1. Chromosomes and Gene Expression
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Live-cell single particle imaging reveals the role of RNA polymerase II in histone H2A.Z eviction

  1. Anand Ranjan
  2. Vu Q Nguyen
  3. Sheng Liu
  4. Jan Wisniewski
  5. Jee Min Kim
  6. Xiaona Tang
  7. Gaku Mizuguchi
  8. Ejlal Elalaoui
  9. Timothy J Nickels
  10. Vivian Jou
  11. Brian P English
  12. Qinsi Zheng
  13. Ed Luk
  14. Luke D Lavis
  15. Timothee Lionnet
  16. Carl Wu  Is a corresponding author
  1. Department of Biology, Johns Hopkins University, United States
  2. Janelia Research Campus, Howard Hughes Medical Institute, United States
  3. Department of Biochemistry and Cell Biology, Stony Brook University, United States
  4. Institute of Systems Genetics, Langone Medical Center, New York University, United States
  5. Department of Molecular Biology and Genetics, Johns Hopkins School of Medicine, United States
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Cite this article as: eLife 2020;9:e55667 doi: 10.7554/eLife.55667

Abstract

The H2A.Z histone variant, a genome-wide hallmark of permissive chromatin, is enriched near transcription start sites in all eukaryotes. H2A.Z is deposited by the SWR1 chromatin remodeler and evicted by unclear mechanisms. We tracked H2A.Z in living yeast at single-molecule resolution, and found that H2A.Z eviction is dependent on RNA Polymerase II (Pol II) and the Kin28/Cdk7 kinase, which phosphorylates Serine 5 of heptapeptide repeats on the carboxy-terminal domain of the largest Pol II subunit Rpb1. These findings link H2A.Z eviction to transcription initiation, promoter escape and early elongation activities of Pol II. Because passage of Pol II through +1 nucleosomes genome-wide would obligate H2A.Z turnover, we propose that global transcription at yeast promoters is responsible for eviction of H2A.Z. Such usage of yeast Pol II suggests a general mechanism coupling eukaryotic transcription to erasure of the H2A.Z epigenetic signal.

Introduction

The H2A.Z variant of canonical histone H2A serves as a key chromatin constituent of the epigenome, providing a unique nucleosome architecture and molecular signature for eukaryotic gene transcription and other chromosome activities (Weber and Henikoff, 2014). H2A.Z is enriched at most promoters and enhancers genome-wide, and plays a role in establishing a permissive chromatin state for regulated transcription (Weber and Henikoff, 2014). H2A.Z is incorporated in nucleosomes flanking DNase hypersensitive, nucleosome-depleted regions (NDRs), especially at the so-called ‘+1 nucleosome’ overlapping with or immediately downstream of the transcription start site (TSS) (Albert et al., 2007; Weber and Henikoff, 2014). The deposition of H2A.Z in budding yeast is catalyzed by the conserved SWR1 chromatin remodeling complex in an ATP-dependent reaction involving exchange of nucleosomal H2A-H2B for H2A.Z-H2B dimers (Mizuguchi et al., 2004).

Genome-wide studies have shown that compared to nucleosomes in the gene body, the +1 nucleosome undergoes higher turnover, which is not correlated with the level of mRNA transcription by Pol II (Dion et al., 2007; Grimaldi et al., 2014; Rufiange et al., 2007). Thus, the disruptive passage of Pol II through +1 nucleosomes during infrequent mRNA transcription is unlikely to account for H2A.Z eviction on a global scale. Biochemical studies have suggested that yeast H2A.Z eviction could be due to chromatin remodeling in reverse mediated by SWR1 itself (Watanabe et al., 2013) or the related INO80 remodeler (Papamichos-Chronakis et al., 2011), but other biochemical studies found no supporting evidence (Luk et al., 2010; Wang et al., 2016). Alternatively, genome-wide assembly of the transcription pre-initiation complex (PIC) has been proposed to evict H2A.Z, but the key event in this multistep process remains elusive (Tramantano et al., 2016). To determine the dominant mechanism of H2A.Z turnover after incorporation, we took an independent approach using single-particle tracking that directly measures the levels of chromatin-free and bound H2A.Z in the physiological environment of living yeast cells, in wild type and conditional mutants for histone eviction.

Single-particle tracking (SPT) of fluorescently tagged proteins in live cells has emerged as a robust imaging technique to determine kinetic behaviors of protein factors (Elf and Barkefors, 2019; Liu and Tjian, 2018). For chromatin-interacting proteins, SPT is complementary to genome-wide chromatin immunoprecipitation-DNA sequencing technologies (ChIP-seq) without the general caveats of chemical fixation and chromatin manipulations. SPT directly measures the fast-diffusing, chromatin-free population as well as the quasi-immobile, chromatin-bound fraction tracking with macroscopic chromosome movements (Liu and Tjian, 2018; Taddei and Gasser, 2012). Here, we combine SPT in yeast with conditional depletion of candidate regulators to study the dynamic deposition and eviction of histone H2A.Z. Our results indicate a prominent role of RNA Polymerase II in the removal of H2A.Z from chromatin.

Results

Tracking single molecules of histones and remodeler SWR1

We fused the self-labeling HaloTag to H2A.Z, H2B, and Swr1 (the catalytic subunit of the SWR1 complex) for sole source expression under native promoter control and validated the function of these fusion constructs (Figure 1—figure supplement 1A). Yeast cultures were fluorescently labeled to saturation with Janelia Fluor 646 (Grimm et al., 2015; Figure 1—figure supplement 1B,C), and movies of single molecules were recorded at high temporal resolution (10 ms exposure) in live cells (Rust et al., 2006; Shim et al., 2012; Videos 13). Single molecule trajectories (n > 1000 and≥6 frames for each trajectory) were obtained from over 50 yeast cells for each strain. The data are presented as histograms of particle frequency over the diffusion coefficient (log D) extracted from mean squared displacements (MSD) (Figure 1A–D, and methods). For a more robust quantitation of diffusive populations, we also applied a kinetic modeling approach (‘Spot-On’) based on single particle displacements (Hansen et al., 2018; Figure 1E,F). We performed Spot-On analysis on single-molecule trajectories (≥3 frames), cite Spot-On values for chromatin-bound and chromatin-free fractions in the text, and provide results from both Spot-On and MSD analyses in all figures.

Figure 1 with 3 supplements see all
Diffusive behaviors of protein fusions to HaloTag (Halo) reveal chromatin-bound and free populations in live yeast.

(A, B) Normalized histograms and two-component Gaussian fits for H2A.Z-Halo (A) and Halo-H2B (B) show the log diffusion coefficient distributions. The Gaussian fit for HaloTag is shown for reference (‘Halo only’ in A). (C, D) Normalized histograms and two-component Gaussian fits for Swr1-Halo in WT cells (C) and the swc2Δ mutant (D). Solid line: sum of two-component fit; dashed line: individual component. Percent value of the slow component along with Bootstrap resampling errors and the number of trajectories (n) are indicated. (E) Cumulative distribution functions (CDF) of 10 ms displacements. (F) Spot-On results with fitting errors showing fractions of chromatin-bound molecules derived from modeling CDFs over 10–50 ms intervals. All molecules tracked with JF646 dye except Halo only, which was imaged with JF552.

Video 1
H2A.Z-Halo single molecules imaged in wild type cells.

Molecules were tracked with JF552 dye. Time is indicated on top in seconds and outline of nuclear regions are marked in yellow.

Video 2
Swr1-Halo single molecules imaged in wild type cells.

Molecules were tracked with JF646 dye. Time is indicated on top in seconds and outline of nuclear regions are marked in yellow.

