The transition from a late 1-cell mouse embryo to a 4-cell embryo, the period when zygotic gene expression begins, is accompanied by an increasing ability to repress the activities of promoters and replication origins. Since this repression can be relieved by either butyrate or enhancers, it appears to be mediated through chromatin structure. Here we identify changes in the synthesis and modification of chromatin bound histones that are consistent with this hypothesis. Oocytes, which can repress promoter activity, synthesized a full complement of histones, and histone synthesis up to the early 2-cell stage originated from mRNA inherited from the oocyte. However, while histones H3 and H4 continued to be synthesized in early 1-cell embryos, synthesis of histones H2A, H2B and H1 (proteins required for chromatin condensation) was delayed until the late 1-cell stage, reaching their maximum rate in early 2-cell embryos. Moreover, histone H4 in both 1-cell and 2-cell embryos was pre-dominantly diacetylated (a modification that facilitates transcription). Deacetylation towards the unacetylated and monoacetylated H4 population in fibroblasts began at the late 2-cell to 4-cell stage. Arresting development at the beginning of S-phase in 1-cell embryos prevented both the appearance of chromatin-mediated repression of transcription in paternal pronuclei and synthesis of new histones. These changes correlated with the establishment of chromatin-mediated repression during formation of a 2-cell embryo, and the increase in repression from the 2-cell to 4-cell stage as linker histone H1 accumulates and core histones are deacetylated.
During mouse development, formation of a 2-cell embryo marks the transition from maternal to zygotic gene dependence (Fig. 1; reviewed by Schultz, 1993; Nothias et al., 1995; Christians et al., 1995). Transcription stops when oocytes undergo meiotic maturation to form unfertilized eggs. Moreover, degradation of maternal mRNA begins and is almost completed by the 2-cell stage, although synthesis of proteins from maternally inherited mRNA can still be detected at the 8-cell stage. Fertilization triggers completion of meiosis and formation of a 1-cell embryo with two haploid pronuclei, one from each parent, but the onset of zygotic gene expression is a time dependent event that is delayed for about 24 hours, thereby beginning after formation of a 2-cell embryo. Transcription of zygotic genes can begin in late 1-cell embryos, but nascent transcripts produced in these embryos are not translated and therefore not productive; translation is coupled to transcription only during the 2-cell stage (Nothias et al., 1996). This delay in zygotic gene expression provides a window of opportunity for remodeling parental chromosomes without accidentally and prematurely expressing their genes. In fact, one or more factors capable of repressing DNA transcription and replication are absent from 1-cell embryos during this remodeling period, and are then produced again prior to the productive transcription of zygotic genes.
Injection of plasmid-encoded reporter genes into the nuclei of oocytes and early embryos has revealed the presence of transacting factors that can repress the activities of promoters (Dooley et al., 1989; Martínez-Salas et al., 1989; Wiekowski et al., 1991, 1993; Majumder et al., 1993; Henery et al., 1995) and replication origins (Martínez-Salas et al., 1988) from 20 to >500-fold. While maternal nuclei in both oocytes and 1-cell embryos exhibit this repression (Martínez-Salas et al., 1989; Wiekowski et al., 1991, 1993), the paternal pronucleus exhibits repression only when 1-cell embryos develop beyond S-phase. Application of nuclear transplantation (Henery et al., 1995) and nuclear alteration (Wiekowski et al., 1993) techniques revealed that this ‘repressor’ is absent from the cytoplasm of early 1-cell embryos (not simply excluded from their paternal pronucleus), and is produced in the cytoplasm sometime between S-phase in a 1-cell embryo and formation of a 2-cell embryo where it can operate within any nucleus, regardless of its parental origin or ploidy. The potency of this repression then increases as development proceeds to the 4-cell stage.
