To my family

List of Papers

This thesis is based on the following papers, which are referred to in the text by their Roman numerals.

I Sundberg*, C., Friman*, T., Hecht, L.E., Kuhl, C., and Solomon, K.R. (2009) Two Different PDGF β-Receptor Cohorts in Hu-man Pericytes Mediate Distinct Biological Endpoints. Am J Pa-thol, 175: 171-189 * indicates equal author contribution.

II Friman*, T., Renata Gustafsson*, R., Burmakin, M., Hellberg, C., Heldin, CH., Oldberg, Å., Rubin, K. (2010) Stromal effects of PDGFR inhibition by STI571 in experimental carcinoma: Al-tered collagen fibrils and reduction of a specific stromal cell population. (Manuscript) * indicates equal author contribution.

III Gustafsson*, R., Friman*, T., Stuhr, L., Chidiac, J., Heldin, NE., Reed, RK., Oldberg Å., Rubin, K. (2010) Integrin αVβ3 re-strains fibrosis and elevated interstitial fluid pressure in synge-neic mouse carcinoma (Manuscript) * indicates equal author contribution.

IV Rodriguez, A., Friman, T., Gustafsson, R., Kowanetz, M., van Wieringen, T., Sundberg, C. (2010) Phenotypical differences in connective tissue cells emerging from microvascular pericytes in response to over-expression of PDGF-B and TGF-β1 in nor-mal skin in vivo. (Manuscript)

Reprints were made with permission from the respective publishers.

Contents

Introduction...................................................................................................11 Loose connective tissue............................................................................11 The tumor microenvironment...................................................................12 The ECM of loose connective tissue........................................................12

Fibrillar components............................................................................12 Non-fibrillar components of the ECM.................................................19

Cells of the loose connective tissue..........................................................21 Cell-ECM interactions..............................................................................24

Integrins ...............................................................................................24 Blood and lymph vessels in loose connective tissue ................................26

The microvasculature...........................................................................26 The lymphatics of loose connective tissue ..........................................29

Growth factors that are important for tissue activation, remodeling and fibrosis......................................................................................................29

Platelet derived growth factor (PDGF)................................................29 Platelet derived frowth factor receptors (PDGFR) ..............................33 Transforming growth factor β (TGF-β)...............................................34

Fluid homeostasis in loose connective tissue ...........................................37 Reactive tissue conditions ........................................................................40

Inflammation .......................................................................................40 Fibrosis ................................................................................................48

The microenvironment of solid tumors ....................................................49 Inflammatory cells in the tumor microenvironment ............................50 The tumor vasculature .........................................................................51 Tumor associated fibroblasts ...............................................................52 The ECM scaffold in tumors ...............................................................55

Present investigation .....................................................................................59 Aims .........................................................................................................59

Paper I..................................................................................................59 Paper II ................................................................................................61 Paper III ...............................................................................................63 Paper IV...............................................................................................65

Future perspectives .......................................................................................67

Acknowledgements.......................................................................................69

References.....................................................................................................71

Abbreviations

ALK Activin like kinase BM Basement membrane BMP Bone morphogenic protein CNS Central nervous system CUB Complement (C1r/C1s), Uegf and Bmp1 EGF Epidermal growth factor EGFR Epidermal growth factor receptor EMT Epithelial to mesenchymal transition FACIT Fibril associated collagens with interupted triple helix FAK Focal adhesion kinase FGF Fibroblast growth factor GAG Glycosaminoglycan HSPG Heparan sulfate proteoglycan IL Interleukin LAP Latency associated peptide LLC Large latent complex LOX Lysyloxidase LTBP Large TGF-β-binding protein MHC II Major histocompability complex II MMP Matrix metalloproteinase PDGF Platelet derived growth factor PDGFR Platelet derived growth factor receptor SLRP Small leucin-rich repeat proteoglycan SPARC Secreted Protein, Acidic and Rich in Cystein s.c. sub cutaneous TAF Tumor associated fibroblast TGF Transforming growth factor TβR Transforming growth factor receptor TNF Tumor necrosis factor TSP Thrombospondin VEGF Vascular endothelial growth factor VEGFR Vascular endothelial growth factor receptor vSMC Vascular smooth muscle cell WT Wild type

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Introduction

Loose connective tissue Connective tissue separates the parenchyma of a tissue from the blood ves-sels that nourishes it. The parenchyma supplies a tissue or organ with its function and there are many different types of parenchyma, such as epithelial glands and linings as well as different types of muscle. The loose connective tissue can also be referred to as the interstitium, although the term inter-stitium or interstitial space by some is defined as the extra-vascular space that is not occupied by cells. The loose connective tissue is situated between the basement membranes (BM) of epithelial parenchyma and blood vessels. The loose connective tissue can also be continuous with areas of dense con-nective tissue, which is seen in the different layers of the dermis. However, the two different types of connective tissue are continuous and not separated from each other by a BM. Apart from blood vessels, nerves and lymphatic vessels also run through the loose connective tissue. Lymphatic vessels most often have a discontinuous BM, whereas large nerves are delimited by a BM and a coat of dense connective tissue, and smaller nerves only have a BM. Connective tissue is made up by a fibrillar matrix network into which cells and ground substance are interspersed. The ground substance comprises the filling material and is made up by different kinds of glycoproteins, pro-teoglycans and glycosaminoglycans (GAG). The major difference between loose and dense connective tissue lie in the amount of the fibrillar compo-nent, which is more abundant in dense connective tissue. The extracellular matrix (ECM) and the different cell types of loose connective tissue will be described in detail below.

At a glance, the obvious function of the loose connective tissue is to provide structure and mechanical support to tissues and organs, but loose connective tissues also regulates the local fluid balance. The stromal cells of the loose connective tissue are reactive to pathological stimuli. Immune reactions are often elicited in this tissue and loose connective tissue constitutes the battle-ground where the body is trying to fight off foreign invaders. This infer that when it comes to pathological stimuli and fluid balance, the loose connective tissue can be viewed as a buffer zone, situated between different compart-ments. As will be discussed below, pathological stimuli and fluid balance in loose connective tissue are tightly interconnected.

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The tumor microenvironment Carcinomas are malignant neoplasias that arise from epithelial cells, and lead to death and morbidity if left untreated. Carcinomas or other solid tumors are comprised of transformed malignant cells embedded in a stroma that conaiata of extracellular matrix (ECM), blood vessels, inflammatory and mesenchy-mal cells[1]. In contrast to normal tissue, the stroma in solid tumors is acti-vated in a way that is reminiscent of chronic inflammatory conditions[2]. The stroma is therefore said to be reactive. Reactive stroma has been shown to facilitate tumor growth and progression. The extensive stroma reaction that is found in many types of carcinomas is referred to as desmoplasia. Even though many tumors are immunogenic, the inflammatory environment in the desmoplastic stroma rather supports tumor growth instead of inhibiting it. This occurs through a diversion of the immune-system, which is driven by a deregulated signaling that occurs between the inhabitants of the tumor micro-environment[3]. The deregulated signaling that help tumors evade directed immune responses also conveys other altered features of the tumor tissue. Like almost all other tissues, tumors need a vascular supply[4]. Blood vessels are formed through angiogenesis, but due to the deregulated signaling in the desmoplastic stroma these have an impaired function and do not provide enough oxygen and nutrients[5]. This will lead to hypoxia, which elicits an even stronger demand for angiogenesis, and hypoxia may also aggravate the malignancy of the tumor cells. Also, the ECM architecture is altered and dis-plays a phenotype that is reminiscent of fibrosis. This together with the dys-functional vasculature have profound effects on fluid balance and penetration of solutes into the tumor tissue[6]. The latter constitutes a problem when try-ing to treat a solid tumor with chemotherapeutic agents[7]. The outcome of a malignant disease will depend on tumor growth rate, metastases formation, and the efficacy of treatment. These features are dependent on all constituents of the tumor microenvironment.

The ECM of loose connective tissue Fibrillar components The backbone of the loose connective tissue is the fibrillar components, which are the major contributors to mechanical structure. There are several types of fibers and fibrils that span the ECM of loose connective tissue and they all have different characteristics and functions, which will be presented below.

The collagens The collagens are a family of proteins that is primarily distinguished by their triple helical structure, which is formed by three polypeptide chains. A pre-

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requisite for the triple helix confirmation is that every third amino acid (aa) must be a Glycine (Gly), thus the collagenous domain has the repeated se-quence X-Y-Gly (where X and Y can be any aa, but one of them is often a proline (Pro)). To date, there are 28 known collagens in mammals, which are made up from polypeptide chains encoded by at least 42 genes[8, 9]. Not all collagens can form fibrils, those who can is called fibrillar collagens and these are collagen types I, II, III, V, XI, XXIV and XXVII. In loose connec-tive tissue collagen type I is the predominant form, type II is found in carti-lage where it is predominant, and type III is found in several tissues, includ-ing loose connective tissue. Collagen type V and XI are minor constituents of collagen I and II fibrils respectively. The type XXIV and XXVII colla-gens are rather novel and not much is currently known about their fibril structure, distribution and function[10]. There are other types of collagens besides the ones that form fibrils:

Network forming collagens (IV, VI, VIII, X).

Fibril associated collagens with interrupted triple helix (FACIT, comprising collagen type IX, XII, XIV, XVI, XIX, XX, XXI and XXII).

Membrane associated collagen with interrupted triple helices (MACIT, com-prising collagen type XIII, XVII, XXIII and XXV).

Multiple triple helix and interruptions (MULTIPLEXIN, comprising colla-gen type XV and XVIII).

Anchoring fibrils forming collagen, comprising only collagen VII [10].

Collagen type I synthesis The collagen type I triple helix contains three polypeptides encoded by two separate genes; COL1A1, which encodes the pro-α1 chain and COL1A2, which encodes the pro-α2 chain. The C- and N-terminal pro-peptides are cleaved off extra-cellularly, leaving short non-collagenous endings, referred to as telopeptides[11]. In the endoplasmic reticulum (ER), Pro residues of pro-collagen can be hydroxylated in the 4 position of the ring structure by the enzyme Prolyl-4-hydroxylase (P4H), in a reaction requiring oxygen (O2), iron ions (Fe2+) and ascorbate (Vitamin C). The hydroxylation of Pro resi-dues is important for the thermal stability of the triple helix, which is only stable up to 24° C when all Pro residues are un-hydroxylated. Typically, around 50 % of the Pro residues in collagen type I are hydroxylated, which results in a triple helix that can withstand body temperature[12]. Moreover, Pro residues can also be hydroxylated in the 3 position by the enzyme 3-prolyl-hydroxylase (3PH), this has only been reported to occur on one Pro residue on collagen I[13, 14]. Lysine (Lys) residues can also be hydroxylated

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by the three enzymes Lysyl hydroxylase 1, 2 and 3 (LH1-3, a.k.a. PLOD1-3, after the gene name Pro-collagen-Lysine, 2-Oxoglutarate 5-Dioxygenase). Further, the hydroxylated Lys residues can be glycosylated and acquire a galactose mono-saccharide or a galactose-glucose di-saccharide[12]. Two pro-α1 and one pro-α2 chain are assembled into a triple helix, which is ex-ported out of the cell[15]. The pro-domains are cleaved off by proteases extracellularly. Bone morphogenic protein 1 (BMP-1) is responsible for cleaving the C-terminal pro-peptide[16] and a disintegrin and metallopro-teinase with thrombospondin motifs 2 (ADAMTS-2) is responsible for cleaving the N-terminal pro-peptide[17, 18]. The reason for the extra-cellular cleavage is that cleaved collagen molecules have the propensity to self as-semble, and this would have detrimental effects if it would occur inside the ER. However, there are reports of fibril assembly in tendons that involves cellular structures referred to as “fibropositors”, which are cell protrusions that deposit small fibrils in the ECM where further growth and maturation of the collagen fibril takes place[19].

Collagen type I, fibrils and fibers Collagen type I is one of the most abundant proteins in loose connective tissue and it forms large supramolecular structures together with other colla-gens and proteoglycans. Collagen V has been proposed to have a role as a nucleator for collagen I fibril assembly. Collagen V is also a fibrillar colla-gen, comprised of one α1(V) chain and two α2(V) chains in most tis-sues[20]. Fibril segments containing collagen I/V fibril fuse with other seg-ments and polymerize into larger fibrils. Fibrils grow both longitudinally and laterally. The lateral growth seems to be regulated by small leucine rich re-peat proteoglycans (SLRPs) and FACIT collagens, which will be further discussed below. Collagen III can also be incorporated into collagen I fibrils and this partly regulates the fibril thickness [21, 22]. The collagen molecules are arranged in a quarter stagger formation in the mature fibril, which gives rise to the characteristic D-band observed in transmission electron images. It is the gap between collagen molecules that give rise to this pattern and the length varies between 64-67 nm depending on the tissue and species[23]. A collagen I molecule is 4.4 D-bands in length, ~300 nm. The thickness of collagen fibrils depends on the needs of the tissue, for example in mice col-lagen fibrils can range from 10 nm in cornea to 150 nm in tendon [24]. Bun-dles containing more than hundred fibrils can assemble in to a collagen fiber, and the fiber is the unit which provide the tissue with its actual strength[25].

Another extracellular event is the cross linking of collagens, which occurs both within and between collagen molecules. Crosslinking between collagen molecules of different type also occur[26]. Lysyl oxidases are responsible for catalyzing the initiation of these events and there are five different en-zymes; lysyl oxidase (LOX) and the four lysyl oxidase like (LOXL) 1-4[27].

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The most studied lysyl oxidase is LOX, which is a pro-enzyme that is cleaved by BMP-1 and mammalian tolloid family of metalloproteinases[28]. These are the same proteases that also cleave the C-terminal of collagen type I and III[16, 29]. Active LOX catalyzes the oxidation of Lys and hydroxy-Lys residues in the helical and telo-peptide regions in collagen. The resulting aldehyde, hydroxal-Lys spontaneously reacts further with other Lys and hydroxy-Lys residues forming intermediate divalent crosslinks. Reactions with a third hydroxy-Lys leads to a mature trivalent crosslink. The role of covalent crosslinks is to stabilize collagen fibrils conferring increased strength. Qualitative and quantitative changes in the crosslinking pattern occur during tissue remodeling, where the amount of immature crosslinks increases during remodeling and then decreases with tissue maturation[30]. The crosslinking pattern is also affected in pathological conditions, e.g. fi-brosis[31, 32] and cancer[33], as will be discussed in more detail below. Crosslinks can also form without the help of enzymes. Sugars[34] and de-rivatives of lipid metabolism[35] can also form crosslinks. These unwanted crosslinks impair the function of the collagen matrix and increase with the age of the collagen molecule. Sugar-derived crosslinks are especially preva-lent in diabetic patients due to elevated glucose levels[30].

Reticular fibers Collagen type III is the major constituent of reticular fibers[36], which are much thinner than regular collagen fibers that mostly consist of collagen I. Collagen III is a homotrimeric collagen encoded by the single gene COL3A1, which encodes the α1(III) polypeptide. The biosynthesis of colla-gen III is essentially similar to that of collagen I, with some exceptions, e.g. additional crosslinking by formation of cystein bridges[37]. The reticular fibers consists of small bundles or single collagen III fibrils that are distinct from but continuous with collagen I fibers. Apart from collagen type III, reticular fibers also comprise fibronectin[38] and collagen V, as well as various proteoglycans[39]. Reticular fibers form intricate networks, which are most abundant just below the BMs of blood vessels and epithelia in loose connective tissue [40]. At this location reticular fibers interact with collagen type VII that forms anchoring fibrils. The N-terminal of the anchoring fibrils interact with BM components, which provides a link between the loose con-nective tissue and the BM. Because reticular fibers are thinner, they are more flexible than the thicker, more rigid collagen fibers. This suggests that reticu-lar fibers could act as a cushion or buffer when tensional forces are imposed on one of two adjacent compartments.

Elastic fibers and microfibrils Elastic fibers are made up from two distinct major components; elastin fi-brils and microfibrils. The morphology of elastic fibers differs between dif-ferent tissues and both fibrillar and lamellar forms exist. In loose connective

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tissue there are only fibrillar elastic fibers present. They appear rough due to their close association with the microfibrils, which entangle the elastic fibril core and project out from the fiber[40]. Microfibrils seems to be the template structure, which the elastic fiber forms around. The role of the elastic fibers is to provide the tissue with an elastic recoil property, enabling stretching of tissues. Microfibrils are composed of fibrilins, and other quantitatively minor components such as microfibril associated glycoproteins (MAGPs), Fibulins and EMILIN-1. Fibrilin-1 and -2 exists in rodents, whereas a third member, fibrilin-3 is presents in humans. Both fibrillin-1 and -2 have affinity for elastin[41]. Except for the close association with elastin in elastic fibers, microfibrils are also important in sequestering of the transforming growth factor β (TGF-β) Large Latent Complex (LLC)[42] in the ECM. TGF-β is one of the single most important growth factors when it comes to inducing expression of ECM molecules (see section below about TGF-β). Fibrilins contain an Arginine-Glycine-Aspartic acid (RGD) sequence, which enables binding between microfibrils and cellular ECM receptors, i.e. integrins[43]. The elastin fibril is composed of the monomer tropo-elastin encoded by a single gene (ELN). Extracellularly, tropo-elastin associates with fibulins and aggregates are formed. The exact role of the associated fibulins is uncertain but deficiency in some fibulins leads to altered microfibril and elastic fiber function and morphology[41]. Fibulin-4 seems to be involved in the regula-tion of the activity of LOX and/or LOXL1 on tropo-elastin[44]. Tropo-elastin has around 40 Lys residues and LOX and/or LOXL1 modify the ma-jority of these, which subsequently form crosslinks. These crosslinks are of another type compared to the crosslinks in collagen, and these crosslinks are also partly responsible for the insolubility of elastic fibers[41]. Aggregates of tropo-elastin, fibulins and possibly other proteins are deposited by cells onto microfibrils, which serves as template for the nascent elastic fiber[45].

Small Leucine-rich Repeat Proteoglycans (SLRPs) As the name suggests, this is a family of ECM proteins that share a charac-teristic C-terminal Leucine (Leu) -rich repeat (LRR), which is responsible for binding with fibrillar collagen. The N-terminal is highly variable and is often subjected to extensive post-translational modifications. The SLRP family is further divided into four classes, of which class I and II are the most studied. It has been shown from studies with knock-out (KO) mice deficient in one or more SLRPs that these proteins have important roles in the regulation of collagen fibril structure and assembly. In the absence of some SLRPs, the collagen fibrils that normally are uniform in diameter, be-come abnormally fused resulting in large irregular fibrils, and an increase in smaller fibrils can be observed in some models. This leads to a heterogene-ous distribution of collagen fibrils and an impairment of tissue function. Some SLRPs can also interact with elastic fibers through binding of tropo-elastin[46] and with FACIT collagens[47, 48]. Another interesting feature of

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some SLRPs is the ability to bind and sequester TGF-β, Thus rendering TGF-β biologically inactive[49, 50]. Additionally, a subset of SLRPs can also act as ligands for receptor tyrosine kinases. For example, decorin binds epidermal growth factor receptor (EGFR) without activating it, and subse-quently induces its internalization and degradation[51]. On the other hand decorin can activate the insulin like growth factor I receptor (IGF-IR)[52]. Biglycan binds and activates Toll like receptor 2 and 4 (TLR-2 & -4)[53] that are important regulators of innate immunity responses. Typical of the latter interactions is that the SLRPs exert their actions at much higher molar ratios than the cognate ligands for these receptors. The SLRPs that are most relevant for loose connective tissue in developmental, normal adult and pathological conditions are presented below.

Class I SLRPs This class consists of asporin, decorin and biglycan. In contrast to biglycan and decorin, the variable N-terminal of asporin contains no GAG chain. In-stead, asporin has a calcium binding poly-Aspartate tail[54]. Asporin is be-lieved to have a role in the calcification of tissues, and calcification of loose connective tissue can be observed during pathological conditions[55]. Asporin and decorin can compete for the same collagen binding site[54], whereas biglycan’s binding to collagen is debated[56]. Decorin deficient mice have revealed that this SLRP is important for the diameter and size distribution of collagen fibrils in connective tissue (dermis). Collagen fibrils from decorin KO mice displayed a broader range in size and there was also an increase in the observed maximal fibril diameter. This indicates that decorin is involved in the regulation of collagen fibril diameter and that its function is unique in dermis, which is indicated by the lack of compensatory rescue from other SLRPs. The disorganized collagen matrix in decorin KO mice leads to lax and fragile skin[57]. The biglycan KO mouse display ab-normal dermal collagen fibrils, with enlarged mean fibril diameter and a broader fibril size range, similar to the decorin KO mouse. However, the abnormal collagen structures in biglycan deficient mice do not lead to any clinical skin phenotype, in contrast to the decorin KO mouse. Instead there is a severe phenotype in the bones of biglycan KO mice[58], which is aug-mented by decorin deficiency[59]. Biglycan and decorin are often expressed in the same tissues during development, but with a temporal delay where expression of biglycan precedes the expression of decorin[60]. This is also observed during inflammatory conditions where biglycan is expressed early and decorin late[61, 62]. However, Keloids which are fibrotic skin lesions have a high expression of biglycan[63], suggesting that biglycan could be a marker of active inflammation. Decorin on the other hand is down-regulated in the fibrotic skin condition Scleroderma[64]. There have been no reports of an asporin deficient mouse model so far.

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Class II SLRPs This class is comprised of fibromodulin, lumican, keratocan, osteoadherin, and PRELP (Proline and Arginine-rich End Leucine-rich repeat Protein). However, only fibromodulin and lumican will be discussed further, because of their roles in loose connective tissue during homeostasis and pathology. Both fibromodulin and lumican have several sulfated (-SO4) tyrosines (Tyr) in their variable N-terminals. Fibromodulin have two collagen binding sites[65], in contrast, to lumican which only possess one[66]. The fibro-modulin KO mice display thinner and abnormal collagen fibrils in ten-dons[67], which seems to be the primary location for fibromodulin expres-sion. Fibromodulin is also expressed in loose connective tissue during reactive conditions[68, 69]. Lumican deficiency in mice leads to increased collagen fibril diameters, which also results in abnormal collagen structures in skin, cornea and tendons. The most severe phenotype is an opaque cornea in the lumican KO mouse and this leads to decreased transparency and sight impairment[24]. Fibromodulin and lumican double KO mice have severe joint problems and display abnormal collagen fibrils. There is an increase in the fraction of fibrils that are large and very large. However, there is no skin phenotype, additional to that seen in lumican KO mice reported in the fibro-modulin and lumican double KO mice[70]. When expressed in the same tissue, fibromodulin and lumican also show a temporal difference similar to biglycan and decorin. Lumican expression peaks before fibromodulin during tendon development and fibromodulin is expressed in the mature tendon[71], alongside decorin. The temporal difference of expression and the difference in collagen affinity, as well the number of binding sites for collagen that differ between fibromodulin and lumican indicates that these two SLRPs have different roles during collagen fibrillogenesis. A theory that has been proposed for tendon development is that lumican could hinder lateral fusion of collagen fibrils through its single binding site on collagen. This would favor the formation of more, but smaller fibrils that probably are more im-portant early in tendon development. Fibromodulin expression coincides with increased fibril diameter, which could imply that fibromodulin with its higher affinity for collagen could compete with lumican for binding and eventually displace lumican from collagen. Fibromodulin could due to its ability to bind discrete sites on collagen, bind two different collagen triple helices and thereby favor lateral fusion[56].

Fibronectin Fibronectin is a large glycoprotein present in plasma in soluble form and as fibers in connective tissue. Fibronectin is encoded by a single gene (FN), but through alternative splicing several isoforms exist. The fibronectin polypep-tide chains contain many different domains or modules, which enables inter-actions with a broad range of other ECM molecules. Notably, native and to a

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larger extent denatured collagen (gelatin); fibrin and fibrinogen; GAGs; and the matricellular protein thrombospondin (TSP)[72]. The amount of alterna-tively spliced fibronectin variants is increased during embryogenesis, but decreases in adulthood, except in conditions of tissue remodeling[72]. This suggests that the alternative spliced variants of fibronectin could be impor-tant for the assembly of the ECM, and it has been shown that alternatively spliced fibronectin containing the type III module extra domain A and B (EDA & EDB) are more readily incorporated into matrices[73]. Fibronectin is a homo-dimeric molecule, which is linked by cystein bridges. Although fibronectin consists of two chains encoded by the same gene, the two chains do not have to be of the same splice variant, which is the case with plasma fibronectin. Plasma fibronectin is produced by hepatocytes in the liver, and the concentration in plasma is 0.33 mg/ml[74]. The other major form of fi-bronectin is cellular fibronectin, which is mainly produced by fibroblasts in loose connective tissue[75]. However, other cells present in the loose con-nective tissue have also been reported to express and secrete fibronectin in vitro, e.g. macrophages[76] and endothelial cells[77]. As several other ECM molecules, fibronectin contains an RGD motif, which enables binding to certain integrins. Through interaction with cell surface integrins, fibroblasts can assemble fibronectin molecules into a fibrillar matrix. These elaborate fibronectin networks are observed both in vitro[78] and in vivo[79]. The fibronectin matrices has been reported to be part of the reticular fiber sys-tem[38] and of the fibrillar collagens fibronectin has the highest affinity for collagen III, which is a major constituent of reticular fibers[80]. Also, both collagen I and collagen III fibers are deposited on to a preformed fibronectin matrix in vitro[78]. Fibronectin fibrils has also been proposed to have elastic features, but how and to what extent fibronectin fibrils have this ability is unknown[81].

Non-fibrillar components of the ECM

The ground substance The ground substance constitutes the filling material that resides within the compartments, which are formed by the fibrillar structures in loose connec-tive tissue. However, the ground substance is far from any inert filling, it has crucial roles in regulating tissue function. The ground substance consists of proteoglycans with covalently attached GAG chains of variable length that consists of repeating disaccharides.

