Synaptonemal complexes in insects

Pergamon

Inr. J. Insect Morphol

& Embrwl., Vol. 25, No. 3, pp. 205-233, 1996 Copyright e 1996 Elsev~er Science Ltd Printed in Great Bntam. All nghts reserved 002&7322/96 $15 OO+O.OO

PII: SOO20-7322(96)00009-S

SYNAPTONEMAL

COMPLEXES

FrantiSek Institute

of Entomology,

Czech Academy

IN INSECTS

Marec*

of Sciences, BraniSovska

31, 370 05 eeskt

Budgjovice,

Czech Republic

(Received 20 March 1996; accepted 15 May 1996) Abstract-The synaptonemal complex (SC) is the key nuclear element formed in meiotic prophase I to join 2 homologous chromosomes at the pachytene bivalent. It is a highly conserved structure that is universally present in eukaryotes. The SC is presented as a tripartite protein structure, which consists of 2 lateral elements and a central region. In insects, the central region is particularly distinct and highly ordered. This made it possible to describe the fine structure of the central region and propose a model of its architecture. Chromatid DNA is arranged in chromatin loops extending radially from the SC. The loops appear to consist ofa basic chromatin fiber with a diameter of 20-30 nm. In many insect species, synaptonemal polycomplexes occur in postpachytene cells. They represent one of the possible ways of SC degradation. Another process, which occurs beyond pachytene, is the formation of proteinaceous chromatid axis, the silver-stained chromatid core. Based on results in insect models, the chromatid cores have been related to the structure and formation of the SC. Research on insect models significantly contributed to understanding individual steps of the SC formation and temporal sequence of chromosome pairing. These include the formation of lateral elements of the SC, pairing initiation, interlocking of chromosomes, and synapsis of homologous chromosomes. Attention is also given to non-homologous pairing, including synaptic adjustment, correction of pairing, and pairing of sex chromosomes. In the next section, chiasmatic and achiasmatic modes of meiosis are compared with respect to the SC formation. In the chiasmatic mode, the SCs display recombination nodules that are believed to mediate the process of recombination. These nodules were discovered in insects, and indirect evidence for their role comes from insects. Two different examples of achiasmatic meiosis, occurring in the heterogametic sex of several insect orders, are given: one involves the SC formation, whereas in the other, SCs are absent. Finally, the potential of SC karyotyping for analysis of the insect genome is discussed. Copyright c> 1996 Elsevier Science Ltd Index descriptors (in addition to those in the title): Meiotic prophase I, chromatin loops, recombination nodule, polycomplexes, chromatid core, chromosome pairing, interlocking, synaptic adjustment, sex chromosomes, achiasmatic meiosis, karyotyping.

INTRODUCTION

Forty years ago, a unique nuclear structure, joining 2 homologous chromosomes at the pachytene bivalent, was independently described by Moses (1956) in crayfish primary spermatocytes and by Fawcett (1956) in spermatocytes of pigeons, cats, and humans. This structure, termed the synaptonemal complex (SC), was later found in a variety of organisms ranging from yeast to man. It has been shown that the SC plays a central role in meiosis,

*Tel: +49-38-817,

Fax: +49-38-43625,

e-mail: [email protected] 205

F. Marec

206

and is highly conserved in structure and dynamics during the meiotic prophase I (von Wettstein et al., 1984; Holm, 1985; John, 1990; Loidl, 1990; Verma, 1990). The SCs play an essential role in the pairing of homologous chromosomes, which is the key event in meiosis of diploid organisms, and on which the subsequent meiotic events, i.e. recombination and chromosome segregation, depend (John, 1990). The SC mediates the intimate association of 2 homologs throughout their entire length. This enables the chromosome regions to come in to close contact and verify their allelic homology that is necessary for performing the meiotic crossing-over. Within the mechanical framework provided by the SC, precise DNA sequence matching takes place, which guarantees reciprocal strand exchanges at exactly corresponding DNA sites in the course of crossing-over (Rasmussen and Holm, 1982; Loidl, 1990, 1994). SC precursors, the axial elements, may play a supportive role in the homology search process (see Loidl, 1994). It has been postulated that SC formation is responsible for detecting and resolving interchromosomal tangles (interlockings) in early meiotic prophase (Rasmussen, 1986). It has also been proposed that the SC participates in organizing chiasmata (see Rasmussen and Holm, 1982; and references therein). Forming bivalents of homologous chromosomes is another important role of SCs. The bivalents are, in later stages of meiotic prophase I, maintained by chiasmata or other means until metaphase I (e.g. Wolf, 1994a) to ensure that homologous chromosomes separate from each other in an orderly manner. This is essential for providing each gamete with one complete haploid chromosome set (Loidl, 1994). Thus, the SC has 2 main functions: (1) it is a prerequisite for reciprocal crossing-over, and (2) it indirectly ensures proper disjunction of homologous chromosomes. However, this classical view is challenged by recent genetic and molecular evidence, mainly from yeast, which suggests that crossovers are initiated prior to synapsis and that they can occur in the absence of SCs. In this model, the SC has the function to stabilize early recombination intermediates and to transform them into functional chiasmata (Loidl, 1994; and references therein). In insects, pioneer contributions to understanding ultrastructural organization and function of SCs were done in Drosophila (Koch et al., 1967; King, 1970) Locusta (Moens, 1969), and Bombyx (King and Akai, 1971). Thus, it appears worthwhile to cover the literature on SCs in insects in our article.

MORPHOLOGY

OF

THE

SYNAPTONEMAL

COMPLEX

Since its discovery in 1956, the SC has been studied by transmission electron microscopy (TEM) through embedded meiocytes. This enabled cytogeneticists to describe its ultrastructural organization (e.g. King and Akai, 1971). Three-dimensional reconstruction of serial sections from meiocytes was then employed to investigate the SC formation as well as its function in chromosome pairing, crossing-over, and conversion into chiasma (for a review see Holm, 1985). The reconstruction of SCs in a nucleus from serial sections represents an extremely laborious and time-consuming process, although it has the great advantage of preserving nuclear topology. The introduction of simpler and more rapid microspreading techniques has made the study of SCs and the pairing process much more straightforward (see Sumner, 1983; John, 1990; Loidl, 1990). First, animal meiocytes were spread on the surface of a

Synaptonemal

Complexes

in Insects

207

saline solution and fixed with formaldehyde (Counce and Meyer, 1973). Then SCs were selectively displayed with phosphotungstic acid on whole mount preparations for TEM. This technique allowed a one-dimensional visualization of whole SCs. Later, a silver staining technique was devised for the surface-spread preparations (e.g. Moses et al., 1979). This technique has been particularly useful for studying the pairing process, including behavior of sex chromosomes in the heterogametic sex, and for detection of pairing abnormalities. Another spreading technique was introduced by Miller and Bakken (1972). Meiocyte nuclei were spread in a detergent, centrifuged onto EM grids, fixed with formaldehyde, and stained either with uranyl acetate or with phosphotungstic acid. The centrifugation spreading technique yields preparations of SCs with associated chromatin. This has proved useful for visualization of the fine structure of heterochromatin (Weith and Traut, 1980) and for investigating the organization ofchromatin loops (Rattner et al., 1980,198l). This technique was also used to visualize the polyfusomes in insect ovaries and testes (Marec et al., 1993). Scanning electron microscopy has been rarely used to study SCs (Barlow et al., 1993; Wolf et al., 1994). Recently, several novel microscopic techniques have been employed in the SC research. For instance, the morphology of SC was studied using atomic force microscopy (Putman et al., 1993). Another technique, electron microscope tomography, has made it possible to produce a three-dimensional picture of the SC (Schmekel et al., 1993a). Immunocytochemistry (immunofluorescent or immunogold labelling) and in situ hybridization (Sumner, 1983) are now also available for analysis of the SC using light and electron microscopy. These techniques enable us to identify the location of specific SC proteins (Dobson et al., 1994; Heng et al., 1994).

Gross morphology and composition The fully formed SC is located between the paired meiotic chromosomes at pachytene as a ribbon-like structure along the entire length of each bivalent (Fig. 1). It consists of 2 lateral elements (LEs), which are considered to be the protein axis of individual homologous chromosomes. At the ends of LEs, terminal points are visible as globular structures. In a medial position between the 2 LEs, there is a central element (CE) that is connected with the LEs by fine transverse filaments (TFs), perpendicularly oriented to the longitudinal axis

Fig. 1. A schematic model of SC organization. CE =central element; Ch =chromatid; GTD = globular terminal differentiation; LE = lateral element; RN = recombination nodule; TF = transversal filament. Chromatin loops of the 2 sister chromatids are associated with each LE.

208

F. Marec

of the SC. In many insects, the CE is relatively wide, highly ordered, and resembles a ladder, whereas in mammals, it appears thinner and amorphous (see Schmekel and Daneholt, 1995). In organisms with the chiasmatic mode of meiosis, one to several small electrondense bodies, the so-called recombination nodules, are associated with the central region of almost each SC (Holm and Rasmussen, 1980; John, 1990). These nodules are roughly spherical or ellipsoidal and vary considerably in size with an average of about 100 nm (Carpenter, 1975b; Holm and Rasmussen, 1980). Chromosomal DNA of each chromatid is folded into discrete chromatin loops that are, at their bases, attached to LEs and extend radially (Fig. 2). Both sister chromatids of each homologous chromosome are associated with one LE. The single-stranded LE is sometimes partly split into 2 strands, particularly in microspread preparations (in insects see, e.g. Weith and Traut, 1980; Wang et al., 1993). This could mean that the LE is composed of 2 filaments corresponding to the sister chromatids (Weith and Traut, 1980; Wahrman, 198 1). Images of the SC, recently obtained in rats using the atomic force microscope favour such a double structure of LEs (Putman et al., 1993). Using different EM techniques, Dietrich et al. (1992) have shown that the LE of rat SCs appears to be subdivided into 3 longitudinal components: 2 major lateral strands and a 3rd thinner strand, localized on the inner side of the LE. The authors suggest that the latter strand could be the connecting element between TFs and LEs. However, numerous ultrastructural investigations in insect models have not disclosed such a triple structure of the LE (cf. King and Akai, 1971; Carpenter, 1975a; Rasmussen, 1976; Schmekel et al., 1993a, 1993b). The centromere region is not so prominent in pachytene bivalents as in metaphase chromosomes, but can be recognized as a lightly stained ball of fine fibrils through which the SC passes uninterrupted (John, 1990). In surface-spread pachytene spermatocyte nuclei of a grasshopper, Chorthippus jacobsi, centromere regions appear as irregular, thickened structures at submedial position in 3 submetacentric bivalents and close to one of the terminal ends in 5 acrocentric bivalents and in the X univalent (Santos et al., 1993). Spread SCs of another grasshopper, Pyrgomorpha conica, with all telocentric chromosomes, show centromeres as terminal knob structures (de1 Cerro et al., 1996). In organisms with relatively large kinetochores (i.e. with holokinetic organization of chromosomes), namely the Lepidoptera, an element, which could be related to the kinetochore region, has not been detected in pachytene bivalents (Rasmussen, 1976; Weith and Traut, 1980). The precise chemical composition of the SC is not fully known. The SC can be completely digested by trypsin, but remains intact after DNase treatment. This is strong evidence that it is predominantly proteinaceous (Westergaard and von Wettstein, 1970). Persistence of SC proteins after removal of DNA and histones from nuclei suggests an association of SCs with the inner nuclear matrix (Comings and Okada, 1976; Raveh and Ben-Ze’ev, 1984). Results of other experimental approaches employing selective staining, differential extraction, and enzyme treatment, indicate a high content of basic arginine-rich, non-histone proteins (Dresser, 1987). Thus, basic non-histone proteins together with a certain fraction of chromosomal DNA probably constitute a large part of the SC. It has also been suggested that the SC contains some RNA (Chevaillier, 1974). A radical change in the research of SC composition has arrived when monoclonal antibodies have been developed, which recognize specific SC-proteins (Moens et al., 1987; Heyting et al., 1987, 1989). Recently, a number of SC-specific and SC-associated proteins have been identified in yeast, Saccharomyces cerevisiae (Sym et al., 1993; and references therein), and in mammals (Dobson et al., 1994; and references therein). The biochemical composition of recombination nodules that are

