University of Groningen Major role of the extracellular ... · Chapter 1 8 The extracellular matrix Within the human body, tissue cells reside in a three-dimensional protein network,

University of Groningen

Major role of the extracellular matrix in airway smooth muscle phenotype plasticityDekkers, Bart Gerrit Jan

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Major role of the extracellular matrix in airway smooth muscle phenotype plasticity

Implications for chronic asthma

Proefschrift

ter verkrijging van het doctoraat in de Wiskunde en Natuurwetenschappenaan de Rijksuniversiteit Groningen

op gezag van de Rector Magnificus, dr. F. Zwarts, in het openbaar te verdedigen op

vrijdag 25 juni 2010 om 16.15 uur

door

Bart Gerrit Jan Dekkers

geboren op 28 januari 1981

te Hardenberg

Promotores: Prof. dr. H. Meurs Prof. dr. J. Zaagsma

Beoordelingscommissie: Prof. dr. A.J. Halayko

Prof. dr. K. Racké Prof. dr. W. Timens

ISBN: 978-90-367-4423-2

Paranimfen: I.S.T. Bos T.J.H. Dekkers

The research project described in this thesis was performed within the framework of the Groningen graduate school for behavioral and cognitive Neurosciences (BCN), and the Groningen research institute for asthma and COPD (GRIAC), and was financially supported by a grant from the Netherlands asthma foundation (grant 3.2.03.36). Printing of this thesis was financially supported by: Cover: ‘The extracellular matrix’ by Jesper Julyan Dekkers. Copyright © 2010 by Bart G.J. Dekkers. All rights reserved. No parts of this book may be reproduced in any manner or by any means without the permission from the publisher. Printing: Printpartners Ipskamp B.V., Enschede.

Table of contents

Chapter 1 General introduction 7

Chapter 2 Airway structural components drive airway smooth muscle remodeling in asthma Proc Am Thoracic Soc (2009) 8:683-692

29

Chapter 3 Extracellular matrix proteins differentially regulate airway smooth muscle phenotype and function Am J Physiol Lung Cell Mol Physiol (2007) 292: L1405-1413

53

Chapter 4 Functional consequences of human airway smooth muscle phenotype plasticity Submitted (2010)

73

Chapter 5 Insulin-induced laminin expression promotes a hypercontractile airway smooth muscle phenotype Am J Respir Cell Mol Biol (2009) 41:494-504

85

Chapter 6 The integrin-blocking peptide RGDS inhibits airway smooth muscle remodeling in a guinea pig model of allergic asthma Am J Respir Crit Care Med (2010) 181:556-565

109

Chapter 7 The laminin �1-competing peptide YIGSR induces a hypercontractile, hypoproliferative airway smooth muscle phenotype in chronic asthma

131

Chapter 8 Signalling pathways of collagen I-induced airway smooth muscle phenotype modulation

151

Chapter 9 Glucocorticosteroids and �2-adrenoceptor agonists synergistically prevent the induction of a proliferative, hypocontractile airway smooth muscle phenotype

171

Chapter 10 General discussion and summary 189

Nederlandse samenvatting 203

Dankwoord 213

Curriculum vitae 215

List of publications 216

List of abbreviations 219

Table of contents

1General Introduction

Chapter

Chapter 1

8

The extracellular matrix Within the human body, tissue cells reside in a three-dimensional protein network, the extracellular matrix (ECM), which is actively secreted, maintained and molded into the intercellular space by the cells residing in it. The ECM surrounds individual cells and plays a key role in determining physical and mechanical properties of tissues and organs, including the lung. The diverse components that comprise the ECM include collagens, elastin, glycoproteins (including laminins and fibronectin) and proteoglycans (like decorin and biglycan) [1]. In addition to its function as a scaffold, the ECM also influences fluid balance, lung compliance and elasticity and stores inflammatory mediators and growth factors, which may be rapidly released upon cleavage by matrix metalloproteinases (MMPs) and serine proteases [2-4]. Importantly, ECM proteins have also been found to influence cellular functions like migration, differentiation, proliferation and apoptosis. The extracellular matrix in the airways Extracellular matrix composition in healthy airways In the airways, the ECM is part of the cartilage, basement membranes, interstitium and provisional matrices. Cartilage surrounds the (central) airways and prevents airway collapse during breathing [5]. Basement membranes surround the structural cells and separate the cells from the adjacent interstitium [6]. Provisional matrices are formed after tissue injury and are replaced when normal tissue function is restored. The ECM is assembled from a large number of different macromolecules of which the fibrous proteins like collagens and elastin, glycoproteins and proteoglycans are the main components.

Collagens comprise a large family of fibrous ECM proteins and are formed from three polypeptide �-chains coiled in a helical structure [7]. The long, stiff chains contain characteristic repeating -glycine-X-Y-sequences in which the X-position is frequently occupied by proline and the Y-position by 4-hydroxyproline [7]. At least 28 collagen subtypes, divided into 8 subfamilies, are formed in humans (see [7] and [8] for detailed review). Collagens are widespread throughout the body and fulfil a variety of biological functions [7]. In the lung, collagens constitute approximately 15% of total lung dry weight [9]. Collagen IV is most prominently expressed in the basement membranes of the epithelium, vasculature and airway smooth muscle (ASM) [10]. Collagens I, III, V are only weakly expressed in the basement membrane, but are predominantly expressed in the interstitium and beneath the subepithelial basement membrane [10,11]. Collagen fibres are often assembled together with elastic fibres, which are formed from an elastin core surrounded by fibrillin-rich microfibrils, which conveys elastic recoil, whereas the fibrous collagens I and III provide tensile strength [12].

General Introduction

9

Laminins are large heterotrimeric glycoproteins comprised of �, � and � chains, which are important components of basement membranes [13]. Sixteen isoforms, assembled from five laminin �, four � and three � chains, have been identified [14]. Laminin �2, �3 and �5 chains have been observed in the epithelial basement membranes of healthy subjects. In addition, all � and � chains are present in the epithelial basement membrane [15-18]. Laminins �4, �5, �1, �2 and �1 have been found in the basement membrane of ASM cells [15,16,19].

The glycoprotein fibronectin can be found in the circulation in its soluble form and within the ECM in its insoluble form [20]. Expression of fibronectin in tissue is increased during cycles of injury and repair. Accordingly, only diffuse staining of fibronectin has been observed in the airways of healthy subjects [10,21]. Other glycoproteins, like vitronectin, tenascin, thrombospondin or SPARC (Secreted Protein Acid and Rich in Cysteine) are often increased in tissue undergoing repair as well. However, little is known about their expression in the airways of healthy subjects [1].

Proteoglycans are macromolecules consisting of a core protein with glycosaminoglycan (GAG) side chains, with the exception of hyaluronan which lacks a core protein. GAGs are divided in two classes, the first class being sulphated GAGs, like chondroitin sulphate, heparan sulphate and heparin, and the second class consisting of non-sulphated GAGs, like hyaluronan. The combination of the various core proteins and GAGs results in a large number of proteoglycans of which the small leucine rich proteoglycans (SLRPs) with only limited numbers of GAG side chains, like decorin, biglycan and lumican, and the modular proteoglycans with multiple GAG side chains, like versican, are best characterized [22]. Proteoglycans have been implicated in the assembly of collagen fibrils, the regulation of water balance and the storage of growth factors, chemokines and cytokines [1]. Decorin, biglycan and lumican have been observed in the subepithelial, ASM and adventitial compartments of the airways, with strong staining for decorin and biglycan in the adventitial compartment and within the ASM bundle [23,24]. Staining for versican is mainly observed beneath the epithelial basement membrane and within the airway ASM bundle [23].

Changes in the extracellular matrix in chronic asthma Asthma is an inflammatory airway disease characterized by airway inflammation, variable airway obstruction and airway hyperresponsiveness (AHR) [25]. AHR is defined by exaggerated airway narrowing in response to a variety of direct (e.g. histamine, methacholine) and indirect (adenosine monophosphate (AMP), fog, cold air, sulphur dioxide and exercise) stimuli [26]. Variable airway hyperresponsiveness occurs after episodic exposure to environmental factors, including allergens, and relates to airway inflammation, whereas persistent airway hyperresponsiveness is considered to reflect structural changes in the airway wall, collectively termed airway remodelling [27,28]. The most striking changes in the structure of the airway wall of asthmatics include epithelial

Chapter 1

10

shedding, increased ASM mass, goblet cell hyperplasia and metaplasia, increased microvasculature, subepithelial thickening and alterations in the ECM composition [29-31]. Mathematical modelling studies have indicated that increased ASM mass is likely to be the most important factor contributing to persistent airway hyperresponsiveness [32,33]. On the other hand, increased ECM deposition beneath the epithelium and within the ASM layer could be protective against airway constriction, as it may stiffen the tissue as a result of decreased elasticity and increased preload, whereas increased ECM deposition in the adventitia may lead to enhanced airway narrowing due to uncoupling of the tissue from the elastic recoil of the surrounding tissue [34].

Studies of endobronchial and post-mortem biopsies on the nature of the subepithelial alterations in the ECM in asthmatics have revealed increased deposition of collagen I, III and V, fibronectin, tenascin, hyaluronan, versican, biglycan, lumican and several laminin chains, including �2, �3, �5, �1, �2 and �1 chains [10,15,21,35-39]. Although weak, staining of laminin �1 chains has also been observed in the airways of allergic asthmatics, while no expression was observed in the airways of non-allergic asthmatics or healthy subjects [39]. In addition, both in allergic and non-allergic asthmatics with compromised epithelial integrity, increased laminin �2 deposition has been observed, which correlated closely with the level of epithelial integrity [39]. Expression of collagen IV, decorin and elastin, on the other hand, was decreased in the airway wall of asthmatics [1,40].

Post-mortem studies on airway tissue from individuals with fatal asthma have indicated that increased ECM deposition is not only present beneath the epithelium, but also inside and outside of the ASM bundle [41]. Increased ECM expression in the ASM bundle has been reported to involve deposition of collagen type I, fibronectin, hyaluronan, versican, biglycan, lumican and elastic fibres [24,42-44]. Increased collagen I and versican deposition was, however, not found in a subsequent study [44]. AHR may be linked to changed ECM deposition as it was shown that in asthmatics, airway responsiveness to methacholine was inversely correlated with elastin expression in the ASM bundle [45]. Mechanisms regulating extracellular matrix composition Tissue and ECM turnover are physiological processes, which are dynamically governed by a balance between matrix synthesis and matrix degradation [9]. In the airways, matrix turnover is estimated to amount more than 10% per day [46]. ECM turnover may be increased in asthmatics, as levels of cellular fibronectin, laminin degradation products and hyaluronan are increased in the bronchoalveolar lavage (BAL) fluid [47-49], of which the hyaluronan levels were correlated with asthma severity [48].


11

Epithelial cells and fibroblasts are considered to be the main source of ECM in the airways. However, ASM cells also produce a large variety of ECM proteins, including collagens, fibronectin, laminins, perlecan, elastin, thrombospondin, versican, decorin and hyaluronan [50-53]. Asthmatic and healthy ASM produce different ECM profiles, as indicated by increased production of collagen I, perlecan and fibronectin, and decreased production of laminin �1, chondroitin sulphate, collagen IV and hyaluronan by asthmatic ASM [50,53,54]. Production of ECM proteins is increased by profibrotic factors like transforming growth factor-� (TGF-�), granulocyte macrophage colony-stimulating factor (GM-CSF), connective tissue growth factor (CTGF) and vascular endothelial growth factor (VEGF) [55]. Moreover, asthmatic ASM cells produce more CTGF in response to TGF-� [56], suggesting that this factor may be involved in changed ECM production by these cells. In addition, exposure of healthy ASM cells to atopic serum increases the production of fibronectin, laminin �1, perlecan and chondroitin sulphate [57].

Degradation of ECM is governed by a variety of proteases and binding proteins, of which the matrix metalloproteinases (MMPs), their inhibitors – the tissue inhibitors of MMPs (TIMPs) – and A disintegrin and metalloproteinases (ADAMs) are best recognized [58,59]. Several studies have shown that in asthma, expression of MMP-2, MMP-3 and MMP-9 is increased, of which MMP-9 is increased most predominantly (reviewed in [59]). In addition, polymorphisms in the ADAM33 gene correlate with asthma incidence, bronchial hyperresponsiveness and lung function decline, although the precise mechanism remains to be determined [60,61]. Extracellular matrix proteins, integrins and airway smooth muscle function Extracellular matrix proteins affect airway smooth muscle function Increased ASM mass is, next to increased and changed ECM deposition, another hallmark of airway remodelling in asthmatics [62-64]. Increased ASM mass may involve increased cell numbers (hyperplasia), cell size (hypertrophy) or a combination of both [62-64]. ASM hyperplasia may, at least partly, be explained by enhanced proliferative responses and in culture ASM cells derived from asthmatics proliferate faster than those obtained from healthy subjects [65,66]. In addition to increased proliferative capabilities, contractile function and synthetic capabilities of ASM cells derived from asthmatics have also been shown to be increased [54,56,67]. ASM may exert these functions as they retain the ability of reversible phenotypic plasticity, enabling them to switch between proliferative, synthetic, contractile and migratory phenotypes [68,69]. In vitro, exposure of ASM cells to mitogens results in the switch from a contractile to a proliferative phenotype, associated with a decreased contractile ability [70] due to decreased expression of contractile proteins [68]. Conversely, removal of

Chapter 1

12

growth factors, in the presence of insulin or TGF-�, results in the reintroduction of a (hyper)contractile phenotype [71,72]. For a detailed review of the literature on ASM phenotype plasticity, the reader is referred to references [72] and [73].

Various studies have addressed the effects of ECM proteins on ASM function, including ASM proliferation, contractile protein expression, maturation, synthetic function, survival and migration (Figure 1). Of the collagens, collagen type I has been most extensively studied. Monomeric collagen I has been shown to enhance basal and growth factor-induced ASM proliferation, ASM survival and synthesis of pro-inflammatory mediators like eotaxin, RANTES (Regulated upon Activation, Normal T-cell Expressed, and Secreted, CCL5) and GM-CSF [54,74-77]. Collagen I concentration-dependently increases ASM migration [78]. The reduction in contractile protein expression induced by platelet-derived growth factor (PDGF) is further reduced by monomeric collagen type I [75]. Surprisingly, although the fibrillar form of collagen I increased cytokine synthesis to a similar extent as did monomeric collagen I, no effects of fibrillar collagen I were observed on ASM cell proliferation [54,77], whereas recently fibrillar collagen I has even been shown to inhibit both basal and growth factor-induced proliferation [79]. In addition, inhibition of collagen degradation by the MMP inhibitor ilomastat further enhanced the growth-attenuating effects of fibrillar collagen I, indicating that degradation of collagen to its monomeric isoform may enhance ASM proliferation [79]. Little is known on the effects of other collagens on ASM function. The studies available indicate that culturing ASM cells on collagen type III does not change PDGF-induced ASM cell proliferation [77], but increases PDGF-induced migration [78]. Collagen type V increased migration, but was without effects on ASM cell survival [76,78]. Finally, collagen IV has been shown to inhibit ASM cell apoptosis [76].

Many effects observed for fibronectin are comparable to those observed for monomeric collagen I. Growth factor-induced proliferation, cytokine synthesis and survival are increased in cells cultured on fibronectin matrices and the effects of PDGF on contractile protein expression are enhanced [54,74-77]. In addition, fibronectin increased migration of ASM cells towards PDGF [78]. Vitronectin also increased growth factor-induced proliferation, although to a lesser extent than observed for fibronectin [77]. No effects of vitronectin were observed on ASM survival [76].


13

ECM

Migration

Cytokine synthesis

Eotaxin, RANTES

Survival

Coll I, III, V, FN

LN-111, LN-211

Coll I, FN

Coll I, FN, VN

LN-1, FN, Coll I, IV

Decorin

Maturation Contractile protein expression

Coll I, FN

Proliferation

LN-111, Heparan, Chondroitin

Figure 1 Extracellular matrix proteins affect airway smooth muscle phenotype and function. ECM proteins differentially regulate ASM contractile protein expression and maturation, proliferation, cytokine synthesis, survival and migration. Changes in the ECM environment surrounding the cell may contribute to ASM abnormalities as observed in asthma. Abbreviations used: Coll, collagen; FN, fibronectin; LN, laminin; VN, vitronectin. �, increased �, decreased. Laminin-111 (laminin-1) or matrigel, a basement membrane extract containing multiple ECM components including laminins [80], reduced growth factor-induced proliferation and prevented growth factor-induced reductions in contractile protein expression [75]. Laminin-111 also increased ASM survival [76]. Moreover, maturation of ASM cells to a contractile phenotype – in the presence of insulin – is accompanied by an increased expression of laminin �2, �1 and �1 chains, found in laminin-211 (laminin-2). Increased expression of these laminin chains was required for the maturation as contractile protein accumulation by serum deprivation is normalized by the laminin competing peptide Tyr-Ile-Gly-Ser-Arg (YIGSR) and the Arg-Gly-Asp (RGD)-containing peptide Gly-Arg-Gly-Asp-Ser-Pro (GRGDSP) [81].

Proteoglycans differentially affect ASM function. Culturing ASM cells on decorin decreased PDGF-induced proliferation and increased apoptotic rates, while biglycan did not have an effect on both parameters [82]. Decorin may also play an important role in ASM maturation by binding of and inhibiting the function

Chapter 1

14

of TGF-�1 [83,84]. Heparin, heparan and chondroitin, other members of the proteoglycan family, also inhibited serum-induced ASM proliferation [85].

Changes in the ECM profile produced by asthmatic ASM may contribute significantly to the changed function of asthmatic ASM cells [50,54]. Both enhanced proliferation and increased eotaxin synthesis by asthmatic ASM cell is dependent on the ECM: healthy ASM cells cultured on an ECM laid down by asthmatic ASM cells, showed increased proliferative rates and increased synthetic capacities and vice versa [50,54], suggesting that in asthma the ECM may be a critical regulator of increased ASM mass and persistent inflammation in the remodelled airway. Integrins and airway smooth muscle function Cells interact with their surrounding matrix mainly through integrins, a group of heterodimeric transmembrane glycoproteins, which interact with specific sequences within the ECM proteins. Eighteen � integrin subunits and eight � integrin subunits forming twenty-four heterodimers have been identified thus far [86]. In culture, ASM cells have been shown to express the �1, �2, �3A, �4, �5, �6A, �6B, �7B, �v, �v�3 and �1 subunits, while the �3B, �7A, �2 and �4 subunits are relatively rare (Table 1)[76,77,87,88].

The majority of the studies on the role of integrins in ECM-induced changes in ASM function have focused on collagens, fibronectin and laminins. Increased synthetic responses of ASM cells in response to IL-1� or IL-13 on collagen I matrices required interaction with the collagen binding integrin �2�1 [54,87]. Similarly, enhancement of growth factor-induced proliferation by monomeric collagen I was inhibited by �2�1 function-blocking antibodies. In addition, blocking of the fibronectin binding integrins �4�1 and �5�1 also prevented the enhancement of growth factor-induced proliferation by monomeric collagen I, suggesting that fibronectin may also be involved in the enhanced proliferation [77]. Attachment of ASM cells to collagen I required the �2�1 integrin and the fibronectin binding integrin �v�3, however [77].

Similar to collagen I, enhancement of growth factor-induced proliferation by fibronectin required interaction with the �2�1, �4�1 and �5�1 integrins [77]. The collagen binding integrin �2�1 and the fibronectin binding integrins �5�1, �v�1 and �v�3 are important in the increased eotaxin production in response to IL-1� by ASM cells cultured on fibronectin [87]. Moreover, peptides containing the RGD peptide sequence, present in the integrin recognition site of several ECM proteins [92,93], also inhibited the enhancement of IL-1�-induced eotaxin release [87]. Enhancement of IL-13-induced eotaxin release, on the other hand, only required interaction with the �5�1 integrin [54]. Attachment of ASM cells to fibronectin is also mediated by �5�1 integrins [77].


15

Table 1: Airway smooth muscle: integrin expression and function. Integrin Expression

level Known functions in ASM

�1 ~30-50% -- �2 ~60-80% Enhanced growth factor-induced proliferation on collagen I

and fibronectin matrices [77], enhanced cytokine release on collagen I and fibronectin matrices [87], attachment to collagen I [77], resistance to glucocorticoid action on collagen I matrices [89].

�3A ~50% Increased expression by PDGF stimulation [77], increased expression during maturation [88], inhibition of �5�1 expression [88].

�3B fraction -- �4 ~20-30% Enhanced growth factor-induced proliferation on collagen I

and fibronectin matrices [77], serum-induced proliferation of nonasthmatic ASM [90].

�5 ~100% Enhanced growth factor-induced proliferation on collagen I and fibronectin [77], serum-induced proliferation of nonasthmatic and asthmatic ASM [91], attachment to fibronectin [77], ASM survival [76], cytokine release on fibronectin and increased cytokine release by asthmatic ASM [54,87], regulation of fibronectin expression [91].

�6A ~40% Increased expression during maturation [88], negative regulation of �5�1 expression [88]

�6B ~30% -- �7A fraction -- �7B ~20% Increased expression during maturation [88], required for

maturation [88] �v ~50-100% Enhanced cytokine release on fibronectin [87], serum-

induced proliferation [90] �v�3 ~50% Attachment to collagen I [77], enhanced cytokine release on

fibronectin matrices [87] During maturation, ASM cells not only increase the expression of contractile marker proteins and laminin �2, �1 and �1 chains [81], but also the expression of the laminin binding integrin subunits �3A, �6A and �7B [88]. Increased expression of the �7 subunits was shown to be restricted to cells acquiring a contractile phenotype. Knockdown of the �7 integrin, but not of the �3 or �6 integrins, fully prevented phenotype maturation, indicating an essential role for this integrin in ASM maturation. Interestingly, knockout of the laminin binding �3 or �6 integrins increased expression of the fibronectin �5 integrin [88], suggesting an inverse relationship between laminin-binding and fibronectin-binding integrins.

Chapter 1

16

Integrins of the �5�1 subtype have been implicated in a number ASM functions. Survival of ASM cells on several matrices is significantly reduced in the presence of �5�1 integrin blocking-antibodies [76]. Moreover, survival of ASM cells was also attenuated in the presence of RGD containing peptides [76]. In addition, �5�1 integrins are important regulators of fibronectin expression and of serum-induced proliferation of both asthmatic and non-asthmatic ASM cells [91]. Preliminary results also indicate that serum-induced proliferation of non-asthmatic ASM cells requires �v�1 and �4�1 integrins, whereas asthmatic ASM cells are unresponsive to inhibition of �4�1 integrins [90]. Moreover, increased eotaxin release by asthmatic ASM is significantly inhibited by antibodies blocking the �5�1 integrin [54]. Integrin-induced signalling Integrins not only provide a physical link between the ECM and intracellular compartment, but may also trigger a large number of intracellular signalling cascades to influence cellular processes, including proliferation, differentiation, migration and apoptosis. Many of these signalling pathways are also activated by receptor tyrosine kinases (RTKs) and G protein-coupled receptors (GPCRs) [94-96].

The biologically active sites in the ECM interacting with the integrins, like for example the RGD binding site in fibronectin, are usually not exposed in mature ECM proteins, but may become exposed after structural or conformational changes (for a detailed review on matricryptic sites see [97]). Many integrins are not constitutively active. However, upon activation, integrins mediate signal transduction through the cell membrane in both directions: binding of ECM proteins to the extracellular domain of integrins elicits signalling cascades in the cell, via the cytoplasmic domains of the integrin (outside-in signalling), whereas the binding between the integrins and the ECM can be activated or enhanced from the inside of the cell (inside-out signalling) [86]. Intracellularly, integrins activate various protein tyrosine kinases, including integrin linked kinase (ILK) and focal adhesion kinase (FAK) [98]. ILK may be activated by beta integrin subunits and has been shown to be important in the expression of ASM contractile proteins. Knock-down of ILK increased gene expression of a number of contractile proteins, including smooth muscle specific myosin heavy chain (sm-MHC) SM22� and calponin, but only increased protein expression of sm-MHC. This increase was associated with a decreased phosphorylation of protein kinase B (PKB/Akt) and increased binding of serum response factor (SRF) to the promoter for the sm-MHC gene. Conversely, overexpression of ILK had the opposite effect on these processes [99].

Activation of FAK is initiated by autophosphorylation of tyrosine 397 (Y397) [94]. Autophosphorylation requires clustering of the integrins, which may occur after binding to the ECM. The integrin clustering triggers a conformational


17

change in the associated FAK that alters the interaction of the FERM domain with the kinase domain [100]. In addition to activation of FAK by integrins, FAK may also be phosphorylated by growth factors receptors and G protein-coupled receptors. Growth factor receptors activate FAK via interaction with the FERM domain [101], whereas G protein-coupled receptors activate FAK via a mechanism which is currently unclear, but appears to involve Rho-dependent signalling pathways [96]. Phosphorylation of Y397 generates a high-affinity binding site for Src (Rous sarcoma oncogene cellular homolog) and results in the recruitment and binding of cellular Src to pY397 [102]. Cellular Src subsequently phosphorylates Y576/Y577 in the kinase domain of FAK, which is essential for maximal FAK kinase activity and activation [103]. Studies on the role of FAK in tracheal smooth muscle have indicated that phosphorylation and activation of FAK can be increased by mechanical strain [104] and by acetylcholine (ACh), in a Ca2+-independent fashion [105]. Stimulation with ACh also increased the membrane localization of FAK and of cytoskeletal linker proteins like paxillin, vinculin, talin and �-actinin [106]. Depletion of FAK from the tracheal tissue decreased KCl- and ACh-induced contraction in tension, myosin light chain phosphorylation and increase in intracellular Ca2+, indicating that FAK plays an important role in ASM contraction [107].

In addition to its role in ASM contraction, activation of FAK is also important in proliferation of various cell types [103]. Upon autophosphorylation and activation, FAK activates multiple signalling cascades, including the phosphatidylinositol 3-kinase (PI3-kinase) and extracellular signal-regulated kinase (ERK) signalling pathways [94]. Although the contribution of these signalling cascades to integrin-mediated changes in ASM function are currently unknown, activation of the PI3-kinase and the ERK signalling pathways has been shown to be critical in the response of ASM cells to peptide growth factors [95]. Thus, activation of PI3-kinases is important in growth factor-induced ASM cell proliferation and hypertrophy [108-110]. Activation of the PI3-kinase is associated with transcriptional activation and protein synthesis leading to ASM cell proliferation and hypertrophy [108,110]. ERK1/2 (or p42/p44 mitogen activated protein kinases (MAPKs)) are involved in the transfer of growth promoting signals to the nucleus and the subsequent induction of ASM proliferation [111]. In addition to p42/p44 MAPKs, p38 MAPK, another member of the MAPK family, is also involved in the regulation of growth factor-induced proliferation of ASM cells [112]. Collectively, these observations suggest that activation of PI3-kinase and MAPK pathways by FAK may contribute to ECM-induced changes in ASM function (Figure 2).

Chapter 1

18

Figure 2: Proposed mechanism by which integrins may activate intracellular signalling cascades and regulate ASM phenotype. Clustering of integrins (ITG) in response to ECM proteins triggers a conformational change in the focal adhesion kinase (FAK), resulting in an autophosphorylation at Y397. In addition, FAK may be activated by receptor tyrosine kinases (RTKs) and G protein-coupled receptors (GPCR) as well. Phosphorylation of FAK on Y397 creates a binding site for Src, which in turn phosphorylates FAK on Y576/Y577, leading to the full activation of the kinase, which may then activate PI3-kinase (PI3K) and mitogen activated protein kinase (MAPK) signalling pathways to regulate ASM phenotype and function. In addition, integrins may activate integrin-linked kinase (ILK) and protein kinase B (PKB/Akt), which inhibit ASM contractile protein expression. See text for detailed description. Effects of extracellular matrix proteins on airway smooth muscle response to respiratory therapeutics ECM proteins not only directly affect ASM function, but also affect the response of ASM towards asthma therapeutics. Studies by Freyer et al, have indicated that changes in the ECM environment may affect the response of ASM cells to �2-adrenoceptor agonists, used for the relief of acute bronchospasm [113,114]. Thus, it was shown that production of the second messenger cAMP in response to �2-adrenoceptor stimulation was increased in cells cultured on fibronectin,

GPCR

FAK

��

ITG

Regulation of ASM phenotype and function

MAPK PI3K

RTK

ECM

Src

P ILK

Akt


19

whereas collagen V and laminin-111 decreased cAMP accumulation. No effects of collagen I or collagen IV on cAMP accumulation were observed [113]. These changes were due to differences in G�i activity but not G�i protein expression, which causes inhibition of adenylyl cyclase activity [113].

Effects of glucocorticosteroids, used for the control of (chronic) asthma symptoms [115], have been investigated on ECM production and proliferation of ASM cells. Production of ECM proteins by ASM cells appears to be insensitive to glucocorticoids and treatment may even increase ECM production [116]. The increased ECM deposition by ASM cells in response to healthy or atopic serum is not affected by treatment with beclomethasone, although treatment did inhibit the increases in ASM cell number [57]. Treatment with beclomethasone even increased production of fibronectin, perlecan and chondroitin sulphate and this increase was larger in cells exposed to atopic serum [57]. In ASM cells and intact bronchial rings, treatment with �2-agonists or glucocorticosteroids did not inhibit TGF-�-induced increases in collagen I, fibronectin or CTGF, whereas the PDE4 inhibitor roflumilast did. Moreover, in the absence of TGF-� glucocorticosteroids increased expression of these proteins [117].

Glucocorticoids have been shown to inhibit human ASM cell proliferation [118-120]. Inhibition of human ASM cell proliferation required downregulation of cyclin D1 and reduced phosphorylation of retinoblastoma protein (pRb)[120]. In human ASM cells cultured on collagen I, however, inhibition of basic fibroblast growth factor (bFGF)-induced proliferation by glucocorticoids is largely abrogated [89,121]. This resistance to glucocorticoid action was no longer observed in the presence of �2�1 function blocking antibodies [89], indicating that this integrin is importantly involved in this process. By contrast, production of the cytokine GM-CSF by human ASM was inhibited by the glucocorticoid dexamethasone independent of the ECM environment, suggesting that proliferation and synthetic functions are differentially regulated by glucocorticoids [121]. On the other hand, no effects of collagen I were observed on glucocorticoid sensitivity in bovine ASM cell proliferation [74]. The �2-agonist salbutamol inhibited bFGF-induced human ASM cell proliferation in cells cultured on both laminin and collagen I, whereas GM-CSF release was not inhibited by salbutamol in cells cultured on collagen I [122]. Collectively, these findings suggest that changes in the ECM deposition in the airway wall, especially in the ASM surroundings, may change the effectiveness of the medication in asthma. Aims of the studies The above mentioned observations suggest that ECM proteins and their integrins may be important mediators of altered ASM function as observed in asthma. The primary aim of this thesis was to investigate the regulation of ASM phenotype and function by ECM proteins and to investigate the potential contribution of these effects to ASM remodelling in asthma.

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Chapter 2 provides a comprehensive review of the abnormalities observed in asthmatic ASM function. In addition, the mechanisms by which structural components of the airway wall may contribute to abnormal ASM behaviour in asthma are discussed. Chapters 3 and 4 address the effects of different ECM proteins on ASM phenotype and function. Chapter 3 describes the effects of prolonged treatment with the ECM proteins collagen type I, fibronectin and laminin-111 on contractility of intact bovine tracheal smooth muscle (BTSM) tissue. In addition, these effects were related to contractile protein expression and cell proliferation, in order to assess phenotype modulation under these conditions. Moreover, the effect of these ECM proteins on PDGF-induced phenotype modulation was investigated as well. In chapter 4, some of the findings observed in BTSM were translated to human tracheal smooth muscle (HTSM). To this aim, the effects of prolonged treatment of intact HTSM strips with collagen I, in the absence and presence of PDGF, were investigated on contractility, contractile protein expression and cell proliferation. Chapter 5 describes the role of laminins in the induction of a hypercontractile ASM phenotype. Previous studies have indicated that long-term exposure of BTSM to insulin induces a functionally hypercontractile, hypoproliferative phenotype [71,123]. The contribution of laminins to the induction of such a hypercontractile, hypoproliferative phenotype was assessed, using the laminin competing peptides Tyr-Ile-Gly-Ser-Arg (YIGSR) and Arg-Gly-Asp-Ser (RGDS). In addition, the effects of insulin on laminin mRNA and protein expression were investigated, as well as the potential signalling mechanisms involved (PI3-kinase and Rho kinase). The potential contributions of ECM proteins and their integrins to airway wall remodelling in vivo were assessed using a guinea pig model of chronic allergic asthma [124]. Chapter 6 describes the effects of the integrin blocking peptide RGDS on parameters of airway remodelling induced by repeated allergen-challenge. Thus, the contribution of RGD-binding integrins to allergen-induced ASM hyperplasia, increased contractile protein expression and hypercontractility as well as inflammation and fibrosis were assessed. Furthermore, to investigate potential underlying mechanisms the effects of RGDS on ECM- and growth factor-induced proliferation and maturation of human ASM cells were assessed in vitro. The effects of the specific laminin competing peptide YIGSR on airway remodelling in vivo are described in Chapter 7. The involvement of pro-mitogenic signalling pathways in collagen I-induced changes of ASM phenotype was studied in Chapter 8. The role of FAK in ASM cell adhesion and in collagen I-induced cell proliferation was assessed by overexpression of the kinase as well as by overexpression of two FAK inhibitors.


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The contribution of the downstream signalling pathways Src, PI3 kinase, p42/p44 MAPK and p38 MAPK to the induction of a proliferative, hypocontractile ASM phenotype by collagen I was investigated using specific inhibitors. Finally, in Chapter 9 the functional impact of glucocorticoids and �2-adrenoceptor agonists on ASM phenotype switching was studied. The effects of the glucocorticosteroids fluticasone, budesonide and dexamethasone and the �2-adrenoceptor agonist fenoterol on the induction of a proliferative, hypocontractile BTSM phenotype by PDGF or collagen I were assessed. As previous studies have indicated that glucocorticosteroids and �2-adrenoceptor agonists synergize to inhibit ASM cell proliferation [119], the functional impact of this synergism on the induction of a proliferative, hypocontractile phenotype was assessed as well. References 1. Fernandes DJ, Bonacci JV, Stewart AG. Extracellular matrix, integrins, and

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119. Roth M, Johnson PR, Rudiger JJ, King GG, Ge Q, Burgess JK, Anderson G, Tamm M, Black JL. Interaction between glucocorticoids and beta2 agonists on bronchial airway smooth muscle cells through synchronised cellular signalling. Lancet 2002; 360: 1293-1299.

120. Fernandes D, Guida E, Koutsoubos V, Harris T, Vadiveloo P, Wilson JW, Stewart AG. Glucocorticoids inhibit proliferation, cyclin D1 expression, and retinoblastoma protein phosphorylation, but not activity of the extracellular-regulated kinases in human cultured airway smooth muscle. Am J Respir Cell Mol Biol 1999; 21: 77-88.

121. Bonacci JV, Harris T, Wilson JW, Stewart AG. Collagen-induced resistance to glucocorticoid anti-mitogenic actions: a potential explanation of smooth muscle hyperplasia in the asthmatic remodelled airway. Br J Pharmacol 2003; 138: 1203-1206.

122. Bonacci JV, Stewart AG. Regulation of human airway mesenchymal cell proliferation by glucocorticoids and beta2-adrenoceptor agonists. Pulm Pharmacol Ther 2006; 19: 32-38.

123. Gosens R, Nelemans SA, Hiemstra M, Grootte Bromhaar MM, Meurs H, Zaagsma J. Insulin induces a hypercontractile airway smooth muscle phenotype. Eur J Pharmacol 2003; 481: 125-131.

124. Meurs H, Santing RE, Remie R, van der Mark TW, Westerhof FJ, Zuidhof AB, Bos IS, Zaagsma J. A guinea pig model of acute and chronic asthma using permanently instrumented and unrestrained animals. Nat Protoc 2006; 1: 840-847.

Airway structural

components drive

Bart G.J. Dekkers Harm Maarsingh Herman Meurs Reinoud Gosens

Proc Am Thoracic Soc (2009) 8:683-692

2Airway structural components drive airway smooth muscle

remodeling in asthma

Chapter

Chapter 2

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Abstract Chronic asthma is an inflammatory airways disease characterized by pathological changes in the airway smooth muscle bundle that contribute to airway obstruction and hyperresponsiveness. Remodeling of the airway smooth muscle is associated with an increased smooth muscle mass, involving components of cellular hypertrophy and hyperplasia, and changes in the phenotype of the muscle that facilitate proliferative, synthetic and contractile functions. These changes are considered major contributing factors to the pathophysiology of asthma, because of their role in exaggerated airway narrowing. The mechanisms that regulate changes in airway smooth muscle mass and phenotype are incompletely understood, but likely involve the regulatory role of mediators and growth factors secreted from inflammatory cells on airway smooth muscle cell proliferation and phenotype. An alternative hypothesis is that cellular and structural components that together constitute the airway wall, such as the airway epithelium, airway nerves, and the extracellular matrix, interact with the airway smooth muscle bundle to facilitate changes in smooth muscle phenotype and function that drive remodeling under inflammatory conditions. This review will discuss the mechanisms by which structural components of the airway wall communicate with the airway smooth muscle bundle to regulate remodeling and discuss these mechanisms in the context of the pathophysiology of asthma.

Introduction

Asthma is a chronic disease of the airways, which is characterized by persistent airway inflammation, reversible airways obstruction, airway remodeling and airway hyperresponsiveness (1). Airway hyperresponsiveness is defined by an exaggerated narrowing of the airways to a variety of chemical, physical and pharmacological stimuli (2). Acute, variable airway hyperresponsiveness has been considered to reflect increased airway smooth muscle contraction associated with airway inflammation and is related to asthma activity and severity, whereas chronic persistent airway hyperresponsiveness may reflect airway remodeling (3). Airway remodeling is characterized by changes in the structure of the airway wall, which include shedding of the epithelium, goblet cell hyperplasia, increased blood vessel number and area, increased and changed deposition of extracellular matrix (ECM) and increased airway smooth muscle (ASM) mass (4-6). Airway remodeling, notably the abnormalities in the ASM that encircles the airways and regulates lumen diameter, may contribute to the pathogenesis and pathophysiology of asthma.

Several studies have indicated that differences in contractile responses exist between ASM cells derived from asthmatics and cells derived from healthy subjects, which could at least in part explain hyperresponsiveness in asthma.

Strcutural components drive ASM remodeling

31

Thus, asthmatic ASM cells contract with greater velocity and maximum shortening capacity compared to healthy ASM (7). These changes may be explained by increases in the expression of smooth muscle myosin light chain kinase (sm-MLCK), transgelin (SM22) and myosin heavy chain (sm-MHC) as reported in asthmatic ASM cells and in asthmatic biopsies (7;8). Moreover, asthmatic ASM appears to express increased levels of the 7 amino acid insert SM-B isoform of sm-MHC, which shows a two-fold greater ATPase activity and shortening velocity compared to the SM-A isoform (8;9), although this latter finding is at odds with the study by Ma et al. who reported no expression of the SM-B isoform in smooth muscle obtained from asthmatic subjects (7). The increase in contractile protein expression may be of clinical relevance as expression levels of sm-MHC, sm-�-actin and desmin correlate with methacholine responsiveness in asthmatics (8;10). Studies using asthmatic and nonasthmatic ASM cultured in collagen gels also showed that maximal histamine-induced condensation of the gel was increased when cells derived from asthmatics were used (11). In addition, ASM relaxation may also be changed as relaxation of passively sensitized ASM is slower compared to controls (12).

Increased ASM mass is one of the most striking features of airway remodeling in asthma. Mathematical modeling studies on the impact of remodeling on airway narrowing indicated that increased ASM mass is likely to be the most important feature in increased airway narrowing in asthma, when assuming that the capacity of the ASM bundle to produce force is proportional to its mass (13;14). This idea is underscored by the fact that asthmatic patients in which the ASM layer has been reduced by bronchial-thermoplasty, show improved asthma control (15).

Several studies have addressed the underlying pathology causing the increased ASM mass in asthma. Ebina et al. (16) examined the ASM layer surrounding the airway lumen in fatal asthma and found two different asthmatic phenotypes, one showing an increased number of ASM cells (hyperplasia), the other showing an increased ASM cell size (hypertrophy). In subsequent studies, Woodruff et al. (17) found evidence for hyperplasia, but not hypertrophy in the ASM layer of mild to moderate asthmatics, whereas Benayoun et al. (18) found ASM hypertrophy, but not hyperplasia in patients with intermittent, mild-to-moderate and severe asthma. The latter group also found a clear correlation between disease severity and the degree of ASM thickening, consistent with a recent study showing that ASM thickening is more significant in fatal asthma as compared to non-fatal asthma (19). The relationship between age and duration of disease and ASM thickening is still subject of debate (18-21). Collectively, these findings suggest that increased ASM mass in asthma may reflect both cellular hyperplasia and hypertrophy, the degree of which depends primarily on asthma severity.

The increased ASM mass may be explained by intrinsic changes in the asthmatic ASM cell that facilitate their proliferative and secretory characteristics.

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Asthmatic ASM produces more pro-inflammatory, pro-angiogenic and pro-remodeling factors including eotaxin, vascular endothelial growth factor (VEGF) and connective tissue growth factor (CTGF) (22-24), and less anti-mitogenic factors like PGE2 (25). Accordingly, asthmatic ASM cells in culture proliferate faster compared to healthy controls (26), which is caused by changes in ECM protein deposition (27), and by enhanced mitochondrial biogenesis and mitochondrial activity that support increased cell growth (28). The exact mechanisms that regulate these responses are still incompletely identified; nonetheless these studies do highlight the importance of the ASM cell as an interactive player in the remodeling process rather than being the passive contractile partner as traditionally proposed. Remodeling of the airway smooth muscle bundle: mechanisms Although the mechanisms that regulate airway wall remodeling have thus far been incompletely identified, there is likely a major role for airway inflammation. Airway inflammation precedes airway remodeling in animal models of asthma (29;30), and ASM is known to proliferate in response to numerous growth factors and mediators that are released during allergic airway inflammation both in vitro and in vivo (31). Nonetheless, both clinical and animal studies indicate that the relationship between inflammation and remodeling is complex, and still incompletely understood. The presence of airway inflammation in patients with asthma is no guarantee at all for the occurrence of airway remodeling, and there is no clear correlation between the degree of inflammation and the degree of remodeling (18). Also, components of remodeling, including smooth muscle thickening appear to be present already in young children (32-34) and there is no clear relationship between age or duration of disease and the extent of ASM thickening (19). Furthermore, although airway inflammation can be resolved upon allergen avoidance in a murine model, remodeling persists, suggesting that ongoing inflammation is not required to support the maintenance of the remodeled airway wall (35). Collectively, although these studies point to an important, probably indispensible, role for airway inflammation in initiating or regulating the remodeling response, these studies also indicate that additional mechanisms exist in the airway wall that are necessary to direct or maintain the remodeling response. Moreover, these studies suggest that targeting inflammation per se may not be sufficient to reverse existing airway smooth muscle remodeling, a contention supported by studies showing that corticosteroid treatment prevents but does not reverse remodeling in allergen challenged rats and mice (36;37).

In the next sections, we will discuss recent findings that underscore the hypothesis that communication between different structural cells and compartments of the airway wall is central to the development of remodeling and may provide useful alternative drug targets for the treatment of smooth muscle


33

remodeling. These mechanisms include communication between the ASM, the airway epithelium, the airway parasympathetic nervous system and the ECM. The airway epithelium The airway epithelium forms the interface between the external environment and the airways (38). In asthma, the epithelial barrier is disrupted which contributes to AHR and inflammation associated with this disease via increased release of pro-inflammatory cytokines. In addition, (damaged) epithelial cells in asthma release a number of growth factors, including epidermal growth factor (EGF), platelet-derived growth factor (PDGF), transforming growth factor-� (TGF-�) and VEGF, as well as acetylcholine (see below), which may contribute to airway remodeling in asthma by inducing ASM growth, ECM deposition and angiogenesis (38-41) (Figure 1). In this section, we will focus on the possible contribution of other epithelial processes, particularly alterations in L-arginine homeostasis, to airway remodeling in asthma.

Thus far, the role of L-arginine homeostasis in airway (patho)physiology has mostly been studied in the context of regulating airway (hyper)responsiveness. The epithelium is an important source of the bronchodilator nitric oxide (NO), which is produced by NO synthase (NOS) from the hydrolysis of L-arginine (42). Three NOS isozymes have been identified: neuronal (nNOS), endothelial (eNOS) and inducible NOS (iNOS). In the airway epithelium, nNOS and eNOS are constitutively expressed, whereas iNOS is particularly induced by proinflammatory cytokines during the late asthmatic reaction (43;44). The NO production is regulated by the substrate availability to NOS and alterations in the L-arginine homeostasis contribute to the pathophysiology of (acute) allergic asthma (45). Although levels of exhaled NO are elevated in asthmatics due the induction of iNOS (43;46;47), it has paradoxically been shown that a deficiency in bronchodilating (epithelium-derived) NO underlies the development of airway hyperresponsiveness in animal models of allergic asthma (48-54) and in asthmatic patients (55;56). This NO deficiency is caused by a decreased bioavailability of L-arginine to NOS isozymes (49;57;58), which also leads to uncoupling of the oxidase and reductase moieties within the iNOS enzyme (50). Uncoupled iNOS not only produces NO but also superoxide, leading to an efficient formation of peroxynitrite (59) (Figure 1), which induces airway hyperresponsiveness, epithelial damage, mucus hypersecretion and inflammation (60-63).

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Figure 1 Putative role of altered L-arginine homeostasis in the airway epithelium in regulating remodeling of the ASM in asthma. The bioavailability of L-arginine to NOS isoforms is decreased is asthma, leading to a deficiency of bronchodilating NO as well as increased formation of procontractile peroxynitrite due to uncoupling of NOS. The NO deficiency may also contribute to the increased ASM mass in asthma, since NO is antiproliferative. Increased formation of peroxynitrite (ONOO-) could also contribute by causing epithelial damage. Reduced L-arginine bioavailability is caused by at least two mechanisms: i) inhibition of the cationic amino acid transporter by eosinophilic polycations, such as major basic protein (MBP), and ii) increased consumption of L-arginine by arginase, which is induced in asthma, presumably due to increased release of Th2-cytokines and growth factors. Increased arginase activity may directly contribute to the increased ASM mass via the production of polyamines and L-proline downstream from L-ornithine. See text for further detail. A deficiency of NO could contribute to airway smooth muscle thickening. It has been shown that NO inhibits mitogen-induced proliferation of cultured human (64-66) and guinea pig (67) ASM cells. Scavenging of superoxide anions, thereby increasing the levels of authentic NO and inhibiting peroxynitrite formation, also decreased mitogen-induced human ASM cell proliferation (66). The downstream mechanisms of NO-mediated inhibition of cell proliferation have been studied in more detail in VSMC and involve cGMP-dependent repression of cell cycle promoting genes, including cyclin D1, and the induction of cell cycle inhibitors, such as p21Waf1/Cip1 (68). Also in VSMC, NO has been shown to inhibit the 26S proteosome, which regulates the degradation of cell cycle proteins, via S-nitrosylation (69), whereas cGMP inactivates p42/p44 MAPK and activates MAPK phosphatase 1 (68). Moreover, NO attenuates PDGF-induced activation of protein kinase B (PKB) and subsequent VSMC proliferation (70) and reduces embryonic fibroblast proliferation by inhibiting EGF receptor tyrosine kinase activity via S-nitrosylation (71;72). Taken together,


35

these findings suggest that a deficiency of (epithelium-derived) NO in asthma may also contribute to increased ASM mass in chronic asthma (Figure 1).

An important mechanism contributing to the L-arginine limitation to NOS and subsequent NO deficiency in asthma is increased consumption of the amino acid by arginase, yielding L-ornithine and urea (Figure 1). Two arginase isoforms have been identified, the cytosolic arginase I and the mitochondrial arginase II, and both are expressed in the airway epithelium (73). Arginase activity and/or expression of particularly arginase I are increased in a number of animal models of acute allergic asthma (for review see: (74)) and in human asthmatics (75-77). Increased arginase activity, which may involve Th2 cytokines (77;78), importantly contributes to the development of allergen-induced bronchial obstructive reactions (58), airway hypersponsiveness in vivo and ex vivo (49;50;58;76;79) and airway inflammation (58). The functional significance of increased arginase activity in asthma is reinforced by reduced L-arginine levels due to inhibition of cellular L-arginine transport by eosinophil-derived polycations (50;80;81) (Figure 1).The release of NO by inhibitory nonadrenergic, noncholinergic (iNANC) neurons is also regulated by endogenous arginase (82). Moreover, the allergen-induced increase in arginase activity attenuates iNANC nerve-mediated NO release and ASM relaxation by limiting the L-arginine bioavailability (49). Thus, in addition to reduced ASM relaxation, alterations in L-arginine homeostasis in the iNANC nervous system may also contribute to airway remodeling via reduced synthesis of NO.

Increased arginase activity could also contribute to the pathophysiology of allergic asthma via increased synthesis of L-ornithine (Figure 1). L-Ornithine is a precursor of the polyamines (putrescine, spermidine and spermine) and L-proline, which are involved in cell proliferation and collagen synthesis, respectively (74;81;83-85). Polyamines induce the expression of genes involved in cell proliferation by promoting histone acetyltransferase activity and chromatin hyperacetylation (86); and polyamine synthesis is initiated by ornithine decarboxylase (ODC), which converts L-ornithine into putrescine (85). Both arginase and ODC are expressed in airway epithelial cells (87) and the expression and activation of both enzymes in the vasculature can be induced by growth factors, leading to increased polyamine levels (87-92). Growth factor-induced activation of ODC has also been observed in the airways (93). Interestingly, animal models demonstrate that arginase activity is increased in chronic asthma, underlying AHR in vivo (76) and ex vivo (94) by limiting the L-arginine availability to NOS. Moreover, elevated levels of polyamines have been detected in lungs of allergen-challenged mice (77) and in serum of asthmatic patients (95). These findings suggest that increased arginase activity may also contribute to the increased ASM mass in asthma via increased polyamine production. In support, transfection of VSMC with arginase I leads to elevated polyamine levels and increased cell proliferation (96). Since NO inhibits ODC via S-nitrosylation (97), allergen-induced deficiency of NO may contribute to the elevated polyamine levels in asthma.

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In conclusion, altered L-arginine homeostasis due to increased arginase activity in the airway epithelium of asthmatics could contribute to airway remodeling via increased production of polyamines and L-proline downstream of L-ornithine as well as by limiting the substrate availability to NOS enzymes, leading to NO deficiency and enhanced peroxynitrite formation (Figure 1). The airway parasympathetic nervous system The airway parasympathetic system, in which acetylcholine is the primary neurotransmitter, has long been recognized for its role in bronchoconstriction and mucus secretion (98;99). However, recent studies in guinea pigs and mice have revealed that acetylcholine, by acting on muscarinic receptors, is involved in the regulation of ASM mass and phenotype, suggesting an important role of the airway cholinergic system in regulating responses associated with remodeling (100-102). Indeed, the regulation of neuronal release of acetylcholine appears to be highly facilitated by eosinophilic airway inflammation and acetylcholine appears to have postjunctional effects on ASM that could explain its action as a regulator of airway wall remodeling, notably thickening of the smooth muscle (103;104) (Figure 2A).

