Advances in Genetics Volume 37

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Advances in Genetics

Volume 37

This Page Intentionally Left Blank

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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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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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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-

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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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Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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

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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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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%).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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: two- armed (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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Same as D. algonquin

Miller and Roy (1964)

Same as D. algonquin

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

Baimai et al. (1986)

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

Baimai et al. (1983)

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Baimai et al. (1983)

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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,

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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
Rsp DNA.

66

I. F. Zhimulev

It was also suggested that the SD phenomenon depends on the interaction of some sort of transposable elements (Sandler and Golic, 1985).

C. The Y chromosome The genetic organization of the Y chromosome in various species of Drosophila has been the subject of many reviews (Williamson, 1976;Leibovitch, 1977;Hess, 1980; Hillikeret al., 1980; Hennig, 1985,1987,1988,1990,1993; Pimpinelli etal., 1986; Hennig et al., 1987;Lifschytz, 1987; Hilliker and Sharp, 1988; Hennig and Ktemer, 1990; Gatti and Pimpinelli, 1992; Zhimulev, 1993; Palumbo et al., 1994a). After Bridges (1916) concluded that the Y chromosome plays a tole in the control of fertility in males of D. melanogaster, nine genes were identified in the entire Y chromosome (Stem, 1929; Neuhaus, 1938; Btosseau, 1960a;Williamson, 1972; Ayles et al., 1973; Sanders and Ayles, 1974; Kennison, 1981; Hazelrigg et al., 1982;Gatti and Pimpinelli, 1983;Hardy etal., 1984;Pimpinelli et al., 1986), which include bobbed, two loci affecting the course of meiosis, and six fertility factors in males (see Figure 36). The bobbed locus constitutes approximately 5% of the total DNA in the Y chromosome (Pimpinelli et al., 1986). The collochore locus is needed for regular pairing and, therefore, disjunction of the sex chromosomes in meiosis (Cooper, 1964). It makes up approximately 7% of the Y chromosome (Pimpinelli et al., 1986). Primary spermatocytes of D. melanogaster males lacking the Y chromosome exhibit proteinaceous crystals (Cox et al., 1976; Hardy et al., 1981, 1984). When chromosome regions occupied by the crystal (cry) locus break (see Figure 36), there is no associated phenotypic expression. However, with deletions in the region, protein-like crystals appear in the primary oocytes, and chromosome segregation is disturbed in meiosis (Hardy et al., 1984). It is noteworthy that, when the normal allele of the Stellate' gene is present in the X chromosome, the crystals are needle shaped, whereas when Ste- is present, they are star shaped (Hardy et al., 1984; see also Meyer et al., 1961). Males carrying cry Y and Ste X are sterile; the cry, Ste+ males are somewhat fertile but show chromosome nondisjunction (Hardy et al., 1984). I t is evident that the cry referred to as Su(Ste) (Livak, 1990) and Ste form a system of interacting genes (Pimpinelli et al., 1986; Gatti and Pimpinelli, 1992; Palumbo et al., 1994a). By genetic methods and in situ hybridization, Ste was mapped to the 12E1-2 band (Palumbo et al., 1994b), and cry to the h l l heterochromatic block of the Y chromosome (see Figure 36) (Hardy et al., 1984; Palumbo et al., 1994a,b). At the molecular level cry is an array of tandemly repeated (80 copies) Cfo I ftagment that is 800 bp long (Livak, 1984). In several populations 240 copies of cry were found (Lyckergaard and Clark, 1989). According to other data (Balakireva et al., 1992; McKee and Satter,

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

67

1996), the 2500- to 2800-bp repeats of the cry sequence consist of three main elements: the region of homology of the Ste gene, which is 800 bp long, an adjacent AT-rich Y chromosome-specificsegment, and a 1360-bp mobile element inserted into the Ste sequence (Balakireva et al., 1992). Some of the Ste sequences share homology with the DNA of the HeT family (Danilevskaya et al., 1991). The DNA of the Ste gene has been cloned (Lovett et al., 1980; Livak, 1984).The Ste DNA contains three classes of tandem repeats of 950, 1100, and 1150 bp after Cfo I digestion; they are partially homologous to the cry repeat (Livak, 1984; Palumbo et al., 1994b), and they are present in about 200 copies in Ste and at an order of magnitude less abundant in Ste' (Livak, 1984, 1990; Palumbo et al., 1994a,b). A part of the Ste sequence is located in the third locus of the genome. Although the majority, if not all, of the 1250-bp Xba I Ste fragments are mapped to the 12E region, almost all 1150-bp Xba I fragments are concentrated in the X chromosome heterochromatin (Shevelyov, 1992). These Ste sequences are located in SCLR elements comprising numerous middle repetitive sequences (Shevelyov, 1992,1993; Nurminsky et al., 1994). The SCRL elements were mapped in the distal region of the X chromosome heterochromatin between the right breakpoint of duplication BSYy and the left breakpoint of Df(l)b-3 (Nurminsky et al., 1994), or in the h27 heterochromatic region (Palumbo et al., 1994a,b) (see Figure 3 2 ) . According to other data (Tulin et al., 1997), one cluster of Stellate sequences is located in h26 and another in h27 heterochromatic blocks. The Ste sequences containing two introns are abundantly transcribed only in testes of cry- males; the corresponding 750-bp poly(A)+ RNA was not found in XY males. This sequence encodes a 19.5-kDa protein sharing homology with the p-subunit of casein-kinase 2 (Livak, 1990).The enzyme may be involved in chromosome condensation and segregation (see Palumbo et al., 1994a, for discussion). Ste' transcripts are detected in testicles. The presence of the normal Su(Ste) allele (=crystal) inhibits the accumulation of Ste' RNA. It has been suggested that Su(Ste) controls the activity of Ste'; removal of the Y chromosome leads to overproduction of Ste+ RNA (Livak, 1990), and, as a result, excess Ste' protein is crystallized in spermatocytes (Coyne, 1986). In analysis of the genetic bearing of the cry-Ste system, it should be borne in mind that Ste protein was not found in normal spermatogenesis (Bozzetti et al., 1995). Moreover, the Ste sequences are present only at reduced copy numbers and only on the Y chromosome in D. simulans and D. mauritiana. The sequence is apparently not present at all in D. erecta, D. teissieri, and D. yakuba (Livak, 1984). In Drosophila melanoguster males deficient for one or more fertility factors or lacking the entire Y chromosome, spermatids undergo an extensive elongation process, but spermiogenesis is not completed and sperm degenerates before maturation (Kiefer, 1966, 1973; Meyer, 1968; Hardy et al., 1981).

68

I. F. Zhlmulev

Of the six fertility factors, three (kl-5, kl-3, and ks-l ) each occupy about 10% (approximately 4000 kb) of the Y chromosome (Gatti and Pimpinelli, 1983; Pimpinelli et al., 1986; Bonaccorsi et al., 1988). A similar principle of genetic organization of the Y chromosome has been detected in D. busckii: removal of the left arm and three-fourths of the right leads only to sterility. The rest of the chromosome contains factors essential for survival (Krivshenko, 1952). The activity of fertility factors is manifested by the Y chromosome in a remarkable way. In 1961, Meyer and colleagues described peculiar stage-specific threadlike structures in the nuclei of D. melanogaster spermatocytes, which have been detected later in more than 50 other Dosophila species (Figure 39). Subsequently, success was achieved in demonstrating that these are decompacted regions of the Y chromosome (“loops”) in which RNA is synthesized and proteins accumulate (Hennig, 1967; Hess, 1973). Each loop in the nucleus is unique with respect to size, ultrastructure, differential staining, and appearance; in addition, the morphology of the loops is in general species specific as well (Meyer, 1963; Hess, 1967, 1973; Yamasaki, 1977, 1981). There is evidence that the loops are formed from the material of the Y chromosome: 1. In males without the Y chromosome (XO),there are no loops, and in XYY males, they are present in two sets (Figure 40). When a part of the Y chromosome is deleted, not all the loops are detected; in strains with duplications, the number of loops correspondingly increases (Hess and Meyer, 1963, 1968; Hess, 1965a, 1974; Bonaccorsi et al., 1988). 2. In interspecific hybrids with a Y chromosome inherited from D. neohydei, the morphology of the loops is the same as in the donor species of the Y chromosome (Hess, 1974). Evidence that the fertility genes are located in loops is as follows: 1. When at least one loop is deleted, the male is rendered sterile (Hess, 1965a).

2. At first, a correlation between the number of genes and loops was revealed (Hess and Meyer, 1963; Hennig et al., 1974; Ayles et al., 1973; Sanders and Ayles, 1974; Leoncini, 1977). Then, with the use of chromosomal rearrangements, a direct correspondence was established between factors and loops: the kl-5 factor to loop A, the kl-3 to loop B, and the ks-l to loop C in D. melanogaster. This correspondence is incomplete, however, because the chromosome regions occupied by the kl-5 and ks-I factors are much longer than the loop-forming site. The minimum size of the kl-5 factor corresponds to the size of the loop-forming site (Bonaccorsi et al., 1988). Deletion or mutation of the loop-forming gene kl-5 results in the absence of outer dynein arms in each peripheral microtubule in the sperm tail axoneme,

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

69

Figure 39. Organization of the Y chromosome in D. hydei. (a) General appearance of a spermatocyte nucleus. The regions occupied by loops are designated (Tr, Th, Ps, CI, Ns); Nu, nucleolus. (b) Scheme of loops location in the nucleus of a spermatocyte. (c and d ) Location and designation of loops ( c ) and also their location on the map of differential staining of the mitotic chromosome (d). After Hennig (1985).

the structure that provides the mechanical force for sperm movement (Hardy et al., 1981, 1984; Goldstein et al., 1982; Hepner and Hays, 1993). The Y chromosome of D. hydei has 7-16 complementation groups need-

70

1. F. Zhimulev

Figure 40. Variation in number and morphology of loops in D. hydei strains with translocarions between the Y chromosome and autosome. A, the autosome, the normal partner of the chromosome with translocation; F L p . A , ,long, and A,.YL”,short elements of translocation; Fd, Fk,p, T,K, loops; N, nucleolus. After Hess (1965a).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

71

ed for male fertility, 5 of which are related with lampbrush loops (Hennig, 1985; Hackstein, 1987). After the generation of clones of DNA from the Y chromosome (Lifschytz, 1979; Vogt er al., 1982; Hennig er al., 1983), analysis of its molecular organization became feasible (see reviews by Hennig, 1985; Hennig et al., 1987). Both the loop-forming (kl-5, kl-3, and ks-I ) and the non-loop-forming (kl-2 and ks-2) fertility genes in D. mehnogmter contain substantial amounts of satellite DNA. Moreover, each fertility region is characterized by a specific combination of satellite sequences (Bonaccorsi and Lohe, 1991). The total length of all the loops in D. hyd-ei makes up approximately 1000 km, or 1/12th of the total length of DNA in the Y chromosome. The functions of the remaining 11/12ths are unknown (Hennig et al., 1974). The DNA of the Y chromosome of D. hydei comprises repetitive sequences of two types:

1. The Y-specific,satellite-like sequences; these occur only in this particular chromosome, and are represented by families of 200-2000 copies. They are organized in clusters of tandemly repetitive units arranged in loops. The ayl family provides the best studied example (Vogt er al., 1982; Vogt and Hennig, 1986a). It is transcribed in the Q fertility gene forming the Nooses lampbrush loop. These repeats do not show protein-coding capacity (Lifschytz et

al., 1983; Vogt and Hennig, 1983;Hennig et al., 1987, 1989; Huijser and Hennig, 1987; Huijser et ul., 1988; Trapitz et al., 1988, 1992; Wlashek et al., 1988). 2. The Y-linked sequences; these occur in both the Y and other chromosomes, but less frequently. Their number in a site varies from several to 50 copies of different lengths (Vogt and Hennig, 1986b; Vogt et al., 1986; Hennig et al., 1987). In at least three of the five loop-forming genes of D. hydei, retrotransposons have been identified as Y-linked loop constituents. Members of the micropia family are located in the fertility genes forming the loop pairs Threads and Pseudonuckolus. In the X chromosomes and autosomes, 2-1 1 copies of the micropia are found, while their number varies from SO to 100 in the Y chromosome. The Y-linked members of the retrotransposon family are defective. Micropiacopies have been found in D. melanogaster (Huijser et al., 1988; D.-H. Lankenau et al., 1988,1989,1990;Lankenau and Hennig, 1990; Lankenau, 1993; S. Lankenau et al., 1994). Retrotransposons of the gypsy family are found in the Nooses lampbrush pair, where they occupy about one-third of the Y-linked DNA of the loop (approximately 40-50 kb of DNA). Most, if not all, gypsy elements within the loop are truncated and defective. The gypsy sequences occur interspersed between the Y-specific loop constituents, the ayl repeats (Hochstenbach et ul., 1993a,b, 1994a-c). Repeats specific to the Y chromosome were found in other dipterans (e.g., Ceratitis capitata (Anleitner and Heymer, 1992). The sequences specific to

72

1. F. Zhimulev

the Y chromosome are not evolutionary conserved because many occur either in closely related species or only in D. hydei (Huijser and Hennig, 1987). The Ylinked sequences are more conserved because many are identifiable in the genomes of remote species (Vogt et al., 1986). In some cases, the Y chromosome contains remarkable repeats; thus, in D. simulans, a genotype was identified lacking the cluster of rRNA genes, yet possessing a repeat composed of a nontranscribed spacer (240 bp), reiterated about 12,500 times, which ultimately generates a tract approximately 3000-kb long (Lohe and Roberts, 1990). Danilevskaya et al. (1991) have sequenced DNA from the cry region, which not only shows homology to Ste but is also related to the HeT family of repetitive sequences (Traverse and Pardue, 1989). HeT-A is a novel transposable element that appears to form part of the telomeres of D. melanogaster chromosomes. It occurs in nontelomeric regions of the heterochromatic Y chromosome in regular tandem repeats of parts of the HeT-A sequence joined to nonrelated sequences (Danilevskaya et al., 1993). Evidence for the genetic activation of loops is RNA synthesis: about 50% of extranuclear RNA is synthesized in the loops. The remaining 50% of the label is incorporated into the transcriptionally active regions of the X chromosome and the autosomes not forming the loops. After pulse incorporation of 3H-uridine, radioactive RNA is retained in loops for 30-40 hr (Meyer and Hess, 1965; Hennig, 1967). Under the effect of actinomycin D (Meyer and Hess, 1965; Hess and Meyer, 1968) or irradiation (Hess, 1965b), the loops disappear. a-Amanitin exerts a similar action (Hennig et al., 1974). The presence of RNA polymerase I1 has been demonstrated with the use of the method of indirect immunofluorescence (Rungger-Brandle et al., 1981). In loop material of D. hydei spread by Miller’s technique, the formation of a gradient of distribution of chromosome material is seen along the loop axis, like that characteristic of transcription visualization (Hennig et al., 1974; Grond et al., 1983). The loop transcripts are usually long. Judging by ultrastructural data, the Nooses, Threads, and Pseudonucleolus loops represent transcription units with lengths of 260 to more than 1500 kb (Meyer and Hennig, 1974; Glatzer and Meyer, 1981; Grond et al., 1983,1984; DeLoos et al., 1985; Hochstenbach et al., 1996). From in situ hybridization of DNA clones isolated from various loops of the Y chromosome of D. hydei with nuclear RNA, each clone was shown to hybridize with its own nuclear domain (Lifschytz et al., 1983; Trapitz et al., 1992; Hochstenbach et al., 1994a, 1996). Different types of repeats, including those composed of satellites, are transcribed in loops. The repeat of sequence AAGAG (the 1.686 satellite) hybridizes in situ and on Northern blots with the RNA of the kl-5 and ks-l loops in D. melanogaster. It is noteworthy in this respect that transcript sizes vary from maximum (with migration at limiting mobility in gels) to less than 1 kb (Bonaccorsi eta[., 1990). Repeats of both types are transcribed in D. hydei, the Y specific and the

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

73

Y linked, with transcripts from 260 to more than 1500 kb identified on Northern blots (Vogt et al., 1982; Lifschytz et al., 1983; Vogt and Hennig, 1983, 1986a,b; Lifschytz and Hareven, 1985; DeLoos et al., 1985; Hareven et al., 1986; Huijser and Hennig, 1987; Huijser et al., 1987, 1988, 1990; Trapitz et al., 1988, 1992; Wlaschek et al., 1988; Hochstenbach et al., 1993a,b, 1994a,c). The mobile micropia and gypsy elements are expressed as constituents of lampbrush loops (S. Lankenau et al., 1994; Hochstenbach et al., 1994b, 1996). In addition, micmpia encodes full-length RNA that extends between the two long terminal repeats of the element (S. Lankenau et al., 1994). Other repeats are transcribed, too. For example, the T14 clone isolated from a cDNA library, derived from testis poly(A)+ RNA of D. hydei, has a multiple location in the Y chromosome and in salivary gland polytene chromosomes (Brand and Hennig, 1989). In spite of the active transcription of loop DNA, sequence determination gives no reason for believing that the loops can encode proteins (Hennig et al., 1987). Large amounts of various proteins, which constitute the bulk of proteins in the nuclei of spermatocytes, accumulate in loops (Grond et al., 1983, 1984; Glatzer, 1984; Hulsebos et al., 1984; Hennig, 1985; Glatzer and Kloetzel, 1986; Tschendorf et al., 1989; Pisano et al., 1993). A part of the proteins is included in RNP particles filling the matrix of the loops (Glatzer and Kloetzel, 1985; Meltzer and Glatzer, 1985; Bonaccorsi et al., 1988). The proteins accumulated in the loops of the Y chromosomes are encoded by the genes of the other chromosomes. The following is adduced as supporting evidence:

1. Autosomal recessive mutations, whose action leads to the disappearance of the corresponding loops, were identified (Hackstein et al., 1982, 1987). 2. Males receiving the Y chromosome from D. neohydei and the autosomes from D. hydei can be obtained from interspecific crosses (Hennig, 1977). The structure of loops in such males resembles that in D. hydei, although it is clear that loop morphology depends not only on protein composition, but also on transcription intensity, transcript size, and morphology of RNP-containing particles (Hennig, 1985). 3. With the use of antibodies, certain loop proteins are identified in spermatocyte nuclei in individuals without the Y chromosome, and, consequently, not forming loops (Kloetzel et al., 1981; Bonaccorsi et al., 1988; Tischendorfet al., 1989; Pisano e t al., 1993).Certain loop-associated proteins are testis specific: 80-kDa testis protein (Hulsebos et al., 1983, 1984) and T53-1 tektinlike protein (Pisano et al., 1993). Antisera raised against proteins from synaptonemal complexes (sc) from rat spermatocytesreact strongly with Y chromosomal lampbrush loops in D. mlanogaster and D. hydei. Each serum recognizes one or two loops (Hennig et al., 1995).

74

1.

F. Zhimulev

Lifschytz (1987) holds the view that the loops may have a dual function: (1) they regulate the development of spermatids, forming the regular nuclear matrix specific to spermatocytes; and (2) loop formation is a consequence of decompaction and transcription of repeats. Gatti and Pimpinelli (1992) speculate that Y loops fulfill protein-binding functions. According to Pisano et al. (1993), loops present only in the Drosophila genus have evolved to facilitate the development of the giant sperm tails (2 mm in D. mehogaster and 1 cm in D. hydei) characteristic of this genus (Gatti and Pimpinelli, 1992). The DNA coding for a set of transcripts expressed in the germline of D. melanogater males has been found in the Y chromosome heterochromatic regions h18-hl9. In contrast to D. mehogaster, where Y-linked copies were found in nine different wild-type strains, no Y-linked copies were found in sibling species (Russell and Kaiser, 1993).

D. Genetic degeneration of the neo-Y chromosome in Drosophila miranda Drosophitamiran& males show only nine chromosomes in metaphase because they are apparently monosomic for one of the autosomes (Figure 41). This autosome has been designated X2 and corresponds to the third chromosome in D. pseudoobscum and 2R in D. melanogater (Dobzhansky, 1935b; MacKnight, 1939; Steinemann, 1982b; Steinemann et al., 1984). The homolog of X2 has been attached to the Y chromosome and is now referred to as the neo-Y chromosome. In situ hybridization of telomeric DNA to male metaphase spreads reveals a massive labeling block within the neo-Y in addition to the labeling sites at chromosome ends, thus implying an “end-to-end” fusion mechanism (Steinemann, 1984). The neo-Y is polytenized in the salivary gland cells (MacKnight, 1939; Steinemann, 1982a).

aX’ \

R4

b X’ \

44

Figure 41. Diagram of the Drosophila miranda female (a) and male (b)

karyotypes. After Steinemann and Steinemann (19911.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

75

The attached autosome is gradually acquiring the features of the Y chromosome, displaying changes in chromosome structure and loss of gene activity. The X2chromosome is evolving into an X chromosome. Certain genes on the X2 are now capable of dosage compensation and enhanced transcription in males that compensate for the inactivity of the homologous gene on the neo-Y. Thus the system is a very good model of genetic degeneration of the Y chromosome. In situ hybridization using cRNA transcribed from unfractionated D. mirun& DNA reveals hybridization to the neo-Y with dispersed label distributed over the entire chromosome. The former partner chromosome, X2, is essentially unlabeled. This suggests that repetitive DNA sequences “invade” the translocated chromosome (Steinemann, 1982a). One of the repeats, a BumHl element, is predominantly associated with degenerating neo-Y chromosome, but absent in the D. melanoguster genome (Ganguly et ul., 1992). The TRAM transposable element was found in neo Y chromosome. Its size is 3,452 bp, including long terminal direct repeat of 372 bp on both sides. In situ hybridization with TRAM subclones showed massive accumulation of TRAM in the neo-Y chromosome (50-60 labeled sites), while the former homologue, the X2 chromosome, contains no TRAM .sites (Steinemann and Steinemann, 1997). Staining D. mirunda with antibodies against isoform of histone H4 acetylated at lysine 16, showed that the X2 shows much brighter staining over 90% chromosome length than neo-Y (Steinemann et al., 1996). Steinemann and Steinemann (1990, 1992) have analyzed the region around the four larval cuticle proteins (Lcp) from the X2and compared it with the homologous region from the neo-Y chromosome. The striking difference between the two chromosome regions is the large number of insertions on the neo-Y but not on the X2. Sequence analysis of Lcp locus cloned from the X2and Y chromosomes reveals a massive accumulation of inserted DNA sequences (ISYs) in the Y chromosome Lcp region (Figure 42). In situ hybridization with the TRIM retrotransposon (Steinemann and Steinemann, 1991) and ISYs insertions show a differential distribution of these transposable elements on the X2 and the neo-Y chromosomes. The transposable elements show heavy hybridization over the formerly autosomal sequences that are now on the neo-Y. In contrast, the X2 homolog shows approximately the same density of sites of hybridization as do the autosomes and the X chromosome (see Figure 42). There are insertions of many other transposing elements in this region (Steinemann and Steinemann, 1992, 1993).Thus, in the neo-Y chromosome, all the four Lcp genes are embedded within a dense cluster of transposable elements. Numerous deletions and a large duplication were found on the neo-Y chromosome in comparison with the X2 chromosomes (Steinemann and Steinemann, 1993). As a result, while the X2 Lcp-l through Lcp4 loci are expressed, the Y chromosomal Lcp-3 locus shows only reduced activity and the Lcpl , Lcp-2, and Lcp-4 are completely inactive. The ob-

76

I. F. Zhimulev

C

x2 Lcp 1

H H C h

1 kb

LCp REGION C

XhH

Lcp 2

R

H

Lep 3

C Sa

B

XhXh

Lcp 4

r----------4

‘

:

DUPLICA

Y

Dv4

la

DV6

Figure 42. Schematic view representing the Lcp region at the X2 and neo-Y chromosomesof D. miranda. Unsequenced DNA sections are indicated by lines flanked by arrowheads at both sides. Large open arrowheads indicate the direction of transcription. neo-Y chromosome sequences (ISYs)additional to X2 chromosomeare shown by stippled boxes. The ISY2X2, ISYZY, and ISY2/3 insertions are indicated by hatched boxes. The large 1SY4 and ISY5 insertions identified as retrotransposons are designated TRIM and TRAM. Missing DNA sequences in the neo-Y chromosomal region are indicated as deletions (DY). The duplication is indicated by a dashed box. After Steinemann and Steinemann (1992,1993) and Steinemann et al. (1993).

tained results suggest that neo-Y Lcp-f and Lcp-3 loci are silenced by neighboring transposable elements (Steinemann and Steinemann, 1992; Steinemann et al., 1993,1996a,b). Thus, in a relatively short evolutionary span from the time the autosome was translocated onto the Y chromosome, the sequence of the homologs has evolved very differently. The sequence on the neo-Y has acquired many more copies of insertions than X2 homolog, and Y chromosome degeneration is driven by the accumulation of transposable elements, especially retrotransposons. In the evolutionary process of Y chromosome degeneration, the mechanism of silencing might be the first step and could even be a prerequisite for degradation (Steinemann et al., 1993; Steinemann and Steinemann, 1997). In other species, Drosophila americanaamen’canu, three loci, coding for the enzymes enolase, phosphoglycerate kinase and alcohol dehydrogenase, are located in the chromosome 4,which forms the neo-Y and neo-X chromosomes. Genes coding for these enzymes carry active alleles on the neo-Y chromosome. These data suggest that the neo-Y chromosome has undergone very little degeneration (Charlesworth et al., 1997).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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E. Other features of the genetic organization of heterochromatin One genetic feature of the heterochromatin of D. melanogaiter is low frequency of crossing over (Muller and Painter, 1932; Mather, 1939; Baker, 1958; Roberts, 1965; Schalet and Lefevre, 1976; Carpenter and Baker, 1982; Charlesworth et al., 1986; Lake, 1986; McKee and Handel, 1993. For example, the distance between the genes car+ (18D1-2 region) and su(f)+, located at the eu- and heterochromatin junction, is 3.54 m.u., while the distance between the su(f>+and they+ genes, transposed to the right arm of chromosome XR, is 0.04 m.u. It should be noted that there are, in all, about 60 bands between the genes in the former case, and the entire heterochromatin (i.e., 40% of all the X chromosome) between su(f)+ and y’ in the latter case (Roberts, 1965). Using a similar approach, Baker (1958) revealed an analogous phenomenon in

D. virilis. A comparison of crossing-over frequencies in blocks of centromeric and intercalary heterochromatin is of interest. Recombination frequency between the bristle’ and light’ genes lying at the boundary of the repeat cluster of the histone genes (approximately 500 kb), was 0.042%, and that between the light’ and straw+ genes (between which the entire block of centromeric heterochromatin of the second chromosome is located) was 0.047%. When pupae ages 126-168 hr (the primary oocytes are layered at the age of 126-132 hr) are treated with high temperature (35”C), crossing-over frequencies increase 30- to 35-fold in both regions (Grell, 1978). The presence of sites of meiotic pairing in the sex chromosomes may be regarded as another feature of the genetic organization of heterochromatin. Gershenzon ( 1933a,b, 1940) established that the heterochromatic (“inert”) regions of the X chromosome are important for meiotic pairing. Cooper (1964) called these loci “collochores” (col). Since the X and Y chromosomes of Drosophita are very different morphologically and functionally, the question of the mechanisms of pairing of these elements at prophase I of meiosis is quite timely. Much evidence exists for the presence of pairing sites in heterochromatin of the X chromosome (Muller and Painter, 1932; Gershenzon, 1933a,b, 1940; Sandler and Braver, 1954; Cooper, 1964; Peacock, 1965; Peacock and Miklos, 1973; Yamamoto and Miklos, 1977; Karpen et al., 1996), particularly in the region of the nucleolar organizer (Cooper, 1964; Merriam, 1968; Sandler, 1975; Peacock et al., 1975; Appels and Hilliker, 1982; McKee, 1984, 1987, 1996; McKee and Lindsley, 1987; Dernburg et al., 1996). McKee and Karpen (1990) demonstrated that proper X-Y recognition and segregation in male meiosis is dependent upon ribosomal DNA repeats. Using P-element-mediated transformation, McKee et al. ( 1992) further demonstrated that this recognition is due to 240-bp repeats in the nontranscribed spacer, which is present in tandemly repeated copies (IGS) between each of the rDNA repeats. While a single copy of the IGS element inserted ectopically into the

78

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F. Zhimulev

X-euchromatin provides a significant rescue of X-Y pairing, proper recognition and segregation improves with increased numbers of these repeats. Internally deleted copies of an IGS with fewer than six repeats of the 240-bp element were unable to ensure X-Y segregation (McKee and Karpen, 1990; McKee et al., 1992). Some cytogenetic data run counter to this hypothesis. According to Park and Yamamoto (1993, the pairing site is in the region distal to the In(l)scv2 breakpoint within the cluster of rRNA genes. Attempts to detect sites of meiotic pairing in the autosomal heterochromatin of Drosophila were unsuccessful (Yamamoto, 1979; Hilliker, 1981a,b; Hilliker et al., 1982; McKee et al., 1993), although heterochromatic homology seems responsible for the segregation of the achiasmate X and fourth chromosomes (Hawley et al., 1993; Irick, 1994). One of the peculiarities of heterochromatin is its tight contact with the nuclear envelope (see Zhimulev, 1993, 1996, for review). DNA sequences associated with nuclear lamina were found within p-heterochromatin of Drosophila melanogaster (20CD, 40EF, and 41AB, 81 and 101 regions). They are represented by two repeated HindIII-EcoRI fragments 1.4 and 1.6 kb long. About 120 copies of these sequences were detected per haploid genome (Baricheva et al., 1995, 1996; Sharakhov, 1995, 1997; Bogachev et al., 1996).

V. DIMINUTION OF CHROMATIN AND CHROMOSOMES Differentiation of the germ and somatic cell lines, associated with loss of a part of the genetic material during early embryonic development (chromatin diminution, chromosome elimination), is quite widespread in nature. Different degrees of diminution occur in certain species of ascarids, cyclops, infusoria, fishs, ticks, beetles, butterflies, and flies (for review, see Wilson, 1936; Beams and Kessel, 1974; John and Miklos, 1979; Stanley et al., 1984; Ammermann, 1985, 1987; Hennig, 1986; Tobler, 1986; Pimpinelli and Goday, 1989; Tobler et al., 1992).

A. Chromatin diminution in ascarids (Nematoda) In Ascaris megalocephalu ( =Parascaris univakns), the zygote contains two chromosomes, a thin centromeric and a thicker terminal region can be identified in each (Figure 43). Boveri (1887) found that, as early as at the second cleavage, the thick ends of the chromosomes in a cell of the cleaving zygote separate from the middle part and, having no centromeres, remain in the regions of the equator of the mitotic spindle, where they eventually degenerate. As a result, a substantial portion of the chromosome is lost, chromatin diminution, as Boveri has termed it, takes place. The cell that underwent diminution gives rise to a cell clone (Figure 44)with much shorter chromosomes. Chromosome diminution does not take place in the second daughter cell, and it will give rise to two new cells: one will

Poiytene Chromosomes, Heterochromatin, and Position Effect Variegation

79

Figure 43. C-banding of the mitotic chromosomes of Parascaris univalens (2n = 2) (a) and P. equorum (2n = 4) (b) before diminution. After Goday et al. (1985).

undergo diminution, the other will not. As a consequence, at the 32-cell stage the embryo consists of two cells with the complete set of DNA sequences (see Figure 44), and these two will form the germlines. The remaining 30 will give rise to all the somatic cells. A similar diminution pattern has been described for another species, A. lumbricoides (Bonnevie, 1902). In 11 other nematode species chromatin diminution occurs during the fifth through eighth cleavage divisions (Tobler et al., 1992). It was shown that in ascarids the centromere is holocentric and the threads of the spindle are attached to numerous regions along the entire length of the chromosome. In presomatic cells of the early cleavage divisions, the microtubes of the spindle are attached only to the euchromatic regions (Goday et al., 1992), and this presumably is the reason why diminution takes place. In P. univukns, the ends of the chromosomes cast off during diminution (up to 80% of all chromatin; see Moritz, 1970a) have the features of heterochromatin, such as H- or C-stainability (Goday and Pimpinelli, 1984, 1986; Goday et al., 1985; Pimpinelli and Goday, 1989; see Figure 43) and the capacity to hybridize in situ with satellite DNA (Goday and Pimpinelli, 1989; Pimpinelli and Gcday, 1989). The amount of heterochromatin can vary, as judged by the fact that eggs obtained from the same female differ considerably in DNA amounts; after the first cleavage division, such differences are found between homologous chromosomes. DNA content in the spermatozoa of different males varies as well (Moritz, 1970b).The central part of the chromosome not eliminated during diminution does not stain for H- and Cheterochromatin, and it does not hybridize with satellite DNA (Goday and Pimpinelli, 1984, 1989;Goday et al., 1985; Pimpinelli and Goday, 1989). In P. equorum (=A. megalocephala bivalens), the terminal portions of the

80

1. F. Zhimulev

GERM LINE

.

Mesoderm 1

I

SOMA CELLS

Figure 44. Schematic representation of the cell lineage and segregation of germline and somatic cells in Ascaris lumbricoides mum. The presumptive primordial germ cells (P,-P,) are indicated by half-filled circles, and primordial germ cells (P4,P5a, P5& giving rise to all germ cells by filled circles. The presomatic cells S,, S,,, and S,-S, undergoing chromatin diminution are represented by open circles surrounded by black dots. These cells eventually form, in a precisely determined manner, the various parts of the embryo that are indicated. After Tobler et al. (1992), modified from Boveri (1904), and Beams and Kessel(1974).

chromosomes are also thicker; in addition, there are numerous tangles of dense material in their middle section (Navashin, 1936).Thickenings of all types show C- and H-banding (see Figure 43); these thickenings are eliminated during diminution (Goday and Pimpinelli, 1984;Goday et al., 1985).The eliminated DNA is enriched in repeats. Thus, in P. equorum, germline cells contain two light satellites, which together make up approximately 85% of zygotic DNA. Precisely these satellites are eliminated from somatic cells (Moritz and Roth, 1976). In Ascaris suum ( = A . lumbn'coides), the amount of eliminated DNA varies 2.5-fold: 22% (Moritz and Roth, 1976),27% (Tobler et al., 1972,1985), 34% (Pastemak and Barrell, 1976),and 56% (Davies and Carter, 1980).It is noteworthy in this respect that, according to the data of Roth and Moritz (Moritz and Roth, 1976;Roth, 1979;Roth and Moritz, 1981),the highly repetitive DNA with repeat units 125 and 134 bp long is mainly eliminated. According to the data of Mueller et al. (1982),repetitive DNA contains a set of different, yet sharing some similarities, sequences with a unit length of 120 bp.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

81

Tasl and Tas2 middle repetitive retrotransposons are present at about 50 copies per haploid germline genome. Approximately 25% of Tasl and 100% of Tas2 are expelled from somatic cell lineages (Aeby et al., 1986; Felder et al., 1994). The ratio of repetitive to unique sequences in the eliminated DNA has been estimated as 1:l (Tobler et al., 1972; Goldstein and Straus, 1978). A cluster containing at least two different single-copy genes eliminated during chromatin diminution was found. One is the ALEP-1 gene encoding the 148-amino-acid protein RpS19G, homologous to the small ribosomal subunit protein S19 of eukaryotes (Etter et al., 1991). The gene is normally expressed in Ascaris germline cells (Etter et al., 1991,1994;Tobler et al., 1992).A t least one more single-copy gene, fert- I , becomes eliminated from all somatic cells during chromatin diminution in A. lumbncoides (Spicher et al., 1994). Approximately 99% of satellite DNA present in the germline cells is eventually eliminated during diminution. The proportion of the 18s and 28s rRNA genes in the pre- and postdiminution nuclei is the same; this indicates that at least large blocks of the rRNA genes are not eliminated (Tobler et al., 1985; Landolt and Tobler, 1988). Clones with small copy number, or unique DNA not present in somatic cells, were isolated from germline cells (Muller et al., 1986). Elimination results in the formation of somatic nuclei consisting of 90% unique sequences and 10% repetitive DNA. Only 0.01-0.05% of DNA is highly repetitive (Roth and Moritz, 1981; Mueller et al., 1982). After removal of heterochromatin fragments, chromosomal fragmentation at CBR (the chromosome break region) is followed by the addition of about 2-4 kb of telomeric TTAGGC repeats (Muller et al., 1991).

B. Chromatin diminution in Cyclops Chromatin diminution was detected in at least 12 species of Cyclops, in addition to ascarids (S. Beermann, 1959, 1960, 1977, 1984; Stich, 1962; Kiknadze, 1972; Akifjev, 1974; Beermann and Meyer, 1980; Akifjev and Grishanin, 1993; Grishanin et d., 1996). The course of diminution is restricted to presumptive somatic cells, and it occurs in various species during the fourth through seventh cleavage divisions. The diploid chromosome number in C. strenuus is 22. It does not change during chromatin diminution, although chromosome diameter becomes twofold shorter (from 0.8 to 0.2 pm). The chromosomes of C. strenuus at the stage of 32-64 blastomeres are 2.5-3.0 times shorter and 3.0-4.0 times thinner than the chromosomes at the first cleavage division (Kiknadze, 1972) (Figure 45). Chromosome length decreases from 11-22 to 2.6-7 pm (Grishanin and Akifjev, 1993; Grishanin, 1995) and diameter of nuclei decreases from 10.6 to 3.4 p m (Grishanin, 1995). In C. kolensis, the time interval between the third (prediminutive) and

Figure 45. Diminution of chromatin in Cyclops furcifer (a and b) and C. strenuus (c-0. (a) The chromosomes before diminution. K, centromeres; T,telomeric regions. (b) Chromatin to be eliminated in the equatorial zone of late anaphase. (c and d) Chromosomes before (c) and after (d) diminution. (e and f) Droplets of eliminated chromatin in the nucleus ( e ) and cytoplasm (0. Pairs a and b, as well as c and d, are at the same magnification. (a and b) after S. Ekermann (1966); (c-f) after Akifjev and Grishanin (1993).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

Figure 45. Continued

83

84

I. F. Zhimulev

fourth cleavage divisions is 6 times longer than that between the first three divisions and 10 times longer than intervals between following divisions (Grishanin, 1996a). The chromosomes at the third cleavage division (prediminutive) show differential banding even after simple acetoorcein staining. In interphase prediminutive nuclei no chromocenters were found, in spite of abundant heterochromatin (Grishanin and Akifjev, 1993). Chromatin is eliminated from different regions of the chromosomes: terminal heteropycnotic fragments in Cyclops divulsus and C. furcifer or intercalary fragments in C. strenuus. It remains as large blocks, drops, or granules of Feulgenpositive material in the equatorial part of the division spindle (see Figure 45). The number of granules at the beginning of diminution is about 600 in C. koknsis and 150 in C. strenuus; subsequently, they fuse and their number reduces to 150. The granules are rounded or elongated structures distinctly separated from cytoplasm by a membrane 50 nm thick. Chromatin is exceptionally condensed. After two to three cell divisions, the drops degenerate. The large number of granules at the early steps of diminution provides evidence for the numerous start points of elimination (Akifjev and Grishanin, 1993; Grishanin and Akifjev, 1993; Grishanin, 1995, 1996a,b). It is believed that the diminution process is generally similar to the excision mechanism of bacteriophages from the Escherichia coli genome, that is, formation of loops and recombination at the point where the bases of the loops come into contact (S. Beermann, 1966; W. Beermann, 1966; Akifjev, 1974). When nuclei at the interphase, the stage preceding division, are spread, chromatin rings composed of fibrils 25-30 nm thick and 0.6-16 km long are identifiable in C. divulsus, and 0.4-40 pm in length in C. furcifer (Beermann and Meyer, 1980; Beermann, 1984).When nucleosomal chromatin isolated at the diminution stage is spread by Miller’s technique, rings ranging in length from 0.6 to 100 pm can be seen (Beermann, 1984). According to cytophotometric data, 65-75% of DNA is eliminated in C. furcifer during diminution (S. Beermann, 1966). The ratio of haploid DNA content in germline cells to DNA content in somatic cells is 2.2:0.9 pg for C. strenuus, 2.9:1.44 for C. furcifer, and 3.1:1.8 for C. divulsus (Beermann, 1977). Grishanin et al. (1994) and Grishanin (1995) have shown that 94% of DNA are eliminated in C. koknsis (the species is C. strenuus in their paper, but the correct species assignment is C. koknsis, according to A. P. Akifjev, personal communication), and in real C. strenuus 75% of DNA is eliminated (Grishanin et al., 1996). Special tests were not done to refer the eliminated parts of the chromosomes to heterochromatin; nevertheless, it is believed that it is precisely heterochromatin that undergoes diminution because these regions are heavily heteropycnotic (S. Beermann, 1966;W. Beermann, 1966) (see Figure 45). According to the view of Prokofyeva-Belgovskaya(19861, the genetically inactivated regions of eu- and heterochromatin are eliminated.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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Akifjev and Grishanin (1993) suggest that the diminution process may arise under the effect of a single mutation activating the diminution enzyme. All the remaining steps of the process, in their view, unfold under normal conditions.

C. Elimination of chromosomes in dipteran insects In 1908, W. Kahle detected chromatin diminution in the gall midge Miastor me~raloasthat, as proved later, is associated with loss of whole chromosomes (Huettner, 193313; Reitberger, 1934, 1940; Kraczkiewicz, 1935, 1936, 1937; Metcalfe, 1935; White, 1946). This phenomenon occurs in the representatives of three families of the Diptera order. A distinction is made between the B chromosomes and the chromosomes restricted to germline cells; the latter are given different names: E chromosomes in the representatives of the Cecidomyiidae, K chromosomes in those of the Chironomidae and Orthocladiinae, and L chromosomes in those of the Sciaridae families. The chromosomes of the main set are termed the S (somatic) chromosomes (White, 1950, 1954). The elimination process may be considered by referring to the exemplary representatives of the Orthocladiinae subfamily, whose differentiation of germline cells has three distinguishing features:

1. Elimination of all the germline-restricted chromosomes takes place at the fifth through seventh cleavage divisions in somatic chromosomes.

2. When division of germline cells resumes after hatching, the K chromosome is reduced to half in the zygotes due to partial losses at anaphases. 3. This decrease in number is compensated for by a specific duplication at the differentiation stage of spermatocytes and oocytes, when all the spermatogonia and oogonia undergo one differential division. The K chromosomes reduplicate at prophase, then, with omission of chromatid division, they assemble at one pole of the spindle. As a result of the next meiosis, one daughter cell has the complete set of the K and the S chromosomes, while the other possesses only the somatic set. One cell will give rise to the oocyte in females and to the spermatozoid in males; the other will give rise to the nurse cell of the oocyte in females and to the degenerating spermatocyte in males (Beermann, 1956). Other features were revealed in the midges: the E chromosomes are completely eliminated as early as during spermiogenesis and partly during oogenesis, and, for this reason, the mature spermatozoon contains only the S chromosomes, and the oocyte contains the S and a part of the E chromosomes (White, 1950, 1973; Geyer-Duszynska, 1961; Stuart and Hatchett, 1988b). Furthermore, it is believed that in the midges the S chromosomes represent the diploid set segregating from the octaploid, and elimination of the E chromosomes means removal of the hexaploid set (Huettner, 1934; Kraczkiewicz, 1935, 1936; Reitberger, 1940; White, 1946,1947; Camenzind, 1966; Panelius, 1971; Bregman, 1975;Sager and

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Kitchin, 1975). It has not been clarified if this is, indeed, the case. The results of cytological analysis of chromosome behavior in midge species argue against this possibility (White, 1950; Nicklas, 1959). Noteworthy in this respect is that Ortiz (see Beermann, 1956) obtained translocations of sections of K chromosomes to the S chromosomes in Memiomemus hygropenicus (Orthocladiinae). Polytene fragments of the K chromosomes were formed in some translocations of this kind. Their banding pattern had no homology with that of the S chromosomes. The other sections of the E chromosomes were completely heterochromatic (for greater detail, see Section X). In contrast, in similar translocations generated in Acricotopus lucidus, nine fragments of the K chromosomes had a banding pattern identical to that shown by the corresponding regions of the S chromosomes (32% of bands of the set) (Staiber, 1 9 9 1 ~ ) . The number of eliminated chromosomes can be large (Table 5). At anaphase of the eliminating division (at the third through seventh cleavage divisions), eight daughter chromatids continue moving to the poles, and the eliminated chromosomes remain at the position they have occupied at the early anaphase, while the S chromosomes, having reached the poles, enter the telomere (Figure 46). According to different estimates, approximately half of chromatin in Sciara copophikz (Du Bois, 1933), and 74% of DNA in Miastor sp. (Nicklas, 19591, is eliminated. In Hetmopeza pygmaea, 15% of genomic DNA is composed of the 1.716 satellite, Complementary RNA, transcribed from satellite DNA, isolated from somatic cells and not containing E chromosomes, hybridizes with centromeric heterochromatin of both types of chromosomes. Counts of silver grains established identical satellite distribution over the chromosomes of both types (Kunz and Eckhardt, 1974). It is unclear whether or not the E chromosome is enriched in any other sequences. Certain findings reveal enrichment of the eliminated chromosomes in heterochromatin. Transpositions of material from the K to the E chromosome in polytene chromosomes frequently transfer giant blocks of a-heterochromatin (see Section X). In the L chromosomes of Sciara coprophila, DNA synthesis proceeds at the end of the S phase (Rieffel and Crouse, 1966). However, the distinction between eu- and heterochromatin is rather tentative in the midges. The point is that in oogonia and spermatogonia-for example, of Trishormomyia helianthi and Miastor-specifically the somatic, and not the eliminated, chromosomes, undergo complete positive heteropycnosis (White, 1946,1950,1973). In Mikiokzfagi, all the S chromosomes in the interphase oogonial nuclei are heteropycnotic (Matuszewski, 1962). In Wachtliella persicaria, the S chromosomes are heteropycnotic in the interphase nuclei, and the E chromosomes are loose and incorporate 3H-uridine (Kunz, 1970; Kunz et al., 1970). In Mayeti& destructor, the E chromosomes are not only entirely heteropycnotic, but also stain completely for C-heterochromatin; the E chromosomes in somatic cells have only blocks ofcentromeric C-heterochromatin (Stuart and Hatchett, 1988a,b).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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Table 5 Occurrence of Germline Limited Chromosomes in Representatives of the Diptera Order

Species Cecidornyiidae Heteropeza pygmaea

Number of germline limited chromosomes

References

Up to 48

Hauschteck (1962). Camenzind (1966, 1974), Panelius (1968, 1971), Kunz and Eckhardt (1974)

Mayetiola destructor

35-45 20-35 28

Bantock (1970) Stuart and Hatchett (1988h) Bregman (1975)

Miastor sp.

30-32

Nicklas (1959)

Mikiola fagi

16

Matuszewski (1962)

Mycophila speyeri

-23

Nicklas (1960)

Oligarces paradoxus Rhabdaphaga batatas

-60 -30

Geyer-Duszynska (1961)

R . saliciperda

8

Wachtliella persicariae

More than 30

-30

Kraczkiewicz (1966) Geyer-Duszynska (1959) Kunz et al. (1970)

Chironomidae Acricotopw lucens

Reitberger (1940)

5-6

Michailova and Dimitrova (1984)

A. lucidw

17-19 4-19

Bauer and Beermann (1952) Staiber and Thudium (1986)

Cardiocladius sp.

Up to 80

Beermann (1956)

Clunio marinus

-8

Bauer and Beermann (1952)

Eucricotopus artritarsis

-9

Bauer and Beermann (1952)

E. silwestris

10-12

Bauer and Beermann (1952)

Limnophyes sp.

-8 12-26

Bauer and Beermann (1952)

M. inopinatus

1-4 6-8

Bauer and Beermann (1952)

M. sp.

2-3

Bauer and Beermann (1952)

Psectrocladius obwiw

1-4 3-5 10-14

Bauer and Beermann (1952)

Metriocnemus cavicola M. hydropeeicus

P. platypus P. rematus P. sp. Smittia parthenogenetica

-6 To 15

Trichocladiw wieipennis

5-7

Sciaridae Sciara coprophila

1-2 3

Bauer and Beermann (1952) Bauer and Beermann (1952)

Bauer and Beermann (1952) Bauer and Beermann ( 1952) Bauer and Beerrnann (1952) Bauer (1970) Bauer and Beermann (1952) Merzand Schmuck (1931), n u Bois (1933) Pavan eta/ (1975)

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F. Zhlmulev

Figure 46. Elimination of the chromosomesduring the third cleavage division in Miastor sp. (a) Anaphase. (b) Telophase. After Nicklas (1959).

D. Elimination of chromatin in infusoria The complex sexual processes of the formation of the micronucleus (Mi) and maturation of the macronucleus (Ma), controlling somatic cell functions as well as DNA elimination in somatic cells, have been described in a number of reviews (Ammermann, 1985; Steinbruck, 1986; Zhimulev, 199213).The micronuclei of all the flagellates contain germline-restricted DNA and the DNA that functions in the somatic macronucleus. The diminution of Ma takes place twice during its maturation: whole chromosomes are removed at the first diminution. Ten to 15 hr after the exconjugants have disjuncted in Stylonichiu lemnae, about 140 chromosomes (ranging from 100 to 180 in different strains) become more compact, pass to the inner membrane of the nuclear envelope, and then degenerate (Ammermann, 1985,1987). The remaining 35-36 chromosomes gradually polytenize, and interchromosomal elimination of DNA takes place a day later: single bands or groups are surrounded by membrane-like material forming the “substance” of the bands. The whole polytene chromosome breaks down into numerous independent vesicles in which a substantial part of DNA is lysed; only the genetically significant sequences remain. Telomeric DNA joins them at both ends and the Ma is con-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

89

verted into a “bag with genes.” As a result of the two diminution events, only 2% of the DNA sequences present in the Mi remain in the mature Ma (Ammermann et al., 1974). A large copy number of repetitive DNA is removed in the meantime. The DNA from Mi S. mytilw consists of 55% repeats; in contrast, repeats make up only 2% of the mature Ma (Ammermann, 1985). Chromatin elimination has been described in many other infusorian species (for review, see John and Miklos, 1979; Ammerrnann, 1985; Steinbruck, 1986; Zhimulev, 199213).

E. Physiological significance of diminution In 1904, Boveri suggested that there may be a correlation between chromation diminution and the structural features of the oocyte and sperm in ascarids. Neifakh and Timofeeva (1977) expressed the view that “the genes located in that part of DNA, which is retained in germ-line cells, are not directly involved in the control of embryogenesis, larval development and regulation of the functions of the adult organism. It appears natural to assume that these genes are involved in the processes of oogenesis and spermatogenesis.” A point of view akin to this is that of Hennig (19861, who believes that heterochromatin should be directly related to particular functions of the germline cells. Prokofyeva-Belgovskaya( 1986) assumed that the heterochromatin regions of the ascarid chromosomes may be needed only for the accomplishment of meiotic processes. However, it appeared unlikely to Akifjev and Grishanin (1993) that 50% of the genome of Cyclops and 98% of that of Stylonichia mytilus encode only functions specific to the sex cells, with the remainder encoding all the rest. Little is known also about the functions of the E chromosome. With respect to Mayetiola and Wachttielh,Bantock ( 1961) and Geyer-Duszynska (1966) showed that germline cells without E chromosomes are smaller, and they do not form gametes in the latter species. The same was detected for Trichocladius triannuhtus (Beermann, 1956). Possibly, the material of the E chromosome can restrain the somatic genes from becoming active in germline cells (Beermann, 1956).White ( 1954) believes that elimination of the E chromosome provides the contribution of a specific nucleoprotein to the formation of segmenting oocyte. In the view of Cavalier-Smith (1978), the eliminated chromosomes are needed excusively for increasing DNA amount in the cell, resulting in a sharp increase in the size of the oocyte nucleus; this, in turn, accelerates maturation of the oocyte. According to Spradling ( 1993), elimination in somatic cells might function in such a manner that repetitive DNA around the active gene regulates its activity. “The amount and nature of the interspersed and flanking repeats are proposed to regulate activity of rDNA at appropriate levels in different cell types.” Changes in the amount of repetitive DNA may result in changes of neighboring gene activity (Spradling, 1993). These are all mere hypotheses.

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VI. CENTROMERIC HETEROCHROMATIN IN POLYTENE CHROMOSOMES A. Morphological characteristics The question of the correspondence of regions of centromeric heterochromatin, identified in mitotic chromosomes, to a particular structure in polytene chromosomes is resolved in its own way for each species. Inasmuch as polytene chromosomes are not involved in cell division, the position of centromeres and, accordingly, of centromeric heterochromatin is not always easy to determine. Position of the centromeric region in polytene chromosomes can be determined according to the arm ratio in the mitotic chromosomes, the presence of bulky masses of heterochromatin and, quite rarely, on chromosomal rearrangements whose breakpoints in mitotic and polytene chromosomes have been localized (Beermann, 1962). Centromeric heterochromatin has been most accurately mapped in Dosophila because, first, localization can be aided by the use of rearrangements, and, second, heterochromatic regions of nonhomologous chromosomes pair ectopically, forming a chromocenter (Figure 47). There are chromocenters in representatives of many families of the Diptera order (for review, see Zhimulev, 1992b). In many other species, although the chromocenter does form, centromeric blocks of heterochromatin, differing in tighter compaction (stainability), nevertheless are frequently connected by long loose strands of ectopic contacts-for example, in Chironomus crmsimunus (Keyl, 1961), Ch. cucini (Martin, 1979), Ch. plumosus (Michailova, 1989a), and Anopheles species (Rishikesh, 1959; Coluzzi and Cancrini, 1971).Occasionally, these contacts are extremely infrequent, as in Ch. thumrni (Badaev et al., 1973). Cases were described in which closely related species differ in chromocenter organization: Eusimulium costaturn is homosequential with E. wemum, but the first of these species is chromocentric, whereas the last is not (Hunter, 1987). Centromeric heterochromatin is often observed to be such bulky blocks (Figure 48) that the heterochromatic nature of this material is not in doubt-for example, in Glyptotendipes barbipes (Bauer, 193613; Basrur, 1957; Michailova, 1993), Orthocladius bipunctellw (Michailova, 1982; Michailova and Belcheva, 1982), and Demeijerea rufipes (Belyanina, 1993). Moreover, these regions are also involved in chromosomal contacts (Wen et al., 1974; Belyanina, 1993). A small band, somewhat more compact than the others or positively staining for C-heterochromatin, quite frequently is mistaken for centromeric heterochromatin (Hagele, 1977a; Belyanina and Sigareva, 1978; Sigareva, 1981; Michailova, 3987a). In many cases, the chromosomes contain no particular bands to which the role of centromeric heterochromatin may be ascribed. Thus, cyto-

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logical studies did not reveal heterochromatin in the polytene chromosomes of many of the 13 studied chironomid species (Keyl, 1960) (see also Michailova, 1989a). In some instances, the position of centromeres is precisely mapped to chromosomal rearrangements, as, for example, in various species of Drosophila or Sciara copophila (Crouse, 1977). Heitz (1934) has found two types of heterochromatin. A very dense, heavily staining vacuolized body refracting light can be seen in the chromocenters (see Figure 47). Heitz has referred to it as a-heterochromatin. Two facts are relevant to the compaction degree of a-heterochromatin. When isolated nuclei of Ch. tentans are treated with salt solutions containing 20-3OmM NaCl, the chromosomes rapidly swell and lose their stainability and banding pattern, and the nucleus becomes homogeneous and structureless. Only a-heterochromatin remains heavily stained (Sass, 1980a). When a squash preparation is maintained in 55% lactic acid (removing RNA and proteins), bands become structureless. Only a-heterochromatin exhibits a condensed, heavily staining structure (Figure 49) (see also Koryakov etal., 1996; Belyaeva etal., 1997). Longterm (several years) keeping of constant preparations of Anopheles messae nurse cell polytene chromosomes stained with orcein results in almost complete loss of staining in all chromosome regions but centromeric heterochromatin (Stegniy and Sharakhova, 1990; Sharakhova et al., 1997). The reticular granular grayish formation constituting the greatest part of the chromocenter and characterized by lower compaction degree (see Figure 47) represents P-heterochromatin (Heitz, 1934). In certain species its content is very high-for example, at the base of the X chromosome of D. funebris (Tinyakov, 1936, 1965; Emmens, 1937b; Perje, 1955) and in the chromosomes of D. oiritis (Fujii, 1942) (see also Figure 47), D. psewloobscura (Bauer, 1936a), D. nebulosa (Pavan, 1946a), and D. orena (Lemeunier and Ashburner, 1984). In D. funebris, P-heterochromatin frequently establishes ectopic contacts with the euchromatic regions of the X chromosomes (Tinyakov, 1965). When these results are taken together, one concludes that there is invariably less a-than P-heterochromatin. Thus electron microscopic (EM) analysis of thin sections revealed that the area of p-heterochromatin is 1639.84 Fm2, whereas that ofa-heterochromatin is 26.48 Fm2 (Lakhotia and Jacob, 1974). According to other data, these values are 1542.06 and 61.66 pm2, respectively. The cross section of an a-heterochromatin block is usually 3-4 pm (Lakhotia, 1974a). Morphology of the chromocentre heterochromatin in the larval salivary gland chromosomes of D. hydei shows remarkable similarity with that of D. melanoguster (Lakhotia, 1974b). In some organisms, in particular Drosophila, both a-and P-heterochromatin are represented in considerably smaller amounts in the polytene than the mitotic chromosomes (Painter, 1933, 1934a,c; Heitz, 1934), that is, heterochromatic regions partially polytenize (for greater detail, see Section V1,D).