The composition of the disaccharide repeat is characteristic for each class of GAG. The major classes are: heparan-sulfate and heparin; chondroitin sul-fate; dermatan sulfate; keratin sulfate; and hyaluronan. Most GAGs are found coupled to a protein core, and they are therefore referred to as pro-

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teoglycans[82]. Hyaluronan which is not covalently linked to any core pro-tein, is the most abundant GAG in loose connective tissue[83]. Hyaluronan consists of one very long GAG chain that can have a molecular weight of several million Daltons. The sugars that build up GAG chains are negatively charged at physiological pH and this contributes to the tertiary structure of GAG chains. Long GAG chains like hyaluronan assume a random coil shape, which make them occupy much more space than the molecular con-stituents actually warrants. However, several hyaluronan molecules can en-tangle with each other, which means that several molecules can share the same space. The negatively charged GAG chain attracts positive counter ions, which will exert an osmotic effect that in turn attracts water[84]. If such GAGs are placed in a water solution, they will form a hydrated gel. In loose connective tissue, the GAGs are under-hydrated due to the tight regu-lation of fluid homeostasis that the loose connective tissue as an organ ex-ert[83], which will be discussed below. However, the GAGs are still in a gel-like state in loose connective tissue, which provide the tissue with compres-sion resistance. When the tissue is compressed, water is pressed out of the gel, whereas upon cessation of compression, water moves back and the tis-sue regains its shape.

Matricellular proteins The matricellular proteins are a heterogeneous class of ECM proteins com-prised of diverse members with some features in common. The matricellular proteins are signified by their location in the ECM, and by their ability to interact with cells and structural components of the ECM, as well as with extracellular enzymes and signaling molecules. The matricellular proteins also have their tempo-spatial expression pattern in common. They are ex-pressed during embryogenesis and in conditions of active tissue remodeling, e.g. wound healing, inflammation and in tumors. Matricellular proteins like the CCNs (abbreviation derived from the three most studied members of the CCN family: Cyr61 a.k.a. CCN1, Connective tissue growth factor a.k.a. CCN2, and Nov a.k.a. CCN3); thrombospondins (TSP) 1 and 2; tenascin-C (TNC); SPARC (Secreted Protein, Acidic and Rich in Cystein); periostin; and osteopontin all have the ability to bind different integrins and other types of cell surface proteins that normally interact with ECM ligands. Most ma-tricellular proteins have a RGD motif that enables binding with αV-integrins. However, this interaction can lead to an altered adhesion, compared to in-tegrin-ligand interactions in the absence of matricellular proteins. Most ma-tricellular proteins like CCNs[85], periostin[86], SPARC, TNC, and TSP-1 & -2[87] have been proposed to induce an intermediate type of adhesion, which could render cells more motile[88]. This would reflect the needs of a remodeling tissue, where firm attachments could constitute a hindrance in the remodeling process. However, some matricellular proteins can also in-duce mature adhesions, with actin stress-fiber formation and focal adhesion

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kinase (FAK) phosphorylation in vitro, which have been reported for osteo-pontin[89, 90] and periostin[91].

In conjunction with the modulation of adhesion, some matricellular proteins are also capable of modulating signaling of certain growth factors and cyto-kines. For instance, CNN2 enhances the effect of EGF and IGF[92]; osteo-pontin potentiates the migratory effect of EGF in epithelial cells[93]; and SPARC can inhibit binding of platelet derived growth factor (PDGF) to its receptor[94]. Importantly, many types of matricellular proteins can bind and regulate the activation and availability of TGF-β[95, 96]. This is especially prominent with TSP-1, which is a direct activator of TGF-β[97].

Additionally, the activity of extracellular proteases can be modulated by matricellular proteins. Several matricellular proteins can also bind structural components of the ECM, such as collagen and fibronectin, and act as linkers between different ECM structures, as well as between ECM and other pro-teins. Even if many matricellular proteins associate with collagen, they are not considered to be a integral part of the fibril. However, like KOs of SLRPs, mice deficient in some matricellular proteins display altered colla-gen fibrillogenesis. Periostin deficient mice are born with reduced collagen fibril diameters in tendons and have a decreased amount of collagen cros-slinks, which affects the biomechanical properties of connective tissues[98]. Periostin KO mice also show decreased survival in a model of acute myo-cardial infarction and this was also attributed to an insufficient collagen ma-trix, which could not support the mechanical stress inherent to the heart[91]. Osteopontin deficient mice display reduced collagen fibril diameters and a disorganized dermal ECM after wound healing[99]. Mice lacking functional SPARC display smaller collagen fibrils and decreased tensile strength of skin[100]. In contrast, TSP-2 null mice display enlarged collagen fibrils with irregular shapes. Also, the gross appearance of the collagen fiber architecture in dermis of TSP-2 KO mice is altered, with irregular, non-parallel collagen fibers[101].

Cells of the loose connective tissue

The fibroblast Fibroblasts are cells of mesenchymal origin, residing in connective tissue. These cells are dispersed in the tissue and appear solitary, but they have di-rect contacts with other cells through long cellular processes ending with gap junctions[102, 103]. Fibroblasts are the main regulators and effectors of ECM turnover, since they are capable of producing and degrading most ECM components found in loose connective tissue. In resting tissues, turn-

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over occur at a low but steady pace that keeps the tissue in balance. If this homeostasis is deregulated, with too much or too little of either synthesis or degradation, it will lead to a dysfunctional tissue as will be discussed below. Further, fibroblasts do not just produce and degrade ECM molecules, they also have the ability to organize the ECM into a ordered structure, which is important for proper tissue function[104, 105]. Moreover, fibroblasts partici-pate in the regulation of other homeostatic processes, such as the fluid bal-ance in connective tissues.

Tissue macrophages Macrophages are myeloid cells derived from the bone marrow. These cells enter tissues from the circulation as monocytes, but differentiate into macro-phages when in the tissue. The name, macrophage means “eat big”, which indeed is a suitable name for these phagocytotic cells. The macrophages are part of the innate immune system and constitute the first line of defense against invasion by foreign organisms, tissue injury and transformed cells. As mentioned earlier, the loose connective tissue constitutes a buffer zone between different body compartments, and any breach in an adjacent com-partment will alarm the immune competent cells in the loose connective tissue.

Macrophages express a broad range of cell surface receptors that allow them to sense if there is any tissue damage or presence of invading microorgan-isms. The pattern recognition receptors (PRRs) are comprised of several families of receptors that recognize pathogen-associated molecular patterns (PAMPs), as well as some endogenous ligands. PAMPs are a collective term for microbial derived molecules, such as peptides, lipids, and nucleic acid. Biglycan, versican, fragments of hyaluronan, and oxidized low density lipo-protein constitute examples of endogenous ligands. The most studied PRR family is the toll like receptors (TLR), which are further divided into two groups that differ in their sub-cellular location and type of ligands. TLR 1, 2, and 4-6 are expressed on the cell surface and these primarily recognize shed membrane components of microbes. In contrast, TLR 3 and 7-9 are located inside intracellular vesicles, where they are exposed to degradation products from phagocytosed microbes. Their primary ligand is microbial DNA or RNA. Another PRR is the RIG-I like receptors (RLRs), which are RNA he-licases that respond to viral RNA[106]. The Nod like receptors (NLR) also belong to the PRRs and they detect PAMPs and endogenous ligands inside the cytosol[107]. Stimulation of PRR induces macrophage activation, which leads to expression of bio-active polypeptides, e.g. interferons (IFN), growth factors, and cytokines, as well as lipid derivatives, such as prostagland-ins[108]. The function of these released factors is to induce an inflammatory response. Macrophages themselves, express receptors for inflammatory me-diators, and macrophages can adopt different phenotypes resulting from in-

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teractions with different kinds of T lymphocytes (T-cells). Stimulation by TH1 cells induces classical activation of macrophages, also called the M1 phenotype, Stimulation by TH2 cells induces an alternative activation of ma-crophages that is also called the M2 phenotype. The M1 phenotype is in-duced by PRR signaling, or activation by IFNγ and tumor necrosis factor alpha (TNFα). The M1 macrophages are pro-inflammatory and secrete fac-tors that augment the inflammation, e.g. TNFα, IL-1 and IL-6. The M2 phe-notype can be divided into subgroups, but they are all considered to be less pro-inflammatory. Instead of secreting large amounts of pro-inflammatory cytokines, they release high levels of anti-inflammatory factors, such as IL-10. Macrophages can acquire the M2 phenotype when exposed to IL-4 or IL-13[109].

Apart from detecting invasion and tissue damage, macrophages can also perform clearance of degraded tissue and foreign invaders through phagocy-tosis. Macrophages use scavenger receptors in order to detect damaged cells and ECM fragments, which are subsequently phagocytosed/endocytosed and degraded in lysosomes[110, 111]. Macrophages express complement recep-tors and IgG receptors (FcγRI-III), which both mediate phagocytosis of whole microorganisms through opsonization. The opsonins IgG or comple-ment factors bind to the surface of microbes. IgG and complement then bind their receptors on macrophages, which phagocytose the microbe. Well inside the macrophage, the phagosomes fuses with lysosomes, which contain de-grading enzymes and reactive oxygen species that destroys the mi-crobe[112].

Macrophages are also antigen-presenting cells (APC) and short peptides are displayed on the macrophage cell surface through the major histocompability complex class II (MHC II). In this way foreign or aberrant endogenous pep-tides become presented to T-cells of the adaptive immune system, which can further regulate the immune response[113]. Macrophages also have a role in the remodeling of ECM during an inflammation, since they both secrete ECM molecules and proteases that can degrade them[76, 114].

Mast cells Like the macrophages, mast cells are derived from the hematopoesis and mature in peripheral tissue. Mast cells are part of the innate immune system and are primarily located in loose connective tissue, often in close proximity to blood vessels and they constitute sentinels that can respond immediately to foreign invaders. Mast cells are particularly well equipped to counter parasitic infections. The cytoplasm of mast cells is filled with granules that contain bio-active compounds, e.g. histamine and proteases. Upon stimula-tion, the content of these granules can be released immediately, causing an instant but local inflammatory response. The inflammatory stimulation is

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also augmented by the production of bio-active lipid derivatives, cytokines and growth factors. Classical mast cell activation is achieved through liga-tion of IgE-antigen complex with the IgE receptor (FcεRI) on mast cells, which leads to subsequent degranulation[115]. However, mast cells like macrophages also express PRRs, complement and IgG receptors. Activation of these receptors can also stimulate mast cells, but do not necessarily lead to degranulation[116].

Dendritic cells Dendritic cells originate from the same myeloid lineage as macrophages and like the macrophages, they arrive from the circulation and becomes resident in a tissue. The dendritic cells have many features in common with macro-phages, however their role as APCs predominates. In contrast to macro-phages, dendritic cells can re-enter the circulation upon stimulation and travel to lymphoid tissues, where they can present antigens to a wider popu-lation of T-cells[117]. Dendritic cells express a broad range of cell surface receptors, similar to macrophages and dendritic cells also become activated by the same type of stimuli as macrophages. In order to obtain material for antigen presentation, dendritic cells also need to phagocytose microbes. However, the purpose of phagocytosis in dendritic cells is not primarily to clear the tissue, rather the aim is to acquire peptides for display through MHCII[118].

Cell-ECM interactions Integrins Cells need receptors for ECM components in order to interact with their environment. Most cells perform their differentiated function when attached to an ECM substratum, and normal cells are obligated to be adherent in order to survive. If normal cells are deprived of ECM contacts, they undergo apop-tosis, a phenomenon known as anoikis (Greek word meaning homelessness). However, many transformed cells independent of lineage, can cope with anoikis and grow without adherence[119]. Cells express a large number of adhesion receptors that can mediate contacts with the ECM. Integrins consti-tute an extensively studied family of such receptors. An integrin receptor is a heterodimer, comprised of one α chain and one β chain, which are encoded by different genes. There are 18 α chains and 8 β chains, which can be com-bined in a number of ways, resulting in 24 known heterodimers[120]. Most integrin heterodimers can be subdivided into three major clusters, based on their composition:

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β1-integrins: Large group, comprising 12 heterodimers, which can bind sev-eral different matrix molecules, e.g. collagen, fibronectin and laminin.

β2-integrins: Comprises four heterodimers, exclusively expressed on leuko-cytes and predominantly bind proteins on other cells, e.g. endothelium. In addition β2-integrins also binds complement factors and this is important for phagocytosis.

αV-integrins (where V stands for vitronectin): This integrin cluster comprises five heterodimers with a broad ligand specificity. All αV-integrins bind the RGD sequence, which is present in many ECM proteins, including fi-bronectin, fibrin, fibrillin, vitronectin, and many matricellular proteins[121]. The expression of αV-integrins are primarily up-regulated in conditions of tissue remodeling, which are conditions in which a wider range of substrates are available. The importance of αV-integrins in tissue remodeling is further substantiated by the ability of some αV-integrins to activate latent TGF-β[122].

Integrin signaling Several integrin heterodimers cluster together at the cell membrane upon ligation of ligands. The clustering of integrins at the cell surface attracts other molecules that link integrins to the cytoskeleton, but also signaling molecules are recruited to these structures, which are referred to as focal adhesions. Integrins lack intrinsic kinase activity, but several kinases are among the molecules that are recruited to focal adhesions. Also growth fac-tor receptors are attracted to focal adhesions as well as their downstream substrates and interaction partners[123]. This implies that focal adhesions can serve as foci for signaling, since many types of signaling pathways con-verge in these structures. This opens up for crosstalk between different types of signaling that may modulate the signaling pathways. Moreover, integrin clustering can induce growth factor receptor phosphorylation in the absence of its cognate ligand[124, 125]. The extent of phosphorylation is further increased by addition of the cognate ligand. The interaction between in-tegrins and receptor phosphorylation is further backed by evidence that show a specific association between integrin αVβ3 and the hyperphosphorylated pool of PDGFRβ[126]. Evidence also put integrins together with growth factor receptors, such as PDGFRβ, in another signaling hot-spot, the caveo-lae[123]. The current and emerging evidence suggests that there is a substan-tial crosstalk between integrin and growth factor signaling, which emphasize that neither of these systems should be studied as separate entities.

Integrins do not only mediate static adherence, they also facilitates migration on a substratum consisting of ligands for the specific integrin heterodimer at interest. The focal adhesions are dynamic structures and focal adhesion ki-

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nase (FAK) which is responsible for initiating many signal transduction events in focal adhesions are also involved in the resolution of focal adhe-sions[127]. The dynamic nature of focal adhesions is a prerequisite for mi-gration, since static adhesions would not permit motility.

Blood and lymph vessels in loose connective tissue The microvasculature Almost all exchange between blood and tissue occurs in loose connective tissue, which embed the microvasculature. The capillaries are the smallest vessels with diameters ranging from 4-12 µm, and the endothelial lining in these vessels are permeable enough to allow exchange of nutrients and waste products take place. The capillaries are continuous with both arterioles that lead oxygenated blood from the arterial side into the tissue, and with venules that leads out of the tissue. This set of vessels comprises the micro vascular unit. There are some differences in vascular morphology between the differ-ent vessels in the microvascular unit, which follow their functional differ-ences. The role of the arteriole is to lead blood into the capillary bed, and therefore arterioles are rather impermeable and covered by a continuous coat of vascular smooth muscle cells (vSMCs) that can maintain blood pressure. The transition from arteriole to the capillary morphology is gradual, and it is emphasized by a gradual loss of vSMCs and reduced luminal diameter. In-stead of vSMCs, the capillary is coated by another type of mural cell, the pericyte. The pericyte coat is less dense compared to the vSMC coat of arte-rioles. The function of capillaries is to allow proper exchange of nutrients over the endothelium and therefore the capillaries are more permeable to molecules. The capillaries then transitions into post-capillary venules, and this is emphasized by an increase in lumen diameter and a denser coat of mural cells. However, the mural cell coat in venules is not as dense as in arterioles, and the type of mural cells are more reminiscent of pericytes than vSMCs[128]. The role of the venules is to transport blood from the capillar-ies into the collecting veins, but venules also have a role in inflammation, where the venules, like capillaries, become dilated and hyperperme-able[129].

The endothelium The endothelium is the continuous luminal lining of blood vessels, and it is comprised of endothelial cells. The role of the endothelium is to constitute a tight barrier that controls the passage of molecules from the blood into the tissue. In addition, the endothelium also shields the clotting factors of the blood from the procoagulant factors that reside on the abluminal side. The endothelial luminal surface is also anti-coagulant during normal homeosta-

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sis, and in addition endothelial cells secrete anti-coagulant factors. Endothe-lial cells form tight junctions between each other, which limits the passage between endothelial cells. However, these junctions are not as tight in the capillaries as in larger vessels, which will allow limited passage of solutes. Endothelial cells of capillaries can also allow passage of molecules through fenestrations in the endothelial plasma membrane, and through vesicular organelles[130].

The vascular basement membrane Endothelial cells are firmly anchored to the vascular basement membrane (BM), which provide structure and support for the vessel, in addition to its role as a physical barrier. The major constituents of BM are collagen IV, laminins, nidogen, perlecan and fibulins. Collagen IV is a network forming collagen, which provide a scaffold for the BM. Collagen IV is made up from six different genes, COL4A1-6. However, only three triple helices are known to exist in vivo. In contrast to the fibrillar collagen, collagen IV have numerous interruptions in its triple helix, which provides flexibility to the molecule and enables collagen IV to adopt a network suprastructure[131]. There are conflicting reports about the nature of the crosslinks in collagen IV, but disulphide bonds are present[132]. Collagen XV and XVIII are also present in the BM, but to a minor extent compared to collagen IV. Collagen XV and XVIII are interesting since proteolytic cleavage of these generates fragments with anti-angiogenic properties[133, 134]. Anti-angiogenic frag-ments derived from collagen IV has also been described[135].

Laminins are heterotrimeric proteins consisting of an α, β and γ chain that form a coiled-coil structured stalk that diverge into a cross-shaped structure. There are five different α-chains, three different β-chains, and three different γ-chains, which can combine into at least 17 hetero-trimers. The laminins can polymerize into network structures in BM, which exist in parallel with the collagen IV network and this further contributes to the structure of BMs[136]. Both endothelial and mural cells express integrins that mediate adhesion to both collagen IV and laminins[137]. The role of nidogen and the heparan sulfate proteoglycan perlecan is to link collagen IV and laminin polymers together, which increases the stability of the BM. Fibulins link laminins, perlcan and nidogen together[138].

The pericyte Pericytes are situated on the abluminal side of the endothelium and are con-tinuous with the BM. Pericytes have their location and some features in common with vSMC, such as the expression of certain markers (desmin and α-SMA (alpha smooth muscle actin)). However, pericytes and vSMCs differ in the type of vessels to which they associate; pericytes are located in capil-laries and even more numerous in post-capillary venules, vSMCs on the

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other hand are located in larger vessels. The morphology are also different, vSMCs wrap around the endothelial tube over a rather short distance, whereas pericytes extend cellular processes that enwrap the endothelium over a longer distance of the vessel, enabling one pericyte to be in contact with several endothelial cells[139]. The transition between arterioles and capillaries, and capillaries and venules are continuous, and so are also the transitions from vSMCs to pericytes and vice versa. In addition, cells with an intermediate phenotype exists in the transitions zones[140].

Apart from the expression of the smooth muscle proteins, desmin and α-SMA, pericytes also express other proteins involved in smooth muscle cell contraction, e.g. myosin and tropomyosin[141]. Indeed, pericytes have been shown to regulate vascular tone and to respond to neurotransmitters[142]. However, the expression of contractile proteins is lower in pericytes com-pared to vSMC[141, 143], which probably reflects the reduced need of vas-cular tone regulation in the types of vessels that are covered with pericytes.

The main function of pericytes seems to be providing structural support to the endothelium, where pericytes are the cellular and dynamic component and the BM provides resilience. The pericyte coverage of microvessels is heterogeneous throughout the body, with the highest density in the CNS, and particularly in the retina[144]. In support for a stabilizing role of pericytes in microvessels is the finding that retinal microvessels are more resistant to the damaging effects of supraphysiological blood pressures (>250 mmHg in rats) compared to microvessels in other parts of the CNS[145]. Pericytes do not only provide physical support, but they also provide molecular cues that are crucial for vessel maintenance and maturation of nascent vessels[146, 147]. This is supported by the vulnerability of vessels that lack or have di-minished pericyte coverage[148, 149]. Pericytes are important for proper vascular development during embryogenesis, where lack of pericyte recruit-ment results in vascular impairment [150].

Besides being similar to vSMCs, in some aspects pericytes also resembles interstitial fibroblasts. The major difference between pericytes and fibro-blasts is the location; pericytes are per definition positioned juxtaposed to the blood vessel, whereas fibroblasts are present in the interstitium. Also, fibro-blasts produce collagen type I, which pericytes do not[143]. Interestingly, isolated pericytes have been shown to differentiate into both fibro-blasts[143], myofibroblasts[151] and vSMCs[152], as well as into other mesenchymal cells, such as chondroblasts and adipocytes[153]; os-teoblasts[154] and Leydig cells[155]. This implies that there are mesenchy-mal progenitor cells within the pericyte population, but a detailed identity of this population is currently unknown.

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The lymphatics of loose connective tissue The lymphatic vessels constitute a draining system for excess fluid in tissue. The lymph is drained into the initial lymphatics, which are blind ended ves-sels present in loose connective tissue. The lymph is then transported via larger collecting vessels into central lymph vessels that eventually return the lymph into the blood circulation. Like blood vessels, lymphatic vessels are lined with endothelial cells. However, the lymphatic endothelium has several differences in morphology and function compared to blood vessels. First, there are no tight junctions between endothelial cells in lymphatic vessels, which reflect the function of lymphatics that is to let fluid, larger molecules and cells out of the tissue. Also, in line with the functional demands of lym-phatic vessels, there is no continuous BM. Instead, the lymphatic endothelial cells are connected to the underlying ECM through anchoring fibrils. The coverage of mural cells is also sporadic, but the larger collecting lymphatic vessels have smooth muscle cells (SMC) associated with certain segments of the vessels, which are the valve portions. The valves are comprised of endo-thelial cell processes that extend out in the lumen, and when the SMC con-tract in order to propel the lymph forward, these flaps block the anterior por-tion of the lumen. This inhibit retrograde flow of lymphatic fluid[156]. Apart from, being the sewage system of tissues, the lymphatic system is also im-portant during inflammatory processes, because APCs can use these vessels for transport to local lymph nodes, where antigen presentation can take place.

Growth factors that are important for tissue activation, remodeling and fibrosis Platelet derived growth factor (PDGF) Four different gene products build up the PDGF family of growth factors. Two of them, PDGF-A and -B have been known for almost 30 years and the other two, PDGF-C and -D were found later. PDGF-A and -B chains form hetero- and homodimers, whereas PDGF-C and -D is only known to form homodimers. The PDGF-AB heterodimer is the most abundant isoform in platelets[157], which was the source from which PDGF was originally iso-lated. Cells that express both PDGF-A and B, also produce both hetero and homodimers without any preference, which indicates that the process proba-bly occurs at random[158]. The PDGF family is one of the best characterized growth factor systems and mesenchymal cells are the main receptor-bearing cells outside of the CNS. Two different genes encode one receptor chain each, α and β, which are transmembrane receptor tyrosine kinases that dimerize upon ligand binding. PDGFRα and PDGFRβ are expressed to-

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gether in most mesenchymal cells and the receptor chains can both hetero and homodimerize, but the significance of PDGFRαβ signaling is less clear compared to the well studied signaling transmitted from homodimers of the α and β receptors. PDGF dimers differ in affinity for the different receptor chain combinations. Both PDGFRα and PDGFRβ have intracellular tyrosine kinase domains that autophosphorylates after dimerization. This elicits an intracellular signaling cascade that leads to certain biological endpoints. The ligand-receptor complex is then internalized and the whole complex is either degraded or the receptor ligand complex dissociates, and the receptor is re-cycled back to the plasma membrane[159].

PDGF-A PDGF-A exists in two isoforms due to alternative splicing. The shorter vari-ant lacks a HSPG binding retention motif, which makes this variant more diffusible. PDGF-A has many functions that are fundamental for normal embryonic development. PDGF-A KO mice die before birth or shortly after, due to inadequate lung alveoli formation, which give the mice a phenotype that resembles lung emphysema[160]. PDGF-A is also important for the proper involution of epithelial tissues during embryogenesis. This is not a direct effect on epithelial cells, since the receptor bearing cells are of mesen-chymal origin located near the epithelial lining[161, 162]. In the CNS, PDGF-A has a role in Oligodendrocyte proliferation and migration during embryonic development. Lack of functional PDGF-A during development also results in hypomyelineation of nerve fibers in the CNS[163] and results in a phenotype characterized by tremor. PDGF-A is expressed by a wide range of cell types and induces cell growth and actin rearrangement in recep-tor bearing cells in vitro[164]. In vivo, PDGF-A is expressed during wound healing[165] and in pathological conditions, such as cancer. For example, PDGF-A acts as an autocrine growth stimulant of glioma cells[166]. Over-expression of PDGF-A in vivo leads to hyperproliferation of mesenchymal and glial cells[167].

PDGF-B PDGF-B also contains a heparin biding motif, which enables binding to HSPGs in the extracellular matrix and on cell surfaces. As with PDGF-A, the mouse knockouts of PDGF-B are lethal[167] and KO mice die due to severe vascular hemorrhage, edema and lack of renal mesangial cells. The hemorrhage is caused by deficient recruitment of vascular mural cells to blood vessels, with the exception of the aorta which has a vSMC coat. In the developing fetus, the endothelial cells express PDGF-B and the high affinity receptor, PDGFRβ, is expressed on pericytes and vSMC. Endothelial de-rived PDGF-B induces both proliferation and migration of receptor bearing cells during embryonic vascular development, which is crucial for proper

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investment of mural cells in patent vessels. Vessels that lack pericytes or vSMC are instable and abnormal[150].

