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Conceived and designed the experiments: YHS YC PJW MLM AR. Performed the experiments: YHS YC SUE SAY MK FY. Analyzed the data: YHS SUE SAY MK FY PJW MLM AR. Contributed reagents/materials/analysis tools: YC PJW AR. Wrote the paper: YHS SUE SAY PJW AR.
Meiosis is unique to germ cells and essential for reproduction. During the first meiotic division, homologous chromosomes pair, recombine, and form chiasmata. The homologues connect via axial elements and numerous transverse filaments to form the synaptonemal complex. The synaptonemal complex is a critical component for chromosome pairing, segregation, and recombination. We previously identified a novel germ cell–specific HORMA domain encoding gene, Hormad1, a member of the synaptonemal complex and a mammalian counterpart to the yeast meiotic HORMA domain protein Hop1. Hormad1 is essential for mammalian gametogenesis as knockout male and female mice are infertile. Hormad1 deficient (Hormad1−/−) testes exhibit meiotic arrest in the early pachytene stage, and synaptonemal complexes cannot be visualized by electron microscopy. Hormad1 deficiency does not affect localization of other synaptonemal complex proteins, SYCP2 and SYCP3, but disrupts homologous chromosome pairing. Double stranded break formation and early recombination events are disrupted in Hormad1−/− testes and ovaries as shown by the drastic decrease in the γH2AX, DMC1, RAD51, and RPA foci. HORMAD1 co-localizes with γH2AX to the sex body during pachytene. BRCA1, ATR, and γH2AX co-localize to the sex body and participate in meiotic sex chromosome inactivation and transcriptional silencing. Hormad1 deficiency abolishes γH2AX, ATR, and BRCA1 localization to the sex chromosomes and causes transcriptional de-repression on the X chromosome. Unlike testes, Hormad1−/− ovaries have seemingly normal ovarian folliculogenesis after puberty. However, embryos generated from Hormad1−/− oocytes are hyper- and hypodiploid at the 2 cell and 8 cell stage, and they arrest at the blastocyst stage. HORMAD1 is therefore a critical component of the synaptonemal complex that affects synapsis, recombination, and meiotic sex chromosome inactivation and transcriptional silencing.
The biology of germ cells is intimately intertwined with meiosis. Meiosis I is a unique biological event, when chromosomes pair, recombine, and segregate. The synaptonemal complex is a protein lattice that enables chromosome pairing and recombination and is unique to meiosis I. Meiosis I requires a subset of factors that are unique to germ cells and meiosis. Germ cell–specific factors are known to play crucial roles during formation of the synaptonemal complex and include synaptonemal complex proteins SYCP1, SYCP2, and SYCP3, among others. We discovered a mouse HORMA domain containing protein, Hormad1 (Nohma), which is germ cell–specific and essential for male and female fertility. Mice deficient in Hormad1 have severe defects in early recombination, synapsis, and segregation—functions attributed to yeast HORMA domain containing protein, Hop1. Moreover, Hormad1 is likely a germ cell–specific component of the meiotic sex chromosome inactivation and transcriptional silencing complex.
Mammalian meiosis is unique to germ cells and a critical step in sexual reproduction. Meiosis reduces the chromosome complement to haploidy in preparation for fertilization. The first meiotic division is unique in pairing of homologous chromosomes, homologous recombination, and formation of chiasmata. The reduction in chromosome numbers happens when homologous chromosomes segregate to opposite poles during the first meiotic division. Proper disjunction (separation) requires crossovers (manifested cytologically as chiasmata). The sister chromatids organize along structures called axial elements (AEs) and transverse elements connect AEs to form the synaptonemal complex (SC) . SC is a proteinaceous structure that connects paired homologous chromosomes during prophase I of meiosis, and SC is critical for wild-type levels of crossovers to occur during meiosis. AEs are critical part of the SCs and mutations in proteins that form AEs disrupt sister chromatid cohesion, recombination, and chromosome segregation –. Proteins with HORMA domain are critical components of the axial elements . HORMA domain proteins are predicted to form globular structure that may sense specialized chromatin states, such as those associated with double strand breaks (DSBs) or other forms of DNA damage . Several mammalian proteins that contain HORMA domain, such as mitotic arrest deficient protein 2, MAD2, are essential for mitosis –. Mice lacking MAD2 unsurprisingly die during early embryogenesis . In lower organisms, several meiotic specific HORMA proteins are known and all are critical for meiosis. These HORMA proteins are: Hop1  and Red1  in yeast; Him-3  in nematodes; and Asy1  in plants. Him-3 localizes to the axial cores of both synapsed and unsynapsed chromosomes. C. elegans Him-3 mutants are deficient in chromosome pairing, synapsis, and the regulation of double strand break repair , –. Synapsis in both male and female Asy1 mutants is disrupted –. In yeast, plants, and nematodes, HORMA domain proteins are critical components of the synaptonemal complex and essential for meiosis I. Others and we identified a previously uncharacterized gene that we named Nohma, later re-named to Hormad1 –. Hormad1 encodes a protein that contains a HORMA domain, and unlike Mad2, Hormad1 expression is germ cell–specific . Mouse and human HORMAD1 are highly conserved and share 77% amino acid identity overall, and share 89% amino acid identity in the HORMA domain. Moreover, mouse and human HORMAD1 HORMA domains share 28% amino acid identity with Hop1 HORMA domain. Hop1 in yeast appears to bind near or at the sites of DSB formation and may modulate the initial DSB cleavage . Hop1 mutants in yeast have reduced number of DSBs , and Hop1 may participate in recruiting DMC1, RAD51 and other proteins that are required for DNA repair during meiotic synapsis and recombination –. Phosphorylation of Hop1 by Mec1/Tel1 yeast kinases is important for interhomologue recombination and prevents DMC1-independent repair of meiotic DSBs . Here we report that HORMAD1 is likely the mammalian counterpart of Hop1, and that HORMAD1 deficiency disrupts mammalian synaptonemal complex formation, meiotic recombination, and chromosome segregation.
We previously showed that Hormad1 RNA expression in testes began at postnatal day 10, with little expression detected at birth or postnatal day 5 . Hormad1 RNA expression pattern coincided with the onset of meiosis, and appearance of primary spermatocytes in the developing testes. In situ hybridization with anti-sense Hormad1 riboprobe revealed that Hormad1 expression was confined to germ cells, and specifically spermatocytes, with no signal detected in spermatogonia or sertoli cells . We generated antibodies against HORMAD1 and studied its protein localization pattern in testes. HORMAD1 localized exclusively in germ cells, specifically in zygotene, and early pachytene spermatocytes as previously described for the RNA expression .
Since HORMAD1 protein showed localization consistent with its potential role in meiosis I and contains the HORMA domain, we disrupted the Hormad1 gene to examine its requirement for germ cell development and meiosis in mouse. Hormad1 is located on chromosome 3 and composed of sixteen exons. We deleted exons 4 and 5 (Figure S1A), and this mutation is predicted to remove 33 amino acids from the highly conserved HORMA domain and to cause a frame shift mutation. Small amounts of truncated Hormad1 RNA transcripts were detectable on RT-PCR, and Western blots on testes extracts showed absence of HORMAD1 protein in knockout mice as expected (Figure S1B and S1C).
