Next Article in Journal
Umbilical Cord Mesenchymal Stromal/Stem Cells and Their Interplay with Th-17 Cell Response Pathway
Next Article in Special Issue
The Role of Fatty Acid Synthase in the Vascular Smooth Muscle Cell to Foam Cell Transition
Previous Article in Journal
Housekeeping Gene Stability in Adipose Mesenchymal Stromal Cells Cultivated in Serum/Xeno-Free Media for Osteoarthritis
Previous Article in Special Issue
AKT Mediates Adiponectin-Dependent Regulation of VSMC Phenotype
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Cells
Volume 13
Issue 2
10.3390/cells13020168
cells-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

SWI/SNF Complex in Vascular Smooth Muscle Cells and Its Implications in Cardiovascular Pathologies

by
Hongyu Liu
1,2,3,
Yang Zhao
1,
Guizhen Zhao
1,
Yongjie Deng
1,
Y. Eugene Chen
1,4,* and
Jifeng Zhang
1,*
1
Department of Internal Medicine, Cardiovascular Center, University of Michigan Medical Center, 2800 Plymouth Road, Ann Arbor, MI 48109, USA
2
Department of Molecular & Integrative Physiology, University of Michigan Medical Center, Ann Arbor, MI 48109, USA
3
Department of Vascular Surgery, The Second Xiangya Hospital, Central South University, Changsha 410011, China
4
Department of Cardiac Surgery, University of Michigan Medical Center, Ann Arbor, MI 48109, USA
*
Authors to whom correspondence should be addressed.
Cells 2024, 13(2), 168; https://doi.org/10.3390/cells13020168
Submission received: 7 November 2023 / Revised: 9 January 2024 / Accepted: 12 January 2024 / Published: 16 January 2024
(This article belongs to the Special Issue Role of Vascular Smooth Muscle Cells in Cardiovascular Disease)
Browse Figures
Review Reports Versions Notes

Abstract

:
Mature vascular smooth muscle cells (VSMC) exhibit a remarkable degree of plasticity, a characteristic that has intrigued cardiovascular researchers for decades. Recently, it has become increasingly evident that the chromatin remodeler SWItch/Sucrose Non-Fermentable (SWI/SNF) complex plays a pivotal role in orchestrating chromatin conformation, which is critical for gene regulation. In this review, we provide a summary of research related to the involvement of the SWI/SNF complexes in VSMC and cardiovascular diseases (CVD), integrating these discoveries into the current landscape of epigenetic and transcriptional regulation in VSMC. These novel discoveries shed light on our understanding of VSMC biology and pave the way for developing innovative therapeutic strategies in CVD treatment.

1. Introduction

Vascular smooth muscle cells (VSMCs) reside in the tunica media of vessels [1] and play critical roles in both vascular homeostasis and the pathogenesis of cardiovascular diseases (CVD). As early as the 1960s, it has been established that VSMC are enriched in atherosclerotic lesions in human specimens [2] and animal models [3,4]. The plasticity of VSMC has been identified as one of the major pathological factors in atherogenesis [5,6]. Key transcription factors and cofactors, like serum response factor (SRF) [7], myocardin [8], and Krüppel-like factor 4 (KLF4) [9], were identified during the investigation of SMC plasticity. Despite their essential role in DNA–protein interactions, the importance of chromatin remodelers in VSMC plasticity has often been overlooked. Summarizing the known and unknown of chromatin remodeling in VSMC research offers valuable insights into this understudied aspect. This review delves into the role of a primary ATP-dependent chromatin remodeler, the SWItch/Sucrose Non-Fermentable (SWI/SNF) complexes, in VSMC biology and its implications for cardiovascular diseases.

2. Roles of VSMCs from Physiology to Pathology

Like skeletal muscle cells and cardiomyocytes, the primary function of VSMCs is contraction. Under precise hormonal and neural control, VSMCs regulate blood distribution and blood pressure. To maintain the contractile phenotype, VSMC undergoes differentiation via the expression of a repertoire of contractile apparatus and regulators [10]. Myosin heavy chain 11 (MYH11), also known as smooth muscle myosin heavy chain (SMMHC), is responsible for encoding SMC-specific myosin. Different from myosins in other muscle tissues, MYH11 lacks intrinsic ATPase activity. Instead, its activity is triggered when Ser 19 on the regulatory myosin light chain becomes phosphorylated. This phosphorylation is precisely controlled by myosin light chain kinase (MYLK) and myosin light chain phosphatase (MLCP). Thin myofilaments are composed of α-actin, also known as α-smooth muscle actin (α-SMA), which is encoded by actin alpha 2 (ACTA2) [11,12]. Of note, variants of those genes have already been well-documented in patients with inherited aortic diseases [13,14]. This underscores the importance of fully functional contractile machinery in VSMCs for maintaining aortic health.
Another important role of VSMCs is to regulate vascular extracellular matrix (ECM) homeostasis through the production of components including elastin, collagen [15], and proteoglycans and regulation of ECM remodelers including matrix metalloproteinase (MMP) and tissue inhibitor of matrix metalloproteinases (TIMPs). These determine the vessel wall’s mechanical strength, compliance, and elastic recoil. As a result, ECM construction and maintenance of VSMC have emerged as central priorities in the field of bioengineering for blood vessel grafts [16].
Interestingly, the two major roles of VSMCs seem to be conflicting. During the initial embryonic development stage, VSMCs exhibit rapid proliferation [17], migrate, and actively secrete ECM components crucial for vasculogenesis. However, mature VSMCs lose those properties and become quiescent with a low proliferation rate and synthetic activity. In the mature aorta, VSMCs possess contractile apparatus and maintain mechanical strength. Nevertheless, during vascular injury, alterations in the environmental cues cause the loss of contractile markers, including MYH11 and α-SMA, coupled with increased proliferation, migration, and protein synthesis in VSMC. This type of dedifferentiated VSMC is essential for vascular repair. The transition between the contractile and synthetic states of VSMCs is termed “phenotypic switch”. It is thus logical to infer that the intrinsic plasticity of VSMCs is beneficial for adaption to complex environments and response to damage. However, substantial changes in diet and lifestyle may hijack this ability and disrupt vascular homeostasis.

3. Epigenetic and Transcriptional Regulation of VSMC Plasticity in Health and Diseases

Vascular smooth muscle cell plasticity in CVD, including atherosclerosis, has been well-documented [2]. Especially in the past decade, the emergence of new technologies, such as lineage tracing, single-cell RNA sequencing (scRNA-Seq), and spatial transcriptomics [18,19,20,21], has largely extended our knowledge of cell origin, characteristics, and transcriptomic profiles in the atherosclerotic lesions. Under pathological conditions, VSMCs transdifferentiate into various cell types such as foam cells [22,23], mesenchymal-stem-cell (MSC)-like cells [24], macrophage-like cells [25,26], adipocyte-like cells [27], osteochondrogenic cells [28,29], fibromyocytes [19]. Multiple reviews thoroughly discuss this phenomenon [5,30,31,32,33,34]. A dramatic shift in gene expression occurs during dedifferentiation or transdifferentiation. Such changes necessitate a sophisticated transcriptional regulation network in VSMCs to precisely control specific gene expression in response to environmental changes [34,35,36].
In differentiated VSMC, the expressions of contractile genes are governed by an array of transcription factors and coactivators [35,36,37] (Table 1; Figure 1 and Figure 2). In 1985, CC(A/T)6GG, also referred to as the CArG element, was identified in the promoter of the human cardiac actin gene [38]. Subsequent research has highlighted the crucial role of the CArG element in regulating muscle-specific genes [39]. Serum response factor (SRF) was then identified to bind with CArG elements and regulate downstream gene expression [7,40]. In addition, the discovery of myocardin [8] and its competition with Elk-1 for SRF interaction [41] revealed the delicate regulation of the VSMC phenotype.
Aside from myocardin/SRF, other transcription factors also play essential roles in regulating contractile genes [35]. The roles of transforming growth factor-β (TGFβ) in VSMC differentiation have been documented [79]. In 1997, the TGFβ control element (TCE), proximal to two CArG elements, was discovered in the promoter of ACTA2 [80]. TGFβ increases contractile gene expression by facilitating the binding of SRF to the CArG elements, possibly through interactions between Smad3 and p300 [81]. Subsequent research on the synergetic function and direct interaction of Smad3 and myocardin has further elucidates the roles of TGFβ signaling in VSMC differentiation [48]. Another transcription factor, GATA binding protein 6 (GATA6), was found to be highly expressed in the VSMCs during development [82], and it was shown to protect against injury-induced VSMC phenotypic switch [50]. Further study has revealed that GATA6, NK3 homeobox 2 (NKX3-2), and SRF form a triad complex to regulate contractile gene expressions [52].
Conversely, in dedifferentiated VSMC, KLF4 plays an essential role [83]. The significance of KLF4 in VSMC was first discovered through yeast one-hybrid cloning against TCE [84]. Subsequent studies have firmly demonstrated that KLF4 was a potent repressor for VSMC differentiation [62,85]. In response to PDGF-BB treatment, Sp1, pELK1, and KLF4 cooperatively bind to the G/C repressor element flanked by two CArG elements in the promoters of contractile genes, including MYH11 and TAGLN. In addition, KLF4 recruits HDACs, reducing histone acetylation in the promoters. Consequently, it diminishes SRF binding and the expression of contractile genes [86]. In the mouse atherosclerosis model, SMC-specific KLF4 knockout showed less mesenchymal-stem-cell- and macrophage-like cells derived from SMC, along with less lesion and increased fibrous cap thickness, underscoring significant roles of KLF4 in the cardiovascular diseases [9].
As we delve deeper into the research on the transcriptional regulation of contractile genes, epigenetic regulation gains increasing interest [87,88]. Structural investigations revealed that SRF does not bind to nucleosomal DNA [89,90,91], implying its exclusive binding to open DNA regions. Meanwhile, several factors affect the binding activity of SRF to the CArG elements [80,86,88]. These findings shed light on the significance of chromatin conformation in the transcriptional regulation of contractile genes. McDonald et al. discovered that H3K4me2, H3K79me2, H3K9Ac, and H4Ac are enriched in the CArG elements in the SMC but not in the non-SMC [86]. In addition, H3K4me2 tethers with SRF and myocardin, and this modification persists even after VSMC dedifferentiation [92]. Loss of H3K4me2 at the CArG elements causes a decrease in contractile gene expression through the reduced TET2 (ten-eleven translocation-2)-mediated DNA demethylation [77,93]. In contrast, in embryonic stem cells or other non-SMCs, H3K9me3 and H3K27me3 govern the repression of contractile gene expression [91]. Lysine demethylase 3A (KDM3A), previously known as JMJD1a, interacts with myocardin and demethylates H3K9me3, thus promoting the expression of contractile genes [69]. In addition, numerous histone modifiers have been identified to regulate contractile genes, including PR/SET domain 6 (PRDM6), SET and MYND domain containing 2 (SMYD2), SUV39H1, polycomb repressive complexes 2 (PRC2), and protein arginine methyltransferase 5 (PRMT5) (Table 1; Figure 1 and Figure 2) [68,71,72,73,74,75,78]. Although previous studies elucidated the intricate regulation of transcription and histone modification, the specific roles of chromatin remodelers in directly executing chromatin transformations require further investigation.

4. Chromatin Remodeling and SWI/SNF Complexes

In eukaryotic cells, DNA wraps around histones, forming the basic structural unit: a nucleosome [94]. In a rigid and delicate manner, DNA accessibility is highly regulated for complex activities, including replication, repair, and transcription. Chromatin remodeling involves altering interactions between histones and DNA, including assembly and disorganization of nucleosomes. Transcription requires DNA free from histones to interact with proteins such as polymerase II and transcription factors. Studies have shown that chromatin remodelers are essential in controlling pluripotency, cell fate, and differentiation [95,96,97]. Four prominent remodeler families have spiked the most study interest to date, including SWI/SNF, Imitation switch (ISWI), chromodomain helicase DNA-binding (CHD), and INOsitol requiring 80/SWI2/SNF2-Related 1 (INO80/SWR1) [98,99]. This review focuses on the SWI/SNF complexes.
The SWI/SNF chromatin remodeling complexes were first discovered in Saccharomyces cerevisiae. SWI stands for ‘switch’ as the relevant genes regulate the HO gene, which is crucial for mating type switching [100]. Similarly, SNF denotes ‘sucrose nonfermenting’ since these genes control the SUC2 gene responsible for sucrose catabolism [101]. Subsequent research has found that some genes from these two screenings overlap, forming a complex that regulates chromatin structure [102]. SWI/SNF complexes use energy from ATP hydrolysis to mediate nucleosome sliding or ejection [103]. The mammalian SWI/SNF complex was purified in 1996 [104]. In mammals, either Brg1 or Brm serves as the ATPase of the complex; the SWI/SNF complex is also named the BRG1/BRM-associated factor (BAF). The initially isolated complex was termed canonical BAF (cBAF) [104]. Subsequent research has identified two more types of SWI/SNF assemblies: polybromo-associated BAF (PBAF) [105] and non-canonical BAF (ncBAF) [106,107,108]. These complexes are assembled by 9–16 subunits, and some subunits consist of several paralogs (Figure 3 and Table 2) [109]. Each subunit conveys unique functions for chromatin remodeling [110]. More than 1000 unique combinations of SWI/SNF subunits form diverse complexes with varied functions [111,112].
Recent advancements have significantly illuminated the structure of SWI/SNF complexes [113,114,115,116]. These findings further deepen our understanding of the chromatin remodeling mechanism of SWI/SNF complexes and the function of their subunits. According to Chen et al., SWI/SNF complexes can be divided into three modules: the motor module, actin-related protein (ARP) module, and substrate recruitment module (SRM) [117]. SRM can be further divided into nucleosome binding lobe (NBL), DNA binding lobe (DBL), and histone-tail binding lobe (HBL) [117]. The central motor module is the ATPase, SMARCA4 (BRG1), and SMARCA2 (BRM), which directly hydrolyze ATP to translocate DNA [118]. The ARP module comprises actin beta (ACTB), BCL7, and actin-like 6 (ACTL6). They serve the dual function of connecting ATPase with SRM and regulating the ATPase activity. Within the nucleosome binding lobe, SMARCB1, coupled with double PHD fingers (DPF) and SMARCC, attaches directly to the nucleosome acidic patch via the C-terminal domain (CTD) [119]. The DNA binding lobe interacts with extranucleosomal linker DNA and various factors. This lobe consists of several DNA binding domains like the HMG-box domain on SMARCE1 and polybromo 1 (PBRM1) [120], AT-rich interaction domain (ARID) on ARID1A/B, and ARID2 [121].
Of note, multiple interactors of SWI/SNF complexes exist to orchestrate gene transcription (some interactions validated by the GST pull-down assay are listed in Table 3). The SMARCD family, including SMARCD1, SMARCD2, and SMARCD3, are reported to mediate the interactions between the SWI/SNF complex and various factors, such as PPARG coactivator 1α (PGC1α) and CCAAT enhancer binding protein ε (CEBPε) [122,123]. Recently, Wolf et al. [124] fused TurboID [125] with SMARCD1 and revealed close associations of the SWI/SNF complex with various transcription factors and epigenetic machinery, including lysine-specific methyltransferase 2 (KMT2) family, nuclear receptor coactivator (NCOA) family and histone acetyltransferases (HATs). The SWI/SNF complex can also recognize histone modifications. The bromodomains within its subunit BRD7/9, SMARCA4, SMARCA2, and PBRM1 facilitate their binding to acetylated histones [126]. The chromodomains on the subunit SMARCC1 and SMARCC2 recognize methylated H3 [127]. Given their sophisticated structure and interactors, it is speculated that the whole complexes remodel the chromatin at precise locations and accurate time points.