The development of glomerular vasculature in the kidney is also affected by the lack of PDGF-B. Insufficient recruitment of mesangial cells, which is the glomerular equivalent of vascular pericytes, give rise to a primitive capillary plexus instead of the highly involuted capillary network that is required for proper glomeruloid function[168], suggesting an important role of mural cells in vascular remodeling. The importance of the matrix binding retention motif for vascular development is underscored by the finding that mutant mice that only lack this motif also exhibit vascular defects due to defective mural cell recruitment[169].

In the adult, PDGF-B has important effects in tissue remodeling and in pa-thological conditions. PDGF-B plays a major part in several steps of tissue remodeling, such as recruitment of leukocytes[170] and activation, migration and expansion of mesenchymal cell populations. Especially, the interstitial mesenchymal population is activated by PDGF-B, and this population is to large extent responsible for the deposition of new ECM during tissue activa-tion. However, PDGF-B by itself has not been reported to be a potent stimu-lator of ECM production in vitro, with the exception of fibronectin[171]. On the other hand, PDGF-B activates many processes in mesenchymal cells, including protein synthesis, which results in increased collagen translation, but no specific induction in collagen transcription[172]. PDGF-B can also elicit angiogenesis in the CAM (chorioallantoic membrane; i.e. chicken egg) assay. Since endothelial cells generally lack receptors for PDGF-B, this ef-fect is probably secondary to stimulation of receptor bearing mesenchymal cells. Indeed, an expansion of the extravascular connective tissue was also reported[173], which suggests that stromal mesenchymal cells also were activated. Even if PDGF-B has been reported to accelerate delayed wound healing[174], most attempts to overexpress PDGF-B in tissues results in fibrosis[175]. Interstitial fluid pressure (IFP) is partly controlled by fibro-blast contraction of the ECM and PDGF-B induces increased contraction when administrated into tissues, and this results in an increased IFP[176].

A viral homologue of PDGF-B, the v-sis oncogene, is capable of transform-ing receptor-bearing cells in vitro and in vivo and PDGF-B is detected in a large number of different malignancies at both mRNA and protein level, but it is mostly in sarcomas that PDGF-B has direct stimulatory effect on the cancer cells[177]. In carcinomas, which also express high levels of PDGF-B, the main target seems to be the tumor stromal cells. These mesenchymal cells respond to PDGF-B and initiate tissue remodeling, which is further regulated in concert with other factors that are present in high levels in any carcinoma[178]. Considering the central role of PDGF-B as an activator of

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mesenchymal cells and their role in tissue remodeling, it is not surprising that PDGF-B also is expressed in a number of other non-neoplastic patho-logical conditions, such as atherosclerosis, rheumatoid arthritis, lung and kidney fibrosis[175].

PDGF-C This PDGF family member has an extra domain called CUB for Comple-ment (C1r/C1s), Uegf and Bmp1, which were the proteins in which the CUB domain was first identified. This domain needs to be cleaved extracellularly in order for PDGF-C to bind and activate its receptor. The CUB domain is also present in PDGF-D but not in PDGF-A and -B. There are basic amino acid residues at the C-terminal end of PDGF-C that facilitates binding to HSPG[179].

Mice that lack PDGF-C die perinatally with a phenotype consisting of a cleft secondary palate and blistering of the skin[180]. However, this does not occur in PDGF-C mice in a mixed genetic background, consisting in SV129 and C57Bl/6[181]. Interestingly, the blistering of the skin was suggested to be derived from lack of anchoring fibrils, which connect the epidermal BM to the connective tissue of the dermis. This could imply that PDGF-C has a role in inducing the expression of specialized ECM components that are part of the anchoring fibrils, e.g. collagen type VII, instead of being an inducer of bulk ECM components, e.g. collagen type I. Alternatively or additionally, PDGF-C could be an inducer of ECM structure organization in the dermal-epidermal junction. PDGF-C has been reported to have pro-angiogenic prop-erties in the cornea pocket and CAM assays[182], which suggests that also PDGF-C could have a general role in tissue remodeling. In consistence with that notion, PDGF-C is upregulated in models of inflammation[181] and carcinongenesis[183], and inhibition of PDGF-C in these models has posi-tive effects on morbidity. In both situations are the stromal reaction reduced, when PDGF-C is inhibited. In agreement with these studies, ectopic overex-pression of active PDGF-C induces inflammation that results in fibrosis[184] and even carcinoma[185]. However, in a model of glomeronephritis, treat-ment with active PDGF-C has an positive effect through its ability to pro-mote angiogenesis[186], which in this model rescues the tissue from col-lapse. The positive effects of PDGF-C treatment are further supported by studies of ischemic tissue, where administration of PDGF-C gives more fa-vorable results[187]. Again, the angiogenic response to PDGF-C rescues the already worn out tissue from further depression.

PDGF-D PDGF-D like PDGF-C contains a CUB domain that needs to be cleaved before it can activate its receptors[188]. PDGF-D has not yet been shown to have any affinity for heparin. The developmental role of PDGF-D is unclear

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since no PDGF-D null mouse has as yet been described, but expression of PDGF-D has been detected in brain, eyes and kidneys during embryogene-sis, which infer a role of PDGF-D in development. In the adult, PDGF-D is constitutively expressed in several tissues and its expression is also upregu-lated in sites of active tissue remodeling[189]. For example, PDGF-D is upregulated in renal fibrosis, with high protein levels in both the kidney and in serum[190]. Like the other PDGFRβ ligand PDGF-B, overexpression of active PDGF-D causes fibrosis in tissues[189]. Many of the effects of PDGF-D overexpression is overlapping with PDGF-B overexpression and PDGF-D also have the ability to transform cells when ectopically ex-pressed[191], which is not surprising since they activate the same receptors. PDGF-D and PDGF-B differ, however, in bioavailability due to the lack of heparin binding motif in PDGF-D. In a transgenic model where PDGF-D is overexpressed in the skin, an increased number of tissue macrophages are noted, but no hyperproliferation or fibrotic response in the dermis was re-ported. However, the interstitial fluid pressure was increased[192], which could be due to increased contraction of the ECM by interstitial fibroblasts in response to PDGF-D, or as a response to other factors released by macro-phages.

Platelet derived frowth factor receptors (PDGFR) PDGFRα All naturally occurring PDGF dimers can bind and activate PDGFRα. As with its ligands, this receptor is crucial for normal embryonic development. PDGFRα null mouse embryos die before birth and the fetuses exhibit a phe-notype that resembles the combined phenotypes of the PDGF-C and PDGF-A KO mouse, which indicate that PDGF-A and PDGF-C have overlapping roles in development[167, 180]. PDGFRα is expressed on a wide variety of mesenchymal and neuroectodermal cells during embryogenesis. This pattern persists in adulthood, but there are reports that PDGFRα is expressed on other cell types as well[159]. The expression of PDGFRα is to a large extent inducible and various conditions and cytokines regulate its expression. PDGFRα signaling is important in inflammations as well as in fibrosis and cancer (see above). Activation of PDGFRα in vitro leads to proliferation and actin rearrangement. Whether PDGFRα activation induces chemotaxis or not is debated[164, 193].

PDGFRβ PDGF-BB and PDGF-DD can bind and stimulate PDGFRβ phosphorylation. PDGF-CC has been shown to bind and induce PDGFRβ phosphorylation in cells that has been transfected with both PDGFRα and PDGFRβ, which would indicate that PDGF-C activates receptor heterodimers[194]. However,

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this has not been confirmed in cells that have an endogenous receptor ex-pression. The PDGFRβ KO mice display the same phenotype as the PDGF-B null mice[167], which suggest a minor role for the other ligand, PDGF-D in development. The expression pattern of PDGFRβ and PDGFRα are to a large extent overlapping, but the expression of PDGFRβ is generally higher than PDGFRα in mesenchymal cells[159]. Platelets and Oligodendrocyte precursors only express PDGFRα. On the other hand, leukocytes and lym-phocytes only express PDGFRβ[159] and this may explain the pro-inflammatory effects of PDGF-B. As with PDGFRα, the expression of PDGFRβ is inducible and the expression is often upregulated in the same conditions as PDGF-B[195]. The actions of PDGF-B is transmitted through PDGFRβ and executed by the receptor bearing cells in these inflammatory and pathological conditions. As mentioned above, PDGFRβ has an impor-tant role in both adult and embryonic angiogenesis, due to the expression of the receptor on pericytes[167, 196, 197].

In vitro effects of PDGFRβ stimulation include proliferation, actin rear-rangement and chemotaxis. The cellular response is in part depending on the concentration of ligand, since partial actin rearrangement is induced at lower levels than proliferation[198]. Full actin rearrangement is achieved at doses that also induce proliferation. Actin rearrangement is a fast process that reaches a maximum change in morphology in less than 1 hour. Proliferation on the other hand is more complex and requires gene transcription, which makes this response slower and it takes approximately 12 hours before the cells enter the S-phase. Most studies of proliferation has been carried out with a continuous stimulation with PDGF-BB, but shorter stimulations can induce proliferation as well[199]. This implies that the intensity of the stimu-lation is important for the biological endpoint and that there could be a thre-shold effect. Stimulation with PDGF-BB downregulates PDGFRβ[200], which further implies that a continuous stimulation may not be needed.

Since PDGF-B has a heparin binding retention motif, it can be presented to its receptor as a soluble ligand as well as immobilized on cell surfaces and on matrix molecules. Carcinoma cell derived PDGF-B stimulates adjacent fibroblast PDGFRβ mainly through cell-cell contacts[201]. Mutant mice lacking the PDGF-B retention motif or the native HSPG binding partner, have impaired pericyte migration[202]. This could be due to improper inter-action of PDGF-B with cell surfaces or that no functional gradients could be achieved along the endothelium.

Transforming growth factor β (TGF-β) The TGF-β superfamily, comprises many members with important roles in development, tissue remodeling and homeostasis in most tissues. TGF-β1,

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TGF-β2 and TGF-β3 are most important for this regulation in loose connec-tive tissue. These TGF-βs are encoded by three different genes and the pro-teins exists as homodimers[203].

The TGF-βs bind to TGF-β type II receptors (TβR-II, a single gene TGFRB2), which subsequently forms a tetrameric receptor complex with two TGF-β type I receptors (TβR-I a.k.a. Activin like kinase: ALK) and an additional TβR-II. Both types of receptors possess serine/threonine kinase activity, and upon formation of the receptor ligand complex, TβR-II phos-phorylates ALK, which in turn activate downstream signaling molecules. There are two types of ALKs that are involved in TGF-β signaling, ALK1 and 5[204]. Endoglin is an auxiliary TGF-β receptor (TβR-III), which is required for binding of TGF-βs to ALK1. Betaglycan is also a TβR-III that specifically facilitates the interaction between TGF-β2 and TβR-II[203]. Receptors for TGF-β are expressed on a wide variety of cells of different origins, including endothelial, epithelial, mesenchymal and hematopo-etic[205].

The importance and functions of the TGF-βs and their receptors during de-velopment has been revealed through studies of KO mice. Any deficiency in TGF-β ligand or receptor results in embryonic or perinatal lethality. Inacti-vation of TGF-β1, TβR-II, ALK1, ALK5 or endoglin results in defects in the vascular development, where both mural and endothelial cells are af-fected[206, 207]. In addition, the TGF-β1 KO also displays, defects in hema-topoesis[208] and increased inflammation in multiple organs[209], which indicate that TGF-β1 also has an immune-regulating effect. Deletion of the TGF-β2 gene results in numerous defects in the formation of different tis-sues, which lead to perinatal death. The TGF-β2 KO display, defects in lung, cardiovascular, skeletal, eye, ear and urogenital development. Most of these defects are the results from impaired recruitment or expansion of cells and failure to deposit ECM[210]. TGF-β3 KO mice also die perinatally, due to immature lungs, in addition they also display a cleft palate[211]. Many of the KO phenotypes displayed by the TGF-β system, have similarities with mice deficient in components of the PDGF system, which indicate that these systems are acting in concert during embryogenesis.

Activation of TGF-β

Before TGF-β has the opportunity to bind its receptors, it has to be activated. TGF-β is secreted as an inactive homodimer, which is associated with the latency-associated protein (LAP) and large TGF-β-binding protein (LTBP 1, 3 or 4). The LAP peptide is actually a part of the TGF-β peptide, which is cleaved by furin convertase and the LAP then bind non-covalently to TGF-β. LAP inhibits binding of TGF-β with its receptors and the TGF-β–LAP com-plex is called small latent complex (SLC). There are few cells that have been

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reported to be able to secrete the SLC, most cells secrete the large latent complex (LLC), which include a LTBP that is covalently attached to the LAP. The LTBPs have affinity for various ECM proteins as mentioned above, in addition to being ligands for αV-integrins. This conveys that most TGF-β that is secreted is inactive and stored in the ECM. There are several known ways by which latent TGF-β can be activated. Proteolytic release of TGF-β from LLC can be mediated by a large number of proteases: BMP1, various MMPs, plasminogen and both types of plasminogen activators (tPA and uPA), thrombin, elastase and cathepsin[122]. In addition, the matricellu-lar protein TSP-1 is also capable of activating latent TGF-β through interac-tion with LAP. A ternary complex between TSP-1, LAP and TGF-β is formed, which allows for receptor activation. This occurs independently of the presence of LTBP[97].

The interaction of LAP with certain integrin heterodimers, can also lead to activation of latent TGF-β. All αV-integrins (αVβ3, αVβ5, αVβ6 and αVβ8) and probably one of the β1 integrins (α5β1 and/or α8β1) that bind LAP, can perform this activation. The evidence for the integrin based activation of TGF-β originates from the αV, β6, and β8 KO mice, which display pheno-types that overlap with the augmented inflammation in the TGF-β KO.

The mechanism for how integrin mediated TGF-β activation is achieved is not totally clear, but there are currently two alternative models. The first suggests that integrin binding to the LLC facilitates the binding or action of proteases. The second model propose that tractional forces from the integrin bearing cells, either releases TGF-β completely or just changes the confor-mation of the LLC in a way that enables receptor activation[122]. The latter can in principle only allow TGF-β to bind to receptors in the absolute vicin-ity, which can be on the activating cell or a cell in close contact. In addition to activate latent TGF-β, integrin αVβ3 associates with TβR-II in lipid raft/caveolae and enhances the signaling leading to proliferation in human fibroblasts, which suggests that integrins and the TGF-β systems crosswalks on several levels[212]. The binding of LLC to the ECM is also of impor-tance, since mutations in fibrillins, which cause improper microfibril assem-bly leads enhanced TGF-β activation. This probably occurs due to that un-anchored LLC can disassemble “spontaneously”[203].

TGF-βs is expressed in almost all types inflammations and fibrotic condi-tions[213], which is reflected by the results of various experiments with TGF-β overexpression. When active TGF-β is overexpressed locally in tis-sues, the result is usually increased ECM production by mesenchymal cells and decreased proliferation of epithelial cells. Hepatic overexpression of TGF-β under the albumin promotor induces high plasma levels of TGF-β that leads to multiorgan fibrosis[214]. The increased ECM production in

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mesenchymal cells is due to direct stimulation of ECM gene transcription by TGF-β signaling[213]. However, TGF-β can further enhance the ECM syn-thesizing effect through the ability of inducing differentiation of mesenchy-mal cells into myofibroblasts, which are α-SMA expressing cells that are present in many inflammatory and fibrotic conditions. Myofibroblasts are also known to have a high collagen type I production[215]. Apart from being the most important inducer of ECM production in fibrosis and inflammation, TGF-β is also a potent suppressor of protease activity, which further aug-ments ECM accumulation[213]. TGF-β can also exert immuno-modulating effects, targeting most immune cells to some extent. For instance, TGF-β is a chemotractant for monocytes and poly-morpho-nuclear (PMN) cells, but many of their mature effector functions are suppressed. In addition, many of the effector functions of T and B cells are also suppressed by TGF-β[216].

TGF-β has a dual role during carcinogenesis, since TGF-β acts as an inhibi-tor of epithelial proliferation, but also as an inducer of tumor progression. TGF-β has through its pro-fibrotic effects an important role in promoting desmoplasia. The fibrotic nature of the microenvironment probably facili-tates the progression of a developing neoplasia into a cancer. Moreover, TGF-β can also induce epithelial to mesenchymal transition (EMT) at later stages of cancer progression. Cancer cells that have acquired mesenchymal features are more prone to metastasize[217]. Also, the propensity of TGF-β to modulate the immune-system favors tumor progression, since anti-tumor effects of the immune system is suppressed[216].

Fluid homeostasis in loose connective tissue The kidneys regulate the overall fluid balance of the mammalian body, but the local fluid balance in tissues and organs is regulated by the loose connec-tive tissue. As mentioned before, the loose connective tissue constitute a border zone between body compartments, and all exchange of nutrients, gases, waste products and fluid takes place in loose connective tissue. Also, all trafficking of cells, that migrate between different body compartments must pass through loose connective tissue at some instance. The fluid bal-ance regulation is comprised of several elements: The vasculature, the ECM of the loose connective tissue, the interstitial fibroblasts, and the lymphatic system.

The role of the vasculature The vasculature constitutes a barrier that contain plasma proteins and blood cells inside the circulation. However, the endothelial lining of blood vessels is semi-permeable; water and other low molecular weight solutes can pass over the vascular wall, whereas the permeability to plasma proteins is more

38

restricted. This infers that there exists a difference in concentration between the blood and the interstitium with regard to plasma proteins, but not for the small solutes. The vasculature also contributes another variable to the sys-tem, the blood fluid pressure, which originates from the beating heart. The fluid pressure drives the flow of fluid along the blood vessel, but it can also drive a flux of fluid across the vessel wall. When regarding the high perme-ability for water, one might wonder what stops all fluid from leaving the circulation and flow out into the interstitium? One contributing factor that prevent this from happen is the concentration difference of plasma proteins, which exert a colloid osmotic (oncotic) pressure that help to retain fluid in-side the vessel[83]. The other factors that prevent fluid loss from the circula-tion are found in the interstitium.

The role of the interstitium Since much of the ECM is composed of supra-molecular complexes that cannot diffuse, the colloid osmotic pressure of the interstitium is lower than one might expect, when regarding the high interstitial protein concentration. This infers that the difference in colloid osmotic pressure favors retention of fluid inside the blood vessel[83]. However, one component of the ECM ac-tually promotes fluid flux out of the circulation, and this is the ground sub-stance. The negative charges on the GAG chains attracts positively charged ions, which in turn are osmotically active and attracts water, and this will result in a swelling of the ground substance[84]. However, at homeostasis, the ground substance is physically restrained by the collagen fiber network, which is further controlled by the extent of contraction exerted by interstitial fibroblasts. The force generated by interstitial fibroblasts on the collagen fibers opposes the osmotic activity of the ground substance, and is referred to as the interstitial fluid pressure (IFP)[218]. However, the IFP is normally slightly negative in loose connective tissue under homeostatic conditions, and this enhances capillary filtration.

Capillary filtration The theory of capillary filtration suggests that there is a pressure difference between the afferent and the efferent portions of the microvascular bed. The higher blood fluid pressure in the afferent portion allows for a flux of fluid and low molecular weight solutes out of the circulation and into the tissue. At the efferent portion of the microvascular bed, the blood fluid pressure is low-er and the pressure gradient favors a fluid flux out of the tissue and into the circulation. This is dependent on that the other parameters only change mar-ginally[219]. This theory provides a plausible explanation for how nutrients and waste products are exchanged in the tissue. Fluid that is not reabsorbed into the venous side of the vasculature is drained via the lymphatic system.

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The Starling equation and regulation of fluid homeostasis A mathematical expression that describes the fluid flux process was formu-lated 1896 by Ernest Henry Starling, and the equation that is consequently referred to as the Starling equation contains many of the variables discussed above. The flux of fluid (JV) over the endothelium is dependent on the capil-lary fluid pressure (PC); the capillary colloid osmotic pressure (COPC); the interstitial fluid pressure (PIF); the interstitial colloid osmotic pressure (CO-PIF), the permeability of water and other small solutes over the vascular wall (LP, hydraulic conductivity); the surface area for hydraulic exchange (S); and the permeability of the vascular wall to plasma proteins (the capillary reflec-tion coefficient, σ).

JV= LPS((PC-PIF) - σ(COPC -COPIF))[219]

In simple terms, the Starling equation postulates that the flow of fluid over a vascular endothelium is dependent of the net pressure gradient and the amount of fluid that can pass in a defined vascular segment. As is shown below.

JV= LPSΔP

When the system is in balance, the rate of fluid flux across the vessel wall is dependent on the difference between the hydrostatic blood pressure and the blood colloid osmotic pressure. An increased fluid flux will be counteracted by a corresponding increase in IPF and increased lymphatic drainage[83].

The most frequent event that leads to disturbance of this homeostasis is in an acute inflammation. During an inflammation, a large set of vasoactive sub-stances are released (see next section), which affect several parameters of the vasculature (Figure 1A & B):

• Vessel dilation leads to increase of the variable S, and results in a larger area of exchange that leads to an enhanced fluid flux.

• Increased vascular permeability to plasma proteins. The reflection coefficient, σ is decreased, which infers that more plasma protein pass over the endothelium. This in turn results in a decrease in blood colloid osmotic pressure and a transient increase in interstitial colloid osmotic pressure, which further enhances fluid flux out of the vessel.

However it is not just the vasculature that is affected by mediators released during an acute inflammation. The interstitial cells are also affected, and what happens is that the contraction of the collagen matrix is relaxed, which

40

allow the under-hydrated ground substance to take up water and swell, i.e. the ground substance is allowed to exert its osmotic potential. This even further enhance the fluid flux out of the blood vessel and into the inter-stitium[218].

The only element that now can prevent the formation of an edema is an un-surpassed increase in the lymphatic drainage, which will not occur due to the limited capacity of the lymphatic system. Eventually, if the inflammatory stress is relieved, the vessel dilation and permeability will be decreased and the fibroblasts will start to contract the fibrillar collagen matrix again. This will lead to a gradual normalization of the interstitial fluid content, where the excess interstitial water will be transported away via the lymphatics[83].

Reactive tissue conditions Inflammation Inflammations are tissue reactions that can be summarized by the classical signs of a cutaneous inflammation; rubor, calor, dolor and tumor. Where rubor stands for the redness and calor the heat caused by vascular dilation and increased blood flow in the inflamed area, dolor stands for pain caused by stimulation of sensory nerve endings by inflammatory mediators and by the tissue swelling, i.e. tumor, which is caused by the leakage of fluid from the vasculature. Many pathological conditions have a pronounced inflamma-tory component. Importantly, almost all inflammatory processes involve the loose connective tissue, since it is situated between many organ parenchyma and the circulation. An inflammation commences as a response to an insult that can be caused by mechanical tissue damage, deregulation of cell or or-gan homeostasis, exposure to harmful substances or foreign microorganisms. The inflammatory response begins with an acute phase, in which immune-competent cells are recruited. The aim of this process is to contain and/or clear the tissue of harmful factors. The next step is the formation of a tempo-rary matrix that facilitates the rebuilding of the tissue. Finally, the tissue is remodeled and left is a scar that bares resemblance to the initial unharmed tissue, but most often the scar has a decreased function compared to the pre-inflamed tissue.

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Figure 1FFigure 111

Hyluronan

Proteoglycans

Endothelial cell

Pericyte

Collagen fiber

Fibroblast

Neutrofil

Mast cell

Carcinoma cell

Macrophage

A B

C

Figure 1.

A) Loose connective tissue during homeostasis. The vasculature is contain-ing plasma proteins and cells and interstitial fibroblasts restrain the ground substance. B) In acute inflammations, the vasculature leaks plasma proteins and leukocytes transmigrate into the interstitium, simultaneously the fibro-blast stops contracting the collagen network, which allows for the ground substance to swell. C) The vasculature in tumors is also leaking, but a dense collagen matrix and tumor associated fibroblasts (TAFs) that contract this matrix counteracts an edema. This partly compensate for the lack of lym-phatics.

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Acute inflammatory phase Upon any insult that is sensed by sentinels in the tissue, an acute inflamma-tory response will be elicited. As mentioned above, cells of the innate im-mune system present in loose connective tissue has the capability to recog-nize and respond to various insults, such as cell and matrix debris from damaged tissue, and intruding microbes. This is not exclusive to the immune cells, but the stimulation of these cells has more potent effects during the initial phase. All three of the immune cells present will respond, but macro-phages and mast cells will be most important during the early phase of in-flammation[220]. Activation of macrophages and mast cells will result in a quick and in the case of mast cell degranulation, massive release of vasoac-tive substances, e.g. vascular endothelial growth factor A (VEGF-A), and histamine. VEGF-A is a very potent inducer of vascular permeability[221], which is released by mast cells[116], macrophages[222] and platelets[223]. In addition to increasing the vascular permeability to plasma proteins, VEGF also induces dilation of post-capillary venules[224]. Histamine is an amine derivative released from mast cells that also induce vascular permeabil-ity[116], although not as potent as VEGF-A[225], histamine is released in large quantities. The actions of VEGF-A and histamine will affect the fluid balance homeostasis, and as aforementioned this will lead to increased flux of fluid into the interstitium. The increased permeability will also lead to an extravascular deposition of plasma proteins, e.g. fibronectin, vitronectin and fibrinogen. Extravascular fibrinogen will subsequently be cleaved and form a extravascular fibrin matrix. Out of the different portions of the microvascu-lar unit, it seems to be the postcapillary venules that are most affected by the activation with vasoactive substances[226].