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Figure 47. a-And P-heterochromatin in D. wirilis (a and b) and D. mekmoguster in XY (c), XO (d), XYY ( e ) ,XX (0,and XXY (g). Chromosomes in c-g were kept in lactic acid ac-

cording to E. S. Belyaeva. (a) Long arrow designates a-heterochromatin; short arrows designate p-heterochromatin. (c-g) Arrows indicate a-heterochromatin. (a) after Heitz (1934); (b) reprinted by permissionfrom Gall et al. (1971); (c-g) after Koryakov et al. (1995).

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Figure 47. Continued

The chromocenter is formed at the very earliest developmental stages, and it can be seen in the nuclei of the smallest salivary gland cells amenable to examination (Frolova, 1938). The chromocenters include the centromeric a-and P-heterochromatic fragments of the chromosomes, as well as the Y chromosome (Painter and Stone, 1935; Prokofyeva-Belgovskaya, 1935a-c, 1937a,b; Kaufmann, 1937). The contribution of any given chromosome to the pericentric heterochromatin formation in polytene sets is different (Heitz, 1934, for D. oin'lis; Pavan, 1946a, for D. nebulosa). Analysis of heterochromatin in the salivary gland cells of XO, XU, and XYY males as well XX and XXY females after aging the squashes in lactic acid

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Figure 48. Pericentric heterochromatin in Chironomus melanotus (a) and Glyptot e n d i p s bnrbiper (b). C, centromere; N, nucleoli; numbers and letters designate chromosomes. Labeling of heterochromatin after application of labeled DNA on the chromosomes treated with DNase is shown in b. (a)

reprinted by permission from Hagele (1977b); (b) reprinted by permission from Schmidt and Key1 (1981).

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Figure 49. Morphology of D. melanogaster a-heterochromatin after lengthy treatment of salivary gland squashes with 50% lactic acid. ( a ) Fresh preparation of salivary gland chromosomes of bwD, containing insertion of pericentric heterochromatin (het) into the 59 region of the 2R chromosome. (h) After 5 days in 50% lactic acid at room temperature. After E. S. Belyaeva (unpublished ohservations).

showed no differences in the amount of a-heterochromatin in all genotypes examined. This indicates that the Y chromosome does not substantially contribute to the formation of a-heterochromatin (Koryakov et al., 1996). In D. pallidtpennis and D. ananassae, the chromocenter contains a microchromosome that resembles p-heterochromatin in structure (Dobzhansky, 1944; Moriwaki and Tobari, 1975). Painter and Stone’s (1935) idea that the heterochromatin of the chromocenter is not associated with individual chromosomes, but rather forms a common mass dipped into the centromeric regions of all the chromosomes, has not been supported. The Y chromosome appears as a dense rounded body composed of a-heterochromatin (Heitz, 1934; Kaufmann, 1937).The Y chromosome in the salivary gland nuclei of Phryne cincta is represented by a rounded and tightly packed formation ofthe same kind (Wolf, 1957,1970). InD. nebulosa,this chromosome cor-

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responds to a series of small, dense, compact granules (Pavan, 1946a). The hypothesis of Prokofyeva-Belgovskaya (1935b) and Frolova (1936a) that a part of the Y chromosome (in the region of the bb locus) has a banding pattern was not confirmed in later studies. In D. busckii, a small fragment of the salivary gland Y-chromosome shows a banding pattern. Its length is 1/24th that of the X chromosome, and it contains 14-15 bands. The vital genes presumably map to this element since XO males die at early developmental stages. A similar fragment is also present in the X chromosome. Presumably, these fragments are homologous to the microchromosomes of the other Drosophila species, and they are irrelevant to heterochromatin (Sirotina, 1938; Krivshenko, 1939, 1941, 1950, 1955). In polytene chromosomes, the boundaries between euchromatin and aand P-heterochromatin are rather blurred. It is not always distinctly seen where a-heterochromatin ends and P-heterochromatin starts, or where p-heterochromatin terminates and the euchromatic part proper first makes its appearance (Figure 50). Moreover, the morphology of bands at the boundary with the p-heterochromatin network is frequently abnormal (diffusion, bends, breakages, reticulation). It is noteworthy in this regard that the boundary between bands and the fbheterochromatin network can pass through a certain region in one preparation and through a region several bands distal or proximal in another preparation (I. F. Zhimulev, unpublished observations).

Figure 50. The chrornocenter in the salivary gland cells of Drosophila according to data from electron microscopy. X, 2L, 2R, chromosome numbers; thin arrows indicates the eu- and heterochromatic junction, bold arrow indicates a-heterochromatin. After Sorsa (1969).

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In an attempt to precisely map the point nearest to the heterochromatin in the D. pallidipennis X chromosome, it was found that its localization varies widely from one cell to another. For this reason, rather extensive regions of the chromosomes occasionally either show a well-defined banding pattern or appear structureless and “foamy” (Dobzhansky, 1944). The chromocenter, and correspondingly P-heterochromatin, were not observed in certain mutants of D. mehogaster: l(3)tl (Kobe1 and Breugel, 1967; Zhimulev etal., 1976) andfs 231 (King etal., 1981). According to Heino (1989), there is no chromocenter in pseudo-nurse cells of the otu mutants in D. mehogaster. However, as shown later, pericentric heterochromatin in these mutants occurs even more extensively than in salivary gland cells (Mal’ceva and Zhimulev, 1993, 1997; Mal’ceva et al., 1995; Koryakov and Zhimulev, 1995; Koryakov et al., 1996, 1997). These data also demonstrate tissue differences in the amount of pericentric heterochromatin. There are differences in chromocentric structure between larval organs. Thus there is no chromocenter in D. melanogarter midgut cells (Hochstrasser and Sedat, 1987a). It was found that, at varying temperatures, different numbers of bands are detectable in the nearest-to-heterochromatin region of chromosome 3R of D. melanogaster: 46% of the nuclei had a more compact shape at 15°C and only 20% at 25”C,whereas a more elongated base with the formation of an additional band formed in the greater part of nuclei (32%) (Lovering, 1986). Taking into account these uncertainties, it may be noted that p-heterochromatin in the polytene chromosomes of D. mehogaster is accurately mapped only to the regions proximal to certain segments: segment 20 in the X chromosome, segment 40C in chromosome 2L, segment 41D in chromosome 2R, segment 80C in chromosome 3L, segment 81F in chromosome 3R, and segment 101 in the fourth chromosome. In certain species of the Calliphoridae, Tephritidae, Glossinidae, and Sarcophagidae families, the X and Y chromosomes are not represented by polytene elements in the bristle-forming cells. There is a granular reticular mass and frequently a dense compact body instead (Figure 51). Such structures were found in Surcophagu bullata (Trepte, 1976), Lucilia cuprha (Childress, 1969; Bedo, 1982b), representatives of the Parasarcophugu genus (Srivastava et al., 1982; Kaul et al., 1983, 1989a), Ceratitis capitata (Bedo, 1986, 1987a; Zacharopoulou et al., 1991a,b), Cochliomyia hominivorax (Dev et al., 1986), Chrysomia species (Puchalla, 1994), and Dacw oka (Mavragani-Tsipidou et al., 1992; Zambetakietal., 1995). In Ceratitis capituta, the granular reticular mass is identified with the X and the dense compact body with the Y chromosome (Bedo, 1987a; Zacharopoulou, 1987; Bedo and Zacharopoulou, 1988; Kerremans et al., 1990; Zacharopoulou et al., 1991a,b; Semeshin et al., 1995). In experimentally produced XXY individuals, one of the Xs is converted into the usual banded polytene chromosome and the other remains reticular (Bedo, 1982b). Tissue specificity in the morphology of the sex chromosomes was found in this species. In trichogen cells, the

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Figure 51. A complex of heterochromatized X and Y chromosomes in the trichogen cells of Ceratitis capitam. S, Y chromosome; G, X chromosome; N, nucleolus (dotted line); C, intranucleolar chromatin. After Bedo (1987a).

Y chromosome is represented by a compact block of chromatin united with the nucleolus (see Figure 51), and the X chromosome is represented by a granular mass brightly fluorescing after staining with quinacrine. Only separate Q' granules are present in the salivary gland chromosomes (Bedo and Zacharopoulou, 1988; Semeshin et al., 1995). In Glossina morsituns and G. austeni, the X chromosome is represented by a polytene element, and the Y and B chromosomes are represented by a pycnotic reticular mass (Southern et al., 1973; Southern and Pell, 1974). A stable chromocenter of a-heterochromatin is formed in Ch. melanotus (see Figure 48) (Keyl, 1961; Hagele, 1977b; Sass, 1980b) and some Yakutian Chironomidae (Kiknadze et al., 1996). The DNA included in the chromocenter of Ch. melanotus is presumably completely polytenized (Steinemann, 1978). In thin EM sections, a-heterochromatin of this species is represented by tightly packed

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and heavily staining chromatin in which 20- to 25-nm fibrils are distinguished (Sass, 1980b), although the high degree of compaction of this material makes it doubtful whether distinction is possible. In other species, centromeric heterochromatin not included in the chromocenters can be represented by a-heterochromatin (e.g., in Glyptotendipes, Orthocladius, and certain midges; Chubareva, 1979) and p-heterochromatin (e.g., in mosquitoes; Moeur and Istock, 1982; Munstermann and Marchi, 1986). In the polytene chromosomes of bean haustorium (Phaseolus coccineus), the chromocenter is not formed; however, compact a-and loose P-heterochromatin is distinguished in the chromosomes (Brady and Clutter, 1974). Heterochromatin and the chromocenters of polytene chromosomes lie at the inner membrane of the nuclear envelope (for review, see Zhimulev, 199213). Presumably, there is an affinity of proteins involved in compaction of DNA sequences making up heterochromatin for components of the membranes.

B. Structural components of centromeric heterochromatin Differences in structure between eu- and heterochromatin were first described in 1950. After treatment of the larvae of D. virilis with ultrasound, the centromeres, which stained purple-red (like the euchromatic regions of the chromosomes), assumed an orange-red color. Forty-five minutes or later after a salivary gland preparation was made, the differences in color were abolished. The causes of differential staining of this kind are unclear (Yasuzumi and Yoshida, 1950). Great progress in studies of DNA and proteins has allowed thorough analysis of the structural organization of centromeric heterochromatin in polytene chromosomes.

1. DNA In the various species of Diptera, satellite and highly repetitive DNAs hybridize

in situ mainly with the regions of the chromocenters, and label is distributed quite uniformly (Hennig et al., 1970; Jones and Robertson, 1970; Botchan et al., 1971; Sederoff et al., 1975a; Cordeiro-Stone and Lee, 1976; Swift et al., 1978; Samols and Swift, 1979a; Chernyshev and Leibovitch, 1981; Ranganath et al., 1982; Lohe and Roberts, 1988; Stegniy and Sharakhova, 1990,1991). In in situ hybridization of DNA from single satellites or cRNA transcribed from them, it was found that chromocenters label nonuniformly. At first for D. virilis (see Figure 60) and then for many other species of the virilis group, it was demonstrated that satellite DNAs do not label the whole chromocenter, only the a-heterochromatic block, and they additionaly label one to two zones in the region of P-heterochromatin (Steinemann, 1976; Cohen and Bowman, 1979;Cohen and Kaplan, 1982; Mezzanotte et al., 1987).

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Likewise, the highest repetitive DNA in D. mehogaster is localized in a-heterochromatin (Rae, 1970; Hennig, 1973). The strictly local regions of the chromocenter label the DNA of the 1.705 satellite (Steffensenet al., 1981). Only the region of the eu- and heterochromatin junction in chromosome 3R is labeled by the DNA of the 1.672 satellite (Steffensen et al., 1981; Donnelly and Kiefer, 1986).A satellite of D. hydei (1.714) hybridizes only with a-heterochromatin, and the 1.696 satellite hybridizes with a-heterochromatin and the base of the X chromosome (Renkawitz, 1978b). What sequences is p-heterochromatin composed of? Centrifugation of the DNA from D. hydei yielded two middle repetitive fractions, one of which (1.702 in a CsCl density gradient) hybridized with the heterochromatin of virtually all the chromosomes,with the exception of the X. This DNA underwent polytenization. The other fraction (1.697) hybridized with the chromocenter and the nucleolar organizer (Renkawitz, 1978a). In D. simulans, the 1.696 satellite uniformly labels the whole chromocenter and, presumably, the base of the arms of all the chromosomes (Appels and Peacock, 1978). Of the middle repeats hybridizing with p-heterochromatin, mobile genetic elements (MGE) play the most important role. In Ashburner’s laboratory handbook, there is information that more than 40 families of the MGE have been identified in Drosophih. Their size varies in the 0.3-8.8 kb range, and a duplication from 4 to 12-14 bp is formed in the insertion sites in chromosomal DNA; copy number in the genomes varies from 0-6 to 50-100 in different strains (Ashburner, 1989). The portion of the genome occupied by the MGE is about 15,000 kb (Ashbumer, 1989), or approximately 10%of the genome (Spradling and Rubin, 1981). A schematic representation of these elements is given in Figure 52. The organizational features of the MGE are considered in greater detail in a number of reviews (Ilyin, 1982; Ananiev, 1984; Finnegan and Fawsett, 1986; Ashbumer, 1989; Blackman and Gelbart, 1989; Bingham and Zachar, 1989; Engels, 1989; Finnegan, 1989; Louis and Yannopoulos, 1991; Busseau et al., 1994; Coen et al., 1994; DiNocera et al., 1994; Kidwell, 1994; Ladeveze et d., 1994; Periquet et al., 1994). In situ hybridization analysis demonstrated that various MGEs, besides many other sites, in the euchromaticregions of the chromosomesof D. melanogmter heavily label chromocenters (Figure 53), as well as the nearest chromosome regions, which are suggested to be p-heterochromatic (regions 20,40,41, 80, and 81) (Wensink et al., 1974; Young, 1979; Kidd and Glover, 1980; Pardue and Dawid, 1981; Dowsett and Young, 1982; Ananiev et al., 1984a; Ananiev and Barsky, 1985; Bucheton et al., 1986; Biemont and Gautier, 1988; Kholodilov et d., 1988; Charlesworth et al., 1994; Coelho et al., 1996). According to Ananiev and Barsky (1985), who have studied the localization of many MGE families, a total of 11%of all the copies of mobile elements

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation +-+

+

+

-7200bp

-----------

101

MDG-1

+++ .--$

copia H.M.S. Beagle

+

Figure 52. A scheme of the structure of mobile genetic elements. 1, Chromosomal DNA; 2, duplication of the chromosome region in the integration site of the mobile element; 3, internal region of the mobile element; 4,direct long terminal repeats; 5, inverted repetitive sequences. After Ananiev (1984).

are assigned to the chromosome regions closely adjacent to the chromocenter (regions 20AB, 4OABC, 41AB, 80ABC, 8lEE and 101EF) (see also Ananiev et al., 1984a-c). They have localized 16 mobile elements in the 20,40, and 41AB regions. p-Heterochromatin labels heavily when DNA from the Stalker genetic element is used as a probe (Georgiev et al., 1990; Simonova, 1992). In the in situ localization experiments of one of the DNA clones isolated, presumably, from the Drosophita Y chromosome (Lifschytz and Hareven, 1982b) in the polytene chromosomes, the following features were identified:

The clone hybridizes with the chromocenter and the regions of the arm base of all the chromosomes showing the usual banding pattern (20AF, 39E-40F, 41AE 80AC, and 81F). The entire fourth chromosome labels. Of the euchromatic regions, the 12E1-2 band remains consistently labeled in all the studied strains. Based on sequence data, it was concluded that the clone includes a mobile element called the “1360 element” (Kholodilov et al., 1987, 1988).

Figure 53. In situ hybridization of element 1360 to polytene chromosomes of gt/gtf37larvae of D. melanogaster. (a) Labeling of chromosomal bases (designated with figures and letters), the chromocenter (cc), chromosome 4, and some sites on the third chromosome. (b) Multiple sites of labeling on chromosome 4.(c-e) Labeled ectopic stretches (arrows)to unlabeled IH region 11A6-9, from labeled regions 12E (c)and 40A-F (d) and from labeled sites on chromosome 4 (e) to the laheled chromocenter (cc). Bar represents3 km (a, c-e) and 1 km (b). Reprinted by permission from Kholodilov et al.

(1988).

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These data provide evidence for homology of P-heterochromatin of all the chromosome arms and the whole fourth chromosome because all have sites of 1360 mobile element. The presence of the 1360 element (or hopel, according to Kurenova et al., 1990; Leibovitch et al., 1991) in repeated units of Ste and Su(Ste) (Balakireva et al., 1992) may be the reason why the euchromatic region 12E1-2, where Ste is located, is labeled (Palumbo et al., 199413). Microdissection and microcloning of section 20 of the X chromosome revealed that 60% of the generated clones contain repetitive elements, when the 19EF region was cloned, 40% contained these elements. The size of the inserts was, on average, 4.5 kb. These repeats showed homology neither with satellite DNA nor with scrambled repeats. In in situ hybridization of one of the clones (Dr.D),as with 1360,the whole chromocenter, the 20AF, 40DF, 4lAF, 80AC, and 81F regions, and the whole fourth chromosome labeled (Miklos et al., 1988). These results provide evidence for the high homology of the chromosome regions composing P-heterochromatin. Several clones taken from microdissection library of the region 20 exist as a 12.5 kb island of unique DNA surrounded by repetitive DNA (Andrewset al., 1996). As for the other species, P-element insertions are mainly restricted to the heterochromatin in four Drosophila species of the willistoni-salturn group (Lansman et al., 1985; Daniels and Strausbaugh, 1986). In D. wirilis, in situ hybridization of clone v68-52 was shown primarily with the chromocenter and the base of chromosome 6 (Lozovskayaet al., 1993). A repetitive clone 2 labels about 60 sites, as well as the telomeric regions and the chromocenter in D. subobscura (Felger and Sperlich, 1989). Insertions of the first type into the 28s rRNA gene are located mainly in the chromocenter in Drosophila. The flanking sequences, isolated and cloned together with the insertions, are the mobile elements themselves (Pardue and Daw-

id, 1981). Data on cloning of the I-element in Drosophila, which is mainly located in the chromocenter, also favor the saturation of p-heterochromatin with mobile elements. I t was found that the DNA sequences flanking the normal I-element are defective, being immobilized copies from representatives of families such as mdg4,297, i 731F, and Doc (Crozatier et al., 1988; Vaury etal., 1989, 1990, 1994). Presumably, the immobilized m d g l elements are located in p-heterochromatin because their copy number is reduced in the DNA of salivary glands. This is indicative of their incomplete polytenization and their resulting disposition in heterochromatin (Shevelyov et al., 1987, 1989). I-elements are located in the pericentric heterochromatin in all the melanogaster subgroup species, in the other melanogaster group species, and in Scaptomyza pallida (Drosophilidae) (Bucheton et al., 1986; Simonelig et al., 1988). In D. subobscura, P-element located in p-heterochromatin is degraded (Paricio et al., 1991, 1994).

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The distal part of the X chromosome heterochromatin contains repeated SCLR elements (Shevelyov, 1992; Nurminsky et al., 1994); scrambled structures built from middle repetitive DNAs, such as copia-like elements (aurora, mdglhet, GATE) (DiNocera et al., 1986; Nurminsky, 1993; Shevelyov, 1993); LINE (G-element, type I rDNA insertions) (Roiha et al., 1981; DiNocera et al., 1986;Jakubczaket al., 1990); and Stellate (Shevelyov, 1992). The first three groups of DNA elements are immobilized in heterochromatin only (DiNocera and Dawid, 1983; Shevelyov et al., 1989; Shevelyov, 1993). A relatively new family of M repeats has been detected in D. melanogaster. These fragments are 200-3000 bp, representing three to four families arranged in blocks with one representative of each family. The blocks have mobile element mdgl at each flank. The latter occasionally inserts into one of the M sequences. The primary structure of the M sequences shows no considerable homology with the known sequences; it contains neither an extensive reading frame nor motifs characteristic of the promoters of RNA polymerase 11. The clusters of M sequences do not polytenize to any great extent (Balakireva et al., 1988). One may suggest that heterochromatin could function either as a trap where transposing elements are immobilized and start to degenerate, or as a chromosome region containing insertional hot spots (Lansman et al., 1987; Bucheton et al., 1992). The above data allow one to share the view of Miklos et al. (1988) that “the bulk of the p-heterochromatin is largely the end product of the insertion, deletion, and amplification of mobile elements into each other, as well as into regions containing unique sequences.” It is unclear, however, to what extent the mobile elements can be the structural components of (3-heterochromatin because of their varying localization. Comparisons of the genomes of D. mehnogaster with those of the sibling species D. simulans revealed that the former contain approximately three times more middle repeats than the latter (Dowsett and Young, 1982). A relevant finding is that a small number of the MGE families detected in D. mehmgaster, occur in other subspecies of the melanogaster subgroup, while D. erecta virtually lacks such elements. However, this species has many other MGE families unidentified in D. melanogaster (Dowsett, 1983). In addition to the mobile elements, labeling sites of some other repeats were found in pericentric heterochromatin. D. melanogaster HeT element is found exclusively in telomeric regions, in proximal heterochromatin, and on the Y chromosome (Traverse and Pardue, 1988; Valgeirsdottir et al., 1990; Danilevskaya et al., 1992, 1993). A middle repeat, present as 200 copies in the D. melanogaster genome in DNA fragments isolated from nuclear lamina, was located in P-het. erochromatin (Baiborodin et al., 1993a,b; Sharakhov et al., 1993, 1997; Baricheva et al., 1995, 1996; Sharakhov, 1995; Bogachev et al., 1996). Unknown repeats were found on squash preparations after digestion with A h I and Hae 111 restric-

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tases (Mezzanotte, 1986).In D. simulans, the insertion of the ribosomal genes of the second type is mapped in the heterochromatin of the 80BC region (Mecheva, 1990). Somewhat distal to P-heterochromatin in the 19E8 region, which already shows a clear-cut banding pattern (the locus of the uncoordinated gene) as a consequence of “walking,”an approximately 60-kb DNA was cloned. The cloned sequences fall distinctly into two classes. The first class includes sequences with small copy number, or that are unique and conserved to a great extent, which is revealed by comparing various strains. The second class is composed of tandemly repetitive units, which are polymorphic both within and between strains. The repetitive unit is separated by insertions of the first type in the ribosomal cistron (Healy et al., 1988). In the 19F-20AB region of the X chromosome of Drosophila, there are sequences homologous to satellite DNAs (1.709) from the C- heterochromatic part of the sex chromosomes of snakes (Singh et al., 1981). The structural unit of the repeat is tetranucleotide GATA. It has a potential open reading frame: three copies of the nucleotide yield four amino acids, Ile-Ser-Leu-Tyr (Singh et al., 1984; Simpson, 1990). In species in whose polytene chromosomes chromocenters are not formed, centromeric heterochromatin is also rich in highly repetitive sequences, for example, in Rhynchosciara americana (Eckhardt and Gall, 1971; Machado-Santelli et al., 1979), Chironomus thummi thummi (Schmidt et al., 1980; Zainiev et at., 1986; see also Section VII,C,6), Glyptotendipes barbipes (Schmidt, 1980), Sciara coprophila (Abbott et al., 1981) and Anopheles messeae (Sharakhova, 1997; Sharakhova et al., 1997).

2. Proteins Using a modified Giemsa staining technique (N-staining), which includes sequential removal of DNA, RNA, and acid-soluble proteins from the chromosomes, C+-banding is, nevertheless, identified at sites where C-staining for constitutive heterochromatin is detected (Matsui and Sasaki, 1973; Matsui, 1974). The nonrandom distribution of nonhistone proteins presumably causes such staining. In the salivary gland nuclei of Drosophila, a-heterochromatin in the chromocenter stains positively with this stain (Matsui, 1974; Hagele, 1977b). When one of the treatments is omitted (tetrachloracetic acid or HCl), the puffs intercalary heterochromatic bands 74A and 75C as well as the stain. However, when treatment with these acids continues for a sufficient span of time, only a-heterochromatin shows N- banding (Hagele, 1977b). It is tempting to suggest that C+-banding or large heterochromatic blocks (Hagele, 1977a; Belyanina and Sigareva, 1978; Sigareva, 1981; Michailova and Belcheva, 1982; Recco-Pimentel and Mello, 1986) may be due to the presence of specific proteins.

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However, in three species of Chironomus (Ch. melanotus, Ch. thummi thumrni, and Ch. th. piger), positive N-staining was detected in the active regions: puffs, Balbiani rings, and nucleoli. The loose heterochromatic chromocenters of Ch. melanotus were completely unstained (Hagele, 197713). Definitive evidence has been provided to demonstrate that the centromeric regions differ in particular protein composition from euchromatin. HI histones included in nucleosomes having satellite DN As are phosphorylated. Certain types of nucleosomes possessing the 1.672 satellite (AATAT) contain protein D1 (see Section IV). Furthermore, the molecules of histone H1 are packaged in a specific manner. Thus, in Glyptotendipes barbipes, histone HI in the giant blocks of centromeric heterochromatin is not identified after application of antibodies to this protein in squash preparations. However, the results of electrophoresis of histones isolated microsurgically from centromeric proteins show that this protein is normally represented (Nonchev et al., 1989). It has been reported that the Cla-repeat-containing centromeres of Ch. th. thummi (see Section VII,C,B) are rich in a particular fraction of histone, HI11, identified with the use of monoclonal antibodies (Mohr et al., 1989; Schulze et al., 1993). Antibodies to HI-I1 were not detected in homologous regions of Ch. th. piger, where these repeats are not identified. A particular set of nonhistone proteins has been identified, in addition to modification of histones in chromocenters. A comparison of protein patterns yielded by the embryonic diploid cells and salivary glands of D. hydei revealed that the latter lack fractions psi, lambda, and kappa3. Presumably, they bind to heterochromatin, which is underreplicated in salivary gland cells (Elgin and Boyd, 1975). Of the nonhistone proteins identified, one with a molecular mass of 38 kDa lies in the chromocenters of Dosophila melanogaster and several intercalary sites. It is noteworthy that the largest amount of protein was in a-heterochromatin (Will and Bautz, 1980). The heterochromatin-specificprotein with a molecular mass of 19 kDa, HP1 (heterochromatic protein), is located in the chromocenters (a-and p-heterochromatin) and also at the bases of the chromosome arms (the 41AF and 80AC regions), in the whole fourth chromosome, and in several other regions (Figure 54) (see Powers and Eissenberg, 1993; Zhimulev, 1996; and Sections XIII,F, and XVII,A, for references). Another protein binding to polytene heterochromatin is Mod, encoded by the Su(var)moduEo gene (Garzino et d., 1992) (see Section XII1,F). There is great affinity between nonhistone proteins of heterochromatin and DNA, and the DNA-protein complex dissociates with difficulty (Hsieh and Brutlag, 1979b; Will and Bautz, 1980; James et al., 1989). This property is presumably retained in vitro, too. In any event, the satellite DNA from the giant heterochromatic proteins of Glyptotendipes barbipes hybridizes with the protein com-

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Figure 54. Location of antibodies on HPl protein (right) in the polytene chromosome of D. melanogaster (left). Labeling of chromocenters in squashed and unsquashed nuclei. (Courtesy of C. Craig and S. C. R. Elgin).

ponent of these blocks (see Figure 48). It should be noted that t3H]DNAbinds to protein only under the condition that chromosomal DNA is digested with DNase I (Schmidt and Keyl, 1981).

3. Structural changes in heterochromatin Several instances are known in which sharp changes in the structure (compaction degree) of chromatin are related to environmental conditions. According to Stich and Naylor (1958), when the larvae of Glyptotendipes barbipes are grown at 6-8"C, no heterochromatic blocks are formed; puffs develop in these chromosomal regions instead. Two to 3 days after transfer of the larvae to a temperature of 18°C

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large heterochromatic blocks are formed in the puffed regions. This is associated with a 10-fold increase in DNA amount. Key1 (1963) observed that, in this species, heterochromatin is expressed as a puff at 18°C and as a compact block at 6°C (Figure 55); there is no associated change in DNA amount during the transition. Puffs in the centromeric heterochromatic blocks of Glyptotendipes burbipes are inducible by heat shock (37°C) and x-ray irradiation (10,000 rad). RNA synthesis was not demonstrated in the formed puffs (Walter, 1973). Puffs arise in the centromeric heterochromatic blocks in Ch. th. thummi (Figure 56) after prolonged maintenance of larvae in a solution of actinomycin D (1-50 p,g/ml) or a-amanitin (for 24 hr and longer) (Kiknadze, 1965,1972; Valeyeva et al., 1979, 1984; Perov and Kiknadze, 1980; Kiknadze and Valeyeva, 1983; Kiknadze et ul., 1983; Valeyeva, 1991). The degree of inducibility is high: 82-100% of glands have puffed chromosomes in the centromeric regions at all developmental stages, except for a 24-hr period after the molt (19%). The percentage of chromosomes with puffs varies from 20% to 91% (with the exception of the 24-hr stage, when it is 9%) (Kiknadze and Valeyeva, 1983). These puffs do not incorporate [3H]uridine (Valeyeva et al., 1979; Kiknadze and Valeyeva, 1983), nor do they bind antibodies to DNA-RNA hybrids

Figure 55. Changes in the morphology of the heterochromatic block in the second chromosome of Glyptotendipes barbipes at different temperatures.

(a) pl appears as a Balbiani ring (18°C).(b) Transitional state at 18°C. (c) p l appears as a dense block (6°C). After Key1 (1963).

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Figure 56. Pseudopuffs (indicated by arrows) arising in blocks ofcentromeric heterochromatin of Ch. th. thummi under long-term inhibition of RNA synthesis by actinomycin D. After Valeyeva (1991).

(Valeyeva, 1991), and this makes their suggested transcriptional activity uncredible. At the electron microscopic level, a specific ultrastructure is detected when these puffs are induced; the material separates into two components in the centromeric region: an electron-dense component similar to band material and a thin fibrillar component (Perov and Kiknadze, 1980). Puffs are formed i n c h . th. thummi only, in hybrids between this species and Ch. th. piger. For this reason, proof of puff formation relies on the heterochromatin amount in a block. No preferential binding of actinomycin D to the heterochromatin of this species was detected (Valeyeva et al., 1984; Stepanova et al., 1985). The suggestion (Kiknadze, 1965) that puffs arise as a result of impaired RNA transport is likely in error (Valeyeva et al., 1979), because puffs develop in a background of completely inhibited RNA synthesis. Presumably, the antibiotic inhibits the synthesis of proteins involved in heterochromatin compaction and, as a consequence, heterochromatin becomes decompacted.

C. Variation in the amount of centromeric heterochromatin A distinguishing feature of heterochromatin is variation in its amount (see also Section 11,F). The variability in polytene chromosomes is manifested as incom-

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plete polytenization of different degrees (underrepresentation) or increased number of copies (amplification) (for review, see Pavan and da Cuhna, 1969; Rudkin, 1969, 1972, 1973; Berendes, 1973; Steinemann, 1978; Richards, 1985; Korge, 1987; Spradling, 1987; Spradling and Orr-Weaver, 1987). After the discovery of polytene chromosomes, it became clear that not all regions polytenize to the same degree. Painter (1933, 1934c), Heitz (1934), Hinton (1942b), and Hannah (1951) noted that the giant fragments of heterochromatin seen in the mitotic chromosome are, in fact, not present in the polytene chromosome of Drosophila. It was a widely held notion that the Drosophila Y chromosome contributes only a little material to the chromocenter (Bridges, 193%; ProkofyevaBelgovskaya, 1935a-c, 1937a, Pavan, 1946a,b), and it is therefore almost come pletely underrepresented. Analysis of flanking DNA after insertion of P-elements into the Y chromosome has shown that all nine Y chromosome insertion sites were underrepresented more than 20-fold in Southern blots of polytene DNA (Zhang and Spradling, 1994, 1995). One arm of chromosome I is not represented in polytene elements in the salivary gland, in larval malpighian tubule cells, or in imago ovarian nurse cells of mosquitoes of the Anopheles genus (Kitzmiller and Baker, 1963; Coluzzi and Kitzmiller, 1975;Stegniy, 1981). In Prodiumesa olioacea, only a part of an arm polytenizes in females, and it is completely underrepresented in males (Zacharias, 1979, 1981). In another chironomid, Pseudodiumesu brunickii, the fourth, nucleolus-forming chromosome is underrepresented. It contains 11.4% of the DNA in metaphase chromosomes and 3.7% in polytene chromosomes (Zacharias, 1984). An interesting case of underrepresentation was found during polytenization of one of the arms of the fourth chromosome of D. rnelanoguster. It has long been recognized that this is a small, one-armed chromosome with 25-50 bands in salivary gland cells (Bridges, 1935c), which starts with the lOlD dense band from the chromocenter (Figure 57). Later, on the basis of genetic data, the conclusion was made that there exists a second arm of the fourth chromosome, not completely polytenized in the salivary gland cells (Panshin and Khvostova, 1938;Griffen and Stone, 1939a, 1940b). Roberts ( 1969a, 1972b,c) generated several remarkable translocations between the third and fourth chromosomes, a break in which was located in the nonpolytenized part of the fourth chromosome proximal to the lOlD band. As a result of transfer of this part of the chromosome from the neighborhood of heterochromatin, additional bands, located to the left of IOID, became visible: 10-12 dark bands in the T(3;4)10 translocation (see Figure 57) and a small light region in T(3;4)83 that even occasionally formed an enormous puff. The lOlC band appeared in another translocation, T(2;4)14. It may be suggested that the fourth chromosome consists of three regions (see Figure 57):

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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1. Heterochromatin, owing to which the dot fourth chromosome in mitotic spreads stains for C+-heterochromatin. It replicates late and contains satellite DNA. 2. The region lying between lOlA and IOlD, that is, presumably between the centromere and the normally polytenizing part. This region seems to underreplicate under the usual conditions because, even in EM sections, it is not seen immersed in the chromocenter. 3. The euchromatic part proper (see Figure 57). Transfer of the 101A-101D region from the vicinity of heterochromatin to euchromatin makes normal replication possible. It is unclear whether the centromere is located in lOlA or 101D. One arm or one and one-half of the fourth chromosome underreplicates depending on its location. Cases in which heterochromatin amount varies widely in the mitotic chromosomes are known. However, the variants do not affect the sizes of the replicating parts of the polytene chromosome. For example, there is three times less heterochromatin in the D. ezoana than the D. virilis genome; however, the sizes of the banded parts of the polytene chromosome, and even the banding patterns (with the exception of inversions), are the same in these species (Holmquist,

1975a). In 14 species of Hawaiian Drosophila, the heterochromatin amount is so enormous that they have become the largest chromosome of the karyotype in several cases. This was not due to change in banding pattern (Yoon and Richardson,

197813). There are well-known instances of underrepresentation of whole chromosomes. For example, the B (see Section IX) or the Y chromosomes of Drosophila and Phryne cincta (Sokoloff and Zacharias, 1977) and of the mosquitoes of the Anopheles genus (Coluzzi and Kitzmiller, 1975) frequently do not polytenize. In D. pallidipennis, five pairs of rodlike chromosomes, a pair of dot microchromosomes, and a pair of giant V-shaped chromosomes the length of which is almost that of the total length of all the other chromosomes are identified in mitotic spreads. Thus the detection of seven large polytene elements would be expected in the salivary gland nuclei. However, there are only five elements; the giant V-shaped chromosome is presumably underrepresented (Dobzhansky, 1944). In D. nasutoides, the fourth chromosome pair contains 65% of all genomic DNA, but it is heteropycnotic and entirely unpolytenized (Zacharias, 1990). In flies of the Calliphoridae, Tephritidae, Glossinidae, and Sarcophagidae families, differences in polytenization degree of the sex chromosome were revealed. The pertinent data are controversial. It has been reported that the X and Y chromosomes are represented by polytene elements in the salivary gland nuclei or bristle-forming cells (Sharma et al., 1976; Handa et al., 1980, 1981). They are

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nonpolytenized, as may be judged from other data (Bultmann, Clever, 1969; Clever et al., 1969; Whitten, 1969; Samols and Swift, 1979a; Kaul and Tewari, 1978; Perkins et al., 1992). Cytophotometric evidence indicates that the same amount of DNA is retained in the heterochromatin of polytene chromosomes as is contained in the diploid nuclei of Drosophila (i.e., 2C-4C) (Rudkin and Schultz, 1961; Rudkin, 1965a,b, 1969; Berendes and KeyI, 1967; Mulder et al., 1968; Zacharias, 1986, 1990,1993),while the polytenization level of euchromatin is 1000-2OOOC. As determined by Southern blotting, restriction fragments comprising mainly centromeric heterochromatin near the euchromatin-heterochromatin breakpoint of the Dp 1187 were drastically reduced (more than 50-fold) in salivary gland and ovarian DNA (Karpenand Spradling, 1990; Spradling et al., 1992). Furthermore, euchromatic sequences even 50 kb or 100 kb from the breakpoint were still underrepresented.

Figure 57. Structure of the fourth microchromosome of D. mefumgaster. (A) Electron microscopic section of a normal chromosome. (B)T(3;4)10.(C and D)T(3;4)83 with puff (shown by arrow in C) and without puff (D). (E) A scheme of the location of nonpolytenizing C+-heterochromatin (1 ) and polytenizing euchromatin ( 3 ) and a region (2) polytenizing in translocations. Numbers and letters designate chromosome regions. (A) after V. E Semeshin (unpublished observations); ( E D ) after Roberts (1972~);(E)after I. E Zhimulev (unpublishedobservations).

2 Figure 57. Continued

3

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-

Based on cytophotometric analysis of Feulgen-stained chromosomes, Dennhofer (1981, 1982a,b) obtained data indicating that all the chromosome regions are polytenized to the same degree. Later, Lamb (1982) found that adult malpighian tubules and midgut nuclei contain DNA doublings of the diploid value during replication. This finding conflicts with the known facts, and is difficult to interpret. It is believed that the three components of the mitotic chromosomethe bulk of the heterochromatin (or a-heterochromatin), the euchromatin, and the P-heterochromatin lying between them (see Figure 64)-behave differently during polytenization: euchromatin completely replicates, a-heterochromatin is represented by the same amount as in the mitotic chromosome, and P-heterochromatin partly replicates (Heitz, 1934; Gall, 1973). The highly repetitive DNAs are strongly underrepresented in the polytene chromosomes of the salivary glands of many dipteran species (Hennig et d., 1970; JonesandRobertson, 1970;Rae, 1970;Botchanetul., 1971; Dicksonetal., 1971;Gall et d., 1971; Blumenfeld and Forrest, 1972;Hennig, 1972a,b;Balsamoet d., 1973a-c; Cordeiro et d., 1975; Spear, 1977; Wollenzien et al., 1977; Peacock et d., 1978; Steinemann, 1978; Miklos and Gill, 1981; Redfem, 1981a; Nazimiec and Beckingham, 1986,1989; Perkins et d., 1992). In the polyploid nurse cells of D. mehnoguster satellite sequence replicate only partially (Hammond and Laird, 1985a). Figure 58 presents an illustrative pattern of the sedimentation profiles of DNA fragments in diploid and polytene tissues. The various satellites in the various tissues underreplicate differently (Figure 59). Comparisons of chromatin composition in the nuclei of salivary glands and embryos demonstrated that several fractions of nonhistone protein are missing in polytene nuclei. These proteins are possibly important components of underrepresented constitutive heterochromatin (Elgin and Boyd, 1975). In situ hybridization experiments have shown that the amount of satellite DNA in diploid cells and a-heterochromatic blocks in the polytene chromosomes of D. virilis is virtually the same (Figure 60, see also Peacock et ul., 1974). This has led Gall et al. (1971) to the conclusion that a-and P-heterochromatin are differentially polytenized. While the amount of a-heterochromatin remained at 2 4 2 , P-heterochromatin polytenized more effectively. The following facts support this conclusion: 1. In an EM autoradiographic study after pulse incorporation of [3H]thymidine, there was fivefold less label per unit area in a-than P-heterochromatin. This is evidence of decreased involvement of the former in replication and of continuing replication in the latter (Lakhotia, 1974a). 2. After treatment of the salivary glands of D. nasuta with 5-bromodeoxyuridine (5-BrdU) during the last one to two replication cycles, only a-heterochromatin (the Hoechst 33258 stain at pH 7.0) fluoresces. Inasmuch as 5-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation M 1.702

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I

Figure 58. Profiles of DNA distribution in D. m u toides in a CsCl density gradient isolated from the cells of brain (a), adult nuclei (b), embryonic nuclei (c), and larval salivary glands (d). M, major fraction; Roman and Arabic numerals designate satellite numbers. Reprinted by permission from Cordeiro et al. (1975).

BrdU can substitute for DNA during replication and H-fluorescence is quenched as a consequence, fluorescence shows that a-heterochromatin is not involved at least during the last replication cycles (Kumar and Lakhotia, 1977). Similar results were obtained in other species of Drosophilu (Lakhotia and Mishra, 1980). 3. In D. muta, the Hoechst 33258-staining a-heterochromatin block is virtually of the same size in the nonpolytene cells of imaginal disc salivary glands and the polytene cells of larval salivary glands and malpighian tubules (Lakhotia, 1984). In the case of long-term incorporation (46 hr) of [3H]thymidine into the cells of imaginal discs, approximately 100% of interphases label; however, the chromocenters remain unlabeled in approximately 50% of nuclei (Lakhotia, 1981).

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Thoracic

Midgut

Molpighion Tubulr

AJL Adult Ovary

Broin

Wholr Adult Pupal Ovary

Figure 59. Representation of satellites in various tissues of adult flies and pupae of D. vinlis. Reprinted by permission from Endow and Gall (1975).

There are data mainly confirming the idea that there is no polytenization in a-heterochromatin, yet they are somewhat at variance from the following facts: (1 ) the areas of the chromocenters in the interphase nuclei are larger in neuroblast cells of third instar larvae than in 6-hr embryos of D. muta (Lakhotia and Kumar, 1980); and (2) t3H]thymidineis incorporated into the chromocenter in some cases (Plaut, 1963, 1968; Swift, 1964), although it is possibly incorporated into the DNA of the ribosomal genes included in heterochromatin. The DNAs enriched in transposing elements and composingmost P-hetetochromatin are underrepresented in the Drosophih mekmogaster polytene chtomosomes as well. Nonmobile heterochromatic mdgl her, SCLR, and aurora are underrepresented in the DNA of salivary gland cells compared with those isolated from flies (Shevelyov et al., 1989; Shevelyov, 1992, 1993).The repeats of the 18s and 28s rRNA genes, which are included in the heterochromatin of the mitotic chromosomes, are considerably underrepresented in salivary gland cells, in addition to satellites. The DNA content of the genes in the highly polytenized cells is 20-28%

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Figure 60. Location of satellite DNA in chromocenters of polytene chromosomes ( a x , e ) and diploid cells of salivary glands (d) and larvae of D. virilis. Reprinted by permission from Gall et al. (1971).

of that in diploid cells (Hennig and Meer, 1971; Spear and Gall, 1973; Gambari-

ni and Lara, 1974; Spear, 1974; Mohan, 1977;Jacobs-Lorena, 1980; Sinibaldi and Cummings, 1981; Lifschytz, 1983; Beckingham and Rubacha, 1984). This is associated with polytenization of the ribosomal DNA of only one of the two X chromosome in females and of only the X chromosome in males (Spear and Gall, 1973; Endow and Glover, 1979; Endow, 1980). The genes, which contain insertions of element I, are mainly underrepresented (Endow and Glover, 1979; Kunz et al., 1981a,b; Beckingham and Thompson, 1982; Kalumuck et al., 1990). Figure 61 is a scheme of the polytenization levels in euchromatin ( I ) , the nucleolar organizers (2), heterochromatin (3), and the telomeres (4). In Ch. tenturn, the location

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1

3

r-

1

2

3

Figure 61. A scheme of polytenization levels in the euchromatic part of the chromosome (11, in the region of the nucleolar organizer (2), in centromeric heterochromatin ( 3 ) , and presumably in the telomeric region (4). After Laird et al. (1974).

site of rRNA genes shows no features of heterochromatin and rDNA is proportionately represented in polytene chromosomes (Hollenberg, 1976). As for unique DNA, which is present in heterochromatin, at least several “hetetochromatic” (It) genes are highly represented in salivary gland cells (Devlin et al., 1990b). The DNA su(f) locus is 80% polytenized with respect to euchromatin (Yamamoto et al., 1990). It has been shown at the cytological and molecular levels that the proximal heterochtomatic gene rolled (see Figure 33) undergoes significant replication during polytenization with respect to the Ban-1 or AAGAG repeats, which are thought to be as underrepresented (Berghella and Dimitri, 1996). The ci gene located near the edge of the mitotic heterochromatin shows a normal level of in situ hybridization in salivary gland chromosomes (Locke and Tartof, 1994). Analysis of DNA sequences flanking 16 P-element insertions into mitotic heterochromatin has shown that in situ hybridization signals in the polytene chromosomes were observed in all of these lines. Nine tested insertion sites in autosomal heterochromatin were represented fully in salivary gland DNA. Moreover, P-elements themselves, after insertion into autosomal heterochromatin, become highly polytenized and form part of the chromocenter. Insertions form unusual shapes for chromocenter structures: bands, clusters of dots, and split bands (Zhang and Spradling, 1995). The accepted model implies that heterochromatin underrepresentation is due to underreplication of heterochromatic DNA (Gall, 1973; Rudkin, 1969). However, more recent experiments suggest that heterochromatin reduction results from DNA elimination rather than underreplication (Karpen and Spradling, 1990; Glaser et al., 1992; Spradling et al., 1992). Repeats behave differently during polytenizaton of the chromosomes of Chironomus. Satellite DNA replicates completely in all six studied species (Ch.

plumosus, Ch. nuditarsis, Ch. th. thummi, Ch. melanotus, Camptochironomus pallidivittatus, and Glyptotendipes barbipes) (Walter, 1973; Steinemann, 1978; Schmidt, 1980), and this seems to explain why there exist large heterochromatin blocks in centromeric regions. In Ch. tentans, the tRNA genes are located in euchromatin of the second chromosome and they replicate proportionately through the entire genome (Hollenberg, 1976).

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In giant cells of mammalian trophoblast containing cryptic polytene chromosomes, satellites also completely replicate (Sherman et al., 1972; Barlow and Sherman, 1974). In plant polytene chromosomes, differential DNA replication may include underrepresentation of pericentric heterochromatin and extra replication of another portion of the genome (see Nagl, 1982a,b, for review). Inasmuch as germline cells execute particular functions, information regarding the replication of repeats in polyploid ovarian nurse cells should be considered separately. It was an early observation that they contain more heterochromatin (Schultz, 1965). The data obtained using molecular biological methods show that underrepresentation is expressed more weakly in these cells. In a number of cases, such as in salivary gland cells, whole chromosomes or their arms underreplicate-for example, the Y chromosome of Drosophih (Schultz, 1956)or the chromosomes ofcerodonta dorsalis (Stalker, 1954),Anopheks stephensi (Redfern, 1981a), and Prodiumesa olivacea (Zacharias, 1979). The chromocenters do not increase in size with increasing polyploidy degree (e.g., in Mwca domestica; Bier, 1961. [jH]thymidine is not actually incorporated into the heterochromatic blocks of the ovarian nurse cells of Brudysia hygida (Sauaia and Alves, 1969). According to cytophotometric data, in females of the wild-type and the 0tu7 mutant of D. mehogaster, DNA content is close to 1024C or 2048C in the nuclei of ovarian nurse cells, although it corresponds to the expected amount provided that replication is complete within the first six replication cycles (Rasch et al., 1984).In the subsequent cycles, DNA content no longer conforms to the expected values; it becomes smaller. It is believed that 10-30% of the genome can underreplicate (Mulligan et al., 1981; Rasch et al., 1984; Hammond and Laird, 1985b; Mulligan and Rasch, 1985). According to other data, underrepresentation in nurse cells occurs neither in strains of the wild type nor in mutants; moreover, overreplication was detected in strains such as dic, spiC, and gt (Bohrmann er al., 1986). The underrepresentation degree of the 1.705 satellite is lower than in the salivary glands in D. mehogaster. Its amount remains at the level of three or four replications in the mitotic cells (Hammond and Laird, 1985a,b). A 30% higher copy number of satellites was detected in the pupal ovaries versus diploid cells of the brain of adults of D. oirilis. The number of satellites in the ovary of the adult fly is the same as in diploid tissue, or somewhat higher (Endow and Gall, 1975). Conversely, it was demonstrated that, in this species, satellite sequences constitute 44% of the genome in diploid cells, but only 16% of the DNA in the ovaries at the age of 2 days (Renkawitz-Pohl and Kunz, 1975). In D. hydei, satellites constitute 13% of DNA in diploid cells and only 2.2% of DNA in nurse cells (Renkawitz and Kunz, 1975).Underreplication of satellite DNA in nurse cells was detected in D. oirilis (Renkawitz-Pohland Kunz, 1975). In the highly polyploid ovarian nurse cells of Culliphoru erythrocephah,

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satellite sequences are completely represented, although they are underreplicated in the polytene chromosomes of trichogen cells (D. Ribbert and G. Dover, 1980, personal communication). A 200-bp sequence of the 3B55 centromeric satellite, which is highly underrepresented in larval salivary glands, is proportionally represented in nurse cell nuclei: there are 15,000, 17,800, and 48 copies/haploid genome in embryonic cells, nurse cells, and salivary gland cells, respectively (Nazimiec and Beckingham, 1986, 1989). Similar results were obtained for Prodiamesa oliuacea: satellites replicate completely in nurse, but not in salivary gland, cells (Zacharias, 1979). P-Heterochromatin in polytene chromosomes of pseudo-nurse cells of Drosophih melanoguster otu mutants is represented in much larger amounts than in larval salivary gland cells of the same mutant. Large new pieces of pericentric p-heterochromatin were found in the X, 3L, and 3R chromosomes of this cell type (Mal’ceva and Zhimulev, 1993; Koryakov and Zhimulev, 1995; Mal’ceva et al., 1995; Koryakov et al., 1996, 1997). The underrepresentation degree of DNA of the 18s and 28s rRNA genes is lower in ovarian nurse than polytene chromosome cells. Accepting the representation of the ribosomal genes in the diploid cells of D. mehnoguster as loo%, the corresponding values are 20% in salivary glands cells and 80-82% (Jacobs-Lorena, 1980; Mohan, 1977; Hammond and Laird, 1985a) in nurse cells. In D. hydei degree of representation of 28s 18s rDNA in the nurse cells may reach 85% (Renkawitz-Pohl, 1978). The DNA of these genes occurs in similar proportions in the ovaries: 135% in Calliphora erythrocephah, 51% in D. hydei, and 47% in Sarcophuga barbata (Renkawitz and Kunz, 1975). In Rhynchosciara angelae, the number of rRNA genes is about 100 in the salivary gland DNA, and it is 220 in the mature ovary tissue (Gambarini and Meneghini, 1972). Both homologous nucleolar organizers replicate in the ovarian nurse cells of Calliphora erythrocephah (Belikoff and Beckingham, 1985). Inconstancy of centromeric heterochromatin amount is not manifest only as incomplete polytenization. Its other manifestations include variation in DNA content along the chromatid and DNA overreplication. Instances of variation in the thickness of centromeric bands or heterochrornatic blocks (Figure 62) are widely known (Table 6). The DNA of heterochromatin occasionally overreplicates. Larvae developing much slower than the rest (3 months instead of the usual 3 weeks) were encountered in a collection of Chironomus melanotus. Material was “released”from the centromeric regions in some salivary gland cells during the fourth larval instar (Figure 63). The “released” heterochromatic bodies (H-bodies) consisted of a dense, rounded centromeric mass surrounded by a concentric circle of condensed chromatin. The H-bodies were not detected in all individuals with extended development, and their number varied in cells. Spectrophotometric measurements in normal larvae of Chironomus

+

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Figure 62. Variation in the size of centromeric heterochromatin in heterozygous individuals of Chirmomus plumosus. Reprinted by permission from Key1 (1965b).

melanotw demonstrated that the proportion of heterochromatin in the chromosomes at this developmental stage varies from 7 to 13%, and this may be the precondition requisite for overreplication (Key1 and Hagele, 1966). Extra droplets of centromeric heterochromatin were described in Chironomw sp. Yul from Yakutian population (Kiknadze et d.,1996). A similar event was detected in the giant foot pad cells of pulvilli of the Sarcophagidae family. Numerous granules are seen even in unsquashed nuclei in Sarcophaga bullata. They lie freely in nucleoplasm, unite into groups, and come into contact with the membrane or many regions of the chromosomes (Whitten, 1964, 1965; Roberts, 1968).The granules have a core composed of DNA, which is surrounded by ribonucleoproteins (staining with Azure B). The diameters of the core vary from 0.445 to 1.389 pm in Tricholioprocta impatiens (B. Roberts, 1975). It is believed that the granules result from (1) overreplication and “budding-off from many chromosomal regions (Whitten, 1965; Roberts et al., 1976); (2) the partly underreplicated sex chromosomes (Trepte, 1976); and (3) amplified ribosomal cistrons (Roberts et al., 1976). The first assumption appears quite unlikely because no homology is found between the DNA granules and the polytene chromosome bands with which they come into contact. When the chromosomes of S. bullata are stained with quinacrine, most granules fluoresce, while the bands contacting the granules

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Table 6 Variation in Amount of Centromeric Heterochromatin in Polytene Chromosomes in Representatives of the Diptera Order Species

Varying heterochromatin

References

Cecidomyiidae Dasyneura mame@ Chironomidae

Baeotendipes taurica

Ch. kiknadzeae

In 14-16% of larvae the size of centromeric heterochromatin in 2 chromosome AB is increased. Variation in thickness of centromere heterochromatic blocks between two karyotypes 1 and 2

Kiknadze et al. (1993)

Variation in centromeric heterochro. matin in the fourth chromosome

Michailova and Belyanina ( 1984) Key1 (1965b), Michailova and Ficher (1984, 1986), Kiknadze et al. (1987b), Michailova and Petrova (1991), Petrova (1991), Ilyinskaya (1992, 1993) Michailova (1987a) Kiknadze and Siirin (1991, 1993)

Variation in C-heterochromatin Sizes of centromeric blocks in each of the four pairs vary; in no examined nuclei were they increased at the same time; for each of the 9 sibling species blocks of the same size class as detected in Ch. plumosus are characteristic

Related species Chironomus of the pseudochummi complex (Ch. cloacalis, Ch. dupkx, Ch. tepperi) Ch. th. thummi x Ch. th. piger

Kiknadze et al. (1991c, 1996)

Heterozygosity in thickness of pericentric heterochromatin blocks in 25-33% of larvae

Ch. plumosus

Hybrids Ch. plumosus x Ch vancouveri

Kiknadze et al. (1991b)

In the former species, the centromeric block and C-heterochromatin are much thicker

Michailova (1987a)

Considerable interspecific differences in the thickness of the centromeric band

Lentzios and Stocker (1979)

In the chromosomes of the former species, Key1 (1965a,b), Hagele there is considerably more (by 16 (1977a), Gunderina and times) DNA including C-heteroAimanova (1996) chromatin In the former species, DNA replication Key1 and Pelling (1963), in centromeric blocks terminates later Hagele (1970, 1976)

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Table 6 Continued Species

Varying heterochromatin

References

Centrometic blocks of heterochromatin of the former species contain considerably more AT repeats

Schmidt (1980, 1981, 1984), Schmidt et al. (1980,

Ch. sp. Yal Ch. sp. Ya2

-

Glyprotendipes barbipesl G . salinus

Twofold excess of DNA in heterochromatin of the former species in comparison with the latter

Kiknadze et al. (1996) Kiknadze et al. (1996) Michailova (1989b), Michailova and Nikolov

G . gripekoveni

-

-

1982)

(1992) Michailova and Petrova

(1984) Culicidae Anopheles messeae

Differences in quality of pericentric C+/- heterochrornatic blocks in different populations of Tomsk

Sharakhova (1997)

-

Bedo (1975b)

Simuliidae Simulium pictipes

do not. Label was observed also over the majority of granules in in situ hybridization of fast renaturating DNA. Labeling sites in the chromosomes were found only in the centromeric heterochromatin of the regions of the C and E chromosomes (Swift et al., 1978; Samols and Swift, 1979b). There are no granules in the giant foot pad cells on the third day of development; Q’ granules occur only in association with Q’ regions of C and E chromosomes on the fourth day, and Q’ granules can contact with Q- bands of polytene chromosomes on the fifth day (Samols and Swift, 197913). The granules are not digested with endonucleases Hue 111, Alu I, or Hind111 in squash preparations. These enzymes do not digest the 1.663, 1.670, or 1.692 satellites (Bultmann and Mezzanotte, 1987). Presumably, overreplication of repetitive DNA is a specific feature of the nuclei of giant pulvillar cells only, inasmuch as there were no reports concerning granules of this kind in cells of other types, especially bristle-forming cells (Ribbert, 1967; Whitten, 1969). There is information that similar granules occur in Drosophila. Outpocketings of the nuclear membranes, which protrude into the cytoplasm (20-200 calculated outpocketings per nucleus, according to calculations), were detected in salivary gland cells. DNA material of the chromosomes “from reverted cytological repeats” (possibly from intercalary heterochromatin) or terminal chromosome regions is frequently found in the formed blebs (Gay, 1956; Kaufmann and Gay, 1958).

