1. Chromosomes and Gene Expression
Download icon

Chromatin structure-dependent histone incorporation revealed by a genome-wide deposition assay

  1. Hiroaki Tachiwana  Is a corresponding author
  2. Mariko Dacher
  3. Kazumitsu Maehara
  4. Akihito Harada
  5. Yosuke Seto
  6. Ryohei Katayama
  7. Yasuyuki Ohkawa
  8. Hiroshi Kimura
  9. Hitoshi Kurumizaka
  10. Noriko Saitoh  Is a corresponding author
  1. Division of Cancer Biology, The Cancer Institute of Japanese Foundation for Cancer Research, Japan
  2. Laboratory of Chromatin Structure and Function, Institute for Quantitative Biosciences, The University of Tokyo, Japan
  3. Division of Transcriptomics, Medical Institute of Bioregulation, Kyushu University, Japan
  4. Division of Experimental Chemotherapy, Cancer Chemotherapy Center, Japanese Foundation for Cancer Research, Japan
  5. Cell Biology Center, Institute of Innovative Research, Tokyo Institute of Technology, Japan
Tools and Resources
  • Cited 0
  • Views 855
  • Annotations
Cite this article as: eLife 2021;10:e66290 doi: 10.7554/eLife.66290

Abstract

In eukaryotes, histone variant distribution within the genome is the key epigenetic feature. To understand how each histone variant is targeted to the genome, we developed a new method, the RhIP (Reconstituted histone complex Incorporation into chromatin of Permeabilized cell) assay, in which epitope-tagged histone complexes are introduced into permeabilized cells and incorporated into their chromatin. Using this method, we found that H3.1 and H3.3 were incorporated into chromatin in replication-dependent and -independent manners, respectively. We further found that the incorporation of histones H2A and H2A.Z mainly occurred at less condensed chromatin (open), suggesting that condensed chromatin (closed) is a barrier for histone incorporation. To overcome this barrier, H2A, but not H2A.Z, uses a replication-coupled deposition mechanism. Our study revealed that the combination of chromatin structure and DNA replication dictates the differential histone deposition to maintain the epigenetic chromatin states.

Introduction

Eukaryotic genomic DNA is packaged into chromatin, in which the basic structural unit is the nucleosome. The nucleosome is composed of around 150 base pairs of DNA and a histone octamer consisting of two copies of each core histone, H2A, H2B, H3, and H4 (Luger et al., 1997). Chromatin is not only the storage form of genomic DNA but also the regulator of DNA-templated processes, such as transcription, replication, repair, and chromosome segregation. To enable these processes, chromatin forms various structures that can be reversibly altered. Historically, cytological studies first identified euchromatin and heterochromatin, which are transcriptionally active and inactive regions, respectively (Passarge, 1979). Subsequent studies revealed that euchromatin coincides with nuclease hypersensitivity, indicating that it forms a more accessible structure (open chromatin) than heterochromatin (closed chromatin) (Garel and Axel, 1976; Spiker et al., 1983; Tsompana and Buck, 2014; Weintraub and Groudine, 1976). Although these chromatin structures are involved in the regulation of DNA template-mediated processes, little is known about how the open and closed chromatin configurations are formed and maintained.

Among the chromatin associated proteins, histones have a significant impact on the chromatin structure. In humans, the canonical histones, H2A, H2B, and H3.1, have non-allelic variants with distinct expression and/or localization patterns (Buschbeck and Hake, 2017; Maehara et al., 2015). Canonical H3.1 and H2A are expressed in S phase and show genome-wide localizations (Buschbeck and Hake, 2017; Wu et al., 1982). In contrast, the histone variants H3.3 and H2A.Z are expressed throughout the cell cycle and concentrated at promoters in both open chromatin and pericentric heterochromatin (Ahmad and Henikoff, 2002; Boyarchuk et al., 2014; Drané et al., 2010; Goldberg et al., 2010; Greaves et al., 2007; Jin and Felsenfeld, 2007; Raisner et al., 2005; Rangasamy et al., 2003; Sarcinella et al., 2007; Wu et al., 1982). In addition, H3.3 localizes at telomeres (Goldberg et al., 2010). Another H2A variant, MacroH2A, is found at transcriptionally suppressed chromatin, such as inactive X chromosomes (Costanzi et al., 2000). Along with the canonical H2A, the localization of H2A.X, which functions in DNA repair, is genome-wide (Yukawa et al., 2014). Thus, in spite of the high-sequence homology between the canonical and variant forms, each histone has specific functions. A previous study identified the six essential residues responsible for the H2A.Z-specific functions, which are located in the αC helix called the M6 region (Clarkson et al., 1999). The H2A.Z swap mutant, in which M6 was replaced with the equivalent residues in H2A, failed to rescue the embryonic lethality of the H2A.Z null mutant in Drosophila melanogaster. Thus, the importance of M6 in the organism was evident; however, its molecular mechanism remained elusive.

Histone deposition and exchange have been studied for more than 30 years (Clément et al., 2018; Jackson, 1990; Jansen et al., 2007; Kimura and Cook, 2001; Louters and Chalkley, 1985; Ray-Gallet et al., 2011; Tachiwana et al., 2010). An in vivo histone deposition study must distinguish the pre-incorporated histones (parental histones) from the newly incorporated histones (new histones). To enable this, initial studies used radio-labeled histones and revealed that histone deposition occurred by both replication-coupled and replication-independent mechanisms (Jackson, 1990; Louters and Chalkley, 1985). In principle, the replication-independent histone deposition would accompany histone eviction from chromatin, in a process described as histone exchange or turnover. Several key proteins involved in these mechanisms have been identified. Anti-silencing function 1 (ASF1), together with chromatin assembly factor 1 (CAF-1), plays indispensable roles in replication-coupled H3.1 deposition (Smith and Stillman, 1989; Tagami et al., 2004; Tyler et al., 1999; Tyler et al., 1996; Tyler et al., 2001). CAF-1 is a heterotrimeric complex composed of p150, p60, and p48, and directly interacts with proliferating cell nuclear antigen (PCNA), leading to the assembly of H3.1 at replicating chromatin (Moggs et al., 2000; Shibahara and Stillman, 1999). The histone regulator A (HIRA) or death-associated protein (DAXX) promotes the accumulation of H3.3 at transcription sites and regulatory elements in a replication-independent manner (Drané et al., 2010; Goldberg et al., 2010; Ray-Gallet et al., 2011; Ray-Gallet et al., 2002; Tagami et al., 2004).

H2A- and H2A.Z-specific chaperones have also been identified (Obri et al., 2014). Facilitates chromatin transcription (FACT), a histone chaperone, may be involved in replication-coupled H2A deposition (Orphanides et al., 1998; Ransom et al., 2010; Wittmeyer and Formosa, 1997). Recent studies revealed that FACT binds to the H3-H4 complex, indicating that FACT functions in H3-H4 depositions as well (Liu et al., 2020; Tsunaka et al., 2016; Wang et al., 2018). ANP32E functions in H2A.Z eviction from chromatin, and the SNF2-related CBP activator protein (SRCAP) chromatin-remodeling subunit YL1 promotes H2A.Z deposition at gene promoters (Latrick et al., 2016; Liang et al., 2016; Mao et al., 2014; Obri et al., 2014; Wong et al., 2007). The histone acetyl transferase (HAT) complex, NuA4/TIP60, also functions in H2A.Z deposition by interacting with pre-deposited H2A.Z (Gévry et al., 2009; Giaimo et al., 2019; Shia et al., 2006).

The precise distribution of histones is crucial for chromatin organization and its epigenetic states. The ChIP-seq analysis method is powerful and useful to visualize steady state histone localizations; however, it does not enable the analysis of the incorporation of each histone into open/closed chromatin or both (genome-wide). To analyze histone incorporations, several approaches using fluorescence imaging, mass spectrometry, inducible tagged proteins, and labeling of newly synthesized proteins have been developed (Deal and Henikoff, 2010). Imaging-based methods, such as fluorescence recovery after photobleaching (FRAP) and SNAP/CLIP-tag technology, were developed to analyze the histone dynamics in living cells. FRAP is suitable for investigating the histone mobility in living cells (Kimura and Cook, 2001; Tachiwana et al., 2010). The SNAP/CLIP-technology is effective for analyzing the histone deposition, as it can distinguish parental histones from new histones in cells with cell-permeable fluorophores that covalently bind to the tag (Clément et al., 2018; Jansen et al., 2007; Ray-Gallet et al., 2011). Although these are powerful methods, they have resolution limitations since they are based on imaging. The SNAP/CLIP-technology led to the establishment of the time-ChIP method, in which a biotin-labeled SNAP-histone is captured (Deaton et al., 2016). The time-ChIP method was developed to measure the stability of parental histones, rather than the histone incorporation, as it has a time lag during the synthesis and labeling of histones, which limits the time resolution (Deaton et al., 2016; Siwek et al., 2018). The tetracycline (Tet) inducible expression system, which induces the expression of epitope-tagged histones, has been applied for analyses of nucleosome turnover/histone exchange. However, this system also has a time resolution limitation (Ha et al., 2014; Yildirim et al., 2014). To uncover the correlation between histone incorporation and open/closed chromatin structures, a new analysis method for the histone incorporation at the DNA sequence level is desired.

Permeabilized cells are useful to dissect the molecular pathways in nuclear events (Adam et al., 1990; Okuno et al., 1993). In assays with such cells, the cellular membranes are permeabilized by a nonionic detergent treatment. In the permeabilized cells, the chromatin and nuclear structures remain intact and react with exogenously added proteins (Kimura et al., 2006; Maison et al., 2002; Misteli and Spector, 1996; Saitoh et al., 2006). A previous study showed that an exogenously added GFP-tagged histone, prepared from cultured human cells, was incorporated into the chromatin of permeabilized cells in the presence of a cellular extract (Kimura et al., 2006). Moreover, permeabilized cells are suitable for monitoring replication timing, by labeling the nascent DNA with exogenously introduced nucleotides (Kimura et al., 2006; Misteli and Spector, 1996).

In the present study, we developed a new method, in which a reconstituted histone complex, instead of a fluorescent protein-tagged histone, was added to permeabilized cells. We named this the RhIP assay (Reconstituted histone complex Incorporation into chromatin of Permeabilized cell). Since the histone complexes are reconstituted in vitro using epitope-tagged recombinant histones, RhIP with sequencing allows the analysis of incorporations at the DNA sequence level, without the need for specific antibodies. We found that the chromatin structure regulates the histone incorporations, which may be necessary for maintaining the epigenetic state of chromatin.

Results

RhIP assay reproduces in vivo histone deposition

To understand how histones are incorporated into chromatin in cells, we developed the RhIP assay, in which an in vitro reconstituted histone complex, nucleotides, and a cellular extract are added to permeabilized cells (Figure 1A). We first confirmed that the RhIP assay can recapitulate the specific histone incorporations observed in cells. The H3.1-H4 incorporation into chromatin is coupled with replication, while the H3.3-H4 incorporation occurs throughout the cell cycle (Ahmad and Henikoff, 2002). We reconstituted H3-H4 complexes in vitro, using recombinant H3.1, H3.3, and H4 (Figure 1B). The recombinant H3.1 and H3.3 were fused to HA and FLAG tags at their C-termini, respectively. The permeabilized cells were then prepared by treating HeLa cells with a nonionic detergent, Triton X-100, and the reconstituted H3-H4 complexes were mixed with the cellular extract and nucleotides. Cy5-dUTP was also added, in order to monitor DNA replication. After the reaction, the exogenously added H3.1 and H3.3 were detected with antibodies against the HA and FLAG tags, respectively (Figure 1C–E). As a result, the H3.1 was detected in the Cy5 positive cells (S phase cells), while the H3.3 was detected irrespective of the Cy5 signal. As the co-incubated H3.1–3HA-H4 and H3.3-3FLAG-H4 complexes showed different staining patterns, they were incorporated into the chromatin by specific mechanisms, rather than non-specifically, in the RhIP assay.

Figure 1 with 4 supplements see all
RhIP (Reconstituted histone complex Incorporation into chromatin of Permeabilized cells) assay recapitulates the replication-coupled H3.1-H4 and -dependent H3.3-H4 depositions.

(A) Schematic representation of the RhIP assay, using reconstituted H3.1-H4 and H3.3-H4 complexes. Permeabilized cells were prepared from HeLa cells treated with non-ionic detergent, to perforate the cellular membranes. The in vitro reconstituted H3-H4 complexes were then added to the cells with the cellular extract and nucleotides. Cy5-dUTP was added to label the nascent DNA, so replication could be monitored. (B) Reconstituted H3.1-H4 and H3.3-H4 complexes were analyzed by SDS-16% PAGE with Coomassie Brilliant Blue staining. The 3HA and 3FLAG tags were fused to the C-termini of H3.1 and H3.3, respectively. Lane one indicates the molecular mass markers, and lanes 2 and 3 indicate the H3.1-H4 and H3.3-H4 complexes, respectively. (C) RhIP-immunostaining of H3.1 and H3.3. Exogenously added H3-H4 complexes were stained with an anti-HA or -FLAG antibody. Cells in S phase were monitored with Cy5-dUTP, which was incorporated into the nascent DNA. Bar indicates 10 μm. (D) Quantification of C. The mean fluorescence intensities (MFI) of H3.1–3HA (left) and H3.3-3FLAG (right) were measured. Nuclei were divided into S phase (Cy5 positive) and out of S phase (Cy5 negative) (n > 50, triplicate). (E) Relative intensity of H3.1 or H3.3 signal in S phase against signal out of S phase. Experiments were repeated three times and averaged data with standard deviations are shown. The two-tailed Student’s t-test was used for the statistical comparisons. (F) Schematic representation of the RhIP-ChIP assay, using the reconstituted H3.1-H4 and H3.3-H4 complexes. The reconstituted H3.1-H4 or H3.3-H4 complex was added to permeabilized cells with the cellular extract and nucleotides. Cy5-dUTP was added to label the nascent DNA. The chromatin was partially digested with micrococcal nuclease (MNase). Chromatin immunoprecipitation was performed with anti-HA magnetic beads. The precipitated DNA was extracted and analyzed by agarose gel electrophoresis. (G) Reconstituted H3.1-H4 and H3.3-H4 complexes were analyzed by SDS-16% PAGE with Coomassie Brilliant Blue staining. A 3HA tag was fused to the C-termini of H3.1 and H3.3. Lane one indicates the molecular mass markers, and lanes 2 and 3 indicate the H3.1-H4 and H3.3-H4 complexes, respectively. (H) The immunoprecipitated DNA was analyzed by 2% agarose electrophoresis. Upper and lower images were obtained from the same gel. The DNA was visualized with SYBR Gold (upper), and the nascent DNA was visualized by detecting the Cy5 signals (lower). Lane 1 indicates the 100 bp DNA ladder. Lanes 2–4 and 5–7 indicate input samples and immunoprecipitated samples, respectively. Each set includes the experiments with no reconstituted histone complex (negative control, lanes 2 and 5), with H3.1–3HA-H4 (lanes 3 and 6), and with H3.3–3HA-H4 (lanes 4 and 7).

To improve the resolution of the analysis, we performed the RhIP assay followed by chromatin immunoprecipitation (RhIP-ChIP) (Figure 1F). The reconstituted H3.1–3HA-H4 or H3.3–3HA-H4 complex was added to permeabilized cells along with the cellular extract and nucleotides, including Cy5-dUTP to label the nascent DNA (Figure 1F and G). After the reaction, the chromatin was partially digested by micrococcal nuclease (MNase), and the nucleosomes containing H3.1–3HA or H3.3–3HA were immunoprecipitated with an antibody against the HA tag. The precipitated DNA was then extracted and analyzed by gel electrophoresis. As shown in Figure 1H, the amounts of precipitated DNA are nearly the same between the H3.1 and H3.3 samples, as judged from the SYBR Gold staining (Figure 1H, upper); however, the amount of nascent DNA labeled with Cy5 is much greater in the H3.1 sample than in the H3.3 sample (Figure 1H, lower). This result indicates that H3.1 is incorporated into replicating chromatin more efficiently than H3.3. We further performed the RhIP-ChIP-seq of H3.3 to examine the distribution of incorporated H3.3 in the RhIP assay (Figure 1—figure supplement 1A). We found that exogenously added H3.3 was preferentially incorporated into transcriptionally active genes, rather than inactive genes. This distribution pattern is consistent with many previous studies (Bachu et al., 2019; Mito et al., 2005; Pchelintsev et al., 2013). The RhIP-ChIP-qPCR analysis also demonstrated the H3.3 enrichment at the transcriptionally active GAPDH, as compared to that at the inactive gene, LNC02199 (Figure 1—figure supplement 1B and C). Thus, our results imply that the reconstituted H3.1-H4 and H3.3-H4 complexes are incorporated into the chromatin of permeabilized cells with the same dynamics as observed in intact cells.