Not only vasoactive substances are released during the initial phase of an acute inflammation, also general acting inflammatory mediators are released, e.g. IL-1, TNFα, and chemokines. The role of the inflammatory cytokines IL-1 and TNFα is to further increase the level of activation of all cells in the tissue[223]. Of importance is their effect on interstitial fibroblasts, which in their presence relieve their contraction of the collagen fibrillar matrix, as mentioned above this allows the extravasated water to be taken up by the ground substance, which will subsequently swell and this results in an ede-ma[227]. Another cell type whose activation also is of outermost importance is the endothelial cell, which has stored cytoplasmic granules that are ex-posed on the luminal surface upon stimulation with IL-1 and TNFα. These granules are referred to as Weibel-Palade bodies and they contain preformed proteins involved in blood clotting and leukocyte adhesion (selectins)[228]. The role of the chemokines is to attract circulating neutrophils and mono-cytes from the blood. Circulating leukocytes are slowed down by the se-lectins exposed on the endothelial surface of post-capillary venules. The

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leukocytes are said to start rolling on the endothelial surface, which not just exposes selectins. Chemokines are also exposed on the endothelial surface in addition to cell-cell adhesion molecules, such as ICAM-1 (Inter Cell Adhe-sion molecule). The chemokines induce the leukocytes to activate β2 in-tegrins, which can bind to ICAM-1. These interactions are more firm and the leukocyte stops rolling and starts to transverse the endothelium in order to migrate out into the inflamed tissue[129] (Figure 1B). Well inside the in-flamed tissue, the extravasated leukocytes continue to follow chemokine gradients. In cutaneous wounds, neutrophils accumulate near the wound surface where they form a barrier against the potential intruders[229]. How-ever, the presence of neutrophils in the tissue also comes with a price, since they release numerous harmful substances that not only target intrud-ers[230].

There is a high level of migration taking place in the inflamed tissue, and this is facilitated by the edema that leads to a more porous ECM. Another feature that favors migration is the partial degradation of ECM because of the increased release of proteases. IL-1 and TNFα are potent inducers of metalloproteinases (MMPs)[223], and some of these have the capability to even break down native collagen, which normally is resistant to most prote-ases. Both mesenchymal and inflammatory cells are sources of MMPs, and inflammatory cells also express other types of proteases, e.g. neutrophil elas-tase and mast cell tryptases. These proteases will modulate the activity of other proteases, e.g. zymogens like MMPs can get activated, whereas other proteases can get inactivated. Moreover, proteases can also modulate the activity and availability of growth factors and cytokines, e.g. TGF-β.

The elevated protease activity in the inflamed area leads to exposure of cryp-tic sites in processed matrix molecules[231], which potentially can induce novel adhesion interactions for the migrating cells[232]. In addition, the matricellular proteins are up-regulated during inflammation, and as afore-mentioned these are known to modulate integrin-ECM interactions. Many of the matricellular proteins modulate adhesion, which probably prevents cells from becoming immobile[85, 87]. The normal reportoire of integrins ex-pressed on endothelial and mesenchymal cells are suited for performing dif-ferentiated tasks during homeostasis. However, during inflammation the ECM displays a wide variety of substrates in the inflamed tissue and migra-tion on these substrates require integrin receptors with broad ligand specific-ity. Integrin heterodimers containing αV fulfills this requirement, especially αVβ3 and αVβ5 are up-regulated in endothelial and mesenchymal cells during inflammatory conditions[233]. The up-regulation of αV-integrins comes timely when regarding that the expression of TGF-β is increased, and that αV-integrins contribute to its activation.

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The release of IL-1 and TNFα induces a catabolic state in the inflamed tissue with breakdown of ECM and formation of edema. The purpose of this re-sponse is to quickly put cells that can counteract infectious agents on loca-tion in the affected area in order to contain a potential infection. However, in paralell are also more anabolic factors released, e.g. PDGF, TGF-β, EGF and fibroblast growth factors (FGFs)[234]. PDGF-B induces contraction of the collagen matrix by interstitial fibroblast, which counteracts the edema. This is a process that requires αVβ3 expression on fibroblasts[176]. More-over, mural pericytes are detached from activated blood vessels during in-flammations[235, 236]. Pericytes express PDGFRβ[196] and PDGF-B has been shown to induce pericyte detachment in activated vessels during em-bryonic development[149]. This pool of detached pericytes most likely con-tributes to the increase in mesenchymal cells in the interstitium of inflamed tissues. PDGF-B is a known mitogen and chemotractant for these cells, and pericytes migrate out from isolated vascular fragments and differentiate into collagen I producing fibroblasts in vitro [143]. Moreover, inhibition of PDGFR signaling by the tyrosine kinase inhibitor Imatinib limits the size of this population in wounds[237]. TGF-β is an inducer of matrix production and synthesis of protease inhibitors, in addition to being immuno-modulating. However, both PDGF-B and TGF-β also contribute to the in-flammatory response by being chemotractants for inflammatory cells. EGF is an important mitogen and migration inducer for epithelial, endothelial and mesenchymal cells. The FGFs is a large family of growth factors, and FGF-2 is of major importance for its role in stimulating growth, migration and sur-vival in endothelial and mesenchymal cells. FGF-7 (a.k.a. keratinocyte growth factor, KGF) has an important role in stimulating epithelial cells[223]. These growth factors collectively counteract the deteriorating effects of the inflammatory response, but still in the early phase of inflam-mation the balance is shifted in favor of the catabolic factors.

The cutaneous wound healing model allow for a temporal observation of the different pahses in inflammatory progression. In this model the epidermis represent the organ parenchyma and the dermis, which contains both dense and loose connective tissue, represents the interstitium, which nerves, blood vessels and lymphatics run through. The wound healing model is appropriate for describing the resolution of inflammations and therefore it will be used to describe these events. There are, however, certain things that one should bare in mind when evaluating results from a wound healing model, which is not applicable to all inflammations. Inflicting a wound in an external surface of the body will never result in a sterile inflammation, which will affect the extent of immune system involvement. Moreover, due to the high vasculari-zation it is hard to inflict a wound without damaging blood vessels, which in turn leads to excessive clotting and platelet degranulation. The platelet gran-ules comprise an additional source of inflammatory mediators and growth

45

factors that will not always be present in other types of inflammations. In-flammations in internal organs, induced by various toxic stimuli constitutes examples of other types of inflammations, which are sterile and do not in-volve blood vessel damage to the same extent as in wound healing. The re-solving phases of the wound healing response will be outlined below.

Granulation tissue formation The next step of the inflammatory response is the formation of a granulation tissue, which is a process that is particularly prominent in wound healing. At this point, if there has been a successful clearance of any invading microbes, there will be a shift in the predominant inflammatory cells present, from neutrophils to macrophages[238]. Lymphocytes will also be present in higher numbers than in the acute phase. As mentioned above, depending on the type of activation, macrophages can assume a M1 or M2 phenotype, and in the initial phase of inflammation the M1 phenotype dominates, whereas in the later phases the M2 phenotypes becomes predominating. The shift from M1 to M2 conveys a difference in which factors that are released and ma-crophage functions. The role of M2 macrophages is to perform debridement of the inflamed tissue, by phagocytosing dead cells and endocytosing mo-lecular debris. M2 macrophages secrete anabolic and anti-inflammatory fac-tors, such as PDGF, TGF-β and IL-10[109]. The latter being an anti-inflammatory cytokine that inhibits the production of pro-inflammatory cy-tokines, e.g. TNFα[239]. The effect of PDGF and TGF-β is to promote ex-pansion of mesenchymal cells, which subsequently migrate and populate the granulation tissue. The exact origin of these cells is currently unknown, as mentioned above these could originate from perivascular cells[151, 240, 241]. However, other origins has also been proposed[242], and most likely several of the proposed sources contibute to the final population. Regardless of their origin, TGF-β will induce them to produce ECM molecules, mostly fibronectin but also collagen I. TGF-β also induce the production of tissue inhibitor of matrix metalloproteinases (TIMPs), which as their name sug-gests inhibit MMPs[243, 244]. This leads to an enhanced ECM deposition, which will fill out the tissue deficit created by the inflammation.

Another important event during the formation of granulation tissue is the formation of new blood vessels, angiogenesis. This is a process in which new blood vessels are formed by expansion of preexisting ones. Growth factors, such as VEGF-A and FGF-2 are important for this process[222]. In addition, hypoxia within the granulation tissue also leads to expression of pro-angiogenic factors, since many genes involved in angiogenesis is under control by hypoxia inducible factors. However, hypoxia is neither evident nor required for the expression of pro-angiogenic growth factors in the initial phases of wound healing[245]. Angiogenesis in adult tissues often originates from the activated post-capillary venules, which in conjunction with discus-

46

sions about angiogenesis also have been referred to as mother vessels (MV). The MVs are activated segments of vessel, which at this time display a loose pericyte coat, since activated pericytes have detached[224]. In addition, leu-kocytes have used these vessels for transmigration, which infer that the BMs around these vessels would be more permissive for migration[246]. Angio-genesis in adult wound healing seems to by a mechanism in which endothe-lial cells and pericytes act in concert to expand the vasculature into the in-flamed, avascular area[247-249]. The leading portions of the growing vasculature display decreased mural cell coverage and discontinuous BMs, which lead to frequent ruptures and leakage of red blood cells[250]. Far less is known about the formation of new lymphatic vessels during inflamma-tions and wound healing. However, abundant nascent lymph vessels are pre-sent within the granulation tissue[251] and growth factors that stimulate this process are available, e.g. VEGF-A[252] and -C[251]. The migration per-formed by endothelial cells, pericytes and other mesenchymal cells during this stage in inflammation, still requires proteases that can pave the way. Also, proteolytic activity is needed to remove unwanted ECM components, such as fibrin.

Mostly so far has the reciprocal signaling and involvement of endothelial, mesenchymal and inflammatory cells been discussed. However, the paren-chymal cells, which most often are epithelial, also participate in regulating the evolving inflammation. This is especially important in wound healing, because the barrier against the outer world is comprised of some kind of epithelium. A wounded tissue that faces the outside world needs to be reepi-thelized before proper healing can be achieved. The epithelial cells need signals that promote growth, survival and migration. Also, epithelial cells needs to receive signals that induce expression of proteases in order to facili-tate migration[234].

Remodeling and contraction When the granulation tissue has filled out the tissue deficit, it contains a surplus of cells, ECM and blood vessels that are immature and poorly organ-ized. During the consecutive phase are differentiation and maturation in-duced, and excess cells and blood vessels will disappear. The result will be a transformation of granulation tissue into scar tissue. This transition is ac-companied by a decrease in growth factor expression[253]. However, TGF-β is still one of the most important factors during this phase, since its impor-tance for the expression of ECM molecules and the enzymes that participate in their remodeling. In addition, TGF-β induces differentiation of fibroblasts into myofibroblasts in the wound area. Myofibroblasts are interstitial cells that unlike resting fibroblasts synthesize high amounts of ECM proteins and express contractile proteins of the same type as SMCs, e.g. α-SMA. The myofibroblasts appear in a healing wound at the same time as wound con-

47

traction occur and it has been proposed that myofibroblasts are the effectors of this contraction, due to their expression of contractile proteins. Moreover, myofibroblasts also synthesize ECM proteins, e.g. collagens, TNC and ED-A fibronectin, the latter is also crucial for the differentiation of fibroblasts to myofibroblasts[254]. The importance of TGF-β for differentiation into myo-fibroblasts was recently shown in mice that have a fibroblast lineage specific deletion of TβR-II. These mice display defective wound contraction in con-junction with an absence of α-SMA expression in wound interstitial cells[255]. This clearly indicates a central role for TGF-β in myofibroblast differentiation, but the tensional state of the ECM also seems to be of impor-tance. In vitro, myofibroblasts cultured under high tension develop static focal adhesions of an increased size, referred to as super-mature focal adhe-sions. These has been suggested to have a role in sensing the mechanical tension in the environment, by inducing or modulating signal transduction events[254]. Another role of the increased tension in the ECM could be that it would favor TGF-β activation by αV-integrins, by the model where cells pull the latent complex apart. The αV-integrins might still be important for TGF-β activation, but the remodeling and interactions with fibrillar collagen requires collagen binding β integrins, predominantly α2β[256]. Although tension is needed for differentiation into myofibroblasts, the myofibroblasts themselves contribute to even more tension. The exact role of this tissue hypertension is unclear, but tension induces expression of collagens and collagen modifying enzymes by fibroblasts that contracts stressed collagen gels in vitro[257]. LOX is among the up-regulated enzymes, and the expres-sion of LOX is high during wound healing, but active LOX is not detected until later timepoints, when ECM remodeling occur[258]. LOX is the major collagen crosslinking enzyme, and an increased number of crosslinks could favor stabilization and maturation of the collagen scaffold. Moreover, cros-slinked collagen has also been shown to induce faster collagen contraction by fibroblasts in vitro[259]. This suggests that tissue hypertension might be important for proper tissue organization and ECM maturation in scar forma-tion. Further, the ECM maturation is also accompanied by a decrease in the ratio of collagen III to collagen I, which indicate that a more dense fibrillar matrix is deposited[260].

The reason for the regression of excess cells and blood vessels in the matur-ing scar is largely unknown, but the rate of apoptosis is increased in both interstitial and vascular cells[261]. The reason for this could be the de-creased expression of growth factors, e.g. VEGF-A, PDGFs and FGFs, at this stage[222, 253, 262]. However, since not all cells and vessels regress upon the decline in growth factor expression, there should be an additional maturation process that some cells and vessels go through that allows them to be independent of external growth factor stimuli. Even if the ultrastructure of the collagen fibers from dermal scars look largely reminiscent of normal

48

skin[263], dermal scars never attain the same tensile strength as unharmed skin[234].

Fibrosis If a scar never matures, the inflammation becomes chronic and results in a fibrosis. Excessive deposition of collagen type I that is alternatively cros-slinked is a hallmark of fibrosis. This process is often irreversible and can occur in almost any organ. Fibrosis in internal organs is a major cause of morbidity. The typical fibrotic sequels of dermal wound healing are hyper-trophic scars and keloids. These two fibrotic conditions have many features in common, but the major difference is that keloids spreads beyond the ini-tial borders of the wound[264]. In both types of skin fibrosis are the scar formation process prolonged, which leads to an excessive ECM produc-tion[265]. The reason for this is unknown but there is clearly a dysregulation of the reciprocal signaling between T-cell, macrophages and fibro-blasts/myofibroblasts. TGF-β and the TH2 cytokines IL-4 and IL-13 are im-plicated to be the main culprits, since all three can be found at elevated lev-els and all of them induce ECM production either directly or via TGF-β[266]. Apart from inducing ECM production, TGF-β also has a role as an immuno-modulator that can suppress immune cell functions. The optimal regulation of the scar formation, probably involves that TGF-β exert both its functions, but if the immune system is unresponsive to TGF-β mediated signals, only the fibrogenic effects will be prevalent. Indeed, regulatory T-cells (Treg) can secrete the anti-inflammatory cytokine IL-10 in response to TGF-β[267], but if these cells are unable to respond or are not present in sufficient numbers, there will be fewer signals that lead to dissolution of the scarring process. Moreover, IFN-γ secreted by TH1 can inhibit the actions of IL-13 on macrophages and fibroblasts[266], but if there has been a polariza-tion of the T-cells into TH2 this action will not either take place. This further suggests that the excessive scarring seen in fibrosis probably is regulated by the reciprocal signaling between cells present in the microenvironment and how they are conditioned to respond to a given signaling mediator.

Hypertrophic scars and keloids display features of the maturing scar, whose progress to resolution has been halted. On the histological level, this can be seen as the increased density of blood vessels and that these have an altered, dilated morphology[268]. There are also more cells present in the inter-stitium than in mature scars, which suggests that the apoptosis that reduces cell numbers in normal scars is inhibited[265]. This could be due to the con-tinued high expression of growth factors in fibrotic lesions[264]. Interest-ingly, unwounded quiscient loose connective tissue display an elaborate network of inter-cellular junctions between interstitial fibroblasts[103], but this network is much less developed in hypertrophic scars and keloids. It has

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been suggested that cell to cell signaling via these junctions could regulate apoptosis[269]. Also the interstitial ECM displays an altered morphology and composition. The collagen III to collagen I ratio is still high[260], and the ultrastructure of the collagen fibril bundles or fibers display a disorgan-ized morphology[263]. The altered collagen scaffold morphology could be due to an altered crosslinking pattern in fibrosis, and the lysylhydroxylase 2 isoform b (LH2b) is specifically expressed in fibrosis. LH2b mediates hy-droxylation of collagen telopeptides and the crosslinks, mediated by LOX, that results from these hydroxylysines are not normally found in loose con-nective tissue. These crosslinks renders collagen more resistant to the action of the native collagenase MMP1[270]. This indicates that the collagen could be too stable and that proper remodeling might not be able to occur. Interest-ingly, LH2b mRNA is up-regulated by all TGF-β isoforms, which is not surprising, but also TNFα and IL-4 up-regulates this enzyme. TNFα is a known suppressor of ECM synthesis, but the positive regulation of LH2b suggests that it participate in diverting the ECM phenotype in fibrosis[271]. The fibrotic tissue also shows an altered expression of SLRPs like decorin, whose expression is decreased. As mentioned above, decorin participates in collagen fibril assembly and low decorin expression could partially contrib-ute to the altered collagen scaffold morphology. In addition, the decreased expression of decorin could infer an increased activation of TGF-β, since decorin binds and sequester TGF-β[49]. In contrast, expression of some other SLRPs like biglycan and fibromodulin is up-regulated in fibrosis[68].

In contrast to fibrosis, fetal wound healing restores the original tissue archi-tecture. This is an intrinsic capacity of the fetal skin and is not attributed to the intra-uterine environment. The amount of scarring in the fetal tissue is proportional to how far the tissue has developed. This implies that as long as the tissue has not completed its development, there is a capacity for total regeneration. The contribution of inflammatory cells is reduced in fetal wound healing, and this would probably also limit the deteriorating effects of an inflammatory response[272].

The microenvironment of solid tumors Solid tumors, like carcinomas, simultaneously display similarities with both acute inflammations and fibrotic lesions, and are sometimes referred to as wounds that do not heal[273]. Many of the cells, growth factors and proc-esses that have described above are present also in the tumor microenviron-ment. Moreover, many aspects of the dysregulation that can be found in fi-brosis and chronic inflammations are also present in carcinomas. The carcinoma stroma is characterized by a substantial inflammatory infiltrate, comprised of immune cells representing both the innate and adaptive im-

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mune system. Further, the interstitial fibroblasts display an activated pheno-type, in many ways reminiscent of myofibroblasts. Angiogenesis is also constantly ongoing within a growing tumor, which leads to activated vessels with poor function. The ECM in the tumor stroma is composed of extrava-sated plasma proteins, due to the leaky vasculature, but also dense collagen scaffolds are present, due to the fibrotic reaction. These features of the tumor stroma profoundly contributes to the nature of the neaoplastic disease, and many aspects of the progression and treatment of cancers are dependent on the non-malignant cell component of the tumor[7]. In fact, a high expression of ECM and stromal related genes in breast carcinoma predicts an increased resistance to pre-operative (neoadjuvant) chemotherapy[274], which indi-cates that a large stromal component either correlates with a more malignant cancer phenotype or that a large stromal component protects the cancer cells. It has also been shown that chronic inflammations can lead to the develop-ment of cancers, mainly through the deteriorating effects of a prolonged exposure to immune activity[2].

Inflammatory cells in the tumor microenvironment The inflammatory infiltrate in carcinomas is reminiscent of that found in chronic inflammations and fibrosis. Apart from stimulating their own growth and activation, tumor cells secrete a vast range of growth factors and cyto-kines that stimulate other cells in the microenvironment, which in turn aug-ment an inflammatory response. The cytokine and growth factors that are present in the tumor microenvironment leads to a polarization of the inflam-matory response towards TH2-cells and M2 macrophages, in addition other inflammatory cell types also display altered features compared with an acute inflammatory response. However, the tumor associated macrophages (TAMs) are not identical to the M2 macrophages found in inflammations, instead they are comprised of features found in both M1 and M2 and should be treated as a separate entity. TAMs have been reported to express M1 cy-tokines like TNFα, IL-1, IL-6 and IL-8 (CXCL8) as well as the gelatinase MMP9. In addition TAMs also express M2 cytokines and growth factors like PDGFs, TGF-β, IL-10 and IL-12, as well as MMP12 and fibronectin[109]. In addition TAMs express MMP8, osteopontin, FGF-2 and VEGF-A, which contributes to angiogenesis. The bivalence of inflammatory signals released by TAMs, i.e. both acute inflammatory and fibrogenic, contributes to the resulting phenotype of the tumor microenvironment. The TAMs constitute a large part of many clinical and experimental carcinomas, and their extent have a positive correlation with bad prognosis[109]. Similar to fibrosis, high expression of TGF-β has profound implications for the regulation of the immune system in tumors. Tumors are immunogenic, i.e. they express aber-rant proteins that can be recognized by the immune system, and cytotoxic T-cells with anti-tumor activity can be found in patients and experimental tu-

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mor models. However, the tumor microenvironment with its high expression of TGF-β suppresses most anti-tumor responses by the immune system. This is in part dependent on a TGF-β mediated up-regulation of Treg, which in turn can secrete more IL-10 and suppress inflammatory responses[3]. Also, the tumor microenvironment inhibits maturation of dendritic cells, which affects antigen presentation. Dendritic cells that are not sufficiently activated when they present antigens to T-cells will not stimulate an activation that elicits a specific immune response. Instead they will signal tolerance, and this is a regulatory mechanism that prohibits autoimmunity[3]. In conclu-sion, tumor cells not only enroll the immune system to serve as microenvi-ronment modifiers that will support their growth and progression, they also suppresses the anti-tumor effects of the immune system.

The tumor vasculature All tissues that are metabolically active need a vascular supply that can pro-vide oxygen and nutrients, as well as removing waste products. Carcinomas as well as other malignant cells often have a higher glucose utilization than normal tissues[275], which even further emphasize the need for a rich vascu-lar supply. Indeed, tumor masses often contain an abundance of vascular structures, but these vessels are poorly perfused and do not contain an cor-rect hierarchy with regards to arteries, veins and capillaries[276]. The reason for the poor vascular function stems from the cytokine and growth factor environment in tumors, which induce a constant vascular activation and an-giogenesis, but at the same time this hampers vascular maturation. Carci-noma cells and infiltrating myeloid cells, like TAMs, secrete large amounts of growth factors that affect the vasculature, e.g. VEGF-A, PDGFs and FGFs. TAMs also contribute by producing proteases that can degrade ECM in order to facilitate angiogenesis. This leads to a tumor vasculature that bare reminiscence to the activated vessels in acute inflammations, which are characterized by pericyte detachment and hyperpermeability, the latter can frequently be detected in tumors by extravascular fibrin deposition[277]. Moreover, the constant angiogenesis give rise to fragile nascent vessels that are prone to rupture, which can lead to intravascular coagulation and fibrin deposition. Angiogenesis seems to originate from the venous vessels and this could explain the lack of a proper vascular hierarchy, and why vessels are dilated with a poor regulation of vascular tone[276]. The constant activation by growth factors in the tumor microenvironment probably inhibits the needed maturation that would produce a correct hierarchy, which can be seen in a mature scar. The low coverage by of mural cells and their loose association to the vasculature is often used as a hallmark for the immature tumor vessels[278]. Here PDGF-B seems to have a dual effect, in retina PDGF-B can induce both pericyte dissociation from activated vessels[149], and recruitment of pericytes to nascent vessels, which in turn can induce

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endothelial and vessel maturation[146]. The effect of PDGF-B on the tumor vasculature is probably dependent on the immediate microenvironment. For instance acute inflammatory mediators like IL-1β and TNFα induces activa-tion of both retinal pericytes and endothelial cells, which leads to increased expression of inflammatory and pro-angiogenic mediators[279]. TGF-β on the other hand is required for maturation of retinal vasculature[280], which probably occurs in concert with PDGF-B. This indicates that the action of PDGF-B on the vasculature is context dependent. In tumors, all these factors are present at the same time, which seems to favor pericyte detachment and activation. Moreover, the effect of PDGF-B on pericytes is probably de-pendent on the cellular source. PDGF-B delivered from endothelial cells in close contact is probably more important for maturation[197], whereas tu-mor and inflammatory cell derived PDGF-B probably give rise to activating responses. The defective vasculature conveys impairment in oxygen supply to the tumor, which leads to a stabilization of the transcription factor hypoxia inducible factor 1α (HIF-1α), which is a stress inducible factor that is in part regulated by the oxygen tension in the tissue. HIF-1α regulates a number of genes with hypoxia responsive elements in their promotor, many of these are pro-angiogenic factors[281]. This will further boost the cytokine storm pre-sent in the tumor microenvironment. Its is not only the oxygen level that is decreased in many tumors, the pH can also be lower than in normal tissues due to the increased glycolysis[275]. The dysfunctional vasculature also impairs the transport of lactate out of the tumor. In conclusion, tumors con-tain abundant vascular structures, but not all are perfused vessels due to im-paired regulation of vascular tone and intravascular coagulation. Thus, tu-mors are capable of inducing angiogenesis, but since the vessels produced are of poor quality, there is a demand to increase the quantity.

