Evidence for the nuclear import of histones H3.1 and H4 as monomers
2018; Springer Nature; Volume: 37; Issue: 19 Linguagem: Inglês
10.15252/embj.201798714
ISSN1460-2075
AutoresMichael James Apta‐Smith, Juan R. Hernández‐Fernaud, Andrew Bowman,
Tópico(s)Radiation Therapy and Dosimetry
ResumoArticle3 September 2018Open Access Source DataTransparent process Evidence for the nuclear import of histones H3.1 and H4 as monomers Michael James Apta-Smith Michael James Apta-Smith Division of Biomedical Sciences, Warwick Medical School, University of Warwick, Coventry, UK Search for more papers by this author Juan Ramon Hernandez-Fernaud Juan Ramon Hernandez-Fernaud Proteomics Research Technology Platform, School of Life Sciences, University of Warwick, Coventry, UK Search for more papers by this author Andrew James Bowman Corresponding Author Andrew James Bowman [email protected] orcid.org/0000-0002-5507-669X Division of Biomedical Sciences, Warwick Medical School, University of Warwick, Coventry, UK Search for more papers by this author Michael James Apta-Smith Michael James Apta-Smith Division of Biomedical Sciences, Warwick Medical School, University of Warwick, Coventry, UK Search for more papers by this author Juan Ramon Hernandez-Fernaud Juan Ramon Hernandez-Fernaud Proteomics Research Technology Platform, School of Life Sciences, University of Warwick, Coventry, UK Search for more papers by this author Andrew James Bowman Corresponding Author Andrew James Bowman [email protected] orcid.org/0000-0002-5507-669X Division of Biomedical Sciences, Warwick Medical School, University of Warwick, Coventry, UK Search for more papers by this author Author Information Michael James Apta-Smith1, Juan Ramon Hernandez-Fernaud2 and Andrew James Bowman *,1 1Division of Biomedical Sciences, Warwick Medical School, University of Warwick, Coventry, UK 2Proteomics Research Technology Platform, School of Life Sciences, University of Warwick, Coventry, UK *Corresponding author. Tel: +44 2476 150220; E-mail: [email protected] The EMBO Journal (2018)37:e98714https://doi.org/10.15252/embj.201798714 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Newly synthesised histones are thought to dimerise in the cytosol and undergo nuclear import in complex with histone chaperones. Here, we provide evidence that human H3.1 and H4 are imported into the nucleus as monomers. Using a tether-and-release system to study the import dynamics of newly synthesised histones, we find that cytosolic H3.1 and H4 can be maintained as stable monomeric units. Cytosolically tethered histones are bound to importin-alpha proteins (predominantly IPO4), but not to histone-specific chaperones NASP, ASF1a, RbAp46 (RBBP7) or HAT1, which reside in the nucleus in interphase cells. Release of monomeric histones from their cytosolic tether results in rapid nuclear translocation, IPO4 dissociation and incorporation into chromatin at sites of replication. Quantitative analysis of histones bound to individual chaperones reveals an excess of H3 specifically associated with sNASP, suggesting that NASP maintains a soluble, monomeric pool of H3 within the nucleus and may act as a nuclear receptor for newly imported histone. In summary, we propose that histones H3 and H4 are rapidly imported as monomeric units, forming heterodimers in the nucleus rather than the cytosol. Synopsis A tether-and-release system engineered to study protein dynamics reveals that histones are kept in a monomeric state in the cytoplasm and only engage histone chaperones after nuclear import. The H3/H4 chaperones ASF1, NASP, HAT1 and RbAp46 are nuclear, but rapidly leak from the nucleus upon sub-cellular fractionation. Post-translational tethering of H3 and H4 in the cytosol show they exist as stable monomeric units. Release of cytosolically tethered histones results in their rapid nuclear import and incorporation into chromatin. Cytosolically tethered H3 and H4 interact stably with Importin-alpha proteins, but not with histone chaperones. sNASP co-purifies with a molar excess of H3 over H4, suggesting the presence of a stable monomeric H3 population within the nucleus. Introduction Each cell division requires the doubling of both DNA and histone content, with half of the histones