Repression of promoter activity at the beginning of mouse development appears to be mediated through chromatin structure, because it can be relieved either by treating the cells with butyrate or, in cleavage stage embryos (≤2 cells), by linking the promoter or replication origin to an embryo-responsive enhancer (Wiekowski et al., 1993; Majumder et al., 1993). Butyrate and other inhibitors of histone deacetylase increase the fraction of hyperacetylated core histones (Yoshida et al., 1995) and thereby stimulate transcription from specific genes (Seigneurin et al., 1995; Yoshida et al., 1995; Arts et al., 1995), consistent with the fact that transcriptionally active eukaryotic genes are generally associated with acetylated core histones (Tazi and Bird, 1990; Hebbes et al., 1988, 1994; Jeppesen and Turner, 1993; Sommerville et al., 1993). In mouse embryos, the fact that butyrate can stimulate promoter activity in the maternal pronucleus (‘repressed’) but not in the paternal pronucleus (‘unrepressed’) of a 1-cell embryo shows that this stimulation results from changes in chromatin structure on the injected plasmid rather than from increased synthesis of transcription factors, a change that would be expected to affect both pronuclei. In fact, inhibition of histone deacetylase simply stimulates synthesis of transcription dependent proteins at the onset of zygotic gene expression without changing the overall pattern of protein synthesis (Wiekowski et al., 1993; Worrad et al., 1995). Enhancers can substitute for butyrate in stimulation of promoter activity only after formation of a 2-cell embryo, because prior to this event, an enhancer specific coactivator activity is missing (Majumder et al., 1997). In vitro, enhancers do not stimulate transcription unless the DNA substrate is organized into chromatin (reviewed by Majumder et al., 1993; Paranjape et al., 1994). In vivo, enhancers have little, if any, effect on promoters injected into cleavage stage embryos treated with butyrate (Wiekowski et al., 1993; Majumder et al., 1993). Therefore, the primary role of enhancers is not simply to provide additional transcription factors to facilitate formation of an active initiation complex, but to relieve the repression of weak promoters by chromatin. Here we report changes in the synthesis and modification of chromatin bound histones at the beginning of mouse development that support the conclusion that formation of a 2-cell mouse embryo is accompanied by changes in chromatin structure that can repress the activity of promoters and replication origins (summarized in Fig. 1). Histone H4 is diacetylated in 1-cell embryos and then progressively deacetylated during early cleavage stages, while synthesis of histones H2A, H2B and H1 does not begin until the late 1-cell stage and relies on maternally inherited mRNA. Somatic histone H1 does not appear until the 4-cell stage (Clarke et al., 1992). These events have parallels in frog development where histone H4 also is stored in eggs as a diacetylated form and then progressively deacetylated after zygotic gene expression begins at the blastula stage (Dimitrov et al., 1993). Histone deacetylase inhibitors can then induce expression of specific genes (Almouzni et al., 1994). The linker histone changes from the maternal histone H1 variant (B4) at the mid-blastula transition to the somatic histone H1 variant at the end of gastrulation, resulting in specific repression of oocyte 5S RNA gene expression (Bouvet et al., 1994; Kandolf, 1994), and the initial activation of H1 synthesis occurs entirely by mobilizing maternal transcripts that then disappear by the early gastrula stage (Woodland et al., 1979).
MATERIALS AND METHODS
Radiolabeling of mouse histones
Oocytes and embryos were cultured as previously described (DePamphilis et al., 1988; Wiekowski et al., 1991, 1993). Mouse oocytes were isolated from 14- to 16-day-old [C57BL/6J × SJL/J]F1 females. Mouse 1-cell embryos were isolated from 8- to 10-week-old [C57BL/6J × SJL/J]F1 females 18 hours after injection of human chorionic gonadotropin (post-hCG) and mating with [C57BL/6J × SJL/J]F1 males. Two-cell embryos were isolated from pregnant females at 38 hours post-hCG. Nascent histones in mouse oocytes and preimplantation embryos were characterized as described in Wiekowski and DePamphilis (1993). In general, nascent histones were labeled with 3H-arginine and 3H-lysine before lysing the cells in the presence of butyrate to inhibit histone deacetylase, and PMSF and bisulfite to inhibit proteases. A similar cell lysate was prepared from mouse fibroblasts and combined with the oocyte or embryo lysate in order to provide an excess of unlabeled (carrier) histones. The nuclei were then isolated, disrupted in the presence of EDTA, low salt, butyrate and protease inhibitors to release chromatin, and the chromatin was washed and then sonicated before extracting histones into acid. The acid soluble fraction was combined with acetone to precipitate histones which were stored at −80°C for not more than 1 week before analysis. Additional details are provided in figure legends. The mobilities and relative amounts of various proteins exhibited some variability among the three to twenty examples obtained for each experiment. Representative patterns are shown in each figure.
Unless otherwise indicated, 1-cell embryos were labeled from 20 to 26 hours post-hCG, and 2-cell embryos from 40 to 46 hours post-hCG. Oocytes were maintained in 100 μg/ml dibutryl-cAMP to arrest them in prophase of meiosis I and labeled for 24 hours. Proliferating 3T3 mouse fibroblasts were cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% calf serum and labeled for 24 hours.
Where indicated, 2.5 mM or 10 mM sodium butyrate was added to the medium 4 hours before addition of labeled amino acids.