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F. Zhimulev

Figure 63. Formation of “drops”ofheterochromatic material released from blocks of centrorneric hetrochromatin (a-c), and freely floating (d) in the nuclei of salivary gland cells of Chiraomus melanotus. Reprinted by permission from Key1 and Hagele (1966).

There is no reasonable argument for suggesting that extrachromosomal bodies may be present in the nuclei of the salivary gland cells of the adults of Lathriopygu rnonticola (the lnsecta class, the Collembola) (Cassagnau, 1968). In fact, the X chromosome in this species can be pompon-like with a large a-heterochromatin block in the center. “Extrachromosomal”DNA and the X chromosome are very similar in morphological appearance, as may be judged by the illustrations (Cassagnau, 1974a).

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In one case variability of the quantity of the pericentric heterochromatin was correlated with resistance to bacterial infection in Anopheks messeae. Midges with smaller blocks of the heterochromatin in ovarian nurse cell polytene chromosomes appeared to be less sensitive to infection by Bacillus thun‘ngiensis (Timofeyeva et al., 1997).

D. Cytogenetics of (Y- and p-heterochromatin In the mid-I 930s, Muller-then working in Russia-suggested that chromosome regions covering the nucleolar organizer may be genetically organized. Although not confirmed later, his idea was a strong impetus to research along this line (for discussion, see Schalet and Lefevre, 1976). Briefly, his idea was that the inert regions are composed of nongenic material. Few in number in a region, the genes are represented by relatively large amounts of chromosomal material. Inertness of a region is manifested as a considerably higher proportion of substances extrinsic to the genes at metaphase in the inert compared to the “active” regions. The chromosome is not inert and the genes in it are normally located, but they are covered with a “crust” that makes their coiling into a helix at mitosis impossible (Muller and Gershenzon, 1935; Muller and Prokofyeva, 1935; Muller et at., 1937). In contrast, Fujii (1938) did not believe that at least p-heterochromatin is an intrinsically inert region (i.e., heterochromatic). Figures 64 and 65 show models of the correspondence between the heterochromatin of mitotic and polytene chromosomes. In the first model (Figure 64), a-and P-heterochromatin are present in both mitotic and polytene chromosomes. In the second (Figure 65), p-heterochromatin completely replicates and is saturated with genes; 6-heterochromatin and euchromatin are functionally indistinguishable. a-Heterochromatin, with the exception of the nucleolar organizer, does not contain any other genes and it does not polytenize (Miklos and Cotsell, 1990). According to the model of Spradling et al. (1992), a-heterochromatin is formed from a large central block including all satellite DNAs. The more

Euchrornatin

Helerochrornatin

CenlrOmere

Figure 64. Distribution of euchromatin and a-and p-hetrochromatin in the mitotic (top) and polytene (bottom) chromosomes of Drosophila. After Gall (1973).

126

1. F. Zhimulev ouchromrlin

mitotic heterochromatin

mitotic

X -chromosome

x .cI%%me

.-

Figure 65. A scheme of the cytogenetic organization of the transitional region from euchromatin to a-heterochromatin in the X chromosome of D. mekmogaster. After Yamamoto et al. (1990).

distant parts of repeated DNA form p-heterochromatin. P-Heterochromatin is densely populated with complementation groups. In large-scale genetic experiments designed to study the most proximal region of the X chromosome, about 40 deletions and duplications were generated; the majority had both breakpoints proximal to the 19D2-El region at the position of the mehizedlike-mell. T h e numerous mutations mapped using these rearrangements were assigned to 15 complementation groups. Five genes identified in section 20 (Schalet and Lefevre, 1973, 1976) showed the features of p-heterochromatin: unusual banding pattern and richness in mobile elements. Based on data obtained subsequently (Lifschytz, 1978; Miklos et al., 1987; Perrimon et al., 1989; Yamamoto et al., 1990), a cytogenetic map was built for the most proximal region (Figure 66). Thirteen loci were detected in section 20. A new gene, flamenco, that seems to control mobility properties of gypsy was found in the 20A1-3 region (Prud’homme et al., 1995). T h e suppressor offorked, su(f), gene is the most proximal gene on the X chromosome. By using a DNA fragment that encompasses the entire su(f), its cytological location was determined in situ to be in the chromocenter of the polytene chromosome, presumably very close to the a-and P-heterochromatin junction region of the X chromosome (Yamamoto et al., 1990). According to in situ hybridization of DNA of this gene in polytene chromosomes of pseudo-nurse cells, su(f) is located in the banded part

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Figure 66. A cytogenetic map of the proximal region of the X chromosome of D. melanogaster. (a) Regions of the X chromosome. (b) Gene symbols. (c) Symbols and extension of deletions (white bands) and duplications (black bands). After Perrimon et nl. (1989).

of the 20th region, in the 20C (D. E. Koryakov, A. A. Alekseyenko, E. S. Belyaeva and I. F. Zhimulev, 1996, unpublished). It was suggested that there is a distinct boundary between the eu- and heterochromatin of the mitotic chromosome passing in the 20DE region in the

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polytene X chromosome (Schalet and Lefevre, 1976). However, Lifschytz (1978) believes that there is no such readily perceived boundary and that the base of the polytene chromosome is composed of heterochromatic segments of different lengths that alternate with euchromatic segments. Thus far, the region of the polytene chromosome to which the distal end of the mitotic heterochromatin corresponds has not been defined. Changes in the banded part of polytene chromosomes were usually not seen when chromosomal rearrangements with one of the breakpoints in centromeric heterochromatin were used (Hinton, 194210; Holm et al., 1969; Hilliker et al., 1980). Section 20 is the most proximal to a-heterochromatin in the X chromosome of Drosophikz. This banded region differs in morphology from the others: it contains bands that are numerous and small, twisted, and often involved in ectopic pairing. The region frequently breaks off from both the chromocenter and euchromatin (Prokofyeva-Belgovskaya, 1 9 3 7 ~ )However, . the wm4 inversion, the proximal breakpoint of which is in heterochromatin (see Figure 32), does not affect any band in section 20 in the polytene chromosome, and hence it has a break residing in the more proximal heterochromatin (Figure 67). In the sc4sc8 chromosomal rearrangement with almost completely removed heterochromatin, section 20 is also completely retained. However, there is some uncertainty as to whether segment 20 is a polytenized fragment of heterochromatin located between the sc4 breakpoint and its distal part (see Figure 32). In reviewing these data, the conclusion is reached that the bulk ofa-heterochromatin of the X chromosome is not represented by bands of the polytene chromosome (Schalet and Lefevre, 1973, 1976). The proximal breakpoint of the Df(ZR)Z4,mapped in the middle of the het46 block (see Figure 33), is located in the 41C region of the polytene chromosome. This means that the most distal heterochromatic region (the middle of the het46 block) corresponds to the banded part of chromosome 2R. The distal part of the het46 block can give only the banded part of the polytene chromosome distal to the 41C region. In such cases there is no material on the distal end of the mitotic 2R heterochromatin to form P-heterochromatin (Koryakov et al., 1996). Data have clarified the organization of a- and P-heterochromatin. Analysis of the distribution of HeT DNA sequences in pericentric heterochromatin of Drosophila chromosomes demonstrated their interspersion with satellite DNAs. During the formation of the chromacenter, the satellite DNAs aggregate into the a-heterochromatin, leaving the interspersed regions of the other sequences, including HeT DNA, looped out of the blocks of a-heterochromatin to form P-heterochromatin (Traverse and Pardue, 1989). This interpretation is supported by the behavior of the rRNA genes also located within heterochromatin. The rDNA is pulled out from the chromocenter into the nucleolus (Gatti and Pimpinelli, 1992).

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Figure 67. Morphology of the closest-to-heterochromatin2OAD region of the X chromosome in wm4/+ (a) and sc4sca/y Drosophila (b). Reprinted by permission from Schalet and Lefevre, 1973).

Numerous nonsatellite sequences including genes were found in heterochromatic regions of mitotic chromosomes (Devlin et al., 1990a,b; Dimitri, 1991; Eberl e t al., 1993; Brunner et al., 1994; Biggs e t al., 1994; Zhang and Spradling, 1995). They form “islands”containing middle-repetitive and/or singlecopy sequences in a “sea” of satellite DNAs (Karpen and Spradling, 1990; Karpen, 1994; Le et al., 1995; Zhang and Spradling, 1995). Unique sequences found in autosomal heterochromatin in sites of P-element insertions are represented fully in the salivary gland polytene chromosomes. These sequences dispersed at multiple sires of mitotic heterochromatin may contribute to p-heterochromatin (Zhang and Spradling, 1995). Underrepresentation of heterochromatic DNA is reduced in the pseudo-

130

1. F. Zhimulev

nurse chromosomes (PNC) of the otu mutant of D. melanogaster. As a result, pheterochromatin is represented much more abundantly in the PNC than in salivary glands cells. The additional blocks of heterochromatin (1 and 2) located proximal to the 80AC region in the 3L chromosome and the one block (3) proximal to 8 1F in 3R are clearly discerned in PNC polytene chromosomes (Mal’ceva and Zhimulev, 1993). There are differences in molecular organization between these three additional blocks of p-heterochromatin. The DNA of the 20p1 .4 clone, which makes contact with nuclear lamina, hybridized in situ only with block 3; the Dm 12 clone DNA (mobile element 1360) hybridized in 80AC and blocks 1 and 2 (Sharakhov, 1995). Morphological analysis of chromocenters and cytogenetic mapping of several inversion breakpoints showed that the second chromosome heterochromatin in the PNC is represented by interspersed meshlike (p-heterochromatin) and compacted (a-heterochromatin) material (Figure 68). Mapping the DNA clones and chromosomal rearrangement breakpoints in the mitotic heterochromatin of the second chromosome demonstrated that the polytene chromosome region 4OD-F corresponds to mitotic heterochromatin h35. The phB block is almost completely formed by the h39-h42B regions (RspVDe2). The proximal half of phC is formed by h43-h44 (VDe2-VDe1). This means that in the PNC proximal mitotic heterochromatin of chromosome 2 appears as nonpolytenized a-heterochromatin and polytenized p-heterochromatin. In the SG chromosomes, however, material of the mitotic regions h43-h44 appears not to be polytenized at all. This was confirmed by dot hybridization of the AAGAC satellite DNA clone 35 36

37

38 C

3940 41

42

A

B

43 44

45

46

B

C

Figure 68. The scheme of polytenization of different parts of pericentric heterochromatin in pseudonurse cells of the otu” mutant in Drosophila melanogarter. ( A ) Heterochromatin map. (B and C ) Location of inversion and clones on mitotic (B) and polytene ( C )chromosome maps. phA, phB, and phC, heterochromatic blocks visible in pseudo-nurse cell chromosomes, forming from deep heterochromatin. After Koryakov and Zhimulev (1995).

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131

1.686 (Koryakov and Zhimulev, 1995; Koryakov et al., 1996), which maps to region h44 of 2Rh and to the Y chromosome (Lohe et al., 1993). These experiments performed using XX females showed similar hybridization rates with DNA from the head, ovaries, and PNCs, and almost no hybridization with SG DNA (Koryakov et al., 1996). Whereas in the SG cells one can see a body (bodies) of a-heterochromatin mainly in the middle of the common chromocenter, the chromosome 2 pericentric region in PNC blocks has the appearance of a linear sequence of structures of a-heterochromatin (phA and constrictions) and p-heterochromatin (blocks phB and phC in Figure 68). When taken together, these data suggest a scheme for the organization of a-and P-heterochromatin and the junction between eu- and heterochromatin (Figure 69). Mitotic heterochromatin is represented as an array of segments, tentatively numbered 1 through 8 in Figure 69a. This heterochromatin obviously performs different functions and, accordingly, shows different replicative and other properties in various cell lines. In the SG chromosomes (Figure 69b), regions 1,

Polytene chromosome in SG 1 2

-

3 4 5 6

7

8

Heterochromatin Euchromatin

Mit4lt.k

a chromosome

1 2 3 4 5 6 7 8

= *

1 2 3 4 5 6

chromosome Polytene in PNC

d

L*

7 8

Figure 69. General scheme of chromocenter organization in Drosophila melanoguster (a) Blocks of pericentric heterochromatin (1-8) in mitotic chromosome. (b) Chromocenter. Fragments of tightly packed heterochromatin (1, 3 , 5, and 7) are underreplicated and collected together, forming a-heterochromatin. Loops of heterochromatin blocks that are less compact are more represented in polytene chromosomes (2,4, and 6 ) . 8, boundary between euchromatin and deep heterochromatin, partly underreplicated. Reprinted by permission from Koryakov et al. (1996h).

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1. F. Zhimulev

3, 5 and 7 remain underpolytenized and compacted. Segment 8 is a euchromatin-heterochromatin transition zone developing the base of the polytene arm similar to the map regions 40 or 41. The compact and underpolytenized segments associate tightly, forming a-heterochromatin (Figure 69c), while the partly polytenized regions loop out to form diffuse p-heterochromatin, as Traverse and Pardue suggested (1989). In PNC (Figure 69d), certain segments (e.g., 5), that are compacted in SGs replicate to some extent and become diffuse; the same region can be present as a type in one tissue (5 in Figure 69b and 69c) and as p type in another (5 in Figure 69d) (Koryakov et al., 1996, 1997). So, the nature of alphaand beta-heterochromatin is dynamic. In general, heterochromatin passes through more endoreplication rounds in the PNCs than in the SG cells (Mal’ceva and Zhimulev, 1993; Mal’ceva et al., 1995). There is a similar view about heterochromatin in the SG chromosomes (Traverse and Pardye, 1989; Berghella and Dimitri, 1996). However, Traverse and Pardue (1989) as well as Devlin et al. (1990b) thought that a-heterochromatin develops from nonreplicated satellites, while p-heterochromatin is formed from other interspersed DNA sequences. It would be more correct to be concerned with sequences having different replication properties rather than with the satellite versus nonsatellite sequences or, for instance, with differently staining regions. Indeed, in the SG chromosomes of D. melanogaster, certain nonsatellite sequences are not polytenized-for example, the mobile elements mdgl (Shevelyov et al., 1989) and aurora (Shevelyov, 1993) and the heterochromatic copies of the Ste gene (Shevelyov, 1992). In contrast, satellite sequences replicate in polyploid nurse cell nuclei of D. virilis (Gall et al., 1971). Certain P-element insertions into pericentric heterochromatin polytenize at the level of euchromatin (Zhang and Spradling, 1995). It was proposed that heterochromatic state (compaction, underpolytenization of a type, or fragility and partial polytenization of p type) depends not only on nucleotide sequence, but also on functional specificities of different heterochromatin regions (Koryakov et al., 1996, 1997). The model accommodates very well the numerous data on transcriptional activity found in chromocentral DNA. As follows from EM autoradiographic data, a-heterochromatin is transcriptionally inactive, and no RNP particles are identified in it (Lakhotia and Jacob, 1974). Furthermore, ultrastructural study of unsquashed nuclei of the salivary glands of Dosophila revealed RNP granules of two types in P-heterochromatin: (1) particles 25 nm in size, making contacts with chromocentral heterochromatin; and (2) particles 45-50 nm in size, organized in clusters and located near or within the transition zones of P-heterochromatin and the chromosome arms (Lakhotia, 1974b; Lakhotia and Jacob, 1974). Granules of two kinds (approximately 35 nm and from 50 to 60 nm) were identified in the EM studies of the chromocenters of Drosophila when the chromosomes were isolated from the nuclei by microdissection (Mott et d., 1980). ---

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133

Incorporation of [3H]uridine was observed in the 80AC region of chromosome 3L of larvae and prepupae and also in the 41AF region of chromosome 2R of O-hr prepupae of D. rnelanogaster (Zhimulev and Lychev, 1972; Zhimulev and Belyaeva, 1974, 1975). Both regions are considered as P-heterochromatin. Incorporation of [3H]uridine into 6-heterochromatin was observed in the EM sections of the chromosomes (Lakhotia and Jacob, 1974; Kerkis et al., 1977). The total area of a-heterochromatin on sections of unsquashed nuclei was 26.48 Fm2, and that of P-heterochromatin was 1639.84 FmZ. No silver grains were detected over a-heterochromatin, and 774 silver grains were observed over p-heterochromatin (Lakhotia and Jacob, 1974). Localization of antibodies on DNA-RNA hybrids supports the notion that P-heterochromatin is transcriptionally active. They tightly bind in segment 40 of chromosome 2L and in 41AB of chromosome 2R (Vlassovaet at., 1984,1985). In in situ hybridization of labeled poly(A)+ RNA isolated from a cell culture of Drosophila, a heavily labeling site, besides the numerous regions in the euchromatic part of the chromosomes as well as regions 41AF, 40AF, 81F, and 19E-20F, were found to be P-heterochromatin (Spradling et al., 1975; Gvozdev et al., 1980a,b). A genomic DNA clone of Drosoghila, 4.4 kb in size and mapped to the 80C region, encodes poly(A)+ RNA of 1.4 kb; it is abundantly represented in cytoplasm (0.6% in clusters of cells and embryonic cells). The RNA encodes a protein with a molecular mass of 16 kDa (Biessmann et al., 1981). The GATA repeat lying in the 19F-20AB region is also transcribed (Singh et al., 1984). The Mts40 gene expressed in primary spermatocytes is located in region 40C a t the base of chromosome 2L, very close to or within P-heterochromatin, or at the junction between eu- and p-heterochromatin (Russell and Kaiser, 1994). Expression was also found for the l(l)B214 locus located in the immediate vicinity of p-heterochromatin of the X chromosome (19Fl-2 region) (Russell et al., 1992). In insects, in which chromocenters are not formed and heterochromatin polytenizes completely (e.g., in sciarids), the nucleolar organizer also lie in the centromeric blocks of heterochromatin (see Section IV,A). In Rhynchosciara a n g e h , the largest blocks of heterochromatin lie in the centromeric regions of chromosomes B, C, and X. After in witro incubation of the salivary glands for several hours to 12 days, puffs arise in 3-15% of cells (depending on larval age) in the heterochromatin of chromosomes A, B, and X (Simoes et al., 1974; Simoes and Cestari, 1982). Puffs similar in morphology and location are induced in cells infected with viruses and microsporidia (Diaz et al., 1969). These so-called heterochromatic puffs are also induced by heat shock when culturing conditions are poor (temperature or medium pH change) (Simoes et al., 1969). The induced puffs intensely incorporate [)H]uridine (Simoes and Cestari, 1982). [3H]uridine is also intensively incorporated into the kinetochore regions

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of the nurse cell chromosomes of Culliphoru erythrocephala (Ribbert, 1979). Nazimiec and Beckingham (1986, 1989) demonstrated that the 3B55 satellite sequence is transcribed in nurse cells but not in 9- to 10-hr embryos in this species. In Cerutitis cupitata, heterochromatic meshwork representing the X chromosome in prepupal orbital trichogen and larval salivary gland cells is surrounded by 25- to 30-nm RNP granules. Similar granules were found around the spherical, compact Y chromosome of this species (Semeshin et al., 1995). As for replication organization in centromeric heterochromatin, it may be noted that, without exception, both centromeric heterochromatin in species lacking the chromocenter and chromocentral p-heterochromatin are late replicating (Pavan, 1959; Key1 and Pelling, 1963; Gabrusewycz-Garcia, 1964; Tulchin et ul., 1967; Hagele, 1970; Gubenko, 1974a,b, 1976a; Mukherjee et al., 1977; Sokoloff, 1977; Stocker et al., 1978; Zacharias, 1979; Steinemann, 1981; Zhimulev et al., 1989a,b; Bolshakov and Zhimulev, 1990). In the polytene chromosomes of bean haustorium (Pheolus coccineus), the chromocenter is not formed, but the chromosomes differ in denser a- and more loosely compacted P-heterochromatin. At the beginning of the S phase, euchromatin replicates first, then P-heterochromatin, and a-heterochromatin replicates last (Brady and Clutter, 1974).

VII. INTERCALARY HETEROCHROMATIN Bridges encountered difficulties in assigning the proximal part of chromosome 2L on the cytological map of polytene chromosomeof Drosophla. The bands of the 32F-33C region were connected with the those of the 34F-35C region by fibers, obscuring the morphology of the chromosomes. There were also a number of ambiguities in mapping the 37F-38A and 39CE regions (Bridges, 1935c, 1936; Morgan et d., 1937). At the distal (telomeric) end of the fourth microchromosome, a region was identified attracted to the proximal (centromeric) region of the same chromosome with the result that the chromosome bent like a horseshoe (Bridges, 193513). In the same year, pairing contacts were detected in D. psewloobscura and Ch. thummi (Bauer, 1935; Tan, 1935).Soon thereafter, it became clear that such regions are present at the ends of the chromosomes in many species and that they can be involved in pairing with the terminal regions of other chromosomes; with the internal sites, lying in different regions of polytene chromosomes; and with pericentromeric heterochromatin (Bauer, 1936a; Tinyakov, 1936;Frolova, 1936b; Prokofyeva-Belgovskaya,193713,1938,1939d).Pairing between the nonhomologous regions of the chromosomes was called ectopic (Slizynski, 1945). In their studies of the distribution of the ends of radiation-induced chromosomal rearrangements, Prokofyeva-Belgovskaya and Khvostova ( 1939) and, independently, Kaufmann ( 1939) demonstrated that the rearrangements occur much more frequently in some regions than others. The localizations of the “hot

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

135

spots” of chromosomal rearrangements in ectopic pairing regions were coincident to a large extent (Slizynski, 1945). By analogy with pericentromericheterochromatin, where chromosomal rearrangements are frequent, Kauhann (1939) suggested that higher fragility of the regions of ectopic contacts is due to the presence of heterochromatin intercalated in euchromatin (i.e., intercalary heterochromatin). Prokofyeva-Belgovskaya(193713) suggested that these regions be called “inert”; however, this term did not take root. In support of the existence of intercalary heterochromatin was the thenwidespread notion that ectopic pairing conforms to the rule “similar with similar” (Prokofyeva-Belgovskaya, 1941), and that the regions making contacts with pericentromeric heterochromatin may be heterochromatic, although intercalary. It became further evident that the notion of intercalary heterochromatin (IH) covers a group of chromosome regions with a large set of properties distinguishing them from the neighboring so-called euchromatic regions and often from one another. In addition to the mentioned ectopic pairing and fragility arising during the formation of rearrangements in these regions, other findings included, as a rule, high compaction level of chromatin, late termination of DNA replication during the S phase, and high frequencies of mechanical breaks. In some cases, these regions stained differently with specific dyes. Discussion of the real existence of IH continued unabated. While in extensive reviews facts have been gleaned in favor of the presence of the IH in chromosomes (Hannah, 1951; Beermann, 1962; Kaufmann and Iddles, 1963; Ananiev et al., 1978; Gvozdev, 1981a,b; Zhimulev et al., 1981, 1982, 1989a,b), in other papers it was in strong doubt. It is Back’s (1976, p. 32) opinion that “the hypothetic intercalary heterochromatin (Hannah, 1951) cannot be demonstrated cytologically. Its existence is deduced from the increased tendency toward breaking of certain chromosome regions of Drosophila.” Spofford (1976) called attenion to the fact that the formation frequency of rearrangements correlates with DNA amount in a region; rearrangements are formed in the large bands. Other authors believe that the ectopic pairing region cannot be called heterochromatic; these regions do not appear as such in the mitotic chromosomes, they are incapable of inducing genetic inactivation resulting from position effect variegation, and satellite DNAs are not detected in them (Hilliker et al., 1980; Hilliker and Sharp, 1988; Miklos and Cotsell, 1990; Gatti and Pimpinelli, 1992). It is Laird’s view (Laird et al., 1987a,b; Laird, 1989) that DNA sequences possessing properties of the IH are underrepresented in polytene chromosomes, they presumably have no other features of centromeric heterochromatin. At least a part of the IH regions represent normal euchromatic DNA the replication cycle of which is delayed in some cell types. For this reason, the new term “fragile sites” can indeed be useful (Laird, 1989). A number of erroneous hypotheses have been proposed to explain the relation of the IH to some of the structural-functional features of polytene chromosomes. Criticism of these hypotheses has also made more vulnerable the whole

136

1. F. Zhimulev

concept of the IH. Thus Bridges (1935c, 1936) suggested that ectopic pairing occurs between identical band sets or cytological repeats, as they are frequently called. Subsequently, various authors have discussed these “repeats” as a diagnostic feature of the IH (Hannah, 1951; Kaufmann and Iddles, 1963; Ananiev et al., 1978, 1979a; Gvozdev, 1981a,b). However, it became still further evident that cytological repeats in the regions Bridges has indicated are nonexistent, being mostly artifacts (for discussion, see Zhimulev, 199213). Likewise, no support was provided for Hannah’s (1951) suggestion that the Minute genes map to heterochromatic regions (see Section VII,C,8). Section VII presents a description of the organizational features of intercalary heterochromatin of various types, and it is also concerned with controversial interpretations.

A. Intercalary a-heterochromatin 1. Dipteran insects a. Phryne cincta In an alpine population of Phryne cincta, individuals were encountered whose male and female X chromosomes had several (one to three) additional blocks of heterochromatin (see a-1, 01-2, and a.3 in Figure 70), appearing as heavily staining heterochromatic vacuolar masses intercalated in the euchromatic parts of the chromosome. Only two a-blocks were sufficient for producing a twofold elongation of the X chromosomes (X1 in Figure 70) as compared to the normal (Xs). Individuals of the most various genotypes (Xl/Xl, Xl/Xs, and Xs/Xs) occur in the alpine population, while there were only Xs/Xs individuals in the Berlin population. Because of incomplete polytenization of intercalary a-heterochromatin in polytene chromosomes, there was almost no difference in X chromosome length. Several small a-blocks were localized in the autosomes (Wolf and Struck, 1960; Wolf, 1963, 1970; Wolf and Wolf, 1969; Wolf and Sokoloff, 1973, 1976). Breaks and morphology of blocks of heterochromatin can vary from exceptionally large to very small. The material can stretch out to form threads or bulky protuberances, which occasionally break loose from the block, freely wandering out of the nucleus (“a-migration”; Wolf, 1970; Wolf and Sokoloff, 1973). Loss of integrity of a-blocks, presumably due to incomplete polytenization, leads to the fragmentation of polytene chromosomes whose frequency correlates with change in temperature: 84% of the chromosomes remained intact at 18”C,and 86% fragmented at 2°C (Wolf, 1970; Wolf and Sokoloff, 1973). Fragmentation is tissue specific: the number of unfragmented X chromosomes was twice as great in the midgut as in salivary glands in the same individuals (Wolf and Sokoloff, 1973).No differences in the frequenciesof fragmented chromosomes were found between males hemizygous for the X chromosome and homozygous females, although a-blocks appear bulkier in males (Wolf and Sokoloff, 1973).

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137

Figure 70. Intercalary a-heterochromatin in the polytene chromosomes of the salivary glands ( a x , f-h) and mitotic (d and e ) chromosomes in Phryne cincta. a-1, a-2, a-3, Blocks of heterochromatin; arrows indicate material extruded from a-blocks (f and g) or a break along the a-block region (0. (d and e ) Numbers designate chromosomes; the X chromosomes are depicted black. After Wolf and Sokoloff (1973).

In addition to compact state, blocks of intercalary a-heterochromatin show features of heterochromatin, such as ectopic pairing (Wolf and Sokoloff, 1973, 1976; Sokoloff, 1977). The considerable decrease in the amount of a-heterochromatin in polytene compared to mitotic chromosomes, and their incorporation of [3H]thymidine, indicate that they are polytenized, but not completely (Sokoloff, 1977). Blocks of intercalary heterochromatin have a strong effect on crossing over. XI/Xs females with autosomes of the wild type were crossed to Xs males homozygous for four inversions in the second and third chromosomes (2a2b/2a2band 3a3b/3a3b). The progeny were heterozygous for these inversions (SS/2a2b, SS/3a3b). Some in-

138

I. F. Zhimulev

Figure 70. Continued

dividuals were homozygous and some heterozygous for Xs/Xs. Females of these types were mated to males without inversions. Evidence for crossing over was the appearance of individuals having only a single inversion on the chromosome. Crossovers for the second chromosome were recovered at a frequency of 2% and those for the third chromosome at 0% among progeny of Xs/Xs females. The respective values were 42% and 13%for progeny of Xl/Xs females (Wolf, 1963, 1970). No incorporationof PHIuridine into a-blockswas detected (Sokoloff,1977).

b. Drosophila heretensis (littoralis) Polymorphism for the heterochromatin region H-2 in the short arm of the second chromosome was detected in some populations of this species (Mitrofanov and Poleuktova, 1982).The majority of individuals (68.9%) of the population of the Mirniy settlement lacks the block; the remaining individuals are homozygous (14.3%) or heterozygous (16.8%). In the authors’ view this region is similar in

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

139

staining and structure to the region of p-heterochromatin (Mitrofanov and Poluektova, 1982), although its general compacted state and deep staining allow it to be referred to as a-heterochromatin. Individuals without a block predominate in natural conditions, however, the frequency of the block is severalfold higher in laboratory conditions. Presumably, this is due to the elimination of unfavorable effects of the laboratory conditions (Poluektovaetd.,1984).It is believed that the presence of blocks of heterochromatin can modify the expression of the activities of certain genes (Korochkin et d., 1983). a-NM4 and a-Ltl 1 DNA clones most strongly hybridize in situ with the DNA of chromocenters, dispersely along chromosome length, and they intensely label the previously mentioned block of heterochromatin (Sayanova et al., 1987).

c. Some species of Chironomus “Dark knobs” are seen in each chromosome of Orthocladizls bipunctellus (2n = 6). These knobs are blocks of strongly staining compacted vacuolar heterochromatin that lie in the middle or at the ends of the chromosomes (Figure 71; see also Figure 120f in Section IX). The blocks show other features of heterochromatin besides morphological ones. For example, ectopic contacts are frequently observed between the ends of the chromosomes where they are located. When stained with quinacrine, these blocks fluoresce brightly, and they stain positively for C+-heterochromatin (Michailova and Belcheva, 1982; Michailova, 1989a). A block of heavily staining material is connected with the nucleolus at one of the ends of the fourth chromosome of Cryptochironomusfidmanae. It contains Q+and C’ material (Michailova and Belcheva, 1982; Michailova, 1989a). Accumulation of heavily staining material, presumably a-heterochromatin, was identified in Ch. nditarsis (Fischer and Ttchy, 1980; Pulver and Fis. cher, 1980).

2. Apterygotan insects In the polytene chromosomes of many species of the Collembola order (see Zhimulev, 1992b, 1996),large blocks of darkly staining, frequently vacuolar chromatin, called a-heterochromatin, were identified. The blocks are located in the centromeric and telomeric regions as well as in different regions of chromosome arms. They make ectopic contacts; for this reason, they can be referred to as sites of intercalary a-heterochromatin. Their number varies widely both in various populations within a species and in species within a genus (Cassagnau, 1968, 1970, 1975, 1976). In Bilobella aurantiaca, site number can vary from 9 ro 32 (Figure 72), and a south-to-north gradient for the amount of a-heterochromatin was detected in a Catalonian population (Spain) (Dalens, 1979). Individuals of the population of southern Italy have smaller amounts of heterochromatin than those of the central part of the country (Dallai, 1979). A similarly wide interpopulational variation in site number of a-heterochromatin was found among the Eu-

140

I. F. Zhlrnulev

Figure 71. Giant blocks of heterochromatin (indicated by arrows) in the polytene chromosomes of Orthocladius bipunctellus. C, the centromere. After Michailova and Belcheva (1982).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

141

IV

I

Ill 01

a

b

7 Y '(1 a

b

a

b

21b

b

25

a

P20 I

22

b

a

b

Figure 72. Differences in the location of regions of intercalary a-heterochromatin in Bilobella aurantiacain the population of Sainte Baume, southern France (a) and Gredos, central Spain (b).I-VII, chromosome numbers; 1-32, sites of intercalary heterochromatin. After Deharveng and Lee ( 1984).

ropean populations of Lathriopygu longisem and BilobeUa grussei (Cassagnau, 1971; Cassagnau et a!., I979), as well as among closely related species of the Virronuru genus from Nepal (Cassagnau and Deharveng, 1981). In Neanuru monticola, an association was disclosed between changes in ecological conditions and the amount of a-heterochromatin in the X chromosome. The amount of heterochromatin is small in individuals living at altitudes below 1000 m and greater in those living at altitudes above 2000 m (Cassagnau, 1974a,b). This species, as a rule, is distributed along the slopes of mountains and, in such a case, it contains little heterochromatin. In individuals occupying the southern slopes, the amount of heterochromatin is maximal (Deharveng, 1976). Blocks of a-heterochromatin can make contacts, this is associated with the formation of true ectopic fibers and somatic pairing of homologs usually not observed in Collemboles (Cassagnau, 1966, 1971; Deharveng and Lee, 1984). In Bilobella aurantimu, the terminal blocks of a-heterochromatin pair with especially high frequencies and the subterminal blocks with lower frequencies. No pairing of centromeric blocks was found in some samples, although they are the largest in the

142

1. F. Zhimulev

karyotype. In the other insect samples of this population, intercalary (interstitial, according to the author’s terminology) regions were frequently involved in pairing with the “homologous”or the “nonhomologous”chromosomes, and contacts were made between the terminal and interstitial blocks (Deharveng and Lee, 1984).

3. Infusoria Large blocks of darkly staining material are identified in the polytene chromosomes of the macronuclear anlage of Stylonychia mytitus (for review, see Zhimulev, 1992b, 1996). The blocks are so large that they protrude beyond the diameter of the other part of the chromosome (Figure 73). The morphology and size of each is very characteristic, so that the block renders chromosomes pattern specific and their individual regions readily recognizable (Jareno et al., 1972; Ammermann et al., 1974; Raikov and Ammermann, 1976; Alonso, 1978; Jareno, 1990). Besides compaction, the regions show at least two other features typical of a-heterochromatin: late termination of replication (see Figure 73) and involvement in ectopic pairing (Jareno, 1990).

Figure 73. Blocks of a-heterochromatin in the polytene chromosomes of the entire set (a) and in an individual chromosome (M) in Stylonychiarnytilus.( candd) Labeling pattern ofthe chromosomes during termination of DNA replication after I3H1thymidineincorporation at the ages of 30-32 (c) and 3 8 4 0 hr (d). Arrows indicate blocks of a-heterochromatin. (a) after Ammermann (1979); (M) reprinted by permission from Ammermann er nl. (1974).

Polytene Chromosomes, Heterochrornatin, and Position Effect Variegation

143

B. Identification of intercalary heterochromatin by differential staining 1. C-heterochromatin The data presented in Table 7 provide evidence for the occurrence of C-heterochromatin-that is, intercalary heterochromatin-in the euchromatic regions of polytene chromosomes. C-staining of the chromosomes of Rhynchosciara hollaenderi during development is revealed in the major blocks of C'material at all developmental stages, although some C+-bands are not consistently seen in the chromosomes with low polyteny degree (Figure 76).

2. Q- and H-heterochromatin Using fluorescent dyes, rather lengthy tracts of repeats rich in AT sequences are identified (see Section 11). The number of fluorescent bands in species can vary widely, from complete absence in D. hydei, D. melanopalpa, D. repkta, D. peninsuhris, D. tumiditarsis, D. hernipem, D. grimshawi (Barr and Ellison, 1971b;

Figure 73. Continued

144

1. F. Zhirnulev

Table 7 Occurrence of Intercalary C-Heterochromatin in Polytene Chromosomes ~

Family, species

Location and characteristics

References

INSECTS Chironomidae Ch. balatonicw

Ch. plumosus

To 30 C-bands in arms A, B, and F Thin bands in the centromeric region, single regions of the fourth chromosome Single regions of the fourth chromosome

Ch. thummi thumrni

Regions of the third chromosome in N-1 (Switzerland) and no staining in individuals of the Durankulak (Bulgaria) population To 15 C+-bands in each chromosome

Ch. luridus

Ch. vancouveri Cryprochirunornus fridmam Endochironornw albipennis, E. impar, E. tendens

To 43 C+-bands in karyotype Several C+-bands in the second and fourth chromosomes Several C+-bands in each chromosome (1982)

Michailova (1987a) Hagele (1977a) Belyanina and Sigareva (1978) Michailova (1987a)

Hagele (1977a), Sigareva (1981) Michailova (1987a) Michailova and Belcheva (1982) Michailova and Gercheva

Orthocladius bipunctellus Besides giant blocks of a-heterochromatin, approximately in 15 intercalary regions

Michailova and Belcheva (1982)

Smittia parthenogenetica

Two intercalary bands in the second chromosome

Hagele (1980)

7-8 bands

Kaul et al. (1989a)

Two bands in the 3B region

Tewari et al. (1983)

40 bands in polytene chromosomes

Stocker et al. (1978)

Several bands

Eastman et al. (1980)

Sarcophagidae Parasarcophaga argyrostoma

P. misera Sciaridae Rhynchosciara hollaenderi Sciara coprophila Simuliidae Simulium melatum S. orlatipes

PLANTS Phaseolw coccinew

-

Tens of C + bands Large number of C+-bands (Figure 74)

Fkdo (1975a, 1978) Bedo (1975a, 1978)

Several C+-bands of intercalary heterochromatin (see Figure 75)

Schweizer (1976)

Figure 74. A comparison of the banding pattern of chromosomes stained by the Giemsa technique (a) and orcein (b). Triangles indicate C+-bands; t, telomeres; 1L and IS, long and short arms of the first chromosome. Reprinted by permission from Bedo (1975a).

Figure 75. Polytene chromosomes of Phareolus coccineus haustorium after staining by Giemsa technique for C+-heterochromatin. After Schweizer (1976).

146

1. F. Zhimulev

a

c

( '--L

Figure 76. The chromosomes of Rhynchosciara h o b n d e r i salivary gland stained for C-heterochromatin in ontogenesis from almost diploid to maximum polyteny level observed in fourth instar larvae. X, A, B, C, designations of the chromosomes. Reprinted by permission from Stocker er al. (1978).

Holmquist, 1975b), D. ananassue (Adkisson et al., 1971), and Drosophila of the montium and ananassue groups (Lakhotia and Mishra, 1980) to 28 bands in Rhynchosciara holtaenderi (Tables 8 and 9 and Figure 77). There are wide variations in the presence of Q-bands. The set of bands is invariant in Drosophila (see Table 9), but they occur in different regions in various strains. The differences are particularly marked between related species. Some Q+-bands of the polytene chromosomes of D. melanoguster, besides Q+-staining,show additional features: the 1.672 satellite (AATAT) is identified in the 81F band with in situ hybridization (Peacock et al., 1978) and antibodies to triplex DNA are localized. The latter is derived from tracts whose one chain contains only pyrimidines and the other only purines. The usual AT and GC pairs of the duplex are retained in the triplex form. A third pyrimidine chain, which lies in the large groove of the DNA molecule, is added. This results in the formation of a triple C-G-C- contact, the purine chain then remains unpaired (Burkholderet d., 1991).

3. Staining with acridine orange When stained with acridine orange, polytene chromosomes fluoresce yellowgreen, with staining generally coincident with banding pattern: the largest bands

Table 8 Occurrence of Q+-Bands in Polytene Chromosomes of Diptera Family, species

Location and characteristics

References

Chironomidae Anatopynia dyari

Bands in the centromeric end of the fourth chromosome, weaker fluorescence in the centromeric regions of the thirdfifth chromosomes

Bedo (1974)

Chironomus thummi

Some regions in chromosome arms (4A2, 1A4a, Flef-F2ab) Approximately 10 bands fluoresce brighter than the others

Badaev et al. (1973)

Several Q+-bands in each chromosome

Michailova and Gercheva ( 1982)

Several bands included in pseudochromocenter (see Section VlI,C,2)

Yoon (1989)

D. flavomontana

A band in the base of the fourth chromosome

Sinibaldi and Ban, (1979)

D. lummei

-

D. ovivororum

A band in the proximal third of the X chromosome

Sinibaldi et al.(1976), Sinibaldi and Barr (1979) Sinibaldi and Barr, 1979

D. virilis

A band in the centromeric region of the fourth chromosome

Adkisson et al. (1971), Bart and Ellison (1971b), Holmquist (1975b), Sinibaldi and Barr (1979)

Parasarcophaga argyrostoma

7-8 bands

Kaul et al. (1989a)

P. misera misera

Two bands in the 3 8 region

Tewari et al. (1983)

Rhy nchosciara hollaenderi

28 Q+- and 17 H+-bands

Stocker et al. (1978)

Sciara copophila

Several bands

Eastman et al. (1980)

Ch. valkanovi Endochironomw albipennis, E. impar, E . tendens

Drosophilidae D. biseriata

Belcheva and Michailova (1974, 1980)

Sarcophagidae

Sciaridae

Simuliidae Bedo (197515)

Simulium pictipes

Tephritidae Ceratitis capitam

4 Q+-hands in autosornes Large number of Q+-bands in polytene chromosomes

Note. Dash indicates that species is only mentioned in reference(s).

Bedo (1986) Zacharopoulou (1990)

148

1. F. Zhimulev

Table 9 Variation in Localization of Q+-Bands in the Polytene Chromosomes of Drosophila of the melanogaster Group _________~

Chromosome region Species, strain

lOlF

102B

lO2D

lO2F

81F

D. melanogaster,

+

+

-

+

+a

“wild strain” D. melanogaster

+

-

+

-

+

-

+

-

+

+

+

-

+

-

+

+

+

+h

D. melanogaster D. melanogasrer

+ +

+

+ +

-

Hochman (1974)

+

D. melanogaster

+

D. simulans

+

-

+=

Levina, (1974, 1975) Lakhotia and Mishra (1980) Adkisson et al. (1971)

D. simulans, yw

+

-

D. simulans,

+

-

D. melanogaster, Swb-9 D. melanogaster,

83E

Vosa (1970)

Oregon-R

-

+

-

-

+

-

-

-

-

-

-

+

+

D. erecta, D.

1-2 bands in the fourth chromosome

mauritiana , D . orem, D. teissieri, D. yakuba

lordansky et al. (1971) Adkisson et al. (1971) Ellison and Barr (1971a), Mayfield and Ellison (1975)

Ellison and Barr (1971b) Ellison and Barr (1971b)

“wild strain” from Honduras

D. simulans

References

-

-

Lakhotia and Mishra, (1980) Lemeunier et al. (1978)

“The author called it 2OD. ”Fluorescencedetected in only some individuals of this strain. The authors designate it as the 83D region, but it is correctly designated as 83E in the paper by Barr and Ellison (1972). ‘4-band is much thiner than in D. melanogaster.

are the most intensely stained, and staining of puffs and Balbiani rings is hardly discernible (Nash and Plaut, 1964; Zelenin et al., 1977). When stained by a modified method (the preparations are exposed to light after staining for 3-5 days), the polytene chromosomes of Drosophila acquire orange-red fluorescence and only certain regions (81F, lOlF, 102D, 102F) still

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

Figure 77. Q-fluorescence of the 81F and 83F regions in D. melanogaster connected by a stretch of ectopic contact. Bold and thin arrows indicate the 81F and 83E regions, respectively. Reprinted by permission from Barr and Ellison (1972).

Figure 78. Differential fluorescence of the polytene chromosomes of Rhynchosciara angehe after alkali treatment and acriflavin-HN02 staining. X, C, B, chromosome designations; AR and AL, right and left arms of chromosome A. Reprinted by permission from Diaz (1973).

149

150

I. F. Zhimulev

show bright yellow fluorescence. These regions fluoresce when stained with quinacrine (Mezzanotte, 1978).

4. Staining with methyl green and pyronine After treatment with alkali and acids at high temperature, the chromosomes in squash preparations cease staining with methyl green and pyronine. However, when conditions for DNA renaturation are provided, stainability is restored in some chromosome regions of Rhynchosciara angelae. These are mainly the centromeric regions of the chromosomes and about 10 neighboring bands (Diaz, 1972). The author believes that differential staining underlies DNA renaturation in the location of repeats; the dye binds renaturated DNA.

5 . Staining with acriflavine-HNOz After treatment with alkali and staining with acriflavine-HNO,, 36 fluorescent bands are identified in the polytene chromosomes of Rhynchosciara angelae (Figure 78). In 30 of these, at least one of the following features of heterochromatin is observed: breaks, ectopic pairing, asynchronous replication, or the previously described differential staining with methyl green and pyronine (Diaz, 1973).

6. Staining with ammoniacal silver Using the ammoniacal silver staining technique of nuclei, the material of nucleoli and micronucleoli is identifiable (Stocker, 1978; Stocker et al., 1978; Ananiev et al., 1981). However, with this method, about 20 bands stain in the polytene chromosomes of Rhynchosciaru hollaenderi (Stocker et al., 1978), and several bands in D. hydei (Banerjee et al., 1988).

7. Other staining methods Besides the previous stains, mention may be found in the literature of the possibility of differential staining with simpler methods. For example, when preparations of the polytene chromosomes of ovarian nurse cells of Anopheles messeue are stored for a long time, the chromosomes, which were squashed in acetoorcein, become colorless and only regions of pericentromeric heterochromatin and some intercalary regions remain stained. The chromosomes of salivary glands cannot show differential loss of stainability (Stegniy and Sharakhova, 1990).

C. Identification of intercalary heterochromatin by morphological criteria I. Chromosomal breaks In his early studies Painter ( 1934a,b) had already observed polytene chromosome regions that frequently break. A great deal of information about this particular chromosomal breakage has been obtained (Table lo), and the identified number

Table 10 Occurrence and Characterization of Breaks in Various Species of Diptera ~~

Breaks expressed Family, species

Strongly

Weakly

References

Agromy zidae Cerodonta (Butomomym) eucaricis

The 56 region of gastric cecum chromosomes

Block (1974)

C. spinata

The 53 region on the chromosome map

Block (1976)

Trilobomyza jlawifrons

Weak spots in the chromosomes vary in different tissues

Block (1976)

Calliphoridae Calliphura erythrocephala

The chromosomes of trichogen cells

Extremely rarely in the chromosomes of ovarian nurse cells

A break detected in unsquashed nucleus

Ribbert (1967, 1979) Ribbert (1967)

c.S t y g i a

In the cells of the salivary glands and fat body, because of the abundant breaks and ectopic pairing, the chromosomes do not spread when squashed

Chrysomya bezziana

Trichogen cells

Ch. pinguis, Ch.rififacies

-

Puchalla (1994)

Cochliomyia hominiworax

Trichogen cells

Dev et al. (1985)

Lucilia cuprina

In the chromosomes of trichogen cells

Childress (1969)

L. sericata Chironornidae Ch. dorsalis, Ch. pseudo-

Trichogen cells

Ribbert (1979)

In the salivary gland chromosomes

Michailova (1989a)

In the trichogen cells

Thompson (1969)

Bedo (1992)

thummi, Endochironomus mpar (continues)

F

ln N

Table 10 (Continued) Breaks expressed

Family, species

Strongly

Weakly

References

Smittia parrhenogenetica

Induction of break formation in salivary gland chromosomesafter y-ray irradiation -

Telmtogeton sp.

In the salivary gland chromosomes

Newman (1977)

In the salivary gland chromosomes Very “fragile”chromosomes of salivary glands

Chaudhry (1981)

Ch. thummi thummi

Culicidae Anopheles hectoris C&x wishnui

No breaks in the chromosomesof salivary glands

Gunderina and Aimanova (1997) Bauer (1970)

Baker et al. (1966)

hsophilidae

D. auraria

Same as C. wishnui

D. b u m u t a

-

D. biseriata D. erecta

Same as C. wishnui

D. funebns

In the salivary gland chromosomes

D. gibberosa

Breaks in the salivary gland cells frequent in the middle part of the second chromosome

Scouras (1984, 1986) Mavragani-Tsipidouet al. (1992)

Break frequencies in female salivary gland chromosomes in 64C+5,67A1-4,67Dt?-13,70A1-2, 7CC-2, 71C1-2, and 75C1-2 regions were (%I: 68.4, 36.1,38.9,0, 61.7,69.0, and 73.8, respectively

Ymn and Richardson (1978a) Zhimulev et d. (1982)

Slizynska and Slizynski (19411, Beermann (1962), Xnyakov (1965) No breaks encountered in the fat body cells in this region; there are three broad dark bands instead

Roberts and MacPhail(1980)

Roberts and MacPhail(1985)

D. guanche D. hydei D .hysaicosa

In the salivary gland chromosomes Same as D. gwnche Same as D. gwnche

Ananiev and Barsky (1982) Yoon and Richardson (1978a)

D.mndeirensis D. maurinhna

Saem as D. guanche

Papaceit and Prevosti (1991)

Break frequencies in female salivary gland chromosomes in 64C4-5,67A1-4,67D%13,70A1-2, 7E1-2, 71C1-2, and 75CI-2 regions were (90): 91.4, 3.6,80.0, 2.9,85.1,90.4, and 95.9, respectively Salivary gland chromosomes

Zhimulev et al. (1982)

D. mehogaster

Moho et al. (1987)

Breaks occurred in 11 regions: 11A, IZDE, 19E, 33A, 35E, 36DE, 42B, 64C, 70BC, 75C, and 89E Breaks in 19 regions Breaks detected in 62 regions In the X chromosomes of salivary glands of females, break frequencies in the 3C, 1 1 A 6 9 , 12E8-9, and 19E1-4 regions were (the results of two experiments expressed as %): 65.446.7,92.9-99.3, 86.2-92.0, and 80.693.8, respectively In the weak spot between the 39D3 and 39E1-2 bands (the histone gene locus), a hne granular network was located

In the same regions of males, break frequecies were 0, 0, 0-1.4, and 2-2.6, respectively

Painter (1934a,b), Bridges (1935c, 1938), Tinyakov (l936), Prokofyeva-Belgovskaya (1939d, 1941). Beermann (1962). Korge (1987). Hochstrasser and Sedat (1987), Hofmann et d. (1987). Sorsa (1988a, b) Kaufmann and Iddles (1963) Lefevre (1976) Zhimulev and Kulichkov (1977), Zhimulev et al. (1982) Zhimulev et al. (1982)

Semeshin et d.(1985)

Table 10 (Continued) Family, species

Breaks expressed Strongly

Weakly

References Shilov and Zhimulev (1986)

In serial sections of 12 unyuashed salivary gland nuclei, 25 breaks were detected Weak points were found in polytene chromosomes of prothoracic gland, hindput, and midgut in the same places as previously mapped in salivary glands; some of the constriction were observed in intact unfixed midgut nuclei Under the effect of modifiers of position effect variegation (temperature 18°C and decrease of heterochromatin amount in the genome), break frequencies increase

Hochstrasarr (1987)

Zhimulev et al. (1989a.b)

In the chromosomes of hindgut break frequencies considerably decreased In the fat body cells breaks occur less frequently than in salivary glands Breaks occur infrequently in the ovarian nurse cells in mutants: fs(Z)B, om', otu' In pseudo-nurse cells of otul I mutants, weak points were found in 45 regions, but their frequency is substantially lower then those in salivary glands

'

Zhimulev and Kulichkov (19771, Zhimulev et al. (1982) Richards (1980)

H. Gyurkovics (1986, personal communication), Heino (1989,1994) Mal'ceva et al. (1995); Mal'ceva and Zhimulev (1997)

Wilson et al. (1969)

D. nasura D. nebulosa D. o r e m

Salivary gland chromosomes

D. pallLirhennis D. puranaensis

Same as D. nasuta Same as D. nasuta

D. pseudoobscuru D. robusta D. simulans

Same as D. m u t a Same as D. nasuta In the female X chromosome of the salivary glands, break frequencies in the 3C, 9A1-4, llA6-9, 12E8-9, and 19E1-4 regions were (%): 70.0,29.7, 82.1,47.4, and 85.0, respectively

D. subobscura

63 break regions in the X chromosomes of salivary glands 7 sites of breaks in the X chromosome

de Frutos and l'ascual (1985), Molto et al. (1987) de Frutos et al. (1987)

D. teissieri

Break frequencies of the female salivary gland chromosomes in the 64C4-5,67A1-4,67D8-13, 70A1-2,7CC-2,71C1-2, and 75C1-2 regions were (%): 87.4, 17.6,30.9,2.2, 51.9, 71.7, and 85.5, respectively

Zhimulev et al. (1982)

D. virilis D. virilis D.willistoni

The salivary gland chromosomes

Fujii (1942)

The salivary gland chromosomes

Hughes (1939)

The salivary gland chromosomes

Dobzhansky (1950)

Same as D. nasuta

Pavan (1946b)

Same as D. nasuta

Lemeunier and Ashbumer (1984) Dobzhansky (1944) 1. F. Zhimulev (unpublished observations) Koller (1935) Carson and Stalker (1947) In the same regions of the male, break frequencies were 1.2,0, O,O, and 1.8. respectively

Zhimulev et al. (1982)

(continues) r

LJl LJl

Table 10

(Continued) ~

Family, species D.Yaw

~

~

~

~

~~

~

Breaks expressed Strongly Break trequencies ot the female salivary gland chromosomesin the 64C4-5,67A1-4,67D%13, 70A1-2,7oC1-2.71C1-2and 75C1-2regions

Weakly

References Lhimulev eta(.

(m

were(%):88.7,37.1,68.5,0,82.1,71.4,and92.4, respectively Muscidae Musca domestics

In the salivary gland cells, because of the numerous

breaks and ectopic pairing, the chromosomesdo not spread when squashed Phryneidae Phryne cincta

The chromosomesof the salivary glands In the chromosomesof the salivary glands, breaks occur in blocks of a-heterochromatin Break frequenciesare modified by temperature: at 2°Cbreaks occur in 86% of the chromosomes; at 18°C they occur in 16%

Sarcophagidae Sarcophaga b u k

In the pupae trichogen cells, the chromosome breaks were less frequent and the preparations were good

In chromosomesin foot pad cells of pupae

In midgut cells breaks occur twofold less frequently

Beermann (1962)(see review in Zhimulev (1992b,1996)

Wolf (1957) Wolfand Sokoloff (1973)

Whitten (1969),Bultmann and Meuanotte (1987)

Sciaridae Rhynchosciara angelne

Rh. hollamderi

T~icho~ia pubescens Stratiomyiidae (representatives of the family) Tephritidae Batrocera okae (Dacus okae) Ceratitis capitata

In supergiant cells developed as a result of intranuclear infection (see review in Zhimulev, 199213, 1996) in chromosome A, there are 5 regions of local DNA underreplication

These “weak spots” are absent in the normal chromosomes of the salivary glands, cecum, and malpighian tubules

Diaz et al. (1969). Pavan et nl. (1971)

Stocker et nl. (1978)

The chromosomes of salivary glands Salivary gland chromosomes

Amabis ( 1983)

O n l y very young larvae have extended chromosomes;

Melland (1942), Maim (1949)

in older larvae the chromosomes are broken down into small fragments as a result of numerous breaks Polytene chromosomes of salivary glands and Malpighian tubes The chromosomes of hindgut, orbital, and trichogen cells

Zambetakiet d., (1995)

Xpulidae (representatives of the family)

Same as in Stratiomyiidae

Melland (1942), Mainx (1949)

Volucellinae (representatives of ‘the subfamily)

Same as in Stratiomyiidae

Maim (1949)

Note. Dash indicates that species is only mentioned in reference(s).

Bedo (1986, 1987a), Zacharopoulou (1987,1990)

158

1.