Synaptonemal

Complexes

in Insects

209

Fig. 2. Microspread SCs of a female moth, Ephestia kuehniella, stained with phosphotungstic acid. Parallel lateral elements are surrounded by a halo of chromatin loops. Bar=3 pm. [Unpublished material.] Fig. 3. (a,b) The sectioned SC in a beetle, Slaps cribrosa. (a) Frontal view of the SC; CE=central element; ch = surrounding mass of chromatin; LE = lateral element. (b) Cross-sectional view with 4 CE layers marked (arrows). Bars= 150 nm. [From Schmekel ef al., 1993a; reprinted with permission of Springer-Verlag, Heidelberg.]

believed to represent multi-enzyme complexes is not known. Their strong staining with phosphotungstic acid indicates a high content of basic proteins (Holm, 1985). Chromatin represents a complex consisting of the DNA molecule and basic proteins, the histones (Verma, 1990). The DNA is arranged in loops extending from the SCs. Only a small amount of DNA is directly associated with the LEs. Rattner et al. (1981) estimated that this variety represents about 0.08% of the DNA in Bombyx mori spermatocytes. Others claimed that about 0.12% of DNA is not in loop form in B. mori oocytes. The value for Drosophila melanogaster oocytes is about 0.2% (table 4.4 in John, 1990). It is an open

210

question whether the fraction or may have special properties

F. Marec

of DNA associated with the SC contains (Holm, 1985; Moens, 1994).

specific sequences

Fine structure

At the ultrastructural level, the SC has been thoroughly studied using three-dimensional reconstructions of serial sections through pachytene meiocytes in several insect species. These are the silkworm B. mori (King and Akai, 1971; Rasmussen, 1976; Rasmussen and Holm, 1982), D. mehnogaster (Carpenter, 1975a, 1975b), and a beetle Slaps cribrosa (Schmekel et al., 1993a). In longitudinal sections through the SC (Fig. 3a), the LEs appear as strands of electron-dense amorphous material, about 3040 nm thick (Rasmussen and Holm, 1982). The LEs are often not clearly distinguishable from the lateral chromatin. Carpenter (1975a) measured the distance between LEs in a large number of SCs in D. melanogaster females, and found that the central region is relatively constant within and between oocytes: 109 f 8 nm wide (n = 97). A similar distance between LEs was reported for B. mori spermatocytes: loo-120 nm (King and Akai, 1971; Rasmussen and Holm, 1982). The central region in B. mori oocytes is, however, narrower: 70-80 nm wide. This difference may be related to the lack of crossing-over in B. mori females (Rasmussen and Holm, 1982). In insects, the CE has a scalariform (King and Akai, 1971) or ladder-like (Schmekel et al., 1993a) substructure, and occupies approximately one-third to one-half of the space between LEs. Schmekel et al. (1993a) reconstructed the entire central region by means of electron microscope tomography in B. cribrosa (Coleoptera) which exhibits a particularly distinct CE and highly ordered TFs (Fig. 3a). In this beetle, the CE is 35-nm wide, and the width of the entire central region is about 90 nm. The total height of the CE is 200 nm. In cross-sectional views, the CE appears subdivided into 3 or 4 layers (Fig. 3b). Both LEs are linked to each other by regularly spaced TFs passing from one LE across the central region and through the CE to the opposite LE. The TFs are thin fibers usually 5 nm in diameter, although they can range from 2 to 10 nm. Moreover, they tend to split into 2 or more thinner fibers, particularly in the vicinity of LEs. Therefore, the authors suggested that the TFs consist of bundles of finer fibers, the elementary fiber being 2-nm thick. In the proposed model of the central region in B. cribrosa (Schmekel et al., 1993a; Schmekel and Daneholt, 1995; Fig. 4) the CE is built like a ladder with 2 longitudinal CE components and a large number of regularly spaced, transverse CE components. The CE is multi-layered. In B. cribrosa, it is composed of 3 to 4 distinct layers, each corresponding to one TF. In a CE layer, short pillar-like structures form the junctions between the longitudinal and transverse CE components. The pillars are oriented perpendicular to a frontal plane through the SC. In addition, the individual layers are connected by occasional vertical fibers linking pillars on the top of each other and keeping the layers in approximate register. This finally results in a three-dimensional, well-defined network. Each TF is associated with 2 pillars of a CE layer. The TF from the LEs, crosses the CE and appears to pass through the center of each pillar. It has been proposed that a TF with 2 associated pillars represents the basic structural unit in the central region of the SC. In another study (Schmekel et al., 1993b), it has been shown that the central region of D. melanogaster contains the same structural unit and exhibits a similar architecture to that of B. cribrosa. The same basic structural units have also been found in the rat. In the mammalian system, however, individual components of the central region of the SC are less frequent and less organized. This gives the entire central region in the rat amorphous and poorly defined

Synaptonemal

Complexes

in Insects

211

Fig. 4. A schematic drawing of the three-dimensional model of the central SC region in a beetle, Blaps cribrosa. The structural unit, consisting of a transverse filament (TF) and 2 symmetrically arranged pillars, is shown in the upper left corner. The central region of the SC is schematically displayed with a large number of structural units organized side by side into 3 layers stacked in register. A fibrous network connects the pillars of the individual units. The central element (CE) is demarcated, and the lateral elements (LEs) and the surrounding chromatin (ch) are depicted as plates. The frontal (F), lateral (L), and cross-section (C) views are indicated. Bar=50 nm. [From Schmekel and Daneholt, 1995; reprinted with permission of Elsevier Trends Journals, Cambridge.]

appearance. Nevertheless, these results region could be more widely valid.

indicate

that the proposed

model

of the central

Polycomplexes Polycomplexes (PCs) are elements that occur in postpachytene cells of various organisms. They may lie in the nucleus or in the cytoplasm. Typically, they consist of stacked SC fragments (Fig. 5). The structure of PCs corresponds to that of SCs, in that they contain alternating LEs and central regions. The latter possess a CE in a medial position. PCs are interpreted as self-assembly products of SC fragments discarded from the pachytene bivalents (for reviews see Goldstein, 1987; Verma, 1990). In late prophase I of meiosis, the SCs disassemble in various ways. One may be the formation of PCs. SCs fragments are freed from the bivalents and may associate to form aggregates, the PCs (John, 1990). It has been suggested that PCs prevent involvement of previously functional SC material in chromosome segregation (Goldstein, 1987). PCs may occur either free or attached to chromatin or other components (see Verma, 1990). Their frequent association with nucleoli in degenerating nuclei of D. melunogaster nurse cells has been taken to indicate that the nucleolus is involved in the synthesis and assembly of the PCs (Rasmussen, 1975). In insects, PCs are frequently found in late prophase I both in oogenesis and spermatogenesis. For instance, they were observed within the nuclei of mosquito oocytes (Fig. 5; Fiil and Moens, 1973). Rasmussen (1976) reported polycomplex-like structures in

212

F. Marec

Fig. 5. EM micrograph of the sectioned polycomplexes in an oocyte nucleus of a mosquito, Aedes aegypti. CE =central element; LE = lateral element. Bar= 0.5 pm. [From Fiil and Moens, 1973; reprinted with permission of Springer-Verlag, Heidelberg.] Fig. 6. EM section through young spermatid nucleus of a cricket, Eneoprera surimmensis, displaying a polycomplex (arrow). N = a nucleolar body. Bar = 1 pm. [Courtesy of K. W. Wolf, Ltibeck, Germany.] Fig. 7. A portion of the young spermatid nucleus of a cricket, Eneoptera surinamemis, displaying a synaptonemal polycomplex with regularly stacked lateral elements (LE) and central regions (CR) of the SCs. Bar=250 nm. [Courtesy of K. W. Wolf, Ltibeck, Germany.] Fig. 8. Silver-stained early metaphase I bivalent of a male grasshopper, Arcyp/eraJiisca. The chromosome cores, each consisting of 2 associated chromatid cores, are visible as deeply stained axes of each homolog. The cores do not reach the chromosome ends, and are enlarged at their distal tips, thus forming telochores (T). In each chromosome, the joined sister kinetochores (K) appear as a unique round structures. Bar= 10 pm. [From Suja and Rufas, 1994; reprinted with permission of Rapid Communications, Oxford.]

degenerating nuclei of nurse cells in B. mori ovaries. In another Lepidoptera species, Ephestia kuehniella, PCs occurred in nurse cell nuclei as well (Raveh and Ben-Ze’ev, 1984). PCs have also been described in male meiosis of the crane fly, Pales ferruginea (Diptera)

Synaptonemal

Complexes

in Insects

213

(Fuge, 1979). A number of studies reported PCs in several Orthoptera species (see Wolf and Mesa, 1993; and references therein). In this group, PCs seem to persist later than in other insects. The PCs are embedded in metaphase I bivalents or found within the cytoplasm of spermatids (Esponda and Krimer, 1979; Moens and Church, 1979). In a recent study, Wolf and Mesa (1993) have shown that PCs regularly persist through both meiotic divisions and come to lie within the nuclei of young spermatids (Figs. 6 and 7) in a cricket, Eneoptera surinamensis (Orthoptera). Transfer of the PCs into the nuclei may be caused by their adhesion to chromosomes. In spermatid nuclei, PCs were often attached to a spherical nuclear body of unknown nature. The significance of the PCs persistency into young spermatids remains unclear, but the authors speculated that disassembly products of the PCs may play a role in spermatid maturation. The persistency of PCs into young spermatids seems to be typical for Orthoptera, but this phenomenon may also occur in other systematic groups. Recently, Dallai and Afzelius (1995) found PCs closely connected to a dense mass of chromatin in young spermatid nuclei of a caddis-fly, Hydroptila aegyptia (Trichoptera). Chromatin organization During meiotic prophase I, the chromatin organization changes dramatically. Starting in leptotene, the chromatid DNA becomes folded into loops that are attached to a protein core (Moens and Pearlman, 1988; Moens, 1994). This core probably represents a matrix for the synthesis of LE proteins. For the study of chromatin organization in insects, a microspreading technique of Miller and Bakken (1972) or its modification (Weith and Traut, 1980) have been used (see introduction to Section 2). The technique basically consists of the treatment of meiocytes with a detergent that dissolves both the cellular and nuclear membranes, and disperses the nuclear content. The spread material is then fixed onto EM grids by centrifugation. The subsequent contrasting either with uranyl acetate or with phosphotungstic acid allows the visualization of SC proteins, chromatin representing the DNA-histone complex as well as ribonucleoproteins (RNPs) (Rattner et al., 1981). It is evident from EM preparations of spread meiocytes of E. kuehniella and B. mori that the chromatin forms a series of closed loops radially extending from the LEs of the SC. In B. mori, the loops consist of a basic chromatin fiber with a diameter of 2&30 nm (Rattner et al., 1980, 1981) while in E. kuehniella, thinner fibers with a diameter of about 13 nm on average were found (Weith and Traut, 1980). The fibers have the typical “beads-on-astring” appearance of nucleosomes on a connecting thin DNA fiber. Conformation of chromatin is, however, dependent upon the preparative conditions (see Rattner et al., 1980). The different spreading procedure used in E. kuehniella could thus result in a relaxation of the basic chromatin fiber (20-30 nm) to a nucleosomal fiber (about 10 nm in diameter, see Verma, 1990). This opinion is based on our experience with spread preparations from 3 Lepidoptera species, E. kuehniella, the wax moth Galleria mellonella and the tobacco hornworm Manduca sexta (Marec, unpublished). In particular, longer treatment of meiocytes with a hypotonic solution or then with a detergent in order to disperse individual SCs may dissociate pachytene bivalents as well as conformation of chromatin. Whereas an optimal treatment produces spread SCs (see Fig. 2) and loops consisting of the basic chromatin fiber. We suggest that such an arrangement of chromatin represents a universal mode of DNA folding during early meiotic prophase I. Most chromatin loops associated with the SC are transcriptionally inactive. In E. kuehniella as well as in B. mori, only a few loops displayed active transcription units. Short

