PRC1 Sustains the Memory of Neuronal Fate Independent of PRC2 Function

bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

Ayana Sawai1, Sarah Pfennig1, Milica Bulajić2, Alexander Miller1, Alireza Khodadadi- Jamayran3, Esteban O. Mazzoni2, Jeremy S. Dasen1*

1Neuroscience Institute, Department of Neuroscience and Physiology, NYU School of Medicine, New York, NY 10016, USA

2Department of Biology, New York University, New York, NY 10003, USA

3Applied Bioinformatics Laboratories, Office of Science and Research, NYU School of Medicine, New York, NY 10016, USA

*Corresponding author: [email protected]

1 Abstract 2 3 Polycomb repressive complexes (PRCs) 1 and 2 maintain stable cellular memories of 4 early fate decisions by establishing heritable patterns of gene repression. PRCs repress 5 transcription through histone modifications and chromatin compaction, but their roles in 6 neuronal subtype diversification are poorly defined. We unexpectedly found that PRC2 is 7 dispensable to preserve the morphogen-induced positional fates of spinal motor neurons (MNs), 8 while PRC1 is essential for the specification of segmentally-restricted subtypes. Mutation of the 9 core PRC1 component Ring1 in mice leads to increased chromatin accessibility and ectopic 10 expression of a broad variety of fates determinants, including Hox transcription factors, while 11 neuronal class-specific features are maintained. Loss of MN subtype identities in Ring1 mutants 12 is due to the suppression of Hox networks by derepressed caudal Hox genes. These results 13 indicate that PRC1 can function independently of de novo PRC2-dependent histone methylation 14 to maintain chromatin topology and transcriptional memory at the time of neuronal 15 differentiation. 16

17 Introduction 18 19 Accurate control of gene expression is essential for the specification and maintenance of 20 neural fates during development. Studies of cell type-restricted transcription factors have 21 illuminated the mechanisms by which spatial and temporal regulation of gene expression gives 22 rise to identifiable neuronal subtypes (Doe, 2017; Fishell and Kepecs, 2020; Hobert and Kratsios, 23 2019; Sagner and Briscoe, 2019; Venkatasubramanian and Mann, 2019). A parallel and critical 24 mechanism of gene regulation is through the post-translational modification of histones, which 25 enables and restricts transcription by modifying chromatin structure (Kishi and Gotoh, 2018; 26 Schuettengruber et al., 2017). The Polycomb group (PcG) is key family of histone-associated 27 proteins that play evolutionarily conserved roles in restricting gene expression during 28 development (Blackledge et al., 2015; Gentile and Kmita, 2020; Simon and Kingston, 2009; 29 Soshnikova and Duboule, 2009). In embryonic stem cells, cell fate determinants are repressed 30 through PRC activities, and PRC-associated histone marks are subsequently removed from loci 31 as cells differentiate (Boyer et al., 2006; Farcas et al., 2012; Tavares et al., 2012). PRC 32 repression is maintained through cell division and after differentiation, and is thought to 33 contribute to stable cellular memories of early patterning events (Ciabrelli et al., 2017; Coleman 34 and Struhl, 2017). In vertebrates, much of our knowledge of how PRCs regulate gene expression 35 has emerged from biochemical studies of PcG proteins or from the activity of these factors in 36 proliferating cells. Despite an in depth understanding of the mechanisms of PRC action, how 37 PcG proteins interact with gene regulatory networks in the CNS remains poorly understood. 38 39 The specification of neuronal fates in the vertebrate spinal cord provides a tractable 40 system to elucidate the function of PcG proteins, as the pathways that determine identities are 41 well-defined, and the molecular signatures of many subtypes are known (Butler and Bronner, 42 2015; Sagner and Briscoe, 2019). One neuronal class where fate specification has been closely 43 examined is the spinal MN. A core set of transcription factors, including Hb9, Isl1/2, and Lhx3, 44 determines class-specific features of MNs, including neurotransmitter identity (Shirasaki and 45 Pfaff, 2002). The subsequent diversification of MNs into hundreds of muscle-specific subtypes is 46 achieved through a conserved network of Hox transcription factors differentially expressed along 47 the rostrocaudal axis (Philippidou and Dasen, 2013). During neural tube patterning, opposing 48 gradients of retinoic acid (RA) and fibroblast growth factors (FGFs) provide spinal progenitors

49 with a positional identity (Bel-Vialar et al., 2002; Dasen et al., 2003; Liu et al., 2001). These 50 morphogens act, in part, by temporally and spatially depleting Polycomb-associated histone 51 marks from Hox clusters (Mazzoni et al., 2013; Narendra et al., 2015). As progenitors exit the 52 cell cycle, MNs continue to express Hox genes, where they regulate repertoires of subtype- 53 specific genes (Catela et al., 2016; Dasen et al., 2005; Mendelsohn et al., 2017). Although the 54 role of Hox proteins in the CNS are well-characterized (Parker and Krumlauf, 2020), and are 55 known targets of PRC activities (Gentile and Kmita, 2020), the specific contributions of PRC1 56 and PRC2 to CNS maturation remain unclear, as few studies have directly compared their 57 functions during embryonic development. 58 59 Polycomb repression is initiated by PRC2, which methylates histone H3 at lysine-27, 60 permitting recruitment of PRC1 through subunits that recognize this mark, leading to chromatin 61 compaction at genes targeted for repression (Margueron and Reinberg, 2011; Schuettengruber et 62 al., 2017). PRC1 and PRC2 can also exist in a variety of configurations, which may contribute to 63 neuronal subtype-specific activities. The core subunit of PRC1, Ring1, binds to one of six 64 Polycomb group Ring finger (Pcgf) proteins (Gao et al., 2012). PRC1 containing Pcgf4 interacts 65 with Cbx proteins (canonical PRC1) which recognize H3K27me3 (Bernstein et al., 2006; Morey 66 et al., 2012). Variant forms of PRC1 containing Rybp can inhibit incorporation of Cbx proteins 67 into PRC1, and bind target loci independent of H3K27me3 (Tavares et al., 2012; Wang et al., 68 2010). We previously found that a PRC1 component, Pcgf4, is required to establish rostral 69 boundaries of Hox expression in differentiating MNs (Golden and Dasen, 2012). By contrast, 70 deletion of a core PRC2 component, Ezh2, from MN progenitors had no apparent impact on fate 71 specification, possibly due to compensation by Ezh1 (Shen et al., 2008), or through use of variant 72 PRC1 containing Rybp. These observations raise the question of what are the specific roles of 73 canonical PRC1, variant PRC1, and PRC2 during MN subtype diversification. 74 75 To determine the function of PRCs during neural differentiation, we removed core 76 components of these complexes from MN progenitors. We found that depletion of Ring1 77 proteins, essential constituents of all PRC1 isoforms, causes pronounced changes in transcription 78 factor expression and a loss of Hox-dependent MN subtypes. Surprisingly, neither PRC2 nor 79 variant PRC1 activities are required for rostrocaudal patterning at the time of MN differentiation. 80 Deletion of Ring1 leads to increased chromatin accessibility and derepression of a broad variety

81 of cell fate determinants, while class-specific features of MNs are preserved. The derepression of 82 caudal Hox genes in Ring1 mutants leads to the suppression of MN subtype diversification 83 programs. These findings indicate that PRC1 can preserve the memory of early patterning events 84 in the CNS. 85 86 Results 87 88 PRC1 is Essential for Rostrocaudal Patterning During Neuronal Differentiation 89 To determine the relative contributions of PRC1 and PRC2 to neuronal specification, we 90 analyzed mice in which core subunit-encoding genes were selectively removed from MN 91 progenitors. Ezh genes encode the methyltransferase activity of PRC2, while Eed is required to 92 enhance this function. We first generated mice in which both Ezh genes are conditionally 93 deleted, by breeding Ezh1 and Ezh2 floxed lines to Olig2::Cre mice, which targets Cre to MN 94 progenitors (EzhMNΔ mice) (Hidalgo et al., 2012; Su et al., 2003). We confirmed MN-restricted 95 loss of PRC2 activity by examining the pattern of H3K27me3 at E11.5, which was selectively 96 depleted from progenitors and post-mitotic MNs (Figure 1A,B). Expression of Hb9 and Vacht, 97 two general markers of MN identity, were grossly unchanged in EzhMNΔ mice. (Figure 1C,D,G). 98 We next analyzed expression of Hox proteins in EzhMNΔ mice and found that the MN columnar 99 subtype determinants Hoxc6, Hoxc9, and Hoxc10, were all expressed in their normal domains 100 (Figure 1E,F, Supplementary Figure 1A-C). Similarly, we observed no changes in Hb9, Hoxc6, 101 Hoxc9, and Hoxc10 expression in MNs after removal of the PRC2 component Eed, using 102 Sox1::Cre (EedNEΔ mice), which targets Cre to neuroectoderm (Supplementary Figure 1D-H) 103 (Takashima et al., 2007; Yaghmaeian Salmani et al., 2018). Depletion of PRC2 activity from 104 neural progenitors therefore does not affect MN class specification or rostrocaudal patterning. 105 106 We next investigated the function of PRC1, which is thought to repress gene expression 107 in a PRC2-dependent manner. We generated mice in which Ring1B is conditionally deleted from 108 MN progenitors using Olig2::Cre in a global Ring1A mutant background (Ring1MNΔ mice) (Cales 109 et al., 2008; del Mar Lorente et al., 2000). In Ring1MNΔ mice, Ring1B protein was selectively 110 removed from progenitors and post-mitotic MNs (Figure 1H,I). The number of Hb9+ MNs and 111 pattern of Vacht expression were similar to controls in Ring1MNΔ mice (Figure 1J,K,N). By 112 contrast, Hox proteins normally expressed by forelimb-innervating brachial MNs were not

