Introduction

The embryonic neural crest is a multipotent progenitor cell population that gives rise to peripheral neurons, glia, Schwann cells, melanocytes, and diverse mesenchymal cells such as osteoblasts, chondrocytes, fibroblasts, cardiac mesenchyme, and cardiomyocytes (Bronner and LeDouarin, 2012; Simões-Costa and Bronner, 2015). Following induction in the neural tube, cranial neural crest cells undergo dorsolateral migration to the pharyngeal arches and differentiate to form craniofacial bones and cartilage (Minoux and Rijli, 2010). To do so, cranial neural crest cells express transcription factors such as Sox9, Sox10, and Twist1 that specify cell fate decisions in derivates of the neural crest (Bertol et al., 2022; Cheung et al., 2005). Pre-migratory neural crest cell specification is partially controlled by the epigenetic regulator Polycomb repressive complex 2 (PRC2), an H3K27 histone methyltransferase that is broadly responsible for chromatin compaction and transcriptional silencing (Margueron and Reinberg, 2011). The catalytic activity of PRC2 is comprised of four core subunits: (1) enhancer of zeste homologue 1 (Ezh1) or Ezh2, (2) suppressor of zeste 12 (Suz12), (3) RBBP4 or RBBP7, and (4) embryonic ectoderm development (Eed) (Piunti and Shilatifard, 2021). Eed binds to H3K27 trimethylation peptides (H3K27me3) and stabilizes Ezh2 for allosteric activation of methyltransferase activity and on-chromatin spreading of H3K27 methylation. Eed is required for stem cell plasticity, pluripotency, and maintaining cell fate decisions, but all PRC2 core subunits are required for embryonic development and loss of any individual subunit is embryonic lethal around gastrulation (Faust et al., 1995; O’Carroll et al., 2001; Pasini et al., 2004). Eed null embryos can initiate endoderm and mesoderm induction but have global anterior-posterior patterning defects in the primitive streak (Faust et al., 1995; Schumacher et al., 1996) and genome-wide defects in H3K27 methylation (Montgomery et al., 2007).

Ezh2 has been studied in the context of pre-migratory neural crest development. In mice, Wnt1-Cre Ezh2^Fl/Fl embryos fail to develop skull and mandibular structures due to de-repression of Hox transcription factors in cranial neural crest cells (Ferguson et al., 2018; Kim et al., 2018; Schwarz et al., 2014), and loss of Ezh2 in mesenchymal precursor cells causes skeletal defects and craniosynostosis in Prrx1-Cre Ezh2^Fl/Fl embryos (Dudakovic et al., 2015). Ezh2 also regulates neural crest specification in xenopus (Tien et al., 2015), and Col2-Cre Eed^Fl/Fl mice have kyphosis and accelerated hypertrophic differentiation due to de-repression of Wnt and TGF-β signaling in chondrocytes (Mirzamohammadi et al., 2016). Genetic and molecular interactions allow Eed to repress Hox genes to maintain vertebral body identity during mouse development (Kim et al., 2006), but the embryonic, cellular, and molecular consequences of Eed activity in craniofacial development are incompletely understood. To address this gap in our understanding of epigenetic mechanisms that may contribute to craniofacial development, we conditionally deleted Eed from the migratory neural crest and its derivatives. Our results suggest that Eed is required for craniofacial osteoblast differentiation from post-migratory neural crest mesenchymal stem cells, and that Eed regulates diverse transcription factor programs that are required for mesenchymal cell proliferation, differentiation, and osteogenesis. More broadly, these data show that Eed is required at early post-migratory stages in neural crest progenitors of craniofacial structures.

Results

Loss of Eed after neural crest induction causes severe craniofacial malformations

