Compensatory functions of histone deacetylase 1 (HDAC1) and HDAC2 regulate transcription and apoptosis during mouse oocyte development

Abstract

Dramatic changes in chromatin structure and histone modification occur during oocyte growth, as well as a global cessation of transcription. The role of histone modifications in these processes is poorly understood. We report the effect of conditionally deleting Hdac1 and Hdac2 on oocyte development. Deleting either gene has little or no effect on oocyte development, whereas deleting both genes results in follicle development arrest at the secondary follicle stage. This developmental arrest is accompanied by substantial perturbation of the transcriptome and a global reduction in transcription even though histone acetylation is markedly increased. There is no apparent change in histone repressive marks, but there is a pronounced decrease in histone H3K4 methylation, an activating mark. The decrease in H3K4 methylation is likely a result of increased expression of Kdm5b because RNAi-mediated targeting of Kdm5b in double-mutant oocytes results in an increase in H3K4 methylation. An increase in TRP53 acetylation also occurs in mutant oocytes and may contribute to the observed increased incidence of apoptosis. Taken together, these results suggest seminal roles of acetylation of histone and nonhistone proteins in oocyte development.

Fig. 1.

HDAC1 and HDAC2 protein expression patterns during oocyte development. Immunocytochemical analysis of HDAC1 or HDAC2 expression during oogenesis. All samples for a given HDAC were processed for immunocytochemistry together, and all images were taken at the same laser power. The experiment was conducted three times (at least three mice per experiment), and at least 25 oocytes were analyzed for each sample. Shown are representative examples. 5d, 5 d postpartum; 12d, 12 d postpartum; GV, full-grown oocyte; MI, metaphase I; MII, metaphase II. DNA was stained with SYTOX green or DAPI (blue). (Scale bar: 35 μm.)

Fig. 2.

Conditional targeting of Hdac1 and Hdac2 in oocytes results in decreased ovary size and defective oogenesis. (A) Ovaries from mice 6 wk of age from the different genotypes were collected, and their weights measured. Data are expressed as mean ± SEM, and six ovaries (one ovary per mouse) per genotype were measured. The other genotypes are described in the text (*P < 0.0001). (B) Ovary morphology from WT and Hdac1:2^−/− mice 6 wk of age. WT ovaries show presence of mature follicles (arrows), whereas such follicles are absent in ovaries from Hdac1:2^−/− mice. (Scale bar: 1 mm.) (C). Full-grown oocytes are recovered from WT mice, whereas only secondary follicles are recovered from Hdac1:2^−/− mice. (Scale bar: 100 μm.)

Fig. 3.

Developmental block at secondary follicle stage in Hdac1:2^−/− mice. (A) Histological analysis of ovaries obtained from WT and Hdac1:2^−/− mice 18 d of age. In WT ovaries, primary (PF), secondary (SF), and antral follicles (AF) are indicated. In ovaries from Hdac1:2^−/− mice, no antral follicles are found, and there is an increase in the number of degenerating follicles (white arrow). (Scale bars: low-magnification image, 0.5 mm; higher-magnification image, 0.17 mm.) (B) Follicle counts from ovaries obtained from WT and mutant mice 18 d of age: primordial (PmF), primary (PF), secondary (SF), antral follicles (AF), and dead/degenerating follicles (DF). Numbers are mean ± SEM of counts of three sequential sections from serially sectioned ovaries. Data are from four biological replicates (*P < 0.05 and **P < 0.01). (C) Ovaries were collected from WT and Hdac1:2^−/− mice 10 d of age to 6 wk of age and weighed. The data are expressed as mean ± SEM of n ≥ 8 mice (*P < 0.05 and **P < 0.01). (D) Histological analysis of ovaries obtained from WT and Hdac1:2^−/− 12 d of age. In ovaries from mutant mice, there is an increase in the number of degenerating follicles (white arrow). (Scale bar: 0.5 mm.) (E) Follicle counts from ovaries obtained from WT and Hdac1:2^−/− mice 12 d of age. Numbers are mean ± SEM of counts of three sequential sections from serially sectioned ovaries. Data are from four biological replicates (*P < 0.05).

