Chapter Two a Novel Role of Vimentin Filaments: Bindiing and Stabilization

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Electronic Theses, Treatises and Dissertations The Graduate School

2011 Understanding the Functions of Vimentin Filaments in Collagen Expression and Targeting Vimentin Filaments for the Treatment of Fibrosis Azariyas Challa

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COLLEGE OF MEDICINE

UNDERSTANDING THE FUNCTIONS OF VIMENTIN FILAMENTS IN COLLAGEN

EXPRESSION AND TARGETING VIMENTIN FILAMENTS FOR THE TREATMENT OF

FIBROSIS

AZARIYAS CHALLA

A dissertation submitted to the Department of Biomedical Sciences in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Degree Awarded: Fall Semester, 2011

Azariyas Challa defended this dissertation on October 28, 2011.

The members of the supervisory committee were:

Branko Stefanovic Professor Directing Dissertation

Hengli Tang University Representative

Choogon Lee Committee Member

James Olcese Committee Member

The Graduate School has verified and approved the above-named committee members, and certifies that the dissertation has been approved in accordance with university requirements.

I dedicate this dissertation to my family.

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ACKNOWLEDGEMENTS

First I would like to thank my major professor Dr. Stefanovic who has opened his door for me and guided me whole heartedly throughout the past 4 years. His exceptional commitment to my projects made my work smooth, with much less moments of frustrations than I expected when I first learnt the reality of what a PhD entails. His work ethic has set the standards towards which I had to always push myself to match. Besides helping me with my dissertation projects, I would like to thank him for going out of his way to help me with my career advancement endeavors. He has consistently encouraged me to simultaneously take the licensing examinations for medical residency. He has also greatly helped me in finding postdoctoral positions by using his personal contacts and by facilitating a trip to potential postdoc mentors for interviews.

Special thanks to my committee members, Dr. Choogon Lee, Dr. James Olcese and Dr. Hengli Tang for their dedication to facilitate my progress and also for their help in my career development. Although not my committee members, Dr. Yanchang Wang and several other faculty members had always been keen to help me with my studies.

My thanks also go to our lab manager, Lela Stefanovic whose excellent technical support and care made life easier in the lab. I was also fortunate enough to work with exceptional lab members, previous and current, Drs. Dilon Fritz, and Le Cai, as well as Dr. Milica Vukmirovic, Zarko Manojlovic, and Hao Wang.

Several important parts of my dissertation were possible only due to the willingness to help and the positive minds of many of our collaborators including, Drs. Robert Singer, Amber Wells, Robert Evans, David Markovitz, and John Blackmon.

I would like to thank God above all who has been with me through the thick and the thin. My deepest thanks go to my dad (Assefa Challa) and my mom (Menen Kassa) for their love and continued support. By their principles of life, they shaped me; they taught me how to be strong, optimistic, and hopeful in the good times as well as in the bad ones. The never-give up attitude they passed to me persisted in me throughout the days I have been away from them. Their unwavering belief in me left me no option but believe in myself. It is, thus, my strongest conviction that I wouldn’t be where I am today, had it not been for these wonderful parents.

My parents blessed me with countless things, of which my siblings Kaleb, Yabetse, Perasim, Elroe, Eyasped and Herma are the six most valuable. Their true friendship and love has been a constant source of joy for me and has sustained me through the difficult times of adapting to a new culture in a new country. I am truly grateful that they are mine.

This work was supported by NIH 5R01DK059466-08 grant to B. S. and American Heart Association grant to A. C.

TABLE OF CONTENTS

List of Tables ...... vii List of Figures ...... viii Abstract ...... x 1. INTRODUCTION ...... 1 1.1- Pathogenesis of fibrosis ...... 2 1.2- Regulation of collagen expression ...... 5 1.3- Vimentin intermediate filaments ...... 9 1.4- Myocardial fibrosis ...... 11 1.5- Treatment of fibrosis ...... 13

2. NOVEL ROLE OF VIMENTIN FILAMENTS: BINDING AND STABILIZATION OF COLLAGEN MESSANGER RNAs ...... 16 2.1- Results ...... 17 2.2- Discussion ...... 39 2.3- Materials and Methods ...... 41

3. THE ANTI-FIBROTIC EFFECTS OF WITHAFERIN-A IN TISSUE CULTURE AND IN A MOUSE MODEL OF MYOCARDIAL FIBROSIS ...... 47 3.1- Results ...... 49 3.2- Discussion ...... 71 3.3- Materials and Methods ...... 76

4. GENERAL CONCLUSION ...... 82

APPENDIX ...... 84

REFERENCES ...... 87

BIOGRAPHICAL SKETCH ...... 106

LIST OF TABLES

1. Major tissues affected by fibrosis and respective diseases with fibrosis as a main feature ...... 1

2. Primers used for RT-PCR ...... 79

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LIST OF FIGURES

Figure- 1. Collagen α1(I) and collagen α2(I) mRNAs interact with vimentin in the 5’SL dependent manner...... 18

Figure-2. Collagen α1(I) and αβ(I) mRNAs co-fractionate with vimentin filaments ...... 20

Figure-3. Collagen mRNAs co-localize with vimentin intermediate filaments ...... 22

Figure-4. LARP6 dependent association of collagen mRNAs with vimentin ...... 24

Figure-5. Interaction of LARP6 with vimentin...... 25

Figure-6. LARP6 interacts with vimentin through the LA domain ...... 28

Figure-7. Decreased collagen synthesis in vimentin knock out cells ...... 30

Figure-8. Disruption of vimentin filaments by ,’- iminoditropionitrile (IDPN) reduces collagen synthesis ...... 32

Figure-9. Disruption of vimentin filaments by a dominant negative mutant of desmin reduces collagen synthesis ...... 36

Figure-10. Vimentin associates with un-translating collagen mRNAs ...... 38

Figure-11. Effect of Withaferin-A on vimentin intermediate filaments in fibroblasts ...... 49

Figure-12. Determination of toxicity of Withaferin-A in cultured fibroblasts ...... 50

Figure-13. Withaferin-A reduces expression of type I collagen in primary human and rodent fibroblasts ...... 53

Figure-14. Withaferin-A destabilizes collagen α1(I) and α2(I) mRNAs in human lung and scleroderma fibroblasts ...... 55

Figure-15. Effects of WF-A on collagen expression in vimentin knock out mouse embryonic fibroblasts ...... 57

Figure-16. Withaferin-A inhibits TGF-1 induced expression of type I collagen ...... 59

Figure-17. WF-A inhibits TGF- signaling ...... 62

Figure-18. Effect of WF-A on quiescent and partially-activated primary rat HSCs ...... 66

Figure-19. Withaferin-A inhibits isoproterenol induced myocardial fibrosis in mice...... 67

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Figure-20. Withaferin-A inhibits expression of α-smooth muscle actin and type I and type III collagens in isoproterenol induced cardiac fibrosis ...... 69

ABSTRACT

The 5’ stem-loop (5’SL) in the 5’ UTR of collagen α1(I) and αβ(I) mRNAs is the key element regulating stability and translation of collagen mRNAs. LARP6 is the RNA binding protein which exhibits high affinity binding to the 5’SL of collagen α1(I) and αβ(I) mRNAs. In the first part of the dissertation, we report that vimentin filaments associate with type I collagen mRNAs in a 5’SL and LARP6 dependent manner. This association is needed for stabilization of type I collagen mRNAs. Our conclusion was based on the following lines of evidence. First, RNA immunoprecipitation and cellular fractionation experiments showed that collagen α1(I) and αβ(I) mRNAs exhibit specific interaction with vimentin intermediate filaments. This was substantiated by RNA-FISH experiments which showed that collagen I mRNAs colocalize with vimentin filaments. Second, we showed that the binding of collagen mRNAs to vimentin intermediate filaments is dependent on the 5’ stem-loop. Collagen mRNAs from mouse embryonic fibroblasts carrying a knock-in mutation in the 5’SL failed to interact with vimentin intermediate filaments. Third, interaction of collagen mRNAs with collagen mRNAs was mediated by the RNA binding protein, LARP6, which specifically binds the 5’SL of type I collagen mRNAs. siRNA knock down of LARP6 abrogated the interaction of type I collagen mRNAs with vimentin intermediate filaments. We also found that LARP6 interacts and colocalizes with vimentin intermediate filaments. Mapping of the domain of LARP6 needed for interaction with vimentin revealed that the La domain of LARP6 is necessary and sufficient to interact with vimentin. Fourth, disruption of vimentin filaments using the vimentin disrupting drug ,’-imminodipropionitrile or by expressing a dominant negative intermediate filament markedly reduced production of type I collagen. The reduction in collagen synthesis was due to decreased stability of collagen mRNAs. Last, but not least, we observed a marked reduction in collagen synthesis from vimentin -/- mouse fibroblasts. This reduced collagen production was also due to reduced stability of type I collagen mRNAs in vimentin deficient fibroblasts. This was consistent with the impaired wound healing phenotype displayed by vimentin deficient mice further verifying the important role of vimentin filaments in collagen synthesis. We concluded that vimentin intermediate filaments play a key role in the development of tissue fibrosis by stabilizing type I collagen mRNAs. This finding served as a basis for targeting vimentin in the investigation of a novel anti-fibrotic therapy.

In the second part of the project, we examined the antifibrotic effects of Withaferin-A. The intermediate filament vimentin is the primary target of Withaferin-A. In light of our finding of the role of vimentin in stabilizing collagen mRNAs, we hypothesized that Withaferin-A may reduce collagen production by disrupting vimentin filaments and decreasing the stability of collagen mRNAs. We thus aimed to determine if Withafrein-A exhibits anti-fibrotic properties in vitro and in vivo and to elucidate the molecular mechanisms by which WF-A exerts its antifibrotic effects. We found that, in tissue culture, Withaferin-A suppresses collagen expression, both at transcriptional and post-transcriptional level, by inhibiting the TGF- signaling pathway and by disrupting vimentin filaments, respectively. Withaferin-A disrupted vimentin filaments, and caused degradation of vimentin in fibroblasts. The toxicity of Withaferin-A is not due to disruption of vimentin filaments as the disruption occurs at concentrations lower than the toxic range. WF-A can potently inhibit the expression of type I collagen in human and rodent fibroblasts. Withaferin-A increases the rate of decay of α1(I) and α2(I) collagen mRNAs. The effect of WF-A on half-life of collagen mRNAs is dependent on the presence of vimentin. In addition to destabilizing collagen mRNAs by disrupting vimentin filaments, WF-A also interferes with TGF- induced transcription of collagen genes. WF-A inhibits both the TGF- induced as well as the culture induced phosphorylation of Smad3. Withaferin-A inhibits in vitro activation of hepatic stellate cells (HSCs) and decreases collagen production by HSCs. In vivo, WF-A inhibits isoproterenol-induced myocardial fibrosis and results in downregulation type I and type III collagen as well as α-smooth muscle actin. Our findings provide a strong evidence base for the further exploration of Withaferin-A as a therapeutic drug against fibroproliferative diseases, including but not limited to cardiac interstitial fibrosis.

Taken together, this dissertation project explores both the basic science and translational aspect of the posttranscriptional regulation of collagen expression by a complex involving vimentin intermediate filaments, LARP6 and the 5’SL of collagen mRNAs. The findings strongly show that, in this complex, vimentin confers stability to collagen mRNAs. Importantly, the dissertation utilizes the collagen stabilizing function of vimentin for targeting by a potential ant-fibrotic drug. We discovered for the first time that Withaferin-A displays anti-fibrotic properties in vitro and in vivo in mouse model of isoproterenol-induced myocardial fibrosis.

CHAPTER ONE

INTRODUCTION

Nearly 45% of all deaths in the developed world are attributed to some type of chronic fibroproliferative disease [1]. Fibrosis affects all major organs (Table 1). Liver fibrosis is a global health problem resulting mainly from viral hepatits, chronic alcohol consumption and complication of obesity [2]. Cardiac fibrosis is an important hallmark of maladaptive hypertrophy and heart failure in hypertensive heart disease and several other cardiac diseases [3]. Interstitial fibrosis of the kidney is a common pathway and a strong predictor of clinical outcome in progressive renal diseases leading to end-stage renal failure regardless of the etiology. [4]. Scleroderma (SSc) is a severe systemic disease affecting the skin and several internal organs; it has a worldwide distribution and estimated 75,000–100,000 individuals in the United States have SSc [5]. Idiopathic pulmonary fibrosis (IPF) is the most common idiopathic interstitial disease with a prevalence of 500,000 individuals in United States and Europe [6]. IPF is a fatal disease with the median survival for patients with IPF being less than 3 years [7].

Table 1: Major tissues affected by fibrosis and respective diseases with fibrosis as a main feature (adapted from [8]) Tissue Diseases with fibrosis as a major feature

Liver Cirrhosis, chronic viral hepatitis, alcoholic liver disease, schistosomiasis, Nonalcholic steatohepatitis Lung Idiopathic pulmonary fibrosis, sarcoidosis, silicosis, collagen vascular diseases including rheumatoid arthritis and systemic sclerosis Kidney Diabetic nephropathy, Chronic glomerulonephritis, Tubulointerstitial nephritis, Hepertensive nephropathy Heart Post MI myocardial fibrosis, Hypertensive heart disease, Valvular heart diseases, chronic ischemic heart disease, the cardiomyopathies Vessels Atherosclerosis, hypertension, restenosis Eye Age related macular degeneration, diabetic retinopathy, vitreoretinopathy, corneal opacification Skin Scleroderma, keloids and hypertrophic scars, burn contractures Pancreas Chronic alcoholic pancreatitis, autoimmune pancreatistis, obstructive chronic pancreatitis Intestine Crohn’s disease Other Alzheimer’s disease, AIDS (brain); ageing, malignancies (bone marrow); PID. 1

1.1- Pathogenesis of tissue fibrosis

Fibrosis can be defined as a wound healing response that has gone out of control [1]. Tissue damage is immediately followed by the repair process with the intent to heal the damaged tissue. The normal wound healing process normally involves three major phases [9]. The first phase is the inflammatory phase, where inflammatory cells such as neutrophils and monocytes are recruited to the area of tissue damage primarily for their innate immunity related functions including phagocytosis of cell debris and invading organisms. Inflammatory cells, particularly macrophages and activated T lymphocytes produce a host of cytokines and growth factors which mediate the next phase of wound healing, the proliferative phase [10]. The proliferative phase is comprised of reepithelization, where injured cells are replaced by cells of the same type; angiogenesis, where new blood vessels sprout from preexisting ones to form the granulation tissue; and fibroplasia, where proliferation and activation of fibroblasts is followed by deposition of extracellular matrix (ECM). Myofibroblasts, often the result of activation of local fibroblasts, are the main cells depositing the ECM. Myofibroblasts also promote the process of wound contraction, setting the stage for the last phase of fibrosis [11]. The final phase of wound healing is the remodeling phase, where there is continued deposition and reorganization of ECM. Remodeling transforms the immature, transient, cell-rich matrix into a mature, permanent, cell- poor matrix [12]. Towards the end of this phase, most vessels, fibroblasts and inflammatory cells disappear from the wound site leaving a highly organized structure made of mainly the ECM, called scar [13].

The wound healing process is physiologic at first and is beneficial for restoration of any damaged tissue; however, this process can become pathologic when it continues unchecked. Pathologic wound healing often occurs in the setting of chronic inflammation and persistent tissue damage, where inflammation, tissue destruction and tissue repair occur simultaneously. Pathologic wound healing leads to fibrosis which is the end result of chronic inflammatory reactions induced by a variety of stimuli including persistent infections, autoimmune reactions, allergic responses, chemical insults, radiation and other tissue injury [8]. In such settings, the transient fibroplasia and ECM deposition observed in normal wound healing is protracted due to persistent activation of myofibroblasts. The state of persistent activation of myofibroblasts is best exemplified by the dermal fibroblasts obtained from patients with scleroderma. These fibroblasts have a constitutively activated myofibroblast-like phenotype, characterized by enhanced ECM

2 synthesis, constitutive secretion of cytokines and chemokines and increased expression of cell surface receptors [14]. In scleroderma as well as other fibroproliferative diseases, persistent activation of myofibroblasts leads to excessive deposition of ECM and extensive scar formation. Extensive scars distort the architecture of the involved organ with subsequent loss of function and eventual organ failure [1, 11, 13].

The key cellular determinants for the development of fibrosis are activated fibroblasts and myofibroblasts [15]. Profibrotic cytokines released from inflammatory cells activate fibroblasts, and activated fibroblasts transform into α-smooth muscle actin (α-SMA) –expressing myofibroblasts. Myofibroblasts serve as the primary collagen producing cells, thus, are at the centers of the pathogenesis of many fibrotic diseases [16]. In addition to activated fibroblasts, local mesenchymal cells are important sources of myofibroblasts. Hepatic stellate cells in liver are the prime examples of local mesenchymal cells that differentiate into myofibroblasts [17]. The process of HSC activation generates an α-SMA positive myofibroblast-like cells from normally quiescent vitamin A storing HSCs. Activated HSCs are proliferative and are responsible for deposition of the majority of excess ECM that forms scar tissue in the fibrotic liver [18]. Mesangial cells are specialized pericytes juxtaposed to the capillaries within the renal glomerulus. In renal diseases, glomerular mesangial cells get activated, acquire a myofibroblast phenotype and synthesize matrix proteins [19]. Myofibroblasts can also be derived from epithelial and endothelial cells which undergo epithelial-mesenchymal transition (EMT) or endothelial-mesenchymal transition (EndMT) [20-23]. Last, myofibroblasts can be generated from circulating fibroblast like cells called fibrocytes recruited from the bone marrow [24].

The key molecular determinant for the development of tissue fibrosis is TGF-. TGF- has been linked to the development of fibrosis in a number of diseases [25-28]. Several cell types produce TGF-β; the monocytes and macrophages are the predominant cellular sources. There are three isotypes of TGF- in mammals, TGF-1, β, and γ; all exhibit similar biological activity. Tissue fibrosis in several organs is primarily attributed to TGF-1. The TGF-1/Smad pathway is the primary signaling pathway for production of ECM proteins. Through this pathway, TGF- 1 increases the synthesis of ECM proteins, such as collagen, fibronectin, CTGF, and PAI-1, a protease inhibitor important for tissue remodeling [8, 29]. TGF-1 inhibits degradation of collagen by reducing production of collagenases and by stimulating the expression of tissue inhibitor of metalloproteinases (TIMP), resulting in a net accumulation of collagen [30] . TGF-

3 is also central to the activation of fibroblasts and their differentiation into myofibroblasts [11]. Furthermore, TGF-1 stimulates its own production by myofibroblasts, establishing an autocrine cycle of fibroblast activation and differentiation. This self-sustaining route, is for instance, underscored by the observation that the autocrine TGF- pathway is involved in promoting and maintaining the constitutively activated myofibroblasts found in the skin of scleroderma patients [14]. Other molecular determinants of tissue fibrosis include angiotensin II, IL4, IL-13, NFkB, PPAR, PDGF, CTGF etc. All major components of the renin-angiotensin-aldosterone system exhibit profibrotic activity; however, angiotensin II (Ang II) appears to be the dominant hormone responsible for fibrosis in the heart, blood vessels, kidney and liver [31-34]. Ang II is produced locally by activated macrophages and fibroblasts and exerts its effects directly by stimulating TGF-1 production and enhancing TGF-1 signaling through up regulation of Smadβ levels [33, 35]. For instance, in vascular smooth muscle cells (VSMCs), Ang II stimulates TGF- mRNA expression and promotes its activation from latent form. Ang II enhances TGF- expression at the level of transcription by PKC and p38 MAPK-dependent pathways. AngII also stimulates Thrombospondin-1 (Tsp-1) which in turn leads to release of active TGF- from the inactive latent complex [36]. In VSMCs and in cultured primary rat cardiac fibroblasts, it has also been reported that Ang II can activate the Smads independent of TGF-. Ang II causes a rapid Smadβ phosphorylation, nuclear translocation of phosphorylated-Smad2 and Smad4, and increased Smad DNA-binding activity. The Ang II induced activation of Smads was not inhibited by neutralizing antibody against active TGF-. Nevertheless, the Smad activating action of Ang II was inhibited by p38MAPK inhibitor suggesting that Ang II may activate the p38MAPK in TGF- independent manner [37-39]. In other cells such as the tubuloepithelial cells, only the short term effects of Ang II were independent of TGF-. Ang II dependent long-term Smad activation in the renal tubuloepithelial cells and differentiation to myofibroblasts was TGF- dependent [40]. The Th2 cytokines IL-4 and IL-13 have roles in the regulation of tissue remodeling and fibrosis. An increased level of IL-4 is found in the bronchoalveolar lavage of fluids of patients with idiopathic pulmonary fibrosis [41]. Receptors of IL-4 are found on fibroblasts and in vitro studies showed that IL-4 stimulation results in marked up regulation of the genes for ECM [42]. Inhibitors of IL-4 were found to reduce dermal fibrosis in a mouse model of scleroderma [43].

Another profibrotic cytokine is IL-13 which exploits the same IL-4Rα/Stat6 signaling pathway as IL-4. IL-13 has been identified the dominant effector cytokine in several experimental models of fibrosis [44-46]. Treatment with anti-IL-13 antibody markedly reduced collagen deposition in the lungs of bleomycin-treated mice [47]. Some studies have shown that HSC activation is associated with elevation of NFkB activity [48]. Persistent elevation of NFkB activity is observed in activated HSCs compared to quiescent cells [49]. The elevated NFkB activity is demonstrated to protect activated HSCs from TNF-α induced apoptosis [50]. In support of this, gliotoxin, inhibitor of NFkB, promotes accelerated recovery from CCl4 induced liver fibrosis in rats via its ability to induce apoptosis of activated HSCs [51]. Contrary to its role in maintaining the pool of activated HSCs, NFkB as transcription factor represses the expression of type I collagen genes. TNF-α is, for instance, reported to inhibit expression of COL1A1 by increasing the activity of NFkB [52, 53]. It is possible that in the presence of the activity of multiple transcription factor, the inhibitory effect of NFkB on collagen expression may not be seen. Expression of PPAR is dramatically reduced in activated HSCs. Several PPAR agonists are shown to block the phenotypic features of activated HSCs, including proliferation, collagen production and, α-SMA expression [54, 55]. 1.2- Regulation of collagen expression Type I collagen is the most abundant protein in human body. It is a heterotrimeric protein composed of two α1(I) polypeptides and one αβ(I) polypeptide. Excessive deposition of type I collagen is the hall mark of all fibroproliferative disorders. The net amount of collagen deposited by fibroblasts is regulated continuously by collagen synthesis and collagen catabolism. Degradation of collagen and other ECM components is controlled by various matrix metalloproteinases (MMPs) and tissue inhibitors of metalloproteinases (TIMPs). In fibrotic diseases, the synthesis of new collagen by fibroblasts exceeds the rate at which it is degraded, resulting in its excessive deposition [56, 57].

Synthesis of type I collagen is regulated at both the transcriptional and posttranscriptional levels.

1.2.1- Transcriptional regulation of type I collagen: Basal as well as cytokine induced transcription of the genes for type I collagen is driven by proximal promoter sequences which bind several ubiquitous transcription factors. The ubiquitous transcription factors involved in the

5 expression of collagen genes include Sp1, Sp3, NF1, AP1, BTEB1, KLF6, CBF1, C/EBPF, p300/CBP and c-Krox. Most of the ubiquitous transcription factors involved in constitutive expression of collagen serve for basal transcription of collagen genes in different tissues. Some of the same transcription factors, however, can also be employed by upstream profibrotic signaling cascades. TGF- enhances the expression of COL1A1 and COL1Aβ genes, through a functional interaction between Sp1, p300/CBP and the Smad3/4 complex. NF1 binds the promoter of mouse COL1A2 gene between -350 and -300bp and is required for TGF- mediated stimulation of the mouse COL1A2 gene [58-60].

Signal transduction mechanisms affecting the expression of type I play important roles in the regulation of expression of type I collagen in collagen producing cells. This signaling molecules include several cytokines, growth factors and others; they can be grouped into profibrotic (TGF-, CTGF, PDGF, FGF, IGF-1, IL-1, IL-4, IL-13), and antifibrotic (TNF-α, IL-10, IFN, PGE2 and corticosteroids). Most of these soluble modulators of collagen expression work mainly by altering transcription from collagen I genes. Cis-acting response elements in the type I collagen genes for some of the main signaling factors have been identified. Examples of known response elements for some of the profibrotic and antibrotic factors are mentioned below.

