Epigenetic Regulation of Craniofacial Development and Disease

Abstract

The formation of the craniofacial complex relies on proper neural crest development. The gene regulatory networks (GRNs) and signaling pathways orchestrating this process have been extensively studied. These GRNs and signaling cascades are tightly regulated as alterations to any stage of neural crest development can lead to common congenital birth defects, including multiple syndromes affecting facial morphology as well as non-syndromic facial defects, such as cleft lip with or without cleft palate. Epigenetic factors add a hierarchy to the regulation of transcriptional networks and influence the spatiotemporal activation or repression of specific gene regulatory cascades, however less is known about their exact mechanisms in controlling precise gene regulation. In this review, we discuss the role of epigenetic factors during neural crest development, specifically during craniofacial development and how compromised activities of these regulators contributes to congenital defects that affect the craniofacial complex.

Introduction

The craniofacial skeleton houses and protects vital sensory organs providing the ability to see, hear, smell, eat and breathe while also providing ways to convey social interactions and communication between others through facial expressions. The formation of the craniofacial complex depends mostly on the proper development of a highly migratory, multipotent stem/progenitor population of cells termed neural crest cells (NCCs) (Bronner & LaBonne, 2012; Bronner & LeDouarin, 2012; Green et al., 2015; Le Douarin & Dupin, 2018; Simoes-Costa & Bronner, 2015). During gastrulation, NCCs are induced at the neural plate border, which separates the neural and non-neural ectoderm, by signaling from BMP and Wnt pathways. Specification involves a complex gene regulatory network (GRN) of transcription factors including the upregulation of the neural crest specifier genes Pax3, Pax7, FoxD3, Sox10 and Snail2. As neurulation occurs, morphogenetic movements resolve the NCCs to the dorsal neural tube. From there, they undergo an epithelial-to-mesenchymal transition (EMT) and delaminate from the neural epithelium and migrate in distinct streams throughout the embryo along the entire anterior-posterior axis. When NCCs reach their destination, they will begin differentiation toward many different derivatives throughout the body including melanocytes, cardiac muscle, neurons and glia, Schwann cells, chondrocytes and osteoblasts. The cranial NCCs subpopulation form the neurons and glia of the peripheral nervous system and the cells that populate the pharyngeal/branchial arches, which interact with endodermal and ectodermal components of the arches. Cranial NCCs require endothelin signaling to differentiate into cartilage and bone to form (Betancur et al., 2010; Sauka-Spengler & Bronner-Fraser, 2008; Sauka-Spengler & Bronner, 2010) the craniofacial structures in the skull, including the maxilla (upper jaw), mandible (lower jaw), frontonasal prominence, cranial base, external ear, middle ear and hyoid bone. All these tissues are innervated with NCC-derived sensory neurons from the trigeminal ganglia.

While the GRNs governing proper NCC development have been extensively studied and characterized, less is known about how the GRNs are precisely spatiotemporally regulated. Growing evidence supports the functional roles of epigenetic factors as important gatekeepers facilitating many aspects of neural crest development, including controlling the timely activation and/or repression of gene expression across developmental time. Importantly, alterations to the spatiotemporal activation or repression of NCC specific GRNs can result in congenital birth defects affecting neural crest derived tissues, as well as non-syndromic defects including cleft lip with or without cleft palate, heart defects, Hirschprung’s disease, and cancers including melanoma, neuroblastoma, and neurofibromatosis. Many congenital birth defects arise from abnormal neural crest development, however, a large majority of their etiologies remain unknown. In addition to genetic mutations, there are environmental factors that can have a significant influence on the genome and epigenome. Diet, for example can directly affect the epigenome, as evidenced by the Dutch famine studies (Heijmans et al., 2008; Tobi et al., 2009). Other environmental factors that can induce epigenomic changes include tobacco and alcohol use, psychological stress, specific minerals, certain medications (i.e. retinoic acid), cigarette smoke and infections (Marsit, 2015). The epigenetic regulators and the epigenome they govern during neural crest development are particularly sensitive due to the rapid transcriptional changes occurring during these stages of development. Teratogens can induce epigenetic changes that in turn affect neural crest development.

In this review, we discuss the role of epigenetic regulators during neural crest development, with specific focus on the formation of the craniofacial skeleton and how alterations to the functions of these factors lead to congenital diseases.

Mechanisms of Epigenetic Regulation

The term epigenetics is defined as mechanisms that lead to changes in gene expression by altering chromosome structure rather than changes in the DNA sequence (Berger, 2007). Negatively charged DNA is tightly wrapped around a core of histones consisting of two H2A histones, two H2B histones, two H3 histones and two H4 histones (Quina et al., 2006). This resulting protein octamer forms a nucleosome, the basic unit of chromatin (Allis & Jenuwein, 2016). How chromatin is arranged within chromosomes determines if the DNA is transcriptionally active. Chromatin packaged loosely indicates transcriptionally active euchromatin while densely bundled chromatin typically indicates a more transcriptionally inactive heterochromatin state (Bannister & Kouzarides, 2011).

Epigenetic modifiers include groups of “writers,” “erasers” and “readers” that change how chromatin is organized. These modifiers function through a variety of different mechanisms: (1) DNA modifications, (2) histone modifications; (3) through the formation of large protein complexes that can regulate higher-order chromatin via nucleosome positioning, chromatin dynamics and interactions; or (4) non-coding RNAs (Bannister & Kouzarides, 2011; Kouzarides, 2007). DNA modifications mostly consist of methylated CpG islands near promoter sequences. Histone modifications occur when different chemical marks are added to the tails extending off histones that influences what type of transcriptional machinery is recruited to that mark. Large protein complexes, for example, either Polycomb Repressive Complex or ATP-dependent chromatin remodeling complexes influence chromatin architecture through nucleosome rearrangements and DNA accessibility. While not specifically discussed further here, non-coding RNAs are functional RNA molecules that are transcribed but not translated into proteins associated with gene silencing. Non-coding RNAs (microRNAs and long non-coding RNAs) participate in DNA methylation, histone modifications, formation of heterochromatin and gene silencing. Across these different epigenetic mechanisms, depending on the specific type of regulator, modification or recruited transcriptional machinery, the outcome will lead to either gene activation or repression.

When epigenetic factors are mutated and their functional activity is compromised, abnormal activation or repression of gene expression can lead to cancers and disease states. Here, we focus on the function of epigenetic factors and their roles in facilitating proper neural crest development, formation of the craniofacial skeleton and neural crest specific diseases presenting with craniofacial anomalies associated with aberrant activity of these factors (Figure 1) and (Table 1). Below we highlight factors involved in DNA methylation, histone modification, and protein complexes that alter chromatin structure.

Figure 1. Epigenetic regulators spatiotemporally control gene regulatory networks during neural crest development and formation of the craniofacial skeleton.

Neural crest cells arise at the neural plate border during gastrulation (A). Through a dynamic process, these cells are specified, undergo epithelial-to-mesenchymal transitions, and migrate out to different areas along the anterior-posterior axis where they begin differentiating to different derivatives (A-B). The neural crest cells that migrate to the pharyngeal arches (PA) will differentiate toward chondrocytes, osteoblasts, to form the cartilage and bone of the facial structures as well as neurons and glia of the peripheral nervous system (B-D). These cell types and tissues integrate with one another to form the craniofacial complex and this process is highly conserved across vertebrates (D). Various gene regulatory networks are important along every step of neural crest and craniofacial development, from neural crest induction and specification to patterning of the pharyngeal arches to differentiation of different derivatives. Different epigenetic regulators have important roles in mediating the activation or repression of these different GRNs. These epigenetic regulators can directly influence the activation or repression of the transcription factors (TFs) in each GRN. For example, EZH2 represses the Hox gene family and activates osteogenic differentiation programs to facilitate bone formation (E). See Table 1 for references for each factor featured above.

Table 1. Epigenetic Factors involved in Craniofacial Development and Disease.

