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.

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.

Figure 7

Download asset Open asset

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.

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.

1. 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.

   https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE130947
2. 1. Tachiwana H

   (2021) NCBI Gene Expression Omnibus

   ID GSE163502. RhIP-ChIP-seq of H2A.Z under ANP32E or ATP depletion condition.

   https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE163502
3. 1. Tachiwana H

   (2020) DNA Data Bank of Japan

   ID DRA009580. RhIP-ChIP-seq of H3.3.

   https://ddbj.nig.ac.jp/DRASearch/submission?acc=DRA009580

1. 1. ENCODE DCC

   (2011) NCBI Gene Expression Omnibus

   ID GSM816643. Duke_DnaseSeq_HeLa-S3.

   https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSM816643
2. 1. ENCODE DCC

   (2012) NCBI Gene Expression Omnibus

   ID GSM1003483. Broad_ChipSeq_HeLa-S3_H2A.Z.

   https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSM1003483

1. 1. 1000 Genome Project Data Processing Subgroup
   2. Li H
   3. Handsaker B
   4. Wysoker A
   5. Fennell T
   6. Ruan J
   7. Homer N
   8. Marth G
   9. Abecasis G
   10. Durbin R

   (2009) The sequence alignment/Map format and SAMtools

   Bioinformatics 25:2078–2079.

   https://doi.org/10.1093/bioinformatics/btp352

   • PubMed
   • Google Scholar
2. 1. Adam SA
   2. Marr RS
   3. Gerace L

   (1990) Nuclear protein import in permeabilized mammalian cells requires soluble cytoplasmic factors

   Journal of Cell Biology 111:807–816.

   https://doi.org/10.1083/jcb.111.3.807

   • Google Scholar
3. 1. Aguilar-Gurrieri C
   2. Larabi A
   3. Vinayachandran V
   4. Patel NA
   5. Yen K
   6. Reja R
   7. Ebong IO
   8. Schoehn G
   9. Robinson CV
   10. Pugh BF
   11. Panne D

   (2016) Structural evidence for Nap1-dependent H2A-H2B deposition and nucleosome assembly

   The EMBO Journal 35:1465–1482.

   https://doi.org/10.15252/embj.201694105

   • PubMed
   • Google Scholar
4. 1. Ahmad K
   2. Henikoff S

   (2002) The histone variant H3.3 marks active chromatin by replication-independent nucleosome assembly

   Molecular Cell 9:1191–1200.

   https://doi.org/10.1016/S1097-2765(02)00542-7

   • PubMed
   • Google Scholar
5. 1. Altaf M
   2. Auger A
   3. Monnet-Saksouk J
   4. Brodeur J
   5. Piquet S
   6. Cramet M
   7. Bouchard N
   8. Lacoste N
   9. Utley RT
   10. Gaudreau L
   11. Côté J

   (2010) NuA4-dependent acetylation of nucleosomal histones H4 and H2A directly stimulates incorporation of H2A.Z by the SWR1 complex

   Journal of Biological Chemistry 285:15966–15977.

   https://doi.org/10.1074/jbc.M110.117069

   • Google Scholar
6. 1. Bachu M
   2. Tamura T
   3. Chen C
   4. Narain A
   5. Nehru V
   6. Sarai N
   7. Ghosh SB
   8. Ghosh A
   9. Kavarthapu R
   10. Dufau ML
   11. Ozato K

   (2019) A versatile mouse model of epitope-tagged histone H3.3 to study epigenome dynamics

   Journal of Biological Chemistry 294:1904–1914.

   https://doi.org/10.1074/jbc.RA118.005550

   • Google Scholar
7. 1. Barski A
   2. Cuddapah S
   3. Cui K
   4. Roh TY
   5. Schones DE
   6. Wang Z
   7. Wei G
   8. Chepelev I
   9. Zhao K

   (2007) High-resolution profiling of histone methylations in the human genome

   Cell 129:823–837.

   https://doi.org/10.1016/j.cell.2007.05.009

   • PubMed
   • Google Scholar
8. 1. Boyarchuk E
   2. Filipescu D
   3. Vassias I
   4. Cantaloube S
   5. Almouzni G

   (2014) The histone variant composition of centromeres is controlled by the pericentric heterochromatin state during the cell cycle

   Journal of Cell Science 127:3347–3359.

   https://doi.org/10.1242/jcs.148189

   • PubMed
   • Google Scholar
9. Software

   1. Broad Institute

   (2019) Picard Toolkit

   Picard Toolkit.

   http://broadinstitute.github.io/picard/
10. 1. Buschbeck M
    2. Hake SB

    (2017) Variants of core histones and their roles in cell fate decisions, development and Cancer

    Nature Reviews Molecular Cell Biology 18:299–314.

    https://doi.org/10.1038/nrm.2016.166

    • PubMed
    • Google Scholar
11. 1. Chen K
    2. Hu Z
    3. Xia Z
    4. Zhao D
    5. Li W
    6. Tyler JK

    (2015) The overlooked fact: fundamental need for Spike-In control for virtually all Genome-Wide analyses

    Molecular and Cellular Biology 36:662–667.

    https://doi.org/10.1128/MCB.00970-14

    • PubMed
    • Google Scholar
12. 1. Clarkson MJ
    2. Wells JR
    3. Gibson F
    4. Saint R
    5. Tremethick DJ

    (1999) Regions of variant histone His2AvD required for Drosophila development

    Nature 399:694–697.

    https://doi.org/10.1038/21436

    • PubMed
    • Google Scholar
13. 1. Clément C
    2. Orsi GA
    3. Gatto A
    4. Boyarchuk E
    5. Forest A
    6. Hajj B
    7. Miné-Hattab J
    8. Garnier M
    9. Gurard-Levin ZA
    10. Quivy JP
    11. Almouzni G

    (2018) High-resolution visualization of H3 variants during replication reveals their controlled recycling

    Nature Communications 9:3181.

    https://doi.org/10.1038/s41467-018-05697-1

    • PubMed
    • Google Scholar
14. 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.

