Origins of Aortic Coarctation: A Vascular Smooth Muscle Compartment Boundary Model

Christina L. Greene; Geoffrey Traeger; Akshay Venkatesh; David Han; Mark W. Majesky

doi:10.20944/preprints202412.2407.v1

Submitted:

26 December 2024

Posted:

30 December 2024

You are already at the latest version

Abstract

Compartment boundaries divide the embryo into segments with distinct fates and functions. In the vascular system, compartment boundaries organize endothelial cells into arteries, capillaries, and veins that are the fundamental units of a circulatory network. For vascular smooth muscle cells (SMCs) such boundaries produce mosaic patterns of investment based on embryonic origins with important implications for the non-uniform distribution of vascular disease later in life. Morphogenesis of blood vessels requires vascular cell movements within compartments as highly-sensitive responses to changes in fluid flow shear stress and wall strain. These movements underlie remodeling of primitive plexuses, expansion of lumen diameters, regression of unused vessels, and building multilayered artery walls. Although loss of endothelial compartment boundaries can produce arterial-venous malformations, little is known about the consequences of mislocalization or failure to form SMC origin-specific boundaries during vascular development. We propose that the failure to establish a normal compartment boundary between cardiac neural crest-derived SMCs of the 6th pharyngeal arch artery (future ductus arteriosus) and paraxial mesoderm-derived SMCs of the dorsal aorta in mid-gestation embryos leads to aortic coarctation observed at birth. This model raises new questions about the effects of fluid flow dynamics on SMC investment and the formation of SMC compartment borders during pharyngeal arch artery remodeling and vascular development.

Keywords:

vascular smooth muscle

;

pharyngeal arch arteries

;

congenital heart disease

;

remodeling

;

ductus arteriosus

Subject:

Biology and Life Sciences - Cell and Developmental Biology

Introduction

Coarctation of the aorta (CoA) is a form of congenital heart disease that is characterized by a structural narrowing of the aorta that restricts blood flow and thereby impairs normal growth and development of the newborn (Figure 1). Blood flow impairment is the result of a constriction of the descending thoracic aorta (AO) usually near the ductal ostia and is thought to be due to vascular smooth muscle cells (SMCs) from the ductus arteriosus (DA) mislocalized within the wall of the AO. CoA is one of the most frequent presentations of congenital heart disease (CHD), and the most common congenital defect of the aorta affecting 1 per 2,500 live births [1,2]. First diagnosed over two hundred years ago, CoA has been surgically repaired for the last 50 years yet we have little understanding of the underlying pathobiology. Genome sequencing of CoA and other CHD patient cohorts over the last decade has identified a handful of associated genes, but this effort has failed to establish a unifying mechanism for CoA [3,4,5,6,7,8].

Development of the Ductus Arteriosus

In searching for the origins of CoA, one must look to the formation and remodeling of the pharyngeal arch arteries (PAAs) in midgestation embryos. This is because the DA arises from flow-sensitive remodeling of the 6th PAA in vascular development [9,10] (Figure 2) and because CoA can be detected by echocardiography in utero [11,12]. Although anatomically continuous with the aortic media, DA-SMCs in the ductal media do not normally extend beyond their junction with the descending thoracic aorta [9,13,14]. As the 6th PAA begins to assemble its complement of SMCs, two types of SMC progenitor cells are present in the pharyngeal arch complex: cardiac neural crest (CNC) and second heart field (SHF) [15]. Endothelial cells of both the 4th arch and 6th arch arteries are derived from SHF progenitors [16]. However, fate mapping studies have shown that medial SMCs of the 6th PAA originate exclusively from cardiac neural crest cells (CNCs) [17]. By contrast, medial SMCs in the dorsal aorta at the level of the 6th PAA junction arise from paraxial mesoderm (PM) [18]. Therefore, a border is formed at the interface between CNC-derived SMCs in the 6th PAA and PM-derived SMCs in the dorsal aorta. Elzenga et al. described the normal DA-AO junction as a “fish tail-like” insertion of DA-SMCs within the AO media involving up to one-third of the AO circumference in a newborn infant [19] (Figure 3). The formation of an insertion junction, therefore, means that the DA-SMC interface with AO-SMCs does not exactly correspond to the anatomical landmark of a DA (or later, LA) connection to the AO. Nevertheless, the juxtaposition of CNC-derived DA-SMCs and PM-derived AO-SMCs appears to result in a SMC origin-dependent border that, in effect, acts as a sorting mechanism keeping DA-SMCs confined to the limits of the ductal insertion and maintaining AO-SMCs in a separate anatomical compartment [19]. The necessity of a sorting mechanism becomes evident when the baby takes its first breath at birth. Loss of placental-derived prostaglandins together with an increase in arterial oxygen tension resulting from nascent blood flow to the lungs initiates closure of the DA, a critical event required for adequate perfusion of the now functioning lungs [14,20,21]. The directly adjacent AO-SMCs in the aortic media exhibit none of these oxygen-dependent closure activities. While the DA closes, the AO media continues to increase circumferential growth in proportion to increasing cardiac output required for postnatal growth of the neonate. The mechanisms that specify 6th arch SMCs to acquire a unique ductal SMC identity while, in close proximity (Figure 2, note the proximity of IV to VI PAAs to each other), dorsal aorta SMCs acquire an entirely different identity and developmental trajectory remain an intriguing mystery yet to be solved.

Compartment Boundaries in Embryonic Development

Formation of tissue boundaries that divide the embryo into compartments with distinct cell fates is one of the most fundamental processes in development [22,23,24,25,26,27]. For example, compartment boundaries separate adjacent rhombomeres during hindbrain development [28,29,30], somites during axial patterning [31,32,33], septation of the cardiac interventricular septum [34,35], pattern dorsal-ventral domains of the limb bud [36,37], and delineate imaginal discs during development and regeneration in Drosophila [38,39]. Compartment boundaries are formed when cells with different identities encounter each other [34,35,38,40]. A lack of mixing at such borders is a direct result of compartment-specific transcriptomes and maintenance of different cell identities and cell fate potentials. Studies of the Drosophila embryo have provided a conceptual framework for morphogenesis that includes segmentation, acquisition of cell identity, compartmentation, and pattern formation [40,41]. An important principle that has emerged is that the early embryo is organized into lineage blocks called compartments [38]. Two different lineage blocks that form adjacent to each other are separated by a compartment border that is maintained as a function of different cell identities on either side of the border [38]. If an imaginal disc compartment border is disrupted, for example, then the two compartments regenerate the missing cells, reestablish compartment-specific cell identity, and reorganize the border between them [42]. What mechanisms actually produce compartment borders have been the subject of much investigation [40]. These studies have produced three general models: the differential cell adhesion model [43,44,45], the actin-myosin cortex contraction model [40,46], and the contact repulsive model focused on heterotypic cell-cell contacts [25,27].

Compartment Boundaries in Vascular Development

In the vascular system, three types of developmental borders have been identified. One is the boundary separating arterial and venous endothelial cells [47,48,49] or arterial and venous SMCs [48,50,51] marked by guidance molecules that mediate contact repulsive signaling between adjacent heterotypic cells. A second type of border separates aortic root endothelium of second heart field origin [16] from coronary artery endothelium of sinus venosus origin [52]. This represents an artery-artery EC boundary and, once established, was found to be stable out to at least postnatal day 28 in the mouse [52]. A third type of compartment boundary separates SMCs of different developmental origins in a common artery wall [53,54,55,56,57,58,59]. Although multiple descriptions of these SMC lineage-specific compartments have been made in the studies cited above and elsewhere, little is known about SMC compartment borders themselves, the molecules & mechanisms that produce these borders, the compartmentalization of SMC functions they maintain, or the consequences of a loss of border function in vascular dysmorphogenesis and disease. Drawing clues from compartment boundary studies in Drosophila and other model organisms [25,60,61], we will discuss these questions with a particular focus on the DA-SMC junction with AO-SMCs in the proximal descending thoracic aorta.