Bolded are those factors discussed more thoroughly in the text.

FACTOR	EPIGENETIC FUNCTION	FUNCTION DURING NCC DEVELOPMENT	DISEASE ASSOCIATION	REFERENCES
ARID1A	Member of SWI/SNF complex, repressor that binds chromatin	potential role in NCC formation, migration, differentiation	Coffin-Siris Syndrome	(Chandler & Magnuson, 2016; Gatchalian et al., 2018; Ho et al., 2009; Pagliaroli et al., 2021)
ARID1B	Member of SWI/SNF complex, repressor that binds chromatin	potential role in NCC formation, migration, differentiation	Coffin-Siris Syndrome	(Gatchalian et al., 2018; Ho et al., 2009; Pagliaroli et al., 2021)
ARID2	Member of SWI/SNF-B complex, repressor that binds chromatin	osteoblast differentiation	Coffin-Siris Syndrome	(Bramswig et al., 2017; Xia et al., 2023; Xu et al., 2012)
ASXL1	Member of Polycomb Repressive Complex	generation of NCC, neuroectoderm specification	Bohring-Opitz syndrome	(Matheus et al., 2019; Zhao et al., 2021)
ASXL2	Member of Polycomb Repressive Complex	generation of NCC	Shashi-Pena Syndrome	(Shashi et al., 2016; Shashi et al., 2017)
ASXL3	Member of Polycomb repressive deubiquitination	generation of NCC, early nervous system development	Bainbridge-Ropers Syndrome	(Dinwiddie et al., 2013; Koboldt et al., 2018; Kuechler et al., 2017; Lichtig et al., 2020)
ATRX	ATP-dependent chromatin remodeling	neural progenitor differentiation	Alpha thalassemia/mental retardation syndrome	(Berube et al., 2005; Bieluszewska et al., 2022; Dyer et al., 2017; Thienpont et al., 2007; Tillotson et al., 2023)
BAF155	Core structural subunit of BAF complex	NCC proliferation and differentiation	Nicolaides-Baraitser Syndrome	(Bi-Lin et al., 2021; Sousa et al., 2014)
BAF170	Core structural subunit of BAF complex	NCC proliferation and differentiation	Nicolaides-Baraitser Syndrome	(Bi-Lin et al., 2021; Sousa et al., 2014)
BAZ1A	ATP-dependent Chromatin Remodeling	NCC formation and maintenance, migration, differentiation	CHARGE syndrome	(Baggiolini et al., 2021; Zaghlool et al., 2016)
BAZ1B/WSTF	ATP-dependent Chromatin Remodeling	NCC formation and maintenance, migration, differentiation	Williams-Beuren Syndrome	(Barnett & Krebs, 2011; Barnett et al., 2012; Culver-Cochran & Chadwick, 2013; Lalli et al., 2016; Torres-Perez et al., 2023; Zanella et al., 2019)
BRD2	BET protein; binds acetylated histones, recruits transcriptional machinery	neural progenitor proliferation and differentiation; NCC migration	neural tube defects and craniofacial malformations	(Garcia-Gutierrez & Garcia-Dominguez, 2015; Garcia-Gutierrez et al., 2014; Gyuris et al., 2009; Shang et al., 2009; Tsume et al., 2012)
BRD4	BET protein; binds acetylated histones, recruits transcriptional machinery	NCC formation and maintenance, migration, differentiation	Cornelia de Lange Syndrome	(Garcia-Gutierrez & Garcia-Dominguez, 2021; Liang et al., 2021; Linares-Saldana et al., 2021)
BRD7	BET protein; binds acetylated histones, recruits transcriptional machinery	NCC formation by regulating NC transcription; controls osteogenic differentiation	craniofacial abnormalities	(Bajpai et al., 2010; Sinha et al., 2020)
BRG1	One of 2 ATPase cores belonging to the BAF complex	NCC induction/specification and neurogenesis	Coffin-Siris Syndrome	(Eroglu et al., 2006; Li et al., 2013; Lou et al., 2015; Tzeng et al., 2014)
CBP	CREB Binding protein nuclear transcriptional co-activator	Expressed in NCC/developing craniofacial tissue	Rubinstein-Taybi Syndrome	(Petrij et al., 1995; Tanaka et al., 1997; Warner et al., 2002)
CHD1	Chromodomain helicase, DNA-Binding protein 1; ability to remodel chromatin structure and influence histone acetylation	NCC formation and differentiation	Pilarowski-Bjornsson Syndrome	(Pilarowski et al., 2018; Wyatt et al., 2021; Zepeda-Mendoza et al., 2019)
CHD2	Chromodomain helicase, DNA-Binding protein 2; ability to remodel chromatin structure and influence histone acetylation	neurogenesis	CHARGE syndrome	(Galizia et al., 2015; Kulkarni et al., 2008; Semba et al., 2017; Suls et al., 2013)
CHD7	Chromodomain helicase, DNA-Binding protein 7; ability to remodel chromatin structure and influence histone acetylation	maintain NCC multipotency, NCC specification	CHARGE syndrome	(Bajpai et al., 2010; Balow et al., 2013; Bosman et al., 2005; Chai et al., 2018; Fujita et al., 2014; Lalani et al., 2006; Sperry et al., 2014; Y. Sun et al., 2022)
CTCF	Zn finger binding protein; regulates 3D organization of the genome; chromatin architecture	NCC proliferation and differentiation to facial derivatives	Neurodevelopmental Disorder with facial dysmorphism	(Adams et al., 2008; Bastaki et al., 2017; Gregor et al., 2013; Hori et al., 2017; Konrad et al., 2019; Min et al., 2019)
DNMT3A	DNA methyltransferase 3A; DNA methylation of CpG Islands	NCC formation; NT-to-NCC fate transition in chick during NCC specification	Tatton-Brown-Rahman Syndrome	(Hu et al., 2012; Okano, Bell, et al., 1999; Smith et al., 2021; Yokoi et al., 2020)
DNMT3B	DNA methyltransferase 3B; DNA methylation of CpG Islands	NCC formation and differentiation of PNS neurons in chick but maybe dispensable in mouse NCC development	Immunodeficiency, centromere instability facial anomalies syndrome	(Ehrlich et al., 2001; Hu, Strobl-Mazzulla, Simoes-Costa, et al., 2014; Jacques-Fricke et al., 2012; Jiang et al., 2005; Jin et al., 2008; Jin et al., 2009; Martins-Taylor et al., 2012; Nowialis et al., 2019; Okano, Bell, et al., 1999; Okano, Takebayashi, et al., 1999; Ueda et al., 2006)
DPF2	BAF complex subunit member double plant homeodomain finger 2	potential role in NCC formation, migration, differentiation	Coffin-Siris Syndrome	(MacDonald et al., 2022; Vasileiou et al., 2018)
EHMT1/GLP/KMT1D	Euchromatin histone methyltransferase 1; dimethylation of H3K9me2	neurogenesis, NCC differentiation	Kleefstra Syndrome	(Balemans et al., 2014; Frega et al., 2019; Kleefstra et al., 2009; Martens et al., 2016)
EHMT2/G9A/KMT1C	Euchromatin histone methyltransferase 2; dimethylation of H3K9me2	neurogenesis; NCC differentiation	Kleefstra Syndrome	(Balemans et al., 2014; Higashihori et al., 2017; Ideno et al., 2020; N. Liu et al., 2015; Rai et al., 2010)
EP300	EP1A binding protein; lysine acetyltransferase (histones and nonhistones)	neurogenesis; NCC differentiation to craniofacial elements	Rubinstein-Taybi Syndrome	(Babu et al., 2018; Bartsch et al., 2010; Bhattacherjee et al., 2009; Hamilton et al., 2016; Saettini et al., 2022; Warner et al., 2002, 2006)