5. SWI/SNF Complex in Cardiovascular Development

Coffin–Siris syndromes (CSSs) represent congenital disorders predominantly linked to mutations in the subunits of SWI/SNF complexes and are characterized by mental retardation [138]. ARID1B is responsible for CSS1, with ARID1A for CSS2, SMARCB1 for CSS3, SMARCA4 for CSS4, SMARCE1 for CSS5, ARID2 for CSS6, DPF2 for CSS7, SMARCC2 for CSS8, SMARCD1 for CSS11, and BICRA for CSS12. Recently, more phenotypes in the cardiovascular system have been noticed in CSS patients [139]. Among fetuses, 67% of patients present cardiac anomalies, and 53% of patients present vascular anomalies [140]. For instance, one case study has reported that one fetus with SMARCC2 deficiency was diagnosed with tetralogy of Fallot, a rare congenital disease caused by a combination of four heart defects [141]. Considering the lethality of cardiovascular anomality and the higher possibility of termination due to poor prognosis, the actual incidence of cardiovascular complications in CSS might be underestimated.
Over decades of research on the mammalian SWI/SNF complex, numerous animal models have been developed. In the mouse model, SMARCA4 null embryo dies at day three due to the arrest of differentiation [142], while SMARCA2 knockout mice are viable [143]. This indicates that the roles of SMARCA4 and SMARCA2 differ in development. As an essential subunit in the SWI/SNF complex, DPF3 is highly expressed in the heart and skeletal muscle and serves as a histone modification reader. DPF3 possesses 2 plant homeodomains (PHDs), which facilitate its binding with acetylated H3 and H4, and H3K4me1/2. Knockdown dpf3 in the zebrafish leads to irregular cardiac morphology and muscular fiber disarray [144]. Constitutive knockout of BIRCA (BRD4 interacting chromatin remodeling complex associated protein) (also known as GLTSCR1 (glioma tumor suppressor candidate region gene 1)) results in embryonic lethality and cardiac defects, including ventricular septal defect, double outlet right ventricle and a thinner ventricular wall [145]. These results indicate the indispensable roles of the SWI/SNF complex in the development of the cardiovascular system.

6. SMARCA4 and SMARCA2 in VSMC Biology and Cardiovascular Diseases

SMARCA4 and SMARCA2 (also called Brg1 and Brm, respectively) are the two mutually exclusive ATPase subunits of the SWI/SNF complex, which hydrolyze ATP to provide energy for the complex to remodel chromatin. Emerging evidence from early cardiovascular studies illuminates the potential significance of the SMARCA family in VSMC. In 2003, Chang et al. [54] found that SMC-enriched protein cysteine and glycine-rich protein (CSRP) 1/2 bridge SRF with GATA4/5/6 to promote SMC contractile protein expression in pluripotent 10T1/2 fibroblasts, while skeletal-muscle-enriched CSRP3 inhibits this transition to VSMC. Further study in 2007 [55] found that CSRP2 overexpression in adult cardiomyocytes promotes SMC-specific marker gene expression. At VSMC marker promoters, such as MYH11 and Calponin (CNN1), an interplay of SRF, p300, and varying histone modifications was observed. Notably, SMARCA4 and SMARCB1, but not SMARCA2, were recruited, underlining the significance of the SWI/SNF complex in SMC marker gene expression in adult cardiomyocytes. Interestingly, the interaction between CSRP and the SWI/SNF complex is also captured via SMARCD1-TurboID in the abovementioned study [124].
Following Chang’s work, Herring and his colleagues conducted a series of studies investigating the roles of SMARCA4 in VSMCs. They employed a special cell line called B22 cells derived from NIH-3T3 fibroblast, which express a dominant-negative SMARCA4 (DN-SMARCA4) induced by tetracycline withdrawal [146]. In this system, the increase in contractile genes via overexpression of myocardin-related transcription factor (MRTFA) is attenuated by DN-SMARCA4. Notably, MRTFA promotes contractile protein expression in cervical cancer HeLa cells but not in adrenal carcinoma SW13 cells, as the latter lacks SMARCA4 or SMARCA2. When SMARCA4 or SMARCA2 is overexpressed in SW13, the ability of MRTFA to promote contractile protein expression is restored. Furthermore, DN-SMARCA4 led to a decrease in contractile protein expression in primary colon SMCs. Through in vitro investigations, it was proved that SMARCA4 directly binds with MRTFA but not with SRF. DN-SMARCA4 significantly interferes with SRF binding [42]. Similarly, SMARCA4 interacts with myocardin and permits its effect on contractile proteins [43]. SMARCA4 also regulates miR-143 and miR-145 expression in conjunction with myocardin or MRTFA. Only when SMARCA4 and myocardin or MRTFA are coexpressed in the SW13 cells can SRF be recruited to CArG elements in the promoter of miR-143/145 [147]. Among all the SRF targets, ACTA2 is a special case that offers profound insight, as it appears to be less susceptible to the influence of SMARCA loss. ACTA2 is already highly expressed in the B22 cells, and SRF binding at its promoters is much higher than that of MYLK and TAGLN. When MRTFA or myocardin is induced, SRF binding at the ACTA2 promoter and ACTA2 expression are increased. DN-SMARCA4 did not attenuate the effect of MRTFA or myocardin for ACTA2. This indicates a possibility that the ACTA2 promoter in B22, fibroblasts, and SMCs remains accessible for SRF binding. The loss of SMARCA4 does not close this region, and MRTFA or myocardin can still effectively promote ACTA2 expression.
Herring and his colleagues further specifically knocked out Smarca4 with or without Smarca2 in SMCs using SMMHC-Cre. About 33% of SMC-Smarca4 knockout mice demonstrate cardiopulmonary abnormalities, including defects in the cardiac outflow tract, patent ductus arteriosus (PDA), and ventricular septal defect (VSD). Those defects are coupled with cyanosis and elevated neonatal mortality rates. In addition, striking phenotypes are noticed in the gastrointestinal (GI) tract. They found that SMC-Smarca4/Smarca2 double knockout mice died within two weeks after birth, with dilated intestines. While SMC-Smarca4 knockout mice consistently showed enlarged intestines, the SMC-Smarca2 knockout mice demonstrated relatively fewer aberrations. Consistently, contractile proteins are significantly decreased in the SMC derived from surviving SMC-Smarca4 knockout and double knockout mice, including telokin, Myh11, Cnn1, Tagln, and Acta2. Moreover, SMCs exhibited increased apoptosis and decreased proliferation [148].
Additionally, compelling evidence suggests that SMARCA4 and SMARCA2 regulate VSMC proliferation and inflammation. Endothelin 1 (ET-1) enhances VSMC contraction, proliferation, and inflammation in VSMCs, playing multifaceted roles in CVD [149]. ET-1 was found to increase the expression of SMARCA4 in rat aortic smooth muscle cells, while this can be attenuated by NaHS. Overexpression of Smarca4 promotes VSMC proliferation and the expression of Ntf3, Pcna, and Pdgfα. Smarca4 binds at the promoters of those genes and modulates DNA accessibility [150]. SMARCA4 and SMARCA2 are critical for activating inflammatory genes, including IL6, IL1β, and CCL2, particularly when stimulated by ET-1 in VSMCs. A luciferase assay revealed that the binding of SMARCA4 and SMARCA2 at the promoters of those genes is increased in VSMC in response to ET-1 [151].
Additional evidence further indicates the roles of SMARCA4 in CVD. Several groups have reported an increase in SMARCA4 in the human standard type A aortic dissection specimens [152,153,154]. To mimic different types of heritable thoracic aortic aneurysm (TAA), two pathologic variants, TGFR2G357W and ACTA2R179H, were, respectively, overexpressed in human aortic VSMC. Consistent with the patient’s specimen carrying that allele, contractile genes, including smoothelin-1 (SMTN1), CNN1, vinculin (VCL1), and Tagln, are decreased, while HDAC9 is increased, especially in the nucleus. In this model, SMARCA4 is indispensable for HDAC9 and EZH2 binding at the promoters of contractile genes, which are then repressed [155]. In the ligation-induced carotid artery model, SMARCA4 is also significantly increased in the adventitial SCA1+ smooth muscle cells (AdvSca1-SM). Oral administration of SMARCA4/SMARCA2 inhibitor PFI-3 can rescue ligation-induced vascular remodeling and inflammatory cell infiltration. Perivascular administration of SMARCA4 shRNA attenuates adventitial expansion and vascular fibrosis. Under normal or TGFβ-stimulated conditions, PFI-3 decreases Myh11, Cnn1, and Acta2 expression, indicating essential roles of SMARCA4/SMARCA2. The cleavage under targets and release using nuclease (CUT&RUN) assay reveals that, in the presence of PFI-3, the binding of SMARCA4 and the level of H3K27Ac at the Acta2 promoter are reduced [156].
A prevailing controversy exists that SMARCA4 augments the expression of contractile proteins in vitro, but SMARCA4 increases with contraction protein decrease in human TAA samples. Since SMARCA4 and SMARCA2 constitute all SWI/SNF complexes, they might collaborate with diverse factors to activate and repress genes [157]. SMARCA may serve as a universal subunit for different regulations in the process of vascular diseases.