A sequence located between -378 and -183 bp in human COL1A2 promoter mediates the transcriptional effects of TGF-. This site binds a large multimeric complex containing R-Smads, Sp1, AP1, and p300/CBP [61-63]. Interleukin-4 (IL-4) induced transcriptional activator STAT6 binds to various sequences within collagen genes distributed up to -1kb for COL1A1 and up to - 3kb for COL1A2. Sequences located b/n -506 and -351 bp of the COL1A2 gene are important for corticosteroid mediated inhibition of transcription. TNF-α response elements are the same cis-acting elements that mediate the effects of TGF- [64]. IF- needs sequences between -129 and -120 to mediate its inhibitory action on collagen expression [65].

Transcription of type I collagen genes is highly cell type specific. Despite the fact that collagen type I is produced in most tissues, it is synthesized by only few discrete cell types; namely, fibroblasts, osteoblasts, odontoblasts and hepatic-stellate cells. Studies have attempted to explain the basis for the specificity of its expression in these restricted cell types. It appears that tissue specificity depends on the cis-acting enhancer elements active in different cells. COL1A1, for instance, has been found to have a modular structure with characteristic regulatory elements active in tissue specific manner. In support of this, DNA sequences between -2.3 and -1.7 kb 6 were required for the transcriptional activity of COL1A1 in bone and tooth, where as in tendons the regulatory elements involved are distributed between -1.7kb to -3.5kb [66]. In a study of transgenic mice harboring 900bp of the proximal promoter of the COL1A1, the expression of the transgene was limited only to the skin. When the length of the promoter was extended to 2.3kb, the transgene began to be expressed in osteoblasts and odontoblasts as well. Further extension to 3.2kb of the proximal promoter resulted in its expression in tendons and facial fibroblasts [67, 68].

1.2.2- Posttranscriptional regulation of type I collagen:

In addition to transcriptional regulation, ample evidence from various tissues has accrued over the last two decades that expression of type I collagen is also regulated at the posttranscriptional level [69-73]. Culturing of NIH 3T3 fibroblasts to confluence increases the half-life of collagen α1(I) from 4hrs to 9hrs, associated with the increase in collagen production. And treatment of these fibroblasts with TGF-1 increases the half-life of collagen α1(I) mRNAs several fold [74, 75]. In human lung fibroblasts, activation of the PI3K pathway led to prolongation of the half- life of collagen α1(I) mRNAs. Conversely, inhibition of PIγK activity led to decreased stability of the mRNAs [76]. Importantly, in activated HSCs, it has been shown that the posttranscriptional regulation is the predominant mechanism by which expression of collagen is up regulated. The steady state of α1(I) collagen mRNA is increased about 50 fold in activated HSCs compared to quiescent HSCs. This increase was due to 3 fold increase in transcription of the gene for collagen α1(I) and 16 fold increase in the stability of collagen α1(I) mRNA. Half- life of collagen α1(I) mRNA is increased from 1.6h in quiescent HSCs to β4hrs in activated cells [77, 78]. Taken together, these studies showed that posttranscriptional regulation is the predominant level at which expression of type I collagen is controlled.

Two sequence specific binding activities have been discovered for collagen α1(I) mRNA; α-CP in the γ’UTR and LARP6 in the 5’UTR [79]. α-CP, also known as hnRNP-E, belongs to the KH-domain family of RNA binding proteins and shuttles between the nucleus and cytoplasm. Binding of α-CP is targeted to the C-rich sequence located 23 nt downstream of the stop codon of collagen α1(I) mRNA and stabilizes the mRNA [80]. The binding of α-CP to the γ’UTR of collagen α1(I) mRNA only occurs in activated HSCs contributing to the prolonged half-life of the mRNA in the activated HSCs compared to quiescent HSCs [81]. α-CP also binds a similar C- rich sequence in the γ’ UTR of other mRNAs including α globin, 15-lipoxygenase and tyrosine 7 hydroxylase. The binding of α-CP to these mRNAs is associated with their increased stability [82, 83].

In the 5’UTR of three collagen mRNAs, α1(I), αβ(I), and α1(III), there is a stem-loop (5’ SL) structure. The 5’SL is located 75 nt from the cap and encompasses the translation initiation codon. The 5’SL is evolutionarily conserved from sea urchins, to zebrafish, to chicks, to frogs, and humans, differing by only two nucleotides in Xenopus and human collagen mRNAs [78, 84, 85]. The evolutionary conservation suggests that the 5’SL has a vital role in regulating the expression of collagen gene. The absence of the 5’SL in collagen mRNAs has been shown to impair the proper assembly of collagen polypeptides into a triple helix [86]. Additional evidence for the important role of the 5’ SL in regulating type I collagen synthesis comes from in vivo study of transgenic mice; mice that have the mutated 5’SL in the endogenous collagen α1(I) gene develop 50% less liver fibrosis than control mice [87]. Steady-state levels of α1(I) collagen mRNA was significantly decreased in MEFs and HSCs derived from mice carrying homozygous mutation in the 5’SL [87].

LARP6 binds the 5’SL of collagen α1(I) and αβ(I) mRNAs in a sequence specific manner and with high affinity. LARP6 interacts with two single-stranded regions of the 5’ stem loop [88]. The Kd for binding of LARP6 to the 5’ SL is 1.4 nM indicating the high affinity of binding. LARP6 doesn’t associate with polysomes; however, overexpression of LARP6 inhibits loading of collagen mRNAs on to polysomes. Similarly, knockdown of LARP6 resulted in significantly reduced collagen protein synthesis, which was associated with decreased loading of collagen mRNAs on to polysomes. Absence of LARP6 also led to diffuse accumulation of procollagens throughout the ER, instead of the discrete focal areas of high concentration of procollagens α1(I) and αβ(I) seen in normal fibroblasts. LARP6 was therefore needed for efficient translation of collagen mRNAs by facilitating the loading of the mRNAs onto polysomes. Additionally, the binding of LARP6 to the 5’SL of collagen mRNAs may serve to prevent the random translation of the mRNA facilitating the coordinated translation on the membrane of the ER to form the collagen heterotrimer [88].

Non-muscle myosin is LARP6 interacting protein involved in translation of collagen mRNAs [89]. In cardiac fibrosis, reexpression of the fetal form of nonmuscle myosin was found only at the sites of focal fibrosis [90]. In vitro, intact non-muscle myosin filaments are required for the synthesis of heterotrimeric type I collagen [89]. LARP6 interacts with nonmuscle myosin 8 through its C-terminal domain and associates collagen mRNAs with the filaments. Dissociation of non-muscle myosin filaments or inhibition of the motor function of myosin results in secretion of collagen α1(I) homotrimer and their increased intracellular degradation. The function of myosin filaments in collagen secretion is dependent on the presence of the 5’SL. Based on these results, it was suggested that the association of collagen mRNAs with non-muscle myosin filaments is necessary to coordinately synthesize collagen α1(I) and αβ(I) polypeptides [89]. 1.3- Vimentin intermediate filaments

In addition to non-muscle myosin, vimentin was also identified by the 5’SL RNA affinity purification as a 5’SL interacting protein [89]. Vimentin is a type III intermediate filament protein with previously unknown function in collagen expression.

Intermediate filaments (IF) are one of the three main cytoskeleton components and are composed of a large family of proteins which have common structural features. The size of their diameter (~ 12 nm) is intermediate between microtubules (~ 25 nm) and actin-microfilaments (~ 6 nm), hence the name [91]. Five different types of intermediate filaments are distinguished on the basis of biological and structural similarity: type I (acidic keratins), type II (basic keratins), type III (desmin, vimentin, peripherin, and GFAP (glial filament acidic protein), type IV (neurofilaments) and, type V (nuclear lamins). Intermediate filaments are expressed in different cell types and are often used as markers for identifying cell types. Vimentin is the major intermediate filament of mesenchymal cells highly conserved in species from Xenopus to humans [92].

Vimentin exists in cells mainly as filamentous polymers, however, a small fraction of soluble tetramers are also found in cells [93]. Similar to all intermediate filament proteins, vimentin contains a central rod domain important for assembly of the filaments, flanked by head and tail regions [94]. The head and tail regions are unique to each intermediate filament (IF) and are believed to be responsible for the special characteristics displayed by each IF protein. Vimentin interacts with structural and signaling molecules through the head and tail domains. In addition, phosphorylation of the head and tail domains is a key regulator of the dynamics of vimentin filament networks [95].

Vimentin filaments are involved in cell motility, maintenance of cell shape, organization of organelles, cell-matrix adhesions and endurance to mechanical stress in mesenchymal cells

[96, 97] [98]. Vimentin -/- mice have morphologically aberrant glial cells leading to cerebellar defect and impaired motor coordination, impaired wound healing due to defects in the capacity of fibroblasts to migrate, decreased flow induced dilation of resistant arteries reflecting their role in mechano-transduction of shear stress, disturbed homing of leukocytes to lymph nodes, and lack of integrity in vascular endothelium [99-103] . In addition to the mechanical and structural roles, recent studies show that vimentin filaments can act as organizers of signaling molecules [104]. Vimentin was identified as a -adrenergic receptor (AR) interacting protein modulating downstream Erk MAPK pathway to initiate lipolysis in adipocytes [105]. Recruitment of Src kinase by vimentin was involved in the activation of AR/Erk pathway. In another study, it has been reported that the head domain of vimentin can interact with 14-3-3 proteins in a phosphorylation dependent manner [106, 107]. Vimentin limits the ability of 14-3-3 to interact with other target molecules such as Raf, thereby modulating signaling pathways requiring 14-3- 3.

Association of mRNA with the cytoskeleton represents a fundamental aspect of mRNA physiology involved in their transport, localization, translation, and turnover [108]. Retention of mRNA by the cytoskeleton was, for instance, proposed to be a mechanism for stabilization of LDL receptor mRNAs [109]. As an important filamentous system in collagen producing cells, the possibility that vimentin intermediate filaments may have roles in regulation of collagen expression is consistent with known mechanisms of mRNA physiology. Moreover, the possible role of vimentin in synthesis of collagen is consistent with the impaired wound healing observed in vimentin knockout mice. In vimentin deficient adult mice, it was reported that wound healing is dramatically impaired at the level of fibroblast invasion into wound site and delayed wound contraction [99] [100]. It has later become evident that initial collagen deposition at the proliferative phase of wound healing is important for the process of fibroblast invasion of the wound site [110]. Nonetheless, there have been no reports that vimentin can contribute to posttranscriptional regulation of collagen expression.

1.4- Myocardial fibrosis

Myocardial fibrosis is a key pathologic feature of a number of chronic cardiovascular diseases including hypertensive heart disease, ischemic heart diseases, dilated cardiomyopathy, hypertrophic cardiomyopathy and valvular heart diseases. In HHD the extent of fibrosis is much more than in the other cardiovascular diseases, where all walls of the LV, the interventricular septum and the RV are affected by the fibrotic process. Recent works support that fibrosis could facilitate the transition from left ventricular hypertrophy to heart failure in patients with HHD suggesting fibrosis as a key prognostic indicator [111, 112]. Evidences for association between myocardial fibrosis and HHD come from several observations. The collagen volume fraction (CVF) is significantly increased in the myocardium of patients with HHD compared with that of normotensive controls. An exaggerated accumulation of collagen type I and III fibers within the myocardium has been consistently found in a number of studies analyzing postmortem human hearts and endomyocardial human biopsies [113-115]. In spontaneously hypertensive rats, transition from left ventricular (LV) hypertrophy to LV diastolic dysfunction was exclusively associated with progressive remodeling of the extracellular matrix of the myocardium [116]. The diastolic dysfunction is due to increased ventricular stiffness caused by the myocardial fibrosis [117, 118]. In patients with HHD, the degree of myocardial fibrosis correlates with deterioration of left ventricular diastolic function [119, 120]. Severe left ventricular diastolic dysfunction of the fibrosed and hypertrophied myocardium eventually leads to development of decompensated congestive heart failure [121, 122]. The excess collagen deposited in these hearts is the result of both increased types I and III collagen synthesis as well as decreased degradation by matrix metalloproteinases [35, 123]. The factors contributing to development of fibrosis in HHD are mechanical stress such as the hemodynamic overload secondary to the hypertension, humoral factors including TGF, components of the renin-angiotensin-aldosterone system (RAAS), and endothelin-1, and genetic predisposition. Hemodynamic overload of the left ventricle secondary to hypertension is an important factor contributing to the development of cardiac fibrosis in patients with HHD [123]. In vitro mechanical stress to cardiac fibroblasts as well as in vivo pressure overload of the left ventricle in mice is associated with increased collagen deposition and reduced activity of collagenases [124, 125]. The increase in the level of collagen deposited towards the inner wall of 11 the ventricle compared to the outer wall in the myocardium of HHD patients may support the gradient of mechanical stress to which the ventricle is exposed [113, 114]. It has also been reported that the CVF correlated with systolic BP and pulse pressure in patients with HHD [126]. Non-hemodynamic factors also contribute to the development of fibrosis in HHD. The important role of non-hemodynamic factors was supported by the observation that fibrosis in HHD affects, besides the left ventricle where the mechanical stress is located, the right ventricle, the interventricular septum, and the left atrium [127, 128]. Furthermore, it has been shown that the ability some hypertensive drugs to reverse fibrosis is not associated with the BP lowering effect of the drugs [129, 130]. Cytokines such as the TGF-, vasoactive substances such as endothelin 1 and effector hormones of the renin-angiotensin-aldosterone system (RAAS) are among the main humoral factors involved in development of cardiac fibrosis. The in vitro profibrotic effects of TGF- in cardiac fibroblasts have been well documented [131]. In patients with HHD, an association between increased activity of TGF- and extent of fibrosis has been reported [132]. In addition, overexpression of TGF- in transgenic mice results in cardiac hypertrophy with significant interstitial fibrosis [133]. Angiotensin II exerts multiple profibrotic effects in the heart; these include inducing proliferation of fibroblasts, promoting the differentiation of fibroblasts to myofibroblasts, activating pathways with downstream effects of increased collagen expression and inhibition of pathways of collagen degradation [134, 135]. Similarly, aldosterone, another component of the RAAS, is also shown to be independently associated with development of cardiac fibrosis [136, 137]. Last, but not least are genetic factors. Genetic markers associated with predisposition to myocardial fibrosis include the rat ACE-1. Rats carrying the B allele develop more extensive ventricular fibrosis following isoproterenol compared to rats carrying the L allele [138]. In addition, the A1166C SNP of the angiotensin II type 1 receptor gene has been found to be correlated with excessive synthesis and unaltered degradation of type I collagen in patients with HHD [139]. Recent studies suggest that monitoring circulating markers of collagen type I turnover in combination with echocardiographic assessment of myocardium may provide important diagnostic information with respect to extent of fibrosis in HHD patients [129, 140-142]. Considering improved diagnostic capabilities and based on the accumulated evidences on the relationship myocardial fibrosis and cardiac dysfunction in HHD, it is suggested that it is

12 necessary to develop new approaches aimed at correcting the imbalance in collagen synthesis and turnover in patients with HHD. Although some studies reported the antifibrotic effects of some drugs, there is currently no proven effective treatment for myocardial fibrosis. It has been shown that treatment of HHD with angiotensin converting enzyme inhibitors or angiotensin receptor blockers resulted in reduced myocardial fibrosis and improved cardiac function independent of blood pressure control and regression of hypertrophy [129, 130, 143]. Alternative potential therapeutic strategies for the treatment of experimental myocardial fibrosis included the use of inhibitors of TGF- and agonists of the peroxisome proliferator-activator receptor-α (PPAR-α). The anti-inflammatory drug tranilast and a natural inhibitor of hematopoetic stem cell proliferation, Ac-SDKP, were shown to reduce myocardial fibrosis in rats with experimental hypertension through inhibition of the TGF- pathway [144, 145]. Similarly fenofibrate, a PPAR-α agonist has been reported to reduce cardiac fibrosis in experimentally induced hypertension in rats [146-149]. 1.5- Treatment of fibrosis

Considering the fact that 45% of all deaths in developed world are attributed to some type of chronic fibroproliferative disease, there is a great demand for antifibrotic drugs that are safe and effective [1]. Several considerations need to be taken into account in the development of anti-fibrotic strategies. An important consideration is whether the antifibrotic strategy will deprive the involved tissue its ability to undergo physiologic healing. In most tissues, repair of damaged tissues solely by regeneration of parenchymal cells without scarring is not possible. Therefore, development of therapeutic strategies that limit the progression of fibrosis without adversely affecting the basic repair process would be ideal [150].

Several strategies for novel antifibrotic agents are being investigated. Perhaps the most effective antifibrotic strategy would be to cure the underlying disease process before advanced fibrosis has developed. Elimination of HBV and HCV in patients with chronic hepatitis was associated with significant regression of fibrosis [151]. Similar observations have also been made in Schistomiasis patients following treatment with praziquantel [152]. The above observations from elimination of the main causative agents are challenging the long-held notion that reversal of advanced fibrosis to restore normal architecture is impossible. Unfortunately, removing the underlying causative agent is often not possible in most fibrotic diseases, and specific antifibrotic therapies are needed [153].

Emerging antifibrotic therapies in animal models of fibrosis act by two main mechanisms: 1- inhibiting the accumulation of collagen producing cells and/or, 2- preventing the deposition of type I collagen [150]. Some of the therapies which showed promise in animal models of fibrosis include TGF-–signaling modifiers, inhibitors of IL-13, specific MMPs, adhesion molecules (e.g. integrins), and inducers of angiogenesis (e.g. VEGF), chemokine and TLR antagonists, angiogenesis inhibitors, antihypertensive drugs, inducers of apoptosis of activated fibroblasts and myofibroblasts, and stem/progenitor cell transplantation technologies [8, 35, 51, 151, 154, 155].

The evidence supporting the antifibrotic effects of TGF-β inhibitors, Ang-II inhibitors and stem cell therapies are discussed below. Numerous anti-fibrotic therapies are focused on inhibiting the TGF-β1 signaling pathway [156]. The blockage of TGF- with neutralizing antibodies [26, 157-159] or use of decorin [160], a scavenger of active TGF-, have been shown to reduce fibrosis in experimental models of renal and cardiac fibrosis. In a different approach, several preclinical studies have shown the antifibrotic effects of shRNAs against TGF-β in mice models of renal and kidney fibrosis [161, 162]. In contrast to the fact that TGF- knockout mouse are lethal, displaying a hyperinflammatory phenotype and severely impaired wound healing [163], long-term exposure of mice either to TGF-β antibody or shRNA is shown to be tolerated in mice [164]. Nevertheless, the issue of safety and effective method of delivery in clinical settings remain challenges to the above approaches. For instance, a TGF-β1 neutralizing antibody (metelimumab) has been shown to have significant adverse effects when given intravenously to patients with Scleroderma [165]. Other approaches to target TGF-β include soluble TGF-β receptors and small molecule inhibitors of TGF-β receptors. Soluble TGFβR3 has demonstrated efficacy against progression of renal fibrosis in diabetic mice [166]. TGF-βR1 small-molecule inhibitors are also showing promise for treating experimental models of liver and kidney fibrosis [167, 168]. Although the specificity of small molecule inhibitors of TGF-β needs further study, the use of these small molecules can circumvent the challenges of safe delivery seen in the other approaches. In all the approaches to interfere with TGF-β, it is clear that more inputs from clinical studies are highly needed. Angiotensin antagonism attenuated TGF- secretion from cardiac fibroblasts and development of myocardial fibrosis [169]. The inhibition of Ang II type I receptors diminishes

14 the activation of the Smad pathway in post MI myocardial fibrosis in rats, in hypertension induced vascular fibrosis, and in experimental models of renal and hepatic fibrosis [34, 37, 39, 170]. Therapies that target the renin-angiotensin-aldosterone system are therefore suggested to be potential strategies to slow the progression of fibrosis in HHD, progressive renal disease and hepatic fibrosis [35, 171]. In agreement with the findings in the experimental models, angiotensin blockers have demonstrated beneficial effects in the treatment of a number of cardiac diseases associated with fibrosis in humans [169]. Last, stem cell therapies have proved successful at restoring cardiac function in fibrosed hearts [155]. Translating the available experimental information into clinically effective drugs is hindered by several hurdles. Some of the obstacles include the invasive nature of some of the treatments in terms of their wider targets, for example TGF-β inhibitors, and lack of well developed non-invasive techniques to monitor progress of fibrosis.

In the subsequent chapters of this dissertation, readers find the description of the novel role of vimentin filaments in the binding and stabilization of collagen mRNAs (chapter-two) and the application of this novel role for the treatment of fibroproliferative diseases (chapter-three). In the last chapter, general conclusion summarizing the important insights gained from the studies is presented.

CHAPTER TWO

A NOVEL ROLE OF VIMENTIN FILAMENTS: BINDIING AND STABILIZATION OF COLLAGEN MESSANGER RNAS

Fibroproliferative disorders are leading causes of morbidity and mortality globally [2] [172] [1] [173] and pose an enormous threat to human health. There are no effective therapies for fibrotic diseases. Excessive production of type I collagen by activated fibroblasts and myofibroblasts is the hallmark of fibroproliferative disorders. Type I collagen is a heterotrimer composed of two α1(I) chains and one αβ(I) chain [174]. The increased collagen synthesis can be due to the increased rate of transcription of collagen genes, increased half-life of collagen mRNAs and their enhanced translation [77] [175] [80, 176]. Increased collagen production by activated fibroblasts is primarily due to an increase in stability of collagen mRNAs [70]. During activation of hepatic stellate cells, which synthesize collagen in liver fibrosis, a 16-fold prolongation of the half-life of collagen α1(I) mRNAs is primarily responsible for its 50 fold increased expression [177]. The transformation of fibroblasts into myofibroblasts is also associated with increased stability of collagen α1(I) mRNA [178]. TGF-beta, the most potent profibrotic cytokine, induces collagen synthesis by prolonging the half life of collagen α1(I) mRNA [175]. Thus, it is now well established that stability of collagen mRNAs is the predominant mechanism regulating collagen expression [76, 179]. Two cis-acting elements were implicated in regulating stability of collagen mRNAs. In the γ’ untranslated region (γ’UTR) of collagen α1(I) mRNA, there is a cytosine rich sequence that interacts with αCP protein; this interaction stabilizes the mRNA [80, 180]. In the 5’ UTR of collagen α1(I) and αβ(I) mRNAs, there is a stem-loop sequence (5’SL) [84] that also regulates stability and translation of collagen mRNAs [85] [88] [89] [78]. The 5’SL is located 75-85 nucleotides from the cap and includes the translation initiation codon. It is well conserved in evolution, differing only by 2 nucleotides in Xenopus and human collagen mRNAs. LARP6 is the protein which specifically binds the 5’SL with high affinity and is required for high expression levels of type I collagen [88]. LARP6 interacts with non-muscle myosin; this interaction is required for coordinated synthesis of α1(I) and αβ(I) polypeptides and their productive folding into heterotrimeric type I collagen[89]. Disruption of non-muscle myosin

16 filaments resulted in secretion of homotrimer of type I collagen composed of three α1(I) chains in lung fibroblasts or in total lack of secretion of type I collagen in scleroderma fibroblasts [89].

Vimentin is a member of type III intermediate filaments and is a marker of cells of mesenchymal origin (fibroblasts and myofibroblasts). Vimentin intermediate filaments are polymers of soluble tetrameric vimentin. Vimentin filaments are involved in motility, maintenance of cell shape, and endurance to mechanical stress of mesenchymal cells [96] [97]. Vimentin deficient mice exhibit a delayed wound healing; the phenotype was attributed to decreased motility of vimentin-null fibroblasts [99]. There have been no reports that vimentin can contribute to post-transcriptional regulation of collagen expression.

The present study describes a novel role of vimentin in stabilization of collagen mRNAs. This stabilization is dependent on the 5’SL and on binding of LARP6 and may contribute to the high level of collagen expression by mesenchymal cells.