113 detected, while thoracic and lumbar Hox proteins were ectopically expressed in more rostral 114 MNs (Figure 1L,M,O-T, Supplementary Figure 1I,J,K). Brachial-level Hox proteins (Hoxc4, 115 Hoxa5, Hoxc6 and Hoxc8), were selectively depleted from MNs, while thoracic and lumbar Hox 116 determinants, (Hoxc9, Hoxc10, and Hoxd10) were derepressed in brachial segments (Figure 1Q- 117 T). At thoracic levels Hoxc9 expression was attenuated and Hoxd10 was ectopically expressed 118 (Supplementary Figure 1J). These results show that loss of Ring1 leads to a marked rostral shift 119 in Hox pattern, without affecting pan-MN molecular features (Figure 1O). 120 121 Canonical PRC1 Regulates Hox Expression in MNs 122 Ring1 can interact with multiple Pcgf proteins, raising the question of which PRC1 123 configuration contributes to MN patterning. PRC1 containing Pcgf4 is required to establish Hox 124 boundaries in MNs (Golden and Dasen, 2012), but can exist in two alternative configurations, 125 depending on mutually exclusive incorporation of Cbx (canonical PRC1) or Rybp (variant 126 PRC1) (Gao et al., 2012; Tavares et al., 2012). We examined the function of PRC1 isoforms first 127 by manipulating Rybp and Cbx expression in MNs. We hypothesized that if canonical PRC1 128 regulates Hox expression, then overexpression of Rybp would inhibit binding of Cbx to Ring1, 129 leading to MN phenotypes similar to Ring1 mutants (Figure 1U). We used chick in ovo neural 130 tube electroporation to express Rybp in postmitotic MNs using the Hb9 promoter (Hb9::mRybp). 131 Expression of mRybp under Hb9 led to ectopic Hoxc9 and Hoxc10 expression at brachial levels 132 (Figure 1V). If Rybp acts by displacing Cbx, then elevating Cbx levels should restore normal 133 Hox expression. To test this, we co-electroporated Hb9::mRybp and Hb9::mCbx2 at equivalent 134 plasmid concentration, and no longer observed ectopic Hoxc9 in brachial segments (Figure 1X). 135 136 These observations are consistent with canonical PRC1 containing Pcgf4, Cbx, and 137 Ring1 restricting Hox expression in MNs. Rybp is expressed by MNs at the time of their 138 differentiation (Supplementary Figure 1L), raising the possibility that Rybp (or its paralog Yaf2) 139 also plays regulatory roles during development. We therefore selectively deleted Rybp from MNs 140 using Olig2::Cre mice in a Yaf2-/- background (Rybp/Yaf2MNΔ mice). Combined deletion of Rybp 141 and Yaf2 did not affect MN generation, Hoxc6, Hoxc9, or Hoxc10 expression (Supplementary 142 Figure 1L). These findings indicate that canonical PRC1 maintains appropriate Hox expression, 143 while variant PRC1 and PRC2 do not contribute to rostrocaudal patterning at the time of MN 144 differentiation.

145 146 PRC1 is Required for MN Subtype Diversification 147 Hox transcription factors play central roles in establishing neuronal subtype identities 148 through regulating expression of subtype-specific genes. To determine the consequences of 149 altered Hox expression in Ring1MNΔ mice, we analyzed the molecular profiles and peripheral 150 innervation pattern of MNs. A key Hox target in MNs is the transcription factor Foxp1, which is 151 essential for the differentiation of limb-innervating lateral motor column (LMC) and thoracic 152 preganglionic column (PGC) neurons (Dasen et al., 2008; Rousso et al., 2008). In Ring1MNΔ 153 mice, Foxp1 expression is lost from brachial and thoracic MNs, and markedly reduced in lumbar 154 MNs (Figure 2A,B,G, Supplementary Figure 2C). By contrast, in EzhMNΔ, EedNEΔ and 155 Rybp/Yaf2MNΔ mice, Foxp1 expression was maintained, consistent with the preservation of 156 normal Hox profiles in these mutants (Supplementary Figure 1L, 2A,B). In Ring1MNΔ mice, 157 expression of the LMC marker Raldh2 was lost at brachial levels, and the thoracic PGC marker 158 nNos was not detected (Figure 2C, D). Moreover, expression of determinants of respiratory 159 phrenic MNs (Scip+ Isl1/2+), was markedly depleted, and likely contributes to the perinatal 160 lethality of Ring1MNΔ mice (Supplementary Figure 2D). These results indicate that Ring1 deletion 161 leads to a loss of genes acting downstream of Hox function in MNs. 162 163 We next assessed the impact of Ring1 deletion on two MN subtypes that are specified 164 independent of Hox function, the hypaxial and median motor columns (HMC and MMC). In 165 Ring1MNΔ mice, MMC neurons (Lhx3+ Hb9+) were generated at normal numbers in thoracic 166 segments, while the number of HMC neurons (Isl1/2+ Hb9+ Lhx3-) was increased in thoracic and 167 brachial segments (Figure 2E,F,H, Supplementary Figure 2E,F). The increase in HMC neurons 168 in Ring1 mutants is likely due to a reversion of presumptive Hox-dependent subtypes to an HMC 169 fate, similar to mice in which Hox function is disrupted (Dasen et al., 2008; Hanley et al., 2016). 170 Mutation in Ring1 therefore depletes Hox-dependent subtypes, with the remaining MNs having a 171 more ancestral axial identity (Figure 2P). 172 173 As we observed a dramatic loss of segmentally-restricted MN subtypes in Ring1MNΔ mice, 174 we next assessed the impact on peripheral innervation pattern. To trace motor axon projections, 175 we crossed Ring1MNΔ mice to an Hb9::GFP reporter, in which all MN axons are labelled with 176 GFP. In control mice there are nine primary trajectories of forelimb-innervating motor axon at

177 E12.5 (Figure 2I,J) (Catela et al., 2016). In Ring1MNΔ mice, only three nerve branches, radial, 178 median, and ulnar were visible but appeared prematurely truncated and unbranched (Figure 2K). 179 In the trunk, innervation of sympathetic chain ganglia was lost, consistent with a loss of PGC 180 fates (Figure 2L,M). Projections to dorsal and ventral axial muscles by MMC and HMC subtypes 181 were maintained in Ring1MNΔ mice, although axons were shorter than in controls (Figure 2N, O). 182 Loss of Ring1 therefore causes severe defects in innervation pattern, while the trajectories of 183 axial MNs are relatively spared. 184 185 Loss of Ring1 Causes Broad Derepression of Developmental Fate Determinants 186 Polycomb proteins regulate diverse aspects of differentiation by restricting gene 187 expression during development. To investigate changes in gene expression after deletion of 188 Ring1 genes in an unbiased manner, we performed RNAseq on MNs isolated from Ring1MNΔ 189 mice. We purified MNs from control and Ring1MNΔ;Hb9::GFP embryos at E12.5 by flow 190 cytometry (n=4 Ring1 mutants, and n=4 Cre- controls), and performed RNAseq. Because Ring1 191 mutants display segment-specific phenotypes, we collected MNs from brachial, thoracic, and 192 lumbar levels and profiled each population independently. 193 194 We identified a total of 1001 upregulated and 641 downregulated genes in Ring1MNΔ mice

195 (log2-FC>2, FDR<0.1), with 391 genes upregulated in all three segmental regions, compared to 196 46 genes that were commonly downregulated (Figure 3A-C, Supplementary Figure 3A,C, Table 197 1). Thus, many upregulated genes are shared between each segment, while downregulated genes

198 tended to be segment-specific. Strikingly, 76% of the top 100 derepressed genes (by log2-FC) at 199 brachial levels encode transcription factors, and include genes normally involved in the 200 specification of spinal interneurons (e.g. Gata3, Shox2, Lhx2), brain regions (FoxG1, Six6, 201 Lhx8), and non-neuronal lineages (Figure 3B). 202 203 Although a variety of cell fate determinants were derepressed in Ring1MNΔ mice, our 204 RNAseq analyses provide further evidence that core molecular determinants of MN class identity 205 are preserved in Ring1 mutants. We found no significant changes in transcription factors (Isl1, 206 Isl2, Hb9, Lhx3, Lhx4), guidance molecules (Slit2, Robo1, Robo2, DCC) and neurotransmitter 207 genes (Chat, Acly) associated with pan-MN features (Figure 3D). There was a modest decrease 208 in expression Slc18a3 (Vacht) (Figure 3D). Expression of genes that mark other excitatory