214

F. Marec

stretches of the loops possessed lateral ribonucleoprotein (RNP) fibrils (Weith and Traut, 1980; Rattner et al., 1980). In another study, Rattner et al. (1981) observed in a bivalent associated with the nucleolus 1 or 2 prominent clusters of loops with adherent RNP transcripts and interpreted them as loops carrying actively transcribed ribosomal DNA cistrons. Tangles of heterochromatin, described in E. kuehniella females, are another prominent differentiation of the chromatin visible in microspread pachytene nuclei (Weith and Traut, 1980; Marec and Traut, 1994). They are exclusively associated with the heterochromatic W chromosome. The chromatin fibers in the tangles are condensed and form several compact electron-dense centers consisting of globular particles with a diameter of about 40 nm. These particles are connected by a thin fiber and are embedded in an amorphous mass. The tangles disperse with longer treatment during spreading procedure. Rattner et al. (1981) attempted to estimate the density of chromatin loops during male meiosis in B. mori. They calculated approximately 7000 loops within the haploid genome. The total length of SCs in B. mori males is about 260 pm, thus there are about 26 loops/pm of the SC. The maximal size of the loops was 2.5 ,um. The authors also found that individual attachment sites on the LEs are separated during early prophase by 0.15-0.20 pm of nucleosomal DNA. An open question is whether similar chromatin organization applies to other insects. We have recently measured the density of chromatin loops in microspread oocytes of E. kuehniella (Marec, 1995; details unpublished). In late zygotene-pachytene nuclei, the loop density varied in the range of 2432 loops per 1 pm of the SC with an average of 29 loops. The loop size varied from 2 to 3 pm. The mean total length of SCs in E. kuehniella females is 160 pm (Marec and Traut, 1993b), thus we estimated about 4600 loops per haploid genome. From these data we may conclude that the loop density as well as the loop size are similar in the 2 Lepidoptera species examined. The B. mori and E. kuehniella genomes differ considerably in the total SC lengths. This difference, however, corresponds to their different genome sizes. The DNA content of B. mori is 0.52 pg/lC DNA (Berry, 1985) while that of E. kuehniella is 0.3 pg/lC DNA (W. Traut, personal communication). The above described organization of chromatin loops in pachytene may be typical, at least for Lepidoptera that possess small chromosomes and a relatively low DNA content. Moens and Pearlman (1988) reported a similar loop size (3 pm) in another moth, Hyalophora columbia, but they highlighted the different loop sizes in the DNA-rich grasshopper, Chloealtis conspersa. In this species, the loops are enormously large: about 14 pm from base to top. Relationship between the synaptonemal complex and chromatid cores In early diplotene, SCs dissociate from the bivalents in different ways (see Holm, 1985; John, 1990). In B. mori males, for example, both LEs and CEs are shed from the bivalents as a structurally amorphous mass that degrades subsequently; only short remnants of intact SCs remain associated with early chiasmata and disappear before diakinesis (Holm and Rasmussen, 1980; Rasmussen and Holm, 1982). The formation of multiple sheets of aggregated SCs, the so-called polycomplexes, is another way that is frequently observed in insects (see section on polycomplexes). Another process, which occurs beyond pachytene, is the formation of a non-histone proteinaceous axis in each chromatid. This axis can be visualized is termed the axial silver-stained by silver impregnation techniques, and therefore, chromatid core. Based on indirect evidence (see Suja et al., 1991) the meiotic chromatid cores have been related to the scaffold observed in histone-depleted mitotic chromosomes.

Synaptonemal

Complexes

in Insects

215

The SCs and the chromatid cores have several features in common. The SCs and the cores are axial chromosome structures that run along the chromosome. The LEs and the cores possess a high content of non-histone proteins, and appear as dark threads in silverstained preparations. Finally, each LE of the SC displays globular terminal structures at the ends. Similar spherical differentiations have recently been described at the distal ends of chromatid cores in grasshopper spermatocytes (Suja and Rufas, 1994). The authors refer to these differentiations as “telochores” (Fig. 8). There are, however, some differences between the SC and the chromatid cores. First, there is one core per chromatid, whereas one LE is always associated with 2 sister chromatids. Secondly, the SC represents a highly organized meiotic organelle consisting of several specific components, whereas the core is relatively simple, uniformly stained and ill-defined. Thirdly, while the SC function is restricted to early meiotic prophase I, the chromatid core seems to have a function in the structure of both the meiotic and mitotic chromosomes (Suja et al., 1991; Rufas et al., 1992). Rufas et al. (1992) analyzed silver-stained SCs and chromatid cores in 2 grasshopper species of the genus Chorthippus, in order to determine the relationship between the SC and the chromatid core. They have proposed a model of meiotic chromosome organization with a possible role of the chromatid core in the formation of the SC (Fig. 9). Providing that the scaffold/radial loop model of chromatin organization applies both to the mitotic and meiotic chromosomes, the cores of sister chromatids would associate intimately during leptotene. The associated cores could act as attachment sites for specific LE proteins. Helicoidal arrangement of the cores would then bring about chromatin condensation throughout the prophase I. The LE proteins would be released throughout late prophase I which would result in the disappearance of the SC. Subsequent chromatid condensation would render the chromatid cores visible.

Fig. 9. A schematic mode1 of the relationship between the chromatid cores and the SC elements. For simplicity, only 1 homolog is drawn. CC=chromatid core; Ch=chromatid; LE=lateral element; RL=radial loop; TF= transversal filament. [From Rufas et al., 1992; with a slight modification; reprinted with permission of NRC Research Press, Ottawa.]

216

F. Marec

FORMATION

OF THE

SYNAPTONEMAL

COMPLEX

Early meiotic prophase

During early meiotic prophase I, the LEs appear as a chromosomal axis made of proteins. Because one LE is formed per leptotene chromosome, both sister chromatids must be associated with it. Heyting et al. (1989) have suggested that LE proteins are newly synthesized and are not assembled by a rearrangement of pre-existing components. In insects, our knowledge about the beginning of SC formation in early prophase I are limited to data obtained by three-dimensional reconstruction of leptotene-mid zygotene nuclei in the silkworm, B. mori (Rasmussen, 1976; Holm and Rasmussen, 1980). In oocytes and spermatocytes of the silkworm, the formation of LEs starts at the telomeres. First, the LEs are recognizable as individual short pieces localized inside the chromatin. Then, the LEs become attached by their telomeres to the inner membrane of the nuclear envelope through cone-shaped modifications. The attachment sites are preferentially located at one pole of the nucleus opposite the nucleolus, and near the 2 pairs of centrioles. In some organisms (e.g. the fungus Sordaria, the plants Lilium IongiJlorum and Zea mays), a continuous LE is assembled before SC formation (see Holm, 1985; and references therein), while in oocytes and spermatocytes of B. mori (Rasmussen, 1976; Rasmussen and Holm, 1979; Holm and Rasmussen, 1980) initiation of pairing and SC formation precede the completion of LE formation. This becomes evident in early zygotene, when the first short segments of SC appear near the nuclear envelope but many LEs are still discontinuous (Rasmussen, 1976). The LEs are almost continuous at mid zygotene, the stage characterized by the appearance of the first completely paired bivalents. During this interval, all the telomeres have moved to a specific region of the nuclear envelope. The polarized distribution of the attachment sites gives rise to a typical chromosome bouquet (Fig. 10). This bouquet configuration is maintained until early pachytene. In mid to late pachytene, the bouquet configuration can no longer be recognized and the attachment sites of the telomeres are redistributed evenly on the nuclear envelope (Rasmussen and Holm, 1979; Holm and Rasmussen, 1980). Rasmussen and Holm (1982) suggested that 2 processes initiated in leptotene, namely, the telomere association with the nuclear envelope and the aggregation of the attachment sites, may facilitate the initial pairing of the homologous chromosomes. The former process reduces the freedom of telomere movement, and the latter ensures that the telomeres, including homologous telomeres, are brought into proximity. The temporal sequence

of chromosome

pairing in zygotene

andpachytene

The zygotene state of meiosis is characterized by initiation of chromosome pairing and subsequent bivalent formation finally resulting in close connection of homologous chromosomes by the SC at the onset of pachytene. Chromosome pairing probably consists of 2 independent processes, namely, the search of chromosomes for their homologous partner and the proper pairing of the homologs (Loidl, 1990). The mechanisms responsible for bringing together the homologous chromosomes and for their homology recognition are largely unknown. There are hints that the homologs are brought into proximity by random rather than by directed movements (see Holm and Rasmussen, 1980; and references therein). Homologous interactions during meiosis may be facilitated by a partial premeiotic alignment of the homologs observed in some eukaryotes, for example in fission yeast, Schizosaccharomyces prombe (Scherthan et al., 1994; and