Repulsive Guidance Molecule Signaling

Chemo- or contact-repulsive signaling is one of the key border-forming mechanisms employed in embryonic development [51,62,63]. Contact-repulsive guidance molecules are expressed by CNC-derived SMC progenitors enabling them to precisely navigate to the 6th PAA during vascular development [51,64,65,66,67,68,69,70]. Likewise, migration of paraxial mesoderm-derived AO-SMC progenitors to the dorsal aorta is also guided by repulsive signaling [31]. During neural crest cell migration in Xenopus embryos, the homophilic calcium-dependent cell-cell adhesion molecule cadherin-11 was found to be necessary for contact inhibition of movement [71]. This may reflect the cooperative activity of cadherin 11-dependent cell-cell adhesion to form close contacts between adjacent cells so that contact-repulsive mechanisms are efficiently engaged by those cells [72]. That engagement may initiate the contact-mediated collapse of cell protrusions oriented in the direction of migration [73]. A similar role for N-cadherin in CNC migration has been reported [74]. Upon contact of DA-SMCs with AO-SMCs, we suggest that engagement of contact-repulsive guidance molecules stops migration of both types of vascular SMCs, initiates border-dependent cytoskeletal reorganization [75], and creates a barrier that prevents mixing of DA-SMCs and AO-SMCs. The abruptness of the DA-AO SMC border [76] (Figure 3) implies a boundary-forming mechanism similar to that which separates arterial and venous endothelial cells in early vascular development [47,49,77,78]. The report of a Semaphorin-3D (SEMA3D) loss of function mutation in human CoA is consistent with this hypothesis [6].

It is instructive to consider SMC border formation at another site in the developing vascular system. Lineage- and fate-mapping studies in mouse embryos have shown that SMCs in the aortic root are derived from progenitor cells of either cardiac neural crest or second heart field origin [79]. Sawada et al. showed that these two SMC origins reside in distinct spatial domains in the ascending thoracic aorta with little or no detectable mixing [56]. Cardiac neural crest-derived SMC progenitors migrate into the pharyngeal arch complex and reach the 4th arch artery wall earlier than second heart field derived SMC progenitors and form the first few inner layers of the aortic root and ascending aorta [79]. Slightly delayed in time, second heart field-derived cells also migrate to the developing aorta and constitute the major SMC type in the aortic root, and form the outer medial layers of the ascending aorta [53,55,56,79]. These two types of aortic SMCs express similar levels of SMC contractile marker genes and cytoskeletal genes but they retain SMC lineage-specific differences in TGF-beta receptor-dependent growth and transcriptional responses [80,81], baseline gene expression patterns [59,82], and accessible chromatin sites identified by ATAC-seq analysis [83]. Likewise, SMCs from different embryonic origins express different levels of extracellular matrix genes in vivo [84] and in vitro [59].

Remodeling of the Pharyngeal Arch Artery Complex

Endothelial cells: Developing blood vessels are highly sensitive to blood flow variations [15,85,86,87,92,95]. Studies in multiple species have shown that changes in blood flow rates are major morphogenic forces acting on endothelial cells directing their growth, migration, and cell-cell interactions [10,85,86,87,88,89,90,91]. Fluid shear stress on the endothelial cell surface regulates the assembly of endothelial cell junctions, cytoskeletal organization, polarity, and arterial-venous identity [85,92,93,94,95]. In particular, the complex remodeling of the PAAs is, in large part, determined by developmental changes in cardiac function, blood flow dynamics, and endothelial shear stress forces [10,89,96]. Using fluorescent dye injection, pulsed Doppler velocity recordings, and micro-CT scans, Wang et al. showed that a shift occurs in flow velocity and wall shear stress (WSS) in the PAA complex between stages HH18 and HH24 in the chick embryo [97]. That shift is from a higher WSS in PAA-3 at HH18 to a higher WSS in PAA-4 at HH24 [87,97,98,99]. These developmental shifts in flow velocity and WSS are recorded as corresponding diameter changes in the respective PAAs during the remodeling period [87,97,99].

There is considerable evidence that defects in heart or valve development and resultant changes in blood flow rate, flow distribution, and endothelial shear stress responses during cardiovascular development produce congenital heart defects [98,99,100]. There is also evidence that both endothelial cells and SMCs are surprisingly motile within the vessel wall during the early stages of vascular development [91,101]. It therefore follows that the positions of the 6th PAA ostia with the dorsal aorta and pulmonary artery, the investment of this vessel with SMCs, or the SMC origin-specific border between 6th PAA SMCs and dorsal aorta (AO) SMCs may be sensitive to disturbances in normal blood flow dynamics in this critical period of PAA remodeling.

Smooth muscle cells: For example, the murine left pulmonary artery (PA) at birth consists of an inner layer of endothelial cells, two layers of SMCs and an outer layer of adventitia [101]. The first layer of PA-SMCs emerges from PDGFRβ-positive mesenchyme surrounding the developing PA wall at around E11.5. As SMC differentiation markers appear in the first layer of SMCs, expression of PDGFRβ is down-regulated in these cells [101]. The second layer of PA-SMCs arises from two sources. The major fraction originates from surrounding PDGFRβ-positive mesenchyme in a sequence much like the first layer. Examination of nuclear morphology and the orientation of SMC mitotic spindles during PA wall formation showed that during E11.5 to E13.5, the axis of division of SMCs in the first layer was predominantly longitudinal (>75%) thus contributing daughter cells to growth in length of the PA. At E14.5, however, the axis of division shifts so that the majority of first layer SMCs divide circumferentially while cells in the second layer of media and in the adventitia continue to divide in the longitudinal orientation. Therefore, circumferential orientation of division of first layer SMCs contributes daughter cells to expand the second layer. Using a single low dose of tamoxifen in Myh11CreERT2; mTmG mice administered at E11.5 to label individual SMCs, the position of clones originating from inner layer SMCs could be mapped in the PA wall from E13.5 to E18.5. These experiments showed that individual cells within a clone often dispersed widely from each other, intermixing with unlabeled cells, and populating both first layer and second layer SMCs. While migrating extensively within the media both longitudinally and circumferentially, labeled SMCs did not enter the adventitia or the intima. These results illustrate two important principles about early vascular development. The first is that cells within nascent blood vessel walls are not stationary, but exhibit considerable motility within the developing vessel or primitive vascular network. The second is that barriers, or borders, exist that define compartments into which entry by cells from outside the compartment is prevented. As described in a previous section, compartments and the borders that define them are fundamental to tissue morphogenesis and it is no surprise to also find them in developing blood vessels.