209 (Slc17a7, Slc17a6) or inhibitory (Slc32a1, Gad2, Gad1,) neuronal classes were not markedly 210 changed in Ring1 mutants (Supplementary Figure 3B). By contrast, expression of genes 211 associated with MN subtype identities was decreased in Ring1MNΔ mice, including genes that 212 mark limb-innervating (e.g Foxp1, Etv4, Raldh2, Runx1), thoracic-specific (Nos1, Cyp26b), and 213 respiratory (Alcam, Ptn) subtypes (Figure 3E). 214 215 To validate these changes in gene expression, we performed mRNA in situ hybridization 216 on a subset of upregulated or downregulated genes. We found that Gata3, Lhx8, and Lhx2 were 217 markedly upregulated in MNs of Ring1MNΔ mice (Figure 4A-F), while other genes (Foxg1, Six6, 218 Pitx2, Shox2, Nkx2.1) showed less prominent, but detectable, ectopic expression (Supplementary 219 Figure 4A-J). We also examined expression of previously uncharacterized genes that were 220 downregulated in all three segmental regions. We found Rgs4, Uts2b, Vat1, and Gabra2 were 221 expressed by MNs of control embryos, and were downregulated in Ring1 mutants (Figure 4G-L, 222 Supplementary Figure 4K,L). These results indicate that despite the derepression of multiple fate 223 determinants in Ring1MNΔ mice, core features of MN identity are preserved, but that subtype 224 diversification programs are selectively disrupted. 225 226 Selective Derepression of Caudal Hox genes in Ring1MNΔ mice 227 As our preliminary analyses revealed altered expression in a subset of Hox genes in 228 Ring1MNΔ mice, we further evaluated Hox genes in our RNAseq dataset. Consistent with the 229 analysis of Hox protein expression, rostral Hox genes were reduced in brachial segments of 230 Ring1MNΔ mice, while Hox10-Hox13 genes were derepressed (Figure 5A, Supplementary Figure 231 5A-C). We observed derepression of caudal Hox genes from each of the four vertebrate Hox 232 clusters, including genes not normally detectable in the ventral spinal cord (e.g HoxB genes) 233 (Dasen et al., 2005) (Figure 5A, Supplementary Figure 5A-C). The extent of caudal Hox 234 derepression correlated with the relative position of genes within a cluster, with Hox13 genes 235 displaying the most pronounced derepression in Ring1MNΔ mice (by FC), relative to other caudal 236 Hox genes (Figure 5B, Supplementary Figure 3C). The marked derepression of Hox13 genes was 237 observed in each of the three segmental levels we analyzed, while Hox10 genes were selectively 238 derepressed in brachial and thoracic segments (Figure 5A, Supplementary Figure 5A-C). 239

240 To further validate these findings, we analyzed Hox13 expression by in situ 241 hybridization. In controls Hoxc13 is restricted to sacral segments, while Hoxb13 is not detected 242 in MNs (Figure 5C). In Ring1MNΔ mice, both Hoxb13 and Hoxc13, were de-repressed in MNs 243 throughout the rostrocaudal axis (Figure 5C). By contrast, in situ hybridization of Hoxc6 and 244 Hoxc9 expression revealed reduced expression at brachial and thoracic levels, respectively 245 (Supplementary Figure 5C). Thus, ectopic expression of Hox13 genes in Ring1 mutants is 246 associated with a loss of rostral Hox gene expression. 247 248 Ring1 is Essential to Maintain MN Chromatin Topology 249 Our findings indicate that in the absence of Ring1 genes, a broad variety of cell fate 250 determinants are ectopically expressed in MNs, while only a subset of caudal Hox genes are 251 derepressed. As PRC1 restricts gene expression through chromatin compaction, we investigated 252 whether removal of Ring1 leads to changes in DNA accessibility at derepressed loci. We used 253 Assay for Transposase-Accessible Chromatin with high-throughput sequencing (ATACseq) to 254 identify genomic regions which have gained or lost accessibility in Ring1MNΔ mice. We purified 255 Hb9::GFP+ MNs at E12.5 from control and Ring1MNΔ embryos at brachial, thoracic, and lumbar 256 levels, and performed ATACseq. In control samples, we observed a progressive opening of 257 caudal Hox genes in more rostral segments. For example, the accessibility of Hox9 genes 258 increases from brachial to thoracic segments, while Hox10 and Hox11 genes are more accessible 259 in lumbar segments (Supplementary Figure 6A, B). 260 261 To determine which genomic regions gained accessibility in Ring1 mutants, we compared 262 ATACseq profiles between MNs of control and Ring1MNΔ mice. We identified a total of 2,305 263 loci that gained accessibility in Ring1MNΔ embryos, 324 (14%) of which were common to all 264 three segmental levels (Figure 6A, Table 2). Common genes included caudal Hox genes 265 (Hoxa13, Hoxc13) and other transcription factors (e.g. Foxg1, Lhx2, Pitx2) that were derepressed 266 in our RNAseq analyses (Figure 6C). We also identified 1,264 loci that lost accessibility in 267 Ring1MNΔ mice, 39 (3%) of which were common to all three segments (Figure 6A). Thus, similar 268 to our RNAseq results, loss of Ring1 leads to increased accessibility in many genes that are 269 shared among each segment, while genes that lose accessibility tend to be segment-specific. 270

271 To assess which genes may be directly repressed by PRC1, we examined the overlap 272 between transcripts that were upregulated and loci that gained accessibility in Ring1MNΔ mice. 273 We found that 18% (148/813) of genes that were ectopically expressed at brachial segments also 274 gained accessibility in Ring1MNΔ mice (Figure 6B). We found 19 genes, including Hoxa13 and 275 Hoxc13, were derepressed and gained accessibly in all three segments (Supplementary Figure 276 6C). Since lumbar segments still retain features of Hox-dependent subtypes, we also compared 277 the overlap between genes that were upregulated and gained accessibility in brachial and thoracic 278 MNs. We identified 73 genes, 56 (77%) of which encode transcription factors, including each of 279 the caudal Hox genes we found by RNAseq (Figure 6B, Supplementary Figure 6C). The gain of 280 accessibility at transcription factor-encoding genes was prominent near transcription start sites 281 (Figure 6C), and regions that gained accessibility in Ring1MNΔ neurons correspond to regions 282 shown to be bound by Ring1B in ES cells (Supplementary Figure 6D) (Bonev et al., 2017). 283 284 In agreement with our RNAseq analysis, Hox13 paralogs showed pronounced increases 285 in accessibility, with each of the four Hox13 genes gaining accessibility in each of the three 286 segmental levels (Figure 6D,E). By contrast, rostral Hox4-Hox8 genes, which are 287 transcriptionally downregulated in Ring1 mutants, were not among that targets that lost 288 accessibility (Figure 6D). These results suggest that reduced expression of rostral Hox4-Hox9 289 genes in Ring1 mutants is not due to a loss in chromatin accessibility. 290 291 Hox13 Paralogs Represses Rostral Hox genes by Engaging Accessible Chromatin Domains 292 As PRC1-mediated chromatin compaction is essential for Hox repression, it is surprising 293 that Hox4-Hox9 genes are diminished in Ring1 mutants, without a noticeable reduction in 294 chromatin accessibility at these loci (Supplementary Figure 6D). Because cross-repressive 295 interactions between Hox genes themselves are an important regulatory mechanism determining 296 Hox boundaries (Philippidou and Dasen, 2013), it is possible that ectopically expressed Hox13 297 genes directly repress multiple Hox genes in Ring1MNΔ mice. To test this, we first analyzed MNs 298 derived from ES cells (ESC-MNs) in which Hoxc13 expression can be induced upon 299 doxycycline (Dox) treatment (Bulajic et al., 2020). ESC-MNs differentiated via RA and Sonic 300 Hedgehog agonist are characterized by expression of Hox4-Hox5 paralogs. RNAseq analyses 301 revealed that Hoxa4, Hoxa5, Hoxc4, and Hoxc5 expression were markedly reduced after Dox- 302 induced Hoxc13 expression in MN progenitors, compared to non-induced ESC-MNs (Figure 7A,

303 Supplementary Figure 7A-C). This repressive effect also appears to be direct, as ChIPseq 304 analysis indicates that Hoxc13 can bind at multiple Hox genes (Figure 7A). 305 306 Hox13 proteins can target inaccessible chromatin (Bulajic et al., 2020; Desanlis et al., 307 2020), while repression by the Drosophila Hox protein Ubx is associated with chromatin 308 compaction (Loker et al., 2021). To examine the possible effects of Hox13 proteins on local 309 chromatin structure, we compared accessibility at Hox loci in both control and Ring1 mutants, 310 relative to the location of Hoxc13 binding sites. At Hox4-Hox5 genes, Hoxc13 sites correspond 311 to regions that are accessible in both control and Ring1 mutant MNs (Figure 7A, Supplementary 312 Figure 7A-C). Hox13 proteins therefore appear to repress rostral Hox genes, in part, through 313 binding at pre-existing accessible regions. 314 315 To test whether Hox13 genes can repress multiple Hox paralogs in vivo, we used chick 316 neural tube electroporation to express Hox13 genes within brachial, thoracic, and lumbar 317 segments, and assessed Hox4-Hox10 gene expression. We electroporated pCAGGs::mHoxa13- 318 ires-nucGFP or pCAGGs::mHoxb13-ires-nucGFP and found that both cell-autonomously 319 repressed expression of brachial, thoracic, and lumbar Hox proteins (Hoxc4, Hoxa5, Hoxc6, 320 Hoxc8, Hoxc9, and Hoxc10) (Figure 7B-D). By contrast, misexpression of Hox13 genes did not 321 affect expression of the general MN markers Isl1/2 and Hb9 (Figure 7B-D). These results 322 indicate that Hox13 proteins can directly repress multiple Hox genes, and likely contribute to the 323 MN fate specification defects of Ring1MNΔ mice. 324 325 Discussion 326 327 The Polycomb group encompass a large and diverse family of proteins essential for 328 maintaining epigenetic memory of early patterning events. Classically, PcG-mediated repression 329 is thought to depend on recruitment of PRC1 through recognition of histone methylation marks 330 deposited by PRC2 activity. Although alternative, H3K27me3-independent, mechanisms of 331 PRC1 repression have been described (Gao et al., 2012; Tavares et al., 2012), the relative 332 contribution of PRC1 and PRC2 to neuronal fate specification have not been resolved. In this 333 study, we found that genetic depletion of PRC2 components has no discernable impact on neural 334 class or subtype differentiation programs at the time of MN differentiation. By contrast, a core

