Growth and Differentiation in Physarum Polycephalum 9781400885886

Table of contents :
Contents
Preface
Abbreviations and Genetic Symbols
Contributors
I. Introduction
II. The Nuclear Replication Cycle in Physarum polycephalum
III. Transcription in the Myxomycete Physarum polycephalum
IV. The Nuclear Proteins of Physarum polycephalum: A Comparative View
V. The Genetic Approach in the Analysis of the Biology of Physarum polycephalum
VI. Developmental Phases in the Life Cycle of Physarum and Related Myxomycetes
Bibliography
Index

Citation preview

Growth and Differentiation in Physarum polycephalum

Growth and Differentiation in Physarum polycephalum Edited by W. F. Dove and H. P. Rusch

PRINCETON, NEW JERSEY PRINCETON UNIVERSITY PRESS

Copyright © 1980 by Princeton University Press Published by Princeton University Press, Princeton, New Jersey In the United Kingdom: Princeton University Press, Guildford, Surrey ALL MGHTS RESERVED

Library of Congress Cataloging in Publication Data will be found on the last printed page of this book This book has been composed in Linotype Cahdonia Clothbound editions of Princeton University Press books are printed on acid-free paper, and binding materials are chosen for strength and durability Printed in the United States of America by Princeton University Press, Princeton, New Jersey

Princeton Legacy Library edition 2017 Paperback ISBN: 978-0-691-61589-9 Hardcover ISBN: 978-0-691-62891-2

Contents

PREFACE

IX

ABBREVIATIONS AND GENETIC SYMBOLS

Xl

CONTRIBUTORS

Chapter I.

XVli

Introduction

By Harold P. Rusch I. The Search II. Description of Physarum polycephalum

1 1 6

Chapter II. The Nuclear Replication Cycle in Physarum polycephalum

By Charles E. Holt I. Introduction II. Chromosomal DNA

9 9 11

III. Nucleolar DNA

51

IV. Concluding remarks

62

Chapter III.

Transcription in the Myxomycete Physarum polycephalum

By Peter W. Melera I. Introduction II. Complexity of the Physarum genome; ΙΌΝΑ

64 64 64

III. DNA-dependent RNA polymerase

68

IV. RNA transcripts

73

V. Ribosomal RNA

73

VI. Transfer RNA and 5S RNA

84

VII. Heterogeneous nuclear RNA and MRNA

86

VIII. Transcription during the mitotic cycle

90

IX. Concluding remarks

96 ν

CONTENTS

Chapter IV.

The Nuclear Proteins of Physarum polycephalum: A Comparative View

By B. W. Walker, M. E. Christensen, A. L. Beyer, and W. M. LeStourgeon I. Introduction II. Physarum histones and chromatin subunit structure

98 98 99

III. Physarum nuclear nonhistone proteins

105

IV. Physarum nucleolus and nucleolus-specific proteins

119

V. Nuclear RNA-binding proteins and mammalian 40S HnRNP particles VI. Concluding remarks Chapter V.

122 127

The Genetic Approach in the Analysis of the Biology of Physarum polycephalum

By Finn B. Haugli, David Cooke, and Peter Sudbery I. Introduction II. Genetic aspects of the Physarum cell nucleus

129 129 130

III. Genetic analysis of biological functions in Physarum polycephalum

133

IV. Methods of genetic analysis

147

V. Concluding remarks

155

Chapter VI. Developmental Phases in the Life Cycle of Physarum and Related Myxomycetes

By Jessica A. Gorman and Adam S. Wilkins I. Introduction II. Alternate states during the life cycle

157 157 158

III. Alternate modes of differentiation: sexual and asexual life cycles

163

IV. The amoebal-plasmodial transition

167

CONTENTS

V. The amoebo-flagellate transformation VI. Encystment: reversible differentiation to a dormant state VII. Sporulation

180 181 192

VIII. Concluding remarks

201

BIBLIOGRAPHY INDEX

203 245

Preface

Physarum polycephalum has lived in the McArdle Laboratory for Cancer Research at the University of Wisconsin for 25 years, spreading to laboratories in a number of countries during that span. We have assembled this group of critical essays to com­ municate the status of experimental analysis of the problems in growth and differentiation presented by Physarum. This mono­ graph does not aim to be comprehensive, either for Physarum or for any of the biological problems engaged here. What it does aim for is a statement of the current status and prospects for experimental analysis. We have felt it important to complement the critical efforts of each contributor by cross-criticism. Physarum research is still in its early stages, so we need to work from consensus. We are grateful to critics who freely offered additions and subtractions to the scientific content of this monograph: V. Allfrey (New York), R. Anderson (Sheffield), R. Braun (Bern), O.R. Collins (Berke­ ley), J. Dee (Leicester), H. and T. Evans (Cleveland), E. Guttes (Dallas), K. Raper, (Madison), W. Sachsenmaier (Innsbruck), H. Sauer (Wiirzburg), W. Schiebel (Miinchen), T. Shinnick (La JolIa), S. Smith (La Jolla), and V. Vogt (Ithaca). In addition, we were glad for active cross-exchanges among the contributors of the essays. Voluntary offerings of unpublished manuscripts were made by a number of active Physarum researchers. This sort of courtesy motivates the informal circular Physarum Newsletter—organized by Thomas Evans of Case Western Reserve University, Cleve­ land, Ohio—by which all Physarum workers help one another to deal with the literature. Several scientists came to the fore to provide illustrative ma­ terial for the monograph: J. Dee, J. Gorman, C. E. Holt, A. Hiittermann (Gottingen), and J. Mohberg (Chicago). We are particularly grateful to Herr Galle, director of the Institut fur den Wissenschaftlichen Film, Gottingen, for material from the films on Physarum under preparation by that institute. The Princeton University Press has helped us throughout this

PREFACE

endeavor. Mr. Edward Tenner encouraged us to initiate this monograph; Mrs. Margaret Case and Ms. Joanna Ajdukiewicz helped throughout the execution of the work; and Ms. Avis KnifiBn helped greatly to improve our style. We end by thanking Dr. Ilse Riegel of the McArdle Labora­ tory, who has played a central role in coordinating the manu­ scripts from which this monograph has been fashioned. With the assistance of Ms. Bette Sheehan, she has served not only to keep pages and references in order, but also to exert a rare sense of judgment over the scientific clarity of each sentence on each page. We hope that this set of critical essays can form the basis for an active growth phase in Physarum research. The authors have agreed to make available, over the next several years, an anno­ tated, up-to-date bibliography. This material, collated and peerreviewed once every year, will be available from the McArdIe Laboratory, University of Wisconsin, Madison, Wisconsin 53706, and will also be reprinted in the Physarum Newsletter. Madison, Wisconsin March, 1980

χ

Abbreviations and Genetic Symbols

ABBREVIATIONS

General 2', 3', and 5'

positions on the ribose or deoxyribose group of nucleosides

C value

the haploid unreplicated DNA level in an organism

CHL

Chinese hamster lung cell line

CHO

Chinese hamster ovary cell line

Cot

a parameter of nucleic acid reassociation ki­ netic measurements: the initial concentration of the rate-controlling species of nucleic acid multiplied by the time of reaction

Cotl/2

the mid-value for reassociation of a particu­ lar species of nucleic acid in a mixture, re­ lated to the recurrence of that species in the mixture

DEAE

a diethylaminoethyl moiety of a polymer used in anion exchange chromatography

DNase

deoxyribonuclease

EcoRI

a restriction endonuclease cleaving at

G/AATTC. CTTAA/G

G1 phase

the period in the cell replication cycle sepa­ rating mitosis from the major period of DNA synthesis

G2 phase

the period in the cell replication cycle sepa­ rating the DNA synthesis period from cell mitosis

ABBREVIATIONS AND GENETIC SYMBOLS

HeLa

a human cell line

HindIII

a restriction endonuclease A/AGCTT cleaving at TTCGA/A

HKC

a histone kinase activity from mammalian liver

HnRNA

heterogeneous (in size) nuclear RNA

HnRNP

heterogeneous nuclear ribonuclear protein complex

L

mouse fibroblast cell line

log-phase

the period in which a population grows with constant doubling time the wavelength of maximal light absorption of a compound