F. Zhimulev

of breaks has continued to grow over the years. Tinyakov (1936) and ProkofyevaBelgovskaya (1939d) described 1-2 regions where breaks occur; later, the identified number was 11 (Kaufmann and Iddles, 1963), then 19 (Lefevre, 1976), then at least 62 (Zhimulev and Kulichkov, 1977; Zhimulev et al., 1982). The breaks are very different in morphological appearance and they have been variously called the “weak spots,” the “offsets” (Bridges, 1935c), “breaks” (Prokofyeva-Belgovskaya, 1941), “narrowed places” (Carson and Stalker, 1947), the “discontinuity,”“ruptures,”“weak opens” (Kaufmann and Iddles, 1963), “constrictions” (Lefevre, 1976), and “sharp constrictions” (Ribbert, 1979). The results of light and electron microscopic studies (Zhimulev et al., 1982) have allowed us to distinguish break regions according to morphology:

1. Constrictions (Figure 79a). These conform to Lefevre’s “attenuation” fiber. The fiber is most frequently derived from the material of a single band (e.g., from the proximal part of 64C4-5; see Figure 79a) or from the facing neighboring bands (e.g., 11A6-7 and 11A8-9; Figure 80b). 2. The true break. This is the same as a constriction except that the broken ends are not connected by a fiber. A break of this type is shown in Figure 80d: the parts of the 11A6-7 and 11A8-9 bands connecting the fiber are hardly discernible. 3. Fissure. An incomplete manifestation of a break, this is the emergence of a cleft within a band, for example, in the 86D1-2,89E1-4, 71C1-2, and 75C1-2 (see Figure 79b and 79d-f) or the 25A1-4 and 11A6-9 (see Figure 80d and 80e) regions. In such cases, bands frequently assume the shape of the Latin letter “V” as Kaufmann and Iddles (1963) observed. Fissures do not damage the nearby chromosomes, so the “weak spot” can be very precisely localized. For example, in the 71C1-2 band (see Figure 79d), its undamaged part is continuous at the breakage site; however, two newly arisen bands are quite distinct. The breaks do not occur between “doublets” as Lefevre (1976) claimed, but rather within a single band. 4. A shift. This is a break in which the broken parts of the chromosomes, although appearing to be reunited, are shifted along the long axis, for example, in the 86D1-2 (see Figure 79b), 12E8-9 (see Figure 79c), and llA6-9 (see Figure 79c) regions. The location of a true break within a band is highly specific; it may be located at one of the edges of a band (75C1-2,64C4-5), in its middle (86D1-2, 7OC1-2,1 lA6-9), or in different regions of the same band (71C1-2 in Figure 79d and 79e). In the latter case, the break can occur to the left or right of the band or in the middle of a band and, what is noteworthy, in the same chromosome (Zhimulev et al., 1982). There are great sex differences in the morphology of break regions in the

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

159

Figure 79. Morphological types of breaks in Batumi-L larvae. (a) Constrictions. (b and c) Breaks of bands with weak (86D1-2 in b) or strong (1288-9 in c) transversal displacement of parts of th chromosome. (c) Complete break in the 1l A 6 9 band. (d) Partial breaks (fissures) of the 89E1-4 and 71C1-2 bands. (e) Multiple breaks distal and proximal to the 71C1-2 band. (0 Break distal to the 75C1-2 band. Arrow in c indicates the stretch of ectopic contact between the 12E1-2 and 12E8-9 bands. The scale is 10 p n (a) and 1 p,m (b-f). Reprinted by permission from Zhimulev et al. (1982).

X chromosome. According to Bridges’ map, there are two “doublets” in the 11A region in females, llA6-7 and 11A8-9, with a break (see Figure 80d) or a fiber (see Figure 80b) between them. In these cases, when there is no break, one thin 11A6-9 band is present in the region. It is much thinner than the neighboring 11A1-2 and 11B1-2 bands (see Figure 80a). The pattern is entirely different in the male. In all the chromosomes studied at the EM level (more than 30), there is a single band between 11A1-2

160

1. F. Zhimulev

Figure 80. Morphology of the 11A6-9 region in females of Batumi-L strain. Reprinted by permission from Zhimulev er at. (1982).

and 11A11-12; it is, however, much more bulky than the llA1-2 and llB1-2 marker bands mentioned previously (Figure 81). Neither two- to threefold stretching nor greater magnification under the electron microscope allow one to discern the two bands in the region, much less the two “doublets” (see Figure 81). Breaks are not detectable in the same region of a chromosome in all the cells of the salivary gland, with their frequencies being characteristic of each region (Kaufmann and Iddles, 1963;Lefevre, 1976; Zhimulev and Kulichkov, 1977). For example, of 200 cases of breaks in the 3C, 11A, and 12Eregions, 53%were assigned

Polytene Chromosomes, Heterochromatin, and Posltlon Effect Variegation

161

Figure 81. Morphology of the 11A6-9 region in males of Butmi-L strain. Reprinted by permission from Zhimulev et al. (1982).

to 11A, 36% to 12E, and 11% to 3C (Kaufmann, 1944). Breaks are nonrandomly distributed within the chromosome, being restricted to specific regions (Figure 82). The data in Table 10 indicate that breaks frequently occur in representatives of different families. Beermann (1962) noted that breaks are not detected in the Chironomus species and the Simuliidae midges. With regard to the latter, no evidence facts of breaks has so far been observed in the Simulid midges, and they have been identified only in rare instances in chironomids despite careful studies (see Table 10). It is Beermann’s opinion that absence of “weak spots” in Chironomids and Simuliidae cannot be taken to mean that they lack intercalary heterochromatin because “stretchability” and “breakability” can be different in different species. The hypothesis has long been held that, by analogy with centromeric heterochromatin, breaks result from local DNA underreplication (Barr and Ellison, 1972;Woodcock and Sibatani, 1975; Lefevre, 1976; Spofford, 1976; Ananiev et ul., 1978). Differences in the thickness of the 1lA6-9 band (see Figures 80 and 81) and, consequently, in DNA amount are obvious so that there is no need for measurements. Counts of silver grains over the chromosome regions in larvae grown in a [3H]thymidine-containingmedium revealed that DNA amount in the 11A6-9 region is approximately fourfold greater in males, where breaks are not identified, than females (Zhimulev et ul., 1982). According to the data from Southern blots, the DNA fragment-located in the middle part of the 11A6-9 band and hybridizing in situ, containing ectopic pairing fibers, and stretching beyond this band in the polytene chromosomes of the salivary glands-constitutes only 25% of its amount in the cells of imaginal discs (Lamb and Laird, 1987). The histone genes in Drosophila are arranged in clusters with a total length of 500 kb; the cluster is divided into two groups of thick bands in the 39D

162

1. F. Zhirnulev

nr160 A

b

1

5

2

?(I

2

Figure 82. Locations of all types of breaks in the X chromosomes of salivary gland larvae of giant strain (a), complete breaks in Batumi-L strain (b), regions of late replication (c), ectopic pairing (d), strong synapsis (e), breakpoints of chromosomal re-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

163

and 39E regions (Pardue et al., 1977). Breaks frequently occur between these groups of bands, that is, in the middle part of the cluster (Zhimulev and Kulichkov, 1977; Zhimulev et al., 1982). Judging by the results of blot hybridization, there are at least twofold fewer (by 50-67%) histone genes in the salivary gland nuclei compared to the diploid cell tissues of pupae (Lifschytz and Hareven, 1982b). Quantitative autoradiography in conjunction with in situ hybridization identified only 18% of copies of the histone genes in larval salivary glands compared to imaginal bands (Hammond and Laird, 1985b). In blot hybridization experiments, the level was 31% (in the 1 8 4 3 % range from one experiment to another). Polytenization level of the histone genes in fat body cells was 34% (Lamb and Laird, 1987). The bithorax (Bx-C)genetic complex is located in the region where the 89E1-4 break regularly occurs (Lindsley and Zimm, 1985). In an early molecular study of this region, Spierer and Spierer (1984) mentioned underreplication of certain bx sequences in the 89E1-4 region. One of the DNA fragments of a Bx-C complex, labeling the thinnest part of the constriction of the 89E1-4 break, consitutes only 22% of DNA in the polytene chromosomes of the salivary glands, and 89% of that in the cells of imaginal discs (Lamb and Laird, 1987). The numerous variations in break frequencies and, hence, in DNA underreplication include

1. Interspecific differences. A comparison of break frequencies in the closely related species of Drosophila of the rnelanogaster group (see Table 10) revealed variations ranging from 8% in D. mehogaster to 37.1% in D. yakuba for the 67A1-4 region, and ranging from 30.9% in D. teissieri to 80% in D. muuritiana for the 67D3-8 region. Variations from 20% to 30% were also found for other regions (Zhimulev et al., 1982). 2. Sex differences. In males of Drosophila breaks virtually do not occur in the X chromosomes (with the exception of the 20AF region, for which there are no quantitative data). It is noteworthy that the two sexes do not differ in break frequencies in the autosomes (Zhimulev et al., 1982, 1989a,b). In the male X chromosomes of D. madeirensis breaks were not found either (Papaceit and Prevosti, 1991). 3. Effect of position effect modifiers. Low temperature considerably promotes arrangements (0,and distribution of silver grains after in situ hybridization with total ( h ) and complementary (g) RNA. Ordinate: (a and b) Break frequencies (n = number ofchromosomes examined). (c) Degree of late replication (for explanation see Section XVI1,c). (d) Number of ectopic contacts in the region (n = total number of ectopic contacts studied). (e) Number of events when asynapsis is ended in the given region (n = total number of asynapsis events in the X chromosome). (f) Number of breaks of chromosomal rearrangements. (g and h ) Average number of silver grains in the region for n chromosomes. Black asterisk, no data; white asterisk, regions where more precise localization is difficult. After Zhimulev and Kulichkov (1977) and Zhimulev et al. (1982).

164

1. F. Zhimulev

the expression of chromosome “fragility” in the weak spots. At 14“C,break frequencies increase at all their location sites in the male X chromosomes. The increase in the 19E region in males is particularly conspicuous: from 5% at 25°C to 55% at 18°C. Decrease in heterochromatin amount (removal of the Y chromosome) in the nucleus produces a sharp increase in break frequencies. The frequencies are highest under the combined effect of removal of the Y chromosome at 18°C (see Figure 154 in Section XVII). There is circumstantial evidence concerning the action of genetic modifiers of position effect. A comparison of strains containing En-uur( 3)2 and Su-vur(3)9 revealed higher break frequencies in larvae having an enhancer (Zhimulev et d., 1989a,b). Exposure of different instar larvae to low and high temperature did not reveal a clear-cut temperature-sensitive period in break formation. Larvae are sensitive to temperature shifts at all the polytenization stages (Zhimulev et uf., 1989a,b). 4. Tissue differences. Table 10 presents numerous examples of tissue differences in break frequencies. In representatives of the Calliphoridae, Sarcophagidae, and Muscidae families, polytene chromosomes of most larval organs break down into pieces when squashed, they do not spread, being interconnected by ectopic pairing fibers. In some cell types (foot pad and trichogen), the chromosomes remain intact and amenable to cytogenetic analysis. In Drosophilidae, breaks occur rarely in organs composed of low polyteny cells (Zhimulev and Kulichkov, 1977). In cells where breaks are less frequent, the bands with breaks are more massive, presumably because they do not underreplicate. Examples are 11A6-9 in the salivary gland chromosomes of males and the bands of chromosome 3L in the hindgut chromosomes (Figure 83). The 89E1-4 region (the location site of the Bx-C) in the fat body chromosomes is represented by a single dense band in contrast to the counterpart region in the salivary glands (Richards, 1980). Molecular biology data also provide evidence that underreplication is tissue specific. The percentages for the histone genes in the salivary glands and fat body chromosomes are almost the same (31% and 34% of that in diploid cells), whereas the underreplication degrees of the Bx-C gene region in these cells are 78% and 11%, respectively (Lamb and Laird, 1987). Histone DNA sequences replicate only partially during polyploid nurse cell growth, while 5s sequences fully replicate (Hammond and Laird, 1985a). Unusual chromosome breaks have also been described. The relation of breaks to heterochromatin is unclear in two cases. In a D. virilis stock that had long been maintained in the laboratory, several individuals with more fragile polytene chromosomes were found (Roca and Rubio, 1984). During the making of squash preparations, slight pressure applied to the glass slide suffices to disrupt the

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

165

Figure 83. Differences in relative thickness of the 7OC-2,

71C1-2, 74A1-5, and 75C1-2 bands in the chromosomes of salivary gland cells ofgiant strain larvae (a) and hindgut of DfC 1 )@/Dpt~+Yy+ strain larvae (b). Reprinted by permission from Zhimulev et d. (1982).

integrity of the chromocenter. Under the pressure usually applied to obtain normal preparations, the chromosomes fall apart into fragments without interconnections. The breakpoints in the chromosomes are randomly distributed. Data on their relation to intercalary heterochromatin have not been provided. Crosses of strains with the frailer chromosomes to normal strains demonstrated that "fra" (frailty) behaves as a recessive and an X-linked trait (Roca and Rubio, 1984). During the making of squash preparations, the chromosomes frequently break down. It was found that, in several strains (y2 sn m, ywsn3, Batumi-L/y2 sn) of D. mehogaster, breaks occur in the region of the 2B puff in 3 8 4 5 % of larvae; the respective values are 13-15% for the other strains. The frequency decreases at low temperature; it is 55% at 30"C, 38.7% at 25"C, and 16.6% at 18°C in y2 sn m males; a similar tendency was observed for y2 sn m/O males.

166

1. F. Zhimulev

At the same temperature, breaks in 2B frequently occur in larvae without the Y chromosome: in 12-16% of larvae in yz sn m/Y and in 33% of larvae in y2 sn m/O at 18”C,and in 38% in the former group and in 53% in the latter group at 25°C (M. L. Balasov, E. S. Belyaeva, and I. E Zhimulev, unpublished observations). It is known that breaks occur in the chromosomes, but they are very rarely inducible. In 10-hr larvae of Ch. th. thummi fed with 5-fluorouracil 2-deoxyriboside, an inhibitor of thymidilate synthetase in fourth instar larvae, breaks and decrease in chromosome diameter occur. Their location is coincident with sites of late replication (Hagele, 1971).

2. Ectopic pairing Bridges (1935c, 1936) noted that the contacts of various chromosome regions giving rise to loops make difficult the analysis of banding patterns shown by chromosome 2L of Drosophila. In nuclei with well-spread loops, the edges of bands involved in contacts spread during the formation of loops, joining with the edges of the other bands (Figure 84a). The length of the ectopic strands can reach several tens of microns, and they are readily identified under the light microscope (Ashburner, 1980; Ananiev et al., 1981; Scouras, 1984). In unsquashed nuclei of the salivary glands of Acricotopus lucidus and Chironornus sp., the ectopic fibers are as long as 45 pm, and are 0.2-0.5 pm thick (Quick, 1980). In some cases, contact nodes are formed. These are temporary tight contacts between some chromosome regions as well as constant contacts. For example, in D. biseriata (Figure 85), seven chromosome regions form a constant contact node, the so-called pseudo-chromocenter, and individuals of this species have two chromocenters, one resulting from pairing of centromeric and the other from pairing of intercalary heterochromatin. In some species, the ectopic contacts are so numerous that the chromosomes cannot be enfolded during the preparation of squashed samples. For example, in D. griseolineatu the whole X chromosome is represented by an irregular chromatin body (Figure 86). In Sciaridae, Calliphoridae, Muscidae, and Culicidae species, the chromosomes in a number of tissues are so firmly held by ectopic pairing fibers that they do not spread when squashed. For this reason, when a squash preparation cannot be made, we may regard ectopic pairing as tight. Ectopic pairing fibers can also form between the nearest neighboring bands, for example, at the 12E8-9 (see Figure 79c) and 98C1-2 breaks (see Figure 84b), and they can be identified with certainty with the aid of the electron microscope. “Cryptic” is a specific type of ectopic contact detectable only in EM sections. The neighboring bands with cryptic contacts are connected with fibers passing within the chromosome (see Figure 84 and also Saura et al., 1989; the 68EF region in Figure 1la).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

Figure 84. Morphologiocal types of ectopic contacts. Black arrows indicate an ectopic stretch (a and e), contacts between neighboring bands (b), and a tangle of ectopic contacts between the 86D1-2, 84D, and 9 4 A 1 4 regions (c). White arrows indicate contacts concealed within the body of the chromosome. ( a x ) reprinted by permission from Zhimulev et al. (1982); (d) reprinted by permission from Berendes (1970); (e) after Scouras and Kastritsis (1988).

167

168

1. F. Zhimulev

Figure 84. Continued

The data of Table 11 support the frequent occurrence of ectopic pairing. In spite of thorough karyological studies in numerous representatives of the Chironomidae and Simuliidae families (see Zhimulev, 1992b), ectopic contacts have rarely if ever been identified. Some remarkable ectopic associations are the contacts of nucleoli or their fragments with chromosome regions, with nucleolar material contacting the

Polytene Chromosomes, Heterochromatin, and Position Elfect Variegation

169

Figure 84. Continued

chromosome side-by-side, hanging on long fibers and entering into chromosome cavities produced by breakages (see Figure 87 and Table 12). The data in Table 12 provide evidence for the numerous contacts of the nucleolar material with the chromosomes, and for the contacts established most frequently with bands of intercalary heterochromatin. The properties of ectopic pairing are as follows:

1. Light (Slizynski, 1945; Kaufmann and Iddles, 1963; Zhimulev and Kulichkov, 1977; Ashburner, 1980; Ananiev et al., 1981;Scouras, 1984), and electron microscopic (Zhimulev et al., 1982; Ananiev and Barsky, 1985; Sorsa, 1988a,b) data converge in demonstrating that the bands involved in ectopic contacts do not differ in morphology from the other bands, with the exception of break occurrence. 2. The location of the ectopic pairing regions in polytene chromosomes is nonrandom. Figure 89 presents a map of ectopic contacts in a chromosome of Drosophila. Maps of this kind (Slizynski, 1945; Kaufmann et al., 1948; Kaufmann and Iddles, 1963; Scouras, 1984; Scouras and Kastritsis, 1988)allow us to see how the ectopic contacting bands are located and also how each band associates within the nucleus. Not every band is involved in ectopic associations. A particular break frequency is inherent in each band (see Figure 82d). A total number of 200-300 bands is involved in ectopic contacts in various species of Drosophila (see Table 11). Presumably these are underestimates, because without additional use of EM it is difficult to reveal the “cryptic” ectopic contacts and the contacts between the adjacent bands. 3. Ectopic pairing occurs in living unsquashed cells (Wolf and Sokoloff, 1973; Quick, 1980; Shilov and Zhimulev, 1986; Hochstrasser and Sedat, 1987a).

Figure 85. Chromocenter (black arrow) and “pseudo-chromocenter” (light arrow) in the nuclei of the salivary glands of lFrosophikz biseriara. X, 2 , 3 , 4 , 5 , 6 ,chromosome numbers. After Yoon and Richardson (1978a) and Yoon (1989).

Figure 86. Numerous ectopic contacts in the X chromosomes of a male (a) and a female (b) in the two closely related species D. mediosmata (a) and D. griseolineata (b). After Kastritsis et al. (1970).

171

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation Table 1 1 Occurrence and Characterization of Ectopic Pairing in Polytene Chromosomes of Various Species Family, species

Occurrence and characterization

References

DIPTERA Calliphoridae Calliphora erythrocephala

Ectopic contacts occur frequently in the chromosomes of trichogen cells, and extremely seldom in ovarian nurse cells

Ribbert (1979)

c.sty&

The chromosomes of larval salivary glands and fat bodies spread poorly because of abundant ectopic pairing; good preparations are made from the chromosomes of trichogen cells

Thomson, 1969

Cochliomyia hominivorax

-

Devetal. (1965)

Lucilia cuprina

Preparations of polytene chromosomes are made only from pupal trichogen cells, not from salivary gland or foot pad cells

Whitten et al. (1975)

Cecidomyiidae Wyattella ussuriensis Chironomiidae Acricotopus lucidus

Grinchuk (1977) In unsquashed nuclei the chromosomes are attached to the nuclear membrane and to each other by stretches 0.24.5 pm in diameter and 45 p m long

Quick (1960)

Chironomus gr. defectus

A remarkable case detected when the position and configuration of all the chromosomes on two squash preparations made over an interval of 7 years were completely identical; this provided evidence for the nonrandom position of the chromosomes in the nucleus, possibly due to ectopic contacts

Miseiko and Popova (1970)

Chironomus sp.

In unsquashed nuclei, numerous stretches of ectopic pairing up to 45 pm in length and 0.2-0.5 p m in diameter

Quick (1980)

Anatopynia dyari

Bedo (1974)

Gavrila (1963)

Ch. species Ch. tentans

lntrachromosomal stretch of contact

ten Tusscher and Derksen (1982)

Ch. thummi

Ectopic contacts between Q+-bands

Badaev et al. (1973)

Ch. uaialkanowi Clinotanypus nerwosus

Belcheva and Michailova (1980) The chromosomes of salivary glands are tied in a tight knot, and they are unraveled with difficulty when squashed

Pavlova and Bely anina (1972)

(continues)

172

1. F. Zhimulev

Table 11 (Continued) Occurrence and characterization

Family, species

References

Endochironomus albipennis

-

Michailova and Gercheva (1982), Petrova and Michailova (1989)

E. impar, E . tendens

-

Michailova and Gercheva (1982)

Glyptotendipes barbipes

Ectopic pairing between chromosome regions hybridizing with 18s and 26s rRNA

Wen et d. (1974)

Orthocladiinae Species (Chironomidae) subfamily

Because of ectopic pairing, the chromosomes quite frequently do not spread; associations identified: telomere-telornere, telomerecentromere, telomere-intercalary regions

Michailova (1989a)

Orthocladius thienemanni

-

Michailova (1985)

Smittia parthenogenetica

No ectopic contacts or, in the rare case, very infrequent such contacts

Bauer (1970)

Telmatogeninae Species (Chironomidae) subfamily

Same as Orthocladiinae

Michailova (1989a)

Newman (1977)

Telmatogetonsp. Culicidae Aedes aegypti

Although polytene chromosomes were identified in many tissues, n o squash preparations were obtained because the chromosomes failed to spread; variations in temperature (1S-ZBoC),addition of yeast or RNA to the medium not helpful

Sr. A. L. Mescher in Rai and Hartberg (1975)

Anopheles stephensi

-

Culex (representatives of the genus)

Polytene chromosomes spread poorly because of numerous “transversal” associations

Barr (1975)

Orthopodmayia pulcripalpis

9 pairs of regions involved in contacts

Munstermann et al. (1985)

Sabethes cyaneus

Mittal and Dev (1979), Redfern ( 1981c)

Munstermann and Marchi (1986)

Drosophilidae

D. ananassae

323 bands can be involved in ectopic pairing; according to preliminary data, 55.7% of puffs (39 of 70) involved in ectopic contacts

Duttagupta et al. (1973)

(continues)

173

Polytene Chromosomes, Heterochromatin, and Position Eflect Variegation Table 11 (Continued) Family, species

Occurrence and characterization

References

D. aurariu

225 bands, as well as both Balbiani rings and the 73B puff in active state, can be involved in ectopic pairing; intercalary heterochromatin of male X chromosome is involved in ectopic pairing

Scouras (19841, Scouras and Kas tritsis (1988)

D. biseriata

Two regions of both the second and the fifth chromosome and three regions of the X chromosome in 100% of cells are involved in tight ectopic contact, forming a “pseudochromocenter” (see Figure 85)

Yoon et al. (1972), Yoon and Richardson (1976, 1978a), Carson and Yoon (1982), Yoon (1989)

D. funebris Approximately 40 regions involved in ectopic contacts Ectopic pairing between two clusters of RNA genes observed in one of the 50 examined nuclei

5s

Slizynska and Slizynski (1941) Tinyakov, 1965 Cohen (1976a)

D. furoifacies

A “pseudo-chromocenter”, like that in D. biseriata, formed in 75% of cells (see earlier)

Yoon and Richardson (1978a)

D. griseolineata

Tight ectopic pairing between the numerous regions of the X chromosome in females, as a result of which it does not spread and lies as a clump in the preparation; in males, ectopic pairing in the chromosome is considerably weaker (see Figure 86)

Kastritsis et al. (1970)

D. gr. guarani

In species of this group the chromosomes do not spread when squashed

King (1947)

D. hysmcosa

“Pseudo-chromocenters” (see D. biseriata, D. furwifucies, earlier) formed in 75% of cells

Yoon and Richardson (1978a)

D. mediosniata D. melanogaster

Same as D. griseolineata

Kastritsis et al. (1970)

Contacts between regions of chromosome 2L

Bridges (1935c, 1936, 1938) Prokofyeva-Belgovskaya (1938, 1939d, 1986)

8 regions of the X chromosome (2B, 3C3,3C6, 4C3-7, 11A, l2EF, 15F, 19E3-4) ectopically pair with telomeres and centromeric heterochromatin 180 regions are involved in contacts with telomeres Intrachromosomal ectopic strand in the 4D region (see Figure 84d)

Hinton (1945) Berendes (1970)

(continues)

174

1. F. Zhlmulev

Table 11 (Continued) Family, species

Occurrence and characterization

References

B. P.Kaufmann and 290-350 bands involved in ectopic contacts; M. K. Iddles in connection of regions in the same chromoKaufmann et al. some occur more frequently than between different chromosomes (1948) Kaufmann and Iddles Numerous ectopic contacts detected in orceinstained unsquashed nuclei (1963) Kaufmann (1965) Ectopic contacts located near puffs; of the 71 studied regions, only in 12 was the region of ectopic contactdistanced from the puff by more than 10 bands In individuals with Q+-band in the 83E region, Barr and Ellison frequency of ectopic pairing of this region (19721, Mayfield and Ellison (1975) with the other Q+-staining 81F hand increases 40-fold compared to individuals without Q+.hand 201 chromosome regions of the 220 letter subdi-- Kulichkov and Zhimulev (1976); visions of the map are involved in ectopic pairZhimulev and Kuing; contacts are most frequent between the close-lying regions of the same chromosome; the lichkov (1977), Zhimulev et al. frequencies of ectopic pairing are considerably reduced in male X chromosome; certain com(1982) combinations of contacts appear with higher frequency than others-these are contact nodes Lefevre (1976), Ashburner (1980), Korge (1987), di Pascuale Paladin0 et al. (1988), Sorsa, 1988a,b), Vagapova (1991), lvashchenko et al. (1991) Richards (1980) No ectopic contact in the proximal part of chromosome 2L found in the fat body cells (for comparison with salivary gland cells, see Figure 89) Ectopic contacts revealed after staining with Ananiev et al. (1981) ammoniacal silver Based on EM results, it was suggested that Ananiev and Barsky ectopic contacts were “produced by recombi(1985, p. 13) nation between homologous DNA sequences located in different chromosomes,” for example, between the 85D and 19E regions; chromomere pattern was found in stretches of ectopic pairing (continues)

175

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 11 (Continued) Family, species

Occurrence and characterization Ectopic contact detected in a section of unsquashed nucleus In unfixed nuclei, ectopic stretches seen in the 11A, 1ZF,35CD, 36CD, 56F, and 89E regions The 10A and 11A regions involved in ectopic contacts at frequencies of 12% and 22% in the agnostic mutants and at 3.5% and 7% in wild Canton-S strain Ectopic conjugation was found in polytene chromosomes of mutant otu’ pseudo-nurse cells, but with much lower frequencies then in salivary gland cells Neo-Y chromosome is strongly bent and twisted, probably due to accumulation of ectopically contacting sites; normal X2 homolog does not show such “stickness”

’

D. miranda

D. mitchelli

References Shilov and Zhimulev (1986) Hochstrasser and Sedat (1987a) Medvedeva and Savvateeva ( 1991)

Mal’ceva et al. (1995), Mal’ceva and Zhimulev (1997) Steinemann (1982a)

“Pseudo-chromocenters”(see D. biseriata earlier) formed in 50% of cells

Yoon and Richardson (1978a)

D. w u t a subgroup D. nebulosa D. orena

“Loops” at the base of chromosome 2R

Wilson et al. (1969)

-

Pavan (1946b)

Very poorly spreading X chromosomes because of abundant ectopic contacts

Lemeunier and Ashburner (1984)

D. subobscura

151 sites of ectopic contacts

de Frutos and Pascual (1985), de Frutos et al. (1987)

D. tumiditarsis

Sinibaldi and Cummings (1981)

Zaprionus multisniatw

Sciandra et al. (1973)

Glossinidae Glossina mursitans morsitans

Southern et aL(1973)

Muscidae Musca domestica

Numerous contacts resulting in unsquashing polytene chromosomes

Mycetophilidae Tipula agmici Phcyneidae Phryne cincta

Vecchi and Ruhini (1973), Sharma et al. (1979),El Agoze er al. (1992) Cavrila (1983)

Ectopic contacts between blocks of a-heterochromatin exist in unsquashed nuclei

Wolf and Sokoloff (1973) (continues)

176

1. F. Zhimulev

Table I1 (Continued) Family, species Psychodidae Lutzomyia longipalgis

Occurrence and characterization The chromosomes are twisted and presumably associated with each other by ectopic pairing; they do not spread when squashed

References White and Killick. Kendrick (1975)

Telmatoscopusalbipunctatus

Troiano (1975)

Telmatoscopus sp.

Amabis and Simoes (1972)

Rhagionidae Atherix ibis

Salivary gland chromosomes do not spread when squashed

Sciaridae Rhynchosciara angelae

Pavan et al. (1971), Diaz (1972)

Sciara coprophila

Gabrusewycz-Garcia (1975) Carson (1944)

S . impatiens S . ocelfaris

Contacts of telomeres with intercalary regions and these regions with each other

S. species Simuliidae (representatives of family) Tephritidae Batrocera okae (Dacus okae) Ceratitis capitam

COLLEMBOLA Bilobelfa aurantiaca

Mainx (1976)

Meti (1936), Metz and Lawrence (1938) Perondini and Dessen (1985) McCarthy (1945a,b)

Ectopic contacts infrequent

Rothfels (1956), Ottonen (1966)

The chromosomes of salivary glands, malpighian tubules, mid and hindgut, and fat body cells of larvae and pupae spread poorly because of numerous ectopic contacts; the chromosomes of trichogen cells of the orbital bristle of pupae spread readily Ectopic contacts between blocks of a-heterochromatin

INFUSORIA Stylonychia mytilus

Zambetaki et at., (1995) Ekdo (1986), Zacharopoulou (1987), Semeshin et al. (1995)

Deharveng and Lee ( 1984) Jareno (1990)

~

Note. Dash indicates that the species is only mentioned in reference(s)

Figure 87. Ectopic contacts of the micronucleoli and nucleoli with the regions of polytene chromosomes. (a) Filling of the break in the 42B1-3 region with nucleolar material. (b) Contact with the region of a complete break in 11A8-9. ( c )Long strand connecting the nucleolus with the 23AC region. (d) “Sticking”of the nucleolus on the 8 9 E 1 4 band. (a and b) Electron microscopy. (c and d) Light microscopy. n, nucleolus. (a and b) reprinted by permission from Zhimulev et al. (1982); (c and d) reprinted by permission from Ananiev et al. (1981).

178

I. F. Zhimulev

Table 12 Occurrence and Characterization of Ectopic Contacts of the Chromosomes and Nucleolar Material in Diptera Family, species Chironomidae Chironomus plumsus

Drosophilidae D. mehnogmter

Occurrence

References

In three larvae of one population, in all salivary gland cells the fourth chromosome was attached to the nucleolar forming zone of one of the arms of long chromosomes 3L, 2L, or 2R

Belyanina (1978a)

Nucleolar-like material attached to breaks of the 11A and 19E regions Presence of nucleolar-like material in the 11A region Contact of nucleolar-like material with band When versene (372 mg/100 g of food) was added to food there formed additional nucleoli frequently attached to regions of intercalary heterochromatin, especially in regions 2,50, 56, and 58; in the X chromosome, nucleoli were seen in regions 1A, 2B, 3C, 4B, 4C, 7A, 7C, 7D, 8B, 8C, 9A, 11A, 11B, 11C, 11E 12E, 13E, 13E 15E 16F, 17A, 19A, 19D, and 19F Nucleolus-like body containing the rRNA genes contacts with the 56F intercalary heterochromatin region

Bridges (1938)

Up to 80 regions contact with the nucleolar material (Figure 88); more than 50% of contacts occur in the intercalary heterochromatin 3C, 4CD, 7AB, 11A, 36AD, 39D, 56EE 70DE, 71C, 76C, 816 84AB, 84D, 87DE, and 89EF regions Contact of the nucleolus with the chromosomescan be achieved through submicroscopicstretches Contacts of micronuclei with the chromosomes

Treatment with EDTA does not produce formation of additional nucleoli that show some aftinity for heterochromatin The number of nucleoli in 733 examined nuclei ranged from 1 to 12; the occurrence frequencies of nuclei were 20.05% with 1 nucleolus, 21.289: with 2 nucleoli, and 21.83% with 3 nucleoli; most irequently they were attached to the chromosomes in the lAC, 2AC, 5D, IOAC, 19F, 20CE 21A5 22A, 25A, 26D, 27AE 30A, 31AF, 33AF, 40AF,

Sutton (1940a) Gay (1956) Khristolyubova (1961,1964), Khristolyubova et al. (1961,1962), Khristolyubovaand Auslender (1967)

Steffensen and Wimber (1972), Wimber, Steffensen (1973) Ananiev et al. (1978, 1981), Ananiev and Barsky (1985), Gvozdev (1978, 1981a,b) Ananiev and Barsky (1985) Hill and Watt (1978), Hochstrasser and Sedat (1987a) Semionov et al. (1978) Michailova et al. (1982)

(continues)

Polytene Chromosomes, Heterochrornatin, and Position Effect Variegation

179

Table 12 (Continued) Family, species

D. melanogaster X D. simulans

D. robusta Zaptionus

indianus Psychodidae Telmatoscopw sp. Sciaridae Rhynchosciara hollaenderi

Sciara coprophila

Tephritidae Ceratitis capitata

Occurrence 41AF, 43E 44A, 46B, 53C, 54A, 55A, 56E 57AE, 58A, 59DE, 60EE 61AC, 62AD, 63AF, 64A, 67AB, 80AE 81E 83AE, 85AE 87A, 89D, 94A, 95A, 96A, 98AE 99DE, lOODE regions; altogether, nearly 390 attachment points of the nucleoli to the chromosomes were localized Tight contact of the nucleolar material with breaks in the 42B1-3 and 11A8-9 regions Nucleoli most frequently attached to the telomeric regions of the chromosomes and regions of intercalary heterochromatin, and quite frequently to centromeric heterochromatin Additional nucleoli, including the 18sand 28s rRNA genes and two types of insertion (ivs-I and ivs-ll), were found attached to the chromosome regions containing the ivs-I insertion Contact of nucleolar-like material with the break site of chromosome 3L According to EM data, nucleolar-like material fills chromosome breaks in the 70A and 70C regions Ribosomal DNA of the nucleolus associated with regions of the polytene chromosomes by ectopic contacts The nucleolus attached to various sites of the chromosomes Contact of the microchromosome with the nucleolus

References

Zhimulev et al. (1982) Genova and Semionov, 1985a,b

Semionov and Kirov ( 1986)

Sorsa (1988a) Saura er af. (1989) Mecheva ( 1990)

Levitan (1970) Gupta and Kumar (1987)

Nucleolus-forming regions contact with heterochromatin

Amabis and Simoes (1972)

In micronucleoli adhered to the chromosomes, rDNA was identified by the method of in situ hybridization Nucleolar material as drops adhered to the chromosomes was detected by staining with ammoniacal silver Approximately 150 chromosome regions contact with micronucleoli; of these, 67 (45%) are regions of late-replicating DNA

Pardue et al. (1970)

Gabrusewycz-Garcia and Kleinfeld (1966)

Ectopic stretches between the nucleolus and “heterochromatin sphere”

Bedo and Webb (1989)

Stockeret al. (1978)

180

I. F. Zhimulev

Figure 88. Contact frequencies of the nucleolar material with the chromosome regions of Drosophila. Long horizontal lines with number subdivisionsrepresent the maps of chromosomesX, 2L, 2R, 3L, and 3R. Short horizontal lines over the map depict the location of nucleoli in the chromosome regions. Reprinted by permission from Ananiev et al. (1981).

4. In unsquashed preparations, considerably more ectopic associations are detected between the neighboring than the remote bands (see Figure 89; see also Kaufmann and Iddles, 1963). Figure 90 presents a matrix of contact involvement for 40 regions with the most prominent property of ectopic pairing. The highest frequency values are distributed along the diagonal of the matrix, and this is evidence for the predominant contacts between the nearest neighboring regions. These facts can be accounted for by the mechanisms of contact formation (accepting that they are randomly established, the nearest bands should be in contact because they are closely adjacent) and the technique of making squash preparations (when the nucleus is squashed, the associations between the remote regions are the most readily severed). Analysis of the matrix in Figure 90 demonstrates that all ectopic contacts are most frequently established at a distance of 1 map unit of Bridges’ map, and thus, for example, between the more or less remote regions. A striking correlation with the distribution of inversion ends was revealed (see Figure 98). Mathematical treatment of the matrix using a method of automatic classification enables us to distinguish groups of bands with the same relation to all the associations of the remaining regions. These groups of bands, in the view of Kulichkov and Zhimulev (1976), form contact nodes involved in the maintenance of the three-dimensional structure of the nucleus.

Polytene Chromosomes, Heterochromatin, and Position Effect Varienation

181

Figure 89. A map of intra- and interchromosomal ectopic associations of chromosome 2L of Drosophifu. The chromosome is depicted as a semicrescent within which contact regions are connected by lines; the region of contacts with other chromosomes are shown outside. After Kaufmann and lddles (1963).

182

met

1. F. Zhimulev

~~

CC

in

Figure 90. A matrix of ectopic contacts of 49 regions of polytene chromosomes of Drosophika. The designations of the regions are given along the vertical and horizontal lines; the occurrence frequencies between the respective regions are given in squares. After Kulichkov and Zhimulev (1976) and Zhimulev et al. (1982).

Tissue specificity was detected for the manifestation of ectopic contacts. For example, in Calliphoridae or Muscidae, squash preparations from the cells of many larval organs cannot be made, while the chromosomes of trichogen cells are easily identified in the chromosomes of ovarian nurse cells (see Table 11, earlier). Kaufmann and Iddles (1963) have posed the question of whether the IH regions can be involved in puff activation. Having compared the location of 71 puffs and the regions of ectopic pairing in D. mekmogaster, they found that the puffs are close to the IH regions: only 12 of the 71 puffs are at a distance of more than 10 bands; the others are either in close proximity or at a distance of not more than 5 bands. It is the authors' view that these correla-

Polytene Chromosomes, Heterochromatin, and Position Effect Varieuation

183

tions can underlie tests of the suggested important role of the IH in puff activation (Kaufmann and Iddles, 1963; Kaufmann, 1965). The estimates obtained for D. ananussue are close: of the 70 known puffs, 47 (67.1%) were localized in the regions of ectopic pairing (Duttagupta et al., 1973). In contrast, Hartmann-Goldstein (1966) revealed a negative correlation between the distribution of puffs and the IH along chromosome length. Finally, having compared the proportions of the chromosomes that are occupied by euchromatin and the IH, and puff distribution, Ashburner (1966) came to the conclusion that the puffs are randomly distributed. Because there are considerably more puffed and intercalary heterochromatin regions in D. mehnogaster than the previously mentioned authors believed-there are approximately 60-70 large and small puffs in each arm (Ashburner, 1972a; Belyaeva et al., 1974; Zhimulev, 1974) and 40-50 bands of ectopic pairing (Kaufmann and Iddles, 1963; Zhimulev and Kulichkov, 1977)-in the case of uniform distribution, the large and small puffs would lie randomly and in the neighborhood, too. Nevertheless, certain facts prove that ectopic pairing does affect at least puff morphology. The 2B3-5 (Bridges, 1938; Belyaeva et al., 1980, 1987) and 85F1-6 (Baricheva et al., 1987) puffs in D. melanoguster and Balbiani rings l and 2 (Scouras, 1984) in D. auraria are bounded at both sides by bands engaged in ectopic contacts within the chromosome body. As a result, the active material of the puff intermingles with the neighboring bands, and this confers a granular appearance to the puffs (Figure 91). It cannot be excluded that the fuzzy, “heterochromatic puffs” in Ohgmia ornata and 0.nolkri are of the same nature (Shcherbakov, 1968). Perhaps the toroid-shaped bands on the chromosome tips (Scouras et al., 1992) results from ectopic contacts between neighboring bands. The mechanism of the formation of ectopic contacts appears to be the most debatable issue in the whole problem of intercalary heterochromatin. In his early studies, Bridges suggested that ectopic contacts may be established within the so-called cytological repeats, that is, in the chromosome regions showing the same banding pattern (they have been considered in greater detail in Zhimulev, 1992b, 1996). It follows from his suggestion that homologous regions pair (Bridges, 1935c, 1936). Prokofyeva-Belgovskaya (1937b, 1941) assumed that IH is a specific region of the chromosome containing many homologous loci “multiplications”, that is, she long ago predicted the existence of repeats in the genome and viewed ectopic pairing as a contact of similar with similar. Later, the idea of DNA homology in regions of ectopic contacts gained many followers (Barr and Ellison, 1971b, 1972; Mayfield and Ellison, 1975; Zhimulev et al., 1982; Ananiev and Barsky, 1985).

184

1. F. Zhlmulev

Figure 91. Morphology of puffs developing between bands establishingectopic contacts with one another. (a) Map of distal part of the X chromosome of D. melanogaster, including the 2B3-5 puff. (b) Scheme of the development of Balbiani rings 1 and 2 (BR, and BR,) in D. nuraria. (a) after Bridges (1938); (b) after Scouras (1984) and Scouras and Kastritsis (1984).

In the 1930s and 1940s’ectopic pairing of the telomeric regions (see Section VIII) was suggested to be due to “nonspecificaffinity”of the terminal chromomeres” or “nonhomologous attraction of heterochromatic chromomeres” (Bauer, 1936a). It is Fujii’s opinion (1942) that ectopic contacts of telomeres are nothing more than secondary associationsof the chromosomes produced by their squashing.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

185

Finally, it has been proposed that a homologous “nucleoprotein” may lie in the regions of ectopic pairing. Its presence implies either (1) a comparable relative proportion of nucleic acids and protein, (2) a similar composition of proteins, or (3) nucleic acids as constituents of the chromosomes (Kaufmann and Iddles, 1963). What data argue in favor of nucleoprotein homology in the regions of ectopic pairing? The presence of the same DNA sequences identified by, for example, in sinc hybridization, or proteins of particular type detected by specific dyes or reaction with antibodies support nucleoprotein homology. Furthermore, information concerning homology can be provided by differential staining of the chromosomes. From in situ hybridization studies, at least two labeling types can be expected: (1)both regions of ectopic interactions and the ectopic fiber between them label, or (2) only one of the IH regions and a DNP fiber extending from this region to another region label (Figure 92). In the former case, the results provide evidence for the presence of homologous DNA of all three components of ectopic contact. Labeling of the second type shows that DNA that is homologous to the DNA of the hybridization probe is present in only one of the pairing chromosome regions. The data in Table 13 indicate that ectopic contacts between chromosome regions with identical features are widespread. Leibovitch ( 1990b) detected a correlation between decrease in the occurrence frequencies of mobile ele-

Figure 92. Labeling of the 84D region and a n ectopic stretch to the 86D region after in situ hybridization of pDm2. The scale is 10 pm. After D. Finnegan and M. L. Pardue; reprinted by permission from Pardue er al. (1977).

186

I. F. Zhlmulev

Table 13 Presence or Absence of DNA Homology in Chromosome Regions Involved in Ectopic Contacts Species

Information on homology in ectopic contacts

References

I. HOMOLOGY Anntopynia dyari

Between Q' centromeric bands of chromosome 3L and of the fourth chromosome

Bedo (1974)

Chironomus thummi

Between Q' centromeric and intercalary bands

Badaev et al. (1973)

Drosophila biseriara At least some regions of the chromosomes included in the pseudo-chromocenter are rich in highly repetitive DNA and are Q+-stained

Yoon (1989)

D. funebris

Two regions of localization of 5 s rRNA genes and ectopic stretch between them are labeled by 5s rDNA after in situ hybridization

Cohen (1976a)

D. melanogaster

In some Oregon-R individuals there is an additional Q+ band in the 83E region; the frequency of ectopic contacts between 81F Q+- and 83E Q+-bands is 36.996, while that between 81F Q+- and 83E Q--bands is only 1.1% Pairs of regions (67BC and 64E 62DE and 93E, 32A and 56F) and stretches between them labeled after in situ hybridization with DNA of h 2 2 5 mobile element The 84A and 19E regions and stretches between them labeled by hybridization with DNA of DmJ 18 mobile element; the same occurred in two unidentified regions after hybridization with DNA of DmlJ 10 cases of labeled stretches connecting labeled bands (DNA horn IOAI-2 band as probe)

Barr and Ellison (1971a, 1972), Mayfield and Ellison (1975) Ananiev et al. (1978). Gvozdev (1981a,b)

Tchurikov et al. (1981)

Kokoza and Zhimulev (1994)

Glyptotendipes barbipes

Two regions of the nucleolar organizers and stretches between them labeled by hybridization with [3H]18S+ 28s rRNA

Wen et al. (1974)

Orthocladius bipunctellus

Between C+-bands at the ends of the chromosomes

Michailova and Belcheva (1982)

Phryne cincta

Contacts between blocks of intercalary a-heterochromatin

Wolf and Sokoloff (1973)

Rhynchosciara angelne

Contacts between stained bands detected after DNA denaturation-renaturtion and staining with methyl green and pyronine

Diaz (1972)

(continues)

187

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 13 (Continued) Family, species

Occurrence

References

11. NO HOMOLOGY Amtopynia dyari

Contacts between Q+-and Q--material in dispersed chromocenter

Bedo (1974)

Chironomus thummi

Between Q+- and Q--intercalary regions

Badaev et d.(1973)

Drosophila melanogasrer

In the case of contact of the 39DE (locus of the histone genes) and the 36D" regions, only 39DE and the stretch of ectopic contact are labeled after in situ hybridization (the histone gene DNA as a probe); labeling of the 84D band and ectopic stretch in contact of the 86Dband 84D regions (pDm2 as a probe) (see Figure 92) Labeled ectopic contact between the labeled 93E and unlabeled 92DE regions (DNA of Dm225 mobile element as a probe) Labeling of histone gene locus and ectopic stretch, but not of the second locus involved in contactthree cases (the histone gene DNA as a probe) Same as in Lifschytz (1983) study for both the 12E and the 40AF regions, ectopically pairing with the 11A6-9 region (DNA of the mobile element 1360 as a probe) Labeling of the 89E1-4 band and ectopic stretch connecting it with an unidentified region (a fragment of the Ubx gene as a probe); the same occurred for the 11A6-9 band (DNA from the 11A region as a probe) Contacts of the 5s rRNA genes with the nucleolus

Pardue et al. (1977)

Labeling of the 10A1-2 band and a stretch of ectopic contact connecting it with the unlabeled regions (KJ72 clone DNA from the 10Al-2 band as a probe) (8 cases) Labeling the region 12E and ectopic thread to unlabeled 11D or chromocenter

D. neohydei

After hybridization of ['HIRNA with the heavy satellite of D. neohydei, the label is present over one of the regions with an ectopic stretch

Ananiev et al. (1978)

Lifschytz ( 1983)

Kholodilov et al. (1988)

Lamb and Laird (1987)

Steffensen and Wimber (1972), Wimber and Steffensen (1973) Kokoza and Zhimulev (1994)

Palumbo et al. (1994b) Hennig et al. (1970)

(continues)

188

1. F. Zhimulev

Table 13 (Continued) Family, species

D.sirnufans

Rhynchosciara angelae Sarcophagidae (representatives of the family)

Sciara coprophila

Occurrence

References

Labeling of the 10A1-2 band and ectopic stretch connecting it with the unlabeled 7B region (DNA from the 10A1-2 band of D. melanogaster as a probe) Contacts between unstained and stained bands detected after DNA denaturation-renaturation and staining with methyl green and pyronine

Kokoza and Zhimulev (1994)

Q+-DNP granules contact with numerous Q--regions of the chromosomesand the nuclear membrane of pulvillar cells; no preferential contacts of granules precisely with intercalary heterochromatic regions was established

See section VI, A-C

Contacts of nucleoli with late-replicatingchromosome regions

Gabrusewycz-Garcia and Kleinfeld (1966)

Diaz (1972)

aThe authors called it the “region more proximal than 37A.” bThe authors designated it as 86C.

ments and rare ectopic pairing in the 14-17 region of the X chromosome of

Drosophika. However, there are many more cases of contact with qualitatively different DNA regions. In addition to those listed inTable 13, there are exemplary contacts between intercalary heterochromatin and nucleolar material (see Table 12, earlier), between the IH and the telomeres. It is known that the 18s and 28s RNA genes, included in the nucleolus, map to (with rare exceptions) the nucleolar organizer, like telomeric DNA, which maps to the telomeres (and centromeric heterochromatin). Consequently, there is no particular reason to suppose that there may be DNA homologies in the region of ectopic contacts of the IH with telomeres and nucleoli. The facts adduced in Table 13, I1 are difficult to explain from the standpoint of DNA homology. The following considerations show that DNA homology is not determinant in establishment of ectopic pairing. Numerous insertion sites of the mobile elements of many families are scattered among the chromosomes of eukaryotes. If merely physical DNA homology were sufficient for the establishment of ectopic contacts, then, first, they would preferentially arise in sites where mobile elements are most frequently located and, second, their location would change after the mobile elements have transposed to another position. Neither of these events is observed, however.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

189

From the previous considerations, it is clear that nucleoprotein homology at the DNA level is not the condition necessary to provide ectopic contact. The presence of weak points (or DNA underreplication) proved to be another condition necessary for their establishment. The lines of evidence are as follows:

1. The highest values for the frequencies of breaks and ectopic contacts in the chromosomes of females were detected in the same regions.

2 . Sharp reduction in the manifestation of breaks in the male X chromosomes have been noted (Kastritsis et al., 1970; Zhimulev et al., 1982, 1989a,b), and, accordingly, a reduction in the frequencies of ectopic pairing (Figure 93).

3. The karyology of the dipteran order has been quite amply studied. However, the Simuliidae and Chironomidae families stand aloof. Polytene chromosomes have been described in hundreds of species of these families, but, by way of contrast, ectopic pairing and breaks are rare findings to the present time. The data on differential staining (see Sections VII,A and VI1,B) and late replication (see section VII,C,3) provide evidence for the presence of IH in the chromosomes, but breaks and ectopic pairing are not manifested in a linked manner in these species. Several models of ectopic pairing have been put forward (Figure 94), each taking into account the various organizational features of the IH. According to the first model, underreplication (breaks) occurs in the cluster of repeated se-

3c

llA6-9

12E8-9

1W1-4

Figure 93. Frequencies of breaks (white columns) and ectopic contacts (black columns) in certain regions of the male and female X chromosomes ofD. melanogaster. The histogram was built on the basis of data published in Zhimulev et al. (1982). Abscissa, designation of region, sex of larva; ordinate, occurrence frequencies (%).

190

1.

F. Zhimulev

a

u

Y

Y

A I

l

& y

Y...Y

Figure 94. Hypotheses of the formation of breaks and ectopic associations in polytene chromosomes. (b) Letters A, C, and D designate the various repeat clusters. (c) Arrows indicate the 5' + 3' direction of the DXA strand. (a) reprinted by permission from Zhimulev et al. (1982); (b) reprinted by permission from Kholodilov et al. (1988); (c) after Ashburner (1980); (d) after Laird et al. (198713).

quences composing the IH bands; "sticky ends" arise and all types of ectopic contacts result (Figure 94a). Thus the model incorporates the correlation between break location and ectopic pairing (see also Scouras and Kastritsis, 1988). A n attempt is made to reconcile the hypothesis of DNA homology con-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

A

A

A

C

C

191

C

C

A Figure 94. Continued

tact sites with the data presented in Tables 12 and 13. If it is true that different clustered sequences (A, C, D) are present in each band, then it seems logical that an ectopic fiber would result from contacts of homologous DNA in each chromosome region involved in contact (cluster “ A in Figure 94b, for example). However, in in situ hybridization of the DNA from clusters “ D or “C,”only the ectopic fiber and one of the chromosome regions label (see Table 13). The third and the fourth models are based on the mechanism of DNA replication the impairment of which produces branch migration. The migrated fragments of the newly replicating DNA of one of the molecules join with the

192

I. F. Zhimulev

Figure94. Continued

counterpart fragments of the other molecule presumably because of the restricted homology of the base pairs, thereby entering into an ectopic association (Figure 94c). When one considers that a branch can migrate only at the end of the cycle in the replication eye, the correlation between the location of late replication in the chromosomes and ectopic pairing becomes reasonable. Moreover, because of local DNA underreplication in the region of migration, there arises a chance for breaks to occur (Ashburner, 1980). Accepting this model, it becomes unclear why ectopic pairing takes place in strictly defined regions of the chromosomes, while the dispersed short repeats, which can provide homologous contacts between DNA fragments, are more or less uniformly distributed throughout the genome. According to Spradling et al. (1992), intertelomeric ectopic fibers are formed at the expense of cleavage of tandemly repeated subtelomeric DNA and its occasional religation between the two chromosomes. The last model (Figure 94d) postulates that, when a replication branch migrates, the unpaired fibers are held together by proteins analogous to the V protein of the F1 phage, which binds together any single-stranded DNA regardless of base specificity. In terms of this model, an ectopic contact can arise in virtually in any chromosome region. The hypothesis that ectopic contacts result from protein homology is consistent with established facts. Accepting that certain fragments of the genome are deeply inactivated and that this state is effected through particular proteins compacting chromatin (see Section XVIII), ectopic contact between these re-

Poiytene Chromosomes, Hetarochromatin, and Position Effect Variegation

193

gions differing in base composition and DNA, albeit having homologous proteins, becomes explicable.

3. Late replication After incorporation of [3H]thymidine, a labeled precursor of DNA synthesis, into the salivary gland cells, different types of labeling are detectable: ( 1) chromosomes free of label, presumably those beyond the S phase; ( 2 ) a general labeling, or continuously labeled chromosomes; and ( 3 ) chromosomes discretely spotted with label. The number of spots varies from one or two to several tens (Key1 and Pelling, 1963; Plaut, 1963,1968,1969; Plaut etal., 1966;Howard and Plaut, 1968;Simoes, 1970). Is replication continuous at first, then discrete, or vice versa? The results of intensive investigation of replication in the chromosomes of various dipteran species allow us to envisage the process as follows. Puffs, interbands, and very thin bands are the first to enter the S phase. The number of labeled regions keeps growing to the stage of complete labeling. The large bands and the centromeric heterochromatin are last to enter the S phase. The difference in time between the beginning of the S phase and entry of the last regions into the phase of DNA synthesis is not great for the large euchromatic bands, but it is quite considerable for the heterochromatic regions. Puffs, interbands, and thin bands are the first to fully replicate and, as a result, the chromosome becomes discretely labeled. By the end of the S phase, oniy some regions (Figure 95), the

Figure 95. Incorporation of [)H]thymidine into chromosomes 3L and 3R at different stages of

termination of the S phase. After Zhimulev and Kulichkov (1977) and Zhimulev et al. (1982).

