Advances in Genetics Volume 37

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Advances in Genetics

Volume 37

This Page Intentionally Left Blank

n

Volume 37

Advances in Genetics Edited by Jeffery C. Hall

Jay C. Dunlap

Department of Biology Brandeis University Waltham, Massachusetts

Department of Biochemistry Dartmouth Medical School Hanover, New Hampshire

Theodore Friedmann

Francesco Giannelli

Department of Pediatrics Center for Molecular Genetics School of Medicine University of California, San Diego La Jolla, California

Division of Medical and Molecular Genetics United Medical and Dental Schools of Guy’s and St. Thomas’ Hospital London Bridge, London, United Kingdom

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Academic Press San Diego

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Copyright 0 1998 by ACADEMIC PRESS 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. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (222 Rosewood Drive, Danvers, Massachusetts 01923). for copying beyond that permitted by Sections 107 or 108 of the US. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-1998 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0065-2660/98 $25.00

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Polytene Chromosomes, Heterochromatin, 1 and Position Effect Variegation I. E Zhimulev 1. General Remarks 1 11. Morphology of the Heterochromatic Regions of the Chromosomes in Dividing Cells 5 111. Repetitive Sequences 43 IV. Genetic Content of Heterochromatic Regions 55 of Mitotic Chromosomes V. Diminution of Chromatin and Chromosomes 78 VI. Centromeric Heterochromatin in Polytene Chromosomes 90 VII. Intercalary Heterochromatin 134 VIII. Telomeric Heterochromatin 238 IX. The B Chromosomes 282 X. Heterochromatin of Chromosomes Restricted to Germline Cells

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XI. Changes in Expression of Genes Dependent on Their Position in the Genome 306 XII. Genetic Inactivation under Position Effect Variegation 309 XIII. Modification of Gene Expression under Position Effect 334 XIV. Time of Genetic Inactivation in Development 355 XV. Unusual Cases of Position Effect 357 XVI. Molecular and Cytogenetic Aspects of Position Effect Variegation

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XVII. Current Concepts of the Mechanism of Position Effect 422 Variegation XVIII. Heterochromatization of Chromosome Regions and Regulation of Gene Activity 435 References

Index

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V


Polytene Chromosomes, Heterochromatin, and Position Effect Variegation 1. F. Zhimulev Institute of Cytology and Genetics Siberian Division of the Russian Academy of Sciences, Novosibirsk 630090 Russia

1. GENERAL REMARKS A. Foreword Problems relating to the organization of heterochromatin are prominent in our general thinking about the organization of genetic material in chromosomes. Although investigators continue to focus attention on these problems, there is, as yet, no comprehensive account of all the facts. It is little wonder that Shah et al. (1973) mention Pontecorvo’s statement made 15 years earlier: “studies on heterochromatin are at the prescientific level . . . and we have no other alternative as to ignore it” (p. 467). In recent years, it has been demonstrated that the features of DNA primary structure, in particular the repetitiveness of nucleotide sequences, underlie heterochromatin structure. Quite reasonably, John and Miklos (1979) commence their review with Walker’s epigraph that we know so much about the structure, variability, and localization of satellite DNA that it becomes all the more remarkable that we know nothing about the origin and functions of these specific DNA sequences (p. 1). In this monograph various aspects of heterochromatin organization in the mitotic and interphase polytene chromosomes are considered. As a rule, a multitude of details pertaining to the organization of heterochromatin at the high resolution achievable with the use of highly decondensed chromosomes are analyzed. The effect of heterochromatin on fragments of euchromatic regions transposed to close proximity under position effect is treated. The book is the second authored by Dr. Zhimulev concerned with polyAdvances in Genetics, Vol. 37 Copyright 0 1998 by Academic Press. All rights of reproduction in any form reserved. 0065-2660198 $25.00

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tene chromosomes. The first volume, “Morphology and Structure of Polytene Chromosomes” (Zhimulev, 1996), has been published as Volume 34 of Advances in Genetics; in addition a series of short versions of these reviews have been published in Russian (Zhimulev, 199213, 1993, 1994). The author is faithful to his style: detailed accounts are combined with extensive references to the treated topics. The accounts are restricted to analysis of facts obtained with animal and plant species possessing polytene chromosomes. This is done so as to make possible comparisons of the results obtained with the compact mitotic and the decondensed interphase (polytene) chromosomes.

B. Introduction Our body of knowledge of heterochromatin was gleaned unhurriedly through a protracted course of time. In the past century, cytologists were aware of the existence of various heavily staining bodies appearing as clumps, rods, or drops in the interphase nuclei (for a more detailed historical account, see Prokofyeva-Belgovskaya, 1986, p. 15-19). In the 1920s and 1930s, having studied the behavior of chromosomes during the cell cycle, E. Heitz demonstrated that these enigmatic bodies are either specific chromosomes or chromosome regions retaining a compact state and, therefore, stronger stainability during the entire cell cycle. Starting from the formation of the nucleus in telophase, they remain compact until the beginning of the successive division, they then decondense for a brief time and assume anew (earlier than the rest of chromosome material) the compact state. By analogy with the accepted term “heterochromosome,” referring to the sex chromosomes, which retained increased compaction and stainability throughout the cell cycle in some insect species, the concept of heterochromatin was suggested for these bodies. The remaining chromatin was called euchromatin, the condensation-decondensationcycle of which underlies mitotic division (Heitz, 1928,1929,1930, 1932, 1933a,b, 1934, 1935). Comparisons of the properties of heterochromatin in mitotic and polytene chromosomes led Heitz to the conclusion that there exist two types of heterochromatin that differ in degree of compaction: a-heterochromatin, the more compact, heavily staining and, presumably, the less decondensed in the mitotic cycle; and P-heterochromatin, the more diffuse, loose, and weakly staining. At first, Heitz made a distinction between heterochromatin and euchromatin relying on the criterion of degree of compaction, or, as he put it, the degree of heteropycnosis. He also suggested that the heterochromatic regions are genetically inert because loss of these regions or decrease in their size was inconsequential. Furthermore, according to the concepts accepted in those years, the chromosomes are “genetically active” only in the nucleus itself, not at mitosis, and, hence, the heterochromatic regions resembling mitotic chromosomes in compaction degree should, in his view, be “genetically inert.” This prediction was

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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brilliantly confirmed by Muller and Painter (1932), who have demonstrated that almost all the then-known genes residing on the X chromosome are located only in its euchromatic portion. Subsequently, it was shown that genes are, nevertheless, present in heterochromatin, although with an occurrence frequency lower by two orders of magnitude than in euchromatin (see reviews in Hilliker and Sharp, 1988; Gatti and Pimpinelli, 1992; Zhimulev, 1993). There followed a continual discovery of new properties ofheterochromatin. In 1959, it became apparent that heterochromatin replicates at the end of the S phase (“late replication”) (Lima-de-Faria, 1959a,b; for review, see Lima-de-Faria and Jaworska, 1968). There is considerably less heterochromatin in the polytene than the mitotic chromosomes (Painter, 1933). This was found to be due to incomplete poly’ tenization of a part of the chromatids constituting heterochromatin during the formation of polytene chromosomes (Rudkin, 1965a, 1969). A property of heterochromatin, such as its “fragility,” may possibly be explained by underpolytenization: the chromosomes are most frequently ruptured in regions of centromeric heterochromatin during the making of squash chromosome preparations. Another property, higher concentration of chromosomal rearrangements, has also been related to heterochromatin (Kaufmann, 1939; Prokofyeva-Belgovskaya and Khvostova, 1939; Mukhina et al., 1981). The finding of repetitive DNA sequences in the eukaryotic genome (Britten and Kohne, 1968)and the method of in situ hybridization (Gall and Pardue, 1969) provided a broader basis for an understanding of heterochromatin structure and detection of its richness in repetitive DNA sequences (Lohe and Roberts, 1988; Lohe et al., 1993). Numerous regions intercalated into euchromatin and showing all the properties of heterochromatin were identified in polytene chromosomes: the socalled intercalary heterochromatin. According to the view taken by Ananiev et al. (1978; Gvozdev, 1981a,b), it consists of material of mobile genetic elements, it is the view of Zhimulev et al. (1982) that it is composed of tandem repeats. The existence of intercalary heterochromatin was even in doubt (Spofford, 1976; Hilliker and Sharp, 1988; Gatti and Pimpinelli, 1992). Specific molecular genetic structures, the telomeres, were detected in the interphase polytene chromosomes at the chromosomes’ tips (Muller, 1932, 1941; Rubin, 1978) that possess all the morphological features of heterochromatin. Many reviews are concerned with the properties of heterochromatin; the following deserve mention: Frolova (1934), Gershenzon (1940), Schultz (1941b, 1943a, 1947, 1956, 1965), Pontecorvo (1944), Prokofyeva-Belgovskaya (1945, 1947, 1977a,b, 1981, 1986), Resende (1949, Barigozzi (1950), Hannah (1951), Alfert (1954, Cooper (1959), Beermann (1962), Rudkin (1965b,1972), Brown (1966), Yunis and Yasmineh (1971), Berendes (1973), Shah et al. (1973), White (1973), Sandler (1975), Back (1976), Spofford (1976), John and Miklos (1979,

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1988), Smirnov and Smaragdov (19791, Stahl and Hartung (1981), Rocchi (1982), Babu and Verma (1987), Hilliker and Sharp (1988),John (1988), Verma (1988), Pardue and Hennig (1990), Belyaeva and Zhimulev (1991), Dryanovska etal. (1991), Bataninetal. (19921,GattiandPimpinelli(19921, Zhimulev (1992a, 1993), Gvozdev (1993), Cook and Karpen (1994), Eissenberg et al. (1995) Lohe and Hilliker (19959, Weiler and Wakimoto (1995), Zacharias (1995), Zuckerkandl and Hennig (1995), and Elgin (1996). A remarkable experimental model for studying the physiological role of heterochromatin is position effect. When a euchromatic region is transposed to heterochromatin, the genes immediately adjacent to it can become inactivated under position effect (Muller, 1930). The degree of genetic inactivation can be modified by various agents, including variation in heterochromatin amount in the cell. Thus, by taking advantage of position effect, the influence of gene activity in both trans and cis positions can be clarified (Belgovsky, 1944;Lewis, 1950; Hannah, 1951; Baker, 1968; Zakharov, 1968; Birstein, 1976; Spofford, 1976; Becker, 1978;Lima-de-Faria, 1986; Babu and Verma, 1987; Eissenberg, 1989; Tartof et al., 1989; Spradling and Karpen, 1990; Henikoff, 1992; Reuter and Spierer, 1992; Zhimulev, 1992a, 1993; Henikoff et al., 1993; Spradling, 1993; Karpen, 1994; Eissenbergetal., 1995;Elgin, 1996). It was Noujdin’s (1947) opinion that, withstudies on position effect, “the time, when heterochromatic (inert) regions of the chromosome were regarded as inactive regions, not playing any role in heredity and development, was left far behind. Research on mosaicism established the exclusive significance of heterochromatic material in the processes of intracellular metabolism, and, concomitantly, in hereditary changes” (p. 192). Studies on the morphological and morphofunctional organization of chromosomefragments in close proximity to heterochromatin demonstrated that genetic inactivation is related to compaction of the chromosome region (Schultz, 1965; Zhimulev et al., 1986; Belyaeva and Zhimulev, 1991) and to the acquisition of the properties of a-heterochromatin. However, the manifestation of the properties of heterochromatin, at least the intercalary type, is modified by the same factors that affect degree of compaction of euchromatin under position effect. This may be evidence for the close similarity between the DNA inactivation mechanisms in heterochromatin and adjacent euchromatin (Zhimulev et al., 1989a,b). A major breakthrough in studies of heterochromatin and position effect came with the discovery of the complex gene system affecting the expression of genetic inactivation and, consequently, the compaction degree of chromatin (Spofford, 1967; Reuter and Wolff, 1981; Sinclair et al., 1983; Eissenberg, 1989; Tartoff et al., 1989; Reuter et al., 1990). Modifiers show dose transitions: a gene can suppress genetic activation in one dose, give rise to the normal phenotype in two doses, and enhance the normal phenotype in three doses. This becomes comprehensible in view of the remarkable consequences of the action of the position


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effect modifier genes on the structure of chromatin proteins. To illustrate, one of these genes affects the acetylation of histones (Dorn et al., 1986),and the product of another gene is a protein contained by heterochromatin and, presumably, providing its compaction (Eissenberg e t al., 1987; James et al., 1989). These data may provide evidence for universality of the mechanisms providing compaction of chromosome material in eu- and heterochromatin, and pose in a new light the problem of what constitutes eu- and heterochromatin. White (1945) holds the view that heterochromatin may be any region of a chromosome that has become heteropycnotic at certain stages of the cell cycle. In contrast, Schultz (1947) defined heterochromatin as chromosome regions that have the specific property of remaining in compact blocks in the intermitotic state. The more profound concept of heterochromatin as both a substance (constitutive heterochromatin) and a condition (facultative heterochromatin) suggested by Brown (1966)was strongly criticized by Prokofieva-Belgovskaya (1977a, 1986). In the meantime, the acquisition of the features of heterochromatin by one of the female X chromosomes has remained beyond doubt (Lyon, 1961). From this brief outline, it is apparent that new information calling for closer scrutiny has accumulated in the research area of heterochromatin.

II. MORPHOLOGY OF THE HETEROCHROMATIC REGIONS OF THE CHROMOSOMES IN DIVIDING CELLS A. Change in compaction degree during the cell cycle It was known in the beginning of this century that particular chromosomes or their fragments appear more condensed and deeply stained than the rest of the chromosome material during the cell cycle. This differential condition was called heteropycnosis (Gutherz, 1907). Heteropycnosis is negative when staining is light, and it is positive when staining is heavy. Heitz (1928) coined the term “heterochromatin” to draw a distinction between the chromosome regions that show positive heteropycnosis at certain stages of the mitotic cycle and the rest of the chromosome material passing through usual compaction-decompaction cycles of cell division, called euchromatin. In the majority of eukaryotic species, the chromosomes contain both euand heterochromatic regions, with the latter constituting, as a rule, the bulk of the genome. Thus, in Drosophila mekmoguster, the male Y chromosome is almost completely heteropycnotic, whereas heterochromatin makes up to 40-50% of the length of the X chromosome (Hannah, 1951; Ananiev et al., 1973) and 29% of that of the third chromosome. Presumably, the fourth chromosome is almost completely heterochromatic. The total percentage of heterochromatin in the karyotype is 33% of chromosome length (Ananiev et al., 1973; Gatti e t al., 1976).

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In several species, this percentage is rather high, being 30% for Dosophila simulans, 52% for D. uirilis, 47% for D. texanu, 30% for D. hydei, 33% for D. ezoana (Gatti et al., 1976; Pimpinelli et al., 1976), 40% for D. na~uta(Lakhotia and Kumar, 1978), 45% for D. grimshawi, and 55-60% for D. cyrtoloma (Clayton, 1985). Concepts of changes in the compaction condition of the heterochromatic regions during the cell cycle, the major feature of heterochromatin accompanying its description, are rather controversial. According to the data of Heitz and other authors, starting from the early prophase, the heterochromatic regions of the chromosomes become readily identified and different in staining from the euchromatic regions (Figure 1).These differences disappear by the end of the rnetaphase. At the next interphase, the heterochromatic regions are represented by numerous heavily staining grains or large blocks of heteropycnotic material (see Figure I), which have long been termed “chromocenters” (Heitz, 1928, 1929, 1930, 1932, 1933a, 193313, 1934, 1935; Kaufmann, 1934; Frolova, 1934; Dobzhansky, 1944). In Glossina austeni, differences in the compaction rate of the usual and supernumerary chromosomes (or the S chromosome, see Section IX), rich in heterochromatin, were detected. The S and Y chromosomes are condensed as early as the beginning of early prophase, and they appear as condensed, heavily staining bodies at the middle of the prophase (Southern et al., 1972). At meiosis, heterochromatic regions become conspicuous as early as the stage of leptotene (Huettner, 1930). Schultz (1947) was thus led to formulate the concept of heterochomatin: chromosome regions that possess the specific property of retaining a blocklike appearance at the intermitotic stage. Euchromatin and heterochromatin differ in cycles of compaction: while the former passes through the entire cycle of compaction-decompaction from interphase, the latter remains in a comparatively compacted state. These differences have long been called allocycly (Darlington and La Cour, 1940). The compaction degree of the packaging of heterochromatin at a particular stage of the cell cycle may be considerable; thus, for example, the integrity and general morphology of the chromocenters in mouse liver cells are retained during preparative isolation involving procedures such as treatment with DNase I1 and ultrasound (Stephanova and Pashev, 1988). However, “permanence” of tight packaging of material in the heterochromatic regions of the chromosomes is relative:

1. Kaufmann (1934) did not detect heavy staining of heterochromatin of the autosomes at the prophase stage, while the heterochromatin of the X and Y chromosomes appearing as heteropycnotic blocks differs in the neuroblast cells in interphase. Permanent heteropycnosis of heterochromatic elements also was not identified in gonial cells (Cooper, 1959). 2. Heteropycnotic regions are not identified in the interphase nuclei of embryos at any of the cleavage stages (see Section 11,G).

Figure 1. Heterochromatin in Drosophila in the early (A, 8,E-G), middle (C, D, H-J, L), and late (K) prophase and metaphase ( M a ) of mitosis, and heterochromatic blocks in interphase (P-R). (A-D) after Kaufmann (1934); (E-L) after Cooper (1959); (M-R) after Heitz (1933a).

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3. In the interphase nuclei of Drosophikz, the number of chromocenters varies from 0 to 5. Their absence may be evidence for complete decompaction of heterochromatin at any of the interphase stages, possibly at the end of the S phase, when chromatin DNA replicates (Smimov and Smaragdov, 1979).

4. Measurements of the lengths of the mitotic chromosomes of Drosophila at different stages of decompaction have demonstrated that the ability of the chromosome to shorten diminishes with increased heterochromatin content in it (Figure 2). Measurements of the total length of heterochromatin in all the chromosomes demonstrated that they vary in the range of 4.7-7.2 pm, while the absolute length of the haploid set lies in the range of 13.5-67 pm (Ananiev et at., 1973). These data support first the concept of the considerable compaction of heterochromatin at the early stages of mitosis and, second, that of impermanent compaction of heterochromatin during the cell cycle. 5. Various factors affecting compaction degree have been identified. These include various DNA ligands binding to the AT-rich regions of double-strand-

-.-.

-*-----------.

d . .

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so il 7a Figure 2. Correlation between changes in the absolute length of the mitotic chromosomes (ordinate) and the total length of the haploid set (abscissa)of the chromosomesof Dosophila neuroblasts. Numbers and letters designate the third, the second, the X, the Y, and the fourth chromosomes.After Ananiev et al. (1973). Il,

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ed DNA (distamycin A, Hoechst 33258, DAPI I), treatment with these ligands in interphase leads to inhibition of compaction at the next prophase.

1. Factors affecting compaction degree All data show that the heterochromatic fragments of chromosomes during the cell cycle appear more condensed than the rest of the chromosome material. Numerous factors can change the degree of compaction.

a. Hoechst 33258 Treatment of Drosophila cells at the subsequent mitotic cycle revealed a delay in compaction (Figure 3 ) of various heterochromatic regions (Pimpinelli et al., 1975; Gatti et al., 1976; Lakhotia and Roy, 1981; Roy et al., 1982; Felcher et al., 1982; Smirnov, 1984; Smirnov et al., 1986). The pattern of compaction delay is species specific, for example, in D. mehnoguster, the distal region of heterochromatin of chromosome 3L and the centromeric block of the Y chromosome, as well as the proximal part of the heterochromatin of the X and Y chromosomes, are the most decompacted. In D. simulans, heterochromatin is almost completely decompacted (Gatti et al., 1976). According to other data, the frequency of compaction delay in D. melamgmer was the highest in the X chromosome, the Y chromosome ranked next, and chromosome 3L was the last (Felcher et al., 1982; Smirnov et al., 1986).To compare, compaction delay was identified in metaphases at frequencies of 48% in D. melanoguster and 2.5% in D. simulans, and it was not identified in the third related species, D. mauritiunu (Smirnov et al., 1986). Only a certain part of heterochomatin is subject to the decompacting effect of the Hoechst stain-for example, only 13% of metaphases of the third chromosome of D. melanoguster. This allowed Felcher et al. (1982) to make the conclusion that “the relation between heterochromatin in interphase and mitosis is much more complicated than appeared to be within the framework of the concept of permanent compaction of heterochromatin during the cell cycle” (p. 1144). As suggested by Felcher et ul., interphase chromocenters do not represent the entire heterochromatin identified in mitosis. The composition of the karyotype affects the frequency of compaction delay produced by the Hoechst 33258 stain. In D. melanoguster, the occurrence frequency of abnormal metaphases is the highest in XX and very low in XO males (Felcher et al., 1982, Smirnov et al., 1986). When the embryonic cells of D. nusuta, were treated with Hoechst in conjunction with 5-bromodeoxyuridine, compaction was inhibited in fewer cells, and heterochromatin was more decompacted in those in which it was. However, treatment of neuroblast cells with 5-bromodeoxyuridine neutralizes the effect of the Hoechst stain (Lakhotia and Roy, 1981).

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When cells are sequentially treated with hypotonic saline and the Hoechst stain, or with other DNA-specific ligands such as distamycin A, netropsin, and olivomycin, interphase heterochromatin sharply condenses (Figure 4). It is known that the Hoechst stain preferentially binds to the AT-rich regions of DNA (see Section 11,B). Delay in decompaction attributable to this agent was detected only in species that have AT-rich satellites (D. melanogaster, D. simulans, D.uin'lis, D.texana). In D. hydei and D. ezoana, weak decompaction was revealed only in the Y chromosome (Pimpinelli et al., 1975; Gatti et al., 1976;Felcher et al., 1982; Smimov et al., 1986). It was suggested that during the S phase the Hoechst stain preferentially binds to the specific regions of the chromosomes interacting with the proteins that determine the mitotic compaction of chromatin (Gatti et al., 1976).

b. Distamycin A In four species of Drosophila, the pattern of compaction delay of the chromosomes from embryonic and neuroblast cells produced by treatment with distamycin A correlated with the distribution of AT-rich regions (Q-bands, see Section II,B) and satellite DNA. The correlation was most clear cut for D. uirilis, in whose chromosomes compaction was incomplete both in the Q+ and Q+ regions. In D. melanogaster, the Q++ region was more frequently and the Q + region was less frequently decompacted; in D. hycki and D. funebns, the correlation between compaction delay and Q+staining was less distinct (Faccio Dolfini and Bonifazio Razzini, 1983; Faccio Dolfini, 1987). +

c. Low temperature It was shown in the 1940s that certain regions of the mitotic chromosomes of plants and amphibia, which appear euchromatic at normal temperature, become heterochromatic when temperature is lowered (Darlington and La Cour, 1940; Callan, 1942). After exposure of the larvae of Drosophila to low temperature (6°C for 24 hr), the metaphase chromosomes sharply shorten while the regions of centromeric Figure 3. Changes in the compaction state of heterochromatin in D. melanogaster under ( A ) the effect of low temperature, (RE)treatment of living material with Hoechst 33258, and (F-K) the mw-lOl" mutation. (A) Karyotype of a female; arrow indicates the decompacted regions of heterochromatin of the X chromosomes. (B and C) Controls: (B) karyotype of a male; (C) karyotype of a female. (D) Karyotype of a male after 4 hr of treatment. Arrows indicate the decompacted regions of an arm of the third chromosome, arrowheads indicate the decompacted regions of the X chromosome, and letters and numbers designate chromosomes. (F) Control (18°C). (G-K) At 29°C; arrows indicate the decompacted regions of heterochromatin in the Y chromosome (G-I), in one of the autosomes (J), and in the X chromosome (K). (A) after Brosseau (1967); ( R E ) after Pimpinelli et nl. (1975); (F-K) after Gatti ec al. (1983).

Figure 4. Changes in the compaction state of heterochromatin under the effect of irradiation with y-rays in Akodon molinae (Rodentia, Cricetidae) (a) and combined treatment with hypotonic saline and Hoechst 33258 in D. nmuta (b-f). (a) Arrow indicates a decompacted chromosome. (b) Control interphase nuclei with large loose chromocenters. (c) Treatment with Hoechst for 4 hr. (d) Treatment with hypotonic saline. (e and f) Compact chromocenters after treatment with hypotonic saline and Hoechst. (a) after Bianchi (1982); (b-f) after Lakhotia and Roy (1981). 12

Polylene Chromosomes, Helerochromatin, and Position Effect Variegation

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heterochromatin remain decompacted (see Figure 3). T h e Occurrence frequency of metaphases with such chromosomes was high, although not 100% (Brosseau,

1967).

d. Anoxia In embryos of Drosophila (at the blastoderm stage) deprived of oxygen, development is rapidly arrested. The interphase chromosomes are compacted, each becoming visible during the arrest (Foe and Alberts, 1985). e. Radiation effect In y-irradiated mammalian cells, single compact chromosomes are identified in 0.33% of metaphase spreads, while the rest of the chromosomes of the set are typically metaphase (see Figure 4).Three hypotheses have been advanced to account for this phenomenon: (1 ) numerous breaks may arise in the DNA molecule during irradiation; (2) the protein scaffold, which is possibly involved in the process of chromosome compaction, may be impaired; and (3) mutations may arise at loci controlling compaction. If the latter hypothesis is correct, the existence of such loci in every chromosome should be accepted (Bianchi, 1982).

f. Genetic control of compaction In homozygotes for the lethal temperature-sensitive mutation mapped to the X chromosome of Drosophila, ms-l Olrsl, all the mitotic chromosomes are normal at 18°C. However, after exposure to high temperature (29°C) for 2 hr, the Y chromosome becomes longer and, in many cases, elongated stretches appear in twothirds of metaphase spreads (see Figure 3). In a small number of cells, incomplete compaction of heterochromatin is revealed in autosomes. After longer exposure, heterochromatin is incompletely compacted in the Y chromosome in almost all the spreads and in all the chromosomes in some spreads. The patterns of compaction delay produced by mutation or treatment with the Hoechst stain are diverse (Gatti et al., 1983). In l(2)gl mutants, compaction of mitotic chromosomes varies in a very wide range from superdecompacted to supershort (Radhakrisnan and Sinha, 1987).

2. Mechanisms of compaction Little is known about the molecular mechanisms of heterochromatin compaction. Muller and Prokofyeva (1935, p. 658) believed that a heterochromatic region “does not coil into a spiral, when the chromosome undergoes changes preparative to mitosis. . . . The chromonema in this region should be, for this reason, amidst a great mass of accessory (non-genic) chromatic material, which makes it about as thick as the coiled into spiral” euchromatic region of the chromosome. T h e view that accumulation of “accessory material” remaining in the heterochromat-

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ic regions is possible even during mitotic compaction of the chromosomes was shared by Cooper (1959). Muller and Prokofyeva ( 1935) believed that, insofar as heterochromatin occupies only a small portion of the polytene chromosomes, there should not be much “genic material” in the mitotic chromosomes either (underreplication of heterochromatic DNA in polytene chromosomes was then not yet established) (see Section VI,C). From their point of view, the comparatively voluminous heterochromatic fragment of the mitotic chromosome can be accounted for by accumulation of accessory material. However, calculations demonstrate that only satellite DNA fills to a great extent the heterochromatic regions of the chromosomes of Drosophila: 77,67, 54,22, and 79% (Smirnov, 1984). What role proteins play in compaction is unclear. Large amounts of nonhistone proteins were not detected in heterochromatin of the embryonic cells of D. virilis (Comings et al., 1977). However, there are suggestions that nonhistone proteins may also be involved in compaction (Burkholder and Weaver, 1977; Burkholder and Duczek, 1982).

8. Differential staining

1. C-staining In 1970 Pardue and Gall (1970) noted that centromeric heterochromatin in mouse chromosomes stained more heavily with Giemsa after DNA denaturation-renaturation in in situ hybridization. Hsu and Arrighi (1971; Arrighi and Hsu, 1971; Hsu, 1973) suggested a technique for treating chromosome preparations permitting differential staining of eu- and heterochromatin. The main steps of the procedure included denaturation of chromosomal DNA by treatment with 0.07M NaOH, renaturation (65”C), and staining with a Giemsa solution. Having thus treated chromosome preparations of 20 mammalian species, they demonstrated that the regions of centromeric heterochromatin more densely stained by the conventional methods also stained heavily with Giemsa, whereas euchromatin remained lightly stained. The investigators called the procedure of staining for constitutive (C) heterochromatin the C-staining technique, or C-banding. The technique has since been modified (Sumner, 1978). C-heterochromatin has been described in dipteran species of the Calliphoridae (Ullerich, 1976, Bedo, 1980, 1991), Cecidomyiidae (Bregman, 19751, Chironomidae (Hagele and Speier, 1988), and Culicidae families (Motara and Rai, 1977, 1978); in D. funebns (Faccio Dolfini, 1987); in D. hydei (Pimpinelli et al., 1976; Beck and Srdic, 1979; Faccio Dolfini, 1987); in D. melanogaster (Hsu, 1971;Pimpinelliet al., 1976; Halfer and Barigozzi, 1977; Faccio Dolfini and Halfer, 1978; Sved and Verlin, 1980; Faccio Dolfini, 1987; Vlassova et al., 1991a,b); in the species of the rnelanoguster group (Lemeunier et al., 1978); in D. m u t a


15

Figure 5. Localization ofeu- and heterochromatin in Drosophifadetected by the criterion ofcompaction (a) and according to the results ofC-banding (band c). Numbers and letters designate chromosomes. Karyotypes: (b) female; ( c )male. Scale is 5 pm. a after Heitz ( 1933b); band c after Faccio Dolfini (1974).

(Wheeler and Altenburg, 1977; Lakhotia and Kumar, 1978); in D. simuhns (Pimpinelli et al., 1976); in the species of the virilis group (Pimpinelli et al., 1976; Faccio Dolfini, 1987); in Hawaiian Drosophih (Clayton, 1985); and in species of the Glossinidae (Davies and Southern, 1976), Sarcophagidae (Kaul et al., 1978, 1989a; Samols and Swift, 1979a), Sciaridae (Eastman et al., 1980), and Tephritidae (Bedo, 1986, 1989) families. There is, as a rule, a good agreement between the location of heterochromatic regions identified by allocycly of compaction (staining with orcein) and C-banding in all the studied species (Figure 5). The mechanism of C-staining is unclear. In the majority of cases, the position of the C-stained regions of the chromosomes is coincident with the location of satellite DNA (see Section 111). There are very important exceptions, for example, the Y chromosome of D. hydei does not contain highly repetitive DNA (Hennig, 1973), yet shows distinct C-staining. Similar inconsistencies were found for the chromosomes of several other species (see Pimpinelli et al., 1976).

2. Q- and H- staining In 1969, in Caspersson’s laboratory, a method of differential staining of chromosomes based on preferential banding of various fluorochromes to chromosomal

16

I. F. Zhirnulev

DNA was developed (Caspersson et al., 1969). Some of the fluorochromes proved to be very effective in identification of heterochromatin. The first fluorochrome used was quinacrine, hence the name Q-staining (Figure 6). In the chromosomes treated with the reagent, fluorescence was revealed in the region of centromeric heterochromatin (see Figure 6)--or not revealed, as, for example, in D. ananassae (Adkisson et al., 1971). Q-staining varies considerably along the length of the metaphase chromosome (Figure 7). Regions that fluoresce brightest with quinacrine have been tentatively designated as Q++,the regions that do not fluoresce as Q--, and the intermediate variants as Q' and Q-. It is believed that quinacrine intercalates in the DNA without any specificity to base pairs; however, fluorescence is detected mainly in the AT-rich regions, because the neighboring GC-rich DNA quenches fluorescence. The following facts support the concept of the specificity of fluorescence in the AT-rich regions:

1. In Samoaia konensis (Drosophilidae), the content of AT pairs is highest in the regions that fluoresce brightly after staining with quinacrine (Ellison and Barr, 1972a,b).

Figure 6. Staining of the male (a and b) and female (c and d) chromosomes of Drosophila with the fluorochrome quinacrine. Numbers and letters designate chromosomes. (a and c) Staining with quinacrine. (b and d) Staining with orcein. After Faccio Dolfini (1976).


17

Figure 7. Fluorescence intensity levels ( b and c ) of the regions of the Y (left) and the second (right) (a) chromosomes of Drosophila. Abscissa, chromosome regions; ordinate, fluorescence intensity; Q+,Q-, and Q--, different fluorescence levels. After Gatti et al. (1976).

a++,

2. In DrosophiIa of the vin'lis group, six types of satellite DNA are detected in the genomes, three large (designated as I, 11, and 111 in Figure 8) and three small, the latter constituting together less than 5% of the genome (designated according to their buoyant density values in Figure 8). When large ATrich satellites are present in the genome, heterochromatin in the metaphase chromosomes and the chromocenters in interphase show heavy staining (see Figure 8). Species that have no satellites do not show Q-staining (Holmquist, 1975a). 3. A similar correlation between the presence of AT-rich satellites and H- or Q-banding of the chromosomes was detected in many species of Drosophila (Gatti et al., 1976; Bonaccorsi and Lohe, 1991; Lohe et al., 1993). 4. With the use of various polynucleotides, it was demonstrated that preparations of poly(dA-dT), poly(dA) x poly(dC), or AT-rich DNA satellites showed fluorescence. However, these single-stranded polynucleotides, as well as poly(dC), poly(rC), and GC-rich satellites, were not fluorescent (Weisblum and de Haseth, 1972, 1973; Comings et al., 1975; Comings and Drets, 1976; Moutschen, 1976; Comings, 1978). 5. In studies of transcription in vino (an Escherichia coli EWA-polymerase system was applied to fixed polytene chromosomes) in the presence of actinomycin

18

1. F. Zhimulev Satellite I

D . virilis

mo~itunu

U

IU

Metaphase

Interphase

1.721 1.712 1676

tt tt tt

-

-

-

tt

-

-

t

-

+

-

- -

-

+ - sQ5 .35

D. virilis Satellites Sequence 1 5' ACAAACT II 5' ATAAACT III 5' ACAAATT

':,Genome 25% 8% 8% 4 1 010

Metaphase chromosomes Stain ",,Genome Q-H27.5% Q*H* 12.5'10 O*H5 - 8 "10 4 5 -48%

Figure 8. Relation between satellite DNAs and heterochromatin in closely related species of Drosophila of the virilis group. (Left) An evolutionary tree of species of the virilis group. (Center) Presence ( + +, + 1 or absence (-) of various satellite DNA in species. (Right) Amount of heterochromatin and degree of its fluorescence in metaphase chromosomes and interphase nuclei. The amount of heterochromatin in Drosophila virilis is taken as unit. aI,proximal, brightly fluorescent block of heterochromatin; a*,distal, nonfluorescent block of heterochromatin. After Holmquisit (1975a).

D binding to GC pairs, 3H-ATPwas shown to incorporate into the Q' regions only (Leibovitch et al., 1974). Heterochromatic regions that fluoresce after staining with quinacrine were detected in many dipteran species of the Calliphoridae (Bedo, 1980, 1991)) Cecidomyiidae (Bregman, 1975)) and Culicidae families (Diaz and Lewis, 1975; Steiniger and Mukherjee, 1975; Tiepolo et al., 1975); in D. melanoguster (Becker, 1970; Vosa, 1970; Adkisson et al., 1971; Ellison and Barr, 1971a; Lewis and Craymer, 1971; Zuffardi et al., 1971; Faccio Dolfini, 1974, 1976; Gatti et al., 1976; Barigozzi et al., 1977;Faccio Dolfini and Halfer, 1978;Gatti and Pimpinelli, 1983); in D. simulans (Adkisson et al., 1971; Barr and Ellison, 1971b; Ellison and Barr, 1971a; Gatti et al., 1976); in the species of the melanogaster group (Lemeunier et al., 1978); in the species of the virilis group (Barr and Ellison, 1971b; Holmquist, 1975a;Gatti etal., 1976;Abrahamet al., 1983); in other species of Drosophila (Barr


19

and Ellison, 1971b; Ellison and Barr, 1972b; Gatti et al., 1976; Bonaccorsi et al., 1981); and in Sarcophagidae (Samols and Swift, 197913; Kaul et al., 1989a). Analysis of the distribution of fluorescent regions in mitotic chromosomes demonstrated that the Q' regions are consistently located in constitutive (C') heterochromatin (Figure 9). However, not all the C' regions show Q-fluorescence. Such fluorescence is related to the presence or absence of AT-rich satellites in particular regions of heterochromatin. There are exceptions, however. In D. nasutoides, the C- regions fluoresce brightly after staining with quinacrine (see Figure 13). Unlike the Q-stain, the H-stain (Hoechst 33258) preferentially binds to AT-rich nucleotide regions owing to external attachment to the DNA double helix (Comings, 1975; Latt and Wohlleb, 1975; Comings and Drets, 1976). According to the model of Mikhailov et al. (1981), the Hoechst molecule is located in a narrow groove of DNA occupying four base pairs. The AT specificity of the binding of this compound to DNA is provided by the formation of hydrogen bonds between the molecules of the stain and AT pairs.

Figure 9. A comparison of Q+ and C' regions in the mitotic chromosomes of Drosophila of the mekmogaster group. X, Y, 4 , 2 , 3 , chromosome numbers; Q and C, Q+-and C+-banding, respectively. After Lemeunier et al. (1978).

20

1. F. Zhimulev

H-heterochromatin has been described in many dipteran species of the Calliphoridae (Bedo, 1980) and Culicidae (Gatti et al., 1977; Bonaccorsi et al., 1980) families; in the Drosophila hydei group (Holmquist, 1975b; Gatti et al., 1976), D. mehogaster (Holmquist, 1975b; Gatti et al., 1976; Smaragdov, 1977, 1978; Smaragdov et al., 1980a; Gatti and Pimpinelli, 1983), D. simulans (Gatti et

Figure 10. A comparison of the localization of (2- (1) and H- ( 2 ) staining in the chromosomes of D. virilis. Numbers and letters designate the chromosomes. ( 3 ) The same chromosomes under phase contrast. Scale is 3.8 km. After Holmquist (197%).


21

Figure 11. Localization of the C-, Q-,and H-bands and satellite in the chromosomes of Drosophila nasutoides. (a) Q- and H-bands. (b) C-banding and localization of the satellite. After Wheeler and Altenburg (1977).

al., 1976; Smaragdov, 1978),D. wirilis (Holmquist, 197513,Gatti et al., 1976; Abraham et al., 1983), and other species of Drosophih (Wheeler and Altenburg, 1977; Lakhotia and Mishra, 1980; Singh and Gupta, 1982), and in the Sarcophagidae (Kaul et al., 1989b). The location of Q- and H-stains is generally coincident (Figure 10); there are differences, however (Figure 11). As in the case of Q- staining, H+ fragments occur mainly in the regions of C-heterochromatin, although not in all cases (Figures 12 and 13).An exception is D. nasutoides (see Figure 11).

3. N-staining After sequential removal of DNA, RNA, and acid-soluble proteins, the region of the nucleolar organizers retains its capacity to stain positively with the Giemsa reagent. In subsequent studies, it was found that this staining technique permits the identification of constitutive heterochromatin as well as the nucleolar organizers (Bianchi et al., 1971; Matsui and Sasaki, 1973; Matsui, 1974; Pimpinellietal., 1976; Hagele, 1977a; Beck and Srdic, 1979; Clayton, 1985;Kaulet al., 198913).These data may indicate that heterochromatic regions are rich in nonhistone proteins. A good correlation was found between the N-banded region and 1.705 repeats (AAGAG) in D. mehogaster (Bonaccorsi and Lohe, 1991; Lohe et al., 1993). Comparison of the location of the N+ regions of the chromosomes (Figure 14) reveals a strong correlation with other kinds of dyes, although not consistently (see also Gatti et al., 1976).

22

1. F. Zhimulev

Figure 12. Localization of H- and Giemsa (G) staining for constitutive heterochromatin bands in mitotic chromosomes of various Drosophifu species. After Lakhotia and Mishra (1980).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation (c+,H--,Q+)

0c+

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

tjH+++ (c- lH+++1Q++++ 1 (c- ,H+++,Q++) Figure 13. A scheme of the distribution of various heterochromatin types (C-,Q-,and H-banding) in a n arm of the fourth chromosome of D. nasutoides. After Wheeler e t nl. (1978).

Hoechst 33258

Quinacrine

1

hr1.696

hr1.714

mr 1.702

I

Figure 14. Localization of regions detected by various differential staining techniques in the mitotic chromosomes of D. hydei. X, Y, A, X and Y chromosomes and autosomes, respectively; G, C, N , AgNO,, Hoechst 33258, quinacrine, stain types; TAG, combined treatment with trichlordcetic acid, AgNO, and Giemsa staining; hr 1.696, hr 1.714 and mr 1.702: localization of the corresponding satellites. After Beck and Srdic (1979).

23

24

4.

1. F. Zhimulev

Differential enzymatic digestion of chromosomes

Heterogeneous distribution of nucleotide pairs in mitotic chromosomes is detected by digestion with restriction enzymes. The repeat-rich regions of the chromosomes not containing restriction sites for particular enzymes remain undamaged, and they are identified with any dye staining DNA, for example, ethidium bromide (Mezzanotte and Ferrucci, 1983; Mezzanotte, 1986). In the chromosomes of Diptera, these regions are coincident with heterochromatic regions (Mezzanotte and Ferrucci, 1983, Mezzanotte, 1986; Mezzanotte et al., 1986; Marchi and Mezzanotte, 1988; Faccio Dolfini, 1990;Tewari and Lakhotia, 1991).

5. Other treatments Among the other agents that are used to identify the heterogeneous distribution of chromosome material, mention may be made of acridine orange (Stockert and Lisanti, 1972; Lakhotia and Kumar, 1978; Mezzanotte, 1978; Smaragdov, 1978; Kaul e t al., 1989b), distamycin A binding to AT sequences (Faccio Dolfini and Bonifazio Razzini, 1983; Bedo, 1989), and diamidinophenylindol (DAPI) (Eastman et al., 1980; Abbott et al., 1981; Abraham et al., 1983; Bedo, 1989), as well as of various agents specific to GC sequences, such as antibodies to GC pairs (Eastman et al., 1980),mitramycin and chromomycin A, (Eastman et al., 1980;Abraham et al., 1983), binding to 3H-actinornycin D (Pirnpinelli et al., 1978), methylation of bases (Bianchi et al., 1986), or combined treatment with these agents (Beck and Srdic, 1979; Lakhotia et al., 1979; Hagele and Ranganath, 1983; Bedo, 1989).

6. Heterogeneity of heterochromatin Use of agents specifically binding to the various components of heterochromatin also reveals the considerable heterogeneity of the chromosome regions that are in a compacted state during most of the mitotic cycle (see Figure 14). Such examples are numerous (Gatti et al., 1976; Mezzanotte et al., 1979a-c; Abraham et al., 1983; Bedo, 1989; Kaul et al., 1989b).

C. Pairing of the heterochromatic regions of chromosomes A specific property of the heterochromatic regions is their capacity to establish contacts with one another.

1. Formation of chromocenters in interphase Data on the structure of interphase nuclei (Vosa, 1970; Ellison and Barr, 1971a, 1972b; Lewis and Craymer, 1971; Lakhotia and Kumar, 1980) provide evidence


25

for the presence of one or several chromocenters that arise by a fusion of the heterochromatic regions (Figure 15). The presence of more than one chromocenter (Figure 16) shows that the heterochromatic regions do not consistently fuse together to form a single chromocenter. In in situ hybridization of clones of DNA from the regions of the centromeric heterochromatin of Drosophikz, up to three labeling sites are detected in interphase nuclei (Lifschytz and Hareven,

198215). The centromeres are usually located at the nuclear envelope (see Figure

15) (Prokofyeva-Belgovskaya, 1965; Diaz and Lewis, 1975; Semionov and Smirnov, 1979,1984).

Figure 15. The chromocenters in the interphase nuclei of the neuroblasts of Drosophila melanogaster (A and B), D. simulans (C and D) and Anopheks aaoparwus (E-G). (A-D) After Ellison and Barr (1971a); (E-G) after Diaz and Lewis (1975). (A-D) Q-staining. (E-G) Phase contrast. (G) The location of the chromocenter at the nuclear envelope is seen. Scale is 10 k m (E-G).

26

1. F. Zhimuleu I

I

4oI

C

1

2

3

4

5

1

2

3

4

5

1

2

3

4

5

1

2

3

4

5

6

1

8

Figure 16. Number of chromocenters in the interphase nuclei of Drosophila neuroblasts. Ordinate, occurrence frequency of nuclei with the respective number of chromocenters (%I; abscissa, number of H’ chromocenters in the nuclei. (I) The preparations were made without the use of hypotonic solutions and colchicine. Total nuclei examined: a, 835 nuclei in XY males; b, 860 nuclei in XX females; c, 1636 nuclei in XO males. (11) The preparations were made with hypotonic solution; 103 nuclei in XX females were examined. After Smaragdov et al. (1980b).

2. Pairing in mitosis In the early prophase, the heterochromatic regions still join together in the chromocenter, subsequently, the euchromatic parts of sister chromatids, which were already tightly paired at the prophase of mitosis, separate (Figure 17), while the heterochromatic regions remain conjugated to the beginning of anaphase (this phenomenon is termed “chromatid apposition,” or adhesion). Chromatids of the chromosomes containing large quantities of heterochromatin (e.g., the Y or the fourth chromosome of D. melanogaster) never disjunct before the beginning of anaphase. Conjugation of chromatids is based on the features of heterochromatin itself but not those of the centromere. This appear to be so because fragments of heterochromatin, when transposed by the In( J)scv2 inversion to the distal end of the D. melanogaster X chromosome, pair exactly as though they are near the centromere (Smaragdov, 1978; Smirnov and Smaragdov, 1979). Data on the induction of mitotic crossing over favor the idea that the heterochromatic regions are in close proximity from the S phase to metaphase. Higher frequencies of mosaics for the genes in the X chromosome were obtained in strains containing more heterochromatin in this chromosome (Becker, 1969). At prophase of mitosis, homologous chromosomes become very closely approximated in Diptera and plants (somatic pairing; see Zhimulev, 1992b), frequently reminiscent of meiotic pachytene. There is no evidence indicating that the eu- and heterochromatic parts of the chromosomes differ in synaptic state (Nichols et al., 1972; Semionov and Smirnov, 1984).


Figure 17. Pairing of heterochromatic regions of sister chromatids in the karyotype ofDsosophilnarhabarca(a) and D. nusum (h). Numbers and letters designate chromosomes. (a) Staining with orcein. (h), C-handing. (a) after Paika and Miller (1974); (h) after Lakhotia and Kumar (1978).

27

28

I. F. Zhimulev

In preparations made with the use of colchicine and hypotonic solution, the heterochromatic regions of the chromosomes do not pair, whereas the condition of tight synapsis is characteristic of euchromatin. The sex chromosomes of females and males containing large amounts of heterochromatin are consistently asynaptic at this stage. With increasing degree of compaction, synapsis of euchromatin weakens, and homologs lie at some distance from each another as early as at prometaphase (Halfer and Barigozzi, 1972, 1973, 1977; Guest, 1975; Smaragdov, 1978; Smaragdov et al., 1980a). In homozygotes for the In(l)scv2 inversion, separating the heterochromatic block of the X chromosome and transposing it to the telomeric tip of the chromosome, the homologous chromosomes are asynapsed at both the distal and proximal heterochromatic blocks (Smaragdov et al., 1980a). Because the association of heterochromatic regions with the membrane persists to prometaphase (Zatsepina et al., 1977), it follows that the surface area of the membrane increases under the effect of hypotony; the widening membrane expands the heterochromatic regions of the chromosomes (Smaragdov et al., 1980a). Heterochromatin of homologous or nonhomologous metaphase chromosomes can associate under normal conditions (Semionov and Smimov, 1979, 1984) and after experimental treatment. When cell cultures of Drosophila are treated with distamycin A, chromocenters occur in 6.5% of metaphases (Faccio Dolfini and Bonifazio Razzini, 1983; Faccio Dolfini, 1987).

3, Pairing in meiosis At prophase I of meiosis, centromeric heterochromatin appears united in a dense, darkly staining body, the chromocenter (Figure 18). Chromocenters were described in the mosquitoes Anopkks atroparvus and Cukx pipiens (Diaz and Lewis, 1975; Fiil, 1978) and in Drosophila (Davring and Sunner, 1976, 1979; Nokkala and Puro, 1976; Chubykin and Chadov, 1987). It is Moens’ (1973) view that the chromocenter is a structure that can play a role in orientation of the chromosomes during the prophase of meiosis. By contrast, Chadov ( 1989) believes that events determining coorientation of the chromosomes do not occur before the chromosomes start to pair. Hence, the role of the chromocenter in meiosis at this stage is unclear. Pairing of nonhomologous X and Y chromosomes during the first division of meiosis in males of Drosophila is provided by specific recognition sites located in heterochromatin (Cooper, 1948,1949,1951,1964; de Marco et al., 1975; Yamamoto and Miklos, 1977, see Gatti and Pimpinelli, 1992, for review). There are specific sites responsible for pairing of nonhomologous sex chromosomes in Lucilia cuprina males (Bedo, 1987b).


29

Figure 18. Integration into the chromocenter of diplotene chromosomes at prophase of meiosis in Drosophila. After Nokkala and Puro (1976).

D. Localization of chromosomal rearrangements Concepts of the important role of heterochromatic regions in karyotype evolution have long been well known. In certain heterochromatic regions, chromosome arms become fused and separate, and this produces change in the morphology and number of chromosomes (Patterson and Stone, 1952; Swanson, 1957; Pathak et al., 1973; White, 1973). Intraspecific karyotype variation is largely the sum total of the contributions of chromosomal rearrangements with breakpoints in heterochromatin. Studies of 14 strains of Anopheles stephenst with chromosomal rearrangements resulted in mapping of three breakpoints to heterochromatin of the Y and second chromosomes (Sakai et al., 1983). The heterochromatic regions of the chromosomes of certain endemic species of Hawaiian Drosophila contain a high proportion of inversion breakpoints (Baimai, 1975a,b, 1977). In cell cultures of Drosophila, spontaneous rearrangements are most frequently formed in heterochromatic regions (Faccio Dolfini, 1976). There are data indicating that the occurrence frequency of breaks arising during the formation of induced chromosomal rearrangements is higher in the heterochromatic regions

30

I. F. Zhlrnulev

(see Hannah, 1951, for review). The suggested explanations are conflicting. Muller (1954) held the view that heterochromatin is more sensitive to the effect of radiation, opposed to this view Kaufmann (1954), who believed that heterochromatin possesses no particular susceptibility to this effect. Both conclusions were based on the results of studies of rearrangements induced in polytene chromosomes (see Section VII,C,4). Experimental data obtained with mitotic chromosomes are very scant. In analysis of metaphase plates of the neural ganglia of D. melanogaster larvae 2-14 hr after irradiation, it was found that the frequencies of terminal and isochromatid deletions are higher in the heterochromatic regions. Because the heterochromatin amount is about 20% in the autosomes and approximately 50% in the X chromosome, break frequencies corresponding to these percentages would be expected. However, the frequencies were 29.7-56.1% and 50.0-70.8% for the heterochromatin of the autosomes and the X chromosomes, respectively, while the frequencies of the formation of rearrangements in the almost entirely heterochromatic Y chromosome were considerably reduced. The authors explain it by the circumstance that heterochromatin in the Y chromosome is different from that in the other chromosomes (Gatti et al., 1974). In somatic cell cultures of D. melanogaster, ultraviolet light (uv)-induced chromosomal rearrangements are formed for the most part in the heterochromatin of the X and Y chromosomes and also in autosomes. The rearrangements induced by methylmethane sulfonate and x-rays are clustered on the heterochromatin of the autosomes and the X chromosome (Pimpinelli et al., 1977). When treating Drosophilidae carrying mutations for disturbed recombination and sensitivity to mutagens, it was found that about 80% of induced breaks map to chromatin in mus 109 mutants. In all the mutants approximately 80% of the “heterochromatic” breaks occur at the junction between eu- and heterochromatin (Gatti, 1979).

E. Late replication In his study of label distribution on autographs after incorporation of 3H-thymidine into the spermatocyte cells of the grasshopper Melanoplus differentialis, Limade-Faria (1959a,b) revealed four types of labeling: (1) no labeling, (2) labeling of only the euchromatin of the autosomes, ( 3 )continuous labeling, and (4) labeling of only a heterochromatic block of the sex chromosomes (Figure 19). Since the testicles of males of this species consist of a group of follicles in which spermatids are united into cysts synchronously passing through the stages of meiosis, Limade-Faria succeeded in demonstrating that the fourth type of labeling corresponds to the late stage of the S phase. The results of studies on many species of animals and plants allowed the general conclusion that the termination of DNA replication in the heterochro-


31

Figure 19. Types of 3H-thymidineincorporation into the nuclei at early pachytene of meiosis in Melanoplus differendis. The sex chromosomes of this species form the chromocenter, which is seen at the lefr part of the nucleus in all the figures. After Lima-de-Faria (1959a).

matic regions of the chromosomes is delayed (Lima-de-Faria and Jaworska, 1968; Shah et al., 1973; Back, 1976). Back (1976) even concluded that late DNA synthesis is the only invariable characteristic of heterochromatizable chromosomes known so far. This conclusion has been confirmed mainly in representatives of the Diptera order. In D. melanogaster, all the regions of heterochromatin (the Y and

32

1. F. Zhimulev

the fourth chromosomes, the proximal region of the X chromosome, and the centromeric regions of the second and the third chromosomes) replicate late. The location sites of Q+, H+, and C+ fragments and those of late replication are coincident (Barigozzi et al., 1966a, b, 1967, 1969, 1977; Barigozzi, 1968; HaIfer et al., 1969, 1970; Dolfini, 1971; Barigozzi and Halfer, 1972). Similar data (Figure 20) were obtained in Samoaia konensis (Ellison and Barr, 1972b), another representative of Drosophilidae. In Anopheles amoparvus, the long arms of the X and Y chromosomes containing Q' and C' fragments replicate late (Tiepolo et al., 1975). In D. virilis, heterochromatin constitutes approximately half of the genome. When 3H-thymidine is incorporated, labeled nuclei of three types are detected: the entire nucleus only, euchromatin, or heterochromatin. It was established that only euchromatin replicates during the first hour of the S phase; then the replication period of both eu- and heterochromatin proceeds for another 8 hr, and only heterochromatin replicates for 3 hr. Thus, the euchromatic part of the genome replicates for 9 hr while the heterochromatic part replicates 2 hr longer; consequently, heterochromatin replicates not only late, but also for a longer period. The heterochromatic regions terminate replicating asynchronously in different heterochromatic regions (Steinemann, 1980). These data provide evidence for a highly significant correlation between the location of heterochromatic regions and late replication.