Identification of mouse histones
Mouse histones were identified by three criteria: their solubility in acid, their migration during electrophoresis in poly-acrylamide gels containing sodium dodecyl sulfate (SDS), and their migration in polyacrylamide gels containing Triton X-100/acetic acid/urea (TAU). Proteins such as histones with a net positive charge are soluble in acid. Therefore, to establish the identity of mouse histones, proteins were extracted from mouse fibroblasts into 0.4 N sulfuric acid and then subjected to SDS-gel electrophoresis in parallel with purified calf thymus histones (Fig. 2A). SDS-gel electrophoresis separates proteins primarily according to their molecular masses, although proteins with a net positive charge (e.g. histones) sometimes migrate anomalously (von Holt et al., 1989). The migration rates of mouse histones (H4>H2A>H2B>H3>H1) were indistinguishable from those of calf histones, and were consistent with previous studies (Bonner et al., 1980). Based on amino acid composition, core histones H2A, H2B, H3 and H4 are between 11 and 14.5 kDa (van Holt et al., 1989) but migrated as though they were 13 to 17 kDa proteins. Histone H1 is 22 kDa, but migrated as though it was a 34 to 38 kDa protein.
TAU-gel electrophoresis fractionates histones according to their charge as well as their molecular mass, thereby allowing identification of post-translational modifications such as acetylation (Lennox and Cohen, 1989). Since the relative migrations of histones during TAU-gel electrophoresis are strongly influenced by the composition of the gel (Zweidler, 1978), optimal conditions were developed for fractionation of mouse histones (Fig. 2B). Under these conditions, the migration pattern of mouse histones (H4>H2B>H3>H1>H2A) was consistent with previous studies (Bonner et al., 1980), and mouse histones co-migrated with their corresponding calf histones.
Acetylation of lysine residues decreases the mobility of histones during TAU-gel electrophoresis, because each acetylation event neutralizes one positive charge on the protein (Ruis-Carrillo et al., 1975). For example, histones extracted from mouse fibroblasts treated with butyrate were separated during TAU-gel electrophoresis into five bands representing histone H4 with 0 to 4 acetyl groups (Fig. 2B). In contrast, histones from butyrate treated cells migrated identically to those from untreated cells during SDS-gel electrophoresis (Fig. 2A). The increased amounts of proteins observed with butyrate treated cells reflected increased protein synthesis (Wiekowski et al., 1993).
Staggered synthesis of core histones in 1-cell embryos
The relatively small size (2.5×10−4 the volume of a frog egg) and low number (∼25 zygotes per pregnant female) of embryos available restricted analysis of histone composition to nascent proteins that could be labeled by culturing oocytes or embryos in the presence of 3H-lysine and 3H-arginine. Since these two amino acids comprise 18% to 21% of the amino acids in each of the five major histones (von Holt et al., 1989), the amount of radiolabel per mole of each histone was nearly equivalent. Histones were then acid-extracted from purified chromatin preparations in the presence of an excess of unlabeled mouse proteins prepared from 3T3 fibroblasts. Gels were stained first with Coomassie Blue to visualize total proteins and then subjected to fluorography to visualize 3H-labeled proteins. Nascent 3H-histones were identified by superimposing the resulting autoradiogram over the stained gel. In addition, 3H-histones from mouse 3T3 fibroblasts were run in parallel lanes. Fractionation of nascent core histones by SDS-gel electrophoresis revealed that the pattern of chromatin bound nascent core histones changed from oocytes to 1-cell embryos to 2-cell embryos (Fig. 3A). Core histones synthesized in 2-cell mouse embryos closely resembled those synthesized in mouse fibro-blasts. However, the fraction of nascent H2A and H2B was over-represented in oocytes and under-represented in 1-cell embryos, suggesting that the pattern of core histone synthesis changed during development of fertilized eggs to 2-cell embryos.
Development of mouse embryos can be synchronized by triggering ovulation through injection of human chorionic gonadotrophin (hCG). Fertilization in vivo occurs ∼12 hours post-hCG, paternal and maternal pronuclei appear ∼18 hours post-hCG, DNA replication begins ∼23 hours post-hCG followed by the first mitosis and formation of a 2-cell embryo at ∼32 hours post-hCG (Fig. 1). In early 1-cell embryos (19-26 hours post-hCG), the amount of nascent histones H3 and H4 was greater than the amount of nascent histones H2A and H2B (Fig. 3B). In 2-cell embryos (40-47 hours post-hCG), the opposite was true: synthesis of H2A and H2B was greater than synthesis of H3 and H4. Occasionally the amount of nascent H2B in mouse fibroblasts was over represented, which may reflect changes in histone synthesis as a function of the fraction of cells in S-phase (Osley, 1991).
TAU-gel electrophoresis of these histone preparations confirmed that early 1-cell embryos synthesized histones H3 and H4, but not H2B (Fig. 4A). Synthesis of histone H2B occurred concurrent with formation of a 2-cell embryo (∼32 hours post-hCG). Histone H2A could not be distinguished from other basic 3H-proteins under these TAU-gel conditions. The 1-cell stage was also characterized by synthesis of two unidentified acid-soluble proteins (X and Y, Fig. 4A).
