1. Chromosomes and Gene Expression
  2. Structural Biology and Molecular Biophysics
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Structural basis for PRC2 decoding of active histone methylation marks H3K36me2/3

  1. Ksenia Finogenova
  2. Jacques Bonnet
  3. Simon Poepsel
  4. Ingmar B Schäfer
  5. Katja Finkl
  6. Katharina Schmid
  7. Claudia Litz
  8. Mike Strauss
  9. Christian Benda
  10. Jürg Müller  Is a corresponding author
  1. Max Planck Institute of Biochemistry, Laboratory of Chromatin Biology, Germany
  2. California Institute for Quantitative Biology (QB3), University of California, California Institute for Quantitative Biology (QB3), Molecular Biophysics and Integrative Bio-Imaging Division, Lawrence Berkeley National Laboratory, United States
  3. University of Cologne, Center for Molecular Medicine Cologne (CMMC), Faculty of Medicine and University Hospital Cologne, Germany
  4. Cologne Excellence Cluster for Cellular Stress Responses in Ageing-Associated Diseases (CECAD), University of Cologne, Germany
  5. Max Planck Institute of Biochemistry, Department of Structural Cell Biology, Germany
  6. Max Planck Institute of Biochemistry, cryoEM Facility, Germany
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Cite this article as: eLife 2020;9:e61964 doi: 10.7554/eLife.61964

Abstract

Repression of genes by Polycomb requires that PRC2 modifies their chromatin by trimethylating lysine 27 on histone H3 (H3K27me3). At transcriptionally active genes, di- and tri-methylated H3K36 inhibit PRC2. Here, the cryo-EM structure of PRC2 on dinucleosomes reveals how binding of its catalytic subunit EZH2 to nucleosomal DNA orients the H3 N-terminus via an extended network of interactions to place H3K27 into the active site. Unmodified H3K36 occupies a critical position in the EZH2-DNA interface. Mutation of H3K36 to arginine or alanine inhibits H3K27 methylation by PRC2 on nucleosomes in vitro. Accordingly, Drosophila H3K36A and H3K36R mutants show reduced levels of H3K27me3 and defective Polycomb repression of HOX genes. The relay of interactions between EZH2, the nucleosomal DNA and the H3 N-terminus therefore creates the geometry that permits allosteric inhibition of PRC2 by methylated H3K36 in transcriptionally active chromatin.

Introduction

Many post-translational modifications on histone proteins are essential for processes in the underlying chromatin. Typically, histone modifications themselves do not alter chromatin structure directly but function by binding effector proteins which alter chromatin or by interfering with such interactions. The histone methyltransferase Polycomb Repressive Complex 2 (PRC2) and its regulation by accessory proteins and histone modifications represent a prime example for understanding these interaction mechanisms (Laugesen et al., 2019; Yu et al., 2019).

PRC2 trimethylates lysine 27 in histone H3 (H3K27me3), a modification that is essential for the transcriptional repression of developmental regulator genes that control cell fate decisions in metazoans (Pengelly et al., 2013; McKay et al., 2015). H3K27me3 marks chromatin for interaction with PRC1 (Fischle et al., 2003; Min et al., 2003), an effector which compacts chromatin (Francis et al., 2004; Grau et al., 2011). H3K27me3 is also recognized by PRC2 itself, and this interaction allosterically activates the PRC2 enzyme complex to facilitate deposition of H3K27me3 across extended domains of chromatin (Hansen et al., 2008; Margueron et al., 2009; Jiao and Liu, 2015).

Genetic studies and subsequent biochemical work established that PRC2 is in addition subject to negative regulation. In particular, the H3K4me3, H3K36me2, and H3K36me3 marks present on nucleosomes in transcriptionally active chromatin directly inhibit H3K27 methylation by PRC2 (Klymenko and Müller, 2004; Schmitges et al., 2011; Yuan et al., 2011; Gaydos et al., 2012; Streubel et al., 2018). Importantly, while stimulation of PRC2 activity by H3K27me3 acts in trans, inhibition of PRC2 by H3K4me3, H3K36me2, and H3K36me3 requires that these modifications are present in cis, that is, on the same H3 molecule containing the K27 substrate lysine (Schmitges et al., 2011; Yuan et al., 2011; Voigt et al., 2012). While recent structural studies have uncovered the allosteric activation mechanism for PRC2 (Jiao and Liu, 2015; Justin et al., 2016), the molecular basis of PRC2 inhibition by active chromatin marks has remained enigmatic. In particular, in nucleosome-binding assays, PRC2–DNA interactions make the largest contribution to the nucleosome-binding affinity of PRC2 (Wang et al., 2017; Choi et al., 2017) and H3K4me3, H3K36me2 and H3K36me3 do not seem to have a major effect on this binding affinity (Schmitges et al., 2011; Guidotti et al., 2019; Jani et al., 2019). Instead, these three modifications were found to reduce the kcat of PRC2 for H3K27 methylation (Schmitges et al., 2011; Jani et al., 2019). Recent cross-linking studies led to the suggestion of a possible sensing pocket for H3K36 on the surface of EZH2 (Jani et al., 2019) but there is no structural data how this proposed interaction might occur. Similarly, a recent structure of PRC2 bound to a dinucleosome revealed how the catalytic lobe of PRC2 contacts nucleosomes through DNA interactions but provided no structural insight into how the H3 N-termini might be recognized (Poepsel et al., 2018).

Here, a refined structure of PRC2 bound to a dinucleosome allowed us to visualize how the histone H3 N-terminus on substrate nucleosomes is threaded into the EZH2 active site. Our analyses reveal that H3K36 assumes a critical position in the PRC2-nucleosome interaction interface that permits the complex to gauge the H3K36 methylation state.

Results

EZH2 interaction with nucleosomal DNA orients the H3 N-terminus for H3K27 binding to the active site

We assembled recombinant full-length human PRC2 in complex with its accessory factor PHF1 (i.e. PHF1-PRC2) (Choi et al., 2017) on a heterodimeric dinucleosome (di-Nuc), which consisted of a ‘substrate’ nucleosome with unmodified histone H3 and an ‘allosteric’ nucleosome containing H3 with a trimethyllysine analog (Simon et al., 2007) at K27, separated by a 35 base pair (bp) DNA linker (Poepsel et al., 2018; Figure 1A,B). Single-particle cryo-electron microscopy analysis yielded a reconstruction of the PHF1-PRC2:di-Nuc assembly with an overall resolution of 5.2 Å (Figure 1—figure supplement 1, Figure 1—figure supplement 2, Figure 1—figure supplement 3). The map showed clear density for the catalytic lobe of PRC2 with similar chromatin interactions and binding geometry as previously described for the catalytic lobe of AEBP2-PRC2 (Poepsel et al., 2018) where PRC2 contacts the two nucleosomes via interactions with the DNA gyres (Figure 1C). Specifically, the substrate nucleosome is bound by the EZH2CXC domain residues K563, Q565, K569 and Q570 (Figure 1D, Figure 1—figure supplement 4A, cf. Poepsel et al., 2018), while the allosteric nucleosome is contacted by EED and by the SBD and SANT1 domains of EZH2 (Figure 1E, Poepsel et al., 2018). We could not detect density for the ‘bottom lobe’ of PRC2 (Chen et al., 2018; Kasinath et al., 2018) or for the N-terminal winged-helix and tudor domains of PHF1 that bind DNA and H3K36me3, respectively (Choi et al., 2017; Li et al., 2017; Ballaré et al., 2012; Cai et al., 2013; Musselman et al., 2013).

Figure 1 with 5 supplements see all
Interaction of the PRC2 catalytic lobe with nucleosomal DNA orients the H3 N-terminus for H3K27 binding to the active site.

(A) Domain organization in the five subunits of PHF1-PRC2. Dashed boxes indicate protein portions visible in the PHF1-PRC2:di-Nuc cryo-EM reconstruction and fitted in the structural model. In PHF1, C corresponds to the short C-terminal fragment used in PHF1C-PRC2. (B) Coomassie-stained SDS PAGE analysis of representative PHF1-PRC2 (upper panel) and Xenopus laevis (X.l.) octamer preparations (lower panel) after size-exclusion chromatography (SEC) purification. Pooled fractions of PHF1-PRC2, incubated with heterodimeric dinucleosomes generated by DNA ligation of a reconstituted unmodified and a H3Kc27me3-modified mononucleosome as described in Poepsel et al., 2018 were used as input material for cryo-EM analysis. (C) Cryo-EM reconstruction of PHF1-PRC2:di-Nuc in two orientations with fitted crystal structures of human PRC2 catalytic lobe (PDB: 5HYN, Justin et al., 2016) and nucleosomes (1AOI, Luger et al., 1997) in a di-Nuc model with 35 bp linker DNA (see also Figure 1—figure supplements 14, Supplementary file 1, Figure 1—video 1, Source code 1). Density is colored as in (A) to show PRC2 subunits, DNA (blue) and octamers of substrate (pink) and allosteric (yellow) nucleosomes. Boxes indicate regions shown in (D), (E) and (F), respectively. (D) Interaction of EZH2CXC residues with the DNA gyres of the substrate nucleosome; residues mutated in PRC2CXC>A are indicated. For the H3 N-terminus (pink), only the peptide backbone is shown in this view (see F). (E) Interface formed by EED and the EZH2 SBD domain with DNA gyres on the allosteric nucleosome; residues mutated in PRC2EED>A are indicated. Asterisk indicates the approximate location of a residue, which is not built in the model. (F) The H3 N-terminus (pink), shown as a pseudoatomic model fitted into the 4.4 Å density map, is recognized by EZH2 through an extensive interaction network (see text). Note the well-defined side-chain density of H3K36 (see also Figure 1—figure supplement 3D and Figure 1—figure supplement 4C–E).

Using particle signal subtraction and focused refinement on the interface of EZH2 and the substrate nucleosome (Figure 1—figure supplement 2, Figure 1—figure supplement 3), we then obtained an improved map at an apparent overall resolution of 4.4 Å which revealed well-defined density for the H3 N-terminus (Figure 1F, Figure 1—figure supplement 3B–D). The visible sidechain density combined with the crystallographic models of the PRC2 catalytic lobe and of the mononucleosome enabled us to build a pseudo-atomic model of the histone H3 N-terminus spanning residues R26 to K37 (Figure 1F). This model revealed that EZH2 recognizes the H3 N-terminus via an extended network of contacts besides the previously described ionic interactions near the active site where H3 R26 interacts with EZH2 Q648/D652, and H3 K27 with the aromatic cage above the EZH2 catalytic center (Justin et al., 2016; Figure 1F). Specifically, our structure suggests two hydrophobic hotspots, the first one involving H3 A29/P30 and EZH2 residues F667, A697, V699, I708 and F724 and the second one involving H3 V35 and F542, F557 and P558 of EZH2 (Figure 1F). H3 G33/G34 is likely not recognized by PRC2 but might act as a flexible hinge between the two hydrophobic interaction sites (Figure 1F). H3K36 is directly juxtaposed to the EZH2CXC-DNA interaction surface and appears to be involved in the EZH2-DNA interface. The side-chain density of H3K36 suggests that the epsilon-amino group of H3K36 engages in a polar interaction with the carbonyl group of Q570 and possibly in long-range electrostatic interactions with the phosphate backbone of the nucleosomal DNA (Figure 1F, Figure 1—figure supplement 4C–E). Taken together, our analyses reveal an extensive network of interactions between EZH2, the nucleosomal DNA and the H3 N-terminus. This complex geometric arrangement orients the H3 N-terminus into an extended conformation, threading H3K27 into the EZH2 active site. In this context, it should be noted that a previously postulated H3K36-binding pocket centered on E579 of EZH2 (Jani et al., 2019) is located approximately 19 Å away from H3K36 in our structure (Figure 1—figure supplement 4F). An interaction of H3K36 with E579 of EZH2 as proposed by Muir and co-workers (Jani et al., 2019) would require a very different binding geometry of PRC2 on the nucleosome and major structural rearrangements of PRC2 or the nucleosome in order to avoid steric clashes.

The EZH2 CXC contact with DNA is essential for H3K27 methylation

We next analyzed how the PRC2 surfaces contacting the substrate and the allosteric nucleosome contribute to the formation of productive PRC2-chromatin interactions. For these experiments, we used PHF1C-PRC2, which contains the minimal 5 kDa PRC2-interaction domain of PHF1 (Figure 1A, Choi et al., 2017; Chen et al., 2020) but lacks the H3K36me3-binding tudor and the DNA-binding winged-helix domains of PHF1 (Musselman et al., 2013; Choi et al., 2017; Li et al., 2017). PHF1C-PRC2 therefore only retains the DNA-binding surfaces of the 4-subunit PRC2 core complex and was used because it generally behaved better in purifications than the 4-subunit PRC2 core complex. For simplicity we shall, in the following, refer to the PHF1C-PRC2 complex as PRC2. We generated three mutant versions of PRC2. In PRC2CXC>A (K563A Q565A K569A Q570A), the EZH2CXC interface is mutated (Figure 1D), in PRC2EED>A (K77A K83A K385A K398A K400A K408A), the EED interface contacting the allosteric nucleosome (Figure 1E), is mutated, and PRC2CXC>A/EED>A carries the combination of these mutations. We first used electromobility shift assays (EMSA) to measure the binding affinity of wild-type and mutant PRC2 complexes on mononucleosomes. These mononucleosomes were assembled on a 215 bp long DNA fragment containing the 147 bp 601 nucleosome-positioning sequence (Lowary and Widom, 1998) in the center and linker DNA on both sides. Wild-type PRC2 bound this mononucleosome with an apparent Kd in the mid-nanomolar range (Figure 2A,B, cf. Choi et al., 2017). The binding affinities of PRC2CXC>A or PRC2EED>A were two- to three-fold lower than that of wild-type PRC2 and that of PRC2CXC>A/EED>A was about five-fold lower compared to the wild-type complex (Figure 2A,B, compare lanes 11–40 with 1–10). The PRC2CXC>A/EED>A complex therefore still binds to nucleosomes with sub-micromolar affinity (Figure 2A, lanes 21–30). Nucleosome binding by the PRC2CXC>A/EED>A complex could in part be due to incomplete disruption of the mutated interfaces but it likely also represents nucleosome binding contributed by the bottom lobe of PRC2 comprising the N-term of SUZ12 and RBBP4 (Chen et al., 2018; Nekrasov et al., 2005). In particular, biochemical studies on Drosophila PRC2 originally found that a minimal complex formed between Su(z)12 and the RBBP4 ortholog Caf1-55 binds to nucleosomes (Nekrasov et al., 2005). Moreover, negative stain EM analyses of human PRC2 bound to a dinucleosome identified several 2D classes where the bottom lobe contacts one or two of the two nucleosomes (Poepsel et al., 2018).

Figure 2 with 1 supplement see all
The EZH2CXC-DNA interaction interface is critical for H3K27 methylation on nucleosomes.

(A) Binding reactions with indicated concentrations of PRC2 (lanes 1–10), PRC2CXC>A (lanes 11–20), PRC2CXC>A/EED>A (lanes 21–30), or PRC2EED>A (lanes 31–40) and 45 nM 6-carboxyfluorescein-labeled mononucleosomes, analyzed by EMSA on 1.2% agarose gels. The EMSA with PRC2EED>A was run on a separate gel. (B) Quantitative analysis of EMSA data in A by densitometry of 6-carboxyfluorescein signals from independent experiments (n = 3); error bars, SEM. (C) Western Blot (WB) analysis of H3K27me1 and H3K27me3 formation in HMTase reactions with indicated concentrations of PRC2 and PRC2CXC>A on 446 nM mononucleosomes (lanes 1–7) or 223 nM dinucleosomes (lanes 8–14). Note that these concentrations result in equal numbers of nucleosomes and therefore equal numbers of H3 substrate molecules in the reactions on mono- and dinucleosomes, as can be seen from the Coomassie-stained gel of the reactions in Figure 2—figure supplement 1B. H4 WB signal served as control for western blot processing.

The binding affinity of the PRC2 core complex for chromatin therefore appears to result from interactions of at least three distinct complex surfaces with nucleosomes, the EZH2CXC domain, the EED nucleosome-binding interface and the SUZ12N:RBBP4 lobe. Considering the architecture of the catalytic lobe (Figure 1C) and of the isolated full PRC2 core complex (Kasinath et al., 2018), it is very unlikely that the EZH2CXC domain and the EED nucleosome-binding interface could simultaneously engage with the same nucleosome at a time. Finally, we note that in EMSAs monitoring binding of PRC2 to a dinucleosome, we observed a complex mixture of slowly migrating species and this has precluded experiments aimed at discriminating between binding events involving specific PRC2 surfaces with dinucleosomes. In conclusion, the structural data (Figure 1C,D and Poepsel et al., 2018) suggest that a key interaction of PRC2 with substrate nucleosomes occurs through contacts of the EZH2CXC domain with the DNA gyres, whereas the biochemical analyses argue that this interaction contributes only modestly to the overall chromatin-binding affinity of the complex.

