973b). Recent molecular analysis of the two alleles involved (Bonas et al., 1984a,b; K. Upadhyaya, personal communication) has shed light on the molecular basis of this phenomenon.

186

PATRICIA NEVERS ET A L .

The unstable niuea-recurrens (niu-53) mutant of A . majus has highly variegated, red and white flowers and bears the 17-kb element Tam1 in the promoter region of the chalcone synthase gene (Bonas et al., 1984a). In contrast the niu-44 allele is stable and characterized by white flowers. This line contains the 5-kb element Tam2 at the first exonhntron junction of the chalcone synthase gene (K. Upakhyaya, personal communication). When these two lines are crossed, the FI is expected to exhibit the highly variegated phenotype of the niu-53 allele. Strangely enough, however, a large fraction of the F1(66-88%) have flowers with an intermediate pink phenotype (Harrison and Carpenter, 1973b). This is summarized in Fig. 13. An occasional full-colored revertant can be found in the FI,indicating that the effect cannot be explained by a simple “diluting” factor in the niv-44 genome (Nevers, unpublished results). The fraction of F1individuals exhibiting the unexpected pink phenotype varies depending upon several parameters. The number of such plants is greater when niv-44 is used as a female as opposed to reciprocal crosses. It is also increased when crosses and seed maturation are carried out at 25 rather than at 15°C. Finally the phenomenon shows some specificity since it occurs only in combination with the niu-44 allele, while another stable niu- allele, niv45, has no effect. Moreover, one allelic variant of niv-53, the allele niu-55, which exhibits flaking only in the marginal area of the petals, seems to be less susceptible to paramutation. When niv-55 is crossed with niv-44, 83% of the F1is flaked (Harrison and Carpenter, 1973). But the most important observation is that individuals with pink flowers that have arisen by paramutation maintain their phenotype for two further generations and in repeated backcrosses. Segregation of the original niv-53 allele is not observed (Harrison and Carpenter, 1973a). Thus it appears that instability of the niv-53 allele can be altered in a genetically heritable fashion by exposure to the Tam2-induced niv-44 allele. The most pressing question now is how niu-44 interacts with niv-53 in this form of paramutation. In considering this question it is important to note that the genetic evidence currently available does not indicate whether the niv-44 allele itself or some other factor(s) in the niv-44 genome is responsible for paramutation. However, in view of the presFig. 13. Paramutation in A . majus: a possible case of interaction between two transposable elements. The niu-53 mutant harbors the transposable element Tam1 within the promoter region of the chalcone synthase gene and has highly variegated flowers. The niu-44 allele is a stable, recessive allele caused by the integration of Tam2 at the first exonhtron junction of the chalcone synthase gene. Flowers of the niu-44 line are white. When niu-53 and niu-44 lines are crossed, the majority of the F, plants have flowers with the intermediate pink phenotype shown here rather than the expected variegated phenotype (Harrison and Carpenter, 1973). See text for details.

187

PLANT TRANSPOSABLE ELEMENTS

.i l P

P1

El

lVS1

EZ

IVS2

€3

P

El

lVSl

E2

niv-44

niv- 53

niv- 53/n iv-44

lVS2

€3

188

PATRICIA NEVERS ET A L .

ence of Tam2 at the niuea locus in niu-44, it is tempting to speculate that this Tam2 element and/or other copies of Tam2 in the niv-44 may be involved. One possible explanation for this form of paramutation is that some product of Tam2 causes a structural alteration in or around the Tam1 on the nivea homologue. Alternatively such a product might influence transcription or excision of Taml without causing a structural alteration. Here it should be recalled that Tam2 and Taml share extensive sequence homology in their termini, implying that certain element-specific proteins may be able to operate on both elements in common. Elucidation of this problem can be expected from the molecular analysis of paramutant F, plants currently in progress in our laboratory. VI. INDUCTION OF GENOMIC INSTABILITIES In Table VIII a series of procedures are listed by which variegation has been evoked in the past. In some instances transposable elements are known to be involved, in others this has not yet been demonstrated conclusively. What is remarkable is the wide variety of means by which variegated plants can be induced. The best characterized transposable elements, the A d D s and Spm systems in maize, arose in material that had experienced repeated chromosomal breakage and healing during development (McClintock, 1942a,b, 1946, 1950a, 1951a). The breakage-fusion-bridge (BFB) cycle was initiated by introducing an altered form of chromosome 9 into the genome. This particular chromosome carries a duplication of part of the short arm in inverse order. As shown in Fig. 14, crossing over between the inverted, duplicated portion of this chromosome and its homologue during meiosis generates a dicentric chromatid that is torn apart at anaphase. New dicentrics are formed by fusion of chromatids with “ragged ends” produced in this manner (see Fig. 14) and the cycle is perpetuated. Mobilization of transposable elements by the breakage-fusion-bridge cycle has been observed in two other instances in maize (Doerschug, 1973; Bianchi et al., 1969). Induction of heritably variegated plants in association with dicentric chromosomes has also been achieved in Nicotiuna (Goodspeed et al., 1933; Saccardo and Devreux, 1968)and in Pisum (Saccardo, 1971). In addition three new mutable systems, Ac2, Mut, and Mrh, were isolated from a line in which dicentric bridge formation led to loss of segments of chromatin from knobbed A chromosomes. The dicentric bridges were proposed to have arisen by delayed replication of the heterochromatic knobs (Rhoades and Dempsey, 1982).

189

PLANT TRANSPOSABLE ELEMENTS

TABLE VIII Induction of Variegation in Plants: Selected Examples

Inductive Measure Chromosome disruption (breakagefusion-bridge cycle) Mutagenesis: Irradiation X-ray and UV irradiation y irradiation Heat Induction of haploidy Interspecific crosses Viral infection

Plant

References

Zea mays

McClintock (1950b; 1951; 1965); Doerschug, (1973); Rhoades and Dempsey (1982, 1983)

Zea mays Zea mays

Peterson (1960) Neuffer (1966)

Petunia hybrida Antirrhinum rnajus

Cornu (1977) Stubbe, cited in Kuckuck ( 1936) Straub (1973) Sand (1957, 1969, 1976); Smith and Sand (1957) Sprague and McKinney (1966, 1971); Friedernann and Peterson (1982); Mottinger et a / . (1984) Robertson (1977); Strommer et ul. (1982); Bennetzen ei al. (1984); Freeling (1984) Lorz and Scowcroft (1983); Lorz (1984) Rhoades, cited in McClintock (1951a) Bianchi et al. (1978) Sastry et a/. (1980) Gonella and Peterson (1976)

Petunia hybrida Nicotiana langsdorfii and N . sanderae Zea mays

Crosses with Robertson's mutator line

Zea mays

Cell culture regeneration

Nicotiana tabacum

Seed aging

Zea mays

None; spontaneous occurrence

Petunia hybridu Impatiens balsaminu Zea mays

In other cases mutable alleles arose after exposure of plants to various forms of mutagenesis. The allele pg-m, for example, was isolated in plant material exposed to atomic radiation subsequent to tests on the Bikini Islands (Peterson, 1960). y-Ray treatment of zygotes of Petunia hybrida as well as other kinds of chemical mutagenesis led to the isolation of mutable alleles of the an2 and r loci (Cornu, 1977). Sometimes y-irradiation elicited cycles of chromosome breakage and fusion as described above (Cornu, 1977). In maize lines bearing the receptor elements of either Dt or Spm at the a locus but no detectable regulator elements, regulator activity was observed after exposure of pollen from these lines to X-ray and UV treatment (Neuffer, 1966). This was manifested by the

190

PATRICIA NEVERS E T A L . MEIOSIS

s h r

&

Prophase 1

Anaphase 1

baabc

Gamete

I

F E RTI L IZATION a) One gamete with

dicentric chromosome MITOSIS

b) Both gametes with dicentric chromosomes

MITOSIS

b

b b

Anaphase

b

aabc

___o__

Telophase

cba

bbaabc

7

1 BFB

c

Prophase

PLANT TRANSPOSABLE ELEMENTS

191

appearance of pigmented spots on kernels of plants that had been pollinated with irradiated pollen. Heating Antirrhinum pollen at 30°C for 108 hr or exposing it to X-ray irradiation also resulted in mutants with variegated flowers (Kuckuck, 1936). Mutable alleles have also been induced by less obvious treatments. Passing Petunia through a haploid stage was accompanied by the occurrence of new mutable alleles manifested by variegated flowers. No such alleles were found in diploids under the same circumstances (Straub, 1973). A cross between two different species of Nicotiana induced new mutable alleles of the u locus (Sand, 1976). Following infection of maize with barley stripe mosaic virus (BSMV) or wheat stripe mosaic virus (WSMV), many new mutants can be found, some of which are unstable. These mutants exhibit aberrant segregation ratios, even in descendants many generations removed where viral proliferation has obviously ceased (Sprague and McKinney, 1966, 1971). Viral sequences cannot be detected in this material, although new mutations continue to arise (Wienand et al., 1983). One mutant isolated from aberrant ratio (AR) lines proved to be unstable and could be attributed to the action of a new, hitherto unknown two-component system called Ubiquitous (Vq) (Friedemann and Peterson, 1982). A series of still other, shrunken mutants have been analyzed by Southern hybridization, and all but one exhibit structural DNA rearrangements at the sh locus, likely to be associated with the presence of insertional elements (Mottinger et al., 1984; S . Dellaporta, personal communication). Five out of eight such sh mutants are unstable, and one of these is the source of the Tz86 element listed in Table V. Moreover, marker loss reminiscent of that generated by BFB cycles can be observed in 0.12 to 4% of the progeny from crosses with aberrant ratio plants, while the frequency in normal stocks is only about (Mottinger et al., 1984). These cases must be distinguished from those discussed in Section I where variegation is caused by direct effects of viral infection. Fig. 14. The breakage-fusion-bridge cycle (BFB) in maize according to McClintock, 1951a. The maize line used to initiate the BFB cycle carries a duplication in inverse orientation in one homologue of chromosome 9 and a normal sequence in the other, as shown in the upper drawing. During meiosis in germinal tissue, classical crossing over generates a dicentric chromatid that is torn apart at the first meiotic anaphase stage. Each such torn chromatid enters a gamete and is replicated, after which the two sister chromatids fuse at the point of lesion. Two different cycles of breakage can be generated depending upon whether one or both gametes that fuse at fertilization contain such a dicentric chromosome. In the first case, shown on the left, the single dicentric chromosome continues to initiate BFB cycles in the endosperm of the kernel but seldom in the embryo. When two dicentric chromosomes are present, as shown on the right, each may be torn apart at mitotic anaphase. Parts from two different tom chromosomes can then fuse, forming a dicentric chromatid that is replicated as a whole. In this case, BFB cycles are observed to continue in the embryo.

192

PATRICIA NEVERS ET A L .

In maize, mutable alleles can be readily induced by crossing normal stocks with mutator lines (Robertson, 1978), which have been referred to at several points in this article. Approximately 35% of the mutations generated in this manner are unstable (Robertson, 1978), and a number of the unstable ones seem to have been caused by insertion of the 1.4-kb element Mu1 (Strommer et ul., 1982; M. Freeling, B. Burr, and F. Burr, cited in Freeling, 1984). Furthermore, only mutator lines contain multiple copies of this element while it is completely missing in nonmutator lines (Bennetzen, 1984). It is still not clear whether most mutants derived from mutator lines are Muf-induced, nor is it certain whether products of MuZ itself are responsible for mobilization of transposable elements in mutator lines. However, several aspects of the inheritance of mutator capacity indicate that a polyfactorial, mobile system is involved. Inheritance is non-Mendelian; in fact, it is almost infectious. When hybrids between mutator and nonmutator lines are backcrossed to a nonmutator line, 87% of the progeny exhibit the mutator trait instead of the expected 50%, and these continue to remain mutators (Robertson, 1983). Thus the mutator trait cannot be diluted out by crosses. It appears that in mutator lines transposable elements may be active that continue to transpose and/or mobilize other elements in subsequent generations. One case has been reported in which it appeared as if allowing maize kernels to age resulted in the induction of new mutable alleles (Rhoades, 1950, cited in McClintock, 1951a). Finally many mutable alleles seem to have arisen spontaneously and have been maintained by selection. Interestingly, selection has not been restricted to scientists concerned with transposable elements. Chance variegated kernels of corn must have caught the fancy of certain American Indians many generations back, who proceeded to propagate them. Thus the FCU system was discovered in a particular population of variegated tribal maize maintained by the Cuna Indians in Colombia (Gonella and Peterson, 1976). Once a transposable element has been released, that is, once a particular mutable allele has arisen in a plant, new instabilities continue to be found at a greater than average rate in its descendants, as discussed in Section II1,G. One question which arises is whether the new instabilities are incited primarily by elements from the same family, or whether there is general mobilization of transposable elements. Southern hybridization analysis revealed that the Antirrhinum element Tumf apparently continues to transpose in revertants of a mutant induced by this element (Bonas, 1984). This observation provides support for the idea that “resurgence of mutability” in A . mujus (Harrison and Carpenter, 1979), as discussed in Section II,D, is due to continued transposition of a previ-

PLANT TRANSPOSABLE ELEMENTS

193

ously excised element. However, the fact that transposable elements can become activated by such widely diverse means as those listed in Table VIII suggests that general mobilization may sometimes occur in response to extreme provocation. McClintock (1978b) proposed that transposable elements constitute an inherent system of the organism for generating genomic diversity and subsequently also adaptability when the need arises. The circumstances under which the potential of transposable elements is released have been summed up in the term “genomic stress.” When an organism is subjected to such stress, it increases its own mutational frequency via transposable elements in an attempt to survive. Thus our attitude toward transposable elements is moving from regarding them simple as genetic delinquents to considering them as genetic lifesavers.

VII. ADDENDUM In preparing this manuscript we have become aware of the inadequate state of plant genetic nomenclature and of a very definite need for all plant geneticists and molecular biologists to come to grips with this problem. We wish to make an urgent plea to reach some kind of consensus on this matter as soon as possible. We ourselves favor the nomenclature used in prokaryotic genetics (see Miller, 1972) and have applied these rules as much as possible in the text of this manuscript. Thus the phenotype is designated by a three-letter abbreviation beginning with a capital letter (Gal+, Niv+) while lower case letters are used for the genotype (gal-301, niu-53).To indicate the presence of a transposable element the symbolism recommended by Campbell et al. (1977) might be applied (e.g., niu53::TumJ). This nomenclature differs somewhat from that recommended by maize geneticists (see the 1975 Maize Genetics Cooperation News Letter, Vol. 49).

ACKNOWLEDGMENTS We wish to thank Frau F. Furkert, Frau M. Kalda, and Herr D. Bock for their help in preparing the drawings and photographs; Frau G. Linde and Frau M. Pasemann for assistance in typing; Frau C. Wienke for library services; Dr. B. Bowman for the computerized “back-search” on mutable alleles; Frau L. Glahn and Herr F. Shepherd for assistance in preparing this manuscript and the many scientists who provided us with preprints and unpublished results. We also wish to thank Prof. P. Starlinger and Dr. H.-P. Doring for critical reading of the manuscript.

194

PATRICIA NEVERS ET AL.

REFERENCES Ah-Khan, S. T. (1971). Can. J. Genet. Cytol. 13, 736-741. Anderson, E. G. (1924). Genetics 9,442-453. Andersson, I . (1923). J . Genet. 13, 1-11. Anderson-Kotto, I. (1930). Z. Indukt. Abst. Vererbungsl. 56, 115-201. Arthur, A., and Sherratt, D. (1979). Mol. Gen. Genet. 175, 267-274. Ashman, R. B. (1960). Genetics 45, 19-34. Ashman, R. B. (1965). Genetics 51, 305-312. Barclay, P. C., and Brink, R. A. (1954). Proc. Natl. Acad. Sci. U.S.A. 40, 1118-1126. Baur, E. (1910a). Z. Indukt. Abst. Vererbungsl. 3, 34-98. Baur, E. (1910b). Z. Induki. Absr. Vererbungsl. 4, 81-102. Baur, E. (191 1). “Einfiihrung in die experimentelle Vererbungslehre.” Gebr. Borntraeger, Berlin Baur, E. (1912). Z.Indukt. Absr. Vererbungsl. 6, 201-216. Baur, E. (1918). Z. Indukt. Abst. Vererbungsl. 19, 177-193. Baur, E. (1924). Bibl. Genet. 4. Baur E. (1926). Z. Indukt. Abst. Vererbungsl. 41,47-53. Baur, E. (1932). Z. Indukt. Abst. Vererbungsl. 60,467-473. Behrens, U., Fedoroff, N., Laird, A., Miiller-Neumann, M., Starlinger, P., and Yoder, J. (1984). Mol. Gen. Genet. 194, 346-347. Bennetzen, J. L., Swanson, J., Taylor, W. C., and Freeling, M. (1984). Proc. Natl. Acad. Sci. U.S.A. 81,4125-4128. Benoist, C., O’Hare, K., Breathnach, R., and Chambon, P. (1980). Nucleic Acids Res. 8, 127-142. Berg, D. E. (1983). Proc. Narl. Acad. Sci. U.S.A. 80, 792-796. Bergfeld, R. (1958). Z. Indukt. Abst. Vererbungsl. 89, 143-160. Bianchi, A., Salamini, F., and Parlavecchio, R. (1969). Genet. Agrar. 22, 335-343. Bianchi, F., de Boer, R., and Pompe, A. J. (1974). Acta Bot. Merl. 23, 691-700. Bianchi, G., Avato, P., and Salamini, F. (1977). Maydica 22, 9-17. Bianchi, F., Cornelissen, P. T. J., Gerats, A. G. M., and Hogervorst, J. M. W. (1978). Theor. Appl. Genet. 53, 157-167. Blakeslee, A. F. (1920). Genetics 5, 119. Bonas, U. (1984). PhD thesis, University of Cologne. Bonas, U., Sommer, H., Harrison, B. J., and Saedler, H. (1984a). Mol. Gen. Genet. 194, 138-143. Bonas, U., Sommer, H., and Saedler, H. (1984b). EMBO J . 3, 1015-1019. B@rresen,S., and Fjeld, A. (1977). Hereditas 85, 215-218. Bresler, S. E., Tamm, S. E., Goryshin, I. Yu.,and Lanzov, V. A. (1983). Mol. Gen. Genet. 190, 139-142. Brink, R. A. (1954). Genetics 39, 724-740. Brink, R. A. (1956). Genetics 41, 872-889. Brink, R. A. (1958). Genetics 43,435-447. Brink, R. A. (1973). Annu. Rev. Genet. 7, 129-152. Brink, R. A., and Barclay, P. C. (1954). Science 119, 577-578. Brink, R. A., and Mikula, B. (1957). Z. Indukt. Absr. Vererbungsl. 89,94-102. Brink, R. A., and Nilan, R. A. (1952). Genetics 37, 519-544. Brink, R. A., and Weyers, W. H. (1957). Proc. Natl. Acad. Sci. U.S.A. 43, 1053-1060. Brink, R. A., and Williams, E. (1973). Genetics 73, 273-296.

PLANT TRANSPOSABLE ELEMENTS

195

Britten, R. J., and Davidson, E. H. (1976). Fed. Proc., Fed. A m . SOC.Exp. Biol. 35, 21512157. Burns, J. A., and Gerstel, D. U. (1967). Genetics 57, 155-167. Burr, B., and Burr, F. A. (1980). Cold Spring Harbor Symp. Quant. Biol. 45,463-465. Burr, B., and Burr, F. (1981a). Stadler Symp. 13, 115-128. Burr, B., and Burr, F. A. (1981b). Genetics 98, 143-156. Burr, B., and Burr, F. A . (1982). Cell 29,977-986. Burr, B., and Burr, F. A. (1983). In “Gene Structure and Regulation in Development,” pp. 185-195. Liss, New York. Calos, M. P., and Miller, J. H. (1980). Cell 20, 579-595. Campbell, A. (1983). In “Mobile Genetic Elements” (J. A. Shapiro, ed.), pp. 65-103. Academic Press, New York. Campbell, A., Berg, D., Botstein, D., Lederberg, E., Novick, R., Starlinger, P., and Szybalski, W. (1977). In “DNA Insertion Elements, Plasmids and Episomes” (A. Bukhari, J. A. Shapiro, and S. L. Adhya, eds.), pp. 15-22. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Christie, W. (1922). Z. Indukt. Abst. Vererbungsl. 27, 134-141. Clutterbuck, A. J. (1970). Genet. Res. (Cambridge) 16, 303-316. Cornu, A. (1977). Mutat. Res. 42, 235-248. Correns, C. (1909). Z. Indukt. Abst. Vererbungsl. 1, 291-329. Correns, C. (1910). Ber. Dtsch. Bot. Ges. 28,418-434. Courage, U., Doring, H. P., Frommer, W.-B., Kunze, R., Laird, A., Merckelbach, A., Miiller-Neumann, M., Riegel, J., Starlinger, P., Tillmann, E., Weck, E., Werr, W., and Yoder, J. (1984). Cold Spring Harbor Symp. Quant. Biol. 49, 329-338. Courage-Tebbe, U., Doring, H.-P., Fedoroff, N., and Starlinger, P. (1983). Cell 34,383-393. Coyne, D. P. (1966). Crop Sci. 6, 307-310. Coyne, D. P. (1967). J . Hered. 58, 146-147. Coyne, D. P. (1969). J . A m . SOC.Hortic. Sci. 94, 488-491. Cuff, E. L., Feenstra, A., Leuders, K., Rechavi, G., Givol, D., and Canaani, E. (1983). Nature (London) 302, 547-548. Darwin, Ch. (1868). “The Variation of Animals and Plants under Domestication.” John Murray, London. Dawson, G. P. W. (1955). Heredity 9, 409-412. De Haan, H. (1930). Genetica 12, 321-439. Delool, R. A. H., and Tilney-Bassett, R. A. E. (1984). Euphytica 33, 169-176. Demerec, M. (1931). J . Genet. 24, 179-193. Dennis, E. S., Gerlach, W. L., Pryor, A. J., Bennetzen, J. L., Inglis, A . , Llewellyn, D., Sachs, M. M., Ferl, R. J., and Peacock, J. W. (1984). Nucleic Acids Res. 12,3983-4000. Derrner, H. (1969). Am. Horric. Mug. 39, 123-173. D’Eustachio, P., and Ruddle, F. H. (1983). Science 220, 919-924. De Winton, D., and Haldane, J. B. S. (1935). J . Genet. 31, 67-100. Doerschug, E. B. (1973). Theor. Appl. Genet. 43, 182-189. Doerschug, E. B. (1976). Theor. Appl. Genet. 48, 119-130. Doodeman, M., Boersma, E. A., Koomen, W., and Bianchi, F. (1984a). Theor. Appl. Genet. 67, 345-355. Doodeman, M., Gerats, A. G. M., Schram, A. W., de Vlaming, P., and Bianchi, F. (1984b). Theor. Appl. Genet. 67, 357-366. Dooner, H. K. (1980). Cold Spring Harbor Symp. Quart. Biol. 45, 457-462. Dooner, H. K., and Kerrnicle, J. L. (1971). Genetics 67,427-436. Dooner, H. K., and Kerrnicle, J. L. (1974). Genetics 78, 691-701.

196

PATRICIA NEVERS ET A L .

Dooner, H. K., and Nelson, 0. E. (1977a). Proc. Nutl. Acad. Sci. U . S . A . 74, 5623-5627. Dooner, H. K., and Nelson, 0. E. (1977b). Biochem. Genet. 15, 509-519. Dooner, H. K. and Nelson, 0. E. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 2369-2371. Doring, H. (1937). Ber. Dtsch. Bot. Ges. 55, 167-182. Doring, H. (1941). Biol. Zentralbl. 61, 65-68. Doring, H.-P., and Starlinger, P. (1984). Cell 39, 253-259. Doring, H.-P., Geiser, M., and Starlinger, P. (1981). Mol. Gen. Genet. 184,377-380. Doring, H.-P., Geiser, M., Weck, E., Werr, W., Courage-Tabke, U., Tillman, E., and Starlinger, P. (1983). In “Genetic Engineering of Plants” (E. Hollander, ed.), pp. 203209. Plenum, New York. Doring, H.-P., Tillman, E., and Starlinger, P. (1984a). Nature (London) 307, 127-130. Doring, H.-P., Fueling, M., Hake, S., Johns, M. A., Kunze, R., Merckelbach, A., Salamini, F., and Starlinger, P. (1984b). Mol. Gen. Genet. 193, 199-204. Dubos, R. J. (1958). In “Perspectives in Virology” (M. Pollard, ed.), Vol. I , p. 291. Wiley, New York. Echt, C. S., and Schwartz, D. (1981). Genetics 99, 275-284. Emerson, R. A. (1914). Am. Nut. 48, 87-115. Emerson, R. A. (1917). Genetics 2, 1-35. Emerson, R. A. (1921). Am. J . Bot. 8, 411-424. Emerson, R. A. (1922). Am. Nat. 56, 64-79. Emerson, R. A. (1929). Genetics 14, 488-511. Emerson, R. A , , and Anderson, E. G. (1932). Genetics 17, 503-509. Engels, J. M. M., Van Kester, W. N. M., Spitters, C. J . T., Vosselman, L., and Zeven, A. C. (1975). Euphytica 24, 1-5. Eyster, W. H. (1924). Genetics 9, 372-404. Eyster, W. H. (1925). Genetics 10, 179-196. Eyster, W. H. (1926). Generics 11, 372-386. Eyster, W. H. (1928). Z . Indukt. Absr. Vererbungsl. Suppl. 1, 666-686. Faberg&, A. C., and Beale, G. H. (1942). J . Genet. 43, 173-187. Farcy, E., and Cornu, A. (1979). Theor. Appl. Genet. 55, 273-278. Fedoroff, N. V. (1983). In “Mobile Genetic Elements” (J. A. Shapiro, ed.), pp. 1-63. Academic Press, New York. Fedoroff, N., Mauvais, J., and Chaleff, D. (1983a). J . Mol. Appl. Genet. 2, 11-29. Fedoroff, N., Wessler, S., and Shure, M. (398313). Cell 35, 235-242. Fedoroff, N., Chaleff, D., Courage-Tebbe, U., Doring, H.-P., Geiser, M., Starlinger, P., Tillman, E., Weck, E., and Werr, W. (1983~).In “Structure and Function of Plant Genomes” (0.Cifem and L. Dure, eds.), pp. 61-72. Plenum, New York. Fedoroff, N., Furtek, D. B., and Nelson, 0. E., Jr. (1984). Proc. Natl. Acad. Sci. U.S.A. 81, 3825-3829. Fincham, J. R. S., and Harrison, B. J . (1964). Heredity 22, 21 1-224. Fincham, J. R. S., and Sastry, G. R. K. (1974). Annu. Reu. Genet. 8, 15-50. Fink, G., Farabaugh, P., Roeder, G., and Chaleff, D. (1980). Cold Spring Harbor Symp. Quant. Biol. 45, 575-580. Flavell, R. B., O’Dell, M., and Hutchinson, J. (1981). Cold Spring Harbor Symp. Quant. Biol. 45, 501-508. Fogel, S. (1950). Rec. Genet. Soc. A m . 19, 105. Fowler, R.,and Peterson, P. A. (1974). Genetics 76, 433-446. Fowler, R. G., and Peterson, P. A. (1978). Genetics 90, 761-782. Fradkin, C. W., and Brink, R. A. (1956). A m . J . Bot. 43,267-273. Freeling, M.(1978). Genetics 89, 211-224.

PLANT TRANSPOSABLE ELEMENTS

197

Freeling, M. (1984). Annu. Rev. Plant Physiol. 35, 277-298. Freeling, M., Cheng, D. S.-K., and Alleman, H. (1982). Deu. Genet. 3, 179-196. Friedemann, P. F., and Peterson, P. A. (1980). Maize Genet. Coop. News Lett. 54, 2-3, Friedemann, P., and Peterson, P. A. (1982). Mol. Gen. Genet. 187, 19-29. Galas, D. J., and Chandler, M. (1981). Proc. Natl. Acad. Sci. U . S . A . 78, 4858-4862. Gavazzi, G. (1977). Stadler Symp. 9, 38-60. Geiser, M., Weck, E., Doring, H.-P., Werr, W., Courage-Tebbe, U., Tillman, E., and Starlinger, P. (1982). EMBO J . 1, 1455-1460. Gerats, A. G. M., Cornelissen, R. T. J., Groot, S., Hagervorst, J. M. W., Schram, A. W., and Bianchi, F. (1982). Theor. Appl. Genet. 62, 199-203. Gerats, A. G. M., Wallroth, M., Donker-Koopman, W., Groot, S. P. C., and Schram, A. W. (1983a). Theor. Appl. Genet. 65, 349-352. Gerats, A. G. M., Groot, S. P. C., Peterson, P. A., and Schram, A. W. (1983b). Mol. Gen. Genet. 190, 1-4. Gerats, A. G. M., Farcy, E., Wallroth, M., Groot, S. R. C., and Schram, A. (1984a). Genetics 106, 501-508. Goldberg, R. B., Hoschek, G., and Vodkin, L. 0. (1983). Cell33, 465-475. Gonella, J. A., and Peterson, P. A. (1977). Genetics 85, 629-645. Gonella, J. A., and Peterson, P. A. (1978). Mol. Gen. Genet. 167, 29-36. Goodspeed, T . H . , and Avery, P. (1933). Genetics 18, 487-520. Gorman, M. B., and Peterson, P. A. (1978). Maydica 23, 173-186. Greenblatt, I. M. (1965). Maize Genet. Coop. News Lett. 39, 119-120. Greenblatt, I. M. (1966). Genetics 53, 361-369. Greenblatt, 1. M. (1968). Genetics 58, 585-597. Greenblatt, I. M. (1974). Generics 77, 671-678. Greenblatt, I. M. (1975). Maize Genet. Coop. News Lett. 49, 23-27. Greenblatt, I. M. (1984). Genetics 108, 471-485. Greenblatt, I. M., and Brink, R. A. (1962). Genetics 47, 489-501. Greenblatt, I. M., and Brink, R. A. (1963). Nature (London) 197, 412-413. Grindley, N., and Sherratt, D. (1978). Cold Spring Harbor Symp. Quant. Biol. 43, 12571261. Gupta, M., Bertram, I., Shepherd, N., and Saedler, H. (1983). Mol. Gen. Genet. 192, 373377. Gupta, M., Shepherd, N . S., Bertram, I., and Saedler, H. (1984). EMBOJ. 3, 133-139. Hagernann, R., and Berg, W. (1977). Biol. Zentralbl. 96, 257-301. Hannah, L. C., and Nelson, 0. E. (1976). Biochem. Genet. 14, 547-560. Hannah, L. C., Tuschall, D. M., and Mans, R. J. (1980). Genetics 95,961-970. Harayama, S . , Oguchi, T., and Iino, T. (1984). Mol. Gen. Genet. 194, 444-450. Harrison, B. (1965). John Innes Inst. Annu. Rep. 15. Harrison, B. J. (1971). Murat. Res. 12, 381-390. Harrison, B. J., and Carpenter, R. (1973a). John Innes Annu. Rep. Harrison, B. J., and Carpenter, R. (1973b). Heredity 31, 309-323. Harrison, B. J., and Carpenter, R. (1977). Heredity 38, 169-189. Harrison, B. J., and Carpenter, R. (1979). Mutat. Res. 63, 47-66. Harrison, B. J., and Carpenter, R. (1980). John Innes Annu. Rep. 71-76. Harrison, B. J., and Fincham, J. R. S. (1963). Annu. Rep. John Innes Inst. 54, 15-17. Harrison, B. J., and Fincham, J. R. S. (1964). Heredity 19, 237-258. Harrison, B. J., and Fincham, J. R. S. (1968). Heredity 23, 67-72. Harrison, B. J., and Stickland, R. G. (1974). Heredity 33, 112-115. Harshey, R. hl., and Bukhari, A. 1. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 1090-1094.