Figure 20. A metaphase plate from Samaia leonensis fluorescent Q-staining, negative (a), and 120 min after incorporation of )H-thyrnidine (b). After Ellison and Barr (1972b).


33

However, the late-replicating regions are exceptionally, not consistent-

ly, heterochromatic (for review, see Lima-de-Faria and Jaworska, 1968). ConverseIy, not a 1 heterochromatic regions are late replicating, for example, the sex chromosomes (ZW) forming the heteropycnotic body replicate early in some snake species. In a number of angiospermous plants and several species of the bryophyte, including members of the Pellia genus, whose heterochromatin additionally shows C-banding (Heitz, 1928), the heterochromatic regions terminate replication early (see references in the review of John, 1988). In John’s opinion (1988) the replication time of chromatin depends on when it becomes decompacted in interphase. For example, in several bryophytes heterochromatin is compact in the G, phase but becomes diffuse from the beginning of the S phase; replication ceases in the middle of the S phase, and then it becomes compact again (John, 1988).

F. Variation in the amount of heterochromatin Two races differing only in the shape of the Y chromosome were identified in populations of D. pseudoobscura;the arms were of unequal length in the representatives of one race and they had the appearance of the Latin letter J, whereas their lengths were almost equal in those of the second race (V-shaped) (Lancefield, 1929). Dobzhansky’s (1935a, 1937) later studies proved that there may be as many as seven morphological types of Y chromosomes in this species and that the length of the chromosome can differ by twofold and more (Figure 21). A definite morphology is characteristic of each strain isolated from natural populations. Based on these differences, the original species was subdivided into two, D. pseudoobscura and D. prsimilis (Dobzhansky and Epling, 1944). At the end of the 1920s and the beginning of the 1930s, intraspecific differences in the sizes of the Y chromosome were detected in D. simulans (Sturtevant, 1929; Heitz, 1933a). A great deal of research has been done on the subject of karyotypes. Data on the karyotypes of 215 species of Drosophilu are given in Patterson and Stone’s (1952), and up to 513 species in Clayton and Wheeler’s (1979, catalogs. The karyotypes of approximately 150 species of Hawaiian Drosophila were described

I

m

P

Figure 21. Seven morphological types (Roman letters) of the Y chromosome of Drosophila pseudoobscura. After Dobzhansky (1937).

34

1. F. Zhimulev

(Clayton, 1968,1969,1971; Clayton etal., 1972; Carson, 1981; Carson and Yoon, 1982). Extensive studies of the karyotypes of other dipteran species are available (Boyes and Wilkes, 1953; Boyes, l953,1954a,b; Boyes and van Brink, 1964,1965, 1967,1970; Boyesetal., 1973; Boyes and Boyes, 1975; Boyes and Shewell, 1975). Four types of chromosomes are distinguished in karyotypes: twoarmed (metacentrics, or V-shaped), chromosomes with arms of unequal lengths (submetacentrics, or J-shaped), one-armed (telocentrics, or rodlike); and microchromosomes (dotlike). Variation in karyotypes is due mainly to centromeric fusion of telocentrics associated with the formation of V- and J-shaped variants and inversions. Wide variations in the sizes of the X, Y and microchromosomes (Figure 22), which are generally believed to be rich in heterochromatin, were concomitantly found (Clayton and Wheeler, 1975). With the use of specific stains, it was shown that the varying component is in fact heterochromatin (Figure 23), and a correlation between varying amount of heterochromatin and satellite DNA was established in some cases (Holmquist, 1975a) (see also Figure 8). In many species of Drosophila, particularly Hawaiian, polytene chromosomes show exactly the same banding pattern (the euchromatic part of the chromosome). They do not differ even in inversions (homosequential species), while exhibiting great differences in heterochromatin amount or satellite DNA (Craddock, 1973; Carson, 1981; White, 1982; Chang and Carson, 1985). The information given in Table 1 shows that this phenomenon is widespread. Data on variations in heterochromatin amount in other species of animals and plants may be found in published reviews (Battaglia, 1964; Brown, 1966; Yunis and Yasmineh, 1972; White, 1973; Evans, 1976; Prokofyeva-Belgovskaya, 197713, 1986; Bostock, 1980). It is unclear why heterochromatin amount is not constant. One cause may be variation in the copy number of repetitive DNAs abundant in the heterochromatic regions. An understanding of the causes may be approached through the correlation established between the formation of inversions with a single breakpoint in a heterochromatic region and heterochromatin amount. The karyotype of

0

St

Mt

Lt Mm Lm Ssm Msm Lsm

Figure 22. Different types of the fourth microchromosome of D. kikkawai occurring in natural populations. Letter designations: D, dot; t, telomeric; sm, subtelomeric; m, metacentric. T h e chromosome sizes of small, medium, and large are designated by S, M, and L, respectively. After Baimai et al. (1986).


35

Figure 23. Differences in the amount of C-heterochromatin in a Chironomw thummi (th) X Ch. th. piger (pi) hybrid. After Hagele and Speier (1988).

Drosophila formella contains an inordinately large amount of heterochromatin in the autosomes, the X and the Y chromosomes. In karyotypes with paracentric inversions, no changes in heterochromatin amount occur; however, once an inversion break occurs in heterochromatin, one might expect a virtually new chromosome arm composed of heterochromatin to arise. Similar correlations were found in D. recticilia (Figure 24) and D. disjuncta (Baimai, 1975a,b, 1977).

G. Formation of heterochromatic regions of chromosomes during deve Iopm e nt Studies carried out in fishes, amphibia, and mammals have led Prokofyeva-Belgovskaya (1960, 1982, 1986) to the conclusion that the early metaphase chromosomes are much different in morphology from those at the later developmental stages: they are more slender, they are decompacted to a very high degree, and they lack heterochromatin blocks. She has called such chromosomes juvenile (Prokofyeva-Belgovskaya 1960, 1982, 1986). In Drosophila, eight synchronous divisions of nuclei occur during early embryonic development; subsequently, the nuclei start to migrate to the egg surface during ongoing division. Later, near the surface, four additional divisions take place (the 10th through 13th cycles), and, as a result, blastoderm is formed from one layer of the nuclei. After the formation of cell membranes around the nuclei in the interphase of the 14th cycle, cell blastoderm is layered. Cell cleavage divisions proceed rapidly. One mitotic cycle takes 10 min, on average (Table 2). Starting from the 12th division, the duration of each cycle increases: from 12.4 min in the 12th division and to 21.1 min in the 13th (Foe and Alberts, 1983). In both the early cytological and the more recent electron microscopic studies on Drosophila, it was noted that there are no nucleoli and chromocenters in

36

1. F. Zhimulev

Table 1 Variation in the Amount of Centromeric Heterochromatin in Mitotic Chromosomes of Diptera Species

Variation types

References

Anopheles complex balabacensis

Variation in the amount of constitutive heterochromatin

Baimai et al. (1981)

A. gambiae and arubiensis

Interspecific differences in the amount of H-heterochromatin in the Y chromosome

Bonaccorsi et al. (1980)

A. dims

Variation in constitutive heterochromatin of the sex chromosomes (5 types of the X chromosome, 4 types of fourth chromosome)

Baimai et al. (1984a) Baimai and Traipakvasin (1987)

A. maculatus

Three types of variation in C-heterochromatin in the X chromosome and 4 types in the Y chromosome

Greenet al. (1985)

Anopheles

Variation in heterochromatin in the sex chromosomes Variation in the size of the X chromosome

Baimai et al., (198413) Boyes and Wilkes (1953)

Variations in the length of the sex chromosomes

Boyes and Shewell (1975)

Large blocks of C-heterochromatin in Ch. th. thummi and almost complete absence in Ch. th. piger (see Figure 23) Variation in morphology of the Y chromosome

Hagele and Speier (1988)

Of 152 studied species, variation in heterochromatin amount was detected in the microchromosomes of 14 species Of 103 species, heterochromatin amount varies in 9

Yoon and Richardson (1978b), Carson and Yoon (1982)

Comparisons of species reveal large differences in the amount of heterochromatin with the same polytene chromosomes

Baimai and Aheam (1978), Aheam and Baimai (1987)

D. afinis

Intraspecific polymorphism for size and morphology of the Y chromosome

Miller and Stone (1962), Miller and Roy (1964)

D. albumicans

lntraspecific variation in the Y and the fourth chromosomes; variation in C-heterochromatin (from complete absence to large block) Various morphological types of the Y chromosome

Wilson et al. (1969), Clyde (1980), Hatsumi (1987)

Species of the Aplomya genus Species of the Calliphoridae family subspecies Chironomus thummi thummi, Ch. th. piger Closely related species Cnephia dacotensis and C. omitophilia Drosophila (Hawaiian species) Drosophila (picturewinged Hawaiian species) D. afinidis-juncta. D. bosqcha, D. disjuncta

D. algonquin

Procunier (1975a)

Clayton (1988)

Miller and Roy (1964)

continued

37

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 1 Continued Species

Variation types

References

D. athabasca

Same as D. algonquin

Miller (19571, Miller and Stone (1962), Miller and Roy (1963)

D. azteca D. bifurca D. birchii


Miller and Roy (1964)


Ward (1949)

Four types of the X, three types and two types of the fourth chromosomes

Baimai (1969a)

D. brunnei-palpa

Very small, almost dotlike Y chromosomes

Dobzhansky and Pavan (1943a, b)

D. cardini

Addition of heterochromatin on the microchromosome

Ward ( 1949)

D. cyrtoknna

Increase in the size of heterochromatic regions of the microchromosomes compared to D. grimshawi

Clayton (1985)

D . disjuncta

Variations in the amount of hererochromatin correlate with the formation of inversions, with breakpoint in heterochromatin

Baimai (1975a)

D. formella D. furvifacies

Same as D. disjuncta

Baimai (197513)

Interspecific variations in the amount of heterochromatin

Yoon et al. (1972)

Variation in the shape and size of the Y chromosomes Variation in heterochromatin of the Y and fourth chromosomes

Mather (1962)

in comparison with related species D. biseriata, D. hysaicosa, D. mitchelli

D. gr. immigrans

Wakahama et al. (1983)

D. gr. melanogaster

Interspecific variation in Q- and C-heterochromatin (see Figure 9)

Lemeunier et al. (1978)

D. hernipem

Less C-heterochromatin in the X chromosome, but more in the microchromosome than in related species D. silwesais, D. heteroneura, D. differens, D. pianitibia

Chang (1984), Chang and Carson (1985)

D. kikkawai

Variation in constitutive heterochromatin and in the microchromosomes (9 types of morphology) and the Y chromosome (4 types) (see Figure 22)

Baimai (1973), Baimai and Chumchong (1980), Baimai et al. (1986)

D. kontia

Variation in constitutive heterochromatin of the microchromosomes and the Y chromosome