2.1- Results

Vimentin specifically interacts with collagen mRNAs in a 5’stem loop dependent manner

In a previous study we observed that vimentin copurified with the 5’SL of collagen mRNA in a RNA-affinity purification method [89]. Therefore, we investigated if collagen mRNAs associate with vimentin filaments in vivo using an RNA-immunoprecipitation assay and cellular extracts prepared by hypotonic and detergent lysis. Vimentin is present mostly in the insoluble cellular fraction (see later), but the combination of hypotonic and detergent lysis resulted in presence of significant amounts of vimentin in the soluble extract (Fig 1D). This allowed us to do the immunoprecipitation with anti-vimentin antibody and analyze collagen mRNAs in the precipitate by RT-PCR (Fig 1). The immunoprecipitation with an anti- vimentin antibody in human lung fibroblasts pulled down both collagen mRNAs, α1(I) and αβ(I), but not fibronectin mRNA (Fig. 1A, lane 1).

A 1 2 3 4 B 1 2

COL α I COL α I

FIB FIB

IP: VIM CON VIM CON MEFS: +/+ -/-

C 1 2 D

COL α I VIM

TUB COL α I

FIB ACT

MEFS: +/+ -/- FRAC: EXT PEL

Figure 1. Collagen α1(I) and collagen α2(I) mRNAs interact with vimentin in the 5’SL dependent manner. A. Pull down of collagen mRNAs with an anti-vimentin antibody. Immunoprecipitation with an anti-vimentin antibody (VIM) followed by RT-PCR analysis of collagen α1(I), collagen α2(I) and fibronectin (FIB) mRNAs in human lung fibroblasts (lane 1) and scleroderma fibroblasts (lane 3). Lanes 2 and 4; control antibody (CON). Radio-labeled PCR products are indicated. B. 5’SL dependent interaction of collagen mRNAs with vimentin. Experiment as in A except extracts of wild type (lane 1, +/+) and mutant mouse embryonic fibroblasts (lane 2, -/-), which carry a mutation of the 5’-stem loop in collagen α1(I) gene, were used. C. Total RNA of wild type (lane 1, +/+) and mutant (lane 2, -/-) mouse embryonic fibroblasts analyzed by RT-PCR for collagen α1(I), collagen α2(I) and fibronectin mRNAs. D. Presence of vimentin in the cell extract prepared by hypotonic and detergent lysis. Extract (EXT) and insoluble pellet (PEL) obtained after cell lysis were analyzed for vimentin, actin and tubulin by western blot.

Neither collagen nor fibronectin mRNAs were pulled down using a control antibody (Fig 1A, lane 2), indicating the specific association of collagen mRNAs with vimentin. To confirm this result in another collagen producing cell, we repeated the experiment in primary scleroderma skin fibroblasts. Again, collagen α1(I) and αβ(I) mRNAs were pulled down with vimentin, suggesting that this interaction could be a general feature of collagen producing cells (Fig1A, lanes 3 and 4).

Since vimentin was initially discovered in a complex copurifying with the 5’SL of collagen mRNAs, we determined if the interaction between vimentin and collagen mRNAs depends on the 5’SL. To this end, we utilized mouse embryonic fibroblasts (MEFs) derived from knock-in mice in which the 5’SL of collagen α1(I) mRNA was mutated (denoted here 5’SL-/- MEFs) [87]. Immunoprecipitation with anti-vimentin antibody using extracts from wild type MEFs (5’SL+/+) pulled down collagen α1(I) and αβ(I) mRNAs (Fig. 1B, lane1). However, only trace amounts of collagen mRNAs were pulled down in 5’SL-/- MEFs (Fig1B, lane2). Fibronectin mRNAs were not precipitated in either cell types. Total steady state levels of collagen and fibronectin mRNAs from the wild type and mutant cells are shown in Fig 1C. The difference in total mRNA levels between 5’SL+/+ and 5’SL-/- MEFs was 2-fold, this difference cannot account for the dramatic difference in the pull down of collagen mRNAs (Fig 1B). In these experiments, collagen αβ(I) mRNA contained the wt 5’ SL while only the 5’ SL of collagen α1(I) was mutated. Still, collagen αβ(I) mRNA failed to interact with vimentin. This suggests that binding of collagen αβ(I) mRNAs to vimentin is coupled to the 5’ SL dependent binding of collagen α1(I) mRNA to vimentin. Collagen mRNAs are enriched in cellular fractions containing vimentin filaments Vimentin exists in the cell in two forms: as soluble tetramers and insoluble filaments [183]. 80% of the cellular vimentin is in the form of filaments and 20% is soluble. In the filamentous form vimentin is one of the most insoluble proteins, precipitating in buffers of high ionic strength and high concentrations of nonionic detergents [184, 185]. Precipitation in these buffers can separate filaments from the soluble form. To assess which form of vimentin binds collagen mRNAs, we fractionated the cell extract into vimentin filament-rich and vimentin filament-poor fractions using high salt and nonionic detergent lysis. This method has been established before for separating intermediate filaments from other soluble cellular components [186, 187]. The fractions were analyzed for the presence of vimentin, tubulin, non-muscle myosin, actin, fibronectin and LARP6 by western blot (Fig 2). Most of the vimentin was found in the insoluble fraction (Fig. 2A, lane 1) whereas tubulin, actin, non-muscle myosin and fibronectin were all exclusively found in the soluble fraction (Fig. 2A, lane 2). Significant amount of LARP6 was found in the insoluble fraction. The soluble and insoluble fractions were then analyzed for the relative abundance of collagen α1(I) and αβ(I) mRNAs by RT-PCR. Actin, GAPDH and fibronectin mRNAs were analyzed as controls. Figure 2B shows that collagen α1(I)

19 and αβ(I) mRNAs were predominantly located in the insoluble fraction (lane 1), whereas actin, GAPDH and fibronectin mRNAs were enriched in the soluble fraction (lane 2). Preferential localization of collagen mRNAs in the detergent insoluble fraction containing vimentin filaments suggests that the filamentous vimentin is the main form involved in the interaction with collagen mRNAs.

A B C 1 2 3 4 1 2 1 2

VIM COL I α COL 1A1

COL 1A2 TUB COL I α

GAPDH MYO FIB IP: VIM CON VIM CON ACT ACT FRACT : INS SOL FIB FIB FRAC: INS SOL LARP6

FRAC: INS SOL

Figure 2. Collagen α1(I) and α2(I) mRNAs co-fractionate with vimentin filaments. A. Insoluble vimentin representing filaments (INS, lane 1) and soluble vimentin (SOL, lane 2) were fractionated by detergent lysis and centrifugation. The fractions were probed for presence of vimentin (VIM), tubulin (TUB), myosin IIB (MYO), actin (ACT), fibronectin (FIB) and LARP6 proteins by western blot. B. Collagen mRNAs segregate with insoluble vimentin. The fractions in A were analyzed by RT-PCR for collagen α1(I) , collagen αβ(I), GAPDH, actin (ACT) and fibronectin (FIB) mRNAs. Pull down of collagen mRNAs with an anti-vimentin antibody. C. Collagen mRNAs in insoluble fraction co-immunoprecipitate

with vimentin. Insoluble fraction (INS) was re-dissolved and immunoprecipitation with vimentin antibody (lane 1) or control antibody (lane 2) was performed. Lanes 3 and 4; the same experiment using the soluble fraction (SOL). The immunoprecipitated material was analyzed by RT-PCR for presence of collagen α1(I), collagen αβ(I) and fibronectin mRNAs.

In order to further verify the association of the filamentous form of vimentin with collagen mRNAs, we performed RNA-immunoprecipitation experiment from the detergent- soluble and detergent insoluble fractions. To this goal, we re-dissolved the insoluble fraction in buffer containing 0.1% SDS. The immunoprecipitation with anti-vimentin antibody from the detergent-insoluble fraction of human lung fibroblast pulled down collagen α1(I) and αβ(I) mRNAs, but not fibronectin mRNA (Fig. 2C, lane 1). Neither collagen nor fibronectin mRNAs were pulled down using a control immunoprecipitation with no antibody (Fig 2C, lane 2). However, immunoprecipitation from the detergent-soluble fraction pulled down only trace amount of collagen α1(I) and αβ(I) mRNAs (Fig βC, lane γ). This suggests that collagen α1(I) and αβ(I) mRNAs found in the insoluble fraction (Fig 2B) are associated with vimentin filaments. Visualization of the interaction between vimentin filaments and collagen α1(I) and αβ(I) mRNAs. To directly show the association of collagen mRNAs with vimentin filaments in vivo, we utilized RNA fluorescence in situ hybridization (RNA-FISH). This method can detect single mRNA molecules in the cytoplasm [188-192]. To visualize collagen mRNAs, we designed multiple short 50-mer oligonucleotide probes that specifically hybridize with their target mRNAs. Each probe was labeled with multiple fluorophores to increase the signal to noise ratio for detection of individual mRNAs: collagen α1(I) probes were labeled with Cyγ and αβ(I) probes with Cy5. RNA-FISH experiments using oligonucleotide probes specific for collagen α1(I) mRNA detected multiple spots distributed both in the nucleus and the cytoplasm of human lung fibroblasts (Fig 3A, upper images). The probes spanned the exons of collagen mRNAs and the signals in the nucleus may be from unspliced mRNA. The cytoplasmic signals appear more concentrated around the perinuclear region, but they can be seen throughout the cytoplasm. The bottom panels of Fig 3A show RNA-FISH using oligonucleotide probes specific for collagen αβ(I) mRNA using a different cell than that shown in the upper panels. The distribution of RNA- FISH signals was similar to what was seen for collagen α1(I) mRNA. To verify that our probes specifically detected collagen mRNAs, we stained HeLa cells, which do not express type I collagen. In HeLa cells neither set of probes showed signal (Fig 3B), indicating that they specifically detected collagen α1(I) and αβ(I) mRNAs.

A Cy5COL α1 DAPI +Cy5COL α1 B Cy5COL α1 DAPI +Cy5COL α1

Cy3-COL α2 DAPI+Cy3-COL α2 Cy3COL α2 DAPI+Cy3COL α2

C DAPI+COL α1 DAPI+VIM DAPI+VIM+COL α1

DAPI+VIM+Cy5COL α1 (3X) VIM/COL α1 COLOCALIZATION COL α1 +COLα1/VIM COLOCALIZATION

D DAPI+Cy5COL α2 DAPI+VIM DAPI + VIM+Cy5COL α2

DAPI+VIM+Cy5COL α2 (3X) VIM/COL α2 COLOCALIZATION COL α2 +COLα2/VIM COLOCALIZATION

Figure 3. Collagen mRNAs co-localize with vimentin intermediate filaments. A. Visualization of collagen α1(I) mRNA (top panels) and collagen αβ(I) mRNA (bottom panels) by RNA-fluorescence in situ hybridization (RNA-FISH). Collagen α1(I) specific probes (labeled with cy5, red) and collagen α2(I) specific probes (labeled with cy3, green) were hybridized to human lung fibroblasts. B. The same experiment in HeLa cells, which do not express collagen α1(I) and α2(I) mRNAs. C. Co-localization of collagen α1(I) mRNA with vimentin. RNA-FISH for collagen α1(I) mRNA (upper left panel), immunostaining of vimentin (upper middle panel) and the overlaid image (upper left panel). 3x magnification of the selected area of the overlaid image is shown in bottom left. The yellow dots indicate co-localization. The image showing only the α1/VIM colocalization is in the bottom middle panel. This image is overlaid with total signal for α1(I) mRNA (upper left) to estimate the fraction of colocalized mRNA and shown in bottom right. Bars indicate 1 µ. D. Co- localization of collagen αβ(I) mRNA with vimentin. The same experiment as in C except collagen αβ(I) mRNA was visualized.

Next, we determined if collagen mRNAs co-localize with vimentin filaments by coupling RNA-FISH with immunostaining for vimentin protein (IF-RNA-FISH). In these experiments, we labeled collagen αβ(I) mRNA with cy5 oligonucleotide probes to avoid any bleed-over of the vimentin immunostaining in the cy2 channels. Figure 3C, upper right panel, shows that a significant proportion of collagen α1(I) mRNAs appears to be co-localized with vimentin filaments. The areas of co-localization (yellow) are seen both around the nucleus and throughout the cytoplasm (high magnification image shown in lower left panel). By overlaying the yellow signal of vimentin associated α1(I) mRNA (lower middle panel, Fig γC) with the total signal for α1(I) mRNA (red, upper left panel, Fig γC), we estimated that about γ0% of collagen α1(I) mRNAs are located at or in close proximity to vimentin intermediate filaments (lower right panel, Fig 3C). Similar results were obtained analyzing multiple cells. Co-localization of αβ(I) mRNA and vimentin is shown in Fig. γD. Collagen αβ(I) mRNA was also co-localized with vimentin filaments; it appears that a higher fraction of this mRNA is found associated with filaments than what was seen for α1(I) mRNA (upper right panel, Fig γD). The areas of co-localization were distributed throughout the cytoplasm and some areas clearly show beads-on-a-string appearance, indicating the association with filaments (higher magnification image, lower left panel, Fig γD). The proportion of αβ(I) mRNAs co-localized with vimentin filaments was estimated to be about 50% (Fig 3D, lower right panel). Thus, the IF- RNA-FISH results provided direct evidence for the in vivo interaction between collagen α1(I) and αβ(I) mRNAs and vimentin filaments. 23

Association of vimentin with collagen mRNAs is LARP6 dependent The experiments in the previous sections could not distinguish whether the interaction between collagen mRNAs and vimentin is direct, or mediated by protein-protein interactions. LARP6 is the protein which directly binds 5’SL [88] and, since collagen mRNAs interact with vimentin in the 5’ SL dependent manner, LARP6 may be bridging collagen mRNAs to vimentin filaments. To assess the role of LARP6, we knocked-down LARP6 using siRNA [88] and analyzed the association of vimentin with collagen mRNAs. The siRNA was able to deplete LARP6 to 30% of the initial level (Fig. 4A) without drastically affecting the expression of collagen mRNAs as assessed from RT-PCR analysis of their mRNA input levels (Fig. 4C).

A B C 1 2 1 2 1 2 COL(I)α1 COL(I)α1 LARP6 COL(I)α2 COL(I) 2 α FIB FIB FIB siRNA: CON LARP6 siRNA: CON LARP6 siRNA: CON LARP6 Figure 4. LARP6 dependent association of collagen mRNAs with vimentin. A. Knock down of LARP6. Control siRNA (lane 1) and LARP6 specific siRNA (lane 2) were expressed in human lung fibroblasts and the level of LARP6 determined by western blot. Fibronectin (FIB) was analyzed as loading control. B. LARP6 knock down reduces association of collagen mRNAs with vimentin. IP with an anti-vimentin antibody was performed in cells expressing control siRNA (lane 1) or LARP6 specific siRNA (lane 2).

Collagen α1(I), collagen α2(I) and fibronectin (FIB) were analyzed in the immunoprecipitate by RT-PCR. C. Analysis of the input levels of mRNAs. Total mRNA before the IP was analyzed for expression of collagen α1(I), collagen α2(I) and fibronectin mRNA by RT-PCR.

When RNA-immunoprecipitation was done with anti-vimentin antibody in LARP6 depleted cells, it pulled down less collagen α1(I) and αβ(I) mRNAs compared to the amount pulled down from cells with control siRNA (Fig. 4B). Considering that not all LARP6 could be depleted (Fig 4A), this suggested that the interaction of collagen mRNAs with vimentin filaments is LARP6 dependent. LARP6 binds vimentin.

To determine if there is a protein-protein interaction between vimentin and LARP6, we performed immunoprecipitation experiments. The anti-vimentin antibody pulled down endogenous LARP6 (Fig. 5A, lane 1), while control antibody had no effect (lane 2). The control protein, fibronectin, was not pulled down suggesting that this interaction is specific for LARP6. We also assessed if the interaction between vimentin and LARP6 is dependent on intact RNA. Treatment with RNase did not release LARP6 from the immunoprecipitate, suggesting that intact RNA is not needed for the interaction between vimentin and LARP6 (Fig. 5A, lane 3).

A 1 2 3 4 B 1 2 3 4

LARP6 VIM IP: HA- HA- HA- HA- LARP6 CON LARP6 CON FIBRO RNASE: -- + + IP: VIM CON VIM CON RNASE: -- + + LARP6

CON VIM VIM

HA- HA- LARP6 CON

C LARP6 LARP6 + VIM

VIM LARP6 + VIM (3X)

Figure 5. Interaction of LARP6 with vimentin. A. Pull down of endogenous LARP6 with anti-vimentin antibody. Immunoprecipitation from lung fibroblasts with anti-vimentin antibody (lanes 1 and 3) or control antibody (lanes 2 and 4). Immunoprecipitates were left untreated (lanes 1 and 2) or treated with RNase A (lanes 3 and 4) prior to washing and western blot analysis with anti-LARP6 antibody and anti-fibronectin antibody. Bottom panel; input levels of vimentin. B. Reverse experiment using HA-tagged LARP6. HA-tagged LARP6 (lanes 1 and 3) or control HA-tagged protein (lanes 2 and 4) were expressed in HEK293 cells and IP performed with anti-HA antibody without (lanes 1 and 2) or with (lanes 3 and 4) RNase A digestion. The immunoprecipitate was analyzed by western blot using anti- vimentin antibody. Bottom panel; the input levels of transfected proteins and endogenous vimentin. C. Co-localization of LARP6 with vimentin in lung fibroblasts. Immunostaining for vimentin (green, upper left panel) and endogenous LARP6 (red, upper right panel) and the merged image (lower left panel). The areas of co-localization are yellow. Lower right panel; higher magnification of the same image.

To further investigate the interaction between LARP6 and vimentin, we overexpressed HA-tagged LARP6 in HEK293 cells and performed immunoprecipitation with anti-HA antibody. Anti-HA antibody pulled down vimentin in LARP6 transfected cells but not in the cells transfected with the HA-tagged control protein, RBMS3 [193] (Fig. 5B, lanes 1 and 2). Treatment with RNase had no effect on the pull down of (Fig. 5B, lane 3). To determine if there is cellular co-localization of vimentin and LARP6, we performed double immunostaining using anti-LARP6 and anti-vimentin antibodies. As previously reported [88], LARP6 exhibited nuclear and cytoplasmic localization (Fig 5C, upper left), whereas vimentin staining was restricted to the cytoplasm (Fig. 5C, lower left). The merged image of these two channels showed a significant degree of co-localization of LARP6 and vimentin filaments (Fig. 5C, right panels). This is consistent with the result of cofractionation of LARP6 into the insoluble cellular material containing vimentin filaments (Fig. 2A). La-domain of LARP6 is required for its interaction with vimentin LARP6 has four domains; the N-terminal domain, the La homology domain [194], an RNA recognition motif (RRM) and the C-terminal domain. The La domain and RRM are needed for binding 5’ SL [88], while the C-terminal domain is required for binding to non-muscle myosin [89]. To determine which domain of LARP6 is involved in the interaction with vimentin, we constructed a series of deletion mutants of LARP6 and evaluated their interaction with vimentin in pull down assays. A schematic representation of full-size LARP6 and LARP6

26 mutants is shown in Fig 6a. The constructs were tagged with an HA-affinity tag and transfected into HEK293 cells. Western blots showed similar expression levels of all LARP6 mutants, except mutant E which was expressed at a lower level. The input levels of vimentin were also comparable (Fig. 6B, lower panel). When immunoprecipitation was performed with an anti-vimentin antibody (Fig. 6B, upper panel), the wild type LARP6 (FS) was pulled down efficiently. Similar co- immunoprecipitation of mutants A and B, which lack the C-terminal and the RRM, respectively, suggested that these domains are not needed for interaction with vimentin. The N-terminal deletion mutants, D and F, also coimmunuprecipitated with vimentin, suggesting that the N- terminus is also dispensable for the interaction. Mutant E, which lacks both the N-terminal and the C-terminal domain showed a similar interaction, albeit the amount pulled down was less which is expected given its lower expression relative to the other mutants. However, mutant C (containing only the C-terminal domain) and mutant G (containing only the RRM-domain), failed to interact with vimentin. Therefore, it appears that all the mutants which contained the La- domain of LARP6 (A, B, D, E,F, H and I) interacted with vimentin, while the mutants lacking the La- domain (C and G) failed to do so. This suggests that the La-domain of LARP-6 is required for the interaction with vimentin. Finally, we expressed the La domain only (Fig 6A, construct I) and performed the immunoprecipitation. The La domain was efficiently pulled down with vimentin (Fig 6D, lane 3), verifying that the La-domain is necessary and sufficient for the interaction.

A N-TER LA RRM C-TER 1 85 183 296 491 FS 1 300 A 1 218 B 296 491 C 46 300 D

85 300 E

1 45 81 300 F

183 300 G 1 45 81 218 H 85 183 I

B C D

1 2 3 4 5 6 7 8 1 2 3 1 2 3

MUT: FS A B C D E F G MUT: FS G H MUT: FS C I IP: VIM IP: VIM IP: VIM

VIM

FS FS FS A F B D C E C H I G G

Figure 6. LARP6 interacts with vimentin through the LA domain. A. Schematic representation of the constructs used. FS, full-length LARP6. The domains of LARP6 are indicated, with amino-acid numbering on the top. A-I, different deletion mutants of LARP6. All proteins had HA-tag at the N-terminus. B. Interaction of FS LARP6 and deletion mutants with vimentin. Top panel, FS LARP6 (lane 1) and the deletion mutants (lanes 2-8) were expressed in HEK293 cells. IP was done with anti-vimentin antibody and western blot with anti-HA antibody. Bottom panel; expression of the proteins in the input material. C. Interaction of the extended LA domain with vimentin. Construct containing the intact La domain with additional amino-acids at the N-terminus (H, lane 3) was analyzed for interaction with vimentin by IP. Lane 1, FS LARP6 as positive control; lane 2, mutant G as negative control. Bottom panel; expression of the proteins in the input material. D. Interaction of the La domain only with vimentin. Construct I, containing only the La domain, was analyzed for IP with vimentin (lane 1). Lane 2, construct C as negative control. Bottom panel; expression of the proteins in the input material.

Vimentin knock out mouse embryonic fibroblasts have decreased collagen synthesis Since LARP6 is needed for high collagen expression [88] and LARP6 and vimentin interact, we determined if collagen expression is altered in cells lacking vimentin. To this end, we measured collagen production from embryonic fibroblasts (MEFs) of vimentin knockout (vim-/-) mice and wild type (vim+/+) mice. (Vim+/+ and Vim-/- MEFs were kind gifts from Dr. Robert Evans, University of Colorado Health Sciences Center). Figure 7A shows that the cellular and the secreted levels of collagen α1(I) polypeptide were decreased 60% , while the levels of αβ(I) polypeptide were decreased 70% (Fig 7A, lanes β and 4; bottom panel). Fibronectin levels in these two cell types were similar, suggesting a specific reduction in collagen expression. The decreased level of collagen protein in vimentin deficient MEFs may be due to either the decreased mRNA expression, decreased translation or increased protein turnover. To distinguish between these possibilities, we determined the steady state levels of collagen mRNAs in vim-/- MEFs and vim+/+ MEFs. The steady state levels of both collagen α1(I) and αβ(I) mRNAs were reduced in vimentin deficient MEFs (Fig 7B, lane 2). The steady state level of GAPDH mRNA was unchanged. Next, we tested if the reduced level was due to an increased turnover of collagen mRNA. We determined the stability of collagen mRNAs by assessing their steady state level at different time points after transcriptional block by Actinomycin D. Collagen mRNAs have a half-life of 12 or more hours in various fibroblasts, therefore, there was little decay in wild type MEFs. However, the decay was much faster in vim-/- MEFS, where the half-life was measured to be 6 hrs. This result suggests that vimentin is needed for stability of collagen mRNAs and that association of vimentin filaments with collagen mRNAs may serve this purpose.