Female and male heterozygote matings produced expected Mendelian ratios, averaged 8.1±2 pups per litter (n=20 breeding pairs) over a 6-month period, and remained fertile for at least 9 months. The litter size was statistically not significantly different from the wild-type average (8.4±2 pups per litter). Male and female mice heterozygous for the mutation (Hormad1+/−) were fertile with grossly normal male and female gonadal morphology and histology. However, both Hormad1−/− males and females were infertile with no pups produced over a period of 6 months from mating with wild-type female and male mice, respectively.
While ovaries showed no gross morphologic differences between the knockout and wild-type mice, Hormad1 knockout adult testes were significantly smaller than the wild-type testes (Figure S2A). Testes in the 7-day-old Hormad1−/− mice were grossly normal and weighed 8.0±2.0 mg/pair and did not significantly differ from the wild-type, 9.0±3.0 mg/pair of testes. By 4 weeks of postnatal life, the knockout testes (47±6 mg/pair) were 50% of the wild-type weight (94±1.7 mg/pair), and by 8 weeks the knockout testes (60±7.7 mg/pair) were 27% of the wild-type weight (225±2.7 mg/pair) (Figure S2B).
Histology at 6 weeks showed hypocellular seminiferous tubules with clumps of sertoli cells in the lumen. We observed spermatogonia and early spermatocytes, but no post-meiotic germ cells such as spermatids or spermatozoa (Figure 1A–1H). We therefore carefully examined spermatogenesis in Hormad1−/− mice. Spermatogenesis is a complex process that involves differentiating and proliferating self-renewing spermatogonia that differentiate into spermatozoa. Type A spermatogonia self-renew and can initiate differentiation into Type B spermatogonia which in turn differentiate into primary spermatocytes. Primary spermatocytes undergo meiosis I to form secondary spermatocytes. Secondary spermatocytes enter meiosis II and divide to produce haploid spermatids. We examined Hormad1 knockout testes histology during gonadal development to determine the stage at which spermatogenesis is disrupted. Identical testes weights at postnatal day 7, and similar histology between Hormad1−/− and wild-type testes argue that pre-spermatogonia in Hormad1−/− testes proliferate into Type A spermatogonia without major disruption. Immunohistochemistry with antibodies directed against PLZF and SOHLH1, markers that identify self-renewing (PLZF) and differentiating spermatogonia (SOHLH1), showed the presence of both proteins in the wild-type as well as the knockout animals, confirming that spermatogonia are unaffected (Figure 1J, 1K, 1N, 1O). At postnatal day 10, testes contain preleptotene/leptotene primary spermatocytes, and there was no gross difference between wild-type and Hormad1−/− testes. At 14 days, testes contain pachytene spermatocytes, and there was a significant difference between the wild-type and Hormad1−/− testes, with many apoptotic cells and few pachytene spermatocytes in Hormad1−/− testes (), and rising apoptotic index with age in the knockout as compared to the wild-type (Figure S3K). We counted leptotene, zygotene and pachytene spermatocytes in 6 week old wild-type and Hormad1−/− testes. Hormad1−/− testes showed declining number of spermatocytes beginning in stages II-III with 28±8 spermatocytes as compared to 52±12 in the wild-type (Figure 1A and 1E). No spermatocytes were noted in stages IV-IX in the Hormad1−/− testes (Figure 1F and 1G), and no significant difference was noted in the spermatocyte number in stages X-XII between the wild-type and knockout testes (Figure 1D and 1H). These results indicate that Hormad1 deficiency in the male gonad caused meiotic arrest at the pachytene stage.
Previous studies on HORMA domain proteins indicate their specific involvement in cell division. MAD2 is a ubiquitously expressed mammalian HORMA domain protein involved in both meiosis and mitosis , while yeast HOP1, RED1, nematode HIM3 and plant ASY1 genes are specifically involved in meiotic segregation, synapsis and recombination , –, , , . No mammalian counterparts to Hop1, Red1, Him3 and Asy1 have been functionally evaluated up to date. We previously hypothesized that HORMAD1 is a functional counterpart to Hop1, Him3 and Asy1 . Critical components of the synaptonemal complex include meiosis specific SYCP1, SYCP2 and SYCP3 proteins. SYCP1 is a major component of the transverse filaments, while both SYCP2 and SYCP3 are components of the axial lateral elements –. To determine HORMAD1 localization during meiosis, and whether HORMAD1 localizes to the axial elements, or transverse filaments, we used antibodies against SYCP1, SYCP2 and SYCP3 to study their respective co-localization with HORMAD1. HORMAD1 co-localized with SYCP3 and SYCP2 but did not co-localize with SYCP1, which indicates that HORMAD1 is located along the axial elements (Figure 2A, 2C, and 2E). Recent studies also show that HORMAD1 localizes to the axial elements –. We also studied whether absence of SYCP2 affected HORMAD1 localization along the chromosomes. HORMAD1 localization is independent of major germ cell–specific components of the axial elements of the synaptonemal complex because neither Sycp2 (Figure 3A–3F) nor Sycp3 mutation affected HORMAD1 localization to the axial elements .
We examined localization of germ cell–specific synaptonemal complex proteins SYCP1, SYCP2 and SYCP3 in Hormad1−/− testes. SYCP1 is a major component of the transverse filaments and is known to form fibrillar structures in Sycp3 mutant spermatocytes . However, the fibers are truncated, contain axial gaps, and do not associate with the centromeres of the meiotic chromosomes. We also observed truncated fibers with anti-SYCP1 antibodies in Hormad1−/− mice (Figure 2B). HORMAD1 is therefore not necessary for SYCP1 binding to the chromatin. We also examined localization of SYCP2 and SYCP3 proteins in Hormad1−/− mice. The localization of SYCP2 and SYCP3 to the chromatin was not significantly affected by the lack of HORMAD1 (Figure 2D and 2F). These results indicate that HORMAD1 is not necessary for SYCP2 and SYCP3 localization to the axial elements.
The deficiency in synaptonemal complex proteins such as SYCP3 is known to affect chromosome synapsis . In order to determine the effect of HORMAD1 deficiency on chromosome synapsis during meiosis I, we utilized CREST sera. CREST sera labels centromeres and allows the determination of the pairing status during meiosis . In the wild type spermatocytes, prior to the synapsis, 40 centromeres are usually observed in the leptotene stage. The number of visible centromeres become reduced as the synapsis of homologues progresses. At the completion of the synapsis in pachytene, 20 centromeric foci are usually observed corresponding to 19 autosomal homologues and partially paired X-Y chromosomes. We examined CREST foci formation in Hormad1−/− spermatocytes. Examination of over 100 Hormad1−/− spermatocytes and oocytes in meiosis I, revealed greater than 20 centromeric foci in both male and female germ cells, most containing 40 CREST foci (Figure 4A, 4D, and data not shown). These results indicate that Hormad1 deficient germ cells cannot complete homologous chromosome pairing, and Hormad1 is therefore critical for chromosome synapsis during meiosis.
