Next Article in Journal
Identification of Blueberry miRNAs and Their Targets Based on High-Throughput Sequencing and Degradome Analyses
Next Article in Special Issue
β4 and β6 Integrin Expression Is Associated with the Subclassification and Clinicopathological Features of Intrahepatic Cholangiocarcinoma
Previous Article in Journal
Identification of Dysregulated microRNA Networks in Schwann Cell-Like Cultures Exposed to Immune Challenge: Potential Crosstalk with the Protective VIP/PACAP Neuropeptide System
Previous Article in Special Issue
Dexamethasone-Mediated Upregulation of Calreticulin Inhibits Primary Human Glioblastoma Dispersal Ex Vivo
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
IJMS
Volume 19
Issue 4
10.3390/ijms19040982
ijms-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

Multifaced Roles of the αvβ3 Integrin in Ehlers–Danlos and Arterial Tortuosity Syndromes’ Dermal Fibroblasts

by
Nicoletta Zoppi
,
Nicola Chiarelli
,
Marco Ritelli
and
Marina Colombi
*
Division of Biology and Genetics, Department of Molecular and Translational Medicine, School of Medicine, University of Brescia, 25123 Brescia, Italy
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2018, 19(4), 982; https://doi.org/10.3390/ijms19040982
Submission received: 28 February 2018 / Revised: 19 March 2018 / Accepted: 24 March 2018 / Published: 26 March 2018
(This article belongs to the Special Issue Integrins and Human Pathologies)
Browse Figures
Versions Notes

Abstract

:
The αvβ3 integrin, an endothelial cells’ receptor-binding fibronectin (FN) in the extracellular matrix (ECM) of blood vessels, regulates ECM remodeling during migration, invasion, angiogenesis, wound healing and inflammation, and is also involved in the epithelial mesenchymal transition. In vitro-grown human control fibroblasts organize a fibrillar network of FN, which is preferentially bound on the entire cell surface to its canonical α5β1 integrin receptor, whereas the αvβ3 integrin is present only in rare patches in focal contacts. We report on the preferential recruitment of the αvβ3 integrin, due to the lack of FN–ECM and its canonical integrin receptor, in dermal fibroblasts from Ehlers–Danlos syndromes (EDS) and arterial tortuosity syndrome (ATS), which are rare multisystem connective tissue disorders. We review our previous findings that unraveled different biological mechanisms elicited by the αvβ3 integrin in fibroblasts derived from patients affected with classical (cEDS), vascular (vEDS), hypermobile EDS (hEDS), hypermobility spectrum disorders (HSD), and ATS. In cEDS and vEDS, respectively, due to defective type V and type III collagens, αvβ3 rescues patients’ fibroblasts from anoikis through a paxillin-p60Src-mediated cross-talk with the EGF receptor. In hEDS and HSD, without a defined molecular basis, the αvβ3 integrin transduces to the ILK-Snail1-axis inducing a fibroblast-to-myofibroblast-transition. In ATS cells, the deficiency of the dehydroascorbic acid transporter GLUT10 leads to redox imbalance, ECM disarray together with the activation of a non-canonical αvβ3 integrin-TGFBRII signaling, involving p125FAK/p60Src/p38MAPK. The characterization of these different biological functions triggered by αvβ3 provides insights into the multifaced nature of this integrin, at least in cultured dermal fibroblasts, offering future perspectives for research in this field.

1. Introduction

1.1. Extracellular Matrix

Connective tissues are a mixture of different cell types and protein components organized in the reticular network of the extracellular matrix (ECM), which ensures tissue differentiation, structure, integrity and elasticity. ECM regulates many physiological cells’ activities, including adhesion, proliferation, survival, migration and apoptosis and it is also involved in phenotypic transition during morphogenesis and wound healing [1,2,3,4]. The ECM in vivo provides the interstitial matrix of tissues or, in basement membranes, pericellular layers binding the parenchymal cells to maintain them closely anchored to the connective tissue. The ECM is a dynamic structure undergoing organization and remodeling where ECM components are deposited, degraded or modified [5,6]. This constant turnover is essential during development and to reorganize the tissue architecture in physiological and pathological conditions [7,8]. The ECM composition is specific for different tissues and their physical properties, and about 300 macromolecules are defined to form the so-called matrisome, including collagens (COLLs), fibronectin (FN), elastin (ELN), ELN-associated proteins, i.e., fibrillins (FBNs), fibulins, and EMILINs, tenascins, laminins, osteonectin, osteopontin, glycosaminoglycans and proteoglycans (PGs) [9,10,11]. These constituents associate with each other to form a hydrated three-dimensional network that is also a reservoir of bioactive molecules and growth factors, such as the epidermal growth factor (EGF) and the transforming growth factor β (TGF-β), which regulate different cells’ activities [1]. The ECM is the result of biosynthetic activity and proteolytic degradation by cells that produce either structural ECM molecules or proteases, such as metalloproteases (MMPs), disintegrin and metalloproteinases, and plasminogen activators [12,13,14,15,16,17]. The unbalance between these two activities elicits an abnormal ECM organization, as observed in stiffness and fibrosis [5,8,18,19,20], or ECM degradation and loss occurring in invasive cancer disease [5,8,21,22] and in several heritable connective tissue disorders (HCTDs) [5,8,9,23,24,25]. Among the numerous ECM structural components, the FN is a high molecular weight (470–500 kDa) heterodimeric glycoprotein that is mainly expressed during embryogenesis and wound healing in a fibrillar form, whereas it is present in a globular form in the bloodstream [26,27,28,29]. The fibrillar FN is encoded by mRNA that contains the EDA segment (EDA+FN), whereas the circulating FN does not (EDA–FN) [30,31,32,33,34]. Other internal regions derived from alternative splicing at two additional sites i.e., EDB and IIICS [35], are known to play a role in tissue differentiation and are modulated in different pathological conditions [31,34,36,37,38,39,40,41].

1.2. Integrins

ECM proteins are connected with the cell cytoskeleton by integrins, a family of 24 heterodimeric transmembrane receptors containing an α and a β subunit, which transduce mechanical forces derived from the ECM into the cell or forces generated by the cytoskeleton, i.e., actin microfilaments, to the extracellular environment. The main integrins’ functions are reviewed elsewhere [42,43,44,45,46,47]. Integrin-mediated mechanotransduction induces cellular responses that drive development, cell movement, proliferation, survival and tissue homeostasis [48]. The binding of integrins to the cytoskeleton orchestrates cell migration [49], cell-ECM interaction, a survival signal that inhibit pro-apoptotic proteins [50,51], and an appropriate ECM releases several growth factors driving the cell cycle [50,52]. Cell-matrix adhesions are integrin-mediated molecular sites where the mechanical cues are converted into biochemical signaling. Different adhesive structures exist, such as nascent, focal, and fibrillar adhesions, which vary in shape, subcellular location, lifetime, functions and protein composition. Movements and interactions of their scaffold and signaling protein components lead to the assembly/disassembly, maturation, and interconversion of these dynamic architectures both in vivo and in vitro [53,54,55]. In cultured fibroblasts, the ECM adhesions can be classified in two major types, recruiting different settings of integrins and cytoskeleton anchor molecules, i.e., focal and fibrillar adhesions. Focal adhesions are transmembrane anchorage sites located at the fibroblast periphery between cells and underlying ECM fibrils. They are associated with the end of stress fibers and usually contain the αvβ3 integrin interacting with a complex pattern of proteins that include vinculin, talin, paxillin, α-actinin, zyxin, p125 focal-adhesion kinase (p125FAK), integrin-linked kinase (ILK), and other phosphotyrosine proteins and kinases [53,56,57]. Fibrillar adhesions, arising from focal adhesions, are elongated structures distributed more centrally on the cell surface, enriched in α5β1 integrins bound to tensin, and directly involved in the FN–ECM organization [53,56,58].

1.3. The αvβ3 Integrin

The αvβ3 integrin, originally named vitronectin receptor [59] because of its predominant ECM ligand, is one of the most promiscuous receptors binding a plethora of at least 21 different ECM proteins, including FN, FBNs, osteopontin, laminin, fibrinogen, von Willebrand factor, thrombospondin, and thrombin [60]. This integrin is widely expressed in endothelial cells (EC) where it is involved in angiogenesis [61,62,63,64], in smooth muscle cells (SMC) [65], myofibroblasts [66,67], osteoclasts, and blood cells, such as monocytes and platelets [59,60]. The recruitment on the cell surface of the αvβ3 integrin is a hallmark of myofibroblasts, specialized cells with both fibroblasts’ and SMCs’ phenotypic characteristics. These cells are activated by inflammatory cytokines and are involved in wound healing [68] and in pathological conditions, such as fibrosis [69] and chronic inflammation [70]. Table 1 summarizes the αvβ3 integrin cell expression, its major ligands and main functions.
As with all integrins, αvβ3 acts as a bidirectional signaling molecule. During “inside-out” signaling, the short cytoplasmic tail of the β3 subunit, through the binding to talin and kindlin, links the integrin to the actin cytoskeleton and elicits conformational changes, e.g., disruption of the intracellular bridges between the cytoplasmic subunits, dissociation of the transmembrane helices, and reorganization of the integrin in a high-affinity binding form that increase the affinity of αvβ3 for the extracellular ligands [71,72,73]. The binding of αvβ3 to the ECM drives the “outside-in” signals by clustering at the plasma membrane of other heterodimers, increase of adhesiveness, and downstream phosphorylation of several kinases for signal transduction. The cytoplasmic domain does not contain intrinsic tyrosine kinase activity and therefore “outside-in” signaling occurs primarily via the recruitment of intracellular signaling kinases, e.g., p125FAK, ILK, Src family kinases, paxillin, and vinculin that are important also for the actin cytoskeleton assembly [74,75]. A selected group of integrins including αvβ3 can stimulate the activation of Ras via interactions of the αv subunits with the adaptor molecule Shc and its association with Grb2 and Sos [76]. These interactions are crucial in adhesion-dependent cell proliferation and survival, as demonstrated by the up-regulation of the antiapoptotic protein Bcl-2 [77]. In addition, the integrin-mediated cell anchorage suppresses the p53 activity in the regulation of apoptosis [78,79]. Besides, the phosphorylation of the mitogen-activated protein kinases (MAPK), phosphoinositide kinase (PI3K)/Akt, and extracellular signal regulated kinase (ERK) is a downstream effect of the αvβ3 integrin activation that regulates cell proliferation, migration/invasion, and cell survival [80].
The αvβ3 integrin’s signaling can act synergistically with several growth factor receptors, such as the EGF receptor (EGFR) [81,82], and the TGF-β receptor (TGFBR) [83], also through the cross-talk with their downstream pathways [84,85,86]. For instance, previous studies have shown a direct interaction between αvβ3 and TGFBRII upon stimulation with active TGF-β [83,87].

1.4. Heritable Connective Tissue Disorders (HCTDs)

HCTDs comprise a wide range of pleiotropic multisystem diseases mainly affecting the connective tissue of various organ systems, including heart, blood vessels, bone, eyes, skin, joints and lungs. HCTDs result from genetic defects that perturb ECM assembly, maintenance, and homeostasis. Defects in the amount or structure of one of the numerous ECM constituents affect the proper organization and structural integrity of the supporting connective tissues and cause the weakness of bones, skin or vascular tissue which characterizes the disease phenotypes of different HCTDs [88]. Indeed, disease-causing mutations in several ECM-related genes or enzymes involved in biosynthesis or processing of ECM proteins, cause a myriad of HCTDs, e.g., Ehlers–Danlos syndromes (EDS), Osteogenesis imperfecta (OI), Marfan syndrome (MFS), Loeys–Dietz syndromes (LDS), arterial tortuosity syndrome (ATS), and numerous skeletal dysplasias [9]. Many of these disorders show some clinical overlap regarding cardiovascular, skeletal, craniofacial, ocular, and cutaneous features reflecting the common denominator of the ECM perturbation [89].

1.5. The Ehlers–Danlos Syndromes (EDS) and Arterial Tortuosity Syndome (ATS)

Among HCTDs, EDS share a variable combination of skin hyperextensibility, joint hypermobility (JHM), and manifestations of generalized connective tissue fragility. The revised 2017 EDS nosology distinguishes 13 different EDS types with 19 causative genes known to date (Table 2) [90]. For a comprehensive clinical and molecular description of all EDS types see the paper by Malfait and coworkers [90].
The classical (cEDS), vascular (vEDS) and hypermobile (hEDS) EDS forms account for more than 90% of EDS patients. Briefly, cEDS is characterized by abnormal skin involvement and generalized JHM (gJHM) [90,91], and is mainly caused by mutations in the COL5A1 and COL5A2 genes encoding the type V collagen (COLLV) [92,93]. vEDS is characterized by a clinical history of arterial rupture, dissection or aneurysm, rupture of the large intestine, and pregnancy complications at young ages [94]. vEDS is caused by mutations in the COL3A1 gene that encodes the type III collagen (COLLIII), which is the major expressed collagen in blood vessels and hollow organs [95]. The clinical criteria according to the revised 2017 EDS nosology suggestive for cEDS and vEDS are shown in Table 3 and Table 4. Confirmatory molecular testing is needed to reach a final diagnosis.
hEDS follows an autosomal dominant inheritance pattern with an unknown molecular basis and is mainly characterized by gJHM, joint instability complications, and minor skin changes [90]. The clinical criteria for a hEDS diagnosis according to the revised 2017 EDS nosology are summarized in Table 5. The phenotypic spectrum of hEDS also includes multiple associated symptoms shared with chronic inflammatory systemic diseases. Many of these features are not sufficiently specific nor sensitive to be included in the formal diagnostic criteria. These include, but are not limited to, sleep disturbance, fatigue, postural orthostatic tachycardia, functional gastrointestinal disorders, dysautonomia, anxiety, and depression [96]. Following the new classification, the term hypermobility spectrum disorders (HSD) is an alternative label for patients with symptomatic JHM who do not meet the new criteria for hEDS [97].
EDS are characterized by huge genetic heterogeneity, wide phenotypic variability between the different forms, and clinical overlap with other HCTDs. Indeed, EDS share with other HCTDs such as ATS, MFS, LDS, and OI, some degree of phenotypical overlap of cardiovascular, cutaneous, and skeletal features. Briefly, ATS is characterized by tortuosity and elongation of large- and medium-sized arteries and is caused by loss-of-function mutations in the SLC2A10 gene encoding the facilitative glucose transporter 10 (GLUT10), which facilitates the uptake of glucose and dehydroascorbic acid (DHA) [98,99]. For an overview of the phenotypic presentation in ATS patients see Table 6. MFS, caused by heterozygous mutations in FBN1 encoding the ECM protein fibrillin 1, is characterized by cardiovascular, ocular, and skeletal manifestations. The most common cardiovascular phenotype involves aortic aneurysm and dissection at the sinuses of Valsalva [89]. LDS, caused by mutations in different components of the TGF-β signaling pathway, i.e., TGFBR1, TGFBR2, SMAD2, SMAD3, TGFB2 and TGFB3, is mainly characterized by a clinical triad including hypertelorism, bifid uvula or cleft palate, and aortic aneurysm with arterial tortuosity [89]. OI comprises a heterogeneous group of diseases characterized by susceptibility to bone fractures with variable severity. OI display different modes of inheritance with autosomal dominant as the predominant inheritance pattern caused by mutations in the COL1A1 and COL1A2 genes (in about 85% of individuals) encoding the α1 and α2 chains of type I COLL (COLLI), respectively [100,101].

