Df31 and chromatin structure in D. melanogaster - Journal of Cell ...

RESEARCH ARTICLE

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Df31 is a novel nuclear protein involved in chromatin structure in Drosophila melanogaster Gilles Crevel, Hella Huikeshoven and Sue Cotterill* Dept Biochemistry and Immunology, St Georges Hospital Medical School, Cranmer Terrace, London SW17 ORE, UK *Author for correspondence (e-mail: [email protected])

Accepted 26 October; published on WWW 11 December 2000 Journal of Cell Science 114, 37-47 © The Company of Biologists Ltd

SUMMARY We originally isolated the Df31 protein from Drosophila embryo extracts as a factor which could decondense Xenopus sperm, by removing the sperm specific proteins and interacting with histones to facilitate their loading onto DNA. We now believe that this protein has a more general function in cellular DNA metabolism. The Df31 gene encodes a very hydrophilic protein with a predicted molecular mass of 18.5 kDa. Immunostaining showed that Df31 was present in a wide range of cell types throughout differentiation and in both dividing and nondividing cells. In all cases the protein is present in large amounts, comparable with the level of nucleosomes. Injection of antisense oligonucleotides to lower the level of Df31 in embryos caused severe disruption of the nuclear structure. Large irregular clumps of DNA were formed, and in most cases the amount of DNA associated with each

clump was more than that found in a normal nucleus. Immunofluorescence, cell fractionation, and formaldehyde cross-linking show that Df31 is associated with chromatin and that a significant fraction of it binds very tightly. It also shows the same binding characteristics when loaded onto chromatin in vitro. Chromatin fractionation shows that Df31 is tightly associated with nucleosomes, preferentially with oligonucleosomes. Despite this no differences were observed in the properties of nucleosomes loaded in the in vitro system in the presence and absence of Df31. These results suggest that Df31 has a role in chromosomal structure, most likely acting as a structural protein at levels of folding higher than that of nucleosomes.

INTRODUCTION

classes each with differing specificities of DNA binding. The class 1 and 2 hmg group proteins are postulated to facilitate histone loading. Other HMG classes, such as 14/17, have been shown to alter chromatin structure directly. These were first observed as abundant chromatin associated proteins (one per two nucleosomes) copurifying through many chromatographic steps with histones. Analysis of their functions suggests that when incorporated in chromatin they produce a more open structure (Paranjape et al., 1995; Trieschmann et al., 1995). However, their exact cellular function is still unclear, especially as deletion of both HMG 14 and 17 from a chicken B cell line has no apparent effect (Li et al., 1997). How the DNA is folded into higher levels of structure is much less well understood. Physically this has been postulated to involve further helical wrappings of the nucleosome structures, and looping of the chromatin, from a number of fixed points to divide each chromosome into a number of domains (Gerdes et al., 1994; Woodcock, 1994; Kimura and Hirano, 1997). Several types of proteins have been identified which are thought to play a role in the higher levels of structure, including DNA topoisomerase II (Adachi et al., 1989; Cardenas and Gasser, 1993; Gimenez-Abian et al., 1995; Hirano and Mitchison, 1993; Swedlow et al., 1993) and the SMC proteins. SMC proteins have been isolated from a wide range of eukaryotes, e.g. S. cerevisiae – smc1 and 2 (Strunnikov et al., 1995), S. pombe – cut3 and 14 (Sutani and Yanagida, 1997), Xenopus – xcap e and c (Hirano and Mitchison, 1994), and

In an interphase eukaryotic chromosome extensive coiling and folding of the DNA reduces its length at least a million times over the possible extended length. Further compaction, twoto fivefold depending on species, also occurs due to the process of condensation prior to mitosis. The formation and maintenance of these condensed structures is vital to the correct functioning of DNA metabolism, and loss of compaction can have catastrophic effects such as inappropriate gene expression, impaired DNA replication, incomplete separation of DNA strands and strand breakage. Formation and maintenance of this remarkable compaction requires the combined action of an array of different proteins – both structural, and catalytic. Of the structural proteins by far the best characterised are the histones, which are responsible for folding the DNA into the primary structural unit, the nucleosome. The properties of the DNA are very dependent on presence or absence of histones and also on their state of modification, and recently it has become clear that a large body of loading unloading and modifying proteins are in place, to help maintain the structure and provide the structural flexibility for processes such as transcription and replication (Hartzog and Winston, 1997; Tsukiyama and Wu, 1997). Another set of conserved proteins which might play a structural role in chromatin are the HMG proteins (Baxevanis and Landsman, 1995; Grosschedl et al., 1994). These are small acid soluble proteins which have been divided into several