Synaptonemal

Complexes

in Insects

217

references therein). However, there are many controversial observations on the concept of premeiotic association as a means of ensuring preferential homologous synapsis (see Loidl, 1990). Loidl and Langer (1993) have constructed mathematical models of homolog search, based on homology recognition sites. These sites could be conceived as specialized DNA sequences distributed throughout the genome. Such a direct DNA-DNA interaction is widely accepted as the mechanism for recognition of homology. In addition, proteins associated with the DNA, which have been identified in various organisms, might play a role in the recognition of homology (for a review see Heyer, 1994). Once homology has been detected, chromosome pairing and SC formation can take place. During this process, 3 subsequent steps can be distinguished: (i) initiation of pairing and SC formation; (ii) interlocking and resolution (see later); and (iii) synapsis of the homologous chromosomes along their entire lengths. In addition, a rough alignment of homologous chromosomes (“presynaptic alignment”) is observed in many organisms prior to the formation of the SC (Holm, 1985; John, 1990; Loidl, 1990). Pairing initiation. In general, organisms with short SCs preferentially exhibit telomeric initiation of pairing, while in species with long SCs, additionally interstitial initiation of pairing is found (Holm, 1985). For example, telomeric initiation of synapsis was found in both B. mori oocytes (Rasmussen, 1976) and spermatocytes (Holm and Rasmussen, 1980); interstitial initiation also occurred and this happened in cases where movements of homologs were impeded by interlockings (Rasmussen, 1986). Telomeric SC initiation was almost exclusively observed in polyploid B. mori meiocytes (Rasmussen, 1977b, 1987; Rasmussen and Holm, 1979). SC initiation near the telomeres was also reported by Jones and Croft (1986) in 2 grasshoppers, Locusta migratoria and Schistocerca gregaria, despite several large chromosome pairs in their genome. A similar situation has recently been described in another grasshopper, Chorthippus jacobsi (Santos et al., 1993). In the submetacentric bivalents, SC formation was initiated from both chromosomal ends, whereas in acrocentric bivalents 2 potential initiation sites were located at the distal and proximal regions of the long arms, the short arms remaining as late pairing regions. Interstitial SC occurred in long and middle bivalents at a relatively low frequency (3.21 interstitial initiations of pairing per nucleus). Rasmussen (1986) studied the process of pairing initiation in spread and silver-stained chromosome complements from diploid B. mori spermatocytes. He found that the initial alignment of homologs, which precedes SC formation, is mediated by a short subterminal segment, the so-called recognition site, at each end of the chromosome. SC formation was initiated at the subterminal regions and then proceeded, in most cases, to the nearest telomere. Less frequently, SC formation proceeded in the opposite direction. He also showed in tetraploid spermatocytes of B. mori (Rasmussen, 1987) that individual chromosomes are capable of associating with more than one homolog at a given recognition site. An earlier study of pairing in triploid B. mori oocytes revealed trivalents, in which one homolog was associated at one or both ends to a fully synapsed pair (Rasmussen, 1977b), thus confirming the existence of 2 recognition sites per chromosome. Taken together, Rasmussen’s observations clearly indicate that the specific pairing of homologous chromosomes, at least in B. mori, is the consequence of initial recognition of short subterminal sites rather than the result of an absolute requirement for homology of chromosome regions. Interlocking and resolution. SC interlocking was found to be a regular feature of zygotene pairing (see Holm, 1985; John, 1990). It occurs when synapsis of 2 homologous chromosomes is initiated at 2 different sites and another chromosome or bivalent is located

218

F. Marec

Fig. 10. Reconstruction of a mid zygotene nucleus from serial sections through Bombyx mori spermatocytes. All LEs and early formed SCs are arranged into a bouquet configuration. Two conspicuous electron-dense structures represent the 2 nucleoli. [From Helm and Rasmussen, 1980; reprinted with permission of Springer-Verlag, Heidelberg.]

between their unpaired regions (Fig. 11). Then the chromosome or bivalent is trapped and an interlocking is formed. Hence, 2 forms of interlockings can be distinguished (Holm and Rasmussen, 1980): (1) chromosome interlocks, where one LE of a partially paired bivalent is trapped between the 2 homologs of an unpaired interstitial segment of another bivalent, and (2) bivalent interlocks, where both the LEs are trapped in another bivalent. The phenomenon was first discovered in B. mori oocytes, where 3 of 4 reconstructed late zygotene nuclei contained a total of 6 interlockings (Rasmussen, 1976; Rasmussen and Holm, 1982). Four interlockings per nucleus were observed in late zygotene spermatocytes (Holm and Rasmussen, 1980). The findings have been confirmed in surface spread preparations: 4.2 per zygotene nucleus. In pachytene nuclei, the frequency of interlockings was very low (Rasmussen, 1986). Similar data have been reported for spermatocytes of Ch. jacobsi (Santos et al., 1993). In this grasshopper, most interlockings were resolved throughout zygotene. The estimated mean number of interlockings in this system was 1.6 at zygotene but only 0.17 at early pachytene. The nearly complete absence of interlockings in pachytene led to a conclusion that interlockings occur at zygotene and get mainly resolved before entering the pachytene stage (Rasmussen, 1986; Santos et al., 1993). It has been determined that the interlockings are resolved by the break-reunion-repair mechanism, not directly related to the DNA break-repair activities (Holm, 1985; Rasmussen and Holm, 1982). This process starts by breaking one of both LEs. The LE ends stay joined, because discontinuous LEs are rarely found in pachytene (Holm and Rasmussen, 1980; Rasmussen, 1986). Rasmussen (1986) suggested that resolution of interlockings occurs in 2 phases: (1) the release/passage of the interlocked LE and the reestablishment of the LE continuity, and (2) the passage of the chromatin of the involved chromosomes. The biochemical control of the break-repair activity capable of resolving chromosome interlockings is not known. It is probable that an ATP-dependent type II topoisomerase, which is known to be able to pass one double helix through another by a

Synaptonemal

Complexes

in Insects

219

Fig. 11. Bivalent interlocking in a microspread spermatocyte nucleus of Ephestia kuehnidlu. A fully paired bivalent (arrow) is entrapped between the unpaired middle region of another bivalent. Bar= 3 pm. [Unpublished material.] Fig. 12. A microspread bivalent of a female nucleus in Ephesfia kuehniellu, displaying the terminal modification (TM) of lateral elements at the distally located NOR segment. The TMs seem to impede full synapsis of the NOR bivalent. Bar = 2 pm. [From Marec and Traut, 1993b; reprinted with permission of Blackwell Scientific Publications, Oxford.]

transient double-strand Rasmussen, 1980).

break,

is responsible

for resolution

of interlockings

(Holm

and

Synaptic speczjicity. To ensure precise site-to-site pairing of homologous chromosomes in the SC, required for subsequent crossing-over, synapsis must be highly specific. Biochemical evidence from the analysis of DNA synthesis during meiotic prophase I in the plant Lilium has suggested that this specificity is mediated by the so-called zygotene DNA (zygDNA). Single copy sequences of this DNA, 5-10 kb in length and with a GC content of 50-60%, are distributed throughout chromosomes, comprising 0.3% of the genome. The zygDNA

220

F. Marec

remains unreplicated during the premeiotic S phase, its replication being delayed until zygotene (Hotta and Stern, 1971). The delayed replication of zygDNA might ensure a specific molecular pairing by complementary base pairing of corresponding zygDNA sequences of aligned homologous segments (Hotta and Stern, 1984; Hotta et al., 1984; see also Holm, 1985; and references therein). The first step in the assembly of the SC in zygotene is the alignment of homologous segments at a distance of 300 nm from each other. Then, the chromatin of one of the segments moves out of the central space between both LEs. Subsequently, precursor material for the central region binds to the exposed LE and is organized into a central region. Upon rotation of the chromatin of the homologous LE, binding of this to the central region completes the SC formation (Fig. 13; Holm, 1985). Based on the analysis of serial sections through B. mori meiocytes (Rasmussen, 1976; Holm and Rasmussen, 1980), the temporal sequence of SC formation has been established using additional cellular and nuclear markers for individual stages. At early zygotene, SC assembly starts in terminal or subterminal parts of the homologs, although the LEs are not yet continuous between the telomere regions. The LEs are nearly continuous at mid zygotene. Afterwards, the SC formation proceeds in all bivalents from telomere to telomere, if not impeded by interlockings, in a zipper-like fashion. At late zygotene, SC formation is completed in most regions of the homologs. All homologs appear synapsed along their entire length at early pachytene. The speed of synapsis is not uniform throughout the chromosome complement. Delayed synapsis of terminal segments was regularly observed in 2 bivalents in B. mori, one had a compact knob and the other carried the nucleolus organizing region (NOR). This suggests that the knob and NOR represent an impediment to synapsis (Rasmussen, 1986). In pachytene nuclei of E. kuehniella, thicker terminal modifications of LEs were observed in 2 bivalents carrying NORs (Fig. 12). Frequently, the SCs opened up in the region of the terminal modifications indicating that these modifications may obstruct perfect synapsis

a

b

C

d

Fig. 13. A schematic presentation of the assembly of the synaptonemal complex in zygotene. CE=central element; ch = chromatin; LE = lateral element; TF = transversal filament. (a) Homologous chromosome segments are parallelly aligned. (b) The chromatin of the right segment rotated out of the central space; formation of the central region starts at the exposed LE. (c) The right segment shows fully formed central region; the chromatin of the homologous segment rotated out of the central space. (d) The homologous LE binds to the central region, thus completing the SC formation. [Modified after Holm, 1985.1

Synaptonemal

Complexes

in Insects

221

(Marec and Traut, 1993b). A considerable delay in synapsis accompanied by incomplete pairing seems to be a typical feature of non-homologous sex chromosomes, W and Z, in females of moths E. kuehniella and Galleria mellonella (Marec and Traut, 1994; Wang et al., 1993).

Synaptic adjustment, non-homologous pairing and correction of pairing Chromosome pairing is a highly specific process as far as structurally normal homologs in diploid organisms are concerned. In heterozygous organisms, polyploids and hybrids, chromosome synapsis appears to take place in 2 phases. In zygotene, only homologous segments pair (homosynapsis). During pachytene, however, pairing may involve nonhomologous regions or entire non-homologous chromosomes (heterosynapsis). An inverted segment, for example, produces an inversion loop in zygotene. The loop is missing in pachytene, and a straight SC with the inverted segment non-homologously-paired in reversed order exists. Another expression of this phenomenon is found in the observation that differences in the length of chromosomes may be adjusted. Consequently, the LEs in a given heterozygous bivalent may attain the same length. The process is referred to as synaptic adjustment, the term coined by Moses and co-workers, who studied SCs in mammals (Moses, 1977; Moses and Poorman, 1981; Moses et al., 1982; for a review see John, 1990). In polyploid B. mori females, multivalent configurations that frequently occur at zygotene may be corrected during pachytene to produce bivalents and univalents (see Rasmussen, 1977b). Several cases of synaptic adjustment and non-homologous pairing have been described in E. kuehniella females (Figs 14 and 15; Weith and Traut, 1986). In bivalents heterozygous for a T(A; W) translocation (i.e. an autosome carries a translocated segment of the W sex chromosome), LEs differing considerably in lengths, were brought to the same length in pachytene. The adjusted length was a compromise between the mutant and the wild-type homolog length (Fig., 14~). A typical example of non-homologous pairing is the W-Z sexchromosome bivalent in E. kuehniella wild-type females. In this case, pachytene pairing extended from two-thirds to the full length of the shorter W chromosome, although the W and Z are largely, if not completely, non-homologous (Weith and Traut, 1980; Marec and Traut, 1994). A rather strange pairing configuration was observed in sex chromosome bivalents containing a deleted W chromosome, Df(W) (Weith and Traut, 1986). In about 30% of pachytene nuclei, the halves of the Z chromosome were synapsed to either side of the Df(W). Thus, in addition to non-homologous pairing, one half of the Z synapsed with the Df(W) in reversed order (Fig. 15e). When supernumerary W segments were introduced into the E. kuehniella genome via the T(A;W) translocation, pairing between the W chromosome proper and the translocated W segment has never been observed. This represents an example of non-pairing of homologous segments, demonstrating that homology itself is not always sufficient for pairing initiation. Theoretically, the lack of homologous pairing could also be explained by dissociation of corresponding pairing configurations during spreading procedure. However, the possibility is less probable in this case. The author’s conclusion is based on a large number of nuclei inspected and, in addition, configurations with non-homologously paired W segments were frequently found. The classical assumption that synaptic adjustment occurs in 2 phases is inconsistent with the results of a recent study carried out on zygotene and pachytene spermatocytes of the grasshopper, Ch. jacobsi. In animals heterozygous for a paracentric inversion, the inverted