1.6. Organization of Fibronectin (FN) and Collagens (COLLs) and their Canonical Integrin Receptors in Dermal Fibroblasts from Different EDS Types and Other HCTDs

The skin is one of the connective tissues affected in all EDS types as well as in other HCTDs, and dermal fibroblasts have been shown to represent an excellent in vitro cell model to study ECM organization and molecular mechanisms involved in the pathophysiology of several HCTDs [102,103,104,105,106,107,108,109,110,111,112,113]. In vitro grown human dermal fibroblasts synthesize and secrete several ECM structural proteins that are deposited on the substrate and organized in a network covering the cell layer. In particular, three days after seeding, control fibroblasts organize the ECM of FN (FN–ECM), with a predominant deposition of the EDA+FN variant, COLLIII, COLLV, and rare fibrils of COLLI ([102,103,104,105,106,107,108,109,110,111,112,113], and Table 7). The main COLLs receptor expressed by control cells is the α2β1 integrin, whereas FN is preferentially bound to the α5β1 integrin [102,103,104]. In vitro cultured dermal fibroblasts derived from patients affected with different EDSs, except spEDS-B3GALT6 [109], exhibit a common cellular phenotype that is characterized by rare FN–ECM fibrils in association with reduced/absent patches in the plasma membrane of the canonical FN receptor the α5β1 integrin., Consequently, these FN–ECM-deficient cells show the preferential expression of the alternative FN receptor, i.e., the αvβ3 integrin, which is organized in linear patches both in fibrillar and focal adhesions ([102,103,106,107,110,111,112,113], and Table 7). This phenotype is also observed in dermal fibroblasts from patients with ATS but not in LDS-TGFBR1, MFS, and OI cells, which organize an abundant FN–ECM and express the α5β1 integrin and not αvβ3 ([105,114], and Table 7). EDS fibroblasts also show a reduced/absent deposition into the ECM of COLLI and COLLIII, in association with a variable organization of COLLV ([102,103,104,106,107,108,109,110,111,112,113], and Table 7). A similar disorganization of the COLLs–ECM is also observed in LDS-TGFBR1, OI, and ATS cells but not in MFS fibroblasts ([105], and Table 7). The abnormal COLLs–ECM depositions observed in all EDS types as well as in OI and ATS cells is associated with the loss of the canonical COLLs α2β1 integrin receptor’s expression ([102,103,106,107,108,109,110,111,112,113], and Table 7). For the genetically defined forms of EDS as well as for OI, the abnormal COLLs–ECM organization and the consequent loss of the α2β1 integrin is easily explained by the underlying molecular defects that include not only anomalies of the collagen primary structure (cEDS, vEDS, and OI), collagen processing (dEDS, aEDS), folding and cross-linking (kEDS), but also defects in glycosaminoglycan biosynthesis (spEDS, mcEDS) that are known to impact COLLs fibril formation and deposition [9]. Concerning hEDS and HSD cells, the lack of COLLs–ECM [107] could partly be explained by the high levels of the active form of the MMP-9 collagenase recently reported in their culture media [106]. In ATS cells, the lack of GLUT10 was shown to affect both the redox homeostasis and the proper processing and secretion of several ECM components, thus accounting for their defective COLLs–ECM [99,105,115,116].
Taken together, the FN–ECM disarray in association with reduced expression of its canonical integrin receptor α5β1 and consequent recruitment of the αvβ3 integrin seems to represent a peculiar in vitro phenotype of the majority of EDS cells as well as ATS fibroblasts (Figure 1).
Herein, we review our previously reported findings that unraveled different biological mechanisms elicited by the αvβ3 integrin in cEDS, vEDS, hEDS, HSD, and ATS dermal fibroblasts.

2. The Pro-Survival Role of the αvβ3 Integrin in Classical EDS (cEDS) and Vascular EDS (vEDS) Dermal Fibroblasts

In cEDS and vEDS fibroblasts the αvβ3 integrin is abundantly clustered both in focal and fibrillar contact sites, where it drives adhesion either to uncoated or purified FN-coated substrates [102,103]. The functional blocking of this integrin receptor with inhibiting antibodies reduces their adhesive potential corroborating that the αvβ3 integrin is the main adhesive ECM receptor in these EDS fibroblasts. In these cells, the αvβ3 integrin is a FN-binding receptor that sustains the FN assembly, since EDS cells grown in the presence of exogenous purified human plasma FN are induced to organize a FN fibrillar matrix without recruiting the canonical FN receptor α5β1 integrin [103]. This is consistent with the well-known capability of the αvβ3 integrin to bind and assemble a FN–ECM [117].
Moreover, the expression of the αvβ3 integrin in cEDS and vEDS fibroblasts is a downstream effect of COLLV and COLLIII deficiency, respectively, which, in turn, also affects the synthesis of the EDA–FN, and its secretion and organization into the ECM [104]. Indeed, the treatment of cEDS and vEDS fibroblasts with purified COLLV and COLLIII respectively restores the COLLs–ECM assembly, induces the up-regulation of the EDA+FN expression in association with its organization in a control-like ECM. This COLLs-mediated ECM rescue is associated with the restoration of a canonical cells’ integrin setting, since the αvβ3 integrin patches disappear and both the lacking COLLs receptor α2β1 integrin and the FN-specific α5β1 integrin are organized on the cell surface [102,104]. In cEDS and vEDS fibroblasts, the αvβ3 integrin is activated, as demonstrated by its tyrosine phosphorylation, and it transduces adhesion signals [103]. The adhesion-dependent fibroblasts’ survival is known to be regulated by ECM assembly and turn over. The lack of cell adhesion to the ECM induces the fibroblasts’ growth arrest and apoptosis/anoikis [118,119,120,121,122,123,124,125]. ECM-deficient cEDS and vEDS fibroblasts proliferate in vitro as well as control fibroblasts. However, the inhibition of the αvβ3 integrin induces cEDS and vEDS fibroblasts to undergo anoikis, suggesting that this receptor plays a key survival role in the rescue of these cells from anoikis activated by the ECM disassembly [103]. The survival function of the αvβ3 integrin has been reported in other cell types, i.e., endothelial [126] and tumor [127] cells. In cEDS and vEDS fibroblasts, the anti-apoptotic transduction pathway activated by the αvβ3 integrin does not involve p125FAK [103], which is known to have a central role in the PKB/Akt-mediated activation of the cell cycle and the inhibition of pro-apoptotic mediators, such as Bad and caspases [128,129]. In cEDS and vEDS cells, the down-regulation of p125FAK should explain the low synthesis of the survival protein Bcl-2 and the activation of caspases, thus leading to a pre-apoptotic cell behavior [103]. The caspases’ proteolytic activity in these EDS fibroblasts should produce the disassembly of actin microfilaments, as previously reported in other cell types [130]. In the absence of p125FAK, the αvβ3 integrin co-immunoprecipitates paxillin, which is distributed both in focal and fibrillar adhesions, as well as the αvβ3 integrin, talin, and vinculin. Since vinculin and paxillin are usually recruited in fibroblasts’ focal adhesion sites together with rare αvβ3 integrin patches [131], whereas fibrillar contacts consist of α5β1 integrin, tensin and talin [58,132], a different type of fibrillar adhesion is organized in cEDS and vEDS fibroblasts. Paxillin is activated trough tyrosine-phosphorylation by the p60Src kinase playing a role in EDS cells’ survival, since p60Src inhibition elicits EDS cells’ anoikis [103]. In these fibroblasts, paxillin is not serine-phosphorylated and does not recruit ILK, which is known to be involved in its activation [133]. The concomitant lack of ILK and p125FAK in EDS fibroblasts might contribute to the low levels of Bcl-2 expression and caspases’ activation. In addition, the αvβ3 integrin-p60Src-paxillin complexes recruit p130Cas at less extent, a docking protein that is physiologically involved in cytoskeleton remodeling [134] and that is degraded by caspases [135]. Therefore, in cEDS and vEDS fibroblasts the low amounts of p130Cas could result from the activity of caspases and could explain the actin cytoskeleton disassembly [103].
Furthermore, in these cells the αvβ3 integrin signaling rescues from anoikis by a cross-talk with EGFR, as demonstrated by the immunoprecipitation of phosphorylated EGFR with the αvβ3 integrin and by the induction of apoptosis observed either after antibody-mediated EGFR or αvβ3 inhibition or both [103], in line with several evidences that reported cross-talk mechanisms between integrins and growth factors’ receptors eliciting cell growth and rescue from apoptosis [81,82,136]. EGF–EGFR can transduce for tyrosine phosphorylation of paxillin [137], which, in turn, can act as an adaptive molecule integrating signals from integrins and growth factor receptors to ensure cell proliferation [138]. In Figure 2 a schematic representation of this survival pathway is shown.
The pre-apoptotic behavior of vEDS fibroblasts has been associated with an abnormal endoplasmic reticulum (ER) homeostasis resulting from intracellular retention of mutant and misfolded COLLIII chains. In vEDS cells, an ER perturbation was suggested by the abnormal ER distribution of the protein disulfide isomerase enzyme and by reduced expression of FKBP22, an ER resident peptidyl-prolyl cis-trans-ER isomerase involved in the folding of COLLIII triple helix [108,139,140]. Since the perturbed ER redox state may influence the down-regulation of Bcl-2 and the activation of the caspase-dependent apoptosis [141], and FKBP22 can have an anti-apoptotic role acting on the expression of Bcl-2 and caspases [142], this aspect merits further studies in vEDS fibroblasts.
In conclusion, in cEDS and vEDS fibroblasts an in vitro survival mechanism supported by an αvβ3 integrin-EGFR cross-talk transducing to paxillin is activated, thus sustaining cell adhesion in the absence of the actin cytoskeleton. Paxillin, activated by p60Src, probably works as a strategic molecule to reinforce the αvβ3 integrin- and EGFR-mediated signaling pathways ensuring the rescue from anoikis.

3. The αvβ3 Integrin Signaling Sustains the Hypermobile EDS (hEDS) and Hypermobility Spectrum Disorders (HSD) Fibroblast-to-Myofibroblast Transition