To determine if Eed is required for the development of neural crest derivatives, homozygous floxed Eed alleles (Eed^Fl/Fl) (Yu et al., 2009) were used to conditionally delete Eed following neural crest cell induction using Cre under control of the Sox10 promoter, which is expressed in the migratory neural crest at embryonic day E8.75 and results in complete recombination in post-migratory neural crest cells by E10.5 (Matsuoka et al., 2004; Niethamer et al., 2020). Sox10-Cre Eed^Fl/Fl mice were not recovered past postnatal day 0, suggesting that loss of Eed following induction of the neural crest is embryonic lethal (Fig. 1a). Sox10-Cre Eed^Fl/Fl embryos were recovered at expected genotypic frequencies from embryonic day E9.5 to E17.5 (Fig. 1a), and there were no differences in the penetrance or severity of Sox10-Cre Eed^Fl/Fl phenotypes whether Cre was maternally or paternally inherited, as has been reported for other phenotypes arising in cells expressing Sox10-Cre (Crispino et al., 2011; Luo et al., 2020). No overt phenotypes were identified at early post-migratory neural crest cell stages E10.5 or E11.5, but craniofacial malformations were seen in Sox10-Cre Eed^Fl/Fl embryos starting at E12.5 that increased in severity throughout the remainder of embryonic development (Fig. 1b). Sox10-Cre Eed^Fl/Fl phenotypes were broadly consistent with impaired development of craniofacial structures (Ferguson et al., 2018; Schwarz et al., 2014), including frontonasal and mandibular hypoplasia, a prominent telencephalon and exencephaly resulting from underdevelopment of the viscerocranium and frontal calvarium, collapsed nasal cavity bones, and microtia compared to controls (Fig. 1c). Sox10-Cre Eed^Fl/Fl embryos had increased pupillary distance and craniofacial width (Supplementary Fig. 1a-d), and whole mount DAPI imaging of Sox10-Cre Eed^Fl/Fl embryos revealed severe frontonasal dysplasia with underdeveloped midface and mandible, and an irregular and corrugated facial structure compared to controls (Fig. 1d). Although PRC2 is ubiquitously expressed in the developing embryo (O’Carroll et al., 2001), Sox10-Cre Eed^Fl/Fl embryos did not have cardiac outflow tract defects or other structural heart malformations that can arise from impaired neural crest differentiation (Lindsay et al., 2001, 1999; Nakamura et al., 2009) (Supplementary Fig. 2a-d and Movie 1, 2). Instead, fetal echocardiography showed subtle structural and functional changes but preserved ejection fraction and fraction shortening in Sox10-Cre Eed^Fl/Fl embryos compared to controls (Supplementary Fig. 2d).

Fig. 1.

Eed regulates craniofacial mesenchymal cell differentiation and proliferation from the early post-migratory neural crest

Skeletal stains, computed tomography (microCT), histology, and immunofluorescence were used to shed light on how loss of Eed impacts craniofacial development. Whole mount alcian blue/alizarin red staining of Sox10-Cre Eed^Fl/Fl embryos revealed hypoplasia of the viscerocranium, including reduced frontal, temporal, maxillary, and mandibular bones, and complete loss of the tympanic ring and premaxillary and nasal bones compared to controls (Fig. 2a, b and Supplementary Fig. 3a). MicroCT validated these findings, showing fragmentation and hypoplasia of the frontal, temporal, maxillary, and mandibular bones, loss of frontal calvarial fusion, and absence of tympanic ring and nasal bones (Fig. 2c-e and Movie 3, 4). H&E histology showed that frontal calvarium reduction resulted in anteriorly displaced brain structures in Sox10-Cre Eed^Fl/Fl embryos compared to controls, with the midbrain and cortex in the same coronal plane as the nasal cavity and developing mandible (Fig. 3a and Supplementary Fig. 3b). Most of the midfacial and mandible were absent in Sox10-Cre Eed^Fl/Fl embryos, and the tongue and masseter muscles were underdeveloped (Fig. 3a).

Fig. 2.

Fig. 3.