Key words: Df31, Chromatin structure, Drosophila

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human (Jessberger et al., 1996). The SMC group has been subdivided into two groups based on their homology to S. cerevisiae smc1 and smc2, which show physical interactions with each other, but do not cross complement. Many of the SMC group proteins were originally isolated through their effects on condensation. Like topoisomerase II they seem to be involved in the formation of the condensed structure, and in fact may act with the topoisomerase in some cases (Saitoh et al., 1995). There is also good evidence that the SMC proteins are involved in the maintenance of the structure, once it has been formed (Strunnikov et al., 1995). The SMC proteins most likely function as part of a complex and recently several other proteins have been identified which interact with them and also seem to be involved in condensation. These include the S. cerevisiae proteins Scc1 (mcd1) and Scc2 (Michaelis et al., 1997; Guacci et al., 1997), the Drosophila protein Barren (Bhat et al., 1996), and its Xenopus homologue xcaph (Hirano et al., 1997; Kimura and Hirano, 1997). Barren has also been shown to interact with topoisomerase II. In addition to these general effect factors there are also a whole range of proteins that have more localised influences on DNA structure. These include the polycomb group (Franke et al., 1992) of 10-15 proteins, which form a large complex needed to maintain repression of specific genes. The action of the polycomb proteins are counterbalanced by proteins of the trithorax group (Gould, 1997; Gerasimova and Corces, 1998). Finally proteins have been characterised that are involved in formation and maintenance of the unique structures at the telomeres and centromeres (Shore, 1997; Straight, 1997). Df31 is a protein originally isolated as a factor promoting decondensation of Xenopus sperm in a Drosophila cell free replication system (Crevel and Cotterill, 1995). It appeared to act by the removal of sperm specific proteins and the loading of histones onto the sperm DNA. Additional experiments showed that Df31 could also load histones onto naked DNA. This led to the suggestion that, like nucleoplasmin it might act as a chaperone for histones during the process of chromatin formation. The characterisation of Df31 presented in this paper strongly suggests that sperm decondensation is not the main function of this protein and that it has a more general involvement in chromatin structure. MATERIALS AND METHODS Cloning and sequencing The gene was cloned using expression screening using a λ zap library purchased from Stratagene, and an antibody made by injection of full length purified protein into rabbits. Short clones produced from this were then used to screen a λ fix library, also from stratagene by DNA screening. DNA sequencing was carried out using the Sequenase kit (US Biochemicals, Cleveland, OH) or samples were sent to Advanced Biotechnology Centre, Charing Cross and Westminster Medical School for ABI sequencing. Nucleotide sequence analysis and amino acid comparisons were performed using Geneworks (IntelliGenetics Inc.) for the Macintosh and the BLAST programmes at NIH. In addition sequences from the Berkeley genome sequencing project were accessed using the flybase resource. Mass spectroscopy was carried out by M scan. In situ chromosome localisation Hybridisation of Df31 cDNA to Drosophila (Oregon R) third instar

larval polytene chromosomes was performed essentially according to published procedures (Ashburner, 1989). Interpretation of results was performed with the assistance of M. Ashburner and colleagues (Cambridge, UK). Immunostaining reagents Df31 antibodies were made in rabbits by injection of the purified Df31. This was carried out either in the lab of Paul Fisher at StonyBrook, or by Neosysteme Strasbourg. Affinity purification of antibodies was carried out by passing the crude antibody over a Df31 column, made by crosslinking purified Df31 onto CNBr activated Sepharose as described previously (Crevel and Cotterill, 1995). The antibodies were eluted with 0.2 M glycine, pH 2.5. Tubulin monoclonal antibodies and labelled wga were from Sigma. Monoclonal ubiquitin and histone antibodies were from Chemicon. Secondary antibodies used for westerns were peroxidase labelled anti-mouse, and anti-rabbit IgG purchased form Pierce. Secondary antibodies for immunostaining were Texas Red labelled anti-mouse and fluorescein labelled anti-rabbit, both of which were purchased from Vector labs. Western blot analysis Western blots were carried out according to standard procedures (Harlow and Lane, 1988). Drosophila from the stages of development as indicated were dechorionated and homogenised directly into standard 2× SDS protein loading buffer. Approximately equal amounts of protein were loaded onto 10% polyacrylamide gels and after electrophoresis, passively transferred onto Hybond ECL membrane (Amersham). This produces 2 blots one on each side. Antiα-Tubulin was used as a standard loading control. The blots were developed using SuperSignal Substrate Western Blotting (Pierce) as per the manufacturer’s instructions. Immunohistochemistry Embryos were fixed as previously described (Ashburner, 1989) and stored in methanol at 4°C. After rehydration and blocking in PBT (PBS, 0.1% Triton X-100, 1% BSA, 0.02% Na azide) for 3 hours at room temperature, the primary antibody incubation was carried out at 4°C overnight. The embryos were washed 3 times each for 30 minutes in complete Pbn (PBS, 0.1% Triton X-100, 0.1% BSA, 2% normal goat serum). Fluorescent secondary antibody was added and incubated at room temperature for 1 hour. The samples were washed 2 times with complete Pbn and five times with Pbt (PBS, 0.1% Tween20), and the Pbt was removed and replaced with 4% propyl gallate in 80% glycerol as an antifading agent before viewing on a Bio-Rad Confocal 600. Subcellular fractionation Dechorionated Drosophila embryos were homogenised in the presence of 3 vols of nuclear prep buffer (15 mM Tris-HCl, pH 7.5, 15 mM NaCl, 60 mM KCl, 0.34 M sucrose, 0.5 mM spermidine, 0.1% β-mercaptoethanol, 2 mM EDTA, 0.5 mM EGTA) plus protease inhibitors, with a loose pestle to help maintain the integrity of the nuclei. The homogenate was filtered through two layers of Miracloth to remove cell debris and centrifuged at 25,000 rpm for 20 minutes in a Beckman TL100. The resulting supernatant was the cytoplasmic fraction. The pellet was washed again with the same buffer and was then subjected to successive washes with buffer plus 0.5% Triton, 0.5% Triton and 250 mM NaCl, 0.5% Triton and 500 mM NaCl, 0.5% Triton and 2 M NaCl. Two washes were carried out with each buffer, and in all cases the pellet was incubated with the buffer for 10 minutes at 4°C, and the pellet and supernatant separated by centrifugation at 25,000 rpm for 10 minutes. Equivalent amounts of each fraction were then passively transferred to nitrocellulose and western blots were carried out as above.