In hEDS and HSD cells, the αvβ3 integrin was shown to be involved in the fibroblast-to-myofibroblast transition by a transduction pathway involving ILK that signals to the transcription factor Snail1 [106]. Contrary to cEDS and vEDS fibroblasts, hEDS and HSD cells exhibit a peculiar in vitro myofibroblast-like phenotype characterized by organization of the α-smooth muscle actin (α-SMA) cytoskeleton, expression of the cadherin-11, and enhanced migratory capability, probably because they synthesize high levels of MMP9, a collagenase also able to digest FN into proteolytic fragments [106].
The implication of αv subunits-containing integrin receptors, including αvβ3, in the promotion of the α-SMA stress fibres assembly and fibroblast-to-myofibroblast transition has been investigated in different in vitro cell types [143,144]. In hEDS and HSD cells, the αvβ3 integrin binds to ILK and the β3 subunit is phosphorylated. These αvβ3 integrin-ILK complexes are recruited and activated in focal adhesion sites, promoting the α-SMA cytoskeleton’s organization, and ensuring the maintenance of the myofibroblast-like phenotype. The ILK inhibition with the liposoluble inhibitor Cpd22 induces both in hEDS and HSD cells the gradual disappearance of αvβ3 integrin and the α-SMA stress fibres disassembly [106]. Snail1, a transcription factor known to be involved in the transdifferentiation mechanisms [145], is one of the downstream effectors of the αvβ3 integrin-ILK axis in hEDS and HSD cells [106]. Indeed, it localizes both at cytoplasmic and nuclear level and immunoprecipitates either with the αvβ3 integrin or with ILK, suggesting a role in the phenotypic switch of these cells. Furthermore, the ILK inhibition delocalizes Snail1 from nuclei, confirming the activation in hEDS and HSD cells of an αvβ3 integrin-ILK-Snail1 axis sustaining the myofibroblasts’ phenotype. This condition is consistent with the role of Snail1 in the induction of myofibroblasts’ markers, i.e., cadherin-11 and α-SMA, as previously described in synovial fibroblasts from rheumatoid arthritis patients [146,147,148]. Although target molecules possibly activated by αvβ3 integrin-ILK complexes that act upstream Snail1 were not investigated in hEDS and HSD cells [106], transcriptome data in these cell models suggest a possible involvement of PI3K/Akt/GSK-3β and NF-kB signaling [107], which plays an important role in transdifferentiation mechanisms [149]. In this regard, ILK may act as kinase downstream the PI3K signaling pathway [150] and could be involved in the negative regulation, through the phosphorylation of a specific serine residue, of the glycogen synthase kinase 3β (GSK-3β) [151]. GSK-3β is a well-known kinase that physiologically controls the Snail1 export from the nucleus to the cytoplasm and its consequent degradation by ubiquitination [150,152,153]. Based on these findings, it reasonable to assume that αvβ3 integrin-ILK complexes in the focal contact sites of hEDS and HSD cells may be involved in the regulation of the GSK-3β function [106]. In hEDS and HSD cells, the ILK-mediated action should repress GSK-3β activity by its phosphorylation, thus resulting in the nuclear Snail1 expression and in the induction of the α-SMA organization. This is also sustained by the reduction of the nuclear Snail1 associated with the α-SMA disassembly observed after inhibition of the ILK kinase activity by Cpd22 in a dose-dependent manner [106], given that Cpd22 is known to prevent the ILK-mediated phosphorylation of GSK-3β [154]. In addition, ILK could phosphorylate Akt that, in turn, promotes NF-κB activation that enhances Snail1’s transcription by binding its promoter [155]. In Figure 3 a schematic representation of the αvβ3 integrin-ILK-Snail1 axis involved in the fibroblast-to-myofibroblast transition is shown.
The αvβ3 integrin’s ligands likely involved in the myofibroblast’s phenotype have not been identified yet, but the ECM disarray and, in particular, the proteolytic fragments generated by MMP9 from structural ECM components such as FN should play a key role in this signaling. Indeed, control fibroblasts, grown in the presence of the hEDS- and HSD-conditioned media, disassemble the FN–ECM and are induced to cluster at membrane level the αvβ3 integrin, which transduces through ILK to the nuclear Snail1 for the α-SMA fibers’ organization [106]. This “reprogramming” of control fibroblasts with hEDS and HSD cells’ media raises the question which factor(s), including not only growth factors and cytokines, e.g., TGF-β and Wnt, but also ECM fragments produced by proteolytic activity of different proteases such as MMP9, might be the primary contributor(s) to this transition. Indeed, it is recognized that intracellular molecules released from damaged tissues, as well as fragments of the ECM released downstream cell injury, can act as damage-associated molecular patterns (DAMPs). These molecules serve as danger signals that can elicit an immune response following tissue injury or in response to the changes in tissue composition and organization [156,157,158,159,160]. In this light, it is reasonable to assume that in hEDS/HSD cells these molecules may act as DAMPs and could be recognized by the αvβ3 integrin or other specialized receptors, which turn on pathways that transduce for the synthesis of a plethora of inflammation molecules. Both in vivo and in vitro, DAMPs could be responsible for the inflammation and chronic pain described in hEDS and HSD patients [161,162]. In this regard, hEDS and HSD cells express high levels of CCN2/CTGF, responsible for myofibroblast’ proliferation and differentiation, and low levels of CCN1/CYR61, involved in the myofibroblast’ apoptosis and resolution of inflammation [163,164]. The unbalanced synthesis of these inflammatory mediators might avoid the hEDS and HSD cells’ apoptotic death and sustain the hEDS and HSD myofibroblast-like phenotype.
Since in hEDS and HSD fibroblasts the serum deprivation does not induce cells to undergo apoptosis, the αvβ3 integrin-ILK-Snail1 pathway should be independent from serum growth factors’ action. Indeed, the contribution of mitogenic molecules in the fibroblast-to-myofibroblast transition is a thrilling topic, since myofibroblast differentiation and α-SMA expression are induced by the TGF-β [165,166]. The enhanced CCN2/CTGF expression observed in hEDS and HSD cells suggests the possible involvement of the TGF-β signaling [106]. Furthermore, it is known that CCN2/CTGF co-distributes and interacts with αv-containing integrins, including αvβ3 [167,168]. The possible involvement of the TGF-β pathway in the maintenance of the high levels of CCN2/CTGF observed hEDS ad HSD cells and, given the putative binding of CCN2/CTGF to the αvβ3 integrin, its possible role in the activation of the αvβ3 integrin-ILK-Snail1 signaling finally leading to the fibroblast-to-myofibroblast transition remain to be clarified. In this view, in different cell models increased Snail1 levels have been associated with the secretion of CCN2/CTGF, which, in turn, induces fibroblast-to-myofibroblast transition [169,170].
In conclusion, the identification of the αvβ3 integrin-ILK-Snail1 transduction pathway in hEDS and HSD cells provides insights into the molecular mechanisms likely involved in the pathophysiology of these neglected disorders. Further additional studies are needed and may represent a starting point for identifying potential therapeutic options. Quantitative protein profiling of hEDS and HSD cells’ media, aimed at clarifying which key factors secreted in their culture media are involved in the fibroblast-to-myofibroblast transition, are ongoing. In addition, the elucidation of the possible ILK-regulated downstream signaling partners, i.e., AKT and GSK-3β, which are likely involved in the transduction pathway that links the αvβ3 integrin, likely through Snail1, to the phenotypic switch of hEDS and HSD fibroblasts requires future in-depth studies.

4. In Arterial Tortuosity Syndrome (ATS) Dermal Fibroblasts the αvβ3 Integrin Is Involved in a Non-Canonical TGF-β Signaling

Among the different HCTDs dermal fibroblasts shown in Table 7, only ATS fibroblasts share with EDS cells the peculiar cellular phenotype characterized by the αvβ3 integrin expression (Figure 1). In addition to the disorganization of their FN- and COLLs–ECMs, these cells also show the lack of decorin (DCN) expression and do not organize FBNs and ELN in a fibrillar network [105]. This abnormal ECM organization and particularly that of FBNs and ELN is consistent with the elastic tissue disarray reported in the arterial wall of ATS patients [98]. The exact pathomechanisms by which GLUT10 deficiency leads to the generalized ECM disarray, which reflects not only the typical ATS vascular anomalies but also the multisystem involvement overlapping with EDSs [171], were debated for long time and remain incompletely explored [98,99,105,115,116,172]. The first study that identified disease-causative variants in SLC2A10 described an up-regulation of TGF-β signaling in arterial wall and ATS dermal fibroblasts [98]. The involvement of this pathway in the molecular pathology of ATS was sustained by the increased levels of CCN2/CTGF expression reported in ATS arterial tissue [98] and by the reduced expression of DCN, which is a TGF-β signaling inhibitor PG, in in vitro-grown patients’ fibroblasts [105]. In the last years, several evidences demonstrated that GLUT10 is localized in the ER and acts as a DHA transporter, which is the oxidized form of the ascorbic acid (AA) [99,115,116]. AA plays a major role in redox homeostasis by reducing reactive oxygen species (ROS) production, thereby protecting cells against oxidative stress. It also acts as a cofactor in reactions catalyzed by prolyl- and lysyl hydroxylases, which are a class of ER-resident enzymes involved in the maturation of COLLs and ELN [99,115,116]. ATS fibroblasts undergo oxidative stress, as shown by their high levels of ROS-mediated lipid peroxidation products [105]. However, in both the “enzyme cofactor” and “antioxidant” models of vitamin C-related pathology, the exact interaction with the TGF-β pathway remains elusive.
Since it is known that the αv-containing integrins, including αvβ3, play a key role in the activation of TGF-β [173] by interacting with the RGD motif present in the latency-associated peptide that results in the activation of the latent TGF-β [143,174,175], we hypothesized that this specific integrin might be involved in the aberrant TGF-β signaling observed in ATS fibroblasts [98,105]. Indeed, we unraveled a cross-talk mechanism between the αvβ3 integrin and a non-canonical TGF-β signaling. Specifically, in these cells the αvβ3 integrin co-immunoprecipitates and co-distributes with CCN2/CTGF, suggesting a direct interaction between αvβ3 and this growth factor, as previously reported in other cell models [168]. In ATS fibroblasts, the αvβ3 integrin transduces to p60Src through the recruitment of p125FAK, a key mediator of the TGF-β signaling [176,177], which is likely involved in the phosphorylation of the β3 integrin subunit [105]. Furthermore, ATS cells express very low levels of the TGFBRI and higher amounts of TGFBRII, which immunoprecipitates with the αvβ3 integrin. TGFBRII is tyrosine phosphorylated and p60Src should play a role in its activation, because of their co-immunoprecipitation with the αvβ3 integrin. This non-canonical pathway not only does not involve TGFBRI but also SMAD2 that is expressed at lower level than in control cells. Furthermore, TGFBRII also recruits p38MAPK that is activated by p60Src, as shown by the PP2-mediated inhibition of p60Src leading to the complete elimination of p38MAPK phosphorylation [105]. p38MAPK is a kinase involved in a wide range of signaling pathways that stimulate a multitude of different biological functions including adhesion, migration, ECM remodelling. Since p38MAPK might also be activated by stimuli other than TGF-β signaling, including an imbalance of redox homeostasis [178], it reasonable to assume that this kinase and its downstream effects play a role in the pathomechanisms of ATS. However, future investigations are needed to elucidate this intriguing aspect.
In ATS cells, the perturbation of redox homeostasis and TGF-β signaling associated with the aberrant ECM organization is dependent on the defective GLUT10-mediated DHA transport. In fact, the stable expression of functional GLUT10 in ATS fibroblasts restores a correct DHA transport activity [99], normalizes the impaired redox homeostasis, and rescues a canonical TGF-β signaling recruiting the TGFBRI/TGFBRII receptor, which transduces to SMAD2, in the absence of αvβ3 integrin, p60Src and p38MAPK. The recovery of both redox homeostasis and canonical TGF-β signaling is associated with re-expression of DCN and, partly, reorganization of their ELN- and FBNs-ECM [105]. These observations agree with previous findings showing that the oxidative stress can induce an aberrant TGF-β signaling resulting in abnormal elastogenesis and consequent ELN disassembly into the ECM [179,180].
Overall, in ATS fibroblasts, GLUT10 deficiency, through a faulty intracellular DHA uptake, leads to redox unbalance and abnormal ECM protein maturation resulting in generalized ECM disorganization and activation of a non-canonical αvβ3 integrin-TGFBRII signaling, involving CCN2/CTGF, p125FAK, p60Src, and p38MAPK.

5. Concluding Remarks

Human dermal fibroblasts from healthy individuals organize in vitro several structural ECM components that recruit specific integrin receptors. FN is assembled in a fibrillar network through the binding with the α5β1 integrin. In the majority of EDS and in ATS fibroblasts carrying mutations in different ECM-related genes, but not in those derived from LDS, MFS and OI patients, the FN–ECM and the α5β1 integrin are disorganized and strongly reduced respectively and, consequently, the alternative FN receptor the αvβ3 integrin, which is almost absent in control fibroblasts, is recruited. The different implications of the αvβ3 integrin identified in cultured cEDS, vEDS, hEDS, HSD, and ATS dermal fibroblasts highlight the multifaced nature of this integrin at least in this in vitro cell model. In cEDS and vEDS cells, the αvβ3 integrin exerts a pro-survival role promoting cell adhesion and preventing anoikis through p60Src-paxillin-mediated signaling and EGFR cross-talking. In hEDS and HSD cells, the αvβ3 integrin is involved in the fibroblast-to-myofibroblast transition through the interaction with ILK that signals to the transcription factor Snail1. In ATS fibroblasts, the αvβ3 integrin cross-talks with a TGFBRII-mediated non-canonical TGFβ signaling.
In conclusion, these data show that αvβ3 integrin, specifically recruited by EDS and ATS cells, participates in different signaling pathways depending on the specific disorder and underlying molecular defects. The αvβ3 integrin is sufficiently multifaced to switch on different signaling cascades downstream of the interplay enrolled with its numerous intracellular interactors and by its cross-talk capability with different growth factor-receptor complexes. These findings add insights into the manifold functions of the αvβ3 integrin, but more effort is required to deepen knowledge on the numerous partners identified in the reported pathways that could shed light on pathomechanisms associated with these HCTDs. In addition, further functional studies are also needed to ascertain whether similar or different αvβ3-mediated signaling mechanisms are elicited in the other EDS fibroblasts expressing this integrin.

Acknowledgments

The authors thank the patients for their kind availability for these studies, the EDS patients’ association C.E.D.I. (Clinici Ehlers–Danlos Italia) Onlus and the Fazzo Cusan family for their generous support. This work was supported by research grants from the Italian Ministry of University (MIUR, Fondo per gli investimenti della ricerca di base 2017).

Conflicts of Interest

The authors declare no conflict of interest.