being of parental origin and half being newly synthesised. Whilst much effort has gone into studying the dynamics of recycled parental histones (Prior et al, 1980; Jackson, 1987, 1990; Katan-Khaykovich & Struhl, 2011; Radman-Livaja et al, 2011; Alabert et al, 2015), less is known about the program for newly synthesised histone incorporation. As they form the stable core of the nucleosome and are the substrates for the majority of post-translational marks, histones H3 and H4 are often at the forefront of these investigations. Unlike recycled histones, newly synthesised histones H3 and H4 must pass through the cytosol before they are incorporated into chromatin. Biochemical isolation of H3.1 (the replication-dependent H3 variant) containing complexes suggests it folds with H4 soon after synthesis, interacting with a number of histone chaperones to form a cytosolic chaperoning network that coordinates nuclear import (Mosammaparast et al, 2002; Campos et al, 2010; Alvarez et al, 2011; Ask et al, 2012). Key cytosolic events in the proposed pathway include H3.1 and H4 forming a heterodimer in the cytosol and interacting with NASP, ASF1, HAT1 and RbAp46 (Mosammaparast et al, 2002; Campos et al, 2010; Alvarez et al, 2011), HAT1 modification of H4 K5 and K12 by acetylation (Alvarez et al, 2011; Parthun, 2011), modification of H3 K9 by methylation (Pinheiro et al, 2012; Rivera et al, 2015) and association with the importin-β protein IPO4 (Imp4b) (Mosammaparast et al, 2002; Blackwell et al, 2007; Campos et al, 2010; Ask et al, 2012; Keck & Pemberton, 2012; Gurard-Levin et al, 2014; Hammond et al, 2017). In addition, a number of importin-β proteins have been suggested to provide chaperoning roles for basic nuclear cargo including ribosomal proteins and linker histones (Jakel et al, 2002), but not, as yet, the core histones. Nuclear import and incorporation of histones into chromatin occurs very rapidly (Ruiz-Carrillo et al, 1975; Bonner et al, 1988), thus following such events in a pulse-chase manner remains challenging. Import rates most likely exceed the folding and maturation kinetics of fluorescent proteins, making the process difficult to study by FRAP, FLIP, photoactivation or their derivative techniques (Reits & Neefjes, 2001; Lukyanov et al, 2005; Ishikawa-Ankerhold et al, 2012). Similar difficulties arise with self-labelling domains such as the SNAP-tag, requiring minutes to hours for quenching, pulsing and labelling steps (Juillerat et al, 2003; Jansen et al, 2007; Crivat & Taraska, 2012; Clement et al, 2016). Metabolic incorporation of radioactive amino acids or functional amino acid derivatives (Dieterich et al, 2007; Deal et al, 2010; Lang & Chin, 2014) affords immediate labelling for biochemical analysis, but presents challenges for imaging proteins due to the requirement for derivatisation of the incorporated functional groups (Lang & Chin, 2014), and are subject to artefacts arising from biochemical purification processes. Thus, many of the current ideas about the nucleo-cytoplasmic chaperoning of histones remain to be tested in a cellular setting. In an attempt to address this, and potentially gain new information regarding the histone import and deposition pathway, we have developed an approach termed RAPID-release (rapamycin activated protease through induced dimerisation and release of tethered cargo) that allows observation of dynamic cellular events in real time in living cells. In this approach, we circumvent the requirement for immediate labelling of newly synthesised histones by first capturing them on the cytosolic face of the outer mitochondrial membrane (OMM). The quiescent histones are then released by concomitant recruitment and activation of a site-specific, viral protease through the addition of the small molecule rapamycin (Stein & Alexandrov, 2014). The histones can be followed by fusion of a fluorescent protein, allowing visualisation of nuclear import and incorporation at replication domains in real time. We apply this approach to investigate the early maturation and import of H3.1 and H4, corroborating our findings through quantitative analysis of histone stoichiometries bound to core chaperoning components. Results Mislocalisation of histone chaperones