CAF-1 and HIRA are key factors for the H3.1 and H3.3 depositions in vivo, respectively (Drané et al., 2010; Goldberg et al., 2010; Ray-Gallet et al., 2002; Ray-Gallet et al., 2011; Smith and Stillman, 1989; Tagami et al., 2004; Tyler et al., 1996; Tyler et al., 1999; Tyler et al., 2001). We therefore examined whether their roles can be recapitulated in the RhIP assay (Figure 1—figure supplement 2). We first analyzed whether CAF-1 and HIRA remain in the permeabilized cells or are extracted during the permeabilization process (Figure 1—figure supplement 2A). Our immunoblot showed that both CAF-1 (p60) and HIRA remained in the permeabilized cells. We then performed the RhIP-immunostaining assay of H3.1 or H3.3 using CAF-1- or HIRA-knockdown cells, respectively (Figure 1—figure supplement 2B). For a precise evaluation, we co-cultured the control and knockdown cells on the same coverslips, and performed RhIP-immunostaining assays of H3.1 or H3.3. The knockdown cells were judged by immunostainings for CAF-1 (p60) (cells with arrowheads in Figure 1—figure supplement 2C) and HIRA (cells with arrowheads in Figure 1—figure supplement 2D). For the detection of H3.1, cells were synchronized to S phase by adding thymidine 18 hr before performing the RhIP assay. As a result, the H3.1 incorporation was detected in CAF-1-positive S phase cells (arrows in Figure 1—figure supplement 2C). Similarly, the H3.3 was detected in HIRA positive cells (arrows in Figure 1—figure supplement 2D). We also investigated whether H3.1 and H3.3 were incorporated into the chromatin or merely bound to histone chaperones, CAF-1 and HIRA, respectively, which are present in the chromatin of the permeabilized cells. After performing the RhIP assay, we washed the cells with PBST containing 300 mM NaCl prior to the fixation and immunostaining processes (Figure 1—figure supplement 3A). We found that that the exogenously added H3.1 or H3.3 was still detected after washing the cells with PBST containing 300 mM NaCl, even though CAF-1 and HIRA were no longer detected (Figure 1—figure supplement 3B–E). Therefore, the exogenously added H3.1 and H3.3 were mostly incorporated into the chromatin of the permeabilized cells in the RhIP assay. These data demonstrated that the H3.1 and H3.3 incorporations into chromatin in cells were correctly recapitulated in the RhIP assay.

We further examined whether the cellular extract is essential or replaceable by the histone chaperones, NAP1 or ASF1 (Figure 1—figure supplement 4A). NAP1 and ASF1 bind to the H2A-H2B and H3-H4 complexes in vivo, respectively, and both promote nucleosome formation in vitro (Ishimi et al., 1984; Munakata et al., 2000; Tachiwana et al., 2008; Tyler et al., 1999). Human NAP1 and ASF1 were purified as recombinant proteins (Figure 1—figure supplement 3B). The H3.1-H4 complex was then added to the permeabilized cells in the absence of the cellular extract or in the presence of NAP1 or ASF1, and the incorporation was analyzed by immunostaining (Figure 1—figure supplement 4C). Without the cellular extract or the histone chaperone, the exogenously added H3.1 was not detected in the permeabilized cells, indicating that no H3.1 incorporation had occurred. NAP1 promoted the promiscuous incorporation irrespective of DNA replication, and ASF1 facilitated H3.1 accumulation in the nucleoli. These data indicated that the functional deposition of the exogenously added histone complex requires the cellular extract, which may contain essential components. Together with the fact that the efficiency of RhIP-ChIP of exogenously added H3.3 is almost the same as that of endogenous H3.3 ChIP (Figure 1—figure supplement 1D and E), we conclude that the RhIP assay reproduces cellular histone deposition and is suitable for analyzing histone incorporation in vitro.

H2A.Z incorporation into chromatin differs from that of H2A and H2A.X

Among the H2A family members, canonical H2A and the H2A.X variant show even and broad genome-wide distributions, but H2A.Z specifically localizes in open chromatin, including promoters and enhancers (Buschbeck and Hake, 2017; Raisner et al., 2005). To test whether this difference reflects their deposition manners, we performed the RhIP assay (Figure 2A). The H2A-H2B and H2A.Z-H2B complexes were reconstituted in vitro using recombinant proteins (Figure 2B) and added to permeabilized cells, which were then immunostained (Figure 2C). The H2A and H2A.Z signals were both observed in the Cy5-negative and -positive permeabilized cells, indicating that their incorporations occur irrespective of DNA replication. We also found that H2A forms foci in the S phase nuclei. We then merged the images of the H2A and Cy5 signals. The replication foci change as cells progress through S phase (Leonhardt et al., 2000). In early S phase, the replication foci are present throughout the nucleoplasm, except for the nucleoli. The foci then accumulate at the nuclear periphery and around the nucleoli. In late S phase, the foci increase in size but decrease in number. We found that the H2A signals overlapped well with the replication foci throughout replication (Figure 2C and D). The H2A.Z signals overlapped with the early replication foci to some extent, but they were clearly eliminated from the late replication foci (Figure 2C and D). In contrast to the difference between H2A and H2A.Z, the H2A and H2A.X signals overlapped well with each other, suggesting that their incorporation mechanism is the same (Figure 2—figure supplement 1). These results indicate that H2A, H2A.X, and H2A.Z can be incorporated into chromatin in a replication-independent manner; however, during S phase, H2A and H2A.X are preferentially incorporated into the chromatin of ongoing replication sites, in contrast to H2A.Z.

Figure 2 with 2 supplements see all
The H2A.Z and H2A deposition patterns are different in the RhIP assay.

(A) Schematic representation of the RhIP assay, using the reconstituted H2A-H2B and H2A.Z-H2B complexes. (B) The reconstituted H2A-H2B and H2A.Z-H2B complexes were analyzed by SDS-16% PAGE with Coomassie Brilliant Blue staining. The 3HA and V5 tags were fused to the N-termini of H2A and H2A.Z, respectively. Lane one indicates the molecular mass markers, and lanes 2 and 3 indicate the H2A-H2B and H2A.Z-H2B complexes, respectively. (C) RhIP-immunostaining of H2A and H2A.Z. Top: Exogenously added H2A-H2B or H2A.Z-H2B complexes were stained with either an anti-HA or -V5 antibody. Cells in S phase were monitored with Cy5-dUTP. Middle and Bottom: merged images of Cy5-dUTP (green) and H2A or H2A.Z (red) in early S (Middle) and late S (Bottom) phase. Bar indicates 10 μm, and r indicates the Pearson’s correlation coefficient. (D) Colocalization analyses of Cy5-dUTP and H2A.Z or H2A (n > 35 cells). Experiments were repeated three times and averaged data are shown. The two-tailed Student’s t-test was used for the statistical comparisons. (E) Reconstituted H2A-H2B and H2A.Z-H2B complexes were analyzed by SDS-16% PAGE with Coomassie Brilliant Blue staining. A 3HA tag was fused to the N-termini of H2A and HA.Z. Lane 1 indicates the molecular mass markers, and lanes 2 and 3 indicate the H2A-H2B and H2A.Z-H2B complexes, respectively. (F) The RhIP-ChIP assay was performed using H2A and H2A.Z, as described in Figure 1F. The immunoprecipitated DNA was analyzed by 2% agarose electrophoresis. Upper and lower images were obtained from the same gel. The DNA was visualized with SYBR Gold (upper), and the nascent DNA was visualized by detecting the Cy5 signals (lower). Lane 1 indicates the 100 bp DNA ladder. Lanes 2–4 and 5–7 indicate input samples and immunoprecipitated samples, respectively. Each set has the experiments with no reconstituted histone complex (negative control, lanes 2 and 5), with 3HA-H2A-H2B (lanes 3 and 6), and with 3HA-H2A.Z-H2B (lanes 4 and 7).

We further analyzed the incorporation of H2A and H2A.Z into replicating chromatin by a RhIP-ChIP assay, as in Figure 1F (Figure 2E and F). The amounts of precipitated DNA are nearly the same between the H2A and H2A.Z precipitants, as judged from the SYBR Gold staining (Figure 2F, upper); however, the amount of nascent DNA labeled with Cy5 is much greater in the H2A precipitant than in the H2A.Z sample (Figure 2F, lower). This result indicates that H2A is incorporated into replicating chromatin more efficiently than H2A.Z.

The replication timing in S phase strongly correlates with the chromatin configurations (Rivera-Mulia and Gilbert, 2016). In general, early and late replicating chromatin regions correspond to open and closed chromatin, respectively. We investigated whether the efficiencies of H2A and H2A.Z incorporation into replicating chromatin change, according to the replication timing (Figure 2—figure supplement 2). For this analysis, the cells were synchronized in early S phase by a double thymidine block, and then early and late S phase cells were collected at 0 and 5 hr post thymidine-release, respectively. The synchronized cells showed the typical early and late replication foci representing nascent DNA labeled with Cy5-dUTP (Figure 2—figure supplement 2A). Using these cells, we performed RhIP-ChIP assays of H2A and H2A.Z. The results revealed that the incorporation efficiencies of both H2A and H2A.Z into replicating chromatin did not change, irrespective of the replication timing in S phase. This implies that the efficiencies of replication-coupled histone deposition are not different between open (early S-replicating) and closed (late S-replicating) chromatin (Figure 2—figure supplement 2B).

Open and closed chromatin structures regulate histone deposition

The RhIP-ChIP assay showed that the H2A.Z deposition on nascent DNAs was constant in the early and late S phases (Figure 2F and Figure 2—figure supplement 2). In contrast, the RhIP-immunostaining revealed that more signals of H2A.Z incorporation were overlapped at the early replicating foci than the late replicating foci (Figure 2C and D). This discrepancy may arise from the lower resolution of the immunostaining imaging. Some of the overlapping signals of H2A.Z and Cy5 in early S phase might represent the replication-independent H2A.Z deposition that occurred close to, but not exactly at, the replication sites in open chromatin. To determine whether the efficiency of the histone deposition depends on the open/closed chromatin configuration, we performed RhIP-ChIP-seq and analyzed the H2A and H2A.Z incorporations in each type of chromatin (Figure 3). First, we investigated the replication-independent histone deposition using asynchronous permeabilized cells, in which the majority of the cells are out of S phase. We found that the RhIP-ChIP-seq profiles of H2A and H2A.Z showed a strong correlation (0.62 Pearson correlation) and specific peaks at megabase resolution, which were not observed in the input samples (Figure 3A). We noticed that these patterns are similar to the DNaseI-seq results, which mapped open chromatin regions (Tsompana and Buck, 2014), suggesting that H2A and H2A.Z are predominantly incorporated into open chromatin regions. We then analyzed the efficiency of histone incorporations into open and closed chromatin using the chromHMM data (Core 15-state model), which classified the open and closed chromatin regions (Ernst and Kellis, 2012; Roadmap Epigenomics Consortium et al., 2015Figure 3B and Figure 3—figure supplement 1A). The ChIP/input ratios of each chromatin region revealed that H2A and H2A.Z were efficiently incorporated into transcriptionally active open chromatin, while their incorporations into closed chromatin were inefficient. These results indicate that histone incorporation occurs mainly in the open chromatin regions in a replication-independent manner, and closed chromatin suppresses histone incorporation.

Figure 3 with 2 supplements see all
The incorporation of histones H2A and H2A.

Z mainly occurs at less condensed chromatin and H2A incorporation into condensed chromatin requires a replication-coupled deposition mechanism. (A) RhIP-ChIP-seq and DNaseI-seq profiles using asynchronous cells were visualized with the Integrative Genomics Viewer (left). From top to bottom, profiles of H2A, H2A.Z, DNaseI-seq (GEO:GSM816643), input (H2A), and input (H2A.Z) are indicated. Scatter plot analyses of the H2A.Z and H2A RhIP-ChIP-seq along the genome (right). (B) Enrichment of incorporated H2A (left) or H2A.Z (right) in asynchronous cells. Each chromatin region was previously annotated by the chromHMM, as follows. TssA: active TSS, TssAFlnk: flanking active TSS, TxFlnk: transcribed state at the 5' and 3' ends of genes showing both the promoter and enhancer signatures, Tx: strong transcription, TxWk: weak transcription, EnhG: genic enhancers, Enh: enhancers, Het: heterochromatin, ReprPC: repressed PolyComb, and ReprPCWk: weak repressed PolyComb (Ernst and Kellis, 2012; Roadmap Epigenomics Consortium et al., 2015). (C) The Pearson’s correlation coefficients between asynchronous and late S cells (two biological replicates each) were calculated from the RhIP-ChIP-seq data at 800 bp intervals. Left and right panels indicate the correlation coefficients of the H2A and H2A.Z data, respectively. (D) Enrichment of incorporated H2A in asynchronous (white boxes) and late S (green boxes) phase cells. Data for asynchronous cells are those shown in Figure 4C.

We then examined whether the efficiency of the histone incorporation into closed chromatin changes during S phase (Figure 3C and D). We performed RhIP-ChIP-seq with cells synchronized at late S phase. We found that the RhIP-ChIP-seq profiles between asynchronous cells and late S phase cells changed for H2A, but not for H2A.Z, indicating that H2A incorporation into chromatin is affected by replication, in contrast to H2A.Z incorporation (Figure 3C and Figure 3—figure supplement 1B). We then investigated how the incorporation efficiency of H2A into open and closed chromatin regions changes between asynchronous and late S phase cells. The results revealed that the efficiency of H2A incorporation into open chromatin decreased in late S phase (Figure 3D, lanes 1–7), while in contrast, the incorporation into closed chromatin increased (Figure 3D, lanes 8–10 and Figure 3—figure supplement 1B). Considering the fact that only closed chromatin is replicated in late S phase, the changes in the incorporation efficiency may be due to the changes in the replicating chromatin. Note that replication does not progress completely in the RhIP assay, and only a small fraction of closed chromatin is replicated under the conditions shown in Figure 4E, which is the reason why the ChIP/input ratio does not exceed one. We concluded that replication allows the incorporation of H2A, but not H2A.Z, in closed chromatin. The frequency of exchange is high in open chromatin and low in closed chromatin for H2A and H2A.Z. As a result, H2A.Z is specifically incorporated within open chromatin.

Figure 4 with 1 supplement see all
H2A.Z is specifically enriched around transcription start sites in the RhIP assay.

(A) Representative profiles of RhIP-ChIP-seq using asynchronous cells and ChIP-seq (GEO:GSM1003483) at chr10: 73,079,443–73,805,171. (B) Aggregation plots of the H2A (RhIP-ChIP, left), H2A.Z (RhIP-ChIP, center), and H2A.Z (ChIP, right) at the center of TSS-flanking 5 Kb regions from the whole genome. Red and blue lines indicate expressed and non-expressed genes, respectively. (C) The correlations of H2A.Z (ChIP) and H2A (RhIP-ChIP) (left), and H2A.Z (ChIP), H2A.Z (RhIP-ChIP) (center) at known H2A.Z sites determined by ChIP-seq analysis. The correlations of H2A.Z (ChIP) and H2A.Z (RhIP-ChIP) at known H2A.Z sites located in gene bodies (right). (D) Heatmaps of H2A.Z and H2A (RhIP-ChIP) at the center of H2A.Z peaks of ChIP-seq flanking 5 Kb regions. The heatmap represents H2A.Z peaks ranked from the strongest to weakest in RhIP-ChIP-seq. Corresponding aggregation plots are at the top. (E) Reversed analysis of (D). Heatmap of H2A.Z (ChIP) at the H2A.Z peaks of RhIP-ChIP-seq. The corresponding aggregation plot is at the top. Asterisk indicates peaks with no read counts of H2A.Z (ChIP), which are approximately 3% of the entire peaks.

The distribution of incorporated H2A.Z in the RhIP assay is similar to the steady-state of H2A.Z localizations

As endogenous H2A.Z predominantly localizes in transcription start sites (TSS) (Barski et al., 2007; Buschbeck and Hake, 2017; Ernst et al., 2011; Jin et al., 2009; Link et al., 2018; Obri et al., 2014; Schones et al., 2008), we further analyzed the H2A.Z distributions in the RhIP assay at kilobase resolution (Figure 4). The alignment of the H2A.Z profiles from RhIP-ChIP-seq and ChIP-seq (ENCODE Project Consortium, 2012) at a representative chromosome position (chr10:73,079,443–73,805,171) highlighted the similarity between the two, but the H2A profiles of the RhIP-ChIP-seq did not (Figure 4A). Moreover, in the RhIP assay the incorporated H2A.Z predominantly accumulated at the TSS of expressed genes, as also observed in the ChIP-seq analysis of H2A.Z, but the incorporated H2A was relatively excluded from the TSS (Figure 4B). A correlation analysis at known H2A.Z sites, determined by a ChIP-seq analysis, showed a moderately positive, linear relationship between H2A.Z of RhIP-ChIP-seq and H2A.Z of ChIP-seq (0.31 Pearson correlation coefficient) and little to no correlation between H2A of RhIP-ChIP-seq and H2A.Z of ChIP-seq (0.20 Pearson correlation coefficient) (Figure 4C). In addition, a correlation analysis at the known H2A.Z sites located in gene bodies showed a fairly linear relationship between H2A.Z of RhIP-ChIP-seq and H2A.Z of ChIP-seq (0.25 Pearson correlation coefficient) (Figure 4C, right). The heatmap revealed that the H2A.Z of RhIP-ChIP-seq accumulates at almost all of the H2A.Z peaks found by ChIP-seq (Figure 4D left and Figure 4—figure supplement 1A). In contrast, the H2A of RhIP-ChIP-seq showed no accumulation at the H2A.Z peaks of ChIP-seq (Figure 4D right). This is consistent with the results that the correlation of RhIP-ChIP-seq of H2A and H2A.Z at the H2A.Z peaks of ChIP-seq is lower than that along the genome (Figure 3A right and Figure 3—figure supplement 2A right). The reversed heatmap analysis again showed the accumulations of the H2A.Z of ChIP-seq at the H2A.Z peaks found by RhIP-ChIP-seq, and only 3% of the H2A.Z peaks of the RhIP assay were absent from those of ChIP-seq (denoted with * in Figure 4E and Figure 4—figure supplement 1B). These data revealed that the distribution of the incorporated H2A.Z in the RhIP assay overlapped well with the steady state localization of H2A.Z determined by the ChIP-seq analysis.