MI, Mil, Mill (or Ml, M2, M3)

the first, second, and third synchronous mi­ tosis of a plasmodium after fusion of microplasmodia

mBNA

messenger RNA

NHP

non-histone nuclear proteins

OD265

optical density (absorbance) at 265 νημ wave­ length

oligo (dT)

oligomer of deoxythymidylate

poly (A)

polymer of adenylate

poly (ADP-ribose)

ribose-linked polymer of adenosine diphos­ phate

poly (d(A-T))

a polymer of alternating deoxyadenylate and deoxythymidylate

TDNA

DNA encoding ribosomal RNA

REM

relative electrophoretic mobility

RNase

ribonuclease

RNP

ribonucleoprotein complex

xii

ABBREVIATIONS AND GENETIC SYMBOLS

RPC

reversed-phase liquid partition chromatog­ raphy

FRNA

ribosomal RNA

S

Svedberg unit of sedimentation velocity

S phase

the period in the cell replication cycle in which most DNA synthesis occurs

5S RNA

a small ribosomal RNA of sedimentation ve­ locity near 5 Svedbergs

Tm

the mid-temperature of a thermal naturation curve

TK

thymidine kinase

tRNA

transfer RNA

tRNA val

the transfer RNA that accepts valine

DNA

de-

ABBREVIATIONS

Chemical

ATP

adenosine triphosphate

BUDR

5-bromodeoxyuridine

C

cytosine

cAMP

cyclic adenosine monophosphate

dAMP

deoxyadenosine monophosphate

dATP

deoxyadenosine triphosphate

dCMP

deoxycytosine monophosphate

dGMP

deoxyguanosine monophosphate

dTMP

deoxythymidine monophosphate

dTTP

deoxythymidine triphosphate

DOPA

dihydroxyphenylalanine

EDTA

ethylenediamine tetraacetic acid

EMS

ethyl methane sulfonate XLLL

ABBREVIATIONS AND GENETIC SYMBOLS

FUDR

5-fluorodeoxyuridine

G

guanine

GTP

guanosine triphosphate

NAD

nicotine-adenosine dinucleotide

NAD+

oxidized form of NAD

NADP

nicotine adenosine disphosphate

NG

N-methyl-N'-nitro-N-nitrosoguanidine

PMSF

phenylmethylsulfonyl fluoride, a protease inhibitor

SDS

sodium dodecyl sulfate detergent

TTP

thymidine triphosphate

UDP

uridine diphosphate

UR

uridine

UTP

uridine triphosphate

GENETIC SYMBOLS

act

resistant to actidione (cycloheximide)

ale

amoeba-less life cycle

apt

deficient in amoebal-plasmodial transition

bur

resistant to BUDR-induced killing

CL

Colonia Leicester

CLd

Colonia Leicester, delayed in plasmodiuni formation

cpf

clonal plasmodium former

dif

deficient in differentiation from amoeba to plasmodium

fus

plasmodial somatic fusion incompatibility

gad

good asexual differentiation

kil, let

lethal interaction upon plasmodial fusion

leu

deficiency in leucine biosynthesis

Iys

deficiency in lysine biosynthesis

xiv

ABBREVIATIONS AND GENETIC SYMBOLS

mt

mating type

mth

"homothallic" mating type, in CL

npf

non-plasmodium former

rac

rapid amoebal crossing

whi

white Plasmodium

Contributors

Chapter I HABOLD P. RUSCH, McArdle Laboratory for Cancer Research,

University of Wisconsin, Madison Chapter Il CHARLES E. HOLT, Department of Biology, Massachusetts In­

stitute of Technology Chapter III PETER W. MELERA, Laboratory of RNA Synthesis and Regulation,

Sloan-Kettering Institute for Cancer Research, Walker Labora­ tory, Rye, New York Chapter IV

B. W. WALKER, M. E. CHRISTENSEN, A. L. BEYER, and W. M. LESTOURGEON, Department of Molecular Biology, Vanderbilt University Chapter V FINN B. HAUGLI, Institute of Medical Biology, University of

Tromso, Norway DAVID COOKE and PETER SUDBERY, Department of Genetics, Uni­

versity of Sheffield, England Chapter Vl JESSICA A. GORMAN, McArdle Laboratory for Cancer Research,

University of Wisconsin, Madison ADAM S. WILKINS, Department of Microbiology and Genetics,

Massey University, New Zealand

Growth and Differentiation in Physarum polycephalum

I.

Introduction HAROLD P. RUSCH

I. THE SEARCH

My interest in the cause and nature of cancer started in 1936 when I began experiments with chemicals and ultraviolet radia­ tion as causative agents. During the next decade many chemicals were identified as carcinogens, and evidence increased that various animal species reacted differently to carcinogens and that a number of factors, such as diet, could influence the re­ sponse of animals to carcinogens. I became deeply involved in experiments with carcinogenesis during the late thirties and throughout the forties. It is a fascinating and fruitful area of research, which has become even more popular recently because of increasing awareness of environmental contaminants. During the forties, three circumstances combined to influence the course of my future research. Elizabeth and James Miller, Charles Heidelberger, and Roswell K. Boutwell came to the McArdle Laboratory for Cancer Research. They were imagina­ tive, enthusiastic, energetic, well-trained biochemists who ini­ tiated highly original, first-rate investigations with chemical car­ cinogens. It soon became obvious that the studies on chemical carcinogens at McArdle were in exceptionally good hands. Sec­ ondly, the experiments I did with carcinogens required the use of mice and rats, to which I developed an allergy. Lastly, I had a growing feeling that we must have a simple model system for biochemical studies on growth and differentiation to obtain a clearer understanding of these two important events in the life cycle of cells. Cancer tissue is composed of a mixture of cancer cells in all stages of the cell cycle plus a variety of other cell types, including those derived from blood vessels, blood, and fibrous tissue. This heterogeneity complicates the interpretation of results obtained from biochemical studies on cancer tissues. Various models have been developed to overcome these problems, such as the use of

RUSCH

synchronized cells grown in culture, but the methods for the employment of such cells did not exist in the late forties. I was convinced, therefore, that it was necessary to study the basic biochemical aspects of growth and differentiation on a simple primitive organism in which these stages of the life cycle were distinct and separate. Where does one look for such an organism? One may ask colleagues or search the literature. I did both, and eventually found a reference to Raper's work on Dictyostelium discoideum, an organism that seemed to be ideal for my purpose. Kenneth Raper (1935) had published a paper on this organism while he was at Harvard University, but when I wrote to him for a specimen, I waited for a considerable period before getting a reply from the Northern Regional Research Laboratory in Peoria, Illinois, where he had gone during World War II to seek new antibiotics from various organisms. After learning that I could obtain a specimen, I drove to Peoria, was graciously re­ ceived, and returned with a specimen of D. discoideum. This was early in 1947 when wartime gas rationing was still in effect, but by skimping on the use of my car for some period, I saved sufficient gas coupons to make the trip from Madison to Peoria at a speed not exceeding 35 miles an hour, which was the regulation at that time. Dictyostelium was grown on bacteria as the source of nutrient, and I spent the next three years attempting to culture it on an axenic medium. I had the loyal and able assistance of Erich Hirschberg. We failed to culture it on an axenic medium but tested the effects of a number of chemicals on the aggregation of previously independent myxamoebae; this aggregation is a prelude to the cellular differentiation exhibited during sporulation (Hirschberg and Rusch, 1950). I was deeply disappointed not to have succeeded in the axenic culture, since having to work with cultures grown on bacteria would have severely com­ plicated any biochemical determinations; even now, axenic cul­ ture of Dictyostelium is not routine and is limited to certain mutants. During this period of frustration, I became better acquainted with the field of mycology and noticed a paper by Howard (1932) that described synchronous mitosis in the plasmodium of Physarum polycephalum, an organism first described by Schweinitz in 1822. At the time we began our studies, this

INTRODUCTION

organism had received little attention for over a hundred years except for a few papers on taxonomy and morphology (de Bary, 1860; Camp, 1937; Gray, 1938) and on protoplasmic streaming (Seifriz, 1936). Except for Loewy's (1952) studies on the pres­ ence of an actomyosin-like protein in the plasmodium, I found no reports of biochemical studies on the organism when I began my investigations in the early fifties. Howard's description of mitotic synchrony stimulated my interest, and I was convinced that P. polycephalum would be ideal for my purpose. I asked Myron Backus, a mycologist at the University of Wisconsin, how to collect a specimen. He described the procedure in detail but then informed me that it was unnecessary to go to the trouble, since he would give me a specimen of Physarum, which he had used in class demonstrations for a number of years. The earlier investigators had grown the organism on various lands of media, but all cultures were grossly contaminated with many other organisms that served as nutrients. Since my planned biochemical studies would require an organism free of con­ taminating microbes, I began the long search for an axenic medium. I tried all the media I had used in my efforts to grow Dictyostelium discoideum (malt liquors, yeast and bacterial extracts, etc.) without success. I was successful, however, in obtaining excellent growth on autoclaved Quaker oatmeal flakes as Camp had done (1937), provided that the oatmeal was of the "old-fashioned" variety and that the flakes were dry before being autoclaved. Heating wet oatmeal destroyed some necessary growth factor, so sterile water was added only after the autoclaving. Using such a medium, I obtained a culture free of bacteria and kept an actively growing culture while the search continued for a chemically more defined medium. In 1953 I offered a postdoctoral fellowship to John Daniel to assist me, since his major professor, Marvin Johnson of the Biochemistry Department, had considerable experience with the culture of microorganisms. We again tried almost every culture medium that had ever been described for the growth of microorganisms. We had no luck until about 1956, when 1 suggested that we add a chick embryo extract to our medium, because such extracts had been used for many years for the growth of cells grown in cul­ ture. To our delight the organism grew (Daniel and Rusch, 1961). We soon learned that the embryo extract could be re­ placed by hematin, and the path was finally clear to conduct