222

F. Marec 14

A

2

TtA;WI

W

la/58 L

[bl _ Df(W1

A 17/M

[cl

Ttkwlfl W [d) A

2

TtA;Wl

W

23158

te1

L& 2 DfCWl

6129

Dftw1

9/29

2/29

2

lf[Wl=[

9/29

Figs. 14 and 15. A schematic presentation structural mutants of the W chromosome oocytes. In both figures, the right-hand

of the pachytene configurations of sex chromosomes and found in microspread preparations of Ephestia kuehniella column gives the observed frequencies of the respective configurations. Fig. 14. The T(A; W) translocation in the heterozygous condition. Z (thin line), W (thick line), structurally normal sex chromosomes; A = structurally normal autosome; T(A; W) = an autosome carrying a translocated segment of the W sex chromosome. (a) The autosomes are homologously paired, while the translocated W chromosome segment remains unpaired, W-Z bivalent shows partial pairing; (b) a quadrivalent is formed, in which the translocated W chromosome segment pairs with the free terminal segment of the Z chromosome; (c) both the A-T(A; W) and W-Z bivalents are completely paired by synaptic adjustment of the length of their axes. Fig. 15. The W chromosome deficiency Df(W). (a) Only short terminal segment of the Df(W)-Z bivalent is synapsed; (b) the Df(W) chromosome is fully synapsed with the Z chromosome but a large segment of the Z remains free; (c) the longer Z chromosome axis is twisted along the shorter Df(W) axis; (d) the Df(W) and the Z chromosome axes were brought to the same length by synaptic adjustment resulting in a completely paired bivalent; (e) the halves of the Z chromosome were non-homologously synapsed to either side of the Df(W). [From Weith and Traut, 1986; reprinted with permission of SpringerVerlag, Heidelberg.]

segments usually heterosynapsed at pachytene without displaying the first phase, i.e. previous homosynapsis at zygotene (Diez and Santos, 1993). In triploid B. mori oocytes (Rasmussen, 1977b), trivalent configurations did occur in the first phase of pairing at zygotene. In contrast, mid to late pachytene nuclei showed a maximum number of homologous bivalents and a number of non-homologous associations in the form of either foldback pairing of univalents or pairing of 2 chromosomes of unequal length. The unstable trivalent configurations entirely disappeared, because the zygotene homosynapsis was followed by correction of pairing in the second phase. Similarly, a relatively high frequency of quadrivalents was found in autotetraploid B. mori oocytes at zygotene (Rasmussen and Holm, 1979). This picture changed by mid to late pachytene, when nearly all chromosomes formed bivalents. Evidently, the quadrivalents represent unstable configuration in the system. These kinds of pairing correction have led to the hypothesis that SCs tend to optimize pairing in the form of bivalents (Rasmussen and Holm, 1982). However, this applies only to female meiosis of B. mori, which is achiasmatic (Rasmussen, 1977a). In autotetraploid B. mori spermatocytes, the number of zygotene quadrivalents was reduced during pachytene but about 50% of the quadrivalents persisted up to metaphase I (Rasmussen, 1987). The author concluded that the occurrence of crossing-over prevents the transformation of multivalents into bivalents in the second phase of pairing.

Synaptonemal Complexes in Insects

223

Pairing behavior of sex chromosomes In the homogametic sex, pairing of the sex chromosomes does not differ from that of autosomal bivalents. The 2 sex chromosomes, either XX or ZZ, form a complete SC. In the heterogametic sex, however, various pairing patterns may occur, depending on the sex-chromosome system and on the degree of homology between the heterologous sex chromosomes. This concerns many insects because of the great diversity of the karyotypes. Unfortunately, only a few ultrastructural studies on sex chromosome pairing in insects are available. Concluding from studies done in male meiosis of mammals (see John, 1990), we can theoretically expect at least 3 different situations in insects: (i) the X and Y or the Z and W are paired and form SCs of various lengths; (ii) pairing is missing between the 2 sex chromosomes; and (iii) species with the X0 (e.g. some Heteroptera males, grasshopper males) or ZO (e.g. Trichoptera females) system should display a sex chromosome univalent. The last situation was demonstrated in X0 males of the grasshopper, C/z. jacobsi, by Santos er al. (1993). In EM surface-spread preparations of pachytene spermatocytes, the X chromosome appeared as a single unpaired axis. Sex chromosome pairing has been thoroughly examined in E. kuehniella, which possess a WZ sex chromosome pair in the female sex. This is typical of Lepidoptera species. In the moth, the W chromosome can be identified in EM spreads by densely-staining heterochromatin tangles associated with the LE (Weith and Traut, 1980). The W and Z chromosomes, although largely non-homologous pair completely. The formation of the WZ bivalent is, however, delayed. In pachytene complements, 23% to 39% partially paired WZ bivalents were found (Weith and Traut, 1986; Marec and Traut, 1994). The latter authors suggested that W-Z pairing in E. kuehniella proceeds in 4 phases. First, the W and Z chromosomes become associated with their telomeric segments (Fig. 16a). The pairing starts at one end only, as reported also for the WZ bivalent of the wax moth, Galleria mellonella (Wang et al., 1993). In the second phase, pairing proceeds until a bivalent is produced in which the shorter axis of the W chromosome is almost fully paired, while the longer axis of the Z chromosome shows a free distal segment (Fig. 16b). In the third phase, contraction and multiple twisting of the Z chromosome along the axis of the W chromosome results in a completely paired WZ bivalent (Fig. 16~). In the last phase, both LEs are adjusted in length (see Weith and Traut, 1986). The delay in synapsis of the W and Z probably results form their lower affinity to one another compared with the specific pairing of autosomes. This conclusion has been supported by the fact that pairing efficiency of the E. kuehniella sex chromosomes was significantly improved in T(W; Z) females when a Z-chromosome segment was translocated onto the W chromosome (Marec and Traut, 1994). Analysis of structural mutants of sex chromosomes in E. kuehniella using the EM microspreading technique revealed that pairing configurations may also influence meiotic segregation. In the so-called ASF (“abnormal segregating females”) lines, non-Mendelian segregation occurs affecting a Z-linked phenotypic marker. ASF females possess in their genome an additional fragment of the Z chromosome (Z+) containing the dominant wildtype allele of the marker gene. This fragment was frequently synapsed with the homologous segment of the regular Z chromosome. Most often a WZZ+ trivalent was visible. It was suggested that this configuration could lead to the segregation of the Z+ fragment together with the W chromosome, thus simulating a W linkage. Concerning the remaining pairing configurations, free or autosynapsed Z+ fragments could segregate randomly, resulting in the occurrence of exceptional phenotypes in progeny (Marec and Traut, 1993a).

224

F. Marec

Fig. 16. Phases of sex chromosome pairing in microspread pachytene oocytes of Ephestia kuehniella. (a) Delayed terminal initiation of W-Z pairing. (b) Partial pairing of W-Z bivalent with lateral elements of unequal length. (c) A typical wild-type sex chromosome SC: the longer Z chromosome axis is twisted along the shorter W axis. In all figures, note that sex chromatin tangles (arrows) decorate the W axis. Bar = 1 pm. [From Marec and Tram, 1994; reprinted with permission of NRC Research Press, Ottawa.] Fig. 17. A microspread male bivalent of Ephestia kuehniella, showing 1 recombination nodule (arrow). Bar = 1 pm. [From Marec and Tram, 1993b; reprinted with permission of Blackwell Scientific Publications, Oxford.] Fig. 18. A detail of a distal segment of a microspread male bivalent in Ephestia kuehniella, showing a recombination nodule placed in the free space between the two LEs. Bar=0.5 pm. [From Marec and Traut, 1993b; reprinted with permission of Blackwell Scientific Publications, Oxford.] Fig. 19. EM micrograph of a cross-section through a metaphase I plate of a Bombyx mori oocyte, showing that the modified SCs have fused into a continuous sheet between the electron-dense chromatin of the homologs. Bar = 2 pm. [From Rasmussen and Holm, 1982; reprinted with permission of Plenum Press, New York.]

225

Synaptonemal Complexes in Insects CHIASMATIC Chiasmatic

VERSUS

ACHIASMATIC

MEIOSIS

meiosis

Most organisms exhibit a chiasmatic mode of meiosis. This means that SCs are, after their degradation during the pachytene-diplotene transition, replaced by chiasmata, which then hold bivalents together until metaphase I. While the SC provides the structural basis for recombination, the actual process of recombination followed by chiasma formation is, probably, mediated by recombination nodules (RNs). These small electron-dense spheres, about 50-100 nm wide, are associated with the central region of the SC. A number of indirect evidence, mostly obtained in insect models, suggests that RNs represent multienzyme sites playing a significant role in exchange between the DNA of non-sister chromatids (Carpenter, 1975b, 1994; Holm, 1985; John, 1990). RNs were first described in D. melanogaster females by Carpenter (1975b), who also proposed that they play a role in meiotic recombination on the basis of a good correlation between RNs and exchange events. A maximum number of 5 RNs per nucleus correlated well with the expected frequency of crossing-over (5.6 per nucleus). The distribution of RNs along the bivalents correlated with the site of crossing-over as well. In addition, the D. melanogaster SCs lacked RNs in the pericentromeric heterochromatin regions, where crossing-over does not occur. The number of RNs is reduced in recombination defective females of the meiand meimutations of D. melanogaster (Carpenter, 1979a). It has been shown using autoradiography that a small amount of DNA synthesis occurs in the vicinity of the RNs in D. melanogaster females after bulk DNA synthesis is over. The DNA synthesis presumably represents DNA repair accompanying recombination (Carpenter, 1981). In wild-type females of D. melanogaster, Carpenter (1979b) discovered 2 sorts of RNs. They differ in their shape and time of appearance and are now referred to as early (ellipsoidal) or late (spherical) nodules. While the late nodules correlate with crossing-over and their probable function is thus clearly defined, we still lack any evidence that the early nodules play any role in meiotic recombination at all. It is speculated that the early nodules, which are always more numerous than late nodules, could mediate gene conversion (Carpenter, 1994). Holm and Rasmussen (1980) studied the evolution of RNs in B. mori spermatocytes. RNs first appeared at early zygotene simultaneously with the initiation of SC formation. Their number gradually increased during zygotene and reached a maximum number of 9 1 per nucleus by late zygotene. This number was, however, reduced to about 60 RNs per 28 bivalents until early pachytene. At all stages, the RNs appeared associated with the LEs through filamentous material. Starting from mid pachytene, RNs were transformed into elements intimately associated with the chromatin. These were termed chromatin nodules (CNs). At the transition from pachytene to diplotene, most nodules resembled the larger CNs, which subsequently evolved into chiasmata. The number and distribution of chiasmata at diplotene-diakinesis corresponded to the number and distribution of RNs at pachytene. Consequently, it has been suggested that crossing-over takes place at pachytene, preferentially at mid pachytene. The authors also found that the distribution of RNs among and along bivalents is a result of an initially random distribution, which is then slightly modified in order to ensure a minimum of one chiasma per bivalent (see also Rasmussen and Holm, 1982; Holm, 1985). Compared with B. mori, much less RNs were found in pachytene nuclei of another moth

