Regulation of an RNA granule during spermatogenesis: acetylation of MVH in the chromatoid body of germ cells

The differentiation program of germ cells is characterized by highly specialized mechanisms of gene expression that operate at transcriptional and post-transcriptional levels (Kotaja and Sassone-Corsi, 2007; Maclean and Wilkinson, 2005; Rolland et al., 2008; Sinha Hikim et al., 2003). Following a wave of post-meiotic transcription, a large proportion of mRNAs encoding products required for late spermatogenesis remain untranslated for 4–5 days. This post-transcriptional regulation includes alteration in poly(A) and 3′-UTR length, a process elicited by regulatory RNA-binding proteins and involving microRNA (miRNA-mediated gene repression (Kashiwabara et al., 2002; Zhong et al., 1999). Interestingly, Dicer-deficient mice show several developmental defects, including spermatogenesis arrest (Hayashi et al., 2008; Kanellopoulou et al., 2005). In addition, another class of small RNAs, named PIWI-interacting RNAs (piRNAs), is germline specific (Siomi and Kuramochi-Miyagawa, 2009). Although evidence indicates that post-transcriptional regulation by small RNAs is essential in germ cell differentiation, the signaling pathways governing these events remains largely unclear.

Germ cells from many organisms have a unique cloud-like structure, often referred to as nuage, where translational regulation is thought to take place (Styhler et al., 1998). Post-meiotic mammalian germ cells have a cytoplasmic, perinuclear structure called the chromatoid body (CB), which is thought to be the counterpart of nuage (Parvinen, 2005). We have previously shown that the CB shares many structural and molecular properties with the somatic processing bodies (P-bodies). Indeed, the CB contains several RNA-induced silencing complex (RISC) components (Kotaja et al., 2006a). Only mature miRNAs, and not pre-miRNAs, are located in the CB, and in accordance Dicer also is located in the CB (Kotaja et al., 2006a). Most importantly, argonaute family proteins, which constitute the central catalytic engine of RISC, and poly(A)-containing mRNAs, are also located in the CB (Kotaja et al., 2006a). Therefore, we have proposed that the CB constitutes a crucial structure where miRNA-mediated gene repression takes place during late stages of spermatogenesis (Kotaja et al., 2006a). In addition to miRNA RISC components, mouse PIWI (MIWI) and MIWI-like (MILI) proteins are also concentrated in the CB (Kotaja et al., 2006b), suggesting that the CB has a role in piRNA-mediated transposable element repression.

The protein Vasa [in the mouse MVH (mouse Vasa homologue)] is expressed exclusively in germ cells and is an essential component of the nuage and CBs. Vasa (MVH) is an ATP-dependent DEAD-box RNA-binding protein able to regulate RNA–protein interactions. Its structure and intracellular location are conserved from Drosophila to mammals (Tanaka et al., 2000). MVH interacts with both miRNA- and piRNA-pathway proteins, such as Dicer, MIWI and MILI (Kotaja et al., 2006a; Kuramochi-Miyagawa et al., 2004). In addition, Mvh has been reported to have a role in piRNA-mediated transposon repression (Kuramochi-Miyagawa et al., 2010). MVH has been postulated to regulate germ-cell-specific small RNA-mediated (miRNA and/or piRNA) gene repression regulation (Kotaja et al., 2006a). MVH is essential for germ cell differentiation as demonstrated by loss-of-function approaches (Tanaka et al., 2000).

We have suggested that the dynamic composition and function of the CB indicates that germ-cell-specific intracellular signaling and post-translational modifications are probably involved in its control (Kotaja and Sassone-Corsi, 2007). Here we show that histone acetyl transferase 1 (Hat1) and its co-factor p46 are specific components of the CB, where they localize in a time-regulated manner. To explore the role of MVH, we have identified 858 mRNAs associated with the MVH–ribonucleoprotein (RNP) complex. Importantly, MVH is acetylated in a Hat1-dependent manner, an event that leads to inhibition of MVH RNA-binding capacity. Our results indicate the presence of a developmentally regulated mechanism in which Hat1 and p46 control the physiology of the CB by acetylation of MVH.