Glossary

Extracellular matrix (ECM)ECM is a three-dimensional structure that encapsulates cells and defines their microenvironment, providing a physical scaffolding for the cellular constituents. ECM is a dynamic structure, constantly undergoing a remodeling process whereby ECM components are deposited, degraded, or modified. ECM dynamics are essential during restructuring of tissue architecture.
IntegrinsA family of cell adhesion receptors that mediate either cell–cell interactions or cell–ECM interactions. Integrins are heterodimers with two distinct subunits, the α-subunit and the β-subunit.
FibroblastsThe major cells responsible for the production of collagens, glycosaminoglycans, and proteoglycans, which are major components of the ECM.
MyofibroblastsSpecialized cells with both fibroblasts’ and SMCs’ phenotypic characteristics. These cells are activated by inflammatory cytokines and are involved in wound-healing mechanisms and in pathological conditions, such as fibrosis and chronic inflammation. They peculiarly express the α-SMA that they organize into the cytoskeleton to generate contractile force and migrate.
Fibroblast-to-myofibroblast transitionPhenotypic conversion of fibroblast into myofibroblast by transdifferentiation mechanisms occurring during wound healing, fibrosis, and inflammation. A well-characterized hallmark of the fibroblast-to myofibroblast transition is the novo formation of α-SMA stress fibers.
Focal adhesionsTransmembrane anchorage sites organized on the lower surface of the cell and in the cell’s periphery to anchor the underlying ECM fibrils. These structures are associated with the end of actin stress fibers and usually contain the αvβ3 integrin that interacts with a complex pattern of proteins including vinculin, talin, paxillin, α-actinin, zyxin, p125FAK, ILK, and other phosphotyrosine proteins and kinases.
Fibrillar adhesionsIntegrin-containing complexes arising from focal adhesions, distributed on the upper cell surface, involved in the ECM fibrils’ organization. These elongated structures are enriched in α5β1 integrins bound to tensin.
AnoikisMechanism of programmed cell death or apoptosis induced by the loss of cell-ECM adhesion. Consequently, the ECM can be considered a survival signal for cells.
Cross-talk mechanismMechanism by which two or more surface receptors or two or more interactors recruited in different signal-transduction pathways affect each other and reinforce the downstream cell response to an extracellular signal.
Damage-associated molecular patterns (DAMPs)Molecules released from damaged tissues, such as components or fragments of the ECM, released downstream of the cell injury. These danger signals bind specific receptors, such as Toll-like receptors, to elicit an immune response following tissue injury or in response to the changes in tissue composition and organization. DAMPs can also play a role in chronic pain conditions.
Tissue homeostasisA homeostatic process involved in the maintenance of an internal steady state within a defined tissue of an organism, including control of cellular proliferation and death and control of metabolic function.
Redox homeostasisBalance between intracellular reactive oxygen species (ROS) generation and elimination.