EZH2	enhancer of zeste homolog 2 (catalytic component of the Polycomb repressor complex; mono-di-tri methylation of H3K27)	NCC differentiation to craniofacial skeletal structures	Weaver syndrome	(Cohen et al., 2016; Dudakovic et al., 2015; H. Kim et al., 2018; Lui et al., 2018; Schwarz et al., 2014; Tatton-Brown et al., 2013)
HDAC1	Histone deacetylase	NCC proliferation and NCC differentiation of skeletal structures; PNS	craniofacial abnormalities with Copy number variations at 1p35	(Ignatius et al., 2013; Milstone et al., 2017; Pillai et al., 2004; Rao & LaBonne, 2018)
HDAC2	Histone deacetylase	NCC proliferation and NCC differentiation of skeletal structures; PNS	craniofacial abnormalities with Copy number variations at 6q21-6q22.1	(Hudson et al., 2014; Milstone et al., 2017)
HDAC3	Histone deacetylase	NCC proliferation and formation	chromatin remodeling disorders associated with craniofacial malformation	(Singh et al., 2013; Singh et al., 2011)
HDAC4	Histone deacetylase	NCC proliferation and differentiation to osteoblasts	Brachydactyly-Mental Retardation Syndrome, non-syndromic oral clefting	(DeLaurier et al., 2012; Ha et al., 2022; Park et al., 2006; Villavicencio-Lorini et al., 2013; Williams, Aldred, et al., 2010)
HDAC8	Histone deacetylase	NCC patterning and differentiation of craniofacial elements	Cornelia de Lange Syndrome; Wilson-Turner Syndrome	(Deardorff et al., 2012; Deardorff et al., 2016; Gao et al., 2018; Haberland et al., 2009; Harakalova et al., 2012; Kaiser et al., 2014; Lucia-Campos et al., 2022; Mio et al., 2021)
KAT2A/GCN5	Lysine histone acetyltransferase	NCC formation and differentiation	chromatin remodeling disorders associated with craniofacial malformation	(Dong et al., 2023; Pezoa et al., 2020; Sen et al., 2018)
KAT2B/PCAF	Lysine histone acetyltransferase	NCC formation and differentiation	chromatin remodeling disorders associated with craniofacial malformation	(Sen et al., 2018)
KAT6A	Lysine histone acetyltransferase	NCC formation, patterning and differentiation of skeletal/craniofacial structures by regulating gene expression patterns	KAT6A syndrome; Arboleda-Tham Syndrome; Pancraniosynostosis	(Arboleda et al., 2015; Bae et al., 2021; Crump et al., 2006; Marji et al., 2021; Miller et al., 2004; Satoh et al., 2017; Tham et al., 2015; Vanyai et al., 2019; Voss et al., 2012; Wiesel-Motiuk & Assaraf, 2020)
KAT6B	Lysine histone acetyltransferase	NCC formation, patterning and differentiation of skeletal/craniofacial structures	Noonan syndrome, Ohdo Syndrome, Genitopatellar Syndrome	(Campeau et al., 2012; Clayton-Smith et al., 2011; Kraft et al., 2011; Sheikh et al., 2012; Simpson et al., 2012; Szakszon et al., 2013; Wiesel-Motiuk & Assaraf, 2020; Zhang et al., 2020)
KDM1A	Lysine histone demethylase	NCC function unknown; adipogenesis, neurogenesis, distinct facial features	Cleft palate, facial dysmorphology; Kabuki syndrome-like	(Chong et al., 2016; Tunovic et al., 2014)
KDM2A	Lysine histone demethylase	with BCOR---inhibits odontogenic differentiation of human stem cells from apical papilla (SCAPS)	no known disease association	(Dong et al., 2013; Fan et al., 2009)
KDM2B	Lysine histone demethylase	unknown; may lead to NT closure defects; with BCOR and recruited to the promoter of AP2alpha for osteogenesis OFCD patients roots of canine teeth	no known disease association	(Fan et al., 2009; Fukuda et al., 2011)
KDM4A	Lysine histone demethylase	NCC specification	no known disease association	(Strobl-Mazzulla et al., 2010)
KDM5B	Lysine histone demethylase	neurogenesis; development of craniofacial structures	Intellectual developmental disorder with facial dysmorphia	(Albert et al., 2013; Faundes et al., 2018)
KDM5C	Lysine histone demethylase	NCC migration and differentiation; neurogenesis	Claes-Jensen syndrome; X-linked intellectual disability with facial dysmorphia	(Hatch & Secombe, 2022; Iwase et al., 2016; Y. Kim et al., 2018; Ounap et al., 2012; Vallianatos et al., 2020)
KDM6A/UTX	Lysine histone demethylase; can interact with TFs, chromatin remodeling enzymes; and chromatin remodeling complex	NCC development post migration and differentiation of skeletal/craniofacial elements	Kabuki Syndrome	(Guo et al., 2022; Khodaeian et al., 2021; Lederer et al., 2012; Lindgren et al., 2013; Miyake, Koshimizu, et al., 2013; Miyake, Mizuno, et al., 2013; Ng et al., 2010; Porntaveetus et al., 2018; Shpargel et al., 2017; Van Laarhoven et al., 2015)
KDM6B	Lysine histone demethylase	NCC differentiation to craniofacial elements; osteogenic commitment; neurogenesis	Neurodevelopmental disorder; facial/skeletal dysmorphology	(Burgold et al., 2008; Estaras et al., 2012; Guo et al., 2022; Xu et al., 2013)
KMT2A	Lysine methyltransferase	NCC function undefined	Wiedemann-Steiner Syndrome	(Jones et al., 2012; Luo et al., 2021; Strom et al., 2014; Sun et al., 2017; Vallianatos et al., 2020)
KMT2D/MLL2	Lysine methyltransferase; can interact with TFs, chromatin remodeling enzymes; and chromatin remodeling complex	NCC formation, cell dispersion prior to migration; later development post migration and differentiation of skeletal/craniofacial elements	Kabuki Syndrome	(Bogershausen et al., 2016; Khodaeian et al., 2021; Lederer et al., 2012; Miyake, Koshimizu, et al., 2013; Ng et al., 2010; Porntaveetus et al., 2018; Schwenty-Lara et al., 2020; Shpargel et al., 2020; Shpargel et al., 2017; Van Laarhoven et al., 2015)
KMT3A/SETD2	Lysine histone methyltransferase	Early NCC specification and migration	Luscan-Lumish Syndrome; Sotos Syndrome	(Lumish et al., 2015; Luscan et al., 2014; Roffers-Agarwal et al., 2021; Sotos et al., 1964; van Rij et al., 2018)
KMT5B	Lysine histone methyltransferase	NCC function undefined; formation of the craniofacial skeletal elements; neurogenesis; differentiation	Intellectual development disorder with facial dysmorphism; overgrowth syndrome; craniosynostosis	(Angerilli et al., 2023; Eliyahu et al., 2022; Faundes et al., 2018; Timberlake et al., 2023)
MBD5	Methyl-CpG-binding domain protein	NCC function undefined; formation of the craniofacial skeletal elements; neurogenesis; differentiation	Intellectual development disorder with facial dysmorphism 2q23.1 microdeletion, MAND syndrome	(Bonnet et al., 2013; Jaillard et al., 2009; Mullegama et al., 2014; Talkowski et al., 2011; van Bon et al., 2010; Williams, Mullegama, et al., 2010)