Fig. 4.

Effect of deleting Hdac1 or Hdac2 or both on Hdac1 and Hdac2 transcript and protein abundance. (A) Relative abundance of Hdac1 and Hdac2 transcripts in oocytes obtained from mice 12 d of age. Data are expressed relative to that in WT oocytes. The experiment was performed four times and the data expressed as mean ± SEM (*P < 0.05 and **P < 0.001). (B) Total protein was extracted from oocytes (n = 300 for HDAC1 and n = 100 for HDAC2) obtained from mice 12 d of age for immunoblotting; at least five mice were used. The amount of HDAC1 and HDAC2 in WT oocytes was set as 100%. Quantification of the band intensities revealed that the amounts of HDAC1 relative to WT were 32%, 223%, and 36% in Hdac1^−/−, Hdac2^−/−, and Hdac1:2^−/− oocytes, respectively, and the amounts of HDAC2 relative to WT were 97%, 0%, and 0% in Hdac1^−/−, Hdac2^−/−, and Hdac1:2^−/− oocytes, respectively. The experiment was performed two times. (C) Immunoblot analysis of HDAC3 and HDAC8 expression in WT and mutant oocytes obtained from mice 12 d of age; 200 oocytes were used and collected from at least five mice. The experiment was conducted two times, and similar results were obtained in each case. In B and C, β-tubulin (TUBB) was used as a loading control.

Fig. 5.

Expression of components of HDAC1- and HDAC2-containing complexes in Hdac1:2^−/− oocytes. (A) Relative amount of NuRD, SIN3A, and COREST complex components RBBP4, RBBP7, MTA1, MTA2, MTA3, CHD4, REST, COREST, MECP2, SIN3A, and LSD1 was determined by immunoblot analysis by using total protein extracts from WT and mutant oocytes obtained from at least five mice 12 d of age. Equal numbers of oocytes were loaded per lane. The TUBB loading control is not shown because the immunoblot is a composite of several experiments for which β-tubulin (TUBB) was used as a loading control for each experiment. The experiment was performed three with two times, and similar results were obtained. (B) Quantification of the data shown in A. Data are expressed as mean ± SEM (*P < 0.05). (C) Immunocytochemical detection of RBBP4, RBBP7, MTA1, MTA2, MTA3, CHD4, REST, COREST, MECP2, SIN3A, and LSD1 in WT and mutant oocytes obtained from mice 12 d of age. At least 20 oocytes for each genotype were analyzed, and the experiment was conducted three times with at least three mice used for each experiment. Shown are representative images, and only the nucleus is shown. (Scale bar: 10 μm.)

Fig. 6.

Increased histone acetylation and decreased global transcription and histone H3K4 methylation in Hdac1:2^−/− oocytes. (A) Different acetylated histones were analyzed by immunocytochemistry using oocytes obtained from WT and mutant mice 12 d of age. For each histone variant, at least 20 oocytes from each genotype were analyzed, and the experiment was conducted three times with at least three mice used for each experiment. Shown are representative images, and only the nucleus is shown. (Scale bar: 10 μm.) (B) Global transcription was assayed by BrUTP incorporation by WT and mutant oocytes isolated from at least three mice 12 d of age. (Scale bar: 10 μm.) (C) Quantification of data shown in A. The relative fluorescence intensities of the nuclei were determined and the average value for WT oocytes was set as 1. The experiment was performed three times, with at least three mice used for each experiment, and data are expressed as mean ± SEM, with at least 60 oocytes analyzed for each group (*P < 0.001). (D) Immunocytochemical detection of different H3K4 methylated species on WT and mutant oocytes obtained from mice 12 d of age; only the nucleus is shown, and at least three mice were used for each experiment. (Scale bar: 10 μm.) Signal intensities relative to WT for H3K4me1–3 are 81 ± 1% (n = 4), 58 ± 12% (n = 5), and 40 ± 8% (n = 5), respectively. (E) Quantification of the data shown in D as well as in Fig. S2 shows staining of other methylated histone species. Nuclear staining intensity of specific methylated lysine in WT oocytes was set to 1, and the data are expressed as mean ± SEM. At least 20 oocytes for each genotype and for each modified histone were analyzed; the experiment was conducted three times and at least three mice were used for each experiment (*P < 0.05 and **P < 0.001). (F) Immunocytochemical detection of CTD phosphorylated on S2 or S5 in WT and mutant oocytes obtained from mice 12 d of age; only the nucleus is shown, and at least three mice were used for each experiment. At least 50 oocytes were analyzed, and representative images are shown. The signal intensity for S2 phosphorylation in mutant oocytes is less than that of WT (mean ± SEM, 64 ± 2%; P < 0.01). (Scale bar: 10 μm.)

