REGULATION of INFLAMMTORY ACTIVATION in ENDOTHELIAL CELLS by PIN1 DISSERTATION Presented in Partial Fulfillment of the Requireme

REGULATION OF INFLAMMTORY ACTIVATION

IN ENDOTHELIAL CELLS BY PIN1

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of

Philosophy in the Graduate School of The Ohio State University

TONGZHENG LIU, M.S

*****

The Ohio State University 2009

Dissertation Committee: Approved by Professor, Dale G. Hoyt, advisor

Professor, Lakhu M. Keshvara, co-advisor

Professor Anthony Young

Professor Terry S. Elton

Advisor Graduate Program in Pharmacy

TONGZHENG LIU

2009

ABSTRACT

Endothelial cells (EC) are a primary target for injury and cardiovascular risk

factors. Endothelial inflammatory activation and dysfunction has been critically

implicated in diverse cardiovascular diseases such as atherosclerosis, sepsis

and heart failures. Inflammatory cytokines and bacterial products such as

lipopolysaccharide (LPS) together with interferon-gamma (IFN-γ), can activate vascular endothelial cells, and induce expression of pro-inflammatory proteins, which essentially mediates inflammatory processes. Two prominent inflammatory processes are the production of nitric oxide (NO) by inducible nitric oxide synthase ( iNOS) and the formation of prostaglandins by cyclooxygenase-2

(COX-2), which may promote endothelial cell injury and play an important role in the pathogenesis of many cardiovascular diseases.

Protein phosphorylation regulates many cellular processes by causing changes in protein conformation. The prolyl isomerase PIN1 has been identified as a novel regulator of phosphorylation signaling that catalyses the conversion of specific phosphorylated motifs between the two completely distinct conformations in a subset of proteins. PIN1 regulates diverse cellular processes including

inflammation and immune responses. Various transcription factors and regulators have been identified as substrates for PIN1. For example, in response to cytokine signals, PIN1 acts on the phospho-p65 subunit of NF-κB and increases

its nuclear accumulation and NF-κB activity. It enhances AP-1 activity via binding

and isomerizing of both c-Jun and c-Fos. Such transcription factors are crucial for

expression of inflammatory genes, including iNOS and COX-2. In addition, PIN1

can also regulate mRNA and protein turnover. However, the role of PIN1 in

expression of these proteins in endothelial cells is unexplored.

Here, the effect of depletion of PIN1 on induction of iNOS by LPS and IFN-γ in murine aortic endothelial cells (MAEC) was determined. Knockdown (KD) of

PIN1 by 85% with a shRNA-lentiviral system enhanced the induction of NO and iNOS protein by LPS/IFN-γ. Similar results were also observed in PIN1 knock out

MAEC and cells transiently transfected with small interfering RNAs. There was

no effect on induction of iNOS mRNA, suggesting a post-transcriptional effect.

The enhanced levels of iNOS protein were functionally significant since LPS/IFN-

γ was cytotoxic to MAEC lacking PIN1 but not MAEC harboring an inactive

control construct, and because cytotoxicity was blocked by the NO synthase

inhibitor N ω-nitro-L-arginine methyl ester. Knockdown of PIN1 increased the

stability of iNOS protein in cycloheximide-treated cells. Furthermore, loss of iNOS

iii

was blocked by the calpain inhibition, but not by the selective proteasome inhibition. Immunoprecipitation and GST-PIN1 pull-down results indicated that

PIN1 can interact with iNOS. The results suggest that PIN1 associates with iNOS and can limit its induction by facilitating calpain-mediated degradation in MAEC

(chapter 2).

In addition, regulation of PIN1 on COX-2 degradation was investigated. Similar to iNOS, knockdown of PIN1 in endothelial cells enhanced the induction of COX-2 and its production of prostaglandin E2 in response to LPS/IFN-γ. The additional

increase in COX-2 protein due to PIN1 depletion was post-transcriptional, as

induction of COX-2 mRNA by LPS/IFN-γ was the same in cells containing or

lacking PIN1. Instead, the loss of COX-2 protein, after treatment with

cycloheximide to block protein synthesis, was reduced in cells lacking PIN1 in

comparison with control cells, indicating that degradation of the enzyme was

reduced. Calpain inhibition also enhanced the induction of COX-2 by LPS/IFN-γ, and slowed its loss in after treatment with cycloheximide. In contrast to iNOS, physical interaction between COX-2 and PIN1 was not detected, suggesting that

PIN1 may regulate COX-2 degradation by affecting calpain, rather than by an effect on COX-2 protein (chapter 3).

Since depletion of PIN1 reduced calpain-mediated turnover of iNOS and COX-2, we hypothesized that PIN1 regulated calpain activity in MAEC. Knockdown of

PIN1 reduced the basal and LPS/IFN-γ stimulated activity of calpain as

compared with MAEC harboring a control small hairpin RNA. However, the

depletion of PIN1 did not affect the levels of the large or small subunits of

ubiquitous, heterodimeric calpains. PIN1 did not interact with these proteins

either. Instead, deletion of PIN1 increased the basal level of calpastatin (CAST),

the endogenous protein inhibitor of heterodimeric calpains. The effect of PIN1 on

CAST was post-transcriptional, as CAST mRNA was not affected by PIN1

depletion. Loss of CAST, after inhibiting protein synthesis with cycloheximide,

was slower in KD than control MAEC, indicating that PIN1 regulated degradation

of CAST. Inhibition of calpain increased CAST protein levels in control MAEC,

whereas a pan-caspase inhibitor, did not, suggesting that calpain, not caspase,

contributed to the turnover of CAST in MAEC. Extracts from KD MAEC treated

with either vehicle or LPS/IFN-γ contained more inhibitory activity towards an exogenous calpain compared to control shRNA MAEC. Co-immunoprecipitation and glutathione-S-transferase-PIN1 fusion protein pull-down results indicated that CAST associated with PIN1. The PIN1 fusion protein also antagonized the ability of CAST to inhibit exogenous calpain in vitro. The results suggest that

PIN1 associated with CAST, either facilitating degradation of CAST or

decreasing its inhibitory activity towards calpain in MAEC. The results suggest that PIN1 normally maintains calpain activity and consequently limits the induction of calpain substrates, such as iNOS and COX-2 in endothelial cells.

The regulation of calpain system by PIN1 in MAEC indicates that PIN1 may play an essential role in various calpain related physiological and pathological processes (chapter 4).

The role of PIN1 in the regulation of calpain activity in vivo was investigated as well. Here, calpain activity and CAST inhibitory activity in various organs from

PIN1 wild type and knock out mice were measured. The regulation of the calpain system by PIN1 in vivo is more complicated and appeared tissue specific: lung and kidney from PIN1 knockout mice contained significantly lower calpain activity which may be due to the higher CAST inhibitory activity in them. However, higher calpain activity and lower CAST inhibitory activity were observed in brains from

PIN1 knockout mice. Liver and heart from PIN1 knock out mice contained significantly lower calpain activity although the CAST inhibitory activitiy was similar to wildtype mice, indicating that mechanisms other than CAST were involved in PIN1-regulated calpain activity in these organs. PIN1 knockout influenced normal cardiac function, causing a significant decrease of stroke volume compared with wildtype (chapter 5).

Overall, the results presented here suggest that PIN1 regulates calpain/CAST system in endothelial cells and various organs, which further modulates the turnover of inflammatory calpain substrates, such as iNOS and COX-2. Thus, the work presented in this thesis will direct our further investigation of the important role of PIN1 in MAEC endothelial activation and dysfunction during inflammatory conditions. These mechanistic insights of endothelial biology will provide new therapy for the calpain/CAST related diseases, such as cardiovascular disease.

vii

ACKNOWLEDGMENTS

I feel great pleasure to acknowledge the contribution of those people whose support and encouragement help me so much on my journey through graduate school here at the Division of Pharmacology, College of Pharmacy.

First, I would like to extend my deepest thanks to my advisors, Drs. Dale G. Hoyt and Lakhu Keshvara, whose consistent guidance, patience and encouragement in my research have been very helpful to my growth as a scientist. The work, time and guidance from my committee members, Drs. Terry Elton and Anthony

Young are greatly appreciated. I also thank all the members of the Division of

Pharmacology for their assistance and advice.

I would like to thank my parents Shilan Zhang and Yanting Liu, whose unconditional love, support and encouragement is always a source of motivate for me and help me through my years at graduate school. I would also like to thank my wife Jie Li for her steadfast love and support through all the roughness of school and my life.

viii

I thank many of my friends (Yongcheng, Vaibhav, Hong, Bin, Liyue, , Vivek,

Raeann, Rachel, Brandon and Zhaogang) in the Department of Pharmacology who have provided so many supports and a lot of fun during my time as a student at the Ohio State University.

VITA

July 18th, 1978 ………………… Born in Shandong China

1995 -1999 ……………………….B.S. in Medicinal Chemistry

Shandong University, Jinan China

1999 - 2002 ………………………M.S. in Pharmacology

China Pharmaceutical University, Nanjing China

2002 - 2003 ………………………Assistant researcher

Peking University, Beijing China

2003-present………………………Graduate Teaching Associate

The Ohio State University, Columbus, Ohio

PUBLICATIONS

RESEARCH PUBLICATIONS

Liu T, Huang Y, Likhotvorik RI, Keshvara L, Hoyt DG. Protein Never in Mitosis

Gene A Interacting-1, PIN1, Regulates Degradation of Inducible Nitric Oxide

Synthase in Edothelial Cells. Am J Physiol Cell Physiol. 2008 Jul 23.

Huang H, Liu T, Rose JL, Stevens RL, Hoyt DG. Sensitivity of mice to

lipopolysaccharide is increased by a high saturated fat and cholesterol diet. J

Inflamm (Lond). 2007 Nov 12;4:22

Huang Y, Shah V, Liu T , Keshvara L. Signaling through Disabled 1 requires

phosphoinositide binding. Biochem Biophys Res Commun. 2005 Jun 17; 331(4):

1460-8.

MEETING ABSTRACTS

Liu T , Huang Y, Likhotvorik RI, Keshvara L, Hoyt DG: Post-transcriptional

regulation of inducible nitric oxide synthase (iNOS) by Protein Interacting with

Never in Mitosis Gene A-1 (PIN1) in murine aortic endothelial cells. FASEB J. 22:

964.34, 2008

Liu T , Huang Y, Likhotvorik RI, Keshvara L, Hoyt DG: Regulation of calpain and

CAST by Protein Interacting with Never in Mitosis Gene A-1 (PIN1) in murine

aortic endothelial cells. FASEB J. 22: 964.35, 2008

Liu T and Hoyt DG: Post-transcriptional regulation of CAST by Protein

Interacting with Never in Mitosis Gene A-1 (PIN1). FASEB J. 22: 1133.4, 2008

Schneider RA, Liu T , Huang Y, Keshvara L, Hoyt DG: Depletion of Protein

Interacting with Never in Mitosis Gene A-1 (PIN1) decreases mammary

adenocarcinoma cell growth in culture, but increases VEGF and tumor growth in

mice. FASEB J. 22: 321.8, 2008

Liu T , Hoyt DG: Hypercholesterolemic Diet Increases Angiotensinogen and

Angiotensin Receptor Type 1a mRNAs in Mouse Lung. Am J Resp Critical Care

Med 175: A912, 2007

Liu T , Hoyt DG: Basic fibroblast growth factor increases the effects of endotoxin

and interferon gamma on iNOS in mouse aortic endothelial cells. FASEB J. 21(5):

A525, 2007

Liu T , Hoyt DG: Src Family Kinases mediate inhibition of interleukin 6 signaling

by fibroblast growth factor-2 in mouse aortic endothelial cells FASEB J. 20(4):

A116, 2006

xii

Huang H, Liu T , Bauer JA, Hoyt, DG: Hypercholesterolemic diet enhances pulmonary inflammation caused by endotoxin in mice Proc. Amer. Thoracic Soc.

3: A20, 2006

FIELDS OF STUDY

Major Field: Pharmacy

xiii

TABLE OF CONTENTS

PAGE

Abstract…………………………………………………………………………… ii

Acknowledgements……………………………………………………………... viii

Vita……………………………………………………………………………….. x

List of Figures……………………………………………………………………. xv

List of Tables……………………………………………………………………. xviii

Abbreviations…………………………………………………………………….. xix

CHAPTERS

1. Introduction……………………………………………………………………….1

2. PIN1 regulates degradation of iNOS in endothelial cells……………………57

3. PIN1 regulates degradation of COX2 in endothelial cells …………..………100

4. PIN1 regulates calpain/CAST system in endothelial cells …….………...... 140

5. PIN1 regulates calpain/CAST system in vivo...…………….….………...... 171

6. Conclusions and Future Directions…………………………………………….191

Bibliography………………………………………………………………………… 202

xiv

LIST OF FIGURES

Figure Page

1.1 Conformational switch of Ser/Thr-Pro and p- Ser/Thr-Pro motifs………….. 5

1.2 PIN1 regulates multiple pro-proliferative and pro-apoptotic pathways……..11

1.3 Role of STAT1 on iNOS induction in ECs …………………………………….30

1.4 PIN1 interacts with STAT1………………..……………………………………..32

1.5 Domain structure of µ- and m-calpain …..……………………………………..50

1.6 Domain structure of CAST ……………………………………………………...54

1.7 CAST specifically binds and inhibits m-calpain..……………………………...55

2.1 Suppression of PIN1 in MAEC………………………………………………….71

2.2 Effect of PIN1 knockdown or genetic deletion on iNOS……………………...74

2.3 Effect of PIN1 knockdown on iNOS and VCAM-1 mRNA and NF-κB………77

2.4 Effect of PIN1 knockdown on NO accumulation and cytotoxicity……………81

2.5 Effect of PIN1 knockdown on iNOS protein stability …..……………………..85

2.6 Effect of calpain inhibition on iNOS…………………………………………….87

2.7 Effect of proteasome inhibition on iNOS……………………………………….91

2.8 PIN1 interacts with iNOS………………………………………………………...93

3.1 Effect of PIN1 knockdown on COX-2 protein and activity………………….110

3.2 Effect of PIN1 knockdown on COX-2 mRNA………………………………..114

3.3 Effect of PIN1 knockdown on COX-2 stability……………………………….116

3.4 Effect of calpain inhibition on COX-2 protein………………………………..118

3.5 Degradation of COX-2 by purified porcine µ-calpain in vitro……………….123

3.6 Cathepsin does not contribute to PIN1-regulated COX-2 degradation …..125

3.7. PIN1 does not interact with COX-2 in MAEC………………………………..127

3.8 Effect of proteasome inhibition on COX-2 mRNA and protein……………..129

3.9 Effect of PIN1 knockdown, calpain and proteasome inhibition on COX-2 fragments……………………………………………………………………………...134

4.1 Effect of PIN1 knockdown on calpain activity………………………………..149

4.2 Effect of PIN1 knockdown calpain 1, 2 and 4 levels…………………..….…151

4.3 Effect of PIN1 knockdown on CAST protein and mRNA…………………...154

4.4 Effect of PIN1 knockdown on CAST protein stability ……………………….158

4.5 Effect of PIN1 knockdown on CAST inhibitory activity……………………...162

4.6 Purified GST-PIN1 decreases CAST inhibitory activity……………………..165

5.1 Effect of PIN1 knockout on calpain and CAST inhibitory activity in lung and kidney…………………………………………………………………..……………..177

5.2 Effect of PIN1 knockout on calpain and CAST inhibitory activity in brain ..180

xvi

5.3 Effect of PIN1 knockout on calpain and CAST inhibitory activity in liver and heart ………………………………………………………………………………..…183

5.4 Effect of PIN1 knockout on cardiac function ………………………………..185

6.1 PIN1 regulates calpain mediated proteolysis………………………………..195

xvii

LIST OF TABLES

Table Page

5.1 Effect of PIN1 on body weight and organ weight……………………………176

xviii

ABBREVIATIONS

Ser/Thr-Pro: serine or threonine residues preceding proline

AD: Alzheimer’s diseases

CDKs: cyclin-dependent protein kinases

ERKs: extracellular signal-regulated kinases

SAPKs/JNKs: stress-activated protein kinases/c-Jun-N-terminal kinases

GSK3: glycogen synthase kinase-3

PPIases: peptidyl prolyl cis-trans isomerases

PIN1: protein interacting with NIMA (never in mitosis A)-1

GM-CSF: granulocyte macrophage colony stimulating factor

SOCS1: suppressor of cytokine signaling 1

MCL1: myeloid cell leukaemia sequence-1

APP: amyloid precursor protein

STAT1: signal transducer and activator of transcription-1

GAS: IFN-γ activation sites

NF-κB: nuclear factor-κB

IRF3: interferon-regulatory factor 3

AP-1: activating protein-1

ARE: adenine-uridine–rich elements

xix

AUF1: AU-rich element-binding protein

APC: adenomatous polyposis coli protein complex

IGF-1: insulin-like growth factor 1

CAST: calpastatin

MAEC: murine aortic endothelial cells

IFN-γ: interferon gamma

LPS: lipopolysaccharide iNOS: inducible nitric oxide synthase

COX-2: cyclooxygenase-2

PG: prostaglandins shRNA: short hairpin RNA

KD: knock down

CHX: cycloheximide

GST: Glutathione S-transferase

NBT: Nitroblue Tetrazolium

SDS-PAGE: Sodium dodecyl sulfate polyacrylamide gel electrophoresis

CHAPTER 1

INTRODUCTION

A novel signaling regulatory mechanism:

PIN1-catalyzed prolyl isomerization after phosphorylation

Phosphorylation of proteins is a common reversible post-translational

modification, which is controlled by kinases and phosphatases. Protein kinases

add a phosphate group to Serine (Ser), Threonine (Thr) and Tyrosine (Tyr)

residues in a protein and phosphotases remove it from proteins. Addition or

removal of phosphate group can cause a dramatic conformational change in the

structure of the protein and affect the properties of target proteins, which make it

as one of the most important signaling mechanisms to regulate many cellular

processes (1). For instance, phosphorylation on Tyr residues in certain proteins may function as specific binding sites for SH2 domain containing protein and allows the formation of signaling complex. Phosphorylation on Ser or Thr residues may also have similar functions (2-5). In addition, phosphorylation on some enzymes or receptors alters conformation, allowing cells to either turn on/off enzymes’ catalytic activity or increase/decreases receptor signaling intensity or duration (6, 7 ).

Protein kinases may phosphorylate their substrates in different consensus sequences. Some Ser/Thr kinases are called proline-directed kinases since they phosphoryalte their substrate motifs (Ser/Thr-Pro motifs) in a proline directed manner. Cyclin-dependent protein kinases (CDKs), extracellular signal-regulated kinases (ERKs), stress-activated protein kinases/c-Jun-N-terminal kinases

(SAPKs/JNKs), glycogen synthase kinase-3 (GSK3) and others are known as to be members of the proline-directed kinase family. Proline-directed phosphorylation is important for regulation of many cellular events (8) and have

been implicated in the pathogenesis of many human diseases, such as cancer

and Alzheimer’s disease (AD) (9-13 ).

Although phosphorylation is believed to regulate protein function via changing protein conformation, it remains largely unknown how conformational changes that occurr after phosphorylation on Ser/Thr-Pro motifs alter protein’s function.

It is known that either trans or cis conformation can be adopted in any peptidyl bond. The trans conformation is much more energetically favored than cis and is more dominant in peptidyl bonds under most conditions. However, this is not the case in proline which contains a five membered ring and makes the free energy difference between cis and trans much smaller. Thus, proline adopts completely

distinct cis and trans conformations and is unique in providing an intrinsic backbone switch (11 ) (See Fig. 1.1). The spontaneous interconversion rate is very low without the assistance of a catalyst. Peptidyl prolyl cis -trans isomerases

(PPIases) can dramatically speed up the conversion rates (14 ). So far, PPIases can be divided into several families: cyclophilins, FK506-binding protein and parvulins. These PPIases are structurally unrelated, have their own conserved amino acid sequence and substrate specificity. Cyclophilins and FK506-binding protein are cellular targets for immunosuppressive drugs cyclosporins and

FK506/rapamycin respectively. Their actions invovles facilitating protein folding or promoting in protein complex formation, while the PPIase activity is not necessary for immunosuppression (8). The biological significance of PPIase activity as an important regulatory mechanism was not known until the discovery of Protein Interacting with NIMA (never in mitosis A)-1 ( PIN1 ).

Human PIN1 was originally identified by its ability to interact with NIMA, a mitotic kinase whose phosphorylation on multiple Ser/Thr-Pro motifs is critical for mitosis in Aspergillus nidulans (15 ). Although PIN1 belongs to parvulin subfamily, its substrate specificity is completely distinct from other parvulin members. PIN1 is the only identified mammalian prolyl isomerase that specifically acts on phosphorylated Ser/Thr-Pro peptide bonds and is highly conserved from yeast to

human. PIN1 contains a C-terminal PPIase domain and an N-terminal domain, characterized by two tryptophan residues separated by 22 amino acids that can bind to phosphorylated serine- or threonine-proline sequences in substrate proteins, known as a WW domain. PPIase domain and WW domain are coordinated to specifically act on phospho-substrates. The WW domain is responsible for binding to phospho-Ser/Thr-Pro–containing proteins. PPIase domain is sufficient to catalyze isomerization at phosphorylated Ser/Thr-Pro motifs and induce isomerization, which triggers various functional outcomes and regulates a wide range of cellular processes.

It is well-known that phosphorylation on Ser/Thr preceding Pro can dramatically reduce the isomerization rate between cis /trans conformation and make the

peptide bond resistant to isomerization catalyzed by other known PPIase.

Furthermore, phosphatases that modify pSer/Thr-Pro motifs and certain proteins

that interact with these motifs are conformation-specific and only act on the trans

conformation. Thus, PIN1 catalyzed isomerization of pSer/Thr-Pro motifs is

extremely important in phosphorylation signaling. Restoration of the equilibrium

between cis and trans would require minutes without a catalyst, while PIN1 can

dramatically accelerate cis to trans or trans to cis isomerization rate and maintain

equilibrium of cis and trans conformations on the millisecond timescale. Such an

acceleration in equilibrium from minutes to millisecond make it more relevant for the regulation of dynamic biological processes (16 ).

S P PPIase S P

cis trans

Proline-directed Protein phosphatase kinase 2A CTD Phosphatase

P PIN1 pS pS P cis trans

Function B Function A

Fig 1.1 Conformational switch of Ser/Thr-Pro and p- Ser/Thr-Pro motifs catalyzed by PPIase.

Accumulating evidence indicates that PIN1-catalyzed isomerization after proline directed phosphorylation regulates a wide variety of key proteins, suggesting that the phosphorylation-specific, PIN1-catalyzed conformational regulation may be a common signaling mechanism. Isomerization of pSer/pThr-Pro motifs by PIN1 has been shown to play an important role in regulating diverse cellular processes, including immune responses, cellular stress responses, cell growth and proliferation and neuronal function. PIN1 is also critically involved in several human diseases including cancer, AD and microbial infections. Some of PIN1 functions in these cellular processes and related diseases are reviewed next.

PIN1 and immune responses

PIN1 may play an important role in immune by acting on Nuclear factor-κB (NF-

κB) and interferon-regulatory factor 3 (IRF3) which are essential mediators in viral infection and other immune responses.

As a central mediator in the immune responses, NF-κB is important in the development of innate immune cells as well as B and T cells, which directly mediate immune responses (17 ). NF-κB also regulates expression of mediators of inflammation includes cytokines (for example, TNF α and IL-1β) and adhesion molecules (for example, ICAM-1). When exposed to cytokine signals, p65

subunit of NF-κB is phosphorylated in its Thr254-Pro motif and interacts with

PIN1, which inhibits p65 interaction with endogenous inhibitory protein I κBα and increases its nuclear accumulation and activity (18 ). Furthermore, the association between PIN1 and p65 also inhibits its degradation by suppressor of cytokine signaling 1 (SOCS1). Deletion of PIN1 renders cells resistant to NF-κB activation by cytokine signals (18 ).

IRF3 has been identified as a PIN1 substrate recently (19 ). IRF3 controls the primary induction of interferon-β (IFN-β), which in turn controls expression of hundreds of genes and involved in viral infection. PIN1 interacts with phospho-

IRF3 at pSer339–Pro340, facilitates IRF3 ubiquitination – proteasome mediated degradation and negatively regulates its activation. Therefore, knockdown or genetic deletion of PIN1 results in enhanced IRF-3-dependent production of interferon-β and reduces virus replication. These results also indicate the important role of PIN1 in the establishment of innate immunity against viral infection.

Recent studies have shown that PIN1 is important in the development of asthma

(20-22 ). The activation of eosinophils in the lung triggers the secretion of granulocyte-macrophage colony-stimulating factor (GM-CSF), an essential survival factor for activated eosinophils. PIN1 isomerase activity is necessary for

GM-CSF mRNA stabilization and its secretion, which in turn increases the survival of activated eosinophils. The action of PIN1 on GM-CSF mRNA stability is through its association with the AU-rich element-binding protein, AUF1, which regulates the decay rate of GM-CSF mRNA. Similar results were also observed in human PBMC and CD4+ lymphocytes. PIN1 activity has been demonstrated to be enhanced in eosinophils obtained from the airways of patients with asthma, suggesting that PIN1 may be a potentially therapeutic target IN allergic asthma.

PIN1 and cellular stress

In response to various stresses, many proline-directed kinases such as JNKs and p38 are activated and phosphorylate their substrate motifs (p-Ser/Thr-Pro), which are putative PIN1 binding sequences. Thus it is important to investigate whether PIN1 is involved in cellular stress. Several key proteins in cellular stress responses, such as p53 and p73 have been identified as PIN1 substrates, and

PIN1-dependent isomerization regulates their function in cellular stress.

Genotoxic stress leads to p53 stabilization and accumulation, which is regulated by proline-directed phosphorylation and MDM-2-ediated degradation. PIN1 regulates p53 function by several different mechanisms (23, 24 ). Firstly, PIN1

acts on phosphorylated p53 and inhibits its binding to MDM2, therefore increaseing p53 protein stability in response to DNA damage. Secondly, PIN1 may also be required for efficient binding of p53 on target promoters, enhancing the DNA-binding activity and transcriptional activity of p53. Thirdly, the association with PIN1 can stimulate binding of the p300 acetyltransferase to p53 and consequently increase p53 acetylation. Fourthly, PIN1 also mediates p53's dissociation from the apoptosis inhibitor, iASPP, promoting cell death (25 ). The stability and activity of p73, a homologue to p53, can be up-regulated by PIN1

(26, 27 ). These results suggest that PIN1 is involved in the response to different cellular stresses by regulating substrate protein function or their interaction with other proteins. PIN1 itself can also be inactivated in oxidative stresses (28 ). Thus, it would be interesting to examine the significance of PIN1 in cellular stresses.

PIN1, growth-signaling pathways and oncogenesis

Proline-directed phosphorylation of proteins on certain Ser/Thr-Pro motifs is an essential signaling mechanism in cell proliferation and transformation in response to growth stimulation. By binding and isomerizing specific pSer/Thr-Pro motifs in many proteins central to tumorigenesis, including β-catenin, NF-κB, cyclinD1 and

p53, PIN1 regulates the duration and outcome of signaling after phosphorylation.

The following section will describe how PIN1 plays a key role in the regulation of the Wnt/ β-catenin, cytokine/ NF-κB and Neu-Raf-Ras-MAP kinase pathways

(See Fig. 1.2).

As a key transcription activator in the Wnt pathway, β-catenin is negatively regulated by the adenomatous polyposis coli protein (APC) complex. APC interacts with β-catenin, holds it in the cytoplasm and triggers its degradation through proteasome pathway. PIN1 specifically binds to pSer 246-Pro of β- catenin and prevents it from binding to APC, which stabilizes β-catenin and increases its translocation in to nucleus (29 ). In the nucleus, β-catenin and its cofactors T-cell transcription factor activates targeted gene expression.

Mutation of the APC or β-catenin for example in their binding sites can disrupt the binding of APC to β-catenin, resulting in subsequent degradation and prolonged activation of β-catenin. However, in many other types of cancer where APC or β- catenin mutations are not common, PIN1 is a key regulator of β-catenin and strongly correlates with its expression levels and nuclear accumulation. For example, overexpression of PIN1 increases β-catenin level, while knockdown or genetic deletion of PIN1 decreases β-catenin level in breast cancer tissues. Thus

PIN1- mediated isomerization is an important mechanism in enhancing β-catenin activity and creates a positive growth signal for tumor cells.

Stress signal Growth factor Wnt cytokine

Ras Raf Kinase? GSK-3β Kinase? MAPK P P P p53 P β-catenin NF-kB Unstable/inactive c-Jun

PIN1 PIN1

Stable/active p53 c-Jun β-catenin NF-kB P P P P

Nuclear translocation

Target

E2F cyclin D1

GSK-3β P cyclin D1 Cell cycle arrest apoptosis PIN1

cyclin D1 P

Target Rb

Cell proliferation and transformation

Fig. 1.2 PIN1 regulates multiple pro-proliferative and pro-apoptotic pathways.

Arrows indicate interaction or stimulation

symbols indicate inhibition

“p” indicates phosphorylation

Considering the importance of NF-κB in oncogenesis and the fact that PIN1 critically regulates NF-κB signaling in response to cytokines (18 ), it is necessary

to investigate the relationship between PIN1 overexpression and NF-κB activation in certain cancers. PIN1 interacts with the p65 subunit of NF-κB and inhibits its interaction with the endogenous inhibitory protein I κBα, which

increases NF-κB nuclear accumulation and activity. Furthermore, PIN1 also inhibits p65 degradation by SOCS1 (18 ). Therefore, PIN1 overexpression would

uncouple the inhibition of NF-κB by I κBα, which might explain why NF-κB signaling is overactivated even though I κBα is also upregulated in certain

cancers.

Another well-known example is the Neu-Raf-Ras-MAP kinase pathway (30 ).

Growth factor receptor Neu activates Raf and Ras in response to growth

stimulation. The MAP kinase cascades are activated as a result. MAP kinases

phosphorylate c-Jun and c-Fos, thereby enhancing AP-1 transcriptional activity toward its downstream targets such as cyclin D1, which positively promotes cell cycle progression (30 ). Furthermore, MAP kinase-mediated phosphorylation of

Raf inactivated Raf kinase, forming a negative feedback loop (31 ).

As an E2F downstream target gene, PIN1-catalyzed isomerization critically

regulates Neu-Raf-Ras-MAPK signaling pathway by acting on multiple targets.

First, PIN1 can directly act on c-Jun/c-Fos and increase AP-1 transcriptional

activity and downstream target gene expression, including cyclin D1 (30, 32 ).

Second, PIN1 directly interacts with cyclin D1 to enhance its stability and nuclear

localization. The transcription factor, E2F, downstream target of cyclin D1, may

be activated and increase PIN1 expression (33 ). Increased PIN1 expression in

turn may enhance growth signaling by increasing cyclin D1 transcription and

stabilization through a positive feedback loop (8). Third, the negative feedback

loop can be blocked by PIN1 via promoting dephosphorylation of Raf kinase,

prolonging the activation of Raf kinase (31 ).

PIN1 is overexpressed in many human cancers, such as malignancies of the breast and prostate (32 ). Ras signaling, β-catenin, and NF-kB play critical roles in oncogenesis. Oncogenic signals, such as Neu activation, induce PIN1

overexpression, which greatly promotes cell transformation by enhancing Ras siganling, β-catenin expression, and NF-kB signaling. In contrast, knockdown of

PIN1 induces apoptosis and inhibits tumorigenesis. PIN1 also enhances downstream signalling by amplifying positive feedback mechanisms (multiple targets: c-Jun, c-Fos, cyclin D1 and E2F) and suppressing negative feedback mechanisms (Raf), synergistically driving the cell in one direction, for instance, to growth promotion or transformation. Collectively, PIN1 plays an essential role in oncogenesis. PIN1 inhibition might be a potentially novel anticancer therapy target.

PIN1 in neuronal function and Alzheimer’s disease

Although it is expressed ubiquitously, PIN1 protein levels and activity vary significantly in various tissues. The PIN1 level in most neurons is relatively high and increases during neuronal differentiation (34, 35 ), suggesting that PIN1 may be invovled in neuronal function. Several key proteins in neuronal function such as Tau (36 ), amyloid precursor protein (APP) (16 ), and myeloid cell leukaemia sequence-1 (MCL1) (37 ) have been reported as PIN1 substrates. Tau is able to bind to microtubules and promote microtubule assembly, which is abolished by

the phosphorylation of tau. PIN1-catalysed isomerization can restore the biological function of phosphorylated tau by different mechanisms (36 ). PIN1 directly interacts with phosphorylated tau and restores the ability of tau to bind to microtubules and promote microtubule assembly. PIN1 can also facilitate dephosphorylation of phosphorylated tau by Pro-directed phosphatase PP2A, which is conformation-specific and effectively dephosphorylates only the trans pSer/Thr-Pro isomer. In the case of APP, PIN1 binds to the phosphorylated Thr

668-Pro motif in APP and regulates its non-amyloidogenic processing, therefore producing neurotrophic α-APPs and reducing neurotoxic amyloid-β (A β) peptides.