    • PubMed
    • Google Scholar
15. 1. Deal RB
    2. Henikoff S

    (2010) Capturing the dynamic epigenome

    Genome Biology 11:218.

    https://doi.org/10.1186/gb-2010-11-10-218

    • PubMed
    • Google Scholar
16. 1. Deaton AM
    2. Gómez-Rodríguez M
    3. Mieczkowski J
    4. Tolstorukov MY
    5. Kundu S
    6. Sadreyev RI
    7. Jansen LET
    8. Kingston RE

    (2016) Enhancer regions show high histone H3.3 turnover that changes during differentiation

    eLife 5:e1002358.

    https://doi.org/10.7554/eLife.15316

    • Google Scholar
17. 1. Drané P
    2. Ouararhni K
    3. Depaux A
    4. Shuaib M
    5. Hamiche A

    (2010) The death-associated protein DAXX is a novel histone chaperone involved in the replication-independent deposition of H3.3

    Genes & Development 24:1253–1265.

    https://doi.org/10.1101/gad.566910

    • PubMed
    • Google Scholar
18. 1. Dunleavy EM
    2. Roche D
    3. Tagami H
    4. Lacoste N
    5. Ray-Gallet D
    6. Nakamura Y
    7. Daigo Y
    8. Nakatani Y
    9. Almouzni-Pettinotti G

    (2009) HJURP is a cell-cycle-dependent maintenance and deposition factor of CENP-A at centromeres

    Cell 137:485–497.

    https://doi.org/10.1016/j.cell.2009.02.040

    • PubMed
    • Google Scholar
19. 1. Egan B
    2. Yuan CC
    3. Craske ML
    4. Labhart P
    5. Guler GD
    6. Arnott D
    7. Maile TM
    8. Busby J
    9. Henry C
    10. Kelly TK
    11. Tindell CA
    12. Jhunjhunwala S
    13. Zhao F
    14. Hatton C
    15. Bryant BM
    16. Classon M
    17. Trojer P

    (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:e0166438.

    https://doi.org/10.1371/journal.pone.0166438

    • PubMed
    • Google Scholar
20. 1. ENCODE Project Consortium

    (2012) An integrated encyclopedia of DNA elements in the human genome

    Nature 489:57–74.

    https://doi.org/10.1038/nature11247

    • PubMed
    • Google Scholar
21. 1. Ernst J
    2. Kheradpour P
    3. Mikkelsen TS
    4. Shoresh N
    5. Ward LD
    6. Epstein CB
    7. Zhang X
    8. Wang L
    9. Issner R
    10. Coyne M
    11. Ku M
    12. Durham T
    13. Kellis M
    14. Bernstein BE

    (2011) Mapping and analysis of chromatin state dynamics in nine human cell types

    Nature 473:43–49.

    https://doi.org/10.1038/nature09906

    • PubMed
    • Google Scholar
22. 1. Ernst J
    2. Kellis M

    (2012) ChromHMM: automating chromatin-state discovery and characterization

    Nature Methods 9:215–216.

    https://doi.org/10.1038/nmeth.1906

    • PubMed
    • Google Scholar
23. 1. Garel A
    2. Axel R

    (1976) Selective digestion of transcriptionally active ovalbumin genes from oviduct nuclei

    PNAS 73:3966–3970.

    https://doi.org/10.1073/pnas.73.11.3966

    • PubMed
    • Google Scholar
24. 1. Gévry N
    2. Hardy S
    3. Jacques PE
    4. Laflamme L
    5. Svotelis A
    6. Robert F
    7. Gaudreau L

    (2009) Histone H2A.Z is essential for estrogen receptor signaling

    Genes & Development 23:1522–1533.

    https://doi.org/10.1101/gad.1787109

    • PubMed
    • Google Scholar
25. 1. Giaimo BD
    2. Ferrante F
    3. Herchenröther A
    4. Hake SB
    5. Borggrefe T

    (2019) The histone variant H2A.Z in gene regulation

    Epigenetics & Chromatin 12:1–22.

    https://doi.org/10.1186/s13072-019-0274-9

    • PubMed
    • Google Scholar
26. 1. Gilbert N
    2. Boyle S
    3. Fiegler H
    4. Woodfine K
    5. Carter NP
    6. Bickmore WA

    (2004) Chromatin architecture of the human genome: gene-rich domains are enriched in open chromatin fibers

    Cell 118:555–566.

    https://doi.org/10.1016/j.cell.2004.08.011

    • PubMed
    • Google Scholar
27. 1. Goldberg AD
    2. Banaszynski LA
    3. Noh KM
    4. Lewis PW
    5. Elsaesser SJ
    6. Stadler S
    7. Dewell S
    8. Law M
    9. Guo X
    10. Li X
    11. Wen D
    12. Chapgier A
    13. DeKelver RC
    14. Miller JC
    15. Lee YL
    16. Boydston EA
    17. Holmes MC
    18. Gregory PD
    19. Greally JM
    20. Rafii S
    21. Yang C
    22. Scambler PJ
    23. Garrick D
    24. Gibbons RJ
    25. Higgs DR
    26. Cristea IM
    27. Urnov FD
    28. Zheng D
    29. Allis CD

    (2010) Distinct factors control histone variant H3.3 localization at specific genomic regions

    Cell 140:678–691.

    https://doi.org/10.1016/j.cell.2010.01.003

    • PubMed
    • Google Scholar
28. 1. Greaves IK
    2. Rangasamy D
    3. Ridgway P
    4. Tremethick DJ

    (2007) H2A.Z contributes to the unique 3D structure of the centromere

    PNAS 104:525–530.

    https://doi.org/10.1073/pnas.0607870104

    • PubMed
    • Google Scholar
29. 1. Gursoy-Yuzugullu O
    2. Ayrapetov MK
    3. Price BD

    (2015) Histone chaperone Anp32e removes H2A.Z from DNA double-strand breaks and promotes nucleosome reorganization and DNA repair