continued

38

1. F. Zhirnulev

~~~~

Table 1 Continued Species

Variation types

References

D. leontia and D . kikkawai D. melanica

Interspecific differences in the Y and microchromosomes

David et al. (1978)

Increase in heterochromatin amount in the microchromosome

Ward (1949)

D. melanagaster

Some Q' fragments lose fluorescence in cell culture during translocation formation Increase in blocks of centromeric heterochromatin after long-term culturing of cells Interstrain differences in Q-heterochromatin amount in the Y (3 types), the second (2 types) and the X (2 types) chromosomes

Zuffardi et al. (1971)

D. mehnura

Shape and size variation in the fourth and the Y chromosomes

Ward (1949)

D. rneridionalis

Differences in the size of blocks of structural heterochromatin of sex chromosomes and/ or microchromosomes


D. micromelanica

Increase in heterochromatin amount in the microchromosomes

Ward (1949)

D. montium

Shape and size variation in the fourth and the Y chromosomes

D. narmagansett

Shape and size variation in the Y-chromosome

Kikkawa (1936), Ward (1949), Baimai (196913) Ward (1949)

D. w u t a

Variation in heterochromatin amount in the X-chromosome Differences in the size of heterochromatic regions of homologous chromosomes of two subspecies Large variation in size and shape of the Y chromosome both between and within species Differences in the length of the Y chromosome in related species

D. m u t a m u t a and D. m u t a albomicana D. of nasufa subgroup

D. pachea, D . acanthoptera and D. nannoptera D. paranaensis

Variation in size of the microchromosomes

D. pseudoobscura

Seven types of morphology and size of the Y chromosome (see fig. 21)

Halfer et d.( 1980)

Halfer (1981)

Lakhotia and Roy (1981) Ranganath and Hagele (1982) 4

Wilson eta[. (1969)

Ward and Heed (1970)

Wasserman and Wilson (1957) Lancefield (1929), Dobzhansky and Boche (1933), Dobzhansky (1935a, 1937) continued


39

Table 1 Continued Species

Variation types

References

D. recticilia

Variation in heterochromatin amount correlates with the presence of inversions having a single breakpoint in heterochromatin

Baimai (1977)

D.serldo

Differences in block size of structural heterochromatin in the sex chromosomes and/or in microchromosomes


D. simuhns

lntraspecific differences in the size of the Y chromosome

Sturtevant (1929), Heitz (1933a)

Glossina mursirans Interspecific variations in the size and mursitans, G . m. subamount ofC-heterochromatin in the morsitans, G . austeni Y chromosomes Parasarcophaga

In five species, enormous differences in the size of the X and Y chromosomes from dot like in P. albicens to giant rods in P. knabi

Pel1 et d. (1972), Davies and Southern (19761, Jordan et al. (1977)

Kaul et al. (1978)

the interphase nuclei involved in the first 11-12 cleavage divisions. Chromatin is represented by a finely dispersed network (Figure 25), and heterochromatin is not detected in mitotic chromosomes (Huettner, 1933a; Rabinowitz, 1941; Sonnenblick, 1950;Mahowald, 1963b, 1968;Illmensee, 1972;Mahowald and Hardy, 1985).

1

V

t

I ~

t

Q S

Figure 24. A scheme demonstrating increase in heterochromatin amount (from a to c) as a result of formation of chromosomal rearrangements in D. recticilia. g, s, v, inversions. After Baimai (1977).

40

1. F. Zhimulev

Nucleoli and chromocenters appear only at the blastoderm stage (Mahowald, 1963a). In blastoderm nuclei transplant into unfertilized eggs, chromocenters and nucleoli rapidly disappear, chromatin is converted into a finely dispersed network (see Figure 25), and cleavage division starts as early as after 15 min (Illmensee, 1972). When stained for C-heterochromatin, the metaphase chromosomes passing through the first four to five divisions (30-60 min after fertilization) differ from those at the blastoderm stage or the neuroblast chromosomes (Figure 26). They have the appearance of long, slender, weakly condensed fibrils. Only the Y chromosome stains quite distinctly in the karyotype, and mere "traces" of stain are seen in the centromeric regions of the other chromosomes. Metaphase chromosomes are in a state of weak compaction for a brief period, and they assume their usual appearance after 4-5 divisions (Vlassova et al., 1991a,b). The distinctive features of differentiation of the chromosomes into euand heterochromatin during early cleavage divisions are presumably due to the specificity of their organization and functioning. It is known that development during early embryogenesis is effected by maternal RNA and proteins stored in the egg. There is no transcriptional activity or it is very weak to the ninth division cycle. At this developmental stage, there are no nucleoli, and there is no incorporation of labeled amino acids and uridine. Chromatin is almost completely switched off from transcription (Mahowald, 1963a; McKnight and Miller, 1976; Zalokar, 1976; McKnight et al., 1978). Transcription is sharply enhanced (about fivefold) during the 10th cycle at the blastoderm stage, when synthesis of ribosomal RNA starts and nucleoli are formed (Mahowald, 1963a;McKnight and Miller, 1976; Zalokar, 1976; McKnight et al., 1978; Anderson and Lengyel, 1979; Foe et al., 1982; Edgar and Schubiger, 1986; Underwood and Lengyel, 1988; Wieshaus and Sweeton, 1988). The start of transcription is coincident with the beginning of the lengthening of cell cycle and differentiation of the chromosomes into zones of strong (hetTable 2 Duration (rnin) of the Mitotic Cycle Stages in the First 11 Cleavage Divisions in D. melanogaster" ~

~

Temperature

Stage of the mitotic cycle

24°C

29°C

30°C

Complete cycle Interphase Prophase Metaphase Anaphase Telophase

9.5 3.4 4.0 0.3 1.0 0.9

8.9 2.5 3.6 0.5 1.4 0.8

8.8 2.7 3.0 0.7 1.1 1.4

"After Rabinowitz (1941).


41

Figure 25. Ultrastructure of the hlastoderm cell nucleus before (a) and 15 min after (h) transplantation into an unfertilized egg of Drosophila. no, nucleus; ch, chromatin, nm,nuclear membrane. After Illmensee (1972).

Figure 26. Constitutive heterochromatin of Drosophila in neuroblasts (a and b), in blastoderm (c and d), and at early stages of cleavage division (e-i). (a, c, e-i) Metaphase. (bd) Prometaphase. ( e and f) G-banding. (g-i) C-banding. Bold arrows indicate the centromeric regions of the chromosomes; thin arrows indicate the Y chromosome. After Vlassova et al. (1991a,b).

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

43

erochromatin) and weak (euchromatin) compaction. In the meantime, changes in chromosome composition take place, unusual histones (juvenile) that stain neither with alkaline fast green, as do the usual histones, nor with bromphenol blue occur in the dividing nuclei up to the end of blastocyst formation. Staining with fast green appears before the actual formation of blastoderm (Das et al., 1964). A similar substitution of lysine-rich histones was detected in the sea urchin (Hieter et al., 1979) and the snail Helix aspersa (Bloch and Hew, 1960) during embryonic development. A nonhistone protein, HP1, that is identified mainly in heterochromatin is not detected in the nuclei up to the 5th-6th division cycles. It is consistently detected at the 10th-1 Ith stages (James et al., 1989; Kellum et al., 1995).This corresponds to the stage of development of increased phosphorylation of HP1 (Eissenberg et al., 1994).

111. REPETITIVE SEQUENCES In eukaryotes, specific portions of a given genome are composed of short, multiply repetitive sequences with base compositions different from that of the bulk of the genome. Repeated DNA sequences can be isolated with the use of two approaches: one based on their exceptionally high renaturation rate and the other on gradient density centrifugation. In the latter case, the greater part of the DNA constitutes the major precipitation band (the main band) and the repetitive fraction, because of its richness in a particular nucleotide set and, consequently, having a different molecular weight, yields a single or several additional (satellite) bands (Figure 27). Problems relating to satellite DNA have been dealt with in many reviews (Bostock, 1971; Walker, 1971; Yunis and Yasmineh, 1971; Chaudhuri, 1975; Lindsley, 1975; Tartof, 1975; Appels and Peacock, 1978; Panitz, 1978; John and Miklos, 1979; Brutlag, 1980; Peacock et al., 1981b; Beridze, 1982; Singer, 1982; Ginatulin, 1984; Hardman, 1986; Miklos, 1987; Lohe and Roberts, 1988; Lohe and Hilliker, 1995). The main information concerning the molecular organization, as well as the chromosomal location of satellite DNA and its relevance to heterochromatin, is presented in this section. From the results of studies on numerous animal and plant species, it may be concluded that, on average, approximately 30% of genomic DNA consists of satellites, although considerable deviations therefrom are known. Satellites have been described in detail in D. mehogaster and closely relatedspecies (Fansleretal., 1970; Rae, 1970; Botchanetal., 1971; Galletal., 1971; Kram et al., 1972; Travaglini et al., 1972; Peacock et al., 1974, 1977, 1978; Brutlag and Peacock, 1975, 1979; Endow et al., 1975; Brutlag et al., 1977, 1978; Carlson and Brutlag, 1977, 1979; Wollenzien et al., 1977; Barnes et al., 1978; Fry and Brutlag, 1979; Hsieh and Brutlag, 1979a; Donnelly and Kiefer, 1986; Abad et al., 1992;Lohe et al., 1993; Lohe and Hilliker, 1995; Makunin et al., 1995, 1996); in Drosophila gr. virilis (Gall et al., 1971, 1974; Blumenfeld and Forrest, 1972; Blu-

44

1.

F. ZhimUlRV

30

I I 692

I

I688

1

1671

": (C)

Dvmh a D anwictm

I691

1 I 687

Buoyant density

Figure 27. Distribution of DNA fractions in D. wirilis (a), D. americana (b), and their hybrid (c) in a neutral CsCl gradient. Abscissa, buoyant density; ordinate, optic density. After Gall and Atherton (1974).


45

menfeld et al., 1973;Blumenfeld, 1974; Gall and Atherton, 1974; Schweber, 1974; Steinemann, 1976; Mullins and Blumenfeld, 1979; Cohen and Kaplan, 1982); in D. hydei and related species (Hennig et al., 1970; Dickson et al., 1971, Hennig, 1972a,b; Renkawitz, 1978a,b); in D. nasutu and related species (Cordeiro et al., 1975; Cordeiro-Stone and Lee, 1976; Lee, 1978; Ranganath et al., 1982); in Hawaiian Drosophila (Miklos and Gill, 1981); in D. quanche (Bachmann et al., 1989); and in Glossina (Amos and Dover, 1981), Heterogera (Kunz and Eckhardt, 1974), Anopheks (Redfern, 1981a), Chironomus (Steinemann, 1978), Rhynchosciaru (Eckhardt and Gall, 1971), Sciara (Abbott et al., 1981), Surcophaga (Bultmann and Mezzanotte, 1987), and Lucilia cuprinu (Perkins et al., 1992). All the data obtained so far (Figure 28) indicate that satellite DNA is located in the regions of centromeric heterochromatin of the metaphase chromosomes (Jones and Robertson, 1970; Gall et al., 1971; Goldring et al., 1975; Peacock et al., 1977, 1978; Perreault et al., 1978; Renkawitz, 1978a,b; Wheeler et al., 1978; Samols and Swift, 1979a; Steffensen et al., 1981; Lifschytz and Hareven, 1982a,b; Ranganath et al., 1982; Bachmann et al., 1989) or in the chromocenters of interphase nuclei (Gall et al., 1971; Kunz and Eckhardt, 1974; Cordeiro-Stone and Lee, 1976). There are satellite DNAs located not only in centromeric heterochromatin, but also in the euchromatic regions. For example, the 1.672, 1.686, and 1.705 satellites of D. melanoguster are located in the 21D region of chromosome 2L (Rae, 1970; Goldring et al., 1975; Sederoff et at., 1975a;Peacock et al., 1978); one of the satellites has an additional location site in telomeres (Peacock et al., 1978; Traverse and Pardue, 1989). Sequences related to the 1.688 satellite (63-81% homologous) and arranged in short (two to four copies) arrays are present in several different regions of the D. melanogaster X chromosome (Waring and Pollack, 1987; DiBartolomeis et al., 1992; Losada et al., 1995; Kokoza et al., 1997). In D. simulans, the 1.696 satellite, consisting of a repeat 15 bp long, maps to three euchromatic regions in addition to the chromocenter (Appels and Peacock, 1978; Lohe and Roberts, 1988). In Anopheks stephemi, satellite I maps to the centromeric regions and the euchromatic part of the third chromosome (Redfern, 1981a). In D. uirilis, certain satellites are also mapped to positions in the euchromatic (polytenizing) regions of the chromosomes (Cohen and Bowman, 1979; Cohen and Kaplan, 1982). In Glossinu awteni a large amount of satellite DNA is detected in the B chromosomes (see Figure 28), in addition to centromeric regions, and in the gall midge Heteropeza pygmeu, satellite DNA is identified in the chromosomes restricted to the germline tissues (see Section v) (Kunz and Eckhardt, 1974). In the polytene chromosomes of Phaseolus coccinew suspensors, satellite DNA hybridizes with the regions of centromeric heterochromatin, blocks of intercalary heterochromatin, and presumably telomeric heterochromatin (Tagliasacchi et al., 1984).

Figure 28. Localization of satellite DNAs in centromeric regions of the metaphase chromosomes of D. mekanogastes (a and b), in centromeric regions and heterochromatic chromosomes of D. nasutoides (c-g), and in centromeric regions of the chromosomes of the main set (h) and in B chromosomes (i) of Glossinu awteni. Numbers and letters designate chromosomes (B, supernumerary chromosomes). (c and e ) T h e chromosomes after C- and G-banding, respectively. The scale is 5 pm. (a and b) after Perreault et al. (1978); (c-g) after Wheeler er al. (1978); (h and i) after Amos and Dover (1981).


47

Figure 28. Continued

In addition to satellites, rDNA (Hilliker and Appels, 1982), rDNA spacers (Lohe and Roberts, 1990), and other repeats (Huijser and Hennig, 1987, Linares et al., 1994) are identified in the regions of centromeric heterochromatin (see Section IV,A). The molecular organization of satellite DNA and its location in the heterochromatin of D. melanogaster and closely related species have been analyzed in detail. When centrifuged in CsCl density gradient with the addition of antibiotics, DNA from diploid cells is separable into a major fraction and five satellites. The satellites are divided into three groups according to base composition and complexity of repeating sequences: 1. The fraction sedimenting at the zone of the 1.679 g/cm3 gradient (or the 1.679 satellite) and consisting of ribosomal DNA sequences (Peacock et al., 1974, 1978). 2. Satellites (1.672, 1.686, 1.705) consisting of multiply repeated short (5-10 bp) fragments (Peacock et al., 1974, 1978; Brutlag and Peacock, 1975; Endow et al., 1975; Sederoff et al., 1875a; Endow, 1977; Leemann and Ruch, 1984; Lohe and Roberts, 1988). 3. The 1.688 satellite, primarily an array of tandem repeats 359 bp long (Hsieh and Brutlag, 1979a; Lohe and Roberts, 1988; Lohe et al., 1993). Data on representation of various fractions in the Drosophila genome are given in Table 3. The total amount of satellite DNA is close to that calculated as constituting centromeric heterochromatin (see Lohe et al., 1993). Cloning of short (300-600 bp) fragments isolated from various satellites (Lohe and Brutlag, 1986), followed by determination of their nucleotide se-

48

I. F. Zhimulev

Table 3 Amount (kb) and Distribution of Satellites in the Chromosomes of D. mekmogasterR Chromosome Satellite 1.672 1.686 1.688 1.697 1.705 1.690 rDNA Total: Calculated amount of DNA in heterochromatin

X

Y

2nd

3rd

4th

?

380 640 6830 4000 660 2800 15,310 13,00016,000

5140 4620 6260 3400 6820 2200 28,440 39,000

75 2220 570 1300 4320 8490 800015,000

300 1660 430 765 1180 -

1650 91 560 130

5100

-

2430 30004500

4340 13,00016,000

-

-

5100

"After Peacock et al. (1978)

quences, demonstrated that each satellite consists of several types of simple sequences multiply repeated in tandem arrays. Eleven sequences were identified in four satellites (Table 4),with each satellite having a major and several minor fractions, which constitute a small portion of the genome. Minor fractions are also repeated in tandem and form long tracts. When centrifuged, these fractions separate as individual satellites (see Table 4). Because only a relatively small number of clones was derived from satelTable 4 Representation (%) of Different Types of Sequences Composing Satellite DNA in the Genomes of D. rnelanogaster and Closely Related Species" Nucleotide sequence of the repetitive unit, 5'-3'

Buoyant density (gm/cm3)

D. rnelanogaster

D. sirnulam

D. erecta

AATAT AATAG AATAC AAGAC AAGAG AACAA AATAAAC AATAG A C AAGAGAG AATAACATAG 359 bp

1.672 1.693 1.680 1.689; 1.701 1.705 1.663 1.669 1.688 1.686 1.688

3.1 0.23 0.52 2.4 5.6 0.06 0.23 0.23 1.5 2.1 5.1

1.9 2.4 0.0065 0.71

0.0088 0.041 0.00 18 0.01 1 0.55 0.0015 0.0016 0.0070 0.0091 0.24

~

~

0.10 0.036 0.074 0.11 ~

"After Lohe and Brutlag (1987a) and Lohe and Roberts (1988).


49

lite DNA, it was expected that a large number of new minor fractions would be identified, up to 100 or more, as estimated by Lohe and Roberts (1988). A new satellite sequence (AACAC) has been described that is located mainly in the 2R heterochromatin and comprise about 1000 kb (Makunin er al., 1995, 1997). Nucleotide composition within cloned sequences is mostly homogeneous (Lohe and Brutlag, 1987a,b; Lohe and Roberts, 1988).A single nucleotide substitution has been identified per l-kb repeat (Figure 29a and c); however, substitution frequency can be 100-fold greater and more in a few clones (see Figure 29b and 29d). The greater part of the 1.688 satellite is represented by a tract 359 bp long. In contrast to satellites composed of short sequences, the repeating units of the 1.688 satellite are dissimilar and show a 4-5% variation in nucleotide composition (Carlson and Brutlag, 1979; Lohe and Brutlag, 1986; Hsieh and Brutlag, 1979a; Lohe and Roberts, 1988). It was demonstrated that the satellite sequences are complex. Retrotransposon Doc was present in three of the eight plasmids chosen from a genomic library of plasmids containing satellite DNA of density 1.688.In one of the plasmids, two copies of Doc were in the opposite orientation at the 3’ end bounding the repeated monomer (359 bp) at both flanks (Slobodkin and Alatortsev, 1992). Transposable element insertions (including copia and 297) are ocassionally found in clones of satellite DNA (Carlson and Brutlag, 1978;Lohe and Brutlag, 1987a,b; Lohe et al., 1993). The AACAC satellite, located within the 2R heterochromatin, is related to the Stalker mobile element (Makuninetal., 1995,1997). Extensive analysis of the distribution of 11 different transposable elements (TEs) of the D. melanogaster mitotic chromosomes has shown that 9 are clustered into one or more discrete heterochromatic regions. The locations of the mobile elements are highly conserved in geographically distant strains. Most of the heterochromatic regions contain one or more TE families. All the heterochromatic blocks are enriched in satellite DNA sequences. Only a few blocks are devoid of TEs (Pimpinelli et al.,1995; Carmena and Gonzalez, 1995). These data show that the DNAs of the mobile elements intersperse the satellite DNAs. The Dp(I ;f) f 187 duplication, a minichromosome, contains the normal components and functions associated with eukaryotic chromosomes. It is small, about 1.3 Mb or 1/30th of the normal X chromosome, including 1 Mb of centric heterochromatin and a 290-kbsegment from the normal tip of the X chromosome. Early restriction mapping suggested that a cluster of restricted sites, termed an “island of complex DNA,” was present within 50-100 kb of the Dp 1 187 euchromatin-heterochromatin junction (Karpen and Spradling, 1990, 1992; Spradling et al., 1992; Spradling, 1994; Thompson-Stewart et al.,1994; Le et al., 1995). Irradiation mutagenesis of Dp J 187 and pulse-field restriction mapping revealed that this part of Drosophila melanogaster (heterochromatin) is organized alternating blocks of complex islands (Tahiti, Moorea, and Bora Bora) and satellite DNA. Each island is hundreds of kilobase pairs in length, constituting approxi-

50

1. F. Zhimulev

a.

b. A A T A AC ATAG

A A T A A C AT A G

50

--G- * -G-

loo

100

d.

C. AATAG

AATAG

-

Figure 29. Nucleotide composition of various DNA clones isolated from the 1.686 (a and b) and 1.672 (c and d) satellites. The adjacent repeats are ordered one under another, and they are represented by a horizontal line. Nucleotide substitutions are designated by letters at the lines. After Lohe and Brutlag (1987b) and Lohe and Roberts (1988).


51

mately one-half of the Dp 1 187 heterochromatin. Cloning and sequencing of a small part of one island, Tahiti,demonstrated the presence of a Doc retroposon connected to a 359-bp satellite (Le et al., 1995). However, analysis of DNA sequences flanking P element insertions into numerous regions of mitotic heterochromatin revealed that middle repetitive or unique sequence DNAs are frequently interspersed with satellite DNAs (Zhang and Spradling, 1994, 1995). Repeated sequences have been found in the most distal part of the X heterochromatin of D. melanoguster within 60 kb of SCLR. The repeated unit contains 1150 bp of Stellate repeat; a copia-like retrotransposon;an element of LINE type, including amplified insertions of type I into the rRNA genes, and fragments of the rRNA genes themselves (Nurminsky and Shevelyov, 1992; Shevelyov, 1992; Nurminsky et d., 1994; Tulin et al., 1997). The heterochromatic h39 region of the second chromosome of D.melanoguster contains Rsp-associated Xba I repeats and tandemly repeated Bari-I mobile elements (Caizzi et al., 1993). The Aurora mobile element inserts into the highly repeated DNA of the Stellate locus (Shevelyov, 1993). A DNA fragment containing four different repeats, including Rsp, Bari1 and AT-rich repeated sequence, Porto-l , is located very close to the centromere of chromosome 2 in D. melanoguster (Coelho et at., 1996). Fine mapping of the location of DNA of the various satellites in the metaphase chromosomes of D. melanoguster was performed with the use of in situ hybridization (Peacock et al., 1977, 1978; Appels and Peacock, 1978; Steffensen et al., 1981; Bonaccorsi and Lohe, 1991; Lohe et al., 1993).Localization was performed with accuracy for several megabases (Figure 30), and the following conclusions may be made:

1. Five major satellites exhibit a multichromosome distribution. Five minor satellites were found in single site of the Y chromosome. 2. The closely related satellites are often located on nearly the same chromosome.

3. About 80% of Y chromosome DNA is composed of nine simple repeated sequences, among them: AAGAC (8 Mb), AAGAG ( 7 Mb), and AATAT (6 Mb). More than 70% of the heterochromatin of the second chromosome is composed of five simple repeating sequences (Bonaccorsi and Lohe, 1991; Lohe et al., 1993). The repeated DNAs are thought to be the most rapidly evolving parts of the genome (Peacock et al., 1981b). Comparisons of the satellite profiles of even very closely related species (see Figure 27) reveal great differences. For example, approximately 40% of DNA is composed of satellites in D. uirilis, and 35% in D. texam, but, with different sedimentation constants, no satellite DNAs were identified in a third closely related species (Gall et al., 1971, 1974; Gall and Atherton,

52

1. F. Zhimulev 26

27 28

X L --

29

30 31

32

3334

XA

-

-

~

1 2 345678

YL-

- -

9 10 11121314

-

15

t.kW N

1617 18 19

rDNh 359bp hAGhG hATAT

20

21 2 2 2 3 2 4 2 5

m -u Y S &TAT - MGhG - MGAC - AhGhGhG

i

-

MThC MThG .UThGhC hAT'4hAC rDNh

35 36 37

Z L ---

383941 41 4344 45 46

Rig--- 2R

IF&#

- -

- .

-

...-...

3L ---

4758495051 52 53 >*q, 6 %4

5455 56 57 58

--

-

-

MGhG AAGhGhG MGhC MThG AAThAChThG R8p AAChC

3R

AATAAChThG MGAG

.L4TL4T MG4G

Figure 30. Map of major satellite locations in D. melanogaster heternchromatin (according to Bonaccorsi and Lohe, 1991; Lohe et al., 1993, modified). Localization of AACAC satellite is according to Makunin et al. (1995, 1997). Numbers and letters designate chromosomes.

1974). Great differences in the profiles of satellites were detected in two subspecies, Dosophila nasuta nasuta. and D. n. albomicuna (Ranganath et al., 1982); in the closely related species of the melanogaster group (Travaglini et al., 1972; Peacock et al., 1974; Lohe, 1981; Barnes et al., 1978; Lohe and Roberts, 1988), and in the hydei group (Hennig et al., 1970). The data in Table 4 allow us to follow how sharply the occurrence of particular sequences making up satellites decreases in the genomes of D. melanogaster,


53

D.simulans, and the more remote species D. erecta. We are dealing here with variation in copy number in related species. In in situ hybridization, the satellites identified in the mehogaster group are also identified in other species, when the preparations are very overexposed. For example, in hybridization of a clone with DNA containing a sequence of the 1.672 satellite, the distribution of label on the metaphase chromosomes of D.mehogaster, D. simuhns, D. mauritiana,D. yakuba, and D. teisszeri was found to be the same (Lohe and Roberts, 1988). Cases of “slipping” are known: certain sequences (5’-AACAA-3’,5‘AAGAC-3‘, 5 ‘-AATAACATAG-3’) of D. mehogaster that are not identified in the closely related species D. simuhns are detectable in D. erecta (see Table 4). Similar results were obtained in an analysis of the occurrence of the polypyrimidine repeat composing the 1.705 satellite. It was found to occur in the closely related species D. melanoguster, D. simufans, and D. mauritiana, as well as in the remote D.varians species, a member of another group (D. anamsue), and not to occur in the rest of the related species (Cseko et at., 1979). In D. gwmche, heterochromatin is composed mainly of a satellite represented by a sequence 290 bp long and repeated about 80,000 times. This sequence is species specific, not occurring in the sibling species D. subobscura and D. madeirensis (Bachmann et al., 1989). Variation in sequence abundance of highly repeated DNA between strains of Aedes albopictus was as great as between it and six other species of the complex (McLain et al., 1987). There is a specificity in the protein composition of chromatin containing satellite DNAs (for review, see Zhimulev, 1992b). The fraction of HI histone that is involved in compaction of chromatin brought about by aggregation of nucleosomes is underrepresented in chromatin at the early stage of embryonic development (Elgin and Hood, 1973), when the chromosomes have not yet differentiated into eu- and heterochromatin. There is reason for believing that the phosphorylated H1 histones can specifically bind the repeated DNA sequences involved in their compaction into heterochromatin. In in vitro experiments, H 1 histone of D. virilis preferentially binds satellite DNAs; the decrease in affinity can be ordered as satellite III>satellite Ibsatellite bmajor DNA peak (Blumenfeld et al., 1978a,b). In the embryonic cells of D. viriiis, 45% of DNA is composed of satellites, and approximately 50% of H1 histones present are phosphorylated. In the polytene chromosomes of this species, satellites and phosphorylated H1 histones constitute 1.0% and less than 10% of DNA, respectively (Billings et d., 1979). In the cells of embryos and adults of D. melanogaster, phosphorylated H1 histone is 30-40%, and this correlates also with the amount of satellite DNAs (Blumenfeld, 1979). Immunolocalization studies using antibodies specific for H4 histone isoforms acetylated at each of four N-terminal lysine residues showed that the pericentric heterochromatin is relatively enriched in the H4 isoform acetylated at lysinel2 (Turner et al., 1992).

54

1.

F. Zhimulev

The nuclear protein D1 (molecular mass 50 kDa), rich in both essential and acid amino acids, predominantly binds the AT-rich heterochromatin regions. D1 makes up 10% of the content of histone H1 in the nucleus (Rodrigues Alfageme et al., 1976,1980; Levinger and Varshavsky, 1982a,b). Protein D1 binds to AT-rich DNA in viwo, and it is the component of nucleosomes containing ATrich satellites in the native nucleus. Binding is the highest to sequence AATAT (the 1.672 satellite); it is lower to the 1.688 satellite and minimal (almost none) to AAGAG (1.705) (Levinger and Varshavsky, 1982b; Levinger, 1985a,b). An acid-soluble protein with a molecular mass of 17.3 kDa was isolated from chromatin containing satellite DNA in D. uirilis. Insofar as the 17.3-kDa protein is present in the core nucleosome, there is reason for supposing that it is a specific variant of the histone binding to satellite DNA, One molecule per 20 nucleosomes occurs in chromatin (Viglianti and Blumenfeld, 1986). A nonhistone protein specifically binding to a restricted DNA region of the 1.688 satellite is characteristic of Drosophila embryos. When a complex of this protein with DNA is formed, temperature and salt composition must be normal. However, the formed complex becomes very stable, and it is not destroyed even at very high salt concentrations (1M NaC1) or low temperatures (Hsieh and Brutlag, 197913).The latter property distinguishes this protein from D1 (Levinger and Varshavsky, 1982b). Probably the best-characterized heterochromatic associated protein in Drosophila is HPl (heterochromatic protein) (see for details sections XIIIF and

XVIIA). An in uiuo cleavage site for topoisomerase I1 has been mapped in the region where a 359-bp satellite is located (Kas and Laemmli, 1992). Additional nonhistone proteins presumably bind to satellite DNAs (Will and Bautz, 1980). Although there is no direct evidence, the assumption appears plausible when considering that the proteins are located in the regions of centromeric heterochromatin on polytene chromosomes. A comparison of the distribution profiles of the nonhistone proteins from the polytene and diploid nuclei of D . mehnogaster revealed that protein bands psi, lambda, and kuppa3 in particular are severely underrepresented in the polytene nonhistone preparations, thereby suggesting that they may preferentially bind to underreplicated heterochromatin (Elgin and Hood, 1973; Elgin et al., 1974; Elgin and Boyd, 1975). The GAGA transcription factor encoded by the Trithorax-like (Trl) gene in D.melamguster (Farkas et al., 19941, and acting as dominant enhancer of position effect variegation, binds to specific heterochromatic regions (Raffet al., 1994). These regions correspond to the sites of localization of the AAGAG and AAGAGAG satellites, according to Lohe et d. (1993). Because GAGA factor binds to GA/CT rich-elements within gene promoters, it has been suggested that the binding of GAGA factor to heterochromatin could be important for the expression of some gene located there, such as rl or kl-s (Raff et d., 1994; Lohe and Hilliker, 1995).

Polylene Chromosomes, Heterochromatin, and Position Effect Variegation

55

Taken together, the data indicate that there are considerable differences in the structures of eu- and heterochomatin. The system providing compaction of heterochromatin includes modified histones, histone-like proteins, and nonhistone proteins.

IV. GENETIC CONTENT OF HETEROCHROMATIC REGIONS OF MITOTIC CHROMOSOMES The different compaction degrees of eu- and heterochromatin prompted Heitz (1929,1932) to conclude that heterochromatin is genetically inert by analogy with the compact mitotic chromosome. However, in the early genetic studies on D. ampebphila (mehogaster), it was established that, in the case of chromosome nondisjunction in meiosis, males without the Y chromosome (XO), which are viable although sterile, appear among progeny (Bridges, 1916). Thus it was demonstrated that the heterochromatic Y chromosome carries factors responsible for fertility. Somewhat later, gene localization on the map of the mitotic X chromosome with the use of chromosomal rearrangements demonstrated that actually all the genetic map fit into the euchromatic part (Figure 31). Bobbed was the only locus mapped to the heterochromatic region of the X chromosome. Relying on these data, the heterochromatic parts of the chromosomes were thought to be genetically inert (Painter and Muller, 1929; Painter, 1931; Muller and Painter, 1932). Still later, it was demonstrated that loss of heterochromatic blocks is lethal (Schultz, 1941b) or leads to consideral derangements of morphology in adult flies (Morganet al., 1941). This was further evidence that genetic factors are present in the heterochromatin regions. Change in heterochromatin dosage produces change in the course of many genetic processes and, consequently, in the expression of phenotypes-in some quantitative traits, for example (Mather, 1941, 1944; Portin et al., 1983; Jokela and Portin, 1991): the phenotypic expression in the podoptera (GoldSchmidt, 1955), scute, Dichaete, Freckled, vestigial (Mampell, 1965b), sparkling (Morgan, 1947), and hairy (Green, 1960) mutations, the size of cells and ommatidia (Barigozzi, 1951); crossing-over frequencies (Schultz and Redfield, 1951); pairing of homologous polytene chromosomes (Gersh, 1959; Zhimulev and Vagapova, 1991); and mutation frequencies in males (Kerschner, 1949). Schultz (1956) has demonstrated that variation in the number of the Y chromosomes has an influence on the shape of the telomeres in males of D. mehogaster, the development of the ovaries in females, and the processes of DNA synthesis, although without effect on DNA content (Patterson et al., 1954). A large excess of Y chromosomes in the Drosophila genome, namely XXZY or X3Y, differentially affects gene expression and leads to various phenotypic effects: mosaicism of eye color, sterilization, irregularity of facets, shortening of leg, and abnormality of wing membrane. This was shown for many unrelated

56

1. F. Zhimulev I1

111

Mutation

Estimated size of

friqiienoy

frrquency

fragmenta

(Linknge)

(New,Ilethocl)

(Cytologicnl)

I Croesovrr

44.

*I-

--

\ \

\

\ \ \

\

\ \

\

\ \

\

Figure 31. A comparison of maps of gene location (I), distribution of mutation frequencies (]I), and a cytological map (111) of the mitotic X chromosome of Drosophila melanogasrer. Numbers at vertical lines designate position of the genes on the genetic map (I) and distribution of mutation frequencies (%) (11). Gene symbols are designated by letters. A-G, breakpoints of translocations and deletions on genetic and cytological maps. After Muller and Painter (1932).


57

strains and the Y chromosomes isolated from various strains (Cooper, 1956). In the normal genotypes, two supernumerary Y chromosomes induce mosaicism (Cooper, 1949) and sterility in males (Schultz, 1941b). In flies with a supernumerary Y chromosome, there presumably occurs an increase in the number of the histone genes in the genome (Chemyshev et al., 1980). There is no associated influence on the activity of single genes of the Y chromosome, since a decrease in the activity of tryptophan pyrrolase was not detected in v+ males without the Y chromosome when compared to XY males (Tobler et al., 1971). In w+/O males, eyes are uniformely red without any phenotypic variation (Tartof et al., 1984). Change in heterochromatin dose also has an enormous influence on the expression of position effect. The following sections present a summary of the relevant studies.

A. The X chromosome The organization of the heterochromatic regions of the X chromosome has been more thoroughly analyzed than those regions on other chromosomes (for reviews, see Sandler, 1975; Hilliker et al., 1980; Hilliker and Appels, 1982; Hilliker and Sharp, 1988, Gatti and Pimpinelli, 1992). This is because a set of inversion sites are available and also because deletions and duplications of various heterochromatic fragments can be generated by crossing over between the chromosomes and the inversions (Gershenzon, 1933a,b, 1940; Sivertzev-Dobzhanskyand Dobzhansky, 1933). The presence of rearrangements makes possible performance of genetic analysis and in situ hybridization of various DNA segments composing heterochromatin (Steffensen et al., 1981; Appels and Hilliker, 1982; Hilliker and Appels, 1982; Lifschytz and Hareven, 198213; Lindsley et al., 1982). A general scheme of the organization of the heterochromatic region of the X chromosome is given in Figure 32. The locus of the X chromosome nearest to the centromere, su(f), is presumed to lie at the eu- and heterochromatin junction (Schalet and Lefevre, 1976; Hilliker et al., 1980; Yamamotoetal., 1990).This gene is transcribed to produce a major 2.6-kb RNA and minor RNAs of 1.3 and 2.9 kb, which are present throughout development and most abundant in embryos, pupae, and adult females. The major predicted gene product is an 84-kDa protein that is a homolog of RNA14 of Sacchurornyces cerevisiae, a vital gene the mutation of which affects mRNA stability (Mitchelson et al., 1993). In heterochromatin proper, bobbed is the only vital locus (Gerschenzon, 1940; Schalet and Lefevre, 1973, 1976); it is a cluster of 150- to 250-fold repeated genes of 18s and 28s rRNAs (Ritossa et al., 1966; Tartof, 1973; Wellauer et al., 1978; Hilliker et al., 1980; Long and Dawid, 1980; Tautz et al., 1988; Hawley and Marcus, 1989). In sciarids, the nucleolus-forming region is also located in a large block of heterochromatin (Gabrusewycz-Garciaand Kleinfeld, 1966; Pardue et al., 1970; Gerbi, 1971; Gambarini and Meneghini, 1972; Pardue and Gall, 1972;

58

I. F. Zhimulev

b C

XL--

26

27 28

- ABO -

scm ste e

29

col

Rex S*X)

30 31

32

33 34

XR

w 359 bp

-. .

Figure 32. Functional sites in Drosophifu mefunogastes X chromosome heterochromatin. (a) Breakpoints of inversions (Lindsley and Zimm, 1992). (b) Cytological map of het-

erochromatic blocks (h26h34) according to differential staining (Gatti and Pimpinelli, 1992). (c) Mapping the functional sites: su(fl, cr, ABO, bb, and col (from review by Gatti and Pimpinelli, 1992); Zhr (Sawamura and Yamamoto, 1993);SCLR sequences (Nurminsky et al., 1994); Ste (Palumbo et al., 1994a); and Rex and Su(Rex) (Rasooly and Robbins, 1991). (d) Regions modifying position effect variegation (Hilliker and Sharp, 1988). (e) Regions influencingendoreplication of rDNA in salivary gland cells (Hilliker and Sharp, 1988). See text for details.

Gabrusewycz-Garcia, 1975; Dessen and Perondini, 1976, 1985; Crouse et al., 1977; Zegarelli-Schmidt and Goodman, 1981). Many 28s rRNA genes contain an insert of element I (R2) at a distance of 1.2 kb from the distal end of the gene (Glover and Hogness, 1977; Pellegrini et al., 1977; Wellauer and Dawid, 1977; Peacock et d., 1981a; Jakubczak et al., 1990; George et al., 1996), with 35% of the genes containing an insert of 4.1-6.5 kb, 16% containing another of 1.5-4.0 kb, and 14% yet another of 0-1.4 kb (Wellauer et al., 1978).Inserts of elements of this family also occur in small amounts in other regions of the genome (Dawid and Botchan, 1977; Kidd and Glover, 1980), for example, in the heterochromatin of the autosomes and in the 102C8-12 region (Peacock et al., 1981a). Insertions in the ribosomal RNA genes were also found in other dipteran species (Rae et al., 1980; Renkawitz-Pohl et al., 1980, 1981a,b; French et al., 1981; Kunz et al., 1981a)b;Beckingham, 1982; Beckingham and Thompson, 1982). The size of the cluster of genes of ribosomal RNA, together with the associated sequences and insertions, constitutes approximately 50% of the DNA of the centromeric heterochromatic region of the X chromosome (Peacock et al., 1978; Hilliker and Sharp, 1988). There are data (Appels and Hilliker, 1982) indicating that the cluster has been mapped between the breakpoints of the In(l )wm4 and In( I )wIn5lh inversions (see Figure 32). Based on cytological data, this block of genes has all the features of heterochromatin, although a small re-

Polytene Chromosomes, Heterochrornatin, and Position Effect Variegation

59

gion in late prophase seems to remain in a decondensed state, forming a secondary constriction (Hannah, 1951). A region of heterochromatin controlling the disproportionate replication of the ribosomal genes, cr (compensation response) is located between the breakpoints of the sc4 and wm4 inversions (see Figure 32) (Procunier and Tartof, 1978).In the salivary glands of XY males and XX females, the 18sand 28s rRNA genes usually constitute 0.08% of the DNA of polytene chromosomes. They make up 0.42% of DNA in diploid cells (Spear and Gall, 1973). When a homolog (XO) containing these genes or a part of the cluster is deleted, the gene number in the remaining polytene homolog is restored to 0.08%. When the cr+ region is transposed to another site of the genome, there is no compensation (Procunier and Tartof, 1978). Rex (Ribosomal exchange) and Su(Rex) elements are located within the nucleolar organizer. Rex induces mitotic exchange between two separated ribosomal DNA arrays on a single chromosome. Exchange takes place in the offspring of Rex mothers and very early, before the third mitotic division (Robbins, 1981; Swanson, 1987; Rasooly and Robbins, 1991; Robbins and Pimpinelli, 1994). Hybrid females from mating of D. simulans females to D. melanogaster males die as embryos, whereas hybrid males from the reciprocal cross die as larvae. A mutation in D. mehogaster was recovered (Zhr; see Figure 32) that rescues the hybrid females from induced embryonic lethality. The mutation is located on the X chromosome at a position near the centromere in the region covered by Dp(l ;f)I 162,not by Dp(I ;flI205. T h e latter chromosome carries a duplication of heterochromatin located distal to the In( I )sc8 heterochromatic breakpoint (Sawamura et al., 1993; Sawamura and Yamamoto, 1993). It would seem that rather large deletions have no genetic function since they are without phenotypic effect even when homozygous. However, one such region proved to be functioning. The autosomal recessive mutation abo (abnormal oocyte), which was mapped to the 31F-32E region, exerts a maternal effect; that is, it decreases the number of d o / + progeny from mating of abolabo females and +/+ males. However, an increasing proportion of abo/+ progeny can be obtained by increasing doses of a small heterochromatin fragment of the X chromosome between sc4 and wm4 (see Figure 32), which the zygote inherits from the male having, for example, attached XY chromosomes. Survival percentage was 70% for X/XY daughters from mating to abolabo, and it was only 6% for X/O males (Sandler, 1970, 1975; Mange and Sandler, 1973).These heterochromatic elements were designated as Xhaho, or ABO (Parry and Sandler, 1974; Yedvobnick et al., 1980; Pimpinelli et al., 1985). The maternal effect mutation abo is less severe when the abo strain is maintained in homozygous condition for a number of generations (Krider and Levine, 1975; Sullivan and Pimpinelli, 1986).Factors affecting i t act zygotically, dominantly, or additively. The X and the second chromosomes (but not the Y and the third chromosomes) isolated from such homozygotes for abo and transmitted

60

1.

F. Zhlmulev

by males are more effective in reducing the mutant effect when isolated from d o / + (Sullivan and Pimpinelli, 1986). ABO maps between the sc4 and wm4breakpoints (see Figure 32; see also Malva et al., 1985). Possibly, there is yet another ABO locus proximal to wm4(Hilliker and Sharp, 1988). Regions homologous to ABO are present in each arm of the Y chromosome and in heterochromatin of chromosome 2R (Sandler, 1977; Pimpinelli et al., 1985, 1986; Tomkiel et al., 1991; Palumbo et al., 1994a). Two ABO doses, one in the heterochromatin of the X chromosome and the other in that of chromosome 2R, suffice to provide normal survival of progeny of abolabo females. A twofold decrease in ABO dose decreases survival probability correspondingly (twofold). The more recent experiments suggest that simultaneous deletion of four ABO loci results in embryonic lethality (Palumbo et al., 1994a). Flies without the ABO have not been detected, this condition is lethal (Pimpinelli et al., 1986). These data indicate that the heterochromatic element abo, as received by the zygote from the father, accomplishes its effect as early as during the first cleavage divisions (i.e., much earlier than the zygotic genome starts to function); this suggests that heterochromatin contains elements functioning when the euchromatic genes are silent (Pimpinelli et al., 1986;Tomkiel et al., 1991). It was Sandler’s view (1970) that components of the abo/ABO system control the quantity of ribosomal RNA in the cells; in fact, there is a line of evidence (Krider and Levine, 1975) indicating that the amount of rRNA genes in each X chromosome increases about threefold with weakening expression of the abo phenotype in homozygotes. This increase in rDNA amount is associated with variation in the restriction pattern of nontranscribed spacers of ribosomal cistrons, presumably due to a selective increase in certain repeats in the block of the ribosomal RNA genes (Graziani et al., 1981). According to another line of evidence (Manzi et al., 1986), when the expression of the abo phenotype is weakened, the amount of rDNA and its restriction patterns remain unaltered. Molecular characterization has shown that the aba gene encodes a protein sharing homologies with the product of DETl , a negative regulator of gene expression in Arabidopsis (Tomkiel et al., 1995). Maternal mutations closely linked to abo and interacting with heterochrornatin were isolated. The mutations are dal, wdl, and hup (Sandler, 1975, 1977; Palumbo et al., 1994a). During polytenization of the chromosomes in the salivary gland cells of at least some of the strains of Drosophila the nucleolar organizer of only the sex chromosome partly polytenizes (Endow and Glover, 1979; Endow, 1980). Later studies demonstrated that the DNA responsible for this dominance of the nucleolar organizer is located between the proximal breaks of the sc4 and wm51binversions. Another function of heterochromatin, modification of the expression of position effect variegation (see Sections XI-XVI), can be mapped by portions. By generating deletions and duplications of various heterochromatin fragments, the

61


heterochromatin amount can be varied in the nucleus, and the expression of position effect can thus be affected. At least three regions of heterochromatin can act independently as modifiers of position effect variegation (see Figure 32).

B. The autosomes As a result of genetic saturation of the regions of the second chromosome deleted from centromeric heterochromatin, 1 13 mutations were identified. After complementation analysis, these were assigned to 13 loci (Figure 33), taking into account the E(SD), Rsp, It, rl, and some other loci, 18 loci in all have been identified in the heterochromatin of the second chromosome (Hilliker and Holm, 1975; Hilliker, 1976; Hilliker et al., 1980; Hilliker and Sharp, 1988; Hearn et al., 1991; Gatti and Pimpinelli, 1992; Eberl et al., 1993; Russel and Kaiser, 1994). The location of functional elements in the heterochromatin of the other chromosomes is schematically represented in Figures 34 through 36. The occurrence frequencies of genes per unit length of DNA in the heterochromatin of the second chromosome make up approximately 1% of those estimated for euchromatin (Hilliker, 1976). Several genes of the heterochromatic region have been characterized: concertina (cta), ci (cubitw interruptus), light, and rolled. The rolled gene encodes a homolog of mitogen-activated protein (MAP) kinase (Biggs e t al., 1994; Brunner et al., 1994), ci occupies about a 13.7-kb DNA region (Orenic e t al., 1990; Locke

3536 37

a

2L

-- Z R

4OFEta 4OFc It 4OFd msQ)H1 dnF~

b

383941 4243 44 45 4 6

--

2.L --

41Ad

4ifi 4iia rl

--

-5, 8 470

4T-h 41Ah

ZR

6

423

1, 11

Figure 33. Functional sites in Drosophila melanogaster second chromosome heterochromatin. (a) Functional sites are situated above and below the heterochromatin map (h35-h46 according to Dimitri, 1991). After Hearn et al. (1991), Gatti and Pimpinelli (1992), Eberl rt al. (1993),and Russell and Kaiser (1994). See text for details. (h) Location of P-element insertions in heterochromatic regions. After Zhang and Spradling (1995).

62

I. F. Zhimulev

9-52 5-84

10-39

a

'-'16

1-16 10-58

8A-80

b

80Fa

2-66

-

l80FclSOFe

9-56

3-9

-

I 80~11

8OFj

4748495051 52

3L

C

47 10-65

81Fa 81F1, 5455 56 57

53

--

58 --

A7148

336

5

d

3R

1

ce&mere

33Jir

Figure 34. Genetic and cytological maps of Drosophila melanogaster third chromosome heterochromatin. (a and b) Mapping the genetic loci (b) within limits of deficiencies (a). After Marchant and Holm (l988a,b). (c) Map of heterochromatin blocks (h47-h58). After Gatti and Pimpinelli ( 1992). The correspondence between the genes and heterochromatin blocks has not been determined. (d) Insertions of P-elements (Zhang and Spradling, 1995).

ci ....

59 60 61

4R

4dTJmlQ---

a

ABO

1 2 3 4 5 678

b C

d

YL,

9

Figure 35. Map of fourth chromosome heterochromatin (h59-h61). After Gatti and Pimpinelli (1992). Tentative location of ci is after Lindsley and Zimm (1992).

-

cry -

MstlY)

AEO

-

10 1 1 1 2 1 3 1 4

15

1 6 1 7 18 19

col bb 20

21 22 23 24 25

YS

[SyKa

k

-

kl-5 kl-5 A teloiiiere

kl-3 kl-3

kl-2

ks-I

kl-I

B

ks-I C

centrzmere

10 16 c2: c3

-

364

121

4

I

. .

.

ks-2

telokere

95-2

-~ 2.12. 13.17.

512

c8 302 Figure 36. The Y chromosome map of Drosophila melanogaster showing the localization of the heterochromatic functional elements. (a) Genetic sites. (b) Heterochromatic blocks (hlLh25). (c) Thick and thin lines show minimal and maximal limits of fertility factors. A, B, C, loop-forming sites of kl-5, kl-3, and k s - I , respectively. (d) P element insertions (Zhang and Spradling, 1995). After Pimpinelli et al. (1985, 1986),Toinkiel et al. (19911, Gatti and Pimpinelli (1992) and Russell and Kaiser (1993).


63

and Tartof, 1994), and cta encodes the a-subunit of a G-protein playing a role in cell-cell communication during embryonic development (Parks and Wieshaus, 1991). The length of the light gene is at least 17 kb (Devlin et al., 1990a,b). Of particular interest is the system of SD-Rsp (see Lyttle, 1991, 1993; Teminetd., 1991; Palumboet al., 1994a; Palopolietal., 1996,for review). The point is that, in the 37DZ-D6 region of one of the second chromosomes isolated from a population by Y. Hiraizumi in 1956, the SD (Segregation Distortion) factor was disclosed (Sandler et al., 1959). Its presence in SdlSd' males leads to a substantial increase in the number of gametes with the Sd mutation compared to the Sd' gametes. Electron microscopic studies have demonstrated that spermiogenesis of Sd' gametes in SdlSd' genotypes is aberrant due to decompaction of chromatin (Sandler et al., 1959; Hartl et al., 1967; Hartl and Hiraizumi, 1976; Tokuyasu er al., 1977; Sandler and Golic, 1985). Segregation distortion is due to the interaction of three elements: Sd itself; Rsp (Responder)and E(Sd) (EnhancerofSd). Two major Rspalleles are known: RspS (sensitive) and Rspi (insensitive). In strains with distorted segregation, the chromosomes have the constitution Sd, Rsp'lSd', RspS (Figure 37). The condition providing aberrant spermiogenesis is the presence of allele SD and heteroallelism in Rsp (Ganetzky, 1977; Brittnacher and Ganetzky, 1983; Sharp et al., 1985). SD chromosomes also carry several linked drive enhancers, such as M(SD) and

sd

SD +

Rsp-i

t sd'

RSP-s

Figure 37. Schematic representation of SD complex. Sd and Sd', distorting and nondistorting alleles of Sd locus, respectively; Rsp-i and Rsps, insensitive and sensitive alleles of the Rsp locus, respectively. Sd renders Rsp-s-bearing sperm nonfunctional during spermatogenesis (postmeiotically). After Doshi et al. (1991).

64

I. F. Zhimulev

S t ( S D ) (Hiraizumi et al., 1980, 1994; Hiraizumi, 1990; Lyttle, 1991). E(SD) not only enhances the action of Sd, but also independently behaves as a segregation distorter (Sharp et al., 1985; Temin, 1991). Based on cytogenetic and genetic data, the Rsp locus has been mapped to the h35 block of the 2R heterochromatin (see Figure 33), and Sd to the euchromatic part of chromosome 2L (the 37D2-D6 region). The third component, E(Sd), was mapped to the heterochromatic part of the 2L heterochromatin (Ganetzky, 1977; Hiraizumi, 1981; Brittnacher and Ganetzky, 1983, 1989; Sharp et al., 1985;Lyttle, 1989; Pimpinelli and Dimitri, 1989). A cluster of Rsp sequences is also located in the third chromosome (80C region) (Moschetti et ul., 1996). The Rsp locus presumably is a region of reiterated DNA with repeating units of 120-bpfragments. These fragments are enriched in AT, with the general structure of satellite DNA, and organized mainly as dimers consisting of two 120bp repeats with dimers delineated by TCTAGA sequences (the Xba I restriction site) at each end. There are central TCTACA sequences that are less frequently cut by Xba I to result in 120-bp monomers (Wu et al., 1988). The repeats usually have a dimeric structure with an average difference of 16-20% between the left and right halves (Cabot et al., 1993; Lyttle, 1993). Rsp repeats isolated from the same chromosome are not more similar than those from different chromosomes. The whole Rsp locus extends over a region of 600 kb on a standard sensitive (Rsps) chromosome. Within the region, Rsp repeat arrays are interspersed with non-Rsp sequences and account for 10-20% of all sequences (Cabot et al., 1993). The XbaI repeat of Rsp exhibits DNA curvature; nucleosomes containing this DNA are spaced at 240 bp (the size of the dimeric repeated unit) rather

Figure 38. Variation of copy numbers of the Rsp repeat in Drosophila melanogasm stocks differing in Rsp sensitivity. Restriction Xba I digest of DNA from supersensitive (lpcb), sensitive (a bw), and partially sensitive (Canton S) stocks, probed with the DNA Ho clone containing XbaI repeated unit. After Wu et d . (1988).


65

than the 190 bp for bulk DNA. Three DNA-nonhistone protein complexes were identified by means of gel-shift assays involving protein extracts from pupal nuclei combined with labeled Rsp DNA (Doshi et al., 1991). The Sd chromosome has a deletion of the Rsp region (Rsp-) and less than 20 copies of a Rsp-associated 120-bp satellite repeat. Sd' chromosomes sensitive to Sd action carry Rsp" with more than 700 repeats (Figure 38). Insensitive Sd' (Rsp') chromosomes carry forked

X: 56.7; 15F1-3

Bristle shape

klgovsky (1938,1944,1946). Noujdin (1946d)

fa,facet

X:3.0; interband 3 C 7 4

Eye structure, wing shape Microchaetae number Microchaetae and bristles

Demerec (1940.1941a1, &hen (1962) Dubinin and Sidorov (1935). Jeffery (1979)

h, harry

3: 26.5; 66D15

Hw, Hairy wings

X: 0.0; 1B1-2

kz, kurr 1( 1)BAl

X: 0.9; 2 U X; 1F3 to 2Al-2 X; 1F3 to 2A1-2 X; 1F3 to 2A1-2 X; 2A3 to 2B3-4 X; 2A3 to 2 8 3 4

l(l)BA5 I( 1)BA9 l(1)BAll 1( 1)BAl2 1(3)S12 m, miniature

mi, minus

Mot-K, mottled 01 Krivshenko N, Notch pic*piccolo PnP

prune

Alikhanyan (1937), Crew and Lamy (1940) Bridges and Brehme (1944). Getsh (1949) Schulu in Gvozdev et al. (1973)

Bristle shape Eye shape

Demakova et al. ( 1988)

Bristle size Bristle shape

Demakova et al. (1988) Demakova et al. (1988)

Wing shape Size of eye, wings

Demakova et d.(1988) Demakova et al. (1988)

3R; 52,87DF

Bristle morphology

Dutton and Chovnick ( 1991)

X: 36.1; 10E1-2 2: 104.7; 5 9 K E 4

Wing shape

Wargent (1971), Spofford (1982)

Size of body and bristles Eye color

Schultz and Dobzhansky (1934) Krivshenko (1954), Cooper (1956)

Wing morphology

Demerec (1941a,b)

3R: 52.1; 87D11-14

Bristle shape

X: 0.8; 2E2-3

Eye color

Clark and Chovnick (1986) Gerasimova et al. (1972), Alatortsev et al. (1982), Alatomev (1986, 1988)

Related to translocation T(2;3)41

X: 3.0; 3C7

rg, rugose

X: 11.0; 4E1-3

Eye structure

Demerec (1940)

TO, TOUgh

3: 91.1 97D1-9

x: 2,2; 3c5

Eye structure Eye structure

Brosseau (1970)

rst, roughest TUX, TOUgheX

X: 15.0; 5C5-D6

Eye structure

Dernerec (1940)

rY, TOSY

3: 52.0; 8 7 ~ a i 2

Eye color

Rushlow and Chovnick (1981)

Sb, Stubble

3: 58.2; 89B9-10

Bristles

Moore et al. (1981), Sinclair et al. (19891, Hayashi et al. (1990)

sc, s a t e

X: 0.0; 1B3

Microchaetae and bristles

Reviews: Bridges and Brehrne (1944), Lindsley and Grell (1968)

snk, snuke

3: 52.1; 8 7 W 1 3 X: 3.0; 3C7 X: 0.3; 2143-4 to 2B3-5 X; 2B6 X: 33.0; 10Al-2

Gene with maternal effect

Clark and Chovnick (1986)

Structure of eyes, bristles

Schultz (1941b), Cohen (1962)

spl, split stn, stubarisra

swi, singed wings

Dernerec and Slizynska (1937), Gruneberg (1937), Kaufmann (1942),Cohen (1962)

Arista shape

Demakova et al. (1988)

Wing structure

Zhimulev et al.( 1986), Demakova et al. (1988)

Eye color

Tobler et al. (1968, 1971), Lefevre (1969)

Wing morphology

Wargent (1972)

vs, eresindated

2: 67.0; 49D2-El X: 16.3; 5D34A2

Wing structure

Demerec (1940)

w, white

X: 1.5; 3C2

Eye color

Muller (19301, Gowen and Gay (1934), Dernerec and Slizynska (1937), Gersh (1963). Spofford (1982), and many others

wap, wings apart

X: 0.06

Wing position

Alatortsev et al. (1982)

X: 0.0; 1BI

Body color

Noujdin (1935, 1936a,b, 1944), Sidorov (1936), Brunstrom (1955), Zhang and Spradling (1993)

v , vermilion

vg, uestigd

(Alatortsev et al., 1982) y, yellow

316

1. F. Zhimulev

(Figure 126A). The eye develops as a part of the head anlage, already recognizable at the end of the embryonic stage. Two anlagen (one on either side), with the exception of the buccal parts, form the whole head of the fly (Becker, 1966). An eye consists of 25,000 cells organized in a complex manner into 800 radial structures, whose one end is connected with the optic nerve while the other terminates at the eye surface as hexagonal facets. The normal color of the ommatidium, and consequently of the eye, is red; when numerous genes controlling the synthesis of eye pigments are inactivated, the eye becomes vermilion, yellow, brown, and other colors; when synthesis and assembly of pigment are completely suppressed, a white eye is formed. For this reason, when genes for eye color are inactivated in stem cells from which other cells later derive, the formation pattern of mosaic patches can be followed; there appear white or colored patches on the background of ommatidia of the normal red color. Genetic inactivation in malpighian tubules is manifested by the appearance of colorless cells on the background of cells containing yellow pigment. The colorless state may be related to the position effect of white+ (Demerec and Slizynska, 1937; Schultz, 1941b; Hartmann-Goldstein, 1967), dor+ (Ananiev and Gvozdev, 1974), or ry+ genes (Rushlow et al., 1984; Daniels et al., 1986).

2. Vital genes If a gene that becomes lethal when inactivated comes to lie in the immediate vicinity of a rearrangement breakpoint, then there is no corresponding class of flies bearing the chromosomal rearrangement because of position effect. The rearrangement breakpoint can serve as an indication of the localization of the inactivated vital genes. Schultz (1941b) discovered that the presence of an additional Y chromosome in the genome reduces lethality; the chromosome with the T(l ;4)wuD3 translocation is not lethal in the homozygote in females and in the hemizygote in XY males; however, it becomes so in XO males. Mass experiments designed to generate Y-suppressed lethals in the X chromosome demonstrated that about bh of such mutations are due to damage of the nucleolar organizer, which is suppressed by the ribosomal EWA genes of the Y chromosome, and gths of the mutations are caused by chromosomal rearrangements producing position effect variegation (Lindsley and Edington, 1957; Lindsley et al., 1960; Traut, 1966; Borisov, 1972). The vital genes involved in inactivation are located in various chromosome regions (Schultz and Catcheside, 1937; Schultz, 1941b; Ratty, 1954; Lindsley etal., 1960; Baker, 1971;Lefevre and Green, 1972;Judd et al., 1972; Barr, 1973; Lefevre, 1981; Alatortsev et al., 1982; Spofford, 1982; Tolchkov et al., 1984; Dimitri and Pisano, 1985; Lefevre and Watkins, 1986; Zhimulev et al., 1986; Demakova et al., 1988). The rearrangement breakpoints causing the Y chromosomesuppressed lethal position effect (Lindsley et al., 1960) are rather uniformly distributed along the entire extent of the X chromosome (Figure 127). Several


317

B

C c 2

3 a

12 13

I

'8

Figure 126. Structure of the compound eye of Drosophila. (A) General appearance of the eye, with hexagonal-shaped facets. (B) Scheme of the disposition of the structures of the optical system. Numbers designate the number of rhabdomeres (rh). ( C ) Scheme of the longitudinal section of a single ommatidium. Dashed lines indicate the level and structure of the transverse sections through the ommatidium of Drosophila. 1, Cornea; 2, bristle; 3, pseudocone; 4, primary pigment cells; 5, secondary pigment cells; 6, cells of the pseudocone; 7, cells of the retinula; 8, rhabdomere; 9, base membrane; 10, postretinal bundles; 1I , postretinal pigment cells; 12, external granular layer; 13,external optic ganglion; 14, nuclei of retinula cells; 15, nuclei of a retinula cell of the seventh rhabdomere; 16, cells of the retinula of the eighth rhabdomere; 17, pigment granules; 18, pigment granules of the secondary pigment cells. (A) after Casteel (1929); (B) after Pak and Grabowski (1978); (C) after Gersh (1952).

318

1. F. Zhimulev

X CHROMOSOME [ f l I I 1 1 I I I I I 1 I I 1 1 I I 1 bd

CHROMOSOME2 hl I I I I I I I I I I 1 I I 1 1 1 I bdb1 I 1 1 1 1 1 1 1 1 I 1 1 1 1 I 1 1 CHROMOSOME3 bl I I 1 1 I I 1 1 1 1 I I I 1 1 I I W i I I I I I 1 1 1 1 1 1 I I 1 1 1 1 1104 CHROMOSOME 4

+IEH

Sfo

Figure 127. Location of breakpoints in rearrangements in the X chromosome of Dosophila whose lethal effect is suppressed by the Y chromosome. A,Inversions in the X chromosome and ringXCchromosome;0,T(J;2);0,T(1;3);A,T(1;4). AfterLindsleyetal. (1960).

other regions of the X chromosome are mentioned in Spofford's (1976) review along with references to personal communications: to the right of sw(wa)+ (R. E. Rayle and M. M. Green), near dor+ (J. C. Lucchesi and M. L. Bischoff), in the zeste+-white+ interval (T. C. Kaufmann), and near ras+ and v+ (Barr). Ben-Zeev and Falk (1966) reported that Y-suppressed lethals could not be induced in the second chromosome. It is Spofford's (1976) view that the failure is due not to the real absence of such rearrangements in the second chromosome, but rather to technical inadequacies of the applied methods.

3. Loci encoding the products identified by biochemical methods In the case when position effect variegation induces inactivation of loci encoding known proteins, the amount of antibodies bound to the gene product, the electrophoretic fraction (Figure 128) encoded by the rearranged chromosome, or the activity of the enzyme in the extract detected by specific staining is decreased. Such loci are listed in Table 27.

4. The genes in transposons A DNA fragment containing the genes under study can be introduced into the Drosophila genome via P-element-mediated transformation. For example, when


319

Figure 128. Electrophoretic detection of the activity of 6-phosphogluconate dehydrogenase in the homogenates oforgans and whole flies with rearrangementspn2, pn3, TE100, and TElOl (A) and organs or larvae with pn2 rearrangement (B). f, whole fly; h, head; a, abdomen; ov, ovary; I, larva; fb,fat body; sg, salivary gland; id, imaginal discs; ng, neural ganglion. After Slobodyanyuk and Serov (1987).

the R401.1 (strain ry42) transposon containing the rosy+ gene is inserted into heterochromatin of the fourth chromosome, rosy+ is partially inactivated; the extent of inactivation can be modified by varying the amount of heterochromatin in the Y chromosome. XO individuals contain much less xanthine dehydrogenase (the product of the rosy gene) than XXY individuals (Spradling and Rubin, 1983; Daniels et al., 1986). Taken together, the considered data show that actually any euchromatic region of the Drosophila genome can undergo genetic inactivation as the result of position effect variegation.

C. Variegation of inactivation The mosaic pattern of genetic inactivation known as position effect variegation may be evidence for a mechanism preferentially inactivating a gene in a cell group

320

I. F. Zhimulev

Table 27. Position Effect of Genes with Known Biochemical Product Symbol of gene, encoded product

References

Location

Adh, alcohol dehydrogenase Amy, a-amylase Acph-I, acid phosphatase

2R: 54555

Bahn (1971)

3R

Frisardi and Maclntyre (1984), Shaffer and MacIntyre (1990)

ry,xanthine dehydrogenase

3R: 52.0; 87DF

Rushlow and Chovnick (1981,1984), Spradling and Rubin (1983), Clark and Chovnick (1986)

bb, ribosomal RNA

X: 66.0; 2OC2-?

Baker (1971), Nix (1973), Zuchowski-Berg (1978)

Sgs4, protein of salivary

X; 3Cll-12

Komher and Kauffman (1986)

pr, sepia-pterine synthetase

2: 54.5; 37B2-40B2

Tobler et d.(1979)

w , tryptophan oxygenase

X: 33.0; 10A1-2 X; 0.65; 2Dl-6

2L

Hisey et al. (1979)

gland secretion

Pgd, phosphogluconate dehydrogenase

Tobler et al. (1968) Gerasimova and Ananiev (1972), Gerasimova et d.(1972), Gvozdev et al. (1973,19741, Alatortsev et d.(19821, Slobodyanyuk (1982,1983), Slobodyanyuk and Serov (1983, 1984,19871, Tolchkov et aE. (1984), Tolchkov and Gvozdev (1984)

and maintaining it in this state. Consequently, study of mosaic gene inactivation would be helpful not only in resolving the broad issues of developmental genetics (see the pioneering studies ofNoujdin, 1936a,b, l945,1946b), but also in providing insight into the more specific mechanisms of genetic inactivation under position effect. Mosaicism has been proved to be tissue and ontogenetically specific. The specificity is manifest as considerable differences in the proportion of tissue cells in which a gene(s) in a rearrangement has become inactivated. This specificity may result from (1) variegation of the inactivation of stem cells giving rise to organs or (2) differences in the rate and frequency of subsequent division of stem cells in which a gene has become inactivated or not under position effect. Differences in the intensity of transcription and posttranscriptional control are conceivable. Nevertheless, the disclosed specificity does exist, and there is reason to consider the pertinent facts. Study of the distribution of staining for the activity of the enzyme 6phosphogluconate dehydrogenase (6-PGD) in organs of larvae and adults of


