Enhancers: bridging the gap between gene control and human disease

Enhancers: bridging the gap between gene control and human disease Abstract Enhancers are a class of regulatory elements essential for precise spatio-temporal control of gene expression during development and in terminally differentiated cells. This review highlights signature features of enhancer elements as well as new advances that provide mechanistic insights into enhancer-mediated gene control in the context of three-dimensional chromatin. We detail the various ways in which non-coding mutations can instigate aberrant gene control and cause a variety of Mendelian disorders, common diseases and cancer. Introduction Precise gene control is central to life. Transcriptional enhancers are cis-regulatory sequences that recruit transcription factors (TFs) and play a fundamental role in regulating which genes cells express, the timing of their expression and the levels of their expression. Most enhancers are located in non-coding regions of the genome, previously considered ‘junk,” DNA, and now widely accepted to be functional. While the number of genes is ∼20 000 in the human genome, there are millions of candidate enhancers (1,2). The enhancerome of any given human cell type occupies 5–10 times more genomic real estate than the exome. Although weakly conserved across species, enhancer elements are among the most highly constrained sequences across humans (3). They are enriched for germline variants associated with both rare and common diseases, and somatic mutations that disrupt enhancer function can cause cancer. Armed with genomic technologies that facilitate interrogation of the enhancer epigenome and 3D chromatin architecture at high resolution, new insights into enhancer–gene control are being uncovered at an unprecedented rate. This review focuses on basic principles of enhancer regulation with an emphasis on human diseases caused by germline and somatic non-coding mutations that disrupt enhancer–gene control via trans or cis mechanisms. Features of Enhancer Elements Enhancer elements are located in DNase I hypersensitive regions of chromatin flanked by histone H3 covalently modified with monomethylation of lysine 4 (H3K4me1) (4,5) (Fig. 1). Histone variants, H2A.Z and H3.3, are also incorporated at active enhancers (6). Covalent histone modifications in addition to H3K4me1 may also be present and the specific composition of histone marks generally correlates with the activity state of the enhancer. Enhancers can exist in decommissioned (no histone modification), poised (H3K4me1 and H3K27me3), primed (H3K4me1) or active (H3K4me1 and H3K27ac) states (7–9). During development and in terminally differentiated cells, enhancers can switch between states. Upon embryonic stem cell differentiation, for example, poised enhancers lose the repressive H3K27me3 mark and gain H3K27ac. The enhancer state change occurs concomitantly with a switch in the expression of the target gene from off to on and the transition in cell state from undifferentiated to differentiated. Enhancer activation is a step-wise process mediated by a variety of sequence-specific TFs chromatin-remodeling complexes and coactivators (10). RNA Polymerase II (Pol II) is also involved, directly binding to enhancer sequences and transcribing them into enhancer RNA (eRNA) (11). p53 may also participate in eRNA production (12). eRNA levels correlate with enhancer activity, and eRNA can be used as a marker for genome-wide identification of active enhancer elements (13). Enhancer transcription can lead to the formation of R-loops (14), which, if left uncorrected, can induce replication stress, genomic instability and other hallmark features of cancer (15). Figure 1. View largeDownload slide Features of active enhancer elements (see text). Transcription factor (TF), mediator (Med), RNA polymerase II (RNA Pol II), bromodomain-containing protein 4 (BRD4), CCCTC-binding factor (CTCF), enhancer RNA (eRNA), lysine methyltransferase 2C (KMT2C), lysine methyltransferase 2D (KMT2D). Figure 1. View largeDownload slide Features of active enhancer elements (see text). Transcription factor (TF), mediator (Med), RNA polymerase II (RNA Pol II), bromodomain-containing protein 4 (BRD4), CCCTC-binding factor (CTCF), enhancer RNA (eRNA), lysine methyltransferase 2C (KMT2C), lysine methyltransferase 2D (KMT2D). Enhancers communicate with genes through physical interactions with their promoters, mediated by Mediator and Cohesin complexes (16) as well as ZNF143 (17) and YY1 (18). The histone methyltransferases (MLL3/4 or KMT2C/B), and the canonical H3K4me1 enhancer mark these enzymes deposit, are also necessary for establishing enhancer–promoter interactions (19). Most enhancer–gene interactions are contained within topologically associated domains (TADs) (20), self-interacting megabase-size genomic regions bordered by the insulator protein CTCF. Within TADs, CTCF can contribute to formation of insulated neighborhoods that further seclude enhancers and their gene targets (21). Enhancers can be located close to or several hundred kilobases away from their gene targets and they do not necessarily regulate their nearest gene. Multiple enhancers often regulate a single gene, and multiple genes can be regulated by a single enhancer. Super Enhancers Super enhancers (SEs) and stretch enhancers are clusters of enhancers found across diverse cell types (22–24). SEs contain supranormal levels of the