RUSCH

biochemical experiments on the organism grown on a defined axenic medium (Daniel et al., 1962). We have since learned that hemin had solved the problem of culturing Tetrahymena (Kidder and Dewey, 1951), but this fact was unknown to us when we first tested it. We continued to be plagued by another problem. We could not control the onset of sporulation (differentiation), and we wanted to investigate the biochemical changes that occur when the organism shifts from growth to differentiation. Previous investigators had noted that growth occurred best in the dark and that the organism would sporulate if subsequently exposed to light, but the exact procedure was not defined (Gray, 1938). We tried various combinations of the lengths of the dark and light periods, with inconsistent results. After about a year of trial and error we found that it was necessary first to starve the Plasmodium in the presence of niacin for a minimum of four days and then to expose it to light for a minimum of two hours (Daniel and Rusch, 1962a, 1962b). We were now ready to initiate studies on the biochemical differences between growing and differentiating (sporulating) axenic cultures of Physarum polycephalum. One of my main hopes was to isolate a fraction or fractions from sporulating Plasmodia that would inhibit growth and induce sporulation. A determination of the nature of such substances could give clues to a class of compounds that might be present in mammalian cells and that would inhibit the growth of cancer. Many attempts in our laboratory to obtain such fractions were unsuccessful. It is encouraging to learn that Wormington and Weaver (1976) recently have reported that sporulation can be induced by injecting an extract of illuminated plasmodia into a starved Plasmodium. Another goal was to learn about the mechanisms that initiated differentiation following starvation. If such an event were con­ fined only to Physarum, it could be dismissed as something pe­ culiar to this organism, but the initiation of differentiation by the depletion of nutrients is ubiquitous. Many microorganisms form spores or undergo other changes to survive when nutrients be­ come limited, and mammalian cells grown in culture may also dif­ ferentiate in response to this condition (Schubert and Lacorbiere, 1976). The limitation of nutrients may also induce differentiation in intact mammalian organs such as skin. Epidermal cells begin

FIG. 1.1. Schematic diagram of the life cycle of Physarum (Drawing by J. A. Gorman.)

polycephalum.

FIG. 1.2. Macroplasmodium. The Plasmodium is seen growing on the surface of an agar plate (Dee and Poulter, 1970) in a standard (9 cm diameter) petri dish. The agar block that bore the inoculum is in the center. The Plasmodium has prominent veins but is continuous, as is typical of a rapidly growing, well-fed plasmodium on synthetic medium. (Courtesy of J. Dee and C. E. Holt)

FIG. 1.4. Sporangium formation. After starvation of the plasmodium and illumination, fruiting bodies form, to be followed by melanization of spores. (Courtesy of H. P. Rusch)

FIG. 1.3. Plasmodia! nuclear division cycle. A field of synchronous nuclei is seen: (a) interphase, duration 480 minutes; (b) early prophase, duration 15-20 minutes; (c) metaphase, duration 7 minutes; (d) reconstruction, duration 75 minutes. (Courtesy of J. Mohberg)

FIG. 1.5. Amoebal mitosis. A growing amoeba has rounded up and is undergoing mitosis. The diameter of the ceil is 13 μπι (a) Metaphase. (b) Anaphase, 1.1 minutes after metaphase; the chromosomes have moved about halfway from the position of the metaphase plate to their final posi­ tions. (c) Telophase, 3.1 minutes after metaphase; chromosomes are still condensed, (d) Cytokinesis, 5.4 minutes after metaphase; the chromosome masses are no longer visible. (C. E. Holt, A. Hiittermann, and H.-K. Galle)

FIG. 1.6. Nuclear fusion in mating. A mixture of amoebae differing at the rnating-type locus were exposed to conditions favoring mating. The binucleate cell in panel (a) was formed by the fusion of two cells approxi­ mately 15 minutes before this frame was taken; (b) nuclei in contact 13 minutes later; (c) onset of nuclear fusion, 30 minutes after (a); (d) nuclear fusion complete, 32 minutes after (a); (e) onset of nucleolar fusion, 52 minutes after (a); (f) nucleolar fusion complete, 55 minutes after (a). The nuclei are about 4 μτη in diameter. (Courtesy of A. Hiittermann, C. E. Holt, and H.-K. Galle)

INTRODUCTION

to keratinize after one of the two daughter cells from a dividing basal cell is squeezed upward because there is no space for the cell either at the basal layer or downward through the basement membrane. Once the cell is squeezed above the basal layer, its relationship to nutrients from the blood supply is altered and differentiation is initiated. The removal of a piece of skin permits basal cells to divide and spread laterally until the defect is cov­ ered, and then differentiation again stops the continued exponen­ tial cell division. The immediate reactions that trigger differen­ tiation following a depletion of nutrients are still a mystery and deserve more attention. Although we failed to accomplish some of my original goals, we did succeed in collecting considerable information about the cytological and biochemical characteristics of growth and differ­ entiation (Rusch, 1970). I was fortunate to be able to attract a considerable number of outstanding postdoctoral fellows, who conducted most of the experiments. These fellows have scat­ tered over this country and abroad, and most of them have con­ tinued research on various aspects of P. polycephalum. Among these many excellent scientists, Joyce Mohberg deserves especial credit. She was a member of my research team for twelve years starting in 1961, and as an increasing amount of my time, over the years, was spent in administration, she became the central source of information and guidance about Physarum. In addition, our laboratory has supplied cultures of the organism to many other investigators. As a result, there are now sufficient people working with the organism to warrant scheduling special con­ ferences both in the United States and abroad. It is also of interest that Jennifer Dee (see Anderson and Dee, 1977) obtained a specimen of Physarum from us to pioneer genetic studies. Her major professor, Guido Pontecorvo, heard about the organism at a conference in Edinburgh in 1957 where I presented a paper (Rusch, 1959). A number of investigators are now working on Physarum genetics, including William F. Dove and his associates in the McArdle Laboratory for Cancer Research. The history of financial (support for this project raises a ques­ tion about assistance for new projects. Such funds were, and still are, difficult to obtain. Fortunately, as Director of the McArdle Laboratory, I had some funds to support the work during the lean years when no papers were published, but it is doubtful whether a younger person could initiate and sustain

RUSCH

such a program at this time over a similar period without pub­ lications on the subject. Applications for initiating new areas are often disapproved on the basis that "the applicant lacks experi­ ence in the field," "exact details are lacking," or that the research is "a fishing expedition." This tends to discourage applications for new approaches and to encourage research that is either repetitious or simply an extension of previous work. II. DESCRIPTION OF Physarum polycephalum

This organism is a plasmodium-forming slime mold of the class Myxomyceteae, phylum Myxomycophyta, and is commonly re­ ferred to as a true slime mold, or myxomycete (Alexopoulos, 1962). The life cycle, illustrated in Fig. 1.1, is in sharp contrast to that of the Acrasiomyceteae (or cellular slime molds), to which Dictyostelium belongs. For example, fructification of Physarum involves compartmentalization of the plasmodial protoplasm, whereas fructification in Dictyostelium is accomplished by the aggregation and differentiation of a community of cells. Because of the appearance of their fructifications, mycologists have gen­ erally classified both types of slime molds as plants. Protozoologists, on the other hand, emphasize the amoeboid vegetative states and include these organisms among the protozoa. However classified, Physarum polycephalum represents a primitive, ex­ perimentally useful eukaryote. A. Vegetative Plasmodial Growth The plasmodium represents the macroscopic vegetative form of Physarum. In this stage of the life cycle, growth occurs in the absence of cell division, giving rise to a large multinucleate structure. Plasmodia can be grown either on surfaces or in shaken culture. On surfaces the multinucleate plasmodium is yellow, flat, disklike, and consists of protoplasm that exhibits a rapid toand-fro streaming (Fig. 1.2). With adequate nutrition and at optimal temperatures of 21°-24°C, mitosis occurs about every eight hours and is synchronous (Rusch, 1970) (Fig. 1.3). Struc­ turally and because of the mitotic synchrony, the plasmodium may be regarded for biochemical purposes as one giant cell. In shaken flasks the plasmodium is fragmented. Each small microplasmodium exhibits synchronous mitosis, but there is no syn­ chrony among the various microplasmodia within the culture

INTRODUCTION

flask. When the culture is transferred from the flask onto a suitable surface, the microplasmodia coalesce into one large Plasmodium, which then establishes nuclear synchrony (Guttes and Guttes, 1964b). B. Sporulation When surface plasmodia are starved by removal of nutrients, they begin to move about and form a network of streaming protoplasm. If food is located, the movement stops until the food is gone, and then the movement starts again. If the starva­ tion is continued in the dark for approximately four days and the Plasmodium is then exposed to daylight or fluorescent light for a minimum of four hours, sporulation is induced (Daniel and Rusch, 1962a). If nutrients are returned to the starving Plas­ modium at any time during the first three hours following the end of illumination, growth resumes. After three hours, the processes leading to sporulation become irreversible, and at least one mitosis occurs during starvation prior to the onset of sporu­ lation (Sauer et al., 1969c). Sporulation consists of the formation of sporangial strands and the development of many pillars by the pulsating protoplasm (Fig. 1.4). Nuclei migrate up the pillars and become concentrated in sporangial caps. Successive cleavage of the cytoplasm results in the formation of uninucleate immature spores. The partitioned nuclei then divide meiotically. Melanin pigmentation occurs in the spore walls at the end of sporulation, the entire process taking about 16 hours from the end of the light period (Guttes et al., 1961). C. Amoebal Growth Under suitable conditions the spores germinate and release amoebae, which divide and multiply when grown on cultures of bacteria. In contrast to the plasmodial stage, amoebal nuclear division is accompanied by cell division (Fig. 1.5). Thus, this vegetative stage consists of small, free-living uninucleate cells. The amoebae may also be grown as axenic cultures, but the growth is not so vigorous nor so rapid as on bacteria (Goodman, 1972; McCullough and Dee, 1976). When amoebae are placed in suspension, they quickly form flagella and move about actively until a solid substrate is encountered, after which they return to the amoeboid form. The change from amoebae to flagellated