Fig. 7.

RNAi-mediated targeting of Kdm5b in Hdac1:2^−/− oocytes leads to an increase in histone H3K4 methylation. (A) Hdac1:2^−/− oocytes obtained from mice 12 d of age were injected with control siRNA or Kdm5b siRNA, and the amount of Kdm5b mRNA relative to that present in the control siRNA-injected oocytes was determined by qRT-PCR 52 h following injection. The experiment was performed three times, with at least three mice used for each experiment, and the data are presented as mean ± SEM (*P < 0.001). (B) Immunocytochemical detection of H3K4me3 was performed on oocytes injected as described in A, and only the nucleus is shown. (Scale bar: 10 μm.) The experiment was performed two times (50 oocytes were analyzed in each group and collected from at least three mice), and quantification of the data revealed a 23 ± 6% increase in intensity of the H3H4me3 signal. (C) WT oocytes obtained from mice 12 d of age were injected with Gfp cRNA (control) or Kdm5b cRNA (Kdm5b-O), and the amount of Kdm5b transcript relative to that present in the control oocytes was determined by qRT-PCR 24 h following injection. The experiment was performed twice and the data are presented as mean ± SEM (*P < 0.05). (D) Immunocytochemical detection of H3K4me3 was performed 50 h after injection; only the nucleus is shown. (Scale bar: 10 μm.) The experiment was performed two times (24 oocytes analyzed in each group), and quantification of the data revealed a 28 ± 5% decrease in intensity of the H3H4me3 signal.

Fig. 8.

Deletion of both Hdac1 and Hdac2 leads to apoptosis and TRP53 acetylation. (A) The relative abundance of antiapoptotic transcripts is decreased and that of proapoptotic transcripts increased in Hdac1:2^−/− oocytes isolated from mice 12 d of age. The experiment was performed three times using at least three mice per group, and the data are expressed as mean ± SEM (*P < 0.05). (B) TRP53K379 acetylation is increased in mutant oocytes. Immunocytochemical detection of TRP53K379 acetylation was performed in WT, Hdac1 and Hdac2 single mutants, and Hdac1:2^−/− oocytes obtained from mice 12 d of age; only the nucleus is shown. For each protein, at least 20 oocytes for each genotype were analyzed, and the experiment was conducted three times using at least three mice of each genotype per experiment. (Scale bar: 10 μm.)

Fig. 9.

TSA induces TRP53 acetylation in WT oocytes and increased expression of proapoptotic genes. (A) Immunocytochemical detection of acetylated TRP53 after TSA treatment of WT oocytes for 24 h and 48 h, and only the nucleus is shown. At least 20 oocytes were analyzed, and the experiment was performed two times using at least three mice per experiment. Shown are representative images. (B) Relative abundance of anti- and proapoptotic transcripts determined by qRT-PCR following TSA treatment for 72 h. The experiment was performed three times using at least three mice per experiment, and the data are expressed as mean ± SEM (*P < 0.05).

Fig. P1.

Schematic diagram depicting how loss of HDAC1 and HDAC2 perturbs the transcriptome (genes transcribed) and induces apoptosis. Absence of HDAC1 and 2 not only affects expression of transcription factors (TFs; proteins that affect the transcribing of genes) but also increases expression of the Kdm5b gene, which in turn decreases H3K4 methylation, an activating mark. This decreases global transcription, which may cause cell death (i.e., apoptosis), which could also result from hyperacetylation of TRP53 observed in these mutants.