Tumor associated fibroblasts The interstitium of carcinomas contain not only inflammatory cells, but also fibroblasts. These fibroblasts often resemble the myofibroblasts seen in ma-turing scars, chronic inflammations and fibrosis. All fibroblasts found in the tumor microenvironment are referred to as tumor associated fibroblasts (TAFs), independent of the extent of myofibroblast marker expression. It has been shown in various experimental models where tumor cell lines has been co-cultured or co-inoculated in mice with different types of fibroblasts, that the stromal cells contribute to an increased tumorogenicity. Co-culture en-hances proliferation of carcinoma cells and co-inoculation increases tumor take and tumor progression[282, 283]. The presence of TAFs also correlates with more malignant phenotype, as seen in a rat carcinoma where α-SMA expressing TAFs were more abundant in a more malignant carcinoma sub-clone[284]. TAFs in carcinomas are induced to perform similar tasks as fi-broblasts in granulation tissue and myofibroblasts in fibrosis. They interact

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with carcinoma cells and inflammatory cells, providing factors for reciprocal signaling loops. TAFs has been implicated to express growth factors that stimulate carcinoma cell proliferation, like EGF and transforming growth factor α (TGF-α), hepatocyte growth factor (HGF), and FGFs. In turn, car-cinoma cells and inflammatory cells express growth factors that activate TAFs. Similar to inflammation, TGF-β and PDGFs and FGFs constitute the most important activators for TAFs in the tumor microenvironment[285]. The effect of growth factors on TAFs is similar to what is induced in fibro-blasts during inflammatory conditions. TGF-β induces matrix production, which in turn regulates the matrix organization[69]. Most likely, TGF-β is also responsible for the high resemblance between some TAFs and myofi-broblasts, due to induction of differentiation. Ectopic overexpression of ac-tive TGF-β by cancer cells leads to an increased content of TAFs and ECM, showing the importance of TGF-β as an inducer of reactive stroma in can-cer[286]. The same study also showed a correlation between the extent of TGF-β, FGF-1 and FGF-2 expression, and the extent of the stromal com-partment, i.e. desmoplasia. The apparent importance of TGF-β for TAF be-havior has lead to the development of strategies to selectively make TAFs unresponsive to TGF-β. In one such approach, transgenic mice that lacked TβR-II expression specifically in fibroblasts was created. These mice dis-played spontaneous epithelial transformation in the prostate and malignant transformation of forestomach epithelium[287]. These results are similar to other TβR-II KOs, where all cells are lacking TβR-II. The general TβR-II KO also displays epithelial hyperplasia or transformation of various epithel-iums. However, the fibroblast specific TβR-II KO show that signaling from the stromal fibroblasts also are important in suppressing proliferation of epi-thelial cells. In another study, isolated fibroblasts from the TβR-II KO mouse were co-inoculated with mammary carcinoma cells in mice. The re-sult showed that carcinoma cells inoculated together with TβR-II KO grew faster compared to carcinoma cells inoculated with wild type (WT) fibro-blasts[288]. A third study showed that co-inoculating prostate carcinoma cells with prostate stromal cells expressing a dominant negative variant of TβR-II, resulted in decreased growth compared to stromal cells that did not express the dominant negative TβR-II. However, the cells expressing non-functional TβR-II still supported tumor initiation to the same extent[282]. These studies indicate that fibroblast TβR-II signaling has a negative effect on tumor initiation, and positive effects on tumor progression, by enhancing growth of established tumors.

TAFs can also respond to PDGFs present in the tumor microenvironment, although not all TAFs express PDGF receptors. The PDGF-B in tumors can be derived from both malignant cells and inflammatory cells, mostly TAMs. However, there is a difference in how PDGF-B is delivered to the recipient PDGFRβ expressing cell, depending on the source. Macrophages secrete

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most of the produced PDGF-B[289], whereas some carcinoma cells retain PDGF-B on their cell surface, and subsequent stimulation of PDGFRβ ex-pressing cells occurs through heterotypic cell-cell contacts[201]. Although, other carcinoma cells lack or have under-sulfated HSPGs[290] and are less likely to deliver PDGF-B through cell-cell contacts. The other PDGFRβ ligand, PDGF-D lacks the retention motif that enables cellular retention through binding of cell surface HSPGs. Ectopic overexpression of PDGF-B by non-malignant human keratinocytes grafted onto mouse dermis showed an increase in extravascular PDGFRβ expression in conjunction with in-creased α-SMA expression in the same compartment[291]. Moreover, if these mice were maintained for a longer period, the keratinocytes underwent a malignant progression[292], which emphasize the role of a reactive stroma in tumor formation. The effect of PDGF-B overexpression in a conventional s.c. xenotransplanted melanoma model did also show increased stroma for-mation[293]. In contrast, reports from other tumor models have only indi-cated vascular effects of PDGF-B and -D overexpression by tumor cells[294, 295]. When regarding that the vasculature is the main site of PDGFRβ ex-pression in clinical cancers[196], it is not surprising that the most conspicu-ous effects of PDGF-B and -D overexpression are found in this compart-ment. Pericytes are a possible source of TAF progenitors, and the role of PDGF-B and -D could be to activate and expand the pericyte derived pro-genitors, which in turn can differentiate into TAFs. Even if there are tumor models that show abundant PDGFRβ expression in the stroma[183], the low expression in other models could indicate that PDGF-B and -D have less effect on already differentiated TAFs.

Also the ligands for PDGFRα, PDGF-A and -C has been shown to induce expansion of the extravascular stromal compartment of various tumors. PDGF-A expression by fibrosarcomas was found to induce a stromal re-sponse, since inhibition of PDGFRα singnaling decreased the stromal con-tent. The TAFs recruited into the tumor stroma was also responsible for pro-duction of the angiogenic growth factor VEGF-A[296]. Later, the same group showed that both PDGF-A and -C had a similar effect in lung carci-noma[297]. Further support for the role of PDGF-C in reciprocal signaling that leads to VEGF-A expression was observed in a study where TAFs iso-lated from anti-angiogenesis resistant tumors, protected tumors that were sensitive to therapy, by an increased expression of VEGF-A[298]. The sig-nificance of TAFs as VEGF-A producing cells was confirmed in a study where green fluorescence protein (GFP) was expressed under the control of the VEGF-A promotor. This study showed selective VEGF-A promotor activity in stromal TAFs[299]. In experimental cervical carcinoma, a similar signaling loop was detected between PDGF-C secreting carcinoma cells and PDGFRα expressing TAFs. This time the TAFs secreted FGF-2 and FGF-7, out of which FGF-2 turned out to be crucial for tumor angiogenesis[183].

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The role of FGF-2 was confirmed in another study in which murine mela-noma cells overexpressed PDGF-C, which induced increased TAF content in tumors. The TAFs expressed FGF-2 and the matricellular protein osteopon-tin[300]. Beside ECM production and organization, the role of TAFs seems to be relaying signals from carcinoma cells to other stromal components, and PDGF-C constitute an important messenger in this process. TAFs has also been shown to produce chemokines, which in turn can affect various cell types[301]. Mammary carcinoma cells co-inoculated with TAFs or normal mammary fibroblasts, displayed differences in tumor growth. The TAFs promoted increased tumor growth and angiogenesis, through production of the chemokine stromal derived factor 1α (SDF-1α a.k.a. CXCL12). SDF-1α stimulated carcinoma cells directly in addition to recruiting endothelial pro-genitor cells, which facilitate angiogenesis[302]. Once again, this emphasize the the importance of reciprocal signaling between malignant and stromal cells. Apart from reciprocal signaling with malignant cells, TAFs can also interact with immune cells, and primary isolates of TAFs from lung cancer displayed enhanced T-cell activation in five out of eight tested isolates. However, three isolates showed a suppressive effect[303], which once again indicates that such interactions are context dependent, especially in the tu-mor microenvironment. Genetic alterations as a consequence of being in the desmoplastic stroma has been detected in stromal cells of prostate carcino-mas[304]. This can in addition to epigenetic changes explain why isolated TAFs have other features in vitro compared to normal fibroblasts, when cul-tured under the same conditions.

The ECM scaffold in tumors Many carcinomas comprise an extensive stromal compartment and the ECM can constitute a large part of the stroma, especially in some carcinomas like breast cancer[305] and pancreas cancer[306]. The stromal compartment of pancreas cancer is very prominent at the same time as it is an aggressive and deadly form of cancer. Mostly due to its resistance to chemotherapy, which has been linked to the extensive stromal compartment. This was shown in a recent study in which an inhibitor of hedgehog signaling reduced the content of reactive stroma, which in turn increased treatment efficacy of chemother-apy[307].

The ECM of carcinomas is characterized by a collagen scaffold composed reticular and collagen fibers. The collagen fibrils from tumors often display an altered morphology compared to fibrils from normal loose connective tissue. Crossections of fibrils found in tumors are not always smooth and circular, instead they can be multilobular and coarse[308]. Moreover, colla-gen fibrils in tumors display a heterogeneous distribution in diameters, and

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often are large diameter fibrils observed, which can further assemble into larger collagen bundles[309].

A reasonable explanation for the altered collagen scaffold in tumors is that the collagen fibril assembly is dysregulated, like most other processes in tumors. TAFs most likely regulate collagen assembly in tumors and as men-tioned above they are quite different from their counterparts in normal tis-sues. Moreover, collagens, collagen modifying enzymes, SLRPs and ma-tricellular proteins are differently expressed in tumors compared to normal loose connective tissue, which also contributes to the altered collagen fibril assembly. The expression of fibrillar collagens are often up-regulated in many carcinomas[310, 311], and this is also observed for the crosslinking enzymes, LOX or LOXLs[312]. By modulating the activity of LOX or LOXLs has the effect of collagen crosslinking on tumor progression in ex-perimental tumors been exposed. Excessive activity of LOX or LOXLs in-creases the collagen scaffold stiffness, which have been shown to increase invasiveness of tumor cells in vitro and in vivo[313]. Inhibition of LOX in a transgenic mouse model of mammary carcinoma, resulted in less crosslinked collagen, prolonged tumor latency, decreased tumor incidence and vol-ume[33]. The effect of matrix stiffness on carcinoma cells was attributed to increased signaling via focal adhesions containing β1-integrins, which in turn increased cellular motility and invasiveness[33]. In a recent study, similar effects were observed when inhibiting LOX in a transgenic lung cancer model. This study also showed a correlation between LOX expression levels and survival, which were further corroborated in patient blood samples. The LOX activity in lung cancer patient serum correlated with high clinical stage and metastatic disease[314]. Recently, a seemingly very effective antibody targeting LOXL2 was presented. Inhibition of LOXL2 induced similar ef-fects as the aforementioned effects of LOX inhibition, but not only in ex-perimental tumor models, significant effects were also seen in experimental models of fibrosis. LOXL2 inhibition was also reported to inhibit TGF-β signaling, which were suggested to occur through decreased activation via αV-integrins[315]. Moreover, the fibrogenic response in tumors can elicit expression of proteins that are not normally expressed in loose connective tissue, such as fibromodulin, which normally is expressed in tendons and other tissues that are exposed for high tensional stress. Fibromodulin have been shown to have a profound effect on collagen fibril diameter[67]. Fi-bromodulin expression was lowered by an inhibitor specific for TGF-β1 and TGF-β3, which resulted in a reduced collagen scaffold density and decreased fibril diameter, suggesting that a looser ECM was formed. Tumors grown in fibromodulin KO mice confirmed these results[69]. The significance of this alteration will be discussed below.

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Many matricellular proteins are also overexpressed in carcinomas and other tumors, but the effect of these proteins on tumor collagen matrix organiza-tion is uncertain. However, both the periostin and SPARC deficient mice display smaller collagen fibril diameter and tumors grown in these mice both display less fibrosis with less collagen deposition[316, 317]. In contrast to the studies showing a pro-tumorigenic effect of a fibrotic stroma, these mod-els showed an increased tumor growth in periostin and SPARC KO mice. This could be attributed to that the peri-tumoral fibrous capsule present in tumors grown s.c. constitutes a hindrance for tumor growth. This capsule is reduced in both KOs, but it is not present in the clinical setting. Moreover, both periostin and SPARC are associated with poor prognosis in some types of cancer[318, 319].

Fluid balance in the tumor stroma Due to the impaired vasculature of tumors, an apparent lack of lymphatic vessels and an altered ECM phenotype, the fluid balance in tumors is not in balance when compared to normal loose connective tissue. Since the vascu-lature leaks plasma proteins, there is no colloid osmotic pressure difference between the interstitium and the blood, and the lack of functional lymphatics results in a lack of drainage of excess interstitial water[320]. The blood fluid pressure is often lower than in normal tissues, but it is still positive; tumor capillary fluid pressure is in the range of 7-31 (mean 17 ± 6) mmHg[321]. Altogether this would lead to a severe edema, if it was not for the fact that most solid tumors has an increased interstitial fluid pressure (IFP). This re-sults in a low transport of water and other low molecular weight molecules, since the rudimentary pressure gradient that is present in the tumor vascula-ture is not enough to drive transport by convection. Instead most transport is propelled by diffusion, which is far less efficient, especially for larger pro-teins. Further, transport of proteins in the interstitium is also hampered by the composition of the ECM. The collagen scaffolds acts as a sieve that ex-clude larger proteins. This can be demonstrated by treating tumor tissue with collagenase that leads to facilitated transport of the plasma protein albu-min[322]. The high IFP confers an obstacle for treatment of solid tumors, since transport of therapeutic substances also are limited[6]. Several strate-gies to circumvent this problem have been proposed, which have targeted both the vasculature and the interstitial stroma. Increasing central blood pressure did not have any effect, since a reciprocal increase in IFP was ob-served when blood fluid pressure was increased by administration of angio-tensin II[323]. Since interstitial fibroblasts seems to be in control of IFP in normal tissues[324], attempts were made to target these cells in tumors. Ad-ministration of prostaglandin E1 (PGE1) that regulates the actin cytoskeleton significantly lowered tumor IFP and a concomitant uptake of the chemo-therapeutic agent 5 fluorouracil (5-FU) could be observed. This lead to an increased treatment efficacy, but only during the transient period when the

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short lived PGE1 had an effect[325]. Concomitant with a decrease in IFP in most models, there is an increase in extracellular water content, which re-flects that the underhydrated ground substance is allowed to swell. Another strategy, which was believed to target collagen network contraction was administration of the PDGFR inhibitor Imatinib. This strategy was based on the observation that PDGF-BB administered during anaphylaxis normalized dermal IFP via increased contraction of the ECM by interstitial fibro-blasts[176]. Consequently, administration of Imatinib lowered tumor IFP, but the effect was first seen after three days of treatment[326], which could imply that this effect was not solely due to decreased contraction of the col-lagen scaffold by TAFs. Either way, treatment with Imatinib increased the uptake and efficacy of various chemotherapeutic agents[326-329]. The ef-fects on the TAF population were not directly assessed in these studies, but the effects on the vasculature were. The reported vascular effects comprised decreased pericyte coverage and a response that was interpreted as vascular maturation[329]. This maturation response would be in line with the notion that excess PDGF-B or -D in tumors would lead to pericyte activation and detachment. If vascular normalization was the main reason behind the effect of Imatinib on IFP is currently not known. Vascular maturation in conjunc-tion with lowering of IFP was also observed in a model where VEGF-A was inhibited with the blocking antibody bevacizumab (Avastin) [148, 330]. Also, a reduction in macrophage activation was observed in this model, which suggested that inhibition of VEGF-A induced a general lowering of the inflammatory stress. Since TGF-β signaling is highly involved in in-flammatory responses, inhibition of TGF-β signaling was also tested as a modality to lower tumor IFP. Treatment with a TGF-β inhibitor was success-ful in lowering IFP and a concomitant increase of anti-tumor treatment effi-cacy was observed[331]. This was attributed, at first, to decreased inflamma-tion only, but an additional study showed that inhibition of TGF-β signaling also reduced the expression of fibromodulin. This lead to an altered, less dense collagen scaffold, as discussed above[69]. This suggests that a dense collagen scaffold also is capable of constraining the underhydrated ground substance from taking up water. Importantly, this study pointed to a link between a fibrotic ECM structure and high IFP in tumor pathologies (Figure 1C). This link adds additional mechanistic insight to the therapy resistance seen in some tumors with a high content of reactive stroma.

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Present investigation

Aims The aim of the studies included in this thesis was to investigate various as-pects of the tumor microenvironment, with emphasis on cellular interactions, remodulation of ECM and tumor fluid balance.

Paper I Results and discussion It was previously shown that some carcinoma cells retain most of the PDGF-B they produce at their cell surface, and that they stimulate PDGFRβ bearing mesenchymal cells in a paracrine fashion that involves cell-cell contacts. Retention of PDGF-B by carcinoma cells has been attributed to the heparin binding motif of PDGF-B, which enables binding to cell surface HSPGs. In this study we dug deeper into this phenomenon and investigated if there were any difference in biological endpoints depending on the mode of ligand delivery: soluble versus membrane bound. PDGF-B added to PDGFRβ bear-ing cells induce the endpoints proliferation and actin reorganiza-tion/migration in vitro. We wanted to elucidate if the mode of ligand presen-tation could modulate the response in the recipient cell. In carcinomas, a high expression of PDGFRβ is found on pericytes[196]. We therefore used primary pericytes isolated from human skin as PDGFRβ bearing cells in this study. We found that pericytes co-cultured together with carcinoma cells were specifically directed towards proliferation, whereas pericytes in mono-culture exposed to recombinant PDGF-B exhibited both proliferation and actin reorganization. Further, we found that this response was attributed to a selective stimulation of two different PDGFRβ cohorts in pericytes. These two cohorts segregated into two different plasma membrane compartments: the lipid raft and the non-lipid raft compartments respectively. The lipid raft cohort of PDGFRβ responded specifically to soluble PDGF-B, whereas the non-lipid raft cohort responded to membrane bound ligand. There was an overlap in the stimulation with soluble PDGF-B, which in fact stimulated both cohorts, but with different phosphorylation kinetics. The lipid raft PDGFRβ cohort displayed a much faster phosphorylation peak compared to the non-lipid raft receptor cohort. The activation of both receptor cohorts might explain why both proliferation and actin reorganization could be ob-

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served with stimulation by soluble PDGF-B. The proliferative response by pericytes in co-cultures was specific to PDGF-B, since both low molecular weight PDGFR specific tyrosine kinase inhibitors and PDGF-B specific, blocking antibodies inhibited proliferation. The inhibition of PDGFR signal-ing in pericytes also revealed a reciprocal signaling loop, since also the car-cinoma cell proliferation was inhibited.

Moreover, there was no actin reorganization observed in co-cultures even after addition of exogenous soluble PDGF-B, which suggested that the car-cinoma cells not only specifically activated one receptor cohort, but also could block signaling from the other receptor cohort. Older literature de-scribed that PDGFRβ signaling from lipid rafts could be inhibited by addi-tion of IL-1β[332, 333]. We therefore applied IL-1β together with PDGF-B to pericytes in mono-culture, which resulted in decreased actin reorganiza-tion. Further, inhibition of IL-1β by a decoy receptor for IL-1 increased actin reorganization in pericyte-carcinoma cell co-cultures. These results infers that carcinoma cells can influence the response in PDGFRβ expressing cells by the mode of ligand delivery, but also by modulating intracellular signal transduction.

Caveolin-1 is a structural protein that gives rise to caveolae, which are flask shaped invaginations of the plasma membrane with a lipid composition simi-lar to lipid rafts. Immuno-histological staining of pericytes stimulated with soluble PDGF-B displayed a marked co-localization of caveolin-1 and acti-vated PDGFRβ and this indicates that they both would be present in caveo-lae. Co-localization of caveolin-1 and activated PDGFRβ was also found in an ex vivo preparation of human skin after injection of PDGF-B but not after injection of carcinoma cells. However, carcinoma cells also induced staining for activated PDGFRβ, which indicates that the carcinoma cells were capa-ble of activating PDGFR ex vivo. The ex vivo experiments confirmed the in vitro data that soluble PDGF-B primarily induced activation of PDGFRβ residing in lipid rafts/caveolae. PDGFRβ is activated in various reactive conditions in vivo, and we therefore investigated the co-localization between activated PDGFRβ and caveolin-1 in biopsies from different human pa-thologies. The results showed that there was a high degree of co-localization between activated PDGFRβ and caveolin-1 in wound healing and rheuma-toid arthritis, whereas there was less co-localization in colon adenocarci-noma. This study shows that carcinoma cells consequently induced activa-tion of PDGFRβ that did not reside in lipid rafts or co-localized with caveolin-1. Further IL-1β was identified as a modulator of the biological response induced by PDGF-B. These kinds of studies when complex culture systems are applied emphasize that all signaling in activated conditions where numerous soluble, membrane and ECM bound mediators are present are highly context dependent.

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Methodological considerations We choose not to try to isolate pericytes from tumors, since the availability of tumor specimens are limited. Further, such specimens would probably display a higher degree of heterogeneity than the skin pericytes we chose to use. Skin pericytes were obtained by isolating microvascular fragments from human foreskins, and cells from several donors were used. There were no large differences in growth or the expression levels of PDGFRβ, which sup-ports that these cells were rather homogenous.

Lipid rafts was separated from the rest of the plasma membrane by a method that was based on the principle of different in detergent solubility between these membrane domains. There are numerous recipes for such separation buffers, and the species and quantity of proteins that separates into either of the two membranes pools will depend on which recipe that is used. We used a buffer system based on the detergents Triton X-100 and octyl-glycoside. Lipid rafts were resistant to solubility by Triton X-100, but not to Triton X-100 in combination with octyl-glycoside when shear force was applied. Caveolin-1was used as a marker for quality control of the separation. In a successful separation, more than 90 % of caveolin-1 should be in the lipid raft fraction. If this was not the case, the separation was considered unsuc-cessful and data from such separations was not used.

The detection of activated PDGFRβ was based on the mouse monoclonal PDGFR-B2 antibody that only detects clustered, i.e. activated, PDGFRβ. This has previously been shown in vitro[334], but in this study we also show that this antibody works on human tissue sections.

Paper II Background In paper I, inhibition of PDGFR signaling in vitro revealed reciprocal sig-naling between PDGFRβ bearing pericytes and PDGF-B producing carci-noma cells. In paper II we applied an inhibitor of PDGFR, STI571 (Glee-vec/Imatinib) to mice bearing KAT-4 carcinomas. Earlier studies have shown that treatment of KAT-4 tumors with STI571 do not affect tumor growth, whereas tumor IFP is lowered and treatment efficacy is in-creased[327, 328]. The background and aims for this experiment was dual. First, we wanted to investigate the mechanism for how STI571 lowered IFP in tumors. The effect of STI571 was believed to be dependent on inhibition of TAF contraction of the collagen matrix. Contraction of collagen matrices reduces the propensity of the interstitium to take up fluid and swell, and this in turn increases the IFP. Emerging data have pointed to a link between high tumor IFP and a dense collagen scaffold, distinguished by a larger collagen

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fibril diameter[69]. Since the cells that are mostly responsible for the pro-duction and assembly of the ECM are mesenchymal cells expressing PDGFRs, we decided to investigate if there were any differences in the col-lagen network structure and other proteins involved in its assembly. Sec-ondly, based on the results from paper I we wanted to investigate the impact of PDGFR inhibition on the mesenchymal cell populations present in carci-nomas, TAFs and pericytes.

Results and discussion We found that treatment with STI571 induced an altered morphology and decreased diameter of collagen fibrils in KAT-4 carcinomas. Moreover, we found that STI571 also induced an increased collagen type I protein synthe-sis ex vivo. The thinner fibrils with distorted morphology combined with the increased synthesis of collagen, lead us to speculate that the collagen fibril assembly process could be affected. We therefore investigated the mRNA expression level of proteins that are involved in collagen metabolism and fibril assembly. However, there were no significant differences at the mRNA level for these genes in ST571 treated mice compared to untreated mice. The degree of inflammatory stimulation inside a tumor can also affect the collagen scaffold and IFP[69], but we found no such differences. In conclu-sion, STI571 treatment altered the collagen fibril diameter and morphology in KAT-4 carcinoma, and earlier studies has shown that STI571 lowers IFP in tumors. This infers that there could be a link between IFP and collagen fibril diameter also in this model. However, we could not elucidate any de-tailed mechanism for how this was achieved. The increased collagen protein synthesis provided a clue that deregulation of collagen fibril assembly could be responsible. Potentially, more collagen protein was produced but less was build into collagen fibrils.

The second aim was to assess the effect of PDGFR inhibition on the mesen-chymal cells in the tumor microenvironment. We found that STI571 treat-ment had no effect on the level of staining for vascular structures and the coverage of such structures by pericytes. However, we detected a change in the amount of extravascular cells that stained positive for the pericyte marker NG2. This population of cells could be pericytes that are just located further away from the vasculature or pericytes that are in transition towards differ-entiation into fibroblasts, which pericytes has been shown to do in vitro [143]. There was no change in the populations of cells that stained positive for α-SMA and reticular fibroblast marker (RFM), which both stain fibro-blasts and pericytes. These results imply that the NG2 positive cells that were not in close contact with the vascular endothelium were more vulner-able to PDGFR inhibition, which could suggests that close contacts with the endothelium is protective for pericytes. If the NG2 positive cells constituted precursors for TAFs in KAT-4 carcinomas, the staining for α-SMA and

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RFM should also decrease over time, if there is a turnover in these popula-tions. A longer treatment period could perhaps shed light on this issue. An-other theory is that STI571 treatment induced differentiation of pericytes that made them lose NG2 expression.

Methodological considerations Recently, questions has been raised over the true origin of the KAT-4 cells, which were first annotated as anaplastic thyroid carcinoma cells[335], but now, indications exist that the KAT-4 cells actually are HT-29 human colon carcinoma[336]. Either way, this has only limited consequences of the ex-periments undertaken in this study, since we make no correlation with other studies comprising experimental or clinical material from either proposed origin.

The KAT-4 carcinomas were inoculated in severe combined immuno-deficient (SCID) mice that lack both functional T and B-cells. This infers that caution has to be applied when interpreting results referring to adaptive immune responses. As mentioned in the introduction, T-cells has a role in mediating inflammatory signaling that affects fibrosis. However, the tumor microenvironment in these mice are populated by innate immune cells and TAFs, which provides a microenvironment that is fairly reminiscent of the clinical situation, but all speculations of immunological responses directed against tumors cells has to be abolished.

Collagen synthesis ex vivo were performed for 6 hours, longer incubations provided inconsistent results that may depend on the high metabolic rate in tumor tissues. Longer ex vivo incubations probably lead to more unregulated protein turnover, i.e. artifacts.

Paper III Background The IFP of loose connective tissues is reduced during acute inflammations allowing for edema to form. The IFP can be restored by administration of PDGF-B, which involves a mechanism that requires integrin αVβ3. This ef-fect was proved in experiments with antibodies that block the function of the integrin subunit β3 or in mice that lack a functional β3 gene[176]. PDGF-B has no effect on IFP in settings where the function of integrin αVβ3 is per-turbed. In contrast to mice that lack the β3 gene, mice deficient in the αV gene are not viable. Integrin β3 deficient mice have a mild phenotype that consists of an impaired blood clotting, due to the lack of the integrin αIIbβ3 in platelets, and osteopetrosis due to lack of αVβ3. High IFP in solid tumors impairs therapeutic efficacy, since it limits the penetration of chemothera-

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peutic agents into the tumor tissue. Since the integrin αVβ3 is expressed in many solid tumors, we undertook an investigation in order to elucidate if αVβ3 also regulates IFP in tumors.