RUSCH

forms (swimmers) is thus readily reversible. Uninucleate amoebae are more amenable to genetic manipulation than is the multinucleate plasmodium, and many studies employing myxamoebae have been reported by J. Dee; J. Gorman and W. Dove; F. Haugli; C.E. Holt; D.N. Jacobson; and others. D. Plasmodium Formation The plasmodial stage is re-established by the differentiation of cells within an amoebal population. Generally this occurs fol­ lowing a mating between two genetically different amoebae (Fig. 1.6). Following syngamy, nuclear division again occurs in the absence of cytoplasmic division, and a multinucleate plas­ modium develops. E. Encystment and Sclerotial Formation Amoebae subjected to adverse conditions encyst and can sur­ vive in such an environment for considerable periods. Surface and shaken plasmodia also develop resistant forms called sclerotia when subjected to unfavorable conditions. The former form large sheetlike sclerotia, and the latter form small rounded sclerotia called microsclerotia or spherules. These dormant forms provide a convenient means of storing the organism over pro­ longed periods. It should be emphasized that when plasmodia are starved in the dark, they form sclerotia. By contrast, plas­ modia starved and exposed to light develop sporangia and pass through sporulation. F. Protoplasmic Streaming One of the more spectacular characteristics of the plasmodium of P. polycephalum is the very active, reciprocal protoplasmic streaming. In addition to the work of Seifriz (1936), early studies about this phenomenon have been made by Kamiya (1960), and by Wohlfarth-Bottermann (1977). Many others have pub­ lished papers about this streaming in recent years. In addition, a considerable number of reports have been published about the chemical composition of the contractile fibers actin and myosin, which closely resemble muscle fibers and are responsible for the movement (Tanaka and Hatano, 1972).

The Nuclear Replication Cycle in Physarum polycephalum CHARLES E. HOLT

I. INTRODUCTION

Studies on nuclear replication in the multinucleate plasmodium of Physarum polycephalum have focused on two broad areas: DNA synthesis and the control of the time of mitosis. The close temporal association of mitosis with the initiation of chromosomal DNA synthesis (Nygaard et al., 1960) and the apparent need of a nucleus to pass through mitosis before beginning DNA synthesis (Guttes and Guttes, 1968) imply the existence of intimate con­ nections between DNA synthesis and mitosis. Studies in the two areas remain, nonetheless, rather separate from one another, and it is a major challenge for future workers to make the con­ nections between the detailed events of DNA synthesis and the higher-level events that result in the well-known synchronous nuclear division of plasmodia. P. polycephalum nuclei contain two distinct DNA fractions— the bulk, or chromosomal, DNA and a minor DNA species. The minor species, which was first observed as a shoulder on the density profile of nuclear DNA (Braun et al., 1965), is synthesized during the G2 period (Holt and Gurney, 1969; Braun and Evans, 1969) as well as most of the S period (Zellweger et al., 1972), is located in the nucleolus (Guttes and Guttes, 1969; Zellweger et al., 1972; Ryser et al., 1973), and codes for TRNA (Sonenshein et al., 1970; Zellweger et al., 1972; Newlon et al., 1973). This review is divided into two major parts corresponding to the two nuclear DNA species. Within each part the discussion of DNAlevel work is followed by discussion of control. The reader will note that the paucity of studies relevant to the control of nucleolar DNA synthesis contrasts with the large number of studies con­ cerned with the timing of mitosis and by implication with the onset of the chromosomal DNA synthesis phase of the cell cycle, S.

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In addition to the two nuclear DNA species, P. polycephalum has been shown to contain mitochondrial DNA (Guttes and Guttes, 1964a). This species represents about 10 percent of the total DNA, has a buoyant density (1.686 g/cnr) lower than that of either nuclear species, and is synthesized throughout the mitotic cycle (T.E. Evans, 1966; Holt and Gurney, 1969; Braun and Evans, 1969). Further information is available on the localiza­ tion (Kuroiwa, 1974a, 1974b) and structure (Evans and Suskind, 1971) of the mitochondrial DNA. The germinal papers in the area of this review are those of Daniel and Rusch (1961) on axenic growth of plasmodia, Guttes et al. (1961) on the cytology of the synchronous nuclear cycle in plasmodia, and Nygaard et al. (1960) on the incorporation of radioactively labeled orotic acid into RNA and DNA at various times of the cell cycle. The incorporation study shows that DNA synthesis in plasmodia occurs mainly during a period immediately following mitosis (Fig. II.1). Thus, there is no G, period, and the S period occupies about one-third of the cell cycle. The ease with which this determination of S period was made spawned the long, still-continuing series of experiments on the relation of DXA synthesis to the cell cycle.

FIG. II.1. Measurement of time of S phase in Physarum polycephalum plasmodia. Below: Appearance of plasmodial culture in section. The dia­ gram shows a growing plasmodium (stippled area) resting on the surface of a disk of filter paper that is supported by glass beads. Liquid culture medium (diagonally hatched area) fills the space between the filter paper and the bottom of the petri dish. Next page: Incorporation of Ce-llCJorotic acid into DNA. Plasmodia like the one shown below were exposed to medium containing [6-1 'C]orotic acid for the three-hour intervals shown. At the end of each of the intervals, the labeled plasmodium was bisected, and half was used for determination of radioactivity in DNA-thymine. Incu­ bation of the other half continued, and it was sampled periodically for determination of time of mitosis. The symbols D1 and D11 on the lower axis indicate times of mitotic division. (From Nygaard et al., I960; re­ printed by permission of the author and publisher.)

NUCLEAR REPLICATION CYCLE

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where η is the degree of repetition of the component in ques­ tion and C0t,/2 (E. coli) is the value of C0t1/2 for E. coli DNA reassociated with the same salt concentration, molecular weight, and temperature. Values for the C0t1/2 of the slowest reannealing

NUCLEAR REPLICATION CYCLE

DNA have ranged from 500-1,100 mol· sec-liter1 and are com­ patible with this component's being single-copy DNA if one esti­

mates the value of C0ti/2 (-E- coli) under the renaturation condi­ tions employed. The repetition frequency (n) for the middle repetitive DNA has not been well established, and the suggestion that a high degree of mismatching occurs should be checked by measuring the melting temperature of the reannealed material (Cordeiro-Stone and Lee, 1976). Inverted-repeat sequences typically comprise several percent of eukaryotic DNA (Deininger and Schmid, 1976; Davidson et al., 1975). Such sequences are detected in DNA that has been sheared to a length of about 5,000 nucleotide pairs, which is about a factor of ten greater than the length used for quantitative reassociation (cot curve) analysis. When nuclear DNA SO sheared is denatured, exposed briefly to renaturing conditions, and examined in the electron microscope, a substantial number of the DNA fragments are found to have a "hairpin" configuration. A "twotailed" hairpin is illustrated schematically in Fig. II.2. Examina­ tion of P. polycephalum nuclear DNA treated in this fashion revealed that 22 percent of the molecules contained one or more double-stranded regions (Hardman and Jack, 1977). Nearly all of the molecules with regions of double helix had forms comFIG. II.2. Nomenclature for two-tailed hairpins. The symbols s and t represent distances; ss and ds mean single stranded and double stranded respectively. (From Hardman and Jack, 1977; used by permission of the author and publisher.)

t

LOOP(ss) STEM (ds) TAIL (ss)

HOLT

patible with their having arisen from intramolecular reassociation. About half of the structures were two-tailed hairpins, which may be assumed to carry intact (unsheared) "stem" (Fig. II.2) regions. About 42 percent of the two-tailed hairpins contained no loop. The remainder had a detectable loop, and the average loop length in these was 1,200 nucleotides. The longest loops, which were rare, were about 5,000 nucleotides long. The average stem length was 340 nucleotide pairs; this average was the same for both looped and unlooped hairpins. The longest stems, which were also rare, were about 3,000 nucleotide pairs long. The ge­ nomic regions classified as inverted repeats might represent a sub­ class of the middle repetitive DNA. However, since we have only preliminary information on the amount and degree of repetition of middle repetitive DNA in P. polycephalum, and no information in this organism on the degree of interspersion of repetitive DNA with single-copy DNA, the possibility cannot be fully evaluated. Whether or not repeated sequences have functional significance is not known for this or any other organism. However, intriguing possibilities have been suggested (Wallace and Kass, 1976; Davidson et al., 1975), and further description of the sequences would be of great interest. 7. DNA coding for tRNA and 5S RNA. The DNA complementary to 5S RNA and tRNA is located in the main band DNA (Hall and Braun, 1977). At hybridization saturation, 0.0064 percent of the DNA binds 5S RNA; this percentage saturation value corresponds to 270 copies per haploid (unreplicated or G1) genome. Satura­ tion values for tRNA are 0.0062 percent (Hall and Braun, 1977) or 0.0043 percent (Newlon et al., 1973), which correspond to 418 and 290 copies per haploid, Gi genome, respectively. For comparison there are about 80 copies of the TRNA genes in such a nucleus (see subsequent section). B. Synthesis 1. Time of initiation of S phase. The onset of DNA synthesis and the completion of mitosis occur at essentially the same time (Fig. II.1). To discuss the relation of the events more precisely, it is necessary to define what is meant by telophase in P. poly­ cephalum. Mohberg and Rusch (1969a) present a series of sketches of various stages, reproduced here as Fig. II.3. The information they collected is particularly useful, since it in­ cludes estimates of the duration and time of the various micro-