H2A.Z deposition in the RhIP assay requires ANP32E and ATP supplements from the cellular extract

As H2A.Z is incorporated into the same regions of the endogenous H2A.Z in the RhIP assay, the pre-incorporated H2A.Z in the chromatin may be dynamically exchanged with the exogenously added H2A.Z in the RhIP assay. The exchange reaction requires the H2A.Z-H2B complex to be evicted from nucleosomes, and ANP32E has this eviction activity (Gursoy-Yuzugullu et al., 2015; Murphy et al., 2020; Murphy et al., 2018; Obri et al., 2014). We then examined whether ANP32E is involved in the deposition of H2A.Z in the RhIP assay. Our fractionation analysis showed that ANP32E was extracted during the permeabilization process; in contrast, the other H2A.Z deposition factors SRCAP and TIP60 remained in the permeabilized cells (Figure 5A). Given that ANP32E exists in the cellular extract used in the RhIP assay (Figure 5B), we prepared cellular extracts from the control and ANP32E-knockdown cells (Figure 5B), and used them for the RhIP-ChIP-seq analysis of H2A.Z (Figure 5C). To allow a quantitative assessment of the ChIP-seq analysis, we combined spike-in controls with the RhIP-ChIP-seq (Chen et al., 2015; Egan et al., 2016; Orlando et al., 2014). The efficiencies of H2A.Z incorporations into active TSS and enhancer regions were decreased upon the ANP32E-knockdown (Figure 5D and Figure 5—figure supplement 1A). Previous reports showed that the endogenous H2A.Z is predominantly detected in TSS and to a lesser extent in enhancers (Barski et al., 2007; Buschbeck and Hake, 2017; Ernst et al., 2011; Jin et al., 2009; Link et al., 2018; Obri et al., 2014; Schones et al., 2008). Together with these observations, our results suggested that the ANP32E-dependent evictions of the pre-incorporated H2A.Z in active TSS and enhancer regions promote the incorporation of exogenously added H2A.Z.

Figure 5 with 1 supplement see all
H2A.Z deposition requires both ANP32E and ATP in the RhIP assay.

(A) Scheme of cell fractionation (left). HeLa cells were treated with non-ionic detergent and the supernatant (soluble fraction), which is removed from permeabilized cells, and the pellet, which corresponds to permeabilized cells, were separated (insoluble fraction). Distributions of SRCAP, TIP60 and ANP32E were investigated by western blotting (right). Histone H3 and GAPDH served as the chromatin and cytoplasmic protein controls, respectively. (B) Preparation of the ANP32E-knockdown cellular extract. HeLa cells were transfected with siRNA against ANP32E. After 72 hr, the ANP32E-knockdown cellular extract was prepared from the cells. The ANP32E-knockdown was confirmed by western blotting (upper). Loading control with the membrane stained with CBB (lower). (C) Scheme of RhIP-ChIP-seq of H2A.Z using the ANP32E-knockdown cellular extract. (D) Enrichment of incorporated H2A.Z with the control or ANP32E-knockdown extract. Each chromatin region was previously annotated by the chromHMM, as follows. TssA: active TSS, Enh: enhancers, and Ins: insulator. (E) Scheme of RhIP-ChIP-seq of H2A.Z using the ATP-depleted cellular extract. (F) Enrichment of incorporated H2A.Z with the ATP-depleted or ATP-supplemented extract. Each chromatin region was the same as in (D).

In addition to eviction activity, chromatin remodeling is a key determinant for the H2A.Z localizations. The SRCAP and TIP60/EP400 complexes are chromatin remodeling factors that replace nucleosomal H2A with H2A.Z. These factors were found in the permeabilized cells, suggesting that chromatin remodeling is also involved in H2A.Z deposition in the RhIP assay (Figure 5A). A series of studies using the yeast SRCAP ortholog SWR1 revealed the mechanism of chromatin remodeling-mediated H2A.Z deposition and showed that ATPase activity is essential for the H2A.Z deposition (Altaf et al., 2010; Luk et al., 2010; Mizuguchi, 2004; Sun and Luk, 2017; Sun et al., 2020). Therefore, we investigated whether ATP is required for the deposition of H2A.Z in the RhIP assay. As cellular ATP is removed during the permeabilization process, the permeabilized cells lack ATP. We then prepared the ATP-reduced cellular extract and used it in a RhIP-ChIP-seq analysis of H2A.Z with or without ATP supplementation (Figure 5E). We found that ATP facilitated H2A.Z deposition at active TSS and enhancer regions as effectively as ANP32E (Figure 5F and Figure 5—figure supplement 1B). This indicated that the H2A.Z deposition in the RhIP assay is a net result of the ANP32E-mediated eviction and deposition of H2A.Z, and the replacement of pre-incorporated H2A with H2A.Z by chromatin remodeling. As ATP is used in multiple biological processes, such as transcription, splicing/translation, molecular chaperone functions, and so on, the possibility that these ATP-dependent reactions affected the H2A.Z incorporation in this assay cannot be excluded. Intriguingly, the efficiencies of H2A.Z incorporations into insulator regions are less affected under ANP32E- or ATP-reduced conditions (Figure 5D and F). This suggests that there may be another mechanism for H2A.Z deposition in insulator regions.

The αc helix of H2A is important for its replication-coupled deposition

As the RhIP assay reproduces the replication-coupled and replication-independent incorporations of H2A and H2A.Z, we then tried to identify the residues responsible for the replication-coupled H2A and replication-independent H2A.Z depositions by a mutant analysis. A previous study showed that swapping the M6 region of H2A.Z with the corresponding H2A residues could not rescue the embryonic lethality of the H2A.Z null mutation in Drosophila melanogaster (Clarkson et al., 1999), suggesting that the region specifies the H2A.Z identity. The M6 region of H2A.Z and the corresponding region of H2A are exposed on the surface of the H2A.Z-H2B or H2A-H2B dimer (Horikoshi et al., 2013; Luger et al., 1997; Suto et al., 2000; Tachiwana et al., 2010Figure 6A, cyan or green). This indicates that another protein can recognize the regions, which may be important for their depositions. To test this idea, we constructed the swapped mutant (H2A.Z_M6) and performed the RhIP assay (Figure 6B–F). Surprisingly, the H2A.Z_M6-H2B signals were observed in late replicating chromatin (Figure 6D and E), and its incorporation pattern in late S phase was more similar to that of H2A-H2B, rather than H2A.Z-H2B (Figure 6F). This indicated that the mutant is no longer H2A.Z, in terms of deposition. Thus, the M6 region of H2A.Z is responsible for the H2A.Z-specific deposition, and the corresponding region (amino acids 89–100) of H2A is responsible for the replication-coupled H2A deposition.

Identification of responsible residues for H2A- and H2A.Z-specific incorporations.

(A) Amino acid alignments of the H2A.Z M6 region and its counterpart in H2A (upper). The structural models of the H2A.Z-H2B and H2A-H2B dimers (PDB IDs: 3WA9 and 3AFA, respectively). The specific residues are highlighted in cyan or green, respectively. All residues are located on the surface of the dimers. (B) Reconstituted H2AZ_M6-H2B, H2A-H2B, and H2A.Z-H2B complexes were analyzed by SDS-16% PAGE with Coomassie Brilliant Blue staining. H2AZ_M6, H2A, and H2A.Z were expressed as N-terminally V5, 3FLAG, and 3HA fused proteins, respectively. Lane 1 indicates the molecular mass markers, and lanes 2–4 indicate the H2A.Z_M6-H2B, H2A-H2B, and H2A.Z-H2B complexes, respectively. (C) Schematic representation of the RhIP assay, using the reconstituted H2A.Z_M6-H2B, H2A-H2B, and H2A.Z-H2B complexes. (D) RhIP-immunostaining images of H2A.Z and H2A.Z_M6. Exogenously added H2A.Z-H2B and H2A.Z_M6-H2B complexes were stained with the anti-V5 or -HA antibody. Cells in S phase were monitored with Cy5-dUTP. Middle: merged images of Cy5-dUTP (green) and H2A.Z_M6 (red) or H2A.Z (red). Bottom: magnified images of boxed areas are shown. Bar indicates 5 μm. (E) RhIP-immunostaining images of H2A and H2A.Z_M6. Exogenously added H2A-H2B and H2A.Z_M6-H2B complexes were stained with the anti-V5 or -FLAG antibody. Cells in S phase were monitored with Cy5-dUTP. Middle: merged images of Cy5-dUTP (green) and H2A.Z_M6 (red) or H2A (red). Bottom: magnified images of boxed areas are shown. Bar indicates 5 μm. (F) Colocalization analyses of Cy5-dUTP and H2A.Z, H2A.Z_M6 or H2A in late S phase (n > 35 cells). Experiments were repeated three times and averaged data are shown. The two-tailed Student’s t-test was used for the statistical comparisons.

Discussion

We established the novel RhIP assay, combining permeabilized cells and reconstituted histone complexes, to analyze histone incorporation. Previous histone incorporation analyses using genetically encoded histone genes have revealed chromatin dynamics, including nucleosome turnover kinetics, but have limitations on the time resolution, as they require time to synthesize and/or label histones (Deal and Henikoff, 2010). In contrast, the RhIP assay can detect histone incorporation with better time resolution, as it does not require histone synthesis or labeling. In fact, we could analyze histone incorporations in the early and late S phases separately, using synchronized cells (Figure 2—figure supplement 2 and Figure 3D). By combining RhIP with ChIP-seq, the RhIP assay enables the analysis of histone incorporation at the DNA sequence level, while ChIP-seq reveals their static presence. Scatter plot analyses of the H2A.Z RhIP-ChIP-seq replicates at known H2A.Z sites showed a quite high correlation (0.95 Pearson correlation coefficient), while the H2A and H2A.Z RhIP-ChIP-seq data showed a weaker correlation as compared with one of the replicates (Figure 3—figure supplement 2A). Thus, the RhIP-ChIP-seq analysis of H2A.Z also showed the reproducibility (Figure 3, Figure 3—figure supplements 1 and 2, Figure 4 and Figure 4—figure supplement 1) and better enrichment (S/N ratio) as compared with ChIP-seq (Figure 3—figure supplement 2B and C), probably due to the high-affinity antibody against the epitope-tag. Together with the fact that the RhIP-ChIP assay requires a small number of cells (approximately 8 × 106 cells), the RhIP assay is suitable for the biochemical analysis of ChIP samples. The effects of post-translational modifications (PTMs) of histones on their incorporation have remained elusive. Methods to produce histones with PTMs in vitro have been developed (Nadal et al., 2018), and thus the analysis of the effects of PTMs on histone incorporation will be the next target for the RhIP assay.

The proper combination of histone chaperones and histone variants ensures correct histone localization. Therefore, if only histone variants are increased, then the excess histone variants may bind to unsuitable histone chaperones, leading to ectopic localization. Indeed, the overexpressed histone H3 variant, CENP-A, in cells aberrantly binds to the H3.3 chaperone, DAXX, leading to ectopic localizations (Lacoste et al., 2014). In the present study, the ectopic localizations of histones were not observed. Therefore, there may be sufficient amounts of free histone chaperones under these conditions. We performed the RhIP assay simultaneously with two different histone variants, which do not share the same chaperone. This suggests that they do not affect each other’s incorporations. Since the amounts of exogenously added histones can be increased in the RhIP assay, it may be suitable for competition assays to analyze the overexpression of a specific histone variant.

The RhIP assay successfully reproduced the replication-coupled H3.1 incorporation and replication-independent H3.3 incorporation throughout the genome. Furthermore, the H3.1 and H3.3 incorporations required CAF-1 and HIRA, respectively, in this assay (Figure 1 and Figure 1—figure supplement 1). Although CAF-1 is present in permeabilized cells, replication-coupled H3.1 deposition required the cellular extract (Figure 1—figure supplement 2). This suggests that one (or more) unknown cytosolic factor(s) couples newly synthesized H3.1 to chromatin-binding proteins, such as the CAF-1 complex.

The present study has revealed the correlation between histone incorporation and chromatin structure, using the RhIP assay. We analyzed the incorporation of the H2A family members, H2A, H2A.X, and H2A.Z. We found that the incorporations of H2A-H2B and H2A.X-H2B into transcriptionally active open chromatin can occur in both replication-coupled and -independent manners, whereas their incorporation into transcriptionally inactive closed chromatin occurs only in a replication-coupled manner in late S phase (Figures 2, 3 and 6). This indicated that histone exchange would rarely occur in closed chromatin. Thus, the genome-wide localization of H2A and H2A.X is achieved by a combination of deposition into open chromatin and replication-coupled deposition into closed chromatin (Figure 7). In contrast, H2A.Z exhibited a much lower frequency of replication-coupled deposition, as compared to H2A (Figure 2F and Figure 2—figure supplement 2). Together with the fact that the amount of H2A.Z is much lower than that of H2A in S phase cells (Wu et al., 1982), we concluded that little to no H2A.Z is incorporated into closed chromatin (Figure 7). This is consistent with previous observations that the H2A.Z and DNA methylation localizations are mutually exclusive (Nothjunge et al., 2017; Zilberman et al., 2008), and that H2A.Z predominantly exists at promoters and enhancers (Barski et al., 2007; Buschbeck and Hake, 2017; Ernst et al., 2011; Jin et al., 2009; Link et al., 2018; Obri et al., 2014; Schones et al., 2008). The means by which H2A.Z becomes enriched at specific regions of open chromatin have remained enigmatic. Our study suggested that the H2A.Z elimination from transcriptionally inactive chromatin is due to the low frequencies of histone exchange and replication-coupled H2A.Z incorporation, which may partially explain the H2A.Z distribution pattern (Figure 7). Moreover, H2A.Z was specifically incorporated into chromatin around the TSS of expressed genes in the RhIP assay, in excellent agreement with the steady state localization of H2A.Z revealed by ChIP-seq (ENCODE Project Consortium, 2012). We also found that the H2A.Z depositions in active TSS and enhancer regions decreased in the ANP32E knockdown cellular extract, indicating that the ANP32E-mediated eviction promotes the new deposition of H2A.Z in the RhIP assay. However, other mechanism(s) may also exist, because the decrease in the H2A.Z deposition was relatively moderate (Figure 5C and D). This is consistent with the results that ANP32E knockout mice did not show any apparent effects on their viability (Reilly et al., 2010), even though proper H2A.Z incorporation is essential for cell survival and proliferation. Our data also showed that ATP promoted the H2A.Z deposition in active TSS and enhancer regions (Figure 5E and F), suggesting that the H2A.Z deposition is an ATP-dependent process. Taken together, the H2A.Z deposition in the RhIP assay is the net result of the ANP32E-mediated eviction and deposition of H2A.Z, and the replacement of pre-incorporated H2A with H2A.Z by ATP-dependent processes. We also found that the correlation between the RhIP-ChIP-seq and ChIP-seq of H2A.Z at known H2A.Z peaks in the gene body is lower, as compared with all known H2A.Z peaks (Figure 4C center and right). This may reflect the fact that the RhIP assay cannot assess histone recycling; That is, the re-deposition of the evicted histone, which is observed during transcription (Jeronimo et al., 2019; Tornà et al., 2020). H2A.Z is also found at pericentric heterochromatin (Boyarchuk et al., 2014; Greaves et al., 2007; Rangasamy et al., 2003). As human pericentric heterochromatin is composed of repetitive DNA sequences (satellite repeats), we could not analyze H2A.Z incorporation into pericentric heterochromatin by the RhIP-ChIP-seq assay (Figure 3). Previous studies showed that the transcriptional activation of pericentric satellites accompanies the structural changes from heterochromatin to euchromatin (Jolly et al., 2004; Rizzi et al., 2004; Saksouk et al., 2015; Valgardsdottir et al., 2005). In addition, the pericentric region of human chromosome nine is heterogeneous, with both heterochromatic and euchromatic regions (Gilbert et al., 2004). Therefore, H2A.Z incorporation into pericentric heterochromatin may occur, as in the open chromatin of the region.

Model of differential histone incorporations into open and closed chromatin.

In open chromatin, the H2A-H2B and H2A.X-H2B complexes are incorporated in replication-independent (RI) and replication-coupled (RC) manners, respectively, while H2A.Z-H2B is incorporated only in a replication-independent manner. In closed chromatin, new histone depositions of H2A and H2A.X, but not H2A.Z, occur only in a replication-coupled manner. This leads to the global localizations of H2A-H2B and H2A.X-H2B, as well as the specific localization of H2A.Z-H2B, including its elimination from closed chromatin.

We found that histone incorporations are regulated by the chromatin structure, which may be important for maintaining the closed chromatin configuration. Histones reportedly form a pre-deposition complex, which includes many transcription-related factors, before their incorporation into chromatin in vivo (Dunleavy et al., 2009; Mao et al., 2014; Obri et al., 2014; Tagami et al., 2004). For instance, the H2A-H2B pre-deposition complex contains Spt16 and SSRP1, which form the heterodimer complex FACT that functions in transcription facilitation, and the pre-deposition H2A.Z-H2B complex also includes a chromatin remodeling factor (SRCAP), a histone acetyltransferase (TIP60), and acetyl-lysine-binding proteins (GAS41, Brd8). If histone exchange usually occurs in closed chromatin, then these transcription-related factors might accumulate in the closed chromatin and alter the epigenetic chromatin states. Therefore, the deficiency of replication-independent histone exchange in closed chromatin may be important for maintaining a transcriptionally inactive state.

In spite of the high-sequence homology between H2A and H2A.Z, their localizations are different. By using the swapping mutant, we analyzed the residues responsible for the specific depositions of H2A and H2A.Z (Figure 6). We identified residues 88–100 of H2A as being responsible for its replication-coupled deposition. Although the means by which this region contributes to the incorporation into replicating chromatin remain unknown, a factor that binds this region and allows H2A to assemble at a replicating site may exist. The major H2A-specific chaperones, Spt16 and Nap1, which are components of the H2A pre-deposition complex, do not bind to this region (Aguilar-Gurrieri et al., 2016; Kemble et al., 2015). Thus, the protein that recognizes these residues is likely to be a chromatin protein involved in replication, rather than a component of the H2A pre-deposition complex. This region is a counterpart of the H2A.Z M6 region, and the H2A.Z_M6 swapping mutant did not compensate for the embryonic lethality of the H2A.Z knockout in Drosophila melanogaster (Clarkson et al., 1999). As this region is essential for the H2A.Z eviction by ANP32E (Gursoy-Yuzugullu et al., 2015; Obri et al., 2014), the incorporated H2A.Z_M6 mutant may not be removed from closed chromatin, thus resulting in aberrant gene expression and impaired embryonic development.

In conclusion, our novel method elucidated the mechanism of histone incorporation at the DNA sequence level, and revealed that the chromatin structure is the first determinant of histone localization.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
AntibodyAnti-HA
(mouse, monoclonal)
Santa Cruzsc-7392
RRID:AB_627809
IF(1:1,000)
AntibodyAnti-HA
(rabbit, monoclonal)
Cell Signaling Technology3724
RRID:AB_1549585
IF(1:2,000) only used in Figure 1—figure supplements 1D and 3D
AntibodyAnti-DDDDK (anti-FLAG) (rabbit, polyclonal)MBLPM020
RRID:AB_591224
IF(1:500)
AntibodyAnti-V5 (chicken, polyclonal)Abcamab9113
RRID:AB_307022
IF(1:1000)
AntibodyAnti-p60
(rabbit, monoclonal)
Abcamab109442
RRID:AB_10861771
IF(1:400)
AntibodyAnti-HIRA
(mouse, monoclonal)
Active MotifWC119.2H11
RRID:AB_10715607
IF(1:200)
AntibodyAnti-HIRA
(rabbit, monoclonal)
Abcamab129169
RRID:AB_11140220
WB(1:500)
AntibodyAnti-p60
(rabbit, monoclonal)
Abcamab109442
RRID:AB_10861771
WB(1:400)
AntibodyAnti-H3 (mouse, monoclonal)MBLMABI0301
RRID:AB_11142498
WB(1:1000)
AntibodyAnti-SRCAP
(rabbit, polyclonal)
KerafastESL103WB(1:1000)
AntibodyAnti-TIP60
(mouse, monoclonal)
Santa Cruzsc-166323
RRID:AB_2296327
WB(1:50)
AntibodyAnti-ANP32E (rabbit, polyclonal)MyBioSourceMBS9214243WB(1:1000)
AntibodyAnti-GAPDH (mouse, monoclonal)MBLML171-3
RRID:AB_10597731
WB(3:1000)
Sequence-based reagentGAPDH_FThis paperPCR primersAAAGGGTGCAGCTGAGCTAG
Sequence-based reagentGAPDH_RThis paperPCR primersTACGAAGCCCTTCCAGGAGA
Sequence-based reagentLINC02199_FThis paperPCR primersCCGGTGTCAAATGTCACAATGAA
Sequence-based reagentLINC02199_RThermo FisherPCR primersGGGGTTTTGAGGATTCCAAAGTG
Sequence-based reagentsi_p60 geneThis papersiRNAAAUGAUAACAAGGAGCCGGAGdTdT
Sequence-based reagentsi_p150 geneThis papersiRNACUGUCAUGUGGGUUCUGACdTdT
Sequence-based reagentON-TARGET SMARTpool siRNA (HIRA gene)DharmaconsiRNAL-013610-00-0005
Cell line
(Homo sapiens)
HeLaPeter R CookN/AA human cervical cancer cell line (female origin)
Cell line
(Homo sapiens)
SF8628MerckSCC127
RRID:CVCL_IT46
Human DIPG H3.3-K27M Cell Line

Cell culture and thymidine block

Request a detailed protocol

HeLa and SF8628 cells were cultured in DMEM supplemented with 10% fetal bovine serum, at 37°C in a 5% CO2 atmosphere. For cell synchronization, HeLa cells were cultured with 2 mM thymidine for 18 hr (Figure 1—figure supplement 2C). For the double thymidine block, the medium was then changed to remove the thymidine. After 9 hr of culture without thymidine, the HeLa cells were cultured with 2 mM thymidine again for 15 hr, for synchronization in S phase (Figure 2—figure supplement 2). All cell lines used in this study had their identities validated by STR profiling and it was also confirmed by Hoechst staining that there was no mycoplasma contamination.

Reconstitution of histone complex

Request a detailed protocol

All histones used in this study were produced in Escherichia coli cells as recombinant proteins. The human H3.1 and H3.3 genes were inserted in the pET21a vector (Novagen), as C-terminal 3HA-His6 or 3FLAG-His6 fused genes. The human H2A, H2A.X, H2A.Z, and H2A.Z_M6 genes were inserted in the pET15b vector (Novagen) as N-terminal His6-3HA, His6-3FLAG or His6-V5 fused genes. All the genes were overexpressed in the BL21(DE3) E. coli strain, by adding 0.5 mM isopropyl-β-D-thiogalactopyranoside. Each histone was then purified as described previously, using Ni-NTA affinity chromatography (Tachiwana et al., 2010). The epitope tag-fused histones were freeze-dried without removing the His6 tags. As the His6 tag was not used to detect or precipitate histones in this study, His6 was not mentioned in the text and figures, so that His6-epitope-tag-histone and histone-epitope-tag-His6 were referred to as epitope-tag-histone and histone-epitope-tag, respectively. Human H2B and H4 were overexpressed and purified after removing the His6 tags, as described previously (Tachiwana et al., 2010).

Freeze-dried H3.1 or H3.3 was mixed with H4 and H2A, H2A.X, H2A.Z, and H2A.Z_M6, along with H2B, in 20 mM Tris-HCl buffer (pH 7.5), containing 7 M guanidine hydrochloride and 20 mM 2-mercaptoethanol, and incubated on ice. After 1 hr, the samples were dialyzed against reconstitution buffer (10 mM Tris-HCl, pH 7.5, and 2 mM 2-mercaptoethanol) containing 2 M NaCl, overnight at 4°C. The NaCl concentration was then decreased by three steps of dialysis against reconstitution buffer containing 1 M NaCl for 4 hr, 0.5 M NaCl for 4 hr, and 0.1 M NaCl overnight. After the dialyses, the precipitants were removed by centrifugation and the supernatants were analyzed by Superdex 200 gel filtration chromatography (GE Healthcare).

Preparation of the cellular extract

Request a detailed protocol

The cellular extract was prepared as previously described, with modifications (Martini et al., 1998). Confluent HeLa cells in twenty-five 15 cm dishes were rinsed twice with ice-cold PBS in a cold room. Then, 10 ml of ice-cold hypotonic buffer (10 mM HEPES-KOH, pH 7.8, 10 mM KCL, 1.5 mM MgCl2, 1 mM dithiothreitol and 1 × proteinase inhibitor cocktail [cOmplete, EDTA-free, Roche]) was added to each dish to swell the cells, and the dishes were incubated for 5 min in a cold room. After repeating this step once, the excess buffer was removed by leaning the dishes against a wall for 5 min. The cells were then scraped off the dish and disrupted with 35 strokes in a 1 ml Dounce homogenizer with a loose-fitting pestle (Wheaton). Nuclei were removed by centrifugation at 1,500 × g for 5 min at 4°C, and the collected supernatant was further centrifuged at 14,000 × g for 5 min at 4°C. The supernatant was then flash-frozen with liquid nitrogen. After 30 min at −80°C, the cellular extract was thawed and the debris was removed. The concentration of the cellular extract was then measured with a protein quantification kit (MACHEREY-NAGEL). Usually, 3–4 ml of 5–8 mg/ml cellular extract were obtained.

For the preparation of the ATP-depleted cellular extract, HeLa cells were grown to confluence in a 15 cm dish, and further incubated for 60 min at 37°C in DMEM without glucose and fetal bovine serum, supplemented with 6 mM 2-deoxy-D-glucose and 10 mM sodium azide (Schwoebel et al., 2002). The cellular extract was then prepared as described above. Since the ATP-depleted cells were prone to detachment, all steps were performed gently.

For the preparation of the knockdown cellular extract, HeLa cells were transfected with siRNAs against the luciferase or ANP32E gene using the Lipofectamine RNAiMAX Transfection Reagent (Thermo Fisher), according to the manufacturer’s instructions. After 72 hr, the cellular extract was prepared as described above. The siRNAs used are as follows: siRNA against luciferase gene: CGUACGCGGAAUACUUCGAdTdT siRNA against ANP32E gene: AUGGAUUUGAUCAGGAGGAdTdT.

RhIP-immunostaining

Request a detailed protocol

HeLa cells were grown on a coverslip in a six well dish to 2.5 × 105 per well. The cells were chilled on ice and rinsed twice with 2 ml of ice-cold PBF (100 mM CH3COOK, 10 mM Na2HPO4, 30 mM KCl, 1 mM dithiothreitol, 1 mM MgCl2, 1 mM ATP, and 5% Ficoll). To permeabilize the cells, 1 ml of PBF containing 0.2% Triton X-100 was added, and the cells were incubated for 5 min on ice and then rinsed twice with 2 ml of ice-cold PBF. As the permeabilized cells were easily detached from the coverslips, all subsequent steps were performed gently. The coverslips with the attached permeabilized cells were moved onto parafilm laid on an aluminum block, and incubated at 30°C. Then, a 50 μl portion of a RhIP-reaction mixture, containing 100 nM histone complex, 3.6 μg/μl cellular extract, 2.5% Ficoll, 100 mM CH3COOK, 10 mM Na2HPO4, 30 mM KCl, 1 mM dithiothreitol, 1 mM MgCl2, 100 μM each of dNTPs, and NTPs (Roche) with or without 250 nM Cy5-dUTP (Enzo Life Sciences), was added to the permeabilized cells, and incubated for 60 min at 30°C. The coverslip was placed in a well of a 12-well plate, and the cells were washed twice for 5 min with 1 ml of PBS containing 0.1% Tween 20 (PBST), at room temperature. The cells were then fixed with 4% PFA (Electron Microscopy Sciences) in PBS for 20 min at room temperature, and rinsed three times with PBS. The cells were treated with 1% BSA in PBST for 1 hr at room temperature, and then incubated for 2 hr at room temperature with one of the following primary antibodies: anti-HA (mouse, 1/1000, Santa Cruz, sc-7392), anti-HA (only used in Figure 1—figure supplements 1D and 3D, rabbit, 1/2,000, Cell Signaling Technology, 3724), anti-DDDDK (FLAG) (rabbit, 1/500, MBL, PM020), anti-V5 (chicken, 1/1000, Abcam, ab9113), anti-p60 (rabbit, 1/400, Abcam, ab109442), and anti-HIRA (mouse, 1/200, Active Motif, WC119.2H11). The cells were washed with PBST three times for 10 min each, and then incubated for 1 hr at room temperature with secondary antibodies: Goat Alexa Fluor 488 or 546-conjugated anti-mouse IgG (1/1000, Life Technologies), or goat DyLight 488- or 550-conjugated anti-rabbit IgG or chicken IgY (1/1000, Thermo Fisher). The cells were washed with PBST three times for 10 min each. DNA was stained with Hoechst 33342. The samples were mounted with ProLong Gold (Life Technologies). The images in Figure 1, Figure 1—figure supplements 2 and 3, and Figure 2—figure supplements 1 and 2 were acquired by using the Deltavision set-up (Cytiva) with an inverted Olympus IX71 microscope, equipped with a CoolSNAP ES2 CCD camera (Photometrics) and a 60×, 1.42 Plan Apo N Olympus oil-immersion objective. Other images were acquired with an LSM 880 inverted confocal microscope (Zeiss), equipped with an AiryScan module and a 63×, 1.40 Plan-Apochromat Zeiss oil objective. All image files were converted to the TIFF format using the ImageJ software (Schneider et al., 2012) and imported into Illustrator (Adobe) for assembly. The co-localization analysis was performed using the ImageJ Colocalization_Finder plugin.

For the RhIP-immunostaining assay with the CAF-1- or HIRA-knockdown, HeLa cells were transfected with siRNAs against the luciferase (negative control), CAF-1 (p60), CAF-1 (p150), or HIRA gene using the Lipofectamine RNAiMAX Transfection Reagent (ThermoFisher), according to the manufacturer’s instructions. To knockdown the CAF-1 complex, the siRNAs against the CAF-1-p60 and CAF-I-p150 genes were added simultaneously. After an incubation for 8 hr, the cells were collected by trypsin/EDTA treatment, and the CAF-1- or HIRA-knockdown cells were mixed with control-knockdown cells and re-plated and co-cultured for 64 hr. For the CAF-1-knockdown experiment, 2 mM of thymidine was added to synchronize the cells in S phase, 18 hr before the permeabilization. Immunostainings were then performed as described above. siRNAs used for this experiment are as follows: siRNA against p60 gene: AAUGAUAACAAGGAGCCGGAGdTdT siRNA against p150 gene: CUGUCAUGUGGGUUCUGACdTdT siRNA against HIRA gene: ON-TARGET SMARTpool siRNA L-013610-00-0005 (Dharmacon).

RhIP-ChIP

Request a detailed protocol

The RhIP-ChIP assays shown in Figures 1H and 2F and Figure 2—figure supplement 2 were performed as follows. Semi-confluent HeLa cells in a 10 cm dish were chilled on ice, and rinsed twice with 5 ml of ice-cold PBF. To permeabilize the cells, 2.5 ml of PBF containing 0.2% Triton X-100 was added and incubated for 5 min on ice. The cells were rinsed twice with 5 ml of ice-cold PBF, and then 2.5 ml of the RhIP-reaction mixture was added to the permeabilized cells. After sealing the lid of the dish with parafilm, the cells were incubated for 60 min at 30°C. The RhIP-reaction mixture was then removed, and the cells were washed twice with 5 ml of PBST at room temperature for 5 min each. Subsequently, native-ChIP was performed as described previously with minor modifications (Tachiwana et al., 2015). The cells were collected in 1 ml of NB buffer (15 mM Tris-HCl, pH 7.5, 15 mM NaCl, 60 mM KCl, 300 mM sucrose and 1 × proteinase inhibitor cocktail (cOmplete, EDTA-free, Roche)). After centrifugation at 1500 × g for 5 min at 4°C, the cells were resuspended in 100 μl of NB buffer. CaCl2 was then added to a final concentration of 2 mM. The cells were treated with 150 mU/μl of micrococcal nuclease (MNase, Worthington) for 5 min at 37°C, to generate soluble chromatin fragments. The reaction was terminated by adding 10 mM EDTA, and the solubilized chromatin fragments were separated from the pellets by centrifugation at 16,000 × g for 5 min at 4°C. The samples were then mixed with 50 μl of anti-HA-tag mAb-Magnetic Beads (MBL International) in 15 mM Tris-HCl, pH 7.5, 300 mM NaCl, and 0.1% NP-40. After an overnight incubation at 4°C with gentle mixing on a wheel, the beads were washed three times each with 500 μl of ChIP buffer (10 mM Tris-HCl, pH 8.0, 200 mM KCl, 1 mM CaCl2, 0.5% NP-40), ChIP buffer containing 500 mM KCl, and TE. The DNA was then eluted with a Proteinase K solution (20 mM Tris-HCl, pH 8.0, 20 mM EDTA, 0.5% sodium dodecyl sulfate (SDS), and 0.5 mg/ml Proteinase K) and extracted with phenol-chloroform. The DNA was precipitated with ethanol and resuspended in 10 mM Tris-HCl, pH 8.0, and 1 mM EDTA. The resulting DNA samples were separated by electrophoresis on a 2% agarose gel (7.4 V/cm, 35 min) (Figures 1H and 2F) or a 1.5% agarose gel (7.1 V/cm, 60 min) (Figure 2—figure supplement 2B) in 1 × TAE, and stained with SYBR Gold (Thermo Fisher). Images of the DNA stained with SYBR Gold and the nascent DNA labeled with Cy5 were obtained with an Amersham Typhoon scanner (Cytiva).

The H3.3 enrichment shown in Figure 1E was analyzed by qPCR with a StepOne Plus system (Applied Biosystems) using SYBR Green fluorescence. The cycle number required to reach the threshold was recorded and analyzed. PCR was performed using the same amounts of the immunoprecipitated DNA (RhIP-ChIP sample) and the input DNA. Values were normalized to the input DNA. Primer sets used for this analysis are as follows:

  • GAPDH forward: AAAGGGTGCAGCTGAGCTAG

  • GAPDH reverse: TACGAAGCCCTTCCAGGAGA

  • LINC02199 forward: CCGGTGTCAAATGTCACAATGAA

  • LINC02199 reverse: GGGGTTTTGAGGATTCCAAAGTG.

RhIP-ChIP-seq

Request a detailed protocol

RhIP-ChIP-seq was performed in the same way as RhIP-ChIP with modifications. HeLa cells were used in Figures 35 and SF2868 cells were used in Figure 1—figure supplement 1A. After the RhIP assay, the permeabilized cells were washed twice with PBST for 5 min at room temperature, and then cross-linked with 3% formaldehyde in PBS for 5 min at room temperature. After fixation, the cells were resuspended with 1 ml of ChIP buffer and collected by centrifugation at 1500 × g for 5 min at 4°C. The cells were then treated with 7.5 mU/μl MNase in 400 μl ChIP buffer for 30 min at 37°C. The MNase reaction was terminated by adding 10 mM EDTA, and the solubilized chromatin fragments were separated from the pellets by centrifugation at 16,000 × g for 5 min at 4°C. The supernatant was incubated with 50 μl of anti-HA-tag mAb-Magnetic Beads, and gently mixed by rotation at 4°C overnight. The beads were then washed three times each with 500 μl of ChIP buffer, ChIP buffer containing 500 mM KCl, and TE. The beads were resuspended in 100 μl of ChIP elution buffer (50 mM Tris-HCl, pH 8.0, 10 mM EDTA, 25 mM NaCl, and 1% SDS), and incubated overnight at 65°C to reverse the cross-links. The DNA was then eluted with 0.4 mg/ml Proteinase K, and further purified with a PCR clean-up kit (MACHEREY-NAGEL). The DNA libraries were prepared using a SMARTer ThruPLEX Tag-seq Kit (Takara Bio) for Figures 3 and 4, or a SMARTer ThruPLEX DNA-seq kit (Takara Bio) for Figure 5. The samples were sequenced on an Illumina HiSeq 1500 system for Figures 3 and 4, and an Illumina HiSeq X Ten system for Figure 5.

For the spike-in normalization performed in Figure 5, one-tenth of the fragmented chromatin was subjected to cross-link reversal prior to the ChIP procedure. After an overnight incubation at 65°C, the resulting DNA was recovered as described above and the concentration was measured. Then, one two-hundredth of Drosophila spike-in chromatin (Active Motif) was mixed with the rest of the chromatin. Two μg of spike-in antibody (Active Motif) per 50 ng of Drosophila spike-in chromatin were mixed with ProteinA/G magnetic beads (Pierce). The spike-in antibody beads were then mixed with 50 μl of anti-HA-tag mAb-Magnetic Beads and the chromatin mixture. Subsequent steps were performed as described above.

ChIP‐seq data analysis

Request a detailed protocol

For ChIP-seq data shown in Figures 3 and 4, the in-read unique-molecular-identifiers (tags) were extracted using UMI-tools (Smith et al., 2017) with the command: umi_tools extract --extract-method=regex --bc-pattern='(?p<umi_1>.{6})(?p<discard_1>.{0,3}GTAGCTCA){s <= 2}'. The extracted reads were mapped to the human genome (GRCh38) using hisat2 (version 2.1.0) (Kim et al., 2015). The PCR-duplicates were removed using umi_tools dedup. The read counts on 15 chromatin states were calculated using BED-Tools (Quinlan and Hall, 2010). The definition of the ChromHMM track was obtained from the consolidated data set of the Roadmap Epigenomics project (E117_15_coreMarks_hg38lift_mnemonics.bed) (Roadmap Epigenomics Consortium et al., 2015). The BED files of enhancer and insulator regions used in Figure 5 were also extracted from ChromHMM track. The overall concentrations of the ChIP signals (log2 ratio) were calculated as the ratio of the proportion of reads in each chromatin state between the ChIP and input DNA data; that is, log2(ChIP/Input) after normalization of the total reads. The signal tracks (bigwig files) were created at 1 bp intervals on the genome, and then the counts were normalized as CPM (Reads Per Million reads), using deepTools (Ramírez et al., 2014). ChIP-seq signals were visualized with the Integrative Genomics Viewer, IGV (Robinson et al., 2011). The Pearson correlation coefficients were calculated throughout 800 bp intervals with the multiBigwigSummary program of deepTools, and plotted with the plotCorrelation program (Ramírez et al., 2014). The multiBigwigSummary program was also used to compute the average scores for RhIP-ChIP-seq data and ChIP-seq data (GEO:GSM1003483) at H2A.Z sites, which were determined with replicates of the ChIP-seq data (GEO:GSM1003483) by the mergePeaks program of HOMER (Heinz et al., 2010). The computeMatrix programs of deepTools were utilized to analyze the peak localizations at the centers of TSS, the known H2A.Z sites, and the H2A.Z peaks of the RhIP-ChIP-seq data. Genes with five or more read counts in the RNA-seq data (GSE140768 for Figure 1—figure supplement 1, GSE123571 for Figure 4B) were defined as expressed genes, and the rest were regarded as non-expressed genes. The H2A.Z peaks of the RhIP-ChIP-seq data were also determined with the mergePeaks program of HOMER. The plotFingerprint program of deepTools was utilized to analyze the enrichment of ChIP signals.

For the spike-in normalization used in Figure 5, the reads were mapped to the human genome (GRCh38) and the fly genome (BDGP Release 6 + ISO1 MT/dm6), using bowtie2 (version 2.3.4.3) (Langmead and Salzberg, 2012). The PCR-duplicates were removed using the MarkDuplicates tool of Picard (Broad Institute, 2019). Normalization factors were then generated using the numbers of mapped reads on the fly genome. The numbers of mapped reads on the human genome were then downsampled, using the view program of SAMtools according to the normalization factors (1000 Genome Project Data Processing Subgroup et al., 2009).

Purification of NAP1 and ASF1

Request a detailed protocol

Human NAP1 was purified as described previously (Tachiwana et al., 2008). The human ASF1 gene was inserted in the pET15b vector (Novagen), in which the thrombin proteinase recognition sequence was replaced by the PreScission protease recognition sequence. Escherichia coli strain BL21-CodonPlus (DE3)-RIL cells (Agilent Technologies) were freshly transformed with the vector, and cultured at 30°C. After the cell density reached an A600 = 0.8, 1 mM isopropyl- β-D-thiogalactopyranoside was added, and the culture was continued at 18°C for 12 hr to induce His6-tagged Asf1 expression. The cells were collected and resuspended in 20 mM Tris-HCl, pH 7.5, containing 500 mM KCl, 10% glycerol, 0.1% NP-40, 1 mM phenylmethylsulfonyl fluoride (PMSF), and 2 mM 2-mercaptoethanol. After cell disruption by sonication, the debris was removed by centrifugation (27,216 × g; 20 min), and the clarified lysate was mixed gently with 4 ml (50% slurry) of nickel-nitrilotriacetic acid (Ni-NTA)-agarose resin (Qiagen) at 4°C for 1 hr. The Ni-NTA beads were washed with 200 ml of 20 mM Tris-HCl, pH 7.5, containing 500 mM NaCl, 10% glycerol, 10 mM imidazole, and 2 mM 2-mercaptoethanol. The His6-tagged Asf1 was eluted by a 100 ml linear gradient of 10 to 500 mM imidazole in 50 mM Tris-HCl buffer (pH 7.5), containing 500 mM NaCl, 10% glycerol, and 2 mM 2-mercaptoethanol. PreScission protease (8 units/mg protein, Cytiva) was added to remove the His6 tag from the ASF1. The sample was dialyzed against 20 mM Tris-HCl, pH 7.5, containing 100 mM NaCl, 1 mM EDTA, 10% glycerol, and 2 mM 2-mercaptoethanol. The ASF1 was further purified by chromatography on a Mono Q (Cytiva) column, eluted with a 25 ml linear gradient of 100–600 mM NaCl in 20 mM Tris-HCl, pH 7.5, containing 1 mM EDTA, 10% glycerol, and 2 mM 2-mercaptoethanol. The eluted Asf1 was further purified by chromatography on a Superdex 75 (Cytiva) column, eluted with 1.2 column volumes of the same buffer containing 100 mM NaCl. The Asf1 was repurified by Mono Q chromatography, concentrated, and dialyzed against 20 mM Tris-HCl (pH 7.5), containing 150 mM NaCl, 1 mM dithiothreitol, 0.5 mM EDTA, 0.1 mM PMSF, and 10% glycerol.

Subcellular fractionation and western blotting

Request a detailed protocol

Semi-confluent HeLa cells in a 10 cm dish were chilled on ice and rinsed twice with 5 ml of ice-cold PBF. To permeabilize the cells, 2.5 ml of PBF containing 0.2% Triton X-100 was added, and incubated for 5 min on ice. The supernatant was collected as the soluble fraction. The permeabilized cells were rinsed twice with 5 ml of ice-cold PBF and scraped off the dish. After centrifugation at 800 × g for 5 min at 4°C, the pellets were resuspended in 250 μl of RIPA buffer (50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 1 × protease inhibitor cocktail and 0.1% SDS) and treated with benzonase. The supernatant was then collected by centrifugation at 16,000 × g for 5 min at 4°C as the soluble fraction. The insoluble fraction was diluted ten-fold with PBF containing 0.2% Triton X-100 to adjust the dilution ratio between the soluble and insoluble fractions.

Samples for western blots were separated by SDS-8% PAGE and transferred onto PVDF membranes (Cytiva) with a semi-dry blotter (Bio-Rad) at 120 mA for 60 min. For SRCAP, samples were separated by SDS-6% PAGE and transferred onto PVDF membranes with a wet blotter (Bio-Rad) in an ice-cold 1 × AquaBlot (Wako) solution containing 0.05% SDS, at 120 V for 60 min. After blotting, the membranes were blocked with blocking One (Nacalai), and then probed with the following primary antibodies: anti-HIRA (rabbit, 1/500, Abcam, ab129169), anti-p60 (rabbit, 1/400, Abcam, ab109442), anti-H3 (mouse, 1/1000, MBL, MABI0301), anti-SRCAP (rabbit, 1/1000, Kerafast, ESL103), anti-TIP60 (mouse, 1/50, Santa Cruz, sc-166323), anti-ANP32E (rabbit, 1/1000, MyBioSource, MBS9214243) or anti-GAPDH (mouse, 3/1000, MBL, ML171-3). For the secondary antibodies, sheep horseradish peroxidase (HRP)-conjugated anti-mouse IgG or -rabbit IgG (1/10,000, Cytiva) was used. Signals were developed using Chemi-Lumi One Super (Nacalai) and were detected by an Amersham Imager 680 (Cytiva).

Data availability

The deep sequencing data in this study are available through the NIH GEO Database under the accession numbers GEO: GSE163502, GSE130947 and the DNA DATA bank of Japan under the accession number DDBJ: DRA009580.

The following data sets were generated
    1. Tachiwana H
    2. Maehara K
    3. Harada A
    4. Ohkawa Y
    (2021) NCBI Gene Expression Omnibus
    ID GSE130947. RhIP-ChIP-seq of H2A and H2A.Z using asynchronous, early S and late S phase cells.
    1. Tachiwana H
    (2021) NCBI Gene Expression Omnibus
    ID GSE163502. RhIP-ChIP-seq of H2A.Z under ANP32E or ATP depletion condition.
The following previously published data sets were used

References

    1. Costanzi C
    2. Stein P
    3. Worrad DM
    4. Schultz RM
    5. Pehrson JR
    (2000)
    Histone macroH2A1 is concentrated in the inactive X chromosome of female preimplantation mouse embryos
    Development 127:2283–2289.
    1. Louters L
    2. Chalkley R
    (1985)
    Histones H1, H2A, and H2B in vivo
    Biochemistry 24:3080–3085.
    1. Passarge E
    (1979)
    Emil Heitz and the concept of heterochromatin: longitudinal chromosome differentiation was recognized fifty years ago
    American Journal of Human Genetics 31:106–115.

Decision letter

  1. Jessica K Tyler
    Senior Editor; Weill Cornell Medicine, United States
  2. Francesca Mattiroli
    Reviewing Editor; Hubrecht Institute, Netherlands

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Acceptance summary:

The method presented in this article, named RhIP, will be of interest for all fields that interface with chromatin dynamics. It provides a new tool to dissect the mechanisms of chromatin assembly and disassembly genome-wide, and to determine how cell cycle and chromatin structure influence these dynamics.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your manuscript to eLife as a Tools and Resources article. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The reviewers read each other's reviews and discussed their comments.

Although the general enthusiasm is high, significant doubts remain with regard to how faithfully RhIP recapitulates endogenous remodeling activities. In addition, it is unclear how feasible it is to use RhIP to dissect mechanism. For example, can a specific remodeling pathway be blocked by factor depletion? Since substantial amounts of additional work are required to substantiate the conclusions claimed in the paper, we are unable to accept the manuscript in the current form. We are however happy to reconsider if the following concerns are addressed.

1. Given the unique genomic localization of H2A.Z, the authors wisely used H2A.Z as a benchmark to assess the robustness of the RhIP approach. Indeed, site-specific deposition was observed when permeablized cells were incubated with recombinant H2A.Z and (cytosolic) extracts. While this is an interesting result, it remains unclear how well the H2A.Z sites observed by RhIP-ChIP-seq correlate with endogenous H2A.Z sites. The authors should report the percentage of H2A.Z sites correctly identified by RhIP. Scatter plots should be used to compare the relative occupancy of H2A.Z between the RhIP-ChIP-seq data and known H2A.Z sites. Are there H2A.Z sites in the RhIP-ChIP-seq data not seen in vivo. When presenting anecdotal evidence as in Figure 4A, endogenous H2A.Z tracks should be included for comparison.

2. Genomic data analyses generally lack rigor. This is epitomized in Figure 4-supplement 2. The fraction-against-rank plots give little information on the concordance of the two replicates. Do the H2A.Z sites identified in the replicates correlate spatially (e.g. along the genome), quantitatively (e.g. peak height), and qualitatively (e.g. width of peaks)? For example, relative H2A.Z occupancy at known H2A.Z sites of the two H2A.Z RhIP-ChIP-seq replicates should be plotted against each other. The H2A RhIP-ChIP-seq datasets could serve as negative control.

3. It is important to assess how individual factors contribute to a particular remodeling pathway. Given that the steady-state H2A.Z occupancy is a net result of deposition and eviction and that multiple remodeling factors are involved in these processes, whether the observed H2A.Z peaks in RhIP-ChIP-seq represent the consequence of one or more of these factors should be evaluated. For example, are SRCAP and p400/TIP60 responsible for different H2A.Z sites across the genome? Is ATP required for H2A.Z deposition? Is ANP32E responsible for removing H2A.Z at enhancers and insulators? Does transcription contribute to H2A.Z eviction? Proper spike-in controls should be incorporated into the RhIP-ChIP-seq samples before and after the addition of factor-depleted extracts to allow quantitative assessment. Since cytosolic extracts were used, it is not surprising that some factors may be under represented. If such biases do exist, the authors should consider using nuclear extracts and/or provide evidence on how much remodeling factors partitioned into the fractions relative to input (e.g. SRCAP western blots).

4. The title suggests that RhIP is generally applicable to all histones. But there was no mention of the H3 results in the Abstract or the Discussion. In addition, while immunostaining shows H3.1 and H3.3 are incorporated into the nucleus of permeabilized cells at the expected cell cycle stages, microscopy lacks the resolution necessary to validate if H3.1 and H3.3 are targeted to the expected genomic locations. For example, is H3.3 more enriched at active genes? CAF1 and HIRA knockdown should be used to verify if H3.1 and H3.3 are indeed deposited by these pathways. Finally, to show that H3.1 and H3.3 are indeed inserted into nucleosomes, the MNase experiment similar to the one in Figure 3C should be performed.

5. Major improvement of the description of methods is required to allow rigorous assessment of the approach. The current methods section lacks critical details necessary to carry out the experiments.

Reviewer #1:

Tachiwana and coworkers have developed a histone deposition assay called RhIP that recapitulates site-specific incorporation of tagged histones into the chromatin of permeabilized cells using cellular extracts. The authors showed that RhIP recapitulates replication-dependent and -independent deposition of histone H3.1 and H3.3, respectively. They also showed that RhIP allows site-specific deposition of H2A, H2A.X and H2A.Z. However, the paper falls short in showing that RhIP can actually be used to dissect molecular mechanisms. For example, if extracts from chaperone/remodeler-depleted cells were used, could a specific histone deposition step be blocked? Could purified proteins then be added back to restore histone deposition activity? In addition, what contribution endogenous factors (i.e. from the permeabilized cells) have on RhIP is unclear.

An eLife's Tools and Resources article does not require to report major new biological insights; however, it must be able to demonstrate such advances can be made by applying the new tools. In this respect, whether the RhIP approach can be used to dissect mechanism cannot be evaluated. Therefore, I do not recommend the manuscript in the current form to be published in eLife.

Other major issues:

1. 'Open chromatin' and 'close chromatin' are loosely defined terms. The authors should use specific histone marks (e.g. H3K4me3 or H3K27me3) as reference for co-localization analysis of the incorporated histones.

2. Based on the data presented in Figure 4, the robustness of site-specific incorporation of H2A.Z using RhIP cannot be rigorously evaluated. For example, while H2A.Z appears to be deposited at the promoters of some genes, there are H2A.Z peaks outside of promoters. Are these mis-incorporated H2A.Z molecules (i.e. RhIP artifact)? Or are they bona fide H2A.Z sites. The authors should provide endogenous H2A.Z ChIP-seq data for comparison and evaluate how well RhIP recapitulates endogenous H2A.Z deposition.

3. Transfection reagents for proteins, such as the Chariot system, have previously been used to deliver chromatin factors into mammalian cells (Larson AG et al. Nature 2017. PMID:28636604). What is the deposition efficiency of RhIP compared to Chariot?

4. A related question is how efficient is RhIP? What is the percentage of nascent histones incorporated by RhIP? The authors should perform western blots to compare the relative amounts of tagged and untagged histones.

Reviewer #2:

The manuscript by Tachinawa et al. presents a new method (called RhIP) that monitors incorporation of histone dimers into permeabilized cells. The authors test H3.1 and H3.3 containing H3-H4 dimers and then they primarily focus on the incorporation of H2A-H2B dimers and their variants containing H2A.X or H2A.Z. The readouts of the assay range from immunofluorescence, to Chromatin IP (ChIP) and ChIP-seq.

Overall, the method is interesting and may become a powerful tool to study chromatin dynamics. However, I have some comments that may improve the current manuscript.