Df31 and chromatin structure in D. melanogaster Chromatin purification Chromatin was prepared by a modification of the method described by Hancock (1974) with very low ionic strength buffers. Drosophila embryos were homogenised with one volume of 0.25 mM EDTA. The filtered homogenate was spun for 10 minutes at 7.5,000 rpm (Sorvall SS 34), the nuclei pellet was washed with 0.25 mM EDTA. The pellet was adjusted to 0.5% NP40, layered on top of solution of 0.1 M sucrose 0.5 mM Tris-HCl, pH 8.5, and spun for 20 minutes at 10,000 rpm. Micrococcal nuclease digestion Chromatin bodies were adjusted to 2 mM CaCl2. Micrococcal nuclease (Sigma) was added at a concentration of 30 units/ml, and incubation was carried out at 15°C for 5 minutes. When indicated oligoncleosomes were further purified on sucrose gradient as described (Ritzi et al., 1998) with the following modifications: after the micrococcal nuclease digestion, the chromatin was adjusted either to 0.5% Triton X-100 (low salt gradient) or 500 mM NaCl plus 0.5% Triton X-100 (high salt gradient). In both cases the sucrose solutions were adjusted to the same detergent or salt concentrations. Formaldehyde cross-linking Chromatin was cross-linked with formaldehyde 1%, exactly as described (Gohring and Fackelmayer, 1997). In vitro experiments For chromatin assembly assays, the extracts were a kind gift of Patrick Varga-Weisz and were prepared as described previously. The DNA substrate was prepared by cutting plasmid pT1 with EcoRI and ClaI and performing an end filling reaction in the presence of biotin dUTP. Biotinylated DNA was bound to streptavidin magnetic beads (Promega) in 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 2 M NaCl. The bound DNA was washed with the same buffer without NaCl. Chromatin was reconstituted on the DNA by incubating it in the chromatin assembly extracts for 5 hours as previously described (Sandaltzopoulos et al., 1994) and the beads were washed with Ex50 buffer (10 mM Hepes, pH 7.6, 50 mM KCl, 1.5 mM EGTA, 10 mM β-glycerophosphate, 10% glycerol and 0.5% NP40). In order to determine the positions of elution of Df31 and histones the beads were washed sequentially with Ex50 containing 100, 250, 500 mM and 2 M NaCl. All fractions and in addition the pellet remaining after the 2 M wash were adjusted to 1× protein loading buffer and subjected to PAGE and western blotting to visualise the positions of the Df31 and histones. Microinjection of embryos Preparation of embryos and microinjection was carried out as previously described (Ashburner, 1989) The antisense oligos were made up in 10 mM Na phosphate, pH 7.8, 5 mM KCl at concentrations of 0.4 mg/ml. The embryos were injected with antisense oligo at 15-45 minutes and then left to age for 2 hours. They were then fixed in 4% formaldehyde/heptane and stained with DAPI to visualise the DNA. Where any further analysis was carried out, embryos after having been selected with DAPI were hand peeled, refixed in 4% formaldehyde for 5 minutes and then stained with antibodies against Df31, wga, and tubulin as described above.

Fig. 1. Protein sequence of Df31. Positive and negative amino acids are indicated by a + or – sign, respectively. The letters in bold represent the amino acid sequence obtained from the peptide sequencing.