335 PRC1 subunit, Ring1, is required to restrict transcription factor expression in the CNS, maintain 336 neuronal subtype-specific chromatin topology, and determine rostrocaudal boundaries of Hox 337 expression. Our findings indicate PRC1 plays an key role in regulating expression of cell fate 338 determinants during MN differentiation, safeguarding neurons from acquiring inappropriate gene 339 regulatory programs. 340 341 PRC Functions in Neural Development 342 Polycomb proteins function in diverse aspects of CNS maturation, including the temporal 343 transition from neurogenesis to gliogenesis, maintenance of adult neural stem cell fates, and 344 restriction of gene expression in mature neurons (Di Meglio et al., 2013; Fasano et al., 2009; 345 Hirabayashi et al., 2009; Molofsky et al., 2003; von Schimmelmann et al., 2016). The role of 346 PRCs in neural subtype diversification has been challenging to study, in part, due the complexity 347 and dynamic temporal regulation of its constituents. By selectively removing core PRC1 and 348 PRC2 subunits from neural progenitors, we determined the relative contributions of these 349 chromatin-associated complexes to neuronal diversification. We found that PRC1 represses a 350 broad variety of cell fate determinants, with the majority of highly derepressed genes in Ring1 351 mutants encoding transcription factors. Ectopic expression of Ring1-regulated targets is 352 associated with a disruption in chromatin topology, resulting in a gain in accessibility of 353 derepressed genes. 354 355 Despite the derepression of multiple fate determinants in the absence of PRC1 function, 356 expression of transcription factors that define MN class identity (e.g. Hb9, Isl1/2, and Lhx3), are 357 largely unchanged, and MNs do not acquire characteristics of other neuronal classes. This 358 observation suggests that once a class identity has been established, ectopic expression of fate 359 determinants is insufficient to respecify basic neuronal features, such as neurotransmitter 360 identity. In C. elegans the ability of transcription factors to reprogram neuronal class identity 361 requires removal of histone deacetylase proteins (Tursun et al., 2011), suggesting that additional 362 regulatory constraints, such as histone acetylation and/or DNA-methylation, restricts the ability 363 of ectopic transcription factors to interfere with class-specific programs in Ring1 mutants. 364 365 While basic elements of MN fate are maintained in the absence of PRC1, their 366 differentiation into molecularly distinct subtypes is severely compromised. The loss of subtype-

367 specific programs in brachial and thoracic segments can be attributed to the attenuation of Hox4- 368 Hox9 paralog expression, as mice lacking these Hox genes show a similar loss in MN subtype 369 features (Jung et al., 2010; Jung et al., 2014; Philippidou et al., 2012). Selective loss of Hox 370 function in MNs also appears to account for the observation that the majority of downregulated 371 genes in Ring1 mutants are segment-specific. 372 373 PRC-Dependent and Independent Mechanisms of Rostrocaudal Patterning in the CNS 374 We found that removal of PRC1 leads to the derepression and increased accessibility of 375 caudal Hox genes. Loss of PRC1 does not result in a sustained derepression of all Hox genes, as 376 Hox genes normally expressed by brachial- and thoracic-level MNs are attenuated in the absence 377 of Ring1. These findings are reminiscent of the function of PRCs in Drosophila, where loss of 378 PRC function leads to ectopic expression of the caudal Hox gene Abd-b and repression of Ubx 379 expression (Struhl and Akam, 1985; Struhl and White, 1985), likely reflecting cross-repressive 380 interactions between caudal Hox proteins and rostral Hox genes. Notably, reduced rostral Hox 381 gene expression in Ring1 mutants is not a reflection of reduced chromatin accessibility at these 382 targets, consistent with a model in which this form of repression is not associated with global 383 changes in chromatin structure. 384 385 In Ring1 mutants, Hox13 genes are derepressed in MNs, and appear to contribute to the 386 loss of MN subtypes. Derepression of Hox13 genes alone does not appear to account for all of 387 the specification defects in Ring1 mutants, as Hox10 genes were expressed normally in lumbar 388 segments. Hoxc9 can also repress Hox4-Hox8 genes in MNs through direct binding (Jung et al., 389 2010), and likely contributes to the Ring1 mutant phenotype. Derepression of multiple caudal 390 Hox paralogs therefore appears to have a cumulative effect on repressing brachial and thoracic 391 Hox genes. In conjunction with studies in Drosophila, these finding indicate a deeply-conserved 392 mechanism of rostrocaudal patterning, in which PRCs establish and maintain rostral Hox 393 boundaries, while Hox cross-repressive interactions define caudal boundaries. 394 395 PRC2-Independent Functions of PRC1 in Maintaining Neuronal Subtype Fate 396 Studies of neuronal and non-neuronal development provide compelling evidence that 397 PRC2 activity is required to restrict gene expression in early development (Ezhkova et al., 2009; 398 Gentile et al., 2019; Snitow et al., 2015; Yaghmaeian Salmani et al., 2018). An unexpected

399 finding from our studies is that MN subtype differentiation and Hox boundaries are maintained 400 after removal of PRC2 activity. Mutation of Ezh genes or Eed depletes H3K27me3 from 401 progenitors and postmitotic neurons, without appreciably affecting MN generation, Hox 402 expression, or downstream Hox effectors. These observations indicate that neural progenitors 403 can acquire independence from continuous PRC2 activity to restrict gene expression, whereas 404 there is an absolute requirement for PRC1 function to maintain appropriate transcription factor 405 expression and chromatin organization. 406 407 How does PRC1 restrict expression of fate determinants in the absence of PRC2 activity? 408 One possibility is that residual H3K27me3, below the threshold of histological detection, is 409 transmitted through cell division and is sufficient to recruit canonical PRC1 and maintain target 410 gene repression. As cells divide, newly synthesized histones are presumably devoid of 411 H3K27me3 in PRC2 mutants, leading to replication-coupled dilution of H3K27me3. During 412 neural differentiation, the rate of cell division decreases (Kicheva et al., 2014; Wilcock et al., 413 2007), potentially limiting H3K27me3 reduction, and enabling PRC1 to bind at target loci, even 414 in the absence of de novo H3K27 methylation. Although we observed no MN phenotypes in Ezh 415 or Eed mutants, later-born oligodendrocytes that derive from the same precursors as MNs have 416 been shown to depend on PRC2 function for their differentiation (Wang et al., 2020). This more 417 pronounced effect on glial development likely reflects further dilution of H3K27me3 through 418 cell division in PRC2 mutants. 419 420 Stabilization of PRC1 at repressed loci could also maintain target repression 421 independently of de novo H3K27 methylation. Repression by canonical PRC1 has been shown to 422 depend on the formation of phase-separated subnuclear structures (Polycomb bodies) assembled 423 through polymerization of Polyhomeotic-like (Phc) and/or Cbx proteins (Isono et al., 2013; Plys 424 et al., 2019; Tsuboi et al., 2018). Mutation of Phc2 in mice leads to ectopic expression of Hox 425 genes, with Hox13 genes among the most robustly derepressed targets (Isono et al., 2013). One 426 possibility is that PcG-mediated repression may not depend on anchoring of PRC1 through 427 H3K27me3, but is maintained through Phc-mediated chromatin compaction at specific loci. 428 429 We suggest that in the early phases of development PRC2 activity defines the sites of 430 maintained H3K27 tri-methylation, initiates PRC1 recruitment, and restricts transcription factor

431 expression in MNs. These activities are likely deployed prior to neural progenitor specification, 432 as recent studies have demonstrated that the establishment of rostrocaudal positional identities 433 can be specified before neural induction (Metzis et al., 2018). Patterning morphogens, such as 434 RA and FGF, may initially act on stem cells to establish the pattern of H3K27me3 at Hox loci 435 prior to neurogenesis. This early rostrocaudal patterning step may ultimately serve to coordinate 436 Hox expression in the neural tube and surrounding mesodermally-derived tissues. The 437 subsequent switch to reliance on PRC1 function could reflect a general mechanism of gene 438 regulation in neuronal subtypes that become terminally differentiated during development. 439