Most HATs are located in the nucleus where they acetylate histones and contribute to chromatin remodeling. A uniquely cytoplasmic B-type HAT is Hat1, the function of which has been linked to acetylation of some cellular proteins, including histone H4 before its translocation to the nucleus. We sought to determine the intracellular location of Hat1 in male germ cells, and performed a detailed analysis at different times of the differentiation program by using Hat1-specific antibodies. The results revealed that Hat1 is strongly concentrated in the CB in a temporally regulated manner, during stages IV to VI of germ cell development (Fig. 1A).

The location of Hat1 prompted us to determine whether CB components are acetylated in a stage-specific manner. Several CB-localized proteins, including native MVH, are acetylated in male germ cells. Western blot analysis (WB) using a pan-acetyl-lysine antibody showed specific acetylation of the immune-purified MVH but not of the control IgG (Fig. 1B, lanes 2 and 3). Because Hat1 is highly concentrated in the CB during stages IV to VI, we examined MVH acetylation in the different developmental stages. MVH was immunoprecipitated from extracts taken at each stage, followed by WB using a pan-acetyl-lysine antibody. Consistent with the Hat1–MVH colocalization, MVH is acetylated specifically at stages IV to VI (Fig. 1C, lane 2, compare with lanes 1, 2 and 4). Finally, we checked developmental expression of MVH, Hat1 the Hat1 activator p46, and cyclic AMP responsive element (CREM). Unlike CREM, the expression of which is developmentally regulated, the levels of these three proteins are mostly constant (Fig. 1C,D). Thus, the developmentally regulated acetylation of MVH appears to be determined by its CB-targeted localization rather than by its expression levels.

The stage-specific colocalization of Hat1 and MVH in the CB strongly suggested that Hat1 is responsible for MVH acetylation. To test this possibility, we used immune-purified FLAG–MVH that was ectopically expressed with or without Myc–Hat1 and/or Myc–p46 in HeLa cells. Our results indicate that FLAG–MVH becomes readily acetylated by the combined expression of Myc–Hat1 and Myc–p46, but not by either Myc–Hat1 or Myc–p46 alone (Fig. 2A, lanes 1–4). We also detected some MVH acetylation in the absence of exogenous Myc–Hat1 or Myc–p46 (Fig. 2, lane 1). This is most probably caused by endogenous Hat1 and p46 and/or other acetyltransferases.

Next, we used the enzymatically inactive mutant Hat1 mt, in which a single, conserved amino acid essential for acetyltransferase activity is mutated. Coexpression of Myc–Hat1 mt was unable to mediate efficient FLAG–MVH acetylation, even in the presence of Myc–p46 (Fig. 2A, lanes 5 and 6). Finally, we also determined that other CB components were not acetylated by Hat1 and p46 (supplementary material Fig. S1), underscoring the specificity of Hat1 in targeting MVH.

To verify whether Hat1 directly acetylates MVH, we performed in vitro acetylation assays with purified proteins. M2 agarose were used to purify FLAG–Hat1 or FLAG–Hat1 mt with or without Myc–p46 after ectopic expression in HeLa cells. The activity of the purified Hat1 and p46 proteins was validated by incubation with core histones in the presence of [¹⁴C]acetyl-CoA. FLAG–Hat1 in combination with Myc–p46 showed a strong HAT activity, whereas the mutant Hat1 mt with Myc–p46 showed no HAT activity (Fig. 2B). Next, immune-purified FLAG–Hat1 and Myc–p46 or FLAG–Hat1 mt and Myc–p46 were incubated with GST–MVH. GST–MVH is readily acetylated by FLAG–Hat1 and Myc–p46 (Fig. 2C, lane 2) but not by FLAG–Hat1 mt and Myc–p46 (lane 3), thus confirming that Hat1 directly acetylates MVH.