198

PATRICIA NEVERS E T AL.

Hayes, H. K. (1917). Genetics 2, 260-283. Heffron, F. (1983). I n “Mobile Genetic Elements” (J. A. Shapiro, ed.), pp. 223-260. Academic Press, New York. Hertwig, P. (1926). Z . Indukt. Abst. Vererbungsl. 41, 42-47. Hondelmann, W. (1959). Z . Indukt. Abst. Vererbungsl. 90, 159-181. Iida, S., Meyer, J., and Arber, W. (1983). I n “Mobile Genetic Elements” (J. A. Shapiro, ed.), pp. 159-221. Academic Press, New York. Ikeno, S. (1929). Jpn. J . Bot. 4, 189-217. Imai, Y. (1921). Bot. Mag. (Tokyo) 35,49-83. Imai, Y. (1926a). Bor. Mag. (Tokyo) 40, 226-235. Imai, Y. (1926b). J. Genet. 16, 77-99. Imai, Y. (1927). J. Genet. 17, 329-348. Imai, Y. (1930). J . Genet. 22, 191-200. Imai, Y. (1931). J. Genet. 24, 203-224. Imai, Y. (1934). J . Coll. Agric. Tokyo 12, 479-523. Imai, Y. (1934-1935). Z . Indukt. Abst. Vererbungsl. 68, 242-264. Imai, Y. (1936). Z . Indukt. Abst. Vererbungsl. 71, 61-83. Imai, Y. (1938). J. Genet. 35, 375-382. Imai, Y., and Kanna, B. (1935). Genetica 17, 27-31. Jeffries, V. E., and Sastry, G. R. K. (1981). Theor. Appl. Genet. 60, 303-311. Jeffries, V. E., and Sastry, G. R. K. (1982). Theor. Appl. Genet. 63, 81-86. Kedharnath, S., and Brink, R. A. (1958). Genetics 43, 695-704. Kemble, R. J., Mans, R. J., Gabay-Laughnan, S . , and Laughnan, J. R. (1983). Nature (London) 304, 744-747. Kermicle, J. L. (1970). Genetics 64, 247-258. Kermicle, J. L. (1973). Brookhaven Symp. Biol. 25, 262-280. Kermicle, J. L. (1980). Science 208, 1457-1459. Kermicle, J. L. (1984). Genetics 107,489-500. Kihara, H. (1940). Bot. Zool. 8, 975-990. Kihara, H. (1979). Seiken ZihB 27-28, 30-36. Kim, B. D., Mans, R. J., Conde, M. F., Pring, D. R., and Levings, C. S., 111(1982). Plasmid 7, 1-14. Kirk, J. T. O., and Tilney-Bassett, R. A. E. (1978). “The Plastids: Their chemistry, Structure, Growth and Inheritance,” 2nd Ed. Elsevier, Amsterdam. Kleckner, N. (1981). Annu. Rev. Genet. 15, 341-404. Kleckner, N. (1983). I n “Mobile Genetic Elements” (J. A. Shapiro, ed.), pp. 261-298. Academic Press, New York. Klein, A. S., and Nelson, 0. E., Jr. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 7591-7595. Kuckuck, H. (1936). Z . Indukt. Abst. Vererbungsl. 71, 429-440. Kuckuck, H., and Schick, R. (1930). Z . Indukt. Abst. Vererbungsl. 5 6 , 5 1 4 3 . Kutscher, L. (1935). Z . Indukt. Abst. Vererbungsl. 68, 454-485. Larson, R. L., and Coe, E. H., Jr. (1977). Biochem. Genet. 15, 153-156. Lawrence, W. J. C. (1931). J. Genet. 24, 257-306. Lawrence, T. (1974). Can. J . Genet. Cytol. 16, 467-468. Levings, C. S., 111, and Pring, D. R. (1979). I n “Physiological Genetics” (J. C. Scandalios, ed.), Vol. V, pp. 171-193. Academic Press, New York. Levings, C. S., 111, Kim, B. D., Pring, D. R., Conde, M. F., Mans, R. J., Laughnan, J. R., and Gabay-Laughnan, S. J. (1980). Science 209, 1021-1023. Levis, R., Dunsmuir, P., and Rubin, G. M. (1980). Cell 21, 581-588. Lilienfeld, F. A. (1929). Bibl. Genet. 13, 1-214. Linnert, G. (1966). Z . Indukt. Abst. Vererbungsl. 98, 25-40. Linnert, G. (1972). Theor. Appl. Genet. 42, 244-249.

PLANT TRANSPOSABLE ELEMENTS

199

Linnert, G. (1978). Biol. Zentralbl. 97, 513-526. Lorz, H . (1984). I n “Genetic Manipulation: Impact on Man and Society” (W. Arber, K. Illmensee, W. J. Peacock, and P. Starlinger, eds.), pp. 103-114. Cambridge Univ. Press, London. Lorz, H., and Scowcroft, W. R. (1983). Theor. Appl. Genet. 66, 67-75. Maizonnier, D., and Cornu, A. (1971). Genetica 42, 422-436. Mather, K. (1933). PhD thesis, University of London. McClintock, B. (1942a). Proc. Natl. Acad. Sci. U.S.A. 28, 458-463. McClintock, B. (1942b). Carnegie fnst. Washington Year Book 41, 181-186. McClintock, B. (1945). Carnegie fnst. Washington Year Book 44, 108-1 12. McClintock, B. (1946). Carnegie fnst. Washington Year Book 45, 176-186. McClintock, B. (1947). Carnegie fnst. Washington Year Book 46, 146-152. McClintock, B. (1948). Carnegie fnst. Washington Year Book 47, 155-169. McClintock, B. (1949). Carnegie fnst. Washington Year Book 48, 142-154. McClintock, B. (1950a). Proc. Natl. Acad. Sci. U.S.A. 36,344-355. McClintock, B. (1950b). Carnegie fnst. Washington Year Book 49, 157-167. McClintock, B. (1951a). Cold Spring Harbor Symp. Quant. Biol. 16, 13-47. McClintock, B. (1951b). Carnegie fnst. Washington Year Book 50, 174-181. McClintock, B. (1952). Carnegie fnst. Washington Year Book 51, 212-219. McClintock, B. (1953a). Genetics 38, 579-599. McClintock, B. (1953b). Carnegie Inst. Washington Year Book 52, 227-237. McClintock, B. (1954). Carnegie fnst. Washington Year Book 53, 254-260. McClintock, B. (1955). Carnegie fnst. Washington Year Book 54, 245-255. McClintock, B. (1956a). Brookhaven Symp. Biol. 8, 58-74. McClintock, B. (1956b). Cold Spring Harbor Symp. Quant. Biol. 21, 197-216. McClintock, B. (1956~).Carnegie fnst. Washington Year Book 55, 323-332. McClintock, B. (1957). Carnegie fnst. Washington Year Book 56, 393-401. McClintock, B. (1958). Carnegie fnst. Washington Year Book 57,415-429. McClintock, B. (1959). Carnegie Inst. Washington Year Book 58,452-456. McClintock, B. (1961a). Am. Nat. 95, 265-277. McClintock, B. (1961b). Carnegie Inst. Washington Year Book 60,469-476. McClintock, B. (1962). Carnegie fnst. Washingion Year Book 61, 448-461. McClintock, B. (1963). Carnegie fnst. Washington Year Book 62,486-493. McClintock, B. (1964). Carnegie fnst. Washington Year Book 63, 592-602. McClintock, B. (1965a). Brookhaven Symp. Biol. 18, 162-184. McClintock, B. (1965b). Carnegie Inst. Washington Year Book 64, 527-534. McClintock, B. (1967a). Dev.Biol. Suppl. 1, 84-112. McClintock, B. (1967b). Carnegie fnst. Washington Year Book 65, 568-578. McClintock, B. (1968). Carnegie fnst. Washington Year Book 66, 20-28. McClintock, B. (1971). Carnegie fnst. Washington Year Book 70, 5-17. McClintock, B. (1978a). Zn “The Clonal Basis of Development” (S. Subtelny and I. M. Sussex, eds.), pp. 217-237. Academic Press, New York. McClintock, B. (1978b). Stadler Symp. 10, 25-48. McWhirter, K. S. (1973). Abstr. Genet. 74, 170. Mechelke, F., and Stubbe, H. (1954). Z. fndukt. Abst. Vererbungsl. 86, 224-248. Messing, J., Geraghty, D., Heidecker, G., Nien-Tai, H., Kridl, J . , and Rubenstein, I. (1983). In “Genetic Engineering of Plants” (T. Kosuge, C. P. Meredith, and A. Hollander, eds.), pp. 21 1-227. Plenum, New York. Meyer, M., Berg, W., and Hagemann, R. (1981). Biol. Zentralbl. lUU, 483-493. Meyer, M . , Berg, W., and Hagemann, R. (1982). Biol. Zentralbl. 101, 503-511. Michaelis, P. (1957). Protoplasma 48, 403-418.

200

PATRICIA NEVERS ET A L .

Miller, J. H. (1982). “Experiments in Molecular Genetics.” Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Mitra, S. K. (1932). Curr. Sci. 1, 102-103. Mitra, S. K., and Ganguli, P. M. (1934). Indian J . Agric. Sci. 4, 537-545. Miyake, K . , and Imai, Y. (1934-1935). Z. Indukt. Abst. Vererbungsl. 68, 300-302. Miyake, K., and Imai, Y. (1934). J . Coll. Agric. Tokyo 13, 27-44. Miyazawa, B. (1929). Jpn. J. Genet. 4, 167-184. Miyazawa, B. (1932). Bull. Miyazaki Coll. Agric. For. 4, 1 1 1-125. Modolell, J., Bender, W., and Meselson, M. (1983). Proc. Nail. Acad. Sci. U . S . A .80, 16781682. Montanelli, C., DiFonzo, N., Marotta, R., Motto, M., Soave, C., and Salamini, F. (1984). Mol. Gen. Genet. 197, 209-218. Mottinger, J. P., Johns, M. A., and Freeling, M. (1984a). Mol. Gen. Genet. 195, 367-369. Mottinger, J. P., Dellaporta, S. L., and Keller, P. B. (1984b). Genetics 106, 751-767. Mulder, R. J. P., Dietrich, A. J. J., Gerats, A. J. M., and Oud, J. L. (1981). Genetica 55, 111-115. Nelson, 0. E. (1968). Genetics 60, 507-524. Neuffer, M. G. (1965). Genetics 52, 521-528. Neuffer, M. G. (1966). Genetics 53, 541-549. Neuffer, M. G., Jones, L., and Zuber, M. S. (1968). “The Mutants of Maize.” Crop Science Society of America, Madison, Wisconsin. Nevers, P., and Saedler, H. (1977). Nature (London) 268, 109-115. Nowick, E. M., and Peterson, P. A. (1981). Mol. Gen. Genet. 183, 440-448. Nuffer, M. G. (1955). Science l21, 399-400. Nuffer, M. G. (1956). Maize Genet. Coop. News Lett. 30, 101. Nuffer, M. G. (1961). Genetics 46, 625-640. Nuffer, M. G. (1962). Maize Genet. Coop. News Lett. 36, 59-60. Orakwue, F. C., and Crowder, L. V. (1983). J. Hered. 74, 65-67. Orton, E. R. (1966). Genetics 53, 17-25. Orton, E. R., and Brink, R. A. (1966). Genetics 53, 7-16. Osterman, J. C., and Schwartz, D. (1981). Genetics 99, 267-273. Peacock, W. J., Dennis, E. S., Gerlach, W. L., Llewellyn, D., Lorz, H., Pryor, A. J., Sachs, M. M., Schwartz, D., and Sutton, W. D. (1983). Proc. Miami Winter Symp., 15th 20, 311-320. Pearson, 0. H. (1968). Hortic. Sci. 3, 271-274. Peterson, P. A. (1953). Genetics 38, 682-683. Peterson, P. A. (1957). Genetics 42, 388-389. Peterson, P. A. (1960). Genetics 45, 115-133. Peterson, P. A. (1961). Genetics 46, 759-771. Peterson, P. A. (1963). Proc. Iowa Acad. Sci. 70. 129-134. Peterson, P. A. (1964). Maize Genet. Coop. News Lett. 38, 80-81. Peterson, P. A. (1965a). Am. Nut. 99, 391-398. Peterson, P. A. (1965b). Genetics 52, 466. Peterson, P. A. (1966). Genetics 54, 249-266. Peterson, P. A. (1968). Genetics 59, 391-398. Peterson, P. A. (1969). Nature (London) 221, 480-481. Peterson, P. A. (1970a). Genetica 41, 33-56. Peterson, P. A. (1970b). Theor. Appl. Genet. 40, 367-377. Peterson, P. A. (1974). 1973 Brookhaven Symp. Biol. 25, 244-261. Peterson, P. A. (1976a). Mol. Gen. Genet. 149, 5-21. Peterson, P. A. (1976b). Genetics 84,469-483.

PLANT TRANSPOSABLE ELEMENTS

20 1

Peterson, P. A. (1977). I n “DNA Insertion Elements, Plasmids and Episomes” (A. Bukhari, J. A. Shapiro, and S. L. Adhya eds.), pp. 429-435. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Peterson, P. A. (1978a). I n “Maize Breeding and Genetics” (D. B. Walden, ed.), Chap. 38, pp. 601-631. Wiley, New York. Peterson, P. A. (197813). Proc. Int. Congr. Genet., 14th Nauka, Moscow Part 1,49. Peterson, P. A. (1980). Cold Spring Harbor Symp. Quant. Biol. 45, 447-455. Peterson, P. A. (1983). Maize Genet. Coop. News Lett. 57, 2. Peterson, P. A., and Friedemann, P. D. (1983). Maydica 28, 213-249. Peterson, P. A., and Weber, C. R. (1969). Theor. Appl. Genet. 39, 156-162. Pohlman, R. F., Fedoroff, N. V., and Messing, J. (1984). Cell 37,635-643. Pollard, M. (1959). “Perspectives in Virology,” Vol. 1, pp. 291. Wiley, New York. Potrykus, I. (1970). Z. Pflunzenziichtg. 63, 24-40. Pring, D. R., Levings, C. S., 111, Hu, W. W. L., and Timothy, D. H. (1977). Proc. Natl. Acad. Sci. U . S . A . 74, 2904-2908. Proudfoot, N. J. (1979). Nature (London) 279, 376. Punnett, R. C. (1922). J. Genet. 12, 255-281. Punnett, R. C. (1932). J . Genet. 26, 97-112. Punnett, R. C. (1935). J. Genet. 32, 171-177. Reddy, A. R., and Peterson, P. A. (1976). Theor. Appl. Genet. 48, 269-278. Reddy, A. R., and Peterson, P. A. (1977). Can. J . Genet. Cytol. 19, 111-117. Reddy, A. R., and Peterson, P. A. (1978). Can. J. Genet. Cytol. 20, 337-347. Reddy, A. R., and Peterson, P. A. (1983). Mol. Gen. Genet. 192, 21-31. Reddy, L. V., and Peterson, P. A. (1984). Mol. Gen. Genet. 194, 124-137. Reif, H . J., and Saedler, H. (1975). Mol. Gen. Genet. 137, 17-28. Rhoades, M. M . (1936). J . Genet. 33, 347-354. Rhoades, M. M. (1938). Generics 23, 377-397. Rhoades, M. M. (1941). Cold Spring Harbor Symp. Quant. Biol. 9, 138-144. Rhoades, M. M. (1945). Proc. Natl. Acad. Sci. U.S.A. 31, 91-95. Rhoades, M. M . , and Dempsey, E. (1982). Maize Genet. Coop. News Lett. 56, 21-26. Rhoades, M. M., and Dempsey, E. (1983). Maize Genet. Coop. News Lett. 57, 15-18. Robertson, D. S. (1978). Mutat. Res. 51, 21-28. Robertson, D. S. (1983). Mol. Gen. Genet. 191, 86-90. Rubin, G. M. (1983). In “Mobile Genetic Elements” (J. A. Shapiro, ed.), pp. 329-361. Academic Press, New York. Rubin, G. M., and Spradling, A. C. (1982). Science 218, 348-353. Saccardo, F. (1971). Caryologia 24, 71-83. Saccardo, F., and Devreux, M. (1967). Caryologia 20, 239-256. Sachs, M. M., Peacock, W. J., Dennis, E. S . , and Gerlach, W. L. (1983). Maydica 28,289303. Saedler, H., Bonas, U., Deumling, B., Gupta, M., Hahlbrock, K., Harrison, B. J., Kreuzaler, F., Peterson, P. A., Reif, J., Schwarz-Sommer, Zs., Shepherd, N . , Sommer, H., Ubben, D., and Wienand, U. (1983). I n “Genetic Rearrangement” (K. F. Chater, C. A. Cullis, D. A. Hopwood, A. A. W. B. Johnston, and H. W. Woolhouse, eds.). John Innes Symp., 5th pp. 107-1 16. Sager, R. (1951). Genetics 36, 510-540. Salamini, F. (1980a). Cold Spring Harbor Symp. Quant. Biol. 45, 457-476. Salamini, F. (1980b). Mol. Gen. Genet. 179, 497-507. Salamini, F., and Borghi, B. (1966). Genet. Agrar. 20, 239-248. Salamini, F., Brememkamp, M., and Marotta, R. (1982). Heredity 49, 1 1 1-1 15. Sand, S. A. (1957). Genetics 42, 685-703.

202

PATRICIA NEVERS ET A L .

Sand, S. A. (1969). Genetics 61, 443-452. Sand, S. A. (1976). Genetics 83, 719-736. Sastry, G. R. K. (1976). Heredity 36, 315-329. Sastry, G. R. K., Aslam, K. M., and Jeffries, V. (1980). Cold Spring Harbor Symp. Quant. Biol. 45, 477-486. Scherz, W. (1927). Z. Indukt. Abst. Vererbungsl. 45, 1-40. Schick, R. (1933). Z . Indukt. Abst. Vererbungsl. 65, 293-313. Schick, R. (1935). Z . Indukt. Abst. Vererbungsl. 69, 345-373. Schick, R., and Stubbe, H. (1932). Z . Indukt. Abst. Vererbungsl. 62, 249-290. Schwartz, D. (1973). Genetics 74, 615-617. Schwartz, D., and Echt, C. S. (1982). Mol. Gen. Genet. 187, 410-413. Schwarz-Sommer, Zs., Gierl, A., Klosgen, R. B., Wienand, U . , Peterson, P. A., and Saedler, H. (1984). EMBO J . 3, 1021-1028. Sears, B. B. (1984). Stadler Symp. 15, in press. Shapiro, J. A. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 1933-1973. Shapiro, J. A., ed. (1983). “Mobile Genetic Elements.” Academic Press, New York. Sheldon, E., Ferl, R., Fedoroff, N., and Hannah, L. C. (1983). Mol. Gen. Genet. 190,421426. Shepherd, N. S., Schwarz-Sommer, Zs., Wienand, U . , Sommer, H., Deumling, B., Peterson, P. A., and Saedler, H. (1982). Mol. Gen. Genet. 188, 266-271. Shepherd, N. S., Schwarz-Sommer, Zs., Blumberg vel Spalve, J., Gupta, M., Wienand, U., and Saedler, H. (1984). Nature (London) 307, 185-187. Sheridan, M. A., and Palmer, R. G. (1977). J . Hered. 68, 17-22. Sherratt, D. J., Arthur, A., and Burke, M. (1981). Cold Spring Harbor Symp. Quant. Biol. 45,275-282. Shure, M., Wessler, S., and Fedoroff, N. (1983). Cell 35, 225-233. Smith, H. H., and Sand, S. A. (1957). Genetics 42, 560-582. Smith, J. D. (1960). PhD thesis, Iowa State University, Ames. Spitters, C. J. T., Vosselman, L., Engels, J. M. M., Van Kester, W. N. M., and Zeven, A. C . (1975). Euphytica 24, 323-332. Spradling, A. C., and Rubin, G. M. (1982). Science 218, 341-347. Sprague, G. F., and McKinney, H. H. (1966). Genetics 54, 1287-1296. Sprague, G. F., and McKinney, H. H. (1971). Genetics 67, 533-542. Sprague, G. F., Brimhall, B., and Hixon, R. M. (1943). J. A m . SOC.Agron. 35, 817-822. Spribille, R., and Forkmann, G . (1982). Phytochemistry 21, 2231-2234. Stem, D. B., and Lonsdale, D. M. (1982). Nature (London) 299, 698-702. Straub, J. (1973). Z . Pflanzenziichtg. 70, 265-274. Strommer, J. N., Hake, S., Bennetzen, J., Taylor, W. C., and Freeling, M. (1982). Nature (London) 300,542-544. Stubbe, H. (1933). Bibl. Genet. 10, 299-356. Stubbe, H. (1935). Nachr. Ges. Wiss. Gottingen, Math. Phys. Kl. VI, N.F. 2, 57-87. Stubbe, H. (1940). Biol. Zentralbl. 60, 113-129. Stubbe, H. (1941). Z . Indukt. Abst. Vererbungsl. 79, 401-443. Stubbe, H. (1966). “Genetik und Zytologie von Antirrhinum L. sect. Antirrhinum.” Fischer. Jena. Stubbe, H. (1974). Die Kulturpflanze 22, 189-213. Stubbe, H., Berg, H., and Hagemann, R. (1981). Biol. Zentralbl. 100, 385-393. Sutton, W. D., Gerlach, W. L., Schwartz, D., and Peacock, W. J. (1984). Science 223, 1265- 1268. Terasawa, Y. (1922). Bot. Mag. Tokyo 36, 75.

PLANT TRANSPOSABLE ELEMENTS

203

Timothy, D. H., Levings, C. S., Hu, W. W. L., and Goodman, M. M. (1983). Maydica 28, 139-149.

Toxop6us, H. J. (1928). Genetica 9, 341. Tsai, C. Y . (1974). Biochem. Genet. 11, 83-96. Tuschall, D. M.,and Hannah, L. C. (1982). Genetics 100, 105-111. Van Schaik, N. W., and Brink, R. A. (1959). Genetics 44, 725-738. Vaughn, K. C., and Wilson, K. G. (1980). J. Hered. 71, 121-123. Vodkin, L. O., Rhodes, P. R., and Goldberg, R. B. (1983). Cell34, 1023-1031. de Vries, H. (1901/1903). “Die Mutations Theorie.” Veit, Leipzig. Weck, E., Courage, U., Doring, H.-P., Fedoroff, N., and Starlinger, P. (1984). EMBO J. 3, 1713-1716.

Weissinger, A. K., Timothy, D. H., Levings, C. S., 111, Hu, W. W. L., and Goodman, M. M. (1982). Proc. Natl. Acad. Sci. U.S.A. 79, 1-5. Weyers, W. H. (1961). Genetics 46, 1061-1067. Wienand, U., Sommer, H., Schwarz, Zs., Shepherd, N., Saedler, H., Kreuzaler, F., Ragg, H., Fautz, E., Hahlbrock, K., Harrison, B. J., and Peterson, P. A. (1982). Mol. Gen. Genet. 187, 195-201. Wienand, U.,Peterson, P. A., and Saedler, H. (1983). Maize Genet. Coop. News Lett. 57, 157- 158.

Williams, E., and Brink, R. A. (1972). Genetics 71,97-110. Williams, W. M., Satyanarayana, K. V., and Kermicle, J. L. (1984). Genetics 107,477-488. de Winton, D., and Haldane, J. B. S. (1935). J . Genet. 31, 67-100. Wood, D. R. and Brink, R. A. (1956). Proc. Natl. Acad. Sci. U.S.A. 42, 514-519.

This Page Intentionally Left Blank

The Dinoflagellate Chromosome

D. C . SIGEE Departments of Botany and Zoology University of Manchester Manchester, England

I. Introduction ......................................... 11. Class Dinophyceae-General 111. DNA Levels and Chromosome Numbers A. Chromosome Numbers .............................................. B. Chromosome DNA Levels ................................... ure . . . . . . . . . . . . . . IV. Dinoflagellate Chromosome Fi A. Light Microscopy.. ..... B. Electron Microscopy . . . . . . . . . . . . V. DNA Transcription and Chro VI. Chromosome Changes during the Cell Cycle .............................. A. Synthesis of Nuclear D N A . . .......................... B. Changes in Chromosome Fine Structure.. ............................. VII. Macromolecular Composition of Dinoflagellate Chromatin . . . . . . . . . . A. Proteins ................................................ B. DNA ...................................................... C. RNA .......................... VIII. Ionic Composition of Di A. X-ray Microanalysis ............... B. Autoradiography .................................. C. Cations and Chromatin Stabilisation ................... IX. General Discussion ................................... A. Dinoflagellate Chromosome Models . . . . . . . . . . . . . . . . . . B. Prokaryote and Eukaryote Affinities ................. C. Intermediate Status and Phylogeny.. . . . . . . . . . . . . . . . . . References ....................................

209 209

222 225

238

248

254

205 ADVANCES I N BOTANICAL RESEARCH, VUL. 12

Copyright 8 1986 by Academic Press Inc. (London) Ltd. All rights of reproduction in any form reserved.

206

D. C. SIGEE

I. INTRODUCTION

Dinoflagellates are single-celled organisms, typically highly motile, and occur as major constituents of both marine and freshwater phytoplankton. As such, they are widespread over the earth’s surface, and have considerable potential importance in aquatic food webs. In addition to this general ecological importance, dinoflagellates are of considerable interest to biologists for a number of specific reasons. One peculiar and well-known feature of marine dinoflagellates, for example, is their ability to become locally abundant under certain conditions (Anderson and Morel, 1978), forming red tides (Murphy et al., 1975) and producing high levels of toxins which kill fish and cause serious illness in humans that consume the poisoned sea food (Braarud, 1955; Tangen, 1977; White, 1978; Carmichael, this volume). Dinoflagellates are also of particular interest to palaeontologists, since they have an abundant fossil record dating back to the Late Triassic, and a single genus, Arpylorus, has been identified in the Late Silurian (Calandra, 1964; Sarjeant, 1978). Dinoflagellate cysts provide useful stratigraphical markers (Brasier, 1980), and the organisms have been major contributors to deposits of liquid fossil fuels. The ancient lineage of this group places its origins right back to the early development (and possible origins) of the eukaryote cell, and raises perhaps the most interesting question that surrounds these organisms-their phylogenetic position within the early eukaryote sequence. Dinoflagellates have also attracted the attention of cell biologists, particularly in relation to the unusual nature of the cell nucleus. This has remarkably high levels of DNA compared to other eukaryote cells (Loeblich, 1976), and is of considerable interest in terms of its fine structure. Early light microscope studies by Entz (1921, 1927), van Goor (1918), and Kohler-Wieder (1937) suggested that the interphase nucleus contained chromatic granules within an achromatic reticulum. Subsequent transmission electron microscope studies by Grell and WollfarthBotterman (1957) and Grasse and Dragesco (1957) showed that the chromatic granules seen previously were chromosomes, which were condensed throughout the cell cycle. In addition to being permanently condensed, dinoflagellate chromatin is also unusual in having a narrow (3-6 nm) diameter to the constituent fibrils (Giesbrecht, 1962; Bouligand et al., 1968) characteristic of naked DNA (Ris, 1962), whirls of fibrils similar to those seen in bacterial nucleoids (Giesbrecht, 1962), permanent attachment of the chromosomes to the nuclear membrane (Soyer and Haapala, 1974a), absence of centromeres and a conventional mitotic spindle (Kubai and Ris, 1969), continuous replication of nuclear DNA (Filfilan and Sigee, 1977; Galleron and Durrand, 1979) and low levels of basic

T H E DINOFLAGELLATE CHROMOSOME

207

protein (Ris, 1962; Dodge, 1964; Rizzo and Nooden, 1973, 1974a) with an apparent absence of histones (Rizzo and Nooden, 1974b). All of these characteristics show some similarity to the situation seen in prokaryote chromatin (Giesbrecht, 1962; Ris, 1962; Dodge, 1965, 1966), and provide support for the initial suggestion of Ris (1962) that the nuclear organisation of dinoflagellates was intermediate between prokaryotes and eukaryotes. Dodge (1965) has since proposed that dinoflagellates should be placed in a separate group-the Mesokaryota. Other features of dinoflagellate chromatin, such as the presence of repeated and highly complex DNA (Allen et al., 1975a), are typical of normal eukaryote cells, while the presence of substantial amounts of hydroxymethyluracil (Rae, 1976) and an unusual RNA polymerase (Rizzo, 1979) appear to be unique to this group. The absence of histones, normally important in the stabilisation of eukaryote chromatin, is of particular interest. This has been related to the unusually high levels of chromatin-associated cations seen in these cells (Kearns and Sigee, 1979, 1980), which may assume the stabilising role normally occupied by the basic (histone) proteins (Sigee, 1982a, 1984a). The purpose of this article is to review these various aspects of dinoflagellate chromosomes, and their constituent chromatin, within the general context of the biology of dinoflagellate cells. The chromosomes of these organisms will be compared to the chromosomes of other eukaryote cells and to bacteria. These comparisons lead to the conclusion that dinoflagellate chromosomes are highly unusual, if not unique, and have phylogenetic implications for the origin of the group and the evolutionary strategy that has given rise to present-day species. 11. CLASS DINOPHYCEAE-GENERAL

CHARACTERISTICS

Dinoflagellates have a number of distinct structural and biochemical characteristics which place them in the class Dinophyceae (Parke and Dodge, 1976), within the phylum Pyrrophyta. Although this phylum has been considered by some taxonomists to contain a multiplicity of classes (see for example Bold and Wynne, 1978), a simpler scheme (Loeblich, 1976; Bujak and Williams, 1981) divides the phylum into two major classesthe Dinophyceae and Syndiniophyceae (containing the single genus Syndinium). Among the characteristics that unite the dinoflagellates into a single group are the possession of (1) two distinct types of flagella, one of which is naked, the other with a single row of long, fine hairs (Leadbeater and Dodge, 1967; Taylor, 1975); (2) a complex type of cell covering-the theca or amphiesma (Dodge and Crawford, 1970); (3) a large and distinc-

208

D. C. SIGEE

tive (mesokaryotic) nucleus; (4) distinctive chloroplasts (Dodge, 1975); (5) ejectile trichocysts, and (6) a pair of complex osmoregulatory organelles termed pusules (Dodge, 1972). Biochemically, dinoflagellates are characterised by the possession of chlorophylls a and d , the carotenoids fucoxanthin and peridinin, and the sterol dinosterol-which is unique to dinoflagellates and may be used as a chemical marker for the group (Jones et al., 1983). Against these unifying characteristics, it is clear that dinoflagellates form a remarkably diverse group of organisms. This is true whether one considers cell structure, mode of nutrition, life cycle, or biochemical characteristics. In terms of structure, there is diversity in the position of flagella, presence or absence of a hard cell wall, variation in the position and number of thecal plates, and in the structure of the pyrenoids. Variations in the life cycle are shown by differences in the duration of the biflagellate phase, the observed occurrence of sexual reproduction in only a few species, and the formation of cysts in only 10% of living species (Bujak and Williams, 1981). The mode of nutrition varies from eutrophic to heterotrophic in free-living organisms, and many non-free living dinoflagellates occur as symbionts and parasites (Loeblich, 1976; Leutenegger, 1977). Variations in biochemical characteristics include the separate occurrence of fucoxanthin and peridinin in different members of the group. The overall picture of the Dinophyceae is thus that of a well-defined taxonomic group containing a wide diversity of organisms. This is consistent with the ancient origins of the group, and the evolutionary diversification that has taken place since Palaeozoic times. Amidst this diversity the genus Oxyrrhis is of uncertain taxonomic status (Triemer, 1982). Although it has various dinoflagellate characteristics, it is atypical in its mode of cell division and in the structure and high basic protein content of its chromosomes. For these reasons this genus will not be included within this review.

