Polytene Chromosomes: a General Model for the Eucaryotic Interphase State

OOZO-7322(95)00023-2

POLYTENE CHROMOSOMES: A GENERAL MODEL FOR THE EUCARYOTIC INTERPHASE STATE

Horst Kress

Institut fur Genetik, Freie Universitat Berlin, Arnimallee 7, 14 195 Berlin, Germany

(Accepted 7 November 1995)

Abstract-The euchromatic structures of insect polytene chromosomes represent an amplified version of the chromomeric organization in diploid chromatin. As such, they are an excellent model system for studying structure/function relationships of eucaryotic interphase chromosomes. Polytenization is accomplished by replication patterns that are different for euchromatic and heterochromatic chromatin and also seem to depend on the functional state of chromatin. In Diptera, polytene chromosomes are characterized by genetically determined discontinuities of DNA packing in bands and interbands that are modified by functional demands. The cytological visualization of proteins that are associated with the compaction and decondensation of chromatin, allows the analysis of the temporal and spatial dynamics of DNA/protein interactions in the context of structure, transcription, and the processing of RNA. Ribonucleoprotein particles may be followed on their way via the nuclear matrix through the pores of the nuclear envelope to the cytoplasmic compartment. Thus, polytene nuclei provide unique opportunities for studying the flow of genetic information from the site of storage to the site of action, i.e. from gene to phenotype. Copyright 0 1996 Elsevier Science Ltd.

Index descriptors (in addition to those in title): Chromatin, replication, transcription, gene expression, RNA processing, RNA transport.

Genetics without an exact knowledge of the eucaryotic chromosome organization is like physics in the time of Maxwell-without a knowledge of the structure and organization of the atom. Lima-de-Faria (1983)

1. INTRODUCTION The presence of polytene chromosomes has been described in a limited number of plants and animals. They are generally found in cells or organs that are engaged in secretory functions, or those that nourish or support the development and differentiation of cells or embryos, i.e. are usually characterized by high rates of RNA and protein synthesis (Table 1). Polytene chromosomes, since their discovery as a cytogenetic tool in Drosophila about 60 years ago, have become important in solving major problems in special fields of biological research. Beermann’s (1952) classical investigations on Chironomus salivary gland chromosomes focused attention on structure/function relationships of gene expression at the chromosome level. He concluded that the structural modifications known as puffs are the consequence of differential gene activity, a view that was supported by the visualization of RNA synthesis at such sites (Pelling, 1955). Becker (1959, 1962) demonstrated that in Drosophila, the ontogenetic changes in puff formation and regression, occurring during the larval/prepupal transition, must

63 64 H. Kress

Table 1. Occurrence of polytene chromosomes in plants and animals*

Species+ Tissue Level of polyteny*

Plants Monocotyls Scilla bifolia (Lil.1) Antipodal cells 1024C Clivia miniata (Lil.) Antipodal cells 128n Allium ursinum (Lil.) Endosperm haustoria

Dicoryls Aconitum ranunculifolium (Ran.) Antipodal cells 128n Papaver rhoeas (Pap.) Antipodal cells 128n Phaseolus coccineus (Fab.) Suspensor 8192n Tropaeolum majus (Ger.) Suspensor 2048C Rhinanthus alectorolophus (Ser.) Endosperm haustoria 384n Thesium alpinum @cr.) Endosperm haustoria 384n Bryonica dioica (Cut.) Anther hairs 256C

Ciliata Stylonychia mytilus (Spi.) Macronucleus anlage 4096C” (transiotory) lnsecta Bilobella massoudi (Co].) Salivery gland Schistocerca gregariaJ3al.) Fat body (enocytes) Bombyx mori (Lep.) Silk gland 524,288C Calliphora erythrocephala (Dip.) Trichogen cells 4096nn Chironomus tentans (Dip.) Salivary gland 32,768n Chironomus thummi (Dip.) Salivary glands 8192C Malpighian tubules 16,394C Sarcophaga bullata (Dip.) Foot pads 2,048C Drosophila melanogaster (Dip.) Salivary gland 1024n; 2048C

Mammalia Micro&s arvalis (Rod.) Trophoblast 2048C Mus musculus (Rod.) Trohoblast 512-1024C Rattus rattus (Rod.) Trophoblast 1024C; 4096C

*Selected from Nag1 (1978, 1981). ‘Cal. = Collembola; Cut. = Cucurbitales; Dip. = Diptera; Fab. = Fabales; Ger. = Geraniales; Lep. = Lepidoptera; Lil. = Liliales; Pap. = Papaverales; Ran. = Ranunculales; Rod. = Rodentia; Sal. = Saltatoria; Ser. = Scrophulariales; Spi. = Spirotrichia. *C values refer to DNA measurements, n values to other methods (chromosome counts, volumetry). OAmmermann et al. (1974). s = Ribbert (1967). be controlled by a humoral factor. It was shown in Chironomus to be the moulting hormone (ecdysone) (Clever and Karlson, 1960). These investigations were of general interest, because they provided the basis for a new concept of steroid hormone action at the chromosome level (Karlson, 1963). Later, the phenomenon of sequential gene activation was discovered (Clever, 1964), and subsequently analyzed in detail in Drosophila by Ashburner and his associates, who finally proposed a formal model of interactions between the ecdysone-receptor complex and early and late target sites (Ashburner et al., 1974). In addition to ecdysone-induced puffing, the formation of puffs by heat shock turned out to be a second gene-expression system of general importance. Originally described in the polytene chromosomes of Drosophila (Ritossa, 1962), the heat-shock response Polytene Chromosomes: Eucaryotic Interphase State Model 65 has been found in a wide variety of pro- and eucaryotes, and is now regarded as a general protection mechanism in living cells against damage caused by stress conditions. The discovery that the expression of several of the heat-shock genes is also controlled by ecdysone (Ireland and Berger, 1982) provided a formal basis for connecting the hormonal regulatory networks of development with those of the maintenance of cell function in the context of thermotolerance. The cloning of ecdysone- and heat-shock-induced genes for elucidating the molecular mechanisms of converting external signals into specific patterns of gene expression, was a logical consequence of these basic cytological investigations. However, for the comprehensive understanding of the orchestration of gene expression, the analyses had to be extended to the nuclear compartment too. The experimental approach to this aspect of gene expression may be traced back to early attempts to isolate RNA from Chironomus Balbiani Rings (gigantic puffs), which subsequently became an important model system for studying the structure of active genes, the formation of ribonucleoprotein (RNP) particles, their transport through the nuclear matrix, and the translocation of transcripts across the nuclear membrane (reviewed by Daneholt, 1974; Mehlin and Daneholt, 1993). During the last decade, a multitude of non-isotopic methods for the detection of nucleic acid and protein targets became available. The application of these methods in whole-mount preparations of nuclei and cells in combination with new microscopical methods (e.g. confocal microscopy) opened a new exciting field of the spatial visualization of events related to gene expression. Although, in that way, the cytological analysis of interphase chromatin and the nuclear matrix has also become possible in diploid cells, polytene nuclei offer the unique opportunity of correlating reporter signals and visible chromosomal structures. In addition, the larger size of these nuclei provides a better spatial resolution of perichromosomaYinterchromosoma1 relationships. This article describes the present view of polytene chromosome structure, their replicative and transcriptional activities, and aspects of nuclear RNA transport. Owing to space limitations, it will focus only on more recent developments. For information on earlier literature, the reader should consult reviews by Alfert (1955), Beermann (1962, 1972), Berendes (1973), Ashburner (1970, 1980), Daneholt (1974, 1975), Nag1 (1978, 1981), Zegarelli-Schmidt and Goodman (1981), Zhimulev et al. (1981), Richards (1985), Hill and Rudkin (1987), Korge (1987), Sorsa (1988) and Hofmann and Korge (1993).