    PNAS 112:7507–7512.

    https://doi.org/10.1073/pnas.1504868112

    • PubMed
    • Google Scholar
30. 1. Ha M
    2. Kraushaar DC
    3. Zhao K

    (2014) Genome-wide analysis of H3.3 dissociation reveals high nucleosome turnover at distal regulatory regions of embryonic stem cells

    Epigenetics & Chromatin 7:38.

    https://doi.org/10.1186/1756-8935-7-38

    • PubMed
    • Google Scholar
31. 1. Heinz S
    2. Benner C
    3. Spann N
    4. Bertolino E
    5. Lin YC
    6. Laslo P
    7. Cheng JX
    8. Murre C
    9. Singh H
    10. Glass CK

    (2010) Simple combinations of lineage-determining transcription factors prime cis-regulatory elements required for macrophage and B cell identities

    Molecular Cell 38:576–589.

    https://doi.org/10.1016/j.molcel.2010.05.004

    • PubMed
    • Google Scholar
32. 1. Horikoshi N
    2. Sato K
    3. Shimada K
    4. Arimura Y
    5. Osakabe A
    6. Tachiwana H
    7. Hayashi-Takanaka Y
    8. Iwasaki W
    9. Kagawa W
    10. Harata M
    11. Kimura H
    12. Kurumizaka H

    (2013) Structural polymorphism in the L1 loop regions of human H2A.Z.1 and H2A.Z.2

    Acta Crystallographica Section D Biological Crystallography 69:2431–2439.

    https://doi.org/10.1107/S090744491302252X

    • Google Scholar
33. 1. Ishimi Y
    2. Hirosumi J
    3. Sato W
    4. Sugasawa K
    5. Yokota S
    6. Hanaoka F
    7. Yamada M

    (1984) Purification and initial characterization of a protein which facilitates assembly of nucleosome-like structure from mammalian cells

    European Journal of Biochemistry 142:431–439.

    https://doi.org/10.1111/j.1432-1033.1984.tb08305.x

    • PubMed
    • Google Scholar
34. 1. Jackson V

    (1990) In vivo studies on the dynamics of histone-DNA interaction: evidence for nucleosome dissolution during replication and transcription and a low level of dissolution independent of both

    Biochemistry 29:719–731.

    https://doi.org/10.1021/bi00455a019

    • PubMed
    • Google Scholar
35. 1. Jansen LE
    2. Black BE
    3. Foltz DR
    4. Cleveland DW

    (2007) Propagation of centromeric chromatin requires exit from mitosis

    Journal of Cell Biology 176:795–805.

    https://doi.org/10.1083/jcb.200701066

    • PubMed
    • Google Scholar
36. 1. Jeronimo C
    2. Poitras C
    3. Robert F

    (2019) Histone recycling by FACT and Spt6 during transcription prevents the scrambling of histone modifications

    Cell Reports 28:1206–1218.

    https://doi.org/10.1016/j.celrep.2019.06.097

    • PubMed
    • Google Scholar
37. 1. Jin C
    2. Zang C
    3. Wei G
    4. Cui K
    5. Peng W
    6. Zhao K
    7. Felsenfeld G

    (2009) H3.3/H2A.Z double variant-containing nucleosomes mark 'nucleosome-free regions' of active promoters and other regulatory regions

    Nature Genetics 41:941–945.

    https://doi.org/10.1038/ng.409

    • PubMed
    • Google Scholar
38. 1. Jin C
    2. Felsenfeld G

    (2007) Nucleosome stability mediated by histone variants H3.3 and H2A.Z

    Genes & Development 21:1519–1529.

    https://doi.org/10.1101/gad.1547707

    • PubMed
    • Google Scholar
39. 1. Jolly C
    2. Metz A
    3. Govin J
    4. Vigneron M
    5. Turner BM
    6. Khochbin S
    7. Vourc'h C

    (2004) Stress-induced transcription of satellite III repeats

    Journal of Cell Biology 164:25–33.

    https://doi.org/10.1083/jcb.200306104

    • PubMed
    • Google Scholar
40. 1. Kemble DJ
    2. McCullough LL
    3. Whitby FG
    4. Formosa T
    5. Hill CP

    (2015) FACT disrupts nucleosome structure by binding H2A-H2B with conserved peptide motifs

    Molecular Cell 60:294–306.

    https://doi.org/10.1016/j.molcel.2015.09.008

    • PubMed
    • Google Scholar
41. 1. Kim D
    2. Langmead B
    3. Salzberg SL

    (2015) HISAT: a fast spliced aligner with low memory requirements

    Nature Methods 12:357–360.

    https://doi.org/10.1038/nmeth.3317

    • PubMed
    • Google Scholar
42. 1. Kimura H
    2. Takizawa N
    3. Allemand E
    4. Hori T
    5. Iborra FJ
    6. Nozaki N
    7. Muraki M
    8. Hagiwara M
    9. Krainer AR
    10. Fukagawa T
    11. Okawa K

    (2006) A novel histone exchange factor, protein phosphatase 2cgamma, mediates the exchange and dephosphorylation of H2A-H2B

    Journal of Cell Biology 175:389–400.

    https://doi.org/10.1083/jcb.200608001

    • PubMed
    • Google Scholar
43. 1. Kimura H
    2. Cook PR

    (2001) Kinetics of core histones in living human cells: little exchange of H3 and H4 and some rapid exchange of H2B

    The Journal of Cell Biology 153:1341–1353.

    https://doi.org/10.1083/jcb.153.7.1341

    • PubMed
    • Google Scholar
44. 1. Lacoste N
    2. Woolfe A
    3. Tachiwana H
    4. Garea AV
    5. Barth T
    6. Cantaloube S
    7. Kurumizaka H
    8. Imhof A
    9. Almouzni G

    (2014) Mislocalization of the centromeric histone variant CenH3/CENP-A in human cells depends on the chaperone DAXX

    Molecular Cell 53:631–644.