active enhancer histone mark H3K27ac and many other enhancer-associated proteins, like Mediator and Pol II. They are typically several kilobases in length, exquisitely cell-type specific, and regulate genes that control cell state. In embryonic stem cells for example, SEs regulate master TFs of pluripotency, Oct4, Sox2 and Nanog (23). Tumor cells often acquire SEs near oncogenes that are known tumor dependency genes, like MYC (25,26). They are also highly enriched for Genome-wide association studies (GWAS)-identified SNPs that predispose to several common diseases. The constituents of SEs physically interact with one another and their gene targets in 3D chromatin and these interactions are likely essential for maintenance of SE function (27). Recent studies have exploited CRISPR-Cas9 editing technologies to dissect SEs and quantify the functional contribution of individual SE constituents to their target gene levels. These studies have consistently shown that SE constituents function independently and all constituents participate in the regulation of their target genes, with some enhancers contributing more than others. There appears to be no common regulatory modality; some act additively like those at the α-globin locus (28), and others work synergistically, like those at the mammary Wap locus (29). Common disease-associated variants in SE constituents can also influence the expression of their target genes via different modalities (30). Recently, a global analysis of SE function found that while all enhancers are necessary for maximal SE activity, there exists a set of ‘hub enhancers’ whose knockout results in particularly profound decrease in SE activity (31). Hub enhancers are enriched for CTCF and cohesin and are critical for maintaining physical interactions between SE constituents as well as other enhancers within the same TAD. A recently proposed ‘phase separation’ model of transcriptional control may explain how SEs promote consistent and high level transcription. In this model, binding of TFs and other regulators to SE constituents stimulates aggregate formation and their subsequent compartmentalization within the cell nucleus (32). Mutations in Enhancer Regulators Several Mendelian diseases are caused by mutations in genes encoding proteins that function to establish, maintain, or interact with enhancers (Table 1). Most are congenital, multiple malformation diseases caused by heterozygous mutations that arise de novo, implicating haploinsufficiency as the pathogenic mechanism. A key player in enhancer activation is the ATP-dependent BRG1/BRM associated factor complex, or BAF, which slides or evicts nucleosomes to make way for TFs and other coactivators. BAF also associates with H3K4me1 at enhancers (33). Mutations in several subunits of the BAF complex cause Coffin-Siris syndrome, a condition characterized by developmental delays and unique skeletal malformations (34,35). CHARGE syndrome (coloboma, heart defects, atresia choanae, growth retardation, genital abnormalities, and ear abnormalities) is another disease caused by mutation of a chromatin remodeler that acts at enhancers, CHD7 (36). A related CHD family member, CHD8, is implicated in autism (37) and may also function in enhancer activation (38). Table 1. Diseases caused by mutations in proteins that function in establishing, maintaining, or regulating enhancers Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  Table 1. Diseases caused by mutations in proteins that function in establishing, maintaining, or regulating enhancers Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  The switching of enhancers between primed and active states is mediated in part by enzymes that deposit or remove covalent histone modifications, or ‘writers’ and ‘erasers’. An illustrative example of the crucial role of these proteins in normal development is Kabuki syndrome (KS), characterized by distinctive facies, intellectual disability, unique skeletal abnormalities and growth delay. KS is caused by mutations in KMT2D, a histone methyltransferase that writes the canonical H3K4me1 enhancer mark on chromatin (39). KS can also be caused by mutation of KDM6A, a histone demethylase that erases the H3K27me3 mark (40). P300 and CBP, coactivators that bind acetylated lysines at active enhancers are mutated in Rubinstein-Taybi syndrome, characterized by short stature, intellectual disability, distinctive facies and broad thumbs and toes (41). Several of the same genes with enhancer-related functions are frequently somatically mutated in cancer. Mutations in genes encoding subunits of the BAF complex occur in 20% of all human cancers (42). KMT2D, KDM6A, ARID1A, KTM2C, P300, CREBBP and KMT2A, are all constituents of machinery cells use to establish enhancers and are among the most frequently mutated genes in bladder cancer (43). Mutations in enhancer-associated histone variant H3.3 genes have been implicated in pediatric brain cancers (44), although the exact mechanism by which the oncohistone promotes tumorigenesis remains unresolved. Although not generally considered ‘enhanceropathies’ like the diseases described above, Cornelia de Lange syndrome (CdLS), Roberts syndrome, and other ‘cohesinopathies’ are caused by mutations in various components of the cohesin complex (SCC2/NIPBL, SMC1A, SMC3, RAD21, ESCO2 and others) that function in chromatin looping and hence play an important role in facilitating enhancer-promoter interactions (45–47). Moreover, mutations