We next analyzed how mutation of the DNA-contacting residues in the EZH2CXC domain affects H3K27 methylation by PRC2. On the same mononucleosomes used above, PRC2CXC>A showed almost no detectable histone methyltransferase (HMTase) activity compared to wild-type PRC2 (Figure 2C, compare lanes 5–7 with 2–4, see also Figure 2—figure supplement 1A). On dinucleosomes, EED binding to one nucleosome might be expected to enable interaction of the mutated EZH2CXC>A domain with the H3 N-termini on the juxtaposed second nucleosome and thereby - at least partially - restore H3K27 methylation. Indeed, on dinucleosomes, the PRC2CXC>A complex does generate H3K27me1 and -me3 although still much less efficiently than wild-type PRC2 (Figure 2C, compare lanes 12–14 with 9–11). When comparing the activity of wild-type PRC2 and the PRC2CXC>A complex, it is important to keep in mind that on dinucleosomes interpretation of H3K27me1 and -me3 formation as read-out for complex activity is considerably more complicated than on mononucleosomes because H3K27me3, once placed on one of the nucleosomes, will allosterically activate PRC2 to methylate H3K27 on the linked second nucleosome (Margueron et al., 2009; Jiao and Liu, 2015).

To complement these experiments, we also compared the HMTase activity of wild-type PRC2 and PRC2CXC>A complex on free histone H318-42 peptides using a mass spectrometry-based assay to detect H3K27 methylation. It is well established that wild-type PRC2 methylates K27 on free histone H3 with much lower efficiency than on H3 in nucleosomes (Cao et al., 2002; Czermin et al., 2002; Kuzmichev et al., 2002; Müller et al., 2002), and this is also recapitulated in our assays on H318-42 peptides where we primarily detect H3K27me1 but no H3K27me3 formation, even after extended incubation of the reaction (see Figure 2—figure supplement 1B and compare with Figure 2C). However, it is important to note that PRC2CXC>A did not show reduced K27 methylation activity compared to wild-type PRC2 on this H3 peptide substrate (Figure 2—figure supplement 1B). The mutations in the EZH2CXC domain therefore do not appear to alter the conformation of EZH2 in a way that would directly interfere with catalysis. Taken together, these observations strongly argue for a mechanism where interaction of the EZH2CXC domain with the DNA on the substrate nucleosomes is a critical step for engaging the H3 N-terminus in a manner that allows effective H3K27 methylation.

Unmodified H3K36 in the EZH2CXC-DNA interaction interface is critical for H3K27 methylation in nucleosomes

The placement of H3K36 in the EZH2CXC-DNA interface (Figure 1F) suggested that even though a tri- or di-methylated K36 side chain could theoretically be accommodated in this interface, these modified side chains might provide a less optimal fit and thereby inhibit H3K27 methylation. In EMSAs, the affinity of PRC2 for binding to mononucleosomes containing a trimethyllysine analog at H3K36 (H3Kc36me3) (Simon et al., 2007) was indistinguishable from that for binding to unmodified mononucleosomes (Figure 3A,B). As previously reported (Schmitges et al., 2011; Yuan et al., 2011), on H3Kc36me3-containing mononucleosomes, H3K27 mono- and trimethylation by PRC2 was more than 10-fold inhibited (Figure 3C, compare lanes 5–7 with 2–4, see also Figure 3—figure supplement 1A). Methylation of H3K27 was also inhibited on mononucleosomes where H3K36 had been mutated to arginine (H3K36R) and, intriguingly, also on H3K36A mononucleosomes (Figure 3C, compare lanes 8–13 with 2–4). PRC2 inhibition on H3K36R and H3K36A mononucleosomes was less severe than on H3Kc36me3 mononucleosomes (Figure 3C, compare lanes 8–13 with 5–7). We note that the quantitative analyses here show inhibition of PRC2 HMTase activity on H3K36A mononucleosomes, consistent with earlier studies (Jani et al., 2019), whereas other studies previously had failed to detect inhibition on H3K36A mononucleosomes (Schmitges et al., 2011). Taken together, productive positioning of H3K27 in the catalytic center of PRC2 appears to be exquisitely sensitive to the chemical nature of the H3K36 side chain. Neither the side chains of trimethyllysine or arginine nor the short apolar side chain of alanine appear to provide the correct fit at the position of residue 36 in H3.

Figure 3 with 1 supplement see all
The unmodified H3K36 side chain in the EZH2CXC-DNA interaction interface is critical for H3K27 methylation on nucleosomes.

(A, B) EMSA analysis and quantification as in Figure 2A and B, using PRC2 and mononucleosomes that were unmodified (lanes 1–10) or contained a trimethyllysine analog at H3K36 (H3Kc36me3, lanes 11–20) (Simon et al., 2007). (C) Western blot (WB) analysis of HMTase reactions with PRC2 as in Figure 2C on unmodified (lanes 1–4), H3Kc36me3 (lanes 5–7), H3K36R (lanes 8–10) or H3K36A (lanes 11–13) mononucleosomes (446 nM). Coomassie-stained gel of reactions is shown in Figure 3—figure supplement 1A. Bottom: quantification of H3K27me3 and H3K27me1 chemiluminescence signals, respectively, by densiometry analysis from three independent experiments. In each experiment, the methylation signal in lane four was defined as 100% and used to quantify the corresponding H3K27 methylation signals in the other lanes on the same membrane. Circles show individual data points and error bars SEM. (D) HMTase reactions monitoring H3K27me1 formation by PRC2 on H318-42 peptides that were unmodified (top) or contained K36me3 (bottom). Left: Deconvoluted ESI-MS spectra from data shown in Figure 3—figure supplement 1B. On both substrates, areas of the four colored peaks of H3K27me1-modified and unmodified substrate peptides were used for quantification of H3K27me1 formation. Right: Symbols represent percentages of peptides carrying H3K27me1 in technical triplicate experiments, error bars show SD; Welch’s t-test showed no significant (ns) difference between H3K27 monomethylation on the two peptide substrates.

To extend these analyses, we compared PRC2 HMTase activity on histone H318-42 peptides that were either unmodified or contained H3K36me3. Importantly, on this free H3 peptide, H3K36me3 did not inhibit H3K27 monomethylation by PRC2 (Figure 3D, Figure 3—figure supplement 1B). This is consistent with previous studies reporting that on peptide substrates H3K36me3 only has a minor effect on the kcat of H3K27 methylation (Schmitges et al., 2011; Jani et al., 2019). The allosteric inhibition of PRC2 by H3K36me3 therefore only occurs in the context of the geometric constraints of the nucleosome.

H3K36me3 inhibits H3K27 methylation by PHF1-PRC2

DNA-binding by the winged-helix domain of PHF1 increases the binding affinity and residence time of PHF1-PRC2 on nucleosomes about two- to three-fold, resulting in more efficient H3K27 methylation by this complex compared to PRC2 (Choi et al., 2017). Furthermore, the PHF1 tudor domain binds to H3K36me3 in the context of a nucleosome (Musselman et al., 2013) and this interaction has been reported to inhibit PHF1-PRC2 from tri-methylating H3K27 on H3K36me3-containing chromatin isolated from yeast cells (Musselman et al., 2012). To analyze how H3K36me3 inhibits PHF1-PRC2 in our fully recombinant system, we compared the HMTase activity of full-length PHF1-PRC2 (Figure 1B) on unmodified and H3Kc36me3 mononucleosomes. H3K27 mono- and tri-methylation by PHF1-PRC2 was strongly inhibited on H3Kc36me3 mononucleosomes (Figure 3—figure supplement 1C). H3K36me3 therefore inhibits H3K27 methylation by PHF1-PRC2 even though this complex has higher binding affinity and a prolonged residence time on nucleosomes (Choi et al., 2017). In Polycomblike, the Drosophila ortholog of PHF1, a region comprising the tudor domain and the adjacent PHD finger has been reported to bind H3K36me3, H3K4me3, H3K9me3 and, more weakly, also H3K14me3 and H3K27me3 (Ballaré et al., 2012). We note, however, that the tudor domain of Polycomblike contains an incomplete aromatic cage and, on its own, is unable to bind methylated lysines (Friberg et al., 2010). Further analyses will be needed to assess whether and how interaction of PHF1 or Polycomblike with H3K36me3 might change H3K27 methylation by PRC2 on more complex oligonucleosome substrates that contain both H3K36me3-modified and unmodified nucleosomes.

Drosophila with H3K36R or H3K36A mutant chromatin arrest development after completion of embryogenesis

The observation that PRC2 is not only inhibited on H3K36me2/3-modified nucleosomes but also on H3K36R and on H3K36A mutant nucleosomes prompted us to investigate how H3K27 trimethylation is affected in Drosophila with H3K36R or H3K36A mutant chromatin. H3K27me3 is primarily found on canonical histone H3 (Pengelly et al., 2013; McKay et al., 2015). We used the following strategy to replace the canonical histone H3 gene copies encoded in the HisC gene cluster with H3K36R or H3K36A mutant versions. Animals that are homozygous for a deletion of the HisC gene cluster (i.e. Df(2L)HisC homozygotes) arrest development at the blastoderm stage after exhaustion of the pool of maternally-deposited histones but transgene cassettes providing 12 copies of the wild-type histone gene unit (12xHisGUWT) rescue Df(2L)HisC homozygotes into viable adults (McKay et al., 2015; Günesdogan et al., 2010). We therefore generated Df(2L)HisC homozygotes carrying 12xHisGUH3K36R or 12xHisGUH3K36A transgene cassettes and shall refer to these animals as H3K36R and H3K36A mutants, respectively. For the analysis of H3K36R mutants, we used a strain generated by Matera and colleagues that carried a single 12xHisGUH3K36R array (McKay et al., 2015). We used the 12xHisGU transgene strategy developed by Herzig and colleagues (Günesdogan et al., 2010) that relies on the use of multiple copies of 3xHisGU miniarrays to build strains that allowed us to generate H3K36A and, as additional control, also H3K36R mutant animals.

Using the H3K36R strain from Matera and colleagues (McKay et al., 2015), we found that H3K36R mutant animals complete embryogenesis and that their cuticle morphology is indistinguishable from wildtype (Figure 4). In agreement with the results from Matera and colleagues (McKay et al., 2015), we found that these animals arrest development during the larval or pupal stages. Specifically, 81% of H3K36R mutant animals arrested development at variable time points during larval growth, 18% developed to the end of third larval instar and formed pupae that died prior to metamorphosis, and 1% developed into late pupae that complete metamorphosis but then arrested as pharate adults (Figure 4). Like Matera and colleagues (McKay et al., 2015), we have not observed any H3K36R mutants that eclose from the pupal case, and both our studies therefore disagree with a report from the Schwartz lab who claimed that H3K36R mutants would be able to develop into adults (Dorafshan et al., 2019). When we dissected the rare H3K36R mutant pharate adults from their pupal cases and examined their epidermal structures, we found that they consistently showed homeotic transformations reminiscent of polycomb group (PcG) mutants. These PcG mutant phenotypes included antenna-to-leg transformations and extra sex comb teeth on meso- and meta-thoracic legs in males (Figure 4). The molecular analysis of these PcG phenotypes will be presented below.

Drosophila with H3K36R or H3K36A mutant chromatin arrest development after completion of embryogenesis.

(A) Ventral views of cuticles from wildtype (wt), H3K36R, or H3K36A mutant embryos. Note that the cuticle pattern of the mutant animals is indistinguishable from that of the wt embryo. Below: for each genotype, the fraction of embryos that developed into larvae, pupae, pharate adults or viable adults is listed. The fraction was determined by monitoring the development of collected hatched 1st instar larvae (wt: n = 300, H3K36R (Matera strain): n = 2000) or unhatched embryos (H3K36R (strain generated in this study): n = 200; H3K36A: n = 200). The GFP marker on the Balancer chromosomes was used for identifying H3K36R and H3K36A mutants. See Materials and methods for further information and discussion. (B) Dorsal views of the posterior portion of the thorax and of the abdomen. From 2000 hatched H3K36R mutant 1st instar larvae, a total of 18 pharate adults was recovered. Most H3K36R mutant pharate adults showed a relatively normal overall body patterning apart from the homeotic transformations illustrated below. (C) Frontal view of adult heads illustrating the antenna-to-leg transformation in H3K36R mutant pharate adults. The antenna-to-leg transformation in H3K36R mutant animals ranged from mild (arrows) to more extensive transformations with formation of leg-like structures such as in this extreme case (arrowheads). (D) The sex comb in males is normally only present on the protoracic (L1) legs (arrowheads). Among the H3K36R mutant pharate adult males recovered (n = 13), five showed one or several extra sex comb teeth (arrow) on the meso- (L2) or meta-thoracic (L3) legs. Extra sex comb teeth in adults are a hallmark phenotype of polycomb mutants.

The H3K36R mutant animals from the strain constructed in this study (i.e. containing four copies of the 3xHisGUH3K36R miniarray) also completed embryogenesis, and their cuticle morphology was also indistinguishable from wildtype (Figure 4). However, 98% of individuals arrested development already at the end of embryogenesis and the 2% of mutant animals that hatched from the eggshell arrested development as first instar larvae (Figure 4). H3K36A mutants, containing 4 copies of a 3xHisGUH3K36A miniarray, also completed embryogenesis and the morphology of their embryonic cuticle also appeared indistinguishable from wildtype (Figure 4). 96% of these H3K36A mutant animals arrested development before hatching from the eggshell and the 4% that hatched died during the first larval instar (Figure 4). The difference in the lethality phase of the H3K36R and H3K36A mutants generated in this study compared to H3K36R mutants in the strain from Matera and colleagues is possibly linked to the histone rescue transgene system used (see Materials and Methods).

Drosophila with H3K36R or H3K36A mutant chromatin show diminished H3K27me3 levels at canonical PcG target genes

We next performed western blot analyses to examine H3K36me2, H3K36me3, and H3K27me3 bulk levels in H3K36R and H3K36A mutant animals. We used total nuclear extracts from late-stage H3K36A or H3K36R mutant embryos and, in the case of the H3K36R strain obtained from Matera and colleagues, we also used extracts from diploid imaginal disc and central nervous system (CNS) tissues dissected from surviving third instar larvae. For the interpretation of the following experiments, it is important to keep in mind that H3K36R and H3K36A zygotic mutant animals initially also contain a pool of maternally-deposited wild-type canonical H3 molecules that, together with H3K36R and H3K36A, become incorporated into chromatin during the pre-blastoderm cleavage cycles, up to and including the S-phase of cell cycle 14. It is only from the S-phase of cell cycle 15 onwards when only transgene-encoded histones then become incorporated into chromatin (Günesdogan et al., 2010). Although the wild-type H3 molecules in chromatin become diluted during every cell cycle and are eventually fully replaced by mutant H3, they are probably still present in the chromatin of late-stage embryos. The effective replacement of persisting wild-type H3 molecules by mutant H3 greatly varies between embryonic tissues because of the different numbers of cell divisions that take place in the different tissues prior to the end of embryogenesis. For example, whereas epidermal cells undergo only two more divisions after S-phase 14, certain cells in the CNS undergo as many as 12 divisions (Bossing et al., 1996). In diploid tissues from H3K36R mutant larvae, replacement of wildtype H3 by H3K36R can be expected to be much more complete because of the extensive cell proliferation that occurs in these tissues that, after metamorphosis, will give rise to the structures of the adult body.

We first performed western blot analyses on H3K36R mutant larvae and found that H3K36me2 and H3K36me3 bulk levels were reduced more than 4-fold compared to wildtype (Figure 5A). The residual H3K36me2 and H3K36me3 signals (Figure 5A, lane 4) probably represent the methylated versions of the histone variant H3.3 that are encoded by the genes H3.3A and H3.3B that are not located in the HisC locus and had not been mutated in these animals. Intriguingly, H3K36R mutant animals also showed an about two-fold reduction in H3K27me3 bulk levels compared to wildtype (Figure 5A, compare lanes 4–6 with 1–3). The reduction of not only H3K36me2 and H3K36me3 but also of H3K27me3 bulk levels in H3K36R mutant larvae had previously also been noted by Matera and colleagues (Meers et al., 2017). H3K27 tri-methylation by PRC2 therefore appears to be compromised in Drosophila chromatin consisting of H3K36R nucleosomes.

Figure 5 with 1 supplement see all
H3K36A and H3K36R mutants show reduced levels of H3K27me3.