7. Roles of the SMARCD Family in VSMC

The human SMARCD family was first cloned and identified in 1996, with specific tissue distributions observed. Of note, SMARCD1 is universally distributed, SMARCD2 is predominantly expressed in the pancreas, and SMARCD3 is primarily found in the heart and skeletal muscle [158]. All three SWI/SNF complexes contain only one of the three SMARCD subunits: ncBAF only has SMARCD1, whereas cBAF and PBAF can contain any one of the three SMARCD members. SMARCD1-3 is one of the core subunits in the SWI/SNF complex. Knockout of SMARCDs disrupts the engagement of the ARID family and ATPase to the complex core [109].
The roles of the SMARCD family in myogenesis are extensively documented [159]. Smarcd3 participates in the early myogenesis in zebrafish [160]. Myoblast determination protein 1 (MyoD) stands as a pioneering transcription factor in myogenesis, initiating muscular cell differentiation, binding to closed chromatin, and aiding subsequent binding of other transcription factors. SMARCD3 and SMARCA4 interact with MyoD and bind at the promoter of myogenin, a muscle-specific transcription factor. The binding of MyoD at the promoter of Myogenin is facilitated by SMARCD3. SMARCD3 phosphorylation mediated by p38α is indispensable for SMARCA4 recruitment and muscle subsequent differentiation [161].
The roles of SMARCD3 in cardiovascular development were first reported in 2004 [162]. Bruneau and his colleague found that SMARCD3 is expressed in the early stages of cardiac development. Smarcd3 KO causes abnormal cardiac development, including shortened outflow tract, hypoplastic atrium and ventricle, and left–right asymmetry [137,162]. In zebrafish, Smarcd3 cooperates with Gata5 (a functional homolog of mammalian Gata4) and T-box transcription factor 5 (Tbx5) to promote cardiac development [163]. Cardiac defects are found in both Smarcd3 constitutive KO and cardiac-specific KO models driven by Nkx2-5 or Myh6 Cre [134].
SMARCD3 interacts with transcription factors to regulate cardiac developmental markers. Knockdown of Smarcd3 reduces natriuretic peptide A (NPPA), a cardiac differentiation marker. Additionally, NPPA–luciferase reporter is synergistically transactivated by TBX5, NKX2-5, GATA4, and SMARCD3, and this activation is blocked when SMARCA4 is depleted. Moreover, the interaction between SMARCA4 and either TBX5 or NKX2-5 is critically dependent on SMARCD3. This aligns with other studies suggesting that SMARCD3 facilitates the interaction between transcription factors and the SWI/SNF complex [162]. Transient induction of Smarcd3 along with Tbx5, Nkx2-5, and Gata4 in the mouse embryo induces ectopic expression of cardiac marker actin alpha cardiac muscle 1 (Actc1), myosin light chain 2 (Myl2), troponin T2 (TNNT2) in the mesoderm non-cardiac cells [164]. Furthermore, ectopic cardiac myocytes are found to be beating, which indicates the existence of a full contraction machinery. Notably, Gata4 and Smarcd3 interaction can induce the expression of Actc1, although the beating phenomenon is not observed. The addition of Tbx5 is required for the ectopic cardiac myocyte to beat. Moreover, Smarcd3 is essential for guiding Gata4 to its subsequent targets [164].
Although several studies have demonstrated the critical roles of SMARCD3 in developing skeletal muscle and cardiomyocytes, its contribution to VSMC biology was first reported in 2012. Sohni et al. [165] used rat multipotent adult progenitor cells (rMAPCs) as a platform. These cells exhibit the capability to differentiate into SMCs in MAPC basal medium supplemented with TGFβ1 and PDGFBB without the presence of Fetal bovine serum (FBS). During this differentiation process, the expression of Smarcd3 and Myocardin is increased, along with the increase in contractile markers, including Cnn1, Tagln, Smtn, Acta2, and Myh11. They found that Smarcd3 is indispensable for the expression of Tagln and Acta2. Moreover, the elevation of Smarcd3 induced by TGFβ1 in rMAPC can be subdued by the Smad3-specific inhibitor and TGFβ receptor inhibitor. When the Smad3 binding element (SBE) upstream of Smarcd3 is mutated, TGFβ can no longer promote Smarcd3-luciferase activity, indicating that Smarcd3 is indeed regulated by TGFβ via Smad3 signaling. Moreover, Smarcd3 can interact with Srf and CArG elements in the promoters of Acta2 and Tagln. This interaction indicates that SMARCD3 serves as a coactivator for SRF to promote the expression of contractile proteins.
Another SMARCD family member, SMARCD1, is also identified in the vascular system [166]. In primary rat VSMCs, free fatty acid (FFA, mixture of oleic acid, and palmitic acid) decreases Smarcd1 in a dose- and time-dependent manner. With gain- and loss-of-function approaches, Smarcd1 was found to promote the expression of clock genes, including Bmal1 (basic helix-loop-helix ARNT like 1), Clock, and Dbp (D-box binding PAR bZIP transcription factor). Notably, both overexpression and knockdown of Smarcd1 only modulate the basal expression levels of these clock genes without affecting their circadian patterns. RORα has been identified to bind with Smarcd1, mediating the expression of Bmal1. Overexpression of Smarcd1 results in increased levels of H3Ac and H3K4me3 and decreased H3K9me2 at the ROR-binding sites (RORE). Furthermore, overexpression of SMARCD1 inhibits FFA-induced Erk phosphorylation and prevents VSMC proliferation and migration [166]. In 2020, we found that SMARCD1 is significantly upregulated in the human abdominal aortic aneurysm (AAA) specimen and AAA tissues from the mouse model [167]. SMC-specific Smarcd1 knockout mice exhibit a dramatic decrease in AAA incidence and maximal diameter in both AAA models induced by elastase and Angiotensin II with hypercholesterolemia. Extracellular matrix (ECM) degradation and related proteases, including cathepsin S, matrix metalloproteinase-2 (MMP2), and MMP9, are significantly reduced in the Smarcd1SMKO AAA specimen. We also observed reduced infiltration of Mac2+ macrophages and decreased levels of MCP-1 and IL-6 in the serum. Knockdown and overexpression of SMARCD1 in the human aortic smooth muscle cell (HASMC) demonstrates that NF-κB downstream inflammatory genes, including TNFα, IL1α, and IL1β, are modulated by SMARCD1. ChIP-seq data showed reduced SMARCA4 binding at the TSS of NF-κB downstream genes, accompanied by decreased active histone markers, H3K9Ac and H3K27Ac. Interaction between p300 and SMARCA4 is also decreased when SMARCD1 is knocked down.
Of note, SMARCD3 exhibits the opposite effects in AAA pathogenesis in mouse models [44]. Microarray data indicate that SMARCD3 is significantly reduced in human AAA samples [168]. In the scRNAseq analysis of the mouse AAA model, Smarcd3 decreased specifically in VSMC [169]. SMC-specific Smarcd3 knockout (Smarcd3SMKO) mice showed higher AAA incidence and larger maximal diameters, accompanied by significant increases in ECM degradation, lymphocyte, and macrophage infiltration in the aortic wall [44]. Consistent with previous studies [165], SMARCD3 is essential for contractile gene expression in the VSMC. SRF and SMARCA4 binding at the promoters and H3K9Ac and H3K27Ac at the TSS of contractile genes are all regulated by SMARCD3. SMARCD3 is also required for interaction among SWI/SNF complex, SRF, and p300.
In addition, SMARCD3 knockdown increases VSMC apoptosis [44]. Knockdown of SMARCD3 decreases the expression of the anti-apoptotic gene BCL2. ChIP-seq and the ChIP assay demonstrate decreased SMARCA4, H3K9ac, and H3K27Ac binding at the KLF5 binding region within the promoter of BCL2. Knockdown of SMARCD3 also reduces KLF5 binding at the promoter and its interaction with SMARCA4. In human pulmonary artery smooth muscle cells, prostate cancer cells, and colorectal cancer cells, KLF5 was found to serve as a protector against apoptosis, which induces antiapoptotic genes, including BCL2, baculoviral IAP repeat containing 5 (BIRC5) and repress pro-apoptotic genes like BCL2 associated X (BAX) [170,171,172]. Our study uniquely highlights the pivotal role of SMARCD3 and the SWI/SNF complex in regulating VSMC apoptosis through KLF5 [44]. In alignment with our findings, SMARCD3 has been identified as a crucial factor for the survival of pancreatic ductal adenocarcinoma (PDAC) stem cells. We speculate that those cancer cells hijack the SMARCD3/KLF5 pathway to resist apoptosis [173].

8. Concluding Remarks and Future Directions

Research on VSMCs has evolved from studying physiological functions and contractile machinery to exploring transcription and epigenetic regulation. With its intricate plasticity mechanisms, this unique cell type likely holds the therapeutic potential for CVD.
The SWI/SNF complex has been identified as a target for an increasing number of drugs in cancer research [174]. For instance, Xiao et al. [175] found that androgen receptor (AR) and forkhead box A1 (FOXA1)-positive prostate cancer cells are sensitive to proteolysis-targeting chimera (PROTAC) degrader against SMARCA4 and SMARCA2. In VSMCs, however, SMARCA4 is essential for simultaneously expressing contractile proteins and inflammatory genes. Therefore, SMARCA4 and SMARCA2 are not ideal targets for cardiovascular disease treatment. Of note, the knockout of SMARCD1 in VSMC prevents inflammation and AAA formation [167]. Several studies have also highlighted the role of BRD9 in inflammation in β cells [176] and macrophages [177,178,179]. BRD9 is a unique subunit in ncBAF, while SMARCD1 is the only SMARCD family member in ncBAF. Since both BRD9 and SMARCD1 promote inflammation, we speculate that ncBAF plays an important role in inflammation in VSMC. Inhibitors targeting BRD9 or SMARCD1 hold promising potential for mitigating inflammation and addressing associated CVD.
It is crucial to comprehensively investigate the functions of specific SWI/SNF subunits or complexes in VSMC. There are three major complexes, encompassing 9–16 subunit families with over 1000 potential combinations. The diversity of the SWI/SNF complex determines the diversity of its targets and functions. As mentioned above, we have demonstrated the opposite functions of SMARCD1 and SMARCD3 in VSMC and AAA formation [44,167]. Contradicting functions between SMARCD1 and SMARCD3 have also been reported in skeletal muscle differentiation and metabolism regulation [180,181,182]. However, the role of the remaining family member, SMARCD2, in VSMC remains to be elucidated. It has been reported that SMARCD2, but not SMARCD1, partially functions as SMARCD3 in cardiomyocyte differentiation [164]. Interestingly, during in vitro differentiation of VSMC and cardiomyocyte, there is a decrease in SMARCD1 and SMARCD2, along with an increase in SMARCD3 [183,184]. It is still unknown whether SMARCD2 shares similar functions with SMARCD3 or SMARCD1 or possesses unique functions in VSMC biology. Further studies are essential for advancing our understanding of the intricate epigenetic regulation governing VSMC biology and associated vascular diseases.

Funding

This research was funded by National Institutes of Health grants, grant numbers HL153710 and HL138139 (J.Z.), HL109946 and HL162294 (Y.E.C.), HL172832 (G.Z.); grants from the American Heart Association, 936135 (G.Z.).