111. DNA LEVELS AND CHROMOSOME NUMBERS One important characteristic of dinoflagellates is the presence of high levels of nuclear DNA. This has been noted previously by numerous authors (see for example Loeblich, 1976) and is shown in Table I, where mean DNA levels per nucleus are given for various dinoflagellates, lower and higher eukaryotes, and a prokaryote. The dinoflagellate DNA levels shown in Table I are for logarithmically growing cells. These are typically twice the levels obtained for stationary

THE DINOFLAGELLATE CHROMOSOME

209

phase cultures and represent the replicated haploid (2n) level of DNA. Considerable variation occurs between species, ranging from 3.2 pgkell for Amphidinium to about 200 pgkell for Gonyaulax. There may also be considerable variation between individual cells of the same species, as shown by the cytophotometric studies of Gavrila et al. (1981) on Peridinium tabulatum. The general levels of DNA occurring in dinoflagellates are clearly much greater than might be expected for a unicellular organism, and are within the range of levels typical of higher eukaryote cells. These high DNA levels relate both to large numbers of chromosomes and high DNA levels per chromosome. A . CHROMOSOME NUMBERS

Chromosome numbers in dinoflagellates are frequently difficult to determine, due to problems in cytological preparation (Holt and Pfeister, 1982) and the large numbers encountered. The range shown in Table I, largely for marine dinoflagellates, varies from 24-1 12 chromosomes per cell. A recent survey of Indian freshwater dinoflagellates (Shyam and Sarma, 1978) gives comparable values, ranging from 56 (Gymnodinium indium) to 220 (Woloszynskia hiemale). These chromosome numbers are considerably higher than those normally occurring in both lower and higher eukaryotes (Table I). Although dinoflagellates are generally regarded as being haploid organisms (see Section VII,B, l), certain genera-particularly Gymnodinium and Peridinium, show evidence of a polyploid or aneuploid progression (Shyam and Sarma, 1978; Holt and Pfeister, 1982). This may account for some of the high chromosome numbers encountered in this group. The variable counts obtained in cultures of some species, for example Crypthscodinium cohnii (Beam et al., 19771, suggest some degree of genetic redundancy. B. CHROMOSOME

DNA

LEVELS

Data on mean levels of DNA in dinoflagellate chromosomes (Table I) are fragmentory , but available evidence suggests they are more comparable to higher eukaryotes rather than lower eukaryotes in this respect. Dinoflagellate chromosomes do not appear, however, to have as much DNA as some of the larger eukaryote chromosomes (e.g. Vicia, and Lilium), and the DNA level is very much less than that of Dipteran polytene chromsomes (e.g. Chironomus). This is particularly interesting in view of the proposal by various workers (see Section IX,A,2) that dinoflagellate chromosomes are polytenic in constitution.

210

D. C. SIGEE

TABLE I Mean DNA Levels in Nuclei and Chromosomes-Comparisons between Dinoflagellates and Other Organisms"

Species Dinoflagellates Prorocentrum micans Amphidinium carterae Gymnodinium breve Gymnodinium nelsonii Gyrodinium resplendens Crypthecodinium cohnii Cachonina niei Gonyaulax polyedra Scrippsiella trochoidea Bacterium Escherichia coli Lower eukaryotes Euglena gracilis (n) Saccharomyces cerevisiae ( 2 4 Neurospora crassa ( n ) Higher eukaryotes Lilium longiflorum ( 4 4 Vicia faba ( 2 4 Lactuca sativa ( 2 4 Drosophila melanogaster ( 2 4 Chironomus sp. (2n) Chironomus (polytene) Homo sapiens ( 2 4

DNA level (pdcell)

Referenceb

Number of chromosomes

Referenceb

DNA level (pg/ chromosome)

65-69 24-46

15 16

0.63 0.13

4,5,17

0.07

42.0 3.2 101.6 143.0 66.0

I 2 3 3 4

6.9

5,6

7.6 200.0 34.0

7 2 6

111-112

18

0.07

44 80-90

15 19

0.40 0.40

0.0044

7

I

24

0.0044

3.0 0.046

8 9

45 20

20 22

0.07 0.002

0.05

10

7

23

0.007

313.4 60.5 14.5 I .o

II 11 11 12

48 12 18 8

21 21 21 12

6.5 5.0 0.8 0.125

0.43 6720.0 5.6

13 13 14

8 4 46

13 13 14

0.054 1680.OO 0.12

99-1 10

The mean DNA levels given for dinoflagellatesare for log-phase cultures. Chromosome numbers in the other part of the table are the haploid, diploid, or tetraploid count-equivalent to the ploidy quoted for the species. Mean levels of DNA (pg) per chromosome are clearly very approximate, particularly in the dinoflagellates, where DNA levels and chromosome numbers are not known with certainty. References: ( 1 ) Haapala and Soyer (1974a); (2) Holm-Hansen (1969); (3) Kim and Martin (1974);(4) Allen et al. (1975a); (5) Roberts et al. (1974); (6) Rizzo and Nooden (1973); (7) Loeblich (1976); (8) Parenti el al. (1967); (9) Bhargava and Halvorson (1971); (10) Holliday (1970); (11) McLeish and Sunderland (1961); (12) Beermann and Pelling (1965); (13) Daneholt and Edstrom (1967); (14) DuPraw (1970); (15) Dodge (1963a); (16) Grasse and Dragesco (1957); (17) Kubai and Ris (1969); (18) Loeblich (1968); (19) Fine and Loeblich (1974); (20) Leedale (1967); (21) Darlington and Wylie (1955); (22) Hawthorne and Mortimer (1960); (23) Fincham and Day (1971); (24) Dyson (1978).

THE DINOFLAGELLATE CHROMOSOME

21 1

The information presented in this section suggests that although both chromosome size and number contribute to the high DNA levels seen in dinoflagellate nuclei, it is the large number of chromosomes which constitutes the most consistent and unusual feature of these cells.

IV. DINOFLAGELLATE CHROMOSOME FINE STRUCTURE Dinoflagellate chromosomes are normally rod-shaped structures, varying in length from about 0.75-10.0 pm. Occasionally they are smaller than this; Shyam and Sarma (1978), for example, report the presence of small (0.2-0.5 pm diameter) spherical chromosomes in a species of Gymnodinium. Dinoflagellate chromosomes have been investigated by both light and electron microscopy. A . LIGHT MICROSCOPY

The most direct way to observe dinoflagellate chromosomes is in intact cells. Although they are not normally visible within the nuclei of living cells by bright field or phase-contrast microscopy they have been observed by interference microscopy. Livolant (1978) noted a marked positive birefringence in the chromosomes of Prorocentrum micans, indicating a preferential orientation of the DNA filaments normal to the chromosome axis within the living cell. Dinoflagellate chromosomes can be observed in stained, fixed, whole-cell preparations, and the observation of ‘chromatic granules’ in such preparations by early microscopists has already been noted. Squash preparations have been routinely used for chromosome counts (Shyam and Sarma, 1978; Loper et al., 1980; Holt and Pfeister, 1982) and typically show the individual chromosomes to consist of two longitudinal threads which are intertwined in a double helix (Grasse and Dragesco, 1957; Grasse et al., 1965b; Dodge, 1966), as shown in Fig. 4b. Grasse and Dragesco (1957) initially used the term ‘chromonema’ to describe the major spiral units of the chromosomes. Whether these chromonema correspond to the chromatids seen in normal eukaryote metaphase chromosomes is a question of major importance, and will be considered subsequently in the account of electron microscopy studies. Haapala and Soyer (1974b) examined both fixed and unfixed preparations of Prorocentrum micans using various ‘banding techniques. ’ They found that these chromosomes did not have any longitudinal differentiation, and were not able to demonstrate Q, G, or C bands. The absence of G bands demonstrates fundamental differences between dinoflagellate

212

D. C . SIGEE

and normal eukaryote chromosomes, since the latter normally have welldefined alternating regions of A-T base pair-rich and G-C base pair-rich DNA (Jorgensen et al., 1978). The absence of centromeric chromatin (C) bands is consistent with the observed absence of well-defined centromeres, and shows that the attachment of dinoflagellate chromosomes to the nuclear membrane is not mediated by a distinct type of heterochromatin. Light microscope observation of sections of fixed, embedded cells typically reveals a prominent nucleus containing nucleolus and numerous discrete chromosomes. These chromosomes typically appear more condensed than in the squash preparations, and in cells fixed with glutaraldehyde/osmium tetroxide they are seen as single threads along which a fine banding pattern may be apparent (Figs. l a and 1Oa). B . ELECTRON MICROSCOPY

Three main experimental approaches have been used in the preparation of dinoflagellate chromosomes for observation under the electron microscope: fixation and sectioning, cryotechniques, and whole-mount observation of chromosomes and chromatin.

1 . Fixation and Sectioning In ultrathin sections of interphase nuclei the chromosomes are typically sectioned in all planes due to random orientation (Fig. lb). The fine structural detail that is observed in ultrathin sections depends very much on the fixation procedure. With 2-hr fixation in glutaraldehyde followed by osmium tetroxide the chromosomes appear electron dense, and have clear transverse banding (Fig. Ib). Increase in the duration of the aldehyde fixation results in a removal of electron-dense protein matrix, revealing a superstructure of chromatin DNA fibrils. The effect of fixation time on observed chromosome fine structure is shown by comparing Fig. l a with Fig. 2a (Glenodinium foliaceum) and Fig. 9 with Fig. 6 (Prorocentrum micam)-and has been correlated with changes in chromatin elemental composition (Sigee and Kearns, 1982a; Sigee, 1983a). The appearance of a dinoflagellate chromosome in longitudinal ultrathin section is shown diagrammatically in Fig. 4c, corresponding to the micrographs shown in Figs. 2a and 6. The chromatin fibrils occur as periodic, transversely oblique arrays (bands), separated by a series of nested arcs. The arrangement of the DNA fibrils which leads to the above appearance under the electron microscope has been the subject of much discussion, with major models being proposed by Grasse and Dragesco (1957), Giesbrecht (1961, 1965), and Grasse et al. (1965a,b) on the basis of ultrathin sections. Some discussion of the details of fibril arrangement will

THE DINOFLAGELLATE CHROMOSOME

213

Fig. 1. Fine structure of the binucleate dinoflagellate Glenodinium foliuceum, fixed in glutaraldehyde (4 hr) followed by osmium tetroxide (2 hr). (la) Light microscope (LM) phase-contrast view of whole cell in 2-pm-thick section, showing dinocaryotic nucleus (d) and lobes of the supernumerary nucleus (s). ( x 2800.) ( I b) Transmission electron microscope (TEM) view of dinocaryotic nucleus, showing large nucleolus (nu) and numerous chromosomes (C) sectioned in various planes. ( X 14,000.)

214

D. C. SlGEE

be presented later, but the major bands of fibrils probably correspond to the major spirals (chromonemata) seen under the light microscope (Fig. 4). Evidence for this is provided by observation of serial sections (Oakley and Dodge, 1979), where the pitch of the oblique transverse bands changes direction from the one side of the chromosome to the other. Examination of the edge of sectioned chromosomes shows that some of the fibrils extend into the nucleoplasm as fine filaments (Fig. 2b). The presence of these filaments, noted previously by Bouligand et al. (1968), Soyer (1969), and Soyer and Haapala (1973), differentiates the dinoflagellate chromosome into two main regions-a central condensed core surrounded by a halo of peripheral fibrils. The functional distinction between these regions will be discussed in Section V. Visualisation of the well-defined arrangement of the chromatin fibrils in the main body of fixed, chemically dehydrated chromosomes is dependent on the stabilising effect of osmium tetroxide, with or without additional aldehyde fixation. Although the general transverse arrangement of fibrils in osmium tetroxide-fixed cells agrees with the orientation deduced by Livolant (1978) in living cells, it should not be assumed that the detailed arrangement of fibrils seen under the electron microscope, with aggregation of 3-nm fibrils into strands of varying thickness, exactly reflects the in uiuo situation. Livolant and Bouligand (1978), using both conventional and high voltage TEM, have suggested that DNA filaments are not genuinely bent to form arcs. They propose that the arcs are an illusion created by overlapping layers of straight filaments, which are normal to the chromosome axis (see Section IX,A,3). In the absence of osmium tetroxide the retention of fine structural detail in dinoflagellate chromosomes is very much reduced. Fixation in glutaraldehyde alone causes gross precipitation of chromatin, which appears as irregular dense bands separated by electron-transparent spaces (Fig. 2c). Fixation in the coagulative fixative acetic/alcohol results in a uniformly dense appearance to the central core, with peripheral fibrils radiating out into an electron-transparent nucleoplasm (Fig. 2d). Variations in Chromosome Structure. Ultrathin sections of glutaraldehyde/osmium tetroxide fixed chromosomes do not always appear as described above (Fig. 4c). Variations occur, particularly in relation to the cell cycle, and include the following: 1. The presence of central axial structures. These have been reported in various dinoflagellates, particularly during cell division, including Amphidinium carterae (Babillot, 1969), Glenodinium foliaceurn (Dodge, 1971), Prorocentrum micans (Soyer, 1977a), and Oodinium dogieli (Cachon and Cachon, 1977). In Peridinium cinctum, axial fibrils are present during the period of chromosome uncoiling in mid-interphase (Spector et al., 1981a; Spector and Triemer, 1981).

Fig. 2. EM details of longitudinally sectioned chromosomes of GIenodinium foliaceum, fixed in various ways. All sections have been stained with uranyl acetate followed by alkaline lead citrate. (a) Glutaraldehyde/osmium tetroxide fixed cell. The arrows mark the position of bands of oblique fibrils which correspond to feature x shown in Fig. 4c. ( ~ 4 3 , 0 0 0 . )(b) Detail from edge of chromosome, showing fine peripheral fibrils (arrows) extending into the surrounding nucleoplasm. (x87,OOO.) (c) Fixation (4 hr) in glutaraldehyde only. The chromatin is precipitated as coarse bands. (x43,OOO.) (d) Fixation (4 hr) in acetic/ alcohol. The coagulative fixative has caused gross precipitation of the chromatin, with peripheral fibrils radiating out into the dense nucleoplasm. ( ~ 6 2 , 0 0 0 . )

216

D. C. SlGEE

2. Chromosome uncoiling and fragmentation. This has been described in various dinoflagellates, at different stages of interphase, and has been related to DNA replication (Spector er 01.. 1981a,b), DNA transcription (Spector and Triemer, lPSl), and the onset of cell division (Filidan and Sigee, unpublished observations). These different aspects will be considered later in the article.

2. Cryotechniques Two main cryotechniques have been used in the examination of dinoflagellate chromosomes-cryosectioning and freezeetching. The transmission electron microscope appearance of a 0.5 t o I-pm-thick fkeze-dried cryosection of Glenodiniumfoliaceum is shown in Fig. 3c. The chromosomes are highly electron dense, with no chromatin fibrils visible, either in the central core or at the periphery. The effect of briefly washing thick cryosections (which had been collected by a sucrose drop technique) is sbown in Fig. 3b. This procedure causes break-up and dispersion of the cryosections, but the chromosomes remain as groups, united by a system of interchromosomal strands (Kearns and Sigee, 1980). These strands probably represent peripheral chromatin fibrils that have been exposed by removal of tbe nucleoplasm during washing. Demonstration of fibrils within the main body of the chromosome requires thinner (lO@nm-tbick) cryosections (Fig.3a), and reveals a general transverse orientation similar to that deduced previously in fixed and living cells (Kearnsand Sigee, 1980). Freezeetch preparations of dinoflagellate cells reveal clear chrome some proides within the nucleus, though the ability to demonstrate chre math fibrils seems variable. This may relate to the metbod of cell preparation (e.g. chemical treatment, extent of etching). Studies by Kearns (1981) on unfixed cells of Amphidinium carterae did not show clear fibrillar structure, though clear banding was observed in tbe chromosome contours. In contrast to this, the published observations of Giesbrecbt (1965) and Soyer and Escaig (1980), using unfixed and glutaraldebyde-fixed cells, reveal a well-ordered array of internal fibrils similar to those observed in ultrathin sections of osmium tetroxide-fixed cells. The results obtained from freeze-etched preparations and ultrathin cryosections thus support and complement the general body of information obtained from chemically treated (fixed, dehydrated, and embedded) cells. In addition to the two major cryotechniques, others have also been used, including resin-infiltration of frozendried cells (Sigee and Kearns, 1982b) and freeze-substitution (Soyer, 1981). The latter has demonstrated the presence of numerous protein cables in tbe nucleoplasm between the chromosomes, whicb may be important in intranuclear movement.

Fig. 3. EM appearance of freeze-dried(a and c) and airdried cryosections. The section thickness quoted is the estimated thickness in the freshly cut frozen state. This will be reduced considerably during dessication. (a). Prorocentrum micans: 100-nm-thickcryosection. Detail of single chromosome showing clear transverse bands of fibrils (iidicated by bars). (x36.000.) (b) Prorocentrum micans: 1-pm-thick cryosection, collected on a sucrose drop and briefly washed in distilled water. C h m m m e profiles are interconnected by a reticulum of fibrils (0 wbich probably represents peripheral DNA strands. (x 18,700.) (c) Gledinium foliaceum: 0.5- to I - p n thick cryosection. The rippled appearance to the section (chatter) suggests that the section has dried completely in the frozen state, with full retention of soluble constituents. The section shows nucleolus (nu) and dense chromosomes (C),which show a clear banding pattern (indicated by bars in C').(x 15,000.)

218

D. C. SIGEE

3 . Whole-mount Observation of Chromosomes and Chromatin The isolation and observation of intact chromosomes and chromatin fractions has provided valuable information on both the gross organisation of dinoflagellate chromosomes and the fine structure of the native fibril. a . Structure of whole-mount chromosomes. Various techniques have been used to obtain preparations of isolated dinoflagellate chromosomes, including cell ultrasonication followed by spreading of contents onto distilled water (Haapala and Soyer, 1973; Soyer and Haapala, 1974b), cell lysis by Triton X- 100 followed by deposition on hydrophilic carboncoated grids (Hamkalo and Rattner, 1977), and direct bursting and collection of cells from a water surface (Oakley and Dodge, 1979). The major advantage in looking at chromosomes in isolation is that the supercoiling of the DNA superstructyre is relaxed, thus allowing a more ready interpretation of the arrangement of the fibrils within the chromosome. This interpretation is further facilitated by a considerable variation in the degree of relaxation observed, either within a single preparation (Haapala and Soyer, 1973) or by controlled alteration of the duration of spreading on the water surface (Oakley and Dodge, 1979). The initial micrographs published by Haapala and Soyer (1973) demonstrated two major features of the dinoflagellate chromosome. Firstly, they are composed of two major bundles of fibrils, arranged in spiral coils of opposite, i.e. left-handed (anorthospiral) and right-handed (orthospiral) directions. Secondly, the constituent fibrils form loops rather than free ends at the tips of the bundles, suggesting continuity from one bundle to the next. The oppositely coiled double-strand configuration can clearly arise by longitudinal alignment and twisting of a single loop, and was the basis of the chromosome model proposed by Haapala and Soyer (1973). In support of this, they demonstrated the presence of large circular DNA molecules (referred to as chromatids), and suggested that these were twisted to give a sequence of ‘figures of 8,’ where each figure of 8 represents a single turn of the double helix. Haapala and Soyer estimated that 1300 circular DNA molecules were present per chromosome of Prorocentrum micans, and 700 in Gyrodinium, and suggested that these molecules were arranged in 3 dimensions to give ‘balls of chromatin’ (Fig. 17a). Subsequent work by Oakley and Dodge (1979) has supported the observations of Haapala and Soyer (1973) in demonstrating continuity of bundles of fibrils at the ends of the chromosomes. Their interpretation of chromosome structure differs from that of Haapala and Soyer in suggesting that the aggregation of fibrils into two major bundles was a genuine in viuo state, and their proposed arrangement of these bundles as toroidal chromonema (Fig. 4a) follows the arrangement of the individual chromatids originally suggested by Haapala and Soyer (1973). Diagrammatic representations of the appearance of whole-mount chromosomes, and the

T H E DINOFLAGELLATE CHROMOSOME

219

Fig. 4. Light and electron microscope appearance of the dinoflagellate chromosome. Diagrammatic representation of (a) the toroidal chromonema model of chromosome structure, taken, with permission, from Oakley and Dodge (1979). In this model, the chromosome consists of a band of DNA fibres (the chromonema) coiled upon itself in a double spiral. (b) Light microscope appearance of isolated chromosome, with two major spirals of the chromonema. (c) T E M appearance of chromosome in section (resin or cryosection) or in freezeetch preparation, with bands of obliquely arranged fibrils ( x ) alternating with regions of nested arcs (y). (d) TEM view of whole-mount preparation of dividing chromosome, showing the two major spirals of the chromonema along the length of the chromosome axis. Diagrams b, c, and d are general interpretations taken from various publications (see text).

double-helically coiled toroidal chromonema model of Oakley and Dodge (1979) are shown in Fig. 4d and a, respectively. Studies of whole-mount interphase chromosomes thus suggest that the main strands (chromonema) of dinoflagellate chromosomes are part of the same looped structure, and are therefore quite different from the sister chromatids of normal eukaryote chromosomes. Further evidence for this

220

D. C. SIGEE

is provided by the observations of Soyer and Haapda (1974b)on dividing chromosomes of whole mounts, where the daughter chromosome segments have the same structure as the parent chromosome, and are not simply formed by lateral separation of the chromonema. 6. Structure of Chromatin Fibrils. Detailed observations on dispersed chromatin fibres were first camed out by Harnkalo and Rattner (Ian),who reported the presence of smooth threads of uniform diameter (5.5-6.5 nm) in Prorocentrum micans with no evidence of beaded fibres (i.e. DNA arranged as nucleosomes). The absence of nucleosomes has since been reported for other dinoflagellates, including Crypthecodiniurn cohnii (Rizzo and Burghardt, I980), Gymnodinium nelsoni (Rizzo and Burghardt, 1982), and Gymnodinium breoe (Rizzo er al., 1982). Absence of nucleosomes in Prorocenfrum micans has been confirmed by Herzog and Soyer (1981), who also showed that digestion of dinoflagellate nuclei with micrococcal endonuclease or DNase I did not lead to partially digested DNA fragments (i.e. nucleosomal DNA sequences). Nucleosomes have been reported for the binucleate dinoflagellate Peridinium balticum (Rizzo and Burghardt, 1980), where they are considered to be derived from the eukaryotic (endosymbiont) nucleus. Recent studies by Herzog and Soyer (1983)on the compaction of chromatin in whole-mount preparations has suggested a complex helical architecture with five distinct organisational levels. It should not be forgotten, however, that the extraction of chromosomes from intact cells will inevitably lead to the loss of key structural components such as proteins and cations (see Section VIII), and the various associations between DNA fibrils in whole-mount preparations may arise, in part at least, as an artefact of preparation.

V. DNA TRANSCRIPTION AND CHROMOSOME STRUCTURE Transcription of both rRNA and other RNAs appears to be closely related to chromosome structure in dinoflagellate nuclei. The occurrence and formation of nucleoli in dinoflagellate nuclei has been reviewed by Rae (1970)and Dodge (1971).The nucleoli of these cells appear to resemble those of typical eukm-yotes in comprising two major zones, an inner fibrous zone and outer granular zone (Rae, IWO),and are typically associated with strands of condensed chromatin. These strands of nucleolar associated chromatin have a fibrillar structure typical of intact chromosomes (Fig. Sa) and presumably represent regions of chromosome fragmentation or extension. Autoradiographic studies on incorporation of [3H]adenine in Prorocentrum micans (Sigee, 1984b) show that

Fig. 5. RNA transcription in Proroce~riunmicans. Hi&-resolution EM autoradiographs of r3H]adenineincorporation in stationary culture cells, fixed in glutaPaldehyde (12 hr) and osmium tetroxide (2 hr). For further details see Sigee (1984b). (a) MicdoCX development. Silver grains occur over the nucleoh region (nu) but not nucleolar associated condensed chromatin (C'). Surrounding chromosomes (C) have few grains. (~25,400.) (b) Physical development. The fine silver grains are almost entirely conlined to the nucleoplasm and are absent from the condensed main body of the chromosome. (~57,000.)

222

D. C. SlGEE

labelling of nascent nucleolar RNA (rRNA) in stationary phase cells is a highly active process. The resulting preparatipns (e.g. Fig. 5a) have a high frequency of silver grains over the main bedy of the nucleolus, but few grains directly over the condensed nucleolar chromatin. Transcription of other RNAs occurs in relation to non-nucleolar chromatin. The presence of peripheral (sometimes referred to as extrachromosomal) chromatin fibrils, extending from the main body of the chromosome into the surrounding nucleoplasm, has already been noted. Electron microscope autoradiographic evidence suggests that DNA transcription occurs entirely on these fibrils, and that condensed DNA in the main body of the chromosome is not transcriptive. This evidence comes initially from the work of Babillot (1970) who showed that incorporation of [3H]uridine into the nucleus of Amphidinium carterae was largely absent from the main body of the chromosomes. This has been subsequently confirmed by the high-resolution autoradiographic studies on the incorporation of [3H]adeninein stationary cultures of Prorocentrum micans mentioned earlier (Sigee, 1984b). In these cultures, where nuclear DNA synthesis was not apparent, the incorporation of adenine into RNA was restricted to the nucleolus (see previously) and the nucleoplasm (Fig. 5b). The conspicuous absence of silver grains in these preparations from any part of the main body of the chromosome suggests that transcription takes place only on chromatin fibrils extending into the nucleolus and nucleoplasm. On this basis, the chromosomes of dinoflagellates consist of two main regions-a central body of condensed, genetically inactive DNA, and a peripheral zone of diffuse, genetically active DNA. It is possible that enhanced transcription of chromosome DNA may occur during the phases of chromosome fragmentation observed by Spector et al. (1981a,b) and Spector and Triemer (1981), though there is no autoradiographic or biochemical evidence as yet to support this. VI. CHROMOSOME CHANGES DURING THE CELL CYCLE Although dinoflagellate chromosomes remain at least partially condensed throughout the cell cycle, alterations do occur in their activity, as shown by certain biochemical changes (notably DNA synthesis) and also changes in fine structure. A. SYNTHESIS OF NUCLEAR

DNA

The synthesis of nuclear DNA in dinoflagellates has attracted considerable interest since the initial prediction of Dodge (1966) that it would resemble prokaryotes in being continuous throughout the cell cycle.