Amyloidogenic APP processing is favored in transgenic mice with PIN1 knockout alone or in combination with overexpression of mutant APP, which results in the accumulation of insoluble Abeta42, a major toxic species in brain, in an age- dependent manner. JNK3 induced MCL1 degradation and apoptosis after spinal cord injury. However, PIN1 binds and stabilizes MCL1, protecting oligodendrocytes from undergoing apoptosis after JNK3 activation (37 ).

The neuropathological hallmarks of Alzheimer's disease are neurofibrillary tangles composed of tau and neuritic plaques comprising Abeta derived from amyloid precursor protein (APP). PIN1-catalysed prolyl isomerization regulates both tangle (phosphorylation and function of Tau) and plaque (APP processing

and Abeta production) pathologies. Notably, PIN1 is depleted and/or inhibited by oxidation in Alzheimer's disease neurons. PIN1 expression is inversely correlated with predicted neuronal vulnerability and neurofibrillary degeneration in

Alzheimer's disease. PIN1 knockout in mice causes motor and behavioral deficits, tau hyperphosphorylation, tau filament formation and other progressive age- dependent neuropathies. Thus, PIN1 is pivotal in protecting against age- dependent neurodegeneration.

Regulation of PIN1

While PIN1 governs various aspects of cell signaling, PIN1 itself is tightly regulated at multiple levels under physiological conditions.

1. Protein level

Although present in all cells, PIN1 expression is strongly correlated with the proliferative activity of normal tissues. PIN1 is readily detectable in actively dividing cells, but is low in most terminally differentiated tissues, with the exception of neurons (38 ). Interestingly, PIN1 itself is also a target of several oncogenic pathways and its expression is significantly upregulated in

transformed cell lines and some human cancer tissues (32 ). For example, PIN1 is a target gene for the E2F family of transcription factors, which can be activated by Neu or Ras activation (33 ). Insulin-like growth factor 1 (IGF-1) induces PIN1 protein expression in human breast cancer cells, which is mediated via the phosphatidylinositol 3-kinase as well as the MAP kinase pathways (39 ).

Expression of PIN1 was decreased in response to ischemia-reperfusion in liver

(40 ).

2. Post-translational modification

Several post-translational modifications including phosphorylation, ubiquitination and oxidation, have been demonstrated to regulate PIN1 enzymatic activity or stability. Two phosphorylation sites have been identified in PIN1 (S16 and S65).

S16 is located in PIN1 WW domain and its phosphorylation abolishes the binding ability of PIN1 to its substrate proteins (41 ). The S16A mutant of PIN1 binds to substrates constitutively, while S16D mutant of PIN1 is unable to bind to substrates. Given that PIN1 interacts with a large number of substrates, PIN1 appears to be predominantly phosphorylated on S16 in normal breast tissues and might prevent PIN1 from acting on unintended substrates. However, overexpressed PIN1 is mainly non-phosphorylated, is hyperactive in certain tumor tissues (32 ). PIN1 can be ubiqutinated and undergo proteasome-mediated

degradation. In addition, PIN1 has been shown to be phosphorylated on Ser65 by Polo-like kinase-1, which does not affect its isomerase activity, but rather contributes to its protein degradation (42 ). Inhibition of Polo-like kinase-1 activity enhances the ubiquitination of PIN1 and its degradation in human cancer cells.

Importantly, PIN1 was found to be oxidatively modified and to have reduced activity in the hippocampus in mild cognitive impairment (MCI) and in the early stages of AD (28 ). However, oxidative modification sites on PIN1 and their effects on PIN1 activity are not known.

3. Subcellular localization

PIN1, a small 18-kDa protein, is able to diffuse easily across nuclear membrane, and its subcellular localization and function are dependent on the phospho- substrate availability. Thus, the localization of target proteins and related proline- directed kinases determine the distribution of PIN1 in either the nucleus or cytoplasm (43 ). For example, PIN1 is redistributed to cytoplasmic tangles by acting on hyperphosphorylated tau in AD neurons (34 ).

Summary

PIN1-mediated, phosphorylation-dependent prolyl isomerization has been identified as a novel regulatory mechanism. Such conformational regulation can have a profound impact on many key proteins in diverse cellular processes.

Importantly, PIN1 levels and function are tightly regulated under physiological conditions and deregulation has a pivotal role in a growing number of pathological conditions. Thus, it is a potential new diagnostic marker of cancers or AD. It would be beneficial to develop drugs that target PIN1 for treating certain human diseases, such as cancer and immune disorders.

Endothelial inflammatory activation

My project is concerned with inflammatory activation of endothelial cells (EC).

The thin monolayer of endothelial cells lines the interior surface of blood vessels in the entire circulatory system, which forms an interface between blood flow in the lumen and the underlying tissues. Rather than being a simple barrier between intravascular and interstitial compartments, the endothelium is a highly specialized and active tissue. Under resting conditions, the vascular endothelium is responsible for maintaining vascular homeostasis by modulating vascular tone and permeability and inhibiting platelet aggregation, leukocyte activation and adhesion (44 ). The endothelium also contributes to the local balance in pro- inflammatory and anti-inflammatory mediators and prevents the proliferation and migration of the underlying smooth muscle cells (44 ).

The role of endothelial cells is not confined to the regulation of the above functions. Because of its strategic anatomic position, EC are not only a primary target but also a mediator of injury. Multiple mechanisms, such as inflammation, oxidative stress, lipid abnormalities and genetic predisposition, can allow EC to participate in the inflammatory response, which is recognized as EC activation.

EC activation has been critically implicated in such as atherosclerosis, sepsis, vascular diabetic complications, and other cardiovascular diseases (45-48 ).

The growing understanding of intracellular signaling has led to the discovery that a common intracellular mechanism involving activation of transcription factors, including NF-κB, contributes to the diverse effects of EC activation (45 ). A wide range of agents such as certain bacteria and viruses, interleukin-1, tumour necrosis factor, oxidative stress, oxidized low density lipoproteins (49 ), and antiendothelial cell antibodies (50 ) are known to activate NF-κB and other important transcription factors. EC activation is not all or none but a graded response. Activation may occur locally, as in transplant rejection (51 ), or systemically, as in septic shock and the systemic inflammatory response.

Loss of vascular integrity, a shift to a pro-thrombotic state and expression of pro- inflammatory mediators are the core changes of endothelial cell activation. Loss of vascular integrity can expose the subendothelium and enhance permeability of the endothelium. The changes in EC integrity can be graded as well: local increases of permeability or major endothelial cell contraction, exposing large areas of subendothelium. The anti-thrombotic state in resting EC can be shifted

to the prothrombotic state with loss of the surface anticoagulant molecules thrombomodulin and heparan sulphate. Activation of EC also leads to the expression of many proinflammatory mediators. For example, induction of adhesion molecules such as E-selectin and ICAM-1 is critical to recruit macrophages and T lymphocytes to endothelium, which then infiltrate into the vessel wall, initiate cytokine synthesis, and modulate further inflammatory processes (52, 53 ). In addition, EC may also contribute to the production of inflammatory cytokines, chemoattractants and tissue factors, such as IL-6 and monocyte chemoattractant protein 1. In addition to secretion of cytokines, EC are also an important target of such cytokines. For example, inflammatory cytokines and bacterial products such as lipopolysaccharide (LPS) together with T- lymphocytes cytokine interferon-gamma (IFN-γ) can activate endothelial cells and induce expression of pro-inflammatory proteins, which essentially mediates inflammatory processes. Changes in expression of many proteins during EC activation of EC are often controlled by changes in transcription. Several transcription factors, such as NF-kB, AP-1 and β-catenin, are known substrates

of PIN1. However, the role of PIN1 in EC has not been investigated. Two

prominent in inflammatory processes are the production of nitric oxide (NO) by

inducible nitric oxide synthase ( iNOS) and the formation of prostaglandins by cyclooxygenase-2 ( COX-2). iNOS and COX-2 are induced by many inflammatory

stimuli, and may promote endothelial cell injury, which is believed to play a role in the pathogenesis of many cardiovascular diseases. Thus, the role of PIN1 in the induction of iNOS and COX-2 in EC is investigated in my study.

Nitric Oxide Biology

Diverse physiological and pathophysiological functions of EC are mediated by

NO (54 ). In mammalian cells, NO can be produced from from l-arginine and

molecular oxygen by 3 isoforms of NOS, two constitutive isoforms, neuronal

(nNOS) and endothelial (eNOS), and one inducible isoform (iNOS). All three

isoenzymes are homo-dimers and share similar cofactors FAD, FMN, BH4, and

heme. nNOS and eNOS are constitutively expressed in neurons and EC,

respectively. They are low output, Ca 2+ -dependent enzymes producing NO in a

pulsative manner. In contrast, iNOS is a high output, Ca 2+ -independent enzyme

whose expression can be induced in a wide range of cells by cytokines and other

agents. After induction, iNOS produces NO continuously until the enzyme is

degraded.

NO is a short-lived free radical and a very small compound that diffuses freely within cells from its site of formation to its site of action. NO regulates blood vessel tone in vascular system. NO derived from eNOS is a powerful vasodilator, which diffuses through cellular membranes from EC to smooth muscle cells

(SMC) and activates guanylate cyclase leading to increased production of cyclic guanosine monophosphate (cGMP), which in turn modulates a wide variety of effectors such as ion channels, phosphodiesterases and protein kinases to decrease intracellular calcium and ultimately relax vascular smooth muscle (55,

56 ). Physiological concentrations of NO derived from eNOS also exert various

vaso-protective effects, such as inhibition of platelet aggregation, suppression of

integrin-mediated adhesion of leukocytes or monocytes to the endothelial

surfaces, inhibition of proliferation and migration of SMC, and attenuation of

proinflammatory-induced gene expression that control inflammation and oxidative

injuries (57-61 ). Deregulation of NO derived from eNOS accompanies the

development of atherosclerosis, vasospasm and thrombosis, and may contribute

to some forms of hypertension and diabetic vascular diseases (54 ).

Inducible Nitric Oxide Synthesis (iNOS)

The role of nitric oxide (NO) generated by the inducible isoform of nitric oxide synthase (iNOS) is very complex. As an important host defense effector, NO derived from iNOS has been described to have beneficial anti-viral, anti-parasite, anti-microbe and anti-tumor effects (62-64 ). On the other hand, large amounts of

NO derived from aberrant expression of iNOS is a marker of inflammation and a mediator of cell injury and activation. For example, induced at the wrong place or at the wrong time, iNOS has dramatic toxic effects on host cells and seems to be involved in the pathophysiology of different human diseases, including septic shock, myocardial ischemia-reperfusion injury, asthma, arthritis, tumor development, transplant rejection and neurodegenerative diseases (54, 64-67 ).

Inhibition of iNOS may be beneficial for the treatment of inflammatory disease

(64, 68, 69 ). Since modulation of iNOS expression is the most important component of iNOS regulation, it is essential to understand how iNOS is regulated normally and deregulated in pathological processes.

iNOS is low or undetectable in resting cell and can be induced by inflammatory cytokines, bacterial products or infection in many cell types, including EC (66 ).

The pathways regulating iNOS expression is complex and varies in different cells or species. For example, either LPS or IFN-γ is sufficient to induce iNOS

expression in macrophages, while only the combination of them can induce iNOS

in EC. iNOS induction can be regulated at both transcriptional and post-

transcriptional levels, which will be reviewed next.

Transcription

Intense studies have been focused on transcription of iNOS. In general,

activation of the transcription factors NF-kB and signal transducer and activator

of transcription STAT-1α and thereby activation of the iNOS promoter seems to

be an essential step in the regulation of iNOS transcription in most cells (63 ).

1. Signal transducer and activator of transcription-1α (STAT-1α)

As a well-known pro-inflammatory cytokine critically involved in the immune

system and in inflammatory disorders, IFN-γ mediates biological responses

mainly via activation of Janus kinase (JAK)-STAT pathway. IFN-γ binding causes

its receptor to dimerize and activates the receptor-associated tyrosine kinase

JAKs, which in turn tyrosine phosphorylate the receptor, and recruit cytoplasmic

STAT1 via SH2-phosphotyrosyl interactions. STAT1 is then phosphorylated on tyrosine 701 by the JAKs, which triggers STAT1 dimerization and translocation into the nucleus. Then, STAT1 dimers bind to IFN-γ activation sites (GAS)

located in the promoters of target genes, inducing their transcription (70, 71 ).

The promoter region of the iNOS gene has been characterized in different

species, including human, rat, and mouse (72-74 ). Sequence analysis of

promoter revealed that several GAS are contained in mammalian iNOS promoter

(75 ). Inhibition of JAK2 by pharmacological inhibitor or expression of dominant

negative mutant JAK2 or STAT1 reduced STAT1 α DNA binding activity and

iNOS expression (76-79 ), suggesting the essential role of STAT-1α to directly

bind the iNOS promoter and induce its expression in many cells including EC

(80-82 ).

Activated STAT-1α also binds to the GAS site of the interferon regulatory factor-

1 (IRF-1) promoter and induces its expression (83, 84 ). Effects of IRF-1 on iNOS

mRNA induction depend on cell types. IRF-1 knock-out macrophages

synthesized undetectable iNOS mRNA in response to stimulation, suggesting

that IRF-1 is essential for iNOS induction (85 ). The binding of IRF-1 to the iNOS promoter is required for the synergistic contribution of IFN-γ to transcription of

iNOS, and mutation of IRF-1 binding sites blocked such synergy (86, 87 ).

However, in certain cell types, such as hepatocytes, genetic deletion of IRF-1 does not influence iNOS induction in response to LPS/IFN-γ, suggesting that the

role of IRF-1 in regulation of iNOS expression is cell or tissue specific (88, 89 ).

Phosphorylation of tyrosine701 regulates the dimerization of STAT1, which is an

essential prerequisite for its transcription activity. In addition, phosphorylation

within a PMS 727 P motif by MAPK, such as ERK1/2 and p38, can also regulate

STAT1 activity, which may be required for its full transcriptional function in most of cases (80, 90, 91 ). Mice expressing a S 727 A mutation were more resistant to

the LPS-induced septic shock, suggesting that this serine phosphorylation in

STAT1 promotes inflammatory responses. Expression of IFN-γ-induced genes

was strongly reduced in macrophages expressing STAT1 S 727 A mutant.

Moreover, a loss of serine phosphorylation results in a decrease in association

with the coactivator p300 acetyltransferase and histone acetylation at the

interferon-responsive promoter. However, it is not clear how S 727 phosphorylation regulates STAT1 transcription activity and protein-protein association.

Our results showed that STAT1 is crucial for iNOS induction since no detectable iNOS were induced in response to LPS/IFN-γ in STAT1 knock-out EC (Fig. 1.3).

Transfection of wt STAT1 in the EC significantly induced iNOS expression.

Y701A mutant had no influence on iNOS induction when compared to GF-2 vector plasmid, wheras S 727 A mutant partially induced iNOS. These results

confirmed that Y701 is an essential prerequisite while S727 regulates its full

transcription activity. Huang et al showed that STAT1 mediated the synergistic

effect of LPS on IFN-γ in the induction of iNOS in EC. This was associated with p38 MAPK dependent phosphorylation of PMS 727 P. It is not known whether PIN1

affects STAT1 α. Thus, I used the methods described in chapter 2 to investigate

the potential for PIN1 to associate with STAT1. As shown in Fig. 1.4, STAT1 co-

immunoprecipitated with PIN1 in LPS/IFN-γ treated MAEC, where phosphorylation of S 727 is known to occur. A glutathione S-transferase PIN1 fusion protein also pulled down STAT1 from extracts of LPS/IFN-γ treated cells.

Thus, it is essential to investigate whether the association of PIN1 regulates

STAT1 function.

• GF2 STAT1 Y701F S727A

iNOS

α-Tubulin

3 + *

2 -Tubulin α α α α

1 iNOS/ Signal Intensity Ratio Signal 0 GF-2 STAT1 Y701F S727A

Fig 1.3 Role of STAT1 on iNOS induction in ECs. A: 2 µg of vector plasmid (GF-

2), wt STAT1 or its mutants Y701A and S727A are transfected in STAT1

knockout murine aortic endothelial cells (MAEC). Cells were treated with LPS and IFN-γ for 24 h. Representative Western blots of iNOS and α-tubulin are

shown. B. densitometric analysis of signal intensities of images was used to

measured iNOS expression. Bars represent means + SE ratio of iNOS/ α-tubulin

from three cultures of each group. +p < 0.05 for comparison with GF-2, *p < 0.05

for comparison between STAT1 and S727A.

A. Vehicle L/I

PIN1 Ab: - + - +

STAT1

PIN1

B. GST GST-PIN1

STAT1

GST-PIN1

GST

Fig 1.4 PIN1 interacts with STAT1. A: MAEC were treated with vehicle or

LPS/IFN-γ for 24 h. Cell extracts were immunoprecipitated without (None) or with anti-PIN1 antibody (PIN1) and analyzed by Western blot analysis with anti-

STAT1 antibody. B: cell extracts after LPS/IFN-γ treatment for 24h were

incubated with GST or GST-wild-type PIN1 fusion proteins, and glutathione

sepharose, and analyzed by Western blotting with anti-STAT1 and anti-GST

antibodies. Blots are representative of 3 independent cultures.

2. NF-κB

The family of NF κB transcription factors is a highly conserved protein complex from Drosophila to human. There are two structural classes of NF-κB proteins:

class I (p50 and p52) and class II (RelA, RelB, and c-Rel). Both classes of

proteins contain a highly conserved Rel homology domain (RH) in their N-termini,

which serves as a DNA-binding domain and also as a dimerization interface to

other NF-κB transcription factors, and the inhibitory IκB protein. Class II but not

class I NF-κB proteins have a transactivation domain in their C-termini (92 ).

NF-κB is found in almost all cell types and plays a pivotal role in regulating the immune responses (17 ). Deregulation of NF-κB has been implicated in many human diseases including to septic shock, viral infection, cancer inflammatory and autoimmune diseases (93 ).

NF-κB can be activated by exposure of cells to LPS or inflammatory cytokines such as TNF or IL-1, viral infection, UV irradiation, and by other physiological and nonphysiological stimuli (94 ). NF-κB proteins can form homodimers or heterodimers in vivo. The activity of NF-κB is tightly regulated by interaction with

inhibitory protein I κB. Several different I κB proteins have been identified and each has different affinities for individual NF-κB dimers. In resting cells, NF-κB

dimers are sequestered in the cytoplasm by interacting with I κB. In response to

extracellular signals such as LPS, the I κB kinase (IKK) is activated, causes

phosphorylation, inducing degradation of I κB, allowing the dissociation of I κB-NF-

κB complex, resulting in NF-κB nuclear translocation. The activated NF-κB binds to κB sites in promoters and activate transcription of genes encoding proteins

involved inmmune or inflammatory responses and in cell growth control.

NF-κB binding sites in the iNOS promoter region have been demonstrated in human (95 ) and rodents (96, 97 ). Activation of NF-kB seems to be an essential mediator of regulation of iNOS transcription in most cells (63 ). Some of the anti- inflammatory agents such as glucocorticoids, TGF-β, aspirin and thiol compounds act through inhibition of NF-kB via either blocking its nuclear translocation (98 ), inhibiting its transactivation activity by direct protein-protein interaction (99 ) or enhancing expression of its specific inhibitor I κB (100, 101 ). As

discussed earlier, PIN1 acts on p65 subunit of NF-κB, reduces p65 interaction with I κBα, increases its nuclear accumulation and activity and inhibits its degradation after ubiquitination by SOCS1. Based on this information, PIN1 may regulate iNOS induction by acting on NF-κB.

3. AP-1 and other transcription factors

The AP-1 transcription factor is a complex composed of proteins belonging to the c-Fos, c-Jun and other families, in which dimerization is required to promote binding of the complex to AP-1 recognition site. AP-1 induces target genes in response to many physiological and pathological stimuli, including cytokines,

growth factors, stress signals, bacterial and viral infections, as well as oncogenic stimuli. The composition of AP-1 dimers, the quality of stimulus, the cell type and

the cellular environment are important in determining the function of AP-1 (102 ).

Two copies of AP-1 binding sites have been identified in 5’-untranslated region of

the human iNOS gene (103 ). These two sites differed greatly in their ability to induce iNOS in response to cytokine mixture (104 ). The upstream AP-1 site is critical for iNOS induction and mutations decreased gene activation by 90%.

However, the downstream AP-1 site is less important and its mutation only

reduced promoter activity by 45%. Functional AP-1 regulatory regions are required for IFN-γ induced activation of the human NOS2 promoter, suggesting a role for AP-1 activation/binding in the IFN-γ induction of genes. c-Fos and STAT-

1 associated after IFN- γ treatment and bind to GAS element in close proximity to

AP-1 sites to regulate transcription of iNOS (105 ). As indicated earlier, PIN1 is

known to regulate AP-1, but the role of potential effects of PIN1 on AP-1 in iNOS

expression is not known.

Several other transcription factors such as cAMP-responsive element binding

protein (CREB) (106 ), CCAAT-enhancer box binding protein (C/EBP) (97 ), β- catenin (107 ), peroxisome proliferator activated receptor (PPAR) (108 ), hypoxia-

induced factor-1 (HIF-1) (109 ), STAT3 (99 ) and p53 (110 ) are involved in

regulating iNOS expression in different cells.

mRNA stability and translation

Post-transcriptional mechanisms, such as mRNA stability and translation are also

critically involved in the regulation of iNOS expression. The steady-state levels of

many mRNAs are determined in part by their decay rates. The interaction

between cis -acting sequence elements in these mRNAs and certain

ribonucleoproteins controlled mRNA stability. In 3’-untranslated region (3’-UTR) of mRNAs encoding cytokine and other inducible proteins, there is one type of cis -acting instability determinant called adenine-uridine–rich elements (AREs).

AREs stabilize or destabilize mRNA by interacting with different RNA binding proteins such as AU binding factor 1 (AUF-1), Hu antigen R (HuR) and other heterogeneous nuclear ribonucleoproteins (hnRNPs) (111-115 ).

Regulation of iNOS mRNA stability also plays a key role in iNOS induction in mammalian cells. For example, significant basal activity of the human iNOS promoter was shown in some human cell lines such as AKN hepatocytes, while no detectable iNOS mRNA was observed (101 ). TGF-β and other agents have been demonstrated to reduce iNOS mRNA by decreasing iNOS mRNA stability

(116 ).

Several AREs in 3’-UTR have been identified in human (117 ) and rodent (118 ) iNOS mRNA, which may interact with several different RNA binding proteins and regulate its stability. AUF1 interacts with AREs and overexpression of AUF1 in

DLD1 cell destabilizes iNOS mRNA (63 ). In contrast, binding of HuR to the 3'-

UTR of iNOS mRNA seems to stabilize this mRNA. Inhibition of HuR reduced the induction of iNOS mRNA by cytokine, whereas overexpression of HuR

potentiated cytokine-induced iNOS expression (119 ). In addition to HuR and

AUF1, other RNA binding proteins also are involved in the regulation of iNOS.

Two hnRNP family members, L and I, were demonstrated to interact with the 3’-

UTR of murine iNOS mRNA, which is modulated by inflammation, indicating their function in the post-transcriptional regulation of murine iNOS expression (120 ).

As we mentioned earlier, PIN1 regulates GM-CSF mRNA stability by acting on

AUF1. Thus, the possibility of PIN1 to regulate iNOS mRNA stability can not be excluded.

In addition, mRNA translation is a regulated step. In human primary cardiomyocytes, cytokine mixtures induced high and persistent levels of iNOS mRNA, but no iNOS protein (121 ). TGF-β can also reduce iNOS expression by decreasing iNOS mRNA translation in primary murine macrophages (116 ). Lee’s results showed that decreasing l-arginine blocked iNOS protein expression and

NO production without affecting induction of iNOS mRNA, or the stability of iNOS protein (122, 123 ). In fact, depletion of l-arginine inhibits the translation of iNOS mRNA, in which eIF2 α phosphorylation and eIF2 alpha kinase are critically involved (122, 123 ).

Posttranslational regulation and degradation

Posttranslational regulation of iNOS protein includes dimerization, phosphorylation, and protein degradation.

Newly synthesized iNOS has to form a homodimer to be functionally active in NO production in the presence of co-factors and substrates. The regulation of iNOS activity through effects on its dimerization has been demonstrated. For example, kalirin associates with iNOS and inhibits iNOS activity by preventing the formation of iNOS homodimers (124 ). Also, NOS-associated protein-110 kDa

(NAP-110) directly interacts with N-terminus of iNOS, and inhibits iNOS catalytic activity by preventing formation of iNOS homodimers (125 ). A series of potent and selective inducible nitric-oxide synthase (iNOS) inhibitors, such as antifungal imidazoles, was shown to prevent iNOS dimerization in cells and inhibit iNOS in vivo (126 ).

iNOS protein has also been shown to undergo phosphorylation in human and mouse cells (127-130 ). The role of phosphorylation under physiological conditions for iNOS is controversial. In murine macrophages, enhancement of

iNOS tyrosine phosphorylation by a protein tyrosine phosphatase inhibitor is associated with increased iNOS activity in both soluble and insoluble subcellular fractions (130 ). However, Src-mediated tyrosine151 phosphorylation in HEK293 cells redistributed iNOS to detergent-insoluble fraction and decreased activity of human iNOS (129 ). Salh et al also reported that iNOS protein was phosphorylated in concordance with the production of NO, and abolition of iNOS phosphorylation by both rapamycin and Ly294002 resulted in almost complete inhibition of NO production (127 ).

iNOS is ubiquitined and degraded by proteasome activity in some cell types

stimulated in different ways (131-137 ) and by calpain in other experimental

situations (134-137 ). Cell type, stimulus, or other factors may affect the iNOS

turnover path that operates in different systems. Before my study, it was not

known whether PIN1 regulated iNOS degradation by either pathway.

Prostaglandin biology and COX-2 regulation

The prostaglandins (PGs) play an essential role in the maintenance of vascular homeostasis. Deregulation of prostaglandins synthesis has been implicated in many cardiovascular diseases, inflammatory disorder, and cancers. For example,

PGI2 is a vasodilator and inhibits platelet aggregation, whereas TXA2 is a vasoconstrictor and enhances platelet aggregation. Thus, it is important to understand factors and conditions that might affect the synthesis of PGs.

Cyclooxygenease-1 (COX-1) and cyclooxygenease-2 (COX-2) catalyze the rate- limiting step in the formation of PGs from arachidonic acid. COX-1 is expressed in most cells all the time, making it a housekeeping molecule since COX-1 promoter lacks TATA and CAAT boxes but is rich in GC content, a feature of a housekeeping genes. COX-1 leads to production of PGs that maintain integrity of the gastric mucosa, mediate normal platelet function, and regulate renal blood flow (138 ).

In contrast, COX-2 is undetectable in most tissues until a stimulus induces it.

Several extra-cellular signals such as bacterial endotoxin, cytokines, or growth factors rapidly induce COX-2 in many cell types including endothelial cells. COX-

2 appears responsible for synthesis of PGs involved in diseases, particularly those related to acute and chronic inflammation.

Similarly to iNOS, COX-2 is controlled at transcriptional and post-transcriptional levels. (i) Transcription of COX-2 is induced rapidly and transiently in response to many stimuli; (ii) COX-2 mRNA stability and translational efficiency are governed by several regulatory elements in the 3'-UTR; (iii) COX-2 activity and turnover are also regulated (139 ).

Transcription

The human COX-2 gene promoter contains several regulatory elements that stimulate COX-2 transcription in different cell types (140 ). Transcription factors, including NF-kB (140-142 ), AP-1 (142-144 ), STAT1 (142 ), CREB (145 ), C/EBP

(146 ), β-catenin-T cell factor-4 (TCF-4) (147 ), nuclear factor activated T cells

(NFAT) (142, 148 ) and nuclear factor for IL-6 expression (NF-IL6) (141, 142 ), regulate COX-2 transcription either alone or synergistically. Transcriptional regulation of the COX-2 gene is very complex in that it can involve numerous signaling pathways, and the mechanism varies depending on the specific stimulus and the cell type. Most of the transcriptional signaling pathways require

activation of some proline-directed kinases including ERK1/2 and p38 (149 ), PKC and GSK-3 (148, 150 ).

mRNA stability

COX-2 expression can also be critically regulated at post-transcriptional level, which is mediated by AREs present within the 3’-UTR of COX-2 mRNA. The

COX-2 ARE associates with different ARE RNA-binding proteins to target the mRNA for either stabilization, rapid decay, or inhibition of translation (151 ). For

example, AREs in the 3’-UTR of COX-2 mRNA cause rapid mRNA decay by

interacting with different RNA binding proteins (152 ). HuR and tristetraprolin

(TTP) binds to the COX-2 AREs and exert opposite effects on COX-2 expression.

In normal colon cells, low levels of nuclear HuR and higher levels of TTP were

responsible for rapid decay of COX-2 mRNA. Increased HuR and decreased TTP,

as detected in colon cancer cells, results in elevated expression of COX-2 (152,

153 ). COX-2 mRNA was also stabilized by activating β-catenin in NIH3T3 and

293T cells, which is mediated by the interaction between β-catenin with the COX-

2 ARE in vitro and in vivo. In addition, β-catenin induced the cytoplasmic localization of HuR, which may bind to β-catenin and facilitate β-catenin-

dependent stabilization of COX-2 mRNA (154 ). The translational repressor

protein, TIA-1, can also interact with COX-2 AREs and inhibit mRNA translation

without affecting COX-2 transcription or mRNA turnover (155 ). Collectively,

rapid mRNA decay and translational inhibition play a significant role in controlling

COX-2 gene expression and if deregulated, allow for overexpression of COX-2,

contributing to tumorigenesis. Although PIN1 regulates ARE binding proteins and

several transcription factors mentioned here, the effect of PIN1 on COX-2 mRNA

has not been described.

Activity and protein degradation

COX-2 enzymatic activity is affected directly by nitric oxide (156 ). NO has high

affinity for the iron heme center in the catalytic domain of COX-2 and stimulates

COX-2. In addition, iNOS specifically binds to COX-2 and the iNOS-derived NO

can S-nitrosylate it, enhancing COX-2 activity. Disruption of iNOS-COX-2 binding

prevented NO-mediated activation of COX-2 (157 ).

COX-2 degradation limits the amount of COX-2 in normal cells. The deregulation

of COX-2 degradation leads to its overexpression in tumors and contributes to

various pathologies such as colon cancer (158, 159 ). Several different

mechanisms have been reported to regulate COX-2 degradation. Proteasome- mediated degradation is initiated by post-translational N-glycosylation at Asn-594.

COX-2 is embedded in the luminal membrane of the endoplasmic reticulum (ER) and the N-glycosyl group is then processed and unfolded, presumably by ER chaperones. Then COX-2 is translocated to the cytoplasm and undergoes ubiquitination and proteolysis by 26S proteasome (160-167 ). Substrate-

dependent suicide inactivation is the second pathway for COX-2 protein

degradation. COX-2 inhibitors retard COX-2 protein degradation in HEK293 cells

expressing COX-2. Although the mechanism of substrate-dependent

degradation is not clear, proteasome activity seemed not be involved (162, 168,

169 ). The results from our lab, and from others, also showed that COX-2 is a

calpain substrate (170, 171 ). In human synovial fibroblasts, cleavage of COX-2

was reduced by a calpain inhibitor, E64d, indicating a role for calpain and/or

cathepsin. In chapter 3, I describe the effects of other selective calpain inhibitors

on COX-2 induction in MAEC. Although PIN1 regulates the susceptibility of

several proteins (for example, cyclin E and c-myc) to ubiquitination and

proteasomal degradation, its effect on COX-2 turnover has not been investigated.

In summary, induction of iNOS and COX-2 can be regulated at multiple

mechanisms: transcription, post-transcription (mRNA stability), translation and

post-translation (phosphorylation and degradation). PIN-1 is the only identified

peptidyl-prolyl isomerase that can catalyze the isomerization of pSer/pThr-Pro motifs and regulate the substrate protein function in a phosphorylation-dependent manner. PIN1 interacts and increases the level or activity of several transcription factors that are critical for iNOS and COX-2 transcription. These transcription factors include NF-κB (18 ), c-Jun (32 ), c-Fos (30 ), β-catenin (29 ), STAT-1α (Fig.

1.4), STAT3 (172 ), C/EBP (173 ), HIF-1α (174 ) and p53 (23, 24 ). Furthermore, mRNA stability is regulated by interaction between AREs in 3’UTR and RNA binding protein including AUF-1, which is a PIN1 substrate (20, 21 ). PIN1 also regulates the turnover of substrates by modulating ubiquitin-dependent protein degradation (29, 32, 175, 176 ). Phosphorylation in Ser/Thr residues has been observed in human and mouse iNOS (127-130 ), which contains several Ser/Thr-

Pro motifs and may be putative PIN1 substrates. Considering these significant results and the critical role of iNOS and COX-2 in vascular diseases, it is necessary to investigate whether, and how, PIN1 regulates iNOS and COX-2 expression in EC.