Immunofluorescence demonstrated disorganized ALPL and Osteocalcin, markers of osteoblasts and mineralizing bone (Liu et al., 2018), in craniofacial tissues from Sox10-Cre Eed^Fl/Fl embryos compared to controls (Fig. 3b and Supplementary Fig. 3c). Immunofluorescence for Sox9, a marker of chondrocytes that are derived from the neural crest (Mori-Akiyama et al., 2003; Yan et al., 2002), was also disorganized in Sox10-Cre Eed^Fl/Fl embryos compared to controls (Fig. 3b). There was a small increase in immunofluorescence labeling index for cleaved Caspase 3 (Supplementary Fig. 3d, e), suggesting that apoptosis may play a subtle role in craniofacial phenotypes from Sox10-Cre Eed^Fl/Fl embryos compared to controls. In contrast to Wnt1-Cre Ezh2^Fl/Fl embryos (Ferguson et al., 2018; Schwarz et al., 2014), the craniofacial region of Sox10-Cre Eed^Fl/Fl embryos had marked decreased immunofluorescence labeling index for Ki67 (Fig. 3c, 3d), a marker of cell proliferation (Gerdes et al., 1984), and decreased immunofluorescence staining for Vimentin (Vim), a marker of mesenchymal cells (Mendez et al., 2010) (Fig. 3c). Immunofluorescence for Runx2, a transcription factor that is required for osteoblast differentiation and proliferation (Kawane et al., 2018; Komori, 2009), was also decreased in craniofacial tissues from Sox10-Cre Eed^Fl/Fl embryos compared to controls (Fig. 3e and Supplementary Fig. 3f). In support of these findings, BrdU labeling index was decreased in the craniofacial region of Sox10-Cre Eed^Fl/Flembryos compared to controls (Supplementary Fig. 4a, b), Eed, Runx2, Ki67, and ALPL were decreased in primary Sox10-Cre Eed^Fl/Flcraniofacial cell cultures compared to controls (Fig. 3f and Supplementary Fig. 4c), and there was no change in immunofluorescence for Sox10 in either primary Sox10-Cre Eed^Fl/Fl craniofacial cell cultures or Sox10-Cre Eed^Fl/Fl embryos compared to controls (Supplementary Fig. 4c, e). These results suggest craniofacial mesenchymal differentiation, proliferation, and osteogenesis are impaired in Sox10-Cre Eed^Fl/Fl embryos (Honoré et al., 2003; Kim et al., 2003).

Eed regulates craniofacial mesenchymal stem cell, osteoblast, and proliferating mesenchymal cell fate from the early post-migratory neural crest

To define cell types and gene expression programs underlying craniofacial phenotypes in Sox10-Cre Eed^Fl/Fl embryos, single-cell RNA sequencing was performed on litter-matched E12.5 Sox10-Cre Eed^Fl/WT and Sox10-Cre Eed^Fl/Fl heads (n=3 biological replicates per genotype). Uniform manifold approximation and projection (UMAP) analysis of 63,730 single-cell transcriptomes revealed 23 cell clusters (C0-C22) that were defined using automated cell type classification (Ianevski et al., 2022), cell signature genes, cell cycle analysis, and differentially expressed cluster marker genes (Fig. 4a, Supplementary Fig. 5, 6, and Supplementary Table 1, 2). Differentiating osteoblasts marked by Runx2 (C0) and proliferating mesenchymal cells marked by Ki67 (C7) were enriched in Sox10-Cre Eed^Fl/WT samples (Fig. 4b-d). Mesenchymal stem cells marked by Col6a3 (C4) or Dcn (C5) were enriched in Sox10-Cre Eed^Fl/Flsamples (Jang et al., 2016; Lamandé et al., 2006) (Fig. 4b-d), suggesting that loss of Eed prevents craniofacial mesenchymal stem cell differentiation. There were subtle differences in the number of interneurons (C17, 1.4% versus 1.9% of cells, p=0.01), Schwann cells (C19, 1.0 versus 1.3% of cells, p=0.03), pericytes (C20, 0.7% vs 0.9%, p=0.05), and spinal neurons (C21, 0.5% vs 0.9% of cells, p=0.003) in Sox10-Cre Eed^Fl/WT versus Sox10-Cre Eed^Fl/Fl samples, but each of these comprised a small minority of the recovered cell types (Student’s t tests) (Supplementary Table 1). There were no differences between genotypes in the number of chondrocytes (C10, 3.1% versus 2.7%, p=0.09), fibroblasts (C11, 2.6% versus 2.7%, p=0.24), endothelia (C16, 1.9% vs 1.6%, p=0.22), hematopoietic cells, or other cell types recovered at E12.5 (Student’s t tests) (Supplementary Table 1).

Fig. 4.

