Unbiased proteomic screen for binding proteins to modified lysines on histone H3

2.2 PHF2 antibody

PHF2 antiserum was generated by immunizing rabbits with purified recombinant GST-PHF2 C-terminal fragment corresponding amino acids 830–1098 of PHF2.

2.3 HeLa nuclear extracts

HeLa S3 cells were purchased from National Cell Culture Center (Minneapolis, MN). The cytoplasmic (S100) and nuclear extracts (NE) were prepared as previously described [20, 21]. Briefly, HeLa cells were washed in cold PBS, and resuspended in five times the packed cell volume (PCV) with hypotonic buffer (10 mM Tris, pH 7.3, 1.5 mM MgCl₂, 10 mM KCl, 10 mM β-mercaptoethanol, and 0.5 mM PMSF). The cells were incubated on ice for 10 min and then pelleted by centrifugation at 1900×g for 10 min. The swollen cell pellet was then resuspended in half the PCV with hypotonic buffer and homogenized with 15 strokes in a glass dounce homogenizer. The lysate was centrifuged at 5000×g for 20 min at 4°C to separate the cytoplasmic proteins from the nuclear pellet. The nuclear pellet volume (NPV) was determined and the pellet was resuspended in 0.5 mL of extraction buffer (20 mM Tris, pH 7.3, 1.5 mM MgCl₂, 0.2 mM EDTA, 25% glycerol, 10 mM β-mercaptoethanol, and 0.5 mM PMSF) per milliliter of NPV. The nuclei pellet was then homogenized by ten strokes in a glass dounce homogenizer. While stirring, 0.5 mL of extraction buffer containing 1.2 M KCl was added per milliliter of NPV drop wise to the homogenized nuclear extract. The extract was further stirred for 30 min at 4°C. The sample was then centrifuged for 30 min at 20 000×g. The NE supernatant was dialyzed against dialysis buffer (20 mM Tris, pH 7.3, 100 mM KCl, 0.2 mM EDTA, 20% glycerol, 10 mM β-mercaptoethanol, and 0.5 mM PMSF) and then centrifuged at 20 000×g for 30 min. The NE supernatant was aliquoted, snap frozen in liquid nitrogen, and stored at −80°C.

2.4 Peptide pull-down

The peptides were prebound to streptavidin agarose beads (Invitrogen, Carlsbad, CA) in 50 μL of NETN buffer (20 mM Tris, pH 8, 1 mM EDTA, and 0.5% NP-40) containing 100 mM NaCl. Five milligrams of HeLa nuclear extracts was added to each of the peptide-bound agarose beads and rotated for 5 h at 4°C. The beads were then washed five times with 1 mL of NETN buffer containing 200 mM NaCl. The washed beads were boiled with 60 μL of Laemmli buffer, analyzed by SDS-PAGE on a 4–20% gel (Invitrogen), and stained with colloidal CBB.

2.5 In-gel digestion and MS

Sample preparation and MS analysis on a Finnigan LTQ mass spectrometer (Thermo Finnigan, San Jose, CA) were as previously described in ref. [22]. Briefly, each lane of the gel was divided into eight equal size gel slices. The gel slices were destained with 50 mM ammonium bicarbonate in 50% methanol and washed overnight with HPLC grade water. The gel slices were then crushed and digested with 200 ng of trypsin in 50 mM ammonium bicarbonate at 37°C for 4 h. Digested peptides were extracted with 200 μL of ACN and dried by speed vac. Each sample was dissolved in 20 μL of 5% methanol/95% water/0.1% formic acid and analyzed on a C₁₈ column (100 mm×75 μm, 300 Å pore diameter, PicoFrit; New Objective, Woburn, MA). The mobile phase A (0.1% formic acid in water) and B (0.1% formic acid in methanol) was used with a gradient of 10–80% mobile phase B over 10 min followed by 80% B for 10 min at a flow rate of 200 nL/min. Peptides were directly electrosprayed into the mass spectrometer using a nano-spray source. The LTQ was operated in a data-dependent mode, acquiring fragmentation spectra of the 20 strongest ions.

2.6 Database search and protein identification

Raw MS data were searched using BioWorks 3.2 Sequest software (Thermo Electron, Waltham, MA) against the NCBI human protein RefSeq database. Results from the eight bands for each pull-down/MS were loaded into the multi-consensus files with the following filter threshold settings: XCorr of 1.50 (1+), 1.80 (2+), and 3.00 (3+) and probability of 1 × 10⁻² for peptides and XCorr of 10.0 and probability of 1× 10⁻³ for proteins. All protein identifications were verified by manual inspection of MS/MS assignments. The number of spectra acquired from the peptide mixture was used as a measure of protein abundance within the protein pull-down as described previously in ref. [23].