(A) Western blot analysis on serial dilutions (4:2:1) of total cell extracts from wing, haltere and 3rd leg imaginal disc tissues dissected from wildtype (wt, lanes 1–3) and H3K36R mutant (lanes 4–6) third instar larvae. Blots were probed with antibodies against H3K36me3, H3K36me2, or H3K27me3; in each case, probing of the same membranes with antibodies against Caf1-55 and H2B served as controls for loading and western blot processing. Note the reduced levels of H3K36me3 and H3K36me2 but also of H3K27me3 in H3K36R mutants compared to wildtype (wt) (see text). See Materials and Methods for details of all genotypes. (B) Western blot analysis on serial dilutions (8:4:2:1) of total nuclear extracts from 21 to 24 hr old wt (lanes 1–4), H3K36A mutant (lanes 5–8) and H3K36R mutant (lanes 9–12) embryos, probed with antibodies against H3K36me3, H3K36me2 or H3K27me3; and with antibodies against Caf1-55 and H2B as controls. Note that H3K36me3 and H3K36me2 levels are reduced in H3K36A mutants but not in H3K36R mutants where they are comparable to wt. Also note that H3K27me3 levels appear undiminished in either mutant (see text). (C) Left, violin plot showing the fold-change of H3K36me2 coverage in H3K36R mutant larvae relative to wt at genes that in wildtype larval CNS and imaginal disc tissues are decorated with H3K36me2 (see Materials and Methods). The dashed line marks the median reduction (4.5-fold), the dotted lines indicate the interval comprising 80% of regions. Right, Bee plot showing the fold-change of H3K27me3 coverage in H3K36R mutant larvae relative to wt at genes that in wildtype larval CNS and imaginal disc tissues are associated with high-level H3K27me3 regions (see Materials and Methods). The dashed line marks the median reduction (2.1-fold), the dotted lines indicate the interval comprising 80% of regions. Note that H3K27me3 coverage at the HOX genes abd-A, Abd-B, Ubx and Antp is between 3- and 4-fold reduced. (D) Analysis and representation as in (C) but showing fold-changes in H3K36me2 and H3K27me3 coverage in H3K36A and H3K36R mutant late-stage embryos relative to wt at genes that in wildtype embryos are decorated with H3K36me2 and H3K27me3, respectively. Note that H3K27me3 coverage at the HOX genes abd-A, Abd-B, Ubx and Antp is about 1.5-fold reduced. See also Figure 5—figure supplement 1. (E) H3K27me3 and H3K36me2 ChIP-seq profiles in larval CNS and imaginal disc tissues from wt (blue) and H3K36R mutant (purple) third instar larvae; in the tracks showing the profiles in the H3K36R mutant, the wt profile is superimposed as reference (see Supplementary file 2 and Materials and Methods for information about normalization). Top: genomic interval containing the Bithorax-Complex harboring the HOX genes Ubx, abd-A and Abd-B; bottom: genomic interval containing the Antennapedia-Complex with the HOX genes lab, pb, Dfd, Scr, and Antp. Note the 3- to 4-fold reduction of H3K27me3 levels across the Bithorax and Antennapedia loci in H3K36R mutants. Also note that for every HOX gene, the analyzed tissues (CNS, thoracic imaginal discs and eye-antenna discs) represent a mixed population of cells with a fraction of cells in which the gene is inactive, decorated with H3K27me3 and repressed by PcG and fraction of cells in which the gene is transcriptionally active and carrying the H3K36me2 modification. (F) H3K27me3 and H3K36me2 ChIP-seq profiles at the Bithorax and Antennapedia loci as in (E) but from wt (blue), H3K36A mutant (yellow) and H3K36R mutant (purple) late-stage embryos with the wt profile superimposed in the tracks showing the profiles in the H3K36A and H3K36R mutants. H3K27me3 levels across the Bithorax and Antennapedia loci in H3K36A and H3K36R mutants are only about 1.5-fold reduced compared to wt.

We then did western blot analyses on extracts from H3K36A and H3K36R mutant embryos obtained from the strains generated in this study. In H3K36A mutants, H3K36me2 and H3K36me3 bulk levels were reduced about 3- to 4-fold compared to wildtype (Figure 5B, compare lanes 5–8 with 1–4). In H3K36R mutants, H3K36me2 and H3K36me3 bulk levels unexpectedly appeared much less severely reduced (Figure 5B, compare lanes 9–12 with 1–4 and 5–8). As discussed above, the residual H3K36me2 and H3K36me3 signal in H3K36A and H3K36R mutant embryos might in part represent modified maternally-deposited canonical wild-type H3, and in part the modified H3.3 variants. However, the reason for the differential reduction of H3K36me2 and -me3 levels in H3K36A and H3K36R mutant embryos remains unclear. In both genotypes, H3K27me3 bulk levels appeared largely unchanged compared to wildtype (Figure 5B, compare lanes 5–8 and 9–12 with 1–4).

We next performed ChIP-seq experiments to examine how the genome-wide profiles of H3K36me2 and H3K27me3 are changed in H3K36R and H3K36A mutants. In the case of H3K36R mutants, we compared these profiles in cells from imaginal disc and CNS tissues dissected from late-stage third instar H3K36R and wildtype larvae. In parallel, we also compared the profiles in late-stage H3K36A, H3K36R and wildtype embryos, for both mutants using the strains generated in this study. As expected from the western blot analyses (Figure 5A), H3K36me2 levels across the genome were strongly diminished in H3K36R mutant larvae (Figure 5C, Figure 5—figure supplement 1, Supplementary file 2). The genome-wide profile of H3K36me2 was also substantially reduced in H3K36A mutant embryos but, as expected from the western blot analyses, the profile was less severely reduced in H3K36R mutant embryos (Figure 5D, Figure 5—figure supplement 1, Supplementary file 2).

The H3K27me3 genomic profile confirmed that the levels of this modification were considerably reduced in the chromatin of late-stage H3K36R mutant larvae (Figure 5C). While the average reduction was only about two-fold (Figure 5C), H3K27me3 levels were particularly strongly diminished at canonical PRC2 target genes such as the HOX genes that in wildtype animals are decorated with high-levels of H3K27me3 (Figure 5C,E, Figure 5—figure supplement 1, Supplementary file 2). Specifically, at the HOX genes Ultrabithorax (Ubx), abdominal-A (abd-A), Abdominal-B (Abd-B) or Antennapedia (Antp), H3K27me3 levels in H3K36R mutants were between three and fourfold lower than in wildtype (Figure 5C,E).

As expected from the western blot analyses (Figure 5B), H3K36A or H3K36R mutant embryos showed no general reduction in their genome-wide H3K27me3 profiles (Figure 5D). However, in both mutants, H3K27me3 levels were about 1.5-fold reduced across the HOX gene loci (Figure 5D,F). In Drosophila with H3K36R or H3K36A chromatin, PRC2 therefore appears to be unable to generate high levels of H3K27me3 at Polycomb target genes.

Polycomb repression of HOX genes is impaired in Drosophila with H3K36R or H3K36A mutant chromatin

The PcG-like phenotypes in the rare H3K36R mutant animals that survive into pharate adults and the reduction of H3K27me3 levels in HOX gene chromatin in these mutants prompted us to analyze whether and how expression of these genes is altered in H3K36R and H3K36A mutants. In a first set of experiments, we analyzed HOX gene expression in embryos. Both mutants showed stochastic misexpression of Abd-B in single cells or pairs of cells in the CNS of late-stage embryos (Figure 6A). Abd-B misexpression in H3K36R and H3K36A mutant embryos was however clearly less widespread than in H3K27R mutant embryos or in embryos lacking the PRC2 subunit Esc that are shown for comparison (Figure 6A). Moreover, we were unable to detect misexpression of Antp or Ubx in H3K36R or H3K36A mutant embryos.

Figure 6 with 1 supplement see all
Drosophila with H3K36R or H3K36A chromatin show defective Polycomb repression at HOX genes.

(A) Ventral views of stage 16 wildtype (wt), H3K36A, H3K36R, H3K27R, or esc (esc) mutant embryos, stained with antibody against Abd-B protein; the esc mutant embryo lacked both maternal and zygotic expression of esc (see Materials and Methods for details of all genotypes). The vertical bar marks the anterior boundary of Abd-B expression in parasegment (ps) 10 in wt embryos. Note the stochastic misexpression of Abd-B protein in single cells or pairs of cells anterior to ps10 in H3K36R and H3K36A mutant embryos (arrowheads). H3K27R and esc mutant embryos show widespread misexpression of Abd-B protein in the head-to-tail pattern characteristic of PcG mutants. For reasons that are not well understood, H3K36A and H3K36R mutants also show partial loss of Abd-B expression in cells in ps10 (empty arrowheads). (B) Larval CNS and brain lobe tissues from wildtype (wt) or H3K36R mutant third instar larvae, stained with antibody against Ubx protein (red) and Hoechst (DNA) to label all nuclei; location of CNS and brain lobes (BL) are indicated in the right panel. The white bars mark the anterior boundary of Ubx expression in ps5 in wt embryos, the asterisk marks the Ubx-expressing cells in the central midline of ps4 that are part of the wild-type Ubx pattern. Note the stochastic misexpression of Ubx protein in many single cells anterior to ps5 in the CNS and in the brain lobes (arrowheads). (C) Imaginal wing (W), haltere (H) and 3rd leg (3L) discs from wildtype (wt) or H3K36R mutant third instar larvae and, as reference, discs from a larvae with clones of H3K27R mutant cells that are marked by the absence of GFP. In all cases, discs were stained with antibody against Ubx protein (red) and Hoechst (DNA) to label all nuclei. In wt animals, Ubx is expressed in the haltere and 3rd leg disc but not in the wing disc where it is repressed by the PcG machinery. Note that in H3K36R mutants, Ubx is misexpressed in small clusters of cells in the wing blade primordium of the wing disc (arrowheads) but remains repressed in the rest of the wing disc. Such misexpression was detected in 50% of wing discs (n = 28). As reference, a wing discs with H3K27R mutant clones is shown, where all cells in the clones in the wing blade primordium (arrowheads) show misexpression of Ubx whereas cells in the notum and hinge primordium show no misexpression (empty arrowheads) (cf. Pengelly et al., 2013). Also note that in H3K36R mutants (n > 30 mutant animals analyzed), Ubx expression in haltere and leg discs appears unperturbed (asterisks). (D) Eye-antennal imaginal discs from wildtype (wt) or H3K36R mutant larvae and below discs from larvae with clones of H3K36A or H3K27R mutant cells that are marked by the absence of GFP. All animals were stained with antibody against Antp protein (red) and Hoechst (DNA) to label all nuclei. Antp is not expressed in the eye-antennal disc of wt animals. Note that in H3K36R mutant discs, Antp is misexpressed in large clusters of cells (arrowheads) in the antenna primordium (Ant). Note that Antp is also misexpressed in H3K36A or H3K27R mutant cell clones in the antenna primordium (arrowheads) and that in these cases misexpression also only occurs in a subset of the mutant cells and not in all clones.

We next analyzed HOX gene expression in imaginal discs and CNS tissues from third instar H3K36R mutant larvae. In the CNS of every single mutant individual, Ubx was widely misexpressed in many single cells in an apparently stochastic pattern (Figure 6B). 50% of the H3K36R mutant larvae also showed stochastic misexpression of Ubx in individual cells in the wing blade primordium of the wing imaginal disc (Figure 6C), the area of this disc where Ubx is most readily de-repressed if PcG function is perturbed (Beuchle et al., 2001). Ubx misexpression in H3K36R mutant wing discs was less widespread than in clones of H3K27R mutant cells that were induced in H3K27R heterozygotes and are shown for comparison (Figure 6C). Finally, we found that 100% of the H3K36R mutant larvae showed misexpression of Antp in the antenna primordium of the eye-anntennal disc (Figure 6D). We also observed this misexpression in clones of H3K36A homozygous cells that we had induced in H3K36A heterozygous animals (Figure 6D) and in H3K27R mutant clones that were induced as control (Figure 6D). Analogous to Ubx in the wing disc, Antp is misexpressed in the antenna primordium, the region of the eye-antenna imaginal discs where Antp is most susceptible to becoming misexpressed if PcG function is compromised. In this context, it should also be emphasized that the reduction of H3K27me3 signal in tissues from third instar H3K36R mutant larvae is quite uniform across the tissues, as illustrated in Figure 6—figure supplement 1. It should also be emphasized that transcriptome analyses on whole H3K36R mutant third instar larvae found no extensive global deregulation of gene transcription (Meers et al., 2017). The most straightforward explanation for the stochastic misexpression of multiple HOX genes in animals with chromatin consisting of H3K36R or H3K36A nucleosomes therefore is that it is caused by defective Polycomb repression as a result of the reduced H3K27me3 levels in HOX gene chromatin.

Discussion

Understanding how PRC2 binds chromatin and how it is regulated is essential for understanding how the complex marks genes for Polycomb repression to maintain cell fate decisions. The work in this study leads to the following main conclusions. First, the structure of nucleosome-bound PHF1-PRC2 allowed us to visualize how interaction of the catalytic lobe of the complex with the substrate nucleosome threads the histone H3 N-terminus into the active site of EZH2 through a relay of contacts. Second, structure-guided mutational analyses showed that DNA-binding by the EZH2CXC domain is critical for productive PRC2-nucleosome interactions. Third, unmodified H3K36 is accommodated in a key position in the EZH2CXC-DNA interface and while H3K36 provides the correct fit, the methylated forms H3K36me2/3, or mutated H3K36R or H3K36A do not seem to fit because they strongly diminish H3K27 methylation. Fourth, H3K36 is also critical for normal H3K27 methylation in vivo because Drosophila with H3K36R or H3K36A mutant chromatin show reduced levels of H3K27me3 and fail to fully maintain Polycomb repression at HOX target genes. In the following, we shall discuss key aspects of these new findings in the context of our previous knowledge of PRC2 regulation and function.

Different forms of PRC2 use the same molecular interactions for binding the H3 N-terminus on substrate nucleosomes

Unlike many other histone-modifying enzymes (e.g. McGinty et al., 2014; Worden et al., 2019), PRC2 does not recognize the nucleosome by docking on its acidic patch (Luger et al., 1997) to engage with the histone substrate. Instead, the complex interacts with chromatin by binding to the DNA gyres on the nucleosome (Poepsel et al., 2018, this study). Prevous studies that had measured the binding affinity and residence time of PRC2 on nucleosomes and free DNA had found that DNA-binding makes the largest contribution to the chromatin-binding affinity of PRC2 (Choi et al., 2017; Wang et al., 2017). The mutational analyses here establish that interaction of highly conserved residues in the EZH2CXC domain with the DNA on the substrate nucleosome is critical for H3K27 methylation (Figure 2C). Moreover, this interaction sets the register for a network of interactions of the H3 N-terminus with the EZH2 surface that permits H3K27 to reach into the active site (Figure 1D,F). Consistent with our findings here, an independent recent study of a cryo-EM structure of PRC2 with co-factors JARID2 and AEBP2 bound to a mononucleosome with monoubiquitylated H2A (Kasinath et al., 2020) identified very similar interactions of EZH2 with the nucleosomal DNA and the H3 N-terminus. Different forms of PRC2 that contain different accessory proteins and dock in different ways on chromatin therefore contact the substrate H3 N-terminus in the nucleosome through similar interactions.

The position of H3K36 in the EZH2CXC-nucleosome interface enables allosteric regulation by H3K36 methylation

Important novel insight from our structure came from the observation that unmodified H3K36 is located in a critical position in the EZH2CXC-DNA interface. Unmodified H3K36 has the right fit for interaction of the H3 N-terminus with the EZH2 surface and placement of H3K27 in the active site. The inhibition of H3K27 mono-, di- and tri-methylation on nucleosomes carrying H3K36me2 or -me3 (Schmitges et al., 2011; Yuan et al., 2011) or on H3K36R or H3K36A nucleosomes (Figure 3C) suggests that even though these side chains could theoretically be accommodated in the EZH2CXC-DNA interface, these alterations of the side chain of residue 36 in H3 chain must somehow impair productive interaction of H3K27 with the catalytic center of EZH2. On isolated H3 N-terminal peptides, H3K36me3 did not inhibit the formation of H3K27me1 (Figure 3D), consistent with earlier findings that on peptide substrates H3K36me3 only has a minor effect on the kcat of H3K27 methylation (Schmitges et al., 2011; Jani et al., 2019). Also, H3K36me3 does not diminish the affinity of PRC2 for binding to mononucleosomes (Figure 3A,B) and does not reduce the residence time of PRC2 on nucleosome arrays (Guidotti et al., 2019). Taken together, a possible scenario would therefore be that within the time frame of the PRC2-nucleosome binding and reaction cycle, docking of the H3K36 side chain in the EZH2CXC-DNA interface is critical for rapid alignment of the H3 N-terminus on the EZH2 surface into a catalytically competent state. According to this view, H3K36me2/3 does not locally disrupt nucleosome binding but allosterically inhibits H3K27 from interacting with the EZH2 active site.

H3K27 methylation and polycomb repression are defective in Drosophila with H3K36R or H3K36A chromatin

The finding that PRC2 is inhibited on H3K36R and H3K36A nucleosomes in vitro had prompted us to use a genetic histone replacement strategy in Drosophila (Günesdogan et al., 2010; McKay et al., 2015) to assess PRC2 inhibition on H3K36R or H3K36A chromatin in vivo. Previous studies had found that Drosophila H3K36R mutants are able to develop into the pupal stages and, consistent with this late developmental arrest, whole third instar larvae were found to show only relatively minor changes in their transcriptome compared to wildtype animals (McKay et al., 2015; Meers et al., 2017). Of relevance here, these transcriptome analyses did not reveal any gross deregulation of HOX or PcG genes (Meers et al., 2017). Here, we found that a few rare H3K36R mutant animals even survive into pharate adults and that these show remarkably little morphological defects apart from homeotic transformations characteristic of Polycomb mutants (Figure 4). We show that these phenotypes are caused by misexpression of multiple HOX genes (Figure 6). HOX gene misexpression in H3K36R or H3K36A mutants is stochastic and not as widespread as in strong PcG mutants but it occurs in cells and tissues where HOX genes also first become misexpressed if PcG function is removed. We found that HOX misexpression is directly linked to reduced levels of H3K27me3 at these genes (Figure 5). A simple straightforward explanation for these phenotypes in H3K36R or H3K36A mutant animals is that PRC2 is unable to effectively deposit high levels of H3K27me3 on the H3K36R or H3K36A nucleosomes, respectively, in their chromatin. Accordingly, H3K27me3 levels at HOX genes are below the threshold needed to stringently maintain Polycomb repression and consequently, HOX genes become stochastically misexpressed in a fraction of cells. Finally, we note that in H3K36R mutant larvae, the experimental setting where we have been able to generate the most complete replacement of H3 by H3K36R, H3K27me3 levels at HOX genes were only about 3- to 4-fold reduced compared to wildtype (Figure 5C). However, as shown in Figure 3C, on nucleosomes in vitro, H3K36me3 inhibited PRC2 more effectively than H3K36R or H3K36A. It therefore seems likely that in contrast to the H3K36R and H3K36A mutants that we have used as proxy, H3K36me2 and H3K36me3 in vivo also inhibit PRC2 more effectively from depositing H3K27me3 on H3K36me2- or H3K36me3-modified nucleosomes in transcriptionally active chromatin.