Differential expression analysis of single cell transcriptomes from differentiating osteoblasts (C0), which were enriched in Sox10-Cre Eed^Fl/WT samples (Fig. 4c), compared to mesenchymal stem cells marked by Col6a3 (C4), which were enriched in Sox10-Cre Eed^Fl/Fl samples (Fig. 4c), showed Sox5 and Sox6 were reduced in mesenchymal stem cells (Fig. 4e, Supplementary Fig. 7a, and Supplementary Table 3). These data are consistent with the known role of Sox transcription factors in craniofacial specification and development (Smits et al., 2001). Iroquois homeobox (Irx) transcription factors 3 and 5, which contribute to craniofacial osteogenesis and mineralization (Bonnard et al., 2012; Cain et al., 2016; Tan et al., 2020), were also suppressed in mesenchymal stem cells compared to differentiating osteoblasts (Fig. 4e, Supplementary Fig. 7a, and Supplementary Table 3).

Differential expression analysis of single cell transcriptomes from differentiating osteoblasts compared to mesenchymal stem cells marked by Dcn (C5), which were also enriched in Sox10-Cre Eed^Fl/Fl samples (Fig. 4c), showed Mecom, Trps1, and Ptch1 were suppressed and Sparc and Tnmd were enriched in mesenchymal stem cells (Fig. 4f, Supplementary Fig. 7b, and Supplementary Table 4). Loss of Mecom causes craniofacial malformations in mice and zebrafish (Shull et al., 2020), Ptch1 is a key component of the Hedgehog pathway that is crucial for craniofacial osteogenesis (Jeong et al., 2004; Metzis et al., 2013), and Trps1 regulates secondary palate and vibrissa development (Cho et al., 2019; Fantauzzo and Christiano, 2011). Consistently, Sox10-Cre Eed^Fl/Fl samples had decreased vibrissa compared to controls (Fig. 1d and Supplementary Fig. 3b). Sparc drives morphogenesis of the pharyngeal arches and inner ear (Rotllant et al., 2008), and Tnmd regulates mesenchymal differentiation (Shukunami et al., 2006). Pax3 and Pax7, key regulators of neural crest migration and development of skeletal structures (Maczkowiak et al., 2010), were also suppressed in mesenchymal stem cells compared to differentiating osteoblasts (Fig. 4f, Supplementary Fig. 7b, and Supplementary Table 4).

Differential expression analysis of single cell transcriptomes from the two clusters of mesenchymal stem cells that were marked by Col6a3 (C4) versus Dcn (C5), both of which were enriched in Sox10-Cre Eed^Fl/Fl samples (Fig. 4c), showed Twist2, Irx3, and Irx5, which regulate mesenchymal differentiation to osteoblasts (Lee et al., 2000; Tamamura et al., 2017; Tan et al., 2020), distinguished mesenchymal stem cell populations (Fig. 4g, Supplementary Fig. 7c, and Supplementary Table 5).

To determine if there were differences in the gene expression programs from differentiating osteoblasts (C0), mesenchymal stem cells (C4, C5), or proliferating mesenchymal cells (C7) in Sox10-Cre Eed^Fl/WT versus Sox10-Cre Eed^Fl/Fl samples, differential expression analysis was performed on single-cell transcriptomes from these clusters between genotypes (Supplementary Table 6). Consistent with craniofacial findings after loss of Ezh2 (Ferguson et al., 2018; Kim et al., 2018; Schwarz et al., 2014) and vertebral body findings after loss of Eed (Kim et al., 2006), Hox transcription factors were broadly de-repressed in mesenchymal cell populations from Sox10-Cre Eed^Fl/Flsamples compared to controls (Fig. 4h). PRC2 targets Sp7 in bone marrow stroma cells (Liu et al., 2013) and regulates Wnt signaling in dental stem cells (Jing et al., 2016). Differential expression analysis showed that Sp7 was enriched in mesenchymal single-cell transcriptomes from Sox10-Cre Eed^Fl/WT samples and the Wnt and stem cell regulator gene Gata4 was enriched in mesenchymal single-cell transcriptomes from Sox10-Cre Eed^Fl/Fl samples (Fig. 4h). Mki67, Pax3, and Pax7 were enriched in mesenchymal single-cell transcriptomes from Sox10-Cre Eed^Fl/WT samples, and the cell cycle inhibitors Cdkn2a and Cdkn2b were enriched in mesenchymal single-cell transcriptomes from Sox10-Cre Eed^Fl/Fl samples (Fig. 4h). In sum, these data show that Eed specifies craniofacial osteoblast differentiation and mesenchymal cell proliferation by regulating diverse transcription factor programs that are required for specification of post-migratory neural crest cells, and that loss of Eed inhibits mesenchymal stem cell differentiation and mesenchymal cell proliferation. These data contrast with the function of Eed in chondrocytes, where genetic inactivation of Eed at a later stage in mesenchymal development accelerates hypertrophic differentiation, leading to hypoxia and cell death (Mirzamohammadi et al., 2016).