Calpain and Calpastatin (CAST)

In chapter 2 and 3, I describe that PIN1 depletion has no effect on iNOS or COX-

2 mRNA. Instead, depletion of PIN1, or calpain inhibitors increased the induction

of iNOS and COX-2. Depletion of PIN1 also reduced calpain activity in MAEC

(chapter 4). Given the potential role for calpain suggested by these studies, I now

provide a review of calpain and its regulation.

Calpains are calcium-dependent neutral cysteine proteases. Some isoforms are

widely expressed, whereas others are only expressed in specific tissues. In

mammals, the two best-characterized members of the calpain family, µ-calpain

and m-calpain are ubiquitously expressed and play crucial roles in many different physiological processes. Transient influx of calcium into cells can activate calpains locally (for example, those close to Ca2+ channels). The activated calpains play a physiological role by catalyzing the controlled proteolysis of its target proteins. Overactivation of µ- and m-calpain has been implicated in a number of pathologies associated with altered calcium homeostasis such as AD, and cataract formation, as well as secondary degeneration resulting from myocardial ischemia, cerebral ischemia, traumatic brain injury and spinal cord injury (177 ). Calcium overload excessively activated calpain in these cases,

which leads to unregulated proteolysis of target protein with consequent irreversible tissue damages.

So far, more than one hundred proteins have been identified as calpain substrates. Several structural proteins such as spectrin (178 ) and talin (179 ) can be cleaved by calpains, indicating the important role of calpains in remodeling the cytoskeletal structures and regulating cell spreading and mobility. Functional cleavage of kinases (protein kinase C (180 ), focal adhesion kinase (181 ) and

GSK-3 (182 )), phosphatases (183 ) and receptors (( β-integrin (184 ), and epidermal growth factor receptor (185 ) and androgen receptor (186 )) allows calpain to regulate signal transduction. Calpains may also regulate gene expression by cleavage of several transcription factors, such as c-Jun, c-Fos

(187, 188 ), c-myc (189, 190 ), β-catenin (191, 192 ), NF-kB (193 ) and p53 (194 ).

Calpain can also trigger apoptosis by acting on Bcl-2 proteins (195 ).

These substrates may contain one or several calpain cutting sites and undergo limited proteolysis by calpains, generating protein fragments with different functions. For example, m-calpain cleaved GSK-3 at the N-terminus and generated two fragments of 40 kDa and 30kDa. Such truncation increased GSK-

3 activity because of the removal of N-terminal inhibitory domain (182 ). Calpain cleaved β-catenin at the N-terminus in neurons and the truncated fragments lost the sensitivity to GSK-3β-directed proteolysis and became more stable (196 ).

Tompa’s study showed calpain substrates have high frequency of Ser/Thr-Pro

motifs (197 ), which is the consensus site of proline directed kinases and the

phosphorylated forms may associate with PIN1. It is known that phosphorylation

in Ser/Thr-Pro motifs is involved in the susceptibility to proteases (12, 198 ). Thus, it is necessary to investigate whether PIN1-catalyzed isomerization after phosphorylation regulates the susceptibility of substrates to calpains.

Domain structure

µ-calpain is a heterodimer of calpain 1 and 4, while m-calpain is a heterodimer of

calpain 2 and 4. Several domains are contained in calpain 1 and 2: 1) a NH2-

terminal sequence; 2) the active domain IIa and IIb; 3) domain III; 4) domain IV.

The amino-terminal sequence may interact with small subunit and be involved in

calpain stability. Domain IIa and IIb contain a catalytic triad formed by Cys- His-

Asn. Domain III is also involved in binding to phospholipids (199 ). Calpain 4, the

common small subunit in µ and m, contains two functional domains: a NH2- terminal hydrophobic domain V and C-terminal domain VI. Both Domain IV from calpain 1/2 and Domain VI from calpain 4 contain penta-EF-hand (PEF) domains.

The heterodimeric formation between the two subunits of µ- and m-calpain is through a pairing of the fifth EF-hand of Domain IV and Domain VI.

In the absence of calcium, µ- and m-calpain are inactive since Cys residue is distant from His and Asn residues within the active site. This crystal structure reveals that binding of calcium to domain IV in calpain 1/2 and domain VI in calpain 4 induces conformational changes, producing a more compact structure.

Critical residues in domain IIa and IIb can move together to form a catalytically active complex (200 ).

I IIa IIb III IV phospholipid Penta-EF-hand CAPN 1 Active domain binding domain CAPN 2 P P P P P P P P P

dimerization

V VI Penta-EF-hand Gly rich CAPN 4 domain

Fig 1.5 Domain structure of µ- and m-calpain.

Regulation of calpain activity

Calpain activity is regulated at multiple levels: expression of the subunits (201,

202 ), calcium requirement (203 ), phosphorylation (204, 205 ) and, most

importantly, by the endogenous inhibitor, calpastatin (CAST) (201-209 ).

Expression of calpains

The 5’-upstream region of both large and small subunit gene lacks a TATA or

CAAT box sequences and is GC rich. As mentioned earlier, this is a feature of

housekeeping genes. However, recent studies showed that calpain 2 can also

be induced by different cellular stimuli. For example, the COX-2 selective

inhibitor, NS-398, decreased mRNA level of calpain 2 in IEC-6 cells (202 ). The

mRNA level of calpain 2 ,but not calpain 1, can be upregualted by treatment with

12-O-tetradecanoylphorbol-13-acetate (TPA) in HeLa cells (210 ) or by eccentric

exercise in skeletal muscles (201 ).

Calcium requirement

The physiological concentration of calcium in living cells is 50-500 nM (206, 211,

212 ), which is sufficient to activate µ-calpain, but much less than the 400-800 µM required to activate m-calapin in vitro. These calcium concentrations may not apply to calpains in vivo since biomolecules in cell that interact with calpains can reduce their calcium requirement. For example, certain phospholipids including phosphatidylinositol 4,5-bisphosphate (PIP2) lower the calcium concentration required for activation of µ- and m-calpain (213, 214 ). Similarly, protein-protein interactions, such as acyl-CoA-binding proteins, may also change the calcium requirement of calpains (215, 216 ).

Phosphorylation

Both calpain 1 and calpain 2 can be phosphorylated at Ser, Thr and Tyr residues.

The phosphorylated residues are mainly located in two areas: at the domain I/II

junction (IIa) and at the N-terminus of domain II (IIb) (206 ). Phosphorylation at

several sites controls the activity of calpain. For example, m-calpain is activated

by phosphorylation of calpain 2 at Ser50 by ERK during migration of fibroblasts

and in keratinocytes stimulated with EGF (204, 205, 217-220 ). Calpain activity can also be inhibited by phosphorylation. Phosphorylation of calpain 2 by protein

kinase A reverts m-calpain to an inactive conformation and blocks EGF-induced activation of m-calpain during fibroblast migration (205 ). Activated protein kinase

Ciota (PKCiota) by nicotine induces phosphorylation of both µ- and m-calpains

and is involved in accelerated migration and invasion of human lung cancer cells

(221 ).

CAST

The activity of calpains is tightly controlled by the endogenous inhibitor CAST.

CAST was originally identified when no calpain activity could be detected in

crude porcine skeletal muscle homogenates until precipitation of calpain,

indicating its efficiency in inhibiting the heterodimeric calpains (222 ). CAST is an

intrinsically unstructured protein, which is converted into a defined structure only

on binding to active calpain in the presence of calcium (200 ). CAST contains an

N-terminal non-inhibitory leader L domain and four conserved inhibitory domain I-

IV. Although the L domain is not inhibitory, it binds to calcium-free calpain, and

may affect the distribution of calpains before activation (223 ). Each inhibitory

domain is able to efficiently and specifically interact with one calpain molecule

and block its activity although the affinity and inhibitory efficiency may be different

(222 ). The specificity of inhibition of heterodimeric µ- and m-calpain and not other

cysteine protease is determined by the unique interaction between CAST and calcium-bound calpain. Two recent crystallographic studies demonstrated how

CAST blocks the activity of the calpain by using a previously undiscovered mechanism. Three subdomains, A, B, and C are contained in each inhibitory domain of CAST. Subdomain A and C is response to binds the first EF-hand sequence of Domain IV (calpain 1/2) and Domain VI (calpain 4) respectively, which increase the affinity of interaction and the inhibitory activity of subdomain B.

B contacts DI-DIII region of calpain 1/2 and blocks the active site. The highly conserved sequence of B generates a well-organized local structural conformation, which allows it wrap to around calpain’s active site and escape from the cleavage by calpains (200, 224, 225 ).

L-domain CID-I CID-II CID-III CID-IV

A B C A B C A B C A B C CAST

Fig. 1.6 Domain structure of CAST

Inactive calpain Active calpain

DII Active site DI DIII Calcium binding

DVI DIV

CID binding C

B B

A C A

Fig 1.7 CAST specifically binds and inhibits m-calpain

Calpains are a major ubiquitous cellular proteolytic system. Disruption of the calpain–CAST balance occurs in several pathological conditions where calpain is over-activated. These diseases include AD, cancer, cataract formation, traumatic

brain and spinal cord injury, ischemia/reperfusion injury, and atherosclerosis

(177 ). Thus maintaining the balance by increasing the expression of CAST

protein or its inhibitory activity is crucial to protect from pathological disorders

associated with excessive calpain activity.

Deregulations of both PIN1 and the calpain system have each been implicated in

certain human diseases. PIN1 may negatively correlate with calpain activity in

human diseases, including AD and spinal cord injury. For example, deletion of

PIN1 and enhanced calpain activity contributes to pathological process of AD (8,

28, 37, 226 ). PIN1 binds and stabilizes phospho-MCL-1, thus protecting

oligodendrocytes from apoptosis after spinal cord injury (37 ). Upregulation of

calpain, has been implicated in apoptosis and tissue degeneration in spinal cord

injury, while overexpression of CAST and calpain inhibitor prevented apoptosis in

spinal cord injury (227-235 ). Collectively, PIN1 appears to protect, while calpain

may contribute to such diseases. In early studies, I found that PIN1 deletion did

not affect iNOS and COX-2 mRNA. Therefore, I investigated the effects of PIN1

on expression and turnover of iNOS and COX-2, and on calpain activity in MAEC.

CHAPTER 2

PIN1 REGULATES DEGRADATION OF iNOS

IN ENDOTHELIAL CELLS

Abstract

The peptidyl-proline isomerase Protein Never in Mitosis Gene A Interacting-1

(PIN1) increases the level or activity of several transcription factors that can

induce iNOS. PIN1 can also regulate mRNA and protein turnover. Here, the effect of depletion of PIN1 on induction of iNOS by LPS/IFN-γ in murine aortic

endothelial cells (MAEC) was determined. Suppression of PIN1 by 85% with small hairpin RNA enhanced the induction of NO and iNOS protein by LPS/IFN-γ.

There was no effect on induction of iNOS mRNA, suggesting a posttranscriptional effect. The enhanced levels of iNOS protein were functionally significant since

LPS/IFN-γ was cytotoxic to MAEC lacking PIN1 but not MAEC harboring an inactive control construct, and because cytotoxicity was blocked by the NO synthase inhibitor Nω-nitro-L-arginine methyl ester. Consistent with posttranscriptional action, knockdown of PIN1 increased the stability of iNOS protein in cycloheximide-treated cells. Furthermore, loss of iNOS was blocked by the calpain inhibitor carbobenzoxy-valinyl-phenylalaninal but not by the selective

proteasome inhibitor epoxomicin. Immunoprecipitation indicated that PIN1 can interact with iNOS. Pull down of iNOS with a wild-type glutathione-S-transferase-

PIN1 fusion protein, but not with a mutant of the amino terminal phospho-

(serine/threonine)-proline binding WW domain of PIN1, indicated that this domain mediates interaction. The results suggest that PIN1 associates with iNOS and

can limit its induction by facilitating calpain-mediated degradation in MAEC.

Introduction

Cis-trans isomerization about the bond between an amino acid and proline

regulates protein conformation, interactions, and susceptibility to enzymes, including kinases, phosphatases, and proteases. Interconversion between the cis and trans forms is slow, and proline (P), preceded by phosphorylated (p) serine

(S) or threonine (T), isomerizes even more slowly than unphosphorylated sequences. Protein Never in Mitosis Gene A Interacting-1 (PIN1) is the only known cis-trans peptidyl-prolyl isomerase that acts on p(Ser/Thr)-Pro (236, 237 ).

PIN1 binds to p(S/T)-P via its amino terminal WW domain. It also catalyzes

proline isomerization with its carboxyl terminal catalytic domain(237, 238 ).

The fundamental effect of PIN1 on protein conformation may account for its

diverse roles in cell cycle progression, proliferation, and transcription (18, 239 ).

PIN1 increases the activity of β-catenin, NF-κB, and AP-1. PIN1 also inhibits degradation of β-catenin and NF-κB p65, increasing their protein level and nuclear accumulation (18, 29, 30 ). These transcription factors are associated with

induction of inflammatory gene products. PIN1 is also known to regulate degradation of other proteins and mRNA stability (8, 21, 240 ).

iNOS is one of the inflammatory genes activated by β-catenin, NF-κB, and AP-1

(63, 72, 107, 241, 242 ). After induction, iNOS can produce large amounts of NO

from l-arginine in many cell types, including endothelial cells (80, 242, 243 ).

Whether induction of iNOS is beneficial or pathogenic depends on additional

factors, such as the concentration of l-arginine, cofactors, inhibitors, and oxidants

(244-248 ). A role for PIN1 in the induction of iNOS has not been determined.

Although PIN1 regulates protein conformation and turnover of many of its identified substrates, its role in NO production has not been investigated. Since

PIN1 regulates factors that could affect iNOS, the effect of PIN1 knockdown in

murine aortic endothelial cells (MAEC) stimulated with Escherichia coli endotoxin

LPS and IFN-γ was determined.

Materials and Methods

Endothelial cell growth supplement, heparin, phenylmethylsulfonyl fluoride,

Bradford reagent, E. coli LPS, serotype 0111:B4, N ω-nitro-L-arginine methyl

ester (L-NAME), and polybrene were purchased from Sigma Chemical (St. Louis,

MO). Recombinant mouse IFN-γ was from R&D Systems (Minneapolis, MN).

Carbobenzoxy-valinyl-phenylalaninal (zVF), also known as calpain inhibitor III or

MDL-28170, PD150606, epoxomicin, and cycloheximide (CHX) were obtained from Calbiochem (La Jolla, CA). Fetal bovine serum was purchased from

Hyclone Laboratories (Logan, UT). Tris-base, ethylenediamine tetraacetic acid,

NaCl, Na3VO4, NaF, Tween 20, sodium dodecyl sulfate, ethidium bromide, and agarose were obtained from Fisher Scientific (Fair Lawn, NJ). Triton X-100 was purchased from Pierce (Rockford, IL). HEK293T cells were obtained from

American Type Culture Collection (Manassas, VA). Silica columns were purchased from Quiagen (Valencia, CA). Fugene 6 transfection reagent was obtained from Roche (Indianapolis, IN). Glutathione-sepharose was purchased from Amersham Biosciences (Uppsala, Sweden). Quickchange mutagenesis and

RNAMaxx in vitro transcription reagents were obtained from Stratagene (La Jolla,

CA). Recombinant dicer was from Ambion (Austin, TX). Dulbecco's minimum essential medium, trypsin, isopropyl-β-d-thiogalactopyranoside, TRIzol,

Superscript Reverse Transcriptase, Platinum PFX and Taq DNA polymerases,

RNAse-free DNAse, deoxynucleotides, recombinant protein G agarose, and plasmid pcDNA 3.1 CTGFP TOPO were purchased from Invitrogen (Carlsbad,

CA). Plasmid pGEX4T-3 was from Pharmacia (Piscataway, NJ). Plasmids for lentivirus production were purchased from Addgene (www.addgene.org), where they were deposited by Drs. Robert Weinberg (pCMV-dR8.2 dvpr and pCMV-

VSV-G; Whitehead Institute for Biomedical Research) and David Root (pLKO.1;

Massachusetts Institute of Technology).

Anti-iNOS antibody was from Transduction Laboratories (Lexington, KY). Anti-

PIN1 was purchased from R&D Systems (Minneapolis, MN). Anti-NFKB p65 antibody is from Rockland Immunochemicals (Gilbertsville, PA). Anti-glutathione-

S transferase (GST) was prepared as described previously (249 ). Horseradish

peroxidase-conjugated goat anti-mouse and goat anti-rabbit secondary

antibodies were from Jackson Immunoresearch Laboratories (West Grove, PA).

Renaissance Enhanced Chemiluminescence Reagent was purchased from New

England Nuclear Life Sciences (Boston, MA). DETA NONOate was from Caymen

Chemical (Ann Arbor, MI).

Cells

MAEC were cultured from aortas as described previously (243 ). The protocol for

cell isolation was carried out under approval of the Ohio State University Animal

Care and Use Committee.

Short hairpin RNA

The oligomer 5'-CCG GCC GGG TGT ACT ACT TCA ATC ACT CGA GTG ATT

GAA GTA GTA CAC CCG GTT TTT G-3' was designed from murine pin1

(Genbank accession NM 023371) to produce a hairpin structure (self-

complimentary sequences underlined) with an intervening loop CTCGAG and

targeting the 5' underlined sequence in the PIN1 mRNA. This oligomer was

annealed to a complimentary oligomer 5'-AAT TCA AAA ACC GGG TGT ACT

ACT TCA ATC ACT CGA GTG ATT GAA GTA GTA CAC CCG G-3' to produce

Age I- and EcoR I-compatible overhangs. The annealed DNA was ligated into

Age I- and EcoR I-restricted pLKO.1 vector with T4 ligase, adjacent to a U6 RNA

polymerase promoter to drive transcription the shRNA (250 ), as described by

Moffat et al (251 ). A mutated sequence was also produced as a control short hairpin RNA (shRNA).

Lentiviral particles were produced as described by Stewart et al. (252 ). In brief, a

mixture of 6 µg pLKO.1 vector, 5 µg pCMV-dR8.2 dvpr, and 2 µg pCMV-VSV-G,

complexed with Fugene 6 in Dulbecco's minimum essential medium, was added

to HEK293T cells in a 75-cm2 flask. Two days after cotransfection, the virus

containing supernatant was collected and filtered (0.45 µm) to remove cell debris.

The supernatant was stored at –80°C until used to treat MAEC.

Stable PIN1 knockdown (KD) or control (C) shRNA cells were generated using

described methods (253 ). MAEC were cultured for 24 h and then infected with

500 µl of lentiviral supernatants and 8 µg polybrene/ml. After 24 h the viral

solution was removed and fresh medium was added. Cells were then grown with

9 µg puromycin/ml. After cell killing and regrowth, cells were transferred to a new

flask with puromycin. This selection was repeated four times. The entire process

was repeated with two additional MAEC preparations, resulting in three

independent stable KD and control shRNA cell cultures.

Small inhibitory RNA

PIN1 small inhibitory RNA (siRNA) was generated by in vitro transcription from a

double-stranded DNA template made by reverse transcription-polymerase chain

reaction of MAEC RNA with platinum PFX polymerase and primers containing

the T7 RNA polymerase promoter sequence at their 5' ends, as described

previously (254 ). PIN1 cDNA was amplified with sense primer 5'-GCG TAA TAC

GAC TCA CTA TAG GGA GAA CGA GGA GAA GCT GCC ACC A and antisense primer 5'-GCG TAA TAC GAC TCA CTA TAG GGA GAT CTG TGC

GCA GGA TGA TAT G based on Genbank accession number NM 023371

(underlined bases). A control sequence from green fluorescent protein was also produced as described previously (254 ). After in vitro transcription of the PCR products with T7 RNA polymerase, long double-stranded RNA was digested with recombinant Dicer. The siRNA was purified and MAEC were transfected by electroporation with 150 nM siRNA, as described (254 ).Treatments of these

MAEC were initiated 24 h after transfection.

Treatments

MAEC were cultured in Dulbecco's minimum essential medium-0.5% fetal bovine serum for 18 h and then treated with this medium alone, which contained 0.54 mM L-arginine, or this medium containing 10 µg LPS and 20 ng IFN-γ per

milliliter, and other agents for various times. Where necessary, protein synthesis

was inhibited with 90 µg cycloheximide/ml. Some cells were treated with DETA

NONOate.

NO accumulation

Medium was collected, and nitrite, reflecting NO, in medium was measured by the Griess reaction, as described previously (243 ).

Cytotoxicity

To assess cell injury, MAEC were cultured in 96-well plates and treated as

described. Cells were rinsed with PBS and fixed with 3% formaldehyde, and

adherent cells were stained with 0.5% crystal violet, solubilized, and A590

measured (255 ).

Superoxide anion assay

Nitroblue tetrazolium (NBT) reduction assay is used to determine the production

of superoxide anion (O2(-)) (256 ). In brief, KD and Control shRNA MAEC were

treated with vehicle or LPS/IFN-γ for 23h, then treated with NBT (0.25 mg/ml) for

1h to allow the formation of NBT formazan deposits, a product of NBT- intracellular O2(-) reaction. Then cells were rinsed with PBS twice and methanol once to remove extracellular NBT. Intracellular NBT formazan deposits were dissolved by potassium hydroxide (2M) and dimethylsulfoxide and then measured its absorbance using a microplate reader at 620 nm.

mRNA levels

Cells were rinsed with phosphate-buffered saline and lysed with TRIzol reagent, and RNA was extracted and recovered by ethanol precipitation. RNA was dissolved in water, and its concentration was determined by absorbance of an aliquot at 260 nm. Three micrograms of RNA were reverse transcribed, and cDNA was subjected to polymerase chain reactions for murine iNOS, VCAM-1, and β-actin as described previously (80, 257 ). Fifty percent of each reaction was subjected to agarose electrophoresis and ethidium bromide staining. Gels were digitally photographed.

GST-PIN1 fusion proteins. Murine PIN1 was cloned by reverse transcription and polymerase chain reaction with Platinum PFX Polymerase followed by addition of

Platinum Taq polymerase to add A to the 3' ends of the expected 504 base reaction product. Sense (5'-GAA GAT GGC GGA CGA GGA GA-3') and antisense (5'-CCT CAT TCT GTG CGC AGG A-3') primers were designed from

Genbank accession number NM 023371. A sample of the reaction was incubated with pcDNA 3.1 CTGFP TOPO as instructed by the manufacturer and then used to transform E. coli DH5 . Colonies were selected and clones confirmed by

complete sequencing. The wild-type plasmid was used for polymerase chain

reaction mutagenesis. Primers to generate the lysine-64 to alanine (K64A)

mutation were 5'-CTC ACA TCT GCT GGT GGC GCA CAG CCA GTC TCG G-3' and 5'-CCG AGA CTG GCT GTG CGC CAC CAG CAG ATG TGA G-3'. Primers to generate the tryptophan-33 to alanine (W33A) mutation were 5'-CAA CGC

CAG CCA GGC GGA GCG GCC CAG C and 5'-GCT GGG CCG CTC CGC CTG

GCT GGC GTT G-3'. After digestion of the wild-type template with DpnI and bacterial transformation, positive clones were selected and sequenced to confirm mutagenesis.

To produce GST fusions of these PIN1 proteins, wild-type W33A and K64A plasmids were used as templates in polymerase chain reactions with Platinum

PFX polymerase, plasmid-specific sense primer, 5'-GAT CGG ATC CAT GGC

GGA CGA GGA GAA GC-3' containing a BamHI restriction site, and antisense primer, 5'-GCA TGC CTG CTA TTG TCT-3', located 3' of the PIN1 stop codon in each plasmid. Each reaction product was digested with BamHI and NotI, isolated from an agarose gel, and ligated into pGEX-4T3 with T4 ligase in frame to the carboxyl terminal of the GST sequence. Ligation reactions were used to transform E. coli, and clones were selected and confirmed by sequencing. GST-

PIN1 fusion proteins were produced from transformed E. coli BL21 cells induced with isopropyl-β-d-thiogalactopyranoside. Bacteria were sonicated in phosphate- buffered saline and extracted with 1% Triton X100, and debris was removed by

centrifugation (16,000 g x 10 min). The extract was incubated with glutathione- sepharose, washed with 0.1 M Tris pH 7 and stored at 4°C (258 ).

Immunoprecipitation, GST pulldown, and Western blot analysis

Cells were washed with phosphate-buffered saline and lysed and sonicated in lysis buffer containing 1% Triton X-100, 50 mM Tris (pH 7.5), 250 mM NaCl, 5 mM ethylenediamine tetraacetic acid, 4 mM Na3VO4, 20 mM NaF, 1 mM phenylarsine oxide, 30 µg/ml aprotinin, and 30 µg/ml leupeptin. Nuclei were first isolated from some cells by centrifugation, as described previously (80 ), and treated with lysis buffer before processing for Western blot analysis. Protein concentration was determined by the method of Bradford (259 ).

For immunoprecipitation, 500 µg of cell lysate protein were incubated with 5 µg anti-PIN1 antibody for 2 h at 4°C with rocking. Recombinant protein G agarose was added, and samples were incubated for 1 h. For GST pulldowns, 500 µg of

MAEC lysate protein were precleared by incubation with glutathione-sepharose at 4°C for 2 h. The supernatant was then incubated with the specific glutathione- sepharose-GST-fusion proteins at 4°C for 2 h.

All samples were then centrifuged 16,000 g x 1 min, and the supernatant was discarded. The agarose or sepharose pellets were washed three times with lysis buffer by resuspension and centrifugation.

For Western blot analysis, 12 µg proteins were mixed with 20 µl denaturing sample buffer (62.5 mM Tris, pH 6.7, 10% glycerol, 2% sodium dodecyl sulfate,

5% 2-mercaptoethanol, and 0.003% bromphenol blue). Rinsed agarose or sepharose pellets from immunoprecipitation or GST-pulldown were resuspended in 30 or 100 µl denaturing sample buffer, respectively. Samples were heated for

10 min at 95°C (260 ). Proteins were separated on 4–20% Tris-gylcine polyacrylamide gels and transferred to nitrocellulose.

After transfer, the blots were blocked with 5% nonfat dry milk in Tris-buffered saline buffer and Tween 20 (0.1% Tween 20, 10 mM Tris, pH 7.5, and 150 mM

NaCl) and probed with primary antibody. After blots were washed, appropriate horseradish peroxidase-conjugated secondary antibody was added and incubated for 1 h. Proteins were visualized by using enhanced chemiluminescent reagents, captured on X-ray film, and scanned to produce digital images.

Data analysis

Protein levels in Western blot analyses and polymerase chain reaction products in images were quantified with Image J 1.22d (NIH). Data were analyzed by

Student's t-test or analysis of variance (ANOVA) with Bonferroni correction for multiple comparisons.

Results

PIN1 targeting shRNA and siRNA efficiently knockdown PIN1 protein

PIN1 was reduced more than 85% with the specifically targeted PIN1 knockdown shRNA vector and not with the inactive control shRNA construct (Fig. 2.1A/B). PIN1 protein level in control shRNA cells was similar to that in primary MAEC, indicating that the control shRNA was inactive (Fig. 2.1B). PIN1 was also suppressed by transient transfection with a siRNA pool produced from a 538-bp reverse transcription/polymerase chain reaction product of PIN1 mRNA that was digested with recombinant Dicer, in comparison with control RNA produced from a green fluorescent protein cDNA (Fig.

2.1C).

5' - ccggCCGGGTGTACTACTTCAATCActc KD ||||||||||||||||||||| gtttttGGCCCACATGATGAAGTTAGTgag

5' - ccggCCGG GTGTACT GCTTCAATCActc Control |||||||-|||-||||||||| gtttttGGCCCAC-TGATGAAGTTAGTgag

B. - PIN1

- α-tubulin

125

100 75

50 -tubulin Signal Ratio α α α α 25 +

0 PIN1/ KD Control None shRNA

Fig 2.1 Suppression of PIN1 in MAEC continued

Fig 2.1 continued

C. KD Control

25 PIN1 20

10 + 5 +

PIN1 Signal Intensity 0 PIN1 Control siRNA

Fig 2.1 Suppression of PIN1 in MAEC. A: sequences encoding PIN1 knockdown

(KD) short hairpin RNA (shRNA) and inactive control shRNA (control) in the lentiviral system. B: KD or control were used to generate stable MAEC clones as described in the MATERIALS AND METHODS. Proteins were extracted from KD shRNA, control shRNA, or uninfected MAEC (None), and Western blotted with antibodies specific for PIN1 and α-tubulin. C: PIN1 was also suppressed using an small inhibitor RNA (siRNA) pool produced by in vitro transcription of long

double-stranded RNA and digestion with recombinant Dicer. Representative blots are shown. Densitometric analysis of signal intensities of images was used to measured PIN1 expression. Bars represent means + SE of 3–4 independent cultures of each group. +p < 0.001 for comparison with between KD and control shRNA.

PIN1 suppression increases iNOS induction in response to LPS and IFN-γ

Previously, it was found that cotreatment with LPS and IFN-γ was required to

induce iNOS in MAEC (80, 243 ). Here, the effect of PIN1 shRNA on iNOS expression was measured in KD and control shRNA cells stimulated with 10 µg

LPS and 20 ng IFN-γ per ml. Western blot analysis demonstrated that iNOS was

induced more in PIN1 KD than in control shRNA cells 24 h after treatment with

LPS/IFN-γ (Fig. 2.2A). Similar results were obtained in two other independent

pairs of cells selected for KD and control shRNA (data not shown). Suppression

of PIN1 by transient transfection with siRNA also increased the induction of iNOS

(Fig. 2.2B). Genetic deletion of PIN1 also increased iNOS induction, as shown in

Fig. 2.2C. Reconstitution of PIN1 plasmid in KD cells decreased iNOS induction

in response to LPS/IFN-γ (Fig. 2.2D). Therefore, induction of iNOS protein was increased by two different methods of PIN1 suppression and PIN1 genetic deletion.

A. KD Control 4 iNOS + 3 α-Tubulin -tubulin 2 α α α α

Signal Ratio Signal 1 iNOS/

0 KD Control shRNA

B. KD 12 + 10 iNOS

2 iNOS Signal Intensity Signal iNOS 0 PIN1 Control siRNA

Fig. 2.2 Effect of PIN1 knockdown or genetic deletion on iNOS continued

Fig. 2.2 continued

ko wt 0.75 iNOS

0.50 -Tubulin α-Tubulin α α α α

Signal Signal Ratio 0.25 iNOS/

0.00 KO WT

GF -2 PIN1 D iNOS 0.2 GF-2 PIN1 α-Tubul in

-Tubulin 0.1 PIN1 α α α α Signal Ratio iNOS /

0.0

Fig. 2.2 Effect of PIN1 knockdown or genetic deletion on iNOS. KD and control

shRNA MAEC ( A), uninfected MAEC electroporated with diced control or PIN1

siRNA ( B) or MAEC from PIN1 knockout (ko) and wildtype (wt) mice were treated with LPS and IFN-γ for 24 h. Representative Western blots are shown in C. Bars represent means + SE ratio of iNOS/ α-tubulin from densitometric analysis of three cultures of each group. +p < 0.05 for comparison between KD and control shRNA. D. 2 µg of empty vector (GF-2) and PIN1 plasmids were transfected in

PIN1 KD MAEC. Cells were starved for 16h and treated with LPS/IFN-γ for 24h.

Representative Western blots are shown.

Knockdown of PIN1 does not influence iNOS and VCAM-1 mRNA

Despite the increases in iNOS protein there was no difference in iNOS mRNA

between KD and control shRNA cells from 1 to 48 h after treatment with

LPS/IFN-γ (Fig. 2.3, A and B). LPS-IFN-γ also induced VCAM-1 mRNA as expected, and, as with iNOS, PIN1 depletion did not increase the levels of this message (Fig. 2.3C). Nuclear NF-κB p65, which may contribute to transcription

of both iNOS and VCAM-1 mRNAs, rose slightly more rapidly in KD MAEC when

compared with that of control, but the peak level was similar in KD and control

shRNA cells (Fig. 2.3D). These results are consistent with post-transcriptional

regulation of iNOS by PIN1 in MAEC.