in BRD4 were recently shown to cause a Cornelia de Lange-like syndrome (48). BRD4 binds to SEs and also engages NIPBL, which is required for loading the cohesin looping complex onto DNA (49). The proposed model is that BRD4 mutation disrupts looping and transcription at SE-associated loci and as such, the authors noted that CdLS can be considered a “transcriptomopathy”. Mutations Disrupting Enhancer–Gene Control There are a variety of ways in which genetic mutation of non-coding regions can disrupt enhancer–gene regulation and cause disease (Table 2). Deletions can result in either gene silencing by removing active enhancers, or gene activation by repositioning them closer to enhancers. Deletions can also remove TAD boundaries, allowing for ectopic interactions between enhancers and promoters normally insulated from one another in adjacent TADs. In a variant of bracydactyly, for example, deletions removing TAD boundaries creates ectopic interactions between the EPHA4 TAD domain and PAX3 (50). Insertions may create novel TF binding sites at enhancer sequences (51). Translocations can breakup SEs, or displace enhancers from their endogenous targets and relocate them to other chromosomes (52). Inversions can force a promoter to adopt a repositioned enhancer, resulting in increased gene transcript output (53). Alternatively, inversions can reposition an enhancer to be out of range of its promoter (54). Duplications can create ectopic enhancer sequences that can act to increase target gene transcription (55). Point mutations can modulate TF affinity at enhancer sequences (56). Virtually all of these types of non-coding mutations can arise somatically and have been implicated in oncogenesis. For example, a translocation that repositions immunoglobulin heavy chain enhancers near the MYC oncogene is a common driver mutation in Burkitt’s lymphoma (57). In a variety of epithelial cancers, copy number gains containing SEs are known to activate oncogenes (58). In T-ALL, driver indel mutations at the TAL1 locus can create binding sites for MYB, a pioneer transcription factor that engages the indel, instigating SE formation and TAL1 activation (59). Microdeletions can disrupt boundaries of insulated neighborhoods, leading to oncogene activation (60). Table 2. Various types of mutations that impact enhancer function and their associated diseases Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  Table 2. Various types of mutations that impact enhancer function and their associated diseases Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  GWAS SNPs and Enhancers GWAS have successfully identified loci associated with several human traits and susceptibility to numerous common diseases, including heart disease, type 2 diabetes (T2D), multiple sclerosis and cancer. Sixty to eighty percent of the associated SNPs map to enhancers and/or SEs (61). There is also a strong correlation between associated SNPs and enhancers active specifically in tissues involved in disease pathology. SNPs associated with T2D for example, often reside within enhancers active exclusively in pancreatic islet cells. SNPs associated with risk for multiple sclerosis, generally considered to be an immune-related disease, lie in enhancers active in immune-related cell types. These findings offer mechanistic insights into the basis of common diseases and provide a potential explanation for why some tissues are affected while others are spared. The prevailing model is that the disease-associated SNPs impact gene regulation by modulating TF binding in cell types where the enhancer is active. There is compelling support for this model. Increased risk for melanoma is conferred by an enhancer indel that increases the affinity of RECQL, leading to increased PARP1 expression and contributing to melanogenesis (62). A risk SNP for prostate cancer increases RFX6 expression by increasing binding of HOXB13 at the RFX6 enhancer (63). SNPs associated with fetal hemoglobin levels map to BLC11A enhancers in human erythroblasts and modulate GATA1 and TAL1 binding (64). Mechanistic studies of GWAS–enhancer SNPs have also highlighted the complexity of enhancer–gene interactions in 3D chromatin. For example, an obesity associated enhancer variant within an intron of FTO, presumed to influence FTO expression, physically interacts with and impacts the expression of an altogether different gene, IRX3, located megabases away (65). Moreover, more than one gene may be targeted. The T2D risk SNP rs7163757 resides within a B-islet cell-specific stretch enhancer and modulates expression of two inflammatory cytokines, C2CD4B and C2CD4A (66). The targets of enhancer variants may not necessarily be protein coding genes, but can also be non-coding genes including miRNAs and lincRNAs. Investigations of GWAS enhancer SNPs have also helped nominate new cell types in common disease pathophysiology. SNPs associated with Alzheimer’s Disease risk correlate not only with fetal brain-specific enhancers as may have been expected, but also immune cell-specific enhancers, thereby implicating immune-related processes in AD predisposition (67). The success of the above mentioned studies suggests that the veritable gap between SNP association and function is steadily closing. Although successful in pinpointing functional variants at GWAS loci, often these studies do not rule out other variants in the region that could also be contributing to the correlation between genotype