32 1

Drosophila revealed that the degree of mosaicism under position effect in the pn2 and pn3 rearrangements (see Ilyina et al., 1980) was higher in adults (the percentage of cells with the inactive 6-Pgd’ gene was estimated as 70-80%) than in larvae (50%). Genetic inactivation was high in neural ganglia, imaginal discs, and hindgut but low in salivary glands, fat bodies, and malpighian tubules. None of the studied rearrangements gave rise to inactivation in the ovaries, although inactivation was encountered in cells of male gonads. However, inactivation of the Pgd’ gene in the TEJOO rearrangement showed neither tissue nor adult (larvaeadult) specificity (Slobodyanuk, 1983; Slobodyanyuk and Serov, 1987). The proportion of cells with the inactivated w+ gene in the compound eye ofD. melanogaster is greater than in testis sheath (Hessler, 1961).The w+ gene is also differently inactivated in D. hydei. While the gene is inactivated in single eye cells in females, the cells of malpighian tubules are completely wild type (van Breugel, 1970). Even in different types of cells in ommatidia, there are different possibilities for the w+ gene becoming inactivated in the T(I ;4)wm258-J8rearrangement. Some of the secondary pigmented cells (see Figure 126) and the majority of postretinal cells can be either completely pigmented, pigmentless, or intermediate. Primary pigmented cell can only be in the pigmented or unpigmented state (Gersh, 1952). Noujdin (1936a,b) noted that, when the yellow gene is variegated in the In( 1 )s8rearrangement, the occurrence probability of yellow bristles is related to the gene’s location in the mesanotum. There is, however, ample evidence indicating that a gene is inactivated in a correlated manner under position effect in cells of different types. It was demonstrated for T( 1;4)wmz58-z1/wthat, when much more than one-half of the area of the eye is of a mutant color, about one-half of the cells of malpighian tubules are also unpigmented (Schultz, 1956). The proportion of salivary gland cells with strongly expressed “heterochromatization” (see Section XV1,C) and that of unpigmented cells of malpighian tubules or eyes were found to be close (Hartmann-Goldstein, 1967; Ananiev and Gvozdev, 1974; Hartmann-Goldstein and Koliantz, 1981; Koliantz et al., 1984). An interesting pattern that was neither confirmed nor refuted later was disclosed by Noujdin (1946a,d, 1947). In studies of variegating gene expression in the chromosomal rearrangements In( I )sc8, PlumD1,f ~ r k e d ~ ’and ~ , yellow”’, he found that, despite differences in the occurrence frequencies of mosaics (ranging from 1% to loo%), their appearance was not of a random nature. The frequencies of mosaicism fall into a series of groups with the same values. Offspring from different matings could be referred to the same group. The frequencies of mosaics within a group, as a rule, did not differ from each other. Groups of the same type were observed among males and females. The mosaicism frequency differencesbetween consecutive groups are multiples of 2, with the result that the whole series

322

1. F. Zhlmulev

of frequencies was a geometrical progression with an index of approximately 2; for example, the following frequencieswere obtained for the sP inversion: 1.32,5.35, 13.16, 26.61, 42.83, and 98.60. The effect of factors modifying the expression of position effect is detected in all cells expressing the normal allele, although not consistently in the same manner. Thus modifiers enhancing the activation of the w+ locus in eye cells also enhance it in testis sheath and malpighian tubules, although there was no direct correlation between the two events (Schultz, 1956; Hessler, 1961). As noted earlier, maintenance of genetic inactivation through successive generations is simplest to observe in groups of the same cells or in the same cell structure. A number of authors (e.g., Surrarrer, 1935; Gersh, 1952; Becker, 1960, 1961) noted that the white patches indicating gene inactivation are not randomly distributed over the area of the eye, being most frequently located in its anteriormost section. The occurrence of pigmented ommatidia in D. melanogaster gradually decreases in the anterior-posterior direction. A similar clonality was observed for D. uin'lis (Baker, 1967) and D. hydei (van Breugel, 1970, 1972). This gradient is not seen or is lacking in the Bar mutants of D. hydei (van Breugel, 1972). According to other data, unpigmented cells occur with the highest frequency in the middle part of the eye (Koliantz et al., 1984). How can this gradient form? To understand events due to position effect, the main data concerning the formation of eye mosaics resulting from mitotic recombination should be considered. When w/wco heterozygotes are irradiated (eye colors: w, white, wco,coral, w/wco, rosy), mitotic crossing over (Figure 129) takes place and mosaic spots arise because regions composed of wlwco, wco/wco,and wlw cells are formed. Most mosaic eyes have twin white and coral spots, because flecks with recombinant chromosomes (w/w and wco/wcoin Figure 129) are derived from neighboring sister cells. Similar cell clones appear when other markers (e.g., zeste) are used (Becker, 1956a-c, 1957,1960,1961,1965,1966,1969). After irradiation of larvae at early developmental stages, a few, albeit large, spots are formed; after irradiation at later stages, the spots are smaller and more numerous, however, Even the largest spots do not cover the whole eye because the horizontal line passing through the middle of the eye usually forms the boundary of the spot. In 17 of 23 spots 220 ommatidia in size, this horizontal line was the upper boundary in the lower part of the eye. The remaining 6 spots extended over both halves of the eye. In addition to the central dividing line, there are others lines that are particularly conspicuous in the lower part of the eye. The positions of these lines are brought into more prominence by supercomposition of the contours of the mosaic spots (Figure 130a). The boundaries of the spots are concentrated in certain regions. They divide the lower part of the eye into sectors (from I to VIII) about 40 ommatidia in size each (Figure 130b). When larvae molting from the first to the second instar

a

\?/

W

0

X-RAY INDUCED

b TWIN SPOT

Figure 129. A scheme of the arising of cell clones of different genotypes (b) in the compound eye of D. melanogasrer ( c ) after radiation induction of somatic crossing over (a). White, gray, and black circles designate the genotypes w/w, w/w'", and afo/wco,respectively. (a and b) after Baker (1965~);(c) after Becker (1957, 1966).

324

I. F. Zhimulev

Figure 130. Location of boundaries of mosaic spots on the lower half of the eye of D. melanogaster (a) and a schematic representation (b) of the most frequently occurring types (Roman numerals) amd location (continuous lines) of spots; boundaries of spots occurring less frequently are represented by dashed lines. After Becker

(1957, 1960).

(48 hr after egg laying) are irradiated, such spots occur. Inasmuch as the whole eye consists of 800 ommatidia, and a sector includes approximately 40, it may be conceded that each cell of the eye anlage is predetermined for the formation of a region of the adult eye at the time of irradiation. Sectors of two- or threefold greater sizes can include neighboring sectors in any combination (I-VIII). This scheme clearly shows that the development of the eye proceeds from the tail to the head. A similar clonality of somatic crossovers was found in analysis of the zestem mutants in Drosophila mehogaster (Becker, 1966) and D. virilis (Baker, 1953). The formation of the eye in the dominant LobeB mutant of D. melanogaster supports the notion of clonality (Becker, 1957). At 18"C, only the upper half of the eye develops and its lower half lacks many fragments, and these sites are overstrewn with cuticule (Figure 131). The smallest missing sectors remarkably resemble the eye sectors found during mitotic recombination. Probably, the mutation arrests development of certain cell clones that may truly exist and become detectable during mitotic recombination (Becker, 1966). Thus the general conclusion can be made that the region of the eye that is of normal color and its unpigmented regions are derived from single cells whose capacity (or incapacity) to synthesize pigment is established before the eye has been formed. The process of the formation of clones of daughter cells, determined in the same way as the initial initiator cell, was called clonal initiation (Gsell, 1971).


325

Figure 13 1. Disturbance of eye morphology in L8 mutants of Drosophila. Development of B/B+ and LEI+ genotypes at 18°C (a-f), at 25°C (g), and at 18°C (h).After Becker, 1957).

In Drosophila carrying the T(l ;4)wm258-18/~ genotype, and when mosaics are formed under position effect, the pattern of red and white sectors in the eye is identical with that produced by sectors I-VIII in Figure 128B (Becker, 1961, 1966, 1978;Baker, 1963, 1965a, 1967). This similarity, in the authors’ view, may indicate that gene inactivation caused by position effect takes place also at the end of the first larval instar. Then, once inactivated, the gene remains in this state through successive generations, presumably owing to epigenetic factors (Baker, 1965b,c; Khesin and Leibovitch, 1976). When a genital imaginal disc mosaic for groups of y and y+ cells is transplanted into the abdomens of adult flies and then retransplanted to another female after 2 weeks for a total of 23 transplantations, and thereafter the imaginal disc is transplanted to a larva, imaginal organs variegate after metamorphosis. This has been taken to mean that the inactivated state is retained for a long time (Hadorn et al., 1970; Gsell, 1971). Experiments with transplantation of imaginal discs demonstrated that the inactivated state of a gene is determined autonomously (Janning, 1971). From the results of thermal treatments, it follows that inactivation is fixed, even irreversible. When Drosophila are treated with low temperatures at the early stage of embryonic development, maximum mosaicism is achieved and subsequent development at higher temperatures does not lead to reactivation of the once-inactivated genes (Schultz, 1956; Hartmann-Goldstein, 1967; Zhimulev et

326

1. F. Zhimulev

al., 1988). Irreversible inactivation seems to be an autonomous property. Transplantation of imaginal discs from w; Dp( J ;3)N2“-58 to C(I)RM, yw larvae with or without the Y chromosome demonstrated that the amount of eye pigment in the eye derived from the transplants was the same in both cases; that is, the effect of the Y chromosome did not intersect the boundaries of the imaginal disc (Gearhart and MacIntyre, 1971). However, the conclusions regarding clonal inactivation under position effect do not seem to consistently hold true, because they disagree with certain facts: 1. In D. hydei, certain rearrangements also show clonal inactivation, however, the mosaic pattern is not identical to the one Becker and Baker described for D. melanogaster and D . virilis. For example, if the shape of large spots produced by the wm COYrearrangement still fits into the scheme, the small spots arising when the w+ gene is inactivated in the wm2 rearrangement are scattered over the whole eye. Yet another type of unusual spots was described for the R(Y)wmrearrangement. Spots of white cells have the appearance of incomplete sectors; that is, starting from the posterior-most section of the eye, they gradually decrease in size without reaching the anterior-most section, being substituted by unpigmented cells. These data indicated that the w+ gene is additionally inactivated in an increasing number of cells with each successive cell division (Beck et al., 1979). In D. melanoguster, groups of unpigmented ommatidia lie surrounded by pigmented ommatidia in the middle part of the eye (Casteel, 1929), which is evidence of late inactivation of the white gene. 2. The clonal pattern of inactivation is possibly tissue specific. If, indeed, clonality is not a rarity during eye formation, as follows from the previous description, it is unclear whether clonal inactivation takes place in malpighian tubules. These organs are represented by a quasi-linear series of cells, and each generation of yellow (w+)and unpigmented or white ( w ) cells is easily distinguished from neighboring cells, allowing a determination of whether clusters of inactivated cells are present. Experiments performed in D. hydei showed that inactivated cells appear randomly, the mean cell number in a “clone” being 1.1 (Gloor et al., 1967; van Breugel 1973; Beck et al., 1979). Nevertheless, a gradient of gene inactivation in malpighian tubules was found both in D. hydei (van Breugel, 1973), and D. melanogaster (HartmannGoldstein and Koliantz, 1981). 3. Certain data obtained with mosaics of Dosophila also provide evidence for reactivation of the inactivated genes. For example, findings of accumulations of pigmented cells on a white background, their occurrence frequencies unknown, have been described in a series of papers (Muller, 1930; Surrarrer, 1935; Demerec and Slizynska, 1937; Spofford, 1976; Block et d., 1990).


327

Spofford (1 976, p. 978) provides a description of a case when pigment was deposited at various positions along the axis of scutellar bristles in In( J )y3p males, thereby suggesting alternation of act ivat ion-inact ivation stages. 4. There is a striking discrepancy between the described clonal inactivation spreading from the posterior to the anterior quarters of the eye and variegated eye pigmentation in transformants. When a transposon containing the w+ gene was inserted into the w mutant genome, w+ was normally expressed in more than 20 different locations (Hazelrigg et al., 1984; Gehring et al., 1984; Rubin et al., 1985; Sun et al., 1995). Flies of the w+ A 4 4 strain [the transposon inserted in the telomeric (100F) region of chromosome 3R] had a uniform yellow eye when grown at 25°C; however, red ( !) spots appeared on a yellow background when grown at 18°C. This pigmentation pattern was more unusual than the one displayed by AR4-3 flies (insertion in the 39E-40F region): the posterior part of the eye, the region of clonal initiation according to Becker, was white, whereas its anterior part had a normal color (Hazelrigg et al., 1984). When the AR4-3 transposon relocated to occupy one of its new positions (the 24CD region; strain AR4-24), there was a concomitant very unusual change in gene activity with the result that the inactivation sector of eye cells seemingly turned around through 90 degrees. The ventral part of the AR4-24 fly eye was darker than the dorsal part (Levis et al., 1985). 5. Along with clonality, in another mosaicism type the inactivated gene w is tiny, possibly even the size of an ommatidium. These clones are surrounded by ommatidia of normal color. As a result, pigmented and pigmentless cells are mixed in small groups in the eye (“pepper-and-salt”) (Morgan et al., 1937; Schultz, 1941b; Gans, 1953; Becker, 1960,1961,1966; van Breugel, 1970; Koliantz et al., 1984; Demakova et at., 1988). In D. hydei, mosaic types of eyes in turnmutants are specific to the various rearrangements: those with breakpoints to the right of the w+ gene show inactivation of the sector type; when the breakpoint is to the left, they have the “pepper and salt” appearance (van Breugel, 1970). This is a line of evidence indicating that the X chromosome is without influence on the expression of the “pepper and salt” variegated type (Gans, 1953). According to other information, the supernumerary chromosome suppresses mosaicism and the eyes are much lighter in XO than XY males (van Breugel, 1970). Low temperature (16°C)produces the same effect as the supernumerary Y chromosome, that is, opposite to the one it exerts on sector-type inactivation (van Breugel, 1970).

D. levels of inactivation The genetic inactivation caused by position effect is manifested in tissues in which the normal allele of the inactivated gene functions. Judgments concerning gene

328

1. F. Zhimulev

activity in tissues can be based on the gene’s RNA or protein products. Protein can be visualized using either a specific histochemical stain for its activity or antibodies against it. Belgovsky (1944) suggested “that biochemical activity of the gene is attenuated under position effect with the result that the amount of its product approaches the critical minimum required for the normal development of a character.” The question is, at what level does attenuation take place? The following factors cause development of the mutant phenotype: (1) a smaller number of cells are formed under position effect; (2) a protein is not synthesized, there is a decrease in its intensity, or, as Fuscaldo and Fox (1962) suggest, a defective protein is synthesized that is devoid of biochemical activity yet retaining antigenic determinants shared in common with native synthesized protein; (3) the RNA product of the gene is not synthesized; and (4) antisense RNA is synthesized (Frankham, 1988). How real is the decrease in cell number in target tissues under position effect? The decrease is possible when inactivation is addressed to an essential gene, whose mutation would cause cell death. From studies in which mosaicism was easily demonstrable (e.g., in the eye of Drosophila), it generally follows that the organ does not become smaller; it is merely that a gene (e.g., w’) is inactivated in all the cells. Histochemical staining for enzyme activity in some organs (Alatortzev et al., 1982; Slobodyanyuk and Serov, 1983; Tolchkov et al., 1984; Rushlow et al., 1984) also reveals mosaicism for cells without an associated decrease in organ size. Synthesis of the gene products-for example, proteins and pigments imparting color to eye and malpighian tubules, as Morgan and others noted (Morgan et al., 1937; Clark and Chovnick, 1986)-was consistent with the “all-ornone” pattern of gene expression; in the case of mosaicism for the white gene, only two states were observed for malpighian tubules: completely pigmented (‘‘all”) and white pigmentless (“none”) cells. No deviation in cell organization was due to position effect in pupae whose chromosomes bear T ( 1 ; 4 ) ~the ~~ struc~~ ture of unpigmented (w) eye cells was of wild type (w+) or of the type observed in homozygotes for bw and q, mutants also producing the w phenotype. Only pigments were lacking (Shoup, 1966). When cells of the fat body were stained with a specific stain to vizualize 6-Pgd activity, variegated staining of chromosomes bearing rearrangements was also consistent with the all-or-none pattern of changes in gene expression; that is, the chromosomes were without gradual transitions from complete absence to heavy staining (Alatortzev et al., 1982; Slobodyanyuk, 1983; Slobodyanyuk and Serov, 1984, 1987; Tolchkov et al., 1984). However, caution should be exercised when interpreting the data: with all the staining methods, the detectability threshold of these cells remains undefined. It is possible that a gene must be at least 90% active for cells to stain. When gene activity is suppressed in the 0-90%

-~~~;

Polytene Chromosomes, Heterochromatin,and Position Effect Variegation

329

range, no staining is detectable, and activation is more apparent than real. In contrast, low activity of the gene could suffice to produce staining and all the gradations of high gene activity would not intensify it. When analysis of gene inactivation is based on electrophoretic detection of enzymatic activity (see Figure 128), the variants located in an R(g+) chromosome are usually poorly detected, as are hybrid fractions when the protein has a subunit structure. It is difficult to decide what makes them less amenable to detection. Is there a causal association with a reduction in the amount of the produced enzyme molecule or in the catalytic activity of the enzyme? Using antibodies against purified 6-phosphogluconate dehydrogenase and xanthine dehydrogenase in two model systems, success was achieved in demonstrating that the amount of antibodies decreases in target cells. In females with a translocation affecting 6-Pgd, the fall in the amount of 6-Pgd antigen (to 70% of normal) was the same as the fall in enzyme activity (Alatortzev et al., 1982). The peptide product of the inactivated allele of xanthine dehydrogenase was qualitatively unaltered, as demonstrated in various experiments (Rushlow and Chovnick, 1981, 1984; Daniels et al., 1986). Schultz (1965) and later Bahn (1971) assumed that mosaicism was due to suppression of transcription rather than translation. Support for this assumption was first provided by the positionaffected rosy+ locus expression. A comparison of rosy-specific poly(A+) RNA transcripts in extracts from normal and mutant genotypes demonstrated no RNA fractions transcribed from chromosome R(g+) in Northern blot analyses. It was concluded that the effect of transfer of the ry+ gene to heterochromatin is pretranslational, being, in all probability, a defect in transcription leading to the production of smaller amounts of ry+ transcripts (Rushlow et d., 1984). In the case of position effect for the Sgs4 gene in the T(l ;4)wm258-21rearrangement, the quantity of specific transcript is reduced about twofold (Komher and Kauffman, 1986). The amount of pn+ mRNAs transcribed from a strongly variegated In( ILR)pn2a chromosome is about 50% of that transcribed from an R+(pn+)chromosome (Frolov and Alatortsev, 1993). Thus, the general conclusion that genetic inactivation under position effect is due to transcription inactivation appears warranted.

E. Genetic inactivation in homo- and heterozygotes for chromosomal rearrangements A number of studies showed that variegation is more strongly expressed in homozygotes than heterozygotes for chromosomal rearrangements (Demerec and Slizynska, 1937; Schultz, 1941b; van Breugel, 1970). In offspring from crosses of yIMu-5 females to Mu-5 males, mosaicism frequency for the y+ gene in the Mu-5 rearrangement was 9.8%, and it was 16.2% in the Mu-5Nu-5 homozygotes (Brunstrom, 1955). Lewis (1950) explains this by postulating closer association

330

I. F. Zhirnulev

with heterochromatin in the homozygote. However, much information has accumulated that is at variance with this view: certain rearrangements in the homozygote produce an almost normal phenotype. An obvious interpretation is that inactivation of a locus in each homolog is an independent event, and the presence of at least one active locus is a warrant of the formation of the normal phenotype (Spofford, 1976). Examples of this are numerous (Dubinin and Sidorov, 1935; Kaufmann, 1942; Slatis, 1955a; Hessler, 1961; Spofford, 1976). In D. hydei, wm2 and wm3 chromosomal rearrangements produce finegrained mosaicism of the “pepper and salt” type, the pattern being specific to each rearrangement, with wm’ producing sectoral mosaicism. The phenotype of wm3/wm2heterozygotes is not much different from that of wm2/w(i.e., wm2dominates). Large sectors are formed in wm3/wm’heterozygotes (i.e., the sectoral type is dominant) (van Breugel, 1970).

F. Spreading of inactivation along the chromosome A remarkable feature of position effect is spreading of inactivation from the actual rearrangement breakpoint along the chromosome length. As follows from the definition of position effect of the mosaic type (see Section XIII,A), in a strain with a rearrangement, the gene remains unaffected; that is, the notation R(g+) implies that the breakpoint of a chromosome should be some distance away from the gene. For this reason, in all the cases, when the mutant phenotype is expressed in R(g+)/g individuals, inactivation spreads from R in the direction of “g+,” and, strictly speaking, the extension of genetic inactivation (spreading effect) is an obligatory feature of position effect. According to Demerec et al. (1941), approximately 10% of all the rearrangements arising close enough to the Notch locus affect its phenotype, and there is no absolute necessity for the breakpoint to be immediately adjacent to Notch. Muller (1930, 1932) was the first to note spreading of inactivation. He generated a deletion of the greater part of the X chromosome the distal end of which was to the right of the facet locus (position 3.0 on the genetic map), and the proximal end in centromeric heterochromatin. As a consequence, the white gene, located somewhat more distally than facet (at position 1.5 on the genetic map), also started to exhibit variegation. Demerec and Slyzynska (1937) obtained convincing evidence that inactivation spreads along the chromosome in the closely lying roughest (rough eye surface) and white genes. The rst+ gene maps closer to the break point in the T(l ;4)w258-’8rearrangement than white. In the mosaic eye, all the white regions are surely rough, while the spots with rough facets were both normally pigmented and white (Figure 132). These experiments may show that the genes lying closer to the breakpoint are the ones most frequently inactivated. Spreading along the chromosome, inactivation can involve an increasing number of new genes; however, the frequency of their inactivation di-

Poiytene Chromosomes, Heterochromatin, and Position Effect Variegation

33 1

Figure 132. Different manifestations of the w and rst mutations in T( 1 ;4)w+,rst+/w, and 1st heterozygotesin the facet cells of the compound eye of Dosophila. Breakpoint of translocation in the X chromosome is mapped proximal to rst+ between the 3C4 and 3C5 bands; the w+ gene is mapped to the 3C2-3 band and rst+ to the 3C4 band. It is seen that rst+ is inactivated in all the facets where w+ is inactive; however, not all the rst facets are of w phenotype. After Demerec and Slizynska (1937).

minishes with increasing distance of the gene from the breakpoint of the chromosome (Demerec and Slizynska, 1937; Demerec, 1941a). Strictly speaking, it is quite a difficult task to determine the extension of inactivation on the genetic map. A long string of adjacent genes is needed for this purpose; however, there may happen to be no genes suitable for mapping in the vicinity of a rearrangement breakpoint. For this reason, estimates of the extension of inactivation are based on the most remote gene, whose (g) mutation is manifested in the R(g+)/gheterozygote. The data on extension of genetic inactivation provided in Table 28 should be reviewed with this in mind. The data indicate that the extension of genetic inactivation due to position effect is presumably specific to each chromosomal rearrangement, and it varies from several (15-25 kb) to 170 bands; accepting that an average band is about 30 kb, this amounts to approximately 5100 kb. Various deformations of chromosome structure, such as “heterochromatization” (see Section XV1,C) and “compaction” (see Section XVI,D), can spread for a long distance as well (see Table 28). It is unclear to what extent inactivation is continuous (i.e., whether loci very remote from the breakpoint are inactivated or not), and it is also unknown how other loci occupying an intermediate position can behave (Hannah, 1951). An instance of discontinuous inactivation has been reported. Clark and Chovnick

332

I. F. Zhimulev

Table 28. Extension of Genetic Inactivation in Certain Chromosome Regions of D. melanogaster under Position Effect Variegation Extension of genetic inactivation, heterochromatinization Genetic inactivation 10 kb

Rearrangement and breakpoint in euchromatin

Most remote of inactivated

genes (bands)

References

ry+ (87DF)

Clark and Chovnick (1986)

15 kb

T(3;4)$” J49 (87E-F) It1(3R)y~~ (87D-F)

pic+ (87DF)

Clark and Chovnick, (1986)

- 1 band 20 kb

In( 1)rst3 (3C3-4) Dfca74 in Dp(3; I )BI52

Aqh-I (99C-D)

25 kb

In(1)wm4 (3C2) T(J ; 4 ) (3C3-3C5)

1-2 bands”

In( 1)wm5 b, In( 1 )wmMC (3C1 to 2-3)

2-8 bands

4 rearrangements in the X chromosome Dp( I ;3)N264-5a(3B3) Tp( 1 ;3)ras” (9E1-3) T(I ;2)dm”ar7( 2 ~ 7 )

8-10 bands 10 bands

-

(3C2-3)

Gersh (1963) Shaffer and MacIntyre (1990) Tartof et al. (1989) ~Demerec ~ and Slizynska (1937)

w+ (3C2)

~

2-5 bands

8 bands

W+

~ W+

~ ~ (3C2-3)

ut+

(3C1 to 2-3)

~

Sinclair et al. (1989)

Lefevre and Watkins (1986) fa+ (3C7)

Demerec (1940) Tobleretal. (1971)

w+ (10A1-2)

I( 1 )BA5+, I( 1)BA J , Zhimulev et al. +

llI )BAS+, K1 )BAS+; (1F3-4 to 2A1-2) 10 bands

Tp(1 ;4)N264-86(3C7) 264-29 (3D4-5)

dm+ (3D5)

12 bands 14 bands

264-55

17 bands

W + (3D5) dm+ (3D5)

-26 bands

Tp(1 ;2;4)N264-85 (3C1) In(lLR)pn2a (2D5-6)

More than 26 bands

N264-100 (3B3)

-50 bands -70 bands

In(1 )N2m92 (3C3-5) Tp(1 ;2;4)N264-85 (6A2) Dp(I;J)pn2b (2EI-2)

rg+ (4E1-3)

75 bands

4 T( J ; (3E5-6)

)

~

rst+

(3C4)

(19861, Demakova et al. (1988)

Demerec ( 1940) Demerec (1941a) Demerec (1941a) Demerec ( 1940)

dm’ (2B7-8)

Alatortsev er al. (1982)

ec+ (3E8-3Fl)

Demerec (1940)

bi+ (4C7-4D2)

Demerec ( 1940) Demerec (1940)

y + (1A5-8)

~BR-C+ ~ (2B3-5) ~ ~

Zhimulev et d . (1989a1, Belyaeva and Zhimulev ( 1991a) ~ Belyaeva ~ and Zhimulev (1991a) (continues)


333

Table 28. (Contmued) ~

Extension of genetic inactivation, heterochromatinization

80 bands

Rearrangement and breakpoint in euchromatin

Dp(J ;f)R (3C)

Most remote of inactivated genes (bands) a+(1A5-8)

SC+

(1B3)

References Lindsley and Grell (1968), E. S. Belyaeva (unpublished results) Gerasimova et al. (1972)

“Heterochromatinization” (see Section XVI,C)

To 6 bands

In(J)wm4 (3C1 to 3C2-3)

Band 3C2-7

67 bands

T( J ;4)wm25B-21

Band 2B14

(3E5-6) “Compaction” (see Section XV1,D) 10 bands In(J)dorYar2 (2B1-2)

-

20 bands 30 bands

T(J;2Mm”ar7(2B7) Dp(l;flJ337 (2B7)

Koliantz and Hartmann-Goldstein (1984) Hartmann-Goldstein (1967)

Band 2B7-10

E. S. Belyaeva and I. E Zhirnulev un(published results)

1El-2 1814

Zhirnulev et al. (1986) Belyaeva and Zhimulev (1991a) Mal‘ceva and Zhimulev (1997), Mal’ceva et al. (1997a,b)

Much shorter compaction in pseudonurse cells of otu mutant 30 bands

Dp(l ;f)dorreu6O-I and 1C Dp(J;f)do~~“~~~~ (2B7)

Zhimulev et al. (1995), Belousova and Pokholkova (1997)

56 bands

Dp(J;J)pnZb(2El-2)

1B14

75 bands

Dp(l ;f)R (3A)

Much shorter compaction in pseudonurse cells of otu mutant 1814

Zhimulev e t al. (1989a), Belyaeva and Zhirnulev (1991a) Mal’ceva and Zhimulev (1997), Mal’ceva et al. (1997a,b) Belyaeva and Zhimulev (1991a)

100 bands

T( I ; ~ ) W ” ~ ~ ~ * ’

1B14

Belyaeva and Zhimulev (1991a)

- 170 bands

T(I;2)dor““45 (2B7-8)

5D

Pokholkova e t al. (1993a.b)

”Band number after Bridges’ revised maps (Lindsley and Grell, 1968). Localization after Lindsley and Grell (1968), Lindsley and Zimm (1985, 1986, 1987, 1990, 1992).

334

1.

F. Zhimulev

( 1986) described the In(3R)ry54rearrangement expressing position effect for the pic gene 15 kb away from the breakpoint, but not for the snk gene, located between the breakpoint and pic, although snk, in principle, can be subject to position effect in other rearrangements. In some cases, when the euchromatic fragment of the chromosome is framed by heterochromatin on either end, inactivation can proceed from both ends toward the center (Panshin, 1938). When inactivation is induced at both ends of an insertion, as, for example, in the Dp(l ;3)N2ff-58, the w+ and fa'genes mapped within it can become independently inactivated. For this reason, all four possible combinations of phenotypes can occur in mosaic spots (w+fa+,w+fa, wfu+, and wfa) (Cohen, 1962). There is information that position effect can also spread over a long distance in D. virilis. In heterozygotes for T(2;3)D178eand T(Y;2)D178btranslocations, the manifestation of the dominant mutation Delta (thickening and broadening into deltas at junction with margin) is considerably enhanced. Both translocations transpose the distal end of the second chromosome to the neighborhood of heterochromatin. Euchromatic breaks map to the 2 1E region for D178' and to the 22C region for D178b. Inasmuch as the D1+ gene maps to the 20A-D interval, it follows that the distance from the heterochromatic junction to the nearest boundary of the location site of Dl+ is minimally 30 bands for D178' and more than 40 bands for D178b.The expression of mutations of the ebony+ gene located distal to the translocation breakpoint is also enhanced (Gubenko and Baricheva, 1982). There are no other genes suitable for estimation of the extension of inactivation in this region, while the 20CD and 20F heat shock puffs to which the latter maps, at least between Dl' and the rearrangement breakpoints, are normally induced (Gubenko, 1984). The unusual influence of temperature should be taken into consideration when interpreting these data: the strongest mutant manifestation was observed at 30°C and the weakest at 16°C (Gubenko, 1982). Furthermore, extension of position effect was not demonstrated for the gene sequence from the euchromatin-heterochromatin junction to Dl+ and, hence, there remains the possibility that, in this case, one is not concerned with position effect variegation, but rather with a radiation-induced modification of the expression of Dl mutation.

XIII. MODIFICATION OF GENE EXPRESSION UNDER POSITION EFFECT A. Temperature It is a long-held notion that a decrease in temperature to 14-19°C usually enhances genetic inactivation considerably, whereas an increase to 25°C suppresses


335

it (Gowen and Gay, 1933a, 1934; Demerec and Slizynska, 1937, Schultz, 1941b; Kaufmann, 1942; Prokofyeva-Belgovskaya, 1947; Chen 1948; Hinton, 1949b; Stern and Kodani, 1955). In this regard, it should be noted that the occurrence of variegated areas of the eye as well as total w-cell area are temperature dependent. Their frequencies are, respectively, 100% and 85% at 18°C. They decrease to almost 0% at 25"C, remaining unaltered with further increase in temperature (to 29°C) (Surrarrer, 1935). Other data indicate that there are more pigmented cells of wild type in the variegated eye at 29°C than at 25°C (Demerec and Slizynska, 1937). Modification of phenotype by temperature is offered as one of the proofs of position effect (see Section XI1,A). The modifying effect of temperature was demonstrated for the great majority of chromosomal rearrangements and for the genes that can be inactivated in them. There are, however, certain temperaturespecific modifications that one must consider when interpreting the results:

1. In the R(g+)/g heterozygote, the (g) mutation itself can be sensitive to cold, and the mutant effect is manifested more strongly at low temperature (Gersh, 1949; Stern and Kodani, 1955; Mampel, 1965b; Spofford, 1976). 2. Inactivation is not always consistently more expressed at low temperature. No direct dependence was found in sc8 at 14, 18,25, and 3OOC: the scutellar bristles were most frequently missing at higher temperatures and the anterior superalar bristles at lower temperatures (Gersh, 1949). Position effect variegation was strongest in the In(2LR)40d rearrangement at temperatures of 17-23"C, when studied in the 15-29°C range (Hinton, 194913)). 3. Finally, more pigment is produced in eye and malpighian tubule cells at high temperatures in D. melanogasm, while the reverse was observed for the two organs in D. hydei (van Breugel, 1970).

9. Heterochromatin

1. The Y chromosome In the great majority of cases, an extra Y chromosome suppresses genetic inactivation due to position effect, while a lesser number ofYs compared to normal enhances it (Gowen and Gay, 1933b, 1934; Dubinin and Heptner, 1934, 1935; Noujdin, 1936a,b, l944,1946d, 1947; Schultz, 1936, 1941b, 1947; Demerec and Slizynska, 1937; Panshin, 1938; Cooper, 1956; Grell, 1958; Lindsley et al., 1960; Baker and Rein, 1962; Schneider, 1962; Gersh, 1963; Mampell, 1965a). The modifying strength of the Y chromosome is great. Noujdin (1935) found that the expression of yellow variegation in the sc8 inversion, expressed as percentage of flies with yellow spots on the body, increases from less than 2.21% in XXY to 37.2% in XO fe-

336

1. F. Zhimulev

males. The rst mutant phenotype in the In(I)rst3 inversion is manifested in XO males only (Gersh, 1963). Introduction of an additional Y chromosome into the genotype with the T(1;4)wm25ai8 chromosomal rearangement leads to a reversion to an almost normal eye color (Demerec and Slizynska, 1937). The total activity of a-amylase mapped to the T(I ;2)0R32chromosomal rearrangement and producing a position effect increases in series as the number of Y chromosomes increases in flies with different numbers of the Y chromosome: XO < XY < XYY in males and XX < XXY in females (Bahn, 1971). The idea of generating Y chromosome lethals, occurring when a chromosomal rearrangement without the Y chromosome is lethal but becomes viable again with it, is based on the “healing” effect of the Y chromosome (Schultz and Catcheside, 1937; Kerschner, 1949) (see also Section XII,A). It was repeatedly shown that the expression of variegation is much more dependent on the Y chromosome than temperature. For example, the phenotype of wmflies was normal at 18°Cwhen the Y chromosome was present (Lewis, 1950); w+ variegated at 18°C in the In( 1 )rs$ inversion and at 22°C in In/Y and IdIn flies, and at both temperatures in h / O (Gersh, 1963). The results were similar for In(l)wm4 (Hartmann-Goldstein and Koliantz, 1981). In a number of Drosophila species, variegation can be also Y suppressed, as shown for the pe gene in D. virilis (Schneider, 1962) and wm2 in D. hydei (Gloor et d., 1967; van Breugel, 1987). Exceptions to this rule are extremely rare (Spofford, 1976); therefore, modification of the variegated expression by altering heterochromatin amount is adduced as a proof of position effect variegation (see Section XI1,A). The In(I)sc4 rearrangement is one of the exceptions: the addition of the Y chromosome enhances rather than suppresses genic inactivation (Mampel, 1965a,b). In the strain with the Plum-DJ dominant mutation causing a uniformly brown eye color, the addition of the Y chromosome gives variegation. Based on this observation, a new genetic effect of the Y chromosome, namely, induction of mosaicism, was inferred (Dubinin and Heptner, 1934,1935). To explain this phenomenon, Noujdin (1936a,b, 1946d) suggested that brown color represents a 100% suppression of the gene, while the addition of the Y chromosome somewhat attenuates genetic inactivation, and, as a consequence, there appear spots of normal color. The other exceptions include no Y effect on the expression of the yellow variegating rearrangement in D. uin’lis (with the presence of the additional Y chromosome controlled cytologically) (Girvin, 1949) and also the rolled and wmciloci (Morgan and Schultz, 1942; Oster, 1957). The modifying effect of the Y chromosome is cell autonomous. Thus, when eye imaginal discs from w;Dp( J ;3)N264-58(w+) larvae were transplanted into C(I)RM, yw larvae with or without a Y chromosome, the amount of drosopterin in the developing eye was the same in both hosts (Janning, 1970; Gearhart and MacIntyre, 1971). This was taken to mean that the effect of the Y chromosome did not cross the boundary of the imaginal disc.


337

Janning and Becker (Janning, 1970; Becker and Janning, 1977) induced twin spots in the eyes of Dp( J ;3)N264-58-bearing larvae and one of the X chromosomes occasionally had the YS or the YLduplication at its end. As a result, the cells of these spots could have two doses of the YS arm and the cells of the other spots none (Figure 133). It proved that the frequencies of clones of pigmented cells derived from cells containing two YS arms are considerably greater than those of the pigmented cells in the rest of the eye. This provided good evidence that the Y chromosome suppresses inactivation associated with position effect variegation. To what extent do the various fragments of the Y chromosome equally modify position effect? In early experiments it was found that even single arms of the Y chromosome can exert a suppressive action (Noujdin, 1938, 1946d). Subsequently, views on the equal contribution of the various fragments of the Y chromosome diverged.

”

v5

..s

Dp/

D P 1 DP + I +

Ys ywarbru?

r b rux OD/*

w’lz

Figure 133. Scheme of the formation of twin clones of the eye cells of Drosophila as a result of somatic crossing over. (A) A heterozygous female one of whose X chromosomes IS marked by @, IF”’, andfmutations and contains the short arm of the Y chromosome (F) in the centromeric region. The second X chromosome is marked with y, Wn, rb, and rux2mutacontions and contains r‘ in the region of the telomere. In addition, Dp(J taining the normal allele of thew+ gene is present in the genome (the mutation symbols are given in the reference book by Lindsley and Grell, 1968).Flies of this genotype have facets of two colors: normal w+ and Wa in those cells where the w+ gene is inactivated in Dp as a result of position effect. In C, they are depicted, respectively, in the lower (“w+”) and upper (“w-”) parts of the eye; (B) When mitotic crossing over takes place in the regions between 17 and the centromere, chromatids of four kinds arise (two contain one arm of the Y chromosome, one contains two arms, and one has no Y-heterochromatin). ( C )If twin cell clones with diploid chromatids of the two latter types lie in the eye region, where w+ is already inactivated (the “w-” zone), cells of one clone will be @ and lz, and they will be rb rux in the other clone cells. Because the Wa, rub combination produces loss of color, the w, rux phenotype is formed. If these clones were formed on the background of w+ activity in Dp (the “w+”zone), one clone will have w+ lr and the other rub, Tux. Examples of some other clones are shown in the middle part of the eye. In the case when heterochromatin has no effect on the inactivation degree of w+, in the cells with two arms of the Y chromosome the ratio of the number of w+/w cells in the twin clones should be the same as the average w+/w in the whole eye. After Becker and Janning (1977).

w-

”

338

I. F. Zhimulev

Many researchers demonstrated that translocations of various fragments of the Y chromosome to other chromosomes, mainly the fourth, can be generated. It is known that about 50 strains contain such translocations (Parker, 1965, 1967). The availability of fragments of the Y chromosome different in length and qualitative composition permitted researchers to take advantage of them in experiments designed to modify variegation. Study of 17 different fragments of the Y chromosome in regard to w+ expression demonstrated that (1) the Y fragments of the same length exert different suppressive effects, (2) in some cases large twoarmed fragments of the Y chromosome were ineffective in suppression, and (3) certain fragments were even more efficient than the whole Y chromosome; hence it appeared that effective suppression was not correlated with the amount of heterochromatin. It followed that heterochromatin of the Y chromosome can be subdivided into two functional units (Baker and Spofford, 1958, 1959; Baker and Rein, 1962). The idea that the Y chromosome contains genetic loci modifying position effect was shared by many investigators believing that there are two such loci, one in each chromosome arm (Brosseau, 1960a,b, 1964; Barr, 1973). There is also evidence against the presence of specific suppressor sites of variegation of the X chromosome (Benner, 1971). This evidence is based on the claim that the Y-suppressed variegation is not related to definite mapped elements of the chromosomes, but rather to the amount of heterochromatin, and that it is consequently a general feature of this chromosome. Oster (1954) indicated that various fragments of the Y chromosome affect bw+ variegation and that suppression effect of the long arm (or the two short ones) and the entire Y chromosome is equivalent. To elucidate the susceptibility to suppression, females bearing In( I ) Y and I ~ ( I ) Y ' ~lethals ~ were crossed to males whose Y chromosomes contained various deletions. The results show that, first, the chromosomes have the same suppression capacities, and, second, suppression effect increases with increasing amount of Y heterochromatin until it constitutes 6040% of the entire Y chromosome; thereafter the effect changes little, if at all (Pisano and Dimitri, 1984; Dimitri and Pisano, 1985, 1989). The wm2phenotype is Y suppressed in D. hydei. Suppression capacity correlates with cytological length in fragments obtained from the same region of the chromosomes. When fragments are obtained from different regions, effective suppression varies considerably, and it does not correlate to cytological length (Hess, 1970a,b). Usually, genetic modifiers such as the Y chromosome, which alters the expression of position effect variegation, are without influence on stable position effect at the Bur locus, for example. However, irradiation of the BS Y translocation generated 5 new Bsv Y translocations, which caused mosaicism at Bur and loss of dominance. The X,B+/BSwY males had normal eyes. The addition of the Y chromosome in X,B+ Y/Bs" Y led to reversion of BS dominance; the addition of

~

~

~

Polytene Chromosomes, Heterochromatin, and Position Effect Varieuation

339

only a single arm of the Y produced an intermediate phenotype. Genetic enhancers of position effect variegation, M(2)SIO and En(war)7, revert the mutant phenotype in Bsw to normal (Brosseau, 1960b). It may be conceded, in this case, that suppression of the activity of the dominant mutant allele yielding a defective product leading to the Bar phenotype restores the normal phenotype. There is information that the Y chromosome exerts a suppressive effect on gene inactivation in a chromosomal rearrangement in the genome of the female parent. It should be noted that the effect was analyzed in offspring not inheriting the Y chromosome. This phenomenon was described by Noujdin (1944,1946d). His exemplary mutations were yellow and achaete in In(l)sc8 and yellow in the ln(l)y3" rearrangements. Thus 40% of sc8/sc8 daughters of a scaly M: mother were mosaic for yellow and achaete, while only 4.3% of sc8/sc8 daughters of an s$/y ac YLand 5.3% of an sc8/y ac rS mother were mosaic. The effect was confirmed in a number of other gene systems in D. mehogaster, D. hydei, and D. wirilis (Spofford, 1959, 1976; Schneider, 1962; Hess, 1970b; Bahn, 1971).The Y chromosome in the maternal genome suppresses variegation of the Dp(I ;3)wvc0 duplication in offspring only when inherited from the female, and not from the male, parent possessing the Y chromosome (Khesin and Bashkirov, 1978, 1979). Spofford (1976) holds the view that the autosomes must be identical (isogenic) to demonstrate this effect. She obtained a clear-cut maternal effect on w+ variegation of the Dp(1 ;3)N264-58duplication in analysis of offspring from C( 1)RM, y w/Y/Dp mothers. The strains were isogenic and the offspring differed only in the mother having or not having the Y chromosome. The daughters of mothers possessing the Y chromosome had more eye pigment than those of mothers lacking it (Spofford, 1976). In Noujdin's (1944) paper, there are also data on the effect of the Y chromosome on the rearrangement in the paternal genotype. When the arms of the Y chromosome (YLor YS) were translocated to the paternal X chromosome (not transmitted to sons), a small number of sons had yellow bristles due to inactivation of the y+ gene in the In(]) y3p chromosome than sons of fathers with the normal X chromosome. Based on rigorous analysis of these data, Baker (1968) casts doubt on the real occurrence of this event.

2. Centromeric regions of the X chromosome and the autosomes As early as the 1930s, there appeared data indicating that not only heterochromatin of the Y chromosome can act as a variegation suppressor, but also notably heterochromatin of the X chromosome (Dubinin and Heptner, 1934,1935;Noujdin, 1936a,b, 1938,1944,1946d; Schultz, 1941b). Panshin (1938) demonstrated that a heterozygous deletion of the greater part of X chromosome heterochromatin enhances the variegated expression of the w+ gene in the T(I ;4)w"" translocation. It should be noted that the effect of the inert region of the X chromosome on mo-

340

1. F. Zhimulev

saicism is as strong as that of any arm of the Y and twice as weak as that of the entire Y chromosome (Noujdin, 1938). Grell(l958) generated a gynandromorph individual whose left side was u/ulY;bweP"/bwand phenotypically female, whereas the right was presumably ulY;bwPeL/bwand phenotypically male. The left half of the eye was fully vermilion (u) insofar as position effect bwuDeL was suppressed by the Y chromosome. The right eye was variegated, with colored fleck on a white background. The white background in homozygotes is caused by two mutations (u and bw). Consequently, bWuDeLis mainly inactivated; the colored spots result from interaction of v and the active bw@' allele. This means that the X is as effective as the Y chromosome in suppressing position effect because somatic loss of the X sharply enhances genetic inactivation (Grell, 1958). Spofford (1976) noted that, if the Y chromosome were a suppressor of position effect, then in females lacking it the inactivating effect on the rearranged chromosomes would be much more strongly expressed than in males. However, this does not holds true in all cases, presumably because heterochromatin of the Y chromosome has the same suppression effect as the Y chromosome (Spofford, 1976). Experiments provided direct support for this. Deletions in the In(l)wmSJb inversion (see Figure 143 in Section XVI) that removed the distal block of heterochromatin, and a part in some cases or almost the whole nucleolar organizer in other cases, was induced by irradiation. These deletions enhanced the expression of position effect of the w+ gene (Hilliker and Appels, 1982; Hilliker and Sharp, 1988). As the result of crossing over between the inversions wm4and wd' (see Figure 143), which delimit the block of the 18s and 28s rRNA genes, respectively, at the distal and proximal ends, chromosomes of two types can be generated: the block will be deleted in some chromosomes and it will be duplicated in others. The activity of the w+ gene is considerably higher in wm4recombinants in which the nucleolar organizer is duplicated, and it is considerably lower in the w d J b chromosome with the deletion (Spofford and DeSalle, 1991). These data clearly show that the block of the repeated genes of ribosomal RNA behaves as "usual heterochromatin" in the sense of modified expression of position effect. The deletion in the region of the X chromosome between the distal ends of the block of heterochromatin and a cluster of the rRNA genes (see Figure 143) also enhances genetic inactivation of the w+ gene due to position effect variegation (Hilliker and Appels, 1982; Hilliker and Sharp, 1988). In the 1930s, however, data appeared indicating that addition of heterochromatin of the fourth chromosome effectively suppresses position effect variegation of the yellow gene (Noujdin, 1938, 1939,1946d). Somewhat later, autosoma1 enhancers, which produce a stronger effect than loss of the Y chromosome, were identified. A deletion of heterochromatin of the second chromosome, presumably of the 41A region, was detected in one of the enhancer strains,


341

Df(2R)MS2lo, a powerful enhancer of genetic inactivation (Morgan et al., 1941; Morgan and Schultz, 1942) of various genes and compaction of chromatin due to position effect variegation (Belyaeva and Zhimulev, 1991). Spofford (1976) in her review presented evidence for the modifjring effect of autosomal heterochromatin on mosaicism. Chromosomal rearrangement In(2RL)ReuB, one of whose breakpoints is at the base of the left arm of the second chromosome (suggestive of the presence of a deletion), increases the frequency and size of mutant spots of the miniature gene in the In( I )mK rearrangement (Wargent et al., 1974). Duplications of the base of chromosome 2R are suppressors of position effect variegation (Hannah, 1951; Grell, 1970). Translocations between the X chromosome and the autosomes with breakpoints in heterochromatin often act as enhancers of variegation (Spofford, 1976).

3. Exogenous DNA An attempt was made to modify the expression of position effect using exogenous DNA of phage T2. The DNA was isolated, hydrodynamically fragmented to a molecular mass of about 500,000 Da of the fragments, then added to the food of the larvae of Drosophila to a concentration of 30 mg DNA/ml of food. As preliminary experiments showed, the DNA ingested with food penetrated into cells and nuclei, and its polynucleotide structure was retained. The results demonstrated that supplementation of food of the larvae with phage T2 DNA resulted in a deceleration of growth rate by 1-2 days, and a sharp decrease in the number and size of spots composed of w cells in the eyes-that is, a suppression of position effect of the w+ gene in Dp(I ;3)wvc0(Khesin and Leibovitch, 1976).

C.

Parental effects on chromosomal rearrangement

The influence of parental genotype on the extent of genetic inactivation in chromosomal rearrangements in offspring is a possibility discussed in several papers. Two groups of effects are distinguishable; one is parental source of the rearrangement, the other is homozygosity versus heterozygosity of the mothers for the variegation causing rearrangement (Spofford, 1976). First, the receiving of a chromosomal rearrangement through egg or sperm can have different effects. In the Dp(I ;3)NZ64”8duplication, inherited with the autosomes, the extent of gene inactivation is much higher when the duplication is received through egg than sperm (Spofford, 1959,1961; Hessler, 1961; Cohen, 1962). Khesin and Bashkirov (1978, 1979) studied the effect of sex on the inactivation of the white gene in rearrangements Dp(I ;3)wvc0, T(I ;4)wm5, and In(l )wm4 in offspring. Parental influence on the rearrangements was differently manifested. Inactivation of the w+ gene in Dp(I;3)wuCowas expressed more weak-

342

1. F. Zhirnulev

ly when offspring received the duplication from the male parent. Such an effect of rearrangement source was not found in T(1;4)eud; it was quite slight in In(f)wm4. It was assumed that differences in the amount of heterochromatin in the X and Y chromosomes between males and females affect the structure of the chromosomal rearrangement, or, as the result of addition of various histones, DNA compaction may be different. Studies on parental effect of the chromosomal rearrangement dorvar7 demonstrated that the expression of mutant phenotypes was much stronger at all temperatures when the translocation was transmitted from the father (Demakova and Belyaeva, 1988). In D. uirilis, the presence of parental effect was tested on several rearrangements causing variegated phenotypic expression of the pe mutation. The results did not provide a straightforward answer (Schneider, 1962), nor did Luning's (1954) results with variegation of the sc gene in inversions. Second, the extent of genetic inactivation in a chromosomal rearrangement depends on whether it was in a homo- or heterozygous condition in the maternal genome. For example, 4.8% of offspring from the sc8/sc8 X +/Y and 40% from the sc8/+ X +/Y crosses showed variegation (Noujdin, 1944). In another experimental system the effect was similar, with the level of eye pigment being higher in sons of mothers homozygous for Dp( f ;3)N264"8 than in sons heterozygous for it (Hessler, 1961). Parental effect has been likewise described in another series of genetic systems (Spofford, 1958, 1976). Spofford and Baker (Spofford, 1962,1966, 1976; Baker, 1968) suggested that a great part of maternal effect of this type may be due to the dominant supduplication. pressor of position effect (Su-V)closely associated with the N264-58 Although the experiments performed to test this type of maternal effect were concurrently controlled by Su-V segregation, attempts to detect it were unsuccessful. Third, the low temperature (16°C) at which the developing female parents bearing Dp( f ;3)w" were maintained considerably enhanced variegation in offspring with the duplication. Temperature at which males developed had no influence on variegation. The effect of temperature at which the parent female developed is expressed also when Dp(1;3)"'"is transmitted from the male to offspring, but it is expressed more strongly when present in the mother exposed to low temperature (Khesin and Bashkirov, 1978, 1979). For discussion of possible mechanisms of the parental effects, see Singh (1994).

D. Chemical modifiers Many attempts were made to modify the expression of position effect by feeding larvae with various agents, including those modifying histones or interfering with mitosis, the biosynthesis of precursors of RNA synthesis, as different stages of DNA replication and translation (Schultz, 1956, 1963). When fed with 5-bromodeoxyuridine (250 or 1000 mg/ml for 4 hr), lar-


343

vae of Drosophila at the age of 52-76 hr, which were heterozygous for translocations causing position effect ci, showed a significant (P > 0.01) reduction of the effect in the strain with the T(Y;4) translocation. The effect was not revealed in other strains carrying T(2;4) and T(3;4)(Dzhataev, 1973; Shavelzon etal., 1973). The most interesting results were obtained by treating larvae with the oleic acid salt sodium butyrate. As demonstrated for various cell lines, when treated with millimolar concentrations of butyrate, cells accumulate hyperacetylated histones; that is, this salt presumably inhibits their deacetylation, which results in drastic alteration of the functional and structural organization of chromatin (Birren et d., 1978; Candido et al., 1978; Riggs et al., 1978; Sealy and Chalkley, 1978; Spirin et al., 1988). Measurements of the activities of histone deacetylases in Freund cell extracts and in Dosophila adults showed that the addition of butyrate (5-2OmM) reduces enzyme activities 1.5- to 5-fold compared to normal (Mottus and Grigliatti, 1979; Mottus et af.,1980). Treatment of In(I)wm4 larvae with butyrate, starting with a 70mM concentration and increasing it, considerably suppresses position effect variegation (i.e., it increases the proportion of cells in which the w+ gene is active). Since acetylated histones are located in the active regions of the chromosomes, butyrate acts through inactivated deacetylases to promote the accumulation of acetylated histones, thereby increasing gene activity (Mottus and Grigliatti, 1979; Mottus et al., 1980). Butyrate has been found to inhibit phosphorylation of histone H1 (Boffa e t al., 1981). In the course of development, the effect of butyrate is accomplished at the embryonic and larval stages, with the highest sensitivity being detected at the end of the first and the beginning of the third larval instars. The pupal stage is completely insensitive to this chemical (Mottus et al., 1982). The carboxylic acid mpropionate is only one carbon atom shorter than butyrate and, hence, both compounds are equally effective in suppressing position effect variegation (Mottus et d., 1980). Camitine was found to suppress position effect variegation in In(] )wm4 and I n ( 2 ) b ~ " ~The ~ . camitine derivatives interact lethally with Su-var(2) 1"' (Fanti et al., 1994), a mutation that induces hyperacetylation of histones (Reuter et al., 1982a). There is information that dimethylsulfoxide (DMSO), an agent known not to mod@ histones, suppresses variegation (Michailidis et al., 1988). According to the data reviewed by Spofford (1976), colchicine suppresses variegation in T(1;4)w258-21.She takes the view that it may be due to dissociation of the mitotic spindle fibers or, quite reasonably, to self-assemblyof molecules, which takes place at some time point of variegation. The latter appears plausible because colchicine interferes with the process. The phenotype of T(I ; ~ ) W " ~ ~ ~ - ~females ' / W was insensitive to analogs of pyrimidine and purine, but it was less mutant under the effect of 2,6-diaminopurine or benzimidazole. Azaserin somewhat inhibits variegation (Schultz, 1956). Schultz also indicates that ametopterine, an agent blocking the methylation of deoxyuridine in the synthesis of deoxymidine, was the strongest modifier

344

1. F. Zhlmuleu ~~

of the w variegation. Addition of deoxythymidine abolished the effect of ametopterine, whereas addition of deoxyadenosine sharply enhanced it (Schultz, 1956). Inhibitors of transcription and translation are without effect. Actinomycin D, puromycin, cycloheximide, and 5-methyltryptophan were each added to first instar larvae bearing 0 ~ ( 1 ; 3 ) N ~ ~ 4without " ~ , exerting an effect on the pigmentation extent of the eyes (Baker, 1967). When allopurinol was added to food, genetic inactivation caused by position effect was enhanced. And this was found for many rearrangements and for various characters (Table 29). A correlation was found between the extent of variegation of the w+ gene in In(l)wrn4and developmental delay in Dosophila. The pH of larvae medium was adjusted to 2.6 by substituting a citrate-phosphate buffer for the water in food. Flies grown at pH 2.6 emerged later and showed enhancement of variegated eye phenotype compared to flies grown at higher pHs. The proportion of pigmented ommatidia in flies whose development was extended to 25-32 days was approximately 20-40%, and it was 50-8096 in those whose development took less than 20 days (Michailidis et al., 1988). In interpreting experiments of this kind, uncertainty arises: is genetic inactivation caused by an increase in development time or by the direct action of the decelerating agent on the cell nucleus? In this regard, it is noteworthy that treatment of embryos and larvae of D. melanoguster with n-butyrate or n-propi-

Table 29. Effect of Allopurinol as Enhancer of Position Effect Variegation

Gene

Rearrangement

Y+

?

Y+

Dp(I ;I )xu'

ty'

?

W+

Under the effect of allopurinol decrease

ford (1982)

Number of y + bristles

-6 fold

Spofford (1982)

Xanthine dehydrogenase activity

-

Rushlow and Chovnik (1981)

Amount of eye pigment

15-20%

Spofford (1982)

m+ area of the wing

l"59

Viability

ac+

Number of normal microchaetae

BS Y

References

J. Fowle (1980) in Spof-

Amount of pigment in bristles

m+

B+

Degree of position effect ehancement

Eye size

-6-fold

Spofford (1982)

- by 70% - 2 fold

Spofford (1982) Spofford (1982)

-

Spofford (1982)


345

onate suppressess the variegation of the w+ gene in In(J)wm4 and increases development time (Mottus et al., 1980;Rushlow et al., 1984). Noujdin's (1935) earlier observation is also pertinent: when development of sc8 flies was much delayed, taking 45 days from egg laying to fly emergence at 9-1 2"C, neither temperature nor retarded development had an influence on variegation.

E. The histone genes Histone gene deletions can considerably affect the structure of nucleosomes due to deficiencies of histones (e.g., see Norris et al., 1988). Khesin and Leibovitch (1976,1978) reported that deficiency of the histone genes resulting from combination of two T(2;Y) translocations suppresses genetic inactivation; that is, deficiency reactivates the white+ gene in the T(l ;3)wvC0translocation. Two issues were criticized in this study. The T(2;Y) translocation used contains substantial portions of heterochromatin of the Y chromosome that could themselves cause suppression of variegation. Second, the breakpoint of the translocated proximal element was cytologically mapped to 39C rather than to the 40 region; that is, it is in the 39D2-3 to 39E1-2 regions, thereby implying that the deletion would not affect the histone gene complex (Moore et al., 1983). Although this early study did not provide definitive evidence, the effect of the histone genes on mosaicism was demonstrated later. The effect of nine deletions of the proximal regions of chromosome 2L on variegation of white+ in the In(l)wm4 inversion or Bar+ in Bs-warwas studied. Of these nine deletions, four removed all and one removed part of the histone gene cluster. When the deletions did not remove the histone genes, variegation was not affected; when they did, there was an increase in the activity of the two variegating genes. The Bs-" allele showed intermediate activity in heterozygotes for the deletion partially the removing gene cluster complex (Moore et al., 1979a-c, 1981, 1983). The effect of deletions on genes mapped to the autosomes was tested, too. The T(2;3)SbUtranslocation transposes a fragment of the chromosome with the Sb allele from the midpoint of chromosome 3R to the centromeric heterochromatin of chromosome 2R. Inactivation of the mutant dominant allele causes the formation of more normal bristles. A smaller number of bristles are of normal length in flies heterozygous for deletions of the histone genes. This implies reactivation of the Sb chromosome (Moore et at., 1981, 1983). Suppression through the histone genes is not modified by the Y chromosome, and an increase in the dose of histones by addition of a duplication does not lead to enhancement of position effect variegation (Moore et a!., 1983).

F. Genetic modifiers The possible existence of various genes altering the expression of position effect has been discussed in the section concerned with the modifying effect of the Y

346

1. F. Zhimuiev