Synthesis of histone H1 begins in late 1-cell embryos
Purified histone H1 from calf thymus migrated as two prominent bands during SDS-gel electrophoresis (Fig. 2A). The same bands were also identified in the acid soluble proteins extracted from mouse 3T3 fibroblasts (Figs 2A, 3A,B), and they were the only acid-soluble proteins that cross-reacted with a monoclonal antibody specific for histone H1 in a western blot (data not shown). These two histone H1 variants were synthesized in mouse oocytes (Fig. 3A). However, following fertilization, synthesis of histone H1 was not evident until formation of a 2-cell embryo, and then only the faster migrating H1 variant was observed (Fig. 3A). When 1-cell embryos were arrested at the beginning of their S-phase with aphidicolin (a specific inhibitor of replicative DNA poly-merases), histone H1 synthesis began between 26 and 40 hours post-hCG, the time when 1-cell embryos normally underwent cleavage into 2-cell embryos (Fig. 3B). This conclusion was confirmed by fractionating nascent histones by TAU-gel electrophoresis. A prominent band of nascent histone H1 appeared between 26 and 30 hours post-hCG (Fig. 4A).
Acid-soluble proteins from early 1-cell embryos (19-26 hours post-hCG) generally contained one or two faint bands that migrated in the vicinity of histone H1 during SDS-gel electrophoresis (Fig. 3) and 8 distinct bands that migrated in the vicinity of histone H1 during TAU-gel electrophoresis (Fig. 4). Thus, early 1-cell embryos either failed to synthesize histone H1, or synthesized their own histone H1 variants. To distinguish between these two possibilities, nascent 3H-histones in embryos from the 1-cell to 8-cell stage were extracted with perchloric acid and then fractionated by TAU-gel electrophoresis in parallel with 3H-histones that had been extracted with sulfuric acid from 2-cell embryos. While all histone subtypes are soluble in sulfuric acid, histone H1 is selectively solubilized in perchloric acid (Johns, 1964; Ohsumi and Katagiri, 1991). The efficiency of this extraction procedure was monitored by staining the gels with Coomassie Blue to visualize total protein (Fig. 5A). Comparison of these data with the results of fluorography of the same gel revealed that synthesis of 3H-histone H1 began in late 1-cell embryos (24-29 hours post-hCG), reaching its maximum rate in early 2-cell embryos (36-41 hours post-hCG). The rate of histone H1 synthesis was reduced in late 2-cell embryos (43-48 hours post-hCG), and then restored again in 4-cell and 8-cell embryos (Figs 5B and 7B). This change in the rate of histone H1 synthesis may reflect its transition from maternal to zygotic gene dependence.
Histone synthesis in fertilized eggs is independent of cell cleavage, DNA replication and DNA transcription
In most somatic cells, histone synthesis is restricted to S-phase, that period during cell proliferation when DNA replication occurs. When DNA replication is interrupted, histone mRNA rapidly disappears from the cytoplasm (Osley, 1991). To determine whether histone synthesis in fertilized mouse eggs was also dependent on DNA replication, 1-cell embryos were cultured in the presence of aphidicolin, a specific inhibitor of the replicative DNA polymerases.
The pattern of histone synthesis was unaffected by aphidicolin. The relative synthesis of core histones in early 1-cell embryos (19-26 hours post-hCG) was unchanged (Fig. 3). Moreover, the pattern of core histone synthesis between 26 and 40 hours post-hCG changed to the pattern observed in 2-cell embryos, even though aphidicolin prevented these cells from undergoing DNA replication and cleavage into 2-cell embryos. Histone H1 synthesis began between 26 and 30 hours post-hCG in both developing (Fig. 4A) and S-phase arrested 1-cell embryos (Fig. 4B). Similarly, synthesis of histone H2B was not evident until the time when 1-cell embryos normally became 2-cell embryos, regardless of the presence or absence of aphidicolin; by ∼30 hours post-hCG, the pattern of nascent core histones in arrested 1-cell embryos was indistinguishable from that in 2-cell embryos (Fig. 4). Histone synthesis was easier to observe in S-phase arrested 1-cell embryos, because they underwent a reduction in overall protein synthesis that begins at 40 hours post-hCG (Wiekowski et al., 1991). These data demonstrate that regulation of histone synthesis in fertilized mouse eggs is independent of DNA replication and cell cleavage. The same was true for protein bands X and Y that were unique to 1-cell embryos.
Addition of α-amanitin, a specific inhibitor of RNA polymerases II and III, to developing 1-cell embryos did not affect the pattern of core histone synthesis (Fig. 4C), although the same concentration of α-amanitin prevented synthesis of the transcription requiring complex of proteins that marks the onset of zygotic gene expression (Nothias et al., 1996). Therefore, synthesis of histones in 1-cell and early 2-cell embryos must be directed by maternal mRNA rather than zygotic mRNA. This would account for the apparent independence of histone synthesis from cell cycle events in fertilized eggs. The same appeared true for all acid-soluble proteins, included bands X and Y that were unique to 1-cell embryos.