A. Time (h): 0 1 2 4 8 18 48

shRNA: KD C KD C KD C KD C KD C KD C KD C iNOS

ß-actin

B. C. 1.5 KD 2.0 * KD Control * Control 1.6 1.0 * *

-Actin 1.2 + β β β β

0.5 0.8 iNOS/ VCAM-1/ß-actin

mRNA mRNA Signal Ratio 0.4 mRNA Ratio Signal mRNA 0.0 0 1 2 4 8 18 48 0.0 0 1 2 4 8 Time (h) Time(h)

Fig. 2.3 Effect of PIN1 knockdown on iNOS and VCAM-1 mRNA and NF-κB

continued

Fig. 2.3 continued

D. shRNA: KD Control Time (min): 0 15 30 60 0 15 30 60

Nuclear

E. 35 KD 30 Control 25 20 15

Nuclear p65 10 (Signal Intensity) 5 0 0 15 30 45 60 Time (min)

Fig. 2.3 Effect of PIN1 knockdown on iNOS and VCAM-1 mRNA and NF-κB. KD and control (C) were treated with LPS/IFN-γ. Total RNA or nuclear protein was

isolated at the indicated times. mRNAs encoding iNOS, VCAM-1, and β-actin were determined by reverse transcription-polymerase chain reaction and agarose

gel electrophoresis. A: representative images of iNOS and β-actin products in ethidium-stained gels. Quantitation of iNOS ( B) and VCAM-1 ( C) mRNA-to-β-

actin signal ratios. Bars represent means + SE ratio of iNOS to β-actin mRNA

from densitometric analysis of images from three independent cultures. *p < 0.05

for comparison with 0 h. +p < 0.05 for comparison between KD and control

shRNA. Nuclear NF-κB p65 was assessed by Western blotting ( D) and (E).

Knockdown of PIN1 increases nitric oxide production and causes NO synthase-

dependent cytotoxicity

The Griess reaction demonstrated a low basal level of nitrite in medium of all unstimulated MAEC. LPS/IFN-γ caused significant elevation of nitrite in the medium in both KD and control shRNA cells after 48 h. However, the nitrite level was significantly higher in KD when compared with that of control shRNA MAEC

(Fig. 2.4A). To determine the functional significance of the enhanced iNOS levels,

KD and control shRNA cells were treated with 1 mM L-NAME, a general NOS inhibitor, and with LPS/IFN-γ. Cytotoxicity was measured by crystal violet staining

of adherent cells remaining in culture after 48 h and 72h. LPS/IFN-γ caused no significant detachment of MAEC harboring the control construct that does not suppress PIN1 (Fig. 2.4B/C). In contrast, LPS/IFN-γ caused significant loss of KD

cells, which was prevented by L-NAME. Thus depletion of PIN1 enhanced the

induction of iNOS by LPS/IFN-γ and promoted NO synthase-dependent

cytotoxicity. Interestingly, DETA NONOate, an NO donor, induced similar

detachment in both KD and control shRNA MAEC only at high millimolar

concentrations (Fig. 2.4D), suggesting other factors were required in LPS/IFN-γ

induced cell detachment. Superoxide anion can interact with NO to form more

toxic agents. PIN1 KD contained significantly higher level of superoxide anion

than that in control shRNA MAEC (Fig. 2.4E), indicating its possible involvement

in cell detachment in response to LPS/IFN-γ.

60 KD * Control 50 + 40 * 30

20 + [Nitrite] (µM) * 10 *

0 Vehicle L/I L/I + L-NAME Treatment

KD + + 150 Control * *

100 * 50 (% of Vehicle) Adherent Cells Adherent

0 Vehicle L/I L/I + L-NAME Treatment

Fig. 2.4 Effect of PIN1 knockdown on NO accumulation and cytotoxicity

continued

Fig. 2.4 continued

KD Control 125

100

50 (% of Vehicle) Adherent Cells Adherent 25 +

0 * Vehicle L/I L/I + L-NAME Treatment

120 KD 90 Control

(% of Vehicle) 30 Adherent Adherent Cells

0 0 250 500 750 1000 DETA NONOate ( µM)

continued

Fig. 2.4 continued

0.3 KD Control * 0.2 + *

0.1 NBT signal NBT

0.0 Vehicle L/I

Fig. 2.4 Effect of PIN1 knockdown on NO accumulation and cytotoxicity. A: KD

and control MAEC were incubated in Dulbecco's minimum essential medium-

0.5% fetal bovine serum overnight and treated with vehicle or 10 µg LPS/ml and

20 ng IFN-γ /ml (L/I) for 48 h. The concentration of nitrite in cell culture

supernatant was measured by the Griess reaction. Bars represent means + SE

of 7 separate determinations. *p < 0.05 for comparison between vehicle and L/I-

treated cells. +p < 0.05 for comparison between KD and control shRNA cells.

PIN1 KD and control shRNA MAEC were pretreated without or with 1 mM L-

NAME for 1 h and then treated vehicle or LPS/IFN-γ for 48 h (B) and 72h (C), or

treated with an NO donor, DETA NONOate, at various concentrations for 72h (D).

After crystal violet staining was completed, the percent viability, with vehicle set

to 100%, was determined. E. PIN1 KD and control shRNA MAEC were treated

with vehicle or LPS/IFN-γ for 23h and were culturled in media containing NBT

(0.25 mg/ml) for 1h. Superoxide anion generation was measured by NBT

reduction assay as described in Material and Methods. Bars represent means +

SE of of 6–8 separate determinations. *p < 0.05 for comparison between vehicle

and L/I-treated cells. +p < 0.05 for comparison between KD and control shRNA

cells.

Knockdown of PIN1 stabilizes iNOS

Since PIN1 knockdown did not affect the induction of iNOS mRNA, degradation of iNOS was investigated. KD and control shRNA cells were treated with 90 µg

CHX/ml 24 h after induction with LPS/IFN-γ to block translation. Proteins were

extracted 0, 8, and 24 h after the addition of CHX and subjected to Western blot

analysis. The level of iNOS fell to less than 50% of the initial value by 8 h after

CHX in the control shRNA cells. In KD cells, a similar decrease in iNOS was

delayed until 24 h after CHX treatment (Fig. 2.5). The half-life of iNOS protein

was about 7 h in control shRNA cells, but it was prolonged to about 24 h in PIN1

KD MAEC.

A shRNA: KD Control Time (h): 0 8 24 0 8 24

iNOS

α-Tubulin

KD 100 + Control 80 + 60

40 -tubulin Signal Ratio α α α α 20 (% of initial value) 0 iNOS/ 0 8 16 24 Time (h)

Fig 2.5 Effect of PIN1 knockdown on iNOS protein stability. KD and control shRNA MAEC were treated with LPS and IFN-γ for 24 h. The protein synthesis

inhibitor cycloheximide (CHX) 90 µg/ml was added, and cell extracts were collected at the indicated times. A: iNOS and α-tubulin were measured by

Western blot analysis. Representative blots are shown. B: iNOS and α-tubulin levels in each sample were determined by image analysis of Western blots.

Ratios of iNOS to α-tubulin were determined, and the average ratio ± SE of 4 independent cultures for each point, as a percentage of the value at 0 h after

CHX treatment, was calculated. The dashed line marks the 50% value. +p < 0.05 for comparison between KD and control shRNA at the indicated time.

Calpain inhibition stabilizes iNOS in MAEC

Others found that calpain degraded iNOS in RAW 264.7 cells and in vitro (137 ).

Here, KD and control shRNA MAEC were treated with vehicle or the calpain inhibitor zVF (25 µM) for 1 h and then stimulated with LPS/IFN-γ for 24 h. zVF

significantly increased induction of iNOS by LPS/IFN-γ in KD and control shRNA cells. Induction of iNOS was again greater in KD when compared with control shRNA cells (Fig. 2.6, A and B). The effect of zVF on degradation of iNOS was next examined by treating control shRNA MAEC with CHX 24 h after induction with LPS/IFN-γ. The drop in iNOS seen after 8 h in vehicle-treated cells was

prevented by zVF. The value only decreased to 59% of the starting value 24 h

after CHX in cells exposed to zVF compared with 14% in cells incubated without

the calpain inhibitor (Fig. 2.6C).

Vehicle zVF shRNA: KD Control KD Control

iNOS

α-Tubulin

Fig. 2.6 Effect of calpain inhibition on iNOS continued

Fig. 2.6 continued

Vehicle * zVF 5 + 4 *

2 + -tubulin Signal Ratio Signal -tubulin α α α α 1

0 iNOS/ KD Control shRNA

120 Vehicle zVF 100 80

60 40 -tubulin Signal Ratio Signal -tubulin + α α α α *

(% of Value at 0 h) 0 at Value of (% 20

iNOS/ 0 0 8 16 24 Time (h)

Fig. 2.6 Effect of calpain inhibition on iNOS. A: PIN1 KD and control shRNA

MAEC were pretreated with vehicle or zVF (25 µM) for 1 h and then treated with

LPS and IFN-γ for 24 h. Representative Western blot analyses of iNOS are shown. B: densitometric analysis of signal intensities of images from three samples per group for the ratio of iNOS to α-tubulin. Bars represent the means +

SE of three independent cultures per group. *p < 0.05 for comparison between vehicle and zVF. +p < 0.05 for comparison between KD and control shRNA. C: control shRNA MAEC were treated as in A and then treated with CHX (90 µg/ml).

Cell extracts were collected at the indicated times after CHX and analyzed by

Western blot analysis for iNOS and α-tubulin. Ratios of iNOS to α-tubulin were

determined. The mean percentage of the value at 0 h after CHX treatment ± SE

of 2–4 independent cultures per group is shown. The dashed line marks the 50%

value. +p = 0.018 and *p = 0.066 for one-tail comparison between vehicle and

zVF at the indicated time.

The proteasome does not contribute to iNOS degradation in MAEC

The ubiquitin-proteasome pathway can target iNOS for degradation, and it affects iNOS transcription by degrading inhibitor of κB (I κB) α/β. The latter effect

increases levels of active NF-κB, which can induce iNOS mRNA (131 ). When KD

and control shRNA MAEC were initially treated with 2 µM epoxomicin, a selective

proteasome inhibitor (261 ), for 1 h and then stimulated with LPS/IFN-γ, induction

of iNOS was prevented (Fig. 2.7A). As expected the blockage of iNOS induction

by epoxomicin was associated with increased levels of I κBα and β in LPS/IFN-γ treated cells (Fig. 2.7B). I κBα/β levels were similar in KD and control shRNA

MAEC. MAEC were then treated with LPS/IFN-γ for 24 h to induce iNOS, followed with epoxomicin and 90 µg CHX/ml for 0, 8, or 24 h. Epoxomicin had no effect on the loss of the induced iNOS protein over time after CHX in control shRNA MAEC (Fig. 2.7C).

A. Vehicle EPO shRNA: KD Control KD Control

iNOS

α-Tubulin

Fig. 2.7 Effect of proteasome inhibition on iNOS. continued

Fig. 2.7 continued

B shRNA: KD Control Vehicle EPO L/I L/I+EPO Vehicle EPO L/I L/I+EPO IκB-α

IκB-ß

α -Tubulin

120 Vehicle 100 EPO

60 40 -tubulin Ratio Signal α α α α

(% at of 0 h) Value 20

0 iNOS/ 0 8 16 24 Time (h)

Fig. 2.7 Effect of proteasome inhibition on iNOS. A: PIN1 KD and control MAEC

were pretreated with vehicle or epoxomicin (EPO, 2 µM) for 1 h and then treated

with LPS and IFN-γ for 24 h. Representative Western blot analyses of iNOS and

α-tubulin in KD and control shRNA. B: MAEC were pretreated with vehicle or

epoxomicin for 1 h and then without or with LPS and IFN-γ (L/I) for 15 min.

Extracts were analyzed by Western blot analysis using anti-IκBα/β antibody. C: control shRNA MAEC were treated LPS and IFN-γ for 24 h, then with epoxomicin for 1 h, and finally with CHX (90 µg/ml). Cell extracts were collected at the indicated times after CHX and analyzed by Western blot analysis for iNOS and α

-tubulin. Ratios of iNOS to α-tubulin were determined. The mean percentage of

the value at 0 h after CHX treatment ± SE of 2–4 independent cultures per group

is shown.

PIN1 interacts with iNOS

The potential for interaction between iNOS and PIN1 was assessed. iNOS was

coimmunoprecipitated with PIN1 in control shRNA MAEC treated with LPS/IFN-γ

(Fig. 2.8A). To investigate requirements for PIN1 domains in interaction with

iNOS, a WW domain mutant, W33A, which does not bind p(S/T)-P sites in

substrates, and a peptidyl-prolyl isomerase catalytic mutant, K64A, were

generated. Samples were incubated with glutathione-sepharose and GST, GST-

PIN1, GST-W33A PIN1, or GST-K64A PIN1. GST failed to bind iNOS, but iNOS

was pulled down by GST-PIN1. GST-W33A PIN1 interacted poorly with iNOS, whereas the K64A catalytic mutant pulled down a detectable level of iNOS (Fig.

2.8B).

A. IP : None PIN1 Blot - iNOS

- PIN1

GST W33A PIN1 K64A

- iNOS

- GST-Fusion

- GST

Fig. 2.8. PIN1 interacts with iNOS. A: control shRNA MAEC were treated with

LPS/IFN-γ for 24 h. Cell extracts were immunoprecipitated without (None) or with

anti-PIN1 antibody (PIN1) and analyzed by Western blot analysis with anti-iNOS antibody. B: cell extracts were incubated with GST or GST-wild-type PIN1, -

W33A or -K64A fusion proteins, and glutathione sepharose, and analyzed by

Western blotting with anti-iNOS and anti-GST antibodies. Blots are

representative of 3 independent cultures.

DISCUSSION

The role of PIN1 in the inflammatory response of endothelial cells has not been

described. PIN1 is known to increase the level and/or activity of several transcription factors that mediate induction of iNOS (18, 30, 32, 72, 80, 107 ). It

was expected that suppression of PIN1 would limit the induction of iNOS by

LPS/IFN-γ in MAEC. However, depletion of PIN1 by 85% increased induction of

iNOS protein, whereas increases in iNOS mRNA caused by LPS/IFN-γ were not

affected (Fig. 2.3, A and B). Experiments in cytokine-treated PIN1 knockout

murine embryonic fibroblasts and in unstimulated siRNA-treated cancer cells indicated that NF-κB activation of a reporter gene was reduced. In cytokine-

stimulated knockout embryonic fibroblasts, NF-κB activity was impaired, probably as a result of increased proteasomal degradation of p65 in the complete absence

of PIN1 (18 ). Partial suppression of PIN1 in MAEC here did not affect the peak level of nuclear NF-κB p65, induction of endogenous iNOS mRNA, or of another p65-regulated mRNA, VCAM-1, in MAEC (Fig. 2.3). We speculate that different transcription factors and proteins may be significantly affected at different levels

of PIN1, and the level of PIN1 required for expression, function, and turnover of

different proteins may vary with cell type and stimulus. The results here

demonstrate that PIN1 depletion with shRNA increased iNOS by a posttranscriptional mechanism in LPS/IFN-γ -treated MAEC.

Consistent with the increased level of iNOS, knockdown of PIN1 increased the accumulation of nitrite in the medium (Fig. 2.4A). The increases in NO production and/or iNOS were functionally significant since LPS/IFN-γ caused cell injury in

KD MAEC, which was antagonized by a NO synthase inhibitor L-NAME (Fig.

2.4B/C). The loss of cell adherence is a delayed response, since none was seen at 24 h and was greater after 48 and 72 h. DETA NONOate, an NO donor,

induced detachment only at high millimolar concentrations, and KD and control

shRNA cells were equally susceptible (Fig. 2.4D). This suggests that additional

factors are required for detachment induced by LPS/IFN-γ. For example, NO from

iNOS may combine with superoxide from iNOS or other sources, or with other molecules, generating increased levels of agents more toxic than NO in KD cells

(262-265 ). In fact, along with NO, superoxide was increased more in KD than

control shRNA cells (Fig. 2.4E). Whereas other mediators of cell detachment and

their sources remain to be determined, the results indicate that PIN1

suppression increased the induction of iNOS and NO synthase-dependent effects in MAEC.

Calpain and proteasomes degrade iNOS in various cell types (131, 134 ). Here,

depletion of PIN1 stabilized iNOS protein in CHX-treated MAEC (Fig. 2.5).

Furthermore, the calpain inhibitor zVF enhanced the induction of iNOS by

LPS/IFN-γ and antagonized the loss of iNOS after inhibition of protein synthesis

(Fig. 2.6). In contrast, the selective proteasome inhibitor epoxomicin (261 ) did not

raise iNOS levels or slow its loss after CHX. Epoxomicin at 2 µM was effective

against proteasome activity, since acute loss of IκB α/β caused by LPS/IFN-γ

and induction of iNOS was partially blocked when it was applied before LPS/IFN-

γ (Fig. 2.7). The degree of prevention of the loss of I κB, however, was apparently sufficient to prevent iNOS induction in epoxomicin-treated cells. The overall

results indicate that depletion of PIN1 inhibited iNOS degradation and that degradation was mainly due to calpain, as opposed to proteasomal activity, in

MAEC treated with LPS/IFN-γ.

iNOS was ubiquitinated in A549 and bronchial epithelial cells treated with

cytokines, in RAW 264.7 cells treated with LPS-IFN-γ, and in HEK293 cells transfected with the human enzyme (131, 132 ). Degradation of iNOS was

antagonized by proteasome inhibitors in HEK293 cells transfected with iNOS,

and in RAW 264.7 cells and RT4 epithelial cells treated with LPS or a cytokine mixture for 48 h (131, 132 ). On the other hand, a glucocorticoid agonist reduced iNOS levels in IFN-γ -treated RAW 264.7 cells via calpain activation (134 ).

Interestingly, exogenous treatment of HEK293 cells with the endogenous calpain

inhibitor protein CAST (206 ) did not affect degradation of transfected iNOS (131 ).

This suggests that proteasomes, but not calpain, contributed to turnover of

transfected iNOS in HEK293 cells. Thus cell type, stimulus, level, or duration of induction, and other factors may affect the iNOS turnover path that operates in different systems.

In vitro proteolysis of iNOS by calpain I is modulated by protein interactions and conformation. Monomers of iNOS were more sensitive to calpain compared with dimers. Degradation was also antagonized by calmodulin, which associates with murine iNOS at a well-characterized site between amino acids 503 and 532 (136,

137 ). Conformation outside this domain is another factor since heating of an iNOS mutant, lacking the calmodulin-binding sequence, reduced its susceptibility

to calpain I, whereas degradation of wild-type iNOS was insensitive to heat

treatment (137 ). PIN1 may regulate iNOS conformation or protein interactions

that affect degradation by calpain.

PIN1 might affect iNOS turnover by association with the enzyme, which is

phosphorylated (129 ), and contains several S/T-P motifs that could be WW ligands. iNOS was coimmunoprecipitated with PIN1 using an anti-PIN1 antibody

(Fig. 2.8A). In vitro pulldowns with GST-PIN1 and domain mutants indicated that the WW domain was critical, whereas the K64A catalytic mutant was able to precipitate detectable iNOS (Fig. 2.8B). Interaction between PIN1 and iNOS

could be required for effects of PIN1 on iNOS turnover. However, PIN1 could

mediate, or regulate, the association of iNOS with itself or other cofactors, like

calmodulin or heat shock protein 90 (137, 266 ). PIN1 could also stimulate calpain or reduce inhibition by endogenous CAST (206 ). These possibilities are under investigation.

PIN1 is known to affect the expression and function of its various substrates by

many mechanisms (236 ). The range of mechanisms include regulation of

transcription, mRNA turnover, enzyme activity, sensitivity to phosphatases, and unbiquitination (20, 21, 29, 30, 175, 267 ). The findings presented here suggest that PIN1 significantly affects iNOS protein turnover in MAEC that apparently rely

on calpain for its degradation. Thus PIN1 may play a significant role in NO synthase-mediated disease or drug actions depending on cell type and stimulus.

In conclusion, suppression of PIN1 increased the induction of iNOS without

affecting the mRNA. Instead, PIN1 knockdown antagonized the degradation of

iNOS in MAEC, which was sensitive to a calpain inhibitor but not a proteasome inhibitor. WW interactions with p(Ser/Thr)-Pro motifs and proline isomerization

may control the sensitivity of iNOS to calpain or the function of the calpain/CAST system in MAEC.

CHAPTER 3

PIN1 REGULATES DEGRADATION OF COX-2

IN ENDOTHELIAL CELLS

Abstract

The peptidyl-proline isomerase, Protein Never in Mitosis Gene A Interacting-1

(PIN1), regulates turnover of inducible nitric oxide synthase (iNOS) in murine aortic endothelial cells (MAEC) stimulated with E. coli endotoxin (LPS) and interferon-γ (IFN-γ). Degradation of iNOS was reduced by a calpain inhibitor,

suggesting that PIN1 may affect induction of other calpain-sensitive inflammatory

proteins, such as cyclooxygenase (COX)-2, in MAEC. MAEC, transduced with

lentivirus encoding an inactive control short hairpin (sh) RNA or one targeting

PIN1 that reduced PIN1 by 85%, were used. Cells were treated with LPS/IFN-γ, calpain inhibitor (carbobenzoxy-valinyl-phenylalaninal, zVF), cycloheximide and

COX inhibitors to determine the effect of PIN1 depletion on COX-2. LPS or IFN-γ alone did not induce COX-2. However, treatment with 10 µg LPS plus 20 ng IFN-

γ per ml induced the COX-2 signal on western blots 10-fold in control shRNA

MAEC. Induction was significantly greater (47-fold) in PIN1 shRNA cells. COX-2- dependent prostaglandin E2 production was also 3 times greater in cells lacking

PIN1 compared to the control cells. The additional increase in COX-2 protein due

100

to PIN1 depletion was post-transcriptional, as induction of COX-2 mRNA by

LPS/IFN-γ was the same in cells containing or lacking PIN1. Instead, the loss of

COX-2 protein, after treatment with cycloheximide to block protein synthesis, was reduced in cells lacking PIN1 in comparison with control cells, indicating that degradation of the enzyme was reduced. zVF also enhanced the induction of

COX-2 by LPS/IFN-γ, and slowed its loss in after treatment with cycloheximide.

In contrast to iNOS, physical interaction between COX-2 and PIN1 was not

detected, suggesting that PIN1 may regulate COX-2 degradation by affecting

calpain, rather than by an effect on COX-2 protein. The results suggest that PIN1

can limit COX-2 induction by facilitating calpain-mediated degradation in MAEC.

Introduction

Protein Never in Mitosis Gene A Interacting-1 (PIN1) is an enzyme that regulates transcription, and turnover of mRNA and proteins. PIN1 is a cis-trans peptidyl- prolyl isomerase that binds to phosphorylated serine- or threonine-proline sequences in substrate proteins via an amino-terminal WW domain, and isomerizes the site with its carboxyl-terminal catalytic domain (8). Isomerization of the phosphorylated serine- or threonine-proline motif has a significant effect on

101

conformation of many phospho-proteins. The conformational switching catalyzed by PIN1 allows it to regulate transcription factors, mRNA stabilization factors, and the susceptibility of a growing list of proteins to post-translational modifications and proteases (8, 18, 20, 21, 29, 30, 268 ). Previously, we found that depletion of

PIN1 and treatment with a calpain inhibitor each reduced the degradation of iNOS in MAEC stimulated with E. coli endotoxin LPS and IFN-γ. PIN1 bound to

iNOS suggesting that it might directly regulate the sensitivity of iNOS to calpain

(135 ). PIN1 may also regulate expression of inflammatory proteins by an effect on calpain.

COX-2 is induced by LPS, IFN-γ, and other factors in endothelial cells cultured from various organs and species (141, 269-275 ). Elevated endothelial COX-2 may contribute to vascular pathogenesis (276, 277 ). This enzyme is also significant for endotoxin action as COX-2 knockout mice are resistant to LPS- induced inflammation and death (278 ). COX-2 has a relatively short half-life, indicating that turnover may effectively control its expression (141 ). While COX-2 and iNOS can be degraded by several processes, calpain inhibitors are known to suppress their cleavage (135, 141, 163, 170, 279 ). Thus, the purpose of this

investigation was to determine whether PIN1 regulates the expression of COX-2,

which is induced by LPS and IFN-γ in MAEC. It was hypothesized that PIN1

102

would associate with COX-2 and that depletion of PIN1 would enhance its induction in MAEC. The impact of PIN1 depletion on calpain activity was also determined.

Materials and Methods

Endothelial cell growth supplement, heparin, phenylmethylsulfonyl fluoride,

Bradford reagent, E. coli LPS, serotype 0111:B4, arachidonic acid and polybrene were obtained from Sigma Chemical Co. (St Louis, MO). Recombinant mouse

IFN-γ was from R&D Systems (Minneapolis, MN). Cycloheximide, carbobenzoxy- valinyl-phenylalaninal (zVF, MDL-28170 or calpain inhibitor III), PD150606, porcine µ-calpain and carboxybenzyl-phenylalanine-arginine-7-amido-4-

methylcoumarin were obtained from Calbiochem (La Jolla, CA). Fetal bovine

serum was from Hyclone Laboratories (Logan, UT). Agarose, ethidium bromide,

ethylenediamine tetraacetic acid, sodium dodecyl sulfate, NaCl, Na3VO4, NaF,

tris-base and tween 20 were obtained from Fisher Scientific (Fair Lawn, NJ).

Triton X-100 was from Pierce (Rockford, IL). Dulbecco’s minimum essential

medium, trypsin, Trizol, Superscript Reverse Transcriptase Taq DNA polymerase,

RNAse-free DNAse, deoxynucleotides, and protein G agarose were purchased

from Invitrogen (Carlsbad, CA). Glutathione-sepharose was purchased from

103

Amersham Biosciences (Uppsala, Sweden). A prostaglandin E2 competition enzyme-linked immunosorbent assay kit was obtained from R and D Systems,

Minneapolis, MN. Anti-COX-2 antibody directed against a 16 amino acid sequence, ending 7 residues from the C-terminus of the protein, SC-560 and NS-

398 were purchased from Cayman Chemical (Ann Arbor, MI). Anti-PIN1 was from R&D Systems (Minneapolis, MN). Horseradish peroxidase-conjugated goat anti-mouse and goat anti-rabbit secondary antibodies were from Jackson immunoresearch Laboratories, Inc. (West Grove, PA). Renaissance Enhanced

Chemiluminescence Reagent was purchased from New England Nuclear Life

Sciences (Boston, MA).

Cells

MAEC were cultured from aortas of mice in accordance with the Guide for the

Care and Use of Laboratory Animals from the U.S. National Institutes of Health

(280 ). As described previously, cells were transduced with short hairpin RNA

(shRNA) to knockdown (KD) PIN1 or with an inactive mutant sequence (control),

and selected for stable modification. This produced KD MAEC with approximately

15% of the level of PIN1 protein found in control and non-transduced MAEC

(135 ).

104

Treatments

KD and control MAEC were incubated in Dulbecco’s minimum essential medium/0.5% fetal bovine serum for 18 h, and then treated with medium or 10 µg

LPS and 20 ng IFN-γ per ml, and other agents for various times. zVF was added

1 h before LPS/IFN-γ to inhibit calpain (281 ). Ninety µg cycloheximide/ml was

used to inhibit protein synthesis after induction of COX-2 with LPS/IFN-γ (135 ).

COX-2-dependent prostaglandin E2 production was measured after incubating cells with LPS/IFN-γ for 24 h. Cells were then incubated in fresh medium containing 20 µM arachidonic acid, LPS/IFN-γ and the COX-1 selective

antagonist, SC-560 (1 µM) (282 ), with or without the COX-2 selective antagonist,

NS-398 (10 µM) (283 ). The medium was collected after 2 h and stored at -80 degrees C. Prostaglandin E2 was measured by competition enzyme-linked immunosorbent assay in comparison with prostaglandin E2 standard by the manufacturer’s instructions.

mRNA levels

RNA was extracted with Trizol, precipitated, and dissolved in water. cDNA was produced from 3 µg of RNA. cDNA was amplified by polymerase chain reaction

for ß-actin as described previously (284 ), and for COX-2. COX-2 primers were

105

sense, 5’-CCG GAC TGG ATT CTA TGG TG, and antisense, 5’-AGG AGA GGT

TGG AGA AGG CT from Genbank accession BC052900, producing a 263 base pair product. Half of each reaction was electrophoresed in 1% agarose. Gels were imaged and analyzed after ethidium bromide staining.

In vitro cleavage by calpain.

After culture, KD and control cells were treated with LPS/IFN-γ for 24h, then were

scraped in calpain assay buffer (50 mmol/L Hepes buffer (pH 7.4), 150 mmol/L

NaCl, 1 mmol/L EDTA, and 1% Triton X-100). Cell lysates were kept in ice for

20min and clarified by centrifugation at 16000g for 20min. In vitro cleavage by

calpain was performed by incubating 44ul of cell soluble fraction with various

amounts of purified porcine µ-calpain and CaCl2 (5mM) at 30 degree for 1h. The reaction was stopped by adding 2 × SDS sample buffer. COX-2 was detected by

western blotting with anti-COX-2 antibody. COX-2 signal intensity was calculated

as a percent of the value with no added calpain are shown.

Immunoprecipitation, Glutathione S-transferase Pulldown, and Western Blotting

As previously described (135), cells were washed, sonicated in lysis buffer, and

protein concentration was measured. For western blotting, 12 µg of sample protein were denatured and separated on 4-20% Trisgylcine, SDS-

106

polyacrylamide gels and transferred to nitrocellulose. For immunoprecipitation,

500 µg of cell lysate protein was incubated with 5 µg anti-PIN1 antibody and

protein G agarose. For pulldown, glutathione S-transferase or glutathione S-

transferase-PIN1 fusion protein was added to 500 µg of cell lysate protein and

glutathione-sepharose. Samples were then denatured for electrophoresis and

western blotting. Blots were immunostained and imaged on X-ray film by

enhanced chemiluminescence. Films were scanned and digital images of

proteins were analyzed.

Cathepsin activity assay

Confluent cells were washed three times with PBS and scraped in 1ml of ice-cold

PBS. Cells were collected by centrifugation at 1500 x g for 2 min at 4 °C.

Collected cells were resuspended in a reaction buffer (50 mM Na-acetate, 1 mM

EDTA and 2 mM dithioerythritol pH 5.5), and were sonicated four times for 10 s

with 1-min breaks. After sonication, the lysate was centrifuged at 1500 x g for 5

min at 4 °C to remove the cell debris, and the supernatant was used for

cathepsin activity measurements. The reaction was started by mixing 20 µM

cathepsin substrate carboxybenzyl-phenylalanine-arginine-7-amido-4-

methylcoumarin, in the reaction containing 2 µg supernatant. 7-amido-4-

methylcoumarin release was monitored at 37°C for 30 min by fluorescence, with

107

excitation at 380 nm and emission at 460 nm, and the initial velocity was determined (285, 286 ).

Data Analysis

Bands in images of polymerase chain reaction gels and scanned western blots were measured withImage J 1.34s (NIH). Prostaglandin E2 concentrations were estimated from a standard curve and calpain activity was indicated by the fluorescence increase per minute. Data were analyzed by Student’s t test or analysis of variance with Bonferroni correction for multiple comparisons (287 ).

108

Results

PIN1 suppression or genetic deletion increases COX-2 induction in response to

LPS and IFN -γ.

First, COX-2 induction in control MAEC was tested when exposed to vehicle,

LPS (10 µg/mL), IFN-γ (20 ng/mL) and combination of LPS/IFN-γ for 24h.

Similarly to iNOS, COX-2 protein was very low in vehicle-treated control MAEC

and incubation with either LPS or IFN-γ alone did not induce it. However, stimulation with 10 µg LPS plus 20 ng IFN-γ per ml for 24 h dramatically increased induction of COX-2 protein in control shRNA MAEC (Fig. 3.1A). COX-

2 was induced more in PIN1 KD than in control shRNA cells 24 h after treatment with LPS/IFN-γ (Fig. 3.1B). However, no significant difference of COX-1 signal

was detected in either vehicle or LPS/IFN-γ treatment (Fig. 3.1B). Genetic

deletion of PIN1 also increased the induction of COX-2 (Fig. 3.1D). Therefore,

induction of COX-2 protein was increased by both PIN1 suppression and deletion.

COX-2- mediated prostaglandin E2 production increased 3-fold in KD MAEC, but

not in the control cells (Fig 3.1E).

109

Vehicle LPS IFN-γ L/I

COX2

α-Tubulin

Vehicle L/I KD C KD C

COX2

COX1

α-Tubulin

Fig. 3.1 Effect of PIN1 knockdown on COX-2 protein and activity

continued

110

Fig. 3.1 continued

8 KD C 5 KD 6 C 4 -Tubulin 4 -Tubulin 3 α α α α α α α α + 2 2 COX-2/ COX-1/ 1 signal intensityratio signal intensity ratio 0 0 Vehicle L/I Vehicle L/I

ko wt 1.5

COX-2 1.0 -Tubulin α α α α α-Tubulin 0.5

COX2/ + Signal Intensity Ratio 0.0 KO WT

continued

111

Fig. 3.1 continued

300 + NS-398 ( µM) 0 200 10

100 [PGE2] (pg/mL)

0 KD C

Fig. 3.1 Effect of PIN1 knockdown on COX-2 protein and activity. A. control (C)

shRNA MAEC were treated with vehicle (culture medium), LPS, IFN-γ or LPS

and IFN-γ (L/I) for 24 h. Representative western blots and analysis of COX-2 expression are shown. B. KD and control (C) shRNA MAEC were treated with

LPS and IFN-γ (L/I) for 24 h and representative western blots of COX-2 and

COX-1 expression are shown in B and analysis of COX-2 and COX-1 expression

are shown in C. D. MAEC from PIN1 knockout (ko) and wildtype (wt) mice were

treated with LPS and IFN-γ (L/I) for 24 h. In C and D, bars represent mean + SE

112

ratio of COX-2/ α-tubulin from densitometric analysis of 3 cultures of each group.