NUCLEAR REPLICATION CYCLE

FIG. II.3. Mitotic stages in the nucleus of the Physarum plasmodium. Alcohol-fixed plasmodial smears were .photographed through a phasecontrast microscope. Representative nuclei on the photographs were traced to give the drawings of the figure. All drawings are at the same magnifica­ tion. On the first line below each row of nuclei is the duration in minutes of each stage of mitosis, and on the second line is the number of minutes before the beginning or after the end of metaphase. (From Mohberg and Rusch, 1969a; reprinted by permission of the author and publisher.) MITOTIC STAGE

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scopically distinguishable phases. (Photomicrographs of the various stages may be found in Guttes et al., 1961, 1968 and Jockusch, 1975). Telophase is given as a stage of 5-min duration during which the nuclei are elongate and contain two phasecontrast-dense areas, which are the masses of chromosomes at the poles of the mitotic spindle. This phase must be terminated by karyokinesis, since the next stage ("reconstruction") observed in the fixed material begins with nuclei that are substantially smaller and display only a single region of phase-contrast-dense material. Mohberg and Rusch also point out that there is de­ tectable, if small, variation in the time of mitosis among various regions of a plasmodium grown in the conventional way in a petri dish. The variation from one edge to the opposite edge was typically 5 min. Although this is a small variation on the scale of the whole mitotic cycle, the degree of asynchrony it represents

HOLT

is not negligible in evaluating the precise time that DNA synthesis starts. Sachsenmaier (1964) exposed samples of plasmodia in different stages of the cell cycle to [3H]-thymidine and measured the incorporation of radioactivity into acid-insoluble material. The duration of exposure to label was 15 min. When the labeling period began at telophase, the extent of incorporation greatly exceeded that in the G2 period; this indicated that DNA synthesis had begun by the end of the time telophase +15 min. When the labeling period terminated at telophase, the extent of labeling was slightly, but significantly, higher than the background (pre­ sumably mitochondrial DNA) labeling seen in G2. Sachsenmaier pointed out that this pretelophase increase in thymidine incor­ poration might have been brought about by a few nuclei that were slightly advanced in the mitotic stage relative to the others, rather than by actual initiation prior to telophase. On the basis of the same type of experiment reported by Sachsenmaier, but using 10-min [3H] thymidine pulses, Braun et al. (1965) found that incorporation reaches its full rate within 10 min of metaphase. They also measured [3H]thymidine incorporation by auto­ radiography of plasmodia pulsed for 20 min, and 100 percent of the nuclei incorporated radioactivity in the first pulse after mitosis. Kessler (1967) reduced the [3H]thymidine pulse time to 5 min and conducted pulses every 4 min through the mitotic period. He reports that DNA synthesis "began in the sample immediately after anaphase"; if one assumes that the Mohberg and Rusch timing data (Fig. II.3) apply, then one would have to conclude that DNA synthesis begins in telophase. In summary, it is clear that DNA synthesis begins either during or within 5 min either way of the 5-min telophase period, but it is not clear how abruptly the synthesis begins in a given nucleus nor precisely when the synthesis begins, relative to the events of chromosome movement, karyokinesis, and the beginnings of chromosome extension. Further investigation, probably with quantitative autoradiography of plasmodia labeled for short time intervals, would be desirable to resolve the ambiguities. It is clear that, in this organism, movement of chromosomes to the poles of the spindle apparatus, karyokinesis, initiation of DNA synthesis, "uncoiling" of the chromosomes, and resumption of RNA synthesis (Kessler, 1967) all occur within a very short period of time. Moreover, it seems likely that all the events are

NUCLEAR REPLICATION CYCLE

more or less directly related to one another. As further informa­ tion accumulates in such areas as the structure of chromatin, the enzymatic basis of eukaryotic DNA synthesis, and the role of microtubules in the movement of chromosomes, P. polycephalum plasmodia should provide excellent material for further investiga­ tion of the degree and nature of the coupling among these con­ current events. The ease of isolation of large amounts of ma­ terial from defined stages and the possibility of genetic analysis (Chapter V) reinforce this view. 2. Time course of S phase. The incorporation of [3H!thymidine in pulses at different times of the cell cycle rises immediately to a plateau level (Braun et al., 1965; Fig. II.4). Other data from FIG. II.4. Incorporation of [3HJthymidine into DNA. The experiment is of the same basic design as that in Fig. II.1 except that a different radio­ active precursor and shorter (10-min) exposure to radioactivity was used. M3 indicates time of mitosis (the third after the cultures were prepared). (From Braun et al., 1965; reprinted by permission of the author and publisher.)

M3

HOURS

similar experiments (Sachsenmaier, 1964; Braun and Wili, 1969) show a less well-defined plateau region, but the general picture of incorporation increasing sharply and maintaining a high level is still confirmed. An important question to ask is whether the pattern of incorporation versus time in the cell cycle corresponds quantitatively to the pattern of DNA synthesis. Only if the average specific activity of intracellular [3HJdTTP is the same for each of the pulse periods is one justified in directly inferring that the pulse incorporation pattern and synthesis rate pattern are equiva­ lent. Sachsenmaier (1964) included 5-fluorodeoxyuridine and uridine with the radioactive thymidine used to pulse label plas­ modia for 15-min intervals. Plasmodia exposed to 5-fluoro-

HOLT

deoxyuridine and uridine require external thymidine for normal growth and DNA synthesis (Sachsenmaier and Rusch, 1964). How­ ever, one cannot tell from the published data whether or not DNA synthesis would become thymidine-dependent during a 15min interval after addition of the agents. Evans et al. (1976) show that the specific activity reached by dTTP in 15-min pulses with [3H] thymidine varies by more than a factor of two during the S phase. Data such as they have collected could be used to provide a more secure conclusion on the rate of DNA synthesis as a function of time. The timing of DNA synthesis may also be determined by mea­ suring the amount of DNA chemically. Such measurements show that DNA per plasmodium increases mainly during the first third of the cell cycle, as expected from data with radioisotopes (Sachsenmaier and Rusch, 1964; Mohberg and Rusch, 1969). Similarly, Feulgen measurements show that DNA per nucleus increases mainly during the first three hours of the mitotic cycle (Bovey and Ruch, 1972). 3. Termination of S phase. Data such as those shown in Fig. II.4 define an S phase of two to three hours. Although several types of experiments show net chromosomal DNA syn­ thesis during the ensuing G2 phase, this G2 synthesis may be confined to a fraction of the nuclei with aberrant characteristics. Thus, net DNA synthesis in normal nuclei may terminate abruptly. This is not, of course, inconsistent with the conclusion that liga­ tion of DNA fragments occurs during G2 (Brewer et al., 1974; Funderud and Haugli, 1975). Chromosomal DNA synthesis during the G2 period was shown clearly by Hall and Turnock (1976), who incubated prelabeled Plasmodia in nonradioactive medium and followed the decay in specific activity of DNA. During the course of an entire cell cycle, the specific activity dropped by 50 percent, showing the expected doubling of DNA. Most of the decrease (80 percent) occurred during the period from mitosis to 1½ hours after mitosis. But the remaining 20 percent decrease occurred only gradually during the remaining six hours before the next mitosis. In addition Bovey and Ruch (1972), using quantitative microspectrophotometry, report that the average DNA per nucleus in­ creases by about 10 percent during G2. Thus, there is a low, but nonnegligible, rate of DNA synthesis outside the main syn­ thesis period. Nuclei that incorporate [3H]thymidine during G2

NUCLEAR REPLICATION CYCLE

were found to be exceptionally large and frequently to contain several small nucleoli rather than the one nucleolus characteristic of normal nuclei (Guttes and Guttes, 1969). Moreover, results with density shift experiments suggest that chromosomal DNA replicated during G2 does not replicate again during the next cycle (Vogt and Braun, 1977). The G2 synthesis seems to occur, then, in nuclei that are morphologically aberrant and that do not continue to synthesize DNA. The proportion of nuclei exhibiting G2 synthesis may vary markedly from plasmodium to plasmodium. Hall and Turnock's data may be taken to suggest that approxi­ mately 20 percent of the nuclei synthesize DNA out of S phase. On the other hand, early autoradiographic work shows a much smaller fraction of G2 nuclei engaged in DNA synthesis (Braun et al., 1965). It would be useful to apply the autoradiographic, morphological, isotope dilution, and density shift approaches to the same plasmodia, to see if the concept of "aberrant nuclei" will withstand quantitative tests. 4. Temporal sequence of replication. Aplasmodiummaintains its synchrony over many successive nuclear division cycles. This distinguishes it from experimental systems in which a group of cells initially in the same phase of the cell cycle by virtue of selection or blockage gradually lose synchrony in successive cell cycles. The sustained synchrony of the plasmodium was ex­ ploited by Braun et al. (1965) and Braun and Wili (1969) to determine if a DNA sequence replicated in a given subinterval of the S period is replicated in the corresponding subinterval of the subsequent S period. The experiments involve the use of [3H] thymidine to prelabel all the DNA of the cell, of [14C] thymi­ dine to label those molecules replicated in a given portion of a particular S phase, and of bromodeoxyuridine (BUDR) to deter­ mine when in the next S phase the [14C]-labeled DNA serves as a template for a new strand of DNA. The results show the following: DNA synthesized during the first 20 min of a given S phase repli­ cates by the time 40 min of the next S phase have elapsed; DNA synthesized from 30 to 50 min after the start of the given S phase replicates by the time 60 min of the next S period have elapsed; and DNA replicated from 60 to 80 min after the start of the given S phase does not replicate by the time 60 min of the next S phase has elapsed. These are exactly the results predicted if the time for synthesis of a given DNA sequence is the same from one S phase to the next.