2. ACQUISITION OF POLYTENY DURING DEVELOPMENT Polytenization of chromosomes is accomplished by “endo-cycles”, which are defined as replication cycles within the nuclear envelope without spindle formation (Geitler, 1953; Nagl, 1978). In such cells, the cell cycle is reduced to an S- and a G-phase without karyo- and cytokinesis. In dipterans, most of the larval cells enter the endo-cycle pathway, and only cells destined to become precursors of imaginal cells and of nerve cells maintain diploidy. We are just beginning to understand the underlying control mechanisms for switching cells from the mitotic cell cycle to its somatic dead-end short-circuit version. In Drosophila melanogusfer, it is now well established that a Gl phase is acquired for the first time in the 17th embryonic cell cycle. During its prolonged duration, a decision is made between continuing mitotic cycles or entering endo-cycles 66 H. Kress

(Edgar and O’Farrell, 1990). Endoreplication commences at cycle 18 in different regional domains of the embryo, and is later on restricted to distinct periods (Smith and Orr-Weaver, 1991). We have to assume that the genes involved in determining the endo-cycle pathway must be expressed according to epigenetically programmed spatiotemporal patterns. In eucaryotic cells, the control of mitosis depends on a universal control mechanism, which is based on the activation of the ~34~~~’protein kinase by binding of GZcyclins and dephosphorylation by Cdc25 (reviewed by Nurse, 1990). The subsequent phosphorylation of cellular substrates by activated ~34”~” leads to chromosome condensation, breakdown of the nuclear envelope and spindle formation, which is followed by mitosis (reviewed by Moreno and Nurse, 1990). In D. melanogaster, a number of cell cycle genes have been cloned (reviewed by Foe et al., 1993), among them the ~34”~‘~ homologue Dm cdc2, cyclins A and B, string (homologue of the Schizosaccharomyces pombe cdc25 gene) and the Gl-cyclin E., While DmcycE plays also an essential role during S-phase in endo-cycling cells (Knoblich et al., 1994), the main regulators of mitosis, namely, the GZcyclins A and B, Dm cdc2 and String, do not (Lehner and O’Farrell, 1990; Stern et al., 1993; Edgar et al., 1994). Obviously, the expression of the corresponding genes is not resumed in these cells during cycle 18, when their zygotic transcripts are accumulating again in prospective imaginal cells. In endo-cycling cells, therefore, there is a bypass of the G2-M checkpoint, allowing these cells to enter a new S-phase without a preceding mitosis. Durinio and O’Farrell (1994) suggested that this could be brought about by the activation of a putative “checkpoint bypass gene”, which should be inactive in mitotic cells. Its activity should be comparable to that of the S. pombe gene rum1 +, whose overexpression leads to successive rounds of replication uncoupled from mitosis, i.e. to endoreplication (Moreno and Nurse, 1994). Formally, the gene escargot (esg), could represent a gene involved in such processes, because its expression prevents cells from entering the endo-cycle (Fuse et al., 1994). The gene encodes a C2-Hz-type zinc-finger protein (Whiteley et al., 1992), which is interpreted to interfere as a monomer with cisltrans interactions of basic Helix-LoopHelix activators of genes, whose products are required to enter the endo-cycle pathway. Although these interpretations are of highly speculative character, they provide the first conceptual framework for genetically dissecting the mechanism of endo-cycle control, which appears to be an active one rather than a default shuttle. This is consistent with the finding that in string mutant embryos, where cell cycle processes are eliminated, endo-replication occurs according to the epigenetically programmed spatiotemporal patterns (Durinio and O’Farrell, 1994). The analysis of Smith and Orr-Weaver (1991) showed that in D. melanogaster, the C-value of endo-cycling cells deviates from the geometrical increase already after the first S-phase, indicating disproportionate representation of different genomic sequences from the very beginning. It had earlier been suggested that the constitutive heterochromatin (cr-heterochromatin) is excluded from replication (Gall et al., 1971; Lakhotia, 1974). Because a-heterochromatin comprises about 30% of the haploid D. melanogaster genome (Rudkin, 1969; Laird, 1974; Gatti and Pimpinelli, 1992), a C-value of 3.4 after the first endo-S-phase and of 6.2 after the 2nd one is expected. This matches exactly the mean C-values reported by Smith and Orr-Weaver (1991) for hindgut (3.4) and salivary gland (6.2) DNA in 13 h embryos. Some reports challenged the model of disproportionate replication of hetero- and euchromatin in polytene nuclei of D. Polytene Chromosomes: Eucaryotic Interphase State Model 67

melanogaster salivary glands (Dennhbfer, 1981, 1982; Lamb, 1982), but could not be confirmed (Lifschytz, 1983; Lakhotia, 1984). Disproportionate representation of heterochromatin in Drosophila salivary gland chromosomes seems to be real, although it may perhaps be more an exception than a rule. Consistent with this latter statement is the fact that in Chironomus salivary gland chromosomes centromer associated satellite DNA is proportionally represented in distinct bands assumed to be the centromeres (Rovira et al., 1993). In the polytene chromosomes of pseudo nurse cells of the D. melanogaster otu’ mutant (such ovarian tumor mutants produce polytene chromosomes in nurse cells of ovarian follicles that usually lack an oocyte; King et al., 1978), which show a banding pattern that is similar to that of salivary gland chromosomes, the absence of constrictions (comprising intercalary heterochromatin) has been described (Heino, 1989, 1994). In addition, these chromosomes are not fused at a common chromocenter (as is typical for Drosophila salivary gland chromosomes), indicating that the replicative state of the centromeric heterochromatin must be affected as well. Southern blot analyses of Calliphora satellite DNA sequences suggest that they are underrepresented in salivary gland DNA, but proportionately represented in nurse cell DNA (Nazimiec and Beckingham, 1986). The authors point to the possibility that there is a positive correlation between proportional replication of heterochromatin and nurse-cell-specific transcriptional activity. It should be mentioned in this context, that interrelation of transcription and replication has been demonstrated unequivocally in the case of D. melanogaster chorion gene amplification, where an enhancer-like c&element activates both transcription and amplification of these genes (reviewed by Orr-Weaver, 1991). It is conceivable, therefore, that the initial underrepresentation of heterochromatic DNA in polytene chromosomes could be compensated by subsequent rounds of regionally restricted amplification in the context of transcription. The transition from disproportionate to proportionate replication during development has also been demonstrated in D. melanogaster wild type nurse cells (Hammond and Laird, 1985). A switch from proportionate to disproportionate replication during the 2nd or 3rd endo-cycle has been proposed for different tissues in D. nasutoides (Zacharias, 1992). To summarize, these findings strongly suggest that there may be species-, tissue-, stage- and endo-cycle-specific discriminative patterns of the regulation of replication in hetero- and euchromatin in endo-cycling nuclei. We do not know the genetic or molecular basis of these patterns. Recently, it has been doubted that differential replication is responsible for underpolytenization. The 2-dimensional gel analysis of replicative intermediates from a 1.3 kb D. melanogaster minichromosome, containing well-defined heterochromatic regions (Karpen and Spradling, 1990), did not reveal the existence of nested replication forks, as they are expected according to the underreplication model (Glaser et al., 1992). The authors consider elimination of DNA from heterochromatic regions, leading to a gradient of polytenization as an alternative model. During post-embryonic development, replication of polytene chromosomes does not occur continuously, but displays periodic waves which may be under hormonal control (for reviews see Rudkin, 1972 and Spradling and Orr-Weaver, 1987). For the individual S-phases, Rudkin (1972) postulated 3 sequential stages: initiation of DNA synthesis at disperse sites (discontinuous), spreading of replication along the full lengths of chromosomes (continuous), and delayed termination at distinct sites (discontinuous). Mukherjee and co-workers advanced a more detailed model of endo-cycle S-phase in Drosophila salivary gland chromosomes (reviewed by Mukherjee et al., 1980, 1989), 68 H. Kress

continuous

Fig. 1. Graphic presentation of the various stages of the Drosophila polytene chromosome S-phase (from reviews by Mukherjee et al., 1980, 1989). The description refers to the patterns of [3H] thymidine incorporation into nascent chromosomal DNA. DD = dispersed discontinuous; C = continuous; D = discontinuous; CL = chromocenter labeling. Chromosomal structures that start to become labeled are indicated. Relative labeling intensities are shown in sections 1-3.