+, p<0.05 for comparison between KD and control cells or between knockout and

wildtype cells treated in the same manner. E. COX-2-dependent prostaglandin

E2 production was measured in cells treated with LPS/IFN-γ, 20 µM arachidonic

acid and 1 µM SC-560, with or without 10 µM NS-398. Bars represent mean +

SE concentration of prostaglandin E2 in medium for 5 cultures in each group. +,

p<0.05 for between KD and control cells treated in the same way.

Knockdown of PIN1 does not influence COX-2 mRNA

LPS/IFN-γ induced COX-2 mRNA in KD and control shRNA cells were measured

by RT-PCR. Despite the increases in iNOS protein, there was no difference in

COX-2 mRNA between KD and control shRNA cells at various periods after treatment with LPS/IFN-γ (Fig. 3.2A/B), suggesting post-transcriptional regulation of COX-2 by PIN1 in MAEC.

113

Time (h): 0 1 4 8 shRNA: KD C KD C KD C KD C COX -2

ß-actin

KD 1.5 * * Control * * * 1.0 *

0.5

COX-2/ß-actin mRNA Signal Intensity Ratio 0.0 0 1 4 8 Time (h)

Fig. 3.2 Effect of PIN1 knockdown on COX-2 mRNA. KD and control (C) shRNA

MAEC were treated with LPS and IFN-γ (L/I) for 0-8 h. mRNAs encoding COX-2 and ß-actin were determined by RTPCR and agarose gel electrophoresis. A representative image of COX-2 and ß-actin PCR products from a single ethidium-

114

stained gel is shown. Bars represent mean + S.E. ratio of COX-2: ß-actin products from densitometric analysis of images from 3 independent cultures run in a single gel. *:p<0.05 for comparison with cells treated for 0 h.

Knockdown of PIN1 stabilizes COX-2

Since PIN1 knockdown did not affect the induction of COX-2 mRNA, the effect of

PIN1 depletion on degradation of COX-2 after induction with LPS/IFN-γ was

assessed. Cells were induced with LPS/IFN-γ and then treated with 90 µg

cycloheximide/ml to block translation. Proteins were extracted 0, 2, and 4 h after the addition of CHX and subjected to Western blot analysis. The level of 72 kDa

COX-2 protein fell to less than 50% of the initial value by 2 h after addition of

cycloheximide to control shRNA cells. A decrease in COX-2 by approximately

50% was delayed until 4 h after CHX treatment in KD MAEC (Fig. 3.3)

115

shRNA: KD Control Time (h): 0 2 4 0 2 4

COX2

α-Tubulin

100 KD Control 80

-tubulin Ratio Signal + α α α α 20 (% at of 0 Value h) +

0 COX2/ 0 1 2 3 4 Time (h)

Fig. 3.3 Effect of PIN1 knockdown on COX-2 stability. KD and Control shRNA

MAEC were treated with with LPS and IFN-γ for 24 h. Cycloheximide (90 µg/ml) was added, and cell extracts were collected at the indicated times and western blotted for COX-2 and α-tubulin. The average COX-2/ α-tubulin signal intensity ratio + SE of 4 independent cultures for each point, as a percent of the value at 0

116

h after cycloheximide treatment is shown. The dashed line marks the 50% value.

+, p<0.05 for comparison between similarly treated KD and control cells at the indicated time.

Calpain inhibition stabilizes COX-2 in MAEC

Similarly to iNOS, COX-2 is also known as a substrate of calpains (170, 171 ). In

human synovial fibroblasts, cleavage of COX-2 was reduced by E64d indicating

a role for calpain and/or cathepsin. Thus, effects of calpain inhibition on COX-2

degradation were measured. PIN1 KD and control shRNA MAEC were

pretreated with vehicle and different doses of a calpain inhibitor, zVF for 1 h, and

then treated with LPS and IFN-γ for 24 h. zVF increased the induction of the 72 kDa COX-2 protein in both KD and control cells in a dose dependent manner (Fig.

3.4 A). Induction of COX-2 was greater in KD when compared with control shRNA cells (Fig. 3.4B). Similar results were also observed by the pretreatment of a calpain/cathepsin inhibitor, E64d (Fig. 3.4C). Calpain can regulate certain gene transcriptions, for example, eNOS in pulmonary artery endothelial cells

(288 ), which may be due to its action on transcription factor substrates, such as

β-catenin, c-Fos, c-Jun and p53. Thus, the effect of calpain inhibition on COX-2

117

transcription was examined here. Such an increased induction of COX-2 by calpain inhibition was not due to transcription, since zVF pretreatment did not affect induction of COX-2 mRNA in the MAEC (Fig. 3.4D). The effect of zVF on degradation of COX-2 was next examined by treating control shRNA MAEC with

CHX 24 h after induction with LPS-IFN-γ. zVF reduced the degradation of COX-2

in control MAEC by an amount similar to PIN1 KD shRNA. COX-2 protein

decreased to 63% of its initial value after 4 h in zVF-treated control MAEC, and to

61% in zVF-treated KD MAEC (Fig. 3.4E).

shRNA: KD Control Z-VF ( µM) : 0 3 6 12 25 0 3 6 12 25

COX-2

α-Tubulin

Fig. 3.4. Effect of calpain inhibition on COX-2 protein continued

118

Fig. 3.4 continued

shRNA: KD Control Treatment: Vehicle zVF Vehicle zVF

COX-2

α-Tubulin

Vehicle 3 * zVF +

2 * -Tubulin Protein -Tubulin α α α α 1 Signal Intensity Ratio Intensity Signal

COX2/ +

0 KD Control

KD Control Vehicle E64d Vehicle E64d

COX-2

α-Tubulin

continued

119

Fig. 3.4 continued

Treatment: Vehicle zVF shRNA: KD C KD C

COX-2

ß-Actin

1.2 KD Control 1.0

0.8

0.6 -actin mRNA -actin β β β β 0.4

0.2 COX-2/ Signal Intensity Ratio Intensity Signal 0.0 Vehicle zVF

Continued

120

Fig. 3.4 continued

shRNA: KD Control Time (h): 0 2 4 0 2 4

COX2

α-Tubulin

100 KD Control 80

60 +

(% ofat 0h) Value 20 -tubulin Intensity Signal Ratio α α α α 0 0 1 2 3 4

COX2/ Time (h)

Fig. 3.4 Effect of calpain inhibition on COX-2 protein. A: PIN1 KD and control (C) shRNA MAEC were treated with vehicle, different doses of zVF for 1 h, and then treated with LPS and IFN-γ for 24 h. B. Bars represent mean + SE ratio of COX-2 to α-tubulin from densitometric analysis of images from 3 independent cultures in

121

each group pretreated with vehicle or 25 µM zVF. *: p <0.05 for comparison between vehicle and zVF. +, p <0.05 for comparison between KD and control shRNA. C. Representative western blots of COX-2 and α-tubulin were shown

when KD and control MAEC were treated with vehicle or 6 µM E64d for 1h and

followed by LPS/IFN-γ for 24h. D. Effect of zVF on COX-2 mRNA. KD and control

(C) shRNA MAEC were treated with vehicle (DMSO) or 25 µM zVF for 1 h, then with LPS/IFN-γ for 1 h. mRNA for COX-2 and ß-actin were assessed as in figure

3. Representative agarose electrophoresis of PCR products is shown. Bars

represent the mean + SE of ratio of COX-2/ß-actin signal intensity + SE of 4

independent cultures. E. KD and control shRNA MAEC were treated with 25 µM

zVF for 1 h, then with LPS and IFN-γ for 24 h. Cycloheximide (90 µg/ml) was

added, and cell extracts were collected at the indicated times and western blotted

for COX-2 and α-tubulin. The average COX-2/ α-tubulin signal intensity ratio +

SE of 4 independent cultures for each point, as a percent of the value at 0 h after cycloheximide treatment is shown. The dashed line marks the 50% value.

COX-2 is cleaved by purified porcine µ-calpain in vitro

The above results indicate that calpain contributes the degradation of COX-2 in

MAEC. Thus, purified calpain was used to test whether COX-2 was an in vitro substrate for calpain. The incubation of COX-2 with purified porcine µ-calpain

122

resulted in the loss of COX-2 in a dose dependent manner (Fig. 3.5). COX-2 from control extracts appeared to more susceptible to porcine µ-calpain than that from

KD, which may be due to different PIN1 level in KD and control shRNA cells.

µ-calpain(units): 0 1.6 4.8 8

Control

KD Control 90 + + 60 +

30 COX2 Signal Intensity Signal COX2 0 (% of no added calpain value) calpain (% of added no 0 2 4 6 8 µ-calpain (units)

Fig. 3.5 Degradation of COX-2 by purified porcine µ-calpain in vitro. PIN1 KD and control shRNA MAEC were treated with LPS/IFN-γ for 24 h. Cell extracts from

123

KD and control were collected and incubated with various amounts of purified porcine µ-calpain for 1h in the presence of Ca 2+ (5 mM). COX-2 was detected by western blotting with anti-COX-2 antibody. Representative western blots of

COX-2 and α-tubulin are shown. The average COX-2 signal intensity + SE of 3 independent reactions for each point, as a percent of the value with no added calpain are shown. +, p <0.05 for comparison between KD and C.

Calpain, but not cathepsin, contributes to PIN1 regulated COX-2 degradation in

MAEC

It was known that E64d and zVF also inhibit cathepsins. Therefore, it is essential to determine whether cathepsins contribute to PIN1 regulated COX-2 degradation. A more specific calpain inhibitor, PD150606 is able to increase COX

-2 induction in both KD and control cells in a dose dependent manner (Fig. 3.6A).

Also cathepsin activity was assayed by using flurorescent substrate Z-Phe-Arg-

AMC in KD and control MAEC. No significant difference was observed between

KD and control in either vehicle or LPS/IFN-γ treatment group, suggesting that

PIN1 regualtes COX-2 degradation mainly via calpain but not cathepsins (Fig.

3.6B).

124

shRNA: KD Control PD150606 ( µM) : 0 3 6 12 25 50 0 3 6 12 25 50

COX-2

α-Tubulin

150000

100000 + Initial 50000 Reaction Velocity Reaction

0 KD Control KD Control Vehicle L/I

Fig. 3.6 Cathepsin does not contribute to PIN1-regulated COX-2 degradation.

A. PIN1 KD and control shRNA MAEC were treated with vehicle, different doses

of calpain inhibitor, PD150606, for 1 h, and then treated with LPS and IFN-γ (L/I)

125

for 24 h. Representative western blots of COX-2 and α-tubulin are shown. B.

Effect of PIN1 knockdown on cathepsin activity. Initial rate of cleavage of the fluorogenic calpain substrate was measured in extracts of KD and control cells treated with LPS/IFN-γ for 24 h. Bars represent mean + SE fluorescence increase/min for 4 independent cultures in each group.

PIN1 does not interact with COX-2

Since PIN1 is known to bind its substrate proteins, its interaction with COX-2 was investigated. COX-2 from extracts of LPS/IFN-γ -treated control cells was not

pulled down with glutathione-S-transferase-PIN1 fusion protein or glutathione-S-

transferase. Immunoprecipitation of PIN1 from extracts of LPS/IFN-γ -treated

control cells did not produce any COX-2 detectable on western blots (not shown).

Therefore, PIN1 may regulate COX-2 degradation via affecting calpain activity.

126

• GST GST-PIN1 cell lysate

• COX-2

• GST-PIN1

• GST

Fig. 3.7 PIN1 does not interact with COX-2 in MAEC. Control shRNA MAEC were treated with LPS-IFN-γ for 24 h. Cell extracts were incubated with GST or GST-

wild-type PIN1 fusion proteins, and glutathione sepharose, and analyzed by

Western blotting with anti-COX-2 and anti-GST antibodies.

Proteasome contributes to COX-2 degradation but is not regulated by PIN1 in

MAEC

COX-2 can also undergoes ubiquitination and proteolysis by 26S proteasome

(160-167 ). Pretreatment of a selective proteasome inhibitor, epoxomicin (2 µM)

127

inhibited degradation of I κB α/β, blocked activation of NF-κB and prevent iNOS induction (135 ). Differently from iNOS, COX-2 induction was not prevented but

increased by epoxomicn pretreatment (Fig. 3.8A). Epoxomicin does not affect

COX-2 mRNA level either, suggesting no involvement of NF-κB in COX-2

transcription in MAEC (Fig. 3.8B). Effects of proteasome inhibition on COX-2

degradation was also examined. COX-2 degradation was severely reduced by

epoxomicin in both control and KD MAEC (Fig. 3.8B). After 4 h, the level of COX-

2 was unchanged in control shRNA cells treated with epoxomicin, compared

with a drop to 31% of its initial level in the absence of epoxomicin. COX-2

decreased only to 90% of its initial level in epoxomicin-treated KD MAEC,

compared with the decrease to 47% of the initial value in vehicle-treated KD cells

(Fig. 3.8C).

128

A. shRNA: KD Control Treatment: Vehicle EPO Vehicle EPO COX-2

α-Tubulin

Vehicle 3 * EPO + 2 * -Tubulin Protein -Tubulin α α α α 1 Signal IntensitySignal Ratio

COX2/ + 0 KD Control

Fig. 3.8 Effect of proteasome inhibition on COX-2 mRNA and protein

continued

129

Fig. 3.8 continued

Vehicle EPO KD C KD C COX2

β-Actin

1.2 KD Control 0.9 -actin β β β β 0.6

COX-2/ 0.3 Signal Signal Intensity Ratio 0.0 Vehicle EPO

continued

130

Fig. 3.8 continued

shRNA: KD Control Time (h): 0 2 4 0 2 4

COX2

α-Tubulin

KD 150 Control

100

50 (% of Value at 0h) at (% Value of -tubulin Signal Intensity Ratio Intensity Signal -tubulin α α α α 0 0 1 2 3 4 COX2/ Time (h)

Fig. 3.8 Effect of proteasome inhibition on COX-2 mRNA and protein. A. PIN1 KD and control (C) shRNA MAEC were pretreated with vehicle or a proteasome

131

inhibitor EPO (2 µM) for 1 h, and then treated with LPS and IFN-γ for 24 h. COX-

2 and α-tubulin are shown and analyzed. Bars represent mean + SE ratio of

COX-2 to α-tubulin from densitometric analysis of images from 3 independent cultures in each group. *: p <0.05 for comparison between 0 and 24 h. +, p <0.05 for comparison between KD and control shRNA. B. mRNA for COX-2 and ß- actin were assessed. Representative agarose electrophoresis of PCR products is shown. Bars represent the mean + SE of ratio of COX-2/ß-actin signal intensity

+ SE of 4 independent cultures. C. KD and control shRNA MAEC were treated with 2 µM EPO for 1 h, then with LPS and IFN-γ for 24 h. Cycloheximide (90

µg/ml) was added, and cell extracts were collected at the indicated times and western blotted for COX-2 and α-tubulin. The average COX-2/ α-tubulin signal intensity ratio + SE of 4 independent cultures for each point, as a percent of the value at 0 h after cycloheximide treatment is shown. The dashed line marks the

50% value.

Calpain and proteasome differently contribute COX-2 degradation

Epoxomicin also increased the levels of several bands smaller than 72 kDa COX-

2 in LPS/IFN-γ treated cells (Fig. 3.9, zones II-IV). The pattern of these C- terminal immunoreactive bands was identical in control and KD shRNA cells

132

treated with epoxomicin or the combination of epoxomicin and zVF. zVF or E64d alone caused the accumulation of several C-terminal COX-2 fragments smaller than 72 kDa that were of different sizes than the bands in epoxomicin-treated cells. These bands in zones II and III also appeared identical in vehicle-, zVF- and E64d-treated KD cells. Although lower in intensity, bands visible in zones II and III in control cells treated with zVF or E64d also appeared similar to bands seen in KD cells treated with vehicle-, zVF- or E64d. The results with zVF and epoxomicin, alone and in combination, were replicated in 3 independently treated cultures of KD and control cells for each condition (not shown). Thus, fragments of COX-2 that accumulated in KD cell extracts were qualitatively similar to those from cells treated with calpain inhibitors, and different from those seen after epoxomicin.

133

Fig. 3.9 Effect of PIN1 knockdown, and calpain and proteasome inhibition on

COX-2 fragments. KD and control shRNA MAEC were treated for 24 h with

LPS/IFN-γ and Vehicle, zVF (zVF, 25 µM), E64d (6 or 25 µM), epoxomicin (EPO,

2 µM), or EPO plus zVF. Samples were processed for western blotting for COX-2.

The blots were immunostained and developed together in the same reagents.

The 72 kDa full length COX-2 is marked with an arrow. Regions I, II III and IV are marked for discussion in the text.

134

Discussion

PIN1 regulates the levels and activity of factors that can affect COX-2 synthesis in various cell types(18, 30, 32, 107, 289-292 ). Here, suppression of PIN1 in endothelial cells did not affect COX-1 but increased the induction of COX-2, and

COX-2-dependent production of prostaglandin E2 by LPS/IFN-γ. Despite a

nearly 5-fold greater induction of COX-2 protein in KD compared with control

MAEC, there was no difference in the induction of COX-2 mRNA. This suggests

that PIN1 regulates COX-2 by a post-transcriptional mechanism. Previously, it

was found that PIN1 knockdown reduced calpain-mediated degradation of iNOS

in MAEC. As with iNOS, the loss of 72 kDa COX-2 in cycloheximide-treated

MAEC was reduced by PIN1 depletion, consistent with post-transcriptional action.

Results from cycloheximide experiments showed that the half life of COX-2 was

4h and 2h in KD and control MAEC respectively.

Since COX-2 has a relatively short half-life, inhibition of turnover could lead to

large, cumulative, post-transcriptional increases after induction with LPS/IFN-γ

(141 ). Others found that cleavage of COX-2 was reduced by the calpain inhibitor,

E64d, in human synovial fibroblasts indicating a role for calpain and/or cathepsin

(170 ). In agreement with this, induction of 72 kDa COX-2 in KD and control

135

MAEC was increased by E64d, and by the calpain inhibitor, zVF. E64d and zVF may also inhibit cathepsin, which appears not to contribute PIN1-regulated COX-

2 degradation since no significant difference in cathepsin activity was observed in

KD and control in basal and LPS/IFN-γ stimulated condition. An even more selective calpain inhibitor PD150606 was able to increase the induction of COX-2 efficiently, suggesting the essential role of calpain but not cathepsins in COX-2 degradation. Calpain inhibition also reduced the rate of loss of 72 kDa COX-2 in cycloheximide-treated control cells to a level similar to that in KD MAEC. These results suggest that PIN1 depletion reduces calpain-mediated turnover of COX-2.

Consistent with this idea, zVF and PIN1 KD caused the appearance of similar C- terminal COX-2 fragments smaller than 72 kDa on western blots, using a COX-2 antibody directed to the C-terminal of COX-2. E64d also induced the accumulation of these COX-2 fragments, further suggesting that they are a consequence of inhibiting calpain. However, a simple interpretation is that inhibition of calpain by zVF and E64d shifted processing of COX-2 to enzymes other than calpain, and that depletion of PIN1 had a similar effect.

COX-2 is also a substrate for proteasomes and PIN1 regulates proteasomal degradation of several other proteins (cyclin E, c-myc and p65). Here, inhibition of proteasomal activity with epoxomicin increased induction of COX-2 by

136

LPS/IFN-γ, with no effect on COX-2 mRNA. This contrasts with the strong inhibition of the induction of iNOS by epoxomicin when it was present before

LPS/IFN-γ. The inhibition was associated with prevention of acute loss of

inhibitors of kappa B α and ß (see chapter 2). This suggests that in MAEC, induction of COX-2 is not regulated primarily by inhibitors of kappa B/NFkB.

Epoxomicin severely inhibited the loss of COX-2 after cycloheximide treatment in

KD and control cells, indicating that the proteasome inhibitor increased COX-2 by inhibiting its degradation in MAEC. This contrasts with the lesser reduction of

COX-2 degradation caused by calpain inhibitor or depletion of PIN1. A striking difference between epoxomicin-treated cells and PIN1 KD, zVF- or E64d-treated cells was the accumulation of a distinct set of C-terminal COX-2 fragments smaller than 72 kDa (zones II and III). Blocking the proteasome probably leads to processing of COX-2 by calpain-independent pathways, since the fragments were similar in cells treated with epoxomicin alone and with epoxomicin plus zVF.

We speculate that products of proteasomal action on COX-2 are subsequently processed by calpain in MAEC. Although the sequence of steps yielding various

COX-2 fragments is not completely delineated by these inhibitor studies, the qualitatively similar effect of PIN1 KD and calpain inhibitors on COX-2 suggests that PIN1 regulates calpain-dependent processing of the enzyme.

137

Previously, we observed that PIN1 depletion and zVF each increased the induction of iNOS, and reduced its degradation. PIN1 physically interacted with iNOS. The WW and catalytic domains of PIN1 appeared to contribute to the association (135 ). This suggested that PIN1 depletion might alter the

susceptibility of its targets to digestion by calpain. For example, PIN1 could

associate with these substrates and catalyze proline isomerization, affecting

protease sensitivity. In contrast to iNOS, however, interaction between COX-2

and PIN1 was not detected here. A role for direct interaction between COX-2 and

PIN1 cannot be completely excluded, however, since association of the proline

isomerase with its putative substrate may be weak or transient. PIN1 could also

affect association of COX-2, or iNOS, with other proteins that may indirectly

regulate proteolysis. It is also possible that PIN1 regulates the induction of COX-

2 and iNOS, by affecting calpain activity in MAEC. This possibility is addressed in

chapter 4.

The effects of COX-2 in acute and chronic inflammatory responses in the

vasculature are complicated by multiple primary and secondary stimuli that may

be present, and by cellular factors, such as supply of arachidonic acid,

complement of various prostaglandin synthases, and expression of prostaglandin

138

receptors (277 ). Thus, it remains to be determined whether selective

manipulation of PIN1 in endothelial cells will be therapeutically useful. In any

case, the results suggest that COX-2 is degraded by calpain, which is regulated

by PIN1 in MAEC.

In summary, suppression of PIN1 increased induction of COX-2 by LPS/IFN-γ by a post-transcriptional mechanism. Consistent with the short lifespan of COX-2 in

MAEC, suppression of PIN1 and calpain inhibitor increased its induction. The similarity in COX-2 degradation in KD MAEC and cells treated with calpain inhibitors, and differences from cells treated with epoxomicin, indicate that PIN1 selectively affects a component of COX-2 turnover mediated by calpain. This previously unknown connection suggests that PIN1 may normally function to restrain the induction COX-2, iNOS, and perhaps other substrates in MAEC in calpain dependent manner. PIN1 is likely to regulate a range of calpain- dependent endothelial activities.

139

CHAPTER 4

PIN1 REGULATES THE CALPAIN/CAST SYSTEM

IN ENDOTHELIAL CELLS

Abstract

Depletion of the peptidyl-proline isomerase, Protein Interacting with Never in

Mitosis Gene A -1 (PIN1), increased the induction of iNOS and COX-2 in MAEC treated with E. coli endotoxin LPS plus IFN-γ. The enhanced induction was by a

post-transcriptional effect associated with reduced degradation of iNOS and

COX-2. Degradation of iNOS and COX-2 was inhibited by the calpain inhibitor,

carbobenzoxy-valinyl-phenylalaninal (zVF). We hypothesized that PIN1

regulated calpain activation or CAST in MAEC. Knockdown of PIN1 with small

hairpin RNA reduced the basal and LPS/IFN-γ stimulated activity of calpain as compared with MAEC harboring a control small hairpin RNA. However, the depletion of PIN1 did not affect the levels of the large or small subunits of ubiquitous, heterodimeric calpains. PIN1 did not interact with these proteins either. Instead, deletion of PIN1 increased the basal level of CAST, the endogenous protein inhibitor of heterodimeric calpains. The effect of PIN1 on

CAST was post-transcriptional, as CAST mRNA was not affected by PIN1

140

depletion. Loss of calpastatin, after inhibiting protein synthesis with cycloheximide, was slower in KD than control MAEC, indicating that PIN1 regulates degradation of CAST. Inhibition of calpain with zVF increased CAST protein levels in control MAEC, whereas a pan-caspase inhibitor, did not, suggesting that calpain, not caspase, contributed to the turnover of basal CAST in MAEC. Extracts from KD treated with either vehicle or LPS/IFN-γ contained

more inhibitory activity towards an exogenous calpain compared to control

shRNA MAEC. Co-immunoprecipitation and glutathione-S-transferase-PIN1

fusion protein pull-down results indicated that CAST associates with PIN1. The

PIN1 fusion protein also antagonized the ability of CAST to inhibit exogenous

calpain in vitro. The results suggest that PIN1 associates with CAST, either

facilitating degradation of CAST or regulating its inhibitory activity towards

calpain in MAEC. PIN1 may normally maintain calpain activity and consequently

limit the induction of calpain substrates, such as iNOS and COX-2 in endothelial

cells. The regulation of calpain system by PIN1 in MAEC indicates that PIN1 may

play an essential role in various calpain related physiological and pathological

processes.

141

Introduction

The peptidyl-prolyl isomerase, PIN1, acts on proline (P) following phosphorylated

(p) serine (S) or threonine (T) (p(S/T)-P) (236, 237 ). PIN1 contains an N-terminal domain, characterized by two tryptophan residues separated by 22 amino acids that can bind to phosphorylated serine- or threonine-proline sequences in substrate proteins, known as a WW domain. PIN1 also isomerizes p(S/T)-P with its carboxyl-terminal catalytic domain (237, 238 ). This conformational change may have profound effects on protein-protein interactions, subcellular location and the susceptibility to enzymes (18, 21, 29, 30, 236, 237, 239, 240, 293 ).

Previously, we found that depletion of PIN1 with small hairpin RNA enhanced the induction of iNOS and COX-2 in MAEC treated with LPS/IFN-γ. Increased expression of iNOS and COX-2 was due to reduced degradation of each protein, which was also antagonized by a calpain inhibitor. Although PIN1 associated with iNOS, no association was detected with COX-2. PIN1 KD and calpain- inhibitors caused production of similar COX-2 fragments, suggesting that PIN1 may regulate calpain and then modulate COX-2 degradation in MAEC.

Calpain is a family of calcium-activated neutral cysteine proteases that have been implicated in many physiological processes such as cytoskeletal

142

remodeling, cell cycle progression, gene expression, apoptosis and tissue injury

(186, 294, 295 ). The major ubiquitous isoforms of the enzyme, micro (µ) and

milli (m), are heterodimers between calpain 1 and 2 and the common small

subunit, calpain 4. Heterodimeric calpain activity is regulated at multiple levels:

expression of the subunits, requirement for calcium, phosphorylation,

autoproteolysis, intracellular distribution, and, most importantly, by the

endogenous inhibitor CAST (201-209 ). Since depletion of PIN1 and calpain

inhibition reduced the turnover of iNOS and COX-2 in MAEC, we hypothesized

that PIN1 either regulates calpain or CAST in these cells (206, 296 ).

Material and Methods

Fetal bovine serum was obtained from Hyclone Laboratories (Logan, UT).

Bradford reagent, endothelial cell growth supplement, serotype 0111:B4, heparin, phenylmethylsulfonyl fluoride, polybrene, and antibodies against calpain 1, calpain 2 and CAST subdomain 4A were purchased from Sigma Chemical Co.

(St Louis, MO). Anti-CAST antibody was purchased from cell signaling

Technology. Dulbecco’s minimum essential medium, trypsin, isopropyl-ß-d- thiogalactopyranoside, Trizol, Superscript Reverse Transcriptase Taq DNA polymerase, RNAse-free DNAse, deoxynucleotides, pcDNA 3 plasmids were from Invitrogen (Carlsbad, CA). Cycloheximide, carbobenzoxy-valinyl-

143

phenylalaninal (zVF, or MDL-28170 or calpain inhibitor III) and [4-((4-

(dimethylamino)phenyl)azo)benzoic acid, succinimidyl ester]-threonine-proline- leucine-lysine~serine-proline-proline-proline-serine-proline-arginine-[ 5-((2- aminoethyl)amino)naphthalene-1-sulfonic acid] and porcine erythrocyte µ-calpain were obtained from Calbiochem (La Jolla, CA). Triton X-100 and enhanced chemiluminescence reagent were obtained from Pierce (Rockford, IL). Tween 20, tris-base, sodium dodecyl sulfate, NaCl, Na3VO4, NaF, ethidium bromide, ethylenediamine tetraacetic acid and agarose were from Fisher Scientific (Fair

Lawn, NJ). Carbobenzoxy-valinyl-alanyl-aspartyl fluormethyl ketone (zVAD) and anti-PIN1 were from R&D Systems (Minneapolis, MN). Horseradish peroxidase- conjugated goat anti-mouse and anti-rabbit secondary antibodies were obtained from Jackson Immunoresearch Laboratories, Inc. (West Grove, PA).

Cells

MAEC, cultured from aortas of mice in accordance with the Guide for the Care and Use of Laboratory Animals from the U.S. National Institutes of Health, selected for stable knockdown (KD) of PIN1 by 85%, and MAEC transduced with an inactive control vector were used. These cells were were cultured and treated with zVF and cycloheximide as described previously (135 ).

144

Calpain activity and CAST inhibitory activity

Cells were washed and scraped in 1 ml of ice-cold PBS, centrifuged (2 min at

1500 x g, 4°C), and resuspended in 100 mM Tris-HCl, 5 mM EDTA, 5 mM benzamidine,1 mM dithiothreitol, 10 mM ß-mercaptoethanol, and 0.5 mM phenylmethylsulfonyl fluoride. Cell extracts were sonicated with four 10 s pulses.

Debris was removed by centrifuging at 15,000 x g for 20 min. Calpain activity was measured using [4-((4-(dimethylamino)phenyl)azo)benzoic acid, succinimidyl ester]-threonine-proline-leucine-lysine~serine-proline-proline- proline-serine-proline-arginine-[ 5-((2-aminoethyl)amino)naphthalene-1-sulfonic acid as a substrate. The reaction contained 15 mM calcium in 50 µl of calpain buffer (10 mM HEPES, 150 mM NaCl, 1 mM EDTA, 5 mM benzamidine, 0.5 mM phenylmethylsulfonyl fluoride, 10mM ß-mercaptoethanol, pH 7.5) and 40 µg extracted protein. Substrate was added to produce 100 µM, and reaction was monitored at 30°C by fluorescence (320 nm excitation and 480 nm emission).

CAST activity was measured by determining the inhibitory effect of cell extracts towards degradation of the calpain substrate by exogenous porcine µ-calpain.

Cell extracts, isolated as described above, were incubated at 37 degrees for 1 h

in the presence of 15 mM Ca 2+ to inactivate endogenous calpain (206 ). 0.8 units of porcine µ-calpain and 60 µg cell extract proteins were combined and adjusted

to 50 µl with calpain buffer. The reaction was started by adding 100 µM calpain

145

substrate and monitored as indicated above. CAST inhibitory activity was expressed as % inhibition of the exogenous calpain (100 x [calpain activity in the absence of extracts minus calpain activity in the presence of extracts] / [calpain activity in the absence of extracts]). Glutathione S-Transferase (GST)-PIN1 fusion protein (1 µg) was added to some reactions to assess its effect on CAST inhibitory activity (135 ).

mRNA Levels

RNA was extracted and analyzed by RT-PCR as described previously(135 ).

CAST primers were designed from Genbank Accession NM 009817 (sense: 5’-

CTT GTC TCA GGA CTT CTC CA, antisense: 5’-TGA GGT CTC TGT AGT TGA

GG). ß-actin was measured as described previously(284 ). PCR products were

electrophoresed in agarose gels and stained with ethidium bromide for digital

imaging.

GST-pull down and Western Blotting

For GST pulldown, 500 µg of MAEC lysate protein were pre-cleared by

incubation with glutathione-sepharose at 4°C for 2 h. The supernatant was then

incubated with a specific GST-PIN1 fusion proteins at 4°C for 2 h. All samples

were then centrifuged 16,000 g x 1 min, and supernatants were discarded. The

146

sepharose pellets were washed three times with lysis buffer by resuspension and centrifugation. Western blots were produced and analyzed as described previously (135 ).

Data Analysis

Digital images of proteins and PCR products were measured with Image J 1.34

(NIH). Data were analyzed by Student’s t test or analysis of variance (ANOVA) with Bonferroni correction for multiple comparisons (297 ).

Results

Knockdown of PIN1 decreases Calpain activity in MAEC

Previous results showed that PIN1 regulated calpain mediated iNOS and COX-2 degradation. For example, PIN1 may associate with iNOS and catalyze proline isomerization, affecting its susceptibility to calpain cleavage. In contrast to iNOS, we have not detected interaction between COX-2 and PIN1 with similar conditions of immunoprecipitation, suggesting that PIN1 may have multiple mechanisms to regulated calpain mediated substrate degradation. The

147

similarities between the effects of PIN1 KD and calpain-inhibitors on COX-2 fragments suggested that PIN1 may regulate calpain.