during fractionation necessitates an in vivo pulse-labelling system During analyses of histone chaperone localisation, we observed a striking discrepancy between biochemical fractionation and immunofluorescence in the sub-cellular localisation of a number of histone chaperones. Whilst s/tNASP, HAT1, RbAp46 and ASF1A appear overwhelmingly cytosolic using a standard NP-40 lysis protocol (Suzuki et al, 2010), with CAF1p60 almost equally split, they appear entirely nuclear when probed by immunofluorescence (compare Fig 1A with B). Pre-blocking of the antibodies with their immunogens specifically reduced the nuclear fluorescence, suggesting the antibodies are specific and the discrepancy is not due to cross-reactivity (Fig 1C, example images in Fig EV1A). Furthermore, the chaperones did not change their localisation between S and G1/G2 phases, as determined by the presence or absence of PCNA foci in the nucleus (Fig EV1B), suggesting the discrepancy cannot be due to cell cycle effects. Figure 1. Sub-cellular localisation of core H3.1 and H4 chaperoning components Fractionation of nuclear and cytosolic compartments using a standard NP-40 lysis protocol and immunoblotting with chaperone specific antibodies. Immunofluorescence of histone chaperones in fixed cells using the same antibodies used in (A). Cells are segregated into those undergoing replication and those that are not. Scale bar represents 5 μm. Quantification of nuclear localisation shown in (B). Boxes represent the lower quartile, median and upper quartile. Whiskers represent 1.5 times the interquartile range. Real-time imaging of cells undergoing biochemical fractionation. sNASP is tagged with mCherry, whilst Lamin A/C is tagged with EGFP. The left column of panels represents a maximum intensity projection. The time course on the right represents a plane in the z dimension reconstructed from 20 z-stacks penetrating 20 μm into the medium. Scale bars represent 5 μm. Quantification of nuclear leakage shown in (D). Z-stacks were flattened using a maximum intensity projection with the nuclear fluorescent signal over time plotted as a percentage of maximum (normalised). Data points represent the mean of 15 measurements with error bars representing the SEM. Source data are available online for this figure. Source Data for Figure 1 [embj201798714-sup-0007-SDataFig1.pdf] Download figure Download PowerPoint Click here to expand this figure. Figure EV1. Related to Fig 1 Immunofluorescence using antibodies pre-blocked with their immunogen. In each case, the specific nuclear fluorescence was decreased to background levels. Scale bar represents 5 μm. Cellular location of transiently expressed EGFP-histone chaperone fusions in replicating and non-replicating cells. Scale bars represent 5 μm. Time series of a cell expressing mCherry-sNASP and EGFP-Lamin A/C undergoing hypotonic lysis. Scale bar represents 5 μm. Download figure Download PowerPoint We reasoned one explanation for the discrepancy could be the leakage of nuclear components into the cytosolic fraction during biochemical isolation. To determine the behaviour of the soluble nuclear chaperones during fractionation, we took mCherry-sNASP as a representative nuclear component of the histone chaperoning pathway and co-expressed it in HeLa cells with EGFP-Lamin A/C to act as a marker of nuclear integrity. Addition of cytosolic extraction buffer (PBS + 0.1% NP-40) caused nuclei to puff-up (Fig 1D) but remain in close proximity to their pre-lysis position allowing us to follow mCherry-sNASP's location. Interestingly, whilst Lamin A/C remained at the nuclear periphery throughout the time course, sNASP rapidly diffused out of the nucleus (Fig 1D and E, Movie EV1). Similar behaviour was seen for hypotonic lysis in which cells were monitored whilst being exposed to water (Fig EV1C). It should be noted that previous studies using Xenopus oocytes have recorded a similar nuclear leakage with regard to total protein levels (Paine et al, 1983, 1992). In summary, we propose that the discrepancy between biochemical observations and in situ observations may be explained in terms of the rapid diffusion of soluble nuclear components during separation of cytoplasmic and nuclear compartments, and that the