Core histones are acetylated in 1-cell and 2-cell embryos
Acetylated core histones are frequently associated with transcriptionally active genes, and the extent of core histone acetylation is reflected by the acetylated state of chromatin bound histone H4, the core histone most easily characterized. TAU-gel electrophoresis fractionated histone H4 into five distinct species containing from zero to four acetyl groups (H4, H4Ac1, H4Ac2, etc.). In order to provide internal standards of acetylated histone H4, a separate group of oocytes and embryos were cultured in the presence of 2.5 mM butyrate (an inhibitor of histone deacetylase) before and during the radiolabeling period, conditions that were previously shown to produce the maximum stimulation of plasmid-encoded promoter activity (Wiekowski et al., 1993). Thus, individual acetylated forms of nascent histone H4 were identified by comparing the 3H-histone H4 pattern in untreated cells with that in butyrate treated cells and by superimposition of fluorograms onto Coomassie Blue stained gels in order to determine the position of 3H-histones with those of the unlabeled carrier histones provided by mouse fibroblasts.
Mouse fibroblasts provided convenient standards for comparison with oocytes and embryos. The bulk of both nascent (3H-labeled protein) and total (Coomassie Blue stained protein) histone H4 in mouse fibroblasts was either H4 or H4Ac1 (Fig. 6), and treatment of fibroblasts with butyrate produced an equal distribution of all five forms of acetylated histone H4 (0-4, Fig. 6). Mouse oocytes were similar to mouse fibroblasts in that oocyte chromatin contained histone H4, H4Ac1 and H4Ac2, but differed from fibroblasts in that butyrate converted all oocyte chromatin bound histone H4 to H4Ac2-4. However, after fertilization, all of the chromatin bound histone H4 synthesized in either 1-cell or 2-cell embryos contained at least 2 acetyl groups, and butyrate treatment produced significant amounts of H4Ac3 and H4Ac4. Therefore, histone H4 synthesized in 1-cell and 2-cell embryos was hyperacetylated relative to histone H4 synthesized in oocytes and fibroblasts. A reduction in histone H4 acetylation began in 4-cell and 8-cell embryos. These changes were quantified by determining the fraction of 3H-histone H4 at various stages in development that contained two or more acetyl groups (Fig. 7B).
To determine whether or not the acetylated state of nascent histone H4 reflected changes in the overall acetylation pattern of histone H4, cells were stained first with a rabbit antiserum directed against acetylated histone H4 and then with FITC-conjugated anti-rabbit IgG to visualize its intracellular localization. Quantifying the relative amounts of fluorescence/μm2 observed in each type of nucleus (Fig. 8) confirmed conclusions derived from analysis of nascent histone H4 (Fig. 7). The nuclear concentration of acetylated histone H4 was essentially the same in 1-cell and 2-cell embryos, and its amount and distribution in paternal and maternal pronuclei in the same 1-cell embryo as well as the remaining polar body were indistinguishable. The concentration of acetylated histone H4 in 1-cell and 2-cell embryos was 2.5-fold greater than in oocytes or fibroblasts. Treatment with butyrate increased the nuclear con-centration of acetylated histone H4 from 1.7-fold to 7.4-fold, and resulted in a uniform distribution throughout the nucleus in each cell type. Moreover, butyrate increased the nuclear concentration of acetylated histone H4 in oocytes and 2-cell embryos about 2.5 times more effectively than in 1-cell embryos. The ability of butyrate to stimulate the activity of a plasmid-encoded promoter (HSV thymidine kinase promoter) injected into the nuclei of oocytes or embryos, or electroporated into fibroblasts paralleled its ability to increase the nuclear concentration of acetylated histone H4 (Fig. 8B), consistent with the hypothesis that butyrate stimulates promoter activity in mouse oocytes and preimplantation embryos by reducing chromatin mediated repression through increased acetylation of core histones.