chromosome (see Section XIII,B,l). Numerous data on the influence of the “genetic background” on variegation indicate the presence of modifier genes in the other chromosomes. Noujdin (1935) was the first to reach the conclusion that there exist autosomal modifiers of position effect of the sc8 inversion. Later on, Dubinin (1936) found that certain chromosomal rearrangements have an influence on the expression of position effect of Plum.Schultz (1941b) found several instances in which a rearrangement between euchromatin and heterochromatin acts as a modifier of variegation. Somewhat later, a similar modifying effect of the In(2LR)ReuBinversion on position effect In(1)mKwas disclosed (Wargent and Hartmann-Goldstein, 1974). In a number of cases, the expression of variegation can be shifted by selection. Isolation of single genes from the “genetic background” was successful. A hereditary factor determining fully white eyes due to position effect in the translocation T(l;4)wm25a18 was identified; the eye was cream or cherry when the factor was missing (Demerec and Slizynska, 1937). A single selection in the sCa strain for mosaic and nonmosaic females affects the expression of sc8 in offspring. Two new stable lines with frequencies of variegation expression of 51.61% in one and 5.52% in the other were developed through systematic selection of individuals. A ci line (see Section XV, A) in which 78.1% of females and 43.7% of males expressed extremely strong variegation was also developed through selection (Sidorov, 1947). From 1%to 10.6% of mosaics for yellow+in the Muller-5 rearrangement can be obtained by substituting autosomes (Brunstrom, 1955). There are other examples of the modifying effect of the genetic background in the literature (Hinton, 194913;Cohen, 1962; Suzuki, 1965). Genetic modifiers were identified in pure strains (see M. L. Belgovsky in Schultz, 1941b, 1950; see also Spofford, 1965,1967, 1969,1973). One of the suppressors found by Henikoff (197913) proved to be a deletion of the 87E2-F2 region. When heterozygous with the normal homolog, the deletion suppresses the heterochromatic position effect at the wm4 (Henikoff, 1979b) and rosy (Rushlow and Chovnick, 1981) alleles. A new period in study of genetic modifiers of position effect started in the 1980s when, as the result of intensive experiments using insertional (P-elementmediated) mutagenesis, as well as treatment with ethyl methane sulfonate (EMS) and x-rays, a large number of mutations modifying the extent of inactivation of a position-affected gene were obtained in the laboratories of Reuter, Grigliatti, and Tartof. Two approaches are mainly used. In the first, mutations modifying position effect were generated in a strain carrying the R(w+)rearrangement in the male X chromosome and causing moderate variegation of eye color, with red and white sectors intermingled. If the suppressor was dominant, inactivation of R(w+)was suppressed, and the eye acquired normal red color. Induction of the enhancer led to the formation of an eye in which the predominant proportion of cells was white. In the second approach, after it was clearly shown that suppression and


347

enhancement of variegation are related, in many cases, to an increase or decrease in the dose of the normal allele (Henikoff, 1979b; Moore et al., 1979a; Reuter and Szidonya, 1983; Duttagupta et al., 1984; Reuter et al., 1987; Locke et al., 1988;Szidonya and Reuter, 1988; Tartof et al., 1989; Wustmannet al., 1989),a method was developed to define regions whose aneuploidy (a deletion or duplication) produces a modifying effect. As the result of a systematic large-scale search, hundreds of mutations affecting from 30 to 43 genes were identified as proved by complementation analysis (Reuter and Wolff, 1981; Sinclair et al., 1983, 1989, 1992; Tartof et al., 1986, 1989; Locke et al., 1988; Wustmann et al., 1989). Their distribution in the Drosophila genome is shown in Figure 134. The majority of position effect modifiers were localized in the second and third chromosomes (Spofford, 1976; Henikoff, 1979b; Reuter and Wolff, 1981; Reuter and Szidonya, 1983; Sinclair et al., 1983; Reuter et al., 1986). As is apparent in the figure, the number of suppressors is considerably larger than the number of enhancers. The occurrence frequency was estimated as 1:lOOO chromosomes for Su mutations and 1:16,000 for En mutations. It is unclear why the estimates are discrepant. One reason is difficulty in making a distinction between the increase in the extent of variegation caused by enhancer from normal variability of position effect (Sinclair et al., 1983). The definitive identification of genes modifying position effect is questionable, and it is unknown how many of them have been described and mapped because the calculated estimates disagree. Having generated two suppressor loci in the small 86-87 region, Henikoff (1979b) believes that the loci of modifiers, being distributed in the Dosophila genome with the same frequency, should occur once per 25 chromomeres, which suggests the existence of about 200 such genes per genome. This conclusion was refuted by Grigliatti’s group owing to the fact that suppressors are actually arranged into two clusters in chromosomes 2L and 2R rather than uniformly distributed in the Drosophila genome. HenikoWs quite reasonable estimate for the genes located, as proved later, in one of the clusters cannot be extrapolated to the entire genome, and the total number of genes is much smaller (Mottus et al., 1982; Sinclair et al., 1983; Grigliatti et al., 1984). Tartof et al. (1985, 1986) believe that 10-20 dominant suppressors or enhancers are identifiable in D. melanogaster; according to other reports (Locke et al., 1988), their number is 20-30. Based on the calculations of Reuter and colleagues (Reuter and Wolff, 1981; Reuter et al., 1987; Szidonya and Reuter, 1988), the number of modifiers varies from 50 to 150. Having generated a series of deletions covering approximately 30% of the chromosome length containing 38 haplo-dependent modifiers, Wustmann et al. (1989) inferred that their total number should be about 130. According to more recent estimates, the number of enchancer genes is about 30 in the third chromosome and between SO and 60 in the whole autosome complement of Drosophila melanogaster (Dorn et al., 1992).

Figure 134. Location of dominant suppressors and enhancers of position effect in the DrosophiLc g m m e . Sections from 21 to 60 and from 61 to 100 correspond to the regions of the second and third chromosomes, respectively; the positions of the centromeres are 40 and 80.Suppressors and enhancers are depicted: black rectangles, deletions; white rectangles, duplications; arrows and parentheses,point mutations. After Tartof et al. (1989).


349

The specificity of the effect of various modifiers can vary; as a rule, suppressors reduce and enhancers promote inactivation of many genes in various chromosomal rearrangements. For example, Su-w(IIIL-4 I ,4) suppresses position effect induced in sc8, y3p, wm4, and rst3 (Spofford, 1965). Universality of modifiers was also demonstrated (Reuter and Wolff, 1981; Reuter et al., 1982b; Sinclair et al., 1983, 1989). Specificity was revealed, too. Three EMS-induced enhancers, E(uar)301, -302, and -303, differently influence the series of variegated rearrangements w". All three enhance wm4 variegation and none affects wm51b; E(var)303 enhances only wdC, wm4, and wmJ; E(uar)301 enhances only w d and wm4; and E(uar)302 enhances only wm4 (Sinclair et al., 1989). There is a unique case in which one allele of Su-Var suppresses the position effect of the t ~ + ,fa+, and dm' genes and its other allele enhances it in the Dp( I ;3)wm264-58arearrangement (Spofford, 1965). In regard to the other features of the modifier genes, it should be noted that the majority of suppressors are recessive lethals (Reuter et al., 1986; Sinclair et al., 1992). These genes act at the earliest stages of development. The gene products contributed by the mother are required to provide normal embryonic development (Szabad et al., 1988; Sinclair et al., 1992). Many of the enhancer mutations display paternal effect (Reuter et al., 1985; Reuter and Spierer, 1992). Twenty-three of 34 mutations show significant paternal effects (Dorn et al., 1992). Dose dependence is the most remarkable property of genetic modifiers. The position effect suppressor Henikoff has described (197913) proved to be a deletion of the 87E2-F2 region. Subsequently, this relationship between change in dose in the chromosome regions containing the normal allele of the suppressor or enhancer and modifying effect has been reported in the literature (Reuter and Szidonya, 1983; Reuter et al., 1986). When a duplication overlapping a deletion was introduced into heterozygotes for the deletion removing the suppressor (i.e., the dose is restored to diploid level), the modifying effect is, to a large extent, abolished (Reuter and Szidonya, 1983;Reuter etal., 1987;Szidonya and Reuter, 1988). When flies heterozygous for the deletion at the Su-war(3)7 locus were transformed by a cloned DNA fragment containing the normal allele of the gene, the suppressor effect was abolished and an enhancer effect the influence of which increased with copy number of inserted DNA was evoked (Gausz et al., 1989; Reuter et al., 1990). The Dp(2;2)Mdh3 duplication containing a small fragment of the second chromosome to which at least seven different suppressors map considerably reduces the effect of any one of the suppressors when introduced into the genotype (i.e., in the SuJSu+/DpSu+combination) (Sinclair et al., 1989). Considerable changes in dose can produce qualitative alterations in the functions of certain modifiers. Two classes of transitions are distinguished: (1) genes that act as enhancers of position effect variegation when duplicated, or as suppressorswhen in a single dose because of mutations or deletions; and (2) genes that enhance the effect when in a single dose and suppress it when in a triple dose

350

1. F. Zhimulev

(Locke et at., 1988;Tartofetat., 1989;Wustmann etal., 1989;Sinclairetal., 1992). According to Reuter and Spierer (1992), gene modifiers of PEV fall into one of the following classes: (1) about 10 haplo-suppressor and triplo-enhancer loci, (2) about 10 haplo-enhancer and triplo-suppressor loci, (3) about 75 haplo-suppressor loci without an opposite effect of three copies; and (4)about 25 haplo-enhancer loci without a triplo-inverse effect. Taking advantage of dosage dependence, experiments were undertaken to identify regions in which changes in dose do not produce the modifying effect (Wustmann et al., 1989). These data show that the division of some modifier genes of position effect variegation into suppressors and enhancers is a relative matter. The question was also raised whether variegation modifiers, such as temperature, the Y chromosome, butyrate, and suppressors and enhancers, interact. Removal of heterochromatin of the Y chromosomes from the genome appears to be the strongest enhancer of position effect variegation. The effect dominates over that of genetic enhancers or suppressors,although the influence of E(uar) and loss of the Y chromosome is equivalent in the case of position effect bw" (Sinclair et al., 1989). The strongest suppressors are genetic, inasmuch as an additional Y chromosome produces a weaker suppression effect than the Su loci (Reuter and Wolff, 1981; Sinclair et al., 1983, 1989). The suppressive effect of an extra Y chromosome is compensated, at least partly, by the effect of E(var)30J and E(uar)302 enhancers (Sinclair et al., 1989), while the E(war)cJo' enhancer dominates over the suppression effect of an extra Y chromosome (Reuter et al., 1983).The mutant alleles at the three Su-uar loci dominate over the strong enhancer effect of complete loss of the Y chromosome (Reuter et al., 1986). Cases were described where genetic combination of two enhancers in the genome elicits modifier interaction suppression of the genetic activity of the y+ gene in the y3p inversion that is twice as strong as in each separately (Locke et al., 1988). It is easy to assume that the products of the suppressor and enhancer genes can manifest in opposite action. Study of the effect of Su and En in the same genome revealed that the level of genetic inactivation of the w+ gene is the same as in Su/+ individuals (i.e., the suppressor dominates over the enhancer) (Sinclair e t al., 1989). It is of interest that introduction of DNA containing the normal allele of the Su-var(3)7 gene into the genome by DNA transformation can exert a normalizing effect not only on Su-uur(3)7 mutants, but also on bearers of other butyrate-sensitive suppressors (Gausz et d., 1989). Certain specific types of interaction proved to be very strong. For example, Su-uar(3)303 carries recessive female sterility; however, it becomes a zygotic lethal in the presence of butyrate or an additional Y chromosome (Szabad et al., 1988). Lethal interaction of a suppressor with butyrate and an additional Y chromosome was observed for other suppressors. It was shown that viability is decreased in XXY females; it is abnormal in XO males, however, in the presence of the Su-uar(2)I suppressor (Reuter et al., 1982a, 1986).


35 1

Genetic and molecular analyses of modifier loci indicate that they constitute a group of genes with diverse functions (Table 30). The suggestion has been repeatedly made that the genes modifying position effect variegation encode proteins included in chromatin and control the process of its formation (Spofford, 1967; Henikoff, 197910; Reuter et al., 1982a; Sinclair et al., 1983; Eissenberg, 1989;Reuter and Spierer, 1992).This contention is well grounded because ample evidence has been obtained:

1. Reuter et al. (1982b) demonstrated that En(var)CJoJenhances mutant phenotypes of several genes subject to position effect variegation, and that there was an instance in which enhancement correlated with an increase in heterochromatization extent of a chromosome segment in the T(I ; 4 ) ~ ~ translocation. 2. The suppressor mutation Su-var(2)1O J shows recessive sensitivity to butyrate, and mutants for this suppressor show significant hyperacetylation of histone H4 (Dorn et al., 1986). 3. It was found that Su-var(2)205, a suppressor of position effect that maps to the second chromosome, is the structural gene for the protein encompassed by pericentromeric and telomeric heterochromatin (James and Elgin, 1986; Eissenberg et al., 1987, 1990, 1992; Eissenberg, 1989; James et al., 1989; Powers and Eissenberg, 1993). The HP1 is localized predominantly to the chromocenter of polytene chromosomes (see Table 30), so it can be a structural component of heterochromatin. During metaphase and anaphase this protein is no longer associated with condensed chromosomes, but instead is dispersed throughout the spindle and again associates with chromosomes at telophase (Kellum et d., 1995).It is required for correct segregation in Dosophila embryos (Kellum and Alberts, 1995). The Su-var(3)-7gene encodes a putative zinc finger protein, and the modulo gene encodes DNA- and RNA-binding protein (Table 30). The Trithmax-like gene encodes the GAGA protein (see Table 30), a transcription factor binding to GA/CT-rich sites near a variety of different promoters (Becker, 1994). 4. The results of two experiments indirectly support the role of the modifier genes in self-assembly of chromatin. One suppressor showed a strong maternal effect: females bearing the Su-uar mutation had an influence on genetic inactivation in offspring not receiving Su-var (Grigliatti et d., 1984). The butyrate-sensitive Su-var(3)3 gene product with maternal effect is required for normal embryonic development (Szabad et al., 1988). All this indicates that the products of the modifier genes are already needed at the early stages of embryonic development. 5. The protein product of the variegation suppressor gene Su-var(3)7 has seven zinc finger domains for binding to DNA (Reuter et al., 1990), and it is identified in the nucleus (E Cleard, V. Garzino, A. Spierer, and P. Spierer in Reuter et al., 1990; Cleard et d . 1995).

~ ~ ~

352

1. F. Zhlrnulev

Table 30. Molecular-Genetic Characteristicsof Position Effect Modifiers in Drosophila melanogaster. ~

~~

Gene symbol (location)

~~

~ ~ _ _ _ _

Characteristics

References

E(var)3-93D, Mutants are sensitive to butyrate; gene encodes formerly E(var)3-3 protein containing a domain common to the transcription regulator rramnuck and product of Broad-Complex; no zinc finger motif was identified; antiserum against E(var)3-93D protein is located in a large subset of sites in polytene chromosomes;protein seems to establish and/or maintain an open chromatin conformation

Dometd. (1993)

modulo (mod) (100F)

Krejci et d.(19891, Garzino et al.( 1992) 0.V. Demakova,N. I. Mal’ceva, J. Pradel, and I. F. Zhimulev (unpublished results) Gerasimova et d.(1995)

Suppressor of variegation; gene contains DNA, RNA, and protein binding domains Antibodies against mod protein bind nucleolus, chromocenters,and bands in both salivary gland and otu PNC chromosomes Mutations of the gene act as enhancers of position effect variegation

Modifierof white (Mow) mus209 Su var(2) 1 , Su var(2)10

Weak suppressor of PEV

Bhadra and Buchler ( 1996)

Suppressor of PEV, encodes homologue of proliferating cell nuclear antigen (PCNA)

Henderson et al. (1994)

Mutations are sensitive to butyrate

Reuter et al. (1986), Dom et al. (1986)

Su-var(2)205 (29A) Haplo-suppressor, triplo-enchancer;codes for heterochromatin-associatedprotein HP1, which is a 206-amino-acid nonhistone protein, multiply phosphorylated; shares a region

(37 amino acids) of striking homology to Polycomb gene (“chromodomain”)

During embryonal development, enrichment of heterochromatin with HPl was found at nuclear cycles 10-14 This time corresponds to the time of increased phosphorylation of HPl Overexpressionof HP1 under heat shockregulated promoter results in significant enhancement of PEV HP1 homologs have been identified in D. virilis, mealybug, mouse, and human

James and Elgin (1986), James et d.(1989), Eissenberg et d.(1990, 1992,1994, 1995), Paro and Hogness (1991),Emnberg and Harcnett (19931, Powers and Eissenberg (1993), Plater0 et al. (1995), Elgin (1996) Kellum et d . (1995)

Eissenberg et al. ( 1994) Eissenberg and Hartnett (1993) Singhetal. (1991), Epstein et d. (1992), (continues)


353

Table 30. (Continued) Gene symbol (location)

Characteristics

At 10th cell cycle, C-banding in heterochromatic regions starts to be easily discernible Protein appears in euchromatin compacted as a result of position effect variegation Su(vur)231 (31E)

DNA sequencing suggests that it binds to DNA and cytoskeleton

References Clark and Elgin (19921, Saunders et al. (1993) Vlassova et al. (1991a,b)

Belyaeva et al. (1993), Demakova et ul. ( 1993)

I. Whitehead and T. Grigliatti in Reuter and Spierer (1992)

Su-vur(3)6 (87B6-12)

Codes for protein phosphatase 1 catalytic subunit 87B; protein phosphorylation possibly regulates condensation state of chromatin in interphase nuclei

Dombradi and Cohen (1992), Baksa et al. ( 1993)

Su-var(3)7 (87E)

Haplo-suppressor,triplo-enchancer, contains seven Reuter et al. (1990); Cleard et al. (1995) widely spaced zinc fingers, each preceded by a tryptophan box

Su-war(3)3-9(88E) Haplo-suppressor,triplo-enchancer; codes for 635-amino-acidprotein with “chromodomain” and region of homology to En(d and trithurax

Tschiersch et al. (1994)

su (2) 5

Larsson et al. (1996)

The gene encodes S-adenosylmethionine synthetase. Mutant alleles show suppression

of wm4 Trithorax-like (Tr-l) Enchancer, which is required for the normal expression of homeotic genes, encodes (70F1-2) GAGA factor, generating nucleosome-free regions of DNA; it contains tramtrack and poly(Q) domains

Farkas et al. (1994)

Weakener of white (Wow) (76D5-76F)

Acts as a suppressor of position effect variegation

Birchler et al. (1994)

zeste

Encodes DNA-binding protein, acting as a transcription factor and mediating transvection phenomena at several loci; null recessive enchancers of position effect variegation Act in cis to suppress position effect variegation

Judd (1995)

Null allele of dE2F enhances PEV when heterozygous

Seum et al. (1996)

Transcriptional enhancers

Walters et al. (1996)

354

1. F. Zhimulav

6. Preliminary data indicate that certain modifiers of position effect variegation control the expression or maintenance of the determined state of the homeotic genes in the bithorux complex (M. Giarre, J. Gausz, and H. Gyurkovics in Reuter et ul., 1990). The Polycomb (Pc) gene acts as a repressor of the homeotic genes at the early stages of embryogenesis. Transcripts of this gene are maximally represented in unfertilized eggs and at the earliest stages of the embryos. A homology was found between 37 amino acid residues at the N-terminus of the molecule with protein encoded by Su(vur)205 (Paro and Hogness, 1991). This parallelism may provide evidence for similarity in the formation and maintenance of the repressed state in the case of position effect and inactivation of the homeotic genes. 7. The action of suppressors and enhancers is expressed autonomously. In( I )wm4 larvae were transplanted with imaginal discs from individuals with the same inversion and with one of the modifiers Su-vur(2)Io',Su-~ur(3)3"~, or E - ~ a r ( 3 ) 2or~ reciprocal ~, transplantations were done. No evidence for the effect of the donor on the recipient, or the reverse, were found in experiments of both types (Reuter and Szabad, 1987). 8. The previously described transitions from suppression to enhancement of position effect (and the reverse), observed when the dose of the modifier gene of position effect variegation is changed, are readily explained assuming that heterochromatin contains proteins compacting and decompacting its structure. When decreased in dose, protein molecules of the first type (Figure 135) should lead to looser compaction of (hetero)chromatin and to enhancement of gene activity. When in three doses, they should make chromatin more

lz?El

Character of protein action

protern

#--1- -

1

2

3

I

I

I

a

l F

E

Gene dosage

l

i

r n m a I

1

I

I

Modifcationof position effect variegation

Gene dosage Character of protein action

Figure 135. Modificationof the manifestation ofpositioneffect dependingongene dosage controlling compaction and decompaction of heterochromatin (see text for explanation). From Zhimulev (1992a, 1993).


355

compact and, consequently, enhance the effect. Similar considerations are applicable to proteins decompacting (hetero)chromatin (see Figure 135). Mutations of the suppressor are more frequent (in a single dose) than mutations of the enhancer, presumably because the number of compacting proteins is larger in heterochromatin. The information about the action of modifiers provided previously pertains to cases of position effect of genes normally positioned in euchromatin. When variegation is caused by transfer of the gene from hetero- to euchromatin (It+ and others), the direction of modification can change; for example, Su(Vur)208 suppressor can become an enhancer. There are indications that genetic activity associated with position effect can be modified in strains carrying chromosomal rearrangements. For example, in R A , w / Y / D ~ w females, ~ ~ ~pigment ~ . ~ ~level ~ of the eyes is twice lower than in RM, w/Y/Dpwm264.58a.Autosomal inversions Cy and Ubx also reduce pigment level (Suzuki, 1965).

XIV. TIME OF GENETIC INACTIVATION IN DEVELOPMENT Evidence bearing upon the particular stages in developmental at which genes become inactivated due to position effect is controversial. In earlier studies, it was claimed that the temperature at which embryonic development takes place is important for genetic inactivation (see J. M. Gowen in Gersh, 1952). Noujdin (1945) found that, "if the first day of development occurs at 25-27"C, and the cultures are later transferred to conditions of lower temperature ( 1 4 ' 0 , then, irrespective of development time (from 1-34 days), the frequencies of variegated individuals do not differ from control." Subsequently, Schultz ( 1956)demonstrated that the temperature-sensitive period of the inactivation of the w+ gene in malpighian tubules is restricted to the first half of the period of embryonic development. If that period of life of embryos of the T(l ;4)wm25a21/w48h genotype proceeds at 18°C and the larval period at 25"C, inactivation extent is the same as though the entire development occurred at 18°C. Later still, using the same rearrangement, Hartmann-Goldstein ( 1967)showed that the period of highest sensitivity to cold lies in the first 4-6 hr of embryonic development. Finally, using T(l ;2)d0ruar7,the BR-C gene was shown to be most effectively inactivated when embryos were exposed to low temperature during the first 3 hr of development (Zhimulev et al., 1988). Analysis of P-element construct containing a Hsp 70 promoter-driven lac2 showed that gene inactivation (silencing) is most extensive at cellular blastoderm, after which islands of inducible lacZ expression begin to emerge (Lu et d., 1996). The following results may be claimed as evidence for the view that genetic activation should occur during the early stages of development. Compaction

356

1. F. Zhimulev

of chromosome regions resulting in gene inactivation with position effect (see Section XVI,D) is considerably enhanced by removal of Y chromosome heterochromatin. Quite apparently, this can be consequential only when the amount of removed heterochromatin is comparable to that present in the other chromosomes; that is, it is most reasonable to think of the effect as occurring at the diploid level prior to polytenization. The earliest stages of development are presumably critical here, possibly before eggs are laid. In fact, to obtain numerous larvae showing maximal variegation, flies are placed in tubes at 14-16°C 48 hr after they have laid all eggs; thereafter, flies that have developed at higher temperature are transferred to new tubes at 14-16”C, and only flies and larvae maintained in these tubes are analyzed (van Breugel, 1970; I. Hartmann-Goldstein, 1980, personal communication). It is Spofford’s (1976) view that the temperature-sensitive period of genetic inactivation is coincident with the time of blastoderm formation. Thus the gene is inactivated in association with position effect variegation long before it is committed to high activity (Brown, 1966). There are, however, indications that other stages of development are temperature sensitive. For example, Surrarrer (1935, 1938) believes that the period between pupation formation (25-35 hr after it) and fly emergence is of importance for the w+ gene subject to position effect. Subsequently, HartmannGoldstein (1967) found that exposure to cold (14°C) after the period of embryonic development has an influence on “heterochromatization” of the 3C1 band, although it is unclear to what extent this notion is consistent with genetic inactivation (see Section XV1,C). In strains with inactivation subject to enhancers of position effect, the temperature-sensitive period continues throughout larval development, judging by the fact that transfer to higher or lower temperature leads to enhancement of variegation (Spofford, 1976). According to Chen’s (1948) and Gersh‘s (1952) data, treatment of wm258-J8 and w~ strains with low temperature (16-17°C) was effective in decreasing the amount of eye pigment only at early pupal and, possibly, embryonic stages of development. In analysis of inactivation of the y+ gene in the y3‘ inversion during the formation of marginal microchaetae of the wing, it was found that the temperature-sensitive period falls at the pupal stage at any shift of temperature (from 28 to 18°C or from 18 to 28°C) (Tartof and Bremer, 1990). Thus very early changes in genetic activity can produce a pattern of clonal activation in eye cells; then an inactivation pattern occurring during the pupal stage of development can superimpose on these changes, resulting in a new pattern not necessarily related to clonality (Gersh, 1952). From the results of experiments with clonal initiation (see Section XI1,C) it was concluded that a third stage critical to inactivation lies at the transition from the first to the second instars, that is, about 45-48 hr after egg laying (Chen, 1948; Becker, 1957, 1960, 1961, 1978; Baker, 1963,1967); and according to other data, it falls in the 39- to 47-hr interval (Janning, 1970).


357

The experiments with radiation-induced somatic recombination are especially relevant. Cell clones arising following irradiation differed in arm number of the Y chromosome: two arms in the cells of one clone and none in the cells of the other (see Figure 133 in Section XIII). Before irradiation, all the cells already contained one arm of the Y, and this amount of heterochromatin had a definite influence on the inactivation extent of the w+ gene subject to variegation. If the extent of inactivation changed in both cell types as the result of Y chromosome arm redistribution (one arm redistributed to one cell, none to the other), this should be taken to mean that this extent can be modified during the first cell division following irradiation. Becker and Janning (1977) precisely demonstrated this: addition of the Y chromosome can modify position effect at the time of clonal initiation of eye cells (i.e., at the end of the first larval instar). In D. hydei, the white+ gene in the R(Y)wmchromosomal rearrangement is differently inactivated in different cells in the course of inactivation. As a result, incomplete sectors of w+ cells are formed, showing that the w+ gene is still active and gives rise to a w+ cell clone during the first division of the eye cells. However, inactivation takes place and w cells are formed at later stages of division (Beck et al., 1979). A particular type of fine mosaicism (“pepper-and-salt”) of the gene lends credence to the idea that the w+ gene is inactivated late, subsequent to eye formation in Drosophih (see Section XI1,C). Evidence for multiple activation-inactivation of the y+ gene in the In(l) y3p inversion is provided by the case of alternating pigmented and unpigmented zones along the length of the bristles of Drosophih (Spofford, 1976).

XV. UNUSUAL CASES OF POSITION EFFECT A. The Dubinin effect Weakened dominance of the normal allele of the ci+ gene conditioned by a rearrangement bringing heterochromatin into close proximity to the allele was first discovered by Dubinin and Sidorov (1934a,b) (see also Dubinin, 1935; Dubinin et al., 1935). The phenomenon was called the “Dubinin effect.” The ci+ locus controls the development of the cubital wing vein, and, when it is inactivated in association with position effect, a break with length related to inactivation extent occurs in the vein (Figure 136). It is difficult to study position effect of ci+, partly because its genetics is specific and also because ci+ is located in a region barely accessible to genetic mapping. The ci+ locus was localized within rather wide cytological limits. Bridges (1935a,b) places ci+ within the Minute3 deletion with limits defined by bound-

1. F. Zhlmulev

358

m

Figure 136. A schematic representation of wings with different degrees of manifestation of the ci trait. After Khvostova (1939).

aries from 1OlD to 102B (the G band on Bridges’ map, and the third thick band in the 102B region on another map). Th limits given in Lindsley and Grell’s (1968) reference book are between 101F2 and 102A1 (102A2-5). Providing data on position effect at the ci+ locus, Stern and Kodani (1955) map ci+ in the interval 102E102C4. Hochman (1965) localized it in the region delimited by the M(4)63a deletion, removing the 101A6-7 to 102A1-2 region, that is, in the lOlF to 102A1-2 interval or in the first two thick bands of the 102B region (Hochman, 1971).Based on the results of in situ hybridization of cloned fragmentsof the ci+ gene (Tartof et al., 1985)with normal strains and containingtranslocations,it was demonstrated that the labeling sites are situated in the 101F-102A or 101F-102B region (Locke and Tartof, 1994),and in lOlF region according to Demakova et al. (1997). Position effect at the ci+ locus does not arises in each translocation of the fourth chromosome. Thus, Dubinin and Sidorov (1934a,b) found weakening of ci+ dominance in only 10 of 19 translocations studied, Dubinin (1935) in 18 of 38, and Sturtevant and Dobzhansky in 6 of 10, although in the remaining 4 some degree of ci+ inactivation was also observed (Dobzhansky, 1936). Further progress in study of position effect of the ci+ gene was promoted by availability of a simple technique for generating translocations of the fourth chromosome giving position effect: irradiated ci+ males were mated to females of a strain homozygous for two recessive mutations, ci and ey. Individuals with the mutant phenotype ci were chosen from offspring; there could occur ci/ci mutants or R(ci+)/ciheterozygotes among them (Dubinin and Sidorov, 1934a,b; Panshin, 1935; Khvostova and Gavrilova, 1935, 1938; Khvostova, 1936,1939; Neuhaus, 1939;Sternet al., 1946b;Stern and Kodani, 1955;Roberts, 1972b,c;Levina, 1974; Panshin, 1992). Somewhat later it was demonstrated that position efect is manifested also in individuals with the genetic constitution R(ci)/ci (Sidorov, 1941b; Stem and Heidenthal, 1944; Stem et al., 1946a,b). This is presumably because the phenotypic expression of venation is very much dependent on the dose of the mutant genes, being almost normal in individuals with three doses. It is therefore reason-


359

ably expected that the expression decreases in ci+>ci>R(ci+) and ci+>ci>R(ci) series (for greater detail, see Stem, 1943,1948; Stem and Heidenthal, 1944; Stem et al., 1946a,b; Lewis, 1950). It is believed that the Dubinin effect concept can be extended to this type of position effect, too (Altorfer, 1967). The variants of position effect R(ci+) and R(ci) are considered together here. The features of position effect of the ci+ gene may be indicated as follows:

1. Localization of breaks in the fourth chromosome occurs. Based on numerous data, position effect is detected only when the chromosome rearrangement break in the fourth chromosome occurs between the centromere and the ci+ gene (Dubinin and Sidorov, 1934a,b; Panshin, 1935; Khvostova, 1939). In 11 translocations not giving position effect, the break in the fourth chromosome was distal to ci+ (Dubinin et al.,1935). At the cytological level, the breakpoint maps to lOlF [76 translocations were studied by Dubinin et al. (1935) and 2 translocations by Grell (1959)]. According to Khvostova and Gavrilova’s (1935) data, in 50 translocations giving position effect, the break in the fourth chromosome occurred in the same site, distal to the most proximal band and two thin ones (presumably, the lOlDF region was implied). Spofford (1976) holds the view that the breakpoint must be proximal to lOlF for position effect of R(ci+)/ci to be expressed. Of 52 translocations causing position effect, 48 had a break in the 101 region (they were not mapped, to be more precise), 1 in 102A, 2 in lOlC, and 1 in lOlA (Roberts, 1969a, 1972a,b). Of 17 R(ci+) translocations, 11 had breaks in the 101D-F region, 2 in the 102,3 in the immediate vicinity of 102B1-2, and 1 in the left arm (i.e., in the lOlAC region) (Stem and Kodani, 1955). Somewhat different results were obtained in localization of the R(ci) translocation. When the breakpoint was in the lOlF region, occasionally in 102A, or even near 102B1-2 (proximal to this band, however), position effect was expressed. When the breakpoints occurred in the 102I3-F region, position effect was barely manifested (Stern and Kodani, 1955). The considered data show that chromosomal rearrangements that map within wide very cytological limits extending from lOlA to 102B cause position effect. If, indeed, inactivation of ci+ occurs because of removal of the locus from the influence of heterochromatin, the obvious suggestion is that the locus in the normal chromosome is under the effect of a-heterochromatin of the left arm of the fourth chromosome, (i.e., distantly located or in the lOlAC centromeric region). 2. A remarkable pattern in second breaks of other than the fourth chromosome was disclosed by Khvostova (1939, 1941) and confirmed by many investigators: position effect is elicited only when the second breakpoints in the other chromosomes are quite distant from centromeric heterochromatin (Figure 137). The data in the figure show that four or five numeral subdivisions near-

360

I. F. Zhimulev 0

X

P 2L

2R

3L

3R V2

m3

-4

*5

0 6

.’

Figure 137. A summarizing scheme of the translocation breakpoint distribution between the fourth and first through third chromosomesresulting in strong position effect manifestation according to the scheme for R(ci+).The horizontal lines represent Bridges’cytologicalmaps with number and letter subdivisions;all the centromeric regions are positioned at the right. Data from (1) Khvostova (1939); (2) Dzhataev (1973), Shavelzon et d.(19731, and Levina (1974) (mapped by I. F. Zhimulev from the photographs of Dzhataev); (3) Roberts (1969a, 1972a,b);(4) Stem and Kodani (1955); (5) Grell(1959); and ( 6 )Yamamoto (1987). Rearrangements producing position effect according to the R(ci) scheme are marked by symbol 7 (Stern and Kodani, 1955).

est the centromere in each chromosome are actually “vacant”; this means that the chromosomal rearrangements they contain do not cause position effect. There are exceptions to this rule. Four breakpoints of translocations were detected in the heterochromatic region 20 of the X chromosome (see Figure 137), which nevertheless gave position effect. Displacement of the fourth chromosome to the heterochromatic Y chromosome also evoked variegation (Dubinin and Sidorov, 1934a,b; Dubinin et al., 1935; Khvostova and Gavrilova, 1935; Panshin, 1936; Neuhaus, 1939; Khvostova, 1939). Mapping Parker’s (1965, 1967) translocations between the Y and fourth chromosomes, D. B. Benner (1970, in Spofford, 1976; Benner, 1971) found that even a part of the long arm of the Y chromosome (the short arm remains intact) distal to the kI-2 gene (see Section IYC) can cause variegation expression for the ci+ gene. However, by far not each and every fragment of the Y chromosome involving the fourth chromosome inactivates ci+. Only 4 of the 20 analyzed translocations showed position effect. It is unclear why the breakpoints in chromosomes 1-3 are so specifi-

Polytene Chromosomes, Heterochromatln, and Position Effect Variegation

361

cally distributed. The “vacant” regions are normally involved in chromosomal rearrangements (Khvostova, 1939) including the fourth chromosome; however, ci+ is barely, if at all, suppressed (Stem and Kodani, 1955). When a “vacant” region is placed by an inversion to a more distal position, and rearrangements are then induced in it, they are formed in the regions and elicite position effect (Khvostova, 1939). The pertinent data on the action of the Y chromosome on the expression of the position effect of ci+ are conflicting. It was found that an extra Y chromosome enhances the variegated expression of the ci+ gene, (i.e., it lengthens the gap in the cubital vein; see Figure 136), and the absence of the Y chromosome in XO individuals attenuates variagation ( Panshin, 1936; Khvostova, 1939; Benner, 1971). However, the situation proved not to be as simple as all that. It was also found that in R(ci+)/cifemales the addition of a single Y chromosome markedly decreases the break in the cubital vein, while the presence of two Ys more severely suppresses position effect. The general conclusion is that the Y chromosome modifies the position effect of ci just as the other genes (Grell, 1956, 1959; Altorfer, 1967). Position effect is enhanced in XY, R(ci)/ci individuals (Altorfer, 1952). It is not entirely clear how low temperature acts on position effect. There is information, however, that the variegated mutant phenotype is more expressed (Stern et al.,1946a), while the expressivity of the ci mutation is sharply increased in ci/ci homozygotes at a temperature of 19°C and lower (Bridges, 1935a; Stem et al., 1946b; Roberts, 1972a). Little is known about the cytological aspects of the Dubinin effect. Levina (1974, 1975) demonstrated that pulse incorporation of [3H]thymidine is decreased in the translocated homolog compared to its normal counterpart of the fourth chromosome. It was also shown that, in translocations with strongly inactivated ci+ (an extensive break in the cubital vein), a very large block of material fluorescent after staining with atebrin, a dye identifying blocks of AT-repeats, is detected in the lOlF region. When the Dubinin effect was weak, this band did not fluoresce. No changes in polytene chromosome structure (e.g., in its compaction) were found in variegating rearrangement both in XYY males at 25°C and in XO males at 14°C (Demakova et al., 1997). How should we envisage the situation when a gene normally functions in the neighborhood of heterochromatin and becomes inactivated after transfer to euchromatin? Cloning the ci locus and analysis of ci mutants revealed that they contain molecular alterations within a 13.7-kb region. Three mutations (ci’, ci361, and ciw) had insertions and one ( ~ i ~had ~ ga )small deletion. The ci’ mutation is related to the gypsy insertion. The dominant mutations ciD and Ce2each contain two insertions within the 13.7-kb region (Locke and Tartof, 1994).

362

1. F. Zhirnulev

A total of 4.8-5.5 kb of RNA is transcribed from the cloned DNA (Orenic et al., 1990; Locke and Tartof, 1994). The DNA sequence of the transcipt suggests that it may be a transcription factor with zinc finger domains (Orenic et al., 1990). Mechanisms of ci position effect variegation are still enigmatic. According to Dubinin and Sidorov (1934a,b) “weakening of dominance of ci+” is a consequence of changes of the gene position in the genome system. However, homozygous R(ci+)/R(ci+) or hemizygous R(ci+)/O rearrangement is phenotypically ci+. Thus this change in ci+ position is not sufficient for the inactivation. As an explanation of ci position effect variegation, Ephrussi and Sutton (1944) pointed out that the disruption of homolog pairing accompanies the reduction of the normal activity of the ci “structural model” (see Henikoff, 1994,for discussion). Locke and Tartof (1994) found that the ci phenotype is seen only when the ci’ allele (which is related to the gypsy insertion) is present [as ci’ or R(ci’)]. They suggested that the ci’ allele, and possibly the other recessive ci alleles, are gain-of-function mutations whose expression can be repressed by pairing with a wild-type homolog. Normal pairing can be disrupted in some translocations with the result that expression of ci’ in ci’/R(ci+) is modified. So, variegation may be a consequence of variation in the degree of ci’ pairing. Bearing in mind that the sup~essor-of-Hairy-oving[su(Hov)]locus suppresses different mutations induced by the gypsy insertion (Modolell et al., 1983), Henikoff (1994) proposed that special properties of DNA-binding su(Hw) protein bound to the gypsy transposon present at ci’ mediate interaction of two chromosome homologs (see Henikoff, 1994, for more detailed discussion). However, this model cannot explain new facts, witnessing that mutations ~ ici57g~and~ciw give ~ rise , to Dubinin effect in heterozygotes with different rearrangements R(ci+) (O.V. Demakova, 1997, unpublished). Comparison of somatic pairing of normal 4th chromosome with its homolog translocated either to vicinity of chromocenter or very distantly to it has shown that frequencies of pairing are much higher in former case than in latter (O.V. Demakova, 1997, unpublished). This fact witnesses for dependence of Dubinin effect on somatic pairing and explains the reason of suppression of the effect in rearrangements with proximal euchromatic breakpoints as it was found by Khvostova (1939).

B. The light (It) locus in D. melanugaster The It+ locus maps to pericentromeric heterochromatin in the mitotic chromosome 2L (Schultz and Dobzhansky, 1934; Schultz, 1936; Hilliker, 1976). Its location in the polytene chromosomes has not been so accurately determined. There is information that It+ is located between the 40B and D bands (E. S. Gersh, in Hessler, 1958) or 40F (Hannah, 1951). In situ hybridization of cloned It+ DNA showed that It+ is located proximal to the region showing distinct banding (i.e., within the re-


363

gion of P-heterochromatin) (Devlin et at., 1990b; Wakimoro and Heam, 1990). Thus the It+ gene functions normally when located in heterochromatin. Seven types of It transcripts (from 1 to 13 kb) were detected in various tissues and at different stages of development: in the ovaries of adults as well as the fat bodies, gut, malpighian tubules, and salivary glands of the larvae (Devlin et al., 1990a). A set of features render variegation at the It' gene exceptional:

1. Plum-2, a radiation-induced mutation (purple-brown eye color), was found to be associated with the inversion between the It (light eyes) and bw (brown eyes) genes. Either the It' gene is transferred to euchromatin or a lengthy region of chromosome 2R is transferred to its immediate vicinity (Figure 138). Regardless of whether the breakpoint of the inversion maps distal or proximal to It+, variegated spots appear in R(lt+)/ltindividuals (Schultz and Dobzhansky, 1934; Schultz, 1936; Morgan et al., 1937). Hessler (1958) generated 35 chromosomal rearrangements with one breakpoint Located in distal heterochromatin and the other in the euchromatic portions of the X, second, and third chromosomes (Figure 139); from his data it is unclear where the

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364

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breakpoint is situated. However, it was shown for at least one chromosomal rearrangement that It' and the block of heterochromatin to which it maps are, indeed, transferred to the distal (96EF) region of chromosome 3R. The entire heterochromatin of the 2L arm is concomitantly transferred. It was thought that position effect was evoked not by displacement of the It' genes from heterochromatin, but rather by their removal from the usual centromeric position (Hilliker, 1975, 1980; Hilliker and Sharp, 1988). Thus the It' gene normally functions when located in heterochromatin, but becomes inactivated when placed in euchromatin. According to proposals of Wakimoto and Heam (1990) and Howe et d. ( 1995), euchromatin-heterochromatin breakpoints cause effects on It' by reducing the amount of heterochromatin surrounding the gene. 2. The breakpoints in the euchromatic regions of the chromosomes map in an exceptionally specific manner to the middle or distal portions (see Figure 139), which is strongly reminiscent of the ci+ case (see Section XIYA). PEV of heterochromatic genes like It may be sensitive to conjugation with heterochromatin because distal euchromatic breakpoints most severely affect the ability of the displaced heterochromatin to contact with nonrearranged centromeric heterochromatin (Weiler and Wakimoto, 1995). 3. The response of genetic inactivation to change in the amount of heterochromatin in the Y chromosome is unusual. Variegation is completely suppressed in XO males and enhanced in individuals possessing one or two additional Ys (Morgan et al., 1932, 1934, 1935, 1941; Schultz, 1936, 1941a,b; Morgan and Schultz, 1942; A. Hedrick, M. Hearn, T. Grigliatti, and B. Wakimoto, in


365

Devlin et al., 1990a). Studies of the suppressive effect of various fragments of the Y chromosome demonstrated a reciprocal effect exerted on variegation expression in direct and reverse position effects. It was shown that fragments that most actively suppress position effect in Dp(1;3)wm264-58aare the most effective enhancers of variegation at It' (Baker and Rein, 1962). 4. Tests of the effects of 14 dominant modifiers of position effect variegation for their action on variegating genes normally located in euchromatin revealed that 8 had no detectable effects and 6 enhanced variegation of the It' gene (i.e., they acted as enhancers). It was suggested that the proteins of some of the modifiers are required for proper function of the It' gene (Grigliatti, 1991; Heam et al., 1991; C. Liep and B. T. Wakimoto, in Weiler and Wakimoto, 1995). 5. No temperature differences (using 17, 25, and 29°C) were found in the phenotypic expression of position effect at It' associated with the T(I ;2)w ~ chromosomal ~ ~ rearrangement ~ - (Gersh, ~ 1949). ~ 6. Genetic inactivation due to position effect of this type presumably does not spread over the nearby euchromatic genes. Full complementaion was demonstrated in heterozygotes for the variegating R( It+) rearrangements and the Df(2L)TW65 and Df(2L)TWI 61 deletions that remove the closely adjacent 37F5 to 39E2-Fl and 38F to 40A4-Bl euchromatic regions (Wakimoto and Heam, 1990). However, according to Hilliker (1980) and Hilliker and Sharp ( 1988), the type of position effect occurring at It' nevertheless cannot be regarded as the inverse euchromatic gene variegation. Moreover, it is unclear to what extent transfer of the gene from hetero- to euchromatin is critical to the expression of position effect of this type. When cosmid DNA 25-40 kb in length that contains the region where the It mutation is normally located was injected into mutant embryos, gene function was somewhat restored: when a single cosmid was injected, pigment (a product of the It' gene) was detected in 12 malpighian tubule cells of 310 larvae; injection of two other cosmids yielded similar results of 10 of 509 and 18 of 441 cells (Devlin et ul., 1990a). These data make it doubtful whether neighboring heterochromatin is necessary for It' to express under the condition that DNA has not inserted into heterochromatin during DNA transformation. It is of interest that the Plum mutation in D. ummsae and D. melanoguster is associated with a rearrangement in heterochromatin and variegated expression (Kikkawa, 1938).

C. Other loci in D. melanogasfer The rolled'(rl) gene, whose mutation causes rolled downward wing edges and small, coarse, dark eyes, lies in the heterochromatin of chromosome 2R (Hilliker

366

1. F. Zhimulev

and Sharp, 1988; Lohe and Hilliker, 1995); in the 41A region of polytene chromosomes (Lindsley and Grell, 1968); in the 41B1 region (Morgan et al., 1941); or, according to more recent data, deeply in heterochromatin (see Figure 33 in Section IV). Position effect is evoked when rl is displaced from the neighborhood of heterochromatin. Removal or addition of the Y chromosome does not modify the expression of rl+ in rearrangement (Morgan and Schultz, 1942). To assay directly the functional requirements of the autosomal heterochromatic genes to reside in heterochromatin, the rolled gene was relocated within small blocks of heterochromatin to a variety of euchromatic regions by series of chromosomal rearrangements. If rearrangements with a break proximal to rl removed the gene in a large block of heterochromatin, no visible position effect was found, but rearrangements reducing the size of the block of heterochromatin comprising the rl gene caused variegation of the gene. Displacement of the small block of heterochromatin containing the rl gene into or near a larger heterochromatic region in further rearrangement reverted gene activity (Eberl et al., 1993). The position effects of rolled are associated with uniformly lowered levels of rl mRNA among the cells of the eye and wing imaginal discs (A.J. Hilliker, in Lohe and Hilliker, 1995). In an early study, Hilliker (1980) indicated that, although transfer of the It’ locus from the centromere to euchromatin results in its variegated expression, when the genes neighboring It’ are transposed at the same time in this same rearrangement, they do not variegate. However, chromosomal rearrangements with breakpoints in heterochromatin of the second chromosome were detected later. Five translocations not fully complemented to the l(2LjEM.56-4 gene mapped between lt’ and the centromere (see Figure 33). The same rearrangement showed variegation at It+. There is therefore a probability that other “heterochromatic” genes can show position effect of the same type as ’It (Hilliker and Sharp, 1988). Subsequently, it was reported that several other genes-l(2L)40Fa, 40Fc, 40Fd, 40Ff, and cta-mapped to the heterochromatin of chromosome 2L can be subject to position effect upon transfer to euchromatin (Wakimoto and Heam, 1990; Eberl e t al., 1993). Modifiers that act as suppressors of the variegation of genes normally located in euchromatin exert an effect on the expression of activity state of the heterochromatic genes similar to modification of R(lt’)/lt; that is, the modifiers enhance position effect variegation instead of suppressing it. Su(v,ar)205enhances genetic inactivation of at least three (40Fa, It, and 40Ff), and Su(var)208 of five (40Fa, 40Fc, cta, It, and 40Ff) genes (Hearn et al., 1991).

D. The peach+ (pe) locus in D. virilis The peach (light eyes) gene is located between the most proximal band of the fifth chromosome and its centromere in D. virilis. It maps to the group of 4 bands nearest the centromere and is presumably located on the distal end of the large block of heterochromatin of this chromosome (Baker, 1952, 1953, 1954).

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mc n w u MCW 1 1 7 6xxSw-c ~ r.uPrIO* 01 d”uro Figure 140. Distribution of breakpoints of 30 chromosomal rearrangements producing position effect at the pe locus in D. uirilis. Hatched areas, centromeric heterochromatin. Numbers designate breakpoints of chromosomal rearrangements. If the vertical line crosses the chromosome, this chromosome region approaches pe; the remaining vertical lines designate the other associated breakpoints. After Baker

(1953).

In his study of 32 strains showing variegation at pe+, Baker (1954) found that 30 had rearranged chromosomes, with one of the breaks mapping to its euchromatic regions. As in the case of position effect of It’ and ci+ in D. melanoguster, the “euchromatic” breakpoints were in the distal regions of the chromosomes (Figure 140). The heterochromatic breakpoints are of interest. They were located between the pe+ locus lying on the distal end of the block of heterochromatin and the centromere limiting the proximal part of the block. Of particular interest was the T(3;5)pem4translocation. It broke off the whole block of heterochromatin at the centromere and brought it to the third chromosome in females. Thus position effect, in this case, resulted from interaction between the translocation break and the pe locus separated by the whole block of heterochromatin, which constituted 30% of metaphase chromosome length (Baker,

1954). There are data (Baker, 1953) indicating that an extra Y chromosome in the set does not appreciably affect variegation, although it can somewhat suppress it. Other data decisively demonstrate that an additional Y reduces position effect as the euchromatic genes do (Schneider, 1962).

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368

1. F. Zhlmulev

E. Male fertility factors As shown in Section IV, the Y chromosome is almost entirely composed of heterochromatin. For this reason, when translocations place the fertility genes composed of fragments of the Y chromosomes into other euchromatic positions in the genome, the genes can become inactivated in association with position effect. Spofford (1976) believes that male bearers, precisely because of position effect of 38 of 46 translocations between the fourth and Y chromosome generated by Neuhaus, are sterile due to variegation. The kl and ks fertility factors are not active in these translocations when transferred to the 101 region of the fourth chromosome (Neuhaus, 1939). Translocations between the fourth and the Y chromosomes in which kl in the YL arm was weakened were examined in another study. The degree of inactivation was modified by temperature in the same direction as in association with the usual position effect (D. B. Benner (1970), in Spofford 197611. In some D. hydei strains bearing translocations of fragments of the Y chromosomes, specific chromosome loops, morphological manifestations of the activity of the Y chromosomes in the spermatocytes of the first order (see Section VIII,A), affect morphology. Some types of changes are correlated with defects of spermiogenesis (Hess, 1970a).

F. The nucleolar organizer It is known that the nucleolar organizer (NO) (a cluster of the 18s and 28s ribosomal RNA genes) is located in the region of X chromosome heterochromatin. A number of inversions, sc8 in particular (see Figure 143 in Section XVI), transfer the NO from heterochromatin to the telomeric region of the X chromosome. The genotype of s$/O males then becomes lethal. Lethality is suppressed by the Y chromosome or its part. Hess (1962) suggested that in such cases the NO is subjected to position effect. Subsequently, Baker (1971) demonstrated that homozygotes for the sc8 and scsl inversions, as well as sc8/Y and scsl/Ymales, are viable. Removal of the Y chromosome is lethal in sc8/0, scLB/O,and scsl/Omales, i.e., variation in the number of the Y chromosomes in these cases produces the same effect as that associated with the usual position effect (see Johnson et al., 1979). Control sc4/0 males in which the inversion moves the distal block of heterochromatin to another position (see Figure 143), without affecting the ribosomal RNA genes, are almost as viable as sc4/Y males. It may be assumed that death is due to gene inactivation at either end of rearrangements. Inasmuch as the second breakpoint maps to the region of the sc+ gene, the possibility of gene activation in this region was tested by introducing a duplication overlapping this breakpoint. It was shown that the Dp ( 1 $1337, sc+ duplication does not affect the viability of bearers of inversions without the Y chromosome: scsl/Olarvae die during early development, only 30-50% of flies emerge, and virtually all die before


369

the end of the third larval instar (Baker, 1968, 1971). In the scs'/O first instar larvae, the amount of RNA constitutes 86% of the level observed in yw/O larvae, as shown by DNA-RNA hybridization. No concomitant decrease in the gene number (DNA amount) is then observed (Nix, 1973), which means that the genes are inactivated, not lost. In the course of generation of gynandromorphs for adult cuticle, small variegated spots of cells with a lost ring X chromosome (i.e., s$/O and scsJ/O)appeared amid the bulk of sc8 orscs' cells in the heterozygote with the R(l)wVC ring chromosome whose NO region was not unaffected. When one considers that loss of the NO causes cell death, the appearance of such gynandromorphs should indicate that either the NO functions nonautonomously, with the result that rRNA migrates from the normal neighboring cells to sc/O cells, or that the rRNA genes are not subject to position effect suppressors (Pyati, 1976). There is other evidence arguing for or against position effect of the ribosomal RNA genes. Zuchowski-Berg ( 1978) reported unintegrated ribosomal RNA genes, that is, rRNA genes lying outside the long DNA molecule in which the rest of the genes are situated. It is Zuchowski-Berg's view that the genes are not at all integrated into the chromosome, and the event may be variegation asociated. It was shown that wild Oregon-R X/O or sc4/0 larvae whose rRNA genes are unaffected had approximately 42% of the genes in unintegrated form, while there are no such short DNA molecules in the diploid nuclei. In scLB/Oand scs'/O individuals, the number of such low-molecular-weight fragments was sharply increased in the diploid cells of the brain and imaginal discs, and it reached approximately 42% (i.e., the same percentage) in the salivary gland chromosomes. The dominant position effect suppressor Su(var) suppresses the appearance of lowmolecular-weight DNAs containing the ribosomal RNA genes. A hypothetical explanation is offered: in the block of ribosomal genes in the salivary gland polytene chromosomes, underreplication produces chromosome breaks (see Sections VI,D and VII,C,2) making the gene block fall apart into a series of smaller molecules. When the Y chromosome is removed, underreplication is enhanced, which leads to the appearance of low-molecular-weight fragments in the diploid cells, too. As already shown in Section VII,C, 1, suppressors of position effect reduce the expression of underreplication so that these fragments disappear. Position effect for the nucleolar organizer may be expected in yet another case. The strong bobbed phenotype is expressed in In(I)BM'/O individuals, while homozygotes for the inversion do not show the mutant phenotype (Schalet, 1969). Retained activity in the nucleolar organizer and absence of appreciable morphological changes in numerous chromosomal rearrangements between hetero- and euchromatin in D. melanogaster and D. hydei seem to argue against the notion that the ribosomal RNA genes can be inactivated when transferred to the euchromatic regions (van Breugel, 1970; Hannah-Alava, 1971).

370

1. F. Zhirnulev

In transfer of a single ribosomal RNA gene into euchromatic locations by P-element-mediated transformation, it was shown that the genes function normally with respect to both transcriptional activity and ability to form a nucleolus. This was evidence that a heterochromatic environment is not the condition necessary for the expression of the ribosomal RNA genes (Karpen et al., 1988). Based on the obtained data, all types of variegation can be subdivided into two groups according to the effect exerted on the genes normally located in euchromatin and heterochromatin. A set of “rules” that seem to hold true for the majority of known cases of position effect was worked out (Schultz, 1941b; Morgan and Schultz, 1942; Lewis, 1950): Genes that are normally located in the euchromatic regions become inactivated when transferred to heterochromatin. Genes that are located in heterochromatin become inactivated when transferred to euchromatin. In the case of the “euchromatic” genes, addition of extra heterochromatin to the genome suppresses position effect and removal of heterochromatin enhances it. In the case of the “heterochromatic” genes, variation in the amount of heterochromatin in the trans position produces the revetse effect. This rule was derived from data on It+, and earlier on ci+ (Panshin, 1938; Schultz, 1941b; Steinberg, 1943; Lewis, 1950). Panshin (1938) believes that an extra Y chromosome extinguishes the effect of the inert region on its neighboring genes, and, in this way, leads to the norm in the variegating euchromatic genes and to a greater departure from normality in the variegating heterochromatic genes. The last rule, however, is supported by facts relating to the It+ locus only. Actually, in all the cases described in Sections XV,A-F, both types of position effects respond similarly to variation in heterochromatin amount. From the data for It+, ci+, and pe+, the most thoroughly studied variegating “heterochromatic genes,” yet another pattern is inferred: genetic inactivation occurs only when the “heterochromatic” gene is relocated at a considerable distance from heterochromatin.

G. Dominant position effects Several genetic systems exist in which the mutant phenotype is expressed in the R(g)/g+ heterozygotes; that is, the mutant allele starts to dominate when a chromosomal rearrangement appears in its surroundings. Some of the earlier descriptions of dominant position effect are summarized in Lewis’ (1950) review. Dominant position effect was detected presumably at the kar (Henikoff, 1979b) and Pu (O’Donell et al., 1989) loci in D. mehnogaster.


37 1

1. The ci+ locus As shown in Section XV,A, a break in the fourth chromosome between pericentromeric heterochromatin and the ci+ gene (i.e., proximal to the gene) is necessary for the Dubinin effect to occur. Information is provided concerning the weaker, yet detectable, position effect associated with the occurrence of a rearrangement in the ci chromosome, that is, in the R(ci)/ci+ situation. In this case, the breakpoint maps somewhat to the left of the 102B1-2 band. However, translocations with breakpoints to the right of 102B1-2 occasionally also gave rise to variegation, when in combination with R(ci)/ci (Stem and Kodani, 1955).The Y chromosome enhances the mutant phenotype produced by one of the rearrangements, giving rise to dominant position effect of ci+ (Altorfer, 1967). It is Spofford's view (1976) that a dominant suppressor of the ci+ gene is located in the 102B1-2 region. The recessive ci phenotype occurs in the case of its inactivation by a chromosomal rearrangement.

2. The bw+ locus Many authors have described the various genetic systems in which dominant position effect in the brown+ gene (brown color of eyes) is manifested. Muller (1930) was the first to demonstrate that the bw mutant phenotype produced by chromosomal rearrangements dominates over the normal allele bw+, even though all bw point mutations, even nulls, were recessive (Slatis, 1955a). Subsequently, the Plum-2 dominant mutation, which causes the formation of purple-brown eyes with darker spots, was induced by x-irradiation of males of Drosophila. The mutation proved to be associated with a chromosomal rerrangement between the light and brown eye color genes. Plum-2 is an allelic mutation of both genes (Schultz and Dobzhansky, 1934; Dobzhansky, 1936),and it exhibits variegation of four types: dominant and recessive eye color (bw),bristle damage, and light eyes (Schultz, 1936). Subsequently, numerous alleles of Plum, which were called bw", were detected. The majority are associated with chromosomal rearrangements. In heterozygotes for many strains, R(bw+)/bw+variegation for red and brown facets was identified; that is, the bw+ gene acts to suppress the normal bw+ allele when inactivated in a rearrangement in some cells. Variegation is suppressed by an extra Y chromosome (Glass, 1933, 1934). The bw dominant variegation was dealt with by Dubinin and Heptner (1934,1935). It produces homogeneous brown color in the heterozygote with the normal allele. When an extra Y chromosome is added (XXY females or XYY males), eye color changes from uniformly dark brown to spotted with irregular flecks scattered all over the red eye. By use of translocations of different regions of the Y chromosome, it was shown that any one of the three parts into which the

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I. F. Zhimulev