Discussion

Epigenetic regulation of the neural crest, which differentiates into diverse mesenchymal derivatives, is incompletely understood. Here we identify the PRC2 core subunit Eed as a potent regulator of craniofacial development after induction of the neural crest. Conditional deletion of Eed using Sox10-Cre causes severe craniofacial malformations that are consistent with impaired differentiation and proliferation of cells arising from the pharyngeal arches (Frisdal and Trainor, 2014). In support of this hypothesis, we observed severe defects in the development of craniofacial mesenchyme-derived tissues (Fig. 1, 2) that were consistent with molecular and cellular findings from Sox10-Cre Eed^Fl/Fl embryos (Fig. 3, 4). Expression of key regulators of craniofacial osteogenesis such as Runx2, Irx3, Irx5, Mecom, Trps1, Ptch1, Pax3, and Pax7 were reduced in Sox10-Cre Eed^Fl/Fl heads and primary craniofacial cell cultures compared to controls, and single-cell RNA sequencing showed reduced craniofacial osteoblast differentiation and reduced proliferation of mesenchymal cells in Sox10-Cre Eed^Fl/Fl heads compared to controls. We also identified subtle cardiac phenotypes and a small increase in apoptosis in Sox10-Cre Eed^Fl/Flembryos. These data suggest that PRC2 may contribute to the development of diverse neural crest-derived tissues but is particularly important for craniofacial mesenchymal cell differentiation and proliferation.

By using Sox10-Cre, which marks the migratory neural crest (Matsuoka et al., 2004), we bypass potential roles of Eed in neural crest cell induction to examine its function in neural crest cell development. The requirement of PRC2 for craniofacial development has been studied in the context of Ezh2 loss using Wnt1-Cre (Ferguson et al., 2018; Schwarz et al., 2014), but to our knowledge, our study is the first report directly linking Eed to differentiation of neural crest-derived mesenchymal cells. To date, fifteen human missense mutations in EED have been reported as pathogenic for Cohen-Gibson syndrome (Cohen and Gibson, 2016; Cohen et al., 2015; Goel et al., 2024). Like Weaver syndrome, which arises from missense mutations in EZH2 (Gibson et al., 2012; Tatton-Brown et al., 2013), and Imagawa-Matsumoto syndrome, which arises from missense mutations in SUZ12 (Imagawa et al., 2018), Cohen-Gibson syndrome is a rare congenital syndrome associated with craniofacial malformations, advanced bone age, intellectual disability, and developmental delay (Spellicy et al., 2019). EED mutations in individuals with Cohen-Gibson syndrome cluster in WD40 domains 3 through 5 and are predicted to (1) abolish interaction with EZH2, (2) prevent histone methyltransferase activity, and (3) inhibit H3K27 trimethylation peptide binding. In support of these hypotheses, functional investigations have shown that single nucleotide variants in the WD40 domain of EED abolish binding to EZH2 in vitro (Denisenko et al., 1998). Mouse models encoding pathogenic EZH2 missense variants, which are predicted to result in loss of function of the PRC2 complex, phenocopy Weaver syndrome and cause excess osteogenesis and skeletal overgrowth (Gao et al., 2023). In this study, we show that loss of Eed after neural crest induction causes the opposite phenotype, resulting in craniofacial hypoplasia due to impairments in osteogenesis, mesenchymal cell proliferation, and mesenchymal stem cell differentiation. The discrepancy of missense variants in EED causing gain of craniofacial osteogenic function in humans versus loss of Eed causing loss of craniofacial osteogenic function in mice warrants further study, including investigation using alternative Cre drivers to target different developmental stages of the neural crest. Despite these discrepancies, it is notable that many of the phenotypes we observed after loss of Eed in the neural crest of mice were similar to previously reported phenotypes after loss of Ezh2 in the neural crest cell in mice, with the exception of proliferation deficits after loss of Eed but not after loss of Ezh2 (Ferguson et al., 2018; Schwarz et al., 2014).