The release of acetylcholine from parasympathetic nerves is enhanced in allergic airway inflammation because several mechanisms exist that allow inflammatory mediators to activate the cholinergic system and because allergic airway inflammation facilitates the output of parasympathetic nerve endings (99;104) (Figure 2A). Afferent sensory nerve fibres, or C-fibres, play an important role in this regard, as they can be triggered by a variety of inflammatory mediators and by non-specific stimuli such as cold air. This results in the local release of tachykinins as well as the activation of a cholinergic reflex mechanism that facilitates the output of the vagal nerve, both centrally and locally in the airway parasympathetic ganglia (105;106). C-fibres are exposed due to epithelial shedding in asthma and have receptors for histamine, prostanoids, thromboxane A2, bradykinin, serotonin and tachykinins (107;108). In conditions of damaged or stressed airway epithelium, the underlying mesenchyme can therefore be activated by this cholinergic reflex pathway, producing not only bronchoconstrictor responses but perhaps also responses associated with remodeling. The importance of this reflex mechanism is suggested by a recent study that showed that the majority of the bronchoconstrictor response to the inhaled thromboxane A2 mimetic U46619 is prevented by vagotomy or by administration of the M3 receptor selective ligand 4-DAMP (109). Furthermore, the airway hyperresponsiveness to inhaled histamine in ovalbumin sensitized guinea pigs after the early asthmatic reaction is markedly reduced by inhaled ipratropium bromide, indicating increased regulation of the cholinergic reflex in allergic airways disease (110). Such studies indicate that inflammatory mediators use the airway cholinergic system


37

to regulate a major part of their bronchoconstrictor response. In addition, several other mechanisms are at work in allergic airways disease that further induce the output of the vagal nerve. Inflammatory mediators including tachykinins, prostaglandins and thromboxane A2 facilitate neurotransmission in nerve endings and in the ganglia through effects on facilitatory prejunctional receptors (105;106). In addition, the prejunctional muscarinic M2 receptor which limits acetylcholine release under physiological conditions, is dysfunctional during allergic airway inflammation (98;111-113). Allergen-induced M2 dysfunction is regulated by eosinophils that are recruited to airway nerves, and secrete the allosteric muscarinic M2 receptor antagonist major basic protein (114). The above mentioned mechanisms collectively could well explain the increased role of the airway cholinergic system in airway hyperresponsiveness that is associated with loss of epithelial integrity. Since the airway cholinergic system also appears to regulate airway remodeling (100;101;115;116), the same may hold true for this pathological response.

Postjunctionally, there indeed are significant functional interactions between acetylcholine and growth factors that support ASM proliferation (Figure 2B). Muscarinic receptor stimulation by itself is not mitogenic to human ASM; however, in combination with growth factors such as PDGF or EGF, muscarinic M3 receptor stimulation augments the proliferative response to those factors, which likely involves the activation of multiple downstream effector pathways (117-119). In agreement with such an interaction, inhaled anticholinergics are effective in reducing ASM mass in guinea pigs that are allergen challenged, during which inflammatory mediators and growth factors are released, but have no effect on ASM mass in saline challenged controls (101). Muscarinic receptor stimulation cooperates with growth factor receptors to synergistically phosphorylate p70S6K and GSK-3, particularly in their late phase phosphorylation (2-4 h after stimulation) (118-121). P70S6K is required for ASM cell proliferation and hypertrophy and is activated upon phosphorylation, whereas GSK-3 is an anti-mitogenic and anti-hypertrophic kinase that is inhibited upon phosphorylation (Figure 2B) (118;122-125). Both actions of muscarinic receptors, which require the respective activation of PKC and �� subunits as signaling intermediates (118;120;121), support therefore ASM growth. Effects of muscarinic receptors on smooth muscle phenotype marker protein expression (e.g. sm-MHC) could also be explained this way, as both p70S6K and GSK-3 regulate the expression of smooth muscle specific proteins (123;126). Direct activation of smooth muscle specific genes by muscarinic receptor stimulation has indeed been demonstrated (127;128), but the underlying signaling events and interactions with growth factors still need to be assessed in future studies. Interestingly, the study by Fairbank et al., (127) showed that amplification of MLCK expression by muscarinic receptor stimulation occurred only in the presence of mechanical strain, which highlights another potentially important mechanism for remodeling that is regulated by mechanical factors. Mechanical strain on ASM will result in ASM cell proliferation

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(127;129) and mechanical compression of the airway wall is sufficient to activate EGF receptors present in the airway epithelium (130;131). These studies raise the real possibility that bronchodilation, for example using anticholinergic agents, reduces airway remodeling at least in part via the reduction of mechanical stress and strain within the airway wall. Figure 2 Interactions between the airway epithelium, the airway cholinergic system and the ASM regulates remodelling of the ASM bundle. A) Chronic airway inflammation facilitates acetylcholine release from the airway parasympathetic nerves, directly and via the activation of cholinergic reflex mechanisms, which are enhanced by the presence of damaged or stressed epithelium. B) As a result, increased acetylcholine release, in combination with growth factors and mediators released during inflammation coordinate cell responses in ASM associated with remodeling including smooth muscle specific gene expression and cell proliferation. The mechanisms responsible for these responses include phosphorylation of downstream signaling intermediates including p70S6K and GSK-3, resulting from muscarinic M3-receptor derived PI3K activation (via �� subunits) and PKC activation respectively. See text for further detail.


39

In conclusion, the above mentioned studies have provided solid support for the hypothesis that airway inflammation can interact with neuronally derived acetylcholine to facilitate bronchoconstriction and ASM thickening. Also non-neuronal acetylcholine, released for example by airway epithelial cells or inflammatory cells may contribute to these process, which needs to be established in future studies (104). Although this process is dependent on airway inflammation, the interactive role of the airway cholinergic system is considerable and appears to play a major role in regulating ASM mass and phenotype (Figure 2). The extracellular matrix The ECM is a dynamic structure that surrounds cells and provides the mechanical support required for airway structure and function. In the airway wall of asthmatics the amount and composition of the ECM is altered compared to healthy subjects. These changes are most eminent beneath the basement membrane and include increased deposition of collagens I, III and V, fibronectin, tenascin, hyaluronan, versican, biglycan, lumican and laminin �2/�2 as well as decreased expression of collagen IV, elastin and decorin (132-137) (Figure 3). Changes in the ECM have also been observed within and surrounding the ASM bundles. In patients with fatal asthma, the total amount of ECM within and surrounding the ASM bundles is increased, which was correlated with severity, but not duration, of asthma (20). This increase involves increased deposition of collagen I, fibronectin, hyaluronan, versican, biglycan, lumican and elastic fibres (138-140). A recent study, however, showed no changes in fractional area of collagens I or versican in the ASM layer (138). Interestingly, an inverse association between elastin expression and methacholine responsiveness has also been observed (10), suggesting that airway hyperresponsiveness is positively linked to the ECM expression in the ASM layer.

ASM cells are a rich source of ECM components, as shown by the production of collagens, fibronectin, laminins, perlecan, elastin, thrombospondin, versican and decorin by ASM (141-143), the expression of which is increased in response to profibrotic factors like TGF-�, CTGF and VEGF (144) (Figure 3). Interestingly, expression of CTGF in response to TGF-� by asthmatic ASM is increased compared to non-asthmatic ASM (22), identifying this factor as a potentially important contributor to ECM production in asthma. Altered ECM production by asthmatic ASM is also supported by findings showing an increased production of collagen I, perlecan and fibronectin (23;27), and a decreased production of laminin �1, chondroitin sulphate, collagen IV and hyaluronan compared to ASM derived from healthy subjects (27;145). In addition, increased expression of fibronectin, laminin �1, perlecan and chondroitin sulphate by non-asthmatic ASM cells was observed after exposure to atopic serum, indicating that plasma leakage may contribute to increased ECM production by ASM in asthma (143).

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Figure 3 ASM and ECM mutually affect each other to support abnormal ASM function as observed in asthma. A) In the asthmatic airways deposition of ECM proteins is increased, not only beneath the basement membrane but also within and surrounding the ASM bundle. Asthmatic ASM creates an altered ECM environment that facilitates increased contractile, proliferative and synthetic capabilities. B) Asthmatic ASM cells deposit increased amounts of fibronectin, which increases ASM synthetic function via a mechanism involving the �5�1 integrin. In addition, increased proliferation of asthmatic ASM has been shown to be dependent on the ECM, potentially also via a mechanism involving the �5�1 integrin. Laminins are critically involved in the expression of smooth muscle contractile proteins via the �7�1 integrin. Expression of this integrin has been shown to be increased by pro-remodeling factors like TGF-�. Altered deposition of ECM proteins may alter mechanical properties of ASM, as well as the transfer of force between the ASM bundle and surrounding tissue (146). Next to their role in structural support, ECM proteins also regulate the function of the cells embedded therein. ECM proteins have been found to differentially regulate survival, migration, cytokine synthesis, maturation, contractility and proliferation of ASM cells (Chapter 3)(31;149). The alterations in the ECM profile produced by asthmatic ASM cells therefore have the potential to influence behavior and characteristics of the ASM cells (Figure 3). In support of this, studies on the effects of asthmatic ECM showed that culture of both healthy and asthmatic ASM cells on an asthmatic ECM enhanced proliferative responses (27). Similarly, increased eotaxin expression by asthmatic ASM is dependent on the ECM produced by these cells (23). This increased secretory response required interaction of the ASM with its ECM via �5�1 integrins (23), an integrin of which the expression can be increased in response to TGF-� in both nonasthmatic and asthmatic ASM (148) (Figure 3B). Preliminary findings from our laboratory also suggest an important role for integrins in ASM remodeling in vivo. Using a guinea pig model of chronic allergic asthma, we found that treatment with the integrin-blocking peptide Arg-Gly-Asp-Ser (RGDS),


41

containing the RGD binding motif present in fibronectin, collagens and laminins (149;150), inhibits allergen-induced ASM hyperplasia, increased contractile protein expression and ASM hypercontractility, without effects on inflammatory responses (151). The normalization of allergen-induced hypercontractility also suggests a role for changes in ECM composition in regulating ASM contractility. Indeed, studies indicated that exogenously applied laminin-111 (laminin-1) maintains contractile ASM phenotype (Chapter 3)(152), whereas endogenously expressed laminin-211 (laminin-2) has been implicated in ASM maturation (153) and the induction of a hypercontractile ASM phenotype (Chapter 5). ASM maturation required activation of the laminin binding integrin �7 (154), of which expression is increased by TGF-� in both non-asthmatic and asthmatic ASM (155).

Collectively, these findings indicate that the ECM is not just an innocent bystander, but a component which can be actively regulated by the ASM, which in turn facilitates the abnormal ASM function as observed in asthma. These changes may be initiated by airway inflammation, but remain present in the absence of persistent inflammation. Intriguingly, these studies point to a dominant role of ASM – ECM interactions in the regulation of ASM remodeling and indicate that the muscle itself is capable of and in part responsible for creating an altered ECM environment that supports and maintains its increased contractile, proliferative and synthetic characteristics (Figure 3). Conclusions From numerous studies it is quite evident that ASM thickening is a prominent pathological feature in asthma, that contributes to an important extent to increased airway reactivity in patients. The mechanisms underlying this response remain elusive. Clearly, evidence is accumulating to indicate that the model, in which remodeling is due solely to the presence of inflammatory cells that secrete mediators and growth factors promoting cell proliferation and hypertrophy, is incomplete. Rather, the ASM layer is part of an active epithelial mesenchymal trophic unit that is activated during tissue injury and repair and driven by both changes in inflammatory cells and damaged epithelium. The damaged and stressed epithelium expresses increased levels of arginase, which reduces the presence of bronchodilatory and anti-proliferative NO, and promotes the presence of amino acids and polyamines that regulate smooth muscle remodeling. Furthermore, the damaged epithelium allows exposure of afferent sensory nerve endings that, together with the ongoing inflammation of the underlying airway wall, promotes the release of acetylcholine that acts as an important regulator of ASM remodeling via its actions on the postjunctional muscarinic M3 receptor. However, although epithelial cell changes and inflammation most likely play a major regulatory or initiating role, the studies summarized above also indicate that it is incorrect to assume that the underlying

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mesenchyme, including the ASM, is a passive partner in the remodeling process. The ASM actively participates in the remodeling process by regulating inflammation through the secretion of chemokines and cytokines, by producing force on the airway wall during periods of inflammation that regulates gene expression and kinase phosphorylation via mechanisms of mechanotransduction, and by producing an ECM that supports its multifunctional role with respect to its proliferative, secretory and contractile capacities. Therefore, these studies call for a model of bi-directional rather than uni-directional communication between components of the airway wall, in which ASM thickening is controlled by several structural components, including the muscle itself. References 1. Bousquet J, Jeffery PK, Busse WW, Johnson M, Vignola AM. Asthma. From

bronchoconstriction to airways inflammation and remodeling. Am J Respir Crit Care Med 2000;161:1720-1745.

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8. Leguillette R, Laviolette M, Bergeron C, Zitouni NB, Kogut P, Solway J, Kashmar L, Hamid Q, Lauzon AM. Myosin, Transgelin, and Myosin Light Chain Kinase: Expression and Function in Asthma. Am J Respir Crit Care Med 2008;179:194-204.

9. Leguillette R, Lauzon AM. Molecular mechanics of smooth muscle contractile proteins in airway hyperresponsiveness and asthma. Proc Am Thorac Soc 2008;5:40-46.

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12. Stephens NL, Fust A, Jiang H, Li W, Ma X. Isotonic relaxation of control and sensitized airway smooth muscle. Can J Physiol Pharmacol 2005;83:941-951.

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144. Burgess JK, Ceresa C, Johnson SR, Kanabar V, Moir LM, Nguyen TT, Oliver BG, Schuliga M, Ward J. Tissue and matrix influences on airway smooth muscle function. Pulm Pharmacol Ther 2009;5:379-387.

145. Klagas I, Goulet S, Karakiulakis G, Zhong J, Baraket M, Black JL, Papakonstantinou E, Roth M. Decreased hyaluronan in airway smooth muscle cells from patients with asthma and COPD. Eur Respir J 2009;34:616-628.

146. An SS, Bai TR, Bates JH, Black JL, Brown RH, Brusasco V, Chitano P, Deng L, Dowell M, Eidelman DH, Fabry B, Fairbank NJ, Ford LE, Fredberg JJ, Gerthoffer WT, Gilbert SH, Gosens R, Gunst SJ, Halayko AJ, Ingram RH, Irvin CG, James AL, Janssen LJ, King GG, Knight DA, Lauzon AM, Lakser OJ, Ludwig MS, Lutchen KR, Maksym GN, Martin JG, Mauad T, McParland BE, Mijailovich SM, Mitchell HW, Mitchell RW, Mitzner W, Murphy TM, Pare PD, Pellegrino R, Sanderson MJ, Schellenberg RR, Seow CY, Silveira PS, Smith PG, Solway J, Stephens NL, Sterk PJ, Stewart AG, Tang DD, Tepper RS, Tran T, Wang L. Airway smooth muscle dynamics: a common pathway of airway obstruction in asthma. Eur Respir J 2007;29:834-860.

147. Fernandes DJ, Bonacci JV, Stewart AG. Extracellular matrix, integrins, and mesenchymal cell function in the airways. Curr Drug Targets 2006;7:567-577.

148. Moir LM, Burgess JK, Black JL. Transforming growth factor beta(1) increases fibronectin deposition through integrin receptor alpha(5)beta(1) on human airway smooth muscle. J Allergy Clin Immunol 2008;121:1034-1039.

149. Aumailley M, Gerl M, Sonnenberg A, Deutzmann R, Timpl R. Identification of the Arg-Gly-Asp sequence in laminin A chain as a latent cell-binding site being exposed in fragment P1. FEBS Lett 1990;262:82-86.

150. Plow EF, Haas TA, Zhang L, Loftus J, Smith JW. Ligand binding to integrins. J Biol Chem 2000;275:21785-21788.

151. Dekkers BG, Bos IS, Gosens R, Halayko AJ, Zaagsma J, Meurs H. Inhibition of Airway Smooth Muscle Remodeling in an Animal Model of Chronic Asthma by the Integrin-Blocking Peptide RGDS. Am.J.Respir.Crit Care Med. 179, A5600. 2009.

152. Hirst SJ, Twort CH, Lee TH. Differential effects of extracellular matrix proteins on human airway smooth muscle cell proliferation and phenotype. Am J Respir Cell Mol Biol 2000;23:335-344.

153. Tran T, McNeill KD, Gerthoffer WT, Unruh H, Halayko AJ. Endogenous laminin is required for human airway smooth muscle cell maturation. Respir Res 2006;7:117.

154. Tran T, Ens-Blackie K, Rector ES, Stelmack GL, McNeill KD, Tarone G, Gerthoffer WT, Unruh H, Halayko AJ. Laminin-binding Integrin {alpha}7 is Required for Contractile Phenotype Expression by Human Airway Myocyte. Am J Respir Cell Mol Biol 2007;37:668-680.

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155. Kanabar V, Karner LC, Simcock DE, Mahn K, Nguyen NM, Cousins DJ, et al. Integrin gene profiling in airway smooth muscle from asthmatics. Proc.Am.Thorac.Soc. 4, A302. 2007.

Bart G.J. Dekkers Dedmer Schaafsma S. Adriaan Nelemans Johan Zaagsma Herman Meurs Am J Physiol Lung Cell Mol Physiol (2007) 292: L1405-1413

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hapter Extracellular matrix proteins differentially regulate airway smooth muscle phenotype and function

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Abstract Changes in the ECM and increased airway smooth muscle (ASM) mass are major contributors to airway remodelling in asthma and chronic obstructive pulmonary disease. It has recently been demonstrated that ECM proteins may differentially affect proliferation and expression of phenotypic markers of cultured airway smooth muscle cells. In the present study, we investigated the functional relevance of ECM proteins in the modulation of ASM contractility using bovine tracheal smooth muscle (BTSM) preparations. The results demonstrate that culturing of BTSM strips for 4 days in the presence of fibronectin or collagen I depressed maximal contraction (Emax) both for methacholine and KCl, which was associated with decreased contractile protein expression. By contrast, both fibronectin and collagen I increased proliferation of cultured BTSM cells. Similar effects were observed for PDGF. Moreover, PDGF augmented fibronectin- and collagen I- induced proliferation in an additive fashion, without an additional effect on contractility or contractile protein expression. The fibronectin-induced depression of contractility was blocked by the integrin antagonist Arg-Gly-Asp-Ser (RGDS), but not by its negative control Gly-Arg-Ala-Asp-Ser-Pro (GRADSP). Laminin, by itself, did not affect contractility or proliferation but reduced the effects of PDGF on these parameters. Strong relationships were found between the ECM-induced changes in Emax in BTSM strips and their proliferative responses in BTSM cells and for Emax and contractile protein expression. Our results indicate that ECM proteins differentially regulate both phenotype and function of intact ASM. Introduction The ECM is an intricate network of macromolecules that surrounds the tissue cells and affects many aspects of cellular behavior. These include migration, differentiation, survival and proliferation of cells originating from a variety of tissues, including airway smooth muscle (ASM) (14).

Biopsy studies have revealed that both the quantity and the composition of the ECM is altered in the airways of chronic asthmatics. Deposition of collagen IV and elastin is decreased in the airway wall of asthmatic patients, whereas collagen I, III ,V, fibronectin, tenascin, hyaluran, versican and laminin �2/�2 chains are increased compared to healthy subjects (1; 15; 16; 24; 25).

Increased ASM mass within the airway wall is a characteristic feature of chronic asthma and may be one of the mechanisms associated with increased airway responsiveness and decline of lung function (4; 13; 26). Increased ASM cell mass is believed to involve both cellular hyperplasia and hypertrophy (6). Mechanisms involved in increased ASM growth in asthma are currently largely unknown; however, changes in the composition of the ECM proteins surrounding the ASM cell might well be involved (12).

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In the airway wall of healthy subjects, the smooth muscle layer consists mainly of differentiated ASM cells, which are characterized by low proliferation rates, low fractions of biosynthetic organelles and relatively high expression levels of contractile proteins, including smooth muscle �-actin (sm-�-actin), calponin and smooth muscle myosin heavy chain (sm-MHC) (13). Importantly, in contrast to skeletal myocytes and cardiomyocytes (17; 19), ASM cells maintain the ability to re-enter the cell-cycle. Thus, exposure to mitogenic stimuli (e.g. PDGF), results in the induction of a more proliferative/synthetic phenotype (13), which is accompanied by a loss of contractile responsiveness (8), presumably as a consequence of decreased contractile protein expression (13). Long term serum deprivation results in the reinduction of a contractile phenotype, underlining the reversible nature of ASM phenotype (18). This phenotypic plasticity might be involved in growth and repair processes of inflamed airways and may contribute to airway remodeling in chronic asthma (12).

Using ASM cells in culture, it has recently been indicated that ECM proteins may differentially affect growth factor-induced phenotypic modulation. Thus, in human ASM cells cultured on fibronectin or collagen I matrices progression towards a proliferative phenotype, induced by either PDGF or �-thrombin, was promoted, whereas culturing on a laminin or matrigel matrix inhibited phenotype switching by these mitogens (12). Enhancement of PDGF-dependent proliferation of human ASM cells on a fibronectin or collagen I matrix has been shown to be dependent on activation of �2�1, �4�1 and �5�1 integrins (22). In vascular smooth muscle (VSM) cells, expression of �1�1 and �7�1 integrins has been correlated with the differentiated smooth muscle phenotype (2; 30). Both integrins are capable of binding laminin, which has been implicated in maintaining contractile VSM phenotype (20).

At present, the functional significance of ASM phenotypic modulation by ECM proteins is unknown. Therefore, we investigated the effects of exogenously applied fibronectin, collagen I and laminin on BTSM strip contractility in relation to the proliferative response of BTSM cells under these conditions.Our results indicated that fibronectin and collagen I induce a less contractile BTSM phenotype, whereas laminin maintains contractility. However, laminin attenuates the suppressive effects of PDGF on contractility as well as PDGF-induced DNA synthesis. Our results demonstrate for the first time a differential effect of ECM proteins on both phenotype and contractile function of intact ASM.

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Materials and methods Tissue preparation and organ culture procedure Bovine tracheae were obtained from local slaughterhouses and rapidly transported to the laboratory in ice-cold Krebs-Henseleit (KH) buffer of the following composition (mM): NaCl 117.5, KCl 5.60, MgSO4 1.18, CaCl2 2.50, NaH2PO4 1.28, NaHCO3 25.00 and glucose 5.50, pregassed with 5% CO2 and 95% O2; pH 7.4. After dissection of the smooth muscle layer and careful removal of mucosa and connective tissue, tracheal smooth muscle strips were prepared while incubated in gassed KH-buffer at room temperature (RT). Care was taken to cut tissue strips of macroscopically identical length (1 cm) and width (2 mm). Tissue strips were washed once in Medium Zero (sterile DMEM (Gibco BRL Life Technologies, Paisley, UK), supplemented with sodium pyruvate (1 mM, Gibco), nonessential amino-acid mixture (1 :100, Gibco), gentamicin (45 �g/ml, Gibco), penicillin (100 U/ml, Gibco), streptomycin (100 �g/ml, Gibco), amphotericin B (1.5 �g/ml, Fungizone, Gibco), apo-transferrin (5 �g/ml, human, Sigma Chemical Co, St. Louis, MO, USA) and ascorbic acid (100 �M, Merck, Darmstadt, Germany)). Next, tissue strips were transferred into suspension culture flasks and a volume of 7.5 ml Medium Zero was added per tissue strip. Strips were maintained in culture in an Innova 4000 incubator shaker (37°C, 55 rpm) under tightly controlled conditions for 4 days. To avoid direct influences of mechanical plasticity during this culture, strips were maintained under unloaded conditions. When used, collagen I (50 �g/ml, monomeric, calf skin, Fluka, Buchs, Switzerland), fibronectin (10 �g/ml, bovine plasma, Sigma), Engelberth-Holm-Swarm (EHS) Sarcoma laminin, consisting of laminin-1 (27) (4 �g/ml, Invitrogen, Grand Island, NY, USA), PDGF-AB (10 ng/ml, human, Bachem, Weil am Rhein, Germany), Arg-Gly-Asp-Ser (RGDS, 0.1 mM, Calbiochem, Nottingham, UK) and/or Gly-Arg-Ala-Asp-Ser-Pro (GRADSP, 0.1 mM, Calbiochem) were present during the entire incubation period. Occasionally, some strips were used for isometric tension measurements directly after preparation. Isometric tension measurements Tissue strips, collected from the suspension culture flasks, were washed with several volumes of KH-buffer pregassed with 5% CO2 and 95% O2, pH 7.4, at 37°C. Subsequently, strips were mounted for isometric recording (Grass force-displacement transducer FT03) in 20 ml water-jacked organ baths, containing KH-buffer at 37°C, continuously gassed with 5% CO2 and 95% O2, pH 7.4. During a 90-min equilibration period, with washouts every 30 min, resting tension was gradually adjusted to 3 g. Subsequently, muscle strips were precontracted with 20 and 40 mM isotonic KCl solutions. Following two washouts, maximal relaxation was established by the addition of 0.1 �M (-)-isoproterenol (Sigma). In most the experiments, no basal myogenic tone was detected. Tension was readjusted to 3 g, immediately followed by two changes with fresh KH-buffer. After another equilibration period of 30 min, cumulative


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concentration response curves (CRCs) were constructed to stepwise increasing concentrations of isotonic KCl (5.6�50 mM) or methacholine (1 nM�100 �M, ICN biomedicals, Costa Mesa, CA, USA). When maximal tension was obtained, the strips were washed several times and maximal relaxation was established using (-)-isoproterenol. Isolation of bovine tracheal smooth muscle (BTSM) cells After the removal of mucosa and connective tissue, tracheal smooth muscle was chopped using a McIlwain tissue chopper, three times at a setting of 500 �m and three times at a setting of 100 �m. Tissue particles were washed two times with Medium Plus (DMEM supplemented with sodium pyruvate (1 mM), nonessential amino-acid mixture (1 :100), gentamicin (45 �g/ml), penicillin (100 U/ml), streptomycin (100 �g/ml), amphotericin B (1.5 �g/ml) and FBS (0.5%, Gibco)). Enzymatic digestion was performed in Medium Plus, supplemented with collagenase P (0.75 mg/ml, Boehringer, Mannheim, Germany), papain (1 mg/ml, Boehringer) and soybean trypsin inhibitor (1 mg/ml, Sigma). During digestion, the suspension was incubated in an incubator shaker (Innova 4000) at 37°C, 55 rpm for 20 min, followed by a 10-min period of shaking at 70 rpm. After filtration of the obtained suspension over a 50 �m gauze, cells were washed three times in Medium Plus, supplemented with 10% FBS instead of 0.5% FBS. Coating of culture plates with extracellular matrix proteins Calf skin collagen I was reconstituted in 10 mM hydrochloric acid at 5 mg/ml before diluting. Bovine plasma fibronectin was reconstituted in sterile PBS (composition mM): NaCl 140.0; KCl 2.6; KH2PO4 1.4; Na2HPO4.2H2O 8.1; pH 7.4. Dilutions of collagen I, fibronectin and mouse laminin were prepared in PBS. Diluted ECM proteins (0.5 ml) were absorbed to 24-well cluster plates overnight and air-dried at RT. Unoccupied protein-binding sites were blocked by a 30 min incubation with a sterile 0.1% bovine serum albumin (BSA, Sigma) solution. Subsequently, plates were washed twice with Medium Zero and dried before further use. [3H]-Thymidine-incorporation BTSM cells were plated on uncoated or ECM-coated 24-well cluster plates at a density of 50,000 cells per well immediately after isolation and were allowed to attach overnight in Medium Plus, containing 10% FBS. Cells were washed twice with sterile PBS and made quiescent by incubation in Medium Zero, supplemented with apo-transferrin (5 �g/ml), ascorbic acid (100 �M) and insulin (1 �M, bovine pancreas, Sigma) for 72 h. Cells were then washed with PBS and incubated with or without PDGF in Medium Zero for 28 h, the last 24 h in the presence of [methyl-3H]-thymidine (0.25 �Ci/ml, Amersham, Buckinghamshire, UK). After incubation the cells were washed twice with 0.5 ml PBS at RT. Subsequently, the cells were treated with 0.5 ml ice-cold 5% trichloroacetic acid on ice for 30 min, and the acid-insoluble fraction was dissolved in 1 ml NaOH (1

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M). Incorporated [3H]-thymidine was quantified by liquid-scintillation counting using a Beckman LS1701 �-counter. MTT assay BTSM cells were plated as described for the [3H]-thymidine-incorporation protocol. Following quiescence, cells were incubated with vehicle or PDGF for 3 days, after which cell number was estimated using the mitochondrial-dependent reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT, Sigma) to formazan. Briefly, cells were washed twice with Medium Zero after which 200 �l medium containing 0.5 mg/ml MTT was added to each well. After 5 h, 200 �l 10% sodium dodecylsulphate in 0.01 N HCl was added, and the cells were solubilized overnight at 37 °C. The amount of formazan in the obtained solution was estimated by measuring optical density at a test wavelength of 550 and a reference wavelength of 655 nm. Western analysis of contractile protein expression After culturing the tissue strips as described above, homogenates were prepared by pulverizing the tissue under liquid nitrogen, followed by sonification in homogenization buffer (composition in mM: Tris-HCl 50 mM, NaCl 150.0, EDTA 1.0, PMSF 1.0, Na3VO4 1.0, NaF 1.0, pH 7.4, supplemented with 10 �g/ml leupeptin (Sigma), 10 �g/ml aprotinin (Sigma), 10 �g/ml pepstatin (Sigma), Na-deoxycholate 0.25 % (Sigma) and 1% Igepal (NP-40, Sigma)). Homogenates were stored at �80 °C until further use. Protein content was determined according to Bradford (5). In total, 30 �g of protein per lane was separated by SDS/PAGE using 6% polyacrylamide gels for smooth muscle myosin (sm-myosin) or 10% polyacrylamide gels for �-actin, smooth muscle �-actin (sm-�-actin) and calponin. Proteins in the gel were then transferred onto nitrocellulose membranes, which were subsequently blocked in blocking buffer (composition: Tris-HCl 50.0 mM; NaCl 150.0 mM; Tween-20 0.1%, dried milk powder 5%) for 90 min at RT. Next, membranes were incubated overnight at 4 °C with primary antibodies (anti-sm-myosin (Neomarkers, Fremont, CA, USA), anti-sm-�-actin (Sigma), both diluted 1:200, anti-�-actin (Sigma), diluted 1:2000, calponin (Neomarkers), diluted 1:400; all dilutions in blocking buffer). Membranes were incubated with antibodies for �-actin to normalize for equal loading of all samples. After three washes of 10 min each, membranes were incubated with horseradish peroxidase-labelled secondary antibodies (dilution 1:3000 in blocking buffer) at RT for 90 min, followed by another three washes. Antibodies were then visualized by enhanced chemiluminescence. Blots were analyzed by densitometry (Totallab). All bands were normalized to �-actin expression. ECM-induced changes in protein abundance were expressed as a percentage of vehicle treated controls run on the same gels.


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Data analysis Data represent means ± S.E.M. or S.D., from n separate experiments. Statistical significance of differences was evaluated by the Student's t-test for paired observations or one-way ANOVA, as appropriate. Differences were considered to be statistically significant when P<0.05. Results Effect of organ culturing on bovine tracheal smooth muscle strip contractility and contractile protein expression Maximal methacholine and KCl-induced contractile force (Emax) of BTSM strips, cultured for 4 days in Medium Zero, was maintained as compared to freshly isolated BTSM strips (Figure 1A, 1B). No changes in sensitivity were observed for methacholine (pEC50 = 6.99 ± 0.17) or KCl ( Ec50 = 26.5 ± 1.0). Expression of the contractile protein sm-myosin in BTSM strip homogenates was preserved after 4 days of organ culturing (94 ± 8% compared with fresh, Figure 1C).

Figure 1: Concentration-response curves of methacholine- (A) and KCl-induced (B) contraction of freshly isolated and 4 day organ cultured BTSM strips. Data represent means ± S.E.M. of 3 experiments. (C) Western analysis of protein expression in fresh and organ cultured BTSM strips. Representative immunoblots of sm-myosin and �-actin are shown.

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Effects of pretreatment with extracellular matrix proteins on bovine tracheal smooth muscle strip contractility In BTSM strips cultured for 4 days in the presence of fibronectin (10 �g/ml) or collagen I (50 �g/ml), maximal methacholine-induced contractile force was significantly reduced compared to strips cultured in the absence of these ECM proteins (Figures 2A, 2B, Table 1). The suppressive effects on Emax were quantitatively similar to the effects observed after pretreatment with PDGF (10 ng/ml). Combined pretreatment of the strips with the ECM proteins (fibronectin and collagen I) and PDGF did not further affect maximal contraction. Unlike fibronectin and collagen I, pretreatment with laminin (4 �g/ml) did not affect Emax of methacholine. Interestingly, however, co-incubation with laminin fully reversed the suppressive effects of PDGF on Emax (Figure 2C, Table 1). Similar effects of ECM proteins and PDGF were obtained for KCl-induced contractions (Figure 3, Table 1). Figure 2: Concentration-response curves of methacholine-induced contraction of BTSM strips pretreated with (A) fibronectin (10 �g/ml), (B) collagen I (50 �g/ml) or (C) laminin (4 �g/ml) in the absence or presence of PDGF (10 ng/ml) for 4 days. Data represent means ± S.E.M. of 6-7 experiments.


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Table 1: Effects of 4 days culturing in the absence (vehicle) or presence of fibronectin (10 �g/ml), collagen I (50 �g/ml) or laminin (4 �g/ml), with or without PDGF (10 ng/ml), and the effects of 4 days culturing in the absence or presence of fibronectin with or without RGDS (0.1 mM) or GRADSP (0.1 mM), on contractile responses of BTSM strips to methacholine and KCl.

Methacholine KCl Emax (g) pEC50 (-log M) Emax (g) EC50 (mM) Vehicle 28.9±1.7 6.88±0.12 19.3±1.3 21.8±1.3

+ PDGF 24.8±1.2* 6.63±0.11 15.5±1.4** 25.7±1.3 Fibronectin 23.4±0.9* 6.53±0.10 15.4±1.5*** 26.1±1.7

+ PDGF 21.6±2.1* 6.60±0.06 14.8±2.0** 25.6±0.7 Vehicle 29.9±1.6 6.64±0.09 20.6±1.6 25.1±1.3

+ PDGF 24.1±1.9*** 6.62±0.11 15.7±1.3*** 24.7±1.1 Collagen I 22.1±2.4* 6.41±0.11 14.4±2.0* 25.8±0.9

+ PDGF 21.5±2.2** 6.50±0.11 14.8±1.0** 26.6±0.9 Vehicle 26.8±1.7 6.74±0.14 19.3±1.5 22.6±1.2

+ PDGF 21.2±1.7** 6.54±0.11 15.5±1.4** 24.5±1.1 Laminin 29.0±2.7## 6.65±0.13 21.7±2.6## 24.0±1.3

+ PDGF 26.8±2.4# 6.71±0.11 18.2±4.1# 24.3±0.8 Vehicle 28.7±1.8 6.68±0.10 19.2±2.1 25.4±0.9

+ GRADSP 28.8±1.6 6.59±0.11 18.6±2.0 27.2±0.7 +RGDS 24.9±2.7 6.67±0.12 16.8±3.2 25.6±1.0

Fibronectin 21.9±1.6* 6.79±0.20 14.8±1.8* 24.7±1.9 + GRADSP 22.7±1.7* 6.58±0.04 14.8±3.1* 26.8±0.6

+ RGDS 31.1±1.5† 6.54±0.19 20.9±2.1†† 25.4±0.8

Data represent means ± S.E.M. from 4-7 experiments. Abbreviations: Emax: maximal contraction; EC50: concentration of agonist eliciting half-maximal response; pEC50 negative logarithm of the EC50 value. *P<0.05, **P<0.01, ***P<0.001 compared with vehicle-treatment. #P<0.05, ##P<0.01 compared with PDGF-treatment. †P<0.05, ††P<0.01 compared with fibronectin.

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C Figure 3: Concentration-response curves of KCl-induced contraction of BTSM strips pretreated with (A) fibronectin (10 �g/ml), (B) collagen I (50 �g/ml) or (C) laminin (4 �g/ml) in the absence or presence of PDGF (10 ng/ml) for 4 days. Data represent means ± S.E.M. of 6-7 experiments. Combined pretreatment of BTSM strips with fibronectin and its blocking peptide Arg-Gly-Asp-Ser (RGDS, 0.1 mM) normalized Emax for both methacholine and KCl (Figure 4, Table 1), whereas no effects of the negative control, Gly-Arg-Ala-Asp-Ser-Pro (GRADSP, 0.1 mM), were observed. BTSM strips pretreated with RGDS or GRADSP in the absence of fibronectin did not show any significant effect on ASM contractility (Table 1). The sensitivity to methacholine or KCl was unaffected by all treatments (Table 1). It could be envisaged that pretreatment of strips with ECM proteins affect contractility by altering smooth muscle stiffness. To address this issue, BTSM strips were incubated with vehicle and collagen I (50 �g/ml)-containing media for 4 days. After this incubation period, strip length and width were assessed just before mounting and at a resting tension of 3 g. No differences between vehicle and collagen I pretreated strips were found for both parameters (Table 2). Table 2: Effect of collagen I (50 �g/ml) pretreatment on BTSM strip length and width before mounting and at resting tension (3 g).

Before mounting At resting tension Pretreatment Length (cm) Width (cm) Length (cm) Width (cm) Vehicle 0.88 ± 0.03 0.30 ± 0.03 1.18 ± 0.23 0.19 ± 0.01 Collagen I 0.89 ± 0.03 0.30 ± 0.03 1.15 ± 0.20 0.19 ± 0.02

Results are means ± S.D. of two separate experiments, each performed in duplicate


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B KClA Methacholine Figure 4: Concentration-response curves of methacholine- (A) and KCl-induced (B) contraction of BTSM strips pretreated with fibronectin (10 �g/ml) in the absence or presence of Arg-Gly-Asp- (RGD; 0.1 mM) or Arg-Ala-Asp- (RAD; 0.1 mM)-containing peptides for 4 days. Data represent means ± S.E.M. of 4-5 experiments. Effects of extracellular matrices on bovine tracheal smooth muscle cell proliferation. To establish the effects of fibronectin, collagen I and laminin on BTSM cell proliferation, both [3H]-thymidine incorporation and mitochondrial-dependent reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) to formazan were assessed. In BTSM cells, basal DNA-synthesis was increased by approximately 1.7-fold in cells grown on fibronectin (10 �g/ml) and by 2.5-fold in cells grown on collagen I (50 �g/ml) matrices compared to cells grown on plastic (control; Figure 5A). Under control conditions, PDGF significantly augmented DNA-synthesis by 2.5-fold (Figure 5A). This response was significantly enhanced in an additive fashion, when cells were grown on a fibronectin or collagen I matrix. In contrast, cells attached to a laminin matrix (4 �g/ml) did not show a significant change in basal DNA synthesis. Growing BTSM cells on laminin, however, resulted in a significant reduction of the PDGF-induced proliferative response, to approximately 0.6-fold of PDGF-induced proliferation in cells grown on plastic (Figure 5A).

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Figure 5: Effects of fibronectin (10 �g/ml), collagen I (50 �g/ml) and laminin (4 �g/ml) matrices on basal (black bars) and PDGF (10 ng/ml)-stimulated (grey bars) BTSM cell DNA synthesis (A) and BTSM cell number (B). Data represent means ± S.E.M. of 3-5 experiments each performed in triplicate. *P<0.05, **P<0.01 compared to basal controls. #P< 0.05, ##P<0.01 compared to PDGF control. †P<0.05, ††P<0.01 compared to ECM protein in the absence of PDGF. Since increased DNA synthesis may reflect both cell hyperplasia and hypertrophy, we also assessed the effects of ECM proteins and PDGF on cell number using the MTT assay. When cells were grown on fibronectin or collagen I, the cell number was augmented by approximately 1.6-fold as compared to cells grown on plastic (control, Figure 5B). Surprisingly, laminin induced a slight, but significant, increase in cell number as well. PDGF significantly augmented cell number to a similar degree as collagen I and fibronectin, and, as observed for DNA-synthesis, PDGF increased the cell number in an additive fashion when cells were grown on fibronectin or collagen I matrices. In line with the effects on DNA-synthesis, laminin significantly reduced the PDGF-mediated increase in cell number (Figure 5B). Relationship between extracellular matrix-induced changes in contractility and proliferation It has been previously established that growth factor-induced changes in maximal contractility of BTSM strips are inversely correlated with changes in the proliferative response of isolated BTSM cells (8). As illustrated by Figure 6, a qualitatively similar relationship exists between the effects of the applied ECM proteins on proliferative potency, as assessed by [3H]-thymidine incorporation in isolated BTSM cells, and the effects on Emax of methacholine- and KCl-induced contraction of BTSM strips.


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Figure 6: Relationships between maximal contraction (Emax) of BTSM strips in response to methacholine (A) or KCl (B) and DNA synthesis of BTSM cells in the presence of laminin (1), vehicle (2), fibronectin (3) or collagen I (4). Data represent means from 5�7 experiments. Effects of extracellular matrix proteins on contractile protein expression The changes in Emax in response to both receptor-dependent (methacholine) and receptor-independent (KCl) stimuli indicate post-receptor changes, which may occur at the level of the contractile apparatus. BTSM strips pretreated with PDGF showed a significant reduction in the expression of sm-myosin, calponin and sm-�-actin as compared to control conditions (Figure 7, Table 3). Similar results were observed after pretreatment with fibronectin or collagen I or the combined pretreatment of these ECM proteins with PDGF (Figure 7, Table 3). In contrast, strips cultured in the presence of laminin showed a significant increase in sm-myosin and calponin expression, whereas the expression level of sm-�-actin was not increased. Interestingly, although laminin normalized the suppressive effects of PDGF on BTSM strip contractility (Figures 2 and 3) and reduced the mitogenic capacity of the growth factor (Figure 5), no significant effects of laminin were observed on the PDGF-induced reduction of sm-myosin, calponin or sm-�-actin protein expression (Figure 7, Table 3).

Chapter 3

66

Figure 7: Western analysis of protein expression in BTSM strips pretreated with vehicle (control), collagen I (50 �g/ml), fibronectin (10 �g/ml) and laminin (4 �g/ml), in the absence or presence of PDGF (10 ng/ml). Representative immunoblots of sm-myosin, calponin, sm-�-actin and �-actin are shown. Table 3: Contractile protein expression in BTSM strips after 4 days of culturing with vehicle (control), collagen I (50 �g/ml), fibronectin (10 �g/ml) and laminin (4 �g/ml), in the absence and presence of PDGF (10 ng/ml).

Protein expression (% of control)

sm-myosin calponin sm-�-actin Control 100 100 100

+ PDGF 47±8** 61±9** 41±11**

Fibronectin 48±13** 64±12* 42±8***

+ PDGF 55±10** 68±13* 39±11**

Collagen I 59±5** 38±5** 33±7**

+ PDGF 56±6** 55±9* 31±3*** Laminin 134±13*,## 142±16*,## 97±15###

+ PDGF 63±5***,†† 80±12† 49±8**,†

Data represent means ± S.E.M. from 3-7 experiments. *P<0.05, **P<0.01, ***P<0.001 compared with vehicle-treatment; ##P<0.01, ###P<0.001 compared with PDGF-treatment. †P<0.05, ††P<0.01 compared with ECM protein in the absence of PDGF. Relationship between extracellular matrix protein- induced changes in contractile protein expression and contractility Strong correlations were observed between the efficacy of the ECM proteins to affect sm-myosin expression and Emax of methacholine (Figure 8A) or KCl (Figure 8B). Similar results were obtained for calponin expression (r=0.983, P=0.017 and r=0.993, P=0.007, respectively; data not shown). For the relationship between ECM effects on sm-�-actin content and Emax a strong


67

Emax (% of vehicle)

60 70 80 90 100 110 120

0

20

40

60

80

100

120

140

160

r = 0.992

P = 0.008

60 70 80 90 100 110 120

Rel

ativ

e pr

otei

n ex

pres

sion

(% o

f veh

icle

)

0

20

40

60

80

100

120

140

160

r = 0.968

P = 0.032

1

2

4

1

2

334

B KClA Methacholine

tendency was observed for both methacholine and KCl-induced contractions (r=0.942, P=0.058, r=0.947, P=0.053, respectively; data not shown). Figure 8: Relationships between maximal contraction (Emax) of BTSM strips in response to methacholine (A) or KCl (B) and sm-myosin expression in the presence of laminin (1), vehicle (2), fibronectin (3) or collagen I (4). Data represent means from 3-7 experiments. Discussion In this study, we demonstrate for the first time that prolonged (4 days) exposure of intact ASM strips to exogenous ECM proteins may differentially regulate ASM phenotype and function. Thus, exogenously applied fibronectin and collagen I induced a functionally hypocontractile ASM phenotype, characterized by a decreased maximal contractile response to both the muscarinic receptor agonist methacholine and the membrane depolarizing agent KCl. These effects are presumably due to phenotype modulation, since no differences were found in contractility between fresh and cultured strips and no effects were observed on equilibration length. Phenotypic modulation is also supported by the observation that the effects of fibronectin were fully normalized in the presence of RGDS, but not its negative control GRADSP. Similar effects of RGD- and RAD-containing peptides on fibronectin-induced eotaxin release have been observed in human ASM cells (23).

In accordance with previous findings (8; 9), 4 days incubation with PDGF also induced a hypocontractile phenotype. No additive effects of combined pretreatment with the growth factor and fibronectin or collagen I were found on maximal contractility. Interestingly, although laminin had no effect on BTSM

Chapter 3

68

contractility, co-incubation with PDGF fully normalized the suppressive effects of the growth factor on Emax, indicating that laminin may be involved in maintaining a (normo)contractile phenotype. It remains to be determined, however, to what extent the ECM proteins penetrate the tissue and to what extent the observed effects of the ECM proteins represent their maximal effect.

Both in vascular (11; 29) and in ASM (3; 12) cells, it has been demonstrated that ECM proteins are capable of differentially influencing the mitogenic capacity of a variety of growth factors, including PDGF, �-thrombin and basic fibroblast growth factor (bFGF). In human ASM cells, Hirst et al (12) showed that mitogen-stimulated, but not basal, proliferation was significantly enhanced after culturing on a collagen I or fibronectin matrix, whereas mitogen-induced proliferation was reduced on laminin-precoated plates (12). Accordingly, we found that culturing BTSM cells on a fibronectin or collagen I matrix significantly augmented proliferation induced by PDGF. Moreover, basal DNA-synthesis and cell number were also significantly increased by these ECM proteins, which is in agreement with previous findings showing that proliferation of bovine ASM cells was increased on a collagen type I matrix as compared to cells grown on a laminin matrix (3). As the enhancement of the PDGF-induced proliferative effects by a fibronectin or collagen I matrix was additive, it can be envisaged that these matrix proteins and PDGF regulate mitogenesis through distinct rather than common pathways. However, little is known about the pathways involved in these processes, which warrents further investigation.

Laminin, by itself, had no effect on DNA synthesis, whereas cell number, as assessed by formazan accumulation, was slightly increased. A possible explanation for this apparent discrepancy might be the anti-apoptotic potency of laminin as observed in human ASM cells (7). This may result in a preservation of the number of viable cells without increasing DNA synthesis. In agreement with previous studies, we found that laminin reduced proliferation induced by PDGF (12). Also, in porcine coronary artery smooth muscle cells, it has been shown that proliferation in response to growth factors was more pronounced in cells grown on fibronectin than on laminin (21). PDGF-mediated activation of ERK1/2 in these cells was not dependent on the matrix present, whereas activation of FAK was more pronounced on a fibronectin matrix (21). Those findings indicate a pivotal role for ECM in controlling intracellular signaling.

Modulation towards a less contractile ASM phenotype is accompanied by a reduced expression of contractile proteins, including sm-MHC, calponin and sm-�-actin (10; 28). Since prolonged (4 days) culturing in the presence of fibronectin or collagen I induced a decline in maximal contraction both in response to a receptor-dependent (methacholine) and a receptor-independent (KCl) stimulus, post-receptor events such as alterations in contractile protein expression are likely to contribute to the observed changes in contractility. Therefore, we assessed the expression of sm-myosin, calponin and sm-�-actin in homogenates prepared from BTSM strips treated with vehicle, collagen I, fibronectin or laminin in the absence or presence of PDGF for 4 days. In


69

accordance with our previous (8; 9) and current observations that PDGF induces a shift towards a more proliferative (hypocontractile) phenotype, the growth factor reduced the expression of all contractile markers studied. Incubation with fibronectin or collagen I resulted in a reduction of contractile protein expression as well, which was both quantitatively and qualitatively similar to that induced by PDGF. These observations, along with the fact that 4 days of organ culturing, by itself, did not affect sm-myosin expression, confirm the assumption that pretreatment with fibronectin or collagen I, in the absence or presence of PDGF, induces a functional hypocontractile phenotype by reducing contractile protein expression. Similar to the effects of combined treatment on Emax, no additional effects were observed on the level of contractile protein expression, indicating that changes in Emax and contractile protein expression are tightly correlated.

In contrast, laminin markedly increased sm-myosin and calponin expression, whereas it did not affect sm-�-actin protein levels. This might be explained, however, by the fact that sm-myosin and calponin are considered to be a more specific markers for mature contractile ASM cells as compared to sm-�-actin, which is a more general marker for lung cells of mesenchymal origin (10). These results suggest that an increase in contractility in the presence of laminin can be envisaged. Indeed, a tendency towards an increased contractility in response to both methacholine and KCl was observed. Moreover, a direct relationship between contractility and contractile protein expression in the presence of different matrix proteins was confirmed by the significant correlation between these two parameters for all contractile proteins.

In human ASM cells, it has been shown by immunocytochemical detection that laminin, by itself, did not affect expression of contractile proteins at all, but normalized the reduction induced by PDGF (12). In the present study, laminin did not completely reverse the effects of PDGF on contractile protein expression, but showed a tendency to attenuate the growth factor-induced suppression to some extent. This may indicate that other factors are involved in the reversal of PDGF-induced hypocontractility. The apparent discrepancy between our findings and those by Hirst et al (12) could possibly be explained by differences in experimental approach. Thus, we determined protein expression in smooth muscle strips, not cultured cells. In addition, species differences cannot be ruled out. Also, homogeneity of contractile protein expression throughout the tissue might possibly represent another variable. It has been established that there is a strong inverse relationship between the effects of peptide growth factors on maximal methacholine- and KCl-induced contraction of BTSM-strips and the proliferative response of BTSM cells to these growth factors (8). In the present study we found a strong correlation between the degree of change of Emax, both with methacholine and KCl, induced by the applied ECM proteins in BTSM strips and the proliferative response by BTSM cells cultured on these proteins. Although determined using different experimental parameters measured in different (ASM tissue and cellular) conditions, this correlation was very striking and highly reminiscent of our

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previous findings (8). Together with the relationship between contractile protein expression and contractility found for the applied ECM proteins, these results indicate that ECM proteins are importantly involved in the regulation of ASM phenotype and function.

In conclusion, our results indicate that ECM proteins differentially regulate BTSM phenotype and function. Fibronectin and collagen type I induce a (functional) hypocontractile phenotype, associated with an increased proliferative response of BTSM cells, whereas laminin inhibits growth factor-induced proliferation and supports a more contractile phenotype. These findings implicate a critical role of ECM changes (1; 15; 16; 24; 25) in altered ASM function in asthma. Acknowledgements The authors would like to thank Dirk Jan Moes, Hoeke A. Baarsma and Anita I. R. Spanjer for their expert technical assistance. Grants This study was financially supported by the Netherlands Asthma Foundation (NAF grant 03.36). References 1. Altraja A, Laitinen A, Virtanen I, Kampe M, Simonsson BG, Karlsson SE,

Hakansson L, Venge P, Sillastu H and Laitinen LA. Expression of laminins in the airways in various types of asthmatic patients: a morphometric study. Am J Respir Cell Mol Biol 15: 482-488, 1996.