Here, the fluorogenic calpain substrate, DABCYL-TPLKSPPPSPR-EDANS was used to measure basal and LPS/IFN-γ stimulated calpain activity in KD and

control MAEC as described by Tompa et al. (197 ). The fluorogenic calpain substrate is cleaved by porcine µ-calpain, which is blocked by a calpain inhibitor, zVF but not by a proteasome inhibitor epoxomicin (EPO) (Fig. 4.1A). Basal calpain activity was 2.2-fold greater in control cells than in KD (Fig. 4.1B).

LPS/IFN-γ stimulation dramatically increased calpain activity in both KD and

control cells while calpain activity was still about 2-fold greater in control cells

than in KD.

148

300

200

100 Relative Velocity Relative

O F S V z EPO DM

1500 + KD * 1200 Control

900 + Initial 600

300 Reaction Velocity Reaction * 0 Vehicle L/I

Fig. 4.1 Effect of PIN1 knockdown on calpain activity. A. Purified porcine µ-

calpain was added into reaction mixture containing DMSO, zVF (25 µM) or

149

epoxomicin (EPO, 2 µM). Calpain activity was measured as initial rate of

cleavage of the calpain substrate (DABCYL)-TPLK~SPPPSPR-(EDANS). B. KD

and control shRNA MAEC were treated with vehicle or LPS/IFN-γ for 24h and calpain activity were measured. Bars represent mean + SE from 4 independent cultures in each group. *, p <0.05 for comparison between KD and control shRNA; +, p <0.05 for comparison between vehicle and LPS/IFN-γ treatment.

Suppression of PIN1 does not affect protein level of calpain 1, 2 and 4 in MAEC

The ubiquitous µ- and m-calpains are heterodimers of calpain small subunit

(calpain 4) with calpain 1 and 2 respectively (206 ). Since a reduction in calpain activity could be due to reduced expression of the major subunits calpain 1, 2 or

4, expression was assessed by western blotting. Suppression of PIN did not affect the protein level of these calpain subunits either in basal and LPS/IFN-γ

stimulated condition (Fig. 4.2A/B). Even though both calpain 1 and calpain 2 can

be phosphorylated in Ser and Thr residues (204-206, 217-220 ), neither of them could be pull down by GST-PIN1 fusion protein, suggesting that PIN1 regulates calpain activity indirectly rather than direct association with calpains (Fig. 4.2C).

150

Vehicle L/I shRNA: KD C KD C

Calpain 1

Calpain 2

α-Tubulin

1.0 KD 1.0 C 0.8 0.8 -Tubulin -Tubulin 0.6 0.6 α α α α α α α α 0.4 0.4 Signal Ratio Signal Signal Ratio Signal 0.2 0.2 Calpain2/ Calpain 1/ Calpain 0.0 0.0 Vehicle L/I Vehicle L/I

Fig. 4.2 Effect of PIN1 knockdown on calpain 1, 2 and 4 protein levels

continued

151

Fig. 4.2 continued

Vehicle L/I shRNA: KD C KD C

Calpain 4

α-Tubulin

10.0 KD C 7.5 -Tubulin α α α α 5.0

Signal Ratio Signal 2.5 Calpain 4/ Calpain 0.0 Vehicle L/I

GST GST -PIN1 Calpain 1

Calpain 2

GST-PIN1

GST

Fig. 4.2 Effect of PIN1 knockdown on calpain 1, 2 and 4 protein levels. PIN1 KD and control (C) MAEC were starved overnight and cell lyses were collected.

152

Representative western blots and analysis of calpain 1, calpain 2 and α-tubulin

levels are shown in panel A and calpain 4 in panel B. Bars represent mean + S.E.

ratio of calpain 1, 2 and 4 to α-tubulin from densitometric analysis of images from

3 independent cultures in each group. C. Cell extracts from control shRNA MAEC were incubated with GST or GST-wild-type PIN1 fusion proteins, and glutathione sepharose, and analyzed by Western blotting with anti-calpain 1, 2 and anti-GST antibodies.

Suppression of PIN1 affects basal CAST protein level of in MAEC

The activity of heterodimeric calpains is tightly controlled by CAST (200 ). Thus the level of CAST in PIN1 KD and control MAEC was examined. The reduced calpain activity of KD cells was associated here with a 2.7-fold increase in CAST protein signal compared with control MAEC (Fig. 4.3A). LPS/IFN-γ treatment for

24h decreased CAST in KD to a similar level to control, which may be due to decreased CAST mRNA level (Fig. 4.3B) or increased calpain activity (Fig. 4.1B).

153

Vehicle L/I

shRNA: KD C KD C

CAST

α-Tubulin

1.2 KD C 0.8 -Tubulin α α α α * # 0.4 CAST/ signal intensity signal ratio 0.0 Vehicle LI

Fig. 4.3 Effect of PIN1 knockdown on CAST protein and mRNA

continued

154

Fig. 4.3 continued

Time(h): 0 1 2 4 8 shRNA: KD C KD C KD C KD C KD C

CAST

ß-actin

KD 1.2 C + * + + * 0.8 * -actin β β β β

0.4 CAST/ Signal Intensity Ratio Intensity Signal 0.0 0 1 2 4 8 Time (h)

Fig. 4.3 Effect of PIN1 knockdown on CAST protein and mRNA. A. KD and

control (C) shRNA MAEC were treated with vehicle or LPS/IFN-γ for 24h. Cell

extracts of were collected and CAST and α-tubulin were measured by western

blotting. Representative blots and analysis are shown. Bars represent mean +

S.E. ratio of CAST/ α-tubulin from densitometric analysis of 3 cultures of each

155

group. #: p <0.05 for comparison between KD and control shRNA. B. KD and control shRNA MAEC were treated with LPS/IFN-γ and mRNAs were collected at

indicated time periods. mRNA encoding CAST and ß-actin were determined by

RT-PCR and agarose gel electrophoresis. Representative images of CAST and

ß-actin PCR products in ethidium-stained gels are shown. Bars represent mean +

S.E. ratio of CAST:ß-actin mRNA from densitometric analysis of images from 3

independent cultures. + and *: p <0.05 for comparison between 2h, 4h, 8h and 0h

in KD and control respectively.

Suppression of PIN1 affects basal CAST stability in MAEC

PIN1 knockdown did not affect CAST mRNA, suggesting a post-transcriptional

effect of PIN1 on CAST expression. Increased expression of CAST could result

from reduced degradation of the protein. The loss of CAST after inhibition of

protein synthesis was therefore investigated. KD and control shRNA cells were

treated with 90 µg CHX/ml to block translation. Proteins were extracted at 0, 7,

24, and 48 h after the addition of CHX and subjected to western blot analysis.

Regression analysis indicated that the rate of loss of CAST was significantly

different between KD and control MAEC. CAST level fell to less than 50% of the

156

initial value by 24 h after CHX in the control shRNA cells. In KD cells, a similar decrease in CAST was not detected even 48 h after CHX treatment. The half-life of CAST protein was about 17 h in control shRNA cells, but it was prolonged to more than 48 h in PIN1 KD MAEC (Fig 4.4A). CAST is susceptible to degradation by calpain (298, 299 ) and by caspases (300-303 ). Thus, the effects of calpain inhibitor, zVF and the general caspase inhibitor, zVAD, on CAST were determined. zVF significantly increased CAST protein in control shRNA cells to an extent in PIN1 KD (Fig 4.4B), while CAST protein level did not differ between vehicle and zVAD treatment in either PIN1 KD or control (Fig 4.4C).

157

shRNA: KD Control Time (h): 0 7 24 48 0 7 24 48

Calpastatin

α-Tubulin

125 KD 100 C

50 -tubulin Ratio Signal α α α α 25 (% of at Value 0 h)

0 CAST/ 0 12 24 36 48 Time (h)

Fig. 4.4 Effect of PIN1 knockdown on CAST protein stability

continued

158

Fig. 4.4 continued

Vehicle zVF shRNA : KD C KD C CAST

α-Tubulin

2.0 KD * C 1.6

1.2 -Tubulin α α α α 0.8 #

CAST/ 0.4 Signal Intensity Ratio Intensity Signal 0.0 Vehicle zVF

Vehicle zVAD shRNA : KD C KD C

CAST

α-Tubulin

2.0 KD 1.6 C

1.2 -Tubulin α α α α 0.8 # #

CAST/ 0.4 Signal Intensity Ratio Intensity Signal 0.0 Vehicle zVAD

Fig. 4.4 Effect of PIN1 knockdown on CAST protein stability. A. The protein synthesis inhibitor cycloheximide (90 µg/ml) was added to KD and control (C)

159

shRNA MAEC, and cell extracts were collected at the indicated times. CAST and

α-tubulin in each sample were determined by image analysis of western blots.

Representative blots are shown. Ratios of CAST to α-tubulin were determined,

and the average ratio + SE of 3 independent cultures for each point, as a percent

of the value at 0 h after cycloheximide treatment, was calculated. The dashed

line marks the 50% value. The solid lines are linear regressions for KD and

control cells. The slopes were significantly different (p<0.05). PIN1 KD and

control shRNA MAEC were treated with vehicle, zVF (25 µM) (B) or a general caspase inhibitor zVAD (25 µM) (C) for 48h. CAST and α-tubulin were

determined by image analysis of western blots. Bars represent the mean + S.E.

of three independent cultures per group. *, p<0.05 for comparison between

vehicle and zVF or zVAD. #, p<0.05 for comparison between KD and control

shRNA.

Suppression of PIN1 affects both basal and LPS/IFN-γ-stimulated CAST

inhibitory activity in MAEC

The inhibitory efficiency of CAST on calpain can also be regulated by post-

translational modifications, including phosphorylation (208, 209 ). Therefore, the inhibitory activity of CAST towards exogenous porcine µ-calpain was examined in

160

PIN1 KD and control cells. Cell extracts from untreated PIN1 KD contained significantly higher CAST inhibitory activity than that in control MAEC (Fig. 4.5A).

Interestingly, even though the protein level of CAST in KD was similar to that in control after LPS/IFN-γ treatment, KD still contained significantly higher CAST inhibitory activity than control shRNA MAEC (Fig. 4.5B). This may explain the lower calpain activity in KD after LPS/IFN-γ treatment. PIN1 might affect CAST inhibitory activity by association with CAST, which is phosphorylated (208, 209 ) and contains several S/T-P motifs. Thus, the potential for interaction between

CAST and PIN1 was assessed. GST-PIN1 but not GST pulled down CAST (Fig.

4.5C), and CAST was coimmunoprecipitated with PIN1 from extracts of control shRNA MAEC (Fig. 4.5D).

161

KD 90 # C

80 -calpain µ µ µ µ

% inhibition of 50

KD 90 # C

80 -calpain µ µ µ µ

% % inhibition of 50

Fig. 4.5 Effect of PIN1 knockdown on CAST inhibitory activity

continued

162

Fig. 4.5 continued

GST GST-PIN1

CAST

GST-PIN1

GST

IP: None PIN1

CAST

PIN1

Fig. 4.5 Effect of PIN1 knockdown on CAST inhibitory activity. Cell extracts of KD and control (C) shRNA MAEC were treated with vehicle (A) or LPS/IFN-γ (B) for

24h and CAST inhibitory activities were measured by using porcine µ-calpain.

163

#: p <0.05 for comparison between KD and control cells. Cell extracts from control shRNA MAEC were incubated with GST or GST-wild-type and glutathione sepharose (C), and immunoprecipitated without or with anti-PIN1 antibody (D).

Western blotting with anti-CAST and anti-PIN1 was shown.

Purified GST-PIN1 decreases CAST inhibitory activity

From the above results, PIN1 may directly associate with CAST and regulate its inhibitory activity. Thus, the effect of direct addition of purified GST, GST-PIN1 and its mutants, W33A and K64A, in the CAST inhibitory activity assay was examined. GST-PIN1 significantly decreased CAST inhibitory activity of KD extracts treated with vehicle or LPS/IFN-γ compared with GST (Fig. 4.6A/B).

Both binding activity and catalytic activity were required for this effect, since

W33A and K64A were unable to decrease CAST activity compared to GST.

164

-calpain 40 µ µ µ µ

+ 20

% inhibition of 0 GST GST-PIN1 W33A K64A

50 -calpain µ µ µ µ

25 +

% % inhibition of 0 GST GST-PIN1 W33A K64A

Fig. 4.6 Purified GST-PIN1 decreases CAST inhibitory activity. KD MAECs were treated with vehicle (A) or LPS/IFN-γ (B) for 24h and cell extracts were collected.

Purified GST, GST-PIN1 and its mutants W33A, K64A (1 µg) were added into

165

CAST inhibitory activity reaction mixture. Bars represent mean + S.E. ratio of percentage of CAST inhibitory activity of 5 cultures of each group +: p <0.05 for comparison between GST-PIN1 and GST, W33A, K64A.

Discussion

We previously found that PIN1 depletion and the calpain inhibitor, zVF, increased the induction of iNOS and COX-2, and reduced their degradation. PIN1 associated with iNOS, suggesting that it might regulate its susceptibility to digestion by calpain. Thus, binding of PIN1 to iNOS, or proline isomerization in iNOS, might regulate the sensitivity of the enzyme to calpain. However, we did not detect any association between PIN1 and COX-2, indicating PIN1 may also regulate COX-2 turnover via activating calpain. Consistent with these observations, depletion of PIN1 by 85% in MAEC decreased calpain activity in vehicle and LPS/IFN-γ stimulated MAEC (Fig 4.1). Decreased calpain activity could result from decreases in levels of calpain subunits in cells lacking PIN1.

However, depletion of PIN1 did not reduce the expression of subunits of ubiquitous heterodimeric µ- or m-calpains, calpain 1, 2 and 4 here (Fig 4.2).

Calpain activity can also be regulated by phosphorylation. For example, phosphorylation on calpain 2 by protein kinase A (PKA) decreased its activity

166

while ERK directly phosphorylates and activates calpain 2 both in vitro and in vivo (204, 205 ). It is possible that PIN1 binds phosphorylated calpains and regulates its activity directly. However, this was not the case in MAEC since association of heterodimeric calpain subunits with PIN1 was not observed.

Calpain activity is controlled by the endogenous protein inhibitor, CAST, which inhibits heterodimeric µ- and m-calpain. Increases in CAST protein are associated with reduced calpain activity in several experimental settings (304,

305 ). Here, depletion of PIN1 increased the basal level of CAST in MAEC (Fig.

4.3A), which may not result from transcription since no significant differences of mRNA were observed in KD and control shRNA cells (Fig. 4.3B). This suggests that PIN1 regulates translation or degradation of CAST. Here, the loss of CAST after treating MAEC with cycloheximide was measured. Although CAST was longer-lived than iNOS or COX-2 as expected, depletion of PIN1 greatly stabilized it (Fig. 4A). Although CAST is susceptible to degradation by calpain and caspases, inhibition of calpains, but not caspases, increased CAST level in control (Fig. 4.4B/C). The results suggest that CAST is normally degraded by calpain in MAEC. In parallel with the CAST protein level, PIN1 KD contained siginificantly higher inhibitory activity (Fig. 4.5A). Interestingly, CAST inhibitory activity was higher in KD compared with control after LPS/IFN-γ treatment. This

167

may explain lower calpain activity in KD, even though the CAST protein level was similar to that in control after LPS/IFN-γ treatment (Fig. 4.5B). Furthermore, purified GST-PIN1 but not mutant W33A and K64A significantly decreased CAST inhibitory activity in both extracts of vehicle and LPS/IFN-γ treated cells (Fig. 4.6).

Phosphorylation of the active CAST by protein kinase C and protein kinase A

decreases its inhibitory activity (208, 209 ). Thus, PIN1 may directly associate with phosphorylated CAST and reduce its inhibitory activity against calpain in

MAEC (Fig. 4.4C/D).

The calpain-CAST system is in a loop, in which heterodimeric calpain is a protease and CAST is its specific endogenous inhibitor. CAST is also a substrate of the enzyme. Two crystallographic studies uncover a novel mechanism of protease inhibition (200, 224, 225 ). Subdomain A and C in CAST inhibitory domain binds to two separate domains, IV and VI, in calpain 2 and calpain 4 respectively, which allows inhibitory subdomain B to wrap around the active site region of the calpain heterodimer and inhibit its activity. In both of these studies, several amino acids adjacent to the active site of calpain in subdomain B form a loop, which is held away from the catalytic cysteine, avoiding proteolysis by calpain. Deletion of one or 2 amino acids in the loop led to cleavage by the calpain. Previous studies have shown that the disordered regions between

168

inhibitory domains can be cleaved by calpain even at relatively high concentrations of CAST (225 ). Thus its binding sequence (subdomain A/C),

inhibitory sequence (subdomain B) and cleavage sequence (region between

inhibitory domains) are located separately in CAST. Ten S/T-P motifs are located

in mouse CAST protein (close to subdomain A, B, C and in region between

Inhibitory domains), some of which are highly conserved in different species, and

might bind PIN1. PIN1 may therefore regulate calpain activity by affecting the

conformation of CAST bound to calpain, for example, modulating the binding

affinity of CAST to calpain (subdomain A/C), inhibitory efficency against calpain

(subdomain B) or susceptibility to calpain (region between inhibitory domains).

The effects of PIN1 on CAST may require their interaction which was observed in

(Fig. 4.4C/D). Association between PIN1 and CAST is consistent with the known

serine/threonine phosphorylation of CAST (208, 306 ), although the specific phosphorylated residues have not been identified. Phosphorylation of CAST at unknown residues also reduces its interaction with calpain (223 ). PIN1 could

indirectly regulate the phosphorylation state of CAST by modulating kinases or

phosphatases. Altered phosphorylation could then affect the inhibitory activity of

CAST. It is not clear whether one or more than one are phosphorylated and

responsible for the association with PIN1. Thus, to identify S/T-P sites necessary

for direct interaction between PIN1 and CAST is essential for understanding how

169

PIN1 regulates calpain system in endothelial cells. Related signaling pathways and the responsible kinases in basal and LPS/IFN-γ stimulated conditions are

equally important to be investigated.

Collectively, PIN1 significantly affects calpain activity by regulating CAST in

MAEC. PIN1 associates with CAST, reducing its protein stability and efficiency to

inhibit calpains. The elevation of basal CAST in cells lacking PIN1 is due to

reduced degradation, which appears to be mediated by calpain. Consistent with

the known phosphorylation of CAST, PIN1 binds to the inhibitor in vitro and

antagonizes its ability to inhibit calpain. Regulation of calpain via CAST by PIN1

may account for its effect on induction of iNOS and COX-2 in MAEC, and it may

affect other calpain substrates and activities of endothelial cells. Alterations in

PIN1 expression or function may significantly affect a broad range of calpain-

dependent processes in EC. Further investigations should be performed to

understand how PIN1 interacts and regulates CAST.

170

CHAPTER 5

PIN1 REGULATES CALPAIN/CAST SYSTEM

IN VIVO

Abstract

Previous studies showed that PIN1 regulated calpain activity by acting on calpastatin (CAST) in MAEC. PIN1 may either facilitate degradation of CAST or reduce its inhibitory activity to maintain calpain activity, and consequently limit the induction of calpain substrates, such as iNOS and COX-2, in endothelial cells.

Whether PIN1 regulates calpain/CAST system in vivo, however, is not clear. In the present study, this role of PIN1 was examined by studying PIN1 knockout and wildtype mice. The regulation of calpain system by PIN1 in vivo is tissue specific: lung and kidney from knockout mice contained significantly lower calpain activity which may be due to the higher CAST inhibitory activity in them.

However, higher calpain activity and less CAST inhibitory activity were observed in brains from PIN1 knockout mice. Liver and heart from PIN1 knockout mice contained significantly lower calpain activity although the CAST inhibitory activity is similar to wildtype mice, indicating mechanisms other than CAST were involved in PIN1-regulated calpain activity in these organs. PIN1 knockout

171

influenced normal cariac function, causing a significant decrease in stroke volume compared with wildtype.

Introduction

Depletion of the peptidyl-proline isomerase, PIN1, increased the induction of iNOS and COX-2 in MAEC treated with E. coli endotoxin LPS plus IFN-γ (135,

171 ). The enhanced induction was a posttranscriptional effect associated with

reduced degradation of iNOS and COX-2. Degradation of iNOS and COX-2 was

inhibited by the calpain inhibitor, carbobenzoxy-valinyl-phenylalaninal (zVF).

Knockdown of PIN1 with small hairpin RNA reduced the basal and LPS/IFN-γ

stimulated activity of calpain as compared with control shRNA MAEC. PIN1 may

either facilitate degradation of CAST or regulate its inhibitory activity to maintain

calpain activity, and consequently limit the induction of calpain substrates, such

as iNOS and COX-2, in endothelial cells. However, the role of PIN1 in the

regulation of calpain activity in vivo has not been investigated yet. Here, calpain

activity and CAST inhibitory activity of different organs including lung, kidney,

brain, liver and heart from PIN1 wildtype and knockout mice were measured.

172

Materials and Methods

Animals

All aspects of animal use were performed in accordance with the guidelines of the National Institutes of Health and under approval of the Ohio State University

Animal Care and Use Committee. Animals were housed in a sterile cage rack system with HEPA-filtered air circulation (approximately 50 air changes/hour,

Allentown Caging Inc., Allentown, PA). Mice were sacrificed with an overdose of pentobarbital sodium.

Calpain activity and CAST inhibitory activity

For calpain assay in vivo, different organs were collected and homogenized in lysis buffer (100 mM Tris-HCl, 5 mM EDTA, 5 mM benzamidine, 1 mM dithiothreitol, 10 mM ß-mercaptoethanol, and 0.5 mM phenylmethylsulfonyl fluoride). Tissue homogenates were sonicated with four 10 s pulses. Debris was removed by centrifuging at 15,000 x g for 20 min. Calpain activity was measured using [4-((4-(dimethylamino)phenyl)azo)benzoic acid, succinimidyl ester]- threonine-proline-leucine-lysine~serine-proline-proline-proline-serine-proline- arginine-[ 5-((2-aminoethyl)amino)naphthalene-1-sulfonic acid as a substrate.

The reaction contained 15 mM calcium in 50 µl of calpain buffer (10 mM HEPES,

173

150 mM NaCl, 1 mM EDTA, 5 mM benzamidine, 0.5 mM phenylmethylsulfonyl fluoride, 10mM ß-mercaptoethanol, pH 7.5) and certain amount of extracted protein. Substrate was added to produce 100 µM, and reaction was monitored at

30°C by fluorescence (320 nm excitation and 480 nm emission). CAST activity was measured by determining the inhibitory effect of tissue homogenates towards degradation of the calpain substrate by exogenous µ-calpain. Clarified tissue homogenates were incubated at 95 degrees for 10 min to inactivate endogenous calpain. 0.8 units of porcine µ-calpain and tissue homogenates were combined and adjusted to 50 µl with calpain buffer. The reaction was started by adding 100 µM calpain substrate and monitored as indicated above.

CAST inhibitory activity was expressed as % inhibition of the exogenous calpain

(100 x [calpain activity in the absence of extracts minus calpain activity in the presence of extracts] / [calpain activity in the absence of extracts]).

Assessment of cardiac function

Comprehensive cardiovascular performance measures were conducted on male

PIN1 wildtype and knockout mice (C57BL/6) (age 8 weeks, n=6) using echocardiography at rest and following acute isoproterenol treatment (ISO,

2.5mg/kg i.p.). At 8 weeks, in vivo cardiovascular function was determined using a Sonos 1000 echocardiography unit (Hewlett-Packard, Andover, MA). Mice

174

were anesthetized by isoflurane inhalation (0.25–1.5% isoflurane in 95/5%

O2/CO2) and normothermia was maintained by a heating pad. A 7.5 MHz pediatric probe (optimized and dedicated to rodent studies) placed in the parasternal, short axis orientation recorded LV systolic and diastolic internal dimensions. Three loops of M-mode data were captured for each animal, and data were averaged from at least 5 beat cycles/loop. Parameters were determined using the American Society for Echocardiography leading-edge technique in a blinded fashion.

Results

Effect of PIN1 on body weight and organ weight

Different organs were collected from five PIN1 wildtype and knockout mice.

Interestingly, PIN1 knockout mice are smaller with decreased body weight and organ weight when compared to wildtype mice.

175

Whole body (g) Lung (mg) Kidney(mg) heart(mg) brain (mg)

+/+ 24.86 ±0.51* 94.20 ±3.50* 136.28 ±4.62* 107.48 ±2.42* 201.70 ±10.15

-/- 20.72 ±0.71 73.28 ±2.67 96.76 ±3.69 92.46 ±5.20 190.1 ±5.32

Table. 5.1 Effect of PIN1 on body weight and organ weight. Five female PIN1 knockout (-/-) and wildtype (+/+) mice were sacrificed at the age of 6 months. The whole body and organs (right lung, right kidney, right brain and whole heart) were weighted. * p<0.05 for comparison between PIN1 -/- and +/+ samples.

PIN1 knockout decreases calpain activity in lung and kidney

Calpain and CAST inhibitory activity was measured in lung and kidney from PIN1 wildtype and knockout mice. Consistently to lower calpain activity in PIN1 KD

MAEC, lung and kidney from knockout mice contained significantly less calpain activity (Fig. 5.1.A/C), which may be due to the higher inhibitory activity of CAST in PIN1 knockout. No significant difference of CAST protein was observed in lung from PIN1 knockout and wildtype mice, indicating that PIN1 may regulate CAST

176

activity (Fig. 5.1.B). CAST activity in kidney may result from other isoforms of

CAST since the long form was not detected (Fig. 5.1.D).

PIN1 -/- PIN1 +/+

# 60 7500

-calpain 40 µ µ 5000 µ µ # Initial 20 2500 Reaction Reaction Velocity % inhibition of 0 0

Fig. 5.1 Effect of PIN1 knock out on calpain and CAST inhibitory activity in lung and kidney

continued

177

Fig. 5.1 continued

PIN1: -/- +/+ MAEC CAST

PIN1 -/- PIN1 +/+

0.8

0.4 CAST Signal Intensity Signal CAST 0.0 PIN1 -/- PIN1 +/+

PIN1 -/- PIN1 +/+

7500 #

5000 -calpain µ µ µ µ Initial 2500 20 # Reaction Reaction Velocity

0 % inhibition of 0

continued

178

Fig. 5.1 continued

PIN1: -/- +/+ MAEC

CAST

Fig. 5.1 Effect of PIN1 knock out on calpain and CAST inhibitory activity in lung and kidney. Lung (A) and kidney (C) from PIN1 knockout (-/-) and wildtype (+/+) mice (n=5) were collected and homogenized. Calpain (left panel) and CAST activity (right panel) were assayed as described in Materials and Methods. #: p

<0.05 for comparison between PIN1 -/- and +/+ samples. Lung (B) and kidney

(D) samples were homogenized and CAST was measured by western blotting.

Representative blots and analysis are shown. Bars represent mean + S.E. of

CAST signal from densitometric analysis of 5 cultures of each group.

179

PIN1 knockout increases calpain activity in brain

Opposite to lung and kidney, brains from PIN1 knockout contained higher calpain activity (Fig. 5.2A) with lower CAST inhibitory activity compared with wildtype.

The lower CAST inhibitory activity may be due to lower CAST protein level in

PIN1 knockout (Fig. 5.2B).

PIN1 -/- PIN1 +/+ 90 # 20000 #

-calpain 60 µ µ 15000 µ µ

10000 Initial 30 5000 Reaction Reaction Velocity

0 % inhibition of 0

Fig. 5.2 Effect of PIN1 knockout on calpain and CAST inhibitory activity in brain

continued

180

Fig. 5.2 continued

PIN1: -/- +/+ MAEC

CAST

PIN1 -/- PIN1 +/+ + 0.8

0.4 CAST Signal Intensity CAST 0.0 PIN1 -/- PIN1 +/+

Fig. 5.2 Effect of PIN1 knockout on calpain and CAST inhibitory activity in brain.

Brains from PIN1 knockout (-/-) and wildtype (+/+) mice (n=5) were collected and homogenized. Calpain (left panel) and CAST activity (right panel) were assayed as described in Materials and Methods. #: p <0.05 for comparison between PIN1

-/- and +/+ samples. B. Brain samples were homogenized and CAST was measured by western blotting. Representative blots and analysis are shown.

181

Bars represent mean + S.E. of CAST signal from densitometric analysis of 5 cultures of each group. +: p <0.05 for comparison between PIN1 -/- and +/+ samples.

PIN1 knockout decreases calpain activity without influencing CAST activity in liver and heart

Liver and heart from PIN1 knockout mice contained significantly less calpain activity although the CAST inhibitory activity is similar to wildtype (Fig. 5.3 D/E), indicating mechanisms other than CAST were involved in PIN1-regulated calpain activity in these organs.

182

PIN1 -/- PIN1 +/+

# 4000 90 -calpain µ µ µ µ 60

2000 Initial 30 Reaction Reaction Velocity

0 % inhibition of 0

PIN1: -/- +/+ MAEC CAST

PIN1 -/- PIN1 +/+ 0.8

0.4 CAST Signal Intensity Signal CAST 0.0 PIN1 -/- PIN1 +/+

Fig. 5.3 Effect of PIN1 knockout on calpain and CAST inhibitory activity in liver and heart. continued

183

Fig. 5.3 continued

PIN1 -/- PIN1 +/+

# 5000

-calpain 60 µ µ µ µ

2500

Initial 30 Reaction Velocity Reaction

% inhibition of 0 0

Fig. 5.3 Effect of PIN1 knockout on calpain and CAST inhibitory activity in liver and heart. Liver (A) and heart (C) from PIN1 knockout (-/-) and wildtype (+/+) mice (n=5) were collected and homogenized. Calpain (left panel) and CAST activity (right panel) were assayed as described in Materials and Methods. #: p

<0.05 for comparison between PIN1 -/- and +/+ samples. Liver (B) samples were homogenized and CAST was measured by western blotting. Representative blots and analysis are shown. Bars represent mean + S.E. of CAST signal from densitometric analysis of 5 cultures of each group.

184

PIN1 knockout influences c ardiac function

High quality cardiac ultrasound imaging was achieved under light inhalation anesthesia as described in the methods section. Although no difference in heart rate was detected, PIN1 knockout mice showed decrease in SV at rest and after acute cardiac stress compared with wildtype mice (Fig. 5.4).

PIN1 +/+ 600 PIN1 -/-

400

HR (bpm) HR 200

0 Basal ISO

Fig. 5.4 Effect of PIN1 knockout on cardiac function continued

185

Fig. 5.4 continued

# 60 PIN1 +/+ # PIN1 -/- * 40 * l) µ µ µ µ

SV ( SV 20

0 Basal ISO

Fig. 5.4. Effect of PIN1 knockout on cardiac function. Comprehensive

cardiovascular performance measures were conducted on male PIN1 wildtype

(+/+) and knockout (-/-) mice (age 8 weeks, n=6) using echocardiography at rest

and following acute isoproterenol treatment (ISO, 2.5mg/kg i.p.). Cardiac

performance parameters heart rate (HR) and stroke volume (SV) were shown in

A and B respectively. Bars represent mean + S.E. of values of 6 animals. #: p

186

<0.05 for comparison between basal and isoproterenol groups, *: p <0.05 for comparison between PIN1 -/- and +/+.

Discussions

Since PIN1 function may be tissue specific, the much more complicated regulation of calpain-CAST system by PIN1 status in vivo may be expected. PIN1 deficiency decreased calpain activity and increased calpastatin activity in lung and kidney, which may be due to the regulation of CAST by PIN1 (Fig. 5.1).

Calpain and CAST levels were highest in the lung (307 ), thus it is necessary to study how calpain/CAST system is involved in various aspects of lung function.

For example, in pulmonary artery endothelial cells, calpain activity is necessary to maintain angiogenesis and eNOS gene transcription, protein content and its activity, while calpain inhibition by increased CAST protein level in response to cigarette smoke extracts impaired angiogenesis, decreased eNOS gene transcription, and acetylcholine-induced endothelium-dependent relaxation of the pulmonary artery (288, 308, 309 ). PIN1 might regulate calpain-mediated angiogenesis and eNOS expression and activity in pulmonary artery endothelial cells.

187

Excessive activation of calpains or loss of CAST and deregulation of PIN1 have been implicated in neurodegenerative diseases, traumatic brain and spinal cord injuries (295 ). For example, genetic deletion of PIN1 and enhanced calpain

activity contributes to the pathological process of AD (8, 28, 37, 226 ). PIN1 binds

and stabilizes phospho-MCL-1, thus protecting oligodendrocytes from apoptosis

after spinal cord injury (37 ). Upregulation of calpain, has been implicated in

apoptosis and tissue degeneration in spinal cord injury, while overexpression of

CAST and calpain inhibitor prevented apoptosis in spinal cord injury (227-235 ).