which is shown in Fig. 1. During the initial discontinuous phase replication starts in interbands, puffs, and small bands and subsequently extends into bands and heterochromatic regions until the euchromatic arms and the chromocenter are continuously labeled. Both phases represent the initiation of replication at primary and secondary sites. The appearance of unlabeled gaps on the chromosomes initiates the 3rd phase (terminal discontinuous), followed by the 4th phase, which is characterized by the exclusive labeling of the chromocenter. Salient features of the S-phase are multiple and asynchronous sites of initiation and termination and the lack of rereplication of individual sites prior to the completion of the whole S-phase, as indicated by the lack of the simultaneous occurrence of early and terminal (e.g. DD and 1D or CL; compare Fig. 1) labeling patterns. Although the dynamics of phases suggests that the onset and probably also the duration of replication at distinct sites positively correlate with DNA content, Htigele’s (1970,1972) observation of delayed replication in a region of low DNA content on Chironomus chromosome II indicates that factors other than mass or compaction may play a role as well. In this context, it could be argued that long-range effects like the synchronous activation of multiple replicons, extending over several joining bands and interbands (super-replicon complexes; Mukherjee et al., 1980), might be involved. Absolute time values for the duration of S-phases in polytene chromosomes have not been reported. In general, the length of S-phases depends on the synchrony of replication fork initiation, interorigin distances, and fork elongation rates. For interorigin distances, values of about 10 kb in embryonic DNA and >lOO kb for polytene chromosomes have been reported in different Drosophila species (see Table 1 in Spradling and Orr-Weaver, 1987). This finding suggests that in polytene chromosomes, the number of replication origins may be reduced by a factor of 10 or more. This would Polytene Chromosomes: Eucaryotic Interphase State Model 69

also imply that in Drosophila, replicons are, on the average, at least 3 times larger than medium-sized bands (30-40 kb). A generalization of this statement is not possible at present; an individual analysis in Chironomus suggests the opposite situation (L&m, 1981). An elongation rate of 2.6 kb mini has been calculated (Blumenthal et al., 1973) for D. melanogaster cleavage nuclei or cultured cells. This comes close to the estimate of 3 kb min-’ for human HeLa cells (Hozak et al., 1993). For salivary gland chromosomes, a value of 0.3 kb min-’ has been reported in D. virifis (Steinemann, 1981) but 2 different ones, namely 2.7 and 0.2 kb min-i for D. nasuta (Lakhotia and Sinha, 1983). It remains an open question as to whether the existence of 2 distinct classes of replicons in the latter case is due to differences of early and late replicating units, or is related to the fact that Drosophila 3rd larval instar salivary glands consist of 2 cell types (collum and corpus cells; Berendes (1965) and von Gaudecker (1972)) that differ in their degree of polytenization (Becker, 1959). The values of 0.2-0.3 kb min-’ indicate at least that the elongation rates in polytene chromosomes may be reduced by a factor of about 10 in relation to diploid cells. This observation suggests that, in combination with the lo-fold reduction in the number of replication origins, the S-phases in polytene chromosomes may be about 100 times longer than those in diploid cells. The mechanisms that control the cyclic replication patterns in polytene chromosomes are still unknown. In principle, there shouid be at least 2 main checkpoints of control: (1) entrance into the S-phase and (2) the suppression of rereplication at individual sites within the same round of endo-replication (for broader reviews see Coverley and Laskey (1994) and Heichmann and Roberts (1994)). The discontinuous replication patterns of polytene chromosomes at the onset of each round of endo-replication indicate that, despite the theoretically simultaneous availability of the replication machinery at all chromosomal sites, the temporal initiation patterns are site-specific. There must be local signals that are responsible for the discrepancies. It has been demonstrated in vertebrate cells that the specific initiation of replication is achieved only in the correct chromosomal and/or nuclear context, while replication of naked DNA starts unspecifically (briefly reviewed by Coverley and Laskey (1994)). On this basis, we have to conclude that specific chromatin domains or chromosomal structures are more important than trans-factors. This holds true also for polytene chromosomes, because the temporal (early vs. late) replication patterns of homologous X chromosomes in D. miranda and D. persimilis hybrids are conserved, despite their co-existence in the same nuclear environment (Mutsiddi et al., 1984). This applies also to the similarity of termination patterns of replication in Anopheles stephensi salivary glands and nurse cells (Redfem, 1981b) or to salivary glands and gastric caecae of D. nasuta (Lakhotia and Tiwari, 1984). So far, there is only one molecular analysis of replication at a distinct site in polytene chromosomes, namely, the DNA puff IU9a in Sciara coprophila. It is one out of 18 sites (Gabrusewycz-Garcia, 1975) that amplify their DNA by up to almost 20 additional rounds of replications after chromosomal replication has come to an end (Crouse and Keyl, 1968; Wu et al., 1993). Two points are of special interest: (1) replication in DNA puffs is induced by ecdysone (Crouse, 1968) and (2) the selective continuation of replication suggests that hormone-dependent factors either activate replication or release its suppression by interacting with site-specific signals. In the case of puff 11/9A, it was reported that ecdysone-induced amplification depends on protein synthesis (Wu et al., 1993). This suggests that according to the concept of sequential gene activation (Clever, 70 H. Kress

1964), products of early induced genes should be responsible for the initiation of DNA amplification. In the case of puff 11/9A, the replication origin could be mapped to a =l kb region 5’ upstream region of the glue protein gene B/9-1 (Liang et al., 1993). Indications that replication starts at multiple sites in this region were confirmed by a subsequent analysis using a novel 3D gel method (Liang and Gerbi, 1994). The observations suggest that replication starts within this region in a bidirectional manner at a single site on each individual DNA strand. In addition, the position of this site may vary within this region for different DNA strands. This is another strong argument in favour of the proposal that it is not a specific DNA sequence, but rather a distinct feature of the local chromatin that is responsible for the initiation of replication. In mitotic cells, the initiation of a new round of replication of a replicated genome normally depends both on its complete duplication and on the breakdown of the nuclear membrane during mitosis (see Coverley and Laskey, 1994; Heichmann and Roberts, 1994). In polytene chromosomes of Drosophila, the suppression of rereplication within the same S-phase occurs in spite of disproportionate polytenization, and the persistence of the nuclear membrane between subsequent rounds of replication. It is obvious, therefore, that other intracellular signals must be effective. However, as long as even in mitotic cells the corresponding signals have to be identified, the control of replication in polytene nuclei will also remain obscure.