2. Belkin VM, Belkin AM and Koteliansky VE. Human smooth muscle VLA-1 integrin: purification, substrate specificity, localization in aorta, and expression during development. J Cell Biol 111: 2159-2170, 1990.

3. Bonacci JV, Harris T and Stewart AG. Impact of extracellular matrix and strain on proliferation of bovine airway smooth muscle. Clin Exp Pharmacol Physiol 30: 324-328, 2003.

4. Bousquet J, Jeffery PK, Busse WW, Johnson M and Vignola AM. Asthma. From bronchoconstriction to airways inflammation and remodeling. Am J Respir Crit Care Med 161: 1720-1745, 2000.

5. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248-254, 1976.

6. Ebina M, Takahashi T, Chiba T and Motomiya M. Cellular hypertrophy and hyperplasia of airway smooth muscles underlying bronchial asthma. A 3-D morphometric study. Am Rev Respir Dis 148: 720-726, 1993.


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7. Freyer AM, Johnson SR and Hall IP. Effects of growth factors and extracellular matrix on survival of human airway smooth muscle cells. Am J Respir Cell Mol Biol 25: 569-576, 2001.

8. Gosens R, Meurs H, Bromhaar MM, McKay S, Nelemans SA and Zaagsma J. Functional characterization of serum- and growth factor-induced phenotypic changes in intact bovine tracheal smooth muscle. Br J Pharmacol 137: 459-466, 2002.

9. Gosens R, Schaafsma D, Meurs H, Zaagsma J and Nelemans SA. Role of Rho-kinase in maintaining airway smooth muscle contractile phenotype. Eur J Pharmacol 483: 71-78, 2004.

10. Halayko AJ, Salari H, Ma X and Stephens NL. Markers of airway smooth muscle cell phenotype. Am J Physiol 270: L1040-L1051, 1996.

11. Hedin U, Bottger BA, Forsberg E, Johansson S and Thyberg J. Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J Cell Biol 107: 307-319, 1988.

12. Hirst SJ, Twort CH and Lee TH. Differential effects of extracellular matrix proteins on human airway smooth muscle cell proliferation and phenotype. Am J Respir Cell Mol Biol 23: 335-344, 2000.

13. Hirst SJ, Walker TR and Chilvers ER. Phenotypic diversity and molecular mechanisms of airway smooth muscle proliferation in asthma. Eur Respir J 16: 159-177, 2000.

14. Johnson PR. Role of human airway smooth muscle in altered extracellular matrix production in asthma. Clin Exp Pharmacol Physiol 28: 233-236, 2001.

15. Laitinen A, Altraja A, Kampe M, Linden M, Virtanen I and Laitinen LA. Tenascin is increased in airway basement membrane of asthmatics and decreased by an inhaled steroid. Am J Respir Crit Care Med 156: 951-958, 1997.

16. Laitinen LA and Laitinen A. Inhaled corticosteroid treatment and extracellular matrix in the airways in asthma. Int Arch Allergy Immunol 107: 215-216, 1995.

17. Lassar AB, Skapek SX and Novitch B. Regulatory mechanisms that coordinate skeletal muscle differentiation and cell cycle withdrawal. Curr Opin Cell Biol 6: 788-794, 1994.

18. Ma X, Wang Y and Stephens NL. Serum deprivation induces a unique hypercontractile phenotype of cultured smooth muscle cells. Am J Physiol 274: C1206-C1214, 1998.

19. MacLellan WR and Schneider MD. Genetic dissection of cardiac growth control pathways. Annu Rev Physiol 62: 289-319, 2000.

20. Moiseeva EP. Adhesion receptors of vascular smooth muscle cells and their functions. Cardiovasc Res 52: 372-386, 2001.

21. Morla AO and Mogford JE. Control of smooth muscle cell proliferation and phenotype by integrin signaling through focal adhesion kinase. Biochem Biophys Res Commun 272: 298-302, 2000.

22. Nguyen TT, Ward JP and Hirst SJ. beta1-Integrins mediate enhancement of airway smooth muscle proliferation by collagen and fibronectin. Am J Respir Crit Care Med 171: 217-223, 2005.

23. Peng Q, Lai D, Nguyen TT, Chan V, Matsuda T and Hirst SJ. Multiple beta 1 integrins mediate enhancement of human airway smooth muscle cytokine secretion by fibronectin and type I collagen. J Immunol 174: 2258-2264, 2005.

24. Roche WR. Fibroblasts and asthma. Clin Exp Allergy 21: 545-548, 1991.

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25. Roche WR, Beasley R, Williams JH and Holgate ST. Subepithelial fibrosis in the bronchi of asthmatics. Lancet 1: 520-524, 1989.

26. Stephens NL, Li W, Wang Y and Ma X. The contractile apparatus of airway smooth muscle. Biophysics and biochemistry. Am J Respir Crit Care Med 158: S80-S94, 1998.

27. Timpl R, Rohde H, Robey PG, Rennard SI, Foidart JM and Martin GR. Laminin--a glycoprotein from basement membranes. J Biol Chem 254: 9933-9937, 1979.

28. Wong JZ, Woodcock-Mitchell J, Mitchell J, Rippetoe P, White S, Absher M, Baldor L, Evans J, McHugh KM and Low RB. Smooth muscle actin and myosin expression in cultured airway smooth muscle cells. Am J Physiol 274: L786-L792, 1998.

29. Yamamoto M, Yamamoto K and Noumura T. Type I collagen promotes modulation of cultured rabbit arterial smooth muscle cells from a contractile to a synthetic phenotype. Exp Cell Res 204: 121-129, 1993.

30. Yao CC, Breuss J, Pytela R and Kramer RH. Functional expression of the alpha 7 integrin receptor in differentiated smooth muscle cells. J Cell Sci 110 ( Pt 13): 1477-1487, 1997.

Bart G.J. Dekkers Johan Zaagsma Herman Meurs

Submitted (2010)

Functional consequences of human airway smooth muscle

phenotype plasticity

4C

hapter

Chapter 5

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Abstract Airway smooth muscle (ASM) phenotype plasticity, characterized by reversible switching between contractile and proliferative phenotypes, is thought to contribute to increased ASM mass and airway hyperresponsiveness in asthma. In addition, increased expression of the extracellular matrix protein collagen I has been observed within the ASM bundle of asthmatics. Previously, we have shown that exposure of intact bovine tracheal smooth muscle to collagen I induces a switch from a contractile to a hypocontractile, proliferative phenotype. The functional relevance of this finding for intact human ASM, however, has not yet been established. In the present study, we demonstrated that prolonged exposure of human tracheal smooth muscle (HTSM) strips to collagen I decreased maximal methacholine- and KCl-induced contractions as well as expression of contractile proteins. Conversely, culturing on collagen I increased proliferation of HTSM cells. Similar effects were observed for the growth factor PDGF. As observed for BTSM, culturing of HTSM cells on collagen I additively increased PDGF-induced proliferative responses, whereas no additional effects were observed on contractility or contractile protein expression. These findings indicate that collagen I and PDGF induce a functionally hypocontractile, proliferative phenotype of human ASM, which may contribute to airway remodelling in asthma. Introduction Structural changes in the airway wall, including increased airway smooth muscle (ASM) mass, are characteristic features of airway remodelling, which may contribute to airway hyperresponsiveness (AHR) and decline in lung function in asthma [1,2]. Increased ASM mass is at least partially caused by hyperplasia, and in vitro ASM proliferation is increased by serum and various growth factors that may synergize with neurotransmitters and inflammatory mediators [3-5]. Exposure of cultured ASM cells to mitogenic stimuli causes switching from a contractile, hypoproliferative to a proliferative, hypocontractile phenotype, as indicated by changes in molecular phenotypic markers [6,7]. The potential impact of these molecular changes on ASM contractile function has recently been demonstrated in intact bovine tracheal smooth muscle (BTSM). Thus, prolonged exposure of BTSM strips to serum or peptide growth factors like platelet-derived growth factor (PDGF) and insulin-like growth factor-1 decreased maximal methacholine- and KCl-induced contractions, which was inversely correlated with the proliferative responses of BTSM cells to these mitogenic stimuli [8](Chapter 3). ASM phenotype switching is reversible, as indicated by the observation that removal of mitogenic stimuli, for example by serum-deprivation, results in the reintroduction of a (hyper)contractile ASM phenotype associated with increased expression of contractile marker proteins like sm-�-

Functional human ASM phenotype plasticity

75

actin, sm-MHC and calponin, which is further enhanced in the presence of insulin or transforming growth factor-� (TGF-�) [9-11](Chapter 5).

Increased deposition of extracellular matrix (ECM) proteins within the airway wall is another hallmark of airway remodelling in asthma [12](Chapter 2) Expression of various ECM proteins, including collagen I, is increased in the subepithelial basement membrane [12,13]. Within the ASM bundle, increased deposition of collagen I has been observed as well [14-16], which may be due to increased production of this matrix protein by asthmatic ASM cells [17]. In addition, increased proliferative responses of asthmatic ASM cells have also been shown, which depends on the ECM produced by these cells [17], indicating that collagen I may have impact on the phenotype of these cells. Indeed, in vitro studies on the effects of collagen I on human ASM cell function have indicated that cells grown on this ECM protein show increased growth factor-induced proliferation, increased synthetic capabilities, decreased apoptosis and reduced expression of contractile proteins [7,18-20]. Using BTSM, we have recently shown that exposure of intact strip preparations to monomeric collagen I induced a hypocontractile state, characterized by decreased contractile responses to methacholine and KCl and reduced expression of contractile marker proteins, which was associated with increased proliferation of cultured BTSM cells (Chapter 3). In addition, PDGF augmented collagen I-induced proliferation in an additive fashion, without an additional effect on contractility or contractile protein expression (Chapter 3). The functional significance of ECM- and growth factor-induced phenotype switching on intact human ASM is currently unknown. Therefore, in the present study we investigated the effects of monomeric collagen I and PDGF on human tracheal smooth muscle (HTSM) strip contractility, contractile protein expression and cell proliferation. Materials and methods Tissue preparation and organ culture Human tracheal sections from anonymized lung transplantation donors were obtained from the Department of Cardiothoracic Surgery, University Medical Centre Groningen and transported to the laboratory in ice-cold Krebs-Henseleit (KH) buffer (composition in mM: NaCl 117.5, KCl 5.60, MgSO4 1.18, CaCl2 2.50, NaH2PO4 1.28, NaHCO3 25.00, and glucose 5.50), pregassed with 5% CO2 and 95% O2 (pH 7.4). HTSM strips were prepared as described for BTSM (Chapter 3 & 5). After dissection of the smooth muscle layer and careful removal of connective tissue and mucosa, strips were cut and cultured for 4 days in Medium Zero (sterile Dulbecco's Modified Eagle's medium (DMEM; Gibco BRL Life Technologies Paisley, UK), supplemented with sodium pyruvate (1 mM; Gibco), nonessential amino acid mixture (1:100; Gibco), gentamicin (45 μg/ml; Gibco), penicillin (100 U/ml; Gibco), streptomycin (100 μg/ml; Gibco), amphotericin B (1.5 μg/ml; Gibco), apo-transferrin (5 μg/ml, human; Sigma, St.

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Louis, MO, USA) and ascorbic acid (0.1 mM)). When used, monomeric collagen I (50 �g/ml, calf skin; Fluka, Buchs, Switzerland) and/or PDGF-AB (10 ng/ml, human; Bachem, Weil am Rhein, Germany) were present during the entire incubation period. Occasionally, strips were used for isometric tension measurements directly after preparation. Isometric tension measurements. Isometric contraction experiments were performed as described previously (Chapter 3 & 5). Briefly, HTSM strips were mounted for isometric recording in organ baths, containing KH buffer at 37°C. During a 90-min equilibration period with washouts every 30 min, resting tension was adjusted to 0.5 g, followed by precontractions with 20 and 40 mM KCl. Following washout, maximal relaxation was established by the addition of (-)-isoproterenol (0.1 �M; Sigma). Tension was readjusted to 0.5 g immediately followed by two changes with KH buffer. After another equilibration period of 30 min, cumulative concentration-response curves were constructed to KCl (5.6-50 mM) or methacholine (1 nM – 0.1 mM; ICN Biomedicals, Costa Mesa, CA, USA). When maximal tension was reached, the strips were washed several times and maximal relaxation was established by using (-)-isoproterenol (10 �M). Contractions were expressed as the percentage of maximal contraction induced by KCl or methacholine in vehicle-treated strips. After the experiment, strips were snap frozen for Western analysis. Western analysis HTSM strip homogenization and Western analysis of sm-�-actin and sm-MHC expression were preformed as described (Chapter 3 & 5). In short, homogenates were prepared by pulverizing the strips under liquid nitrogen, followed by sonification in homogenization buffer (composition in mM: Tris-HCl 50 mM, NaCl 150.0, EDTA 1.0, PMSF 1.0, Na3VO4 1.0, NaF 1.0, pH 7.4, supplemented with leupeptin 10 �g/ml, aprotinin 10 �g/ml, pepstatin 10 �g/ml, Na-deoxycholate 0.25% and Igepal 1%; all Sigma). Equal amounts of protein were subjected to sodium dodecyl sulfate/polycrylamide gel electrophoresis and transferred onto nitrocellulose membranes, followed by standard immunoblotting techniques. Antibodies (anti-sm-�-actin (Sigma) and anti-sm-MHC (Neomarkers, Fremont, CA, USA) were visualized on film using enhanced chemiluminescence reagents (Pierce, Breda, NL) and analyzed by densitometry (TotallabTM, Nonlinear dynamics, Newcastle, UK). Bands were normalized to �-actin expression. HTSM cell culture HTSM, prepared free of mucosa and connective tissue, was chopped using a McIlwain tissue chopper. Tissue slices were washed once with Medium Plus (DMEM, supplemented with sodium pyruvate (1 mM), nonessential amino acid mixture (1:100), gentamicin (45 �g/ml), penicillin (100 U/ml), streptomycin (100 �g/ml), amphotericin B (1.5 �g/ml) and fetal bovine serum (FBS, 10%; Gibco)), placed in culture flasks and allowed to adhere. Medium was refreshed every 48-


77

72 h. Upon reaching confluency, cells were passaged by trypsinization. Cells from passages 1-5 were used for the present study. Coating of culture plates with collagen I Collagen I was reconstituted in hydrochloric acid (10 mM) at 5 mg/ml and diluted in PBS to a final concentration of 50 �g/ml. Diluted collagen I (0.5 ml) was adsorbed to culture plates overnight and air-dried at room temperature. Unoccupied protein-binding sites were blocked by 30 min incubation with 0.1% bovine serum albumin solution. Subsequently, plates were washed twice with DMEM and dried before further use. [3H]-thymidine-incorporation [3H]-Thymidine-incorporation was performed as described previously (Chapter 3 & 5). HTSM cells were plated on uncoated or collagen I-coated 24-well culture plates at a density of 30,000 cells/well and allowed to attach overnight in Medium Plus. Cells were washed with PBS and made quiescent by incubation in Medium Zero, supplemented with 1% ITS (Insulin, Transferrin and Selenium; Gibco) for 72 h. Subsequently, cells were washed and incubated in the absence or presence of PDGF (10 ng/ml) in Medium Zero for 28 h, the last 24 h in the presence of [methyl-3H]-thymidine (0.25 μCi/ml). After incubation, the cells were washed with PBS at room temperature. Subsequently, the cells were treated with ice-cold 5% trichloroacetic acid on ice for 30 min, and the acid-insoluble fraction was dissolved in NaOH (1 M). Incorporated [3H]-thymidine was quantified by liquid-scintillation counting using a Beckman LS1701 �-counter. Data analysis Data represent means ± SEM from n separate experiments. Statistical significance of differences was evaluated by paired Student's t-test. Statistical significance was reached at P<0.05.

Chapter 5

78

[Methacholine] (- log M)

3456789

Con

tract

ion

(%)

0

25

50

75

100

125Control PDGFCollagen I + PDGF

[KCl] (mM)

0 10 20 30 40 50

0

25

50

75

100

125Control PDGFCollagen I + PDGF

A B

Results Collagen I and PDGF decrease HTSM strip contractility To assess whether exposure to collagen I or PDGF induces a functionally hypocontractile ASM phenotype, HTSM strips were cultured in the absence and presence of monomeric collagen I (50 �g/ml) and/or PDGF (10 ng/ml) for 4 days. Culturing HTSM strips in the presence of collagen I or PDGF for 4 days significantly (P<0.05) reduced maximal methacholine- and KCl-induced contractile force (Emax) compared to vehicle-treated control strips (Figure 1, Table 1). Figure 1: Concentration-response curves of (A) methacholine- and (B) KCl-induced contractions of HTSM strips, pretreated with vehicle (control) or collagen I (50 �g/ml) in the absence or presence of PDGF (10 ng/ml) for 4 days. Data represent means ± SEM of 3-7 experiments performed in duplicate. No additive effects were observed after combined treatment. The sensitivity to both contractile stimuli was unaffected by all treatments. No significant change in methacholine-induced contractility was observed after 4 days of culturing in vehicle compared to freshly isolated strips (Emax=2.89±0.68 g and 2.50±0.32 g, -logEC50=5.95±0.10 and 5.77±0.14, respectively, n=4-7, P>0.05 Student’s t-test for unpaired observations).


79

A B

Basal Collagen I

sm-M

HC

abu

ndan

ce

(% o

f bas

al)

0

25

50

75

100

125BasalPDGF

**

*

**

��actin ��actin

sm-MHC sm-�-actin

Basal Collagen I

sm-�

-act

in a

bund

ance

(%

of b

asal

)

0

25

50

75

100

125

***

*

- +- +- +- +- + - + - + - +

Collagen IPDGF

Collagen IPDGF

Table 1: Contractile responses of HTSM strips to methacholine or KCl after 4 days of culturing in the absence or presence of collagen I (50 �g/ml) and/or PDGF (10 ng/ml).

Methacholine KCl Emax (%) pEC50 (-log M) Emax (%) EC50 (mM) Control 100±0 5.77±0.14 100±0 34.6±0.8 + PDGF 64.8±5.3** 5.66±0.14 61.8±5.2* 34.7±1.1 Collagen I 61.0±6.0** 5.99±0.18 56.9±9.0*** 35.2±1.4 + PDGF 56.7±4.8* 5.69±0.29 60.6±8.6*** 35.6±1.2

Data represent means ± SEM of 3-7 experiments, performed in duplicate. Abbreviations: Emax: maximal contraction; EC50: contraction of agonist eliciting half-maximal response; pEC50: negative logarithm of the EC50 value. *P<0.05, **P<0.01, ***P<0.001 compared to control. In agreement with the reduced contractility, collagen I and PDGF also decreased the protein expression of sm-�-actin and sm-myosin heavy chain (P<0.05 all, Figure 2). No additive effects were observed after combined treatment. Figure 2: Western analysis of (A) sm-MHC and (B) sm-�-actin expression in HTSM strips treated with vehicle (control) or collagen I (50 �g/ml) in the absence (basal) or presence of PDGF (10 ng/ml) for 4 days. Means ± SEM of 3-5 experiments are shown. *P<0.05, **P<0.01 compared to basal control. Representative immunoblots of sm-MHC, sm-�-actin and �-actin are shown.

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Control Collagen I

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Collagen I and PDGF increase proliferation of HTSM cells To assess whether the collagen I- and PDGF-induced decreases in Emax were associated with proliferative changes, [3H]-thymidine-incorporation was assessed using primary HTSM cell cultures. Proliferation of these cells was increased by PDGF (P<0.05) as well as by culturing on a collagen I matrix (P<0.05, Figure 3). PDGF-induced DNA synthesis was significantly enhanced, when the cells were cultured on collagen I-coated instead of uncoated matrices (P<0.05). Discussion In the present study, we demonstrated that prolonged exposure of intact HTSM tissue to the ECM protein collagen I and to the peptide growth factor PDGF induces a functionally hypocontractile, proliferative phenotype. Thus, pretreatment of HTSM strips with collagen I and with PDGF decreased maximal contractions in response to the receptor-dependent agonist methacholine and the receptor-independent stimulus KCl associated with decreased contractile protein expression, whereas both stimuli increased proliferation of cultured HTSM cells. Culturing the HTSM cells on collagen I additively increased the proliferative response to PDGF, whereas no additional effects were observed on contractility or contractile protein expression, suggesting differential regulation of these processes by the combined treatment. Our findings are well in line with previous observations that exposure of human bronchiole ring segments to serum – containing pro-proliferative factors – reduced carbachol-, histamine- and KCl-induced contractions and decreased expression of the contractile protein calponin [21].

Figure 3: Basal and PDGF (10 ng/ml)-stimulated DNA synthesis of HTSM cells cultured on uncoated plastic (control) or collagen I (50 �g/ml)-coated matrices. Data represent means ± SEM of 4 experiments performed in triplicate. *P<0.05 compared to basal control. #P<0.05 compared to PDGF, control.


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ASM cells display phenotype plasticity, characterized by reversible and dynamic changes in the expression of contractile and proliferative markers that may be governed by a variety of growth factors and ECM proteins present in the asthmatic airway wall, including collagen I and PDGF [5,6]. The peptide growth factor PDGF is a well-characterized proliferative stimulus for ASM, which may be released from inflammatory and structural cells within the airway, including eosinophils, macrophages and fibroblasts [22-24]. Exposure of human ASM cells to PDGF increases proliferation and the expression of the proliferative marker Ki67, whereas the expression of the contractile markers sm-�-actin and sm-MHC is reduced [7]. In the present study, we demonstrated the functional relevance of these findings in intact HTSM tissue in which cell-to-cell contacts and endogenous ECM components are preserved, the results being highly reminiscent of previous findings in BTSM [8](Chapter 3).

Collagens are widespread throughout the body, provide structural support and fulfil a variety of biological functions [25]. In vitro, it has been found that culturing of human ASM cells on monomeric collagen I augmented growth factor-induced proliferation and enhanced the reduction in contractile marker expression by PDGF [7]. In the airways of asthmatics, deposition of collagen I is increased in the extracellular microenvironment of the ASM cells [12,14]. In addition, ASM cells from asthmatic patients produce more collagen I than to those obtained from healthy subjects, which, via an autocrine mechanism, could contribute to the increased proliferation of asthmatic ASM cells [17].

Interestingly, culturing of ASM cells on fibrillar collagen I instead of monomeric collagen I did not promote growth factor-induced ASM proliferation [18], whereas recently fibrillar collagen I has even been shown to inhibit both basal and growth factor-induced proliferation [26]. In addition, inhibition of collagen degradation by the MMP inhibitor ilomastat further enhanced the growth-attenuating effects of fibrillar collagen I, indicating that degradation of collagen to its monomeric isoform may enhance ASM proliferation [26]. In BTSM cells, monomeric collagen I has been shown to increase basal as well as PDGF-induced proliferation [27](Chapter 3), which was tightly correlated with decreased contractility of intact BTSM strips (Chapter 3). We now show that these findings can be translated to HTSM. Interestingly, these observations also suggest that BTSM is a representative experimental model for human ASM phenotype plasticity.

Collectively, the findings described above indicate that changes in the extracellular environment surrounding the ASM may contribute to ASM accumulation in asthma. Indeed, using a guinea pig model of allergic asthma, we have recently shown that ASM remodelling induced by repeated allergen-challenges was inhibited by the integrin-blocking peptide Arg-Gly-Asp-Ser (RGDS), containing the RGD binding motif, which also inhibits human ASM cell proliferation induced by monomeric collagen I (Chapter 6).

Increased ASM mass is considered to be a major factor contributing to airway hyperresponsiveness and decline in lung function in asthmatics [28,29].

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Our present findings demonstrating that exposure of intact HTSM preparations to collagen I and PDGF has impact on contractile function provides more insight in the functional consequences of phenotype switching. Next to increased mass, however, asthmatic ASM also shows increased expression of contractile proteins [30], suggesting an increased rather than decreased contractile function. Phenotypic plasticity, however, is a dynamic and reversible process and in patients, episodes with increased levels of growth factors, Gq-coupled neurotransmitters such as acetylcholine and inflammatory mediators - which may (synergistically) promote a proliferative, hypocontractile phenotype [6,8] - alternate with episodes of reduced levels. In vitro, the latter is mimicked by serum deprivation of cultured ASM cells, particularly in the presence of insulin or TGF-�, which redirects the hypocontractile phenotype to a (hyper)contractile state [10,11](Chapter 5). Similar processes could contribute to ASM hypercontractility and AHR in asthma.

In conclusion, our findings indicate that collagen I and PDGF induce a shift of human ASM phenotype to a hypocontractile, proliferative state, which has functional impact on the muscle and may contribute to airway remodelling in asthma. Acknowledgements This study was financially supported by the Netherlands Asthma Foundation (Grant NAF 3.2.03.36). We are grateful to the department of Cardiothoracic Surgery of the University Medical Centre Groningen for providing human tracheal sections. The authors wish to thank Carolina Elzinga for expert technical assistance. References 1. Meurs H, Gosens R, Zaagsma J. Airway hyperresponsiveness in asthma: lessons

from in vitro model systems and animal models. Eur Respir J 2008; 32: 487-502. 2. Bousquet J, Jeffery PK, Busse WW, Johnson M, Vignola AM. Asthma. From

bronchoconstriction to airways inflammation and remodeling. Am J Respir Crit Care Med 2000; 161: 1720-1745.




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5. Gosens R, Roscioni SS, Dekkers BG, Pera T, Schmidt M, Schaafsma D, Zaagsma J, Meurs H. Pharmacology of airway smooth muscle proliferation. Eur J Pharmacol 2008; 585: 385-397.




9. Ma X, Wang Y, Stephens NL. Serum deprivation induces a unique hypercontractile phenotype of cultured smooth muscle cells. Am J Physiol 1998; 274: C1206-C1214.


11. Hirota JA, Nguyen TT, Schaafsma D, Sharma P, Tran T. Airway smooth muscle in asthma: Phenotype plasticity and function. Pulm Pharmacol Ther 2009; 22: 370-378.

12. Fernandes DJ, Bonacci JV, Stewart AG. Extracellular matrix, integrins, and mesenchymal cell function in the airways. Curr Drug Targets 2006; 7: 567-577.

13. Roche WR, Beasley R, Williams JH, Holgate ST. Subepithelial fibrosis in the bronchi of asthmatics. Lancet 1989; 1: 520-524.







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20. Peng Q, Lai D, Nguyen TT, Chan V, Matsuda T, Hirst SJ. Multiple beta 1 integrins mediate enhancement of human airway smooth muscle cytokine secretion by fibronectin and type I collagen. J Immunol 2005; 174: 2258-2264.

21. Moir LM, Ward JP, Hirst SJ. Contractility and phenotype of human bronchiole smooth muscle after prolonged fetal bovine serum exposure. Exp Lung Res 2003; 29: 339-359.

22. Ohno I, Nitta Y, Yamauchi K, Hoshi H, Honma M, Woolley K, O'Byrne P, Dolovich J, Jordana M, Tamura G, . Eosinophils as a potential source of platelet-derived growth factor B-chain (PDGF-B) in nasal polyposis and bronchial asthma. Am J Respir Cell Mol Biol 1995; 13: 639-647.

23. Taylor IK, Sorooshian M, Wangoo A, Haynes AR, Kotecha S, Mitchell DM, Shaw RJ. Platelet-derived growth factor-beta mRNA in human alveolar macrophages in vivo in asthma. Eur Respir J 1994; 7: 1966-1972.

24. Fabisiak JP, Absher M, Evans JN, Kelley J. Spontaneous production of PDGF A-chain homodimer by rat lung fibroblasts in vitro. Am J Physiol 1992; 263: L185-L193.

25. Prockop DJ, Kivirikko KI. Collagens: molecular biology, diseases, and potentials for therapy. Annu Rev Biochem 1995; 64: 403-434.

26. Schuliga M, Ong SC, Soon L, Zal F, Harris T, Stewart AG. Airway smooth muscle remodels pericellular collagen fibrils: implications for proliferation. Am J Physiol Lung Cell Mol Physiol 2010.




30. Leguillette R, Laviolette M, Bergeron C, Zitouni NB, Kogut P, Solway J, Kashmar L, Hamid Q, Lauzon AM. Myosin, Transgelin, and Myosin Light Chain Kinase: Expression and Function in Asthma. Am J Respir Crit Care Med 2008; 179: 194-204.

Bart G.J. Dekkers Dedmer Schaafsma Thai Tran Johan Zaagsma Herman Meurs

Am J Respir Cell Mol Biol (2009) 41:494-504

Insulin-induced laminin expression promotes a hypercontractile airway

smooth muscle phenotype

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Abstract Airway smooth muscle (ASM) plays a key role in the development of airway hyperresponsiveness and remodeling in asthma, which may involve maturation of ASM cells to a hypercontractile phenotype. In vitro studies have indicated that long-term exposure of bovine tracheal smooth muscle (BTSM) to insulin induces a functional hypercontractile, hypoproliferative phenotype. Similarly, the extracellular matrix protein laminin has been found to be involved in both the induction and maintenance of a contractile ASM phenotype. Using BTSM, we now investigated the role of laminins in the insulin-induced hypercontractile, hypoproliferative ASM phenotype. The results demonstrate that insulin-induced hypercontractility after 8 days of tissue culture was fully prevented by combined treatment of BTSM-strips with the laminin competing peptides Tyr-Ile-Gly-Ser-Arg (YIGSR) and Arg-Gly-Asp-Ser (RGDS). YIGSR also prevented insulin-induced increases in sm-myosin expression and abrogated the suppressive effects of prolonged insulin treatment on PDGF-induced DNA-synthesis in cultured cells. In addition, insulin time-dependently increased laminin �2, �1 and �1 chain protein, but not mRNA abundance in BTSM strips. Moreover, as previously found for contractile protein accumulation, signaling through PI3-kinase and Rho kinase dependent pathways was required for the insulin-induced increase in laminin abundance and contractility. Collectively, our results indicate a critical role for �1-containing laminins, likely laminin-211, in the induction of a hypercontractile, hypoproliferative ASM phenotype by prolonged insulin exposure. Increased laminin production by ASM could be involved in the increased ASM contractility and contractile protein expression in asthma. Moreover, the results may be of interest for the use of inhaled insulin administrations by diabetics. Introduction Remodeling of the airway wall is a feature of chronic airway inflammation and may importantly contribute to airway hyperresponsiveness and decline of lung function, as observed in patients with severe and persistent asthma (1-3). Airway wall remodeling in these patients is, amongst others characterized by increased airway smooth muscle (ASM) mass (4) and changes in extracellular matrix (ECM) (5, 6).

Increased ASM mass may result from both cellular hyperplasia and hypertrophy of individual ASM cells (4). In keeping with hyperplasia, ASM cells have been found to retain the ability to alter their phenotype in response to a broad range of environmental cues, varying from cell-cell contacts and mechanical strain to growth factors and ECM components (7). Modulation to a proliferative phenotype results from exposure of ASM cells to mitogenic stimuli leading to increased proliferative, synthetic and migratory capabilities and

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decreased contractile function (Chapter 3)(8-10). In asthmatic airways, accumulation of ASM cells with increased contractile properties and high expression levels of contractile proteins, but with low synthetic and proliferative capabilities has been described (11-14). In culture, induction of a (hyper)contractile ASM phenotype occurs as a consequence of growth arrest by serum deprivation, leading to augmented abundance of contractile proteins and contraction regulatory proteins (15, 16). Recently it has become apparent that insulin promotes ASM maturation to a greater extent than can be attained in its absence during serum deprivation. Long-term treatment (8 days) with insulin has been demonstrated to increase expression of specific contractile phenotype markers in bovine tracheal smooth muscle (BTSM) cells and strips (17), which is accompanied by decreased mitogenic responses and induction of a functionally hypercontractile phenotype (18). In addition, Rho kinase- and PI3-kinase-dependent signaling pathways were found to mediate insulin-induced ASM cell maturation (17).

Laminins are ECM proteins commonly found in basement membranes (19). Currently, five laminin �-, four �- and three �-chains, forming at least 15 different laminin isoforms, have been identified in mammals (20). Laminin-111 (laminin-1) and laminin-211 (laminin-2) are produced in murine developing lung and found to be importantly involved in pulmonary branching and differentiation of naïve mesenchymal cells (20-22). Compared to healthy controls, increased expression of laminin �2 and �2 chains has been observed in the airways of asthmatics (23). Moreover, studies on the role of laminins in ASM function revealed that ASM cells, grown on an exogenously applied laminin-111 matrix, are retained in a hypoproliferative phenotype associated with increased contractile protein abundance (24). This inhibition of mitogenic responsiveness has been found of functional relevance, as recently demonstrated in this laboratory by the observation that the induction of a hypocontractile phenotype in BTSM strips during prolonged treatment with platelet-derived growth factor (PDGF) was prevented by combined treatment with exogenous laminin-111 (Chapter 3). A role for endogenously expressed laminin-211 in regulating ASM phenotype was recently suggested by Tran and colleagues, who found that increased expression of laminin �2, �1 and �1 chains is required for maturation of human ASM cells under serum-deprived and insulin-supplemented conditions (25).

To elucidate whether laminins are functionally involved in the induction of a hypercontractile, hypoproliferative ASM phenotype, we now investigated the effects of insulin incubation in the absence and presence of the laminin competing peptides Tyr-Ile-Gly-Ser-Arg (YIGSR) and Arg-Gly-Asp-Ser (RGDS) on BTSM strip contractility. In addition, we assessed the effects of YIGSR on the insulin-induced sm-myosin accumulation in these strips and on the inhibitory effects of insulin stimulation on proliferative responses of isolated primary BTSM cells. Western analysis was carried out to study the effects of insulin stimulation on laminin �1, �2, �1 and �1 chain abundance in BTSM strip homogenates.

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Since PI3-kinase and Rho kinase have been associated with insulin-induced ASM maturation (17), the effects of the selective PI3-kinase inhibitor LY-294002 and the selective Rho kinase inhibitor Y-27632 on laminin-211 chain protein and mRNA abundance were investigated as well. Moreover, the roles of PI3-kinase and Rho kinase signaling in the induction of a functional hypercontractile phenotype by insulin were assessed. Our results indicate that the insulin-induced hypercontractile ASM phenotype is functionally dependent on signaling via �- and �1-containing laminins and that insulin increases laminin �2, �1 and �1 chain protein abundance, via PI3-kinase- and Rho kinase-dependent signaling pathways. Accordingly, inhibition of both PI3-kinase and Rho kinase signaling normalized the insulin-induced hypercontractility. Increased laminin production by ASM could be involved in the increased ASM contractility and contractile protein expression as observed in asthma (13, 23). The results may also be of interest for the ongoing discussion on the treatment of diabetes mellitus type 1 and 2 by inhaled insulin administrations (26, 27). Materials and methods Tissue preparation and organ-culture procedure.Bovine tracheae were obtained from local slaughterhouses and rapidly transported to the laboratory in ice-cold Krebs-Henseleit (KH) buffer of the following composition (mM): NaCl 117.5, KCl 5.60, MgSO4 1.18, CaCl2 2.50, NaH2PO4 1.28, NaHCO3 25.00 and glucose 5.50, pregassed with 5% CO2 and 95% O2; pH 7.4. After dissection of the smooth muscle layer and careful removal of mucosa and connective tissue, tracheal smooth muscle strips were prepared while incubated in gassed KH-buffer at room temperature. Care was taken to cut tissue strips of macroscopically identical length (1 cm) and width (2 mm). Tissue strips were washed once in Medium Zero (sterile DMEM, supplemented with sodium pyruvate (1 mM), non-essential amino-acid mixture (1 :100), gentamicin (45 �g/ml), penicillin (100 U/ml), streptomycin (100 �g/ml), amphotericin B (1.5 �g/ml), apo-transferrin (5 �g/ml, human) and ascorbic acid (0.1 mM)). Next, tissue strips were transferred into suspension culture flasks and a volume of 7.5 ml Medium Zero was added per tissue strip. Strips were maintained in culture in an Innova 4000 incubator shaker (37°C, 55 rpm), under tightly controlled conditions, for 2, 4 or 8 days, refreshing the medium on day 4. When used, insulin (1 �M), Tyr-Ile-Gly-Ser-Arg (YIGSR, 100 �M), Arg-Gly-Asp-Ser (RGDS, 100 �M), Gly-Arg-Ala-Asp-Ser-Pro (GRADSP, 100 �M), Y-27632 (1 �M) or LY294002 (10 �M) were present during the entire incubation period. Effective concentrations of the antagonists were as per previous reports (Chapter 3)(17, 18). For determination of laminin-chain mRNA or protein expression some strips were flash frozen in liquid nitrogen directly after preparation.


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Isometric tension measurements. Tissue strips, collected from the suspension culture flasks, were washed with several volumes of KH-buffer pregassed with 5% CO2 and 95% O2, pH 7.4 at 37°C. Subsequently, strips were mounted for isometric recording (Grass force-displacement transducer FT03) in 20 ml water-jacked organ baths, containing KH-buffer at 37°C, continuously gassed with 5% CO2 and 95% O2, pH 7.4. During a 90-min equilibration period, with washouts every 30 min, resting tension was gradually adjusted to 3 g. Subsequently, muscle strips were precontracted with 20 and 40 mM isotonic KCl solutions. Following two washouts, maximal relaxation was established by the addition of 0.1 �M (-)-isoproterenol. In > 95 % of the experiments no basal myogenic tone was detected. Tension was readjusted to 3 g, immediately followed by two changes with fresh KH-buffer. After another equilibration period of 30 min, cumulative concentration response curves (CRCs) were constructed to stepwise increasing concentrations of isotonic KCl (5.6�50 mM) or methacholine (1 nM � 0.1 mM). When maximal tension was reached, the strips were washed several times and maximal relaxation was established using (-)-isoproterenol (10 �M). Coating of culture plates with laminin. Mouse laminin, purified from Engelberth-Holm-Swarm (EHS) sarcoma, consisting mainly of laminin-111 (laminin-1) (28), was diluted in PBS to a final concentration of 10 �g/ml. Diluted laminin (0.5 ml) was adsorbed to 24-well cluster plates overnight and air-dried at room temperature. Unoccupied protein-binding sites were blocked by a 30 min incubation with a sterile 0.1% bovine serum albumin (BSA) solution. Subsequently, plates were washed twice with unsupplemented DMEM and dried before further use. Isolation of bovine tracheal smooth muscle (BTSM) cells. Bovine tracheal smooth muscle cells were isolated as described previously (29). In short, after the removal of mucosa and connective tissue, tracheal smooth muscle was chopped. Tissue particles were washed and enzymatic digestion was performed in Medium Plus (DMEM supplemented with sodium pyruvate (1 mM), non-essential amino-acid mixture (1 :100), gentamicin (45 �g/ml), penicillin (100 U/ml), streptomycin (100 �g/ml), amphotericin B (1.5 �g/ml) and FBS (0.5%)), supplemented with collagenase P (0.75 mg/ml), papain (1 mg/ml) and soybean trypsin inhibitor (1 mg/ml). After digestion, cells were filtered and washed three times in Medium Plus, supplemented with 10% FBS instead of 0.5% FBS. [3H]-Thymidine-incorporation. BTSM cells were plated on uncoated or laminin-coated 24-well culture plates at a density of 50,000 cells per well immediately after isolation and were allowed to attach overnight in Medium Plus, containing 10% FBS. Cells were washed twice with sterile PBS and made quiescent by incubation in Medium Zero,

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supplemented with 0.1% FBS, apo-transferrin (5 μg/ml), ascorbic acid (0.1 mM), in the absence or presence of YIGSR (100 �M), for 7 days. When the effects of insulin were studied, 0.1% FBS was replaced by insulin (1 μM). After 7 days, cells were washed twice with PBS and incubated with or without PDGF (10 ng/ml) in Medium Zero for 28 h, the last 24 h in the presence of [methyl-3H]-thymidine (0.25 μCi/ml). After incubation the cells were washed twice with 0.5 ml PBS at room temperature. Subsequently, the cells were treated with 0.5 ml ice-cold 5% trichloroacetic acid on ice for 30 min, and the acid-insoluble fraction was dissolved in 1 ml NaOH (1 M). Incorporated [3H]-thymidine was quantified by liquid-scintillation counting using a Beckman LS1701 �-counter. Western analysis. To obtain whole BTSM tissue homogenates, tissue strips were cultured as described above. Homogenates were prepared by pulverizing the tissue under liquid nitrogen, followed by sonification in homogenization buffer (composition in mM: Tris-HCl 50 mM, NaCl 150.0, EDTA 1.0, PMSF 1.0, Na3VO4 1.0, NaF 1.0, pH 7.4, supplemented with 10 �g/ml leupeptin, 10 �g/ml aprotinin, 10 �g/ml pepstatin, Na-deoxycholate 0.25 % and 1% Igepal (NP-40)). Equal amounts of protein were subjected to electrophoresis and transferred onto nitrocellulose membranes. Membranes were subsequently blocked in blocking buffer (composition: Tris-HCl 50.0 mM; NaCl 150.0 mM; Tween-20 0.1%, dried milk powder 5%) for 60 minutes at room temperature. Next, membranes were incubated overnight at 4 °C (anti-sm-myosin) or for 90 minutes at room temperature (anti-laminin-1 and anti-merosin (laminin �2)) with primary antibodies (anti-sm-myosin and anti-laminin-1 (�1, �1 and �1), both diluted 1:200, anti-merosin and anti-�-actin, diluted 1:2000, all dilutions in blocking buffer). Specificity of the rabbit anti-laminin-1 (polyclonal) antibody for �1, �1 and �1 chains was assessed using mouse EHS laminin (laminin-111) as a control (data not shown). After three washes with TBS-Tween 20 (0.1% TBST, containing 50.0 mM Tris-HCl, 150.0 mM NaCl and 0.1% Tween 20) of 10 min each, membranes were incubated with horseradish peroxidase-labelled secondary antibodies (dilution 1:2000 in blocking buffer) at room temperature for 90 min, followed by another three washes with 0.1% TBST. Antibodies were then visualized on film using enhanced chemiluminescence reagents and analyzed by densitometry (TotallabTM). All bands were normalized to �-actin expression. Real-Time PCR To obtain BTSM mRNA, tissue strips were cultured as described above and total RNA was extracted using the Qiagen RNeasy Mini Kit according to the manufacturer’s protocol. Total RNA (1 �g) was reverse transcribed using AMV transcriptase, incubated for 10 min at 25 °C followed by 45 min at 42 °C and 5 min at 99°C. cDNA was diluted 1:10 with RNase-free water. cDNA was then subjected to real time PCR, performed with an iQ5 Real-Time PCR Detection System (Biorad) and iQ5 SYBR Green supermix. Assay were performed in 25 �l


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volumes in duplicate using the primer pairs listed in Table 1. Cycle parameters were: denaturation at 94 °C for 30 s, annealing at 60 °C for 30 s and extension at 72 °C for 30 s for 40 cycles. Real time PCR data were analyzed using the comparative cycle threshold (CT = amplification cycle number) method. The amount of target gene normalized to an endogenous reference (18s rRNA, designated as CT) and relative to a calibrator (day 8, control) is given by the equation 2-CT. Table 1: Primers for laminin chains and 18S rRNA used in real time PCR.

Laminin chain

NCBI accession number Primer sequence

Laminin �2 XM_001787958. Forward 5’ GGA TCA ACC ACG CTG ATT TT 3’ Reversed 5’ ATT GAT TTT GGT GGG GAT CA 3’ Laminin �1 XM_598260 Forward 5’ ATG GTG GTT CGA GGA AAC TG 3’ Reversed 5’ TTG GTG TTA TGC CTG CAC AT 3’ Laminin �1 BC105436 Forward 5’ ATT GAG CCA TCC ACT GAA GG 3’ Reversed 5’ TAG CCG TGT CAG GTT TAC CC 3’ 18s rRNA AF176811. Forward 5’ AAA CGG CTA CCA CAT CCA AG 3’ Reversed 5’ TCG CGG AAG GAT TTA AAG TG 3’

Materials. Dulbecco’s modification of Eagle’s medium (DMEM), fetal bovine serum, sodium pyruvate solution (100 mM), non-essential amino acid mixture, gentamicin solution (10 mg/ml), penicillin/streptomycin solution (5000 U/ml; 5000 �g/ml) and amphotericin B solution (250 �g/ml, Fungizone) were obtained from Gibco BRL Life Technologies (Paisley, U.K.). Bovine serum albumin, mouse monoclonal anti-�-actin, apo-transferrin (human), soybean trypsin inhibitor, insulin (from bovine pancreas), leupeptin, aprotinin, pepstatin, Na-deoxycholate, Igepal (NP-40) and (-)-isoproterenol hydrochloride were obtained from Sigma Chemical Co. (St. Louis, MO, U.S.A.). Mouse monoclonal anti-sm-myosin and polyclonal rabbit anti-laminin-1 were from Neomarkers (Fremont, CA, U.S.A.). Mouse monoclonal anti-merosin (laminin �2 chain) was from Chemicon (Temecula, CA, U.S.A.). [methyl-3H]-Thymidine (specific activity 25 Ci/mmol) and Lumigen PS-3 detection reagent were from Amersham (Buckinghamshire, U.K.). Platelet derived growth factor (PDGF-AB) was from Bachem (Weil am Rhein, Germany). Methacholine was obtained from ICN Biomedicals (Costa Mesa, CA, U.S.A.). Tyr-Ile-Gly-Ser-Arg (YIGSR), Arg-Gly-Asp-Ser (RGDS) and Gly-Arg-Ala-Asp-Ser-Pro (GRADSP) were obtained from Calbiochem (San Diego, CA, USA). Qiagen RNeasy Mini Kit was from Qiagen (Mississauga, ON, USA). AMV transcriptase was from Promega (Madison, WI, USA). iQ5 SYBR Green supermix was from Biorad (Hertfordshire, UK). PCR primers (Table 1) and EHS laminin were from

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Invitrogen (Grand Island, NY, USA). Collagenase P and papain were from Boehringer (Mannheim, Germany). L(+)ascorbic acid was from Merck (Darmstadt, Germany). (+)-(R)-trans-4-(1-aminoethyl)-N-(4-pyridyl) cyclohexane carboxamide (Y-27632) and 2-(4-morpholinyl)-8-phenyl-4H-1-benzopyran-4-one (LY-294002) were from Tocris Bioscience (Ellisville, MO, USA). All other used chemical were of analytical grade. Data analysis. All data represent means ± SEM from n separate experiments. Statistical significance of differences was evaluated using one-way ANOVA for repeated measures, followed by a Bonferroni’s multiple comparisons test. Differences were considered to be statistically significant when P<0.05. Results Effect of the laminin competing peptides YIGSR and RGDS on insulin-induced hypercontractility. To assess whether laminin � and �1 chains were required for the induction of a functional hypercontractile phenotype, BTSM strips were incubated with insulin in the absence and presence of the laminin competing peptides Tyr-Ile-Gly-Ser-Arg (YIGSR, 100 �M), Arg-Gly-Asp-Ser (RGDS, 100 �M) or the negative control Gly-Arg-Ala-Asp-Ser-Pro (GRADSP, 100 �M) for 8 days. In accordance with previous studies (18), we found that culturing of BTSM strips in the presence of insulin (1 �M) significantly increased maximal contractile force (Emax) to both methacholine and KCl (P<0.05, both) compared to vehicle treated controls (Figure 1, Table 2). Prevention of laminin binding by the laminin �1 chain competing peptide YIGSR normalized the insulin-induced increase in Emax for both stimuli to control levels, whereas no apparent effects of pretreatment with YIGSR were observed under control (insulin-deficient) conditions. The laminin competing peptide RGDS, containing the RGD sequence found in the laminin � chains (30), but not its negative control GRADSP, also prevented the induction of a hypercontractile phenotype by insulin. Of note, we have reported previously that treatment with the RGDS peptide in the absence of other stimuli did not affect contractility (Chapter 3). The sensitivity (pEC50) for either contractile stimulus was unaffected by all treatments. Collectively, these results indicate that endogenously expressed or synthesized laminin � and �1 chains are required for the induction of a functional hypercontractile phenotype by insulin.


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Figure 1: The induction of a hypercontractile phenotype by insulin is fully prevented by combined treatment with the laminin competing peptides YIGSR (A,B) or RGDS (C,D). Concentration-response curves of methacholine- (A,C) and KCl (B,D)-induced contractions of BTSM strips, pretreated with vehicle (control) or insulin (1 �M) in the absence or presence of the laminin �1 chain competing peptide YIGSR, the laminin � chain competing peptide RGDS or the negative control peptide GRADSP (100 �M) for 8 days. Data represent means ± SEM of 3-4 independent experiments each performed in triplicate.

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Table 2: Contractile responses of BTSM strips to methacholine or KCl after 8 days of culturing in the absence or presence of insulin (1 �M), with or without the laminin blocking peptides YIGSR (100 �M), RGDS (100 �M) or the negative control GRADSP (100 �M).

Methacholine KCl Emax (g) pEC50 (- log M) Emax (g) EC50 (mM) Vehicle (control) 19.2 ± 2.6 6.81 ± 0.03 12.2 ± 1.2 20.3 ± 1.4

+ YIGSR 17.6 ± 2.4 6.84 ± 0.12 13.6 ± 1.0 19.3 ± 1.6 Insulin 24.4 ± 1.8** 6.71 ± 0.07 17.1 ± 0.4** 19.0 ± 2.3

+ YIGSR 16.9 ± 1.8### 6.81 ± 0.13 11.2 ± 1.6## 20.4 ± 1.0 Vehicle (control) 17.8 ± 2.3 6.68 ± 0.11 13.3 ± 1.6 25.1 ± 0.8 Insulin 24.1 ± 3.9* 6.79 ± 0.15 19.0 ± 2.7* 23.6 ± 0.5

+ GRADSP 23.9 ± 3.0* 6.76 ± 0.11 18.4 ± 2.1* 22.9 ± 2.1 + RGDS 14.1 ± 1.3## 7.02 ± 0.14 13.7 ± 2.2# 22.8 ± 1.5

Data represent means ± SEM of 3-4 independent experiments each performed in triplicate. Abbreviations: Emax: maximal contraction; EC50: contraction of agonist eliciting half-maximal response; pEC50: negative logarithm of the EC50 value. *P<0.05, **P<0.01 compared to vehicle-treated. #P<0.05, ##P<0.01, ###P<0.001 compared to insulin-treated. Effects of YIGSR on insulin-induced sm-myosin accumulation. The changes in Emax in response to both receptor-dependent (methacholine) and receptor-independent (KCl) stimuli suggest post-receptor changes, which may occur at the level of the contractile apparatus. To assess these changes, we determined the expression of the sm-myosin, a specific marker for contractile ASM (16). In accordance with our previous studies (17), it was found that 8 days of insulin (1 �M) treatment increased sm-myosin protein expression in BTSM strips by approximately 2-fold as compared to controls (Figure 2). Combined treatment with YIGSR (100 �M) completely abrogated the insulin-induced increase in sm-myosin expression, in full agreement with the findings on contractility. No effects of the peptide were observed under vehicle-treated conditions. Interestingly, sm-myosin abundance in strips pretreated with both insulin and YIGSR appeared to be even lower compared to that in strips that were kept under insulin deficient conditions; however, this did not reach statistical significance. These results implicate that the insulin-induced increase in sm-myosin abundance is strongly dependent on the abundance of endogenous �1-containing laminins.