226

F. Marec

E. kuehniella, because the bivalents were shorter (Marec and Traut, 1993b). Any of the 30 bivalents was associated with only one RN. Higher numbers were rare. In contrast to the random distribution observed in B. mori (Holm and Rasmussen, 1980), RNs were preferentially located in distal positions in E. kuehniella (Figs. 17 and 18). According to Traut (1977) individual bivalents in E. kuehniella males contain only 1 chiasma, and many of the chiasmata are distally located. This corroborates again an interdependency between RNs and cross-over sites. RNs were missing in female meiosis but not in male meiosis of the wax moth, Galleria mellonella (Wang et al., 1993) B. mori (Rasmussen, 1976; Holm and Rasmussen, 1980) and E. kuehniella (Marec and Traut, 1993b). The Lepidoptera females lack crossing-over, the absence of RNs in females is thus in keeping with their role in meiotic recombination. Further indirect evidence in favour of a role of RNs in meiotic recombination comes from the comparison of 2 Orthoptera species. In Chloealtis conspersa, RNs are associated with the central region of the SCs exactly at the position of localized chiasma in the distal SC ends, but they are not localized in Locusta migratoria, which do not have localized chiasmata (Bernelot-Moens and Moens, 1986). D. melanogaster oocytes develop in sibling clusters of 16 interconnected cells, from which the 2 pro-oocytes proceed into the pachytene stage, but only 1 develops further into a mature oocyte. The remaining cells (pro-nurse cells) also enter meiosis and occasionally show incomplete SC segments, but then develop into nurse cells. Recently, so-called solitary RNs were found in pro-nurse cells of D. melanogaster ovaries, in addition to the SCassociated RNs (Schmekel et al., 1993~). Most RNs of the pro-nurse cells remained solitary. Their role is not known. Achiasmatic meiosis In a number of organisms, crossing-over and the subsequent chiasma formation do not take place in meiosis of the heterogametic sex. In spite of the fact that one sex has lost recombination, chromosome pairing, and segregation are still regular (White, 1973; Wolf, 1994a). In insects, achiasmatic meiosis is widespread in males of Diptera species. Achiasmatic meiosis is also common in females of 2 closely related orders, Trichoptera and Lepidoptera (Suomalainen, 1966, 1969; Traut, 1977, 1991). The achiasmatic association has been intensively studied in D. melanogaster males. In this system, SCs and chiasmata are missing. When the chromosomes first become visible in early meiotic prophase, the homologs are already paired along their entire length and remain so until their segregation in anaphase. The heterologous sex chromosomes pair at discrete, heterochromatic sites termed collochores. The collochores seem to consist, at least in part, of the NORs that mediate X-Y meiotic chromosome pairing. Each NOR contains some 200 copies of rDNA that provides homology between the otherwise largely nonhomologous X and Y chromosomes. The 240 bp intergenic spacer repeats associated with rRNA genes have been implicated, because of their strong pairing ability, as the essential DNA sequences for XY pairing. Thus X-Y pairing ability is probably based on underlying DNA homology. Recently, a general model for achiasmatic pairing in D. melanogaster males has been proposed. The model is based on the formation of non-recombination heteroduplex in sites of DNA homology by the combined action of a strand transferase and topoisomerase I (see McKee et al., 1992; and references therein). A recent result in males of Drosophila simulans (Ault and Rieder, 1994) supports the hypothesis that synapsis of XY bivalent is intimately associated with 240 bp non-transcribed spacer repeats. In this

Synaptonemal

Complexes

in Insects

221

species, however, the Y chromosome has few, if any, rRNA genes. The XY bivalent of D. simuluns males exhibits no material resembling collochores that have been proposed to hold X and Y chromosomes together in D. melanogaster males (McKee et al., 1992). In the Y chromosome of D. simuluns, the spacer repeats form a large block at the distal end of its long arm, and the X chromosome indeed pairs with this distal region of the Y chromosome. In addition, the synaptic regions of the XY and autosomal bivalents appear similar. Therefore, Ault and Rieder (1994) suggest, in contrast to McKee et al. (1992) that the conjunctive mechanism holding the XY and autosomal bivalents is identical. Achiasmatic meiosis in Lepidoptera females differs from that in D. melunogaster males in many respects. In Lepidoptera, oocytes develop in sibling clusters of 8 interconnected cells (Telfer, 197.5; Rasmussen, 1977a; Marec et al., 1993). All the cells enter meiosis and develop in parallel until the end of pachytene. In this stage, all cells display SCs but RNs are missing (Rasmussen, 1976; Wang et al., 1993; Marec and Traut, 1993b). It is impossible to distinguish the future oocyte from the nurse cells according to the pachytene SC complements (Marec and Traut, 1993b). Beyond pachytene, the oocyte and the 7 nurse cells develop differently. The oocyte nucleus completes meiosis. In the nurse cells, the SCs detach from the bivalents and form aggregates that disappear later. The nurse cell nuclei degenerate (Traut, 1977; Rasmussen, 1977a; Rasmussen and Holm, 1982). The further development of the oocyte nucleus was followed in B. mori (Rasmussen, 1977a). The SCs remain associated with the bivalents, and are retained in a modified form until metaphase I. After pachytene, LEs grow and form a solid rod between the homologous chromosomes. Subsequently, the chromatin condenses, all bivalents align in the metaphase plate, and the modified SCs fuse into a continuous plate (Fig. 19). Finally, the homologs dissociate from the modified SCs, which persist in the metaphase plate, while homologous chromosomes segregate to opposite poles in anaphase I (see results of Wolf, 1993, 1994b, in E. kuehniellu oocytes). That is why several previous authors referred to this material as “elimination chromatin” (e.g. Vereiskaya, 1975). Rasmussen (1977a) suggested that the preservation of SCs until metaphase I has evolved as a mechanism to substitute for the lack of crossing-over and chiasma formation in order to ensure regular disjunction of homologous chromosomes.

SYNAPTONEMAL

COMPLEX

KARYOTYPING

In some insect orders, mitotic chromosome preparations are inadequate for karyotyping, because the chromosomes are small and fail to show distinct morphological characteristics. In such cases, the pachytene bivalents can be exploited for chromosome mapping by a light microscopical analysis of the chromomere pattern or by analyzing the SCs at the electron microscopical level. This approach has been successfully used in Lepidoptera, insects where numerous small chromosomes are the rule. The chromosomes do not possess a primary constriction that could serve as a morphological marker (Suomalainen, 1969). Pachytene mapping using light microscopy was, for example, applied in B. mori females. Six of 28 bivalents were identified according to their distinct chromomere pattern, among them 1 bivalent containing the NOR (Traut, 1976). Similarly, the light microscopic analysis of 30 pachytene bivalents in E. kuehniellu females enabled Schulz and Traut (1979) to recognize 4 bivalents according to conspicuous structural landmarks. One was the WZ bivalent characterized by the heterochromatic W chromosome. Two other bivalents carried the NORs, and the fourth had a distal segment nearly devoid of chromomeres. At the EM level, the chromomere pattern is lost. To differentiate individual chromosomes,

228

F. Marec

the following characteristics can be used: (i) relative sizes of bivalents within a pachytene complement; (ii) morphological landmarks such as heterochromatin tangles, NORs and electron-dense knobs; (iii) position of the centromere in species with monocentric chromosomes; and (iv) chromosomal rearrangements in mutant strains. In Lepidoptera, the wildtype SC complements of 4 species have been analyzed. These were B. mori (Rasmussen, 1976; Holm and Rasmussen, 1980) E. kuehniella (Weith and Traut, 1980; Marec and Traut, 1993b), G. mellonella (Wang et al., 1993) and the pink bollworm, Pectinophora gossypiella (Bartlett and Del Fosse, 1991). In all species, relative sizes of SCs within a complement formed a nearly continuous range. Therefore, individual bivalents could not be unambiguously identified by their lengths. Pachytene oocytes and spermatocytes showed 1 or 2 SCs carrying the NOR and 1 SC with a knob, only in G. mellonella a specific autosomal SC could not be recognized. In E. kuehniella and G. mellonella females, the WZ bivalent was regularly identified. However, the authors failed to find a heteromorphic bivalent in the female karyotype of the 2 remaining species. It seems clear that, at least in insects with “holokinetic” organization of chromosomes (i.e. with relatively large kinetochores and without a primary constriction), SC karyotyping at the EM is a limited option. The EM analysis of spread SC complements has, however, been shown to be particularly useful for mapping various gamma-ray induced chromosome aberrations in E. kuehniella. With this approach, it was not only possible to characterize morphologically individual translocations, whole chromosome fusions, larger deficiencies, and chromosome fragments, but also to test properties of the pairing process, such as synaptic adjustment and nonhomologous pairing (see Section 3.3; Weith and Traut, 1986; Traut et al., 1986; Marec and Traut, 1994) and to analyze impact of their synaptic pairing configurations on chromosome segregation (see section 3.4; Marec and Traut, 1993a). SC karyotyping appears much easier in species with monokinetic chromosomes as demonstrated by Santos et al. (1993) in a grasshopper, Ch. jacobsi. Surface-spread spermatocyte nuclei of the grasshopper displayed 8 autosomal SCs (3 long metacentric, 4 medium acrocentric and 1 short acrocentric) and an unpaired axis of the X chromosome. At pachytene, all autosomal bivalents could be identified with certainty on the basis of SC arm index and relative lengths. In conclusion, the SC karyotyping seems to be a prospective approach in insects with small chromosomes as long as mutants are available. The rapid advances achieved in molecular cytogenetics and the use of chromosome specific probes render the analysis of wild-type karyotypes feasible.