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Experiments with other means of characterizing DNA se­ quences support the view that a given sequence has a defined synthesis time in each S phase. Braun and Riiedi-Wili (1971) showed that chromosomal DNA replicated from 10 to 20 min after the start of an S phase has a perceptibly (ca. 0.003 g/cm3) higher buoyant density than the bulk of the DNA. In addition, Fouquet et al. (1974) showed that DNA replicated in the first quarter of S phase is devoid of repetitive DNA. In contrast, DNA replicated in the third and fourth quarters of the S phase dis­ plays renaturation kinetics very similar to that of bulk nuclear DNA. Finally, Fouquet and Sauer (1976) treated DNA labeled during particular subintervals of the S phase with the restriction enzyme EcoRI. The resulting digests were displayed on polyacrylamide gels, and the distribution of radioactivity was mea­ sured. In each case, all, or nearly all, of the labeled DNA frag­ ments migrated as a broad zone on the gel, as would be ex­ pected. The gel patterns also showed that, in some cases, a small percentage of the applied radioactivity was contained in shorter, more defined pieces. Presumably these shorter pieces are derived from repetitive DNA. Shorter fragments were labeled in Plas­ modia exposed to [ 1H ] thymidine for from 30-45 percent and 50-100 percent of the S phase; the labeled shorter fragments were absent in DNA incubated with the labeled precursor for the first 15 percent of the S phase. The results support the con­ clusion that repetitive DNA is not synthesized during early S phase. The results of these anniversary experiments with BUDR, and the demonstration of differences in synthesis time for DNAS of different density and repetitiveness, strongly support the general conclusion that chromosomal DNA is replicated in a defined temporal sequence. 5. Timing of DNA methylation. The Evanses, with S. Littman, have obtained interesting results on the timing of DNA methyla­ tion. Four types of experiments were performed: 1. Surface plasmodia in different phases of the cell cycle were incubated with [methyl-3H] methionine for 1-hour intervals (Evans and Evans, 1970). The amount of 3H incorporated into DNA 5-methylcytosine was approximately the same in all periods of the cell cycle except possibly for an increase by a factor of two during S phase.

NUCLEAR REPLICATION CYCLE

2. The number of 5-methylcytosine molecules completed during a single S phase was measured by precursor-product mea­ surements (Evans et al., 1973). The exogenously supplied radioactive precursor in this case was [methyl-3H]S-adenosylmethionine, the presumed immediate precursor to the modi­ fying methyl group. The precursor was added to cultures in the beginning of the S phase. At four times during the same S phase, samples of cultures were removed for measurements of the specific activity of intracellular S-adenosylmethionine and of radioactivity in DNA 5-methylcytosine. The results showed that, by the end of the S phase, about one molecule of DNA 5-methylcytosine was synthesized per 500 DNA bases. This means that the methyl groups added during one S phase amount to only 1 per 100 cytosine residues, in contrast to the fact that about 1 cytosine in every 24 carries a methyl group. Thus it appears that methylation of newly synthesized DNA strands is not completed immediately. 3. Using [11Cjdeoxycytidine as a precursor, Evans et al. (1973) labeled plasmodia during an S phase, washed away the pre­ cursor, and then measured the ratio of radioactivity in DNA 5-methylcytosine to radioactivity in both forms of DNA cytosine as a function of time after the S phase. The ratio, which began as expected at about 1 percent, continued to rise for about two doubling times, after which it remained constant at about 5 percent. Thus, a portion of the cytosine moieties built into a DNA strand in one S period continues to receive methyl groups for about two doubling times subsequent to their incorporation. The precise kinetics of methyl addition during these two doubling times is not known, but one may estimate that roughly one-third of the methyl groups that are ultimately added to a given DNA strand are added during the S period of its synthesis and that about one-third are added during each of the next two mitotic cycles. This is shown cartoon-fashion in Fig. II.5. 4. The conclusion that methyl groups continue to be added for two doubling times was further tested with density-labeling experiments (Evans et al., 1973). The results show that DNA classified as parental in one round of replication continues to acquire methyl groups for as long as two generations after the round. Labeling in the last generation was minimal, and the results seem compatible with the hypothesis that the labeling

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FIG. II.5. Kinetics of methylation of cytosine residues in chromosomal A strand of DNA, synthesized during the S period designated Si, continues to receive methyl groups for two cell cycle times, when it reaches its maximum methylation of about one methyl group for every 20 cytosine residues. (Data from Evans and Evans, 1970, and Evans et al., 1973; used by permission of the author and publisher.) DNA of growing Plasmodia.

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STRAND SYNTHESIS AND METHYLATION

METHYL GROUP

METHYLATION

METHYLATION

TIME PERIODS OF DNA SYNTHESIS

is nearly complete two doubling times after a strand is syn­ thesized. The physiological role of DNA methylation in eukaryotes is unknown. The information on methylation in P. polycephalum suggests several approachable questions. Is the extent of methyla­ tion the same in amoebae, plasmodia, spores, and spherules? Does the DNA, which in a growing culture lacks all the methyl groups it could conceivably bear, become more completely methylated when growth slows? Are the DNA sequences that are methylated as soon as they are synthesized always the same from one S period to another? Are the methyl groups selectively located in repetitive or unique DNA? 6. Intermediates in DNA synthesis—Okazaki fragments. Several recent studies on chromosomal DNA synthesis in P. polycephalum have been concerned with the· mechanism of synthesis. In prokaryotes, a surprisingly complex picture is emerging from studies on the mechanism of DNA synthesis (Alberts and Sternglanz, 1977). Eukaryotes undoubtedly share in this complexity, but far less is known here. As will be noted below, P. polycephalum has sev­ eral special advantages for further research in this area. One of the two new strands at a replication fork must increase, overall, in a 3' to 5' direction. As no enzyme capable of adding nucleotides to a 5' terminus has been found in any organism,

NUCLEAR REPLICATION CYCLE

it is assumed that synthesis of this "lagging" strand always occurs by 5' to 3' growth of short pieces of DNA initiated at various points along the parental template strand. The assumption is supported by the finding that in both prokaryotes and eukaryotes a substantial fraction of newly incorporated deoxynucleotides are located in short, so-called Okazaki DNA fragments. There is no apparent necessity to postulate the existence of frequent ini­ tiations on the "leading" strand: the leading strand could be synthesized continuously. In P. polycephalum, Brewer (1975) has proposed that chromosomal DNA is synthesized continuously on the leading strand and discontinuously on the lagging strand. Funderud and Haugli (1975), on the other hand, suggest that synthesis is discontinuous on both strands. The data from the two laboratories are not as different as the two interpretations. Funderud and Haugli (1975) have pulse labeled plasmodia in the S phase with [3H] thymidine. All pulses began 30 min after metaphase, and the pulses varied in length from 15 sec to 9 min. The pulses were terminated by immersing plasmodia in liquid nitrogen. (Immersion of plasmodia in ice-cold nuclear isolation medium was ineffective in stopping synthesis completely.) Nuclear DNA was extracted, denatured, and sedimented on alka­ line sucrose gradients. The results, shown in Fig. II.6, reveal the existence of a very slowly sedimenting DNA species (labeled "I") in the cultures pulsed for the shortest times; the species is identifiable in the gradients of DNA labeled for 15, 30, 60, and 90 sec. Gradients of longer-labeled DNA show no such definable species, presumably because of the predominance of label in larger DNA molecules. In a subsequent publication (Funderud and Haugli, 1977b), the authors estimate that species I sediments at 4.5 S, which corresponds to a molecular weight of 6 x 10' d. This early labeled, low-molecular-weight species is an excellent candidate for precursor to at least some of the mature DNA. The molecular weight of the species is in the range for polynucleotides designated Okazaki fragments" in other eukaryotes (Edenberg and Huberman, 1975) and will be so designated here. Waqar and Huberman (1975) subjected a plasmodium in early S phase to a 2-min pulse of [3H] thymidine. The labeled single strands of DNA isolated from the plasmodium sedimented as a broad peak; the maximum of the peak occurred slightly above the position to which 300 nucleotide (IO5 d) marker DNA sedi­ mented. Thus, the Okazaki fragments seen by these investigators

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FIG. II.6. Alkaline sucrose gradient sedimentation profiles of DNA pulse labeled with [3H]thymidine for time periods indicated in the graphs. Values are given as percent of total radioactivity. Fractions are numbered from top to bottom of gradient. Arrow indicates position of [14C]-labeled P2-phage DNA. (From Funderud and Haugli, 1975; reprinted by permission of the author and publisher.) 14