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Effects of YIGSR on inhibition of growth factor-induced proliferation by laminin. As we previously demonstrated that laminin-111 inhibits BTSM cell proliferation in response to PDGF (Chapter 3), we were interested to learn whether this effect was dependent upon ligation of laminin with its receptors. Therefore, the effects of YIGSR on basal and growth factor-induced DNA-synthesis ([3H]-thymidine-incorporation) were examined, using freshly isolated BTSM cells cultured on plastic or on laminin matrices. As observed previously (Chpater 3)(24), culturing of cells on the exogenously-applied laminin matrix significantly reduced DNA-synthesis in response to PDGF, whereas no significant changes were observed on basal proliferative responses, both on plastic or laminin-coated surfaces (Figure 3). Similar effects were observed for cell number (data not shown). Treatment with YIGSR, by itself, had no effect on basal or growth factor-induced DNA-synthesis in cells grown on plastic. By contrast, the suppressive effects of laminin on the PDGF-induced DNA-synthesis were fully prevented in the presence of YIGSR, indicating that EHS laminin inhibits growth factor-induced DNA-synthesis through ligation with its receptors, which can be inhibited by the laminin �1 competing peptide YIGSR.

Figure 2: YIGSR attenuates the insulin-induced increase in expression of sm-myosin in BTSM strips. Western analysis of sm-myosin protein expression in homogenates of BTSM strips pretreated with vehicle (control) or insulin (1 �M) in the absence or presence of YIGSR (100 �M) for 8 days. (A) Graph shows means ± SEM of 4 independent experiments after densitometric analysis. *P<0.05 compared to control. ††P<0.01 compared to insulin. (B) Representative immunoblots of sm-myosin (upper panel) and �-actin (lower panel).

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Effects of YIGSR on inhibition of growth factor-induced proliferation by insulin. As pretreatment with insulin has been shown to decrease the mitogenic capacity of BTSM cells (18) and as our current study using YIGSR implies a role for �1-containing laminins in the induction of a hypercontractile phenotype by insulin (Figures 1 and 2), we next investigated the role of laminin signaling in the inhibitory effects of insulin pretreatment on growth factor-induced proliferation. To this aim, freshly isolated BTSM cells were cultured on plastic in the absence or presence of insulin, with or without YIGSR (100 �M) for 7 days. After pretreatment, the cells were exposed to either vehicle or PDGF (10 ng/ml). Consistent with our previous findings (18), we found that pretreatment with insulin for 7 days almost completely inhibited the mitogenic effects induced by PDGF (Figure 4). Importantly, this attenuation was fully prevented by combined YIGSR treatment, suggesting that signaling via �1-containing laminins is pivotal for the insulin-induced hypoproliferative BTSM phenotype. Of note, no effects of YIGSR or insulin pretreatment were found on basal DNA-synthesis or cell number (data not shown).

Figure 3: Reduced growth factor-induced DNA-synthesis by BTSM cells grown on a laminin-111 matrix is prevented by YIGSR treatment. Basal and PDGF (10 ng/ml)-stimulated [3H]-thymidine-incorporation in isolated BTSM cells cultured on plastic or laminin-111 (10 �g/ml) matrices in the absence or presence of YIGSR (100 μM) for 7 days. Data represent means ± SEM of 5 independent experiments, each performed in triplicate. ***P<0.001 compared to basal. #P<0.05 compared to PDGF-induced DNA-synthesis in cells grown on plastic (control). †††P<0.001 compared to PDGF-induced DNA-synthesis on laminin-111 matrices in the absence of YIGSR.


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Time-dependent effects of insulin on the accumulation of laminin protein expression in bovine tracheal smooth muscle strips. As our results with the YIGSR peptide indicate that signaling via �1-containing laminins is required for the insulin-induced hypercontractile, hypoproliferative phenotype, we next investigated the effects of insulin on laminin protein accumulation. Previous studies have shown that mRNA and protein levels of laminin-211 chains, but not of laminin-111, are increased during ASM maturation (25). Therefore we investigated the effects of insulin on laminin �1, �2, �1 and �1 chain protein accumulation in homogenates from BTSM strips cultured for up to 8 days. Insulin exposure led to a significant increase in the abundance of laminin �2, �1 and �1 chains, whereas no effects were observed for �1 chain expression after 8 days of stimulation (Figure 5). Expression of laminin �2 chains increased rapidly and peaked at day 4 with an increase of approximately 2-fold compared to freshly isolated tissue, after which expression remained elevated. Expression of laminin �1 and �1 chains was significantly increased by approximately 3-fold for both chains after 8 days of insulin exposure compared to freshly isolated tissue. No significant increases were observed under insulin-deficient conditions for all chains at any time point. These findings suggest that the accumulation of laminin �2, �1, and �1 chains in BTSM tissue is increased after insulin stimulation.

Figure 4: The insulin-induced hypoproliferative BTSM phenotype is fully prevented by combined YIGSR treatment. Basal and PDGF (10 ng/ml)-stimulated [3H]-thymidine-incorporation of isolated BTSM cells cultured on plastic and pretreated with vehicle (control) or insulin, in the absence or presence of YIGSR (100 �M) for 7 days. Data represent means ± SEM of 5 independent experiments each performed in triplicate. *P<0.05, **P<0.01, ***P<0.001 compared to basal. #P<0.05 compared to PDGF-induced DNA-synthesis in cells grown in the absence of insulin. †††P<0.001 compared to PDGF-induced DNA-synthesis in cells pretreated with insulin in the absence of YIGSR.

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A B

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Figure 5: Insulin time-dependently increases laminin �2, �1 and �1 chain expression in BTSM strips. Western analysis of laminin �1, �2, �1 and �1 chain expression in homogenates of BTSM strips pretreated with vehicle (control) or insulin (1 �M). (A) Representative immunoblots of laminin �1, �2, �1 and �1 chains and corresponding �-actin. Graphs show means ± SEM of 4 independent experiments after densitometric analysis of (B) laminin �2, (C) laminin �1, and (D) laminin �1 chains. *P< 0.05, **P<0.01 compared to vehicle treatment (control). #P<0.05, ##P<0.01 compared to laminin chain expression in homogenates of freshly isolated BTSM strips. Intracellular signaling associated with the insulin-induced laminin �2-, �1- and �1-chain accumulation. Both PI3-kinase- and Rho kinase-dependent signaling pathways have been shown to underlie the insulin-induced accumulation of the contractile phenotype marker proteins sm-myosin and calponin in ASM cells (17). To study the role of these enzymes in the accumulation of laminin �2, �1, and �1 chains, we co-incubated BTSM strips with insulin and the selective pharmacological inhibitors of PI3-kinase (LY294002, 10 �M) and Rho kinase (Y27632, 1 �M) for 8 days. Inhibition of PI3-kinase and Rho kinase significantly reduced laminin �2-chain abundance and even fully prevented an increase in laminin �1 and �1 chain expression in BTSM strip homogenates cultured in the presence of insulin (Figure 6). The inhibitors did not affect the expression of the laminin chains in


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strips cultured under insulin-deficient conditions. Collectively, these results suggest a key role for Rho kinase and PI3-kinase dependent signaling pathways in the excessive accumulation of laminin �2, �1 and �1 chains by insulin.

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Figure 6: PI3-kinase and Rho kinase signaling are required for the insulin-induced laminin chain accumulation. Western analysis of laminin �2, �1 and �1 chain expression in homogenates of BTSM strips pretreated with vehicle (control) or insulin (1 �M) in the absence or presence of Y-27632 (1 μM) or LY-294002 (10 μM) for 8 days. Graphs show means ± SEM of 4 independent experiments after densitometric analysis of (A) laminin �2, (B) laminin �1 and (C) laminin �1 chains. *P< 0.05, **P<0.01, ***P<0.01 compared to unstimulated controls. †P<0.05, ††P<0.01 compared to 8 days of insulin treatment. (D) Representative immunoblots of laminin �2 chains (left, upper panel), laminin �1 and �1 chains (right, upper panel) and corresponding �-actin (lower panels). Effects of insulin treatment on laminin mRNA expression. As our studies indicated that abundance of laminin �2, �1, and �1 chains in BTSM tissue is increased by insulin treatment and that these increases were dependent on signaling pathways associated with increased transcription (Rho kinase) (31) or translation (PI3-kinase) (32) in ASM cells, we next assessed whether the increased laminin chain expression was dependent on changes in mRNA abundance. Real time PCR analysis, performed on mRNA from BTSM strips cultured for 8 days in the absence and presence of insulin, revealed that insulin treatment did not affect mRNA levels for the �2, �1 or the �1 laminin chain (Figure 7). Co-incubation of BTSM strips with LY294002 did not affect laminin chain mRNA abundance in the absence and presence of insulin either. Y27632 decreased laminin chain mRNA both under unstimulated (control) and

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insulin-stimulated conditions. These findings suggest that increased expression of the laminin chains after insulin stimulation is independent of changes in gene transcription. Rho-kinase dependent signaling pathways, however, are important for the maintenance of laminin �2, �1, and �1 chain mRNA abundance.

Figure 7: Effects of insulin treatment on accumulation of laminin chain mRNA in BTSM strips. Real time PCR analysis of laminin �2, �1, and �1 chain mRNA in BTSM strips pretreated with vehicle (control) or insulin (1 �M), in the absence or presence of Y-27632 (1 μM) or LY-294002 (10 μM) for 8 days. Graphs show changes in (A) laminin �2, (B) laminin �1 and (C) laminin �1 chain mRNA compared to vehicle (Control). Data represent means ± SEM of 5 independent experiments, each performed in duplicate. *P< 0.05, **P<0.01 compared to unstimulated controls.

Effects of PI3-kinase and Rho kinase inhibition on insulin-induced hypercontractility. The requirement of endogenously expressed laminin in the induction of a functional hypercontractile phenotype and the role of PI3-kinase- and Rho kinase-dependent signaling pathways in the insulin-induced accumulation of laminin �2, �1, and �1 chains, suggest that inhibition of these pathways would normalize the insulin-induced hypercontractile phenotype. Furthermore, we have previously shown that the effects of insulin on contractile protein accumulation were abrogated by combined pretreatment with LY294002 or Y27632 (17). To assess the role of these pathways in the induction of a functional hypercontractile BTSM phenotype, strips were incubated with insulin in the absence and presence of LY294002 or Y27632 for 8 days. Inhibition of PI3-kinase signaling by LY294002 fully normalized the insulin-induced increase in Emax for both methacholine and KCl to control levels, whereas only a small effect of LY294002 was observed under control conditions (Figure 8). In agreement with previous findings (33), we found that treatment with Y27632 in the absence of insulin for 8 days decreased maximal contractility in response to methacholine, whereas for KCl-induced contractions a trend towards a decreased contractility

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was observed. Inhibition of Rho kinase by Y27632 also completely abrogated the insulin-induced increase in Emax for both agonists. The sensitivity (pEC50) for either contractile stimulus was unaffected by all treatments (Table 3). Collectively, these findings indicate a key role for PI3-kinase and Rho kinase dependent signaling pathways in the induction of a hypercontractile phenotype by insulin. Figure 8: Inhibition of the insulin-induced hypercontractile BTSM phenotype by combined treatment with the PI3-kinase inhibitor LY294002 (A,B) or the Rho kinase inhibitor Y27632 (C,D). Concentration-response curves of methacholine- (A,C) and KCl-induced (B,D) contractions of BTSM strips pretreated with vehicle (control) or insulin (1 �M), in the absence or presence of LY294002 (10 μM) or Y27632 (1 �M) for 8 days. Data represent means ± SEM of 4 independent experiments, each performed in duplicate.

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Table 3: Contractile responses of BTSM strips to methacholine or KCl after 8 days of culturing in the absence or presence of insulin (1 �M) with or without the Rho kinase inhibitor Y-27632 (1 μM) or the PI3-kinase inhibitor LY-294002 (10 μM).

Methacholine KCl Emax (g) pEC50 (- log M) Emax (g) EC50 (mM) Vehicle (control) 20.6 ± 2.4 6.71 ± 0.15 15.2 ± 1.9 24.7 ± 1.9

+ LY294002 18.3 ± 3.0 6.80 ± 0.11 13.2 ± 1.1 20.8 ± 1.3 +Y27632 15.1 ± 2.2* 6.70 ± 0.11 10.9 ± 0.9 23.4 ± 0.5

Insulin 25.8 ± 2.8* 6.92 ± 0.11 20.5 ± 2.3* 21.9 ± 1.5 + LY294002 16.3 ± 1.3### 6.81 ± 0.04 11.2 ± 1.0### 20.3 ± 1.7

+Y27632 16.2 ± 2.2### 6.57 ± 0.10 12.9 ± 1.8## 25.0 ± 1.3

Data represent means ± SEM of 4 independent experiments each performed in duplicate. Abbreviations: Emax: maximal contraction; EC50: contraction of agonist eliciting half-maximal response; pEC50: negative logarithm of the EC50 value. *P<0.05 compared to vehicle-treated. ##P<0.01, ###P<0.001 compared to insulin-treated. Discussion In this study, we demonstrate for the first time that induction of a functional hypercontractile and hypoproliferative ASM phenotype by insulin is dependent on the interaction of laminin with its receptors. Moreover, 8 days of insulin exposure resulted in a time-dependent increase in the abundance of laminin �2, �1 and �1 chains in isolated BTSM strip preparations, a process relying on PI3-kinase- and Rho kinase-mediated signaling. Inhibition of these pathways also fully inhibited the insulin-induced hypercontractile BTSM phenotype.

Accumulation of contractile ASM cells has been described in the airways of asthmatics (13, 14). In vitro, a functional hypercontractile phenotype can be induced by culturing isolated BTSM strips in the presence of insulin for 8 days. This hypercontractile phenotype is typically characterized by an increased maximal contractile response to both the muscarinic receptor agonist methacholine and the membrane-depolarizing agent KCl (18). The increased maximal contraction to both receptor-dependent and receptor-independent stimuli suggests post-receptor changes, which can occur at the level of the contractile apparatus. Accordingly, in the present study we have shown that increased contractility of BTSM strip preparations is associated with increased expression of the contractile phenotype marker protein sm-myosin. Moreover, we found that the insulin-induced hypercontractile BTSM phenotype and increased sm-myosin expression in BTSM strips was fully prevented by the YIGSR peptide, an antagonistic penta-peptide derived from the amino acid sequence of the �1-chain, which contains the receptor binding site in laminins (34). Similarly, treatment with the RGDS peptide, which contains the RGD


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binding motif present in several ECM proteins (35), including the laminin � chain (30), also fully normalized induction of a functional hypercontractile phenotype by insulin. These findings are consistent with the important role for laminin signaling in human ASM maturation (25), and show that interaction of laminin � and �1 chains with its receptors is pivotal for the induction of a functional hypercontractile ASM phenotype by insulin.

Changes in the maximal contractility of cultured BTSM strips induced by growth factors and ECM proteins have been shown to correlate inversely with the proliferative responses in isolated BTSM cells (Chapter 3)(8). In agreement with studies in cultured human (24) and bovine (Chapter 3) ASM cells, we found that culturing of cells on a laminin-111 matrix inhibited proliferation induced by the growth factor PDGF and maintained the cells in a hypoproliferative phenotype. This suppressive effect on PDGF-induced proliferation was fully prevented by YIGSR treatment, indicating that interaction of laminin-111 with its receptors is required for this inhibitory effect. The insulin-induced hypercontractile phenotype has previously been shown to be associated with a decreased mitogenic capacity as well (18). Similar to those findings, pretreatment of BTSM cells for 7 days with insulin did not affect basal [3H]-thymidine-incorporation or cell number, but did significantly attenuate the proliferative effects of PDGF-stimulation. We now show that co-incubation with the laminin �1 chain competing peptide YIGSR during insulin treatment normalized PDGF-induced DNA-synthesis to levels observed in cells grown in the absence of insulin, indicating that laminins containing the �1 chain, in addition to their important role in the induction of a hypercontractile phenotype, are also important for the induction of a hypoproliferative phenotype by insulin. The exact mechanism by which YIGSR inhibits laminin-111 signaling in PDGF-induced proliferation of BTSM cells is currently unknown. It may be envisaged that YIGSR interferes with existing interactions of immobilized laminin with its integrins. This interference would, however, occur without changes in cellular attachment, since we did not detect any changes in cell number or thymidine-incorporation on plastic or laminin in the presence of YIGSR. These observations are supported by previous findings (25), showing that YIGSR did not induce cellular detachment, changes in ASM morphology or cellular toxicity, but did inhibit laminin signaling in human ASM cells. Stimulation with PDGF could also possibly enhance expression of laminin-binding integrins, which would be blocked by YIGSR. In support of such a mechanism Nguyen et al found that culturing human ASM cells on plastic in the presence of PDGF specifically increased the expression of the laminin binding integrin �3 (36).

Increased expression of the contractile proteins desmin and calponin after serum deprivation in the presence of insulin has been shown to be associated with increased expression of laminin �2, �1 and �1 chain mRNA and protein by human ASM cells (25). In different cell types, increased expression of several laminin chains has been shown to be dependent on insulin signaling as well (37-39). Further expanding those findings we found that prolonged insulin

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exposure augments the protein abundance of laminin �2, �1 and �1 chains, but not �1 chains, in a time-dependent fashion, implying the involvement of the laminin-211, but not the laminin-111 isoform in the induction of a hypercontractile, hypoproliferative ASM phenotype. Indeed, previous findings have shown that culture of BTSM strips in the presence of exogenously applied laminin-111 only induced a small increase in BTSM contractility and contractile protein expression (Chapter 3), suggesting that laminin-111, by itself, is not able to induce a hypercontractile BTSM phenotype.

Little is known on the signaling pathways involved in the biosynthesis of laminins. However, an essential role for the PI3-kinase-Akt1 pathway in insulin-regulated laminin �1-chain gene transcription (38) and/or translation (39) has been suggested. Insulin-induced �1-chain transcription in Chinese Hamster Ovary cells was inhibited by expression of dominant negative Akt/PKB (38), whereas in murine proximal tubular epithelial cells activation of the PI3-kinase-Akt-mTOR pathway was found to be important for the translation of laminin �1-chains. In these cells insulin increased phosphorylation of the translational protein 4E-BP1 (PHAS-1), which released eukaryotic initiation factor 4E (eIF4E) and subsequently increased laminin �1-chain translation (39). In ASM cells activation of PI3-kinase and downstream pathways was found to be required for protein synthesis, maturation and hypertrophy (32), also in response to ligands such as insulin (17). In particular, accumulation of SM22 and sm-MHC in canine ASM cells has been shown to be dependent on PI3-kinase-mediated signaling pathways, involving Akt1, mTOR and p70S6K (32); pathways which can be activated by insulin in BTSM cells as well (17). In full agreement with these findings, we now demonstrate that inhibition of PI3-kinase reduced the accumulation of laminin �2, �1 and �1 chains in BTSM tissue and normalized insulin-induced hypercontractility. In addition, laminin �2, �1 and �1 chain mRNA abundance in BTSM strips was not affected after insulin stimulation, both in the absence or presence of the PI3-kinase inhibitor LY294002, suggesting that the increased laminin chain protein expression was dependent on post-transcriptional mechanisms.

A signaling pathway shown to be essential for ASM maturation by regulating transcription of smooth muscle-specific genes is the Rho/Rho kinase pathway (3, 17, 31). Activation of this pathway promotes actin polymerization, which governs nuclear translocation and activation of serum response factor (SRF), ultimately resulting in the transcription of smooth muscle specific genes (31). Rho kinase-dependent signaling has recently been shown to be activated by insulin to promote accumulation of contractile proteins in BTSM cells (17). To our knowledge, no reports on the role of Rho kinase signaling in laminin chain expression have been published thus far. Interestingly, we found a significant reduction in the expression of �2, �1 and �1 laminin chains after Rho kinase inhibition, indicating that pathways involved in the induction of contractile proteins and (hyper)contractility may similarly be involved in the expression of laminin chain-isoforms by ASM. In addition, we found that pharmacological


105

inhibition of Rho kinase decreased laminin �2, �1 and �1 chain mRNA levels both under control and insulin stimulated conditions, indicating that although insulin is capable of activation Rho kinase dependent signaling pathways in BTSM (17), activation of this pathway by insulin is not directly required for the increase in laminin protein abundance. These findings further suggest that Rho kinase is necessary for the maintenance of laminin �2, �1 and �1 chain mRNA abundance under basal conditions, which is conditional for increased protein translation after insulin stimulation. Clearly, further in depth investigation is warranted to more definitively determine the role of Rho kinase in regulating the expression of laminin chains.

No significant effects of inhibition of laminin signaling by YIGSR were observed on both contractile and mitogenic responsiveness under vehicle treated (control) conditions. Similarly, no changes in the expression of the contractile proteins desmin and calponin were found in human ASM after 7 days culture in the presence of YIGSR (25).Moreover, exogenously applied laminin-111, by itself, has no effects on maximal ASM contractility (Chapter 3) or contractile protein expression (25). Collectively, these findings imply that additional factors are likely to be required for the induction of a hypercontractile phenotype by insulin. One of these factors could be the laminin-binding integrin �7B, which has recently been shown to be involved in ASM maturation (40).

Adverse effects of inhaled insulin administrations for treatment of diabetes type 1 and 2 have been reported, including decline in lung function in some of the patients (26). Acutely, insulin has been shown to have procontractile effects on ASM (41). Although the exposure of the ASM cells to insulin after inhaled administration is likely to differ from our experimental setup, insulin concentrations in the ASM compartment could reach high levels due to deposition in the bronchial tree. Bronchial deposition results in a low bioavailability (~10-20%) thus necessitating relatively high insulin doses for satisfactory glycemic control (42). Collectively, these findings may particularly be important for diabetic patients with asthma or COPD, in which lung function is already compromized and who, due to poor absorption, may require higher dosages (26).

In conclusion, the current study provides new insight in the development of a hypercontractile and hypoproliferative ASM phenotype induced by insulin. This phenotype modulation involves insulin-induced laminin �2, �1 and �1 chain expression through PI3-kinase and Rho kinase dependent pathways, which may subsequently cause the induction of contractile proteins and inhibit ASM proliferation. Moreover, our findings indicate that increased laminin expression in the airways of asthmatic patients may contribute to the increased ASM contractility and contractile protein expression observed in these patients, on airway smooth muscle phenotype and function. In addition, the induction of laminin expression by insulin may limit the use of insulin as a substituent in serum-free media, as applied in cell culturing experiments in vitro.

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Acknowledgements The authors would like to thank Mariët van der Meer and Jeroen Lubbers for their expert technical assistance. Bart Dekkers is supported by a grant from the Netherlands Asthma Foundation (grant 3.2.03.36). Dedmer Schaafsma is the recipient of a CIHR/HSFC IMPACT Strategic Training Program Fellowship in Pulmonary and Cardiovascular Research. Thai Tran is supported by the Ministry of Education’s Academic Research Fund – Tier 1 (T13-0702-P26). References 1. Bousquet J, Jeffery PK, Busse WW, Johnson M, Vignola AM. Asthma. From

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2. Vignola AM, Kips J, Bousquet J. Tissue Remodeling As a Feature of Persistent Asthma. J Allergy Clin Immunol 2000;105:1041-1053.

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4. Ebina M, Takahashi T, Chiba T, Motomiya M. Cellular Hypertrophy and Hyperplasia of Airway Smooth Muscles Underlying Bronchial Asthma. A 3-D Morphometric Study. Am Rev Respir Dis 1993;148:720-726.

5. Parameswaran K, Willems-Widyastuti A, Alagappan VK, Radford K, Kranenburg AR, Sharma HS. Role of Extracellular Matrix and Its Regulators in Human Airway Smooth Muscle Biology. Cell Biochem Biophys 2006;44:139-146.

6. Fernandes DJ, Bonacci JV, Stewart AG. Extracellular Matrix, Integrins, and Mesenchymal Cell Function in the Airways. Curr Drug Targets 2006;7:567-577.

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8. Gosens R, Meurs H, Bromhaar MM, McKay S, Nelemans SA, Zaagsma J. Functional Characterization of Serum- and Growth Factor-Induced Phenotypic Changes in Intact Bovine Tracheal Smooth Muscle. Br J Pharmacol 2002;137:459-466.

9. Howarth PH, Knox AJ, Amrani Y, Tliba O, Panettieri RA, Jr., Johnson M. Synthetic Responses in Airway Smooth Muscle. J Allergy Clin Immunol 2004;114:S32-S50.

10. Hirst SJ, Walker TR, Chilvers ER. Phenotypic Diversity and Molecular Mechanisms of Airway Smooth Muscle Proliferation in Asthma. Eur Respir J 2000;16:159-177.

11. Halayko AJ, Solway J. Molecular Mechanisms of Phenotypic Plasticity in Smooth Muscle Cells. J Appl Physiol 2001;90:358-368.

12. Halayko AJ, Camoretti-Mercado B, Forsythe SM, Vieira JE, Mitchell RW, Wylam ME, Hershenson MB, Solway J. Divergent Differentiation Paths in Airway Smooth Muscle Culture: Induction of Functionally Contractile Myocytes. Am J Physiol 1999;276:L197-L206.


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15. Ma X, Wang Y, Stephens NL. Serum Deprivation Induces a Unique Hypercontractile Phenotype of Cultured Smooth Muscle Cells. Am J Physiol 1998;274:C1206-C1214.

16. Halayko AJ, Salari H, Ma X, Stephens NL. Markers of Airway Smooth Muscle Cell Phenotype. Am J Physiol 1996;270:L1040-L1051.

17. Schaafsma D, McNeill KD, Stelmack GL, Gosens R, Baarsma HA, Dekkers BG, Frohwerk E, Penninks JM, Sharma P, Ens KM, et al. Insulin Increases the Expression of Contractile Phenotypic Markers in Airway Smooth Muscle. Am J Physiol Cell Physiol 2007;293:C429-C439.

18. Gosens R, Nelemans SA, Hiemstra M, Grootte Bromhaar MM, Meurs H, Zaagsma J. Insulin Induces a Hypercontractile Airway Smooth Muscle Phenotype. Eur J Pharmacol 2003;481:125-131.

19. Patarroyo M, Tryggvason K, Virtanen I. Laminin Isoforms in Tumor Invasion, Angiogenesis and Metastasis. Semin Cancer Biol 2002;12:197-207.

20. Nguyen NM, Senior RM. Laminin Isoforms and Lung Development: All Isoforms Are Not Equal. Dev Biol 2006;294:271-279.

21. Schuger L, Skubitz AP, Zhang J, Sorokin L, He L. Laminin Alpha1 Chain Synthesis in the Mouse Developing Lung: Requirement for Epithelial-Mesenchymal Contact and Possible Role in Bronchial Smooth Muscle Development. J Cell Biol 1997;139:553-562.

22. Relan NK, Yang Y, Beqaj S, Miner JH, Schuger L. Cell Elongation Induces Laminin Alpha2 Chain Expression in Mouse Embryonic Mesenchymal Cells: Role in Visceral Myogenesis. J Cell Biol 1999;147:1341-1350.

23. Altraja A, Laitinen A, Virtanen I, Kampe M, Simonsson BG, Karlsson SE, Hakansson L, Venge P, Sillastu H, Laitinen LA. Expression of Laminins in the Airways in Various Types of Asthmatic Patients: a Morphometric Study. Am J Respir Cell Mol Biol 1996;15:482-488.

24. Hirst SJ, Twort CH, Lee TH. Differential Effects of Extracellular Matrix Proteins on Human Airway Smooth Muscle Cell Proliferation and Phenotype. Am J Respir Cell Mol Biol 2000;23:335-344.

25. Tran T, McNeill KD, Gerthoffer WT, Unruh H, Halayko AJ. Endogenous Laminin Is Required for Human Airway Smooth Muscle Cell Maturation. Respir Res 2006;7:117.

26. McMahon GT, Arky RA. Inhaled Insulin for Diabetes Mellitus. N Engl J Med 2007;356:497-502.

27. von Kriegstein E, von Kriegstein K. Inhaled Insulin for Diabetes Mellitus. N Engl J Med 2007;356:2106.

28. Timpl R, Rohde H, Robey PG, Rennard SI, Foidart JM, Martin GR. Laminin--a Glycoprotein From Basement Membranes. J Biol Chem 1979;254:9933-9937.

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30. Aumailley M, Gerl M, Sonnenberg A, Deutzmann R, Timpl R. Identification of the Arg-Gly-Asp Sequence in Laminin A Chain As a Latent Cell-Binding Site Being Exposed in Fragment P1. FEBS Lett 1990;262:82-86.

31. Liu HW, Halayko AJ, Fernandes DJ, Harmon GS, McCauley JA, Kocieniewski P, McConville J, Fu Y, Forsythe SM, Kogut P, et al. The RhoA/Rho Kinase Pathway Regulates Nuclear Localization of Serum Response Factor. Am J Respir Cell Mol Biol 2003;29:39-47.

32. Halayko AJ, Kartha S, Stelmack GL, McConville J, Tam J, Camoretti-Mercado B, Forsythe SM, Hershenson MB, Solway J. Phophatidylinositol-3 Kinase/Mammalian Target of Rapamycin/P70S6K Regulates Contractile Protein Accumulation in Airway Myocyte Differentiation. Am J Respir Cell Mol Biol 2004;31:266-275.

33. Gosens R, Schaafsma D, Meurs H, Zaagsma J, Nelemans SA. Role of Rho-Kinase in Maintaining Airway Smooth Muscle Contractile Phenotype. Eur J Pharmacol 2004;483:71-78.

34. Graf J, Ogle RC, Robey FA, Sasaki M, Martin GR, Yamada Y, Kleinman HK. A Pentapeptide From the Laminin B1 Chain Mediates Cell Adhesion and Binds the 67,000 Laminin Receptor. Biochemistry 1987;26:6896-6900.

35. Plow EF, Haas TA, Zhang L, Loftus J, Smith JW. Ligand Binding to Integrins. J Biol Chem 2000;275:21785-21788.

36. Nguyen TT, Ward JP, Hirst SJ. Beta1-Integrins Mediate Enhancement of Airway Smooth Muscle Proliferation by Collagen and Fibronectin. Am J Respir Crit Care Med 2005;171:217-223.

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38. Li X, Talts U, Talts JF, Arman E, Ekblom P, Lonai P. Akt/PKB Regulates Laminin and Collagen IV Isotypes of the Basement Membrane. Proc Natl Acad Sci U S A 2001;98:14416-14421.

39. Mariappan MM, Feliers D, Mummidi S, Choudhury GG, Kasinath BS. High Glucose, High Insulin, and Their Combination Rapidly Induce Laminin-Beta1 Synthesis by Regulation of MRNA Translation in Renal Epithelial Cells. Diabetes 2007;56:476-485.

40. Tran T, Ens-Blackie K, Rector ES, Stelmack GL, McNeill KD, Tarone G, Gerthoffer WT, Unruh H, Halayko AJ. Laminin-Binding Integrin {Alpha}7 Is Required for Contractile Phenotype Expression by Human Airway Myocyte. Am J Respir Cell Mol Biol 2007;37:668-680.

41. Schaafsma D, Gosens R, Ris JM, Zaagsma J, Meurs H, Nelemans SA. Insulin Induces Airway Smooth Muscle Contraction. Br J Pharmacol 2007;150:136-142.

42. Heinemann L, Pfutzner A, Heise T. Alternative Routes of Administration As an Approach to Improve Insulin Therapy: Update on Dermal, Oral, Nasal and Pulmonary Insulin Delivery. Curr Pharm Des 2001;7:1327-1351.

Bart G.J. Dekkers I. Sophie T. Bos Reinoud Gosens Andrew J Halayko Johan Zaagsma Herman Meurs

Am J Respir Crit Care Med (2010) 181:556-565

The integrin-blocking peptide RGDS inhibits airway smooth muscle remodeling in a guinea pig model of allergic asthma

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Abstract Rationale: Airway remodeling, including increased airway smooth muscle (ASM) mass and contractility, contributes to airway hyperresponsiveness in asthma. The mechanisms driving these changes are, however, incompletely understood. Recently, an important role for extracellular matrix proteins in regulating ASM proliferation and contractility was found, suggesting that matrix proteins and their integrins actively modulate airway remodeling. Objectives: To investigate the role of RGD (Arg-Gly-Asp)-binding integrins airway remodeling in an animal model of allergic asthma. Methods: Using a guinea pig model of allergic asthma, the effects of topical application of the integrin-blocking peptide RGDS (Arg-Gly-Asp-Ser) and its negative control GRADSP (Gly-Arg-Ala-Asp-Ser-Pro) were assessed on markers of ASM remodeling, fibrosis and inflammation induced by repeated allergen-challenge. In addition, effects of these peptides on human ASM proliferation and maturation were investigated in vitro. Measurements and main results: RGDS attenuated allergen-induced ASM hyperplasia and hypercontractility as well as increased pulmonary expression of smooth muscle myosin heavy chain and the proliferative marker PCNA. No effects were observed for GRADSP. The RGDS effects were ASM-selective, as allergen-induced eosinophil and neutrophil infiltration as well as fibrosis were unaffected. In cultured human ASM cells, we demonstrated that collagen I-, fibronectin-, serum- and platelet-derived growth factor-induced proliferation requires signaling via RGD-binding integrins, particularly of the �5�1 subtype. In addition, RGDS inhibited smooth muscle �-actin accumulation in serum-deprived ASM cells. Conclusions: This is the first study indicating that integrins modulate ASM remodeling in an animal model of allergic asthma, which can be inhibited by a small peptide containing the RGD motif. Introduction Asthma is a chronic inflammatory airways disease which is characterized by reversible airway obstruction, persistent airway inflammation and airway hyperresponsiveness (AHR) to a variety of stimuli (1,2,3). Structural changes in the airway wall of asthmatics are thought to contribute importantly to decreased airway diameter and AHR (4). These structural changes include accumulation of airway smooth muscle (ASM) (5) and changes in the amount and composition of the extracellular matrix (ECM) (6,7). Mechanisms driving airway remodeling are poorly understood; however, interactions between ASM cells and the ECM may be important.

The ECM is a dynamic macromolecular structure that surrounds tissue cells and provides structural support. In asthmatics, the deposition of ECM

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beneath the basement membrane is increased (7). Moreover, in patients with fatal asthma, the total amount of ECM in the ASM bundles is increased (8), which includes deposition of collagen I and fibronectin (9,10). Next to providing structural support, ECM proteins may regulate the function of cells embedded therein (7). Thus, proliferative responses to mitogens are markedly increased when ASM cells are cultured on collagen type I or fibronectin (11)(Chapter 3), whereas laminin, which does not affect ASM cell proliferation by its own, inhibits the induction of a proliferative phenotype by growth factors (11)(Chapter 3). Changes in the ECM may be an important determinant of changes in ASM mass in asthma, as proliferation of healthy ASM cells is increased when cultured on a matrix laid down by asthmatic cells (12). Interaction of cells with their surrounding matrix is mainly mediated through integrins, a group of heterodimeric transmembrane glycoproteins (13). Inhibition of the �5�1 integrins attenuates serum-induced proliferation and normalizes enhanced growth factor-induced proliferation of cells cultured on collagen I- or fibronectin-matrices (14,15), suggesting that �5�1 integrins are important regulators of ASM cell proliferation.

In asthmatic airways, accumulation of ASM cells with increased contractile properties and high expression levels of contractile proteins has been demonstrated (16,17). In culture, growth arrest in insulin-supplemented media increases the abundance of contractile proteins and contraction regulatory proteins (18,19,20). During this phase, ASM cells increase the expression of laminin chains �2, �1 and �1, which is essential for the induction of a functionally contractile phenotype (21)(Chapter 5).

The aim of the present study was to explore the potential role of integrins in allergen-induced airway remodeling. Using a guinea pig model of allergic asthma, we investigated the effects of topical treatment of the airways with the integrin-blocking peptide RGDS (Arg-Gly-Asp-Ser), containing the RGD binding motif present in collagen I, fibronectin and laminin (22,23), on ASM remodeling, fibrosis and inflammation induced by repeated allergen-challenge. In addition, the effects of RGDS and integrin function-blocking antibodies were assessed on human ASM cell proliferation and maturation in vitro. Some of the results have been previously reported in the form of an abstract (24).

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Materials and methods Additional detail on the methods is provided in an online data supplement, which is accessible from this issue’s table of content online at www.atsjournals.org. Animal experiments All protocols described in this study were approved by the University of Groningen Committee for Animal Experimentation. Outbred male, specified pathogen-free Dunkin Hartley guinea pigs (Harlan, Heathfield, UK) weighing 150–250 g were sensitized to ovalbumin using Al(OH)3 as an adjuvant, as described previously (25). Guinea pigs were challenged with aerosolized ovalbumin solutions until airways obstruction once weekly, for 12 consecutive weeks (26,27). Saline challenges served as controls. Animals were treated with saline, RGDS (H-Arg-Gly-Asp-Ser-OH, Calbiochem, Nottingham, UK) or GRADSP (H-Gly-Arg-Ala-Asp-Ser-Pro-OH, Calbiochem) by intranasal instillation (2.5 mM, 200 �l), 0.5 h prior to and 5.5 h after each challenge and sacrificed at 24 h after the last challenge. The lungs were resected and kept on ice until processing. The trachea was removed and transferred to a Krebs–Henseleit solution, pregassed with 5% CO2 and 95% O2, pH 7.4 at 37°C. Isometric tension measurements Isometric contraction experiments were performed as described previously (26,27). Briefly, epithelium-denuded, single open-ring tracheal preparations were mounted for isometric recording in organ baths, containing Krebs–Henseleit solution. After equilibration, resting tension was adjusted to 0.5 g, followed by precontractions with 20 mM and 40 mM KCl. Following washouts and another equilibration period of 30 min, cumulative concentration-response curves were constructed to KCl or methacholine. Histochemistry Transverse cross-sections of the main bronchi from both lung lobes were used for morphometric analysis. Sections were stained for smooth-muscle-specific myosin heavy chain (sm-MHC; Neomarkers, Fremont, CA). Primary antibodies were visualized using horseradish-peroxidase-linked secondary antibodies and diaminobenzidine. In addition, haematoxylin-stained nuclei within the ASM bundle were counted. Eosinophils were identified in haematoxylin- and eosin-stained lung sections. Neutrophils were identified by morphologic criteria and tissue non-specific alkaline phosphatase enzyme histochemistry (28). Airways within sections were digitally photographed and subclassified as cartilaginous or non-cartilaginous. All immunohistochemical measurements were carried out digitally using quantification software (ImageJ).


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Hydroxyproline assay Lungs were analyzed for hydroxyproline as an estimate of collagen content. Lung homogenates were prepared by pulverizing tissue under liquid nitrogen and sonification in 10 ml PBS. Homogenates were incubated with 1.25 ml 5% trichloroacetic acid on ice for 20 min. Samples were centrifuged and the pellet was resuspended in 12 N hydrochloric acid (10 ml) and heated overnight at 110 °C. The samples were dissolved in 2 ml water by incubating for 72 h at room temperature, applying intermittent vortexing. In a 96 well plate, samples (5 μl) were incubated with 100 μl chloramine T (1.4% chloramine T in 0.5 M sodium acetate/10% isopropanol) for 30 min at room temperature. Next, 100 μl Ehrlich’s solution (1.0 M 4-dimethylaminobenzaldehyde in 70% isopropanol/30% perchloric acid) was added and samples were incubated at 65 °C for 30 min. Samples were cooled to room temperature and the amount of hydroxyproline was quantified by colorimetric measurement (550 nm, Biorad 680 plate reader). Cell culture Three human bronchial smooth muscle cell lines, immortalized by stable expression of human telomerase reverse transcriptase (hTERT), were used (29,30). For all experiments, passage 18-40 myocytes grown on uncoated plastic dishes in DMEM (GIBCO BRL Life Technologies, Paisley, UK), supplemented

with 50 U/ml streptomycin, 50 μg/ml penicillin (Gibco) and 10% vol/vol FBS (Gibco) were used. In addition to the cell lines, primary human tracheal cells were used. Tracheal sections from anonymous donors were obtained from the Department of Cardiothoracic Surgery, University Medical Centre Groningen. The tracheal smooth muscle was prepared free of mucosa and connective tissue, and chopped using a McIlwain tissue chopper. Tissue particles were washed once with DMEM supplemented with 1 mM sodium pyruvate (Gibco), 1:100 nonessential amino-acid mixture (Gibco), 45 �g/ml gentamicin (Gibco), 50 �g/ml penicillin (Gibco), 50 U/ml streptomycin (Gibco), 1.5 �g/ml amphotericin B (Gibco) and 10% FBS (Gibco), placed in 25 cm2 culture flasks and allowed to adhere. Medium was refreshed every 48-72 h. Upon reaching confluency, cells were passaged by trypsinization. Cells from passages 2-3 were used for the

present study. Alamar blue proliferation assay ASM proliferation was assessed as described previously (30). Cells were plated on uncoated or ECM-coated culture plates, allowed to attach overnight and maintained in DMEM supplemented with antibiotics and 1% ITS (Insulin, Transferrin and Selenium) for 3 days. Coated culture plates were prepared as described previously (Chapter 3). Cells were then incubated with or without PDGF-AB (10 ng/ml, human, Bachem, Weil am Rhein, Germany) or FBS (10%) for 4 days. Thereafter, cells were incubated with HBSS containing 5% Alamar blue solution for 30 minutes (Biosource, Camarillo, CA). Proliferation was

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assessed by conversion of Alamar blue, as indicated by the manufacturer. In some experiments, cells were treated with RGDS (1-100 �M), GRADSP (100 �M), integrin function–blocking monoclonal anti-�5 (10 �g/ml, clone P1D6, Chemicon, Chandler's Ford, UK) and/or anti-�1 (10 �g/ml, clone 6S6, Chemicon) antibodies or mouse IgG control antibodies (10 �g/ml, Chemicon) 30 min before and during stimulation with mitogens. Western analysis Lung homogenates were prepared as described previously (26,27). Equal amounts of protein were subjected to sodium dodecyl sulfate/polycrylamide gel electrophoresis and transferred onto nitrocellulose membranes, followed by standard immunoblotting techniques. Antibodies were visualized on film using enhanced chemiluminescence reagents (Pierce, Breda, Netherlands) and analyzed by densitometry (TotallabTM, Nonlinear dynamics, Newcastle, UK). All bands were normalized to �-actin expression. Statistics All data represent means ± SEM from n separate experiments. Statistical significance of differences was evaluated using one-way ANOVA, followed by a Newman–Keuls multiple comparisons test. Differences were considered to be statistically significant when P<0.05. Results RGD-binding integrins mediate allergen-induced accumulation of ASM in a guinea pig model of allergic asthma. To explore the potential role of RGD-binding integrins in airway remodeling, we evaluated the effects of treatment with the integrin antagonist RGDS in a guinea pig model of allergic asthma. This model is characterized by allergen-induced early and late asthmatic reactions, airway inflammation, airway hyperresponsiveness and airway remodeling (25). We first investigated the increase in ASM mass by examining lung sections stained for sm-MHC. Repeated ovalbumin challenge increased the sm-MHC-positive area predominantly in the cartilaginous airways, by 1.9±0.1 fold (P<0.001) compared to saline-treated, saline-challenged controls (Figure 1A). Topical treatment of the airways with intranasally instilled RGDS 0.5 h prior to and 5.5 h after each allergen challenge reduced the ovalbumin-induced increase in sm-MHC positive area by 73±7% (P<0.01). In contrast, treatment with the control peptide GRADSP had no effect. Moreover, in saline-challenged animals, neither RGDS nor GRADSP significantly changed the sm-MHC positive area (Figure 1A).


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Figure 1: ASM hyperplasia after repeated allergen challenge in vivo is inhibited by topical treatment with RGDS. To assess the role of RGD-binding integrins in ASM hyperplasia in asthma, the effects of treatment with RGDS or GRADSP were evaluated in a guinea pig model of allergic asthma. (A) Treatment with RGDS inhibited ovalbumin-induced increase in ASM area in the cartilaginous airways. (B) Changes in ASM content were dependent on changes in cell number, as the cell size was unchanged. (C) Expression of the proliferative marker PCNA in lung homogenates was increased by repeated ovalbumin challenges. Treatment with RGDS partially reversed the ovalbumin-induced increase in PCNA expression. Representative blots of PCNA and �-actin are shown. No effects were observed for GRADSP on any of the parameters. ***P<0.001 compared to saline-treated, saline-challenged controls; ##P<0.01 compared to saline-treated, ovalbumin-challenged controls. Data represent means ± S.E.M of 6-8 animals. BM = basement membrane. To determine whether allergen-induced changes in ASM content were associated with changes in cell number and/or changes in cell size, the numbers of nuclei within the ASM layer were counted and expressed relative to total ASM area. With this data, the apparent cell volume of the ASM cells was also calculated. Repeated ovalbumin challenge did not change the number of nuclei

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per mm2 of smooth muscle area (Figure 1B) or cell volume (Figure E1), indicating that the cell size is unchanged and allergen-induced increases in ASM mass are caused by an increased cell number. In addition, both in the ovalbumin- and in the saline-treated animals RGDS and GRADSP did not cause a change in cell size. Collectively, these observations indicate that the inhibitory effect of RGDS on ovalbumin-induced ASM growth is caused by inhibition of ASM cell hyperplasia induced by the repeated allergen exposure.

To investigate the potential role of cell proliferation in the allergen-induced ASM hyperplasia, the expression of PCNA, an auxiliary factor for DNA-replication and repair (31), was measured in whole lung homogenates by Western analysis. A considerable increase (4.1±0.2 fold, P<0.001) in PCNA expression was observed after repeated ovalbumin-challenge compared to saline-treated, saline-challenged controls (Figure 1C). Treatment with RGDS attenuated the ovalbumin-increase in PCNA expression by 58±9% (P<0.01), whereas treatment with GRADSP had no effect. In saline-challenged animals, no effects of peptide treatment on PCNA expression were observed at all. Unfortunately, specific characterization of the proliferating cells in guinea pig lung sections by immunohistochemistry was not possible with the antibody used for the Western analysis mentioned above. Collectively, these data indicate that RGD-binding integrins are involved in allergen-induced proliferative responses in the lung, underlying ASM hyperplasia. RGDS treatment inhibits allergen-induced contractile protein accumulation and hypercontractility The present study and previous observations from our laboratory (26,27) indicate that repeated allergen exposure leads to increased expression of the contractile protein sm-MHC in the guinea pig lung. To investigate the role of integrin signaling in sm-MHC accumulation, Western analysis was used to determine sm-MHC expression in whole lung homogenates from saline-, RGDS- and GRADSP-treated animals. In saline-treated animals, repeated ovalbumin challenges increased sm-MHC expression by 2.5±0.1 fold (P<0.001) compared to saline-challenged controls (Figure 2A). RGDS treatment inhibited the ovalbumin-induced increase in sm-MHC expression by 60±11% (P<0.01), whereas no inhibitory effect of GRADSP was observed. In saline-challenged animals, treatment with RGDS or GRADSP did not affect sm-MHC expression.

Consistent with the increased expression of sm-MHC and with previous findings (26,27), maximal methacholine- and KCl-induced isometric contractions of epithelium-denuded, tracheal open-ring preparations from ovalbumin-challenged animals were significantly enhanced (1.7±0.2 fold and 1.7±0.1 fold, respectively, P<0.001; Figure 2B, Table 1). Notably, in vivo treatment with RGDS completely prevented allergen exposure-induced hypercontractility for both stimuli (P<0.001). No effects on ASM contractility were observed for GRADSP in saline- or allergen-challenged animals. The sensitivity (pEC50) of methacholine- or KCl-induced contraction was unaffected by all treatments


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(Table 1). These data demonstrate that RGD-binding integrins have a key role in the concomitant increase in contractile protein expression and ASM contractility induced by repeated allergen exposure in vivo. Figure 2: Increased contractile protein accumulation and ASM contractility after repeated allergen challenge are inhibited by topical treatment with RGDS in the airways. (A) Treatment with RGDS inhibited ovalbumin-induced accumulation of sm-MHC in the guinea pig lung, while treatment with GRADSP had no effect. Representative western blots of sm-MHC and �-actin are shown. (B) Treatment with RGDS but not with GRADSP fully normalized the ovalbumin-induced enhancement of maximal methacholine-induced isometric contraction of epithelium-denuded tracheal open-ring preparations. ***P<0.001 compared to saline-treated, saline-challenged controls; ##P<0.01 and ###P<0.001 compared to saline-treated, ovalbumin-challenged controls. Data represent means ± S.E.M of 6-8 animals.

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Table 1: Contractile responses of epithelium-denuded, tracheal open-ring preparations after repeated saline or ovalbumin challenges of saline-, RGDS- or GRADSP-treated guinea pigs.

Methacholine KCl Treatment Challenge

Emax (g) pEC50 (-logM) Emax (g) EC50 (mM)

n

Saline Saline 1.42 ± 0.09 6.55 ± 0.18 1.02 ± 0.06 23.7 ± 0.9 6 RGDS Saline 1.46 ± 0.07 6.54 ± 0.16 1.11 ± 0.07 24.4 ± 1.6 6 GRADSP Saline 1.28 ± 0.04 6.46 ± 0.13 0.95 ± 0.04 24.6 ± 1.7 5 Saline Ovalbumin 2.43 ± 0.22*** 6.28 ± 0.11 1.73 ± 0.13*** 23.7 ± 1.2 7 RGDS Ovalbumin 1.22 ± 0.10### 6.30 ± 0.08 0.82 ± 0.07### 23.6 ± 1.1 8 GRADSP Ovalbumin 2.55 ± 0.27*** 6.29 ± 0.07 1.88 ± 0.25*** 25.7 ± 1.0 6

Data represent means ± SEM. Abbreviations: Emax: maximal contraction; EC50: concentration of agonist eliciting half-maximal response; pEC50: negative logarithm of the EC50 value. ***P<0.001 compared to saline-treated, saline-challenged; ###P<0.001 compared to saline-treated, ovalbumin-challenged. Effects of RGDS treatment on allergen-induced airway inflammation Previous in vitro studies have shown that secretion of eotaxin by ASM cells is enhanced when the cells are grown on a fibronectin or collagen type I matrix (32). Moreover, the increased secretory response required interaction of the cells with fibronectin via RGD-binding integrins. In addition, enhanced eotaxin secretion by asthmatic ASM cells has also been reported, and this appears to be largely dependent on the RGD-binding �5�1 integrin (33). To investigate whether RGDS treatment inhibited eosinophilic airway inflammation in our guinea pig model, we assessed the number of eosinophils in different airway compartments. As we have observed previously (26), repeated ovalbumin challenge increased the number of eosinophils in the submucosal and adventitial compartments of the cartilaginous airways (P<0.001, Figures 3A and 3B). However, treatment with the RGDS or GRADSP did not significantly affect the eosinophil influx into these compartments.

RGDS might also inhibit infiltration of neutrophils into the airways, by inhibition of �5�1 integrins present on these cells (34). Repeated ovalbumin challenges increased the infiltration of neutrophils in the submucosa and adventitia of the cartilaginous airways (P<0.05, Figure 3C and 3D). However, as observed for eosinophils, no effects of RGDS or GRADSP treatment were found on the neutrophil infiltration after repeated allergen challenge. Collectively, these findings indicate that, despite its inhibitory effects on ASM remodeling in the cartilaginous airways, RGDS does not suppress eosinophilia and neutrophilia in these airways. This indicates that RGD-binding integrins are not central regulators of allergen-induced infiltration of eosinophils or neutrophils into the airways. Interestingly, these results also suggest that inhibition of ASM


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remodeling in the cartilaginous airways by RGDS is not related to an effect on airway inflammation. Figure 3: RGDS does not affect allergen-induced airway inflammation. Eosinophil numbers after saline or ovalbumin challenges in the submucosal (A) and adventitial (B) compartments of the cartilaginous airways were unaffected by RGDS or GRADSP treatment. Similarly, neutrophilic inflammation in the submucosal (C) and adventitial (D) compartments was unaffected by the treatments. *P<0.05, **P<0.01 and ***P<0.001 compared to saline-treated, saline-challenged controls. Data represent means ± S.E.M of 6-8 animals. BM = basement membrane Effect of RGDS treatment on allergen-induced fibrosis Increased deposition of ECM proteins in the airway wall, including collagens, is a characteristic feature of chronic asthma (35,36). To assess the potential effect of RGDS treatment on allergen-induced collagen production, guinea pig lungs were analyzed for hydroxyproline content, as an estimate of total collagen. Repeated ovalbumin challenges increased hydroxyproline content by 2.2±0.2-fold (P<0.001) compared to saline-challenged controls (Figure 4). However, treatment with RGDS or with GRADSP did not significantly affect hydroxyproline content both in ovalbumin- and in saline-challenged animals.