CONCLUDING

REMARKS

Insect models have frequently been used for the study of SCs because of several advantages in comparison with other groups of animals. Many species can be easily handled and cheaply reared in the laboratory. Some insect models develop fast, and can produce several generations per year. For instance, D. melanogaster requires only lo-12 days for each generation when kept at 25°C. Furthermore, insects are available in great numbers. Another advantage is the large diversity among insects. This applies to chromosome numbers and kinetic organization of chromosomes. We can, for example, use insects with small or high numbers of bivalents, such as D. melanogaster (Diptera; n = 4) or E. kuehniella (Lepidoptera; n = 30). We find species having either monocentric chromosomes (e.g. Orthoptera, Diptera, Coleoptera) or chromosomes with relatively large kinetochores, the so-called holokinetic

Synaptonemal

Complexes

in Insects

229

chromosomes (e.g. Hemiptera, Trichoptera, Lepidoptera). Among insects, we can find various modifications of the sex-chromosome determination including male (XYjXX or X0/Xx) and female heterogamety (ZWjZZ or ZOjZZ), and a number of derived systems (see Blackman, 1995). Different modes of meiosis, such as chiasmatic and achiasmatic, which may or may not involve the SC formation, enable us to study the various roles of SCs in recombination and chromosome disjunction. The relatively low DNA content in most insect species is advantageous when the organization of chromatin loops associated with the SC is studied. In the past 20 years, enough basic information about the SC and related aspects has been obtained using insect models. The architecture of early meiotic prophase I nuclei including the formation of SCs, their interlocking, and the relation between SC, recombination and chiasma formation, have been characterized in detail using the silkworm, B. mori (for a review see Rasmussen and Holm, 1982). In B. mori females, modified synaptonemal complexes have been described as specific structures that substitute chiasmata in the achiasmatic mode of meiosis (Rasmussen, 1977a). Easy identification of the WZ bivalent owing to the sex-specific heterochromatin in the Mediterranean flour moth, E. kuehniella, has made it possible to study non-homologous pairing of sex chromosomes and their synaptic adjustment (Weith and Traut, 1980, 1986; Marec and Traut, 1994). Various sex-chromosome mutants of this moth were useful in the analysis of pairing specificity and pairing abnormalities (Traut et al., 1986; Weith and Traut, 1986; Marec and Traut, 1993a). Based on the correlation between recombination nodules, associated with SCs, and exchange events in D. melanogaster, Carpenter (1975b) first suggested a role of these nodules in meiotic recombination. Various meiotic mutants of D. melanogaster represent excellent models for studying genetic and molecular control of meiotic events (for a review see John, 1990). Grasshoppers and crickets (Orthoptera) are favourable objects of cytogenetic studies due to their large chromosomes, which render chiasmata particularly conspicuous. In addition, the B chromosomes occur frequently. Orthoptera chromosomes have been particularly useful to examine the relation between recombination and chiasma formation (see John, 1990). Recently, based on findings in a grasshopper, a model has been proposed for the role of chromatid cores in the formation of SCs (Rufas et al., 1992). Another interesting phenomenon in Orthoptera is the persistence of the so-called synaptonemal polycomplexes beyond meiotic prophase I, which seems to be characteristic for this order (Wolf and Mesa, 1993). Finally, the architecture of the central region of the SC has recently been discovered in B. cribrosa. It was possible because this beetle has an exceptionally distinct ladder-like and multi-layered central element and highly ordered transversal filaments (Schmekel et al., 1993a). The proposed model of the central region also applies to D. melanogaster. It has also been found to be applicable for the rat, although the central element in this mammalian system is superficially quite different (Schmekel et al., 1993b; Schmekel and Daneholt, 1995). Insect models have a few drawbacks. With the exception of D. melanogaster, formal genetics is poorly developed in most species. Compared to vertebrates, insect gonads are relatively small and it may be difficult to collect enough material for immunocytochemical studies or for construction of cDNA expression libraries. In this respect, mammalian systems are, for example, more convenient for analysis of SC proteins or for localization of specific DNA sequences on SCs (e.g. Dobson et al., 1994; Heng et al., 1994). On the other hand, the SC is morphologically highly conserved and also basic features of SC

230

F. Marec

formation in insects are very similar to that of vertebrates. Thus, many data on SC biology obtained using insect models apply to other eukaryotes and vice versa. Acknowledgemenrs-The

technical assistance of MS Ivana Kollarova is gratefully acknowledged. Preparation of this paper was supported by grant no. 607105 of the Czech Academy of Sciences (Prague) and by the research contract 7161/RB of the International Atomic Energy Agency (Vienna, Austria). This review could not have been compiled without the long-term support of the Alexander von Humboldt Foundation (Bonn, Germany).

REFERENCES Ault, J. G. and C. L. Rieder. 1994. Meiosis in Drosophila males I. The question of separate conjunctive mechanisms for the XY and autosomal bivalents. Chromosoma 103: 352-56. Barlow, A. L., G. Jenkins and I. ap Gwynn. 1993. Scanning electron microscopy of synaptonemal complexes. Chromosome Res. 1: 9913. Bartlett, A. C. and F. E. Del Fosse. 1991. The pachytene karyotype of the pink bollworm (Lepidoptera: Gelechiidae). Southwestern Entomol. 16: 223-35. Bemelot-Moens, C. and P. B. Moens. 1986. Recombination nodules and chiasma localization in two Orthoptera. Chromosoma

93: 22&26.

In G. A. Kerkut and L. I. Gilbert (eds.) Comprehensive Insect Vol. 10. Pergamon Press, Oxford. Blackman, R. L. 1995. Sex determination in insects, pp. 57794. In S. R. Leather and J. Hardie (eds.) Insect Reproduction. CRC Press, B&a Raton, FL. Carpenter, A. T. C. 1975a. Electron microscopy of meiosis in Drosophila melanogasfer females. I. Structure, arrangement, and temporal change of the synaptonemal complex in wild-type. Chromosoma 51: 157-82. Carpenter, A. T. C. 1975b. Electron microscopy of meiosis in Drosophila melunogaster females: II: The recombination nodule-a recombination-associated structure at pachytene? Proc. Nut1 Acad. Sci. U.S.A. 72: Berry, S. J. 1985. Insect nucleic acids, pp. 219-53. Physiology,

Biochemistry

and Pharmacology,

3186-9.

Carpenter, A. T. C. 1979a. Recombination nodules and synaptonemal complex in recombination defective females of Drosophila melanogaster. Chromosoma 75: 259-92. Carpenter, A. T. C. 1979b. Synaptonemal complex and recombination nodules in wild-type Drosophila melanogaster females. Genetics 92: 5 1141. Carpenter, A. T. C. 1981. EM autoradiographic evidence that DNA synthesis occurs at recombination nodules during meiosis in Drosophila melunogaster females. Chromosoma 83: 59-80. Carpenter, A. T. C. 1994. The recombination nodule story-seeing what you are looking at. BioEssays 16: 69-74. de1 Cerro, A. L., A. Fernandez and J. L. Santos. 1996. Chiasma localization, heterochromatin and synaptonemal complexes in the grasshopper Pyrgomorpha conica. Chromosome Res. 4: 69-76. Chevaillier, P. 1974. Recherches sur le complexe synaptonematique I. Mize en evidence des constituants nucleoproteiques du complexe synaptonematique par methode de Bernhard. J. Microsc. 19: 147754. Comings, D. E. and T. A. Okada. 1976. Nuclear proteins, III. The fibrillar nature of the nuclear matrix. Exp. Cell Res. 103: 341-60.

S. J. and G. F. Meyer. 1973. Differentiation of the synaptonemal complex and the kinetochore in Locusta spermatocytes studied by whole mount electron microscopy. Chromosoma 44: 231-53. Dallai, R. and B. A. Afzelius. 1995. Sperm structure OfTrichoptera. II. The aflagellated spermatozoa of Hydropfila, Orthotrichia and Stacfobia (Hydroptilidae). Int. .I. Insect Morphol. Embryol. 24: 161-70. Diez, M. and J. L. Santos. 1993. Synapsis in a paracentric inversion heterozygote of Chorthippus jacobsi (grasshopper). Heredity 70: 231-36. Dietrich, A. J., J. van Marle, C. Heyting and A. C. G. Vink. 1992. Ultrastructural evidence for a triple structure of the lateral element of the synaptonemal complex. J. Struct. Biol. 109: 196200. Dobson, M. J., R. E. Pearlman, A. Karaiskakis, B. Spyropoulos and P. B. Moens. 1994. Synaptonemal complex proteins: occurrence, epitope mapping and chromosome disjunction. J. Cell Sci. 107: 2749-60. Dresser, M. E. 1987. The synaptonemal complex and meiosis: An immunocytochemical approach, pp. 245-74. In P. B. Moens (ed.) Meiosis. Academic Press, New York. Esponda, P. and D. P. Krimer. 1979. Development of the synaptonemal complex and polycomplex formation in three species of grasshoppers. Chromosoma 73: 23745. Fawcett, D. W. 1956. The fine structure of chromosomes in the meiotic prophase of vertebrate spermatocytes. J. Counce,

Biophys.

Biochem.

Cytol. 2: 403-6.

Fiil, A. and P. B. Moens. 1973. The development, structure and function of modified synaptonemal complexes in mosquito oocytes. Chromosoma 41: 37-62. Fuge, H. 1979. Synapsis, desynapsis, and formation of polycomplex-like aggregates in male meiosis of Pales ferruginea (Diptera, Tipulidae). Chromosoma 70: 353-73.

Synaptonemal Goldstein,

P. 1987. Multiple

synaptonemal

complexes

Complexes

in Insects

(polycomplexes):

Origin,

231 structure

and function.

Cell Biol.

Int. 11: 759-96.

Heng,

H. H. Q., L. C. Tsui and P. B. Moens. 1994. Organization of heterologous DNA inserts on the mouse meiotic chromosome core. Chromosoma 103: 401-7. Heyer, W. D. 1994. The search for the right partner: Homologous pairing and DNA strand exchange proteins in eukaryotes. Experientia 50: 223-33. Heyting, C., A. J. J. Dietrich, P. B. Moens, R. J. Dettmers, H. H. Offenberg, E. J. W. Redeker and A. C. G. Vink. 1989. Synaptonemal complex proteins. Genome 31: 81-7. Heyting, C., P. B. Moens, W. van Raamsdonk, A. J. J. Dietrich, A. C. G. Vink and E. J. W. Redeker. 1987. Identification of two major components of the lateral elements of synaptonemal complexes of the rat. Eur. 1. Cell Biol. 43: 148854. Holm, P. B. 1985. Ultrastructural characterization of meiosis. Biol. Skif. (Copenhagen) 25: 39-90. Holm, P. B. and S. W. Rasmussen. 1980. Chromosome pairing, recombination nodules and chiasma formation in diploid Bombyx males. Curlsberg Res. Commun. 45: 483-548. Hotta, Y. and H. Stern. 1971. Analysis of DNA synthesis during meiotic prophase in Lilium. J. Mol. Biol. 55: 337-55.

Hotta,

Y. and H. Stern. Chromosoma

Hotta, John, Jones, King, King, Koch,

1984. The organization

of DNA segments

undergoing

repair synthesis

during

pachytene.

89: 127-37.

Y., S. Tabata and H. Stern. 1984. Replication and nicking of zygotene DNA sequences. Control by a meiosis-specific protein. Chromosoma 90: 243-53. B. 1990. Meiosis. Cambridge University Press, Cambridge. G. H. and J. A. Croft. 1986. Surface spreading of synaptonemal complexes in locusts II. Zygotene pairing observations. Chromosoma 93: 489-95. R. C. 1970. The meiotic behaviour of the Drosophila oocyte. Inf. Rev. Cytol. 28: 125-68. R. C. and H. Akai. 1971. Spermatogenesis in Bombyx mori. II. The ultrastructure of synapsed bivalents. J. Morphol. 134: 18 l-94. E. A., P. A. Smith and R. C. King. 1967. The division and differentiation of Drosophila cystocytes. J. Morphol.

121: 55-70.

Loidl, J. 1990. The initiation of meiotic chromosome pairing: the cytological view. Genome 33: 759-78. Loidl, J. 1994. Cytological aspects of meiotic recombination. Experientia 50: 285-94. Loidl, J. and H. Langer. 1993. Evaluation of models of homologue search with respect to their efficiency on meiotic pairing. Heredity 71: 342-5 1. Marec, F. 1995. Chromatin organization and the length of synaptonemal complex complements in relation to genome size. Chromosome Res. 3 (Suppl. 1): 117. Marec, F., J. Leutelt, W. Traut and K. W. Wolf. 1993. Visualization ofpolyfusomes in gonads of a moth, Ephestia kuehniella Z. (Lepidoptera: Pyralidae), by a microspreading technique and electron microscopy. Int. J. Insect Morphol.

Marec,

Marec, Marec, McKee,

Miller,

Moens, Moens, Moens, Moens, Moens, Moses, Moses,

Embryol.