core histone chaperoning proteins NASP, ASF1A, RbAp46 and HAT1 reside predominantly in the nucleus throughout the cell cycle. The RAPID-release technique allows observation of nascent cytosolic histones and their translocation to the nucleus Taking into account the problem of nuclear leakage detailed above, we wondered whether it is possible to probe cytosolic histones in living cells using fluorescence microscopy. Whilst fluorescent microscopy has been used extensively to study histone turnover in chromatin, the kinetics with which histones are incorporated into chromatin after synthesis (Ruiz-Carrillo et al, 1975; Bonner et al, 1988) is likely an order of magnitude greater than that of fluorescent protein folding or SNAP/HALO-tag labelling, and thus a histone will be incorporated into chromatin before it is observed. To circumvent this, we pursued a cytosolic tether-and-release strategy termed RAPID-release (rapamycin activated protease through induced dimerisation and release of tethered cargo). In this approach, histones are tethered to the cytosolic face of the outer mitochondrial membrane (OMM) and are held in a quiescent state whilst the fluorescent fusion protein matures. Detethering is triggered by addition of rapamycin, which concomitantly recruits and activates an auto-inhibited TVMV protease (Stein & Alexandrov, 2014), thus allowing observation of nuclear import and chromatin deposition. To assess the feasibility of the approach, we fused EGFP to the FKBP12-rapamycin-binding (FRB) domain of mTOR, followed by the mitochondrial tail-anchoring sequence of OMP25 (Horie et al, 2002), with two TVMV cleavage sites separating the EGFP and FRB-OMP25 domains (EGFPTVMVx2-FRB-OMP25) (Fig 2A). The TVMV protease containing a C-terminal auto-inhibitory (AI) peptide and an N-terminal FK506-binding domain (FKBP12) (Stein & Alexandrov, 2014) was fused to the C-terminus of mCherry, creating the construct mCherry-FKBP12-TVMV-AI (Fig 2A). Addition of rapamycin to HeLa cells co-transfected with EGFPTVMVx2-FRB-OMP25/mCherry-FKBP12-TVMV-AI resulted in the recruitment of mCherry-FKBP12-TVMV-AI to mitochondria and release of the EGFP cargo from its tether (Fig 2B). Analysis of the cleavage (as the change in maximum pixel intensity of the cytoplasm) revealed a fit to an exponential decay model (Fig 2C). Plotting the rate constant of fit against the relative expression level of the protease (measured as the ratio of mCherry:EGFP signal) for each cell revealed a strong positive correlation, suggesting the cleavage rate is dependent on the level of protease (Fig 2D). At the highest ratios of protease to substrate, a half-maximal cleavage of 2.5 min was achieved. Removal of the two TVMV cleavage sites from EGFP inhibited cleavage, whilst removal of the AI peptide from the TVMV protease resulted in constitutive activity without recruitment (Fig EV2A and B), demonstrating the specificity of the protease and the importance of the AI peptide fusion, respectively. Figure 2. A tether-and-release approach for studying histone dynamics in living cells Schematic representation of the RAPID-release technique. Addition of rapamycin results in recruitment and activation of an auto-inhibited TVMV protease, leading to the release of EGFP-labelled histones from their mitochondrial tether. TVMV—tobacco vein mottling virus, FKBP—FK506-binding protein (FKBP12), FRB—FKBP12-rapamycin-binding domain, OMM—outer mitochondrial membrane, OMP25—25 kDa outer-membrane immunogenic protein (C-terminal helix), AI—auto-inhibitory peptide. A representative cell showing release of tethered EGFP. Scale bar represents 5 μm. Quantification of EGFP release. Single cells are represented as grey traces. The cell displayed in (B) is highlighted with red circles. The blue line represents a fit to an exponential decay model. Rate constants from exponential decay functions of the traces shown in (C) plotted against the expression level of protease relative to EGFP substrate. The blue line represents a linear regression model with the R2 value shown. Representative cells showing release of H4-EGFP and H3.1-EGFP. Scale bar represents 5 μm. Nuclear import rate and cleavage rate of H3.1-EGFP and H4-EGFP. Single cells are represented as grey traces. Traces