Methods

Mice

Mice were maintained in the University of California San Francisco (UCSF) pathogen-free animal facility in accordance with the guidelines established by the Institutional Animal Care and Use Committee (IACUC) and Laboratory Animal Resource Center (LARC) protocol AN191840. Mice were maintained in a 70°F, 50% humidity temperature-controlled barrier facilities under a 12-12h light cycle with access to food and water ad libitum. Sox10-Cre and Eed^Fl mice were obtained from the Jackson Laboratory (B6;CBA-Tg(Sox10-cre)1Wdr/J, 025807 and B6;129S1-Eed^tm1Sho/J, 022727, respectively). The presence of the floxed Eed allele was determined through standard PCR genotyping using the following primers: 5’ GGGACGTGCTGACATTTTCT 3’ (forward) and 5’ CTTGGGTGGTTTGGCTAAGA 3’ (reverse). To generate embryos at specific time points, Sox10-Cre^tg+ Eed^Fl/WT mice were bred overnight with Eed^Fl/Fl mice. Females were checked for copulation plugs in the morning and the presence of a vaginal plug was designated as E0.5.

Mouse fetal echocardiography

Ultrasound studies were acquired with a Fujifilm Vevo 2100 Imaging system, and instrument specifically designed for lab animal imaging studies. Pregnant females were anesthetized using an isoflurane/oxygen mixture with an isoflurane concentration of 3% during imaging. The pregnancy date (at least p17) was checked to observe the correct development of fetal hearts. Isoflurane was increased until 5% and the abdomen of pregnant females was open surgically to expose the gravid uterus. The number of fetuses was counted, and craniofacial architecture was evaluated for phenotypic confirmation. Once each fetus was defined as mutant or wild type, echocardiography was performed to evaluate left ventricle anatomy and physiology, including left ventricle size and contractility. Three different measurements were obtained in B and M modes, and functional parameters, including heart rate, fraction shortening, ejection fraction, and left ventricle mass were obtained.

Whole-mount embryo staining

Embryos were stained as previously described (Sandell et al., 2018). In brief, embryos were harvested, and heads were fixed overnight in 4% PFA. Embryos were dehydrated through a methanol series up to 100% before being treated with Dent’s bleach (4:1:1, MeOH, DMSO, 30% H2O2) for 2hr at 25°C. Embryos were serially rehydrated in methanol up to 25%, washed with PBST, and stained with DAPI solution (1:40,000, Thermo Scientific, cat# 62248) overnight at 4°C. Samples were mounted in low melt agarose for imaging.

Whole-mount skeletal staining

Embryos were stained as previously described (Rigueur and Lyons, 2013). In brief, embryos were dissected in cold PBS and scalded in hot water to facilitate the removal of eyes, skin, and internal organs. Embryos were fixed in 95% ethanol overnight then subsequently incubated in acetone overnight. Embryos were stained with Alcian Blue solution (Newcomer Supply, cat# 1300B) for 8hr, washed in 70% ethanol, and destained in 95% ethanol. Embryos were cleared in 95% ethanol and 1% KOH, and stained in Alizarin red S solution (SCBT, cat#130-22-3) for 4hr. Embryos were stored in glycerol for imaging.

Micro-computed tomography (microCT)

microCT studies were acquired with a Perkin Elmeŕs Quantum GX2 microCT Imaging system, an instrument specifically designed for lab animal imaging studies. The samples were imaged in Eppendorf tubes containing preservative media, which were placed on the scanning bed and fixed with tape to minimize movement. Acquisition parameters were set as follows: FOV 10mm, acquisition time 14 minutes, current voltage 70KV, amperage 114uA. Each study was composed by a 512×512×512 voxels matrix with a spatial resolution of 0.018mm³. Study reconstruction was based on Feldkamp’s method using instrument dedicated software. For image analysis, tridimensional renders were obtained using 3D SLICER software. The threshold range applied for bone segmentation was 300-1400 Hounsfield Units. For the segmentation and visualization of individual frontal bones, cranial base, and mandible, Avizo Lite software (v9.1.1) was used. The threshold range applied was 570-1570 Hounsfield Units.

Cryosectioning and H&E staining

Embryos were fixed in 4% PFA overnight at 4°C and subjected to a sucrose gradient from 15% to 30% before embedding in OCT (Tissue-Tek, cat#4583) and storage at -80°C. Embryos were sectioned at 25µm on a Leica CM1850 cryostat. Slides were immediately subjected to H&E staining as previously described (Cardiff et al., 2014), or stored at -80°C for immunofluorescence.