A 1 2 3 4 B 1 2 COL(I)α1 COL 1A1 COL(I)α2 COL 1A2 FIB

CELLS: VIM++ VIM-- VIM++ VIM-- GAPDH CELL SEC CELLS: VIM++ VIM--

1.2 1.4 COL(I)α1 COL(I)α2 1 1.2 VIM++ COL/FIB 1 0.8 -- VIM++ COL/GAPDH VIM 0.8 VIM-- 0.6 0.6 0.4 EXPRESSION: 0.4

EXPRESSION: 0.2 0.2 0 0 COL1A1 COL1A2 CELL SEC CELL SEC p=0.0029 p=0.0033 p=0.0001 p=0.0042 p=0.0002 p=0.0004

C D 1 2 3 4 COL1A1 1.2 COL 1A1

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0.4 ACT % RNAREMAINING% 0.2 CELLS: VIM++ VIM++ VIM-- VIM-- 0 FRAC: SOL INS SOL INS ACT-D (h): 0 6 12 24 100

COL1A2 ++ 1.2 80 VIM VIM -- 1 60 0.8

% COL INS COL % 40 0.6

% RNAREMAINING% 0.4 20

0.2 0 0 COL1 A1 COL1 A2 ACT-D (h): 0 6 12 24 p= 0013 p= 0.001 30

Figure 7. Decreased collagen synthesis in vimentin knock out cells. A. Expression of type I collagen in embryonic fibroblasts from wt (VIM++) and vimentin knockout mice (VIM--). Upper panel; a representative western blot of cellular (CELL, lanes 1 and 2) and secreted levels of collagen (I) α1 and αβ polypeptides (SEC, lanes γ and 4) of VIM++ cells (lanes 1 and 3) and — VIM cells (lanes 2 and 4). Loading control; fibronectin (FIB). Bottom panel; quantitation of expression from three independent experiments. Expression of collagen polypeptides was ++ normalized to expression of fibronectin and arbitrarily set as 1 for VIM cells. Error bars of +/- 2 S.E.M. and statistical significance are indicated. B. Reduced steady state levels of collagen — α1(I) and αβ(I) mRNAs in VIM . Upper panel; representative RT-PCR analysis of collagen α1(I), collagen αβ(I) and GAPDH mRNA levels in VIM++ (lane 1) and VIM— (lane 2) mouse embryonic fibroblasts. Bottom panel; quantification of mRNA levels in three independent experiments. Expression of collagen mRNAs was normalized to expression of GAPDH mRNA and arbitrarily set as 1 for VIM++ cells. Error bars of +/- 2 SD and statistical significance are ++ — indicated. C. Decay rate of collagen α1(I) and αβ(I) mRNAs in VIM and VIM cells. Transcription was blocked by Actinomycin D in VIM++ and VIM— cells and collagenα1(I) and αβ(I) mRNA levels were measured at 0h 6h, 12h and 24h hrs after the block by RT-PCR. GAPDH mRNA was measured as a loading control. Expression of collagen mRNAs was ++ normalized to expression of GAPDH mRNA and set as 1 for VIM cells. The values for α1(I) mRNA (left panel) and αβ(I) mRNA (right panel) were plotted for each time point and error bars of +/- 2SD are shown. D. Segregation of collagen mRNAs with insoluble vimentin. `Top panel; vimentin wt fibroblasts (VIM++, lanes 1 and 2) and vimentin knock-out fibroblasts (VIM--, lanes 3 and 4) were fractionated into soluble (SOL, lanes 1 and 3) or insoluble (INS,

lanes β and 4) fractions. The fractions were analyzed for the presence of collagen α1(I), collagen αβ(I) and actin mRNAs by RT-PCR. Bottom panel; percentage of collagen mRNAs found in the insoluble fractions of VIM++ and VIM—cells, as estimated by three independent experiments. Statistical significance and error bars of +/- 2SD are shown.

Vimentin is the only cytoplasmic intermediate filament system in fibroblasts [195]. Since the insoluble fraction of human lung fibroblasts is enriched in collagen mRNAs (Fig 2), the insoluble fraction from vim-/- cells should not contain the majority of collagen mRNAs. We prepared detergent insoluble fractions from vim+/+ MEFs and vim-/- MEFs and estimated the abundance of collagen α1(I) and αβ(I)mRNAs in these fractions. In vimentin deficient MEFs more collagen mRNAs were found in the soluble fraction (Fig 7D, lanes 3 and 4), in wild type MEFs the opposite was found (Fig. 7D, lanes 1 and 2). The total level of collagen mRNAs was lower in vim-/- MEFS, as shown before in Fig 7B. The distribution of actin mRNA into insoluble and soluble fractions was identical in the Vim+/+ and Vim-/- cells (Fig. 7D, lower panel). This finding further corroborates the specific association of vimentin with collagen α1(I) and αβ(I) mRNAs.

Disruption of vimentin filaments decreases expression of type I collagen. Since the absence of vimentin in fibroblasts resulted in destabilization of collagen mRNAs, we wanted to know if similar effects can be achieved by disrupting vimentin filaments. .’-Iminoditropionitrile (IDPN) is a drug that specifically disrupts vimentin intermediate filaments [196]. When human lung fibroblasts were treated with .’-Iminoditropionitrile (IDPN), complete collapse of the vimentin network was observed 24h after the treatment (Fig. 8A). Treatment with IDPN also reduced cellular and secreted levels of collagen α1(I) and αβ(I) polypeptides. Here, the effect was more pronounced on α1(I) polypeptides than αβ(I) polypeptides, while there was no effect on the control protein, fibronectin (Fig 8B). Treatment with IDPN significantly decreased the steady state levels of collagen α1(I) and αβ(I) mRNAs, reaching the maximal effect after 12 hrs after (Fig. 8C). In order to determine if this effect is due to a faster decay of collagen mRNAs, we measured the stability of collagen mRNAs after 12hrs of IDPN treatment. Figure 8D shows that collagen α1(I) and αβ(I) mRNAs decayed with half-lives of 5h and 8h in cells treated with IDPN compared to 18h and 24h in untreated cells. These half-lives are similar to that obtained in vimentin knockout MEFs (Fig. 7). The drawback of using drugs to perturb any cellular function is their pleiotropic effect. So, it is possible that IDPN may have had non-specific effects in our experiments. To verify that the effect of IDPN on collagen production is due to disruption of vimentin filaments, we treated vim+/+ MEFs and vim-/- MEFs with IDPN. If the effect of IDPN is vimentin specific, then only vim+/+ MEFs should show reduced collagen expression, while

CONTROL IDPN

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CELL SEC GAPDH IDPN (h): 0 12 24 30 36 COL(I)α 1 COL(I)α 2 1.2 1.4 1.2 1 COL1A1 CON COL1A2

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EXPRESSION: 0.4

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0 0 CELL SEC CELL SEC IDPN (h): 0 12 24 30 36 0 12 24 30 36 p=0.0019 p=0.0048 p=0.0036 p=0.014

D COL1A1 COL1A2 1.2 1.2 CON CON 1 1 IDPN IDPN

0.8 0.8

0.6 RNAREMAINING% 0.6 % RNAREMAINING% 0.4 0.4 0.2 0.2 0 0 ACT-D (h): 0 6 12 18 24 ACT-D (h): 0 6 12 18 24

E 1 2 3 4 5 6 7 8

COL(I)α1

FIB

IDPN CON IDPN CON IDPNCON IDPN CON CELLS: VIM++ VIM++ VIM-- VIM--

CELL SEC CELL SEC

Figure 8. Disruption of vimentin filaments by β,β’- iminoditropionitrile (IDPN) reduces collagen synthesis. A. Collapse of vimentin filaments after IDPN treatment. Untreated HLF (control) and HLF treated with 1% IDPN were immunostained with anti-vimentin antibody. Bars represent 1 µM. B. IDPN reduces collagen protein. Top panel; cellular (CELL, lanes 1 and2) and secreted (SEC, lanes 3 and 4) levels of collagen α1(I) and αβ(I) polypeptides from control cells (lanes 1 and 3) or IDPN treated cells (lanes 2 and 4) were analyzed by western blot. Loading control; fibronectin (FIB). Bottom panel; collagen expression was normalized to fibronectin expression and plotted from three independent experiments. Statistical significance and error bars of +/- 2SD are shown. C. IDPN decreases the steady state level of collagen mRNAs. Top panel; collagen mRNAs from control cells and cells treated with IDPN for the indicated time periods were analyzed by RT-PCR. Loading control; GAPDH. Bottom panel; expression of collagen mRNAs was normalized to expression of GAPDH mRNA and plotted from three independent experiments. Error bars; +/- 2 SD. D. Decay of collagen α1(I) mRNA (left panel) and collagen αβ(I) mRNA (right panel) in control and IDPN treated HLF. Transcription was blocked with actinomycin D and the level of collagen mRNAs was estimated by RT-PCR at the indicated time periods. The expression at time 0 was set as 1. Error bars; +/- 2 S.E.M. estimated from three independent experiments. E. IDPN affects collagen expression only in vimentin expressing cells. Cellular (CELL) and secreted (SEC) levels of collagen α1(I) polypeptide were measured in VIM++ fibroblasts (lanes 1-4) and in VIM—fibroblasts (lanes 5-8) by western blot. The cells were treated with IDPN or left untreated, as indicated. Loading control; fibronectin (FIB). there should be no effect on vim -/- MEFs. Treatment of vim+/+ MEFs with IDPN reduced both, cellular and secreted amounts of collagen protein (Fig. 8E, lanes 1-4), but the level in vim-/- MEFs was unchanged (Fig 8E, lanes 5-8). Since IDPN had no effect in the absence of vimentin, we concluded that its effect on collagen expression was specifically due to the disruption of vimentin filaments. Dominant negative form of desmin reduces collagen synthesis. We also used an alternative approach to disrupt vimentin filaments by overexpressing a dominant negative form of desmin. Desmin is closely related to vimentin and forms filaments, but it is not expressed in fibroblasts [195]. Desmin monomers truncated at the amino-acid 263 acts as a dominant negative protein that interferes with the organization of desmin, as well as vimentin intermediate filaments [197-199]. Therefore, we constructed a recombinant adenovirus expressing this dominant negative form of desmin. Transduction of human fibroblasts with this adenovirus disrupted vimentin filaments, as assessed by immunostaining (Fig. 9A). We observed granular aggregates of vimentin (Fig. 9A, left panel), instead of the normal filamentous distribution of vimentin network (Fig 9A, right panel). To verify that the truncated desmin was integrated into the collapsed vimentin filaments, we fractionated the cell lysates into detergent

34 soluble and insoluble fractions as described for Fig 2. Most of the transfected desmin co- fractionated into the insoluble fraction together with vimentin (Fig. 9B). Its presence in the insoluble fraction strongly suggests that it has been incorporated into the endogenous vimentin network. We then assessed the effect of the dominant-negative desmin on collagen expression in two collagen producing cell types. In human lung fibroblasts the dominant negative desmin significantly decreased cellular and secreted levels of collagen α1(I) and αβ(I) polypeptides, as compared to cells infected with control virus (Fig. 9C, compare lanes 1 and 3 to lanes 2 and 4). The level of fibronectin was not affected, suggesting the specificity of the effect on collagen production. The steady state levels of both collagen mRNAs, α1(I) and αβ(I), were dramatically reduced in lung fibroblasts expressing the dominant negative desmin (Fig. 9D). In order to determine if this is due to a faster decay of collagen mRNAs, we measured the stability of collagen α1(I) and αβ(I) mRNAs in cells expressing dominant negative desmin. Figure 9E shows that collagen α1(I) and αβ(I) mRNAs decayed with half-lives of 5h and 9h, respectively, compared to 18h and 24h in control cells. This result is similar to the result obtained with vimentin knockout MEFs (Fig. 7) and IDPN treated cells (Fig. 8). We also tested the effect of dominant negative desmin in scleroderma skin fibroblasts. Markedly reduced cellular and secreted levels of collagen α1(I) and αβ(I) polypeptides were found (Fig. 9F, compare lanes 2 and 4 with lanes 1 and 3). Cellular and secreted levels of fibronectin were again unaffected. Decreased collagen protein production was correlated with the decreased levels of collagen mRNAs (Fig. 9G). Taken together, these results strongly indicate that the integrity of vimentin filaments is necessary for stability of collagen α1(I) and αβ(I) mRNAs.

A DN-DES CON B 1 2 3 4

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VIRUS: DN- DN- CON CON DES DES FRACT: INS SOL INS SOL

C D 1 2 1 2 3 4 COL Iα COL1A1 I COL Iα COL1A2 I FIB GAPDH VIRUS: DN- DN- CON CON VIRUS: DN- CON DES DES DES CELL SEC

1.4 COL Iα COL Iα 1.2 COL1A1 I COL1A2 I 1.2 CON 1 DN-DES COL/FIB 1 CON 0.8

0.8 COL/GAPDH DN-DES 0.6 0.6

EXPRESSION: 0.4 0.4 0.2 0.2 EXPRESSION: 0 0 CON DN-DES CELL SEC CELL SEC CON DN-DES p=0.0041 p=0.0001 p=0.0025 p=0.0037 p=0.0093 p=0.0024

E 1.2 COL1A1 COL1A2 1.2 CON 1 CON 1 DN-DES DN-DES 0.8 0.8 0.6 0.6

RNAREMAINING% % RNAREMAINING% 0.4 0.4

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0 0 ACT-D (h): 0 6 12 18 24 ACT-D (h): 0 6 12 18 24

F G 1 2 3 4 1 2

COLI α COL1A1 I

COL Iα COL1A2 I

FIB GAPDH VIRUS: DN- CON DN- CON DES DES VIRUS: DN - CON CELL SEC DES Figure 9. Disruption of vimentin filaments by a dominant negative mutant of desmin reduces collagen synthesis. A. A dominant negative mutant of desmin disrupts vimentin filaments. HLF were transduced with control adenovirus (right panels) or adenovirus expressing dominant negative mutant of desmin (DN-DES, left panels). The cells were immunostained with anti-vimentin antibody. Lower images show a higher magnification. B. DN-DES co-fractionates with insoluble vimentin. The cells in A were fractionated into vimentin soluble and insoluble fraction and analyzed by western blot using anti-vimentin, anti- tubulin and anti-FLAG antibodies, which recognizes DN-DES. C. Top panel; collagen α1(I) and αβ( I) polypeptides from HLF transduced with control (lanes 2 and 4) or DN-DES (lanes 1 and 3) adenovirus were analyzed by western blot. Loading control; fibronectin (FIB). Bottom panel; collagen expression was normalized to fibronectin expression and shown from three independent experiments. Expression in control cells was set as 1 and statistical significance and error bars of +/- 2SD are shown. D. DN-DES decreases steady state level of collagen mRNAs. Top panel; the cells in C were analyzed for expression of collagen α1(I), collagen αβ(I) and GAPDH mRNAs by RT-PCR. Bottom panel; expression of collagen mRNAs was normalized to expression of GAPDH and plotted from three independent experiments. Expression in control cells was set as 1 and statistical significance and error bars of +/- 2SD are shown. E. The levels of collagen α1(I) mRNA (left panel) and collage αβ(I) mRNA (right panel) were estimated in control and in DN-DES expressing cells after transcriptional block with actinomycin D for the indicated time periods. The expression at time 0 was set as 1. Error bars; +/- 2SD estimated from three independent experiments. F. DN-DES decreases collagen expression in scleroderma fibroblasts. Cellular (lanes 1 and 2) and secreted (lanes 3 and 4) levels of collagen polypeptides of cells expressing DN-DES (lanes 1 and 3) or control protein (lanes 2 and 4), analyzed by western blot. Fibronectin (FIB) is shown as loading control. G. Reduced collagen mRNA levels in scleroderma fibroblasts expressing DN-DES. Total RNA from cells expressing DN-DES (lane 1) or control protein (lane 2) were analyzed by RT-PCR for collagen α1(I), collagen αβ(I) and GAPDH mRNAs.

Vimentin filaments stabilize collagen mRNAs that are not actively engaged in translation. Our results suggest that vimentin filaments regulate stability of collagen mRNAs, but do not indicate if these mRNAs are translated or not. When we fractionated polysomes on sucrose

37 gradients [88] and analyzed the fractions for vimentin by western blot, we could not detect any vimentin in the fractions containing polysomes (not shown). To further corroborate the absence of ribosomes on vimentin filaments, we fractionated cells into soluble and insoluble fractions as described in Fig. 2 and assessed if any ribosomal RNA can be found in the insoluble fraction, which contains majority of vimentin. No ribosomal RNA was found in the insoluble fraction (Fig 10A), suggesting no association of ribosomes with vimentin filaments and that the fraction of collagen mRNAs which associates with these filaments is not translated. If vimentin filaments preferentially bind collagen mRNAs that are not being translated, then dissociation of polysomes should increase their loading onto vimentin filaments. To assess this, we performed immunoprecipitation with vimentin antibody in cells treated with puromycin and cycloheximide and analyzed for pull down of collagen mRNAs. Twice as much collagen mRNA was pulled down from the puromycin treated cells, compared to the cells treated with cycloheximide or untreated cells (Fig10C, compare lane 2 with lanes 1 and 4). The total level of collagen mRNAs was unaffected by cycloheximide and puromycin treatment, suggesting the redistribution of collagen mRNAs onto vimentin filaments upon dissociation of polysomes. Together, the results suggest that vimentin filaments stabilize collagen mRNAs that are not actively engaged in translation.

A 1 2 B C 1 2 3 1 2 3 4 28S COL 1A1 COL 1A1 18S COL 1A2 COL 1A2 INS SOL

FIB FIB PUR CHX - IP: VIM VIM CON VIM CHX PUR --

Figure 10. Vimentin associates with un-translating collagen mRNAs. A . Absence of ribosomal RNA in the insoluble fraction containing filamentous vimentin. Soluble (SOL, lane 2) and insoluble (INS, lane 1) fractions were prepared, total RNA extracted and analyzed by agarose gel electrophoresis and ethidium bromide staining. B. Dissociation of polysomes increases association of collagen mRNAs with vimentin. Immunoprecipitation with anti-vimentin antibody (lanes 1, 2 and 4) or control antibody (lane 3) from human lung fibroblasts treated with cycloheximide (CHX, lane 1) or puromycin (PURO, lane 2) or from untreated cells (-, lanes 3 and 4). The immunoprecipitated material was analyzed for

collagen α1(I), collagen α2(I) and fibronectin mRNAs by RT-PCR. C. Cycloheximide and puromycin do not change the total level of collagen mRNAs. RT-PCR analysis of collagen α1(I), collagen αβ(I) and fibronectin mRNAs in cells treated with puromycin (PUR, lane 1) or cycloheximide (CHX, lane 2) or untreated cells (-, lane 3).

2.2- Discussion Recent work from our lab reported that binding of LARP6 to the conserved 5’SL of collagen mRNAs associates collagen mRNAs with filaments composed of nonmuscle myosin and regulates translation [89]. In the present study, we show that: 1. collagen mRNAs that are not translating associate with vimentin intermediate filaments; 2. this association is through the interaction of the La domain of LARP6 with the filaments; 3. the association with vimentin filaments stabilizes collagen mRNAs; 4. one of the roles of vimentin in mesenchymal cells may be to promote collagen synthesis by increasing the level of collagen mRNAs. There is now substantial evidence that in different cell types mRNAs associate with the cytoskeleton [108]. Of the three major filamentous systems, the association of mRNAs with microfilaments and microtubules is well established. There is some evidence that mRNAs may also associate with intermediate filaments. Ultra-structural in situ hybridization study of the distribution of poly(A)+ mRNAs showed about 15% of total poly(A)+ are found close to or associated with vimentin filaments [200]. In another study, mRNAs for calcitonin gene-related peptide in axons of neurons and IL-1 in monocytes were found associated with vimentin filaments [201, 202]. Unlike that of microtubules and microfilaments, however, the functional significance of association of mRNAs with intermediate filaments is not understood. This study gives the first example of a functionally relevant interaction of collagen mRNAs with vimentin intermediate filaments. Vimentin binding to and stabilizing collagen mRNAs was supported by the evidence obtained using different approaches. First, using RNA-immunoprecipitation and cellular fractionation methods, we showed that collagen mRNAs exhibit specific interaction with vimentin filaments (Fig. 1 and 2). Second, using immunoflourescence coupled with RNA-FISH, we showed that a significant proportion of collagen mRNAs are co-localized with vimentin filaments (Fig. 3). Third, fibroblasts from vimentin knockout mice have reduced collagen synthesis due to the decreased half-life of collagen mRNAs in (Fig. 7). Fourth, disruption of vimentin filaments using either a drug or a dominant negative mutant of desmin resulted in a faster decay of collagen mRNAs (Figs. 8 and 9). We also provided insight into the mechanism of the interaction between collagen mRNAs and vimentin filaments. The key feature of this interaction is that it is dependent on a cis-acting RNA element of collagen mRNAs, the 5’SL and its cognate RNA binding protein, LARP6. Possession

39 of cis-acting RNA elements governing interactions with the cytoskeleton is an important characteristic of all mRNAs that localize with the cytoskeleton [203]. These cis acting elements are almost always located in the γ’ UTR. Collagen mRNAs are unique in this respect, because their cis acting sequence is in the 5’ UTR. That collagen mRNAs require LARP6 for binding vimentin filaments was corroborated by multiple lines of evidence. LARP6 coprecipitates and colocalizes with vimentin; knock down of LARP6 decreases the association of collagen mRNAs with vimentin; the interaction of LARP6 with vimentin is 5’SL dependent. Thus, this is a novel function of LARP6 in posttranscriptional regulation of collagen expression. The lack of association of polysomes with vimentin and the absence of ribosomal RNA in the insoluble fraction supports the notion that vimentin filaments preferentially bind non-translating collagen mRNAs. Collagen mRNAs that are released upon dissociation of polysomes seem to be targeted to vimentin filaments (Fig 10). Thus, vimentin filaments could play a role in storage of untranslated collagen mRNAs. We estimated that 30-50% of collagen mRNAs co-localize with vimentin. Based on the polysomal profile of collagen mRNA [88, 89], about 50% of collagen mRNAs are engaged in translation, thus, the rest of collagen mRNAs appear to be primarily stored on vimentin filaments. We mapped the domain of LARP6 that binds vimentin to be the La domain. The La domain is well conserved in all LARPs [194], so it is possible that other LARPs also bind vimentin filaments. This remains to be verified, but it may provide insight into the function of other LARP family members. However, no other LARP family member has a high affinity mRNA target. The most relevant feature of the interaction of collagen α1(I) and αβ(I) mRNAs with vimentin filaments is that this interaction stabilizes collagen mRNAs. Although regulation of the stability of collagen mRNAs has emerged as the predominant means for high-level synthesis of type I collagen, the mechanism of stabilization was only partially understood [70] [85] [77][204]. α-CP binds the C-rich region in the γ’UTR of collagen α1(I) mRNA and contributes to its prolonged half-life [80, 180] [77]. This study suggests that an additional mechanism of stabilizing collagen α1(I) and αβ(I) mRNAs may utilize the 5’UTR. The precise mechanism by which vimentin filaments stabilize collagen α1(I) and αβ(I) mRNAs is not clear yet. One possibility is that vimentin filaments prevent access of the RNA degradation machinery to collagen mRNAs.

Our finding that vimentin deficient fibroblasts exhibited markedly reduced collagen synthesis is consistent with the impaired wound healing observed in vimentin knockout mice. In vimentin deficient mice, wound healing is dramatically impaired due to defective fibroblast invasion into the wound, with the resultant delayed wound contraction [99] [100]. Based on our results, we speculate that reduced collagen production by vimentin deficient fibroblasts may have also contributed to the observed delayed would-healing phenotype. Congruent with our hypothesis, the initial collagen deposition in the proliferative phase of wound healing is an important step for the subsequent fibroblast invasion [110]. Wound healing experiments using vimentin deficient mice, with an emphasis on the amount of collagen deposited, are needed to verify this hypothesis. From a therapeutic point of view, considering the central role of the increased collagen expression in the development of tissue fibrosis, vimentin filaments may be a target for antifibrotic therapy. The potential utility of targeting vimentin is highlighted by our findings that disruption of vimentin reduced synthesis of type I collagen. In conclusion, we propose a model for post-transcriptional regulation of collagen expression where LARP6-bound collagen mRNAs are stored and stabilized by vimentin filaments and are available for recruitment by the translational machinery. 2.3- Materials and Methods Cells and transfections. HEK293 cells and human lung fibroblasts immortalized by expression of telomerase reverse transcriptase [205] were grown under standard conditions. HEK293 cells were transfected with 1 μg of plasmid per γ5 -mm dish using 293TransIT reagent (Mirus). Transduction of lung fibroblasts with adenoviruses was done by adding adenoviruses at a multiplicity of infection (MOI) of 100. With this MOI, between 95% and 100% of the cells were transduced, as judged by expression of GFP viral marker. The cells were harvested for analysis 2-5 days after viral delivery. Scleroderma fibroblasts derived from skin of a scleroderma patient were purchased from the European collection of cell cultures (cell line BM0070). MEFs were derived from knock-in mice in which the 5’SL of collagen α1(I) gene has been mutated and their wild type littermates [87]. All cells were cultured in Dulbecco’s Modified Eagle’s Medium supplemented with 10% fetal bovine serum for up to 10 passages.