NSD1/KMT3B	Lysine histone methyltransferase	NCC migration	Sotos Syndrome; some cases of Weaver syndrome	(Douglas et al., 2003; Hirai et al., 2011; Jacques-Fricke & Gammill, 2014; Makeyev & Bayarsaihan, 2013; Rayasam et al., 2003; Rio et al., 2003; Turkmen et al., 2003; Visser et al., 2012)
NSD2/KMT3G/WHSC1	Lysine histone methyltransferase	NCC migration; patterning of craniofacial structures	Wolf-Hirschhorn syndrome; Rauch-Steindl Syndrome	(Boczek et al., 2018; Jiang et al., 2019; S. Liu et al., 2015; Mills et al., 2019, 2020; Rauch et al., 2001; Zanoni et al., 2021)
NSD3	Lysine histone methyltransferase	NCC formation, specification and migration	chromatin remodeling disorders associated with craniofacial malformation	(Jacques-Fricke & Gammill, 2014; Jacques-Fricke et al., 2021)
PHC1/RAE28	Member of the Polycomb Group of genes	NCC formation and craniofacial patterning	CATCH-22 Syndrome; DiGeorge Syndrome-like	(Isono et al., 2005; Shirai et al., 2002; Takihara et al., 1997; Wilson et al., 1993)
PHC2	Member of the Polycomb Group of genes	NCC formation and craniofacial patterning	Non-syndromic Cleft lip/palate	(Isono et al., 2005; Marc et al., 2023; Shirai et al., 2002; Takihara et al., 1997)
PHF6	Plant homeodomain finger protein; chromatin reader	NCC function undefined; expressed in pharyngeal arches; neurogenesis	Coffin-Siris Syndrome; Borjeson-Forssman-Lehmann Syndrome	(Brunskill et al., 2014; Carter et al., 2009; Garcia-Melendo et al., 2021; Kosho, Okamoto, et al., 2014; Lower et al., 2002; Todd et al., 2015; Zweier et al., 2014)
PHF8	Plant homeodomain finger protein; chromatin reader; demethylase	NCC formation and differentiation toward craniofacial skeletal derivatives; neurogenesis	Siderious X-linked intellectual disability syndrome with craniofacial dysmorphology	(Han et al., 2015; Qi et al., 2010; Siderius et al., 1999; Sobering et al., 2023; Sobering et al., 2022)
PHF12	Plant homeodomain finger protein; histone deactylase complex member	NCC EMT and migration	no known disease association	(Adams et al., 2008; Strobl-Mazzulla & Bronner, 2012; Yochum & Ayer, 2001)
POGZ	Pogo transposable element with Zinc Finger Domain	NCC function undefined; neurogenesis	White-Sutton Syndrome	(Assia Batzir et al., 2020; Assia Batzir et al., 1993; Markenscoff-Papadimitriou et al., 2021; Stessman et al., 2016; X. Sun et al., 2022; White et al., 2016; Ye et al., 2015)
PRDM3/MECOM	Histone methyltransferase	NCC craniofacial development; differentiation of NCC derivatives; chondrocyte differentiation	Non-syndromic Cleft lip/palate	(Ding et al., 2013; Shull et al., 2022; Shull et al., 2020)
PRDM16	Histone methyltransferase	NCC craniofacial development; differentiation of NCC derivatives; chondrocyte differentiation	Non-syndromic Cleft lip/palate; Pierre-Robin Syndrome-like; neurogenesis	(Bjork et al., 2010; Ding et al., 2013; Liu et al., 2012; Motch Perrine et al., 2020; Shaffer et al., 2016; Shull et al., 2022; Shull et al., 2020; Strassman et al., 2017; Warner et al., 2013; Wilderman et al., 2018)
RING1B/RNF2	Polycomb repressive complex 1	craniofacial chondrocyte differentiation		(van der Velden et al., 2013)
SATB2	Special AT-rich sequence binding protein 2; binds nuclear matrix attachment regions; chromatin reader	NCC craniofacial patterning and differentiation of osteoblasts	SATB2-associated syndrome	(Britanova et al., 2005; Britanova et al., 2006; Dobreva et al., 2006; Dobreva et al., 2003; FitzPatrick et al., 2003; Huang et al., 2022; Kikuiri et al., 2018; Rainger et al., 2014; Scott et al., 2019; Urquhart et al., 2009; Zarate et al., 2019; Zarate & Fish, 2017)
SETBP1	SET Binding Protein; chromatin reader; recruits complexes	NCC function undefined; neurogenesis; formation of craniofacial elements	Schinzel-Giedion Syndrome	(Acuna-Hidalgo et al., 2017; Cardo et al., 2023; Hoischen et al., 2010; Leone et al., 2020; Liu et al., 2018; Piazza et al., 2018)
SETD1B	SET Domain Containing 1B; Histone Lysine methyltransferase	NCC function undefined; neurogenesis; formation of craniofacial elements	Intellectual development disorder with facial dysmorphia	(Hiraide et al., 2019; Hiraide et al., 2018; Labonne et al., 2016; Roston et al., 2021; Weerts et al., 2021)
SETD5	SET Domain Containing 5; Histone Lysine methyltransferase	NCC formation; formation of the craniofacial skeletal elements	Intellectual development disorder with facial dysmorphia; KBG Syndrome	(Crippa et al., 2020; Deliu et al., 2018; Fernandes et al., 2018; Gabellini et al., 2022; Garcia-Gutierrez & Garcia-Dominguez, 2021; Grozeva et al., 2014; Kellogg et al., 2013; Kuechler et al., 2015; Nakagawa et al., 2022; Nakagawa et al., 2020; Parenti et al., 2017; Powis et al., 2018; Rawlins et al., 2017; Sessa et al., 2019; Szczaluba et al., 2016; Zaghi et al., 2023)
SIN3A	Swi-independent protein 3A; Scaffolding regulator complexes with HDAC	NCC EMT; neurogenesis; formation of craniofacial elements	Witteveen-Kolk Syndrome	(Balasubramanian et al., 2021; Grzenda et al., 2009; Narumi-Kishimoto et al., 2019; Strobl-Mazzulla & Bronner, 2012; Witteveen et al., 2016; Yochum & Ayer, 2001)
SMARCA2	Member of SWI/SNF complex, repressor that binds chromatin; chromatin reader	NCC function undefined; neurogenesis	Nicolaides-Baraitser Syndrome; Blepharophimosis Intellectual Disability Syndrome	(Cappuccio et al., 2020; Chater-Diehl et al., 2019; Sanchez & Rojas, 2017; Sousa et al., 2014; Tang et al., 2017; Van Houdt et al., 2012)
SMARCB1	Member of SWI/SNF complex, repressor that binds chromatin; chromatin reader	NCC formation; neurogenesis; formation of the craniofacial skeletal elements	Coffin-Siris Syndrome 3	(Diets et al., 2019; Kosho, Miyake, et al., 2014; Kosho, Okamoto, et al., 2014; Santen et al., 2013; Tsurusaki et al., 2012; Tsurusaki et al., 2014; Vitte et al., 2017; Wieczorek et al., 2013)
SMARCE1	Member of SWI/SNF complex, repressor that binds chromatin; chromatin reader	NCC function undefined; potentially involved in NCC differentiation via TWIST1	Coffin-Siris Syndrome; Oculoauriculofrontonasal syndrome	(Kosho, Okamoto, et al., 2014; Miyake et al., 2014; Tsurusaki et al., 2012; Yano et al., 2018)
WSTF	Williams Syndrome Transcription Factor; subunit of ATP-dependent chromatin remodelers WINAC, WICH, B-WICH	NCC formation, NCC migration and maintenance	Williams Syndrome	(Barnett & Krebs, 2011, 2015; Barnett et al., 2012; Culver-Cochran & Chadwick, 2013; Cus et al., 2006; Lu et al., 1998)