As p46 is required for MVH acetylation by Hat1, we explored whether p46 also colocalizes with Hat1 and MVH in the CB. Our analysis showed that p46 is highly concentrated in the CB at stages IV to VI, in a coordinated manner with MVH and Hat1 (Fig. 1A, Fig. 2D). The colocalization is consistent with the timing of MVH acetylation (Fig. 1C). We quantified both Hat1 and p46 signals (supplementary material Fig. S2). Thus, our results support a scenario in which the combined action of Hat1 and p46 within the CB leads to a stage-specific acetylation of MVH.

We sought to identify the lysine(s) in MVH that are acetylated by Hat1. To do so, we first generated various MVH deletion mutants and then examined their acetylation state upon coexpression with Hat1 and p46 (Fig. 3A). The 1–496 mutant, but not the 1–346 or the Δ284–502 mutants, was significantly acetylated by Hat1. The efficacy of interaction of both the 1–346 and Δ284–502 mutants to Hat1 and p46 is similar to the full length MVH and to the 1–496 mutant (Fig. 4E), implying that the acetylated lysine(s) is located within this region (amino acids 347–496; Fig. 3A). This region contains nine lysine residues. We generated individual lysine to arginine mutants for each of the nine potential target residues. Among these mutants, only FLAG–MVH K405R showed no acetylation in the presence of Hat1 and p46 (Fig. 3B), indicating that K405 is a prominent target for Hat1-directed acetylation. Quantification further determined that acetylation of FLAG–MVH was substantially induced by Hat1, whereas FLAG–MVH K405R was not (Fig. 3B). Sequence alignment analysis revealed that K405 and its surrounding amino acids are conserved in all sequenced Vasa proteins, except for the Drosophila counterpart (Fig. 3C). Although more experiments are needed to elucidate the situation in the fly, it is noteworthy that human Vasa is listed in a high-throughput global analysis of 3600 lysine acetylation sites obtained by high-resolution mass spectrometry on 1750 proteins. The human Vasa is listed as acetylated at K431, which is the site corresponding to mouse K405 (Choudhary et al., 2009), thus validating our mutagenesis analysis. Finally, we generated an antibody that recognizes MVH only when acetylated in position K405 (anti-AcMVH). Specificity of this antibody was demonstrated by detection of acetylated FLAG–MVH wild-type (wt) upon coexpression in cultured cells with Myc–Hat1 and p46, and subsequent immunoprecipitation. A FLAG–MVH K405R mutant protein monitored under the same conditions was not recognized by the AcMVH antibody (supplementary material Fig. S3). Using this antibody we also confirmed that MVH is acetylated specifically at stages IV to VI (Fig. 3D). Thus, our data strongly indicate that K405 is the major Hat1-dependent acetylation site on MVH.

The acetylation of MVH suggested that it could physically interact with the Hat1–p46 complex. First, we sought to determine whether the endogenous proteins in the testis would be associated in a complex. To explore this possibility, we performed coimmunoprecipitation assays using testis extracts and demonstrated that native Hat1 and MVH proteins readily interact (Fig. 4A, lane 3, compare with lane 2). As native p46 has the same mobility as IgG heavy chain, the in vivo interaction between p46 and MVH could not be assessed. Moreover, we transiently expressed and immune-purified FLAG–MVH, Myc–Hat1 and/or Myc–p46 from cultured cells. We found that FLAG–MVH co-precipitated with Myc–Hat1 or Myc–p46 using an anti-Myc antibody for the pull-down (Fig. 4B, lanes 2 and 3, compare with lane 1). Next, we wanted to confirm that both Myc–Hat1 and Myc–p46 reciprocally coimmunoprecipitate with FLAG–MVH. Indeed, both Myc–Hat1 and Myc–p46 co-precipitated with FLAG–MVH (Fig. 4C, lanes 5 and 6, compare with lane 4). Interestingly, Myc–Hat1 appears to inhibit the interaction between FLAG–MVH and Myc–p46 (Fig. 4C, lane 8), an effect not observed using an enzymatically inactive mutant of Myc–Hat1 (Fig. 4D, lane 3, compare with lanes 2 and 4). Finally, we used a series of MVH deletion mutants (Fig. 3A) to identify the regions of MVH involved in the Hat1 and p46 interactions. We found that the MVH 1–346, 1–496 and Δ284–502, but not 1–199, bind to both of Myc–Hat1 and Myc–p46 (Fig. 4E, lanes 4–6, compare with lanes 1–3).