The Role of Hemodynamics in PAA Remodeling

The shrunken head (shru) mutation was identified in a forward genetic screen in mice and found to be localized within the titin gene [10]. Shru produced a hypomorphic allele and embryos were defective in development of myocardial cells as revealed by greatly reduced numbers of myofibrils in these cells, reduced cardiac contractile activity, and pronounced delay in the onset of circulation through the vascular system. Shru mutants had little or no blood flow from E8.5 to E9.5 and then exhibited a weak initiation of blood flow from E9.5 to embryonic death at E11.5 [10]. A variety of embryonic and extra-embryonic vascular defects were observed in shru mutant embryos including defective endothelial cell-cell junctions, abnormal endothelial cell protrusions, intermittent lack of lumen formation, extensive areas of hypoxia, and increased VEGFA gene expression. As titin is expressed in cardiac myocytes but not in endothelial cells, the vascular defects in shru embryos are most likely indirect and due to greatly reduced or absent blood flow through the developing vascular system during angiogenesis, plexus reorganization, and PAA remodeling stages. The authors conclude that fluid biomechanical forces produced from the very onset of cardiac contractions are required to orchestrate multiple aspects of endothelial structure and function as well as vascular network formation and remodeling [10]. It will be of interest to learn if the formation or position of SMC origin-specific borders is disrupted in shru embryos.

The outcome of PAA remodeling is also a reflection of the overall control of left-right asymmetry (laterality) during embryonic development [98,102,103]. Control of laterality for internal organs is a function of nodal signaling [102]. Nodal induces the expression of PITX2, a transcription factor that controls left-right asymmetry in cardiac development and aortic arch remodeling [103,104]. Asymmetric expression of Pitx2 is controlled by the asymmetric enhancer element (ASE) in the Pitx2 locus [102,105]. Mice lacking the ASE develop right isomerism, cardiac developmental defects, and randomized laterality of PAAs [105]. While asymmetry of 6th PAA remodeling is randomized in mice lacking the ASE, the 4th PAA was not affected [98]. Laterality of the aortic arch is also randomized in these mice and is dependent upon which side of the paired 6th PAAs undergoes regression. In wild-type embryos, echocardiography studies showed that at E11.5 blood flow through the left dorsal aorta was similar to that in the right dorsal aorta, whereas by E12.0 blood flow was significantly higher in the left compared to the right-side dorsal aorta [106]. Surprisingly, Pitx2 is not expressed in or near the 6th PAA or the dorsal aorta. Rather, it is strongly expressed in the second heart field, the myocardium, and the left wall of the outflow tract [98,103]. In normal heart development, formation of the spiral septum of the outflow tract aligns the ventricles with the great arteries and rotation of the arterial pole produces changes in the right 6th arch artery structure that reduces blood flow and favors regression of the right side [107]. In ASE-deficient mice, defects in outflow tract morphogenesis result in altered blood flow distribution to the right and left sides of the 6th PAA leading to randomized laterality of the 6th PAA [98,99]. These studies show how closely related rotational development of the outflow tract is to normal asymmetric remodeling of the 6th PAA and ultimate formation of the DA. As indicated above for shru embryos, it will be interesting to determine if the position of the SMC origin-dependent border is disrupted if blood flow distribution through the right versus left 6th PAAs is altered due to defects in cardiac valve or outflow tract development.

Role of Hemodynamics in Vascular Smooth Muscle Cell Investment

As discussed above, PAA remodeling is highly sensitive to changes in blood flow. Evidence from multiple experimental models shows that investment of nascent endothelial tubes with mural cells (SMCs and pericytes) is also a blood flow-responsive process [108,109,110]. The release of PDGFB, among other factors, by endothelial cells is stimulated by increased fluid shear stress acting on the endothelial cell surface [111,112]. The abluminal secretion of PDGFB [113] and the activation of nearby PDGFRβ-expressing SMC progenitor cells are key steps in the investment process [114,115,116,117,118]. The effects of PDGFRβ signaling involve stimulation of SMC progenitor cell chemotaxis [112], migration [111], and localized cell proliferation in vivo [114]. Further analysis suggests two models for the investment process: (a) de novo formation of SMCs from surrounding undifferentiated mesenchymal progenitor cells, or (b) migration of SMCs from a pre-existing pool of SMCs usually upstream of the site of investment [110,114,115,116,119,120,121]. One question that is not addressed by these previous studies is how SMC investment proceeds at specific sites in the vascular system where SMCs arising from two different embryonic lineages are involved. What cell dynamics occur when cardiac neural crest-derived SMCs and paraxial mesoderm- or second heart field-derived SMCs encounter each other during assembly of the vessel wall? In any case, investment models must account for the apparent lack of mixing at SMC origin-dependent borders [53,54,122].

Interactions Between Different Types of Vascular SMC Progenitors

Investment of PAAs with SMCs exhibits the ability to compensate for defects in the number of available CNC-derived SMC progenitor cells. Alexander et al., reported that in mice made Smad4-deficient in the CNC-lineage using Sox10iCre, most of the CNC-derived SMC progenitors in the pharyngeal arch mesenchyme were lost due to cell death [123]. In the absence of this dominant source of PAA SMCs, a non-CNC source of locally-available SMC progenitors was observed to rescue the investment process so that the 4th and 6th PAAs developed and remodeled normally [123]. Although not tested directly, it is reasonable to assume that this non-CNC source of SMC progenitors was second heart field-derived progenitor cells which are also present in the PA mesenchyme [123]. This finding suggests the possibility that CNC-derived SMC progenitors normally suppress the SMC fate of available non-CNC progenitors providing a glimpse of the normal cross-talk between SMCs of different embryonic origins at compartment borders. If investment itself is a blood flow-dependent process, then the communication between SMC progenitors as well as the final structure or position of SMC origin-dependent borders may also be influenced by blood flow dynamics.

A Smooth Muscle Compartment Boundary Model for CoA

We propose a vascular SMC compartment boundary model for the pathogenesis of CoA. This model suggests that CoA is the consequence of a failure to form a SMC origin-specific compartment border in its normal position at the ductal ostia (Figure 4). We propose that disturbed blood flow during remodeling of the 6th PAA, either due to direct changes (somatic genetic or epigenetic) in the 6th PAA cells themselves, or indirect hemodynamic effects due to altered cardiac contractile activity, cardiac valve structure or function, or outflow tract defects could lead to an altered position or absence of the DA-SMC junction with AO-SMCs at the ductal ostia thus allowing DA-SMCs to invade the aortic wall (Figure 4). This would explain the localization of a majority of CoA lesions to the ductal-aorta junction and the clinical observation that CoA can be detected by fetal echo before birth [11,12]. It would also explain the frequent co-occurrence of CoA with bicuspid aortic valve and other cardiac congenital defects that would disturb blood flow during PAA remodeling [1,10,98,124,125]. This model for the origin for CoA raises important new questions: (1) if the DA-AO compartment border is the consequence of an interface of SMCs with different identities, similar to other developmental borders, then what molecules and mechanisms are responsible for the specification of distinct identities of DA-SMCs vs AO-SMCs during vascular development? (2) what cell surface molecules and signaling pathways normally act to prevent mixing of DA-SMCs and AO-SMCs at the ductal-aorta interface? and (3) what mechanisms disrupt the expression, function, or localization of these border-forming molecules during the pathogenesis of CoA? Given that vascular SMC origins are far more diverse than just CNC and SHF, the answers to these questions and the molecular insights gained may provide a better understanding of vascular pathologies at other sites within the mosaic vascular system as a whole.

Author Contributions

MWM and CLG wrote the article, GT contributed to preparation of the figures, and all authors made critical input and edits during preparation of the manuscript.