Negative regulation by DNA Methylation

DNA methylation occurs when cytosine nucleotides in cytosine-guanine sequences (CpGs) are methylated (Cheng & Blumenthal, 2008, 2022). These areas are surrounded by other CpGs, creating CpG islands and are often located within promoter regions of genes. Methylated CpGs recruit transcriptional repressors, preventing transcriptional activation by inhibiting DNA-transcription factor interactions (Moore et al., 2013). During development, DNA methylation of specific gene regulatory networks or pathways is critical during cell fate determination, specification and lineage commitments (reviewed in (Altun et al., 2010)).

The process of DNA methylation occurs through the activity of DNA methyltransferases including DNMT1, DNMT3A and DNMT3B. DNMT1 serves as a maintenance methyltransferase by maintaining the patterns of methylation on newly synthesized DNA during DNA replication and repair. DNMT3A and DNMT3B establish a de novo methylation patterns by generating newly methylated CpGs, at promoter regions of genes. Several animal studies have shown DNMT3A and DNMT3B are responsible for suppressing tissue specific gene regulatory networks to facilitate cell fate decisions. In mice, Dnmt3a and Dnmt3b are highly expressed in early undifferentiated embryonic stem cells and their expression gradually decrease as cells become more differentiated (Okano, Bell, et al., 1999; Okano et al., 1998). DNMT3A functions in chick development to repress neural fates (by suppressing neuronal genes Sox2 and Sox3) to promote neural crest specification (Hu et al., 2012). Zebrafish DNMT3B, in cooperation with G9a, regulates neurogenesis and formation of the craniofacial skeletal elements, brain and retina (Rai et al., 2010). Further, studies in human embryonic stem cells have shown that knockdown of DNMT3B accelerates neural and neural crest differentiation by increasing expression of neural crest specifier genes (Pax3, Pax7, FoxD3, Sox10 and Snail2) (Martins-Taylor et al., 2012).

While mutations in DNMT3A have been associated with acute myeloid leukemia, mutations in DNMT3A have now been identified in patients presenting with congenital syndromes including Tatton-Brown-Rahman syndrome (TBRS) (also termed DNMT3A overgrowth syndrome (DOS)) (Tatton-Brown et al., 2013). This autosomal dominant disease manifests with overgrowth, macrocephaly, characteristic facial features or facial dysmorphism and intellectual disability and autism (Tatton-Brown et al., 2013; Yokoi et al., 2020). A recent study characterized the functional and epigenetic phenotypes of patients presenting with DNMT3A-associated TBRS/DOS (Smith et al., 2021). Through whole genome bisulfite sequencing on patient derived peripheral blood cells, they found a focal hypomethylation phenotype. Complementary to their human data analysis, the authors generated a mouse model that phenocopied the disease aspects of the human TBRS/DOS syndrome including hypomethylation (Smith et al., 2021). Patients with mutations in DNMT3B present with immunodeficiency-centromeric instability-facial anomalies (ICF), characterized by a widened nasal bridge and hypotelorism, and neurological dysfunction (Ehrlich et al., 2001; Jiang et al., 2005; Jin et al., 2008; Tuck-Muller et al., 2000). These DNMT3B mutations are conserved, as mouse models carrying homologous mutations to those found in patients with ICF develop similar phenotypes including distinct craniofacial abnormalities (Ueda et al., 2006).

Though numerous studies support a strong role for DNMT3B in neural crest and craniofacial development, Jacques-Fricke et al demonstrated that conditional loss of DNMT3B in the murine neural crest with either the Wnt1-Cre or Sox10-Cre does not cause craniofacial phenotypes, only mild neural crest migration defects (Jacques-Fricke et al., 2012). This discrepancy could be because DNMT3B is required much earlier in neural crest development than the Cres used are active, or because DNMT3B is also strongly expressed in other non-neural crest tissues including the neural and embryonic ectoderm. These primary effects of DNMT3B loss in other tissues is having a secondary effect on neural crest development. A recent study by Nowialis et al suggests DNMT3B carries dual functionality in mediating methylation without its catalytic domain and can function as an accessory cofactor supporting the catalytic activities of other DNMTs, including DNMT3A’s enzymatic activities (Nowialis et al., 2019). Investigating the cell type-specific nature of specialized dual roles of DNMT3B and its accessory function are needed to determine its functions in non-neural crest tissues or whether its function in neural crest development is cell autonomous.

Dual Activating and Repressing Functions of Histone Modifications

The core histones making up the nucleosome have tails that can be modified through post-translational modifications, including methylation, acetylation, deacetylation, phosphorylation, ubiquitination and sumoylation (Bannister & Kouzarides, 2011; Berger, 2007; Kouzarides, 2007). These modifications facilitate either chromatin compaction or relaxation. The histone marks present during neurulation and neural crest development dynamically change over the course of developmental time. This correlates with the specification and lineage commitments that progenitor cells accrue as their cell fates are established (reviewed in (Hu, Strobl-Mazzulla, & Bronner, 2014)). The functions of the proteins that facilitate dynamic histone modifications varies spatiotemporally across different cell types during development.

Depending on the histone protein, the residue, or the number of methyl groups being catalytically added, histone methylation is associated with either activation or repression of gene expression. For example, in most cases H3K4me1 marks poised enhancers/promoters and transcriptional activation; H3K4me3 marks active promoters and transcriptional activation; H3K27me3 indicates inactive promoters and transcriptional repression; H3K9me1 and H3K9me3 mark heterochromatin and are associated with transcriptional repression (reviewed in (Bannister & Kouzarides, 2011)). The addition of methyl groups is achieved through the activity of histone methyltransferases while histone demethylases remove them. Below are some examples of histone methyltransferases and demethylases associated with congenital craniofacial defects and neural crest development.

KDM4A, a member of the Jumonji family of demethylases, catalytically reverts histone trimethylation of H3K9me3 and H3K36me3 (Tan et al., 2008). In chick, KDM4A directly interacts with promoter regions of Sox10 and Snail2 that are occupied by H3K9me3 to ensure neural crest specification (Strobl-Mazzulla et al., 2010). While no congenital craniofacial anomalies have been associated with KDM4A yet, human variants in another demethylase, KDM5B, have been associated with intellectual developmental disorder and facial dysmorphism (Faundes et al., 2018). In mice, loss of Kdm5b results in disorganized cranial nerves, defects in eye development, incidences of exencephaly as well as homeotic skeletal transformations as a result of aberrant accumulation of H3K4me3 at normally inactive genes encoding developmental regulators (Albert et al., 2013). Mutations in another histone demethylase, KDM5C which catalyzes the demethylation of H3K4me2/3, has been associated with Claes-Jensen X-linked intellectual disability syndrome with facial dysmorphia (Hatch & Secombe, 2022; Ounap et al., 2012; Vallianatos et al., 2020). In Xenopus laevis, KDM5C is important for neural crest specification and migration as well as neurogenesis (Y. Kim et al., 2018).

Patients with mutations in plant homeodomain finger protein, PHF8, have Siderious X-linked intellectual disability and craniofacial dysmorphology (Sobering et al., 2023; Sobering et al., 2022). PHF8 is a histone demethylase that targets reversal of H4K20me1 and H3K9me1 at transcription start sites to initiate transcription. In zebrafish, PHF8 is necessary for NCC formation and differentiation of craniofacial skeletal derivatives by directly regulating expression the transcription factor, MsxB (Qi et al., 2010). Another PHF family member, PHF12, promotes neural crest cell migration by establishing the NCC epithelial-to-mesenchymal transition (Adams et al., 2008; Strobl-Mazzulla & Bronner, 2012; Yochum & Ayer, 2001), yet no known human congenital disease has been associated with PHF12 as of yet.