    https://doi.org/10.1016/j.molcel.2014.01.018

    • PubMed
    • Google Scholar
45. 1. Langmead B
    2. Salzberg SL

    (2012) Fast gapped-read alignment with bowtie 2

    Nature Methods 9:357–359.

    https://doi.org/10.1038/nmeth.1923

    • PubMed
    • Google Scholar
46. 1. Latrick CM
    2. Marek M
    3. Ouararhni K
    4. Papin C
    5. Stoll I
    6. Ignatyeva M
    7. Obri A
    8. Ennifar E
    9. Dimitrov S
    10. Romier C
    11. Hamiche A

    (2016) Molecular basis and specificity of H2A.Z-H2B recognition and deposition by the histone chaperone YL1

    Nature Structural & Molecular Biology 23:309–316.

    https://doi.org/10.1038/nsmb.3189

    • PubMed
    • Google Scholar
47. 1. Leonhardt H
    2. Rahn HP
    3. Weinzierl P
    4. Sporbert A
    5. Cremer T
    6. Zink D
    7. Cardoso MC

    (2000) Dynamics of DNA replication factories in living cells

    Journal of Cell Biology 149:271–280.

    https://doi.org/10.1083/jcb.149.2.271

    • PubMed
    • Google Scholar
48. 1. Liang X
    2. Shan S
    3. Pan L
    4. Zhao J
    5. Ranjan A
    6. Wang F
    7. Zhang Z
    8. Huang Y
    9. Feng H
    10. Wei D
    11. Huang L
    12. Liu X
    13. Zhong Q
    14. Lou J
    15. Li G
    16. Wu C
    17. Zhou Z

    (2016) Structural basis of H2A.Z recognition by SRCAP chromatin-remodeling subunit YL1

    Nature Structural & Molecular Biology 23:317–323.

    https://doi.org/10.1038/nsmb.3190

    • Google Scholar
49. 1. Link S
    2. Spitzer RMM
    3. Sana M
    4. Torrado M
    5. Völker-Albert MC
    6. Keilhauer EC
    7. Burgold T
    8. Pünzeler S
    9. Low JKK
    10. Lindström I
    11. Nist A
    12. Regnard C
    13. Stiewe T
    14. Hendrich B
    15. Imhof A
    16. Mann M
    17. Mackay JP
    18. Bartkuhn M
    19. Hake SB

    (2018) PWWP2A binds distinct chromatin moieties and interacts with an MTA1-specific core NuRD complex

    Nature Communications 9:4300.

    https://doi.org/10.1038/s41467-018-06665-5

    • PubMed
    • Google Scholar
50. 1. Liu Y
    2. Zhou K
    3. Zhang N
    4. Wei H
    5. Tan YZ
    6. Zhang Z
    7. Carragher B
    8. Potter CS
    9. D'Arcy S
    10. Luger K

    (2020) FACT caught in the act of manipulating the nucleosome

    Nature 577:426–431.

    https://doi.org/10.1038/s41586-019-1820-0

    • PubMed
    • Google Scholar
51. 1. Louters L
    2. Chalkley R

    (1985)

    Histones H1, H2A, and H2B in vivo

    Biochemistry 24:3080–3085.

    • Google Scholar
52. 1. Luger K
    2. Mäder AW
    3. Richmond RK
    4. Sargent DF
    5. Richmond TJ

    (1997) Crystal structure of the nucleosome core particle at 2.8 A resolution

    Nature 389:251–260.

    https://doi.org/10.1038/38444

    • PubMed
    • Google Scholar
53. 1. Luk E
    2. Ranjan A
    3. Fitzgerald PC
    4. Mizuguchi G
    5. Huang Y
    6. Wei D
    7. Wu C

    (2010) Stepwise histone replacement by SWR1 requires dual activation with histone H2A.Z and canonical nucleosome

    Cell 143:725–736.

    https://doi.org/10.1016/j.cell.2010.10.019

    • PubMed
    • Google Scholar
54. 1. Maehara K
    2. Harada A
    3. Sato Y
    4. Matsumoto M
    5. Nakayama KI
    6. Kimura H
    7. Ohkawa Y

    (2015) Tissue-specific expression of histone H3 variants diversified after species separation

    Epigenetics & Chromatin 8:35.

    https://doi.org/10.1186/s13072-015-0027-3

    • PubMed
    • Google Scholar
55. 1. Maison C
    2. Bailly D
    3. Peters AH
    4. Quivy JP
    5. Roche D
    6. Taddei A
    7. Lachner M
    8. Jenuwein T
    9. Almouzni G

    (2002) Higher-order structure in pericentric heterochromatin involves a distinct pattern of histone modification and an RNA component

    Nature Genetics 30:329–334.

    https://doi.org/10.1038/ng843

    • PubMed
    • Google Scholar
56. 1. Mao Z
    2. Pan L
    3. Wang W
    4. Sun J
    5. Shan S
    6. Dong Q
    7. Liang X
    8. Dai L
    9. Ding X
    10. Chen S
    11. Zhang Z
    12. Zhu B
    13. Zhou Z

    (2014) Anp32e, a higher eukaryotic histone chaperone directs preferential recognition for H2A.Z

    Cell Research 24:389–399.

    https://doi.org/10.1038/cr.2014.30

    • PubMed
    • Google Scholar
57. 1. Martini E
    2. Roche DM
    3. Marheineke K
    4. Verreault A
    5. Almouzni G

    (1998) Recruitment of phosphorylated chromatin assembly factor 1 to chromatin after UV irradiation of human cells

    Journal of Cell Biology 143:563–575.

    https://doi.org/10.1083/jcb.143.3.563

    • PubMed
    • Google Scholar
58. 1. Misteli T
    2. Spector DL

    (1996) Serine/threonine phosphatase 1 modulates the subnuclear distribution of pre-mRNA splicing factors

    Molecular Biology of the Cell 7:1559–1572.