References

  1. Hynes, R.O. The extracellular matrix: Not just pretty fibrils. Science 2009, 326, 1216–1219. [Google Scholar] [CrossRef] [PubMed]
  2. Frantz, C.; Stewart, K.M.; Weaver, V.M. The extracellular matrix at a glance. J. Cell Sci. 2010, 123, 4195–4200. [Google Scholar] [CrossRef] [PubMed]
  3. Theocharis, A.D.; Gialeli, C.; Hascall, V.C.; Karamanos, N.K. Extracellular matrix: A functional scaffold. In Extracellular Matrix: Pathobiology and Signaling; Karamanos, N.K., Ed.; Walter de Gruyter GmbH & Co. KG: Berlin, Germany; Boston, MA, USA, 2012; pp. 3–20. [Google Scholar]
  4. Clause, K.C.; Barker, T.H. Extracellular matrix signaling in morphogenesis and repair. Curr. Opin. Biotechnol. 2013, 24, 830–833. [Google Scholar] [CrossRef] [PubMed]
  5. Karsdal, M.A.; Nielsen, M.J.; Sand, J.M.; Henriksen, K.; Genovese, F.; Bay-Jensen, A.C.; Smith, V.; Adamkewicz, J.I.; Christiansen, C.; Leeming, D.J. Extracellular matrix remodeling: The common denominator in connective tissue diseases. Possibilities for evaluation and current understanding of the matrix as more than a passive architecture, but a key player in tissue failure. Assay Drug Dev. Technol. 2013, 11, 70–92. [Google Scholar] [CrossRef] [PubMed]
  6. Vanakker, O.; Callewaert, B.; Malfait, F.; Coucke, P. The Genetics of Soft Connective Tissue Disorders. Annu. Rev. Genom. Hum. Genet. 2015, 16, 229–255. [Google Scholar] [CrossRef] [PubMed]
  7. Rozario, T.; DeSimone, D.W. The extracellular matrix in development and morphogenesis: A dynamic view. Dev. Biol. 2010, 341, 126–140. [Google Scholar] [CrossRef] [PubMed]
  8. Bonnans, C.; Chou, J.; Werb, Z. Remodelling the extracellular matrix in development and disease. Nat. Rev. Mol. Cell Biol. 2014, 15, 786–801. [Google Scholar] [CrossRef] [PubMed]
  9. Bateman, J.F.; Boot-Handford, R.P.; Lamande, S.R. Genetic diseases of connective tissues: Cellular and extra cellular effects of ECM mutations. Nat. Rev. Genet. 2009, 10, 173–183. [Google Scholar] [CrossRef] [PubMed]
  10. Hynes, R.O.; Naba, A. Overview of the matrisome—An inventory of extracellular matrix constituents and functions. Cold Spring Harb. Perspect. Biol. 2012, 4, a004903. [Google Scholar] [CrossRef] [PubMed]
  11. Theocharis, A.D.; Skandalis, S.S.; Gialeli, C.; Karamanos, N.K. Extracellular matrix structure. Adv. Drug Deliv. Rev. 2016, 97, 4–27. [Google Scholar] [CrossRef] [PubMed]
  12. Puente, X.S.; Sanchez, L.M.; Overall, C.M.; Lopez-Otin, C. Human and mouse proteases: A comparative genomic approach. Nat. Rev. Genet. 2003, 4, 544–558. [Google Scholar] [CrossRef] [PubMed]
  13. Lopez-Otin, C.; Bond, J.S. Proteases: Multifunctional enzymes in life and disease. J. Biol. Chem. 2008, 283, 30433–30437. [Google Scholar] [CrossRef] [PubMed]
  14. Visse, R.; Nagase, H. Matrix metalloproteinases and tissue inhibitors of metalloproteinases: Structure, function, and biochemistry. Circ. Res. 2003, 92, 827–839. [Google Scholar] [CrossRef] [PubMed]
  15. Nagase, H.; Visse, R.; Murphy, G. Structure and function of matrix metalloproteinases and TIMPs. Cardiovasc. Res. 2006, 69, 562–573. [Google Scholar] [CrossRef] [PubMed]
  16. Edwards, D.R.; Handsley, M.M.; Pennington, C.J. The ADAM metalloproteinases. Mol. Asp. Med. 2008, 29, 258–289. [Google Scholar] [CrossRef] [PubMed]
  17. Svineng, G.; Magnussen, S.; Hadler-Olsen, E. Plasmin and the plasminogen activator system in health and disease. In Extracellular Matrix: Pathobiology and Signaling; Karamanos, N.K., Ed.; Walter de Gruyter GmbH & Co. KG: Berlin, Germany; Boston, MA, USA, 2012; pp. 261–290. [Google Scholar]
  18. Wynn, T.A.; Ramalingam, T.R. Mechanisms of fibrosis: Therapeutic translation for fibrotic disease. Nat. Med. 2012, 18, 1028–1040. [Google Scholar] [CrossRef] [PubMed]
  19. Duffield, J.S.; Lupher, M.; Thannickal, V.J.; Wynn, T.A. Host responses in tissue repair and fibrosis. Annu. Rev. Pathol. 2013, 8, 241–276. [Google Scholar] [CrossRef] [PubMed]
  20. Bhattacharyya, S.; Tamaki, Z.; Wang, W.; Hinchcliff, M.; Hoover, P.; Getsios, S.; White, E.S.; Varga, J. Fibronectin EDA promotes chronic cutaneous fibrosis through Toll-like receptor signaling. Sci. Transl. Med. 2014, 6. [Google Scholar] [CrossRef] [PubMed]
  21. Lu, P.; Weaver, V.M.; Werb, Z. The extracellular matrix: A dynamic niche in cancer progression. J. Cell Biol. 2012, 196, 395–406. [Google Scholar] [CrossRef] [PubMed]
  22. Pickup, M.W.; Mouw, J.K.; Weaver, V.M. The extracellular matrix modulates the hallmarks of cancer. EMBO Rep. 2014, 15, 1243–1253. [Google Scholar] [CrossRef] [PubMed]
  23. Kim, H.E.; Dalal, S.S.; Young, E.; Legato, M.J.; Weisfeldt, M.L.; D’Armiento, J. Disruption of the myocardial extracellular matrix leads to cardiac dysfunction. J. Clin. Investig. 2000, 106, 857–866. [Google Scholar] [CrossRef] [PubMed]
  24. Bondeson, J.; Wainwright, S.; Hughes, C.; Caterson, B. The regulation of the ADAMTS4 and ADAMTS5 aggrecanases in osteoarthritis: A review. Clin. Exp. Rheumatol. 2008, 26, 139–145. [Google Scholar] [PubMed]
  25. Sorokin, L. The impact of the extracellular matrix on inflammation. Nat. Rev. Immunol. 2010, 10, 712–723. [Google Scholar] [CrossRef] [PubMed]
  26. Kornblihtt, A.R.; Vibe-Pedersen, K.; Baralle, F.E. Human fibronectin: Cell specific alternative mRNA splicing generates polypeptide chains differing in the number of internal repeats. Nucleic Acids Res. 1984, 12, 5853–5868. [Google Scholar] [CrossRef] [PubMed]
  27. Tamkun, J.W.; Hynes, R.O. Plasma fibronectin is synthesized and secreted by hepatocytes. J. Biol. Chem. 1983, 258, 4641–4647. [Google Scholar] [PubMed]
  28. Pankov, R.; Yamada, K.M. Fibronectin at a glance. J. Cell Sci. 2002, 115, 3861–3863. [Google Scholar] [CrossRef] [PubMed]
  29. Moretti, F.A.; Chauhan, A.K.; Iaconcig, A.; Porro, F.; Baralle, F.E.; Muro, A.F. A major fraction of fibronectin present in the extracellular matrix of tissues is plasma-derived. J. Biol. Chem. 2007, 282, 28057–28062. [Google Scholar] [CrossRef] [PubMed]
  30. Kornblihtt, A.R.; Umezawa, K.; Vibe-Pedersen, K.; Baralle, F.E. Primary structure of human fibronectin: Differential splicing may generate at least 10 polypeptides from a single gene. EMBO J. 1985, 4, 1755–1759. [Google Scholar] [PubMed]
  31. Ffrench-Constant, C.; Van De Water, L.; Dvorak, H.F.; Hynes, R.O. Reappearance of an embryonic pattern of fibronectin splicing during wound healing in the adult rat. J. Cell Biol. 1989, 109, 903–914. [Google Scholar] [CrossRef] [PubMed]
  32. Kornblihtt, A.R.; Pesce, C.G.; Alonso, C.R.; Cramer, A.; Srebrow, A.; Werbajh, S.; Muro, A.F. The fibronectin gene as a model for splicing and transcription studies. FASEB J. 1996, 10, 248–257. [Google Scholar] [CrossRef] [PubMed]
  33. Colombi, M.; Moro, L.; Zoppi, N.; Ghinelli, A.; Barlati, S. Altered fibronectin mRNA splicing in skin fibroblasts from Ehlers-Danlos syndrome patients: In situ hybridization analysis. Cell Biol. Int. Rep. 1991, 15, 1195–1206. [Google Scholar] [CrossRef]
  34. Muro, A.F.; Chauhan, A.K.; Gajovic, S.; Iaconcig, A.; Porro, F.; Stanta, G.; Baralle, F.E. Regulated splicing of the fibronectin EDA exon is essential for proper skin wound healing and normal lifespan. J. Cell Biol. 2003, 162, 149–160. [Google Scholar] [CrossRef] [PubMed]
  35. Gutman, A.; Kornblihtt, A.R. Identification of a third region of cell-specific alternative splicing in human fibronectin mRNA. Proc. Natl. Acad. Sci. USA 1987, 84, 7179–7182. [Google Scholar] [CrossRef] [PubMed]
  36. Clark, R.A.; Winn, H.J.; Dvorak, H.F.; Colvin, R.B. Fibronectin beneath reepithelializing epidermis in vivo: Sources and significance. J. Investig. Dermatol. 1983, 80, 26s–30s. [Google Scholar] [CrossRef] [PubMed]
  37. Colombi, M.; Barlati, S.; Kornblihtt, A.; Baralle, F.E.; Vaheri, A. A family of fibronectin RNAs in human normal and transformed cells. Biochim. Biophys. Acta 1986, 868, 207–214. [Google Scholar] [CrossRef]
  38. Norton, P.A.; Hynes, R.O. Alternative splicing of chicken fibronectin in embryos and in normal and transformed cells. Mol. Cell. Biol. 1987, 7, 4297–4307. [Google Scholar] [CrossRef] [PubMed]
  39. Dufour, S.; Duband, J.L.; Kornblihtt, A.R.; Thiery, J.P. The role of fibronectins in embryonic cell migrations. Trends Genet. 1988, 4, 198–203. [Google Scholar] [CrossRef]
  40. Ffrench-Constant, C. Alternative splicing of fibronectin—Many different proteins but few different functions. Exp. Cell Res. 1995, 221, 261–271. [Google Scholar] [CrossRef] [PubMed]
  41. Serini, G.; Bochaton-Piallat, M.L.; Ropraz, P.; Geinoz, A.; Borsi, L.; Zardi, L.; Gabbiani, G. The fibronectin domain ED-A is crucial for myofibroblastic phenotype induction by transforming growth factor-β1. J. Cell Biol. 1998, 142, 873–881. [Google Scholar] [CrossRef] [PubMed]
  42. Schwartz, M.A. Integrins and extracellular matrix in mechanotransduction. Cold Spring Harb. Perspect. Biol. 2010, 2, a005066. [Google Scholar] [CrossRef] [PubMed]
  43. Campbell, I.D.; Humphries, M.J. Integrin structure, activation, and interactions. Cold Spring Harb. Perspect. Biol. 2011, 3, a004994. [Google Scholar] [CrossRef] [PubMed]
  44. Geiger, B.; Yamada, K.M. Molecular architecture and function of matrix adhesions. Cold Spring Harb. Perspect. Biol. 2011, 3, a005033. [Google Scholar] [CrossRef] [PubMed]
  45. Huttenlocher, A.; Horwitz, A.R. Integrins in cell migration. Cold Spring Harb. Perspect. Biol. 2011, 3, a005074. [Google Scholar] [CrossRef] [PubMed]
  46. Watt, F.M.; Fujiwara, H. Cell-extracellular matrix interactions in normal and diseased skin. Cold Spring Harb. Perspect. Biol. 2011, 3, a005124. [Google Scholar] [CrossRef] [PubMed]
  47. Wickström, S.A.; Fässler, R. Regulation of membrane traffic by integrin signaling. Trends Cell Biol. 2011, 21, 266–273. [Google Scholar] [CrossRef] [PubMed]
  48. Sun, Z.; Guo, S.S.; Fässler, R. Integrin-mediated mechanotransduction. J. Cell Biol. 2016, 215, 445–456. [Google Scholar] [CrossRef] [PubMed]
  49. Horwitz, A.; Duggan, K.; Buck, C.; Beckerle, M.C.; Burridge, K. Interaction of plasma membrane fibronectin receptor with talin—A transmembrane linkage. Nature 1986, 320, 531–533. [Google Scholar] [CrossRef] [PubMed]
  50. Meredith, J.E.; Fazeli, B.; Schwartz, M.A. The extracellular matrix as a cell survival factor. Mol. Biol. Cell 1993, 4, 953–961. [Google Scholar] [CrossRef] [PubMed]
  51. Schellenberg, B.; Wang, P.; Keeble, J.A.; Rodriguez-Enriquez, R.; Walker, S.; Owens, T.W.; Foster, F.; Tanianis-Hughes, J.; Brennan, K.; Streuli, C.H.; et al. Bax exists in a dynamic equilibrium between the cytosol and mitochondria to control apoptotic priming. Mol. Cell 2013, 49, 959–971. [Google Scholar] [CrossRef] [PubMed]
  52. Ruoslahti, E.; Giancotti, F.G. Integrins and tumor cell dissemination. Cancer Cells 1989, 1, 119–126. [Google Scholar] [PubMed]
  53. Zaidel-Bar, R.; Cohen, M.; Addadi, L.; Geiger, B. Hierarchical assembly of cell-matrix adhesion complexes. Biochem. Soc. Trans. 2004, 32, 416–420. [Google Scholar] [CrossRef] [PubMed]
  54. Rossier, O.; Giannone, G. The journey of integrins and partners in a complex interactions landscape studied by super-resolution microscopy and single protein tracking. Exp. Cell Res. 2016, 343, 28–34. [Google Scholar] [CrossRef] [PubMed]
  55. Harjanto, D.; Zaman, M.H. Matrix mechanics and receptor-ligand interactions in cell adhesion. Org. Biomol. Chem. 2010, 8, 299–304. [Google Scholar] [CrossRef] [PubMed]
  56. Zamir, E.; Katz, M.; Posen, Y.; Erez, N.; Yamada, K.M.; Katz, B.Z.; Lin, S.; Lin, D.C.; Bershadsky, A.; Kam, Z.; et al. Dynamics and segregation of cell-matrix adhesions in cultured fibroblasts. Nat. Cell Biol. 2000, 2, 191–196. [Google Scholar] [CrossRef] [PubMed]
  57. Zamir, E.; Geiger, B. Molecular complexity and dynamics of cell-matrix adhesions. J. Cell Sci. 2001, 114, 3583–3590. [Google Scholar] [PubMed]
  58. Pankov, R.; Cukierman, E.; Katz, B.Z.; Matsumoto, K.; Lin, D.C.; Lin, S.; Hahn, C.; Yamada, K.M. Integrin dynamics and matrix assembly: Tensin-dependent translocation of α5β1 integrins promotes early fibronectin fibrillogenesis. J. Cell Biol. 2000, 148, 1075–1090. [Google Scholar] [CrossRef] [PubMed]
  59. Horton, M.A. The αvβ3 integrin “vitronectin receptor”. Int. J. Biochem. Cell Biol. 1997, 29, 721–725. [Google Scholar] [CrossRef]
  60. Dejana, E.; Raiteri, M.; Resnati, M.; Lampugnani, M.G. Endothelial integrins and their role in maintaining the integrity of the vessel wall. Kidney Int. 1993, 43, 61–65. [Google Scholar] [CrossRef] [PubMed]
  61. Brooks, P.C.; Clark, R.A.; Cheresh, D.A. Requirement of vascular integrin αvβ3 for angiogenesis. Science 1994, 264, 569–571. [Google Scholar] [CrossRef] [PubMed]
  62. Moiseeva, E.P. Adhesion receptors of vascular smooth muscle cells and their functions. Cardiovasc. Res. 2001, 52, 372–386. [Google Scholar] [CrossRef]
  63. Sadeghi, M.M.; Bender, J.R. Activated αvβ3 integrin targeting in injury-induced vascular remodeling. Trends Cardiovasc. Med. 2007, 17, 5–10. [Google Scholar] [CrossRef] [PubMed]
  64. Robinson, S.D.; Hodivala-Dilke, K.M. The role of β3-integrins in tumor angiogenesis: Context is everything. Curr. Opin. Cell Biol. 2011, 23, 630–637. [Google Scholar] [CrossRef] [PubMed]
  65. Kokubo, T.; Uchida, H.; Choi, E.T. Integrin αvβ3 as a target in the prevention of neointimal hyperplasia. J. Vasc. Surg. 2007, 45, A33–A38. [Google Scholar] [CrossRef] [PubMed]
  66. Hinz, B.; Phan, S.H.; Thannickal, V.J.; Prunotto, M.; Desmoulière, A.; Varga, J.; De Wever, O.; Mareel, M.; Gabbiani, G. Recent developments in myofibroblast biology: Paradigms for connective tissue remodeling. Am. J. Pathol. 2012, 180, 1340–1355. [Google Scholar] [CrossRef] [PubMed]
  67. Hinz, B. Myofibroblasts. Exp. Eye Res. 2016, 142, 56–70. [Google Scholar] [CrossRef] [PubMed]
  68. Gabbiani, G. The myofibroblast in wound healing and fibrocontractive diseases. J. Pathol. 2003, 200, 500–503. [Google Scholar] [CrossRef] [PubMed]
  69. Kajihara, I.; Jinnin, M.; Honda, N.; Makino, K.; Makino, T.; Masuguchi, S.; Sakai, K.; Fukushima, S.; Inoue, Y.; Ihn, H. Scleroderma dermal fibroblasts overexpress vascular endothelial growth factor due to autocrine transforming growth factor β signaling. Mod. Rheumatol. 2013, 23, 516–524. [Google Scholar] [CrossRef] [PubMed]
  70. Flavell, S.J.; Hou, T.Z.; Lax, S.; Filer, A.D.; Salmon, M.; Buckley, C.D. Fibroblasts as novel therapeutic targets in chronic inflammation. Br. J. Pharmacol. 2008, 153 (Suppl. S1), S241–S246. [Google Scholar] [CrossRef] [PubMed]
  71. Tadokoro, S.; Shattil, S.J.; Eto, K.; Tai, V.; Liddington, R.C.; de Pereda, J.M.; Ginsberg, M.H.; Calderwood, D.A. Talin binding to integrin β tails: A final common step in integrin activation. Science 2003, 302, 103–106. [Google Scholar] [CrossRef] [PubMed]
  72. Ma, Y.Q.; Qin, J.; Wu, C.; Plow, E.F. Kindlin-2 (Mig-2): A co-activator of β3 integrins. J. Cell Biol. 2008, 181, 439–446. [Google Scholar] [CrossRef] [PubMed]
  73. Moser, M.; Legate, K.R.; Zent, R.; Fässler, R. The tail of integrins, talin, and kindlins. Science 2009, 324, 895–899. [Google Scholar] [CrossRef] [PubMed]
  74. Giancotti, F.G.; Ruoslahti, E. Integrin signaling. Science 1999, 285, 1028–1032. [Google Scholar] [CrossRef] [PubMed]
  75. Qin, J.; Vinogradova, O.; Plow, E.F. Integrin bidirectional signaling: A molecular view. PLoS Biol. 2004, 2, e169. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Wary, K.K.; Mainiero, F.; Isakoff, S.J.; Marcantonio, E.E.; Giancotti, F.G. The adaptor protein Shc couples a class of integrins to the control of cell cycle progression. Cell 1996, 87, 733–743. [Google Scholar] [CrossRef]
  77. Matter, M.L.; Ruoslahti, E. A signaling pathway from the α5β1 and αvβ3 integrins that elevates bcl-2 transcription. J. Biol. Chem. 2001, 276, 27757–27763. [Google Scholar] [CrossRef] [PubMed]
  78. Strömblad, S.; Becker, J.C.; Yebra, M.; Brooks, P.C.; Cheresh, D.A. Suppression of p53 activity and p21WAF1/CIP1 expression by vascular cell integrin αvβ3 during angiogenesis. J. Clin. Investig. 1996, 98, 426–433. [Google Scholar] [CrossRef] [PubMed]
  79. Zhang, Y.; Lu, H.; Dazin, P.; Kapila, Y. Squamous cell carcinoma cell aggregates escape suspension-induced, p53-mediated anoikis: Fibronectin and integrin αv mediate survival signals through focal adhesion kinase. J. Biol. Chem. 2004, 279, 48342–48349. [Google Scholar] [CrossRef] [PubMed]
  80. Nieberler, M.; Reuning, U.; Reichart, F.; Notni, J.; Wester, H.J.; Schwaiger, M.; Weinmüller, M.; Räder, A.; Steiger, K.; Kessler, H. Exploring the Role of RGD-Recognizing Integrins in Cancer. Cancers 2017, 9, 116. [Google Scholar] [CrossRef] [PubMed]
  81. Yamada, K.M.; Even-Ram, S. Integrin regulation of growth factor receptor. Nat. Cell Biol. 2002, 4, E75–E76. [Google Scholar] [CrossRef] [PubMed]
  82. Schwartz, M.A.; Ginsberg, M.H. Networks and crosstalk: Integrin signaling spreads. Nat. Cell Biol. 2002, 4, E65–E68. [Google Scholar] [CrossRef] [PubMed]
  83. Galliher, A.J.; Schiemann, W.P. β3 integrin and Src facilitate transforming growth factor-β mediated induction of epithelial-mesenchymal transition in mammary epithelial cells. Breast Cancer Res. 2006, 8, R42. [Google Scholar] [CrossRef] [PubMed]
  84. Guo, W.; Giancotti, F.G. Integrin signaling during tumour progression. Nat. Rev. Mol. Cell Biol. 2004, 5, 816–826. [Google Scholar] [CrossRef] [PubMed]
  85. Yeh, Y.Y.; Chiao, C.C.; Kuo, W.Y.; Hsiao, Y.C.; Chen, Y.J.; Wei, Y.Y.; Lai, T.H.; Fong, Y.C.; Tang, C.H. TGF-β1 increases motility and αvβ3 integrin up-regulation via PI3K, Akt and NF-kappaB-dependent pathway in human chondrosarcoma cells. Biochem. Pharmacol. 2008, 75, 1292–1301. [Google Scholar] [CrossRef] [PubMed]