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Fraction number (from top to bottom). are very similar to those seen by Funderud and Haugli (1975). Although the 2-min pulse time is longer than the ones giving distinguishable Okazaki fragments in Funderud and Haugli's experiments, differences in the strains, culture conditions, label­ ing methods, and centrifugation conditions could readily account for the minor discrepancy. Although Brewer and co-workers (Brewer, 1972a, 1975; Brewer et al., 1974; Evans et al., 1975, 1976) have looked only for frag­ ments labeled in 4-min or longer pulses, Brewer (1975) has observed low-molecular-weight fragments from isolated nuclei incubated with [3H]dATP. He suggests that these are indeed

NUCLEAR REPLICATION CYCLE

Okazaki fragments and that they may be observable in nuclei labeled for a relatively long time (45 min) because the joining of the fragments into larger pieces of DNA may be delayed in vitro. The labeled peak is described as sedimenting at 10S; since the peak is near the top of the gradient, the estimated sedimentation coefficient may not be distinguishable from the 4.5 S value assigned to Okazaki fragments by Haugli and co­ workers. In summary, it seems clear that DNA single strands with a molecular weight of 6 X IO4 d are a major early product in DNA synthesis in P. polycephalum. The question whether they are derived from one or both growing strands is not as readily resolved. Brewer (1975) prefers the hypothesis that they arise from only one of the two strands because he sees another labeled peak representing about the same amount of radioactivity as in Okazaki fragments and sedimenting somewhat more rapidly. However, the longer-labeled molecules could have arisen by the joining of Okazaki fragments. Funderud and Haugli favor the notion that both DNA strands result from the ligation of the Okazaki fragments, because after very short pulses such frag­ ments represent the major type of labeled DNA seen on the sucrose gradients. Although their point of view is a reasonable one, the fact that about one-third of the radioactivity in the 15-sec pulse is in a faster sedimenting peak (labeled "V" in Fig. II.6) and on the bottom of the gradient means that one must leave open the possibility of at least two classes of initial precursors. In any case, it is important to bear in mind that the existence of Okazaki fragments on the leading strand would not demonstrate frequent reinitiation. At least under certain conditions in bacterial cells, such fragments probably arise by endonucleolytic attack on a continuously synthesized strand (Tye et al., 1978), and such action might also occur in eukaryotes. 7. RNA primers. Studies with cell-free eukaryotic systems indicate that the initiation of synthesis of a DNA strand requires "priming" by RNA synthesis (Reichard et al., 1974; Tseng and Goulian, 1975). In this scheme, the first event in initiation is the polymerization of a small number of RNA nucleotides on the DNA template; DNA nucleotides are then added, with the first of them covalently attached to the RNA "primer." Definitive evi­ dence for the presence in vivo of RNA nucleotides attached to nascent DNA has been obtained in P. polycephalum (Waqar and

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Huberman, 1975). The experiments were made possible by the ease with which solutions may be injected directly into plasmodia. The experiments involve injection of 32P-Iabeled deoxyribonucleoside triphosphates, such as [a-32P]dGTP, and analysis of the distribution of radioactivity in newly synthesized DNA mole­ cules. The locations of the labeled phosphorus atoms in the region of the RNA-DNA link are illustrated in Fig. II.7. The FIG. II.7. Position of labeled phosphorus atoms (P") in part of an RNA-DNA intermediate from plasmodia injected with [a-32]dGTP (i.e. deoxyguanosine(5')-P*-P-P). One labeled phosphate group is linked to the 3' position of an RNA nucleotide. Base sequence shown is for illustration only. (Based on Waqar and Huberman, 1975; used by permission of the author and publisher.) A

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DNA NUCLEOTIDES

phosphate group connecting the stretch of RNA nucleotides to the DNA molecule is bound in a 3' ester linkage to a ribose and a 5'-ester linkage to a deoxyribose. The phosphate and other 5'dGMP phosphates in the DNA portion of the molecule are radio­ active. When the 32P-Iabeled, recently synthesized nucleic acid molecules were hydrolyzed to 5'-nucleotides, the 32P appeared only in the 5'-deoxyribonucleotide (e.g. 5'-GMP) of the type with which it was originally associated. The results show that the label was not transferred to other precursors (such as dATP or ATP) during the labeling period. When the labeled nucleic acid molecules were treated with alkali, radioactivity was released in 2',3'-ribonucleotides. Thus, the labeled ribonucleotides were adjacent to labeled deoxynucleotides. All four labeled deoxyribonucleoside triphosphates were effective as precursors, and each was incorporated adjacent to all four ribonucleosides; ini­ tiations do not occur, then, at a specific dinucleotide sequence. Label was transferred to ribonucleotides much more effectively from dGTP than from the other triphosphates; this may indicate a preference for initiations at G:C pairs. The observed incor-

NUCLEAR REPLICATION CYCLE

poration occurred only during the S period, thus being almost certainly into chromosomal DNA. The injection seemed to affect the length of RNA strands attached to DNA, as judged by a com­ parison of the buoyant densities of newly synthesized molecules in injected and noninjected plasmodia. The reason for this effect is not known. The Okazaki fragments labeled in the 2-min period following injection were smaller than those formed in a 2-min pulse with externally supplied [3H]thymidine; possibly this is due to different effective pulse times, or to the postulated dif­ ferences in the length of RNA attached to the DNA. Although further experimentation will be required to answer the questions raised by these observations, it appears unlikely that the major conclusion regarding RNA priming will have to be altered. 8. Intermediates in DNA synthesis—later stages. To complete the synthesis of DNA, the Okazaki fragments must be freed of RNA primer, and the gaps between Okazaki fragments must be filled and sealed. In addition, since growing forks are initiated at a number of points along a given DNA molecule, the ligation of adjacent pieces of DNA, themselves the result of the joining of many Okazaki fragments, is required. In general terms, one expects to find that Okazaki fragments labeled at one particular time will find themselves covalently bound to larger and larger DNA molecules. The data in Fig. II.6, as well as data in several other papers (Brewer, 1972; Brewer et al., 1974; Evans et al., 1975; Funderud and Haugli, 1977b), show that the length of labeled DNA molecules continues to increase for at least two hours. The way in which labeled molecules increase in size during a period of chasing will depend on the rate of nucleotide addition, the time required for gap filling and ligation, and the spacing of the primary initiation events along the whole DNA molecule. Interpretation of relevant data will also depend on knowledge of degrees of degradation of DNA during extraction, effectiveness of chases of radioactivity, and other factors. It seems likely that it will be necessary to employ fiber autoradiog­ raphy, as well as sucrose gradients, to obtain a complete picture. An extensive set of pulse and pulse-chase experiments analyzed with alkaline sucrose gradients has recently been conducted (Funderud et al., 1978). The results indicate that replicons have a size of 1.1-2.2 X IO7 d and that the synthesis of the replicons initiated early in S phase is completed after 40-50 min. These replicon-sized pieces of DNA then seem to maintain their size for

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about 30 min, after which they are ligated into pieces three to four times larger. Another abrupt step in ligation occurs about 120 min after the beginning of S phase. 9. Effect of cycloheximide. The drug cycloheximide inhibits DNA synthesis in P. polycephalum, and interest in this phenome­ non has been considerable. The drug, which is an inhibitor of eukaryotic protein synthesis, causes immediate inhibition of amino acid incorporation and cessation of DNA synthesis after a delay (Cummins and Rusch, 1966). An experiment indicating the existence of a whole series of replicons whose staging depends on sequential protein synthesis stirred a great deal of interest (Muldoon et al., 1971). Cycloheximide was shown to act at the ribosomal level by Haugli et al. (1972). Synthesis of proteins in extracts from wild-type plasmodia was strongly inhibited by cycloheximide. Since a mutant resistant to cycloheximide was shown to have ribosomes resistant to cycloheximide, it seems likely that elimination of the effect of the drug on protein syn­ thesis is sufficient to restore normal growth. The effects of cycloheximide on the increase in size of DNA fragments in wild type are not seen in the cycloheximide-resistant mutant; thus, these effects in particular seem to result from the inhibition of protein synthesis (Funderud and Haugli, 1977b). Nevertheless, cycloheximide has now been shown to have several other effects, and these may or may not be sequelae of its effect on ribosomes (Evans et al., 1976; Wendelberger-Schieweg et al., 1978). 10. Cell-free synthesis. Several methods for the cell-free syn­ thesis of chromosomal DNA have been reported for P. polycepha­ lum. In the cell-free systems that have been described fully, intact nuclei are incubated with a radioactive deoxyribonucleoside triphosphate precursor and other factors. The labeled product has been shown to be degraded by DNase (Schiebel and Schneck, 1974; Brewer, 1975) but not by RNase, pronase, or alkali (Schiebel and Schneck, 1974). The properties of an α-type DNA polymerase have been described recently (Baer and Schiebel, 1978). A method for the cell-free synthesis of mitochondrial DNA is also available (Brewer et al., 1967). Nuclei may be prepared from plasmodia in different phases of the cell cycle and used in cell-free incorporation experiments. Such studies have shown that S phase, but not G2 phase, nuclei are active in DNA synthesis. The studies also show that nuclei from different subphases of the S phase differ from one another