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Pease, D.C.; Paule, W.J. Electron microscopy of elastic arteries; the thoracic aorta of the rat. J. Ultrastruct. Res. 1960, 3, 469–483. [Google Scholar] [CrossRef] [PubMed]
  2. Geer, J.C.; McGill, H.C.; Strong, J.P. The fine structure of human atherosclerotic lesions. Am. J. Pathol. 1961, 38, 263–287. [Google Scholar] [PubMed]
  3. Still, W.J.; O’Neal, R.M. Electron microscopic study of experimental atherosclerosis in the rat. Am. J. Pathol. 1962, 40, 21–35. [Google Scholar] [PubMed]
  4. Parker, F. An Electron Microscopic Study of Experimental Atherosclerosis. Am. J. Pathol. 1960, 36, 19–53. [Google Scholar]
  5. Basatemur, G.L.; Jorgensen, H.F.; Clarke, M.C.H.; Bennett, M.R.; Mallat, Z. Vascular smooth muscle cells in atherosclerosis. Nat. Rev. Cardiol. 2019, 16, 727–744. [Google Scholar] [CrossRef] [PubMed]
  6. Bennett, M.R.; Sinha, S.; Owens, G.K. Vascular Smooth Muscle Cells in Atherosclerosis. Circ. Res. 2016, 118, 692–702. [Google Scholar] [CrossRef] [PubMed]
  7. Landerholm, T.E.; Dong, X.R.; Lu, J.; Belaguli, N.S.; Schwartz, R.J.; Majesky, M.W. A role for serum response factor in coronary smooth muscle differentiation from proepicardial cells. Development 1999, 126, 2053–2062. [Google Scholar] [CrossRef]
  8. Wang, D.; Chang, P.S.; Wang, Z.; Sutherland, L.; Richardson, J.A.; Small, E.; Krieg, P.A.; Olson, E.N. Activation of cardiac gene expression by myocardin, a transcriptional cofactor for serum response factor. Cell 2001, 105, 851–862. [Google Scholar] [CrossRef]
  9. Shankman, L.S.; Gomez, D.; Cherepanova, O.A.; Salmon, M.; Alencar, G.F.; Haskins, R.M.; Swiatlowska, P.; Newman, A.A.; Greene, E.S.; Straub, A.C.; et al. KLF4-dependent phenotypic modulation of smooth muscle cells has a key role in atherosclerotic plaque pathogenesis. Nat. Med. 2015, 21, 628–637. [Google Scholar] [CrossRef]
  10. Owens, G.K. Regulation of differentiation of vascular smooth muscle cells. Physiol. Rev. 1995, 75, 487–517. [Google Scholar] [CrossRef]
  11. Brozovich, F.V.; Nicholson, C.J.; Degen, C.V.; Gao, Y.Z.; Aggarwal, M.; Morgan, K.G. Mechanisms of Vascular Smooth Muscle Contraction and the Basis for Pharmacologic Treatment of Smooth Muscle Disorders. Pharmacol. Rev. 2016, 68, 476–532. [Google Scholar] [CrossRef] [PubMed]
  12. Milewicz, D.M.; Trybus, K.M.; Guo, D.C.; Sweeney, H.L.; Regalado, E.; Kamm, K.; Stull, J.T. Altered Smooth Muscle Cell Force Generation as a Driver of Thoracic Aortic Aneurysms and Dissections. Arterioscler. Thromb. Vasc. Biol. 2017, 37, 26–34. [Google Scholar] [CrossRef] [PubMed]
  13. Isselbacher, E.M.; Lino Cardenas, C.L.; Lindsay, M.E. Hereditary Influence in Thoracic Aortic Aneurysm and Dissection. Circulation 2016, 133, 2516–2528. [Google Scholar] [CrossRef] [PubMed]
  14. Pinard, A.; Jones, G.T.; Milewicz, D.M. Genetics of Thoracic and Abdominal Aortic Diseases. Circ. Res. 2019, 124, 588–606. [Google Scholar] [CrossRef]
  15. Sibinga, N.E.; Foster, L.C.; Hsieh, C.M.; Perrella, M.A.; Lee, W.S.; Endege, W.O.; Sage, E.H.; Lee, M.E.; Haber, E. Collagen VIII is expressed by vascular smooth muscle cells in response to vascular injury. Circ. Res. 1997, 80, 532–541. [Google Scholar] [CrossRef]
  16. Niklason, L.E.; Lawson, J.H. Bioengineered human blood vessels. Science 2020, 370, eaaw8682. [Google Scholar] [CrossRef]
  17. Cook, C.L.; Weiser, M.C.; Schwartz, P.E.; Jones, C.L.; Majack, R.A. Developmentally timed expression of an embryonic growth phenotype in vascular smooth muscle cells. Circ. Res. 1994, 74, 189–196. [Google Scholar] [CrossRef]
  18. Pan, H.; Xue, C.; Auerbach, B.J.; Fan, J.; Bashore, A.C.; Cui, J.; Yang, D.Y.; Trignano, S.B.; Liu, W.; Shi, J.; et al. Single-Cell Genomics Reveals a Novel Cell State During Smooth Muscle Cell Phenotypic Switching and Potential Therapeutic Targets for Atherosclerosis in Mouse and Human. Circulation 2020, 142, 2060–2075. [Google Scholar] [CrossRef]
  19. Wirka, R.C.; Wagh, D.; Paik, D.T.; Pjanic, M.; Nguyen, T.; Miller, C.L.; Kundu, R.; Nagao, M.; Coller, J.; Koyano, T.K.; et al. Atheroprotective roles of smooth muscle cell phenotypic modulation and the TCF21 disease gene as revealed by single-cell analysis. Nat. Med. 2019, 25, 1280–1289. [Google Scholar] [CrossRef]
  20. Chappell, J.; Harman, J.L.; Narasimhan, V.M.; Yu, H.; Foote, K.; Simons, B.D.; Bennett, M.R.; Jørgensen, H.F. Extensive Proliferation of a Subset of Differentiated, yet Plastic, Medial Vascular Smooth Muscle Cells Contributes to Neointimal Formation in Mouse Injury and Atherosclerosis Models. Circ. Res. 2016, 119, 1313–1323. [Google Scholar] [CrossRef]
  21. Mizrak, D.; Zhao, Y.; Feng, H.; Macaulay, J.; Tang, Y.; Sultan, Z.; Zhao, G.; Guo, Y.; Zhang, J.; Yang, B.; et al. Single-Molecule Spatial Transcriptomics of Human Thoracic Aortic Aneurysms Uncovers Calcification-Related CARTPT-Expressing Smooth Muscle Cells. Arterioscler. Thromb. Vasc. Biol. 2023, 43, 2285–2297. [Google Scholar] [CrossRef] [PubMed]
  22. Allahverdian, S.; Chehroudi, A.C.; McManus, B.M.; Abraham, T.; Francis, G.A. Contribution of intimal smooth muscle cells to cholesterol accumulation and macrophage-like cells in human atherosclerosis. Circulation 2014, 129, 1551–1559. [Google Scholar] [CrossRef] [PubMed]
  23. Wang, Y.; Dubland, J.A.; Allahverdian, S.; Asonye, E.; Sahin, B.; Jaw, J.E.; Sin, D.D.; Seidman, M.A.; Leeper, N.J.; Francis, G.A. Smooth Muscle Cells Contribute the Majority of Foam Cells in ApoE (Apolipoprotein E)-Deficient Mouse Atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 2019, 39, 876–887. [Google Scholar] [CrossRef] [PubMed]
  24. Majesky, M.W.; Horita, H.; Ostriker, A.; Lu, S.; Regan, J.N.; Bagchi, A.; Dong, X.R.; Poczobutt, J.; Nemenoff, R.A.; Weiser-Evans, M.C. Differentiated Smooth Muscle Cells Generate a Subpopulation of Resident Vascular Progenitor Cells in the Adventitia Regulated by Klf4. Circ. Res. 2017, 120, 296–311. [Google Scholar] [CrossRef]
  25. Rong, J.X.; Shapiro, M.; Trogan, E.; Fisher, E.A. Transdifferentiation of mouse aortic smooth muscle cells to a macrophage-like state after cholesterol loading. Proc. Natl. Acad. Sci. USA 2003, 100, 13531–13536. [Google Scholar] [CrossRef] [PubMed]
  26. Feil, S.; Fehrenbacher, B.; Lukowski, R.; Essmann, F.; Schulze-Osthoff, K.; Schaller, M.; Feil, R. Transdifferentiation of vascular smooth muscle cells to macrophage-like cells during atherogenesis. Circ. Res. 2014, 115, 662–667. [Google Scholar] [CrossRef]
  27. Davies, J.D.; Carpenter, K.L.H.; Challis, I.R.; Figg, N.L.; McNair, R.; Proudfoot, D.; Weissberg, P.L.; Shanahan, C.M. Adipocytic differentiation and liver x receptor pathways regulate the accumulation of triacylglycerols in human vascular smooth muscle cells. J. Biol. Chem. 2005, 280, 3911–3919. [Google Scholar] [CrossRef] [PubMed]
  28. Durham, A.L.; Speer, M.Y.; Scatena, M.; Giachelli, C.M.; Shanahan, C.M. Role of smooth muscle cells in vascular calcification: Implications in atherosclerosis and arterial stiffness. Cardiovasc. Res. 2018, 114, 590–600. [Google Scholar] [CrossRef]
  29. Proudfoot, D.; Skepper, J.N.; Shanahan, C.M.; Weissberg, P.L. Calcification of human vascular cells in vitro is correlated with high levels of matrix Gla protein and low levels of osteopontin expression. Arterioscler. Thromb. Vasc. Biol. 1998, 18, 379–388. [Google Scholar] [CrossRef]
  30. Sorokin, V.; Vickneson, K.; Kofidis, T.; Woo, C.C.; Lin, X.Y.; Foo, R.; Shanahan, C.M. Role of Vascular Smooth Muscle Cell Plasticity and Interactions in Vessel Wall Inflammation. Front. Immunol. 2020, 11, 599415. [Google Scholar] [CrossRef]
  31. Miano, J.M.; Fisher, E.A.; Majesky, M.W. Fate and State of Vascular Smooth Muscle Cells in Atherosclerosis. Circulation 2021, 143, 2110–2116. [Google Scholar] [CrossRef] [PubMed]
  32. Allahverdian, S.; Chaabane, C.; Boukais, K.; Francis, G.A.; Bochaton-Piallat, M.L. Smooth muscle cell fate and plasticity in atherosclerosis. Cardiovasc. Res. 2018, 114, 540–550. [Google Scholar] [CrossRef] [PubMed]
  33. Grootaert, M.O.J.; Bennett, M.R. Vascular smooth muscle cells in atherosclerosis: Time for a re-assessment. Cardiovasc. Res. 2021, 117, 2326–2339. [Google Scholar] [CrossRef]
  34. Chen, Y.E. Vascular cell lineage determination and differentiation. Arterioscler. Thromb. Vasc. Biol. 2011, 31, 1467–1468. [Google Scholar] [CrossRef]
  35. Mack, C.P. Signaling mechanisms that regulate smooth muscle cell differentiation. Arterioscler. Thromb. Vasc. Biol. 2011, 31, 1495–1505. [Google Scholar] [CrossRef] [PubMed]
  36. Alexander, M.R.; Owens, G.K. Epigenetic control of smooth muscle cell differentiation and phenotypic switching in vascular development and disease. Annu. Rev. Physiol. 2012, 74, 13–40. [Google Scholar] [CrossRef] [PubMed]
  37. Xie, C.; Ritchie, R.P.; Huang, H.; Zhang, J.; Chen, Y.E. Smooth muscle cell differentiation in vitro: Models and underlying molecular mechanisms. Arterioscler. Thromb. Vasc. Biol. 2011, 31, 1485–1494. [Google Scholar] [CrossRef] [PubMed]
  38. Minty, A.; Kedes, L. Upstream regions of the human cardiac actin gene that modulate its transcription in muscle cells: Presence of an evolutionarily conserved repeated motif. Mol. Cell. Biol. 1986, 6, 2125–2136. [Google Scholar] [CrossRef]
  39. Sartorelli, V.; Kurabayashi, M.; Kedes, L. Muscle-specific gene expression. A comparison of cardiac and skeletal muscle transcription strategies. Circ. Res. 1993, 72, 925–931. [Google Scholar] [CrossRef]
  40. Boxer, L.M.; Prywes, R.; Roeder, R.G.; Kedes, L. The sarcomeric actin CArG-binding factor is indistinguishable from the c-fos serum response factor. Mol. Cell. Biol. 1989, 9, 515–522. [Google Scholar] [CrossRef]
  41. Wang, Z.; Wang, D.Z.; Hockemeyer, D.; McAnally, J.; Nordheim, A.; Olson, E.N. Myocardin and ternary complex factors compete for SRF to control smooth muscle gene expression. Nature 2004, 428, 185–189. [Google Scholar] [CrossRef] [PubMed]
  42. Zhang, M.; Fang, H.; Zhou, J.; Herring, B.P. A novel role of Brg1 in the regulation of SRF/MRTFA-dependent smooth muscle-specific gene expression. J. Biol. Chem. 2007, 282, 25708–25716. [Google Scholar] [CrossRef]
  43. Zhou, J.; Zhang, M.; Fang, H.; El-Mounayri, O.; Rodenberg, J.M.; Imbalzano, A.N.; Herring, B.P. The SWI/SNF chromatin remodeling complex regulates myocardin-induced smooth muscle-specific gene expression. Arterioscler. Thromb. Vasc. Biol. 2009, 29, 921–928. [Google Scholar] [CrossRef] [PubMed]
  44. Zhao, G.; Zhao, Y.; Lu, H.; Chang, Z.; Liu, H.; Wang, H.; Liang, W.; Liu, Y.; Zhu, T.; Rom, O.; et al. BAF60c prevents abdominal aortic aneurysm formation through epigenetic control of vascular smooth muscle cell homeostasis. J. Clin. Investig. 2022, 132. [Google Scholar] [CrossRef] [PubMed]
  45. Zhou, J.; Herring, B.P. Mechanisms responsible for the promoter-specific effects of myocardin. J. Biol. Chem. 2005, 280, 10861–10869. [Google Scholar] [CrossRef] [PubMed]
  46. Wang, D.Z.; Li, S.; Hockemeyer, D.; Sutherland, L.; Wang, Z.; Schratt, G.; Richardson, J.A.; Nordheim, A.; Olson, E.N. Potentiation of serum response factor activity by a family of myocardin-related transcription factors. Proc. Natl. Acad. Sci. USA 2002, 99, 14855–14860. [Google Scholar] [CrossRef] [PubMed]
  47. Yoshida, T.; Gan, Q.; Shang, Y.; Owens, G.K. Platelet-derived growth factor-BB represses smooth muscle cell marker genes via changes in binding of MKL factors and histone deacetylases to their promoters. Am. J. Physiol. Cell Physiol. 2007, 292, C886–C895. [Google Scholar] [CrossRef] [PubMed]
  48. Qiu, P.; Ritchie, R.P.; Fu, Z.; Cao, D.; Cumming, J.; Miano, J.M.; Wang, D.-Z.; Li, H.J.; Li, L. Myocardin enhances Smad3-mediated transforming growth factor-beta1 signaling in a CArG box-independent manner: Smad-binding element is an important cis element for SM22alpha transcription in vivo. Circ. Res. 2005, 97, 983–991. [Google Scholar] [CrossRef]