3.2 Identification and quantification of proteins bound to modified histone H3 peptides

The 40 tryptic peptide samples were analyzed individually by HPLC-MS/MS on an LTQ mass spectrometer. To assess the extent of nonspecific binding, we also analyzed the streptavidin beads control pull-down. The majority of proteins that bound to the control agarose beads consisted mainly of common and abundant nonspecific proteins that we often identify in immunoprecipitation assays (see Table 1 of Supporting Information). All proteins identified in the four H3 peptide pull-downs are listed in Table 1.

It has been demonstrated that stable isotope labeling with amino acids in cell culture (SILAC) in combination with high-resolution MS can provide an accurate method for quantifying the relative amount of each protein identified in the pull-downs [25–27]. Indeed, such an approach has been used to identify histone modification-specific binding proteins and protein interaction partners [28, 29]. However, a major limitation of the isotopic amino acid labeling method is the limited number of samples that can be used for relative quantification, which requires pair wise comparisons. Therefore, it is not convenient to use such methods to assess the binding of proteins in the five pull-downs conducted in this study. Instead, we opted for spectral counting as described by Yates and coworkers [23] which serves as a simple and convenient method to estimate the abundance of each protein. We combined the results of the eight MS analyses for each pull-down and the total number of spectra were tallied, Table 1.

To compare and contrast the composition of proteins pulled down by the various modified histone H3 peptides, we used hierarchical clustering to analyze and heat maps to visualize the spectral counts as simple estimates of the relative abundance of each protein (Fig. 3) [24, 30]. As expected, the streptavidin beads pull-down contained the least amount of proteins overall. In a few instances, a significant number of peptides for a protein were detected in the control pull-down, such as in CHD3, CHD4, and TRIM28; however, the marked increase in the peptide spectral counts in the H3 pull-downs suggests that these proteins likely also bound to H3 or to the modified H3 sites (Table 1). In all, we identified 86 distinct proteins that preferentially bound to at least one of the four H3 peptides examined relative to the control beads. Of these 86 proteins, 65 were not recovered in the control beads pull-down; for the 21 proteins that were detected in the control pull-down, they were significantly enriched with greater than 70% increase in the number of peptides identified in at least one of the H3 pull-downs (Table 1).

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Figure 3

Cluster analysis of protein abundance among modified H3 pull-downs. For each protein, spectral counts from Table 1 are scaled to Z-scores, calculated across the various H3 modifications and control beads listed below. As shown in the legend, red indicates abundance that is above average among the eight pull-downs; green indicates below-average abundance, with various shades indicating intermediary abundance levels. To highlight proteins present at similar levels in the same samples, Z-scores for each protein were clustered along the horizontal axis.

3.3 Recovery of known H3 tail binding partners

If our approach captured bona fide interactions, we would expect to identify known binders to the H3 peptides used in this study. Indeed, our screen identified seven proteins previously shown to recognize H3 tail directly including known readers of H3K4me2, H3K9me2, and unmodified H3 tail (Tables 2 and ​and3).3). The major protein complex isolated with the unmodified H3 peptide is the NuRD/MeCP1 protein complex (Fig. 2B and Table 2), as previously reported [31–37]. In addition, we also recovered known interacting proteins of these direct effectors, suggesting that a subset of the detected interactions is indirect. These data are discussed below in more detail (Section 3.4). Remarkably, 33 proteins (more than one-third of the proteins identified in our screen) were previously reported to bind directly or associate with direct binders to the H3 tails examined. These results support the validity of our approach and strongly suggest that the remaining 55 proteins are relevant, novel binding partners that directly or indirectly associate with the H3 peptides examined in this study.

3.4 Known effectors of H3K9me2 and H3K4me2 identified

The three isoforms of HP1 (α, β, and γ), which bind specifically to H3K9me2, were identified in our screen. The chromodomain of HP1 mediates the interaction with H3 di- and trimethylated at K9 for gene silencing and maintenance of heterochromatin. Two of the HP1 isoforms (α and β) displayed strong binding preference for H3K9me2 as previously reported [15], whereas the HP1γ homolog (LOC653972) also bound to unmodified H3 and H3K9ac to a lesser degree. Accordingly, our screen identified proteins known to associate with HP1, including a component of the CAF1 complex CHAF1B [38], and corepressor protein TRIM28/KAP-1 [39]. CHAF1B and TRIM28/KAP-1 exhibited preferential binding to the H3K9me2 peptide, suggesting that the association is HP1-dependent. It is also noteworthy that TRIM28/KAP-1 contains a PHD finger that may contribute to its specificity of binding to H3K9me2.