To determine the role of Hat1-mediated acetylation, we explored whether it affected the RNA binding, ATPase activity and ATP binding of MVH. We used acetylated GST–MVH or acetylated GST–MVH Δ1–199 (Fig. 2C). GST–MVH Δ1–199 or acetylated GST–MVH Δ1–199 was incubated with [α-³²P]ATP followed by UV cross-linking. The protein–[α-³²P]ATP mixtures were separated by SDS-PAGE. The ATP-binding activity of MVH Δ1–199 was unaffected by Hat1 (Fig. 5A, lane 4, compare with lane 3). Notably, FLAG–Hat1 with Myc–p46 did not alter ATP-binding activity of the GST–MVH Δ1–199 K405R mutant (lanes 5 and 6) or that of GST–MVH Δ1–199. Unlike ATP-binding activity, acetylated GST–MVH showed a decreased RNA-binding activity when compared with unacetylated GST–MVH (Fig. 5A, lane 4, compare with lane 3). By contrast, FLAG–Hat1 did not inhibit RNA-binding activity of the GST–MVH K405R mutant (lanes 5 and 6). Next, we measured ATPase activity by coupled pyruvate kinase–lactate dehydrogenase assay. It is known that the ATPase activity of many DEAD-box RNA-binding proteins is stimulated by RNA (Liang et al., 1994). Drosophila Vasa, however, was reported to contain RNA-independent ATPase activity (Liang et al., 1994). Thus, we speculated that MVH could have RNA-independent ATPase activity, and FLAG–Hat1-mediated acetylation might not inhibit GST–MVH ATPase activity. Indeed, FLAG–Hat1 failed to inhibit GST–MVH wt and K405R ATPase activity (Fig. 5B). Quantification of three independent experiments are shown (Fig. 5C). To confirm the RNA dependency of GST–MVH ATPase activity, we measured GST–MVH ATPase activity in the presence or absence of yeast tRNA. The results showed that yeast tRNA fails to activate GST–MVH ATPase activity (Fig. 5D). These data support the observation that Hat1-mediated MVH acetylation has no effect on its ATPase activity. Taken together, we demonstrated that Hat1-mediated MVH acetylation causes a considerable decrease in RNA-binding potential, but has no effects on its ATP binding and ATPase activity.