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RGD-binding integrins are involved in human ASM cell proliferation in vitro As RGDS inhibited allergen-induced increase in ASM mass and contractility in our guinea pig model, we investigated its direct effects on human ASM cell responses in vitro. First, we investigated the effects of RGDS on ECM-induced ASM cell proliferation. Culturing of human ASM cells on monomeric collagen type I or fibronectin increased proliferation, as shown by a 1.7±0.1 fold (P<0.01) increase in cell number for both ECM proteins. Treatment with RGDS fully inhibited both monomeric collagen type I- and fibronectin-induced ASM proliferation in a concentration-dependent fashion (P<0.01, Figure 5A). The concentration of RGDS required for 50% inhibition (IC50) of ECM-induced proliferation was not significantly different for cells cultured on monomeric collagen I or fibronectin (5.2±2.4 �M and 6.7±2.0 �M, respectively). No effects were observed for GRADSP. In addition, no effects of RGDS or GRADSP were observed in cells cultured on uncoated surface. >> Figure 5 (next page): ECM- and growth factor-induced human ASM cell proliferation are dependent on RGD-binding integrins. (A) Increased ASM cells numbers were observed after culturing on collagen I or fibronectin matrices compared to uncoated surface. Increases in proliferation were concentration-dependently inhibited by RGDS. (B) ECM-induced proliferation required interaction with the RGD-binding integrin �5�1. (C) PDGF- and (E) FBS-induced proliferation were dependent on RGD-binding integrins. The additive effects of collagen type I or fibronectin were fully normalized by RGDS. (D) Inhibitory effects of RGDS on PDGF-induced proliferation required integrin �5�1. (F) Growth-attenuating effects of laminin-111 were not affected by RGDS treatment. **P< 0.01, ***P<0.001 compared to cells (control) on uncoated surface. #P<0.05, ##P<0.01, ###P<0.001 compared to cells grown on ECM matrices and/or stimulated with mitogens in the absence of inhibitors. †P<0.05, ††P<0.01, †††P<0.001 compared to mitogen-stimulated cells grown on uncoated surface. Data represent means ± SEM of 6 independent experiments of three donors, performed in duplicate.

Figure 4: RGDS treatment does not affect allergen-induced fibrosis in guinea pig lung. Hydroxyproline content in guinea pig lung after repeated saline or ovalbumin challenges in saline-, RGDS- and GRADSP-treated animals. ***P<0.001 compared to saline-treated, saline-challenged controls. Data represent means ± S.E.M of 6-8 animals.


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To identify the integrin inhibited by RGDS, monoclonal anti-�5 and anti-�1 function-blocking antibodies were used. Treatment with the anti-�1 blocking antibody reduced both monomeric collagen type I- and fibronectin-induced proliferation by 54±4% and 55±4%, respectively (P<0.05 both, Figure 5B), whereas the anti-�5 blocking antibody only inhibited the fibronectin-induced increase in cell number (51±9%, P<0.05). Notably, combining the anti-�5 and the anti-�1 blocking antibodies fully abrogated the proliferation induced when cells were grown on either matrix (P<0.001). Treatment with non-immune control IgG antibodies did not affect ASM cell number and no effects of the blocking antibodies were observed in cells cultured on uncoated surface (Figure 5B).

The effects of RGDS on FBS (10%)- as well as on PDGF (10 ng/ml)-induced proliferation were also investigated. Both PDGF- and FBS-induced proliferation were concentration-dependently inhibited by the peptide (IC50 = 4.4±1.6 �M and 4.0±1.6 �M, respectively, Figures 5C and 5E). Full inhibition of PDGF-induced proliferation of ASM cells grown on uncoated plastic was observed (P<0.001, Figure 5C), whereas FBS-induced proliferation on this matrix was inhibited by 37±7% (P<0.01, Figure 5E). The additive effects of monomeric collagen I- or fibronectin-matrices on PDGF- and FBS-induced proliferation were also fully normalized by RGDS treatment (P<0.001). The IC50 values for the inhibition of the effects of monomeric collagen type I or fibronectin on growth factor-induced ASM proliferation were not significantly different (7.3±2.7 �M and 5.0±1.4 �M, respectively, for PDGF-induced proliferation, and 5.1±3.5 �M and 2.6±1.1 �M, respectively, for FBS-induced proliferation). As with RGDS, PDGF-induced proliferation was fully normalized by culturing in the presence of �5 and �1 function-blocking antibodies (P<0.01, Figure 5D).

To assess the role of RGDS-binding integrins in signaling by growth-attenuating matrix proteins, cells were cultured on laminin-111. FBS-induced proliferation of ASM cells was inhibited on laminin-111, which was not influenced by treatment with RGDS (Figure 5F). Collectively, the results indicate that pro-mitogenic responses of ASM are dependent on signaling by RGDS-binding integrins, in particular the �5�1 integrin. RGD-binding integrins are involved in contractile protein accumulation by ASM cells To assess the effects of RGDS on human ASM cell maturation, accumulation of the contractile protein sm-�-actin was induced by serum-deprivation in insulin-supplemented media, in the presence of different concentrations of the RGDS peptide. Serum-deprivation for 7 days markedly increased the expression of sm-�-actin by 1.79±0.10 fold (P<0.01). Treatment with RGDS dose-dependently inhibited the accumulation of sm-�-actin with an IC50-value of 0.47±0.15 �M and a maximal inhibition of 72±14% (P<0.05). No effects were observed for GRADSP (Figure 6). Collectively, our in vitro findings demonstrate that proliferation and accumulation of contractile proteins by human ASM cells can be concentration-dependently inhibited by a RGD-containing peptide.


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Discussion Using a well-established guinea pig model of allergic asthma, we for the first time demonstrated that RGD-binding integrins mediate allergen-induced ASM remodeling. It was shown that topical administration of the integrin-blocking peptide RGDS in the airways attenuated ASM proliferation, hyperplasia, increased contractile protein expression and hypercontractility induced by repeated allergen challenge. Furthermore, in vitro experiments using human ASM cells indicated that collagen I, fibronectin and laminin, which all contain the RGD binding motif (22,23), could be intrinsic regulators of these processes.

Accumulation of ASM and increased deposition ECM proteins, including collagen type I, fibronectin and laminin �2/�2, in the airway wall are characteristic features of remodeling in asthma (7,37). Previous reports have indicated that monomeric collagen type I and fibronectin increase basal ASM proliferation in cells of bovine origin (38)(Chapter 3). In the present study, we show that basal proliferation of human ASM cells is also increased by culturing on these ECM proteins and that these proliferative responses can be dose-dependently inhibited by RGDS. A number of integrins recognize the RGD-sequence within ECM proteins (22), of which the �3�1, �5�1, �v�1 and �v�3 integrins are expressed by ASM cells in culture (14,39). Previous studies using human ASM indicated that enhanced PDGF-induced proliferation on monomeric

Figure 6: RGD-binding integrins are involved in contractile protein accumulation in serum-deprived human ASM cells. Human ASM cells were grown to confluence in the presence of 10% FBS. Subsequently, ASM cells were kept in serum-free medium for 7 days in the absence or presence of different concentrations of the RGDS peptide or GRADSP. RGDS concentration-dependently inhibited sm-�-actin accumulation. Representative western blots of sm-�-actin and �-actin are shown. **P<0.01 compared to day 0 (10% FBS). #P<0.05 compared to sm-�-actin accumulation in the absence of inhibitors. Data represent means ± SEM of 4 independent experiments of three different donors.

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collagen I and fibronectin matrices is inhibited by function-blocking antibodies directed against �5�1, but not �3�1, �v�1 or �v�3 integrins (14). Using function-blocking antibodies we now established that increased basal ASM proliferation on both collagen and fibronectin-matrices also primarily requires interaction with the RGD-sensitive, fibronectin-selective integrin �5�1. Involvement of the �5�1 integrin in monomeric collagen type I-induced proliferation may be unexpected, as it has low affinity for collagens. Nonetheless, there are some reports indicating that this integrin is important for enhanced PDGF-induced proliferation by both monomeric collagen type I and fibronectin matrices (14), suggesting that the proliferative responses share a common mechanism. Indeed, studies in vascular smooth muscle cells have indicated that culturing on monomeric collagen type I markedly increased the expression of other ECM proteins (40), including fibronectin, suggesting that the proliferative responses on monomeric collagen type I could be influenced by autocrine fibronectin deposition and integrin �5�1 activation.

Not only ECM-induced proliferation but also PDGF- and FBS-induced proliferation of human ASM cells were inhibited by RGDS. These findings are in agreement with previous findings by Moir et al (41), who showed that FBS-induced proliferation of ASM cells obtained from both nonasthmatic and asthmatic patients was reduced by anti-�5 and anti-�1 function-blocking antibodies. The present study showed that also the potentiating effects of monomeric collagen type I and fibronectin on FBS- and PDGF-induced proliferation are fully normalized by RGDS treatment, which indicates that not only function-blocking antibodies (14), but also small peptides can inhibit the synergistic effects of monomeric collagen type I and fibronectin matrices on PDGF-induced proliferation. Similar to the effects described above for the bronchial smooth muscle cell lines, RGDS also inhibited fibronectin and PDGF-induced proliferation in primary human tracheal smooth muscle cells (Figure E2). No differences in the IC50 values of the inhibitory effects by the RGDS peptide toward the different stimuli were observed, strongly suggesting the involvement of a single integrin in these processes. The inhibitory effects of RGDS were specific for pro-proliferative processes, as the peptide did not normalize the inhibitory effects of laminin-111, previously shown to decrease proliferative responses in ASM cells (11)(Chapter 3). Collectively, these findings identify the �5�1 integrin, which is universally expressed on ASM cells (14,39), as an important regulator of ASM proliferation in vitro, which can be inhibited by a small peptide containing the RGD binding motif..

In patients with asthma, airway wall remodeling likely contributes to reduced airway lumen size, fixed airway obstruction, and persistent airway hyperresponsiveness (4,42). Thus far, increased ASM mass has primarily been attributed to the action of growth factors, inflammatory mediators and neurotransmitters (43). From animal models of allergic asthma it has been shown that TGF-�, cysteinyl leukotrienes and acetylcholine are important endogenous mediators of ASM remodeling (44). The role of endogenous ECM


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proteins and integrins in ASM remodeling in asthma has thus far been postulated, but not functionally investigated. Using our guinea pig model of allergic asthma, we found that ASM hyperplasia after repeated allergen-challenge is largely inhibited by topical treatment with RGDS, indicating that RGD-binding integrins are important regulators of increased ASM mass in vivo. The increased mass was due to ASM cell hyperplasia and accompanied by an increased expression of the proliferative marker PCNA in the lung. Although the specific contribution of the different lung cell types to increased PCNA expression could not be established, the data clearly demonstrate that RGDS reduces allergen-induced cell proliferation in total lung. In saline-challenged animals, RGDS treatment did not change sm-MHC positive area, indicating that the peptide does not promote loss of ASM cells per se. Hence, its effect may be to attenuate or prevent integrin activation occurring under pathological conditions, such as the repeated allergen exposure protocol we employed in our study. Increased integrin activation under these conditions may be the result of inflammation-induced changes in the ECM composition or of increased exposure of the integrins to RGD recognition sites within the ECM proteins. This may involve increased ECM synthesis, increased recruitment of plasma-derived ECM proteins to the ASM layer or alterations in the existing ECM within the airway, exposing binding sites which are normally hidden within the proteins (45,46). Indeed, previous studies by Nguyen et al (14) showed that proliferative responses of ASM cells are increased on monomeric, but not on fibrillar collagen type I, suggesting that degradation of the triple helical fibrillar form of collagen may be necessary for the binding to growth-promoting integrins.

Exposure of ASM cells to mitogenic stimuli not only increases proliferation but also results in the switching of the ASM phenotype from a contractile to a hypocontractile phenotype, characterized by increased proliferative rates and low expression levels of contractile proteins (47). In asthmatic airways, however, accumulation of ASM cells with increased contractile properties and high expression levels of contractile proteins has been described (16,17). These findings may be explained by the notion that phenotype plasticity is a reversible process. Thus, in vitro, proliferative hypocontractile ASM cells may be redirected to a (hyper)contractile state by serum deprivation, which can be further enhanced by the presence of insulin in the culturing medium (20). Previous studies have shown that increased expression of laminin-211 chains by ASM cells is required for maturation to a (hyper)contractile phenotype and that this effect may be attenuated by RGD-containing peptides (21)(Chapter 5). In agreement with these findings, we now demonstrate that accumulation of the contractile protein sm-�-actin by serum deprivation in human ASM cells was inhibited by RGDS, in a dose-dependent fashion. The IC50 values for the inhibitory effect of RGDS on sm-�-actin accumulation were clearly lower than observed for proliferation, suggesting that another integrin, potentially the �7 integrin, may be involved. Previous studies have indicated the �7 integrin is an important regulator of ASM maturation

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indeed (48). Collectively, our in vitro findings indicate that RGD-binding integrins are not only importantly involved in ASM proliferation via the �5�1 integrin, but also in ASM maturation to a (hyper)contractile phenotype, presumably via a mechanism involving the �7�1 integrin.

In our guinea pig model, a marked increase in sm-MHC (~2.5-fold) expression after repeated allergen-challenge was observed, which exceeds the increase in ASM mass (~1.75-fold) indicating that ASM thickening is accompanied by phenotype maturation of the ASM cells. Treatment with RGDS strongly inhibited allergen-induced increases in sm-MHC expression and fully normalized allergen-induced hypercontractility, indicating that RGD-binding integrins are important in phenotype maturation in vivo, which may be dependent on increased laminin signaling.

Airway inflammation is a characteristic feature of allergic asthma and is generally believed to contribute substantially to airway remodeling (42). Eotaxin is a key chemokine for the recruitment of eosinophils (49) and ASM cells derived from asthmatic patients release enhanced levels of this chemokine, presumably as a result of enhanced fibronectin deposition and subsequent �5�1 integrin activation (33). In addition, integrin-blocking peptides containing the RGD sequence inhibit the enhanced eotaxin release from healthy ASM cells cultured on fibronectin matrices (32). Treatment with RGDS may therefore potentially inhibit the influx of eosinophils to the airways by inhibiting the release of eotaxin from ASM. However, in our guinea pig model we found that administration of RGDS during repeated allergen challenge had no significant effect on the allergen-induced infiltration of eosinophils to the airways. Moreover, no change in the recruitment of neutrophils, which express the �5�1 on their cell membrane (34), was observed either. Quite remarkably, these findings suggest that treatment with RGDS prevents ASM remodeling without affecting allergic airway inflammation, although it should be noted that not all indices inflammation were examined. No effects of RGDS treatment were found on airway fibrosis either, indicating that the observed effects are probably downstream of changes in the ECM. Our findings are in line with a previous study in mice, using intraperitoneally administered anti-TGF-� antibodies during allergen-challenge, which showed that inhibition of peribronchiolar ECM deposition is associated with a normalization of ASM cell proliferation, without affecting airway inflammation in BAL or in tissue sections (50). Collectively, these findings indicate that changes in the ECM composition and deposition are important in ASM hyperplasia in vivo.

Integrins have previously been investigated as potential targets in asthma. Thus, in animal models it was shown that integrins, especially of the �4�1 subtype, are importantly involved in the recruitment of inflammatory cells to the airways (for detailed review see (34)). However, thus far phase II clinical trials with small integrin �4�1 antagonists have met limited success in affecting lung function and asthma exacerbations (51), suggesting that other strategies are warranted. Mathematical modeling studies have indicated that increased


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ASM mass is likely to be the most important abnormality responsible for increased airway resistance and airway hyperresponsiveness in asthma (52,53). Our current findings, identifying RGD-sensitive integrins as active regulators of ASM hyperplasia and hypercontractility in a guinea pig model of allergic asthma, may therefore unravel novel targets for the treatment of this disease

In conclusion, our results indicate a significant role for RGD-binding integrins in modulating allergen-induced ASM remodeling in an animal model of allergic asthma. The findings provide insight in the consequences of integrin activation on allergen-induced ASM proliferation and hyperplasia as well as increased contractile protein expression and ASM hypercontractility in the pathophysiology of chronic allergic airways disease. Based on these findings, RGD-binding integrins may represent a novel target in the treatment of airway remodeling in asthma. Acknowledgements This work was financially supported by the Netherlands Asthma Foundation, grant NAF 03.36. RG is the recipient of a Veni fellowship from the Dutch Organization for Scientific Research (916.86.036). We are grateful to Dr. W.T. Gerthoffer (University of Nevada-Reno) for preparation of the hTERT cell lines used in the study. We are grateful to the department of Surgery of the University Medical Centre Groningen for supply of the human tracheal sections. References 1. Bousquet J, Jeffery PK, Busse WW, Johnson M, Vignola AM. Asthma. From

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37. Ebina M, Takahashi T, Chiba T, Motomiya M. Cellular hypertrophy and hyperplasia of airway smooth muscles underlying bronchial asthma. A 3-D morphometric study. Am Rev Respir Dis 1993;148:720-726.

38. Bonacci JV, Harris T, Stewart AG. Impact of extracellular matrix and strain on proliferation of bovine airway smooth muscle. Clin Exp Pharmacol Physiol 2003;30:324-328.

39. Freyer AM, Johnson SR, Hall IP. Effects of growth factors and extracellular matrix on survival of human airway smooth muscle cells. Am J Respir Cell Mol Biol 2001;25:569-576.

40. Ichii T, Koyama H, Tanaka S, Kim S, Shioi A, Okuno Y, Raines EW, Iwao H, Otani S, Nishizawa Y. Fibrillar collagen specifically regulates human vascular smooth

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muscle cell genes involved in cellular responses and the pericellular matrix environment. Circ Res 2001;88:460-467.

41. Moir LM, Burgess JK, Black JL. Transforming growth factor beta(1) increases fibronectin deposition through integrin receptor alpha(5)beta(1) on human airway smooth muscle. J Allergy Clin Immunol 2008;121:1034-1039.

42. Cockcroft DW, Davis BE. Mechanisms of airway hyperresponsiveness. J Allergy Clin Immunol 2006;118:551-559.

43. Hirst SJ, Martin JG, Bonacci JV, Chan V, Fixman ED, Hamid QA, Herszberg B, Lavoie JP, McVicker CG, Moir LM and others. Proliferative aspects of airway smooth muscle. J Allergy Clin Immunol 2004;114:S2-17.

44. Meurs H, Gosens R, Zaagsma J. Airway hyperresponsiveness in asthma: lessons from in vitro model systems and animal models. Eur Respir J 2008;32:487-502.

45. Davis GE, Bayless KJ, Davis MJ, Meininger GA. Regulation of tissue injury responses by the exposure of matricryptic sites within extracellular matrix molecules. Am J Pathol 2000;156:1489-1498.

46. Black JL, Burgess JK, Johnson PR. Airway smooth muscle--its relationship to the extracellular matrix. Respir Physiol Neurobiol 2003;137:339-346.

47. Hirst SJ, Walker TR, Chilvers ER. Phenotypic diversity and molecular mechanisms of airway smooth muscle proliferation in asthma. Eur Respir J 2000;16:159-177.

48. Tran T, Halayko AJ. Extracellular matrix and airway smooth muscle interactions: a target for modulating airway wall remodelling and hyperresponsiveness? Can J Physiol Pharmacol 2007;85:666-671.

49. Ponath PD, Qin S, Ringler DJ, Clark-Lewis I, Wang J, Kassam N, Smith H, Shi X, Gonzalo JA, Newman W and others. Cloning of the human eosinophil chemoattractant, eotaxin. Expression, receptor binding, and functional properties suggest a mechanism for the selective recruitment of eosinophils. J Clin Invest 1996;97:604-612.

50. McMillan SJ, Xanthou G, Lloyd CM. Manipulation of allergen-induced airway remodeling by treatment with anti-TGF-beta antibody: effect on the Smad signaling pathway. J Immunol 2005;174:5774-5780.

51. Woodside DG, Vanderslice P. Cell adhesion antagonists: therapeutic potential in asthma and chronic obstructive pulmonary disease. BioDrugs 2008;22:85-100.

52. Lambert RK, Wiggs BR, Kuwano K, Hogg JC, Pare PD. Functional significance of increased airway smooth muscle in asthma and COPD. J Appl Physiol 1993;74:2771-2781.

53. Oliver MN, Fabry B, Marinkovic A, Mijailovich SM, Butler JP, Fredberg JJ. Airway hyperresponsiveness, remodeling, and smooth muscle mass: right answer, wrong reason? Am J Respir Cell Mol Biol 2007;37:264-272.

Bart G.J. Dekkers I. Sophie T. Bos Andrew J. Halayko Johan Zaagsma Herman Meurs.

The laminin �1-competing peptide YIGSR induces a hypercontractile, hypoproliferative airway smooth muscle phenotype in chonic asthma

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Abstract Background: Airway remodelling, including increased airway smooth muscle (ASM) mass and contractility, contributes to airway hyperresponsiveness in asthma. In vitro studies have shown that maturation of ASM cells to a (hyper)contractile phenotype is dependent on laminin, which can be inhibited by the laminin-competing peptide Tyr-Ile-Gly-Ser-Arg (YIGSR). The role of laminins in ASM remodelling in chronic asthma in vivo, however, has not yet been established. Methods: Using an established guinea pig model of allergic asthma, we investigated the effects of topical treatment of the airways with YIGSR on features of airway remodelling induced by repeated allergen challenge, including ASM hyperplasia and hypercontractility, inflammation and fibrosis. Human ASM cells were used to investigate the effect of YIGSR on ASM proliferation in vitro. Results: Topical administration of YIGSR attenuated allergen-induced ASM hyperplasia and pulmonary expression of the proliferative marker proliferating cell nuclear antigen (PCNA). Treatment with YIGSR also increased the expression of sm-MHC and ASM contractility, both in saline- and in allergen-challenged animals, suggesting that treatment with the laminin-competing peptide YIGSR mimics rather than inhibits laminin function in vivo. In addition, treatment with YIGSR increased allergen-induced fibrosis and submucosal eosinophilia. Culturing ASM cells on immobilized YIGSR in vitro concentration-dependently reduced PDGF-induced proliferation to a similar extent as with laminin. Remarkably, the effects of both immobilized YIGSR and laminin were antagonized by soluble YIGSR. Conclusion: These results indicate that the laminin-competing peptide YIGSR promotes a contractile, hypoproliferative ASM phenotype in vivo, which may depend on the microenvironment of the peptide. Background Airway inflammation, airway obstructive reactions and development of transient airway hyperresponsiveness are primary features of acute asthma [1,2]. In addition, structural changes in the airway wall are thought to contribute to a decline of lung function and development of persistent airway hyperresponsiveness in chronic asthma [1,3]. These structural changes include goblet cell metaplasia and mucous gland hyperplasia, increased vascularity, altered deposition of the extracellular matrix (ECM) proteins and accumulation of contractile airway smooth muscle (ASM) cells [1,4-6](Chapter 2). ASM cells may importantly contribute to airway remodelling as they retain the ability for reversible phenotypic plasticity, enabling them to switch between contractile, proliferative, migratory and synthetic states [7,8]. In vitro, switching to a proliferative phenotype results from exposure of ASM cells to mitogenic stimuli,

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leading to increased proliferative activity and decreased contractile function [9,10](Chapter 3). Removal of growth factors, for example by serum deprivation in the presence of insulin, results in maturation of the cells to a contractile phenotype, characterized by increased expression of contractile protein markers, increased contractile function and increased expression of laminin �2, �1 and �1 chains [7,11,12](Chapter 5).

Laminins are basement membrane ECM components composed of heterotrimers of �, � and � chains. Five laminin �-, three �- and three �-chains have been identified in mammals, which form at least 15 different laminin isoforms [13]. Various laminin chains are expressed in the lung and expression appears to be tissue- and developmental stage-dependent [14]. In adult asthmatics, expression of laminin �2 and �2 chains in the airways is increased [15,16]. In addition, asthmatics with compromised epithelial integrity show increased laminin �2 chain expression in the airways [16].

Studies on the function of laminins have shown that laminins are essential for lung development and ASM function. Laminin �1 and �2 chains are required for pulmonary branching and differentiation of naïve mesenchymal cells into ASM [13,17,18]. Primary ASM cells cultured on laminin-111 (laminin-1) are retained in a hypoproliferative phenotype, associated with high expression levels of contractile proteins [19]. This is of functional relevance as the induction of a hypocontractile ASM phenotype by PDGF was prevented by co-incubation with laminin-111 (Chapter 3). Increased expression of endogenous laminin-211 (laminin-2) has been found to be essential for ASM cell maturation [12], and studies from our laboratory have shown that laminin-211 is essential for the induction of a hypercontractile, hypoproliferative ASM phenotype by prolonged insulin exposure (Chapter 5).

Recently, we have shown that in an animal model of chronic allergic asthma ASM remodelling can be inhibited by the integrin-blocking peptide Arg-Gly-Asp-Ser (RGDS) (Chapter 6), containing the RGD-binding motif present in ECM proteins like fibronectin, collagens and laminins [20,21]. The specific role of laminins in ASM remodelling in vivo, however, remains to be determined. Therefore, using a guinea pig model of chronic asthma, we explored the role of laminins in ASM remodelling in vivo, by treating the animals with the specific laminin-competing peptide Tyr-Ile-Gly-Ser-Arg (YIGSR), a binding motif present in the �1 chain of laminins [22].

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Methods Animals All protocols described in this study were approved by the University of Groningen Committee for Animal Experimentation. Outbred, male, specified pathogen-free Dunkin Hartley guinea pigs (Harlan, Heathfield, UK) weighing 150–250 g were sensitized to ovalbumin (Sigma Chemical Co., St. Louis, MO, USA), using Al(OH)3 as adjuvant, as described previously [23]. In short, 0.5 ml of an allergen solution containing 100 �g/ml ovalbumin and 100 mg/ml Al(OH)3 in saline was injected intraperitoneally, while another 0.5 ml was divided over seven intracutaneous injection sites in the proximity of lymph nodes in the paws, lumbar regions and the neck. The animals were group-housed in cages in climate controlled animal quarters and given water and food ad libitum, while a 12-hour on/ 12-hour off light cycle was maintained. Provocation Procedures Four weeks after sensitization, allergen-provocations were performed by inhalation of aerosolized solutions of saline (control) or ovalbumin as described previously [23]. Aerosols were produced by a DeVilbiss nebulizer (type 646, DeVilbiss, Somerset, PA, USA). Provocations were carried out in a specially designed Perspex cage (internal volume 9 L), in which the guinea pigs could move freely. Before the start of the experimental protocol, the animals were habituated to the provocation procedures. After an adaptation period of 30 min, three consecutive provocations with saline were performed, each provocation lasting 3 min, separated by 7 min intervals. Ovalbumin challenges were performed by inhalation of increasing concentrations of ovalbumin (0.5, 1.0, or 3.0 mg/ml) in saline. Allergen inhalations were discontinued when the first signs of respiratory distress were observed. No anti-histaminic was needed to prevent the development of anaphylactic shock. Study design Guinea pigs were challenged with either saline or ovalbumin once weekly for 12 consecutive weeks, as described previously [24,25](Chapter 6). Animals were treated with saline or YIGSR (Calbiochem, Nottingham, UK) by intranasal instillation (2.5 mM, 200 �l), 0.5 hr prior to and 5.5 hr after each challenge with saline or ovalbumin. Treatment groups were as follows: saline-treated, saline-challenged controls (n=6); YIGSR-treated, saline-challenged animals (n=5); saline-treated, ovalbumin-challenged animals (n=7) and YIGSR-treated, ovalbumin-challenged animals (n=7). Data for the saline-treated animals (controls) have been published previously as part of a simultaneous parallel study (Chapter 6). During the 12-week challenge protocol, guinea pig weight was monitored weekly and no differences in weight gain between different treatment groups were found


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Tissue acquisition Guinea pigs were sacrificed by experimental concussion, followed by rapid exsanguination 24 h after the last challenge. The lungs were immediately resected and kept on ice for further processing. The trachea was removed and transferred to a Krebs–Henseleit (KH) buffer of the following composition (mM): 117.5 NaCl, 5.60 KCl, 1.18 MgSO4, 2.50 CaCl2, 1.28 NaH2PO4, 25.00 NaHCO3, and 5.50 glucose, pregassed with 5% CO2 and 95% O2, pH 7.4 at 37°C. Lungs were divided into three parts and weighed. One part was snap frozen in liquid nitrogen for the measurement of hydroxyproline content. One part was frozen at -80 ºC in isopentane and stored at -80 ºC for histological purposes. The remaining part was snap frozen in liquid nitrogen and stored at -80 ºC to be used for Western analysis. Isometric tension measurements Isometric contraction experiments were performed as described previously [24,25](Chapter 6). Briefly, the trachea was prepared free of connective tissue. Single open-ring, epithelium-denuded preparations were mounted for isometric recording in organ baths, containing KH buffer at 37 °C, continuously gassed with 5% CO2 and 95% O2, pH 7.4. During a 90-min equilibration period, resting tension was gradually adjusted to 0.5 g. Subsequently, muscle strips were precontracted with 20 mM and 40 mM KCl. Following washouts, maximal relaxation was established by the addition of 0.1 �M (-)-isoproterenol (Sigma). After washout and another 30 min equilibration period, cumulative concentration-response curves were constructed using stepwise increasing concentrations of KCl (5.6�50 mM) or methacholine (1 nM � 0.1 mM). When maximal tension was reached, the strips were washed several times and maximal relaxation was established using 10 �M (-)-isoproterenol. Histochemistry Immunohistochemistry was performed as described previously [24,25](Chapter 6). Transverse cross-sections (8 �m) of the main bronchi from both right and left lung lobes were used for morphometric analyses. To identify smooth muscle, the sections were stained for smooth-muscle-specific myosin heavy chain (sm-MHC). Sections were dried, fixed with acetone and washed in phosphate-buffered saline (PBS). Subsequently, sections were incubated for 1 h in PBS supplemented with 1% bovine serum albumin (BSA, Sigma) and anti-sm-MHC (diluted 1:100, Neomarkers, Fremont, CA, USA) at room temperature. Sections were then washed with PBS, after which endogenous peroxidase activity was blocked by treatment with PBS containing 0.075% H2O2 for 30 min. Sections were washed with PBS, after which the horseradish peroxidase (HRP)-linked secondary antibody (rabbit anti-mouse IgG, Sigma, diluted 1:200) was applied for 30 min at room temperature. After another three washes, sections were incubated with diaminobenzidine (1 mg/ml) for 5 min in the dark, after which sections were washed and stained with haematoxylin. After rinsing with water

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the sections were embedded in Kaisers glycerol gelatin. Airways within sections were digitally photographed and subclassified as cartilaginous or non-cartilaginous. All immunohistochemical measurements were carried out digitally, using quantification software (ImageJ). For this purpose, digital photographs of lung sections were analyzed at a magnification of 40-100x. For both types of airways, sm-MHC positive areas were measured by a single observer in a blinded fashion. In addition, haematoxylin-stained nuclei within the ASM bundle were counted. Of each animal, 4 lung sections were prepared per immunohistochemical staining, in which a total of 4 to 5 airways of each classification were analyzed. Eosinophils were identified in haematoxylin-and-eosin-stained lung sections. Western analysis Lung homogenates were prepared as described previously [24,25](Chapter 6). Equal amounts of protein were subjected to electrophoresis and transferred onto nitrocellulose membranes, followed by immunoblotting for sm-MHC and PCNA (Neomarkers), using standard techniques. Antibodies were visualized on film using enhanced chemiluminescence reagents (Pierce, Rockford, IL, USA) and analyzed by densitometry (TotallabTM, Nonlinear dynamics, Newcastle, UK). All bands were normalized to �-actin expression. Hydroxyproline assay Lungs were analyzed for hydroxyproline, an estimate of collagen content, as described previously (Chapter 6). In short, total lung homogenates were prepared by pulverizing tissue under liquid nitrogen and sonification in PBS. Homogenates were incubated with 1,25 ml 5% trichloroacetic acid on ice for 20 min, after which the samples were centrifuged. The pellet was resuspended in 12 N hydrochloric acid (10 ml) and heated overnight at 110 °C. The samples were dissolved in 2 ml water by incubating for 72 h at room temperature. To determine hydroxyproline concentrations, samples were incubated with 100 �l chloramine T (1.4% chloramine T in 0.5 M sodium acetate/10% isopropanol) for 30 min at room temperature. Next, 100 �l Ehrlich’s solution (1.0 M 4-dimethylaminobenzaldehyde in 70% isopropanol/30% perchloric acid) was added and samples were incubated at 65 °C for 30 min. Samples were cooled to room temperature and hydroxyproline concentrations were quantified by colorimetric measurement (550 nm, Biorad 680 plate reader). Cell culture Three human bronchial smooth muscle cell lines, immortalized by stable expression of human telomerase reverse transcriptase (hTERT), were used for all experiments. The primary cells used to generate each cell line were prepared as we have described previously [26-28]. All procedures were approved by the Human Research Ethics Board of the University of Manitoba. For all experiments, passages 26–34 myocytes grown on uncoated plastic dishes in Dulbecco's


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Modified Eagle's Medium (DMEM, Gibco BRL Life Technologies, Paisley, U.K.) supplemented with 50 U/ml streptomycin, 50 μg/ml penicillin, (Gibco) and 10%

vol/vol Foetal Bovine Serum (FBS, Gibco) were used. Coating of culture plates with laminin and integrin-blocking peptides Dilutions of mouse Engelberth-Holm-Swarm (EHS) laminin-111 (10 �g/ml, Invitrogen, Grand Island, NY, USA), YIGSR (1-100 μM), Arg-Gly-Asp-Ser (RGDS, 100 �M, Calbiochem) and Gly-Arg-Ala-Asp-Ser-Pro (GRADSP, 100 �M, Calbiochem) were prepared in PBS and absorbed to 24-well culture plates overnight. Unoccupied protein-binding sites were blocked by a 30-min incubation with 0.1% BSA in PBS. Subsequently, plates were washed twice with plain DMEM and dried before further use. [3H]-Thymidine incorporation Cells in DMEM supplemented with streptomycin, penicillin and 10% FBS were plated on uncoated or coated 24-well culture plates at a density of 20,000 cells per well and allowed to attach overnight. Subsequently, cells were maintained in serum-free DMEM supplemented with antibiotics and 1% ITS (Insulin, Transferrin and Selenium, Gibco) for 3 days. Cells were then incubated with or without PDGF-AB (10 ng/ml, human, Bachem, Weil am Rhein, Germany) for 28 h, the last 24 h in the presence of [methyl-3H]-thymidine (0.25 μCi/ml) in DMEM supplemented with antibiotics. After incubation, the cells were washed twice with 0.5 ml PBS at room temperature. Subsequently, the cells were treated with 0.5 ml ice-cold 5% trichloroacetic acid on ice for 30 min, and the acid-insoluble fraction was dissolved in 1 ml NaOH (1 M). Incorporated [3H]-thymidine was quantified by liquid-scintillation counting using a Beckman LS1701 �-counter. Statistics All data represent means ± SEM from n separate experiments. Statistical significance of differences was evaluated using one-way ANOVA, followed by a Newman–Keuls multiple comparisons test. Differences were considered to be statistically significant when P<0.05.

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Results The laminin �1-competing peptide YIGSR inhibits allergen-induced ASM accumulation in a guinea pig model of chronic allergic asthma. In our guinea pig model, repeated ovalbumin-challenge increased the sm-MHC-positive area, predominantly in the cartilaginous airways by 1.9±0.1-fold (P<0.001) compared to saline-treated, saline-challenged controls (Figure 1A). Topical treatment of the airways with intranasally instilled YIGSR 0.5 h prior to and 5.5 h after each ovalbumin-challenge almost fully inhibited the ovalbumin-induced increase in ASM area (by 96±3%, P<0.001). No significant effect of YIGSR treatment was observed in saline-challenged animals.

To determine whether the changes in ASM content were associated with changes in cell number and/or cell size, the number nuclei within the ASM layer were counted and expressed relative to total ASM area. Repeated ovalbumin challenge did not change the number of nuclei per mm2 of smooth muscle area (Figure 1B), indicating that the cell size is unchanged and ovalbumin-induced increases in ASM mass were caused by an increased cell number. YIGSR treatment did not change ASM cell size in saline-challenged animals; however, a small, but significant (P<0.05) decrease in the number of nuclei/mm2 was observed in ovalbumin-challenged animals (Figure 1B), suggesting a slight increase in cell size.

To assess whether the changes in ASM area were associated with changes in proliferative responses, Western analysis was used to determine expression of the proliferative marker PCNA in whole lung homogenates. After repeated ovalbumin-challenge, a considerable increase (4.2±0.2-fold, P<0.001) in PCNA was observed compared to saline-treated, saline-challenged controls (Figure 1C). Treatment with YIGSR fully normalized the ovalbumin-induced increase in PCNA, when compared to saline-challenged controls (P<0.001). In the saline-challenged animals, no significant effect of YIGSR treatment on PCNA expression was observed. Unfortunately, specific characterization of the proliferating cells in guinea pig lung sections by immunohistochemistry was not possible with the antibody used. Collectively, these in vivo data indicate that YIGSR treatment inhibits allergen-induced ASM hyperplasia as well as proliferative responses that may underlie ASM hyperplasia.


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Figure 1: Increased ASM mass after repeated allergen challenge in vivo is inhibited by topical treatment with YIGSR. To assess the role of laminins in increased ASM mass in asthma, the effects of treatment with YIGSR were evaluated in a guinea pig model of chronic allergic asthma. (A) Treatment with YIGSR fully inhibited ovalbumin-induced increase in sm-MHC positive area in cartilaginous airways. (B) Changes in ASM mass were mainly dependent on changes in ASM cell number, only a small increase in cell size was observed for the YIGSR-treated, ovalbumin-challenged animals. (C) Increased pulmonary expression of the proliferative marker PCNA after repeated ovalbumin-challenges, was almost fully reversed by YIGSR. Representative blots of PCNA and �-actin are shown. No effects of YIGSR were shown in saline-challenged animals for any of the parameters. *P<0.05, ***P<0.001 compared to saline-treated, saline-challenged controls. ###P<0.001 compared to saline-treated, ovalbumin-challenged controls. Data represent means ± SEM of 5-7 animals. YIGSR treatment increases contractile protein accumulation and ASM contractility. Previously, we have shown that repeated ovalbumin-exposure increased maximal methacholine- and KCl-induced isometric contractions of epithelium-denuded, tracheal smooth muscle preparations ex vivo [24,25](Chapter 6). Interestingly, treatment with the YIGSR peptide augmented the ovalbumin-induced increases in maximal methacholine- and KCl-induced contractions further (1.33±0.08-fold (P<0.001) and 1.28±0.11-fold (P<0.05), respectively, compared to saline-treated, ovalbumin-challenged controls, Figure 2A, Table 1). Similarly, in saline-challenged animals YIGSR treatment increased methacholine- and KCl-induced contractions (1.29±0.03-fold and 1.39±0.04-fold (P<0.05), respectively, compared to saline-treated, saline-challenged animals).

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Previously, we have found that increased ASM contractility induced by repeated ovalbumin challenge is associated with increased pulmonary sm-MHC expression [24,25](Chapter 6). In saline-treated animals, repeated ovalbumin-challenge increased sm-MHC by 2.5±0.1-fold compared to saline-challenged controls (P<0.001, Figure 2B). In line with the increased methacholine- and KCl-induced contractions, treatment with YIGSR increased pulmonary sm-MHC expression in saline-challenged animals (2.40±0.28-fold, P<0.001), whereas in ovalbumin-challenged animals the increase in sm-MHC was increased further (1.37±0.08-fold compared to ovalbumin-challenged controls, P<0.01). Collectively, these data indicate that in vivo treatment with the laminin-competing peptide YIGSR increases ASM contractility and contractile protein expression both in saline- and allergen-challenged animals. Figure 2: Topical treatment of the airways with YIGSR increases ASM contractility and contractile protein accumulation. (A) Treatment with YIGSR enhanced the maximal methacholine-induced isometric contraction of epithelium-denuded tracheal smooth muscle preparations both in saline- and in ovalbumin-challenged animals. (B) Treatment with YIGSR increased pulmonary expression of sm-MHC, both in saline- and in ovalbumin-challenged animals. Representative blots of sm-MHC and �-actin are shown. ***P<0.001 compared to saline-treated, saline-challenged controls. ##P<0.01 compared to saline-treated, ovalbumin-challenged controls. Data represent means ± SEM of 5-7 animals.


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Table 1: Contractile responses of epithelium-denuded, tracheal smooth muscle preparations after repeated saline or ovalbumin challenge of saline- or YIGSR-treated guinea pigs.

Methacholine KCl Treatment Challenge

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Saline Saline 1.42±0.09 6.55±0.18 1.02±0.06 23.7±0.9 6 YIGSR Saline 1.84±0.04 6.82±0.13 1.41±0.04* 20.4±2.2 5 Saline Ovalbumin 2.33±0.22*** 6.28±0.11 1.73±0.13** 23.7±1.2 7 YIGSR Ovalbumin 3.11±0.18***, ### 6.61±0.08 2.12±0.19***,# 24.5±1.1 7

Data represent means ± SEM. Abbreviations: Emax: maximal contractile effect; EC50: concentration of the stimulus eliciting half-maximal response; pEC50: negative logarithm of the EC50 value. *P<0.05, **P<0.01, ***P<0.001 compared to saline-treated, saline-challenged animals. #P<0.05, ###P<0.001 compared to saline-treated, ovalbumin-challenged animals. Effects of YIGSR treatment on allergen-induced airway inflammation. Infiltration of eosinophils into the airways is a characteristic feature of allergic asthma and is generally considered to contribute to airway remodelling [2]. As observed previously [25](Chapter 6), repeated ovalbumin challenge increased the number of eosinophils in the submucosal and adventitial compartments of the airways (P<0.001 both, Figures 3A and 3B). No significant effect of YIGSR on the increased adventitial eosinophil number after ovalbumin-challenge was observed (Figure 3B). Treatment with YIGSR did not affect eosinophil numbers in the adventitial compartment of saline-challenged animals. Remarkably, however, treatment with YIGSR significantly increased the number of eosinophils in the submucosal airway compartment after repeated allergen challenge (P<0.05, Figure 3A). Similarly, although not statistically significant, a increase in eosinophil numbers in saline-challenged animals was observed.

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A BSubmucosa Adventitia Figure 3: YIGSR treatment increases allergen-induced eosinophilic inflammation in the submucosal airway compartment. (A) Ovalbumin-induced eosinophil numbers in the submucosal compartment are increased by YIGSR treatment. (B) YIGSR treatment does not affect eosinophilic cell number in the adventitial compartment. No effects of YIGSR were found in saline-challenged animals for any of the conditions. ***P<0.001 compared to saline-treated, saline-challenged controls. #P<0.05 compared to saline-treated, ovalbumin-challenged animals. Data represent means ± SEM of 5-7 animals. Effects of YIGSR treatment on allergen-induced fibrosis. Aberrant deposition of ECM proteins, including collagens, in the airway wall is another characteristic feature of chronic asthma [29,30]. In line with previous studies (Chapter 6), we demonstrated that lung hydroxyproline content, as an estimate of collagen, is increased after repeated ovalbumin challenge (P<0.001, Figure 4). Treatment with YIGSR of the ovalbumin-challenged animals further augmented the hydroxyproline content (P<0.01). YIGSR did not change the hydroxyproline content in saline-challenged animals. Collectively, these findings indicate that YIGSR treatment increases allergen-induced submucosal airway eosinophilia as well as collagen deposition in the lung.


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Immobilized YIGSR inhibits ASM cell proliferation in vitro. Paradoxically, previous in vitro studies have indicated that soluble YIGSR inhibits ASM cell maturation and the development of a hypercontractile, hypoproliferative ASM phenotype induced by insulin [12](Chapter 5). To further investigate this apparent paradox, the effects of immobilized and soluble YIGSR on basal and growth factor-induced ASM cell proliferation in vitro were compared. First, human ASM cells were cultured on 24 well plates coated with increasing concentrations of YIGSR (1-100 μM) and stimulated with PDGF (10 ng/ml). Surprisingly, culturing the cells on immobilized YIGSR concentration-dependently inhibited PDGF-induced DNA synthesis compared to culturing on uncoated plates (Figure 5A). No effect of YIGSR was observed on basal DNA synthesis. Similar effects were observed for cell number (data not shown). By contrast, culturing on immobilized RGDS (100 �M) or its negative control Gly-Arg-Ala-Asp-Ser-Pro (GRADSP, 100 �M) did not affect basal or PDGF-induced proliferation (Figure 5B).

To assess the effects of soluble YIGSR on proliferative responses of human ASM, cells were cultured on immobilized laminin-111 (10 �g/ml) or YIGSR (100 �M). Subsequently, the cells were stimulated with vehicle or PDGF in the absence or presence of soluble YIGSR. As observed previously (Chapter 3 & 5), we found that culturing on laminin-111 inhibited PDGF-induced DNA-synthesis (by 56±11%, P<0.05, Figure 5C). This inhibitory effect was fully normalized by soluble YIGSR. Surprisingly, the inhibitory effect of coated YIGSR on PDGF-induced proliferation was also fully normalized by soluble YIGSR. Similar results were obtained for cell number (data not shown). Collectively, these results indicate that the effects of the laminin-competing peptide YIGSR on ASM proliferative responses may depend on the microenvironment of the peptide.

Figure 4: YIGSR treatment increases allergen-induced fibrosis in the guinea pig lung. Hydroxyproline content in guinea pig lung after repeated saline- or ovalbumin-challenges in saline- and YIGSR-treated animals. ***P<0.001 compared to saline-treated, saline-challenged controls. ##P<0.01 compared to saline-treated, ovalbumin-challenged animals. Data represent means ± SEM of 5-7 animals.

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Figure 5: Effects of immobilized and soluble YIGSR on basal and PDGF-induced human ASM cell proliferation. (A) Culturing of human ASM cells on immobilized YIGSR matrices inhibits PDGF-induced thymidine-incorporation in a YIGSR concentration-dependent fashion. Under unstimulated (Basal) conditions, no effects of immobilized YIGSR were observed. (B) Immobilized RGDS or its negative control GRADSP did not affect basal or PDGF-induced thymidine-incorporation. (C) The inhibitory effects of immobilized laminin-111 and YIGSR matrices on PDGF-induced thymidine-incorporation were normalized by soluble YIGSR. ***P<0.001 compared to thymidine-incorporation of unstimulated cells (basal) cultured on uncoated matrices (plastic). #P<0.05 and ##P<0.01 compared to PDGF-induced thymidine-incorporation of cells cultured on uncoated matrices. Data represent means ± SEM of 4-5 independent experiments of 3 different donors, performed in duplicate.


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Discussion In the current study, we demonstrate that treatment with the laminin �1 chain-competing peptide YIGSR promotes the formation of a hypercontractile, hypoproliferative ASM phenotype in an animal model of chronic asthma. Thus, it was shown that topical application of YIGSR to the airways inhibited ASM hyperplasia induced by repeated allergen challenge, whereas ASM contractility and contractile protein expression were increased, both under basal and under allergen-challenged conditions. Apparently, these results are in contrast to previous in vitro studies, demonstrating that YIGSR inhibited maturation of human ASM cells to a hypercontractile, hypoproliferative ASM phenotype [12](Chapter 5).

Accumulation of ASM in the airway wall is a characteristic feature of asthma, which may be due to an increase in cell number (hyperplasia) [31,32] as well as an increase in cell size (hypertrophy) [31,33]. Switching of the ASM phenotype from a contractile to a proliferative state is thought to contribute to the increased ASM mass in asthma [8]. In support, various mitogenic stimuli, including growth factors and ECM proteins, have been shown to induce a proliferative ASM phenotype in vitro [9,10](Chapter 3), which may be inhibited by culturing the cells on immobilized laminin-111 [19](Chapter 3 & 6) or by endogenously produced laminin-211 (Chapter 5). These inhibitory effects are reversed, when cells are cultured in the presence of soluble YIGSR (Chapter 5), a binding motif present in the laminin �1 chain [22]. In accordance with these findings, we found that culturing human ASM cells on laminin-111 reduced proliferation of these cells induced by PDGF, which effect was fully normalized in the presence of soluble YIGSR. Surprisingly, although culturing of human ASM cells on immobilized YIGSR had no effect on basal proliferation, growth factor-induced proliferation of these cells was concentration-dependently inhibited on this matrix, to a similar extent as with laminin-111. These effects were specific for the YIGSR peptide, since culturing on RGDS, containing the RGD binding motif found in fibronectin, collagens as well as laminins [20,21], or its negative control GRADSP did not affect basal or PDGF-induced proliferation. Our observations are in agreement with previous findings, showing that immobilized YIGSR promoted attachment of various cells to a similar extent as laminin-111 [22,34,35], whereas addition of soluble YIGSR to the culture medium blocked the attachment to laminin-111 [34] or matrigel [35]. In our study, addition of soluble YIGSR to the culture medium also normalized the effects of immobilized YIGSR, which corresponds to previous findings in alveolar cells, using the Ser-Ile-Asn-Asn-Asn-Arg (SINNNR) sequence derived from the laminin � chain [36]. Collectively, these findings suggest that the laminin-competing peptide YIGSR may either promote or inhibit ASM proliferative responses, depending on the microenvironment of the peptide. The mechanisms underlying these differential effects are currently unknown. However, our results, as that of others [39], may suggest that bridging of the 67 kDa laminin receptor LAMR1, that has high

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affinity to the YIGSR binding epitope [37], could be important. Previous studies have found similar effects for fibronectin and the �5�1 integrin, showing that monovalent ligand binding to this integrin resulted in the targeting of the receptor to the focal contacts, whereas multivalent ligand binding resulted in reorganization of the cytoskeleton and activation of protein tyrosine phosphorylation [38].

In addition to ASM accumulation, increased expression of expression of contractile proteins and ASM contractility, as well as increased ECM deposition, including laminins, are also characteristic features of airway remodelling in asthma (Chapter 2). Thus, in the airways of asthmatics, increased expression of laminin �2 and �2 chains has been observed [15,16], whereas expression of laminin �2 chains inversely correlated with epithelial integrity [16]. Laminins have not only been shown to inhibit ASM proliferation, but also to be critically involved in both the maintenance and induction of a (hyper)contractile ASM phenotype. Thus, culturing of ASM cells on a laminin-111 matrix inhibits proliferation and maintains contractile protein expression in the presence of various growth factors [19], and prevents the induction of a functionally hypocontractile ASM phenotype by PDGF (Chapter 3). Induction of a contractile ASM phenotype by serum deprivation is associated by increased expression of laminin �2, �1 and �1 chains, all found in the laminin-211 isoform [12]. Similarly, the induction of a hypercontractile, hypoproliferative ASM phenotype by insulin is associated with increased laminin �2, �1 and �1 chain expression (Chapter 5). The increased expression of endogenous laminin is required for the observed ASM phenotype maturation, as the laminin competing peptides YIGSR, GRGDSP and RGDS prevented the induction of contractile protein expression and hypercontractility [12](Chapter 5 & 6). Recently, using the same guinea pig model of chronic asthma, we have shown that in vivo treatment with the RGD-containing peptide RGDS largely inhibits allergen-induced ASM hyperplasia and hypercontractility (Chapter 6). The RGD sequence, however, can be found in several ECM proteins [20,21], and the specific contribution of laminins to ASM remodelling in chronic asthma, is unknown thus far. In present study we found that in vivo treatment with YIGSR inhibited allergen-induced ASM hyperplasia, but increased both the expression of sm-MHC and ASM contractility in allergen-challenged as well as in control animals. In addition, a small increase in cell size in the allergen-challenged YIGSR treated animals was observed suggesting that hypertrophy may also have played a role in the observed effects. Collectively, our results indicate that treatment with YIGSR inhibits allergen-induced ASM hyperplasia and increases ASM contractility in vivo, suggesting that YIGSR mimics rather than inhibits laminin function under this condition.