Our experimental evidence strongly suggests that HORMAD1 localizes to the axial core and is yet another critical component of the synaptonemal complex. To determine the effect of Hormad1 deficiency on the structure of the synaptonemal complex, we visualized synaptonemal complexes during meiosis I in wild-type and Hormad1−/− spermatocytes using electron microscopy. In the wild-type, synaptonemal complexes were well visualized during the pachytene stage of meiosis (Figure 4B and 4C). Electron microscopy examination of one hundred and ten Hormad1−/− spermatocytes from three independent experiments revealed the lack of the typical tripartite synaptonemal complex structure (Figure 4E and 4F). Persistence of pre-synaptic number of centromeric foci (CREST staining) in Hormad1−/− spermatocytes, as well as non-visualization of the tripartite synaptonemal complex structure by electron microscopy, demonstrate that HORMAD1 is essential for chromosomal synapsis.
Previous studies have indicated that Sycp3 deficiency has subtle effects on meiotic recombination . Early recombination events do not seem to be disrupted in Sycp3, as similar number of DMC1 foci are present in Sycp3 mutant and wild-type meiosis . DMC1 is a meiotic specific recombinase that together with ubiquitously expressed RAD51 catalyzes homologous pairing and DNA strand exchange –. These early steps in recombination are critical for establishing the physical connections between homologous chromosomes during meiosis. Hop1 has been implicated in modulating the formation and processing of double stranded breaks . We examined formation of DMC1, RAD51, and RPA foci in zygotene stage Hormad1−/− spermatocytes. There is a dramatic decrease in the number of DMC1 foci as compared to the wild-type (Figure 5A and 5B). We counted a total of 98.9±28.2 DMC1 foci in the wild-type spermatocytes (n=50) and 9.28±3.9 DMC1 foci in the Hormad1−/− spermatocytes (n=45). The number of RAD51 foci was also decreased from 189.3±31.8 in the wild-type spermatocytes (n=50), to 69.3±34.5 in the Hormad1−/− spermatocytes (n=40) (Figure 5C and 5D).
Following double-strand breaks formation by SPO11, RPA is recruited together with RAD51 to the single stranded DNA regions . We counted a total of 194.5±64.5 RPA foci in the wild-type spermatocytes (n=30) as compared to 70.1±38.5 foci in the Hormad1−/− spermatocytes (n=20) (Figure 5E and 5F). These results indicate that early meiotic recombinational events were disrupted and not surprisingly, MLH1, a protein that forms foci in later stages of recombination and required for the formation of most of the crossovers (chiasmata) observed in mice , was dramatically reduced in Hormad1−/− spermatocytes (data not shown).
We also examined DMC1, RAD51 and RPA foci formation in female meiocytes at embryonic day 15.5 (E15.5). Embryonic ovaries contain zygotene to early pachytene oocytes at E15.5 . We counted a total of 208.7±117.1 DMC1 foci in the wild-type E15.5 oocytes (n=50), and 79.1±81.5 foci in the Hormad1−/− oocytes (n=30) (Figure 6A and 6B), a total of 197.9±46.0 RAD51 foci in the wild-type oocytes (n=50) versus 85.1±37.6 foci in the Hormad1−/− oocytes (n=40) (Figure 6C and 6D) and a total of 317.16±135.3 RPA in the wild-type oocytes (n=50) and 51.7±48.8 in the Hormad1−/− oocytes (n=50) (Figure 6E and 6F). DMC1, RAD51 and RPA foci are therefore, similar to our observations in spermatocytes, significantly decreased in Hormad1 deficient female meiocytes.
These results indicate that homologous recombination is significantly affected in Hormad1−/− mammalian germ cells, as previously reported for HOP1 . We also observed effects of Hormad1 deficiency on γH2AX staining (a phosphorylated form of histone H2AX), a well known surrogate marker for DSB formation . In the leptotene stage, phosphorylation of H2AX is induced by SPO11 catalyzed DSBs in meiotic DNA, and γH2AX appears as large, cloud-like patterns thatdisappearat the pachytene stage . At the leptotene stage, γH2AX staining in Hormad1−/− spermatocytes was significantly decreased (76% decrease in signal intensity) as compared to the wild-type (Figure 7A–7D). γH2AX staining was also significantly decreased in Hormad1−/− fetal oocytes (71% decrease in signal intensity) (Figure 6G and 6H). These results suggest that similar to Hop1 mutants, DSBs do not efficiently form in Hormad1 mutants.
HORMAD1 protein localization along autosomes was previously shown to be transient  (Figure 7B and 7F). HORMAD1 staining was highest along unsynapsed chromosome axis in the zygotene to pachytene stage and diminished significantly along autosomes in the pachytene – (Figure 7F and 7I). Interestingly, HORMAD1 localized strongly along desynapsed autosomes in diplotene meiocytes . During the pachytene stage, HORMAD1 is faintly visible along the autosomes, but co-localizes strongly with γH2AX on the XY chromosomes, and specifically along the axial elements –. γH2AX is a phosphorylated form of histone H2AX, and a marker for DSBs , . H2AX is phosphorylated throughout the chromatin in leptotene spermatocytes and by the pachytene stage –, γH2AX staining is undetectable on autosomes and restricted to the sex body (Figure 7E and 7F) . H2ax knockout shows the essential role for H2AX in sex body formation and meiotic sex chromosome inactivation . Meiotic sex chromosome inactivation also involves ATR and BRCA1 dependent phosphorylation of H2AX , . Interestingly, Hormad1 deficiency, similar to H2ax deficiency, abolishes the formation of the sex body (Figure 7G and 7H, and data not shown). The lack of sex body formation is most likely due to the disruption of Hormad1−/− spermatocytes prior to the pachytene. We examined the γH2AX localization in the wild-type and Hormad1−/− spermatocytes. Chromatin in Hormad1−/− spermatocytes stained with anti-γH2AXantibodies, but no preferential localization to the sex chromosomes was observed (Figure 7G and 7H). BRCA1 and HORMAD1 have recently been shown to co-localize in the sex body . We determined whether BRCA1 localization is dependent on HORMAD1. BRCA1 protein could not be localized in Hormad1−/− spermatocytes (Figure 7I–7J, 7M). This finding is unlike H2ax knockout, where BRCA1 is still detected on the sex chromosomes despite the lack of the sex body . We also examined ATR localization in wild-type and Hormad1−/− testes. In wild-type testes, HORMAD1 and ATR co-localized in the sex body, but we could not detect ATR in Hormad1−/− spermatocytes (Figure 7K–7L, 7N). Above data suggest that HORMAD1 may be involved in the recruitment of BRCA1, ATR and γH2AX to the sex chromosome.