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RESULTS Isolation and characterisation of a novel gene, Df31 Antibodies raised against purified DF 31 were used to screen a lambda expression library. The short clone obtained was used for further screening to isolate several much longer clones. Combining our data with EST sequences from the Drosophila genome sequencing project data suggests that the Df31 protein is coded for by a message of at least 1.6 kb. Translation of the RNA (Fig. 1) reveals an open reading frame of 183 amino acids which is sufficient to code for a protein of approximately 18.5 kDa. The identity was confirmed by carrying out proteolysis on purified Df31 and finding a 20 out of 20 match in amino acid sequence in a fragment bounded by a chymotrypsin cleavage site. Furthermore, 85% of the primary amino acid sequence deduced from the cDNA sequence was confirmed by MALDI-TOF mass spectrometry analysis of a V8 protease digest of purified DF31 (data not shown). We mapped the gene to position 39e on chromosome 2. The deduced amino acid sequence of Df31 shows no homology to any other sequences currently in the databanks, and no known motif apart from the presence of a putative bipartite nuclear signalling sequence at position 95-108. The amino acid composition reveals a very high proportion of hydrophilic amino acids (>65%), and a theoretical PI of 4.3. The predicted secondary structure of Df31 is largely α helical, and the N and C termini are likely to be involved in the formation of inter- or intra-molecular coiled coil structures. The calculated mass of 18 kDa for Df31 differs significantly from the molecular mass of Df31 deduced from the electrophoretic migration of the protein which was between 26 and 33 kDa. However, mass spectroscopic analysis (not shown) of the purified Df31 suggested that the actual molecular mass of Df31 was near 18 kDa. The difference between the actual molecular mass and its apparent mass upon gel electrophoresis can be explained in part by the low proportion of hydrophobic amino acids, the main sources of interaction with SDS. However, this is not a complete explanation since coupled in vitro transcription translation of Df31 produced a protein migrating at 20-21 kDa on SDS-PAGE (data not shown). This suggests some contribution from modification of the protein. In agreement with data obtained previously, the MALDI-TOF analysis showed that DF31 was phosphorylated at a site between amino acids 4 and 44. There are only three possible positions of modification in this fragment and only one of these corresponds to a known phosphorylation consensus site, a casein kinase II site at serine 41. In addition the mass spectroscopic analysis suggested that other unidentified modifications were also present on the protein. DF31 protein is expressed throughout the life cycle Df31 is very abundant in early embryos, comprising

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approximately 0.1% of the total cellular protein. Analysis of the expression of the protein throughout the life cycle should tell whether it has a role only in early embryos or if it has a more widespread role. An example of a developmental western profile is shown in Fig. 2. Averaging the results from ten such blots suggests that the expression of Df31 is roughly constant from early embryos through to pupae. The level is slightly lower in males (~twofold) and slightly higher in females (~threefold). The generally high expression of this protein at all times suggests that Df31 has a generalised role in all cell types, not just those which are actively dividing. Microinjection of antisense Df31 oligonucleotides causes clumping of chromatin To investigate the cellular function of Df31 we microinjected Df31 antibodies into embryos. This had no effect, probably because Df31 is such an abundant protein. We therefore used the alternative approach of reducing the levels of the protein by interfering with the metabolism of the RNA message with antisense oligonucleotides. We designed antisense oligonucleotides from two separate regions of the gene (one at the N terminus and one at the C terminus) predicted not to be involved in secondary structure. A control oligonucleotide had the same base composition as one of these but a jumbled base order. In all cases the oligonucleotides were short (14-17 bases in length) to reduce the amount of non specific interactions. All dG bases were replaced by 2′Omethyl rG to limit degradation (Fig. 3A). Embryos (15 to 45 minutes old) were injected with antisense oligonucleotide, left to develop for 2 hours, and then stained for DNA using DAPI. This experiment was carried out on three independent occasions, and in each case >100 embryos were injected for each of the oligonucleotides. Injection with control oligonucleotide showed only small numbers of embryos which could score as abnormal after injection (6, 9 and 8%, respectively, for the three experiments). However, both antisense oligonucleotides caused raised levels of abnormalities in the localisation of the DNA in injected embryos as revealed by DAPI staining (oligo1 – 18, 24 and 22%, oligo2 – 26, 33 and 17% in each of the three experiments, respectively). The abnormalities seen in this case were also far more severe and examples of these are shown in Fig. 3B. This is probably an underestimate of the frequency as only embryos with a pronounced phenotype were classified as abnormal. Embryos with a slight abnormality were counted as normal, as these might have been injection, fixation or staining artefacts, although very few of them were observed in the control embryos. In addition a higher percentage of the antisense injected embryos were devoid of nuclei, but again since these were occasionally seen in control injections these were also classed as normal. In most cases defects were characterised by large irregular clumps of chromatin. These were much fewer in number than the expected number of nuclei, and in most cases the DAPI staining was more intense than that expected from a single nucleus. In a few embryos more regular figures that look like groups of three or four distinct nuclei were also seen. In these cases the overall arrangement of the nuclei was more normal than those where large clumps of chromatin were seen. It is therefore possible that these represent earlier stages in the degeneration process.