Aphidicolin suppresses expression of repressor activity
Previous studies (see Introduction) have shown that repressor is absent from the paternal pronucleus and cytoplasm of S-phase arrested 1-cell embryos. The maternal pronucleus exhibits some repressor activity that presumably is inherited from the oocyte nucleus. Since repressor is present in both nuclei and cytoplasm of 2-cell embryos, arresting DNA synthesis in 1-cell embryos appears to prevent expression of repressor activity. To test this hypothesis, plasmids encoding the firefly luciferase gene were injected into the paternal pronucleus of 1-cell mouse embryos cultured either in the absence or presence of aphidicolin, and the amount of luciferase mRNA produced was measured as a function of time elapsed. The luciferase gene was linked either to the herpes simplex virus thymidine kinase (tk) promoter alone (ptkluc) or to the tk promoter with the F101 polyomavirus enhancer 600 bp upstream (pF101tkluc). Luciferase gene expression is dependent on a linked promoter, and the F101 enhancer produces the highest levels of luciferase enzyme activity in mouse cleavage stage embryos (Nothias et al., 1995; Majumder and DePamphilis, 1995). A quantitative assay for luciferase mRNA (Nothias et al., 1996) revealed directly the effect of embryo development on luciferase gene transcription. Controls were carried out to confirm that deletion of the tk promoter eliminated luciferase gene transcription, and that the F101 enhancer stimulated tk promoter activity when plasmids were injected into 2-cell embryos (Nothias et al., 1996, and data not shown), consistent with previous studies in which luciferase enzyme activity was measured (Henery et al., 1995).
In the absence of aphidicolin (Fig. 9A), most (∼90%) of the injected 1-cell embryos developed into 2-cell embryos and luciferase gene transcription was strongly repressed, regardless of the presence or absence of the F101 enhancer which functions efficiently in 2-cell embryos. In contrast, both ptkluc and pF101tkluc were actively transcribed in 1-cell embryos that did not develop into 2-cell embryos. These results were consistent with previous studies in which luciferase gene expression was monitored by luciferase enzyme activity (Henery et al., 1995), and were interpreted as ‘irreversible’ repression that occurred before enhancer activation factors were present in sufficient quantity to prevent repression. Since plasmid DNA does not replicate in these embryos, the repressed state cannot be reprogrammed at a later time.
In the presence of aphidicolin (Fig. 9B), most of the injected 1-cell embryos remained as 1-cell embryos. These S-phase arrested 1-cell embryos also actively transcribed the injected luciferase gene, but they did so about 20 times more efficiently than 1-cell embryos in the absence of aphidicolin. Moreover, the small fraction of 2-cell embryos that formed in the presence of aphidicolin (<10%) also transcribed the injected gene and to the same extent as did the S-phase arrested 1-cell embryos, regardless of the presence or absence of the F101 enhancer.
These data are consistent with the hypothesis that aphidicolin prevented expression of repressor activity and thus allowed transcription, regardless of whether or not cell cleavage occurred.
Aphidicolin suppresses histone synthesis
Experiments described above reveal that synthesis of histones H1, H2A and H2B begins in late 1-cell embryos, and that deacetylation of core histones begins in late 2-cell or 4-cell embryos (Fig. 6), suggesting that one or both events contributes to appearance of repressor activity upon formation of 2-cell and 4-cell embryos (Fig. 7). Moreover, repressor was not produced when 1-cell embryos were cultured in the presence of aphidicolin, regardless of whether or not they underwent cleavage, suggesting that aphidicolin may affect histone synthesis or modification. However, no changes were detected in either the types or relative amounts of histones (or other basic proteins) synthesized in 1-cell embryos cultured in the presence of aphidicolin (Figs 3, 4). What did change in the presence of aphidicolin was the total amount of protein synthesis. The translational capacity of 1-cell embryos cultured in aphidicolin rapidly decreased between 35 and 40 hours post-hCG (Fig. 10A), that time when early 2-cell embryos had formed and zygotic gene expression had begun (Fig. 1). By the time development would have reached the 4-cell stage, protein synthesis was effectively stopped.
The effect of aphidicolin on total protein synthesis was also evident with histone synthesis. For example, the amount of histone H1 synthesized in 1-cell embryos that developed into early 2-cell embryos in the absence of aphidicolin was ∼4-fold more than the amount synthesized in S-phase arrested 1-cell embryos cultured for the same length of time in the presence of aphidicolin (Fig. 10B and C). This H1 synthesis resulted from maternally inherited mRNA, since it was not inhibited by α-amanitin (Fig. 10C). Increased H1 synthesis in the presence of α-amanitin presumably resulted from increased stability of histone mRNA.
Chromatin structure can repress transcription, and the extent of this repression depends on at least three parameters: the presence of a complete histone octamer, low acetylation of core histones, and the presence of a linker histone such as H1 (Paranjape et al., 1994). Although H3 and H4 alone can organize DNA into nucleosome-like structures, the DNA remains accessible to transcription factors. Addition of H2A and H2B begins to mask the DNA from transcription components such as RNA polymerase II (Baer and Rhodes, 1983) and TFIIIA (Hayes and Wolffe, 1992), but addition of histone H1, which requires the prior addition of H2A and H2B (Hayes et al., 1994), condenses chromatin into a 30 nm fiber that represses transcription (Bouvet et al., 1994; Juan et al., 1994; Kandolf, 1994; Paranjape et al., 1994; O’Neill et al., 1995). Both acetylation of core histones (Lee et al., 1993; Vettese-Dadey et al., 1996) and removal of histone H1 (Juan et al., 1994; Ura et al., 1995) can facilitate the ability of some transcription factors to bind to chromatin, and together these two parameters strongly facilitate transcription (Ura et al., 1997).