440

441 Materials and Methods 442 443 Mouse Genetics 444 Ezh1 flox (Hidalgo et al., 2012), Ezh2 flox (Su et al., 2003), Olig2::Cre (Dessaud et al., 2007), 445 Hb9::GFP (Arber et al., 1999) mice have been previously described. Ring1A-/-; Ring1B flox 446 (Cales et al., 2008; del Mar Lorente et al., 2000) and Rybp flox; Yaf2-/- (Hisada et al., 2012; Rose 447 et al., 2016) mouse lines were generated by microinjection of the mouse ESCs expressing these 448 constructs into the blastocyst followed by implantation of these eggs into pseudopregnant foster 449 female mice. Generation of EzhMNΔ mice was performed by crossing Ezh1 flox, Ezh2 flox and 450 Olig2::Cre. EzhMNΔ mice are viable at birth but do not survive beyond P20. Generation of 451 Ring1MNΔ mice was performed by crossing Ring1A-/-; Ring1B flox and Olig2::Cre. Ring1MNΔ 452 mice perish at birth. Generation of Ring1MNΔ;Hb9::GFP mice was performed by crossing 453 Ring1A-/-; Ring1B flox, Olig2::Cre mice to Hb9::GFP mice. Animal work was approved by the 454 Institutional Animal Care and use Committee of the NYU School of Medicine in accordance to 455 NIH guidelines. 456 457 Slide Immunohistochemistry 458 Embryos were fixed in 4% PFA for 1.5-2 hours at 4C, washed 5-6 times in cold PBS for 15-30 459 minutes each wash, and incubated overnight in 30% sucrose. Tissue was embedded in OCT, 460 frozen in dry ice, and sectioned at 16 µm on a cryostat. For antibody staining of sections, slides 461 of cryosections were placed in PBS for 5 minutes to remove OCT. Sections were then transferred 462 to humidified trays and blocked for 20-30 minutes in 0.75 ml/slide of PBT (PBS, 0.1% Triton) 16

463 containing 1% Bovine serum albumin (BSA). The blocking solution was replaced with primary 464 staining solution containing antibodies diluted in PBT with 0.1% BSA. Primary antibody 465 staining was performed overnight at 4C. Slides were then washed three times for 5 minutes each 466 in PBT. Fluorophore-conjugated secondary antibodies were diluted 1:500-1:1000 in PBT and 467 filtered through a 0.2 µm syringe filter. Secondary antibody solution was added to slides (0.75 468 ml/slide) and incubated at room temperature for 1 hour. Slides were washed three times in PBT, 469 followed by a final wash in PBS. Coverslips were placed on slides using 110 µl of Vectashield 470 (Vector Laboratories). 471 472 Antibodies against Hox proteins and MN subtypes were generated as described (Dasen et al., 473 2008; Dasen et al., 2005). Additional antibodies were used as follows: goat anti-Ring1B (Abcam, 474 1:2000), rabbit anti-Ring1B (Abcam, 1:5000), rabbit anti-Rybp (Abcam1:2000), rabbit anti-Yaf2 475 (Abcam, 1:2000), rabbit anti-Cbx2 (Bethyl, 1:5000) , rabbit anti-H3K27me3 (Cell Signaling, 476 1:2000).

477 478 In situ mRNA Hybridization 479 Probe templates were generated by RT-PCR and incorporated a T7 promoter sequence in the 480 antisense strand. Total RNA was first extracted from eviscerated E12.5 embryos using TRIzol 481 (Invitrogen). Genes of interest were amplified with the One Taq One-Step RT-PCR kit (NEB) 482 using 1 µg of RNA. After amplification by RT-PCR, a second PCR was performed to 483 incorporate a T7 promoter sequence. Antisense riboprobes were generated using the 484 Digoxigenin-dUTP (SP6/T7) labeling kit (Sigma-Aldrich). For in situ hybridization, sections 485 were first dried for 10-15 minutes at room temperature, placed in 4% PFA, and fixed for 10 486 minutes at room temperature. Slides were then washed three times for 3 minutes each in PBS, 487 and then placed in Proteinase K solution (1 µg/ml) for 5 minutes at room temperature. After an 488 additional PFA fixation and washing step, slides were treated in triethanolamine for 10 minutes, 489 to block positive charges in tissue. Slides were then washed three times in PBS and blocked for 490 2-3 hours in hybridization solution (50% formamide, 5X SSC, 5X Denhardt’s solution, 0.2 491 mg/ml yeast RNA, 0.1 mg/ml salmon sperm DNA). Prehybridization solution was removed, and 492 replaced with 100 µl of hybridization solution containing 100 ng of DIG-labeled antisense probe. 493 Slides were then incubated overnight (12-16 hours) at 72C. After hybridization, slides were

494 transferred to a container with 400 ml of 5X SSC and incubated at 72C for 20 minutes. During 495 this step, coverslips were removed using forceps. Slides were then washed in 400 ml of 0.2X 496 SSC for 1 hour at 72C. Slides were transferred to buffer B1 (0.1 M Tris pH 7.5, 150 mM NaCl) 497 and incubated for 5 minutes at room temperature. Slides were then transferred to staining trays 498 and blocked in 0.75 ml/slide of B1 containing 10% heat inactivated goat serum. The blocking 499 solution was removed and replaced with antibody solution containing 1% heat inactivated goat 500 serum and a 1:5000 dilution of anti-DIG-AP antibody (Roche). Slides were then incubated 501 overnight at 4C in a humidified chamber. The following day, slides were washed 3 times, 5 502 minutes each, with 0.75 ml/slide of buffer B1. Slides were then transferred to buffer B3 (0.1 M

503 Tris pH 9.5, 100 mM NaCl, 50 mM MgCl2) and incubated for 5 minutes. Slides were then 504 developed in 0.75 ml/slide of B3 solution containing 3.5 µl/ml BCIP and 3.5 µl/ml NBT for 12-

505 48 hours. After color development, slides were washed in ddH20 and coverslipped in Glycergel 506 (Agilent). A more detailed in situ hybridization protocol is available on our lab website 507 (http://www.dasenlab.com). 508 509 Wholemount Immunohistochemistry 510 For wholemount immunohistochemistry embryos were fixed in PFA for 2 hours, then bleached

511 for 24 hours at 4C in a 10% H2O2, 10% DMSO solution prepared in methanol. Embryos were 512 washed three times for 10 minutes each in methanol, followed by five washes for 10 minutes in 513 PBS. Primary antibodies were diluted in staining solution (5% BSA, 20% DMSO in PBS) and 514 specimens were incubated in staining solution on a rotator overnight at room temperature. 515 Samples were then washed three times for 5 minutes each in PBS, followed by four 1 hour 516 washes in PBS. Specimens were then incubated in secondary antibodies diluted in staining 517 solution overnight at room temperature. Samples were then washed three times for 5 minutes 518 each in PBS, followed by four 1 hour washes in PBS, a single 10 min wash in 50% methanol, 519 and three 20 minute washes in 100% methanol. Samples were transferred to glass depression 520 slides and tissue was cleared by incubating samples in BABB solution (1-part benzyl alcohol: 2- 521 parts benzyl benzoate). Confocal images of embryos were obtained from Z-stacks using Zen 522 software (Zeiss). Further details of wholemount staining protocols are available on our lab 523 website: (http://www.dasenlab.com).

524

525 In ovo chick electroporation 526 In ovo electroporation were performed on Hamburger Hamilton (HH) stage 13-14 chick embryos 527 and analyzed at HH stage 24-25. Fertilized chicken eggs (Charles River) were incubated in a 528 humidified incubator at 39C for 40-48 hours until they reached HH13-14. The top of the egg 529 shell was removed and a 1 µg/µl DNA (150-500 ng/µL expression plasmid and pBKS carrier 530 DNA) containing ~0.02% Fast green was injected into the central canal of the neural tube using a 531 sharpened glass capillary tube. Electrodes (Platinum/Iridium (80%/20%), 250 μm diameter, 532 UEPMGBVNXNND, FHC Inc.) were placed on both sides of the neural tube (4 mm separation) 533 and DNA was electroporated using an ECM 830 electroporator (ECM 830, BTX; 25V, 4 pulses, 534 50 ms duration, 1 second interval). Eggs were sealed with parafilm and incubated for 48 hours 535 prior to fixation. Results shown in figures are representative of at least 3 electroporated embryos 536 from two or more experiments in which electroporation efficiency in MNs was above 60%. 537 538 RNA preparation and Library Preparation 539 RNA was extracted from FACS purified MNs dissected from E12.5 mouse embryo (10-20,000 540 cells/segment), using the Arcturus Picopure RNA isolation kit. For on-column DNase treatment, 541 Turbo DNase was used (ambion, AM2238). Each samples used separated bar code for libraries. 542 RNA quality and quantity were measured with an Agilent Picochip using a Bioanalyzer, all 543 samples had quality scores between 9-10 RIN. For library preparation 10ng of total RNA was 544 used to generate cDNA, which was amplified with SMARTer Stranded RNA-Seq kit. 100ng of 545 cDNA were used as input to prepare the libraries (Takara, #634837), and amplified by 10 PCR 546 cycles. Samples were run in four 50-nucleotide paired-end read rapid run flow cell lanes with the 547 Illumina HiSeq 4000 sequencer. 548 549 RNAseq Data Analyses 550 Sequencing reads were mapped to the assembled reference genome (mm10) using the STAR 551 aligner (v2.5.0c) (Dobin et al., 2013). Alignments were guided by a Gene Transfer Format (GTF) 552 file. The mean read insert sizes and their standard deviations were calculated using Picard tools 553 (v.1.126) (http://broadinstitute.github.io/picard). The read count tables were generated using 554 HTSeq (v0.6.0) (Anders et al., 2015), normalized based on their library size factors using 555 DEseq2(Love et al., 2014), and differential expression analysis was performed. The Read Per 556 Million (RPM) normalized BigWig files were generated using BEDTools (v2.17.0) (Quinlan and 19

557 Hall, 2010) and bedGraphToBigWig tool (v4). To compare the level of similarity among the 558 samples and their replicates, we used two methods: principal-component analysis and Euclidean 559 distance-based sample clustering. All the downstream statistical analyses and generating plots 560 were performed in R environment (v3.1.1) (https://www.r-project.org/ ).