To identify the mRNAs targeted by MVH we purified the MVH–RNP complex using anti-MVH antibodies, followed by co-precipitation of its associated RNAs. An equal amount of IgG was used as a negative control. Co-precipitated RNAs were analyzed by DNA chip. We identified 858 distinct mRNAs (with a score at least twofold as compared with control IgG) that were classified into functional groups using GeneCodis (http://genecodis.dacya.ucm.es/; Fig. 6A; supplementary material Table S1) (Nogales-Cadenas et al., 2009). Of these, 34 corresponded to genes known to play a crucial role in spermatogenesis and/or testis specific expression. Although the spermatogenesis category is not the largest, it is the one with best score of statistical value by hypergeometric distribution. For example, DDX25 mRNA, which co-precipitates with MVH–RNP (Fig. 6B), is essential for mouse spermatogenesis (Tsai-Morris et al., 2004). Additional transcripts encode proteins that could be implicated in the process of spermatogenesis. For example, ~10% of the mRNA targets correspond to genes involved in energy metabolism, including the gene encoding hexokinase 1 (HK1) that phosphorylates glucose during the initial step of glycolysis (Fig. 6B) (Kodde et al., 2007). Another interesting example is carnitine acetyltransferase (Crat), the activity of which has been correlated with sperm fertility (Golan et al., 1984). Most interestingly, many translational regulators that undergo strict regulation at late stages of spermatogenesis (Kimura et al., 2009) were identified as MVH target mRNAs, such as eukaryotic initiation factor 4B (eIF4B) and signal recognition particle 54 (Srp54). The ATPase activity of eIF4A is stimulated by eIF4B, which thus facilitates the translation of mRNAs encoding unique secondary structures within their 5′-UTRs (Shahbazian et al., 2010). Srp54 contributes to signal peptide recognition when the peptide is released from ribosomes and enhances its translocation to the endoplasmic reticulum (Wild et al., 2004). Interestingly, Tnp1 mRNA, well known to be translationally arrested, did not co-precipitated with MVH, suggesting that MVH selectively binds to mRNAs (Fig. 6B). Next, we examined the developmental association between MVH and its target mRNAs. Association of eIF4B mRNA with MVH is developmentally regulated, being low at stages IV to VI compared with stages I to III, whereas expression was significantly (P<0.05) increased at stages IV to VI compared with stages I to III (Fig. 6C; supplementary material Fig. S4). This observation could reflect the high acetylation levels of MVH at stages IV to VI, which parallels the decrease in RNA binding (Fig. 1C; Fig. 5A). However, association with Srp54 mRNA was constant as was the mRNA expression (Fig. 6C; supplementary material Fig. S4). This could indicate that MVH acetylation affects MVH RNA-binding selectively. Considering that mRNAs in RNA granules are not actively translated and that MVH is strongly concentrated in the CB (Anderson and Kedersha, 2009), the weaker association between MVH and eIF4B mRNA could explain why eIF4B is actively translated at later stages. To examine this, we collected tubules at various developmental stages and examined protein expression. Although eIF4B levels increase at stages VII to IX, Srp54 levels were stable (Fig. 6D). Finally, we compared previously reported translationally arrested genes and MVH targets. As shown in Fig. 6E, among 652 known translationally arrested genes, with expression that was detectable at all stages [detectable at 17 days (post partum), 22 days and in adults, by DNA chip], only 30 genes (3.56%) were overlapping with MVH targets. However, among the 34 meiotic (detectable at 17 days and 22 days by DNA chip; e.g. Pgk2 and Crat; Fig. 6B; supplementary material Table S1) and 96 post-meiotictranslationally arrested genes (detectable at 22 days and in adults by DNA chip), 8 (23.5%) and 14 (10.4%) are MVH target mRNAs, respectively. Thus, our data suggest that MVH selectively contributes to translation efficiency of many mRNAs, a regulation that seems to depend on its degree of acetylation (e.g. eIF4B translation) at later stages of spermatogenesis.

The involvement of RNA granules in the control of gene expression is demonstrated (Balagopal and Parker, 2009), but little is known about how components of RNA granules might be able to integrate physiological responses and intracellular signaling. The case of germ cells is particularly intriguing because modulation of the CB could have consequences for the timing of translational repression that characterizes these cells post-meiotically. The DEAD-box RNA-binding protein MVH is a crucial component of the CB, is expressed uniquely in germ cells and is essential for germ cell differentiation (Tanaka et al., 2000). Our findings indicate that acetylation of MVH constitutes an important control mechanism.