Vim+/+ and Vim-/- MEFs derived from wild type and vimentin knockout mice were described previously [206]. (they were kind gifts from Dr. Robert Evans, University of Colorado.) Chemicals. Beta,beta’-iminodipropionitrile (IDPN) was from Acros Organics (Morris Plains, NJ). Actinomycin D, puromycin and cycloheximide were purchased from Sigma. For treatment with IDPN, cells were incubated with 1% IDPN for 24hrs before analysis. Actinomycin at 10µg/ml was added and cells were collected at the indicated time points. Puromycin at 100µg/ml and cycloheximide at 100µg/ml were added to cells for 30 minutes before the cells were fixed for immunostaining or proteins were extracted for immunoprecipitation. Plasmid constructs and adenovirus preparation. HA-tagged wild type LARP6 and some deletion constructs have been described previously [88]. Additional deletion constructs were made by PCR amplification of the regions of LARP6 and cloning the PCR products into pCDNA3 vector. Adenovirus expressing dominant negative desmin was constructed by PCR amplification of the human desmin, region from amino-acid 1 to amino-acid 263, and cloning the PCR product into pAdCMVTRACK vector, followed by recombination with pADEasy vector, as described previously [207]. Adenovirus was amplified in HEK293 cells and purified by a Virapure kit (Clontech). Expression of the construct was verified by western blot analysis. This virus expressed both, the test protein and the green fluorescent protein (GFP), which is encoded by an independent transcription unit [207]. Expression of GFP served as control for viral transduction. Control adenovirus expressed only GFP. LARP6 siRNA. An siRNA effective against LARP6 with the sequence, 5’- UCCAACUCGUCCACGUCCU-γ’, was previously described [88]. This siRNA sequence was expressed as shRNA from adenovirus [88]. A control adenovirus contained shRNA with a scrambled sequence (GGAGGGCUUCGAGUUAGGA). Efficacy of LARP6 knockdown was assessed by western blot. Reverse transcription-PCR analysis. Total cellular RNA was isolated using an RNA isolation kit (Sigma). RT-PCRs were performed with 100ng of total RNA or using rTth reverse transcriptase (Boca Scientific, Boca Raton, FL). [32P]dATP was included in the PCR step to label the products, which were resolved on sequencing gels, as described previously [85]. The number of cycles was adjusted within the linear range of the reaction. The primers used for RT-PCR were as follows: human-collagen α1(I), 5’primer AGAGGCGAAGGCAACAGTCG and γ’ primer GCAGGGCCAATGTCTAGTCC; human-collagen αβ(I), 5’ primer CTTCGTG CCTAG

CAACATGC and γ’ primer TCAACACCATCTCTGCCTCG; human-fibronectin, 5’ primer ACCAACCTACGGATGACTCG and γ’ primer GCTCATCATCTGGCCATTTT; human- GAPDH, 5’ primer : ACCGGTTCCAGTAGGTACTG and γ’ primer CTCACCG TCACTACC GTACC; human-actin, 5’ primer GTGCGTGACATTAAGGAGAAG and γ’ primer GAAG GTAGTTTCGTGGATGCC; mouse-collagen α1(I), 5’primer GAGCGGAGAGTACTGGATCG and γ’ primer TACTCGAACGGGAATCCATC; mouse-collagen αβ(I), 5’ primer CTTCGTGC CTAGCAACATGC and γ’ primer TCAACACCATCTCTGCCTCG; mouse-actin, 5’ primer CGTGCGTGACATCAAAGAGAAGC and γ’ primer TGGATGCCACAGGATTCCATACC. Antibodies. Anti-HA and anti-vimentin antibodies were obtained from Sigma. Anti-collagen α1(I) antibody was obtained from Rockland; anti-collagen αβ(I) antibody specific for human polypeptide was from Cell Signaling; the anti-collagen αβ(I) antibody to detect mouse polypeptide was obtained from Santa-Cruz Biotechnology; anti-fibronectin and anti-tubulin antibodies were from BD Biosciences; anti-LARP6 antibody was obtained from Abnova; anti- actin antibody was from Novus Biological and anti-MYH10 antibody was from Hybridoma bank, University of Iowa. Western blot analysis. Protein concentration was estimated by the Bradford assay, with bovine serum albumin as the standard. Fifty (50) µg of total cellular protein was typically used for western blot analysis. For western blot analysis of secreted proteins, equal numbers of cells were seeded and after 24-48 hrs, serum-free medium was added to the cells and incubation continued for 3hr. The medium was collected, and an aliquot was analyzed directly by western blot analysis. Use of serum free medium in collecting secreted proteins was essential, as fetal calf serum contains substantial amounts of collagen and fibronectin. Immunostaining. For immunostaining the cells were grown on glass coverslips. After treatment, the cells were fixed with 4% formaldehyde for 30min at room temperature and permeabilized with 0.5% Triton X-100 in phosphate buffered saline (PBS) for 10min. Blocking was done with 10% goat serum/5% bovine serum albumin in PBS for 1hr at room temperature, followed by incubation with primary antibody overnight at 40C. After washing, proteins were visualized with AlexaFluor594- , Cy-2-or Cy5- conjugated secondary antibodies. The cells were mounted using PROLONG mounting solution containing 4’,6’-diamidino-2-phenylindole (DAPI) from Invitrogen. Images were taken with the Leica TCS SP2 AOBS laser confocal microscope

43 equipped with a Chameleon Ti-Sapphire multiphoton laser. Optical sections were processed with LCSLite software and single plane confocal images are shown. Immunoprecipitations. Cell extracts were prepared in lysis buffer [10mM KCl, 1.5 mM MgCl2, 10mM Tris-HCl (PH 7.5), 0.5% NP-40 and 170 µg/ml phenylmethylsulfonyl fluoride]. After removal of nuclei by centrifugation, the cleared lysate was incubated with 1µg of antibody for 1hr at 40C. Thirty micro liters of equilibrated protein A/G agarose plus beads (Santa Cruz Biotechnology) were added, and incubation continued for an additional 3hrs. After the beads were washed three times with PBS, immunoprecipitated complexes were analyzed by SDS- PAGE and western blotting. For analysis of immunoprecipitated RNA, total RNA was extracted from the immunoprecipitated material. In reactions in which RNase A digestion of the precipitate was performed, 0.2µg/µl RNase A was added and incubated with A/G agarose plus beads for 15minutes at room temperature and then the samples were washed two times in PBS before loading into the gel. Determination of RNA stability. Cells were treated with Actinomycin D (10 μg/ml) for 6, 1β, or 24h. After the Actinomycin D incubation period, the cells were scraped and total RNA was extracted and analyzed by RT-PCR. RNA extracted from cells at time point 0 (immediately after addition of Actinomycin D) was used as the initial level of mRNA and arbitrarily set as 100%. The results shown are from 3 independent experiments. Fractionation of cell extracts into intermediate filament (IF) rich fractions. Fractionation of cells into IF-rich (detergent insoluble), and IF-poor (detergent soluble) fractions was done as described [186]. Briefly, cells were homogenized in a buffer containing 1% Nonidet P-40, 10% glycerol, 20 mM Hepes, pH 7.6, 150 mM NaCl, and protease inhibitors (2mM benzamidine, 0.5 mM aprotinin and 1 mM phenylmethylsulfonyl fluoride). The homogenates were incubated in the same buffer at 37°C for 30 min. The soluble fraction (supernatant) and the insoluble fraction (pellet) were collected after centrifugation at 5,200 rpm, 4°C for 30 min. The volume of both fractions was made the same by adding SDS sample buffer. Equal volumes of the samples were analyzed by western blot. For measurement of the distribution of collagen mRNAs into IF-rich and IF-poor cellular fraction, total RNA was extracted after fractionation and analyzed by RT- PCR. For immunoprecipitations of collagen mRNAs from the detergent-soluble and detergent- insoluble fractions, the soluble fraction was prepared as above and insoluble fraction was

44 dissolved in equal volume of buffer containing 0.1%SDS, 10mM Tris-HCl (pH=6.8), 10% glycerol, and protease inhibitors. Then, the immunoprecipitation was carried out as described for cell extracts. RNA Fluorescent In Situ Hybridization (RNA FISH). For the detection of collagen α1(I) and αβ(I)) mRNAs, five fluorescently labeled DNA oligos were designed for each mRNA. Design of the oligonucleotide probe was done as described before [189]. Probes were 50 nucleotides long and contained four or five amino-modified nucleotides (amino-allyl T) coupled with cy3 or cy5. For Collagen-α1(I) (accession number: NM-000088), the sequences of the probes (with modified T’s in bold type) and the nucleotides in the mRNA they are complementary to (in parentheses) were: GTTCTGTACGCAGGTGATTGGTGGGATGTCTTCGTCTTGGCCCTCGACTT (255- 206), GCAGTTCTTGGTCTCGTCACAGATCACGTCATCGCACAACACCTTGCCG (370- 321), TGGAGGGAGTTTACAGGAAGCAGACAGGGCCAACGTCGAAGCCGAATTCC (4446-395), GATGATGGGCAGGCGGGAGGTCTTGGTGGTTTTGTATTCAATCACTGT CTT (4531-4482) and TGGAATCCATCGGTCATGCTCTCGCCGAACCAGACATGCCTCTT GTCCTT (5030-4981). For Collagen-αβ(I) (accession number: NM-000089), the sequences of the probes and the nucleotides they are complementary to are: GGACGTGGACACTTTTGAG GCTTTCAAGGGGAAACTCTGACTCGTGTCT (120-70), CCGATGTCCAAAGGTGCAA TATCAAGGAAGGGCAGGCGTGATGGCTTATT(4520-4471), GGGCCAAGTCCAACTCCT TTTCCATCATACTGAGCAGCAAAGTTCCCACC(740-691), CATCCAGACCATTGTGTCC CCTAATGCCTTTGAAGCCAGGAAGTCCAGGA(1030-981) and AGAAGTCTCCATCGTAA CCAAAGTCATAACCACCACCGCTTACACCTGGA (3820- 3771). RNA-FISH and IF-RNA FISH were done as described previously [189]. Briefly, human lung fibroblasts were grown on cover slips for 24 hrs, followed by fixation with 4% formaldehyde in PBS for 30 min. Cells were rinsed in PBSG two times for 5 min. each, followed by permeabilzation with 0.5% Triton X-100 for 10 min at room temperature. Cells were washed twice in PBSG and equilibrated in pre-hybridization buffer (50% formamide/2X SSC) for 10 minutes. For the hybridization step, cover slips were incubated with fluorescently labeled oligos in hybridization buffer (2× SSC, 10% dextran sulfate, 50% formamide, 1mg/ml BSA, 1mg/ml Sheared salmon sperm DNA, 1 mg/ml E. coli RNase-free t-RNA, and 10 ng/μl of oligonucleotide probe) for 2hrs at 37°C. After hybridization, cells were washed twice in 2× SSC/50% formamide at 370c for 20 min, in 1× SSC/50% formamide at 370c for 30 min, then in

1xSSC at room temperature for 15 min and finally in 0.5xSSC at room temperature for 15 min. Coverslips were mouned with PROLONG anti-fade mounting medium containing DAPI (Invitrogen). For IF-RNA FISH experiments, human lung fibroblasts were fixed and permeabilized as above followed by blocking with FISH compatible blocking solution (CAS-block) (Invitrogen). Cover slips were then processed for immunofluorescence using anti-vimentin antibodies diluted in CAS block containing 10mM RNase inhibitor (RVC). After overnight incubation, cells were washed with PBS containing 0.1% Tween 20 (PBST) and primary antibody was fixed with 4% paraformaldehyde in PBS for 10 min. After fixation, cells were washed with PBST for 10 minutes followed by FISH hybridization and washing as above. Following the last wash, coverslips were incubated with cy5-labeled anti-rabbit secondary antibody diluted in CAS block with 10mM RVC. After washing with PBST for 30 min, mounting was done as above. Fluorescent images were acquired with The DeltaVision image restoration (deconvolution) microscope system (Applied Precision) using a 60X objective (1.42 NA; Olympus). Digital images were acquired using a charge-coupled device camera (Coolsnap HQ; Photometrics), and stacks of 64 images were taken with a Z-step size of 0.β μm using DeltaVision software and filter sets for DAPI, Cy3, and Cy5. The three-dimensional (3D) stacks were deconvolved to quantitatively improve the signal to noise ratio using the SoftWorx (Applied Precision) constrained iterative deconvolution process. Fractionation for polysomes. Polysomes were fractionated from 2x107 cells on linear 15-45% sucrose gradients as described [88, 89]. In some experiments, puromycin at 100 µM was used to disrupt polysomes. Fractions (0.5 ml) were collected and RNA was extracted with phenol- chlorophorm and precipitated with isopropanol. For analysis of the distribution of vimentin and non-muscle myosin, total protein from the fractions were precipitated with 0.05% deoxycholate and 6.5% trichloracetic acid, washed with acetone, and analyzed by Western blot. Statistical analysis. Densitometric analysis of scanned images from gels was performed by Image J software. Results were analyzed for statistical significance according to the student’s t-test. Statistical values of p< 0.05 were considered to be significant. Data are presented as mean ± S.E.M.

CHAPTER THREE

THE ANTI-FIBROTIC EFFECTS OF WITHAFERIN-A IN TISSUE CULTURE AND IN A MOUSE MODEL OF MYOCARDIAL FIBROSIS

Fibroproliferative disorders are major causes of morbidity and mortality globally [2, 173]. Fibroproliferative disorders affect all tissues and organ systems, and include liver cirrhosis, interstitial lung diseases, chronic renal diseases, and several cardiovascular diseases [3, 181, 182]. In addition to their high prevalence, fibrotic diseases typically have severe and progressive nature [8]. Despite the huge impact of these diseases on human health, there are currently no anti-fibrotic therapies approved for use in humans [16]. Excessive collagen deposition is the hall mark of all fibroproliferative disorders [1]. Activated fibroblasts and myofibroblasts are the most important cells depositing type I collagen in all tissues. Increased activity of profibrotic cytokines such as TGF-1 and IL-13, are implicated in the activation and differentiation of fibroblasts in to myofibroblasts, as well as in mediating the up regulation of type I collagen in these cells [208]. Increased expression of type I collagen from activated fibroblasts and myofibroblasts is regulated both at the level of transcription and post-transcriptionally [176]. Transcription of collagen genes increases markedly in activated fibroblasts [74]. The increase in the stability of collagen mRNAs during activation contributes even more to the high expression. For instance, the dramatic increase in steady state level of collagen mRNAs during activation of hepatic stellate cells is mainly attributed to significant prolongation of the half lives of collagen mRNAs [77, 180]. The increased production of collagen by skin fibroblasts from scleroderma patients is also, primarily, due to an increase in stability of type I collagen mRNAs [209].

The stem-loop of the 5’ untranslated region (UTR) of collagen α1(I) and αβ(I) mRNAs (5’SL) is the key element regulating their stability and translation. LARP6 binds the 5’SL of collagen mRNAs with high affinity and specificity [88]. We recently identified vimentin as key molecule involved in posttranscriptional regulation of collagen expression [210]. We showed that vimentin filaments bind collagen mRNAs in a LARP6 dependent manner and that the

47 integrity of these filaments is crucial for stability of type I collagen mRNAs. The absence of vimentin in mouse embryonic fibroblasts led to significantly decreased collagen production, due to the decreased half life of collagen mRNAs. Likewise, disrupting vimentin filaments by overexpression of dominant negative desmin mutant or by treatment of cells with IDPN, led to a marked reduction in collagen synthesis. Based on these results, we suggested targeting vimentin filaments can be an effective therapy. Withaferin-A is a steroidal lactone and the principal active ingredient of the herbal plant, Withania sominifera [211]. The intermediate filament vimentin is the primary target of Withaferin-A. Withaferin-A (WF-A) binds vimentin and covalently modifies a conserved cysteine residue located in the α-helical rod 2B domain of vimentin [212]. Withaferin-A treatment results in disruption of the vimentin filament network in endothelial cells. In light of our finding of a novel role of vimentin in collagen production, we hypothesized that Withaferin A may reduce collagen production by disrupting vimentin filaments and decreasing the stability of collagen mRNAs. No previous study investigated the effect of Withaferin-A on collagen synthesis and/or fibrosis. The aim of the present study was to determine if Withafrein-A exhibits anti-fibrotic properties in vitro and in vivo and to elucidate the molecular mechanisms by which WF-A exerts its anti-fibrotic effects. We report that, in tissue culture, Withaferin-A suppresses collagen expression, both at transcriptional and post-transcriptional level, by inhibiting the TGF- signaling pathway and by disrupting vimentin filaments, respectively. Importantly, in vivo, we demonstrate that WF-A attenuates cardiac fibrosis in a mouse model of isoproterenol-induced myocardial fibrosis.

3.1- Results Withaferin-A disrupted vimentin filaments and caused degradation of vimentin in fibroblasts It has been reported that WF-A causes disruption of the vimentin network in different cell types including endothelial cells, astrocytes and vimentin positive breast cancer cell lines [212- 214]. However, the effect of WF-A on the integrity of vimentin network in fibroblasts has not been studied. We, thus, determined if the vimentin disrupting effect of WF-A also holds true in fibroblasts. Treatment of human lung fibroblasts (HLFs) with 1.0µM Withaferin-A caused complete collapse of the vimentin network observed 2hr after treatment (Fig. 1A, upper image). Similar results were obtained in scleroderma fibroblasts (SCL) (Fig. 1A, lower images), rat cardiac fibroblasts (RCF) (Fig. 1B, upper image) and mouse embryonic fibroblasts (MEFs) (Fig. 1B, lower images) suggesting that WF-A can disrupt vimentin filaments probably in all fibroblasts.

A B C RCF VIM HLF

ACTIN

WF-A (µM) 0 0.5 1.0 SCL MEF ……………………………………………

DMSO WITHAFERIN-A DMSO WITHAFERIN-A

Figure 11: Effect of Withaferin-A on vimentin intermediate filaments in fibroblasts. A) Collapse of vimentin filaments in human fibroblasts treated with WF-A. Primary human lung fibroblasts (HLF, upper) and Scleroderma fibroblasts (SCL, lower) treated with DMSO or 1.0µM of WF-A for 2hrs were immunostained with anti-vimentin antibodies. Bars, 1µm. B) Collapse of vimentin filaments in rodent fibroblasts treated with Withaferin-A. Primary rat cardiac fibroblasts (RCF, upper) and mouse embryonic fibroblasts (MEF, lower) were immunostained with antivimentin antibodies 2hrs after treatment with DMSO or 1.0µM of WF-A. Bars, 1µm. C) Degradation of soluble vimentin by Withaferin-A. Western blot analysis of HLFs showed that WF-A caused dose dependent increase in degradation products of vimentin 2hrs after treatment.

In addition to disrupting the vimentin filaments, in endothelial and glial cells, WF-A was reported to cause degradation of soluble vimentin [212, 213]. We tested if WF-A can cause degradation of soluble vimentin in human lung fibroblasts. Separation of vimentin into the soluble fraction was undertaken as described before [210]. WF-A caused appearance of lower molecular products of soluble vimentin in dose dependent manner, suggesting that WF-A can cause degradation of vimentin in fibroblasts (Fig. 1C).

Study of the toxicity of Withaferin-A in cultured fibroblasts Withaferin-A has previously been reported to cause apoptosis in various normal and cancer cell types, particularly at higher concentrations [214, 215]. Therefore, we next determined the concentrations of WF-A that induce apoptosis in human and rodent collagen producing cells. Cells were cultured the presence of various concentrations of WF-A (0.25-3.0 µM) for 24 hrs and viability of cells was determined using the caspase glo 3/7 assay kit. Viability of primary human lung fibroblasts (HLF) 24hrs after treatment with the indicated concentrations of Withaferin-A is shown in Fig. 2A. Withaferin-A in concentrations less than 1.5µM did not cause significant cell death in HLFs (Fig. 2A). Concentrations above 1.5µM, however, caused increased loss of cells due to apoptosis in dose dependent manner. At 2.0µM concentration of Withaferin-A, approximately 10% of HLFs were lost by apoptosis, whereas the 3.0µM concentration of WF-A caused 15% loss (Fig. 2A). This results were consistent with what has been reported for human dermal fibroblasts where only doses greater than 1.5µM were considered to be toxic [216]. In Scleroderma fibroblasts, the sub-toxic concentration was below 1.0µM (Fig. 2B).

A HLF SCL HLF B SCL 100 100

95 95

90 90

85 cells Viable (%) 85 Viable cells Viable (%)

80 80 0 0.25 0.5 1 1.5 2 3 0 0.25 0.5 1 1.5 2 3 WF-A (µM) WF-A (µM) 50

MEF VIM+/+ RCF C 100 D VIM-/- 100 RCF

95 95

90 90

85 85 Viable cells Viable (%)

cells Viable (%) 80 80 0 0.25 0.5 1 1.5 2 3 0 0.25 0.5 1 1.5 2 3 WF-A (µM) WF-A (µM)

Figure 12: Determination of toxicity of Withaferin-A in cultured fibroblasts. Cells were seeded in 96-well plates and cultured in triplicate in the presence of various concentrations of WF-A (0.25-3.0 µM) for 24 hrs and apoptosis was determined using the caspase glo 3/7 assay kit. Relative luminescence values for treated cells were converted to % viable cells. A) Viability of primary human lung fibroblasts (HLF) 24hrs after treatment with the indicated concentrations of WF-A (0.25-3.0 µM). B) Viability of Scleroderma cells (SCL) 24hrs after treatment with the indicated concentrations of WF-A (0.25-3.0 µM) is shown as percent of control cells. C) Viability of wild type mouse embryonic fibroblasts (VIM+/+ MEFs) and mouse embryonic fibroblasts from vimentin knockout mouse (VIM-/- MEFs) 24hrs after treatment with the indicated concentrations of WF-A (0.25-3.0 µM) is shown as percent of control cells. D) Viability of rat cardiac fibroblasts (RCF) 24hrs after treatment with the indicated concentrations of WF-A (0.25-3.0 µM) is shown as percent of control cells.

We also tested the toxicity of WF-A in wild type (VIM+/+) and vimentin deficient (VIM -/-) mouse embryonic fibroblasts (MEF). Concentrations of WF-A greater than 1.0µM were found to be toxic in VIM+/+ as well as VIM-/- MEFs (Fig. 2C). This indicates that toxicity of Withaferin-A isn’t due to disruption of vimentin filaments. Last, WF-A at concentrations below 1.5µM didn’t significantly affect the viability of primary rat cardiac fibroblasts (Fig. βD). Withaferin-A significantly reduces expression of type I collagen in fibroblasts To evaluate the effect of WF-A on collagen expression, we used only the sub-toxic concentrations which have negligible cytotoxicity based on the result of the apoptosis assay shown in Fig. 2. We first tested the effect of WF-A on collagen expression in cultured primary human lung fibroblasts (HLF) and primary fibroblasts from skin of scleroderma patients (SCL). These cells were treated with WF-A at low concentrations for 24hrs (Fig. 3A) and cell extracts

51 were analyzed for collagen α1(I) and αβ(I) polypeptides by Western blot. Withaferin-A dose dependently reduced production of both α1(I) and αβ(I) polypeptides from HLFs (Fig. γA, left panel) as well as from SCL fibroblasts (Fig. 3A, right panel). Actin levels weren’t affected and are shown as loading control. Densitiometric analysis of the bands is shown in the lower panel of Fig 3A. At 1.5µM, WF-A in HLFs, approximately β fold and γ fold reduction in α1 and αβ(I) collagen polypeptides is observed, respectively. Similarly, in SCL cells 1.0µM of WF-A resulted in approximately 4 fold reduction of both α1 and αβ(I) collagen polypeptides. Total RNA from the HLFs and SCL cells was also analyzed for the steady state levels of collagen mRNAs. Congruent with the effect on the polypeptides, WF-A significantly decreased the steady state levels of collagen α1(I) and αβ(I) mRNAs in a dose-dependent manner in both HLFs (Fig. 3B, left panel) and SCL fibroblasts (Fig 3B, right panel). The steady state of the control -actin mRNA was unaltered. Densitiometric analysis of the bands standardized for expression of actin mRNA is shown in the lower panel of Fig 3B. WFA reduced the steady state level of α1 collagen mRNA approximately γ fold, whereas it resulted in a 4 fold reduction of αβ collagen mRNA in both HLFs and SCL cells. To confirm the above results in cells of different tissue of origin, we prepared activated hepatic stellate cells (HSC) from rat livers [217] and cardiac fibroblasts (RCF) from rat hearts [218]. Activated HCSs were treated with WF-A at the indicated concentrations for 24hrs and whole cell extract and total RNA were analyzed for western blot and RT-PCR analysis. Withaferin-A caused 3-4 reductions in the expression levels of type I collagen both at the level of protein and mRNA (Fig.3C and D. left panels). Cardiac fibroblasts were isolated from rat’s heart and [218] and at the 3rd passage, the cells were treated with Withaferin-A at concentration of 0-1.5μM for β4hrs. The treatment of primary rat cardiac fibroblasts with Withaferin-A reduced the expression levels of type I collagen both at the level protein and mRNA 3-4 fold (Fig. 3C and D, right panels). From the results using different primary collagen producing cells, we concluded that WF-A can potently inhibit the expression of type I collagen regardless of the tissue of origin.