561 562 ATACseq 563 ATACseq was performed following previously described protocols(Buenrostro et al., 2015). 564 DNA was extracted from purified dissected mouse embryonic MNs. Cells were aliquoted and 565 washed twice in ice-cold 1× PBS. Cell pellets were resuspended in 10 mM Tris (pH 7.4), 10 mM 566 NaCl, 3 mM MgCl2, 0.1% NP-40 (v/v), 0.1% tween20, 0.01% Digitonin and 1% BSA, 567 centrifuged at 500 g for 5 min at 4°C. Pellets were resuspended in 12.5 μl of 2× tagmentation 568 DNA buffer, 1.25 μl Tn5 (Nextera DNA Sample Preparation Kit, FC-121–1030) and 11.25 μl of 569 water, and incubated at 37°C for 30 min. The sample was purified using the MinElute PCR 570 Purification Kit (Qiagen, 28004). PCR enrichment of the library was performed with custom- 571 designed primers and 2× NEB Master Mix. A qPCR reaction with 1× SYBR Green (Invitrogen), 572 custom-designed primers and 2× NEB Master Mix (New England Labs, M0541) was performed 573 to determine the optimal number of PCR cycles (one third of the maximum measured 574 fluorescence)(Buenrostro et al., 2013). The libraries were purified using the AMPure XP beads 575 (Beckman Coulter, A63880). High Sensitivity DNA ScreenTape (Agilent, 5067-5584) was used 576 to verify the fragment length distribution of the library. Library quantification was performed 577 using the KAPA Library Amplification kit on a Roche LightCycler 480. The libraries were 578 sequenced on an Illumina NovaSeq (100 cycles, paired-end). 579 580 ATACseq data analysis 581 All of the reads from the Sequencing experiment were mapped to the reference genome (mm10) 582 using the Bowtie2 (v2.2.4)(Langmead and Salzberg, 2012) and duplicate reads were removed 583 using Picard tools (v.1.126) (http://broadinstitute.github.io/picard/). Low quality mapped reads 584 (MQ<20) were removed from the analysis. The read per million (RPM) normalized BigWig files 585 were generated using BEDTools (v.2.17.0) (Quinlan and Hall, 2010) and the 586 bedGraphToBigWig tool (v.4). Peak calling was performed using MACS (v1.4.2)(Zhang et al., 587 2008) and peak count tables were created using BEDTools. Differential peak analysis was

588 performed using DESeq2 (Love et al., 2014). ChIPseeker (v1.8.0)(Yu et al., 2015) R package 589 was used for peak annotations and motif discovery was performed using HOMER (v4.10)(Heinz 590 et al., 2010). ngs.plot (v2.47) and ChIPseeker were used for TSS site visualizations and quality 591 controls. KEGG pathway analysis and Gene Ontology (GO) analysis was performed using the 592 clusterProfiler R package (v3.0.0)(Yu et al., 2012). To compare the level of similarity among the 593 samples and their replicates, we used two methods: principal-component analysis and Euclidean 594 distance-based sample clustering. The downstream statistical analyses and generating plots were 595 performed in R environment (v3.1.1) (https://www.r-project.org/).

596 Statistics 597 Samples sizes were determined based on previous experience and the number of animals and 598 definitions of N are indicated in the main text and figure legends. In figures where a single 599 representative image is shown, results are representative of at least two independent experiments, 600 unless otherwise noted. No power analysis was employed, but sample sizes are comparable to 601 those typically used in the field. Data collection and analysis were not blind. Graphs of 602 quantitative data are plotted as means with standard error of mean (SEM) as error bars, using 603 Prism 8 (Graphpad) software. Unless noted otherwise, significance was determined using 604 unpaired t-test in Prism 8 software, or using adjusted p-values. Exact p-values are indicated, 605 where appropriate, in the main text, figures, and figure legends. 606 607 Data Availability 608 RNAseq and ATACseq data are available through GEO (GSE175503). 609

610 Acknowledgements 611 We thank Kristen D’Elia, Sara Fenstermacher, Jessica Treisman, and Ed Ziff for discussion and 612 comments on the manuscript, and Rachel Kim and Orly Wapinski for assistance. We thank 613 Isabel Hidalgo and Susana Gonzalez for providing Ezh1flox mice, Alexander Tarakhovsky for 614 Ezh2 flox mice, Stefan Thor for Eed mutant embryos, Haruhiko Koseki for Ring1A-/-;Ring1B 615 flox ES cells, and Robert Klose for Rybp flox; Yaf2-/- ES cells. We also thank Genome 616 Technology Center and Cytometry and Cell Sorting Laboratory at NYU Langone. This work was 617 supported by NIH NINDS grants T32 GM007238, F31 NS087772 to AS, R01NS100897 to 618 EOM, R35 NS116858, R01 NS062822 and R01 NS097550 to JD.

619 620 Author Contributions 621 A.S. and J.S.D. designed experiments, interpreted data, and wrote the paper. S.P. performed in 622 situ hybridizations. A.M. and A.S. performed chick neural tube experiments. M.B. and E.O.M 623 designed and performed mouse ESC experiments. A.S. performed all mouse genetic analyses 624 and performed RNAseq and ATACseq. A. K.-J. analyzed ATACseq and RNAseq data. 625 626 Declaration of Interests 627 The authors declare no competing interests. 628

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Figure 3. Ring1 is required to restrict transcription factor expression in spinal MNs. (A) Venn diagrams showing the number of upregulated (left) and downregulated (right) genes in bracial (Br), thoracic (Th), and lumbar (Lu) MNs upon loss of Ring1 from MNs by RNAseq (log2-FC>2, FDR<0.1). (B) Heat map of top 100 upregulated genes (by log2-FC) in Ring1MNΔ mice and control MNs. Genes shown in blue are Hox genes and other transcription factor are shown in red. (C) Heat map of 45 genes downregulated in Ring1MNΔ mice in comparison to control MNs. (D) Plots of DESeq2 counts of genes associated with MN class identity. Each data point shows DESeq2 counts for each sample, and segment-specific counts are plotted together. Expression of Slc18a3 (Vacht) is reduced (padj.=0.026477) in Ring1MNΔ. (E) DESeq2 counts of genes associated with specific MN subtype identities were reduced in Ring1MNΔ mice. Counts for LMC and PMC markers are from Br segments, PGC from Th segments. Black bars shown in graphs indicate mean± SEM. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001. See also Supplementary Figure 3. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

upregulated downregulated Gata3 Rgs4 A brachial thoracic lumbar B G brachial thoracic lumbar H 5 Control 4 Control Ring1 MN∆ Ring1 MN∆ 10 10 4 3 Control Control 3

2 2 DESeq2 Counts Log Counts DESeq2 DESeq2 Counts Log Counts DESeq2 1 1 Ring1MN ∆ Ring1MN ∆ Br Th Lu Br Th Lu

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1 DESeq2 Log Counts DESeq2 DESeq2 Log Counts DESeq2 0 2 Ring1MN ∆ Ring1MN ∆ Br Th Lu Br Th Lu

Figure 4. Analyses of misregulated genes in Ring1 mutants. (A,C,E) In situ mRNA hybridization of selected upregulated genes from Ring1MNΔ RNAseq. Images show sections of brachial, thoracic, and lumbar segments from E12.5 control and Ring1MNΔ mice. (B,D,F) Graphs of DESeq2 counts for each upregulated gene in each segment. Data points show DESeq2 counts from MNs of individual animals from indicated segments. (G, I, K) Analysis of downregulated genes by in situ hybridization. Rgs4 and Uts2b displayed elevated expression in specific segmental levels in controls, suggesting that a subset of the commonly-downregulated genes are also Hox-dependent. Rgs4 expression is normally elevated in LMC neurons (panel G), while Uts2b is elevated in thoracic segments of controls (panel I) (H,J,L) Graphs of DESeq2 counts for each downregulated gene in each segment. See also Supplementary Figure 4. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

A HoxC genes HoxA genes HoxB genes 5 ** Control 5 5 Ring1 MN∆ ** * ns * *** 4 *** * *** 4 4 ns ns 10 ns ns * ns ns **** ** **** *** **** * ns 3 3 *** 3 * ns ** **** *** ns *** brachial 2 2 2 DESeq2 Counts Log

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Hoxc4Hoxc5Hoxc6Hoxc8Hoxc9 Hoxa1Hoxa2Hoxa3Hoxa4Hoxa5Hoxa6Hoxa7Hoxa9 Hoxc10Hoxc11Hoxc12Hoxc13 Hoxa10Hoxa11 Hoxa13 Hoxb1Hoxb2Hoxb3Hoxb4Hoxb5Hoxb6Hoxb7Hoxb8Hoxb9 Hoxa11os Hoxb13

B C brachial thoracic lumbar sacral Hox13 750 Control Hoxb13 500 Ring1 MN ∆ Hox12 250 FC DESeq2 Counts FCDESeq2 Hox11 Control Hox10 Hox9 Hoxc13 0

Hoxa9Hoxb9Hoxc9Hoxd9 Ring1 MN ∆ Hoxa10Hoxc10Hoxd10Hoxa11Hoxc11Hoxd11Hoxc12Hoxa13Hoxb13Hoxc13Hoxd13