T H E DlNOFLAGELLATE CHROMOSOME

223

Experiments to investigate this have involved a variety of approaches, both in terms of experimental material and the technique used to monitor DNA increase, and are summarised in Table 11. The majority of investigators have analysed DNA synthesis in bulk populations of cells, using techniques such as spectrophotofluorometry (Allen et al., 1975b; Karentz, 1983) or scintillation counting (Franker, 1971; Franker et al., 1974; Galleron and Durrand, 1979; Spector et al., 1981a). These bulk sample studies generally involve the use of synchronous cell populations, obtained either by selection synchrony (Franker, 1971; Franker et al., 1974) or induction synchrony (Loeblich, 1976; Galleron and Durrand, 1979; Spector et al., 1981a). Karentz (1983) has recently investigated DNA levels in phased cultures of marine dinoflagellates over multiple light/dark cycles. The majority of investigations using bulk analysis have measured increases in the level of total DNA, without distinguishing between the nuclear and cytoplasmic components. Galleron and Durrand (1979), however, monitored changes in both nuclear and plastid DNA in their studies on Amphidinium carterue, obtaining closely similar results for the two constituents. The alternative approach to bulk sample analysis is to determine DNA changes in individual cells, using techniques such as autoradiography (Filfilan and Sigee, 1977). This approach has the advantage that synchronised cell populations are not required, and the location of the newly synthesised DNA within the cell is precisely known. Various radioactive DNA precursors have been used in scintillation counting or autoradiography. In some cases (Franker, 1971; Galleron and Durrand, 1979; Spector et al., 1981a) this has been radioactive thymidine, though Allen et al. (1975b) and Filfilan and Sigee (1977) have noted that some dinoflagellates do not readily incorporate this molecule into their DNA. Alternative precursors have included 32P-orthophosphate(Franker et al., 1974), [3H]adenine (Allen et al., 1975a), and r3H]thyrnine (Filfilan and Sigee, 1977). The above studies collectively show that the synthesis of dinoflagellate (nuclear) DNA has a clear and distinct peak at some stage during the cell cycle. This peak frequently occurs early in the cycle (Allen et al., 1975a; Loeblich, 1976; Galleron and Durrand, 1979), and may even occur in the newly formed daughter cells immediately after cytokinesis (Franker, 1971; Franker et al., 1974). Peridinium cinctum appears to be unusual in having a peak of DNA synthesis late in the cell cycle (Spector et al., 1981a). In addition to a peak of DNA synthesis, Filfilan and Sigee (1977) and Galleron and Durrand (1979) have also detected a continuous low level of' precursor uptake in their respective studies on Prorocentrum micuns and Amphidinium carterae. Recent work by Karentz (1983) on a variety of

TABLE I1 Recent Studies on the Timing of DNA Synthesis in Dfnoflagellates

Reference

Material

Technique

DNA

Synchrony

Result

Peak synthesis During cytokinesisin cyst In mitotic cyst

Franker (1971)

Endomic dinoflagellate

146-scintillation counting

Whole cell

Selection synchrony

Discrete S phase

Franker et al. (1974) Allen et al. (1975b)

Crypthecodinium cohnii

32P-scintiUation counting Spectrofluorometry

Whole cell

Selection synchrony No synchrony

Discrete S phase Discrete S phase

Loeblich (1976)

Cachonina niei

Whole cell

FiWan and Sigee (1977)

Prorocentrum micans

Spectrofluorometry [5H]thymine autoradiography

Induction synchrony Phased culture

Oalleron and Durrand (1979)

Amphidinium carterae

[3H]scintillation counting

Nuclear

Induction synchrony

Spector et al. (1981a) Karentz (19883)

Peridinium cinctum

[fH]scintillation counting Spectrophotometry

Whole cell

Induction synchrony Phased cultures

Discrete S phase Continuous with peak synthesis Continuous with peak synthesis Discrete S phase Discrete S phase or continuous

Gymnodinium, Crypthecodinlum, Gyrodinium

Six species of marine dinoflagellates

Whole cell

Nuclear/ chromosomal

Whole cell

Shortly after mitosis Early in interphase During interphase Early interphase Late interphase Variable

THE DINOFLAGELLATE CHROMOSOME

225

marine dinoflagellates has also demonstrated continuous DNA synthesis in two particular species-Prorocentrum triestinum and Gonyaulux tamarensis. The contention by Spector and Triemer (1981) that continuous DNA synthesis does not occur in dinoflagellates may thus apply to some species, but does not appear to be true for all. In the autoradiographic studies of Filfilan and Sigee (1977) the phase of peak synthesis is shown by heavy labelling over some cells, and the continuous synthesis is demonstrated by the presence of at least some silver grains over all other interphase nuclei examined. The cell shown in Fig. 6a is a lightly labelled cell that is not undergoing a major synthetic phase. At the level of the electron microscope (Fig. Sb), silver grains were present over both the central condensed bodies of chromosomes and the surrounding nucleoplasm. The latter probably represents synthesis of DNA on the peripheral chromatin fibrils, as well as scatter of sensitisation from the main part of the chromosome. The studies of Galleron and Durrand (1979) suggest that the peak and continuous synthesis of DNA represent two distinct and superimposed components. The peak synthesis is inhibited by ethidium bromide, while the continuous phase is not. This biochemical distinction does not appear to parallel any distinction in the location of replicating DNA, since the electron microscope autoradiographs of Filfilan and Sigee (1977) do not show any clear differences between heavily labelled and poorly labelled cells in terms of silver grain distribution. Differences do occur, however, between actively synthetic and poorly synthetic chromosomes in terms of fine structure, and will be discussed in the following section. B. CHANGES IN CHROMOSOME FINE STRUCTURE

The degree of condensation and the general appearance of dinoflagellate chromosomes shows considerable variation between cells at different stages of the cell cycle. Some indication of this is given by the early studies of Haller er ul. (1964) on Amphidinium carterue, where considerable difference in chromosome structure was found between cells of the same asynchronous population. The fine structure of dinoflagellate chromosomes has now been widely investigated in relation to the cell cycle, though relatively few workers have attempted to determine a complete sequence of events over the entire cycle. Matthys-Rochon (1979) has studied fine structural events during the cell cycle in synchronised cultures of Amphidinium carrerue, and Sigee and Filfilan have investigated chromosome changes in phased populations of Prorocentrum micuns. A summary of the fine structural changes that occur during the vegeta-

226

D. C. SIGEE

Fig. 6. DNA replication in Prorocenrrum micans. Cells have been labelled with [3H]thymine and prepared as detailed in Filfilan and Sigee (1977). The cells shown here are not in the peak phase of DNA synthesis. (a) LM autoradiograph under bright field. The cell has a scattering of silver grains over the central region of chromosomes (C). ( ~ 5 2 0 0 . (b) ) EM autoradiograph. Details of nucleus, showing localised occurrence of silver grains over the main body of the chromosomes. ( ~ 9 5 0 0 . )

THE DINOFLAGELLATE CHROMOSOME

227

tive cycle of the free-living dinoflagellate Prorocentrum micans is shown in Fig. 7, and will serve as a basis for considering the changes that occur in dinoflagellates generally. In a laboratory culture of Prorocentrum, interphase stages (Fig. 7, stages a-d) occur as motile cells in suspension, while mitotic cells (Fig. 7, stages e-h) appear to have lost their flagella and sunk to the bottom of the culture vessel. A similar alternation has been noted in Amphidinium carterae (Galleron and Durrand, 1979). Chromosome changes during interphase and cell division will be considered separately.

1 . Interphase Cells Interphase cells of Prorocentrum micans show variation in the appearance of the chromosomes at both the light and electron microscope level.

Fig. 7. Morphological changes during the cell cycle of Prorocenfrum micans. The diagram is based on transmission light and electron microscope observations on cells taken from phased cultures (Filfilan and Sigee, 1977). (a) Naked daughter cells-dense chromosomes. (b) Early to mid-interphase cell. Chromosomes appear pale within a dense nucleoplasm under LM phase contrast. Electron-transparent zones under the EM. Peak phase of DNA replication. (c) Mid to late interphase. Dense chromosomes. (d) Late interphase. Regions of chromosome ‘fragmentation’ seen under EM. (e) Early cell division. Dividing chromosomes. (f) Cell division-karyokinesis. (8) Cell division-cytokinesis. (h) Final separation of daughter cells with loss of mother cell wall.

228

D. C. SIGEE

Direct comparison of these images for individual cells has been achieved by resectioning light microscope sections for electron microscopy, using the resin slide technique (Sigee,1976). The sequence of changes shown in Fig. 7 contains information obtained from both types of section, and relates to cells fixed in glutaraldehyde followed by osmium tetroxide (Ffilan and Sigee,1977). Naked daughter cells (Fig. 7a) appear generally pale, and contain dense chromosomes when observed by phasecontrast and electron microscopy. These cells increase in sue, develop a cell wall, and the chromosomes become pale within a dense nucleoplasm when observed under phase contrast (Fig. 7b). This stage corresponds to the phase of peak DNA synthesis (Sigee, 1976; Filfilan and Sigee, 1977). Under the electron microscope (Fig. 8a) the chromosomes appear quite different from those at other stages of the cell cycle in that they contain prominent electron-transparent regions. These regions may correspond to localised areas of chromatin decondensation, where maximal replication of DNA is taking place. With continuing progression through interphase the chromosomes recondense, becoming dense under phase contrast and losing their electron-transparent regions under TEM (Fig. 7c). The chromosomes remain condensed for the rest of interphase, but show signs of terminal fragmentation (Figs. 7d and 8b) as the onset of cell division approaches. The chromosome changes that occur during interphase in Prorocentrum micans have two stages of particular interest-the phase of partial decondensation (Fig. 7b) and the phase of chromosome fragmentation (Fig. 7d). Corresponding stages have been observed during interphase in other dinoflagellates. Partial decondensation in relation to DNA replication has been observed in Peridinium cincturn by Spector et a f .(1981a). In this species the chromosome uncoiling that occurs during the peak phase of [31-Ilthymidine incorporation represents a much more extensive h e structural change than the decondensation observed in Prorocentrum. The phase of chromosome fragmentation observed in Prorocentrum micans at the end of interphase does not relate to any peak of DNA synthesis, and electron microscope autoradiographs of cells labelled with ['Hlthymidine do not show any localisation of silver grains to regions of fragmentation within the chromosome complement (FilGlan and Sigee, unpublished observations). The significance of this fi-agmentation, which appears to involve a terminal splitting of the main body of the chronosome (Fig. 8b), is not understood. It does seem, however, to be an essential pre-requisite for chromosome division, and continues into the early stages of mitosis. Other investigators have also observed chromosome changes at this point in the dinoflagellate cell cycle. Soyer (1978a,b) noted in Prorocentrum micans a disorganisation in the pattern of DNA fibrils

Fig. 8. Interphase chromosomes during the cell cycle of Pvorocentrum miccrns. (a) Stage of peak DNA synthesis (Fig. 7b). with partial decondensation of chromatin to form electrontransparent regions within the chromosomes. lntranuclear cytoplasmic channels (Ch) contain various organelles, including trichocysts (t). Fixation in glutaraldehyde (2 hr) followed by osmium tetroxide (2 hr). (~20,400.)(b) Late interphase (Fig. 7d) with chromosomes showing regions of terminal ‘fragmentation’ (arrows), which probably represent regions of chromatin uncoiling. Fixation as for (a). ( X 18.300.)

230

D. C. SIGEE

prior to chromosome division, which was associated with the presence of axial fibres. Matthys-Rochon (1979) observed a ‘premitotic phase’ in Amphidinium carterae showing chromosome changes very similar to those seen in Prorocentrum (Fig. 8b), and in the same species Oakley and Dodge (1979) reported the presence of chromosome protrusions during early prophase.

2 . Cell Division a. Mitosis. It is not the purpose of this article to include a detailed review of the process of mitosis in dinoflagellates, and for this the reader is referred to articles by Kubai and Ris (1969), Oakley and Dodge (1976, 1979), and Spector and Triemer (1981). Mitosis has now been investigated in a wide variety of dinoflagellates, and the process in these organisms is quite atypical of other eukaryote cells in a number of important respects. These include the persistence of the nuclear membrane throughout division, the absence of a typical eukaryote metaphase plate, chromosome attachment to the nuclear membrane during segregation, absence of a centriole and centromeric chromatin, and the presence of a permanently extranuclear spindle. Changes in the general appearance of cells during mitosis in Prorocentrum micans are shown in Fig. 7e-h. Two particular aspects of this will be considered, the division of chromosomes and the migration of the daughter chromosomes. The term ‘chromatid’ will not be used here in reference to the two chromosome division products since this implies homology to the chromatids of other eukaryote cells and the term has also been separately used to define the individual constituent DNA molecules (Haapala and Soyer, 1973; Soyer and Haapala, 1974a). i. Chromosome division. The first stage of cell division in dinoflagellates involves longitudinal splitting of the chromosomes. This was initially observed under the light microscope by Kohler-Wieder (1937) in studies on Glenodinium pulvisculus and Peridinium willei, and by Skoczylas (1958) for Ceratium spp. Subsequent electron microscope studies by Soyer and Haapala (1974a) have revealed fine structural details of the dividing Y-shaped chromosomes, as seen in ultrathin section. The division of chromosomes in Prorocentrum micans appears to coincide with a loss of cell motility (Fig. 7e) since these cells are not detected in the supernatant. Y-shaped chromosomes can be clearly seen at the level of the light microscope (Fig. 9a), where they are randomly orientated within the nucleus. Transmission electron micrographs of these cells show that the two daughter chromosome units are typically associated with nuclear membranes, either with intranuclear membrane channels (Fig. 9b) or with the nuclear envelope. The model of a dividing chromo-

Fig. 9. Dividing chromosomes during the cell cycle of Prorocenirum micans (Fig. 7e). The ultrathin section shown in (b) is derived from the 2-pm-thick section photographed in (a), using the resin slide technique (Sigee, 1976). (a) Phase-contrast micrograph. Section of whole cell containing large nucleus in which chromosomes are undergoing division and ) TEM detail of appear as Y-shaped structures (arrows). Fixation as for Fig. 8a. ( ~ 2 5 0 0 .(b) dividing chromosome, consisting of mother chromosome (C) and two daughter units (C'). Two large intranuclear cytoplasmic channels (Ch) are closely associated with the chromosome. One of these extends into the division fork, where there appears to be direct chromatin-membrane contact (arrow). Fixation as for Fig. 8a. (X45.900.)

232

D. C. SlGEE

Fig. 10. Three-dimensional reconstruction of dividing chromosome of Prorocenrrum micans, from serial sections of the nucleus shown in Fig. 9, with direct contact between the chromosome fork (C') and the nuclear envelope (Ne). Several intranuclear channels (Ch) occur in close approximation. The non-dividing part of the chromosome (C) does not have any membrane associations. (Regions of channel and chromosome cut in the uppermost section of the series are coded black and white, respectively.)

some illustrated in Fig. 10, derived from serial sections, shows direct association of the division fork with nuclear envelope, and also the presence of various non-associated nuclear channels in the vicinity of the chromosome. Attachment of dividing chromosomes to the nuclear membrane in dinoflagellates was initially noted by Kubai and Ris (1969) for Gyrodinium and Soyer (1969) for Blustodinium. The close association of the daughter units with nuclear membranes would suggest that separation is mediated at least partly by membrane flow, just as the separation of daughter genomes is membrane mediated in bacteria (Kubai and Ris, 1969). Other structures may also be important in the process of chromosome division, including the presence of central axial structures within the dividing chromosome (Soyer, 1977b, 1978a,b). Longitudinal splitting of the chromosome often leads to a point of ter-

THE DINOFLAGELLATE CHROMOSOME

233

minal attachment to the nuclear envelope, as observed in Blastodinium (Soyer, 1969) and Prorocentrum micans (Soyer and Haapala, 1973, 1974a), resulting in a final V-shaped configuration. This is not always the case, however, as shown by Amphidinium carterae (Matthys-Rochon, 19791, where chromosomes are attached to the nuclear membrane centrally by a kinetochore and splitting occurs from both ends to this point, which is the last part of the chromosome to divide. The process by which the complex fibrillar superstructure of the chromosome separates into two daughter units is not understood, though Haapala and Soyer (1973) have proposed a mechanism based on their initial polytenic model of chromosome structure. Whatever the process, the daughter chromosome units have the same fibrillar architecture as the mother chromosome when observed in ultrathin section (Fig. 9b) and in whole-mount preparations (Soyer and Haapala, 1974b). ii. Chromosome migration. Early studies by Dodge (1963b, 1966) involving the effect of mitotic inhibitors such as colchicine and 8-hydroxyquinone, and the effect of X-ray treatment, suggested that integrity of a nuclear spindle was not required for chromosome separation in Prorocentrum micans and Peridinium trochoideum. Studies on Woloszynskia miera (Leadbeater and Dodge, 1967), Crypthecodinium cohnii (Kubai and Ris, 1969), and Amphidinium carterae (Oakley and Dodge, 1974, 1976) have since shown that chromosome separation in dinoflagellates is mediated by intranuclear channels, which typically form at the beginning of mitosis. The nuclear envelope remains intact throughout cell division, but a nuclear spindle is present as discrete microtubules within the nuclear channels (Leadbeater and Dodge, 1967; Oakley and Dodge, 1976; Tippit and Picket-Heaps, 1976; Mattys-Rochon, 1979). Although chromosomes are not directly attached to microtubules in dinoflagellate nuclei, the latter are generally regarded as being involved in chromosome migration. In some species, including Amphidinium carterae (Oakley and Dodge, 1974, 1976), Oodinium dogieli (Cachon and Cachon, 1974), Apodinium spp. (Cachon and Cachon, 1979), and Polykrikos spp. (Spector and Triemer, 1981), chromosomes are associated with a distinct kinetochore, which consists of a small pad attached to the inner nuclear membrane of the intranuclear channel, with a single microtubule terminating at the pad of the kinetochore. These specialised attachment zones are not present, however, in Crypthecodiniurn cohnii and Peridiniurn cinctum (Spector and Triemer, 1981) and should not therefore be regarded as a universal characteristic of this group. In addition to the probable involvement of nuclear membranes and extranuclear microtubules in chromosome migration during karyokinesis, it has been suggested that intranuclear cables are also involved (Soyer,

234

D. C. SIGEE

1981). These structures have been demonstrated within the nucleus of Prorocentrum micans (Soyer, 1981) using a freeze-substitution technique. Prorocentrum micans is unusual in that intranuclear channels are not restricted to the period of mitosis (see, for example Fig. 8a) and the presence of nuclear channel microtubules requires special fixation procedures (Soyer, 1977a,b). Conventionally fixed cells undergoing karyokinesis (Fig. 7f) and cytokinesis (Fig. 7g-h) typically appear pale when observed in resin blocks (Fig. 1la). The light micrograph in Fig. 1l b shows a late stage of karyokinesis, with V-shaped profiles of migrating chromosomes still apparent. The electron micrograph in Fig. 1l c shows details of chromosomes from this cell, and it is interesting to note the presence of a halo of dispersed fibrils around the central condensed body, just as in interphase cells. The clarity of these peripheral fibrils probably arises due to a decrease in the density of nucleoplasm in fixed cells at this stage of the cell cycle. 6 . Meiosis. Sexual reproduction is now known to occur in dinoflagellates far more commonly than was originally believed. Early reports of sexual life histories in Ceratium (Zederbauer, 1904) and Glenodinium fubiniensiforme (Diwald, 1937) have now been supplemented by more recent studies which show that sexual life histories (including meiosis) occur in at least 21 dinoflagellate species, including 10 genera (von Stosch, 1973; Pfeister, 1977; Spector et al., 1981b; Pfeister et al., 1984). With the exception of Noctiluca all these life histories are haplontic (Steidinger, 1975). Available evidence on the process of meiosis in dinoflagellates is fragmentory and is based on genetic and light microscope studies. There are indications that meiosis in these cells may show a number of important differences from normal eukaryote cells. Genetic analysis, involving 16 non-allelic mutants, has been carried out on Crypthecodinium cohnii by Himes and Beam (1975) and Beam et al. (1977) and reveals an absence of second division segregation and an almost completely independant assortment of genes. They conclude that meiosis in this species is a onedivision process, which is corroborated by the observation of two-celled zygotic cysts. In contrast to Crypthecodinium, recent light microscope studies by Pfeister et al. (1984) have demonstrated a two-stage division process in Peridinium inconspicuum. No details of chromosome morphology or disposition are presented. Although there have apparently been no transmission electron microscope studies on the process of meiosis in dinoflagellates, Spector et af. (1981b) have investigated the preceding stages of gamete formation and fertilisation in Peridinium cinctum. The structure of the chromosomes during gamete formation is similar to that of vegetative cells, but follow-

Fig. 11. Dividing cells of Prorocenrrum micnns. (a) Light micrograph of whole cells at surface of resin block. Interphase cells appear very dense, but dividing cells, including cells undergoing cytokinesis (arrows, see Fig. 7h) and newly formed (laughter cells (d, see Fig. 7a), appear pale. (b). Light micrograph of early stage in cytokinesis (Fig. 7g). Within the two daughter nuclei (A and B), the V-shaped profiles of chromosomes that have been recently pulled apart are clearly visible. Fixation as for Fig. 8a. ( x 1900.) (c) TEM detail of two chromosomes in ultrathin section, taken from the thick section shown in (b), using the resin slide technique. The main body of the chromosome (C) has the same fibrillar structure as in interphase cells, but the peripheral fibrils (0 appear much clearer due to the electron transparency of the nucleoplasm. Fixation as for Fig. 8a. ( ~ 1 6 , 5 0 0 . )

236

D. C. SIGEE

ing liberation of the gametes the chromosomes become partially uncoiled and show fibrous extensions. Fusion of the gametes leads to the formation of a zygote which subsequently develops a secondary wall, and becomes a dormant cyst. Chromosomes within the zygote are no longer uncoiled and fibrous but have returned to the fully condensed structure typical of vegetative cells.

VII. MACROMOLECULAR COMPOSITION OF DINOFLAGELLATE CHROMATIN One of the earliest histochemical studies on the nuclei of dinoflagellates was made by Ris (1962), who showed that the chromosomes of Amphidinium carterae stained uniformly with the Feulgen reaction for DNA, but did not stain with the protein-specific reagents alkaline fast green, alkaline eosin, and mercuric bromophenol. Ris concluded that the chromosomes were primarily, if not exclusively, composed of DNA. Similar results were obtained by Dodge (1964) for Prorocentrum micans, with histochemical detection of DNA but not RNA or proteins in the chromosomes. These findings led Dodge (1965, 1966) to propose that dinoflagellates constitute a separate mesokaryote group, intermediate between prokaryotes and eukaryotes. Electron microscope examination of nuclei extracted by DNase (Leadbeater, 1967) revealed specific removal of chromatin fibrils and supported the idea that the chromosomes of dinoflagellates are completely composed of DNA. Subsequent electron microscope studies by Soyer and Haapala (1974b,c), however, suggested that RNA and protein are present in dinoflagellate chromatin as an integral structural component. The X-ray microanalytical detection of substantial levels of sulphur in the chromosomes of fixed and unfixed cells by Kearns and Sigee (1979, 1980) is consistent with the in situ presence of proteins, while the occurrence of RNA has been verified by the presence of a residual phosphorus peak after DNase digestion (Sigee and Kearns, 1981c). In recent years, biochemical techniques have been used to extract and characterise chromatin components. Mendiola et al. (1966) isolated large numbers of nuclei of Gymnodinium nelsoni and showed that the average composition of the nucleus was 49% DNA, 44% protein, and 7% RNA. The characterisation of these three components will be considered separately.

THE DINOFLAGELLATE CHROMOSOME

237

A. PROTEINS

The overall fraction of protein determined by Mendiola et al. (1966) is much lower than in higher eukaryotes (DuPraw, 1970). Rizzo and Nooden (1972, 1974a,b) analysed the extracted chromatin from isolated intact nuclei of Crypthecodinium cohnii, Gymnodinium nelsoni, Peridinium trochoideum, and P . cinctum. This chromatin contained substantial levels of acidic (acid insoluble) protein. Subsequent studies by Franker (1972) and Franker et al. (1974) using DNA-cellulose chromatography have confirmed the presence of DNA-binding proteins. In contrast to acidic proteins, the level of basic proteins is very low in dinoflagellate chromatin (Rizzo and Nooden, 1973). Rizzo (1981) has shown that the mean basic protein/DNA ratio in a range of dinoflagellates varies from 0.022-0.13, while corresponding values for acidic protein vary from 0.29-1.22. The mean ratio of acididbasic protein varies from 5.6-15.3. The levels of acidic and basic proteins are comparable to the situation in prokaryotes, but are in marked contrast to the situation in eukaryote chromatin. Rizzo and Nooden (1974a,b) isolated a single basic (acid soluble) protein from dinoflagellate chromatin, with a molecular weight of 16,000. Although generally similar in its properties to eukaryote histones, Rizzo and Nooden (1974b) considered that this protein should not be designated a true histone, and reported a complete inability to detect any histone proteins in dinoflagellate chromatin. This apparent absence of histones is in contradiction with the immunofluorescence studies of Stewart and Beck (1967), which claimed positive identifications of DNA-histone complexes in five dinoflagellate species. Rizzo (1976) has claimed that the techniques used in the immunofluorescence work were not histone specific. Since the initial studies of Rizzo and Nooden on Crypthecodinium and Peridinium, histone-like proteins have been identified in other dinoflagellates (Rizzo, 1981; Rizzo and Burghardt, 1982; Herzog and Soyer, 1981), where they are present either as one or two types of molecule per cell. The proteins from different dinoflagellates are similar but not identical (Rizzo, 1981), reflecting the great diversity of attributes within this group. Gymnodinium breve, for example, resembles Prorocentrum micans in having two histone-like proteins of molecular weights 12,000 and 13,000, but these do not co-migrate in SDS gels. The two histone-like proteins of Gymnodinium breve also differ from those of Prorocentrum micans in their extractibility with dilute acid (Rizzo et al., 1982). The initial studies of Rizzo and Nooden also suggested that basic proteins were only present in log-phase cells. Rizzo et al. (1984), however, have recently described an improved method for the isolation of dinoflagellate nuclei, resulting in

238

D. C. SlGEE

a more sensitive detection of basic chromatin protein, and have shown that histone-like proteins are present in stationary as well as log-phase cells. Relative proportions of the different histone-like proteins do appear to vary during the period of culture of a particular dinoflagellate. In Gymnodinium nelsoni, for example, there is a major component at 10,000 Da which is invariably present, but minor components at 13,000 and 17,000 Da are highly variable in occurrence (Rizzo and Burghardt, 1982). The distinction between acidic and basic proteins in dinoflagellate chromosomes has recently been investigated at the fine structural level by Sigee (1983b, 1984a) using buffer extraction of briefly fixed whole cells of Glenodinium foliaceum and Amphidiniurn carterae. Glutaraldehyde fixation in pH 5.0 phosphate buffer, and fixation followed by 72 hr treatment in the same buffer (Fig. 12a), results in major retention of the electrondense chromosome matrix. Fixation in neutral (pH 7.0) buffer results in the retention of matrix, but this is largely lost in cells treated for a further 72 hr with buffer alone (Fig. 12b). Fixation in an alkaline (pH 9.0) Tris buffer rapidly extracts the chromosome matrix. After a further 72-hr incubation in buffer alone the whole chromosome appears electron transparent and the chromatin has become very irregular (Fig. 12c), possibly due to DNA destabilisation after protein removal. B.

DNA

The high levels of DNA in dinoflagellates have already been noted (Table I). Characterisation of isolated DNA from these organisms is of potential importance in providing some explanation for these high levels, giving further insight into prokaryote/eukaryote affinities of the group and possibly revealing unique features which are peculiar to dinoflagellates generally. The characterisation of this DNA has concentrated on two main features, renaturation kinetics, and the presence of modified bases. 1 . Renaturation Kinetics of Dinoflagellate DNA Studies by Allen et al. (1975a) on the renaturation kinetics of Crypthecodinium cohnii DNA revealed high levels (55-60%) of repeat sequence DNA interspersed with highly complex DNA. This places dinoflagellate affinities very firmly with the eukaryotes, since prokaryotes possess very small amounts of repeat sequence DNA by comparison (Britten and Kohne, 1968). The results obtained by Allen et al. (1975a) suggest that the highly complex DNA is largely unique DNA, which is consistent with some genes being present in a single copy, or haploid condition. This is in agreement with other studies which suggest that the vegetative cells of dinoflagellates contain a haploid genome. Evidence for this

Fig. 12. Buffer extraction of the chromosome martix in Amphidinium currerue. For each buffer, cells were fixed in 2.5% buffered glutaraldehyde, washed, treated for a further 72 hr in buffer at 20"C, then postfixed in osmium tetroxide (Sigee, 1984b). (All X54,000.) (a) pH 5.0 (Sorensen's phosphate buffer). The chromosome is very electron dense due to maximal retention of protein matrix. (b) pH 7.0 (Sodium cacodylate buffer). The chromosome has become electron transparent due to major loss of protein matrix, revealing a residual framework of DNA fibrils. Some peripheral fibrils can be seen extending into the surrounding nucleoplasm. (c) pH 9.0 (Tris buffer). The chromosomes appear electron transparent within a dense nucleoplasm. The DNA fibrils form an irregular reticulum, possibly due in part to a destabilisation caused by complete removal of acidic protein matrix.

240

D. C. SIGEE

comes from the work of von Stosch (1973) who has demonstrated a haplontic life cycle in various free-living dinoflagellates, and also from the mutation studies of Roberts et al. (1974). By inducing mutations in Crypthecodinium cohnii and scoring their frequencies, Roberts et al. (1974) were able to show that many genes were present only as a single copy in the chromosome complement. Evidence for genetic recombination in crosses between mutant strains of Crypthecodinium cohnii has been obtained by Tuttle and Loeblich (1974a,b, 1975), and other workers (Beam and Himes, 1974; Himes and Beam, 1975) have shown that chromosomes are functionally single strands at the time of crossing over. 2 . ModiJed Bases in Dinojagellate DNA All groups of organisms appear to contain in their DNA bases other than adenine, guanine, cytosine, and thymine (Shapiro, 1976). These modified bases include 5-methylcytosine, N6-methyladenine, and 5-hydroxyuracil, and can be found in varying amounts in DNA of viral, prokaryote, and eukaryote origin. Since the initial studies of Franker (1970), demonstrating a small fraction of 5-methylcytosine in the dinoflagellate symbiont of Anthopleura, the general presence of modified bases in dinoflagellate DNA has been established by Rae (1973, 1976), Allen et al. (1975a), and Herzog and Soyer (1982). A general survey of the nucleotide composition of DNA from dinoflagellates is presented by Rae and Steele (1978). The most conspicuous feature of dinoflagellate DNA, and the one which distinguishes it from other eukaryotes (Rae, 1973), is the presence of high levels of the base 5-hydroxymethyluracil. This base has been found elsewhere only in viral DNA (Kallen et al., 1962) and has the effect of raising the density and lowering the thermal stability of the DNA in which it is contained (Rae, 1976). The non-randomness in the distributions of hydroxymethyluracil and thymine in Crypthecodinium (Rae, 1976) suggests that the hydroxymethyluracil arises due to modification of thymine in short, specific sequences of DNA. The amount of hydroxymethyluracil varies considerably, varying from 12% of the thymine in Exuviella to 68% in Peridinium (Rae and Steele, 1978). Rae and Steele (1978) have suggested that this base has no contemporary role in modern dinoflagellates, either in terms of chromosome structure or nuclear metabolism, but was probably important as part of a genetic modification-restriction system in the earliest dinoflagellates. Such a system protects the cell from the consequences of invasion by foreign DNA (Meselson et al., 1972).

THE DINOFLAGELLATE CHROMOSOME c.