The transition from a late 1-cell mouse embryo to a 4-cell embryo, the period wherein zygotic gene expression begins, is accompanied by an increasing ability to repress the activities of promoters and replication origins (see Introduction). This repression can be relieved by either butyrate or enhancers, suggesting that it is mediated through chromatin structure. In fact, changes in the synthesis and modification of chromatin bound histones at the beginning of mouse development (summarized in Fig. 1) are consistent with this hypothesis: repression was greatest when chromatin contained all five newly synthesized histones and core histones were minimally acetylated (growing oocytes and ≥late 2-cell embryos).
Histone synthesis in mouse preimplantation embryos
All five major histones were synthesized in mouse oocytes, but only histones H3 and H4 were synthesized in early 1-cell embryos. Synthesis of histones H2A, H2B and H1 did not resume until the late 1-cell/early 2-cell stage. Histone synthesis in 1-cell and early 2-cell mouse embryos was independent of both DNA replication, DNA transcription and cell cleavage (Figs 3, 4). Therefore, these histones were translated from mRNA inherited from the oocyte. Histone H1 synthesized in 1-cell and 2-cell embryos was identified by its mobility during SDS and TAU gel electrophoresis that was indistinguishable from histone H1 isolated from mouse fibroblasts (Figs 3, 4), and by the fact that histone H1 could be selectively extracted with perchloric acid (Fig. 5). Intriguingly, this early form of histone H1 was not detected using anti-H1 antibodies until formation of a 4-cell embryo during either mouse (Clarke et al., 1992) or bovine (Smith et al., 1995) development. Since these antibodies were made against somatic cell histone H1, these results suggest the existence of both maternal and zygotic histone H1 variants. In other studies, addition of α-amanitin to late 2-cell embryos (G2-phase) prevented expression of somatic histone H1 at the 4-cell stage (Clarke et al., 1992), suggesting that synthesis of somatic histone H1 begins during the second phase of zygotic gene activation (Nothias et al., 1995). Moreover, synthesis of somatic histone H1 was sensitive to inhibition of DNA replication at the 4-cell stage, consistent with coupling of somatic histone gene expression to DNA replication during the third cell proliferation cycle (Clarke et al., 1992). These results reveal that histone H1 is first expressed from maternally inherited mRNA in the late 1-cell and early 2-cell stages of mouse development, followed by a transition to expression from zygotic genes in the late 2-cell to 4-cell stages. Consistent with a transition from maternal to zygotic histone gene expression, mRNAs for histones H2A, H2B and H3 have been identified in fertilized mouse eggs, and the levels of these mRNAs have been observed to decrease ∼10-fold by the mid 2-cell stage (Giebelhaus et al., 1983; Graves et al., 1985). In somatic mammalian cells (Osley, 1991), histone synthesis is regulated by activating translation of histone mRNA present in G1-phase by the onset of S-phase, by coupling synthesis of new histone mRNA to DNA replication, and by coupling the stability of histone mRNA to DNA replication (histone mRNA is rapidly degraded after completion of S-phase). Therefore, translation of maternally inherited histone mRNA may be activated by S-phase in 1-cell embryos, while transcription of histone genes in the zygote may be coupled to S-phase in the 2-cell embryo. During the long G2-period in 2-cell embryos, maternally inherited histone mRNA will be degraded, and the 4-cell stage will be dependent primarily on transcription of zygotic histone genes. This would account for the observation that histone synthesis occurs independently of DNA replication in 2-cell embryos (Figs 3, 4; Kaye and Church, 1983), but that these two events are coupled by the blastocyst stage (Kaye and Church, 1983).
Establishment of chromatin mediated repression at the beginning of mouse development
Virtually all of the chromatin bound nascent histone H4 in fertilized mouse eggs and 2-cell embryos was diacetylated (Figs 6, 7), and immunofluorescence staining with antibodies directed against acetylated histone H4 revealed that the concentrations of total nuclear H4Ac2-4 in 1-cell and 2-cell embryos was at least twice those in oocytes or fibroblasts (Fig. 8). Moreover, synthesis of histones H2A, H2B and H1 did not begin until the late 1-cell/early 2-cell stage and somatic histone H1 was not detected by immunofluorescence staining until the 4-cell stage in either mouse or bovine embryos (Clarke et al., 1992; Smith et al., 1995). Since hyperacetylated core histones can faciliate assembly of nucleosomes onto nonreplicating DNA (Cotten and Chalkley, 1985), and H4Ac2 is used specifically in the assembly of nucleosomes at replication forks (Sobel et al., 1995; Kaufman et al., 1995), fertilized eggs are poised to both remodel and replicate parental chromosomes. However, DNA injection experiments (see Introduction) have shown that chromatin assembled under these conditions (early (G1-phase) 1-cell embryos) exhibit reduced levels of chromatin mediated repression, consistent with the combined effects of the presence of H4Ac2 and the absence of histone H1 on transcription of chromatin assembled in vitro (Ura et al., 1997). Therefore, prior to zygotic gene expression, newly assembled chromatin must be modified in order to repress transcription so that genes can be activated selectively during the subsequent phases of development.
DNA injection experiments reveal that repression is reestablished following S-phase in 1-cell embryos and increases as development proceeds to the 4-cell stage (see Introduction). This appears to be accomplished in two stages. First, the appearance of chromatin bound nascent histones H2A, H2B and H1 began just prior to formation of a 2-cell embryo. Thus, although nuclei in both 1-cell and 2-cell embryos contained similar concentrations of H4Ac2-4 (Fig. 8), repression of transcription will be brought on by the increasing presence of histone H1 (Ura et al., 1997). Second, the level of acetylation in chromatin bound histone H4 began to decrease in 4-cell embryos and was nearly complete in 8-cell embryos (Fig. 6). This reduction in core histone acetylation should facilitate establishment of a repressive chromatin structure. In fact, increasing the concentration of H4Ac2-4 by inhibition of histone deacetylase with butyrate, trichostatin A or trapoxin also increases the amount of hyperacetylated histone H4 in 2-cell to 8-cell embryos (Fig. 8; Worrad et al., 1995; Thompson et al., 1995), consistent with the effect of these inhibitors on stimulating promoter activity in either injected plasmids (Wiekowski et al., 1993; Majumder et al., 1993) or transgenes (Thompson et al., 1995). Stimulation was greatest in 4-cell embryos. When the rate of protein synthesis, including histone synthesis, was suppressed by arresting 1-cell embryos at the beginning of S-phase (Fig. 10), their ability to repress transcription from an injected gene did not materialize, even when some embryos slipped through their check point controls and formed a 2-cell embryo (Fig. 9). Therefore, protein synthesis is required in late 1-cell embryos in order to establish chromatin mediated (i.e. butyrate sensitive) transcription repression in cleavage stage embryos. The fact that synthesis of histones H2A, H2B and H1 did not begin until the late 1-cell/early 2-cell stage was consistent with their role in chromatin-mediated repression, although other proteins also may be involved. Thus, the extent of transcription repression (Fig. 7A) is increased as the amount of linker histone is increased (Fig. 7C) and the amount of diacetylated core histone is decreased (Fig. 7B; Fig. 8, top). Together, these changes in histone synthesis and modification are consistent with a developmental acquisition of chromatin mediated repression which, in the mouse and perhaps other mammals as well, begins in the late 1-cell embryo and is completed by the 4-cell stage.
One puzzle is why the maternal pronucleus in an S-phase arrested 1-cell embryo can repress transcription from an injected promoter, but not the paternal pronucleus (Wiekowski et al., 1993; Nothias et al., 1996). This difference appears to be due to chromatin-mediated repression because butyrate can stimulate promoter activity in the maternal pronucleus to the level observed in the paternal pronucleus, but butyrate does not stimulate promoter activity in the paternal pronucleus (Wiekowski et al., 1993). Surprisingly, both pronuclei contain similar concentrations of H4Ac2-4 (Fig. 8). Therefore, the maternal pronucleus must inherit one or more additional proteins from the oocyte, such as histones H2A, H2B and H1, that allow repression to occur. Since the paternal pronucleus is derived from the sperm, it must be provided with histones that are either inherited from the oocyte or synthesized after fertilization from maternal mRNA. Based on synthesis of histone H4, sufficient core histones are synthesized in oocytes to support two or three rounds of DNA replication (Wassarman and Mrozak, 1981). However, this free histone pool may be sequestered within the maternal nucleus by chaperone proteins used in chromatin assembly, as appears to be the case with the much larger histone pool in amphibian oocytes (Patterton and Wolffe, 1996). Thus, following meiotic maturation and fertilization, the fraction and composition of the maternal histone pool available to the paternal pronucleus remains to be determined. Chromatin assembled in the two pronuclei could differ in composition and thereby provide the fertilized egg with an opportunity for genomic imprinting by masking sequences in one genome but not in the other.
Monoclonal antibodies directed against mammalian histone H1 were provided by John Brenneman (University of California, Davis) and Missag Parseghian (University of California, Irvine). Anti-serum against acetylated histone H4 was provided by Bryan Turner (University of Birmingham Medical School, Birmingham, UK).