Previous studies have shown that treatment with YIGSR inhibits tumour growth [39,40], which may involve blockade of the laminin receptor on endothelial cells, leading to inhibition of angiogenesis and thereby limiting the blood supply to the tumour [35]. This mechanism may also have contributed to the effects of YIGSR described in the present study, as the allergen-induced


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increase in sm-MHC positive microvasculature in the airway wall (1.33±0.17-fold increase, P<0.05 compared saline-treated, saline-challenged controls, data not shown), an indication for the accumulation of stabilized blood vessels [41], was significantly inhibited by YIGSR treatment (90±27% reduction, P<0.05, data not shown). The potential role of angiogenesis in increased ASM growth and contractility in asthma, however, is completely unknown and would warrant further investigation.

Eosinophils express a number of integrins, of which the �6�1 integrin mediates adhesion to laminin, but not to collagen type I or type IV [42,43]. Eosinophils isolated from allergic donors also showed a higher adhesion to laminin than those isolated from healthy subjects [43]. In addition, migration of eosinophils through matrigel, a basement membrane extract containing laminin-111, required interaction with �1-integrins [43]. These findings suggest that laminin-competing peptides could affect allergen-induced infiltration of inflammatory cells to the airways. Thus far, no reports on the effects of YIGSR on eosinophil migration are available. In the current study, we observed that YIGSR increased allergen-induced eosinophil cell numbers in the submucosal compartment, without affecting eosinophil numbers in the adventitial compartment. The increased number of eosinophils in the submucosal compartment suggests that, rather than, infiltration, retention time of the eosinophils in the compartment is increased.

Increased and altered deposition of ECM proteins, including laminins and collagens, in the airway wall is another characteristic feature of remodelling in chronic asthma [29,30]. The effects of laminins on the deposition of collagens and other ECM components by fibroblasts and other structural cells is currently unknown. However, increased ECM deposition may be secondary to prolonged airway inflammation [2] and therefore increased allergen-induced airway fibrosis in YIGSR-treated animals could also indirectly result from increased eosinophilia. Conclusions Our results indicate that the laminin-competing peptide YIGSR promotes a contractile, hypoproliferative ASM phenotype in vivo, which may depend on the microenvironment of the peptide. In addition, treatment with YIGSR increased allergen-induced fibrosis and submucosal eosinophilia. Collectively, our data indicate that YIGSR mimics rather than competes with laminin function in vivo. Competing interests The authors declare that they have no competing interests

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Authors’ contributions BGJD: design of the study, acquisition of data, data analysis and interpretation, manuscript writing; ISTB: design of the study, acquisition of data, data analysis and interpretation; AJH: preparation of ASM cell lines and critical revision of the MS; JZ: design of the study, data interpretation and critical revision of the MS; HM: design of the study, data interpretation and critical revision of the MS. All authors have read and approved the manuscript. Acknowledgements This work was financially supported by the Netherlands Asthma Foundation, grant NAF 3.2.03.36. We are grateful to Dr. W.T. Gerthoffer (University of Nevada-Reno) for preparation of the hTERT cell lines used in the study. References 1. Bousquet J, Jeffery PK, Busse WW, Johnson M, Vignola AM: Asthma. From

bronchoconstriction to airways inflammation and remodeling. Am J Respir Crit Care Med 2000, 161:1720-1745.

2. Cockcroft DW, Davis BE: Mechanisms of airway hyperresponsiveness. J Allergy Clin Immunol 2006, 118:551-559.

3. Davies DE, Wicks J, Powell RM, Puddicombe SM, Holgate ST: Airway remodeling in asthma: new insights. J Allergy Clin Immunol 2003, 111:215-225.

4. Dunnill MS, Massarella GR, Anderson JA: A comparison of the quantitative anatomy of the bronchi in normal subjects, in status asthmaticus, in chronic bronchitis, and in emphysema. Thorax 1969, 24:176-179.

5. Parameswaran K, Willems-Widyastuti A, Alagappan VK, Radford K, Kranenburg AR, Sharma HS: Role of extracellular matrix and its regulators in human airway smooth muscle biology. Cell Biochem Biophys 2006, 44:139-146.

6. Fernandes DJ, Bonacci JV, Stewart AG: Extracellular matrix, integrins, and mesenchymal cell function in the airways. Curr Drug Targets 2006, 7:567-577.

7. Halayko AJ, Salari H, Ma X, Stephens NL: Markers of airway smooth muscle cell phenotype. Am J Physiol 1996, 270:L1040-L1051.

8. Halayko AJ, Tran T, Ji SY, Yamasaki A, Gosens R: Airway smooth muscle phenotype and function: interactions with current asthma therapies. Curr Drug Targets 2006, 7:525-540.

9. Gosens R, Meurs H, Bromhaar MM, McKay S, Nelemans SA, Zaagsma J: Functional characterization of serum- and growth factor-induced phenotypic changes in intact bovine tracheal smooth muscle. Br J Pharmacol 2002, 137:459-466.

10. Gosens R, Roscioni SS, Dekkers BG, Pera T, Schmidt M, Schaafsma D, Zaagsma J, Meurs H: Pharmacology of airway smooth muscle proliferation. Eur J Pharmacol 2008, 585:385-397.

11. Schaafsma D, McNeill KD, Stelmack GL, Gosens R, Baarsma HA, Dekkers BG, Frohwerk E, Penninks JM, Sharma P, Ens KM et al.: Insulin increases the


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12. Tran T, McNeill KD, Gerthoffer WT, Unruh H, Halayko AJ: Endogenous laminin is required for human airway smooth muscle cell maturation. Respir Res 2006, 7:117.

13. Nguyen NM, Senior RM: Laminin isoforms and lung development: all isoforms are not equal. Dev Biol 2006, 294:271-279.

14. Virtanen I, Laitinen A, Tani T, Paakko P, Laitinen LA, Burgeson RE, Lehto VP: Differential expression of laminins and their integrin receptors in developing and adult human lung. Am J Respir Cell Mol Biol 1996, 15:184-196.

15. Altraja A, Laitinen A, Virtanen I, Kampe M, Simonsson BG, Karlsson SE, Hakansson L, Venge P, Sillastu H, Laitinen LA: Expression of laminins in the airways in various types of asthmatic patients: a morphometric study. Am J Respir Cell Mol Biol 1996, 15:482-488.

16. Amin K, Janson C, Seveus L, Miyazaki K, Virtanen I, Venge P: Uncoordinated production of Laminin-5 chains in airways epithelium of allergic asthmatics. Respir Res 2005, 6:110.

17. Schuger L, Skubitz AP, Zhang J, Sorokin L, He L: Laminin alpha1 chain synthesis in the mouse developing lung: requirement for epithelial-mesenchymal contact and possible role in bronchial smooth muscle development. J Cell Biol 1997, 139:553-562.

18. Relan NK, Yang Y, Beqaj S, Miner JH, Schuger L: Cell elongation induces laminin alpha2 chain expression in mouse embryonic mesenchymal cells: role in visceral myogenesis. J Cell Biol 1999, 147:1341-1350.

19. Hirst SJ, Twort CH, Lee TH: Differential effects of extracellular matrix proteins on human airway smooth muscle cell proliferation and phenotype. Am J Respir Cell Mol Biol 2000, 23:335-344.

20. Plow EF, Haas TA, Zhang L, Loftus J, Smith JW: Ligand binding to integrins. J Biol Chem 2000, 275:21785-21788.

21. Aumailley M, Gerl M, Sonnenberg A, Deutzmann R, Timpl R: Identification of the Arg-Gly-Asp sequence in laminin A chain as a latent cell-binding site being exposed in fragment P1. FEBS Lett 1990, 262:82-86.

22. Graf J, Ogle RC, Robey FA, Sasaki M, Martin GR, Yamada Y, Kleinman HK: A pentapeptide from the laminin B1 chain mediates cell adhesion and binds the 67,000 laminin receptor. Biochemistry 1987, 26:6896-6900.

23. Meurs H, Santing RE, Remie R, van der Mark TW, Westerhof FJ, Zuidhof AB, Bos IS, Zaagsma J: A guinea pig model of acute and chronic asthma using permanently instrumented and unrestrained animals. Nat Protoc 2006, 1:840-847.

24. Gosens R, Bos IS, Zaagsma J, Meurs H: Protective effects of tiotropium bromide in the progression of airway smooth muscle remodeling. Am J Respir Crit Care Med 2005, 171:1096-1102.

25. Bos IS, Gosens R, Zuidhof AB, Schaafsma D, Halayko AJ, Meurs H, Zaagsma J: Inhibition of allergen-induced airway remodelling by tiotropium and budesonide: a comparison. Eur Respir J 2007, 30:653-661.

26. Gosens R, Stelmack GL, Dueck G, McNeill KD, Yamasaki A, Gerthoffer WT, Unruh H, Gounni AS, Zaagsma J, Halayko AJ: Role of caveolin-1 in p42/p44 MAP kinase activation and proliferation of human airway smooth muscle. Am J Physiol Lung Cell Mol Physiol 2006, 291:L523-L534.

27. Gosens R, Dueck G, Gerthoffer WT, Unruh H, Zaagsma J, Meurs H, Halayko AJ: p42/p44 MAP kinase activation is localized to caveolae-free membrane domains in

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airway smooth muscle. Am J Physiol Lung Cell Mol Physiol 2007, 292:L1163-L1172.

28. Gosens R, Dueck G, Rector E, Nunes RO, Gerthoffer WT, Unruh H, Zaagsma J, Meurs H, Halayko AJ: Cooperative regulation of GSK-3 by muscarinic and PDGF receptors is associated with airway myocyte proliferation. Am J Physiol Lung Cell Mol Physiol 2007, 293:L1348-L1358.

29. Jeffery PK: Remodeling in asthma and chronic obstructive lung disease. Am J Respir Crit Care Med 2001, 164:S28-S38.

30. Postma DS, Timens W: Remodeling in asthma and chronic obstructive pulmonary disease. Proc Am Thorac Soc 2006, 3:434-439.

31. Ebina M, Takahashi T, Chiba T, Motomiya M: Cellular hypertrophy and hyperplasia of airway smooth muscles underlying bronchial asthma. A 3-D morphometric study. Am Rev Respir Dis 1993, 148:720-726.

32. Woodruff PG, Dolganov GM, Ferrando RE, Donnelly S, Hays SR, Solberg OD, Carter R, Wong HH, Cadbury PS, Fahy JV: Hyperplasia of smooth muscle in mild to moderate asthma without changes in cell size or gene expression. Am J Respir Crit Care Med 2004, 169:1001-1006.

33. Benayoun L, Druilhe A, Dombret MC, Aubier M, Pretolani M: Airway structural alterations selectively associated with severe asthma. Am J Respir Crit Care Med 2003, 167:1360-1368.

34. Graf J, Iwamoto Y, Sasaki M, Martin GR, Kleinman HK, Robey FA, Yamada Y: Identification of an amino acid sequence in laminin mediating cell attachment, chemotaxis, and receptor binding. Cell 1987, 48:989-996.

35. Grant DS, Tashiro K, Segui-Real B, Yamada Y, Martin GR, Kleinman HK: Two different laminin domains mediate the differentiation of human endothelial cells into capillary-like structures in vitro. Cell 1989, 58:933-943.

36. Matter ML, Laurie GW: A novel laminin E8 cell adhesion site required for lung alveolar formation in vitro. J Cell Biol 1994, 124:1083-1090.

37. Nelson J, McFerran NV, Pivato G, Chambers E, Doherty C, Steele D, Timson DJ: The 67 kDa laminin receptor: structure, function and role in disease. Biosci Rep 2008, 28:33-48.

38. Miyamoto S, Akiyama SK, Yamada KM: Synergistic roles for receptor occupancy and aggregation in integrin transmembrane function. Science 1995, 267:883-885.

39. Sakamoto N, Iwahana M, Tanaka NG, Osada Y: Inhibition of angiogenesis and tumor growth by a synthetic laminin peptide, CDPGYIGSR-NH2. Cancer Res 1991, 51:903-906.

40. Michigami T, Nomizu M, Yamada Y, Dunstan C, Williams PJ, Munday GR, Yoneda T: Growth and dissemination of a newly-established murine B-cell lymphoma cell line is inhibited by multimeric YIGSR peptide. Clin Exp Metastasis 1998, 16:645-654.

41. Conway EM, Collen D, Carmeliet P: Molecular mechanisms of blood vessel growth. Cardiovasc Res 2001, 49:507-521.

42. Barthel SR, Johansson MW, McNamee DM, Mosher DF: Roles of integrin activation in eosinophil function and the eosinophilic inflammation of asthma. J Leukoc Biol 2008, 83:1-12.

43. Georas SN, McIntyre BW, Ebisawa M, Bednarczyk JL, Sterbinsky SA, Schleimer RP, Bochner BS: Expression of a functional laminin receptor (alpha 6 beta 1, very late activation antigen-6) on human eosinophils. Blood 1993, 82:2872-2879.

Bart G.J. Dekkers Robert D. van der Schuyt Anita I.R. Spanjer Willem Jan Kuik Johan Zaagsma Herman Meurs

Signalling pathways of collagen I-induced airway smooth muscle

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Abstract Increased extracellular matrix (ECM) deposition and airway smooth muscle (ASM) mass are major contributors to airway remodelling in asthma. Increased expression of the ECM protein collagen I has been observed surrounding asthmatic ASM as well. Recently, we have demonstrated that collagen I induces a proliferative, hypocontractile ASM phenotype. Little is known, however, on the signalling pathways involved. Using bovine tracheal smooth muscle (BTSM), we now investigated the role of focal adhesion kinase (FAK) and downstream signalling pathways in collagen I-induced phenotype modulation. Phosphorylation of FAK was increased during adhesion to plastic or collagen I, without differences between these matrices. No differences between cellular adhesion were found either. Inhibition of FAK activity, by overexpression of the FAK deletion mutants FAT (focal adhesion targeting domain) and FRNK (FAK-related non-kinase), attenuated adhesion. After attachment, FAK phosphorylation was time-dependently increased in cells cultured on a collagen I matrix, whereas no activation was found on an uncoated plastic matrix. In addition, collagen I time- and concentration-dependently increased BTSM cell proliferation, which was inhibited by FAT and FRNK. In the presence of specific pharmacological inhibitors of p38 MAPK (SB203580) and Src-kinase (PP2) collagen I-induced proliferation was fully inhibited, while partial inhibition was observed by inhibition of PI3-kinase (LY294002) and MEK (U0126). The inhibition of cell proliferation by the inhibitors was associated with attenuation of the collagen I-induced hypocontractility. Collectively, the results indicate that induction of a proliferative, hypocontractile ASM phenotype by collagen I involves p38 MAPK, MEK, PI3-kinase and Src-mediated signalling pathways downstream of FAK. Introduction Airway hyperresponsiveness (AHR), persistent airway obstruction and decline in lung function are characteristic features of chronic asthma [1]. Airway remodelling, characterized by structural changes in the airway wall architecture, including increased airway smooth muscle (ASM) mass and altered deposition of extracellular matrix (ECM) proteins, is considered to contribute to these features [1,2]. Increased ASM mass may comprise hyperplasia as well as hypertrophy of ASM cells [3]. ASM cells may contribute to ongoing airway remodelling as they display phenotype plasticity and retain the ability to re-enter the cell-cycle [4]. Thus, exposure of ASM to mitogenic stimuli results in the induction of a proliferative phenotype, characterized by increased proliferation and decreased contractile function [5,6](Chapter 3). Phenotype plasticity is a reversible process and removal of mitogenic stimuli, for example by serum

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deprivation in the presence of insulin, results in the reintroduction of a (hyper)contractile ASM phenotype [7,8](Chapter 5).

From biopsy studies, it has become apparent that ECM deposition, including collagens, fibronectin and laminin �2/�2, is increased beneath the epithelial basement membrane of asthmatics [9,10]. In patients with asthma, the total amount of ECM in the microenvironment of the ASM is increased as well [11], and may involve increased deposition of collagen I, fibronectin, hyaluronan, versican, biglycan, lumican and elastic fibres [12-14]. The ECM surrounding the ASM cell plays a key role in determining physical and mechanical properties. In addition, ECM proteins have the capacity to affect ASM phenotype. Thus, growth factor-induced phenotype switching of ASM cells is inhibited by culturing the cells on laminin-111 resulting in the maintenance of a contractile ASM phenotype [15](Chapter 3). Conversely, culturing of ASM cells on monomeric collagen type I or fibronectin enhances growth factor-induced proliferation as well as growth factor-induced reductions of contractile marker proteins [16] (Chapter 3). Moreover, collagen I and fibronectin induce a hypocontractile, proliferative ASM phenotype in bovine tracheal smooth muscle (BTSM) strips by themselves [17] (Chapter 3). ASM cells obtained from asthmatics produce more collagen I and fibronectin compared to cells from healthy subjects [18,19]. In addition, nonasthmatic ASM cells cultured on an ECM laid down by asthmatic ASM cells proliferate more rapidly and vice versa [18], suggesting that changes in the ECM profile may contribute to enhanced asthmatic ASM growth in situ.

Integrins consist of a group of heterodimers linking the ECM to the intracellular compartment [20]. The collagen-binding integrin �2�1 is the main integrin involved in collagen I-induced ASM cell attachment, increased ASM cell proliferation and cytokine production by these cells, and glucocorticosteroid resistance [16,21]. In addition, the fibronectin-binding integrins �4�1 and �5�1 appeared important in the enhancement of PDGF-induced proliferation by collagen I, whereas the fibronectin-binding integrin �v�3 was also required for attachment to collagen I [16]. Recently, we have demonstrated that the �5�1 integrin is also of major importance in collagen I-induced increases of basal proliferation (Chapter 6).

No information is yet available on the signalling pathways of ECM-integrin interactions in ASM cells. From other cell types it is known that most integrins activate focal adhesion kinase (FAK), which results in autophosphorylation at Tyr397 and generates a binding site for Src, which then phosphorylates a number of other tyrosine residues on FAK [20,22,23]. FAK may subsequently activate downstream signalling cascades, including the PI3-kinase and MAPK pathways [20], which are importantly involved in ASM proliferation [24].

The aim of the present study was to explore the role of FAK and downstream signalling pathways in collagen I-induced ASM phenotype switching. Using BTSM cells, we examined the effects of collagen I on FAK phosphorylation during adhesion and proliferation. The role of FAK in these

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processes was assessed by overexpression of FAK and of the FAK deletion mutants FAT (derived from the focal adhesion targeting (FAT) domain of FAK) and FRNK (FAK-related non-kinase). FRNK is a alternative transcript of FAK, which contains the C-terminal domain but not the kinase domain [23,25]. FAT is a region present in both FAK and FRNK which is necessary for targeting of FAK to focal adhesion sites [26]. Both proteins inhibit FAK localization to the focal adhesions and FAK activation [25,26].In addition, by pharmacological inhibition of Src, mitogen activated protein kinase kinase (MEK), PI3-kinase and p38 mitogen activated protein kinase (MAPK), we investigated the contribution of these pathways to collagen I-induced BTSM proliferation and hypocontractility. Materials and methods Tissue preparation and organ-culture procedure. BTSM strips were prepared as described (Chapter 3). Tissue strips were washed in Medium Zero (sterile DMEM, supplemented with sodium pyruvate (1 mM), non-essential amino-acid mixture (1 :100), gentamicin (45 μg/ml), penicillin (100 U/ml), streptomycin (100 μg/ml), amphotericin B (1.5 μg/ml), apo-transferrin (5 μg/ml, human) and ascorbic acid (0.1 mM)) and transferred into suspension culture flasks. Strips were maintained in culture in an Innova 4000 incubator shaker (37°C, 55 rpm) for 4 days. When applied, collagen type I (50 μg/ml), PP2 (10 �M), U0126 (3 �M), LY294002 (10 �M) and/or SB203580 (10 �M) were present during the entire incubation period. Isometric tension measurements. Isometric tension measurements were performed as described (Chapter 3). In short, BTSM strips were washed with Krebs Henseleit (KH) buffer (composition (mM): NaCl 117.5, KCl 5.60, MgSO4 1.18, CaCl2 2.50, NaH2PO4 1.28, NaHCO3 25.00 and glucose 5.50, pregassed with 5% CO2 and 95% O2; pH 7.4 at 37°C). Subsequently, strips were mounted for isometric recording in organ baths. During a 90-min equilibration period resting tension was gradually adjusted to 3 g. Subsequently, BTSM strips were precontracted with 20 and 40 mM KCl solutions. Following washout, maximal relaxation was established by the addition of (-)-isoproterenol (0.1 �M; Sigma). Tension was readjusted to 3 g and after another equilibration period of 30 min cumulative concentration response curves were constructed to methacholine. When maximal tension was reached, the strips were washed and maximal relaxation was established using isoproterenol (10 �M).

Isolation of bovine tracheal smooth muscle cells. BTSM cells were isolated as described (Chapter 3). In short, tracheal smooth muscle was chopped, tissue fragments were washed in Medium Plus (DMEM supplemented with sodium pyruvate (1 mM), non-essential amino-acid mixture


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(1 :100), gentamicin (45 μg/ml), penicillin (100 U/ml), streptomycin (100 μg/ml), amphotericin B (1.5 μg/ml) and FBS (0.5%)). Enzymatic digestion was performed in Medium Plus, supplemented with collagenase P (0.75 mg/ml), papain (1 mg/ml) and soybean trypsin inhibitor (1 mg/ml). The suspension was incubated in an incubator shaker (Innova 4000) at 37°C, 55 rpm for 20 min, followed by a 10-min period of shaking at 70 rpm. After filtration of the obtained suspension over a 50 �m gauze, the cells were washed in Medium Plus, supplemented with 10% FBS. For all protocols, cells were used in passage 1-2. Transfection of BTSM cells with GFP expression vectors BTSM cells were plated at a density of 30.000 cells/well in 24-well culture plates and allowed to attach overnight, or grown to 95% confluency in 100 mm culture dishes. Subsequently, cells were washed twice with phosphate-buffered saline (PBS). Transfections in 24-well culture plates were performed using a mixture of 2 �l lipofectamine 2000 and 0.1 μg expression vector (GFP or GFP-FAK) or 0.8 μg expression vector (GFP, GFP-FAT or GFP-FRNK) for 6 h in 120 μl plain DMEM without serum and antibiotics. For transfections in the 100 mm dishes, a mixture of 60 μl lipofectamine 2000 and 3 μg or 24 μg of GFP expression vector, respectively, in 3.6 ml of DMEM were used. After 6 h, cells were washed twice with PBS and medium was replaced by DMEM Zero supplemented with 0.1% FBS. Preliminary results indicated that transfection efficiency for GFP reached 30 ± 4% (n=3). Cell adhesion assay Collagen-coated (50 μg/ml) culture plates were prepared as described (Chapter 3). The method for measurement of cell adhesion was adapted from [27]. Untransfected or GFP-, GFP-FAK-, GFP-FAT- or GFP-FRNK-transfected BTSM cells were harvested from 100 mm dishes by trypsinization. Cells were washed, resuspended in Medium Plus and transferred into uncoated or collagen-coated 24-well culture plates at a density of 50.000 cells/well and placed back in the incubator. At varying time intervals, plates were removed from the incubator and overlying medium was removed by gentle aspiration. After washing with 0.5 ml PBS at 37 °C, cells were fixed with 70% ethanol for 15 minutes at 4 °C. Subsequently, the plates were air dried for at least 30 min at 37 °C and stained for 25 minutes at room temperature using 0.1% crystal violet in water (0.3 ml/well). Cells were rinsed briefly with water and air dried. The stain was solubilized at room temperature using 10% acetic acid in water (0.5 ml/well) and quantified by colorimetric analysis (550 nm, Biorad 680 plate reader). Western analysis For the measurement of the phosphorylation of FAK, BTSM cells were cultured on uncoated or collagen I (50 μg/ml)-coated surfaces for varying periods of time. Cells were lysed in homogenization buffer (composition in mM: Tris-HCl 50 mM, NaCl 150.0, EDTA 1.0, PMSF 1.0, Na3VO4 1.0, NaF 1.0, pH 7.4, supplemented

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with leupeptin 10 μg/ml, aprotinin 10 μg/ml, pepstatin 10 μg/ml, Na-deoxycholate 0.25 % and Igepal 1% (NP-40)). Equal amounts of protein were subjected to electrophoresis and transferred onto PVDF membranes. Membranes were subsequently blocked in blocking buffer (composition: Tris-HCl 50.0 mM; NaCl 150.0 mM; Tween-20 0.1%, dried milk powder 5% (FAK) or BSA 5% (pFAK)) for 60 min at room temperature. Next, membranes were incubated overnight at 4 °C with primary antibodies (anti-FAK 1:2000 and anti-pFAK 1:1000, dilutions in blocking buffer containing BSA 5% or 3% BSA, respectively). After three washes with TBS-Tween 20 (TBST 0.1%, containing Tris-HCl 50.0 mM, NaCl 150.0 mM and Tween 20 0.1%) of 10 min each, membranes were incubated with horseradish peroxidase-labelled secondary anti-rabbit antibodies (dilution 1:2000 in blocking buffer containing 5% or 3% BSA, respectively) at room temperature for 90 min, followed by another three washes with TBST 0.1%. Antibodies were then visualized on film using enhanced chemiluminescence reagents and analyzed by densitometry (TotallabTM). Alamar blue proliferation assay BTSM cells were plated on uncoated or collagen I (1-100 μg/ml)-coated 24-well culture plates at a density of 30,000 cells/well and were allowed to attach overnight in Medium Plus, containing 10% FBS. The next day, cells were washed twice with PBS and made quiescent by incubation in Medium Zero, supplemented with 0.1% FBS for 3 days. Cells were then incubated with or without PDGF-AB (10 ng/ml) for 4 days in Medium Zero. Thereafter, cells were washed two times with PBS and incubated with HBSS containing 5% (vol/vol) Alamar blue solution. Conversion of Alamar blue into its reduced form by mitochondrial cytochromes was quantified by fluorimetric analysis, as indicated by the manufacturer. When applied PP2 (10 �M), U0126 (3 �M), LY294002 (10 �M) or SB203580 (10 �M) were present during the entire incubation period. For overexpression of GFP, GFP-FAK, GFP-FAT or GFP-FRNK, BTSM cells were transfected with the vectors after attachment, and subsequently cells were made quiescent as described above. [3H]-thymidine-incorporation [3H]-Thymidine-incorporation was performed as described previously (Chapter 3 and 5). BTSM cells were plated on uncoated or collagen I-coated 24-well culture plates at a density of 30,000 cells/well and allowed to attach overnight in Medium Plus. The next day, cells were transfected with the GFP, GFP-FAK, GFP-FAT or GFP-FRNK, washed with PBS and made quiescent by incubation in Medium Zero, supplemented with 1 �M insulin for 72 h. Subsequently, cells were washed and incubated in the absence or presence of PDGF (10 ng/ml) in Medium Zero for 28 h, the last 24 h in the presence of [methyl-3H]-thymidine (0.25 μCi/ml). After incubation, the cells were washed with PBS at room temperature. Subsequently, the cells were treated with ice-cold 5% trichloroacetic acid on ice for 30 min, and the acid-insoluble fraction was


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dissolved in NaOH (1 M). Incorporated [3H]-thymidine was quantified by liquid-scintillation counting using a Beckman LS1701 �-counter. Materials Dulbecco’s modification of Eagle’s medium (DMEM), FBS, sodium pyruvate solution, non-essential amino acid mixture, gentamicin solution, penicillin/streptomycin solution and amphotericin B solution (Fungizone) were obtained from Gibco BRL Life Technologies (Paisley, U.K.). Bovine serum albumin, apo-transferrin (human), leupeptin, aprotinin, pepstatin, soybean trypsin inhibitor, insulin (bovine pancreas) and (-)-isoproterenol hydrochloride were obtained from Sigma Chemical Co. (St. Louis, MO, U.S.A.). Platelet-derived growth factor (human, PDGF-AB) was from Bachem (Weil am Rhein, Germany). Methacholine was obtained from ICN Biomedicals (Costa Mesa, CA, U.S.A.). Anti-FAK was from Cell Signalling (Boston, MA, USA). Anti-FAK [pY397] and Alamar blue were from Biosource (Camarillo, CA, USA). Collagenase P and papain were from Boehringer (Mannheim, Germany). Monomeric collagen type I (calf skin) was from Fluka (Buchs, Switzerland). Lipofectamine was from Invitrogen (Paisley, UK). L(+)-ascorbic acid was from Merck (Darmstadt, Germany). SB203580 (4-[5-(4-Fluorophenyl)-2-[4-methylsulphonyl)phenyl]-1H-imidazol-4-yl]pyridine), LY294002 (2-(4-morpholinyl)-8-phenyl-4H-1-benzopyran-4-one), U0126 (1,4-diamino-2,3-dicyano-1,4-bis [2-aminophenylthio]butadiene) and PP2 (4-Amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine) were obtained from Tocris Cookson (Bristol, UK). pEGFP expression plasmids (Clontech) encoding FAK, and the FAK deletion mutants FAT and FRNK coupled to green fluorescent protein (GFP) were kindly provided by Dr. B. van de Water and Dr. S.E. Le Dévédec from the Division of Toxicology, Leiden Amsterdam Center for Drug Research [28,29]. All used chemicals were of analytical grade. Data analysis Data represent means ± SEM, from n separate experiments. Statistical significance of differences was evaluated by the Student's t-test for repeated measurements. Differences were considered to be statistically significant when P<0.05.

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Results Role of focal adhesion kinase in bovine tracheal smooth muscle cell adhesion.To investigate the effects of collagen I on ASM cell adhesion, BTSM cells were removed from the culture dish by trypsinization and replated onto uncoated plastic or collagen I (50 �g/ml)-coated culture plates. BTSM cells adhered to both substrates within 8 hours, without differences between plastic or collagen I (Figure 1A). To assess changes in FAK activation during adhesion, BTSM cells were plated and at 1, 2, 4, 6 and 24 h non-adhered cells were removed, adhered cells were lysed and FAK phosphorylation was determined. A significant increase (P<0.05) in FAK phosphorylation was observed in the cells adhered to plastic and collagen I compared to cells in suspension (Figure 1B). No differences in FAK phosphorylation were observed between both substrates. No FAK phosphorylation was observed in the non-adhered cells (not shown).

Figure 1: Focal adhesion kinase is activated during BTSM cell adhesion to plastic and collagen I. (A) Adhesion of BTSM cells to plastic and collagen I. BTSM cells were allowed to adhere either directly onto the plastic surface of culture plates or to collagen I (50 μg/ml)-coated surfaces. Data represent means ± SEM of 4 experiments performed in triplicate. (B) Focal adhesion kinase is activated during adhesion, without differences between plastic and collagen I matrices. Data are expressed as percentage of the maximal FAK phosphorylation observed between 1-24 h. Data represent means ± SEM of 4 experiments after densitometric analysis. Representative immunoblots of


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Y397 pFAK (upper panel) and total FAK (lower panel) are shown. *P<0.05, **P<0.01 compared to cells in suspension.To investigate the role of FAK in ASM cell adhesion, BTSM cells were transfected with GFP expression vectors encoding GFP (control), GFP-FAK or the FAK deletion mutants GFP-FAT and GFP-FRNK. In successfully transfected cells, expression of GFP-FAK, GFP-FAT and GFP-FRNK was detected in the focal adhesion sites (Figure 2), which is in correspondence with previous findings in rabbit primary synovial fibroblasts [29]. By contrast, expression of GFP was observed diffusely throughout the cytoplasm. Cells expressing GFP, GFP-FAT and GFP-FRNK remained elongated, whereas cells overexpressing GFP-FAK showed increased cell surface areas. Figure 2: Cellular expression of GFP fusion proteins in BTSM cells transfected with expression vectors encoding for GFP (control, upper left panel), GFP-FAK (upper right panel) and the FAK deletion mutants GFP-FAT (lower left panel) and GFP-FRNK (lower right panel). To assess whether FAK activation was required for ASM cell adhesion, BTSM cells were transfected with the expression vectors, trypsinized and replated onto plastic. No effects of overexpression of GFP-FAK on cell adhesion were observed, whereas overexpression of GFP-FRNK or GFP-FAT significantly reduced cell adhesion (Figure 3). Cell adhesion was maximally reduced at t=24

GFP-FAKGFP

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h, reaching 62±8% (P<0.01) in GFP-FRNK transfected cells and 67±12% (P<0.05) in GFP-FAT transfected cells compared to GFP transfected cells. Figure 3: Focal adhesion kinase is required for BTSM cell adhesion. (A) Overexpression of GFP-FAK did not affect BTSM cell adhesion. BTSM cells were cultured in 100 mm dishes and transfected with expression constructs for GFP (control) or GFP-FAK. After 6 hours, medium was replaced by Medium Zero + 0.1% FBS and cells were allowed to express fusion proteins for 3 days, after which cells were trypsinized and replated onto plastic. Data represent means ± SEM of 5 experiments each performed in triplicate. (B) Overexpression of the FAK deletion mutants GFP-FAT and GFP-FRNK decreases BTSM cell adhesion to plastic. Data represent means ± SEM of 5 experiments, each performed in triplicate. #P<0.05, ##P<0.01 compared to GFP controls. Role of focal adhesion kinase in bovine tracheal smooth muscle cell proliferation Culturing of BTSM cells on collagen I, concentration-dependently increased cell number (P<0.05, Figure 4A). The concentration of collagen required for 50% increase (EC50) in cell number was 14.1±1.8 �g/ml. Collagen I-induced increases in BTSM proliferation were also time-dependent, reaching 153±12% at day 4 (P<0.01, Figure 4B). To assess whether increases in cell number were associated with FAK activation, BTSM cells were cultured on uncoated plastic or collagen I (50 �g/ml) and cells were lysed after 1, 2, 3 or 4 days of culture and analyzed for FAK phosphorylation. Culturing on collagen I increased FAK phosphorylation, at days 2, 3 and 4 (P<0.05, Figure 4C). In correspondence with the findings for FAK phosphorylation during cell adhesion, no significant collagen I-induced phosphorylation was observed after 1 day.


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C Figure 4: Collagen I increases BTSM proliferation and FAK phosphorylation in a concentration- and time-dependent fashion. (A) BTSM cell number after culturing for 4 days on increasing concentrations of collagen I (0-100 μg/ml). Data represent means ± SEM of 4 experiments performed in triplicate. (B) BTSM cell number after culturing on uncoated plastic or collagen I (50 μg/ml) for 2 or 4 days. Data represent means ± SEM of 6 experiments performed in triplicate (C) Phosphorylation of FAK is time-dependently increased by culturing on collagen I (50 μg/ml), but not on uncoated plastic. Data represent means ± SEM of 4 experiments after densitometric analysis. Representative immunoblots of Y397 pFAK (upper panel) and total FAK (lower panel) are shown. *P<0.05, **P<0.01, ***P<0.001 compared to cells cultured on plastic. To investigate the role of FAK in BTSM cell proliferation, cells were plated on plastic or collagen I (50 μg/ml), allowed to attached overnight and transfected with GFP, GFP-FAK, GFP-FAT and GFP-FRNK. Subsequently, cells were serum deprived and stimulated with vehicle or PDGF-AB (10 ng/ml) for 4 days. Although not statistically significant, overexpression of GFP-FAK tended to decrease proliferation induced by both PDGF and collagen I (Figure 5A). Moreover, no effects of GFP-FAK overexpression were observed on DNA synthesis induced by PDGF, collagen I or the combination of both (Figure 5B). Overexpression of GFP-FAT or GFP-FRNK fully inhibited the increase in BTSM cell number induced by collagen I, whereas no significant effects of the inhibitory proteins were observed on PDGF-induced proliferation (Figure 5C). PDGF-induced proliferation on collagen I was normalized to the level observed for PDGF-induced proliferation in cells cultured on plastic. Similar effects were observed for GFP-FAT and GFP-FRNK on DNA synthesis (Figure 5D).

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Figure 5: Activation of FAK is required for collagen I-induced BTSM cell proliferation. (A) Overexpression of FAK tends do decrease BTSM cell number. BTSM cells were plated in 24-well culture plates and allowed to adhere. After adhesion, cells were transfected with GFP or GFP-FAK and subsequently serum deprived in Medium Zero + 0.1% FBS for 3 days. Cells were stimulated with or without PDGF-AB (10 ng/ml) for 4 days and cell number was assessed. (B) No effects of GFP-FAK overexpression were observed on DNA synthesis in BTSM cells. (C) Overexpression of GFP-FAT and GFP-FRNK decreases collagen I-induced increases in BTSM cell number. (D) Overexpression of GFP-FAT and GFP-FRNK decreased collagen I-induced DNA synthesis. Data represent means ± SEM of 7 experiments performed in triplicate. *P<0.05, **P<0.01, ***P<0.001 compared to GFP-transfected control cells cultured on uncoated plastic. #P<0.05, ##P<0.01 compared to GFP-transfected cells grown on collagen I and/or stimulated with PDGF.


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Role of Src, MEK, PI3-kinase and p38 MAPK in the induction of a proliferative, hypocontractile ASM phenotype by collagen I. To determine the contribution of downstream signalling pathways of FAK in the induction of a proliferative, hypocontractile ASM phenotype by collagen I, BTSM cells were cultured on plastic or collagen I in the absence and presence of specific pharmacological inhibitors of Src (PP2, 10 μM), MEK (U0126, 3 μM), PI3-kinase (LY294002, 10 μM) and p38 MAPK (SB203580, 10 μM). Collagen I-induced proliferation was inhibited by all inhibitors investigated, while basal cell numbers were significantly decreased by U0126 and significantly increased by SB203580 (Figure 6A). To investigate whether these pathways were involved in collagen I-induced hypocontractility as well, BTSM strips were cultured in the absence and presence of collagen I (50 �g/ml) and the inhibitors for 4 days. As observed for proliferation, collagen I-induced hypocontractility was normalized by all inhibitors investigated (Figure 6B, Table 1). The sensitivity (pEC50) for methacholine was unaffected by all treatments. Figure 6: Src-kinase, MEK, PI3-kinase and p38 MAPK are required for collagen I-induced BTSM cell proliferation and hypocontractility. (A) Effects of pharmacological inhibitors of Src-kinase (PP2, 10 μM), MEK (U0126, 3 μM), PI3-kinase (LY294002, 10 μM) and p38 MAPK (SB203580, 10 μM) on basal and collagen I (50 �g/ml)-induced changes in cell number. Data represent means ± SEM of 6 experiments performed in triplicate. *P<0.05, **P<0.01 compared to untreated control cells cultured on plastic. #P<0.05, ##P<0.01 compared to cells grown on collagen I in the absence of inhibitors. (B) Effects of the inhibitors on collagen I-induced hypocontractility. Data represent means ± SEM of 5-10 experiments performed in duplicate. *P<0.05, **P<0.01, ***P<0.001 compared to vehicle-treated control. #P<0.05, ##P<0.01 compared to collagen I-treated control.

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Table 1: Contractile responses of BTSM strips to methacholine after 4 days of culturing in the absence or presence of collagen I (50 �g/ml), with or without the pharmacological inhibitors of Src (PP2, 10 �M), MEK (U0126, 3 �M), PI3-kinase (LY294002, 10 �M) or p38 MAPK (SB203580, 10 �M)

Control Collagen I

Emax pEC50 (-log M) Emax pEC50 (-log M) Vehicle 100±0 6.82±0.12 77±4*** 6.72±0.11 PP2 91±11 6.75±0.23 111±13# 6.72±0.15 U0126 99±8 6.76±0.11 94±5# 6.94±0.12 LY294002 85±6 7.21±0.14 114±2# 7.04±0.16 SB203580 104±5 6.83±0.18 101±5# 6.84±0.24

Data represent means ± SEM of 5-10 independent experiments, each performed in duplicate. Abbreviations: Emax : maximal contraction; EC50: contraction of agonist eliciting half-maximal response; pEC50: negative logarithm of the EC50 value. ***P<0.01 compared to vehicle-treated control. #P<0.05 compared to collagen I-treated control (vehicle). Discussion In the current study, we demonstrate for the first time that the induction of a proliferative ASM phenotype by collagen I is dependent on the activation of FAK and downstream signalling pathways. The results indicate that FAK is activated during BTSM cell adhesion and that overexpression of two FAK deletion mutants FAT and FRNK, which compete with endogenous FAK for localization to the focal adhesions, inhibited cell adhesion. Moreover, FAK was activated during and required for collagen I-induced BTSM cell proliferation. Pharmacological inhibition of Src, MEK, PI3-kinase and p38 MAPK signalling pathways, which may be activated downstream of FAK, normalized the induction of a proliferative and hypocontractile phenotype induced by collagen I.

AHR is a characteristic feature of asthma and is defined by an exaggerated airway narrowing in response to either direct (histamine, methacholine) or indirect (exercise, cold air, hyperventilation) stimuli [30]. Variable AHR is observed after allergen exposure and is considered to reflect airway inflammation, whereas persistent AHR is considered to relate to structural changes in the airway wall, collectively termed airway remodelling [31,32]. Increased ASM mass, as a feature of airway remodelling, is considered to be the most important factor contributing to AHR and decline in lung function in asthmatics [3,33,34]. Previously, we and others have shown that changes in the ECM environment surrounding the ASM may contribute to ASM accumulation [17,18](Chapters 3 and 5). Thus, culturing of ASM on collagen I matrices increased proliferative responses and decreased contractile function, indicating that collagen I modulates the ASM phenotype to a proliferative,


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hypocontractile phenotype [17](Chapter 3) . Little is known, however, on the signalling pathways involved in this process. In the current study, it was found that culturing of BTSM cells on collagen I time-dependently increased phosphorylation of FAK, a cytoplasmic protein tyrosine kinase activated by most integrins [20]. Activation of FAK was found to be important in the BTSM cell proliferation as overexpression of FAT and FRNK, which inhibit FAK translocation to the focal adhesions and subsequent activation of the enzyme [25,26], also attenuated collagen I-induced proliferation. Activation of FAK was observed during adhesion of BTSM cells to uncoated plastic and collagen I matrices as well, without differences between the two matrices. FAK activation was also required for BTSM cell adhesion. The effects of overexpression of FAT and FRNK on collagen I-induced changes in BTSM cell number as mentioned above are unlikely to be due to changes in cell adhesion, as overexpression of these proteins only inhibited the collagen I-induced proliferative responses, whereas no effects were observed on basal and PDGF-induced increases in cell number. No effects of overexpression of FAK were observed on the parameters assessed, suggesting that the endogenous expression of the kinase is sufficient and not the limiting activation of downstream signalling pathways.

Changes in FAK activation during the proliferative phase only became apparent after 2 days of culture on collagen I, suggesting that FAK is not directly activated by collagen I-binding integrins, but that additional processes may be required. Indeed, studies in vascular smooth muscle cells have indicated that culturing on monomeric collagen type I increased the expression of other ECM proteins, including fibronectin [35], suggesting that the activation of FAK could be due to autocrine ECM deposition. This notion is also supported by previous findings showing that collagen I-induced increases in basal and growth factor-induced ASM proliferation required not only the collagen-binding integrin �2�1, but also the fibronectin binding integrins �4�1 and �5�1 [16](Chapter 6).

No effects of FAT or FRNK were observed on PDGF-induced proliferation, although in fibroblasts activation of the PDGF receptor has been shown to induce FAK phosphorylation [36]. Also in BTSM, PDGF increased FAK phosphorylation (not shown). The lack of effect of the deletion mutants, however, may be explained by the fact that activation of FAK by PDGF requires interaction of the receptor with the FERM domain, which is localized at the N-terminus of the kinase [23,36]. Both deletion mutants, however, are derived from the C-terminal domain and inhibit FAK localization to the focal adhesions, which is required for FAK activation by integrins [25,26], but do interfere with the activation of FAK via the FERM domain, providing an explanation for the lack of effect on PDGF-induced proliferation.

Phosphorylation of FAK at Tyr397, generates a high affinity binding site for Src, which then in turn fully activates FAK by phosphorylating Tyr576 and Tyr577 in the kinase domain [22,23]. Previous studies have found a critical role for Src in growth factor-induced ASM proliferation [37]. In agreement of the involvement of Src in collagen I-induced phenotype modulation, pharmacological

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inhibition of Src fully normalized both collagen I-induced proliferation and hypocontractility. Upon full activation, FAK can initiate a number of other signalling pathways, including the PI3-kinase and MAPK signalling pathways [20]. Activation of these pathways has been found to be important in the response of ASM cells to growth factors. PI3-kinase activation has been associated with transcriptional activation and protein synthesis leading to ASM cell proliferation and hypertrophy [38,39]. Integrins may not only activate PI3-kinase through FAK, but also via integrin linked kinase (ILK), another cytoplasmic protein tyrosine kinase, which is activated by the � subunit of integrins [40]. ILK has also been shown to be important in the regulation of contractile protein expression by human ASM. Knock-down of ILK increased mRNA and protein expression of smooth muscle-specific myosin heavy chain (sm-MHC), via regulation of Akt, which is downstream of PI3-kinase [41]. In the present study, inhibition of both PI3-kinase and FAK prevented collagen I-induced proliferation and hypocontractility, indicating the involvement of the FAK-PI3-kinase pathway in collagen I-induced BTSM proliferation.

ERK1/2 or p42/p44 mitogen activated protein kinases (MAPKs) transfer growth promoting signals to the nucleus and subsequently increase ASM proliferation [24]. In addition, p38 MAPK is involved in the regulation of growth factor-induced proliferation in ASM as well [42]. Inhibition of the MAPK signalling pathways, either directly (p38 MAPK) or by inhibiting MEK, which is upstream of p42/p44 MAPK, also inhibited collagen I-induced BTSM proliferation and hypocontractility. Collectively, these findings suggest that collagen I-induced activation of FAK results in activation of Src and, subsequently, of PI3-kinase and MAPK signalling pathways, which are all involved in collagen I-induced BTSM cell proliferation and hypocontractility (Figure 7).

Next to its important role in ECM-induced phenotype switching, FAK is also involved in acute ASM contractile responses. Thus, phosphorylation and membrane localization of the kinase is increased by mechanical strain and by contractile agonists [43-45]. Knock-out of FAK in human tracheal smooth muscle strips decreased tension development, myosin light chain phosphorylation and calcium signalling in response to the muscarinic receptor agonist acetylcholine and the membrane depolarizing stimulus KCl [46], suggesting an important role of FAK in smooth muscle contraction. These and our current findings, suggest that modulation of FAK activity in asthma may be an important new target in the treatment of ASM responsiveness and proliferation.

In conclusion, the present study provides new insights in the signalling events leading to ASM phenotype modulation by collagen I. These signalling pathways involve activation of FAK and downstream activation of Src-kinase, MEK, PI3-kinase and p38 MAPK. Moreover, our results indicate that modulation of FAK activity may be a new target in the treatment of both variable and persistent AHR in asthmatics.


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Figure 7: Proposed mechanism by which collagen I may affect ASM phenotype. Exposure of integrins (ITG) to collagen I results in the activation focal adhesion kinase (FAK). Autophosphorylation of FAK on Tyr397 creates a binding site for Src, which in turn phosphorylates FAK on Tyr576 and Tyr577. This leads to the full activation of the kinase, which may then activate PI3-kinase (PI3K) or mitogen activated protein kinase kinase (MEK) and mitogen activated protein kinase (MAPK) signalling pathways that regulate ASM phenotype and function. Alternatively, PI3-kinase may be activated by integrin-linked kinase (ILK) which is important in the regulation of contractile protein expression. See text for further detail. Acknowledgements This work was financially supported by the Netherlands Asthma Foundation, grant NAF 3.2.03.36. We are grateful to Dr. B. van de Water and Dr. S.E. Le Dévédec from the Division of Toxicology, Leiden Amsterdam Center for Drug Research for the GFP, GFP-FAK, GFP-FAT and GFP-FRNK expression plasmids.

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36. Sieg DJ, Hauck CR, Ilic D, Klingbeil CK, Schaefer E, Damsky CH, Schlaepfer DD. FAK integrates growth-factor and integrin signals to promote cell migration. Nat Cell Biol 2000; 2: 249-256.

37. Krymskaya VP, Goncharova EA, Ammit AJ, Lim PN, Goncharov DA, Eszterhas A, Panettieri RA, Jr. Src is necessary and sufficient for human airway smooth muscle cell proliferation and migration. FASEB J 2005; 19: 428-430.

38. Walker TR, Moore SM, Lawson MF, Panettieri RA, Jr., Chilvers ER. Platelet-derived growth factor-BB and thrombin activate phosphoinositide 3-kinase and protein kinase B: role in mediating airway smooth muscle proliferation. Mol Pharmacol 1998; 54: 1007-1015.

39. Halayko AJ, Kartha S, Stelmack GL, McConville J, Tam J, Camoretti-Mercado B, Forsythe SM, Hershenson MB, Solway J. Phophatidylinositol-3 kinase/mammalian target of rapamycin/p70S6K regulates contractile protein accumulation in airway myocyte differentiation. Am J Respir Cell Mol Biol 2004; 31: 266-275.

40. Liu S, Calderwood DA, Ginsberg MH. Integrin cytoplasmic domain-binding proteins. J Cell Sci 2000; 113 ( Pt 20): 3563-3571.

41. Wu Y, Huang Y, Herring BP, Gunst SJ. Integrin-linked kinase regulates smooth muscle differentiation marker gene expression in airway tissue. Am J Physiol Lung Cell Mol Physiol 2008; 295: L988-L997.

42. Fernandes DJ, Ravenhall CE, Harris T, Tran T, Vlahos R, Stewart AG. Contribution of the p38MAPK signalling pathway to proliferation in human cultured airway smooth muscle cells is mitogen-specific. Br J Pharmacol 2004; 142: 1182-1190.

43. Smith PG, Garcia R, Kogerman L. Mechanical strain increases protein tyrosine phosphorylation in airway smooth muscle cells. Exp Cell Res 1998; 239: 353-360.

44. Tang D, Mehta D, Gunst SJ. Mechanosensitive tyrosine phosphorylation of paxillin and focal adhesion kinase in tracheal smooth muscle. Am J Physiol 1999; 276: C250-C258.


46. Tang DD, Gunst SJ. Depletion of focal adhesion kinase by antisense depresses contractile activation of smooth muscle. Am J Physiol Cell Physiol 2001; 280: C874-C883.