and target gene expression. There may be more than one casual SNP, particularly at loci where multiple disease-associated SNPs lie in SE constituents that physically interact with one another and their gene targets in 3D chromatin. As recently demonstrated, the SNPs at SE-associated loci may collude to influence target gene expression and the clinical risk to disease, and the influential SNPs may not necessarily be in linkage disequilibrium with one another (30,68). Furthermore, given that enhancers respond to specific physiological, pathological or environmental conditions, the regulatory effects of enhancer SNPs may be context specific (69). While considerable progress has been made, there is a clear need for functional studies that go beyond the standard eQTL and reporter assays to nominate causal variants. Strategies in which risk SNPs are directly edited in the genome are needed. Concluding Remarks As highlighted above, a variety of Mendelian diseases and cancers are caused by mutations in genes encoding proteins that function in trans in global transcriptional control either through regulating enhancer activity or enhancer–gene interactions via chromatin looping. These have been termed enhanceropathies and cohesinopathies, but given that the true pathology lies in transcription, ‘transcriptomopathies’ might be more precise descriptor for these diseases. Because clinical sequencing has largely focused on the exome, the number of genetic diseases due to non-coding mutations that disrupt enhancer–gene control in cis is not yet known. Given the sizeable percentage of Mendelian diseases currently explained by mutations in the exome (30%), the proportion of diseases due to mutations in non-coding regions is probably much smaller, even in spite of the fact that the enhancerome is 5–10× larger than the exome. Consistent with this prediction, whole genome sequence analysis of a large cohort of patients with developmental delay found that 1–3% without a diagnostic coding mutation had de novo mutations in regulatory elements (compared to 42% in the exome) (70). However, with respect to risk to common diseases, the proportion of causal variants in noncoding regions likely dwarfs that of the exome. Similarly, in cancer, and in particular cancers with high mutation rates, excessive aneuploidy and other structural rearrangements, functional mutations in non-coding regions are likely to be fairly common. Whole genome sequence analysis of tumor genomes paired with epigenome analysis should help shed light on the prevalence of noncoding cancer driver mutations. These studies are essential for the full potential of targeted therapies and precision medicine to be realized. Acknowledgements We thank Zachary Faber and Andrew Morton for critically reading the manuscript and their insightful comments. Conflict of Interest statement. None declared. Funding This work was supported by the National Institutes of Health [R01DA043980, R01CA193677, R01CA204279, R01CA143237 to P.C.S.]. References 1 ENCODE Project Consortium. ( 2012) An integrated encyclopedia of DNA elements in the human genome. Nature , 489, 57– 74. CrossRef Search ADS PubMed  2 Roadmap Epigenomics Consortium, Kundaje A., Meuleman W., Ernst J., Bilenky M., Yen A., Heravi-Moussavi A., Kheradpour P., Zhang Z., Wang J., Ziller M.J. ( 2015) Integrative analysis of 111 reference human epigenomes. Nature , 518, 317– 330. Google Scholar CrossRef Search ADS PubMed  3 di Iulio J., Bartha I., Wong E.H.M., Yu H.-C., Lavrenko V., Yang D., Jung I., Hicks M.A., Shah N., Kirkness E.F. et al.   ( 2018) The human noncoding genome defined by genetic diversity. Nat. Genet ., 50, 333– 337. 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Enhancers: bridging the gap between gene control and human disease

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Oxford University Press
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© The Author(s) 2018. Published by Oxford University Press. All rights reserved. For permissions, please email: journals.permissions@oup.com
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0964-6906
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1460-2083
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10.1093/hmg/ddy167
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Abstract

Abstract Enhancers are a class of regulatory elements essential for precise spatio-temporal control of gene expression during development and in terminally differentiated cells. This review highlights signature features of enhancer elements as well as new advances that provide mechanistic insights into enhancer-mediated gene control in the context of three-dimensional chromatin. We detail the various ways in which non-coding mutations can instigate aberrant gene control and cause a variety of Mendelian disorders, common diseases and cancer. Introduction Precise gene control is central to life. Transcriptional enhancers are cis-regulatory sequences that recruit transcription factors (TFs) and play a fundamental role in regulating which genes cells express, the timing of their expression and the levels of their expression. Most enhancers are located in non-coding regions of the genome, previously considered ‘junk,” DNA, and now widely accepted to be functional. While the number of genes is ∼20 000 in the human genome, there are millions of candidate enhancers (1,2). The enhancerome of any given human cell type occupies 5–10 times more genomic