3. BASIC STRUCTURE AND ORGANIZATION Polytene chromosomes are dynamic structures that, starting from an oligonemic state, gradually gain their final size, shape, and texture in the course of development. In addition, they may regionally change their morphology under the influences of functional demands, involving differential replication and/or transcription in already terminally differentiated organs or tissues. Differing degrees of polytenization and of synapsis of individual chromatids result in a continuum of diverse chromosomal bodies, ranging from the diffusely banded and fibrillate structures in Cecidomyiidae (Bier, 1961; brief review by Ashburner, 1980) to the compact, rod-like chromosomes in Chironomidae or Sciaridae with levels of polyteny up to 32.768 (Beermann, 1959). The most striking feature of polytene chromosomes is the banding pattern, which is clearly detectable in higher Diptera. For the last 4 decades, there has been a controversial debate as to whether the banding pattern, which is most probably the result of discontinuities of DNA packing in bands and interbands along the chromosome axis (Spierer and Spierer, 1984; Ananiev and Barsky, 1985), is (a) an inherent character of DNA or (b) the consequence of local decondensations of DNA in the context of transcription (for reviews on that particular issue see Zhimulev et al., 1981; Hill and Rudkin, 1987), which leads to the formation of interbands and puffs. The careful comparison of the banding patterns and their variations on polytene chromosomes from different larval and/or adult tissues of the same stock or strain in different dipteran species (e.g. Beermann, 1952; Richards, 1980; Redfern, 1981a; Zhimulev et al., 1982; Hochstrasser, 1987) supported the former alternative (genetic hypothesis; Ashburner, 1980). However, under artificial conditions, exceptions have been reported (Ribbert, 1979). Local deviations may be effected by epigenetic modulations of compaction or polytenization that are imposed on the basic pattern by cell or tissue specific functional differences. This is impressively demonstrated in the case of the already mentioned Poiytene Chromosomes: Eucaryotic Interphase State Model 71

Table 2. Evaluations of DNA compaction in various structures of Drosophila salivary gland chromosomes*

Bands+

Puff Interband+ Small Large Reference

23 Laird (1980) c 23 -+ 180 Spierer et al. (1983) lo-17t Kress et al. (1985) 2.ti.2 Udvardy et al. (1985) 836 151-161 Kozlova et al. (1994) 1.4-3.5 11 30-50 Shemeshin et al. (1989) 7-8 Rykowski et al. (1988) 18t -_, Friedman et al. (1991) (;‘,” ;I) Requena et al. (1987) 30 170 Means (7.3 ::.6) Means including Drosophila hydei 0.2 0.4 1 5.4 relation to small bands (30 nm fiber)

*Values were calculated by the authors (see references) on the basis of different criteria and preparative conditions. Consequently the means should be regarded only as non-committal informative. ++n+ average compaction of interbands and small bands. *On surface spread chromosomes the average compaction of 3.4 for puff 68C DNA was calculated. Given an extension factor for spreading of 3-5 a value of about l&17 would result for squashed reparations. gCalculated from Fig. 2 of this reference.

ovarian tumor (otu) mutations in D. melunoguster, which lead to the abnormal formation of polytene chromosomes in the pseudo nurse cells of adult ovaries. The banding patterns observed in these germ line cells are basically similar to those of somatic larval salivary gland chromosomes, but exhibit tissue-specific differences in puffing patterns (Sinha et al., 1987; Heino, 1989, 1994). What are the basic differences in chromatin structure in bands and interbands? The published values of packing ratios in various structures of Drosophila salivary gland chromosomes (Table 2) suggest that there are 2 clearly distinct ranges. Interbands, puffs, and small bands represent a continuum of values. Their upper limit of about 30 suggests that the highest degree of chromatin organization in these structures is the 30-nm fiber (packing ratio = 30-40; Mirkovitch et al., 1987). This is the canonical solenoid structure, which is demonstrable in interphase and metaphase chromosomes of almost all eucaryotic chromosomes (Wagner et al., 1993). Chromatin in interbands and puffs may be organized at lower levels like the non-beaded 5-nm fiber, the open lo-nm fiber, which exhibits a compaction of 2-3 in the beads-on-a-string configuration, or the closed lo-nm fiber with a compaction of about 7. Intermediate values could be explained by varying contributions of these various structural conditions at a given chromosomal site, as it has actually been described for Balbiani Ring chromatin in native Chironomus tentans polytene chromosomes (Bjorkroth et al., 1988). Consistent with this view is the presence of the linker histone Hl, which is responsible for the transition from the open to the closed lo-nm fiber and its solenoidal folding to the 30-nm fiber, in Balbiani Rings (Ericsson et al., 1990) and in puffs and interbands of native D. melunogaster chromosomes (Hill et al., 1989). 72 H. Kress

In large D. melanogaster bands, the packing ratios are between 150 and 180 (Table 2), i.e. about l/lOth of the packing ratio assumed for the average mammalian interphase 240-nm chromatin fiber (Manuelidis and Chen, 1990). This is thought to be achieved by the attachment of radial loops of 30-nm fibers to a central scaffold of non-histone proteins (NHPs) via specific scaffold-attaching regions (SARs, for a recent review see Laemmli et al., 1992). The reported values of DNA compaction in D. melanogaster bands are too low to substantiate this type of organization for polytene bands. Whether this holds true for all large bands in that species is not known, and comparable data for other species are presently not at hand. NHPs in chromomeres should serve at least two different structural functions: (a) increase the compaction of DNA beyond the level of the 30-nm fiber (in D. melanogaster by a factor of at least 5, see Table 2) and (b) adhere the supercoiled chromatids aligned in parallel in order to form stable chromomeres. The data available on the distribution of NHPs on polytene chromosomes do not allow any distinct conclusions on the actual composition of bands (for review see, for example, James et al., 1988). The existence of chromomere-specific self-adhesion proteins that are responsible for the synapsis of chromatids had been postulated earlier by Mayfield and Ellison (1975). Poorly characterized band-specific proteins have been described by Saumweber et al. (1980), Alfageme et al. (1980), and Kuo et al. (1982). The association of ubiquitin, which modifies histones H2A and H2B, with band structures has been suggested (Izquierdo, 1994) and High Mobility Group protein I (HMGla) was identified on condensed regions of Chironomus thummi chromosomes (Wisniewski and Schulze, 1992). For the same species, it was demonstrated that in 2 subspecies some homologous bands differ significantly in the accumulation of the histone Hl variant I-l (Mohr et al., 1989). The variant protein is characterized by a specific N-terminal reiterated insertion, which was proposed to confer the formation of a condensed subtype of chromatin (Schulze et al., 1993). In D. melanogaster, Fleischmann et al. (1987) found a protein associated with condensed bands and centric heterochromatin. This location suggests a role in the packaging of chromatin. In the same species, Frasch (1991) identified a band-specific protein (NJl), which exhibits homologies with the protein encoded by the vertebrate gene Regulator of Chromatin Condensation, RCCl. It could be associated with members of the Polycomb group (PC-G) proteins, which repress the expression of homeotic and of other developmental genes (reviewed by Epstein, 1992). The Polycomb (PC) protein and the polyhomeotic (ph) protein co-localize at about 100 chromosomal sites, and co-precipitate with at least 10-15 other proteins (Franke et al., 1992). This led to the hypothesis that they are constituents of multimeric complexes that repress gene activity. Although the binding of PC and ph encoded proteins is sequence-specific, neither of the 2 proteins binds directly to DNA, and it is more probable that they do not work at the transcriptional level, but rather work at the chromatin level via protein/protein interactions. Whether PC-G proteins play a genuine structural role in specifically packaging DNA containing inactive developmental genes into chromomeres remains to be established. Circumstantial evidence for such a function is provided by the fact that the PC protein shares a conserved domain (chrome domain) with the heterochromatin-associated protein (HPl; see Eissenberg and Elgin, 1991), which is also associated with DNA compaction in euchromatic regions (Belyaeva et al., 1993). The values shown in Table 2 suggest that the band/interband ratio of DNA compaction Polytene Chromosomes: Eucaryotic Interphase State Model 73