Funding

This work was supported by a Programs of Distinction grant from the Heart Center, Division of Pediatric Cardiology, Seattle Children’s Hospital, Seattle, WA. Additional support was provided by the Loie Power Robinson Stem Cell & Regenerative Medicine fund, Seattle Children’s Research Institute, and the Seattle Children’s Hospital Foundation. MWM received partial support from NIH grant HL-121877.

Acknowledgements

We thank our colleagues in the Heart Center, Seattle Children’s Hospital, for helpful discussions. We are grateful for help with preparation of this manuscript from Abigail Tsai.

Conflicts of Interest

There are no conflicts of interest to disclose.

References

Hoffman JI, Kaplan S. The incidence of congenital heart disease. J Am Coll Cardiol 2002, 39, 1890–1900. [Google Scholar] [CrossRef] [PubMed]
Mai CT, Isenburg JL, Canfield MA et al. National population-based estimates for major birth defects, 2010-2014. Birth Defects Res 2019, 111, 1420–1435. [Google Scholar] [CrossRef] [PubMed]
Tagariello A, Breuer C, Birkner Y et al. Functional null mutations in the gonosomal homologue gene TBL1Y are associated with non-syndromic coarctation of the aorta. Curr Mol Med 2012, 12, 199–205. [Google Scholar] [CrossRef] [PubMed]
Freylikhman O, Tatarinova T, Smolina N et al. Variants in the NOTCH1 gene in patients with aortic coarctation. Congenit Heart Dis 2014, 9, 391–396. [Google Scholar] [CrossRef] [PubMed]
Moosmann J, Uebe S, Dittrich S, Rüffer A, Ekici A, Toka O. Novel loci for non-syndromic coarctation of the aorta in sporadic and familial cases. PLoS One 2015, 10, e0126873. [Google Scholar] [CrossRef] [PubMed]
Sanchez-Castro M, Pichon O, Briand A, Poulain D, Gournay V, David A, Le Caignec C. Disruption of the SEMA3D gene in a patient with congenital heart defects. Human Mutation 2015, 36, 30–33. [Google Scholar] [CrossRef] [PubMed]
Bjornsson T, Thorolfsdottir RB, Sveinbjornsson G et al. A rare missense mutation in MYH6 associates with non-syndromic coarctation of the aorta. Eur Heart J 2018, 9, 3243–3249. [Google Scholar] [CrossRef] [PubMed]
Donadille B and Christin-Maitre, S. Heart and Turner syndrome. Ann Endocrinol (Paris) 2021, 82, 135–140. [Google Scholar] [CrossRef] [PubMed]
Hiruma T, Nakajima Y, Nakamura H. Development of pharyngeal arch arteries in early mouse embryo. J Anat 2002, 201, 15–29. [Google Scholar] [CrossRef] [PubMed]
May SR, Stewart NJ, Chang W, Peterson AS. A Titin mutation defines roles for circulation in endothelial morphogenesis. Dev Biol 2004, 270, 31–46. [Google Scholar] [CrossRef] [PubMed]
Franklin O, Burch M, Manning N, Sleeman K, Gould S, Archer N. Prenatal diagnosis of coarctation of the aorta improves survival and reduces morbidity. Heart 2002, 87, 67–69. [Google Scholar] [CrossRef] [PubMed]
Houshmandi MM, Eckersley L, Fruitman D, Mills L, Power A, Hornberger LK. Fetal diagnosis is associated with improved perioperative condition of neonates requiring surgical intervention for coarctation. Pediatr Cardiol 2021, 42, 1504–1511. [Google Scholar] [CrossRef] [PubMed]
Waldo KL, Kirby ML. Cardiac neural crest contribution to the pulmonary artery and sixth aortic arch artery complex in chick embryos aged 6 to 18 days. Anat Rec 1993, 237, 385–399. [Google Scholar] [CrossRef] [PubMed]
Bökenkamp R, DeRuiter MC, van Munsteren C, Gittenberger-de Groot AC. Insights into the pathogenesis and genetic background of patency of the ductus arteriosus. Neonatology 2010, 98, 6–17. [Google Scholar] [CrossRef] [PubMed]
Gittenberger-de Groot AC, Peterson JC, Wisse LJ, Roest AAW, Poelmann RE, Bökenkamp R, Elzenga NJ, Kaxekamp M, Bartelings MM, Jongbloed MRM, DeRuiter MC. Pulmonary ductal coarctation and left pulmonary artery interruption; pathology and role of neural crest and second heart field during development. PLoS One 2020, 15, e0228478. [Google Scholar] [CrossRef] [PubMed]
Wang X, Chen D, Chen K, Jubran A, Ramirez A, Astrof S. Endothelium in the pharyngeal arches 3,4 and 6 is derived from the second heart field. Dev Biol 2017, 421, 108–117. [Google Scholar] [CrossRef] [PubMed]
Jiang X, Rowitch DH, Soriano P, McMahon AP, Sucov HM. Fate of the mammalian cardiac neural crest. Development 2000, 127, 1607–1616. [Google Scholar] [CrossRef] [PubMed]
Wasteson P, Johansson BR, Jukkola T, Breuer S, Akyürek LM, Partanen J, Lindahl P. Developmental origin of smooth muscle cells in the descending aorta in mice. Development 2008, 135, 1823–1832. [Google Scholar] [CrossRef] [PubMed]
Elzenga NJ, Gittenberger-de Groot AC. Localised coarctation of the aorta. An age dependent spectrum. Br Heart J 1983, 49, 317–323. [Google Scholar] [CrossRef] [PubMed]
Rabinovitch, M. Cell-extracellular matrix interactions in the ductus arteriosus and perinatal pulmonary circulation. Semin Perinatol 1996, 20, 531–541. [Google Scholar] [CrossRef] [PubMed]
Bentley RET, Hindmarch CCT, Dunham-Snary KJ, Snetsinger B, Mewburn JD, Thébaud A, Lima PD, Thébaud B, Archer SL. The molecular mechanisms of oxygen-sensing in human ductus arteriosus smooth muscle cells: A comprehensive transcriptome profile reveals a central role for mitochondria. Genomics 2021, 113, 3128–3140. [Google Scholar] [CrossRef] [PubMed]
Dahmann C, Basler K. Compartment boundaries: at the edge of development. Trends Genet 1999, 15, 320–326. [Google Scholar] [CrossRef] [PubMed]
Brand-Saberi B, Christ B. Evolution and development of distinct cell lineages derived from somites. Curr Top Dev Biol 2000, 48, 1–42. [Google Scholar] [CrossRef] [PubMed]
Groves AK, LaBonne C. Setting appropriate boundaries: fate, patterning and competence at the neural plate border. Dev Biol 2014, 389, 2–12. [Google Scholar] [CrossRef] [PubMed]
Fagotto, F. The cellular basis of tissue separation. Development 2014, 141, 3303–3318. [Google Scholar] [CrossRef] [PubMed]
Wang J, Dahmann C. Establishing compartment boundaries in Drosophila wing imaginal discs: An interplay between selector genes, signaling pathways and cell mechanics. Semin Cell Dev Biol 2020, 107, 161–169. [Google Scholar] [CrossRef] [PubMed]
Fagotto, F. Cell sorting at embryonic boundaries. Sem Cell Dev Biol 2020, 107, 126–129. [Google Scholar] [CrossRef] [PubMed]
Maves L, Jackman W, Kimmel CB. FGF3 and FGF8 mediate a rhombomere 4 signaling activity in the zebrafish hindbrain. Development 2002, 129, 3825–3837. [Google Scholar] [CrossRef] [PubMed]
Cooke JE, Kemp HA, Moens CB. EphA4 is required for cell adhesion and rhombomere-boundary formation in the zebrafish. Curr Biol 2005, 15, 536–542. [Google Scholar] [CrossRef] [PubMed]
Kiecker C, Lumsden A. Compartments and their boundaries in vertebrate brain development. Nat Rev Neurosci 2005, 6, 553–564. [Google Scholar] [CrossRef] [PubMed]
Tannahill D, Cook GM, Keynes RJ. Axon guidance and somites. Cell Tissue Res 1997, 290, 275–283. [Google Scholar] [CrossRef] [PubMed]
Takahashi Y, Koizumi K, Takagi A, Kitajima S, Inoue T, Koseki H, Saga Y. Mesp2 initiates somite segmentation through the Notch signalling pathway. Nat Genet 2000, 25, 390–396. [Google Scholar] [CrossRef] [PubMed]