  86. Kwakwa, K.A.; Sterling, J.A. Integrin αvβ3 Signaling in Tumor-Induced Bone Disease. Cancers 2017, 9, 84. [Google Scholar] [CrossRef] [PubMed]
  87. Scaffidi, A.K.; Petrovic, N.; Moodley, Y.P.; Fogel-Petrovic, M.; Kroeger, K.M.; Seeber, R.M.; Eidne, K.A.; Thompson, P.J.; Knight, D.A. αvβ3 Integrin interacts with the transforming growth factor β (TGFβ) type II receptor to potentiate the proliferative effects of TGFβ1 in living human lung fibroblasts. J. Biol. Chem. 2004, 279, 37726–37733. [Google Scholar] [CrossRef] [PubMed]
  88. Van Laer, L.; Proost, D.; Loeys, B.L. Educational paper. Connective tissue disorders with vascular involvement: From gene to therapy. Eur. J. Pediatr. 2013, 172, 997–1005. [Google Scholar] [CrossRef] [PubMed]
  89. Meester, J.A.N.; Verstraeten, A.; Schepers, D.; Alaerts, M.; Van Laer, L.; Loeys, B.L. Differences in manifestations of Marfan syndrome, Ehlers-Danlos syndrome, and Loeys-Dietz syndrome. Ann. Cardiothorac. Surg. 2017, 6, 582–594. [Google Scholar] [CrossRef] [PubMed]
  90. Malfait, F.; Francomano, C.; Byers, P.; Belmont, J.; Berglund, B.; Black, J.; Bloom, L.; Bowen, J.M.; Brady, A.F.; Burrows, N.P.; et al. The 2017 international classification of the Ehlers-Danlos syndromes. Am. J. Med. Genet. C Semin. Med. Genet. 2017, 175, 8–26. [Google Scholar] [CrossRef] [PubMed]
  91. Bowen, J.M.; Sobey, G.J.; Burrows, N.P.; Colombi, M.; Lavallee, M.E.; Malfait, F.; Francomano, C.A. Ehlers-Danlos syndrome, classical type. Am. J. Med. Genet. C Semin. Med. Genet. 2017, 175, 27–39. [Google Scholar] [CrossRef] [PubMed]
  92. Symoens, S.; Syx, D.; Malfait, F.; Callewaert, B.; De Backer, J.; Vanakker, O.; Coucke, P.; De Paepe, A. Comprehensive molecular analysis demonstrates type V collagen mutations in over 90% of patients with classic EDS and allows to refine diagnostic criteria. Hum. Mutat. 2012, 33, 1485–1493. [Google Scholar] [CrossRef] [PubMed]
  93. Ritelli, M.; Dordoni, C.; Venturini, M.; Chiarelli, N.; Quinzani, S.; Traversa, M.; Zoppi, N.; Vascellaro, A.; Wischmeijer, A.; Manfredini, E.; et al. Clinical and molecular characterization of 40 patients with classic Ehlers-Danlos syndrome: Identification of 18 COL5A1 and 2 COL5A2 novel mutations. Orphanet J. Rare Dis. 2013, 8, 58. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Byers, P.H.; Belmont, J.; Black, J.; De Backer, J.; Frank, M.; Jeunemaitre, X.; Johnson, D.; Pepin, M.; Robert, L.; Sanders, L.; et al. Diagnosis, natural history, and management in vascular Ehlers-Danlos syndrome. Am. J. Med. Genet. C Semin. Med. Genet. 2017, 175, 40–47. [Google Scholar] [CrossRef] [PubMed]
  95. Pepin, M.; Schwarze, U.; Superti-Furga, A.; Byers, P.H. Clinical and genetic features of Ehlers-Danlos syndrome type IV, the vascular type. N. Engl. J. Med. 2000, 342, 673–680. [Google Scholar] [CrossRef] [PubMed]
  96. Tinkle, B.; Castori, M.; Berglund, B.; Cohen, H.; Grahame, R.; Kazkaz, H.; Levy, H. Hypermobile Ehlers-Danlos syndrome (a.k.a. Ehlers-Danlos syndrome type III and Ehlers-Danlos syndrome hypermobility type): Clinical description and natural history. Am. J. Med. Genet. C Semin. Med. Genet. 2017, 175, 48–69. [Google Scholar] [CrossRef] [PubMed]
  97. Castori, M.; Tinkle, B.; Levy, H.; Grahame, R.; Malfait, F.; Hakim, A. A framework for the classification of joint hypermobility and related conditions. Am. J. Med. Genet. C Semin. Med. Genet. 2017, 175, 148–157. [Google Scholar] [CrossRef] [PubMed]
  98. Coucke, P.J.; Willaert, A.; Wessels, M.W.; Callewaert, B.; Zoppi, N.; De Backer, J.; Fox, J.E.; Mancini, G.M.; Kambouris, M.; Gardella, R.; et al. Mutations in the facilitative glucose transporter GLUT10 alter angiogenesis and cause arterial tortuosity syndrome. Nat. Genet. 2006, 38, 452–457. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  99. Németh, C.E.; Marcolongo, P.; Gamberucci, A.; Fulceri, R.; Benedetti, A.; Zoppi, N.; Ritelli, M.; Chiarelli, N.; Colombi, M.; Willaert, A.; et al. Glucose transporter type 10-lacking in arterial tortuosity syndrome-facilitates dehydroascorbic acid transport. FEBS Lett. 2016, 590, 1630–1640. [Google Scholar] [CrossRef] [PubMed]
  100. Van Dijk, F.S.; Sillence, D.O. Osteogenesis imperfecta: Clinical diagnosis, nomenclature and severity assessment. Am. J. Med. Genet. A 2014, 164, 1470–1481. [Google Scholar] [CrossRef] [PubMed]
  101. Marini, J.C.; Forlino, A.; Bächinger, H.P.; Bishop, N.J.; Byers, P.H.; Paepe, A.; Fassier, F.; Fratzl-Zelman, N.; Kozloff, K.M.; Krakow, D.; et al. Osteogenesis imperfecta. Nat. Rev. Dis. Primers 2017, 3, 17052. [Google Scholar] [CrossRef] [PubMed]
  102. Zoppi, N.; Gardella, R.; De Paepe, A.; Barlati, S.; Colombi, M. Human fibroblasts with mutations in COL5A1 and COL3A1 genes do not organize collagens and fibronectin in the extracellular matrix, down-regulate α2β1 integrin, and recruit αvβ3 instead of α5β1 integrin. J. Biol. Chem. 2004, 30, 18157–18168. [Google Scholar] [CrossRef] [PubMed]
  103. Zoppi, N.; Barlati, S.; Colombi, M. FAK-independent αvβ3 integrin-EGFR complexes rescue from anoikis matrix-defective fibroblasts. Biochim. Biophys. Acta 2008, 1783, 1177–1188. [Google Scholar] [CrossRef] [PubMed]
  104. Zoppi, N.; Ritelli, M.; Colombi, M. Type III and V collagens modulate the expression and assembly of EDA+ fibronectin in the extracellular matrix of defective Ehlers-Danlos syndrome fibroblasts. Biochim. Biophys. Acta 2012, 1820, 1576–1587. [Google Scholar] [CrossRef] [PubMed]
  105. Zoppi, N.; Chiarelli, N.; Cinquina, V.; Ritelli, M.; Colombi, M. GLUT10 deficiency leads to oxidative stress and non-canonical αvβ3 integrin-mediated TGFβ signaling associated with extracellular matrix disarray in arterial tortuosity syndrome skin fibroblasts. Hum. Mol. Genet. 2015, 24, 6769–6787. [Google Scholar] [CrossRef] [PubMed]
  106. Zoppi, N.; Chiarelli, N.; Binetti, S.; Ritelli, M.; Colombi, M. Dermal fibroblast-to-myofibroblast transition sustained by αvβ3 integrin-ILK-Snail1/Slug signaling is a common feature for hypermobile Ehlers-Danlos syndrome and hypermobility spectrum disorders. Biochim. Biophys. Acta 2018, 1864, 1010–1023. [Google Scholar] [CrossRef] [PubMed]
  107. Chiarelli, N.; Carini, G.; Zoppi, N.; Dordoni, C.; Ritelli, M.; Venturini, M.; Castori, M.; Colombi, M. Transcriptome-wide expression profiling in skin fibroblasts of patients with joint hypermobility syndrome/Ehlers-Danlos syndrome hypermobility type. PLoS ONE 2016, 11, e0161347. [Google Scholar] [CrossRef] [PubMed]
  108. Chiarelli, N.; Carini, G.; Zoppi, N.; Ritelli, M.; Colombi, M. Transcriptome analysis of skin fibroblasts with dominant negative COL3A1 mutations provides molecular insights into the etiopathology of vascular Ehlers-Danlos syndrome. PLoS ONE 2018, 13, e0191220. [Google Scholar] [CrossRef] [PubMed]
  109. Ritelli, M.; Chiarelli, N.; Zoppi, N.; Dordoni, C.; Quinzani, S.; Traversa, M.; Venturini, M.; Calzavara-Pinton, P.; Colombi, M. Insights in the etiopathology of galactosyltransferase II (GalT-II) deficiency from transcriptome-wide expression profiling of skin fibroblasts of two sisters with compound heterozygosity for two novel B3GALT6 mutations. Mol. Genet. Metab. Rep. 2014, 2, 1–15. [Google Scholar] [CrossRef] [PubMed]
  110. Baumann, M.; Giunta, C.; Krabichler, B.; Rüschendorf, F.; Zoppi, N.; Colombi, M.; Bittner, R.E.; Quijano-Roy, S.; Muntoni, F.; Cirak, S.; et al. Mutations in FKBP14 cause a variant of Ehlers-Danlos syndrome with progressive kyphoscoliosis, myopathy, and hearing loss. Am. J. Hum. Genet. 2012, 90, 201–216. [Google Scholar] [CrossRef] [PubMed]
  111. Burkitt Wright, E.M.M.; Spencer, H.L.; Daly, S.B.; Manson, F.D.C.; Zeef, L.A.H.; Urquhart, J.; Zoppi, N.; Bonshek, R.; Tosounidis, I.; Mohan, M.; et al. Mutations in PRDM5 in brittle cornea syndrome identify a pathway regulating extracellular matrix development and maintenance. Am. J. Hum. Genet. 2011, 88, 767–777. [Google Scholar] [CrossRef] [PubMed]
  112. Porter, L.F.; Gallego-Pinazo, R.; Keeling, C.L.; Kamieniorz, M.; Zoppi, N.; Colombi, M.; Giunta, C.; Bonshek, R.; Manson, F.D.; Black, G.C. Bruch’s membrane abnormalities in PRDM5-related brittle cornea syndrome. Orphanet J. Rare Dis. 2015, 10, 145. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  113. Janecke, A.R.; Li, B.; Boehm, M.; Krabichler, B.; Rohrbach, M.; Müller, T.; Fuchs, I.; Golas, G.; Katagiri, Y.; Ziegler, S.G.; et al. The phenotype of the musculocontractural type of Ehlers-Danlos syndrome due to CHST14 mutations. Am. J. Med. Genet. A 2016, 170, 103–115. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  114. Gardella, R.; Zoppi, N.; Assanelli, D.; Muiesan, M.L.; Barlati, S.; Colombi, M. Exclusion of candidate genes in a family with arterial tortuosity syndrome. Am. J. Med. Genet. A 2004, 126, 221–228. [Google Scholar] [CrossRef] [PubMed]
  115. Segade, F. Glucose transporter 10 and arterial tortuosity syndrome: The vitamin C connection. FEBS Lett. 2010, 584, 2990–2994. [Google Scholar] [CrossRef] [PubMed]
  116. Gamberucci, A.; Marcolongo, P.; Németh, C.E.; Zoppi, N.; Szarka, A.; Chiarelli, N.; Hegedűs, T.; Ritelli, M.; Carini, G.; Willaert, A.; et al. GLUT10-Lacking in Arterial Tortuosity Syndrome—Is Localized to the Endoplasmic Reticulum of Human Fibroblasts. Int. J. Mol. Sci. 2017, 18, 1820. [Google Scholar] [CrossRef] [PubMed]
  117. Wu, C.; Hughes, P.E.; Ginsberg, M.H.; McDonald, J.A. Identification of a new biological function for the integrin αvβ3: Initiation of fibronectin matrix assembly. Cell Adhes. Commun. 1996, 4, 149–158. [Google Scholar] [CrossRef] [PubMed]
  118. Folkman, J.; Moscona, A. Role of cell shape in growth control. Nature 1978, 273, 345–349. [Google Scholar] [CrossRef] [PubMed]
  119. Re, F.; Zanetti, A.; Sironi, M.; Polentarutti, N.; Lanfrancone, L.; Detona, E.; Calotta, F. Inhibition of anchorage-dependent cell spreading triggers apoptosis in cultured endothelial cells. J. Cell Biol. 1994, 127, 537–546. [Google Scholar] [CrossRef] [PubMed]
  120. Frisch, S.M.; Francis, H. Disruption of epithelial cell matrix interactions induces apoptosis. J. Cell Biol. 1994, 124, 619–626. [Google Scholar] [CrossRef] [PubMed]
  121. Frisch, S.M.; Ruoslahti, E. Integrin and anoikis. Curr. Opin. Cell Biol. 1997, 9, 701–706. [Google Scholar] [CrossRef]
  122. McGill, G.; Shimamura, A.; Bates, R.C.; Savage, R.E.; Fischer, D.E. Loss of matrix adhesion triggers rapid transformation-selective apoptosis in fibroblasts. J. Cell Biol. 1997, 138, 901–911. [Google Scholar] [CrossRef] [PubMed]
  123. Sechler, J.L.; Schwarzbauer, J.E. Control of cell cycle progression by fibronectin matrix architecture. J. Biol. Chem. 1998, 273, 25533–25536. [Google Scholar] [CrossRef] [PubMed]
  124. Stupack, D.G.; Cheresh, D.A. Get a ligand, get a life: Integrins, signaling and cell survival. J. Cell Sci. 2002, 115, 3729–3738. [Google Scholar] [CrossRef] [PubMed]
  125. Gu, J.; Fujibayashi, A.; Yamada, K.M.; Sekiguchi, K. Laminin-10/11 and fibronectin differentially prevent apoptosis induced by serum removal via phosphatidylinositol 3-kinase/Akt- and MEK1/ERK-dependent pathways. J. Biol. Chem. 2002, 277, 19922–19928. [Google Scholar] [CrossRef] [PubMed]
  126. Brooks, P.C.; Montgomery, A.M.P.; Rosenfeld, M.; Reisfeld, R.A.; Hu, T.; Klier, G.; Cheresh, D.A. Integrin avb3 antagonists promote tumor regression by inducing apoptosis of angiogenic blood vessels. Cell 1994, 79, 1157–1164. [Google Scholar] [CrossRef]
  127. Brooks, P.C.; Stromblad, S.; Klemke, R.; Visscher Sarkar, D.F.H.; Cheresh, D.A. Antiintegrin αvβ3 blocks human breast cancer growth and angiogenesis in human skin. J. Clin. Investig. 1995, 96, 1815–1822. [Google Scholar] [CrossRef] [PubMed]
  128. Khwaja, A. Apoptosis: Akt is more than just a Bad kinase. Nature 1999, 401, 33–34. [Google Scholar] [CrossRef] [PubMed]
  129. Rosen, I.; Rak, J.; Leung, T.; Dean, N.M.; Kerbel, R.S.; Filmus, J. Activated Ras prevents down regulation of Bcl-X(L) triggered by detachment from the extracellular matrix. A mechanism of Ras-induced resistance to anoikis in intestinal epithelial cells. J. Cell Biol. 2000, 149, 447–456. [Google Scholar] [PubMed]
  130. Kook, S.; Kim, D.H.; Shim, S.R.; Kim, W.; Chun, J.S.; Song, W.K. Caspase-dependent cleavage of tensin induces disruption of actin cytoskeleton during apoptosis. Biochem. Biophys. Res. Commun. 2003, 303, 37–45. [Google Scholar] [CrossRef]
  131. Schaller, M.D. Paxillin: A focal adhesion-associated adaptor protein. Oncogene 2001, 20, 6459–6472. [Google Scholar] [CrossRef] [PubMed]
  132. Cukierman, E.; Pankov, R.; Yamada, K.M. Cell interaction with three-dimensional matrices. Curr. Opin. Cell Biol. 2002, 14, 633–639. [Google Scholar] [CrossRef]
  133. Nikolopoulos, S.N.; Turner, C.E. Integrin-linked kinase (ILK) binding to paxillin LD1 motif regulates ILK localization to focal adhesions. J. Biol. Chem. 2001, 276, 23499–23505. [Google Scholar] [CrossRef] [PubMed]
  134. Turner, C.E. Paxillin and focal adhesion signaling. Nat. Cell Biol. 2000, 2, E231–E236. [Google Scholar] [CrossRef] [PubMed]
  135. Bannerman, D.D.; Sathyamoorthy, M.; Goldblum, S.E. Bacterial lipopolysaccharide disrupts endothelial monolayer integrity and survival signaling events through caspase cleavage of adherens junction proteins. J. Biol. Chem. 1998, 273, 35371–35380. [Google Scholar] [CrossRef] [PubMed]
  136. Moro, I.; Venturino, M.; Bozzo, C.; Silengo, L.; Altruda, F.; Beguinot, L.; Tarone, G.; Defilippi, P. Integrins induce activation of EGF receptor: Role in MAP kinase induction and adhesion-dependent cell survival. EMBO J. 1998, 17, 6622–6632. [Google Scholar] [CrossRef] [PubMed]
  137. Tapia, J.A.; Camello, C.; Jensen, R.T.; Garcia, L.J. EGF stimulates tyrosine phosphorylation of focal adhesion kinase (p125FAK) and paxillin in rat pancreatic acini by a phospholipase C-independent process that depends by phosphatidylinositol 3-kinase, the small GTP-binding protein, p21rho, and the integrity of the actin cytoskeleton. Biochim. Biophys. Acta 1999, 1448, 486–499. [Google Scholar] [PubMed]
  138. Brown, M.C.; Turner, C.E. Paxillin: Adapting to change. Physiol. Rev. 2004, 84, 1339–1351. [Google Scholar] [CrossRef] [PubMed]
  139. Ishikawa, Y.; Bächinger, H.P. A substrate preference for the rough endoplasmic reticulum resident protein FKBP22 during collagen biosynthesis. J. Biol. Chem. 2014, 289, 18189–18201. [Google Scholar] [CrossRef] [PubMed]
  140. Gjaltema, R.A.; Bank, R.A. Molecular insights into prolyl and lysyl hydroxylation of fibrillar collagens in health and disease. Crit. Rev. Biochem. Mol. Biol. 2017, 52, 74–95. [Google Scholar] [CrossRef] [PubMed]
  141. Szegezdi, E.; Logue, S.E.; Gorman, A.M.; Samali, A. Mediators of endoplasmic reticulum stress-induced apoptosis. EMBO Rep. 2006, 7, 880–885. [Google Scholar] [CrossRef] [PubMed]
  142. Lu, M.; Miao, Y.; Qi, L.; Bai, M.; Zhang, J.; Feng, Y. RNAi-Mediated Downregulation of FKBP14 Suppresses the Growth of Human Ovarian Cancer Cells. Oncol. Res. 2016, 23, 267–274. [Google Scholar] [CrossRef] [PubMed]
  143. Asano, Y.; Ihn, H.; Yamane, K.; Jinnin, M.; Mimura, Y.; Tamaki, K. Increased expression of integrin αvβ3 contributes to the establishment of autocrine TGF-β signaling in scleroderma fibroblasts. J. Immunol. 2005, 175, 7708–7718. [Google Scholar] [CrossRef] [PubMed]
  144. Sarrazy, V.; Koehler, A.; Chow, M.L.; Zimina, E.; Li, C.X.; Kato, H.; Caldarone, C.A.; Hinz, B. Integrins αvβ5 and αvβ3 promote latent TGF-β1 activation by human cardiac fibroblast contraction. Cardiovasc. Res. 2014, 102, 407–417. [Google Scholar] [CrossRef] [PubMed]
  145. Dave, I.; Guaita-Esteruelas, S.; Gutarra, S.; Àlex Frias, M.; Beltran, S.; Peiró García de Herreros, A. functional cooperation between Snail1 and twist in the regulation of ZEB1 expression during epithelial to mesenchymal transition. J. Biol. Chem. 2011, 286, 12024–12032. [Google Scholar] [CrossRef] [PubMed]
  146. Kasperkovitz, P.V.; Timmer, T.C.; Smeets, T.J.; Verbeet, N.L.; Tak, P.P.; van Baarsen, L.G.; Baltus, B.; Huizinga, T.W.; Pieterman, E.; Fero, M.; et al. Fibroblast-like synoviocytes derived from patients with rheumatoid arthritis show the imprint of synovial tissue heterogeneity: Evidence of a link between an increased myofibroblast-like phenotype and high-inflammation synovitis. Arthritis Rheum. 2005, 52, 430–441. [Google Scholar] [CrossRef] [PubMed]
  147. Song, H.Y.; Kim, M.Y.; Kim, K.H.; Lee, I.H.; Shin, S.H.; Lee, J.S.; Kim, J.H. Synovial fluid of patients with rheumatoid arthritis induces α-smooth muscle actin in human adipose tissue-derived mesenchymal stem cells through a TGF-β1-dependent mechanism. Exp. Mol. Med. 2010, 42, 565–573. [Google Scholar] [CrossRef] [PubMed]
  148. Chen, S.Y.; Shiau, A.L.; Li, Y.T.; Lin, C.C.; Jou, I.M.; Liu, M.F.; Wu, C.L.; Wang, C.R. Transcription factor snail regulates tumor necrosis factor α-mediated synovial fibroblast activation in the rheumatoid joint. Arthritis Rheum. 2015, 67, 39–50. [Google Scholar] [CrossRef] [PubMed]
  149. Xu, W.; Yang, Z.; Lu, N. A new role for the PI3K/Akt signaling pathway in the epithelial-mesenchymal transition. Cell Adhes. Migr. 2015, 9, 317–324. [Google Scholar] [CrossRef] [PubMed]
  150. Hannigan, G.; Troussard, A.A.; Dedhar, S. Integrin-linked kinase: A cancer therapeutic target unique among its ILK. Nat. Rev. Cancer 2005, 5, 51–63. [Google Scholar] [CrossRef] [PubMed]