WDR5 was identified to preferentially bind to H3K4me2 using a similar peptide pull-down assay [40]. In agreement with this result, WDR5 was significantly enriched in the H3K4me2 pull-down in comparison to the unmodified H3 tail. WDR5 contains a WD40 domain and is a common component of the MLL family of H3K4 methyltransferase complexes. WDR5 associate with RbBP5, ASH2 and a SET domain protein to form the “core complex” that is sufficient for methylation on nucleosomal substrates [41]. RbBP5, which is known to tightly associate with WDR5, was also recovered in our study, and although less abundant in terms of peptide recovery, it exhibited binding specificity closely mimicking that of WDR5. Thus, it is likely that RbBP5 binding to H3K4me2 peptide is via interactions with WDR5.

CHD1 is an ATP-dependent chromatin remodeling protein that binds to H3 tails containing di- or trimethylated K4 via its tandem chromodomains [42]. We confirmed CHD1 interaction with H3K4me2 in our peptide pull-downs (Tables 1 and ​and3).3). We also detected weak binding of CHD1 to the H3K9ac peptide, although the biological significance of this interaction is not clear.

Interestingly, we also identified multiple subunits of PAF1 elongation complex: PAF1, LEO1, CTR9, CDC73 that preferentially associated with H3K4me2 tail, although binding to other H3 peptides was also observed. PAF1 promotes transcriptional elongation and regulates methylation of H3 at K4. Preference for H3K4me2 may be explained by the previously reported association of the PAF1 complex with CHD1 and H3K4 methyltransferase core complex [43, 44]. Alternatively, one or more PAF1 components can directly bind to the histone tail.

Finally, we wish to comment on the lack of recovery of BPTF, ING2, and TAF3, three proteins that have been reported to recognize methylated H3K4 via PHD finger motif. The PHD fingers of these proteins show a higher preference for binding to trimethyllysines over dimethyllysines in pull-down assays, which likely accounts for their absence in our H3K4me2 pull-down [45, 46]. In future studies, we will perform similar proteomic screen to search for novel proteins associated with specific mono-, and trimethyllysines, which will expand our knowledge on how the different methylation states may be distinguished by the downstream effectors.

3.5 Known binders discriminating against H3K4me2 identified

Histone modifications can exert their function not only via recruitment of specific readers, but also through exclusion of certain effector molecules from the histone tails. One example includes the NuRD complex, whose binding to the H3 tail is perturbed by methylation at K4 [47, 48]. In agreement with these reports, our results indicate that the binding of multiple components of the NuRD complex, and the related MeCP1 complex, are dramatically reduced in the H3K4me2 pull-down, however to a much lesser degree, H3K9me2 also reduced binding of this complex (Fig. 2A and Table 1).

3.6 Other identified proteins previously implicated in association with H3 tail

Our study also identified two proteins previously reported to directly bind H3, albeit without clearly defined modification-dependent specificity. The histone chaperone SET/TAF-1β was shown to directly recognize unmodified and methylated H3 tails [49]. However in our assay, SET/TIF-1β does not significantly bind to the unmodified H3 tail, but instead strongly associates with H3K4me2, H3K9me2 and, to a lesser extent, with H3K9Ac.

The second protein, ERCC6, is an ATP-dependent chromatin remodeler involved in DNA excision repair and was previously reported to bind histone tails [50]. In our assay ERCC6 exhibited the highest spectral counts to the unmodified H3 and H3K9me2 peptides; however binding to H3K4me2 was not detectable, whereas binding to the H3K9Ac peptide was very weak, Table 1.

In sum, our screen identified the NuRD/MeCP1 complex and seven proteins (HP1α, HP1β, HP1γ, CHD1, WDR5, PAF1, SET/TAF-1β, and ERCC6) which were previously shown to directly associate with modified or unmodified H3 tail. These direct binders and their associated proteins constituted 33 of the 86 proteins identified in these four peptide pull-down assays. Importantly, the protein partners that associate with the direct binders displayed similar binding preference to that of the direct binders. Notably however, not all protein partners of known binders had analogous association patterns. For example, ANP32A and ANP32B, which associate with SET/TIF-1β in the INHAT complex, appear to bind specifically to H3K9me2, whereas SET/TIF-1β recognizes both methylated peptides equally well. These data indicate that different complexes formed by a given protein may exhibit different histone-binding properties.

Taken together, results discussed above (section 3.3–3.6) demonstrate not only the feasibility of the approach undertaken in this study, but also suggest that the majority of the remaining 53 identified proteins represents novel direct and indirect H3 tail binders.

We thank Joanna Wysocka and Eric Chan for critical comments on the manuscript. J.W. is partially supported by the Research Platform of Cell Signaling Networks from the Science and Technology Commission of Shanghai Municipality. This work was supported by National Institutes of Health Grant 1R43CA132680-01 to D.W.C.