Since ATR, BRCA1 and γH2AX are involved in the transcriptional silencing of sex chromosomes, we examined whether Hormad1 deficiency affects transcriptional repression. Previous studies have shown that H2ax and Brca1 deficiencies individually, lead to the over-expression of genes exclusively expressed from the X or Y chromosome . We examined whether X-linked germ cell–specific genes were over-expressed in Hormad1−/− testes compared to wild-type testes. We performed quantitative real-time PCR on select autosomal genes (Hormad2, Rnh2 and Mov10l1) as well as X-chromosome derived genes (Usp26, Fthl17, Pramel3, Tex11, and Tex13) on wild-type and Hormad1−/− testes (Figure 8A–8H). Rnh2 and Mov10l1 are germ cell–specific transcripts, derived from autosomes, and were not differentially expressed between wild-type and Hormad1−/− testes (Figure 8B and 8C). In contrast, all of the germ cell–specific transcripts transcribed from the X chromosome were significantly elevated in Hormad1−/− testes over the wild-type. These include Usp26 (4.5 fold increase), Fthl17 (6.5 fold increase), Pramel3 (3.5 fold increase), Tex11 (2.2 fold increase), and Tex13 (2.8 fold increase) (Figure 8D–8H). Moreover, RNA expression microarray analyses comparing two week old Hormad1 deficient testes with corresponding wild-type, indicate that almost 20% of the up-regulated genes derive from the X chromosome (Figure 8I). Our results are remarkably similar to transcriptional de-repression observed in H2ax and Brca1 mutants –, and indicate that HORMAD1 is a germ cell and meiosis specific factor critical in meiotic sex chromosome inactivation and transcriptional silencing.
HORMAD1 is a likely mammalian counterpart to the yeast HORMA domain meiotic protein, Hop1. Hop1 is phosphorylated by Mec1/Tel1, the budding yeast homologue to the mammalian ATR and ATM kinases, and this phosphorylation is thought to play an important role in inter-homologous recombination . We therefore examined HORMAD1 expression in Atm deficient mice as well as ATM expression in Hormad1−/− animals. ATM is a serine/threonine-specific protein kinase that has been associated with cell cycle regulation, apoptosis, and response to DNA damage repair. ATM kinase activation is associated with increased auto-phosphorylation of ATM at multiple sites including serine 1981 . We examined HORMAD1 protein expression in testes between postnatal days 5–21 (Figure 9A). Eight to ten day old testes contain spermatogonial cells as well as preleptotene/leptotene spermatocytes . At postnatal day 14–18, testes contain pachytene spermatocytes and visible sex bodies . HORMAD1 is known to be phosphorylated  (Figure 9B). Western blot analysis on testes extracts detected phospho-HORMAD1 beginning at postnatal day 14 (Figure 9A). Phospho-HORMAD1 protein was decreased in postnatal day 18 and 21 wild-type testes (Figure 9A). Phospho-HORMAD1 appearance correlates temporally with sex body formation. HORMAD1 phosphorylation was not affected by Atm deficiency (Figure 9A). These results indicate that ATM is not responsible for HORMAD1 phosphorylation. We also examined HORMAD1 localization in Atm−/− spermatocytes. We observed HORMAD1 localization to chromosomal axes in Atm deficient spermatocytes, as previously described by others  (Figure 9C and 9D). Since synaptonemal complexes do not form in Atm mutants, HORMAD1 association with unsynapsed chromosomes does not require ATM. The anti-ATM phospho-S1981 antibody did not detect phosphorylated ATM in Hormad1−/− testes (Figure 9E and 9F). These results suggest that HORMAD1 is upstream of ATM auto-phosphorylation, and therefore likely upstream of ATM kinase activation.
We have previously shown that Hormad1 RNA expression in the ovary was confined to the germ cell . Meiosis I in the female gonad commences circa E13.5 and most oocytes arrest at the dictyate stage by the time of birth. Antibodies against HORMAD1 recognized HORMAD1 protein at E14.5 (leptotene) and E18.5 (arrest in diplotene) oocytes (Figure 10A and 10B), but little HORMAD1 protein was detected in the newborn ovary oocytes (Figure 10C), at the time when oocytes are arrested in diplotene. Deficiency in genes critical in meiosis can disrupt early ovarian development, as is the case for Dmc1, Msh5, Spo11 and Atm –.
We therefore examined ovarian development in Hormad1−/− females. Antibodies against germ cell–specific transcriptional regulator SOHLH1  stained wild-type and knockout oocytes throughout embryonic gonadal development with no significant differences noted (Figure 10A–10F). Moreover, the numbers of primordial, primary and secondary follicle counts did not significantly differ between the mutant and wild-type ovaries at post-natal day 8 (Figure S4). These results indicate that Hormad1 deficiency does not affect embryonic ovarian development, germ cell cyst breakdown, and primordial follicle formation. We also examined the histology of wild-type and Hormad1−/− ovaries between 2 and 30 weeks of life. Mice reach sexual maturity around 6 weeks of life, and mouse ovaries at this time consist of all of the follicular types including corpora lutea, an indication that the ovaries are ovulating. Postnatal Hormad1−/− ovaries were grossly indistinguishable from wild-type mice between 2 and 30 weeks of life, with abundant corpora lutea in the Hormad1−/− ovaries indicating that the normal process of oocyte maturation was not disrupted, and ovulation has occurred (Figure 11A–11H). We induced superovulation in knockout and wild-type mice with exogenous gonadotropins to determine whether subtler ovarian defects contributed to infertility in Hormad1−/− mice. Hormad1−/− females super-ovulated 28±11 eggs (n=22), while wild-type animals superovulated 29±14 eggs (n=13). We therefore did not observe significant difference between the number of eggs superovulated from wild-type versus knockout mice. These results indicate that ovarian development is grossly normal in Hormad1−/− mice, and that ovarian defects are unlikely to account for observed infertility.
We studied early embryonic development of fertilized Hormad1−/− oocytes because ovarian defects were unlikely to explain observed infertility. We recovered embryos from wild-type male matings with Hormad1−/− females at E0.5, E1.5, E2.5, and E3.5. Comparable numbers of morphologically indistinct 1-cell zygotes were recovered from oviducts of control and mutant female mice at E0.5 (Figure 12A–12C), and little difference was noted at the 2-cell stage, except for an increased number of 1-cell embryos in the knockouts, indicating a lag in the progression from the 1-cell to 2- cell stage (Figure 12D–12F). By E2.5, the number of normal appearing 8-cell stage embryos was significantly less in Hormad1−/− fertilized eggs as opposed to the wild-type (Figure 12G–12I), and no morphologically normal blastocysts were observed in the Hormad1−/− fertilized eggs at E3.5. It is interesting to note that at E3.5, a significant number of 4 and 8 cell stage embryos were observed in the knockout while only blastocysts were observed in the wild-type E3.5 embryos (Figure 12J–12L). We also tested, using Chicago Sky Blue 6B dye (Sigma, MO, USA) injection into the tail vein , whether blastocysts derived from Hormad1−/− fertilized eggs could implant. We did not detect implantation of Hormad1−/− fertilized eggs (Figure 12M). These results indicate that a defect in early embryogenesis led to premature loss of embryos.