Concluding remark

The structural, biochemical and genetic work reported in this study shows that it is the exquisite geometry formed by a relay of interactions between the PRC2 enzyme, nucleosomal DNA and the H3 N-terminus that enable the histone methylation marks H3K36me2 and H3K36me3 in transcriptionally active chromatin to allosterically prevent PRC2 from depositing the repressive histone methylation mark H3K27me3 at transcribed genes.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Strain, strain background D. melanogasterOregon-RFlybase
Strain, strain background D. melanogasterw; Df(2L)HisC FRT40A/Df(2L)HisCFRT40A; 12xHisGUwt/12 xHisGUwtMcKay et al., 2015
Strain, strain background D. melanogasterw; Df(2L)HisC FRT40A/CyO ubi-GFP; 12xHisGUH3K36R/TM6BMcKay et al., 2015
Strain, strain background D. melanogasterw; Df(2L)HisC FRT40A/CyO twi:Gal4 UAS:GFP; 3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)/3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)This studyAvailable on request
Strain, strain background D. melanogasterw; Df(2L)HisC FRT40A/CyO twi:Gal4 UAS:GFP; 3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb)/3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb)This studyAvailable on request
Strain, strain background D. melanogasterw; Df(2L)HisC FRT40A/CyO ubi:GFP; 3xHisGUH3K27R(68E) 3xHisGUH3K27R (86Fb)/3xHisGUH3K27R (68E) 3xHisGUH3K27R (86Fb)Pengelly et al., 2013
Strain, strain background D. melanogasterw hs-flp; w; hs-nGFP FRT40A/hs nGFP FRT40; 3xHisGUH3K27R(68E) 3xHisGUH3K27R(86Fb)/3xHisGUH3K27R(68E) 3xHisGUH3K27R (86Fb)Pengelly et al., 2013
Strain, strain background D. melanogasterw hs-flp; M(2)25A ubi-GFP FRT40A/CyOMüller lab stocksAvailable on request
Strain, strain background D. melanogasteryw; esc6 b pr/CyO, P[esc+]Struhl laboratory
Struhl, 1981
Strain, strain background D. melanogasterIn(2LR) Gla/CyO, esc2Struhl laboratory
Struhl, 1981
Strain, strain background D. melanogasterw hs-flp; hs-nGFP FRT2A/hs nGFP FRT2ABeuchle et al., 2001
Strain, strain background (Escherichia coli)BL21(DE3)Sigma-AldrichCMC0016Electrocompetent cells
Cell line (Trichoplusia ni)HighFive cell line for expressionInvitrogenProduct nr.: B85502
BTI-Tn-5B1-4 (RRID:CVCL_C190)
Protein expression
Cell line (Spodoptera frugiperda)Sf21 cell line for Baculovirus productionInvitrogenProduct nr.: 1149701 (RRID:CVCL_0518)Baculovirus production for protein expression
AntibodyH3K27me3
Rabbit monoclonal antibody
Cell Signaling
Technology
Cell Signaling Technology #9733IF (1:50)
WB (1:2000)
ChIP (1:500)
AntibodyH3K27me3
Rabbit polyclonal antibody
MilliporeMillipore
#07–449
WB (1:1000)
AntibodyH3K27me1
Rabbit polyclonal antibody
MilliporeMillipore
#07–448
WB (1:6000)
AntibodyH3K36me3
Rabbit monoclonal antibody
Cell Signaling
Technology
Cell Signaling Technology #4909WB (1:750)
AntibodyH3K36me2
Rabbit monoclonal antibody
Cell Signaling
Technology
Cell Signaling Technology #2901WB (1:250)
AntibodyH3K36me2
Rabbit monoclonal antibody
Abcam#9049ChIP (1:300)
AntibodyH2B
Rabbit polyclonal antibody
This studyRaised against full-length recombinant Drosophila H2BWB (1:10000)
Available on request
AntibodyH4
Rabbit polyclonal antibody
AbcamAbcam
#10158
WB (1:200000)
AntibodyCaf1
Rabbit polyclonal antibody
Gambetta et al., 2009Müller labWB (1:10000)
AntibodyAbd-B
Mouse monoclonal antibody
DHSBDSHB (1A2E9)IF (1:300)
AntibodyAntp
Mouse monoclonal antibody
DHSBDSHB (8C11)IF (1: 100)
Recombinant DNA reagentpfC31-attB-3xHisGU.H3K36AThis studySee Materials and Methods
Available on request
Recombinant DNA reagentpfC31-attB-3xHisGU.H3K36RThis studySee Materials and Methods
Available on request
Recombinant DNA reagentnucleosome-positioning sequence 601
(147 bp + linker version)
Lowary and Widom, 1998, Nekrasov et al., 2005
Recombinant DNA reagentpFB-EZH2Choi et al., 2017N-terminal 6xHis-tag
Recombinant DNA reagentpFB-EZH2CXC>AThis studyN-terminal 6xHis-tag
See Materials and Methods
Available on request
Recombinant DNA reagentpFB-EEDChoi et al., 2017N-terminal 6xHis-tag
Recombinant DNA reagentpFB-EEDEED>AThis studyN-terminal 6xHis-tag
See Materials and Methods
Available on request
Recombinant DNA reagentpFB-SUZ12Choi et al., 2017N-terminal 6xHis-tag
Recombinant DNA reagentpFB-RBBP4Choi et al., 2017N-terminal 6xHis-tag
Recombinant DNA reagentpFB-PHF1Choi et al., 2017N-terminal twin-strep and 6xHis-tag (SHT)
Recombinant DNA reagentpFB-PHF1C
(PHF1515-567)
Choi et al., 2017N-terminal twin-strep and 6xHis-tag (SHT)
Peptide, recombinant proteinH318-42 peptideMPIB core facilty
Peptide, recombinant proteinH318-42K36me3 peptideMPIB core facilty

Protein expression and purification

Human PHF1-PRC2 wild-type (wt) complex was expressed and purified as previously described (Choi et al., 2017). In brief, an optimized ratio of the baculoviruses (produced in Sf21 cells, (Invitrogen 1149701)) for the different PHF1-PRC2 subunits was used to infect Trichoplusia ni High Five insect cells (Invitrogen B85502). The Sf21 and High Five cells were authenticated by genotyping (Eurofins) and tested negative for mycoplasma contamination (LookOut Mycoplasma PCR Detection Kit, Sigma-Aldrich). Cells were lysed using a glass Dounce homogenizer and the complex was purified using affinity chromatography (Ni-NTA and Strep-tag), followed by simultaneous TEV mediated protease tag cleavage and Lambda Phosphatase treatment (obtained from the MPI of Biochemistry Protein Core facility) and a final size-exclusion chromatography (SEC) step in a buffer containing 25 mM Hepes, pH 7.8, 150 mM NaCl, 10% glycerol, 2 mM DTT.

PRC2CXC>A, PRC2EED>A and PRC2CXC>A/EED>A mutants were generated by PCR with primers containing the desired mutations, subsequent ligation and transformation. Expression and purification were performed as above.

Xenopus laevis (X.l.) and D. melanogaster (D.m.) histones were expressed in E.coli strains BL21 and purified from inclusion bodies as described in Luger et al., 1999. To mimic the inhibitory mark H3K36me3 or the allosteric activating mark H3K27me3, the cysteine side chain of a mutated D.m. histone H3C110A K36C or X.l. histone H3C110A K27C was alkylated with (2-bromoethyl) trimethylammonium bromide (Sigma-Aldrich) as described previously (Simon et al., 2007). Nucleosomes containing these modifications are abbreviated with e.g. H3Kc36me3.

For histone octamers, equimolar amounts of histones H2A, H2B, H4 and H3 (wt, H3K36A, H3K36R, H3Kc27me3 or H3Kc36me3) were mixed and assembled into octamers in high salt buffer containing 10 mM Tris-HCL pH 7.5, 2 M NaCl, 1 mM EDTA, 5 mM β-mercaptoethanol. Subsequent SEC was performed to separate octamers from H3/H4 tetramers or H2A/H2B dimers (Luger et al., 1999).

Reconstitution of nucleosomes

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For X.l. and D.m mononucleosomes used in biochemical assays, 6-carboxyfluorescein (6-FAM)-labeled 215 bp 601 DNA (Lowary and Widom, 1998) was PCR amplified from the p601 plasmid, purified on a MonoQ column (GE Healthcare), precipitated with ethanol and dissolved in the same high salt buffer used for octamers. Optimized ratios of octamer to DNA (usually ranging between 0.8–1.3: 1) were mixed and nucleosomes were reconstituted by gradient and stepwise dialysis against low salt buffers to a final buffer containing 25 mM Hepes, pH 7.8, 60 mM NaCl, 2 mM DTT.

X.l. asymmetrical dinucleosomes for cryo-EM studies containing one unmodified substrate nucleosome and one H3Kc27me3-modified (allosteric) nucleosome connected with a 35 bp linker DNA were reconstituted using the protocol described in Poepsel et al., 2018. In brief, substrate nucleosomes and allosteric nucleosomes were separately assembled on the respective DraIII digested nucleosomal DNA. The latter was generated by PCR with primers introducing the desired linker and DraIII recognition sites and purified as described above. The assembled nucleosomes were purified on a preparative native gel system (Biorad 491 prep cell). After ligation using T4 ligase (Thermo Fisher Scientific) the resulting dinucleosomes were purified from aberrant or non-ligated mononucleosomes by a second preparative native gel system (Biorad 491 prep cell). In contrast to Poepsel et al., 2018, the dinucleosome DNA used in this study contained an additional 30 bp overhang on the substrate nucleosome, thus resulting in the following DNA sequence:

5′–601 binding (allosteric nucleosome) – agcgatctCACCCCGTGatgctcgatactgtcata – 601 binding (substrate nucleosome) – atgcatgcatatcattcgatctgagctcca –3’ (after DraIII digestion, assembly of substrate/allosteric nucleosome and ligation to dinucleosomes).

X.l. symmetrical unmodified dinucleosomes used for the HMTase assays with the PRC2CXC mutants were obtained by reconstituting octamers with a 377 bp DNA containing two 601 sequences connected by a 35 bp linker DNA. A vector containing the 377 bp sequence was ordered from Invitrogen GeneArt and was used for PCR resulting in:

5′–atatctcgggcttatgtgatggac – 601 binding (substrate nucleosome 1) – agcgatctcaacgagtgatgctcgatactgtcata – 601 binding (substrate nucleosome 2) – gtattgaacagcgactcgggatat–3′.

The PCR products were purified as described above. Optimized ratios of octamer: DNA (usually ranging between 1.8–2.3: 1) were mixed and nucleosomes were reconstituted by gradient and stepwise dialysis against low salt buffers to a final buffer containing 25 mM Hepes, pH 7.8, 60 mM NaCl, 2 mM DTT.

Cryo-EM data acquisition

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Complexes of PHF1-PRC2 and asymmetrically modified 35 bp dinucleosomes were assembled and grids were prepared as described previously, with the difference of using 0.005% NP40 instead of 0.01% (Poepsel et al., 2018). Cryo-EM data were collected on an FEI Titan Krios microscope operated at 300 kV and equipped with a post-column GIF and a K2 Summit direct detector (Gatan) operated in counting mode. A total of 3467 movies were collected at a nominal magnification of 81,000x (1.746 Å/pixel) at the specimen level using a total exposure of 53 e-/ Å2 distributed over 60 frames and a target defocus range from 1.5 to 3 µm. Data acquisition was carried out with SerialEM.

Cryo-EM data processing

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Movies were aligned and corrected for beam-induced motion as well as dose compensated using MotionCor2 (Zheng et al., 2017). CTF estimation of the summed micrographs was performed with Gctf (Zhang, 2016) and particles were picked in Gautomatch (http://www.mrc-lmb.cam.ac.uk/kzhang/ K. Zhang, MRC LMB, Cambridge, UK) using templates created from the AEBP2-PRC2-dinucleosome cryo-EM structure (EMD-7306, Poepsel et al., 2018). All subsequent image processing steps were performed in Relion 3.0 (Zivanov et al., 2018) as shown in Figure 1—figure supplement 2. A total of 1,028,229 candidate particles were subjected to two rounds of initial 3D classification against a reference map (AEBP2-PRC2-dinucleosome low-pass filtered to 60 Å) and the Bayesian fudge factor (T value) set to 8. 330,482 remaining particles were subjected to two more rounds of 3D classification, this time using the best 3D model from the previous run as reference. Finally, the two best 3D models were 3D refined and further classified into 10 classes without translational and rotational sampling, using a T value of 4. From this run, the best 3D classes with the highest nominal overall resolution and rotational and translational accuracies were subjected to iterative rounds of 3D refinement, this time applying a soft mask for solvent flattening, per particle CTF refinement and Bayesian polishing. The highest nominal resolution was only achieved by combining several classes from the previous 3D run, likely due to missing particle views in one or the other individual class. The final map after postprocessing had an overall nominal resolution of 5.2 Å, as determined from the gold-standard FSC criterion of 0.143 (Rosenthal and Henderson, 2003; Figure 1—figure supplement 1D). The density (Overall PHF1-PRC2:di-Nuc) with fitted models is shown in Figure 1C and in Figure 1—figure supplement 1E using UCSF ChimeraX (Goddard et al., 2018). Local resolution estimation was performed in Relion 3.0 and is shown in Figure 1—figure supplement 1B. The spherical angular distribution of all particles in the final model is shown in Figure 1—figure supplement 1C.

To further improve the resolution and map details of the region around the H3 N-terminus, particle subtraction and focused 3D refinement was applied (Bai et al., 2015; Zhou et al., 2015; Ilca et al., 2015). Using a mask generated with UCSF Chimera (Pettersen et al., 2004) and Relion 3.0 the signal of the allosteric nucleosome as well as parts of PRC2 (EED and EZH2allo) was subtracted from all particle images. These signal subtracted particles were then subjected to focused 3D refinement using a soft mask around the substrate nucleosome and EZH2sub. This yielded a 4.4 Å map (EZH2sub-Nucsub) as determined from the gold-standard FSC criterion of 0.143 (Rosenthal and Henderson, 2003Figure 1—figure supplement 3B). Local resolution estimation is shown in Figure 1—figure supplement 3A. For model building and depiction, the final density was further sharpened (applied b – factor: - 66) using the Multisharpen function in Coot (Emsley et al., 2010) (e.g. in Figure 1F, Figure 1—figure supplement 3D).

To confirm the side-chain information visible in the Coot sharpened map, Phenix Resolve density modification was run on the two half maps generated from the 3D refinement of the EZH2sub-Nucsub map (Terwilliger et al., 2020). The resolution of the map according to Phenix cryo-EM density modification output improved to 4 Å and the resulting map was used as an additional guideline for model building as well as for depiction (in Figure 1—figure supplement 4A-D).

Cryo-EM data fitting, modeling and refinement

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Available crystal structures were fitted into the final maps using rigid-body fitting in UCSF Chimera and all manual remodeling, morphing and building was performed in Coot. For PRC2, the crystal structure of the catalytic lobe of human PRC2 (PDB: 5HYN Justin et al., 2016; and comparing the fitted model to the cryo EM model of AEBP2-JARID2-PRC2 PDBs: 6C23 and 6C24 Kasinath et al., 2018) was used. Since the SBD helix and the SANT1 helix bundle of the crystal structure was not accommodated well by the corresponding EM density, this region was fitted separately. A model of a dinucleosome with linker DNA (Supplementary dataset one in Poepsel et al., 2018, including crystal structures of nucleosomes, PDB 3LZ1, Vasudevan et al., 2010, also PDB 1AOI, Luger et al., 1997, also PDB 6T9L, Wang et al., 2020, was fitted.