The epigenetic etiologies of Kabuki Syndrome are of interest due to the identification of two histone methylation modifiers associated with the disease, histone demethylase KDM6A and histone methyltransferase KMT2D (also known as MLL2) (Bogershausen et al., 2016; Lederer et al., 2012; Miyake, Koshimizu, et al., 2013; Miyake, Mizuno, et al., 2013; Ng et al., 2010). Kabuki syndrome is a multiple congenital anomaly syndrome characterized by distinctive facial features, developmental delay, intellectual disability and cardiovascular and musculoskeletal abnormalities (Lederer et al., 2012; Miyake, Koshimizu, et al., 2013; Miyake, Mizuno, et al., 2013; Ng et al., 2010). Most Kabuki syndrome cases have been associated with KMT2D which catalyzes the methylation of H3K4 to activate transcription. Though less common, some Kabuki syndrome patients harbor mutations in KDM6A which functions to demethylate H3K27 and remove repressive marks (Miyake, Koshimizu, et al., 2013). Both kmt2d morphant and mutant zebrafish develop phenotypes to those observed in Kabuki Syndrome patients including microcephaly, palate defects, abnormal ear development and cardiac defects, while kdm6a zebrafish morphants only develop craniofacial phenotypes (Serrano et al., 2019; Van Laarhoven et al., 2015). In mice, KDM6A, mainly functions through demethylase-independent transcriptional mechanisms to maintain neural crest stem-cell progenitor states and NCC viability to promote neural crest differentiation toward osteochondroprogenitor cell fates (Shpargel et al., 2017). Loss of KMT2D causes similar craniofacial malformations as those in KDM6A mutant mice, with some differences, such as a fully penetrate cleft palate, mandibular hypoplasia, and defects in cranial base ossification, suggesting KMT2D functions in later stages of NCC differentiation (Schwenty-Lara et al., 2020; Shpargel et al., 2020).

Two euchromatin histone methyltransferases, EHMT1 (also known as GLP) and EHMT2 (also known as G9a) are associated with Kleefstra syndrome (Kleefstra et al., 2009). These two histone methyltransferases promote the demethylation of H3K9 to H3K9me2. EHMT1 mutant mice phenocopied the disease characteristics of Kleefstra syndrome patients, particularly in the development of the brachycephalic crania, shorter or bent nose, and hypotelorism (Balemans et al., 2014). EHMT1 controls H3K9me2 to repress osteoblastic developmental programs (Balemans et al., 2014; N. Liu et al., 2015). Like EHMT1, EHMT2/G9a also regulates osteogenic differentiation during craniofacial development, though through opposing mechanisms. Neural crest specific ablation of G9a in mice causes craniofacial defects mimicking those present in Kleefstra syndrome, including hypoplasia of the lower jaw (Higashihori et al., 2017). G9a catalyzes the methylation of Twist to repress expression of progenitor state genes and facilitate osteogenic programs (Higashihori et al., 2017). Additionally, G9a directly activates Runx2 to promote osteoblast differentiation (Ideno et al., 2020). G9a can also cooperate with other epigenetic regulators and transcription factors to facilitate spatiotemporal activation and repression of developmental programs (Rai et al., 2010). The changes in function or gene targets of G9a depend on timing, tissue specificity, and the expression of other binding partners.

One other member of the histone methyltransferase family that has been studied more recently in neural crest development are the PRDM or positive regulatory domain methyltransferases, PRDM3 (also called MECOM) and PRDM16. Both PRDM3 and PRDM16 have been associated with development of non-syndromic cleft lip and palate as well as variation in normal facial morphology (Jugessur et al., 2010; Liu et al., 2012; Shaffer et al., 2016). PRDM16 is also considered a Pierre Robin Syndrome (PRS) loci, based on phenotypes observed in mutant mouse models that closely resemble the phenotypes observed in patients with PRS (mandibular hypoplasia, tongue displacement and cleft palate) (Bjork et al., 2010; Shull et al., 2022; Shull et al., 2020; Strassman et al., 2017). PRDM3 and PRDM16 are expressed in neural crest derived tissues, including the facial processes and exhibit intrinsic methyltransferase activity by modifying H3K9me3 and H3K4me3 (Baizabal et al., 2018; Pinheiro et al., 2012; Shull et al., 2020; Zhou et al., 2016). In zebrafish, PRDM3 and PRDM16 drive craniofacial chondrocyte differentiation, albeit through opposition functions, whereby PRDM3 acts a traditional repressor of gene expression while PRDM16 serves more as an activator of gene expression (Shull et al., 2022). Both share similar gene targets, including factors involved in Wnt/β-catenin signaling (Shull et al., 2022). An epigenomic atlas of human embryonic craniofacial development and identified a significantly enriched super-enhancer region encompassing the PRDM16 gene suggesting many important functions of this gene (and its paralog, PRDM3) in contributing to the spatiotemporal regulation of many gene regulatory networks in craniofacial development (Wilderman et al., 2018).

Histone acetylation, or the addition of acetyl groups to lysine residues on histone tails through the activity of histone acetyltransferases (HATs), allows for chromatin relaxation, increased DNA accessibility to transcription factors and active transcription. Histone deacetylases (HDACs) directly antagonize the activity of HATs by removing acetyl groups causing chromatin compaction and repression of gene expression. With their corresponding effects on gene expression, HATs have also been identified as co-transcriptional activators, while HDACs have been identified as transcriptional co-repressors (Bannister & Kouzarides, 2011; Kouzarides, 2007). As with the other histone modifiers described above, the dynamic nature of HATs and HDACs and their influences on gene expression suggests their importance in spatiotemporally controlling gene regulatory networks necessary for developmental processes.

The lysine acetyltransferase, KAT6A was identified with patients presenting with Arboleda-Tham Syndrome (also termed KAT6A syndrome), characterized by developmental delay, intellectual disability, and distinct facial features (Arboleda et al., 2015; Bae et al., 2021; Satoh et al., 2017; Tham et al., 2015). De novo mutations in KAT6A have also been associated with pan-craniosynostosis (Marji et al., 2021). Kat6a mutant mice develop phenotypes similar to those observed in humans, including palate defects and abnormal facial structures (Voss et al., 2012). KAT6A acetylates H3K9 and controls the expression of the transcription factor TBX1 and subsequently neural crest migration (Voss et al., 2012). Loss of Kat6a in mice alters expression of craniofacial patterning genes, including members of the DLX gene family (Dlx1/2/3/4/5), genes regulated by Dlx factors (Barx1, Gbx2, Osr2, and Sim2), and genes important for osteoblast differentiation (Runx2, Pax9 and Nkx3-1) (Vanyai et al., 2019). In zebrafish, kat6a (also called moz), along with hox genes, specifies and patterns neural crest cells toward specific skeletal structures (Crump et al., 2006). KAT6A’s paralog, KAT6B, is associated with several congenital diseases with facial dysmorphia (Noonan Syndrome, Ohdo Syndrome, and Genitopatellar Syndrome) (Campeau et al., 2012; Clayton-Smith et al., 2011; Simpson et al., 2012; Szakszon et al., 2013; Wiesel-Motiuk & Assaraf, 2020). Like KAT6A, KAT6B activates osteogenic differentiation by interacting with RUNX2 through the C-terminal serine/methionine domains (Pelletier et al., 2002). Two other histone acetyltransferases, KAT2A (also known as GCN5) and KAT2B (also known as PCAF) control craniofacial chondrocyte differentiation and maturation programs through direct histone acetylation of H3K9 as well as through non-histone acetyltransferase activity and indirect control of the mTORC1 pathway (Dong et al., 2023; Pezoa et al., 2020; Sen et al., 2018).

Like HATs, histone deacetylases, HDACs, have important roles in controlling neural crest EMT and migration. The transcriptional repressor, Snail2, interacts with an HDAC/Sin3A repressive complex via an adaptor protein PHD12, to repress expression of the adhesion molecule Cad6B in premigratory neural crest cells (Strobl-Mazzulla & Bronner, 2012). HDACs are important later in development in regulating downstream differentiation of neural crest cells toward different craniofacial derivatives. In zebrafish, loss of hdac1 results in decreased hoxb3a, dlx2, and dlx3-expressing posterior pharyngeal arch precursors that are specified due to increased apoptosis of these cells. Anterior pharyngeal arch precursors survive but have limited differentiation capacity as evidenced by failure of chondrocyte precursors in these regions to terminally differentiate (Ignatius et al., 2013; Pillai et al., 2004). In mice, HDAC1 and HDAC2 are necessary for neural crest proliferation, development in the pharyngeal arches and formation of the craniofacial structures by repressing cyclin-dependent kinase inhibitors (Milstone et al., 2017). These studies provide mechanistic insight into the etiology of some patients with craniofacial abnormalities with unknown origin in chromosomal regions. For example, copy number variations at 1p35, a region encompassing HDAC1 and 6q21-22, a region encompassing HDAC2 were found in patients with craniofacial malformations, perhaps due to altered neural crest cell proliferation and differentiation potential (Hudson et al., 2014; Rao & LaBonne, 2018).