  49. Charron, F.; Paradis, P.; Bronchain, O.; Nemer, G.; Nemer, M. Cooperative interaction between GATA-4 and GATA-6 regulates myocardial gene expression. Mol. Cell. Biol. 1999, 19, 4355–4365. [Google Scholar] [CrossRef]
  50. Mano, T.; Luo, Z.; Malendowicz, S.L.; Evans, T.; Walsh, K. Reversal of GATA-6 downregulation promotes smooth muscle differentiation and inhibits intimal hyperplasia in balloon-injured rat carotid artery. Circ. Res. 1999, 84, 647–654. [Google Scholar] [CrossRef]
  51. Wada, H.; Hasegawa, K.; Morimoto, T.; Kakita, T.; Yanazume, T.; Sasayama, S. A p300 protein as a coactivator of GATA-6 in the transcription of the smooth muscle-myosin heavy chain gene. J. Biol. Chem. 2000, 275, 25330–25335. [Google Scholar] [CrossRef] [PubMed]
  52. Nishida, W.; Nakamura, M.; Mori, S.; Takahashi, M.; Ohkawa, Y.; Tadokoro, S.; Yoshida, K.; Hiwada, K.; Hayashi, K.; Sobue, K. A triad of serum response factor and the GATA and NK families governs the transcription of smooth and cardiac muscle genes. J. Biol. Chem. 2002, 277, 7308–7317. [Google Scholar] [CrossRef] [PubMed]
  53. Kurz, J.; Weiss, A.C.; Lüdtke, T.H.; Deuper, L.; Trowe, M.O.; Thiesler, H.; Hildebrandt, H.; Heineke, J.; Duncan, S.A.; Kispert, A. GATA6 is a crucial factor for Myocd expression in the visceral smooth muscle cell differentiation program of the murine ureter. Development 2022, 149. [Google Scholar] [CrossRef] [PubMed]
  54. Chang, D.F.; Belaguli, N.S.; Iyer, D.; Roberts, W.B.; Wu, S.P.; Dong, X.R.; Marx, J.G.; Moore, M.S.; Beckerle, M.C.; Majesky, M.W.; et al. Cysteine-rich LIM-only proteins CRP1 and CRP2 are potent smooth muscle differentiation cofactors. Dev. Cell 2003, 4, 107–118. [Google Scholar] [CrossRef]
  55. Chang, D.F.; Belaguli, N.S.; Chang, J.; Schwartz, R.J. LIM-only protein, CRP2, switched on smooth muscle gene activity in adult cardiac myocytes. Proc. Natl. Acad. Sci. USA 2007, 104, 157–162. [Google Scholar] [CrossRef]
  56. Yoshida, T.; Hoofnagle, M.H.; Owens, G.K. Myocardin and Prx1 contribute to angiotensin II-induced expression of smooth muscle alpha-actin. Circ. Res. 2004, 94, 1075–1082. [Google Scholar] [CrossRef] [PubMed]
  57. Shang, Y.; Yoshida, T.; Amendt, B.A.; Martin, J.F.; Owens, G.K. Pitx2 is functionally important in the early stages of vascular smooth muscle cell differentiation. J. Cell Biol. 2008, 181, 461–473. [Google Scholar] [CrossRef] [PubMed]
  58. Kawai-Kowase, K.; Kumar, M.S.; Hoofnagle, M.H.; Yoshida, T.; Owens, G.K. PIAS1 activates the expression of smooth muscle cell differentiation marker genes by interacting with serum response factor and class I basic helix-loop-helix proteins. Mol. Cell. Biol. 2005, 25, 8009–8023. [Google Scholar] [CrossRef]
  59. Katoh, Y.; Molkentin, J.D.; Dave, V.; Olson, E.N.; Periasamy, M. MEF2B is a component of a smooth muscle-specific complex that binds an A/T-rich element important for smooth muscle myosin heavy chain gene expression. J. Biol. Chem. 1998, 273, 1511–1518. [Google Scholar] [CrossRef]
  60. Noseda, M.; Fu, Y.; Niessen, K.; Wong, F.; Chang, L.; McLean, G.; Karsan, A. Smooth Muscle alpha-actin is a direct target of Notch/CSL. Circ. Res. 2006, 98, 1468–1470. [Google Scholar] [CrossRef]
  61. Tang, Y.; Urs, S.; Boucher, J.; Bernaiche, T.; Venkatesh, D.; Spicer, D.B.; Vary, C.P.; Liaw, L. Notch and transforming growth factor-beta (TGFbeta) signaling pathways cooperatively regulate vascular smooth muscle cell differentiation. J. Biol. Chem. 2010, 285, 17556–17563. [Google Scholar] [CrossRef]
  62. Liu, Y.; Sinha, S.; McDonald, O.G.; Shang, Y.; Hoofnagle, M.H.; Owens, G.K. Kruppel-like factor 4 abrogates myocardin-induced activation of smooth muscle gene expression. J. Biol. Chem. 2005, 280, 9719–9727. [Google Scholar] [CrossRef] [PubMed]
  63. Yoshida, T.; Gan, Q.; Owens, G.K. Kruppel-like factor 4, Elk-1, and histone deacetylases cooperatively suppress smooth muscle cell differentiation markers in response to oxidized phospholipids. Am. J. Physiol. Cell Physiol. 2008, 295, C1175–C1182. [Google Scholar] [CrossRef] [PubMed]
  64. Salmon, M.; Gomez, D.; Greene, E.; Shankman, L.; Owens, G.K. Cooperative binding of KLF4, pELK-1, and HDAC2 to a G/C repressor element in the SM22alpha promoter mediates transcriptional silencing during SMC phenotypic switching in vivo. Circ. Res. 2012, 111, 685–696. [Google Scholar] [CrossRef] [PubMed]
  65. Liu, Y.; Sinha, S.; Owens, G. A transforming growth factor-beta control element required for SM alpha-actin expression in vivo also partially mediates GKLF-dependent transcriptional repression. J. Biol. Chem. 2003, 278, 48004–48011. [Google Scholar] [CrossRef] [PubMed]
  66. Cao, D.; Wang, Z.; Zhang, C.L.; Oh, J.; Xing, W.; Li, S.; Richardson, J.A.; Wang, D.Z.; Olson, E.N. Modulation of smooth muscle gene expression by association of histone acetyltransferases and deacetylases with myocardin. Mol. Cell. Biol. 2005, 25, 364–376. [Google Scholar] [CrossRef] [PubMed]
  67. Chakraborty, R.; Ostriker, A.C.; Xie, Y.; Dave, J.M.; Gamez-Mendez, A.; Chatterjee, P.; Abu, Y.; Valentine, J.; Lezon-Geyda, K.; Greif, D.M.; et al. Histone Acetyltransferases p300 and CBP Coordinate Distinct Chromatin Remodeling Programs in Vascular Smooth Muscle Plasticity. Circulation 2022, 145, 1720–1737. [Google Scholar] [CrossRef] [PubMed]
  68. Zhou, Y.; Sharma, S.; Sun, X.; Guan, X.; Hou, Y.; Yang, Z.; Shi, H.; Zou, M.H.; Song, P.; Zhou, J.; et al. SMYD2 regulates vascular smooth muscle cell phenotypic switching and intimal hyperplasia via interaction with myocardin. Cell. Mol. Life Sci. 2023, 80, 264. [Google Scholar] [CrossRef]
  69. Lockman, K.; Taylor, J.M.; Mack, C.P. The histone demethylase, Jmjd1a, interacts with the myocardin factors to regulate SMC differentiation marker gene expression. Circ. Res. 2007, 101, e115–e123. [Google Scholar] [CrossRef]
  70. Gan, Q.; Thiébaud, P.; Thézé, N.; Jin, L.; Xu, G.; Grant, P.; Owens, G.K. WD repeat-containing protein 5, a ubiquitously expressed histone methyltransferase adaptor protein, regulates smooth muscle cell-selective gene activation through interaction with pituitary homeobox 2. J. Biol. Chem. 2011, 286, 21853–21864. [Google Scholar] [CrossRef]
  71. Li, N.; Subrahmanyan, L.; Smith, E.; Yu, X.; Zaidi, S.; Choi, M.; Mane, S.; Nelson-Williams, C.; Behjati, M.; Kazemi, M.; et al. Mutations in the Histone Modifier PRDM6 Are Associated with Isolated Nonsyndromic Patent Ductus Arteriosus. Am. J. Hum. Genet. 2016, 98, 1082–1091. [Google Scholar] [CrossRef] [PubMed]
  72. Davis, C.A.; Haberland, M.; Arnold, M.A.; Sutherland, L.B.; McDonald, O.G.; Richardson, J.A.; Childs, G.; Harris, S.; Owens, G.K.; Olson, E.N. PRISM/PRDM6, a transcriptional repressor that promotes the proliferative gene program in smooth muscle cells. Mol. Cell. Biol. 2006, 26, 2626–2636. [Google Scholar] [CrossRef] [PubMed]
  73. Chatterjee, P.; Chakraborty, R.; Xie, Y.; Sizer, A.; Hwa, J.; Martin, K.A. The Histone Methyl Transferase (SUV39H1) Promotes Smooth Muscle Cell Dedifferentiation. In Arteriosclerosis, Thrombosis, and Vascular Biology; 2022. [Google Scholar]
  74. Liang, J.; Li, Q.; Cai, W.; Zhang, X.; Yang, B.; Li, X.; Jiang, S.; Tian, S.; Zhang, K.; Song, H.; et al. Inhibition of polycomb repressor complex 2 ameliorates neointimal hyperplasia by suppressing trimethylation of H3K27 in vascular smooth muscle cells. Br. J. Pharmacol. 2019, 176, 3206–3219. [Google Scholar] [CrossRef] [PubMed]
  75. Lino Cardenas, C.L.; Kessinger, C.W.; MacDonald, C.; Jassar, A.S.; Isselbacher, E.M.; Jaffer, F.A.; Lindsay, M.E. Inhibition of the methyltranferase EZH2 improves aortic performance in experimental thoracic aortic aneurysm. JCI Insight 2018, 3. [Google Scholar] [CrossRef] [PubMed]
  76. Ostriker, A.C.; Xie, Y.; Chakraborty, R.; Sizer, A.J.; Bai, Y.; Ding, M.; Song, W.L.; Huttner, A.; Hwa, J.; Martin, K.A. TET2 Protects Against Vascular Smooth Muscle Cell Apoptosis and Intimal Thickening in Transplant Vasculopathy. Circulation 2021, 144, 455–470. [Google Scholar] [CrossRef] [PubMed]
  77. Liu, R.; Jin, Y.; Tang, W.H.; Qin, L.; Zhang, X.; Tellides, G.; Hwa, J.; Yu, J.; Martin, K.A. Ten-eleven translocation-2 (TET2) is a master regulator of smooth muscle cell plasticity. Circulation 2013, 128, 2047–2057. [Google Scholar] [CrossRef] [PubMed]
  78. Zhu, N.; Guo, Z.-F.; Kazama, K.; Yi, B.; Tongmuang, N.; Yao, H.; Yang, R.; Zhang, C.; Qin, Y.; Han, L.; et al. Epigenetic regulation of vascular smooth muscle cell phenotypic switch and neointimal formation by PRMT5. Cardiovasc. Res. 2023, 119, 2244–2255. [Google Scholar] [CrossRef] [PubMed]
  79. Shah, N.M.; Groves, A.K.; Anderson, D.J. Alternative neural crest cell fates are instructively promoted by TGFbeta superfamily members. Cell 1996, 85, 331–343. [Google Scholar] [CrossRef]
  80. Hautmann, M.B.; Madsen, C.S.; Owens, G.K. A transforming growth factor beta (TGFbeta) control element drives TGFbeta-induced stimulation of smooth muscle alpha-actin gene expression in concert with two CArG elements. J. Biol. Chem. 1997, 272, 10948–10956. [Google Scholar] [CrossRef]
  81. Janknecht, R.; Wells, N.J.; Hunter, T. TGF-beta-stimulated cooperation of smad proteins with the coactivators CBP/p300. Genes Dev. 1998, 12, 2114–2119. [Google Scholar] [CrossRef]
  82. Narita, N.; Heikinheimo, M.; Bielinska, M.; White, R.A.; Wilson, D.B. The gene for transcription factor GATA-6 resides on mouse chromosome 18 and is expressed in myocardium and vascular smooth muscle. Genomics 1996, 36, 345–348. [Google Scholar] [CrossRef] [PubMed]
  83. Yap, C.; Mieremet, A.; de Vries, C.J.M.; Micha, D.; de Waard, V. Six Shades of Vascular Smooth Muscle Cells Illuminated by KLF4 (Kruppel-Like Factor 4). Arterioscler. Thromb. Vasc. Biol. 2021, 41, 2693–2707. [Google Scholar] [CrossRef] [PubMed]
  84. Adam, P.J.; Regan, C.P.; Hautmann, M.B.; Owens, G.K. Positive- and negative-acting Kruppel-like transcription factors bind a transforming growth factor beta control element required for expression of the smooth muscle cell differentiation marker SM22alpha in vivo. J. Biol. Chem. 2000, 275, 37798–37806. [Google Scholar] [CrossRef] [PubMed]
  85. Yoshida, T.; Kaestner, K.H.; Owens, G.K. Conditional deletion of Kruppel-like factor 4 delays downregulation of smooth muscle cell differentiation markers but accelerates neointimal formation following vascular injury. Circ. Res. 2008, 102, 1548–1557. [Google Scholar] [CrossRef]
  86. McDonald, O.G.; Wamhoff, B.R.; Hoofnagle, M.H.; Owens, G.K. Control of SRF binding to CArG box chromatin regulates smooth muscle gene expression in vivo. J. Clin. Investig. 2006, 116, 36–48. [Google Scholar] [CrossRef]
  87. Manabe, I.; Owens, G.K. Recruitment of serum response factor and hyperacetylation of histones at smooth muscle-specific regulatory regions during differentiation of a novel P19-derived in vitro smooth muscle differentiation system. Circ. Res. 2001, 88, 1127–1134. [Google Scholar] [CrossRef]
  88. Yoshida, T.; Sinha, S.; Dandré, F.; Wamhoff, B.R.; Hoofnagle, M.H.; Kremer, B.E.; Wang, D.Z.; Olson, E.N.; Owens, G.K. Myocardin is a key regulator of CArG-dependent transcription of multiple smooth muscle marker genes. Circ. Res. 2003, 92, 856–864. [Google Scholar] [CrossRef]
  89. Pellegrini, L.; Tan, S.; Richmond, T.J. Structure of serum response factor core bound to DNA. Nature 1995, 376, 490–498. [Google Scholar] [CrossRef]
  90. Luger, K.; Mader, A.W.; Richmond, R.K.; Sargent, D.F.; Richmond, T.J. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 1997, 389, 251–260. [Google Scholar] [CrossRef]