We have shown that MVH is acetylated by Hat1 at Lys405 in a p46-dependent manner (Figs 2, 3), and that this modification results in inactivation of its RNA-binding activity, whereas ATP binding and ATPase activity remain unaltered (Fig. 5). In addition, both Hat1 and p46 colocalize with MVH in the CB at seminiferous epithelium cycle stages IV to VI, when MVH is coordinately acetylated (Figs 1, 3). Because the CB concentrates a remarkable array of components (Kotaja and Sassone-Corsi, 2007), we speculate that a high-order organization of RNA–protein complexes must exist within the CB. MVH is likely to play a key role in the organization of the CB. Indeed, MVH is essential in the control of germ cells differentiation (Tanaka et al., 2000) and has been shown to associate with several CB components (see below). Among these, MVH physically interacts with MIWI, and mutation of the Miwi gene in the germline results in disruption of the CB organization (Kotaja et al., 2006a; Kuramochi-Miyagawa et al., 2004).

Our findings pave the way to further experimentation to elucidate the role of MVH in germ cells, and specifically to the regulatory function of its acetylation. The CB contains several RNA-induced silencing complex (RISC) components such as miRNAs, Dicer and Argonaute 2 (Ago2), in addition to mRNAs (Kotaja et al., 2006a). Interestingly, MVH interacts with Dicer, AU-rich elements (ARE) binding protein (HuR) and ARE-containing mRNA (Kotaja et al., 2006a; Nguyen Chi et al., 2009). This raises the possibility that acetylation modulates MVH function at various levels.

In addition to its implication in the miRNA pathway, MVH has been reported to interact with PIWI family proteins MIWI and MIWI-like (MILI), both in mammals and in Drosophila (Kuramochi-Miyagawa et al., 2004; Megosh et al., 2006) and it interacts with piRNA in mouse testis (Kirino et al., 2010). The role of PIWI proteins is carried out through the biogenesis of a specialized class of small RNA, the piRNAs (Siomi and Kuramochi-Miyagawa, 2009). piRNA-mediated control is thought to repress transposable elements in the germline (Siomi and Kuramochi-Miyagawa, 2009). Interestingly, Mvh-deficient mice show impaired piRNA expression and de-repressed transposable elements (Kuramochi-Miyagawa et al., 2010). Considering that both MILI and MIWI are localized to the CB, it is conceivable that MVH might operate as a putative piRNA pathway component in an acetylation-dependent manner.

To determine the function of MVH in the CB, we identified 858 target mRNAs by DNA chip. An important fraction of these is either directly or indirectly involved in the spermatogenesis process. There is evidence to indicate that mRNAs located in RNA granules are in an inactive state for translation and/or stored to be translated later (Balagopal and Parker, 2009). mRNAs in the MVH–RNP complex in the CB also seem to be stored without entering translation, a process followed by the release of the mRNA from the MVH–RNP in the CB to the cytoplasmic polysome. Indeed, 652 mRNAs detectable at all stages (17 days, 22 days and adult) show significant shift from translationally inactive RNPs to translationally active polysomes as spermatogenesis (Iguchi et al., 2006) and 30 of the translationally arrested genes (4.6%) do indeed overlap with the MVH target (Fig. 6E), in accordance to the above hypothesis. Indeed, Crat, which is a translationally arrested gene, is an MVH target in vivo (Fig. 6B) (Iguchi et al., 2006). Notably, among 34 meiotic (detectable at 17 days and 22 days by DNA chip) and 96 post-meiotic translationally arrested genes (detectable at 22 days and in adult animals by DNA chip), 8 (23%) and 14 (14%) genes are targets of MVH (Fig. 6E). Importantly, the translationally arrested Tnp1 transcript is not an MVH target (Fig. 6B), suggesting that MVH binds meiotic and/or post-meiotic translationally arrested mRNAs in a selective manner.