Results and discussion We implanted syngeneic CT26 colon and LM3 mammary carcinomas into WT and integrin β3-deficient mice. We found that deficiency in integrin β3 did not affect tumor growth or the density and function of tumor blood ves-sels. However, we did find that integrin β3-deficient mice had a significantly higher tumor IFP, which was surprising when considering the role of this integrin in acute inflammations. We found a plausible explanation for the high tumor IFP when we examined the ECM scaffold in tumors that were grown in integrin β3 deficient mice. These mice had a denser collagen matrix and the collagen fibrils were also thicker than the WT counterparts. Thicker collagen fibrils and a dense collagen matrix have previously been shown to correlate with high tumor IFP in the KAT-4 model. In this model, treatment with an inhibitor of TGF-β lowered inflammatory parameters, tumor IFP and decreased the thickness of collagen fibrils[69]. In contrast to the previous study, we observed no increase in inflammatory parameters in conjunction with the increased IFP and collagen fibril thickness. Instead, there was a significantly lower content of macrophages and a decreased level of inflam-matory transcripts. This suggests that the mechanism for increasing IFP and collagen thickness was not dependent on inflammation in the integrin β3-deficient mice. Earlier studies of wound healing in mice deficient in integrin β3 has shown that macrophage content in granulation tissue is de-creased[337], which suggests that macrophage infiltration could be partly regulated by integrin β3. Our study adds data that support a link between increased tumor IFP and a fibrotic ECM phenotype in tumors.

Methodological considerations When studying the tumor stroma, a model with a substantial stromal com-partment is desired, but these are hard to find and especially syngeneic tumor models have a limited stromal compartment compared to clinical carcino-mas. However, electron microscopy revealed that the investigated syngeneic carcinomas had an extensive collagen fiber network, which gives legitimacy to the use of these models for studying the tumor stroma.

The electron microscopy technique applied in this study enables the selective visualization of collagen fibrils, all other matter is dissolved by alkali treat-ment. This means that the only fibrils that are visual on the images obtained from the tumors constitute collagen fibers and reticular fibers.

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Paper IV Background PDGF-B and TGF-β constitutes two important growth factors that affect the stromal compartment of solid tumors and other inflammatory pathologies. In this study we investigated the impact of forced expression of these growth factors in mouse skin of the ears. This model is artificial but offers an oppor-tunity to study the role of each growth factor separately. Our primary aim was to investigate if these factors elicited an activation of connective tissue cells and if questions regarding their origin and fate could be answered. Fur-ther, we wanted to investigate if the different growth factors induced activa-tion in separate types of connective tissue cells or if there was any differ-ences in the resulting phenotype of these cells. We also wanted to investigate how these cells contributed to a possible fibrosis.

Results and discussion Adenoviruses that either expressed PDGF-B (AdPDGF-B) or an active re-combinant variant of TGF-β1 (Ad TGF-β1) were used. An adenovirus en-coding the gene for green fluorescent protein (AdGFP) was used as a con-trol. Both AdPDGF-B and AdTGF-β1 evoked a tissue response consisting of vascular activation, expansion of connective tissue cells and infiltration of macrophages. No such responses were observed in the AdGFP treated skin. The vascular responses consisted of detachment of mural pericytes and ves-sel dilation. These activated vessels showed resemblance to mother vessels that has been reported to be initiation points for angiogenesis[226]. How-ever, no increase in blood vessels was detected in either of the treatments. Although AdPDGF-B transiently induced vascular malformations that were reminiscent of glomeruloid bodies, which are microvascular proliferations found in certain pathologies such as cancers. Over time these structures dis-appeared. Subsequently to pericyte detachment could an increase of perivas-cular connective tissue cells be observed and after 7 days there was an in-crease in cellularity throughout the dermis of the affected area. The cells that constituted the hypercellular dermis were connective tissue cells and macro-phages, which stained positively for reticular fibroblast marker and F4/80 respectively. These cells did not just populate the dermis but also the sub-dermal muscular layer, which resulted in altered tissue architecture in that area. There was a marked increase in dermal thickness concomitant with the increase in cellularity. As our aim was to elucidate the origin and phenotype of the emerging connective tissue cell population, we stained the tissue for different markers of pericytes, fibroblasts and myofibroblasts/ smooth mus-cle cells. Skin treated with AdPDGF-B showed a marker profile that sug-gested a pericyte origin of the connective tissue cells since they had a high expression of the pericyte marker NG2. AdTGF-β1 induced expansion of connective tissue that displayed less NG2 staining, but more staining with

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the myofibroblast marker α-SMA. This is consistent with previous studies showing that TGF-β is an inducer of fibroblast to myofibroblast differentia-tion[254]. Fewer cells displayed α-SMA staining in AdPDGF-B treated skin. At the last time point, 14 days, could a small difference in tissue histology be seen. The involvement of the muscular layer had decreased in AdPDGF-B treated skin, whereas in AdTGF-β1 treated skin, this area was still populated by an excess of connective tissue cells and macrophages. This suggested that the reactive response could be subsiding in AdPDGF-B treated skin. More-over, we also investigated the mRNA expression at day 7 of a number of ECM-related transcripts and these showed a marked difference in expression levels depending on the treatment. AdTGF-β1 treated skin displayed a high expression of most ECM related transcripts, such as fibrillar collagens, col-lagen modifying enzymes, SLRPs, matricellular proteins and fibronectin. In contrast, no fibrillar collagens or collagen crosslinking enzymes were up-regulated by AdPDGF-B. However, AdPDGF-B increased expression of a few ECM-related genes that are associated with granulation tissue. This sug-gests that the ECM phenotype induced by AdPDGF-B was more immature, not involving formation of dense fibrillar structures like collagen fibers.

Methodological considerations Adenoviruses are powerful expression vectors that can be administrated to a number of tissues. They create a transient response that allows for temporal studies. However, they are also immunogenic and this could create difficul-ties in interpreting the results, especially when inflammatory processes are being studied. This was circumvented in the present study by using athymic mice that lack functional T-cells. However, this creates another drawback when studying inflammatory and fibrotic conditions, since T-cells also are involved in the tuning of inflammatory responses, even if there is no activa-tion by antigens.

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Future perspectives

Since treatment specifically targeting the malignant cells in carcinoma is currently not available, other strategies has to be explored. The tumor stroma constitutes an attractive target because of the additional treatment modalities it confers. Much additional work in this field is at hand.

Additional experiments regarding paper I could clarify if the two different cohorts also are unequally sensitive to different concentrations of PDGF-B. Indications for this notion was found in paper I, where different endpoints selectively were induced depending on the concentration of added PDGF-B. Moreover, it would be interesting to investigate which kind of intracellular signal transduction response that is elicited from each receptor cohort and how IL-1 induced signal transduction interferes. This task will be hard when regarding that the co-cultures consist of two cell types that both express the same signal transduction molecules. However, membrane inserts that sepa-rates the two celltypes but still allows cell-cell contacts trough pores might be an alternative. Another strategy could be to use magnetic bead separation technique already employed in paper I. Since this separation takes quite some time, the risk of introducing artifacts is increased. In situ techniques could be a third option, the challenge would be to achieve enough spatial resolution in the co-cultures that would certify that only one and the same cell is examined. The reciprocal signaling loop between carcinoma cells and pericytes may confer a protection from cell death for carcinoma cells. We have already begun to test this hypothesis by studying therapeutic efficacy in co-cultures. Preliminary data suggests that mesenchymal cells confer some protection.

Paper II revealed a difference in the collagen scaffold between tumor-bearing mice treated with STI571 or vehicle. The mechanism for this effect should be elucidated and a plausible approach is to further investigate the role of SLRPs and matricellular proteins. So far has no explanation for this difference been found on the transcriptional level. However, evidence for a differently regulated expression of collagen I was found on the protein level. Thus, further studies should focus on investigating differences on the protein level in STI571 treated tumors. It would be interesting to analyze the amount denatured collagen, which will not be incorporated into fibrils and fibers. This will clarify if the collagen that is produced in STI571 treated tumors is

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incorporated into fibrils. The extent of crosslinking would also be interesting to investigate, since this also contributes to collagen scaffold quality. The challenge here would be to acquire enough material in order to run this type of analysis. Currently, efforts are being made to optimize mass spectrometry methods in order to study collagen crosslinks, which would profoundly fa-cilitate this kind of analyses. Longer treatment periods would also be inter-esting to perform, both regarding the ECM phenotype and the fate of the NG2 positive stromal cells.

It would be interesting to treat tumor bearing integrin β3 deficient mice with agents with known effects on tumor IFP. One such agent could be STI571, but also prostaglandin E1 and TGF-β inhibitors could be used. This would presumably aid in the elucidation of the mechanism behind the effects seen in this system, when regarding the ECM phenotype and IFP. Our laboratory has found a bacterial peptide that inhibits integrin αVβ3 mediated collagen contraction, and this peptide would be interesting to apply in tumor experi-ments.

The extent of fibrosis needs to be determined in paper IV. The affected areas in the mouse ears are to small for a quantitative protein analysis, with classi-cal methods. Thus, he collagen content is hard to assess and use as an evi-dence for fibrosis. We can still use other markers for fibrosis, and we have already observed some potential markers on the mRNA level, e.g. pro-collagens, biglycan and fibromodulin. We can also employ electron micros-copy in order to evaluate the collagen scaffold and collagen fibril diameter. We would presumably also need to obtain observations from more distant timepoints, in order to investigate a possible resolution of the lesions, or if these lesions becomes chronic. When regarding the connective tissue cells, their expansion requires proliferation and it would be interesting to investi-gate where this proliferation takes place in situ, perivascular or deeper into the interstitium. A perivascular proliferation would support that these cells originate from the vascular wall.

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Acknowledgements

I would like to thank my supervisors Christian Sundberg and Kristofer Rubin for passing some of your knowledge and insights of the biomedical universe on to me. I will do my best to make use of it all.

I would also like to thank my examiner Staffan Johansson for help, fruitful discussions and a good colaboration in teaching the integrin lab. In addition, I would like to thank you for providing sources of energy for hungry stu-dents outside your office.

Further, I would like to thank the co-authors of my manuscripts: Renata Gus-tafsson, Alejandro Rodriguez, Carina Hellberg, Mikahail Burmakin, Marcin Kowanetz, Tijs van Wieringen, Jean Chidiac, Rolf Reed, Linda Stuhr, Åke Oldberg, Nils-Erik Heldin, Carl-Henrik Heldin, Keith Solomon, Lea Hecht and Christine Kuhl. I would like thank my colleagues in the Sundberg group: Jakob; the Rubin group: Vahid, Lena and Marcus; the Olsson group: Anna-Karin, Else, Julia, Jessica and Maria; the Åbrink group: Ananya and Magnus; Johansson group: Anjum, Birgitta, Kathrin and Rajesh. I would like to thank Joachim Gullbo, Clinical Pharmacology at UAS.

I will also give a thank you to all present and past colleagues at IMBIM, and a special thanks to Marissa, Åsa Lidén, Ann-Marie Gustafsson, Mårten Lars-son, Anh-Tri Do, Jimmy Larsson, Chi, Per Jemth, Kajsa, Lena Möller, Johan Heldin, Johnny Wernersson, Dorothe Spillman, Jens, Olav Nordli, Ted Pers-son and the virus girls: Monika, Anna, Heidi, Ellenor, Cecilia and Sara.

The DIA lab with all of its members: Anna, Linus, Jocke, Peter, Maria, Chris, Marlene, Erik and Lisa.

Mina kära vänner från biomedicinarprogrammet: Micke och Peter, ett stort tack skall ni ha för er vänskap.

Tack till Övergrans skjutvapens och rusdryckssällskap/ c.o. Bålstamilisen: Jocke, Nicke, Jocke och Peter. Tack för en livslång vänskap.

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Jag vill även tacka kamraterna i Alpha Sierra, att vara bland er är som bal-sam för själen. Jag vill även tacka krutgubben och livscoachen Erik U.M. Klintskog, du är en kär vän.

Ett stort tack till familjen Koskiniemi, Sundsvalls-filialen: Marrku och Si-nikka. Umeå-filialen: Koskiniemi-Niemi, Sattu, Fredrik, Wilma och Ebba. Bålsta-filialen (ej att förväxla med Bålstamilisen): Matti, Janette och min gudson Jimi. Det är alltid lika trevligt att träffa er.

Ett stort varmt tack till mina föräldrar, Ulla-Britt och Börje för allt stöd, kär-lek och hänsyn. Ni är bäst. Tack också till mormor Eva och farmor Lisa.

Mest av allt vill jag tacka navet som min värld kretsar kring, min älskade Sanna. Tack för att du lyser upp mitt liv, jag älskar dig.

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References

1. Liotta, L.A. and E.C. Kohn, The microenvironment of the tumour-host interface. Nature, 2001. 411(6835): p. 375-9.

2. Coussens, L.M. and Z. Werb, Inflammation and cancer. Nature, 2002. 420(6917): p. 860-7.

3. Flavell, R.A., et al., The polarization of immune cells in the tumour envi-ronment by TGFbeta. Nat Rev Immunol. 10(8): p. 554-67.

4. Hanahan, D. and R.A. Weinberg, The hallmarks of cancer. Cell, 2000. 100(1): p. 57-70.

5. Jain, R.K., Normalization of tumor vasculature: an emerging concept in antiangiogenic therapy. Science, 2005. 307(5706): p. 58-62.

6. Heldin, C.H., et al., High interstitial fluid pressure - an obstacle in cancer therapy. Nat Rev Cancer, 2004. 4(10): p. 806-13.

7. Tredan, O., et al., Drug resistance and the solid tumor microenvironment. J Natl Cancer Inst, 2007. 99(19): p. 1441-54.

8. Veit, G., et al., Collagen XXVIII, a novel von Willebrand factor A domain-containing protein with many imperfections in the collagenous domain. J Biol Chem, 2006. 281(6): p. 3494-504.

9. Brinckmann, J., Collagens at a Glance. Top Curr Chem, 2005. 247: p. 1-6. 10. Sylvie Ricard-Blum, F.R., Michel van der Rest., The Collagen Superfamily.

Top Curr Chem, 2005. 247: p. 35-85. 11. Prockop, D.J. and K.I. Kivirikko, Collagens: molecular biology, diseases,

and potentials for therapy. Annu Rev Biochem, 1995. 64: p. 403-34. 12. Myllyharju, J., Intracellular Post-Translational Modifications of Colla-

gens. Top Curr Chem, 2005. 247: p. 115-147. 13. Cabral, W.A., et al., Prolyl 3-hydroxylase 1 deficiency causes a recessive

metabolic bone disorder resembling lethal/severe osteogenesis imperfecta. Nat Genet, 2007. 39(3): p. 359-65.

14. Vranka, J.A., L.Y. Sakai, and H.P. Bachinger, Prolyl 3-hydroxylase 1, enzyme characterization and identification of a novel family of enzymes. J Biol Chem, 2004. 279(22): p. 23615-21.

15. Bonfanti, L., et al., Procollagen traverses the Golgi stack without leaving the lumen of cisternae: evidence for cisternal maturation. Cell, 1998. 95(7): p. 993-1003.

16. Kessler, E., et al., Bone morphogenetic protein-1: the type I procollagen C-proteinase. Science, 1996. 271(5247): p. 360-2.

17. Kuno, K., et al., Molecular cloning of a gene encoding a new type of metal-loproteinase-disintegrin family protein with thrombospondin motifs as an inflammation associated gene. J Biol Chem, 1997. 272(1): p. 556-62.

72

18. Colige, A., et al., Human Ehlers-Danlos syndrome type VII C and bovine dermatosparaxis are caused by mutations in the procollagen I N-proteinase gene. Am J Hum Genet, 1999. 65(2): p. 308-17.

19. Canty, E.G., et al., Coalignment of plasma membrane channels and protru-sions (fibripositors) specifies the parallelism of tendon. J Cell Biol, 2004. 165(4): p. 553-63.

20. Birk, D.E., Type V collagen: heterotypic type I/V collagen interactions in the regulation of fibril assembly. Micron, 2001. 32(3): p. 223-37.

21. Spiess, K. and T.M. Zorn, Collagen types I, III, and V constitute the thick collagen fibrils of the mouse decidua. Microsc Res Tech, 2007. 70(1): p. 18-25.

22. Liu, X., et al., Type III collagen is crucial for collagen I fibrillogenesis and for normal cardiovascular development. Proc Natl Acad Sci U S A, 1997. 94(5): p. 1852-6.

23. Brodsky, B., et al., Variations in collagen fibril structure in tendons. Bio-polymers, 1982. 21(5): p. 935-51.

24. Chakravarti, S., et al., Lumican regulates collagen fibril assembly: skin fragility and corneal opacity in the absence of lumican. J Cell Biol, 1998. 141(5): p. 1277-86.

25. Silver, F.H., J.W. Freeman, and G.P. Seehra, Collagen self-assembly and the development of tendon mechanical properties. J Biomech, 2003. 36(10): p. 1529-53.

26. Henkel, W. and R.W. Glanville, Covalent crosslinking between molecules of type I and type III collagen. The involvement of the N-terminal, nonheli-cal regions of the alpha 1 (I) and alpha 1 (III) chains in the formation of in-termolecular crosslinks. Eur J Biochem, 1982. 122(1): p. 205-13.

27. Lopez, B., et al., Role of lysyl oxidase in myocardial fibrosis: from basic science to clinical aspects. Am J Physiol Heart Circ Physiol. 299(1): p. H1-9.

28. Uzel, M.I., et al., Multiple bone morphogenetic protein 1-related mammal-ian metalloproteinases process pro-lysyl oxidase at the correct physiologi-cal site and control lysyl oxidase activation in mouse embryo fibroblast cul-tures. J Biol Chem, 2001. 276(25): p. 22537-43.

29. Prockop, D.J., A.L. Sieron, and S.W. Li, Procollagen N-proteinase and procollagen C-proteinase. Two unusual metalloproteinases that are essen-tial for procollagen processing probably have important roles in develop-ment and cell signaling. Matrix Biol, 1998. 16(7): p. 399-408.

30. Bailey, A.J., R.G. Paul, and L. Knott, Mechanisms of maturation and age-ing of collagen. Mech Ageing Dev, 1998. 106(1-2): p. 1-56.

31. Moriguchi, T. and D. Fujimoto, Crosslink of collagen in hypertrophic scar. J Invest Dermatol, 1979. 72(3): p. 143-5.

32. Bailey, A.J., et al., Characterization of the collagen of human hypertrophic and normal scars. Biochim Biophys Acta, 1975. 405(2): p. 412-21.

33. Levental, K.R., et al., Matrix crosslinking forces tumor progression by enhancing integrin signaling. Cell, 2009. 139(5): p. 891-906.

73

34. Sady, C., S. Khosrof, and R. Nagaraj, Advanced Maillard reaction and crosslinking of corneal collagen in diabetes. Biochem Biophys Res Com-mun, 1995. 214(3): p. 793-7.

35. Slatter, D.A., et al., Reactions of lipid-derived malondialdehyde with colla-gen. J Biol Chem, 1999. 274(28): p. 19661-9.

36. Fleischmajer, R., et al., Immunoelectron microscopy of type III collagen in normal and scleroderma skin. J Invest Dermatol, 1980. 75(2): p. 189-91.

37. Sykes, B., et al., The estimation of two collagens from human dermis by interrupted gel electrophoresis. Biochem Biophys Res Commun, 1976. 72(4): p. 1472-80.

38. Unsworth, D.J., et al., Studies on reticulin. I: Serological and immunohis-tological investigations of the occurrence of collagen type III, fibronectin and the non-collagenous glycoprotein of Pras and Glynn in reticulin. Br J Exp Pathol, 1982. 63(2): p. 154-66.

39. Montes, G.S., et al., Histochemical and morphological characterization of reticular fibers. Histochemistry, 1980. 65(2): p. 131-41.

40. Ushiki, T., Collagen fibers, reticular fibers and elastic fibers. A compre-hensive understanding from a morphological viewpoint. Arch Histol Cytol, 2002. 65(2): p. 109-26.

41. Wagenseil, J.E. and R.P. Mecham, New insights into elastic fiber assembly. Birth Defects Res C Embryo Today, 2007. 81(4): p. 229-40.

42. Hynes, R.O., The extracellular matrix: not just pretty fibrils. Science, 2009. 326(5957): p. 1216-9.

43. Bax, D.V., et al., Cell adhesion to fibrillin-1 molecules and microfibrils is mediated by alpha 5 beta 1 and alpha v beta 3 integrins. J Biol Chem, 2003. 278(36): p. 34605-16.

44. McLaughlin, P.J., et al., Targeted disruption of fibulin-4 abolishes elasto-genesis and causes perinatal lethality in mice. Mol Cell Biol, 2006. 26(5): p. 1700-9.

45. Kozel, B.A., et al., Elastic fiber formation: a dynamic view of extracellular matrix assembly using timer reporters. J Cell Physiol, 2006. 207(1): p. 87-96.

46. Reinboth, B., et al., Molecular interactions of biglycan and decorin with elastic fiber components: biglycan forms a ternary complex with tropoe-lastin and microfibril-associated glycoprotein 1. J Biol Chem, 2002. 277(6): p. 3950-7.

47. Font, B., et al., Characterization of the interactions of type XII collagen with two small proteoglycans from fetal bovine tendon, decorin and fibro-modulin. Matrix Biol, 1996. 15(5): p. 341-8.

48. Font, B., et al., Binding of collagen XIV with the dermatan sulfate side chain of decorin. J Biol Chem, 1993. 268(33): p. 25015-8.

49. Yamaguchi, Y., D.M. Mann, and E. Ruoslahti, Negative regulation of transforming growth factor-beta by the proteoglycan decorin. Nature, 1990. 346(6281): p. 281-4.

50. Hildebrand, A., et al., Interaction of the small interstitial proteoglycans biglycan, decorin and fibromodulin with transforming growth factor beta. Biochem J, 1994. 302 ( Pt 2): p. 527-34.

74

51. Zhu, J.X., et al., Decorin evokes protracted internalization and degradation of the epidermal growth factor receptor via caveolar endocytosis. J Biol Chem, 2005. 280(37): p. 32468-79.

52. Schonherr, E., et al., Decorin, a novel player in the insulin-like growth factor system. J Biol Chem, 2005. 280(16): p. 15767-72.

53. Schaefer, L., et al., The matrix component biglycan is proinflammatory and signals through Toll-like receptors 4 and 2 in macrophages. J Clin Invest, 2005. 115(8): p. 2223-33.

54. Kalamajski, S., et al., Asporin competes with decorin for collagen binding, binds calcium and promotes osteoblast collagen mineralization. Biochem J, 2009. 423(1): p. 53-9.

55. Tse, G.M., et al., Calcification in breast lesions: pathologists' perspective. J Clin Pathol, 2008. 61(2): p. 145-51.

56. Kalamajski, S. and A. Oldberg, The role of small leucine-rich proteogly-cans in collagen fibrillogenesis. Matrix Biol.

57. Reed, C.C. and R.V. Iozzo, The role of decorin in collagen fibrillogenesis and skin homeostasis. Glycoconj J, 2002. 19(4-5): p. 249-55.

58. Xu, T., et al., Targeted disruption of the biglycan gene leads to an osteopo-rosis-like phenotype in mice. Nat Genet, 1998. 20(1): p. 78-82.

59. Corsi, A., et al., Phenotypic effects of biglycan deficiency are linked to collagen fibril abnormalities, are synergized by decorin deficiency, and mimic Ehlers-Danlos-like changes in bone and other connective tissues. J Bone Miner Res, 2002. 17(7): p. 1180-9.

60. Zhang, G., et al., Genetic evidence for the coordinated regulation of colla-gen fibrillogenesis in the cornea by decorin and biglycan. J Biol Chem, 2009. 284(13): p. 8888-97.

61. Westergren-Thorsson, G., et al., Altered expression of small proteoglycans, collagen, and transforming growth factor-beta 1 in developing bleomycin-induced pulmonary fibrosis in rats. J Clin Invest, 1993. 92(2): p. 632-7.

62. Krull, N.B., T. Zimmermann, and A.M. Gressner, Spatial and temporal patterns of gene expression for the proteoglycans biglycan and decorin and for transforming growth factor-beta 1 revealed by in situ hybridization dur-ing experimentally induced liver fibrosis in the rat. Hepatology, 1993. 18(3): p. 581-9.

63. Hunzelmann, N., et al., Co-ordinate induction of collagen type I and bigly-can expression in keloids. Br J Dermatol, 1996. 135(3): p. 394-9.

64. Gambichler, T., et al., Differential expression of decorin in localized scle-roderma following ultraviolet-A1 irradiation. J Am Acad Dermatol, 2007. 56(6): p. 956-9.

65. Kalamajski, S. and A. Oldberg, Fibromodulin binds collagen type I via Glu-353 and Lys-355 in leucine-rich repeat 11. J Biol Chem, 2007. 282(37): p. 26740-5.

66. Kalamajski, S. and A. Oldberg, Homologous sequence in lumican and fibromodulin leucine-rich repeat 5-7 competes for collagen binding. J Biol Chem, 2009. 284(1): p. 534-9.

75

67. Svensson, L., et al., Fibromodulin-null mice have abnormal collagen fi-brils, tissue organization, and altered lumican deposition in tendon. J Biol Chem, 1999. 274(14): p. 9636-47.

68. Venkatesan, N., et al., Alterations in large and small proteoglycans in bleomycin-induced pulmonary fibrosis in rats. Am J Respir Crit Care Med, 2000. 161(6): p. 2066-73.