Immunofluorescence

Immunofluorescence of cryosections was performed on glass slides. Immunofluorescence for primary cells was performed on glass cover slips. Primary cells were fixed in 4% PFA for 10 minutes at room temperature. Antibody incubations were performed in blocking solution (0.1% Triton X-100, 1% BSA, 10% donkey serum) for 2hr at room temperature or overnight at 4°C. Slides or cover slips were mounted in ProLong Diamond Antifade Mountant (Thermo Scientific, cat# P36965). Primary antibodies used in this study were anti-Sox9 (1:500, Proteintech, cat# 67439-1-Ig), anti-Sox9 (1:500, Abcam, cat# ab185230), anti-Runx2 (1:500, Cell Signaling, cat# D1L7F), anti-Ki67 (1:2000, Abcam, cat# ab15580), anti-Vimentin (1:1000, Abcam, cat# ab8069), anti-BrdU (1:1000, Abcam, cat# ab6326), anti-Sox10 (1:500, Cell Signaling, cat# D5V9L), anti-ALPL (1:1000, Invitrogen, cat# PA5-47419), anti-Eed (1:1000, Cell Signaling, cat# E4L6E), and anti-Cleaved Caspase-3 (Asp175) (1:500, Cell Signaling, cat#9661). Secondary antibodies used in this study were Donkey anti-Rabbit IgG (H+L) Alexa Fluor Plus 555 (1:1000, Thermo Scientific, cat# A32794), Donkey anti-Rabbit IgG (H+L) Alexa Fluor 488 (1:1000, Thermo Scientific, cat# A21206), and Donkey anti-Rabbit IgG (H+L) Alexa Fluor Plus 568 (1:1000, Thermo Scientific, cat# A10042). DAPI solution (1:5000, Thermo Scientific, cat# 62248) was added to secondary antibody solutions. Slides or cover slips were mounted in ProLong Diamond Antifade Mountant (Thermo Scientific, cat# P36965) and cured overnight before imaging.

BrdU staining

Timed mating dams were subjected to intraperitoneal injection of 100mg/kg sterile BrdU in PBS (Abcam, cat# ab142567). After 4hr, mice were euthanized, and embryos were harvested in cold PBS. Embryos were fixed in 4% PFA overnight at 4°C and cryosectioned. Sections were incubated with 1M HCl for 2hr then 0.1M sodium borate buffer for 15min before being subjected to standard immunofluorescence.

Microscopy and image analysis

Whole-mount skeletal and H&E stains were imaged using a Zeiss Stemi 305 Stereo Zoom microscope running Zeiss Blue v2.0. Whole-mount embryo DAPI stains were imaged using an upright Zeiss Axio Imager Z2 running AxioVision v4.0. Immunofluorescence images were obtained using a Zeiss LSM800 confocal laser scanning microscope running Zen Blue v2.0. Quantification of embryo measurements and immunofluorescence staining intensities was performed in ImageJ using standard thresholding and measurement of signal integrated density. Signal to nuclei normalization was performed by dividing signal intensity by DAPI signal per image. Quantification was performed using n>6 images per embryo. Proliferation index was calculated by dividing the number of Ki67+ or BrdU+ nuclei by the total number of DAPI stained nuclei per image.

Primary craniofacial cell culture

Timed mating embryos were dissected on ice cold PBS and heads were removed. Brains and eyes were removed from each head, and the remaining craniofacial structures were minced with a sterile blade on a glass surface. Tissue was enzymatically dissociated in 3mg/ml Collagenase Type 7 (Worthington, cat# CLS-6) in HBSS for 2-3hr at 37°C with frequent trituration using an Eppendorf ThermoMixer. Dissociated cells were filtered through a 70uM MACS smart strainer (Miltenyi Biotec, cat#130-110-916) and either plated on cover slips or cell culture dishes. Primary craniofacial cells were grown in DMEM with 10% FBS. For RNA extraction and immunofluorescence, cells were harvested one day following initial dissociation.