24 1

RNA

The early studies of Rizzo and Nooden (1972, 1973, 1974a,b) on isolated chromatin demonstrated an RNA/DNA ratio of about 0.04, which is similar to that described for mammalian cells. Biochemical studies on extracted dinoflagellate RNA and associated enzymes (polymerases) have tended to emphasise homologies with higher eukaryote rather than prokaryote cells. Hinnebusch et al. (1981) have recently examined partial nucleotide sequences for 5 S and 5.8 S rRNAs from Crypthecodiniumcohnii. The 5 S RNA sequence showed least homology with prokaryote sequences, and most homology (75%) with the 5 S sequences of higher eukaryotes. Hinnebusch et al. concluded that the evolutionary affinities of the dinoflagellates were firmly with higher eukaryotes. More recently, Reddy et al. (1983) have presented evidence for the presence of six capped small nuclear RNAs (snRNAs) in Crypthecodinium cohnii. The six snRNAs were shown to be closely similar to the snRNAs (designated uI-u6) of higher eukaryotes, as determined by (1) the presence of trimethylguanosine cap structure in UI-Us RNAs, (2) sequence homology between rat and dinoflagellate Uz, Us, and u6 RNAs, (3) presence of other post-transcriptional modifications such as sugars and base modifications, and (4) association of Sm antigen with five of the six RNAs. This is the first report of such U-snRNAs in a unicellular organism, and suggests (Reddy et al., 1983) a close affinity of this group with higher eukaryotes, where they have been identified in insects, birds, rodents, and mammals. The intranuclear location of these RNAs in dinoflagellates has not been determined. They are most likely, however, to be chromatin associated rather than part of the chromatin structure, since they occur within ribonucleoprotein particles in higher eukaryotes. Work on RNA polymerases has been carried out by Rizzo (1979), who has shown the presence in Crypthecodinium cohnii of an enzyme which is typical of eukaryotes in its sensitivity to a-amanitin but is atypical in its inhibition by Mn2+ ions.

VIII. IONIC COMPOSITION OF DINOFLAGELLATE CHROMATIN

One conspicuous feature of dinoflagellate chromosomes is their high cation content. This was initially demonstrated by Hamkalo and Rattner

242

D. C. SIGEE

(1977), who used positive group-specific stains to locate cationic moieties in isolated chromatin fibres. The localised occurrence of cations in dinoflagellate nuclei has more recently been investigated by X-ray microanalysis and autoradiography, and has been the subject of several reviews (Sigee, 1983a, 1984a). A . X-RAY MICROANALYSIS

Initial electron probe X-ray microanalytical studies by Kearns and Sigee (1979) revealed the presence of a range of insoluble cations in the chromatin of fixed, dehydrated cells, including both marine (Prorocentrum micans, Amphidinium carterae, Peridinium faroense, Glenodinium foliaceum) and freshwater (Ceratium hirundinella) species. Fig. 13 shows an X-ray emission spectrum taken from a glutaraldehyde-fixed chromo-

CI

I

1

I

1

I I

I I

I I

VI

1

E

I

X

I I I I

I

I

I I I I

1 I I I

I

r

I

1

1

c

I

I I

RESIN CONTROL

currr

X-ray energy IkeV)

Fig. 13. X-ray emission spectrum of glutaraldehyde-fixed cell of Glenodinium foliaceum. Characteristic spectra are shown for probe areas in the section of chromatin and pure resin adjacent to the cell. Prior to fixation, the cells had been cultured long term in very dilute (10% of normal) growth medium (Sigee and Kearns, 1981b) and the spectrum is unusual in having substantial amounts of K and C1. The resin section control spectrum has a clear Ti peak (derived from the nylon grid material) but no other extraneous peaks.

THE DINOFLAGELLATE CHROMOSOME

243

some of Glenodiniumfoliaceum, with clear peaks of P (nucleic acid), S (proteins), Ca, and the transition metals Fe, Cu, and Zn. This spectrum is unusual in the presence also of K, which normally occurred only as a soluble cation, and was not characteristically detected in fixed chromatin. Emission spectra from the nucleoplasm showed smaller, more variable peaks of these elements, with the exception of Ni, which was only detected in chromosomes (Sigee and Kearns, 1980). P, S, and Ca were routinely present in the cytoplasm, but transition metals were not normally detected (Kearns and Sigee, 1979). This qualitative work on fixed cells was followed by examination of cryoprepared material, with subsequent quantitative determinations of elemental mass fractions in cells prepared and treated in various ways. 1. Cryopreparations X-ray emission spectra from ultrathin frozen-dried cryosections of Prorocentrum micans (Fig. 3a) showed peaks of the soluble cations Na and K, with small peaks of Ca and transition metals, Fe and Cu (Kearns and Sigee, 1980). Cation peaks have also been obtained from washed, dispersed cryosections (Kearns and Sigee, 1980) and from sections of resininfiltrated, frozen-dried cells (Sigee and Kearns, 1982b). In all of these cases the cells were unfixed, so that the detection of divalent cations in dinoflagellate chromatin cannot be attributed to fixation artefact. Divalent cation peaks were, however, generally less prominent in cryoprepared material compared to fixed cells, and this is particularly evident in recent work (Harrison and Sigee, unpublished observations) on 200-nm-thick frozen-dried cryosections of Glenodiniumfoliaceum. These sections (Fig. 3c) are considerably thicker than those obtained previously with Prorocentrum micans and the rippled appearance indicates that they completely freeze dried in the solid (frozen) state. The X-ray emission spectra (Fig. 14) have prominent peaks of P, S, K, and C1 with a small Ca peak (which is partly obscured by the KK, peak), but no detectable transition metals. The difference between the spectra of frozen (unfixed) and chemically processed (fixed and dehydrated) cells can be accounted for in terms of loss of chromatin constituents during chemical preparation. In the living cell, and resulting cryosection, the chromosomes contain substantial levels of soluble protein and soluble monovalent ions, including K and C1. Divalent cations are present but difficult to detect. Fixation and dehydration remove soluble components, leaving only insoluble constituents such as nucleic acids, high-molecular-weight proteins, and bound divalent cations. The ability to detect divalent cations, as with other elements, in X-ray microanalysis appears to depend on two factors: (1) the total X-ray signal being generated from the element, and (2) the

244

D. C . SIGEE P

S C I

K

CHROMATIN

FORMVAR CONTROL

I

I I

5 X-ray energy (keV)

Fig. 14. X-ray emission spectrum from freeze-dried cryosection of Glenodinium foliaceum. The chromatin spectrum is taken from a thick (0.5-1 .O pm) cryosection, similar to that shown in Fig. 3c. The high level of K results in a substantial K peak, which largely obscures a small peak of detectable Ca. The control spectrum is taken from a region of clear formvar adjacent to the cryosection, and shows a single extraneous peak of Al, derived from the grid material.

elemental mass fraction. In frozen sections, these chromatin-bound cations are not at sufficiently high concentration to have reached the threshold of detectability. In chemically processed cells, however, tissue shrinkage results in increased concentration and the large-scale loss of soluble chromosome matrix increases the bound cation mass fraction. Both of these factors render divalent cations more detectable in the chromatin of chemically processed cells. 2. Mass Fraction Levels in Different Species The mass fraction of cations within the chromatin can be calculated from the X-ray emission spectra, using Hall (1971) quantitation for ultrathin specimens, and by reference to known standards. The mean values obtained (for 25-30 cells) show considerable variation between cultures of different dinoflagellate species (Kearns and Sigee, 1980). Considerable variation also exists between cultures of the same species (Sigee and

10

THE DINOFLAGELLATE CHROMOSOME

245

Kearns, 1981a,b), however, so that the pattern of cation occurrence in dinoflagellate chromatin is not species specific. The overall presence of cations can be quantitatively related to particular macromolecules by reference to marker elements. Thus the levels of cations might be related to nucleic acid levels by comparing the cation mass fraction with that of P (Kearns and Sigee, 1980), or to sulphurcontaining proteins by comparing to the mass fraction of S (Sigee, 1984a). Although there is considerable variation in the occurrence of particular cations between cultures, when the overall levels are expressed in relation to P the situation becomes uniform, with 40-50 divalent cations present per 100 atoms of P (Kearns and Sigee, 1980). This is equivalent to one divalent cation per two P atoms, or per two nucleotides. This precise stoichiometry suggests that divalent cations are directly associated with nucleic acids, particularly DNA. Corresponding calculations with reference to 100 atoms of S (Sigee, 1984a) give quite different, highly variable results, suggesting no close correlation between mean levels of cations and overall level of S-containing proteins. 3. Elemental Occurrence in Chromatin-extracted Cells The association of cations with particular macromolecules can be further investigated by extraction of specific chromatin constituents. Extraction of DNA by DNase (Sigee and Kearns, 1981~)results in visible removal of chromatin fibrils, and a corresponding decrease in the P peak. The level of cations falls but is not eliminated. Specific extraction of RNA by RNase also brings about a decrease in the level of P and a partial release of cations. These experiments suggest that although cations are bound to both DNA and RNA, they are also associated with other chromatin constituents, particularly proteins. The residual cations left after nuclease treatment do not simply relate to residual nucleic acid (DNA or RNA), since the ratio of cations/P is very much increased by the enzyme treatment (Sigee and Kearns, 1981~). Extraction of the protein matrix can be achieved by prolonged fixation in buffered glutaraldehyde (Sigee and Kearns, 1982a). Fixation times in excess of 4 hr result in a decrease in the overall electron density of the chromosomes, with a correlated drop in the mean S mass fraction. This loss in protein matrix is associated with a fall in the level of divalent cations, but no decrease in the level of P (i.e. no extraction of nucleic acid). The results obtained from the chromatin extraction experiments are consistent with the divalent cations being associated with both nucleic acid and protein matrix. It has been suggested (Kearns and Sigee, 1980) that they may act as bridging molecules between the two major chromatin constituents.

246

D . C. SIGEE

4 . Chromatin Cations and External Availability The high levels of cations found in dinoflagellate chromatin might arise as a cellular response to abnormally high levels in the culture medium. Such a response has been obtained in other cells (Silverberg et al., 1976; Skaar et al., 1973), where electron-dense deposits of metal within the nucleus were the direct result of abnormally high external levels. In the case of dinoflagellate cells the presence of metal atoms within the chromatin did not appear to correlate with any electron-opaque deposit. The possibility that the presence of metal ions in dinoflagellate chromatin might arise as a pollution effect was tested by growing cells of Glenodinium foliaceum in culture media at various dilutions (Sigee and Kearns, 1981b). This species was particularly suitable for this study, since it can tolerate a wide range of salinity (Silva, 1962). Dilution of the medium down to 30% of the original concentration has no effect on cell growth and multiplication, and cells are able to survive in a stationary phase at dilutions down to 10% normal strength. At these various dilutions, the mass fraction of the chromatin divalent cations within chemically processed cells was not reduced. This shows that the presence of this chromatin constituent is not simply a sequestration response to high external cation levels, but is an important and integral part of the chromatin structure. In the case of cells grown in very dilute (10%) medium, where external levels of cations are clearly very limiting, the chromatin was more condensed (higher P mass fraction), and the number of divalent cations per 100 atoms of P dropped to 30. The decreased proportion of divalent cations seemed to be compensated by the presence of bound monovalent cations, since K then appeared as a substantial peak in the fixed cells (Fig. 13). B. AUTORADIOGRAPHY

The uptake and occurrence of divalent cations in dinoflagellate chromosomes has been investigated by autoradiography using radioactive nickel, 63Ni2+(Sigee, 1982), and radioactive calcium, 45Ca2+(Sigee, 1983b). These isotopes are suitable for autoradiography since they have long half-lives (92 years and 164 days, respectively) and are both emitters of low-energy radiation. Light microscope autoradiographs of 63Ni2+-labelledcells of Glenodinium foliaceum show a general scatter of silver grains over and around the whole cell (Fig. 15a,b). With the higher resolution of electron microscope autoradiography (Sigee, 1982) the incorporated nickel is localised to the nucleus, and within this to chromosomes. In the case of cells incubated with 45Ca2+,the nuclei dinoflagellate

Fig. 15. Uptake of 63Ni2+in Glenodinium foliaceum. Cells were treated with radioisotope for 4 hr prior to fixation in glutaraldehyde and osmium tetroxide, as detailed in Sigee (1982). (a) Phase-contrast micrograph of cell group in autoradiograph. ( X 1100.) (b) Bright-field micrograph of cells shown in (a). There is a wide scatter of grains over and around the cells. ( X 1100.) (c) TEM autoradiograph showing detail from nucleus. Silver grains typically showed a clear localisation to the nucleus, and within this, to the main bodies of the chromosomes. ( X 15,000.)

248

D . C. SIGEE

nuclei appear well labelled under the light microscope (Fig. 16a and b). Electron microscope autoradiographs of 45Ca-labelled cells in ultrathin section did not prove successful, and the absence of silver grains in these preparations probably results from the higher energy of the isotope (254 keV for 45Ca, compared to 67 keV for 63Ni and 18 keV for 3H). Some indication that 45Ca2+is largely incorporated into the chromosomes is given, however, by the results of buffer extraction of labelled cells. This results in a specific visible extraction of chromosome matrix and a largescale loss of 45Ca2+(Fig. 16c). The autoradiographic results suggest that both radioactive cations are actively taken up into the dinoflagellate nucleus, and become bound to the chromatin as an insoluble constituent. Experimental variation in the duration of labelling indicates that differences in uptake between the two cations do occur, with 63Niincorporation being continuous over an extended time period (Sigee, 1982), while 45Ca2+rapidly saturates nuclear affinity sites within a period of 1 hr. With both radioisotopes, heavy labelling was detected in cells processed in various fixatives, both additive (glutaraldehyde, paraformaldehyde) and coagulative (acetic/alcohol). The binding of radioactive cations to these cells could not therefore be attributed to specific fixation artefact similar to that occurring in other systems with labelled amino acids (Peters and Ashley, 1967). The localised occurrence of incorporated radioactive nickel and calcium obtained with the autoradiography was consistent with the X-ray microanalytical data. C. CATIONS AND CHROMATIN STABILISATION

The potential importance of cations in the stabilisation of dinoflagellate chromatin structure was recognised in the studies of Hamkalo and Rattner (1977) on isolated chromatin fibres, and Kearns and Sigee (1980) on sectioned chromosomes. Kearns and Sigee (1980) proposed that this stabilisation would be of particular importance in dinoflagellate chromatin, where there is an absence of the histones that are normally present in eukaryote chromatin. Sigee (1984a) referred to this as a ‘cationic nonhistone stabilisation system’ to distinguish it from normal eukaryotes, where cation-mediated condensation of DNA requires the presence of H1 histone (Thoma et al., 1979). Sigee (1983a) proposed a model (Fig. 17c) in which monovalent and divalent cations are primarily associated with the outside of the DNA molecule, acting both as bridging molecules between the DNA and surrounding acidic protein matrix and neutralising the highly negatively charged phosphate groups. The proposed role of cations in dinoflagellate chromatin is consistent with the work of Eickbush and

Fig. 16. Uptake of 45Ca2+in Gknodiniurn foliuceum. Cells were labelled for 24 hr, followed by glutaraldehyde fixation, as detailed in Sigee (1983b). (a) Phase-contrast micrograph of autoradiograph, showing prominent dinocaryotic nuclei in the central region of the cells. ( X 1500.) (b) Bright-field micrograph of same area as (a). Silver grains are clearly localised to the dinocaryotic nuclei, with only a light scattering over other parts of the cell, including supernumerary nucleus and cytoplasm. ( x 1500.) (c) Phase-contrast micrograph of autoradiograph of buffer-extracted (36 hr, pH 7.2, sodium cacodylate buffer) cell. The nucleus shows specific loss of matrix from the chromosomes (C), which appear very pale, but not the nucleolus (nu). In buffer-extracted cells the labelling of the cells was very much reduced. ( x 2800.)

250

D. C. SIGEE

a

b

c

o* 9 P P P

tlo 0

P

P P

P P

?P

P P

b

*O

+O? P

P

LPO” I<EY TO SYMBOLS DNA

. .

-

P

P

D

U

Matrix Protein - 0 Divalent Cations

--

Monovalent Cations

+

Fig. 17. Models of chromosome structure. This figure shows particular models of chromosome structure in relation to three different levels of organisation. (a)Fibril arrungemenr. Polytenic chromatid model of Haapala and Soyer (1973). reproduced with permission of the authors. The model shows the linear arrangement of two complete “circular” chromatids that have been twisted to give a spiral configuration. The distance p represents one complete twist of the spiral (enclosing a “figure of 8’7, and the arrows define the polarity of the DNA in one of the chromatids. The central segment shows the spread of a multiplicity of chromatid fibrils to give a “ball of chromatin.” (b) Liquid crystal. Interpretation of chromatin fibrillar patterns (shown in offset rectangle) in terms of a cholesteric liquid crystal model based on alternating layers of regularly arranged DNA strands. Taken from Bouligand er a / . (1968) and Livolant and Bouligand (1978), and reproduced with permission of the authors. (c) Cation associarions. Simple representation of the association of divalent and monovalent cations with the DNA strands and part of the protein matrix. The very high levels of monovalent ions that are seen in cryosections-probably as part of the general chromatin matrix-are not shown in this model. The figure is taken from a larger diagram (Sigee, 1983a) which relates chromatin preparation to microanalytical data.

Moudrianakis (1978) on in uitro stabilisation of eukaryote DNA by salts, which concludes that these act primarily by neutralisation of phosphate groups, mimicking the action of histones and polyamines in uiuo. Recent studies by Herzog and Soyer (1983) suggest that the helical compaction of isolated dinoflagellate chromatin is closely related to the level of Ca2+and Mg2+in the local environment.

THE DINOFLAGELLATE CHROMOSOME

25 1

IX. GENERAL DISCUSSION The preceeding account serves to emphasise the variety of experimental approaches that have been adopted in the study of dinoflagellate chromosomes and their constituent chromatin. The results obtained demonstrate that these structures have a number of remarkable features, and are different in a number of respects from the chromosomes of other eukaryote cells. These different aspects will be considered under three main headings: (1) dinoflagellate chromosome models, (2) prokaryote and eukaryote affinities, and (3) intermediate status and phylogeny. A. DINOFLAGELLATE CHROMOSOME MODELS

Models of dinoflagellate chromosome structure are derived in part from microscopical examination, but also relate to studies on cell cycle, macromolecular composition and characterisation, genetic activity, and X-ray microanalysis. These models fall into four main categories, depending on the level of organisation to which they apply. 1. Chromosome General Structure The toroidal chromonema model of Oakley and Dodge (1979) represents perhaps the simplest and most basic interpretation of light microscope and electron microscope section and whole-mount images of typical interphase chromosomes, and has been described previously (Section IV,B,3,a; Fig. 4a). Other models of general chromosome structure have also been proposed, including one by Spector et al. (1981a) for chromosome organisation in the uncoiled state at the peak of DNA replication. Spector et al. (1981a) and Spector and Triemer (1981) suggest that observation of dinoflagellate chromosomes in the unwound state provides a particularly useful opportunity to decipher structure, and propose a model consisting of a central core of 9.0-nm fibres, surrounded by two major helices which are connected by a series of fibres.

2 . DNA Fibril Arrangement Several models have been proposed to account for the observed chromosome structure in terms of constituent DNA fibrils. These have been summarised by numerous authors, including Livolant and Bouligand (1978), Oakley and Dodge, (1979), and Spector and Triemer (1981). The earliest model, proposed by Grasse and Dragesco (1957), considered the dinoflagellate chromosome to be composed of DNA fibrils ar-

252

D. C . SIGEE

ranged in major and minor helices. This model was modified by Giesbrecht (1961) by proposing that the minor spiral of the previous model was composed of fibrils which themselves spiraled. Giesbrecht (1965) amended his earlier model to propose a serpentine coiling model, in which the chromosome contains a backbone to support the loops of DNA comprising the chromonema. Grasse et al. (1965a) proposed a further model in which the fibrils making up the chromosome were regarded as individual units (chromonemata) arranged in a hemihelical manner. The development of a whole-mount technique lead Haapala and Soyer (1973) to propose a model consisting of a large number of circular chromatids. This model has been described in Section IV,B,3,a and is shown in Fig. 17a. The arrangement of DNA fibrils within the dinoflagellate chromosome has thus been the subject of wide and varied interpretation. In so far as these models are based on the detailed arrangement of fibrils within fixed embedded material, or extracted whole-mount preparations, the problem of possible artefact should be borne in mind. The various models fall into two main categories, depending on whether the dinoflagellate chromosome is envisaged as being composed of a multiplicity of separate DNA strands (polytenic) or a single (or few) strands. The models of Grasse et al. (1965a,b) and Haapala and Soyer (1973) are polytenic, while Oakley and Dodge (1979) propose that the toroidal chromonema is composed of just 1-5 strands. The elegant studies of Haapala and Soyer (1973), with the clear demonstration of circular chromatids, provide firm support in favour of polyteny. Apart from this, however, most of the evidence would oppose the concept of polyteny, including evidence for haploid DNA (Section VII,B), substantial levels of complex (non-repeated) DNA (Section VII,B), and viscoelastic measurements made by Allen et al. (1974) which suggest only 1-5 molecules of DNA per chromosome. The question of polyteny versus non-polyteny is not resolved at the present time, though most authors consider the evidence against polyteny to be conclusive. The most compelling evidence against polyteny is the haploid nature of the genome. If the genetically active DNA is restricted to peripheral fibrils, however, then the major part of the chromosome could be organised on a polytenic basis. If this interpretation were correct, then the haploid system in dinoflagellates would conceal a polytenic organisation of the condensed, genetically inactive DNA, and the chromosome as a whole could be regarded as partially polytenic. The various models that have been proposed for DNA fibril arrangement appear to relate particularly to the main body of the chromosome and tend to ignore the peripheral fibrillar system.

T H E DINOFLAGELLATE CHROMOSOME

253

3. Liquid Crystal State of Dinoflagellate Chromatin In the previous section, models of chromosome structure attempted to define the linear arrangement of DNA molecules in the dinoflagellate chromosome in relation to the fibrillar patterns observed in electron microscope images. Bouligand et al. (1968) adopted a different approach, by carrying out a mathematical analysis of serial sections, and proposing a twisted filament model in which the chromatin was viewed as a crystalline structure. They therefore interpreted the fibrillar arrangement more in terms of a physical state than in terms of gross morphology. According to Bouligand et al. (1968) and Livolant and Bouligand (1978), the nested arcs seen in longitudinal sections arise from a continuous twist in a system of superimposed layers of parallel filaments. A diagram of how this might occur is shown in Fig. 17b. On this interpretation, the arcs of fibrils seen in ultrathin section (e.g. Fig. 4c) are an illusion created by overlapping filaments. The DNA filaments are not genuinely bent, but are straight within the body of the chromosome. Evidence in support of the twisted filament model is provided by goniometric analysis of ultrathin sections, with demonstration of inversion of the arcs in relation to the angle of tilt, and analysis of fibrillar orientation in relation to chromosome defects (Livolant and Bouligand, 1978). Although Livolant and Bouligand (1978) initially believed that the question of filament topology could only be resolved by analysis of spread chromosomes, they have since considered (Livolant and Bouligand, 1980) that such preparations have a structure which is profoundly different from that present in the intact nucleus. Whether or not the interpretation of fibrillar arrangement in fixed chromatin by Livolant and Bouligand (1978) is correct, the proposal that dinoflagellate DNA occurs in a crystalline state is persuasive, and is supported by analysis of fresh unfixed chromosomes (Livolant, 1978, 1984a) and comparison with in uitro studies on isolated DNA (Livolant, 1984a). The crystalline condition of dinoflagellate chromatin is referred to as a cholesteric liquid crystal, and is typical of mixtures formed by stiff, elongated molecules. Cholesteric DNA in viuo appears to be particularly characteristic of situations where histones wre absent, including bacteria, dinoflagellates, mitochondria (Livolant, 1984a), and mammalian sperm (Livolant, 1984b). The concept of a tightly packed crystalline dinoflagellate chromosome is particularly attractive, since it relates directly to the fully hydrated in uiuo condition of the chromatin, is in accordance with the absence of transcription in the main body of the chromosome, and, by analogy to in uitro studies (Livolant, 1984a) on DNA-salt interactions, explains why cations are a major integral structural component.

254

D. C. SIGEE

4 . Cation-Chromatin Interactions The X-ray microanalytical studies of Sigee and Kearns (Section VII) have led to the formulation of several simple models to explain the associations of cations with dinoflagellate chromatin DNA and protein matrix under various conditions. These models propose a stabilising, structural role for the cations, and have been proposed in relation to the effects of nuclease enzymes on the ionic composition of chromatin (Sigee and Kearns, 1981~) and the effects of various preparative treatments (Sigee, 1983a). The model shown in Fig. 17c is part of a larger scheme (Sigee, 1983a)to explain variations in X-ray microanalytical data and appearance in unfixed and fixed chromatin. It is based on a simple concept, envisaging a bridging (DNA-protein) role for divalent cations and a purely electrostatic role for monovalent cations. B . PROKARYOTE A N D EUKARYOTE AFFINITIES

Dinoflagellate chromatin shows a number of specific and separate similarities to the chromatin of prokaryotes and other eukaryotes, as noted in the introduction and at various points in the article. These have been widely discussed in the literature.

1 . Prokaryote Afjnities The significance of some of the similarities with bacterial chromatin have been viewed critically by various authors. The kinetic role and significance of nuclear membranes in chromosome division and migration has been questioned by Soyer (1981), for example, with the demonstration of intranuclear cables in freeze-substituted material. The significance of low levels of basic protein, and the apparent absence of histones, has been discussed by Spector and Triemer (1981), who point out that low levels also occur in several fungi and that histones were initially reported absent from this group. The absence of histones in dinoflagellates is now, however, well documented and does appear to represent a genuine difference between dinoflagellates and other eukaryote cells. The only other situation where eukaryote chromatin lacks histones is in the highly condensed chromatin of sperm cells, where histones have been replaced by histone-derived protarnines (Kolodny, 1980; Balhorn, 1982). The absence of histones in dinoflagellates is correlated with an absence of nucleosomes in chromatin that has been extracted and visualised by conventional techniques (see Section IV,B,3,b). This is similar to the situation in bacteria (Stonington and Pettijohn, 1971; Worcell and Burgi, 1972). It is interesting to note, however, that nucleosome-like structures

255

THE DINOFLAGELLATE CHROMOSOME

have now been observed in the chromatin of partially disrupted bacterial cells (Griffith, 1976; Pettijohn, 1982), and it is possible that similar highly unstable structures may be demonstrated in dinoflagellates with more careful preparation techniques. The suggestion by Kearns and Sigee (1980), that divalent cations have a special role (in the absence of histones) in the stabilisation of dinoflagellate DNA, has been noted earlier and may represent a further similarity with prokaryote cells. Similarities certainly exist in the localisation of bound cations to the chromatin in both types of cells. Recent autoradiographic studies on the uptake of 63Ni2+in the bacterium Pseudomonas tabaci (Al-Rabaee and Sigee, 1984; Sigee et al., 1984) reveal a localised occurrence of silver grains over centrally located masses of condensed DNA. X-Ray microanalytical studies on extracted bacterial chromatin (Fig. 18) show a similar range of bound cations to that seen in the chromatin of dinoflagellates (El-Masry and Sigee, unpublished data). Evidence that divalent cations may be important in the stabilisation of bacterial chromatin is provided by the in uiuo experiments of Whitefield and Murray (1956), who showed that nucleoid condensation could be P

K I

I

Ca

CHROMATIN

I I I I

Fe I I I

I

Cu I I

Zn I 1

I

I

I

1

FORMVAR CONTROL

&."-4-d

A + + , 5

X-ray energy (keV1

Fig. 18. X-ray emission spectrum from extracted chromatin of Pseudomonas tabaci. Characteristic X-ray emission spectrum of supercoiled bacterial DNA, extracted using a modification of the Sat0 and Miura technique (Eiochim. Eiophys. Acta 72, 619-629). (ElMasry and Sigee, unpublished observations.)

10

256

D. C. SIGEE

sntrolled by ionic manipulation. Indirect evidence is also provided be the classical studies of Ryter and Kellenberger (Kellenberger, 1962; Kellenberger et al., 1958; Ryter and Kellenberger, 1958) with the demonstration that addition of cations during fixation maintained the fibrils in a dispersed, non-precipitated state. The in uiuo general organisation and activity of the bacterial genome also shows close similarities to that of a single dinoflagellate chromosome (Al-Rabaee and Sigee, 1984), as illustrated in Fig. 19. Ryter and Chang (1975) used high-resolution autoradiography to show that DNA transcription does not occur in the central condensed nucleoid region, but is restricted to the peripheral loops which extend into the surrounding ribosomal protoplasm (Fig. 19a). This division of function in terms of transcriptive and non-transcriptive DNA parallels the situation seen in the dinoflagellate chromosome (Fig. 19b). The 63Ni2+autoradiographic studies of Al-Rabaee and Sigee (1984) also suggest a localisation of cations within the bacterial genome to the central nucleoid core. This corresponds to the localised occurrence of cations seen within the main body of

central nucleoid I

ribosomal protoplasm

I I

I

a BACTERIAL CHROMOSOME

/

,

Central Core - condensed DNA - non-active in transcription - high cation content

.

\

Peripheral Strands - diffuse DNA - active in transcription I I

DINOFLAGELLP

22irq

~

CHROMOSOME

Fig. 19. General organisation of the bacterial (a) and dinoflagellate (b) chromosome. The bacterial chromosome is shown within the context of the whole bacterial cell, the dinoflagellate chromosome in isolation.

THE DINOFLAGELLATE CHROMOSOME

257

the dinoflagellate chromosome, and suggests that the central, genetically inactive core of both structures is stabilised by a cationic system.