from cells displayed in (E) are highlighted with circles. Import rate plotted against cleavage rate. Values represent those shown in (F), with the cells shown in (E) highlighted with red circles. Linear regression models are shown with blue lines. The mean R2 and slope (m) are shown for the 11 cells quantified in (F), +/− relates to standard deviation. Download figure Download PowerPoint Click here to expand this figure. Figure EV2. Related to Fig 2 Cleavage site deletion mutant. The auto-inhibited TVMV protease is recruited to the tethered EGFP, but is unable to release it due to the absence of a TVMV cleavage site separating EGFP from its tether. Scale bar represents 5 μm. Auto-inhibitory (AI) deletion mutant. Deletion of the AI domain from the mCherry-FKBP12-TVMV-AI construct results in constitutively active protease that cleaves the tethered EGFP independent of recruitment. Scale bar represents 5 μm. Histogram of a time course for EGFP or H4-EGFP release. Pixels from the whole cell (grey), the cytosol (blue) and the nucleus (red) plotted for the cells are shown in Fig 2E. Whilst the modal value from the whole cell significantly shifts to the right after the release of EGFP, with overlapping cytosol and nuclear signals, the modal value for H4-EGFP remains at a similar level, whilst the nuclear and cytosolic signals form discrete populations. Grey lines represent modal pixel values before rapamycin addition. Scale bar represents 5 μm. Download figure Download PowerPoint Next, we used the RAPID-release system to observe the dynamics of histone H3.1 and H4 upon release from the OMM. In this instance, tail anchoring (Suzuki et al, 2002), in contrast to N-terminal anchoring, permitted C-terminal tagging of histones, allowing us to avoid N-terminal fusions that have previously been shown to affect chromatin incorporation dynamics (Kimura & Cook, 2001). Transient transfection of H3.1 or H4 fused to the N-terminus of the EGFPTVMVx2-FRB-OMP25 construct did not observably affect mitochondrial or cellular morphology. However, a small amount of background nuclear fluorescence was observed (Fig 2E, Movies EV2 and EV3). Release of H3.1/H4-EGFP from the cytosolic tether resulted in rapid nuclear localisation (measured as the median nuclear fluorescence) (Fig 2E and F) at a rate mirrored by the kinetics of cleavage (measured as the S.D of the cytoplasm) (Fig 2G). Standard deviation of the cytoplasm was used instead of maximum pixel intensity as it was less affected by sub-cellular partitioning. Plotting the nuclear import rate against the cleavage rate resulted in a strong fit to a linear model (Fig 2G), revealing nuclear import of histones occurs at a rate greater than proteolytic cleavage and in excess of our sampling rate. Confirming this, the modal value of the partitioned cytoplasmic signal, representing the portion of the cytoplasm outside of the mitochondrial network, did not increase over the cleavage period (Fig EV2C) as it did for freely diffusing EGFP. In summary, the RAPID-release technique allows observation of histone nuclear import in living cells and provides a pulse-chase strategy with significantly improved kinetics compared to currently available techniques. Histones released from their tether incorporate rapidly at actively replicating domains In order to validate the tether-and-release approach in studying histone deposition, we tested cells for their ability to incorporate released histones into their chromatin. Mammalian genomes are organised into topological domains (TADs), which have been suggested to correlate with stable units of replication, or replication domains (RDs) (Pope et al, 2014; Rivera-Mulia & Gilbert, 2016). In a subset of asynchronously dividing cells, we observed foci forming in the nucleus soon after histone release (Fig 3A). To determine whether these foci represent histone incorporation at RDs, we co-transfected H3.1-EGFPTVMVx2-FRB-OMP25 and TagBFP-FKPB-TVMV-AI fusions with a PCNA-VHH-TagRFP chromobody to mark active replication (Burgess et al, 2012) (the TagBFP channel was used to identify expressing cells, but was not imaged to minimise bleaching of the other fluorophores). Fixing and imaging cells 30 min after rapamycin addition showed that nuclei