194

1. F. Zhimulev

chromocenter in Drosophih and centromeric heterochromatin in Chironomus, still label (Pavan, 1959, 1965;Keyl and Pelling, 1963; Plaut, 1963;Gabrusewycz-Garcia, 1964; Plaut et al., 1966; Tulchin et al., 1967; Arcos-Teran and Beermann, 1968;Nash and Bell, 1968; Rodman, 1968; Hagele, 1970; Lakhotia, 1970; Chatterjee and Mukherjee, 1971, 1973, 1975; Arcos-Teran, 1972; Tiepolo and Laudani, 1972; Mukherjee and Mitra, 1973; Hagele and Kalisch, 1974, 1980; Gubenko, 1976a; Kalisch and Hagele, 1976; Meer, 1976; Ilyinskaya and Martynova, 1978; Mukherjee et al., 1980). The timetable for the complete replication of a single region of the chromosome is mainly invariable (Keyl and Pelling, 1963; Plaut et d., 1966; ArcosTeran and Beermann, 1968; Rudkin, 1972, 1973). DNA replication is also temporally ordered in the salivary gland chromosomes of closely related species (Gubenko, 1976~). The behavior of the X chromosome is unusual in the males of Drosophila: the X starts DNA replication somewhat earlier (Hagele, 1973) and finishes it much earlier than the autosomes. This has been demonstrated for D. hydei (Berendes, 1966; Mulder et al., 1968), D. melanogaster (Rodman, 1968; Lakhotia and Mukherjee, 1969, 1970; Hagele and Kalisch, 1974), D. anuwsae (Duttagupta et d.,1973), D. virilis (Gubenko, 1976b), and hybrids from D. azteca X D. athabasca matings (Meer, 1976). Thus, in salivary glands of D. hydei, in 55% of nuclei whose autosomes are at the stage of continuous labeling, the male X chromosome already shows discrete labeling, never observed in the female X (Berendes, 1966). In nuclei exhibiting discrete labeling, the number of replicated regions of the male X chromosome is consistently smaller than in the female X chromosome (Berendes, 1966; Mukherjee et al., 1980), it is, for example, twofold smaller in D. virilis (Gubenko, 197613).The same regions are referred to as late replicating in males and females (Berendes, 1966). From the data being considered here, it is obvious that the degree to which chromosome regions lag behind in completion of replication can be tentatively determined from the labeling frequencies of a single chromosome region. In early experimentswith small segments of the X chromosome and chromosomes 3L and 2R, and with the entire X chromosome of Drosophih, it was shown that the regions referred to as intercalary heterochromatin by other criteria labeled most frequently (Plaut et al., 1966; Arcos-Teran and Beermann, 1968; Arcos-Teran, 1972). Comprehensivedata on the late-replicatingregions in Drosophila and on their relation to the IH have been obtained by Zhimulev and Kulichkov (1977). Chromosomes at the stage of continuous labeling, with the number of discretely labeling regions ranging from 1-2 to 30, were graphically represented (see Figure 95). The number of labeling regions and their localizations were determined. The chromosome regions were plotted along the abscissa. Columns of arbitrary units corresponding to single chromosome, where they occurred most frequently, were

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

195

plotted along the ordinate. The labeled sites of another chromosome with a smaller number of labeled sites were plotted, one by one, from bottom to top. Ultimately, the location of the replicating regions of the chromosome with the smallest number of such regions was plotted at the top of the histogram. The tallest columns (see Figure 82c) represented the greatest delay in replication completion in a site. Generally, the number and location of the late-replicating regions and regions of ectopic pairing in the chromosome of Drosophila are closely correlated. Both regions are much more numerous than those of breaks, which are located in the late-replicating regions only. Moreover, it should be emphasized that silver grains during late labeling are located precisely over the band, which is subjected to breaks regardless of where they occur-in the distal or proximal edges or in the middle of the band. Not all late+replicating regions were detected in breaks: they were not identified at all in the 41AD, 56F, and 80AC regions and others, and in exceptional cases in the 9A1-4 and 63El-5 regions. The almost complete absence (except for chromosome 2R) of late replication in telomeric regions was an unexpected finding. In D. nasuta, the nucleolar DNA included in the nucleolus is late replicating (Lakhotia and Roy, 1979). In D. oririlis, the late-replicating regions are presumably rich in repetitive sequences. Third instar larval salivary glands 98% of whose chromosomes are mainly at the end of the S phase were incubated with ['Hlthymidine for 15 min. As a result of analysis of renaturation kinetics, it was demonstrated that the DNA replicating during that time reassociated more rapidly than the major DNA peak, (i.e., it contains more repeated nucleotide seqeunces) (Levin et ul., 1977; Evgen'ev et ul., 1977). The graphic representation of results (see Figure 82c) brings out the temporal discoordination uncoupling of replication completion in a region of different chromosomes. As indicated earlier, replication is completed by all the chromosome regions in a strictly temporal order; however, exceptions are possible for virtually every region. For example, 3C (eight chromosomes) is the last to complete replicating in 1A-3C of the X chromosome of D. melunogustm; however, 1AB is the last replicating in a single chromosome (Plaut et al., 1966). The 11A region is the last to fully replicate in the entire X chromosome, although single nuclei in which the late-replicating region was 3C were detected (Arcos-Teran and Beermann, 1968; Arcos-Teran, 1972). In the asynaptic region of the third chromosome of Drosophila (Hagele and Kalisch, 1978) or Simulium ornatipes (Bedo, 1982a),distinct regions occasionally replicate asymmetrically (i.e., only in one of the homologs). The discontinuities in the columns in Figure 82c and figures in many previously published papers indicate that the completion of replication is temporally uncoupled in many regions. To illustrate, replication is completed by some 2R chromosomes in the 42B region by the stage of 25 replication spots, while others

196

1. F. Zhlrnulev

in this region still label at the stage of 9 replication spots. This may indicate either different replicating rates in chromosome regions or the containing of replicating material, From Figure 1 it is immediately apparent that the event is widespread, and not only in D. mehogaster (see also Mishra and Lakhotia, 1982b). A similar pattern was observed for the chromosome of the larval salivary glands of D. nasutu (Lakhotia and Tiwari, 1984), Simulium ornatipes (Bedo, 1982a), and the chromosomes of the ovarian nurse cells of the mosquito Anopheles stephensi (Redfern, 1981b). The data on tissue differences in the rate of replication completion are controversial. In A. stephensi, no substantial differences in the labeling pattern of the chromosome regions at the end of the S phase were found between larval salivary gland and ovarian nurse cells (Redfem, 1981~).In the ota3 mutant of D. mehogaster, late-replicating sites in the 21DE, 25A, 25EE 32F, 33AC, 34EF, 35CE, 36CD, and 39-40 regions were identified in chromosome 2L of ovarian pseudo-nurse cells (Sinha et al., 1987)-that is, in the same regions as in the salivary gland chromosomes (Zhimulev et al., 1982). Comparisons of the same feature in fragments of the X and second chromosomes of the salivary glands and gastric cecum in D. hydei disclosed differences in the chronology of replication completion in 8 of the 62 studied regions of the X chromosome and in 11 of the 55 studied regions of the second chromosome (Tiwari and Lakhotia, 1984). A similar comparison of the fragments of the second and third chromosomes of D. nasutu revealed great differences in the temporal order of replication completion in chromosome 2R (5 regions of 22) and in 5 of the 38 regions of the third chromosome (Figure 96). The data on differences in the amount of late-replicating material between closely related species are extensive. The differences are very conspicuous in hybrids when one of the homologs has late-replicating material and the homolog received from the other species does not-for example, in Anopheles atroparvus X A. labranhia (liepolo et al., 1974), Drosophila virilis X D. littoralis (Evgen’ev and Gubenko, 1977), and Chironomus thummi thummi X Ch. th. piger (Keyl, 1965a,b, 1966; Hagele, 1970, 1976). In the latter case, variation in latereplicating material and in DNA amount is coincident. Considerable prolongation of larval life in two strains (gtand Oregon-R) of D. mehnogaster was achieved by transfer to low temperature (10°C). This implies that the S phase may be much prolonged, because of the asynchronous entry and completion of replication in distinct sites of the chromosomes. However, no differences were found in the location of sites and the degree to which replication completion was delayed between larvae grown at 10°C and those grown at 24°C (Mishra and Lakhotia, 1982a,b). What may be the causes of late completion of DNA replication by the regions of intercalary heterochromatin? It may be suggested that replication finishes late because large bands contain many replicons and, as a consequence of

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

197

Figure 96. A comparison of labeling frequencies of certain regions of chromosomes 2R (a) and 3 (b) in the nuclei of salivary gland and gastric cecum of Drosophila nasuta. Abscissa, chromosome regions; ordinate, labeling frequency of regions. Reprinted by permission from Lakhotia and Tiwari (1984).

their asynchronous activation, some enter the synthesis phase very late. However, the degree to which replication completion was delayed did not correlate with band size (Ananiev and Barsky, 1975). However, it was shown that heterochromatic regions enter the S phase much later than interbands and puffs; for example, replication does not begin in large blocks of centromeric heterochromatin in Ch. melanow even at the stage of continuous synthesis (Hagele, 1970). Lakhotia and Sinha (1983) have found two types of replicons in the polytene chromosomes of the salivary glands of D. mum: (1) long (64 p m on average), with high migration rate of replication fork (approximately 0.95 pmlmin); and (2) short (approximately 20 pm) and replicating considerably more slowly (0.07 p.m/min). Replicons of the first type are active at the beginning and those of the second type at the end of the S phase. For this reason, it may be suggested that the latter are located in the late-replicating region. Before making one more assumption, it is expedient to recall certain

198

1. F. Zhlrnulev

facts: (1) the earlier completion of replication by the male X chromosome, which is more loosely packaged than the autosomes or the two female X chromosomes; and (2) a corollary of the replication scheme, namely the decompacted chromosome regions, puffs, and interbands replicate earliest, and become compacted only later. Therefore, it is conceivable that the compaction degree of chromatin affects replication intensity, presumably because molecular processes are hindered in the most compact chromosome regions.

4. Chromosomal rearrangements In early studies on induction of chromosomal rearrangements in Drosophih, it was found that the distribution of breakpoints along the length of polytene chromosome is not specific (Kaufmann and Demerec, 1937). Later analysis of 1038 radiation-induced rearrangement breakpoints demonstrated that they are distributed uniformly among the chromosomes but disproportionately among the proximal hetetochromatic regions (Bauer et al., 1938; Bauer, 1939). The rearrangements were mapped only to the precision of numbered map units, which leveled out the differences between the smaller (lettered) fragments of the chromosomes. From a survey of the data in Bauer’s paper, it follows that the rearrangements occurred more often in the subdivisions at which IH is mapped: the 11, 12,35,42,56,59, 62,64,67, 75,86,94,96, and 98 regions. Using mapping of higher resolution (to the letter subdivision), nonrandom formation of rearrangements in the X chromosome was demonstrated in two other studies. In a sample of 141 breakpoints, 37 fall in regions 11 and 12, where the “inert” 11A, IlCD, and 12EF are located. In another four regions (2B, 4C, 7C, and 19E), the rearrangements occur more frequently than might have been expected if they had a random distribution (Prokofyeva-Belgovskayaand Khvostova, 1939). Mapping of the breakpoints of 627 x-irradiation-induced chromosomal rearrangements revealed that 28% are located in the 20AF region of proximal heterochromatin. Break frequencies were also exceptionally high in the 1lA, 12D, 12E, and 19E regions. It was inferred that heterochromatic material may be present in these intercalary regions (Kaufmann, 1939, 1941a,b). Preferential location of breakpoints was found for other chromosomes, too. Dubinin et al. (1941) detected disproportionately high break frequencies in the 21D, 22A, 22B, 22D, 245 57F, and 59D regions of the second chromosome and in the 61C, 61D, 615 62B, 64B, 89DE, 93EF, 99AB, and 99F regions of the third chromosome. Somewhat later, Kaufmann (1944, 1946) reported that, in an analysis of approximately 1400 breakpoints, 1048 were localized within the euchromatic portion of the X chromosome. Peaks of the highest frequencies were again observed mainly in the IH regions: 12E, 11A, 12D, 1F, 7B, 19E, 9A, 16F, 4E, and 3C.

Table 14 List of Chromosomal RearrangementsTaken into Account in Building of Histograms of Distribution Frequencies of Breaks in Polytene X Chromosome of Drosophila melanogastef

Rearrangements

I. Spontaneously

Deletions

Duplications

28

6

Various rearrangements, including those with Translocations and imprecise localization Breaks produced in the first generation Inversions transpositions of one of breaks

34

1

22 52

8

17

References Lindsley and Grell(l968)

arisen

1

43 2

8

13 23 6 6 6 20 4 13

15

3 30

2 10 2 58 1

Oshima et al. (1964) Yamaguchi et al. (1976) Stalker (1976) Grossman (1967) Pipkin et al. (1976)

Rim et d. (1986)

2

Dubinin and Sokolov (1940) Zacharopoulou (1974b) dos Santos et al. (1991) Koryakov et al. (1994) Yang et al. ( 197I ) lnoue (1988) Zacharopoulou (1974a) Zacharopoulou and Pelecanos (1980) Paik et al. (1969) Roca et al. (1982) Mukai et al. (1970) Paik (1979) Koliantz (1971) (continues)

C

.d

-4-

I

s

m

W

m 0

-

m

N

*

c1

m

m

1

1

48

1

7

17 1

43

5 4

1

29

13

14 11 38 18

35

10

12 167

20

50 77 5

69

Mukhina and Zhimulev ( 1980) Mamon et al. (1977) Lefevre (1974) Hilliker (1985) Ashbumer (1972b) Reuter et al. (1985) Omelianchuk et af. (1991) Volkova and Buzykanova

(1991) Denell et al. (1978) Tiusis and Hilliker (1984), Hilliker and Trusis-Coulter (1987),Hilliker et d.

3

(1991)

1

Roberts (1972a) Yamamoto (1987) Woodruff and Ashbumer

6

(1978) 170

Total (111) Total number of rearrangements

59 92

12

107 626

1091 1176

(1-111) aAfter Mukhina et al. (1981) and Koryakov and Zhimulev (1996).

130 176

1400 77 1639 1639

Prokofieva-Belgovskaya and Khvostova (1939) Kaufmann (1946) Demakova et af. (1994)

202

1. F. Zhimulev

During the ensuing years drosophilists have described a great variety of new chromosomal rearrangements,and the pertinent data are summarized in Table 14. In tabulating the data, those studies in the extensive literature were included in which rearrangements in populations were described without determining their causes (i.e., rearrangements tentatively called spontaneous), or were radiation-induced or transposable elements, without any special selection. Data on the location of 1639chromosome breaks and 2082 rearrangements were gleaned in this way. Figure 97 presents information regarding the location of rearrangement breakpoints in Drosophila. It is quite obvious that the major frequency peaks fall in the IH regions. This is most conspicuous in the histogram for the X chromosome, for which the most complete samples were obtained. Although scantier, the data for the other chromosomes confirm this conclusion. Nonrandomness of rearrangements along chromosome length has been demonstrated for many dipteran species (Helfer, 1941; Koller and Ahmed, 1942; Nicoletti and Lindsley, 1960; Kunze-Muhl, 1961; Rabbani et al., 1977; Sharma et al., 1978; Pleshkova, 1983; Naveira and Fontdevila, 1985; Gunderina and Aimanova, 1997). Chromosomal rearrangements, mainly inversions, are widespread in dipteran populations (for review, see DaCunha, 1955; Sperlich and Pfriem, 1986). They are frequently referred to as spontaneously arisen, although transposition of mobile elements often underlies their emergence. Comprehensive studies of nonrandom location of the ends of chromosomal rearrangements identified in populations have been performed (Dobzhansky and Sturtevant, 1938; Dobzhansky and Epling, 1944; Novitsky, 1946; Acton, 1955; Kunze-Muhl and Sperlich, 1955; Rothfels and Fairlie, 1957; Gupta and Bihari, 1987; Ilyinskaya et al., 1988). Spontaneous rearrangements more frequently arise in regions of intercalary heterochromatin (see Figure 97). In 43 of the 83 rearrangements induced by the 31.1.MRF factor, breaks were localized in the intercalary heterochromatin regions, as determined by the presence of late replication and ectopic pairing (Yannopoulos and Zacharopoulou, 1980). Some features of the formation mechanism of chromosomal rearrangements presumably underlie their nonrandom distribution. Two conditions must be met: (1) occurrence of breaks in distinct regions of the same or different chromosomes, and (2) spatial approximation of the broken ends. It is no wonder that these conditions would be most likely met if the chromosomes were not apart from each other, but instead in contact in certain regions. Having presented data on the formation of complex chromosomal rearrangements, Dubinin and Khvostova (1935) concluded that there are contact nodes in nuclei. When nuclei are irradiated, breaks arise, the chromosome fragments are reunited, and, as a result, rearrangements are formed. Prokofyeva-Belgovskaya ( 1939a, 1941) has suggested that “chromosomal rearrangements may result from pairing of inert regions located in the same chromosome (inversions) or in nonhomologous chromosomes ( translocations).”

B 30 T

20 16 10

6 0

D

1

3

6

7

B

11

13

3

6

7

B

11

13

I6

17

E 100

Bo 80 70 80

-

.. ..

.. .'

so ..

40 .. 30 20 10 0 1

16

17

Figure 97. Distribution of the ends of chromosomal rearrangements in the X chromosome of D. mefunogasrer according to the data of Table 14. (A) Spontaneous inversions. (B) Induced inversions. (C) Translocations and transpositions. (D) Sum of the data of Kaufinann (1946) and Prokofyeva-Belgovskaya and Khvostova (1939) indicated in the Table 14. (E) Sum of A-D plus deficiencies (data from Koryakov and Zhimulev, 1996). Abscissa, chromosome regions according to Bridges' revised maps; ordinate, number of breaks in the letter subdivision of the map.

204

1. F. Zhimulev

The idea that chromosomal rearrangements most frequently arise in chromosome regions held together by ectopic pairing has long appeared attractive. To substantiate this idea, Kulichkov and Zhimulev (1976) suggested a version of mathematical treatment of data on ectopic pairing allowing them to identify contact nodes and the included chromosome regions. However, using a similar treatment of chromosomal rearrangements, similar nodes were not identified (V. A. Kulichkov, 1978, personal communication). Nevertheless, Yoon and Richardson (1978a) and Yoon (1989) have detected a consistently occurring node of ectopic contacts in a number of Drosophita species. They called it the pseudo-chromocenter. They also found several chromosomal rearrangements between chromosome regions comprised by the node. Having analyzed the available data on ectopic contacts (Kulichkov and Zhimulev, 1976) and on inversion formation (Hilliker, 1985), Vagapova (1991) found that both are most frequently formed between regions that are at a distance of approximately two number sections on the map of polytene chromosomes, for example, between the 22A and 24A regions (Figure 98). These correlations suggest the existence of a chromosomal helix in which regions two sections apart in the linear chromosome are approximated and thus retained owing to ectopic pairing. Exchanges can occur between them, and inversions form. There are drawbacks in the model proposed for the relation of ectopic nodes to the formation of chromosomal rearrangements. First, a correlation between the formation frequencies of ectopic pairing and breaks was demonstrated. The latter were most frequently detected at a high degree of polyteny. For this reason, it is unknown how frequently ectopic contacts establish in a diploid (haploid) nucleus in which chromosomal rearrangements are formed. Second, it follows that the degree to which ectopic pairing is manifested is tissue specific. The manifestation is particularly reduced in ovarian nurse cells, those closest to the germline cells. There is disagreement about the published information regarding the hypothetical mechanisms keeping DNA fragments in spatial proximity during break formation. Thus, according to Soyfer and Akifjev (1977), a temporary duplex polynucleotide structure may be formed at the G, and G, phases of the cell cycle, when DNA is corrected. The size of the duplex is restricted by the complementary (repetitive) DNA sequences. Any breakage at a duplex site induces a chromosome breakage and, if the complementary broken ends interact, an exchange chromosomal rearrangement may arise. The site of DNA homology can be approximately 200 bp (Belyaev and Akifjev, 1988). In Romanov’s (1980) opinion, the DNA molecules cannot come into contact in a repeat region; rather they come into contact in the chromosome loop-forming palindromes. Association of transposable elements with chromosomal rearrangement breakpoints has been frequently noted, including roo-Bt 04 (McGinnis and Beckendorf, 1983; Alatortsev, 1988; Tschudi et al., 1992), hobo (Lim et al., 1983; Lim,

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation 250

205

a

200

150

1W

50

0

0

1

2

3

4

5

8

7

8

9

10 11

12

13 14

15

16

17

18

1

2

3

1

5

6

7

6

9

10 11

12

13

15

16

17

18

14

Figure 98. Variation in the distances between the chromosome regions involved in ectopic pairing (a) and regions of inversion formation (b). Abscissa, distance between regions of ectopic pairing (a) or breakpoints of inversion (b) expressed as units of Bridges’ chromosome map; ordinate, number of events. (a) after lvaschenko et al. (1991); (b) after Hilliker (1985) and Vagapova (1991).

1988))P-element (Engels and Preston, 1981a,b, 1984),foldback (Bingham, 1981; Levis et al., 1982), and various other elements (Aleksandrova et al., 1989). The sharp increase in the frequencies of chromosomal rearrangements in wild populations and laboratory stocks of Dosophila occasionally appeared inex-

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plicable (Novitski, 1961; Levitan, 1963; Yamaguchi and Mukai, 1974; Hinton, 1979; Yannopoulos and Zacharopoulou, 1980; Yannopoulos et al., 1982). Transposable elements have been shown to cause this increase (Yannopoulos et al., 1987).Thus the presence of the hobo element in every one of the 29 rearrangement breakpoints was inferred from the results of in situ hybridization experiments in an unstable X chromosome line of D. melanoguster. The hobo transposon differed qualitatively in its ability to be involved in rearrangement formation: when lying in the same orientation, the two hobos produced a deletion; when in different orientations, they produced an inversion (Lim, 1988). However, the high frequencies of chromosomal rearrangements in the regions of intercalary heterochromatin are difficult to account for by mobile elements because ( 1) there is, nevertheless, no correlation between peaks of rearrangements (see Figure 82) and the location of mobile elements; and (2) the locations of intercalary heterochromatin are invariable, while those of mobile elements are not. Lee (1975) found a strong correlation between the location of rearrangements in the chromosomes and tandem repeats. Similar correlation was found as a result of comparison of chromosome rearrangement frequencies induced by gamma-ray irradiation in sibling species, c h . th. thummi and Ch. th. piger. The former species contains bulky of tandem copies of repeats located in pericentric heterochromatic regions. Majority of chromosome rearrangements were induced in these regions. Ch. th. pigo almost does not contain repeats, and no rearrangements were induced in this species (Gunderina and Aimanova, 1997). Repetitive DNA was detected in the chromosome regions of D. biseriata involved in ectopic contacts during the formation of a chromocenter and in which inversions are formed (Yoon, 1989). Thus, in compliance with all three considered hypotheses implying the involvement of two DNA molecules in exchange, there must be homology between nucleotide sequences: mobile elements, palindromes, or tandem repeats. Finally, there is one more hypothesis. In the view of Zhimulev et al. ( 1982), conceding that tandem repeats are located in the regions of IH, breaks in the regions of repeat do not affect viability as strongly as damage of unique DNA fragments. For this reason, rearangements in the IH regions are more readily identified.

5 . Somatic pairing I t is known that homologous polytene chromosomes in Diptera pair mainly together, thereby reducing the number of visible chromosomes in nuclei to haploid. The synaptic state of the chromosomes in a part of the nuclei is impaired, and the homologs become disposed independently in them (for details, see Zhimulev, 199213, 1996). The transition points of asynaptic to synaptic state in polytene chromosomes are not randomly distributed, being restricted to particular regions (Ribbert, 1967, 1979; Polyanskaya, 1974; Kulichkov and Belyaeva, 1975). It should be noted that the dense black bands in these border regions of

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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homologous chromosomes pair, while the neighboring thinner bands are already in an asynaptic state; that is, the dense bands are less frequently asynaptic. These dense bands (see Figure 82) were referred to as intercalary heterochromatin by other criteria (Kulichkov and Belyaeva, 1975; Polyanskaya, 1975a,b, 1976; Evgen’ev and Polyanskaya, 1976; Lapta, 1977, 1980; Polyanskaya and Samokish, 1978; Vagapova, 1991). The IH regions produce the impression of being more “adhesive,” that is, pairing not only with “nonhomologous” (ectopically) but also with homologous regions of the chromosomes. Modifiers of position effect, such as variation in heterochromatin amount or temperature, affect the synaptic state of the polytene chromosomes. At the low temperature of 14-1 8”C, asynapsis frequency is considerably lower in individuals without the Y chromosome (XO) than in those with two Ys ( X U ) grown at 25°C (Zhimulev and Vagapova, 1991).Asynapsis frequency is appreciably higher in females with two Y chromosomes (XXYY) than in normal females (XX) (Gersh, 1959). By analogy with the events of position effect variegation, it may be suggested that particular site-nonspecific proteins uniformly distributed along the DNA length may be involved in somatic synapsis of the chromosomes. They are greater in number in the IH regions because they contain more DNA (as a rule, large bands), which makes the IH more adhesive, and they are synapsed less frequently. When an additional dosage of heterochromatin is introduced into the genome, these proteins are redistributed, their amount per DNA length unit in the IH regions decreases, and somatic pairing weakens (Zhimulev and Vagapova, 1991).

6. Repetitive sequences Repeats of various kinds have been localized in the euchromatin of polytene chromosomes: dispersed, such as transposable elements or the tRNA genes; and tandem, such as the histone or the ribosomal RNA genes, satellite DNA, and oligonucleotide tracts. At least 50% of the eukaryotic genome is an alternating interspersion of short (300-bp) repeats adjacent to longer stretches of nonrepetitive sequences approximately 1500 bp long. This pattern of sequence arrangement has been found in Xenopus and it is named the “Xenopus pattern” of interspersion. Other organisms have the “Drosophila pattern” of interspersion, with much longer repeats (5.6 kb) adjacent to unique sequences longer than 10 kb (Davidson et al., 1973; Manning et al., 1975; Crain et al., 1976; Samols and Swift, 1979a). This section is aimed at clarifying the relation of the repeat location in a chromosome region to the manifestation of the properties of the intercalary heterochromat in. a. tRNA genes The Drosophila genome presumably contains 590-670 tRNA genes, approximately 60 families. The organization of the families is complex because they are

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not clustered, nor are they scattered throughout the genome as are transposable elements. More than 30 chromosome regions, which contain from 1 to 18 tRNA genes, were detected instead. The number and types of genes, the orientation of transcription, and the size of spacers between the genes in such “nests” are different (Spradling and Rubin, 1981). Analysis (Zhimulev et al., 1982) revealed that, of the 35 regions of polytene chromosomes where tRNAs were identified, features of the IH (e.g., late replication) were detected in only 16 regions. It is noteworthy that two clusters of tRNA location (42A and 56F) are diametrically opposite for this feature: complete absence of late replication at the 42A and extremely delayed replication completion at the 56F region. It should be noted that the tandemly repetitive 5s rRNA genes are located in the latter region (see later).

b. Repeats included in the genes Comparatively short sequences making up a part of the structure of many genes were identified in the eukaryotic genomes:

1. The “suffix” was originally derived from the cut gene. It is a 265+nucleotide, highly conserved sequence located at the end of many genes. The repeat is conserved; it contains a short (26-bp) intron and a variable part including a signal and a polyadenylation site. It is present as 300-400 copies in the Dosophila genome. Sequences homologous to the suffix were detected in DNA prepared from representatives of 17 taxonomic groups (the ant, molluscs, worms, the sea urchin, the clawed frog, the lizard, and humans, among others) (Tchurikov et al., 1982, 1986; Kolesnikov and Tchurikov, 1991). The central domain of the suffix is homologous to the 16s ribosomal sequence from the endosymbiotic organisms. The opposite DNA strand of the element encodes C domains of reverse transcriptase in F- and Doc-elements (Tchurikov, 1996). 2. The “opa” family was originally derived from the Notch gene. Opa is 93 bp long, and it is present in many developmentally regulated genes. The sequence is composed mainly of the triplets CAG and CAA. At least 30 location sites were identified by in situ hybridization. The number of silver grains in the sites varies (Wharton et al., 1985). 3. “Pen” repeat sequences are present in many cDNA clones. They are clusters of GGN triplets, where N can be any nucleotide. The repeat encodes a glycine-rich domain homologous to the rat DNA helix-destabilizing protein (Haynes et al., 1987). 4. According to repeat mapping at the early ecdysone puffs 63F of D. melanoguster and 90B of D. hydei, there are up to 15 repeats consisting of 229 nucleotides in the locus. Homologous repeats are present in the genomes of amphibia, reptiles, and mammals (Izquierdo et al., 1984).

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5 . The homeobox is a short (180-bp) sequence, a structural component of many genes active during early embryogenesis. It is represented by multiple copies in the genome (McGinnis et al., 1984; Gehring, 1985). 6. The GC-rich repeat (consists of GC by 75%) was found in the Dosophila genome. It has been localized in six regions of the genome (15EF, 50DE, 57BC, 59C5, 79B, and 100DE). It has been suggested that these sequences encode the glycine-rich domains in the proteins of Dosophila (Flavell et al.,

1987). 7. Tandem repeats consisting of four GATA nucleotides were detected in the genomes of the calliphorids Chrysomya rufifucies and Calliphora erythrocephula. They are actively transcribed in poly(A)+ RNA. Several heavily and many weakly labeled regions were revealed by in s i t u hybridization. Comparisons of genomic DNA from embryos and adults have disclosed appreciable differences in the length of the repeat (Kirchhoff, 1988). There is no evidence indicating that there is a relation between the location of the repeats and intercalary heterochromatin. Two genes containing the homeoboxes Bx-C and Antp-C are located in the intercalary heterochromatic 89E and 83AB regions. However, it is difficult to believe that the 0.18-kb DNA of the homeobox might have conferred heterochromatic properties to large bands, or even to band sets.

c. Mobile elements of the genome From their early studies, Georgiev and Gvozdev (1980) inferred that the mobile elements are predominantly located in the IH regions. Ilyin et al. (1978) have adopted the view that “mobile elements are predominantly concentrated in the IH regions which may be regarded as chromosomal loci where the strongly expressed repetitive genes, which readily transpose from site to site, are located” (p. 969). Ananiev et al. (1979a) have established that the “location of the Dm 225 and Dm 234B genes may serve as an additional criterion for the identification of the IH regions” (p. 793). Based on the first mapping results of transposable elements on polytene chromosomes, it was claimed that intercalary heterochromatin may be regarded as centers (“nests”) containing several types of actively transcribed, tandemly repeated genes encoding nonspecialized cell functions and products the cell needs in large amounts (Ananiev et al., 1979a,b; Gvozdev, 1981a,b). It is the view ofTchurikov et al. (1981) that various mobile elements “are located in close vicinity, thus forming long sequences from middle repetitive DNA in the Dosophila genome. Such regions with informational content vary in different individuals, and thus probably, correspond to regions of the IH” (p. 658). Ananiev and Barsky (1985) conceded that “accumulation of mobile genetic ele-

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ments at a concentration exceeding the critical leads to the acquirement of features of intercalary heterochromatin by these regions” (p. 21). The location of various mobile elements in polytene chromosomes of Drosophila strains has been the topic of intense investigation (Ananiev et al., 1978, 1979a,b, 1984a-c; Young, 1979; E. Sp. Belyaevaet al., 1981,1982, 1984a,b, 1989; Gvozdev et al., 1981; Pardue and Dawid, 1981; Pierce and Lucchesi, 1981; Tchurikov et al., 1980, 1981; Young and Schwartz, 1981; E. Sp. Belyaeva, 1984; Pasyukova et at., 1984, 1986; Ananiev and Barsky, 1985; Biemont et at., 1985; Pasyukova and Gvozdev, 1986; Vasilyeva et al., 1987, 1988; Biemont and Gautier, 1988; Kholodilov et al., 1988; Sorsa, 1988b; Aleksandrova et al., 1989; Leibovitch, 1990a,b;Glushkova et al., 1991; Goryachkovskayaand Vasilyeva, 1991; Kolesnikova et al., 1991; Ratner et al., 1992a,b). As a result more than 300 location sites have been identified (Ananiev et al., 1984a). Four general conclusions are relevant:

1. Virtually every individual possesses its own unique set of insertion sites. 2. The mobile element is detected in insertional “hot spots” within particular chromosome regions more frequently in one Drosophila strain compared to others. The assignment of a region to a “hot spot” is quite tentative: for example, there were no differences for 97 location sites of h 2029 in 10 giant larvae, (i.e., the occurrence was 100%) (Ananiev et al., 1984a). In contrast, a chromosome region in which a mobile element occurs at a frequency not lower than one-half of individuals is viewed as a hot spot (Leibovitch, 1990a) or a location “in which insertions are more frequent than in the other regions” (Biemont and Aouar, 1987, p. 40). In certain populations, variations in location in the X chromosome were so wide that it was not feasible to identify the preferential location sites of mobile elements (Montgomery and Langley, 1983; Leigh Brown and Moss, 1987). However, investigations of Pasyukova and Gvozdev (1986) and Leibovitch (1990a) have revealed, along with high heterogeneity, hot spots as well. 3. Comparisons of various strains revealed coincident location of mobile elements precisely in the hot spots regions (Figure 99), presumably owing to the presence of specific nucleotide sequences needed for insertion. The mobile element mgdl inserts into the same region, 93EE to a precision of several tens of nucleotide pairs, in Oregon-R, Swedish-b, GB-39, and 171 strains, as ~~

Figure 99. Location of four mobile elements. The cytological maps of the chromosomes of Drosophila are represented as five horizontal lines in each figure. Numbers designate the subdivisions of maps. Black circles designate the presence of a mobile element in one of the strains. Elements in the chromosomes of inbred strains: (a) mdgl, (b) copia, (c) I, and (d)

P. After Biemont and Gautier (1988).

L

-K

L

e

X

L

T

'Tt

X

e

C X

a

f

d X

a

Figure 99. Continued

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

2 13

demonstrated by analysis of DNA from this region (Shevelyov and Gvozdev,

1987).

4. Mobile elements can insert in certain chromosome regions. For example, insertions of mdgl , mdg4, D m 2027, Dm 2029, Drn 2066, D m 2068, Dm 2074, Dm 2078, and the copia type have been detected at the 41AB region of the gt strain (Ananiev et al., 1984a). Regions where three to four elements are located are not exceptional. Several lines of evidence suggest that there may be a relation between the location site of a mobile element and intercalary hetrochromatin: For example, mapping of mdgl , one of the earliest elements used in transposable element localizations, in many regions of intercalary heterochromatin in the giant strain; and instances in which ectopic contacts (see Table 13) and regions of chromosome breakage label (Ananiev et al., 1978, 1979a; Gvozdev, 1981a,b; Gvozdev et al., 1981; Tchurikov et al., 1981; E. Sp. Belyaeva et al., 1984a,b). It is true that, of the 95 insertion sites of mdgl , 57 were mapped within the regions of intercalary heterochromatin. However, some of the typically intercalary regions of heterochromatin, such as 1lA, 35EE and 75C, were not “filled.” These data are subject to criticism. The insertion sites and intercalary heterochromatin are not always precisely mapped. They have been associated simply because they are neighbors. Localization of Dm 47 with accuracy to the region 59ABC has allowed it to be ascribed to intercalary heterochromatin (Tchurikov et al., 1981), although it has been precisely localized at the neighboring 59D1-2 band and 59AC shows no features ofheterochromatin (Zhimulev and Kulichkov, 1977). The localization of mdgl is very often unacceptably imprecise; for example, it has been localized at the 12CDE band, ignoring the fact that only bands of the 12E region exhibit the properties of intercalary heterochromatin. The other regions possess other properties, although lying in close proximity. Intercalary heterochromatin has not been identified at 27AB, 29EF, or 49EE which are the location sites of Dm 47; nonetheless, the regions are referred to intercalary heterochromatin. There occasionally were failures to make proper distinctions in comparisons of the location sites of mobile elements with other chromosome features. Thus bands 3C, 7B, 9A, 11A, 13A, 19F, 20C, 22A, 25A, 30A, 32F, 33C, 35C, 36CD, 39D, 42B, 56F, 59D, 64C, 67DE, 70C, 71C, 75C, 81F, 83DE, 84D, 86D, 87F, 89E, 92C, 94A, 95A, and 98C have been referred to regions of intercalary heterochromatin in Figure 1 and another group, 7A, 7E, 19F, 23A, 30A, 33C, 34D, 34EF, 35D, 41CD, 56F, 59D, 65A, 67BC, 75F, 82DE, 84A, 88E, 90A, 96BC, 98BC, 99A, 99D, 99F, and lOOF, have been referred to the same regions in Figure 2 by E. Sp. Belyaeva (1984). There is an overlap between the lists for only six regions.

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When the mdgl localizations for strains other than giant are considered, the association with intercalary heterochromatin becomes rather tenous. In analysis of 20 strains, only five mdgl elements have been found in only 33 intercalary regions (E. Sp. Belyaeva, 1984). In the D32 strain, only eight mdgl sites are identified in the two arms of the second chromosome, and only 35D and 58A have properties of heterochromatin (Aleksandrova et al., 1989). Studies of the localization of mdgl in the chromosomes of 17 inbred strains of Drosophila (see Figure 99) revealed 102 insertion sites in which an insertion occurred in at least one strain (Biemont and Gautier, 1988). Analysis of these data, in conjunction with those on late replication [154 late replication regions have been identified in the Drosophila genome (Zhimulev et al., 1982)], has led to the conclusion that there are 49 regions containing mdgl and late-replicating DNA, 53 containing only mdgl , and about 100 containing only the latereplicating DNA in these 17 strains (Table 15).These data hardly prove that mdgl is a diagnostic feature. Mention has been made in the literature that there is a relation between the location of certain other mobile elements and IH. For example, a part of the locations of the hot spots of the DNA fragments for Dm 85, S34, Dm 67, and Dm 132 containing mobile elements coincide with the IH regions, with these coincidences being appreciably higher ( 1 9 4 6 % ) in the high-altitude than the coastal regions of Azerbaidjan (21-29%) (Leibovitch, 1990a). Hybridization of Dm 25 with the chromosomes of 15 isogenic and laboratory stocks revealed 38 insertion sites (Pierce and Lucchesi, 1981); of these, 17 sites in one to six strains were coincident with the late-replicating regions. Obviously, the great majority of the late-replicating sites do not contain insertions. In 17 inbred strains, the localization of four mobile elements was studied: the already mentioned mdgl, copia, and I- and P-elements (see Table 15). There was no relation between the IH and mobile elements. As for the P-element, having examined 272 independent cases of transformation, Spradling ( 1986) concluded that the P-derived transposon inserts randomly. Analysis of a great number of various mobile elements in polytene chromosomes revealed no specificity in their location (Zhimulev et al., 1982). In a very precise analysis of the localization of mobile elements of 12 families carried out on stretched polytene chromosomes, approximately 300 of their location sites were identified, and there was no indication that IH is enriched in mobile elements; at any rate, the authors did not raise the issue (Ananiev et al., 1984a-c). In another report based on the same material, it was mentioned that approximately 25% of all the hybridization sites of mobile elements fall in the IH regions, mdgl most frequently maps to these regions, and the small multiple copy elements of the Dm 2029 kind are the least frequently mapped there (Ananiev and Barsky, 1985).

215

Poiytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 15 Relation of Localization of Late-Replicating Regions and Insertions of Four Mobile Elements in Chromosomes of Drosophila Number of sites Late replicating and containing insertions

O n l y late replicating

copia

49 63

I-element P-element

55 54

105 91 99 101

Mobile element d g J

O n l y insertionb 53

77 77 127

“Data on the localization of late replication from Zhimulev et al. (1982). q o t a l number of sites in 17 inbred strains (Biemont and Gautier, 1988).

A comparison of the location frequencies of mobile elements with relative amounts of DNA in particular chromosome regions demonstrated that in regions in which the IH, as a rate, occurs, such as 7B, 7C, 9A, 11A, 12EF, 39E, 53A, 57A, 57B, 58A, 60F, 64C, 70‘2, 71C, 75C, and 89E, there is no specificity. The frequencies were high in a number of regions in only one (giant) of the two studied strains (Bolshakov et al., 1985a,b). In an analysis of 10,911 location sites of 21 mobile elements of the polytene chromosomes of D. melanogaster, it was demonstrated that their number, when estimated on the basis of DNA content in a chromosome region, varies slightly from one region to another. Mobile elements are inserted less frequently in the 14+15 and 16+17 regions of the X chromosome (Leibovitch, 1990b; see also Ronserray and Anxolabehere, 1986; Biemont and Gautier, 1988), in the 26+27 and 28+29 regions of chromosome 2L, and in the 54+55 region of chromosome 2R. Insertion takes place somewhat more frequently in the 18+ 19 regions (Leibovitch, 1990b). Data are available for the location of transposable elements in the chromosomes of other dipteran species. Hybridization of DNA from the mobile element pDv I I I with the polytene chromosomes of D.eririlis (Evgen’ev et at., 1982a,c) revealed more than 170 insertion sites, which included telomeric regions of particular chromosomes, the heat shock puffs, breaks (weak spots), and the break sites of single chromosomal rearrangements. To compare the location sites of IH and a mobile element, the authors determined the degree to which replication was delayed in the insertion sites of the X chromosome and established four types of regions: (1) hybridization in situ with pDv 1 I I and late replication in both males and females (14 regions); (2) the same with the difference that replication is late in females only (24 regions); (3) no late replication, insertions detected (20 regions); and (4) late replication, no insertions (6 regions) (Gubenko and

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Evgen’ev, 1986). Thus the ratio of insertions to late replication is the same, in principle, as in D. melanogaster (see Table 15). In the pCtCl clone of Chironomus thummi thummi, 2HR hybridizes with a total number of 90 sites (27-39 sites in a laboratory strain, 22-45 in wild populations), such as the centromeric and telomeric regions, a region of intercalary heterochromatin, Balbiani rings, and small puffs (Kiknadze et al., 1987c; see also Wobus et al., 1990). Thus the information examined in this section hardly gives reason to believe that the mobile elements of the genome determine the properties of the IH.

d. Tandem repeats An approach to the detection of repeats in polytene chromosomes is in situ hybridization of labeled DNA or cRNA transcribed in vitro from the DNA of the entire genome, a major fraction, or satellites. In such a case, hybridization efficiency is related to reiteration degree of copy number in the solution used for hybridization and also to copy multiplicity in the DNA preparation, and this ultimately makes possible identification of tandem repeats. As experiments in D. melanogaster demonstrated, in hybridization of cRNA transcribed from the fast-reassociating DNA, the regions of centromeric heterochromatin and the 21D1-2 region labeled first (the authors of the two papers did not identify the 21D1-2 region and called it the end of chromosome 3L) (Rae, 1970; Gall et al., 1971). Furthermore, weak diffuse labeling is discerned in all the chromosome arms (Rae, 1970; Gall et al., 1971). The X chromosome labels somewhat stronger than the autosomes (Gall et al., 1971). The 21D1-2 region labels, as became obvious later (Peacock et al., 19781, owing to the hybridization of the 1.705 satellite, which includes a polypyrimidine tract (see later). Late replication and ectopic pairing are the IH features observed for this band (Plaut et al., 1966; Zhimulev and Kulichkov, 1977; Zhimulev et al., 1982). In D. hydei and related species, as well as in Rhynchosciara hollaenderi, [3H]RNA, complementary satellite DNA, and centromeric heterochromatin weakly label the regions of polytene chromosomes. Label is homogeneous with some concentrations (Hennig et al., 1970; Eckhardt and Gall, 1971). In D.oirilis the [)H]RNA transcribed from middle repeats (Cot100) is detected along the length of the chromosomes (Evgen’ev and Shilov, 1977). The bulkof [3H]RNAtranscribed from the fractionated satellites of D. virilis is mapped to the chromocenter. A small amount of label is detected in the euchromatic parts of the chromosomes: the IH satellite in two regions in D. americanu americana and in four regions in D. novamexicana. A small number of hybridization sites are also observed for satellites I1 and 111 (Cohen and Bowman, 1979). In hybridization of a repeat 15 bp long (the 1.696 satellite) of D. simulans with polytene chromosomes of the same species, label is detected in the chromocenter and three regions in the chromosome arms (Lohe and Roberts, 1988).

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The cRNA transcribed from total DNA, as well as from the three DNA fractions of the salivary glands of Rhynchosciara americana, the fast, intermediate, and slow reassociating, preferentially hybridize with centromeric heterochromatin; however, intermediate and middle repeats also label the euchromatic parts of the chromosomes (Machado-Santelli et al., 1979). In hybridization of [3H]RNA transcribed from the bulk (or the whole) genome of various species, polytene chromosomes usually label quite heavily (Hennig et al., 1970; Jones and Robertson, 1970; Eckhardt and Gall, 1971; Gall et al., 1971; Berendes et al., 1974; Rudkin and Tartof, 1974; Evgen’ev and Gauze, 1975; Alonso et al., 1977b; Machado-Santelli et al., 1979; Samols and Swift, 1979a,b;Gvozdev et al., 1980a,b; Sharakhova, 1990). Silver-grain counts in chramosome regions performed in some of these studies demonstrated that hybridization is not random; rather, it is preferential in certain regions (Rudkin and Tartof, 1974; Alonso et al., 1977b; Gvozdev et at., 1980a,b). Analysis of small chromosome fragments of Drosoghila (1A-13F of the X chromosome and 54A-60F of chromosome 2R) following hybridization with cRNA demonstrated strong labeling in the regions of the chromocenter, the nucleolar organizer, 56EE IAB, 3C, and 12CE However, intensity of specific (i.e., per unit of DNA mass) binding of cRNA to these regions did not differ significantly from the mean distribution along the chromosome (Rudkin and Tartof, 1974). Based on this observation, the conclusion was that no chromosome regions are preferentially enriched in repetitive sequences. Relying on these data, the authors assumed correlation between the degree of enrichment in repeats and IH features, such as late replication and ectopic pairing, and they found that the assumption does not hold true (Rudkin and Tartof, 1974). However, relying on these same data, Lee (1975) disclosed a strong correlation between the frequency of chromosomal rearrangements induced by Kaufmann (1946). In Rudkin and Tartof’s study (1974), grain number was not counted over bands in a region, but rather over whole sections of the map (e.g., 11A-F) after in situ hybridization, and this surely failed to brinb into prominence the exclusive band of intercalary heterochromatin in the 11A6-9 band. Hybridization of cRNA with the polytene chromosomes of D. melanogaster, followed by localization of the label in strictly defined sites, has demonstrated strong labeling in the 3C, 9A, 12E, 13A, 19AC, 19DE, 39DE, 42B, 44D, 50A, 50C, 53AC, 56E 57A, 61CD, 63E, 67C, 67D, 68C, 82E, 83DE, and lOOC regions (see Figure 82) (Gvozdev et al., 1980a,b). Specific binding intensity in the IH regions has been shown to exceed random values (Bolshakov et al., 1985a,b). The previously considered data of Evgen’ev et al. indirectly support the idea that the IH regions are enriched in repeats (Evgen’ev et al., 1977; Levin et al., 1977). The data are consistent, therefore, in indicating that clusters of tandem

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repeats are present in polytene chromosomes. Some of these repeats are characterized at length in this section. (1). The rRNA genes The repeated 18s and 28s rRNA genes comprise approximately 2% of the Drosophila genome (or 20% of middle repeats). They are concentrated in the nucleolar organizers of the Y and X chromosomes. The number of cistrons in these genes varies, being, on average, 250 in the X chromosome and occurring in somewhat smaller numbers in the Y chromosome (for further details see Spradling and Rubin, 1981; Leibovitch, 1984; George et al., 1996). Replication is completed later by the regions of the nucleolar organizer in Phryne, D. nusuta, and Simulium ornutipes than by the rest of the DNA of the chromosomes (Sokoloff, 1977; Lakhotia and Roy, 1979; Bedo, 1982a), alttaugh in these cases it cannot be excluded that precisely the heterochromatic blocks surrounding the nucleolar organizer are late-replicating.In Rhynchosciarahollaenderi, the location region of rRNA is late-replicating (Pardue et al., 1970), satellite DNA and tracts of poly(A) sequences were identified in it (Stocker et al., 1978). In the ovarian nurse cells of Calliphora erythrocephala, D. hydei, and Sarcophaga barbata, the percentages of the rRNA genes are 135,51 and 47 of that in diploid cells (Renkawitz and Kunz, 1975). In Chironomus tentans, the location site of rRNA genes shows no features of heterochromatin and rDNA is proportionately represented without underreplication (Hollenberg, 1976). Every one of the 160 5s rRNA genes in Drosophila consists of an encoding region 135 nucleotides long (15 nucleotides at the 3' end are removed later, thereby yielding a total number of 120 nucleotides) and a spacer 238 nucleotides in length. The size of the spacer can vary (for comprehensive reviews, see Spradling and Rubin, 1981; Leibovitch, 1984). This cluster has been localized to the 56F section (Wimber and Steffensen, 1970; Grigliatti et al., 1974), which is the latest replication in chromosome 2R and is quite frequently involved in ectopic contacts (Zhimulev et al., 1982). Underrepresentation (Szabo et al., 1977; Renkawitz-Pohl, 1979) and breaks (Zhimulev and Kulichkov, 1977) have not been found to occur in the region of this cluster. The 5s rRNA genes have been mapped to the large dense 2-23B1-2 band (Alonso and Berendes, 1975) in D. hydei and to the 97E region in D. otirilis (Cohen, 197613; Wimber and Wimber, 1977; Kress et al., 1985). In D. hydei, the underreplication degree in the 5s rRNA genes is 21% in salivary gland and 28% in ovarian nurse cells (Renkawitz-Pohl, 1978). The location sites of RNA in the D. psewloobscura XL chromosome exhibit weak points (Steinemann, 1982b).The location sites of the 5s rRNA genes in Glyptotendipesbarbipes, Chironomus tentans, and Ch. thummi and in Drosophila miranda show no characteristics allowing them to be recognized as heterochromatic (Wen et al., 1974; Wieslander, 1975; Wieslander et al., 1975; Baumlein and Wobus, 1976; Wenand Hague, 1979;Steinemann, 1982b).

5s

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

219

In Lucilia cuprina, a block of the 5s rRNA genes has also been localized at a single site (Bedo and Webb, 1990), also showing no features of heterochromatin. (2.) The genes encoding histones In D. melanoguster, the genes encoding these chromosomal proteins are arranged in a cluster with a total length of about 500 kb (100 gene sets of about 5 kb each) (Lifton et al., 1978), and they are located in the 39DE region of chromosome 2L (Bimstiel et al., 1973; Wimber and Steffensen, 1973; Pardue et al., 1977; Fitch et at., 1990).The middle part of the 39E1-4 band does not polytenize fully (Lifschytz, 1983),and, as a consquence, breaks have been shown to occur frequently in this region (Zhimulev and Kulichkov, 1977). In Ch. th. thummi, five histone genes span a DNA fragment of 6262 bp, which hybridizes in polytene chromosomes with five bands in arm D (Hankeln and Schmidt, 1990). A cluster of the histone genes (of 5153 bp and reiterated 120-140 times) maps to the 50A band in the third chromosome of D. hydei (Fitch et al., 1990; Kremer and Hennig, 1990), and to two regions of polytene chromosomes of D. virilis and D. huwaiiensis (Anderson and Lengyel, 1984; Domier et al., 1986; Fitch et al., 1990), and it maps to a single band of D. pseudoobscura (Steinemann, 198210).There is no information that these bands show features of intercalary heterochromatin. This may be due to the species-dependent and wide variation in copy number per haploid genome in tandem repeats: from 100 to 110 in D. melanoguster, 20 in D. huwaiiensis, 5-10 in D. hydei, and approximately 50 in D. virilis (for reference, see Fitch et al., 1990). (3.) Polypyrimidines The pyrimidine strand can be isolated following depurination of ['HIDNA with formic acid with diphenylamine. However, some DNA fractions are resistant to such treatment, and, as a result, whole strands consisting of pyrimidines can be liberated. In Drosophila, up to 3% of [3H]thymidine label is detected in such strands. They are of variable length, from 100 to 1000 nucleotides (750 nucleotides on average). Polypyrimidines make up a considerable part of the cryptic satellite in total DNA. In CsCl this satellite yields density gradients with a 1.707 peak, clearly distinguishable from the major 1.702 peak. The DNA of the cryptic satellite is composed of 34.5% deoxycytidine and 65% thymidine (see Cseko et al., 1979). The RNA transcribed from these template RNAs hybridizes very efficiently to Drosophila DNA at very low Cot values; it is located mainly in centromeric heterochromatin, and to a lesser extent in the chromosome arms, where a band is identified at the 21D1-2 region (Bimboim and Sederoff, 1975; Sederoff etal., 1975a,b). It was found that this band is 2101-2 (see earlier) in Chironomids. (4.) Cla-elements in Chironumus thummi The genomes of the chironomid subspecies Chironomus thummi thummi and Ch. th. piger differ in DNA amounts: there is 27% more DNA in the former (Keyl, 1965b). It should be noted that the band sequence is the same in the two species (Keyl and Strenzke, 1956). However, in Ch. th. thummimany bands and centromeric heterochromatin

220

1. F. Zhimulev

blocks are much thicker, they contain more DNA presumably owing to duplications. The ratios between successive DNA amounts in at least 20 homologous bands in Ch. th. thummi are in geometrical progression: 1:2, 1 4 , 1:8, and 1:16 (Keyl and Strenzke, 1956; Keyl, 1957, 1962, 1964a, 1965a,b, 1966). This difference in DNA amount is due to the additional content of a fastdenaturating, AT-rich DNA fraction (Keyl, 1964b, 1965a; Schmidt et al., 1980) represented by a sequence approximately 120 bp long tandemly reiterated up to 50 times (Schmidt et al., 1982). A restriction site for Cla I endonuclease is present in it; 5.5 times more Cla-elementsoccur in the Ch. th. thummi than in the Ch. th. piger genome (Schaeferand Schmidt, 1981;Schmidt, 1981,1989;Vistorin and Schmidt, 1983; Zainiev et al., 1986).This repeat is located mainly in the block of centromeric heterochromatin in Ch. th. piger (Figure loo), while being present in more than 200 chromosome regions in Ch. th. thummi (Schmidt, 1980, 1981, 1984, 1988, 1989; Hankeln et al., 1989). These elements have been identified even in the nontranscribed spacers between the ribosomal DNA genes (Schmidt et al., 1982; Israelewski and Schmidt, 1982; Israelewski, 1983). It has been suggested that Cla-elements may be mobile because they are flanked by duplications from 5 to 12 bp in size (Schmidt and Godwin, 1983,1987; Schmidt, 1984). Many location sites of Cla repeats stain positively for C-heterochromatin (Hagele, 1977a) and replicate late (Keyl and Pelling, 1963; Keyl, 1966; Hagele, 1970, 1976, 1977a). A specific variant of histone, H1-11, has been detected in bands in Ch. th. thummi (Mohr et al., 1989). Cla repeats are weakly represented in Ch. dorsalis. In Ch. halophilis, the DNA hybridizing with the Cla-element in Southern blot analysis varies in size from more than 25 to only 2 kb, which indicates that a specific restriction site has been lost (Hankeln et at., 1989; Hankeln, 1990). In Ch. luridus, a smear is also detected on blots, and a 90-bp fragment is the repeat unit (Hankeln et al., 1989; Hankeln, 1990). In addition to the family of Cla repeats, the Alu family of repeats (8000 copies/genome),which are organized in clusters of various lengths, were identified in Ch. th. thummi. A 170-bpfragment consisting of 70% AT is its repeat unit. I t is weakly represented in other species. Hinf-elements (repeat unit of 110 bp, consisting of 70% AT) are presumably restricted to the Ch. th. thummi and Ch. th. piger genomes. In situ hybridization reveals 35 sites in both species. The repeats are arranged in clusters (Hankeln et al., 1989; Hankeln, 1990). 15.) Poly[A) nucleotide complexes Poly(A)-containingtracts are detected by various methods; for example, [3H]polyuridylicacid hybridizes with the polytene chromosomes at approximately 10-1 2 loci, mainly in the centromeric regions, in telomeres, and in several intercalary heterochromatic regions in Rhynchosciura angelae (Joneset al., 1973).There are no data indicating that these tracts are related to the IH. In D. hydei, [3H]poly(U)hybridizes with polytene chromo-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

221

Figure 100. In situ hybridization of Cka-element on the polytene chromosomes of Chironomus thummi thummi (th) X Ch. th. piger (pi). Black arrows, position of centromeres; white arrows,

labeled sites in Ch. th. thummi. After Schmidt (1984).

somes quite uniformly (Alonso et at., 1977a). Poly(AT) tracts are readily identified after Q- and H-banding in many dipteran species (see Sections I1 and XVI). When an E. coli RNA polymerasemixture containing [3H]UTPis applied to fixed polytene chromosomes, label is preferentially incorporated into the 8 lF, lOlE and 102DE regions. In another experiment, actinomycin D was added to the [3H]ATP-containingreaction mixture, which binds to GC pairs and halts transcription where these nucleotides are present. After this treatment, the chromosomes do not label along their length, except for the same 81F, lOlF, and 102DE regions (Leibovitch et al., 1974), which are Q' and H' (see Section VII,B,2).