NUCLEAR REPLICATION CYCLE

in ( a ) the rate at which they carry out cell-free DNA synthesis, (.b) the degree to which DNA synthesis in them is stimulated by spermine (Brewer and Rusch, 1966), and (c) the size of the labeled DNA they synthesize in a 45-min incubation with [3H] dATP (Brewer, 1975). The fact that synthesis in S phase nuclei greatly exceeds that in G2 phase nuclei indicates both that nor­ mal cellular controls operate in the cell-free system and that the observed incorporation represents normal DNA synthesis. The ease with which one may obtain nuclei from different subphases of the S period is likely to be a special advantage in further studies on cell-free chromosomal DNA synthesis in Physarum. Nuclei isolated from early S phase plasmodia do not support the highest observed incorporation of precursors into DNA. This observation contrasts with the results of thymidine incorporation in intact plasmodia; in these, the maximum rate of incorporation is observed at the onset of the S phase. Nuclei show the greatest cell-free incorporation when isolated from plasmodia 30-60 min (Brewer and Rusch, 1965; Schiebel and Schneck, 1974; Brewer and Ting, 1975; Funderud and Haugli, 1977a) or 90 min (Brewer and Rusch, 1966) into the S phase. The last cited paper shows a marked peak of incorporation at 90 min in nuclei with added DNA and spermine; since the other references concern synthesis unprimed by added DNA, it is tempting to conclude that late-peaking synthesis differs from the synthesis that peaks earlier. However, the Brewer and Rusch (1966) paper also seems to show unprimed synthesis peaking at 90 min; thus, the reason the peak is so late in this work remains obscure. The incorporation of precursors into DNA during a relatively short incubation of isolated nuclei might result from the con­ tinued replication of replicons initiated in vivo; alternatively, the incorporation might reflect anachronous synthesis, the repair of DNA fragmented during nuclear isolation, or other abnormal processes. Schiebel and Schneck (1974) distinguished the two alternatives by the use of a density label. In one experiment, plasmodia were exposed briefly (less than 20 min) to BUDR at the beginning of the S phase. Nuclei were isolated from the plasmodia and incubated with [3H]TTP. At the end of the incubation, DNA was isolated from the nuclei, reduced to a size of 1-3 X IO6 d by sonication, and analyzed on CsCl gradients. Nearly all the bulk DNA was of normal density. The incorporated [3H], on the other hand, was associated with dense (hybrid)

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DNA. The association was complete on neutral gradients. On alkaline gradients, more than half the [3H]-labeled DNA was dense; the alkaline profile was heterogeneous, as would be ex­ pected. Additional experiments show that the cell-free synthesis is semiconservative and that [3H] incorporated into isolated nuclei is not associated with dense DNA strands synthesized in the presence of BUDR some 45 min before the nuclei are iso­ lated and incubated with [3H]TTP. Similar results have been obtained by Funderud and Haugli (1977a). Thus, the DNA syn­ thesis in the cell-free system is a continuation of synthesis begun in the intact plasmodium and not aberrant synthesis induced by the isolation and incubation conditions. The earlier cell-free system used by Brewer and Rusch (1965, 1966), Brewer's later cell-free system (Brewer and Ting, 1975; Brewer, 1975), and the system developed by Schiebel and Schneck (1974) and modified by Funderud and Haugli (1977a) differ from one another in various details. Potential users are advised to study all the papers for relevant variations. Some salient points are as follows. With all the systems, DNA synthesis depends on Mg++ and all four deoxyribonucleoside triphosphates. The chelating agent EGTA stimulates incorporation, presumably by removing Ca++. Spermine, which is inhibitory in systems from other organisms, is stimulatory in P. polycephalum. Triton X-100, which is sometimes used to isolate nuclei, is somewhat inhibitory (Brewer and Ting, 1975). Dithiothreitol reduces incorporation (Brewer and Rusch, 1965); mercaptoethanol is used in one of the systems (Schiebel and Schneck, 1974). The optimal temperature of incubation is about 35°, which is significantly higher than the maximum temperature (31°) for growth of P. polycephalum. Dextran is stimulatory (Brewer, 1975); whether its effect is dif­ ferent from the effect of spermine cannot be inferred from the available results. Brewer (1975) calculates that the initial rate of synthesis in the dextran-containing system is 25 percent of the rate in vivo and that in a two-hour incubation 15 percent of the genome is replicated.

C. Control of Synthesis 1. Relationship between DNA synthesis and mitosis. In numer­ ous experiments, which are reviewed above, the onset of DNA synthesis has been seen to coincide with the end of nuclear mitosis. This coincidence suggests that DNA synthesis is triggered

NUCLEAR REPLICATION CYCLE

by some event in the mitotic sequence. An elegant set of experi­ ments by Guttes and Guttes (1968) confirms this view and indi­ cates in addition that DNA synthesis is autonomous within a nucleus once started. The investigators studied the synthesis of DNA in nuclei transplanted from their original plasmodium into another plasmodium at a different phase of the cell cycle. The methods used in the work are as follows. Pairs of plasmodia in different phases of the cell cycle were allowed to expand into contact with one another. Cell fusion, or "coalescence," followed contact, and the vigorous protoplasmic streaming, which trans­ ports nuclei as well as cytoplasm, began to mix the contents of the plasmodia. The connections between the two plasmodia were severed approximately 15 min after fusion. Each of the resulting plasmodia contained, at separation, a few nuclei from the other. The transplanted nuclei were identified and enumer­ ated either by their appearance in the microscope or by virtue of one of the plasmodia having been labeled with [3HJthymidine before coalescence. The transplanted nuclei represented 1-5 percent of the total nuclei. Labeling with [3H] thymidine auto­ radiography was used to detect DNA synthesis. The results showed: (a) G2 phase nuclei transplanted to an S phase plas­ modium do not begin DNA synthesis during that S phase; (b) the presence of these G2 nuclei and associated cytoplasm does not interfere with the ongoing DNA synthesis in the host nuclei; (c) transplanted G2 nuclei do seem to synthesize DNA after having undergone a synchronous division with the host nuclei; (d) S phase nuclei transplanted to a G2 phase plasmodium continue synthesizing DNA in an apparently normal manner; and (e) the presence of these S phase nuclei and associated cytoplasm does not trigger DNA synthesis in the G2 phase host nuclei. In sum­ mary, it appears that chromosomal DNA synthesis is triggered only by the passage of a nucleus through mitosis and that, once syn­ thesis has started in a nucleus, synthesis runs its course regard­ less of the phase of the cytoplasm in which the nucleus finds itself. The synthesis of DNA in fused plasmodia has also been studied by Wille and Kauffman (1975). They asked a somewhat dif­ ferent question, namely, Can late S cytoplasm stimulate pre­ mature synthesis of late S DNA in early S nuclei? Late S DNA was labeled by exposing plasmodia to [3HJthymidine during the latter part of one S period. The resynthesis of this DNA in the

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next S period was detected by the addition of BUDR and mea­ surement of the amount of labeled DNA shifted to higher buoyant density. For equivalent treatments with BUDR, more DNA seemed to be shifted in early S nuclei transplanted to a late S cytoplasm than in control untransplanted early S nuclei. Thus, the authors conclude that late S cytoplasm contains signals that stimulate premature synthesis of DNA normally synthesized in late S only. The data are consistent with the conclusion, but as the difference in amount of shift between the control and experimental groups is not large, and as the overlap of shifted and unshifted DNA peaks interferes with the assessment of their relative heights, the work should not be regarded as definitive. I review next a series of investigations in which observation of nuclear mitosis, rather than measurement of DNA synthesis, is the principal means of monitoring the progress of plasmodia through the mitotic cycle. The central concern remains, nonetheless, with the question of control. 2. The asynchrony of shaker cultures. When plasmodia are grown in agitated suspension culture, the mold is maintained as small, irregular, but still multinucleate pieces. Guttes et al. (1961) and Guttes and Guttes (1964b) report that these small "microplasmodia" in a single culture vessel are not in mitotic synchrony with one another. Nuclear division is synchronous within a single microplasmodium. These important observations show that whatever is responsible for synchrony in a single Plas­ modium is not transmitted to other nearby plasmodia. I note that the asynchrony of microplasmodia has never been described quantitatively; it is not known, for example, whether the distri­ bution function derived by Puck and Steffen (1963) for asyn­ chronously growing animal cells is applicable. Possibly, microplasmodial cultures will be found to be partially synchronized. In any case, the basic conclusion is not in doubt, and it is clear that one must look inside the cell for the major factors that cause synchrony. 3. Fusion of asynchronous microplasmodia. A number of in­ vestigators have addressed the question, What is the timing of mitosis in a plasmodium constructed by fusing plasmodia in different phases of the cell cycle? The first experiments of this sort involved mass fusions of asynchronous microplasmodia to form individual plasmodia. In such an experiment, the micro­ plasmodia are pipetted onto the surface of a filter paper or a

NUCLEAR REPLICATION CYCLE

nitrocellulose filter, allowed to fuse with one another (for a period up to 1½ hours), and then fed by addition of liquid medium under the paper (see Fig. II.1). Fusion was "followed by a period of 6-7 hours during which no mitosis occurred. . . . At the end of this period, all nuclei divided simultaneously with one another through the following divisions until the Plas­ modium reached a diameter of 5 cm" (Guttes et al., 1961). The nuclei at the first postfusion mitosis, MI, were not uniform. A few were smaller than normal, and some (