is about 10: 1. The band/interband ratio of dry masses in D. mefanogaster polytene chromosomes has been estimated to be less than 4:l (Laird, 1980). This suggests that in relation to DNA, there should be more protein in interbands than in bands. High relative protein concentrations might prevent interband chromatin from aggregating, a tendency that appears to be a natural property of the 30-nm fiber, at least in vitro (Widom and Klug, 1985) and thus explains the lack of association of interchromomeric chromatids (Ananiev and Barsky, 1985). However, evidence for the presence of NHPs in interbands is scarce (Saumweber et al., 1980; Kuo et al., 1982). The single characterized proteins localized in interbands so far are RNA polymerase II (Sass and Bautz, 1982) and protein B52, which is homologous to the human pre-mRNA splicing factor ASF/SF2 (Champlin et al., 1991). The presence of U2 snRNA, a component of snRNPs in interbands, has also been reported (Zachar et al., 1993). The identification of proteins involved in transcription and RNA processing in interbands leads us to the long-standing issue of the relative distribution of genes in bands and interbands. Starting with the concept that chromomeres are morphological units that represent informative units (Beermann, 1962), and its initial support by a fair 1: 1 relationship between genetic units and bands (Judd et al., 1972), subsequent more elaborate genetic studies disclosed the lack of coincidence between the numbers of bands and complementation groups in the 3Al-3C7 region of the D. melanogaster X- chromosome (Young and Judd, 1978). That the original model extremely simplified matters was evidenced when studies of gene organization at the DNA level became possible, and revealed that the chromosomal organization of eucaryotic genes is unexpectedly complicated. This is demonstrated by the fact that opposite strands at the same site may encode different products, or that genes may be localized in introns of other genes. There are genes that encompass 100 kb or more, which corresponds to several contiguous polytene chromosome bands and interbands. In D. melanogaster, several independent chromosome walks opened first insights into the diversity of eucaryotic chromosome organization. Two walks that each extend over more than 300 kb of DNA in the 87DE (Spierer et al., 1983; Bossy et al., 1984) and in the 9F/lOA regions (Kozlova et al., 1994) provided the unexpected information that in these regions genes may preferably be localized in chromosomal regions where smaller bands and interbands dominate, while in conspicuous bands gene frequencies are remarkably reduced. This notion is corroborated by another study, covering 38 kb in the 28Cl/C&5 region, where at least 6 transcription units have been identified in 2 faint bands with flanking interbands (Friedman et al., 1991). The survey of about 70 kb in the 59F region revealed the existence of nested gene organization in the small band 59F5 and its surroundings (Kurzig-Dumke et al., 1995). Larval glue protein genes are also localized in regions of small bands and interbands rather than in large bands: Sgs-l in 3Cll-12 (Korge, 1977), Sgs-3, -7 and -8 in or around 68C6 (Kress et al., 1985) and the D. virilis glue protein genes Lgp-1 and Lgp-3, which are related to the D. melanogaster 68C genes (Swida et al., 1990), in or close to the small band 17Al (Fig. 2A). Therefore, accumulating evidence suggests that interbands and small bands are preferred sites of transcription. This is consistent with the reported presence of RNA polymerase II (Sass and Bautz, 1982), of RNA/DNA hybrids (Requena et al., 1987), of RNP particles (Ananiev and Barsky, 1985; Mott and Hill, 1986 and literature therein) in interbands and with the in situ hybridization of salivary gland-specific cDNA to interband DNA (Kress et al., 1985; Kress, 1986). H. Kress

Fig. 2. Localization of genes in small bands and interbrand regions. (A) In situ localization of D. virifis larval glue protein gene Lgp-I in the region of the small band 17Al by hybridization of a 1 kb Lgp-I specific biotinylated DNA probe to a moderately surface spread preparation (Kress et al., 1985). Signal visualization (arrow-heads) by a horse-radish peroxidase reaction (Thiiroff et al., 1992) in conventional phase contrast (upper panel) and reflection phase contrast (centrepanel). The small arrows demarcate the bands 16F3 and 17A2 (band assignments according to Kress, 1993) bordering the interband that is decorated by the signal. The lower panel shows this interband in a squash preparation of an untreated chromosome. Scale bars = 5 pm. (B) Visualization of distinct transcription units in interbands of an unidentified region of D. virilis chromosome 2. Gold-labeling (arrow-heads) of hybridized biotinylated cDNA prepared from 3rd larval instar salivary-gland poly(A) RNA (Kress ef al., 1985). Note that the signals accumulate at the borders of chromomeres. The small arrows indicate clumps of chromatin that may represent a small band. Scale bar = 2 pm. (From Kress, 1986, Naturwtirenschufe~, 73, 180-187, by copyright permission of Springer-Verlag, Heidelberg.)

In an effort to determine the interphase state of an individual gene, Rykowski et al. (1988) studied the relative positions of 5’ flanking and coding sequences of the D. melanogaster Notch gene in the 3C6/7 region. They interpreted their results as the promoter region being localized in the C6-7 interband (lo-nm fiber), while the coding portion, which is not transcribed in salivary glands, is found in band C7, probably as 30-nm fiber. A similar crossing of the band/interband border by gene sequences has been proposed for C. tentans Balbiani Ring 2 genes, which encode glue proteins (Sass, 1984). In generalizing this situation, we would expect that upon activation such genes should become part of interbands, a conclusion that is supported by a number of indications of transcriptional activity at interbandhand junctions (Requena et al., 1987; Ananiev and Barsky, 1985; Kress et al., 1985; Kress, 1986). Consequently, interbands should,be composite structures that are formed by 2 different factors: sequence inherent structural forces and the consequences of transcriptional activity (see Fig. 2B). Polytene Chromosomes: Eucaryotic Interphase State Model 75

4. STRUCTURE-FUNCTION RELATIONSHIPS The first step in the expression of genetic information is transcription, and its appropriate substrate in eucaryotes is chromatin in the interphase state. It is generally assumed that active genes require an “open” configuration of chromatin structure, which allows access of the components of the transcription machinery to enhancer and promoter regions. Indicative of open chromatin is the presence of DNase hypersensitive sites (DHsites; for review see Cartwright and Elgin, 1988). These are characterized by histone Hl depletion (Manuelidis and Chen, 1990), which opens the 30-nm fiber and exposes nucleosome-packed DNA. It has been proposed that there are preset promoters, which are partially or completely nucleosome-free, and remodeling promoters that are fully packed with nucleosomes, their array being specifically perturbed during gene activation. As a rule, the first type is found in ubiquitously inducible genes, constitutively expressed genes, and cell cycle dependent genes, while the second type is represented by tissue and/or stage specifically expressed genes (Wallrath et al., 1993). The 3-dimensional manifestations of such chromatin transitions have been studied extensively on polytene chromosomes. In Drosophila, there is a number of genes that are well characterized at the genetic and molecular level. Their activation exhibits distinct changes of chromatin morphology: the heat-shock genes comprise preset promoters and the larval glue-protein genes represent the remodeling promoter type. The latter encode proteins, which are synthesized in huge amounts in the salivary glands. In D. mehnoguster, such genes are localized in intermolt puffs and their expression is restricted to the 2nd half of the 3rd larval instar (reviewed by Meyerowitz et al., 1987). In accordance with this tissue and stage-specific pattern of expression, the appearance of 3 DH sites in the distal promoter region of the Sgs-4 gene (between -330 and -480) has been demonstrated and, in various strains carrying deletions of this region, Sgs-l expression is severely reduced (Shermoen and Beckendorf, 1982). A deletion of 52 bp, covering the most proximal DH site, prevents binding of the highly basic nuclear protein Bx42 (Frasch and Saumweber, 1989; Wieland et al., 1992) to the puff 3C region harbouring the gene (Saumweber et al., 1990). In addition to the binding of this factor, that of proteins of the Broad-Complex (Kalm et al., 1994) as well as the binding of ecdysone receptor (EcR) and of secretion enhancer binding proteins SEBP2 and 3 (Lehmann and Korge, 1995) has been reported. This qualifies that region at an extent of about 130 bp as a distal regulatory element, which enhances ecdysone-induced Sgs-4 expression by the interaction of &-elements with EcR and several other truns-factors. This is a convincing example for the existence of an open chromatin configuration at a site of regulatory significance on polytene chromosomes. A comparable situation, although less characterized, is found in the promoter of the Sgs-3 gene, whose expression is cooperatively controlled by a distal enhancer and a proximal promoter element (Roark et a(., 1990). Prior to gene activation, the distal site is maintained inactive by a single nucleosomal core particle, which is flanked by 2 DH sites that are already present in embryonic DNA. Gene activation involves the modification of this site (Ramain et al., 1988; George1 et al., 1991). HOW is puffing related to the activation of promoters? The initiation of puff formation by the incorporation of acidic proteins under conditions of inhibited RNA synthesis had been demonstrated by Berendes (1968) in D. hydei. In D. melunoguster, the puff 68C, which harbours the Sgs-3 gene, is observed in a mutant background that prevents the 76 H. Kress

