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Gamete Biology |
Departments of Obstetrics and Gynecology5
Biology,6 McGill University, Montréal, Québec, Canada H3A 1A1
Department of Cell Biology,7 Albert Einstein College of Medicine, Yeshiva University, Bronx, New York 10461
INSERM U309,8 Institut Albert Bonniot, Grenoble, France
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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developmental biology, early development, gamete biology, oocyte development, ovum
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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All histones are encoded by multiple genes; however, the H1 histones show greater variability than the core histones. Six H1 subtypes have been identified in mammalian somatic cells [5, 6]. Five of these share a highly conserved central globular domain, although their N- and C-terminal tails vary somewhat, and are termed the somatic H1 subtypes. These constitute the major linker histones of proliferating cells, although their relative proportions vary among cell types. Whether the individual subtypes are functionally different remains unresolved [7]; there is evidence for their differential distribution on chromatin [8], yet mice lacking either H1c, H1d, or H1e show no phenotypic abnormalities [9]. The sixth subtype, H10, differs from the somatic subtypes in both its globular domain and flanking tails [10]. Additionally, in contrast with the somatic H1 mRNAs, which accumulate specifically during S-phase, H10 mRNA is present throughout the cell cycle [7]. Consequently, H10 accumulates on the chromatin of many cell types after they exit the mitotic cell cycle [11]. The high sequence-conservation of H10 across mammalian species suggests a specific function for this subtype, yet mice lacking H10 are apparently normal [12].
In many nonmammalian organisms, oocytes contain a complement of linker histones that differs from that present in somatic cells. Sea urchin oocytes contain a cleavage-stage histone H1 (csH1) that continues to be synthesized from maternal transcripts during the early embryonic cleavage divisions [13, 14]. Xenopus oocytes contain an evolutionarily related linker histone, termed B4, which persists until about the midblastula stage, when it is replaced by the somatic H1 subtypes [15, 16]. Similarly, oocytes of several marine invertebrates express unique H1 subtypes [17, 18], and H1 appears to be replaced by the high-mobility group protein D (HMG-D) in oocytes and preblastula embryos of Drosophila [19]. In Caenorhabditis elegans, one of the somatic H1 subtypes is specifically required for gene silencing during germ-cell development [20]. CsH1, B4, and H1oo are all larger than the somatic H1 subtypes, which may be relevant to their biological function. Furthermore, experimentally altering the timing of the switch from B4 to somatic H1 in Xenopus embryos correspondingly shifts the period of time during which the embryo is competent to receive mesoderm-inducing signals [21]. Thus, developmentally regulated changes in the H1 complement may play a key role in establishing the normal structure of embryonic chromatin.
The presence of stage-specific H1 subtypes in nonmammalian oocytes and embryos, together with its role in regulating chromatin activity, suggests that H1 may play a central role in the nuclear reprogramming that is thought to be required for successful animal cloning by nuclear transplantation. To evaluate this possibility, it is essential to define the H1 subtypes that are present on mammalian oocyte and embryonic chromatin. Antibodies raised against rat somatic H1 failed to detect H1 in murine and bovine oocytes or early embryos, whereas it became detectable during the early embryonic cleavage divisions [22, 23]. The mRNA encoding H10 was detected in these cells, raising the possibility that the H1 complement of mammalian oocytes might resemble that of nondividing somatic cells in that somatic H1 became depleted and H10 became enriched [24]. Subsequently, however, proteins comigrating with somatic H1 were detected in mouse oocytes and early embryos using radiolabeling [25] and antibodies raised against Xenopus somatic H1 [26]. In contrast, H10 was not detected [27]. Most recently, a mammalian homologue of the oocyte-type H1 of frog and sea urchin, termed H1oo, has been identified [28]. H1oo is present in oocytes and one-cell embryos but is undetectable in blastomeres at the two-cell stage and beyond.