Video 3
H2A.Z-Halo single molecules imaged after anchor-away of swc5.

Molecules were tracked with JF552 dye. Time is indicated on top in seconds and outline of nuclear regions are marked in yellow.

The SPT profiles for H2A.Z and H2B were best fitted by a simple model comprised of two diffusive populations—a major, slow-diffusing chromatin-bound fraction (H2A.Z: 82%, H2B: 76%, average D: 0.03 μm2s−1), and a minor, fast-diffusing chromatin-free fraction (H2A.Z: 1.18 μm2s−1, H2B: 1.29 μm2s−1) (Figure 1E,F and Figure 1—figure supplement 2A,B,E). Additional minor populations of H2A.Z and H2B with distinct diffusive values are not excluded. The fraction of chromatin-bound H2A.Z was consistent with a previous estimate by in vivo cross-linking (Mohan et al., 2018), and the D value of bound yeast H2B was also consistent with that of mammalian H2B (0.02 μm2s−1) in a previous report (Hansen et al., 2018). The ‘free’ H2A.Z fraction represents soluble H2A.Z-H2B dimers biochemically associated with histone chaperones, in addition to a minor population in complex with the ~1 MDa SWR1 complex (Luk et al., 2007). We observed similar frequencies of chromatin-bound and free H2A.Z in cells growing synchronously after release from G1 arrest into S phase (Figure 1—figure supplement 3).

In contrast to behaviors of the histones, the SWR1 complex (Swr1-Halo subunit) showed more chromatin-free diffusion. In addition, deletion of Swc2, a key subunit involved in the recruitment of SWR1 to gene promoters (Ranjan et al., 2013), substantially reduced the chromatin-bound fraction from 47% to 21% (Figure 1C,D,F). (Our imaging regime captures both stable and transiently bound SWR1 in the slow-diffusing population; the remaining 21% of slow molecules for the swc2∆ mutant may be largely attributed to transient binding). With these validations, we proceeded to investigate regulators of H2A.Z dynamics, based on the fractional changes in chromatin-bound and free H2A.Z. Notably, while the aforementioned labeling of HaloTag was adequately conducted with the JF646 dye, a superior flurophore JF552 became available in the course of this work, prompting its use in subsequent experiments for improved signal to noise (Zheng et al., 2019; Figure 1—figure supplement 1F).

Eviction of H2A.Z upon SWR1 depletion in live yeast

The steady-state chromatin occupancy for H2A.Z is a function of competing deposition and eviction pathways. To highlight H2A.Z eviction in live cells, we blocked the H2A.Z incorporation pathway at gene promoters by conditional ‘anchor-away’ (AA) depletion of the Swc5 subunit, which is not required for Swr1 recruitment (Figure 2—figure supplement 1), but essential for SWR1 activity (Haruki et al., 2008; Sun and Luk, 2017; Tramantano et al., 2016). In the AA system, rapamycin mediates heterodimerization of FRB and FKBP12 moieties fused to Swc5 and the ribosomal protein RPL13A, respectively (i.e. Swc5-FRB and RPL13A-FKBP12), thus depleting Swc5 from the nucleus along with pre-ribosomal subunit export (Haruki et al., 2008). The ‘wild type’ AA yeast strain, mutated for TOR1 and the FK506-binding protein, is physiologically resistant to rapamycin and displays normal growth phenotype and normal H2A.Z and Pol II distributions (Tramantano et al., 2016; Wong et al., 2014).

Upon Swc5 AA, we found the expected decrease of chromatin-bound H2A.Z from 79% to 49% (Figure 2A–C and rapamycin time course in Figure 2—figure supplement 2), consistent with ChIP-seq results showing genome-wide reduction of H2A.Z at +1 nucleosomes under similar conditions (Tramantano et al., 2016). The remaining chromatin-bound H2A.Z may be due to residual H2A.Z at the +1 nucleosome or to histone chaperone-mediated H2A.Z deposition in nucleosomes over the entire genome, as suggested by in vivo cross-linking studies (Mohan et al., 2018; Tramantano et al., 2016). Our live-cell findings are thus consistent with the SWR1 requirement for promoter-specific H2A.Z deposition.

Figure 2 with 2 supplements see all
H2A.Z chromatin binding is substantially reduced upon abrogation of the deposition pathway by SWR1 inactivation.

(A) Time course of H2A.Z-Halo labeling, rapamycin treatment and image acquisition in Swc5-AA cells. Rapamycin treatment for an hour before SPT, and imaging performed in continued presence of rapamycin. (B, C) Normalized histograms and two-component Gaussian fits for H2A.Z-Halo imaged in the Swc5-AA cells. Imaging data were acquired in absence of rapamycin (B) or presence of rapamycin (C). Spot-On results show that Swc5 depletion causes a reduction in chromatin-bound H2A.Z. (E) Overlay of tracks, color-coded according to log diffusion coefficients, obtained from representative nuclei. Number of tracks (n) is indicated for each nucleus. All molecules tracked with JF552 dye.

Role of RNA Pol II in H2A.Z eviction

To identify H2A.Z eviction factors, we tested candidates that could mitigate the loss of chromatin-bound H2A.Z when both deposition and eviction factors were co-depleted in a double AA experiment. As the transcription PIC is constitutively enriched at the majority of NDRs (Rhee and Pugh, 2012) and has been causally linked to H2A.Z eviction (Tramantano et al., 2016), we first imaged the distribution of H2A.Z after nuclear depletion of both Swc5 and the Rpb1 catalytic subunit of Pol II. When Swc5 and Rpb1 are co-depleted by double AA, the chromatin-bound H2A.Z fraction increased (66%) relative to Swc5 AA alone (49%) (compare Figure 3A to Figure 2C and Figure 3A to Figure 2D). A complete restoration to the wild type level of H2A.Z is not anticipated because co-depletion of Swc5 also reduces H2A.Z deposition by SWR1 during anchor away. Fluorescence microscopy confirmed relocation of Swc5 to the cytoplasm in double AA cells, excluding inefficient nuclear depletion as a caveat (Figure 3—figure supplement 1C,D). These results indicate that Pol II indeed plays a major role in H2A.Z eviction. (Single AA of Rpb1 in rapamycin-treated cells showed a marginal increase from 84% to 87% of the bulk chromatin-bound H2A.Z over the untreated control [Figure 3—figure supplement 2A–C]).

Figure 3 with 2 supplements see all
RNA polymerase II is critical for H2A.Z eviction.

(A) Normalized histograms and two-component Gaussian fits for H2A.Z-Halo imaged in cells co-depleted for Rpb1 and Swc5. (B) H2A.Z-Halo distributions in cells co-depleted for Ino80 and Swc5. (C) Spot-On results showing co-depletion of Rpb1 along with Swc5 inhibits H2A.Z eviction. All molecules tracked with JF552 dye.

INO80 is not required for eviction of H2A.Z

To examine the role of the INO80 remodeler in H2A.Z eviction, we analyzed the H2A.Z distribution for Swc5 and Ino80 co-depletion by double AA and found no rise in bound H2A.Z compared to the single AA of Swc5 (compare Figure 3A to Figure 2C and Figure 3C to Figure 2D). (We observed no change in chromatin-bound H2A.Z for single AA of Ino80 (Figure 3—figure supplement 2D–F)). Taken together, we conclude that Pol II, but not the INO80 remodeler, has a major role in H2A.Z eviction. Contributions by other factors such as the ANP32E histone chaperone found in mammalian cells are not excluded (Mao et al., 2014; Obri et al., 2014).