    https://doi.org/10.1091/mbc.7.10.1559

    • PubMed
    • Google Scholar
59. 1. Mito Y
    2. Henikoff JG
    3. Henikoff S

    (2005) Genome-scale profiling of histone H3.3 replacement patterns

    Nature Genetics 37:1090–1097.

    https://doi.org/10.1038/ng1637

    • PubMed
    • Google Scholar
60. 1. Mizuguchi G

    (2004) ATP-Driven exchange of histone H2AZ variant catalyzed by SWR1 chromatin remodeling complex

    Science 303:343–348.

    https://doi.org/10.1126/science.1090701

    • Google Scholar
61. 1. Moggs JG
    2. Grandi P
    3. Quivy JP
    4. Jónsson ZO
    5. Hübscher U
    6. Becker PB
    7. Almouzni G

    (2000) A CAF-1-PCNA-mediated chromatin assembly pathway triggered by sensing DNA damage

    Molecular and Cellular Biology 20:1206–1218.

    https://doi.org/10.1128/MCB.20.4.1206-1218.2000

    • PubMed
    • Google Scholar
62. 1. Munakata T
    2. Adachi N
    3. Yokoyama N
    4. Kuzuhara T
    5. Horikoshi M

    (2000) A human homologue of yeast anti-silencing factor has histone chaperone activity

    Genes to Cells 5:221–233.

    https://doi.org/10.1046/j.1365-2443.2000.00319.x

    • PubMed
    • Google Scholar
63. 1. Murphy PJ
    2. Wu SF
    3. James CR
    4. Wike CL
    5. Cairns BR

    (2018) Placeholder nucleosomes underlie Germline-to-Embryo DNA methylation reprogramming

    Cell 172:993–1006.

    https://doi.org/10.1016/j.cell.2018.01.022

    • PubMed
    • Google Scholar
64. 1. Murphy KE
    2. Meng FW
    3. Makowski CE
    4. Murphy PJ

    (2020) Genome-wide chromatin accessibility is restricted by ANP32E

    Nature Communications 11:5063.

    https://doi.org/10.1038/s41467-020-18821-x

    • PubMed
    • Google Scholar
65. 1. Nadal S
    2. Raj R
    3. Mohammed S
    4. Davis BG

    (2018) Synthetic post-translational modification of histones

    Current Opinion in Chemical Biology 45:35–47.

    https://doi.org/10.1016/j.cbpa.2018.02.004

    • Google Scholar
66. 1. Nothjunge S
    2. Nührenberg TG
    3. Grüning BA
    4. Doppler SA
    5. Preissl S
    6. Schwaderer M
    7. Rommel C
    8. Krane M
    9. Hein L
    10. Gilsbach R

    (2017) DNA methylation signatures follow preformed chromatin compartments in cardiac myocytes

    Nature Communications 8:1667.

    https://doi.org/10.1038/s41467-017-01724-9

    • PubMed
    • Google Scholar
67. 1. Obri A
    2. Ouararhni K
    3. Papin C
    4. Diebold ML
    5. Padmanabhan K
    6. Marek M
    7. Stoll I
    8. Roy L
    9. Reilly PT
    10. Mak TW
    11. Dimitrov S
    12. Romier C
    13. Hamiche A

    (2014) ANP32E is a histone chaperone that removes H2A.Z from chromatin

    Nature 505:648–653.

    https://doi.org/10.1038/nature12922

    • PubMed
    • Google Scholar
68. 1. Okuno Y
    2. Imamoto N
    3. Yoneda Y

    (1993) 70-kDa heat-shock cognate protein colocalizes with karyophilic proteins into the nucleus during their transport in vitro

    Experimental Cell Research 206:134–142.

    https://doi.org/10.1006/excr.1993.1129

    • PubMed
    • Google Scholar
69. 1. Orlando DA
    2. Chen MW
    3. Brown VE
    4. Solanki S
    5. Choi YJ
    6. Olson ER
    7. Fritz CC
    8. Bradner JE
    9. Guenther MG

    (2014) Quantitative ChIP-Seq normalization reveals global modulation of the epigenome

    Cell Reports 9:1163–1170.

    https://doi.org/10.1016/j.celrep.2014.10.018

    • PubMed
    • Google Scholar
70. 1. Orphanides G
    2. LeRoy G
    3. Chang CH
    4. Luse DS
    5. Reinberg D

    (1998) FACT, a factor that facilitates transcript elongation through nucleosomes

    Cell 92:105–116.

    https://doi.org/10.1016/S0092-8674(00)80903-4

    • PubMed
    • Google Scholar
71. 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.

    • PubMed
    • Google Scholar
72. 1. Pchelintsev NA
    2. McBryan T
    3. Rai TS
    4. van Tuyn J
    5. Ray-Gallet D
    6. Almouzni G
    7. Adams PD

    (2013) Placing the HIRA histone chaperone complex in the chromatin landscape

    Cell Reports 3:1012–1019.

    https://doi.org/10.1016/j.celrep.2013.03.026

    • PubMed
    • Google Scholar
73. 1. Quinlan AR
    2. Hall IM

    (2010) BEDTools: a flexible suite of utilities for comparing genomic features

    Bioinformatics 26:841–842.

    https://doi.org/10.1093/bioinformatics/btq033

    • PubMed
    • Google Scholar
74. 1. Raisner RM
    2. Hartley PD
    3. Meneghini MD
    4. Bao MZ
    5. Liu CL
    6. Schreiber SL
    7. Rando OJ
    8. Madhani HD

    (2005) Histone variant H2A.Z marks the 5' ends of both active and inactive genes in euchromatin

    Cell 123:233–248.

    https://doi.org/10.1016/j.cell.2005.10.002

    • PubMed
    • Google Scholar
75. 1. Ramírez F
    2. Dündar F
    3. Diehl S
    4. Grüning BA
    5. Manke T

    (2014) deepTools: a flexible platform for exploring deep-sequencing data

    Nucleic Acids Research 42:W187–W191.