  151. Delcommenne, M.; Tan, C.; Gray, V.; Rue, L.; Woodgett, J.; Dedhar, S. Phosphoinositide-3-OH kinase-dependent regulation of glycogen synthase kinase 3 and protein kinase B/AKT by the integrin-linked kinase. Proc. Natl. Acad. Sci. USA 1998, 95, 11211–112116. [Google Scholar] [CrossRef] [PubMed]
  152. Zhou, B.P.; Deng, J.; Xia, W.; Xu, J.; Li, Y.M.; Gunduz, M.; Hung, M.C. Dual regulation of snail by GSK-3β-mediated phosphorylation in control of epithelial-mesenchymal transition. Nat. Cell Biol. 2004, 6, 931–940. [Google Scholar] [CrossRef] [PubMed]
  153. Serrano-Gomez, S.J.; Maziveyi, M.; Alahari, S.K. Regulation of epithelial-mesenchymal transition through epigenetic and post-translational modifications. Mol. Cancer 2016, 15, 18. [Google Scholar] [CrossRef] [PubMed]
  154. Lee, S.L.; Hsu, E.C.; Chou, C.C.; Chuang, H.C.; Bai, L.Y.; Kulp, S.K.; Chen, C.S. Identification and characterization of a novel integrin-linked kinase inhibitor. J. Med. Chem. 2011, 54, 6364–6374. [Google Scholar] [CrossRef] [PubMed]
  155. Wu, Y.; Zhou, B.P. TNF-α/NF-κB/Snail pathway in cancer cell migration and invasion. Br. J. Cancer 2010, 102, 639–644. [Google Scholar] [CrossRef] [PubMed]
  156. Rosin, D.L.; Okusa, M.D. Dangers within: DAMP responses to damage and cell death in kidney disease. J. Am. Soc. Nephrol. 2011, 22, 416–425. [Google Scholar] [CrossRef] [PubMed]
  157. Anders, H.J.; Schaefer, L. Beyond tissue injury-damage-associated molecular patterns, toll-like receptors, and inflammasomes also drive regeneration and fibrosis. J. Am. Soc. Nephrol. 2014, 25, 1–14. [Google Scholar] [CrossRef] [PubMed]
  158. Tajerian, M.; Clark, J.D. The role of the extracellular matrix in chronic pain following injury. Pain 2015, 156, 366–370. [Google Scholar] [CrossRef] [PubMed]
  159. Kato, J.; Svensson, C.I. Role of extracellular damage-associated molecular pattern molecules (DAMPs) as mediators of persistent pain. Prog. Mol. Biol. Transl. Sci. 2015, 131, 251–279. [Google Scholar] [PubMed]
  160. Miller, R.E.; Belmadani, A.; Ishihara, S.; Tran, P.B.; Ren, D.; Miller, R.J.; Malfait, A.M. Damage-associated molecular patterns generated in osteoarthritis directly excite murine nociceptive neurons through toll-like receptor 4. Arthritis Rheumatol. 2015, 67, 2933–2943. [Google Scholar] [CrossRef] [PubMed]
  161. Castori, M.; Morlino, S.; Celletti, C.; Ghibellini, G.; Bruschini, M.; Grammatico, P.; Blundo, C.; Camerota, F. Re-writing the natural history of pain and related symptoms in the joint hypermobility syndrome/syndrome, hypermobility type. Am. J. Med. Genet. Part A 2013, 161, 2989–3004. [Google Scholar] [CrossRef] [PubMed]
  162. Syx, D.; De Wandele, I.; Rombaut, L.; Malfait, F. Hypermobility, the Ehlers-Danlos syndromes and chronic pain. Clin. Exp. Rheumatol. 2017, 107, 116–122. [Google Scholar]
  163. Lau, L.F. CCN1/CYR61: The very model of a modern matricellular protein. Cell. Mol. Life Sci. 2011, 68, 3149–3163. [Google Scholar] [CrossRef] [PubMed]
  164. Borkham-Kamphorst, E.; Schaffrath, C.; Van de Leur, E.; Haas, U.; Tihaa, L.; Meurer, S.K.; Nevzorova, Y.A.; Liedtke, C.; Weiskirchen, R. The anti-fibrotic effects of CCN1/CYR61 in primary portal myofibroblasts are mediated through induction of reactive oxygen species resulting in cellular senescence, apoptosis and attenuated TGF-β signaling. Biochim. Biophys. Acta 2014, 1843, 902–914. [Google Scholar] [CrossRef] [PubMed]
  165. Carthy, J.M.; Garmaroudi, F.S.; Luo, Z.; McManus, B.M. Wnt3a induces myofibroblast differentiation by upregulating TGF-β signaling through SMAD2 in a β-catenin-dependent manner. PLoS ONE 2011, 6, e19809. [Google Scholar] [CrossRef] [PubMed]
  166. Blyszczuk, I.; Müller-Edenborn, B.; Valenta, T.; Osto, E.; Stellato, M.; Behnke, S.; Glatz, K.; Basler, K.; Lüscher, T.F.; Distler, O.; et al. Transforming growth factor-β-dependent Wnt secretion controls myofibroblast formation and myocardial fibrosis progression in experimental autoimmune myocarditis. Eur. Heart J. 2017, 38, 1413–1425. [Google Scholar] [CrossRef] [PubMed]
  167. Babic, A.M.; Chen, C.C.; Lau, L.F. Fisp12/mouse connective tissue growth factor mediates endothelial cell adhesion and migration through integrin αvβ3, promotes endothelial cell survival, and induces angiogenesis in vivo. Mol. Cell Biol. 1999, 19, 2958–2966. [Google Scholar] [CrossRef] [PubMed]
  168. Nakerakanti, S.S.; Bujor, A.M.; Trojanowska, M. CCN2 is required for the TGF-β induced activation of Smad1-Erk1/2 signaling network. PLoS ONE 2011, 6, e21911. [Google Scholar] [CrossRef] [PubMed]
  169. Lee, S.W.; Won, J.Y.; Kim, W.J.; Lee, J.; Kim, K.H.; Youn, S.W.; Kim, J.Y.; Lee, E.J.; Kim, Y.J.; Kim, K.W.; et al. Snail as a potential target molecule in cardiac fibrosis: Paracrine action of endothelial cells on fibroblasts through snail and CTGF axis. Mol. Ther. 2013, 21, 1767–1777. [Google Scholar] [CrossRef] [PubMed]
  170. Wójcik-Pszczoła, K.; Jakieła, B.; Plutecka, H.; Koczurkiewicz, P.; Madeja, Z.; Michalik, M.; Sanak, M. Connective tissue growth factor regulates transition of primary bronchial fibroblasts to myofibroblasts in asthmatic subjects. Cytokine 2018, 102, 187–190. [Google Scholar] [CrossRef] [PubMed]
  171. Colombi, M.; Dordoni, C.; Chiarelli, N.; Ritelli, M. Differential diagnosis and diagnostic flow chart of joint hypermobility syndrome/ehlers-danlos syndrome hypermobility type compared to other heritable connective tissue disorders. Am. J. Med. Genet. C Semin. Med. Genet. 2015, 169, 6–22. [Google Scholar] [CrossRef] [PubMed]
  172. Lee, Y.C.; Huang, H.Y.; Chang, C.J.; Cheng, C.H.; Chen, Y.T. Mitochondrial GLUT10 facilitates dehydroascorbic acid import and protects cells against oxidative stress: Mechanistic insight into arterial tortuosity syndrome. Hum. Mol. Genet. 2010, 19, 3721–3733. [Google Scholar] [CrossRef] [PubMed]
  173. Annes, J.P.; Rifkin, D.B.; Munger, J.S. The integrin αvβ6 binds and activates latent TGFβ3. FEBS Lett. 2002, 511, 65–68. [Google Scholar] [CrossRef]
  174. Munger, J.S.; Sheppard, D. Cross talk among TGF-β signaling pathways, integrins, and the extracellular matrix. Cold Spring Harb. Perspect. Biol. 2011, 3, a005017. [Google Scholar] [CrossRef] [PubMed]
  175. Conroy, K.P.; Kitto, L.J.; Henderson, N.C. αv integrins: Key regulators of tissue fibrosis. Cell Tissue Res. 2016, 365, 511–519. [Google Scholar] [CrossRef] [PubMed]
  176. Liu, S.; Xu, S.W.; Kennedy, L.; Pala, D.; Chen, Y.; Eastwood, M.; Carter, D.E.; Black, C.M.; Abraham, D.J.; Leask, A. FAK is required for TGFβ-induced JNK phosphorylation in fibroblasts: Implications for acquisition of a matrix-remodeling phenotype. Mol. Biol. Cell 2007, 18, 2169–2178. [Google Scholar] [CrossRef] [PubMed]
  177. Leask, A. Focal adhesion kinase: A key mediator of transforming growth factor β signaling in fibroblasts. Adv. Wound Care 2013, 2, 247–249. [Google Scholar] [CrossRef] [PubMed]
  178. Valko, M.; Leibfritz, D.; Moncol, J.; Cronin, M.T.; Mazur, M.; Telser, J. Free radicals and antioxidants in normal physiological functions and human disease. Int. J. Biochem. Cell Biol. 2007, 39, 44–84. [Google Scholar] [CrossRef] [PubMed]
  179. He, T.; Quan, T.; Shao, Y.; Voorhees, J.J.; Fisher, G.J. Oxidative exposure impairs TGF-β pathway via reduction of type II receptor and SMAD3 in human skin fibroblasts. Age 2014, 36, 9623. [Google Scholar] [CrossRef] [PubMed]
  180. Larroque-Cardoso, P.; Mucher, E.; Grazide, M.H.; Josse, G.; Schmitt, A.M.; Nadal-Wolbold, F.; Zarkovic, K.; Salvayre, R.; Nègre-Salvayre, A. 4-Hydroxynonenal impairs transforming growth factor-β1-induced elastin synthesis via epidermal growth factor receptor activation in human and murine fibroblasts. Free Radic. Biol. Med. 2014, 71, 427–436. [Google Scholar] [CrossRef] [PubMed]
Ijms 19 00982 g001 550
Figure 1. Organization of FN–ECM and expression of the α5β1 and αvβ3 integrin receptors in control and HCTDs dermal fibroblasts. Among the analyzed HCTDs (Table 7), all EDS fibroblasts, except for spEDS-B3GALT6, and ATS cells show the FN–ECM disarray, the reduced expression of the canonical FN integrin receptor α5β1 and the consequent recruitment of the αvβ3 integrin. This peculiar phenotype is not observed in cells derived from MFS, LDS, and OI patients. Dermal fibroblasts were immunoreacted with antibodies against FN, α5β1, and αvβ3 integrins as previously described [102,106,107]. Scale bar: 10 μm.
Figure 1. Organization of FN–ECM and expression of the α5β1 and αvβ3 integrin receptors in control and HCTDs dermal fibroblasts. Among the analyzed HCTDs (Table 7), all EDS fibroblasts, except for spEDS-B3GALT6, and ATS cells show the FN–ECM disarray, the reduced expression of the canonical FN integrin receptor α5β1 and the consequent recruitment of the αvβ3 integrin. This peculiar phenotype is not observed in cells derived from MFS, LDS, and OI patients. Dermal fibroblasts were immunoreacted with antibodies against FN, α5β1, and αvβ3 integrins as previously described [102,106,107]. Scale bar: 10 μm.
Ijms 19 00982 g001
Ijms 19 00982 g002 550
Figure 2. Schematic representation of the αvβ3 integrin- and EGFR-mediated signaling pathways ensuring the rescue from anoikis in cEDS and vEDS fibroblasts.
Figure 2. Schematic representation of the αvβ3 integrin- and EGFR-mediated signaling pathways ensuring the rescue from anoikis in cEDS and vEDS fibroblasts.
Ijms 19 00982 g002
Ijms 19 00982 g003 550
Figure 3. Schematic representation of the αvβ3 integrin-ILK-Snail1 transduction pathway involved in the myofibroblast-like phenotype of hEDS and HSD cells.
Figure 3. Schematic representation of the αvβ3 integrin-ILK-Snail1 transduction pathway involved in the myofibroblast-like phenotype of hEDS and HSD cells.
Ijms 19 00982 g003
Table
Table 1. Cell expression, major ligands, and main functions of the αvβ3 integrin.
Table 1. Cell expression, major ligands, and main functions of the αvβ3 integrin.
Cell typesEndothelial cells, smooth muscle cells, myofibroblasts, osteoclasts, monocytes, platelets, fibroblasts, tumor cells (melanoma, glioblastoma, pancreatic, prostate, ovarian, breast tumor cells), placenta
LigandsVitronectin, fibrinogen, von Willebrand factor, thrombospondin, prothrombin, fibronectin, fibrillins, laminin, osteopontin, bone sialoprotein
FunctionsCell adhesion, cell migration, cell survival and proliferation, growth factor deprivation-induced apoptosis rescue, anoikis rescue, angiogenesis, hemostasis, platelet aggregation, wound healing, fibrosis, inflammation, tumor cells’ invasion and metastasis, restenosis, bone resorption, activation of latent TGF-β, embryonic development
Table
Table 2. Ehlers–Danlos syndromes (EDS) types according to the 2017 revised nosology.
Table 2. Ehlers–Danlos syndromes (EDS) types according to the 2017 revised nosology.
EDS TypeIPGeneProtein
Classical EDS (cEDS)ADMajor: COL5A1, COL5A2
Rare: COL1A1		Type V collagen
Type I collagen
Classical-like EDS (clEDS)ARTNXBTenascin-X
Cardiac-valvular EDS (cvEDS)ARCOL1A2Type I collagen
Vascular EDS (vEDS)ADCOL3A1Type III collagen
Hypermobile EDS (hEDS)ADUnknownUnknown
Arthrochalasia EDS (aEDS)ADCOL1A1, COL1A2Type I collagen
Dermatosparaxis EDS (dEDS)ARADAMTS2ADAMTS-2
Kyphoscoliotic EDS (kEDS)ARPLOD1
FKBP14		LH1
FKBP22
Brittle cornea syndrome (BCS)ARZNF469
PRDM5		ZNF469
PRDM5
Spondylodysplastic EDS (spEDS)ARB4GALT7
B3GALT6
SLC39A13		β4GalT7
β3GalT6
ZIP13
Musculocontractural EDS (mcEDS)ARCHST14
DSE		D4ST1
DSE
Myopathic EDS (mEDS)AD/ARCOL12A1Type XII collagen
Periodontal EDS (pEDS)ADC1R
C1S		C1r
C1s
IP: inheritance pattern; AD: autosomal dominant; AR: autosomal recessive.
Table
Table 3. Clinical criteria for classical Ehlers-Danlos syndrome (cEDS) according to the revised 2017 EDS nosology.
Table 3. Clinical criteria for classical Ehlers-Danlos syndrome (cEDS) according to the revised 2017 EDS nosology.
Major CriteriaMinor Criteria
1. Skin hyperextensibility and atrophic scarring
2. Generalized joint hypermobility (BS ≥ 5)		1. Easy bruising
2. Soft, doughy skin
3. Skin fragility (or traumatic splitting)
4. Molluscoid pseudotumors
5. Subcutaneous spheroids
6. Hernia (or history thereof)
7. Epicanthal folds
8. Complications of joint hypermobility (e.g., sprains, luxation/subluxation, pain, flexible flatfoot)
9. Family history of a first degree relative who meets clinical criteria
Minimal criteria suggestive for cEDS: Major criterion (1) Plus either: major criterion (2) and/or: at least three minor criteria.
Table
Table 4. Clinical criteria for vascular Ehlers-Danlos syndrome (vEDS) according to the revised 2017 EDS nosology.
Table 4. Clinical criteria for vascular Ehlers-Danlos syndrome (vEDS) according to the revised 2017 EDS nosology.
Major CriteriaMinor Criteria
1. Family history with documented COL3A1 variant
2. Arterial rupture at a young age
3. Spontaneous sigmoid colon perforation
4. Uterine rupture during the third trimester in the absence of previous C-section and/or severe peripartum perineum tears
5. Carotid-cavernous sinus fistula formation in the absence of trauma		1. Bruising unrelated to identified trauma
2. Thin, translucent skin with increased venous visibility
3. Characteristic facial appearance
4. Spontaneous pneumothorax
5. Acrogeria
6. Talipes equinovarus
7. Congenital hip dislocation
8. Hypermobility of small joints
9. Tendon and muscle rupture
10. Keratoconus
11. Gingival recession/fragility
12. Early-onset varicose veins
Minimal criteria suggestive for vEDS: Family history of the disorder, arterial rupture or dissection in individuals less than 40 years of age, unexplained sigmoid colon rupture, or spontaneous pneumothorax in the presence of other features consistent with vEDS.
Table
Table 5. Clinical criteria for hypermobile Ehlers-Danlos syndrome (hEDS) according to the revised 2017 EDS nosology.
Table 5. Clinical criteria for hypermobile Ehlers-Danlos syndrome (hEDS) according to the revised 2017 EDS nosology.
The Clinical Diagnosis of hEDS Needs the Simultaneous Presence of Criteria 1 and 2 and 3
Criterion 1Criterion 2Criterion 3
Two or more among the features A–C must be presentAll must be met
1. Generalized joint hypermobility:
BS ≥ 6 for pre-pubertal children and adolescents;
BS ≥ 5 for pubertal men and women up to the age of 50;
BS ≥ 4 for those >50 years of age		SIGN A (a total of five must be present):
1. Unusually soft or velvety skin
2. Mild skin hyperextensibility
3. Unexplained striae
4. Bilateral piezogenic papules of the heel
5. Recurrent or multiple abdominal hernia(s) (e.g., umbilical, inguinal, crural)
6. Atrophic scarring involving at least two sites and without the formation of truly papyraceous and/or hemosiderotic scars
7. Pelvic floor, rectal, and/or uterine prolapse in children, men or nulliparous women
8. Dental crowding and high or narrow palate
9. Arachnodactyly
10. Arm span-to-height ≥ 1.05
11. Mitral valve prolapse mild or greater based on strict echocardiographic criteria
12. Aortic root dilatation with Z-score  >  +2
SIGN B:
Positive family history, with one or more first degree relatives independently meeting the diagnostic criteria for hEDS
SIGN C (at least one):
1. Musculoskeletal pain in two or more limbs, recurring daily for at least 3 months
2. Chronic, widespread pain for ≥3 months
3. Recurrent joint dislocations or frank joint instability, in the absence of trauma 		1. Absence of unusual skin fragility, which should prompt consideration of other types of EDS
2. Exclusion of other HCTDs, including autoimmune and rheumatologic conditions
3. Exclusion of alternative diagnoses that include joint hypermobility by means of hypotonia and/or connective tissue laxity
Table
Table 6. Overview of the clinical features of arterial tortuosity syndrome (ATS).
Table 6. Overview of the clinical features of arterial tortuosity syndrome (ATS).
Craniofacial: Aged appearance; long face; hypertelorism; downslanting palpebral fissures; beaked nose; cleft palate/bifid uvula; high arched palate; micrognathia; sagging cheeks
Ocular: Keratoconus; keratoglobus; myopia
Cutaneous: Velvety texture; thin skin; hyperextensible skin; cutis laxa
Skeletal: Pectus deformity; scoliosis; arachnodactyly; joint hypermobility and pain
Cardiovascular: Aortic tortuosity; tortuosity of other arteries; abnormal implantation of the aortic branches; aortic root aneurysm; other arterial aneurysms; arterial dissections; stenosis of the pulmonary arteries; aortic stenosis
Other manifestations: Diaphragmatic hernia; inguinal hernia; respiratory symptoms; urogenital abnormalities; autonomic dysfunction
Table
Table 7. Organization of FN and COLLs and their canonical integrin receptors in control, EDS, and other HCTDs dermal fibroblasts.
Table 7. Organization of FN and COLLs and their canonical integrin receptors in control, EDS, and other HCTDs dermal fibroblasts.
ECM Components and Integrins #Control FibroblastscEDS
COL5A1
COL5A2		vEDS
COL3A1		hEDS
HSD
Unknown		kEDS
FKBP14		kEDS u
PLOD1		dEDS u
ADAMTS2		aEDS u
COL1A2
ex6		BCS
PRDM5
ZNF469		mcEDS
CSHT14		spEDS
B3GALT6		LDS u
TGFBR1		MFS u
FBN1		OI u
COL1A1
COL1A2		ATS
SLC2A10
FN++++++++++++++++++++
α5β1***/-*/-*/-----*/--*********,u
αvβ3-********,u********,u**,u-,u---**
COLLI++/---------+++--
COLLIII++-----------+--
COLLV++-++/+-++---++++++++/-+/-
α2β1**----------na*--,u
#: detected by immunofluorescence analyses, ++: abundant and fibrillar ECM, +: rare ECM fibrils, **: abundant patches in plasma membrane, *: reduced patches in plasma membrane, -: negligible amounts/absent, na: not analyzed, u: unpublished. The genes underlying the different HCTDs are reported in italics.