Kin28/CDK7 affects H2A.Z eviction

Transcription by Pol II is a complex process involving PIC assembly, Pol II initiation, promoter escape, productive elongation and termination (Jonkers and Lis, 2015; Sainsbury et al., 2015). Given that site-specific phosphorylation of the Rpb1 subunit of Pol II regulates the progression of transcription, targeted depletion of transcriptional kinases provides an opportunity to identify the step involved in H2A.Z eviction. A key post-initiation step involves Serine five phosphorylation (Ser5-P) of heptapeptide repeats on the C-terminal domain (CTD) of Pol II (Rpb1) (Corden, 2013; Harlen and Churchman, 2017). Ser5-P is catalyzed by the yeast Kin28/Cdk7 kinase, a component of the kinase module (Kin28-Ccl1-Tfb3) of TFIIH, and is linked to capping of nascent RNA, Pol II release from the Mediator complex, promoter escape and early elongation. Recently, the Bur1/Cdk9 kinase was shown to phosphorylate the Rpb1 linker just upstream of CTD, at residues Thr 1471 and Ser 1493 (Chun et al., 2019), facilitating Pol II transition from early elongation to productive elongation. Furthermore, the Ctk1/Cdk12 kinase mediates Ser2 phosphorylation of the CTD associated with productive elongation through protein-coding regions (Corden, 2013; Harlen and Churchman, 2017; Wong et al., 2014). To investigate which phosphorylated state of Pol II is linked to H2A.Z eviction, we examined H2A.Z distributions in double AA cells conditionally deficient for Swc5 in combination with each of the three CTD kinases. Only Kin28 is required for H2A.Z eviction, as indicated by 65% chromatin-bound H2A.Z in the Kin28 and Swc5 double AA relative to 49% in the single AA of Swc5 (compare Figure 4B to Figure 2C and Figure 4F to Figure 2D). Consistent with its role in H2A.Z eviction, depletion of Kin28 alone showed an increase in chromatin-bound H2A.Z, though marginal (from 78% to 82%, Figure 4—figure supplement 1A–C). In contrast, double AA of Swc5 and the Bur1 kinase did not inhibit loss of chromatin-bound H2A.Z, nor did double AA of Swc5 and the Ctk1 kinase (Figure 4C, D and F).

Figure 4 with 1 supplement see all
Kin28 phosphorylation of RNA polymerase II CTD is critical for H2A.Z eviction.

(A) Schematic representation shows the three Pol II kinases Kin28, Bur1 and Ctk1 recruited at initiation, early-elongation and elongation phases respectively of Pol II and corresponding phosphorylation of indicated Rpb1 CTD sites. Set1 is the first of the three RNA capping enzymes; it removes γ-phosphate from the RNA 5’end to generate 5’ diphosphate. (B, C, D, E) Normalized histograms and two-component Gaussian fits for H2A.Z-Halo imaged in cells co-depleted for Swc5 along with Kin28 (B), Bur1 (C), Ctk1 (D) and Cet1 (E). (F) Spot-On results show Kin28 is required to evict H2A.Z. All molecules tracked with JF552 dye.

H2A.Z removal does not require RNA capping

In the wake of Pol II initiation, nascent RNA is co-transcriptionally capped by the sequential activity of three enzymes—Cet1, Ceg1 and Abd1—and is completed when RNA reaches ~100 nt (Lidschreiber et al., 2013). Capping of the 5’ end of nascent RNA is initiated by the Cet1-Ceg1 complex, which recognizes the 5’ triphosphate on the RNA and Ser5-P on the Pol II CTD (Martinez-Rucobo et al., 2015). To examine whether RNA capping or associated activities are required for H2A.Z eviction, we performed double AA of Swc5 and Cet1, and found no increase in chromatin-bound H2A.Z compared to single AA of Swc5 (compare Figure 4E to Figure 2C and Figure 4F to Figure 2D). Thus, H2A.Z eviction is not dependent on RNA capping. Likewise, we found no increase of chromatin-bound H2A.Z upon double AA of Swc5 and Rrp6, the 3’−5’ exonuclease responsible for degradation of noncoding RNA (Figure 4—figure supplement 1E,F). Taken together, we conclude that an early stage of transcription elongation closely linked to Pol II CTD Ser5 phosphorylation by Kin28 is required for robust eviction of chromatin-bound H2A.Z.

Discussion

Transcription of most yeast genes is infrequent and nucleosome turnover along gene bodies is low, but the +1 nucleosome constitutively turns over at a 3-fold higher rate (Dion et al., 2007; Grimaldi et al., 2014; Yen et al., 2013). Similarly, H2A.Z is constitutively displaced from +1 nucleosomes for both active and rarely transcribed genes, on a timescale of <15 min (Tramantano et al., 2016). The live-cell SPT approach shows that Pol II rather than the INO80 sub-family of remodelers plays a key role in H2A.Z eviction. The INO80 sub-family of remodelers comprise of SWR1 and INO80 remodeling complexes, with similarities in the catalytic split-ATPase domain and four shared subunits (Rvb1, Rvb2, Act1 and Arp4). INO80 has been previously suggested to mediate the reversal of SWR1-catalyzed H2A.Z deposition (Papamichos-Chronakis et al., 2011). However, in addition to this study, there are several publications reporting that INO80 displays no detectable H2A.Z eviction from chromatin in either yeast or mammalian cells (Au-Yeung and Horvath, 2018; Tramantano et al., 2016; Jeronimo et al., 2015). Although evidently uninvolved in H2A.Z eviction, the INO80 complex is known to mediate nucleosome repositioning in vitro and in vivo (Kubik et al., 2019; Shen et al., 2003).

Our results confirm the previous finding that the Pol II PIC plays an important role in H2A.Z eviction. Furthermore, the observed dependence on Kin28/Cdk7 kinase for robust H2A.Z displacement suggests that PIC assembly per se, that is the recruitment of general transcription factors and Pol II to promoter, is not sufficient for H2A.Z eviction but Kin28-dependent phosphorylation of Ser5 of the CTD heptapeptide repeats is important (Figure 5, box). A role for Kin28 in Ssl2-facilitated TSS scanning by Pol II is not excluded for H2A.Z eviction, although it has been shown that depletion of Kin28 by AA does not alter TSS usage (Murakami et al., 2015).

Cycle of H2A.Z eviction and deposition.

RNA polymerase II assembled genome-wide in the PIC and Rpb1 CTD Ser5 phosphorylated by Kin28 constitutively transcribes short noncoding RNAs (with m7G cap) and evicts H2A.Z-H2B dimers from the +1 nucleosome prior to termination. H2A.Z eviction should also occur in the course of mRNA transcription. Additional factors may be necessary for displacement of H3-H4 tetramer. The directional arrow indicates the annotated transcription start site. The gap is filled by histone chaperone-mediated deposition of canonical histones to reform an H2A-containing +1 nucleosome, which is positioned by chromatin remodelers and sequence-specific transcription factors, maintaining the NDR. This recruits SWR1 which is activated upon recognition of H2A-nucleosome and H2A.Z-H2B dimer substrates to activate one or two rounds of H2A.Z deposition. See text for discussion.

Inhibition of H2A.Z eviction upon depletion of Kin28/Cdk7 kinase, but not Bur1/Cdk9 or Ctk1/Cdk12 kinases narrows the relevant state of Pol II to early elongation after promoter escape, but not to productive elongation. (We note that the PIC remains largely intact upon Kin28/Cdk7 depletion, as shown by accumulation of TFIID, Mediator and Pol II at gene promoters [Knoll et al., 2019; Wong et al., 2014]).The exclusive dependence on Kin28/Cdk7 is further underscored by no reduction of H2A.Z eviction on depletion of Cet1, the 5’ RNA capping enzyme, or depletion of Rrp6, the 3’−5’ exonuclease for noncoding RNA degradation. Thus, H2A.Z eviction is independent of RNA modifying and metabolizing activities just downstream of CTD Ser5 phosphorylation.