expression of the Sgs-3 and of 2 neighbouring Sgs-7 and -8 genes (Crowley et al., 1984). In Sgs-4 transformants, puffs were observed at the sites of integrations without concomitant gene expression (Korge et al., 1990). Puff formation without transcription was also found for fusion constructs that comprised only 2.5 kb of Sgs-4 upstream flanking regions. These results suggest that puffing depends on the presence of distinct Sgs-l upstream sequences, and could represent a morphological correlate of chromatin modification related to the opening of promoters. However, the observation that Sgs-3 as well as Sgs-4 transgenes may exhibit faithful expression without the formation of puffs (Crowley et al., 1984; Korge et al., 1990 and literature therein) indicates that transcription may also be independent of puffing. Thus, we are left with the rather paradoxical situation that glue protein gene expression can be uncoupled from puffing, a fact that renders the puffing phenomenon a rather obscure character, at least in the context of the expression of these genes in Drosphilu. It is possible that puff formation is dispensable for these genes, because they are localized in interbands or small bands (see Section 3) and produce small transcripts (about 1 kb or less) at perhaps moderate rates, while the transcription of genes at high rates (e.g. heat-shock genes, Simon et al., 1985) or the formation transcripts longer than 1 kb (Shemeshin et al., 1989) as well as extensive processing (ecdysone-induced early genes; see Thummel, 1990 and literature therein), causes a longer retention of nascent RNA in the perichromatin of these sites, which leads to puffing. Owing to the ubiquitous character of heat-shock induction, cytological data obtained on polytene chromosomes and molecular data obtained on diploid culture cells can be combined, and a general scenario for chromatin transition can be designed. It was found in 2 D. melunoguster hsp70 genes, localized in opposite orientation in the 87A7 chromomere (Fig. 3A), that the distribution of nucleosomes on the genes and their flanking sequences is modified. Upon heat-shock activation, the major domains of topoisomeraseI1 (topoI1) activity, which were at first predominantly found in the common promoter region and close to the transcription start sites, are shifted in a downstream direction into the coding regions. They also increase in number close to 2 chromosomal domains, scs and scs’ (specialized chromatin structure), in the 3’ flanking regions of the genes (Udvardy and Schedl, 1993). In situ hybridization of fragments of a chromosome walk on polytene chromosomes suggests that the positions of the 2 elements coincide with the borders of the 87A7 chromomere, which harbours about 15 kb of DNA (Fig. 3A). It was hypothesized that the scs’ and scs elements act as swivel points for relaxing the DNA during the decondensation of the chromomere and hsp70 transcription, thus defining the borders of a chromatin domain as a functional unit (for review, see Eissenberg and Elgin, 1991). The redistribution of topoII-sensitive sites during heat shock activation is temporally accompanied by the redistribution of the B52 protein. It colocalizes with RNA polymerase II (PolII) at the genes at low heat-shock temperatures, but is dislodged towards the 3’ flanking regions at full heat shock (Champlin and Lis, 1994). A similar “bracketing” effect of the B52 protein was observed at other sites, too (Fig. 3B). The protein is a member of the group of SR proteins, which are generally involved in splicing processes (for review, see Zahler et al., 1992). The position of the protein in 3’ flanking sequences of the intronless hsp70 genes and its binding to DNA (Champlin et al., 1991) suggest that SR proteins may also serve other functions. They could be involved in condensation/decondensation transitions of 30-nm chromatin in flanking regions of Polytene Chromosomes: Eucaryotic Interphase State Model 77 A

87A6 87A7 87A8 ...... i ...... ,......

...... 15kb ......

...... B52 ......

scs SCS’

Yll 111 III II II Ill 11111 HS B C RNA pol II B52 hrp36 hrp48

87C 87A

Fig. 3. (A) Topographical features of the D. melunoguster 87A7 chromomere. The center line represents genomic DNA in the 87A6-87A8 region, the hatched bars on top represent the corresponding chromomeres. The positions of the 2 hsp70 genes on the DNA and the direction of their transcription are indicated by the open symbols. The small hatched boxes demarcate the special chromatin structures scs and scs’, which border the 87A7 chromomere. The large blocks indicate the positions of the bound B52 protein (see B). The sites of top011 activity prior to (C) and during heat shock (HS) are indicated by vertical bars. (Modified from Udvardy et al., 1985; Champlin ef al., 1991 and Udvardy and Schedl, 1993.) (B) Comparison of heat-shock puffs 87A (left arrows) and 87C (right arrows) stained with antibody to RNA polymerase II and B.52. On the bottom maps, the heat-shock genes responsible for the formation of each puff are indicated. (C) Differential association of the hrp36 and hrp48 hnRNP proteins at the 87A and 87C loci after heat shock. Double-label immunofluorescence with Texas red (hrp36, A) and FITC (hrp48, B) conjugated monoclonal anti-hrp antibodies. For further details see text. (Panel B from Champlin et al., 1991, Genes & Development, 5, 1611-1621, by copyright permission of the Cold Spring Harbor Laboratory Press; panels C from Matunis et al., 1993, J. Cell. Biol., 121, 219-228, by copyright permission of the Rockefeller University Press.) 78 H. Kress genes being transcribed at high rates. The fact that null mutants of the gene encoding B52 are lethal (Ring and Lis, 1994) indicates that its function is indispensable during development and cannot be substituted by other SR proteins. The visible physical separation of PolII, and of the B52 protein at the 87A puff and at others, highlights the power of polytene chromosomes for analyzing the spatial architecture of active chromatin and for gaining access to functional identities of individual components. For example, it has been demonstrated that ecdysone-induced sites are transcribed by the hyperphosphorylated Pol 110 form, while at heat-shock puffs, the hypophosphorylated Pol IIA is also engaged in transcript elongation (Weeks et al., 1993). Prior to heat shock, unphosphorylated Pol IIA seems to be arrested at the promoter, and the binding of heat-shock factors to the corresponding heat-shock elements may activate the arrested enzyme by phosphorylation. There are many other sites that are occupied by Pol IIA only and, since they lack nascent RNA and hnRNP proteins, it is possible that storage of inactive Pol IIA on chromosomes is a feature of distinct sites. This would also imply, for instance, that the presence of Pol II in interbands is not necessarily an indication of ongoing transcription. A number of hnRNP proteins have been isolated which are common to most, if not all, active sites on D. melunoguster polytene chromosomes (Matunis et al., 1992 and references therein). On the other hand, site selectivity has been reported for several NHPs (Kabisch and Bautz, 1983; Hill et al., 1986), including the PEP (protein on ecdysone puffs) zinc finger protein, which specifically binds to puffs harbouring ecdysone-regulated genes (Amero et al., 1991,1993). Matunis et al. (1993) demonstrated clearcut differences between heat-shock puffs and other active sites concerning the presence of the hrp36 hnRNP protein (Fig. 3C). These results strongly implicate the existence of transcript class-specific hnRNP particle-forming proteins, whose differential deposition could channel subsequent splicing reactions of pre-mRNAs. Co-transcriptional splicing has been demonstrated unequivocally for the large transcripts in the ecdysone-controlled puff 74 EF (60 kb primary transcripts) in D. melunoguster (LeMaire and Thummel, 1990) and in Balbiani Ring 1 (40 kb primary transcripts) in C. tentuns (Bauren and Wieslander, 1994), which should be indicated by the formation of snRNP particles in puffs, too. Accordingly, Ul- and U2snRNPs have been identified in puffs of C. tentuns (Sass and Pederson, 1984). In D. melunoguster, these 2 types of particles are deposited independently from A/B type hnRNPs at different active sites (Amero et al., 1992). These results probably exclude the possibility that hnRNPs and snRNPs are components of unitary compaction/processing complexes, but rather mirror the functional separation of pre-mRNA packaging (dependent on RNA mass) and processing (dependent on the number of introns) events. The morphological aspects of RNP particle formation in puffs have been analyzed in detail by biochemical and EM studies of C. tentuns Balbiani Ring RNP samples. Primary transcripts were found as ribbon-like RNP fibers, 7 nm in diameter, which are supposed to be formed by a helical arrangement of RNA around a cylindrical core of proteins (presumably core-type hnRNP proteins) leading to a compaction of 9.6 (Lonnroth et al., 1992). The stability of the filament depends on the presence of a critical segment of the BR transcript. There is evidence that the 7-nm filament is formed prior to splicing and that the RNA in released BR particles is fully, or almost fully, spliced (Mehlin and Daneholt, 1993). The presence of snRNPs in the fibrillar RNP (Vazquez- Nin et al., 1990) suggests that splicing starts early during RNP formation. While still Polytene Chromosomes: Eucaryotic Interphase State Model 79