Bart G.J. Dekkers Adnan Pehli Herman Meurs Johan Zaagsma

Bart G.J. Dekkers Adnan Pehli Herman Meurs Johan Zaagsma

Glucocorticosteroids and �2-adrenoceptor agonists synergistically prevent the induction of a proliferative, hypocontractile airway smooth

muscle phenotype

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Abstract Airway smooth muscle (ASM) accumulation in the airway wall may contribute to increased airway narrowing in asthma. ASM accumulation is in part explained by hyperplasia, and a number of mitogenic stimuli - including the growth factor PDGF and the extracellular matrix protein collagen I - induce a proliferative ASM phenotype, characterized by increased proliferation and decreased contractile function. Glucocorticosteroids and �2-adrenoceptor agonists have been shown to synergistically increase glucocorticosteroid receptor translocation in ASM cells, but the functional impact of this synergism with regard to phenotype modulation remains to be established. Using bovine tracheal smooth muscle, we investigated the effects of the glucocorticosteroids fluticasone, budesonide, dexamethasone, the �2-agonist fenoterol and the combination of 100-fold lower concentrations of fluticasone and fenoterol on the induction of a proliferative, hypocontractile phenotype by 4 days exposure to PDGF or collagen I. The results demonstrate that the glucocorticosteroids inhibited phenotype switching, the effects induced by collagen I being less susceptible to glucocorticosteroid action than those of PDGF. Treatment with fenoterol also inhibited proliferation induced by both stimuli. Fenoterol decreased the sensitivity and maximal contraction in response to methacholine and KCl, by itself, which was not further affected by PDGF and collagen I. At 100-fold lower concentrations, fluticasone and fenoterol synergistically prevented the induction of a hypocontractile, proliferative phenotype by both mitogens, and reversed the collagen I-induced glucocorticosteroid insensitivity. Collectively, our results indicate that glucocorticosteroids and �2-agonists synergize to inhibit ASM phenotype modulation, which may contribute to the therapeutic effectiveness of combined treatment with these drugs. Introduction Allergic asthma is a chronic inflammatory airways disease, associated with allergen-induced early and late bronchial obstructive reactions, airway hyperresponsiveness (AHR) and airway remodelling [1]. Airway remodelling is characterized by changes in the airway tissue architecture, including increased extracellular matrix (ECM) deposition and airway smooth muscle (ASM) accumulation [2-4]. Increased ASM mass is considered to be the most important factor contributing to AHR and to decline in lung function in asthma [5,6], and may comprise ASM hypertrophy and/or hyperplasia [7-9]. In keeping with hyperplasia, proliferation of ASM cells in vitro is increased by various mitogenic stimuli, including growth factors and ECM proteins [10]. Prolonged exposure of ASM to mitogens also induces a switch from a contractile to a hypocontractile phenotype, associated with increased proliferative capability [11,12](Chapter 3). Removal of mitogenic stimuli, in the presence of insulin or TGF-�, results in the

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reintroduction of a (hyper)contractile phenotype [13,14](Chapter 5), emphasizing the reversible nature of phenotype plasticity.

Inhaled glucocorticosteroids and �2-agonists are currently the most effective therapy for asthma control [15,16]. Moreover, combined treatment with glucocorticosteroids and �2-agonists results in better therapeutic management compared to monotherapy [17,18]. In addition to their potent ant-inflammatory effects, there is evidence that glucocorticosteroids may inhibit ASM remodelling in allergic asthma [19]. In line, in vitro studies have indicated that glucocorticosteroids may inhibit ASM cell proliferation [20-22]. In ASM cells, glucocorticosteroids accelerate the nuclear translocation of the glucocorticosteroid receptor and CCAAT/enhancer binding protein � (C/EBP�) and subsequently increase the expression of the cell cycle inhibitor p21waf1/cip1 [21]. Inhibition of ASM cell proliferation is also associated with the downregulation of growth factor-induced increases in cyclin D1 expression and phosphorylation of retinoblastoma protein (pRb) [22]. In ASM cells cultured on the ECM protein collagen type I, however, inhibition of proliferation by glucocorticosteroids is hampered [23-25], suggesting that changes in the ECM environment may impair glucocorticosteroid action. In addition to their anti-mitogenic effects, glucocorticosteroids inhibit TGF-�-induced sm-�-actin mRNA translation [26], indicating that glucocorticosteroids may also affect ASM contractile properties.

�2-Adrenoceptor agonists have also been shown to attenuate ASM cell proliferation induced by various stimuli [27], via a mechanism involving �2-adrenoceptor activation and subsequent activation of adenylyl cyclase, which triggers the cAMP/protein kinase A (PKA) signalling cascade [28]. Recently, however, it was suggested that not PKA, but another downstream effector of cAMP, exchange protein directly activated by cAMP (Epac) was responsible for the inhibition of ASM proliferation [29]. When combined, �2-agonists and glucocorticosteroids synergize and synchronize nuclear translocation of the glucocorticosteroid receptor and C/EBP�, resulting in a faster and longer activation of p21waf1/cip1 and inhibition of ASM proliferation [21].

To elucidate whether glucocorticosteroids and �2-agonists synergize to modulate ASM phenotype plasticity, we investigated the effects of the glucocorticosteroids fluticasone, budesonide and dexamethasone, the �2-agonist fenoterol and the combination of fluticasone and fenoterol on the induction of a hypocontractile bovine tracheal smooth muscle (BTSM) phenotype by PDGF and collagen I. The effects of the glucocorticosteroids and �2-agonists were also assessed on PDGF- and collagen-induced BTSM cell proliferation.

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Materials and methods Tissue preparation and organ-culture procedure. Bovine tracheal smooth muscle (BTSM) strips were prepared as described (Chapter 3 and 5). Tissue strips were washed in Medium Zero (sterile DMEM, supplemented with sodium pyruvate (1 mM), non-essential amino-acid mixture (1 :100), gentamicin (45 μg/ml), penicillin (100 U/ml), streptomycin (100 μg/ml), amphotericin B (1.5 μg/ml), apo-transferrin (5 μg/ml, human) and ascorbic acid (0.1 mM)) and transferred into suspension culture flasks. Strips were maintained in culture in an Innova 4000 incubator shaker (37°C, 55 rpm) for 4 days. When applied, PDGF-AB (10 ng/ml) and collagen type I (50 μg/ml) were present during the entire incubation period. Fluticasone (100 pM – 10 nM), budesonide (30 nM), dexamethasone (100 nM-1 μM) and/or fenoterol (10 nM – 1 μM) were applied 1 hr before and during stimulation with mitogens. Isometric tension measurements. Isometric tension measurements were performed as described (Chapter 3 and 5). In short, tissue strips were washed with several volumes of Krebs Henseleit (KH) buffer (composition (mM): NaCl 117.5, KCl 5.60, MgSO4 1.18, CaCl2 2.50, NaH2PO4 1.28, NaHCO3 25.00 and glucose 5.50, pregassed with 5% CO2 and 95% O2; pH 7.4 at 37°C). Subsequently, strips were mounted for isometric recording in organ baths containing KH buffer. During a 90-min equilibration period, with washouts every 30 min, resting tension was gradually adjusted to 3 g. Subsequently, BTSM strips were precontracted with 20 and 40 mM isotonic KCl solutions. Following washout, maximal relaxation was established by the addition of (-)-isoproterenol (0.1 �M). Collectively, the total washout period before the start of the isometric tension experiments was at least 3 h. After washing, tension was readjusted to 3 g and cumulative concentration response curves were constructed to stepwise increasing concentrations of isotonic KCl (5.6-50 mM) or methacholine (1 nM – 0.1 mM). When maximal tension was reached, the strips were washed and maximal relaxation was established using isoproterenol. Isolation of bovine tracheal smooth muscle cells. BTSM cells were isolated as described (Chapter 3 and 5). In short, after removal of the mucosa and connective tissue, tracheal smooth muscle was chopped. Tissue particles were washed and enzymatic digestion was performed in Medium Plus (DMEM supplemented with sodium pyruvate (1 mM), non-essential amino-acid mixture (1 :100), gentamicin (45 μg/ml), penicillin (100 U/ml), streptomycin (100 �g/ml), amphotericin B (1.5 μg/ml) and FBS (0.5%)), supplemented with collagenase P (0.75 mg/ml), papain (1 mg/ml) and soybean trypsin inhibitor (1 mg/ml). After digestion, cells were filtered and washed three times in Medium Plus, supplemented with 10% FBS. For all protocols, cells were used in passage 1-3.


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Alamar blue proliferation assay Collagen I-coated 24-well culture plates were prepared as described (Chapter 3). BTSM cells were plated on uncoated or collagen I-coated plastic culture plates at a density of 30,000 cells/well. Cells were washed twice with PBS and made quiescent by incubation in Medium Zero, supplemented with insulin (1 �M) for 3 days. Cells were then incubated with or without PDGF-AB (10 ng/ml) for 4 days in Medium Zero. Thereafter, cells were washed twice with PBS and incubated with HBSS containing 5% vol/vol Alamar blue solution. Conversion of Alamar blue into its reduced form by mitochondrial cytochromes was quantified by fluorimetric analysis, as indicated by the manufacturer. When used, cells were pretreated for 1 hr with fluticasone, budesonide, dexamethasone and/or fenoterol before stimulation with PDGF-AB. Western analysis of contractile protein expression To obtain whole BTSM tissue homogenates, tissue strips were cultured as described above. Western analysis was performed as described (Chapter 5). In short, homogenates were prepared by pulverizing the tissue under liquid nitrogen, followed by sonification in homogenization buffer. Equal amounts of protein were subjected to electrophoresis and transferred onto nitrocellulose membranes. Membranes were subsequently blocked in blocking buffer for 60 minutes at room temperature. Next, membranes were incubated overnight at 4 °C with primary antibodies (anti-sm-�-actin 1:2000, GAPDH 1:400, all dilutions in blocking buffer). After three washes with tris-buffered saline + 0.1% tween (0.1% TBST) of 10 min each, membranes were incubated with horseradish peroxidase-labelled secondary antibodies (dilution 1:2000 in blocking buffer) at room temperature for 60 min, followed by another three washes with 0.1% TBST. Antibodies were then visualized on film using enhanced chemiluminescence reagents and analyzed by densitometry (TotallabTM). All bands were normalized to GAPDH expression. Materials Platelet-derived growth factor (human, PDGF-AB) was from Bachem (Weil am Rhein, Germany). Monomeric collagen type I (calf skin) was from Fluka (Buchs, Switzerland). Dulbecco’s modification of Eagle’s medium (DMEM), FBS, gentamicin solution, non-essential amino acid mixture, penicillin/streptomycin solution, sodium pyruvate solution and amphotericin B solution (Fungizone) were obtained from Gibco BRL Life Technologies (Paisley, U.K.). Anti-sm-�-actin, apo-transferrin (human), bovine serum albumin, dexamethasone, fluticasone, insulin (bovine pancreas), (-)-isoproterenol hydrochloride and soybean trypsin inhibitor were obtained from Sigma Chemical Co. (St. Louis, MO, U.S.A.). Anti-GAPDH was from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Alamar blue was from Biosource (Camarillo, CA, USA). Methacholine was obtained from ICN Biomedicals (Costa Mesa, CA, U.S.A.). Collagenase P and papain were from Boehringer (Mannheim, Germany). Fenoterol was from

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Boehringer Ingelheim (Ingelheim, Germany). Budesonide was a gift of Prof. H.W. Frijlink, University of Groningen. All chemicals used were of analytical grade. Data analysis All data are presented as mean ± SEM. Statistical differences between means were calculated using one-way ANOVA for repeated measures, followed by a Newman–Keuls multiple comparisons test. Significance was reached at P<0.05. Results Effects of glucocorticosteroids on PDGF- and collagen I-induced BTSM hypocontractility and proliferation. To assess whether glucocorticosteroids inhibit the induction of a hypocontractile ASM phenotype, BTSM strips were incubated with PDGF or collagen I in the absence and presence of fluticasone, budesonide or dexamethasone. In accordance with previous studies [11](Chapter 3), we found that culturing of strips in the presence of PDGF-AB (10 ng/ml) or collagen I (50 μg/ml) significantly (P<0.05) decreased maximal contractile force (Emax) to methacholine compared to vehicle-treated controls (Figures 1-3). Figure 1: The induction of a hypocontractile, proliferative phenotype induced by PDGF or collagen I is inhibited by fluticasone. (A,B) Concentration-response curves of methacholine-induced contractions of BTSM strips, pretreated with vehicle (control), (A) PDGF (10 ng/ml), or (B) collagen I (50 �g/ml), in the absence or presence of fluticasone (10 nM) for 4 days. Data represent means ± SEM of 5 independent experiments, each performed in duplicate. (C) Effects of fluticasone (10 nM) on basal and PDGF (10 ng/ml) or collagen I (50 �g/ml)-stimulated increases in cell number of cultured BTSM cells. Data represent means ± SEM of 7 independent experiments each performed in triplicate. **P<0.01, ***P<0.001 compared to vehicle-treated (control). #P<0.05, ##P<0.01, ###P<0.001 compared to mitogen in the absence of fluticasone.


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Fluticasone (10 nM, Figure 1) and budesonide (30 nM, Figure 2) both inhibited the decrease in Emax induced by PDGF-AB (P<0.01, both glucocorticosteroids) and collagen I (P<0.05, both glucocorticosteroids). Figure 2: The induction of a hypocontractile, proliferative phenotype induced by PDGF or collagen I is inhibited by budesonide. (A,B) Concentration-response curves of methacholine-induced contractions of BTSM strips, pretreated with vehicle (control), (A) PDGF (10 ng/ml), or (B) collagen I (50 �g/ml), in the absence or presence of budesonide (30 nM) for 4 days. Data represent means ± SEM of 5-6 independent experiments each performed in duplicate. (C) Effects of budesonide (30 nM) on basal and PDGF (10 ng/ml), or collagen I (50 �g/ml)-stimulated increases in cell number of cultured BTSM cells. Data represent means ± SEM of 6 independent experiments each performed in triplicate. *P<0.05, **P<0.01, ***P<0.001 compared to vehicle-treated (control). #P<0.05, ##P<0.01, ###P<0.001 compared to mitogen in the absence of budesonide. 100 nM Dexamethasone attenuated the decrease in Emax induced by PDGF (Figure 3A, P<0.001), whereas the decrease induced by collagen I was not affected (Figure 3B). At 1 μM, however, dexamethasone did inhibit the decrease in Emax induced by collagen I (Figure 3D, P<0.01). No apparent effects of the glucocorticosteroids were observed under control conditions. Similar effects were observed for KCl-induced contractions (data not shown). The sensitivity for both contractile stimuli was unaffected by all treatments.

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Figure 3: Effects of dexamethasone on the induction of a hypocontractile, proliferative phenotype by PDGF or collagen I. (A,B) Concentration-response curves of methacholine-induced contractions of BTSM strips, pretreated with vehicle (control), (A) PDGF (10 ng/ml), or (B) collagen I (50 �g/ml), in the absence or presence of 100 nM dexamethasone for 4 days. Data represent means ± SEM of 4-6 independent experiments, each performed in duplicate. (C) Effects of 100 nM dexamethasone on basal and PDGF (10 ng/ml) or collagen I (50 �g/ml)-stimulated increases in cell number of cultured BTSM cells. Data represent means ± SEM of 5 independent experiments each performed in triplicate. (D) Concentration-response curves of methacholine-induced contractions of BTSM strips, pretreated with vehicle (control) or collagen I (50 �g/ml) in the absence or presence of 1 �M dexamethasone for 4 days. Data represent means ± SEM of 5 independent experiments, each performed in duplicate. (E) Effects of 1 �M dexamethasone on basal and PDGF (10 ng/ml), or collagen I (50 �g/ml)-stimulated increases in cell number of cultured BTSM cells. Data represent means ± SEM of 6 independent experiments, each performed in triplicate. *P<0.05, **P<0.01, ***P<0.001 compared to vehicle-treated (control). #P<0.05, ##P<0.01, ###P<0.001 compared to mitogen in the absence of dexamethasone.


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As we have previously shown that the decrease in Emax in response to methacholine and KCl was inversely correlated with the proliferative responses of BTSM cells to growth factors or ECM proteins [11](Chapter 3), we also investigated whether glucocorticosteroids inhibit the proliferative responses in BTSM cells. PDGF and collagen I matrices increased BTSM cell number (Figures 1-3, P<0.01). Both fluticasone (Figure 1C) and budesonide (Figure 2C) inhibited the increase in cell number induced by PDGF (P<0.001, both glucocorticosteroids) and collagen I (P<0.05 and P<0.001, respectively). In agreement with the effects on contractility, PDGF-induced proliferation was decreased by 100 nM dexamethasone (P<0.05), but collagen I-induced proliferation was not affected by this concentration of the glucocorticosteroid (Figure 3C). At a concentration of 1 �M, however, dexamethasone inhibited proliferation induced by both PDGF and collagen I (P<0.01 and P<0.05, respectively, Figure 3E). No effects of the glucocorticosteroids were observed on basal cell number. Effects of the �2-adrenoceptor agonist fenoterol on PDGF- and collagen I-induced hypocontractility and proliferation. To assess whether �2-adrenoceptor activation affects the induction of a hypocontractile BTSM phenotype, strips were incubated with PDGF or collagen I in the absence and presence of fenoterol. Treatment with fenoterol (1 μM) for 4 days, followed by prolonged (approximately 3 h) and repeated washout, significantly (P<0.01) decreased the Emax to methacholine and reduced the sensitivity to the agonist in control strips (pEC50=6.92±0.01 (control) and 6.53±0.08 (fenoterol), P<0.05, Figure 4A and 4B). Both PDGF and collagen I also reduced the Emax to methacholine, however, without an effect on the sensitivity (Figure 4A and 4B), as also shown in Figure 1-3. Combined treatment of PDGF or collagen I with fenoterol did not further decrease maximal methacholine-induced contractions or sensitivity. Similar effects were observed for KCl-induced contractions (data not shown). To investigate whether changes in contractility were accompanied by changes in contractile protein abundance, expression of sm-�-actin was determined. In accordance with previous findings (Chapter 3), it was found that 4 days of treatment with PDGF decreased sm-�-actin expression by approximately 40% (Figure 5). Treatment with fenoterol (1 �M) tended to decrease sm-�-actin expression by about 25% compared to controls; however, this did not reach statistical significance. Combined treatment with PDGF did not further decreased sm-�-actin expression.

Treatment of BTSM cells with fenoterol fully inhibited both PDGF- and collagen I-induced increases in cell number (P<0.001, both mitogens, Figure 4C), whereas no effects were observed on cell number in the absence of mitogenic stimuli.

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Figure 4: Effects of fenoterol on the induction of a hypocontractile, proliferative phenotype induced by PDGF or collagen I. (A,B) Concentration-response curves of methacholine-induced contractions of BTSM strips, pretreated with vehicle (control), (A) PDGF (10 ng/ml), or (B) collagen I (50 �g/ml), in the absence or presence of fenoterol (1 �M) for 4 days. Data represent means ± SEM of 5 independent experiments, each performed in duplicate. (C) Effects of fenoterol (1 �M) on basal and PDGF (10 ng/ml), or collagen I (50 �g/ml)-stimulated increases in cell number of cultured BTSM cells. Data represent means ± SEM of 7 independent experiments, each performed in triplicate. **P<0.01, ***P<0.001 compared to vehicle-treated (control). ###P<0.001 compared to mitogen in the absence of fenoterol. Figure 5: Fenoterol decreases the expression of the contractile protein sm-�-actin. Western analysis of sm-�-actin protein expression in homogenates of BTSM strips pretreated with vehicle (control) or PDGF (10 ng/ml) in the absence or presence of fenoterol (1 �M) for 4 days. (A) Graph shows means ± SEM of 4 independent experiments after densitometric analysis. *P<0.05 compared to control. (B) Representative immunoblots of sm-�-actin (upper panel) and GAPDH (lower panel).


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Fenoterol and fluticasone synergize to prevent the induction of a hypocontractile, proliferative phenotype by PDGF and collagen I. To assess whether �2-agonists and glucocorticosteroids synergize to inhibit the induction of a hypocontractile ASM phenotype, BTSM strips were incubated with PDGF or collagen I in the absence and presence of 100-fold lower concentrations of fenoterol (10 nM), fluticasone (100 pM) or the combination of both. At these concentrations, fenoterol and fluticasone, by themselves, only slightly diminished the PDGF- and collagen I-induced decrease of the maximal methacholine-induced contractions, which did not reach statistical significance. However, combined treatment with fenoterol and fluticasone fully inhibited the reduction in Emax induced by PDGF and collagen I (P<0.01 and P<0.05, respectively, Figure 6A and 6B). Of note, under control conditions no effects were observed for fenoterol, fluticasone or the combination (data not shown). For KCl-induced contractions, the reduction in Emax by both agonists appeared to be additive. Moreover, fenoterol significantly reduced the effects of collagen I (P<0.05). The sensitivity for either contractile stimuli was unaffected by all treatments. Figure 6: Low concentrations of fenoterol and fluticasone synergistically prevented the induction of a hypocontractile, proliferative phenotype by PDGF or collagen I. (A,B) Concentration-response curves of methacholine-induced contractions of BTSM strips, pretreated with vehicle (control), (A) PDGF (10 ng/ml), or (B) collagen I (50 �g/ml), in the absence or presence of fenoterol (10 nM), fluticasone (100 pM) or the combination of both for 4 days. Data represent means ± SEM of 5 independent experiments, each performed in duplicate. (C) Effects of fenoterol (10 nM), fluticasone (100 pM) or the combination of both on basal and PDGF (10 ng/ml), or collagen I (50 �g/ml)-stimulated increases in cell number of isolated BTSM cells. Data represent means ± SEM of 5 independent experiments, each performed in triplicate. *P<0.05 **P<0.01, ***P<0.001 compared to vehicle-treated (control). #P<0.05, ##P<0.01, ###P<0.001 compared to mitogen in the absence of fluticasone and fenoterol.

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Treatment of BTSM cells with fenoterol (10 nM) did not inhibit PDGF- or collagen-induced proliferation (Figure 6C). Fluticasone (100 pM) significantly (P<0.01) reduced the effects of PDGF on BTSM cell proliferation, while no effects of were observed on collagen I-induced proliferation. Combined treatment with both fenoterol and fluticasone completely prevented the increase in cell number induced by both mitogens (P<0.001, both), in case of collagen I in a synergistic fashion. No effects of fenoterol, fluticasone or the combination were observed under control conditions. Discussion In the present study, we demonstrate for the first time that glucocorticosteroids and �2-agonists synergize in preventing ASM phenotype switching. It was found that the induction of a proliferative, hypocontractile ASM phenotype by prolonged exposure to PDGF and collagen I was inhibited by glucocorticosteroids fluticasone, budesonide and dexamethasone. The effects of collagen I were relatively resistant to glucocorticosteroid action seen both in intact ASM tissue (contractility) and ASM cells (proliferation). Treatment with fenoterol also inhibited PDGF- and collagen I-induced BTSM cell proliferation. When applied in low concentrations, fenoterol and fluticasone synergized to prevent ASM phenotype switching by both mitogens.

ASM accumulation is a characteristic feature of airway wall remodelling in asthma, which is considered to be a major contributor to AHR and to decline in lung function [5,6]. Exposure of ASM to mitogens in vitro induces a switch from a contractile to a hypocontractile phenotype, characterized by increased proliferative rates and decreased contractile responses [11](Chapter 3). Glucocorticoids have been shown to inhibit ASM proliferation [20-22], but the effects on ASM contractility remained to be determined. In our current studies, we found that glucocorticosteroids inhibit PDGF-induced hypocontractility of intact BTSM strips as well as BTSM cell proliferation, indicating that ASM phenotype switching by the growth factor was inhibited. Similarly, collagen I-induced BTSM hypocontractility and proliferation were inhibited, although these effects were more resistant to glucocorticosteroid treatment as indicated by the reduced effects of dexamethasone (100 nM) and fluticasone (100 pM), compared to the effects on PDGF-induced phenotype switching. These findings correspond to previous observations, showing that bFGF-induced proliferation of human ASM cells cultured on collagen I, but not on laminin matrices, was resistant to dexamethasone (100 nM) and fluticasone (1 nM) [23,24]. In a subsequent study, it was demonstrated that the failure of dexamethasone to inhibit ASM proliferation was associated with a failure to decrease cyclin D1 expression and pRb levels [25]. In bovine ASM, however, dexamethasone did inhibit bFGF-induced proliferation on collagen I [30], suggesting that ASM cells of bovine origin may remain more sensitive to steroid treatment.


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In addition to their anti-mitogenic effects, glucocorticoids have also been shown to inhibit sm-�-actin accumulation in response to TGF-� [26]. In the current study, glucocorticosteroids prevented BTSM hypocontractility and proliferation without effects in the absence of mitogenic stimuli. Collectively, these findings suggest that glucocorticoids inhibit switching both towards a hypocontractile and towards a hypercontractile ASM phenotype, which may both be present in asthmatics [7,31], to maintain a (normo)contractile ASM phenotype. In support of the maintenance of such a phenotype in vivo, budesonide has been shown to inhibit both ASM hyperplasia and hypercontractility in a guinea pig model of chronic allergic asthma [19].

�2-Adrenoceptor agonists are the primary treatment for the relief of bronchospasm in asthma [32]. In addition, �2-agonists attenuate ASM cell proliferation in response to various stimuli in vitro [27,32]. In the present study, we found that both PDGF- and collagen I-induced BTSM cell proliferation were inhibited by the �2-agonist fenoterol. Interestingly, pretreatment of BTSM strips with fenoterol for 4 days decreased maximal contractions induced by the receptor-dependent agonist methacholine and the receptor-independent stimulus KCl, and reduced the sensitivity to both stimuli. These effects are unlikely to be due to the presence of residual fenoterol as the strips have been washed for at least 3 h, but may be partly explained by increased constitutive �2-adrenoceptor activity, as reported previously by this laboratory [33,34]. In these studies, pretreatment with fenoterol time- and concentration-dependently increased constitutive �2-adrenoceptor activity, resulting in decreased maximal contractile responses as well as reduced sensitivity of the tissue in response to KCl [33]. However, although in these studies methacholine sensitivity after 18 h of fenoterol pretreatment was attenuated, no effects were observed on maximal contractile responses [34], suggesting that additional effects may underlie this decrease. Indeed, treatment of BTSM strips for 4 days with fenoterol tended to decrease sm-�-actin expression, which corresponds with previous findings showing that treatment of human ASM with the long acting �2-agonist salmeterol reduced sm-�-actin expression [26].

Combined treatment with glucocorticosteroids and �2-agonists results in more effective therapeutic management of asthma and COPD than monotherapy [17,18]. In human lung, glucocorticoids have been shown to increase transcription of the �2-adrenoceptor gene [64]. In ASM glucocorticosteroids and �2-agonists synergize to accelerate nuclear translocation of the glucocorticoid receptor and C/EBP�, resulting in the synergistic activation of the cell cycle inhibitor p21waf1/cip1 [21]. In addition, �2-agonists synergistically enhance glucocorticosteroid response element (GRE)-dependent transcription, amplifying the transcription of anti-inflammatory genes, including mitogen activated protein kinase phosphatase (MKP-1) and the cyclin-dependent kinase inhibitor p57KIP2 [35], of which the latter has been shown to be involved in the antiproliferative effects of glucocorticosteroids [36].

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In our current study, we showed that combined treatment with low concentrations of fenoterol and fluticasone synergistically inhibited PDGF- and collagen I-induced phenotype switching. No effects of combined treatment were observed in the absence of mitogenic stimuli, suggesting that fenoterol increased the activity of the glucocorticosteroid and not vice versa, as this would have resulted in decreased maximal contractions and reduced sensitivity in response to the contractile stimuli. These findings are in agreement with clinical studies, in which monotherapy with �2-agonists did not suppress inflammation, but enhanced the anti-inflammatory effects of inhaled glucocorticosteroids [17]. Studies on the contribution of the different aspects of airway remodelling in asthmatics have indicated that increased ASM mass is likely to be the most important factor in increased airway resistance and persistent AHR [5,6]. Our current studies may contribute to the increased therapeutic efficacy of combined treatment with �2-agonists and glucocorticosteroids [17], by effectively reducing the increase in ASM mass and development of persistent AHR.

In addition to increased ASM mass, airway remodelling in asthmatics is characterized by increased deposition of ECM proteins, including collagen I, beneath the epithelial basement membrane and surrounding the ASM bundles [37,38]. Collagen I may not only contribute to ASM accumulation by increasing proliferative responses [30,39](Chapter 3), but also because it renders ASM cells resistant to the anti-mitogenic actions of glucocorticosteroids [23,24]. This could provide an explanation, why a subgroup of severe asthmatics, is poorly controlled by glucocorticosteroids [40]. Our current findings, showing that combined treatment with fluticasone and fenoterol synergistically normalizes not only PDGF-, but also collagen I-induced ASM phenotype switching, suggests that combination therapy may be not only be more beneficial in the therapeutic management of asthma by increasing asthma control and duration of bronchodilation, but also by normalizes the sensitivity of the ASM to lower doses of glucocorticosteroid.

In conclusion, our results indicate that glucocorticoids and �2-agonists synergize to prevent PDGF- and collagen I-induced ASM phenotype switching. As increased ASM mass is considered to contribute importantly to AHR in asthma, these findings may explain the enhanced efficacy of �2-adrenoceptor agonist/glucocorticosteroid combination therapy in controlling asthma. Acknowledgements This work was financially supported by the Netherlands Asthma Foundation, grant NAF 3.2.03.36.


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References 1. Bousquet J, Jeffery PK, Busse WW, Johnson M, Vignola AM. Asthma. From



3. Lloyd CM, Robinson DS. Allergen-induced airway remodelling. Eur Respir J 2007; 29: 1020-1032.

4. Postma DS, Timens W. Remodeling in asthma and chronic obstructive pulmonary disease. Proc Am Thorac Soc 2006; 3: 434-439.




8. Benayoun L, Druilhe A, Dombret MC, Aubier M, Pretolani M. Airway structural alterations selectively associated with severe asthma. Am J Respir Crit Care Med 2003; 167: 1360-1368.







15. Barnes PJ. Corticosteroids: the drugs to beat. Eur J Pharmacol 2006; 533: 2-14. 16. Penn RB. Embracing emerging paradigms of G protein-coupled receptor agonism

and signaling to address airway smooth muscle pathobiology in asthma. Naunyn Schmiedebergs Arch Pharmacol 2008; 378: 149-169.

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17. Giembycz MA, Kaur M, Leigh R, Newton R. A Holy Grail of asthma management: toward understanding how long-acting beta(2)-adrenoceptor agonists enhance the clinical efficacy of inhaled corticosteroids. Br J Pharmacol 2008; 153: 1090-1104.

18. Barnes PJ. Scientific rationale for inhaled combination therapy with long-acting beta2-agonists and corticosteroids. Eur Respir J 2002; 19: 182-191.

19. Bos IS, Gosens R, Zuidhof AB, Schaafsma D, Halayko AJ, Meurs H, Zaagsma J. Inhibition of allergen-induced airway remodelling by tiotropium and budesonide: a comparison. Eur Respir J 2007; 30: 653-661.







26. Goldsmith AM, Hershenson MB, Wolbert MP, Bentley JK. Regulation of airway smooth muscle alpha-actin expression by glucocorticoids. Am J Physiol Lung Cell Mol Physiol 2007; 292: L99-L106.

27. Tomlinson PR, Wilson JW, Stewart AG. Inhibition by salbutamol of the proliferation of human airway smooth muscle cells grown in culture. Br J Pharmacol 1994; 111: 641-647.

28. Hirst SJ, Martin JG, Bonacci JV, Chan V, Fixman ED, Hamid QA, Herszberg B, Lavoie JP, McVicker CG, Moir LM, Nguyen TT, Peng Q, Ramos-Barbon D, Stewart AG. Proliferative aspects of airway smooth muscle. J Allergy Clin Immunol 2004; 114: S2-17.

29. Kassel KM, Wyatt TA, Panettieri RA, Jr., Toews ML. Inhibition of human airway smooth muscle cell proliferation by beta 2-adrenergic receptors and cAMP is PKA independent: evidence for EPAC involvement. Am J Physiol Lung Cell Mol Physiol 2008; 294: L131-L138.


31. Leguillette R, Laviolette M, Bergeron C, Zitouni NB, Kogut P, Solway J, Kashmar L, Hamid Q, Lauzon AM. Myosin, Transgelin, and Myosin Light Chain Kinase:


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Expression and Function in Asthma. Am J Respir Crit Care Med 2008; 179: 194-204.

32. Deshpande DA, Penn RB. Targeting G protein-coupled receptor signaling in asthma. Cell Signal 2006; 18: 2105-2120.

33. De Vries B, Meurs H, Roffel AF, Elzinga CR, Hoiting BH, de Vries MM, Zaagsma J. Beta-agonist-induced constitutive beta(2)-adrenergic receptor activity in bovine tracheal smooth muscle. Br J Pharmacol 2000; 131: 915-920.

34. De Vries B, Roffel AF, Zaagsma J, Meurs H. Effect of fenoterol-induced constitutive beta(2)-adrenoceptor activity on contractile receptor function in airway smooth muscle. Eur J Pharmacol 2001; 431: 353-359.

35. Kaur M, Chivers JE, Giembycz MA, Newton R. Long-acting beta2-adrenoceptor agonists synergistically enhance glucocorticoid-dependent transcription in human airway epithelial and smooth muscle cells. Mol Pharmacol 2008; 73: 203-214.

36. Samuelsson MK, Pazirandeh A, Davani B, Okret S. p57Kip2, a glucocorticoid-induced inhibitor of cell cycle progression in HeLa cells. Mol Endocrinol 1999; 13: 1811-1822.




40. Barnes PJ. New drugs for asthma. Nat Rev Drug Discov 2004; 3: 831-844.

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General discussion and summary 10C

hapter

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Regulation of airway smooth muscle phenotype and function by extracellular matrix proteins Shedding of the epithelium, goblet cell hyperplasia, increased blood vessel formation, enhanced deposition of extracellular matrix (ECM) proteins and increased airway smooth muscle (ASM) mass are characteristic aspects of airway wall remodelling in asthma [1-3]. Increased ASM mass may be explained by ASM hyperplasia as well as by hypertrophy [4] and is considered to be a major factor contributing to airway hyperresponsiveness and decline in lung function in asthmatics [5,6]. In keeping with hyperplasia, ASM cells, in contrast to skeletal myocytes and cardiomyocytes [7,8], retain their ability to enter the cell-cycle and change their phenotype (phenotypic plasticity) [9]. In vitro, exposure of ASM cells and tissue to serum and various growth factors results in a switch from a contractile to a proliferative phenotype. This phenotype is characterized by increased expression of proliferative markers such as Ki67, increased proliferation, decreased expression of contractile marker proteins like smooth muscle �-actin (sm-�-actin), calponin and smooth muscle myosin heavy chain (sm-MHC), and decreased contractile function [10,11]. ASM phenotype switching is reversible, as indicated by the observation that removal of mitogenic stimuli, for example by serum deprivation of the culture medium in the presence of insulin or TGF-� results in the reintroduction of a contractile ASM phenotype, associated with increased contractile protein expression and increased contractile function [12,13].

Increased ECM deposition is another characteristic feature of airway remodelling in asthma. Studies on the nature of ECM changes in the airway wall of asthmatics have revealed increased subepithelial deposition of collagens I, III and V, fibronectin, tenascin, hyaluronan, versican, biglycan, lumican and several laminin chains (�2, �3, �5, �1, �2 and �1) [14-21]. Patchy staining for laminin �1 chains has also been observed in the airways of allergic asthmatics, whereas no staining was observed in the airways of non-allergic asthmatics or healthy subjects [21]. On the other hand, expression of collagen IV, decorin and elastin in the airway wall of asthmatic patients is decreased [22,23]. The composition of ECM in the microenvironment surrounding the asthmatic ASM cell is changed as well, and is characterized by increased deposition of collagen I, fibronectin, hyaluronan, versican, biglycan, lumican and elastic fibres [24-26]. In vitro, ASM cells produce a variety of ECM proteins, including collagens, fibronectin, laminins, perlecan, elastin, thrombospondin, versican, decorin, chondroitin sulphate and hyaluronan [27-30]. ECM synthesis by asthmatic ASM cells is changed compared to that of healthy cells and is characterized by increased production of collagen I, perlecan and fibronectin, and decreased production of laminin �1, chondroitin sulphate, collagen IV and hyaluronan [27,30,31]. This may contribute to the changed ECM microenvironment of the ASM cells and has been shown to have an impact on ASM function, potentially underlying intrinsic differences observed between healthy and asthmatic ASM cells (Chapter 2).

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Thus, ASM cells obtained from asthmatics proliferate faster than those obtained from healthy subjects [32]. whereas proliferation of healthy ASM cells is increased when cultured on a matrix secreted by asthmatic ASM cells and vice versa [27]. Similarly, increased production of eotaxin by asthmatic ASM cells, a feature of increased synthetic function, is also dependent on the ECM proteins produced by these cells [31]. Recent reports have demonstrated that changes in the ECM environment of ASM cells may induce phenotype switching as well. Thus, culturing of human ASM cells on the ECM proteins fibronectin and collagen I increased proliferative responses to the growth factors thrombin and platelet-derived growth factor (PDGF), whereas the PDGF-induced reduction in protein expression of sm-�-actin, calponin and sm-MHC was further enhanced by culturing the cells on these ECM proteins [33]. On the other hand, culturing ASM cells on laminin-111 or matrigel decreased the proliferative responses and maintained contractile protein expression in the presence of growth factors [33].

Altered deposition of ECM proteins in the airway wall could also alter mechanical properties of the ASM by phenotypic modulation, as has been observed previously for growth factors [11]. Thus, changes in proliferation of cultured bovine tracheal smooth muscle (BTSM) cells in response to peptide growth factors were found to be tightly correlated with growth factor-induced changes in contractility of intact strips, in which endogenous ECM constituents and cell-to-cell contacts are still intact [11]. Using the same approach, we now investigated the functional impact of ASM exposure to fibronectin, collagen I and laminin-111 (Chapter 3). In this study, it was demonstrated that culturing BTSM cells on fibronectin and collagen I matrices increased the proliferation of these cells, whereas prolonged exposure (4 days) of BTSM strips to these ECM proteins attenuated the maximal contractile responses to the receptor-dependent agonist methacholine and the receptor-independent stimulus KCl. These changes were associated with decreased protein expression of sm-�-actin, calponin and sm-MHC in the strips. Collectively, these effects were similar to the phenotypic changes observed for PDGF. Moreover, culturing of ASM cells on fibronectin or collagen I augmented PDGF-induced proliferation in an additive fashion; however, without additional effects on contractility or contractile protein expression. The fibronectin-induced depression of contractility was blocked by the integrin antagonist Arg-Gly-Asp-Ser (RGDS), but not by its negative control Gly-Arg-Ala-Asp-Ser-Pro (GRADSP), indicating that interaction of the ECM protein with its integrin receptors was required. Laminin-111 did not affect contractility on its own, but reduced PDGF-induced hypocontractility. Similarly, PDGF-induced proliferation was reduced when cells were cultured on laminin-111. Strong correlations were observed between ECM-induced changes in BTSM strip contractility and sm-�-actin, calponin and sm-MHC expression, indicating that changes in contractile protein expression underlie the changes in contractile force. As observed for peptide growth factors [11], strong inverse

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correlations between altered contractility and mitogenic responses were observed as well, suggesting that the changes in contractility were indeed due to ASM phenotype switching. Moreover, our findings clearly suggest that ECM proteins differentially affect ASM phenotype and function.

To determine the impact of phenotype switching on contractile function in human ASM, a similar approach was used by Moir et al [34]. In this study, exposure of intact human bronchiole ring segments to serum for 3 or 6 days did not affect ASM area or protein expression of sm-�-actin and sm-MHC. However, 6 days of culturing in the presence of serum reduced carbachol-, histamine- and KCl-induced contractions and decreased calponin expression, suggesting that the ASM phenotype - at least to some extent - had switched to a hypocontractile state [34]. In Chapter 4, we addressed the impact of ECM- and growth factor-induced phenotype switching on the function of intact human ASM. In this chapter, it was demonstrated that prolonged exposure (4 days) of human tracheal smooth muscle (HTSM) strips to collagen I or PDGF decreased the maximal contractions induced by methacholine or KCl, which was associated with decreased expression of the contractile proteins sm-�-actin and sm-MHC. In addition, both collagen I and PDGF increased proliferation of cultured primary HTSM cells. As observed for BTSM (Chapter 3), it was found that culturing of HTSM cells on collagen I additively increased the proliferative responses by PDGF, whereas no additional effects of combined treatment were observed on contractility or contractile protein expression. Moreover, the results presented in Chapter 6 indicate that ECM-induced phenotype switching of human ASM cells may occur not only after exposure to collagen I, but also after exposure to fibronectin, as culturing of these cells on fibronectin increased their proliferation as well. In conclusion, these results indicate that ECM- and growth-factor-induced phenotype modulation is of relevance to human ASM function. Interestingly, our findings also indicate that BTSM is a representative experimental model for human ASM phenotype plasticity. In addition to increased ASM mass, increased expression of contractile marker proteins has been reported in asthmatic ASM as well (Chapter 2) [35]. In vitro, expression of contractile marker proteins is increased by serum deprivation, which is further augmented in the presence of insulin or TGF-� [12,13]. Studies on the role of ECM proteins in this process, showed that expression of laminin �2, �1 and �1 chains is increased during serum deprivation [36]. Increased laminin expression was required for ASM maturation as the laminin �1 chain competing peptide Tyr-Ile-Gly-Ser-Arg (YIGSR) and the RGD containing peptide Gly-Arg-Gly-Asp-Ser-Pro (GRGDSP) both inhibited the increase in contractile protein expression [36]. Moreover, ASM cells increase expression of the laminin binding integrins �3, �6 and �7 during maturation, of which the �7 integrin was found essential for ASM maturation [37].

Exposure of BTSM strips and cells to insulin results in the induction of a hypercontractile, hypoproliferative phenotype, characterized by increased


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contractile protein expression, increased contractile responses and decreased proliferation in response to growth factors [13,38]. The role laminins in the induction of a hypercontractile, hypoproliferative ASM phenotype by insulin are described in Chapter 5. The results presented in this chapter demonstrate that insulin-induced hypercontractility is inhibited by the laminin competing peptides YIGSR and RGDS, while no effects were observed for GRADSP. In addition, increased expression of the contractile protein sm-MHC and decreased proliferative responses after insulin treatment were normalized by YIGSR as well. The insulin-induced increase in BTSM contractility was associated with increased protein expression of laminin �2, �1 and �1 chains, suggesting that increased laminin-211 expression is required for the induction of a hypercontractile ASM phenotype by insulin. As observed previously for contractile protein accumulation [13], PI3-kinase and Rho kinase signalling pathways were required for the insulin-induced increase in laminin expression and hypercontractility. Collectively, these results indicate a critical role for laminins in the induction of a hypercontractile, hypoproliferative ASM phenotype by prolonged insulin exposure. Increased laminin expression in the airways of asthmatic patients could be involved in the increased contractility and contractile protein expression of asthmatic ASM. Moreover, the results may have important implications for the use of inhaled insulin formulations in diabetes mellitus. As also indicated above, ASM cells and ECM proteins communicate with each other mainly via integrins, a group of heterodimeric, transmembrane glycoproteins [39]. In vitro studies have indicated that integrins are importantly involved in ECM-induced changes in ASM cell adhesion, maturation, synthetic function, survival and proliferation [23,36]. Various integrins have been implicated in ASM proliferation. Thus, the collagen-binding �2�1 integrin as well as the fibronectin-binding �4�1 and �5�1 integrins were shown to be required for the enhanced growth factor-induced proliferation on collagen I and fibronectin [40]. The �5�1 integrin has also been found to be important in serum-induced proliferation of both nonasthmatic and asthmatic ASM [41]. Moreover, increased eotaxin secretion by asthmatic ASM largely required the �5�1 integrin [31].

Although integrins have been shown to modulate ASM function in vitro, the importance these ECM receptors in allergen-induced ASM remodelling in vivo has not yet been explored. Using a guinea pig model of chronic allergic asthma [42], the contribution of integrins and ECM proteins to allergen-induced airway remodelling in vivo was investigated using the integrin-blocking peptide RGDS, which contains the RGD binding motif present not only in fibronectin, but also in collagens and laminins and which blocks the binding of these ECM proteins to multiple integrins [43,44], including the �3�1, �5�1, �v�1 and �v�3 integrins which are expressed by ASM cells in culture [40,45]. The results described in Chapter 6 demonstrate that RGD-binding integrins and their endogenous agonists are important contributors to ASM remodelling in chronic allergic asthma. It was shown that topical treatment of the airways with RGDS

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inhibited ASM hyperplasia, pulmonary expression of the proliferative marker proliferating cell nuclear antigen (PCNA), increased contractile protein expression and ASM hypercontractility induced by repeated allergen challenge once a weekly for 12 weeks. No effects were observed for the negative control peptide GRADSP. RGDS did not affect allergen-induced fibrosis or inflammation, which may suggest that the peptide directly influenced the ASM cell. Fully in line, it was found that proliferative responses induced by collagen I, fibronectin, serum and PDGF in human ASM cells required signalling via RGD-sensitive integrins, particularly of the �5�1 subtype. In line with the results described in Chapter 5, RGDS also dose-dependently inhibited sm-�-actin accumulation induced by serum deprivation in the presence of insulin. Collectively, these findings for the first time indicate that integrins modulate ASM remodelling in allergic asthma, which can be inhibited by a small peptide containing the RGD motif.

No effects of RGDS were observed in saline-challenged guinea pigs (Chapter 6), suggesting that integrin activation occurs only under pathological conditions. In allergen-challenged animals, integrin activation may result from synthesis of new ECM proteins, recruitment of plasma-derived ECM proteins to the ASM layer or alterations in the existing ECM, exposing the matricryptic integrin binding sites which are normally hidden within the proteins [46,47]. In support of such a notion, denatured monomeric collagen I, but not fibrillar collagen I, increased growth factor-induced proliferation of human ASM, suggesting that changes in the configuration of the ECM are required for the induction of proliferative signals [40]. Similarly, no effects of RGDS or the function-blocking antibodies were observed on ASM cell numbers (Chapter 6) or BTSM strip contractility (Chapter 3) under unstimulated (control) conditions, suggesting that exposure of RGD-binding integrins to their recognition sites within the ECM proteins is limited under these conditions. Previous studies [36] and the results described in Chapter 5 suggest that laminins are important in a shift of the ASM to a hypercontractile, hypoproliferative phenotype. Therefore, the effects of the specific laminin-competing peptide YIGSR on airway remodelling were also investigated in our guinea pig model of chronic allergic asthma (Chapter 7). Surprisingly, topical administration of YIGSR in the airways attenuated allergen-induced ASM hyperplasia and pulmonary PCNA expression. Treatment with YIGSR also increased pulmonary sm-MHC expression and ASM contractility, both in saline and allergen-challenged animals, suggesting that treatment with the peptide increased rather than decreased laminin function. In vitro, culturing of human ASM cells on immobilized YIGSR concentration-dependently reduced PDGF-induced proliferation of the cells to a similar extent as culturing of the cells on laminin-111. Remarkably, the effects of both immobilized YIGSR and laminin were antagonized by soluble YIGSR. Collectively, these results indicate that in vivo treatment with YIGSR mimics rather than inhibits laminin function, which


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seems to depend on the physicochemical microenvironment of the peptide. The mechanisms underlying these differential effects are currently unknown. Collectively, the findings presented in Chapters 3-7 point to an important functional role of cell – matrix interactions in the regulation of ASM phenotype. Changes in the extracellular microenvironment surrounding the asthmatic ASM cells may therefore importantly contribute to the increased contractile, proliferative and synthetic characteristics of these cells. The ASM itself may be importantly involved in creating an ECM environment that supports these abnormal ASM functions. Extracellular matrix-induced signal transduction in airway smooth muscle In several cell types it has been established that specific integrin – ECM interactions result in the activation of various intracellular signalling cascades required for changes in proliferation, migration, differentiation and survival induced by ECM proteins [48]. Various integrins activate focal adhesion kinase (FAK), which then subsequently activates downstream signalling pathways, including PI3-kinase- and MAPK-dependent pathways [48]. The results described in Chapter 8 indicate an important role for the activation of FAK and downstream signalling pathways in the induction of a proliferative, hypocontractile ASM phenotype by collagen I. Using BTSM cells in vitro, it was shown that FAK is activated during adhesion to an uncoated plastic matrix and to a collagen I matrix, without differences between the two conditions. Activation of FAK was required for cell adhesion, as adhesion was inhibited by overexpression of the FAK deletion mutants FAT (derived from the Focal Adhesion Targeting (FAT) domain of FAK) and FRNK (FAK-Related Non-Kinase), which both inhibit FAK localization to the focal adhesions and subsequent FAK activation [49,50]. FAK activation was further increased by culturing on collagen I for 2-4 days, but not by culturing on plastic. The delayed activation suggests that FAK is not directly activated by collagen I, but that additional secondary processes may be required. In support of this notion, monomeric collagen I has been shown to increase the expression of other ECM components, including fibronectin, in vascular smooth muscle cells [51]. Moreover, the collagen I-induced increase in basal (Chapter 6) and growth factor-dependent [40] proliferation required interaction with the fibronectin-binding integrin �5�1.

In our experiments, collagen I increased BTSM cell proliferation in a concentration- and time-dependent fashion, which was inhibited by overexpression of FAT and FRNK. Pharmacological inhibition of the Src, MEK, PI3-kinase and p38 MAPK signalling cascades inhibited collagen I-induced proliferation and hypocontractility, suggesting that activation of downstream

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signalling pathways of FAK is required for collagen I-induced phenotype modulation. Next to its role in collagen I-induced ASM phenotype modulation, FAK has also been shown to be directly involved in contractile responses, as indicated by the observation that downregulation of FAK in HTSM strips decreased tension development, myosin light chain phosphorylation and calcium signalling in response to acetylcholine and KCl [52], suggesting that FAK may pose not only a target in the treatment of persistent but also in acute airway hyperresponsiveness. Effects on glucocorticosteroids and �2-adrenoceptor agonists on ASM phenotype switching Glucocorticosteroids and �2-adrenoceptor agonists are currently the most effective therapy for asthma control based on their anti-inflammatory and bronchodilating actions, respectively [53,54]. Treatment with glucocorticosteroids has been shown to prevent increases in ASM mass in animal models of asthma [55], suggesting that these drugs also affect airway remodelling. In vitro, both glucocorticosteroids and �2-adrenoceptor agonist have been shown to inhibit ASM proliferation [56-59]. However, anti-mitogenic actions of glucocorticosteroids were shown to be reduced when ASM cells are cultured on collagen I matrices [60-62]. This indicates that increased deposition of collagen I surrounding the ASM may not only increase ASM proliferation, but may also contribute to steroid resistance as observed in severe asthmatics. Clinical studies have indicated that combined treatment with glucocorticosteroids and �2-agonists results in a better asthma control than monotherapy with each of the drugs [63]. In human lung, glucocorticoids have been shown to increase transcription of the �2-adrenoceptor gene [64] and in ASM cells, �2-agonists have been shown to synergistically increase glucocorticosteroid receptor translocation [57]. However, the impact of this synergism on ASM phenotype switching remains to be established. The results described in Chapter 9 demonstrate that the glucocorticosteroids fluticasone, budesonide and dexamethasone inhibited both PDGF- and collagen I-induced BTSM switching to a proliferative, hypocontractile phenotype, although the collagen I-induced phenotype switch was less sensitive to glucocorticosteroid action. Proliferative responses induced by both mitogens were inhibited by the �2-agonist fenoterol as well. When applied in 100-fold lower concentrations, that were not or only partially effective, fluticasone and fenoterol synergized to inhibit ASM phenotype switching induced by PDGF and collagen I. As increased ASM mass is likely to be the most important factor of increased airway resistance and AHR in asthma [5,6], these findings suggest that combined treatment may effectively reduce decline in lung function and persistent AHR in chronic asthma. Since not only growth factor-, but also collagen I-induced phenotype switching was inhibited by the combined treatment, it can be envisaged that, in addition to increasing anti-


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inflammatory, bronchodilating and anti-proliferative effects [63], normalizing steroid sensitivity by �2-agonists contributes to the enhanced asthma control by the combination therapy. Collectively, these findings may contribute to the enhanced efficacy of �2-adrenoceptor agonist/glucocorticosteroid combination therapy in controlling asthma. Taken together the studies described in this thesis reveal that: ECM proteins differentially regulate ASM phenotype and contractile function.