Transcriptional elongation by Pol II is known to cause displacement of nucleosomal histones in biochemical assays (Lorch et al., 1987), providing a mechanism for H2A.Z turnover at the +1 nucleosome in the process of transcription through protein-coding regions. We propose a similar mechanism for genes that do not engage in productive transcription of mRNA, but exhibit genome-wide, constitutive transcription of noncoding RNAs which are prematurely terminated by the Nrd1-Nab3-Sen1 pathway in budding yeast (Schaughency et al., 2014). The early elongation activity of Pol II would dislodge H2A.Z-H2B dimers from the histone octamer of the +1 nucleosome. Displacement of the more stably bound H3-H4 tetramer likely requires assistance from histone chaperones and/or other remodelers (Figure 5).

After displacement of core histones, reassembly of a canonical nucleosome on gapped chromatin should occur, mediated by the mass action of the predominating H2A-H2B histone pool and histone chaperones, nucleosome positions being reset by chromatin remodelers such as ISWI, RSC, and INO80 (Lai and Pugh, 2017 Figure 5). Maintenance of a NDR of sufficient length (>60 bp DNA) by remodelers and subsequent histone acetylation recruits SWR1 to canonical +1 nucleosomes, the essential substrate for SWR1 (Ranjan et al., 2013). Stimulation of the catalytic Swr1 ATPase by nucleosome and H2A.Z-H2B dimer substrates then triggers histone dimer exchange (Luk et al., 2010; Ranjan et al., 2013), completing the cycle of H2A.Z/H2A replacement (Figure 5).

We envision that H2A.Z eviction is coupled to transcription not only from protein-coding genes transcribed by Pol II but also ribosomal, 5S and tRNA genes transcribed by Pol I and Pol III. Because H2A.Z eviction is not correlated with mRNA transcription by Pol II (Tramantano et al., 2016), the constitutive global transcription of noncoding RNA by Pol II is additionally coupled to H2A.Z eviction. There is substantial evidence for low-level, heterogenous transcripts of several hundred nucleotides, initiating from multiple start-sites within yeast NDRs (Pelechano et al., 2013). For budding yeast, these noncoding RNA transcripts evidently result from Pol II initiation without substantial pausing (Booth et al., 2016). At metazoan promoters, turnover of H2A.Z enriched in +1 nucleosomes may be similarly coupled to transcription in the process of Pol II pausing and release (Tome et al., 2018). Likewise at metazoan enhancers, infrequent Pol II transcription of eRNAs (Tippens et al., 2018) could be responsible for eviction of H2A.Z, representing erasure of a permissive histone variant mark on the epigenome. Much remains to be learned about the functional significance of this process and its relationship to productive mRNA transcription, presenting an outstanding problem for future studies of chromatin dynamics in eukaryotic gene regulation.

Materials and methods

Yeast strains and plasmids

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The plasmid for HaloTag (Halo) fusions was generated by cloning HaloTag (Promega) in the pBluescript SK (-) vector followed by insertion of a KanMx cassette (Kanamycin) or NatMx cassette (Nourseothricin), following standard procedures (Gelbart et al., 2001). PCR amplification and standard yeast transformation methods were used for tagging the protein of interest at the C-terminus, with a serine-glycine (SG4)2 linker to HaloTag.

For Halo-H2B, plasmid HTA1-SNAP-HTB1 (pEL458, gift from Ed Luk) was modified to replace the SNAP coding sequence with HaloTag, with a four amino acid GA3 linker between HaloTag and the N-terminus of H2B. The plasmid expressing Halo-H2B was introduced in the FY406 strain (gift from Fred Winston) by the plasmid shuffle procedure (Hirschhorn et al., 1995). The endogenous H2B promoter drives expression of Halo-H2B as the sole gene copy in cells.

Free HaloTag was fused at the N-terminus to a bipartite SV40 NLS (KRTADGSEFESPKKKRKV, where two clusters of basic residues are underlined) (Hodel et al., 2006) and expressed from the pRS416 vector. Plasmid pAC1056 expressing BPSV40 NLS-GFP (gift from Anita Corbett) was modified for free Halo expression.

Strains and plasmids used for anchor-away studies were obtained from Euroscarf.

The transporter gene PDR5 was deleted in all strains for retention of HTL-dye conjugate in live yeast cells. Strain genotypes are listed in Supplementary file 1.

Flow cytometry analysis

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Cells were fixed by adding two volumes of 100% ethanol and stored for one hour at 4C. Cells were washed with 50 mM Tris-HCl (pH 7.5) buffer and digested with RNase (1 mg/ml) and RNase A (0.2 mg/ml) overnight at 37C on a rotator. Proteins were digested with Proteinase K (1 µg/µl) at 50C for 30 min. Cells were stained with 2 µM SYTOX (Tris buffer) at 4C for 4 hr and sonicated on Diagenode Biorupter 300 for 10 s at high setting. Cells were scanned on LSR II FACS instrument.

Cell culture and labeling

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Cells were grown and imaged in CSM media (Complete Supplement Mixture) supplemented with 40 mg/L adenine hemisulfate. The JF646-HaloTag ligand was synthesized as previously described (Grimm et al., 2015). The new JF552 dye has a higher signal to noise ratio and is more photostable than JF646. The JF552 dye is a modification of JF549, with similar brightness, but enhanced cell permeability that allows its use for SPT in yeast (Zheng et al., 2019). For in vivo labeling, early log phase cells (O.D600 0.2) were labeled with JF-HaloTag ligand (10 nM for JF646 and 20 nM for JF552) for two hours at 30 degrees in suspension culture. Cells were washed four times with CSM to remove free dye.

Prior to use, 0.17 mm coverslips (Ø 25 mm, Electron Microscopy Services) were flamed to remove punctuated surface auto-fluorescence and to suppress dye binding, and coated with Concanvalin A (2 mg/ml) for 30 min at room temperature, and air-dried for one hour. Coverslips were assembled in a Ø 35 mm Attofluor chamber (Invitrogen). A 1 ml cell suspension was immobilized for 10 min and live cells were imaged in CSM media at room temperature. For anchor away experiments, rapamycin (1 µg/ml) was added one hour prior to imaging, and cells were imaged in the presence of rapamycin. Two biological replicates were performed for each experimental condition.

Cell cycle synchronization

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Cells were synchronized in G1 by adding α factor for 2 hr (3 µg/ml at 0 min and additional 2 and 1 µg/ml at 60 and 90 min respectively). High autofluorescence did not allow SMT in presence of α factor, which was removed by replacing culture medium. Cells released from G1 at room temperature took 40 min to enter S phase. Both for Pre-S and S phase SMT, cells were stained and synchronized in suspension culture and immobilized right before SMT.