    https://doi.org/10.1093/nar/gku365

    • PubMed
    • Google Scholar
76. 1. Rangasamy D
    2. Berven L
    3. Ridgway P
    4. Tremethick DJ

    (2003) Pericentric heterochromatin becomes enriched with H2A.Z during early mammalian development

    The EMBO Journal 22:1599–1607.

    https://doi.org/10.1093/emboj/cdg160

    • PubMed
    • Google Scholar
77. 1. Ransom M
    2. Dennehey BK
    3. Tyler JK

    (2010) Chaperoning histones during DNA replication and repair

    Cell 140:183–195.

    https://doi.org/10.1016/j.cell.2010.01.004

    • PubMed
    • Google Scholar
78. 1. Ray-Gallet D
    2. Quivy JP
    3. Scamps C
    4. Martini EM
    5. Lipinski M
    6. Almouzni G

    (2002) HIRA is critical for a nucleosome assembly pathway independent of DNA synthesis

    Molecular Cell 9:1091–1100.

    https://doi.org/10.1016/S1097-2765(02)00526-9

    • PubMed
    • Google Scholar
79. 1. Ray-Gallet D
    2. Woolfe A
    3. Vassias I
    4. Pellentz C
    5. Lacoste N
    6. Puri A
    7. Schultz DC
    8. Pchelintsev NA
    9. Adams PD
    10. Jansen LE
    11. Almouzni G

    (2011) Dynamics of histone H3 deposition in vivo reveal a nucleosome gap-filling mechanism for H3.3 to maintain chromatin integrity

    Molecular Cell 44:928–941.

    https://doi.org/10.1016/j.molcel.2011.12.006

    • PubMed
    • Google Scholar
80. 1. Reilly PT
    2. Afzal S
    3. Wakeham A
    4. Haight J
    5. You-Ten A
    6. Zaugg K
    7. Dembowy J
    8. Young A
    9. Mak TW

    (2010) Generation and characterization of the Anp32e-deficient mouse

    PLOS ONE 5:e13597.

    https://doi.org/10.1371/journal.pone.0013597

    • PubMed
    • Google Scholar
81. 1. Rivera-Mulia JC
    2. Gilbert DM

    (2016) Replication timing and transcriptional control: beyond cause and effect-part III

    Current Opinion in Cell Biology 40:168–178.

    https://doi.org/10.1016/j.ceb.2016.03.022

    • PubMed
    • Google Scholar
82. 1. Rizzi N
    2. Denegri M
    3. Chiodi I
    4. Corioni M
    5. Valgardsdottir R
    6. Cobianchi F
    7. Riva S
    8. Biamonti G

    (2004) Transcriptional activation of a constitutive heterochromatic domain of the human genome in response to heat shock

    Molecular Biology of the Cell 15:543–551.

    https://doi.org/10.1091/mbc.e03-07-0487

    • PubMed
    • Google Scholar
83. 1. Roadmap Epigenomics Consortium
    2. Kundaje A
    3. Meuleman W
    4. Ernst J
    5. Bilenky M
    6. Yen A
    7. Heravi-Moussavi A
    8. Kheradpour P
    9. Zhang Z
    10. Wang J
    11. Ziller MJ
    12. Amin V
    13. Whitaker JW
    14. Schultz MD
    15. Ward LD
    16. Sarkar A
    17. Quon G
    18. Sandstrom RS
    19. Eaton ML
    20. Wu YC
    21. Pfenning AR
    22. Wang X
    23. Claussnitzer M
    24. Liu Y
    25. Coarfa C
    26. Harris RA
    27. Shoresh N
    28. Epstein CB
    29. Gjoneska E
    30. Leung D
    31. Xie W
    32. Hawkins RD
    33. Lister R
    34. Hong C
    35. Gascard P
    36. Mungall AJ
    37. Moore R
    38. Chuah E
    39. Tam A
    40. Canfield TK
    41. Hansen RS
    42. Kaul R
    43. Sabo PJ
    44. Bansal MS
    45. Carles A
    46. Dixon JR
    47. Farh KH
    48. Feizi S
    49. Karlic R
    50. Kim AR
    51. Kulkarni A
    52. Li D
    53. Lowdon R
    54. Elliott G
    55. Mercer TR
    56. Neph SJ
    57. Onuchic V
    58. Polak P
    59. Rajagopal N
    60. Ray P
    61. Sallari RC
    62. Siebenthall KT
    63. Sinnott-Armstrong NA
    64. Stevens M
    65. Thurman RE
    66. Wu J
    67. Zhang B
    68. Zhou X
    69. Beaudet AE
    70. Boyer LA
    71. De Jager PL
    72. Farnham PJ
    73. Fisher SJ
    74. Haussler D
    75. Jones SJ
    76. Li W
    77. Marra MA
    78. McManus MT
    79. Sunyaev S
    80. Thomson JA
    81. Tlsty TD
    82. Tsai LH
    83. Wang W
    84. Waterland RA
    85. Zhang MQ
    86. Chadwick LH
    87. Bernstein BE
    88. Costello JF
    89. Ecker JR
    90. Hirst M
    91. Meissner A
    92. Milosavljevic A
    93. Ren B
    94. Stamatoyannopoulos JA
    95. Wang T
    96. Kellis M

    (2015) Integrative analysis of 111 reference human epigenomes

    Nature 518:317–330.

    https://doi.org/10.1038/nature14248

    • PubMed
    • Google Scholar
84. 1. Robinson JT
    2. Thorvaldsdóttir H
    3. Winckler W
    4. Guttman M
    5. Lander ES
    6. Getz G
    7. Mesirov JP

    (2011) Integrative genomics viewer

    Nature Biotechnology 29:24–26.

    https://doi.org/10.1038/nbt.1754

    • PubMed
    • Google Scholar
85. 1. Saitoh N
    2. Uchimura Y
    3. Tachibana T
    4. Sugahara S
    5. Saitoh H
    6. Nakao M

    (2006) In situ SUMOylation analysis reveals a modulatory role of RanBP2 in the nuclear rim and PML bodies