The above described overall model was then used as a starting model for fitting and building EZH2sub-Nucsub into the focused map. Where possible, missing parts in the model were built de-novo, that is the H3 N-terminal tail (residues 30–37) between the catalytic site of PRC2 and the substrate histone. Available information from crystal/cryo EM structures was used as a guide (PRC2 with H3 peptide bound: PDB: 5HYN Justin et al., 2016 and cryo EM model of AEBP2-JARID2-PRC2 PDBs: 6C23 and 6C24 Kasinath et al., 2018), and high-resolution structures of nucleosomes (PDB 1AOI and PDB 6T9L) (Luger et al., 1997; Wang et al., 2020). Parts of EZH2sub-Nucsub model were then fitted using the morph fit routine in Coot or manually (Casañal et al., 2020). Secondary structure restraints for real-space refinement were generated automatically with phenix.secondary_structure_restraints (Sobolev et al., 2015) and manually curated. Hydrogens were added and the model was real-space refined with a resolution- cutoff of 4.4 Å with Phenix (Afonine et al., 2018) (phenix-1.18rc1-3777), using reference structures (PDB 6T9L, Wang et al., 2020, and PDB 1AOI, Luger et al., 1997 for nucleosome and one copy of the human PRC2 crystal structure generated from PDB 5HYN (Justin et al., 2016 ), applying strict secondary structure and Ramachandran restraints.

Our final model includes the modeled side chains of the fitted crystal/cryo-EM structures. This is in our opinion supported by the data as the substrate nucleosome protein core is resolved to app. 4 Å (Figure 1—figure supplement 3A) and the map in these regions shows clear bulky side-chain information (Figure 1—figure supplement 3D). The EZH2 density is of worse quality however even at lower resolution side chains likely contribute to the signal in the particle images and thereby an overall good model to map fit (in our case given by the high CC values as well as FSCmodelvsmap) is arguably only ensured in the presence of side chains. However we caution readers against in interpreting our model at side-chain resolution in poorly resolved regions.

Structures were visualized with UCSF ChimeraX (Goddard et al., 2018) and PyMOL2 (https://pymol.org/2/).

Electrophoretic mobility shift assay (EMSA)

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EMSAs on a 1.2% agarose gel in 0.4x TBE Buffer with 45 nM 6-FAM - labeled mononucleosomes (unmodified wt X.l. for bandshifts with the PRC2CXC mutants, unmodified wt D.m. and D.m H3Kc36me3 Simon et al., 2007) trimethyllysine analog containing nucleosomes) and increasing PRC2 concentrations (concentrations indicated in the figures above the gels) were performed in triplicates as described in Choi et al., 2017. A Typhoon FLA 9500 scanner and the Fiji software was used for densitometric analysis of the 6-FAM signal (Schindelin et al., 2012). Background correction and calculation of the fractions of bound nucleosomes was performed with R using tidyverse (https://www.r-project.org/). In detail: two parts were boxed out in each lane: 1. unbound nucleosomes (‘unbound’ box) and 2. shifted nucleosomes (‘bound’, everything above ‘unbound’). The boxed-out signals were integrated and background corrected by subtracting the respective control (‘bound’ background of lane one for ‘bound’ boxes and ‘unbound’ background of lane 10 for ‘unbound’ boxes). To calculate the fraction of bound vs. unbound nucleosomes, the value for ‘bound’ nucleosome in each lane was divided by the total signal (sum of bound and unbound) of the same lane. Hill function fitting and illustration of the plot were subsequently performed with Prism 8 (GraphPad).

Histone methyltransferase (HMTase) assay

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For all HMTase assays, 446 nM of mononucleosomes or 223 nM of dinucleosomes were incubated with indicated amounts of the different PRC2 complexes, in a reaction buffer containing 20 mM HEPES pH 7.8, 50 mM NaCl, 2.5 mM MgCl2, 5% glycerol, 0.25 mM EDTA, 0.5 mM DTT and 80 μM S-adenosylmethionine (SAM). Reactions were allowed to proceed for 90 min at RT before quenching by the addition of 1x (final concentration) SDS loading buffer and heat inactivation at 95°C for 5 min. Proteins were separated by electrophoresis on a 16% (w/v) SDS gel, transferred to a nitrocellulose membrane and probed with antibodies against H3K27me3 (Millipore, 07–449), H3K27me1 (Millipore, 07–448) and H4 (Abcam, ab10158). For quantification, HMTase reactions and the corresponding western blots on D.m. unmodified, H3Kc36me3, H3K36A/R mononucleosomes were performed in triplicates and subjected to densitometric analysis (Chemiluminescence signal, ImageQuant LAS 4000). The integrated densitometric signal (band) in each lane was background corrected against the control lane (lane 1, no PRC2 in the reaction) and normalized with respect to the lane containing the highest amount (i.e. 100%) of PRC2 on unmodified nucleosomes (lane 4). The relative amounts of trimethylation/monomethylation for all other lanes were calculated with respect to lane 4. Graphical representations were made with Prism 8 (GraphPad).

Mass spectrometry (MS)

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500 nM of PRC2 or PRC2CXC>A were incubated with 2 µM of either unmodified or H318-42 peptide containing the K36me3 modification in HMTase reaction buffer (described above) and methyltransferase activity was allowed to proceed over night at RT. Reactions were then quenched with 1% trifluoroacetic acid (TFA). Home-made stage tips with poly(styrenedivinylbenzene) copolymer (SDB-XC) were used to remove PRC2 from the reactions (Rappsilber et al., 2007). First, stage tips were washed with methanol, followed by a second wash with buffer B (0.1% (v/v) formic acid, 80% (v/v) acetonitrile). The SDB-XC material was then equilibrated with buffer A (0.1% (v/v) formic acid) and 40 µl of sample was applied and washed several times. Finally, samples were eluted using buffer B and introduced into the Bruker maXis II ETD mass spectrometer by flow injection of 20 µl sample using an Agilent HPLC at a flow rate of 250 µl/min and 0.05% TFA in 70% acetonitril:H2O as solvent for ESI-MS time-of-flight analysis. Peptides were ionized at a capillary voltage of 4500 V and an end plate offset of 500 V. Full scan MS spectra (200–1600 m/z) were acquired at a spectra rate of 1 Hz and a collision cell energy of 15 eV.

Raw data files were processed using Bruker Compass DataAnalysis. The m/z spectra were deconvoluted (maximum entropy method) with an instrument resolving power of 10,000 and the resulting neutral spectra peaks were integrated. For quantification, the experiment was performed in triplicates. The sum of the monomethylation peak areas was divided by the sum of the first 4 peaks of the input peptide together with the sum of the monomethylation peak areas. Illustration of the quantification was subsequently performed with Prism 8 (GraphPad). A Welch’s t-test was calculated to show the nonsignificant difference between the activity of PRC2 on unmodified or H3K36me3 peptide.

Construction of histone transgenes to generate H3K36A and H3K36R strains

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Site directed mutagenesis on pENTR221-HisGU.WT, pENTRL4R1-HisGU.WT and pENTRR2L3-HisGU.WT (Günesdogan et al., 2010) was used to mutate histone H3K36 to alanine or arginine. The final constructs pfC31-attB-3xHisGU.H3K36A and pfC31-attB-3xHisGU.H3K36R were generated by Gateway LR recombination of above vectors and integrated at attP sites VK33 (BDSC 9750) and 86Fb (BDSC 130437). The full genotypes of animals used in the study are described below.

Drosophila strains and genotypes

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The following strains were used in this study:

  • Oregon-R

  • w; Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 12xHisGUwt/12 xHisGUwt (McKay et al., 2015)

  • w; Df(2L)HisC FRT40A/CyO ubi-GFP; 12xHisGUH3K36R/TM6B (McKay et al., 2015)

  • w; Df(2L)HisC FRT40A/CyO twi:Gal4 UAS:GFP; 3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)/3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb) (generated in this study)

  • w; Df(2L)HisC FRT40A/CyO twi:Gal4 UAS:GFP; 3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb)/3xHisGUH3K36R(VK33) .3xHisGUH3K36R(86Fb) (generated in this study)

  • w; Df(2L)HisC FRT40A/CyO ubi:GFP; 3xHisGUH3K27R(68E) 3xHisGUH3K27R (86Fb)/3xHisGUH3K27R (68E) 3xHisGUH3K27R (86Fb) (Pengelly et al., 2013)

  • w hs-flp; w; hs-nGFP FRT40A/hs nGFP FRT40; 3xHisGUH3K27R(68E) 3xHisGUH3K27R(86Fb)/3xHisGUH3K27R(68E) 3xHisGUH3K27R (86Fb) (Pengelly et al., 2013)

  • w hs-flp; M(2)25A ubi-GFP FRT40A/CyO yw; esc6 b pr/CyO, P[esc+]

  • In(2LR) Gla/CyO, esc2

  • w hs-flp; hs-nGFP FRT2A/hs nGFP FRT2A

The following genotypes were used for the experiments shown in:

  • Figure 4

  • wt: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUwt(VK33)/12xHisGUwt(VK33)

  • H3K36R: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUH3K36R(VK33)/TM6B

  • H3K36R: w; Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb)/3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb)

  • H3K36A: Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)/3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)

  • Figure 5A, C, E

  • wt: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUwt(VK33)/12xHisGUwt(VK33)

  • H3K36R: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUH3K36R(VK33)/TM6B

  • Figure 5B, D, F

  • wt: Oregon-R

  • H3K36A: w; Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)/3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)

  • H3K36R: w; Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb)/3xHisGUH3K36R(VK33). 3xHisGUH3K36R(86Fb)

  • Figure 6A

  • wt: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUwt(VK33)/12xHisGUwt(VK33)

  • H3K36R: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUH3K36R(VK33)/TM6B

  • H3K36R: w; Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb)/3xHisGUH3K36R(VK33) 3xHisGUH3K36R(86Fb).

  • H3K36A: w; Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb)/3xHisGUH3K36A(VK33) 3xHisGUH3K36A(86Fb).

  • H3K27R: w; Df(2L)HisC FRT40A/Df(2L)HisC FRT40A; 3xHisGUH3K27R(68E) 3xHisGUH3K27R (86Fb)/3xHisGUH3K27R (68E) 3xHisGUH3K27R (86Fb)

  • esc: esc6 b pr/CyO, esc2 (escmat- zyg- obtained as progeny from esc6 b pr/CyO, esc2 parents).

  • Figure 6B

  • wt: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUwt(VK33)/12xHisGUwt(VK33)

  • H3K36R: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUH3K36R(VK33)/TM6B

  • Figure 6C

  • wt: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUwt(VK33)/12xHisGUwt(VK33)

  • H3K36R: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUH3K36R(VK33)/TM6B

  • H3K27R: w hs-flp; Df(2L)HisC FRT40A/hs-nGFP FRT40A; 3xHisGUH3K27R(68E)3xHisGUH3K27R(86Fb)/3xHisGUH3K27R(68E)3xHisGUH3K27R(86Fb)

  • Figure 6D

  • wt: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUwt(VK33)/12xHisGUwt(VK33)

  • H3K36R: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUH3K36R(VK33)/TM6B

  • H3K36A: w hs-flp; Df(2L)HisC FRT40A/M(2)25AubiGFP FRT40; 3xHisGUH3K36A(VK33) 3xHisGUH3K36R(86Fb)/ +

  • H3K27R: w hs-flp; Df(2L)HisC FRT40A/M(2)25A ubi-GFP FRT40; 3xHisGUH3K27R(68E) 3xHisGUH3K27R (86Fb)/ +

  • Figure 6—figure supplement 1

  • wt: w hs-flp; hs--nGFP FRT2A/hs nGFP FRT2A

  • H3K36R: Df(2L) HisC FRT40/Df(2L) HisC FRT40; 12xHisGUH3K36R(VK33)/TM6B

  • H3K27R: w hs-flp; Df(2L)HisC FRT40A/hs-nGFP FRT40A; 3xHisGUH3K27R(68E)3xHisGUH3K27R(86Fb)/3xHisGUH3K27R(68E)3xHisGUH3K27R(86Fb)

Comparison of the lethality of H3K36R and H3K36A mutants

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The difference in the lethality phase of the H3K36R mutants generated in this study compared to H3K36R mutants in the strain from Matera and colleagues was unexpected because both strains the Df(2L)HisC homozygotes carry 12 copies of the HisGUH3K36R cassette (i.e. four 3x HisGUH3K36R arrays in our strain and a single 12x HisGUH3K36R array in the strain from Matera and colleagues). A possible explanation for the poorer survival of H3K36R mutants in the strain generated here could be that histone transgene expression from the 3xHisGUH3K36R miniarrays is for some reason less effective that in the case of the 12x HisGUH3K36R array. We also note that a recent study reported that among Df(2L)HisC homozygotes that carry 20 HisGUH3K36A copies, about 50% of the mutant animals develop up to the pupal stages (Zhang et al., 2019). Zhang et al have not analyzed their H3K36A mutants any further but it is possible that the higher copy number of the HisGUH3K36A cassette accounts for the better survival compared to the H3K36A strain generated in this study. Further studies will be needed to explore whether H3K36R and H3K36A mutants show comparable phenotypes in larvae.

Immunohistochemistry and immunofluorescence stainings

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Embryos of the appropriate genotypes listed above were identified by the lack of GFP marked balancer chromosomes, fixed and stained with Abd-B antibody, following standard protocols. Imaginal discs from third instar larvae were stained with Antp and Cy3-labeled secondary antibodies following standard protocols. For clonal analysis (Figure 3D), clones were induced 96 hr before analyses by heat-shocked induced expression of Flp recombinase in the genotypes listed above.

ChIP-seq analysis in Drosophila embryos and in larval tissues

Embryo collection, chromatin preparation, and ChIP

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21-24 hr old wt, H3K36A embryos (see above for details of genotypes) were dechorionated, quick-frozen in liquid N2 and stored at -80°C. 5 µl of thawed embryos were homogenized in 5 mL of fixing solution (60 mM KCl, 15 mM NaCl, 4 mM MgCl2, 15 mM Hepes pH 7.6, 0.5% Triton X-100, 0.5 mM DTT, protease inhibitors, 0.9% Formaldehyde) at r.t. The homogenate was filtered through a strainer (Greiner Bio-One, EASYstrainer 100 µm, #542 000) and incubated for 10 min with frequent gentle shaking. Cross-linking was stopped by the addition of 450 µl of 2.5 M Glycine. Fixed nuclei were washed with 1 ml of buffer A1 (60 mM KCl, 15 mM NaCl, 4 mM MgCl2, 15 mM Hepes pH 7.6, 0.5% Triton X-100, 0.5 mM DTT, protease inhibitors), washed with 1 ml of pre-lysis buffer (140 mM NaCl, 15 mM Hepes pH 7.6, 1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.5 mM DTT, 0.1% Na Deoxycholate, protease inhibitors), resuspended in 1 ml of lysis buffer (140 mM NaCl, 15 mM Hepes pH 7.6, 1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.5 mM DTT, 0.1% Na Deoxycholate, protease inhibitors, 0.1% SDS, 0.5% N-laurylsarcosine), incubated at least 10 min at 4°C with shaking, and transferred into milliTUBES 1 ml AFA Fiber (100) (Covaris, #520130) for sonication. Sonication was performed in a Covaris S220 AFA instrument using the following setup: 140W (peak incident power) / 5% (duty cycle) / 200 (cycle per burst) / 15 min. Insoluble material was removed by centrifugation in an Eppendorf centrifuge at 14000 rpm (10 min at 4°C). Input chromatin was quantified by measuring DNA concentration after decrosslinking using Qubit (Thermo Scientific) and 250 ng of chromatin were used for each ChIP experiment. 250 ng of an independently prepared batch of D. pseudoobscura chromatin were spiked-in in each ChIP experiment for subsequent normalization of the ChIP-seq datasets. The rest of the ChIP protocol was performed as described in Bonnet et al., 2019. For each condition, the ChIP experiment was performed in duplicates from two biologically independent chromatins. ChIP on hand-dissected CNS and imaginal disc tissues from 3rd instar wt or H3K36R homozygous larvae (see above for details on genotypes) was performed as described Laprell et al., 2017 with the difference D. pseudoobscura chromatin was spiked in at a 1:1 ratio of dm / dp chromatin.

Library preparation and sequencing

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Library preparation for sequencing was performed with TruSeq kits from Illumina. Illumina systems (NextSeq 500) were used for paired-end DNA sequencing. All reads were aligned using STAR (Dobin et al., 2013) to the D. melanogaster dm6 genome assembly (dos Santos et al., 2015) and to the D. pseudoobscura dp3 genome assembly (Nov. 2004, FlyBase Release 1.03). Only sequences that mapped uniquely to the genome with a maximum of two mismatches were considered for further analyses.

Identification of H3K36me2 and H3K27me3 enriched regions

The Bioconductor STAN-package (Zacher et al., 2017) was used to define the location of H3K36me2-enriched regions. The seven chromosome arms (X, 2L, 2R, 3L, 3R, 4 and Y) defined in the dm6 genome assembly were segmented in 200 bp bins. STAN annotated each of these bins into 1 of 3 ‘genomic states’ based on the number of H3K36me2 ChIP-seq reads and the number of input reads overlapping with each bin in wildtype embryos or larvae. These 3 ‘genomic states’ corresponded to: ‘H3K36me2 enriched’ regions; ‘low or no H3K36me2’ regions and ‘no input’ regions. The Poisson Lognormal distribution was selected and fitting of hidden Markov models was performed with a maximum number of 100 iterations. Stretches of consecutive bins annotated as ‘H3K36me2 enriched’ regions were sometimes separated by a few bins showing another type of annotation (i.e. ‘no input’). To define a relevant set of H3K36me2 enriched regions, we considered that if stretches of consecutive bins annotated as ‘H3K36me2 enriched’ regions are not separated by more than 7 Kb, they can be fused. High-level H3K27me3 domains previously defined using the same Bioconductor STAN-package in Bonnet et al., 2019 were used in this study.

Normalization and visualisation of H3K27me3 and H3K36me2 ChIP-Seq datasets

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The proportion of D. pseudoobscura reads as compared to D. melanogaster reads in input and in samples was used to normalize the H3K36me2 and H3K27me3 ChIP-seq datasets from H3K36A and H3K36R mutants to the corresponding wildtype H3K36me2 and H3K27me3 ChIP-seq datasets respectively (see Supplementary file 2). Chip-seq tracks shown in Figure 4 show the average of the two replicates that were performed for each condition. Y-axes of ChIP-seq tracks correspond to normalized numbers of mapped reads per million reads per 200 bp bin.

Calculation of read coverage

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In wildtype and H3K36A and H3K36R mutant conditions, H3K36me2 and H3K27me3 ChIP-seq read coverages across gene bodies were computed on genomic intervals starting 750 bp upstream transcription start sites and ending 750 bp downstream transcription termination sites. Read coverage is defined as the normalized number of mapped reads per million reads from a ChIP-seq dataset divided by the number of mapped reads per million reads from the corresponding input dataset across a genomic region. Among the D. melanogaster Refseq genes, approximately 10800 and 9200 are overlapping with H3K36me2 enriched regions, approximately 1030 and 1030 genes are overlapping with high-level H3K27me3 domains and 5400 and 6300 are localized in other genomic regions in embryos and larvae, respectively.

Drosophila nuclear and cell extracts for western blot analysis

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For embryonic total nuclear extracts, nuclei from 21 to 24 hr old wt, H3K36A or H3K36A mutant embryos were purified and quantified as described in Bonnet et al., 2019. Pellets of nuclei were resuspended in appropriate volumes of SDS sample buffer proportional to the number of nuclei in each pellet. Extracts were then sonicated in a Bioruptor instrument (Diagenode) (eight cycles (30 s ON/30 s OFF), high power mode), incubated at 75 °C for 5 min and insoluble material was removed by centrifugation at 14000 rpm for one mn at r.t.

Total cell extracts from imaginal disc tissues were prepared by resuspending hand-dissected disc tissues in SDS sample buffer. Extracts were then sonicated, incubated at 75 °C for 5 min and insoluble material was removed by centrifugation.

Antibodies

For ChIP analysis:
Rabbit monoclonal anti-H3K27me3Cell Signaling Technology #9733
Rabbit polyclonal anti-H3K36me2Abcam #9049
For western blot analysis on embryonic and larval extracts:
Rabbit monoclonal anti-H3K27me3Cell Signaling Technology #9733
Rabbit polyclonal anti- H3K27me3Millipore #07-449
Rabbit polyclonal anti-H3K27me1Millipore #07-448
Rabbit monoclonal anti-H3K36me3Cell Signaling Technology #4909
Rabbit monoclonal anti-H3K36me2Cell Signaling Technology #2901
Rabbit polyclonal anti-H2B(against full-length recombinant D.m. H2B)
Rabbit polyclonal anti-H4Abcam #10158
Rabbit polyclonal anti-Caf1Gambetta et al., 2009
For immunohistochemistry and immunofluorescence analysis:
Mouse monoclonal anti-Abd-BDSHB (1A2E9)
Mouse monoclonal anti-AntpDSHB (8C11)
Rabbit monoclonal anti-H3K27me3Cell Signaling Technology #9733

Data availability

The sequence datasets generated in this study have been deposited in GEO (accession number: GSE148254). The protein structure data reported in this study have been deposited in PDB under the accession code 7AT8 and in the EMDB under the accession codes EMD-11910 and EMD-11912.

The following data sets were generated
    1. Müller J
    2. Bonnet J
    (2020) NCBI Gene Expression Omnibus
    ID GSE148254. Inhibition of PRC2 activity by H3K36 methylation.
    1. Finogenova K
    2. Benda C
    3. Schäfer IB
    4. Poepsel S
    5. Strauss M
    6. Müller J
    (2020) RCSB Protein Data Bank
    ID 7AT8. Histone H3 recognition by nucleosome-bound PRC2 subunit EZH2.
    1. Finogenova K
    2. Benda C
    3. Schäfer IB
    4. Poepsel S
    5. Strauss M
    6. Müller J
    (2020) Electron Microscopy Data Bank
    ID EMD-11910. Histone H3 recognition by nucleosome-bound PRC2 subunit EZH2.
    1. Finogenova K
    2. Benda C
    3. Schäfer IB
    4. Poepsel S
    5. Strauss M
    6. Müller J
    (2020) EMDataBank
    ID EMD-11912. Cryo-EM map of PHF1-PRC2 on a heterodimeric dinucleosome.

References

    1. Beuchle D
    2. Struhl G
    3. Müller J
    (2001)
    Polycomb group proteins and heritable silencing of Drosophila Hox genes
    Development 128:993–1004.
    1. Emsley P
    2. Lohkamp B
    3. Scott WG
    4. Cowtan K
    (2010) Features and development of coot
    Acta Crystallographica. Section D, Biological Crystallography 66:486–501.
    https://doi.org/10.1107/S0907444910007493

Decision letter

  1. Cynthia Wolberger
    Senior Editor; Johns Hopkins University School of Medicine, United States
  2. Jerry L Workman
    Reviewing Editor; Stowers Institute for Medical Research, United States
  3. Jerry L Workman
    Reviewer; Stowers Institute for Medical Research, United States

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

Acceptance summary:

This manuscript provides a structural explanation for how histone H3K36 methylation, an active transcription mark, prevents H3K27 methylation by Polycomb complexes which repress gene expression. These results provide substantial insights into crosstalk among histone modifications.

Decision letter after peer review:

Thank you for submitting your article "Structural basis for PRC2 decoding of active histone methylation marks H3K36me2/3" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Jerry L Workman as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Cynthia Wolberger as the Senior Editor.

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

As the editors have judged that your manuscript is of interest, but as described below that additional experiments are required before it is published, we would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). First, because many researchers have temporarily lost access to the labs, we will give authors as much time as they need to submit revised manuscripts. We are also offering, if you choose, to post the manuscript to bioRxiv (if it is not already there) along with this decision letter and a formal designation that the manuscript is "in revision at eLife". Please let us know if you would like to pursue this option. (If your work is more suitable for medRxiv, you will need to post the preprint yourself, as the mechanisms for us to do so are still in development.)

Summary:

In this manuscript, the authors determine cryo-EM structures of human PRC2 on dinucleosomes. Several such structures have been published in recent years. The novel aspects presented in this study, are focused on DNA-interacting interphases of EZH2 and EED as well as a cis-acting role of unmodified H3K36 in facilitating efficient H3K27 methylation by PRC2.

H3K36me2/3 has been proposed to influence PRC2 function in several ways: H3K36me3 has been shown to inhibit PRC2 catalytic activity in vitro, but has also been suggested as a potential recruitment mechanism to transcribed chromatin via recognition by the Tudor domains of PCL proteins. Here, the authors demonstrate, through in vitro and in vivo approaches, that the substitution of H3K36 with H3K36R or H3K36A, and therefore the absence of an unmodified H3K36, leads to global reduction of H3K27me3. Moreover, they show that PRC2 binds to mononucleosomes with H3K36me3, however, that they are poor PRC2 substrates. Taken together with the presented cryo-EM data, the authors provide a plausible mechanism for the inhibitory effects of H3K36me2/me3 on PRC2 activity.

Overall, this is a well-performed study, which addresses an open question in the field. The manuscript is well-written and the conclusions are generally well-supported – however, some additional experiments and alternative interpretations should be taken into consideration and incorporated in the manuscript (see specific comments below).

Essential revisions:

1) One of the primary conclusions reached by the authors is that the cleft which H3K36 binds at the EZH2-nucleosomal DNA interface does not fit H3K36A and H3K36R mutations or methylated forms of H3K36 (i.e., H3K36me2 and H3K36me3), leading to allosteric inhibition of PRC2. The structure of the nucleosome-PRC2 complex (Figure 1—figure supplement 4E) illustrates that the side chain of K36 engages in few interactions in this cleft and does not contact the nucleosomal DNA. Furthermore, there is a relatively large unoccupied space in the cleft between the K36 side chain and the DNA, suggesting that amino acids larger than lysine can fit within it. Modeling of an K36R mutation indicates that the arginine side chain can be accommodated in the K36 binding cleft without clashes. Given their similar size, methylated forms of K36 could also fit within the H3K36 binding cleft. The claim that the cleft at the EZH2-DNA interface fits lysine (K36) but cannot accommodate larger amino acids is unconvincing in the absence of additional data.

2) The mechanism by which H3K36 binding at the EZH2-nucleosomal DNA interface allosterically regulates PRC2 enzymatic activity is not clearly established. The nucleosome-PRC2 structure shows one potential polar interaction between the K36 side chain and the backbone of Gln570 in EZH2, implying a lack of specificity in recognizing K36. In contrast, other proteins that specifically recognize unmodified lysines in the histone H3 N-terminal tail have been shown to do so through a network of hydrogen bonds and/or salt bridge interactions (e.g., H3K4 recognition by the BHC80 PHD domain; Lan et al. (2007) Nature 448, 718-22). It is unlikely that a single (potential) polar interaction allosterically regulates PRC2 methyltransferase activity.

3) The authors attribute the loss of H3K27me3 in the Drosophila H3K36A and H3K36R mutants to allosteric inhibition of PRC2. However, these mutations also diminish H3K36me3 in developing Drosophila embryos and larvae. The Drosophila PRC2 accessory subunit Pcl has been shown to bind H3K36me3, analogous to its mammalian homolog PHF1 (Ballare et al., 2012; Musselman et al., 2012). Thus, the global loss of H3K36me3 may result in either diminished recruitment or mislocalization of PRC2 in chromatin. The authors should examine whether the H3K36A and H3K36R mutations affect PRC2 localization within chromatin relative to wild type Drosophila.

4) The authors use full length human PRC2 in a complex with PHF1 for the Cryo-EM studies and in nucleosome binding experiments. In the Cryo-EM studies, they are not able to detect the density for the N-terminal part of SUZ12, RBBP4 and the N-terminal part of PHF1. In several publications these parts of the PRC2 complex have been shown to be important for the localization of PRC2 to specific sites in the genome. Although, the authors do not test this directly, the N-terminal part of SUZ12 and RBBP4 are most likely important for the binding (affinities) that the authors measure in EMSA. If this indeed is correct, the authors should take this into their discussion of the relative contribution of the various parts of PRC2 to DNA and nucleosome binding affinities.

5) The description in the text states that PRC2CXC>A and PRC2EED>A have similar binding strength deficits in the EMSA assays, which is also shown in the quantification (Figure 2B) – however, based on the gels presented (Figure S5A), the EED mutant appears to perform similarly to the PRC2 WT? Either it is a poorly chosen example (of the three replicates used for quantification) or we would question whether the gel image is of sufficient quality for this quantification.

6) The EMSA experiments (Figure 2A and S5A) are performed solely on mononucleosomes, presumably meaning that only one of the studied PRC2 binding surfaces (CXC vs EED) can be engaged at a time. In this scenario, mutation of one binding surface presumably would allow the other surface to "take over". Given the different enzymatic activities observed for PRC2CXC>A on mono- vs. di-nucleosomes, it would be relevant to know how well this mutant is binding di-nucleosomes. Alternatively, that the authors discuss this aspect in the manuscript.

7) Overall, one limitation in this experimental approach, is that we cannot adequately distinguish between the effect of mutating the specific sites vs. how the mutations may confer overall conformational changes in PRC2 contributing to the altered binding strength. Furthermore, the concluding statement does not take into consideration that the introduction of mutations in the CXC domain may well alter the conformation of EZH2 (or core PRC2) in a way that directly affects the catalytic activity – so loss of H3K27me3 in Figure 2C might also stem from an overall reduction in enzymatic activity rather than (only) from reduced nucleosome binding through this specific surface, especially given the rather modest impact on binding strength shown in Figure 2A. To distinguish between effects on overall catalytic activity and reduced binding, the authors could perform in vitro methyltransferase assays on peptide substrates (as in Figure 3D), using the CXC domain mutant.

8) The interpretations of Figures 4A vs 4B are difficult to follow from the presented data: the Western blot presented in Figure 4B does not strongly support the conclusion of a 3-4-fold global reduction of H3K36me2/me3 (the bands are quite “washed out” in lanes 5-8). The authors further state that the reduction in H3K36me2/me3 is less pronounced in H3K36A embryos vs. H3K36R larvae – and that this might represent methylated H3.3 (which would presumably also be the case for H3K36R), but also maternally-deposited wildtype canonical H3. To substantiate this speculation, the authors should compare the different genotypes in the same developmental stages – thus, a meaningful comparison should include H3K36R mutant embryos (since the H3K36A mutants die before the larval stage).

9) Similarly, it would be interesting to see the ChIP data performed in H3K36R embryos to better evaluate whether the apparently absent (by WB, Figure 4B) or very modest (by ChIP, Figure 4D) reduction of H3K27me3 in H3K36A mutant embryos represents a mechanistic difference between H3K36A vs. H3K36R or simply a difference in developmental stages.

10) How do the authors explain/speculate about the absent/modest effect on H3K27me3 in H3K36A mutant embryos despite the severity of the phenotype for these mutants? Here, again, a side-by-side comparison of H3K27me3 levels in H3K36R and H3K36A mutant embryos would be relevant. It would also be relevant to factor in knowledge about the number of cell divisions and maternal histone load in order to estimate if H3K36A is also impacting H3K27 methylation in vivo.

11) The statement in the Figure 4E legend that "For each HOX gene there is a substantially larger proportion of cells in which the gene is decorated with H3K27me3 and repressed by Polycomb" is probably not correct: While the analyzed tissues are surely a mixed population with few cells showing expression of the tested HOX genes (Figure 5), it is unlikely that the larger proportion of genes show normal H3K27me3 – if this were the case, the ChIP data in Figure 4 would not show the large reduction in H3K27me3 at the tested loci. Thus, the data likely reflects that H3K27me3 is reduced in most of the cell types – yet the de-repression is only seen in a few cells – to show this, immunostaining for H3K27me3 should be included in Figure 5 (discussed further below).

12) The authors state that "the most straightforward interpretation" of the observed misexpression of Polycomb target genes in the H3K36R larvae is that it results from reduced H3K27me3 at these loci. While this may well be the case in this study, loss of H3K27me3 can also be secondary to gene activation – a distinction that is overlooked in many studies. In this case, the question is whether loss of H3K27me3 is a direct effect of the H3K36 substitution or secondary to gene expression changes or altered/delayed development. In this context, the authors could draw more strongly on the previous observation (McKay et al., 2015; Meers et al., 2017) that the H3K36R mutant does not show massive deregulation of gene expression (as mentioned in the Discussion), and, most importantly, should emphasize their own observations that global H3K27me3 is reduced (Figure 4A) and H3K27me3 at Polycomb target genes is strongly reduced (Figure 4C) despite the low percentage of cells assayed showing misexpression of these particular genes (Figure 5B-D). Along with the in vitro data presented in Figure 2, this provides stronger evidence for a direct effect of an “intact” H3K36 in preserving normal PRC2 activity.

13) Along the same lines, it would be interesting to include immunostaining for H3K27me3 in Figures 5B-D to learn whether the loss of H3K27me3 is more prominent in the cells showing mis-expression and to substantiate the data from Figure 4 showing global reduction of H3K27me3.

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

Thank you for submitting your article "Structural basis for PRC2 decoding of active histone methylation marks H3K36me2/3" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Jerry L Workman as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Cynthia Wolberger as the Senior Editor.

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

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, we are asking editors to accept without delay manuscripts, like yours, that they judge can stand as eLife papers without additional data, even if they feel that they would make the manuscript stronger. Thus the revisions requested below only address clarity and presentation.

The authors' revisions have addressed the majority of the comments on the original manuscript, and the additional discussion has clarified their model for how H3K36 interactions with PRC2 may influence its activity. However, there is a remaining point that should be addressed:

The authors' assertion that Drosophila Pcl does not bind H3K36me2 or H3K36me3 is not entirely correct. Although the Tudor domain of Pcl does not recognize H3K36me2/3, Ballare et al. demonstrated that a Pcl Tudor-PHD1 construct does bind to H3K36me2 and H3K36me3 (Ballare et al., 2012). This finding is also explicitly stated in their article: "Nevertheless, using a Pcl Tudor-PHD1 construct, we were able to rescue the binding to H3K36me2 and H3K36me3 (Figure 2B)." Furthermore, Musselman et al., 2012, investigated the interactions between the human PHF1 Tudor domain and H3K36me3 but did not test the binding of the Drosophila Pcl Tudor-PHD1 domains to H3K36me2 or H3K36me3 (Musselman et al., 2012). In sum, the authors should amend their manuscript to acknowledge that the Drosophila Pcl Tudor-PHD1 domains have been shown to bind to H3K36me2 and H3K36me3.

https://doi.org/10.7554/eLife.61964.sa1

Author response

Essential revisions:

1) One of the primary conclusions reached by the authors is that the cleft which H3K36 binds at the EZH2-nucleosomal DNA interface does not fit H3K36A and H3K36R mutations or methylated forms of H3K36 (i.e., H3K36me2 and H3K36me3), leading to allosteric inhibition of PRC2. The structure of the nucleosome-PRC2 complex (Figure 1—figure supplement 4E) illustrates that the side chain of K36 engages in few interactions in this cleft and does not contact the nucleosomal DNA. Furthermore, there is a relatively large unoccupied space in the cleft between the K36 side chain and the DNA, suggesting that amino acids larger than lysine can fit within it. Modeling of an K36R mutation indicates that the arginine side chain can be accommodated in the K36 binding cleft without clashes. Given their similar size, methylated forms of K36 could also fit within the H3K36 binding cleft. The claim that the cleft at the EZH2-DNA interface fits lysine (K36) but cannot accommodate larger amino acids is unconvincing in the absence of additional data.

2) The mechanism by which H3K36 binding at the EZH2-nucleosomal DNA interface allosterically regulates PRC2 enzymatic activity is not clearly established. The nucleosome-PRC2 structure shows one potential polar interaction between the K36 side chain and the backbone of Gln570 in EZH2, implying a lack of specificity in recognizing K36. In contrast, other proteins that specifically recognize unmodified lysines in the histone H3 N-terminal tail have been shown to do so through a network of hydrogen bonds and/or salt bridge interactions (e.g., H3K4 recognition by the BHC80 PHD domain; Lan et al. (2007) Nature 448, 718-22). It is unlikely that a single (potential) polar interaction allosterically regulates PRC2 methyltransferase activity.

We agree with the comment that the side chain of an arginine or a tri-methylated lysine or an alanine could theoretically all be accommodated in the cleft between the EZH2CXC domain and the DNA. Our wording in parts of the text that we used in the original manuscript (i.e. the implication that bulkier side chains inhibit) was not well chosen and we replaced the first sentence in the section “Unmodified H3K36 in the EZH2CXC-DNA interaction interface is critical for H3K27 methylation in nucleosomes“ by writing “The placement of H3K36 in the EZH2CXC-DNA interface (Figure 1F) suggested that even though a tri- or di-methylated K36 side chain could theoretically be accommodated in this interface, these modified side chains might provide a less optimal fit and thereby inhibit H3K27 methylation.”

The reviewer wrote: “K36 engages in few interactions in this cleft and does not contact the nucleosomal DNA.” We are certainly aware that electrostatic interactions between amino acids in proteins are usually considered to require distances of less than 4Å. However, we would like to be cautious and would prefer not to exclude possible (long-range) electrostatic interactions of the K36 side chain with the phosphate backbone of the nucleosomal DNA. First, it should be kept in mind that the global resolution of our structure is in the range of 4.4 Å. Second, what may appear as a “static geometry” in the structure, is certainly not static in solution and the binding geometry may well change the angle of the EZH2 DNA interaction slightly, depending on length of the nucleosomal linker DNA, the sequence of the DNA around the histone octamer etc. We changed the sentence in the section where we describe the structure to include the word “possibly”, i.e. it now reads: “The side chain density of H3K36 suggests that the epsilon-amino group of H3K36 engages in a polar interaction with the carbonyl group of Q570 and possibly in long-range electrostatic interactions with the phosphate backbone of the nucleosomal DNA (Figure 1F, Figure 1—figure supplement 4 C-E).”

It should be emphasized that it is DNA binding that is the driving force for the interaction of PRC2 with nucleosomes (Choi et al., 2017; Wang et al., 2017). The contribution of histone tail interactions to the nucleosome-binding is not really measurable because it is in a completely different range of binding affinity. The Kd of PRC2 for binding to nucleosomes is about 60 nM. The Kd of PRC2 for binding to an unmodified H3[21-41] tail peptide is 3 orders of magnitude higher (29 µM) and for binding to the matching H3K36me3 peptide it is 18 µM (Jani et al., 2019).

We found that PRC2 binds unmodified and H3K36me3-modified nucleosomes with comparable affinity (Figure 3) but that H3K27 methylation on H3Kc36me3 nucleosomes is about 10- to 20-fold less effective than on unmodified nucleosomes. In other words, K27 on K36me3-modified H3 must still be able to engage in a productive manner with the catalytic center, but it is possibly the frequency of productive interactions that is 10- to 20-fold lower. The same is of course true for H3K36R or H3K36A where the inhibition of H3K27me3 formation is about 5-fold (i.e. as shown in Figure 3). So while we agree that H3K36me3, H3K36R or H3K36A can likely be accommodated in the same cleft like H3K36, we however imagine that the residence time of this interaction is diminished and, hence, H3K27 engages with the active site at lower frequency.

In summary, we think that a possible scenario could be that the presence of an unmodified native lysine side chain at position 36 of histone H3 is important within the dynamics of the reaction cycle which involves (i) docking of the complex on the nucleosome via EZH2CXC-DNA contacts, (ii) alignment of the H3 N-terminus on the EZH2 surface (iii) docking of the H3K27 side into the catalytic center for methylation of its side chain. This is what we proposed in the Discussion section: “Taken together, a possible scenario would therefore be that within the time frame of the PRC2-nucleosome binding and reaction cycle, docking of the H3K36 side chain in the EZH2CXC-DNA interface is critical for rapid alignment of the H3 N-terminus on the EZH2 surface into a catalytically competent state.”

3) The authors attribute the loss of H3K27me3 in the Drosophila H3K36A and H3K36R mutants to allosteric inhibition of PRC2. However, these mutations also diminish H3K36me3 in developing Drosophila embryos and larvae. The Drosophila PRC2 accessory subunit Pcl has been shown to bind H3K36me3, analogous to its mammalian homolog PHF1 (Ballare et al., 2012; Musselman et al., 2012). Thus, the global loss of H3K36me3 may result in either diminished recruitment or mislocalization of PRC2 in chromatin. The authors should examine whether the H3K36A and H3K36R mutations affect PRC2 localization within chromatin relative to wild type Drosophila.

The statement “The Drosophila PRC2 accessory subunit Pcl has been shown to bind H3K36me3, analogous to its mammalan homolog PHF1 (Ballare et al., 2012; Musselman et al., 2012).” is incorrect. The structure of the tudor domain of Drosophila Pcl was determined in 2010 and revealed that Pcl tudor lacks an aromatic cage and does not bind to methylated lysines or arginines (Friberg et al., 2010). The Musselman et al. and Ballare et al. studies in 2012 showed that while the PHF1 tudor domain does bind to H3K36me3 peptides, the tudor domain of Drosophila Pcl does not bind H3K36me3 (Ballare et al., 2012, Figure 2B) and that mutation of two aromatic residues in the PHF1 tudor domain that are absent in the Pcl tudor domain abolishes H3K36me3-binding by the PHF1 tudor domain (Musselman et al., 2012).

To remind the reader out this difference, we have added a sentence at the end of the paragraph where we describe the results of HMTase assays with PHF1-PRC2 on H3Kc36me3 nucleosomes:

“In this context it should also be noted that in Polycomblike, the Drosophila ortholog of PHF1, the tudor domain contains an incomplete aromatic cage and is unable to bind methylated lysines (Friberg et al., 2010; Musselman et al., 2012; Ballare et al., 2012).”

It should also be kept in mind that, in Drosophila, PRC2 is recruited to Polycomb Response Elements (PREs). PRC2 targeting to PREs requires Pho (Wang et al., Mol Cell 2004; Frey et al., G&D 2006) and Pcl (Nekrasov et al., 2007; Savla et al., Development 2008). Moreover, PREs are depleted of nucleosomes (Papp and Müller, G&D 2006; Schwartz et al. JBC 2006 and many more studies since). So there is little foundation for a scenario where H3K36me2/3 would be involved in targeting PCl-PRC2.

The reviewer requested to generate PRC2 binding profiles in Drosophila with H3K36R or H3K36A chromatin. With the recent worsening of the pandemic situation in Germany, we have not been able to collect the large amount of imaginal disc material that would be needed for perform ChIP-seq analyses for comparing genome-wide binding profiles of PRC2 subunit in wild-type and H3K36R mutant larvae. It is unlikely that we will be able to get that experiment done anytime soon. Considering this and considering that we are in a competitive situation with a manuscript from Eva Nogales’ lab that also reports the H3 binding mode and the role of H3K36, we would ask the reviewers not to insist on including this experiment.

4)The authors use full length human PRC2 in a complex with PHF1 for the Cryo-EM studies and in nucleosome binding experiments. In the Cryo-EM studies, they are not able to detect the density for the N-terminal part of SUZ12, RBBP4 and the N-terminal part of PHF1. In several publications these parts of the PRC2 complex have been shown to be important for the localization of PRC2 to specific sites in the genome. Although, the authors do not test this directly, the N-terminal part of SUZ12 and RBBP4 are most likely important for the binding (affinities) that the authors measure in EMSA. If this indeed is correct, the authors should take this into their discussion of the relative contribution of the various parts of PRC2 to DNA and nucleosome binding affinities.

This is a helpful comment. We extended our discussion of this issue. The text now reads:

“Nucleosome binding by the PRC2CXC>A/EED>A complex could in part be due to incomplete disruption of the mutated interfaces but it likely also represents nucleosome binding contributed by the bottom lobe of PRC2 comprising the N-term of SUZ12 and RBBP4 (Chen et al., 2018; Nekrasov et al., 2005). […] In conclusion, the structural data (Figure 1C, D and Poepsel et al., 2018) suggest that a key interaction of PRC2 with substrate nucleosomes occurs through contacts of the EZH2CXC domain with the DNA gyres, whereas the biochemical analyses argue that this interaction contributes only modestly to the overall chromatin-binding affinity of the complex.”

5) The description in the text states that PRC2CXC>A and PRC2EED>A have similar binding strength deficits in the EMSA assays, which is also shown in the quantification (Figure 2B) – however, based on the gels presented (Figure S5A), the EED mutant appears to perform similarly to the PRC2 WT? Either it is a poorly chosen example (of the three replicates used for quantification) or we would question whether the gel image is of sufficient quality for this quantification.

We agree that the gel in Figure S5A comparing nucleosome binding by PRC2EED>A and the wild-type PRC2 control next to it was not the nicest gel but it was the only gel where we had run the wild-type control directly next to the mutant.

We have removed Figure S5A and now show the EMSA with PRC2EED>A in Figure 2A. The gel image of the PRC2EED>A EMSA (lanes 31-40) has the same experimental setup as in the left panel (lanes 1-30) but was run on a separate gel.

6) The EMSA experiments (Figure 2A and S5A) are performed solely on mononucleosomes, presumably meaning that only one of the studied PRC2 binding surfaces (CXC vs EED) can be engaged at a time. In this scenario, mutation of one binding surface presumably would allow the other surface to "take over". Given the different enzymatic activities observed for PRC2CXC>A on mono- vs. di-nucleosomes, it would be relevant to know how well this mutant is binding di-nucleosomes. Alternatively, that the authors discuss this aspect in the manuscript.

We had performed EMSAs with the aim to analyze binding of PRC2 to dinucleosomes. However, we observed a complex mixture of slowly migrating species and this has precluded further experiments aimed at discriminating between binding events involving specific PRC2 surfaces using dinucleosomes as binding substrates.

We discuss this aspect more extensively in the revised version of the manuscript:

Nucleosome binding by the PRC2CXC>A/EED>A complex could in part be due to incomplete disruption of the mutated interfaces but it likely also represents nucleosome binding contributed by the bottom lobe of PRC2 comprising the N-term of SUZ12 and RBBP4 (Chen et al., 2018; Nekrasov et al., 2005). In particular, biochemical studies on Drosophila PRC2 originally found that a minimal complex formed between Su(z)12 and the RBBP4 ortholog Caf1-55 binds to nucleosomes (Nekrasov et al., 2005). Moreover, negative stain EM analyses of human PRC2 bound to a dinucleosome identified several 2D classes where the bottom lobe contacts one or two nucleosomes of the dinucleosome (Poepsel et al., 2018). The binding affinity of the PRC2 core complex for chromatin therefore appears to result from interactions of at least three distinct complex surfaces with nucleosomes, the EZH2CXC domain, the EED nucleosome-binding interface and the SUZ12N:RBBP4 lobe. Considering the architecture of the catalytic lobe (Figure 1C) and of the isolated full PRC2 core complex (Kasinath et al., 2018), it is very unlikely that the EZH2CXC domain and the EED nucleosome-binding interface could simultaneously engage with the same nucleosome at a time. Finally, we note that in EMSAs monitoring binding of PRC2 to a dinucleosome, we observed a complex mixture of slowly migrating species and this has precluded experiments aimed at discriminating between binding events involving specific PRC2 surfaces with dinucleosomes. In conclusion, the structural data (Figure 1C, D and Poepsel et al., 2018) suggest that a key interaction of PRC2 with substrate nucleosomes occurs through contacts of the EZH2CXC domain with the DNA gyres, whereas the biochemical analyses argue that this interaction contributes only modestly to the overall chromatin-binding affinity of the complex.

7) Overall, one limitation in this experimental approach, is that we cannot adequately distinguish between the effect of mutating the specific sites vs. how the mutations may confer overall conformational changes in PRC2 contributing to the altered binding strength. Furthermore, the concluding statement does not take into consideration that the introduction of mutations in the CXC domain may well alter the conformation of EZH2 (or core PRC2) in a way that directly affects the catalytic activity – so loss of H3K27me3 in Figure 2C might also stem from an overall reduction in enzymatic activity rather than (only) from reduced nucleosome binding through this specific surface, especially given the rather modest impact on binding strength shown in Figure 2A. To distinguish between effects on overall catalytic activity and reduced binding, the authors could perform in vitro methyltransferase assays on peptide substrates (as in Figure 3D), using the CXC domain mutant.

This is a good control and we have now added this experiment in the revised version. PRC2CXC>A did not show reduced K27 methylation activity compared to wild-type PRC2 on the H3 peptide substrate. The new data are shown in Figure 2—figure supplement 1B and the results are discussed in the text:

“To complement these experiments, we also compared the HMTase activity of wild-type PRC2 and PRC2CXC>A complex on free histone H318-42 peptides using a masspectrometry-based assay to detect H3K27 methylation. It is well known that wild-type PRC2 methylates K27 on free histone H3 with much lower efficiency than on H3 in nucleosomes (Cao et al., 2002; Czermin et al., 2002; Kuzmichev et al., 2002; Muller et al., 2002), and this is also recapitulated in our assays on H318-42 peptides where we primarily detect H3K27me1 but no H3K27me3 formation, even after extended incubation of the reaction (see Figure 2—figure supplement 1B and compare with Figure 2C). However, it is important to note that PRC2CXC>A did not show reduced K27 methylation activity compared to wild-type PRC2 on this H3 peptide substrate (Figure 2—figure supplement 1B). The mutations in the EZH2CXC domain therefore do not appear to alter the conformation of EZH2 in a way that would directly interfere with catalysis.”

8) The interpretations of Figures 4A vs 4B are difficult to follow from the presented data: The Western blot presented in Figure 4B does not strongly support the conclusion of a 3-4-fold global reduction of H3K36me2/me3 (the bands are quite “washed out” in lanes 5-8). The authors further state that the reduction in H3K36me2/me3 is less pronounced in H3K36A embryos vs. H3K36R larvae – and that this might represent methylated H3.3 (which would presumably also be the case for H3K36R), but also maternally-deposited wildtype canonical H3. To substantiate this speculation, the authors should compare the different genotypes in the same developmental stages – thus, a meaningful comparison should include H3K36R mutant embryos (since the H3K36A mutants die before the larval stage).

9) Similarly, it would be interesting to see the ChIP data performed in H3K36R embryos to better evaluate whether the apparently absent (by WB, Figure 4B) or very modest (by ChIP, Figure 4D) reduction of H3K27me3 in H3K36A mutant embryos represents a mechanistic difference between H3K36A vs. H3K36R or simply a difference in developmental stages.

We originally had actually analyzed bulk levels of H3K36me2/me3 and H3K27me3 in H3K36R mutant embryos in parallel with the analysis in H3K36A mutant embryos (run the same gel) and we had also performed the H3K36me2 and H3K27me3ChIP-seq analyses in H3K36R mutant embryos. See https://www.biorxiv.org/content/10.1101/2020.04.22.054684v1.full.pdf). We had then performed the analyses in H3K36R mutant larvae where the replacement of wt H3 by H3K36R is more complete and – for simplicity, but perhaps not wisely – had removed the data in H3K36R mutant embryos in the submission to eLife. We have now added the western blot and ChIP-seq analysis in H3K36R mutant embryos in Figure 5B, D and F.

Regarding the comment that the “The Western blot presented in Figure 4B does not strongly support the conclusion of a 3-4-fold global reduction of H3K36me2/me3 (the bands are quite “washed out” in lanes 5-8)”, we added Author response image 1 with overexposed uncropped images of this western. As illustrated in that figure, there indeed is a clear reduction of H3K36me2 and -me3 in extracts from H3K36A mutant embryos, the western against H2B of the same membrane shown in Author response image 1 serves as control. We estimate the reduction in H3K36me2 and -me3 levels in H3K36A mutant embryos (Figure 4B, lanes 5-8) compared to wild-type embryos (Figure 4B, lanes 1-4) to be about 3-4-fold.

Author response image 1

Unexpectedly, we found that in H3K36R mutant embryos the reduction of H3K36me3 is less severe and in the case of H3K36me2 we could not detect a clear reduction (Figure 5B, lanes 9-12). We used the H3K36R strain that we had generated in this study (4 copies of the 3xHisGUH3K36R miniarray) and the effects are therefore directly comparable to the effects observed in H3K36A mutant embryos that carried 4 copies of the 3xHisGUH3K36A miniarray. The less severe reduction in H3K36R mutant embryos compared to H3K36A mutant embryos is also paralleled in the genome-wide H3K36me2 profiling data in these embryos (Figure 5D, violin plots on the left). A simple technical reason such as cross-reactivity of the H3K36me2 and H3K36me3 antibodies specifically with H3K36R seems unlikely given the strong reduction of the H3K36me2 and H3K36me3 signals in extracts from H3K36R mutant larvae (shown in Figure 4A). We currently have no explanation why H3K36me2 and H3K36me3 levels appear to be only very slightly diminished in H3K36R mutant embryos and state this in the text. However, in the ChIP-seq analyses, we nevertheless observe that H3K27me3 levels at HOX are reduced in H3K36R mutant embryos (Figure 4F). Hence, at the HOX genes, H3K27me3 deposition appears to be similarly impaired in H3K36A and H3K36R mutant embryos.These results are described and discussed:

“In H3K36A mutants, H3K36me2 and H3K36me3 bulk levels were clearly reduced compared to wildtype (Figure 5B, compare lanes 5-8 with 1-4). In H3K36R mutants, H3K36me2 and H3K36me3 bulk levels unexpectedly appeared much less severely reduced (Figure 5B, compare lanes 9-12 with 1-4 and 5-8). As discussed above, the residual H3K36me2 and H3K36me3 signal in H3K36A and H3K36R mutant embryos might in part represent modified maternally-deposited canonical wild-type H3 and in part the modified H3.3 variants. However, the reason for the differential reduction of H3K36me2 and -me3 levels in H3K36A and H3K36R mutant embryos remains unclear. In both genotypes, H3K27me3 bulk levels appeared largely unchanged compared to wildtype (Figure 5B, compare lanes 5-8 and 9-12 with 1-4).”

“As expected from the Western blot analyses (Figure 5B), H3K36A or H3K36R mutant embryos showed no general reduction in their genome-wide H3K27me3 profiles (Figure 5D). However, in both mutants, H3K27me3 levels were about 1,5-fold reduced across the HOX gene loci (Figure 5D, F).”

10) How do the authors explain/speculate about the absent/modest effect on H3K27me3 in H3K36A mutant embryos despite the severity of the phenotype for these mutants? Here, again, a side-by-side comparison of H3K27me3 levels in H3K36R and H3K36A mutant embryos would be relevant. It would also be relevant to factor in knowledge about the number of cell divisions and maternal histone load in order to estimate if H3K36A is also impacting H3K27 methylation in vivo.

As discussed in our response to comments 8 and 9, for the western blot and ChIP-seq analyses in embryos, we used the H3K36A and H3K36R strains that we had generated in this study, i.e. Df HisC homozygotes rescued by 4 copies of the 3xHisGUH3K36A miniarray and by 4 copies of the 3xHisGUH3K36R miniarray, respectively. So the effects on H3K36me2 and -me3 are therefore directly comparable.

We discuss the difference in lethality in the different mutant strains in the main text:

“The H3K36R mutant animals from the strain constructed in this study (i.e. containing 4 copies of the 3xHisGUH3K36R miniarray) also completed embryogenesis, and their cuticle morphology was also indistinguishable from wildtype (Figure 4). However, 98% of individuals arrested development already at the end of embryogenesis and the 2% of mutant animals that hatched from the eggshell arrested development as first instar larvae (Figure 4). H3K36A mutants, containing 4 copies of a 3xHisGUH3K36A miniarray, also completed embryogenesis and the morphology of their embryonic cuticle also appeared indistinguishable from wildtype (Figure 4). 96% of these H3K36A mutant animals arrested development before hatching from the eggshell and the 4% that hatched died during the first larval instar (Figure 4). As discussed in the Materials and methods section, the difference in the lethality phase of the H3K36R and H3K36A mutants generated in this study compared to H3K36R mutants in the strain from Matera and colleagues is possibly linked to the histone rescue transgene system used.”

The Materials and methods section contains a more extensive discussion of this point:

“Comparison of the lethality of H3K36R and H3K36A mutants: The difference in the lethality phase of the H3K36R mutants generated in this study compared to H3K36R mutants in the strain from Matera and colleagues was unexpected because in both strains the Df(2L)HisC homozygotes carry 12 copies of the HisGUH3K36R cassette (i.e. four 3x HisGUH3K36R miniarrays in our strain and a single 12x HisGUH3K36R array in the strain from Matera and colleagues). A possible explanation for the poorer survival of H3K36R mutants in the strain generated here could be that histone transgene expression from the 3xHisGUH3K36R miniarrays is for some reason less effective that in the case of the 12x HisGUH3K36R array. We also note that a recent study reported that among Df(2L)HisC homozygotes that carry 20 HisGUH3K36A copies, about 50% of the mutant animals develop up to the pupal stages (Zhang et al., 2019). Zhang et al. have not analyzed their H3K36A mutants any further but it is possible that the higher copy number of the HisGUH3K36A cassette accounts for the better survival compared to the H3K36A strain generated in this study. Further studies will be needed to explore whether H3K36R and H3K36A mutants show comparable phenotypes in larvae.”

In the main text, we also expanded our explanation about the issue of perdurance of maternally-deposited wild-type histone H3 in H3K36R and H3K36A mutants. The relevant section now reads:

“For the interpretation of the following experiments, it is important to keep in mind that H3K36R and H3K36A zygotic mutant animals initially also contain a pool of maternally-deposited wild-type canonical H3 molecules that, together with H3K36R and H3K36A, become incorporated into chromatin during the pre-blastoderm cleavage cycles, up to and including the S-phase of cell cycle 14. It is only from the S-phase of cell cycle 15 onwards when only transgene-encoded histones then become incorporated into chromatin (Gunesdogan et al., 2010). Although the wild-type H3 molecules in chromatin become diluted during every cell cycle and are eventually fully replaced by mutant H3, they are probably still present in the chromatin of late-stage embryos. The effective replacement of persisting wild-type H3 molecules by mutant H3 greatly varies between embryonic tissues because of the different numbers of cell divisions that take place in the different tissues prior to the end of embryogenesis. For example, whereas epidermal cells undergo only two more divisions after S-phase 14, certain cells in the CNS undergo as many as 12 divisions (Bossing et al., 1996). In diploid tissues from H3K36R mutant larvae, replacement of wildtype H3 by H3K36R can be expected to be much more complete because of the extensive cell proliferation that occurs in these tissues that, after metamorphosis, will give rise to the structures of the adult body.”

11) The statement in the Figure 4E legend that "For each HOX gene there is a substantially larger proportion of cells in which the gene is decorated with H3K27me3 and repressed by Polycomb" is probably not correct: While the analyzed tissues are surely a mixed population with few cells showing expression of the tested HOX genes (Figure 5), it is unlikely that the larger proportion of genes show normal H3K27me3 – if this were the case, the ChIP data in Figure 4 would not show the large reduction in H3K27me3 at the tested loci. Thus, the data likely reflects that H3K27me3 is reduced in most of the cell types – yet the de-repression is only seen in a few cells – to show this, immunostaining for H3K27me3 should be included in Figure 5 (discussed further below).

We agree and modified the text to remove any allusion about the proportion of cells where HOX genes are inactive and active. The text now reads: “For every HOX gene, the analyzed tissues (CNS, thoracic imaginal discs and eye-antenna discs) represent a mixed population of cells with a fraction of cells in which the gene is inactive, decorated with H3K27me3 and repressed by PcG and fraction of cells in which the gene is transcriptionally active and carrying the H3K36me2 modification.”

We have added a new data figure (Figure 6—figure supplement 1) that documents that the reduction of H3K27me3 immunofluorescence signal in wing imaginal discs from wild-type and H3K36R mutant larvae. As expected, in H3K36R mutant larvae, the reduction of H3K27me3 is quite uniform across the different cells that form the wing disc. As control for the specificity of the H3K27me3 IF signal, we also show an image of a wing disc with clones of H3K27R mutant cells.

12) The authors state that "the most straightforward interpretation" of the observed misexpression of Polycomb target genes in the H3K36R larvae is that it results from reduced H3K27me3 at these loci. While this may well be the case in this study, loss of H3K27me3 can also be secondary to gene activation – a distinction that is overlooked in many studies. In this case, the question is whether loss of H3K27me3 is a direct effect of the H3K36 substitution or secondary to gene expression changes or altered/delayed development. In this context, the authors could draw more strongly on the previous observation (McKay et al., 2015; Meers et al., 2017) that the H3K36R mutant does not show massive deregulation of gene expression (as mentioned in the Discussion), and, most importantly, should emphasize their own observations that global H3K27me3 is reduced (Figure 4A) and H3K27me3 at Polycomb target genes is strongly reduced (Figure 4C) despite the low percentage of cells assayed showing misexpression of these particular genes (Figure 5B-D). Along with the in vitro data presented in Figure 2, this provides stronger evidence for a direct effect of an “intact” H3K36 in preserving normal PRC2 activity.

This is a good point and we modified the main text to discuss the result from the Meers et al., 2017 study: “these transcriptome analyses did not reveal any gross deregulation of HOX or PcG genes”. We also point out that HOX gene misexpression in H3K36R or H3K36A mutants is stochastic and not as widespread as in strong PcG mutants. Specifically, we explain that the stochastic misexpression of Ubx in individual cells in the wing blade primordium occurs in an “area of this disc where Ubx is most readily de-repressed if PcG function is perturbed (Beuchle et al., 2001)”. Similarly, we explain that “Antp is misexpressed in the antenna primordium, the region of the eye-anntenna imaginal discs where Antp is most susceptible to becoming misexpressed if PcG function is compromised.”

13) Along the same lines, it would be interesting to include immunostaining for H3K27me3 in Figures 5B-D to learn whether the loss of H3K27me3 is more prominent in the cells showing mis-expression and to substantiate the data from Figure 4 showing global reduction of H3K27me3.

As mentioned in our response to Comment 11, we have added a new data figure (Figure 6—figure supplement 1) that documents that the reduction of H3K27me3 immunofluorescence signal in wing imaginal discs from wild-type and H3K36R mutant larvae. As expected, in H3K36R mutant larvae, the reduction of H3K27me3 is quite uniform across the different cells that form the wing disc. As control for the specificity of the H3K27me3 IF signal, we also show an image of a wing disc with clones of H3K27R mutant cells.

We also explain in the text that the stochastic misexpression of Ubx in individual cells in the wing blade primordium occurs in an “area of this disc where Ubx is most readily de-repressed if PcG function is perturbed (Beuchle et al., 2001)”. Similarly, we explain that “Antp is misexpressed in the antenna primordium, the region of the eye-anntenna imaginal discs where Antp is most susceptible to becoming misexpressed if PcG function is compromised.”

In conclusion, the misexpression of HOX genes is not linked to a particularly severe reduction of H3K27me3 in specific sets of cells or parts of tissues but it simply occurs in stochastic manner and most readily occurs in those cells and tissues where HOX genes also first become de-repressed when Polycomb function is removed (i.e. as illustrated in Beuchle et al., 2001).

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

The authors' revisions have addressed the majority of the comments on the original manuscript, and the additional discussion has clarified their model for how H3K36 interactions with PRC2 may influence its activity. However, there is a remaining point that should be addressed:

The authors' assertion that Drosophila Pcl does not bind H3K36me2 or H3K36me3 is not entirely correct. Although the Tudor domain of Pcl does not recognize H3K36me2/3, Ballare et al. demonstrated that a Pcl Tudor-PHD1 construct does bind to H3K36me2 and H3K36me3 (Ballare et al., 2012). This finding is also explicitly stated in their article: "Nevertheless, using a Pcl Tudor-PHD1 construct, we were able to rescue the binding to H3K36me2 and H3K36me3 (Figure 2B)." Furthermore, Musselman et al., 2012, investigated the interactions between the human PHF1 Tudor domain and H3K36me3 but did not test the binding of the Drosophila Pcl Tudor-PHD1 domains to H3K36me2 or H3K36me3 (Musselman et al., 2012). In sum, the authors should amend their manuscript to acknowledge that the Drosophila Pcl Tudor-PHD1 domains have been shown to bind to H3K36me2 and H3K36me3.

We have edited the text to address the remaining point that the reviewers had raised on the revised version.

Specifically, we have clarified what is known about Polycomblike and H3K36me2/3 binding in Drosophila. The text now reads:

“In Polycomblike, the Drosophila ortholog of PHF1, a region comprising the tudor domain and the adjacent PHD finger has been reported to bind H3K36me3, H3K4me3, H3K9me3 and, more weakly, also H3K14me3 and H3K27me3 (Ballaré et al., 2012). We note, however, that the tudor domain of Polycomblike contains an incomplete aromatic cage and, on its own, is unable to bind methylated lysines (Friberg et al., 2010). Further analyses will be needed to assess whether and how interaction PHF1 or Polycomblike with H3K36me3 might change H3K27 methylation by PRC2 on more complex oligonucleosome substrates that contain both H3K36me3-modified and unmodified nucleosomes.”

https://doi.org/10.7554/eLife.61964.sa2

Article and author information

Author details

  1. Ksenia Finogenova

    Max Planck Institute of Biochemistry, Laboratory of Chromatin Biology, Martinsried, Germany
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7512-8637
  2. Jacques Bonnet

    Max Planck Institute of Biochemistry, Laboratory of Chromatin Biology, Martinsried, Germany
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  3. Simon Poepsel

    1. California Institute for Quantitative Biology (QB3), University of California, California Institute for Quantitative Biology (QB3), Molecular Biophysics and Integrative Bio-Imaging Division, Lawrence Berkeley National Laboratory, Berkeley, United States
    2. University of Cologne, Center for Molecular Medicine Cologne (CMMC), Faculty of Medicine and University Hospital Cologne, Cologne, Germany
    3. Cologne Excellence Cluster for Cellular Stress Responses in Ageing-Associated Diseases (CECAD), University of Cologne, Cologne, Germany
    Contribution
    Methodology
    Competing interests
    No competing interests declared
  4. Ingmar B Schäfer

    Max Planck Institute of Biochemistry, Department of Structural Cell Biology, Martinsried, Germany
    Contribution
    Validation, Methodology
    Competing interests
    No competing interests declared
  5. Katja Finkl

    Max Planck Institute of Biochemistry, Laboratory of Chromatin Biology, Martinsried, Germany
    Contribution
    Methodology
    Competing interests
    No competing interests declared
  6. Katharina Schmid

    Max Planck Institute of Biochemistry, Laboratory of Chromatin Biology, Martinsried, Germany
    Contribution
    Methodology
    Competing interests
    No competing interests declared
  7. Claudia Litz

    Max Planck Institute of Biochemistry, Laboratory of Chromatin Biology, Martinsried, Germany
    Contribution
    Methodology
    Competing interests
    No competing interests declared
  8. Mike Strauss

    Max Planck Institute of Biochemistry, cryoEM Facility, Martinsried, Germany
    Present address
    McGill University, Department of Anatomy and Cell Biology, Montreal, Canada
    Contribution
    Methodology
    Competing interests
    No competing interests declared
  9. Christian Benda

    Max Planck Institute of Biochemistry, Department of Structural Cell Biology, Martinsried, Germany
    Contribution
    Validation, Methodology
    Competing interests
    No competing interests declared
  10. Jürg Müller

    Max Planck Institute of Biochemistry, Laboratory of Chromatin Biology, Martinsried, Germany
    Contribution
    Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    For correspondence
    muellerj@biochem.mpg.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2391-4641

Funding

Deutsche Forschungsgemeinschaft (SFB1064)

  • Jürg Müller

Max-Planck-Gesellschaft

  • Jürg Müller

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

Acknowledgements

We thank Eva Nogales for generous advice, sharing of expertise and for hosting KF for grid preparation. We thank Tom Cech for stimulating discussions. We thank JR Prabu for excellent computing support, S Uebel, E.Weyher, R Kim and A Yeroslaviz of the MPIB core facilities and Martin Spitaler and Giovannie Cardone of the MPIB imaging facility for excellent technical support and S Schkoelziger and S Schmähling for help with some of the experiments. This work was supported by the Deutsche Forschungsgemeinschaft (SFB1064) and the MPG. ChIP-seq data have been deposited in GEO (accession number: GSE148254). The structural data are deposited in the EMDB (EMD-11910 and EMD-11912) and PDB (7AT8).

Senior Editor

  1. Cynthia Wolberger, Johns Hopkins University School of Medicine, United States

Reviewing Editor

  1. Jerry L Workman, Stowers Institute for Medical Research, United States

Reviewer

  1. Jerry L Workman, Stowers Institute for Medical Research, United States

Publication history

  1. Received: August 10, 2020
  2. Accepted: November 18, 2020
  3. Accepted Manuscript published: November 19, 2020 (version 1)
  4. Version of Record published: December 9, 2020 (version 2)
  5. Version of Record updated: December 11, 2020 (version 3)

Copyright

© 2020, Finogenova et al.

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

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