Wide-field single molecule imaging with epi-Illumination microscope

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Single-molecule imaging was conducted on a Zeiss Observer Z1 microscope with a Zeiss Plan-Apochromat 150X/1.35 glycerin-immersion objective. Cells of interest were identified under infrared illumination (750 nm, 10 nm FWHM) using a near IR-CCD camera (IDS UI-3370CP-NIR-GL) and Semrock 743 nm/25 nm FWHM filter. A 555 nm (Crystalaser) or 637 nm (Vortran) laser was used for dye excitation, typically at 100 mW total power (TTL pulsed). All laser beams were spectrally filtered and combined using a custom beam combiner (by J.W., details available upon request). A Semrock FF01-750/SP filter was included at the output to remove any residual near infrared emission from lasers. Combined laser beams were collimated into a 2m-long Qioptic fiber (kineFLEX-P-2-S-405.640–0.7-APC-P2) with output through a 12 mm EFL reflective collimator (Thorlabs). The resulting Ø6mm Gaussian beam was introduced into the back port of the microscope. The following cubes were utilized in the microscope turret to direct excitation light towards the sample and filter fluorescence: 1) for JF646 - 648 beamsplitter and 676/29 nm filter, 2) for JF552 - 561 beamsplitter and 612/69 nm filter. Images were acquired with a Hamamatsu C9100-13 back-illuminated EM-CCD camera through additional FF01-750/SP and NF03-405/488/561/635E quad-notch filters. The camera was operated at −80°C with a typical EM gain of 1200 and directly controlled by laser emission via the TTL signal.

Image acquisition

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Images were obtained using either 637 nm laser (JF646) or 555 nm laser (JF552), of excitation intensity ~1KW/cm2 and for each field of view ~7000 frames were captured. Single molecules were tracked using DiaTrack Version 3.05 software, with the following settings; remove blur 0.1, remove dim 70–100, maximum jump six pixels. Single molecule images were collected after pre-bleach of initial intense fluorescence (glow). While imaging with JF646, a 405 nm laser excitation (1–10 mW/cm2, TTL pulses 2–5 ms per frame) was triggered to maintain single fluorophore detection density. Immobilized cells in CSM media were imaged over a 90 min imaging session.

Analysis of single-molecule images

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Movies with two dimensional single molecule data were analyzed by DiaTrack Version 3.05 (Vallotton and Olivier, 2013), which determines the precise position of single molecules by Gaussian intensity fitting and assembles particle trajectories over multiple frames. In Diatrack remove blur was set to 0.1, remove dim set at 70 and max jump set at five pixels, where each pixel was 107 nm. Trajectory data exported from Diatrack was further analyzed by a custom computational package ‘Sojourner’ (by S.L.). The package is available on Github (https://rdrr.io/github/sheng-liu/sojourner/). The Mean Squared Displacement (MSD) was calculated for all trajectories six frames or longer. Diffusion coefficients for individual molecules were calculated by unconstrained linear fit (R2 >0.8) of the MSD values computed for time lags ranging from 2 dt to 5 dt, where dt = 10 ms is the time interval between frames, and slope of linear fit was divided by 4 (pre-factor for 2-dimensional Brownian motion) (Qian et al., 1991). The histogram of log converted diffusion coefficients was fitted with double gaussian function from the ‘mixtools’ package (Benaglia et al., 2009) to estimate the fraction of chromatin-bound molecules (mean range between 0.050–0.112 µm2 s−1). Standard error on the mean of each gaussian fit parameter was estimated using a bootstrap resampling approach (Efron, 1979).

The Spot-On analysis was performed on trajectories three frames or longer using the web-interface https://spoton.berkeley.edu/ (Hansen et al., 2018). The bound fractions and diffusion coefficients were extracted from the CDF of observed displacements over different time intervals. For Brownian motion in two dimensions, the probability that a particle starting from origin will be found within a circle of radius r at time interval ∆𝜏 is given as follows.

Pr,τ=r2Dτe-r24Dτ

where D is diffusion coefficient. In Spot-On, the cumulative displacement histograms were fitted with a 2-state model.

pr,τ=F1 r2(D1τ+σ2)e-r24(D1τ+σ2)+ZCORR(τ,Z,D2)F2r2(D1τ+σ2)e-r24(D1τ+σ2)

where F1 and F2 are bound and free fractions, σ is single molecule localization error, D1 and D2 are diffusion coefficients of bound and free fractions, and ZCORR is correction factor for fast molecules moving out of axial detection range (Hansen et al., 2018). The axial detection range for JF646 on our setup is 650 nm. The following settings were used on the Spot-On web interface: bin width 0.01, number of time points 6, jumps to consider 4, use entire trajectories-No, Max jump (µm) 1.2. For model fitting the following parameters were selected: Dbound (µm2/s) min 0.001 max 0.1, Dfree (µm2/s) min 0.15 max 5, Fbound min 0 max 1, Localization error (µm)- Fit from data-Yes min 0.01 max 0.1, dZ (µm) 0.65 for JF646 and dZ 0.6 for JF552, Use Z correction- Yes, Model Fit CDF, Iterations 3.

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Decision letter

  1. Geeta J Narlikar
    Reviewing Editor; University of California, San Francisco, United States
  2. Jessica K Tyler
    Senior Editor; Weill Cornell Medicine, United States
  3. Geeta J Narlikar
    Reviewer; University of California, San Francisco, United States
  4. John T Lis
    Reviewer; Cornell University, United States
  5. Xavier Darzacq
    Reviewer; University of California, Berkeley, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

Your work makes a significant addition to existing models of how histone variant deposition and Pol II transcription are linked.

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 work entitled "Live-cell single particle imaging reveals the role of RNA polymerase II in histone H2A.Z eviction" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: John T Lis (Reviewer #3).

As you will see from the individual reviews attached below all the reviewers found that the single-molecule experiments were carefully carried out and that the data is of high quality. However, in the discussion amongst the reviewers it was also agreed that the findings do not go sufficiently beyond the work in Tramantano et al., 2016. The reviewers felt that if additional experiments can be carried out to identify where in the transcription cycle H2A.Z is evicted, then the results would contribute both important and novel insights. These experiments as suggested by reviewers 2 and 3, could involve looking at RRP6 and XRN1 mutants and inhibitors of Cdk9. Such experiments would not only provide an important technologically distinct confirmation of previous conclusions from Tramantano et al., 2016 but provide sufficient new mechanistic insight to pass the bar for eLife publication.

As it is unclear how long it would take to acquire the necessary data, the reviewers concluded that the current manuscript should be rejected. We would however welcome a resubmission should additional data become available that addresses the reviewer concerns. Please feel free to communicate with us if you would like any clarification.

Reviewer #1:The histone variant H2AZ is often present near transcription start sites in yeast. How the steady state levels of this variant are maintained and the consequences of regulating H2AZ dynamics are areas of active study. While some in vitro studies have suggested that IN080 plays a role in evicting H2AZ, others have not. Further, previous work using ChIP-based methods (Tramantano et al., 2016) has indicated that rather than INO80, the Pol II transcription machinery plays a major role in evicting H2AZ histones from the +1 position at transcription start sites. Here the authors use live cell single particle tracking microscopy to directly study the effects of depleting different transcription regulators on H2AZ dynamics. Using the anchor-away (AA) method they find that H2AZ retention on chromatin (as assessed by the slower diffusing fraction) is decreased upon Swc5-AA and restored by simultaneous Rpb1-AA. In contrast, Ino80-AA does not substantially restore the H2AZ chromatin bound levels that are depleted by Swc5-AA. They further find that inhibition of Kin28 also restores a substantial portion chromatin bound H2AZ that is lost upon Swc5-AA. The authors interpret these results to suggest that H2AZ eviction is driven by the early elongating Pol II and therefore could reflect the consequences of nascent transcription of non-coding RNAs.