69. Oldberg, A., et al., Collagen-binding proteoglycan fibromodulin can de-termine stroma matrix structure and fluid balance in experimental carci-noma. Proc Natl Acad Sci U S A, 2007. 104(35): p. 13966-71.

70. Jepsen, K.J., et al., A syndrome of joint laxity and impaired tendon integrity in lumican- and fibromodulin-deficient mice. J Biol Chem, 2002. 277(38): p. 35532-40.

71. Ezura, Y., et al., Differential expression of lumican and fibromodulin regu-late collagen fibrillogenesis in developing mouse tendons. J Cell Biol, 2000. 151(4): p. 779-88.

72. Hynes, R.O., Fibronectins. New York: Springer-Verlag, 1990. 73. Guan, J.L., J.E. Trevithick, and R.O. Hynes, Retroviral expression of alter-

natively spliced forms of rat fibronectin. J Cell Biol, 1990. 110(3): p. 833-47.

74. Mosesson, M.W. and R.A. Umfleet, The cold-insoluble globulin of human plasma. I. Purification, primary characterization, and relationship to fi-brinogen and other cold-insoluble fraction components. J Biol Chem, 1970. 245(21): p. 5728-36.

75. Matsuda, M., et al., Distribution of cold-insoluble globulin in plasma and tissues. Ann N Y Acad Sci, 1978. 312: p. 74-92.

76. Johansson, S., et al., In vitro biosynthesis of cold insoluble globulin (fi-bronectin) by mouse peritoneal macrophages. FEBS Lett, 1979. 105(2): p. 313-6.

77. Kramer, R.H., et al., Synthesis of extracellular matrix glycoproteins by cultured microvascular endothelial cells isolated from the dermis of neona-tal and adult skin. J Cell Physiol, 1985. 123(1): p. 1-9.

78. Velling, T., et al., Polymerization of type I and III collagens is dependent on fibronectin and enhanced by integrins alpha 11beta 1 and alpha 2beta 1. J Biol Chem, 2002. 277(40): p. 37377-81.

79. Couchman, J.R., et al., Fibronectin distribution in epithelial and associated tissues of the rat. Arch Dermatol Res, 1979. 266(3): p. 295-310.

80. Engvall, E., E. Ruoslahti, and E.J. Miller, Affinity of fibronectin to colla-gens of different genetic types and to fibrinogen. J Exp Med, 1978. 147(6): p. 1584-95.

81. Erickson, H.P., Stretching fibronectin. J Muscle Res Cell Motil, 2002. 23(5-6): p. 575-80.

82. Comper, W.D. and T.C. Laurent, Physiological function of connective tissue polysaccharides. Physiol Rev, 1978. 58(1): p. 255-315.

83. Aukland, K. and R.K. Reed, Interstitial-lymphatic mechanisms in the con-trol of extracellular fluid volume. Physiol Rev, 1993. 73(1): p. 1-78.

76

84. Granger, H.J., S.H. Laine, and G.A. Laine, Osmotic pressure exerted by entangled polysaccharide chains. Microcirc Endothelium Lymphatics, 1985. 2(1): p. 85-105.

85. Leask, A. and D.J. Abraham, All in the CCN family: essential matricellular signaling modulators emerge from the bunker. J Cell Sci, 2006. 119(Pt 23): p. 4803-10.

86. Gillan, L., et al., Periostin secreted by epithelial ovarian carcinoma is a ligand for alpha(V)beta(3) and alpha(V)beta(5) integrins and promotes cell motility. Cancer Res, 2002. 62(18): p. 5358-64.

87. Murphy-Ullrich, J.E., The de-adhesive activity of matricellular proteins: is intermediate cell adhesion an adaptive state? J Clin Invest, 2001. 107(7): p. 785-90.

88. Chung, C.Y., J.E. Murphy-Ullrich, and H.P. Erickson, Mitogenesis, cell migration, and loss of focal adhesions induced by tenascin-C interacting with its cell surface receptor, annexin II. Mol Biol Cell, 1996. 7(6): p. 883-92.

89. Liu, Y.K., et al., Osteopontin involvement in integrin-mediated cell signal-ing and regulation of expression of alkaline phosphatase during early dif-ferentiation of UMR cells. FEBS Lett, 1997. 420(1): p. 112-6.

90. Chellaiah, M.A., et al., Rho-A is critical for osteoclast podosome organiza-tion, motility, and bone resorption. J Biol Chem, 2000. 275(16): p. 11993-2002.

91. Shimazaki, M., et al., Periostin is essential for cardiac healing after acute myocardial infarction. J Exp Med, 2008. 205(2): p. 295-303.

92. Grotendorst, G.R. and M.R. Duncan, Individual domains of connective tissue growth factor regulate fibroblast proliferation and myofibroblast dif-ferentiation. Faseb J, 2005. 19(7): p. 729-38.

93. Tuck, A.B., et al., Osteopontin-induced migration of human mammary epithelial cells involves activation of EGF receptor and multiple signal transduction pathways. Oncogene, 2003. 22(8): p. 1198-205.

94. Raines, E.W., et al., The extracellular glycoprotein SPARC interacts with platelet-derived growth factor (PDGF)-AB and -BB and inhibits the bind-ing of PDGF to its receptors. Proc Natl Acad Sci U S A, 1992. 89(4): p. 1281-5.

95. Bornstein, P., Thrombospondins function as regulators of angiogenesis. J Cell Commun Signal, 2009. 3(3-4): p. 189-200.

96. Francki, A., et al., SPARC regulates TGF-beta1-dependent signaling in primary glomerular mesangial cells. J Cell Biochem, 2004. 91(5): p. 915-25.

97. Murphy-Ullrich, J.E. and M. Poczatek, Activation of latent TGF-beta by thrombospondin-1: mechanisms and physiology. Cytokine Growth Factor Rev, 2000. 11(1-2): p. 59-69.

98. Norris, R.A., et al., Periostin regulates collagen fibrillogenesis and the biomechanical properties of connective tissues. J Cell Biochem, 2007. 101(3): p. 695-711.

99. Liaw, L., et al., Altered wound healing in mice lacking a functional osteo-pontin gene (spp1). J Clin Invest, 1998. 101(7): p. 1468-78.

77

100. Bradshaw, A.D., et al., SPARC-null mice display abnormalities in the der-mis characterized by decreased collagen fibril diameter and reduced ten-sile strength. J Invest Dermatol, 2003. 120(6): p. 949-55.

101. Kyriakides, T.R., et al., Mice that lack thrombospondin 2 display connec-tive tissue abnormalities that are associated with disordered collagen fi-brillogenesis, an increased vascular density, and a bleeding diathesis. J Cell Biol, 1998. 140(2): p. 419-30.

102. Furuya, S. and K. Furuya, Subepithelial fibroblasts in intestinal villi: roles in intercellular communication. Int Rev Cytol, 2007. 264: p. 165-223.

103. Langevin, H.M., C.J. Cornbrooks, and D.J. Taatjes, Fibroblasts form a body-wide cellular network. Histochem Cell Biol, 2004. 122(1): p. 7-15.

104. Stopak, D. and A.K. Harris, Connective tissue morphogenesis by fibroblast traction. I. Tissue culture observations. Dev Biol, 1982. 90(2): p. 383-98.

105. Meshel, A.S., et al., Basic mechanism of three-dimensional collagen fibre transport by fibroblasts. Nat Cell Biol, 2005. 7(2): p. 157-64.

106. Kawai, T. and S. Akira, The role of pattern-recognition receptors in innate immunity: update on Toll-like receptors. Nat Immunol. 11(5): p. 373-84.

107. Franchi, L., et al., The inflammasome: a caspase-1-activation platform that regulates immune responses and disease pathogenesis. Nat Immunol, 2009. 10(3): p. 241-7.

108. Hu, X., S.D. Chakravarty, and L.B. Ivashkiv, Regulation of interferon and Toll-like receptor signaling during macrophage activation by opposing feedforward and feedback inhibition mechanisms. Immunol Rev, 2008. 226: p. 41-56.

109. Van Ginderachter, J.A., et al., Classical and alternative activation of mo-nonuclear phagocytes: picking the best of both worlds for tumor promotion. Immunobiology, 2006. 211(6-8): p. 487-501.

110. Poon, I.K., M.D. Hulett, and C.R. Parish, Molecular mechanisms of late apoptotic/necrotic cell clearance. Cell Death Differ. 17(3): p. 381-97.

111. Pluddemann, A., C. Neyen, and S. Gordon, Macrophage scavenger recep-tors and host-derived ligands. Methods, 2007. 43(3): p. 207-17.

112. Flannagan, R.S., G. Cosio, and S. Grinstein, Antimicrobial mechanisms of phagocytes and bacterial evasion strategies. Nat Rev Microbiol, 2009. 7(5): p. 355-66.

113. Ramachandra, L., D. Simmons, and C.V. Harding, MHC molecules and microbial antigen processing in phagosomes. Curr Opin Immunol, 2009. 21(1): p. 98-104.

114. Newby, A.C., Metalloproteinase expression in monocytes and macro-phages and its relationship to atherosclerotic plaque instability. Arterio-scler Thromb Vasc Biol, 2008. 28(12): p. 2108-14.

115. Kalesnikoff, J. and S.J. Galli, New developments in mast cell biology. Nat Immunol, 2008. 9(11): p. 1215-23.

116. Marshall, J.S., Mast-cell responses to pathogens. Nat Rev Immunol, 2004. 4(10): p. 787-99.

117. Banchereau, J., et al., Immunobiology of dendritic cells. Annu Rev Immu-nol, 2000. 18: p. 767-811.

78

118. Blander, J.M., Phagocytosis and antigen presentation: a partnership initi-ated by Toll-like receptors. Ann Rheum Dis, 2008. 67 Suppl 3: p. iii44-9.

119. Frisch, S.M. and E. Ruoslahti, Integrins and anoikis. Curr Opin Cell Biol, 1997. 9(5): p. 701-6.

120. Hynes, R.O., Integrins: bidirectional, allosteric signaling machines. Cell, 2002. 110(6): p. 673-87.

121. Ruoslahti, E., RGD and other recognition sequences for integrins. Annu Rev Cell Dev Biol, 1996. 12: p. 697-715.

122. Wipff, P.J. and B. Hinz, Integrins and the activation of latent transforming growth factor beta1 - an intimate relationship. Eur J Cell Biol, 2008. 87(8-9): p. 601-15.

123. Giancotti, F.G. and E. Ruoslahti, Integrin signaling. Science, 1999. 285: p. 1028-1032.

124. Sundberg, C. and K. Rubin, Stimulation of beta1 integrins on fibroblasts induces PDGF independent tyrosine phosphorylation of PDGF beta-receptors. J Cell Biol, 1996. 132(4): p. 741-52.

125. Cabodi, S., et al., Integrin regulation of epidermal growth factor (EGF) receptor and of EGF-dependent responses. Biochem Soc Trans, 2004. 32(Pt3): p. 438-42.

126. Schneller, M., K. Vuori, and E. Ruoslahti, Alphavbeta3 integrin associates with activated insulin and PDGFbeta receptors and potentiates the biologi-cal activity of PDGF. Embo J, 1997. 16(18): p. 5600-7.

127. Ilic, D., et al., Reduced cell motility and enhanced focal adhesion contact formation in cells from FAK-deficient mice. Nature, 1995. 377(6549): p. 539-44.

128. Cleaver, O., Krieg, P.A., Molecular Mechanisms of Vascular Development, in Heart Development, R.P. Harvey, Rosenthal, N., Editor. 1999, Academic Press: San Diego. p. 221-244.

129. Yadav, R., et al., Migration of leukocytes through the vessel wall and be-yond. Thromb Haemost, 2003. 90(4): p. 598-606.

130. Tse, D. and R.V. Stan, Morphological heterogeneity of endothelium. Semin Thromb Hemost. 36(3): p. 236-45.

131. Birk, D.E., Bruckner, P., Collagen Suprastructures. Top Curr Chem, 2005. 247: p. 185-205.

132. Hudson, B.G., et al., Alport's syndrome, Goodpasture's syndrome, and type IV collagen. N Engl J Med, 2003. 348(25): p. 2543-56.

133. O'Reilly, M.S., et al., Endostatin: an endogenous inhibitor of angiogenesis and tumor growth. Cell, 1997. 88(2): p. 277-85.

134. Sasaki, T., et al., Endostatins derived from collagens XV and XVIII differ in structural and binding properties, tissue distribution and anti-angiogenic activity. J Mol Biol, 2000. 301(5): p. 1179-90.

135. Kalluri, R., Basement membranes: structure, assembly and role in tumour angiogenesis. Nat Rev Cancer, 2003. 3(6): p. 422-33.

136. Scheele, S., et al., Laminin isoforms in development and disease. J Mol Med, 2007. 85(8): p. 825-36.

79

137. Rodriguez, A., et al., Integrin alpha1beta1 is involved in the differentiation into myofibroblasts in adult reactive tissues in vivo. J Cell Mol Med, 2009. 13(9B): p. 3449-62.

138. de Vega, S., T. Iwamoto, and Y. Yamada, Fibulins: multiple roles in matrix structures and tissue functions. Cell Mol Life Sci, 2009. 66(11-12): p. 1890-902.

139. Allt, G. and J.G. Lawrenson, Pericytes: cell biology and pathology. Cells Tissues Organs, 2001. 169(1): p. 1-11.

140. Diaz-Flores, L., et al., Pericytes. Morphofunction, interactions and pathol-ogy in a quiescent and activated mesenchymal cell niche. Histol Histopa-thol, 2009. 24(7): p. 909-69.

141. Joyce, N.C., M.F. Haire, and G.E. Palade, Contractile proteins in pericytes. I. Immunoperoxidase localization of tropomyosin. J Cell Biol, 1985. 100(5): p. 1379-86.

142. Peppiatt, C.M., et al., Bidirectional control of CNS capillary diameter by pericytes. Nature, 2006. 443(7112): p. 700-4.

143. Sundberg, C., et al., Stable expression of angiopoietin-1 and other markers by cultured pericytes: phenotypic similarities to a subpopulation of cells in maturing vessels during later stages of angiogenesis in vivo. Lab Invest, 2002. 82(4): p. 387-401.

144. Sims, D.E., Recent advances in pericyte biology--implications for health and disease. Can J Cardiol, 1991. 7(10): p. 431-43.

145. Laties, A.M., S.I. Rapoport, and A. McGlinn, Hypertensive breakdown of cerebral but not of retinal blood vessels in rhesus monkey. Arch Ophthal-mol, 1979. 97(8): p. 1511-4.

146. Uemura, A., et al., Recombinant angiopoietin-1 restores higher-order ar-chitecture of growing blood vessels in mice in the absence of mural cells. J Clin Invest, 2002. 110(11): p. 1619-28.

147. Gerhardt, H., H. Wolburg, and C. Redies, N-cadherin mediates pericytic-endothelial interaction during brain angiogenesis in the chicken. Dev Dyn, 2000. 218(3): p. 472-9.

148. Tong, R.T., et al., Vascular normalization by vascular endothelial growth factor receptor 2 blockade induces a pressure gradient across the vascula-ture and improves drug penetration in tumors. Cancer Res, 2004. 64(11): p. 3731-6.

149. Benjamin, L.E., I. Hemo, and E. Keshet, A plasticity window for blood vessel remodelling is defined by pericyte coverage of the preformed endo-thelial network and is regulated by PDGF-B and VEGF. Development, 1998. 125(9): p. 1591-8.

150. Hellstrom, M., et al., Lack of pericytes leads to endothelial hyperplasia and abnormal vascular morphogenesis. J Cell Biol, 2001. 153(3): p. 543-53.

151. Rajkumar, V.S., et al., Shared expression of phenotypic markers in systemic sclerosis indicates a convergence of pericytes and fibroblasts to a myofi-broblast lineage in fibrosis. Arthritis Res Ther, 2005. 7(5): p. R1113-23.

152. Nehls, V. and D. Drenckhahn, The versatility of microvascular pericytes: from mesenchyme to smooth muscle? Histochemistry, 1993. 99(1): p. 1-12.

80

153. Farrington-Rock, C., et al., Chondrogenic and adipogenic potential of mi-crovascular pericytes. Circulation, 2004. 110(15): p. 2226-32.

154. Doherty, M.J., et al., Vascular pericytes express osteogenic potential in vitro and in vivo. J Bone Miner Res, 1998. 13(5): p. 828-38.

155. Davidoff, M.S., et al., Progenitor cells of the testosterone-producing Ley-dig cells revealed. J Cell Biol, 2004. 167(5): p. 935-44.

156. Schmid-Schonbein, G.W., Microlymphatics and lymph flow. Physiol Rev, 1990. 70(4): p. 987-1028.

157. Hammacher, A., et al., A major part of platelet-derived growth factor puri-fied from human platelets is a heterodimer of one A and one B chain. J Biol Chem, 1988. 263(31): p. 16493-8.

158. Hammacher, A., et al., A human glioma cell line secretes three structurally and functionally different dimeric forms of platelet-derived growth factor. Eur J Biochem, 1988. 176(1): p. 179-86.

159. Heldin, C.H. and B. Westermark, Mechanism of action and in vivo role of platelet-derived growth factor. Physiol Rev, 1999. 79(4): p. 1283-316.

160. Bostrom, H., et al., PDGF-A signaling is a critical event in lung alveolar myofibroblast development and alveogenesis. Cell, 1996. 85(6): p. 863-73.

161. Karlsson, L., C. Bondjers, and C. Betsholtz, Roles for PDGF-A and sonic hedgehog in development of mesenchymal components of the hair follicle. Development, 1999. 126(12): p. 2611-21.

162. Karlsson, L., et al., Abnormal gastrointestinal development in PDGF-A and PDGFR-(alpha) deficient mice implicates a novel mesenchymal structure with putative instructive properties in villus morphogenesis. Development, 2000. 127(16): p. 3457-66.

163. Fruttiger, M., et al., Defective oligodendrocyte development and severe hypomyelination in PDGF-A knockout mice. Development, 1999. 126(3): p. 457-67.

164. Eriksson, A., et al., PDGF alpha- and beta-receptors activate unique and common signal transduction pathways. Embo J, 1992. 11(2): p. 543-550.

165. Pierce, G.F., et al., Detection of platelet-derived growth factor (PDGF)-AA in actively healing human wounds treated with recombinant PDGF-BB and absence of PDGF in chronic nonhealing wounds. J Clin Invest, 1995. 96(3): p. 1336-50.

166. Hermanson, M., et al., Association of loss of heterozygosity on chromosome 17p with high platelet-derived growth factor alpha receptor expression in human malignant gliomas. Cancer Res, 1996. 56(1): p. 164-71.

167. Betsholtz, C., Insight into the physiological functions of PDGF through genetic studies in mice. Cytokine Growth Factor Rev, 2004. 15(4): p. 215-28.

168. Leveen, P., et al., Mice deficient for PDGF B show renal, cardiovascular, and hematological abnormalities. Genes Dev, 1994. 8(16): p. 1875-87.

169. Lindblom, P., et al., Endothelial PDGF-B retention is required for proper investment of pericytes in the microvessel wall. Genes Dev, 2003. 17(15): p. 1835-40.

170. Deuel, T.F., et al., Chemotaxis of monocytes and neutrophils to platelet-derived growth factor. J Clin Invest, 1982. 69(4): p. 1046-9.

81

171. Liden, A., et al., A secreted collagen- and fibronectin-binding streptococcal protein modulates cell-mediated collagen gel contraction and interstitial fluid pressure. J Biol Chem, 2008. 283(3): p. 1234-42.

172. Ivarsson, M., et al., Type I collagen synthesis in cultured human fibro-blasts: regulation by cell spreading, platelet-derived growth factor and in-teractions with collagen fibers. Matrix Biol, 1998. 16(7): p. 409-25.

173. Risau, W., et al., Platelet-derived growth factor is angiogenic in vivo. Growth Factors, 1992. 7(4): p. 261-6.

174. Uhl, E., et al., Influence of platelet-derived growth factor on microcircula-tion during normal and impaired wound healing. Wound Repair Regen, 2003. 11(5): p. 361-7.

175. Bonner, J.C., Regulation of PDGF and its receptors in fibrotic diseases. Cytokine Growth Factor Rev, 2004. 15(4): p. 255-73.

176. Liden, A., et al., Platelet-derived growth factor BB-mediated normalization of dermal interstitial fluid pressure after mast cell degranulation depends on beta3 but not beta1 integrins. Circ Res, 2006. 98(5): p. 635-41.

177. Smits, A., et al., Expression of platelet-derived growth factor and its recep-tors in proliferative disorders of fibroblastic origin. Am J Pathol, 1992. 140(3): p. 639-48.

178. Pietras, K., et al., PDGF receptors as cancer drug targets. Cancer Cell, 2003. 3(5): p. 439-43.

179. Dijkmans, J., et al., Characterization of platelet-derived growth factor-C (PDGF-C): expression in normal and tumor cells, biological activity and chromosomal localization. Int J Biochem Cell Biol, 2002. 34(4): p. 414-26.

180. Ding, H., et al., A specific requirement for PDGF-C in palate formation and PDGFR-alpha signaling. Nat Genet, 2004. 36(10): p. 1111-6.

181. Eitner, F., et al., PDGF-C is a proinflammatory cytokine that mediates renal interstitial fibrosis. J Am Soc Nephrol, 2008. 19(2): p. 281-9.

182. Cao, R., et al., Angiogenesis stimulated by PDGF-CC, a novel member in the PDGF family, involves activation of PDGFR-alphaalpha and -alphabeta receptors. Faseb J, 2002. 16(12): p. 1575-83.

183. Pietras, K., et al., Functions of paracrine PDGF signaling in the proangio-genic tumor stroma revealed by pharmacological targeting. PLoS Med, 2008. 5(1): p. e19.

184. Ponten, A., et al., Transgenic overexpression of platelet-derived growth factor-C in the mouse heart induces cardiac fibrosis, hypertrophy, and di-lated cardiomyopathy. Am J Pathol, 2003. 163(2): p. 673-82.

185. Campbell, J.S., et al., Targeting stromal cells for the treatment of platelet-derived growth factor C-induced hepatocellular carcinogenesis. Differen-tiation, 2007. 75(9): p. 843-52.

186. Boor, P., et al., PDGF-C mediates glomerular capillary repair. Am J Pa-thol. 177(1): p. 58-69.

187. Li, X., et al., Revascularization of ischemic tissues by PDGF-CC via effects on endothelial cells and their progenitors. J Clin Invest, 2005. 115(1): p. 118-27.

188. Bergsten, E., et al., PDGF-D is a specific, protease-activated ligand for the PDGF beta-receptor. Nat Cell Biol, 2001. 3(5): p. 512-6.

82

189. Ponten, A., et al., Platelet-derived growth factor D induces cardiac fibrosis and proliferation of vascular smooth muscle cells in heart-specific trans-genic mice. Circ Res, 2005. 97(10): p. 1036-45.

190. Taneda, S., et al., Obstructive uropathy in mice and humans: potential role for PDGF-D in the progression of tubulointerstitial injury. J Am Soc Nephrol, 2003. 14(10): p. 2544-55.

191. Li, H., et al., PDGF-D is a potent transforming and angiogenic growth factor. Oncogene, 2003. 22(10): p. 1501-10.

192. Uutela, M., et al., PDGF-D induces macrophage recruitment, increased interstitial pressure, and blood vessel maturation during angiogenesis. Blood, 2004. 104(10): p. 3198-204.

193. Rosenkranz, S., et al., Identification of the receptor-associated signaling enzymes that are required for platelet-derived growth factor-AA-dependent chemotaxis and DNA synthesis. J Biol Chem, 1999. 274(40): p. 28335-43.

194. Gilbertson, D.G., et al., Platelet-derived growth factor C (PDGF-C), a novel growth factor that binds to PDGF alpha and beta receptor. J Biol Chem, 2001. 276(29): p. 27406-14.

195. Rubin, K., et al., Induction of B-type receptors for platelet-derived growth factor in vascular inflammation: possible implications for development of vascular proliferative lesions. Lancet, 1988. 1(8599): p. 1353-6.

196. Sundberg, C., et al., Microvascular pericytes express platelet-derived growth factor-beta receptors in human healing wounds and colorectal ade-nocarcinoma. Am J Pathol, 1993. 143(5): p. 1377-88.

197. Abramsson, A., P. Lindblom, and C. Betsholtz, Endothelial and nonendo-thelial sources of PDGF-B regulate pericyte recruitment and influence vas-cular pattern formation in tumors. J Clin Invest, 2003. 112(8): p. 1142-51.

198. Rankin, S. and E. Rozengurt, Platelet-derived growth factor modulation of focal adhesion kinase (p125FAK) and paxillin tyrosine phosphorylation in Swiss 3T3 cells. Bell- shaped dose response and cross-talk with bombesin. J Biol Chem, 1994. 269(1): p. 704-10.

199. Jones, S.M. and A. Kazlauskas, Growth-factor-dependent mitogenesis requires two distinct phases of signalling. Nat Cell Biol, 2001. 3(2): p. 165-72.

200. Sorkin, A., et al., Effect of receptor kinase inactivation on the rate of inter-nalization and degradation of PDGF and the PDGF beta-receptor. J Cell Biol, 1991. 112(3): p. 469-78.

201. Sundberg, C., et al., Tumor cell and connective tissue cell interactions in human colorectal adenocarcinoma. Transfer of platelet-derived growth factor-AB/BB to stromal cells. Am J Pathol, 1997. 151(2): p. 479-92.

202. Abramsson, A., et al., Defective N-sulfation of heparan sulfate proteogly-cans limits PDGF-BB binding and pericyte recruitment in vascular devel-opment. Genes Dev, 2007. 21(3): p. 316-31.

203. ten Dijke, P. and H.M. Arthur, Extracellular control of TGFbeta signalling in vascular development and disease. Nat Rev Mol Cell Biol, 2007. 8(11): p. 857-69.