real estate than the exome. Although weakly conserved across species, enhancer elements are among the most highly constrained sequences across humans (3). They are enriched for germline variants associated with both rare and common diseases, and somatic mutations that disrupt enhancer function can cause cancer. Armed with genomic technologies that facilitate interrogation of the enhancer epigenome and 3D chromatin architecture at high resolution, new insights into enhancer–gene control are being uncovered at an unprecedented rate. This review focuses on basic principles of enhancer regulation with an emphasis on human diseases caused by germline and somatic non-coding mutations that disrupt enhancer–gene control via trans or cis mechanisms. Features of Enhancer Elements Enhancer elements are located in DNase I hypersensitive regions of chromatin flanked by histone H3 covalently modified with monomethylation of lysine 4 (H3K4me1) (4,5) (Fig. 1). Histone variants, H2A.Z and H3.3, are also incorporated at active enhancers (6). Covalent histone modifications in addition to H3K4me1 may also be present and the specific composition of histone marks generally correlates with the activity state of the enhancer. Enhancers can exist in decommissioned (no histone modification), poised (H3K4me1 and H3K27me3), primed (H3K4me1) or active (H3K4me1 and H3K27ac) states (7–9). During development and in terminally differentiated cells, enhancers can switch between states. Upon embryonic stem cell differentiation, for example, poised enhancers lose the repressive H3K27me3 mark and gain H3K27ac. The enhancer state change occurs concomitantly with a switch in the expression of the target gene from off to on and the transition in cell state from undifferentiated to differentiated. Enhancer activation is a step-wise process mediated by a variety of sequence-specific TFs chromatin-remodeling complexes and coactivators (10). RNA Polymerase II (Pol II) is also involved, directly binding to enhancer sequences and transcribing them into enhancer RNA (eRNA) (11). p53 may also participate in eRNA production (12). eRNA levels correlate with enhancer activity, and eRNA can be used as a marker for genome-wide identification of active enhancer elements (13). Enhancer transcription can lead to the formation of R-loops (14), which, if left uncorrected, can induce replication stress, genomic instability and other hallmark features of cancer (15). Figure 1. View largeDownload slide Features of active enhancer elements (see text). Transcription factor (TF), mediator (Med), RNA polymerase II (RNA Pol II), bromodomain-containing protein 4 (BRD4), CCCTC-binding factor (CTCF), enhancer RNA (eRNA), lysine methyltransferase 2C (KMT2C), lysine methyltransferase 2D (KMT2D). Figure 1. View largeDownload slide Features of active enhancer elements (see text). Transcription factor (TF), mediator (Med), RNA polymerase II (RNA Pol II), bromodomain-containing protein 4 (BRD4), CCCTC-binding factor (CTCF), enhancer RNA (eRNA), lysine methyltransferase 2C (KMT2C), lysine methyltransferase 2D (KMT2D). Enhancers communicate with genes through physical interactions with their promoters, mediated by Mediator and Cohesin complexes (16) as well as ZNF143 (17) and YY1 (18). The histone methyltransferases (MLL3/4 or KMT2C/B), and the canonical H3K4me1 enhancer mark these enzymes deposit, are also necessary for establishing enhancer–promoter interactions (19). Most enhancer–gene interactions are contained within topologically associated domains (TADs) (20), self-interacting megabase-size genomic regions bordered by the insulator protein CTCF. Within TADs, CTCF can contribute to formation of insulated neighborhoods that further seclude enhancers and their gene targets (21). Enhancers can be located close to or several hundred kilobases away from their gene targets and they do not necessarily regulate their nearest gene. Multiple enhancers often regulate a single gene, and multiple genes can be regulated by a single enhancer. Super Enhancers Super enhancers (SEs) and stretch enhancers are clusters of enhancers found across diverse cell types (22–24). SEs contain supranormal levels of the active enhancer histone mark H3K27ac and many other enhancer-associated proteins, like Mediator and Pol II. They are typically several kilobases in length, exquisitely cell-type specific, and regulate genes that control cell state. In embryonic stem cells for example, SEs regulate master TFs of pluripotency, Oct4, Sox2 and Nanog (23). Tumor cells often acquire SEs near oncogenes that are known tumor dependency genes, like MYC (25,26). They are also highly enriched for Genome-wide association studies (GWAS)-identified SNPs that predispose to several common diseases. The constituents of SEs physically interact with one another and their gene targets in 3D chromatin and these interactions are likely essential for maintenance of SE function (27). Recent studies have exploited CRISPR-Cas9 editing technologies to dissect SEs and quantify the functional contribution of individual SE constituents to their target gene levels. These studies have consistently shown that SE constituents function independently and all constituents participate in the regulation of their target genes, with some enhancers contributing more than others. There appears to be no common regulatory modality; some act additively like those at the α-globin locus (28), and others work synergistically, like those at the