Overall the studies are technically of high quality and the experiments are carried out in a well-controlled manner. The main conclusion that H2AZ eviction depends on both Pol II and its Ser5 phosphorylated state would be of interest to the chromatin community. However, as is, the work does not go substantially beyond the insights obtained from Tramantano et al. At the same time, the hypothesis of non-coding RNA transcription driving H2AZ turnover is attractive and novel. If the authors could provide some test of this hypothesis, this would substantially raise the significance of the work.

Reviewer #2:

The manuscript described the role of Pol2 in H2A.Z dynamics by using single-molecule tracking. Authors measure H2A.Z bound fractions in various depletion conditions, concluding that H2A.Z eviction is dependent on Pol2-CTD serine-5p rather than INO80 complex. Although single-molecule tracking for H2A.Z in the various conditions is informative to the field, the biological significance seems to be a large repetition of the results in Tramantano, 2016, with different aspects of technique. It would be nice if authors could more clearly explain the novelty of their work and how it extends beyond Tramantano, 2016.

We think the Tramantano, 2016 paper is pretty tightly controlled and well-executed, even though its findings are largely correlative. Their correlations tell them that both Pol II and PIC are important for H2A.Z turnover at the +1 nucleosome, but the mechanism is unknown and which part of the PIC is necessary is unclear. So the major contribution from this paper is that a defective TFIIH can restore H2A.Z fraction bound to near WT levels in a Swc5-depleted background. Going back to the paper from which the Wu lab got the Kin28is mutant puts a question mark on their proposed mechanism of H2A.Z turnover. They found that in Kin28is cells treated with CMK, Pol II accumulates at the +2 nucleosome. So the Wu hypothesis that it is passage of Pol II through the +1 nucleosome that displaces H2A.Z doesn't make sense. It would be nice to have the authors comment on this and help clarifying their point.

-The key conclusion in the manuscript is that Pol2-CTD-S5p associates with H2A.Z eviction based on the kin28is model. However, it is problematic to disrupt the TFIIH helicase activity for this purpose, in which this would have nothing to do with phosphorylation but shutdown entire transcription.

-Authors used the time course with dye staining before anchoring protein away. This resulted in the possibility that a portion of SPT measurements might be derived from the "WT" condition. Could authors also demonstrate a time course with dye staining after anchoring target away, or staining with different dyes before and after adding rapamycin?

-Authors did not have any data supporting the role of ncRNA here. Authors need to provide the evidence by measuring H2A.Z dynamics in RRP6 or XRN1 mutants.Reviewer #3:

This paper from Carl Wu's group evaluates the mechanisms in vivo that lead to rapid exchange dynamics of the H2A.Z histone variant, which is known to occupy nucleosomes adjacent to active or activatable promoters. The H2A.Z variant is highly conserved across species and is known to be a critical: mutants produce slow growth phenotypes in yeast and are lethal in higher organisms. Understanding H2Az is important to evaluate of mechanisms of interplay of chromatin and promoters during gene expression.

This paper begins by emphasizing that the conserved SWR1 chromatin remodeling complex is responsible for exchanging nucleosomal H2A-H2B for H2A.Z-H2B dimers onto the +1 nucleosome in budding yeast. The +1 nucleosome undergoes much higher turnover than other nucleosomes and the mechanism by which H2A.Z is evicted and replaced by H2A had not been resolved. The authors point out that H2A.Z eviction could be due to chromatin remodeling in reverse mediated by SWR1 itself or the related INO80 remodeler, but a study (Wang et al., 2016) found no supporting evidence for either model. Pol II activity itself is an attractive candidate for H2A.Z eviction, as Pol II transcription can disrupt nucleosome structure, but the amount of H2A.Z eviction from genes does not correlate with the level of mRNA accumulation.

To assess what is the dominant mechanism of H2A.Z turnover after incorporation, the authors used single particle tracking (SPT) to directly measure the levels of chromatin-free and bound H2A.Z in living yeast cells, in WT and conditional mutants (or in cells treated with a specific Cdk7 inhibitor) of candidates/processes that are hypothesized to be involved in H2A.Z eviction. "SPT measures the fast diffusing, chromatin-free population as well as the quasi-immobile, chromatin-bound fraction tracking with macroscopic chromosome movements". From these studies, the authors conclude that "H2A.Z eviction is dependent on RNA polymerase II (Pol II) bearing Serine-5 phosphorylation of carboxy-terminal repeats, linking H2A.Z eviction to transcription initiation, promoter escape and early elongation" in yeast, suggesting a

general mechanism by which noncoding transcription at promoters can lead to H2A.Z erasure.

The paper is clearly written, the experiments appear to be well performed, and the results are intelligently and clearly presented. A few concerns are cited below, but in my opinion this paper is suited for publication once the points are addressed or an additional experiment is performed using the authors established methods.

1) In Figure 1, the results of tracking >1000 molecules as 2D projections are presented as histograms. Unfortunately, the expected two populations, chromatin-bound and unbound, are not cleanly resolved. Nonetheless, the peak of diffusion coefficients is best fit computationally by a model that is comprised of two populations (chromatin-bound and unbound). This is not completely satisfying, but I do point out that the authors go on to makes a fairly compelling case that their modeling is correct. First, videos are provided to allow the reader to appreciate the high quality of the single-particle tracking. Second, the authors argue that the amount of H2AZ bound is consistent with previous estimates by in vivo crosslinking, and D values are consistent with previous SPT measurements of H2B. The authors could calculate the diffusion coefficients expected based on size of the free H2A.Z and SWR1 complex and perhaps estimate the values for bound forms based on binding constants. In any case, some additional discussion of this limitation of the resolution is warranted, and perhaps some acknowledgement that there could be additional states that are also not resolved. This should require only minor editing.

2) The amount of mRNA accumulation does not correlate H2A.Z eviction, so the authors evoke the idea that early elongation in terms of short non-coding RNAs at yeast promoters may be responsible for the eviction. Can the authors take existing estimates of this short transcription genome-wide and assess if it correlates with H2A.Z eviction?

3) Does the eviction of H2A.Z happen in the presence of inhibitors of transcription that act further downstream of Cdk7, for example of the yeast equivalent of Cdk9 (Bur1)? This could serve to pinpoint the eviction at very early steps in the transcription cycle. (These inhibitors may not stop Pol II elongation, but are likely to decrease elongation rates given recently published results in Pombe (Booth et al. 2018 PMID:29416031).

4) In the Discussion, the authors state "At metazoan enhancers and promoters, turnover of H2A.Z enriched in +1 nucleosomes may be similarly coupled to pervasive ncRNA transcription likely after release from sites of Pol II pausing (Tome et al., 2018)." However, there may not be a lot of ncRNA transcription in paused regions of metazoans, as many estimates of paused Pol II half-life are relatively long and there is not much actual ncRNA transcription that goes through the first nucleosome not destined for pre-mRNA production. Perhaps the presence of a nearby transcriptionally-engaged Pol II (promoter-proximal paused Pol II) is sufficient to lead to the destabilization of the adjacent nucleosome – especially considering there is some forward motion, backtracking and TFIIS RNA realignment cleavage taking place (Nechaev et al. 2010, PMID:20007866). Also, there are factors associated with Pol II at this stage that might somehow stimulate eviction activity.

https://doi.org/10.7554/eLife.55667.sa1

Author response

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

Reviewer #1:

[…] Overall the studies are technically of high quality and the experiments are carried out in a well-controlled manner. The main conclusion that H2AZ eviction depends on both Pol II and its Ser5 phosphorylated state would be of interest to the chromatin community. However, as is, the work does not go substantially beyond the insights obtained from Tramantano et al. At the same time, the hypothesis of non-coding RNA transcription driving H2AZ turnover is attractive and novel. If the authors could provide some test of this hypothesis, this would substantially raise the significance of the work.