Watanabe T, Sato Y, Saito D, Tadakoro R, Takahashi Y. EphrinB2 coordinates the formation of a morphological boundary and cell epithelialization during somite segmentation. Proc Natl Acad Sci USA 2009, 106, 7467–7472. [Google Scholar] [CrossRef] [PubMed]
Franco D, Meilhac SM, Christoffels VM, Kispert A, Buckingham M, Kelley RG. Left and right ventricular contributions to the formation of the interventricular septum in the mouse heart. Dev Biol 2006, 294, 366–375. [Google Scholar] [CrossRef] [PubMed]
Kathiriya IS, Dominguez MH, Ro KS, Muncie-Vasic JM, Devine WP, Hu KM, Hota SK, Garay BI, Quintero D, Goyal P, Matthews MN, Thomas R, Sukonnik T, Miguel-Perez D, Winchester S, Brower EF, Forjaz A, Wu PH, Wirtz D, Kiemen AL, Bruneau BG. A disrupted compartment boundary underlies abnormal cardiac patterning and congenital heart defects. bioRxiv, 5789. [CrossRef] [PubMed]
Kimmel RA, Turnbull DH, Blanquet V, Wurst W, Loomis CA, Joyner AL. Two lineage boundaries coordinate vertebrate apical ectodermal ridge formation. Genes Dev 2000, 14, 1377–1389. [Google Scholar] [CrossRef] [PubMed]
Qiu Q, Chen H, Johnson RL. Lmx1b-expressing cells in the mouse limb bud define a dorsal mesenchymal lineage compartment. Genesis 2009, 47, 224–233. [Google Scholar] [CrossRef] [PubMed]
Morata G, Herrera SC. Cell reprogramming during regeneration in Drosophila: transgression of compartment boundaries. Curr Opin Genet Dev 2016, 40, 11–16. [Google Scholar] [CrossRef] [PubMed]
Wang J, Dahmann C. Establishing compartment boundaries in Drosophila wing imaginal discs: An interplay between selector genes, signaling pathways and cell mechanics. Semin Cell Dev Biol 2020, 107, 161–169. [Google Scholar] [CrossRef] [PubMed]
Sharrock TE, Sanson B. Cell sorting and morphogenesis in early Drosophila embryos. Semin Cell Dev Biol 2020, 107, 147–160. [Google Scholar] [CrossRef] [PubMed]
Lawrence, PA. The making of a fly: the genetics of animal design. Blackwell Scientific Publications, Oxford, UK; 1992.
Worley MI, Setiawan L, Hariharan IK. Regeneration and transdetermination in Drosophila imaginal discs. Annu Rev Genet 2012, 46, 289–310. [Google Scholar] [CrossRef] [PubMed]
Townes PL, Holtfreter J. Directed movements and selective adhesion of embryonic amphibian cells. J Exp Zool 1955, 128, 53–120. [Google Scholar] [CrossRef]
Nose A, Nagafuchi A, Takeichi M. Expressed recombinant cadherins mediate cell sorting in model systems. Cell 1988, 54, 993–1001. [Google Scholar] [CrossRef] [PubMed]
Steinberg, MS. Differential adhesion in morphogenesis: a modern view. Curr Opin Genet Dev 2007, 17, 281–286. [Google Scholar] [CrossRef] [PubMed]
Brodland, GW. The differential interfacial tension hypothesis (DITH): a comprehensive theory for the self-rearrangement of embryonic cells and tissues. J Biomech Eng 2002, 124, 188–197. [Google Scholar] [CrossRef] [PubMed]
Wang HU, Xhen ZF, Anderson DJ. Molecular distinction and angiogenic interaction between embryonic arteries and veins revealed by ephrin-B2 and its receptor Eph-B4. Cell 1998, 93, 741–753. [Google Scholar] [CrossRef] [PubMed]
Gale NW, Baluk P, Pan L, Kwan M, Holash J, DeChiara TM, McDonald DM, Yancopoulos GD. Ephrin-B2 selectively marks arterial vessels and neovascularization sites in the adult, with expression in both endothelial and smooth muscle cells. Dev Biol 2001, 230, 151–160. [Google Scholar] [CrossRef] [PubMed]
Adams RH, Eichmann A. Axon guidance molecules in vascular patterning. Cold Spring Harb Perspect Biol 2010, 2, a001875. [Google Scholar] [CrossRef] [PubMed]
Shin D, Garcia-Cardena G, Hayashi S, Gerety S, Asahara T, Stavrakis G, Isner J, Folkman J, Gimbrone MA Jr, Anderson DJ. Expression of ephrinB2 identifies a stable genetic difference between arterial and venous vascular smooth muscle as well as endothelial cells, and marks subsets of microvessels at sites of adult neovascularization. Dev Biol 2001, 230, 139–150. [Google Scholar] [CrossRef] [PubMed]
Finney AC, Orr AW. Guidance molecules in vascular smooth muscle. Front Physiol 2018, 9, 1311. [Google Scholar] [CrossRef] [PubMed]
Tian X, Hu T, Je L, Zhang H, Huang X, Poelmann RE, Liu W, Yang Z, Yan Y, Pu WT, Zhou B. Peritruncal coronary endothelial cells contribute to proximal coronary artery stems and their aortic orifices in the mouse heart. PLoS One 2013, 8, e80857. [Google Scholar] [CrossRef] [PubMed]
Jiang X, Rowitch DH, Soriano P, McMahon AP, Sucov HM. Fate of the mammalian cardiac neural crest. Development 2000, 127, 1607–1616. [Google Scholar] [CrossRef] [PubMed]
Majesky, MW. Developmental basis of vascular smooth muscle diversity. Arterioscler Thromb Vasc Biol 2007, 27, 1248–1258. [Google Scholar] [CrossRef] [PubMed]
Passman JN, Dong XR, Wu SP, Maguire CT, Hogan KA, Bautch VL, Majesky MW. A sonic hedgehog signaling domain in the arterial adventitia supports resident Sca1+ smooth muscle progenitor cells. Proc Natl Acad Sci USA 2008, 105, 9349–9354. [Google Scholar] [CrossRef] [PubMed]
Sawada H, Rateri DL, Moorleghen JJ, Majesky MW, Daugherty A. Smooth muscle cells derived from the second heart field and cardiac neural crest reside in spatially distinct domains in the media of the ascending aorta – brief report. Arterioscler Thromb Vasc Biol 2017, 37, 1722–1726. [Google Scholar] [CrossRef] [PubMed]
Lin CJ, Hunkins B, Roth R, Lin CY, Wagenseil JE, Mecham RP. Vascular smooth muscle cell subpopulations and neointimal formation in mouse models of elastin insufficiency. Arterioscler Thromb Vasc Biol 2021, 41, 2890–2905. [Google Scholar] [CrossRef] [PubMed]
Sawada H, Katsumata y, Higashi H, Zhang C, Li Y, Morgan S, Lee LH, Singh SA, Chen JZ, Franklin MK, Moorleghen JJ, Howatt DA, Rateri DL, Shen YH, LeMaire SA, Aikawa M, Majesky MW, Lu HS, Daugherty A. Second heart field-derived cells contribute to angiotensin II-mediated ascending aortopathies. Circulation 2022, 145, 987–1001. [Google Scholar] [CrossRef] [PubMed]
Pedroza AJ, Dalal AR, Shad R. Yokoyama N, Nakamura K, Cheng P, Wirka RC, Mitchel O, Baiocchi M, Hiesinger W, Quertermous T, Fischbein MP. Embryologic origin influences smooth muscle cell phenotypic modulation signatures in murine Marfan syndrome aortic aneurysm. Arterioscler Thromb Vasc Biol 2022, 42, 1154–1168. [Google Scholar] [CrossRef] [PubMed]
Basler K, Struhl G. Compartment boundaries and the control of Drosophila limb pattern by hedgehog protein. Nature 1994, 368, 208–214. [Google Scholar] [CrossRef] [PubMed]
Lawrence PA, Struhl G. Morphogens, compartments, and pattern: Lessons from Drosophila? Cell 1996, 85, 951–961. [Google Scholar] [CrossRef] [PubMed]
Tessier-Lavigne M, Goodman CS. The molecular biology of axon guidance. Science 1996, 274, 1123–1133. [Google Scholar] [CrossRef] [PubMed]
Zuhdi N, Ortega B, Giovannone D, Ra H, Reyes M, Asención V, McNicoll I, Ma L, de Bellard ME. Slit molecules prevent entrance of trunk neural crest cells in developing gut. Int J Dev Neurosci 2015, 41, 8–16. [Google Scholar] [CrossRef] [PubMed]
Lepore JJ, Mericko PA, Cheng L, Lu MM, Morrisey EE, Parmacek MS. GATA-6 regulates semaphorin 3C and is required in cardiac neural crest for cardiovascular morphogenesis. J Clin Invest 2006, 116, 929–939. [Google Scholar] [CrossRef] [PubMed]