2 . Eukaryote Aflnities Dinoflagellates are eukaryote cells, as defined by the presence of a nuclear membrane (Dougherty, 1957), and in terms of their general size and complexity. Other eukaryote characteristics include the arrangement of the chromatin into discrete chromosomes, the presence of substantial amounts of repeat-sequence DNA, the presence of UI-U6 snRNAs, homology in 5 S RNA sequences, and the presence of a distinct peak or ‘S’ phase of DNA synthesis during the cell cycle. These features have persuaded various authors that dinoflagellates and their constituent chromatin are fundamentally eukaryote in type (Loeblich, 1976; Spector and Triemer, 1981; Hinnebusch et al., 1981; Reddy et al., 1983). Important differences, particularly involving characteristics referred to earlier as prokaryote in type, do however occur. The chromatin-associated proteins are quite different, and related to this, the ability to extract the chromatin matrix from intact cells with buffer appears to be a specific dinoflagellate characteristic. Attempts to extract the matrix from condensed chromatin of eukaryote cells such as Euglena uiridis and Prymnesium paruum (Sigee, unpublished observations) have so far failed. Divalent cations, present and important in the stabilisation of bacterial and dinoflagellate chromatin, also have a major structural role in eukaryote cells. X-ray microanalytical studies on intact eukaryote cells, and analysis of extracted chromatin, have generally suggested (Kearns and Sigee, 1980; Sigee, 1984a) lower levels of cations and less variety compared to dinoflagellates. The importance of cations in controlling the conformation of eukaryote chromatin is shown by the experiments of Cole (1967) on isolated chromosomes and nuclei, and the work of Pruitt and Grainger (1980) on the reversible disassembly of thick chromatin fibres into single strands at low cation levels. The stabilising role of cations in normal eukaryote cells appears to operate mainly at the higher order, three-dimensional level. This has been shown by the work of Finch and Klug (1976) and Renz et al. (1977) with the demonstration that the strand of nucleosomes can be compacted at low levels of Mg2+ions, or higher levels of monovalent cations. The stabilising role of cations in normal eukaryote cells would appear to be quite distinct from that in dinoflagellates and bacteria, since the cation-mediated condensation of eukaryote DNA requires the presence of H1 histone. Although superficially similar, the general structure of the dinoflagellate chromosome is fundamentally different from that of normal eukaryote

258

D. C. SIGEE

cells in terms of banding characteristics, birefringence properties, and chromonemal organisation (see Section IV).

C. INTERMEDIATE STATUS AND PHYLOGENY

It is now generally accepted that dinoflagellates should be regarded as true eukaryote cells, which are unusual in the possession of a number of prokaryote characteristics. Controversy arises as to whether dinoflagellates should be regarded as a primitive or a degenerate eukaryote group. As primitive eukaryote cells, they are considered to have evolved at an early stage of the phylogeny of the eukaryote cell (Allen et al., 1975a; Loeblich, 1976; Oakley and Dodge, 1976; Rae and Steele, 1978; Rizzo and Burghardt, 1982). Such a divergence would have occurred prior to the evolution of histones, but after the evolution of eukaryote-type 5 S RNA, U I - U ~snRNAs, repeat-sequence DNA (Allen et al., 1975a), and the nucleolar system. As degenerate eukaryote cells, they are considered to have evolved at a late stage in the evolution of the eukaryote cell, by secondary loss of such features as the presence of histones and correlated chromatin ultrastructure (Hinnebusch et al., 1981; Reddy et al., 1983). The viewpoint reached in this article is that they are most probably primitive rather than degenerate eukaryote cells. Support for this is based on four main points.

1. Prokaryote-type nuclear characteristics involve a range of features relating to the nature of the chromatin and cell division. Many of these would have to be derived independently during phylogeny from a normal ancestral eukaryote on the degeneracy theory, which seems most unlikely. 2. The dinoflagellate chromosome appears to be so fundamentally different from the normal eukaryote chromosome that it is difficult to see how there can be any homology, which is implied by the late origin of the group in the degeneracy theory. 3. In other eukaryote organisms, there appears to be a universal occurrence of histones and stable nucleosomes (Kornberg, 1977; Horgen and Silver, 1978; Rizzo and Burghardt, 1980). In view of this, and the highly conserved nature of the nucleosome throughout the eukaryote kingdoms, it seems most unlikely that evolution of just one eukaryote group would involve complete secondary loss of these characters. 4.The fossil record (Schopf, 1968), and diversity of present day dinoflagellate species, are both consistent with an early rather than a late origin of this group during evolution.

THE DINOFLAGELLATE CHROMOSOME

259

The evolution of the dinoflagellate genome from that of a primitive eukaryote ancestor may have involved a number of distinct features, including the following: 1 . A considerable increase in the level of nuclear DNA. This major phylogenetic tendency is a particular feature of the dinoflagellates [Sigee, 1984b), with the attainment of exceptionally high levels of genetically inactive DNA, which forms the major structural component of dinoflagellate chromosomes. Other adaptations towards high DNA levels in this group include the characteristic early replication of DNA in the cell cycle, the phylogenetic incorporation of a non-dinoflagellate symbiont in several species (Glenodiniumfoliaceum and Peridinium balticum), and the apparent formation of polyploid or aneuploid series in some genera (see Section I11,A). The significance of these high DNA levels may simply be that they reflect a particular type of evolutionary selection. This has been referred to as K selection (MacArther and Wilson, 1967; Cavalier-Smith, 1978), and leads to large cells, long life cycles. It is typical of organisms that are adapted to a stable, crowded environment, such as the plankton-rich surface waters of lakes and oceans. 2. The permanent incorporation of modified bases such as 5-hydroxymethyluracil within the DNA as part of a modification-restriction system to protect against the consequences of invasion by foreign DNA. 3. Retention of a primitive cationic non-histone stabilisation system in the chromatin. In other eukaryote cells, the evolution of histones was correlated with the evolution of a completely new type of chromatin packaging which was capable of reversible condensatioddecondensation during the cell cycle. This permitted a temporal separation between metabolic functions (DNA replication and transcription during interphase) and genome separation (during cell division), and was a major step in eukaryote evolution.

In dinoflagellates the chromatin condensation is not reversible, though a limited uncoiling occurs in some species during the peak of DNA replication. The permanent separation into condensed core and peripheral fibrils throughout the whole of the cell cycle means that the major distinction between genetically active and nonactive DNA is fixed in relation to chromosome structure and not temporal activity. Perhaps the closest approximation to dinoflagellate chromosomes in the rest of the eukaryote kingdom is the lampbrush chromosomes of amphibian oocytes, where actively transcriptive loops extend from a central, genetically inactive axis of tightly folded (condensed) chromatids (Callan, 1963). This comparison is entirely by analogy, however, since lampbrush

260

D. C. SIGEE

chromosomes must be regarded as a relatively recent evolutionary adaptation towards a phase of transcription within an extended meiotic prophase. The general conclusion reached in this article is that dinoflagellates are a genuinely primitive eukaryote group and that their chromosomes are fundamentally different from those of all other eukaryotes. On this interpretation, the evolution of the dinoflagellate chromosome represents part of an alternative phylogenetic route towards the attainment of the present day eukaryote state.

ACKNOWLEDGEMENTS I gratefully acknowledge the contribution of the work of S . Filfilan, L. P. Kearns, R. AlRabaee, H. El-Masry, and N . Harrison to the studies carried out at Manchester and quoted in this article. I am also grateful to Prof. J. D. Dodge and Drs. Y . Bouligand, 0. K . Haapala, F. Livolant, and M. 0. Soyer for generously giving me permission to reproduce previously published material.

REFERENCES Allen, J. R., Roberts, T. M., Tuttle, R. C., Klotz, L. C., and Loeblich, A. R. (1974). J . Protozool. 21, 426. Allen, J. R., Roberts, T. M., Loeblich, A. R., and Klotz, L. C. (1975a). Cell 6, 161-169. Allen, J. R., Tuttle, R. C., Hedberg, M. F., Klotz, L. C., and Loeblich, A. R. (1975b). J . Phycol. (Suppl.) 11, 15. Al-Rabaee, R. H., and Sigee, D. C. (1984). J . Cell Sci. 69, 87-105. Anderson, D. M., and Morel, F. M. (1978). Limnol. Oceanogr. 23, 283-295. Babillot, C. (1969). J . Microsc. (Paris) 9, 485-502. Babillot, C. (1970). C . R . Acad. Sci. (Paris) 271, 828-841. Balhorn, R. (1982). J . CellBiol. 93, 298. Beam, C. A. and Himes, M. (1974). Nature (London) 250, 435-436. Beam, C. A., Himes, M., Himmelfarb, J., and Link, C. (1977). Genetics 87, 19-32. Beermann, W., and Pelling, C. (1965). Chromosoma 16, 1-21. Bhargava, M. M., and Halvorson, H. 0. (1971). J . Cell Biol. 49, 423-429. Bold, H.C., and Wynne, M. J. (1978). “Introduction to the Algae.” Prentice Hall, New York. Bouligand, Y . , Soyer, M. O., and Puiseux-Dao, S. (1968). Chromosomu 24, 251-287. Braarud, T. (1955). Verh. Int. Ver. Limnol. 12, 811-813. Brasier, M. D. (1980). “Microfossils.” Allen & Unwin, London. Britten, R. J., and Kohne, D. E. (1968). Science 161, 529-540. Bujak, J. P., and Williams, G . L. (1981). Cun. J . Bot. 59, 2077-2087. Cachon, J., and Cachon, M. (1974). C . R . Acad. Sci. (Paris) 278, 1735-1737. Cachon, J., and Cachon, M. (1977). Chromosoma (Berlin) 60, 231-251. Cachon, J. and Cachon, M. (1979). Protistenkunde 122, 267-284.

T H E DINOFLAGELLATE CHROMOSOME

26 1

Calandra, F. (1964). C.R. Acad. Sci. (Paris) 258, 4112-4114. Callan, H. G. (1963). I n t . Rev. Cytol. 15, 1-34. Cavalier-Smith, T. (1978). J . Cell Sci. 34, 247-278. Cole, A. (1967). I n “Theoretical and Experimental Biophysics” (A. Cole, ed.). Dekker, New York. Daneholt, B., and Edstrom, J. E. (1967). Cytogenetics 6, 350. Darlington, C. D., and A. P. Wylie (1955). “Chromosome Atlas.” Allen & Unwin, London. Diwald, K. (1937). Nora 132, 174-192. Dodge, J. D. (1963a). Bot. Mar. 5, 121-127. Dodge, J. D. (1963b). Arch. Protistenkd. 106, 442-452. Dodge, J. D. (1964). Arch. Mikrobiol. 48, 66-80. Dodge, J. D. (1965). Exerpta Med. Int. Congr. Ser. 91, 399. Dodge, J. D. (1966). In “The Chromosomes of the Algae” (M. B. Godward, ed.). Arnold, London. Dodge, J. D. (1971). Bot. Reu. 37, 481-508. Dodge, J. D. (1972). Protoplasma 75, 285-302. Dodge, J. D. (1975). Phycologia 14, 253-263. Dodge, J. D., and Crawford, R. M. (1970). Bot. J . Linn. SOC.63, 53-67. Dougherty, E. C. (1957). J . Prototool. 4 (Suppl.), 14. DuPraw, E. J. (1970). “DNA and Chromosomes.” Holt, New York. Dyson, R. D. (1978). “Cell Biology: A Molecular Approach.” Allyn & Bacon, Boston. Eickbush, T. H., and Moudrianakis, E. N. (1978). Cell 13, 295-306. Entz, G. (1921). Arch. Protistenkd. 43, 415-430. Entz, G. (1927). Arch. Protistenkd. 58, 344-440. Filfilan, S. A., and Sigee, D. C. (1977). J . Cell Sci. 27, 81-90. Finch, J. T., and Klug, A. (1976). Proc. Natl. Acad. Sci. U . S . A . 73, 1897-1901. Fincham, J. R.,and Day, P. R. (1971). “Fungal Genetics.” Bot. Mon. 4. Blackwell, Oxford. Fine, K. E., and Loeblich, A. R. (1974). J . Phycol. lO(Suppl.), 13-14. Franker, C. K. (1970). J. Phycol. 6, 299-305. Franker, C. K. (1971). J . Phycol. 7, 165-169. Franker, C. K. (1972). J. Phycol. 8, 264-268. Franker, C. K., Sakhrani, L. A., Prichard, C. D., and Lamdan, C. A. (1974). J. Phycol. 10, 91-94. Galleron, C., and Durrand, A. M. (1979). Protoplasma 100, 155-165. Gavrila, L., Soran, V., Ahmad, S., Bercea, V., Spirchez, C. T., and Tacina, F. (1981). Cytologia 46, 1-14. Giesbrecht, P. (1961). Zbl. Bakt. 1 . Abt. Or&. 183, 1-44. Giesbrecht, P. (1962). Zbl. Bakt. Parasitenkunde 187, 452-498. Giesbrecht, P. (1965). Z . Naturforsch. 20, 927-930. Grasse, P. P., and Dragesco, J. (1957). C.R. Acad. Sci. (Paris) 245, 1447-1451. Grasse, P. P., Hollande, A., Cachon, J., and Cachon-Enjumet, M. (1965a). C.R. Acad. Sci. (Paris) 260, 1743-1747. Grasse, P. P., Hollande, A., Cachon, J., and Cachon-Enjumet, M. (1965b). C.R. Acad. Sci. (Paris) 260, 6975-6977. Griffith, R. G . (1976). Proc. Natl. Acad. Sci. U . S . A . 73, 563-567. Grell, K. G., and Wollfarth-Botterrnann, K. E . (1957). Z . Zellforsch. Mikrosc. Anat. 47, 717. Haapala, 0. K., and Soyer, M. 0. (1973). Nature (London) New Biol. 244, 195-197. Haapala, 0. K., and Soyer, M. 0. (1974a). Hereditas 76, 83-90. Haapala, 0. K., and Soyer, M. 0. (1974b). Hereditas 78, 141-145.

262

D. C. SIGEE

Hall, T. A. (1971). In “Physical Techniques in Biological Research” (G. Oster, ed.). Academic Press, New York. Haller, G., de, Kellenberger, E., and Rouiller, C. (1964). J. Microsc. (Paris) 3, 627-642. Hamkalo, B. A., and Rattner, J. B. (1977). Chromosoma (Berlin) 60, 39-47. Hawthorne, D. C., and Mortimer, R. K. (1960). Genetics 45, 1085-1 110. Herzog, M., and Soyer, M. 0. (1981). Eur. J. Cell Biol. 23, 295-302. Herzog, M., and Soyer, M. 0. (1982). Eur. J . Cell B i d . 27, 151-155. Herzog, M., and Soyer, M. 0. (1983). Eur. J . Cell Biol. 30, 33-41. Himes, M., and Beam, C. A. (1975). Proc. Natl. Acad. Sci. U.S.A.72,4546-4549. Hinnebusch, A. G., Klotz, L. C., Blanken, R., and Loeblich, A. R. (1981). J. Mol. Evol. 17, 334-347. Holliday, R. (1970). Symp. SOC. Gen. Microbiol. 20, 359-380. Holm-Hansen, 0. (1969). Science 163,87-88. Holt, J. R., and Pfeister, L. A. (1982). Am. J. Bot. 69, 1165-1168. Horgen, P. A., and Silver, J. C. (1978). Annu. Rev. Microbial. 32, 249-284. Jones, G. J., Nichols, P. D., and Johns, R. B. (1983). J . Phycol. 19,416-420. Jorgensen, K. F., van de Sande, J. H., and Lin, C. C. (1978). Chromosoma (Berlin)68,287302. Kallen, R. G., Simon, M., and Marmur, J. (1962). J. Mol. Biol. 5 , 248. Karentz, D. (1983). J . Protozool. 30, 581-588. Kearns, L. P. (1981). Ultrastructural and quantitative X-ray microanalytical studies on selected dinoflagellates. PhD thesis, University of Manchester. Kearns, L. P., and Sigee, D. C. (1979). Experientia 35, 1332-1333. Kearns, L. P., and Sigee, D. C. (1980). J . Cell Sci. 46, 113-127. Kellenberger, E. (1962). In “The Interpretation of Ultrastructure” (R. J. Harris, ed.). Academic Press, New York. Kellenberger, E., Ryter, A., and Sechaud, J. (1958). J . Biophys. Biochem. Cytol. 4, 671678. Kim, Y.S., and Martin, D. F. (1974). Phytochernistry 13, 533-538. Kohler-Wieder, R. (1937). Ost. Bot. Z. 86, 198-220. Kolodny, G. M. (1980). “Eukaryote Gene Regulation,” Vol. 2. CRC Press, Boca Raton, Florida. Kornberg, R. D. (1977). Annu. Rev. Biochem. 46,931-954. Kubai, D. F., and Ris, H. (1969). J . Cell Biol. 40, 508-528. Leadbeater, B. (1967). An ultrastructural study of the morphology and division of some marine dinoflagellates. PhD thesis, University of London. Leadbeater, B., and Dodge, J. D. (1967). J . Gen. Microbiol. 46, 305-314. Leedale, G. F. (1967). “Euglenoid Flagellates”. Prentice Hall, New York. Leutenegger, S. (1977). Mar. Biol. 44, 157-164. Livolant, F. (1978). Chromosoma (Berlin) 68, 45-58. Livolant, F. (1984a). Eur. J . Cell Biol. 33. 300-311. Livolant, F. (1984b). Tissue Cell 16, 535-555. Livolant, F., and Bouligand, Y.(1978). Chromosoma (Berlin) 68, 21-44. Livolant, F., and Bouligand, Y. (1980). Chromosoma (Berlin) 80, 535-555. Loeblich, A. R. (1968). Proc. N . Am. Paleontol. Convention, Chicago 705-788. Loeblich, A. R. (1976). J. Protozool. 23, 13-28. Loper, C. L., Steidinger, K. A., and Walker, L. M. (1980). Trans. A m . Microsc. SOC. 99, 343-346. MacArthur, R. H., and Wilson, E. 0. (1967). “The Theory of Land Geography.” Princeton Univ. Press, Princeton, N.J.

THE DINOFLAGELLATE CHROMOSOME

263

McLeish, J., and Sunderland, N. (1961). Exp. Cell Res. 24, 527-540. Matthys-Rochon, E. (1979). B i d . Cell. 35, 313-320. Mendiola, L. R., Price, C. A., and Guillard, R. R. (1966). Science 153, 1661-1663. Meselson, M., Yuan, R., and Heywood, J. (1972). Annu. Rev. Biochem. 41, 447. Murphy, E. B., Steidinger, K. A., Roberts, B. S., Williams, J . , and Jolley, J. W. (1975). Limnol. Oceanogr. 20,481-486. Oakley, B. R., and Dodge, J. D. (1974). J . Microsc. 108, 79-87. Oakley, B. R., and Dodge, J. D. (1976). Cytobios 17, 35-46. Oakley, B. R., and Dodge, J. D. (1979). Chromosoma (Berlin) 70, 277-291. Parenti, F., Brawerman, G., Preston, J. F., and Eisenstadt, J. M. (1967). Biochim. Biophys. A c f a 195, 234-243. Parke, M., and Dodge, J. D. (1976). J. Mar. Biol. Assoc. U . K . 56, 527-594. Peters, T., and Ashley, C. A. (1967). J. Cell Biol. 33, 53-60. Pettijohn, D. E. (1982). Cell 30, 667-669. Pfeister, L. A. (1977). J. Phycol. 13, 92-95. Pfeister, L. A,, Timpano, P., Skvarla, J. J., and Holt, J. R. (1984). Am. J . B o f . 71, 11211127. Pruitt, S. C., and Grainger, R. M. (1980). Chromosoma (Berlin) 78, 257-274. Rae, P. M. (1970). J . CellBiol. 46, 101-113. Rae, P. M. (1973). Proc. N a f l . Acad. Sci. U . S . A . 70, 1141-1145. Rae, P. M. (1976). Science 194, 1062-1064. Rae, P. M., and Steele, R. E. (1978). BioSystems 10, 37-53. Reddy, R., Spector, D., Henning, D., Liu, D., Liu, M.-H., and Busch, H. (1983). J. B i d . Membr. 258, 13965-13969. Renz, M., Nehls, P., and Hozier, J. (1977). Proc. Nufl. Acad. Sci. U . S . A . 74, 1879-1883. Ris, H. (1962). I n “The Interpretation of Ultrastructure” (R. J . Harris, ed.), pp. 69-88. Academic Press, New York. R~ZZO, P. J. (1976). J . Mol. Evol. 8, 79-94. Rizzo, P. J. (1979). J. Profozool. 26, 290-294. Rizzo, P. J. (1981). BioSystems 14, 433-443. Rizzo, P. J. (1982). J . Profozool. 29, 98-103. Rizzo, P. J., and Burghardt, R. C. (1980). Chromosoma (Berlin) 76, 91-99. Rizzo, P. J., and Burghardt, R. C. (1982). BioSysfems 15, 27-34. Rizzo, P. J., and Nooden, L. D. (1972). Science 178,796-797. Rizzo, P. J., and Nooden, L. D. (1973). J. Prorozool. 20, 666-672. Rizzo, P. J., and Nooden, L. D. (1974a). Biochim. Biophys. A c f a 349, 402-414. Rizzo, P. J., and Nooden, L. D. (1974b). Biochim. Biophys. A c f a 349, 415-427. Rizzo, P. J., Choi, J., and Morris, R. L. (1984). J . Phycol. 20, 95-100. Rizzo, P. J., Jones, J., and Ray, S. M. (1982). J . Prorozool. 29, 217-222. Roberts, T. M., Tuttle, R. C., Allen, J. R., and Loeblich, A. R. (1974). Nature (London)248, 446-447. Ryter, A. (1968). Bacferiol. Rev. 32, 39-54. Ryter, A., and Chang, A. (1975). J. Mol. Biol. 98, 797-810. Ryter, A., and Kellenberger, E. (1958). Z. Nafurforsch. 13, 597-609. Sarjeant, W. H. (1978). Palynology 2, 167-197. Schopf, J. W. (1968). J. Paleontol. 42, 651-688. Shapiro, H. S. (1976). In “Handbook of Biochemistry and Molecular Biology” (G. V. Fasman, ed.), pp. 241-283. CRC Press, Cleveland. Shyam, R., and Sarma, Y. S. (1978). Bot. J. Linn. Soc. 76, 145-159. Sigee, D. C. (1976). J . Microsc. 108, 325-329.

264

D. C . SIGEE

Sigee, D. C. (1982). Protoplasma 110, 112-120. Sigee, D.C. (1983a). Scan. Electron Microsc. IV, 1779-1793. Sigee, D. C. (1983b). Protoplasma 117, 185-195. Sigee, D.C. (1984a). Bot. J. Linn. SOC.88, 127-147. Sigee, D. C. (1984b). BioSystems 16, 203-210. Sigee, D.C., and Kearns, L. P. (1980). In “Nickel Toxicology” ( S . Brown and W. Sunderman, eds.), pp. 163-166. Academic Press, New York. Sigee, D. C., and Kearns, L. P. (1981a). Protoplasma 105, 213-223. Sigee, D.C., and Kearns, L. P. (1981b). Tissue Cell 13,441-451. Sigee, D.C., and Kearns, L. P. (1981~).Cytobios 31, 49-65. Sigee, D.C., and Kearns, L. P. (1982a). Cytobios 33, 51-64. Sigee, D.C., and Kearns, L. P. (1982b). J. Biochem. Biophys. Methods 6,23-30. Sigee, D. C., Al-Rabaee, R. H., and El-Masry, H. (1984). I n “Nickel Toxicology” (S. Brown and W. Sunderman, eds.). Academic Press, New York, in press. Silva, E. S. (1962). Protistologica 14, 113-119. Silverberg, B. A., Stokes, P. M., and Ferstenberg, L. B. (1976). J. Cell Biol. 69, 210-214. Skaar, H., Phus, E., and Gullvag, B. M. (1973). Nature (London) 241, 215-216. Skoczylas, On (1958). Arch. Protistenkd. 103, 197-225. Soyer, M. 0. (1969). C.R. Acad. Sci. 268, 2082-2084. Soyer, M. 0. (1977a). Biol. Cell. 30, 297-300. Soyer, M. 0. (1977b). Biol. Cell. 29, 26a. Soyer, M. 0. (1978a). BioSystems 10, 55-58. Soyer, M. 0. (1978b). Vie Milieu 28, 461-472. Soyer, M. 0. (1981). BioSystems 14,299-304. Soyer, M. O., and Escaig, J. (1980). Protistologica 16,485-495. Soyer, M. O., and Haapala, 0. K. (1973). J . Microsc. 18, 267-270. Soyer, M. O., and Haapala, 0. K. (1974a). J. Microsc. 19, 137-146. Soyer, M.O., and Haapala, 0. K. (1974b). Chromosoma (Berlin) 47, 179-192. Soyer, M.O., and Haapala, 0. K. (1974~).Histochemistry 42, 239-246. Spector, D.L., and Triemer, R. E. (1981). BioSystems 14, 289-298. Spector, D.L., Vasconcelos, A. C., and Triemer, R. E. (1981a). Protoplusma 105, 185-194. Spector, D.L., Pfeister, L. A., and Triemer, R. E. (1981b). Am. J. Bot. 68, 34-43. Steidinger, E. (1975). Enuiron. Lett. 9, 129-139. Stewart, J. M., and Beck, J. S. (1967). J. Protozool. 14, 225-231. Stonington, 0.G., and Pettijohn, D. E. (1971). Proc. Natl. Acad. Sci. U . S . A . 68, 6-9. Tangen, K. (1977). Sarsia 63, 123-133. Taylor, F. J. (1975). Phycologia 14,45-47. Thoma, F., Koller, Th., and Klug, A. (1979). J . Cell Biol. 83, 403-426. Tippit, D. H., and Pickett-Heaps, J. D. (1976). J . Cell Sci. 21, 273-289. Triemer, R. E. (1982). J. Phycol. 18, 399-411. Tuttle, R. C., and Loeblich, A. R. (1974a). Science 185, 1061-1062. Tuttle, R. C., and Loeblich, A. R. (1974b). J . Phycol. lO(Suppl.), 16. Tuttle, R. C., and Loeblich, A. R. (1975). J . Phycol. ll(Suppl.), 15 van Goor, A. C. (1918). Arch. Protist. 39, 147-208. von Stosch, H. A. (1973). Br. Phycol. J. 8, 105. White, A. W. (1978). J. Phycol. 14, 475-479. Whitefield, J. F., and Murray, R. G . (1956). Can. J. Microbid. 2, 245-260. Worcel, A., and Burgi, E. (1972). J . Mol. Biol. 71, 127-147. Zederbauer, E. (1904). Verh. Dtsch. Bot. Ges. 22, 1-18.

AUTHOR INDEX

The numbers in italics indicate the pages on which names are mentioned in the reference lists.

A Abad, M. A,, 86, 87, 88, 96, 97 Adachi, R., 63, 93 Adams, W. H., 65, 70, 77, 100 Adelman, W. J., Jr., 69, 93 Ahmad, S., 209,261 Aitken, A., 19, 42 Akau, C. K., 87, 89, 97 Alam, M., 69, 82, 93, 99 Albuquerque, E. X., 68, 100 Ali-Khan, S. T., 111, 194 Alleman, H., 197 Allen, J. R., 207, 210, 222, 224, 238, 240, 252, 258, 260, 263 Allen, L. H., Jr., 4, 46 Al-Rabaee, R. H., 255, 256, 260, 264 Alscher, R., 5 , 22, 38, 39, 44 Alscher-Herman, R., 38, 39, 44 Amundson, R. G., 38,44 Anderson, D. M., 59, 93, 206, 260 Anderson, E. G., 123, 124, 194, 196 Anderson, H., 88, 97 Anderson, L. E., 2, 3, 5 , 6, 7, 8, 9, 10, 12, 14, 17, 22, 23, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38,39, 40, 41.42, 43, 44,45 Andersson, I., 109, 194 Andersson-Kotto, I., 109, 194 Andreo, C. S., 33, 35, 42 Anet, E. F. L. J., 64,75, 94 Arana, J. L., 5 , 46 Arber, W., 164, 198 Archer, B. G., 69, 93 Arnon, D. I., 14, 20,41 Arthur, A., 144, 171, 194, 202

Ashley, C. A., 248,263 Ashman, R. B., 130, 135, 183, 184, 194 Ashton, A. R., 2, 3, 8, 12, 17, 21, 22, 24, 25, 26, 27, 30, 31, 33, 35, 36,40 Aslam, K. M., 111, 120, 189, 202 Astrachan, N. B., 69, 93 Avato, P., 194 Avery, P., 188, 197 Avron, M., 5 , 6, 7, 8, 38, 39, 40

B Babillot, C., 214, 222, 260 Baccarini-Melandri, A,, 35, 43 Baden, D. G., 57, 86, 88, 89, 90, 93, 96 Badger, M., 5 , 40 Bagnis, R., 62, 63, 86, 87, 88, 93, 94, 95, 97, 101 Balange, A. P., 4,40 Baldwin, I. T., 91, 93, 99 Balhorn, R., 254,260 Bamberger, E. S., 36,40 Bandoni, R. J., 48, 99 Banner, A. H., 62, 80, 86, 87, 88, 93, 96, 101 Barchi, R. L., 85, 95 Barclay, P. C., 123, 133, 141, 194 Barr, M.,88, 93 Bartolini, G., 80, 95 Bassham, J. A., 5 , 23, 43, 44 Bates, H. A., 68, 93 Baur, E., 110, 117, 120, 121, 194 Beale, G. H., 116, 142, 196 Beam, C. A., 209, 234, 240,260, 262 Beasley, V. R., 50, 94

265

266

AUTHOR INDEX

Beck, E., 22, 45 Beck, J. S., 237,264 Beerrnann, W., 210,260 Beevers, H., 4, 44 Behrens, U., 129, 152, 161, 194 Ben-Bassat, D., 2, 3, 8, 23, 26, 27, 30, 34, 36,39, 40 Bender, W., 167, 200 Bennett, J., 4, 40, 43 Bennetzen, J. L., 131, 152, 155, 159, 160, 163, 168, 181, 189, 192, 194, 195, 202 Benoist, C., 162, 194 Bent, P. E., 94 Bercea, V., 209, 261 Beresford, A. M., 74, 95 Berg, D. E., 105, 144, 193, 194, 195 Berg, H., 121, 138, 185,202 Berg, W., 121, 146, 184, 185, 197, 199 Berger, L. R., 78, 97 Bergfeld, R., 118, 194 Bergmann, J. S., 87, 94 Berkowitz, G. A., 38, 40 Berry, J. A., 4, 45 Berstermann, A , , 18, 40 Bertram, I., 160, 197 Bhargava, M. M., 210,260 Bianchi, A., 123, 188, 194 Bianchi, F., 113, 114, 138, 139, 140, 142, 146, 167, 189, 194, 195, 197 Biggs, D. F., 68, 69, 94 Billings, W. H., 81, 94 Bishop, C. T., 64,75, 94 Bismuth, E., 5 , 6, 7, 41 Bjorkman, O., 5,40 Blackman, A. J., 79, 98 Blake, P. A., 88, 98 Blakeslee, A. F., 116, 194 Blanken, R., 241, 257, 258,262 Block, R. E., 57, 89, 90, 93 Blumbergvel Spalve, J., 155, 156, 158, 160, 202 Boag, S., 40 Boersma, E. A., 113, 114, 138, 167, 195 Bold, H. C., 207, 260 Bonas, U., 152, 155, 156, 159, 160, 161, 165, 166, 172, 177, 185, 186, 192, 194, 201 Borbely, G., 23, 30, 45 Borghi, B., 126, 201 Bgirresen, S., 109, 194