Examples of the embryos previously selected as having both normal and abnormal phenotypes were fixed and hand devitellinised, and then stained with antibodies against Df31, tubulin and fluorescently labelled wga to visualise the positions of the nuclear pores. The recovery of embryos from this procedure was poor, as they were significantly more fragile than uninjected embryos treated in the same way. However, while the ‘normal’ embryos which survived the procedure showed the expected levels of df31 the ‘abnormals’ had little or no Df31 staining. Many of the abnormal DNA figures were surrounded by a wga staining structure suggesting that they were surrounded by a nuclear membrane (Fig. 3C). In addition for a few embryos staining with anti-tubulin antibodies revealed the presence of abnormal spindles (data not shown). Df31 is associated with chromosomal DNA throughout the cell cycle The dramatic effect caused by the reduction of the levels of Df31 in the nucleus could indicate a defect at the level of the chromatin structure. However, it could also be explained by failure in other processes, e.g. mitosis. The cellular structures with which a protein interacts should reflect the processes in which it is involved. We therefore analysed the cellular location of Df31using affinity purified antibodies to carry out immunostaining of Drosophila fixed at different points throughout the life cycle. In newly deposited embryos large amounts of staining were seen throughout the embryo. Post cellularisation the staining was largely confined to the chromatin (Fig. 4). The association with chromatin was maintained at all stages of the cell cycle. Association of Df31 with chromatin was also seen at later stages of the life cycle in the imaginal discs, the gut, the testes and associated glands, the ovaries and the brain (data not shown). Df31 was also observed to be associated with all regions of the polytene chromosomes, although it was

Fig. 2. Western blot to show the developmental expression of Df31 protein. Total cell extracts from each of the stages were prepared as described in the Materials and Methods. In order to calculate relative amounts in each lane the gel was also stained with tubulin control. Embryonic stages loaded on the gel are shown at left.

Df31 and chromatin structure in D. melanogaster

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Fig. 3. The effect of injection of Df31 antisense oligonucleotides into embryos. (A) The sequences of the antisense oligonucleotides that were injected into embryos. The first 2 sequences are antisense to df31, the third shows the sequence of the control which has the equivalent numbers of each base to oligo 1 but which are re-ordered. m marks a base that was incorporated in a methylated form to protect against degradation. (B) Examples of the phenotypes obtained by injection. After injection the embryos were aged for 2 hours, then fixed and the DNA visualised with DAPI. n, normal embryo of expected age p1, p2 and p3 – the different types of aberrations observed with oligos 1, 1 and 2, respectively. Inset on p1 is an enlargement of one of the abnormal nuclei. (C) The levels of df31 are reduced on injection of antisense but not control oligos. Fixed and DAPI stained embryos were hand devitellinised and then stained for Df31 and wga as shown. (a and b) Embryos injected with control oligos. (c to f) Antisense injected embryos, (c and d) oligo 1, (e and f) oligo 2. For each oligo examples are shown of one abnormal region, and one relatively normal region proximal to an abnormal region.

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JOURNAL OF CELL SCIENCE 114 (1) particularly concentrated in the interband regions (data not shown).

Fig. 4. Analysis of the subcellular location of Df31 by fractionation. Fractions were prepared from 0-5 hour Drosophila embryos as described in Materials and Methods and equivalent amounts of each were subjected to PAGE and western blotting with Df31 antibodies. (a) Cytoplasmic fraction, (b) 0.5% Triton wash, (c) 0.5% Triton plus 250 mM NaCl, (d) 0.5% Triton plus 500 mM NaCl, (e) 0.5% Triton plus 2 M NaCl, (f) pellet remaining after all washes. Size markers (kDa) are shown at left.

The association of Df31 with chromatin resists high salt extraction To measure the strength of binding of Df31 to chromatin we separated cells into cytoplasmic and nuclear fractions and performed sequential extraction with increasing salt concentrations. Fig. 5 shows such an analysis on 0-5 hour embryos. Some of the Df31 was extracted in the cytoplasmic wash (lane a). Much of this probably came from the

Fig. 5. Immunolocalisation of Df31 in Drosophila embryos. Early embryos were fixed as described and then stained with antibodies against Df31 and tubulin (a), or Df31 and histone (b). The panels are representative of the type of staining observed in different stages of the cell cycle, and show that Df31 appears to be associated with the DNA all the way through these early cycles. Df31 staining is less distinct than that of the monoclonal antibodies used to stain the histone and tubulin, this may be explained by the observation that Df31 is more labile during the fixing process than either of the other antigens. (c) Merged images.


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10 kDa larger than df 31. Since this cross reacts with antibodies affinity purified against Df31 we believe this to be a modified form of Df31. This assertion is also supported by the observation that the ratio of the smaller and larger protein bands is variable between different fractionations. Occasionally a third band a further 7-10 kDa higher is also detected (e.g. see Figs 6, 8). These bands do not represent differentially phosphorylated forms of the protein (data not shown), however, at this point the exact nature of the modification is not clear.

Fig. 6. Demonstration of the association of Df31 with chromatin. (a) Chromatin was prepared by the Hancock procedure and subjected to digestion with micrococcal nuclease as described in Materials and Methods. The supernatant released at each of the times of digestion as shown (minutes) was run on a polyacrylamide gel, blotted and analysed for the presence of Df31 by western blotting. Size markers ( kDa) are shown at left. (b) Chromatin prepared as above was exposed to 1% formaldehyde for the indicated periods of time (minutes). Cross linked DNA proteins complexes were then prepared over CsCl gradients as described in Materials and Methods. Equal quantities of each reaction (as judged by the concentration of DNA) were boiled to break the crosslinks and then analysed by polyacrylamide gel electrophoresis and western blotting for the presence of rpa and Df31 and polyacrylamide gel electrophoresis for the presence of histones.

precellularisation embryos that were present in the 0-5 hour sample, or from accidental rupturing of nuclei during homogenisation. A further fraction of the protein was washed out in the nucleoplasmic fraction (lane b). Most of the Df31 could only be washed out by the inclusion of salt in addition to Triton in the wash buffers, and a significant amount of Df31 still remained in the nuclear pellet even after treating the nuclei with 2 M NaCl (lane f). Therefore in agreement with the immunofluorescence data, a large percentage of the Df31 is associated with the chromatin containing fractions of the nucleus. In addition a subpopulation of the Df31 associated with the chromatin shows resistance to salt extraction. An estimate of the total amount of Df31 associated with the chromatin, was obtained by performing quantitative western blots using purified Df31 as a standard. Df31 was present at levels approximately one quarter those of the core histones. Interestingly fractions binding at higher salt concentrations, particularly those most tightly associated with the nuclear pellet, contain an additional cross reacting band migrating 7-

The association with the salt resistant nuclear pellet is mediated via the DNA The resistance of Df 31 to salt extraction could be explained by its association with the nuclear matrix. To investigate this possibility we purified chromatin from Drosophila embryos, according to the Hancock procedure (Hancock, 1974) and subjected it to micrococcal nuclease treatment. Proteins associated with chromatin have been shown to be released into the supernatant following such a treatment (Ritzi et al., 1998). Fig. 6a shows that micrococcal nuclease treatment promotes the release of Df 31 in the supernatant. The rate of release is the same as that observed for histones. Consistent with the salt fractionation it was the slow migrating form of Df31 which was most prominent in the fractions released. An association with chromatin rather than with nuclear matrix, is also suggested by the observation that Df 31 can be cross-linked to chromatin by formaldehyde treatment. For this experiment purified chromatin was subjected to formaldehyde cross-linking and the cross-linked chromatin was further purified on caesium chloride gradients. As shown in Fig. 6b Df31 cross-links rapidly to DNA and after 8 minutes of treatment most of the Df31 is bound. Although fast, Df31 cross linking is less efficient than that of either histones, or RPA (a single stranded DNA binding protein involved in DNA replication). This suggests that the interaction of the Df31 with the DNA is not direct. Consistent with this, extensive treatment of the chromatin with uv, which should only fix proteins directly in contact with the DNA, does not cause covalent attachment of the Df31 onto DNA. Under the same conditions both histones and rpa are efficiently (>5%) attached (data not shown). Df 31 co-fractionates with mono- and oligonucleosomes The data presented above suggests that Df31 could be involved with higher order structures of chromatin. We were therefore interested in understanding the relationship between Df31 binding and nucleosome structure. To do this we investigated the association of Df 31 with oligonucleosomes on a sucrose gradient of micrococcal nuclease treated chromatin. The sucrose gradient separates the oligonucleosomes according to their size. The size distribution of the oligonucleosomes can be assessed by observing the sizes of the DNA fragments found associated with the histones (Fig. 7). In gradients run in low salt (Fig. 7a) Df31 is found throughout the gradient both in fractions free of DNA and also in those containing both mono and oligo nucleosomes. If, however, the gradient is run in high salt conditions, by adjusting the micrococcal nuclease reaction to 500 mM NaCl prior to loading onto the sucrose gradient (Fig. 7b), Df 31 elutes in two very distinct peaks. The first one

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Fig. 7. Analysis of association of Df31 with oligonucleosomes by sucrose gradient sedimentation. Chromatin was prepared and subjected to micrococcal nuclease digestion as described in Materials and Methods. The reaction mixture was adjusted to 0.5% Triton (a) and 0.5% Triton plus 500 mM NaCl (b) and then loaded onto a 5 to 40% sucrose gradient. The nature of the DNA present in each fraction was analysed on 1.5% agarose gel. The histone and Df31 contents of the same fractions were visualised by polyacrylamide gel electrophoresis and in the case of Df31 by western blotting.

centred around fraction 6 corresponds to free Df 31. The second which peaks around fraction 18 corresponds to oligonucleosomes. The smallest pieces that Df31 is associated with correspond to approximately 600 bp pieces of DNA or 3 to 4 nucleosomes. The Df31, however, is associated more strongly with fractions where the average size of the DNA is around 1 kb, which should contain more than 5 nucleosomes. In vitro loading of Df31 onto chromatin In order to characterise further the association of Df31 with chromatin we investigated the association of Df31 with plasmid DNA that had been formed into chromatin in an in vitro reaction. Chromatin was assembled in vitro from 0-100′ Drosophila embryo extracts as previously described (Sandaltzopoulos et al., 1994). The association of Df31 with this structure was then analysed by subjecting the in vitro formed chromatin to increasing levels of NaCl (Fig. 8). Consistent with our fractionation studies Df 31 was loaded onto chromatin in the in vitro reaction, and was present in the final structure at a ratio of at least one molecule of Df31 per 2/3 nucleosomes. Again a significant fraction of the Df31 remained bound to the chromatin even after it had been subjected to 2 M NaCl. As observed in the embryo fractionation, those populations of Df31 that showed the

tightest binding were greatly enriched in antibody cross reacting bands that were 7-10 and 14-20 kDa higher in molecular mass. To investigate the effects that Df31 might have on nucleosomes formed in the in vitro reaction, we compared the properties of chromatin formed in normal extracts and in those that had been depleted for Df31. Chromatin formed in the absence of Df31 was unaffected with regard to the stability of the histones to salt extraction, or susceptibility to restriction endonuclease or micrococcal nuclease digestion (data not shown). In addition we could detect no differences in the sizes of the nucleosomes generated by micrococcal nuclease digestion of either reaction (data not shown). DISCUSSION Df31 was originally isolated as a protein that effected sperm decondensation in vitro. The analysis presented in this paper suggests that this is not its major function in vivo, but that its actual role is in the general maintenance of the structure of chromatin. There are two main lines of reasoning that lead us to this conclusion: DF31 shows a strong association with chromatin both in vivo and in vitro.


Fig. 8. Analysis of the association of Df31 with in vitro assembled chromatin. Chromatin loading onto plasmid pT1 was carried out as described in Materials and Methods and subjected to increasingly stringent washes in NaCl. The amount of Df31 released at each salt concentration was analysed by PAGE and western blotting. (a) Levels of Df31 present in the in vitro chromatin loading extract, (b) 100 mM NaCl wash, (c) 250 mM NaCl wash, (d) 500 mM NaCl wash, (e) 2 M NaCl wash, (g) Df31 associated with the pellet remaining after washing in 2 M NaCl. (f) The control column where no DNA has been loaded. Size markers (kDa) are shown at left.

If Df31 were simply a sperm decondensation protein, it would only be required in early embryonic stages. However, western blotting of fractions from different stages of the life cycle show that the protein is present at constant high level in all stages. Immunostaining of early embryos indicates that Df31 does not visibly dissociate from the chromatin at any point throughout the cell cycle. This suggests that it is required throughout the cycle not just in one defined stage. It is also associated with chromatin in cells that have left the mitotic cycle, which suggests a general role rather than one specific to cell division. A chromatin association for Df31 is also supported by biochemical observations from cellular fractionation, micrococcal nuclease digestion, formaldehyde cross linking and sucrose density gradient separation of nucleosomes. These experiments further reveal that Df31 is very abundant on DNA, at levels comparable with that of H1 (one half to one third as abundant as nucleosomes), and that at least part of the Df31 binds tightly into the nucleus. The generality of the association with chromatin is underscored by the in vitro loading of Df31onto chromatin. Since the substrate used was plasmid DNA containing largely bacterial DNA it is unlikely to contain intrinsic chromatin structure. Despite this the ratio of Fig. 9. Model to suggest how lack of Df31 might cause the phenotype observed after injection of the antisense oligonucleotides. (a) Represents what happens when Df31 levels are normal. (b) The level of Df31 is reduced, the increase in the amount of DNA occurring during replication therefore results in inadequate amounts of Df31 being present on the chromatin. This alters the structure of the chromatin (represented by the dotted line) resulting in incomplete replication in the S phase period. Since Drosophila embryos have no checkpoint control they proceed into the next mitosis. This results in incomplete separation of the DNA, and by iteration of the process eventually produces the large masses of DNA observed in the injected embryos.

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Df31:core histones is comparable to that of chromatin extracted from Drosophila embryos. The in vitro loading together with the formaldehyde cross linking also exclude the possibility that the observed tight associations are with other large nuclear structures. In both cases the protein DNA complexes are separated from large protein complexes not containing DNA, either by labelling of the DNA and the use of magnetic beads or by gradient separation of the different density complexes. Although chromatin association is mediated via the DNA we feel that it is unlikely that Df31 makes direct contact with the nucleotide chains. The protein does not to bind to either ss or ds DNA cellulose columns (GC unpublished results). More importantly even with prolonged exposure to uv light we have been unable to cross link detectable levels of Df31 to the DNA. It is therefore likely that Df31 is attached to the DNA via other proteins. We had previously shown that Df31 interacted with core histones in vitro. This together with the almost stoichiometric abundance of the 2 groups of proteins on DNA make the core histones good candidates for the sites of Df31 attachment. However, other proteins may also be involved. Stringent washing of Df31 immunoprecipitates from 0-5 hour embryos reveals strong interactions with several as yet unidentified proteins (HH unpublished observations), and we are presently attempting to identify these and determine their relevance for the association of Df31 with chromatin. One interesting observation made during our studies on the chromatin association of Df31 is that, both in vivo and in vitro, the fractions that are the most tightly bound into the nucleus are considerably enriched in a modified form of the protein. This modification is in addition to the phosphorylation and other unidentified modifications observed by mass spectroscopy, since that analysis was carried out on a homogenous population of lower molecular mass. The exact relevance of this observation to Df31 function is not yet clear, however, it is tempting to suggest that the modification is involved in controlling the interaction of Df31 with chromatin. Lowering the concentration of Df31 in vivo causes disruption of the chromatin structure The other observation which strongly suggests that the Df31 has a general structural role is the serious disruption of nuclear structure observed on injection of antisense oligonucleotides

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against the protein into embryos. Injected embryos contain large irregular masses of DNA, and most of these distorted nuclei contain more than one genomes worth of DNA. There is also a much higher incidence of annucleate embryos. The appearance of the DNA here is somewhat reminiscent of the structures generated by some mitotic mutants of Drosophila and by depletion of factors involved in the condensation of chromatin prior to mitosis (Hirano and Mitchison, 1994). It is therefore likely that they are generated by related procedures, i.e. the failure to separate chromosomes properly during anaphase. Since Drosophila has no checkpoints at this stage of the life cycle linking replication and mitosis, the cycle would carry on with unseparated masses of DNA generating the observed large structures containing several genomes worth of DNA. The appearance of multiple spindles in some of the embryos also suggests a failure of mitosis. It is possible that the more regular groups of nuclei that we see represent earlier stages in the process. In support of this where both aberrations are seen in the same nucleus the more regular figures are always found closer to regions with normal nuclear structure. How might a lack of Df31 generate these structures? Its localisation on the DNA at all times makes it unlikely that it is having its effects on the spindle or spindle poles. The SMC proteins produce a similar effect by failing to condense the chromatin prior to mitosis (Hirano et al., 1995), and it is possible that Df31 could function in a similar way. However, SMC proteins are expressed and localised to the chromatin for restricted periods of the cell cycle – many of them are absent in interphase- whereas Df31 appears to be present and bound to the DNA at all times. In addition a role for Df31 in mitotic chromosome condensation would not explain the requirement for Df31 in cells that have left the mitotic cycle. It is more likely that Df31 has a general function in maintaining the structure of chromatin. This is consistent with its expression profile. It would equally explain the phenotype observed (Fig. 9) since any disruption of chromatin structure might lead to incomplete replication and a subsequent disruption of mitosis. Since Df31 can load histones onto DNA in vitro it could act by facilitating the loading of histones during replication and/or transcription. However, it has not been detected as part of any of the known Drosophila nucleosome manipulation complexes (Ito et al., 1997; Kamakaka et al., 1996; Tsukiyama and Wu, 1995; Varga-Weisz et al., 1997). Furthermore Df31 is not more abundant at times of high DNA replication in the life cycle, and is not concentrated in structures such as puffs where large amounts of transcription are taking place. In addition we have been unable to detect any effect at the level of basic nucleosome structure on in vitro chromatin assembled in the absence of Df31. The explanation that we favour therefore is that Df31 is a novel type of chromatin structural protein. This would explain the requirements for a high abundance of the protein. Intriguingly Df31 maps close to a histone cluster on chromosome 2, which also raises the possibility that these two adjacent regions could be subject to co-ordinate control. It is also more consistent with the predicted secondary structure of Df31 since large amounts of α helix and coiled coil regions are common elements in structural proteins. The in vitro interaction of Df31with core histones further suggests that it may function at an early stage in the generation of structure.

This is further supported by the observed association of the protein with nucleosomes on sucrose gradients. In this case the relatively tighter association with oligo- rather than mononucleosomes suggests that the protein is bound in a structure requiring contacts with several nucleosomes. It may therefore play some role in the higher order associations of nucleosomes. Conclusions The properties of Df31 presented in this paper suggest that it is a novel chromatin structural protein. The identification of this protein therefore represents an important step towards our understanding of the structure of chromatin and the ways in which its structure can be modified. It will be interesting to see whether Df31 has homologues in other organisms. There does not appear to be a yeast homologue – notable since the chromatin structure of yeast is not as compact. In addition nothing with homology from any other organism has yet turned up on database searches, however, the classification of Df31 as a possible fast evolving gene (Schmid and Tautz, 1997) might necessitate the search for homologues in a functional rather than a strictly sequence dependent way. We thank Rick Wood, Steve West, Patrick Varga-Weisz and Isabelle Crevel for critical reading of the manuscript, and Helen Bates, Baz Smith and David Loebel for helpful discussions. This work was supported by the Marie Curie Cancer Foundation.

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