Single cell RNA-seq

Matched littermate timed embryos were dissected on ice cold PBS. Embryo heads were removed and minced using sterile razor blades on a glass surface. Tissue was enzymatically dissociated in 3mg/ml Collagenase Type 7 (Worthington, cat# CLS-6) in HBSS for 2-3hr at 37°C with frequent trituration using an Eppendorf ThermoMixer. The quality of the dissociation was frequently monitored by looking at the cell suspension on an automated cell counter. Cells were spun down at 350rcf, resuspended in MACS BSA stock solution (Miltenyi Biotec, cat# 130-091-376), and serially filtered through 70µm and 40µm MACS smart strainers (Miltenyi Biotec, cat# 130-110-916). Dissociation quality checks, cell viability, and counting was performed using an Invitrogen Countess 3 automated cell counter. 10,000 cells were loaded per single-cell RNA sequencing sample. Single-cell RNA sequencing libraries were generated using the Chromium Single Cell 3’ Library & Gel Bead Kit v3.1 on a 10x Genomics Chromium controller using the manufacturer recommended default protocol and settings.

Single-cell RNA sequencing analysis

Library demultiplexing, read alignment to the mouse reference genome mm10, and unique molecular identifier (UMI) quantification was performed in Cell Ranger v7.2.0. Cells with greater than 200 unique genes were retained for analysis. Data were normalized and variance stabilized by SCTransform in Seurat v5.0 (Hao et al., 2024). UMAP and cluster analysis were performed using the Seurat function RunUMAP with parameters of mindist=0.7, res=0.4, dims=1:30. Cluster markers were identified using Seurat function FindAllMarkers with parameters min.pct = 0.25, thresh.use = 0.25. Differential gene expression analysis was performed using DElegate v1.1.0 (Hafemeister and Halbritter, 2023) using the DEseq2 method, with biological replicate animals (n=3 per genotype) as the replicate_column parameter. Featureplots were generated using SCPubr v1.1.2 (Blanco-Carmona, 2022) and scale bars represent log2 UMI’s corrected by SCTransform.

Statistics

All experiments were performed with independent biological replicates and repeated, and statistics were derived from biological replicates. Biological replicates are indicated in each figure panel or figure legend. No statistical methods were used to predetermine sample sizes, but sample sizes in this study are similar or larger to those reported in previous publications. Data distribution was assumed to be normal, but this was not formally tested. Investigators were blinded to conditions during data collection and analysis. Bioinformatic analyses were performed blind to clinical features, outcomes, and molecular characteristics. The samples used in this study were nonrandomized with no intervention, and all samples were interrogated equally. Thus, controlling for covariates among samples was not relevant. No data points were excluded from the analyses.

Additional information

Competing interests statement

The authors declare no competing interests.

Author contributions

All authors made substantial contributions to the conception or design of the study; the acquisition, analysis, or interpretation of data; or drafting or revising the manuscript. All authors approved the manuscript. All authors agree to be personally accountable for individual contributions and to ensure that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved and the resolution documented in the literature. T-C.C. and D.R.R. conceived and designed the study. Experiments were performed by T.C-C., C.T., J.A.C.S., and Y-G.J. Data analysis was performed by T.C-C., S.J.L., and D.R.R. S.J.L. performed bioinformatic analyses. The study was supervised by H.N.V., J.O.B., and D.R.R. The manuscript was prepared by T.C-C. and D.R.R. with input from all authors.

Funding

This study was supported by funding from NIH grant T32 CA151022 to T.C-C., NIH grant R35 DE031926 to J.O.B, and NIH grant R21 HD106238 and DOD grant NFRP NF200021 to D.R.R. Sequencing was performed at the UCSF CAT, supported by UCSF PBBR, RRP IMIA, and NIH 1S10OD028511-01 grants.

Data availability

Single-cell RNA sequencing data that were reported in this manuscript have been deposited in the NCBI Sequence Read Archive under PRJNA1077750 (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA1077750).

Code availability

No custom software, tools, or packages were used. The open-source software, tools, and packages used for data analysis in this study are referenced in the methods where applicable and include Cell Ranger v7.2.0, Seurat v5.0, DElegate v1.1.0, and SCPubr v1.1.2.

Supplementary figures

Supplementary Fig. 1.

Supplementary Fig. 2.

Supplementary Fig. 3.

Supplementary Fig. 4.

Supplementary Fig 5.

Supplementary Fig. 6.

Supplementary Fig. 7.