We have determined that MVH acetylation determines the efficacy of RNA binding (Fig. 5A). This finding parallels the developmental correlation between MVH acetylation, eIF4B mRNA release and active translation (Fig 1C; Fig. 6C,D; supplementary material Fig. S4). Regulation appears to be specific, because interaction with Srp54 mRNA remained constant during development. These observations fit the scenario in which release of eIF4B mRNA from MVH–RNPs is followed by active translation. Therefore, we propose the following temporally regulated processes: (1) MVH binds to many mRNAs, including translationally arrested mRNA, to be stored as inactive RNPs in the CB (stages I to III and/or before); (2) a first subset of MVH target mRNAs are released from MVH–RNP and/or the CB to allow translation, a process dependent on MVH acetylation (stage IV to VI); (3) the rest of the MVH target mRNAs are released from MVH–RNP, their translation being independent of MVH acetylation (stage VII to X); (4) released mRNAs are actively translated (stage VII to XII).

Our findings reveal a previously unforeseen level of regulation within an RNA granule. Importantly, acetylation of MVH occurs in a highly controlled temporal manner (Fig. 1), suggesting the possibility that it is directed by specific pathways of intracellular signaling. It is noteworthy that Hat1, the acetyltransferase implicated in modifying MVH, has been previously shown to be involved in stress responses and specific metabolic pathways (Benson et al., 2007; Pogribny et al., 2007). This could suggest that Hat1 function in germ cells is under endocrine or metabolic control, possibly through somatic cells-originated pathways that are known to contribute to the germ cell differentiation program (Sofikitis et al., 2008).

pGEX4T1-MVH, pcDNA3-FLAG-MVH and deletion mutants were described elsewhere (Kotaja et al., 2006a). A BamHI–XhoI fragment of MVH mutant (1–346) was inserted in the corresponding sites of pcDNA3-FLAG, generating pcDNA3-FLAG-MVH 1–346. An EcoRI–NotI fragment of MVH Δ1–199 wt and K405R was inserted into the corresponding site of pGEX4T3, generating wt and K405R mutants of pGEX4T3-MVH Δ1–199. Point mutated pcDNA3-FLAG-MVH mutants were created using the QuickChange site-directed mutagenesis kit (Stratagene). Western blot analysis was described elsewhere (Kotaja et al., 2006a) with a following modification: coimmunoprecipitated Myc–p46 was probed with a Myc antibody and an HRP-conjugated secondary antibody. The HRP on the secondary antibody was inactivated by sodium azide. The same blot was probed with an anti-Hat1 antibody. Anti-MVH and anti-CREM antibodies were previously described (Kotaja et al., 2006a). Anti-pan-acetyl-lysine antibody and anti-MVH K405 acetylated antibody (Millipore), anti-Hat1 antibody and anti-p46 antibody (Santa Cruz Biotechnologies), anti-FLAG M2 antibody (Sigma), anti-Myc antibody (Abcam) were used following the manufacturer's indications. Immunofluorescence was described elsewhere (Kotaja et al., 2006a).

HeLa cells were transfected with the indicated plasmids using BioT (Bioland Scientific). The cells were collected 2 days after transfection and lysed in TNEN [50 mM Tris-HCl pH 8.0, 150 mM NaCl, 5 mM EDTA, 0.5%NP40 1 mM dithiothreitol (DTT), supplemented with protease inhibitor cocktail (Roche), 5 mM NaF, 5 mM PMSF and HDAC inhibitors]. Cell lysates were mixed with M2 agarose (Sigma) or Myc antibody with protein G agarose (Sigma) and, after centrifugation, were incubated overnight at 4°C. The immunoprecipitated products were separated by SDS-PAGE followed by western blotting with the indicated antibodies after extensive washes. Lysates from decapsulated testes were mixed with protein A agarose (Sigma) and unbound fractions were mixed with anti-MVH antibody or equivalent control rabbit IgG. After overnight incubation, MVH antibody was precipitated with protein A agarose. After extensive washes, the immunoprecipitated products were separated by SDS-PAGE followed by western blotting with the indicated antibodies. GST–MVH proteins were expressed in BL21 and then purified on glutathione–Sepharose 4 beads (Amersham). After three washes with cold TNEN, GST–MVH proteins were eluted with elution buffer [5 mM reduced glutathione (Sigma) in TNEN]. Eluted GST–MVH proteins were mixed with equal amounts glycerol and stored at −30°C. All animal experiments were performed according to approved guidelines.