We thank the reviewer for appreciating the application of direct single molecule imaging to study H2A.Z dynamics in living yeast cells. To raise the significance of the work and pass the bar for eLife publication as noted by the senior editor, we have performed additional experiments to identify where in the transcription cycle H2A.Z is evicted. Accordingly, besides Kin28/Cdk7 and Ctk1/Cdk12, we have tested whether H2A.Z eviction is inhibited by 3 additional enzymes acting at different transcription stages (Bur1/Cdk9 kinase for Pol II, Cet1 RNA capping enzyme, and Rrp6 the 3’-5’ exonuclease for noncoding RNA degradation). We have found that among the 5 candidates, only Kin28 affects removal of H2A.Z. Thus, H2A.Z eviction is independent of RNA modifying and metabolizing activities just downstream of CTD Ser5 phosphorylation. Our findings considerably narrow the relevant stage of the transcription cycle where histone eviction occurs to early Pol II elongation after promoter escape, but not to productive elongation. We believe that these additional experiments provide sufficient new mechanistic insight to the study. (Testing the hypothesis of non-coding RNA transcription driving H2AZ turnover is an important issue, but beyond the scope of the current work. As noted below in our response to reviewer #3, point #2, a bioinformatic approach is currently unfeasible for technical reasons).

Reviewer #2:

The manuscript described the role of Pol2 in H2A.Z dynamics by using single-molecule tracking. Authors measure H2A.Z bound fractions in various depletion conditions, concluding that H2A.Z eviction is dependent on Pol2-CTD serine-5p rather than INO80 complex. Although single-molecule tracking for H2A.Z in the various conditions is informative to the field, the biological significance seems to be a large repetition of the results in Tramantano, 2016, with different aspects of technique. It would be nice if authors could more clearly explain the novelty of their work and how it extends beyond Tramantano, 2016.

We concur and new experiments in the revised manuscript addresses the reviewer’s point. Copied below is our response to the same concern from reviewer #1.

To raise the significance of the work and pass the bar for eLife publication as noted by the senior editor, we have performed additional experiments to identify where in the transcription cycle H2A.Z is evicted. Accordingly, besides Kin28/Cdk7 and Ctk1/Cdk12, we have tested whether H2A.Z eviction is inhibited by 3 additional enzymes acting at different transcription stages (Bur1/Cdk9 kinase for Pol II, Cet1 RNA capping enzyme, and Rrp6 the 3’-5’ exonuclease for noncoding RNA degradation). We have found that among the 5 candidates, only Kin28 affects removal of H2A.Z. Thus, H2A.Z eviction is independent of RNA modifying and metabolizing activities just downstream of CTD Ser5 phosphorylation. Our findings considerably narrow the relevant stage of the transcription cycle where histone eviction occurs to early Pol II elongation after promoter escape, but not to productive elongation. We believe that these additional experiments provide sufficient new mechanistic insight to the study.

We think the Tramantano, 2016 paper is pretty tightly controlled and well-executed, even though its findings are largely correlative. Their correlations tell them that both Pol II and PIC are important for H2A.Z turnover at the +1 nucleosome, but the mechanism is unknown and which part of the PIC is necessary is unclear. So the major contribution from this paper is that a defective TFIIH can restore H2A.Z fraction bound to near WT levels in a Swc5-depleted background. Going back to the paper from which the Wu lab got the Kin28is mutant puts a question mark on their proposed mechanism of H2A.Z turnover. They found that in Kin28is cells treated with CMK, Pol II accumulates at the +2 nucleosome. So the Wu hypothesis that it is passage of Pol II through the +1 nucleosome that displaces H2A.Z doesn't make sense. It would be nice to have the authors comment on this and help clarifying their point.

We appreciate the reviewer’s comment on interpretation of the effects of the Kin28is mutant on Pol II. We also wrestled with the published model for the Kin28is mutant, and recognized that the presented model highlighting Pol II accumulation between +1 and +2 nucleosomes after CMK inhibition of Kin28 kinase does not entirely reflect the actual data in the publication, which shows Pol II accumulation in a broader region at the promoter from approx. -150 to + 170 (Figure 5A, C) (Rodriguez-Molina et al., 2016)). Irrespective, we have performed a further control analyzing effects of CMK-treated Kin28is without Swc5-AA. This revealed that Kin28is actually affects H2A.Z deposition (reason unclear), thus limiting the utility of this mutant for testing H2A.Z eviction. Hence, we used an alternative Kin28-AA mutant in a double Kin28-AA; Swc5-AA experiment to unveil the eviction pathway in the new experiment shown in Figure 4. (Importantly, the single Kin28-AA depletion did not adversely affect H2A.Z deposition (Figure 4—figure supplement 1A-C). We have removed the original data, replacing it with new findings using the Kin28-AA strain (Figure 4).

-The key conclusion in the manuscript is that Pol2-CTD-S5p associates with H2A.Z eviction based on the kin28is model. However, it is problematic to disrupt the TFIIH helicase activity for this purpose, in which this would have nothing to do with phosphorylation but shutdown entire transcription.

We thank the reviewer for drawing our attention to the possibility that Kin28 inactivation or now, depletion, could perturb the helicase activity of TFIIH. Accordingly, the following sentence is added in the revised Discussion: “A role for Kin28 in Ssl2-facilitated TSS scanning by Pol II is not excluded for H2A.Z eviction, although it has been shown that depletion of Kin28 by AA does not alter TSS usage (Murakami et al., 2015).”

-Authors used the time course with dye staining before anchoring protein away. This resulted in the possibility that a portion of SPT measurements might be derived from the "WT" condition. Could authors also demonstrate a time course with dye staining after anchoring target away, or staining with different dyes before and after adding rapamycin?

All SPT data was acquired after one hour of cell growth in presence of rapamycin. This is the standard AA protocol in the field under which maximum depletion is achieved. To be explicit, we have inserted this sentence in the Figure 2 legend: ‘Rapamycin treatment for an hour before SPT, and imaging was performed in continued presence of rapamycin’.

-Authors did not have any data supporting the role of ncRNA here. Authors need to provide the evidence by measuring H2A.Z dynamics in RRP6 or XRN1 mutants.

As stated in the response above, we have performed the requested experiment and present new data examining role of Rrp6 on H2A.Z eviction (Figure 4—figure supplement 1).

Reviewer #3:

[…] The paper is clearly written, the experiments appear to be well performed, and the results are intelligently and clearly presented. A few concerns are cited below, but in my opinion this paper is suited for publication once the points are addressed or an additional experiment is performed using the authors established methods.

1) In Figure 1, the results of tracking >1000 molecules as 2D projections are presented as histograms. Unfortunately, the expected two populations, chromatin-bound and unbound, are not cleanly resolved. Nonetheless, the peak of diffusion coefficients is best fit computationally by a model that is comprised of two populations (chromatin-bound and unbound). This is not completely satisfying, but I do point out that the authors go on to makes a fairly compelling case that their modeling is correct. First, videos are provided to allow the reader to appreciate the high quality of the single-particle tracking. Second, the authors argue that the amount of H2AZ bound is consistent with previous estimates by in vivo crosslinking, and D values are consistent with previous SPT measurements of H2B. The authors could calculate the diffusion coefficients expected based on size of the free H2A.Z and SWR1 complex and perhaps estimate the values for bound forms based on binding constants. In any case, some additional discussion of this limitation of the resolution is warranted, and perhaps some acknowledgement that there could be additional states that are also not resolved. This should require only minor editing.