    Experimental Cell Research 312:1418–1430.

    https://doi.org/10.1016/j.yexcr.2006.01.013

    • Google Scholar
86. 1. Saksouk N
    2. Simboeck E
    3. Déjardin J

    (2015) Constitutive heterochromatin formation and transcription in mammals

    Epigenetics & Chromatin 8:3.

    https://doi.org/10.1186/1756-8935-8-3

    • PubMed
    • Google Scholar
87. 1. Sarcinella E
    2. Zuzarte PC
    3. Lau PN
    4. Draker R
    5. Cheung P

    (2007) Monoubiquitylation of H2A.Z distinguishes its association with euchromatin or facultative heterochromatin

    Molecular and Cellular Biology 27:6457–6468.

    https://doi.org/10.1128/MCB.00241-07

    • PubMed
    • Google Scholar
88. 1. Schneider CA
    2. Rasband WS
    3. Eliceiri KW

    (2012) NIH image to ImageJ: 25 years of image analysis

    Nature Methods 9:671–675.

    https://doi.org/10.1038/nmeth.2089

    • PubMed
    • Google Scholar
89. 1. Schones DE
    2. Cui K
    3. Cuddapah S
    4. Roh T-Y
    5. Barski A
    6. Wang Z
    7. Wei G
    8. Zhao K

    (2008) Dynamic regulation of nucleosome positioning in the human genome

    Cell 132:887–898.

    https://doi.org/10.1016/j.cell.2008.02.022

    • Google Scholar
90. 1. Schwoebel ED
    2. Ho TH
    3. Moore MS

    (2002) The mechanism of inhibition of Ran-dependent nuclear transport by cellular ATP depletion

    Journal of Cell Biology 157:963–974.

    https://doi.org/10.1083/jcb.200111077

    • PubMed
    • Google Scholar
91. 1. Shia WJ
    2. Li B
    3. Workman JL

    (2006) SAS-mediated acetylation of histone H4 lys 16 is required for H2A.Z incorporation at Subtelomeric regions in Saccharomyces cerevisiae

    Genes & Development 20:2507–2512.

    https://doi.org/10.1101/gad.1439206

    • PubMed
    • Google Scholar
92. 1. Shibahara K
    2. Stillman B

    (1999) Replication-dependent marking of DNA by PCNA facilitates CAF-1-coupled inheritance of chromatin

    Cell 96:575–585.

    https://doi.org/10.1016/S0092-8674(00)80661-3

    • PubMed
    • Google Scholar
93. 1. Siwek W
    2. Gómez-Rodríguez M
    3. Sobral D
    4. Corrêa IR
    5. Jansen LET

    (2018) time-ChIP: a method to determine Long-Term Locus-Specific nucleosome inheritance

     Methods in Molecular Biology  1832:131–158.

    https://doi.org/10.1007/978-1-4939-8663-7_7

    • Google Scholar
94. 1. Smith T
    2. Heger A
    3. Sudbery I

    (2017) UMI-tools: modeling sequencing errors in unique molecular identifiers to improve quantification accuracy

    Genome Research 27:491–499.

    https://doi.org/10.1101/gr.209601.116

    • PubMed
    • Google Scholar
95. 1. Smith S
    2. Stillman B

    (1989) Purification and characterization of CAF-I, a human cell factor required for chromatin assembly during DNA replication in vitro

    Cell 58:15–25.

    https://doi.org/10.1016/0092-8674(89)90398-X

    • PubMed
    • Google Scholar
96. 1. Spiker S
    2. Murray MG
    3. Thompson WF

    (1983) DNase I sensitivity of transcriptionally active genes in intact nuclei and isolated chromatin of plants

    PNAS 80:815–819.

    https://doi.org/10.1073/pnas.80.3.815

    • PubMed
    • Google Scholar
97. 1. Sun L
    2. Pierrakeas L
    3. Li T
    4. Luk E

    (2020) Thermosensitive nucleosome editing reveals the role of DNA sequence in targeted histone variant deposition

    Cell Reports 30:257–268.

    https://doi.org/10.1016/j.celrep.2019.12.006

    • PubMed
    • Google Scholar
98. 1. Sun L
    2. Luk E

    (2017) Dual function of Swc5 in SWR remodeling ATPase activation and histone H2A eviction

    Nucleic Acids Research 45:9931–9946.

    https://doi.org/10.1093/nar/gkx589

    • PubMed
    • Google Scholar
99. 1. Suto RK
    2. Clarkson MJ
    3. Tremethick DJ
    4. Luger K

    (2000) Crystal structure of a nucleosome core particle containing the variant histone H2A.Z

    Nature Structural Biology 7:1121–1124.

    https://doi.org/10.1038/81971

    • PubMed
    • Google Scholar
100. 1. Tachiwana H
     2. Osakabe A
     3. Kimura H
     4. Kurumizaka H

     (2008) Nucleosome formation with the testis-specific histone H3 variant, H3t, by human nucleosome assembly proteins in vitro

     Nucleic Acids Research 36:2208–2218.

     https://doi.org/10.1093/nar/gkn060

     • Google Scholar
101. 1. Tachiwana H
     2. Kagawa W
     3. Osakabe A
     4. Kawaguchi K
     5. Shiga T
     6. Hayashi-Takanaka Y
     7. Kimura H
     8. Kurumizaka H

     (2010) Structural basis of instability of the nucleosome containing a testis-specific histone variant, human H3T

     PNAS 107:10454–10459.

     https://doi.org/10.1073/pnas.1003064107

     • PubMed
     • Google Scholar
102. 1. Tachiwana H
     2. Müller S
     3. Blümer J
     4. Klare K
     5. Musacchio A
     6. Almouzni G

     (2015) HJURP involvement in de novo CenH3(CENP-A) and CENP-C recruitment

     Cell Reports 11:22–32.

     https://doi.org/10.1016/j.celrep.2015.03.013

     • PubMed
     • Google Scholar
103. 1. Tagami H
     2. Ray-Gallet D
     3. Almouzni G
     4. Nakatani Y