A HLF SCL B HLF SCL

COL α1 COL α1

COL α2 COL α 2

ACTIN ACTIN

WF-A (µM) 0 0.75 1.5 0 0.5 1.0 WF-A (µM) 0 0.75 1.5 0 0.5 1.0

COL α1 COL α2 COL α1 COL α2 1.2 1.2 1 HLF 1 HLF 0.8 * SCL 0.8 * SCL * 0.6 ** * * 0.6 * * 0.4 0.4 * * * * 0.2 0.2 * PROTEINCOL/ACTIN LEVEL:

0 COL/ACTIN EXPRESSION: 0 WF-A (µM) 0 low high 0 low high WF-A (µM) 0 low high 0 low high

C HSC RCF D HSC RCF

COL α1 COL α1

COL α2 COL α2

ACTIN ACTIN

WF-A (µM) 0 0.75 1.5 0 0.5 1.0 WF-A (µM) 0 0.75 1.5 0 0.5 1.0

COL α1 COL α2 COL α1 COL α2 1.2 1.2

1 1 * RCF HSC 0.8 * 0.8 * * * * RCF HSC 0.6 * 0.6 * * * * * 0.4 * 0.4 * * 0.2 0.2 *

PROTEINCOL/ACTIN LEVEL: 0 COL/ACTIN EXPRESSION: 0 0 low high 0 low high WF-A (µM) 0 low high 0 low high WF-A (conc)

Figure 13. Withaferin-A reduces expression of type I collagen in human and rodent fibroblasts. A. WF-A reduces collagen protein in human fibroblasts. HLF (left panel) and SCL fibroblasts (right panel) were treated WF-A and cellular levels of collagen α1(I) (COLα1) and αβ(I) (COLα1) polypeptides were analyzed by Western blotting. Loading control; actin. Bottom panels; the expression of collagen polypeptides was normalized to that of actin and plotted from three independent experiments. Open bars: HLF, black bars: SCL fibroblasts. Low concentrations of WF-A were 0.75 µM for HLF and 0.5 µM for SCL fibroblasts and high concentrations were 1.5 µM for HLF and 1.0 µM for SCL fibroblasts. The error bars representing +-1 SEM are shown. B. WF-A decreases the steady-state levels of collagen mRNAs in human fibroblasts. Experiment as in A, except mRNAs were analyzed by RT-PCR. C. WF-A reduces collagen protein in primary HSCs and primary cardiac fibroblasts. Rat hepatic stellate cells (HSC) and rat cardiac fibroblasts (RCF) were isolated from rat liver and heart, respectively, and treated with the indicated amounts of WF-A. Cellular levels of collagen α1(I) and αβ(I) polypeptides in HSCs (left panel) and RCF (right panel) were analyzed by Western blotting. Loading control, actin. Bottom panels: levels of collagen α1(I) (COLα1) and collagen αβ(I) (COLαβ) polypeptides were normalized to actin expression and plotted from three independent experiments. Open bars: HSC, black bars: RCF. Low concentrations of WF-A were 0.75 µM for HSC and 0.5 µM for RCF and high concentrations were 1.5 µM for HSC and 1.0 µM for RCF. The error bars representing +-1 SEM are shown. D. WF-A decreases the steady-state levels of collagen mRNAs in HSC and RCF. Experiment as in C, except mRNAs were analyzed by RT-PCR.

Withaferin-A increases the rate of decay of α1(I) and α2(I) collagen mRNAs Since WF-A disrupted vimentin filaments, we tested if the reduced level of collagen mRNAs following treatment with WF-A is due to their increased turnover. We determined the stability of collagen mRNAs after WF-A treatment and transcription block by actinomycin-D. Collagen mRNAs have half-lives between 18-27hrs in normal collagen producing cells [77]. In the DMSO treated HLFs, the half life of collagen α1(I) mRNA was estimated to be around β4hrs (Fig 4A, left panel). The left lower panel shows quantification densitiometric scan of the gels. The half life of collagen αβ(I) mRNA was estimated to be more than 24hrs, as the level drops to about 60% 24hrs after transcription block (Fig 4A, left panel and right lower panel for densitiometric analysis). However in the HLFs treated with 1.5 µM WF-A, the half-life of collagen α1(I) mRNA was reduced three fold to approximately 8hrs (Fig. 4A, right upper panel and left lower panel for quantification). The half-life of collagen αβ(I) mRNA was reduced 4 fold to about 6hrs. Similar results were obtained in scleroderma fibroblasts where the half-lives of both collagen α1(I) and αβ(I) mRNAs were reduced from β4hrs to about 9hrs (Fig. 4B).

A DMSO WF-A (1.5µM) COL α1

COL α2

ACTIN

Act-D (Hrs) 0 6 12 24 0 6 12 24

1.2 COLα1(I) 1.2 COLα2(I) DMSO WF-A WF-A 1 DMSO 1

0.8 0.8 * 0.6 0.6 * 0.4 * 0.4 * * % RNA RemainingRNA % Remaining RNA % 0.2 * 0.2

0 0 Act-D (h) 0 6 12 24 Act-D (h) 0 6 12 24

B DMSO WF-A (1.0µM) COL α1

COL α2

ACTIN Act-D (Hrs) 0 6 12 24 0 6 12 24

1.2 COLα1(I) 1.2 COLα2(I) DMSO DMSO 1 1 WF-A WF-A 0.8 0.8 * * 0.6 0.6

0.4 * 0.4

0.2 * 0.2 % RNA Remaining RNA % RemainingRNA % 0 0 Act-D (h) 0 6 12 24 0 6 12 24 Act-D (h) Figure 14: Withaferin-A destabilizes collagen 1(I) and 2(I) mRNAs in human lung and α α scleroderma fibroblasts. A. Stability of type I collagen mRNAs in Withaferin-A treated HLF. HLFs treated with DMSO (left panel) or 1.5µM WF-A (right panel) for 24h, transcription was blocked with Act D and collagen α1(I) and αβ(I) mRNAs were measured by RT-PCR at the indicated times after the transcription block. Actin mRNA was measured as a loading control. Bottom panels: decay rates of collagen α1(I) and αβ(I) mRNAs were estimated in DMSO and WF-A treated cells in three independent experiments. Expression of collagen mRNAs was normalized to the expression of actin mRNA and plotted as function of time after the transcription block. The levels at time 0 were arbitrarily set as 1. Error bars represent +- 1SEM. B. Stability of collagen mRNAs in WF-A treated SCL fibroblasts. The experiments as in A, except SCL fibroblasts were used.

We, therefore, demonstrated that Withaferin-A reduces stability of collagen mRNAs in two types of fibroblasts and that the downregulation of collagen expression by WF-A is, at least in part, mediated by its ability to reduce the half life of collagen mRNAs. The above results are consistent with the report that cells in which vimentin filaments are disrupted, have greatly reduced half-life of collagen mRNAs [210]. It is therefore likely that the destabilizing effect of Withaferin-A on the collagen mRNAs is mediated by its ability to disrupt vimentin filaments. The Effect of Withaferin-A on collagen mRNA stability is dependent on its effect on vimentin Withaferin-A may have variety of intracellular effects not related to disrupting vimentin filaments as suggested by its ability to induce apoptosis at higher concentrations. Although our results clearly show that WF-A reduced the stability of collagen mRNA, it is possible that the effect of WF-A on half-life of collagen mRNAs may be independent of the integrity of vimentin filaments. To test if targeting vimentin is a prerequisite for effects of WF-A on stability of collagen mRNAs, we compared effect of WF-A on collagen expression in VIM+/+ MEFs and VIM-/- MEFs. In VIM+/+ MEFs, WF-A caused a dose dependent reduction in the levels of collagen α1(I) and αβ(I) polypeptides (Fig. 5A, left panel). Densitiometric analysis showed that at 1.0µM of WF-A, there was about 80-90% reduction. In VIM-/- MEFs, at 1.0µM of WF-A, there was 60-70% reduction (Fig. 5A). This result suggested that WF-A reduces collagen expression in VIM+/+ cells to a greater extent than in VIM-/- cells. Similar result was obtained when collagen mRNAs were analyzed, suggesting some of the effects of WF-A on collagen expression are dependent on presence of vimentin filaments (Fig. 5B). We then wanted to determine whether transcription or mRNA stability is affected in WF- A treated VIM-/- MEFs. To this end, we compared the effect of WF-A on stability of collagen mRNAs in VIM-/- MEFs to that of VIM+/+ MEFs. In VIM+/+ MEFs treated with 1.0 µM WF- A, the half life of both α1(I) and αβ(I) collagen mRNAs was reduced by about 50% (Fig. 5C). However, in a similar experiment in VIM-/- cells, WF-A did not affect the half life of either collagen mRNA (Fig. 5D). Since WF-A didn’t alter half-lives of collagen mRNAs in the absence of vimentin, we concluded that the effect of WF-A on stability of collagen mRNAs is dependent on its effect on vimentin filaments and that this is a reason why we observed a greater effect on collagen expression in Vim+/+ cells.

A VIM+/+ VIM-/- VIM+/+ VIM-/- B Col-α1 Col-α1

Col- 2 Col-α2 α

Actin Actin

WF-A 0 0.5 1.0 0 0.5 1.0 WF-A 0 0.5 1.0 0 0.5 1.0

COLα1(I) COLα2(I) 1.2 1.2 COLα1(I) COLα2(I) 1 1 Vim+/+ VIM+/+ 0.8 0.8 * * Vim-/- VIM-/- * * 0.6 * 0.6 * * * * 0.4 * 0.4 * * * * 0.2 0.2 *

PROTEINCOL/ACTIN LEVEL: 0 COL/ACTIN EXPRESSION: 0 WF-A (µM) 0 0.5 1.0 0 0.5 1.0 WF-A (µM) 0 0.5 1.0 0 0.5 1.0

VIM -/- VIM +/+ D C DMSO WF-A DMSO WF-A COL α1 COL α1 COL α2 COL α2 ACTIN ACTIN Act-D (Hrs) 0 12 24 0 12 24 Act-D (Hrs) 0 12 24 0 12 24

1.2 1.2

VIM +/+ 1 1 VIM -/-

0.8 0.8

* 0.6 0.6 * 0.4 0.4 * % RNA Remaining RNA %

% RNA Remaining RNA % 0.2 0.2 Col-a1(I)-DMSO Col-a1(I)-WF-A * Col-a1(I)-DMSO Col-a1(I)-WF-A Col-a2(I)-DMSO Col-a2(I)-WF-A Col-a2(I)-DMSO Col-a2(I)-WF-A 0 0 12 24 0 0 12 24 Act-D (h) Act-D (h)

Figure 15. Effects of WF-A on collagen expression in vimentin knock out mouse embryonic fibroblasts. A. Vimentin knock out fibroblasts are less responsive to the effects of WF-A. Left panel: wt mouse embryonic fibroblasts (MEFs) (VIM+/+) and vimentin knock out MEFs (VIM-/-) were treated with WF-A and collagen α1(I) and αβ(I) polypeptides analyzed by Western blotting. Loading control: actin. Bottom panel: the expression of collagen α1(I) and αβ(I) polypeptides from VIM+/+ MEFs (open bars) and VIM-/- MEFs (black bars) were normalized to actin expression the results from three independent experiments were plotted. The error bars represent +-1 SEM. B. Effect of WF-A on expression of collagen mRNAs in VIM+/+ and VIM-/- MEFs. Experiment as in A, except mRNAs were analyzed by RT-PCR. C. WF-A destabilizes collagen mRNAs in VIM+/+ MEFS. VIM+/+ MEFs were treated with DMSO (left panel) or 1.0µM WF-A (right panel) for 24h. Transcription was then blocked by actinomycin D (Act-D) and collagen α1(I) and αβ(I) mRNAs were measured at the indicated times after the block by RT-PCR. Actin mRNA was measured as a loading control. Bottom panel: VIM+/+ MEFs were treated with WF-A (full lines) or DMSO (broken lines) and the levels of collagen α1(I) mRNA (circles) and αβ(I) mRNA (squares) were normalized to actin mRNA and plotted for the indicated time points after the transcription block. The level at time point 0 was arbitrarily set as 1. The error bars represent +-1 SEM, estimated from three independent experiments. D. WF-A has no effect on stability of collagen mRNAs in VIM-/-

MEFs. Experiment as in C, except in VIM-/- MEFs were used.

Withaferin-A inhibits the TGF-1 induced type I collagen expression and COL1Aβ promoter activity The observation that WF-A affected, the expression of collagen in VIM-/- MEFS, although to a lesser extent, suggested that the mechanism by which Withaferin-A decreases expression of collagen mRNAs is not only limited to decreasing the half-lives. Another mechanism could also contribute to the effect, thus, we decided to investigate if WF-A exerts its anti-fibrotic effects also at the level of transcription. Transcription of collagen genes can be stimulated by TGF- through the TGF-/Smad signaling pathway. Stimulation of HLFs by TGF-1 increased levels of collagen α1(I) and αβ(I) polypeptides by 3.0 and 2.8 fold, respectively (Fig. 6A). However, TGF-1 induced increase in expression of collagen α1(I) and αβ(I) polypeptides was abolished in cells pretreated with 1.5µM WF-A (Fig. 6A, lane3). We observed that WF-A, not only abolished the induction of collagen by TGF-, but also resulted in a further decrease to the level below that of control cells. Consistent with the effect of TGF- on the polypeptide levels, RT-PCR analysis revealed 2.5 and 2.0 fold increase in the steady state levels of collagen mRNAs after TGF- stimulation. This induction

58 was completely abolished in cells pretreated with WF-A (1.5µM) (Fig. 6B, lane 3) and WF-A pretreatment reduced TGF-induced expression to a level lower than that of unstimulated cells. TGF- stimulates the transcription of collagen αβ(I) gene (COL1Aβ) by inducing binding of Smad-containing complex to the TGF- response element (TbRE), located between nucleotides - 313 and -250 of the promoter. To determine if WF-A can inhibit the TGF-1 stimulated transcriptional activity of COL1A2 promoter, primary HLFs were transiently transfected with a -378COL1A2/LUC reporter gene. This gene was driven by the wild type COL1A2 promoter (Col-α2(wt)-Luc) and control gene had mutated TbRE element of the COL1A2 promoter (Col- αβ(mut)-Luc). These reporter genes have been described before[219]. Twenty-four hrs after transfection, cells were treated with WF-A (1.0µM) followed by stimulation with TGF-1 (5ng/ml). The mutant COlαβ(I) promoter failed to respond to TGF- (Fig. 6C), expression of LUC protein was the same in stimulated and unstimulated cells. TGF-1 stimulated the transcriptional activity of the wild type COL1A2 promoter by 3.5 fold compared to unstimulated cells (Fig. 6C). Treatment of cells with 1.0µM of WF-A completely inhibited the stimulation of transcription from the wild type COL1A2 promoter construct.

A 1 2 3 B 1 2 3 COL α1 COL α1

COL α 2 COL α2

ACTIN ACTIN

TGFβ (5ng/ml) - + + TGFβ (5ng/ml) - + + WF-A (1.5µM) - - + WF-A (1.5µM) - - +

3.5 * * 3 3 2.5 * 2.5 * COL-a1(I) 2 COL-a1(I) COL-a2(I) 2 COL-a2(I) 1.5 1.5 # 1 # 1 ## ** * 0.5 0.5 * EXPRESSION: COL/ACTIN EXPRESSION: PROTEINCOL/ACTIN LEVEL: 0 0 TGFβ - + + TGFβ - + + WF-A - - + WF-A - - +

C 4.5 4

gal) * Col-a2(I)-wild β 3.5 Col-a2(I)-mut 3 2.5 2 # 1.5 1 0.5 Fold inductionFold (Luc/ 0 WF-A(1.0µM) - - + + - - + + Tgfβ (5ng/ml) - + - + - + - +

D 9 8 * DMSO gal) β 7 WF-A (0.75) 6 # WF-A (1.5) 5 4 * # 3 * 2 1 Fold inductionFold (Luc/ 0 Withaferin-A - - + + + + Tgfβ (5ng/ml) - + - + - +

Figure 16: Withaferin-A inhibits TGF-β1 induced expression of type I collagen. A. WF-A inhibits TGF-1 induced increase in collagen polypeptides. HLFs were treated with DMSO (lanes 1 and 2) or 1.5µM WF-A (lane 3) for 1h and then treated with TGF- 1 (lanes β and γ) for β4h. Expression of collagen α1(I) (COLα1) and collagen αβ(I) (COLαβ) polypeptides in cell lysates was measured by Western blot. Loading control, actin. Bottom panel: levels of collagen α1(I) and αβ(I) polypeptides were normalized to that of actin and plotted from three independent experiments. The error bars represent +-1 SEM. B. WF-A inhibits the induction of collagen mRNAs by TGF-1. Experiment as in A, except collagen mRNAs were analyzed by RT-PCR. Loading control, actin. Bottom panel: levels of collagen α1(I) and αβ(I) mRNAs were normalized to that of actin mRNA and plotted from three independent experiments. The error bars represent +-1 SEM. C. WF-A inhibits transcriptional activation of collagen αβ(I) promoter. Luciferase reporter gene containing wild type collagen αβ(I) promoter (open bars) or collagen αβ(I) promoter with mutated TGF--responsive element (black bars) were transfected into HLF. After treatment with WF-A and TGF-1, luciferase activity was normalized to the contransfected internal control gene -GAL and plotted from three independent transfections performed in duplicate. Error bars represent +-1 SEM. D. WF-A inhibits transcriptional activation of SMAD responsive promoter. Experiment as in C, except reporter gene containing (CAGA)12MLP promoter was used and WF-A and was used at 0.75µM (gray bars) and 1.5µM (black bars).

To confirm the above result, we used another TGF- responsive reporter construct, containing 12 repeats of the Smad-binding element CAGA, (CAGA)12MLP-Luc. TGF- induced this promoter 6fold in HLFs, but the induction was inhibited by about 50% in cells treated with 0.75 µM of WF-A and by about 75% with 1.5µM of WF-A (Fig. 6D). This result suggests that, in addition to destabilizing collagen mRNAs by disrupting vimentin filaments, WF-A also interferes with TGF- induced transcription of collagen genes. Withaferin-A blocks Smad3 phosphorylation To gain insight in to the step at which WF-A interrupts TGF- signaling, we assessed the effect of WF-A treatment on the phosphorylation of Smad3. Phosphorylation of Smad3 was assessed by Western blot using phospho-Smad3 specific antibody that detects phosphorylation at S423/S425. Treatment of HLFs with 5ng/ml TGF-1 induced phosphorylation of Smadγ within 30 minutes after treatment and the phosphorylation persisted for 60 minutes (Fig. 7A, left panel). WF-A pretreatment abrogated TGF-1 induced Smadγ phosphorylation by ~ 50% at γ0 minutes (Fig 7A, right panel) and the effect was slightly more at 60 minutes. The level of total Smad3 was not altered by WF-A. The decreased Smad3 phosphorylation following treatment with WF- A suggests that WF-A can interferes with activation of TGF- transcription factors. This was already suggested using reporter gene with COL1A2 promoter (Fig 6). In cultured fibroblasts, there is constitutive phosphorylation of Smad3, although to a much less extent than seen in cells stimulated by exogenous TGF- [220]. To determine if WF- A can reduce the level of constitutive p-Smad3 in cultured fibroblasts, we treated HLF with increasing concentrations of WF-A and analyzed pSmad3 by western blot. WF-A reduced the level of p-Smad3 in cultured HLFs in dose dependent manner (Fig. 7B). The total amount of Smad3 remained unaltered 24hrs after treatment of HLFs with the same concentrations of WF-A (Fig. 7B). Taken together, WF-A inhibited both the basal, as well as TGF-1 stimulated phosphorylation of Smad3, indicating that it interferes with the TGF- signaling pathway. Our data strongly suggests that WF-A inhibits the TGF- signaling pathway upstream of Smadγ phosphorylation. Together with the disruption of vimentin filaments, this may be an additional mechanism by which WF-A reduces collagen expression.

A DMSO WF-A B pSMAD3 pSMAD3

SMAD3 SMAD3

ACTIN ACTIN Time after WF-A (µM) 0 0.5 0.75 1.0 1.25 1.5 TGFβ 6 6 D C COL α1(III) 4.5 FIB 4 TGF-β bioassay 3.5 Smad2 UNCOND 3 WF-A 2.5 TGF-b1 TGFβ1

2 TGF R1 FOLD CHANGE CHANGE FOLD 1.5 β 1 TGF R2 β 0.5 ACTIN 0 WF-A (µM) - 0 0.5 0.75 1.0 1.25 1.5 - - - - - WF-A (µM) 0 0.75 1.5 TGF-β1 (ng/ml) ------0.05 0.1 0.2 0.4 0.8

E F 1 2 3 4 1 2 3 4 COL α1 COL α1

COL α2 COL α2

ACTIN ACTIN P38i - + - + P38i - + - + WF-A - + + - WF-A - - + +

1.2 1.2

1 COLa1 1 COLa1 COLa2 COLa2 0.8 0.8 * 0.6 * 0.6 * * * * 0.4 * 0.4 * # # # # 0.2 ** 0.2 ** ** EXPRESSION: COL/ACTIN EXPRESSION: EXPRESSION: COL/ACTIN EXPRESSION: ** 0 0 P38i - + - + P38i - + - + WF-A - - + + WF-A - - + + 62

Figure 17. WF-A inhibits TGF-β signaling. A. WF-A inhibits TGF-1 induced Smadγ phosphorylation. HLF were treated with DMSO (left panel) or WF-A at 1.5 µM (right panel) and stimulated by TGF-1. Phospho-Smad3 and total Smad3 were measured by western blot after the indicated time periods.. Loading control: actin. B. WF-A inhibits constitutive Smad phosphorylation in a dose dependent manner. HLF were treated with the indicated

concentrations of WF-A and phospho-Smad3 and total Smad3 were measured by western blot. Loading control: actin. C. WF-A does not alter the amount of active TGF- produced by HLFs. HLFs were treated with the indicated concentrations of WF-A for 24h and the conditioned medium was added to Mink lung epithelial cells (MLEC) stably transfected with a luciferase reporter gene driven by TGF- responsive promoter (PIA-Luc). Luciferase activity was measured and normalized to… Gray bar; unconditioned medium, black bars; HLF conditioned medium, white bars: MLEC treated with the indicated amounts of pure TGF-1. Each bar represents three independent experiments analyzed in duplicate. Error bars respresent + 1SEM. D. Effect WF-A on expression of other genes. Type III collagen (COL α1(III)), fibronectin (FIB), Smad2, TGF-1, TGF- receptor 1 (TGFR1), TGF- receptor β (TGFRβ) and connective tissue growth factor (CTGF) mRNAs from control cells and HLFs treated with 0.75 or 1.5µM of WF-A were analyzed by RT-PCR. Loading control: actin. E. Inhibition of p38 MAPK augments the antifibrotic effect of Withaferin-A. HLF were treated with p38 inhibitor SB203580 (10µM, lane 2), WF-A (1.5µM, lane 3) or both (lane 4). Lane 1 are control DMSO treated cells. Expression of collagen α1(I) (COLα1) and αβ(I) (COLαβ) mRNAs was analyzed after 24h by RT-PCR. Loading control: actin mRNA. Bottom panel: expression of collagen α1(I) mRNA (open bars) and αβ(I) mRNA (black bars) were normalized to that of actin mRNA and plotted from three independent experiments. The error bars represent +-1 SEM. F. p38 MAPK inhibitor complements the antifibrotic effect of Withaferin-A in SCL fibroblasts. Similar experiment as E, except scleroderma fibroblasts were used and the concentration of WF-A was 1.0 µM.