  91. McDonald, O.G.; Owens, G.K. Programming smooth muscle plasticity with chromatin dynamics. Circ. Res. 2007, 100, 1428–1441. [Google Scholar] [CrossRef]
  92. Gomez, D.; Shankman, L.S.; Nguyen, A.T.; Owens, G.K. Detection of histone modifications at specific gene loci in single cells in histological sections. Nat. Methods 2013, 10, 171–177. [Google Scholar] [CrossRef]
  93. Liu, M.; Espinosa-Diez, C.; Mahan, S.; Du, M.; Nguyen, A.T.; Hahn, S.; Chakraborty, R.; Straub, A.C.; Martin, K.A.; Owens, G.K.; et al. H3K4 di-methylation governs smooth muscle lineage identity and promotes vascular homeostasis by restraining plasticity. Dev. Cell 2021, 56, 2765–2782.e10. [Google Scholar] [CrossRef] [PubMed]
  94. Richmond, T.J.; Davey, C.A. The structure of DNA in the nucleosome core. Nature 2003, 423, 145–150. [Google Scholar] [CrossRef]
  95. de la Serna, I.L.; Ohkawa, Y.; Imbalzano, A.N. Chromatin remodelling in mammalian differentiation: Lessons from ATP-dependent remodellers. Nat. Rev. Genet. 2006, 7, 461–473. [Google Scholar] [CrossRef] [PubMed]
  96. Lessard, J.A.; Crabtree, G.R. Chromatin regulatory mechanisms in pluripotency. Annu. Rev. Cell Dev. Biol. 2010, 26, 503–532. [Google Scholar] [CrossRef]
  97. Singhal, N.; Graumann, J.; Wu, G.; Araúzo-Bravo, M.J.; Han, D.W.; Greber, B.; Gentile, L.; Mann, M.; Schöler, H.R. Chromatin-Remodeling Components of the BAF Complex Facilitate Reprogramming. Cell 2010, 141, 943–955. [Google Scholar] [CrossRef] [PubMed]
  98. Cairns, B.R. The logic of chromatin architecture and remodelling at promoters. Nature 2009, 461, 193–198. [Google Scholar] [CrossRef]
  99. Clapier, C.R.; Cairns, B.R. The biology of chromatin remodeling complexes. Annu. Rev. Biochem. 2009, 78, 273–304. [Google Scholar] [CrossRef]
  100. Stern, M.; Jensen, R.; Herskowitz, I. Five SWI genes are required for expression of the HO gene in yeast. J. Mol. Biol. 1984, 178, 853–868. [Google Scholar] [CrossRef]
  101. Neigeborn, L.; Carlson, M. Genes affecting the regulation of SUC2 gene expression by glucose repression in Saccharomyces cerevisiae. Genetics 1984, 108, 845–858. [Google Scholar] [CrossRef]
  102. Winston, F.; Carlson, M. Yeast SNF/SWI transcriptional activators and the SPT/SIN chromatin connection. Trends Genet. 1992, 8, 387–391. [Google Scholar] [CrossRef] [PubMed]
  103. Kwon, H.; Imbalzano, A.N.; Khavari, P.A.; Kingston, R.E.; Green, M.R. Nucleosome disruption and enhancement of activator binding by a human SW1/SNF complex. Nature 1994, 370, 477–481. [Google Scholar] [CrossRef]
  104. Wang, W.; Côté, J.; Xue, Y.; Zhou, S.; Khavari, P.A.; Biggar, S.R.; Muchardt, C.; Kalpana, G.V.; Goff, S.P.; Yaniv, M.; et al. Purification and biochemical heterogeneity of the mammalian SWI-SNF complex. EMBO J. 1996, 15, 5370–5382. [Google Scholar] [CrossRef] [PubMed]
  105. Lemon, B.; Inouye, C.; King, D.S.; Tjian, R. Selectivity of chromatin-remodelling cofactors for ligand-activated transcription. Nature 2001, 414, 924–928. [Google Scholar] [CrossRef] [PubMed]
  106. Alpsoy, A.; Dykhuizen, E.C. Glioma tumor suppressor candidate region gene 1 (GLTSCR1) and its paralog GLTSCR1-like form SWI/SNF chromatin remodeling subcomplexes. J. Biol. Chem. 2018, 293, 3892–3903. [Google Scholar] [CrossRef] [PubMed]
  107. Michel, B.C.; D’Avino, A.R.; Cassel, S.H.; Mashtalir, N.; McKenzie, Z.M.; McBride, M.J.; Valencia, A.M.; Zhou, Q.; Bocker, M.; Soares, L.M.M.; et al. A non-canonical SWI/SNF complex is a synthetic lethal target in cancers driven by BAF complex perturbation. Nat. Cell Biol. 2018, 20, 1410–1420. [Google Scholar] [CrossRef] [PubMed]
  108. Wang, X.; Wang, S.; Troisi, E.C.; Howard, T.P.; Haswell, J.R.; Wolf, B.K.; Hawk, W.H.; Ramos, P.; Oberlick, E.M.; Tzvetkov, E.P.; et al. BRD9 defines a SWI/SNF sub-complex and constitutes a specific vulnerability in malignant rhabdoid tumors. Nat. Commun. 2019, 10, 1881. [Google Scholar] [CrossRef] [PubMed]
  109. Mashtalir, N.; D’Avino, A.R.; Michel, B.C.; Luo, J.; Pan, J.; Otto, J.E.; Zullow, H.J.; McKenzie, Z.M.; Kubiak, R.L.; St Pierre, R.; et al. Modular Organization and Assembly of SWI/SNF Family Chromatin Remodeling Complexes. Cell 2018, 175, 1272–1288.e20. [Google Scholar] [CrossRef]
  110. Tang, L.; Nogales, E.; Ciferri, C. Structure and function of SWI/SNF chromatin remodeling complexes and mechanistic implications for transcription. Prog. Biophys. Mol. Biol. 2010, 102, 122–128. [Google Scholar] [CrossRef]
  111. Wu, J.I.; Lessard, J.; Crabtree, G.R. Understanding the words of chromatin regulation. Cell 2009, 136, 200–206. [Google Scholar] [CrossRef]
  112. Otto, J.E.; Ursu, O.; Wu, A.P.; Winter, E.B.; Cuoco, M.S.; Ma, S.; Qian, K.; Michel, B.C.; Buenrostro, J.D.; Berger, B.; et al. Structural and functional properties of mSWI/SNF chromatin remodeling complexes revealed through single-cell perturbation screens. Mol. Cell 2023, 83, 1350–1367.e7. [Google Scholar] [CrossRef]
  113. Mashtalir, N.; Suzuki, H.; Farrell, D.P.; Sankar, A.; Luo, J.; Filipovski, M.; D’Avino, A.R.; St Pierre, R.; Valencia, A.M.; Onikubo, T.; et al. A Structural Model of the Endogenous Human BAF Complex Informs Disease Mechanisms. Cell 2020, 183, 802–817.e24. [Google Scholar] [CrossRef] [PubMed]
  114. He, S.; Wu, Z.; Tian, Y.; Yu, Z.; Yu, J.; Wang, X.; Li, J.; Liu, B.; Xu, Y. Structure of nucleosome-bound human BAF complex. Science 2020, 367, 875–881. [Google Scholar] [CrossRef] [PubMed]
  115. Yuan, J.; Chen, K.; Zhang, W.; Chen, Z. Structure of human chromatin-remodelling PBAF complex bound to a nucleosome. Nature 2022, 605, 166–171. [Google Scholar] [CrossRef] [PubMed]
  116. Wang, L.; Yu, J.; Yu, Z.; Wang, Q.; Li, W.; Ren, Y.; Chen, Z.; He, S.; Xu, Y. Structure of nucleosome-bound human PBAF complex. Nat. Commun. 2022, 13, 7644. [Google Scholar] [CrossRef]
  117. Chen, K.; Yuan, J.; Sia, Y.; Chen, Z. Mechanism of action of the SWI/SNF family complexes. Nucleus 2023, 14, 2165604. [Google Scholar] [CrossRef]
  118. Clapier, C.R.; Iwasa, J.; Cairns, B.R.; Peterson, C.L. Mechanisms of action and regulation of ATP-dependent chromatin-remodelling complexes. Nat. Rev. Mol. Cell. Biol. 2017, 18, 407–422. [Google Scholar] [CrossRef]
  119. Valencia, A.M.; Collings, C.K.; Dao, H.T.; St Pierre, R.; Cheng, Y.C.; Huang, J.; Sun, Z.Y.; Seo, H.S.; Mashtalir, N.; Comstock, D.E.; et al. Recurrent SMARCB1 Mutations Reveal a Nucleosome Acidic Patch Interaction Site That Potentiates mSWI/SNF Complex Chromatin Remodeling. Cell 2019, 179, 1342–1356.e23. [Google Scholar] [CrossRef]
  120. Stros, M.; Launholt, D.; Grasser, K.D. The HMG-box: A versatile protein domain occurring in a wide variety of DNA-binding proteins. Cell. Mol. Life Sci. CMLS 2007, 64, 2590–2606. [Google Scholar] [CrossRef]
  121. Kim, S.; Zhang, Z.; Upchurch, S.; Isern, N.; Chen, Y. Structure and DNA-binding sites of the SWI1 AT-rich interaction domain (ARID) suggest determinants for sequence-specific DNA recognition. J. Biol. Chem. 2004, 279, 16670–16676. [Google Scholar] [CrossRef]
  122. Priam, P.; Krasteva, V.; Rousseau, P.; D’Angelo, G.; Gaboury, L.; Sauvageau, G.; Lessard, J.A. SMARCD2 subunit of SWI/SNF chromatin-remodeling complexes mediates granulopoiesis through a CEBPvarepsilon dependent mechanism. Nat. Genet. 2017, 49, 753–764. [Google Scholar] [CrossRef] [PubMed]
  123. Li, S.; Liu, C.; Li, N.; Hao, T.; Han, T.; Hill, D.E.; Vidal, M.; Lin, J.D. Genome-wide coactivation analysis of PGC-1alpha identifies BAF60a as a regulator of hepatic lipid metabolism. Cell Metab. 2008, 8, 105–117. [Google Scholar] [CrossRef] [PubMed]
  124. Wolf, B.K.; Zhao, Y.; McCray, A.; Hawk, W.H.; Deary, L.T.; Sugiarto, N.W.; LaCroix, I.S.; Gerber, S.A.; Cheng, C.; Wang, X. Cooperation of chromatin remodeling SWI/SNF complex and pioneer factor AP-1 shapes 3D enhancer landscapes. Nat. Struct. Mol. Biol. 2023, 30, 10–21. [Google Scholar] [CrossRef] [PubMed]
  125. Branon, T.C.; Bosch, J.A.; Sanchez, A.D.; Udeshi, N.D.; Svinkina, T.; Carr, S.A.; Feldman, J.L.; Perrimon, N.; Ting, A.Y. Efficient proximity labeling in living cells and organisms with TurboID. Nat. Biotechnol. 2018, 36, 880–887. [Google Scholar] [CrossRef] [PubMed]
  126. Haynes, S.R.; Dollard, C.; Winston, F.; Beck, S.; Trowsdale, J.; Dawid, I.B. The bromodomain: A conserved sequence found in human, Drosophila and yeast proteins. Nucleic Acids Res. 1992, 20, 2603. [Google Scholar] [CrossRef] [PubMed]
  127. Jacobs, S.A.; Khorasanizadeh, S. Structure of HP1 chromodomain bound to a lysine 9-methylated histone H3 tail. Science 2002, 295, 2080–2083. [Google Scholar] [CrossRef] [PubMed]
  128. Hong, C.Y.; Suh, J.H.; Kim, K.; Gong, E.-Y.; Jeon, S.H.; Ko, M.; Seong, R.H.; Kwon, H.B.; Lee, K. Modulation of androgen receptor transactivation by the SWI3-related gene product (SRG3) in multiple ways. Mol. Cell. Biol. 2005, 25, 4841–4852. [Google Scholar] [CrossRef] [PubMed]
  129. Naidu, S.R.; Love, I.M.; Imbalzano, A.N.; Grossman, S.R.; Androphy, E.J. The SWI/SNF chromatin remodeling subunit BRG1 is a critical regulator of p53 necessary for proliferation of malignant cells. Oncogene 2009, 28, 2492–2501. [Google Scholar] [CrossRef]
  130. Hsiao, P.-W.; Fryer, C.J.; Trotter, K.W.; Wang, W.; Archer, T.K. BAF60a mediates critical interactions between nuclear receptors and the BRG1 chromatin-remodeling complex for transactivation. Mol. Cell. Biol. 2003, 23, 6210–6220. [Google Scholar] [CrossRef]
  131. Debril, M.B.; Gelman, L.; Fayard, E.; Annicotte, J.S.; Rocchi, S.; Auwerx, J. Transcription factors and nuclear receptors interact with the SWI/SNF complex through the BAF60c subunit. J. Biol. Chem. 2004, 279, 16677–16686. [Google Scholar] [CrossRef]
  132. Ito, T.; Yamauchi, M.; Nishina, M.; Yamamichi, N.; Mizutani, T.; Ui, M.; Murakami, M.; Iba, H. Identification of SWI.SNF complex subunit BAF60a as a determinant of the transactivation potential of Fos/Jun dimers. J. Biol. Chem. 2001, 276, 2852–2857. [Google Scholar] [CrossRef] [PubMed]
  133. Pal, S.; Yun, R.; Datta, A.; Lacomis, L.; Erdjument-Bromage, H.; Kumar, J.; Tempst, P.; Sif, S. mSin3A/histone deacetylase 2- and PRMT5-containing Brg1 complex is involved in transcriptional repression of the Myc target gene cad. Mol. Cell. Biol. 2003, 23, 7475–7487. [Google Scholar] [CrossRef]
  134. Sun, X.; Hota, S.K.; Zhou, Y.Q.; Novak, S.; Miguel-Perez, D.; Christodoulou, D.; Seidman, C.E.; Seidman, J.G.; Gregorio, C.C.; Henkelman, R.M.; et al. Cardiac-enriched BAF chromatin-remodeling complex subunit Baf60c regulates gene expression programs essential for heart development and function. Biol. Open 2018, 7, bio029512. [Google Scholar] [CrossRef] [PubMed]
  135. Flajollet, S.; Lefebvre, B.; Cudejko, C.; Staels, B.; Lefebvre, P. The core component of the mammalian SWI/SNF complex SMARCD3/BAF60c is a coactivator for the nuclear retinoic acid receptor. Mol. Cell Endocrinol. 2007, 270, 23–32. [Google Scholar] [CrossRef] [PubMed]
  136. Belandia, B.; Orford, R.L.; Hurst, H.C.; Parker, M.G. Targeting of SWI/SNF chromatin remodelling complexes to estrogen-responsive genes. EMBO J. 2002, 21, 4094–4103. [Google Scholar] [CrossRef] [PubMed]
  137. Takeuchi, J.K.; Lickert, H.; Bisgrove, B.W.; Sun, X.; Yamamoto, M.; Chawengsaksophak, K.; Hamada, H.; Yost, H.J.; Rossant, J.; Bruneau, B.G. Baf60c is a nuclear Notch signaling component required for the establishment of left-right asymmetry. Proc. Natl. Acad. Sci. USA 2007, 104, 846–851. [Google Scholar] [CrossRef]
  138. Tsurusaki, Y.; Okamoto, N.; Ohashi, H.; Kosho, T.; Imai, Y.; Hibi-Ko, Y.; Kaname, T.; Naritomi, K.; Kawame, H.; Wakui, K.; et al. Mutations affecting components of the SWI/SNF complex cause Coffin-Siris syndrome. Nat. Genet. 2012, 44, 376–378. [Google Scholar] [CrossRef]