Figure 5. Derepression of caudal Hox genes in Ring1 mutants. (A) DESeq2 counts of HoxC, HoxA, and HoxB cluster genes in brachial segments in control and Ring1MNΔ mice showing derepression of caudal Hox genes. Grey shaded regions highlight Hox genes that are derepressed in Ring1MNΔ mice. Hoxc9 does not show significant derepression, likely because it is normally expressed by caudal brachial MNs. Black bars shown in graphs indicate mean± SEM. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001. (B) Comparison of Hox9-Hox13 paralog gene derepression in brachial segments. Graph shows absolute fold changes of DESeq2 counts, showing a marked increase for caudal Hox13 paralogs in Ring1MNΔ mice. Each data point shows individual counts for Ring1 mutants/average of controls. (C) In situ of Hoxb13 and Hoxc13 mRNA transcripts in E12.5 embryos. Hoxb13 is normally not detectable in spinal cord, but is derepressed in MNs in Ring1MNΔ mice. Hoxc13 transcripts are normally restricted to sacral segments but derepressed in rostral segments in Ring1MNΔ mice. See also Supplementary Figure 5. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

A ATACseq gain ATACseq loss B

Br Br ATACseq Th ATACseq Br 1016 235 Br RNAseq 823 Th RNAseq 975 15 0 714 18 3 common: 264 101 Hoxa11 73 Hoxa13 593 85 Hoxb13 103 70 324 5 5 Hoxc11 39 5 42 Hoxc12 65 Hoxc13 Lu 171 29 156 23 391 Hoxd11 222 Lu Hoxd13 Th Th

C Hoxc13 Foxa2 Foxd1 Foxg1 Lhx2 Pitx2 Six2 Six6 [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] RNAseq

[0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] Control ATACseq

[0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] [0 - 0.88] RNAseq

[0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] [0 - 1.76] RIng1MN ∆ ATACseq

Hoxc13 Foxa2 Foxd1 Foxg1 Lhx2 Pitx2 Six2 Six6

D Hoxc13 Hoxa13 Hoxb13 Hoxd13 Hoxc6 E [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] 43 kb RNA Control ATAC brachial

[0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] RNA RIng1MN ∆ ATAC Control thoracic Hoxc13 Hoxc12 Hotair Hoxc11 [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] 21 kb RNA lumbar Control ATAC [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] RNA RIng1MN ∆ ATAC brachial [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] Hoxa11 Hoxa11os Hoxa13 92 kb RNA thoracic RIng1MN ∆ [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] [0 - 1.05] Control ATAC RNA lumbar RIng1MN ∆ ATAC Hoxc13 Hoxa13 Hoxb13 Hoxd13 Hoxc6 Hoxb13 Gm11538 Hoxb9

Figure 6. Ring1 is essential for maintaining chromatin topology of cell fate determinants. (A) Proportional Venn diagram showing the number of genes that gain (left) or lose (right) chromatin accessibility in brachial, thoracic and lumbar segments in Ring1MNΔ mice. (B) Venn diagram of upregulated genes from RNAseq and genes that gained accessibility in ATACseq of brachial and thoracic segments. (C) IGV browser views of selected genes that are upregulated and gained chromatin accessibility in controls and Ring1MNΔ mice in brachial segments. (D) IGV browser views of ATACseq tracks in controls and Ring1MNΔ mice in brachial, thoracic, and lumbar segments. Hox13 genes gain chromatin accessibility in each segment, while reduced brachial Hoxc6 expression is not associated with a loss of chromatin accessibility. (E) Comparison of IGV browser views of RNAseq and ATACseq tracks of caudal HoxC, HoxA and HoxB genes in brachial segments. See also Supplementary Figure 6. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

127 kb A [0 - 32] RNAseq ESC-MNs [0 - 32] iHoxc13 RNAseq

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C Hoxc4 Hoxa13/GFP Hoxa5 Hoxa13/GFP 1.0 B 0.8

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0.4 Hoxa13-GFP total Hoxa13-GFP Hoxc6 Hoxa13/GFP Hoxc8 Hoxa13/GFP Hoxa13-GFP(+) marker(+) Hoxa13-GFP(+) 0.2

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Hoxc4 Hoxa5 Hoxc6 Hoxc8 Hoxc9 Hoxc10Hb9 Isl1 Br Th Lu

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0.0

Hoxc4 Hoxa5 Hoxc6 Hoxc8 Hoxc9 Hoxc10Hb9 Isl1 Br Th Lu

Figure 7. Hox13 proteins repress multiple Hox4-Hox10 paralogs. (A) Effects of Hoxc13 induction in ESC-MNs. Top panels show IGV browser views (log scale) of RNAseq (red) and ChIPseq tracks (green) at the HoxC cluster. Induced Hoxc13 binds near multiple HoxC genes and represses expression of Hoxc4 and Hoxc5. Bottom panels show ATACseq in brachial control and Ring1 mutant MNs at the HoxC cluster (blue). (B) Misexpression of mHoxa13-ires-nucGFP represses more rostral Hox genes. Hoxc4, Hoxa5, Hoxc6 and Hoxc8 were analyzed in brachial segments, Hoxc9 in thoracic segments, and Hoxc10 in lumbar segments. Hb9 and Isl1 images are from brachial segments. (C,D) Quantification of percentage of electroporated cells which retained the expression of indicated Hox protein and MN markers upon Hoxa13 or Hoxb13 misexpression. Quantified electroporated cells were selected from the ventrolateral spinal cord, where MNs reside. Hb9 and Isl1/2 quantification shows percentage of GFP+ neurons that express either marker, and are shown for Br segments in panel B, and Br and Th segments in C. Data in graphs shown as mean± SEM, averaged from n=3 animals, 4 sections each. See also Supplementary Figure 7. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license. Hoxc6 Hoxc9 Hoxc10 Hb9 H3K27me3 A B c D E Control Control Th Th Lu Br Eed NE ∆ Ezh MN ∆ Th Th Lu Hoxc6 Hoxc9 Hoxc10 F G H Control Control Br Th Br Th Lu Eed NE ∆ Eed NE ∆ Br Th Br Th Lu Hoxc4 Hoxa5 Hoxd10 Hoxc9 Hoxd10 Hoxc10 I J K Control Br Br Br Th Th Lu

Ring1 MN ∆ Br Br Br Th Th Lu Rybp Hb9 Hoxc6 Hoxc9 Hoxc10 Foxp1 L Control Br Br Th Br Th Th Lu Br

Br Br Br Th Lu Br Rybp/Yaf2 MN ∆ Rybp/Yaf2

Supplementary Figure 1. Effects of PRC mutations on MN differentiation. (A-C) Expression of Hoxc9 in thoracic segments and Hoxc10 in lumbar segments is normal in E11.5 EzhMNΔ mice (Eed flox/flox, Sox1::Cre/+), and no ectopic thoracic Hoxc6 expression is detected. (D,E) In EedNEΔ mice (H3K27me3 is depleted throughout the spinal cord at E12.5, while Hb9 expression is unaffected. Some H3K27me3 is detected in the floor plate of EedNEΔ mice (boxed region). (F-H) Brachial Hoxc6, thoracic Hoxc9, and lumbar Hoxc10 are detected normally in EedNEΔ mice. (I) Brachial Hoxc4 and Hoxa5 are lost from MNs in Ring1MNΔ mice, and Hoxd10 is ectopically expressed in brachial MNs. (J) Hoxc9 expression is reduced in thoracic segments of Ring1MNΔ mice, and Hoxd10 is expressed. (K) Expression of Hoxc10 is retained at lumber levels of Ring1MNΔ mice. (L) Analysis of E12.5 Rybp/Yaf2MNΔ mice (Rybp flox/flox; Yaf2-/-; Olig2::Cre/+). Rybp expression is selectively depleted from MNs. Hb9 and indicated Hox proteins are expressed normally. (which wasnotcertifiedbypeerreview)istheauthor/funder,whohasgrantedbioRxivalicensetodisplaypreprintinperpetuity.Itmade m l I e ( SupplementaryFig bioRxiv preprint umbar segments of segments of umbar sl1/2 A,B) Foxp1 expression Foxp1atthoracic, A,B) expression is brachial, andunaffected lumbar in levels xpression xpression in lumbar segment of ice. ice. +

) in in ) rostral segments ofbrachial doi:

Hb9 Isl1/2 Hb9 Isl1/2 Scip Isl1/2 Foxp1 Foxp1 https://doi.org/10.1101/2021.08.09.455667 ure Ring1 Lu Br Br Lu C Lu Th Br A D F

Control Control 2. Analyses 2. subtypes mice. MN of PRC mutant in Control Control MN Δ mice. (F) Expression of HMC markers (Hb9, Isl1/2) in brachial and lumbar segments of of Expression (Hb9, (F) Isl1/2) HMC and inlumbar brachial segments of markers mice. Ring1 Ring1 MN available undera Ring1 MN∆ Ring1 MN∆ Ring1 MN∆ Ezh MN∆ Δ mice. (D) Reduced (D) (PMC) (Scip mice. of expression column phrenicmotor markers MN Δ mice. (E) Expression of (E) MMC Expression mice. (Hb9, Lhx3) in andmarkers brachial neuron CC-BY 4.0Internationallicense ; this versionpostedAugust11,2021.