We hypothesized that aneuploidy is the major cause of embryo wastage in Hormad1−/− females. Unlike meiosis in testes, errors during oocyte meiosis have milder effect on oocyte loss and apoptosis, as has been observed in Sycp3 and Smc1β knockouts , , . The presence of growing oocytes in the Hormad1−/− ovaries, as opposed to early germ cell loss observed in testes during pachytene stage of meiosis, indicates greater tolerance of Hormad1 deficiency in the oocytes as compared to spermatocytes. The whole range of ovarian follicle types are present in the Hormad1−/− ovaries, including primordial follicles, primary, secondary and antral follicles. In contrast to other critical genes during meiosis, such as Dmc1, Msh5, Spo11 and Atm that cause early ovarian failure due to rapid loss of oocytes following disruption of meiosis I –, Hormad1 deficiency does not activate apoptotic pathways and does not lead to gross premature loss of oocytes. We used chromosome specific mouse BAC (bacterial artificial chromosome) clones to perform fluorescent in situ hybridization (FISH) on germinal vesicle (GV) oocytes, 2 cell and 8 cell stage embryos to determine whether aneuploidy is significantly higher in Hormad1−/− embryos. Wild-type GV oocytes are arrested in meiosis I and contain bivalent chromosomes that consist of four chromatids. Examination of 58 GV oocytes from three independent knockout animals and 112 GV oocytes from three wild-type animals with BAC probes specific for chromosomes 19, 18 and X, revealed no significant differences, with presence of four chromatids for each of the chromosome examined as expected (Figure 13A–13C). Following fertilization and completion of meiosis II, each cell in the developing embryo should contain two chromosomes except for the sex chromosomes. We examined by FISH, mouse chromosomes 6 and X in the 2-cell wild-type and Hormad1−/− embryos. Sixty three cells examined for chromosome 6 in the Hormad1−/− 2-cell embryos revealed that 22% of the cells had 4 signals corresponding to chromosome 6, 35% had 3 signals corresponding to chromosome 6, 24% had 1 signal, and 5% had no signal. Wild-type 2-cell embryos were also examined by chromosome specific FISH, and 44 cells examined out of 46 showed only 2 signals, as expected. In 2-cell Hormad1−/− embryos, out of sixty three cells examined for chromosome X, 44% showed four, three or no signal. Wild-type 2-cell stage embryos, as expected, showed only 2 or 1 signal, consistent with either XX or XY sex of the cell. These results indicate widespread hypo and hyperdiploidy in 2-cell embryos (Figure 13D–13F).
We also examined cells from the 8-cell stage embryos. A total of 84 cells from the Hormad1−/− embryos were examined with FISH probes specific for chromosomes 11, X and 2. Hormad1−/− cells from the 8-cell stage, hybridized with BAC specific to chromosome 11, showed 4 signals in 10% of the cells, and 3 signals in 55% of the cells. Therefore, a total of 65% of the cells were either monosomic or trisomic for chromosome 11. In the wild-type 8 cell stage embryo, a total of 96 cells were scored for chromosome 11, and only 2 out of 96 (2%) cells showed a single signal, while 94 out of 96 cells (98%) showed two signals as expected. Similar results were obtained for chromosome 2 (Figure 13G–13I). Our results indicate widespread aneuploidy involving different chromosomes in both 2-cell and 8-cell embryos. Such aneuploidy causes early embryo demise and failure of implantation.
We also visualized second metaphase (M2) spindle using anti-β-tubulin antibodies in Hormad1−/− oocytes. During M2, sister chromatids migrate to the opposite pole to form functional, haploid gametes. Sister chromatids are bi-oriented on the spindle and congregate in the center of the spindle prior to separation (Figure 13J). Each sister kinetochore in a pair is attached to the opposite spindle, and such arrangement generates sufficient tension to cause proper segregation. M2 in Hormad1−/− oocytes is grossly disrupted with mis-orientation of chromatid attachment to the spindle observed in all M2 oocytes examined (Figure 13K) (n=50). These results indicate importance of Hormad1 in proper spindle formation and segregation.
We discovered Hormad1 (Nohma), using in silico screen to identify germ cell–specific transcripts critical for gonadal development . We showed that Hormad1 expression was confined to germ cells, and that Hormad1 transcripts were mostly concentrated in the pachytene spermatocytes and early oocytes. Here we show that Hormad1 is essential for both male and female fertility in the mouse. HORMA domain-containing proteins interact with chromatin, particularly chromatin associated with DNA adducts, and are critical mitotic spindle checkpoint proteins . HORMA domain-containing proteins are critical regulatory proteins for mitosis as exemplified by MAD2 protein, and meiosis as exemplified by Hop1p , and Red1p  in yeast, Him-3  in nematodes, and Asy1  in plants. The 205-aa HORMA domain in the mouse HORMAD1, shares highest similarity to the Saccharomyces cerevisiae Hop1 protein (28% amino acid similarity). Yeast Hop1 mutants have defects in chromosome condensation, synapsis and recombination , , and Hop1p binds DSBs during meiotic prophase and appears to play an important role in interhomologue recombination. We previously hypothesized, based on homology to Hop1 in yeast, that Hormad1 plays an important role in mammalian meiosis. Our results indicate that HORMAD1 is the mammalian homologue to the Hop1 protein in yeast.
Our current studies, and those of others, have shown that HORMAD1 protein is confined to germ cells and co-localizes with synaptonemal complex proteins SYCP2 and SYCP3 on the chromatin as part of the axial core , . Synaptonemal complex is a complex structure composed of multiple germ cell–specific and ubiquitously expressed proteins that connect paired homologous chromosomes. Similar to railroad tracks, the synaptonemal complex axial lateral elements (SYCP2, SYCP3) are connected to each other by proteins known as transverse filaments (SYCP1, TEX12) . The axial/lateral elements play critical roles in chromosome condensation, pairing, and repress recombination pathways that involve sister chromatids . Synaptonemal complex formation is associated with HORMAD1 depletion from the axes –. Our data show that HORMAD1 phosphorylation peaks at the time when HORMAD1 localization shifts to the sex body. ATM and ATR do not appear to be involved in HORMAD1 phosphorylation and binding to chromosomal axes. Previous study showed that Trip13, a homologue of yeast Pch2 kinase, may be involved in HORMAD1 depletion from synapsed chromosome axes . Kinases, such as Mec1/Tel1 in yeast are important to effect inter-homologous recombination  by phosphorylating Hop1, but not much is known regarding HORMAD1. Future studies are necessary to understand HORMAD1 phosphorylation sites and responsible kinases.