  139. van der Sluijs, P.J.; Jansen, S.; Vergano, S.A.; Adachi-Fukuda, M.; Alanay, Y.; AlKindy, A.; Baban, A.; Bayat, A.; Beck-Wödl, S.; Berry, K.; et al. The ARID1B spectrum in 143 patients: From nonsyndromic intellectual disability to Coffin-Siris syndrome. Genet. Med. 2019, 21, 1295–1307. [Google Scholar] [CrossRef]
  140. van der Sluijs, P.J.; Joosten, M.; Alby, C.; Attié-Bitach, T.; Gilmore, K.; Dubourg, C.; Fradin, M.; Wang, T.; Kurtz-Nelson, E.C.; Ahlers, K.P.; et al. Discovering a new part of the phenotypic spectrum of Coffin-Siris syndrome in a fetal cohort. Genet. Med. 2022, 24, 1753–1760. [Google Scholar] [CrossRef]
  141. Sun, H.; Zhang, S.; Wang, J.; Zhou, X.; Zhang, H.; Yang, H.; He, Y. Expanding the phenotype associated with SMARCC2 variants: A fetus with tetralogy of Fallot. BMC Med. Genom. 2022, 15, 40. [Google Scholar] [CrossRef]
  142. Bultman, S.; Gebuhr, T.; Yee, D.; La Mantia, C.; Nicholson, J.; Gilliam, A.; Randazzo, F.; Metzger, D.; Chambon, P.; Crabtree, G.; et al. A Brg1 null mutation in the mouse reveals functional differences among mammalian SWI/SNF complexes. Mol. Cell 2000, 6, 1287–1295. [Google Scholar] [CrossRef] [PubMed]
  143. Reyes, J.C.; Barra, J.; Muchardt, C.; Camus, A.; Babinet, C.; Yaniv, M. Altered control of cellular proliferation in the absence of mammalian brahma (SNF2alpha). EMBO J. 1998, 17, 6979–6991. [Google Scholar] [CrossRef] [PubMed]
  144. Lange, M.; Kaynak, B.; Forster, U.B.; Tönjes, M.; Fischer, J.J.; Grimm, C.; Schlesinger, J.; Just, S.; Dunkel, I.; Krueger, T.; et al. Regulation of muscle development by DPF3, a novel histone acetylation and methylation reader of the BAF chromatin remodeling complex. Genes Dev. 2008, 22, 2370–2384. [Google Scholar] [CrossRef] [PubMed]
  145. Han, F.; Yang, B.; Chen, Y.; Liu, L.; Cheng, X.; Huang, J.; Zhou, K.; Zhang, D.; Xu, E.; Lai, M.; et al. Loss of GLTSCR1 causes congenital heart defects by regulating NPPA transcription. Angiogenesis 2023, 26, 217–232. [Google Scholar] [CrossRef] [PubMed]
  146. de La Serna, I.L.; Carlson, K.A.; Hill, D.A.; Guidi, C.J.; Stephenson, R.O.; Sif, S.; Kingston, R.E.; Imbalzano, A.N. Mammalian SWI-SNF complexes contribute to activation of the hsp70 gene. Mol. Cell. Biol. 2000, 20, 2839–2851. [Google Scholar] [CrossRef] [PubMed]
  147. Chen, M.; Herring, B.P. Regulation of microRNAs by Brahma-related gene 1 (Brg1) in smooth muscle cells. J. Biol. Chem. 2013, 288, 6397–6408. [Google Scholar] [CrossRef] [PubMed]
  148. Zhang, M.; Chen, M.; Kim, J.-R.; Zhou, J.; Jones, R.E.; Tune, J.D.; Kassab, G.S.; Metzger, D.; Ahlfeld, S.; Conway, S.J.; et al. SWI/SNF Complexes Containing Brahma or Brahma-Related Gene 1 Play Distinct Roles in Smooth Muscle Development. Mol. Cell. Biol. 2011, 31, 2618–2631. [Google Scholar] [CrossRef]
  149. Jankowich, M.; Choudhary, G. Endothelin-1 levels and cardiovascular events. Trends Cardiovasc. Med. 2020, 30, 1–8. [Google Scholar] [CrossRef]
  150. Li, L.; Liu, D.; Bu, D.; Chen, S.; Wu, J.; Tang, C.; Du, J.; Jin, H. Brg1-dependent epigenetic control of vascular smooth muscle cell proliferation by hydrogen sulfide. Biochim. Biophys. Acta 2013, 1833, 1347–1355. [Google Scholar] [CrossRef]
  151. Yang, Y.; Cheng, X.; Tian, W.; Zhou, B.; Wu, X.; Xu, H.; Fang, F.; Fang, M.; Xu, Y. MRTF-A steers an epigenetic complex to activate endothelin-induced pro-inflammatory transcription in vascular smooth muscle cells. Nucleic Acids Res. 2014, 42, 10460–10472. [Google Scholar] [CrossRef]
  152. Yuan, Y.; Wang, C.; Xu, J.; Tao, J.; Xu, Z.; Huang, S. BRG1 overexpression in smooth muscle cells promotes the development of thoracic aortic dissection. BMC Cardiovasc. Disord. 2014, 14, 144. [Google Scholar] [CrossRef] [PubMed]
  153. Liao, W.L.; Tan, M.W.; Yuan, Y.; Wang, G.K.; Wang, C.; Tang, H.; Xu, Z.Y. Brahma-related gene 1 inhibits proliferation and migration of human aortic smooth muscle cells by directly up-regulating Ras-related associated with diabetes in the pathophysiologic processes of aortic dissection. J. Thorac. Cardiovasc. Surg. 2015, 150, 1292–1301.e2. [Google Scholar] [CrossRef] [PubMed]
  154. Wang, S.; Zhang, X.; Yuan, Y.; Tan, M.; Zhang, L.; Xue, X.; Yan, Y.; Han, L.; Xu, Z. BRG1 expression is increased in thoracic aortic aneurysms and regulates proliferation and apoptosis of vascular smooth muscle cells through the long non-coding RNA HIF1A-AS1 in vitro. Eur. J. Cardiothorac. Surg. 2015, 47, 439–446. [Google Scholar] [CrossRef] [PubMed]
  155. Lino Cardenas, C.L.; Kessinger, C.W.; Cheng, Y.; MacDonald, C.; MacGillivray, T.; Ghoshhajra, B.; Huleihel, L.; Nuri, S.; Yeri, A.S.; Jaffer, F.A.; et al. An HDAC9-MALAT1-BRG1 complex mediates smooth muscle dysfunction in thoracic aortic aneurysm. Nat. Commun. 2018, 9, 1009. [Google Scholar] [CrossRef] [PubMed]
  156. Jolly, A.J.; Lu, S.; Dubner, A.M.; Strand, K.A.; Mutryn, M.F.; Pilotti-Riley, A.; Danis, E.P.; Nemenoff, R.A.; Moulton, K.S.; Majesky, M.W.; et al. Redistribution of the chromatin remodeler Brg1 directs smooth muscle-derived adventitial progenitor-to-myofibroblast differentiation and vascular fibrosis. JCI Insight 2023, 8, e164862. [Google Scholar] [CrossRef] [PubMed]
  157. Weber, C.M.; Hafner, A.; Kirkland, J.G.; Braun, S.M.G.; Stanton, B.Z.; Boettiger, A.N.; Crabtree, G.R. mSWI/SNF promotes Polycomb repression both directly and through genome-wide redistribution. Nat. Struct. Mol. Biol. 2021, 28, 501–511. [Google Scholar] [CrossRef] [PubMed]
  158. Wang, W.; Xue, Y.; Zhou, S.; Kuo, A.; Cairns, B.R.; Crabtree, G.R. Diversity and specialization of mammalian SWI/SNF complexes. Genes Dev. 1996, 10, 2117–2130. [Google Scholar] [CrossRef] [PubMed]
  159. Puri, P.L.; Mercola, M. BAF60 A, B, and Cs of muscle determination and renewal. Genes. Amp Dev. 2012, 26, 2673–2683. [Google Scholar] [CrossRef]
  160. Ochi, H.; Hans, S.; Westerfield, M. Smarcd3 regulates the timing of zebrafish myogenesis onset. J. Biol. Chem. 2008, 283, 3529–3536. [Google Scholar] [CrossRef]
  161. Forcales, S.V.; Albini, S.; Giordani, L.; Malecova, B.; Cignolo, L.; Chernov, A.; Coutinho, P.; Saccone, V.; Consalvi, S.; Williams, R.; et al. Signal-dependent incorporation of MyoD-BAF60c into Brg1-based SWI/SNF chromatin-remodelling complex. EMBO J. 2012, 31, 301–316. [Google Scholar] [CrossRef]
  162. Lickert, H.; Takeuchi, J.K.; Von Both, I.; Walls, J.R.; McAuliffe, F.; Adamson, S.L.; Henkelman, R.M.; Wrana, J.L.; Rossant, J.; Bruneau, B.G. Baf60c is essential for function of BAF chromatin remodelling complexes in heart development. Nature 2004, 432, 107–112. [Google Scholar] [CrossRef]
  163. Lou, X.; Deshwar, A.R.; Crump, J.G.; Scott, I.C. Smarcd3b and Gata5 promote a cardiac progenitor fate in the zebrafish embryo. Development 2011, 138, 3113–3123. [Google Scholar] [CrossRef]
  164. Takeuchi, J.K.; Bruneau, B.G. Directed transdifferentiation of mouse mesoderm to heart tissue by defined factors. Nature 2009, 459, 708–711. [Google Scholar] [CrossRef] [PubMed]
  165. Sohni, A.; Mulas, F.; Ferrazzi, F.; Luttun, A.; Bellazzi, R.; Huylebroeck, D.; Ekker, S.C.; Verfaillie, C.M. TGFbeta1-induced Baf60c regulates both smooth muscle cell commitment and quiescence. PLoS ONE 2012, 7, e47629. [Google Scholar] [CrossRef]
  166. Chen, S.; Ding, Y.; Zhang, Z.; Wang, H.; Liu, C. Hyperlipidaemia impairs the circadian clock and physiological homeostasis of vascular smooth muscle cells via the suppression of Smarcd1. J. Pathol. 2014, 233, 159–169. [Google Scholar] [CrossRef] [PubMed]
  167. Chang, Z.; Zhao, G.; Zhao, Y.; Lu, H.; Xiong, W.; Liang, W.; Sun, J.; Wang, H.; Zhu, T.; Rom, O.; et al. BAF60a Deficiency in Vascular Smooth Muscle Cells Prevents Abdominal Aortic Aneurysm by Reducing Inflammation and Extracellular Matrix Degradation. Arterioscler. Thromb. Vasc. Biol. 2020, 40, 2494–2507. [Google Scholar] [CrossRef] [PubMed]
  168. Hinterseher, I.; Erdman, R.; Elmore, J.R.; Stahl, E.; Pahl, M.C.; Derr, K.; Golden, A.; Lillvis, J.H.; Cindric, M.C.; Jackson, K.; et al. Novel pathways in the pathobiology of human abdominal aortic aneurysms. Pathobiology 2013, 80, 1–10. [Google Scholar] [CrossRef]
  169. Zhao, G.; Lu, H.; Chang, Z.; Zhao, Y.; Zhu, T.; Chang, L.; Guo, Y.; Garcia-Barrio, M.T.; Chen, Y.E.; Zhang, J. Single-cell RNA sequencing reveals the cellular heterogeneity of aneurysmal infrarenal abdominal aorta. Cardiovasc. Res. 2021, 117, 1402–1416. [Google Scholar] [CrossRef]
  170. Shen, X.; Zhang, Y.; Xu, Z.; Gao, H.; Feng, W.; Li, W.; Miao, Y.; Xu, Z.; Zong, Y.; Zhao, J.; et al. KLF5 inhibition overcomes oxaliplatin resistance in patient-derived colorectal cancer organoids by restoring apoptotic response. Cell Death Dis. 2022, 13, 303. [Google Scholar] [CrossRef]
  171. Li, X.; He, Y.; Xu, Y.; Huang, X.; Liu, J.; Xie, M.; Liu, X. KLF5 mediates vascular remodeling via HIF-1alpha in hypoxic pulmonary hypertension. Am. J. Physiol. Lung Cell Mol. Physiol. 2016, 310, L299–L310. [Google Scholar] [CrossRef]
  172. Li, Y.; Zhang, B.; Xiang, L.; Xia, S.; Kucuk, O.; Deng, X.; Boise, L.H.; Dong, J.T. TGF-beta causes Docetaxel resistance in Prostate Cancer via the induction of Bcl-2 by acetylated KLF5 and Protein Stabilization. Theranostics 2020, 10, 7656–7670. [Google Scholar] [CrossRef] [PubMed]
  173. Ferguson, L.P.; Gatchalian, J.; McDermott, M.L.; Nakamura, M.; Chambers, K.; Rajbhandari, N.; Lytle, N.K.; Rosenthal, S.B.; Hamilton, M.; Albini, S.; et al. Smarcd3 is an epigenetic modulator of the metabolic landscape in pancreatic ductal adenocarcinoma. Nat. Commun. 2023, 14, 292. [Google Scholar] [CrossRef] [PubMed]
  174. Centore, R.C.; Sandoval, G.J.; Soares, L.M.M.; Kadoch, C.; Chan, H.M. Mammalian SWI/SNF Chromatin Remodeling Complexes: Emerging Mechanisms and Therapeutic Strategies. Trends Genet. 2020, 36, 936–950. [Google Scholar] [CrossRef] [PubMed]
  175. Xiao, L.; Parolia, A.; Qiao, Y.; Bawa, P.; Eyunni, S.; Mannan, R.; Carson, S.E.; Chang, Y.; Wang, X.; Zhang, Y.; et al. Targeting SWI/SNF ATPases in enhancer-addicted prostate cancer. Nature 2022, 601, 434–439. [Google Scholar] [CrossRef]
  176. Wei, Z.; Yoshihara, E.; He, N.; Hah, N.; Fan, W.; Pinto, A.F.M.; Huddy, T.; Wang, Y.; Ross, B.; Estepa, G.; et al. Vitamin D Switches BAF Complexes to Protect beta Cells. Cell 2018, 173, 1135–1149.e15. [Google Scholar] [CrossRef]
  177. Ahmed, N.S.; Gatchalian, J.; Ho, J.; Burns, M.J.; Hah, N.; Wei, Z.; Downes, M.; Evans, R.M.; Hargreaves, D.C. BRD9 regulates interferon-stimulated genes during macrophage activation via cooperation with BET protein BRD4. Proc. Natl. Acad. Sci. USA 2022, 119, e2110812119. [Google Scholar] [CrossRef]
  178. Du, J.; Liu, Y.; Wu, X.; Sun, J.; Shi, J.; Zhang, H.; Zheng, A.; Zhou, M.; Jiang, X. BRD9-mediated chromatin remodeling suppresses osteoclastogenesis through negative feedback mechanism. Nat. Commun. 2023, 14, 1413. [Google Scholar] [CrossRef]
  179. Wang, L.; Oh, T.G.; Magida, J.; Estepa, G.; Obayomi, S.M.B.; Chong, L.W.; Gatchalian, J.; Yu, R.T.; Atkins, A.R.; Hargreaves, D.; et al. Bromodomain containing 9 (BRD9) regulates macrophage inflammatory responses by potentiating glucocorticoid receptor activity. Proc. Natl. Acad. Sci. USA 2021, 118, e2109517118. [Google Scholar] [CrossRef]
  180. Saccone, V.; Consalvi, S.; Giordani, L.; Mozzetta, C.; Barozzi, I.; Sandona, M.; Ryan, T.; Rojas-Munoz, A.; Madaro, L.; Fasanaro, P.; et al. HDAC-regulated myomiRs control BAF60 variant exchange and direct the functional phenotype of fibro-adipogenic progenitors in dystrophic muscles. Genes Dev. 2014, 28, 841–857. [Google Scholar] [CrossRef]
  181. Zhang, P.; Li, L.; Bao, Z.; Huang, F. Role of BAF60a/BAF60c in chromatin remodeling and hepatic lipid metabolism. Nutr. Metab. 2016, 13, 30. [Google Scholar] [CrossRef]
  182. Meng, Z.X.; Tao, W.; Sun, J.; Wang, Q.; Mi, L.; Lin, J.D. Uncoupling Exercise Bioenergetics From Systemic Metabolic Homeostasis by Conditional Inactivation of Baf60 in Skeletal Muscle. Diabetes 2018, 67, 85–97. [Google Scholar] [CrossRef] [PubMed]