1. As this has been submitted as a Tool and Resource article, I was expecting a much clearer explanation of how the RhIP is carried out. In the method section, every step of the procedure need to be spelled out more clearly. Currently one has to go back to older publications and it becomes quite difficult to recapitulate what has been done in details.

2. If I understand correctly, the authors co-incubate different histone dimers In Figure 1-2 and 5, but not in Figure 3 (and 4 perhaps). This means that the Immunofluorescence results depend on the "competition" between the H2A and the H2A.Z (or h3.1-H3.1) dimers, while the ChIP and ChIP-seq data in figure 3 and 4 only look at incorporation of a single dimer type (without competition for binding between different variants). This makes the comparison of the results quite challenging. What would the result of Figure 1-2 and 5 be if the different histone dimers would be added individually, rather than as a mixture? This would greatly clarify what are the properties that control dimers incorporation in this assay.

3. The authors do not really acknowledge that the incorporation observed in their studies is the net result of assembly AND disassembly pathways. This needs to be part of the narrative, because the experiments do not address which of these pathways is involved and to what extent. For example, on page 11, lines 10-14, or in page 12, line 1-2 or line 18-19, only suppression of deposition mechanisms are taken into account in the interpretation of the data, but a strong disassembly activity here may result in the same effect.

4. The MNase digestions in Figure 3 are very interesting. Can the authors include a higher exposure gel to show that the overall incorporation of Cy5dUTP is the same for H2A and H2A.Z input samples?

5. In Figure 4 C and E (and supplement 1), are the differences between samples statistically significant, or are these only trends? And what about Figure 1D?Please clarify this.

6. The title and the discussion suggest that the RhIP method is generally applicable and validated both for "histones" (as I understand it, it is H3-H4 and H2A-H2B (and variants thereof)). I found the H3-H4 part rather thin compared to the H2A-H2B (and variant) analysis. This is to the point that H3-H4 are completely omitted from the final model of the manuscript. Can the author address this with a title and discussion that focuses on H2A-H2B rather than "histones"?

Reviewer #3:

The manuscript "Chromatin structure-dependent histone incorporation revealed by a genome-wide deposition assay" by Tachiwana et al. put-forward a novel method, termed RhIP that allows a detailed investigation of mechanisms fostering histone deposition. In this approach the authors substitute permeabilized cells with recombinantly expressed tagged histones and cytosolic extracts (or chaperone proteins), and then measure histone chromatin deposition using various methods, ranging from immunofluorescence microscopy to RhIP/ChIP-sequencing. While this is an interesting method, some technical questions remain that need to be addressed and clarified. Most importantly, the authors need to show that the factors (chaperones) responsible for a faithful temporal and spatial histone variant deposition (or ejection) are actually still present in the permeabilized cell nucleus or available in the cell extract. The observation that H2A.Z is deposited into open but not condensed chromatin, could be explained by lack of (one) H2A.Z-specific chaperone/remodelling complex(es) after permeabilization. The authors need to demonstrate that these observations are not caused by a rather artificial experimental system that might not resemble the in vivo situation.

My specific concerns are listed in more detail below:

1. There are some clarifications on technical aspects of the RhIP assay missing. As I understood it, this assay was performed with a cytosolic extract, at least this is mentioned in the Materials and methods section. It would be interesting to see whether the known H2A.Z-specific chromatin remodeller complexes, such as SRCAP, p400/TIP60, ANP32E, INO80, etc. that are mostly nuclear proteins and required for H2A.Z deposition/ejection, are also present in this extract. The extracts as well as the cells after permeabilization and reconstitution should be immunoblotted for these chaperones to verify their presence. Additionally, have the authors performed these experiments in the presence or absence of ATP?

2. Figure 2: The authors have shown in Figure 2E that the H2A.Z signal overlaps with the dUTP signal in early but not in late S-phase cells. This result is in contrast to other studies using EdU-staining and microscopy in living cells that observed H2A.Z signals only overlapping with late (constitutive heterochromatin) replicating cells (e.g. Boenisch et al., 2012) and other studies showing H2A.Z levels at transcriptional start sites (euchromatin, early replicating) to be decreased during S-phase (e.g. Nekrasoph et al., 2012, NSMB). This discrepancy needs to be explained and discussed.

3. Figure 5: This is a relevant experiment, and a RhIP-ChIP-seq experiment should be included to show in greater detail that H2A.Z-M6 resembles more H2A than H2A.Z when looking at TSS (see Figure 4B).

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for submitting your article "Chromatin structure-dependent histone incorporation revealed by a genome-wide deposition assay" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Jessica Tyler as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential Revisions:

Here we share detailed comments to tackle the limitations of the study:

1. While replication-dependent mechanisms are well captured by RhIP, it is less clear if transcription and chromatin remodeling is functional in this system and thus if transcription-dependent nucleosome exchange processes are faithfully recapitulated. It is important to improve the comparison of RhIP with 'in vivo' (i.e. existing ChIP-seq datasets) localisation and explicitly develop hypotheses why in some cases the data matches the 'in vivo' situation and in others not.

a. Lines 279-301 and Figure 4. There is moderate correlation between genome-wide H2A.Z RhIP-ChIP-seq and H2A.Z ChIP-seq (Figure 4C). However, it seems H2A.Z RhIP-ChIP might be selectively detecting TSS specific signals rather than gene body specific associations (Figure 4A and 4E). It would be helpful to comment on the genetic elements (e.g. gene body) at which H2A.Z RhIP-ChIP and H2A.Z ChIP show low or no correlation and make a comparison. This way strengths and weaknesses of the assay can be better evaluated and compared to already existing protocols. The image resolution of Figure 4A should be improved.

b. Suppl. Figure 1.1 H3.3 distribution across the active and inactive genes were plotted in Suppl Figure 1. However, how active and inactive status of genes was defined, and which data was utilized for this analysis were not indicated under the Methods. These parameters and data source should be included. Transcript levels for GAPDH and LNC02199 should also be indicated or the reader should be referred to a table containing transcription data (i.e. RNA-Seq). I also think that it would be helpful if the authors can comment on the efficiency of H3.3-HA ChIP compared to pull down of endogenous H3.3 by comparing RhIP-ChIP-seq and previously reported H3.3 ChIP-seq analyses.

c. The genomics analysis presented in Figure 3 is somewhat rudimentary, the signals from H2A and H2A.Z appear overall very correlated (Figure 3A, B), albeit the correlation genome-wide calculated in Figure 3-S2 is 'medium'. The scatter plot could be shown better to see potentially different 'population of loci, some that correlate well and 'outliers'. Judging from Figure 3B, it appears that there is a good correlation across broad chromatin states with the notable excavation of active TSS. This is in line with the known H2A.Z mechanism, but could be elaborated in plots a bit clearer. How do the profiles look across highly expressed and non-expressed genes?

The cell-cycle-synchronized plots are only summarized in the heatmap in Figure 3C,D. Example loci could be shown as in Figure 3A.

2. It would be helpful to improve the interpretation of the data to include all existing caveats to the assay setup.

a. The new ATP addition experiments (figure 5E-F) are interpreted as if the only effect of ATP is on chromatin remodellers. It is well possible that by adding ATP in this conditions, additional factors (i.e. transcription, splicing/translation, molecular chaperones, etc) are being affected and responsible for the effect.

b. The authors alternate setups where they either co-incubate different histone dimers, or they use a single histone dimer isoform. It is really unclear to the readers what is the reason for this and how does this affect the results/interpretation. In cases where a mixture of histone dimers were present (i.e. Figures 1A-D, 2, 6), competition between the H2A/H2A.Z-X or H3.1/H3.3 dimers may take place, while in the others this competition is not present. This makes the comparison of the results quite challenging, and their conclusions unclear. This should be discussed at the very least, as it has implications on understanding the properties that control histone incorporation during RhIP.

c. The RhIP-ChIP-seq protocol uses formaldehyde crosslinking instead of native mononucleosomal IP as used in Figure 1H, leaving room for interpretation if the pull-down histone is indeed incorporated into chromatin. Immunofluorescence also does not necessarily prove incorporation of histones into chromatin, since i.e. the fractionation in Figure 1-S2 shows that HIRA and CAF1 remain tightly bound to the isolated nuclei. Thus, accumulation of the epitope-tagged H3 in the nucleus of cells could merely mean that is bound to the histone chaperones present in the nucleus (competing away endogenous histones). It is not possible to discern from the data what the incorporated amount versus the chaperone-bound fraction of recombinant histone is.

d. The functional assessment of ANP32E knockdown lacks statistical analysis – clearly the quite robust knockdown (Figure 5B) does not majorly abrogate TSS incorporation of H2A.Z. Is the difference statistically significant? What is the explanation for the relatively small effect size?

e. Lines 275-277. Please revisit the conclusion and make the statement clearer on H2A.Z incorporation and elimination.

We suggest adding these references:

The authors make a thorough effort to place their method in the context of existing pulse-labeling methods, but Tet-ON based pulsing methods, which do give similar results have not been discussed: Mito, Y., Henikoff, J. G. and Henikoff, S. Genome-scale profiling of histone H3.3 replacement patterns. Nat. Genet. 37, 1090-1097 (2005); Yildirim, O. et al. A system for genome-wide histone variant dynamics in ES cells reveals dynamic MacroH2A2 replacement at promoters. PLoS Genet. 10, e1004515 (2014); Ha, M., Kraushaar, D. C. and Zhao, K. Genome-wide analysis of H3.3 dissociation reveals high nucleosome turnover at distal regulatory regions of embryonic stem cells. Epigenetics Chromatin 7, 38 (2014).

Additional notes (not needed for revision):

Finally we list additional suggestions that were made by the reviewers. These comments are shared with you, but the reviewers do not expect these experiments to be performed in the revised version of the manuscript. We hope these will help you improve the discussion part and future studies.

Figure 1

Deposition of H3.3 seems rather inefficient as judged from the genome-wide profiling RhIP-ChIP-seq experiment in Figure 1-S1, and deposition appears to only occur at TSS but not across the gene bodies of active genes as observed in cells (please make sure to provide primary data on GEO for a revised submission). As I understand, the RhIP-ChIP-seq protocol uses formaldehyde crosslinking instead of native mononucleosomal IP as used in Figure 1H, leaving room for interpretation if the pull-down histone is indeed incorporated into chromatin. Immunofluorescence also does not necessarily prove incorporation of H3.1/H3.3 into chromatin, since the fractionation in Figure 1-S2 shows that HIRA and CAF1 remain tightly bound to the isolated nuclei. Thus, accumulation of the epitope-tagged H3 in the nucleus of cells could merely mean that is bound to the histone chaperones present in the nucleus (competing away endogenous histones). It is not possible to discern from the data what the incorporated amount versus the chaperone-bound fraction of recombinant histone is. Potentially, the product of the RhIP reaction could itself be fractionated using e.g. a salt gradient, which would provide evidence if the retained H3 histones are nucleosomal (retained up to ~500mM NaCl) or non-nucleosomal and dissociating at lower salt concentrations.

Further, in Figure 1H shoes that Cy5-labeled mononucleosomes are at very low abundance within the input (almost undetectable Cy5 signal in lanes 2,3,4) but highly enriched in both H3.1 and H3.3 IP. The fraction of replicated chromatin in each lane can be estimated by the Cy5/SYBR gold ratio. Assuming completely random (i.e. replication-independent) incorporation of recombinant histones, the Cy5/SYBR gold ration after IP (lanes 6,7) should be equivalent to that ratio before IP (lanes 4,5,6). Yet, in lanes 6,7, there's much more Cy5 signal thus, a strong enrichment for replicating chromatin. This suggest that not only H3.1 but also a considerable fraction of H3.3 is incorporated replication-dependent in this assay. This is entirely possible given the known role of HIRA/H3.3 in gap-filling after replication fork, especially if normal replication-dependent pathways are impaired (Ray-Gallet et al., Mol Cell, 2011). Again it needs to be stressed that the IF experiment Figure 1C do not necessarily prove that H3.3 is incorporated into nucleosomes in cells not undergoing replication.

Lastly, the knockdown experiments in Figure 1-S2 appear to confirm known roles of CAF-1 and HIRA, but important controls have and opportunities to both validate the system and learn something new have been missed: following the flowchart of Figure 1-S2B, it is evident that there's ample opportunity to test the authors hypothesis that RhIP assay indeed recapitulates the assembly pathways known in cells. For example, CAF-1 and HIRA knockdown should each be tested for epitope tagged H3.1 and H3.3, to exclude the possibility that what we are looking at here is chromatin assembly and not mere binding to nuclear chaperones. Furthermore, replication can only proceed in the presence of all four deoxynucleotides whereas chromatin remodeling requires only ATP. Thus by adding, or not, nucleotides, it can be explicitly tested if active replication is needed. Consider following setup and expected results:

no siRNA + dNTPs + ATP → nuclei are positive for H3.1 and H3.3 (shown in Figure 1-S2)

no siRNA – dNTPs + ATP → no replication! So nuclei should be negative for H3.1 unless the H3.1 signal is actually coming from H3.1 binding to CAF1. But H3.3 incorporation should be unaffected if truly replication-independent!

no siRNA – dNTPs – ATP → in the absence of nucleotides and ATP there is no energy to remove existing nucleosomes. Since we have to assume that DNA in isolated nuclei is normally chromatinized, chromatin assembly with epitope-tagged histones H3 would require prior nucleome eviction. This cannot happened without energy. Thus, in this control nuclei should be negative for H3.3. If they are still positive, then H3.3 is most likely bound by nuclear chaperones but not incorporated.

siCAF + dNTPs + ATP → nuclei are negative for H3.1 (shown in Figure 1-S2) but should be positive for H3.3 (not assayed). If gap-filling assembly via HIRA is active in RhIP assay, than H3.3 signal should even increase upon CAF1 depletion!

siHIRA + dNTPs + ATP → nuclei are negative for H3.3 (shown in Figure 1-S2) but should be remain for H3.1 when in replication (not assayed).

Figure 1 figure supplement 1B: It would make the argument stronger if the authors could show that this difference is not observed for H3.1-containing dimers.

Figure 1 figure supplement 1B: It would make the argument stronger if the authors could show that this difference is not observed for H3.1-containing dimers.

Figure 2

Since there is no negative control present for H2A, H2A.X, H2A.Z incorporation (all nuclei are positive irrespective of cell cycle), there remains an more pertinent concern that some or most of the fluorescent signal observed after RhIP could arise from non-incorporated histones that accumulate with chaperones in the nucleus. Again, a biochemical fractionation could cleanly show this, potentially also a pre-extraction protocol before immunostaining. A negative control (H2A or H2A.Z chaperone knockdown that abrogates fluorescent signal) should be established, or a -ATP control as used for the genomics experiment in Figure 5.

In Figure 2F, as with Figure 1H, it is evident that replicated chromatin is overrepresented in both H2A and H2A.Z pulldowns, suggesting that both are deposited also in a replication-dependent manner (also supported by Figure 2D image quantification). Comparison to the very similar looking fold-enrichment from pure synchronized S phase population Figure 2-S2 actually may suggest that most incorporation into proper nucleosomes is replication-dependent in this assay.

Figure 5

Is ANP32E acting catalytically (i.e. small amounts would be sufficient for keeping up eviction activity)? What's maybe more surprising is that ATP depletion is not really effective. Histone eviction requires ATP. Thus this mean there are many pre-existing nucleosome-free regions? Or could this suggest that much of the signal observed does not correspond to properly assembled nucleosomes?

Given the hypothesis put forward from Figure 5 (ATP-dependent remodeling and ANP32E together facilitate H2A.Z incorporation), then it should be tested if combined ANP32E + ATP depletion has an synergistic, additive or no additional effect beyond the single treatments.

Figure 6

The image-based analysis does support the conclusion that the M6 mutant behaves H2A-like, but it is unclear what the mechanistic insight gained from this experiment is, again acknowledging that this mutant has been constructed according to known reports.

Figure 1H: beyond demonstrating that the epitope-tagged histone associates with nucleosomal-sized DNA, a protein gel or blot would be revealing to demonstrate a stoichiometric relationship of H3, H4, H2A, H2B. A very interesting question could be addressed by assessing the ratio of epitope-tagged H3.1/3 to untagged H3 in these nucleosomal fragments: do replication-dependent and independent deposition pathways assemble homotypic H3/H4 tetramers with only exogenous (tagged) H3.1, H3.3? Or is the tagged histone H3 mixed equally with endogenous (untagged) histone H3, thus suggesting that one new H3/H4 dimer is paired with an existing H3/H4 dimer during the assembly process.

I would highly recommend performing time-course experiments to really understand the kinetics, dynamic range and saturation of the system.

https://doi.org/10.7554/eLife.66290.sa1

Author response

[Editors’ note: the authors resubmitted a revised version of the paper for consideration. What follows is the authors’ response to the first round of review.]

Although the general enthusiasm is high, significant doubts remain with regard to how faithfully RhIP recapitulates endogenous remodeling activities. In addition, it is unclear how feasible it is to use RhIP to dissect mechanism. For example, can a specific remodeling pathway be blocked by factor depletion? Since substantial amounts of additional work are required to substantiate the conclusions claimed in the paper, we are unable to accept the manuscript in the current form. We are however happy to reconsider if the following concerns are addressed.

1) Given the unique genomic localization of H2A.Z, the authors wisely used H2A.Z as a benchmark to assess the robustness of the RhIP approach. Indeed, site-specific deposition was observed when permeablized cells were incubated with recombinant H2A.Z and (cytosolic) extracts. While this is an interesting result, it remains unclear how well the H2A.Z sites observed by RhIP-ChIP-seq correlate with endogenous H2A.Z sites.

Thank you for this comment. We agree that it is important to clarify how well the H2A.Z sites observed by RhIP-ChIP-seq correlate with the endogenous H2A.Z sites detected by ChIP-seq. Accordingly, we have reanalyzed our data, and made new figures in our revised manuscript. To clearly answer this question, we separated your comments and answered each one, as below.

The authors should report the percentage of H2A.Z sites correctly identified by RhIP. Are there H2A.Z sites in the RhIP-ChIP-seq data not seen in vivo.

According to this comment, we visualized the H2A.Z sites of the RhIPChIP-seq read density across the endogenous H2A.Z peaks determined by ChIP-seq, using heatmaps (new Figure 4D and Figure 4—figure supplement 1A). We found that the H2A.Z incorporated by the RhIP assay exists in almost all endogenous H2A.Z peaks, while the H2A is excluded from the H2A.Z peaks. The reversed heatmap, which depicts the H2A.Z of the ChIP-seq read density across the H2A.Z peaks determined by the RhIP-ChIP-seq, showed that only 3% of the H2A.Z peaks are specific for the RhIP assay (new Figure 4E and Figure 4—figure supplement 1B). We described these analyses on page 15, lines 297-303.

Scatter plots should be used to compare the relative occupancy of H2A.Z between the RhIP-ChIP-seq data and known H2A.Z sites.

We drew scatter plots showing the comparison between the RhIP-ChIP-seq and ChIP-seq data at known H2A.Z sites. We found a moderately linear relationship between H2A.Z (RhIP-ChIP-seq) and H2A.Z (ChIP-seq), but little to no correlation between H2A (RhIP-ChIP-seq) and (ChIP-seq) (new Figure 4C). We described this on page 15, lines 293-297.

2) Genomic data analyses generally lack rigor. This is epitomized in Figure 4-supplement 2. The fraction-against-rank plots give little information on the concordance of the two replicates. Do the H2A.Z sites identified in the replicates correlate spatially (e.g. along the genome), quantitatively (e.g. peak height), and qualitatively (e.g. width of peaks)? For example, relative H2A.Z occupancy at known H2A.Z sites of the two H2A.Z RhIP-ChIP-seq replicates should be plotted against each other. The H2A RhIP-ChIP-seq datasets could serve as negative control.

Thank you for this suggestion. In the revised manuscript, we confirmed the reproducibility of the RhIP-ChIP-seq of H2A.Z according to this suggestion. We plotted the H2A.Z RhIP-ChIP-seq replicates at known H2A.Z sites, which showed a quite high correlation (0.95 Pearson correlation coefficient), while the H2A and H2A.Z RhIP-ChIP-seq data showed a weaker correlation, as compared to one of the replicates (new Figure 3—figure supplement 2A). Moreover, the aggregation plots of the two H2A.Z RhIP-ChIP-seq replicates at known H2A.Z sites showed similar accumulation patterns (new Figure 4D and new Figure 4—figure supplement 1A). In contrast, the aggregation plot of the H2A RhIP-ChIPseq showed the exclusion of H2A at the known H2A.Z sites (new Figure 4D). We described these findings on page 18, lines 377-382.

3) It is important to assess how individual factors contribute to a particular remodeling pathway. Given that the steady-state H2A.Z occupancy is a net result of deposition and eviction and that multiple remodeling factors are involved in these processes, whether the observed H2A.Z peaks in RhIP-ChIP-seq represent the consequence of one or more of these factors should be evaluated. For example, are SRCAP and p400/TIP60 responsible for different H2A.Z sites across the genome? Is ATP required for H2A.Z deposition?

According to these suggestions, we first examined the permeabilized cells for the presence of SRCAP, TIP60 and ANP32E. The fractionation and following western blot analysis confirmed that SRCAP and TIP60 are still in the permeabilized cells, suggesting that the chromatin remodeling activity may be involved in the H2A.Z deposition in the RhIP assay. Since chromatin remodeling requires ATP, we then prepared the ATP-reduced cellular extract and tested it in the RhIP assay with or without ATP supplementation (new Figures 5E and F). As a result, ATP promoted the H2A.Z depositions in active TSS and enhancer regions, suggesting that the H2A.Z deposition in the RhIP assay requires a chromatin remodeling activity that hydrolyzes ATP. We described these new results on pages 1617, lines 329-341.

Is ANP32E responsible for removing H2A.Z at enhancers and insulators? Does transcription contribute to H2A.Z eviction?

Our fractionation and western blot analysis showed that ANP32E is removed from permeabilized cells during the permeabilization process (new Figure 5A). As ANP32E is present in the cellular extract, which is used in the RhIP assay, we prepared the ANP32E-knockdown cellular extract and performed a RhIP-ChIP-seq analysis of H2A.Z (new Figures 5B-D). As a result, the efficiency of H2A.Z incorporations into active TSS and enhancer regions, where endogenous H2A.Z is predominantly localized, was decreased upon the ANP32E-knockdown (new Figure 5D and Figure 5—figure supplement 1A). This indicates that the ANP32Edependent evictions of the pre-incorporated H2A.Z in active TSS and enhancer regions promotes the incorporation of exogenously added H2A.Z in the RhIP assay. We described these data on page 16, lines 313-328.

We note that a previous paper reported that ANP32E-knockout mouse embryonic fibroblast (MEF) cells showed an increased number of H2A.Z peaks in insulator regions. In the RhIP assay using the ANP32Eknockdown cellular extract, the changes in the efficiency of the H2A.Z deposition in the insulator regions were less than those in the active TSS and enhancers (new Figure 5D). This discrepancy may arise from the fact that the H2A.Z distribution pattern of the knockout MEFs represents chromatin reorganization over a long time during embryogenesis, thus facilitating the adaptation to the ANP32E-knockout condition. This may include compensation for the ANP32E-mediated H2A.Z evictions. In contrast, our RhIP-seq analysis using the ANP32E-knockdown cellular extract instead represents the chromatin dynamics after the acute depletion of ANP32E. Consistent with our data, another previous paper showed that the introduction of recombinant ANP32E to zebrafish embryos resulted in the global loss of H2A.Z from the nucleus (P. J. Murphy, Wu, James, Wike, and Cairns, 2018).

Murphy, P. J., Wu, S. F., James, C. R., Wike, C. L., and Cairns, B. R.

(2018). Placeholder Nucleosomes Underlie Germline-to-Embryo DNA

Methylation Reprogramming. Cell, 172(5), 993–1006.e13. http://doi.org/10.1016/j.cell.2018.01.022

Proper spike-in controls should be incorporated into the RhIP-ChIP-seq samples before and after the addition of factor-depleted extracts to allow quantitative assessment.

Thank you for this critical comment. Accordingly, we performed the ATPand ANP32E-depleted RhIP-ChIP-seq analysis using the spike-in controls (new Figures 5C-F and Figure 5—figure supplement 1) (Chen et al., 2015; Egan et al., 2016; Orlando et al., 2014).

Chen, K., Hu, Z., Xia, Z., Zhao, D., Li, W., and Tyler, J. K. (2015). TheOverlooked Fact: Fundamental Need for Spike-In Control for Virtually All Genome-Wide Analyses. Molecular and Cellular Biology, 36(5), 662–667. http://doi.org/10.1128/MCB.00970-14

Egan, B., Yuan, C.-C., Craske, M. L., Labhart, P., Guler, G. D., Arnott, D., et al. (2016). An Alternative Approach to ChIP-seq Normalization Enables Detection of Genome-Wide Changes in Histone H3 Lysine 27 Trimethylation upon EZH2 Inhibition. PLoS ONE, 11(11), e0166438. http://doi.org/10.1371/journal.pone.0166438

Orlando, D. A., Chen, M. W., Brown, V. E., Solanki, S., Choi, Y. J., Olson, E. R., et al. (2014). Quantitative ChIP-seq normalization reveals global modulation of the epigenome. Cell Reports, 9(3), 1163–1170. http://doi.org/10.1016/j.celrep.2014.10.018

Since cytosolic extracts were used, it is not surprising that some factors may be under represented. If such biases do exist, the authors should consider using nuclear extracts and/or provide evidence on how much remodeling factors partitioned into the fractions relative to input (e.g. SRCAP western blots).

As mentioned above, our fractionation analysis indicated that SRCAP and TIP60, components of major H2A.Z deposition complexes, were retained in the permeabilized cells. Even though ANP32E was extracted during the permeabilization process, ANP32E was added back to the RhIP reaction as the ANP32E present in the cellular extract. These data indicate that these factors are not underrepresented in the RhIP assay.

Together with the results mentioned above, we conclude that the H2A.Z deposition in the RhIP assay is a net result of the ANP32E-mediated eviction and deposition of H2A.Z and the replacement of pre-incorporated H2A with H2A.Z by chromatin remodeling, and that the cellular H2A.Z deposition pathways are recapitulated in the RhIP assay.

4) The title suggests that RhIP is generally applicable to all histones. But there was no mention of the H3 results in the Abstract or the Discussion.

Thank you for this advice. We added sentences regarding the H3 results in the Abstract and Discussion on page 2, lines 6-8 and page 19, lines 390-396.

In addition, while immunostaining shows H3.1 and H3.3 are incorporated into the nucleus of permeabilized cells at the expected cell cycle stages, microscopy lacks the resolution necessary to validate if H3.1 and H3.3 are targeted to the expected genomic locations. For example, is H3.3 more enriched at active genes?

We agree with this comment. We performed the RhIP-ChIP-seq analysis of H3.3 and found its enrichment at active genes (new Figure 1—figure supplement 1A). In addition, the RhIP-ChIP-qPCR analysis showed higher enrichment of H3.3 in the active gene (GAPDH) than the inactive gene (LINC02199) (new Figure 1—figure supplement 1B). We described these data on pages 8 and 9, lines 143-162.

CAF1 and HIRA knockdown should be used to verify if H3.1 and H3.3 are indeed deposited by these pathways.

We confirmed the CAF-1 and HIRA requirements for the H3.1 and H3.1 depositions in the RhIP assay by immunostaining (new Figure 1—figure supplement 2), since previous SNAP-tag based analyses clearly showed their requirement by imaging (Ray-Gallet et al., 2011). First, we checked for the existence of CAF-1 and HIRA in permeabilized cells. Our fractionation followed by western blotting analysis showed that these factors were retained in permeabilized cells (new Figure 1—figure supplement 2A). We then prepared the CAF-1- or HIRA-knockdown permeabilized cells and performed the RhIP-immunostaining assay of H3.1 or H3.3 (new Figure 1—figure supplements 2B-D). In this assay, we co-cultured the control and knockdown cells on the same coverslips for a precise evaluation. The knockdown cells were judged by immunostainings for CAF-1 (p60) and HIRA. As a result, we found that the intensities of both the incorporated H3.1 and H3.3 were significantly decreased in the CAF-1- and HIRA-knockdown cells, respectively. Therefore, H3.1 and H3.3 are deposited by the CAF-1- and HIRA-dependent pathways in the RhIP assay, respectively. We described these results on pages 9 and 10, lines 163-180.

Ray-Gallet, D., Woolfe, A., Vassias, I., Pellentz, C., Lacoste, N., Puri, A., et al. (2011). Dynamics of histone H3 deposition in vivo reveal a nucleosome gap-filling mechanism for H3.3 to maintain chromatin integrity. Molecular Cell, 44(6), 928–941.

Finally, to show that H3.1 and H3.3 are indeed inserted into nucleosomes, the MNase experiment similar to the one in Figure 3C should be performed.

According to this comment, we performed an MNase experiment similar to that in the previous Figure 3C (currently Figure 2F), using H3.1 and H3.3 (new Figures 1F-H). As a result, we confirmed their incorporations into the chromatin of permeabilized cells. Moreover, we found that the amount of nascent DNA in the H3.1 RhIP-ChIP sample was more than that in the H3.3 RhIP-ChIP sample. This is consistent with the results of the RhIP immunostaining assay shown in the new Figures 1C-E. We described these data on pages 8 and 9, lines 143-154.

5) Major improvement of the description of methods is required to allow rigorous assessment of the approach. The current methods section lacks critical details necessary to carry out the experiments.

We rewrote the methods section on page 23, lines 480-482, pages 24-30, lines 495-637, page 31, lines 658-672 and page 32, lines 701-724 in the revised manuscript. We hope the new methods section is sufficient for readers to perform the RhIP assay. If more information is needed, we will be happy to provide any details. We then express this in Data availability as If more information, or tips are needed, we will/are willing to provide upon reasonable requests.

[Editors’ note: what follows is the authors’ response to the second round of review.]

Essential Revisions:

Here we share detailed comments to tackle the limitations of the study:

1. While replication-dependent mechanisms are well captured by RhIP, it is less clear if transcription and chromatin remodeling is functional in this system and thus if transcription-dependent nucleosome exchange processes are faithfully recapitulated. It is important to improve the comparison of RhIP with 'in vivo' (i.e. existing ChIP-seq datasets) localisation and explicitly develop hypotheses why in some cases the data matches the 'in vivo' situation and in others not.

We thank the reviewers for this important comment. Based on the suggestions, we performed additional analyses as described below, which allowed us to hypothesize that the difference between the RhIP-ChIP-seq and ChIP-seq may reflect the recycling frequency of the evicted histone complexes.

a. Lines 279-301 and Figure 4. There is moderate correlation between genome-wide H2A.Z RhIP-ChIP-seq and H2A.Z ChIP-seq (Figure 4C). However, it seems H2A.Z RhIP-ChIP might be selectively detecting TSS specific signals rather than gene body specific associations (Figure 4A and 4E). It would be helpful to comment on the genetic elements (e.g. gene body) at which H2A.Z RhIP-ChIP and H2A.Z ChIP show low or no correlation and make a comparison. This way strengths and weaknesses of the assay can be better evaluated and compared to already existing protocols.

According to this suggestion, we have re-analyzed our data and created a new scatter plot between the RhIP-ChIP-seq and ChIP-seq data of H2A.Z at known H2A.Z sites in gene bodies, exclusively (new Figure 4C right). As this reviewer pointed out, we found that there is a fairly linear relationship between them in the gene bodies, indicating that the RhIP-ChIP-seq preferentially detects TSS-specific incorporation. This may have originated from the fact that H2A.Z can be exchanged more dynamically at TSS, as compared to gene bodies. Alternatively, as the histones that are evicted during transcription can be recycled (1, 2), the lower correlation between RhIP-ChIP and ChIP of H2A.Z in gene bodies may reflect the lower frequency of de novo H2A.Z deposition and the higher frequency of H2A.Z recycling during transcription. We described these analyses on page 15, lines 301-304 and discussed them on page 22, lines 453-457.

1. Torné J, Ray-Gallet D, Boyarchuk E, Garnier M, Le Baccon P, Coulon A, Orsi GA, Almouzni G. (2020) Two HIRA-dependent pathways mediate H3.3 de novo deposition and recycling during transcription. Nature Structural Molecular Biology 27(11):1057-1068. doi: 10.1038/s41594-020-0492-7.

2. Jeronimo, C., Poitras, C., and Robert, F. (2019). Histone Recycling by FACT and Spt6 during Transcription Prevents the Scrambling of Histone Modifications.

Cell Reports, 28(5), 1206–1218.e8. http://doi.org/10.1016/j.celrep.2019.06.097

The image resolution of Figure 4A should be improved.

We improved the image resolution of Figure 4A, from 144 dpi (previous) to 300 dpi (current). Please note that all of the bigwig files shown in this study were created with 1 bp intervals on the genome and not smoothed. This may be the reason why the image appears to have low resolution as compared with the smoothed bigwig files.

b. Suppl. Figure 1.1 H3.3 distribution across the active and inactive genes were plotted in Suppl Figure 1. However, how active and inactive status of genes was defined, and which data was utilized for this analysis were not indicated under the Methods. These parameters and data source should be included.

We thank the reviewer for this comment. We defined genes with 5 or more read counts in the RNA-seq data (GSE140768) as expressed genes and the rest as non-expressed genes. We now describe these parameters in the Methods on page 33, lines 698-700.

Transcript levels for GAPDH and LNC02199 should also be indicated or the reader should be referred to a table containing transcription data (i.e. RNA-Seq).

We have created the suggested table, and now show it in the new Figure 1—figure supplement 1C.

I also think that it would be helpful if the authors can comment on the efficiency of H3.3-HA ChIP compared to pull down of endogenous H3.3 by comparing RhIP-ChIP-seq and previously reported H3.3 ChIP-seq analyses.

According to this suggestion, we assessed the signal-to-noise ratios as part of the efficiency of the methods. We have calculated and compared the S/N ratios of RhIP-ChIP-seq and the previously reported ChIP-seq of H3.3, and created new figures (new Figure 1—figure supplements 1D and E). We found that the S/N ratios are almost the same. We now describe these results on page 11, lines 199-201.

c. The genomics analysis presented in Figure 3 is somewhat rudimentary, the signals from H2A and H2A.Z appear overall very correlated (Figure 3A, B), albeit the correlation genome-wide calculated in Figure 3-S2 is 'medium'. The scatter plot could be shown better to see potentially different 'population of loci, some that correlate well and 'outliers'.

We thank the reviewer for this important comment. First, we would like to explain that the analysis presented in Figure 3A shows the entire genome-wide pattern. On the other hand, the scatter plots in Figure 3—figure supplement 2A were calculated at H2A.Z peaks detected in ChIP-seq. As this reviewer pointed out, H2A.Z and H2A are differently incorporated at H2A.Z peaks locally, as the correlation is moderate (Pearson correlation is 0.48). This is consistent with the results shown as the heatmap in Figure 4D, in which the H2A of RhIP-ChIP-seq showed no accumulation at the H2A.Z peaks of ChIP-seq. On the other hand, they are globally incorporated with certain similarity (Pearson correlation is 0.62, new Figure 3A right). We describe this on page 13, line 257 and page 16, lines 307-309.