whole Y chromosome have been fragmented possessed this effect. It appears that Plum is the bwVJallele (see Henikoff and Dreesen, 1989). Noujdin's (1946d) explanation is given in Section XII1,B. Chromosomal rearrangements causing position effect at the bw+ locus were induced in other studies. Seven such rearrangements were recovered in crosses of irradiated bw+ males to bw+ females [i.e., R(bw+)/bw+heterozygotes], and 12 in crosses of irradated bw+ males to bw females [i.e., R(bw+)/bw].In all the cases of position effect, chromosomal rearrangements were detected in strains having one breakpoint lying within the 59C6-59F3 interval-that is, in the vicinity of the bw gene, which maps to the 59D4-59E1 region (Mickey, 1959; Lindsley and Grell, 1968)-and the other lying in pericentromeric heterochromatin of the Y and second through fourth chromosomes. Dominant variegation is enhanced with increasing distance of breakpoints from the putative location of the bw+ gene, and the "strength" is maximal in rearrangements with breakpoints six bands away from bw+ on either side (Slatis, 1955a). The dominant bd' allele discovered by Hinton was studied by Slatis (Hinton, 1940,1942a; Hinton and Goodsmith, 1950;Slatis, 1955b). In bwDpolytene chromosomes, inserted material was first thought to be a duplication of the 59El-2 band; it was later inferred to be insertion of a fragment of centromeric heterochromatin (Hinton, 194613;Hinton and Goodsmith, 1950; Slatis, 1955b). According to Slatis (1955b), there are three thick bands at bwD. The chromosomes with the insertion show breaks in 30% of cells in the bwD region (Slatis, 195515). Variegation is presumably due to adjacency to the insert, since the phenotype is inseparable from the material reverting to normal upon its loss (Hinton and Goodsmith, 1950). Recent studies on metaphase chromosomes of bwD homozygotes demonstrated that there are two blocks in the second chromosome after C-staining, one near the centromere and the other in the distal part. There is a large compact block of heterochromatin not showing a banding pattern in the 59E1-2 region of the salivary gland chromosomes (see Figure 49 in Section VI). HPl protein was found in this block (Belyaeva et al., 1997). The size of the heterochromatic insertion is about 2000 kb (Henikoff et al., 1995). Position effect in this system is best seen in individuals of the R(bw+)lbw+,st/st genotype in which inactivation of the bw+ gene, owing to interaction with the st mutation, leads to the formation of unpigmented cells of eyes (Slatis, 195513). For this reason, variegation is manifested as light brown facets on a white background. While white facets are related to inactivation of both bw+ and st+, light-brown pigmentation of the facets appears at the expense of some bw+ activity. Usually increasing the number of pigmented facets coincides with increasing the background pigmentation of the eyes from almost white to light yellow and orange (E. S. Belyaeva, unpublished). Chromosome rearrangements with strong bw position effect variegation result in lethality of homozygotes and heterozygote combination with each other. Only bwDdoes not affect viability (Lindsley and Zimm, 1992;Belyaevaet al., 1997).


373

Cis-effect (i.e., bw+ variegation) in rearranged chromosomes can be observed in R(bw+)/R+(bw)heterozygotes where R is a euchromatin-heterochromatin rearrangement. Trans-effect (i.e., bw' variegation) in normal homologs occurs in R(bw+)/R+(bw+). In this case degree of variegation is a sum of both cis- and trans-effects. Modifiers of recessive position effect variegation include variation of temperature, amount of heterochromatin, and influence the trans- and cis-inactivation in the same direction (for discussion, see Valencia, 1947; Brosseau, 1959). Cis- and, to a lesser extent, trans-inactivation in strains with inversions was shown to be increased at low ( 18°C) temperature, with a temperature-sensitive period before the third larval instar. Temperature influence on trans-inactivation in bwDlbw+was not found (Belyaeva et al., 1997). The E(uar)302 genetic enhancer increases inactivation degree. The level of bw+ activity is 22% and 31% of normal in females and males, respectively, in the strain with the beyVDe2rearrangement, while it is 9% and 12%, respectively, in the strain with the rearrangement and the enhancer. In the eight suppressors studied, all increase bw+ activity to 39-105% (Hayashi et al., 1990). A total of 150 dominant suppressors and 2 enhancers of bwD variegation were induced with EMS. They fall into two classes: unlinked suppressors suppressing both bwD and variegation of other mutations, and linked suppressors affecting bwD only. Of 111 suppressors tested, 87 suppressed the &inactivation of bw+ and 24 suppressed trans-inactivation. The Su(bwD)suppressorsstudied do not suppress telomeric variegation. Cytological analysis of the 16 suppressorslinked to bwDhas shown that 5 of them have breakpoints in 59E, 6 are translocations to the X chromosome, and 4 are translocations to the third chromosome with breakpoints clustered in the 52D-57D interval. One line did not show chromosomal rearrangements. In the X chromosome, breakpoints were scattered between 2B and 17E; the four breakpoints on the third chromosome are at the distal tips. All enhancers were related to translocations to heterochromatin vicinity (Talbert et at., 1994). Chromosomal rearrangements that move the bevD allelle further from centric heterochromatin suppress its trans-inactivation ability, whereas those that move bwD nearer enhance trans-inactivation. In such distance-enhanced lines, the beoD locus associates more frequently with the chromocenter in polytene salivary gland nuclei. In the interfase nuclei of larval neuroblasts, the 59E and heterochromatin are closer to each other in the W nuclei than in the wild-type nuclei (Csink and Henikoff, 1996). Enhancers of bevD were obtained when the Dp(2;2)59E, Byron duplication, containing a tandem repeat of bwD heterochromatin insertion and bw+ copy after irradiation, were relocated closer to autosomal heterochromatin. Conversely, 38 suppressors contain chromosome rearrangements moving this duplication further away from heterochromatin. The authors propose that W fails to coalesce with the chromocenter when its position along the chromosome places it beyond a threshold distance from heterochromatin (Henikoff et al., 1995). Lewis (1950) postulates a mechanism operating in dominant position ef-

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fect: inactivation of a gene by a rearrangement leads to accumulation of the precursor-substrate normally utilized by the gene, and the excess produces a dominant change in phenotype. Thus a reconcilation between dominant variegation and other variegated types of position effect as due to inactivation of the gene or its product would be reached. Spofford (1976) takes the view that the two loci affecting bw+ function lie to the tight or to the left of bw' in the rearranged regions producing maximum dominance effect. The Su(bw"') gene that maps to 10 bands distal to bw+ may be one such locus (see Lindsley and Grell, 1968). A new impetus to studies of position-affected bw gene came with the availability of molecular biological methods. The insertion causes a null mutation of brown. Using Northern blot hybridization, it was shown that RNA is not detected from both homologs in R(bw+)/bw+heterozygotes, thereby indicating that the rearrangement can exert an inactivating effect on the genes in both cis and trans positions. Trum-inactivation of bw shows remarkable strength: the eye of bwD/bw+flies has more than 98% brown ommatidia (Dreesen et al., 1988;Henikoff and Dreesen, 1989; Henikoff et al., 1993).The hypothesis that RNA is synthesized, but degrades as the result of interaction with bw+ antisense RNA (Frankham, 1988), is incompatible with experimental data. In fact, transcripts were not detected in strongly variegated rearrangements (Henikoff and Dreesen, 1989). An important finding was that heterozygotes for various rearrangements differ in inactivation degree exerted on the bw+ from homozygotes for a single rearrangement. The effect was markedly reduced in the former. Since chromosome pairing is considerably more disrupted in the genome when heterozygous with two different rearrangements than homozygous with one, the dominant position effect at the bw+was suggested to be due to disruption of precise pairing (Henikoff and Dreesen, 1989;Henikoff, 1990;Henikoff et al., 1993). In bwD/Su(Pm) heterozygotes, the small duplication of the bw region causes disruption of pairing in the region (Henikoff and Dreesen, 1989; Dreesenetul., 1991; Henikoff et al., 1993). It is the authors' view that a pairing-sensitivegenetic element (a transceiver) in the immediate vicinity of bw+ makes possible transmission of inactivation produced by juxtaposition of heterochromatin from the rearranged homolog to its paired counterpart. When one copy of the bou gene is cis-inactivated by heterochromatin, the regulator of bou expression on the other homolog might make frequent contact with heterochromatic proteins and thus be prevented from normal functioning. This hypothesis explains both the current data and those from the 1930s indicating that dominance of position effect at the bw+ gene is suppressed in interspecifichybrids between D. mehmguster and D. s i m h m hybrids (Morganet al., 1937) in which chromosome pairing is disrupted (for review, see Zhimulev, 199213, 1996). It is of interest that the beOD is unique among brown position effect alleles because an insertion of heterochromatin causes almost no disruption of somatic pairing of salivary gland and pseudo-nurse cell polytene chromosomes (Slatis, 195513; Belyaeva et d., 1997). This hypothesis did receive support from experiments with 35 transformants with full copies of the bw+ inserted at various ectopic sites of the Drosophi-


375

la genome. These experiments proved that copies of bw+ transposed to ectopic sites were not trans-inactivated by rearrangements affecting the endogenous gene (Dreesen et al., 1991). In cases when the bw position effect variegation was induced in an ectopic copy by new chromosome rearrangement, this copy became inactivated, whereas other copies of transposed bw' were not. In one line, a V21 bw transgene was inserted into the 92B region 55-70 kb away from the euchromatin-heterochromatin breakpoint. In this case the bw transgene showed "classic" type of PEV, that is, PEV resulting from the inactivating influence of nearby heterochromatin. PEV was enhanced when the transgene copy number or orientation changed. Removal of the bulk of centric heterochromatin from the vicinity of the P(bw+)array, (e.g., by inversion) fully reverts a variegated phenotype to normal (Sabl and Henikoff, 1996). Deletion experiments delineated the transceiver region to a 3.8-kb fragment comprising the bw gene open reading frame and about 1.2 kb of 5' flanking sequences (Martin-Morris et al., 1993). The bw gene in D. virilis have been cloned. It shows 86% identity of aligned residues with predicted D. melanogaster protein. The 9 kb of D. virilis genomic DNA containing the bw gene fully rescues D. melanogaster bw mutations (Martin-Morris et al., 1993; Martin-Morris and Henikoff, 1995).This 9-kb D. virilk fragment contains sequences required for trans-inactivation in D. melanogaster bwD (Martin-Morris and Henikoff, 1995). Cytological analysis of polytene chromosomes at the insertion site of the euchromatin-heterochromatin junction in the chromosome with bw" rearrangements showed that the neighborhood of heterochromatin leads to compacting of the chromosome region 59C to 59El-2 (in some cases to 59A). A heterochromatic insert into the bw gene location in the bwD strain also evokes compaction spreading proximally to the 59El-2 band that results in fusion of this band and the insert into a single block. When PEV is weakened, the 59El-2 band can be seen as a separate structure. The separate 59El-2 band is seen in part of the larval salivary gland cells of XYY males at 29°C and always in pseudo-nurse cell polytene chromosomes of otul I/otu'I bwD/+flies. Differences in degree of the genetic bw variegation, both cis- and truns-, in different rearrangements follow the sequence bwVDe'>bwVDeZ>bwVK. A correlation between level of cytological compaction and genetic variegation was found. At the same time there was no heterochromatization of bands in trans, that is, on the normal homologous chromosome with the bw+ gene. No HP1 protein was found in the 59El-2 region on normal chromosomes. This indicates that a block of heterochromatin does not produce visible changes of chromosome structure in the trans position (E. S. Belyaeva, unpublished; Belyaeva et al., 1997; Belyaeva and Zhimulev, 1997). These data support the idea of Henikoff and his colleagues (discussed earlier) on trum-sensitivity of the b w' regulatory element to the heterochromatin protein responsible for bw' trans-inactivation (i.e., for dominant variegation of the bw gene).

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3. Position effect of In(2LR)40d A case was described in which change in eye phenotype was associated with the inactivation of at least two unidentified loci controlling dominant dark eye color and modified structure of facets. An inversion with breakpoints in the 26D region of chromosome 2L and in the 41AB centromeric region of chromosome 2R was identified in the strain. Variegation is more strongly expressed at low temperature (Hinton, 1942a, 1949a; Demerec et al., 1941).

XVI. MOLECULAR AND CYTOGENETIC ASPECTS OF POSITION EFFECT VARIEGATION A. Historical consideration of hypotheses for the mechanism of gene inactivation under position effect variegation Belgovsky (1944, 1946) made the first attempt to classify the hypotheses put forward to explain position effect variegation. He assigned them all to two groups. The first group included the hypotheses attributing absence of genetic function to loss of the gene or its severe impairment. Patterson (1932b, 1933) introduced the concept that regions lose a chromosome fragment during the formation of variegated tissue not expressing gene activity because of instability of the chromosomal rearrangement. Noujdin (1935, 1938) supported this hypothesis at first, but he rejected it later. Schultz suggested that ring structures can result from position effect during gene replication. Rings can lose the genes they contain (Morgan et al., 1936, 1937, 1938; Schultz, 1936; see also Romanov, 1980). Subsequently, Schultz (1941b) abandoned the simplified explanation of gene loss because he thought that the chromosome regions containing the genes underreplicate in polytene chromosome for unknown reasons. Development of such hypotheses came more recently. Transposition of mobile elements, including those causing position effect occurring on the ends of chromosomal rearrangements, may be activated. Neighboring sequences can be carried away in the process of transposition, and deletions can form in the adjacent genes, and this may manifest as variegation of the genes (Spofford and DeSalle, 1991) Stern’s ( 1935) hypothesis postulates structural changes in the chromosomes brought about by somatic crossing over. Arisen mutations or small chromosomal rearrangements were proposed as causes of genetic inactivation (Muller, 1932; Sidorov, 1936, 1940, 1941a; Demerec and Slizynska, 1937; Belgovsky and Muller, 1938). These hypotheses imply a chimeric organism resulting from the presence of qualitatively different cells of various types. The second group of hypotheses postulates that cells are genotypically identical, with variegation producing a change in gene state only, The causes of variegation Belgovsky ( 1938) envisaged include a reduction in the biochemical

.

Polytene Chromosomes, Helerochromatin, and Position Effect Variegation

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activity of the gene placed closely adjacent to the “inert region,” and an increase in the “lability” of biochemical reactions evoked by juxtaposition of the gene to heterochromatin and due to variations in heterochromatization degree of the corresponding chromosome. Prokofyeva-Belgovskaya( 1937b, 1939a) holds the view that the chromosome region transposed to heterochromatin together with the inactivated gene undergoes the structural changes typically found in the “genetically inert” heterochromatin; that is, the gene is subject to “heterochromatization.” The concepts of genetic inactivation and heterochromatization were combined (Schultz, 1965; Gersh, 1973) to work out a mechanism of gene inactivation operating by compaction of the chromosome. DNA underreplication (DNA loss), heterochromatization, and compaction are considered in Sections

XVI, C-E Other hypotheses are known in which the effect of a chromosomal rearrangement impairing the structural-functional integrity in euchromatin or heterochromatin underlies a model of genetic inactivation. Dubinin (1939, Muller (1935,1938a), Offerman (1935), and Stem (1948) independently suggested that variegation may be interpreted as a consequence of changes in interchromosomal or intergenic processes resulting in gene product interaction of a new type in various somatic cells. This view was shared by Sakharov (1936), who assumed that variegation of the eye may be due to different position of the white+ locus with respect to the rest of the chromosomal material in various somatic cells. Dubinin (1936; see Belgovsky, 1944) regards variegation as a consequence of specific gene interaction between a certain gene and “inert” material (i.e., heterochromatin). It is Goldschmidt’s (1946) notion that the mutant phenotype results from a rearrangement break in a chromosome neighboring a normal locus. Koltzoff (1938) adopts the view that different genes are engaged in joint processing of substances that are their products in each chromomere. Hence, insofar as gene surroundings are changed by variegation, increased supply of these products is disrupted. Panshin (1938) came to the conclusion that somatic variegation can result from chance fluctuations in the concentration of heterochromatic products in the region of the w+ gene in different cells, presumably because heterochromatin is differently disposed in w+. Jeffery (1979)takes the view that, in cases of rearrangements giving position effect, a system of modifiers and polygenes affecting the gene of interest can be altered in any rearrangement, and the gene becomes inactivated. Hypotheses are known that explain variegation by changes in chromosome structure in a rearrangement breakpoint. According to one such hypothesis, somatic pairing in the chromosome regions adjacent to the breakpoint is disrupted. The forces leading to pairing are redistributed in new directions at either side of a break, bringing the gene into a “stressed state” (Muller, 1941, 1947; Ephrussi and Sutton, 1944; Gersh and Ephrussi, 1946; see also discussion in Hinton, 1946a; Gotdschmidt, 1952). Another hypothesis accounts for

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position effect by change in the spatial organization of the nucleus. This hypothesis implies that, because the spatial distribution of the chromosomes in the nucleus is invariable (for greater detail, see Zhimulev, 1992b, 1996) on the one hand, and because there is a gradient in the distribution of molecules activating the genes and maintaining them in a differentiated state on the other hand, a chromosome region does not necessarily have to be compacted (subject to position effect). Change in its position in the nucleus is sufficient for the production of variegation (Spofford and DeSalle, 1991) . Herskowitz ( 1961) associates genetic inactivation with a rearrangement break of particular nucleotide sequences, while Taylor (1964) suggests that DNA structure in euchromatin is specific in that it is covered by “activating” and “repressive”histones in heterochromatin and euchromatin transferred to it. Most hypotheses are of interest in historical retrospect because many of the molecular and genetic mechanisms disclosed later could not then be incorporated. In spite of this lack of information, analysis of the models appears worthwhile, reducing the search for the causal mechanisms of position effect to two lines of pursuit: (1) detection of molecular and cytogenetic features of heterochromatic regions to which the chromosomal rearrangement transfers the gene in question, and (2) elucidation of what might actually be happening to the gene (at the cytogenetic and molecular levels). The majority of these hypotheses now have only historical interest. Modem ideas are considered in the next sections.

B. Potential inactivation capacities of various regions of heterochromatin 1. Pericentromeric heterochromatin As Muller (1930) observed in his first paper bearing on variegated position effect, and as confirmed later by many investigators, variegated expression is associated with gene transfer to the region of pericentromeric heterochromatin by a chromosomal rearrangement (see Section XII). Thus comparison of two groups of translocations (seven in each) moving the w+ gene to eu- or heterochromatin, respectively, revealed complete absence of variegation in the first and its presence in the second group (Demerec, 1941a,b). Based on analysis of 312 chromosomal rearrangements known to cause variegation, Baker (1968) came to the conclusion that the two breakpoints are located beyond heterochromatin in only five strains and that these exceptions are due to inaccurate localization (see next section of this chapter and Table 32 for discussion). He also concluded that it is insufficient just to transpose the gene to the neighborhood of heterochromatin for a rearrangement to evoke variegation. The requirement of a rearrangement is a broken heterochromatic region. Attempts were made to discredit the role of heterochromatin in induc-


3 79

tion of genetic inactivation (Griffen and Stone, 1938, 1940a). When a fragment of the X chromosome stretching from the telomere to 3C2 is brought to the fourth chromosome by the w* translocation, there is no visible, at least, contact with heterochromatin. In w* retranslocations obtained by x-ray mutagenesis, heterochromatin also was not seen in the new breakpoints, although the w+ gene still variegated. Subsequently, complete absence of heterochromatin at the junction between w+ and the fourth chromosome was not confirmed, and the possibility that a small heterochromatic fragment was transposed together with the site of the w+ gene in the retranslocations cannot be overlooked (see discussion in Kaufmann, 1942). Relevant data can be obtained by studying revertants for position effect. In irradiated bearers of a chromosomal rearrangement causing inactivation, the phenotype can revett to normal. Cytological analysis of these strains demonstrated the following: 1. Irradiation of T(I ;4)wm” in which the white gene is transferred to heterochromatin of the short arm of the fourth chromosome induced 8 complete and 30 partial reversions. The w+ gene was retranslocated to euchromatin in 7 of the 8 analyzed strains and to the distal heterochromatin of the Y chromosome in one strain. In the case of partial reversion (variegated eye color only decreased, without being entirely abolished), retranslocations also arose in euchromatin. In generated revertants toward stronger inactivation of the white+ gene, additional heterochromatin appeared in the new rearrangements (Panshin, 1938). 2. Transposition of a rearrangement breakpoint to a new position was also detected in every one of the more than 40 revertants variegating to normal at the w* gene (Griffen and Stone, 193913,1940a). 3. In analysis of 17 complete or partial In(l)rst3 reversions, transfer of the gene from hetero- to euchromatin was found in all cases (Kaufmann, 1942). 4. In the majority of 24 irradiation-induced In(ZLR)40d male revertants, a new chromosomal rearrangement appeared. When it did not appear, the expression of position effect variegation was unaltered. The author concludes that heterochromatin is the principal, if not the sole, factor causing position effect (Hinton, 1948, 1949a, 1950). 5. In 5 1 lines in which revertants were generated by irradiation in In(]) wm4, variegation of the w+ reverts to normal, although the cytological pattern differs from that described previously. Thirty-three lines were reinversions in which the w+ had been brought to a new position in X euchromatin, 4 were translocations, of which two were transferred to the 40-41 heterochromatic region of the second chromosome; and cytological changes were not seen in 14 strains (Reuter et al., 1985). 6 . Revertants were generated by irradiation of T(I ;2)hrvaT7causing variega-

380

7.

8.

9.

10.

11

12.

1. F. Zhimulev

tion of all the genes distal to the 2B7-8 region of the X chromosome. In two strains, the 1A to 2B7-8 fragment was prevented from making contact with the main block of heterochromatin and joined to the euchromatic 19A [Dp(l ;f)dOrTev2’ and Dp(1 ;f)dorrev226] regions. The reversions to normal expression were stable, even when induced by various enhancers of position effect (Pokholkova et al., 1993a,b). The ca74 deletion, which exhibits position effect variegation at the Acph-I gene, brings it into the vicinity of the heterochromatic part of the chromosome enriched with highly repetitive or satellite DNA (Frisardi and MacIntyre, 1984; Shafer and MacIntyre, 1990). The PZ transposons containing a P-element-lac2 fusion gene that functions as an “enhancer trap” (O’Kane and Gehring, 1987) do not show any expression of P-galactosidase in salivary gland cells when inserted into chromocentral DNA of D. melanogaster (Zhang and Spradling, 1995). An obvious inactivating effect of heterochromatin was demonstrated in transformation experiments. When w+ transgene was inserted at particular heterochromatic positions of the w strain of Drosophila, the activity of the normal allele was considerably suppressed, and this conferred lighter than normal eye color. The w+ gene was then not affected, but rather profoundly inactivated; when the transposon was moved to new positions, w+ activity was restored provided that the transposon had inserted into euchromatin (Levis et al., 1985). The rosy gene was partly inactivated when the transposon R401.1 was inserted into heterochromatin of the fourth chromosome (Spradlingand Rubin, 1983; Daniels et al., 1986). The minichromosome Dp(3;f)Th in D. melanogaster, which is mitotically unstable, was generated by x-ray mutagenesis at the expanse breakpoint deep in the pericentric heterochromatin within or very near to the DNA sequences essential for centromeric function. Nondisjunction of this minichromosome is mosaic and is possibly related to interaction of heterochromatic sequences and sequences important in centromere function (Wines and Henikoff, 1992). When fragments of centromeric heterochromatin are transposed to euchromatin, they can exert an inactivating influence on any genes that happen to be in the vicinity. It is believed that, when chromosomal rearrangements are reversed, a small region of heterochromatin can be carried away and placed at a new position (Panshin, 1938; Griffen and Stone, 1940a; Kaufmann, 1942;Jeffery, 1979). In the case of direct reversions, the transposed heterochromatin cannot affect variegation, however; potential inactivation capacities are detected only when rearrangements are repeatedly generated in these strains (McClintock, 1951),or when position effect is intensified by an enhancer (Reuter et al., 1985). Study on the bwD strain causing position effect at the bw+ locus revealed an


381

insertion of heterochromatin in the 59E region (for greater detail, see Sec. tion XV,G,2). In the In(2LRj40d inversion, the heterochromatin of the base of chromosome 2R was transferred to the 27 region of chromosome 2L. A new inversion, In(2LR)I ICQ, was derived from it. Its heterochromatin, located between bands of the 23A and 23B regions, exerts an inactivating effect on the nearby (5 cM away) rubroad (rub) gene (Xnderholt and Hinton, 1956). A minor position inactivation was reported for the A&+ gene in whose vicinity fragments of Y heterochromatin were inserted in T ( 2 ; Y ) translocation sites of D. melanogaster (Hisey et al., 1979). 13. In a population of D. ananassue, chromosomal rearrangements were detectable in one or two salivary gland chromosomes in some individuals. It was found that this mosaicism correlates with the presence of a large block of heterochromatin in the proximity of a puff. The appearance of the block led to its inactivation in one case. It was not inactivated in the second puff (Mukherjee and Dutta Gupta, 1966). However, puff inactivation was unreliably documented in the former case. 14. A population of D. imeretensis (lummei) composed mostly of individuals heterozygous for an inversion, and partly of individuals (31%) also heterozygous for an inversion of a large block of heterochromatin in the same chromosome arm, was identified. The block and the inversion were maintained in a balanced state in a lethal system (Mitrofanov and Poluektova, 1982; Poluektova et al., 1984). The presence of a block of heterochromatin was associated with a decrease in Jh- and P-esterase activities, probably encoded by the genes located in regions adjacent to the block (Korochkin and Evgen'ev, 1982; Korochkin et al., 1983). The data in question provide evidence for the exceptional role of heterochromatin in gene inactivation. It was shown that pericentromeric heterochromatin in all the chromosomes is potentially capable of inactivation. Of the 30 independently generated chromosomal rearrangements causing position effect variegation, the w+ gene was transposed to heterochromatin of the X and 3R chromosomes in six rearrangements and to the 2L heterochromatin of chromosomes 2L, 2R, 3L, and 4 in six other rearrangements (Demerec, 1941a,b). It was demonstrated that the Y chromosome possesses an inactivation capacity (Slatis, 1955a; Mickey, 1959; Lindsley et al., 1960). How is the potential inactivation capacity of heterochromatin manifested? Genetic inactivation extent may be dependent on heterochromatin amount. Figure 141 presents the structure of the wm+ll translocation and its various derivatives. It is seen that the inactivation extent of the w+ gene correlates with heterochromatin amount. The eyes of flies with the structure depicted in Figure 141a are of normal (Figure 141b) or almost normal (Figure 141c) color, are strongly variegated (Figure 141e), or are intermediate between these types (Fig-

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1. F. Zhlmulev

a

*W I I

I

C

6

m e 2

tw

C

7 .E

d XP

e,

P 9

4CX

h Figure 141. A scheme illustrating proportionality between heterochromatin amount in the neighborhood of the W+ gene and activity of the gene (see text for explanation). After Panshin (1938, 1941).

ure 141d); eyes can also be almost unpigmented (Figure 141f and 1419) or they can be rendered completely white (Figure 141h) (Panshin, 1938, 1941). There is other evidence for the dependence of genetic inactivation on the amount of adjacent heterochromatin (Kaufmann, 1942; Hinton, 1950). Although any large block of heterochromatin can inactivate the genes transposed to it, since Demerec's time (Demerec, 1941a,b) it has been recognized that certain parts of heterochromatin are more effective in inducing variegation than others. There is every reason for believing that the qualitative composition of heterochromatin is of greatest importance in inactivation of the transposed gene.


383

The abundant results indicating that different regions of heterochromatin possess different potential capacities to inactivate the genes are considered next. The so-called “specificity” or chromosomal rearrangement can significantly differ in “inactivation strength,” the degree to which the gene is turned off and inactivation extended. For example, position effect can spread to include more than tens of bands in some rearrangements (see Table 28, in Section XII), while inactivation in In(] )rs$ does not extend further away than the w+ gene (the distance of one band), even in the case of strong enhancement in XO males (Gersh, 1963). Furthermore, in two chromosomal rearrangements numbered 264-29 and 264-55, one euchromatic breakpoint maps to the 3D4-5 region and the second, respectively, to the 3L and 3R heterochromatin. The mt+, fa+,and dm’ genes (i.e., from 3D4-5 to 3C4 on the cytological map) are inactivated in the 264-29 rearrangement, while the inactivated genes are w+, rst+,fa+,and dm+ (i.e., from 3D4-5 to 3C2) in the other rearrangement (Demerec, 1941a,b). Of the four inversions with a heterochromatic breakpoint in the region of the white+ locus, the heterochromatic breakpoint was distal to the nucleolar organizer in two inversions and proximal in the other two. The former showed variegation; the latter two did not (Demerec, 1941a,b). Two different inversions in almost the same region of the X chromosome, Dp( J ;3)N264-58and Dp( I ; 3 )~ ~ appreciably 4 ~ differ ~ in the , inactivation extent of the vital genes located in them. This is presumably associated with the particular heterochromatic site into which these fragments were inserted. The first duplication mapped between the 80C and D bands of chromosome 3L and the second to the base of the 81 region of chromosome 3R (Ratty, 1954). Two similar inversions with “euchromatic” breaks in the neighboring bands, In(lLR)pn2a and In(ILR)pnndb, considerably differ in inactivation extent of the genes located in the neighborhood of the breakpoint (Tolchkov et al., 1984). When a duplication of the terminal region of the X chromosome containing the sc+ gene was formed in the short arm of the Y chromosome, bringing the gene into direct contact with the bobbed locus (the NO) and the entire Y heterochromatin, the expression of sc+ was not altered (Crew and Lamy, 1940). In the reinversion no great differences in localization from the original In(] )wm4 were found, although the extent of genetic inactivation at the w+ locus was considerably increased (Schultz, 194313). In a large-scale experiment, 577 T(Y;A) translocations between the autosomes and the Y chromosome were generated. Actually any euchromatic region of the second and third chromosomes in any one of the translocations was brought to the proximity of the large fragment of the Y chromosome. No special analysis was performed to detect position effect or its enhancement; there is no information on the inactivation of any genes happening to be in the vicinity of the Y heterochromatin (Lindsley et al., 1972). Seventy-five translocations were generated

384

1. F. Zhimulev

between the X and the Y chromosomes in another study. There is also no information as to whether position effect was detected for any of these (Stewart and Merriam, 1973). In a random screen of x-ray-induced mutants of Dosophila, 1527 lethal X chromosomes were cytologically analyzed. Of these, 40 had rearrangements between eu- and heterochromatin, and only one showed variegation under standard conditions (i.e., when temperature and heterochromatin amount were not varied) (Lefevre, 1981) . It might have been expected that, when deletions are formed between centromeric heterochromatin and the euchromatic regions, the euchromatic genes would become inactivated when brought closer to heterochromatin by deletions. Four deletions with the proximal breakpoint located between the "bobbed" locus and the centromere were generated. There is also no information on position effect exerted on the genes adjacent to the distal breakpoint of these deletions (Lefevre, 1981). Material of the distal end of the X chromosome was moved to the Y chromosome to generate translocations. No clear-cut case of position effect was found in any of the 19 strains (E. S. Belyaeva et al., 1982). Evidence that the various heterochromatic regions are qualitatively different in their capacities to induce genetic inactivation was provided by Pokholkova et aE. (1993a,b). The chromosome with the T(l ;2)durvar7 translocation in which the 1A to 2B7-8 fragment contacting the 2R heterochromatin and the genes located in the 2AB regions became inactivated were irradiated, and reversions were generated. The following types of revertants could be then recovered:

1. The 1A to 2B7-8 fragment was transposed to the euchromatic 19A region [Dp(l ;f)Mu27 and Dp(I ;f)dOrreu226],and position effect was no longer produced even under the influence of enhancers. The transposed fragment care ried about 20 kb of DNA with it from its old position in heterochromatin, and was therefore incapable of causing variegation at a new position. Dp(I ;f)dOr.lev4", Dp(I ;f)dorreu6",Dp(1 ; f ) d ~ l ~ ~ " ~ ~ , 2. In five cases, Dp(1 ;f)Mu3, , the 1A to 2B7-8 fragment together with the 20-kb and Dp(l DNA from heterochromatin was placed into the X heterochromatin. The bulk of the block of the X heterochromatin then remained unaltered, as did the whole block of the rev60 duplication. Also, in spite of this, position effect arose anew only under the influence of enhancers. 3. In T(1;2)d~r""'~, heterochromatin of the second chromosome was transferred to the 2B7-8 region of the X chromosome. Compaction and genetic inactivation, as noted earlier, spread toward the telomere and occasionally the centromere. When a reversion in T(l;2)d0rreu4' resulted from two inversions with a heterochromatic breakpoint in chromosome 2R, one of the regions of

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation _____

4.

385 ~

the 2R heterochromatin near 1A to 2B7-8 in the T(J ;2)dorYQr7 chromosome was transposed to the 2B7-8 to 7A fragment. The distal fragment ceased compacting and the proximal compaction was very strongly expressed. Thus it may be concluded that the heterochromatic center of compaction has moved from one position to another in T( J ; 2 ) d o ~and ~ ~reversion ~ ~ ~ , of position effect to normal in the distal region of the X chromosome has resulted from removal of the compaction center from its neighborhood (Pokholkova et al., 1993a,b). In other stildies, revertants were obtained after irradiation of Dp( J ;j9dmreo6O. Using Southern blot analysis, it was shown that a heterochromatic sequence at least 17 kb long joining euchromatin in “parental” Dp(l;f)durreY6Ois still present in revertants (Belousova and Pokholkova, 1997).

Data on breaks in heterochromatin followed by analysis of the potential capacity to inactivate the broken ends, for example, when small fragments of euchromatin are inserted in heterochromatin, are of interest. In such cases, inactivation can spread from the terminal regions of the insertion toward its center (Panshin, 1938; Demerec, 1941b; Cohen, 1962). This indicates that heterochromatin has the potential capacity to inactivate at either side of the break. In other cases, inactivation is detected not on both ends, but only on one. For example, the w+ gene, located in the middle part of the 2E14A1 fragment comprised from both ends by heterochromatin of the second chromosome [ T ( J ; ~ ) w ~ is ’ ~not ~~] activated. This indicates that either there is no inactivation or it is too weak to reach the w+ gene (Lefevre, 1970).The R40J . J transposon contains the DNA of two genes, l(3)S J 2’ and rosy+. Upon insertion into heterochromatin of the fourth chromosome (Spradling and Rubin, 1983), the rosy gene is to a large extent inactivated and l(3)S12 is not (Clark and Chovnick, 1986). In translocations or inversions causing position effect, heterochromatin, by definition, is broken; it joins euchromatin with the result that two euchromatic-heterochromatic junctions arise. Heterochromatin at one junction frequently has an inactivating influence on euchromatin, whereas that at the other junction does not. For example, inactivation in In(JLR)pnnZa and In( JLR)pn2b (Tolchkov et al., 1984) and T(J;2)doroQr7(0.V. Demakova and E. S. Belyaeva, unpublished observations) spreads in one direction. In a comparison of severity of PEV in seven translocations carrying w+ transgenes inserted at the euchromatin-heterochromatin junction, it was shown that strength of the position effect exerted on the transgene was not correlated with the quantity of the heterochromatin retained at the junction (Howe et al., 1995). After removal of the majority of centric heterochromatin from vicinity of the largest P(bw+) array located in 92C and showing strong inactivation of bw+, a full revertant to normal (X98), insensitive to enhancer E(var)66, was obtained. In salivary gland polytene chromosomes of this “healed” line, heterochromatin is

386

1. F. Zhlmulev

still cytologically visible in the 92B5-10 region (Sabl and Henikoff, 1996). Probably this fraction of heterochromatin lost its ability to induce position effect variegation. In agreement with Lewis (1950), there is reason to conclude that gene transposition to heterochromatin is a necessary, albeit not consistently a sufficent, condition for the variegated phenotype to arise. This raises the question: Is there specificity in the structure of the heterochromatic DNA regions closely relocated to the genes transposed together with them? Spofford (1976) thought that a tentative subdivision of sequences is possible between the bases of the metacentric autosomes and the fourth chromosome, between the regions proximal and distal to the nucleolar organizer in the X chromosome [see earlier discussion of Demerec's (1941a,b) work for information on inversions], and between the regions of the Y chromosomes. However, there are facts supporting the idea that the inactivating property is continuously distributed in heterochromatin. There are many rearrangement breakpoints in rst3, wm4, wmy, m4, and sc4, among others, within heterochromatin of the X chromosome (Gersh, 1963; Spofford, 1976). Moreover, the disposition of a number of genes has been physically mapped. This makes it possible to map the distribution of the potential inactivation capacity in a block of heterochromatin (Figure 142; Table 31). True, systematic studies of the potential capacity to inactivate the various heterochromatic fragments were not performed, and there are too many blanks in Table 3 1. Nevertheless, analysis of the presented data allows us to make at least two conclusions: (1) all the heterochromatic fragments exert position effects on the neighboring genes, and (2) the capacity to inactivate is most frequently distributed on either side of the break.

Figure 142. Cytogenetic map of centromeric heterochromatin of the X chromosome of Dosophila

and certain genes subjected to position effect variegation (original scheme). (a) At the left is shown the map position on the X chromosome of the y+, a+, and sc+ genes (slanted hatching) and breakpoints of inversions:I n ( l ) ~In(1)scv2, ~~, In(l)sB, I n ( l ) s P , and In(1)sc'f (Campuzano et d., 1985). In the middle part of the chromosome is shown the location of the w+ gene (double hatching) and chromosomal rearrangements In( 1)wm4, In( 1)wd b, and In( 1)wmMc. ANOWS determine the size of the w+ gene and the direction of the transcription (Tartofet d.,1984, 1989). Locations of rst (dotted) and In(l)rst3 (Emmens, 1937a) are not designated on the molecular map. In the right part, the heterochromatic block (black) and breakpoints of the rearrangements are depicted. The disposition of the various DNA sequences with respect to breakpoints of rearrangements are shown: insertions in the ribosomalcistron of type I, the 18s and 28s rRNAs, and the 1.688, 1.705, and 1.672 satellites. The numbers over the chromosomes mark the heterochromatin fragment number between the breakpoints of inversions [according to the data of Hilliker and Sharp (1988) and Tartof et al., (1989)l. (b-h) The locations of the same DNA fragments in strains with y3p (b), scvz (c), SB (d), rsc3 ( e ) ,wm4 (0,wrnsJb(g), and wmMc (h) rearrangements.

388

1. F. Zhimulev

Table 31. Inactivating Potential of Various Fragments of Heterochromatin (Based on Figure 142)

Heterochromatic blocks

Rearrangement

1-2

rsP

1-3

wm4,

1-4 1-5 1-6

Contact of blocks with the nearest genes from the proximal side

W+

w*

Data on gene inactivation in contact sites of euand heterochromatid w + - o n l y of X / O indi-

viduals (Gersh, 1963; Lindsley and Grell, 1968)*

*

scv2

Genes distal w+ ac+ from the proximal side

ac+ (Lindsleyand Grell, 1968)*

wm51b

Genes distal to w+

*

Y3p

It is unclear whether y + is transposed to heterochromatin or remains

y + (Lindsley and Greil,

1968)*

in the telomeric region 1-6

sc8

ac+

y+, a+(Lindsleyand

3-8

rs$

rst+

rst+ (Lindsley and Grell,

W+

**

Grell, 1968)* 1968)** 4-8 5-8

wm4,

scv2

sc+

sc+ (Lindsley and Grell, 1968)**

6-8

Wm51b

W+

w+ (Lindsley and Grell,

wd=

1968;Tartof et al., 1989)

7-8

Y3p

7-8

SC?

sc+ (?)

a+, y + (Lindsley and Grell, 1968)** sc+ (Lindsley and Grell, 1968)**

Note. There are no data on inactivation of genes distal (one asterisk) or proximal (two asterisks) to contact site.

The influence of distinct fragments may be considered separately. Information pertaining to the nucleolus (a block of repetitive rRNA genes) as a putative inducer of position effect has been accumulating for years. Demerec (1941b) was the first to report about the 265-52 and 264-84 rearrangements that had a heterochromatin break and were in close proximity to the NO and started to exhibit variegation. The view was held that the NO, when transposed and carrying some amount of heterochromatin to the ct or Ir genes with it, can cause their variegation (Hannah-Alava, 1971). In D. hydei, when a fragment of the chromosome

389


containing the w+ gene is transposed, inactivation takes place between the two parts of the nuclear organizer in the ring Y chromosome R(Y)wm (Beck et al.,

1979). Arguments in favor of the potential inducive capacity of the NO and the chromosomes of D. mehogaster appear convincing at first glance. For example, the wm4or wdlb inversions place thew+ gene right into the NO (see Figure 142), and, breaking the middle of the NO block, the scv2 inversion transposes sc+ to the site where it directly makes contact with the block of the rRNA genes; the other half of NO contacts with the K + gene. Both sc+ and uc+ are then inactivated. As seen in Figure 142, the NO is the nearest neighbour of only the inactivated genes in all cases. It therefore cannot be ruled out that the inactivating effect is exerted by the whole block of heterochromatin that includes the NO. Finally, the short arm of the X chromosome of Drosophila, also encompassed by the block of heterochromatin, produces an exceptionally strong effect on variegation at the gene in the 3C3-6 region included in the 1 n ( l L R ) l - ~in’~~ version (Gersh, 1965). To gain insight as to how the genes are inactivated, the structure of DNA immediately adjacent to the breaks must be known, among other things. Studies on 50-kb DNA in which the white+ gene and the breakpoints for the variegating wm4, wdfb, and wmMcinversions are located (Figure 143) revealed that all three inversions are clustered in a 3-kb range (between -24.5 and -21.3 kb), at a distance of about 25 kb downstream from the promoter w+ gene (between -2 and + 4 kb). The only repeated sequences in the studied region (from -35 to 15 kb) lie in the breakpoint site (between -23.5 and -24.3 kb), and they represent approximately 2.5 copies of the 1.688 satellite repeat usually mapped to the chromocenter. It was suggested that the X chromosome in the spermatocytes is folded in such a way that copies of the 1.688 satellite in the 3C region and heterochromatin are brought into proximity to each other, thereby creating preconditions for inversion formation (Tartof et al., 1984, 1989). When the so-called heterochromatic breakpoints of the wm4 and wmMc are included within a mobile element type I insert into an rRNA gene, wm4 DNA becomes contiguous with the left end of the element and wmMcDNA with its right end. The breakpoint of wdlb becomes contiguous with another mobile element, which additionally has 16 location sites in the euchromatic arms of the other chromosomes. Thus, although making contact with similar heterochromatic fragments, wm4 and wmMcshow different variegating patterns (sectored in wm4 and “peppered” in wmMc)(Tartof et ul., 1984, 1989). The B1 O4(roo) mobile element was detected in the Zn(lLR)pnnZa inversion causing mosaic expression on both ends (Alatortsev, 1986, 1988). Revertants can be used in another approach toward an understanding of the role of DNA structure in genetic activation due to position effect. The three reinversions of wm4 induced by by x-ray mutagenesis did not express the mutant

+

390

1. F. Zhimulev m51 b wm4v WmMc

I

S

b

I

EE I t

t': t

wrn4

BBS I II

I

I

I

Wf

I

I

I

I

I

TRANSCRIPT

...

WmMc

,mSlh

C

S S S EE

.*

,,'"'3

H

\,' lIlM(.

Sma

H

+... H

H

+E

,('Ill5

Ih

BBS

H

Figure 143. Molecular-genetic mapping of the breakpoints of the w"'+, wrnMc,and wZnSlb chromosome rearangements. (a) T h e gray arrow indicates the direction of transcription of the w+ gene. (b) Black rectangle indicates the location of the 1.688 satellite. (c) Restriction map; S designates restriction site SalGI; E, EcoRl; B, BamHI; H, HindlII; Sma, SmaI. After Tartof et al. (1989).

phenotype, although mobile DNA and at least 3 kb of heterochromatic DNA were transferred together with the w+ gene to a new location. On the basis of this evidence, it appears that the DNA of X-heterochromatic sequences more distant from the rearrangement breakpoint plays the major role in causing position effect (Tartof et al., 1984, 1989). Using the same chromosomal rearrangement of the wm4 gene, Reuter et al. ( 1985) obtained 37 reinversions, each transposing w+ from heterochromatin to euchromatin and thereby restoring its normal function. When the strong en-


391

hancer E - ~ a r ( 3 ) 2was ~ ~ introduced into the genome, many revertants started to variegate anew. This clearly indicated that the sequences immediately adjacent to the gene are decisive in inactivation, when brought to heterochromatin. The generated reversions to wild type not associated with cytological reinversions support this conclusion. After x-ray mutagenesis of wm4, w+ remained approximated to heterochromatin, although its activity was restored in 14 lines (Reuter et al., 1985). The possible cause of this was removal of the heterochromatin lying closest to w+ by irradiation. The question was raised whether not only pericentromeric heterochromatin itself, but also some of the regions adjoining it, have an inactivating capacity associated with position effect variegation. It is unclear if variegation is induced by breaks in the 20 region of the X chromosome. Many of the early data supporting this notion proved to be incorrect (Spofford, 1976). The generation of reinversions to normal with transpositions of the genes from heterochromatin in the 18E 19E, or 20 regions (Panshin, 1938; Kaufmann, 1942; Pokholkovaetal., 1993 a,b) are indirect proofs that the region lying close to heterochromatin, although not included in it, has no potential inactivation capacity.

2. Intercalary heterochromatin There is evidence indicating that intercalary heterochromatin (IH) can cause variegation. Lindsley and Grell(l968) provided data on position effect variegation in five chromosomal rearrangements between euchromatic regions. One of them is T(l ;2)w13G2.The possibility that the w+ gene may be inactivated when transferred to the IH of the 56EF region by the T(l ; Z ) W ' translocation ~~~ [see Spofford's (1976) review and also Gvozdev, 1981a,b] is subject to revision. It will be recalled that two elements with the sequence of the lA-3C3/41-56F/41-21 and 20-3C5/56F-60 regions result from this translocation, and, moreover, it is readily apparent that the w+ region (3C2-3) establishes contact with pericentromeric heterochromatin (the 41 region) (Lindsley and Zimm, 1992). Information about four other rearrangements is given in Table 32. Since reversions induced by irradiation of a chromosomal rearrangement causing position effect are incomplete, partial reversions are associated with the effect exerted by IH at new positions (Kaufmann, 1942). However, there is no direct evidence for this. It was reported that reinversions place the w+ gene from heterochromatin itself to mainly the IH (Reuter et al., 1985). It is not clear whether its blocks have an influence on inactivation of the w+ gene. A relation between the IH properties and position effect variegation was found at the cut locus. Three of five mutants had euchromatic-heterochromatic chromosomal rearrangements; however, there were no rearrangements in the two strains, and the frequency of ectopic pairing in the 7B region (where the cut locus is located) was considerably higher than usual (Hannah, 1949; Lindsley and Grell, 1968).This effect can be

392

1. F. Zhimuisv

Table 32. Cases of Position Effect Variegation Potentially Evoked by Intercalary Heterochromatin (IH)

Rearrangement of region

T( I ;

3

)

~

Results

~ Translocation ~ ~ ~ between ~ w+ region and 10CC3-4; variegation of w + gene (Lindsley and Zimm, 1992)

Characteristics of the IH region according to Zhimulev et al. (1982) 1OOC3-5 is a region of late replication

T(2,3)MV

Translocation between 43E and 75C; eye color variegation (Lindsley and Zimm, 1992)

75C is one of the most typical region of IH

Jn(2L)53d

Inversion between 25A and 29F (Lindsley and Zimm, 1992)

25A is typical region of IH

T(2;3)dpw'

Not mapped exactly (Lindsley and Grell, 1968), apparently variegated for dp

Insertion of AR4-24 transposon into 24CD region

More pigmented ventral part of eye than dorsal part, because of w+ inactivation (Hazelrigg et al., 1984; Rubin et al., 1985; Hazelrigg and Petersen, 1992); insertion is mapped in 24D1-2 (Hazelrigg and Petersen, 1992)

The 24D1.2 band is a site of

Insertion of ZQ transposon in the 84DE region

Variegation of w+ in transposon; inactivation is enhanced in XO males (K.Ahmad and K. K. Golic, 1995, personal communication)

84D1-2 is a region of typical IH

Insertion of hsp26pt-T; hsp70-white transposon into 2R pericentric region

Inactivation of w+ gene in transposon (Wallrath and Elgin, 1995; Wallrath et al., 1996) and about 30% underrepresentation of hsp26-pt-T DNA in salivary gland chromosomes strain 39C2)

The transposon is inserted in the 42B1-4 band (mapped by 1. E Zhimulev., 1995, unpublished results from the photograph in Wallrath et al., 1996); the band is most typical 1H region

Nine insertions of hsp26-pt-T; hsp70-white in pericentric, telomeric and fourth chromosome regions

Variegation of white gene, about 60-20% DNA underrepresentation in salivary gland chromosomes; PEV is suppressed by the Y chromosome and Su(uar) mutations (Wallrath and Elgin, 1995; Wallrath et al., 1996)

The fourth chromosome is

Mutation w ~ ~ ~ There ' ~ is~inversion ' between 3C and 4B or shows reproducible 4C (R. Levis, 1995, personal commosaic pattern of munication)

Moderately late replication in 4AC was found

7

late replication

enriched with middle repetitive DNA (Kholodilov et al., 1987, 1988) and binds antibodies against HP1 (James et al., 1989; Belyaeva et al., 1993).

(continues)


393

Table 32. (Continued) Rearrangement of region

Results

Characteristics of the IH region according to Zhimulev et nl. (1982)

eye color (Rubin et al., 1985) Thirty-one P[lacW] insertions (Sun et al., 1985)

Twelve distinct patterns of w+ variegation. Insertions are in the regions: 4C34 (2 cases)

5D34 7A6-8

17A1-6 40A 1-4 ( 9 cases) 26A (2 cases) 33F-34A

-

45A 48A (3 cases)

Weak IH region Very weak IH region Late replication region

55C1-5

-

67D8-12

Typical IH region

67E1-4 69C (3 cases)

8481-6 85C1-3 86C1-4 86E6-8 Insertions of AR4-24 transposon into 7 polytene chromosome sites gives rise to mosaic W + expression (Balasov and Zhimulev, 1997)

Near 4CD, typical IH region Late replicating region Typical 1H region Typical 1H region Typical IH region

102A: “salt and pepper” variegation

4OC: “salt and pepper” variegation 41F1-2: “salt and pepper” variegation 41AB: “salt and pepper” variegation 39A: “salt and pepper” variegation; influence of temperature and Y chromosome was found 25A: “salt and pepper” variegation; influence of temperature and Y chromosome was found 31F: sectorial variegation

-

Late replication region Late replication region Region is close to centromeric heterochromatin, but not in it The same The same The same The same

In 1H

Very closely to IH

explained as a consequence of the strong manifestation of the properties of the IH, as well as of insertion of pericentromeric heterochromatin into the 7B region. The euchromatic regions of the hairy gene can exert a suppressive effect (Jeffery, 1979). In D. hydei, the forked and Zebra mutations were associated with chromosomal rearrangements within euchromatin. The phenotypic expression of

394

1. F. Zhimulev

variegating forked is suppressed at lower temperatures and enhanced at higher temperatures. However, an extra Y chromosome shifts the expression of the mutant allele to normal. In Zebra flies reared at low temperature, pigmented spots were larger and the number of variegated spots increased when an extra Y chromosome was added (van Breugel, 1988). From consideration of all these data, it is evident that position effect modifiers have an influence on the expression of the putative “euchromatic” position effect variegation that is different from the “heterochromatic.” Neither Jeffery nor van Breugel dismissed the possibility that variegation may be associated with transposition of small heterochromatic fragments in inversion breakpoints. P-element inserts gave a new opportunity for analysis of position effect variegation evoked by IH. In the AR4-24 transposon inserted into a late-replicating region, the w gene expression variegates (see Table 32). Similar variegation was found for the reporter genes inserted into such typical IH sites as 42B1-4, 84D1-2, or in 25A (see Table 32). More difficult to explain are cases when transposons insert in euchromatin but into neither IH nor a-and P-heterochromatin, for example, insertion of 118E-25 into basement of the X chromosome, or 118E-3 into 102AB in the fourth chromosome (Wallrath et al., 1996),or insertions into 102A, 40C, 41F1-2, 41AB, 39A, and 31F (see Table 32). Although no clear cases of the inactivating action of the IH are known, it is premature to deny this possibility. Inasmuch as the IH regions are incomparably smaller than blocks of centromeric heterochromatin, it may be assumed that their potential inactivation capacity may be weaker. For this reason, specific conditions are necessary to detect them, for example, generation of chromosomal rearrangements on the background of the action of powerful enhancers of position effect. However, no such experiments have been performed so far.

3. Telomeric heterochromatin Rather numerous cases are known in which telomeric DNA exerts an inactivating effect: the w+ gene becomes inactivated when the functional P(wvaT)transposon is inserted into the telomeric region of chromosome 2L (Gehring et al., 1984) and when the P((w,ry)A)4-4 transposon is inserted into the end of chromosome 2R. In the latter case, flies grown at 25”C,whose only source of the w+ gene product was the transposon gene, had uniformly nonpigmented eyes. This was evidence that gene inactivation was due to proximity to telomeric DNA. In flies grown at 18”C,reddish spots appeared on the light background of the eye. This was clear-cut evidence for weakened variegated eye pigmentation at the w+ gene (Hazelrigg et al., 1984). When transposon A 4 4 was moved from the telomeric region and inserted into euchromatic positions, the wild-type expression of the w+ gene was restored. When inserted in the telomeric tip of the X chromosome in transposon 4-1 6, the A& gene showed very low-level expression only in

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation ~

~~

-~

395

~________

adults and no activity in larval organs (Kirkpatrick and Martin, 1992). A variegating w+ transgene was inserted about 16 kb away from the 3R terminus, within a series of tandem 1-kb repeats (Levis, 1989; Levis et al., 1993). After mobilization of P-element located in the Dp(1 ;f)1 187 minichromosome, several terminal deletions were obtained. Removal of a minichromosome part between the y gene and the telomere results in joining of the telomere and this gene. Dramatic increases in y+ variegation were associated with these terminal deletions in comparison with the original Dp(l;fl1187 (Spradling, 1993; Zhang and Spradling, 1993; Karpen, 1994). Twenty-six transposons showing variegated expression of the ry+ gene were inserted into a 5-kb region located about 40 kb from the Dp 1187 minichromosome telomere (Karpen et al., 1988). The rosy+ genes within transpositions inserted near telomeric sequences were inhibited by position effect variegation (Karpen and Spradling, 1992;Tower et al., 1993). Four telomeric insertions of the hsp26-pt-T;hsp70white transposon showed position effect variegation of the hsp70white gene and about 10% to 50% hsp26pt-T underrepresentation in polytene chromosomes (Wallrath and Elgin, 1995; Wallrath et al., 1996). In another study two independent insertions of the AR4-24 transposon were found in the 2R and one in 4th chromosome telomeric regions. In at least one case, expression of inactivation was modified by temperature and variation of Y chromosomes (Balasov and Zhimulev, 1997). Variegating insertion of P[lacW] into 1AB was found by Sun et al. (1995). Modifiers of PEV (Y chromosome dosage or genetic modifiers) usually do not affect telomeric position effect variegation (Talbert et at., 1994; R. Levis in Weiler and Wakimoto, 1995; Wallrath and Elgin, 1995). Among the exceptions are cases of suppression of variegation of the ry+ transgenes inserted in Dp 1 187 by the Y chromosome (Karpen et al., 1988), suppression of hsg-w' transgenes by Su(var)mutations (Wallrath and Elgin, 1995), and suppression of w+ transgenes by temperature and the Y chromosome (Balasov and Zhimulev, 1997). In some cases, regulatory properties of P-elements at 1A can be inhibited by some of the alleles of the Su(var)205 (Ronsserayet al., 1996).More details can be found in the review by Weiler and Wakimoto (1995). A n inactivation effect of telomeric DNA was found in Saccharornyces cerevisiae (Gottschling et at., 1990; Sandell and Zakian, 1992; Aparicio and Gottschling, 1994; Tartof, 1994; Shore, 1995). The silent mating loci in Saccharomyces cerevisiae adjacent to telomeres show features similar to heterochromatin. Inactivation of transcription in these regions depends on silent information regulators SIR3 and SIR4. These SIRSinteract with specific silencing domains of the H3 and H4 histones (Hecht et al., 1995).

C. Heterochromatizationaccording to Prokofyeva-Belgovskaya Studies of the sc8 strain where an inversion transposes the heterochromatic 20BC region of the X chromosome to the euchromatic 1ABl region of the chromosome

396

1. F. Zhimulev

b

a

C

"

Figure 144. Types of morphology of the 1ABl region and the closely located 2OBC in the inverted In(l)sBchromosome in three cells of the salivary gland. (a) "Euchromatic" state. Homologous chromatidsconjugate forming bands well expressed in lABl and dotted in the 20BC region. (b) Partial "heterochromatization" of both regions. Instead of some bands, their composing chromomeres are seen. (c) Complete "heterochromatization". After Prokofyeva-Belgovskaya ( 1945, 1986).

4

i

e Figure 145. Changes in the morphology of the 3C region (a-fl when juxtaposed into heterochromatin in the w* translocation.After Prokofyeva-Belgovskaya(1939b).


397

encompassing the y+ and SC+ genes revealed that the distinctive pattern of banding varies considerably depending on conditions in the cell (Figure 144). Euchromatin was the name given to “morphotypes” showing normal stainability, both distinct and integral (Figure 144a). In some cases, the pairing properties of chromatids composing the polytene chromosome are affected and, as a consequence, the bands become looser and lose stainability and the whole chromosome region (1ABl and 20BC) is converted into a diffuse network resembling p-heterochromatin; that is, the region acquires a “chromocentral structure” (Figure 144b and 144c) (Prokofyeva-Belgovskaya, 1937a, 1939a,b, 1941, 1945, 1947, 1965, 1986; Noujdin, 1946~). The same was observed for the euchromatic 3C region [the w+ locus in the T(1;4)wd rearrangement (Figure 145)]. This loosening is much less extensive in the In(l)wm4 inversion (Prokofyeva-Belgovskaya, 1939b, 1945, 1986). It is believed that the degree of band development in the region transferred to heterochromatin in the sc8 inversion can be different in two homologs (“heterocyclicity”) and differently manifested in the absence of their pairing (ProkofyevaBelgovskaya, 1947). “Heterochromatization” was the name she gave to the acquisition of a morphology outwardly resembling centromeric p-heterochromatin by a euchromatic region. The term has become common usage in the literature, and the various structural derangements due to position effect variegation were subsequently covered by this term. The heterochromatization Prokofyeva-Belgovskayadescribed has correlations, albeit not consistently direct, with genetic inactivation due to position effect:

1. In some cases, heterochromatization can extend for more than 28-30 bands (to 6 pm) from the breakpoint (i.e., it can be regarded as a spreading effect) (Belgovsky, 1944, Prokofyeva-Belgovskaya, 1986). It should be noted that this conclusion has not been documented by figures or photographs. 2. The state of heterochromatization can be modified, with the resulting modification not always being the same as when these very modifiers act on genetic inactivation. Prokofyeva-Belgovskaya takes the view that, because of the presence of the Y chromosome, the lABl region takes on an abnormal morphological appearance in 84% of cells in males but in only 34.5% of cells in females. An extra Y chromosome in scs/y ac o, f Y’ females somewhat decreases the percentage of heterochromatinized lAB1-2OABC regions compared to sc8/y ac v larvae (13% versus 20%). 3. The effect of temperature was unusual: heterochromatization was observed in 71% of larvae teared at 25°C. In larvae cultured at 14 or 30°C, with the exception of the first 6 hr of embryonic development (2S°C), the percentages were 33% and 37%, respectively. 4. Heterochromatization degree in sc8/y ac o, heterozygotes is greatly dependent