222

1. F. Zhirnulev

A specificity after treatment of a squash preparation of polytene chromosomes with restriction enzymes, followed by staining with undegraded DNA, was observed for these regions. These were chromosome regions from which restriction sites for the given restrictase were extensively missing (because of repetivity). In D. melanoguster, regions 81F, 83E, 101, 102C, 102F, and 56EF remained unstained after treatment with the enzyme Alu I. The list of those regions unstained after treatment with Hue 111was similar, but not completely identical: 101, 102C, 102F, the telomeres of the X chromosomes, 2R, 3R, and several unidentified regions (Mezzanotte, 1986). There are sites specific to restrictases in the D. virilis and Sarcophagabullata genomes, for example, blocks of the histone genes and telomeres of two chromosomes in D. virilis (Bultmann and Mezzanotte, 1987; Mezzanotte et al., 1987; Lopez-Femandes et al., 1991). A tandem repeat with a unit of 279 bp has been identified in the chromosomes of the foot pad cells of the pupae of Sarcophaga bullata. A motif of 17 bp identical to the sequence reiterated five times in the satellite DNA of the Bermuda crab was identified within it. The repeat is mapped in situ to the Q+-centromeric regions of the C and E chromosomes, in the euchromatic arms of the chromosomes. When the chromosomes are asynapsed, one or two homologs can label (Hershfield and Swift, 1990). (6.) Repeats specific to the D. melanogaster X chromosomes A sequence 372 bp long and rich in AT nucleotides, which is reiterated300400 times, has been isolated from the D. melanoguster genome. As a result of in sit% hybridization, a labeling site in the autosomes (the 26A region), as well as approximately 10 major and 20 minor labeling sites, were identified in the central part of the X chromosome between the 4 and 14A regions in polytene chromosomes (Waring and Pollack, 1987). Judging by the figure in that paper, the most typical IH region (11A) as well as some of the late-replicating bands (7E, 8B, 9A, 10A) label heavily. (See also Kokoza et al., 1997, for details.) (7.) Mono- and dinucleotide repeats Three repeat types (dCdA)n.(dT-dG)n, (dC-dT)n.(dA-dG)n, and (dC)n.(dG)n, frequently occur in eukaryotes, and also in Diptera (Tautz and Renz, 1984). Using in situ hybridization, six Drosophila species (D. melanoguster, uirilis, miramla, simulans, pseudoobscura, and hydei) were shown to be uniformly distributed among all the autosomes (except for the microchromosomes and centromeric heterochromatin, where they are not identified). The level of labeling is approximately twofold higher in the X chromosomes (Huijser et al., 1987; Pardue et a/., 1987; Vashakidze et al., 1988a,b, 1990; Lowenhaupt et al., 1989; Miklos and Cotsell, 1990; Pardue, 1990). In males of D. melanogaster, the sole X chromosome in the karyotype functions twice as actively as each of the two chromosomes in the female (i.e., “dosage compensation” takes place). The increase in the amounts of mono- and

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

223

dinucleotide repeats in the dosage-compensated chromosome is, possibly, in some way related to the maintenance of hyperactivity. It was shown that certain proteins with a molecular mass of 67-72 kDa preferentially bind to DNA of this type during embryonic development (Vashakidze et al., 1988a). In D. mzra&, one of the ancestral autosomes has fused with the Y chromosome, thereby giving rise to a new element, the neo-Y, in which the former autosome at the given step of evolution has acquired features of heterochromatin, and numerous repetitive sequences, for example, appeared in it (Steinemann, 1982a). Mono- and dinucleotide sequences do not emerge in the neo-Y, as they do in the other heterochromatic regions (Lowenhaupt et al., 1989). In D. subobscuru, 14 DNA clones were isolated. Characteristic of these are 100-300 hybridization sites in the euchromatic parts of the chromosomes; furthermore, they occasionally label the telomeres and centromeric regions. Several clones predominantly labeled the X chromosome (Felger and Sperlich, 1989), thereby suggesting the presence of mono- and dinucleotide sequences. (8.) Repeats of the “pDw” family Repetitive elements pDv, which map to approximately 200 sites of polytene chromosomes, were identified in Dosophila of the virilis group. They are characterized by a rather stable location in the D. virilis genome, intense transcription, partial polytenization in the salivary gland chromosomes, and, moreover, their ability to transpose in interspecific hybrids. Based on determination of primary sequences, it was shown that the repeats contain tandem arrays of 36-bp repeats flanked by incomplete terminal repeats of approximately 80 bp. The 36-bp repeats can be arranged in three different orientations: head to head, tail to tail, and tail to head. These repeats can be included in a domain 5-6 kb in length. In situ hybridization revealed that grain number in a labeling site can vary presumably because of the variation in the number of 36-bp fragments in each element (Evgen’evet al., 1982a-c; Evgen’ev and Zelentsova, 1984; Zelentsova et al., 1986; Vashakidze et al., 1987, 1989; Sayanova et al., 1987). No information about the relation of the repeats to the IH has been provided. (9.) Scrambled repeats Scrambled repeats constitute an extreme paradigm of middle repeats (Wensink et al., 1979). The repetitive elements are arranged in clusters of 300-1000 bp. Each cluster covers a region several thousand base pairs long and contains sequences of various kinds. The individual repeats are frequently reiterated within a cluster, occasionally in inverted orientation. The elements of a cluster can be found in several thousands of other clusters located in centromeric heterochromatin and chromosome arms. A detailed analysis of four clusters revealed that they contain a minimum number of 17 of different families. It is conceivable that about one-fourth of all middle repetitive sequences are composed of clusters of repeats of this type (Spradling and Rubin, 1981).

224

1. F. Zhimulev

Some middle repeats do not conform to the structural paradigm described previously. For example, the y-element is represented by 30 copies per haploid genome (Lis et al., 1978). One half of these copies are located in different chromosome regions, and the other half are located at the 87C region. The tandemly arranged copies are similar; they are scattered and dispersed to a large extent.

7. Contacts with the nuclear membrane It has long been known that the chromosomes make contact with the nuclear envelope (see the review by Zhimulev et al., 1992b, 1996). Kaufmann and Gay (1958) published an EM photograph of a section through the salivary gland nucleus of Drosophila. The contact between the nuclear envelope and a group of bands showing established ectopic contacts is clearly seen. There is information that ectopic pairing fibers frequently contact the envelope (Holmquist and Steffensen, 1973). However, mapping of regions involved in contacts became feasible only after development of optical section methods with computer treatment of the results. It was found that about 30 regions lie in the nuclear periphery in the cells of gut, salivary and prothoracic glands much more frequently than the others. These are lAF, 2B, 3C, 4D, 5EF, 9A, 12E, 12F, 16D, 18F, and 20CF in the X chromosome; 21A, 22A, 25DE, 35AE, 36D, and 40F in chromosome 2L; 41AE 42AD, 57AC, and 59D in chromosome 2R; 61A, 64C, 70C, 71C, and 79-80 in chromosome 3L; and 81,82AC, 83DE, 88AC, 89E, and lOOF in chromosome 3R (Mathog et al., 1984; Hochstrasser et al., 1986; Hochstrasser and Sedat, 1987b). Obviously, the majority, although not all, of the listed regions are intercalary heterochromatic. However, a number of regions showing very prominent features of intercalary heterochromatin (11A, 19E, 39E, 67D, 75C, and others, for example) are not included in this list.

8. Other features a. The Minute (M) mutations This is a class of genes whose heterozygosity (or hemizygosity) for mutations produces a characteristic syndrome in Drosophila: delayed development rate and formation of thin bristles. There are several tens or so of these genes in the genome. Hannah (1951) brought attention to correlations between the location of the M genes and intercalary (centromeric) heterochromatin. For example, 9 of the 23 M loci known for chromosome 2 lie not more than 5 cM away from the centromere. Approximately 30% of the M loci are confined within these genetic limits in the third chromosome. Another series of M loci, in her view, maps to the IH regions. The possible relation between the M genes and the IH is discussed in later papers, too (Ananiev et al., 1978, 1979a; Gvozdev, 1981a,b).

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

225

Without going into the details of organization and function, it should be noted that, of the 32 M genes mapped cytogenetically (Lindsley and Zimm, 1990), only 86D14 and 30A coincide rather well with IH localization. A single M gene is mapped to the region of p-heterochromatin, and another 10 genes are located in a wide cytologicai range within which the IH is mapped, too; for example, M(2)53 is located in the 52D-53E region and IH lies at 53AC. The coincident location may be purely statistical. Even random coincidences must be frequent at high saturation of the chromosomes with both the IH bands and the M genes. Therefore, there is no good reason to believe that the IH regions are the cytogenetic bases for the M genes. According to modern knowledge, Minute mutations in Drosophih affect ribosomal protein genes (see Saboe-Larssen and Lambertsson, 1996).

b. Cytological repeats Bridges' (1935c, 1938)suggestion that there may exist repeats-that is, fragments in different regions of the chromosomes showing similar banding patterns in polytene chromosomes (for review, see Zhimulev, 1992b, 1996)-has been followed by attempts to take advantage of them as criteria for localization of heterochromatin (Hannah, 1951; Ananiev et al., 1978,1979a; Gvozdev, 1981a,b). In conclusion, the evidence available indicates that internal repeats in the polytene chromosomes of Drosophih are rarities and are encountered in certain species only. Moreover, Bridges' first descriptions of internal repeats were erroneous (for review, see Zhimulev, 199213, 1996).The whole question of the relationship between internal repeats and intercalary heterochromatin was dismissed.

D. Correlations in the manifestation of the properties of intercalary heterochromatin Sections VI1,A-C have provided evidence that the numerous bands showing features of centromeric heterochromatin and identified in the metaphase chromosomes are present in polytene chromosomes of various dipteran species. These features include stainability with various dyes, enhanced ability to be involved in ectopic and homologous pairing, the presence of repetitive sequences, late replication, and frequent rearrangement breaks. However, not all the features are inherent in each IH band. Table 16 presents data on 48 regions of polytene chromosomes of Rhynchosciura hoknderi. Twenty-one combinations of intercalary heterochromatin features are distinguished. The combinations differ from each other and the majority of bands in four differential stainings, late replication, repetitive sequences identified in in situ hybridization of DNA from the major fraction, satellites, and nucleotide tracts. There is no information on ectopic pairing, break localization, and chromosomal rearrangements for Rhynchosciura hollaenden'. However, it is apparent how diverse the sets of heterochromatin features are in the various bands.

226

1. F. Zhimulev

Table 16 Occurrence of Bands with Various Features of Heterochromatin in Polytene Chromosomes of Rhynchosciara hollaenderi" In situ hybridization with various DNA fractions and presence of late replicationb

Number of bands with various combinations of differential staining' A

B

C

D

E

F

G

H

I

J

-

MB MB, LR Sat, MB, LR Poly(A), MB, LR Sat, MB, poly(A) Sat, MB, poly(A), LR Sat, MB, POMA), rDNA, MB

8 5 4

1

1

1

1 1 1 1

1 2 2 8

3

1 1

3

1

1

"After Stocker et al. (1978). 'MB, repeats of DNA from the major fraction; LR, late replication; Sat, satellite RNA; poly(A), poly(A) nucleotide tracts. 'Staining combinations: A, C+Q-H-Ag-; B, C+Q+H-Ag-; C,C+Q+H+Ag-; D, C+Q+H+Ag+; E, C+Q-H-Ag+; F, C+Q-H-Ag+; G, C-Q+H+Ag+; H, C-Q+H+Ag-; I, C-Q+H-Ag+; J, C-Q+H-Ag-.

In D. mehogaster, the regions where breaks and ectopic pairing occur are late replicating and greater in number than the break regions (see Figure 82). Likewise, in D. ananassue, the majority of the break regions also replicate late (Duttagupta et al., 1973). It is noteworthy that the polytene chromosomes do not show C+-bands, the number of Q'-bands is not more than six, and, moreover, their manifestation varies from one strain to another (see Table 9, earlier). Information regarding the various IH features of polytene chromosomes in other species is more fragmentary. In Orthocladiw bipunctellus, the "dark blocks" (see Figure 7 1) stain positively for constitutive heterochromatin (C+),while not staining for quinacrine (Q-). Certain C+-bands are involved in ectopic pairing (Michailova and Belcheva, 1982). In Cryptochironomus fridmanae, certain regions are both Q' and C+ while others are either only Q' or only C' (Michailova and Belcheva, 1982). In Ch. thummi, at least three distinct types of intercalary heterochromatin are recognized: (1) strongly Q+-fluorescent, capable of ectopic pairing, and late replicating; (2) less brightly fluorescent (Q+)and capable of ectopic pairing; and (3) not brightly Q+-fluorescent,although involved in ectopic pairing (Badaev et al., 1973). In certain species (Chironomidae, Simuliidae), breaks occur very rarely (Beermann, 1962), while the frequency of late-replicating regions is the same as in other species.

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

227

Despite the great diversity of the types of intercalary heterochromatin, the correlation coefficients between the manifestation of ectopic pairing, breaks, and late replication are high in Dosophila (Table 17). The correlations are also high between late replication and cRNA hybridization (i.e., with repeats), as well as between the strongly synaptic and the late-replicating regions. The correlation coefficients (see Table 17) among some of the features are low presumably because the data are incomplete. How can we explain the consistent differences in the features of the IH regions? It may be assumed that their higher frequencies are due to organizational specificities of chromatin composing the IH bands. According to another assumption, all the DNA sequences equally possess these properties, and their frequencies increased in regions referred to intercalary heterochromatin on the grounds that bands contain more DNA there (see Spofford, 1976). In fact, regions or bands with high DNA content incorporate [3H]thymidine more intensely when labeling is continuous. These very regions are late replicating. However, the correlation often does not hold: replication is started early by some puff groups and, conversely, some late-replicating puffs are thin (Hagele, 1970, 1976; Ananiev and Barsky, 1975; Gubenko, 1976a,b). This raised the question as to whether the properties of chromatin of the IH regions are indeed specific. To obtain a straightforward answer, the frequencies of the manifestation of the IH properties were related to local DNA content along the polytene chromosomes (Zhimulev et al., 1982; Bolshakov et al., 1985a,b). It was found that the correlation between the manifestation of intercalary heterochromatin features and DNA amount in a region is very weak and nonsignificant (P > 0.05) for all the IH features in all the chromosomes. In an exceptional case, the location of 12 families of mobile elements in chromosome 2R conformed to DNA distribution. In the majority of regions that were assigned to the IH by morphological criteria, increased specific frequencies of the manifestation of at least three features were detected. Some drawbacks of the method may be mentioned by way of stipulation. The expected frequencies of the analyzed features were calculated from DNA content in polytene chromosome regions, accepting that the break regions contain less DNA because of underreplication. Therefore, lower expected frequencies were calculated for these features in these regions in comparison with the fully replicated regions. This makes probable overestimates of chromosomal rearrangement and strong homologous pairing presumably occurring at the diploid level, when all the DNA sequences are present in the chromosomes. There was no such overestimate for late replication, breaks, ectopic pairing and hybridization with RNA already taking place at the level of polytene chromosomes. In contrast, even certain fragments of a band can possess IH features, for example, breaks. DNA content was determined in chromosome regions even larg-

~

00 N

Table 17 Correlation Coefficients between the Manifestation Frequenciesof Intercalary Heterochromatin Features in D. mehgmtera

Chromosome

X

2L

Features of intercallaryheterochromatin

Ectopic pairing

Late replication

Breaks Ectopic pairing Late replication Hybridization intensity With cRNA With total RNA Chromosomal rearrangements

0.77

0.61 0.65

BR&

0.72

0.54 0.62

Breaks Ectopic pairing Late replication Hybridization intensity With cRNA With total RNA

0.54

0.40 0.67

BR&

0.83

0.62 0.59

0.78

0.58 0.60

Ectopic pairing Late replication

2R

3L

Ectopic pairing Late replication

3R

Breaks Ectopic pairing Late replication

Hybridization intensity with cRNA

Total RNA

Chromosomal rearrangement

Tight synapsis

0.49 0.39 0.55

0.48 0.43 0.58

0.47 0.55 0.34

0.49 0.50 0.59

0.77

0.40 0.36

0.56 0.64 0.50

0.21 0.27 0.24 0.33 0.53 0.68

0.27 0.41 0.67

0.08*

0.80

0.22 0.30 0.24 0.35

0.17 0.32

0.20

Note. Asterisks indicate correlation coefficient with nonsignificant values (P 2 0.05). “Data from Zhimulev and Kulichkov (1977), Mukhina et d . (1981),and Zhimulev et al. (1982).

0.06 0.16 0.25

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

229

er than a whole band in the lettered map subdivisions containing, as a rule, several bands. This can lead to underestimation of the degree to which many features with restricted location are manifested (Bolshakov et al., 1985a,b). Whatever the stipulation, the results of this analysis justify the conclusion that the unusual properties of the IH regions are reflections of the organizational specificities of chromatin composing the bands. However, features sets vary almost from one band to another. This posed the question: what are the key diagnostic features of IH?At the end of the 193Us, “fragility” arising during the formation of chromosomal rearrangements and ectopic pairing were thought to be the sought-after key characteristics (Kaufmann, 1939; Prokofyeva-Belgovskaya and Khvostova, 1939; Slizynski, 1945), and later papers make reference to ectopic pairing only (Kaufmann and Iddles, 1963); tandem repeats (Zhimulev et al., 1982); mobile elements (E. Sp. Belyaeva et al., 1984a,b); ectopic pairing, breaks, late replication, and repeats (Bolshakov et al., 1985a,b);and late replication (Lairdet al., 1987a). Laird (1989) believes that solely late replication imparts all the properties of heterochromatin to the usual euchromatic genes. The deciphering of the molecular-genetic organization of these loci may be crucial to an understanding the problem of IH. To the present time, there are data on several regions to which genetic functions, or particular repeats, map. It is premature to base conclusions on these data. In fact, only some functions have been determined and we do not know how many are yet to be revealed. The identified regions include the 3C, 10A1-2, 11A6-9, 21D1-2, 25A1-4, 39DE, 56F, 81F, 83E, 87C1-3,89E1-4, IOlF, and 102D regions in Drosophila. is a late-replicating DNA fragment that is thought to be inert because it is located between the w and rst genes, (3C2-3 and 3C5-6), comprising no other genes. It lacks even the 3C4 band (Beermann, 1972). The 3C4-6 region exerts an influence on the course of meiotic recombination (Hawley, 1980).

The 3C region-This

The 10AI-2 bad-This

is characterized by moderately delayed completion of replication (see Figure 82) and occurrence of ectopic pairing (Lefevre, 1976; Kokoza and Zhimulev, 1994). Several genes are located in the band, and several families of repeats in its middle part (Kokoza, 1994; Kokoza et al., 1997).

The J JA6-9 bad-A

site affecting the course of meiotic recombination is located in the band (Hawley, 1980). Males bearing a T( 1;Y) mutation with breakpoints in the 11A6-9 region are all viable and fertile, implying that the rearrangements do not disrupt some essential genetic loci (Lefevre, 1981; Bolshakov and Zhimulev, 1987).

The 21D1-2 band-A tract consisting of polypyrimidines is located in the band (see Section VII,C,6).

230

1. F. Zhimulev

The 25AI-4 band-Sites

of weak points (see Figure 80), ectopic pairing, and late replication are located in the band (Zhimulev et al., 1982).

The 25AI-2 band-The

dumpy gene is located in the distal part of the band (Lindsley and Zimm, 1992).

The 39DE region-This

chromosomal segment is presumably filled completely with a block of tandem repeats of the histone genes (Pardue et al., 1977; Lifton et al., 1978).

The 56F region-This

is a late-replicating region consisting of four bands. A cluster of the 5s ribosomal RNA genes, the genes of five species of tRNA, are located in it (for review, see Ananiev et al., 1978; Zhimulev et ul., 1982).

The bands ofthe 81F, 83E, IOIF, and 1OZD regions-These peats (see Section VI1,B).

contain poly(A) re-

The 87CI-3 band-Replication

is much delayed for this region; it contains a varying number of structural genes encoding the heat shock proteins (Leigh Brown, 1981) and dispersed repeats of two types, including the y-element organized as a tandem repeat in the region. No mutations of essential genes have been identified within the region (Gausz et al., 1979, 1981).

The 89E1-4 band-This

contains bithorax-complex gene (e.g., Ubx). The p3206 clone containing the 5’ exon of this gene maps right to the site of the break (Lamb and Laird, 1987).

A remarkable complexity was found for Rhynchosdaru. Some bands contain strong and moderate repeats together (Machado-Santelli et al., 1979). There are bands staining positively with four differential dyes (C, H, Q, and Ag) and with different combinations of dyes (e.g., C, Q, and Ag but not H, and other combinations) (Stocker et al., 1978). In Chironomus thummi thummi, the intercalary heterochromatic regions are rich in Cla repeats (see Section VII,C,6). A hypothetical scheme may be envisaged for the organization of the intercalary heterochromatin band, reconciling its various properties and offering an explanation for its various types. 1. DNA structure. The facts considered in this section allow the tentative conclusion that either the noncoding sequences of the poly(A) tract type or loci whose activities are needed only during restricted spans of development [e.g., early embryonic development (bithorax) or meiosis (the 3C and 11A6-9) regions] are located in the bands of intercalary heterochromatic regions. Blocks of tandem repeats composed of the 5s rRNA genes and histones are located in two other regions. Their number is excessive, and, for this reason,

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

23 1

the greater part of these genes is transcriptionally inactive and located in compact bands. It cannot be excluded that the complete set of the gene copies is transcribed only during a short span of development-maturation of the egg, for instance. Thus, despite all the diversity, the DNA types composing the IH bands all remain in a profoundly inactive state during the greater part of development. 2. Compaction of chromatin. DNP compaction is the mechanism providing genetic inactivation. It may be effected via special proteins, such as HP1, or some other heterochromatin-specific macromolecules leading to hypercondensation (Laird, 1989). 3. Late replication. Replication is hindered in the tightly compacted bands, and, as a result, its completion is delayed (Laird, 1989; Zhimulev et al., 1989a,b). Variations in the delay produce great diversification of morphological manifestations of intercalary heterochromatin. 4. Breaks. When delayed replication leads to incomplete polytenization, breaks form; when polytenization is complete (e.g., in the male X chromosome of D. melanoguster owing to its looser structure), breaks are not formed. Presumably, because of the structural features of chromatin, replication is complete in the polytene chromosomes of the representatives of the Chironomidae and Simuliidae families, in which breaks do not occur. Treatment of Chironomus larvae with FudR, delaying replication, can give rise to the formation of weak spots in the late-replicating regions (Hagele, 1971). The marked tissues differences in the manifestation of breaks (see Table 10, earlier) can also be explained in terms of developmentally conditioned DNP compaction. It suffices to concede that compaction degree varies in tissues in the course of ontogeny. Change in the manifestation of break frequencies brought about by position effect modifiers is best explained as follows. When the Y chromosome is removed from the genome, the earlier associated excessive “compacting molecules” are redistributed, their amount in the IH regions increases, compaction is enhanced, and break frequency rises (Zhimulev et al., 1989a,b). 5. Ectopic pairing. For unclear reasons the formation of ectopic contacts is correlated with the presence of breaks; thus this feature is also a “derivative” of late replication (Zhimulev et al., 1982). The superposition of yet another feature with varying manifestations on the whole spectrum of IH diversity displayed by bands possessing various features increases band polymorphism. However, the proposed scheme allows us to identify a sequence of events in the formation of the morphological features of the IH, reflecting the essence of the phenomenon and readily recorded in experiments. There appears to be good reason to regard late replication as the key event (Laird,

232

1. F. Zhimulev

1989; Zhimulev et ul., 1989a,b). The preceding fixation of hypercompacted molecules on DNA and tight compaction are events leading to late replication, and it underlies subsequent breakage and ectopic pairing. It will be recalled that rigorous criteria for “late replication” have not, as yet, been established.

E. Variation in the amount of intercalary heterochromatin Metz and Gay (1934) have detected in the polytene chromosomes of Sciuru ocelhris “asymmetrical” or “heterozygous” bands, which represent variations in the thickness of their counterparts in homologous chromosomes (Figure 101). In their view, a deletion occurs in the homolog with the thin band and a duplication in the homolog with the thick one. This polymorphism is of a stable kind, and it has been maintained for more than 50 years in laboratory strains (Perondini and Dessen, 1988). Asymmetry is more readily identified in heterozygotes from crosses between individuals of closely related species or remote populations. In Chironomus thummi thummi X Ch. th. piger hybrids, the banding pattern shown by homologous chromosomes is generally identical, with the difference that many bands are much thicker in Ch. th. thummi than Ch. th. piger. This is at least partly due to the amplification of the Cla repeat in the chromosomes of Ch. th. thummi. When increased in size, the bands acquire properties of heterochromatin, such as C+-banding and late replication (see Section VII,C,6). In populations of Simulium m t i p e s , there frequently occur individuals having heterozygous “additional” bands whose partners are not found in the homologous chromosome (possibly because of their submicroscopic sizes). All the “additional” bands usually replicate late, in which case the chromosomes of heterozygotes (Figure 102) are “asynchronous,” or late replicating; many of the “additional bands” stain positively for C-heterochromatin (Bedo, 1975a, 1977, 1978, 1982a). Data on asynchronous replication, heterozygosity for size, and differential staining (see Table 18) show that the phenomenon is quite widespread. All explanations of the causes of the emergence of asymmetrical bands converge in admitting possible variation in DNA amount in a band due to the formation of deletions or duplications (Metz, 1937a,b, 1947; Carson, 1944; Keyl, 1957, 1964a,b, 1966), or to multiple amplification of the original sequence (Pavan and da Cunha, 1969; Gabrusewycz-Garcia, 1971). It has also been suggested that the chromomere (band) may be a complex structure; for example, it may comprise gene(s) and additional DNA sequence(s) functionally not associated with it. This sequence can be amplified or removed by a deletion. Band size increases in the former and decreases in the latter case, although the band is not lost completely (Zhimulev et ul., 1981; Perondini and Dessen, 1988).

Figure 101. “Asymmetrical” bands in Sciara ocelhris ( a x ) , Acricotopus lucidus (d), and Chironomus thummi thummi x Ch. th. piger (e) hybrids. Arrows in a-c indicate the thick (a) and thin (c) bands in the homozygote and the same in the heterozygote (b). (a-c) after Perondini (1979); (d) reprinted by permission from Panitz (1965); ( e ) after Key1 (1965a).

234

I. F. Zhimulev

Figure 102. “Asynchronous”termination of replication in the polytene chromosomes of Simulium ornatipes. Arrows indicate regions of asynchronously terminating replication; NO, nucleolar organizer; C, centromere. After Bedo (1982a).

Table 18 Occurrence of Bands with Varying Manifestations of Features of Heterochromatin in Diptera

Family, species

Characterization

References

Agromyridae Melanagrmnyza tropicalis

Variation in band thickness

A. B. da Cunha in Perondini and Dessen (1988)

Calliphoridae Calliphora eryrhrocephala

Same as Agromyzidae

Ribbert (1967)

Chironomidae Acricotopus lucidus

Same as Agromyzidae

Chironomw obtusidens

Same as Agromyzidae

Mechelke (1960), Beermann (1962), Panitz (1965) Key1 (1961,1964b)

Ch. plumosus

Same as Agromyzidae

Key1 (1964b, 1965b)

Ch. plumosus X Ch. vnncouueri hybrids

Large differences in the number of intercalary C+-bands

Michailova (1987a)

Ch. species

Variation of band thickness

Goldschmidt (1942) (continues)

235

Poiytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 18 (Continued)

Family, species

Characterization

References

Ch. tentans

Variation of the thickness of homologous bands in populations

Beermann (1952), Acton (1959)

Chironomw thummi thummi

Variaton of thickness of the C l c band and delay of DNA replication in the homolog with thicker band Variation of band thickness Heterozygosity for different thickness of the ll-B5p band occurs in 8.9-33.3% of larvae in different populations Heterozygosity for labeling intensity between two chromosome homologs after in situ hybridization with DNA of 5s rRNA genes Interspecific comparison revealed variation in thickness of homologous bands

Hagele (1970)

Ch. thummi in comparison with Ch. parathummi and Ch. obtusidem Ch. thummi thummi X Ch. th. piger hybrids

Beermann (1973) Kiknadze er al. (1988)

Baumlein and Wobus (1976)

Key1 (1961)

See section XVII,F,3; in the D3C band of thummi chromosome there are considerably more copies of the TF BJ mobile element inserting into the histone genes HI-H3 than in the homologous band of piger chromosome

Hankeln and Schmidt ( 1990)

Considerable increase in the size of one of the darkly stained bands

Kiknadze et al. (1990)

Interspecific differences in the amount in late-replicating DNA

Tiepolo et al. (1974)

Polymorphism for thickness and stainability of homologous bands

Munstermann and Marchi (1986)

X-located band amplifies its DNA content in males in comparison with females

Bicudo (1983)

D. grimshawi

Variation in band thickness of the third chromosome in various populations

Stuart etal. (1981)

D. mehogaster

Variation in the amount of late-replicating material compared to asynaptic homologs in 86C-89F and 94A-98A regions; variation in amount of material hybridizing in situ with RNA synthesized in cell culture in the 50D, 51D, 61C, 83DE, and 97A regions of asynaptic homologs

Hagele and Kalisch (1978), Gvozdev et al. (1980a,b)

Glyptotendrpes paripes

Culicidae Anopheles anopurvus X

A. lnbranhiae hybrids Sabethes cyaneus Drosophilidae Drosophiln arizonensis

(continues)

236

I. F. Zhlmulev

Table 18 (Continued) ~

Family, species

References

Characterization

di Pascuale Paladin0 et al. (1988) authors' view: there is an additional band in another homolog) See Table 9 earlier Variation in number and location of Q+-band Differences in thickness of homologous bands Pacau (1935)

Loss of the 25A1-4 band in tu-pb stock (the

Drosophila of the melanogaster group

D. mefanogaster X D. simulans hybrids D. flllsuta albomicana D. wirilis and D. hicola Drosophila of the wirilis group

Glossinidae Glossina msitans cenrralis, Glossina morsitans X G. m. morsitans X G. m. subnwrsitanshybrids

Variation in the amount of heterochromatin at the base of the microchromosome

Hagele and Ranganath (1982)

Differencesin thickness of some homologous Patterson and Stone (1952) bands Sinibaldi and Barr Four of the 12 studied species have Q+-bands (1979) in the arms of polytene chromosomes:a toromere on the sixth chromosome in D. lummei, a band in the proximal third of the X chromosome in D. wowororurn, a band at the base of the fourth chromosome in D. wirilis, and a band in the Y i-j in D. flawomontana; no Q+-bands were found in the euchromatin regions of the chromosomes in D. nowamexicana, D. americana texana, D. a. americana, D. littoralis, D. ezoana, D. montana, D. borealis, or D. kicola Variation in thickness of homologous chromosomes

Southern and Pel1 (1973)

Psychodidae Telmatoscopw albipunctatus

Heterozygosity for band thickness

Troiano (1975)

Te4matoscopw sp.

Same as T. albipunctatus

Amabis and Simoes (1972)

Sarcophagidae Parasarcophaga m'sera Sciaridae Rhynchosciara americana

Variation in the presence of one of the C+-bands

a+-, Tewari etal.(1983)

A heterozygous band in chromosome A in larvae of the LII mutant stock

Terra et al. (1976)

Sciara impatiens

Variation in band thickness

Sciara

Same as S. imptiens

Carson (1944) McCarthy (1945a,b) (continues)

23 7

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 18 (Continued) Family, species Sciara ocellaris

Characterization 24 cases of heterozygosity for hand thickness described

References Metz and Gay (1934), Metz (1935, 1937a,b, 1939, 1941, 1947). Metz and Lawrence (1938), Rohm (1947), Pavan and Perondini (1967), Pavan et d.(1968), Perondini (1979), Perondini et al. (1983), Perondini andOtto (1991) Perondini and Dessen ( 1988)

19 cases of size variation of homologous bands described; Occurrence frequency in box population is 4.95 heterozygous bands per larva In situ hybridizing of cRNA transcribed from R. A Eckhardt, E. M. B. Dessen, and A. L. P the sciarid genome was detected in the Perondini in Peronthickened bands under conditions allowing dini and Dessen the authors to identify only repeats (1988) Variation in band size

Gabrusewycz-Garcia (1971)

Cnephia

Same as S. pauciseta

Procunier (1975a)

Eusimulium aureum

Same as S . pauciseta

Dunbar (1958, 1959)

E. nyophilum, E. vernuum

Same as S. pauciseta

Hunter (1987)

odagmiamta

Same as S. pauciseta

Shcherbakov (1968)

Prosimuliumpeaosum

Same as S. pauciseta

Ralcheva (1974)

Simulium nolleri

Same as S. pauciseta Numerous cases of polymorphism for occurrence and thickness of C+-hands Differences in amount of late-replicating DNA in identical loci of homologous chromosomes

Shcherbakov (1968)

Simulium

Variation in sizes of bands

Pastemak (1964), Vajime and Dunhar (1975)

S . pictipes

Heterozygosity of hand size in the 3 1C4 and 63A4 regions; thin band is intensely Q+-stained, thick band fluoresces weakly

Bedo (1975b)

SciaTa pauciseta

Simuliidae

S. m t i p e s

Bedo (1975a, 1977, 1978) Bedo (1982a)

(continues)

238

I. F. Zhfmulev

Table 18 (Continued) Family, species

Characterization

S . vittatum

Variations in band size

Twinnia nova

Same as S. vittatum

Tephritidae Ceratitis capitata

The heterozygosity for Q+-band detected in 1 of 5 individuals

References Rothfels and Dunbar (1953), Rothfels (1979) Rothfels and Freeman (1966) Bedo (1986)

Vlll. TELOMERIC HETEROCHROMATIN A. The telomere concept Based on study of the formation of radiation-induced chromosomal rearrangements, Muller came to important conclusions regarding the behavior of the distal ends of the chromosomes: (1) they never unite with the chromosome ends resulting from induced breaks, which can unite only with the similary broken ends of other chromosomes; and (2) the most distal fragments are virtually never transposed by rearrangements to occupy an interstitial position in the chromosome. To account for these facts, Muller suggested that the gene on the free end of the chromosome should form a peculiar self-perpetuating structure, the “telomere,” which he thought was unipolar (i.e., establishing contact with a single gene only, lying proximally, closer to the centromere) (Muller, 1932, 193813, 1941; Muller and Herskowitz, 1954). The significance of the property of unipolarity has been treated at length by McClintock. In her hypothesis, the inert behavior of the telomere in union with the other chromosome regions was assigned an exclusive role: it prevented their sticking end-to-end, thereby promoting preservation of chromosome individuality and of the karyotype as a whole (McClintock, 1938,1939,1940,1942). Thus, the telomere concept came into being and was elaborated (see also White, 1954,1973; Blackbum and Szostak, 1984; Prokofyeva-Belgovskaya,1986; Hilliker and Appels, 1989; Zakian et al., 1990; Biessmann and Mason, 1992, 1994; Blackbum, 1994; Pardue, 1994, 1995; Blackbum and Greider, 1995; Gall, 1995; Mason and Biessmann, 1995; Biessmann et al., 1996; Greider and Blackburn, 1996; Pardue et al., 1996). Replication completion at the ends of linear DNA molecules posed challenging problems for molecular biology in the 1970s (Figure 103). Single-stranded gaps remain in the newly synthesized strand after removal of RNA primers dur-

Polytene Chromosomes, Hetetochromatin, and Position Effect Variegation

239

-fly7 Figure 103. * Schemes of the replication of the end of the linear DNA molecule (facing the right side of the figure). Formation of -1 gaps after removal of RNA primers (sections with wavy

4

lines). Internal gaps are filled with the involvement of the

3’ ends (arrows) of newly synthesized DNA fragments as primers. The terminal gap o n the 5’ end of the newly synthesized DNA strand has no primer, and it is therefore not filled. After Dancis and Holmquist (1977, 1979).

ing replication. They are filled with DNA polymerase using the 3’ ends of the previously synthesized DNA fragments as primers. Since there is no primer for the gap at the extreme end, the newly synthesized strand proves to be a few nucleotides shorter than the initial strand. As a consequence, the DNA molecules keep becoming shorter with each replication cycle (Watson, 1972; Olovnikov, 1973).An obvious solution to this problem would be the formation of a special structure at the end of the linear DNA molecule (the chromosome), for example, an added peculiar telomeric DNA or fused ends of the molecules (or the chromosomes) making a junction so as to give rise to palindromes and hairpins, among other structures (Dancis and Holmquist, 1977, 1979; Hastie et al., 1990; Murray,

1990). The results of high-resolution mapping of rearrangements using polytene chromosomes and a better understanding of the structure of chromosome ends provided by molecular-genetic methods stimulated further development of the telomere concept. It was shown that (1) the chromosomes possibly can manage without telomeres and (2) telomeres of the chromosomes unite in some cases. Finally the structure of telomeric DNA was deciphered. According to the telomere concept, all the chromosomal rearrangement breakpoints lie proximal to the end of the chromosome, with the telomeric chromosome regions remaining at a position distal to the breakpoint. However, irradiation was found to induce terminal chromosomal rearrangements or they were found in populations of various dipteran species (Kaufmann, 1936;Kikkawa, 1938;Helfer, 1941;Dobzhansky and Dteyfus, 1943;Carson, 1944;Lea and Catcheside, 1945; McCarthy, 1945a,b; Pavan, 194613; Carson and Stalker, 1947; White, 1954; Blaylock

240

1. F. Zhimulev

Figure 104. Variations in the telomere morphology of chromosomes 2L ( a 4 and 2R (e-h) in DrosoQhilnu m ~ s s a e(a . and e) Homozygotes for deletions of telomeres. (b, c, f, and g) Heterozygotes for normal and mutant morphotypes. (d and h) Homozygotes for presence of telomeres. After Kikkawa (1938).

and Koehler, 1969), specifically, those in which telomere material distal to breakpoints has not been identified. Furthermore, cases are now widely known in which the so-called terminal deletions remove one or several polytene chromosome bands (Figures 104 and 105) at the extreme tip of the chromosome (Table 19). The results, however, are hardly explicable. First, as White (1954) pointed out, even when using polytene chromosomes in a real experiment, it may be difficult to make the distinction as to whether the end region of the chromosome has been translocated to a new site, or a break has occurred in close proximity to the telomere and a part of the chromosome (too small to be seen in the microscope) still remains distal to the breakpoint, joining with the transposed nascent neighboring material (Kossikov and Muller, 1935; White, 1954). Second, the telomeric structures may possibly be of nonpolytene nature and escape detection (P. A. Roberts, 1975). Third, according to certain data, the rearrangements are terminal deletion mimics: even when the most conspicuous terminal bands at the extreme end of the chromosome are lost, cloudy greyish material, which is present in the normal chromosomes, is still noticeable. This fact may indicate that the deletions are, indeed, subterminal (P. A. Roberts, 1975). It is not known whether analysis of telomeric material was just as thorough in all the cases listed in Table 19, and it is also not clear what the structure of the material might have been like (see Section VIl1,C). In any event, at least some of the material has the appearance of telomeric deletions (Table 19). The presence of viable telomeric deletions of a telomere from a Drosophi-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

241

Figure 105. Variability of telomere morphology of the polytene chromosomes of D. melanogaster. After Lefevre (1976).

la chromosome was demonstrated in molecular biological studies (Mason et al., 1984; Biessmann and Mason, 1988; Levis, 1989; Wurgler, 1990; Karpen and Spradling, 1992; McKee et al., 1992; Tower et al., 1993; Zhang and Spradling, 1993). When the P-element transposon was induced to transpose from the telomeric region of chromosome 3R of Drosophifu, terminal deletions removing the most distal DNA sequences could be recovered. In subsequent generations, strains with such deletions progressively lost terminal DNA fragments (from the telomere to the centromere) at a rate of 50-100 (75 on average) base pairs per fly generation (Levis et d., 1987;Levis, 1989; Biessmann et al., 1996). In other studies, terminal deletions of the X chromosome were generated by irradiating a “mutator” strain of Drosoghila. In such cases, telomeric DNA sequences were also progressively lost (Figure 106) at a rate of 75 bp per genera-

242

1. F. Zhirnulev

Figure 105. Continued

tion (Biessmann and Mason, 1988; Biessmann et al., 1990a). It is believed that the size of the lost fragment correlates with that of the octonucleotide RNA primer (Kitani et al., 1984) initiating the synthesis of Okazaki fragments during DNA replication (Biessmann et al., 1990a). Despite the absence of telomeric DNA in strains with terminal deletions, the chromosomes do not stick together to form junctions; that is, loss of the telomeres in such cases does not destabilize the karyotype. Numerous instances are now known in which the telomeric tips of the chromosomes ectopically pair with the regions of intercalary heterochromatin and pericentric heterochromatin with the telomeres of the other chromosomes. These ectopic pairings are not at variance with the telomere concept because there is no proof that specifically DNA is involved in these contacts. Analysis of the organization of protozoan and infusorian genomes allowed the first isolations and deciphering of telomeric DNA. In the maturing macronucleus of many infusorian species, polytene chromosomes fragmented into thousands of single genes, with each reiterated about 1000 times due to polyteny. Telomeric DNA sequences were added at either side of each fragment and, as a consequence, the telomeric fragment in the mature macronucleus becomes reit-

243

Polytene Chromosomes, Heterochromatln, and Position Effect Variegation Table 19. Occurrence of Terminal Deletions in Polytene Chromosomes of Diptera Family, species Drosophilidae D. ananassae

Localization, brief description Deletions in chromosomes 2L, 2R, and 3R; no effect of deletions on viability and morphophysiological traits identified -

D. nrizonensis

-

D. melanogarter

The X chromosome; terminal deletion Df(1)260-1 has no effect on imago viability and morphology; fertility is somewhat decreased in females Deleted material partly restored after several years Deficiences in the X chromosome induced by radiation The X chromosome; population and strain differences The X chromosome; radiation induced Deletions in chromosomes X, 2L, 2R, and 3R arise, possibly, during the making of squashed preparations The X chromosome; deficiency in the distal half of the 1A region is, presumably, related to fusion of bands The X chromosome, terminal deficiencies, arisen during irradiation of “mutator strain”

Large differences in banding pattern at the ends of chromosomes X, 2L, 2R, 3L, 3R, and 4 in different strains In T-32 strain, resistant to high (32°C) temperature, unusually long telomeres in 2L and X chromosomes; in the “long” telomeres more copies of DNA of the telomeric repeat (the Dm 665 clone)

D. nasuta Related species: D. disjuncdD. bosnychaJD.

Deletions of 1-3 bands at the ends of chromosomes ZL, 2R, and 4 Comparison of the chromosomes of these species revealed small additional bands in the sixth chromosome of D. disjuncta

References Kikkawa (1938)

Dobzhansky and Dreyfus (1943) Bicudo (1981) Dernerec and Hoover (1936)

Roberts (1969b, 1976) Alikhanyan (1937) Prokofyeva-Belgovskaya (1938, 1986) Sutton (1940b) Goldschmidt and Kodani ( 1943) Kodani (1947)

Green and Lefevre (1972), Mason and Strobel (1982), Mason et al. (1984, 1986), Beissmann et cd (1990b; 1996) Lefevre (1976)

Danilevskaya and Lapta (1990, 1991)

Kumar and Gupta (1986) Aheam and Baimai (1987)

affinidisjuncta

(continues)

244

I. F. Zhimulev

Table 19. (Continued) Family, species

Localization,brief description

References

Related species: Comparison of chromosomesrevealed deficiencies in terminal bands in 2L and 2R D. melanogasterl chromosomesof D. melanogaster and in 3L of D. simulans D. simulans Simuliidae

Ashbumer (1969)

OdagmiamOnti-

Chubareva and Ralcheva (1974)

-

cola

Sciaridae Sciara impatiens

In populations 9 cases of terminal deletions revealed in autosomes;deletions not lethal

Carson (1944)

~

Note. Dash indicates that species is only mentioned in reference(s).

erated several million times so that it could be readily isolated (for detailed review, see Blackburn and Gall, 1978; Gall, 1995). In Stylonychia and Oxycricha, the structure of terminal sequences consisting of tandem repeats can be written

5 -C4A4C4A,C4-3 3 -G4T4G4T4G4T4G4T4G4-5 ’

L I

0

I

6

I

I

I2

16

1

24 Monlhs

1

I

1

30

36

42

Figure 106. Decrease in the size of the terminal DNA fragment in the X chromosome of Drosophila in strains with telomere deletions RT276, RT473, RT624, and RT733. Abscissa, time from the beginning of observation; ordinate, the size of the end PstI fragment in the chromosomeswith deletions. After Biessmann and Mason (1988).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

245

It is seen that the main repetitive unit is the octonucleotide T4G4on the DNA strand, whose 3’ terminus is the end of the chromosome (the lower line in the scheme) (see Stoll et al., 1991, 1993, for rewiew). In ascarids (Ascaris lumbncoides), chromosome ends become “unprotected” as a result of diminution of heterochromatic fragments. The telomeric sequence TTAGGC appears on them presumably owing to the activity of telomerase (Muller et al., 1991). Information concerning telomeric repeats in other organisms is summarized in Table 20. The telomeric repeat has the structure 5’-T,14)/AG(l-8)-3’. The number of such blocks at each end of the chromosome corresponds to the balance established between growth rate and copy number loss. The structure of the overhanging 3’ end of the DNA loop enriched with guanine and thymidine is peculiar. Since this single-stranded DNA is not digested with nucleases, including DNase S1, it was often assumed that it is packaged in a complex manner into a double-stranded structure akin to the hairpin and node, with the majority of models implying non-Watson-Crick interaction of bases, mainly guanine with guanine. The structural variants of the hairpin at the 3’ termini of DNA strands are presented in Figure 107. In the triplex model of the telomere, the chromosome ends form an intramolecular pyrimidine-purine-purine triplex: the single-stranded G-rich overhang folds back, lying into the major groove of the terminal duplex forming the CGG base triads (Veselkov et al., 1993). There are data indicating that the overhang of the 3’ telomeric terminus is bound to specific proteins. Proteins with molecular masses of %(a) and 41 (fl) kDa, which protect the end of the DNA molecule from digestion with exonuclease Bal-31, DNaseI and micrococcal nuclease, were isolated from Oxytrichu nova macronuclei DNA. It was concluded that the proteins are specific precisely to the single-stranded 3’ repeat T4G, (Gottschling and Zakian, 1986; review: Fang and Cech, 1995). The two subunits are very tightly bound to both DNA and each other (Price and Cech, 1989; Zakian, 1989; Fang and Cech, 1993a-c). The proteins bind either to the two T4G4repeats at the 3’ terminus or to the two internal repeats (Raghuraman and Cech, 1989). A 85- to 130-bp DNA is included in the DNA-protein complexes from Euplotes crassus (Price, 1990). The protein encoded by the RAP1 gene in Sacchuromyces cerevisiae binds in viao to several sequences, including the repeat T(G)2-3(TG)l-6 repeat found in yeast telomeres (Conrad et al., 1990). The terminal tracts of C,_,A repeats assume a nonnucleosomal chromatin structure (Wright et al., 1992). There is as yet no unified idea of what the mechanism of telomere formation at the molecular ends may be like. It has been suggested that the telomeres may result from recombination between the tips of the chromosomes (Plu-

246

1. F. Zhimulev

Table 20. Nucleotide Sequences in Tandem Telomeric Repeats in Different Organisms

Organisms

Sequence

References

Infusoria

Tenahymena GLrWOtW

Paramecium

Stylonychia

TTGGGG TTGGGG TTGGGG alternates, TTTGGG TT(T/G)GGG TTTTGGGG

Oxynicha, Euplotes

Review: Blackburn (1984) Review: Blackburn (1990)

J. Forney and E. H. Blackburn in Blackburn and Karrer (1986) Blackburn (1990) Review: Blackhum (1984, 1990), Stoll et d. (1991,1993) Blackbum and Szostak (1984), Boeke (1990), Raghuraman and Cech (1990)

Coccids

FtaSmodium Flagellates Trypanosoma, Leptomonas,

Blackbum (1990) TTAGGG

Leishmania Crithidia

Blackbum (1984), Blackhum and Challoner (1984), van der Ploeg et al. (1984), Blackbum (1990)

TTGGG

Fungi

Physarum Dictyostelium

Blackburn (1984, 1990)

Saccharomycescerevisiae

Review: Henderson (1995)

Schizosaccharomycespombe

Blackburn (1990)

Neurospora

Blackburn (1990)

Blackburn (1984, 1990)

Algae

Chhmydomonas

TMTAGGG

Blackburn (1990)

TTTAGGG

Richards and Ausubel(1988), Blackburn (1990)

Ascaris lumbricoides

TTACGG

Mulleret al. (1991)

Insects

TTAGG

Biessmann and Mason (1994)

TTTAGGG

Riethman et al. (1989), Williamson et al. (1989), Wilkie et al. (19901, Blackbum (1990)

Higher plants

Arabidopsis Roundworms

Mammals

Homo rapiens

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

247

Figure 107. Various models of the organization of telomeric DNA. (a) The telomere hairpin is stabilized by the interaction of CC' and A.A+,while the guanine-rich strand is virtually not involved in the interaction. (b and c) Models of terminal oligonucleotides in Oxytricha (b) and Temahyrnena (c) with two (b) and three (c) G-quartet nucleotides that are opposite each other in a horizontal plane. (d) A hairpin formed of two DNA strands. (e) Variants of non-WatsonCrick interaction of guanine in the telomeric repeat. (a) after Lyamichev et al. (1989); (b and c) after Williamson et al. (1989); (d) after Sundquist and Klug (1989); (e) after Henderson et al. (1987).

248

1. F. Zhimulev

e)

T T GoG G- G G*G G- G T* T T- T G- G G- G G* G G * G, /T

G

T

G.

G G. G G*

T* G T. T G* T G -G G* G G- G T- G T '31

3'

5'

/

....

G G- T G- G C *G T- G TG= GGG*G T* G T.

T G

!

/

5'

....

T GGGG-T-1-GGGG T GGGG T

T GGGG-T-T $1

3'

T-T-

\

3'

....

GGGG,

-3'

5' Figure 107. Continued

ta and Zakian, 1989) or from a de now synthesis with the involvement of the enzyme of terminal transferase (telomerase). Besides the protein component, the enzyme has an RNA molecule approximately 150 bp long containing 1.5 copies of the corresponding telomeric repeat. The RNA component can become the template for completion of synthesis by telomeric DNA (Blackbum, 1984, 1990, 1991; Blackbum and Karrer, 1986; Sprangler et al., 1988; Lamond, 1989; Shippen-Lentz and Blackburn, 1989,1990; Boeke, 1990; Greider, 1995). A hypothetical scheme for the synthesis and formation of hairpins from newly synthesized telomeric DNA is given in Figure 108. The scheme depicts DNA crawling like a caterpillar along the telomerase molecule. Numerous models of developmentally programmed healing of chromosomes, i.e., adding the telomeres, are considered by Blackbum (1995). In the higher eukaryotes, there were several cases in which chromosome regions lacking telomeric DNA have acquired it. Breaks were induced and rings opened by irradiation of the ring X chromosome of Drosophila (Lea and Catcheside, 1945). The formation of new telomeric structures at either side of breaks is expected to provide normal functioning of such chromosomes. In fact, when ring chromosome C(l ) A in the 13E region is opened (Figure 109), material homologous to a telomeric repeat (see later), which hybridizes in situ, appears at either side of the breakpoint (Traverse and Pardue, 1988; Pardue, 1990). Blocks of telomeric repeats can be added to the chromosomes having terminal deletions in Drosophila. In this case, loss of 75 bp of terminal DNA per generation ceases (Biessmann et d., 1990b).

B. Telomere-associated sequences In addition to short, simple repeats located at the extreme ends of the chromosomes and presumably required for replication completion, blocks of larger tandemly repeated DNA sequences lying closer to the terminal repeat have been identified (for review, see Blackburn and Szostak, 1984; Biessmann and Mason, 1992; Henderson, 1995).

Polytene Chromosomes, Heterochromatln, and Position Effect Variegation

249

1. Bind primer

2. Polymerize

3. Translocation and Rehybridization

tt t t 9 9

. ‘3

5-

-

I

Figure 108. Scheme of the synthesis of telomeric DNA. ( 1) Telomeric primer attaches to telomerase due to WatsonCrick interaction of N42-47 nucleotides from RNA included in telomerase. (2) Elongation of DNA strand to N35 nucleotide. (3) Formation of hairpin and GG base interaction. After Blackburn (1990).

The regions of telomere-associated sequences (TAS) vary in length from chromosome to chromosome occupying some 100 kb in Plasmodium up to 2000 kb in humans and may account for about 12-18% of total DNA in Secak cereak (reviews: Biessmann and Mason, 1992; Henderson, 1995). In the human chromosomes, 2 to 20 kb of the tandemly repeated sequences AGGGTT are interspersed with various repeats. A family of unrelated repetitious motifs lies proximally (for review, see Wilkie et al., 1990)). In rye, the

250

1. F. Zhimulev

Figure 109. Appearance of a subtelomeric DNA repeat at the ends (the 13E region) of an opened ring chromosome [C(l)A] of D. melanogaster. C, chromocenter. After Traverse and Pardue (1988) and Pardue (1990).

repetitive units are 120, 480, and 610 bp in size (Jones and Flavell, 1983). A scheme for the organization of telomeric repeats in yeast is shown in Figure 110. In Stylonichia lemnae the telomeric repeats are contained in a very conserved element consisting of two 2-kb direct repeats flanking a 2.6-kb sequence. The 18 mer (C,A4C4A4C,) of the telomeric sequence is immediately adjacent to one of the repeats (Stoll et al., 1991, 1993). The data on the organization of telomeric repeats in organisms with

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

25 1

TE L

CEN c ~ 1types -

x ;

u

Y

I

I

I

X-type

1A

AI A

NAn

X

0.3-3.7kb

Figure 110. Two types of chromosomes of the yeast Saccharomyces cerevisisea differing in the structure of the telomeric repeat, which may consist of the X and Y sequences or of only the X. Black triangles, terminal telomeric repeats; A, autonomously replicating sequence. After Chan and Tye (1983) and Jager and Philippsen (1989).

polytene chromosomes presented in Table 21 allow to make the following conclusions. 1. The DNA located on the ends of the chromosomes can comprise sequnces of various types: blocks of telomeric repeats, which are located only in the telomeres or in the telomeres and pericentromeric heterochromatin. 2. There presumably are several families of telomeric repeats interspersed with each other. A survey of the distribution of the location sites of the TTAGGG sequence for 100 vertebrate species revealed that, besides the telomeric sites, this sequence was most frequently located in the pericentric regions of the chromosomes, and in heterochromatin in many species (Meyne et al., 1990). 3. The representatives of this single family are frequently arranged in tandem arrays (Figure 1 11). Unusual telomeres were found in Drosophila melanogaster. “It is ironic . . . that the first study on telomere behavior-the one that led to the basic concept of a cap on the end of the chromosome-was made on Drosophila, an organism that, in fact, has highly unusual telomeres” (Gall, 1995, p. 3). While the telomeres of most eukaryotes contain short, simple, and highly conserved repeats, Drosophila chromosomes carry retrotransposable elements of two families, HeT and TART. A heterogeneous family of complex repetitive sequences known as H e T was cloned from Drosophila melanogaster and shown to hybridize in situ to all the tips

Table 21. Location of Various DNA Sequences in the Tebmeric Regions of the Polytene Chromosomes ln N

w

DNA type, Species

Copy number

sue

Other data

Anopheles gambiae

820 bp minisatellite

-

Located exclusivelyat the tip of 2L

Anopheksmess~

-

-

In hybridization of total genomic DNA with polytene chromosomes, some increased labeling detected in telomeres

Chirmomus

Fragment, 340 bp

-300 kb in each telo-

Identified in telomeres and ectopic strands between them; transcribed in RNA of -20 kb Four subfamiliesof repeats (MI, D1, D2, and D3) revealed; they are identical in 90% of length and greatly differ in the remaining 10%

mere (1.2% of the genome)

pallidiuittatus

Ch. tentans

Fragment, 350 bp

-

Ch. thummi pigm

176 bp from the Ch.th. hummi genome (Carmona et d., 1985)

-

Ch. th. thummi

176 bp

- loo0 copies in telomere

Drosophila melanogmter

Fragment, 1.4-3.0 kb

-

References Graziosi et d.(1990), Biessmann et d.(1996) Stegniy and Sharakhova (1990) Saiga and Edstrom (1985)

Cohn and Edstrom (1991, 1992a,b)

Repeat consists of two subunits of 185 and 165 bp Detected in all the telomeres of Ch. th.thummi X Ch. th. pigo hybrids; transcribed in the heat shock BR (TBRIV)

Nielsen et d.(199O),Nielsen and Edstrom (1993) Morcillo et d.(1988)

Tandem repeat, shares homology with RNA from the heat shock BRIII; has two hybridization sites with intercalary regions of the second chromosome Not identified in D. melanogmer and Ch. pallrdruinatus Identified in telomeres and ectopic contacts between them

Carmona et d.(1985)

Saiga and Edstrom (19851, Carmona et d. (1985) Rubin (1978), E. Strobel, A. J. Flavell, H. Beck, B. Backner and G. M. Rubin in Spradling and Rubin (1981)

wl N W

Satellite 1.688

-

Fragment, 12.6 kb “He-T D N A

-

Fragment, 2.3 kb

-

Dm 665, 2.4 kb

2OC-250 per haploid set (60-80 copies per telomere)

Satellite 1.672

-

0.8 kb

-

Besides centromeric regions, it is located in the telomere of the X chromosome Fragments included in this clone repeated nontandemly in the genome, mapped in telomeres, bases of chromosome arms, in the Y chromosome Hybridizes with DNA of D. simufam, D. mauritiana,D. ewcta, D. teissiri, D. yucuba, D. miranda, but not D. virilis Located in the 44D region, chromocenter and tips of the chromosomes in proportions: X:2L:2R:3L:3R = 1:3.4:1.9:0:2.7;not identified in the fourth chromosome; transcribed in poly(A) RNA DNA fragment including telomeres 2L. 2R, the fourth, and X chromosomes possesses activity of autonomously replicating sequences (ARS) of yeast; the sequence in cell culture is virtually not transcribed.; copies occur in the Y chromosome The clone contains a 650-bp region homologous to the SteUate Hybridization with the chromosomes of D. simulans is weak, no hybridization with D. virilis; Polymorphism for hybridization intensity in strains and different cells of the same gland In some fragments of Dm 665 clone, insertions of copia and hoppel identified In O r e g o n 4 strain mapped to the lOOF and 81F regions Repeat in the vicinity of the Kruppel gene, mapped to the telomeric region of chromosome 2R

Peacock et al. (1978) Young et al. (1983), Traverse and Pardue (1989). Pardue, (1990), Valgeirsdottir et al. (1990) Young et al. ( 1983)

Renkawitz-Pohl and Bialojan (1984)

Danilevskaya et al. (1984, 1990, 1991). Gragerov et al. (1988a,b)

Danilevskaya et al. (1991) Leibovitch et al. (1990, 1991)

Kurenova et al. (1990) Steffensen (1985) Preiss et al. (1985)

(continues)

Table 21. (Continued) m N

DNA type,

P

Species

Copy number

size

-

-

1176 bp (element 1360)

Not less than 50

References

D m K172

-

1.8 kb, telomereassociated sequence (TAS) in Dp I 187 minisatellites in 3R subtelomeres minisatellites in 2L subtelomeres

-

After treatment of squash preparation with resnictase Hae 111, ethidium bromide stains the telomeres of chromosomesX, 2R, and 3R Mapped at the rips of the X, 2R, and 3L chromosomes, at the bases of arms, in the fourth and X chromosomes “Element 1360” is homologous to the hoppel element Clone C18 obtained bv microdissection of the telomere of one of the chromosomeshybridizes with many regions of the genome X chromosome: Subtelomericregion, the 10A1-2 band and bands in region 9. No labeling in chromocenter. Similar labeling was found in D. s i m h X chromosome: Subtelomeric region, the 10A1-2 band and bands in regions 7.8, 9, 11, 12, 13 The TAS contains 173-bpsubrepeats

-

-

Levis et al. (1993)

-

-

Walter et ol. (1995)

-

-

In most stocks 2L minisatellite is confined to the 2L, in some stocks it also hybridizes with the 3L tip

Mechler et al. (1985), Walter et al. (1995)

Fragment, 4.4 kb

20 in females, 90 in males

Hybridization intensity with telomeres of different chromomosomesvaries widely, from

Nauber and Steinemann (1983), Steinemann

-

D m 191

D. mirnnda

Other data

Mezzanotte (1986)

Kholodilov et al. (1987, 1988) Kurenova et al. (1990) Ponelies et al. (1989)

Kokoza ( 1994)

Kokoza (1994)

Karpen and Spradling (1992), Spradling (1994)

D. subobscura

D. nistis

-

Fragment, 181 bp

-82,000

D. virilis

-

Rhynchosciara angelae R. hollaenderi

-

Sciara coprophifa

Satellite 11

Stylonychra lemnae (Ciliate)

Telomeric repeat

1O W 1 500

h,

L4l L4l

Note. Dash indicates that species is only mentioned in reference(s).

hardly discernible labeling in X'L to extremely strong in the fifth chromosome; heavy labeling of the male Y chromosome; does not hybridize with the chromosomes of D. mefanogaster

(1984), Steinemann and Nauber (1986)

-300 location sites in polytene chromosomes including the chromocenter and telomeres; hybridizes with the chromosomes of D. guanche and D. d r e n s i s , but less intensely (by 2-10 times) Telomeres of chromosomes3 and 4 contain a repeat without Alu 1 restriction site

Felger and Sperlich (1989)

Mezzanotte et al. (1987)

In telomeres of all the chromosomesexept the Y;hybridizes with DNA from D. ambigua, does not hybridize with DNA of other closely related species and D. mehogaster

Bachmann et d.(1990)

The aNM4 DNA fragment from the D.novamexicnna genome hybridized with chromocenter, dispersed throughout the chromosomes and in telomeres of chromosomes 1.3, 4, and 6 of D. win'lis

Sayanova etal. (1987)

Telomere ends hybridize in situ with ['HIeoly(U) Telomere of chromosome B hybridizes with cRNA transcribed from satellite DNA

Jones etd. (1973)

Hybridizes with bands of centromeric heterochromatin and the X, the second, and both telomeres of the fourth chromosomes

Abbott e t a l . (1981)

Located in internal regions of polytene chromosomes

Stolletd. (1991, 1993)

Eckhardt and Gall (1971)

256

1004

-

1091 m

1094

-i

;

:

-.