204. Shi, Y. and J. Massague, Mechanisms of TGF-beta signaling from cell membrane to the nucleus. Cell, 2003. 113(6): p. 685-700.

83

205. Massague, J., et al., TGF-beta receptors and TGF-beta binding proteogly-cans: recent progress in identifying their functional properties. Ann N Y Acad Sci, 1990. 593: p. 59-72.

206. Goumans, M.J. and C. Mummery, Functional analysis of the TGFbeta receptor/Smad pathway through gene ablation in mice. Int J Dev Biol, 2000. 44(3): p. 253-65.

207. Larsson, J., et al., Abnormal angiogenesis but intact hematopoietic poten-tial in TGF-beta type I receptor-deficient mice. Embo J, 2001. 20(7): p. 1663-73.

208. Dickson, M.C., et al., Defective haematopoiesis and vasculogenesis in transforming growth factor-beta 1 knock out mice. Development, 1995. 121(6): p. 1845-54.

209. Letterio, J.J., et al., Autoimmunity associated with TGF-beta1-deficiency in mice is dependent on MHC class II antigen expression. J Clin Invest, 1996. 98(9): p. 2109-19.

210. Sanford, L.P., et al., TGFbeta2 knockout mice have multiple developmental defects that are non-overlapping with other TGFbeta knockout phenotypes. Development, 1997. 124(13): p. 2659-70.

211. Kaartinen, V., et al., Abnormal lung development and cleft palate in mice lacking TGF-beta 3 indicates defects of epithelial-mesenchymal interac-tion. Nat Genet, 1995. 11(4): p. 415-21.

212. Scaffidi, A.K., et al., alpha(v)beta(3) Integrin interacts with the transform-ing growth factor beta (TGFbeta) type II receptor to potentiate the prolif-erative effects of TGFbeta1 in living human lung fibroblasts. J Biol Chem, 2004. 279(36): p. 37726-33.

213. Leask, A. and D.J. Abraham, TGF-beta signaling and the fibrotic response. Faseb J, 2004. 18(7): p. 816-27.

214. Bottinger, E.P. and J.B. Kopp, Lessons from TGF-beta transgenic mice. Miner Electrolyte Metab, 1998. 24(2-3): p. 154-60.

215. Gabbiani, G., The myofibroblast in wound healing and fibrocontractive diseases. J Pathol, 2003. 200(4): p. 500-3.

216. Li, M.O., et al., Transforming growth factor-beta regulation of immune responses. Annu Rev Immunol, 2006. 24: p. 99-146.

217. Pardali, K. and A. Moustakas, Actions of TGF-beta as tumor suppressor and pro-metastatic factor in human cancer. Biochim Biophys Acta, 2007. 1775(1): p. 21-62.

218. Reed, R.K., A. Liden, and K. Rubin, Edema and fluid dynamics in connec-tive tissue remodelling. J Mol Cell Cardiol, 2010. 48(3): p. 518-23.

219. Michel, C.C., Starling: the formulation of his hypothesis of microvascular fluid exchange and its significance after 100 years. Exp Physiol, 1997. 82(1): p. 1-30.

220. Miller, L.S. and R.L. Modlin, Toll-like receptors in the skin. Semin Im-munopathol, 2007. 29(1): p. 15-26.

221. Nagy, J.A., et al., Vascular permeability, vascular hyperpermeability and angiogenesis. Angiogenesis, 2008. 11(2): p. 109-19.

84

222. Nissen, N.N., et al., Vascular endothelial growth factor mediates angio-genic activity during the proliferative phase of wound healing. Am J Pathol, 1998. 152(6): p. 1445-52.

223. Barrientos, S., et al., Growth factors and cytokines in wound healing. Wound Repair Regen, 2008. 16(5): p. 585-601.

224. Sundberg, C., et al., Glomeruloid microvascular proliferation follows ade-noviral vascular permeability factor/vascular endothelial growth factor-164 gene delivery. Am J Pathol, 2001. 158(3): p. 1145-60.

225. Nagy, J.A., A.M. Dvorak, and H.F. Dvorak, VEGF-A and the induction of pathological angiogenesis. Annu Rev Pathol, 2007. 2: p. 251-75.

226. Pettersson, A., et al., Heterogeneity of the angiogenic response induced in different normal adult tissues by vascular permeability factor/vascular en-dothelial growth factor. Lab Invest, 2000. 80(1): p. 99-115.

227. Nedrebo, T., et al., A novel function of insulin in rat dermis. J Physiol, 2004. 559(Pt 2): p. 583-91.

228. Rondaij, M.G., et al., Dynamics and plasticity of Weibel-Palade bodies in endothelial cells. Arterioscler Thromb Vasc Biol, 2006. 26(5): p. 1002-7.

229. Gillitzer, R. and M. Goebeler, Chemokines in cutaneous wound healing. J Leukoc Biol, 2001. 69(4): p. 513-21.

230. Dovi, J.V., A.M. Szpaderska, and L.A. DiPietro, Neutrophil function in the healing wound: adding insult to injury? Thromb Haemost, 2004. 92(2): p. 275-80.

231. Mott, J.D. and Z. Werb, Regulation of matrix biology by matrix metallopro-teinases. Curr Opin Cell Biol, 2004. 16(5): p. 558-64.

232. Montgomery, A.M., R.A. Reisfeld, and D.A. Cheresh, Integrin alpha v beta 3 rescues melanoma cells from apoptosis in three-dimensional dermal col-lagen. Proc Natl Acad Sci U S A, 1994. 91(19): p. 8856-60.

233. Clark, R.A., et al., Transient functional expression of alphaVbeta 3 on vascular cells during wound repair. Am J Pathol, 1996. 148(5): p. 1407-21.

234. Singer, A.J. and R.A. Clark, Cutaneous wound healing. N Engl J Med, 1999. 341(10): p. 738-46.

235. Diaz-Flores, L., R. Gutierrez, and H. Varela, Behavior of postcapillary venule pericytes during postnatal angiogenesis. J Morphol, 1992. 213(1): p. 33-45.

236. Cavallo, T., et al., Ultrastructural autoradiographic studies of the early vasoproliferative response in tumor angiogenesis. Am J Pathol, 1973. 70(3): p. 345-62.

237. Rajkumar, V.S., et al., Platelet-derived growth factor-beta receptor activa-tion is essential for fibroblast and pericyte recruitment during cutaneous wound healing. Am J Pathol, 2006. 169(6): p. 2254-65.

238. Meszaros, A.J., J.S. Reichner, and J.E. Albina, Macrophage-induced neu-trophil apoptosis. J Immunol, 2000. 165(1): p. 435-41.

239. Lee, T.S. and L.Y. Chau, Heme oxygenase-1 mediates the anti-inflammatory effect of interleukin-10 in mice. Nat Med, 2002. 8(3): p. 240-6.

240. Sundberg, C., et al., Pericytes as collagen-producing cells in excessive dermal scarring. Lab Invest, 1996. 74(2): p. 452-66.

85

241. Hadfield, G., The Tissue of Origin of the Fibroblasts of Granulation Tissue. Br J Surg, 1963. 50: p. 870-81.

242. Kisseleva, T. and D.A. Brenner, Mechanisms of fibrogenesis. Exp Biol Med (Maywood), 2008. 233(2): p. 109-22.

243. Madlener, M., W.C. Parks, and S. Werner, Matrix metalloproteinases (MMPs) and their physiological inhibitors (TIMPs) are differentially ex-pressed during excisional skin wound repair. Exp Cell Res, 1998. 242(1): p. 201-10.

244. Overall, C.M., J.L. Wrana, and J. Sodek, Independent regulation of colla-genase, 72-kDa progelatinase, and metalloendoproteinase inhibitor ex-pression in human fibroblasts by transforming growth factor-beta. J Biol Chem, 1989. 264(3): p. 1860-9.

245. Haroon, Z.A., et al., Early wound healing exhibits cytokine surge without evidence of hypoxia. Ann Surg, 2000. 231(1): p. 137-47.

246. Wang, S., et al., Venular basement membranes contain specific matrix protein low expression regions that act as exit points for emigrating neu-trophils. J Exp Med, 2006. 203(6): p. 1519-32.

247. Crocker, D.J., T.M. Murad, and J.C. Geer, Role of the pericyte in wound healing. An ultrastructural study. Exp Mol Pathol, 1970. 13(1): p. 51-65.

248. Del Monte, M.A., et al., Sorbitol, myo-inositol, and rod outer segment phagocytosis in cultured hRPE cells exposed to glucose. In vitro model of myo-inositol depletion hypothesis of diabetic complications. Diabetes, 1991. 40(10): p. 1335-45.

249. Wakui, S., et al., Transforming growth factor-beta and urokinase plasmi-nogen activator presents at endothelial cell-pericyte interdigitation in hu-man granulation tissue. Microvasc Res, 1997. 54(3): p. 262-9.

250. Cliff, W.J., Observations on healing tissue: A combined light and electron microscopic investigation. Philological Transductions of the Royal Society (London) B, 1963. 246: p. 325.

251. Ji, R.C., et al., Expression of VEGFR-3 and 5'-nase in regenerating lym-phatic vessels of the cutaneous wound healing. Microsc Res Tech, 2004. 64(3): p. 279-86.

252. Nagy, J.A., et al., Vascular permeability factor/vascular endothelial growth factor induces lymphangiogenesis as well as angiogenesis. J Exp Med, 2002. 196(11): p. 1497-506.

253. Buckmire, M.A., et al., Temporal expression of TGF-beta1, EGF, and PDGF-BB in a model of colonic wound healing. J Surg Res, 1998. 80(1): p. 52-7.

254. Hinz, B., Formation and function of the myofibroblast during tissue repair. J Invest Dermatol, 2007. 127(3): p. 526-37.

255. Denton, C.P., et al., Inducible lineage-specific deletion of TbetaRII in fi-broblasts defines a pivotal regulatory role during adult skin wound healing. J Invest Dermatol, 2009. 129(1): p. 194-204.

256. Eckes, B., et al., Mechanical tension and integrin alpha 2 beta 1 regulate fibroblast functions. J Investig Dermatol Symp Proc, 2006. 11(1): p. 66-72.

86

257. Lambert, C.A., et al., Coordinated regulation of procollagens I and III and their post-translational enzymes by dissipation of mechanical tension in human dermal fibroblasts. Eur J Cell Biol, 2001. 80(7): p. 479-85.

258. Szauter, K.M., et al., Lysyl oxidase in development, aging and pathologies of the skin. Pathol Biol (Paris), 2005. 53(7): p. 448-56.

259. Woodley, D.T., et al., Collagen telopeptides (cross-linking sites) play a role in collagen gel lattice contraction. J Invest Dermatol, 1991. 97(3): p. 580-5.

260. Hayakawa, T., et al., Changes in type of collagen during the development of human post-burn hypertrophic scars. Clin Chim Acta, 1979. 93(1): p. 119-25.

261. Desmouliere, A., et al., Apoptosis mediates the decrease in cellularity dur-ing the transition between granulation tissue and scar. Am J Pathol, 1995. 146(1): p. 56-66.

262. Kibe, Y., H. Takenaka, and S. Kishimoto, Spatial and temporal expression of basic fibroblast growth factor protein during wound healing of rat skin. Br J Dermatol, 2000. 143(4): p. 720-7.

263. Knapp, T.R., R.J. Daniels, and E.N. Kaplan, Pathologic scar formation. Morphologic and biochemical correlates. Am J Pathol, 1977. 86(1): p. 47-70.

264. Shih, B., et al., Molecular dissection of abnormal wound healing processes resulting in keloid disease. Wound Repair Regen. 18(2): p. 139-53.

265. van der Veer, W.M., et al., Potential cellular and molecular causes of hy-pertrophic scar formation. Burns, 2009. 35(1): p. 15-29.

266. Wynn, T.A., Fibrotic disease and the T(H)1/T(H)2 paradigm. Nat Rev Immunol, 2004. 4(8): p. 583-94.

267. Kitani, A., et al., Transforming growth factor (TGF)-beta1-producing regu-latory T cells induce Smad-mediated interleukin 10 secretion that facilitates coordinated immunoregulatory activity and amelioration of TGF-beta1-mediated fibrosis. J Exp Med, 2003. 198(8): p. 1179-88.

268. Amadeu, T., et al., Vascularization pattern in hypertrophic scars and kelo-ids: a stereological analysis. Pathol Res Pract, 2003. 199(7): p. 469-73.

269. Lu, F., et al., Variations in gap junctional intercellular communication and connexin expression in fibroblasts derived from keloid and hypertrophic scars. Plast Reconstr Surg, 2007. 119(3): p. 844-51.

270. van der Slot-Verhoeven, A.J., et al., The type of collagen cross-link deter-mines the reversibility of experimental skin fibrosis. Biochim Biophys Acta, 2005. 1740(1): p. 60-7.

271. van der Slot, A.J., et al., Elevated formation of pyridinoline cross-links by profibrotic cytokines is associated with enhanced lysyl hydroxylase 2b lev-els. Biochim Biophys Acta, 2005. 1741(1-2): p. 95-102.

272. Buchanan, E.P., M.T. Longaker, and H.P. Lorenz, Fetal skin wound heal-ing. Adv Clin Chem, 2009. 48: p. 137-61.

273. Dvorak, H.F., Tumors: wounds that do not heal. Similarities between tumor stroma generation and wound healing. N Engl J Med, 1986. 315(26): p. 1650-9.

87

274. Farmer, P., et al., A stroma-related gene signature predicts resistance to neoadjuvant chemotherapy in breast cancer. Nat Med, 2009. 15(1): p. 68-74.

275. Kroemer, G. and J. Pouyssegur, Tumor cell metabolism: cancer's Achilles' heel. Cancer Cell, 2008. 13(6): p. 472-82.

276. Less, J.R., et al., Microvascular architecture in a mammary carcinoma: branching patterns and vessel dimensions. Cancer Res, 1991. 51(1): p. 265-73.

277. Dvorak, H.F., Rous-Whipple Award Lecture. How tumors make bad blood vessels and stroma. Am J Pathol, 2003. 162(6): p. 1747-57.

278. Bergers, G. and S. Song, The role of pericytes in blood-vessel formation and maintenance. Neuro Oncol, 2005. 7(4): p. 452-64.

279. Nehme, A. and J. Edelman, Dexamethasone inhibits high glucose-, TNF-alpha-, and IL-1beta-induced secretion of inflammatory and angiogenic mediators from retinal microvascular pericytes. Invest Ophthalmol Vis Sci, 2008. 49(5): p. 2030-8.

280. Walshe, T.E., et al., TGF-beta is required for vascular barrier function, endothelial survival and homeostasis of the adult microvasculature. PLoS One, 2009. 4(4): p. e5149.

281. Pouyssegur, J., F. Dayan, and N.M. Mazure, Hypoxia signalling in cancer and approaches to enforce tumour regression. Nature, 2006. 441(7092): p. 437-43.

282. Verona, E.V., et al., Transforming growth factor-beta signaling in prostate stromal cells supports prostate carcinoma growth by up-regulating stromal genes related to tissue remodeling. Cancer Res, 2007. 67(12): p. 5737-46.

283. Olumi, A.F., et al., Carcinoma-associated fibroblasts direct tumor progres-sion of initiated human prostatic epithelium. Cancer Res, 1999. 59(19): p. 5002-11.

284. Lieubeau, B., et al., The role of transforming growth factor beta 1 in the fibroblastic reaction associated with rat colorectal tumor development. Cancer Res, 1994. 54(24): p. 6526-32.

285. Micke, P. and A. Ostman, Tumour-stroma interaction: cancer-associated fibroblasts as novel targets in anti-cancer therapy? Lung Cancer, 2004. 45 Suppl 2: p. S163-75.

286. Lohr, M., et al., Transforming growth factor-beta1 induces desmoplasia in an experimental model of human pancreatic carcinoma. Cancer Res, 2001. 61(2): p. 550-5.

287. Bhowmick, N.A., et al., TGF-beta signaling in fibroblasts modulates the oncogenic potential of adjacent epithelia. Science, 2004. 303(5659): p. 848-51.

288. Hembruff, S.L., et al., Loss of transforming growth factor-beta signaling in mammary fibroblasts enhances CCL2 secretion to promote mammary tu-mor progression through macrophage-dependent and -independent mecha-nisms. Neoplasia. 12(5): p. 425-33.

289. Doxey, D.L., et al., Diabetes-induced impairment of macrophage cytokine release in a rat model: potential role of serum lipids. Life Sci, 1998. 63(13): p. 1127-36.

88

290. Blackhall, F.H., et al., Heparan sulfate proteoglycans and cancer. Br J Cancer, 2001. 85(8): p. 1094-8.

291. Lederle, W., et al., Platelet-derived growth factor-BB controls epithelial tumor phenotype by differential growth factor regulation in stromal cells. Am J Pathol, 2006. 169(5): p. 1767-83.

292. Skobe, M. and N.E. Fusenig, Tumorigenic conversion of immortal human keratinocytes through stromal cell activation. Proc Natl Acad Sci U S A, 1998. 95(3): p. 1050-5.

293. Forsberg, K., et al., Platelet-derived growth factor (PDGF) in oncogenesis: development of a vascular connective tissue stroma in xenotransplanted human melanoma producing PDGF-BB. Proc Natl Acad Sci U S A, 1993. 90(2): p. 393-7.

294. Xu, L., et al., Blocking platelet-derived growth factor-D/platelet-derived growth factor receptor beta signaling inhibits human renal cell carcinoma progression in an orthotopic mouse model. Cancer Res, 2005. 65(13): p. 5711-9.

295. Furuhashi, M., et al., Platelet-derived growth factor production by B16 melanoma cells leads to increased pericyte abundance in tumors and an associated increase in tumor growth rate. Cancer Res, 2004. 64(8): p. 2725-33.

296. Dong, J., et al., VEGF-null cells require PDGFR alpha signaling-mediated stromal fibroblast recruitment for tumorigenesis. Embo J, 2004. 23(14): p. 2800-10.

297. Tejada, M.L., et al., Tumor-driven paracrine platelet-derived growth factor receptor alpha signaling is a key determinant of stromal cell recruitment in a model of human lung carcinoma. Clin Cancer Res, 2006. 12(9): p. 2676-88.

298. Crawford, Y., et al., PDGF-C mediates the angiogenic and tumorigenic properties of fibroblasts associated with tumors refractory to anti-VEGF treatment. Cancer Cell, 2009. 15(1): p. 21-34.

299. Fukumura, D., et al., Tumor induction of VEGF promoter activity in stro-mal cells. Cell, 1998. 94(6): p. 715-25.

300. Anderberg, C., et al., Paracrine signaling by platelet-derived growth fac-tor-CC promotes tumor growth by recruitment of cancer-associated fibro-blasts. Cancer Res, 2009. 69(1): p. 369-78.

301. Franco, O.E., et al., Cancer associated fibroblasts in cancer pathogenesis. Semin Cell Dev Biol. 21(1): p. 33-9.

302. Orimo, A., et al., Stromal fibroblasts present in invasive human breast carcinomas promote tumor growth and angiogenesis through elevated SDF-1/CXCL12 secretion. Cell, 2005. 121(3): p. 335-48.

303. Nazareth, M.R., et al., Characterization of human lung tumor-associated fibroblasts and their ability to modulate the activation of tumor-associated T cells. J Immunol, 2007. 178(9): p. 5552-62.

304. Hill, R., et al., Selective evolution of stromal mesenchyme with p53 loss in response to epithelial tumorigenesis. Cell, 2005. 123(6): p. 1001-11.

89

305. Ronnov-Jessen, L., O.W. Petersen, and M.J. Bissell, Cellular changes in-volved in conversion of normal to malignant breast: importance of the stromal reaction. Physiol Rev, 1996. 76(1): p. 69-125.

306. Kleeff, J., et al., Pancreatic cancer microenvironment. Int J Cancer, 2007. 121(4): p. 699-705.

307. Olive, K.P., et al., Inhibition of Hedgehog signaling enhances delivery of chemotherapy in a mouse model of pancreatic cancer. Science, 2009. 324(5933): p. 1457-61.

308. Eyden, B. and M. Tzaphlidou, Structural variations of collagen in normal and pathological tissues: role of electron microscopy. Micron, 2001. 32(3): p. 287-300.

309. Hull, M.T. and K.A. Warfel, Ultrastructure of abnormal collagen in human tumors. Ultrastruct Pathol, 1986. 10(4): p. 293-301.

310. Egeblad, M., M.G. Rasch, and V.M. Weaver, Dynamic interplay between the collagen scaffold and tumor evolution. Curr Opin Cell Biol.

311. Zhu, G.G., et al., Immunohistochemical study of type I collagen and type I pN-collagen in benign and malignant ovarian neoplasms. Cancer, 1995. 75(4): p. 1010-7.

312. Kirschmann, D.A., et al., A molecular role for lysyl oxidase in breast can-cer invasion. Cancer Res, 2002. 62(15): p. 4478-83.

313. Paszek, M.J., et al., Tensional homeostasis and the malignant phenotype. Cancer Cell, 2005. 8(3): p. 241-54.

314. Gao, Y., et al., LKB1 inhibits lung cancer progression through lysyl oxi-dase and extracellular matrix remodeling. Proc Natl Acad Sci U S A.

315. Barry-Hamilton, V., et al., Allosteric inhibition of lysyl oxidase-like-2 im-pedes the development of a pathologic microenvironment. Nat Med. 16(9): p. 1009-17.

316. Shimazaki, M. and A. Kudo, Impaired capsule formation of tumors in pe-riostin-null mice. Biochem Biophys Res Commun, 2008. 367(4): p. 736-42.

317. Puolakkainen, P.A., et al., Enhanced growth of pancreatic tumors in SPARC-null mice is associated with decreased deposition of extracellular matrix and reduced tumor cell apoptosis. Mol Cancer Res, 2004. 2(4): p. 215-24.

318. Koukourakis, M.I., et al., Enhanced expression of SPARC/osteonectin in the tumor-associated stroma of non-small cell lung cancer is correlated with markers of hypoxia/acidity and with poor prognosis of patients. Can-cer Res, 2003. 63(17): p. 5376-80.

319. Erkan, M., et al., Periostin creates a tumor-supportive microenvironment in the pancreas by sustaining fibrogenic stellate cell activity. Gastroenterol-ogy, 2007. 132(4): p. 1447-64.

320. Fukumura, D. and R.K. Jain, Tumor microvasculature and microenviron-ment: targets for anti-angiogenesis and normalization. Microvasc Res, 2007. 74(2-3): p. 72-84.

321. Boucher, Y. and R.K. Jain, Microvascular pressure is the principal driving force for interstitial hypertension in solid tumors: implications for vascular collapse. Cancer-Res, 1992. 52(18): p. 5110-4.

90

322. Alexandrakis, G., et al., Two-photon fluorescence correlation microscopy reveals the two-phase nature of transport in tumors. Nat Med, 2004. 10(2): p. 203-7.

323. Zlotecki, R.A., et al., Effect of angiotensin II induced hypertension on tu-mor blood flow and interstitial fluid pressure. Cancer Res, 1993. 53(11): p. 2466-8.

324. Rodt, S.A., et al., A novel physiological function for platelet-derived growth factor-BB in rat dermis. J Physiol, 1996. 495 ( Pt 1): p. 193-200.

325. Salnikov, A.V., et al., Lowering of tumor interstitial fluid pressure specifi-cally augments efficacy of chemotherapy. Faseb J, 2003. 17(12): p. 1756-8.

326. Pietras, K., et al., STI571 enhances the therapeutic index of epothilone B by a tumor-selective increase of drug uptake. Clin Cancer Res, 2003. 9(10 Pt 1): p. 3779-87.

327. Pietras, K., et al., Inhibition of platelet-derived growth factor receptors reduces interstitial hypertension and increases transcapillary transport in tumors. Cancer Res, 2001. 61(7): p. 2929-34.

328. Pietras, K., et al., Inhibition of PDGF receptor signaling in tumor stroma enhances antitumor effect of chemotherapy. Cancer Res, 2002. 62(19): p. 5476-84.

329. Klosowska-Wardega, A., et al., Combined anti-angiogenic therapy target-ing PDGF and VEGF receptors lowers the interstitial fluid pressure in a murine experimental carcinoma. PLoS One, 2009. 4(12): p. e8149.

330. Salnikov, A.V., et al., Inhibition of carcinoma cell-derived VEGF reduces inflammatory characteristics in xenograft carcinoma. Int J Cancer, 2006. 119(12): p. 2795-802.

331. Salnikov, A.V., et al., Inhibition of TGF-beta modulates macrophages and vessel maturation in parallel to a lowering of interstitial fluid pressure in experimental carcinoma. Lab Invest, 2005. 85(4): p. 512-21.

332. Liu, P. and R.G. Anderson, Compartmentalized production of ceramide at the cell surface. J Biol Chem, 1995. 270(45): p. 27179-85.

333. Liu, P., et al., Localization of platelet-derived growth factor-stimulated phosphorylation cascade to caveolae. J Biol Chem, 1996. 271(17): p. 10299-303.

334. Tingstrom, A., et al., Expression of platelet-derived growth factor-beta receptors on human fibroblasts. Regulation by recombinant platelet-derived growth factor- BB, IL-1, and tumor necrosis factor-alpha. J Immu-nol, 1992. 148(2): p. 546-54.

335. Ain, K.B. and K.D. Taylor, Somatostatin analogs affect proliferation of human thyroid carcinoma cell lines in vitro. J Clin Endocrinol Metab, 1994. 78: p. 1097-1102.

336. Schweppe, R.E., et al., Deoxyribonucleic acid profiling analysis of 40 hu-man thyroid cancer cell lines reveals cross-contamination resulting in cell line redundancy and misidentification. J Clin Endocrinol Metab, 2008. 93(11): p. 4331-41.

337. Reynolds, L.E., et al., Accelerated re-epithelialization in beta3-integrin-deficient- mice is associated with enhanced TGF-beta1 signaling. Nat Med, 2005. 11(2): p. 167-74.