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on the paternal or maternal origin of the sc’ chromosome. The percentage of nuclei with a chromosome region in a “heterochromatic” state is 18.3% when the origin is paternal and 71% when it is maternal (Prokofyeva-Belgovskaya, 1945, 1947,1986). The assumption is then made that the “parental effect” is associated with an event preceding the formation of sperm nuclei. This event is heteropycnosis of the X chromosome at all the stages of spermiogenesis. The X chromosome received from the father passes as a compact body all the meiotic stages, possibly because it “remembers” its past and also because heteropycnosis tends to persist in offspring (Prokofyeva-Belgovskaya, 1965).

A seemingly opposite case is, however, essentially similar to heterochromatization: “euchromatization” of heterochromatin transposed to euchromatic surroundings. In rst3, s 8 , wmI4, and wd strains, as the result of rearrangement, a part of the inert material usually lying in heterochromatin was found to be inserted between the active regions. The inert region then passes almost entirely to the euchromatic state and became indistinguishable from the adjacent active regions (Prokofyeva-Belgovskaya, 1941, 1945). However, the author provides no documentary evidence for this conclusion. In another series of studies concerned with heterochromatization, euchromatic regions were shown to lose stainability, and accordingly banding pattern, when transposed to heterochromatin (Schultz and Caspersson, 1939; Cole and Sutton, 1941); in contrast, stainability of the bands then increases (Schultz, 1941a). Schultz (1941b, 1956) also notes that, at maximally expressed variegation at the w+ gene in one of the chromosomal rearrangements [v-D3, subsequently called T(f ; 4 ) ~ ” ~ ~ * -with ~ ’ ] a breakpoint in the 3E5-6 region in the salivary gland chromosomes, the w+ gene (the 3C2 band) immediately borders the chromocenter. This may taken to mean that the bands between the w+ locus and the 3F1 band are completely heterochromatic in some cases, and they are indistinguishable as individual bands in others or, presumably, simply lost. The author has observed the same effect in several translocations. Other authors have also observed the effect, namely, a chromatin-associated band other than the one with a rearrangement break, and more distally located in the T( 1 ;4)wm258-2 translocation (Hartmann-Goldstein, 1967; Wargent and Hartmann-Goldstein, 1976; Reuter et al., 1982b). Other structural modifications of the chromosomes, also called heterochromatization, were detected. A zone was consistently delineated from the borderline between euchromatin and heterochromatin, where heterochromatin stretched distally along the X chromosome and bands were hard to identify. This might have been due to thinning of the chromosome, stretching, loss or deeper staining, or loss of bands. The part of the X chromosome displaced to T(I ; 4 ) ~ frequently ~ ~ ~ appeared ~ - ~shorter ~ than the homologous section of the nontranslocated homolog (Hartmann-Goldstein, 1967; Hartmann-Goldstein


399

and Wargent, 1975; Wargent and Hartmann-Goldstein, 1976; Reuter et al., 1982b; Koliantz and Hartmann-Goldstein, 1984). The double-inversion heterozygote with In( 1)mK,the breakpoints of which are located in euchromatin at 10E4-5 and in heterochromatin proximal to 20B1-2, and with In(2LR)ReuB (breakpoints at 52D5 and 40F, respectively) showed variation in heterochromatization at 14°C. By definition, all states intermediate with respect to “typical euchromatization and frank heterochromatin” were regarded as heterochromatic. The regions adjacent to the breakpoint varied in appearance from darkly staining to a loose granular meshwork showing indistinct banding (Hartmann-Goldstein and Wargent, 1975). These morphotypes also resemble phenotypic variegation in exhibited features. It was shown that the heterochromatized region in the chromosome with the T ( 1 ; 4 ) ~rearrangement ~ ~ ~ ~ - ~lengthens, ~ and it can cover a chromosome region from the breakpoint in 3E5-3F1 to the bands in the 2B region at low temperature ( 14°C). The frequency of cells showing such heterochromatization was also maximal (63%) at 14°C. The first hours of embryonic development are critical to the formation of these morphotypes. Individuals exposed to low temperature after development at 25°C within the first 3 hr of embryonic stage did not show an increase in the amount of heterochromatization (Hartmann-Goldstein,

1967). Ananiev and Gvozdev (1974) described two morphotypes arising in the Dp( 1 ;OR duplication during heterochromatization. They compared two states contrasting in variegated expression:X/Y/Dp (25°C)and X/O/Dp (18°C).The first group of larvae varied in morphology of the 1A-2B region at a frequency not exceeding 1%; the findings for the second group included ( 1) a strong modification of duplication morphology in 40% of the nuclei, making it impossible to identify individual bands; (2) missing bands (e.g., 2E and 2DE) in a number of cases; and ( 3 )unusually thick bands in about half (in the IDE and 2CD regions) or a “modified set of puffs” (not documented). There were no differences in the morphology of the homologous regions between the duplication and the X chromosome in only 30% of nuclei. The total number of silver grains appearing over the duplication area, when it incorporated [3H]uridine, decreased during heterochromatization, and DNA replication in the bands of the 1DEF and 2AB regions was occasionally delayed (Ananiev and Gvozdev, 1974). Thus there is ample evidence indicating that variegating morphology is of diverse kind in the rearranged chromosome. The expression of heterochromatization and genetic inactivation is modified in a correlated manner by the same factors. Nevertheless, it remains unclear whether or not they are related. Does heterochromatization lead to genetic inactivation? After consideration of the problem, Baker (1968) concluded that the chromosomes are unsuitable for analysis in the tissue showing variegation. Indeed, it cannot be observed whether heterochromatization of the rearranged chromosome bearing the wild-type allele occurs in the mutant part of the variegated eyes of adult flies. Likewise, the variants

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of “heterochromatization” in which gene inactivation occurs are unknown. Thus Baker (1968) took the view that “Explanations of the reason a gene is not producing its product (or at least, a normal one) in a given region are still put in terms of ‘heterochromatization’, or ‘compaction’, terms that, in reality, expose our ignorance rather than our understanding” (p. 134). It is Baker’s view that it would be advantageous to study change in chromosome morphology of all types whose genetic inactivation can be detected by relying on the presence (or absence) of puffs or bands in the salivary gland chromosomes. If such a variegation were detected, and if the criteria of variegations of position effect were fulfilled, comparisons of the course of events for biochemical products would become feasible under normal conditions and position effect (Baker, 1968).

D. Chromatin compaction Starting in the 1930s and 1940s, changes in the morphology of chromosome regions transposed to heterochromatin and containing inactivated genes were described. In addition to the previous information about “heterochromatization,” a body of knowledge of another sort was accumulating. Variegation was found to correlate with loss of bands nearest to the heterochromatic junction and also with darkening of the remaining bands, brought closest to heterochromatin after “deletion” of the most proximal bands. When an extra Y chromosome is added to the genome, variegation reduces. This is manifested as shortening of the cytologically visible “deletion,” and chromosomes with the deletion are less frequently encountered (Morgan et al., 1938; Schultz, 1941b, 1943a, 1947). As Schultz (1956) observed, with enhancing variegation, the chromosome regions transposed to heterochromatin keep becoming more similar to heterochromatin itself, and, in extreme manifestations, the bands as such are not identified. In a study of a series of variegated chromosomal rearrangements, Sutton ( 1940a, 1941) found change in the structure of the chromosomes manifesting as an increase in staining intensity of the nearest-to-heterochromatin band only in the 2B region of the In(1)313.25.36 inversion between the 2B and the 20 regions of the X chromosome. This deeper staining was variegated in 10 of the 28 examined nuclei. The bands of the 3C region in the respective rearrangements showed no superstaining. Extensive citations from Schultz (1965) shed light on the notions of the mechanisms of genetic inactivation: “We are now in a position to make a straightforward interpretation, based on the concepts already developed. We now know that heterochromatic regions are late in their replication, and that transcription to RNA does not occur in compacted regions. . . . Thus, in the rearrangements, we see evidence that the delayed replication of the heterochromatic regions influences its neighbors. In extreme cases, these fall completely into the cycle of the heterochromatic regions; in the less extreme cases, they would be able to undergo some replication. . . . The cytological “heterochromatization” by which this delay in replication is accomplished would thus be a compaction of the chromo-


401

some template. The template remaining compact should not function in transcription” (p. 141). Thus Schultz came close to an explanation of genetic inactivation due to position effect through compaction of euchromatic material transposed to the vicinity of heterochromatin. According to Sutton-Gersh‘sconcepts (Gersh, 1973), the DNA in the chromocentral heterochromatic regions of the chromosome may be kept in an extremely condensed state by cross-linkingproteins and, when a free fragment of the chromosome is brought to such a region, the cross-linker effect spreads over the fragment, thereby compacting the most proximal bands. What facts support the concept of compaction? The earliest studies on position effect (Emmens, 1937a) demonstrated that, in the In(l)rst3 rearrangement causing position effect, as a consequence of transposition of the 1A to 3C4-5 region to heterochromatin, distinct bands become recognizable in some cases only in the distal part of the 3B region and beyond the limits of the region; that is, a block of compacted chromatin, presumably including bands of the proximal part of the 3B and 3C1-3 regions, lies closer to heterochromatin. The author did not explain why banding pattern changes in this region. A case of compaction was undoubtedly being dealt with. It was shown that the 3D puff lying at the boundary of heterochromatin is inactivated, and that its material is compacted, in the T(1;4)w258-21translocation (Schultz, 1965; Rudkin, 196513). It should be noted that the puff is missing in T/X individuals at 18°C at all deveopmental stages, and that its activity does not differ from normal in T/XY individuals (25OC) (M. Kemrer, in Schultz, 1965).Another study of this translocation demonstrated that there were no deviations from the norm in the region of the 3C11-12 puff in 46 of the 87 examined nuclei. The whole 3C1-3E5 region stained more heavily and was as compacted as heterochromatin in one of the nuclei. The 3C11-12 puff was missing from these nuclei (HartmannGoldstein, 1966; see also Kornher and Kauffman, 1986). Inactivation of the 87C heat shock puff in translocation on the Y chromosome was detected by Henikoff (1979a, 1981, 1990). It is noteworthy that the heat shock puffs are not inactivated in all the rearrangements in the Y chromosome (Ellgaard and Brosseau, 1969). “Intensified staining” and loss of bands nearest to heterochromatin (Morgan et al., 1938) (see earlier in this section) can be made compatible by imagining that material of the bands becomes compacted by progressively involving the distant band and formation of a single block of compacted material. In such an event, the bands closest to heterochromatin would disappear first, and another band would form instead; it would be thicker because it incorporates the material of the thinner band, and it would stain deeper because its compaction would be associated with tighter packaging. The chromosome can concomitantly shorten. Accepting this, it may be stated that compaction was described by almost all researchers who have studied position effect at the cytological level. Thus, in the 2B region of the In(1)313.25.36 inversion (Sutton, 1940a,b, as seen in plate 2K), there is no darkening; rather, there is formation of a block of compacted mate-

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1. F. Zhimulev

rial. The photographs in the earlier publications demonstrate the presence of blocks in 3C-E (Hartmann-Goldstein, 1967, plate 1C) and in 3AE (Wargent and Hartmann-Goldstein, 1976, Figure 3) in the T(l ;4)w25s2’ translocation. The blocks are presumably formed from the proximal regions in the translocated elements in the T(Y;3;4)1 9 0 c ~and ~ ~T(Y;3;4) j 19Ocu7Of translocations in D. gseudoobscuru (Gersh, 1973). Changes in the chromosomes corresponding to compaction of the proximal parts of the translocated elements can be found in illustrations accompanying several other papers (Ananiev and Gvozdev, 1974, Figure 3; Reuter et al., 198213, Figure 1).Reuter et al. (1982b) note that compaction is quite rare. In some instances, darkening of bands in which the inactivated w+ gene is located is seen in photographs of the wml rearrangement in D. hydei (van Breugel, 1970, Figure 3b). Baker’s (1968) question as to the extent to which “heterochromatization” and “compaction”are related to variegation expression remained unanswered until a gene system allowing us to follow genetic inactivation and alteration in polytene cytology in the same cell became available. In this respect, the system of ecdysterone-stimulated puffs of the salivary gland chromosomes proved to be unique. It is well established that a pattern of changes in the activities of about 120 large puffs controlled by the ecdysterone occur within several hours before puparium formation in the salivary gland cells. Puffs active before exposure to the hormone become inactive (the intermolt puffs), and the inactive loci are concomitantly activated, with some loci (the early ecdysterone) becoming active some minutes later on induction by the hormone and others (the late ecdysterone) becoming active after a delay of several hours. The late ecdysterone puffs eventually reach highest development by the time of puparium formation (Ashburner, 1972a). It was found that 2B3-5, one of the earliest ecdysterone puffs, is the key in the hormone-induced puffing cycle: mutations or deletions at the ecs (BR-C) locus in this puff cause complete loss of cell ability to respond to the hormone and form puffs (E. S. Belyaeva et ul., 1981, 1989). Therefore, absence of the ecdysterone puff is a readily detectable consequence of genetic inactivation of the BRC+ locus in the rearranged chromosome in BR-C+/BR-C heterozygotes. For this reason, puffs associated with genetic inactivation can be identified by following cytological changes in the region where the R(BR-C+) chromosome joins with heterochromatin in cells without puffs. In the T( I ;2)dorvar7strain, which involves position effect at the ecs+ 10cus, several types of deviations from normal morphology can be found at the heterochromatic junction: breaks can occur in the puffed region, and the translocated element can be deeply embedded in the chromocenter so that its most proximal bands are imperceptible for unclear reasons-either because of a superimposed chromocenter or underreplication of these regions. In the 2B region, there is a wedgelike constriction instead of a puff in some cells, and a block of compact stainable material replacing characteristic distinct bands in the other cells (Figure 146). The blocks were considerably more compacted in females than males (Zhimulev et al., 1986, 1988) (see Figure 146).


403

Figure 146. Electron microscopic sections of the 28 region (marked by a square bracket) in the translocated elements in T(1;2)dcirWr7/FM6females (a-d) and dmuar7/0 males ( e and fl. Scale is 1 Fm. Reprinted by permission from Zhimulev et d.(1988).

In heterozygous R(BR-C+)/BR-C females, the ecdysterone puffs did not develop only in those cells in R(BR-C+) chromosome whose 283-5 puff material became compacted and converted into a block. Thus, of all the morphological alterations (“heterochromatization”) caused by position effect, only compaction is associated with genetic inactivation. There are exceptions as in all generalities. In some cells the puffing pattern conforms to normal despite the present blocks. Patterns are intermediate in some cells; both the early and late puffs occur in cells with blocks. The frequency of such cells is 1% (Zhimulev et al., 1986).

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I. F. Zhimulev ~~

Compaction of the 2B region in the looser blocks in T(I ;2)d0PQT7males generally does not prevent the formation of puffs both in X/Y and X/O males. Puffs develop normally in most nuclei in viwo and when the glands are incubated with the hormone. Puffs do not develop at all or develop with some delay in a small number of cells (Zhimulev et al., 1988). During the formation of a block of compacted chromatin in females, transcription of the earlier active material ceases: the blocks do not incorporate [3H]uridine,and hybridization intensity of the clone of DNA from the ecs+ locus (Dmp20.5) with RNA on chromosome preparations falls sharply (no more than 6% puff labeling remains). There were sex differences corresponding to differences in morphology and inactivation ability of the blocks: [3H]uridine incorporation into the blocks of doTvaT7 males was not inhibited (Zhimulev et al., 1988; Vlassova et al., 1 9 9 1 ~ )So, . here we find a very good correlation between degree of compaction of a euchromatic region evoked by position effect variegation and genetic activity. Good correlation was found between cytological compaction and genetic inactivation of the bw+ gene when chromosome rearrangements giving rise to the cis-effect of the bw+ gene were compared (Belyaeva et al., 1997).Therefore, reasons for such conclusions as “compaction is neither sufficient for, nor strictly correlated with inactivation of the ecs gene” and “compaction, gene inactivation . . . may be independent consequences of chromosome rearrangements” (Weiler and Wakimoto, 1995, p. 588) are unknown. The are plausible reasons for the differences in the inactivating strength of compaction between males and females in the doTvar7 translocation. First, it is unknown to what extent blocks of the types differ in chromatin inactivation. Since judgments about inactivation are made on the basis of puff development, the threshold for the induction of the ecdysterone puffs must be determined. Puff induction is not affected in BR-C+/Dffemaleswhen the BR-C+product is in one dose, and, hence, 50% of the product suffices. Therefore, in cells of doTvar7BR-C+/ doruar7+BR-C1t435females, with only one dose of the BR-C+gene product, a belowthreshold level should be reached more easily than in males. Thus dosage compensation may decreases position effect inactivation of the BR-C+gene. Second, dosage compensation possibly affects the degree to which the whole chromosome is compacted. The two processes, dosage compensation with loosening of the entire chromosome and a compaction of a particular region caused by variegation resulting in the formation of looser blocks than in females, may superimpose in the regulation of the activity of the male X chromosome. Finally, it is unclear whether inactivation (compaction) of the material of the 2 8 puff is always less effective in males than females. The latter explanation seems to hold true for viable dorvQT7/Y and dorunr7/0males at 25°C because compaction is accessible to analysis in most of their salivary gland cells. However, viability of X/O males is much reduced, even at 25°C and particularly at 1 8 T , so that only 2 6 4 2 % of larvae manage to survive by the end of the third instar. Variegation-


405

associated inactivation may be expressed more in dying larvae. For this reason, individuals with stronger expressed position effect, and hence compaction, most likely do not survive to the time when analysis is feasible; accordingly, individuals weakly expressing variegation survive and are accessible to analysis (Zhimulev et al., 1988). In Dp(f ;I)pn2b-, Dp(f ;f)f337-, and Dp(1;f)R-bearing strains, there are no differences in morphology and ability to inactivate BR-C’ of compacted material between XO males and females (Zhimulev et ul., 1989a,b; Belyaeva and Zhimulev, 1991a). This is possibly because they do not represent the entire X chromosome, as they do in dOruar7,but rather its small fragment in the presence of the whole chromosome. The formation of blocks of compacted material at the euchromatinheterochromatin junction is a feature common to numerous rearrangements studied in this respect (Belyaeva and Zhimulev, 1991a, Belyaeva et al., 1993, 1997a,b; Pokholkova et al., 1993 a,b; Belousova and Pokholkova, 1997a,b; Belyaeva and Zhimulev, 1997a,b; Mal’ceva et al., 1997a,b). The extension of the heterochromatin is specific to each rearrangement; that is, it presumably depends on the properties of DNA at the junction between euchromatin and heterochromatin. The distance from the breakpoint to the most distal band that can be compacted varies from 10 to 170 hands on Bridges’ cytological map (see Table 28, in Section XII). Compaction as well as genetic inactivation tends to spread from the heterochromatic junction in the distal direction. For this reason, different segments of euchromatin are included in compact blocks in different cells, and they can be arranged in a series in which a chromosome region showing a well-defined banding pattern becomes shorter as the block increases in size, incorporating increasingly longer chromosome regions (Figure 147).

Figure 147. Formation of a block of compact chromatin in a part of the Dp(1;I)pnZb duplication proximal to heterochromatin ( a ) and at different steps of compaction (b-f). Scale is 5 pm. After Zhimulev et al. (1989a); reprinted by permission from Zhimulev et al. (1989a).

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I. F. Zhimulev

Varying extents of cytological compaction include tissue specificity. Earlier it was found that, in pseudonurse cell (PNC) polytene chromosomes of the otu mutant, heterochromatic characteristics such as underrepresentation of pericentric and intercalary heterochomatic DNA and formation of ectopic fibers are not manifested as strongly as in somatic cells [larval salivary gland (SG) cells] (Mal'ceva and Zhimulev, l993,1997a,b; Mal'ceva et al., 1995, 1997a,b;Koryakov et al., 1996). A comparative study of the manifestation of position effect variegation for the polytene chromosomes of salivary gland cell and pseudonurse cell nuclei was made using Dp( I ;1 )pn2B and Dp(I ;f) 1337 rearrangements. Both the spreading and frequency of compaction were significantly lower in the polytene chromosomes of the PNC. Thus the percentage frequencies of block formation in SG and PNC nuclei for the Dp(1;l )pn2B rearrangement were 92.6% and 15.8%, respectively. For Dp(l;f)1337these values were 56.8% and 9.7% (Mal'ceva et al., 1997; Mal'ceva and Zhimulev, 1997). Cytogenetic analysis of dominant position effect variegation of the bw gene in salivary gland chromosomes carrying rearrangements In(2R) bwVDe', and In(2R)bwVKshown that, in rearranged homologs, typical comIn(2R)bwVDe2, paction of the 59E1-2 to 59D region and proximal neighboring bands took place. However, the compaction was never observed in normal homologs (Belyaeva e t al., 1997; Belyaeva and Zhimulev, 1997). In another case of unusual position effect, in ci gene variegation, which is a kind of transvection phenomena, no cytologically visible compaction of chromosomes was found on either rearranged or normal homologs (Demakova et al., 1997). Compaction of the 3CE region in In( 1) wm4 is susceptible to the effect of genetic modifiers. The normal morphology of this region is to a large measure restored when Su(uar) is introduced into the genome (Hayashi e t al., 1990). Discontinuous compaction (Belyaeva and Zhimulev, 1991a; Belyaeva et al., 1993) was revealed in addition to the continuous type: not one, but several, compaction zones separated by well-identifiable stretches with normal morphology and puff-forming ability are detected in the part of the chromosome translocated to heterochromatin. As a result, the regions located further away from the breakpoint can more frequently occur in a compacted state than the ones closer to heterochromatin (Figures 148-151). A group of regions with the highest compaction frequency in all the studied regions are then detected. These are lD, 2B1-12, and 2CD in the distal part of the X chromosome. The lowest frequencies are for the 2B13-18,2C1-2,2E, and 2F regions (Figure 152). Examination of Figure 153 shows that there are particular groups of regions involved together in compaction. When compaction is discontinuous, the genes in the block are inactivated precisely in the same way as when compaction is continuous. For example,

Polytene Chromosomes, Heterochromatln, and Position Effect Varlegation

407

Figure 148. Compaction of material of the duplication of the X chromosome (designated by brackets) in X/O/Dp(l;f)1337 males. (a) Normal chromosome. (b-e) Continuous compaction. (f-h) Discontinuous compaction. In brackets are the 2B1-7 (a and b), 2AB (c), and 1C-2B7 (d-h) regions. Reprinted by permission from Belyaeva and Zhimulev (1991a).

when the 2B1-12 region is in the state of a block, the BR-C+ locus is inactivated, with the result that the cell does not respond to ecdysterone. In such a case, the 2C1-2 and 2E bands lying closer to the breakpoint can retain normal morphology and remain decompacted (Belyaeva and Zhimulev, 1991a). Discontinuous compaction generally occurs more rarely than does the continuous kind: in 10% of the chromosomes showing blocks of compacted chromatin in X/O/Dp(l;f) 1337, in 25% in X/O/Dp(f ;f )j~ndb,and in 100% of nuclei with the T(f ;4)wm258-21 and Dp(l ;f)R rearrangements. The degree to which the blocks are expressed-that is, the frequencies of chromosomes with blocks, as well as extension of a chromosome region compacted in the block-is affected by modifiers of variegation (Figure 154), such as temperature and presence of heterochromatin in the cell. The lower the temperature is and the smaller the heterochromatin amounts are, the more frequently blocks occur and the larger is the number of bands they include. There are grounds for believing that the block frequencies are related to the total gene pool of a strain. Since compaction depends, to a large measure, on temperature, there

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1. F. Zhimulev

Figure 149. Discontinuous compaction of the duplicated material (appears as a ring) of the X chromosome (designated by brackets), ectopic contacts, and breaks in X/O/Dp(l;f)R males. (a) Normal chromosome. ( b e ) Variants of compaction designated by brackets: 1D1-2 (a); l C , lD, 1EE 2A-B12, and 2CD (b); 1B2B12, 2CD, and 2E1-2 (c); 1G2B12 and 2C (d); 1C-2B12 and 2C4-3A (e).Big arrow, ectopic stretches; two small arrows, breaks; asterisk, 2C1-2 p u e circle, 2EF puff. Reprinted by permission from Belyaeva and Zhimulev ( 1991a).

is the possibility of determining the temperature-sensitive period critical to the formation of blocks. In studies of d0rvar7and Dp( 1 ;f) 1337, it was found that exposure to 18°C during the first 6 hr of embryo life (Figure 155) also affects the frequency of block formation, as does exposure to 18°C throughout development (Zhimulev et al., 1988, 1989a,b). It will be recalled that the first 3 hr of embryo life are critical to “heterochromatization,” too (Hartmann-Goldstein, 1967). Parental effects on the compaction degree of material in doYar7were described in 13 larvae with the maternally derived translocation; material of the


409

Figure 150. Discontinuous compaction of material (designated by breaks), ectopic contacts,

and breaks in T( 1; 4 ) ~ " ' ~ ~ Df(ZR)MSZ'O/+ ~~'/+; females. (a) General appearCompaction in 2B1-12 (b, d, and e); 2E1-2 (c); ance of the translocation. (bf) and ICD, IE, and 2B1-12 (0.Designation of ectopic contacts, breaks, and puffs are the same as in Figure 149. Reprinted by permission from Belyaeva and Zhimulev (1991a).

2B1-2 to 2B7-8 band was included in about 28% of the nuclei in the blocks, and compaction involved the 2A1-2 to 287-8 region in the other 3% of the nuclei. In 19 larvae with the paternally derived rearrangement, about 58% of the nuclei have compacted blocks of heterochromatin; one-fourth include the 2A1-2, and even the 1El-4 regions (Demakova and Belyaeva, 1988).Parental effects on compaction were not detected in the other rearrangements (E. S. Belyaeva, 1990, personal communication).

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Figure 151. Discontinuous compaction of the duplicated material of the X chromosome (designated by brackets), ectopic stretches, and breaks in WO/Dp(l;I )pn2b males. (a) Normal duplication pairs with the normal X chromosome;2E1-2 junction point with heterochromatin (het). (b and c) Proximal regions of duplication are represented by a series ofcompact unidentified bands (designated by brackets); 2B1-12 and 2CE (d); 1E-2B12 and 2CE (e); 1C-F, 2B1-12, and 2E (0;l C 2 B l 2 and 2E (g-i). Designationsof ectopic contacts, breaks, and puffs are the same as in Figure 149. Reprinted by permission from Belyaeva and Zhimulev (1991a).

The properties of the involved chromosome regions change during the formation of the blocks of heterochromatin. For example, late-replicating sites, ectopic contacts with the other regions of the genome, and breaks do not normally occur in the 2B region (see Section VII); however, they express themselves as the result of compaction (see Figures 149-151). Earlier studies on some of the chromosomal rearrangements causing position effect revealed that several chromosome regions became later replicating when transferred to centromeric heterochromatin (Ananiev and Gvozdev, 1974; Wargent and Hartmann-Goldstein, 1976). In the case Wargent and HartmannGoldstein described, having become late replicating, the heterochromatized zone covers the 3C1-7 region, which contains a fragment normally late replicating and not causing variegation; it is therefore.difficult to make judgments about late replication in the ‘‘heterochromatized” region. In Ananiev and Gvozdev’s (1974)

np1I;llpnZb

Dpll ; f l R n=207

Figure 152. Compaction frequencies of the chromosome sites in one of two chromosome regions in three duplications. (a) X/O/@(l;I)pn2b(25°C). (b) X/O/Dp(l;f)R (18°C). (c) T(I ; 4 ) ~ " ' ~ ~ ~ -Df(2R)SM2l0/+ ~'/+; (14°C).Abscissa, the chromosome regions; ordinate, occurrence frequencies of the region in the compacted state. Reprinted by permission from Belyaeva and Zhimulev (1991a).

412

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Figure 153. Location of compaction zone in WO/Dp(J ;OR. Height of rectangle reflects number of events ac. cording to the scale given at the bottom right. Length of the rectangle correspondsto the length of the compacted duplication region (abscissa, designation of the regions). Reprinted by permission from Belyaeva and Zhimulev (1991a).

study, the 1DE region, quite distant from contact with heterochromatin, became late replicating. Hence a meaningful interpretation of the data needs analysis of chromosome morphology. Studies on the d0Yur7translocation demonstrated that the same 2B1-7 region differs, in principle, in the time of replication completion when residing in the puff versus the block state. It is known that in both males and females the 2B region completes replicating quite early, together with the completion of continuous labeling, when virtually all the X chromosome regions replicate. In d0Fr7/FM6 females, the late-replicating nuclei in the 2B region with normal morphology then cease incorporating [3H]thymidine. When a block of compact material is formed in this region, it becomes one of the most late replicating in both durvar7/Y and hUQr7/O (Figure 156).


413

d w-

xlDp ~n2wY,~25

c

Figure 154. Effect of the Y chromosome heterochromatin and tem-

perature on the break frequency in the X chromosome (e-h) and extension of compaction ( a d ) in Dp(l ; I )pn2b duplication. Abscissa for a d , compaction state from the 2E1-2 band to the indicated region; abscissca for e-h, breaks in the 3C, 11A6-9, and 19E regions; ordinate, occurrence frequency; N, normal chromosome. After Zhimulev et al. (1989b); reprinted by permission from Zhimulev et al. (1989a).

As shown earlier, chromosome compaction can for this reason variegate

lDE, the late-replicatingregion (Ananiev and Gvozdev, 1974), quite distant from the heterochromatic junction. This region is, presumably, associated with one of the regions of discontinuous compaction.

414

I. F. Zhimulev

7

5, n 27

a

m 21

3

-I

1.

I

Figure 155. Block formation frequency in the 2B region indorvm7/0males developed at different temperatures: (a) 25°C; (b) 18°C; (c) hrst 6 hr after egg laying at 18°C and then at 25°C. Abscissa, mean frequency of nuclei with compact blocks in a larva; ordinate, number of larvae; n, number of studied larvae; m, mean frequency of blocks in the experiment Reprinted by permission from Zhimulev et nl. (1988).

(a).

E. DNA underrepresentation The idea that DNA underreplication is caused by position effect has been widely discussed in the literature (see Section XVI,A). What facts have accumulated since the middle of the 1930s! Caspersson and Schultz (1938), and subsequently Cole and Sutton ( 1941), spectrophotometrically measured relative DNA contents in bands placed close to heterochromatin by a chromosomal rearrangement. In the T( 1 ; 4 ) ~ translocation, ~ ~ ~ ~ whose - ~ junction ~ with the X chromosome heterochromatin is between the 3E5 and 3E6 bands, the inactivation elicited by position effect spreads toward the 3C2-3 band (the w+ locus). Genetic inactivation does not spread toward the bands of the 3F region. The amount of DNA, which was inexplicably measured in the 3F1 and 3F3 bands, proved to be even greater

c

d

n=22

n=27

var7/0

mwt‘m

t-t-t-Ft-r-5 e

n=9

c-t-r-c-4

I

8

f

n-16

Figure 156. Replication of the 2B region in the translocated T(1;2)dorUm7 element in normal (a, c, and e) and compacted (b, d, and f) states in war7/FM6 females (a and b) and var71y (c and d) and var7/0 (e and f) males. Late-replicating regions of the X chromosome and the 75C-80Csegment ofchromosome 3L are designated. In building the histograms, the presence of label in each region was noted; then the nuclei were ranged according to decreasing number of labeled sites in the given regions. Each horizontal line corresponds to an analyzed nucleus, where the symbol “x” designates labeling and a blank space indicates absence of data on labeling of the given nucleus. Nuclei with a minimum number of labeled sites are at very top of the histograms, and therefore, at the latest replication stages. n, number of nuclei. After Bolshakov and Zhimulev (1990); reprinted by permission from Zhimulev et nl. (1989a).

416

I. F. Zhirnulev

than in the chromosome without the translocation. According to other data, there is no stable difference in the content of the absorbing agent in the 3E1 and 3C2-3 bands (i.e., in the direction of genetic inactivation) between the normal and the translocated chromosomes. Scanning microdensitometry of salivary gland chromosomes showed a reduction of 30% in the ratio of the Feulgen-stained DNA contents in the 10D1-2 band relative to that in the 10A1-2 and ?El-2 bands in the single inversion In(l)mK chromosome, which became heterochromatized to some extent at 14°C. The Feulgen-stained DNA contents decreased only when the second In(2)RevB inversion was present in the genome (Wargent et al., 1974). Microdensitometric measurements in the 3D1-3ED region closely adjacent to the heterochromatic junction demonstrated a decrease of 10% in Feulgen-stained DNA content when the strain had the T(l ;4)wm258-21translocation. In the strain in which the region was more distant from the breakpoint (3C1-10 region), the Feulgen-stained DNA content was somewhat higher than normal; the 2E1-3A4 region did not differ in Feulgen-stained DNA ratios form the control (Hartmann-Goldstein and Cowell, 1976; Cowell and Hartmann-Goldstein, 1980a). When position effect was modified by low temperature (15”C), the relative DNA content was unaltered, although enhanced heterochromatization was expected. An extra Y chromosome in the females had no observable effect on the DNA content (Cowell and Hartmann-Goldstein, 1980b). It was not specified whether the measured DNA values were for “heterochromatized” or the nonaffected chromosomes. No relationship was found between variegation and total DNA content in malpighian tubule cells (Hartmann-Goldstein, 1981). Based on the photometric results, it is difficult to make judgments concerning the possibility that DNA underreplication may result from position effect. In studies of the sal locus in D. pseudoobscura, Gersh (1973) found that, when the tip of the third chromosome with the sal+allele is transferred to the heterochromatic chromosome in the T(x3;4) 19Ocv7Of translocation, the number of individual bands reduces from 15 to 4-5 and the whole fragment is occasionally not visible in the nucleus. According to Ananiev and Gvozdev’s (1974) data, the Dp( 1;f)R duplication is identifiable in only 28% of the salivary gland nuclei in X/O/Dp(1 ;j)R males grown at 18”C, and in 90% of X/Y/Dp(I ;f)R males grown at 25°C. Furthermore, measurements of relative DNA content based on long-term [3H]thymidine incorporation demonstrated a 20% decrease in DNA content in the whole duplication compared to the homologous chromosome region in X/O/Dp males ( 18°C). According to other data, “breaks”are formed in chromosome regions the compaction of which is due to position effect (see Section XVI,D), which is also an indication of DNA underreplication in these regions. Arguing for underreplication of the 3C-E region in T(l ;4)wm25s21, Reuter et al. (198213) provided photographs of thin strands in the 3C region where a “weak spot” is located (i.e., the


41 7

region of the underrepresented DNA in wild strains) (see Section VII,C,). This raises the question as to whether polytenization in this region is associated with position effect variegation or with the expression of the normal gene activity. The results obtained with in situ hybridization were confusing. The T(Y;3)A78 translocation, which places the 87C heat shock puff next to Y heterochromatin, does not affect the puffs. There was no puffing or labeling of nuclei from the salivary gland chromosomes of heat-shocked larvae. When the puff was not induced in a smaller proportion of cells, nascent RNA chains did not accumulate. Label was not seen when the hp70 heat shock gene was hybridized on such nuclei. There was, however, a threefold increase in grain count at the puffed 87C locus versus at the same sites in the control nonrearranged homologs. The author attributed the decrease in labeling to the DNA being in a state of compaction, which is less hybridizable than puffed DNA (Henikoff, 1979a, 1981). In hybridization of the pA54 cosmid DNA, which contains a DNA fragment mapped next to the In(1LR)pnZa breakpoint, with squash preparations of the chromosomes of gn2u/+ heterozygotes, labeling level was the same in the nuclei of both homologs. Indirect judgments about cell variegation were based on a decrease in 6-phosphogluconate dehydrogenase (6PGD) activity (56% of normal activity) in another group of cells treated histochemically (Alatortsev, 1986, 1988).Alatortsev (1986,1988) inferred that, genomic DNA corresponding to the pA54 fragment surely is subject to position effect because the 6PGD gene is distal to the pA54 fragment and it is not inactivated. The data on discontinuous compaction (Belyaeva and Zhimulev, 1991a) weaken his inference. There was no control of genetic inactivation in the region (for blocks of compact DNA) at the chromosomal level. Comparisons of the hybridization level of P1471 (see Belyaeva et al., 1987) with the T( I ;2)d43roar7chromosome, the genomic DNA of which, as in the P1471 clone, is in the state of compacted blocks, as well as comparisons of this clone and the normal chromosomes, revealed no in situ hybridization in the former experiment (Zhimulev et d., 1986). Using the T(l ;2)hrVar7rearrangement, the dependence of in situ hybridization on distance of the DNA fragment from the translocation breakpoint and compaction degree of DNP blocks was examined. It was found that clones mapped at a distance from 30 to 150 kb showed no labeling in the case of hybridization with the compacted blocks of heterochromatin of females. The P266 clone located further, at a distance of more than 200 kb from the breakpoint, hybridizes with chromosomal DNA and somewhat variegates: there was no hybridization with the clone in some larvae and only slight hybridization in others. Hybridization intensity was indistinguishable from normal in yet other larvae. Blocks of compact material were present in the chromosomes in all three cases (Umbetova et al., 1991). There is no straightforward explanation for the results obtained in the two latter cases: the DNA in the compact blocks becomes completely inaccessible to heterochromatin because chromatin is

I. F. Zhimulev

418 ~

very tightly packaged. It is of interest that even the clones nearest to the rearrangement breakpoint hybridize to the loose blocks in the dor”“’ chromosomes of males (see Section XV1,D) (Umbetova et al., 1991). In situ hybridization of the DNA of the w+ gene with the In(l)wm4 inversion is considerably less intense (a reduction to 11-1496) of that found in the normal chromosome or the inversion when the Su(vur)323 gene suppressed variegation (Hayashi et al., 1990). Southern blot analysis could be decisive in resolving the question of underreplication. However, the results on this issue were conflicting. Determination of DNA content in the fat body cells of R(ry+”)/ry2 larvae with the ry+’ chromosomal rearrangement exerting rosy+ position effect demonstrated that DNA fragments of the ry+ locus from both homologous chromosomes (?+*I and ry2; 7.2 and 11.5 kb, respectively) occur in the same proportions regardless of the presence of the Y chromosome. Genetic inactivation of the chromosome region was controlled by changes in the activity of the xanthine dehydrogenase (XDG) gene mapped to this region. It should be noted that XDG activity was reduced up to sevenfold in WO; R(ry+”)/ryz individuals compared to individuals with the Y chromosome. Similar results were obtained with Southern blot analysis of RNA extracted from larval malpighian tubules and salivary glands (Rushlow et al., 1984). The w+ gene was inactivated when In(l)wm4 transposed to heterochromatin; w+ DNA content decreased neither in salivary gland nor in adult fly cells (Hayashi et ul., 1990). Another study involved salivary gland cells of larvae heterozygous for T( I ;4)wm258-21exerting position effect variegation on the w+ gene and also on Sgs-4, an adjacent gene encoding a glycoprotein component of salivary gland secretion. It was shown that the DNA from the Sgs-4 gene is polytenized approximately by 30% in 17°C culture when compared to normal. This is associated with a substantial decrease in the amount of RNA and protein synthesized by the gene (Kornher and Kauffman, 1986). There is uncertainty about the “compaction” of the 3C puff region. True, band frequencies decrease at 3C, D, and E in the translocated homolog at 17°C. According to Kornher and Kauffman ( 1986) “compaction” is demonstrated in their Figure 2a. However, the XD4p is not the fragment seen, as stated in the legend to the figure; rather, it is XBer4Dbecause the band sequence is normal to 4C1-2 (to the right of the arrow with the asterisk) and there can be no variegation at Sgs-4 in this chromosome. Material of the fourth chromosome should immediately follow the 3E5-6 band. It is therefore unclear what the authors meant by compaction. In the Dp(1 ;f) 1 187duplication, the euchromatic sequences 1.9 kb away from the euchromatin-heterochromatin junction are severely (39-fold) underrepresented in polytene chromosomes relative to diploid DNA of X/O/Dp males. The sequences, which are at a distance of 54 and 103 kb, are underreplicated 8-


419

and 2.4-fold, respectively. Underrepresentation is weaker in individuals with a Y chromosome, and there is a reduction in copy number values for the three fragments varying from normal to 4.7-fold for the one lying closer to the breakpoint (Karpen and Spradling, 1990; Spradling et al., 1992). Studies were performed in which Southern blot hybridization was combined with control of genetic inactivation on cytological preparations based on the formation of blocks of compacted chromatin (see Section XVI,D). A genetic system Dp( 1 ;I )pn2b and clones of the BR-C gene were used; the rearrangement breakpoint and the gene were quite distant, being separated by 29 bands on Bridges' map. To estimate DNA underreplication, Southern blot hybridization, allowing the comparison of copy number in a fragment of the ecs gene in various populations subject to maximal and minimal position effect, was used. To distinguish the DNA of this gene in the duplication, where it is position affected, from the non-position-affected chromosome, In(I) brLr103was used. The inversion breaks one of the fragments of the HindIIl gene of 4.4 kb (Figure 157) into two parts, with the result that non-position-affected DNA, homologous to the 4.4 kb fragment, is represented by two new fragments of 5.5 and 7.5 kb, while the same

Figure 157. Southem blot hybridization of the 4.4-kh Hind111 fragment with DNA from males of third instar larvae (a and b), salivary gland ( C and d), fat bodies ( e and f), and head complexes (g and h). (a) X/Y/pnZb (25°C). (h) bTltio3/Y (25°C). (c,

e,andg) bSt'03/0/pnZb(18" (d,f, C).andh) btl'03fl/pn2b(25"At C)the . left are markers of molecular length (kb). After Umbetova et al. (1990);reprinted by permission from Umbetova et al. (1991).

420

1. F. Zhimulev

DNA in the duplication (i.e., the one undergoing variegation) is represented by a single fragment of 4.4 kb. This makes it possible, within each variant, to compare DNA amount of a position-inactivated chromosome with the normal (the non-position-inactivated)chromosome on the same lane of the electrophoretic gel (see Figure 157). When 14 different transformed lines with variegating P-element inserts were used to examine the DNA levels, Southern blot analyses showed that the heterochromatic hsp26 transgenes were underrepresented 1.3- to 33-fold in polytene salivary gland chromosomes relative to the endogenous euchromatic hsp26 DNA (Wallrath et al., 1996). Distances between the euchromatin-heterochromatin breakpoint and the DNA site that is underrepresented can be very long: 50 or 100 kb (Karpen and Spradling, 1990; Spradling et al., 1992). These distances can be many hundreds of kilobase pairs between BR-C in the 2B3-5 region and Dp(l;l)pn2bin the 2E1-2 region (Umbetova et al., 1991) or even thousands of kilobase pairs when the whole fragment between 287-8 and 5D is lost in polytene chromosomes (Pokholkova et al., 1993a). It was shown that the underrepresentation level is very much dependent on polyteny degree in the cells. In females, whose salivary gland chromosomes are maximally polytenic, DNA underreplication was most expressed in approximately 80-90% of cells (Figure 157c and 157d). In the chromosomes of the fat body, where polyteny degree is much lower, DNA underrepresentation is manifested considerably more weakly, and there is no underrepresentation in the diploid cells of the cephalic complex (see Figure 157e-h) (see, as well, Henikoff, 1979a, 1981; Rushlow et al., 1984). In lines with variegating P-element inserts, the heterochromatic hsp26 transgenes are present in approximately the same copy number as endogenous euchromatic hsp26 genes in diploid tissue (Wallrath et al., 1996). This conclusion is consistent with the earliest data on immutability of the genes subject to position effect variegation (recovery of gene activity after reversal of a chromosomal rearrangement or removal of the gene from the neighborhood of heterochromatin by crossing over), and it is also consistent with the more recent data obtained by P-element-mediated transformation (see Section XII,A). Clearly, only gene inactivation, expressed as compaction, seems to occur in diploid cells. Underreplication seems to be a feature of mainly polytene chromosomes falling into the highest polyteny class level. The compaction degree of position-affected chromatin plays the major role here (see Section XVI,D), and this may be a reason why different conclusions were made for the possible underrepresentation. It may be conceded that the lowest level of genetic inactivation due to unfeasible puff induction or inactivation does not lead to underrepresentation, too. The formation of a loose block in male polytene chromosomes, which also does not lead even to genetic inactivation, as shown by in situ hybridization (Umbetova et al., 1991), is also not caused by incomplete polytenization of the region.


42 1

No differences in restriction fragment length (no underrepresentation of DNA) were found between regions from normal (non-position-affected) or rearranged (position-affected) chromosomes at either side of h ( l ) w r n 4which , variegates for the white gene (Locke, 1993). In a small proportion of cells of dOrUar7/FM6females, ostensibly compacted blocks are formed; however, these do not completely suppress the activity of the BR-C locus they comprise (Zhimulev et ul., 1986). In the most compacted block in which DNA is stably inactivated, the inactivation may be different because the chromosome region involved in compaction extends for greater distances. It may be assumed that the detectable genetic inactivation caused by position effect may be the consequence of both compaction and underreplication. It appears a reasonable assumption that, spreading from the heterochromatic junction, compaction carries an increasing number of new regions with it, and DNA underreplicates after delay. For this reason, some regions will be lost because of underreplicated, while others will still be present. Since compaction can be discontinuous (Belyaeva and Zhimulev, 19911, the pattern described in the preceding section may prove to be much more complex. There is the probability of variegated disposition of the quite extensive fragments of the compact chromosomes, with endoreplication differently expressed in each. With this in mind, the diverse results and views relating to DNA replication resulting from position effect cause no surprise.

F. Change in the pairing properties of chromatids-formation of “pompons” By irradiation of BurM2males, Belgovsky has induced mutants that, although not expressing the Bur character, showed a variegated forked phenotype: fBl.5, fB27, fB.59, andfBl68. The original BM2chromosome had an inversion withone breakpoint between f and B and the other proximal to bb, although distal to the centromere. The arising reinversions presumably have a breakpoint not quite identical with the original, and, as a consequence, the forked locus remains in chromocentral heterochromatin and the 15E1-Fl bands are accordingly lost from the euchromatic 15 region, and there appears a small fragment of heterochromatin, despite a generally recovered banding pattern (Belgovsky and Muller, 1938; Belgovsky, 1944, 1946). It is likely for this reason that ectopic contacts between the 16A region and centromeric heterochromatin frequently occur in thefB59strain (Bose and Duttaroy, 1986). It is of interest that the fB59 chromosome becomes “infected” with variegation after it has resided in the heterozygote with the d149 inversion, and that in spite of suppressed crossing over (Belgovsky, 1944). Unusual changes in the structure of polytene chromosomes were observed both in the original BM2 inversion and in one of the f B I 5 reinversions.

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F. Zhimulev

“Pompon”-likechromosomes are formed in 36% of nuclei in BM2 larvae reared at 10°C (see Zhimulev, 1992b, 1996, for a characterization of the chromosomes of this type). No changes in incorporation intensity of t3H]uridine or [3H]thymidine were observed (Lakhotia and Mishra, 1982). InfBJ5/Yindividuals, at three different temperatures (18,24, and 28”C), 39.81%, 34%, and 30.21% of cells of the X chromosome, respectively, had a pompon-like appearance. Removal of the Y chromosome in fBf5/0 larvae caused an increase reaching 75.1 7%, 43.85%, and 80.5%, respectively, in the proportions of cells with such chromosomes. The transcriptional activity of the “pomponized” chromosome does not alter as in the original BM2 chromosome. Introduction of a duplication into the genome in the 16A-20F region completely eliminates the appearance of pompons. The addition of a single 16A region has almost the same effect (4% of cells with pompons). When a set of deletions partly remove the histone genes, the frequencies of cells with pompon-like X chromosomes varies in the 17.02-30.6% range, and their frequencies fall to 9.35% only infBJ5/O;Df(2L)6/+ genotype (Bose and Duttaroy, 1986). It cannot be ruled out that the gene encoding the protein responsible for chromatid pairing in the polytene X chromosome or associated, in some way, with dosage compensation may become inactivated because of position effect of this type.

XVII. CURRENT CONCEPTS OF THE MECHANISM OF POSITION EFFECT VARIEGATION A. Change in the state of chromatin due to position effect variegation There is a fairly widespread conviction that the passage of a euchromatic region to an inactivated state caused by position effect variegation is associated with change in the condition of chromatin: its compaction degree, and the rate and order at which the nonhistone and histone proteins, the major DNA components, are self-assembled (Schultz, 1965; Spofford, 1967; Sutton, 1972; Gersh, 1973; Zuckerkandl, 1974; Henikoff, 1979b; Reuter et d.,1982a, 1990; Sinclair et al., 1983; Zhimulev et al., 1989a,b; Belyaeva and Zhimulev, 1991a; Spofford and DeSalle, 1991). In Section XVI,D, it was clearly shown that compaction of a chromosome region in which an inactivated gene resides is the mechanism whereby position effect variegation modifies phenotype and, as a consequence, transcripts and protein product of this gene are not observed in cells. Passage to the compact state is associated with considerable structural and morphological alteration in a chromosome region acquiring properties of heterochromatin, such as late replication, incomplete polytenization, breaks, and ectopic contacts. There are presumably also differences in the level of genetic inac-


423

tivation between the compact chromosome region and an inactive band. If the DNA condensed to a band can be activated by an appropriate inducer, then irreversible inactivation presumably follows during compaction resulting from transposition to heterochromatin, and the genes cease responding to the inducer. Quite obviously, new molecular structures must correspond to a new state of heterochromatin, presumably to the newly appearing compacting proteins. The relevant concepts have been discussed for quite a long time. Sutton (1972) suggested that a certain fraction of chromatin (e.g., that containing repetitive DNA) can be somewhat supercompacted, even “crystallized,” and thus subject to complete inactivation by modified histones. Zuckerkandl ( 1974) proposed that hypothetical locking proteins lock euchromatin into heterochromatin conformation and maintain it in an inactive state. When euchromatin is brought close to variegation-affected heterochromatin, the formerly euchromatic regions specifically interact with the locking molecules that lock them. Many nonhistone proteins capable of binding to DNA have been described, and amino acid sequencing has revealed patterns of their structural organization. Small domains in the protein molecule containing up to 100 amino acids are sufficient to provide specific binding to DNA, and create a tertiary structure of the molecule that already interacts with DNA. The following four types of domains are distinguished:

1. The helix-turn-helix motif. Two a-helices separated by a p-turn underlie the structure (Figure 158a). Amino acids directly make contacts in the major groove delimited by the DNA molecule in one “recognition helix.” The other helix lies across the major groove and makes nonspecific contacts with DNA. In the eukaryotes this structural motif occurs in the protein family of Drosophila and, with the participation of these proteins, makes key decisions regarding the entire development of the organism. The proteins contain a highly conserved region consisting of a 60-amino-acid homeodomain. 2 and 3. Cysteine-histidine and cysteine-cysteine “zinc-fingers.”A unusual structure of TFlIlA was found: a protein factor required for transcription of the SSRNA gene by RNA polymerase 111. This molecule contains 9, on average (ranging from 2 to more than lo), repeating units of approximately 30 amino acid residues; these comprise 7-10 zinc atoms (see Figure 158b and 158c). Each unit comprises two invariable pairs of properly spaced cysteines and histidines providing coordinate binding to zinc atoms. Zinc finger motifs are known to occur in various eukaryotic proteins, from yeast to human. Some of the zinc finger motifs are activators of trancription, whereas others are of importance in development and sex determination. 4. The “leucine zipper.” This motif is found in several transcription factors.

424

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Cys-Cys zinc finger

F. Zhirnulev

leucine zipper

Figure 158. Models of various domains in protein molecules binding DNA. (a) Helix-tum-helix; arrows indicate the direction of the spiral of protein. (b) Cysteine-histidine “Zinc finger” (CH). (c) Cysteine-cysteine zinc finger. (d) Leucine-leucine (L) “tipper”. Rectangle, contact region with DNA. After Struhl(1989).

Four to five leucine residues are spaced exactly seven amino acid residues apart and hence could by viewed as being repeated every two turns of an ahelix. In this way, the leucine molecules in the a-helices of various proteins occupy the same position (see Figure 158d). The presence of another such sequence allows them to interlock like a zip fastener (Berg, 1988; Evans and Hollenberg, 1988; Struhl, 1989). Silencing is a process that establishes and maintains repressed transcriptional domains. Properties of Drosophila chromosomes suggest that heterochromatin and repressed transcriptional stages of inactive domains are formed through a spreading mechanism from inactivation centers (see Tartof et al., 1984, 1989; Balasov and Makunin, 1994,1996). Silencing is a particular type of transcriptional repression characterized by the formation of a heritable genetically repressed state of chromatin (for review, see Rivier and Rine, 1992; Shaffer et al., 1993; Becker, 1994;Moehrle and Paro, 1994; Pirrotta and Rastelli, 1994;Rivier and Pillus, 1994, Eissenberg et al., 1995; Shore, 1995; Holmes and Broach, 1996; Zhimulev, 1996).

425

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation ~

~

~

_

Several groups of proteins have been found that can be involved in the silencing process, including HP1, an abundant heterochromatin-associated protein (see Table 30 in Section XIII); and the polycomb group proteins, the products of a class of genes involved in maintaining the repressed status of the homeotic genes (Eissenberg, 1989; Zink et al., 1991; DeCamillis et al., 1992; Messmer et al., 1992; Reuter and Spierer, 1992; Fauvarque and Dura, 1993; Orlando and Paro, 1993; Powers and Eissenberg, 1993;Schlossher et al., 1994;Eissenberget al., 1995; Lohe and Hilliker, 1995;Weiler and Wakimoto, 1995; Elgin, 1996). Critical to this interpretation is the notion that heterochromatin formation, as assayed by position effect variegation, is sensitive to the dosage of any one of 30-50 factors. According to Locke et al. (1988),equilibrum exists between a population of constituent molecules and the formation of a heterochromatic structure formed by the association of these components. Reduction in the dose of any component would shift the equilibrum away from complex formation. Partial deletions of the histone cluster result in suppression of PEV (Moore et al., 1983); similar effect is obtained when larval growth lasts in the presence of butyrate, an inhibitor of histone deacetylation (Mottus et al., 1980). The suppression effect of Su-var(2)lo’ was associated with a state of hyperacetylation of histone H4 (Dom et d., 1986). Analysis of the heterochromatic properties of yeast telomeres has shown that effective silencing, in that case, appears to require localization to the periphery of the yeast nucleus; this position requires RAPl, SIR3 and SIR4, the latter two proteins interacting with the silencing domains of histones H3 and H4 (see Elgin, 1996, for review). Mutations in the highly conserved amino-terminal tail of histone H4 in yeast lead to decreases in telomeric silencing (see Shore, 1995; Turner, 1995, for reviews). Conversely, ample evidence indicates that there are heterochromatic proteins that are specific to it only (Rodrigues Alfageme et al., 1980; Will and Bautz, 1980; Levinger and Varshavsky, 1982a,b; James and Elgin, 1986; James et al., 1989; Reuter et al., 1990). One such protein (HP1) is encoded by the suppressor gene Su-var(2)205 (James et al., 1989). Point mutations in the HP1 chromodomain abolish the ability of HP1 to promote gene silencing (Plater0 et al., 1995). This is a 30-50 amino acid domain that is conserved in several eukaryotic chromatin binding proteins such as HP1, Polycomb (PC), Su(var)3-9, malespecific lethal-3 in Drosophila, their mammalian homologs and fission yeast SW16. HP1 and PC proteins bind to numerous, specific, non-overlapping loci in chromatin (see Koonin et al., 1995, for review). Many more details on heterochromatin and silencing proteins can be found in review by Eissenberg et al. (1995). Based on cDNA sequencing, it was deduced that the protein of the gene Su-var(3)7 has 932 amino acid residues with seven zinc finger domains of cysteine2-histidine, (see Figure 158b). The zinc finger proteins are not contiguous,

_

_

426

1. F. Zhimulev

being separated from each other by 40-107 amino acids. This may provide conditions requisite for making contacts with large domains of DNA and, consequently, for more extensive compaction and genetic inactivation (Reuter et d., 1990; Cleard et al., 1995). It was reported that a protein containing a homeodomain and regulating a particular state of differentiation shares a homologous domain with proteins that are suppressors of variegation (Paro, 1990; Par0 and Hogness, 1991). Direct localization of antibodies against protein HP1 in the chromosomes of the Dp(1; 1 )pn2B strain demonstrated that they occupy blocks of compact heteochromatin resulting from position effect variegation. The protein was then detected both in “continuous” blocks, when compaction started from the euchromatin-heterochromatin junction and spread distally, and in blocks arising during variegated compaction (Belyaeva et al., 1993; Demakova et d., 1993). Drosophila and yeast have no known covalent DNA modification. Introduction of the dam methyltransferase gene from E. coli and induction of its expression giving rise to lo6 methyl groups per genome does not affect the development of fly. Use of PEV line for methylase assays has shown differences in inactivation of marker gene before and after differentiation (Wines et at., 1996). Is there any specificity in DNA sequences of euchromatin subject to the influence of heterochromatin and to which compacting proteins bind? From general considerations it follows that there is no discernible specificity at the level of the macrostructure of the chromosome. The BR-C region in D. melanogaster was cloned for a distance of about 250 kb and structural features of DNA, including long tracts of repeats, were not found in the region (Belyaeva et al., 1987). However, when transferred to the neighborhood of heterochromatin, these regions became compact owing to binding to the compaction HP1 protein (Belyaeva et d., 1993). As shown in a very interesting model experiment, repetitive sequences induce heterochromatin formation, possibly because these sequences can pair with one another (Dorer and Henikoff, 1994). Arrays of three or more mini-white transgenes, P(hW), produced phenotypes similar to classical heterochromatin-induced position effect variegation. These phenotypes were even modified by PEV modifiers, such as the Y chromosome or Su(var)205. The PEV mutant phenotype strengthened with increasing copy number of the repeated transposon. The authors propose that pairing of repeats underlies heterochromatin formation and is responsible for gene silencing. The proposal is similar to an idea of Pontecorvo (1944) that heterochromatin is derived simply from the repetition of sequences. Since the P(lacW) transposons do not contain heterochromatin-specificsequences, it appears that this “heterochromatization” results from the repeated nature of the transgenes (Dorer and Henikoff, 1994; Henikoff, 1994). Similar “heterochromatization”was found for three and more copies of the transposon P(bw+) (Sabl and Henikoff, 1996). The general model for position effect variegation is given in Figure 159. When a chromosomal rearrangement moves a gene close to heterochromatin,