Collectively, PIN1 protects, and calpain contributes to the process of these

diseases. Therefore, it is essential to find out how calpain-CAST system is

regulated by PIN1 in brain. The effect of PIN1 on calpain-CAST system in brain

is quite different from that in lung and kidney: PIN1 knockout mice contained

more calpain activity, while lower CAST inhibitory activity was observed (Fig. 5.2).

This result appears to verify the negative correlation between PIN1 and calpain

activity. Lower CAST activity in brains from PIN1 knockout mice may be due to

its decreased protein level (Fig. 5.2).

Although excessive activation of calpain contributes to cardiovascular remodeling

induced by angentensin II (310 ), myocardial dysfunction during sepsis (311 ) and

188

during myocardial ischemia and reperfusion (312 ), the physiological function of calpain activity in normal, unstressed hearts was not clear until recently. Galvez et al demonstrated that loss of calpain activity in mouse heart results in progressive dilated cardiomyopathy with decreased ventricular ejection performance (313 ). Basal cardiac calpain activity is necessary for ubiquitin- dependent degradation of specific cardiac proteins, whose abnormal accumulation causes cardiomyocyte degeneration and heart failure (313 ).

Interestingly, PIN1 knockout mice showed significant decrease in stroke volume,

both at rest and after acute cardiac stress with isoproterenol (Fig. 5.4), indicating

a potential role of PIN1 in normal cardiac function. PIN1 may regulate cardiac

function by modulating calpain activity in heart (Fig. 5.3). However, no difference

of CAST level and its activity was observed in hearts from PIN1knockout and

wildtype mice, suggesting that PIN1 may act on calpain interacting proteins

other than CAST, or directly on calpains, to regulate its activity in the heart. PIN1

may also regulate other calpains , such as calpain 3, which is found in skeletal

and cardiac muscles, but which is not inhibited by CAST (206, 313 ).

Regulation of calpain/CAST by PIN1 is more complicated in vivo than in MAEC.

CAST activity is controlled by phosphorylation. For example, two forms of CAST

are present in brains. One form is efficient to inhibit calpain, while the second

189

form is almost completely inactive. Dephosphorylation of the second form by phosphatases can convert it into an active form. Phosphorylation of the active form by PKC and PKA decreased its inhibitory efficiency (208 ). Different

phosphorylation patterns or interconversion between two forms may be involved

in the distinct regulation of CAST activity by PIN1 in lung/kidney and brain. Also

CAST expression patterns are different in various organs. For example, western

blotting results showed that CAST in brain samples contained two bands. The

band with low molecular weight was predominant. It was not the case in lung,

since the band with high molecular weight was predominant. These two bands

were not detected in kidney samples, and CAST in livers showed three bands.

These different bands may represent various isoforms of CAST or degradation

fragments of the long isoform. Thus phosphorylation in various CAST isoforms

and their association with PIN1 should be investigated in MAEC and various

organs.

190

CHAPTER 6

CONCLUSIONS AND FUTURE DIRECTIONS

Conclusions

Endothelial inflammatory activation and dysfunction have been critically implicated in diverse cardiovascular diseases such as atherosclerosis, sepsis and heart failure. Inflammatory cytokines and bacterial products such as LPS and

IFN-γ, can activate vascular endothelial cells, induce the expression of iNOS and

COX-2. They are similarly induced by many inflammatory stimuli, produce NO and various prostaglandins and are believed to play a role in the pathogenesis of many cardiovascular diseases.

Both iNOS and COX-2 are regulated by multiple mechanisms including transcription, mRNA stability, translation and post-translation. Many key proteins in these steps are regulated in a phosphorylation and/or conformation dependent manner. Although PIN1-mediated conformational change of pSer/Thr-Pro motifs is extremely important in phosphorylation signaling, the role of PIN1 on iNOS and

COX-2 induction had been unexplored. Considering the importance of iNOS and

COX-2 in endothelial activation, our investigations were carried out to understand

191

the effect of PIN1 suppression on iNOS and COX-2 induction and to elucidate the underlying molecular mechanisms in endothelial cells.

Stable suppression of PIN1 was generated by using shRNA-lentiviral based knock-down technique, which efficiently decreased PIN1 protein level by 85%.

Here we reported for the first time, the critical involvement of PIN1 in the induction of these proteins. Suppression of PIN1 enhanced the induction iNOS and COX-2 protein by LPS-IFN-γ, which were confirmed by knock-down cells by small interfering RNAs and MAEC isolated from PIN1 wildtype and knockout mice. Both iNOS and COX-2 were functional in producing NO and proglandin E2.

There was no effect on induction of iNOS and COX-2 mRNA, suggesting a posttranscriptional effect. Next, we began using cycloheximide-treated cells to study PIN1 function on their protein stability. It turned out that knockdown of PIN1 increased the stability of iNOS and COX-2 protein. Since both iNOS and COX-2 undergo calpain and proteasome dependent degradation (131-137, 160-167, 170,

171 ), Pharmacological inhibitors of calpain and proteasome were used to elucidate the underlying molecular mechanisms. My results showed that calpain inhibition blocked the degradation of iNOS and COX-2, which is regulated by

PIN1 status. Immunoprecipitation and GST-PIN1 pull-down results indicated that

PIN1 may directly interact with iNOS and regulate its susceptibility to calpain

192

cleavage. However, no direct interaction were detected between PIN1 and COX-

2, suggesting that PIN1 may regulate calpain mediated degradation of COX-2 by different mechanisms, for example, influencing calpain activity. Accordingly, we investigated whether PIN1 regulated calpain activity in endothelial cells. Calpains are Ca2+-regulated cysteine proteases. µ- and m-calpain, two well characterized isoforms, are ubiquitously expressed, catalyze limited proteolysis of substrates and control various cellular functions. Although calpains have been intensely studied for more than 30 years, it is still not very clear how calpain activity is regulated in living cells. Our results demonstrated for the first time PIN1 can regulate calpain activity. In MAEC, knockdown of PIN1 decreased calpain activity in both basal and LPS/IFN-γ-stimulated conditions. Furthermore, PIN1 neither

affected calpain 1, 2 and 4 protein levels nor directly associated with them.

CAST, an endogenous protein inhibitor, is capable of specifically and reversibly

binding and inhibiting calpain activity. The rapidly accumulating structural

information makes it possible to understand how m-calpain is activated by

calcium and how CAST inhibits its activity specifically and potently, as described

in chapter 1 and 4 (200, 224, 225 ). My results demonstrated that the inhibition of

calpain by CAST is critically regulated by PIN1 in endothelial cells. PIN1

knockdown enhanced CAST inhibitory activity in both basal and LPS/IFN-

193

stimulated condition, while purified GST-PIN1, but not its mutants, W33A and

K64A, significantly decreased it compared to GST. Furthermore, PIN1 knockdown could stabilize basal CAST in a calpain-dependent manner.

Immunoprecipitation and GST-PIN1 pull-down results indicated that PIN1 directly interacted with CAST and regulated either its susceptibility to calpain cleavage or its inhibitory activity on calpain. Furthermore, the regulation of calpain-CAST activity was demonstrated in vivo. In lung, kidney, liver and heart, PIN1 knockout may decrease calpain activity compared with PIN1 wildtype mice, by acting on

CAST or other calpain interacting proteins. In brain, PIN1 knockout increased calpain activity with decreased CAST protein level and activity compared with

PIN1 wildtype.

194

Calpain substrates iNOS/COX-2 PIN1

? ?

?pS/T -P ? kinase ?phosphatase Calpains CAST

Degradation

Figure 6.1 PIN1 regulates calpain mediated proteolysis. Calpains catalyze limited cleavage of substrates, which can be regulated by PIN1 via several different mechanisms. First, PIN1 may directly interact with calpain substrates (for example, iNOS) and influence the susceptibility to calpains. Second, PIN1 may regulate calpain activity by acting on its endogenous inhibitor, CAST as in MAEC, lung and kidney. PIN associates with CAST, either regulating CAST protein

195

stability or the inhibitory activity. Third, calpain interacting proteins other than

CAST may be PIN1 targets, as in liver and heart.

Arrows indicate interaction or stimulation

symbols indicate inhibition

“p” indicates phosphorylation

Question mark indicates unknown factors or components

Future directions

Our studies indicated that PIN1 normally limits iNOS and COX-2 induction by

facilitating calpain-mediated degradation in MAEC. In addition, many PIN1

substrates, such as β-catenin, c-Fos, c-Jun and p53, are also calpain substrates.

Thus, it would be interesting to investigate whether it is a common mechanism that PIN1-catalyzed conformational changes regulate calpain mediated degradation.

Our studies indicated that PIN1 may bind to its substrate, iNOS, and regulate its calpain-mediated degradation in EC. Although in vitro studies can provide useful

196

information, it is important that they are verified by in vivo investigations. Thus, induction of iNOS and COX-2 in vivo, especially in EC should be examined in

PIN1 wildtype and knockout mice. PIN1 functions may be cell or tissue specific.

Thus different cell types may have different responses to PIN1 knockdown in vitro and in vivo. For example, induction of iNOS and COX-2 in various cell types other than EC (for example, neural and immune cells), should be investigated since both PIN1 and iNOS are critically involved in Alzheimer’s disease.

Considering the involvement of iNOS and COX-2 in various diseases including cardiovascular, neurodegenerative diseases and cancers, different disease models should be investigated PIN1 wildtype and knockout mice.

COX-2 degradation in MAEC is controlled by calpain and proteasome pathways.

Epoxomicin completely inhibited the rate of loss of 72 kDa COX-2 in cycloheximide treated KD and control cells, while calpain inhibition only partially reduced it. COX-2 fragments were similar in cells treated with epoxomicin alone and with epoxomicin plus zVF, suggesting proteasome inhibition may also block the processing of COX-2 by calpains. All these results indicated that proteasome and calpain pathways are not completely separated and may sequentially contribute to the degradation of COX-2. Further investigations are needed to test this idea.

197

Our investigations also identified PIN1 as a key regulator of the calpain system and subsequent calpain-mediated substrate degradation. However, it leaves several questions that need to be resolved to better understand the function of

PIN1 in calpain related physiological and pathological processes. Although PIN1 regulated calpain activity, and CAST protein level or inhibitory activity in MAEC, further study is necessary to investigate whether regulation of calpain by PIN1 is caused by its action on CAST. In this context, the effect of PIN1 on calpain activity should be determined when CAST is overexpressed or deleted in EC.

Although the direct interaction between PIN1 and CAST is demonstrated in

MAEC, we have not yet characterized phosphorylation sites which mediated the association. CAST in brain tissues can be phosphorylated by PKC and PKA, which promotes a decrease in its inhibitory efficiency (208, 209 ). Several S/T-P

motifs are located close to subdomain A/C (binding to calpains), subdomain B

(inhibiting calpains), and regions between Inhibitory domains (calpain cleavage

sites). Thus it is essential to identify S/T-P sites which are necessary to form

direct interaction between PIN1 and CAST. Signaling pathways and the

responsible kinases are equally important to be identified. Only the long isoform

of CAST was investigated in my study. More than 10 isoforms of CAST have

been reported. All of them contain inhibitory domain and are able to efficiently

198

inhibit calpains. Thus phosphorylation in various CAST isoforms and their association with PIN1 should be investigated in MAEC and various organs.

Regulation of calpain activity in vivo is more complicated than in MAEC. PIN1 knockout may decrease calpain activity in lung, kidney, liver and heart, by acting on CAST or other calpain interacting proteins. Calpains are important to regulate physiological processes, for example, protecting against cardiomyocyte degeneration and heart failure induced by abnormal accumulation of specific cardiac proteins (313 ). Overactivation of calpains or loss of CAST and deregulation of PIN1 have been implicated in neurodegenerative diseases, traumatic brain and spinal cord injuries (8, 28, 37, 226-235 ). Therefore, it will be necessary to investigate how calpain-CAST system is regulated by PIN1 and how this interaction contributes to these diseases, which will provide valuable information to develop drugs that target PIN1 and calpains. For example, does

PIN1 regulate normal cardiac function by modulating calpain activity? PIN1 protects, and calpain contributes to the process of neurodegenerative diseases.

My study showed that PIN1 knockout increased calpain activity with decreased

CAST protein level and activity in brain compared with wildtype, which may indicate that PIN1 negatively correlates with calpain activity in these diseases.

Compared with wildtype mice, PIN1 knockout decreased calpain activity in liver

199

and heart without influencing CAST inhibitory activity, suggesting calpain interacting proteins other than CAST are regulated by PIN1. In this context, it would be interesting to identify these factors in liver and heart.

Although we found that PIN1 interacted with STAT1 in MAEC, the functional consequence was not known. Expression of SOCS-1 or STAT1-driven luciferase in a reporter plasmid in response to different stimuli should be examined in either

PIN1 knockdown or overexpression condition. Other members of STAT family, especially STAT3 and STAT5 contain similar PMSP site in their transactivation domain. Thus, we can not exclude the possibility that PIN1 regulates their activity, especially STAT3. Mice lacking STAT3 Ser727 phosphorylation are smaller in size with less weight in many organs (314 ), which is similar to PIN1 knockout

mice in my study and from others’ results (38 ). Mice lacking STAT3 Ser727

phosphorylation had lower serum levels of insulin-like growth factor 1 (IGF-1)

(314 ), whereas deletion of STAT5 a/b in skeletal muscle reduced IGF-1 level

locally (315 ). Insulin-like growth factor 1 (IGF-1) signaling crucially control organ size and weight (316, 317 ), which is decreased by aging and implicated in degenerative diseases including AD (318-320 ). Considering the role of PIN1 in aging related disease, it would be interesting to study whether PIN1 regulates

IGF-1 signaling by acting on STAT3, 5 or certain signaling components. It is still

200

possible that IGF-1 signaling regulates PIN1 expression or activity and contributes to organ control and development of degenerative diseases.

Although PIN1 is ubiquitous, and may act on many proteins to regulate transcription, mRNA stability, translation, protein turnover and protein function, my thesis shows that the calpain/CAST system is regulated by PIN1.

Furthermore, inflammatory activation in endothelial cells appears to be greatly affected by calpains. This work suggests that PIN1, calpain and CAST might be targeted to manipulate endothelial function in diseases.

201

BIBLIOGRAPHY

1. Pawson, T., and Scott, J. D. (2005) Trends Biochem Sci 30 , 286-90.

2. Sadowski, I., Stone, J. C., and Pawson, T. (1986) Mol Cell Biol 6, 4396-

408.

3. McBride, K. M., and Reich, N. C. (2003) Sci STKE 2003 , RE13.

4. Pestka, S. (1997) Semin Oncol 24 , S9-18-S9-40.

5. Muslin, A. J., Tanner, J. W., Allen, P. M., and Shaw, A. S. (1996) Cell 84 ,

889-97.

6. Cole, P. A., Shen, K., Qiao, Y., and Wang, D. (2003) Curr Opin Chem Biol

7, 580-5.

7. Frame, S., Cohen, P., and Biondi, R. M. (2001) Mol Cell 7, 1321-7.

8. Lu, K. P., and Zhou, X. Z. (2007) Nat Rev Mol Cell Biol 8, 904-16.

9. Blume-Jensen, P., and Hunter, T. (2001) Nature 411 , 355-65.

10. Nigg, E. A. (2001) Nat Rev Mol Cell Biol 2, 21-32.

202

11. Lu, K. P., Liou, Y. C., and Vincent, I. (2003) Bioessays 25 , 174-81.

12. Plotnikov, A., Li, Y., Tran, T. H., Tang, W., Palazzo, J. P., Rui, H., and

Fuchs, S. Y. (2008) Cancer Res 68 , 1354-61.

13. Li, Y., Clevenger, C. V., Minkovsky, N., Kumar, K. G., Raghunath, P. N.,

Tomaszewski, J. E., Spiegelman, V. S., and Fuchs, S. Y. (2006)

Oncogene 25 , 1896-902.

14. Lu, K. P., Finn, G., Lee, T. H., and Nicholson, L. K. (2007) Nat Chem Biol

3, 619-29.

15. Lu, K. P., Hanes, S. D., and Hunter, T. (1996) Nature 380 , 544-7.

16. Pastorino, L., Sun, A., Lu, P. J., Zhou, X. Z., Balastik, M., Finn, G., Wulf,

G., Lim, J., Li, S. H., Li, X., Xia, W., Nicholson, L. K., and Lu, K. P. (2006)

Nature 440 , 528-34.

17. Hayden, M. S., West, A. P., and Ghosh, S. (2006) Oncogene 25 , 6758-80.

18. Ryo, A., Suizu, F., Yoshida, Y., Perrem, K., Liou, Y. C., Wulf, G., Rottapel,

R., Yamaoka, S., and Lu, K. P. (2003) Mol Cell 12 , 1413-26.

203

19. Saitoh, T., Tun-Kyi, A., Ryo, A., Yamamoto, M., Finn, G., Fujita, T., Akira,

S., Yamamoto, N., Lu, K. P., and Yamaoka, S. (2006) Nat Immunol 7,

598-605.

20. Shen, Z. J., Esnault, S., and Malter, J. S. (2005) Nat Immunol 6, 1280-7.

21. Esnault, S., Shen, Z. J., Whitesel, E., and Malter, J. S. (2006) J Immunol

177 , 6999-7006.

22. Anderson, P. (2005) Nat Immunol 6, 1211-2.

23. Zheng, H., You, H., Zhou, X. Z., Murray, S. A., Uchida, T., Wulf, G., Gu, L.,

Tang, X., Lu, K. P., and Xiao, Z. X. (2002) Nature 419 , 849-53.

24. Zacchi, P., Gostissa, M., Uchida, T., Salvagno, C., Avolio, F., Volinia, S.,

Ronai, Z., Blandino, G., Schneider, C., and Del Sal, G. (2002) Nature 419 ,

853-7.

25. Mantovani, F., Tocco, F., Girardini, J., Smith, P., Gasco, M., Lu, X., Crook,

T., and Del Sal, G. (2007) Nat Struct Mol Biol 14 , 912-20.

26. Oberst, A., Rossi, M., Salomoni, P., Pandolfi, P. P., Oren, M., Melino, G.,

and Bernassola, F. (2005) Biochem Biophys Res Commun 331 , 707-12.

204

27. Mantovani, F., Piazza, S., Gostissa, M., Strano, S., Zacchi, P., Mantovani,

R., Blandino, G., and Del Sal, G. (2004) Mol Cell 14 , 625-36.

28. Butterfield, D. A., Abdul, H. M., Opii, W., Newman, S. F., Joshi, G., Ansari,

M. A., and Sultana, R. (2006) J Neurochem 98 , 1697-706.

29. Ryo, A., Nakamura, M., Wulf, G., Liou, Y. C., and Lu, K. P. (2001) Nat Cell

Biol 3, 793-801.

30. Monje, P., Hernandez-Losa, J., Lyons, R. J., Castellone, M. D., and

Gutkind, J. S. (2005) J Biol Chem 280 , 35081-4.

31. Dougherty, M. K., Muller, J., Ritt, D. A., Zhou, M., Zhou, X. Z., Copeland, T.

D., Conrads, T. P., Veenstra, T. D., Lu, K. P., and Morrison, D. K. (2005)

Mol Cell 17 , 215-24.

32. Wulf, G. M., Ryo, A., Wulf, G. G., Lee, S. W., Niu, T., Petkova, V., and Lu,

K. P. (2001) Embo J 20 , 3459-72.

33. Ryo, A., Liou, Y. C., Wulf, G., Nakamura, M., Lee, S. W., and Lu, K. P.

(2002) Mol Cell Biol 22 , 5281-95.

205

34. Lu, P. J., Wulf, G., Zhou, X. Z., Davies, P., and Lu, K. P. (1999) Nature

399 , 784-8.

35. Hamdane, M., Dourlen, P., Bretteville, A., Sambo, A. V., Ferreira, S., Ando,

K., Kerdraon, O., Begard, S., Geay, L., Lippens, G., Sergeant, N.,

Delacourte, A., Maurage, C. A., Galas, M. C., and Buee, L. (2006) Mol

Cell Neurosci 32 , 155-60.

36. Zhou, X. Z., Kops, O., Werner, A., Lu, P. J., Shen, M., Stoller, G., Kullertz,

G., Stark, M., Fischer, G., and Lu, K. P. (2000) Mol Cell 6, 873-83.

37. Li, Q. M., Tep, C., Yune, T. Y., Zhou, X. Z., Uchida, T., Lu, K. P., and

Yoon, S. O. (2007) J Neurosci 27 , 8395-404.

38. Liou, Y. C., Ryo, A., Huang, H. K., Lu, P. J., Bronson, R., Fujimori, F.,

Uchida, T., Hunter, T., and Lu, K. P. (2002) Proc Natl Acad Sci U S A 99 ,

1335-40.

39. You, H., Zheng, H., Murray, S. A., Yu, Q., Uchida, T., Fan, D., and Xiao, Z.

X. (2002) J Cell Biochem 84 , 211-6.

206

40. Kuboki, S., Okaya, T., Schuster, R., Blanchard, J., Denenberg, A., Wong,

H. R., and Lentsch, A. B. (2007) Am J Physiol Gastrointest Liver Physiol

292 , G201-7.

41. Lu, P. J., Zhou, X. Z., Liou, Y. C., Noel, J. P., and Lu, K. P. (2002) J Biol

Chem 277 , 2381-4.

42. Eckerdt, F., Yuan, J., Saxena, K., Martin, B., Kappel, S., Lindenau, C.,

Kramer, A., Naumann, S., Daum, S., Fischer, G., Dikic, I., Kaufmann, M.,

and Strebhardt, K. (2005) J Biol Chem 280 , 36575-83.

43. Ranganathan, R., Lu, K. P., Hunter, T., and Noel, J. P. (1997) Cell 89 ,

875-86.

44. Goligorsky, M. S. (2005) Am J Physiol Renal Physiol 288 , F871-80.

45. Hunt, B. J., and Jurd, K. M. (1998) Bmj 316 , 1328-9.

46. Katz, S. D., Hryniewicz, K., Hriljac, I., Balidemaj, K., Dimayuga, C.,

Hudaihed, A., and Yasskiy, A. (2005) Circulation 111 , 310-4.

47. Hoare, G. S., Birks, E. J., Bowles, C., Marczin, N., and Yacoub, M. H.

(2006) J Appl Physiol 101 , 1466-73.

207

48. Berliner, J. A., Navab, M., Fogelman, A. M., Frank, J. S., Demer, L. L.,

Edwards, P. A., Watson, A. D., and Lusis, A. J. (1995) Circulation 91 ,

2488-96.

49. Rajavashisth, T. B., Yamada, H., and Mishra, N. K. (1995) Arterioscler

Thromb Vasc Biol 15 , 1591-8.

50. Meroni, P. L., Khamashta, M. A., Youinou, P., and Shoenfeld, Y. (1995)

Lupus 4, 95-9.

51. Bach, F. H., Robson, S. C., Winkler, H., Ferran, C., Stuhlmeier, K. M.,

Wrighton, C. J., and Hancock, W. W. (1995) Nat Med 1, 869-73.

52. Luscinskas, F. W., Kansas, G. S., Ding, H., Pizcueta, P., Schleiffenbaum,

B. E., Tedder, T. F., and Gimbrone, M. A., Jr. (1994) J Cell Biol 125 , 1417-

27.

53. Springer, T. A. (1994) Cell 76 , 301-14.

54. Dusting, G. J. (1995) J Vasc Res 32 , 143-61.

55. Behrendt, D., and Ganz, P. (2002) Am J Cardiol 90 , 40L-48L.

56. Ignarro, L. J. (1992) Biochem Soc Trans 20 , 465-9.

208

57. Radomski, M. W., Palmer, R. M., and Moncada, S. (1990) Proc Natl Acad

Sci U S A 87 , 5193-7.

58. Radomski, M. W., Palmer, R. M., and Moncada, S. (1987) Br J Pharmacol

92 , 639-46.

59. Nunokawa, Y., and Tanaka, S. (1992) Biochem Biophys Res Commun

188 , 409-15.

60. von der Leyen, H. E., Gibbons, G. H., Morishita, R., Lewis, N. P., Zhang,

L., Nakajima, M., Kaneda, Y., Cooke, J. P., and Dzau, V. J. (1995) Proc

Natl Acad Sci U S A 92 , 1137-41.

61. Aktan, F. (2004) Life Sci 75 , 639-53.

62. MacMicking, J., Xie, Q. W., and Nathan, C. (1997) Annu Rev Immunol 15 ,

323-50.

63. Kleinert, H., Pautz, A., Linker, K., and Schwarz, P. M. (2004) Eur J

Pharmacol 500 , 255-66.

64. Bogdan, C. (2001) Nat Immunol 2, 907-16.

209

65. Whicher, J., Biasucci, L., and Rifai, N. (1999) Clin Chem Lab Med 37 , 495-

503.

66. Alderton, W. K., Cooper, C. E., and Knowles, R. G. (2001) Biochem J 357 ,

593-615.

67. Dawn, B., and Bolli, R. (2002) Ann N Y Acad Sci 962 , 18-41.

68. Aktan, F., Henness, S., Roufogalis, B. D., and Ammit, A. J. (2003) Nitric

Oxide 8, 235-42.

69. Kroncke, K. D., Fehsel, K., and Kolb-Bachofen, V. (1998) Clin Exp

Immunol 113 , 147-56.

70. Schindler, C., and Darnell, J. E., Jr. (1995) Annu Rev Biochem 64 , 621-51.

71. Milocco, L. H., Haslam, J. A., Rosen, J., and Seidel, H. M. (1999) Mol Cell

Biol 19 , 2913-20.

72. Xie, Q. W., Whisnant, R., and Nathan, C. (1993) J Exp Med 177 , 1779-84.

73. Chu, S. C., Wu, H. P., Banks, T. C., Eissa, N. T., and Moss, J. (1995) J

Biol Chem 270 , 10625-30.

210

74. Eberhardt, W., Kunz, D., Hummel, R., and Pfeilschifter, J. (1996) Biochem

Biophys Res Commun 223 , 752-6.

75. Gao, J., Morrison, D. C., Parmely, T. J., Russell, S. W., and Murphy, W. J.

(1997) J Biol Chem 272 , 1226-30.

76. Doi, M., Shichiri, M., Katsuyama, K., Ishimaru, S., and Hirata, Y. (2002)

Atherosclerosis 160 , 123-32.

77. Kleinert, H., Euchenhofer, C., Fritz, G., Ihrig-Biedert, I., and Forstermann,

U. (1998) Br J Pharmacol 123 , 1716-22.

78. Kleinert, H., Wallerath, T., Fritz, G., Ihrig-Biedert, I., Rodriguez-Pascual, F.,

Geller, D. A., and Forstermann, U. (1998) Br J Pharmacol 125 , 193-201.

79. Ganster, R. W., Taylor, B. S., Shao, L., and Geller, D. A. (2001) Proc Natl

Acad Sci U S A 98 , 8638-43.

80. Huang, H., Rose, J. L., and Hoyt, D. G. (2004) Mol Pharmacol 66 , 302-11.

81. Sagoo, P., Chan, G., Larkin, D. F., and George, A. J. (2004) Invest

Ophthalmol Vis Sci 45 , 3964-73.

211

82. Wagner, A. H., Schwabe, O., and Hecker, M. (2002) Br J Pharmacol 136 ,

143-9.

83. Seidel, H. M., Milocco, L. H., Lamb, P., Darnell, J. E., Jr., Stein, R. B., and

Rosen, J. (1995) Proc Natl Acad Sci U S A 92 , 3041-5.

84. Decker, T., Lew, D. J., Mirkovitch, J., and Darnell, J. E., Jr. (1991) Embo J

10 , 927-32.

85. Kamijo, R., Harada, H., Matsuyama, T., Bosland, M., Gerecitano, J.,

Shapiro, D., Le, J., Koh, S. I., Kimura, T., Green, S. J., and et al. (1994)

Science 263 , 1612-5.

86. Martin, E., Nathan, C., and Xie, Q. W. (1994) J Exp Med 180 , 977-84.

87. Spink, J., and Evans, T. (1997) J Biol Chem 272 , 24417-25.

88. Blair, L. A., Maggi, L. B., Jr., Scarim, A. L., and Corbett, J. A. (2002) J Biol

Chem 277 , 359-65.

89. Shiraishi, A., Dudler, J., and Lotz, M. (1997) J Immunol 159 , 3549-54.

90. Varinou, L., Ramsauer, K., Karaghiosoff, M., Kolbe, T., Pfeffer, K., Muller,

M., and Decker, T. (2003) Immunity 19 , 793-802.

212

91. Blanchette, J., Jaramillo, M., and Olivier, M. (2003) Immunology 108 , 513-

22.

92. Gilmore, T. D. (2006) Oncogene 25 , 6680-4.

93. Baeuerle, P. A., and Henkel, T. (1994) Annu Rev Immunol 12 , 141-79.

94. Baldwin, A. S., Jr. (1996) Annu Rev Immunol 14 , 649-83.

95. Taylor, B. S., de Vera, M. E., Ganster, R. W., Wang, Q., Shapiro, R. A.,

Morris, S. M., Jr., Billiar, T. R., and Geller, D. A. (1998) J Biol Chem 273 ,

15148-56.

96. Goldring, C. E., Reveneau, S., Algarte, M., and Jeannin, J. F. (1996)

Nucleic Acids Res 24 , 1682-7.

97. Eberhardt, W., Pluss, C., Hummel, R., and Pfeilschifter, J. (1998) J

Immunol 160 , 4961-9.

98. Jeon, Y. J., Han, S. H., Lee, Y. W., Yea, S. S., and Yang, K. H. (1998)

Biochem Mol Biol Int 45 , 435-41.

99. Yu, Z., Zhang, W., and Kone, B. C. (2002) Biochem J 367 , 97-105.

213

100. Saura, M., Zaragoza, C., Diaz-Cazorla, M., Hernandez-Perera, O., Eng, E.,

Lowenstein, C. J., Perez-Sala, D., and Lamas, S. (1998) Kidney Int 53 ,

38-49.

101. de Vera, M. E., Shapiro, R. A., Nussler, A. K., Mudgett, J. S., Simmons, R.

L., Morris, S. M., Jr., Billiar, T. R., and Geller, D. A. (1996) Proc Natl Acad

Sci U S A 93 , 1054-9.

102. Hess, J., Angel, P., and Schorpp-Kistner, M. (2004) J Cell Sci 117 , 5965-

73.

103. Chu, S. C., Marks-Konczalik, J., Wu, H. P., Banks, T. C., and Moss, J.

(1998) Biochem Biophys Res Commun 248 , 871-8.

104. Marks-Konczalik, J., Chu, S. C., and Moss, J. (1998) J Biol Chem 273 ,

22201-8.

105. Xu, W., Comhair, S. A., Zheng, S., Chu, S. C., Marks-Konczalik, J., Moss,

J., Haque, S. J., and Erzurum, S. C. (2003) Am J Physiol Lung Cell Mol

Physiol 285 , L137-48.

106. Bhat, N. R., Feinstein, D. L., Shen, Q., and Bhat, A. N. (2002) J Biol Chem

277 , 29584-92.

214

107. Du, Q., Park, K. S., Guo, Z., He, P., Nagashima, M., Shao, L., Sahai, R.,

Geller, D. A., and Hussain, S. P. (2006) Cancer Res 66 , 7024-31.

108. Boyault, S., Simonin, M. A., Bianchi, A., Compe, E., Liagre, B., Mainard,

D., Becuwe, P., Dauca, M., Netter, P., Terlain, B., and Bordji, K. (2001)

FEBS Lett 501 , 24-30.

109. Jung, F., Palmer, L. A., Zhou, N., and Johns, R. A. (2000) Circ Res 86 ,

319-25.

110. Ambs, S., Ogunfusika, M. O., Merriam, W. G., Bennett, W. P., Billiar, T. R.,

and Harris, C. C. (1998) Proc Natl Acad Sci U S A 95 , 8823-8.

111. Wagner, B. J., DeMaria, C. T., Sun, Y., Wilson, G. M., and Brewer, G.

(1998) Genomics 48 , 195-202.

112. Xu, N., Chen, C. Y., and Shyu, A. B. (2001) Mol Cell Biol 21 , 6960-71.

113. Brennan, C. M., and Steitz, J. A. (2001) Cell Mol Life Sci 58 , 266-77.

114. Blaxall, B. C., Pende, A., Wu, S. C., and Port, J. D. (2002) Mol Cell

Biochem 232 , 1-11.

215

115. Misquitta, C. M., Iyer, V. R., Werstiuk, E. S., and Grover, A. K. (2001) Mol

Cell Biochem 224 , 53-67.

116. Vodovotz, Y., Bogdan, C., Paik, J., Xie, Q. W., and Nathan, C. (1993) J

Exp Med 178 , 605-13.

117. Geller, D. A., Lowenstein, C. J., Shapiro, R. A., Nussler, A. K., Di Silvio, M.,

Wang, S. C., Nakayama, D. K., Simmons, R. L., Snyder, S. H., and Billiar,

T. R. (1993) Proc Natl Acad Sci U S A 90 , 3491-5.

118. Galea, E., Reis, D. J., and Feinstein, D. L. (1994) J Neurosci Res 37 , 406-

14.