Hb9 Lhx3 Hb9 Lhx3 Foxp1 Lu Br E Lu Th Br B Control Control

Ezh . MN Δ and Ring1 MN∆ Eed NE∆ The copyrightholderforthispreprint Eed NE Δ mice.Reduced (C) Foxp1 Ring1 + , MN Δ

A B Neurotransmitter genes 5

4 10

group: Control 3

Ring1 MN∆ *** 2 condition: brachial

thoracic Log Counts DESeq2 1 lumbar Control Ring1 MN∆ 0

Gad2 (Gad65)Gad1 (Gad67) Slc17a7Slc17a6 (vGlut1)Slc32a1 (vGlut2) (vGAT)

Hoxc13

Hoxc12

Hoxa13

Supplementary Figure 3. RNAseq analyses of Ring1 mutant mice. (A) PCA plot of RNAseq samples. (B) Plot of neurotransmitter encoding genes in control and Ring1MNΔ mice. Commonly used gene names are shown in parenthesis. Expression of Slc17a7 is increased in Ring1MNΔ mice, although the absolute DESeq2 counts were still relatively low (46.3 +/- 7.2 in controls versus 111.1 +/- 9 in Ring1MNΔ mice, p=0.000012). Elevated expression of Gad1 in controls and mutants may be due to presence of spinal interneurons in sorted samples. (C) Volcano plot of upregulated (red) and downregulated (blue) genes. Three caudal Hox genes are indicated. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

upregulated upregulated Foxg1 Shox2 A brachial thoracic lumbar B G brachial thoracic lumbar H Control 4 Control 3 Ring1 MN∆ Ring1 MN∆ 10 10

2 3 Control Control

1 2 DESeq2 Counts Log Counts DESeq2 DESeq2 Counts Log Counts DESeq2 0 1 Ring1MN ∆ Ring1MN ∆

Br Th Lu Br Th Lu Six6 Nkx2.1 C brachial thoracic lumbar D Control I brachial thoracic lumbar J Control Ring1 MN∆ 3 Ring1 MN∆ 3 10 10

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1 1 DESeq2 Counts Log Counts DESeq2 DESeq2 Counts Log Counts DESeq2 0 0 Ring1MN ∆ Ring1MN ∆ Br Th Lu Br Th Lu downregulated E Pitx2 F K Gabra2 L brachial thoracic lumbar brachial thoracic lumbar 4 Control 4 Control Ring1 MN∆ Ring1 MN∆ 10 10 3 Control Control 2 3

1 DESeq2 Counts Log Counts DESeq2 DESeq2 Counts Log Counts DESeq2 0 2 Ring1MN ∆ Ring1MN ∆ Th Lu Br Th Lu Br

Supplementary Figure 4. Validation of downregulated genes from Ring1MNΔ RNAseq. (A,C,E,G,I) In situ hybridization of selected upregulated genes from the Ring1MNΔ RNAseq. Images show sections at brachial, thoracic, and lumbar levels from E12.5 control and Ring1MNΔ mice. (B,D,F,H,J) Graphs of DESeq2 counts for each upregulated gene in each segments. (K) Analysis of the downregulated gene Gabra2 by in situ hybridization. (L) Graph of DESeq2 counts for Gabra2 in each segment. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

A HoxD genes D 5 Control Hoxc6 Hoxc9 Hoxc9 Ring1 MN∆ Br Br Th

3 Control brachial 2 Br Br Th

1 Ring1 MN ∆

Hoxd1Hoxd3Hoxd4Hoxd8Hoxd9 Hoxd10Hoxd11Hoxd12Hoxd13 B HoxA genes HoxB genes HoxC genes HoxD genes 5 5 5 5

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thoracic 2 2 2 2 DESeq2 Counts Log

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Hoxa1Hoxa2Hoxa3Hoxa4Hoxa5Hoxa6Hoxa7Hoxa9 Hoxb1Hoxb2Hoxb3Hoxb4Hoxb5Hoxb6Hoxb7Hoxb8Hoxb9 Hoxc4Hoxc5Hoxc6Hoxc8Hoxc9 Hoxd1Hoxd3Hoxd4Hoxd8Hoxd9 Hoxa10Hoxa11 Hoxa13 Hoxb13 Hoxc10Hoxc11Hoxc12Hoxc13 Hoxd10Hoxd11Hoxd12Hoxd13 Hoxa11os C 5 HoxA genes 5 HoxB genes 5 HoxC genes 5 HoxD genes

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Supplementary Figure 5. Derepression of caudal Hox genes in Ring1MNΔ mice. (A-C) DESeq2 counts of HoxA, HoxB, HoxC, and HoxD gene clusters in brachial (A), thoracic (B), and lumbar (C) segments. Gray-shaded regions highlight Hox genes that are derepressed in MNs of Ring1MNΔ mice. In lumbar segments, expression of Hox10 genes is not reduced in Ring1 mutants, although Hox13 genes are derepressed. (D) In situ hybridization of Hoxc6 and Hoxc9 in Ring1MNΔ mice at brachial (Br) and thoracic (Th) levels at E12.5. In brachial segments, expression of Hoxc6 is lost from MNs, and Hoxc9 is expressed, similar to the analyses of Hox protein expression. In thoracic segments, Hoxc9 expression is reduced in MNs. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

A [0 - 1.37] 133 kb brachial

[0 - 1.37] thoracic

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Hoxc13 Hoxc12 Hotair Hoxc11 Hoxc10 Hoxc9 Hoxc8 Hoxc6 Hoxc4

B [0 - 1.80] 111 kb brachial

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[0 - 1.80] lumbar

Hoxa1 Hoxa2 Hoxa3 Hoxaas3 Mira Hoxa9 Hoxa10 Hoxa11 Hoxa13

C Common RNAseq ATACseq Br RNAseqUP Th RNAseqUP All segements (19): Br and Th segements (73) 75 9230102O04Rik 5033428I22Rik Foxb2 Lbx1 Six3os1 94 25 Cdkn2a 9230102O04Rik Foxc1 Lhx2 Six6 0 75 Lu RNAseqUP Foxb2 Arhgef3 Foxd1 Mical2 Skor2 214 Foxc1 Atoh1 Foxg1 Nkx2-1 Smad3 50 0 Foxg1 C130021I20Rik Gata3 Nkx2-2 Sox1 2 Hoxa13 C1ql1 Gata4 Nkx2-3 Svil 3 Hoxc13 Cdkn2a Grb14 Npr3 Tal1 9 19 Lbx1 Cdkn2b Hlx Nr2e1 Tbx18 0 Lhx2 Col4a2 Hmx1 Olig2 Tbx2 3 97 Nkx2-3 Hotair Osr2 Tbx3 917 Crabp1 528 1 Otx1 Dlx1 Hoxa11 Otx1 Tlx1 Pax9 Dlx2 Hoxa11os Pax9 Tlx3 62 Pitx2 Hoxa13 Phox2b Vax2os Br ATAC UP Dmrt3 Six2 Hoxb13 Pitx2 Zfpm1 22 212 1 Dmrta2 Six3 E030030I06Rik Hoxc11 Rbpms Zic1 Serpine2 Lu ATAC UP Tal1 Emx2 Hoxc12 Tlx1 En2 Hoxc13 Six1 Vax2os Foxa1 Hoxd11 Six2 Zic1 Foxa2 Hoxd13 Six3

Th ATAC UP

D Hoxc13 Foxa2 Foxd1 Foxg1 Lhx2 Pitx2 Six2 Six6 [0 - 2.46] Control

[0 - 2.46] ATACseq Ring1MN ∆

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Hoxc13 Foxa2 Foxd1 Foxg1 Lhx2 Pitx2 Six2 Six6

Supplementary Figure 6. Integrative analysis of RNA and ATACseq results (A,B) IGV browser view of ATACseq tracks of the HoxC cluster (A) and HoxA cluster (B) in control mice. The peaks highlighted with green box indicate regions that gain accessibility in more caudal segments. (C) Venn diagram of upregulated genes (RNAseqUP) and genes that gain chromatin accessibility (ATACseqUP) in all segments. Genes that are upregulated and gained accessibility for all segments (19 genes) or Br and Th segments (73 genes) are listed on the right. Hox genes are highlighted in blue, other transcription factors in red. (D) IGV browser view comparing ATACseq tracks from brachial segments (control and Ring1MNΔ mice) with Ring1B ChIPseq tracks from ES and neural progenitor cells (NPC). Regions that gain accessibility in Ring1 mutants correspond to regions bound by Ring1B in ES cells. bioRxiv preprint doi: https://doi.org/10.1101/2021.08.09.455667; this version posted August 11, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

A 111 kb

[0 - 19] RNAseq ESC-MNs

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Hoxc5 Gm30528 Hoxc4

Supplementary Figure 7. Comparison of Hoxc13 binding and accessibility in control and Ring1MNΔ MNs (A) Effects of Hoxc13 expression in ESC-MNs at the HoxA locus. Top panels show IGV browser views (log scale) of RNAseq (red) and ChIPseq tracks (green). Induced Hoxc13 represses expression of Hoxa1-Hoxa5 genes and binds near multiple HoxA genes. Bottom panels show ATACseq in brachial control and Ring1 mutant MNs at the HoxA cluster (blue). Bottom panels show ATACseq in brachial control and Ring1 mutant MNs at the HoxA cluster. (B,C) Magnified views of Hoxc13 binding, RNA expression, and ATACseq results at Hox4-Hox5 genes in the HoxA and HoxC clusters.