-

Polytene Chromosomes, Heterochromatln, and Position Effect Variegation

Figure 159. Model of position effect inactivation. (Top) The repeat organization of a chromosome in the vicinity of euchromatin-heterochromatin rearrangement. T h e solid line represents single-copy DNA in euchromatin (eu) and the striped and shaded lines represent two different families of repetitive sequences in heterochromatin (het) and scattered in euchromatin. (Bottom) Heterochromatin forms as a result of local pairing between homologous double-stranded DNA sequences forming hairpin, loop, and more complex structures (magnified in inset), which participate in the formation of a chromocenter (gray oval) on the nuclear envelope (dotted line). Silencing of a gene by PEV can occur when a dispersed repeat nearby pairs with a homologous sequence in a block, sequestering the gene into a heterochromatic environment. Reprinted by permission from Sabl and Henikoff (1996).

427

428

I. F. Zhimulev

then pairing between any element near the gene and similar sequences clustered in heterochromatin bring the gene into the heterochromatic compartment, causing inactivation (Sabl and Henikoff, 1996). Therefore, data on repeats distribution are of importance. Of interest in this regard are blocks of mono- and dinucleotide repeats, whose sizes vary in the range of 10-60 base pairs. Three types of repeats, (dCdA)n.(dT-dG)n, (dC-dT)n.(dA-dG)n, and (dC)n*(dG)n,most frequently occurring in the eukaryotes, are detected in Diptera (see Section VII,C,6). It was shown that certain proteins preferentially bind to the DNA of this type (Vashakidze et al., 1988a,b). The distribution of protein HP1 in polytene chromosomes of D. melanogaster, the product of the Su-war(2)205 gene, is completely complementary to the location of the repeat (dA-dC)n.(dT-dG)n (E. B. Kokoza, 0. V. Demakova, E. S. Belyaeva, and 1. E Zhimulev, unpublished observations). In D. melanogaster, protein HP1 is present in the region of a-and p-heterochromatin, in the fourth chromosome, in the telomeres, and in the 31A-F region (Jameset aI., 1989). The dA-dC repeat is undetectable precisely in these regions. Thus protein HP1 does not bind in chromosome regions containing at least simple dinucleotide repeats AC-GT, and these regions are presumably not subject to deep inactivation by proteins compacting Su-uar(2)205 (Belyaeva et al., 1993). It was suggested that absence of AGGT repeats from heterochromatin affects the general structure of chromatin and, consequently, the degree of genetic inactivation caused by position effect variegation (Huijser et al., 1987). AC-GT repeats were found around breakpoints of about half of chromosome rearrangements in the 2B region of the D. melamgaster X chromosome. Analysis of sequences neighboring the breakpoints of the rearrangement T(l ;2)dWar7,evoking position effect variegation, demonstrated that the breaks between two sequences of AC-GT repeats occurred at distances of 80 and 156 bp. The pentanucleotide repeat (CTGTT),, is located at a distance of 660 bp from the breakpoint. It differs by one nucleotide from the sequence of the GTGTT heterochromatic satellite attached to the 1A-2B fragment joining to heterochromatin (Makunin et al., 1996). Some middle-repetitive heterochromatic sequences were found at PEV breakpoints (Tartof et ul., 1984). One family of repeated DNA, a GC-rich 11-12 mer repeat found in heterochromatin, is widely dispersed within higher eukaryotes (Abad et al., 1992). According to another hypothesis, specificity at the DNA level is provided by inter- and intragenic noncoding sequences. Specifically,they bind to particular regulatory proteins (Zuckerkandl, 1981). Selection of individual nucleotides is not the condition necessary for the existence and conservation of function in the majority of noncoding sequences. Indeed, proteins can bind with relatively low affinity and specificity to multiple sites. However, specificity of protein binding to DNA is achieved through cooperative effects (Zuckerkandl and Villet, 1988).


429

B. Modification of compaction degree Elaborating the notion of locking proteins, Zuckerkandl ( 1974) suggested that competition of locking molecules for binding sites of DNA underlie modification of heterochromatin amount. Obviously, the more heterochromatin in a cell, the larger the number of compacting molecules binding to it to leave the nuclear pool and the smaller the number of loci that can be activated. In fact, Khesin and Leibovitch (1976) demonstrated that even heterologous DNA can be a “magnet” attracting locking proteins. The basic mechanism providing the effect of modifier genes on variegation expression is also explicable (see earlier). It is much more difficult to explain how low temperature enhances genetic inactivation. It is pertinent to note that reduced temperature significantly affects development: the majority of, if not all, poikilotherms reach larger body size when developing at lower temperatures (Sokoloff and Zacharias, 1979). The morphology of polytene chromosomes is greatly affected by low temperature (15°C). The proportion of nuclei falling into the highest polytenic classes was higher and nuclear dry mass was greater in larvae reared at 15°C than in those grown at 24°C (Hartmann-Goldstein and Goldstein, 1979a,b). Having reared Drosophila for two generations at 16°C (the controls were cultured at 25°C) and, then, having determined the amount of satellite and ribosomal DNAs in adult cells, Chernyshev and Leibovitch (1981) found a small (from 1% to 18%) excess in the amount of these DNAs in the cold-reared flies compared to the controls in all the experiments. It is not clear how the increase in the number of sequences composing the bulk of heterochromatin may be related to enhanced inactivation at low temperature.

C. Initiation of inactivation A central unresolved issue in the problem of position effect variegation is: Why are the euchromatic genes inactivated when transposed to heterochromatin? If the effect of heterochromatin is not specific, then why are the genes at the euchromatin-heterochromatin junction in the normal chromosomes not inactivated? An answer to the second question may be found by envisioning the following. A string of regions, including the more compacted a-heterochromatin and the looser compacted p-heterochromatin, retaining a banding pattern may be located between the euchromatic part itself and the centromere. Perhaps the gradient in compaction from euchromatin to a-heterochromatin, stretching along the enormous length of the block of heterochromatin, is coincident with the distribution gradient in compacting proteins, with the result that protein concentration may be the same on either side of the heterochromatin-euchromatin junction. The possibility cannot be dismissed that there is a particular nucleotide sequence that accomplishes boundary functions. If so, all the chromosomal rearrangements with breakpoints distal to the sequence would not cause position ef-

430

1. F. Zhimulev

fect, while those breaking the block of heterochromatin located between the boundary-developing sequence and the centromere would cause it. Such is not the case. Examples abound of rearrangements breaking heterochromatin and not causing position effect varigation (see Section XVI,B). Of relevance here is Baker's (1968) inference that not only transfer of a gene to the neighborhood of heterochromatin, but also a broken heterochromatin sequence, is required for position effect variegation to occur. This inference possibly prompted Tartof and his group to develop models of domain compaction. According to the first extensive model, sequences that are true satellite DNAs (e.g., Figure 160a) must be present in heterochromatin. Position effect arises when the organizational features of heterochromatin are transferred to sequences nearby these regions. It is Tartoff's (1984) view that the hypothesis does not explain how inactivation can extend over the long distance of a 60-band region. The boundary model is an alternative (see Figure 160b). Its only requirement is determination of the edges of a heterochromatic domain. The edges may begin with an initiator (i) and end with a terminator (t) site. All the DNA located between the two sites is necessarily compacted and, thus, becomes heterochromatic. A variant of the model implies absence of a terminator, if the initiator sites are in reverse order (Figure 160c). Genetic inactivation due to position effect is readily explained in terms of this model: a chromosomal rearrangement breaks the domain and transfers the gene to the sphere of influence of initiation. Having started from the initiator, compaction propagates itself into the euchromatic part of the chromosome, too (Tartof et al., 1984,1989). Experimental generation of revertants in the T(I ;2)dorvar7 strain was described earlier. In another experimental series, heterochromatic segments of chromosome 2R were transferred from the neighborhood of 1A to 287-8 to the vicinity of 2B7-8 to 7A of the X chromosome of Drosophila T(1 ;2)dor'e*5. This reversed the variegated expression of genes located in the 1A to 2B7-8 region and produced heavy compaction of the 2B7-8 to 7A region. This particular heterochromatic fragment may be an initiating compacting domain (Pokholkova et al., 1993a,b). The recognition came early in the 1940s that not every chromosomal rearrangement with a single heterochromatic breakpoint is subject to position effect variegation. When taken together, the pertinent data indicate that the various heterochromatic segments are not identical in respect to the degree of variegation they induce. This suggests that either the compaction-initiating domains vary considerably in their inactivation potentials or they alternate with heterochromatic regions that are incapable of eliciting compaction, that is, the initiating domains are not tandemly arranged as depicted in Figure 160. Accepting the existence of compaction domains, it is not clear how inactivation can spread at a considerable distance. Several hypotheses may be offered for examination:

1. Ptashne (1986) takes the view that there exist proteins that hold together the bases of large DNA loops and thereby compact long stretches of chromatin.


COEXTENSIVE MODEL M,

S'S'S'

s s s

BOUNDARY MODEL N

+

C

Figure 160. Models of gene inactivation under position effect. (a) A heterochromatic block is composed of DNA sequences that are true heterochrornatin. A gene (e.g., w') being transfered there could be involved in compaction with heterochromatin. (b) DNA located between the sites of initiation (i) and termination (t) of compaction becomes heterochromatic. (c) The termination site is not required if the initiators face one another. After Tartof et

al. (1984, 1989).

43 1

43 2

1. F. Zhimulev

2. The protein product of the Su-var(3)7 gene suppressor has unusual zinc fingers separated by amino acid residues not comprised by the finger itself, and this circumstance promotes contact of protein with the more distant sites on the DNA (Reuter et al., 1990). 3. Zuckerkandl(1974) proposes that the length of a compacted region is related to compaction time. His idea is that inactivation spreads only during DNA replication and a particular DNA fragment is subject to inactivation at each cell division. Several cell divisions are necessary before inactivation reaches its limit. While explaining quite well how continuous compaction enfolds, the hypothesis of compaction domains does not shed light on the mechanism of discontinuous compaction (Belyaeva and Zhimulev, 1991a; Belyaeva et al., 1993). Compaction is possibly continuous at the start, and then reactivation of some chromosome regions follows. It is also conceivable that the chromosome is arranged in loops and that compaction propagates from one loop base to another, skipping over the loops themselves. Finally, it is plausible that compaction can be initiated from many start points. The latter assumption is not consistent with the concept of involvement of euchromatin in a compaction domain of heterochromatin. To account for discountinous compaction, a model was proposed based on the concept of a statistical distribution of silencer protein (or compaction protein; CP) molecules around compaction initiation centers (CICs). It was assumed that the CICs are present in both hetero- and euchromatin, and different CP molecules interact not only with DNA but also with each other, forming a multimeric complex. When a certain level of DNA-protein binding is exceeded, heterochromatic domains are formed. A new PEV model of chromatin compaction based on the statistical distribution of the molecules of compaction proteins around the special centers along the chromosomes was proposed by Balasov and Makunin (1994,1996). This model requires assumptions such as the existence of (1) compaction initiation centers in both hetero- and euchromatin regions and (2) different compaction proteins having a statistical distribution around the compaction initiation center (Figure 161). According to the model the compaction is revealed only in pericentric heterochromatin (see Figure 1611) if the centers are separated by a region of euchromatin (limited by dots in Figure 1611) without such centers. In the case of the direct contact of the centers when euchromatin is removed (Figure 161II), compaction proteins from euchromatin form multilamellar complexes with proteins of heterochromatin. If the compaction ability of such complexes is weak, the compaction is revealed only in heterochromatin (see Figure 161IIa); if the ability is strong, this results in continuous compaction (Figure 16111~).Molecules of compaction protein can interact not only with centers of initiation, but also with each other to form a multimeric complex. The nonuniform distribution of these centers in euchromatin can result in discontinuous compaction (Figure 161IIb).


I

433

Thl

II

11 ...-......:'.. ..__. '_ -.: ....i.. +

:. . . .

+-:

:

b

:

.

.. . . . . . .

.

:

'

:

I

. ..

_ _ _ _ _ _ ._.-. .-. -. .-. .-.-. .-. -. .

.........

., ..

.

1 2 3

.

8

.

.:.,,.

. .

.

.. .

.. .

C

Figure 161. Types of compaction under PEV. (I) Normal chromosome. (11) Chromosome with rearrangement. 1-7, compaction centers in euchromatin; 8-10, compaction centers in heterochromatin; arrows, chromosome rearrangement breakpoints; a, continuous compaction of heterochromatin; b, discontinuous compaction; c, continuous compaction of eu- and heterochromatin; h', level of vizualization of compaction. Reprinted by permission from Balasov and Makunin (1994).

The question is, if a chromosomal rearrangement and a break in a heterochromatic domain it produces exist in all the cells, why are the genes in the rearrangement inactivated in only a part of the cells?The reasonable explanation appears to be Zuckerkandl's (1974), according to which the locking proteins in the cleaving syncytium of the embryo ate randomly distributed. Therefore, there would be more of them in some cells and less in others. Presumably, the genes transferred to heterochromatin would therefore be inactivated in some nuclei and not in others. This state of activation-inactivation can become stable, inherited in subsequent cell generations and giving rise to a pattern. One experimental approach to mosaicism would be through analysis of chromatin compaction in several chromosomal rearrangements in a single nucleus. Morphological study of chromosome regions in two inversions of a nucleus gave ambiguous results (Hartmann-Goldstein and Wargent, 1975) because the rearrangements were complex. In a stock containing two variegating rearrangements in In( J )y3p and T(2;3)SbV,inactivation of two genes in the same cell spread independently of each other (Bishop, 1992). In another study (Belyaevaet al., 1993), compaction was found to occur independently in each chromosome in the Dp(1 ;f) J337/T(f ;2)duPr7and T(2;4)ast"/Dp(I ;f)1337 rearrangement pairs. This, in turn, shows that cells may differ slightly in the amount of protein compacting molecules during the initiation of position effect in early embryogenesis. For this reason, the probability of euchromatin compaction becoming real is random in a cell. Two variegating rearrangements can display different HP1-binding be-

434

1. F. Zhimulev

havior in the same cell. This suggests that position effect variegation can be associated with stochastic behaviors of individual chromosomes, rather than with protein content in individual cells. In another study of the nine pairwise combinations between different variegating rearrangements, all but two or three exhibited interaction (Lloyd et al., 1997).

D. Maintenance of the inactivated state In the light of evidence presented in Section XI, it appears that establishment of the inactivated state throughout development is associated with at least: the blastoderm formation, the terminal stage of rapid mitoses; growth of imaginal discs in larvae; and the pupal stage. Despite dissimilarities,the stages have a common feature, being all associated with periods of increased mitotic activity. In all such cases, the gene is inactivated long before gene action becomes apparent, as a rule, in the adult. Thus the early achieved state of gene inactivation is correctly maintained through subsequent cell divisions, possibly by epigenetic factors (Baker, 196513; Khesin and Leibovitch, 1976). It would seem that the concept is consistent with the facts. However, Becker and Janning (1977) have shown that the addition of extra Ys heterochromatin or its removal from a cell chromosome in an established state of inactivation leads to change in variegation expression. This suggests that the state of inactivation can alter somewhat through each successive cell division. Theoretically, this appears plausible. In his first papers, Heitz (1932, 1933a) noted that heterochromatin undergoes decompaction during the cell cycle, although for a short span when locking proteins are redistributed (i.e., when the compaction state changes). In addition to Becker and Janning’s data, those of Beck and colleagues may be adduced to support this suggestion. Beck et al. (1979) have shown that the number of eye cells with the normally functioning w+ gene decreases with passage through cell divisions. However, these hypotheses neither refute nor elaborate the concept of the stable and long-continuing compaction of heterochromatin. Two levels of DNA packing may be envisioned: the usual chromosome organization allowing the genes to be normally activated or inactivated. Other proteins providing irreversible inactivation possibly superimpose at this level. Since DNA fragments from the euchromatic region that have undergone transitions from a compacted to a decompacted state and the reverse become profoundly inactivated in normal chromosomes, fixation of the locking proteins at the second level is not related to specificity of DNA sequences. In all probability, the histones on which the compacting proteins bind are modified in the inactivated regions. However, all this is in the realm of speculation. There are several examples of stably and heritably ,inactive transcription in cell generations. In the yeast Saccharomyces cereuisiue, the mating genes HML and HMR


435

are not expressed even though all the signals for expression are present at the loci. This kind of repression is known as silencing (see Shore, 1995; Holmes and Broach, 1996). Repression of these loci is an active process since mutation of any one of four trans-acting regulators, products of the SIR (silence information regulator) genes, results in derepression of both loci. Specific silencers are required for the stable inheritance of a repressed state (Holmes and Broach, 1996). In Drosophila the genes of the Polycomb-group (Pc-G) are responsible for maintaining the inactive expression state of homeotic genes. Mutations in at least fifteen genes have been found to result in complete derepression of homeotic genes (see Eissenberg et al., 1995 for review). These genes are collectively called the Polycomb group. They act through specific cis-regulatory DNA elements (PC-G Response Elements, PRE). Multimeric complexes containing the Pc-G proteins bind the PREs resulting in deep target gene inactivation (Zink and Paro, 1995).

XVIII. HETEROCHROMATIZATION OF CHROMOSOME REGIONS AND REGULATION OF GENE ACTIVITY A. Properties of heterochromatized regions of chromosomes Brown (1966)has justly noted that “the problem of heterochromatin is one of the most difficult and diffuse in modem biology” (p. 424). In fact, it is Hsu’s view (1971) that the term “heterochromatin” is applied to describe different events. Prokofyeva-Belgovskaya ( 1941) holds the view that “heterochromatin and euchromatin are not special substances, any segment of the chromosome can acquire a ‘euchromatic’ or a ‘heterochromatic’ state” (p. 34). White (1948) defines heterochromatin as any region of the chromosome that becomes heteropycnotic at some stages of the cell cycle. Spofford ( 1976)supposes that a chromosome region can become heterochromatic in one cells group and euchromatic in another during early embryonic development, for example. Some investigators believe that heterochromatin does not exist as a specific structure of the chromosome; rather, it is a specific state regarded as heterochromatin (Cooper, 1959; Commoner, 1964). These ideas were reasonably elaborated by Brown (1966), who stated that heterochromatin is the visible expression of gene inactivation during development and evolution. To make a distinction between heterochromatin as a temporary state of the inactivated chromosomes or their regions and constant heterochromatization (heteropycnosis), Brown introduced the concept of “facultative” heterochromatin for the former and “structural” (constitutive) for the latter. However, Prokofyeva-Belgovskaya (1977a, 1986) doubted that the hypothesis of facultative heterochromatin was correct because advances in molecular biology made it obvious that heterochromatin and euchromatin are, in principle, different with respect to the molecular organization of the DNA forming

436

1.

F. Zhimulev

these regions; facultative heterochromatin is not heterochromatin, but rather condensed and, consequently, genetically inactivated euchromatin. Zhimulev et al. (1982),who have suggested that the regions of intercalary heterochromatin of polytene chromosomes consist of tandemly repetitive DNA segments, support the concept of the specific structural organization of heterochromatic DNA. To resolve ambiguities, these following questions must be answered:

1. Is there a correspondence between the set of morphological features of the heterochromatic regions of the chromosomes and a particular genetic constitution of DNA? 2. What might be the mechanism behind the conversion of a chromosome segment into heterochromatin? 3. What might be the genetic consequences of this conversion? Analysis can be carried out in such a way that the main property of heterochromatin, compaction as defined by Heitz, is taken as a reference and all the other properties of the compacted segments of the genome are then described. It is well to keep in mind that the various types of heterochromatin have been studied to different degrees and that information is, as yet, incomplete. The regions of pericentromeric heterochromatin of the mitotic chromosomes have been most thoroughly studied. From a survey of Table 33, it is evident that the regions possess virtually all the features of heterochromatin. However, there are at least three segments included in heterochromatin that considerably differ functionally at the level of their genetic organization: satellite DNA, for the most part, does not encode genetic functions; the second segment in the Dosophila genome consists of a block of the repetitive 18s and 28s rRNAs genes; and, finally, the mendelian genes were found in pericentromeric heterochromatin. The differences in the genetic organization of intercalary heterochromatin of polytene chromosomes are still greater. The intercalary heterochromatic regions generally possess the same properties as centromeric heterochromatin (see Table 33). The nature of the included DNA is known for certain regions of intercalary heterochromatin showing these features: a block of the repetitive histone genes, the ribosomal 5s and 18+288 RNAs, the polypyrimidine repeat, the poly(AT) repeat, the bithorax gene active during early embryonic development, and the 11A region controlling chromosome pairing at meiosis. The morphological appearance of the region encompassed by the telomeric repeat (HeT repeat) is peculiar. In Dosophila, whose telomeric regions in the polytene chromosomeshave been studied most, these regions consist, as a rule, of diffuse, faintly staining material that Lefevre (1976) has even called puffs. Hence it may be assumed that these regions may be compacted more loosely than the others, or perhaps a specific compacting protein (HP1 in those regions or in 31AF) imparts an unusual appearance to the telomere (see Table 33).


437

The morphology of the regions of euchromatin drastically changes when juxtaposed to intercalary heterochromatin (the regions are position affected). Regions not possessing any of the heterochromatic features under normal conditions acquire them when brought into a state of compaction (see Table 33). Thus various types of DNAs (satellites, middle repeats, the repetitive and the unique genes) acquire the properties of heterochromatin as the result of compaction caused by genetic inactivation. Other examples are known in which usually euchromatic segments of the genome acquire heterochromatic features:

. In coccids, a single complete set of the chromosomes (paternal) is compacted during early development in the male and remains so throughout development. The maternal (euchromatic) and paternal (heterochromatic) chromosome sets differ considerably in compaction degree, and do not pair during meiosis. The eu- and heterochromatic sets pass toward the opposite poles during the second meiotic division. The sperm is formed only from the euchromatic chromosome set. The heterochromatized chromosomes slowly degenerate in the meantime (Schrader, 1929; Brown, 1966,1969; Nur, 1967a). According to Schrader’s ( 1929) hypothesis, the heterochromatic set is genetically inert; that is, the genes transmitted by the father to his son are not expressed. In fact, the compact heterochromatized blocks in the interphase nuclei of the mealy bug Psewlococcous obscurus did not label with [3H]uridine (Berlowitz, 1965). The heterochromatized chromosomes are not identified in the interphase nuclei of some larval tissues of coccid larvae. Nur (196713, 1990) suggested that the paternal set is euchromatized in these tissues, thereby providing a unique opportunity to test the hypothesized correlation between compaction and gene inactivation. Paternal x-irradiation, even with very high doses, had no effect on progeny, presumably because the paternal set is not expressed; marked dominant changes in phenotype arise in tissues where the set is euchromatized (e.g., in malpighian tubules). This is a clear-cut example of the relation between the state of compaction and genomic expression. 2. In 1949, a structure consisting of compact chromatin was revealed in the nuclei of nerve cells of female cats that takes the name from its discoverer (Barr and Bertram, 1949). Later, it was found that one of the X chromosomes is inactivated in mammalian females, no matter if the Ban body is present or not. The whole chromosome becomes inactive to compensate for differences in the number of the X chromosomes in males and females. This is how the X chromosome is dosage compensated in mammals (Lyon, 1961, 1962; Verma and Babu, 1989). To explain these facts, Brown introduced the concept of constitutive and facultative heterochromatin. 3. Similar behavior was observed for an 11,000-kbDNA fragment containing

Table 33. Properties of Heterochromatinized Regions the Chromosomes Differing in DNA Composition (according to data of previous sections).

Types of chromosome regions, DNA

Properties of heterochromatinized regiono

4

5

6

7

8

+

+

+

+

+

+

+

1

2

3

Mitotic Chromosomes Pencentric heterochromatin (satellite DNA)

+

+

The 18s and 28s rRNA

+

9

12

13

+

+

+

+

+

+

+

10

11

genes

B chromosomes Chromosomes restricted by germlines Polytene Chromosomes Pericentric a-and p-heterochromatin Intercalary heterochromatin, including: Histone genes 5s rRNA genes

18s and 28s rRNA genes The B x C gene Polypyrimidine repeat, the 21D region

+ + +

+

+

+

+

+

+

+

+

+ + + + +

+

-t

-

+ +

-

+ +

+

+

+

+

+ + + +

+

+

i

+

+

?

+

+

+

-

Telomeric heterochromatin, the HeT repeat

?

+

-

The 3 1A-F region

?

-

-

B chromosomes

+

+

Poly (AT) repeat, the 81F region

+

Chromosomes restricted by the germlines

+

Euchromatic Regions under Position Effect Variegation Compact fragments of euchromatin

+

Inactivated Ban body

+

+

+

+ + t

+

+

+

+ +

+

+

Inactivated Paternal Genome in Coccids Compact inactivated chromosomes

+

Transposons in Transgenic Mammals The tandemly repeated p-globin gene of 11,000kb

+

?

+

+

+

X Chromosome in Mammals

+

+ +

+ +

+ +

+

"1, compaction; 2, late replication; 3, C-banding; 4,underreplication;5, variation in amount; 6, ectopic contacts; 7, chromosomal rearrangements; 8, contact with membrane; 9, location of HPI; 10, induction of position effect variegation; 11, ability to compact by modifiers of position effect variegation; 12, modification of position effect variegaion; 13,compaction during early embryogenesis.

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tandem repeats of the P-globin gene and DNA of the pBR plasmid transformed in the mouse genome. In cells of the nonerythroid series (highly differentiated neurons) not expressing the P-globin gene, transposon DNA assumed characteristic features of heterochromatin: it is converted into a block of structurally condensed material closely associated with the nuclear membrane, (ectopically) tightly pairing with the regions of intercalary heterochromatin in the nucleolus (Manuelidis, 1991). 4. Two closely related species of mosquitoes, Aedes aegypti and A. mascurensis, were found to differ in the C-banding pattern of the sex chromosome pair. In A. aegypti, a centromeric C' block and another interstitial block are seen in the sex chromosomes of females. In A. mascurensis, only the centromeric block appears. The interstitial C' block disappears from hybrids of both sexes. The block reappears in backcross progeny of the F1 hybrids to A. aegypti (Motara and Rai, 1977). 5. Chromosomes of two types were identified in representatives of the three families of Dptera:the chromosomes restricted to the germline (E chromosomes) and eliminated from the somatic cells during the early cleavage stages; and the chromosomes of somatic cells (S chromosomes). As a rule in representatives of these families of Dipterans in germ line cells namely S chromosomes are in totally heteropicnotic condition (White, 1946, 1973; Matuszewski, 1962). In Wachtliella persicu&, S and E chromosomes are differing beginning with oogonial stage. S chromosomes in interphase nuclei are heteropicnotic, but E chromosomes are decondensed and look like interphase chromosomes (Kunz et ul., 1970). In Muyetiola destructor, the set of somatic chromosomes received from the female is compacted during spermatogenesis, and it entirely stains for C- heterochromatin. Only the centromeric regions of these chromosomes remain C-heterochromatized in the somatic cells of cerebral ganglia of larvae (Stuart and Hatchett, 1988a,b).Probably E chromosomes are necessary for development of egg and spermatozoids. During oocyte growth of Wachtliella E chromosomes are decondensed and incorporate [3H]uridine.S chromosomes are not active and compact (Kunz, 1970; Kunz et af., 1970). The facts just stated prove that transition from a euchromatic to a heterochromatic state and vice versa is possible for various DNA types with respect to both functional genetic features (various types of sequences) and size (chromosome segments, single chromosomes, transposons, whole genomes).

B. Supercompaction of heterochromatic regions as a mechanism of DNA inactivation in development Compaction of chromosome material is the basis underlying heterochromatization. Which of the DNA segments are compacted? Examination of the data in Table 33 brings to attention mainly four types of DNA sequences:


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1. Inactivation of the noncoding DNA segments of satellite type. It is pertinent to recall Swanson’s (1957) view that the major function of heterochromatin is a general one, concerned with cell division, cellular growth, and embryonic differentiation. In fact, the available evidence indicates that the DNA amount in satellites characteristic of a species remains relatively invariant only in germline cells; the number of repetitive copies can vary in somatic cells. In terms of ontogenesis, this means that a certain amount of satellites are possibly required for some processes of gametogenesis, egg maturation, and the first cleavage divisions to occur. Satellite DNAs must be securely excluded from function at all the other developmental stages, and heterochromatization (compaction) is the mechanism producing inactivation. Regions of intercalary heterochromatin (1 1A, for example) controlling pairing in the chromosome during meiosis may be most likely referred to DNA of this kind. 2. Repetitive genes. Genes such as the histone, 5S, 18S, and 28s rRNA genes are present as many copies in the genome. However, gene copies are, for the most part, excluded from transcription in somatic cells (e.g., in salivary glands). The regions of the rRNA 5s genes and the histone genes in polytene chromosomes consist of blocks of compactly organized chromatin material. Neither puffs nor noticeable incorporation of [3H]uridine is observed in these regions. Since it is hard to see how the very important “housekeeping” genes of the cell can be inactivated, apparently a small portion of the copies, presumably located at the edges of the clusters where micropuffs are formed, must function. The other copies are inactivated, being in a compacted (heterochromatic) state. It is not clear whether all the gene copies are used in ontogenesis. Possibly they are used during maturation of the oocyte, when the gene products are synthesized in large amounts. Great quantities of compact heterochromatin are also detected in the nucleolar organizer, where the 18s and 28s rRNA genes reside. 3. The unique genes for early development. The bithorax gene is an early acting gene and is later inactivated. The band where this gene is located is converted into a region of intercalary heterochromatin. 4. Euchromatic fragments of the chromosomes, single chromosomes, or whole genomes. The previously presented facts (see Table 30 in Section XIII), on heterochromatization of the euchromatic mammalian X chromosome, or of the whole coccid genome, provide evidence that compaction is a universal mechanism of genetic inactivation of an extensive section of the genome at appropriate stages of ontogenesis. The mechanism starts to operate at the stages of embryonic development.

Compaction and the associated genetic inactivation resulting from position effect may be regarded as similar events. As Becker ( 1 9 5 6 ~put ) the matter,

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“position effect is a system of somatic control over gene functioning, which is similar to cell differentiation in many features” (p. 149). Once compacted, the regions transmit the state through successive cell divisions. It may be suggested that heterochromatin assumes a compact state in early embryogenesis, when the genome starts to be differentially active and all the DNA sequences not required functionally are profoundly repressed. There is reason to believe that specific compacting proteins appear at that time. Thus the formation of heterochromatic structures is an ontogenetic pattern safeguarding cells through profound inactivation from the expression of redundant genetic material. Brown took the view (1966) that heterochromatin and the mitotic chromosomes are compacted to the same degree. The incorrectness of Brown’s view was shown by Prokofyeva-Belgovskaya(19861, who indicated that, in such a case, the whole chromosome set would become heterochromatized from late prophase; however, the fact that even the metaphase chromosomesdifferentiate into C-negative and C-positive regions is consistently revealed by conventional C-banding procedures. Presumably, a heavier compaction of chromatin in the heterochromatized chromosome regions is provided by modification of histones (Blumenfeld et al., 1978a,b) and binding of specific nonhistone proteins (Sutton, 1972; James et al., 1989). It is of interest that the proteins that bind to DNA and are involved in inactivation of the homeotic genes, and also the compacting proteins of the HP1 type, are all similar in primary structure (Paro, 1990). The following functional differences between the organization of compacted euchromatin and heterochromatin may support the idea that a supercompaction of chromatin due to a different protein composition occurs in heterochromatin:

1. The genes in the inactivated mammalian X chromosome, in the coccid genomes, and in the decompacted segments of euchromatin are not reactivated by variegation inducers activating the genes in the nonheterochromatized homologous chromosomes of the same nucleus. 2. DNA replication is naturally hindered in the supercompacted material, and, for this reason, the heterochromatized regions are late replicating in all cases. The simply compacted regions of the chromosomes complete replicating earlier. 3. Polytenization occurring in the supercompacted regions of intercalary and pericentromeric heterochromatin is incomplete because of late replication. 4. Presumably, some feature, such as ectopic pairing when, as a rule, nonhomologous DNA of different heterochromatized regions makes contact, is provided by the molecules of compacting proteins in these regions.


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5 . Possibly, the continuous contacts of heterochromatin with the nuclear envelope are also provided by affinity of the compactor proteins for the proteins of the internal membrane. How long will the problem of heterochromatin be still shrouded in mystery (for perspectives, see Spradling and Karpen, 1990)?It may be appropriate to recapitulate the general conclusion based on this analysis of data from Heitz (1993a,b). Heterochromatin (or the heterochromatized state) may be viewed as representing a portion of the genome that is in a state different from that the remainder of the genome and profoundly inactivated because it is supercompacted. It may be noted that all the heterochromatized structures follow the germline pathway of development; that is, they are fully represented in meiosis and at the early cleavage divisions. Many of the genomic segments are underrepresented in somatic cells. Consequently, heterochromatin is absolutely necessary for germ cells to function, and it may be lost from somatic cells. This difference between cells of the two lines may provide evidence for the fundamental importance of heterochromatin in the sexual process.

Ac know Ie d0ments The writing of this book was a time- and labor-consuming endeavor not only for the author, but also for the many colleagues involved. It is the author’s pleasure to acknowledge his gratitude to them all. Every page of the monograph has been written after in-depth discussions with E. S. Belyaeva. T h e thought-provoking discussions with the late A. A. Prokofyeva-Belgovskayaat a time when the concept of intercalary heterochromatin was coming into being are unforgettable. N. P. Dubinin and I. B. Panshin read and provided comments on the section dealing with position effects. Invaluable assistance in compilation of the literature and preparation of the manuscript was provided by V. E. Graphodatskaya, E. A. Dolbak, E. B. Kokoza, N. P. Korol’kova, D. E. Koryakov, N. Yu. Kuznetsova, 1. V. Makunin, N. I. Mal’ceva, V. A. Prasolov, G. Richards, I. P. Selivanova, V. K. Vasilyeva, and V. V. Volkovintser. Original illustrations were gifts of D. Bedo, E S. Valeyeva, M. Gatti, A. K. Grishanin, C. D. K. Castritsis, I. E. Kerkis, P. V. Michailova, A. P. Akifjev. M. L. Pardue, P. Roberts, M. Hatsumi, K. Hagele, S. Henikoff, J. S. Yoon, S. C. R. Elgin, L. A. Chubareva, and N. A. Petrova. Previously unpublished photographs and materials have also been kindly supplied by R. B. Vagapova, E. B. Kokoza, and V. F. Semeshin. Much material in the form of reprints and preprints has been obtained from A. P. Akifjev, M. Ashburner, H. Biessmann, C. Craig, J. Eissenberg, S. C. R. Elgin, J. G . Gall, S. M. Gershenzon, R. Hill, A. Hilliker, S. C. Lakhotia, J.-A. Lepesant, G. L. G. Miklos, P. V. Michailova, M.-L. Pardue, M. Steinemann, and L. Wallrath. T h e author expresses his deep gratitude to all of the geneticists mentioned above. I am especially grateful to A. N. Fadeeva for translating the manuscript into English. This work was supported by grants of the Frontier Program in Genetics of the Russian Federation, the Russian Fund for Fundamental Research (Grant 95-04-12695), the Soros Fund, and the INTAS program.

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abo gene, X chromosome, 59-60 Accessory chromosomes, see B chromosomes Acridine orange stain, intercalary heterochromatin identification, 143-145 Acriflavine-nitrate, intercalary heterochromatin identification, 146-149 Allopurinol, heterochromatin gene expression modification, 344 Ammoniacal silver stain, intercalary heterochromatin identification, 150 Anopheles chromosomal rearrangement localization, 29 late replication study, 32 Anoxia, heterochromatin compaction, 13 Antibodies, positional effect gene expression inactivation, 329 Apterygotan insects, see also specificspecies intercalary a-heterochromatin, 139-142 Ascaris megalocephala, chromatin diminution, 7W1 Autosomes heterochromatin gene expression modification, 339-341 mitotic heterochromatin genetic content,

60-65

B chromosomes characteristics, 282-304 fitness association, 302-303 heterochromatin association, 300-301 number, 295-297 occurrence, 288-294 pairing, 299-300 properties, 438 supernumerary effects, 301-304 designation symbols, 282 heterochromatin detection, 6 Belgovskaya-Prokofyeva, heterochromatization theory, 395-399 Bilobella auranriaca, intercalary a-heterochromatin, 141

Breaks, see Chromosomes, breaks 5-Bromodeoxyuridine, heterochromatin gene expression modification, 342-343 bw' gene, position effects, 371-376

Cell cycle, dividing cell heterochromatin morphology, 5-43 chromosomal rearrangement localization, 29-30 compaction degree, 5-14 anoxia effects, 13 distamycin A effects, 11 genetic control, 13 Hoechst 33258 effects, 9-1 1 mechanisms, 13-14 overview, 5-6 permanence, 6-9 radiation effects, 12-13 temperature effects, 11-13 developing animal heterochromatin formation, 3543,440-443 differential staining, 14-24 C-staining, 14-15 enzymatic chromosome digestion, 24 heterochrornatin heterogeneity, 24 H-staining, 15-2 1 N-staining, 21-23 Q-staining, 15-21 heterochromatin quantity variation, 33-35 late replication, 30-33 pairing, 24-28 interphase chromocenter formation, 8, 24-25 meiosis, 28 mitosis, 26-28 Centromeres heterochromatin gene expression modification, 339-341 pericentromeric heterochromatin inactivation, 378-391 heterochromatin amount effects, 381-382

557

558

Index

inactivation strength, 383-385 rearrangement breaks, 385-390 revertant strain cytologicalanalysis, 379-381,389-390 polytene chromosome heterochromatin, 89-133 cytogenetics, 125-134 morphology, 90-99 quantity variations, 3639,109-124 structural components, 99-109 DNA, 99-105 proteins, 105-107 structural changes, 107-109 Chironomw Cla-element identification, 106, 2 19-220 heterochromatin puff induction, 107-109 intercalary a-heterochromatin. 138 Chromatids,position effect, pompon formation, 421-422 Chromatin diminution, 78-89 Ascaris megabcephala, 78-81 Cyclops, 81-85 dipteran insects, 85-87 infusoria, 88-89 physiological significance,89 position effect compaction, 400-413,429 characteristics, 402-404 discontinuous compaction, 406-407,410 formation, 404406,410 historical perspectives, 400-401 late-replication,410 occurrence frequencies, 41 1 4 1 3 parental effects, 408-409 temperature effects, 407-408 state changes, 422-428 Chromocenter centromeric heterochromatin DNA hybridization, 99-105 formation interphase heterochromatin pairing, 8, 24-25 meiotic pairing, 28 mitotic pairing, 26-28 polytene sets, 95, 98 morphologicalanalysis,heterochromatin cy. togenetics, 129-132 Chromosomes B chromosomes,282-304 designation symbols, 282

fitness association, 302-303 heterochromatin association, 300-301 heterochromatin detection, 6 number, 295-297 occurrence, 288-294 pairing, 299-300 properties, 436 supernumeraryeffects, 301-304 breaks, intercalary heterochromatin identification, 150-166 constrictions, 150, 158 fissures, 158 frequency, 161-166 shift, 158 elimination, dipteran insects, 84-86 germline cell heterochromatin, 304-306 position effect, inactivation spreading, 311, 329-330 rearrangement gene expression inactivation initiation, 429434 position effect,310-311,329-330 vital genes, 316-318 intercalary heterochromatin identification, 198-206 parental genotype effects, 339,341-342 sex-determiningfactors, 306 ci+ gene Dubunin effect, 357-362 position effects, 371 Cla-element, identification, 105, 219-220 Colchicine, heterochromatin gene expression modification, 343 Compaction dividing cell heterochromatin anoxia effects, 13 distamycin A effects, 11 genetic control, 13 Hoechst 33258 effects, 9-1 1 mechanisms, 13-14 overview, 5-6 permanence, 6-9 radiation effects, 12-13 supercompaction, 440-443 temperature effects, 11-13 position effect, 400-413, 429 characteristics,402-404 discontinuous compaction, 406-407,410 formation, 404-406, 410 historical perspectives, 4 W 0 1 late-replication,4 10

Index occurrence frequencies, 41 1 4 13 parental effects, 408409 temperature effects, 407-408 Crossing over, frequency, 7 6 7 7 C-staining differential heterochromatin staining, 14-1 5 intercalary heterochromatin identification, 141 Cyclops, chromatin diminution, 81-84 Cytological repeats, intercalary heterochromatin identification, 225

Development, see Embryonic development Diminution, chromatin loss, 78-89 Ascaris megalocephala, 78-81 Cyclops, 81-85 dipteran insects, 85-87 infusoria, 88-89 physiological significance, 89 Dipteran insects, see also specificspecies band characteristics, 234-238 B chromosomes number, 295-297 occurrence, 288-294 centromeric heterochromatin amount variation mitotic chromosomes, 36-39 polytene chromosomes, 121-122 DNA structure, 99 chromatin diminution, 84-86 chromosomal break characterization, 151-1 5 7 ectopic pairing nucleolar material, 178-1 79 polytene chromosomes, 171-176, 266-276 germline limited chromosome occurrence, 87 intercalary a-heterochromatin, 135-138 polytene chromosomes centromeric heterochromatin amount variation, 121-122 ectopic pairing, 171-176, 266-276 Q+-bandoccurrence, 147-148 terminal deletion occurrence, 243-244 Distamycin A, heterochromatin compaction, 1 1 Distortion factor, autosomes, 61-65 DNA centromeric heterochromatin structure, 98-105

559

heterochromatin gene expression modification, 341 repetitive sequences, 43-54 composition, 53-54 location, 45-46, 51-52 in situ hybridization, 3,51-53,99-101 slipping, 53 types, 4 7 4 9 underrepresentation intercalary heterochromatin identification, ectopic pairing, 189-193 position effect, 414-421 telomeric heterochromatin, 252-255, 280 Drosophih formella, heterochromatin amount variation, 35 Drosophila hydei position effect clonal inactivation, 326-327 eye mosaics, 322 Y chromosome, 68-72 Drosophila imetensis, intercalary ct.heterochromatin, 138-139 Drosophilu m e h c a s t e r chromosomal rearrangement frequency, 199-201 localization, 29-30 compound eye structure, 3 17 differential heterochromatin staining, Qstaining, 18, 20 DNA satellite repeating sequences, 4 7 4 9 intercalary heterochromatin features, 228 late replication study, 31-32 mitotic pairing, 26-28 mobile element location, 209-216 polytene chromosome Q+-bandoccurrence, 148 position effect bw' locus, 371-376 ci' locus, 371 clonal inactivation, 326-327,332-333 eye mosaics, 322-325 In(2LRj40d locus, 376 light locus, 362-365 rolled+ gene, 362-365 variegation control genes, 3 13-3 15 telomeric heterochromatin, DNA underrepresentation, 280 transposon genes, 318-3 19 X chromosome, 59-60, 102-104,222 Y chromosome, 65-68, 102

560

Index

Drosophila mirunda, neo-Y chromosome degeneration, 74-76 Drosophila preudoobscura, heterochromatin amount variation, 33-34 Drosophila virilis chromosomal rearrangement parental effects, 342 differential heterochromatin staining, Qstaining, 17, 21 late replication study, 32 position effect clonal inactivation, 326-327 peach+ locus, 366-367 Dubunin effect, ci' gene dominance, 357-362

Ectopic pairing characteristics, 134 intercalary heterochromatin identification chromosomalrearrangement frequency, 204 DNA underreplication, 189-1 93 formation mechanisms, 183-184 nucleolar contacts, 168-169, 179 nucleoprotein association, 185 overview, 166-169 polytene chromosomes, 171-1 76 properties, 168, 180-182 telomeric heterochromatin, 266-279 characterization,268-275 properties, 267, 276-279 Embryonic development gene inactivation, 355-357 heterochromatin association formation, 35-43 supercompaction, 440-443 Enzymes, differential heterochromatin staining, chromosomedigestion, 24 Escherichia coli, differential heterochromatin staining, Q-staining, 17 Ethidium bromide, differential heterochromatin staining, chromosome digestion, 24 Euchromatin definition, 2-3 he terochromatization, 437-440

Fertility factors, position effects, 368

Gene expression inactivation embryonic development, 355-357, 440-443 historical perspectives, 376-378 initiation, 429-433 maintenance, 434435 position effect biochemically identified loci, 318 chromosomal rearrangements, 3 10-3 11, 329-330 genes affected, 3 12-3 19 inactivation levels, 327-329 phenotype development control genes, 312-316 position effect variegation characteristics, 309-312 spreading, 31 1,330-334 transposon genes, 318-319 variegation inactivation, 319-327 vital genes, 316-318 modification conditions, 334-355 chemical modifiers, 342-345 genetic modifiers, 345-355 chromatin formation control, 351, 351-355 dose dependence, 34%350 identification, 346-348 molecular-genetic characteristics, 352-353 P-element-mediatedmutagenesis, 346-347 heterochromatin, 335-341 autosomes, 339-341 exogenous DNA, 341 X chromosome, 339-341 Y chromosome, 335-339 histone genes, 345 parental effects, 339,341-342 temperature, 325,334-335,342 position effect, 306-309 Glyptotendipes barbipes, heterochromatin puff induction, 106

Heterochromatin B chromosome association, 300-301 centromeric heterochromatin in polytene chromosomes,90-134

index cytogenetics, 124-133 morphology, 90-99 quantity variations, 109-124 structural components, 99-109 DNA, 99-105 proteins, 105-107 structural changes, 107-109 characteristics, 434-440 definition, 2,5 gene expression modification, 335-341 autosomes, 339-341 exogenous DNA, 341 X chromosome, 339-341 Y chromosome, 335-339 germline cell chromosomes,304-306 a-heterochromatin, 136-142 apterygotan insects, 139-142 Bilobela aurantiaca, 141 centromeric heterochromatin in poiytene chromosomes, 124-134 Chironomus, 139 dipteran insects, 136-139 Drosophila imeretensis, 138-139 infusoria, 142 Neanura monticola, 141-142 Phryne cincta, 136-138 f3-heterochromatin cytogenetics, centromeric heterochromatin in polytene chromosomes, 125-134 inactivation initiation, 429434 intercalary heterochromatin, 391-394 maintenance, 433434 pericentromericheterochromatin, 378-391 heterochromatin amount effects, 38 1-382 inactivation strength, 383-385 rearrangement breaks, 385-390 revertant strain cytological analysis, 379-381,389-390 Prokofyeva-Belgovskaya heterochromatization theory, 395-399 telomeric heterochromatin, 394-395 intercalary heterochromatin differential staining identification methods, 142-150 acridine orange stain, 146150 acritlavine-nitrate, 150 ammoniacal silver stain, 150

56 1 C-staining, 143 H-staining, 143-146 methyl green stain, 150 pyronine, 150 Q-staining, 143-146 a-heterochromatin, 136-1 42 apterygotan insects, 139-142 BilobeUa aurantiaca, 141-142 Chironomw, 139 dipteran insects, 136-139 Drosophila imeretensis, 138-139 infusoria, 142 Neanura monticola, 141-142 Phryne cincta, 136-138 morphological identification methods, 150-225 chromosomal breaks, 150-166 chromosomal rearrangements, 198-206 cytological repeats, 225 ectopic pairing, 166-193 gene repeat sequences, 208-209 late replication, 193-198 minute mutations, 224-225 mobile genome elements, 209-216 nuclear membrane contacts, 224 repetitive sequences, 207-224 somatic pairing, 206207 tandem repeats, 216-224 tRNA genes, 207-208 overview, 133-135 property manifestation correlations, 225-232 quantity variations, 232-238 mitotic genetic content, 54-78 autosomes, 60-65 crossing over frequency, 7 6 7 7 neo-Y chromosome degeneration, 74-76 overview, 54-56 X chromosome, 57-60 Y chromosome, 65-74 Drosophila hydei, 68-72 Drosophila melanogaster, 6 5 4 8 protein accumulation, 73-74 in situ hybridization,66-72 Ste gene, 6 6 4 8 morphology, 5 4 3 chromosomal rearrangement localization, 29-30 compaction degree, 5-14 anoxia effects, 13

562

Index distamycin A effects, 11 genetic control, 13 Hoechst 33258 effects, 9-1 1 mechanisms, 13-1 4 overview, 5-6 permanence, 5 4 radiation effects, 12-13 temperature effects, 11-13 developinganimal heterochromatin formation, 3543 supercompaction,440-443 differential staining, 14-24 C-staining, 14-15 enzymatic chromosome digestion, 24 heterochromatin heterogeneity, 24 H-staining, 15-21 N-staining, 21-23 Q-staining, 15-21 late replication, 30-33 pairing, 24-28 interphase chromocenter formation, 8,

24-2s meiosis, 28 mitosis, 26-28 quantity variation, 33-35 research, historical perspectives, 2-5 sex-determining factors, 306 telomeric heterochromatin characteristics, 259-282 change manifestations, 281 differential staining, 279-280 ectopic pairing, 266279 heterochromatin proteins, 280 overview, 238-248,259-266 repeats, 248-259 Histone genes heterochromatin gene expression modification, 345 intercalary heterochromatin identification, tandem repeats, 219 Historical perspectives chromatin compaction, position effect,

400-401 heterochromatin research, 2-5 definitions, 2-3 position effect, 4-5 positional gene inactivation, 376-378 Hoechst 33258 differentialheterochromatin staining, 15-21 heterochromatin compaction effects, 9-1 1

intercalary heterochromatin identification,

141-143

Inactivation, see Gene expression, inactivation Infusoria chromatin diminution, 86-88 ectopic pairing, polytene chromosomes, 176 intercalary a-heterochromatin, 142 Jn(2LR)40d locus, position effects, 376 Insects, see specifictypes In situ hybridization DNA structure analysis centromeric heterochromatin, 99-1 01 repetitive sequence Localization, 3, 51-53 underrepresentation, 41 7-418 Y chromosome, 6672,102 Intercalary heterochromatin differential staining identification methods,

142-150 acridine orange stain, 146-150 acriflavine-nitrate, 150 ammoniacal silver stain, 150 C-staining, 143 H-staining, 143-146 methyl green stain, 150 pyronine, 150 Q-staining, 143-146 a-heterochromatin, 136-142 apterygotan insects, 139-142 BilobeUu aurantiaca, 141-142 Chironomus, 139 dipteran insects, 136-139 Drosophila inmetensis, 138-139 infusoria, 142 Neanura monticola, 141-142 Phrynecincra, 136-138 heterochromatin inactivation, 391-394 morphological identification methods, 150-225 chromosomal breaks, 150-166 constrictions, 150, 158 fissures, 158 frequency, 161-166 shift, 158 chromosomal rearrangements, 198-206 cytologicalrepeats, 225 ectopic pairing, 166-193 gene repeat sequences, 208-209

Index late replication, 193-198 minute mutations, 224-225 mobile genome elements, 209-216 nuclear membrane contaccs, 224 repetitive sequences, 207-224 somatic pairing, 206-207 tandem repeats, 216-224 tRNA genes, 207-208 overview, 133-135 property manifestation correlations,

225-232 quantity variations, 232-238 Interphase, chromocenter formation, heterochromatin pairing, 8, 24-25

Late replication dividing cell heterochromatin morphology, 30-33 intercalary heterochromatin identification, 193-198 light locus, position effect, 362-365

Male fertility factors, position effects, 368 Meiosis, see also specificphases heterochromatin pairing, 28 Melanoplus diferenhhlis, late replication study,

30 Methyl green stain, intercalary heterochromatin identification, 146 Minute mutations, see also B chromosomes intercalary heterochromatin identification,

224-225 Mitotic heterochromatin genetic content, 55-78 autosomes, 61-66 crossing over frequency, 77-78 neo-Y chromosome degeneration, 74-76 overview, 55-57 X chromosome, 57-61 Y chromosome, 65-74 heterochromatinized region properties, 436 pairing, 26-28 Mosaicism clonal inactivation, 323-328 hereditary change association, 4 specificity, 320-322

563

Neanura monticola, intercalary a-heterochromatin, 140-141 Neo-Y chromosome, mitotic heterochromatin degeneration, 74-76 N-staining centromeric heterochromatin nonhistone protein analysis, 105-106 differential heterochromatin staining, 2 1-23 Nuclear membrane, intercalary heterochromatin identification, contact points, 224 Nucleolar organizer heterochromatin cytogenetics, 124-133 intercalary heterochromatin identification, ectopic pairing, 166, 178-179 position effects, 368-370 Nucleotides, tandem repeats, intercalary heterochromatin identification dinucleotide repeats, 222-223 mononucleotide repeats, 222-223 nucleotide sequences, 246 poly(A) nucleotide complexes, 220-222

Pairing B chromosome homology, 299-300 dividing cell heterochromatin morphology, 24-28 interphase chromocenter formation, 8, 24-25 meiosis, 28 mitosis, 2 6 2 8 ectopic strands characteristics, 134 intercalary heterochromatin identification, 166-193 DNA underreplication, 189-193 formation mechanisms, 183-184 nucleolar contacts, 166, 178-1 79 nucleoprotein association, 184-185 polytene chromosomes, 171-1 76 properties, 168-170, 180-182 telomeric heterochromatin, 266-279 characterization, 268-275 properties, 267, 276-279 intercalary heterochromatin identification ectopic pairing, 166-193 somatic pairing, 206-207 Parascaris uniudens, chromatin diminution, 78-81

564 Parental effects chromatin compaction, 408-409 gene expression modification, 339,341-342 peach+ locus, position effect, 366-367 P-element gene expression modification, mutagenesis, 346-347 position effect, 308-309,311 Phenotype development, position effect, 3 12-3 16 Phryne cincta, intercalary a-heterochromatin, 135-1 37 Polypyrimidines, intercalary heterochromatin repetitive sequences, 219 Polytene chromosomes centromeric heterochromatin, 90-134 cytogenetics, 125-134 morphology, 90-99 quantity variations, 109-125 structural components, 99-109 DNA, 99-105 proteins, 105-107 structural changes, 107-109 heterochromatinized region properties, 438 intercalary heterochromatin identification, ectopic contacts, 171-1 76 Q+-band occurrence, 147-148 telomeric heterochromatin DNA sequence locations, 252-255 ectopic pairing characterization,266-276 terminal deletion occurrence, 243-244 Position effect chromatid pompon formation, 421422 chromatin compaction, 400413,429 characteristics,402404 discontinuous compaction, 406407, 410 formation, 404-406,410 historical perspectives, 400-401 late-replication, 410 occurrence frequencies, 41 1 4 1 3 parental effects, 408-409 temperature effects, 407408 state changes, 422428 DNA underrepresentation,414-42 1 dominant position effects, 370-376 bw' locus, 371-376 ci+locus, 371 In(2LR)40d locus, 376

index Dubunin effect, 357-362 gene expression changes, 306-309 gene expression modification conditions, 334-355 chemical modifiers, 342-345 chromatin formation control, 351, 353-355 dose dependence, 349-350 genetic modifiers, 345-355 heterochromatin, 335-341 autosomes, 339-341 exogenous DNA, 341 X chromosome, 339-341 Y chromosome, 335-339 histone genes, 345 identification, 346-348 molecular-genetic characteristics,352-353 parental effects, 339,341-342 P-element-mediatedmutagenesis,346-347 temperature, 325,334-335,342 gene inactivation chromosomal rearrangements, 3 10-31 1, 316-318,329-330 embryonic development, 355-3 57, 44c-443 genes affected, 312-319 biochemically identified loci, 318 phenotype development control genes, 312-3 16 transposon genes, 318-319 vital genes, 316-318 historical perspectives, 376-378 inactivation levels, 327-329 initiation, 429434 intercalary heterochromatin, 391-394 maintenance, 434435 model, 427 pericentromeric heterochromatin, 378-391 heterochromatin amount effects, 381-382 inactivation strength, 383-385 rearrangement breaks, 385-390 revertant strain cytological analysis, 379-381,389-390 position effect variegation characteristics, 309-3 12 spreading, 31 1,330-334 telomeric heterochromatin, 394-395 variegation inactivation, 3 19-327

Index historical perspectives, 4-5,376-378, 400-40 1 light locus, 362-365 male fertility factors, 368 nucleolar organizer, 368-370 peach' locus, 366-367 Prokofyeva-Belgovskaya heterochromatization theory, 395-399 rolled' gene, 365-366 Prokofyeva-Belgovskaya, heterochromatization theory, 395-399 Prophase, heterochromatic staining, 6 n-Propionate, heterochromatin gene expression modification, 343 Protein centromeric heterochromatin structure, 105- 106 intercalary heterochromatin identification, ectopic pairing association, 184-185 telomeric heterochromatin protein characteristics, 280 Y chromosome mitotic heterochromatin, protein accumulation, 73-74 Pseudo-nursechromosomes, heterochromatin DNA underrepresentation, 128-130 Pyronine, intercalary heterochromatin identification, 146

Q-staining differential heterochromatin staining, 15-2 1 Drosophifa mefanogaster, 18, 20 Drosophila ViTilis, 17, 2 1 Escherichia coli, 17 Samoaia konenis , 16 intercalary heterochromatin identification, 141-143,147-148

Radiation eye mosaics, 322 heterochromatin compaction, 12-13 Repetitive sequences heterochromatin induction, 426-428 intercalary heterochromatin identification, 207-224 cytological repeats, 225 gene repeats, 208-209

565

mobile genome elements, 209-216 tandem repeats Chironomus chummi Cla-elements, 105, 219-220 dinucleotide repeats, 222-223 Drosophila melanogasrer X chromosome repeats, 222 histone encoding genes, 219 mononucleotide repeats, 22 2-223 pDv family repeats, 223 poly(A) nucleotide complexes, 220-222 polypyrimidines,219 rFWA genes, 218-219 scrambled repeats, 223-224 tRNA genes, 207-208 satellite DNA, 43-54 composition, 53-54 location, 45-46, 51-52 in siru hybridization, 51-53 slipping, 53 types, 47-49 telomeric heterochromatin, 248-259 Restriction enzymes, differential heterochromatin staining, chromosome digestion, 24 rolkd+ gene, position effect, 365-366

Samoaia konenis, differential heterochromatin staining, Q-staining, 16 Segregation distortion factor, autosomes, 61-65 Sex-determiningfactors, germline cell chromosomes, 304-306 Silencing, transcriptional domain repression maintenance, 424-426 Silver stain, intercalary heterochromatin identification, 150 Simulium morsitans, B chromosome studies, 283, 286,293-299 Slipping, satellite DNA, 53 Sodium butyrate, heterochromatin gene expression modification, 343 Somatic pairing, intercalary heterochromatin identification, 206-207 Southern blot, DNA underrepresentation analysis, 418-419 Staining methods, differential heterochromatin staining acridine orange stain, 146-150 acriflavine-nitrate, 150

566 ammoniacal silver stain, 150 C-staining, 14-15, 143 enzymatic chromosome digestion, 24 heterochromatin heterogeneity, 24 H-staining, 15-21, 143-146 methyl green stain, 150 N-staining, 21-23 pyronine, 150 Q-staining, 15-21 Drosophila melanogaster, 18,20 Drosophila wirifis, 17, 2 1 Escherichia cob, 17 intercalary heterochromatin, 143-146 Samoaia konenis , 16 telomeric heterochromatin identification, 279-280 Ste gene, Y chromosome, 66-68 Supernumerary chromosomes, see B chromosomes Suppressor genes, position effect modification, 350-355

Index polytene chromosome DNA sequence locations, 252-255 repeats, 248-259 Temperature embryonic development, 355-357 gene expression modification, 325,334-335, 342 heterochromatin compaction, 11-13, 281,

407408 Transposon genes, position effect inactivation, 318-319 P-element transformation, 308-309,3 11 tRNA genes, intercalary heterochromatin identification, 207-208

Vital genes, gene expression inactivation, position effect, 3 1 6 3 1 8

X chromosome Tandem repeats, intercalary heterochromatin identification, 2 16-224 Chironomus thummi Cla-elements, 105, 219-220 dinucleotide repeats, 222-223 Dsosophila melamgaster X chromosome repeats, 222 histone encoding genes, 219 mononucleotide repeats, 222-223 nucleotide sequences, 246 pDw family repeats, 223 poly(A) nucleotide complexes, 220-222 polypyrimidines, 219 rRNA genes, 218-219 scrambled repeats, 223-224 Telomeres, heterochromatin characteristics, 259-282 change manifestations, 281 differential staining, 279-280 ectopic pairing, 266-279 heterochromatin proteins, 280 DNA underrepresentation, 252-255, 280 inactivation, position effect, 394-395 overview, 238-248,259-266

centromeric heterochromatin DNA structure, 102-104 a-heterochromatin cytogenetics, 127-1 28 P-heterochromatin cytogenetics, 124-126 heterochromatin gene expression modification, 339-341 intercalary heterochromatin identification, tandem repeats, 222 mitotic heterochromatin characteristics, 57-60

Y chromosome centromeric heterochromatin DNA structure, 102 heterochromatin gene expression modification, 335-339 mitotic heterochromatin characteristics, 65-74 Drosophila hydei, 68-72 Drosophila melanogaster, 65-68 protein accumulation, 73-74 in situ hybridization, 6 6 7 2 Ste gene, 6 6 4 8


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