Botes, D. P., 56, 65, 75, 76, 77, 94, 99 Botstein, D., 105, 193, 195 Bouligand, Y.,206, 2 14, 250, 25 1, 253, 260, 262 Bourke, A. T. C., 74, 94 Bowden, J. P., 82, 99 Bowes, G., 4, 46 Boyer, G. L., 82, 83, 84, 94, 100 Boylan, D. B., 87, 96 Braarud, T., 206, 260 Bradbeer, J. W., 32,46 Brasier, M. D., 206, 260 Brawerman, G., 210, 263 Breathnach, R., 162, 194 Breazeale, V. D., 14, 45 Brememkamp, M., 126,201 Brennan, T., 2, 8, 12, 17, 26, 27, 29, 30, 36,40 Bresler, S. E., 144, 194 Brimhall, B., 202 Brinckmann, E., 4,46 Brink, R. A., 123, 125, 130, 131, 133, 141, 144, 148, 149, 181, 185, 194, 196, 198, 200, 203 Britten, R. J., 160, 195, 238, 260 Buc, J., 11, 16, 17, 18, 26, 28, 32, 40, 42, 43, 44, 45 Buchanan, B. B., 2, 3, 5 , 8, 10, 11, 14, 15, 16, 17, 18, 19, 20, 21, 25, 26, 30, 32, 40, 41, 42, 43, 44, 45, 46 Buck, W., 50, 94 BucMey, L. J., 82, 94 Bujak, J. P., 207, 208, 260 Bukhari, A. I., 171, 198 Burghardt, R. C., 220, 237, 238, 258, 263 Burgi, E., 254,264 Burke, M., 202 Burnell, J. N., 8, 21, 22, 40, 41 Bums, J. A., 141, 195 Burr, B., 127, 128, 139, 141, 159, 167, 195 Burr, F. A., 127, 128, 139, 141, 159, 167, 195 Busch, H., 241, 257, 258,263

C

Cachon, J., 211, 212, 214, 233, 252, 260, 261 Cachon, M., 214, 233,260

267

AUTHOR INDEX

Cachon-Enjumet, M., 21 1, 212, 252, 261 Calandra, F., 206,261 Callan, H. G., 259, 261 Calo, N., 36, 42 Calos, M. P., 153, 195 Campbell, A., 105, 173, 193, 195 Campbell, H. F., 68, 94 Canaani, E., 158, 195 Cao, R. Q . , 16,41 Capindale, J. B., 75, 98 Cardellina, J. H., 11, 78, 80, 94 Carlson, D. C., 10, 11, 15, 19, 21, 41, 46 Carlson, D. M., 50, 94 Carlson, R. D., 63, 85, 94, 100 Carlson, R. E., 85, 94 Carmichael, W . W . , 48, 52, 56, 57, 61, 64, 65, 66, 68, 69, 70, 78, 81, 83, 94, 95, 100 Carpenter, R., 120, 135, 139, 140, 142, 184, 185, 186, 192, 197 Carroll, C. R., 91, 94 Carter, N., 52, 94 Cates, R. G., 92, 98 Cavalier-Smith, T., 259,261 Center for Disease Control, Foodborne Disease Surveillance, 94 Cerff, R., 32,41 Chaleff, D., 127, 128, 152, 155, 158, 159, 162, 164, 167, 196 Chambon, P., 162, 194 Champigny, M. L., 5, 6, 7, 31, 39, 41 Chan, L. S., 61, 84, 98 Chandler, M., 171, 197 Chang, A., 258,263 Chanteau, S., 62, 87, 93 Chapman, K. S . R., 21, 41 Charles, S. A., 6, 26, 29, 36, 41 Chastain, C. J., 4, 41 Chehebar, C., 25,46 Cheng, D. S.-K., 197 Cheuk, C. E., 79, 98 Chew, F. S., 48, 91, 94 Chin, H. M., 5, 31, 40 Chretien, J. H., 88, 94 Christie, W., 110, 195 Chu, G . W., 78, 97 Chungue, E., 87, 94, I01 Clardy, J., 57, 79, 89, 90, 91, 98, 99, 100, 101 Clement-Metral, J. D., 4, 19, 41 Clutterbuck, A. J., 109, 195

Codd, G . A., 57, 64, 65, 66, 69, 81, 95, 100 Coe, E. H., Jr., 198 Cohen, S . A., 85, 95 Cole, A., 257, 261 Conde, M. F., 108, 198 Cooper, M. J . , 80, 95 Coppock, R. W., 50, 94 Corley, R. A., 50, 94 Cornelissen, P. T. J., 113, 135, 139, 142, 167, 184, 189, 194, 197 Cornu, A., 114, 139, 142, 189, 195, 196, 199 Cornwell, K. L., 16, 42 Correns, C., 112, 195 Cossar, J. D., 27, 41 Courage, U., 127, 128, 143, 152, 155, 156, 159, 162, 163, 166, 168, 170, 172, 195, 196, 197, 203 Coyne, D. P., 115, 195 Craig, J. C., 79, 98 Crawford, N. A., 5, 10, 11, 16, 17, 18, 19, 21, 32,41, 42, 43, 46 Crawford, R. M.,207, 261 Crowder, L. V . , 110,200 Cstke, C., 23, 30,45 Cuff, E. L., 158, 195 Cullimore, J . V . , 5, 41

D Dabell, P. E. R., 59, 100 Dale, B., 59, 95, I01 Dalzoppo, D., 19, 45 Daneholt, B., 210, 261 Darbre, A., 75, 98 Darlington, C. D., 210,261 Darwin, Ch., 109, 195 Davenport, J. W., 31,43 Davidson, E. H . , 160, 195 Davio, S. R., 86, 95 Dawson, G. P. W., 111, 195 Day, P . R., 210, 261 DeAmbrosis, W., 98 de Boer, R., 113, 114, 139, 142, 167, 189, I94 De Haan, H., 114, 195 de la Torre, A., 10, 11, 14, 16, 19, 21, 41, 43, 46 Dellaporta, S. L., 128, 200

268

AUTHOR INDEX

Delool, R. A. H., 112, 185, 195 Dembert, M. L., 88, 95 Demerec, M., 111, 135, 184, 195 Dempsey, E., 123, 124, 128, 129, 133, 137, 146, 182, 188, 189,201 Dennis, E. S., 155, 159, 161, 162, 163, 166, 170, 172, 173, 195, 200, 201 Derrner, H., 139, 195 Deumling, B., 152, 155, 156, 158, 160, 201, 202 D’Eustachio, P., 160, 195 de Vlaming, P., 114, 139, 140, 146, 195 Devlin, J. P., 68, 95 Devreux, M., 188, 201 deVries, H., 203 Dewar, H. A., 85, 97 De Winton, D., 116, 142, 195 Dickey, R. W., 63, 85, 86, 87, 94, 95, 97, 100 Dietrich, A. J. J., 113, 135, 200 DiFonzo, N., 126,200 Divan, C. L., 83, 100 Diwald, K., 234,261 Dodge, J. D., 207, 208, 210, 214, 218, 219, 220, 222, 230, 233, 236, 251, 258, 261, 262,263 Doerschug, E. B., 124, 148, 189, 195 Donker-Koopman, W., 113,197 Doodeman, M., 113, 114, 138, 139, 140, 146, 167, 195, 197 Dooner, H. K., 128, 130, 162, 167, 195, I96 Doring, H. P., 118, 120, 121, 122, 127, 128, 143, 152, 155, 156, 158, 159, 162, 163, 164, 168, 169, 170, 172, 177, 182, 195, 196, 197 Dougherty, E. C., 257,261 Dragesco, J., 206, 210, 211, 212, 251, 261 Droux, M., 15, 16, 41, 42 Dubos, R. J., 104, 196 Dufva, E., 88, 95 Dugdale, R. C., 60, 95 Duggan, J. X.,6, 35, 38, 39, 41 Dunsmuir, P., 158, 198 DuPraw, E. J., 210, 237, 261 DuPuy, J. L., 61, 95 Duraini, M. O., 2, 36, 40 DuRandt, W. C., 71, 96 Durrand, A. M., 206, 223, 224, 225, 227, 261 Dyson, R. D., 210, 261

E Echt, C. S., 129, 162, 167, 196, 202 Edstrom, J. E., 210, 261 Edwards, G. E., 5 , 22, 24, 28, 37, 44, 46 Edwards, 0. E., 68, 94, 95 Eickbush, T. H., 250,261 Eisenstadt, J. M., 210,263 Elder, J. W., 68, 94 Elleman, T. C., 75, 95 Ellis, R. J., 4, 43 El-Masry, H., 255, 264 Eloff, J. N., 65, 75, 76, 95 Ely, R., 50, 94 Emerson, R. A., 123, 124, 148, 196 Engels, J. M. M., 112, 135, 196, 202 Enriquez, M. B., 88, 97 Entz, G., 206, 261 Escaig, J., 216, 264 Euler, K. L., 69, 93 Evans, M. H., 82, 85, 95 Eyster, W. H., 116, 123, 196 F

Fabergt, A. C., 116, 142, 196 Faggiani, R., 32, 44 Falconer, I. R., 70, 71, 72, 73, 74, 75, 95, 96, 98, 99 Fallon, W. E., 82, 84, 99 Farabaugh, P., 167, 196 Farber, J. L., 71, 95 Farcy, E., 113, 114, 142, 196, 197 Farkas, G. L., 23, 30, 45 Fautz, E., 124, 165, 203 Federoff, N. V., 106, 127, 128, 129, 131, 133, 143, 149, 151, 152, 155, 158, 159, 160, 161, 162, 163, 164, 166, 167, 170, 172, 194, 195, 196, 202, 203 Feenstra, A., 158, 195 Fed, R. J., 163, 195, 202 Fermaglich, J., 88, 94 Ferstenberg, L. B., 246, 264 FertB, N., 28, 33, 42, 43 Fickenscher, K., 32,45 Fiffilan, S. A . , 206, 223, 224, 225, 226, 227, 228,261 Finch, J. T., 258,261 Fincham, J. R. S . , 106, 116, 120, 132, 133, 135, 138, 142, 149, 184, 196, 197, 210, 261

269

AUTHOR INDEX

Fine, K. E., 210, 261 Fink, G., 167, 196 Fitzgerald, G. P., 64, 95 Fjeld, A., 109, 194 Flavell, R. B., 160, 196 Fogel, S., 130, 196 Fohlmeister, J. F., 69, 93 Follmann, H., 16, 18, 40, 45 Forkmann, G., 202 Fowler, R. G., 106, 125, 142, 147, 196 Foxall, T. L., 57, 66, 69, 71, 73, 95, 96, 99 Foyer, C. H., 42 Fradkin, C. W., 123, 196 Fraenkel, G. S., 91, 95 Franker, C. K., 223, 224, 237, 240, 261 Freeling, M., 131, 152, 153, 155, 160, 163, 168, 189, 191, 192, 194, 196, 197, 200, 202 Friedemann, P. F., 124, 133, 136, 181, 189, 191, 197, 200, 201 Frommer, W.-B., 156, 168, 195 Fueling, M., 127, 163, 170, 182, 196 Fujiki, H., 80, 95, 96, 98, 99 Fukuyo, T., 63, 93 Fullrner, C. S., 31, 44 Furtek, D. B., 151, 152, 155, 196 Fusetani, N., 87, 94, 95 G Gabay-Laughnan, S., 108, 198 Gadal, P., 15, 16, 17, 18, 32, 41, 42, 43 Galas, D. J., 171, 197 Gallagher, T. F., 4, 43 Galleron, C., 206, 223, 224, 225, 227,261 Gaionnier, M., 87, 88, 97 Ganguli, P. M., 113,200 Garagusi, V. F., 88, 94 Gardemann, A., 35, 37, 41 Gamier, R., 6, 7, 8, 43 Gavalas, N. A., 5, 23, 24, 34, 43 Gavazzi, G., 185, 197 Gavrila, L., 209,261 Geiser, M., 127, 128, 152, 159, 162, 196, 197 Genenah, A. A., 62, 84, 95, 99 Gentile, J. H., 48, 66, 69, 95, 96, 99 Geraghty, D., 162, 199 Gerats, A. G. M., 113, 114, 128, 135, 139, 140, 142, 143, 146, 167, 184, 189, 194, 195, 197, 200

Gerlach, W. L., 152, 155, 158, 159, 161, 162, 163, 166, 170, 172, 173, 195, 200, 201, 202 Gerloff, G. C., 64, 95 Gerstel, D. U., 141, 195 Ghazarossian, V. E., 82, 95, 99 Gibbs, M., 5 , 6, 7, 8, 36, 38, 40, 42, 43 Gierl, A., 129, 152, 155, 156, 159, 162, 165, 202 Giesbrecht, P., 206, 207, 212, 216, 252, 261 Gilfillan, E. S., 61, 62, 95, 96 Givol, D., 158, 195 Gleason, F. K., 19,42, 46 Godeh, M. M., 23, 30,45 Goldberg, R. B., 152, 155, 156, 197, 203 Gonella, J. A., 130, 163, 181, 189, 192, 197 Gontero, B., 26, 28, 42, 43 Goodman, M. M., 108,203 Goodspeed, T. H., 188, 197 Gopichand, Y., 86, 100 Gorham, P. R., 50, 56, 64, 65, 66, 68, 69, 75, 94, 95, 96, 97 Gorman, M. B., 123, 127, 197 Goryshin, I. Yu.,144, 194 Grabow, W. 0. K., 71, 96 Grainger, R. M., 257,263 Granade, H. R., 87, 96 Grasse, P. P., 206, 210, 211, 212, 251, 252, 261 Greenblat, I. M., 123, 125, 148, 149, 150, 197 Grell, K. G., 206, 261 Griffith, R. G., 255,261 Grigor’yeva, L. V., 73, 96 Grindley, N., 144, 170, 171, 197 Groot, S. R. C., 113, 128, 135, 139, 142, 143, 184, 197 Guillard, R. R. L., 60, 96, 236, 237, 263 Guire, P. E., 63, 85, 94 Gupta, M., 155, 160, 197, 201 Gupta, V. K., 5 , 6, 7, 31, 40, 42

H Haapala, 0. K., 206, 210, 211, 214, 218, 220, 230, 233, 236, 250, 252, 261, 264 Hadfield, M., 79, 100 Hagemann, R., 121, 138, 146, 184, 185, 197, 199, 202 Hahlbrock, K., 124, 155, 165, 201, 203

270

AUTHOR INDEX

Hake, S., 127, 131, 163, 168, 170, 182, 189, 192, 196, 202 Hakii, H., 80, 95 Haldane, J . B . S., 116, 142, 195, 203 Hall, S., 57, 60, 61, 82, 84, 92, 96, 97 Hall, T . A., 244, 262 Haller, G., 225, 262 Halliwell, B., 6, 26, 29, 36, 42, 42 Halstead, B . W., 62, 87, 88, 96 Halvorson, H. O., 210,260 Hamkalo, B. A., 218, 220, 241, 248,262 Hammel, K. E., 16, 42 Hammer, U. T . , 64,65, 95 Hammond, S. J., 56, 75, 76, 77, 94, 99 Hannah, L. C., 124, 162, 163, 167, 197, 202, 203 Hanno, H. A., 88, 96 Hansen, M. J., 6, 42 Hansen, S. A., 61, 95 Harada, H., 80, 99 Harada, T., 52, 82, 96, 98 Harayama, S., 144, 197 Harrison, B. J . , 106, 116, 117, 120, 124, 132, 133, 135, 138, 139, 140, 142, 146, 149, 152, 155, 161, 165, 184, 185, 186, 192, 194, 196, 197, 201, 203 Harshey, R. M., 171, 198 Hartley, M. R., 4, 43 Hartman, F. C., 33,44 Harvey, P., 98 Hashimoto, Y.,78, 79, 86, 96, 101 Hatch, M . D., 5 , 6, 7, 8, 21, 22, 24, 25, 28, 35, 40, 41, 42, 43, 45 Hauman, J . H., 67, 96 Hawes, R. B., 74, 94 Hawthorne, D. C., 210, 262 Hayes, H. K., 123, 198 Heber, U., 26,43 Hedberg, M. F., 222, 224, 260 Heffron, F., 164, 198 Heidecker, G., 162, 199 Heldt, H. W., 6, 35, 36, 37,41, 42, 43, 46 Henning, D., 241, 257, 258,263 Hertig, C., 25, 42, 46 Hertwig, P., 117, 198 Herzog, M., 220, 237, 240, 250, 262 Heuer, B., 6, 42 Heywood, J., 240, 263 Himes, M., 209, 234, 240, 260, 262 Himmelfarb, J., 209, 234, 260

Hindin, E., 69, 101 Hinnebusch, A. G., 241, 257, 258, 262 Hirata, Y.,80, 99 Hixon, R. M . , 202 Hoffman, C . A., 91, 94 Hoffman, P. A., 87, 96 Hogervorst, J. M . W., 113, 135, 139, 142, 167, 184, 189, 194, 197 Hokama, Y.,86, 87, 88, 96, 97 Hollande, A., 211, 212, 252, 261 Holliday, R., 210, 262 Holligan, P. M., 60, 96 Holmgren, A., 4, 17, 19, 41, 42 Holm-Hansen, O., 210,262 Holmstedt, B., 88, 95 Holt, J. R., 209, 211, 234, 262, 263 Hondelmann, W., 110, 138, 142, 198 Hong, S. K . , 88, 99 Horgen, P. A., 258, 262 Horwitz, W., 85, 96 Hoschek, G., 152, 156, 197 Hoshino, H., 80, 95, 96 Houser, L. S., 85, 96 Hozier, J . , 257, 263 HSU,C. -P., 82, 83, 84, 96, 99 Hu, W. W. L., 108, 201, 203 Huber, C . S., 68, 96 Huber, S. C . , 27, 36, 42 Hughes, E. O., 64,96 Hughes, J . M . , 85, 96 Hunter, N. R., 68, 95 Hurst, J. W., 59, 61, 62, 95, 96, 101 Hutcheson, S . W., 10, 11, 15, 18, 25, 42 Hutchinson, J., 160, 196 Huttermann, A., 5 , 4 5 Huynh, V. L., 75, 95 I

Ichihara, N., 62, 99 Iglesias, A. A , , 33, 35, 42 Iida, S., 164, 198 Iino, T., 144, 197 Ikawa, M., 57, 66, 69, 82, 93, 94, 96, 99 Ikeno, S., 115, 116, 198 Imai, Y.,110, 113, 114, 115, 135, 184, 198, 200 Incze, L., 59, 60, 101 Ingham, H. R., 85, 97

AUTHOR INDEX

Inglis, A., 163, 195 Ip, S. -M., 19, 42 Iwaoka, W. T., 86, I00 J

Jablonski, P., 9, 10, 12, 40 Jackim, E., 69, 96 Jackson, A. R. B., 70, 72, 75, 95, 96 Jacquot, J., 15, 16, 41 Jacquot, J. -P., 3, 10, I I , 16, 17, 18, 20, 24, 31, 32, 42, 43, 45 James-Kracke, M. R., 84, 96 Janzen, D. H., 61, 91, 96 Jeffries, V. E., 111, 120, 184, 189, 198, 202 Jenkins, G. I., 4 , 4 3 Johns, M. A., 127, 163, 170, 182, 189, 191, 196, 200 Johns, R. B., 208,262 Johnson, H. S., 5 , 4 3 Johnson, T. C., 16, 19, 41 Jolley, J. W., 206, 263 Jones, C. L. A., 52, 61, 66, 70, 72, 81, 83, 94, 96 Jones, G. J., 208,262 Jones, L., 126, 131, 200 Jorgensen, K. F., 212, 262 Jornvall, H., 19, 42 Juhasz, A., 23, 30, 45 K Kachru, R. B., 6, 43 Kagawa, T., 4, 28, 43, 44 Kalberer, P. P., 14, 20, 32, 41 Kallen, R. G., 240,262 Kamiya, H., 78, 79, 83, 96 Kanna, B., 115, 198 Kao, C. Y., 84, 85, 96 Kao, P. N., 84. 96 Karabourniotis, G., 5 , 24, 34, 43 Karentz, D., 223, 224, 262 Kashiwagi, M., 78, 80, 97, 98 Kato, Y., 78, 79, 97 Kearns, L. P., 207, 212, 216, 236, 242, 243, 244, 245, 246, 248, 254, 255, 257, 262, 264 Kedharnath, S., 123, 198 Kellenberger, E., 225, 256, 258, 262, 263

27 1

Keller, P. B., 128, 200 Kemble, R. J., 32, 46, 108, 198 Kermicle, J. L., 130, 185, 195, 198, 203 Ketchum, S. R., 31, 43 Kihara, H., 110, 112, 198 Kikuchi, H., 86, 100 Kim, B. D., 108, 198 Kim, V. S., 68, 97 Kim, W. K., 64,65, 96 Kim, Y. S., 210, 262 Kimura, L. H., 86, 87, 88, 96, 97 Kirk, J. T. O., 107, 108, 110, 111, 113, 115, 116, 141, 198 Kirpenko, N. I., 73, 78, 97 Kirpenko, Yu, A., 73, 78, 96, 97 Kleckner, N., 153, 164, 198 Kleczkowski, L. A., 5 , 4 3 Klein, A. S., 128, 162, 163, 198 Klosgen, R. B., 129, 152, 155, 156, 159, 162, 165, 202 Klotz, L. C., 207, 210, 222, 224, 238, 240, 241, 252, 257, 258, 260, 262 Klug, A., 248, 258, 261, 264 Kobayashi, M., 62, 99 Kobayashi, Y., 26, 28, 43, 45 Koehn, F. E., 82, 97 Koghn, F., 84, 96 Kohler-Wieder, R., 206, 230, 262 Kohne, D. E., 238,260 Koller, Th., 248, 264 Kolodny, G. M., 254, 262 Kolt, R. J., 68, 94 Koomen, W., 113, 114, 138, 167, 195 Kornberg, R. D., 258, 262 Kosaki, T., 88, 97 Kotaki, Y., 52, 83, 98, I01 Kow, Y.W., 5 , 4 3 Koyanagi, L. M., 86, 98 Kozakai, H., 68, 97 Kramer, E., 28, 45 Kreuzaler, F., 124, 155, 165,201, 203 Kridl, J., 162, 199 Kruger, H., 56, 75, 77, 94, 99 Ku, M. S. B., 37, 46 Kubai, D. F., 206, 210, 230, 232, 233, 262 Kuckuck, H., 117, 118, 119, 120, 191, 198 Kung, J . E., 16, 41 Kunisawa, R., 64,76, 100 Kunze, R., 127, 156, 163, 168, 170, 182, 195, 196

272

AUTHOR INDEX

Kutscher, L., 118, 198 Kycia, H., 65, 75, 76, 95 Kycia, J. H., 70, 72, 100

L Ladha, J. K., 5 , 44 Laing, W., 36, 42 Laing, W. A., 6, 43 Laird, A., 129, 152, 156, 161, 168, 194, 195 Lambert, C., 4,40 Lambert, W., 92, 97 Lamdan, C. A., 223, 224, 237, 261 Langley, J., 70, 95 Lanzov, V. A., 144, 194 Lara, C., 10, 11, 14, 15, 16, 19, 21, 41, 43, 46 Larson, R. L., 198 Latzko, E., 6, 7, 8, 37,43, 46 Laughnan, J. R., 108, 198 Lawrence, D. N., 88, 97 Lawrence, T., 11 1, 198 Lawrence, W. J. C., I I I , 198 Leadbeater, B., 207, 233, 236, 262 Lederberg, E., 105, 193, 195 Lee, B. W., 63, 85, 94 Lee, J. S., 62, 98 Leedale, G. F., 210,262 Leegood, R. C., 5 , 6, 7, 24, 26, 28, 36,43 Legrand, A. M., 87, 88, 97 Lendzian, K. J., 6, 23, 43 Leuders, K., 158, 195 Leutenegger, S., 208, 262 Lever, M. L., 63, 85, 94 Levin, D. A., 91, 97 Levings, C. S., 111, 108, 198, 201, 203 Levis, R., 158, 198 Lewin, P., 88, 98 Lewis, R., 98 Li, K., 88, 97 Lierop, P., 113, 139, 142, 197 Lightner, D. V., 79, 97 Lilienfeld, F. A., 112, 198 Lim, T. C., 31,39 Lin, C. C., 212, 262 Link, C., 209, 234, 260 Linnert, G., 119, 184, 198, 199 Liston, J., 86, 100 Liu, D., 241, 257, 258, 263

Liu, M. -H., 241, 257, 258,263 Livolant, F., 211, 214, 250, 251, 253, 262 Llewellyn, D., 159, 161, 162, 163, 195, 200 Loeblich, A. R., 59, 99, 206, 207, 208, 210, 222, 223, 224, 238, 240, 241, 252, 257, 258, 260, 261, 262, 263, 264 Loison, G., 88, 95 Lonsdale, D. M., 202 Loper, C. L., 211, 262 Lopez-Gorge, J., 19, 44 Lonmer, G. H., 4, 36,42, 44 Lorz, H., 159, 161, 162, 189, 199, 200 Losada, M., 3,43 Loussan, E., 62, 87, 93 Louw, P. G. J., 75, 97 Lukina, L. F., 78, 97 Lumish, R. M., 88, 97 Lynch, J. M., 82, 84, 99

M MacArthur, R. H., 259, 262 McCarty, R. E., 31,43, 44 McClintock, B., 105, 106, 123, 124, 125, 126, 127, 130, 135, 137, 138, 139, 141, 142, 143, 145, 146, 147, 148, 163, 164, 170, 176, 181, 182, 183, 188, 189, 191, 192, 193, 199 McCollum, J. P. K., 85, 97 Maceo, A., 88, 97 McInnes, A., 72, 96 McKay, R., 98 McKey, D., 91, 92, 97 McKinley, K. R., 59, 99 McKinney, H. H., 189, 191,202 McLachlan, J., 64, 65, 95, 97 McLeish, J., 210, 263 McMillan, J. P., 87, 96 McWhirter, K. S.,116, 199 Maeda, K., 10, 11, 17, 18, 19, 20, 45 Mague, F. C., 60, 101 Mahmood, N. A., 48, 52, 56, 57, 61, 66, 69, 70,81, 83, 94 Maizonnier, D., 114, 199 Makao, T., 77, 100 Malkin, R., 11, 16, 21, 41, 43 Maloney, T. E., 66, 95 Mandel, M., 64,76, 100 Manetas, Y.,4, 5 , 6, 23, 24, 34, 43 Mans, R. J., 108, 162, 197, 198

273

AUTHOR INDEX

Maranda, L., 61, 100 Marchand, A,, 83, 84, 96 Marmur, J., 240,262 Marner, F. J., 78, 80, 94 Marotta, R., 126, 200, 201 Marques, I. A., 34, 35, 36, 43 Martin, D. F., 210,262 Mather, K., 120, I99 Matsumoto, G. K., 79, 98 Matsumoto, K., 57, 82, 84, 89, 90, 91, 99, 101

Matthys-Rochon, E., 225, 230, 233,263 Mauvais, J., 127, 128, 152, 155, 158, 164, 167, 196 Maze, J. R., 48, 99 Mechelke, F., 118, 199 Meier, P. G., 98 Meirnanis, S., 26, 43 Melandri, B. A., 35, 43 Mende, T. J., 57, 89, 90,93 Mendiola, L. R., 236, 237, 263 Merckelbach, A., 127, 156, 163, 168, 170, 182, 195, 196 Merson, M. H., 85, 96 Meselson, M., 167,200, 240, 263 Messing, J., 129, 155, 158, 162, 164, 166, 167, 172, 199, 201 Meunier, J. C., 11, 16, 17, 18, 26, 28, 32, 36, 40, 42, 43, 44, 45 Meyer, J., 164, 198 Meyer, K. F., 82, 97 Meyer, M., 121, 146, 184, 199 Michaelis, P., 111, 199 Miginiac-Maslow, M., 15, 16, 20, 31, 42 Mikula, B., 130, 133, 141, 194 Miller, D. M., 86, 87, 95, 97 Miller, J. H., 153, 193, 195, 200 Mills, J. D., 7, 37, 44 Mitchell, P., 37, 44 Mitra, S. K., 113, 200 Miura, I., 82, 84, 99 Miwa, M., 80, 96 Miyachi, S., 5 , 44 Miyahara, J. T., 87, 89, 97 Miyake, K., 110, 115, 200 Miyazawa, B., ll5.200 Miziorko, H. M., 4, 44 Modolell, J., 167, 200 Mohamed, A. H., 2, 3, 8, 9, 12, 26, 27, 30, 36, 40, 44

Moikehai, S. N., 78, 97 Mold, J. D., 82, 99 Monahan, B., 31,45 Montanelli, C., 126, 200 Morre, R. E.,48, 56, 66, 78, 79, 80, 95, 97, 98 Morel, F. M., 206, 260 Morey-Gaines, G., 52, 55, 63, 94, 98, I00 Mod. M., 80, 95, 98, 99 Moroney, J. V., 31, 44 Moms, J. G., Jr., 88, 98 Mortimer, R. K., 210,262 Mottinger, J. P., 128, 189, 191,200 Motto, M., 126,200 Moudnanakis, E. N., 250, 261 Mulder, R. J. P., 113, 135, 200 Miiller, B., 23, 44 Muller, H., 82, 98 Miiller-Neumann, M., 129, 152, 156, 161, 168, 194, 195 Murakami, Y.,63, 98, 101 Murphy, D. J., 26, 37, 46 Murphy, E. B., 206,263 Murray, R. G., 255, 264 Murthy, J. R., 75, 98 Muschinek, G., 38,44 Muto, S. , 5 , 44 M v t Akamba, L., 7 , 4 4 Mynderse, J. S., 78, 79, 80, 97, 98

N Nakajima, I., 63, 98, 101 Nakamoto, H., 11, 22, 24, 28, 44 Nakamura, Y.,4, 44 Nakanishi, K., 82, 84, 99 Nakayasu, M., 80, 95, 98, 99 Nalin, C . M., 31, 44 Nechay, B. R., 87, 94 Nehls, P., 257, 263 Nehrlich, S. C., 23, 31, 39 Neilson, A . , 74, 94 Nelson, 0. E., 128, 129, 151, 152, 155, 162, 163, 167, 196, 197, 198, 200 Nerad, T. A., 98 Neuffer, M. G., 123, 124, 126, 132, 181, 189,200 Neve, R. A., 82, 96 Nevers, P., 133, 139, 200