High F, Epstein JA. Signaling pathways regulating cardiac neural crest migration and differentiation. Novartis Found Symp 2007, 283, 152–161. [Google Scholar] [CrossRef] [PubMed]
Toyofuku T, Yoshida J, Sugimoto T, Yamamoto M, Makino N, Takamatsu H, Takegahara N, Suto D, Hori M, Fujisawa H, Kumanogoh A, Kikutani H. Repulsive and attractive semaphorins cooperate to direct the navigation of cardiac neural crest cells. Dev Biol 2008, 321, 251–262. [Google Scholar] [CrossRef] [PubMed]
Scholl AM, Kirby ML. Signals controlling neural crest contributions to the heart. WIRES 2009, 1, 220–227. [Google Scholar] [CrossRef] [PubMed]
Schulz Y, Wehner P, Optiz L, Salinas-Riester G, Bongers EMHF, van Ravenswaaij-Arts CMA, Wincent J, Schoumans J, Kohlhase J, Borchers A, Pauli S. CHD7, the gene mutated in CHARGE syndrome, regulates genes involved in neural crest cell guidance. Hum Genet 2014, 133, 997–1009. [Google Scholar] [CrossRef] [PubMed]
Kodo K, Shibata S, Miyagawa-Tomita S, Ong SG, Takahashi H, Kume T, Okano H, Matsuoka R, Yamagishi H. Regulation of Sema3c and the interaction between cardiac neural crest and second heart field during outflow tract development. Sci Rep 2017, 7, 6771. [Google Scholar] [CrossRef] [PubMed]
Schussler O, Gharibeh L, Mootoosamy P, Murith N, Tien V, Rougemont AL, Sologashvili T, Suuronen E, Lecarpentier Y, Ruel M. Cardiac neural crest cells: their rhombomeric specification, migration, and association with heart and great vessel anomalies. Cell Mol Neurobiol 2021, 41, 403–429. [Google Scholar] [CrossRef] [PubMed]
Becker SFS, Mayor R, Kashef J. Cadherin-11 mediates contact inhibition of locomotion during Xenopus neural crest cell migration. PLoS One 2013, 8, e85717. [Google Scholar] [CrossRef] [PubMed]
Carmona-Fontaine C, Matthews HK, Kuriyama S, Moreno M, Dunn GA, Parsons M, Stern CD, Mayor R. Contact inhibition of locomotion in vivo controls neural crest directional migration. Nature 2008, 456, 957–961. [Google Scholar] [CrossRef] [PubMed]
Mayor R, Carmona-Fontaine C. Keeping in touch with contact inhibition of locomotion. Trends Cell Biol 2010, 20, 319–328. [Google Scholar] [CrossRef] [PubMed]
Theveneau E, Marchant L, Kuriyama S, Gull M, Moepps B, Parsons M, Mayor R. Collective chemotaxis requires contact-dependent cell polarity. Dev Cell 2010, 19, 39–53. [Google Scholar] [CrossRef] [PubMed]
Willecke M, Hamaratoglu F, Sansores-Garcia L, Tal C, Halder G. Boundaries of Dachsous Cadherin activity modulate the Hippo signaling pathway to induce cell proliferation. Proc Natl Acad Sci USA 2008, 105, 14897–14902. [Google Scholar] [CrossRef] [PubMed]
Huang J, Cheng L, Li J, Chen M, Zhou D, Lu MM, Proweller A, Epstein JA, Parmacek MS. Myocardin regulates expression of contractile genes in smooth muscle cells and is required for closure of the ductus arteriosus in mice. J Clin Invest 2008, 118, 515–525. [Google Scholar] [CrossRef] [PubMed]
Yancopoulos GD, Klagsbrun M, Folkman J. Vasculogenesis, angiogenesis, and growth factors: ephrins enter the fray at the border. Cell 1998, 93, 661–664. [Google Scholar] [CrossRef] [PubMed]
Wu MF, Liao CY, Wang LY, Chang JT. The role of Slit-Robo signaling in the regulation of tissue barriers. Tissue Barriers 2017, 5, e1331155. [Google Scholar] [CrossRef] [PubMed]
Waldo K, Hutson M, Ward C, Zdanowicz M, Stadt H, Kumiski D, Abu-Issa R, Kirby M. Secondary heart field contributes myocardium and smooth muscle to the arterial pole of the developing heart. Dev Biol 2005, 281, 78–90. [Google Scholar] [CrossRef] [PubMed]
Topouzis S, Majesky MW. Smooth muscle lineage diversity in the chick embryo. Two types of aortic smooth muscle cell differ in growth and receptor-mediated transcriptional responses to transforming growth factor-beta. Dev Biol 1996, 178, 430–445. [Google Scholar] [CrossRef] [PubMed]
MacFarlane EG, Parker SJ, Shin JY, Kang BE, Ziegler SG, Creamer TJ, Bagirzadeh R, Bedja D, Chen Y, Calderon JF, Weissler K, Frischmeyer-Guerrerio PA, Lindsay ME, Habashi JP, Dietz HC. Lineage-specific events underlie aortic root aneurysm pathogenesis in Loeys-Dietz syndrome. J Clin Invest 2019, 129, 659–675. [Google Scholar] [CrossRef] [PubMed]
Shukla S, Jana S, Sanford N, Lee C, Liu L, Cheng P, Quertermous T, Dichek DA. Single cell transcriptomics identifies selective lineage-specific regulation of genes in aortic smooth muscle cells in mice. Arterioscler Thromb Vasc Biol.
Weldy CS, Cheng PP, Guo W, Pedroza AJ, Dalal AR, Worssam MD, Sharma D, Nguyen T, Kundu R, Fischbein MP, Quertermous T. The epigenomic landscape of single vascular cells reflects developmental origin and identifies disease risk loci. The epigenomic landscape of single vascular cells reflects developmental origin and identifies disease risk loci. bioRxiv 2022, 492517. [Google Scholar] [CrossRef]
Jaffe M, Sesti C, Washington IM, Du L, Dronadula N, Chin MT, Stolz DB, Davis AC, Dichek DA. Transforming growth factor-β signaling in myogenic cells regulates vascular morphogenesis, differentiation, and matrix synthesis. Arterioscler Thromb Vasc Biol 2012, 32, e1–e11. [Google Scholar] [CrossRef] [PubMed]
Le Noble F, Moyon D, Pardanaud L, Yuan L, Djonov V, Matthijsen R, Bréant C, Fluery V, Eichmann A. Flow regulates arterial-venous differentiation in the chick embryo yolk sac. Development 2004, 131, 361–375. [Google Scholar] [CrossRef] [PubMed]
Lucitti JL, Jones EAV, Huang C, Chen J, Fraser SE, Dickinson ME. Vascular remodeling of the mouse yolk sac requires hemodynamic force. Development 2007, 134, 3317–3326. [Google Scholar] [CrossRef] [PubMed]
Kowalski WJ, Dur O, Wang Y, Patrick MJ, Tinney JP, Keller BB, Pekkan K. Critical transitions in early embryonic aortic arch patterning and hemodynamics. PLoS One 2013, 8, e60271. [Google Scholar] [CrossRef] [PubMed]
Langille, BL. Arterial remodeling: relation to hemodynamics. Can J Physiol Pharmacol 1996, 74, 834–841. [Google Scholar] [CrossRef] [PubMed]
Bi W, Drake CJ, Schwarz JJ. The transcription factor Mef2c-null mouse exhibits complex vascular malformations and reduced cardiac expression of angiopoietin-1 and VEGF. Dev Biol 1999, 211, 255–267. [Google Scholar] [CrossRef] [PubMed]
Isogai S, Lawson ND, Torrealday S, Horiguchi M, Weinstein BM. Angiogenic network formation in the developing vertebrate trunk. Development 2009, 130, 5281–5290. [Google Scholar] [CrossRef] [PubMed]
Udan RS, Vadakkan TJ, Dickinson ME. Dynamic responses of endothelial cells to changes in blood flow during vascular remodeling of the mouse yolk sac. Development 2013, 140, 4041–4050. [Google Scholar] [CrossRef] [PubMed]
Langille, BL. Morphologic responses of endothelium to shear stress: reorganization of the adherens junction. Microcirculation 2001, 8, 195–206. [Google Scholar] [CrossRef] [PubMed]
Tzima E, Irani-Tehrani M, Klosses WB, Dejana E, Schultz DA, Engelhardt B, CaoG, DeLisser H, Schwartz MA. A mechanosensory complex that mediates the endothelial cell response to fluid shear stress. Nature 2005, 437, 426–431. [Google Scholar] [CrossRef] [PubMed]