     (2004) Histone H3.1 and H3.3 Complexes Mediate Nucleosome Assembly Pathways Dependent or Independent of DNA Synthesis

     Cell 116:51–61.

     https://doi.org/10.1016/S0092-8674(03)01064-X

     • Google Scholar
104. 1. Tornà J
     2. Ray-Gallet D
     3. Boyarchuk E
     4. Garnier M
     5. Le Baccon P
     6. Coulon A
     7. Orsi GA
     8. Almouzni G

     (2020) Two HIRA-dependent pathways mediate H3.3 de novo deposition and recycling during transcription

     Nature Structural & Molecular Biology 27:1057–1068.

     https://doi.org/10.1038/s41594-020-0492-7

     • Google Scholar
105. 1. Tsompana M
     2. Buck MJ

     (2014) Chromatin accessibility: a window into the genome

     Epigenetics & Chromatin 7:33.

     https://doi.org/10.1186/1756-8935-7-33

     • Google Scholar
106. 1. Tsunaka Y
     2. Fujiwara Y
     3. Oyama T
     4. Hirose S
     5. Morikawa K

     (2016) Integrated molecular mechanism directing nucleosome reorganization by human FACT

     Genes & Development 30:673–686.

     https://doi.org/10.1101/gad.274183.115

     • Google Scholar
107. 1. Tyler JK
     2. Bulger M
     3. Kamakaka RT
     4. Kobayashi R
     5. Kadonaga JT

     (1996) The p55 subunit of Drosophila chromatin assembly factor 1 is homologous to a histone deacetylase-associated protein

     Molecular and Cellular Biology 16:6149–6159.

     https://doi.org/10.1128/MCB.16.11.6149

     • Google Scholar
108. 1. Tyler JK
     2. Adams CR
     3. Chen S-R
     4. Kobayashi R
     5. Kamakaka RT
     6. Kadonaga JT

     (1999) The RCAF complex mediates chromatin assembly during DNA replication and repair

     Nature 402:555–560.

     https://doi.org/10.1038/990147

     • Google Scholar
109. 1. Tyler JK
     2. Collins KA
     3. Prasad-Sinha J
     4. Amiott E
     5. Bulger M
     6. Harte PJ
     7. Kobayashi R
     8. Kadonaga JT

     (2001) Interaction between the Drosophila CAF-1 and ASF1 Chromatin Assembly Factors

     Molecular and Cellular Biology 21:6574–6584.

     https://doi.org/10.1128/MCB.21.19.6574-6584.2001

     • Google Scholar
110. 1. Valgardsdottir R
     2. Chiodi I
     3. Giordano M
     4. Cobianchi F
     5. Riva S
     6. Biamonti G

     (2005) Structural and Functional Characterization of Noncoding Repetitive RNAs Transcribed in Stressed Human Cells

     Molecular Biology of the Cell 16:2597–2604.

     https://doi.org/10.1091/mbc.e04-12-1078

     • Google Scholar
111. 1. Wang T
     2. Liu Y
     3. Edwards G
     4. Krzizike D
     5. Scherman H
     6. Luger K

     (2018) The histone chaperone FACT modulates nucleosome structure by tethering its components

     Life Science Alliance 1:e201800107.

     https://doi.org/10.26508/lsa.201800107

     • PubMed
     • Google Scholar
112. 1. Weintraub H
     2. Groudine M

     (1976) Chromosomal subunits in active genes have an altered conformation

     Science 193:848–856.

     https://doi.org/10.1126/science.948749

     • Google Scholar
113. 1. Wittmeyer J
     2. Formosa T

     (1997) The Saccharomyces cerevisiae DNA polymerase alpha catalytic subunit interacts with Cdc68/Spt16 and with Pob3, a protein similar to an HMG1-like protein

     Molecular and Cellular Biology 17:4178–4190.

     https://doi.org/10.1128/MCB.17.7.4178

     • Google Scholar
114. 1. Wong MM
     2. Cox LK
     3. Chrivia JC

     (2007) The Chromatin Remodeling Protein, SRCAP, Is Critical for Deposition of the Histone Variant H2A.Z at Promoters*

     Journal of Biological Chemistry 282:26132–26139.

     https://doi.org/10.1074/jbc.M703418200

     • Google Scholar
115. 1. Wu RS
     2. Tsai S
     3. Bonner WM

     (1982) Patterns of histone variant synthesis can distinguish go from G1 cells

     Cell 31:367–374.

     https://doi.org/10.1016/0092-8674(82)90130-1

     • Google Scholar
116. 1. Yildirim O
     2. Hung JH
     3. Cedeno RJ
     4. Weng Z
     5. Lengner CJ
     6. Rando OJ

     (2014) A system for genome-wide histone variant dynamics in ES cells reveals dynamic MacroH2A2 replacement at promoters

     PLOS Genetics 10:e1004515.

     https://doi.org/10.1371/journal.pgen.1004515

     • PubMed
     • Google Scholar
117. 1. Yukawa M
     2. Akiyama T
     3. Franke V
     4. Mise N
     5. Isagawa T
     6. Suzuki Y
     7. Suzuki MG
     8. Vlahovicek K
     9. Abe K
     10. Aburatani H
     11. Aoki F

     (2014) Genome-wide analysis of the chromatin composition of histone H2A and H3 variants in mouse embryonic stem cells

     PLOS ONE 9:e92689.

     https://doi.org/10.1371/journal.pone.0092689

     • PubMed
     • Google Scholar
118. 1. Zilberman D
     2. Coleman-Derr D
     3. Ballinger T
     4. Henikoff S

     (2008) Histone H2A.Z and DNA methylation are mutually antagonistic chromatin marks

     Nature 456:125–129.

     https://doi.org/10.1038/nature07324

     • PubMed
     • Google Scholar

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.

[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.

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

   "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

   "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

   "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

• 2,869

  Page views

• 331

  Downloads

• 4

  Citations

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