Although HORMAD1 is not essential for the binding of well-characterized germ cell–specific synaptonemal complex proteins such as SYCP1, SYCP2 and SYCP3, HORMAD1 deficient spermatocytes are defective in synapsis and do not form recognizable synaptonemal complexes. Hormad1 deficiency causes a male meiotic arrest, that is similar to male meiotic arrests observed in other components of the synaptonemal complex and recombination including: Atm, Spo11, Sycp1, Sycp2, Sycp3 and Dmc1 , –, –. These findings are not surprising as HORMAD1 co-immunoprecipitates with known axial proteins, SYCP2, SYCP3, REC8, and SMC1β . Whether these interactions are biologically significant, and whether HORMAD1 interacts with other members of the synaptonemal complex, including newly discovered HORMAD 2 protein, is unclear. Interestingly, Hop1 interacts with another phosphoprotein, Red1 , , and whether HORMAD2 is the mammalian counterpart of Red1 remains to be seen.
Like its counterpart in yeast, Hop1, Hormad1 deficiency disrupts early and later stages of recombination as shown by the drastic diminution in the formation of DMC1, RAD51, RPA and MLH1 foci. These findings differ from the Sycp3 mutant, which affects later stages of recombination . DMC1 is a meiotic specific recombinase that together with ubiquitously expressed RAD51, catalyzes homologous pairing and DNA strand exchange, and marks earlier stages of recombination than MLH1 foci . HORMAD1 expression is not significantly affected in Dmc1 mutant spermatocytes . These results indicate that unrepaired DSBs and synaptonemal complexes between non-homologous chromosomes seen in Dmc1 mutants, do not affect HORMAD1 localization. In Hop1 mutants, DMC1 foci are only faintly detected when compared to controls . These findings have led to a suggestion that Hop1 binds at or near the sites of DNA double strand break and modulates the action and perhaps recruitment of recombinases such as DMC1 and RAD51 . Moreover, Hop1 appears to be part of the meiosis specific surveillance system that monitors meiotic double stranded break repair . Our observations support the possibility that HORMAD1 affects early recombination and may therefore perform similar functions in the mammalian meiosis.
During mammalian meiosis, X and Y chromosomes are inactivated and transcriptionally silenced (meiotic sex chromosome inactivation, MSCI) and chromatin condenses to form a sex (XY) body. The epigenetics of MSCI involves γH2AX, BRCA1 and ATR at a minimum. The most intriguing finding regarding HORMAD1 is its co-localization with BRCA1, ATR and γH2AX in the sex chromosomes during pachytene . Hormad1 deficiency, similar to H2ax deficiency, abrogates formation of the sex body, as well as γH2AX localization to the sex chromosomes. Hormad1 deficiency also abrogates BRCA1 and ATR localization to the sex chromosomes. Interestingly, H2ax deficiency does not affect BRCA1 localization to the sex chromosome, despite anomalous sex body, while BRCA1 deficiency does affect both ATR and γH2AX localization to the sex body. These findings indicate that HORMAD1 is involved in the initial recruitment of BRCA1, ATR and γH2AX to the sex body. The mechanism whereby HORMAD1 affects ATR, γH2AX and BRCA1 localization is unclear, but our results are consistent with the conclusion that HORMAD1 is upstream of ATR, γH2AX and BRCA1, and establishes HORMAD1 as an essential component of the MSCI complex. Moreover, we have shown here that Hormad1 deficient testes preferentially over-express X-linked genes as observed in mice deficient for other components of the MSCI complex such as γH2AX and BRCA1 . Almost 20% of genes preferentially up-regulated from the Hormad1−/− testes are derived from the X chromosome. Hormad1 is therefore the first germ cell–specific component known to play a role in MSCI.
Unlike males, meiosis I in females arrests during the embryonic development at the diplotene stage and is completed upon ovulation. The lack of dramatic germ cell apoptosis during ovarian follicle development has also been observed for Sycp2 and Sycp3 deficient mice , , and reiterates the laxity of oocytes to early meiotic errors, as opposed to spermatogenesis, where meiotic check point causes the dramatic phenotype observed in male gonads. Hormad1 deficiency does not affect folliculogenesis nor ovulation. These findings are unusual, given that Hormad1 affects recombination, and recombination associated proteins DMC1 and MLH1 are important in early oogenesis. Unrepaired recombination intermediates or defects in homologous chromosome pairing and synapsis are likely causes of early oocyte loss . Spo11 deficient animals can ameliorate observed oocyte losses in recombination defective mutants, presumably due to the failure of Spo11−/− deficient mice to form DSBs that initiate meiotic recombination. Spo11 deficiency does not affect HORMAD1 association with the chromosome axes , a finding similarly observed for Hop1 localization in Spo11−/− yeast . Moreover, Hop1 mutants are required for full levels of DSBs formation , . It is therefore possible that early loss of oocytes in Hormad1 deficient animals does not occur in part because unrepaired DSBs do not form. Reduced staining of γH2AX in Hormad1 mutant leptotene stage oocytes argues that unrepaired DSB formation is indeed reduced. However, unlike Spo11−/− ovaries that have abnormal folliculogenesis, Hormad1−/− ovaries are not defective in folliculogenesis and other mechanisms must protect Hormad1 mutant oocytes from early loss.
Infertility in Hormad1−/− females is due to early embryo demise. This is in stark contrast to males, where spermatocytes are eliminated due to the pachytene defect. The greater ability of oocytes to tolerate meiosis I errors is well known , and exemplified by Sycp3 and Smc1β knockouts , , which have defects in synapsis and recombination. Mechanisms that underlie oocyte's insensitivity to such errors are not well understood . Embryos derived from Hormad1−/− oocytes fertilized with wild-type sperm, cannot implant. Examination of post-fertilization events in fertilized Hormad1−/− oocytes, showed significant abnormalities in early embryo development, including growth lag, and severe disruption in development prior to the blastocyst stage. Very few blastocysts were produced in fertilized Hormad1−/− oocytes, and most were abnormal in morphology. We utilized FISH hybridization against specific chromosomes to show widespread hypo and hyperdiploidy during early embryonic development. These results are reminiscent of Sycp3 and Smc1β knockout embryos. SYCP3 and SMC1β are involved in the synaptonemal complex formation, are germ cell–specific, and implicated in human aneuploidy. Sycp3 deficient females are subfertile, but one third of the embryos die in utero due to aneuploidy . Smc1β encodes a meiosis specific component of the cohesin complex, important in sister chromatid cohesion and recombination . Smc1β deficient females are sterile, like Hormad1−/− females, and meiosis continues until metaphase II .