:

;:

:=

:

T

A

A

1 -

A

I.-.; AT .. I- --

1.1-.

. _ .-

-.-. A .AT -

.-

A

”.- .

1

A

T

T

A T

.-.

A.

AT -

A -.-

._ . .--

T

A

AT -..

-

.. --

AT

z

1095

..-

T

: A‘U+O

--

1092 D 1093

1. F. Zhimulev

I.-.-

T

.

”’-

T

-.-

A

A

7

-

1096 m ~ -

1097

-,

-. A , - .

T-

; ::=: 1204 m ;

1090-

1208

I

P .

A ’ -

- AT

. . .

I

A

.. -- ..

AT

_..

.

.

l

.

1

-

L

.

A

’.

4 -

I.-

.

.

.

.

.

Sill EcoRI A BamHl I Xhol x Hind 111 8 Bgl 11 : KpnI T

T

a

A

._

A

m

..-

AT T

-..

..-

T

;::=:

:=:

__

T . .-. 1“-.

-

I

Figure 111. Repetitive DNA of telomeric repeat in D. m i r a d . Distribution of EcoR1, BamHI and Sal I restriction sites in the inserts of various phages (the arms of the phages are shown by thick horizontal lines, wrih phage number to the left) clearly shows the presence of a repetitive unit of 4.4 kb. Polymorphism for the Xho I and Hind111 sites indicates some heterogeneity of the repeat. After Steinemann and Nauber (1986).

of the salivary gland chromosomes, proximal heterochromatin, and the Y chromosome (Rubin, 1978; Young et al., 1983; Traverse and Pardue, 1988, 1989; Valgeirsdottir et al., 1990; Biessmann et al., 1992b; Danilevskaya et al., 1993). For example, the “He-T DNA” fragment is identified by in situ hybridization in telo-meres and pericentromeric heterochromatin of Drosophila as early as 7 days after exposure (Figure 112). No additional hybridization sites appear when exposure is prolonged to 228 days (Traverse and Pardue, 1988). HeT DNA is a complex family of repeated DNA. Much of the HeT DNA family appears to be a mosaic of several different classes of large sequence elements arranged in a scrambled array. The HeT-A element is about 6 kb in size (Biessmann et al., 1994; Danilevskaya et al., 1994a; Pardue, 1995; Biessmann et al., 1996; Pardue et al., 1996). The coding region has retrotransposon-like overlapping open reading frames (ORFs) with a “- 1”frameshift and a long noncoding region 3’ of the ORFs that makes up about half (2.4 kb) of the element (Danilevskaya et al., 1994a).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

257

Figure 112. Insitu hybridization of"He-T D N A with the ends of long chromosomes (arrows), the strand of ectopic pairing between them, the chromocenter (C),and the base of the fourth chromosome (4). after Young et al. (1983) and Pardue (1990).

HeT-A has an oligo(A)3' tail characteristic of elements that transpose through an RNA intermediate, but lacks sequences conserved in the RT domains of all retrotransposons. Some HeT-A elements are associated with coding sequences homologous to the gag-like region of LINE, supporting the classification of HeTA as LINE (Biessmann et al., 1990b, 1992a,b, 1993; Valgeirsdottir et al., 1990; Danilevskaya et al., 1992). The structure of the HeT-A element suggests that transposition of the element is mediated by a polyadenylated RNA intermediate that is very tightly, possibly covalently, associated with protein (Danilevskaya et al., 1994b). Another telomere-specific retrotransposon, TART (telomere-associated retrotransposon) has been shown to be confined to the telomere regions of Drosophila. TART has an ORF with homology to the RT domain of LINES. There is a second ORF with gag homology in longer TART elements, and it has an exceptionally long (5.1 kb) 3'UTR (Levis et al., 1993; Sheen and Levis, 1994). A novel mechanism by which telomeric DNA might be elongated is suggested by the occasional addition of DNA to broken chromosome ends (Traverse and Pardue, 1988; Levis, 1989; Biessmann et a!., 1990b, 199213). Addition of the HeT element occurs onto roughly 1% of the receding ends per generation (Biessmann et al., 1990b, 1992b). Terminal additions of HeT-A have been seen as a part of a process specific for the repair of broken ends or as having the potential to elongate all Drosophila telomeres. According to the model, the Gag-like protein is required in cis and may be responsible for targeting the RNA intermediate to chromosome

258

1.

F. Zhimulev

ends (Biessmann and Mason, 1992; Biessmann et al., 1992a, 1993; Mason and Biessmann, 1995). It is not known how elements such as HeT and TART recognize chromosome termini (for discussion, see Biessmann and Mason, 1994; Spradling, 1994). D. melanogaster telomeric DNA does not share the telomeric repeat sequences of another invertebrate, Ascaris lumbricoides (Levis, 1993). Copy number in a telomeric repeat can vary widely both in different polytene chromosomes of a nucleus (Figure 113) and in homologous chromosomes of a gland (Leibovitch et al., 1990) or within a strain (Young et al., 1983). Telomeric regions retain close homology to the DNA of very closely related species only. This is evidence that there is no considerable conservation (see Table 21). Interestingly, in spite of the spatial disassociation of blocks of telomeric repeats, the nucleotide sequences they contain can be generally similar, and this may be evidence of concerted changes in their structure (Valgeirsdottir et al., 1990). Other repeat types, in particular satellite DNAs, mobile elements, and middle repeats, are occasionally located at the ends of the chromosomes (see Table 21).

Figure 113. Variation in the copy number of the subtelomeric repeat in telomeric ends (arrows) of the polytene chromosomes of D. mirandu. 2 , 4 , 5 , X2,

X'L, X'R, chromosome numbers. After Steinemann and Nauber ( 1986).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

259

The Dm 665 clone of Drosophila possesses the activity of ARS elements (see Figure 1lo), which are AT-rich sequences several tens of nucleotides long comprising an 11-member conserved sequence necessary, albeit insufficient, for efficient replication and also an enhancer, an additional sequence. It is believed that the ARS isolated from the yeast genome has regions of DNA replication initiation. In Drosophila, a characteristic sequence of 11 nucleotides, 5'TAAATATAAAT, .and an enhancer sequence of 92 bp are detected in the ARS element (Danilevskaya et al., 1984, 1990; Gragerov et al., 1988a,b). It is of interest that reunion of a chromosome ring that was broken in the zone of a telomeric repeat, making a junction with the other end, is not forbidden. In entire compounds of the chromosomes of Drosophila whose distal ends of chromosome 2L unite with the pericentromeric region of 2R (Novitski et al., 1981a,b), the DNA of the telomeric repeat is detected in the junction by in situ hybridization (Goldstein et al., 1984). In D. miranda, during the formation of the neo-Y chromosome resulting from the fusion of the ancestral autosome with the Y-chromosome, two chromosomes unite, making a junction in the zone of telomeric heterochromatin (Steinemann, 1984).

C. Characterization of telomeric heterochromatin The notion that the telomeric tips of the polytene chromosomes have a heterochromatic nature has gained wide recognition. Indeed, blocks of heterochromatin, the sizes of which can vary even within a population, are present on the ends of the mitotic chromosomes in many animal and plant species (for review, see Mik10s and Nankivell, 1976; John and Miklos, 1979; Prokofyeva-Belgovskaya, 1986). It was seen that studies of telomeres in polytene chromosomes (with highresolution capacity) would provide a better understanding of telomere organization. Based on the available information, which region of DNA at the end of a linear chromosome can be defined as the telomere is difficult to determine because the structure of the DNA termini is so diverse. This emphasizes the uniqueness of this region of DNA. The stipulation will be recalled, however, that the telomere itself can be nonpolytene, that is, underreplicated in polytene chromosomes, and all the terminals may have all the structural features of the chromosomes of the subtelomeric regions, not those of the telomeres (I? A. Roberts, 1975). Another structural feature of the telomeric tips is the frequent occurrence of short, terminal deletions removing several of the most distal bands (see Figures 104 and 105, Table 19). The idea of the phenomenon is that a single or several terminal bands are missing in certain strains or in one closely related strains or species yet not in the other(s), with the difference readily detectable in the heterozygote. In Roberts' (1979) view, there are at least three hypotheses explaining why terminal bands are missing. First, there may be authentic deletions following the germline development. Second, there may be pseudo-deletions, artifacts pro-

260

1. F. Zhimulev ~

duced when squash preparations are made (Goldschmidt and Kodani, 1943). Stretching of the chromosome, even by a micromanipulator, disrupts its telomeric associations with difficulty, but the terminal bands are spared (Figure 114). Finally, variation in banding pattern can be related to the changes in compaction of a constant amount of chromatin in the same or several bands. Several variants of banding patterns were revealed on the end of the chromosome. When a single

-@J/j'j

e2\"

.v'

3L TIP

/

C

Figure 114. Stretching of the telometic contact between chromosomes 2R and 3L of Bosophilu with needles of a manipulator. (A and C) Squashed preparation before (A) and after (C)stretching. (B) Stretching and break of the contact region. 1-4, sequential steps of stretching. (D) Morphology and banding pattern at the ends of the chromosomes. After Hinton (1945).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

261

band occurred in the chromosome, it was thick and stained well; the greater the number of detected bands, the thinner they were (Kodani, 1947). Several types of morphology of polytene chromosome tips are described. The tips of the telomeres of the chromosomes of Trichotanypus pectinatus contain characteristic fanlike blocks of granular vacuolized heterochromatin (Bauer, 1936b). A dense heterochromatic “knob” was found in the telomere regions of Chironomus nuditarsus (Zhirov and Petrova, 1993) and Cryptochironomus obreptans (Morozova, 1993). The structure of the most distal regions of the chromosomes of Drosophila is different. In D. hydei, the terminal regions of the chromosomes, engaged in ectopic pairing, are represented by a block of compact, heavily staining material (Figure 115) that is connected by 10- to 40-nm diameter fibrils to the more proximal regions of the chromosome (Berendes and Meyer, 1968). However, in the chromosomes flattened out and sharply reduced in length and width by Miller spreading on a water surface (for review, see Zhimulev, 1992b), the terminal material loosens and acquires a filamentous texture (Figure 116). Chromosome ends can be different in D. melanoguster: they are dense black bands (Figure 117c-e), loose stained masses (Figure 117c), or the previously mentioned gray filamentous mass in other instances (Figure 117a, 117e, and 117f). It is of interest that chromosomes with both dense bands and a grayish mass on their ends can be engaged in ectopic pairing (see Figure 117e and 117f). Analysis demonstrates that such a mass is irregularly present, however, in all the chromosomes with missing terminal deletions (see Figure 105). Sutton (1940b) believed that this structure is nucleolar-like; it has the appearance of material seemingly seeping out of the end of the chromosomes. Lefevre (1976) called these masses “puffs,” although [3H]uridine did not incorporate into the most terminal regions of the chromosomes (Berendes and Meyer, 1968). This gray material differs from the rest of the chromosome in a specific set of features. For example, the polytene chromosomes are converted into lampbrush chromosomes after treatment with NaOH-urea mixtures, (for review, see Zhimulev, 1992b, 1996).The blocks of heterochromatin on the chromosome tips are decompacted last (Kodani, 1942). Interesting data were obtained when using needles to stretch telomeric ectopic contacts on squash preparations of the salivary glands of Drosophila (see Figure 114). Of 28 stretching performed, the associations between the distal ends of the chromosomes were severed in 16, and then, even after severance, the structure and banding pattern of the chromosome ends were not altered (see Figure 114B). An exceptionally great stretching of the material connecting the chromosome ends preceded the severance (see Figure 114B). In another 6 cases, the telomeric contacts could not be severed, and, on release from load, the chromosomes assumed their initial state with no traces of stretching remaining. In yet another 6 cases, the telomeric contacts were so tenacious that the chromosome regions proximal to the telomeres broke (Hinton, 1945).

Figure 115. Association of the telomeric ends of D. hydei under the electron microscope (a and b) and of D. oirilis under the light microscope (c). B, E band numbers; C, heavily staining body in the contact region of two (a) and three (b) telomeres of the chromosomes. (a and b) reprinted by permission from Berendes and Meyer (1968); (c) after Poluektova (1973).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

263

Figure 115. Continued

Transitions from a structurelessgray mass to a banded structure can occur. The fourth chromosome of Drosophila terminates with a band; when this region is transposed by a translocation to the middle part of the X chromosome, the telomeric end sharply elongates and acquires a filamentous texture (Lefevre, 1976). It will also be recalled that precisely cloudy material of this kind was consistently present on chromosome ends, including the chromosomes with teminal deletions. This allowed P. A. Roberts (1975) to adduce experimental data supporting the telomere concept.

264

1. F. Zhimulev

Figure 116. Morphology of the telomeric contact of three polytene chromosomes of D. hydei spread on a water surface and photographed under the electron microscope. Arrows, chromatin strands connecting the chromosome ends; arrowheads, fibrous gray mass located in the region of telomeric contact. The scale is 2.5 p,m. After Alanen (1986).

Figure 117. Ultrastructure of the telomeres of the polytene chromosomes of D. mefunogaster. (a and b) Chromosome 2L. (c and d) Chromosome 3R. (e) Telomeric contacts between 2L and 2R chromosomes. (f) Telomeric contacts between 2L and 3R chromosomes. Strains: (a and d) Batumi-L; (b) 28X-B(30A); (c) Oregon-R; (e and 0 In(l)br1"03 Bold arrows, fibrous structure at the end of the chromosomes and in ectopic contacts; asterisks, dense bands. After V. F. Semeshin (1990, unpublished observations).

Polyfene Chromosomes, Heferochromafin, and Position Effect Variegation

265

266

1.

F. Zhimulev

Based on these considerations, it may be concluded that one morphological manifestation of telomeric heterochromatin is capping of the tips of the chromosomes by this gray loose material in Drosophila. No such specialized structures were detected on the chromosome tips in many other representatives of Diptera, and, even when ectopic pairing of the telomeres happened to be disclosed, the extreme bands were involved in it, as, for example, in Chironornus tentuns (Beermann, 1952,1962;tenTusscher and Derksen, 1982), Ch. sdinurius (Michailova, 1973), and Ch. hummi (Kiknadze et al., 1976). In the midges of the Simuliidae family, the telomeric terminals of the chromosomes can be composed of a series of thin weakly and intensely staining bands (Chubareva, 1977b). In the gall midge Aphidaktes a ~ ~ m y dense ~ a , bands, regarded as telomeres (Chubareva and Kozlova, 1980),were found to cap the ends of all the chromosomes. Information concerning the location of active genes at the ends of the chromosomes is extensive: for example, Balbiani rings in the ovarian nurse cells of Calliphura erythrocephala (Ribbert, 1979); and various loosenings, such as in Chironomus behningi (Belyanina and Kolosova, 1979) or Ch. tenmns (Sass, 1981), Pseudodiumesa niuosu, Ps. branickii, and Promnipus gr. morio (Kuberskaya, 1979). At the extreme tips a heat shock-induced Balbiani ring in Ch. thummi (Carmona et al., 1985; Morcillo et al., 1988), a nucleolus in Phytomyza abdominalis (Block, 1969) and Pseudodiamesa gr. branickii (Michailova and Petrova, 1989), and micronucleoli in the polytene chromosomes of plants (Schweizer, 1976) (see Figure 75 in Section VII) were detected. RNA polymerase I11 is detected at the tip of the telomere of chromosome 3L of Drosophila (Kontermann et al., 1989). The toromeres are intensely staining bodies in which the material is not distributed as in the usual band, but rather as rings (tori) with cavities in the middle. These darkly staining structures were first identified in chromosome 3L of D. melanogaster (Barr and Ellison, 1973). A toromeric structure was later detected in the telomeric region of the microchromosome of D. lummei (Sinibaldi et al., 1976) and in the various chromosomes of D. aurariu, D. triaururia, and D. quadruria (Scouras and Kastritsis, 1985; Scouras, 1986). It is therefore difficult to conclude whether heterochromatin is present in the telomeric regions because their structure is enormously diverse. When a chromosomal structure is referred with some certitude to a telomere (e.g., as judged by the gray filamentous cloudiness at the chromosome tips of Drosophila), the structure lacks the two features usually assigned to heterochromatin: tight packaging and, as a consequence, heavily staining material. Numerous features of heterochromatin have concomitantly been revealed in the telomeres. These are discussed in the following sections.

1. Ectopic pairing Associations of the telomeric regions are widespread in representatives of Diptera (Table 22). Such contacts of the telomeres were found in chromosomes of divid-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

267

ing cells also-for example, in the grasshoppers (White, 1961) or polyploid plants (Sved, 1966). From a survey of the data compiled in Table 22 and those in the literature, conclusions regarding the main properties of ectopic pairing of the telomeres are as follows.

1. The ability of telomeres to enter into associations is related to the existence of a specific structure on the very end of the chromosome. This appears plausible because the chromosomes with terminal deletions are not involved in the associations (Kikkawa, 1938; Lapta and Shakhbazov, 1986). In Drosophila, the association frequency correlates with the size of the gray filamentous terminal structure (Roberts, 1979). The chromosomes retain their inherent frequencies of ectopic pairing when translocated to another genotype. More than that, when a recombinant chromosome is generated in a heterozygote for chromosomes with different frequencies of telomere ectopic pairing, its ability to enter into associations corresponds to that of the parent from which the telomeric tip was derived (Hinton, 1945). In D. melanoguster (Hinton and Atwood, 1941) and D. hydei (Berendes and Meyer, 1968),the involvement frequency of the polytene X chromosome in telomeric associations is lower in males than in females; in D. virilis, D. nouamexicana, D. texanu, D. littoralis, and D. imeretensis the frequencies are 1.2- to 4-fold lower (Poluektova, 1975a). This is presumably also related to a decrease in the amount of material involved in ectopic contact because the polytene X chromosome is composed of a single element in the male and of two in the female. 2. The strength of ectopic associations varies widely. In certain species, the association can be very weak, and, in Bauer’s (1936a) view, it depends on the extent to which the nucleus was squashed. According to other data, even when the chromosomes are stretched by manipulator needles, the telomeric associations hold faster than the subtelomeric regions of the chromosomes (Hinton, 1945). Telomeric asociations are not found in some species (e.g., in certain chironomids and mosquitoes of the Anopheles genus) presumably due to this variation (Trosch and Lindemann, 1973; Stegniy, 1979, 1987; Glazko and Zainiev, 1985). They are found in other representatives of these very families. In some species (e.g., Prodiamesa oliuacea), the contacts of the telomeres are so tenacious that, when cells are squashed, the end-to-end connections are not destroyed, and the chromosomes frequently have the appearance of a continuous long cable. The single chromosomes comprising the cable lose their “individuality,”thereby making difficult their identification and the building of chromosome maps (Bauer, 1936b; Petrova and

268

1. F. Zhirnulev

Table 22. Occurrence and Features of Ectopic Pairing in Telomeres of Polytene Chromosomes Characterization of telomeric contacts

Family, species

SALIVARY GLANDS OF DIPTERA Cecidomyiidae Aphidoktes sp. Chironomidae Ch. abewatus Ch. agilis

Ch. pllidivittatus Ch. plumosus Ch. salinarius Ch. species Ch. species Ch. species Ch. sp. Ya3 Ch. tentans

References

Grinchuk (1974) Belyanina (1989)

-

Kiknadze et al. (1996)

Telomeric DNA labels ectopic associations of telomeres -

Saiga and Edstrom (1985) Belyanina (1976)

-

Michailova (1973) Gavrila (1983) Belyanina (1989) Stoian et d.(1983)

-

Kiknadze et al. (1996) Telomeric associations identified in 10% of saliBeermann (1952) vary gland nuclei, frequently in malpighian tubules, more frequently in midgut where in 26 cases of associations chromosome 1L was involved in 13 and chromosome 3R in 11 cases; in nuclei of malpighian tubules in 24 associations, chromosomes 3L and 3R were involved 11 times

Ch. thummi

DNA from the telomeric Balbiani ring TBRIII labels ectopic associations of telomeres

Bauer (1935) Carmona et al. (1985)

Clinotanypus

One of the telomeres of the each of the long chromosomes connected with each other to form a loose granular “chromocenter”

Belyanina and Sigareva (1981)

Endochironomus albipennis

Telomeres contact with other chromosome telomeres and with the internal regions of the chromosomes

Michailova and Gercheva (1982)

E. impr

-

Michailova and

nervosus

Gercheva (1982)

E. tendens

Chromosome 3 is most frequently involved in as- Belyanina (1978b) sociations: arm S with arm lL, arm L with arm 2 s Telomeres contact with telomeres of the other Michailova and chromosomes and with the internal regions Gercheva (1982) of the chromosomes

Lipiniella arenicola, L. prima, L. moderata

Strong telomeric contacts result in pour spread chromosome preparation

Kiknadze et al. (1989), Shilova et d.(1992)

(continues)

269

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

Table 22. (Continued _______~~ ~

Family, species Species of the Orthocladiinae subfamily Orthocladius bipunctellus Prodiamesa olivacea

~

Characterization of telomeric contacts

References Michailova (1989a)

Ectopic contacts between C + telomeres

Michailova and Belcheva (1982)

Chromosomes 3 and 4 are almost constantly united by telomeres; chromosome 1 frequently joins them, forming “branching chromosomes” Of 187 nuclei, there were no associations of telomeres in only 7; of 10 possible variants of telomere association, the proportions were 54% for 1-2-3-4, 9.6% for 1-3-4, 14.9% for 1-2, and 21.3% for 3-4 associations

Bauer (1936b)

-

Petrova (1986), Chubareva (1986b) Chubareva (1980b, 1984, 198613)

Petrova and Chubareva (1978)

Psectrotanypus variw

Tight telomere associations

Smittia parthenogenerica Syndiamesa nivosa

Telomeric contacts occur frequently

Bauer (1970)

Telomeric contacts occur in 124 of 222 nuclei forming 34 variants: most frequently occurring were 1-4 (13.05% of nuclei), 2-5 (9.45%), 3-4 (25.65%), and 3-4-7 (4.95%)

Petrova (1983)

Attachment of the chromosomes in associations tighter than between centromeric regions; free telomeric ends infrequently occur Telomeres of the chromosome arms frequently associate

Dennhofer (1974)

Culicidae Culex pipiens

Orthopodomyia pulcripalpis Sabethes cyaneus Drosophilidae D.ananassae

D. funebris D. hydei

-

Chromosomes with terminal deletions not involved in telomeric associations and in contacts with intercalary heterochromatin

Munstermann et a[. (1985) Munstermann and Marchi (1986) Kikkawa (1938)

Bauer (1936a). Tinyakov (1936) Bauer (1936a), Fiala and Neubert (19521, Berendes (1963), Ananiev and Barsky (1982) (continues)

270

1. F. Zhlmulev

Table 22. (Continued) Characterization of telomeric contacts

Family, species

Telomeric associations in 91% of nuclei; of 498 associations: two chromosomes involved in 21.6%, three in 36%, four in 31%, and five in 2.4%; of 26 possible variants of telomeric associations, the following occurred most frequently: 2-3, 2-4,2-5,2-3-4, 2-3-5, 2-4.5, 2-3-4-5 chromosomes, making up 78.2% of all the associations; contacts between X-3, X-4, X-2-3, and X-4-5 chromosomes were not found. Telomeres frequently contact with subtelomeric regions

References Berendes and Meyer

(1968)

J. Derksen in ten Tusscher and Derksen

(1982)

D. imeretensis

Telomere of the second chromosome of gastric cecum cells involved in ectopic contacts less frequently than in salivary glands

Tiwari and Lakhotia

Of 15 possible combinations of telomeric con-

Poluektova (1975a ,b)

(1984)

tacts in males, more frequently occurring than others were 2-3 (27%), 2-4 (15.1%), and 3-4 (15.5%); in females: 1-4 (8.6%), 2-3 (24.7%), 2-4 (15.7%), and 3-4 (12.9%); occurrence frequencies of certain chromosomes in associations in females were 1 in 12.8%, ZL in 27.7%, in 7.2%, 3 in 27.7%, 4 in 18.9%, 5 in 5.8%, and 6 in 0.2%; in males, 1 in 7.8%; 2L in 26.7%;

2s

ZSin8.5%,3in29.8%;4in19.9%,5in6.8%, and 6 in 0.5%

D. littoralis

D. melanogaster

Of 15 possible variants, most frequently occurring in males were 2-3 (10.2%), 2-4 (9.4%), 3-5 (16.2%), and 4-5 (17.8%); in females, 3-4 (10.4%), 3-5 (15.8%), 4-5 (21.5%) -

Poluektova (1975a)

N. P. Dubinin, N. N. Sokolov and G. G . Tinyakov in Tinyakov, (1936), Goldschmidt and Kodani (1942,

1943) Telomeric region of the X chromosome frequently contacts with intercalary regions

Of 107 cases of telomeric associations in the Swedisch strain, the following chromosome contacts occurred: 2L-3L in 36 cases, X-3L in 18, X-2L in 15, ZL-3R in 11, 2L-2R in 10,

Prokofyeva-Belgovskaya

(1937b, 1938,1939d), Kaufmann et al. (1948) Hinton and Atwocd

(1941)

(continues)

271

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 22. (Continued)

Family, species

Characterization of telomeric contacts

2R-3L in 9, 3L-3R in 8, X-3R in 6,2R-3R in 3, and X-2R once of 10 possible pair combinations of telomere contacts, 4 combinations were significantly more frequent than the others, and 5 occurred considerably less frequently than expected In a study of 1060 cases of telomeric associations, it was found that the occurrence frequences of each of the 10 possible combinations in two strains were different; 40 triple and 7 quadruple contacts, as well as contacts with intercalary heterochromatin, were identified Associations of two chromosomes occurred most frequently; less frequent were associations of three, four, and five chromosomes Contacts detected between telomeres and with intercalary regions of the chromosomes In one strain: of 416 nuclei, telomere associations were observed in 129 (31%); the highest occurrences were observed for X-2L (16 times), X-3L (411, X-3R (15), 2R-3R (141, and 3L-3R (10); associations with the chromocenter and fourth chromosome occurred In another strain: of 400 nuclei, associations of 31 types were observed in 193 (48.3%), more frequently X-2L (26 times), X-3L (581, X-3R (25), and 2R-3R (19) in contact with chromocenter and the fourth chromosome; when the fourth chromosome is translocated to the end of chromosomes 2L and 2R, it establishes ectopic contacts with other chromosomes (e.g., 59% of cases with the X chromosome in both strains) Q+-bands at the telomeric and centromeric ends of the fourth chromosome pair ectopically Terminal ectopic contacts between chromo. s o m a 3L and 3R only when they contain a telomere Telomeres contact both with each other and regions of intercalary heterochromatin In gt strain, among 550 cases of ectopic pairing, chromosome 3R was not involved in telomeric associations; 3L made contact with telomeres of 2R, 2L, and X; 2R made contact

References

Hinton and Sparrow (1941), Hinton (1945)

Hinton and Atwood (1941, 1942), Hinton

(19451

Warters and Griffen (1950) Kaufmann and Iddles

(1963) Kaufmann (1969),Kaufmann and Gay (1969)

Barr and Ellison (1971b) Sauer (1971)

Lefevre (1976) Kulichkov and Zhimulev (1976),Zhimulev et al.

(1982)

(continues)

272

1.

F. Zhimulev

~~

Table 22. (Continued) ~~

Family, species

Characterization of telomeric contacts with 3L, 2L, X, and 8 intercalary regions; 2L made contact with 3L, 2R, X, and 4 intercalary regions; and the fourth chromosome made contact with centromeric heterochromatin and the proximal regions of the other chromosomes Labeled stretches of ectopic pairing between telomeres after in situ hybridization of telomeric DNA

Associations in Urbana-S (CalTech) strain identified in 80% of nuclei (300 examined) mainly between the X chromosome and 2L and 3R: X-2L (22%), X-3R (25%), X-2L-3R (34%), 2L.3R (9%),X-2R (7%), 2L-2R (less than I%), and X-3L (3%); in Urbana-S (Bowling Green) there were associations in only 5% of nuclei, mainly between 2L and 2R, and in hybrids between strains in 50% of nuclei Nucleoli frequently adjacent to telomeres

In the 0 3 2 strain, of 10 possible types of pair associationsof telomeres, most frequently occurring in 1693 nuclei were X-3R (11.8%), 2L-X (3.2%), X-3L (3%), and 3L-3R (2.9%); the rest occurred at a frequency lower than 2% In D32 x Swedisch hybrids the highest association frequency (1.6% in 1585 cells) was de, tected between the X and 3L chromosomes; in Sw X 0 3 2 between the X and 3R (3.6% in 573 nuclei); in the 0 3 2 X Batumi hybrid between 2R and 3R (6.5% in 1061 nuclei); in Batumi X 0 3 2 hybrids, the frequency was 1.6% in 732 nuclei

References

Rubin (19781, Young et al. (1983), RenkawitzPohl and Bialojan (1984), Hammond and Laird (1985a,b), Traverse and Pardue (1988), Danilevskaya et al. (19901, Pardue ( 1990), Karpen and Spradling (1992), Spradling et al. (1992) Roberts (1979)

Genova and Semionov (1985a,b) Ananiev and Barsky (1985) Lapta and Shakhbazov (1986)

(continues)

273

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 22. (Continued) Characterization of telomeric contacts

Family, species

D. novamexicana

References

In the yMRwMRctMKpNfY strain the frequency of paired telomeric contacts was 26.25%, of triple contacts 3.10%, and ofquadruple 0% in 419 nuclei; of 20 types of telomeric associations, X-2R occurred in 10.2696, X-3L in 5.0196, and 2R-3L in 8.35% of nuclei

Vagapova ( 1991)

Of 15 possible telomeric associations the follow-

Poluektova (1975a)

ing occurred more frequently than others in males: 4-X (13.2%), 2-4 (15.6%), 2-5 (8.7%), 3-4 (22.1%), 3-5 (12.4%), and4-5 (9.5%); in females: 3-4 (8.5%),4-X (18.8%), 2-4 (11.8%), 3-4 (17.0%), 3-5 (8.7%), and4-5

(9.1%) D. pseudoobscura

Occurrence of telomere associations of different chromosomes: XL-XR in 55 nuclei, XL-2 in 11, XL-3 in 4, XL-4 in 16, XR-2 in 36, XR-3 in 11, XR-4 in 27, 2-3 in 25,2-4 in 46, and 3-4 in 20

T. Dobzhansky in Hinton and Atwood (1941)

D. repleta D. quadraria

-

Frolova (1936b)

Associations of the telomeres of 3L or 3R chromosomes

Scouras (1986)

D. texana

Of 15 possible types of associations, the follow-

Poluektova (1975b)

ing occurred more frequently than others in males: 2-3 (9.9%), 2-4 (21.6%), 2-5 (12%), 3-4 (l2%), 3-5 (10.8%) and 4-5 (17.2%); in females: 2-X (8.7961, 2-3 (9.4%), 2-4 (15.6%), 2-5 (9.2%), 3-4 (10.7%), 3-5 (10.2%) and

4-5 (15.8%) Frequencies of association involvement by chromosome in f e d in male

X 2 3 4 5 6

D. virilis

13.3% 20.7% 18.2% 24.2% 22.1% 0.5%

6% 23.7% 18.9% 28.0% 22.7% 0.7%

In 40 “not strongly squashed” nuclei, 35% of the chromosome telomeres are united into groups of two to four chromosomes More frequently associations of two, less frequently of three to five chromosomes

Bauer (1936a)

Warters and Griffen

(1950) (continues)

274

1. F. Zhimulev

Table 22. (Continued) Characterization of telomeric contacts

Family, species

Of 5577 cases of ectopic contacts of telomeres, 89% were formed of two, 10.6% of three, and 0.4% of four chromosomes; the larvae did not differ in frequencies of three-chromosomal associations during the last day of larval development (72-99 h); the female X chromosome was involved in associations 1.5-2.0 times

References Poluektova (1973)

more frequently than that of males; in autosomes, association frequencies were higher in males at all developmental stages (see Figure

118) Of 15 possible combinations of telomere con-

Poluektova (1975a)

tacts, the following occurred more frequently in strain N9 in males: 5-X (8.9%), 2-3 (20.1%), 2-4 (10.2%), 3-4 (18.5%), and 3-5 (9%); in females: 2-4 190.2%), 3-4 (9.7%),

4-X (8.1%), 5-X (10.7%), 2-3 (14.7%), 2-4 (14.7%), and 3-4 (14.1)% Frequencies of association involvement by chromosome: in female Strain 9 (strain 151)

X 2 3 4 5 6

D. virilis X D.

19.4 (11.8) 22.5 (18.1) 21.8 (13.0) 21.1 (15.2) 14.7 (12.9) 0.4 (3.2)

Poluektova (197513)

in male Strain 9 (strain 151)

10.5 (5.2) 23.3 (13.7) 27.0 (13.5) 22.8 (19.8) 15.7 (12.6) 0.7 (3.0)

Asynchronous replication termination in the 20A telomeric region

Gubenko ( 1976b)

Mycetophilidae Epuh agarici

-

Gavrila (1983)

Sciaridae Rhynchosciara angelae

Contact of telomeres with intercalary regions

Diaz (1972)

littoralis hybrids

(continues)

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

275

Table 22. (Continued) Characterization of telomeric contacts

Family, species

References

R. hollaenderi

Nucleolar material detected with ammoniacal silver makes contacts with telomeres

Stocker et al. (1978)

Sciara impatiens S. ocellaris

-

Carson (1944)

Associations of telomeres with each other and intercalary regions

Perondini and Dessen (1985)

-

Chuhareva and Shcher. bakov (1963), Simonenko (1966), Chuhareva (1980a) Shcherbakov (1968)

Simuliidae Ewimulium securifm

Telomeric regions of the salivary gland chromo, somes in constant ectopic association -

Petrova (1981)

-

Ralcheva (1974)

Wilhelmia equina

-

Tephritidne

In collections made in May, ectopic contacts of telomeres detected not more than in 4% of larvae; in those made in July to September, there was detected in 14-32% of larvae -

Grinchuk (1968), Knoz et al. (1975) Grinchuk (1969a)

Metacnephia variafiuis Prosimulium petrosum

Mavragani-Tsipidou et al. (1992)

Dacw okae

SUSPENSOR OF PLANTS Phaseolw coccinew Micronucleoli are frequently attached to Q+telomeric hands of polytene chromosomes

Schweizer (1976)

Chubareva, 1978; Chubareva, 1986a,b). Such cases were also described in D. melanoguster (Lefevre, 1976). In Psecworunypus oarius, the telomeric associations are so tenacious that it is occasionally hard to decide whether the telomeres of two chromosomes have united or represent a single two-armed chromosome with a distinct centromere (Chubareva, 1980b, 1986a,b). In Cukx pipiens, the telomeric associations hold faster than the centromeric (Dennhofer, 1974). 3. Observing telomeric associations in D. funehis, Tinyakov (1936) noted that “the association of the telomeres with each other is very weak and inconstant because the same ‘ends’ enter into different associations in different nuclei.” Since then, many investigators have demonstrated that the as-

276

1. F. Zhimulev

sociations are not constant, and particular chromosomes show high preference in entering into associations. Theoretically, a great number of ectopic contacts between telomeres may be envisaged; 10, for example, for 5 chromosomes associating in pairs. The greater the number of chromosomes entering associations, the more diverse are the ectopic contacts. However, the data of Table 22 indicate that only certain combinations occur virtually in all species, while others are very rare or do not occur. 4. The chromosomes mostly associate in groups of two; groups of three chromosomes occur much more rarely. The data obtained in D. hydei (see Table 22) are quite exceptional. 5. There are great interstrain differences in the involvement of particular chromosomes in associations. The pertinent data (Table 23) obtained in a study performed by the same investigator (Hinton, 1945) are also entered into Table 22. The values are intermediate for the telomeres from crosses between strains differing in involvement frequencies of telomeres in associations. 6. It is unclear whether there are any differences between the sexes. For example, frequencies of association by a particular autosome are higher (Figure 118),and the associations frequencies of three chromosomes are somewhat lower, in males than in females of D. vin'lis (Poluektova, 1975a).

Table 23. Frequencies (%) of Ectopic Contacts between Telomeres of Two Strains of D. melanogaster in Different Years" Strain of DrosophiLc Variant of ectopic contact

x-2L X-2R x-3L X-3R 2L-2R 2L-3L 2L-3R 2R-3L 2R-3R 3L-3R TOTAL

Swedisch-b

Oregon-R JUh

1940 17.3 3.1 17.3 16.3 3.1 3.1 5.1 6.1 16.3 12.2 100

"After Hinton (1945).

711 1 1943

11/17 1943

21.0 12.0 5.0 20.0 2.0 1.o

20.0 4.0 8.0 24.0 4.0 2.0 1.o 1.o 30.0 6.0 100

1.o

1.o 31.0 6.0 100

Iulv 1941 14.0 0.9 16.8 5.6 9.3 24.3 10.0 8.4 2.8 7.5 100

11/28 1943

11/30 1943

3.0 10.0

1.o 14.0 0.0 21.0 2.0 3.0 1.0 3.0 50.0 5.0 100

8.0

9.0 1.o 3.0 1.o 5.0 52.0 8.0 100

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

277

30 25

20 15 10

5 7;

78

I4

90 .46 96-99

Figure 118. Involvement frequencies of the distal end of the third chromosome of D. uirilis in different telomeric associations. Abscissa, developmental stage; ordinate, involvement frequencies; hatched columns, association frequencies in the salivary glands nuclei of females, open columns, same in males. After Poluektova (1973).

7. The various features of ectopic pairing of the telomeres do not change during development (see Figure 118,Table 22). 8. Cytoplasm has no effect on the frequencies of ectopic pairing of the telomeres, since association frequencies in offspring from Drosophila strains differing in the degree of expression of this character are not dependent on the direction of the cross (Hinton, 1945). Similar results were obtained in interspecific crosses of Drosophila of the virilis group: the frequencies of telomeric associations of both single chromosomes and chromosomes associated in groups were not related to the direction of the cross (Poluektova,

1975b). 9. It is of interest that Hinton’s observation that the frequencies of telomeric associations change with time (see Table 23) has been neither proved nor disproved. However, Roberts’ finding that the terminal deletions in D. mehogaster, which Bridges as well as Demerec and Hoover described in the 1930s and 1940s, cease to be such with time argue in favor of this conclusion. Thus, in the 1960s, several thin bands not homologous to the other parent chromosome were found to appear at the end of chromosome Df(J)260-1 instead of the terminal deletion (Roberts, 1969b, 1976). There prsumably occurs an addition of a telomere repeat (see earlier) after its removal by a deletion. Additional bands appear with time at the end of chromosome 2R (Roberts, 1974, 1979).

278

1. F. Zhimulev

10. The types of ectopic contacts of the telomeres and the composition of their associations are not affected by temperature. However, telomeric contact frequencies are higher at low temperatures. Thus, the percentages of nuclei with ectopic contacts were 14-26% in larvae of Drosophila cultured at 17°C and 9-17% in those cultured at 23°C (Hinton, 1945). 11. A combination of the position effect modifiers,variation in temperature, and heterochromatin amount demonstrated a modifier effect. The frequencies of nuclei with telomeric associations were 25.04% in X/O larvae of Drosophila melanogaster at 18°C and 3.46% in XW larvae at 25°C (Vagapova, 1991). 12. Information was obtained on the sensitive periods of the formation of telomeric contacts: the frequencies of telomeric contacts were the same in larvae of Drosophila kept at 23°C during the first 24,48, or 72 hr and then transferred to 17°C to develop to pupariation as in larvae left at 17°C. However, when larvae developed at high temperature for 4 days, telomeric contact frequencies were the same as at 23°C. This was thought to be related either to a particular temperature sensitivity during the fourth day of life or to a cumulative effect of temperature (Hinton, 1945). 13. The telomeres establish ectopic contacts not only with one another, but also with many regions of the chromosomes (see Table 22). Hinton (1945) described 180 contacts in D. melanoguster. It was found that the intercalary regions forming associations with the telomeres are nonrandomly distributed along the chromosome length. The most distal regions of the chromosomes-2B, 3C, 21-22,56,58,99-100, and 100CD, for example-and 46 other regions make contacts more frequently. In the X chromosome, the highest frequencies of ectopic contacts with the telomeres were for the

lCD, 2B, 2D, 3B, 3C, 3D, 4B, 5,5-6,7B, 8F, 8-9,10A, 11A, 12D, 14, 15-16, 16E, 18EE and 19E regions. The contact frequencies were very different, with involvement in ectopic contacts occurring 11 times for region 2B, 22 times for region 3C, and 1-3 times for the rest of the chromosome regions. It is easy to see that the regions of intercalary heterochromatin establish contacts in the majority of cases (see Sections VII,C,l-6). The contact frequencies of the telomeric regions with the intercalary heterochromatin regions of various chromosomes proved to be very different; for example, in the Oregon-R strain, the X chromosome entered into associations eight times more frequently than chromosome 2L and 13 times more frequently than chromosome 3R. In contrast, the telomeres of the X and 3R chromosomes did not differ in their ability to enter associations with those of the other chromosomes, and the frequencies were only twofold less in chromosome 2L (Hinton, 1945). 14. The spatial disposition of the chromosomes in the nucleus is of great importance in establishment of telomeric ectopic contacts. The microchromosomes of Drosophila, lying at the centromeric pole of the nucleus, hardly ever enter into associations, so there is no information on this point (see

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

279

Table 22). However, when a microchromosome is transposed to the end of the long chromosomes, its telomere is normally involved in associations. 15. How real is the existence of telomeric associations in the intact nucleus?The contacts remaining undisrupted even in squash preparations make conceivable the arrangement of the telomeres in associations in the whole telomere. However, as early as in the first schemes of the spatial distribution of the chromosomes in the nucleus proposed by Frolova and Emmens (see Zhimulev, 1992b, 1996),the telomeres were not tied into a single knot, although they were depicted close to each other. Bauer (1936a) regarded the postmortem changes in the structure of the nucleus as associations. In Fujii’s (1942) view, fusion of the telomeres is possibly nothing more than a secondary association of the chromosomes, which have been initially independent of each other. Decisive evidence might have been obtained on nuclear sections because they demonstrate that the distal ends lie independently. Three to five labeled sites were detected by in situ hybridization of telomeric DNA with the unsquashed nuclei of the salivary glands of Drosophila (Hammond and Laird, 1985b). The same was demonstrated with optic sections: the telomeres lie independently, although not far apart, and they are randomly distributed in the nuclear volume in the midgut of Drosophila (Mathog et al., 1984; Hochstrasser et al., 1986; Hochstrasser and Sedat, 1987a).No telomeric associations were identified in 150 unsquashed nuclei of Chironomus thummi studied by scanning microscopy (Trosch and Lindemann, 1973).

2. Differential staining Staining of the salivary gland chromosomes of Chironomus thummi thummi revealed weak C+-bands at the ends of some chromosomes. The telomeres of the closely related Ch. thummi piger subspecies stain heavier (Hagele, 1977a; Sigareva, 1981).The chromosome ends ofCh. plumosus (Belyanina and Sigareva, 1978), Clinotanypus nervosus (Belyanina and Sigareva, 1981), Orthocladius bipunctellus (Michailova and Belcheva, 1982), and Chironomus balatonicus (Michailova, 1987a) stain weakly with this technique. In Simulium omatipes and S. melatum, some chromosomes have very prominent C+-bandsat the tips of the telomeres, while other chromosomes lack them together (Bedo, 1975a). In Endochironomus impm, the telomeres of the first through third chromosomes stain for constitutive heterochromatin (C+);the telomere of the left arm of the first chromosome is Q’ (Michailova and Gercheva, 1982). In D. hydei, the compact blocks at the tips of the chromosomes (see Figure 115) and the ectopic fibers between them form groups of regions showing DNA late replication of the continuous type (Berendes and Meyer, 1968). In D. lummei, the toromeres (see earlier) fluoresce after staining with quinacrine (Q+), by the Giemsa method. They are rich in repeats (Cot 10-1-10-2) and are late replicating (Sinibaldi and Barr, 1979; Evgen’ev et al., 1982a, 1983).

280

I. F. Zhimulev

3. Underrepresentation of DNA Roberts (1979) described a Drosophila strain in which the structure of the tips varies among the cells of the same gland, for example, the morphology of the features of the 60F region in Oregon-RC or the end of the X chromosome of the Urb a n d strain. This is most readily accounted for by the incomplete polytenization of telomeric DNA sequences in one cell and a more complete polytenization in another. Similar variations among nuclei were revealed by in situ hybridization of the labeled fragments of telomeric DNA (Leibovitch et aE., 1990, 1991). High temperature (32°C) causes structural instability of the telomeres of polytene chromosomes in E-32 D. melanoguster strains. Changes in telomere length are retained for several generations at 23°C. In in situ hybridization of the Dm 665 clone containing a telomeric repeat, considerably more label concentrates in the “long” than the “short” telomeres (Danilevskaya and Lapta, 1990). Probably, there is variability in the length of telomere DNA between chromosomes. In D. melanoguster the X chromosome telomere is as much as 100 kb on Dp I187 (Karpen and Spradling, 1992; Spradling, 1994). In contrast, the first gene lies only about 15 kb from the tip of chromosome 3R (Levis et al., 1993). There is no agreement between the representation of DNA amount in the haploid genome and in the salivary gland chromosomes. In Ch. palliditittatus a telomeric sequence 340 bp long constitutes 1.2% of the genome; that is, there are approximately 300 kb in each telomere (Saiga and Edstrom, 1985). This amount should be DNA compacted into a very large band. Such has not been found at the chromosome ends. In the haploid set of D. melanoguster,there should be 200-250 copies of the Dm 665 subtelomeric fragment (60-80 copies per telo-mere) (Danilevskaya et al., 1990); that is, large bands 140-200 kb in size might be expected at the tips of the telomeres. This has not been found to be the case. Comparison of the amount of telomeric DNA in the salivary glands and brain of larvae on genomic blots demonstrates that several fractions underreplicate in polytene chromosomes (Figure 119). In situ hybridization of cloned probes for telomere sequences shows that they replicate only partially during polyploid nurse cell growth (Hammond and Laird, 1985a). The subtelomeric region in Dp I187 is underrepresented in D. mehogaster salivary gland chromosomes (Karpen and Spradling, 1992; Spradling et al., 1992). However, unlike the salivary gland cells, endopolyploid nurse cells did not show significant changes in the copy number of the telomeric regions in the minichromosome (Spradling et al., 1992).

4. Heterochromatin proteins Protein H 1, preferentially binding to heterochromatin, is frequently detected in the telomeres as well (James et al., 1989). The inconstancy in its manifestation is no surprise when one considers the variation in heterochromatin.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

28 1

Figure 119. Various representations of telomeric DNA in the diploid nuclei of brain (D) and in the polytene chromosomes of salivary glands cells (P) in DTosophila larvae. ( A ) Hybridization with labeled DNA from the 848 a-tubulin gene. (B) The same filter after hybridization with the telomeric DNA. After Young et al. (1983).

5. Changes in telomeric heterochromatin manifestation A characteristic feature of heterochromatin is change in its manifestation due to modifiers of position effect variegation. There is evidence indicating that the structure of the chromosome ends may be changed by altering the dose of the Y chromosome in D. melanoguster. For example, 6.4% of the 2L arms had a constriction at the end in males with the constitution XO, and 98.7% of 2L arms showed a constriction in XYY males (Schultz, 1947). In D. lummei, the size and occurrence of the toromeres are strongly temperature dependent: only 0.5-1.4% of salivary gland cells have toromeres at 25°C and 18°C; the respective percentages are 93.2% and 99% at 15°C and 12°C. A similar dependence was also found for the number of toromeres in the nucleus. Only 6.1% of cells have more than one toromere at 15"C, and 63.5% at 12°C (Sinibaldi et al., 1976). According to other data, the toromeres in larvae reared at normal temperature either are not detected or they have the appearance of a small dot; they are seen as large bright Q' bodies in larvae reared at 12°C (Evgen'ev et al., 1982a, 1983). Temperature can also affect chromosome structure; for example in the blood sucker midge Wihlmia equina, the short end of chromosome 3 becomes wider and it looks like a fan in winter and spring, whereas it is the same diameter as the rest of the chromosome in summer (Grinchuk, 1969a,b). The end of chromosome 3 undergoes similar changes in another midge, Boophthora ery throcephala (Petrukhina, 1970). In Ch. balatonicus,heterochromatin is more distinctly seen at the telomeric tip near the nucleolus, particularly in larvae c,ollectedduring the winter season (Kiknadze and Kerkis, 1986). Data indicating that the manifestation of ectopic pairing can be modified were considered.

282

1. F. Zhimulsv

6. Telomeres and nuclear envelope The telomeric regions are located at the nuclear envelope as the other kinds of heterochromatin are (Bauer, 1935; Kimoto 1958; Sved, 1966; Gruzdev and Kiknadze, 1970; Skaer and Whytock, 1975; Fiil, 1978; Hill and Watt, 1978; Quick, 1980;Agard and Sedat, 1983; Mathogetal.! 1984;Thermanand Murashige, 1984; Petrova et af,, 1985; Hochstrasser et al., 1986;Hochstrasser and Sedat, 1987a; Hill and Whytock, 1993; Demburg et al., 1995) or are associated with nuclear matrix (de Lange, 1992). Thus telomeric DNA in polytene chromosomes forms heterochromatin, which has a number of features, primarily structural, distinguishing it from the other types of heterochromatin.

IX. THE B CHROMOSOMES Chromosomes that are different in structural and functional features from the other chromosomes and present in addition to the chromosomes of the regular (A) set are referred to as supernumeraries, or the B chromosomes. The major features of the B chromosomes are (1) a much smaller size than the chromosomes of the regular set, (2) a genetic constitution that produces little physiological effect in the organism whether present or not, (3) variation in number in different cells of an organism and in different individuals of a population, (4)no pairing with the chromosomes of the regular set in meiosis and ( 5 ) chromatids frequently not disjuncting at the anaphase of mitosis. These features are not always consistently observed (Battaglia, 1964). About 15 different terms have been coined for the chromosomes of this type; “accessory chromosomes,” “additional chromosomes,’’ and “B chromosomes” are the most frequently used. The other terms encountered in the literature include “diminutive chromosomes,” “extra-diminutive” and “diminutive Bchromosomes,” “extra-” and “fragmented chromosomes,” “microchromosomes” (incorrect), and “minute accessories.” In addition, the supernumeraries are designated by different symbols: (1) B (C, D, E, F); (2) B, and B,; (3) fl, f2,fl, fs, F, and F1; (4)e; and ( 5 ) E, e, and ee (Battaglia, 1964). The B chromosomes have been found in 5 10 species of dicotyledonous plants (2.6%) among species whose karyotypes have been described, and in 1007 species of monocotyledonous plants (3.6%). There is scattered information in the literature indicating that the B chromosomes occur in gymnosperms, ferns, mosses, and lichens. In animals, the number of species with identified B chromosomes is mainly dependent on the specific interest of cytologists in a representative of a particular taxon. For example, of 263 animal species with B chromosomes, more than 40% are insects, according to Jones and Rees (1982).