Judging from Figure 3B, it appears that there is a good correlation across broad chromatin states with the notable excavation of active TSS. This is in line with the known H2A.Z mechanism, but could be elaborated in plots a bit clearer. How do the profiles look across highly expressed and non-expressed genes?

Thank you for this suggestion. We have recalculated our data and revised the aggregation plot to show the expressed and non-expressed genes separately (new Figure 4B). It is now clear that H2A.Z predominantly accumulated at the TSS of active (expressed) genes. We described this on page 15, line 296.

The cell-cycle-synchronized plots are only summarized in the heatmap in Figure 3C,D. Example loci could be shown as in Figure 3A.

In the revised manuscript, we show the example loci in the new Figure 3—figure supplement 1B.

2. It would be helpful to improve the interpretation of the data to include all existing caveats to the assay setup.

Thank you for this suggestion. We addressed this comment, as described below.

a. The new ATP addition experiments (figure 5E-F) are interpreted as if the only effect of ATP is on chromatin remodellers. It is well possible that by adding ATP in this conditions, additional factors (i.e. transcription, splicing/translation, molecular chaperones, etc) are being affected and responsible for the effect.

We agree with this comment. We discussed the possible effects of ATP addition, on page 18 lines 353-355.

b. The authors alternate setups where they either co-incubate different histone dimers, or they use a single histone dimer isoform. It is really unclear to the readers what is the reason for this and how does this affect the results/interpretation. In cases where a mixture of histone dimers were present (i.e. Figures 1A-D, 2, 6), competition between the H2A/H2A.Z-X or H3.1/H3.3 dimers may take place, while in the others this competition is not present. This makes the comparison of the results quite challenging, and their conclusions unclear. This should be discussed at the very least, as it has implications on understanding the properties that control histone incorporation during RhIP.

We agree with this reviewer’s concern. We understand that the proper combination of histone chaperones and histone variants ensures the correct localizations of histone variants. However, if one histone variant is increased, then the excess histone variant may bind to unsuitable histone chaperones, leading to ectopic localization. Indeed, the overexpressed histone H3 variant, CENP-A, in cells aberrantly binds to the H3.3 chaperone, DAXX, leading to ectopic localizations (1). In this study, we performed the RhIP assay with two different histone variants simultaneously, which do not share the same chaperone. Therefore, if the amounts of added histone complexes are appropriate, then they would only interact with their defined chaperones and be deposited in a specific manner. As a result, we observed different staining patterns in the same nucleus when H3.1 and H3.3 or H2A/H2A.X and H2A.Z were co-incubated (Figures 1A-D, 2, 6), which proved that they were incorporated into the chromatin by specific mechanisms, rather than non-specifically incorporated. These results also indicate that there is little to no competition under these conditions. Therefore, the results are unlikely to change if one or two histone variant(s) was/were used for the RhIP assay. However, as the reviewer mentioned, the competition assay using this system is worth trying, to analyze the overexpression situations of a specific histone variant. We added this discussion on page 20, lines 402-412.

1. Lacoste N, Woolfe A, Tachiwana H, Garea AV, Barth T, Cantaloube S, Kurumizaka H, Imhof A, Almouzni G (2014) Mislocalization of the centromeric histone variant CenH3/CENP-A in human cells depends on the chaperone DAXX Molecular Cell 53(4):631-44. doi: 10.1016/j.molcel.2014.01.018.

c. The RhIP-ChIP-seq protocol uses formaldehyde crosslinking instead of native mononucleosomal IP as used in Figure 1H, leaving room for interpretation if the pull-down histone is indeed incorporated into chromatin. Immunofluorescence also does not necessarily prove incorporation of histones into chromatin, since i.e. the fractionation in Figure 1-S2 shows that HIRA and CAF1 remain tightly bound to the isolated nuclei. Thus, accumulation of the epitope-tagged H3 in the nucleus of cells could merely mean that is bound to the histone chaperones present in the nucleus (competing away endogenous histones). It is not possible to discern from the data what the incorporated amount versus the chaperone-bound fraction of recombinant histone is.

Thank you for this comment. This point is critical and must be clarified experimentally. We therefore performed an additional RhIP-immunostaining assay. We treated the cells with PBST containing 300 mM NaCl after the RhIP reaction, to wash out the nuclear proteins that are associated with chromatin, and then fixed the cells. In fact, CAF-1 and HIRA were no longer detected by the immunostaining; however, the exogenously added H3.1 and H3.3 were still detected in the permeabilized cells. This result indicates that the exogenously added H3.1 and H3.3 were mostly incorporated into the chromatin of the permeabilized cells in the RhIP assay, as shown by the native ChIP experiment. We added these data in the new Figure 1—figure supplement 3 and described them on page 10, lines 175-184.

d. The functional assessment of ANP32E knockdown lacks statistical analysis – clearly the quite robust knockdown (Figure 5B) does not majorly abrogate TSS incorporation of H2A.Z. Is the difference statistically significant?

We thank the reviewer for pointing out this issue. We think the TSS incorporation of H2A.Z. significantly changed upon the ANP32E knockdown, with a p-value below 10-50. We would like to be cautious about mentioning this, because we included more than forty thousand loci for each in our analysis. With such a large number of samples, the p-value tends to be close to zero; therefore, it does not necessarily show practical significance (1).

1. Lin, M., Lucas, H. C., Jr., and Shmueli, G. (2013). Too big to fail: Large samples and the p-value problem. Information Systems Research, 24(4), 906–917. https://doi.org/10.1287/isre.2013.0480

What is the explanation for the relatively small effect size?

We agree with the reviewer that the reduction of TSS incorporation of H2A.Z is relatively small in the ANP32E knockdown cellular extract. We suggest the presence of a redundant mechanism, which is consistent with the previous report that ANP32E knockout mice did not exhibit any apparent effects on their viability (1), even though proper H2A.Z incorporation is essential for cell survival and proliferation. We also found that ATP promotes the H2A.Z depositions in the RhIP assay. Since ATP is essential for many biological processes, such as ATP-dependent chromatin remodeling and transcription, the H2A.Z deposition in the RhIP assay may require ATP-dependent processes. Taken together, the H2A.Z deposition in the RhIP assay is the net result of the ANP32E-mediated eviction and deposition of H2A.Z, and the replacement of pre-incorporated H2A with H2A.Z by ATP-dependent processes. This is our explanation for the limited effects of the ANP32E knockdown on the H2A.Z incorporation. We discussed this on pages 21-22, lines 443-457.

1. Reilly PT, Afzal S, Wakeham A, Haight J, You-Ten A, Zaugg K, Dembowy J, Young A, Mak TW. (2010) Generation and characterization of the Anp32e-deficient mouse PLoS One, 5(10):e13597. doi: 10.1371/journal.pone.0013597.

e. Lines 275-277. Please revisit the conclusion and make the statement clearer on H2A.Z incorporation and elimination.

According to this comment, we rewrote the conclusion, on page 15, lines 284-286.

We suggest adding these references:

The authors make a thorough effort to place their method in the context of existing pulse-labeling methods, but Tet-ON based pulsing methods, which do give similar results have not been discussed: Mito, Y., Henikoff, J. G. and Henikoff, S. Genome-scale profiling of histone H3.3 replacement patterns. Nat. Genet. 37, 1090-1097 (2005); Yildirim, O. et al. A system for genome-wide histone variant dynamics in ES cells reveals dynamic MacroH2A2 replacement at promoters. PLoS Genet. 10, e1004515 (2014); Ha, M., Kraushaar, D. C. and Zhao, K. Genome-wide analysis of H3.3 dissociation reveals high nucleosome turnover at distal regulatory regions of embryonic stem cells. Epigenetics Chromatin 7, 38 (2014).

Thank you for this suggestion. We have already cited the first paper on page 9 line 155, to refer to H3.3 incorporation in the genome. We newly added the second and third references to introduce the tetracycline-based methods, on page 6, lines 99-101.

https://doi.org/10.7554/eLife.66290.sa2

Article and author information

Author details

  1. Hiroaki Tachiwana

    Division of Cancer Biology, The Cancer Institute of Japanese Foundation for Cancer Research, Tokyo, Japan
    Contribution
    Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    hiroaki.tachiwana@jfcr.or.jp
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9227-7653
  2. Mariko Dacher

    Laboratory of Chromatin Structure and Function, Institute for Quantitative Biosciences, The University of Tokyo, Tokyo, Japan
    Contribution
    Formal analysis, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  3. Kazumitsu Maehara

    Division of Transcriptomics, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan
    Contribution
    Formal analysis, Funding acquisition, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  4. Akihito Harada

    Division of Transcriptomics, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan
    Contribution
    Formal analysis, Funding acquisition, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  5. Yosuke Seto

    Division of Experimental Chemotherapy, Cancer Chemotherapy Center, Japanese Foundation for Cancer Research, Tokyo, Japan
    Contribution
    Formal analysis, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  6. Ryohei Katayama

    Division of Experimental Chemotherapy, Cancer Chemotherapy Center, Japanese Foundation for Cancer Research, Tokyo, Japan
    Contribution
    Formal analysis, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  7. Yasuyuki Ohkawa

    Division of Transcriptomics, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan
    Contribution
    Funding acquisition, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6440-9954
  8. Hiroshi Kimura

    Cell Biology Center, Institute of Innovative Research, Tokyo Institute of Technology, Yokohama, Japan
    Contribution
    Conceptualization, Funding acquisition, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0854-083X
  9. Hitoshi Kurumizaka

    Laboratory of Chromatin Structure and Function, Institute for Quantitative Biosciences, The University of Tokyo, Tokyo, Japan
    Contribution
    Conceptualization, Funding acquisition, Validation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  10. Noriko Saitoh

    Division of Cancer Biology, The Cancer Institute of Japanese Foundation for Cancer Research, Tokyo, Japan
    Contribution
    Conceptualization, Funding acquisition, Validation, Writing - review and editing
    For correspondence
    noriko.saito@jfcr.or.jp
    Competing interests
    No competing interests declared

Funding

Japan Society for the Promotion of Science (JP17H05013)

  • Hiroaki Tachiwana

Japan Society for the Promotion of Science (JP16K14785)

  • Hiroaki Tachiwana

Japan Society for the Promotion of Science (JP20K06496)

  • Hiroaki Tachiwana

Japan Society for the Promotion of Science (JP20H05397)

  • Hiroaki Tachiwana

Japan Society for the Promotion of Science (16H06279 (PAGS))

  • Hiroaki Tachiwana

Japan Society for the Promotion of Science (JP18H05531)

  • Noriko Saitoh

Japan Society for the Promotion of Science (JP18K19310)

  • Noriko Saitoh

Japan Society for the Promotion of Science (JP19H04970)

  • Kazumitsu Maehara

Japan Society for the Promotion of Science (JP19H03158)

  • Kazumitsu Maehara

Japan Society for the Promotion of Science (JP20H05393)

  • Kazumitsu Maehara

Japan Science and Technology Agency (JPMJPR2026)

  • Kazumitsu Maehara

Japan Society for the Promotion of Science (JP18K19432)

  • Akihito Harada

Japan Society for the Promotion of Science (JP19H05425)

  • Akihito Harada

Japan Society for the Promotion of Science (JP19H03211)

  • Akihito Harada

Japan Society for the Promotion of Science (JP20H05368)

  • Akihito Harada

Japan Science and Technology Agency (JPMJPR19K7)

  • Akihito Harada

Japan Society for the Promotion of Science (JP17K19356)

  • Yasuyuki Ohkawa

Japan Society for the Promotion of Science (JP17H03608)

  • Yasuyuki Ohkawa

Japan Society for the Promotion of Science (JP18H05527)

  • Yasuyuki Ohkawa
  • Hiroshi Kimura

Japan Science and Technology Agency (JMJCR16G1)

  • Yasuyuki Ohkawa
  • Hiroshi Kimura

Japan Society for the Promotion of Science (JP18H04802)

  • Yasuyuki Ohkawa

Japan Society for the Promotion of Science (JP19H05244)

  • Yasuyuki Ohkawa

Japan Society for the Promotion of Science (JP20H00456)

  • Yasuyuki Ohkawa

Japan Society for the Promotion of Science (JP20H04846)

  • Yasuyuki Ohkawa

Japan Agency for Medical Research and Development (JP20ek0109489h0001)

  • Yasuyuki Ohkawa

Japan Society for the Promotion of Science (JP18H05534)

  • Hitoshi Kurumizaka

Japan Science and Technology Agency (JPMJER1901)

  • Hitoshi Kurumizaka

Japan Agency for Medical Research and Development (JP20am0101076)

  • Hitoshi Kurumizaka

Japan Society for the Promotion of Science (JP20H00449)

  • Hitoshi Kurumizaka

Japan Society for the Promotion of Science (JP17H01417)

  • Hiroshi Kimura

Japan Society for the Promotion of Science (JP20H03520)

  • Noriko Saitoh

Nakajima Foundation

  • Hiroaki Tachiwana

Takeda Science Foundation

  • Noriko Saitoh

Vehicle Racing Commemorative Foundation

  • Noriko Saitoh

Princess Takamatsu Cancer Research Fund

  • Noriko Saitoh

Japan Science and Technology Agency (CREST JPMJCR16G1)

  • Yasuyuki Ohkawa
  • Hiroshi Kimura

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported by JSPS KAKENHI Grant Numbers JP17H05013, JP16K14785, JP20K06496, JP20H05397, 16H06279 (PAGS) (to HT), JP18H05531, JP18K19310, JP20H03520 (to NS), JP19H04970, JP19H03158, JP20H05393 (to KM), JP18K19432, JP19H05425, JP19H03211, JP20H05368 (to AH), and JP17K19356, JP17H03608, JP18H05527, JP18H04802, JP19H05244, JP20H00456, JP20H04846 (to YO), JP17H01417 and JP18H05527 (to HKimura), JP18H05534, JP20H00449 (to HKurumizaka), JST PRESTO JPMJPR2026 (to KM), JPMJPR19K7 (to AH), AMED JP20ek0109489h0001 (to YO), JST CREST grants JPMJCR16G1 (to YO and HKimura), JST ERATO JPMJER1901, AMED JP20am0101076 (to HKurumizaka). We thank Drs. Yuma Ito at Tokyo Institute of Technology and Kazuhiko Uchida at the Cancer Institute of JFCR for technical advice. We also thank Dr. Crawford at Duke University for the DNaseI-seq data (GEO:GSM816643), Broad Institute for the H2A.Z ChIP-seq data (GEO:GSM1003483) and the ENCODE Consortium. HT is supported by The Nakajima Foundation. NS is supported by the Takeda Science Foundation, the Vehicle Racing Commemorative Foundation and a Research Grant from the Princess Takamatsu Cancer Research Fund.

Senior Editor

  1. Jessica K Tyler, Weill Cornell Medicine, United States

Reviewing Editor

  1. Francesca Mattiroli, Hubrecht Institute, Netherlands

Publication history

  1. Received: January 6, 2021
  2. Accepted: April 5, 2021
  3. Version of Record published: May 10, 2021 (version 1)

Copyright

© 2021, Tachiwana et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

Metrics

  • 855
    Page views
  • 105
    Downloads
  • 0
    Citations

Article citation count generated by polling the highest count across the following sources: Crossref, PubMed Central, Scopus.

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Download citations (links to download the citations from this article in formats compatible with various reference manager tools)

Open citations (links to open the citations from this article in various online reference manager services)

Further reading

    1. Chromosomes and Gene Expression
    2. Plant Biology
    Alexander R Leydon et al.
    Research Article Updated

    The plant corepressor TOPLESS (TPL) is recruited to a large number of loci that are selectively induced in response to developmental or environmental cues, yet the mechanisms by which it inhibits expression in the absence of these stimuli are poorly understood. Previously, we had used the N-terminus of Arabidopsis thaliana TPL to enable repression of a synthetic auxin response circuit in Saccharomyces cerevisiae (yeast). Here, we leveraged the yeast system to interrogate the relationship between TPL structure and function, specifically scanning for repression domains. We identified a potent repression domain in Helix 8 located within the CRA domain, which directly interacted with the Mediator middle module subunits Med21 and Med10. Interactions between TPL and Mediator were required to fully repress transcription in both yeast and plants. In contrast, we found that multimer formation, a conserved feature of many corepressors, had minimal influence on the repression strength of TPL.

    1. Cancer Biology
    2. Chromosomes and Gene Expression
    Shou Liu et al.
    Research Article Updated

    ARID1A is one of the most frequently mutated epigenetic regulators in a wide spectrum of cancers. Recent studies have shown that ARID1A deficiency induces global changes in the epigenetic landscape of enhancers and promoters. These broad and complex effects make it challenging to identify the driving mechanisms of ARID1A deficiency in promoting cancer progression. Here, we identified the anti-senescence effect of Arid1a deficiency in the progression of pancreatic intraepithelial neoplasia (PanIN) by profiling the transcriptome of individual PanINs in a mouse model. In a human cell line model, we found that ARID1A deficiency upregulates the expression of aldehyde dehydrogenase 1 family member A1 (ALDH1A1), which plays an essential role in attenuating the senescence induced by oncogenic KRAS through scavenging reactive oxygen species. As a subunit of the SWI/SNF chromatin remodeling complex, our ATAC sequencing data showed that ARID1A deficiency increases the accessibility of the enhancer region of ALDH1A1. This study provides the first evidence that ARID1A deficiency promotes pancreatic tumorigenesis by attenuating KRAS-induced senescence through the upregulation of ALDH1A1 expression.