However, unlike Smc1β−/−, Hormad1−/− ovaries do not lose oocytes by 6 months of age, and postnatal oocyte counts do not differ significantly from the wild type. Sycp3−/− ovaries show significant decline in the primordial follicle pool at postnatal day 8, indicating accelerated apoptosis of oocytes as germ cell clusters break down to form primordial follicles . DSBs, as indicated by the assembly of the histone variant γH2AX, form in Smc1β−/− and Sycp3−/− meiocytes and may lead to higher rate of oocyte loss in these mutants , . Hormad1 deficient ovaries do not display loss of primordial follicles. Hormad1−/− ovaries may not be losing oocytes at an accelerated rate due to our observations that DSBs, as assessed by γH2AX staining, are substantially reduced in Hormad1 mutants. Similar to Hop1 mutants in yeast, lack of HORMAD1 may derepress Dmc1-independent inter-sister repair pathway, resulting in efficient DNA break repairs . The reduced DSBs in Hormad1−/− meiocytes, and de-repression of Dmc1-independent inter-sister repair pathways, may not affect regular apoptotic pathways that eliminate substantial number of oocytes during germ cell clusters breakdown to form primordial follicles at the time of birth, resulting in no decrease in Hormad1−/− primordial oocyte numbers.
All murine experiments were carried out on the 129S7/SvEvBrd x C57BL/6 hybrid background. Litters were weaned at 3 weeks and breeding pairs set up at 6 weeks of age. One mating pair was placed per cage and inspected daily for presence of litters. All experimental and surgical procedures complied with the Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Agricultural Animal Care and Use of Committee of Baylor College of Medicine and the Institutional Animal Care and Use of Committee of University of Pittsburgh.
Targeting construct, electroporation and ES cell selection was done as previously described –. Exons 4 and 5 of the mouse Hormad1 gene were replaced with a neomycin resistance gene flanked by 5.3 kb (5′) and 6.2 kb (3′) homologous sequences (Figure S1A). Targeted ES cells were verified by Southern blotting, and microinjected into the C57BL/6 blastocysts to produce chimeric mice that carried the mutation into the germ line. These mice were mated with C57BL/6 wild-type mice to generate Hormad1 heterozygous animals that were subsequently crossed to produce F2 offspring for functional analysis. PCR genotyping was performed using the following primers: WT-forward (5′- TCAAGACCAACCTGGGCTAC -3′) and WT-reverse (5′- CCATGTGGGTTGTAGGGAGT -3′) to amplify a 196-nucleotide wild-type band, and KO- forward (5′- TCAAGACCAACCTGGGCTAC -3′) and KO- reverse (5′- GGGGAACTTCCTGACTAGGG -3′) to amplify a 505-nucleotide mutant band (Figure S1A).
Testes were fixed in Bouin's solution (Sigma-Aldrich, MO, USA) and ovaries were fixed in 10% buffered formalin or 4% paraformaldehyde. Fixed tissues were embedded in paraffin, serially sectioned (5 µm) and stained with hematoxylin and eosin or with Periodic Acid Schiff (PAS). At least five pairs of testes and ovaries of each genotype were subjected to gross and microscopic analyses for each time point. Germ cell cysts, primordial, primary, and secondary follicles were defined as described . We used antibodies against SOHLH1 , PLZF ( sc-22839, Santa Cruz, CA,USA), DDX4 (ab13840, Abcam, MA, USA), ATM phospho-S1981 (05-740, Millipore, CA, USA), and HORMAD1 proteins. Affinity purified, anti-HORMAD1 guinea pig and rabbit antibodies, were generated against part of HORMAD1 protein (amino acids 23-373) at Cocalico Biologicals (Lancaster, PA, USA).
Oocytes and spermatocytes for chromosome analyses were prepared essentially as described previously –. Ovaries and testes were incubated in trypsin-EDTA solution at 37°C for 15 min and washed briefly in PBS. Trypsinized ovaries and testes were pipetted repeatedly and centrifuged, followed by resuspension in PBS. Cell suspensions were placed on poly-L-lysine-coated slides containing 120 mM sucrose solution and 0.05% Triton X-100. The slides were fixed in 2% paraformaldehyde and 0.02% SDS for 1 hour at room temperature, washed in distilled water and air dried, and stored at −80°C before use. Immunostaining was preformed as described –. Slides were incubated with the primary antibody overnight at 4°C, washed with PBS and incubated with Alexa-488 and Alexa-595 (Invitrogen, CA, USA) secondary antibody (1300 dilutions) for 1 hour at room temperature. After washing with PBS, slides were mounted using VECTASHIELD medium with 4,6-diamidino-2-phenylindole (DAPI) (Vector Laboratories, CA, USA). SYCP2 rabbit and guinea pig immunoaffinity-purified antibody , DMC1 (sc-8973, Santa Cruz, CA, USA), RAD51 (ab1837, Abcam, CA, USA) and RPA (ab87272, Abcam, CA, USA) were used. Dr. Christer Höög (Karolinska Institutet, Sweden) kindly provided SYCP1 and SYCP3 rabbit immunoaffinity-purified antibodies. Dr. William R. Brinkley (Baylor College of Medicine, TX, USA) kindly provided anti-CREST human serum.
Wild-type and mutant oocyte and spermatocyte spreads were stained at the same time with the same mixture of antibodies. In each experiment, when comparing wild type and mutants, imaging of the cells was performed on the same day with the same microscope and camera settings. PerkinElmer Volocity software 5.3 was used to control for possible changes in illumination during the course of imaging and measurement of the immunofluorescence. We measured total immunofluorescence of γH2AX in identical-sized rectangles that were placed over the cell boundaries. Fifty individual leptotene stage spreads from wild type, mutant spermatocytes and oocytes were subjected to immunofluorescence analysis. A two-tailed non-parametric Wilcoxon–Mann–Whitney two-sample rank-sum test was used for sample comparison.
Testes were dissected into ~0.25 cm pieces and fixed in 1x PBS with 2% formaldehyde and 3% glutaraldehyde (Ted Pella Inc., CA, USA), for 2 hr at room temperature. Samples were treated with 0.5% uranyl acetate and osmium tetraoxide, dehydrated with ethanol, and embedded in LX-112 medium (Ladd Research Industries, VT, USA). The samples were polymerized in a 70°C oven for 2 days. Ultrathin sections (70 –100 nm) were cut in a Leica Ultracut microtome (Leica, IL, USA), stained for 5 min in 1% aqueous uranyl acetate and 2 min in 1% aqueous lead citrate at room temperature in a Leica EM Stainer, and examined by a JEM 1010 transmission electron microscope (JEOL, MA, USA) at an accelerating voltage of 80 kV. Digital images were obtained using AMT Imaging System (Advanced Microscopy Techniques Corp, MA, USA).
Western blot analysis was performed essentially as described previously . Postnatal day 5, 8, 14, 18 and 21 testes were collected and lysed in RIPA buffer (50 mM Tris-HCl, pH 7.4, 1% NP-40, 0.25% sodium deoxycholate, 150 mM NaCl, 1 mM EDTA with or without protease inhibitor cocktail (Roche)). Total protein (20 µg) was loaded onto the SDS–PAGE gel, transferred to nitrocellulose membrane, and immunoblotted with HORMAD1 rabbit antibody (15000 dilutions). Varying amounts of protein phosphatase I (Sigma P7937, MO, USA) were incubated with 20 µg of 18 day old wild-type total testes lysates, for 30 min on ice. Phosphatase treated testes lysates were electrophoresed, transferred to nitrocellulose membrane and immunoblotted with anti-HORMAD1 rabbit antibody.