274

AUTHOR INDEX

Nichols, P. D., 208, 262 Nien-Tai, H., 162, 199 Nilan, R. A., 123, 148, 181, 194 Nishiyama, A., 85, 96 Nishizawa, A. N., 5 , 17, 18, 25, 26, 30, 32, 41, 43, 44, 46 Nitta, I., 80, 99 Nooden, L. D., 207, 210, 237, 241,263 Norton, T. R., 78, 80, 97, 98 Novick, R., 105, 193, 195 Nowick, E. M., 124, 148, 200 Nozawa, K., 78, 79, 96 Nukina, M., 86, 88, 96, 98

0

Oakley, B. R., 214, 218, 219, 230, 233, 251, 258, 263 O’Dell, M., 160, 196 Ogren, W. I., 4, 44 Ogren, W. L., 4, 41 Oguchi, T., 144,197 O’Hare, K., 162, 194 Ohizumi, Y.,87, 100 Ohshika, H., 89, 98 Oishi, S., 77, 100 Omnaas, J., 33, 44 Orakwue, F. C., 110,200 Orcutt, J. D., 92, 98 Orlovskiy, V. M., 73, 78, 96, 97 Orrenius, S., 71, I00 Orton, E. R., 123, 131, 148,200 Oshima, Y.,52, 57, 63, 68, 82, 83, 89, 90, 91, 96, 97, 98, 99, 101 Ostensvik, O., 70, 72, 98 Osterman, J. C., 123, 162, 182, 200 Oud, J. L., 113, 135, 200 Oyama, M., 87, 97

P Padilla, G. M., 68, 97 Palmer, R. G., 111, 202 Parenti, F., 210, 263 Parke, M., 207, 263 Parlavecchio, R., 123, 188, 194 Peacock, J. W., 152, 155, 158, 159, 161,

162, 163, 166, 170, 172, 173, 195, 200, 201, 202 Pearn, J. H., 88, 95, 98 Pearson, 0. H., 139, 200 Pearson, R. C. M., 85, 97 Peek, C. A., 66, 98 Pelling, C., 210, 260 Perlmuter, M. E., 25, 46 Peters, T., 248, 263 Peterson, M. A., 68, 94 Peterson, P. A., 106, 123, 124, 125, 127, 128, 129, 130, 131, 133, 136, 139, 142, 143, 144, 146, 147, 148, 155, 156, 159, 161, 162, 163, 165, 181, 183, 189, 191, 192, 196, 197, 200, 201. 202, 203 Pettijohn, D. E., 255,263 Pezzanite, J. O., 82, 99 Pfeister, L. A., 209, 211, 216, 222, 234, 262, 263, 264 Phinney, H. K., 66, 76, 98, 100 Phus, E., 246, 264 Pickering, R. P., 86, 95 Pickett-Heaps, J. D., 233, 264 Pike, R. K., 68, 95 Pitout, M. J., 75, 100 Pla, A., 19, 44 Pohlman, R. F., 129, 155, 158, 164, 166, 167, 172, 201 Poli, M. A., 57, 89, 90, 93 Pollard, M., 104, 201 Pompe, A. J., 113, 114, 139, 142, 167, 189, 194 Porter, K. G., 92, 98 Porter, M. A., 33,44 Portis, A. R., 4, 38, 40, 44 Pradel, J., 11, 16, 17, 28, 32, 36,43, 44, 45 Preston, J. F., 210, 263 Price, C. A., 236, 237,263 Price, R. J., 62, 98 Prichard, C. D., 223, 224, 237, 261 Pring, D. R., 108, 198, 201 Procter, N. H., 61, 84, 98 Proudfoot, N. J., 162, 201 Provasoli, L., 63, 98 Prozesky, 0. W., 71, 96 Pruitt, S. C., 257, 263 Pryor, A. J., 159, 161, 162, 163, 195, 200 Puiseux-Dao, S., 206, 214, 250, 253, 260 Punnett, R. C., 112, 135, 184, 201 Pupillo, P., 32, 35, 43, 44

AUTHOR INDEX

R

275

Rowell, P., 5 , 19, 27, 41, 42, 44 Rubenstein, I., 162, 199 Rubin, G. M., 158, 181, 198, 201 Rudd, T. P., 22,44 Ruddle, F. H., 160, 195 Rukuyo, Y., 63, 101 Runnegar, M. T., 70, 71, 72, 73, 74, 75, 95, 96, 98, 99 Ryter, A., 256, 258, 262, 263 Ryther, J. H., 60, 96

Rabin, R., 75, 98 Rae, P. M., 207, 220, 240, 258, 263 Ragelis, E. P., 52, 57, 62, 98 Ragg, H., 124, 165,203 Randall, D. D., 5 , 43 Randall, J., 62, 86, 98, 101 Ransom, R. E., 92, 98 Rao, I. M., 5 , 6, 37, 38, 44 Rapoport, H., 68, 82, 84, 93, 98, 99 Rattner, J. B., 218, 220, 241, 248, 262 Ravazzini, R. A., 5 , 46 S Rayner, M. D., 88, 89, 98, 99 Raziuddin, S., 82, 98 Saccardo, F., 188,201 Rechavi, G., 158, 195 Sachs, M. M., 155, 159, 161, 162, 163, 166, Reddy, A. R., 124, 125, 127, 137, 142, 143, 170, 172, 173, 195, 200, 201 183,201 Saedler, H., 124, 129, 133, 139, 152, 155, Reddy, L. V., 135,201 156, 158, 159, 160, 161, 162, 165, 166, Reddy, R., 241, 257, 258,263 168, 172, 177, 185, 186, 194, 200, 201, Reichardt, P. B., 57, 82, 96, 97 202, 203 Reif, H. J., 155, 168, 201 Sager, R., 129,201 Renz, M., 257, 263 Sakabe, N., 80, 99 Rhoades, D. F., 48, 92, 98 Sakai, H., 80, 100 Rhoades, M. M., 123, 124, 128, 129, 133, Sakamoto, H., 80, 99 137, 142, 146, 181, 182, 184, 188, 189, Sakhrani, L. A., 223, 224, 237, 261 192, 201 Salamini, F., 123, 126, 127, 163, 170, 182, Rhodes, P. R., 155, 156,203 188, 194, 196, 200, 201 Ricard, J., 11, 16, 17, 26, 28, 32, 33, 36, Salvucci, M. E., 4, 44 41, 43, 44, 45 Sampaio, M. J. A., 5,44 Riegel, B., 82, 99 Sand, S. A., 113, 135, 138, 142, 184, 189, Riegel, J., 156, 168, 195 191, 202 Riel, P., 82, 99 Santikam, S., 56, 75, 76, 77, 94, 99 Ris, H., 206, 207, 210, 230, 232, 233, 236, Sajeant, W. H., 206,263 262, 263 Sarma, Y. S., 209, 211, 263 Riviere, M., 17, 18, 40 Sasner, J. J., Jr., 57, 66, 69, 71, 73, 82, 93, Rizzo, P. J., 207, 210, 220, 237, 238, 241, 94, 95, 96, 99 258,263 Sastry, G. R. K., 106, 111, 120, 133, 149, Roberts, B. S., 206, 263 184, 189, 196, 198, 202 Roberts, T. M., 207, 210, 222, 238, 240, Satyanarayana, K. V., 130, 203 252, 258,260, 263 Sawyer, P. J., 69, 93, 99 Robertson, D. S., 131, 181, 189, 192, 201 Scagel, R. F., 48, 99 Robinson, S. P., 36, 44 Schantz, E. J., 82, 83, 84, 85, 94, 95, 99, Rodman, J. E., 48, 91, 94 loo Roeder, G., 167, 196 Scheibe, R., 2, 3, 8, 17, 20, 22, 24, 26, 27, Rogers, R. S., 84, 98 28, 29, 30, 32, 33, 36, 40, 44, 45 Rosa, L., 25, 44 Scheitler, B., 4, 46 Roughan, P. G., 4,44 Scherz, W., 118, 202 Rouiller, C., 225, 262 Scheuer, P. J., 78, 79, 86, 88, 96, 97, 98, Rouse, G. E., 48, 99 99, 100

276

AUTHOR INDEX

Schick,R., 117, 118, 119, 198, 202 Schmidt, R. J . , 59, 99 Schmidt-Clausen, H. J., 5, 18, 23, 28, 45, 46 Schmitz, F. J., 86, 100 Schneider, R., 88,98 Schnoes, H. K., 82, 84, 94, 95, 96, 97, 99, 100 Schoenholz, P., 82, 97 Schofield, W. B., 48, 99 Schopf, J. W., 258,263 Schram, A. W., 113, 114, 128, 135, 139, 140, 142, 143, 146, 184, 195, 197, 202 Schriek, U.,23, 45 Schuett, W., 82, 99 Schultz, J. C., 91, 93, 99 Schwartz, D., 123, 129, 152, 158, 159, 161, 162, 166, 167, 182, 196, 200, 202 Schwartz, L. D., 48, 94 Schwarz-Sommer, Zs., 124, 129, 152, 155, 156, 158, 159, 160, 162, 165, 201, 202, 203 Schwenn, J., 23, 45 Schiirmann, P., 2, 10, 11, 14, IS, 17, 18, 19, 20, 26, 28, 30, 31, 32, 34, 41, 42, 45 Scott, W. E., 71, 75, 96, 100 Scowcroft, W. R., 189, 199 Sears, B. B., 108,202 Sechaud, J . , 256,262 Seeman, J. R., 4, 45 Seliger, H. H., 59, 99 Selinioti, E., 5, 43 Setliff, J. A., 88, 99 Shapiro, H. S., 240,263 Shapiro, J. A., 144, 158, 170, 171, 202 Shavel, J . , 82, 99 Sheldon, E., 163,202 Shepherd, N., 124, 152, 155, 156, 158, 160, 165, 197,201, 202, 203 Sheridan, M. A., 111,202 Sherratt, D., 144, 170, 171, 197 Shibata, S., 89, 99 Shilo, M . , 48, 56, 58, 59, 67, 68, 99, 100 Shimizu, Y.,62, 69, 82, 83, 84, 86, 88, 89, 93, 95, 96, 99 Shure, M., 127, 129, 158, 159, 160, 162, 163, 167, 196, 202 Shyam, R., 209, 211,263 Siegelman, H. W., 65, 70, 75, 76, 77, 82, 95, 98, 100

Sigee, D. C., 206, 207, 209, 211, 212, 216, 220, 221, 222, 223, 224, 225, 226, 227, 228, 231, 236, 238, 239, 242, 243, 244, 245, 246, 247, 248, 249, 254, 255, 256, 257, 259,260, 261, 262, 263, 264 Silva, E. S., 246, 264 Silver, J . , 70, 71, 99 Silver, J. C., 258, 262 Silverberg, B. A., 246, 264 Simon, M., 240,262 Simm, J . , 50, 94 Sims, G. G., 83, 84, 96 Sirenko, L. A., 73, 97 Skaar, H., 246,264 Skoczylas, O., 230,264 Skoog, F., 64,95 Skulberg, 0. M . , 57, 65, 66, 69, 72, 98, 100 Skvarla, J. J . , 234, 263 Slack, C. R., 5 , 42 Slatkin, D. N., 65, 70, 72, 77, 100 Slovacek, R., 31, 34, 45 Smith, C. W., 88, 98 Smith, H. H., 113, 189, 202 Smith, J. D., 202 Smith, M. T., 71, 100 Smith, R. J., 56, 75, 76, 77, 94, 99 Smyth, D. A., 5 , 43 Snell, T. W., 92, 100 Soave, C., 126,200 Soh, N. E., 98 Solomon, A. E., 79, 100 Sommer, H., 82, 97, 99, 124, 152, 155, 156, 158, 159, 160, 161, 165, 166, 172, 177, 185, 186, 192, 194, 201, 202, 203 Soran, V., 209, 261 Soulik, J.-M., 11, 16, 17, 28, 32, 36, 43, 44, 45 Soyer, M. O., 206, 210, 211, 214, 216, 218, 220, 228, 230, 232, 233, 234, 236, 237, 240, 250, 252, 253, 254,260, 261, 262, 264 Spaulding, B. C., 85, 100 Spector, D., 214, 216, 222, 223, 224, 225, 228, 230, 233, 234, 241, 251, 254, 257, 258, 263, 264 Spirchez, C. T., 209, 261 Spitters, C. J. T., 112, 135, 196, 202 Spivak, C. E., 68, 100 Spradling, A. C., 181, 201, 202

277

AUTHOR INDEX

Sprague, G. F., 189, 191,202 Spribille, R., 202 Springer, J. P., 82, 99 Srivastava, D. K., 31, 32, 33, 45 Stahl, E., 47, 100 Stallman, N. D., 74, 94 Stanier, R. Y.,64,76, 100 Stankevich, V. V., 73, 96, 97 Starlinger, P., 105, 127, 128, 131 , 143, 152, 155, 156, 158, 159, 162, 163, 164, 166, 168, 170, 172, 177, 182, 193, 194, 195, 196, 197, 203 Stavric, B., 68, 95 Steele, R. E., 240, 258, 263 Steidinger, K., 89, 99 Steidinger, K. A., 206, 211, 234, 262, 263, 264 Steiger, E., 2, 5 , 45 Stein, J. R., 48, 99 Steinmuller, K., 4, 45 Stem, D. B., 202 Stewart, J. M., 237,264 Stewart, W. D. P., 5 , 19, 27, 41, 42, 44 Stickland, R. G., 197 Stitt, M., 6, 35, 36, 37, 41, 42, 43, 46 Stoner, R. H., 65, 70, 77, 100 Stokes, P. M., 246, 264 Stranger, D. W., 82, 99 Straub, J., 189, 191,202 Strommer, J. N., 131, 163, 168, 189, 192, 202 Strong, F. M., 82, 95, 99 Stroughton, R. B., 79, 100 Stubbe, H., 117, 118, 119, 120, 121, 122, 138, 185, 199, 202 Suave, P., 33, 42 Suganuma, M., 80, 95 Sugimura, T., 80, 95, 96. 98, 99 Sugiyama, T., 8, 11, 44, 45 Sullivan, J. J., 86, 100 Sunderland, N . , 210, 263 Suske, G., 16,45 Sutton, C. W., 19, 41 Sutton, W. D., 152, 158, 159, 161, 162, 166, 200, 202 Suzuki, A., 15, 16, 41 Swanson, J., 152, 155, 163, 189, 194 Switzer-Dunlap, M., 79, 100 Szekerczes, J., 89, 98 Szybalski, W., 105, 193, 195

T Tachibana, K., 86, 89, 98, 99, 100 Tacina, F., 209, 261 Tajiri, M., 83, 101 Takahashi, M., 87, 100 Takahashi, W., 86, 99 Takashima, M., 80, 100 Takayama, S., 80, 95 Tamm, S. E., 144, 194 Tanaka, T., 115, 203 Tangen, K., 206,264 Taylor, F. J., 207, 264 Taylor, F. J. R., 59, 89, 100 Taylor, W. C., 131, 152, 155, 163, 168, 189, 192, 194, 202 Terada, M., 80, 95 Terasawa, Y.,110, 203 Theiss, W. C., 52, 61, 66, 70, 72, 78, 81, 83, 100 Thevenin, S., 62, 87, 93 Thoma, F., 248, 264 Thomas, D. A., 4,46 Thor, H., 71, 100 Tillmann, E., 127, 128, 152, 155, 156, 158, 159, 162, 164, 168, 195, 196, 197 Tilney-Bassett, R. A. E., 107, 108, 110, 111, 112, 113, 115, 116, 141, 185, 195, 198 Timothy, D. H., 108,201, 203 Timpano, P., 234, 263 Tindall, D. R., 63, 86, 87, 94, 95, 97, I00 Tippit, D. H., 233, 264 Tischner, R., 5 , 45 Toerien, D. F., 75, 100 Tomiie, Y.,80, 99 Tornabene, T. G., 82, 98 ToxopCus, H. J., 203 Treichel, S., 4, 46 Trevor, A. J., 61, 84, 98 Triemer, R. E., 208, 214, 216, 222, 223, 224, 225, 228, 230, 233, 234, 251, 254, 257,264 Tsai, C. Y.,203 Tsugita, A., 10, 11, 17, 18, 19, 20, 45 Tsukitani, Y.,86, 100 Tsutsumi, J., 86, 99 Tuinman, A. A., 56, 75, 77, 94, 99 Turpin, D. H., 59, 100 Tuschall, D. M., 124, 162, 167, 197, 203

278

AUTHOR INDEX

Tuttle, R. C., 210, 222, 224, 240, 252, 260, 263, 264 Twarog, B. M., 62, 85, 100 Tyler, M. A., 59, 99

U Ubben, D., 155,201 Udvardy, J., 23, 30, 45 Ulitzur, S., 67, 68, 100 Usuda, H., 5 , 37, 44, 46

V Vallejos, R. H., 5 , 46 van der Laan, J., 113, 139, 142, 197 van de Sande, J. H., 212,262 Van Engen, D., 86, 100 van Goor, A. C., 206,264 Van Kester, W. N. M., 112, 135, 196, 202, 203 Van Schaik, N. W., 123, 148, 203 Vasconcelos, A. C., 214, 216, 222, 223, 224, 228, 251, 264 Vaughn, K. C., 111,203 Vaughn, S., 31, 34,45 Vayvada, G., 82, 84, 99 Vidal, J., 17, 42 Viljoen, C. C., 56, 75, 77, 94, 99 Vodkin, L. O., 152, 155, 156, 197, 203 Vogt, K., 18,40 Volk, S . L., 76, 100 von Stosch, H. A., 234, 240,264 Von Willert, D. T., 4, 46 Vosselman, L., 112, 135, 196, 202 Vu, J. C. V., 4, 46

W Wagner, W., 16, 45 Walker, D. A., 5 , 6, 7, 24, 26, 28, 36, 43, 44, 46 Walker, L. M., 211,262 Walker, S. E., 84, 85, 96 Wall, D., 59, 93

Wallroth, M., 113, 142, 197 Wara-Aswapati, O., 32, 46 Warncke, K., 31,43 Watanabe, M. F., 77, 100 Watson, W. H., 69, 99 Weber, C. R., 1 1 1 , 139,200 Weck, E., 127, 128, 143, 152, 155, 156, 158, 159, 162, 163, 166, 168, 170, 172, 195, 196, 197, 203 Wegener, K., 66, 69, 96 Weinstein, I. B., 80, 99 Weissinger, A. K., 108,203 Wekell, M. M., 86, 100 Werr, W., 127, 128, 152, 156, 159, 162, 168, 195, 196, 197 Wessels, P. L., 56, 75, 77, 94, 99 Wessler, S . , 127, 129, 158, 159, 160, 162, 163, 167, 196, 202 Weyers, W. H., 130, 194, 203 Whatley, F. R., 25,44 White, A. W., 61, 100, 206, 264 Whitefield, J. F., 255, 264 Whittaker, M. M., 19, 42, 46 Wichmann, C. F., 82, 97 Wichmann, C. R., 83, 100 Wienand, U., 124, 129, 152, 155, 156, 158, 159, 160, 162, 165, 191,201, 202, 203 Wildner, G. F., 7, 46 Williams, D. H., 56, 75, 76, 77, 94, 99, I01 Williams, E., 130, 131, 144, 194, 203 Williams, G. L., 207, 208, 260 Williams, J., 206, 263 Williams, W. M., 130, 203 Wilson, E. O., 259,262 Wilson, K. G., 111, 203 Winter, K., 23, 34, 35, 46 Wirtz, W., 36, 42, 46 Witkop, B., 68, 100 Wollfarth-Bottermann, K. E., 206, 261 Wolosiuk, R. A., 2, 5 , 10, 11, 14, 16, 17, 18, 25, 26, 30, 32, 34, 41, 42, 44, 45, 46 Wood, D. R., 123, 181,203 Wood, P. C., 85, 97 Woodard, R. W., 79, 98 Woodrow, I. E., 5 , 26, 37,46 Wong, S. H., 69, 101 Worcel, A., 254, 264 Wyler, R. S., 82, 99 Wylie, A. P., 210, 261 Wynne, M. J., 207,260

279

AUTHOR INDEX

Y Yarnada, M . , 4, 44 Yarnaguchi, H . , 62. 85, 100 Yamazato, K . , 78, 79, 96 Yasumoto, T., 52, 57, 63, 68, 83, 86, 87, 88, 89, 90, 91, 93, 95, 96, 97, 98, 100, 101

Yee, B. C., 10, 11, 15, 17, 18, 19, 21, 32, 41, 43, 46 Yentsch, C . M . , 59, 60, 61, 62, 95, 96, IOI Yoder, J., 129, 152, 156, 161, 168, 194, 195 Yokochi, L., 86, 88, 96 Yoshida, T., 86, 99

Yoshioka, M., 62, 99 Yuan, R., 240, 263

Z

Zederbauer, E., 234, 264 Zehnder, A., 64, 96 Zetsche, K . , 4, 45 Zeven, A., 112, 135, 196, 202 Ziegler, H . , 2, 5, 23, 38, 43, 44, 45, 46 Ziegler, I., 2, 5, 6, 23, 38, 44, 45, 46 Zilbois, F., 19, 45 Zuber, M. S . , 126, 131,200

This Page Intentionally Left Blank

SUBJECT INDEX

A Algal toxins, 47- 101 characterization, 67-91 Chrysophyta, 48, 52-59 Cyanophyta, 48, 64-67 dinoflagellate ciguatera toxins, 86-89 dinoflagellate paralytic shellfish poisons (PSP), 82-86 dinoflagellate toxins, 61, 86-91 environmental role of, 91-93 freshwater cyanophyte toxins, 68-78 growth, 52-67 isolation, 67-91 marine cyanophyte toxins, 78-82 naming, 52 occurrence, 52-67 prymnesium toxins, 67-68 Pyrrophyta, 48, 59-64 toxicity, 52-67 toxinology, 67-91 Amanita phalloides, 72 A mphidium , 209 A . carterae, 216, 222, 223, 236 Anabaena, 15, 16, 18-19, 48, 68 A. cylindrica, 19 A . flos-aquae, 65, 68-69 Anacystis nidulans, 23 Anatoxins, 68 Anthopleura, 240 Antimycin, 7 Antirrhinum majus, 138, 142, 156, 185-188 Aphanizornenon, 48 Aph. flos-aquae, 65-66 Aphantoxins, 69 Apodinium S D D . . 231

B Biotoxins, 48 Blastodiniurn, 232 Bromoaplysiatoxin, 80

C

Callistephus chinensis, 138, 142 Candida, 73 Ceratium spp., 230 Chaetomorpha h u m , 63 Chironomus, 209 Ciguatera, 62-63, 86-89 Ciguatoxin (CTX), 86-89 Cloning, I51 153 Corynebacterium nephridii, 19 Crypthecodinium cohnii, 209, 233, 234, 237, 238 CTX, see Ciguatoxin Cucurbita moschata, 91

-

D Debromoaplysiatoxin, 78-79, 80 Delphinium ajacis, 184 Diarrhetic shellfish poisoning (DSP), 89-91 Dictyota dichotoma, 63 Dinoflagellate chromosome, 205-264 central axial structures, 214 chromatin stabilization, 248-250 chromosome changes during cell cycle, 222-236 chromosome numbers, 209 28 1

282

SUBJECT INDEX

DNA fibrils, 251-252 DNA levels, 209-211 DNA transcription, 220-222 electron microscopy, 2 12-220 eukaryote affinities, 257-258 fine structure, 21 1-220 intermediate status and phylogeny, 258-260 ionic composition, 241-250 light microscopy, 21 1-212 macromo~ecu~ar compos~t~on, 236-241 models, 251-254 prokaryote affinities, 254-257 synthesis of nuclear DNA, 222-225 Dinophyceae, 207-208 Dinosterol, 208 Drosophila melanogaster, 158 DSP, see Diarrhetic shellfish poisoning Duboisia myoporoides, 88

E Enterococcus, 73 Escherichia coli, 18, 19 Exuviella, 240

G . catenatum, 52 G . indicum, 209 G . nelsoni, 236, 237 Gyrodinium, 218, 232

H Hepatoenteritis, 74 High-performance liquid chromatography (HPLC), 69 HPLC, see High-performance liquid chromatography

J Jania sp., 52, 83

K Kalanchoe, 15, 16, 18

L F Ferralterin, 15, 21 Ferredoxin, 8, 12-13 Ferredoxin-thioredoxin reductase, 14- 16 Fructose-biphosphatase, 4, 6, 23, 25-26

G Gambierdiscus, 52 G . toxicus, 63, 86 Glenodiniumfoliaceum, 212, 214, 216-217, 238, 242-243, 246-249 G . lubiniensiforme, 234 G . pulvisculus, 230 Glucose-6-phosphate dehydrogenase, 27 Glyceraldehyde-3-P dehydrogenase, 6, 27 Gonyaulax, 209, see also Protogonyaulax G. tamarensis, 225 Gonyautoxin, 83 Gymnodium breve, 237

Lathyrus, 184 LEM (light effect modulation) system, 8-13 Lighddark modulation, 1-46 CF,-CFo Mg ATPase, 37 changes in target enzyme, 30-36 kinetic parameters, 34-36 as result of dithioreitol treatment, 32-34 as result of light modulation, 30-32 as result of thioredoxin-DTTcatalyzed modulation, 32 cytosolic enzymes, 23-24 dark modulation, 28-30 dark reversal of light modulation, 28-29 enzyme activity, 4-6 ferralterin, 15, 21 ferredoxin-thioredoxin reductase, thioredoxin system, 14-20 fructose-biphosphatase, 4, 6, 23, 25-26 function of light modulation, 36-38 glucose-6-phosphate dehydrogenase, 27

283

SUBJECT INDEX

glyceraldehyde-3-phosphate dehydrogenase, 27 inhibitor experiments, 6-7 LEM system, 8-13 mechanisms, 6-24 NADP-linked malic dehydrogenase, 24 occurrence of, 3-6 osmotic stress, 38 photosynthetic carbon dioxide, 36-37 pyruvic, P, dikinase, 4,21-22 regulation of dark modulation, 30 regulation of light modulation, 24-27 ribulose-5-phosphate kinase, 27 sedoheptulose-biphosphatase, 6,26-27 sulfur dioxide, 38 thioredoxin in, 8,17-20,28-29 Lilium, 209 Lyngbya, 48 L . majuscula, 78-80 Lyngbyatoxin, 78 M

Maitotoxin (MTX), 86-89 Maize, 16,18 Mercenaria mercenaria, 62 Mesembryanthemum crystallinum, 34-35 Methylviologen, 7 Microcystitis, 48,69-72,75 M. aeruginosa, 64,69 Mirabilis jalapa, 137,185 MTX, see Maitotoxin Mutable alleles, 109 Mutations due to transposable elements,

109-15 1

Mya arenaria, 62 Myrothamnus Jlabellifolia (resurrection plant), 2 Mytilus edulis (blue mussel), 61-62

N Nicotiana, 138 Nodularia, 48 0 Oodinium dogieli, 233 Oscillatoria, 48 Oxyrrhis, 208

P Paralytic shellfish poisons (PSP), 52,59,

82-86 Paramutation, 185 Penaeus sfylirostris (blue shrimp), 79 P-enolpyruvate carboxylase, 4 Peridinium cinctum, 223,228,237 P . inconspicuum, 234 P . tabdatum, 209 P . trachoideum, 233,237 P . willei, 230 Petunia hybrida, 139-140,142,167 Phalloidin, 72 Pharbitis nil, 184 Phaseolus vulgaris (bean), 17 Pisum sativum (pea), 3-4 Placopecten magellanicus, 62 Plant transposable elements, 103-203 chromosomal anomalies, 141 cloning, 151-153 definition of, 105 dosage effects, 181-183 genomic instabilities, 188-193 excision of, 171-172 gene expression, 161-164 integration of, 172 interaction between, 177-188 large-scale rearrangements, 176-177 at molecular level, 151-170 mutable alleles, 109 mutations, 109-15 1 multiple copies of, 159-160 origin of receptor elements, 137 proteins encoded by, 164 reversion events, 137-139 segregation ratios, 109,133-137 specificity of integration, 177 structural features, 153-159 transposition, 144-151 variegation, 104-108 Polykrikos spp. 233 Procentrum, 52 P . concavum, 63 P . mexicanum, 63 Prokaryotes, 153,170-171 Prorocentrum micans, 21 1-212,218,

220-222,223,225-237

cell division, 230-236 interphase cells, 227-230 P . triestinum, 225

284

SUBJECT INDEX

Protogonyaulax, 52, 59 P. excauata, 60 P. (Gonyaulax) catenella, 59 P. polyedra, 59 Prymnesium paruum, 48, 52,58-59, 67-68 Pseudomonas tabaci, 255 PSP, see Paralytic shellfish poisons Ptychodiscus, 89 Pyocyanin, 7 Pyrodinium bahamense var. compressa, 52 Pyruvate Pi dikinase, 4, 21-22

R Red tides, 59, 206 Reverse variegation, 139-141 Reversion events, 137-139 at molecular level, 165-167 Rhodopseudomonas sphaeroides, 19 Ribulose-biphosphate carboxylase, 3-4 Ribulose-Sphosphate kinase, 27

S

Salmonella typhimurium, 73 Salsola soda, 34 Saxidomus gigentus, 82 Saxitoxin (STX), 82-86 Scaritoxin, 87 Scenedemus, 18 Schizothrix, 48 S.calcicola, 80 Secondary biosynthesis, 47-48 Sedoheptulose-biphosphatase,6, 26-27 Sedum praealtum, 34 Segregation ratios, 109, 133-137 Shigella Jexneri, 73 Siganus fuscescens (rabbitfish), 79 Spinach, 14, 22-23 Staphylococcus aureus, 73 Streptomyces mediocidicus, 80 STX, see Saxitoxin

Stylocheilus longicauda (sea hare), 79 Sulfite, 8, 38 Sulfur dioxide, 38 Swimmer’s itch, 48, 66, 78 Synechococcus, 18

T Teleocidin, 80

Tetramethylethylenediamine,7 Thioredoxin, 8, 17-20, 28-29 Transposition, 144-151 model, 170-177 at molecular level, 160-161 of Mp(Ac) in detail, 148-151 of receptor element, 145-146 of regulatory element, 146-148 replicative, 144 TST, see Turban shell toxin Turban shell toxin (TST), 83 Turbinaria turbinata, 63

V Variegation, 104-108 chloroplast mutations, 107-108 chromosomal anomalies, 141 mitochondria1 mutations, 107-108 reverse, 139-141 Vicia, 209 Vicinal dithiol, 8 W

Woloszynskia hiemale, 209 W . micra, 233 Z

Zea mays, 132-133, 156 Zwittergent, 9