119. Rodriguez-Pascual, F., Hausding, M., Ihrig-Biedert, I., Furneaux, H., Levy,

A. P., Forstermann, U., and Kleinert, H. (2000) J Biol Chem 275 , 26040-9.

120. Soderberg, M., Raffalli-Mathieu, F., and Lang, M. A. (2002) Mol

Pharmacol 62 , 423-31.

121. Luss, H., Li, R. K., Shapiro, R. A., Tzeng, E., McGowan, F. X., Yoneyama,

T., Hatakeyama, K., Geller, D. A., Mickle, D. A., Simmons, R. L., and

Billiar, T. R. (1997) J Mol Cell Cardiol 29 , 1153-65.

216

122. Lee, J., Ryu, H., Ferrante, R. J., Morris, S. M., Jr., and Ratan, R. R. (2003)

Proc Natl Acad Sci U S A 100 , 4843-8.

123. Petrov, T., Underwood, B. D., Braun, B., Alousi, S. S., and Rafols, J. A.

(2001) J Neurotrauma 18 , 799-812.

124. Ratovitski, E. A., Alam, M. R., Quick, R. A., McMillan, A., Bao, C.,

Kozlovsky, C., Hand, T. A., Johnson, R. C., Mains, R. E., Eipper, B. A.,

and Lowenstein, C. J. (1999) J Biol Chem 274 , 993-9.

125. Ratovitski, E. A., Bao, C., Quick, R. A., McMillan, A., Kozlovsky, C., and

Lowenstein, C. J. (1999) J Biol Chem 274 , 30250-7.

126. Blasko, E., Glaser, C. B., Devlin, J. J., Xia, W., Feldman, R. I., Polokoff, M.

A., Phillips, G. B., Whitlow, M., Auld, D. S., McMillan, K., Ghosh, S.,

Stuehr, D. J., and Parkinson, J. F. (2002) J Biol Chem 277 , 295-302.

127. Salh, B., Wagey, R., Marotta, A., Tao, J. S., and Pelech, S. (1998) J

Immunol 161 , 6947-54.

128. Vodovotz, Y., Russell, D., Xie, Q. W., Bogdan, C., and Nathan, C. (1995) J

Immunol 154 , 2914-25.

217

129. Hausel, P., Latado, H., Courjault-Gautier, F., and Felley-Bosco, E. (2006)

Oncogene 25 , 198-206.

130. Pan, J., Burgher, K. L., Szczepanik, A. M., and Ringheim, G. E. (1996)

Biochem J 314 ( Pt 3) , 889-94.

131. Musial, A., and Eissa, N. T. (2001) J Biol Chem 276 , 24268-73.

132. Kolodziejski, P. J., Musial, A., Koo, J. S., and Eissa, N. T. (2002) Proc Natl

Acad Sci U S A 99 , 12315-20.

133. Felley-Bosco, E., Bender, F. C., Courjault-Gautier, F., Bron, C., and Quest,

A. F. (2000) Proc Natl Acad Sci U S A 97 , 14334-9.

134. Walker, G., Pfeilschifter, J., and Kunz, D. (1997) J Biol Chem 272 , 16679-

87.

135. Liu, T., Huang, Y., Likhotvorik, R. I., Keshvara, L., and Hoyt, D. G. (2008)

Am J Physiol Cell Physiol 295 , C819-27.

136. Smallwood, H. S., Shi, L., and Squier, T. C. (2006) Biochemistry 45 , 9717-

26.

218

137. Walker, G., Pfeilschifter, J., Otten, U., and Kunz, D. (2001) Biochim

Biophys Acta 1568 , 216-24.

138. Crofford, L. J. (1997) J Rheumatol Suppl 49 , 15-9.

139. Harper, K. A., and Tyson-Capper, A. J. (2008) Biochem Soc Trans 36 ,

543-5.

140. Chun, K. S., and Surh, Y. J. (2004) Biochem Pharmacol 68 , 1089-100.

141. Kang, Y. J., Mbonye, U. R., DeLong, C. J., Wada, M., and Smith, W. L.

(2007) Prog Lipid Res 46 , 108-25.

142. Chen, C. C. (2006) Curr Pharm Des 12 , 3497-508.

143. Xie, W., and Herschman, H. R. (1995) J Biol Chem 270 , 27622-8.

144. Kang, Y. J., Wingerd, B. A., Arakawa, T., and Smith, W. L. (2006) J

Immunol 177 , 8111-22.

145. Tsatsanis, C., Androulidaki, A., Venihaki, M., and Margioris, A. N. (2006)

Int J Biochem Cell Biol 38 , 1654-61.

219

146. Zhu, Y., Saunders, M. A., Yeh, H., Deng, W. G., and Wu, K. K. (2002) J

Biol Chem 277 , 6923-8.

147. Araki, Y., Okamura, S., Hussain, S. P., Nagashima, M., He, P., Shiseki, M.,

Miura, K., and Harris, C. C. (2003) Cancer Res 63 , 728-34.

148. Tanabe, T., and Tohnai, N. (2002) Prostaglandins Other Lipid Mediat 68-

69 , 95-114.

149. Wheeler-Jones, C. P. (2008) Pharmacol Rep 60 , 109-18.

150. Wang, Q., Zhou, Y., Wang, X., and Evers, B. M. (2006) Oncogene 25 , 43-

50.

151. Dixon, D. A. (2004) Curr Pharm Des 10 , 635-46.

152. Dixon, D. A., Kaplan, C. D., McIntyre, T. M., Zimmerman, G. A., and

Prescott, S. M. (2000) J Biol Chem 275 , 11750-7.

153. Young, L. E., Sanduja, S., Bemis-Standoli, K., Pena, E. A., Price, R. L.,

and Dixon, D. A. (2009) Gastroenterology .

154. Lee, H. K., and Jeong, S. (2006) Nucleic Acids Res 34 , 5705-14.

220

155. Dixon, D. A., Balch, G. C., Kedersha, N., Anderson, P., Zimmerman, G. A.,

Beauchamp, R. D., and Prescott, S. M. (2003) J Exp Med 198 , 475-81.

156. Kurumbail, R. G., Kiefer, J. R., and Marnett, L. J. (2001) Curr Opin Struct

Biol 11 , 752-60.

157. Kim, S. F., Huri, D. A., and Snyder, S. H. (2005) Science 310 , 1966-70.

158. Kargman, S. L., O'Neill, G. P., Vickers, P. J., Evans, J. F., Mancini, J. A.,

and Jothy, S. (1995) Cancer Res 55 , 2556-9.

159. Shao, J., Sheng, H., Aramandla, R., Pereira, M. A., Lubet, R. A., Hawk, E.,

Grogan, L., Kirsch, I. R., Washington, M. K., Beauchamp, R. D., and

DuBois, R. N. (1999) Carcinogenesis 20 , 185-91.

160. Neuss, H., Huang, X., Hetfeld, B. K., Deva, R., Henklein, P., Nigam, S.,

Mall, J. W., Schwenk, W., and Dubiel, W. (2007) J Mol Med 85 , 961-70.

161. Mbonye, U. R., Wada, M., Rieke, C. J., Tang, H. Y., Dewitt, D. L., and

Smith, W. L. (2006) J Biol Chem 281 , 35770-8.

162. Mbonye, U. R., Yuan, C., Harris, C. E., Sidhu, R. S., Song, I., Arakawa, T.,

and Smith, W. L. (2008) J Biol Chem 283 , 8611-23.

221

163. Rockwell, P., Yuan, H., Magnusson, R., and Figueiredo-Pereira, M. E.

(2000) Arch Biochem Biophys 374 , 325-33.

164. Shearer, A. G., and Hampton, R. Y. (2004) J Biol Chem 279 , 188-96.

165. McCracken, A. A., and Brodsky, J. L. (2003) Bioessays 25 , 868-77.

166. Tsai, B., Ye, Y., and Rapoport, T. A. (2002) Nat Rev Mol Cell Biol 3, 246-

55.

167. DeMartino, G. N., and Slaughter, C. A. (1999) J Biol Chem 274 , 22123-6.

168. Song, I., Ball, T. M., and Smith, W. L. (2001) Biochem Biophys Res

Commun 289 , 869-75.

169. Mevkh, A. T., Miroshnikov, K. A., Igumnova, N. D., and Varfolomeev, S. D.

(1993) FEBS Lett 321 , 205-8.

170. Mancini, A., Jovanovic, D. V., He, Q. W., and Di Battista, J. A. (2007) J

Cell Biochem 101 , 425-41.

171. Liu, T., Schneider, R., Shah, V., Huang, Y., Likhotvorik, R. I., Keshvara, L.,

and Hoyt, D. G. (2009) Journal of Inflammation , submitted.

222

172. Lufei, C., Koh, T. H., Uchida, T., and Cao, X. (2007) Oncogene 26 , 7656-

64.

173. Miller, M. (2006) Cell Cycle 5, 2501-8.

174. Kim, M. R., Choi, H. S., Heo, T. H., Hwang, S. W., and Kang, K. W. (2008)

Biochem Biophys Res Commun 369 , 547-53.

175. Yeh, E., Cunningham, M., Arnold, H., Chasse, D., Monteith, T., Ivaldi, G.,

Hahn, W. C., Stukenberg, P. T., Shenolikar, S., Uchida, T., Counter, C. M.,

Nevins, J. R., Means, A. R., and Sears, R. (2004) Nat Cell Biol 6, 308-18.

176. van Drogen, F., Sangfelt, O., Malyukova, A., Matskova, L., Yeh, E., Means,

A. R., and Reed, S. I. (2006) Mol Cell 23 , 37-48.

177. Saez, M. E., Ramirez-Lorca, R., Moron, F. J., and Ruiz, A. (2006) Drug

Discov Today 11 , 917-23.

178. Warren, M. W., Larner, S. F., Kobeissy, F. H., Brezing, C. A., Jeung, J. A.,

Hayes, R. L., Gold, M. S., and Wang, K. K. (2007) Acta Neuropathol 114 ,

277-86.

223

179. Yan, B., Calderwood, D. A., Yaspan, B., and Ginsberg, M. H. (2001) J Biol

Chem 276 , 28164-70.

180. Kennett, S. B., Roberts, J. D., and Olden, K. (2004) J Biol Chem 279 ,

3300-7.

181. Li, J. J., and Xie, D. (2007) Biochem Biophys Res Commun 357 , 648-54.

182. Goni-Oliver, P., Lucas, J. J., Avila, J., and Hernandez, F. (2007) J Biol

Chem 282 , 22406-13.

183. Pasquet, J. M., Dachary-Prigent, J., and Nurden, A. T. (1998) Biochem J

333 ( Pt 3) , 591-9.

184. Xi, X., Flevaris, P., Stojanovic, A., Chishti, A., Phillips, D. R., Lam, S. C.,

and Du, X. (2006) J Biol Chem 281 , 29426-30.

185. Miloso, M., Mazzotti, M., Vass, W. C., and Beguinot, L. (1995) J Biol

Chem 270 , 19557-62.

186. Yang, H., Murthy, S., Sarkar, F. H., Sheng, S., Reddy, G. P., and Dou, Q.

P. (2008) J Cell Physiol 217 , 569-76.

224

187. Jariel-Encontre, I., Salvat, C., Steff, A. M., Pariat, M., Acquaviva, C.,

Furstoss, O., and Piechaczyk, M. (1997) Mol Biol Rep 24 , 51-6.

188. Pariat, M., Salvat, C., Bebien, M., Brockly, F., Altieri, E., Carillo, S., Jariel-

Encontre, I., and Piechaczyk, M. (2000) Biochem J 345 Pt 1 , 129-38.

189. Niapour, M., Yu, Y., and Berger, S. A. (2008) J Biol Chem 283 , 21371-81.

190. Small, G. W., Chou, T. Y., Dang, C. V., and Orlowski, R. Z. (2002) Arch

Biochem Biophys 400 , 151-61.

191. Lynch, D. R., and Gleichman, A. J. (2007) Neuron 53 , 317-9.

192. Benetti, R., Copetti, T., Dell'Orso, S., Melloni, E., Brancolini, C., Monte, M.,

and Schneider, C. (2005) J Biol Chem 280 , 22070-80.

193. Chen, F., Demers, L. M., Vallyathan, V., Lu, Y., Castranova, V., and Shi, X.

(2000) Am J Physiol Cell Physiol 279 , C709-16.

194. Gonen, H., Shkedy, D., Barnoy, S., Kosower, N. S., and Ciechanover, A.

(1997) FEBS Lett 406 , 17-22.

195. Gil-Parrado, S., Fernandez-Montalvan, A., Assfalg-Machleidt, I., Popp, O.,

Bestvater, F., Holloschi, A., Knoch, T. A., Auerswald, E. A., Welsh, K.,

225

Reed, J. C., Fritz, H., Fuentes-Prior, P., Spiess, E., Salvesen, G. S., and

Machleidt, W. (2002) J Biol Chem 277 , 27217-26.

196. Abe, K., and Takeichi, M. (2007) Neuron 53 , 387-97.

197. Tompa, P., Buzder-Lantos, P., Tantos, A., Farkas, A., Szilagyi, A.,

Banoczi, Z., Hudecz, F., and Friedrich, P. (2004) J Biol Chem 279 , 20775-

85.

198. Basu, A., and Haldar, S. (2002) Int J Oncol 21 , 597-601.

199. Tompa, P., Emori, Y., Sorimachi, H., Suzuki, K., and Friedrich, P. (2001)

Biochem Biophys Res Commun 280 , 1333-9.

200. Hanna, R. A., Campbell, R. L., and Davies, P. L. (2008) Nature 456 , 409-

12.

201. Vissing, K., Overgaard, K., Nedergaard, A., Fredsted, A., and Schjerling, P.

(2008) Eur J Appl Physiol 103 , 323-32.

202. Raveendran, N. N., Silver, K., Freeman, L. C., Narvaez, D., Weng, K.,

Ganta, S., and Lillich, J. D. (2008) J Pharmacol Exp Ther 325 , 389-99.

226

203. Melloni, E., Michetti, M., Salamino, F., Minafra, R., and Pontremoli, S.

(1996) Biochem Biophys Res Commun 229 , 193-7.

204. Glading, A., Bodnar, R. J., Reynolds, I. J., Shiraha, H., Satish, L., Potter,

D. A., Blair, H. C., and Wells, A. (2004) Mol Cell Biol 24 , 2499-512.

205. Shiraha, H., Glading, A., Chou, J., Jia, Z., and Wells, A. (2002) Mol Cell

Biol 22 , 2716-27.

206. Goll, D. E., Thompson, V. F., Li, H., Wei, W., and Cong, J. (2003) Physiol

Rev 83 , 731-801.

207. Wendt, A., Thompson, V. F., and Goll, D. E. (2004) Biol Chem 385 , 465-

72.

208. Salamino, F., Averna, M., Tedesco, I., De Tullio, R., Melloni, E., and

Pontremoli, S. (1997) FEBS Lett 412 , 433-8.

209. Averna, M., De Tullio, R., Salamino, F., Melloni, E., and Pontremoli, S.

(1999) FEBS Lett 450 , 13-6.

210. Hata, A., Ohno, S., and Suzuki, K. (1992) FEBS Lett 304 , 241-4.

211. Jacquemond, V. (1997) Biophys J 73 , 920-8.

227

212. Maravall, M., Mainen, Z. F., Sabatini, B. L., and Svoboda, K. (2000)

Biophys J 78 , 2655-67.

213. Cong, J., Goll, D. E., Peterson, A. M., and Kapprell, H. P. (1989) J Biol

Chem 264 , 10096-103.

214. Saido, T. C., Shibata, M., Takenawa, T., Murofushi, H., and Suzuki, K.

(1992) J Biol Chem 267 , 24585-90.

215. Melloni, E., Averna, M., Salamino, F., Sparatore, B., Minafra, R., and

Pontremoli, S. (2000) J Biol Chem 275 , 82-6.

216. Melloni, E., Michetti, M., Salamino, F., and Pontremoli, S. (1998) J Biol

Chem 273 , 12827-31.

217. Satish, L., Blair, H. C., Glading, A., and Wells, A. (2005) Mol Cell Biol 25 ,

1922-41.

218. Glading, A., Lauffenburger, D. A., and Wells, A. (2002) Trends Cell Biol 12 ,

46-54.

219. Glading, A., Uberall, F., Keyse, S. M., Lauffenburger, D. A., and Wells, A.

(2001) J Biol Chem 276 , 23341-8.

228

220. Glading, A., Chang, P., Lauffenburger, D. A., and Wells, A. (2000) J Biol

Chem 275 , 2390-8.

221. Xu, L., and Deng, X. (2006) J Biol Chem 281 , 4457-66.

222. Dayton, W. R., Goll, D. E., Zeece, M. G., Robson, R. M., and Reville, W. J.

(1976) Biochemistry 15 , 2150-8.

223. Melloni, E., Averna, M., Stifanese, R., De Tullio, R., Defranchi, E.,

Salamino, F., and Pontremoli, S. (2006) J Biol Chem 281 , 24945-54.

224. Moldoveanu, T., Gehring, K., and Green, D. R. (2008) Nature 456 , 404-8.

225. Mellgren, R. L. (2008) Nature 456 , 337-8.

226. Higuchi, M., Iwata, N., and Saido, T. C. (2005) Biochim Biophys Acta 1751 ,

60-7.

227. Sribnick, E. A., Matzelle, D. D., Banik, N. L., and Ray, S. K. (2007)

Neurochem Res 32 , 2210-6.

228. Momeni, H. R., Azadi, S., and Kanje, M. (2007) Funct Neurol 22 , 105-10.

229. Momeni, H. R., and Kanje, M. (2006) Neuroreport 17 , 761-5.

229

230. Das, A., Sribnick, E. A., Wingrave, J. M., Del Re, A. M., Woodward, J. J.,

Appel, S. H., Banik, N. L., and Ray, S. K. (2005) J Neurosci Res 81 , 551-

62.

231. Ray, S. K., Matzelle, D. D., Sribnick, E. A., Guyton, M. K., Wingrave, J. M.,

and Banik, N. L. (2003) J Chem Neuroanat 26 , 119-24.

232. Ray, S. K., and Banik, N. L. (2003) Curr Drug Targets CNS Neurol Disord

2, 173-89.

233. Ray, S. K., Matzelle, D. D., Wilford, G. G., Hogan, E. L., and Banik, N. L.

(2001) Brain Res 916 , 115-26.

234. Ray, S. K., Hogan, E. L., and Banik, N. L. (2003) Brain Res Brain Res Rev

42 , 169-85.

235. Ray, S. K., Matzelle, D. D., Wilford, G. G., Hogan, E. L., and Banik, N. L.

(2000) Neurochem Res 25 , 1191-8.

236. Wulf, G., Finn, G., Suizu, F., and Lu, K. P. (2005) Nat Cell Biol 7, 435-41.

230

237. Yaffe, M. B., Schutkowski, M., Shen, M., Zhou, X. Z., Stukenberg, P. T.,

Rahfeld, J. U., Xu, J., Kuang, J., Kirschner, M. W., Fischer, G., Cantley, L.

C., and Lu, K. P. (1997) Science 278 , 1957-60.

238. Ilsley, J. L., Sudol, M., and Winder, S. J. (2002) Cell Signal 14 , 183-9.

239. Lu, K. P., Liou, Y. C., and Zhou, X. Z. (2002) Trends Cell Biol 12 , 164-72.

240. Yeh, E. S., and Means, A. R. (2007) Nat Rev Cancer 7, 381-8.

241. Mei, J. M., Hord, N. G., Winterstein, D. F., Donald, S. P., and Phang, J. M.

(2000) Faseb J 14 , 1188-201.

242. Morikawa, A., Koide, N., Kato, Y., Sugiyama, T., Chakravortty, D., Yoshida,

T., and Yokochi, T. (2000) Infect Immun 68 , 6209-14.

243. Huang, H., McIntosh, J., and Hoyt, D. G. (2003) In Vitro Cell Dev Biol

Anim 39 , 43-50.

244. Cuzzocrea, S., Mazzon, E., Di Paola, R., Esposito, E., Macarthur, H.,

Matuschak, G. M., and Salvemini, D. (2006) J Pharmacol Exp Ther 319 ,

73-81.

245. Gross, S. S., and Wolin, M. S. (1995) Annu Rev Physiol 57 , 737-69.

231

246. Hemmrich, K., Kroncke, K. D., Suschek, C. V., and Kolb-Bachofen, V.

(2005) Nitric Oxide 12 , 183-99.

247. Kroncke, K. D., Fehsel, K., and Kolb-Bachofen, V. (1997) Nitric Oxide 1,

107-20.

248. Shelton, J. L., Wang, L., Cepinskas, G., Sandig, M., Scott, J. A., North, M.

L., Inculet, R., and Mehta, S. (2007) Microvasc Res 74 , 23-31.

249. Huang, Y., Magdaleno, S., Hopkins, R., Slaughter, C., Curran, T., and

Keshvara, L. (2004) Biochem Biophys Res Commun 318 , 204-12.

250. Janas, J., Skowronski, J., and Van Aelst, L. (2006) Methods Enzymol 406 ,

593-605.

251. Moffat, J., Grueneberg, D. A., Yang, X., Kim, S. Y., Kloepfer, A. M., Hinkle,

G., Piqani, B., Eisenhaure, T. M., Luo, B., Grenier, J. K., Carpenter, A. E.,

Foo, S. Y., Stewart, S. A., Stockwell, B. R., Hacohen, N., Hahn, W. C.,

Lander, E. S., Sabatini, D. M., and Root, D. E. (2006) Cell 124 , 1283-98.

252. Stewart, S. A., Dykxhoorn, D. M., Palliser, D., Mizuno, H., Yu, E. Y., An, D.

S., Sabatini, D. M., Chen, I. S., Hahn, W. C., Sharp, P. A., Weinberg, R. A.,

and Novina, C. D. (2003) Rna 9, 493-501.

232

253. Lundberg, A. S., Randell, S. H., Stewart, S. A., Elenbaas, B., Hartwell, K.

A., Brooks, M. W., Fleming, M. D., Olsen, J. C., Miller, S. W., Weinberg, R.

A., and Hahn, W. C. (2002) Oncogene 21 , 4577-86.

254. Rose, J. L., Reeves, K. C., Likhotvorik, R. I., and Hoyt, D. G. (2007) J

Pharmacol Exp Ther 321 , 318-26.

255. Kok, R. J., Schraa, A. J., Bos, E. J., Moorlag, H. E., Asgeirsdottir, S. A.,

Everts, M., Meijer, D. K., and Molema, G. (2002) Bioconjug Chem 13 , 128-

35.

256. Choi, H. S., Kim, J. W., Cha, Y. N., and Kim, C. (2006) J Immunoassay

Immunochem 27 , 31-44.

257. Huang, H., Liu, T., Rose, J. L., Stevens, R. L., and Hoyt, D. G. (2007) J

Inflamm (Lond) 4, 22.

258. Shen, M., Stukenberg, P. T., Kirschner, M. W., and Lu, K. P. (1998)

Genes Dev 12 , 706-20.

259. Bradford, M. M. (1976) Anal Biochem 72 , 248-54.

260. Laemmli, U. K. (1970) Nature 227 , 680-5.

233

261. Meng, L., Mohan, R., Kwok, B. H., Elofsson, M., Sin, N., and Crews, C. M.

(1999) Proc Natl Acad Sci U S A 96 , 10403-8.

262. Javesghani, D., Hussain, S. N., Scheidel, J., Quinn, M. T., and Magder, S.

A. (2003) Shock 19 , 486-93.

263. Muzaffar, S., Jeremy, J. Y., Angelini, G. D., Stuart-Smith, K., and Shukla,

N. (2003) Thorax 58 , 598-604.

264. Pacher, P., Beckman, J. S., and Liaudet, L. (2007) Physiol Rev 87 , 315-

424.

265. Xia, Y., and Zweier, J. L. (1997) Proc Natl Acad Sci U S A 94 , 6954-8.

266. Yoshida, M., and Xia, Y. (2003) J Biol Chem 278 , 36953-8.

267. Esnault, S., Braun, R. K., Shen, Z. J., Xiang, Z., Heninger, E., Love, R. B.,

Sandor, M., and Malter, J. S. (2007) PLoS ONE 2, e226.

268. Esnault, S., Braun, R. K., Shen, Z. J., Xiang, Z., Heninger, E., Love, R. B.,

Sandor, M., and Malter, J. S. (2007) PLoS ONE 2, e226.

269. Tan, X., Essengue, S., Talreja, J., Reese, J., Stechschulte, D. J., and

Dileepan, K. N. (2007) J Immunol 179 , 7899-906.

234

270. Taubert, A., and Hermosilla, C. (2008) Vet Res Commun 32 , 35-47.

271. Ruetten, H., and Thiemermann, C. (1997) Br J Pharmacol 121 , 695-704.

272. Swierkosz, T. A., Mitchell, J. A., Warner, T. D., Botting, R. M., and Vane, J.

R. (1995) Br J Pharmacol 114 , 1335-42.

273. Yang, T., Sun, D., Huang, Y. G., Smart, A., Briggs, J. P., and Schnermann,

J. B. (1999) Am J Physiol 277 , F10-6.

274. Zhang, S., Thomas, K., Blanco, J. C., Salkowski, C. A., and Vogel, S. N.

(2002) J Endotoxin Res 8, 379-88.

275. El-Achkar, T. M., Plotkin, Z., Marcic, B., and Dagher, P. C. (2007) Am J

Physiol Renal Physiol 293 , F1187-96.

276. Flavahan, N. A. (2007) Trends Pharmacol Sci 28 , 106-10.

277. Bishop-Bailey, D., Mitchell, J. A., and Warner, T. D. (2006) Arterioscler

Thromb Vasc Biol 26 , 956-8.

278. Ejima, K., Layne, M. D., Carvajal, I. M., Kritek, P. A., Baron, R. M., Chen,

Y. H., Vom Saal, J., Levy, B. D., Yet, S. F., and Perrella, M. A. (2003)

Faseb J 17 , 1325-7.

235

279. Chen, G., Wood, E. G., Wang, S. H., and Warner, T. D. (1999) Life Sci 64 ,

1231-42.

280. Huang, H., McIntosh, J., and Hoyt, D. (2003) In Vitro Cell Dev Biol Anim

39 , 43-50.

281. Mehdi, S. (1991) Trends Biochem Sci 16 , 150-3.

282. Smith, C. J., Zhang, Y., Koboldt, C. M., Muhammad, J., Zweifel, B. S.,

Shaffer, A., Talley, J. J., Masferrer, J. L., Seibert, K., and Isakson, P. C.

(1998) Proc Natl Acad Sci U S A 95 , 13313-8.

283. Futaki, N., Takahashi, S., Yokoyama, M., Arai, I., Higuchi, S., and Otomo,

S. (1994) Prostaglandins 47 , 55-9.

284. Huang, H., Liu, T., Rose, J. L., Stevens, R. L., and Hoyt, D. G. (2007) J

Inflamm (Lond) 4, 22.

285. Yih, L. H., and Lee, T. C. (1994) Biochem Biophys Res Commun 202 ,

1015-22.

286. Werle, B., Staib, A., Julke, B., Ebert, W., Zladoidsky, P., Sekirnik, A., Kos,

J., and Spiess, E. (1999) Biol Chem 380 , 1109-16.

236

287. Snedecor, G., and Cochran, W. (1980) Statistical Methods , 7th ed., Iowa

State University Press, Ames.

288. Cui, Z., Han, Z., Li, Z., Hu, H., Patel, J. M., Antony, V., Block, E. R., and

Su, Y. (2005) Am J Respir Cell Mol Biol 33 , 513-20.

289. Prescott, S. M., and White, R. L. (1996) Cell 87 , 783-6.

290. Howe, L. R., Subbaramaiah, K., Chung, W. J., Dannenberg, A. J., and

Brown, A. M. (1999) Cancer Res 59 , 1572-7.

291. Mei, J. M., Hord, N. G., Winterstein, D. F., Donald, S. P., and Phang, J. M.

(1999) Carcinogenesis 20 , 737-40.

292. Kim, S. J., Im, D. S., Kim, S. H., Ryu, J. H., Hwang, S. G., Seong, J. K.,

Chun, C. H., and Chun, J. S. (2002) Biochem Biophys Res Commun 296 ,

221-6.

293. Finn, G., and Lu, K. P. (2008) Curr Cancer Drug Targets 8, 223-9.

294. Li, Y., Arnold, J. M., Pampillo, M., Babwah, A. V., and Peng, T. (2009)

Free Radic Biol Med 46 , 51-61.

237

295. Mehendale, H. M., and Limaye, P. B. (2005) Trends Pharmacol Sci 26 ,

232-6.

296. Carragher, N. O. (2006) Curr Pharm Des 12 , 615-38.

297. Snedecor, G. a. C. W. (1980) Statistical Methods , 7th ed., Iowa State

University Press, Ames.

298. Sorimachi, Y., Harada, K., Saido, T. C., Ono, T., Kawashima, S., and

Yoshida, K. (1997) J Biochem (Tokyo) 122 , 743-8.

299. Blomgren, K., Hallin, U., Andersson, A. L., Puka-Sundvall, M., Bahr, B. A.,

McRae, A., Saido, T. C., Kawashima, S., and Hagberg, H. (1999) J Biol

Chem 274 , 14046-52.

300. Kato, M., Nonaka, T., Maki, M., Kikuchi, H., and Imajoh-Ohmi, S. (2000) J

Biochem (Tokyo) 127 , 297-305.

301. Barnoy, S., and Kosower, N. S. (2003) FEBS Lett 546 , 213-7.

302. Wang, K. K., Posmantur, R., Nadimpalli, R., Nath, R., Mohan, P., Nixon, R.

A., Talanian, R. V., Keegan, M., Herzog, L., and Allen, H. (1998) Arch

Biochem Biophys 356 , 187-96.

238

303. Barnoy, S., and Kosower, N. S. (2007) Int J Biochem Cell Biol 39 , 253-61.

304. Wingrave, J. M., Sribnick, E. A., Wilford, G. G., Matzelle, D. D., Mou, J. A.,

Ray, S. K., Hogan, E. L., and Banik, N. L. (2004) J Neurotrauma 21 , 1240-

54.

305. Wei, W., Fareed, M. U., Evenson, A., Menconi, M. J., Yang, H., Petkova,

V., and Hasselgren, P. O. (2005) Am J Physiol Regul Integr Comp Physiol

288 , R580-90.

306. Adachi, Y., Ishida-Takahashi, A., Takahashi, C., Takano, E., Murachi, T.,

and Hatanaka, M. (1991) J Biol Chem 266 , 3968-72.

307. Blomgren, K., Nilsson, E., and Karlsson, J. O. (1989) Comp Biochem

Physiol B 93 , 403-7.

308. Su, Y., Cao, W., Han, Z., and Block, E. R. (2004) Am J Physiol Lung Cell

Mol Physiol 287 , L794-800.

309. Qiu, K., Su, Y., and Block, E. R. (2006) Mol Cell Biochem 292 , 69-78.

310. Letavernier, E., Perez, J., Bellocq, A., Mesnard, L., de Castro Keller, A.,

Haymann, J. P., and Baud, L. (2008) Circ Res 102 , 720-8.

239

311. Li, X., Li, Y., Shan, L., Shen, E., Chen, R., and Peng, T. (2009)

Cardiovasc Res .

312. Khalil, P. N., Neuhof, C., Huss, R., Pollhammer, M., Khalil, M. N., Neuhof,

H., Fritz, H., and Siebeck, M. (2005) Eur J Pharmacol 528 , 124-31.

313. Galvez, A. S., Diwan, A., Odley, A. M., Hahn, H. S., Osinska, H.,

Melendez, J. G., Robbins, J., Lynch, R. A., Marreez, Y., and Dorn, G. W.,

2nd. (2007) Circ Res 100 , 1071-8.

314. Shen, Y., Schlessinger, K., Zhu, X., Meffre, E., Quimby, F., Levy, D. E.,

and Darnell, J. E., Jr. (2004) Mol Cell Biol 24 , 407-19.

315. Klover, P., and Hennighausen, L. (2007) Endocrinology 148 , 1489-97.

316. Stratikopoulos, E., Szabolcs, M., Dragatsis, I., Klinakis, A., and Efstratiadis,

A. (2008) Proc Natl Acad Sci U S A 105 , 19378-83.

317. Wang, J., Zhou, J., Powell-Braxton, L., and Bondy, C. (1999)

Endocrinology 140 , 3391-4.

318. Breese, C. R., Ingram, R. L., and Sonntag, W. E. (1991) J Gerontol 46 ,

B180-7.

240

319. Fontana, L., Weiss, E. P., Villareal, D. T., Klein, S., and Holloszy, J. O.

(2009) Aging Cell .

320. Pollak, M. (2009) Aging Cell 8, 214; author reply 215.

241