growing, the fiber is further compacted and bent to form a ring-like toroid. The 3D order of the structure is well-defined with 4 domains, the 5’ end of the packaged transcript being located in domain 1 and its 3’ end in domain 4 (Mehlin and Daneholt, 1993). After completion, the 50-nm particle is released from perichromatin and transported to the nuclear membrane.

5. NUCLEAR RNA TRANSPORT Three-dimensional reconstructions of D. melunogaster polytene nuclei from various tissues revealed that in general (exceptions can be explained by exogenous physical constraints) the chromosomes occupy separate spatial territories in the nuclear periphery. They display the so-called Rabl configuration (telomeres and centromeres at opposite sides of the nucleus) of embryonic cells, which persists during many rounds of replication throughout postembryonic development (Mathog et al., 1984; Hochstrasser et al., 1986; Hochstrasser and Sedat, 1987a). In salivary-gland nuclei several factors could contribute to create a rather rigid physical frame for positioning the chromosomes in distinct subnuclear areas. These are the common chromocenter, multiple points of contact between sites of intercalary heterochromatin and the laminar network of the inner nuclear membrane, associations of chromosomes with nucleolar structures (Ananiev et al., 1981; Hill and Whytock, 1993), or interstitial or telomeric ectopic fibers (reviewed by Ashburner, 1989). Accordingly, profound changes in transcriptional patterns, as they are induced by heat shock or ecdysone, do not significantly displace the chromosomes from their original positions (Hochstrasser and Sedat, 1987b). This suggests that irrespective of the transcriptional patterns, the transport of processed transcripts from perichromatin to the nuclear membrane could occur on established routes. Two modes of transport can be envisaged: (i) RNP particles could migrate along physical structures of the nuclear matrix or (ii) they could freely diffuse, being driven by a concentration gradient originating at the site of synthesis. While the former alternative has been propagated for diploid eucaryotic nuclei in a controversial model proposing transport along “tracks” (for reviews, see Xing and Lawrence, 1993; Rosbash and Singer, 1993), experimental evidence from both Drosophila and Chironomus polytene nuclei suggests a simple diffusion mechanism. Zachar et al. (1993) analyzed nuclear pre-mRNA metabolism in D. melunoguster using a transgene construct comprising the salivary gland specific Sgs-4 promoter, a segment of the SU(W~) gene containing 3 introns, followed by the 1acZ gene and the SV40 polyadenylation signal. The expression of this gene in 3rd larval instar salivary glands leads to steady state levels of transcripts with highly reproducible distribution patterns. High concentrations of nascent or of very young pre-mRNAs were found in close association with the perichromatin of the transcribed gene, producing a “primary zone”. Throughout the nucleoplasm, which is constituted by the space excluded from chromosomes and the nucleolus (referred to as the interchromosomal channel network (ICN), Kramer et al., 1994), pre-mRNAs were uniformly distributed, their concentrations depending on the rate of processing. From their results and a number of theoretical considerations, the authors concluded that pre-mRNAs, processed at different extents, move through the ICN by isotropic diffusion. Retention of incompletely spliced RNAs at the nuclear membrane leads to their uniform distribution in the ICN. The inability 80 H. Kress

to detect mature transcripts neither of the fusion gene nor of the endogenous intronless Sgs-4 gene was interpreted as being caused by their rapid export into the cytoplasm. Owing to the rather artificial nature of the fusion transcripts and of their processing (e.g. regulated splicing of the SU(@) introns instead of constitutive splicing), it is questionable as to whether these results are representative for the metabolism of native salivary gland transcripts. The observations in this context are contradictory. Transcript accumulation in a band that is comparable to the postulated primary zone has been described for the ecdysone-induced D. melunoguster E74A early gene (Boyd et al., 1991) and for the 2 D. virilis genes Egp-1 and Egp-2 (Thiiroff et al., 1992). We have to conclude from the fact that in the latter cases the transcripts are small (cl kb) and intronless, that even the presence of small nascent RNA that is not spliced is sufficient to cause the formation of a detectable primary zone. The induction of the D. virilis larval glue protein genes Lgp-1 and Lgp-3 in mid-3rd larval instar salivary glands is also indicated by a primary zone of transcripts developing at a single chromosomal site (Fig. 4A). On the basis of the results presented in the preceding section, we conclude that the small primary transcripts containing a single 59 nt intron (Swida et al., 1990), must be processed in this zone and should leave it in their mature state. As Fig. 4B demonstrates, the Lgp-3 transcripts become homogenously dispersed throughout the whole nucleus. With ongoing transcription, there is further accumulation of RNA in the nucleus, a clear border being formed by the nuclear envelope (Fig. 4C) and it is only after some delay that transcripts start to appear in the cytoplasm, where they concentrate in huge amounts later on (Fig. 4D). In accordance with Zachar et al. (1993), we conclude from these observations that the transcripts can move freely within the nucleus by diffusion. However, the results do not support the notion of these authors that the export of mature glue protein gene transcripts (in their case Sgs-4) into the cytoplasm is too rapid, as to allow their intranuclear detection. On the contrary, we assume that mature transcripts are also retained at the nuclear envelope. Our interpretation is corroborated by earlier EM studies in Chironomus salivary glands, which suggest even distribution of single and independent Balbiani Ring RNP particles in all areas of the nucleoplasm, but possible accumulation at the nuclear envelope. There, they undergo a morphological change from a spherical to a rod-like conformation during the passage through the nuclear pores (Stevens and Swift, 1966). Daneholt and colleagues, who confirmed and extended these investigations (reviewed by Mehlin and Daneholt, 1993), could demonstrate that translocation through the nuclear pore complex always starts with the 5’ end of the transcript, which is packaged in domain 1 (see previous section) of the completed BR RNP particles. Within the nuclear pore complex, the particles unfold, the 5’ end presumably becoming engaged in protein synthesis, while the 3’ end is still in the complex.