mammary Wap locus (29). Common disease-associated variants in SE constituents can also influence the expression of their target genes via different modalities (30). Recently, a global analysis of SE function found that while all enhancers are necessary for maximal SE activity, there exists a set of ‘hub enhancers’ whose knockout results in particularly profound decrease in SE activity (31). Hub enhancers are enriched for CTCF and cohesin and are critical for maintaining physical interactions between SE constituents as well as other enhancers within the same TAD. A recently proposed ‘phase separation’ model of transcriptional control may explain how SEs promote consistent and high level transcription. In this model, binding of TFs and other regulators to SE constituents stimulates aggregate formation and their subsequent compartmentalization within the cell nucleus (32). Mutations in Enhancer Regulators Several Mendelian diseases are caused by mutations in genes encoding proteins that function to establish, maintain, or interact with enhancers (Table 1). Most are congenital, multiple malformation diseases caused by heterozygous mutations that arise de novo, implicating haploinsufficiency as the pathogenic mechanism. A key player in enhancer activation is the ATP-dependent BRG1/BRM associated factor complex, or BAF, which slides or evicts nucleosomes to make way for TFs and other coactivators. BAF also associates with H3K4me1 at enhancers (33). Mutations in several subunits of the BAF complex cause Coffin-Siris syndrome, a condition characterized by developmental delays and unique skeletal malformations (34,35). CHARGE syndrome (coloboma, heart defects, atresia choanae, growth retardation, genital abnormalities, and ear abnormalities) is another disease caused by mutation of a chromatin remodeler that acts at enhancers, CHD7 (36). A related CHD family member, CHD8, is implicated in autism (37) and may also function in enhancer activation (38). Table 1. Diseases caused by mutations in proteins that function in establishing, maintaining, or regulating enhancers Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  Table 1. Diseases caused by mutations in proteins that function in establishing, maintaining, or regulating enhancers Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  Disease  Gene (protein) implicated  Function  Reference  Kabuki syndrome  KMT2D (KMT2D)  Histone methyltransferase that deposits H3K4me1 on histones adjacent to enhancers.  (39)  KDM6A (KDM6A)  Histone demethylase that removes H3K27me3 mark.  (40)  Rubinstein-Taybi syndrome  EP300 (P300)  Histone acetyltransferase and enhancer coactivator.  (71,72)  CREBBP (CBP)  Histone acetyltransferase and enhancer coactivator.  (41,73)  CHARGE syndrome  CHD7 (CHD7)  ATP-dependent chromatin remodeler that functions at enhancers.  (74,75)  Coffin-Siris syndrome  ARID1A (ARI1A)  Subunits of the BAF (SWI/SNF), chromatin remodeling complex that function in establishing enhancers.  (35,76)  ARID1B (ARI1B)  SMARCA4 (BRG1)  SMARCB1 (SMARCB1)  SMARCE1 (SMARCE1)  Schwannomatosis  SMARCB1 (SMARCB1)  See above.  (77)  Potocki-Shaffer syndrome  PHF21A (PF21A)  Component of BRAF-histone deacetylase complex.  (78)  Cornelia de Lange-like syndrome  BRD4 (BRD4)  BRD4 binds to H3K27ac and mediates RNA polymerase II pause-release.  (48)  The switching of enhancers between primed and active states is mediated in part by enzymes that deposit or remove covalent histone modifications, or ‘writers’ and ‘erasers’. An illustrative example of the crucial role of these proteins in normal development is Kabuki syndrome (KS), characterized by distinctive facies, intellectual disability, unique skeletal abnormalities and growth delay. KS is caused by mutations in KMT2D, a histone methyltransferase that writes the canonical H3K4me1 enhancer mark on chromatin (39). KS can also be caused by mutation of KDM6A, a histone demethylase that erases the H3K27me3 mark (40). P300 and CBP, coactivators that bind acetylated lysines at active enhancers are mutated in Rubinstein-Taybi syndrome, characterized by short stature, intellectual disability, distinctive facies and broad thumbs and toes (41). Several of the same genes with enhancer-related functions are frequently somatically mutated in cancer. Mutations in genes encoding subunits of the BAF complex occur in 20% of all human cancers (42). KMT2D, KDM6A, ARID1A, KTM2C, P300, CREBBP and KMT2A, are all constituents of machinery cells use to establish enhancers and are among the most frequently mutated genes in bladder cancer (43). Mutations in enhancer-associated histone variant H3.3 genes have been implicated in pediatric brain cancers (44), although the exact mechanism by which the oncohistone promotes tumorigenesis remains unresolved. Although not generally considered ‘enhanceropathies’ like the diseases described above, Cornelia de Lange syndrome (CdLS), Roberts syndrome, and other ‘cohesinopathies’ are caused by mutations in various components of the cohesin complex (SCC2/NIPBL, SMC1A, SMC3, RAD21, ESCO2 and others) that function in chromatin looping and hence play an important role in facilitating enhancer-promoter interactions (45–47). Moreover, mutations in BRD4 were recently shown to cause a Cornelia de Lange-like syndrome (48). BRD4 binds to SEs and also engages NIPBL, which is required for loading the cohesin looping complex onto DNA (49). The proposed model is that BRD4 mutation disrupts looping and transcription