It has previously been suggested that WF-A may decrease the release of cytokines from macrophages and other inflammatory cells [221]. It is, thus, possible that WF-A may interfere with synthesis, release and activation of latent-TGF- from the cultured fibroblasts. In order to exclude this alternative explanation, we compared the amount of active TGF-1 secreted from control and WF-A treated fibroblasts. To this end, we utilized the TGF- bioassay, where conditioned medium from fibroblasts is transferred to ming lung epithelial cells (MLEC) stably expressing TGF- inducible Luc reporter gene. This assay is an established way of measuring the activity of TGF- and has been described before [222]. As a test of functionality of the assay, we treated MLECs with increasing concentrations of pure TGF-1. Adding exogenous TGF- at low concentrations increased luciferase expression up to 4 fold (Fig 7C, white bars). The conditioned medium of control cells or cells treated with WF-A in concentrations from 0 to 1.5µM increased the reporter expression 2 fold compared to the unconditioned medium (Fig. 7, black bars). Since

63 there was no difference whether the cells were treated with WF-A or not, this excludes the possibility that WF-A inhibits activity or secretion of the TGF- from cultured fibroblasts. Another possible mechanism by which WF-A can inhibit phosphorylation of Smad3 is to down regulate the expression of TGF- receptors, Smadβ or even TGF- itself. RT-PCR analysis showed that WF-A didn’t affect the levels of expression of TGF-1, TGF-R1, TGF-Rβ and Smad2 (Fig. 7D), ruling out the possibility of indirect effect by down-regulating the expression of molecular components of the TGF- signaling pathway. To further confirm that WF-A inhibits TGF- signaling pathway, we evaluated if WF-A alters the expression of 2 other genes that are targets of the TGF- signaling pathway, type III collagen and fibronectin. The steady state levels of type III collagen and fibronectin, were significantly reduced (Fig. 7D). The selective effect on extracellular matrix genes that are regulated by TGF- pathway suggests the relatively specific interruption of the TGF- signaling pathway by WF-A. In addition to the canonical TGF-/Smad pathway, TGF- also can induce the activation of the p38 MAPK pathway mediated by the upstreatm TGF--activated kinase 1(TAK1), a member of the MAPKKK family [204]. The TGF-/pγ8-MAPK signaling pathway has been implicated in TGF- induced collagen expression. Based on our finding that Withaferin-A inhibited the TGF-/smad pathway, we surmised that WF-A may also inhibit the TGF- /pγ8MAPK pathway. To test this, we treated HLF with 10µM SBβ0γ580, a specific pγ8/MAPK inhibitor [223], for 1hr before WF-A was added into the medium for additional 24hrs. The combination of p38-MAPK inhibitor (SB203580) and WF-A reduced collagen synthesis more than either compound individually (Fig. 6E, lane4), suggesting that WF-A does not affect the TGF-/pγ8MAK pathway. The combination resulted up to 10 fold reduction in the steady state levels of collagen α1(I) and αβ(I) mRNAs (Fig. 7E). Similar experiments in SCL fibroblasts confirmed the potent additive effect of a combination of WF-A and p38MAPK inhibitor on collagen expression (Fig. 7F). Taken together, our results show that WF-A inhibits the TGF-/Smad arm of the TGF- signaling pathway at a level upstream of Smad3 phosphorylation and that a combination of WF- A with p38MAPK inhibitor confers an additive effect on collagen expression.

Withaferin-A inhibits in vitro activation of HSCs and decreases collagen production from HSCs Following liver injury, hepatic stellate cells undergo activation, which refers to a transition from quiescent vitamin A-rich cells into proliferative, fibrogenic, and contractile myofibroblasts. Activated hepatic stellate cells are the sources of excessive ECM in chronic liver diseases and are, thus, the most relevant cell types for development of hepatic fibrosis [16, 178]. The process of activation of HSCs is mediated by several cytokines among which the most important being TGF-[224]. Since our results demonstrated that WF-A can inhibit the TGF- signaling pathway, we further examined if WF-A can also inhibit the culture activation of quiescent hepatic stellate cells isolated from rat liver. After isolation, HSCs were cultured for two days, then DMSO or WF-A (0.25µM) were added to the medium. After incubation for additional 4 days, cells were harvested and total RNA analyzed for α-smooth muscle actin (α- SMA). α-SMA is a marker of activation of HSCs. Fig. 8A shows that WF-A at 0.25µM significantly down regulated the expression of α-SMA in cultured primary HSCs suggesting that WF-A inhibited the activation of quiescent HSCs in culture. WF-A treated HSCs also expressed dramatically lower levels of collagen α1(I) and αβ(I) compared to the controls cells (Fig. 8A). To determine if WF-A can inhibit the expression of type I collagen in partially activated HSCs, rat HSCs were cultured for 4 days and then treated with WF-A at the indicated concentrations (Fig. 8B). After 2 additional days, the cells were harvested for mRNA and protein analysis. RT-PCR analysis for α-SMA revealed that the level in WF-A treated cells was comparable to that of control cells, suggesting that WF-A may inhibit the activation process on at an early stage. However, WF-A was still able to significantly decrease the levels of mRNAs for both collagen α1(I) and αβ(I), with the more noticeable effect on αβ(I) mRNA (Fig. 8B). Protein analysis by western blot showed similar reduction in collagen expression as the RT-PCR analysis (Fig. 8C). From the experiments on isolated rat HSCs, we concluded that WF-A inhibits the activation of quiescent hepatic stellate cells in culture, consequently preventing the upregulation collagen in these cells. Treatment of partially activated HSCs with WF-A also inhibits collagen production without affecting the expression level of α-SMA.

A B C Col- 1 α Col-α1 Col-α1

Col-α2 Col-α2 Col-α2 α-SMA -SMA α-SMA α GAPDH

Actin Actin Actin

WF-A 0 0.25 0.5 WF-A 0 0.25 0.5 WF-A 0 0.25 th th (at 4 day) (at 2nd day) (at 4 day)

Figure 18. Effect of WF-A on quiescent and partially-activated primary rat HSCs. A) WF- A delays activation of quiescent HSCs. The isolation of the rat HSCs was described in the methods. Isolated cells cultured for 2days were treated either with DMSO or WF-A (0.25µM). At the 6th day, cells were harvested and total RNA analyzed for collagen α1(I), collagen αβ(I), and α-smooth muscle actin (α-SMA) mRNAs by RT-PCR. Loading control: actin mRNA. B) WF-A decreases collagen expression from partially-activated primary rat HSCs. Isolated cells cultured for 4days were treated with DMSO or with the indicated concentrations of WF-A. At the 6th day, cells were harvested and total RNA analyzed for collagen α1(I), collagen αβ(I), and α-smooth muscle actin (α-SMA) mRNAs by RT-PCR. Loading control: actin mRNA. C) WF-A deceases collagen protein from partially activated primary HSCs. Same as B, except that lysates were analyzed for collagen α1(I) and αβ(I) polypeptides and α-smooth muscle actin (α-SMA) by Western blot. Loading control: actin.

Withaferin-A inhibited isoproterenol-induced myocardial fibrosis in mice To validate if WF-A has an effect on fibrosis in vivo, we investigated if Withaferin-A treatment can attenuate fibrosis in a mouse model of isoproterenol induced myocardial fibrosis. Isoproterenol is a well known -adrenergic agonist that has been used to induce experimental myocardial injury and fibrosis in mice and rats [225-228]. Twelve weeks old (25-30 grams) 129Svev male mice (n=24) were randomly divided into four groups; Isoproterenol (ISO) group (n=6), Isoproterenol and Withaferin-A (ISO+WF-A) group (n=6), Withaferin-A (WF-A) group (n=6) and vehicle (CONT) group (n=6). Isoproterenol (50mg/kg) was injected subcutaneously to

66 the ISO and the ISO+WF-A groups for 2 consecutive days, whereas mice in the control groups received subcutaneous saline injections. WF-A (4mg/kg) was injected intraperitoneally daily for 14days, where as the vehicle was injected into the control groups. Twenty four hrs after the last injection, hearts were removed, weighed, fixed and processed for histopathologic analysis. In addition, piece of heart tissue was snap frozen for analysis of protein and RNA. The heart weight to body weight ratio (HW/BW ratio) didn’t significantly differ among the groups (Fig. 9A). However, the isoproterenol administration resulted in mild to moderate fibrosis. Two representative sections, one from the middle of the heart around the level of papillary muscles, and the other from the apex stained with Masson’s trichrome are shown for each group (Fig. 9B). WF-A only group showed no fibrosis and histology was indistinguishable from the control group (not shown). The percent of fibrosis area was determined from the extent of the Masson’s trichrome staining by one of the authors (J. Blackmon), blinded as to the groups. The percentage of fibrosis was also analyzed by image-J software. Treatment of mice with Withaferin-A (4mg/kg for 14 days) significantly ameliorated the development of isopoterenol- induced myocardial fibrosis (Fig. 7B, compare the (ISO) and (ISO+WF-A) sections). The % of fibrosis area estimated from 6 animals was ~7%, compared to ~ 3.5% in the ISO+WF-A groups (p<0.001) (Fig. 9C). This result verified that Withaferin-A can reduce cardiac fibrosis by 50%.

A ISO ISO+WFA WF-A CONT (n=6) (n=6) (n=6) (n=6) BW (g) 26.5 + 2.5 26.0 + 1.5 25.5 + 1.5 27.0 + 2.0 HW (mg) 140 + 5 135+ 5 140 + 5 145 + 5 HW:BW 0.189 0.192 0.175 0.186

B CONT ISO ISO+WF-A

2.0 mm APEX SECTIONAPEX SECTION MID

2.0 mm C 9 8 * 7 6 5 * # 4 3 2 FIBROSIS AREA% 1 0 ISO I SO+WF-A WF-A VEHIC

Figure 19. Withaferin-A inhibits isoproterenol-induced myocardial fibrosis in mice. A. Mice were injected with isoproterenol (ISO), isoproterenol + WF-A (ISO + WF-A), WF-A alone (WF-A) or vechicle (CON). Heart weight (HW), body weight (BW) and the ratio of heart B. weight to body weight (HW:BW) is shown as the mean +_1 SEM. WF-A decreased isoproterenol induced myocardial fibrosis in mice. Masson’s trichrome staining of the base of the ventricle sections of hearts of mice treated with ISO and with ISO + WF-A. Two representative sections are shown. Scale bar: 2mm. C. Morphometric analysis of the percentage of fibrotic area of the hearts. Fibrosis areas (blue) within the sections were quantified using Image J software. The percentage of fibrosis was estimated from ventricular and apex sections of each heart. Data represented mean +-1 SEM of six mice per group. 68

To further corroborate the effect if WF-A on cardiac fibrosis, we analyzed expression of type I and type III collagen, and α-SMA in tissue extracts by western blot and RT-PCR. Collagen α1(I), collagen α2(I), and collagen α1(III) mRNAs were reduced 2 fold in the ISO+WF-A group compared with the ISO group (Fig. 10A). Expression of α-SMA was also decreased in WF- A+ISO treated animals compared to ISO-treated animals and was similar to that of control animals (Fig 10.B). This suggested that WF-A almost completely prevented differentiation of myofibroblasts in heart tissue, in agreement with the inihibition of differentiation of HSC (Fig. 8). Western blot analysis for type I and type III collagen showed similar results as that obtained from the mRNA analysis (Fig.10C). Collagen polypeptides were decreased by ~50%. This biochemical analysis verified a significant effect of WF-A on fibrosis in vivo and showed that WF-A can suppress collagen expression in vivo, as observed in the cultured cells. αSMA expression was also downregulated in the ISO+WF-A group compared to the ISO group at the protein level (Fig. 10C).

CONTROLS A ISO ISO + WF-A WF-A VEHICLE

COLα1(I)

COLα2(I)

ACTIN

1 2 3 4 5 6 1 2 3 4 5 6 1 2 3 1 2 3

2.5 *

2 * * COL a1(I) * COL a2(I) 1.5 # # 1

0.5

EXPRESSION: COL:ACTIN EXPRESSION: 0 ISO ISO+WF-A WF-A VEHIC

CONTROLS B ISO ISO + WF-A WF-A VEHICLE

COLα1(III)

α-SMA

ACTIN

1 2 3 4 5 6 1 2 3 4 5 6 1 2 3 1 2 3 4 3.5 *

3 * 2.5 * COLa1(III) 2 * # α-SMA # 1.5

0.5

EXPRESSION:ACTIN 0 ISO ISO+WF-A WF-A VEHIC

CONTROLS C ISO ISO + WF-A WF-A VEHICLE COL α1(I)

COL α1(III)

α-SMA

ACTIN 1 2 3 4 5 6 1 2 3 4 5 6 1 2 3 1 2 3 4.5 4 * 3.5 Col-a1(I) 3 * * Col-a1(III) 2.5 * a-SMA * * # 2 # # 1.5 1 0.5 0 ISO ISO+WF-A WF-A VEHIC

Figure 20. Withaferin-A inhibits expression of α-smooth muscle actin and type I and type III collagens in isoproterenol induced cardiac fibrosis. A. Withaferin-A decreased

expression of collagen α1(I) and αβ(I) mRNAs in experimental cardiac fibrosis. Total RNA was extracted from the heart tissue of the study groups and analyzed by RT-PCR for expression of collagen α1(I) mRNA (COLα1), collagen αβ(I) mRNA (COLαβ) and α-smooth muscle actin mRNA (α-SMA). Loading control: actin mRNA. The numbers indicate individual animals in each experimental group. Bottom panel: Expression of collagen α1(I) mRNA (open bars) and

collagen αβ(I) mRNA (black bars) was normalized to actin mRNA and plotted for each group. Error bras represent +-1 SEM. B. Withaferin-A abrogated expression of collagen α1(III) and α- smooth muscle actin mRNAs in isoproterenol induced cardiac fibrosis. Experiment as in A, except collagen α1(III) mRNa (COLIII) and α-smooth muscle actin mRNAs (α-SMA) mRNA were analyzed. C. Withaferin-A reduced the isoproterenol induced increase in level of of

collagen α1(I) and α1(III) polypeptides and α-smooth muscle actin protein. Total protein extracted from the hearts of the study groups was analyzed for collagen α1(I) polypeptide, collagen α1(III) polypeptide and α-SMA protein by western blot. Loading control: actin. The numbers represent individual animals in the experimental groups. Bottom panel: protein expression was normalized to actin expression and plotted for the groups. Collagen α1(I) polypeptide; open bars, collagen α1(III) polypeptide; black bars and α-SMA protein; gray bars. Error bars: +-1 SEM.

3.2- Discussion Increasing understanding of the key factors involved in the regulation of normal and excessive collagen expression is helping lay out the frame work of potential targets for discovery of anti-fibrotic drugs. However, the discovery of antifibrotic interventions hasn’t kept pace with the progresses made in the understanding of molecular mechanisms that mediate the initiation and progression of fibrosis. Currently, no antifibrotic agents are licensed for use in humans [79, 150, 153, 208, 229, 230]. In fibroproliferative disorders, a profibrotic tissue injury is associated with activation of fibroblasts, which precedes the excessive synthesis and deposition of type I collagen in these disorders [182]. In activated fibroblasts, the increased expression of collagen is accountable to the increased rate of transcription of collagen genes as well as to the increased half-life of collagen mRNAs [85]. Efforts at targeting molecules involved in transcriptional and or posttranscriptional of regulation of expression of type I collagen could lead to potential antifibrotic therapies [79].

We recently reported that vimentin filaments bind and stabilize type I collagen mRNAs [210]. Withaferin-A, a bioactive substance obtained from a widely used Indian herbal plant Withania sominifera, was reported to disrupt vimentin intermediate filaments in endothelial cells, astrocytes and other vimentin containing cancer cell lines [212-214]. Based on our recent finding that the integrity of vimentin filaments is needed for stability of type I collagen mRNAs, we hypothesized that Withaferin-A treatment may reduce collagen expression in vitro as well as in vivo. No prior study has examined the effects of Withaferin-A on collagen synthesis or in animal models of fibrosis. In this study, we tested the potential anti-fibrotic effect in a tissue culture and animal model of tissue fibrosis. Our results from the tissue culture experiments using fibroblasts from different tissues (lung, skin, liver, heart) demonstrated that WF-A exhibits a potent antifibrotic activity that was achieved at a concentration lower than its minimal toxic effect. Importantly, our results provided insights into the mechanism of action of WF-A. Withaferin-A reduced the production of collagen from cultured fibroblasts through multiple mechanisms. First, WF-A caused decreased stability of collagen mRNAs by disrupting the vimentin filaments in fibroblasts. Second, WF-A inhibited the TGF- induced transcription activity of TGF--responsive promoters including that of COL1A2. This was associated with inhibition TGF- induced phosphorylation of Smad3. Last, WF-A also interfered with activation of quiescent hepatic stellate cells. The fact that Withaferin- A acted at multiple levels in the expression of collagen in vitro suggested that Withaferin-A may potentially be a robust anti-fibrotic agent in vivo. Regarding the effect on the stability of collagen mRNAs, the WF-A caused reductions in half lives of collagen mRNAs depended on the presence of vimentin. This was evidenced by the lack of effect of WF-A on collagen mRNA stability in VIM-/- fibroblasts. We reported that lack of vimentin or disruption of vimentin using dominant-negative desmin in fibroblasts resulted in decreased stability of type I collagen mRNAs [210]. The destabilizing effect WF-A on collagen mRNAs was, therefore, consistent with the key role of vimentin in posttranscriptional regulation of collagen expression. TGF- is synthesized as an inactive protein, named latent TGF- that contains a major region and latency associated peptide (LAP). LAP interacts with latent TGF- binding proteins (LTBP) and is anchored in the ECM. Activation of TGF- is possible by proteolytic cleavage, thrombospondin-1 (Tsp-1), plasmin, acidic microenvironments, 6 integrin and matrix

72 metalloproteinases [29, 231]. Three TGF- isoforms have been described, TGF-1, TGF-β, and TGF-γ. TGF-1 is the most important isoform in the cardiovascular system. It is expressed in endothelial cells, vascular smooth muscle cells, myofibroblats, and macrophages [232]. TGF-1 is the single most potent profibrogenic factor involved in the initiation and maintenance of fibrogenesis [131, 178, 224, 233-235]. TGF-1 predominantly transmits signals through the Smad proteins which translocate into the nucleus to activate transcription of target genes. Based on their function, the Smads are classified as receptor-activated Smads (R-Smads), common- partner Smads (Co-Smads) and Inhibitory Smads (I-Smads). R-Smads include Smad2, Smad3, Smad5 and Smad8, Co-Smads include Smad4, and I-Smads include Smad6 and Smad7. Smads 2 and 3 are specific mediators of TGF- signaling, whereas Smads 1, 5, and 8 are involved in BMP signaling. Smad4 continuously shuttles between the cytoplasm and the nucleus, and forms hetero-oligomers with R-Smads and is a common mediator of TGF- and BMP signaling. Smad7 binds to activated type I receptor, preventing phosphorylation of Smad2/3. Smad7 can also recruit the ubiquitin ligases Smurf1 and Smurf2 to induce proteasomal degradation of the receptor complexes [236]. Activation of Smads by TGF- plays an important role in cardiac remodeling and heart failure [237]. Approaches aimed at interfering with the TGF1 signaling pathway are being studied for a potential cure for cardiac fibrosis and fibrosis of other organs [150, 229, 230]. In vivo experiments using different strategies to block TGF1 have demonstrated significant antifibrotic effects in several organ systems including the liver, lung and heart [168, 238, 239]. Using luciferase reporter assays for assessing transcriptional activity of collagen genes in TGF- stimulated cells, we demonstrated that WF-A reduced the TGF- induced increase in transcription of collagen genes in fibroblasts resulting in downregulation of collagen expression both at the levels of mRNA and protein synthesis. We further investigated if repression the TGF- induced collagen expression was due to direct interruption of the TGF- signaling pathway. Our results confirmed that there is markedly decreased TGF- induced phosphorylation of Smad3 in cells pretreated with WF-A. Moreover, even in unstimulated fibroblasts, we showed that WF-A abolished the level of constitutive phospho-Smad3 in a dose dependent manner suggesting interruption of the autocrine TGF- signaling pathway. Yet another effect of WF-A was at the level of activation of quiescent cells. Our studies demonstrated that WF-A treatment abrogated activation of quiescent HSCs as indicated by the

73 reduced expression of α-smooth muscle actin (α-SMA) 7 days after isolation from rat liver compared to control cells. Since TGF- has a vital role in activation of quiescent fibroblasts [224], the delay in activation of HSCs corroborates our finding that WF-A inhibits the TGF- signaling pathway. There are two main TGF- receptors: TGF-RI and TGF-RII. Both are transmembrane receptors with serine-threonine kinase activity. Active TGF- binds to TGF-RII, causing a change in the receptor, which induces dimerization with TGF-RI and its phosphorylation. The dimerized complex transmits TGF- signaling into the cell [240]. In our efforts to exclude alternative explanations for the observed TGF- inhibiting effect of WF-A, we showed that WF- A didn’t significantly influence the level of mRNA, amount or activity of TGF- produced form fibroblasts. This was shown by the TGF- bioassay and RT-PCR analysis for TGF-1 transcripts. Elevated Smad expression has been described in different cardiovascular diseases including cardiac fibrosis [37, 38]. We therefore measured if the WF-A down regulated the level of expression of Smad2 in cultured fibroblasts. Our results showed level of Smad2 was not affected. Neither were mRNA levels of TGF- receptors altered, suggesting the effect is likely to be a direct one on the TGF-/smad pathway upstream of Smadγ phosphorylation instead of being mediated by blockage of another pathway. In line with the interruption of the TGF-/Smad pathway, the combination of WF-A with p38MAPK inhibitor (SB32580) afforded additive antifibrotic effects. Myocardial fibrosis, excessive accumulation of collagen fibers within the cardiac interstitium, occurs in several important cardiac diseases. Hypertensive heart disease (HHD) being the main one, others include myocardial infarction, hyperterophic cardiomyopathy, idiopathic intersitial cardiac fibrosis, and decompensated congestive heart failure of any ethiology [123, 241-243]. In hypertensive heart disease, type I and type III collagen content of the myocardium is significantly increased as a consequence of a number of hemodynamic, neuro-hormonal and cellular factors [111]. Two patterns of fiber accumulation are seen in HHD. One is a diffuse increase in density of collagen fibers referred to as interstitial fibrosis; the second pattern is localized deposition including perivascular fibrosis and microscopic scarring [58]. As a stiff fibrillar protein, excessive deposition of type I collagen contributes to the severe ventricular diastolic dysfunction seen in patients with hypertensive heart disease[35]. Importantly, the presence of fibrosis is associated with deterioration of cardiac function and

74 transition to heart failure in these patients [3]. In support of this, some antihypertensive drugs such as Lisinopril (ACE inhibitor) and Losartan (Aldosterone receptor blocker) have been shown to improve left ventricular function better than other drugs independent of their efficacy to lower blood pressure[130, 244]. This was attributed to the ability of these drugs to reduce myocardial fibrosis in addition to their antihypertensive effects. It has now become apparent that novel drugs and approaches to slow the progression of fibrosis in HHD patients are desirable [245]. In recognition of the importance of treating fibrosis in heart diseases, we studied the effect of WF-A administration on the development of isoproterenol induced myocardial fibrosis. Isoproterenol is a well-known -adrenergic agonist that produces positive inotropic and chrontropic effects. Single subcutaneous injection of isoproterenol (85mg/kg) in rats produced myocardial fibrosis[226]. Repeated daily injection of isoproterenol with wide range of doses (1- 100mg/kg) induces substantial LV fibrosis in mice [225]. We used a dose of 50mg/kg subcutaneous Isoproterenol for two days. With regards to the dose of Withaferin-A, previous studies have used 4-30 mg/kg WF-A, with no apparent toxicity to the animals [216, 246, 247]. In this study, the animals treated were treated with WA (4 mg/kg) for 2 weeks. WF-A treatment resulted in a significant reduction in the degree of myocardial fibrosis as assessed by histology (nearly 50%). Consistent with the histologic findings, analysis of tissue for proteins and mRNAs revealed abrogation of the induction of collagen α1(I), αβ(I), and α1(III) compared to mice treated with isoproterenol alone (ISO). In addition, there was a significant reduction in the level of α-SMA in WF-A treated mice compared to the ISO group. The significant inhibition of the induction in expression of α-SMA was in agreement with our in vitro studies. Our data thus demonstrated that WF-A has a strong and significant in vivo antifibrotic activity against isoproterenol induced cardiac fibrosis. The benefits of using natural products for therapies include history of safe use, and potential multiple mechanisms with possible synergistic outcomes [248]. Withaferin-A, as an ingredient of the common herbal plant, Indian winter cherry clearly offers the advantage of safety. Our results underscored the multiple steps at which WF-A can act to inhibit the initiation and progression of the process of fibrogenesis. Moreover, since Withaferin-A has been reported to display anti-inflammatory properties [249], its use may prove even more beneficial in fibrotic diseases with underlying chronic inflammation.