6. SYNOPSIS AND OUTLOOK During interphase, the physical entity and the biochemical composition of a chromosome are the main determinants in executing the 2 major cellular functions: replication and transcription. Our understanding of these processes essentially depends on the degree of our knowledge of the specific and intricate interactions between DNA and chromosomal proteins. The corresponding available data obtained for polytene chromosomes are rather promising, and predispose this system for future successful Polytene Chromosomes: Eucaryotic Interphase State Model 81

Fig. 4. mRNA distribution in the nuclear and cytoplasmic compartments of individual cells from D. virilis mid-3rd larval instar salivary glands, starting with Lgp-3 glue protein gene expression. Transcripts were hybridized with a Digoxigenin-labeled probe and visualized by an alkaline phosphatase reaction (Thiiroff et al., 1992). (A) Appearance of a single strong signal (arrow-head) in the nuclear periphery, where the chromosomes (c) are localized. It is interpreted as representing the primary zone (Zachar et al., 1993). n = nucleolus. Scale bar = 5 pm. (B) Homogeneous accumulation of transcripts throughout the nucleus. (C) Later stage of transcript accumulation. Note the sharp drop of staining at the nuclear envelope, suggesting the retention of transcripts. (D) Nuclear + cytoplasmic export of transcripts appears to start simultaneously at all points of the nuclear envelope as suggested by the even cytoplasmic staining between the basal (b), lateral, and apical (a) sides of the cell. investigations, especially in the case of Drosophila, where genetic approaches are also feasible. It deserves, however, some deliberation on whether a polytene nucleus is representative of the somatic diploid interphase state and whether the results obtained with polytene nuclei are of general validity. There is accumulating evidence in D. melanogaster that favours the comparability of the systems. From transcript mapping data in the 87DE region of embryos, larval, and adult tissues it was concluded that “the banding pattern seen in polytene chromosomes reflects a level of organization which is not confined to these giant structures” (Bossy et al., 1984). Champlin et al. (1991) demonstrated that the binding of the B52 protein to subregions of the heat-shock puff 87AC is similar in tissue-culture cells and polytene chromosomes, and the results of Udvardy et al. (1985) suggest that the 2 nuclease-hypersensitive sites (scs and scs’) flanking the hsp70 genes, which they detected in chromatin from embryonic and cultured 82 H. Kress

cells, “define the proximal and distal borders of the 87A7 chromomere” in salivary polytene chromosomes. These findings substantiate the notion of Hochstrasser and Sedat (1987a) that “the banded structure of the polytene chromosome represents a direct geometrical amplification of a chromomeric organization in diploid chromatin”. Although this statement may not apply to features that are peculiar to polytene chromosomes (e.g. underpolytenized heterochromatin), the basic concepts for the elaboration of euchromatic structures in the inactive and active state either during replication or transcription should be the same. According to the current ideas in diploid interphase nuclei, replication occurs simultaneously at a few hundred foci, each containing multiple replication forks (see Coverley and Laskey, 1994). It is thought that “replication factories” (Hozak et al., 1993) are anchored in the nuclear matrix and that the chromosomal DNA is spooled through these fixed sites. In addition, the chromosomes themselves appear to be anchored at specific sites at the inner surface of the nuclear envelope, which represent focal points for decondensation and condensation of chromatin in the context of interphase activities (Hiroaka et al., 1989). The question of whether a similar situation exists in polytene nuclei cannot be answered at present. The problem may paradigmatically be discussed in the case of Drosophila, where the most relevant information is available. The positions of salivary gland chromosomes seem to be stabilized by multiple points of contacts to other components of the nucleus. Accordingly, replication must take place in chromatin, which is immobile at the chromosome level, but requires mobile elements at the subchromosomal level. In the case that there are fixed replication factories working on polytene chromosomes and the DNA is mobile, the spooling of hundreds of chromatids oriented in parallel alignment, even in small elements like replicons, appears to be a formidable task, because not only the DNA must be duplicated, but also parental and new nucleosomes and NHPs must be redistributed or be newly assembled under conditions that do not significantly alter the physical morphology of bands and interbands. This might represent one of the parameters causing the slow rates of replication in polytene chromosomes mentioned earlier. Alternatively, the mobile entity of replication could be formed by the replication factories, which would imply that in diploid and polytene nuclei different mechanisms of replication should work. Consequently, the developmental switch to endo-cycles would involve not only cell cycle controls but also a modification of the replication machinery. Interbands appear to be the “living” portions of the interphase chromosome, which are preferred sites of transcription, while bands may either bury silent genes or contain non-essential sequences. Puffs or Balbiani Rings may be interpreted as special forms of interbands, being produced by the above-average accumulation of proteins or of RNPs, which are effected either by high rates of transcription (e.g. at heat-shock loci) or by the prolonged retention of transcripts due to their extremely long size and/or processing time (e.g. early ecdysone-induced genes). The demonstrated, initiation of replication in interbands, the possible localization of promoter regions in these substructures and their low level of DNA compaction (compare Table 2), which facilitates the targeted access of truns-factors, suggest that they may also harbour the sites of initiation of replicative and transcriptional processes that are propagated into large chromomeres with high levels of compaction. Small chromomeres, whose compaction suggests their organization as the 30-nm solenoid (compare Table 2), may also contain sites in the open chromatin configuration, Polytene Chromosomes: Eucaryotic Interphase State Model 83 as it is exemplified by the 87A7 chromomere prior to heat-shock induction. It displays distinct patterns of nuclease and top011 cleavage sites in the central domain that encompasses the common promoter region of the 2 hsp70 genes (see Fig. 3A). Whether this preset domain functions as the initiator of chromomere decondensations during the heat-shock response, or whether the chromomere responds as a whole, remains to be elucidated. The identification of the scs and scs’ elements at the borders of this chromomere, which were interpreted as swivel points for chromatin transitions, would define the chromomere as a unit for decondensation/condensation reactions. This would represent the first example for the genetic demarcation of a chromomere as a functional unit of regulated packaging. However, Shemeshin et al. (1982) showed that, upon heat-shock, the 87A7 chromomere decondensates in concert with 3 flanking bands (87A6-9), which do not contain heat-shock genes. This and other cases (Shemeshin et al., 1985) suggest that domains of regulated packaging may comprise several contiguous bands, the problem of boundary formation being raised to the level of long-range effects. Chromatin boundaries that insulate long regulative units have been proposed for homeotic gene complexes (for a brief review, see Eissenberg and Elgin, 1991), and the binding patterns of PC-G proteins to corresponding sites that extend over several chromomeres (e.g. Zink and Paro, 1989) corroborate such ideas. It should also be noted that PC-G-related proteins involved in the long-range phenomenon of position effect variegation (PEV) have been interpreted as opening or closing large DNA domains (Reuter et al., 1990). The super replicons proposed by Mukherjee et al. (1980) could also be interpreted in these terms. The exciting results of the pilot studies on the D. melunoguster hsp70 heat-shock genes, and the demonstration of specific hnRNP or snRNP proteins or of the PEP protein at individual chromosomal sites, convincingly display the full power of polytene chromsomes for analyzing the temporal and spatial dynamics of the interactions of proteins with DNA and RNA in the context of transcription and RNA processing. We can anticipate that specific labeling patterns of polytene chromosomes will help to elucidate poorly characterized specific functions of other proteins in active and inactive chromatin. This applies, for example, to the roles of different classes of HMG proteins or to variants of histone Hl. In D. melunoguster, histone H4 molecules, acetylated at different lysine residues, exhibit differential distributions in eu- and heterochromatic regions, suggesting differing roles in packaging (Turner et al., 1992). Most interestingly, the preferential localization of H4 acetylated at lysine 16 to the X-chromosome of males coincides with the exclusive presence of a putative helicase (the muleless protein) at hundreds of sites on that chromosome (Kuroda et al., 1991). This is the first example to show that there is a correlation between the chromatin configuration of a whole chromosome (leading to hyperexpression of its genes), and the concomitant presence of distinct histone and non-histone proteins. In continuing these investigations, at both the genetic and cytological levels, it should be possible to elucidate the mystery of the functional relation between dosis compensation of sex chromosomal genes and chromatin structure. Another challenge for the future is the unraveling of the functional cooperativity of chromosomes and nucleus, which has attracted increasing attention during the past few years. For a long time, investigations on nuclei of vertebrate cells were hampered by the lack of intranuclear landmarks that allow the unambiguous fixing of the spatial coordinates for the positions of chromosomes and for RNA metabolism. They may have 84 H. Kress gained a new impetus now, by the postulation of chromosomal territories that are separated by the interchromosome domain compartment (ICD), where mRNA processing and transport are thought to occur (Zirbel et al., 1993; Cremer et al., 1993). The rather similar picture of the compartmentalization of the polytene nucleus in polytene chromosomes and the ICN, provides a promising formal basis to initiate comparative studies for testing the validity of the interpretations in diploid and polytene nuclei, and thus to clarify the present track vs. diffusion controversy.

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