We thank the reviewer for his thoughtful comments, and wish to point out that the quality of population histograms are in line with publications in the field. We have revised our text to emphasize that we have chosen the simplest model to fit data and do not exclude the presence of additional minor populations with distinct diffusive values. Regarding the values for free diffusion, because of constraints of motion blurring and focal depth and nuclear membrane confinements, it is challenging to precisely measure the D for free molecules. Nonetheless, it is useful to note that the average D for free H2A.Z (chaperoned H2A.Z-H2B dimer, 1.17 μm2s-1) and D for free Swr1 (Swr1 complex, 0.62 μm2s-1) reflects their different molecular sizes.

2) The amount of mRNA accumulation does not correlate H2A.Z eviction, so the authors evoke the idea that early elongation in terms of short non-coding RNAs at yeast promoters may be responsible for the eviction. Can the authors take existing estimates of this short transcription genome-wide and assess if it correlates with H2A.Z eviction?

The eviction rate of H2A.Z at yeast promoters can be estimated, in theory, based on the depletion rate of H2A.Z upon Swc5 depletion using the ChIP-seq data from Tramantano et al., 2016. However, since different promoters have different initial steady-state levels of H2A.Z prior to Swc5 perturbation, the slope of H2A.Z depletion is influenced by prior H2A.Z deposition, preventing a direct measurement of eviction. To our knowledge, there is currently no good genome-wide dataset for the rates of H2A.Z eviction. Therefore, at this time, we cannot confidently compare noncoding transcription with H2A.Z eviction.

3) Does the eviction of H2A.Z happen in the presence of inhibitors of transcription that act further downstream of Cdk7, for example of the yeast equivalent of Cdk9 (Bur1)? This could serve to pinpoint the eviction at very early steps in the transcription cycle. (These inhibitors may not stop Pol II elongation, but are likely to decrease elongation rates given recently published results in Pombe (Booth et al. 2018 PMID:29416031).

We have imaged H2A.Z in cells with double AA of Bur1/Cdk9 along with Swc5. We find, that unlike Kin28, depletion of Bur1 does not inhibit H2A.Z eviction (Figure 4). As noted above, we also examined whether H2A.Z eviction is inhibited by Cet1 RNA capping enzyme, and Rrp6 the 3’-5’ exonuclease for noncoding RNA degradation, and found them lacking effects on H2A.Z eviction. Thus, H2A.Z eviction is independent of RNA modifiying and metabolizing activities just downstream of CTD Ser5 phosphorylation.

4) In the Discussion, the authors state "At metazoan enhancers and promoters, turnover of H2A.Z enriched in +1 nucleosomes may be similarly coupled to pervasive ncRNA transcription likely after release from sites of Pol II pausing (Tome et al., 2018)." However, there may not be a lot of ncRNA transcription in paused regions of metazoans, as many estimates of paused Pol II half-life are relatively long and there is not much actual ncRNA transcription that goes through the first nucleosome not destined for pre-mRNA production. Perhaps the presence of a nearby transcriptionally-engaged Pol II (promoter-proximal paused Pol II) is sufficient to lead to the destabilization of the adjacent nucleosome – especially considering there is some forward motion, backtracking and TFIIS RNA realignment cleavage taking place (Nechaev et al. 2010, PMID:20007866). Also, there are factors associated with Pol II at this stage that might somehow stimulate eviction activity.

We take note of reviewer’s comment and have rephrased our Discussion to accommodate potential effects of paused Pol II on H2A.Z eviction. The revised sentence reads “At metazoan promoters, turnover of H2A.Z enriched in +1 nucleosomes may be similarly coupled to transcription in the process of Pol II pausing and release (Tome et al., 2018). “

https://doi.org/10.7554/eLife.55667.sa2

Article and author information

Author details

  1. Anand Ranjan

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Conceptualization, Resources, Data curation, Software, Formal analysis, Supervision, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6071-6017
  2. Vu Q Nguyen

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Resources, Data curation, Formal analysis, Investigation, Visualization, Writing - review and editing
    Competing interests
    No competing interests declared
  3. Sheng Liu

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Software
    Competing interests
    No competing interests declared
  4. Jan Wisniewski

    Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, United States
    Present address
    Experimental Immunology Branch, National Cancer Institute, Bethesda, USA
    Contribution
    Conceptualization, Resources, Supervision, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  5. Jee Min Kim

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Data curation, Formal analysis
    Competing interests
    No competing interests declared
  6. Xiaona Tang

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Software
    Competing interests
    No competing interests declared
  7. Gaku Mizuguchi

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
  8. Ejlal Elalaoui

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Data curation
    Competing interests
    No competing interests declared
  9. Timothy J Nickels

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Data curation
    Competing interests
    No competing interests declared
  10. Vivian Jou

    Department of Biology, Johns Hopkins University, Baltimore, United States
    Contribution
    Data curation
    Competing interests
    No competing interests declared
  11. Brian P English

    Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4037-6294
  12. Qinsi Zheng

    Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
  13. Ed Luk

    Department of Biochemistry and Cell Biology, Stony Brook University, Stony Brook, United States
    Contribution
    Resources, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6619-2258
  14. Luke D Lavis

    Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
  15. Timothee Lionnet

    Institute of Systems Genetics, Langone Medical Center, New York University, New York, United States
    Contribution
    Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  16. Carl Wu

    1. Department of Biology, Johns Hopkins University, Baltimore, United States
    2. Department of Molecular Biology and Genetics, Johns Hopkins School of Medicine, Baltimore, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    wuc@jhu.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6933-5763

Funding

National Institutes of Health (GM125831)

  • Carl Wu

National Institutes of Health (GM127538)

  • Timothee Lionnet

National Institutes of Health (GM104111)

  • Ed Luk

HHMI (Transcription Imaging Consortium, Janelia Research Campus)

  • Luke D Lavis
  • Timothee Lionnet
  • Carl Wu

Damon Runyon Cancer Research Foundation

  • Vu Q Nguyen

Johns Hopkins University (Bloomberg Distinguished Professorship)

  • Carl Wu

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

Acknowledgements

This work is dedicated to the memory of Maxime Dahan, former project leader of the HHMI-Janelia Transcription Imaging Consortium. We thank Anita Corbett for reagents, Zhe Liu, Brian Mehl, Aseem Ansari and Herve Rouault for discussions, Felix Wu for image processing, Anders Hansen, Maxime Woringer, and Xavier Darzacq for consultation on the Spot-On program, Prashant Mishra and Munira Basrai for assistance with FACS analysis, and James Brandt and Yumi Kim for deconvolution microscopy, Debbie Wei for making yeast strains, Sun Jay Yoo and Taibo Li for computational assistance. The study was supported by HHMI-Janelia Transcription Imaging Consortium funding to CW, TL, and LL, the Damon Runyon Cancer Research Foundation (V.N.), the Johns Hopkins Bloomberg Distinguished Professorship (CW), a grant to EL from the National Institutes of Health (GM104111), a grant to TL from National Institutes of Health (GM127538), and a grant to CW from the National Institutes of Health (GM125831).

Senior Editor

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

Reviewing Editor

  1. Geeta J Narlikar, University of California, San Francisco, United States

Reviewers

  1. Geeta J Narlikar, University of California, San Francisco, United States
  2. John T Lis, Cornell University, United States
  3. Xavier Darzacq, University of California, Berkeley, United States

Publication history

  1. Received: January 31, 2020
  2. Accepted: April 24, 2020
  3. Accepted Manuscript published: April 27, 2020 (version 1)
  4. Version of Record published: May 29, 2020 (version 2)

Copyright

This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.

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