Fibronectin and collagen I induce a functionally hypocontractile ASM phenotype, associated with increased proliferative responses, whereas laminin-111 maintains a functionally contractile phenotype and inhibits growth factor-induced proliferation (Chapters 3 & 6).

Collagen I and PDGF induce a proliferative, hypocontractile phenotype in human ASM, which is of relevance to ASM function (Chapter 4).

Insulin-induced laminin �2, �1 and �1 chain expression, mediated by PI3-kinase- and Rho kinase-dependent signalling pathways, contributes to the induction of a hypercontractile, hypoproliferative ASM phenotype. Increased laminin expression in the airway wall could contribute to the increased ASM contractility and contractile protein expression as observed in asthma (Chapter 5).

Induction of contractile protein and laminin expression in response to insulin may limit the use of inhaled insulin formulations in diabetes mellitus (Chapter 5).

Endogenous activation of RGD-binding integrins modulates allergen-induced ASM remodeling in an animal model of allergic asthma, which can be inhibited by topical application of RGDS, a small peptide containing the RGD motif. Based on these findings, RGD-binding integrins may represent a novel target in the treatment of airway remodeling in asthma (Chapter 6).

The laminin �1 chain-competing peptide YIGSR inhibits the induction of a hypercontractile, hypoproliferative ASM phenotype in vitro (Chapter 5). By contrast, in vivo YIGSR promoted a contractile, hypoproliferative ASM phenotype, which could depend on the physicochemical microenvironment of the peptide (Chapter 7).

Activation of FAK is obligatory for ASM cell adhesion both to plastic and to collagen I matrices and mediates collagen I-induced proliferation of these cells (Chapter 8).

Phenotypic modulation of ASM cells by collagen I is dependent on Src, MEK, p38 MAPK and PI3-kinase, which may be activated downstream of FAK (Chapter 8).

Glucorticosteroids inhibit ASM phenotype switching induced by PDGF and collagen I, the effects of collagen I being more resistant to glucocorticosteroid action. Low doses of glucocorticoids synergize with low

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doses of �2-adrenoceptor agonists to prevent the induction of a proliferative, hypocontractile ASM phenotype, without differences between the two mitogens. This synergism could contribute to enhanced control of asthma by combination therapy (Chapter 9).

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Nederlandse samenvatting Astma is een chronische inflammatoire luchtwegaandoening die wordt gekenmerkt door een variabele luchtwegobstructie en een verhoogde reactiviteit van de luchtwegen voor bronchusobstructieve prikkels, zoals koude lucht, mist, rook, chemische irritantia, en farmacologische prikkels, zoals histamine en methacholine. Deze verhoogde reactiviteit wordt luchtweghyperreactiviteit genoemd. In de meeste gevallen heeft astma een allergische oorsprong, waarbij de patiënt tevens overgevoelig is voor specifieke allergenen, zoals huisstofmijt, gras- en boompollen, schimmels en epitheel van huisdieren. Allergische reacties in de luchtwegen zijn een belangrijke oorzaak voor luchtwegontsteking, waarbij allergische mediatorstoffen, cytokines en neurotransmitters vrijkomen, die – direct of indirect – bronchusobstructie en luchtweghyperreactiviteit kunnen induceren. Factoren die een rol spelen bij de ontwikkeling van luchtweghyperreactiviteit zijn veranderingen in de neuronale en niet-neuronale regulatie van de luchtweg-gladde spierfunctie, een toegenomen gevoeligheid van het gladde spierweefsel voor contractiele prikkels, en fysische veranderingen in de luchtwegen, zoals oedeem en luchtwegremodelling.

Luchtwegremodelling wordt onder meer gekenmerkt door schade aan het luchtwegepitheel, verhoogde aantallen slijmproducerende cellen, toename van het aantal bloedvaten (angiogenese), verhoogde en veranderde depositie van extracellulaire matrix (ECM) eiwitten in de luchtwegwand, en toename van de luchtweg-gladde spiermassa. De toename van de spiermassa wordt met name veroorzaakt door een toename van het aantal gladde spiercellen (hyperplasie), hoewel vergroting van gladde spiercellen (hypertrofie) ook wordt waargenomen.

De luchtweg-gladde spiercel draagt in belangrijke mate bij aan luchtwegremodelling, en aan luchtweghyperreactiviteit, dankzij de fenotypische plasticiteit van deze cel, waarbij een reversibele overgang van een contractiel naar een proliferatief fenotype mogelijk is. In vitro is aangetoond dat luchtweg-gladde spiercellen na blootstelling aan serum of sommige groeifactoren een proliferatief, hypocontractiel fenotype kunnen aannemen. Dit fenotype wordt gekenmerkt door een toegenomen celdeling, geassocieerd met een verhoogde expressie van proliferatieve markers, zoals bijvoorbeeld Ki67 en PCNA (proliferating cell nuclear antigen) – en een afgenomen contractiliteit die gepaard gaat met een verlaagde expressie van contractiele eiwitten – zoals smooth muscle �-actine (sm-�-actin), calponine en smooth muscle myosine heavy chain (sm-MHC). Omgekeerd kan een contractiel, hypoproliferatief fenotype (weer) worden geïntroduceerd, wanneer de proliferatieve stimuli worden verwijderd, bijvoorbeeld door middel van serumdeprivatie. In aanwezigheid van insuline of transforming growth factor-� (TGF-�) kan zelfs een hypercontractiel fenotype worden geïnduceerd, gekenmerkt door een hoge expressie van contractiele eiwitten en een toegenomen contractiele functie.


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Verhoogde en veranderde depositie van ECM eiwitten in de luchtwegwand is een ander karakteristiek kenmerk van luchtwegremodelling in astma. Onder de epitheliale basaalmembraan van astmapatiënten is de expressie van collageen I, III en V, fibronectine, tenascine, hyaluronan, versican, biglycan, lumican en verschillende laminine ketens (�2, �3, �5, �1, �2 en �1) is verhoogd. Verder wordt verspreid door de luchtwegwand van deze patiënten heen de laminine �1 keten aangetroffen, terwijl deze in de luchtwegen van gezonde personen afwezig is. Collageen IV, decorine en elastine daarentegen, worden verminderd tot expressie gebracht in de luchtwegen van astmapatiënten.

Rondom de luchtweg-gladde spierbundel van astmapatiënten is de samenstelling van de ECM ook veranderd en wel door een verhoogde depositie van collageen I, fibronectine, hyaluronan, versican, biglycan, lumican en elastine-positieve vezels. In vitro produceren luchtweg-gladde spiercellen een scala aan ECM eiwitten, waaronder collagenen, fibronectine, laminines, perlecan, elastine, thrombospondine, versican, decorine, chondroïtine sulfaat en hyaluronan. De productie van ECM eiwitten door astmatische gladde spiercellen is veranderd in vergelijking met gezonde spiercellen en wordt gekenmerkt door een toegenomen productie van collageen I, perlecan en fibronectine, alsmede een afgenomen productie van laminine �1, chondroïtine sulfaat, collageen IV en hyaluronan. Deze veranderde productie draagt mogelijk bij aan de veranderde ECM samenstelling rondom de astmatische luchtweg-gladde spier en heeft invloed op het gedrag van de spiercel, wat mogelijk de intrinsieke verschillen tussen gezonde en astmatische spiercellen verklaart (Hoofdstuk 2). Zo neemt de proliferatieve respons van gezonde gladde spiercellen toe wanneer deze worden gekweekt op een matrix geproduceerd door astmatische cellen en vice versa. Ook een toegenomen eotaxine productie door de astmatische gladde spiercel – wat in verband wordt gebracht met een toegenomen synthetische functie – is afhankelijk van de ECM geproduceerd door deze cellen. Recent is gevonden dat veranderingen in de ECM rondom de luchtweg-gladde spiercel het fenotype van deze cel kunnen beïnvloeden. Wanneer humane luchtweg-gladde spiercellen worden gekweekt op collageen I of fibronectine reageren deze sterker op de proliferatieve effecten van de groeifactoren thrombine en platelet-derived growth factor (PDGF). Tevens wordt de PDGF-geïnduceerde afname van de sm-�-actine, calponine en sm-MHC expressie versterkt. Het kweken van luchtweg-gladde spiercellen op laminine-111 of matrigel, een basaalmembraanextract rijk aan laminine-111, verlaagt daarentegen de proliferatieve effecten van thrombine en PDGF, en handhaaft de expressie van de contractiele eiwitten.

Veranderde depositie van ECM eiwitten rondom de luchtweg-gladde spiercel zou ook de mechanische eigenschappen van de spier kunnen veranderen – door fenotype modulatie, zoals eerder ook is gevonden voor groeifactoren. Voorgaand onderzoek aan ons laboratorium heeft namelijk aangetoond dat er een sterke inverse relatie bestaat tussen de effecten van


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groeifactoren op de proliferatieve activiteit van geïsoleerde gladde spiercellen van de rundertrachea en de contractiliteit van het intacte spierpreparaat. In Hoofdstuk 3 worden de effecten van fibronectine, collageen I en laminine-111 op de contractiele functie van rundertracheale gladde spierstrippen beschreven in relatie tot de proliferatieve respons van gladde spiercellen, gekweekte op deze ECM eiwitten. Aangetoond werd dat het kweken van rundertracheale gladde spiercellen op fibronectine- en collageen I-matrices de proliferatie van deze cellen bevorderde, terwijl langdurige blootstelling (4 dagen) van intacte rundertracheale spierstrippen aan deze eiwitten een afname veroorzaakte van de maximale contractie, in respons op zowel de receptorafhankelijke agonist methacholine als de receptoronafhankelijke stimulus KCl. De afname in contractiliteit was geassocieerd met een verlaagde expressie van contractiele eiwitten (sm-�-actine, calponine en sm-MHC). Vergelijkbare effecten werden waargenomen voor PDGF. Bovendien werd de PDGF-geïnduceerde celproliferatie additief verhoogd door fibronectine en collageen I, terwijl er geen additionele effecten werden waargenomen op de contractiliteit of contractiele eiwitexpressie. De fibronectine-geïnduceerde afname in contractiliteit was afhankelijk van interacties tussen het ECM eiwit en zijn integrinereceptoren, aangezien deze afname werd genormaliseerd door de integrine-antagonist Arg-Gly-Asp-Ser (RGDS), en niet door zijn negatieve controle Gly-Arg-Ala-Asp-Ser-Pro (GRADSP). Het kweken van spierstrippen in aanwezigheid van laminine-111 had zelf geen effecten op de contractiliteit, maar voorkwam de door PDGF geïnduceerde afname in de maximale contractie. Ook de door PDGF geïnduceerde proliferatie was verlaagd in cellen die werden gekweekt op laminine-111. Sterke correlaties werden gevonden tussen de effecten van de ECM eiwitten op de contractiliteit en de sm-�-actine, calponine en sm-MHC eiwitexpressie. Dit betekent dat de veranderingen in contractiliteit veroorzaakt worden door een veranderde contractiele eiwitexpressie. Tevens werd een sterke inverse relatie gevonden tussen de celproliferatie en de contractiliteit, hetgeen suggereert dat de waargenomen veranderingen in gladde spiercontractiliteit het gevolg zijn van discrete fenotypeveranderingen. Samenvattend geven deze resultaten aan dat verschillende ECM eiwitten verschillende effecten kunnen hebben op het fenotype en de contractiele functie van de luchtweg-gladde spier. In hoofdstuk 4 werd onderzocht of ECM- en groeifactor-afhankelijke fenotypeveranderingen ook de functie van de humane luchtweg-gladde spier beïnvloeden. Gevonden werd dat 4 dagen blootstelling van humane tracheale gladde spierpreparaten aan collageen I of PDGF de maximale methacholine- en KCl-geïnduceerde contracties verlaagde, hetgeen geassocieerd was met een afgenomen expressie van de contractiele eiwitten sm-�-actine en sm-MHC. Ook verhoogden zowel collageen I als PDGF de proliferatie in primaire humane tracheale luchtweg-gladde spiercellen. Net als bij de rundertracheale gladde spier (Hoofdstuk 3), werd de PDGF-geïnduceerde celproliferatie additief


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verhoogd door collageen I, terwijl er geen additieve effecten werden gevonden op contractiliteit of contractiele eiwitexpressie. De resultaten gepresenteerd in hoofdstuk 6 suggereren dat deze resultaten mogelijk niet alleen gelden voor collageen I maar ook voor fibronectine, aangezien kweken van humane luchtweg-gladde spiercellen op fibronectine ook de proliferatie verhoogde. Deze resultaten tonen aan dat ECM-geïnduceerde fenotypemodulatie ook de contractiele functie van humaan glad spierweefsel beïnvloedt, en tevens dat rundertracheaal glad spierweefsel een representatief experimenteel model vormt voor de fenotypische plasticiteit van humaan weefsel. Naast een toename van de gladde spiermassa is ook de expressie van contractiele eiwitten toegenomen in de astmatische luchtweg-gladde spier (Hoofdstuk 2). In vitro neemt tijdens serumdeprivatie de expressie van contractiele eiwitten toe, wat nog versterkt wordt in aanwezigheid van insuline of TGF-�. Eerder onderzoek heeft aangetoond dat daarnaast ook de expressie van de laminine �2-, �1- en �1-ketens toeneemt tijdens serumdeprivatie. Deze toename was nodig voor de toegenomen contractiele eiwitexpressie, aangezien de laminine-blokkerende peptides Tyr-Ile-Gly-Ser-Arg (YIGSR) en Gly-Arg-Gly-Asp-Ser-Pro (GRGDSP) de toename in contractiele eiwitexpressie remden. Tijdens serumdeprivatie nam ook de expressie van de laminine-bindende integrines �3, �6 en �7 toe, waarvan het �7 integrine noodzakelijk was voor de toename in contractiele eiwitexpressie.

In ons laboratorium was eerder aangetoond dat blootstelling van rundertracheale spierstrips aan insuline resulteert in de inductie van een hypercontractiel, hypoproliferatief fenotype, gekarakteriseerd door toegenomen contractiliteit, toegenomen contractiele eiwitexpressie en een verlaagde proliferatieve respons. Hoofdstuk 5 beschrijft de rol van laminines aan de inductie van een hypercontractiel, hypoproliferatief fenotype door insuline. De resultaten laten zien dat de toename in methacholine- and KCl-afhankelijke contracties door insuline wordt geremd door YIGSR en RGDS, terwijl GRADSP geen effecten had. Ook de toegenomen sm-MHC-expressie en de afgenomen proliferatieve respons na insulinestimulatie werden genormaliseerd door YIGSR. In strips behandeld met insuline nam de expressie van de laminine �2, �1 en �1 ketens tijdsafhankelijk toe, wat suggereert dat laminine-211 belangrijk is voor de inductie van een hypercontractiel fenotype door insuline. In overeenstemming met eerdere waarnemingen voor de toename van contractiele eiwitexpressie, waren activatie van zowel de PI3-kinase als de Rho-kinase signaaltransductieroute ook belangrijk voor de toename in de laminine-expressie en de hypercontractiliteit onder invloed van insuline. De resultaten tonen aan dat laminines een kritische rol spelen in de inductie van een hypercontractiel, hypoproliferatief fenotype door insuline. De toegenomen laminine-expressie in de luchtwegen van astmapatiënten zou dus kunnen bijdragen aan de toegenomen contractiele eiwitexpressie en contractiliteit van de astmatische luchtweg-gladde spier. Daarnaast zouden deze bevindingen consequenties


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kunnen hebben voor het gebruik van geïnhaleerd insuline bij de behandeling van diabetes mellitus. De acute en chronische effecten van insuline op de functie en het fenotype van de humane luchtweg-gladde spier zouden kunnen leiden tot een afname van de longfunctie, zowel onder fysiologische als onder pathofysiologische omstandigheden. Zoals eerder beschreven, communiceren luchtweg-gladde spiercellen en de ECM met elkaar via integrines, een groep heterodimere transmembraaneiwitten. Celkweekstudies hebben aangetoond dat integrines belangrijk zijn voor de adhesie, maturatie, synthetische functie, overleving en proliferatie van luchtweg-gladde spiercellen. Diverse integrines zijn belangrijk voor de proliferatie van luchtweg-gladde spiercellen . Het collageen-bindende integrine �2�1 en de fibronectine-bindende integrines �4�1 en �5�1 zijn betrokken bij de toename van de groeifactor-geïnduceerde proliferatie op zowel collageen I als fibronectine. Daarnaast speelt het �5�1 integrine ook een rol bij de serum-geïnduceerde proliferatie van zowel astmatische als niet-astmatische gladde spiercellen. Ook de verhoogde synthetische functie van de astmatische gladde spiercel is afhankelijk van het �5�1 integrine.

Hoewel integrines dus belangrijk zijn voor de luchtweg-gladde spiercel functie in vitro, is de rol van integrines aan de luchtweg-gladde spierveranderingen bij astma in vivo nog niet onderzocht. In een caviamodel voor chronisch allergisch astma werd de rol van ECM eiwitten en van integrines in luchtwegremodelling onderzocht met behulp van het integrine-blokkerende peptide RGDS. Dit peptide bevat de RGD sequentie, die voorkomt in fibronectine, collagenen en laminines, en blokkeert de interactie tussen RGD-bindende integrines en de ECM eiwitten. De resultaten in Hoofdstuk 6 tonen aan dat RGD-bindende integrines en hun endogene agonisten een belangrijke rol spelen in de remodelling van de luchtweg-gladde spier bij chronisch allergisch astma. Lokale toediening van RGDS via de neus remde de gladde spiercelhyperplasie, de toegenomen pulmonale expressie van de proliferatieve marker PCNA en van contractiele eiwitten, en de hypercontractiliteit van de luchtweg-gladde spier, alle het gevolg van herhaalde allergeenblootstelling; het negatieve controlepeptide GRADSP had geen effect. De remmende effecten van RGDS werden waarschijnlijk veroorzaakt door een direct effect op de gladde spiercel, aangezien de allergeen-geïnduceerde luchtwegfibrose en inflammatie niet werden beïnvloed. In humane luchtweg-gladde spiercellen remde RGDS de proliferatie geïnduceerd door collageen I, fibronectine, PDGF en serum, voornamelijk door remming van het �5�1 integrine. In overeenstemming met de resultaten beschreven in Hoofdstuk 5, remde RGDS concentratieafhankelijk de toename in sm-�-actine-expressie, geïnduceerd door serumdeprivatie in aanwezigheid van insuline. Deze resultaten tonen voor het eerst aan dat activatie van integrines betrokken is bij de veranderingen in de luchtweg-gladde spier bij allergisch astma, en dat dit geremd kan worden door een klein peptide dat het RGD-bindingsmotief bevat.


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De resultaten in Hoofdstuk 6 toonden eveneens aan dat activatie van de RGD-bindende integrines het gevolg is van allergeenprovocatie, aangezien geen effecten van RGDS werden waargenomen in controledieren. Activatie van integrines na allergeenprovocatie kan het gevolg zijn van de synthese van nieuwe ECM eiwitten, het lekken van ECM eiwitten uit het plasma naar de gladde spierlaag of door veranderingen in de reeds aanwezige matrix, waardoor zogenaamde ‘matricryptische’ integrine-bindende aminozuursequenties, die normaal verborgen zijn in de matrixeiwitten, bloot komen te liggen. In overeenstemming hiermee was eerder gevonden dat gedenatureerd monomeer collageen I, in tegenstelling tot fibrillair collageen I, de groeifactor-geïnduceerde proliferatie verhoogt, wat suggereert dat veranderingen in de structuur van het matrixeiwit noodzakelijk zijn voor de inductie van proliferatie.

Ook werden er in vitro geen effecten van RGDS of integrine-blokkerende antilichamen gevonden op de proliferatie van gladde spiercellen (Hoofdstuk 6) of de contractiliteit van glad spierweefsel (Hoofdstuk 3) onder ongestimuleerde controleomstandigheden, wat suggereert dat interactie tussen de RGD-bindende integrines en de extracellulaire matrix beperkt is onder deze omstandigheden. De resultaten beschreven in Hoofdstuk 5 suggereren dat laminines mogelijk een belangrijke rol spelen in de verhoogde contractiele eiwitexpressie en hypercontractiliteit van de luchtweg-gladde spier bij astma. In Hoofdstuk 7 werden daarom de effecten van het laminine-blokkerende peptide YIGSR onderzocht in het caviamodel voor chronisch allergisch astma. In tegenstelling tot onze verwachtingen, verlaagde lokale toediening van YIGSR in de luchtwegen de allergeen-geïnduceerde gladde spiercelhyperplasie en pulmonale PCNA expressie. Bovendien verhoogde de behandeling met YIGSR zowel bij de controledieren als bij de allergeen-geprovoceerde dieren de pulmonale contractiele eiwitexpressie en de contractiliteit van de luchtweg-gladde spier. Deze waarnemingen suggereren dat in vivo behandeling met YIGSR de effecten van laminine eerder bevordert dan vermindert. In vitro verlaagde het kweken van humane gladde spiercellen op geïmmobiliseerd YIGSR de PDGF-geïnduceerde proliferatie van deze cellen in vergelijkbare mate als op laminine-111. Opmerkelijk genoeg werden de effecten van zowel geïmmobiliseerd YIGSR als laminine-111 geantagoneerd door YIGSR dat in opgeloste vorm aanwezig was in het kweekmedium. Geconcludeerd wordt dat in vivo behandeling met YIGSR de effecten van laminine bevordert in plaats van remt, wat mogelijk wordt veroorzaakt door de fysisch-chemische micro-omgeving van het peptide. De mechanismen die hieraan ten grondslag liggen zijn echter niet bekend. In diverse celtypen is aangetoond dat specifieke ECM – integrine interacties resulteren in de activatie van intracellulaire signaaltransductieroutes die belangrijk zijn voor de veranderingen in proliferatie, migratie, differentiatie en apoptose van de cel. Verschillende integrines activeren focal adhesion kinase (FAK), dat vervolgens andere intracellulaire signaaltransductieroutes kan


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activeren, zoals de phosphoinositide 3-kinase (PI3-kinase) en de mitogen-activated protein kinase (MAPK) routes. De resultaten beschreven in hoofdstuk 8 tonen aan dat activatie van FAK en de daaraan gekoppelde signaaltransductieroutes een belangrijke rol spelen in de inductie van een proliferatief, hypocontractiel fenotype door collageen I. In rundertracheale gladde spiercellen werd aangetoond dat activatie van FAK in even sterke mate plaats vindt bij adhesie van de cellen aan een ongecoate plastic matrix als aan een collageen I matrix. Activatie van FAK is noodzakelijk voor de celadhesie, aangezien deze werd geremd door overexpressie van de FAK deletiemutanten FAT (het ‘focal adhesion targeting’ (FAT) domein van FAK) en FRNK (het ‘focal-adhesion related non-kinase’ domein), die beide de translocatie van FAK naar de focal adhesion sites en de daaropvolgende activatie van het enzym remmen. De activatie van FAK nam vervolgens gedurende 2-4 dagen verder toe in cellen gekweekt op monomeer collageen I, maar niet in cellen gekweekt op plastic. De vertraagde activatie suggereert dat activatie van FAK door collageen I niet direct, maar via een secundair proces plaats vindt. Een mogelijke verklaring hiervoor kan worden afgeleid uit waarnemingen bij vasculaire gladde spiercellen, waarin de expressie van andere ECM eiwitten, inclusief fibronectine, toeneemt als de cellen worden gekweekt op monomeer collageen I. Ook is de toename in de basale (Hoofdstuk 6) en de groeifactor-geïnduceerde proliferatie van luchtweg-gladde spiercellen op collageen I afhankelijk van het fibronectine-bindende integrine �5�1.

In rundertracheale gladde spiercellen nam de collageen I-geïnduceerde proliferatie zowel concentratie- als tijdafhankelijk toe. Ook deze toename werd geremd door overexpressie van FAT en FRNK. Farmacologische remming van Src, mitogen-activated extracellular kinase (MEK), PI3-kinase en p38 MAPK remde ook de collageen I-geïnduceerde proliferatie en hypocontractiliteit, hetgeen suggereert dat activatie van FAK en daardoor geactiveerde signaaltransductieroutes noodzakelijk zijn voor de collageen I-afhankelijke fenotypeveranderingen van de luchtweg-gladde spiercel. Eerder onderzoek heeft aangetoond dat FAK ook direct betrokken is bij contractie van de gladde spiercel. Uitschakeling van FAK-expressie in humane tracheale gladde spierstrippen resulteerde in een afgenomen krachtsontwikkeling, een verminderde fosforylering van de myosine lichte keten en een afgenomen calcium respons na stimulatie met KCl en acetylcholine. Deze waarnemingen suggereren dat remming van FAK mogelijk een interessant target is voor de behandeling van zowel acute als persistente luchtweghyperreactiviteit. Samengevat geven onze bevindingen aan dat interacties met de ECM belangrijk zijn voor (de regulatie van) het fenotype van de luchtweg-gladde spiercel. Veranderingen in het extracellulaire milieu rond de astmatische gladde spiercel dragen daardoor mogelijk in belangrijke mate bij aan de toegenomen contractiele, proliferatieve en synthetische kenmerken van deze cel. Daarnaast


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is de luchtweg-gladde spiercel mogelijk zelf verantwoordelijk voor het creëren van een ECM milieu dat deze abnormale functies ondersteunt. Effecten van glucocorticosteroïden en �2-adrenoceptor agonisten op fenotypeveranderingen van de luchtweg-gladde spier Glucocorticosteroïden en �2-adrenerge receptoragonisten zijn op dit moment de meest effectieve middelen voor de behandeling van astma, vanwege hun anti-inflammatoire respectievelijk bronchusverwijdende eigenschappen. Daarnaast is in diermodellen van astma aangetoond dat behandeling met glucocorticosteroïden de toename in spiermassa voorkomt, wat suggereert dat deze middelen ook luchtwegremodelling kunnen beïnvloeden. In vitro remmen beide geneesmiddelen de groei van luchtweg-gladde spiercellen. Wanneer de cellen echter worden gekweekt op collageen I zijn de remmende effecten van glucocorticosteroïden verminderd. Dit suggereert dat de toename van collageen I-expressie rond de astmatische luchtweg-gladde spier niet alleen de proliferatie bevordert, maar ook kan bijdragen aan steroïdresistentie, zoals waargenomen in een subgroep van de patiënten met ernstig astma. Combinatietherapie met glucocorticosteroïden en �2-adrenerge receptoragonisten leidt over het algemeen tot een effectievere astmabehandeling dan monotherapie met elk van de middelen afzonderlijk. In humaan longweefsel is aangetoond dat glucocorticosteroïden een verhoogde gentranscriptie van de �2-adrenerge receptor induceren, terwijl in humane luchtweg-gladde spiercellen was waargenomen dat glucocorticosteroïden en �2-agonisten een synergistisch effect hebben op de translocatie van de glucocorticosteroid receptor. De impact van dit synergisme op fenotypeveranderingen van de luchtweg-gladde spier zijn echter nog niet onderzocht. De resultaten beschreven in Hoofdstuk 9 tonen aan dat de glucocorticosteroïden fluticason, budesonide en dexamethason zowel de PDGF- en collageen I-geïnduceerde proliferatie als de afname in de contractiliteit remmen, hoewel de effecten van collageen I minder gevoelig voor de glucocorticosteroïden waren. De effecten van PDGF en collageen I op de proliferatie werden ook geremd door de �2-agonist fenoterol. In honderdvoudig lagere concentraties, waarbij niet of nauwelijks meer een effect van deze middelen afzonderlijk werd waargenomen, remde de combinatie van fluticason en fenoterol synergistisch zowel de PDGF- als de collageen I-geïnduceerde fenotypeveranderingen. Aangezien toegenomen gladde spiermassa waarschijnlijk een belangrijke bijdrage levert aan de toegenomen luchtwegweerstand en persistente luchtweghyperreactiviteit bij chronisch astma, suggereren deze waarnemingen dat de combinatie van glucocorticosteroïden en �2-agonisten de progressie van de ziekte zouden kunnen dempen. Dat de groeifactoren collageen I-geïnduceerde fenotypeveranderingen even effectief door de glucocorticosteroïd-�2-agonist combinatie werden geremd suggereert dat naast het verhogen van de anti-inflammatoire, bronchusverwijdende en


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antiproliferatieve effecten, combinatietherapie met deze middelen ook de ontwikkeling van steroïdresistentie zou kunnen afremmen. Conclusies De belangrijkste conclusies van dit proefschrift zijn: ECM eiwitten hebben differentiële effecten op het fenotype en de

contractiele functie van de luchtweg-gladde spier. Collageen I and fibronectine induceren een functioneel hypocontractiel fenotype, geassocieerd met een toegenomen celproliferatie, terwijl laminine-111 een contractiel fenotype bevordert en groeifactor-geïnduceerde proliferatie remt (Hoofdstuk 3 & 6).

Net als in rundertracheaal spierweefsel induceren collageen I en PDGF een proliferatief en functioneel hypocontractiel fenotype in humaan tracheaal-glad spierweefsel (Hoofdstuk 4).

De insulinegeïnduceerde toename van laminine �2-, �1- en �1-ketens, gemedieerd door activatie van PI3-kinase en Rho-kinase, draagt bij aan de inductie van een hypercontractiel, hypoproliferatief fenotype van de luchtweg-gladde spier. Toegenomen laminine-expressie in de luchtwegwand zou bij kunnen dragen aan de toegenomen contractiliteit van de luchtweg-gladde spier en contractiele eiwitexpressie bij astma (Hoofdstuk 5)

De toegenomen expressie van contractiele eiwitten en laminine als gevolg van blootstelling aan insuline maakt de toepassing van geïnhaleerd insuline bij de behandeling van diabetes mellitus problematisch (Hoofdstuk 5).

Endogene activatie van RGD-bindende integrines draagt bij aan de door allergeen geïnduceerde luchtweg-gladde spierveranderingen in een caviamodel voor allergisch astma. Dit proces kan worden geremd door lokale toediening van RGDS, een klein peptide dat het RGD-bindingsmotief in verschillende ECM-eiwitten bevat. Gebaseerd op deze waarnemingen zouden RGD-bindende integrines een nieuwe strategie kunnen vormen voor de behandeling van luchtwegremodelling bij astma (Hoofdstuk 6).

Het laminine blokkerende peptide YIGSR remt de inductie van een hypercontractiel, hypoproliferatief fenotype van de luchtweg-gladde spier in vitro (Hoofdstuk 5). Daarentegen bevordert in vivo behandeling met YIGSR de inductie van een contractiel, hypoproliferatief fenotype, hetgeen afhankelijk zou kunnen zijn van de fysisch-chemische micro-omgeving van het peptide (Hoofdstuk 7).

Activatie van FAK is noodzakelijk voor de adhesie van luchtweg-gladde spiercellen aan zowel plastic als collageen I, alsmede voor collageen I-geïnduceerde proliferatie van deze cellen (Hoofdstuk 8)

Fenotypische modulatie van luchtweg-gladde spiercellen door collageen I is afhankelijk van Src, MEK, p38 MAK en PI3-kinase, die allen geactiveerd kunnen worden door FAK (Hoofdstuk 8)


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Glucocorticosteroïden remmen PDGF- en collageen I-geïnduceerde fenotypeveranderingen van de luchtweg-gladde spier, hoewel de effecten van collageen I minder gevoelig zijn voor glucocorticosteroïden dan die van PDGF. In lage concentraties remmen glucocorticosteroïden en �2-adrenerge receptoragonisten synergistisch de inductie van een proliferatief, hypocontractiel fenotype door zowel PDGF als collageen I, zonder een verschil tussen beide mitogenen. Dit synergisme zou kunnen bijdragen aan de verhoogde effectiviteit van de combinatietherapie bij astma, mede door vermindering van de glucocorticosteroïdresistentie geïnduceerd door collageen I (Hoofdstuk 9).

Dankwoord

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Dankwoord Een promotieonderzoek samenvatten in slechts een paar hoofdstukken was al een uitdaging, maar iedereen bedanken die hierbij betrokken is geweest in slechts een paar pagina’s lijkt helemaal onmogelijk. Maar uiteraard wil ik iedereen bedanken die dit proefschrift direct of indirect heeft mogelijk gemaakt. Een aantal mensen wil ik in het bijzonder noemen.

Ten eerste mijn promotores Prof. Dr. H. Meurs en Prof. Dr. J. Zaagsma. Beste Herman, zonder jouw enthousiasme en kennis van zaken rondom de luchtwegfarmacologie was dit proefschrift niet mogelijk geweest. Na een bespreking met jou liep ik altijd vol met ideeën en nieuwe vragen de deur weer uit. Ook wil ik mijn dank uitspreken voor de vrijheid die ik heb gekregen om vorm en richting aan het onderzoek te geven. Hoewel in eerste instantie wat confronterend, heeft jouw aandacht voor details en schrijfstijl er toe geleid dat mijn schrijfvaardigheid sterk verbeterd is en het eindresultaat mag er dan ook zijn. Ik heb de laatste vijf jaar veel van je geleerd en ik ga ervan uit dat dit de komende jaren ook zo zal blijven.

Beste Hans, toen ik nog farmacie studeerde was jij nog het hoofd van de Basiseenheid Moleculaire Farmacologie. Jij was dan ook degene die mij het eerst benaderde voor dit project. Je vroeg of ik interesse had in het doen van een promotieonderzoek bij de vakgroep en of je mij mocht bellen als je financiering had. Na een positief antwoord van mijn kant werd ik vervolgens enige maanden later – vlak voor de kerst – tijdens mijn apotheekstage in Assen gebeld en zat ik vervolgens tijdens de facultaire kerstborrel met jou en Herman rond de tafel om het project door te nemen. Dat was 4 ½ jaar geleden, nu ben je met emeritaat, maar je staat nog (bijna) elke dag klaar om mij met je kennis als ‘functionele/klassieke’ receptorfarmacoloog op te voeden. Hans bedankt voor je waardevolle bijdrage! Ook wil ik Prof. Dr. M. Schmidt en Dr. S.A. Nelemans bedanken. Lieber Martina, hoewel jij minder direct betrokken bent bij mijn onderzoek, weet jij wel altijd de juiste vragen te stellen en een twist te geven aan de discussies. Beste Ad, jouw enthousiasme tijdens mijn afstudeerproject heeft er voor gezorgd dat ik zelf ook enthousiast werd voor het farmacologisch onderzoek. I would also like to thank the members of the reading committee, Prof Dr. A.J. Halayko, Prof Dr. K. Racké and Prof Dr. W. Timens. Professor Halayko and Professor Racké, thank you for being part of the reading committee and for taking the time and effort to carefully read my manuscript. Professor Timens, bedankt voor uw bijdrage in het beoordelen van mijn proefschrift.

Verder wil ik Sophie Bos nog speciaal bedanken. Sophie, jouw bijdrage als analist was van onschatbare waarde, niet alleen voor mijn introductie in het dierexperimentele werk, maar ook bij tal van andere (vervolg)experimenten. Ook onze gezellige etentjes en te gezellige borrels, zoals bij de FIGON dagen, zal ik niet snel vergeten. Bedankt dat je mijn paranimf wil zijn!

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Natuurlijk wil ik ook alle (ex)collega’s van de Basiseenheid Moleculaire Farmacologie bedanken. Anita, Annet, Anouk, Carolina, Dedmer, Efi, Harm, Hoeke, Jacques, Janneke, Jelte, Marieke, Mark (2x), Reinoud, Sara, Tjitske en Tonio: bedankt voor jullie interesse en discussies en voor de gezelligheid tijdens de koffiepauzes en lunches! Dedmer, ik zou jou speciaal willen bedanken, jouw hulp en ondersteuning tijdens het opstarten van dit project heeft er voor gezorgd dat het project een vliegende start kreeg, wat resulteerde in een snelle publicatie. Ook de ‘vrijdagmiddagdiscussies’ samen met Tonio in de Groote Griet of de Pintelier zal ik niet snel vergeten.

Daarnaast hebben verschillende studenten farmacie en farmaceutische wetenschappen in het kader van hun afstudeerproject bijgedragen aan dit proefschrift: Abdullah, Adnan, Anita, Dirk Jan, Erwin, Hans, Robert en Willem Jan, het was erg inspirerend om met jullie onderzoek te doen, bedankt voor jullie inzet!

Experimenteel onderzoek is alleen mogelijk bij de gratie van anderen. Allereerst wil ik het Astma fonds bedanken voor het financieren van dit project. Ook wil ik graag de medewerkers van de slachthuizen in Groningen, Surhuisterveen, Eelde en Tolbert bedanken voor het leveren van de noodzakelijke rundertrachea’s. De afdeling thoraxchirurgie wil ik bedanken voor het beschikbaar stellen van humaan luchtwegweefsel. De medewerkers van de campusstore, de instrumentmakerij en de centrale dienst proefdieren (CDP) wil ik ook van harte bedanken voor de ondervonden medewerking. Tenslotte wil ik de Graduate school voor Cognitive and Behavioral Neurosciences (BCN) en de Nederlandse Stichting voor Farmacologische Wetenschappen (NSFW) bedanken voor de financiële ondersteuning voor de deelname aan congressen.

Uiteraard wil ik ook mijn familie en vrienden bedanken. Jullie interesse in mijn onderzoek, ook al was het maar moeilijk te begrijpen, en jullie goede raad en tips hebben mij erg geholpen. Pap en mam bedankt voor jullie onvoorwaardelijke steun, liefde, geduld en vertrouwen. Ton bedankt voor alle lol die we altijd samen hebben gehad, vooral tijdens het drinken van een biertje. Ik vind het ontzettend leuk dat je mijn paranimf wil zijn.

Tenslotte wil ik mijn lieve vriendin Femke bedanken. Woorden kunnen niet beschrijven wat je voor mij betekent. Samen met Jesper en Camyl zorg je er voor dat er buiten het werk ook nog een wereld bestaat waar ik tot mezelf kan komen en ik me kan ontspannen – ook al is dit er de laatste tijd wat minder van gekomen. Ik weet zeker dat we in de toekomst onze weg zullen vinden.

Bart Groningen, juni 2010

Curriculum Vitae

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Curriculum Vitae The author of this thesis was born in Hardenberg, The Netherlands, on the 28th of January 1981. After finishing his pre-university education (Gymnasium, Almelo) in 1999, he moved to Groningen and started his Pharmacy study at the University of Groningen. During his studies he was a studentassistent in a practical analytical chemistry course for year 1 pharmacy students (2002-2003) and a pharmacological course for year 3 pharmacy students (2003) at the University of Groningen. He performed his graduation project on the effects of (endo)cannabinoids on [Ca2+]i and identification of Ca2+-channels in human bronchial epithelial cells and smooth muscle cells at the Department of Molecular Pharmacology and graduated cum laude in March 2004. He continued his studies at the University of Groningen to become a pharmacist and graduated in August 2005. After graduation, he initiated his PhD-study at the Department of Molecular Pharmacology, where he worked on a research project funded by the Netherlands Asthma Foundation (3.2.03.36), entitled: ‘Effects of extracellular matrix proteins on airway smooth muscle contractility and proliferation in chronic asthma’ , the results of which are presented in this thesis. In the forthcoming three years he will be active as a lecturer and a post-doc at the same department.

List of publications

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List of publications Full papers BGJ Dekkers, J Zaagsma, H Meurs. Functional consequences of human airway smooth muscle phenotype plasticity. Submitted 2010 SS Roscioni, BGJ Dekkers, AG Prins, H Meurs, M Schmidt, H Maarsingh. cAMP inhibits airway smooth muscle phenotype modulation via Epac and PKA. Submitted 2010 H Maarsingh, BGJ Dekkers, AB Zuidhof, IST Bos, J Zaagsma, H Meurs. Increased arginase activity contributes to airway remodeling, inflammation and hyperresponsiveness in chronic allergic asthma. Submitted 2010 BGJ Dekkers, IST Bos, R Gosens, AJ Halayko, J Zaagsma, H Meurs. Activation of RGD-binding integrins regulates allergen-induced airway smooth muscle remodeling. Am J Respir Crit Care Med, 2010, 181(6):556-565. BGJ Dekkers, H Maarsingh, H Meurs, R Gosens. Airway structural components drive airway smooth muscle remodeling in asthma. Proc Am Thorac Soc, 2009, 6(8):683-692. BGJ Dekkers, D Schaafsma, T Tran, J Zaagsma, H Meurs. Insulin-induced laminin expression promotes a hypercontractile airway smooth muscle phenotype. Am J Respir Cell Mol Biol, 2009, 41(4):494-504. E Gkoumassi, BGJ Dekkers, MJ Dröge, CRS Elzinga, RE Hasenbosch, H Meurs, SA Nelemans, M Schmidt, J Zaagsma. (Endo)cannabinoids mediate different Ca2+ entry mechanisms in human bronchial epithelial cells. Naunyn Schmiedebergs Arch Pharmacol, 2009, 380(1):67-77. R Gosens, SS Roscioni, BGJ Dekkers, T Pera, M Schmidt, D Schaafsma, J Zaagsma, H Meurs. Pharmacology of airway smooth muscle proliferation. Eur. J. Pharmacol.,2008, 13:585(2-3):385-97. BGJ Dekkers, D Schaafsma, SA Nelemans, J Zaagsma, H Meurs. Extracellular matrix proteins differentially regulate airway smooth muscle phenotype and function. Am. J. Physiol. Lung Cell. Mol. Physiol. 2007, 292:L1405-13. D Schaafsma, KD McNeill, GL Stelmack, R Gosens, HA Baarsma, E Frohwork, BGJ Dekkers, JM Penninks, KM Ens, SA Nelemans, J Zaagsma, AJ Halayko, H Meurs. Insulin increases expression of contractile phenotypic markers in airway smooth muscle. Am. J. Physiol Cell Physiol. 2007, 293:C429-39.


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E Gkoumassi, MJ Dröge, BGJ Dekkers, CRS Elzinga, M Schmidt, H Meurs, J Zaagsma, SA Nelemans. (Endo)cannabinoids modulate cAMP production and IL-8 release in human bronchial epithelial cells. Br. J. Pharmacol. 2007, 151:1041-8. DG Demuth, E Gkoumassi, MJ Droge, BGJ Dekkers, HJ Esselink, RM van Ree, ME Parsons, J Zaagsma, A Molleman, SA Nelemans. Arachidonic acid mediates non-capacitative calcium entry evoked by CB1-cannabinoid receptor activation in DDT1 MF-2 smooth muscle cells. J. Cell Physiol. 2005, 205: 58-67. Published abstracts BGJ Dekkers, A Pehli, H Meurs, J Zaagsma. Glucocorticosteroids and �2-adrenoceptor agonists synergistically prevent the induction of a hypocontractile, proliferative airway smooth muscle phenotype. Am J Respir Crit Care Med 2010, 181, A5308 BGJ Dekkers, J Zaagsma, H Meurs. Functional consequences of human airway smooth muscle phenotype plasticity,. Am J Respir Crit Care Med 2010, 181, A2133 SS Roscioni, BGJ Dekkers, AG Prins, G Oldenbeuving, KME Pool, CRS Elzinga, H Meurs, M Schmidt. Epac and PKA Inhibit PDGF-Induced Airway Smooth Muscle Phenotype Modulation. Am J Respir Crit Care Med 2010, 181, A2142 H Maarsingh, AB Zuidhof, IST Bos, BGJ Dekkers, J Zaagsma, H Meurs. Arginase inhibition protects against allergen-induced airway remodeling, hyperresponsiveness and inflammation in chronic asthma. Am J Respir Crit Care Med 2010, 181, A2307 BGJ Dekkers, A Pehli, H Meurs, J Zaagsma. Glucocorticosteroids and �2-adrenoceptor agonists synergistically inhibit airway smooth muscle phenotype modulation. Naunyn-schmiedeberg Archives of Pharmacology 2010, 381, 250 H Maarsingh, BGJ Dekkers, AB Zuidhof, IST Bos, J Zaagsma, H Meurs. Increased arginase activity underlies allergen-induced airway remodeling, fibrosis, inflammation and hyperresponsiveness in chronic asthma. Naunyn-schmiedeberg Archives of Pharmacology 2010, 381, 251 M Schmidt, SS Roscioni, BGJ Dekkers, AG Prins, H Meurs. Epac and PKA inhibit PDGF-induced airway smooth muscle phenotype modulation. Naunyn-schmiedeberg Archives of Pharmacology 2010, 381, 252 BGJ Dekkers, AIR Spanjer, J Zaagsma, H Meurs. Role for proliferative signalling pathways in collagen I-induced phenotype modulation of airway smooth muscle. Naunyn-schmiedeberg Archives of Pharmacology 2009, 380, 259


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BGJ Dekkers, IST Bos, AJ Halayko, J Zaagsma and H Meurs. The laminin �1-competing peptide YIGSR induces a hypercontractile, hypoproliferative airway smooth muscle phenotype in an animal model of chronic asthma. Am J Respir Crit Care Med 2009, A5056 BGJ Dekkers, IST Bos, R Gosens, AJ Halayko, J Zaagsma and H Meurs. Inhibition of airway smooth muscle remodeling in an animal model of chronic asthma by the integrin-blocking peptide RGDS. Am J Respir Crit Care Med 2009, A5600 BGJ Dekkers, D Schaafsma, J Zaagsma, H Meurs. Insulin-induced laminin expression promotes a hypercontractile airway smooth muscle phenotype. Naunyn-schmiedeberg Archives of Pharmacology 2009, 379:202 BGJ Dekkers, D Schaafsma, T. Tran, J Zaagsma, H Meurs. Increased expression of laminin is required for the insulin-induced hypercontractile airway smooth muscle phenotype. Am J Respir Crit Care Med 2008, A327 BGJ Dekkers, AIR Spanjer, J Zaagsma, H Meurs. Signaling pathways of collagen I-induced phenotype modulation of airway smooth muscle. Am J Respir Crit Care Med 2008, A489 BGJ Dekkers, D Schaafsma, SA Nelemans, J Zaagsma, H Meurs. Extracellular matrix proteins differentially regulate airway smooth muscle phenotype and function. Naunyn-schmiedeberg Archives of Pharmacology 2007, 375:148 BGJ Dekkers, D Schaafsma, SA Nelemans, J Zaagsma, H Meurs. Functional role for extracellular matrix proteins in phenotypic modulation of intact bovine tracheal smooth muscle. Proc Am Thor Soc 2007, A302 E Gkousmassi, MJ Dröge, CRS Elzinga, BGJ Dekkers, H Meurs, J Zaagsma, SA Nelemans. Dual modulation of adenylyl cyclase by cannabinoids in human bronchial epithelial cells. 16th Symposium on Cannabinoids, International Canabinoid Research Society (ICRS) symposium 2006, Tihany, Hungary, P94. E Gkoumassi, MJ Dröge, BGJ Dekkers, CRS Elzinga, H Meurs, J Zaagsma, SA Nelemans. Cannabinoid signaling in human bronchial epithelial cells. 3rd conference on Cannabinoids in Medicine, International Association for Cannabis as Medicine (IACM) Meeting 2005, Leiden, The Netherlands. E Gkoumassi, MJ Droge, BGJ Dekkers, CRS Elzinga, H Meurs, J Zaagsma, SA Nelemans. Cannabinoid signaling in human bronchial epithelial cells. Proc Am Thor Soc 2005, 2: A754 E Gkoumassi, MJ Dröge, BGJ Dekkers, HJ Esselink, H Meurs, J Zaagsma and SA Nelemans. Cannabinoid signal transduction and cytokine release. Fundam Clin Pharmacol, P09.03

List of abbreviations

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List of abbreviations ACh Acetylcholine ADAM A Disintegrin And Metalloproteinase AHR Airway hyperresponsiveness ANOVA Analysis of variance ASM Airway smooth muscle AMP Adenosine monophosphate ATP Adenosine triphosphate BAL Bronchoalveolar lavage bFGF Basic fibroblast growth factor BSA Bovine serum albumin BTSM Bovine tracheal smooth muscle cAMP Cyclic adenosine monophosphate CRC Concentration response curve C/EBP� CCAAT/enhancer binding protein � cGMP Cyclic guanosine monophosphate Coll Collagen COPD Chronic obstructive pulmonary disease CTGF Connective tissue growth factor DMEM Dulbecco’s modified eagle medium ECM Extracellular matrix EGF Epidermal growth factor EHS Engelberth-Holm-Swarm eIF4E Eukaryotic initiation factor 4E EC50 Concentration of the stimulus eliciting half-maximal response Emax Maximal contraction eNOS Endothelial NOS Epac Exchange protein directly activated by cAMP ERK Extracellular signal-regulated kinase FAK Focal adhesion kinase FAT Focal adhesion targeting FBS Foetal bovine serum FN Fibronectin FRNK FAK-related non-kinase GAG Glycosaminoglycan GFP Green fluorescent protein GM-CSF Granulocyte macrophage colony-stimulating factor GPCR G protein coupled receptor GRADSP Glycine-Arginine-Alanine-Aspartic acid-Serine-Proline GRE Glucocorticosteroid response element GRGDSP Glycine-Arginine-Glycine-Aspartic acid-Serine-Proline GSK-3 Glycogen synthase kinase 3 HBSS Hank’s buffered salt solution HRP Horseradish peroxidase hTERT Human telomerase reverse transcriptase HTSM Human tracheal smooth muscle Ig Immunoglobulin iNANC Inhibitory nonadrenergic, noncholinergic

List of abbreviations

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iNOS inducible NOS IL interleukin ILK Integrin linked kinase ITG Integrin ITS Insulin, transferrin and selenium KH Krebs-Henseleit LN Laminin MAPK Mitogen activated protein kinase MBP Major basic protein MMP Matrix metalloproteinase mTOR Mammalian target of rapamycin MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide nNOS neuronal NOS NO Nitric oxide NOS NO synthase ODC Ornithine decarboxylase PBS Phosphate-buffered saline PCNA Proliferatng cell nuclear antigen PDGF Platelet-derived growth factor pEC50 Negative logarithm of the EC50 value. PGE2 Prostaglandin E2 P70S6K p70 ribosomal S6 kinase PI3-kinase Phosphatidyl inositol 3-kinase PKA Protein kinase A PKB Protein kinase B PKC Protein kinase C RANTES Regulated upon Activation, Normal T-cell Expressed and Secreted Rb Retinoblastoma protein RGD Arginine-Glycine-Aspartic acid RGDS Arginine-Glycine-Aspartic acid-Serine RT Room temperature RTK Receptor tyrosine kinase SDS/PAGE Sodium dodecyl sulfate/polycrylamide gel electrophoresis SLRP Small leucine rich proteoglycan SINNNR Ser-Ile-Asn-Asn-Asn-Arg sm22 Transgelin sm-�-actin Smooth muscle �-actin sm-MHC Smooth-muscle-specific myosin heavy chain sm-MLCK Smooth muscle myosin light chain kinase SPARC Secreted protein acid and rich in cysteine SRF Serum response factor TBST Tris-buffered saline with tween TGF-� Transforming growth factor-� TIMP Tissue Inhibitor of MMPs YIGSR Tyrosine-Isoleucine-Glycine-Serine-Arginine VEGF Vascular endothelial growth factor VN Vitronectin VSMC Vascular smooth muscle cell

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University of Groningen Major role of the extracellular ... · Chapter 1 8 The extracellular matrix Within the human body, tissue cells reside in a three-dimensional protein network,