positive for PCNA foci were also positive for H3.1-EGFP foci, whereas nuclei negative for PCNA foci were also negative for H3.1-EGFP foci (Fig 3B). Cells absent of PCNA foci, and by deduction not replicating their genomes, demonstrated a nucleolar enrichment of H3.1-EGFP, which has previously been shown to be an artefact of excess soluble histones in the nucleus (Musinova et al, 2011; Safina et al, 2017). Figure 3. Incorporation of released histones into chromatin at sites of active replication Accumulation of nuclear H3.1-EGFP and the formation of distinct foci. Scale bar represents 5 μm. Representative cells showing two different subpopulations of released H3.1-EGFP staining and their colocalisation with PCNA-positive foci. PCNA was detected with a PCNA-VHH-TagRFP chromobody. Scale bar represents 5 μm. Representative cells showing points from a time course of H3.1-EGFP and H4-EGFP release. Scale bar represents 5 μm. Colocalisation of PCNA and H3.1/H4-EGFP signals at various time points after release. A Wilcoxon rank sum test was carried out for each time point versus the 30-min time point, with those scoring a P-value of < 0.001 indicated by brackets. Released H3.1/H4-EGFP colocalise with mitotic chromosomes 48 h post-release. Scale bar represents 5 μm. Download figure Download PowerPoint To analyse the incorporation dynamics more quantitatively, we released histones at four time points prior to fixation and colocalisation analysis. Cells in mid/late S-phase were chosen, identified from their peripheral PCNA staining pattern (Burgess et al, 2012), as these cells would have been in S-phase as histones were released. Within 30 min, formation of foci occurred in the EGFP channel that colocalised to replication domains marked by the PCNA chromobody (Fig 3C and D). Colocalisation continued for 1 h, then decreased dramatically at 2.5 h, with a slight anti-correlation seen at 5 h post-release (Fig 3C and D). We interpret this as the pulse of released histones entering the soluble pool and being incorporated at actively replicating domains, up until the fluorescent pool of histones is depleted and replication moves on to neighbouring domains. To further verify that released histones are incorporated into chromatin, we imaged cells in mitosis 48 h post-release and found the H3.1-/H4-EGFP signal localised to the condensed, mitotic chromosomes, providing further evidence that released H3.1-/H4-EGFP are deposited into chromatin and stably retained through cell division (Fig 3E). Together, these experiments demonstrate that released H3.1-/H4-EGFP enter the histone chaperoning pathway and are incorporated into chromatin in a similar fashion to endogenous histones, and validate the RAPID-release system as a method for investigating chromatin assembly and the histone chaperoning pathway. Cytosolically tethered H3.1 and H4 are monomeric and do not detectibly associate with endogenous NASP, ASF1A, RbAp46 or HAT1 H3.1 and H4 exist as an obligate heterodimer in chromatin (Luger et al, 1997) and are also found as a dimer when bound to a number of histone chaperoning proteins, such as ASF1A/B, s/tNASP, RbAp46, HAT1 and the CAF1 complex (Tagami et al, 2004). However, the two histones are synthesised separately and must fold at a point prior to entry into the chromatin deposition pathway. We reasoned that if H3.1 and H4 fold in the cytoplasm, we would expect to see enrichment of the endogenous partner on the mitochondrial network. Similarly, endogenous histone chaperones that interact with H3.1 and H4 should also be enriched at the mitochondrial network (Fig 4A). To test this, we carried out immunofluorescence to probe for the co-occurrence of endogenous histone binding partners (Figs 4B and EV3A and B). Pearson's correlation coefficients of cytosolic regions encompassing the mitochondrial network were calculated for the EGFP and immunofluorescent channels (Figs 4C and EV3A and B). Interestingly, whilst we could detect the tethered histones, we could not detect enrichment of their orthologous binding partners, nor could we detect any of the histone chaperones (Fig 4C), most likely due to their nuclear localisation (Figs 1 and EV3B). Figure 4. Interaction profiling of tethered cytosolic histones Schematic