Jones EAV, le Noble F, Eichmann A. What determines blood vessel structure? Genetic prespecification vs hemodynamics. Physiology 2006, 21, 388–395. [Google Scholar] [CrossRef] [PubMed]
Baeyens N, Bandopadhyay C, Coon BG, Yun S, Schwartz MA. Endothelial fluid shear stress sensing in vascular health and disease. J Clin Invest 2016, 126, 821–828. [Google Scholar] [CrossRef] [PubMed]
Kowalski WJ, Pekkan K, Tinney JP, Keller BK. Investigating developmental cardiovascular biomechanics and the origins of congenital heart disease. Front Physiol 2014, 5, 408. [Google Scholar] [CrossRef] [PubMed]
Wang Y, Dur O, Patrick MJ, Tinney JP, Tobita K, Keller BB, Pekkan K. Aortic arch morphogenesis and flow modeling in the chick embryo. Ann Biomed Eng 2009, 37, 1069–1081. [Google Scholar] [CrossRef] [PubMed]
Yashiro K, Shiratori H, Hamada H. Haemodynamics determined by a genetic programme govern asymmetric development of the aortic arch. Nature 2007, 450, 285–288. [Google Scholar] [CrossRef] [PubMed]
Snider P, Conway SJ. Developmental biology: The power of blood. Nature 2007, 450, 180–181. [Google Scholar] [CrossRef] [PubMed]
Karakaya C, Goktas S, Celik M, Kowalski WJ, Keller BB, Pekkan K. Asymmetry in mechanosensitive gene expression during aortic arch morphogenesis. Sci Rep 2018, 8, 16948. [Google Scholar] [CrossRef] [PubMed]
Greif DM, Kumar M, Lighthouse JK, Hum J, An A, Ding L, Red-Horse K, Espinoza FH, Olson L, Offermanns S, Krasnow MA. Radial construction of an arterial wall. Dev Cell 2012, 23, 482–493. [Google Scholar] [CrossRef] [PubMed]
Hamada H, Meno C, Watanabe D, Saijoh Y. Establishment of vertebrate left-right asymmetry. Nat Rev Genet 2002, 3, 103–113. [Google Scholar] [CrossRef] [PubMed]
Liu C, Liu W, Palie J, Lu MF, Brown NA, Martin JF. Pitx2c patterns anterior myocardium and aortic arch vessels and is required for local cell movement into atrioventricular cushions. Development 2002, 129, 5081–5091. [Google Scholar] [CrossRef] [PubMed]
Hill MC, Kadow ZA, Li L, Tran TT, Wythe JD, Martin JF. A cellular atlas of Pitx2-dependent cardiac development. Development 2019, 146, dev180398. [Google Scholar] [CrossRef] [PubMed]
Shiratori H, Sakuma R, Watanabe M, Hashiguchi H, Michida K, Sakai Y, Nishino J, Saijoh Y, Whitman M, Hamada H. Two-step regulation of left-right asymmetric expression of Pitx2, initiation by nodal signaling and maintenance by Nkx2. Mol Cell 2001, 7, 137–149. [Google Scholar] [CrossRef] [PubMed]
Phoon CK, Aristizabal O, Turnbull DH. 40 MHz doppler characterization of umbilical and dorsal aortic blood flow in the early mouse embryo. Ultrasound Med Biol. 2000, 26, 1275–1283. [Google Scholar] [CrossRef] [PubMed]
Kirby, ML. Cardiac development. Oxford University Press, Oxford OX2 6DP, United Kingdom, 2007. ISBN: 9780195178197.
Chen X, Gays D, Milia C, Santoro MM. Cilia control vascular mural cell recruitment in vertebrates. Cell Rep 2017, 18, 1033–1047. [Google Scholar] [CrossRef] [PubMed]
Padget RL, Mohite SS, Hoog TG, Justis BS, Green BE, Udan RS. Hemodynamic force is required for vascular smooth muscle cell recruitment to blood vessels during mouse embryonic development. Mech Dev 2019, 156, 8–19. [Google Scholar] [CrossRef] [PubMed]
Cheng S, Xia IF, Wanner R, Abello J, Stratment AN, Nicoli S. Hemodynamics regulate spatiotemporal artery muscularization in the developing circle of Willis. eLife 2024, 13, RP94094. [Google Scholar] [CrossRef] [PubMed]
Palumbo R, Gaetano C, Antonini A, Pompilio G, Bracco E, Rönnstrand L, Heldin CH, Capogrossi MC. Different effects of high and low shear stress on platelet-derived growth factor isoform release by endothelial cells: consequences for smooth muscle cell migration. Arterioscler Thromb Vasc Biol 2002, 22, 405–411. [Google Scholar] [CrossRef] [PubMed]
Dardik A, Yamashita A, Aziz F, Asada H, Sumpio BE. Shear stress-stimulated endothelial cells induce smooth muscle cell chemotaxis via platelet-derived growth factor-BB and interleukin-1alpha. J Vasc Surg 2005, 41, 321–331. [Google Scholar] [CrossRef] [PubMed]
Zerwes HG, Risau W. Polarized secretion of a platelet-derived growth factor-like chemotactic factor by endothelial cells in vitro. J Cell Biol 1987, 105, 2037–2041. [CrossRef] [PubMed]
Hellström M, Kalén M, Lindahl P, Abramsson A, Betsholtz C. Role of PDGF-B and PDGFR-beta in recruitment of vascular smooth muscle cells and pericytes during embryonic blood vessel formation in the mouse. Development 1999, 126, 3047–3055. [Google Scholar] [CrossRef] [PubMed]
Stratman AN, Pezoa SA, Farrelly OM, Castranova D, Dye LE, Butler MG, Sidik H, Talbot WS, Weinstein BM. Interactions between mural cells and endothelial cells stabilize the developing zebrafish dorsal aorta. Development 2017, 144, 115–127. [Google Scholar] [CrossRef] [PubMed]
Leonard EV, Figueroa RJ, Bussmann J, Lawson ND, Amigo JD, Siekmann AF. Regenerating vascular mural cells in zebrafish fin blood vessels are not derived from pre-existing mural cells and differentially require Pdgfrβ signaling for their development. Development 2022, 149, dev199640. [Google Scholar] [CrossRef] [PubMed]
Siekmann, AF. Biology of vascular mural cells. Development 2023, 150, dev200271. [Google Scholar] [CrossRef] [PubMed]
Lee J, Goeckel ME, Levitas A, Colijn S, Shin J, Hindes A, Mun G, Burton Z, Chintalapati B, Yin Y, Abello J, Solnica-Krezel L, Stratman A. CXCR3-CXCL11 signaling restricts angiogenesis and promotes pericyte recruitment. Arterioscler Thromb Vasc Biol. [CrossRef] [PubMed]
Grazioli A, Alves CS, Konstantopoilos K, Yang JT. Defective blood vessel development and pericyte/pvSMC distribution in alpha 4 integrin-deficient mouse embryos. Dev Biol 2006, 293, 165–177. [Google Scholar] [CrossRef] [PubMed]
Ando K, Fukuhara S, Izumi N, Nakajima H, Fukui H, Kelsh RN, Mochizuki N. Clarification of mural cell coverage of vascular endothelial cells by live imaging of zebrafish. Development 2016, 143, 1328. [CrossRef] [PubMed]
Padget RL, Mohite SS, Hoog TG, Justis BS, Green BE, Udan RS. Hemodynamic force is required for vascular smooth muscle cell recruitment to blood vessels during mouse embryonic development. Mech Dev 2019, 156, 8–19. [Google Scholar] [CrossRef] [PubMed]
Sinha S, Iyer S, Granata A. Embryonic origins of human vascular smooth muscle cells: implications for in vitro modeling and clinical application. Cell Mol Life Sci 2014, 71, 2271–2288. [Google Scholar] [CrossRef] [PubMed]
Alexander BE, Zhao H, Astrof S. SMAD4, a critical regulator of cardiac neural crest fate and vascular smooth muscle development. Dev Dyn 2024, 253, 119–143. [Google Scholar] [CrossRef] [PubMed]
Yokoyama U, Ichikawa Y, Minamisawa S, Ishikawa Y. Pathology and molecular mechanisms of coarctation of the aorta and its association with the ductus arteriosus. J Physiol Sci 2017, 67, 259–270. [Google Scholar] [CrossRef] [PubMed]
Sinning C, Zengin E, Kozlik-Feldman R, Blakenberg S, Rickers C, von Kodolitsch Y, Girdauskas E. Bicuspid aortic valve and aortic coarctation in congenital heart disease. Cardiovasc Diagn Ther 2018, 8, 780–788. [Google Scholar] [CrossRef] [PubMed]