Polytene Chromosomes, Heterochromatin, and Position Effect Varieoation

283

This unusual type of chromosome has long attracted the attention of investigators, and it is not surprising that it has been described in many extensive compilations, reviews, and monographs (Muntzing, 1954,1966; Swanson, 1957; Battaglia, 1964; White, 1959,1973; Hewitt, 1973;Volobuev, 1978;John and Miklos, 1979; Jones and Rees, 1982; Chubareva and Petrova, 1984; Ilyinskaya and Petrova, 1985; Prokofyeva-Belgovskaya, 1986; Bell and Burt, 1990; and others). Most students of B chromosomes believe that they are composed mainly of heterochromatin. If such is the case, study of the interphase polytene nuclei of insects offers the possibility of analyzing in detail the organization of heterochromatin of another type in the supernumerary B chromosomes. Figure 120 presents various types of diploid (in metaphase spreads) and polytene (from salivary gland cells) supernumerary B chromosomes. In the dividing cells of gonads and neural ganglia, centromeric regions, even arms, are recognizable in the B chromosomes, although these chromosomes are much smaller than the other members of the chromosome set (Chubareva, 1985).In the larvae of Simulium rnorsitans, the supernumeraries are rounded at metaphase, and occasionally have the appearance of elongated commas (Kudule and Zitzer, 1975). It is characteristic that nuclei with polytene chromosomes in the supernumeraries tend to vary greatly in morphology. These can be structureless clumps of loose (see Figure 120) or compact chromatin resembling P-heterochromatin (Chubareva, 1971; Valkanov and Michailova, 1974; Michailova, 1976; Petrova et al., 1981; Michailova and Petrova, 1989). In some cases (Phryne cincta), these are small, rounded (Wolf, 1970) or elongated (e.g., in Chironomus aberratus; Belyanina, 1989), strongly heteropycnotic bodies resembling a-heterochromatin. In Ch. behningi, two types of B chromosomes are most frequently encountered: one appearing as darkly staining a-heterochromatic bodies and the other as a network of a rare heterochromatic accumulations. Each type is observed in different individuals and within the salivary gland cells of an individual (Belyanina and Kolosova, 1979). Supernumerary chromosomes showing banding patterns have been described in many papers (Chubareva and Petrova, 1968; Chubareva, 1974; Chubareva and Ralcheva, 1974; Belyanina, 1975; Michailova, 1985, 1987a,b). The number of bands can be large: 11 bands in the left and 15 in the right arm of Glyptotendipes p i p e s (Miseiko et al., 1971). In the Simuliidae midges, one or two very large blocks of compact material strongly resembling a-heterochromatin are identified in the supemumeraries showing a banded structure (Chubareva and Shcherbakov, 1963; Shcherbakov, 1965; Chubareva, 1974; Chubareva and Kachvoryan, 1974). In Odagmia ornata, one arm of the supernumerary chromosome exhibits a banding pattern while the other arm is uncoiled to a great extent, imparting a fanlike shape to the chromosome. This is associated with a very large, heavily staining “centromeric” band (Chubareva, 1979). In some cases only one arm may be heterochromatic (e.g., in Eusimulium inflatum) (Chubareva, 1974). In Simulium mor-

284

I. F. Zhimulev

Figure 120. Variety of the morphology of mitotic (m) and polytene (a-I) B chromosomes (indicated by arrows). ( a 4 1 B chromosomes of Chiraomw plumosw. (d) The pattern of replication termination. Arrow indicates the B chromosome with nucleolus. ( e )OTthOcladiw bipuncteuw. (f) Accumulation presumably of B chromosomes in the salivary gland cells of Chironomw gr. thummi. (g and h) Loop-shaped B chromosomes in Ch. bunus. (i) Odagmiu m t a . ( j and k) Sulcicnephia outshinnibti. (I) Cneta zachuriensis. (m) Giemsa-stained (C+-banding)metaphase chromosomesof

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

285

Figure 120. Continued

a Drosophila albomicaw male with a single B chromosome. X, Y, 2, 3 , 4, designations of the chromosomes. reprinted by permission from ( a d ) Key1 and Hagele (1971); (e) after Michailova (1982); (f) after I. E. Kerkis (1990, unpublished); (g and h) after Kerkis et d.(1989); (j-I) after L. A. Chubareva and N. A. Petrova (personal communication); (m) after Hatsumi (1987).

286

I. F. Zhlmulev

Figure 120. Continued

sitans, the B chromosomes are of two kinds: (1) those composed of “heterochromatin,” that is, of a reticular substance exhibiting one or two bands; and (2) fanlike bodies with a clump of heterochromatin at the narrow ends and a set of six to seven bands at the wide ends of the bodies. In some larvae, both types are found together (Kudule, 1974). In Ch. bonus, the B chromosome is represented by a dense heterochromatic body in all the studied cells of malpighian tubules. Vari-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

287

ous types were found in the salivary glands, from a dense block of heterochromatin to a banded structure with stray chromatin fibers scattered throughout the nucleus (see Figure 120). The morphology can vary in different salivary gland cells (Kerkis et al., 1989). Cases are known in which morphology varies considerably. In Ch. melanotus, the B chromosomes of only malpighian tubule cells display banding patterns; “heterochromatic” bodies are seen in the salivary gland cells (Keyl and Hagele, 1971). According to Ityinskaya and Petrova (1985), in Chironomus plumosus, the size, shape, and extent of heterochromatization, or distinctness of the banding pattern, of the B chromosomes vary not only from one larva to another and in different populations, but in different cells of a gland. Puffs are sometimesfound in the B chromosomes (Chubareva, 1984) [the data given in Shcherbakov and Chubareva’s (1966) paper are not reliably documented], as are nucleolar organizers on which additional nucleoli are formed, but in a tissue-specific manner: in malpighian tubules but not in salivary gland cells in Ch. plumosus (Keyl and Hagele, 1971). The B chromosomes incorporate [3H]uridine in this species (Ilyinskaya e t al., 1991). Nucleolar organizers were also found in the B chromosomes of Cnephia dacotensis and C. ornithophila (Procunier,

197513). Measurements of the DNA content in the nuclei on Feulgen-stained squashes of the B chromosomes of Ch. plumosus demonstrated that they contain from 1.6% to 5.9% of total DNA content in the nuclei (Ilyinskaya and Petrova, 1985). No agreement between polyteny degree of the supernumerary chromosomes and a definite ploidy class was found. This was accounted for by the different replication from one region to another (Keyl and Hagele, 1971; Ilyinskaya and Petrova, 1985). Data on the occurrence of the B chromosomes in representatives of the Diptera order are presented in Table 24. Wide variations in the frequencies were found both between and within species. It cannot be excluded that the ability to form B chromosomes is species specific, being possibly specific even to distinct populations. For example, karyotype data on 138 species of the Simuliidae species are given in Chubareva and Petrova’s (1979) compilation. In spite of studies on this large number of species, supernumerary chromosomes have been identified in only 17. There are no B chromosomes in three populations of Sulcicnephia o w shinnibti of the highly elevated reservoirs of Tadjikistan, whereas 76.2% of individuals of a Kirghizia population were found to possess supernumeraries (Chubareva and Petrova, 1984). The dipteran species appear to obey White’s (1973) rule that the higher the numbers of larger B chromosomes, the less frequently larvae having them occur in a population (see Table 24). The highest number of supernumeraries generally is not more than 12, although it is 45 in exceptional instances (see Tables

288

1. F. Zhlmulev

Table 24. Occurrence of B Chromosomes in Representatives of the Diptera Order

Percentage of individuals with B chromosomes

Number of Mitotic (M) B chromosomes or polytene in individuals (P)

-

1-7

M, P

Wolf (1950, 1954, 1962,1970)

Hykmia ulna

-

1

M

Boyes and van Brink (1965)

H. cilimura

19.2

1-3

M

Boyes (1954a,b),

Family, species

References

Anisopodidae

Phryne cincta Anthomyiidae (Muscidae)

Boyes and van Brink (1965) Chamaemyiidae Leucopis obscura

-

3-5

M

Boyes and b y e s (1975)

-

P P

Belyanina (1989)

P

Kiknadze et al. (1993) Ilyinskaya (1994, 1995)

Chironomidae

Chironomus &atus

1.11 2.26

-

(different populations)

Ch. ngilis Ch. agilis (from Yakutia) Ch. bafatonicus Ch. behningi

17-33 -

-

5.4 38.1

-

-

1

P

P

Kiknadre et al. (1996)

Up to 80

Petrova (1991) Belyanina and Kolosova (1979) Belyanina (1986) Belyanina (1983), Kerkis et al. (1989)

Ch. bonus

99.6

Ch. bmoknsis

5 -

-

P

-

31.8

1-2

15.00 -

1 -

P P P P P

6.0 1.2

1 1

P, M

Key1 and Hagele (1971)

-

Kiknadze et al. (1987a)

11.0 9.09 2.9 1.5

1 1 1 1

P, M

Key1 and Hagele (1971) Belyanina (1975) Belyanina (1976) Maksimova (1976)

Ch. entis Ch. gr. salinarius Ch. heterodentatus Ch. jonmrtini Ch. mefanotus Ch. nudivenms Ch. plumosaa

-

P

P P

Kerkis et al. (1988) Kiknadze et al. (1996) Kiknadze et al. (1991a) Chubareva (1971) Belyanina (1975) Kiknadze et al. (1996)

(continues)

289

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

Table 24. (Continued) Family, species

Percentage of individuals with B chromosomes

Number of B chromosomes in individuals

Mitotic (M) or polytene

(P)

References

0.77 8.1 3.22

Belyanina (1977) Belyanina (1978a) Belyanina and Mociyash

16.21

Belyanina and Mosiyash

9.1 8 1-8, o n

Petrova etd. (1981) Sigareva (1985) Kiknadze et al. (198713)

(1980) (1980)

average 2.1

Ch. plumosw (different populations)

3.3-12.0 21

Ilyinskaya et af. (1988) Petrova (1991)

1.66 2.27 2.00 3.80 15.40 9.15 1.22 9.61 4.17

Belyanina er al. (1983)

Ilyinskaya and Petrova

(1985) 5.90 4.81 20.80 17-33 Ch. sp.

Kiknadze et al. (1993) Belyanina (1989)

0.97 24-50

Kiknadze et al. (1996)

Ch. sp. Ya3

349

Kiknadze et al. (1996)

Endochironomw tendens

-

Michailova and Gercheva

Ch. sp. Yal

Glyptotendipes gripekoveni

G . paripes

(1982) 8.33

1

P

Belyanina (1975)

15.9 16

-

P P

Miseikoetd. (1971) Miseiko and Minsarinova

-

(1974) 3.57

11.1 4.0 Orthocladiw

-

1 -

P -

Michailova and Petrova (1984), Petrova and Feger ( 1985) Michailova (1987~) Kiknadze et af. (1990) Petrova (1980), Petrova

(1987) (continues)

290

1. F. Zhimulev

Table 24. (Continued) Family, species OrthOCk7diW bipunctellus 0.olivacew 0. rubicundus

Number of Mitotic (M) Percentage of individuals with B chromosomes or polytene (P) B chromosomes in individuals

References

2.2

1

P

Michailova (1982)

3.3

1

P

5

M

Michailova (1985) Michailova (1985)

-

Michailova and Petrova (1989)

Pseudodiamesa gr. branickii

12

1 2

TMassayia frauenfefdi

100

1

P, M

Valkanov and Michailova (1974), Michailova (1976,1989a)

In 5.35% of females and 22.35% of males

1

M

Belcheva and Michailova (1971)

In 17.64%of females and 20.00% of males

1

M

Belcheva and Michailova (1971)

D. albomicans

7.4-26.9 69.1

1-8 1-6

M M

Hatsumi (1987) Ling Fayao and Kitagawa (1991)

D. fivopilosa D. nasuta

-

1 1-8

M

67-95

Brncic (1962) 0. Kitagawa in Ramachandra and Ranganath (1985a), Ramachandra and Ranganath (1985a,b, 1987)

Glossinidae Glossina austeni

-

a12

M

G. morsirans

-

2-7

M

Culicidae Anopheles mulipennis

A. messae

Drosophilidae

M

albomicana

morsirans

hard (1970), Maudlin (1970), Southern et al. (1972), Southern and Pel1 (1973,1974), Davies and Southern (1976, 19771, Amos and Dover (1981) ltard (1966, 1970, 1973), Southern et al. (1973), Davies and Southern (1977) (continues)

29 1

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 24. (Continued)

Family, species

Percentage of individuals with B chromosomes

G. m. submmsitans G. pallidipes

Number of Mitotic (M) B chromosomes or polytene in individuals (P)

3-6

References Pel1 eta!. (1972), Southern and Pel1 (1973) Itard (1970), Maudlin (1970), Amos and Dover (1981)

-

-

loo(?)

7-8(!)

Boyes (1953), Boyes and van Brink (1965)

Sarcophagidae Pseudosarcophaga afinis Simuliidae Cnephia dacorensis

5.76-44.44

1-6

Procunier (197%. 1982)

C . hpponica

-

-

Procunier (197%)

C . mitophih C. saskntchewana

15.0-52.15 -

1-8 -

Procunier (1975b, 1982)

Cnethu crmsa C . chubarevne

34.88 87.6

-

Chubareva (1974) Kachvoryan et al. (1993)

C . djafarovi

78

1-4

C . species

32.26 -

-

C. zakhariensis Eusimulium (Cnetha) costacum

-

2.8

2 2

-

-

39.5 E. crassum E. (Montisimulium) 36.4

1-3 1

inflatum

E. gr. latipes E. vernum E. (Cnetha)

lepnevae

Kachvoryan (1988, 1990), Kachvoryan et al. (1993) Chubareva (1974) Chubareva (1984) Knoz and Chubareva (1974), Chubareva and Petrova (1984) Hunter (1987) Chubareva (1974) Chubareva (1974), Chubareva and Petrova ( 1984) Chubareva, (1974)

32.2 -

1-5 -

27.8

1-4

-

-

Rothfels (1979)

-

-

Chubareva and Perrova (1979)

zakharienze

Memphia borealis Montisimulium

Madahar (19671, Procunier (197%)

Hunter (1987) Kachvoryan and Chubareva (1974), Chubareva (1974), Chubareva and Kachvoryan (1974)

(continues)

292

1. F. Zhirnulev

Table 24. (Continued)

Family, species

Number of Mitotic (M) Percentage of individuals with B chromosomes or polytene B chromosomes in individuals (PI

M.octofiliatum

33.33

M.species

60

Odagmia maxima

59.0

1-5

P, M

0.monticola

35.7

1-3

P

0. obreptanr

33.3

2-3

P

0.omam

-

2

P

22.3 30.42 22.3

1-8 1-8 1-6

P M, P

19.2

1-8

P,M

Chubareva and Petrova (1979, 1984) Chuhareva and Petrova (1984), Chuhareva (1985)

31.3

Chubareva and Ralcheva (1974) Knoz and Chubareva (1974) Knoz and Chubareva (1974) Chubareva and Shcherbakov (1963) Shcherhakov (1965) Shcherbakov (1966h) Shcherbakov and Chubareva (1966) Chubareva and Petrova (1968), Chubareva et al. (1971) Chubareva (1979) Chubareva and Petrova (1984) Chubareva (1985)

1.6 3.33 34.0 -

Chubareva (1974) Chubareva (1974) Chuhareva (1974) Rothfels (1979)

M,P

23.88

0. omam (different populations)

0. tuberosum 0.wariegata

References

Armenia

17.1

1-2

P,M

Bulgaria

61.7

1-7

P, M

Simuliidae unidentified species

-

Simuliurn austeni

22.2

2

P

Chubareva and Petrova (1968), Chuhareva and Ralcheva (1974) Chubareva and Ralcheva (1974) K.H. Rothfels in Dunbar (1966) Knoz and Chubareva (1974) (continues)

293

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation Table 24. (Continued) Percentage of individuals with B chromosomes

Number of Mitotic (M) B chromosomes or polytene in individuals (PI

S. fividum

90.0

-

-

S. latipes

-

-

-

Procunier (197%)

S. mmsitans

56

1-5

-

1-5

P P, M

Chubareva (1974) Kudule (1973,1974, 1976), Kudule and Zitzer (1975)

S. venustum

-

-

-

S. verecundum

8.2

1-3

P

15.6

1 4

P

-

-

-

S. complex wmum

-

-

P

Rothfels et al. (1978) Chubareva and Petrova (1968) Chubareva and Petrova (1968) Rothfels et d.(1978) Brockhouse et al. (1989)

Sulcicnephia ovtshinnikovi Tetisimulium alajense T. condici

76.2

-

-

23.8

1-2

P

-

-

P

Chubareva and Kachvoryan ( 1993)

Bacchn ulicia B. virginio

-

3 8

Eristalis abusivus

-

10-16

M M M

Boyes et al. (1973)

-

M

Boyes and van Brink (1964) Boyes and van Brink (1970) Boyes and van Brink (1970)

Family, species

References Chubareva and Petrova (1984),Chubareva (1985)

Chubareva and Petrova (1984) Chubareva and Petrova (1968)

Syrphidae

E. arbustorum Mabta marquerita

-

7-8

M

Merodon avidus

-

5-6

M

M

Platycheirus afbimnus

Somula decora

-

8

M

Boyes et al. (1973) Boyes and van Brink (1964,1970)

b y e s and van Brink

(1964); not detected subsequently (Boyes and van Brink (1970) Boyes and van Brink (1967, 1970) (continues)

294

1. F. Zhimulev ~~

Table 24. (Catinued)

Family, species

Sphaerophoria sp.

Percentage of individuals with B chromosomes

Number of B chromosomes in individuals

Mitotic (M) or polytene

-

6

M

Boyes and van Brink (1970)

Complexes of microchromosomes

M

Boyes and van Brink (1970)

sphecomyia vespiformis

(PI

References

VoluceUa ekgans

-

35-45

M

Boyes and van Brink (1970)

V. pellucens

-

8

M

Boyes and van Brink (1964, 1970)

V. zonaria

-

About 30

M

Boyes and van Brink 1970

Xylota nemorum

-

About 20

M

Boyes and van Brink (1967)

11

1-2

M

Boyes and Wilkes (1953)

-

0-4

M

Bauer ( 1931)

Tachinidae Ceromasia

auricaudatn Xpulidae Tipuh paludosa

Note. Dash indicates no relevant data were provided in reference(s).

24 and 25). In his studies, Shcherbakov (1965,196613) disclosed higher frequencies in individuals with even numbers of B chromosomes; however, the total number for all these species (Figure 121) does not allow us to infer that individuals with two B chromosomes are in excess. There is seasonal variability in the number of B chromosomes. For example, 23.3% of larvae had the B chromosomes in spring and 61.8% in autumn in one population of Odagmiaomta (Shcherbakov, 1965,1966b). Such variations were not found for other populations and species (Chubareva and Petrova, 1968; Ilyinskaya et al., 1988). It has been reported that the number of B chromosomes in different tissues and different cells of an individual is invariable (Chubareva and Petrova, 1968, 1984; Belyanina et al., 1983; Chubareva, 1984, 1986a). Because supemumerary chromosomes occur in larvae of both sexes-for example, in Cnephia dacotensis, C. ornithophila, Sirnuliurn rnorsitans,Chironornus plurnosus, Cnetha djafaroui, and Drosophila nasuta albornicana (Procunier, 197513; Kudule and Zitzer, 1975; Chubareva and Petrova, 1984; Ilyinskaya and Petrova, 1985; Kachvoryan, 1988;

Table 25. Number of B Chromosomes in Representativesof the Diptera Order Family, species Chironomidae Chironomus gr.

Number of individuals with different chromosome number (1-12)

0

1

2

3

15

6

1

13 33 9 13

9 5 4 9

10 1 10 11

33

36

26

5

5

32

38

21

4

5

6

7

8

9

1 0 1 1 1 2

References Chubareva (1971)

SaliWriUS

Drosophilidae D. albomicana (different strains)

D. nasuta albomicana (different years) Glossinidae Glossinu ausreni Glossina m.

1

4 5

1 2 1

1

3

1

Hatsumi (1987) Ramachandra and Ranganath (1985a) Ramachandra and Ranganath (1985a) Ramachandra and Ranganath (1985a)

1

1

Ramachandra and Ranganath (1985a)

11

24

25

13

4

SouthemandPell(l973)

9

30

26

8

Southern and Pel1 (1973)

2

7

7

5

1

Southem and Pel1 (1973)

1 2

3

morsitnns

1

Glossim m. submorsitans

Simuliidae Cnephiu ducotensis c.omimphila

1085

213

93

13

17

168

67

61

10

15

1

Procunier (1975b) 0

3

Procunier (1975b) (continues)

Table 25. (Continued)

Family, species

Number of individuals with different chromosome number (1-12)

0

Cnetha djafarovi

18

Eusimulium costaturn

35

E. crassurn E. injkztum E. gr. latipes

E. znkharienze

1

2

12

48

4

2

4

5

6

7

8

9

1 0 1 1 1 2

References Kachvoryan (1988)

1

Knoz and Chubareva (1974)

8

2

Chubareva (1974)

28

7

7 21

4

5

2

1

1

249

54

36

4

2

48 9

24

31

6

6

1

3

1

Knoz and Chubareva (1974)

1

1

Knoz and Chubareva (1974)

76 91 219 2 28 22 11

5 22 42

22

13 2 1

6 5

4

(Populations of Leningrad region)

3

394 446 2206 145 287 65 132

5 17 226 2 23 16 16

Chubareva (1974)

55 27

Chubareva (1974)

1

Chubareva (1974). Chubareva and Kachvoryan (1974) Chubareva and Ralcheva (1974)

1

0 2 6

4 7 1

0 0 1

1

2

1

1

1

1 1

Shcherbakov (1965), Shcherbakov (1966b), Chubareva and Petrova ( 1968) Chubareva (1985) Chubareva (1985) Chubareva (1985) Chubareva (1985)

223 184 89 83

31 29 19 24

31 29 19 19

0 . UQriegata Armenia

68

4

10

Bulgaria

193

99

163

10

4 12

Simulium alcrreni

S. morsimns

S. morsimns (different populations) S . verecundum Sulcicnephia ovuhinnikowi

Tetisimulium

14 22 (78.1) (72.0) (86.0) (40.0) 97 144 25 32

(18.1) (20.0) (12.0) (24.0) 9 10

(3.3) (4.0) (2.0) (16.0)

5 2

18

30

7

3

dajense

Note. Data in parentheses are percentages.

Chubareva (1985) Chubareva (1985) Chubareva (1985) Chubareva (1985)

1

24

20

2

2

1

Chubareva and Petrova (1968), Chubareva and Ralcheva (1974) Chubareva and Petrova (1968) Chubareva and Ralcheva (1974) Knoz and Chubareva (1974)

4

(2.0) (16.0)

2 1 22

1

Chubareva (1974) Kudule (1974) Kudule and Zitzer (1975) Kudule and Zitzer (1975) Kudule and Zitzer (1975) Chubareva and Petrova (1968) Chubareva and Petrova (1984) Chubareva and Petrova (1968)

I. F. Zhimulev

298

0

1

2

3

4

5

6

7

8

9

1

0

1

1

1

2

Figure 121. Total distribution of the occurrence of individuals with different numbers of B chromosomes: a synopsis of information given in Table 25. Abscissa, number of B chromosomes; ordinate, number of events.

Ramachandra and Ranganath, 1987)-the conclusion was made that their occurrence does not depend on sex either. However, there are descriptions of at least three types of mosaic distribution of the B chromosomes: (1) their presence in dividing cells and absence in polytene chromosomes in the representatives of the Glossinidae family (Glossinaausteni, G. morsitaru morsitans, G. m. submorsimns), in Dosophila nasuta albomicana (Ramachandra and Ranganath, 1985b,1987), and in Orthocladius rubicundus (Michailova, 1985); (2) mosaic representation in dividing cells, for example, in the testicles and in salivary glands of Ch. melanotus (Key1 and Hagele, 1971); and ( 3 ) mosaic representation of polytene supemumeraries in different salivary gland cells, for example, in larvae of the Kurshskaya population of Ch. plumosus (Figure 122). A similar mosaicism has been observed for Pseudodiamesa gr. branickii (Michailova and Petrova, 1989).

Polytene Chromosomes, Heterochromatin, and Positlon Effect Variegation

1 2

3

4

5 6

299

7 8 9 1011 1 2 1 3 1 4 1 5

Figure 122. Occurrence frequencies of nuclei with different numbers of B chromosomes in a single individual of Ch.

plumosus. Abscissa, number of B chromosomes in the nucleus; ordinate, number of nuclei. After Ilyinskaya and Petrova (1985).

Pairing-related data are also in favor of homology of the B chromosomes. In Simuliidae, the B chromosomes of the same nucleus pair with together, with a ring forming from them in some cases (Shcherbakov, 1966a; Procunier, 1975b). These chromosomes consistently lie side by side at metaphase, “displaying a tendency to somatic pairing” (Shcherbakov and Chubareva, 1966). Globules of heteropycnotic material occasionally form from the B chromosomes (Shcherbakov, 1967). In the interphase nuclei of the cells of larvae with microchromosomes, a heterochromatic body causing the nuclear membrane to bulge and bearing a resemblance to microchromosomes is noticeable. This body is always single irrespective of the number of B chromosomes. Based on this feature, microchromosoma1 karyotypes are readily identifiable by studying only interphase nuclei (Shcherbakov and Chubareva, 1966; Chubareva, 1985). In Chironomus plumosus, the B chromosomes lie at the nuclear membrane (Ilyinskaya and Petrova, 1985). In the chironomid Thalassomiyafiauenfeldi, the B chromosome is also located at the periphery of the nucleus, close to the surface (Michailova, 1976). In Glossina austeni and G.morsitans, the B chromosomes form an enormous, compact, heavily staining body at meiosis. The Y chromosome is also included in this body in the former species. When this mass falls apart into distinct elements, the B chromosomes usually lie side by side, but they do not pair with one another (Southern and Pell, 1973; Southern et al., 1973). In Phryne cincta, the supernumerary chromosomes in salivary gland nuclei fuse together or with other a-heterochromatic regions of the autosomes, usually with the proximal segments of autosome 4.When not uniting, they are small and very different in size (Wolf, 1962, 1970). These data may indirectly provide evidence for homology of the B chromosomes included in a nucleus. In contrast, it was shown that the B chromosomes

300

1. F. Zhimulev

of a single individual differ in size and morphology. In Sulcicnephia outshinnikoui, two sorts of supernumerarieswere found, disk-shaped with a block of compact heterochromatin and amorphic, which can occur in all combinations in a nucleus (Chubareva and Petrova, 1984). Numerous instances are known in which supernumerary chromosomes pair with single regions of the chromosomes of the regular set. Thus the B chromosomes of Chironomus plumosus frequently pair with the centromeric region of the fourth chromosome (Key1and Hagele, 1971; Sigareva, 1985).The contact frequencies in different populations vary in the range of 19.85-69.53%. The B chromosome was associated with a group of other chromosomes, which included the fourth chromosome, in another 6.24-22.39% of nuclei (Ilyinskaya and Petrova, 1985; Petrova, 1986). This suggested that supernumerary B chromosomes might have arisen from the fourth chromosome, the source of breaks in the centromeric region in this species. In Orthocladius bipunctellus, the B chromosome in the salivary gland cells is ectopically associated with the first and second chromosomes (Michailova, 1982). In Ch. heterodentatus, another chironomid species, the supernumerary microchromosomeswere found to be associated with the distal end of one of the long chromosomes in several salivary gland cells (Belyanina, 1975). In Cn. dacotensis, the B chromosomesestablish contacts with the regions of the second and third chromosomes (Procunier, 1975b). The contacts are reminiscent of the ectopic pairing discribed in Section VII,C, and they indicate that homology may exist between the material of the chromosomal elements at the contact sites. It has been reported that polytene B chromosomes are not identical in two related midge species, Cnephia dacotensis and C. urnitophilia (Procunier, 1975b). The relation of the B chromosomes to heterochromatin is ambiguous. At any rate, they cannot be called a priori heterochromatic. In some cases (e.g., Phryne cincta), it was demonstrated that the supernumerary chromosomes are tightly compacted and they strongly resemble a-heterochromatin, although polytenizing in salivary gland cells. In other cases, the B chromosomes are represented by a reticular structureless block presumably composed of P-heterochromatin. In many species, the B chromosomes have the appearance of polytene chromosomes showing a distinctive pattern of banding, yet containing many large blocks of compact and heavily staining material (see Figure 120). Finally, a type of B chromosome that occurs in metaphase spreads and is not represented in cells with polytene chromosomes is distinguishable. The B chromosomes of plants polytenize infrequently. Among species of the Rhinuntus genus, supernumerary chromosomes nevertheless undergo polytenization. In Rh. alectrolophus,polytene chromosomes characteristic of plants are formed (for review, see Zhimulev, 199213, 1996). In other species, the number of

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

30 1

B chromosomes increases with each successive endoreplication cycle because the products of endomitosis separate into distinct chromosomes (Tschermak-Woess and Hasitschka-Jenschke, 1963; Tschermak-Woess, 1967). In Drosophila nasutu albomicana, these chromosomes show moderate C+-banding (i.e., weaker than Cheterochromatin in the A chromosomes) in metaphase. According to estimates, the amount of heterochromatin in individuals with eight B chromosomes in the nucleus constituted 46% of the total chromatin amount in males and 39% in females, when the B chromosomes are missing, the amounts are 34% and 27%, respectively. Silver staining (N-banding) of these chromosomes was also positive (Ramachandra and Ranganath, 1986a, 1987). In D. albomicana, staining of the metaphase chromosomes by the Giemsa procedure revealed only a small block of C+-heterochromatin in the B chromosomes in some cases (see Figure 120), while they were entirely C+-positive in other cases (Ling Fayao and Kitagawa, 1991). In Glossinidae, the unpolytenized supernumerary chromosomes show C+-banding (Davies and Southern, 1976), and satellite DNA is revealed in them by in situ hybridization (Amos and Dover, 1981). In Chironomus plumosus, C-banding is detected in the centromeric band of the B chromosome (Hagele, 1977a). According to Sigareva’s data (1989, about half of the polytene B chromosomes show positive C-banding in this species. Staining intensity of the heterochromatic part of the supernumerary chromosomes represented by a whole block or separate clumps was weaker than in centromeric regions of the other chromosomes. It was found that the heavily staining segments of the B chromosomes of Ch. plumosus and Ch. melanotus are late replicating, like heterochromatin (Key1 and Hagele, 1971). Finally, the heterocyclicity of the heterochromatin of the B chromosome should be noted. The supernumerary chromosomes remain in a compacted state longer, and they pass to the poles earlier, than the A chromosomes at anaphase (Michailova, 1976; Chubareva, 1985). It remains unclear whether B chromosomes affect the timing of physiological functions. No differences were found in the external morphological features between individuals with and without supernumerary chromosomes (Wolf, 1962, 1970; Shcherbakov, 1968; Chubareva and Petrova, 1968, 1984; Chubareva, 1974; Belyanina, 1975, 1978a; Belyanina and Kolosova, 1979; Petrova et al., 1981; Michailova, 1982; Ilyinskaya and Petrova, 1985). The different possible effects of the supernumeraries on the various aspects of the vital functions have been the subject of much discussion: 1. It is Chubareva’s view (1977a) that pairing of the homologous chromosomes occurs in the midge forms with the B chromosome karyotypes.

2. In the presence of the B chromosomes in the karyological pool of chironomids and simuliids, inversion polymorphism is slight, and it is completely

302

1.

F. Zhimulev

~~

lacking when their frequencies are high. In contrast, in species with high inversion polymorphism, no supernumerary chromosomes were detected (Belyanina, 1986; Chubareva, 1986a). In populations of Cnetha djufurovi, “additional nucleoli” were found only in individuals possessing B chromosomes (Kachvoryan, 1990). In the simuliids, the frequencies of the B chromosomes are positively correlated with the extent to which reservoirs are contaminated (Chubareva and Petrova, 1968). Individuals carrying the B chromosomes are more numerous in localities where rivers have copious streams and ecological niches are more diverse than in those where brooklets are stagnant and the streams of rivulets are small (Chubareva, 1974). B chromosome-bearing individuals of Odagmiu m t u are less widespread in a woody locality compared to denser populated human habitats (Chubareva, 1986a). It has been suggested that the B chromosomes also widen the range of genotypic variability of the population and of the whole species. Increasing polymorphism for the number and frequency of the accessory chromosomes may provide the population with better chances of surviving under extreme conditions (Chubareva, 1974,1985; Chubareva and Kachvoryan, 1974; Chubareva and Ralcheva, 1974; Belyanina, 1986).The higher occurrence frequencies of supernumeraries in contaminated water reservoirs may be differently explained if we accept the possibility that a heavily contaminated environment, being more mutagenic, causes more frequent breakages of the hereditary machinery in its inhabitants. Finally, there appeared to be no way in which the B chromosomes might have arisen in Ch. plumosus, inasmuch larvae with the B chromosomes also occurred in a relatively clean water reservoir (Ilyinskaya and Petrova, 1985). In Cnephiadacotensis, females carrying supernumerary chromosomes are, as a rule, somewhat smaller than males, and those possessing the supernumeraries develop slower during the cycle from egg to early pupa (Procunier, 1975b, 1982). In Sulcicnephiu ovtshinnikooi, females with B chromosomes occur 1.5 times more frequently than males carrying them (Chubareva and Petrova, 1984). There is evidence indicating that the B chromosomes can adversely affect total fitness. An attempt was made to determine population fitness in strains of Drosophila nasutu albomicuna with and without B chromosomes. Development rate and fitness from egg to adult, as well as total adaptivity, were estimated in the two strains at 18, 22, and 26°C. It was demonstrated that (1) eggs of flies with B chromosomes develop faster at 26”C, while fitness and adaptivity were highest at 22”C, (2) strains without the B chromosomesshow the highest development rate, fitness, and adaptivity at 20, 18, and 22”C, respectively; and (3) adult eclosion is

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

303

delayed and adaptivity is reduced in the two strains at 18°C.Thus, whether the B chromosomes were present or not, the ecological optimum temperature for the two strains of D. n. albomicana was in the range of 22°C (Ramachandra and Ranganath, 198613). Flies of this species not carrying the B chromosomes live longer, are more fit, and lay more eggs than those carrying it in mixed populations (Ramachandra and Ranganath, 1986a). The problem of the functional significance of the B chromosomes should be viewed broadly. The data on position effect variegation (see later sections XI-XVII) show that variation in the amount of heterochromatin has a strong effect on gene activity. Since most supernumerary chromosomes are heterochromatic, it may be expected that variation in the number of the B chromosomes would have a great influence on genome activity, and this might be demonstrated in an appropriate experimental model. Having established an inverse relation between the number of B chromosomes and nuclear organizer expression in the A chromosome set, Procunier (1982) obtained support for this line of reasoning. Little is known about the origin of supernumerary chromosomes not only in Diptera but also in broad outlines. According to White's (1959) concept, the B chromosomes operate as the major mechanism in increasing chromosome number in the set. A small additional chromosome element with a centromere is formed first; then, if it is stabilized, other parts of the chromosomes can be translocated on it. Chubareva (1985) holds the view that the B chromosomes have undoubtedly derived from the chromosomes of the regular set and, as believed, from the sex chromosomes (Boyes et al., 1973). A hypothetical variant of the origin of supernumerary chromosomes in the tsetse fly is depicted in Figure 123. As shown in Figure 123 it is conceivable that (a) parts of a regular A

a

stage 1

Figure 123. A hypothetical scheme of the two-step formation of the supernumerary chromosome in Glossinkfa (see text for explanation). Reprinted by permission from Amos and Dover (1981).

304

I. F. Zhlrnulev

chromosome and of a centromere have duplicated for whatever reason (stage 1). This may be followed by (b) transposition of minority sequences of the A chromosome and duplication, presumably as a consequence of interchromosome exchange; amplification (c) of transposed telomeric sequences of the A chromosome; and duplication (e) and formation of satellites (stage 2). If the already existing telomeric sequences are amplified, transposition of minority repeats (b) is no longer needed, and heterochromatization (d) of the sequences composing the duplication (the B chromosome only) follows. Another variant of supernumerary A chromosomes is conceivable: arising as the result of deletions, they lose the greater part of heterochromatin and convert into B chromosomes. In a general way, it can be also imagined that the bulk of the heterochromatin can be removed by a deletion from accessory A chromosomes resulting from trisomy (Ved Brat and Rai, 1975) and that such accessories can be transformed into B chromosomes. According to interesting speculations of Hackstein et al. (1996) the Y chromosomes of most Drosophila species share substantial affinities with B chromosomes. The Y chromosomes evolved as specialized supernumeraries similar to classical B chromosomes.

X. HETEROCHROMATIN OF CHROMOSOMES RESTRICTED TO GERMLINE CELLS A. Chromosomes of germline cells As mentioned in Section V, germline cells contain-in addition to the regular, or somatic, S set of chromosomes-a set of K chromosomes eliminated from the presumptive cells at the early cleavage divisions in representatives of the Orthocladiinae (Chironomidae) family. Irradiation of newly hatched larvae can induce material of the K chromosomes to transpose to the S chromosomes and, in this way, to achieve their polytenization (Bauer, 1970; Staiber and Thudium, 1986; Staiber, 1991~). In the polytene chromosomes of salivary glands, the translocated parts of the K chromosomes have the appearance of large vacuolated globules of chromatin (Figure 124) showing all the features of a-heterochromatin, such as heavy stainability presumably due to high compaction and absence of banding pattern (Bauer, 1970; Hagele, 1980; Staiber and Thudium, 1986). In Smittia parthenogenetica, this insertion of a-heterochromatin stains with Giemsa and quinacrine; it is the latest Figure 124. Morphology (a-0, C- and Q-banding (g and h) and late termination of replication (i) in transpositions of heterochromatin from the K to the S chromosomesin Acricotopus luc i d (a-e) ~ and Smittia parthenogenetica (f-i). Arrows in figures indicate translocated heterochromatin. N, nucleolus; BRI and BR2, Balbiani rings 1 and 2; C and arrowhead, centromere. (a-e) after Staiber and Thudium (1986); (f-i) reprinted by permission from Hagele (1980).

306

1. F. Zhlrnulev

replicating element of the chromosomes not actively engaged in transcription (see Figure 124). The pattern obtained by N-staining is more complex: the bulk of the globule is N-, and an N’ body is identified in the center (Hagele, 1980). There is evidence of a banding pattern exhibited by the K chromosomes. When transpositions from the K to the S chromosomes are induced in A. lucidus, small segments of the chromosomes containing both a block of heterochromatin and banded fragments, which somatically pair with the chromosomes of the S set, are seen in the salivary gland cells (Staiber and Thudium, 1986; Staiber, 1991b,c). Additional chromosomes that also contain a small block of heterochromatin 1.5-2.0 Fm long and composed of up to 70 bands also pair with the polytene chromosomes of the S set. In contrast to induced transpositions, spontaneous supernumerary chromosomes are seen to variegate in organ cells, for example, in 27,4,4,3, 19 and 8 cells of 70-80 salivary gland cells (Staiber, 1987).

B. Sex-determining factors Chironomids usually have no visible sex chromosomes, and karyotypes differ in inversions only when there are sex differencesbetween individuals (Beermann, 1955). However, sex-linked variation in karyotype is nevertheless found in certain species. For example, in Polypedilum nubif0 (Diptera, Nematocera), the heterochromatic fragment at the end of the fourth chromosome was found to be heteromorphic. Associated findings were female heterozygosity for the block and males without the block (Martin, 1966). In Telmtoscopus ulbigunctutus (Diptera, Psychodidae), heteromorphism was found for the large block of heterochromatin in the 7C region of the fourth chromosome (Figure 125); it is noteworthy that males were heterozygous for the block and females were homozygous for it (Amabis, 1977). The former provide evidence for female, and the latter for male, heterogamety. Sex-related polymorphism for the size of the block of heterochromatin was also revealed for certain species of Chironomidae (Rosin and Fischer, 1972) and Culicidae (Rothfels and Mason, 1975).

XI. CHANGES IN EXPRESSION OF GENES DEPENDENT ON THEIR POSITION IN THE GENOME Several types of changes in the expression of genes dependent on their position in the genome are known. From Sturtevant’s early investigations in Drosophih, it became apparent that two Bur alleles had a stronger effect on the mutant phenotype when present on the same chromosome than on different ones. Sturtevant has suggested that the function of genes can be differently manifested dependent on their position with respect to neighboring genes, and he suggested calling this phenomenon the position effect (Sturtevant, 1925, 1928). At present, the term “position effect’’ (Ephrussi and Sutton, 1944) is

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

307

Figure 125. Heteromorphism of the heterochromatic block in the 7C region of the fourth chromosome of Telmurorcopus albipunctatw, presumably due to sex. a, b, c, homozygote for the thin band and heterozygote and homozygote for the thick band, respectively. Reprinted by permission from Amabis (1977).

broadly applied to all cases of variation in the activity of genes that occur when their position in the genome is altered. The position effect may be stable (Lewis, 1950) when its manifestation changes little from one somatic cell to another or within a single genetic strain, and when it is affected by neither environmental nor genetic factors. A stable position effect arises when both chromosome breaks involved in rearrangements are mapped to euchromatin. Position effect of the stable type has been observed for the Bar (Sturtevant, 1925; Dobzhansky, 1936; Muller et al., 1936; Dubinin and Volotov, 1940; Rapoport, 1940; Sutton, 1943; Panshin, 1992) and the Star-asteroid (Lewis, 1945, 1950) loci. The various instances of the stable-type position effect are readily accounted for by intragenic shuffling of material produced by small rearrangements. The following pivotal observations suggest explanations for stable position effect. The DNA in the transposon-generated Bar (B) breakpoint causing the B mutant phenotype has been cloned; the results were interpreted as indicating that the B phenotype results from juxtaposition of the coding regions of the B locus to new controlling elements (Tsubota et al., 1989). The mutationfacet-strawbeny (faswb)may provide another case of stable position effect. It is caused by a tiny deletion (of approximately 0.8 kb) at the 5' end of the Notch locus, continuously disturbing transcription (Welshons and Welshons, 1985, 1986).A case in point also appears to be ocellikss, a mutation induced by a small inversion with a

308 ~~

1. F. Zhlmulev ~

breakpoint 1-3 kb upstream of the s36-1 and s38-1 chorionic genes, adjacent to the chromosomal sequence undergoing amplification; as a consequence, copy number is greatly altered in the sequence (Spradling and Mahowald, 1981). Lewis (1950) cautioned that it is difficult to make a distinction between position effect and intragenic mutation in analysis of stable changes in gene function. Position effect of the stable type has scarcely been covered in reviews. It is of no relevance to the functional role of heterochromatin, the subject matter of this book. Cases of changes in puff activity during the emergence of chromosomal rearrangements are known: larvae with a subterminal inversion in the AB region of the first chromosome were found in a natural population of Acricotopus lucidus. An additional Balbiani ring (BR) formed in the middle of the inverted region; interestingly, it had the appearance of a large puff in heterozygotes that was the shape and size of a BR in homozygotes for the inversion (Mechelke, 1960).In the same species, the BR was derived from a part of the centromeric heterochromatic band broken by an inversion, which transposed it to a euchromatic part not associated with heterochromatin (Staiber, 1982). In D. subobscura,two puffs, 61AC and 67AB, lying at the end of an inversion are strongly inactivated compared to the same puffs in the strain without the inversion. Suppression of the activity of these puffs is not due to heterochromatin (de Frutos et al., 1987). In P-element transposons resulting from transformations and containing various genes, the activity of the gene varies in a manner dependent on its new location site (e.g., Goldberg et al., 1983; Scholnick et al., 1983; Spradling and Rubin, 1983; de Cicco and Spradling, 1984; Gehring et al., 1984; Hazelrigg et al., 1984; Krumm et al., 1985; Levis et al., 1985; Daniels et al., 1986; Karpen e t al., 1988; Wilson et al., 1990; Dutton and Chovnick, 1991). For example, among 36 strains of Drosophih containing transposons comprising the rosy+ gene, the strains differ almost fivefold in the specific activity of the xanthine dehydrogenase encoded by the rosy gene (Spradling and Rubin, 1983). All these examples illustrate position effects of the stable type. Transvection is another phenomenon related to change in gene expression dependent on position in nuclear volume, particularly on pairing of homologous chromosomes in the interphase nucleus (Lewis, 1954;Jack and Judd, 1979; Gelbart and Wu, 1982; Judd, 1988; Ashbumer, 1989; Tartof and Henikoff, 1991; Henikoff et al., 1995; Milot e t al., 1996). By the end of the 1920s and the beginning of the 1930s, the numerous papers of Demerec, Sturtevant, Patterson, Casteel, Muller, Gowen, and Gay, concerned mainly with mosaicism of the eyes of Drosophih, stirred interest unabated for years (see references in Noujdin, 1935).Mosaicism was accounted for by causes including high gene mutability and gene loss through successive cell divisions. In 1930, Muller disclosed that, in Drosophila heterozygous for the normal and mutant alleles of the w+ gene, a white sector appears in a normal red background in the eye composed of 700-800 ommatidia of the same type; that is, the mutant phe-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

309

notype is manifested in a part of the cells of w/+ heterozygotes. Somatic instability of gene action arising in association with a chromosomal rearrangement and transferring a gene to heterochromatin, was given the name “eversportingdisplacement” by Muller (1930,1932), Schultz (1936) termed it “variegation.”Dobzhansky (1936) has called it position effect; that is, he applied the term used for changes of the stable type. Finally, Lewis (1950) introduced the term “position effect variegation.” The mysteries of position effect variegation have long included debatable issues. Lewis (1950) thought that the position effect is “fundamental in genetic theory, shedding light on the organization of the eukaryotic chromosome.” Expressing the alternative view, Crick (1979) preferred not to consider the event of position effect variegation as an extraordinary or incomprehensible consequence of chromosome structure as a whole. The literature concerned with analysis of position effect of precisely this type is voluminous (for review, see Kerkis, 1934; Bauer, 1937; Muller, 1941; Belgovsky, 1944,1946; Goldschmidt, 1946; Lewis, 1950; Hannah, 1951; Baker, 1968; Zuckerkandl, 1974; Birstein, 1976; Spofford, 1976; Becker, 1978; Lima-de-Faria, 1986; Prokofyeva-Belgovskaya, 1986; John, 1988; Eissenberg, 1989; Henikoff, 1990, 1994, 1995a,b, 1996a,b; Spradling and Karpen, 1990; Tartof and Bremer, 1990; Belyaeva and Zhimulev, 1991b; Grigliatti, 1991; Reuter and Spierer, 1992; Zhimulev, 1992a, 1993; Henikoff et al., 1993; Spradling, 1993; Karpen, 1994; Lohe and Hilliker, 1995; Weiler and Wakimoto, 1995).These phenomena and issues are considered in the next sections. Inactivation of repeated white genes in P-transposons is a position effect of another kind. Arrays of three or more transgenesproduced phenotypes similar to classical heterochromatin-inducedposition effect variegation. This w+ inactivation is affected by PEV modifiers (Dorer and Henikoff, 1994). Partial inactivation of genes transferred by a translocation to the inactivated mammalian X chromosome also stands close to the group of position effects (for review, see Baker, 1968, Zakharov, 1968; Eicher, 1970; Cattanach, 1974; Riggs, 1975; Birstein, 1976; Gartler and Riggs, 1983; Zakiyan and Nesterova, 1992; Nesterova and Zakiyan, 1994; Willard, 1996). From the previous analysis, it follows that changes in the activity of genes brought about by changes in their position in the genome may be differently caused and manifested. Hence clear-cut criteria are needed to distinguish one type of position effect from another.

XII. GENETIC INACTIVATION UNDER POSITION EFFECT VARIEGATION A. Features of position effect variegation The variegated phenotype is manifestated by the appearance of qualitatively different cells of the same genotype. Groups of cells with the mutant phenotype can

3 10 ~~

1.

F. Zhirnulev

~

arise on the background of a normal phenotype, this leads to mosaic expression of the gene. However, the mutant phenotype can result from genetic events irrelevant to position effect itself. For example, two “mosaics” for eye color are known. The genetic structure is different in distinct parts of the eye in one of the mosaics. Differences can arise at early stages of development due to (1) mutations of the normal allele, (2) loss of the chromosome with the normal allele, or (3)somatic crossing over. In the other mosaic, phenotypically different parts of the eye are of the same genotype, and this is due primarily either to incomplete expression of the normal dominant allele, or to a chromosomal rearrangement placing a gene into a heterochromatic position (i.e., position effect variegation) (Becker, 1966). The other causes include mutational instability of genes (Demerec and Slizynska, 1937) and gene interaction (white’ with other genes; Oster, 1957). The causes of mosaicism are unclear in some other cases, and what is then known is that mosaicism is not associated with chromosomal rearrangements (Muller, 1946). For this reason, variegation observed as obvious phenotypic mosaicism has been referred to as real position effect variegation provided that proofs (or features) were adduced (Baker, 1963,1968; Becker 1966,1978; Spofford, 1976).There has been subjective bias dominated by the effort to demonstrate that all the cells are of the same genotype and that only gene activity is altered in cells expressing the mutant phenotype. The criteria establishing these position effects vary. The list below surveys the cogent criteria. Muller (1930, 1935, 1938a, 1941), Patterson and Painter (Patterson and Painter, 1931; Patterson, 1932a,b, 1933), and later other investigators (Offermann, 1935; Sakharov, 1935, 1936; Dubinin, 1936; Schultz, 1936; Demerec and Slizynska, 1937; Hannah, 1949; among many others) demonstrated that position effect variegation is consistently associated with a chromosomal rearrangement whose one breakpoint lies in centromeric heterochromatin and the other in euchromatin nearby the gene showing mosaic expression. Exceptions to this rule are (a) the genes normally mapped to heterochromatin, whose position effect arises when translocated to euchromatin; and (b) rearrangements between euchromatic regions. These two exceptions are considered in greater detail in Sections XV and XVI,B, respectively. Gene inactivation takes place in a rearranged chromosome (Muller, 1930), with the normal alleles losing their dominance (Dobzhansky and Sturtevant, 1932). According to the proposed notations (Stem and Heidenthal, 1944; Stem et al., 1946a,b; Lewis, 1950; Baker, 1965c, 1968), the symbol R( +) or R(g+) is used to designate the chromosome with rearrangement ( R ) causing position effect and containing the normal allele of the gene (g+), and the symbol R( )/g or R(g+)/gis used for heterozygotes under study. Lengthy genotype notations are used, such as bp g+/bp+g, where bp is the rearrange-

+

Polytene Chromosomes, Heterochromatin, and Posttion Effect Variegation

3 11

ment breakpoint (Spofford, 1976). Spofford regards the expressed mutant phenotype in the bp g+/bp+g configuration (or R(g+)/g) as the basis of the proof of the position-effect nature of variegation, and she calls it cis-dominance. Exceptions to this rule are very rare, and they concern only the dominant R(g)/g+ position effect (see Section XV). The normal phenotype is restored in R(G)/G+ heterozygotes under position effect for dominant mutations. For example, in R(Sb)/Sb+ flies, in the case of absence of position effect the bristles are shortened, which is observed in the case when the Sb mutation is expressed. Under a strong position effect, and hence an inactivation of the dominant mutant allele, bristle length tends to be restored to normal (Moore et al., 1981, 1983; Sinclair et d., 1989). The normal morphology of the eye is similarly restored in R(Bs)/B+ heterozygotes (Brosseau, 1960b). 3. Many genes adjacent to the breakpoint can be inactivated in the chromosome bearing the R(g+) rearrangement. The phenomenon has been termed “spreading effect.” The inactivaing effect of heterochromatin diminishes with increasing distance of the gene from the breakpoint (for greater detail, see Section XII,F). 4. A gene is not lost upon inactivation; instead, its state is altered. Evidence for this came from experiments. As a result of crossing over between the hairy locus of the R(h+)/h translocation+induced breakpoint, Dubinin and Sidorov (1935) generated two chromosomes, one with the normal allele h’ and the other with R(h). Similar results were obtained for the curled gene (Panshin, 19351, the white gene (Judd, 1955), and the peach locus in D. oirilis (Baker, 1954). Reversion of chromosomal rearrangements-transposing a gene from the neighborhood of heterochromatin to a new euchromatin positionreactivates the gene, and position effect is completely eliminated (Gruneberg, 1936, 1937; Emmens, 1937a; Panshin, 1938; Kaufmann, 1942; Hinton, 1950; Novitski, 1961). These data show that, before the reversion has occurred, a normal allele of the gene was present in the rearranged chromosome. Similar results were obtained in experiments with insertion of a transposon generated via P-element-mediated transformation. When a transposon is inserted into heterochromatic regions, the genes contained in the transposon become inactivated. Mobilization of a transposon and its removal from heterochromatin restores gene activity (see Section XVI,B). 5. There is proof that the extent of genetic inactivation under position effect can change because it is dependent on various external and internal factors. Addition of heterochromatin of the Y chromosome reduces the amount of variegation and restores the normal phenotype, while removal of the Y induces reversion to the mutant phenotype (see Section XII1,B). Likewise, an increase in developmental temperature leads to partial normalization of the

312

I. F. Zhimuleu

phenotype and its decrease results in enhancement of mosaicism. Genetic and chemical modifiers are also known (see Section XIII). These and other features of position effect variegation are considered in this section. Other events, frequently also referred to as position effects (e.g., Goldschmidt et al., 1939; Raffel and Muller, 1940; Ratner and Furman, 1978) but not showing the usual mosaic position effect variegation, are not dealt with here. For example, two closely linked unstable mutations, singed and club wings, were found to revert together in the X chromosome isolated from a wild population. It was demonstrated that the frequency of the mutant phenotype clw decreased at a lower temperature (18°C) while clw expression normalized at a higher temperature (25°C). However, the effect of the Y chromosome was entirely different. The mutant expression of clw is 1.5-2.0 times more frequent in XY than XO males. A paternal effect of the Y chromosome was established: the presence of the paternal Y, together with the clw in the genome, increased the frequency of the mutant expression of the character in offspring (Zakharov and Golubovsky, 1980).

6. Genes affected by position effect variegation Analysis of the available data demonstrated that actually any gene mapped to any part of the Dosophila genome can be subject to inactivation, when brought into association with heterochromatin. The following group of genes subject to position effect can be tentatively distinguished.

1. Genes controlling development of phenotypic characters These genes include those controlling color of cuticule, bristles, wings, color and structure of eyes, and position and structure of wings (Table 26). Obviously, this mosaicism is not restricted to particular genes, organs, or cells. The ebony gene has long been regarded as an exception to the general rule, since attempts to observe position effect in rearrangements translocating it to heterochromatin were unsuccessful (Brosseau, 1970; Spofford, 1976). However, such rearrangements were eventually generated (Henikoff, 1980). Lewis (1950) takes the view that mosaicism is most readily observed when, first, the expression of the gene is autonomous (i.e., the product is synthesized in a given cell, not transported from other cells), and, second, a great number of cells of the same type are present. For this reason, few genes are suitable for detailed analysis. The white+ gene has been the most extensively used. Variegation of the white locus is very distinctive, showing light or white patches on a red background within the eye. The w+ gene is the most widely used in research to the present time. Hence it is expedient to describe the compound eye of Drosophila in greater detail

Table 26. Genes Controlling the Development of Morphological Traits and Position Effect Variegation in D. melunogastes

Gene, symbol

Gene position on genetic and cytological maps (Lindsleyand Zimm, 1992)

Controlled trait

References

abb, abbreviated

2: 105.5; 59E24OB10

ac, achaete

X: 0.0: 1B1-2

Size of body and bristles Microchaetae and bristles

Schultz and Dobzhansky (1934) Noujdin (1935,1944), Crew and Lamy (1940), Brunstrom (1955) Spofford (1982)

B, Bar bi, bifid bw,brown

X: 57.0; 16A1-2 X: 6.9; 4C5-6

Eye shape Wing venation

Belgovsky (1938), Dubinin and Volotov (1940) Demerec (1940, 1941a), Sutton (1943)

2: 104.5; 59D1-2 to E2-3; 59136-1 1 (Belyaeva et d., 1997)

Eye color

Glass (1933), Schultz and Dobzhansky (1934), Dubinin and Heptner (1935), Oster (1954)

cW, canopy wing

X: 2.5 (between w and vt) 3: 50.0; 86Dl-Q X: 13.7; 5B X; 2B7-11 X: 13.6; 5A-B X 4.0; 3D5 X: 0.3; 2B6-7

Shape of wing Shape of wings and bristles

Sinclair et al. (1989)

cu, curkd cv, cross-veinkss cwi, crumpkd wings cx, curkx dm, diminutive

dor, deep orange

Wing venation

Panshin (1935) Demerec (1940)

Structure of wings Shape of wings

Zhimulev et al. (1986), Demakova et d.(1988) Demerec (1940)

Size of body and bristles

Demerec (1940,1941a, b), Ratty (1954), Spofford (1973) Bridges and Brehme (1944), Schultz in Gvozdev et al. (1973), Clancy (1964), Alatortsev et al. (1982), Zhimulev er al. (1986)

Eye color, wing shape (E. S. Belyaeva et al., 1982)

e , ebony

2: 13.0; 25A1-2 3: 70.7; 93D2-6

See Lindsley and Zimm (1992) Kotarski et al. (1983) Body color Henikoff ( 1980)

ec, echinus

X: 5.5; 3E8 or 3F1

Facet size

4, dumpy

Demerec (1940, 1941a) (continues)

Table 26. (Continued)

Gene, symbol

Gene position on genetic and cytological maps (Lindsley and Zimm, 1992)

References

Controlled trait

f>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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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)

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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,

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

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-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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)

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Poiytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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)

Polytene Chromosomes, Heterochromatin,and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

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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-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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Hilliker (1975); and (3) Wakimoto and Hearn (1990).

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

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

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).

N

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

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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.

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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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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).

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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-

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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

.

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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-

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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-

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9.

10.

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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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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-

382

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.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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)

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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 ~

~~

-~

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~________

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

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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).

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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

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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-

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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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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).

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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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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-

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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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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

408

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

410

1. F. Zhlmulev

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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F. Zhimulev

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).

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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).

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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.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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“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-

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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 ~

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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,

_

_

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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

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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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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-

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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.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

COEXTENSIVE MODEL M,

S'S'S'

s s s

BOUNDARY MODEL N

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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

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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).

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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-

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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

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

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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

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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).

Polytene Chromosomes, Hetarochromatin, and Position Effect Variegation

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.

440

1. F. Zhimulev

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:

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

441

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,

442

1. F. Zhlmulsv

“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.

Polytene Chromosomes, Heterochromatin, and Position Effect Variegation

443

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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