Overexpressed FLAG–Hat1 and Myc–p46 were immune-purified using M2 agarose and then mixed with GST–MVH Δ1–199 or full length or core histones in the presence of 0.1 mM acetyl-CoA or [¹⁴C]acetyl-CoA, respectively, at 33°C for 1 hour. Acetylated histones were spotted on p81 filter paper and activity was detected by scintillation counting. Acetylated GST–MVH was separated by SDS-PAGE followed by western blotting with pan-acetyl-lysine antibody.

Acetylated or unacetylated MVH Δ1–199 or full length proteins were prepared as shown above. For ATP-binding assay, GST–MVH Δ1–199 wt and K405R was incubated with [α-³²P]ATP in the presence of 2 mM MgCl₂ and 100 ng/ml yeast tRNA for 30 minutes on ice. This protein–[α-³²P]ATP mixture was crosslinked on Parafilm by UV radiation (780 mJ/cm²). Crosslinked protein mixtures were separated using 6% SDS-PAGE. The gel was dried then exposed to X-ray film. For RNA-binding assays, 5′-extended RNA was transcribed using the Megashortscript kit (Ambion) and purified by phenol–chloroform extraction and ethanol precipitation. Purified 5′ extended RNA (Jaramillo et al., 1991) was dephosphorylated with calf intestinal alkaline phosphatase (Fisher Scientific) and then end-labeled using polynucleotide kinase (New England Biolabs) with [γ-³²P]ATP. RNA 5′ extension was perfumed as described elsewhere (Jaramillo et al., 1991). Acetylated or unacetylated GST–MVH proteins were incubated with end-labeled 5′-extended RNA in the presence of 2 mM MgCl₂ and 0.5 mM ATP for 30 minutes at room temperature. These protein–RNA mixtures were crosslinked and exposed to X-ray film. ATPase assays were also performed on these mixtures, as previously described (Sengoku et al., 2006).

Two testes were sheared using 18 G and 25 G needles in 1 ml PBS(–). After centrifugation at 1000 g (4°C) the supernatant was discarded. The testicular cell pellets were lysed in 1 ml of MVH immunoprecipitation (IP) buffer [10 mM Tris pH 7.5, 200 mM NaCl, 2.5 mM MgCl₂, 0.5% NP40, 0.1% Triton X-100, supplemented with protease inhibitor cocktail (Roche), 1 mM DTT, 5 mM NaF and 40 U/ml RNasin) and rocked for 30 minutes at 4°C. After centrifugation at 16,200 g for 15 minutes (4°C), the supernatant was mixed with 20 μl of protein A–Dynal beads (Invitrogen) for 1 hour at 4°C. An aliquot of the supernatant (400 μl) was then mixed with 10 μl normal rabbit IgG or 10 μl anti-MVH/DDX4 antibody (Abcam). After rocking overnight at 4°C, 20 μl of protein A–Dynal beads were added and rocking continued for 1 hour at 4°C. After five washes with 1 ml MVH IP buffer for 5 minutes each, co-precipitated RNA was extracted by TRIzol reagent. DNA chip analysis was performed by TORAY (TORAY, Tokyo, Japan). cDNA was reverse transcribed using SuperScript II with oligo(dT) primer, followed by ethanol precipitation. The cDNA was used as a template for qPCR.

Squash preparations of cells were fixed in 4% paraformaldehyde, and immunofluorescence was performed using specific polyclonal antibodies and Alexa Fluor 488 and Cy3 secondary antibodies (Molecular Probes).

We thank Bruce Stillman (Cold Spring Harbor Laboratory) and Witold Filipowicz (FMI, Basel) for the p46 and the GST–Dicer expression vectors, respectively. We also thank A. Annunziato (Boston College), D. Mishra Prasad, Noora Kotaja and all colleagues in the Sassone-Corsi laboratory for discussions and help.