Share and Cite

MDPI and ACS Style

Zoppi, N.; Chiarelli, N.; Ritelli, M.; Colombi, M. Multifaced Roles of the αvβ3 Integrin in Ehlers–Danlos and Arterial Tortuosity Syndromes’ Dermal Fibroblasts. Int. J. Mol. Sci. 2018, 19, 982. https://doi.org/10.3390/ijms19040982

AMA Style

Zoppi N, Chiarelli N, Ritelli M, Colombi M. Multifaced Roles of the αvβ3 Integrin in Ehlers–Danlos and Arterial Tortuosity Syndromes’ Dermal Fibroblasts. International Journal of Molecular Sciences. 2018; 19(4):982. https://doi.org/10.3390/ijms19040982

Chicago/Turabian Style

Zoppi, Nicoletta, Nicola Chiarelli, Marco Ritelli, and Marina Colombi. 2018. "Multifaced Roles of the αvβ3 Integrin in Ehlers–Danlos and Arterial Tortuosity Syndromes’ Dermal Fibroblasts" International Journal of Molecular Sciences 19, no. 4: 982. https://doi.org/10.3390/ijms19040982

APA Style

Zoppi, N., Chiarelli, N., Ritelli, M., & Colombi, M. (2018). Multifaced Roles of the αvβ3 Integrin in Ehlers–Danlos and Arterial Tortuosity Syndromes’ Dermal Fibroblasts. International Journal of Molecular Sciences, 19(4), 982. https://doi.org/10.3390/ijms19040982

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