Figure 1. CT image of a constriction site (arrow) in the region where the DA joins the descending AO in a neonatal patient with CoA.

Figure 2. Vascular cast of pharyngeal arch arteries (PAAs) in an E12.0 mouse embryo visualized by scanning electron microscopy. Note the close proximity of PAAs IV and VI during the period of SMC investment and flow-dependent remodeling of the PAA complex. Reprinted with permission from [9].

Figure 3. “Fish tail-like” insertion junction (between the asterisks) of human ductus arteriosus SMCs (LA) into the AO media (reprinted with permission from [19]. Note the absence of organized elastic fibers in the ductal insertion. LA – ligamentum arteriosum.

Figure 4. A smooth muscle compartment boundary model of CoA. (A) Illustration of a common site of aortic coarctation where the ductus arteriosus (purple) joins the descending thoracic aorta (red). Boxed area is illustrated in more detail in panel B. (B, top) Normally, cardiac neural crest-derived DA-SMCs (DA, purple) and paraxial mesoderm-derived descending Ao-SMCs (dAo, red) fail to mix and a compartment boundary is formed at their interface (see insertion junction, Figure 3). (B, bottom) In coarctation of the aorta, loss of the compartment boundary results in DA-SMCs (DA, purple) invading the aortic media (dAo, red) producing an aortic constriction upon birth of the neonate. (C) Potential mechanisms for the pathogenesis of CoA. The normal DA-SMC interface with Ao-SMCs is established during SMC investment of pharyngeal arch artery (PAA) walls in mid-gestation (see, Figure 2). Remodeling of the paired PAAs is highly sensitive to changes in blood flow rate and flow distribution through the right and left sides of the PAAs [10,15,98,99]. Either disruptions in SMC cell-cell interactions that normally prevent mixing across SMC origin-dependent compartment boundaries (C1), or defects in cardiac contractile activity (C2), valve morphogenesis (C3), or outflow tract formation/rotation (C4), can alter the hemodynamic shear stress distribution during PAA remodeling. We propose that the formation of the normal DA-Ao SMC interface (B) is also a flow-sensitive process during PAA remodeling and disturbed patterns of SMC investment during PAA remodeling can produce CoA observed in the neonate.

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.

© 2024 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

Copyright: This open access article is published under a Creative Commons CC BY 4.0 license, which permit the free download, distribution, and reuse, provided that the author and preprint are cited in any reuse.