In light of these reports, we have reexamined whether histone H10 and somatic H1 subtypes are expressed in mouse oocytes and embryos. We used highly specific monoclonal antibodies and metabolic labeling to study the presence and synthesis of these H1s at various stages. We also employed mice lacking the H10 gene [12] as a rigorous approach to assess the specificity of these assays. Our results indicate that H10 is expressed in oocytes and that its abundance relative to somatic H1 declines during the postfertilization cleavage divisions. A subset of the somatic H1 subtypes is synthesized by oocytes and the full complement in embryos. Taken together with the previous data, these results imply that the H1 complement in mammalian chromatin changes during late oogenesis and early embryogenesis.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Experiments were performed using oocytes and embryos collected from CD-1 mice (Charles River Canada, St-Constant, QC, Canada), mice lacking a functional H10 gene, and wild-type mice of the same genetic background as the H10-null mice. The H10-null mice were originally produced by Sirotkin et al. [12]. Both the null and wild-type derived from the founders were crossed into CD-1 mice to produce H10-null and wild-type mice of mixed genetic background. Oocytes and embryos were collected and cultured as described [22].
H10 Genotyping
Two-millimeter tail samples were cut into several pieces and incubated overnight at 60°C with gentle agitation in 10 mM Tris (pH 7.5), 150 mM NaCl, 50 mM EDTA, 1% SDS, and 100 µg/ml proteinase K. DNA was extracted with a fresh 1:1 solution of phenol and chloroform and precipitated with a solution of 80% isopropanol, 20% 3 M sodium acetate, and 0.2% 0.5 M EDTA. DNA was centrifuged, washed with 70% ethanol, dried, and resuspended in 25 µl of sterile water. Three hundred to 1000 ng of DNA was amplified in a buffer containing 1.5 mM MgCl2, 0.2 mM dNTPs, 120–180 pmol of each primer, 5% DMSO, and 1.25 units Taq polymerase using the following conditions: 5 cycles 94° (30 sec), 57° (45 sec), 72° (75 sec); and then 39 cycles of 94° (30 sec), 60° (45 sec), 72° (75 sec). The following primers flank the neomycin cassette and therefore amplified both wild-type and mutated H10 alleles:
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Reverse Transcription and Polymerase Chain Reaction Amplification
RNA extraction and cDNA synthesis were performed essentially as previously described [29]. To test for the presence of H10 mRNA, RNA was extracted from 100 oocytes or embryos and resuspended in 10 µl of water. Random hexamers and M-MLV reverse transcriptase were used for the reverse-transcription reaction, which was done on 40 oocyte/embryo-equivalents. Nine equivalents were subjected to polymerase chain reaction (PCR) using the H10 primers shown below and the following conditions: 96°, 2 min; then 36 cycles of 94° (1 min), 58° (45 sec), 72° (45 sec); and then 72°, 5 min. Products were electrophoresed in 4% agarose gels and stained using ethidium bromide. Each oocyte or embryo stage was analyzed at least twice.
To test for changes in the length of the poly(A) tail, RNA was extracted from 50–75 oocytes or embryos, and 20–30 equivalents was reverse-transcribed using AMV and the oligo-d(T) primer-adapter not labeled with Cy5. Eight to 10 equivalents were subjected to PCR using the appropriate mRNA-specific 5'-primer, the Cy5-labeled oligo-d(T) primer-adapter, and the following conditions: 93° (5 min); then 28 cycles of 93° (30 sec), 60° (1 min), 72° (1 min), + 5 sec/cycle; and then 72° (7 min). A portion of the products was electrophoresed in 4% agarose gels and stained using ethidium bromide. Another portion was electrophoresed through a 6% agarose denaturing gel and the fluorescence quantified using an ALFexpress (Amersham Pharmacia, Montréal, QC, Canada). Each oocyte or embryo stage was analyzed at least twice.
Primers
The primers used were as follows:
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Antibodies
Somatic subtypes of H1 were detected using a rabbit polyclonal antibody raised against mouse H1 and affinity-purified against calf thymus H1 [22], diluted 1:50 for immunofluorescence and 1:500 for immunoblotting. H10 was detected using a mouse monoclonal antibody (clone 34 [30]) raised against H10, used undiluted for immunofluorescence and 1:100 for immunoblotting.
Immunofluorescence
Zonae pellucidae were dissolved by a brief exposure to warmed acidified Tyrode medium (pH 2.5, containing 0.1% PVP; Sigma-Aldrich, Oakville, ON, Canada). The oocytes or embryos were then fixed in fresh 2% para-formaldehyde in PBS for 10 min at room temperature with gentle agitation. Fixed cells were blocked in PBS, 3% BSA, 0.1% Triton X-100 for 30 min at room temperature with gentle agitation. Cells were deposited into 15-µl drops of primary antibody placed in 72-well trays and incubated in a humidified chamber overnight at 4°C with gentle agitation. The following day, cells were washed three times for 5 min each in blocking buffer, then transferred to 15-µl drops of the appropriate fluorescence-conjugated secondary antibody (Jackson ImmunoResearch, West Grove, PA) diluted 1:250 and incubated for 1 h at room temperature with gentle agitation. Cells were washed three times in blocking buffer and mounted on silicon-coated microscope slides in Moviol medium (Hoechst, Montréal, QC, Canada) supplemented with 0.4 µg/ml of the DNA-binding dye DAPI (Roche, Montréal, QC, Canada) and 2.25% of the antifade agent DABCO (Sigma-Aldrich). At least three replicates of 10 specimens per oocyte or embryo stage were stained with the antisomatic H1 and anti-H10 antibodies. Stained and mounted cells were visualized using an Olympus BX60 fluorescent microscope and images were captured with CytoVision System software (Applied Imaging, Santa Clara, CA).
Electrophoresis and Immunoblotting
Cells were deposited within a minimal drop of media into a sterile, siliconized 0.5-ml PCR tube, frozen immediately on dry ice, and then stored at -80°C. Cells were thawed in a triple lysis buffer containing 50 mM Tris-HCl (pH 8.0), 500 mM NaCl, 0.1% SDS, 0.6 mM PMSF, 1 µg/ml of aprotinin, leupeptin, and N-tosyl-L-phenylalanine chloromethyl ketone, 1.0% NP-40, and 0.5% sodium deoxycholate, vortexed, and placed on ice for 15 min. Samples were centrifuged at 14 000 rpm at 4°C for 30 min, the supernatant was retained, and the pellet was discarded. A 1:1 volume of SDS-PAGE sample buffer of 15.6 mM Tris-HCl (pH 6.8), 5.0% glycerol, 0.5% SDS, 360 mM ß-mercaptoethanol, and bromophenol blue was added and samples were denatured for 5 min at 95°C. Samples were electrophoresed through a standard 12% Tris/glycine SDS-polyacrylamide gel for 1
h at 100 V.
Proteins were transferred for 2 h at 70 V onto a nitrocellulose membrane (Amersham) in 25 mM glycine (pH 9.7), 25 mM ethanolamine, 10% methanol, and 0.1% SDS. Membranes were then blocked in a Tris-buffered saline (TBS) solution containing 5% milk for 30 min at room temperature with gentle agitation. Antibodies were diluted in the same block solution. Membranes were incubated in primary antibody overnight at 4°C with gentle agitation. Following three washes in TBS, membranes were then incubated in the appropriate biotin-conjugated secondary antibody diluted 1:5000 in TBS for 1 h at room temperature, then washed as above and incubated in streptavidin-HRP diluted at 1:1000 in TBS for 30 min at room temperature. Protein was revealed on the membrane using ECL+ (Amersham).
Radiolabeling and Autoradiography
Up to 50 cells were incubated in a 20-µl drop of medium that contained 100 µCi/ml each of [3H]-lysine and [3H]-alanine (Amersham) for 24 h (growing oocytes) or 6 h (embryos) at 37°C in 5% CO2. The embryos were labeled during periods of time expected to coincide with DNA replication, indicated here as the number of hours after hCG injection: one-cell, 26–32 h; two-cell, 36–42 h; four-cell, 54–60 h; eight-cell, 70–76 h; morula, 80–86 h; blastocyst, 95–111 h. After labeling, cells were washed three times in Hepes-buffered media, transferred to a sterile, siliconized PCR tube in a small drop of media, and immediately frozen on dry ice for storage at -80°C.
Total histones were acid extracted as described [31]. Triple lysis buffer was added to frozen cells, which were vortexed briefly and set on ice for 15 min. Five micrograms of calf thymus histone were then added to the samples. Sulfuric acid was added to a final concentration of 0.4 N in 5–6 times the original volume, and the samples were incubated for 1 h on ice. Tubes were centrifuged at 14 000 rpm for 30 min at 4°C, and the supernatant was transferred to a fresh siliconized tube. To precipitate the histones, 10 volumes of 100% cold acetone were added to the supernatant and the mix vortexed and incubated overnight at -20°. The precipitate was centrifuged at 14 000 rpm for 30 min at 4°C, and the pellet obtained was washed twice in cold 80% acetone and then dried in a Savant Speed Vac (Fisher, Montréal, QC, Canada) with heat for about 1 h, until the pellet turned white and powdery. Proteins were resuspended in distilled water and stored at -20°C for up to 1 wk before electrophoresis.
To extract H1 selectively, cells were processed as above except that sulfuric acid was replaced by perchloric acid, which added to a final concentration of 5% in dH2O. The samples were incubated on ice for 1 h. Supernatant was separated from cell debris and acid-insoluble proteins as described above, and HCl was added to the supernatant at a final concentration of 0.1 N. Histones were then precipitated in 100% cold acetone, and the tube was mixed continuously overnight at 4°C on a nutator. The following day, the histone pellet was centrifuged down and acetone was removed. The pellet was then washed twice in cold acidified acetone and the pellet dried and resuspended as described above.
For autoradiography, the samples were electrophoresed through a 12% PAGE gel as described above. The gel was then stained with Coomassie blue dye for 3–4 h, then destained overnight in a solution containing 5% methanol and 7.5% glacial acetic acid. The following morning, proteins in the gel were fixed by incubating the gel in a solution of 30% methanol and 10% glacial acetic acid for 1 h with gentle agitation. The gel was then impregnated with EN3HANCE autoradiography enhancing solution for 1 h. Next, an excess of cold water was added to the tray containing the gel, and the tray was gently agitated at 4°C. The gel was then placed on two pieces of Whatman paper and dried at 70°C for 2 h in a slab gel drying apparatus (Bio-Rad, Oakville, ON, Canada) under vacuum. Fluorescent proteins in the gel were visualized by exposing the gel to Kodak BioMax MS film. Radiolabeling experiments were conducted at least twice. The gels and autoradiograms were digitized, and data shown are representative composites.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To test whether mRNA encoding histone H10 was present in oocytes and preimplantation embryos, RNA was extracted, reverse transcribed using random hexamers, and PCR-amplified using primers derived from mouse H10 mRNA sequence that were expected to generate a 148-nt product. PCR product of the expected size was obtained from germinal vesicle (GV)-stage and metaphase II oocytes as well as from embryos at all stages of preimplantation development (Fig. 1A and data not shown). These results confirmed that mRNA encoding H10 is present in oocytes and early embryos.
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We then tested whether the extent of polyadenylation of H10 mRNA changed during this period of development. We employed a modified PCR procedure known as the poly(A) test (PAT) developed by Strickland and colleagues [32], in which cDNA is synthesized using a primer adapter consisting of oligo-d(T) coupled at its 5' end to an arbitrary sequence, and the cDNA is amplified using the primer adapter and a gene-specific 5' primer. As the primer adapter can anneal anywhere along the poly(A) tail during reverse transcription, the PCR reaction is expected to produce products of different lengths, the average size of which will increase when the poly(A) tail of the mRNA is lengthened. We first confirmed that we could detect the polyadenylation of mRNA encoding tPA that occurs during meiotic maturation and which has been detected using the PAT [32, 33]. As shown in Figure 1, B (left) and C, metaphase II oocytes generated a range of PCR products of which the maximum length was longer than that generated from GV-stage oocytes. These results confirmed that the PAT could detect changes in the state of polyadenylation of an mRNA.
The PAT method was then used to examine changes in the polyadenylation state of H10 mRNA during maturation and after fertilization. In contrast with tPA, the abundance and average length of the PCR products corresponding to polyadenylated H10 mRNA were not longer in metaphase II oocytes than in GV oocytes (Fig. 1, B [right] and C). Very little polyadenylated H10 mRNA was detected at the one-cell stage, but it then increased in abundance and average length beginning at the four-cell stage. These results indicate that H10 mRNA does not become polyadenylated during meiotic maturation, although it becomes polyadenylated in cleavage-stage embryos.
Expression of H10 Protein in Oocytes and Early Embryos
To examine whether H10 protein was present in oocytes and early embryos, we employed a monoclonal antibody that is highly specific for H10 and has been used to follow expression of H10 in differentiating somatic cells [30]. To verify its specificity, we compared wild-type animals and animals lacking a functional copy of the H10 gene. To confirm the genotype of the H10 knockout animals, we developed a PCR test in which primers lying outside the region modified by homologous recombination are used to amplify an 1142-nt fragment from the wild-type allele and a 1590-nt fragment from the mutant allele (Fig. 2A). This allowed us to unambiguously verify the genotype of each animal used in these experiments. By immunoblotting, the anti-H10 antibody recognized a single band of the expected size in somatic cell extracts from wild-type animals but not from H10 -/- animals, whereas an antibody recognizing the somatic H1 recognized bands of the expected size in both wild-type and H10 -/- animals (Fig. 2B). These results confirm that the antibody specifically recognizes histone H10.
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We then stained wild-type and H1(0) -/- fully grown GV-stage oocytes with either the antisomatic H1 or anti-H10 antibody. Using the antisomatic H1 antibody, very weak or absent nuclear staining was detected in both wild-type and H10 -/- oocytes (Fig. 3). In contrast, the anti-H10 antibody produced distinct nuclear staining in the wild-type oocytes but not in the -/- oocytes. These results indicate that histone H10 is present in the nuclei of GV-stage oocytes.
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To determine whether H10 remains present following meiotic maturation and fertilization, oocytes at metaphase II and embryos at different stages of preimplantation development were stained using these antibodies. In wild-type embryos, somatic H1 could be detected at the two-cell stage but was much stronger at the four-cell stage (Fig. 4, upper). Histone H10 was also detectable in embryos at the same stages. Unlike the case for somatic H1, however, the staining did not become stronger in more advanced embryos. In H10 -/- embryos, no staining was observed using the anti-H10 antibody, confirming the specificity of the staining in wild-type embryos. The antisomatic H1 antibody occasionally stained the pronuclei of -/- one-cell embryos (see Fig. 4), but this was observed in less than 10% of the cases and we were unable to establish conditions under which a higher fraction of one-cell embryos were stained. These results indicate that H10 is present in the nuclei of oocytes and early embryos and further suggest that its abundance relative to somatic H1 decreases during preimplantation development.
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Synthesis of Histone H1 Subtypes in Oocytes and Early Embryos
To identify linker histone subtypes that are synthesized by oocytes and early embryos, we radiolabeled wild-type oocytes and embryos and extracted the acid-soluble proteins. The results indicate qualitative changes in acid-soluble protein synthesis during this period but do not provide quantitative information because labeling periods differed, the amino acid pools are unknown, and equal radioactive counts were not loaded into each lane. Labeled species comigrating with somatic histone H1 were not detected in growing or fully grown immature oocytes but were readily detectable in mature metaphase II oocytes of both wild-type and H10 -/- animals (Fig. 5A, closed arrow). These species were also synthesized in embryos. In one- and two-cell embryos, as in mature oocytes, the H1-like proteins migrated as a single species. Beginning at the four-cell stage, however, a doublet was detected (Fig. 5B, closed arrow). Typically, somatic H1 migrates as a doublet during SDS-polyacrylamide gel electrophoresis, with subtypes H1b, H1d, and H1e present in the slow-migrating upper band and H1a and H1c present in the fast-migrating lower band [6]. Furthermore, the species comigrating with somatic H1 were soluble in perchloric acid (data not shown), which is a characteristic property of histone H1. Taken together, these results may suggest that synthesis of a subset of the somatic H1 variants is up-regulated in oocytes beginning at meiotic maturation and that additional variants are synthesized in embryos beginning at the four-cell stage.
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In addition to species comigrating with somatic H1, several prominent acid-soluble proteins were synthesized by oocytes and embryos. Notably, synthesis of a species migrating at about 28 kD was detectable beginning at the four-cell stage (Fig. 5, open arrow). This species migrates near the expected position of histone H10 and is perchloric acid soluble. However, H10 -/- embryos synthesize a protein migrating at the same position, which indicates that the 
28-kD protein observed in wild-type oocytes and embryos is not H10.
Two other species migrating near the 34.5-kD marker were also prominent. The smaller of these (Fig. 5, A and B, asterisk) was weakly detectable in growing and maturing oocytes. It was prominent in one- and two-cell embryos of both wild-type and H10 -/- genotype but was not detected in more advanced embryos. Synthesis of the larger protein (Fig. 5A, arrowhead) was detectable in growing and maturing oocytes but was greatly reduced in embryos. Although this could correspond to the minor species migrating at Mr 37 kD detected by antibodies against the oocyte-type H1oo [28], we did not detect synthesis of a species corresponding to the major Mr 52-kD species detected by this antibody and the acid solubility of H1oo has not been reported. Thus, the identity of these proteins and their relationship to histones remain unknown.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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We also found that oocytes and embryos synthesize acid-soluble proteins that comigrate with the somatic histone H1 subtypes. These were not detectable in GV-stage oocytes but began to be synthesized during meiotic maturation, and synthesis continued following fertilization. Interestingly, the slow-migrating form of the typical somatic H1 doublet was not detected in oocytes but appeared at the four-cell stage. Previous studies of somatic H1 synthesis have reported apparently conflicting results. An antibody raised against rat somatic H1 did not detect the protein by immunoblotting or immunofluorescence in oocytes and one-cell embryos, although it was detectable at subsequent preimplantation stages [22, 34]. In contrast, an antibody raised against Xenopus H1 detected a very weak signal in GV oocytes, a modest signal in metaphase II oocytes, and a stronger signal in preimplantation embryos [27]. In addition, when oocytes or embryos were radiolabeled and the chromatin-associated proteins analyzed by electrophoresis, proteins comigrating with somatic H1 were synthesized by both oocytes and embryos [25].
These differences may be reconciled, however, as also noted by Adenot et al. [27]. The five somatic H1 subtypes migrate during SDS-PAGE as two bands—the upper (slow-migrating) band contains H1b, H1d, and H1e; the lower (fast-migrating) band contains H1a and H1c [6]. Among these, only the mRNA for H1c is known to become polyadenylated; consequently, this subtype tends to persist in nondividing cells. Ours and previous metabolic labeling studies [25] indicate that, in oocytes and one- and two-cell embryos, most somatic H1 synthesized is of the fast-migrating type. Comparison of the different antibodies used reveals that the anti-rat H1 preferentially recognizes the slow-migrating H1 species, which are not synthesized until the four-cell stage (Fig. 5), whereas the anti-Xenopus H1 antibody preferentially recognizes the fast-migrating H1 species [27], which are synthesized by maturing oocytes and one-cell embryos. Thus, it may be that oocytes and early embryos synthesize primarily the fast-migrating histone H1 subtypes, composed primarily of H1c. Beginning at the two- to four-cell stage and possibly linked to activation of embryonic transcription, the slow-migrating species, corresponding to H1b, H1d, and H1e, are also synthesized.
Taken together with the recent identification of H1oo, these data indicate that multiple H1 subtypes are present on oocyte and early embryonic chromatin. Their relative abundance, however, changes during this period. For example, the immunofluorescence staining pattern produced by the anti-H1oo antibody, which stains oocyte and one-cell nuclei but only the second polar body chromatin at subsequent stages [28], is the converse of the pattern produced by the anti-mouse H1 antibody, which stains embryonic nuclei beginning at the two-cell stage but does not stain polar body chromatin at any stage [22]. In addition, when somatic cell nuclei are transplanted into newly fertilized eggs, their somatic H1 rapidly becomes undetectable [27, 36], possibly because of its replacement by H1oo. Finally, histone H10, while present on oocyte chromatin, appears to decline during early embryogenesis. Based on these results, we propose that oocyte chromatin contains a mixture of H1oo, H1c, and H10. By analogy with nonmammalian species, H1oo may be the dominant linker histone at this stage. During the first two embryonic cell cycles, the complement changes so that by the four-cell stage the chromatin contains most or all of the five somatic subtypes as well as some H10. The apparently normal fertility of mice lacking H10 implies that the H10 present in oocytes and embryos does not serve an indispensable role. It may be that, as occurs in somatic cells [12], synthesis of the other linker histones is up-regulated in its absence. As multiple H1 subtypes are present on the chromatin, it is possible that they could be targeted to specific chromatin domains. H1 in somatic cells rapidly shuttles on and off chromatin [37, 38], however, and the loss of detectable somatic H1 from transplanted somatic nuclei suggests that shuttling occurs in oocyte and embryonic chromatin also. Thus, if the different H1 subtypes associate with distinct chromatin domains, a mechanism must exist to assure specificity of binding.
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FOOTNOTES |
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2 Correspondence: H.J. Clarke, Room F3.50, Royal Victoria Hospital, 687 av. des Pins O., Montréal, Québec, Canada H3A 1A1. FAX: 514 8431662; hugh.clarke{at}muhc.mcgill.ca
3 Current address: University of California at Santa Barbara, Santa Barbara, CA
4 Current address: Department of Biochemistry and Biophysics, Tehran University, Tehran, Iran
Received: 14 October 2002.
First decision: 29 October 2002.
Accepted: 18 November 2002.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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