RT-PCR and quantitative real time PCR were performed as described previously , . We used previously published oligonucleotide sequences corresponding to Rnh2, Mov10l1, Pramel3, Usp26, Fthl17, Tex11, Tex13 , and Hormad1 (forward 5′ CTGCTGACACCAAGAAAGCA 3′- and reverse 5′- CCTGGTGGTTGGTAATCTGG -3′) and Hormad2 (forward 5′ GCTCATCAGGGGCTAGACTG 3′- and reverse 5′- TGGTTCGCTGACCTTCTTCT -3′) primers. Q-PCR was performed using the SyberGreen PCR Master Mix (Bio-Red, CA, USA) and probe set specific for each gene under investigation. Each sample was analyzed in triplicate from at least three independent wild type and Hormad1−/− testes cDNA samples. The relative amount of transcripts was calculated by the ΔΔCT method as described by Applied Biosystems, and normalized to β-actin. The average and standard errors were calculated for the triplicate measurements, and the relative amount of target gene expression for each sample was plotted. Student's t test was used to compute the P values. Significance was defined as a P value<0.05.
RNA expression arrays on wild-type and Hormad1−/− 2 week old testes were performed on the Illumina BeadChip MouseWG-6 2.0 arrays and analyzed as previously described (GEO accession numbers, GSE21524) –.
Mouse oocytes and pre-implantation embryos were collected by using standard protocols for timed mating . Briefly, 4 to 6 weeks female mice were superovulated by injection of 5 IU of pregnant mare serum gonadotropin (PMSG) (Prospec, Rehovot, Israel) and 48 h later 5 IU of human chorionic gonadotropin (hCG) (Sigma, MO, USA). Females were placed individually with 10-week old males immediately after the injection of hCG. Collection of one cell, two cell, four cell, eight cell, and blastocyst stages was performed at 24 h, 42 h, 68 h, and 96 h after hCG injection, by flushing the oviducts and uterine horns with HEPES-buffered media under the microscope. Removal of the cumulus cells was achieved in HEPES-buffered media containing 1 mg/ml hyaluronidase (Sigma, MO, USA).
Oocytes and embryos were exposed to a hypotonic solution containing 0.1% sodium citrate, 0.1% bovine serum albumin for about 15 min and gently transferred onto the slides. Cells were fixed with fresh methanol: acetic acid (31) solution dispensed carefully over the surface of the slide. The slides were air-dried and dehydrated at room temperature in a series of 70%, 85% and 100% ethanol solutions before FISH analysis.
Twenty BAC (bacterial artificial chromosomes) clones from a mouse RPCI-23 library representing sequences unique to specific chromosomes were selected from the UCSC Genome browser (http://genome.ucsc.edu) and NCBI (http://www.ncbi.nlm.nih.gov) databases (Table S1). Probe DNA extraction was performed according to the standard alkaline lysis method and labeled by standard nick translation with Spectrum Orange- or Spectrum Green-dUTP using a commercially available kit (Abbott/Vysis, IL, USA). Sequential fluorescence in situ hybridization experiments using a mixture of three probes were performed according to manufacturer's instructions (Abbott/Vysis, IL, USA). Each mixture contained a red, green and yellow (combined green/red) fluorescent probes. Probes were applied to slides, hybridized for 20 hr at 37°C, washed with 0.4×SSC/0.3% NP-40 for 2 minutes at 73°C and with 2×SSC/0.1% NP-40 for 1 minute at room temperature, and counterstained with DAPI. Digital FISH images were captured by a Power Macintosh G3 System using MacProbe software version 4.4 (Applied Imaging, CA, USA).
Generation of Hormad1−/− Mice. (A) Disruption of the Hormad1 locus. The targeting construct and schematized genomic locus of Hormad1 are shown. The neo cassette replaced HORMA domain encoding highly conserved exons 4 and 5, and introduced an Xba I restriction enzyme site into the locus. This Xba I site was used as a diagnostic for Southern blot analysis of ES cells electroporated with the targeting vector (data not shown). Arrows indicate genotyping primers used to distinguish wild-type and mutant alleles in the transgenic animals. (B) RT-PCR analysis of Hormad1 knockout mice shows lack of transcript corresponding to Hormad1 in the knockout animals. A faintly visible lower molecular weight band in the knockout (−/−), corresponds to a mutant transcript without exons 4 and 5. Removal of exons 4 and 5 causes a frameshift mutation. (C) Total protein was isolated from testes of 2 week old wild-type (+/+), Hormad1−/− (−/−) and Hormad1+/− (+/−) mice and Western blot analysis was performed with anti-HORMAD1 specific antibody. No significant amount of HORMAD1 was detected. The β-actin signal serves as a loading standard.
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Adult testes are atrophied in Hormad1−/− Mice. (A) Reduced testicular size and weight in Hormad1−/− (KO) as compared to the wild-type (WT) testes from 4 week old male siblings. (B) Ratio of testes/body weight from wild-type (WT) and Hormad1−/− (KO) testes. Error bars represent the standard error of mean. Student's t test was used to calculate P values.
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Excess apoptosis and increased apoptotic index in Hormad1−/− testes. (A-E) TUNEL assay in 1, 2, 3, 4, and 6 week old wild-type mouse testes. (F-J) TUNEL assay in 1, 2, 3, 4, and 6 week old Hormad1−/− mouse testis. (K) Apoptotic index was calculated in wild-type (WT) and Hormad1−/− mouse testis (KO) mice. Knockout testes had significantly higher apoptotic index at 2 weeks of post-natal life and beyond. Error bars represent the standard error of the mean. Error bars represent the standard error of the mean. Student's t test was used to calculate P values. P value was less than 0.001 at every time point except one week. Scale bar: 100 µM.
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No germ cell loss in the Hormad1−/− ovary. At post-natal day 8, primordial, primary, and secondary follicles were detected in wild-type (A) and the Hormad1−/− ovaries (B). Anti-Lhx8 antibody was used to detect germ cells (brown stain). Histogram represents primordial follicles (PF), primary follicles (PrF), and a total number of secondary follicles (SF) in the wild-type (WT) and mutants (KO) (C). Every fifth section of wild-type (n=6) and Hormad1−/− ovaries (n=5) were counted. Error bars represent the standard error of the mean. Fisher's exact t test was used to calculate P values. P value between mutant and wild-type primordial follicles, primary, and secondary follicles was >0.5, and therefore not statistically significant. Bars, 50 µm.
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BAC clones used in FISH experiments.
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We thank Hyo won Ahn for technical assistance, Hitomi Suzuki and Krishna Jagarlamudi for critical thinking, Tianjiao Chu for statistical analysis of the RNA expression array data, and Christer Höög and William R. Brinkley for supplying antibodies.
The authors have declared that no competing interests exist.
This work was supported in part by the National Institutes of Health Grant HD054829 to AR (http://grants.nih.gov/grants/oer.htm). The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.