at SE-associated loci and as such, the authors noted that CdLS can be considered a “transcriptomopathy”. Mutations Disrupting Enhancer–Gene Control There are a variety of ways in which genetic mutation of non-coding regions can disrupt enhancer–gene regulation and cause disease (Table 2). Deletions can result in either gene silencing by removing active enhancers, or gene activation by repositioning them closer to enhancers. Deletions can also remove TAD boundaries, allowing for ectopic interactions between enhancers and promoters normally insulated from one another in adjacent TADs. In a variant of bracydactyly, for example, deletions removing TAD boundaries creates ectopic interactions between the EPHA4 TAD domain and PAX3 (50). Insertions may create novel TF binding sites at enhancer sequences (51). Translocations can breakup SEs, or displace enhancers from their endogenous targets and relocate them to other chromosomes (52). Inversions can force a promoter to adopt a repositioned enhancer, resulting in increased gene transcript output (53). Alternatively, inversions can reposition an enhancer to be out of range of its promoter (54). Duplications can create ectopic enhancer sequences that can act to increase target gene transcription (55). Point mutations can modulate TF affinity at enhancer sequences (56). Virtually all of these types of non-coding mutations can arise somatically and have been implicated in oncogenesis. For example, a translocation that repositions immunoglobulin heavy chain enhancers near the MYC oncogene is a common driver mutation in Burkitt’s lymphoma (57). In a variety of epithelial cancers, copy number gains containing SEs are known to activate oncogenes (58). In T-ALL, driver indel mutations at the TAL1 locus can create binding sites for MYB, a pioneer transcription factor that engages the indel, instigating SE formation and TAL1 activation (59). Microdeletions can disrupt boundaries of insulated neighborhoods, leading to oncogene activation (60). Table 2. Various types of mutations that impact enhancer function and their associated diseases Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  Table 2. Various types of mutations that impact enhancer function and their associated diseases Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  Mutation type  Disease  Enhancer defects  References  Insertion/deletion      X-linked deafness type 3 (DFN3)  Multiple deletions 900 kb from POU3F4.  (79)  Pierre Robin sequence  Deletions 1–1.5 Mb upstream and downstream of Sox9 on chromosome 17.  (52)  Split-hand-split food malformation (SHFM)  7q21.3 deletion affecting enhancer sequences within DYNC1I1.  (80,81)  Craniosynostosis  Multiple deletions upstream of Indian hedgehog (IHH).  (82)  Autosomal dominant adult-onset demyelinating leukodystrophy  Deletion eliminating TAD, allowing for enhancer adoption of LMNB1.  (83)  Axenfeld-Rieger syndrome  Deletion of up to 1.1 Mb upstream of PITX2.  (84)  Syndromic unspecified intellectual disability  Microdeletions in STAG1, which makes up the core cohesin complex.  (85)  F-syndrome  Deletions removing TAD and allowing aberrant interaction between EPHA4 and WNT6.  (50)  Brachydactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and PAX3.  (50)  Polysyndactyly  Deletions removing TAD and allowing aberrant interaction between EPHA4 and IHH  (50)  Preaxial polydactyly  13 bp insertion in the zone of polarizing activity regulatory sequence (ZRS) affecting sonic hedgehog (SHH) expression.  (51)  Translocation      Aniridia  Involves 11p13, downstream of PAX6.  (86)  Pierre Robin sequence  1 Mb away from SOX9. Abrogates binding of MSX1 in in vitro studies.  (52)  Split-hand syndrome  t(2; 7)(p25.1; q22), separates limb enhancers in DYNC1I1 from DLX5/6.  (81)  Inversion      Limb syndactyly  Enhancer adoption by SHH induced by a 7q inversion.  (53)  Hand-foot-genital syndrome  Chromosome 7 inversion causing a HOXA13 enhancer delocalization.  (54)  Duplication      Disorders of sex development (DSD)  16p13.3 duplication of GNG13 and SOX8 enhancers.  (87)  600 kb upstream of SOX9.19  (55)  Haas-type polysyndactyly and Laurin-Sandrow syndrome  Microduplications in SHH limb enhancer ZRS.  (88)  Syndactyly  Multiple targets upstream of IHH.  (82)  Keratolytic Winter Erythema  Duplication of enhancer upstream of CTSB.  (89)  Preaxial polydactyly  Microduplications in 7q36.3 affecting ZRS.  (90)  Point mutation      Aniridia  De novo mutation 150 kb downstream of PAX6 disrupting PAX6 autoregulation.  (91)  Holoproscencephaly  460 kb upstream of SHH resulting in loss of SHH brain enhancer-2 activity.  (92)  Pancreatic agenesis  25 kb downstream of PTF1A.  (56)  Van der Would syndrome  Mutation in IRF6 enhancer, abrogating p63 and E47 binding.  (93)  Hirschsprung disease  Common non-coding variant within an enhancer-like sequence in RET intron 1.  (94)  Isolated congenital heart defects  Mutation 90 kb dowstream of TBX5.  (95)  Preaxial polydactyly  Various point mutations in the ZRS enhancer (e.g. 295 T>C)  (96,97)  GWAS SNPs and Enhancers GWAS have successfully identified loci associated with several human traits and susceptibility to numerous common diseases, including heart disease, type 2 diabetes (T2D), multiple sclerosis and cancer. Sixty to eighty percent of the associated SNPs map to enhancers and/or SEs (61). There is also a strong correlation between associated SNPs and enhancers active specifically in tissues involved in disease pathology. SNPs associated with T2D for example, often reside within enhancers active exclusively in pancreatic islet cells. SNPs associated with risk for multiple sclerosis, generally considered to be an immune-related disease, lie in enhancers active in immune-related cell types. These findings offer mechanistic insights into the basis of common diseases and provide a potential explanation for why some tissues are affected while others are spared. The prevailing model is that the disease-associated SNPs impact gene regulation by modulating TF binding in cell types where the enhancer is active. There is compelling support for this model. Increased risk for melanoma is conferred by an enhancer indel that increases the affinity of RECQL, leading to increased PARP1 expression and contributing to melanogenesis (62). A risk SNP for prostate cancer increases RFX6 expression by increasing binding of HOXB13 at the RFX6 enhancer (63). SNPs associated with fetal hemoglobin levels map to BLC11A enhancers in human erythroblasts and modulate GATA1 and TAL1 binding (64). Mechanistic studies of GWAS–enhancer SNPs have also highlighted the complexity of enhancer–gene interactions in 3D chromatin. For example, an obesity associated enhancer variant within an intron of FTO, presumed to influence FTO expression, physically interacts with and impacts the expression of an altogether different gene, IRX3, located megabases away (65). Moreover, more than one gene may be targeted. The T2D risk SNP rs7163757 resides within a B-islet cell-specific stretch enhancer and modulates expression of two inflammatory cytokines, C2CD4B and C2CD4A (66). The targets of enhancer variants may not necessarily be protein coding genes, but can also be non-coding genes including miRNAs and lincRNAs. Investigations of GWAS enhancer SNPs have also helped nominate new cell types in common disease pathophysiology. SNPs associated with Alzheimer’s Disease risk correlate not only with fetal brain-specific enhancers as may have been expected, but also immune cell-specific enhancers, thereby implicating immune-related processes in AD predisposition (67). The success of the above mentioned studies suggests that the veritable gap between SNP association and function is steadily closing. Although successful in pinpointing functional variants at GWAS loci, often these studies do not rule out other variants in the region that could also be contributing to the correlation between genotype and target gene expression. There may be more than one casual SNP, particularly at loci where multiple disease-associated SNPs lie in SE constituents that physically interact with one another and their gene targets in 3D chromatin. As recently demonstrated, the SNPs at SE-associated loci may collude to influence target gene expression and the clinical risk to disease, and the influential SNPs may not necessarily be in linkage disequilibrium with one another (30,68). Furthermore, given that enhancers respond to specific physiological, pathological or environmental conditions, the regulatory effects of enhancer SNPs may be context specific (69). While considerable progress has been made, there is a clear need for functional studies that go beyond the standard eQTL and reporter assays to nominate causal variants. Strategies in which risk SNPs are directly edited in the genome are needed. Concluding Remarks As highlighted above, a variety of Mendelian diseases and cancers are caused by mutations in genes encoding proteins that function in trans in global transcriptional control either through regulating enhancer activity or enhancer–gene interactions via chromatin looping. These have been termed enhanceropathies and cohesinopathies, but given that the true pathology lies in transcription, ‘transcriptomopathies’ might be more precise descriptor for these diseases. Because clinical sequencing has largely focused on the exome, the number of genetic diseases due to non-coding mutations that disrupt enhancer–gene control in cis is not yet known. Given the sizeable percentage of Mendelian diseases currently explained by mutations in the exome (30%), the proportion of diseases due to mutations in non-coding regions is probably much smaller, even in spite of the fact that the enhancerome is 5–10× larger than the exome. Consistent with this prediction, whole genome sequence analysis of a large cohort of patients with developmental delay found that 1–3% without a diagnostic coding mutation had de novo mutations in regulatory elements (compared to 42% in the exome) (70). However, with respect to risk to common diseases, the proportion of causal variants in noncoding regions likely dwarfs that of the exome. Similarly, in cancer, and in particular cancers with high mutation rates, excessive aneuploidy and other structural rearrangements, functional mutations in non-coding regions are likely to be fairly common. Whole genome sequence analysis of tumor genomes paired with epigenome analysis should help shed light on the prevalence of noncoding cancer driver mutations. These studies are essential for the full potential of targeted therapies and precision medicine to be realized. Acknowledgements We thank Zachary Faber and Andrew Morton for critically reading the manuscript and their insightful comments. 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Human Molecular GeneticsOxford University Press

Published: May 2, 2018

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