22: 481-96.

F. and W. Traut. 1993a. Analysis of structural rearrangements of Lepidoptera chromosomes using the centrifugation spreading technique, pp. 243-50. In P. Howard-Kitto, R. F. Kelleher and G. V. Ramesh (eds.) Management qf Insect Pests: Nuclear and Related Molecular and Genetic Techniques. IAEA, Vienna. F. and W. Traut. 199313. Synaptonemal complexes in female and male meiotic prophase of Ephestia kuehniella (Lepidoptera). Heredity 71: 394404. F. and W. Tram. 1994. Sex chromosome pairing and sex chromatin bodies in W-2 translocation strains of Ephestia kuehniella (Lepidoptera). Genome 37: 426-35. B. D., L. Habera and J. A. Vrana. 1992. Evidence that intergenic spacer repeats of Drosophila melanogaster rRNA genes function as X-Y pairing sites in male meiosis, and a general model for achiasmatic pairing. Genetics 132: 52944. 0. L. Jr and A. H. Bakken. 1972. Morphological studies of transcription, pp. 155-77. In E. Diczfalusy (ed.) Karolinska Symp. Res. Meth. Reprod. Endocrinology. 5th Symp., Acta Endocrinol., Suppl. 168, Copenhagen. P. B. 1969. The fine structure of meiotic chromosome polarization and pairing in Locusta migratoria spermatocytes. Chromosoma 28: l-25. P. B. 1994. Molecular perspectives of chromosome pairing at meiosis. BioEssays 16: 101-6. P. B. and K. Church. 1979. The distribution of synaptonemal complex material in metaphase I bivalents of Locusta and Chloealtis (Orthoptera: Acrididae). Chromosoma 73: 247-54. P. B. and R. E. Pearlman. 1988. Chromatin organization at meiosis. BioEssays 9: 151-3. P. B., C. Heyting, A. J. J. Dietrich, W. van Raamsdonk and Q. Chen. 1987. Synaptonemal complex antigen location and conservation. J. Cell Biol. 105:933103. M. J. 1956. Chromosomal structures in crayfish spermatocytes. J. Biophys. Biochem. Cytol. 2: 215-18. M. J. 1977. Microspreading and the synaptonemal complex in cytogenetic studies. Chromosomes Today 6: 71-82.

Moses, M. J. and P. A. Poorman. 1981. Synaptonemal complex analysis of mouse chromosomal II. Synaptic adjustment in a tandem duplication. Chromosoma 81: 519-35.

rearrangements

232 Moses,

F. Marec

M. J., P. A. Karatsis and A. E. Hamilton. 1979. Synaptonemal complex analysis of heteromorphic trivalents in Lemur hybrids. Chromosoma 70: 141-60. Moses, M. J., P. A. Poorman, T. H. Roderick and M. T. Davisson. 1982. Synaptonemal complex analysis of mouse chromosomal rearrangements IV. Synapsis and synaptic adjustment in two paracentric inversions. Chromosoma 84: 45774. Putman, C. A. J., A. J. J. Dietrich, B. G. de Grooth, J. van Marle, C. Heyting, N. F. van Hulst and J. Greve. 1993. An atomic force microscopical study of the synaptonemal complex. Micron 24: 273-77. Rasmussen, S. W. 1975. Synaptonemal polycomplexes in Drosophila melanogaster. Chromosoma 49: 321-31. Rasmussen, S. W. 1976. The meiotic prophase in Bombyx mori females analyzed by three-dimensional reconstructions of synaptonemal complex. Chromosoma 54: 245593. Rasmussen, S. W. 1977a. The transformation of the synaptonemal complex into the “elimination chromatin” in Bombyx mori oocytes. Chromosoma 60: 205-2 1. Rasmussen, S. W. 1977b. Chromosome pairing in triploid females of Bombyx mori analyzed by three-dimensional reconstructions of synaptonemal complexes. Carlsberg Res. Commun. 42: 163-97. Rasmussen, S. W. 1986. Initiation of synapsis and interlocking of chromosomes during zygotene in Bombyx spermatocytes. Curlsberg Res. Commun. 51: 401-32. Rasmussen, S. W. 1987. Chromosome pairing in autotetraploid Bombyx males. Inhibition of multivalent correction by crossing over. Carlsberg Res. Commun. 52: 21 l-42. Rasmussen, S. W. and P. B. Helm. 1979. Chromosome pairing in autotetraploid Bombyx females. Mechanism for exclusive bivalent formation, Curlsberg Res. Commun. 44: 10 l-25. Rasmussen, S. W. and P. B. Holm. 1982. The meiotic prophase in Bombyx mori, pp. 61-85. In R. C. King and H. Akai (eds.) Insect Ultrastructures, Vol. 1. Plenum Publ. Corp., New York. Rattner, J. B., M. Goldsmith and B. A. Hamkalo. 1980. Chromatin organization during meiotic prophase of Bombyx mori. Chromosoma 79: 2 15-24. Rattner, J. B., M. R. Goldsmith and B. A. Hamkalo. 1981. Chromosome organization during male meiosis in Bombyx mori. Chromosoma 82: 341-5 1. Raveh, D. and A. Ben-Ze’ev. 1984. The synaptonemal complex as part of the nuclear matrix of the flour moth, Ephestia kuehniella. Exp. Cell. Res. 153: 99-108. Rufas, J. S., J. L. Santos, M. Diez and J. A. Suja. 1992. Meiotic chromosome structure: relationship between the synaptonemal complex and the chromatid cores. Genome 35: 105461. Santos, J. L., A. L. de1 Cerro and M. Diez, M. 1993. Spreading synaptonemal complexes from the grasshopper Chorthippus jacobsi: pachytene and zygotene observations. Hereditas 118: 23541. Scherthan, H., J. Bahler. and J. Kohli. 1994. Dynamics of chromosome organization and pairing during meiotic prophase in fission yeast. J. Cell Biol. 127: 273-85. Schmekel, K. and B. Daneholt. 1995. The central region of the synaptonemal complex revealed in three dimensions. Trends Cell Biol. 5: 239-42. Schmekel, K., U. Skoglund and B. Daneholt. 1993b. The three-dimensional structure of the central region in a synaptonemal complex: a comparison between rat and two insect species, Drosophila melanogaster and Blaps cribrosa. Chromosoma 102: 682-92. Schmekel, K., J. Wahrman and B. Daneholt. 1993~. Solitary and synaptonemal complex-associated recombination nodules in pro-nurse cells during oogenesis in Drosophila melanogaster. Chromosoma 102:396402. Schmekel, K., J. Wahrman, U. Skoglund and B. Daneholt. 1993a. The central region ofthe synaptonemal complex in Blaps cribrosa studied by electron microscope tomography. Chromosoma 102: 669-8 1. Schulz, H. J. and W. Traut. 1979. The pachytene complement of the wildtype and a chromosome mutant strain of the flour moth, Ephestia kuehniella (Lepidoptera). Genetica 50: 61-6. Suja, J. A. and J. S. Rufas. 1994. The telochore: a telomeric differentiation of the chromosome axis. Chromosome Res. 2: 361-8. Suja, J. A., J. de la Torre, J. F. Gimenez-Abian, C. Garcia de la Vega and J. S. Rufas. 1991. Meiotic chromosome structure. Kinetochores and chromatid cores in standard and B chromosomes of Arcypterafuca (Orthoptera) revealed by silver staining. Genome 34: 19-27. Sumner, A. T. 1983. New approaches to the study of chromosome organization. Sci. Prog. Oxford 68: 543-64. Suomalainen, E. 1966. Achiasmatische Oogenese bei Trichopteren. Chromosoma 18: 201-7. Suomalainen, E. 1969. Chromosome evolution in the Lepidoptera. Chromosomes Today 2: 132-8. Sym, M., J. A. Engebrecht and G. S. Roeder. 1993. ZIP1 is a synaptonemal complex protein required for meiotic chromosome synapsis. Cell 72: 365578. Telfer, W. H. 1975. Development and physiology of the oocyte-nurse cell syncytium. Adv. Insect Physiol. 11:223319. Traut, W. 1976. Pachytene mapping in the female silkworm, Bombyx mori L. (Lepidoptera). Chromosoma 58: 275-84. Traut, W. 1977. A study of recombination, formation of chiasmata and synaptonemal complexes in female and male meiosis of Ephestia kuehniella (Lepidoptera). Geneiica 47: 13542. Traut, W. 1991, Chromosomen. Klassiche und molekulare Cytogenetik. Springer-Verlag, Berlin.

Synaptonemal

Complexes

in Insects

233

W., A. Weith. and G. Tram. 1986. Structural mutants of the W chromosome in Ephestiu (Insecta Lepidoptera). Genetica 70: 69-79. Vereiskaya, V. N. 1975. A cytochemical study of the elimination chromatin in the silkworm (Bombyx mori L.) meiosis. Tsitologiya (in Russian) 17: 603-6. Verma, R. S. 1990. The Genome. VCH Publ., New York. Wahrman, J. 1981. Synaptonemal complexes-origin and fate. Chromosomes Today 7: 105-13. Wang, Y. X., F. Marec and W. Traut. 1993. The synaptonemal complex complement of the wax moth, Galleria meilonella. Hereditas 118: 113-19. Weith, A. and W. Traut. 1980. Synaptonemal complexes with associated chromatin in a moth, Ephestia kuehniella Z. The fine structure of the W chromosomal heterochromatin. Chromosoma 78: 275-91. Weith, A. and W. Traut. 1986. Synaptic adjustment, non-homologous pairing, and non-pairing of homologous segments in sex chromosome mutants of Ephestia kuehniellu (Insecta Lepidoptera). Chromosoma 94: 12s 31. Westergaard, M. and D. von Wettstein. 1970. Studies on the mechanism of crossing over IV. The molecular organization of the synaptinemal complex in Neottiella (Cooke) Saccardo (Ascomycetes). C.R. Trav. Lab. Carlsberg 37: 239-68. von Wettstein, D., S. W. Rasmussen and P. B. Holm. 1984. The synaptonemal complex in genetic segregation. Annu. Rev. Genet. 18: 331413. White, M. J. D. 1973. Animal Cytology and Evolution, 3rd edn. Cambridge University Press, Cambridge. Wolf, K. W. 1993. Centripetal movement of homologs occurs at the onset of anaphase A in primary oocytes of Ephesria kuehniella Z. (Pyralidae, Lepidoptera). Cell Motil. Cytoskel. 24: 2004. Wolf, K. W. 1994a. How meiotic cells deal with non-exchange chromosomes. BioEssays 16: 107-14. Wolf, K. W. 199413. The spindle apparatus in early embryonic divisions of Ephestiu kuehniella Z. (Pyralidae, Lepidoptera) is formed by alignment of minispindles. Zygote 2: 87-95. Wolf, K. W. and A. Mesa. 1993. Synaptonemal polycomplexes in spermatids: a characteristic trait of Orthoptera? Chromosome Res. 1: 181-8. Wolf, K. W., R. L. Blackman and A. T. Sumner. 1994. Scanning electron microscopy of insect chromosomes: Schistocercagregaria (Acrididae, Orthoptera), Megaselia scalaris (Phoridae, Diptera), and Myzuspersicae (Aphididae Hemiptera). J. Submicrosc. Cylol. Pathol. 26: 79-89.

Tram,