HDAC4 has been associated Brachydactyly-Mental Retardation Syndrome and non-syndromic clefts (Park et al., 2006; Villavicencio-Lorini et al., 2013; Williams, Aldred, et al., 2010). In zebrafish, hdac4 is highly expressed in migratory cranial neural crest cells and knockdown of hdac4 reduces cranial neural crest cell populations that reach the pharyngeal arches which ultimately impairs differentiation and formation of the cartilaginous craniofacial structures (DeLaurier et al., 2012). In mice, loss of Hdac4 alters frontal bone formation due to decreased proliferative capacity of osteoblast precursors as a result of dysregulated cell cycle-related genes (Ha et al., 2022). Similarly, loss of Hdac8, both globally and specifically in murine neural crest cells impairs cranial morphogenesis with ossification defects in the frontal and interparietal bones (Haberland et al., 2009). HDAC8 represses aberrant expression of homeobox transcription factors Otx2 and Lhx1 necessary for patterning portions of the neural crest derived calvaria (Haberland et al., 2009). Mutations in HDAC8 have been identified in patients with Cornelia de Lange Syndrome and Wilson-Turner Syndrome, both associated with slow and delayed developmental growth and abnormal formation of the craniofacial complex (Deardorff et al., 2012; Deardorff et al., 2016; Gao et al., 2018; Harakalova et al., 2012; Kaiser et al., 2014; Lucia-Campos et al., 2022; Mio et al., 2021; Shaffer et al., 2016).

In summary, histone modifications, either through lysine methyltransferases, lysine demethylases, HATs or HDACs, have important roles in controlling methylation and acetylation patterns of different histone marks necessary to facilitate proper spatiotemporal activation or repression of neural crest gene regulatory networks. As described above, their activities have numerous effects on neural crest development from proliferation, specification, migration and differentiation. Alterations in their activity can hinder neural crest development and result in congenital craniofacial birth defects.

Large Protein Complexes Influence Chromatin Structure: Polycomb Repressive Complex

The polycomb repressive complex, PRC1 and PRC2, are well studied transcriptional repressor complexes. Both PRC1 and PRC2 are multimeric protein complexes that function to suppress gene expression (Piunti & Shilatifard, 2021, 2022). One well known protein component of PRC2 is EZH2 or enhancer of zest homolog 2. EZH2 carries the catalytic activity of PRC2 by initiating mono-di-tri methylation of H3K27 (O’Carroll et al., 2001; Rea et al., 2000). Conditional ablation of Ezh2 in pre-migratory murine neural crest cells leads to a de-repression of Hox genes in cranial neural crest cells (H. Kim et al., 2018; Schwarz et al., 2014). This misexpression of Hox genes maintains cranial neural crest cells in a pre-differentiation state, thereby suppressing activation of osteochondroprogenitor programs, affecting subsequent cartilage and bone formation (Schwarz et al., 2014). Numerous other studies have demonstrated the importance of EZH2 in regulating osteochondroprogenitor differentiation programs (Camilleri et al., 2018; Dudakovic et al., 2018; Dudakovic et al., 2015; Dudakovic et al., 2020). Mutations in EZH2 have been identified in patients with Weaver Overgrowth Syndrome, a genetic condition that causes bone overgrowth (Cohen et al., 2016; Lui et al., 2018; Tatton-Brown et al., 2013).

Ring1b/Rnf2, which functions as an E3 ubiquitin ligase in the PRC1 complex, controls craniofacial chondrocyte differentiation (van der Velden et al., 2013). Zebrafish ring1b mutants do not develop cranial cartilage or bone due to impaired chondrocyte differentiation. Other PRC proteins, PHC1 and PHC2, are important in neural crest cell formation and patterning craniofacial skeletal elements by regulating Hox expression (Isono et al., 2005; Marc et al., 2023). These two factors have also been associated with congenital diseases including CATCH-22 syndrome (PHC1) as well as patients presenting with DiGeorge Syndrome like phenotypes (PHC1) and non-syndromic cleft lip and palate (PHC2) (Marc et al., 2023; Shirai et al., 2002; Takihara et al., 1997; Wilson et al., 1993). ASXL proteins (Additional sex combs-like) ASXL1/2/3, which serve as scaffolds in the PRC complex, have important roles in early neural crest development, from generation of neural crest to delamination and emigration (Lichtig et al., 2020; Matheus et al., 2019). Patients with mutations in ASXL genes have been linked to Bohring-Opitz Syndrome (ASXL1), Shashi-Pena Syndrome (ASXL2) and Bainbridge-Ropers Syndrome (ASXL3) (Dinwiddie et al., 2013; Koboldt et al., 2018; Kuechler et al., 2017; Shashi et al., 2016; Shashi et al., 2017; Zhao et al., 2021). These neurodevelopmental disorders are different from one another, but share similar phenotypes including dramatic craniofacial defects, microcephaly and developmental delay and severe intellectual disability. In Xenopus laevis, knockdown of ASXL3 disrupts neural cell fate specification and the phenotype of the mutant animals resembles the phenotypes observed in Bainbridge-Ropers patients (Lichtig et al., 2020).

Large Protein Complexes Influence Chromatin Structure: ATP-Dependent Chromatin Remodeling Complexes

ATP-dependent chromatin remodeling complexes control gene expression by altering the position or structure of chromatin in an ATP-dependent manner. These complexes remodel and reposition chromatin to create nucleosome free regions that allow transcription factors access to the DNA and initiate transcription (Bannister & Kouzarides, 2011; Kouzarides, 2007). One ATP-dependent chromatin remodeling complex, the SWI/SNF complex consists of several components that function together to drive gene expression, these include ARID1A and ARID1B, BRG1 (SMARCA4), BRM, BAF155, BAF157, as well as SMARCA2, SMARCB1 and SMARCE1, among others (Cenik & Shilatifard, 2021; Sokpor et al., 2017). Independent loss of each separate component of the complex has profound effects on neural crest development and formation of the craniofacial complex. The majority of patients presenting with Coffin-Siris Syndrome carry mutations in one of the SWI/SNF chromatin remodeling complex subunits, particularly in the subunits found in ARID1A and ARID1B (Diets et al., 2019; Kosho, Miyake, et al., 2014; Kosho, Okamoto, et al., 2014; Miyake et al., 2014; Pagliaroli et al., 2021; Santen et al., 2013; Tsurusaki et al., 2012; Tsurusaki et al., 2014; Tzeng et al., 2014; Wieczorek et al., 2013; Xia et al., 2023). This disease is characterized by developmental disability, distinct facial features and abnormalities of the fifth fingers or toes (Tsurusaki et al., 2012; Vasileiou et al., 2018). Loss of Arid1a in neural crest cells causes severe craniofacial defects in mice, including shortened snouts, low set ears and reduced ventral cranial bones (Chandler & Magnuson, 2016). Recently, Pagliaroli et al utilized ARID1B haploinsufficient Coffin-Siris patient-derived iPSCs to model cranial neural crest development. In this study, ARID1B was active only during the first stage of neural crest formation, coinciding with neuroectoderm specification (Pagliaroli et al., 2021). ARID1B regulates the exit from pluripotency and lineage commitment by inactivating NANOG and SOX2 gene regulatory networks. In control iPSCs, ARID1A maintains the pluripotency state but at the onset of differentiation, cells transition from ARID1A to ARID1B to trigger the pluripotency exit and begin differentiation toward specific cell types. However, in iPSCs derived from Coffin-Siris patients, the cells fail to perform the ARID1A/ARID1B switch and instead maintain an ARID1A-dependent pluripotency state throughout all stages of cranial neural crest formation, impairing differentiation of neural crest derivatives (Pagliaroli et al., 2021).