In summary, our studies reveal for the first time in vitro and in vivo evidences for the anti-fibrotic activities of Withaferin-A and its mechanisms of action. Our results show that WF- A possesses a potent in vitro, and in vivo antifibrotic activity. These findings provide a strong pre-clinical evidence base for the further exploration of Withaferin-A as a therapeutic drug against fibroproliferative diseases including but not limited to cardiac interstitial fibrosis either in preventive and/or curative settings. 3.3- Materials and Methods: Cells Primary human lung fibroblasts immortalized by expression of telomerase reverse transcriptase [205] were grown under standard conditions. Scleroderma fibroblasts derived from skin of a scleroderma patient were purchased from the European collection of cell cultures (cell line BM0070). Mouse embryonic fibroblasts derived from wild type (Vim+/+ MEF) and vimentin knockout mice (Vim-/- MEFs) were described before [206] and were kind gifts from Dr. Robert Evans, University of Colorado. Cardiac fibroblasts were isolated from 200–250 g female Sprague–Dawley rats as described [218] and cultured for 5days. They were then passaged 2-3 times before use for the experiments. All cells were cultured in Dulbecco’s Modified Eagle’s Medium, supplemented with 10% fetal bovine serum. Isolation and culture of hepatic stellate cells Rat HSCs were isolated by perfusion of the liver with collagenase and pronase, followed by centrifugation over a Nycodenz gradient, as described[217]. Isolated HSCs were cultured in DMEM supplemented with 10% FBS. Growth medium was changed every day until two days later when Withaferin-A was added to medium. Cells were harvested after additional 4days with total number of days in culture being 6. For some experiments, WF-A was added at day 4 when the cultured HSCs were partially activated. To obtain fully activated HSCs, cells were trypsinized after 7 days in culture and grown for additional three passages before being used for the experiments. Chemicals For the tissue culture experiments, Withaferin-A (Chromadex, Santa Ana, CA) was dissolved in DMSO and stored in -20 0C. Cultured cells were treated with DMSO or different concentrations of Withaferin-A (0.25-3.0µM) for 24hrs. For the in vivo experiments, Withaferin-A (Chromadex Santa Ana, CA) was dissolved in a vehicle containing 10% Ethanol, 40% DMSO and 50%

Cremephor as previously described [216]. Isoproterenol (Sigma-Aldrich) was dissolved in 0.9% saline for use in the in-vivo experiments. SB203580, a specific MAPK inhibitor [223], was obtained from Sigma and was used at a concentration of 10µM. Cell viability assay Apoptosis in fibroblasts following WF-A treatment was assessed by measuring Caspase 3 and 7 activities using a Caspase-Glo 3/7 assay kit (Promega, Madison, WI). Experiments were carried out by following the manufacturer's recommended procedures. Briefly, 2x104 cells/well were plated into white-walled 96-well plates and incubated for 24hrs under normal growth conditions. Subsequently, cells were treated with DMSO or various concentrations of WF-A (0.25-3.0µM), and the plates were incubated for additional 24hrs. After the 24 hrs incubation, the Caspase-Glo 3/7 reagents were added to the wells, and 1 h later, luminescence was measured with a plate-reading luminometer. Three separate experiments each in duplicates were performed and the data were expressed as relative light units and converted to % relative to untreated cells. TGF- stimulation of cultured cells HLFs were cultured in a low serum medium (1% FBS) for 24hrs. Cells were treated with DMSO or Withaferin-A(1.0-1.5µM) 1 hr before adding TGF-1 (R&D Systems, Minneapolis, MN) at a concentration of 5ng/ml. Incubation continued for additional 18-24hrs. Luciferase assay The wild type COL1A2-Luc construct is TGF- inducible Luciferase reporter that contains the sequence γ76 bp of the αβ(I) collagen (COL1Aβ) promoter and 58 bp of the transcribed sequence fused to the luciferase (Luc) reporter. The mutant COL1A2-Luc harbors a mutation in the Smad binding element within the TGF-beta response element. Both plasmids were kind gifts from Dr. Francesco Ramirez, Mount Sinai School of Medicine [219]. Another TGF- responsive plasmid (CAGA)12MLP-Luc, containing 12 CAGA boxes cloned upstream of the initiator sequence of the adenovirus major late promoter (MLP) was a kind gift of Dr. Peter ten Dijnke [250]. Primary HLFs were seeded at a density of 10,000 cells/cm2 and cultured in 1% FBS. Cells were transiently co-transfected using Lipofectamine β000 (Invitrogen) with 0.βµg of - galactosidase plasmid and 0.8µg of the Luciferase reporter plasmids described above. TGF-1 (5ng/ml) was added 24hrs after transfections and luciferase readings were taken 16-24hrs after later. In order to standardize results for transfection efficiency, the -galactosidase assay readings at 420nm were used.

TGF- bioassay Mink Lung Epithelial Cells (MLECs) stably transfected with TGF- responsive PAI-1-Luc (plasminogen activator inhibitor-1) were kind gifts of Dr. Daniel Rifkin, NYU. The use of these cells for measuring TGF-beta activity has been described [222]. Briefly, MLECs were seeded at 60,000 cells/cm2 and serum starved for 24hrs. The cells were then incubated with conditioned medium collected from cultured HLFs which had been treated with varying concentrations of WF-A. TGF- activity was assessed 16hrs later in cell lysates by measuring the luciferase activity using Luminometer. MLECs incubated with unconditioned medium and MLECs treated directly with increasing concentration of TGF-1 were measured as negative and positive controls, respectively. The results were performed in triplicate. Western blot analysis Cells were lysed using RIPA buffer containing 50mM Tris PH 7.4, 150mM NaCl, 0.5% sodium deoxycholate, 1% NP-40, 0.1% SDS, 1mM EDTA, and mix of protease inhibitors. Protein concentration was estimated by the Bradford assay, with bovine serum albumin as the standard. Forty to fifty (40-50) µg of total cellular protein was typically used for western blot analysis. Anti-Smad3 and anti-phospho-Smad3 Ser423/425 monoclonal antibodies were obtained from Cell Signaling. Anti-collagen α1(I) antibody was obtained from Rockland; anti-collagen αβ(I) antibody specific for human polypeptide was from Cell Signaling; the anti-collagen αβ(I) and the anti-collagen α1(III) antibodies to detect mouse polypeptides were obtained from Santa-Cruz Biotechnology. Anti-vimentin antibody, anti-smooth muscle actin and anti-actin antibodies were obtained from Abnova. For determination of PhosphoSmad3, cells were lysed with lysis solution containing also phosphatase inhibitors (NaF and Na3VO4). For samples from tissues, TRI-reagent (Sigma) was used to lyse and extract proteins from the snap-frozen heart tissue specimens. Reverse transcription-PCR analysis Total cellular RNA was isolated using an RNA isolation kit (Sigma) except for the in vivo studies where extraction was done by Tri-reagent (Sigma). RT-PCRs were performed with 100ng of total RNA and using rTth reverse transcriptase (Boca Scientific, Boca Raton, Fl). [32P]dCTP was included in the PCR step to label the products, which were resolved on sequencing gels, as described previously [77]. The number of cycles was adjusted within the linear range of the reaction. The primers used for RTPCR are shown in table-2.

Table 2: Primers used for RTPCR h-collagen α1(I) F: AGAGGCGAAGGCAACAGT nd, R: GCAGGGCCAATGTCTAGTCC h-collagen αβ(I) F: CTTCGTGCCTAGCAACATGC and, R: TCAACACCATCTCTGCCTCG h-collagen α1(III) F: ATCTTGGTCAGT CTATGCGG and, R: GCAGTCTAATTCTTGATCGTCA h-actin F: GTGCGTGACATTAAGGAGAAG and, R: GAAGGTAGTTTCGTGGATGCC h-fibronectin F: ACCAACTACGGATGACTCG and, R: GCTCATCATCTGGCCATTTT h-TGF-1 F: GGGACTATCATCCACCTGCAAGA and, R: CCTCCTT GCGTAGTAGTCG h-TGFR1 F: GGGGA ACAATACTGGCTGA and, R: GAGCTCTTGAGGTCCCTGTG h-TGFRII F: GCAAGTTTTGCGATGTGAGA and, R: GGCATCTTCCAGAGTGAAGC h-CTGF F: CGTACTCCAAAATCTCCA and, R: GTAATGGCAGGCACAGGTCT m-collagen α1(I) F: GAGCGGAGAGTACTGGATCG and, R: TACT GAACGGGAATCCATC m-collagen αβ(I) F: CTTCGTGCCTAGCAACATGC and, R: TCAACACCATCTCTGCCTCG m-collagen α1(III) F: ACGTAAGCACTGGTGGACAG and, R: AGCTGCACATCAACGACATC m-actin F: CGTGCGTGACATCAAAGAGAAGC and, R: TGGATGCCACAGGATTCCATACC m-fibronectin F: AATGGAAAA GGGAATGGAC and, R: CTCGGTTGTCCT CTTGCTC m-αSMA F: CTGACAGAGGCACCACTGAA and, R: CATCTCCAGAGTCCAGCACA m-TGF-1 F: TTGCTTCAGCTCCACAGAGA and, R: TGGTT T TGGGCA GGAC m-TGF-2 F: CCGGAGGTGATTTCCATCTA and, R: GCGGACGATTCTGAAGTAGG m-TGF-3 F: GATGAGCACATAGCCAAGCA and, R: GTGACATGGACAGTGGATGC m-TGFRI F: GGCGAAGGCATTACAGTCTT and, R: TGCACATACAAATCGCCTGT m-TGFRII F: GCAAGTTTTGCGATGTGAGA and, R: GGCATCTT C AGTGAA C m-smad2 F: GGAACCTGCATTCTGGTGTT and, R: ACGTTGGAGAGCAAGCCTAA m-smad7 F: CAGCTCAATTCGGACAACAA and, R: AACCAGGGAACACTTTGTGC m CTGF F: CAAAGCAGCTGCAAATACCA and, R: GGCCAAATGTGTCTTCCAGT r-collagen α1(I) F: TGAGCCAGCAGATTGAGAAC and, R: TGATGGCATCCAGGTTGCAG r-collagen αβ(I) F: CTCACTCCTGAAGGCTCTAG and, R: CTCCTAACCAGACATGCTTG r-actin F: CGTGCGTGACATTAAAGAGAAGC and, R: TGCATGCCACAGGATTCCATACC r-GAPDH F: ACCGGTTCCAGTAGGTACTG and, R: CTCACCGTCACTACCGTACC r-α-SMA F: ACAGAGAGAAGATGACGCAG and, R: GGAAGATGATGCAGCAGTAG F, forward primer; R, reverse primer; h, human; m, mouse; r,rat; and SMA, smooth muscle actin.

Determination of RNA stability Cells were treated with actinomycin D (10µg/ml) for 6, 12 and 24 hrs. After the actinomycin D incubation periods, the cells were scraped, an total RNA was extracted and analyzed by RT- PCR. RNA extracted from cells at time point 0 (immediately after the addition of actinomycin D) was used as the initial level of mRNA and arbitrarily set as 100%. Results from three independent experiments are used to plot the histograms. Immunostaining Cells were seeded onto glass coverslips. After treatment, the cells were fixed with 4% formaldehyde for 30min at room temperature and permeabilized with 0.5% Triton X-100 in phosphate buffered saline (PBS) for 10min. Blocking was done with 10% goat serum/5% bovine serum albumin in PBS for 1hr at room temperature, followed by incubation with anti-vimentin antibody overnight at 40c. After washing, cells were incubated with AlexaFluor594- conjugated secondary antibody diluted in a blocking solultion. Cells were, then, mounted using Prolong mounting solution containing 4’,6’-diamidino-2-phenylindole (DAPI) (Invitrogen). Images were taken by the Leica TCS SP2 AOBS laser confocal microscope equipped with a Chameleon Ti- Sapphire multiphoton laser. Optical sections were processed with LCSLite software and single plane confocal images are shown. In vivo experiment (study groups) Twelve weeks old (25-30 grams) 129Svev male mice (n=24) were obtained from the Charles River (MA, US). The mice were housed in a standard condition with 12 h dark-light cycle and were given standard diet with ad libitum access to tap water. The mice were divided into four groups randomly, Isoproterenol (ISO) (n=6), Isoproterenol and Withaferin-A (ISO+WF-A) (n=6), Withaferin-A (WF-A) (n=6) and vehicle (CONT) (n=6). The protocol for this study was approved by the Florida State University Animal Care and Use Committee. And all animal procedures and experiments were performed under the NIH guidelines for animal care and use. Isoproterenol (50mg/kg) was subcutaneously injected to the ISO and the ISO+WF-A groups for 2 consecutive days, whereas mice in the WF-A and CONT groups received subcutaneous 0.9% saline injections. WF-A (4mg/kg) was injected intraperitoneally to the ISO+WF-A group and to the WF-A group for 14days, whereas vehicle IP injections were given to the mice in the ISO and CONT groups.

Tissue sampling Twenty four hrs after the last injection (or 15days after the first injection), mice were sacrificed using overdose of pentobarbital (50mg/kg ip). The heart was exposed and removed via thoracotomy and placed in ice-cold normal saline. The great vessels were trimmed before the heart was weighed. The weight was used to determine heart weight to body weight (HW/BW) ratio. Then, the atria were removed and a transverse section at the level of the papillary muscles was snap-frozen in Liquid Nitrogen for western blot and RT-PCR analysis. The remaining ventricle including the apex was fixed overnight with 10% buffered formalin for histopathology analysis. Histopathology analysis Fixed heart tissue was embedded in paraffin and transverse sections (5 µm) from the apex of the ventricle as well as from the middle zone of the ventricle were placed on microscope slides. Slides were processed for Masson’s Trichrome staining. To determine the degree of cardiac fibrosis, lower magnification (20X) images were analyzed using image J soft ware with Threshold-color plug-in. The area of blue stained region (fibrosis) was divided by the total area to obtain % cardiac fibrosis. Image quantification Densitiometric analysis of scanned images from films (western blots) was performed by Image J soft ware. For the radio-labeled RT-PCRs, reactions run in 6% urea gel were exposed for autoradiography and quantified using typhoon phosphorimager. Statistical analysis Data is presented as mean + SEM. For the in vitro experiments, results from a minimum of three independent experiments were analyzed. Differences between two groups were analyzed for statistical significance using the Student’s t test. Comparisons among groups (i.e. more than two groups) were made by ANOVA method. Statistical values of p< 0.05 were considered to be significant.

CHAPTER FOUR GENERAL CONCLUSION

Withaferin-A reduced the production of collagen from cultured fibroblasts through multiple mechanisms. First, WF-A caused decreased stability of collagen mRNAs, this effect is due to the ability of WF-A to disrupt the vimentin filaments. WF-A induced destabilization of collagen mRNAs was seen only in fibroblasts containing vimentin. In VIM-/- fibroblasts, the half-life of collagen mRNAs wasn’t affected by WF-A. The destabilizing effect of WF-A on collagen mRNAs is consistent with the role of vimentin in posttranscriptional regulation of collagen expression. Second, WF-A inhibits the TGF- induced transcriptional activation of TGF--responsive promoters, including that of COL1A2. WF-A directly interrupted the TGF- signaling pathway, as demonstrated by the markedly decreased TGF- induced phosphorylation of Smad3. Moreover, even in unstimulated fibroblasts, WF-A abolished the level of constitutive phosphorylation of Smad3 in a dose dependent manner, suggesting interruption of the autocrine TGF- signaling pathway. Third, WF-A treatment abrogated culture activation of quiescent 82

HSCs into activated HSCs, as indicated by the reduced expression of the marker of activation, α- smooth muscle actin (α-SMA). The in vitro effect of Withaferin-A on collagen expression was reproduced in vivo in a mice model of isoproterenol-induced myocardial fibrosis. WF-A treatment resulted in a 50% reduction in the degree of myocardial fibrosis as assessed by evaluation of Masson’s trichrome stained heart sections. Use of WF-A, and other approaches to disrupt vimentin filaments, can be a promising therapeutic avenue for the treatment of fibrotic disorders.

APPENDIX A

ACUC APPROVAL

APPENDIX B

LIST OF ABBREVIATIONS

5’SL= 5’ stem loop α-SMA= alpha-smooth muscle actin HBV= Hepatitis B Virus HCV= Hepatitis C Virus HHD= Hypertensive heart disease HLF= Human Lung Fibroblasts HSC= Hepatic stellate cells IL-13= Interleukin-13 MEF= Mouse embryonic fibroblasts MMP= Matrix metalloproteinase LARP6= La ribonucleoprotein domain family 6 LV= Left ventricle RCF= Rat cardiac fibroblasts RNA-FISH= RNA- fluorescent in situ hybridization RV= Right ventricle SCL= Scleroderma TGF-= Transforming growth factor beta UTR= Untranslated region WF-A= Withaferin-A

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BIOGRAPHICAL SKETCH

PRESENT CONTACT ADDRESS: - 1115 W. Call St., Biomedical Sciences, College of Medicine, Florida State University, Tallahessee, FL 32306 DATE AND PLACE OF BIRTH: - 10/07/1981 G.C. Debrebzeit, Ethiopia. EDUCATION: - MD (Doctor of Medicine) (1998-2004), Gondar College of Medical Sciences, University of Gondar, Ethiopia - PhD (Biomedical sciences), (2007-2011), Department of Biomedical sciences, College of Medicine, Florida State University. EMPLOYMENT HISTORY: - Graduate Research Assistant (2007-2011): Biomedical sciences, College of medicine, Florida State University - Lecturer of Epidemiology and research methodology (2005 – 2007): College of Health Sciences, Mekelle University - General practitioner of medicine and Assistant lecturer of Pathology (2004-2005): Gondar University Hospital - Intern Physician (2003-2004): Internal medicine, Surgery, Obstetrics/Gynecology and Pediatrics, Gondar University Hospital, Gondar, Ethiopia. RESEARCH GRANTS: - Predoctoral Research Fellowship (2009-2011); American Heart Association. Targeting vimentin filaments for prevention and treatment of myocardial fibrosis: an in vivo study. 10PRE4490011 ($42,540.00 USD) - NORAD Young Researcher award (2005-2006); Norwegian Government. Hepatitis B in Pregnancy: the role of perinatal transmission in the acquisition of Hepatitis B. (PI) - Mekelle University Junior Investigator Grant (2005-2006). Seroprevalence of HBsAg and HBeAg among pregnant woman admitted to Obstetric unit, Mekelle Hospital, Ethiopia. (PI) PROFESSIONAL AND HONOR SOCIETY MEMBERSHIPS:

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- American Association for Advancement of Science (AAAS) (2009 – present) - Golden Key Honor society (2008 - present) - Ethiopian Medical Association (2005 - 2007) TEACHING EXPERIENCE: - Teaching assistant (2009-2011): Research techniques in biomedical sciences to 1st year PhD students (included giving class lecture, facilitating lab sessions). - Small group facilitator (2008-2009): Medical Microbiology, Clinical anatomy, and Pharmacology to Year I and Year II Med students; College of Medicine, FSU. - Lecturer (2005-2007): Epidemiology and Research methodology to Year II Med students, College of health sciences, Mekelle University, Ethiopia. - Lecturer (2005-2007): Epidemiology to Health science students, College of health sciences, Mekelle University, Ethiopia. - Assistant Lecturer (2004-2005): General Pathology to Year II Med students, Gondar College of Medical Sciences, University of Gondar, Ethiopia. AWARDS AND HONORS: - Outstanding graduate student award (2011), Department of biomedical sciences, Florida State University) for exemplary performance as a graduate student. - Honorable mention (2010), for a poster at Gordon Intermediate Filaments Research conference, Tilton, NH. - Shell Centenary Chevening Scholarship. (2007) for Masters of Public health at Edinburgh University, England. (country wide competition) (offer declined) - Best intern of the year (2004), Gondar University Hospital for outstanding performance as an intern physician (2 recipients among 60 practicing interns) - Other past awards: best freshman student of the year (1999) for highest academic score (4 recipients among 600 students); Very Great Distinction on Ethiopian School Leaving Certificate Examination (1998), Winner of regional Q and A competition (1997) LEADERSHIP ROLES AND COMMITTEE MEMBERSHIPs: - Head, Office of Community Based Training Program (CBTP) Coordination , Mekelle University, Ethiopia (2005-2007).

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- Focal person, Medical Students Exchange Program between Mekelle University and Texas Tech University, USA (2006-2007). - Committee member, Collaboration between Mekelle University and Tigray Regional Health Buerro (2006-2007) ORAL PRESENTATIONS AT SCIENTIFIC MEETINGS: 1- Biomedical Sciences, Departmental Seminar (2010): College of Medicine, Florida State University. Topic: Vimentin intermediate filaments and the regulation of collagen synthesis. 2- Annual Research Review (2007): College of Health Sciences, Mekelle University. Topic: Hepatits B in Pregnancy. 3- 15th Annual Staff and Student’s Research Conference (β005): University of Gondar. Topic: Bacterial meningitis beyond the neonatal period in Gondar University Hospital: retrospective study. Gondar, Ethiopia. 4- 1γth Annual Staff and Student’s Research Conference (β004): University of Gondar. Oral. Topic: Knowledge, Attitude and Practice of high-school students concerning tuberculosis in Dabat, North Gondar, Ethiopia. POSTER PRESENTATIONS AT SCIENTIFIC MEETINGS: 1- Keystone. Molecular cardiology and disease mechanisms (2011): Keystone, Colorado Title: Vimentin’s role in ECM production by cardiac fibroblasts. (abstract published) 2- Life science symposium (2010): Florida State University. Poster. Title: Posttranscriptional regulation of type I collagen expression. (abstract published) 3- Gordon Intermediate Filaments Research Conference (2010): Tilton, NH. Title: Vimentin binds and stabilizes type I collagen mRNAs. (abstract published) 4- Annual Research Fair (2009): College of Medicine, Florida State University. Poster. Title: Role of vimentin intermediate filaments in synthesis of type I collagen.

PUBLICATIONS: 1- Challa AA*, Stefanovic B. (2011) A novel role of vimentin filaments: binding and stabilization of collagen mRNAs. Mol Cell Biol. 2011; 31, 18: 3773–3789. (Image chosen for journal cover)

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2- Challa AA*, Blackmon J, Stefanovic B. (2011) Vimentin knockout mice are protected from isoproterenol induced myocardial fibrosis. (in preparation for publication) 3- Challa AA*, Stefanovic B. (2011) Withaferin A, as an antifibrotic agent in a mouse model of isoproterenol-induced myocardial fibrosis. (submitted for publication) 4- Azariyas A*, Awala E, Seifu H. (2007) Lecture note on Principles and Concepts of Epidemiology for Health Science students. Mekelle University Press. 5- Azariyas A*, Awala E, Seifu H, Dawit S. (2007) Manual on Investigation and Management of Epidemic Prone Diseases in Ethiopia. Carter Center, USAID. 6- Azariyas Assefa*, Asfawossen Gebreyohannes. (2006) A case report of laboratory acquired visceral leishmaniasis in a pathologist working in Gondar University Hospital. Focus in GCMS, Vol 17, 21-24. 7- Amsalu Solomon, Assefa Azariyas*. (2005) Meningitis in children beyond the neonatal period, in Gondar University Hospital, Ethiopian Med J. Jul: 43(3) : 175- 80. ______//______

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