representation of the fluorescence-2-hybrid approach for analysing interaction with endogenous proteins. Example of interaction screening using an α-H3 antibody against tethered EGFP (EGFP-OMP25), tethered H3.1 (H3.1-EGFP-OMP25) and tethered H4 (H4-EGFP-OMP25). Background-corrected, single Z-slices of representative cells are shown with 2D histograms displayed on the right. Nucleus and cytoplasm are portioned with a white line. Pearson's coefficients (R) for the cytosolic region of the depicted cells are shown in the histogram inset. Scale bar represents 10 μm. A boxplot of Pearson's coefficients between tethered EGFP, H3.1-EGFP or H4-EGFP and endogenous histone counterparts or known histone chaperones. P-values of < 0.001 from a Wilcoxon rank sum test versus the EGFP alone control are indicated by brackets. Whiskers extend to 1.5 times the IQR. Schematic representation of a fluorescence-2-hybrid assay using mCherry-tagged, forced cytosolic chaperone. Example of interaction screening as in (B), but using sNASP-dNLS against EGFP (EGFP-OMP25), tethered H3.1 (H3.1-EGFP-OMP25) and tethered H4 (H4-EGFP-OMP25). Scale bar represents 10 μm. A boxplot of Pearson's coefficients between tethered EGFP, H3.1-EGFP or H4-EGFP and cytosolically forced chaperones. P-values of < 0.001 from a Wilcoxon rank sum test versus the EGFP control are indicated by brackets. Whiskers extend to 1.5 times the IQR. Crystal structures of RbAp46-H4, HAT1-H4 and ASF1A-H3-H4 complexes. H3 is shown in red, and H4 is shown in blue. Download figure Download PowerPoint Click here to expand this figure. Figure EV3. Related to Fig 4 An example cell expressing tethered EGFP and probed with an anti-H3 antibody, demonstrating the image processing workflow. Raw images are shown on top, and background-corrected images shown below. The manually partitioned cytosolic region encompassing the mitochondrial network is outlined in red. A bisection of the cell is indicated in the images and the corresponding profiles of each channel shown in the third column. The Pearson's correlation coefficient of the raw and corrected images is shown, demonstrating the artifactually high colocalisation from the gradient of background staining in the raw image coinciding with the preference of mitochondria to reside around the microtubule organising centre of the cell, juxtaposed to the nucleus. Representative images for cells quantified are shown in Fig 4C. Scale bar represents 5 μm. Representative images from cells quantified are shown in Fig 4E. Scale bar represents 5 μm. Download figure Download PowerPoint To determine whether lack of binding was due to the nuclear partitioning of histone chaperones, rather than tethered histones adopting an unfavourable conformation that prevents binding, we expressed forced cytosolic chaperones that were mCherry-tagged. To achieve cytosolic localisation, chaperones were either mutated in their nuclear localisation sequence (ΔNLS), where a defined NLS existed (as for NASP; Kleinschmidt & Seiter, 1988; O'Rand et al, 1992), or engineered with a strong nuclear export signal (NES) (Henderson & Eleftheriou, 2000) where no known NLS was present (as for RbAp46, HAT1, ASF1A) (Fig 4D). This effectively drove the cytosolic location of all histone chaperones tested (Fig EV3C). The rationale behind the experiment was as follows: as RbAp46 and HAT1 bind to H4 epitopes within the H3.1-H4 heterodimer (Murzina et al, 2008; Song et al, 2008) (Fig 4G), if tethered histones were folded with their endogenous counterpart, we would expect to see the recruitment of RbAp46 and HAT1 to both tethered H3.1 and tethered H4, whereas if histones were monomeric, we would expect to see recruitment to tethered H4, but not H3.1. Conversely, as sNASP interacts directly with H3 as a monomer and as an H3.1-H4 heterodimer (Bowman et al, 2016, 2017), we would expect to see recruitment to both tethered H3.1 and H4 if a heterodimer was present, but only to tethered H3.1 if the histones were monomeric. As ASF1 contacts both H3 and H4 through independent binding sites (English et al, 2005; Natsume et al, 2007), we may expect to see recruitment to both independent of the histone's oligomeric status.
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