Mutations in BRG1 is also associated with Coffin-Siris Syndrome (Miyake et al., 2014; Tsurusaki et al., 2012; Tsurusaki et al., 2014; Tzeng et al., 2014). BRG1 is the ATPase subunit of the SWI/SNF Brg1/Brahman associated factor (BAF) complex. Loss of Brg1 in neural crest cells leads to aberrant survival, migration and differentiation. Brg1 is necessary for maintaining a multipotent reservoir of neural crest cells primed for differentiation (Li et al., 2013). In zebrafish brg1 is necessary for neural crest induction and specification as expression of neural crest specifiers are severely affected in brg1 morphants, while mutants have more pronounced craniofacial abnormalities (Eroglu et al., 2006). Additionally, Brg1 may control chondrocyte differentiation through interactions between Med14 a component of the Mediator complex and BAF complex (Lou et al., 2015).

Loss of BAF155/BAF170, the core subunits required for the assembly, stability, and function of the BAF complex in neural crest cells, leads to a variety of neural crest defects, including cardiac outflow tract defects and craniofacial abnormalities by controlling expression of pathways essential for neural crest survival, migration, proliferation and differentiation (Bi-Lin et al., 2021). Though their functions are currently less defined, new evidence suggests different members of the SWI/SNF complex that can bind and read chromatin, SMARCA2, SMARCB1 and SMARCE1 control neural crest formation and differentiation of different craniofacial structures (Diets et al., 2019; Kosho, Okamoto, et al., 2014; Sanchez & Rojas, 2017; Sousa et al., 2014; Tang et al., 2017; Van Houdt et al., 2012; Vitte et al., 2017; Yano et al., 2018).

Another ATP-dependent chromatin domain helicase DNA-binding domain member, CHD7 has been well characterized in neural crest development. Mutations in CHD7 are present in patients with CHARGE syndrome (Lalani et al., 2006). CHD7 cooperates with PBAF (polybromo- and BRG1-associated factor-containing complex) to promote neural crest gene expression (activation of Sox9, Twist, and Slug) and neural crest cell migration (Bajpai et al., 2010). Knockdown or overexpression of CHD7 in Xenopus embryos recapitulates the major phenotypes in CHARGE syndrome (Bajpai et al., 2010). Loss of Chd7 in mice leads to craniofacial defects including frontal and occipital bone dysplasia, hypoplasia of the maxillary shelves and mandible, and a cleft palate (Bosman et al., 2005; Hurd et al., 2010; Sperry et al., 2014). A zebrafish chd7 genetic mutant recapitulates the craniofacial and cardiovascular defects that co-occur in patients with CHARGE syndrome and further showed disrupted chondrocyte cellular organization and stacking within cartilage structures (Y. Sun et al., 2022). The variety of CHD7 mutant animal models for CHARGE syndrome will help delineate the molecular functions of CHD7 and how alterations to its activity contribute to the etiology of CHARGE syndrome.

The Function of Other Epigenetic Regulators

There are other epigenetic regulators that are important for craniofacial development and do not fall into the above categories. For example, the special AT-rich sequence binding protein 2 is a chromatin reader that binds nuclear matrix-attachment regions to activate transcription (Britanova et al., 2005; Dobreva et al., 2003). In humans, translocations that involve the chromosomal region 2q32-q33 have been found to encompass the SATB2 gene (Urquhart et al., 2009; Zarate & Fish, 2017). These translocations are associated with cleft palate (FitzPatrick et al., 2003). Other mutations in SATB2 have now been associated with SATB2-associated syndrome which is characterized by intellectual disability, developmental delay with craniofacial and dental abnormalities (Huang et al., 2022; Kikuiri et al., 2018; Rainger et al., 2014; Scott et al., 2019; Urquhart et al., 2009; Zarate et al., 2019; Zarate & Fish, 2017). Satb2 mutant mice have both craniofacial abnormalities that resemble the phenotypes of humans with translocations in SATB2, as well as defects in osteoblast differentiation (Dobreva et al., 2006). SATB2 regulates skeletal patterning by controlling expression of Hox genes and later drives osteogenic programs by directly regulating the expression of Runx2 and ATF4 as well as other craniofacial patterning genes (Pax9, Msx1, Alx4, and Lhx7). These studies implicate two functions of SATB2 during neural crest development, one early to promote neural crest cell survival and later, to drive osteogenic differentiation programs during the formation of craniofacial structures.

Conclusions and Future Challenges for the Field

The formation of the craniofacial complex relies on precise activation and or repression of different gene regulatory network and signaling cascades during cranial neural crest cell development. Epigenetic factors control the upstream transcriptional networks that govern the complex process of craniofacial development. Because these factors control genomic access to important developmental transcriptional and signaling genes, alterations to the timely orchestration of these events can compromise neural crest cell development and ultimately contribute to the etiology of congenial birth defects that affect the craniofacial skeleton. These birth defects include, but are not limited to the ones highlighted here, CHARGE Syndrome, Coffin-Siris syndrome, Keefstra Syndrome, Weaver Syndrome, Kabuki Syndrome, as well as non-syndromic craniofacial abnormalities including cleft lip and palate, among others. While there are other factors associated with the development of these diseases, there is growing evidence suggesting the importance of epigenetic regulators and how their altered activity contributes to these congenital diseases. In this review, we discussed the functions of epigenetic modifiers and the vastly growing number of syndromes with mutations specifically associated with these chromatin remodelers. We also discussed the ongoing work to delineate the functions of these factors during normal development and how alterations to their activities contributes to developmental disorders affecting neural crest development and formation of the craniofacial skeleton.

The influence of environmental factors on the genome and epigenome during embryonic and fetal development can have drastic consequences on the development of highly sensitive tissues including the neural crest and subsequently neural crest derived tissues, particularly the craniofacial skeleton. Understanding how such environmental stressors impact the epigenome is necessary to understand how they influence craniofacial development. Defining what extent environmental and teratogenic factors have on neural crest and craniofacial development will be important for determining how outside factors can influence the epigenome, which could in turn be used to provide therapeutic or environmental intervention during a pregnancy. While this represents a challenge it is also presents a therapeutic opportunity. Treatment options that modify methylation (i.e. folic acid) and other modifications can be delivered to the pregnant mother to provide a positive effect on the developing embryo and fetus. While outside stressors influence embryonic and fetal development, it is also interesting to think about how environmental factors impact facial morphology later in life, especially given the changes in facial structure that can occur from early postnatal development, adolescence and into aging adulthood.

Given the widespread and varying functionality of epigenetic factors across different stages of neural crest development, that correlate with the gene expression changes needed to facilitate the transitions from early neural crest specification to differentiation of different derivatives, it makes sense that different progenitor cell populations, cell lineages and tissues are associated with different combinations of epigenetic modifications over the course of their differentiation trajectories. The dynamic changes in and the factors that control the deposition of the marks in neural crest cells and the craniofacial specific derivatives have only started to be elucidated. Further investigating these functions will provide us with a greater understanding of how alterations to these factors and their epigenetic activities leads to congenital craniofacial birth defects. New advances in epigenetic technologies will allow for a deeper investigation into the mechanisms of epigenetic function, including Cleavage Under Targets and Release Using Nuclease (CUT&RUN), single cell Multiomics, ATAC-seq, and HiC-seq to assess enhancer-promoter interactions and 3D chromatin architecture in the context of neural crest development and craniofacial skeletal biology. These sequencing technologies can be used in both human control samples versus syndromic patient-derived samples, as well as with animal models (genetic mutants as compared to wildtype). These studies will greatly expand our understanding of how epigenetic regulators control these gene regulatory networks from a mechanistic standpoint.