  183. Patsch, C.; Challet-Meylan, L.; Thoma, E.C.; Urich, E.; Heckel, T.; O’Sullivan, J.F.; Grainger, S.J.; Kapp, F.G.; Sun, L.; Christensen, K.; et al. Generation of vascular endothelial and smooth muscle cells from human pluripotent stem cells. Nat. Cell Biol. 2015, 17, 994–1003. [Google Scholar] [CrossRef] [PubMed]
  184. Hota, S.K.; Johnson, J.R.; Verschueren, E.; Thomas, R.; Blotnick, A.M.; Zhu, Y.; Sun, X.; Pennacchio, L.A.; Krogan, N.J.; Bruneau, B.G. Dynamic BAF chromatin remodeling complex subunit inclusion promotes temporally distinct gene expression programs in cardiogenesis. Development 2019, 146, dev174086. [Google Scholar] [CrossRef] [PubMed]
Cells 13 00168 g001
Figure 1. Transcriptional regulation at the CREs (cis-regulatory elements) of VSMC contractile genes. This diagram showcases the intricate network of interactions involving transcription regulators, such as chromatin remodelers, histone modifiers, transcription factors, and cofactors, at the CREs of the VSMC contractile genes. Chromatin remodelers that expose DNA regions by unwinding nucleosomes are central to the regulation. Several CArG elements, present at promoters and introns, bind to two SRFs and interact with myocardin. Within this region, there are binding sites for Smad2/3, NICD/RBPJ, and GATA. Retinoic acid signaling, crucial for VSMC differentiation, facilitates interactions between RAR/RXR and the SMARCD subunit of the SWI/SNF complexes. Other key contributors to VSMC differentiation include Prx1, Nkx3-2, PITX2, MEF2, and PIAS1. In addition, chromatin remodelers and transcription factors interact with histone modifiers, such as p300 and members of the NCOA family. Such complex interactions determine delicate and complicated regulation of contractile genes.
Figure 1. Transcriptional regulation at the CREs (cis-regulatory elements) of VSMC contractile genes. This diagram showcases the intricate network of interactions involving transcription regulators, such as chromatin remodelers, histone modifiers, transcription factors, and cofactors, at the CREs of the VSMC contractile genes. Chromatin remodelers that expose DNA regions by unwinding nucleosomes are central to the regulation. Several CArG elements, present at promoters and introns, bind to two SRFs and interact with myocardin. Within this region, there are binding sites for Smad2/3, NICD/RBPJ, and GATA. Retinoic acid signaling, crucial for VSMC differentiation, facilitates interactions between RAR/RXR and the SMARCD subunit of the SWI/SNF complexes. Other key contributors to VSMC differentiation include Prx1, Nkx3-2, PITX2, MEF2, and PIAS1. In addition, chromatin remodelers and transcription factors interact with histone modifiers, such as p300 and members of the NCOA family. Such complex interactions determine delicate and complicated regulation of contractile genes.
Cells 13 00168 g001
Cells 13 00168 g002
Figure 2. Temporal regulation of VSMC contractile gene expression. (A) Stem cell chromatin state: specific DNA regions are tightly wrapped into nucleosomes in stem cells. These regions are marked by repressive histone modifications, notably H4K20me3, H3K9me3, and H3K27me3, which play pivotal roles in maintaining gene silencing. (B) Activation of chromatin regions: during the transition to an active chromatin state, repressive histone marks are displaced, and active modifications emerge, such as H4Ac, H3K4Me2, H3K9Ac, H3K14Ac, and H3K79Me. Key players in this process include JMJD1a (H3K9me removal), SMYD2 (H3K4Me induction), and p300 (H3Ac and H4Ac induction). As histone acetylation weakens histone-DNA binding, pioneer factors can bind with nucleosomal DNA, subsequently recruiting either chromatin remodelers like the SWI/SNF complex or additional histone modifiers. (C) Transcription: the SWI/SNF complex introduces nucleosome-free regions, enabling TET2 to demethylate DNA. As a result, specific transcription factors bind to their respective elements. Subsequently, cofactors, including myocardin and CSRP, are recruited, promoting interactions between transcription regulators to promote transcription. (D) Dedifferentiation and transcriptional inactivation: in response to dedifferentiation signals like platelet-derived growth factor BB (PDGF-BB), the transcription of contractile genes was suppressed. In this situation, KLF4 binds to the G/C elements, while SRF partners with Elk1 rather than myocardin. Concurrently, HDACs and PRC are also recruited to deacetylate and induce methylation of H3K27. Additionally, PRDM6, PRMT5, and SUV39H1 mediate methylation at H4K20, H4R3 and H3R8, and H3K9, respectively. The loss of required transcription factors and DNA accessibility cease contractile gene transcription.
Figure 2. Temporal regulation of VSMC contractile gene expression. (A) Stem cell chromatin state: specific DNA regions are tightly wrapped into nucleosomes in stem cells. These regions are marked by repressive histone modifications, notably H4K20me3, H3K9me3, and H3K27me3, which play pivotal roles in maintaining gene silencing. (B) Activation of chromatin regions: during the transition to an active chromatin state, repressive histone marks are displaced, and active modifications emerge, such as H4Ac, H3K4Me2, H3K9Ac, H3K14Ac, and H3K79Me. Key players in this process include JMJD1a (H3K9me removal), SMYD2 (H3K4Me induction), and p300 (H3Ac and H4Ac induction). As histone acetylation weakens histone-DNA binding, pioneer factors can bind with nucleosomal DNA, subsequently recruiting either chromatin remodelers like the SWI/SNF complex or additional histone modifiers. (C) Transcription: the SWI/SNF complex introduces nucleosome-free regions, enabling TET2 to demethylate DNA. As a result, specific transcription factors bind to their respective elements. Subsequently, cofactors, including myocardin and CSRP, are recruited, promoting interactions between transcription regulators to promote transcription. (D) Dedifferentiation and transcriptional inactivation: in response to dedifferentiation signals like platelet-derived growth factor BB (PDGF-BB), the transcription of contractile genes was suppressed. In this situation, KLF4 binds to the G/C elements, while SRF partners with Elk1 rather than myocardin. Concurrently, HDACs and PRC are also recruited to deacetylate and induce methylation of H3K27. Additionally, PRDM6, PRMT5, and SUV39H1 mediate methylation at H4K20, H4R3 and H3R8, and H3K9, respectively. The loss of required transcription factors and DNA accessibility cease contractile gene transcription.
Cells 13 00168 g002
Cells 13 00168 g003
Figure 3. Composition of mammalian SWI/SNF complexes. The mammalian SWI/SNF chromatin remodeling complexes exist in three distinct types of assemblies: canonical BAF (cBAF), non-canonical BAF (ncBAF), and polybromo-associated BAF (PBAF). All three share several common subunits, such as the ATPase SMARCA2/4 and the ARP module (which includes ACTB, ACTL6A, and BCL7A/B/C). There are several differences between three types of assemblies. ARID1A/B and DPF1/2/3 only exist in cBAF. There are ARID2, BRD7, PHF10, and PBRM1 but no SS18/SS18L in PBAF. Non-canonical BAF possesses BICRA, BICRAL, and BRD9, and lacks SMARCC2, SMARCD2/3, SMARCB1, and SMARCE1.
Figure 3. Composition of mammalian SWI/SNF complexes. The mammalian SWI/SNF chromatin remodeling complexes exist in three distinct types of assemblies: canonical BAF (cBAF), non-canonical BAF (ncBAF), and polybromo-associated BAF (PBAF). All three share several common subunits, such as the ATPase SMARCA2/4 and the ARP module (which includes ACTB, ACTL6A, and BCL7A/B/C). There are several differences between three types of assemblies. ARID1A/B and DPF1/2/3 only exist in cBAF. There are ARID2, BRD7, PHF10, and PBRM1 but no SS18/SS18L in PBAF. Non-canonical BAF possesses BICRA, BICRAL, and BRD9, and lacks SMARCC2, SMARCD2/3, SMARCB1, and SMARCE1.
Cells 13 00168 g003
Table 1. Regulators of contractile gene transcription.
Table 1. Regulators of contractile gene transcription.
GeneCategoryFunctionEffect on Contractile Gene TranscriptionReference
SMARCA4Chromatin remodeler Mediate chromatin accessibilityDependent on interactors [42,43]
SMARCD3Chromatin remodelerMediate chromatin accessibilityPromotion[44]
SRFTranscription factorBind at CArG elementsDependent on interactors[8]
MyocardinCofactorInteract with SRFPromotion[8,41,45]
MRTFA/BCofactorInteract with SRFPromotion[41,46,47]
Smad2/3Transcription factorBind DNAPromotion[48]
GATA4/6Transcription factorBind DNAPromotion[49,50,51,52,53]
CSRP2CofactorInteract with SRF, GATA6Promotion[54,55]
NKX3-2Transcription factorBind DNAPromotion[52]
Prx1Transcription factorBind DNAPromotion[56]
PITX2Transcription factorBind DNAPromotion[57]
PIAS1Transcription factorBind DNAPromotion[58]
MEF2Transcription factorBind DNAPromotion[59]
Notch/RBPJTranscription factorBind DNAPromotion[60,61]
KLF4Transcription factorBind G/C repressor elementRepression[9,62,63,64,65]
Elk1CofactorInteract with SRFRepression[47,63,64]
p300Histone acetyltransferaseIncrease histone acetylationPromotion[51,66,67]
HDACHistone deacetylaseDecrease Histone modificationRepression[47,63,64]
SMYD2Histone lysine methyltransferaseIncrease H3K4me1, H3K4me3Promotion[68]
JMJD1AHistone demethylaseDecrease H3K9me2Promotion[69]
WDR5CofactorIncrease H3K4me1, H3K4me3Promotion[70]
PRDM6Histone lysine methyltransferaseIncrease H4K20me2Repression[71,72]
SUV39H1Histone lysine methyltransferaseIncrease H3K9me3Repression[73]
EZH2Histone lysine methyltransferaseIncrease H3K27me3Repression[74,75]
TET2Methylcytosine dioxygenaseDNA demethylationPromotion[76,77]
PRMT5Histone arginine methyltransferaseH3R8me2, H4R3me2Repression[78]
Table 2. SWI/SNF complex subunit HUGO name and common name.
Table 2. SWI/SNF complex subunit HUGO name and common name.
HUGO NameCommon Name
SMARCC1BAF155, SRG3
SMARCC2BAF170
SMARCD1/2/3BAF60A/B/C
SMARCB1BAF47, INI1
SMARCE1BAF57
ARID1A/BBAF250A/B
ARID2BAF200
PHF10BAF45A
DPF1/2/3BAF45B/D/C
BICRA/BICRALGLTSCR1/GLTSCR1L
SMARCA4BRG1
SMARCA2BRM
ACTL6A/BBAF53A/B
PBRM1BAF180
Table 3. SWI/SNF subunit-interacting proteins validated by GST pull-down assay.
Table 3. SWI/SNF subunit-interacting proteins validated by GST pull-down assay.
InteractorSWI/SNF SubunitReference
ARSMARCC1[128]
CBPSMARCA4[129]
ERαSMARCD1[130]
ERαSMARCD3[131]
FOSSMARCD1[132]
JUNSMARCD1[132]
JUNSMARCD3[131]
MYCSMARCA2[133]
MYCSMARCA4[133]
MYCSMARCB1[133]
MYCSMARCE1[133]
MyocardinSMARCD3[134]
NCOA1ARID1[135]
NCOA1SMARCC1[128]
NCOA1SMARCE1[135,136]
Nkx2-5SMARCD3[134]
NR3C1/GRSMARCD1[130]
NR3C1/GRSMARCE1[130]
PCG1αSMARCD1[123]
PPARγSMARCD1[123]
PPARγSMARCD3[131]
PRMT5SMARCB1[133]
PRMT5SMARCE1[133]
RARSMARCD3[135]
RBP-JSMARCD3[137]
RORαSMARCD3[131]
RXRSMARCD3[131,135]
SREBP1αSMARCD3[131]
Tbx5SMARCD3[134,137]
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Liu, H.; Zhao, Y.; Zhao, G.; Deng, Y.; Chen, Y.E.; Zhang, J. SWI/SNF Complex in Vascular Smooth Muscle Cells and Its Implications in Cardiovascular Pathologies. Cells 2024, 13, 168. https://doi.org/10.3390/cells13020168

AMA Style

Liu H, Zhao Y, Zhao G, Deng Y, Chen YE, Zhang J. SWI/SNF Complex in Vascular Smooth Muscle Cells and Its Implications in Cardiovascular Pathologies. Cells. 2024; 13(2):168. https://doi.org/10.3390/cells13020168

Chicago/Turabian Style

Liu, Hongyu, Yang Zhao, Guizhen Zhao, Yongjie Deng, Y. Eugene Chen, and Jifeng Zhang. 2024. "SWI/SNF Complex in Vascular Smooth Muscle Cells and Its Implications in Cardiovascular Pathologies" Cells 13, no. 2: 168. https://doi.org/10.3390/cells13020168

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop